Bacillus: Cellular and Molecular Biology [3 ed.] 9781910190586, 9781910190579

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Bacillus Cellular and Molecular Biology (Third edition)

Edited by

Peter L. Graumann

Caister Academic Press

Bacillus

Cellular and Molecular Biology (Third edition) https://doi.org/10.21775/9781910190579

Edited by Peter L. Graumann Faculty of Chemistry and SYNMIKRO (Centre for Synthetic Microbiology) University of Marburg Marburg Germany

Caister Academic Press

Copyright © 2017 Caister Academic Press Norfolk, UK www.caister.com British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-1-910190-57-9 (paperback) ISBN: 978-1-910190-58-6 (ebook) Description or mention of instrumentation, software, or other products in this book does not imply endorsement by the author or publisher. The author and publisher do not assume responsibility for the validity of any products or procedures mentioned or described in this book or for the consequences of their use. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior permission of the publisher. No claim to original U.S. Government works. Cover design adapted from images provided by Felix Dempwolff (Indiana University, Bloomington, IN, USA), images of different membrane proteins from Bacillus subtilis taken with STED microscopy. Ebooks Ebooks supplied to individuals are single-user only and must not be reproduced, copied, stored in a retrieval system, or distributed by any means, electronic, mechanical, photocopying, email, internet or otherwise. Ebooks supplied to academic libraries, corporations, government organizations, public libraries, and school libraries are subject to the terms and conditions specified by the supplier.

Contents

Prefacev 1

Replication of the Bacillus subtilis Chromosome1

2

Dynamics of DNA Double-strand Break Repair in Bacillus subtilis35

3

Chromosome Arrangement and Segregation

67

4

Cell Division

89

5

The Organization of Transcription and Translation

6

RNA-mediated Regulation in Bacillus subtilis155

7

General and Regulatory Proteolysis in Bacillus subtilis191

8

The Actin-like MreB ‘Cytoskeleton’

223

9

Ins and Outs of the Bacillus subtilis Membrane Proteome

263

Heath Murray, Tomas T. Richardson, Patrice Polard, Philippe Noirot and Marie-Françoise Noirot-Gros Begoña Carrasco, Paula P. Cárdenas, Ester Serrano, Rubén Torres, Elena M. Seco, Silvia Ayora and Juan C. Alonso Peter L. Graumann

Frederico Gueiros-Filho Peter Lewis and Xiao Yang

127

Wade C. Winkler Kürşad Turgay

Rut Carballido-López

Jan Maarten van Dijl, Annette Dreisbach, Marcin J. Skwark, Mark J.J.B. Sibbald, Harold Tjalsma, Jessica C. Zweers and Girbe Buist

10

The Cell Wall of Bacillus subtilis295

11

Genomics and Cellular Biology of Endospore Formation

12

Multicellularity and Social Behaviour in Bacillus subtilis367

Danae Morales Angeles and Dirk-Jan Scheffers Patrick Eichenberger

José Eduardo González-Pastor

333

iv  | Contents

13

Competence and Transformation

14

Swimming, Swarming and Sliding Motility in Bacillus subtilis415

15

Nucleotide Second Messengers: (p)ppGpp and Cyclic Dinucleotides

Berenike Maier

395

Anna C. Hughes and Daniel B. Kearns

Danny K. Fung, Brent W. Anderson, Jessica L. Tse and Jue D. Wang

439

Index467

Preface

Bacillus subtilis is one of the best understood prokaryotes in terms of molecular biology and cell biology. Its superb genetic amenability and relatively large size have provided powerful tools to investigate a bacterium in all possible aspects. Recent improvements in fluorescence microscopy techniques have provided novel and amazing insight into the dynamic structure of a single-cell organism. Research on B. subtilis has been at the forefront of bacterial molecular biology and cytology, and the organism is a model for differentiation, gene/ protein regulation and cell cycle events in bacteria. The aim of this book is to present an overview of the most recent exciting new research fields, and to provide a picture of the major cytological aspects of a bacterium, many of which are highly relevant for a wide variety of bacteria. Bacillus subtilis is a ubiquitous soil bacterium that can be easily isolated from soil, using starch as an energy source and relatively high salt concentration. Ideally, the soil sample is heated up to 100°C for 30 min, allowing only for enduring spores to be cultured from the sample. B. subtilis is unique in that it can choose between at least three different genetic programmes when nutrients or other resources become scarce and/or cell density reaches a critical threshold. To survive or adapt to adverse condition, cells can either enter stationary phase, which is characterized by the formation of single motile cells (exponentially growing cells contain a mixture of mostly nonmotile chains of cells and a few motile single cells), can differentiate into enduring and metabolically inactive spores, or, thirdly, can become competent and take up DNA from the environment for acquisition of new genetic material. In all three cases,

strikingly different genetic programs are turned on that guide the cell through the differentiation processes. In addition to this, B. subtilis shows social behaviour, in that the cells communicate with each other, and form multicellular structures in the form of swarming cells and biofilms. Two component systems, cascades of different sigma factors, regulatory RNAs and specific proteolysis of target proteins form an intricate regulatory network, which is beginning to be unravelled, not only in terms of specific modules, but also in terms of whole complex processes that are connected with each other. Specific mono- and di-nucleotides have gained a lot of interest, as they regulate key steps in bacterial growth and physiology. Most strikingly, it has become clear that many proteins have specific subcellular addresses in bacterial cells. These findings have established the field of ‘bacterial cell biology’, and B. subtilis has been a forerunner in this field. Many vital processes are disturbed if proteins lose their specific localization, but the fundamental question of how proteins are targeted and specifically located in a call lacking intracellular compartments is still unclear for most cases. Therefore, it has become important to also study proteins in terms of their localization within the cell, in addition to analysing their biochemistry and regulation. This book is intended to show that we are beginning to understand the way a bacterial cell functions as a whole entity and in 3D, i.e. how it is spatially organized, and even how bacteria talk to each other, or give their life for the sake of the whole community. In this book, we will take and inside out approach to look at Bacillus, starting with duplication of the chromosome, cell cycle and transcriptional regulation, following its MreB cytoskeleton underneath

vi  | Preface

the cell membrane, through the membrane, to the cell wall. Finally, we will consider the multiarchitectural processes of biofilm formation and sporulation that embrace many of cytological and genetical aspects throughout the cell. New added chapters of the third edition cover the important aspects of motility (Chapter 14) and regulation through nucleotides (Chapter 15). As will become apparent to the reader, many chapters overlap in a variety of aspects, which is due to the fact that most processes addressed in the book are interconnected with each other. For example, the specific localization of the replication machinery (Chapter 1) is an important aspect in DNA repair (Chapter 2) and in ordered chromosome segregation (Chapter 3), and also touches aspects of cell division (Chapter 4). Amazingly, the actin-like MreB cytoskeleton (covered in depth in Chapter 8) is essential for viability, for the ordered insertion of cell wall material (Chapter 10). The structure of short and dynamic filaments appears to connect and coordinate cell cycle events and rod-shaped cell growth, showing that the cytoskeleton was actually a functional prokaryotic invention. Many processes thought to occur throughout the cell or all over the membrane (Chapter 9) have been found to be spatially confined to discrete regions, which has shed light onto processes such as cell division, replication, cell growth and sporulation. Even though transcription and translation are coupled in prokaryotes, these processes occur at defined places within the cells (Chapter 5), apparently facilitating ordered chromosome segregation and efficient synthesis of highly expressed proteins and of stable RNA. Regulation of transcription through RNA molecules (Chapter 6) and regulation of protein activity through proteolysis (Chapter 7) have only recently been recognized as major factors affecting bacterial physiology, and are intertwined with the organization of transcription (Chapter 5), sporulation (Chapter 11) and cell division (Chapter 4). Like many bacteria. B. subtilis cells form biofilms, which contain several distinct subpopulations sharing different kinds of labour, and which act rather as a multicellular organism. This new concept in bacteriology is described in Chapter 12. In the second edition, several important recent findings in the rapidly moving fields of research have been included, and the book has had the addition of Chapter 13 dealing with the

ability of B. subtilis cells to take up DNA from the environment and incorporate it into the chromosome, when sufficient homology exists. This developmental state is called ‘competence’ and is described in detail. Importantly, the new chapter also explains the molecular basis and mathematics of the phenomenon called bistability, in which two interchangeable B. subtilis subpopulations exist in parallel that have distinct physiological states and genetic programmes. This ability allows bacterial populations to do ‘bet hedging’ and to be able to respond to environmental conditions that may (or may not) change in the near future. The third edition has now captured important new developments from the recent 4 years, and has won two important aspects in the life of a bacterium, motility (Chapter 14) and the regulation of cellular signalling pathways via small nucleotides, of which the important functions played by cyclic di-AMP and cyclic di-GMP have only recently been recognized, and are only slowly being understood at a molecular level. Small nucleotides also affect the lifestyle decision of bacteria whether to become motile or stay sessile, and also motility is under bistable control in B. subtilis, tying together several chapters of this new edition. Clearly, different fields in bacterial cell biology and molecular biology are growing together, providing a more and more integral view of the bacterial cell. My thank goes out to Gert Bange, who has helped me in editing several chapters of this third edition and to all authors, who have done a great job updating the chapters. Useful links Several useful sites on the internet exist that provide tools to study B. subtilis in more depth. Foremost, the Bacillus sequencing consortium has set up a site in which the whole genome of B. subtilis is accessible in a superb way. Subtiwiki is a new site in which all genome data and gene expression analyses can be obtained. Also, genotypes of many strains with gene deletions are accessible via two websites. Finally, most authors of this book have websites for the interested reader to find further information on the various research areas covered in this book.

Preface |  vii

B. subtilis gene expression database http://subtiwiki.uni-goettingen.de/wiki/index.php/ Main_Page B. subtilis genome genolist.pasteur.fr/SubtiList/ B. subtilis stock centre www.bgsc.org/

W. Winkler, regulatory RNAs http://www.wadewinkler.com P. Lewis, organization of transcription https://www.newcastle.edu.au/profile/peter-lewis K. Turgay, proteolysis https://www.ifmb.uni-hannover.de/turgay.html

List of essential genes www.pnas.org/cgi/content/full/100/8/4678

R. Carballido-Lopez, cytoskeleton https://www.micalis.fr/Poles-et-Equipes/Pole-Biosys/ ProCeD-Carballido-Lopez

H. Murray, replication http://www.ncl.ac.uk/camb/staff/profile/heathmurray. html#background

J.M. van Dijl, membrane proteins w w w.r ug.nl/umc g/fac u lte it/d i sc ipl inegroepen/ medischemicrobiologie/index

J.C. Alonso, DNA repair http://www.cnb.csic.es/index.php/en/research/researchdepartments/microbial-biotechnology/genetic-stability

D.-J. Scheffers, cell wall http://www.rug.nl/staff/d.j.scheffers/

P.L. Graumann, DNA segregation htt p://s y nmi kro.com/de/for schung/zel lulaereorganisation/peter-graumann.html F. Gueiros Filho, cell division http://www2.iq.usp.br/docente/?id=fgueiros

P. Eichenberger, sporulation http://biology.as.nyu.edu/object/PatrickEichenberger E. Gonzales Pastor, biofilm formation  https://cab.inta-csic. es/en/investigadores/343/jose-eduardo-gonzalez-pastor

Current Books of Interest The CRISPR/Cas System: Emerging Technology and Application2017 Brewing Microbiology: Current Research, Omics and Microbial Ecology2017 Metagenomics: Current Advances and Emerging Concepts2017 Bacillus: Cellular and Molecular Biology (Third Edition)2017 Cyanobacteria: Omics and Manipulation2017 Foot-and-Mouth Disease Virus: Current Research and Emerging Trends2017 Brain-eating Amoebae: Biology and Pathogenesis of Naegleria fowleri2016 Staphylococcus: Genetics and Physiology2016 Chloroplasts: Current Research and Future Trends2016 Microbial Biodegradation: From Omics to Function and Application2016 Influenza: Current Research2016 MALDI-TOF Mass Spectrometry in Microbiology2016 Aspergillus and Penicillium in the Post-genomic Era2016 The Bacteriocins: Current Knowledge and Future Prospects2016 Omics in Plant Disease Resistance2016 Acidophiles: Life in Extremely Acidic Environments2016 Climate Change and Microbial Ecology: Current Research and Future Trends2016 Biofilms in Bioremediation: Current Research and Emerging Technologies2016 Microalgae: Current Research and Applications2016 Gas Plasma Sterilization in Microbiology: Theory, Applications, Pitfalls and New Perspectives2016 Virus Evolution: Current Research and Future Directions2016 Arboviruses: Molecular Biology, Evolution and Control2016 Shigella: Molecular and Cellular Biology2016 Aquatic Biofilms: Ecology, Water Quality and Wastewater Treatment2016 Alphaviruses: Current Biology2016 Thermophilic Microorganisms2015 Flow Cytometry in Microbiology: Technology and Applications2015 Probiotics and Prebiotics: Current Research and Future Trends2015 Epigenetics: Current Research and Emerging Trends2015 Corynebacterium glutamicum: From Systems Biology to Biotechnological Applications2015 Advanced Vaccine Research Methods for the Decade of Vaccines2015 Antifungals: From Genomics to Resistance and the Development of Novel Agents2015 Bacteria-Plant Interactions: Advanced Research and Future Trends2015 Aeromonas2015 Antibiotics: Current Innovations and Future Trends2015 Full details at www.caister.com

Replication of the Bacillus subtilis Chromosome Heath Murray1*, Tomas T. Richardson1, Patrice Polard2, Philippe Noirot3 and Marie-Françoise Noirot-Gros3

1

1

Centre for Bacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle University, Newcastle Upon Tyne, UK. 2 Centre National de la Recherche Scientifique, Laboratoire de Microbiologie et Génétique Moléculaires, Université de Toulouse, Toulouse, France. 3 Argonne National Laboratory, Argonne, IL, USA. Correspondence: [email protected] https://doi.org/10.21775/9781910190579-01

Abstract Eubacteria have evolved multicomponent protein machines, termed replisomes, which duplicate their chromosomes rapidly and accurately. Extensive studies in the model bacteria Escherichia coli and Bacillus subtilis have revealed that in addition to the core replication machinery, other proteins are necessary to form a functional replication fork. Specific subsets of proteins mediate (a) assembly of the replisome at the chromosomal origin of replication [initiation]; (b) progression of the replication forks along the chromosome [elongation] and their maintenance by providing solutions for replication restart, which are adapted to overcome possible ‘roadblocks’ encountered on the DNA template; and (c) physiological arrest of replication when chromosome duplication is completed [termination]. This review summarizes recent knowledge about chromosomal replication in Bacillus subtilis and related Gram-positive bacteria. It is focused on the events governing the assembly and fate of the replication fork, describes protein networks connected with the replisome, and emphasizes several novel aspects of DNA replication in this group of bacteria. Introduction Bacillus subtilis duplicates its circular chromosome by initiating DNA synthesis at a single locus, the

replication origin (oriC, 0°; see Fig. 3.1). Replication proceeds bidirectionally with the two replication forks progressing in the clockwise and counter-clockwise directions along the chromosome halves (Lemon et al., 2002). Chromosome replication is completed when the forks reach the terminus region (180°; see Fig. 3.1), which is positioned opposite the origin on the chromosome map and contains several short DNA sequences (ter sites) that promote replication arrest (Duggin and Wake, 2002). Specific proteins mediate all of the steps in the DNA replication pathway. The comparison between the sets of proteins involved in chromosomal DNA replication in B. subtilis and in E. coli reveals both similarities and differences (Table 1.1). Although the basic components promoting initiation, elongation, and termination of replication are well conserved, some important differences can be found (e.g. essential proteins in one bacterium missing from the other). These differences underline the diversity in the mechanisms and strategies that various bacterial species have adopted to carry out the duplication of their genomes. This review summarizes recent knowledge about chromosomal replication in B. subtilis and related Gram-positive bacteria. Detailed understanding of the events governing the fate of a replication fork in E. coli will be briefly described and used as

2  | Murray et al.

Table 1.1 Comparison of the B. subtilis and E. coli proteins involved in different aspects of chromosomal DNA replication B. subtilis

E. coli

Protein functiona

References

DnaA

DnaA

Master initiator of DNA replication, oriC binding at DnaA-boxes and DnaA-trios, local unwinding of the DNA duplex, and recruitment of DnaD

Kornberg and Baker, 1992b; Krause et al., 1997; Moriya et al., 1990; Richardson et al., 2016

DnaB

–

Initiation at oriC, component of the replication restart primosome, co-loader of the DnaC helicase

Bruand et al., 1995; Bruand et al., 2001; Velten et al., 2003

DnaC

DnaB

Replicative helicase

Velten et al., 2003

DnaD

–

Initiation at oriC, recruitment of DnaB, and component of the replication restart primosome

Bruand et al., 2001; Bruand et al., 2005; Rokop et al., 2004

DnaE

DnaE b

Class C DNA polymerase essential for replication fork progression, lagging strand polymerase, involved in error-prone synthesis and lesion bypass

Bruck et al., 2003; Dervyn et al., 2001; Inoue et al., 2001; Le Chatelier et al., 2004; Sanders et al., 2010

DnaG

DnaG

DNA primase, primer synthesis on lagging strand template

Bird et al., 2000

DnaI

–

Co-loader of the DnaC helicase and component of the replication restart primosome

Bruand et al., 2001; Velten et al., 2003

DnaN

DnaN

β-sliding clamp, processivity factor of the replicative polymerases and binding hub for recruitment of alternative polymerases and repair enzymes

Bruck and O’Donnell, 2000; Ogasawara et al., 1986

–

DnaC

Loader of the replicative helicase

Davey and O’Donnell, 2003

–

DnaQ

ε subunit of DNA polymerase III, proofreading (3′–5′) exonuclease

Scheuermann and Echols, 1984

–

DnaT

PriA-dependent replication restart primosome

Sandler and Marians, 2000

DnaX

DnaX

τ subunit of DNA polymerase III, clamp loader, coupling of leading and lagging strand synthesis

Lemon and Grossman, 1998; McHenry, 2003

GyrA

GyrA

DNA gyrase subunit A, DNA breakage and rejoining

Orr and Staudenbauer, 1982

GyrB

GyrB

DNA gyrase subunit B, ATP hydrolysis

Orr and Staudenbauer, 1982

–

Hda

Regulatory inactivation of DnaA, hydrolysis of DnaA-bound ATP in the presence of sliding clamp loaded onto DNA

Kato and Katayama, 2001; Su’etsugu et al., 2004

HolA

HolA

δ subunit of DNA polymerase III, clamp loader wrench

Bruck and O’Donnell, 2000; Noirot-Gros et al., 2002

HolB

HolB

δ′ subunit of DNA polymerase III, clamp loader

Bruck et al., 2005; Bruck and O’Donnell, 2000

–

HolC

χ subunit of DNA polymerase III

Johnson and O’Donnell, 2005

–

HolD

ψ subunit of DNA polymerase III

Johnson and O’Donnell, 2005

–

HolE

θ subunit of DNA polymerase III, dispensable

Johnson and O’Donnell, 2005

LigA

Lig

NAD-dependent ligase

Petit and Ehrlich, 2000

PcrA

UvrD, Rep b

DNA helicase, essential for DNA replication and repair

Petit et al., 1998; Petit and Ehrlich, 2002

PolA

PolA

DNA polymerase I, removal of RNA primers by 5′–3′ exonuclolysis, gap filling and translesion DNA synthesis

Duigou et al., 2005; Yasuda and Okazaki, 1985

PolC

–

Replicative DNA polymerase with proofreading (3′–5′) exonuclease activity

Barnes et al., 1992; Bruck and O’Donnell, 2000; Sanjanwala and Ganesan, 1991

PolY1

DinB b

Y-family DNA polymerase, translesion synthesis in association with PolA

Duigou et al., 2004, 2005

PolY2

UmuC b SOS-induced Y-family DNA polymerase, translesion synthesis in association with PolA, UV-induced mutagenesis

Duigou et al., 2004, 2005

Construction and Maintenance of a Replication Fork |  3

Table 1.1 Continued B. subtilis

E. coli

Protein functiona

References

PriA

PriA

Replication restart primosome assembly, recognition of forked DNA substrates, DNA helicase

Marsin et al., 2001; Polard et al., 2002

–

PriB

PriA-dependent replication restart primosome

Sandler and Marians, 2000

–

PriC

PriC-dependent replication restart primosome, recognition of forked DNA substrates

Heller and Marians, 2005a; Sandler and Marians, 2000

–

SeqA

Regulation of initiation of chromosome replication by sequestration of hemimethylated GATC sites

Lu et al., 1994

SirA

–

Negative regulator of DnaA during sporulation, bind to domain I

Jameson et al., 2014; Rahn-Lee et al., 2009; Rahn-Lee et al., 2011; Wagner et al., 2009

–

Dam

Methylation of GATC sites

Lobner-Olesen et al., 2005

–

DiaA

Positive regulator of DnaA

Ishida et al., 2004

–

IciA

Negative regulator of initiation of chromosome replication

Thony et al., 1991

Soj

–

DnaA regulator, monomer inhibits DnaA filament formation, Murray and Errington, 2008; dimer activates DnaA-dependent replication initiation Scholefield et al., 2012

SSB

SSB

Single-strand binding protein

Polard et al., 2002

YabA

–

Regulation of DNA replication initiation, complex formation with DnaA and DnaN, inhibits DnaA filament formation

Noirot-Gros et al., 2002; Noirot-Gros et al., 2006; Scholefield and Murray, 2013

YpcP

–

Potential 5′–3′ exonuclease, paralogous to the N-terminal exonuclease domain of PolA, co-essential with PolA

Duigou et al., 2005

aFor

proteins present in B. subtilis, function is summarized from studies performed in B. subtilis or related Gram-positive bacteria. For proteins missing in B. subtilis, the function of the E. coli protein is indicated. bThe E. coli protein is not the strict functional homologue of the B. subtilis protein.

a framework to compare and to highlight the differences in molecular mechanisms. The process of DNA replication will also be contextualized with other cellular pathways. The basics of DNA replication in Escherichia coli Replication fork assembly and progression The mechanics of DNA replication have been the focus of several reviews (Langston et al., 2009; McHenry, 2003, 2011; O’Donnell, 2006). The construction of replication forks is a highly regulated process mediated by protein-protein and protein–DNA interactions, through which a dynamic molecular machine able to replicate both DNA strands in a coordinated manner is assembled. The E. coli replisome machinery comprises more than a dozen proteins. Extensive biochemical, structural, and biophysical studies of the E. coli DNA polymerase III holoenzyme (Pol III HE) have led

to a detailed molecular understanding of the different steps and activities that are necessary for its assembly and for the progression of the replication fork. Briefly, the E. coli Pol III HE consists of 10 subunits that assemble into a tripartite complex composed of the catalytic DNA polymerase, the β-clamp processivity factor DnaN, and the clamp loader DnaX (with subunits τ/γ, δ, δ′, ψ and χ). The DnaX complex assembles the dimeric β-clamp ring so that it encircles the DNA template. Binding of the β-clamp to the polymerase creates a tether that holds the enzyme on the DNA. The DnaX complex also coordinates the active DNA polymerases, thus physically coupling the synthesis of the leading (continuous) and lagging (discontinuous) strands. Importantly, the leading strand DNA polymerase is coupled to the helicase DnaB via a bridge formed by the τ subunit. Helicase stimulates the efficiency of DNA duplex unwinding and accelerates the rate of fork progression. However, unwinding of the DNA template by the replicative helicase generates topological constrains that must be removed to allow complete duplication (and segregation) of

4  | Murray et al.

the newly replicated chromosomes; this is achieved by the actions of DNA topoisomerases and DNAcondensing proteins (see Chapter 3) (Hardy et al., 2004; Schvartzman and Stasiak, 2004). The DnaG primase interacts transiently with the DnaB helicase and synthesizes RNA primers every 1–2 kb for initiation of lagging-strand synthesis. The association of the helicase–primase complex with the polymerase holoenzyme at the replication fork completes the replisome. As the fork progresses, ssDNA is generated on the lagging-strand template and is coated with the single-stranded binding protein (SSB) to inhibit secondary structures and protect from nucleolytic attack. The RNA oligoribonucleotides deposited by primase are elongated by DNA polymerase III up to the 5′ end of the next RNA primer to form Okazaki fragments (Kornberg and Baker, 1992b). Adjacent Okazaki fragments are separated by a single-strand interruption (nick). DNA polymerase I extends synthesis from the 3′ end of the nick and simultaneously degrades the downstream RNA primer with its 5′–3′ exonuclease activity. After removal of the RNA primer, the nick is sealed by DNA ligase. The lagging strand polymerase must rapidly dissociate from DNA upon completing each Okazaki fragment, unbinding from the used β-clamp before associating with a new one. As the replication fork advances the lagging strand polymerase is coupled to the leading strand polymerase, resulting in the formation of a DNA loop (‘trombone structure’) that repeatedly grows and subsequently dissolves upon completion of each Okazaki fragment. Advances in our understanding of the dynamics of DNA replication loops at the fork have been reviewed (Geertsema and van Oijen, 2013; Hamdan and van Oijen, 2010; Robinson and van Oijen, 2013). Initiation of DNA replication at the chromosome origin Within the cell formation of an active replication fork is a tightly controlled process. At the chromosomal origin, oriC, initiation occurs only once per cell cycle to ensure that the number of chromosomes remains constant during exponential growth (Boye et al., 2000). The replication origin, oriC, is specifically bound and unwound by the replication initiator protein DnaA (Kornberg and Baker, 1992a; Messer, 2002; Mott and Berger, 2007). Bacteria have evolved diverse regulatory

mechanisms to control chromosomal replication at the initiation step, which mainly regulate the activity of DnaA and/or the accessibility of its DNA sequence targets within oriC (Katayama et al., 2010). Protein interactions between DnaA and the DnaB helicase in complex with the DnaC protein promote the loading of DnaB onto the unwound region (Marszalek et al., 1996; Sutton et al., 1998). The subsequent association of the DnaG primase with the helicase results in the synthesis of an RNA primer on the DNA template. The Pol III HE binds to the primer end, and the DnaX–DnaB interaction completes replication fork assembly. Restart of blocked replication forks As the replication fork progresses along the chromosome it can encounter damage on the DNA template. Some DNA damage can be skipped over by the replisome, leaving the lesion behind in a ssDNA gap for later recombinational repair (see Chapter 2) (Langston and O’Donnell, 2006). However, some DNA damage sites have the potential to stop replication and act as ‘roadblocks’ (McGlynn and Lloyd, 2002; Michel et al., 2004). The persistence of such blocks would prevent completion of chromosome replication and consequently would be lethal. Thus, in order to survive the cell must activate damage-tolerance mechanisms, which involve either bypass of the replicative block or its repair, followed by the restart of replication. Bypass of a replication block can be performed by specialized low-fidelity DNA polymerases that are capable of incorporating nucleotides opposite damaged sites (Friedberg et al., 2005; Tippin et al., 2004). Upon completion of this translesion synthesis, which is often mutagenic (because an incorrect base is inserted), the replicative polymerase resumes DNA synthesis with its usual high fidelity. The shift from replicative to translesion synthesis at sites of blocked forks involves the polymerase sliding clamp (β2) being able to accommodate more than one polymerase at the same time (Friedberg et al., 2005; Indiani et al., 2005). The central role of the sliding clamp in modulating DNA polymerase switching highlights the dynamic nature of the replication machinery. Alternatively, the E. coli replisome may stall upon collision with a DNA binding protein such as a co-transcribing RNA polymerase, but instead of collapsing the replication fork remains intact and

Construction and Maintenance of a Replication Fork |  5

later resumes synthesis (Pomerantz and O’Donnell, 2010). Single-molecule analysis has revealed the capacity of the replicative DNA polymerase to hop from one β clamp to another without leaving the replication fork. Relatedly, it has been demonstrated that mRNA transcripts can also be extended by the leading strand polymerase, as occurs during lagging strand replication (Pomerantz and O’Donnell, 2008). These observations support the notion that DNA synthesis is likely to be discontinuous on both strands. In cases where the replisome encounters a blocking DNA adduct or lesion that causes the fork to disassemble, this deleterious event is overcome by eliminating the cause of the arrest and then directing the re-assembly of the replisome. Replication restart (RR) is mediated by specific proteins that associate in various combinations to act on different types of arrested forks (Marians, 2004; Michel et al., 2004). For instance, recombination intermediates can be used to generate forked DNA onto which the replisome is reassembled (see also Chapter 3). A key protein in this process is the PriA helicase, which specifically binds forked-DNA and ultimately promotes the recruitment of the DnaB helicase and its loading onto ssDNA, followed by the subsequent assembly of a functional replisome (Kogoma, 1997; Michel et al., 2004; Sandler and Marians, 2000; Xu and Marians, 2003). Termination of replication Replication forks end their progression along the E. coli chromosome at physiological arrest sites in the terminus region. The termination protein Tus binds to ter sites on the DNA and the Tus-ter complex acts by blocking the replication fork approaching from one direction but not from the other (Neylon et al., 2005). This way, a fork that has transversed the 180° point will be arrested on the other chromosome arm, until the slower fork arrives. The mechanism that determines the polarity of the Tus-ter block is a molecular mousetrap operating at the nonpermissive face of the Tus–ter complex (Berghuis et al., 2015; Elshenawy et al., 2015; Mulcair et al., 2006). The trap is set by the binding of Tus to the ter site and is sprung by the incoming DnaB helicase, which separates the strands within the Ter site and triggers the formation of a locked Tus–ter complex upon unwinding of a strictly conserved G-C base pair. On the strand displaced by the DnaB helicase,

the conserved cytosine residue in ter interacts with a cryptic cytosine-specific site on Tus to form the lock. The locked Tus–ter complex is extremely stable and halts progression of the DnaB helicase. However, the locked Tus-ter can be readily dissociated by a fork approaching from the permissive side of the complex, thus allowing complete replication of the terminus region (Mulcair et al., 2006). Chromosomal DNA replication in Bacillus subtilis The DNA replication machinery The replicases B. subtilis DNA polymerase III, encoded by the polC gene, is essential for chromosome replication (Gass and Cozzarelli, 1973). The PolC polypeptide carries a DNA polymerase active site in its C-terminal domain and a 3′–5′ proofreading exonuclease in its N-terminal domain (Barnes et al., 1992; Sanjanwala and Ganesan, 1989, 1991). This is in contrast to the E. coli DNA polymerase III in which the polymerase and the proofreading exonuclease activities are encoded by the dnaE (α subunit) and dnaQ (ε subunit) genes, respectively (Table 1.1). PolC also contains a zinc finger-like structure, not present in the Gram-negative α subunit, that tightly binds zinc and is essential for polymerase function (Barnes et al., 1998). Genome sequence analyses revealed that PolC is highly conserved in low G+C Gram-positive bacteria (Lemon et al., 2002). The PolC holoenzyme replicases from four Gram-positive organisms, B. subtilis, Streptococcus pyogenes, Staphylococcus aureus and Mycobacterium tuberculosis have been reconstituted in vitro from purified subunits (Bruck et al., 2005; Bruck and O’Donnell, 2000; Gu et al., 2016; Sanders et al., 2010). The polC gene (or dnaE1 and dnaQ genes for M. t.), as well as the widely conserved dnaX, holA, holB, dnaN and ssb genes, encoding the subunits τ, δ, δ′, β and SSB, respectively, were expressed in E. coli and the resulting proteins purified. The dnaX genes from all Gram-positive species produce only the full-length protein τ. This is in contrast to E. coli dnaX, which encodes τ, the full-length product, and γ, a shorter product generated by translational frameshifting. Although the B. subtilis τ protein has a similar domain organization to E. coli τ (Haroniti

6  | Murray et al.

et al., 2004; Martinez-Jimenez et al., 2002), atomic force microscopy (AFM) revealed that it forms a crescent-shaped structure that resembles the overall configuration of the E. coli γ complex (Haroniti et al., 2003, 2004;). The τδδ’ complex uses the energy of ATP hydrolysis to open the β ring and load it as a functional sliding clamp around the DNA. No detectable homologues of the clamp loader ψ and χ subunits, which increase the affinity of τ(γ) for δ and δ′ in the DnaX complex of E. coli (McHenry, 2003), have been found in the genomes of Grampositive bacteria (Lemon et al., 2002). However, it is possible that orthologous proteins could act to stabilize the interactions within the clamp loader of these organisms. In combination, the clamp loader complex and the β clamp endow PolC with a high speed (~700 nucleotides/second) and processivity that is comparable to that of the E. coli Pol III holoenzyme. Thus, the reconstituted polymerase complexes from B. subtilis, S. pyogenes, S. aureus, and M. tuberculosis share the characteristics of previously characterized Gram-negative replicases (Bruck et al., 2005; Bruck and O’Donnell, 2000; Gu et al., 2016). In addition, the pair wise interactions between PolC, τ, δ, δ′ and β identified in these reconstitution studies are also found between the B. subtilis subunits (with the exception of the δ-δ′ interaction) using a yeast two-hybrid assay (NoirotGros et al., 2002). This further underscores the conserved organization of the PolC replicases in Gram-positive bacteria. A pentameric recognition sequence (consensus: QL[S/D]LF) termed the clamp-binding motif mediates the interaction of many proteins with the β clamp (Dalrymple et al., 2001; Wijffels et al., 2004) and is present at the C-terminus of virtually all PolC proteins from low G+C Gram-positive bacteria (Wijffels et al., 2005). Interestingly, the E. coli α (polymerase) subunit contains two β-binding motifs (Wijffels et al., 2005); one at the C-terminus that is important for coordinating interactions between β and τ (Lopez de Saro et al., 2003), and one located internally that promotes β-binding in vitro and is critical for replication in vivo (Dohrmann and McHenry, 2005). In addition to PolC, many Gram-positive bacteria also encode a second DNA polymerase, DnaE, related to the E. coli Pol III α subunit. DnaE is essential for chromosome replication in B. subtilis and S. aureus (Dervyn et al., 2001; Inoue et al., 2001).

Similarly to PolC, DnaE is required for the elongation phase of chromosome replication (Dervyn et al., 2001). In DnaE-depleted cells, a plasmid with unidirectional replication specifically produces a single-stranded DNA corresponding to the leading strand, indicating that DnaE is involved in lagging strand synthesis. In addition, the subcellular localization of DnaE is similar to that of PolC, suggesting that both polymerases are functioning at the B. subtilis replication fork (Dervyn et al., 2001). Recent work based on the in vitro reconstitution of an active B. subtilis replication fork brought significant advances to the understanding of the essential roles played by the two replicases during DNA synthesis (Sanders et al., 2010). It was found that B. subtilis PolC is the major DNA replicative enzyme of the leading strand while both PolC and DnaE are required for lagging-strand synthesis. In the presence of the full complement of proteins that compose the replisome machinery the rate of synthesis by the PolC holoenzyme correlated well with the progress of the replication fork in vivo (about 500 nt/s), while the reaction supported by DnaE alone was slow (25 nt/s), even in the presence of τ and β2. The authors also observed that both PolC and DnaE could efficiently elongate a DNA primer, but that under the conditions used only DnaE could elongate an RNA primer. Finally, strong evidence was provided for a handoff mechanism between DnaE and PolC, whereby RNA primers are first extended by DnaE and are then elongated by PolC during lagging strand replication (Sanders et al., 2010). Hence, the Bacillus replication machinery assembled at active replication forks likely contains two distinct replicative DNA polymerases, PolC and DnaE. This architecture differs from that of E. coli where identical PolIII enzymes are assembled asymmetrically to efficiently replicate the two DNA strands, but it is similar to the eukaryotic replisome where a combination of Pol α and Pol δ are required for lagging strand synthesis (Table 1.2). Thus, it appears that the need for specialized leading and lagging strand polymerases might be evolutionarily conserved (McHenry, 2003). The purified B. subtilis DnaE enzyme alone lacks an associated proofreading 3′–5′ exonuclease activity but can synthesize DNA on an SSB-coated ssDNA template at low speed (Bruck and O’Donnell, 2000; Le Chatelier et al., 2004; Sanders et al., 2010). The presence of the β sliding

Construction and Maintenance of a Replication Fork |  7

Table 1.2  Differences in replication forks composition in E. coli, B. subtilis and yeast Replisome factor

E. coli

B. subtilis

Eukaryotic

Leading strand polymerase

Pol III

Pol C

Pol δ, Pol ε

Lagging strand polymerase

Pol III

DnaE + Pol C

Pol α + Pol δ

Clamp

[τ/γ]δδ′ψ χ

β2

PCNA

Clamp loader

β2

τδδ′

RFC

Helicase

DnaB

DnaC

MCM2–7

Helicase loader

DnaC

DnaB/I (+ DnaD)

Cdc6/Cdt1

Primase

DnaG

DnaG

Primase

SSB

SSB

SSB

RPA

clamp increases the processivity of DnaE but not its intrinsic speed (Bruck and O’Donnell, 2000). Another striking property of the DnaE polymerase is its capacity to bypass many types of DNA lesions in vitro that generally block other replicative polymerases (Bruck et al., 2003; Le Chatelier et al., 2004). The bypass is efficient, highly error-prone, and takes place either by extension of misaligned 3′ termini (thereby generating single base deletions in the nascent strand) or by direct extension of mispaired termini, depending on the nature of the lesion (Bruck et al., 2003; Le Chatelier et al., 2004). Notably, the expression of DnaE is induced about 3-fold upon treatment of B. subtilis cells with DNAdamaging agents (Le Chatelier et al., 2004), and the SOS repressor DinR binds to the dnaE promoter region (Au et al., 2005). Although the expression of DnaE is SOS-controlled (see Chapter 3 for more details), DnaE is unlikely to function as an error-prone polymerase in vivo because its overproduction does not increase the rate of spontaneous mutagenesis (as occurs following overproduction of an error-prone Y-family polymerase; see below) (Le Chatelier et al., 2004). A potential role of DnaE in DNA repair and mutagenesis remains to be investigated. The primosome The primosome is a complex between helicase and primase that is required for primer synthesis during DNA replication. Although primase alone can catalyse oligoribonucleotide synthesis, this activity is increased 300-fold in the presence of helicase (Tougu et al., 1994). The ordered assembly of the primosome onto the DNA is directed at specific sites, such as the chromosomal replication origin

or at arrested forks, by the master initiator protein, DnaA, and the helicase, PriA, respectively (see above). In E. coli, the loading of the DnaB helicase at oriC occurs through the formation of a DnaB6– DnaC3 complex in which the N-terminal domain of DnaC is physically associated with the C-terminal domain of DnaB (Makowska-Grzyska and Kaguni, 2010) (Table 1.1). The transition from initiation to elongation is triggered by the recruitment of the primase, DnaG, to the N-terminus of DnaB and concomitant dissociation of DnaC, to promote the synthesis of RNA primers. This reaction of ‘general priming’ is inhibited in the presence of SSB (Marians, 2000). In the absence of PriA E. coli cells are deficient in the restart of arrested replication forks and rapidly accumulate mutations in dnaC that restore proficiency for restart. The DnaC810 suppressor protein has gained the capacity to load DnaB onto SSB-coated DNA. These findings indicate that in the cell, primosome assembly onto ssDNA has to overcome SSB coating that acts as a safeguard to prevent random initiation events at single-stranded DNA regions in the chromosome (Marians, 2000; Sandler and Marians, 2000). In B. subtilis the dnaC and dnaG genes encode the functional counterparts of the E. coli DnaB helicase and DnaG primase, respectively, and the initiator proteins DnaA and PriA are also conserved (Table 1.1). However, the DnaD, DnaB and DnaI proteins, which are components of the B. subtilis primosome (Bruand et al., 1995; Bruand et al., 2001), are not conserved in E. coli and conversely the E. coli PriA partners, PriB, PriC and DnaT, are not present in the B. subtilis genome (Lemon et al., 2002). The loading of the E. coli replicative helicase DnaB1 involves a homo-oligomeric DnaC complex

8  | Murray et al.

(Marians, 2000). In contrast, DnaD, DnaB, and DnaI are all required to load the helicase, DnaC,1 at oriC or at stalled forks during replication restart in B. subtilis. The B. subtilis helicase loader, DnaB, is multifunctional; biochemical characterization has shown that it cooperates with DnaI for the loading of the helicase, DnaC, in vitro (Velten et al., 2003). Evidence suggests that both DnaB and DnaD are necessary to promote helicase loading at oriC and that DnaB is recruited to the oriC–DnaA–DnaD complex by interaction with DnaD (Rokop et al., 2004; Smits et al., 2010). DnaB has been shown to undergo proteolysis at its C-terminus in a growth phase-dependent manner, which could play a role regulating its interaction with DnaD and hence its recruitment to oriC (Grainger et al., 2010). The observation of structural similarities between DnaB and DnaD is reflected in their ability to oligomerize upon binding to DNA. Furthermore, DnaD possesses DNA remodelling activity on supercoiled DNA, forming protein-DNA scaffolds that could facilitate DnaA-dependent duplex unwinding at oriC (Zhang et al., 2008). B. subtilis DnaC belongs to the F4 family of ATP-dependent annular hexameric helicases (Gorbalenya, 1993). Purified DnaC self-assembles into a homohexamer in the presence of ATP (Velten et al., 2003). However, neither the DnaC monomer nor the hexamer exhibits helicase activity in vitro. This is in sharp contrast to the replicative helicases of E. coli and Bacillus stearothermophilus which alone display helicase activity in vitro (Bird and Wigley, 1999; Davey and O’Donnell, 2003). In B. subtilis, as well as in Geobacillus kaustophilus, the ATPase DnaI interacts with DnaC in the presence of ATP and promotes the formation of a dodecameric complex containing 6 units of each protein (Tsai et al., 2009; Velten et al., 2003). The C-terminal domain of DnaI belongs to the structurally well-characterized AAA+ superfamily of ATPases (see Chapter 7), while the N-terminal domain adopts a novel zinc-binding fold important for interaction with DnaC (Loscha et al., 2009). Interestingly, evidence suggests that DnaI and DnaB act together to mediate the functional

loading of DnaC onto ssDNA by promoting its hexamerisation around the ssDNA (Velten et al., 2003). The mutant protein DnaB75 (DnaBS371P), which suppresses the defects of a priA null mutant (Bruand et al., 2001), binds with higher affinity to a forked DNA substrate and stimulates the DnaIdependent helicase activity of DnaC more than wild-type DnaB (Velten et al., 2003). Together with the observation that the conversion of circular ssDNA into dsDNA is highly efficient in dnaB75 cells (Bruand et al., 2001), these findings are consistent with the notion that the DnaB75 protein has gained the capacity to promote helicase loading onto SSB-coated ssDNA molecules (Velten et al., 2003). Thus, as in E. coli, the coating of ssDNA by SSB in wild-type B. subtilis cells may act as a safeguard to prevent random initiation events, and the capacity to deliver the replicative helicase onto SSB-coated ssDNA likely is a general property of helicase loading systems in bacteria. The DnaG primase associates with the translocating replicative helicase and deposits RNA primers along the ssDNA template. This critical interaction is mediated by the carboxy-terminal domain of the primase (DnaG-Cter) both in E. coli and in B. stearothermophilus2 (Soultanas, 2005). The three-dimensional structures of the E. coli and B. stearothermophilus DnaG-Cter domains reveal a similar architecture composed of two subdomains; a large helix bundle followed by a smaller helix hairpin (Oakley et al., 2005; Syson et al., 2005). The two subdomains have distinct functions; the helix hairpin module alone is sufficient for strong binding to the replicative helicase, but the larger subdomain is required to stimulate helicase ATPase activity. Remarkably, the DnaG-Cter helix bundle is structurally homologous to the N-terminal domain of E. coli DnaB helicase and superposition of the domains reveals a network of spatially conserved surface residues that are largely invariant in the replicative helicases of 14 bacterial species (Soultanas, 2005). The creation of chimeric proteins in which the C-terminal region of DnaG was swapped with the N-terminal region of DnaB revealed that these domains are not only structrural but also functional homologues (Chintakayala et

1 It is important to note the nomenclature here; in E. coli and B. stearothermophilus the replicative helicase is DnaB, whereas in B. subtilis it is DnaC.

2  Importantly, note that the nomenclature of the replicative helicase is DnaB in E. coli and B. stearothermophilus whereas it is DnaC in B. subtilis.

Construction and Maintenance of a Replication Fork |  9

al., 2007). In B. stearothermophilus the DnaG primase interacts with the flexible linker that connects the N- and C-terminal domains of the helicase, inducing the hexameric helicase to adopt a 3-fold symmetrical conformation (a trimer of dimers) with three primase molecules bound to the helicase ring (Soultanas, 2005; Thirlway et al., 2004). The crystal structure of hexameric DnaB helicase complexed with the helicase binding domain (HBD) of DnaG revealed a distinct two-layered ring structure formed by the two domains of DnaB (Bailey et al., 2007). Binding of B. stearothermophilus DnaG to the replicative helicase reduces the length of synthesized primers (Thirlway and Soultanas, 2006). Some helicase mutants bind to DnaG but fail to modulate the length of primers whereas other helicase mutants inhibit primase activity. Taken together, these observations indicate that within the replisome helicase and primase are capable of modulating each other’s activities (Chintakayala et al., 2008; Soultanas, 2005; Thirlway and Soultanas, 2006). The assembly of the replication machinery is complete when the primosome connects with the DNA polymerase holoenzyme via an interaction between the τ subunit and helicase. The B. subtilis τ protein interacts with the B. stearothermophilus DnaB replicative helicase through its C-terminal domain. This interaction stimulates helicase activity and is essential for rapid fork progression in E. coli (see above). The architecture of the complex revealed by atomic force microscopy suggests that a crescent-shaped τ pentamer interacts with the bell-shaped DnaB hexamer (Haroniti et al., 2003, 2004). The τ–helicase complex plays an additional role by allosterically modulating primase activity through a τ–DnaB−DnaG ternary complex (Chintakayala et al., 2009). The role of SSB at the replicative fork During fork progression, ssDNA corresponding to the lagging strand template is generated by the activity of the replicative helicase and subsequently is coated by single-stranded DNA binding (SSB) protein. SSBs protect exposed ssDNA and prevent the formation of secondary structures at the fork to promote replisome progression. SSB assembles into a stable homotetramer and binds ssDNA through its N-terminal OB (Oligonucleotide/

Oligosaccharide-Binding) domain (Raghunathan et al., 2000). Beyond its role in DNA binding, SSB has been shown to physically interact with various replication proteins which facilitates their recruitment to forks. For instance, E. coli SSB interacts with the χ subunit of the clamp loader complex and with DnaG (Benkovic et al., 2001). These interactions require a short evolutionarily conserved amphipathic signature motif of about 10 residues located at the tip of the SSB C-teminal domain (Shereda et al., 2009). In B. subtilis there are two SSB paralogues, SSB (also called SsbA) and YwpH (SsbB), encoded by the ssb and ywpH genes, respectively. The two proteins share 63% identity in the N-terminal DNA binding domain, but SsbB lacks the C-terminal domain of SSB. SSB is essential for viability whereas SsbB is dispensable (Kobayashi et al., 2003). Transcription of ssb is high in fast growing cells and is induced during the SOS response, whereas ywpH is strongly induced during competence development (Berka et al., 2002; Hahn et al., 2005; Lindner et al., 2004). Thus, in B. subtilis, SSB displays a housekeeping role during chromosome replication and genome maintenance, whereas SsbB acts during transformation, possibly by binding the incoming ssDNA and facilitating its recombination into the chromosome (see Chapter 13). The role of B. subtilis SSB during progression of the fork can be gauged by its binding partners. Currently, the SSB C-terminus interaction network is composed of 12 proteins involved in various DNA metabolic pathways (including DNA repair), all of which are enriched at the replicative fork via the C-teminal domain of SSB (Costes et al., 2010). Interestingly, it was found that DnaE-GFP did not accumulate at the replisome in a strain expressing an SSB derivative lacking its C-terminal interaction motif. This suggests that DnaE accumutates at active chromosomal forks through a physical interaction with SSB whereas PolC is known to bind to other components of the replisome machinary. Bacillus SSB also binds the primary DNA replication restart protein PriA and the DNA recombination effectors RecG and RecQ (Lecointe et al., 2007). These three DNA helicases were shown to be continually associated with active replication forks. In the case of PriA, repair activity at arrested forks was found to require interaction with the SSB C-terminus binding motif (Lecointe et al., 2007). Thus, it appears

10  | Murray et al.

that SSB acts as a platform for the recruitment of DNA-processing enzymes in anticipation of their role in replication restart. It has been found that both SSB and SsbB can be phosphorylated on the conserved residue, tyrosine-82, by the protein-tyrosine kinase, YwqD, and dephosphorylated by the phosphatase, YwqE (Mijakovic et al., 2006). Phosphorylation of SSB appears to increase its affinity for ssDNA and is reduced upon treatment of cells with the DNA damaging agent, mitomycin C. Tyrosine phosphorylation of SSB also occurs in E. coli and in Streptomyces coelicolor, indicating that it is an evolutionarily conserved process. These findings suggest that SSB phosphorylation may act to regulate some as yet unknown aspects of DNA metabolism (Mijakovic et al., 2006). Other replication proteins In B. subtilis Okazaki fragments are processed by DNA polymerase I, using its 5′–3′ exonuclease activity to remove RNA primers and 5′–3′ polymerase activity to simultaneously fill the resulting gaps (Okazaki et al., 1968; Tamanoi et al., 1977). Nicks are subsequently sealed by a DNA ligase. The B. subtilis chromosome encodes two ligases: the NAD-dependent ligase, LigA (YerG), and the ATP-dependent ligase, LigB (YkoU). LigA is essential for viability and fully complements the growth and UV sensitivity defects of an E. coli ligts mutant (Petit and Ehrlich, 2000), indicating that LigA acts to seal Okazaki fragments processed by DNA Pol I. LigB is not essential and participates in the nonhomologous end joining (NHEJ) pathway to repair double-strand breaks (see Chapter 2) (Weller et al., 2002). In B. subtilis removal of the topological constraints generated as the replicative helicase unwinds the DNA template is achieved by: the three essential DNA topoisomerases Topoisomerase I (TopA), DNA gyrase (GyrAB) and Topoisomerase IV (ParCE); the general DNA binding protein HBsu (Kohler and Marahiel, 1997); and the SMCScpAB condensin complex which is required for chromosome organization and segregation (see Chapter 3) (Lindow et al., 2002; Mascarenhas et al., 2002; Nolivos and Sherratt, 2014; Soppa et al., 2002).

The DNA replication factory The visualization of replisome proteins (e.g. PolC, DnaE, τ, δ, δ′, β, SSB) in living cells, using functional fusions with green fluorescent protein (GFP), revealed localization predominantly at or near midcell (Fig. 1.1) (Dervyn et al., 2001; Lemon and Grossman, 1998; Meile et al., 2006). This midcell localization of the replisome depends upon DNA replication elongation and when replication is blocked at a specific chromosomal locus this site colocalizes with the DNA polymerase at midcell (Lemon and Grossman, 1998, 2000). Upon release of the block and resumption of DNA replication the duplicated chromosome sites are segregated towards opposite cell poles away from the centrally located replisome. These data indicate that in B. subtilis active DNA replication forks are predominantly maintained near midcell, forming a replication factory. The replication factory contains at least two replisomes assembled at two forks, each replicating one half of the chromosome (Lemon and Grossman, 1998). Although it most frequently appears in discrete positions at midcell, time-lapse microscopy has revealed that the factory is mobile around the cell centre, suggesting that the factory is not rigidly fixed at a specific midcell position (Migocki et al., 2004). Also, a PolC-GFP focus can split into two foci that will merge again later, suggesting that the two replisomes in the factory are able to separate and come back together during a given round of replication (Migocki et al., 2004). Interestingly, the master replication initiation protein, DnaA, was found to be associated with the replication machinery during most of the cell cycle (Soufo et al., 2008). Likewise YabA and SirA, which are negative regulators of DnaA, localize at the replication factory (Hayashi et al., 2005; Jameson et al., 2014; Noirot-Gros et al., 2006) (Fig. 1.1). In addition DNA repair enzymes, such as MutS (mismatch repair) and SbcC, are dynamically associated with the factory during replication (Fig. 1.1) (Meile et al., 2006; Simmons et al., 2008; Mascarenhas et al., 2006). Taken together, these findings suggest that many auxiliary proteins are recruited to the replication factory as and when required. These findings also support the accumulating evidence that chromosomal and extrachromosomal elements act at

Construction and Maintenance of a Replication Fork |  11

Figure 1.1  Subcellular localization of proteins at the replication factory. (A) Left: Localization of key components of the replication machinery: the replicative polymerase α-subunit, PolC; the τ-subunit, DnaX; the single strand binding protein, SSB; and the C-family DNA polymerase DnaE. Green fluorescent protein (GFP)-fusions were expressed in living cells, which were subsequently mounted on agarose slides and observed by fluorescent microscopy. All four fusion proteins localize at the replication factory with typical single or dividing foci located at or near the cell centre. Right: The localization of the β-clamp subunit, DnaN, and the Rad50-homologue, SbcC, at the replication factory depends upon active DNA replication. Using a strain in which the expression of the DnaA initiator is controlled by the IPTG-inducible promoter, Pspac, the formation of GFP-DnaN and GFP-SbcC foci requires the presence of IPTG (Meile et al., 2006). (B) Left: Domain organization of DnaA and a crystal structure showing the DnaA filament bound to ssDNA through the AAA+ domain (PDB: 3R8F). Right: DnaA assembles into a filament specifically on a single DNA strand containing an array of DnaA-trios. DnaA filament formation can be captured by cross-linking and visualized on SDS-PAGE as higher-order oligomeric species. Assembly of the DnaA filament requires loading from double-stranded DnaA-boxes, is dependent upon ATP and the ssDNA-binding residue Ile190, and occurs specifically on repeating trinucleotide sequences termed DnaA-trios (3′-GAT-5′) (Richardson et al., 2016). (C) Living cells expressing various GFP-fusion proteins involved in genome maintenance (indicated on the top of each set of panels) in ssb3+ cells (isogenic wild-type) or ssbΔ35 cells (possessing a truncation mutant of SSB lacking the C-terminal 35 amino acids). White arrowheads point to visible GFP foci. The loss of distinct foci in the ssbΔ35 strain reveals that the SSB C-terminus is required for localization of the DnaE (but not PolC) replicase at the fork, and for localization of SbcC and RecJ (but not YabA) (Costes et al., 2010).

12  | Murray et al.

specific intracellular locations in bacteria (Bravo et al., 2005). Initiation of DNA replication at the chromosomal origin DnaA and oriC The ubiquitous bacterial DNA replication initiator DnaA is a multifunctional protein composed of four distinct domains that act in concert to promote opening of the DNA duplex and deposition of the DNA replication machinery (Fig. 1.1B) (Mott and Berger, 2007). The C-terminal domain IV contains a helix–turn–helix motif that specifically recognizes nine base-pair duplex DNA sequences called ‘DnaA-boxes’ (consensus 5′-TTATCCACA-3′). Domain IV is connected to domain III by a flexible α-helix, which is observed to adopt alternative conformations in different crystal structures of DnaA proteins. Domain III is composed of an initiator specific AAA+ motif, conserved amongst DNA replication initiator complexes. The AAA+ motif is capable of binding and hydrolysing ATP, and it acts as the primary oligomerization determinant for the protein. Additionally, domain III contains the residues required for DnaA to interact with single-stranded DNA. Domain II tethers domains III/IV to domain I, and domain I acts as a protein interaction hub that promotes DnaA dimerization and facilitates loading of the replicative helicase. A comparison of 104 different DnaA proteins has shown that domains III and IV are the most similar (Messer, 2002), indicating that the ATP-dependent assembly of DnaA oligomers is likely to be a conserved activity across bacterial species. Oligomeric structures of DnaA in complex with the non-hydrolysable ATP analog AMP-PCP have been determined in the presence and absence of single-stranded DNA (Duderstadt et al., 2011; Erzberger et al., 2006). In both structures DnaA assembles into a right-handed helical filament that is built upon interactions between AAA+ motifs and that requires a conserved ‘arginine finger’ to contact the γ-phosphate of the nucleotide bound by the adjacent protein. The structure of the DnaAAMP-PCP:ssDNA complex revealed that each DnaA protein contacts the phosphate backbone of three nucleotides through the AAA+ motif. It has been proposed that the DnaA filament stretches the

ssDNA to promote opening of the duplex replication origin. Archetypical bacterial genomes have a unique replication origin that contains several specific binding sites for DnaA. The mechanism of DnaAmediated initiation at oriC has been historically studied in E. coli (Messer, 2002), with significant insights into DnaA structure coming from more recent studies using the Aquifex aeolicus homologue (Duderstadt et al., 2010, 2011; Erzberger et al., 2002, 2006). DnaA first binds to high affinity DnaA-boxes within oriC using its dsDNA binding motif in domain IV. Following ATP-binding, DnaA assembles co-operatively onto lower affinity sites, ultimately triggering DNA duplex unwinding (Krause et al., 1997). Recently, a new replication origin element called the DnaA-trio (3′-GAT-5′) was identified within B. subtilis oriC (Richardson et al., 2016). The DnaA-trio is a repeating trinucleotide motif that directs assembly of the ATP-dependent DnaA filament onto one strand of the DNA duplex promote open complex formation (Fig. 1.1B). DnaA-mediated unwinding of the origin is a crucial step in initiation as it triggers loading of the helicase onto ssDNA. In E. coli a direct interaction between DnaA and the helicase/DnaC complex promotes the loading of helicase onto the unwound DNA (Marszalek et al., 1996; Sutton et al., 1998). In B. subtilis three initiation proteins, DnaD, DnaB, and DnaI are required to assist DnaA in the recruitment and loading of the replicative helicase, DnaC, at oriC (Fig. 1.2) (Gross et al., 1968; Imai et al., 2000; Karamata and Gross, 1970) (see above). DnaD directly interacts with DnaA and DnaB (Bruand et al., 2005; Ishigo-Oka et al., 2001; Rokop et al., 2004) and co-operates with DnaB and DnaI to orchestrate the loading of the replicative helicase, DnaC, around ssDNA (Smits et al., 2010; Velten et al., 2003). It has been found that these proteins are recruited to oriC in a hierarchical order, with DnaD requiring DnaA, DnaB requiring DnaA/DnaD, and helicase requiring DnaA/DnaD/DnaB (Smits et al., 2010). The assemblage of DnaD, DnaB and DnaI is called the prepriming complex and, in contrast to E. coli, these proteins are also components of the PriAdependent replication restart system (Bruand et al., 1995, 2001).

Construction and Maintenance of a Replication Fork |  13

A

Replication Restart

Initiation

Arrested and repaired fork

oriC

Lagging strand

SSB

DnaA boxes Leading strand

DnaA

DnaD

(i)

PriA (i)

(ii) DnaI

DnaB

(iii) DnaC Helicase Helicase loading by ring-assembly

B

Figure 1.2  The DnaA and PriA primosomes in B. subtilis. (A) The DnaA and PriA primosomes are two multiprotein complexes that promote initiation of chromosome replication by the assembly of a pair of replisomes at oriC (left), and replication restart by the assembly of a single replisome on arrested and repaired chromosomal forks (right), respectively. DnaA and PriA specifically recognize loci at which primosome assembly is needed: DnaA binds to an array of short dsDNA motifs (DnaA-boxes) within oriC; PriA targets forked DNA substrates with a double-stranded leading-strand arm and a single-stranded lagging-strand arm. Once specifically bound to their respective substrates, DnaA and PriA direct the recruitment and loading of the ring-shaped hexameric replicative helicase, DnaC, around ssDNA at the tip of the fork. Importantly, DnaC is loaded on the lagging-strand template where DNA replication is discontinuous. In subsequent steps (not shown) the loaded helicase recruits the other components of the replisome. A hallmark of the B. subtilis DnaA and PriA primosomes is that they both rely on the same triad of proteins to recruit and load DnaC, i.e. the essential DnaD, DnaB and DnaI proteins, which are highly conserved in Gram-positive bacteria. Bi-directional arrows indicate the characterized physical interactions between proteins, leading to the representation of two primosomal cascades initiated by DnaA and PriA binding to their specific DNA substrates (unidirectional arrows). The cascades involve a series of protein interactions: (i) first, DnaD is contacted either by DnaA or PriA bound to DNA; (ii) next, DnaB is contacted by DnaD and helps overcome the SSB barrier; (iii) then, DnaC is dually contacted by DnaB and DnaI, which leads to the functional loading of the helicase as a hexamer encircling the ssDNA. The loading takes place through a ring-assembly mechanism, which relies on the recruitment of individual monomers of DnaC by DnaI and DnaB on ssDNA to promote DnaC hexamerization around the ssDNA template. Thick arrowheads indicate the direction of replication. (B) SSBCter interactome at the active fork. SSB tetramers act as a platform for recruiting partners involved in fork restart upon adventitious blockage. The replisome machinery re-assembles on the branched DNA backbone of the fork. The recruitment of PriA is crucial and takes place when arrest results from replisome dismantling. In more complex situations other actions, aimed at protecting and/or clearing the fork, may occur via individual or concerted actions of the many members of the SSBCter interactome.

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The E. coli replication origin contains specific binding sites for accessory proteins like IHF and Fis that facilitate origin unwinding (Messer, 2002). In B. subtilis it is not known whether sequence-specific DNA binding proteins other than DnaA associate with oriC during initiation. Potentially the DNA-remodelling activities of DnaD and/or DnaB could play roles analogous to IHF/Fis to facilitate open complex formation (Carneiro et al., 2006; Turner et al., 2004; Zhang et al., 2005). The control of initiation In bacteria, chromosome replication is tightly regulated to ensure that initiation at the chromosomal origin takes place only once per cell cycle (Boye et al., 2000). This regulation involves a balance between triggering initiation at a specific time of the cell cycle and repressing it to prevent extra initiation events. Different initiation control pathways have evolved to regulate the activity and the availability of DnaA during the cell cycle (Katayama, 2001; Marczynski and Shapiro, 2002). In E. coli, the paradigm for initiation studies, the predominant mechanism that prevents overinitiation is the regulatory inactivation of DnaA, termed RIDA, which acts after initiation to promote the switch from active ATP-DnaA to the inactive ADP-DnaA form. RIDA is mediated by Hda, a protein that is paralogous to DnaA and whose activity requires interaction with the β clamp (Camara et al., 2005; Katayama et al., 1998; Kato and Katayama, 2001; Su’etsugu et al., 2005). A second level of regulation is the sequestration of newly replicated oriC regions by the SeqA protein (Lu et al., 1994; Taghbalout et al., 2000; von Freiesleben et al., 1994). SeqA binds with high affinity to hemimethylated dam sites (GATC) found throughout the genome and especially enriched at oriC and within the dnaA promoter. In doing so, SeqA temporarily restrains re-initiation both by preventing DnaA access to oriC and by inhibiting DnaA expression (Campbell and Kleckner, 1990). The third level of initiation control involves the titration of large amounts of DnaA protein to the datA locus (Kitagawa et al., 1998), which stimulates hydrolysis of the DnaA-bound ATP and lowers the concentration of active DnaA in the cell (Kasho and Katayama, 2013; Ogawa et al., 2002).

DiaA is a positive regulator of DnaA and is required to ensure timely initiation of chromosome replication (Ishida et al., 2004). The structure of DiaA and functional analysis of defective mutants revealed that it binds as a tetramer to multiple DnaA molecules to stimulate inter-DnaA interactions (Keyamura et al., 2007). DiaA is proposed to positively regulate ATP–DnaA assembly, thus promoting the formation of the open complex at oriC (Keyamura et al., 2009). In addition, two different mechanisms have been described that regenerate ADP-DnaA into active ATP-DnaA. One relies on an interaction of ADP-DnaA with acidic phospholipids present in the membrane (Boeneman and Crooke, 2005) and the other involves two specific chromosomal loci called DnaA-reactivating sequences (DARS1 and DARS2) (Fujimitsu et al., 2009). All together this network of control systems contributes to proper regulation of DNA replication in order to maintain a once-per-cellcycle initiation rate in E. coli cells. Although well characterized, the conservation of these regulatory proteins is limited to the Enterobacteriaceae and other bacteria appear to have developed distinct mechanisms for initiation control. In B. subtilis there are no homologues of E. coli Hda, SeqA, DiaA or Dam. Nonetheless, DNA replication initiation is regulated as tightly as in E. coli (Lemon et al., 2002; Moriya et al., 1999) and several mutants affecting the regulation of initiation have been isolated. The B. subtilis initiation regulator, YabA, was revealed in a yeast two-hybrid screen to identify the protein interaction network of the DNA replication machinery (Noirot-Gros et al., 2002). A ΔyabA mutant exhibits overinitiation and replication asynchrony phenotypes, indicating that YabA acts as a negative regulator of initiation (Noirot-Gros et al., 2002). YabA assembles into a tetramer mediated through its N-terminal domain and interacts with both DnaA and the β clamp through its C-terminal domain (Felicori et al., 2016; Noirot-Gros et al., 2006). Although YabA and E. coli Hda share the capacity to interact with both DnaA and DnaN, YabA is distinct from Hda in that it has no homology with DnaA and does not contain a consensus motif for binding to DnaN (Kurz et al., 2004). This indicates that YabA lacks the key structural elements of Hda required to promote DnaA-ATP hydrolysis, as occurs in the RIDA system (Su’etsugu et al.,

Construction and Maintenance of a Replication Fork |  15

2005). Thus, although YabA regulates initiation through a novel mechanism, the involvement of interactions with DnaA and DnaN suggests a conserved coupling between initiation and elongation of replication. Live cell fluorescence microscopy has shown that YabA localizes at the replication factory during most of the cell cycle through its interaction with DnaN (Hayashi et al., 2005; Noirot-Gros et al., 2006). Moreover, YabA was shown to confine GFPDnaA at the replisome during most of the cell cycle (Soufo et al., 2008), leading to the model that YabA prevents untimely reinitiation by mediating spatial sequestration of DnaA away from oriC (NoirotGros et al., 2006; Soufo et al., 2008). However, the sequestration model predicts that YabA would titrate DnaA away from other chromosomal loci known to bind DnaA, yet global gene expression in a yabA null mutant revealed that the expression of genes controlled by DnaA was not altered (Goranov et al., 2009) and the absence of YabA did not increase the binding efficiency of DnaA at DnaA-boxes throughout the chromosome (Cho et al., 2008). These results suggest that the effect of YabA on DnaA activity is somehow localized to oriC. Mutagenesis indicates that YabA interacts with the AAA+ oligomerizaton domain of DnaA (Cho et al., 2008). Moreover, it has been found that YabA inhibits DnaA filament formation, which could explain this oriC-specific regulation (Scholefield and Murray, 2013). Soj is a dynamic Walker-type ATPase that is directly regulated by the DNA-binding protein Spo0J (Soj and Spo0J are members of the ParA and ParB family of partition proteins). Soj oligomerization state is regulated by ATP (Leonard et al., 2005). Monomeric Soj (Soj-ADP) has been found to physically interact with domain IIIb of DnaA and to inhibit DNA replication initiation by blocking DnaA filament formation (Murray and Errington, 2008; Scholefield et al., 2012). In the absence of Spo0J, or when expressed as a mutated form deficient for ATP hydrolysis (SojD40A), SojATP dimerizes and DnaA-dependent initiation is stimulated (Lee et al., 2003; Murray and Errington, 2008; Ogura et al., 2003). It has been shown that Spo0J stimulates the ATPase activity of Soj, driving it from a dimer to a monomer and triggering a switch from an activator to an inhibitor of DNA replication initiation (Scholefield et al., 2011).

However, it remains unknown when this switch is activated during the B. subtilis cell cycle. Genome-wide chromatin immunoprecipitation assays have revealed that DnaA is significantly enriched at six regions on the chromosome that contain clusters of DnaA-box sequences (Breier and Grossman, 2009; Ishikawa et al., 2007). A deletion mutant lacking these six DnaA–box clusters (DBC) overinitiated DNA replication (Okumura et al., 2012), similar to the datA locus in E. coli. Reintroduction of a single DBC at an ectopic position near oriC, or reintroduction of multiple DBCs at an ectopic position near terC, suppressed the overinitiation phenotype, suggesting that a critical dosage of DBCs are required to regulate DnaA activity. However, a multicopy plasmid harbouring a DBC did not complement the mutant phenotype, indicating that the DBC acts in cis. The intriguing mechanism underlying DBC activity is yet to be determined. Replication restart after adventitious arrest In B. subtilis and in E. coli the genome is a large circular molecule of over 4 million basepairs and because of their single replication origin each fork must travel more than 2 Mbp to achieve the complete duplication of the chromosome. Consequently, any accidental arrest suffered by one of the forks represents a threat to cell cycle completion, and thus cellular pathways have evolved to facilitate the restart of arrested forks (Michel et al., 2004). Intensive studies conducted in E. coli have uncovered multiple responses to overcome the different types of fork arrest that can be provoked by a variety of events (DNA damage, DNA-bound proteins, and higher order DNA structures). These responses range from simple pausing in the case of a protein block, to induction of appropriate repair pathways followed by re-activation of the fork and resumption of chromosome replication. Replication fork re-activation most often relies on the re-assembly of the replisome on a processed arrested fork. This event, termed ‘replication restart’ (RR), is directed by a specific set of RR proteins (McGlynn and Lloyd, 2002; Sandler and Marians, 2000). The pathways of replication restart In E. coli, the RR proteins PriA, PriB, PriC and DnaT drive origin-independent initiation at sites

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of collapsed replication forks (Table 1.1) (Marians, 2000). PriA plays a central role in replication fork reactivation by triggering the serial assembly of the downstream RR factors into a multiprotein complex at the stalled fork. This RR prepriming complex recruits the DnaB replicative helicase in association with its loader DnaC to promote the subsequent assembly of the replisome. There are three pathways for restart in vivo based on the composition of the prepriming complex: PriA–PriB–DnaT, PriA–PriC, and PriC–Rep. The different complexes recognize structurally distinct forked DNA molecules (Heller and Marians, 2005a). The assembly of the replisome onto forked DNA molecules mediated by RR proteins parallels the pathway mediated by DnaA at oriC, where the replicative helicase is the first partner to be recruited and loaded onto the lagging strand template at the forefront of the replication fork (Marians, 2000). However, whereas DnaA binds specifically to repeated DNA motifs within the origin, RR proteins recognize specific forked DNA substrates. In the cell such structures originate from randomly arrested replication forks that can be processed by various DNA repair pathways and from DNA recombination intermediates (Fig. 2.2) (Michel et al., 2004). Thus, in contrast to DNA replication initiation at oriC, the RR apparatus has to deal with many different types of forked DNA molecules. A key distinction between the various forked DNA substrates is whether or not the leading and lagging strand arms of the fork are single-stranded. PriA is a DNA helicase belonging to the large superfamily 2 (SF2) (Gorbalenya, 1993). PriA preferentially unwinds forked DNA substrates with a double-stranded leading-strand template to produce the ssDNA needed for delivery of the replicative helicase, a fork remodelling activity that is crucial in the PriA and PriC pathways (Heller and Marians, 2005b). A common characteristic between the PriAdependent RR systems of E. coli and B. subtilis is that PriA is not essential for growth. However, E. coli and B. subtilis priA null mutant cells are poorly viable and exhibit severe defects that can all be explained by their failure to efficiently restart arrested forks. Indeed, the defects of priA null mutants can be almost fully suppressed by mutations that enhance the loading apparatus of the replicative helicase, DnaC in E. coli and DnaB in B. subtilis (Bruand et al., 2001; Sandler et al., 1996). This indicates that

PriA controls the major RR pathway in wild-type cells in both bacteria. In E. coli, the combination of priA and priC null mutations is lethal, providing strong evidence that RR is essential for cell survival (Sandler, 2000). In B. subtilis, there is no PriC homologue encoded in the genome, and only the PriA-dependent RR pathway is well characterized. However, genetic evidence supports the notion that DnaD, which is a direct partner of PriA, promotes RR in the absence of PriA (Bruand et al., 2001). The presence of redundant RR pathways not only provides a ‘backup’ system in case the initial system fails, but also facilitates the targeting and efficient processing of the distinct forked DNA structures that can be produced upon replication arrest (Heller and Marians, 2005a,b, 2006). PriA-dependent replication restart With the exception of PriA, the B. subtilis RR proteins are distinct from those of E. coli (Table 1.1). Notably, B. subtilis PriA and DnaA use the same essential DnaD, DnaB and DnaI proteins to recruit and load the DnaC replicative helicase onto ssDNA (Fig. 1.2A) (Bruand and Ehrlich, 1995; Bruand et al., 1995, 2001). B. subtilis PriA is functionally similar to its E. coli counterpart, as it is a forked DNA-binding helicase that acts as the primary RR protein in vivo (Marsin et al., 2001; Masai et al., 1999; Polard et al., 2002). However, B. subtilis PriA cannot complement an E. coli priA mutant (Masai et al., 1999; Polard et al., 2002). B. subtilis PriA initiates a cascade of events starting with the recognition of arrested forks in the chromosome, the sequential recruitment of DnaD, DnaB, DnaI and finally the functional loading of the DnaC helicase onto ssDNA (Fig. 1.2) (Bruand et al., 2005; Marsin et al., 2001; Velten et al., 2003). A remarkable feature of the DnaD dimer and DnaB tetramer is that both are non-specific DNA binding proteins (Marsin et al., 2001; Zhang et al., 2005). Compelling evidence indicates that the essential activities of DnaD and DnaB during both chromosomal replication initiation and RR rely on their distinct DNA binding activities at loci targeted by DnaA and PriA. The association of DnaD and DnaB with chromosomal regions bound by DnaA, including oriC, was confirmed by chromatin immunoprecipitation (ChIP) (Smits et al., 2011). Furthermore, a genome-wide analysis of DnaD and DnaB binding sites by ChIPchip indicated that the helicase loader proteins can

Construction and Maintenance of a Replication Fork |  17

be associated with the highly transcribed regions of rRNA (rrn) genes. The association of DnaD, DnaB, and DnaC with rrn loci depends on transcription and on the replication restart protein PriA (Merrikh et al., 2011), strongly suggesting that DnaD and DnaB are acting to reload the helicase for replication restart at forks stalled upon encountering RNA polymerases. DnaD and DnaB proteins have an architectural role on DNA that facilitates the co-ordination, and perhaps the licensing, of DnaI-dependent loading of DnaC onto ssDNA (Bruand et al., 2005; Rokop et al., 2004). The affinity of DnaB for ssDNA is enhanced upon its interaction with DnaD (Marsin et al., 2001) and the dnaB75 mutation suppresses the thermosensitivity of the dnaD23 mutant strain (Bruand et al., 2005; Rokop et al., 2004), suggesting that these proteins act in combination with one another. Consistent with this notion, the DnaD23 protein displays a diminished and thermosensitive ssDNA binding activity, whereas the DnaB75 mutant protein exhibits a higher affinity for ssDNA than the wild-type protein (Bruand et al., 2005). Moreover, while neither DnaD nor DnaB nor DnaB75 alone can interact with ssDNA coated by SSB, which is known to create a physical barrier to the loading of the helicase in the E. coli system (see above) (Marians, 2000), together DnaD and DnaB proteins can associate with SSB-coated ssDNA and this binding is more efficient in the presence of the DnaB75 derivative (Bruand et al., 2005). In addition, DnaB promotes DnaI-dependent helicase loading and again this stimulation is more efficient for the DnaB75 protein (Velten et al., 2003). Thus, the concerted actions of DnaD and DnaB likely provide a way to overcome the SSB barrier and facilitate the subsequent DnaI-dependent loading of the helicase on ssDNA (Bruand et al., 2005). This effect could account for the observation that the DnaD-, DnaB-, DnaI- and DnaC-dependent replication of a ssDNA circular template produced by a rolling circle plasmid is more effective in B. subtilis cells expressing the mutant DnaB75 protein than the wild type DnaB protein (Bruand et al., 2001). Finally, DnaD and DnaB can interact in a distinctive manner with supercoiled DNA in vitro, producing open and compacted DNA structures, respectively (Carneiro et al., 2006; Zhang et al., 2005). These opposite DNA-remodelling

activities of DnaD and DnaB could modulate local chromosome architecture to facilitate the DnaA- and PriA-mediated initiation events. SSB as an organizer of the replication fork maintenance complex As stated above, SSB plays a pivotal role in the recruitment of accessory proteins to active forks. The E. coli SSB interactome is currently estimated to comprise 14 proteins involved in various DNA pathways. Parallels can be drawn with B. subtilis where SSB is described as a protein-hub able to target 12 known players at active replication forks (Costes et al., 2010). In addition to PriA, RecG and RecQ helicases, SSB also interacts with the RecQhomologue RecS in complex with YpbB and with the putative helicase/nuclease, YrrC. The need for PriA to interact with SSB in order to act efficiently in fork restart has been demonstrated (Lecointe et al., 2007). Similarly, a physical interaction of E. coli RecG with SSB was shown to stabilize its binding to various DNA substrates mimicking forked DNA structures that are produced upon replication arrest (Buss et al., 2008). Two other exonucleases, RecJ and XseA (ExoVII subunit) were found localized by SSB at chromosomal forks in B. subtilis. In E. coli, the SSB partners RecQ and RecJ were found to cooperate at stalled replication forks, degrading nascent DNA prior to resumption of replication (Courcelle and Hanawalt, 1999; Shereda et al., 2007). The propensity of SSB proteins to recruit various DNA helicases and exonucleases correlates with the requirement for DNA unwinding and degradation functions to remodel a disassembled fork. Similarly, SSB’s interactions with RarA and RecO, two proteins involved in the loading of RecA (the central player in homologous recombination; see Chapter 2) at arrested forks in E. coli (Lestini and Michel, 2007), also illustrate the need for RecAmediated fork rescue in recovering replication after accidental arrest. Finally, the physical interaction of SSB with the essential polymerase DnaE, recently shown to extend RNA primers laid down by DnaG at forks (Sanders et al., 2010), suggests that DnaE may also fulfil additional tasks in specialized repair pathways. This dual role of DnaE as a DNA polymerase during replication initiated at oriC and in replication restart at arrested forks is intriguing and remains to be investigated.

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Translesion DNA synthesis Living cells are constantly exposed to endogenous and exogenous DNA damaging agents and they cope with DNA lesions by employing various DNA repair pathways. In detrimental conditions E. coli and B. subtilis cells activate a co-ordinated cellular response, termed SOS, which controls the expression of several proteins involved in DNA repair, recombination and mutagenesis (see Chapter 2) (Becherel and Fuchs, 1999; Courcelle et al., 2001; Friedberg, 1995). Although efficient repair processes act to remove most of the lesions from DNA in an error-free manner, the persistence of unrepaired damage may result in arrest of the DNA replication fork as replicative DNA polymerases are often unable to incorporate nucleotides opposite non-instructive damaged bases. The Y-family DNA polymerases In SOS response-induced E. coli and B. subtilis cells, mutagenesis at both damaged and undamaged sites on the DNA template is increased (Becherel and Fuchs, 1999; Duigou et al., 2004; Friedberg, 1995). This results from translesion DNA synthesis (TLS), a process mediated by specialized DNA polymerases that are capable of synthesis through blocking DNA lesions (Friedberg et al., 2005; Prakash et al., 2005). Lesion bypass is often errorprone, thus producing mutations in the nascent DNA strand opposite the damaged site. The vast majority of TLS enzymes belong to the Y-family of DNA polymerases, which is widely spread among living organisms. Rev1p and Pol η in yeast, or Pol IV and Pol V in E. coli, are prototypical members of this family (Ohmori et al., 2001). The Y-polymerases display low fidelity DNA synthesis, low processivity, and lack 3′–5′ proofreading exonuclease activity (Kokoska et al., 2002; Kunkel and Bebenek, 2000). These enzymes adopt the ‘right hand’ structure typical of DNA polymerases, albeit with a more flexible active site that can accommodate various distorting lesions (Friedberg et al., 2001; Ling et al., 2001). Depending on the context and nature of the damaged site, bypass involves the recruitment of one or more specialized DNA polymerases (Friedberg et al., 2005). B. subtilis possesses two Y-family members, PolY1 and PolY2, encoded by the yqjH and yqjW genes, respectively (Duigou et al., 2004; Ohmori et al., 2001). When overproduced PolY1 promotes

untargeted mutagenesis, a process that requires its binding to the β-clamp (Duigou et al., 2004). Furthermore, PolY1 acts in adaptive mutagenesis (Sung et al., 2003). These characteristics are reminiscent of E. coli Pol IV, encoded by dinB, which confers a mutator phenotype when overproduced (Kim et al., 2001; Wagner et al., 1999; Wagner and Nohmi, 2000) and also requires interaction with the β-sliding clamp (Becherel et al., 2002; Lenne-Samuel et al., 2002). The second B. subtilis Y-polymerase, PolY2, is responsible for most UVinduced mutagenesis in B. subtilis, and like PolY1 also promotes untargeted mutagenesis when overproduced (Duigou et al., 2004). PolY2-mediated mutagenesis on undamaged DNA requires the presence of RecA in addition to interaction with the β-clamp (Duigou et al., 2004, 2005). Based on these properties, PolY2 appears to be the functional homologue of E. coli Pol V. However, whereas Pol V exists as a heterotrimeric complex termed UmuCD2, B. subtilis PolY2 likely acts as a single protein (no umuD homologue can be detected in the B. subtilis genome) to carry out TLS across UVinduced lesions. Evidence also suggests that both PolY1 and PolY2 may participate in TLS during sporulation, enhancing the survival of sporulating cells exposed to genotoxic agents (Rivas-Castillo et al., 2010). Considering the mutagenic potential of Y-family polymerases, clearly their intracellular levels must be tightly controlled. In E. coli Pol IV is constitutively expressed at a low level, Pol V is strongly repressed under normal growth conditions, and both polymerases are induced during the SOS response (Courcelle et al., 2001). In M. tuberculosis the two Y-family polymerases DinP and DinX are expressed constitutively and are not up-regulated after DNA damage (Boshoff et al., 2003; Brooks et al., 2001). In B. subtilis PolY1 is expressed at a low constitutive level and does not belong to the SOS regulon, suggesting that it may act during normal DNA replication, whereas PolY2 is expressed only upon SOS induction (Duigou et al., 2004). It is conceivable that PolY1 may be up-regulated under specific environmental stress or at a specific stage of development, consistent with its involvement in adaptive mutagenesis (Sung et al., 2003). Thus, it appears that different bacteria have evolved different strategies to regulate the expression of their Y-family polymerases, which might reflect

Construction and Maintenance of a Replication Fork |  19

pathway for TLS in B. subtilis is a dual polymerase mechanism involving PolY1 or PolY2 together with DNA polymerase I (Duigou et al., 2005). Interestingly, TLS in eukaryotes is often promoted by the concerted action of two polymerases, one acting as an ‘inserter’ at the damaged site and the other one as an ‘extender’. However, such a TLS mechanism is not present in E. coli (a comparison of the E. coli and B. subtilis Y-family polymerases properties is summarized in Table 1.3). Although the bypass of some bulky lesions in particular DNA contexts may require the concerted action of two DNA polymerases in E. coli (Wagner et al., 2002), the major pathway is a one-enzyme process carried out by Pol V in the presence of RecA (Fujii and Fuchs, 2004; Pham et al., 2002; Tang et al., 2000). Pol IVmediated mutagenesis is similarly independent of both Pol I and Pol II (Wagner and Nohmi, 2000). Alignment of the B. subtilis Pol I sequence with orthologues from various eubacteria reveals that it belongs to a Gram-positive subfamily of A-polymerases. Polymerases of this subfamily have a vestigial 3′–5′ exonuclease active site which lacks the residues involved in co-ordination of the divalent cations required for exonuclease activity (Duigou et al., 2005). Indeed it has been found that B. subtilis Pol I is deficient for exonuclease activity (Kiefer et al., 1997), which is consistent with its role in TLS (i.e. – proofreading activity would abrogate

adaptation to their own environmental growth conditions to minimize the detrimental effect of DNA damage and increase their fitness and survival. B. subtilis Pol I assists translesion synthesis catalysed by Y-family polymerases Surprisingly, the vast majority (> 90%) of UVinduced mutagenesis observed in B. subtilis upon PolY2 overproduction depends on the presence of catalytically proficient DNA polymerase I (Duigou et al., 2005). It was found that Pol I directly interacts with PolY2 in a yeast two-hybrid assay, suggesting that Pol I may assist PolY2 in TLS across UV lesions. The remaining ( 75%) of the cells, and GFPSsbA or SsbA-YFP formed a variable number of foci (from 1 to more than 5) (Meile et al., 2006; Jers et al., 2010). RecN does not co-localize with the replisome The essential SsbA protein is the key factor that links active chromosomal replication forks with DNA recombination functions (Lecointe et al., 2007; Costes et al., 2010). SsbA co-localized with PriA, RecG, RecJ, RecQ, RecS-YpbB, RecO, SbcC, RecD2, DnaE and RarA proteins in vivo (Lecointe et al., 2007; Costes et al., 2010). On the other hand, RecN neither interacts (Sanchez et al., 2008) nor competes with SsbA bound to ssDNA. This is consistent with the observation that the single RecN-YFP focus was often present close to the mid-cell, but rarely co-localized with the replisome, as judged by DnaX-CFP localization (Kidane and Graumann, 2005a). Damage-induced RecN foci and replisome foci respond differently to increased MMC dose. Indeed, when exponentially growing cells were treated with a MMC dose that impairs growth of ~50% of the cells, ~55% of the cells showed clear RecN–YFP foci, and ~60% of the cells formed one single DnaX-CFP focus at midcell. The DnaX foci at quarter-cell position were not observed, indicating that the replisome does not disassemble in a high proportion of cells, and that the nucleoids fuse (Kidane et al., 2004; Mascarenhas et al., 2006). At a MMC dose which impairs growth of ~75% of the cells, ~70% of the cells showed clear RecN–YFP

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foci and ~40% of the cells contained two or more DnaX-CFP foci, rather than one (45%), and ~15% of the cells contained no signal (Kidane et al., 2004; Mascarenhas et al., 2006). However, when a lethal dose of MMC, which impairs growth of ~99% of the cells was used, ~72% of the cells showed clear RecN–YFP foci, but the DnaX-CFP foci were no longer apparent in most cells (Mascarenhas et al., 2006). At stalled replication forks, caused by shifting dnaA or dnaB thermosensitive mutants to non-permissive temperature, an extensive reorganization of the nucleoids was observed. When DNA replication is not impeded, ~15% of total cells contained a discrete RecN-YFP or YFP-RecA focus per nucleoid (Mascarenhas et al., 2006; Krishnamurthy et al., 2010; Simmons et al., 2007). However, when the fork was halted by inactivation of DnaA or DnaB (temperature shift up) and additional treatment with MMC, discrete RecN foci were observed in ~75% of the cells (Krishnamurthy et al., 2010), but RecA foci were present in only ~15% of total cells 30 min after MMC addition (Simmons et al., 2007). One possibility to reconcile these apparent contradictions is that when DNA replication is stalled and DNA damage is generated two different DNA repair events, DSG and one-ended DSB, overlap. It is likely that (i) a block in replication generated by the inactivation of DnaA or DnaB would lead to DSG repair, but a fraction of one-ended DSBs to be recognized by both RecN and RecA in ~15% of the cells (Simmons et al., 2007); and (ii) inactivation of DNA replication and addition of low or moderate MMC doses do not increase the number of RecA foci, but RecN foci formation accumulates in ~75% of the cells within the first 30 min, with DNA damage-induced RecA foci and threads forming later (30 to 60 min; Fig. 2.3, stage d) (Kidane et al., 2004; Simmons et al., 2007). At collapsed replication forks generated by addition of a MMC lethal dose that impairs growth of ~99% of the cells, the RecN focus does not seem to localize with the replication forks, and the DnaX-CFP focus is no longer present in most cells (Kidane et al., 2004; Kidane and Graumann, 2005a; Mascarenhas et al., 2006). These experiments show that cells still form a RC, while the replisome disintegrates. Similarly, after depletion of the replication initiator protein DnaA, the foci of DnaX-GFP, GFP-DnaN and GFP-SsbA, all of which co-localize,

become dispersed in the cytoplasm (Berkmen and Grossman, 2006; Meile et al., 2006). DNA end-processing The termini of one-ended or two-ended DSBs must be resected to leave a 3′-terminated ssDNA to which RecA can polymerize (Figs. 2.2 and 2.4). This end resection process occurs in two steps, basal and long-range resection. In the first step PNPase, which co-purified with RecN in B. subtilis or with RecA in E. coli, is required for the formation of RecN-promoted discrete RC upon DNA DSBs induction, where it provides the RecN-associated 3′ → 5′ ssDNA exonucleolytic activity (Cárdenas et al., 2009, 2011). Unlike E. coli, which has different 3′-ssDNA exonucleases (e.g. ExoI, ExoVII, ExoIX, ExoX and ExoXI), B. subtilis may lack genes encoding such activity (Cárdenas et al., 2009). PNPase, or another uncharacterized exonuclease, may catalyse a limited end resection (basal resection) to generate the proper substrate for AddAB or RecJ long-range end processing or to remove any ‘dirty’ 3′-end (Cárdenas et al., 2009, 2011). Alternatively, PNPase-mediated DNA polymerization is modulated in vitro by RecN, RecA, and SsbA (Cárdenas et al., 2011). The pnpA/pnp phenotype is epistatic to addAB/recB, and ruvAB/ruvA both in B. subtilis and E. coli cells (Cárdenas et al., 2009; Rath et al., 2012). The basal resection processing is followed by a processive long-range end resection. The functions involved in long-range end resection vary among different bacteria. In E. coli, ~99% of the recombination events occurring at DSBs require the trimeric RecBCD complex (functional counterpart of Firmicutes AddAB or Actinobacteria AdnAB) for the resection of the broken ends. In the recBC sbcA or recBC sbcB sbcCD context end-processing requires the ‘activation’ of RecF pathway to promote end resection (Kowalczykowski et al., 1994; Spies et al., 2005; Dillingham and Kowalczykowski, 2008; Yeeles and Dillingham, 2010; Carrasco et al., 2014). Here, resection by the activated RecF pathway requires the 5′ → 3′ exonuclease, RecJ, the 3 → 5′ RecQ helicase and the SSB protein (counterpart of B. subtilis SsbA) (Handa et al., 2009). Among the > 820 genomes examined of free-living bacteria > 85% contained a recJ gene, and ~80% of these genomes contained recN and recQ (or recS) genes. However, < 50% of those genomes examined contained the recBCD (addAB or adnAB) genes, suggesting that

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Figure 2.4  One-ended DSB repair by HR. A collapsed fork, during DNA replication, generates a one-ended DSB that can be repaired by HR. (a) RecN, in concert with PNPase, recognizes and processes the damaged ends. If the ends already have ssDNA regions, SsbA might bind to them. In the absence of RecN, SbcEF forms DNA damage-induced foci. (b) The 5′-ends are resected either by the AddAB complex or by RecJ-RecQ-like (RecQ or RecS)-SsbA. RecN tethers the DNA ends and promotes loading of the RecA mediators. (c) The RecO(RecR)– SsbA complex, at a RecN RC, could promote the loading of RecA onto the ssDNA. In vitro RecO-SsbA alone promote RecA polymerization onto SsbA-coated ssDNA. The RecA nucleoprotein filaments (RecA threads) promote the search for a homologous template and strand invasion. RecA modulators (RecF, RecX, PcrA, RecU, etc.) control RecA by modulation of RecA filament extension. (d) The invading strand provides the template for primosome loading. The primosome assembly machinery then loads the replisome. (e–g) D-loop dissociation and in concerted action with DNA synthesis generates a nicked HJ or an extended D-loop. Then, a disruptase (e.g. MutS2) might generate an active NCO. (h) Reversal (also termed regression) of the D-loop by RuvAB or RecG converts the D-loop into a HJ. (i–j) Once the RecU recognition site is exposed, RecU, aided by RuvAB or RecG, resolves the recombination intermediate. The resolution of the HJ might generate CO or NCO products (Fig. 2.2), The undesired dimers (CO products) are converted into monomers by the tyrosine site-specific recombinase in concert with a translocase (Fig. 2.2).

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this E. coli long-range end resection pathway is not broadly conserved. In B. subtilis, long-range end resection involves redundant avenues, which work with similar efficiency; the AddAB complex and the ssDNA exonuclease (RecJ) that works in concert with a RecQ-like DNA helicase (RecQ and RecS [no counterpart in E. coli]) and SsbA (Fig. 2.3, stage c, and Table 2.1) (Fernández et al., 1998; Sanchez et al., 2006). Indeed, deletion of the recJ gene causes substantial defects to a wide spectrum of DNAdamaging agents, and the addA or addB genes cause substantial defects in DSB repair (Alonso et al., 1988, 1993; Sanchez et al., 2006). The ∆recQ or ∆recS mutations are epistatic with ∆recJ, but they show additive sensitivity to DNA damaging agents in the addAB context. Finally, ∆recJ in combination with ∆addAB shows a synergistic loss of survival, to sensitivity levels similar to that of ∆recA cells (Sanchez et al., 2006). Analysis of RecN-YFP localization in the absence of AddA and RecJ showed that RecN-YFP foci are still formed in response to DNA damage, yet RecN-YFP forms multiple small foci rather than a single large focus (Sanchez et al., 2006), for as yet unknown reasons. The AddAB enzyme comprises a single helicase motor and two exonuclease domains that are dedicated to the cleavage of each of the unwound DNA strands (‘destructive mode’) (Carrasco et al., 2014; Yeeles and Dillingham, 2010). AddAB, as well as the RecBCDEco complex, undergoes an allosteric conformational change (‘transient stalled mode’) after binding to the canonical five-(5′-AGCGG-3′) and eight-base (5′-GCTGGTGG-3′) χ sequences, respectively (Yeeles et al., 2011; Carrasco et al., 2013, Gilhooly and Dillingham, 2014, Yang et al., 2012). Upon binding to the χ sequence the AddAB complex pauses, decreases the rate of translocation and only the 3′-terminated strand is sequestered (Carrasco et al., 2013; Gilhooly and Dillingham, 2014). The χ-terminated end escapes degradation by the AddA exonuclease domain, thereby the 3′-terminated strand loops upstream the χ sequence (Saikrishnan et al., 2012). The AddAB complex continues unwinding several kilobases beyond the χ sequence, and degrades the 5′-terminated strand (‘recombinogenic mode’) (Carrasco et al., 2014; Dillingham and Kowalczykowski, 2008; Yeeles and Dillingham, 2010). This end resection avenue, which does not seem to play a role in DSG repair, generates a duplex molecule with a 3′-ssDNA tail

terminating with a χ sequence (Carrasco et al., 2014; Dillingham and Kowalczykowski, 2008; Yeeles and Dillingham, 2010). In vivo, functional AddA-GFP or AddB-YFP localizes throughout the B. subtilis cytosol, and both functions fail to organize into discrete foci upon MMC addition (Mascarenhas et al., 2006). In a reconstituted reaction with E. coli proteins, RecJ in concert with SSB was shown to generate sufficient ssDNA to promote RecA-catalysed strand invasion, although the extent of degradation and joint molecule formation was less than that observed in the presence of RecQ (Handa et al., 2009). The B. subtilis RecJ 5′ → 3′ ssDNA exonuclease, in concert with any of the RecQ-like DNA helicases (RecQ or RecS) that unwind duplex DNA with 3′-overhangs, and with SsbA that covers generated ssDNA streches, degrades the 5′-terminated strand, resulting in a 3′-ssDNA overhang. SsbA bound to ssDNA further stimulates the helicase and exonuclease activities by a direct protein–protein interaction (Tables 2.1 and 2.2). Genetic and cytological studies have revealed that (i) RecN, in concert with PNPase, appear to organize the assembly of a single network of protein–protein; (ii) the AddAB and RecJ nucleases play a redundant role in vivo; (iii) AddAB and/or RecJ are not necessary for the formation of RecN repair foci, but for the generation of one discrete RecN RC; (iv) concomitant with end-processing, RecN binds to the ssDNA tail of the duplex molecule at the DSB, protects the 3′-OH end, and facilitates the tethering of these DNA ends together to form mainly one discrete focus or RC; and (v) SsbA binds to the ssDNA without displacing RecN from the ssDNA end (Kidane et al., 2004; Kidane and Graumann, 2005a; Sanchez and Alonso, 2005). It is likely therefore, that end processing takes place between the damage recognition by the RecN protein and RecA loading (Figs. 2.2 and 2.4). In eukaryotes, HR is generally restricted to the S and G2 phases of the cell cycle, when a sister chromatid or a homologous chromosome is available as a repair template, whereas NHEJ is the predominant pathway outside of S phase. The choice of pathways is to a significant extent not stochastic but a function of the cell cycle (Ferretti et al. 2013). Reduced resection in G1 results from low CDK (Cdc28) activity and Ku binding to DNA ends. Indeed, elimination of Ku or Dnl4 or

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activation of Cdc28 partially restores end-resection initiation in G1-phase cells, but extensive resection is still partially defective (Clerici et al., 2008). Furthermore inhibition of Cdc28 in G2 cells, blocks end-resection (Clerici et al., 2008). In contrast, in B. subtilis HR is the predominant pathway in the presence of an intact DNA template. In exponentially growing cells, commitment to HR is still a poorly understood process and its physiological importance remains to be addressed. Hence we favour a scenario where a highly complex multistep repair by HR response is finely orchestrated towards different error-free recombinational repair avenues by multiple redundant functions. In the absence of an homologous template or when there is a defect in the early steps of HR, two-ended DSBs might be processed by NHEJ (Alonso et al., 2013, Schiller et al., 2014). Previously, it was shown that (i) the Mycobacterium Ku protein protects DNA ends from resection by the AdnAB helicase–nuclease complex (Sinha et al., 2009), and (ii) B. subtilis ku is not epistatic with addAB, recA, recO or recN, but that ku, pnpA and sbcC are epistatic to one another (Weller and Doherty, 2001; Mascarenhas et al., 2006; Cárdenas et al., 2009; Vlasic et al., 2014). These experiments show that NHEJ may serve as a backup system for two-ended DSB repair in stationary phase cells or in exponentially growing cells with a defect in the early stages of HR, and suggest that SbcC and PNPase might also play a role in NHEJ (Mascarenhas et al., 2006). In B. subtilis, sbcC is also epistatic with recA (Mascarenhas et al., 2006), and both SbcC and SbcE proteins may play a more active role in the absence of RecN (Mascarenhas et al., 2006; Krishnamurthy et al., 2010). RecA loading The mechanism by which the recombinase is delivered to the 3′-tailed duplex is of universal importance. In E. coli cells, the RecBCD complex, through a RecB·RecA interaction, directly loads RecA onto newly generated 3′-ended naked ssDNA during continuous unwinding (Anderson and Kowalczykowski, 1997, Anderson et al., 1999, Spies et al., 2005; Spies and Kowalczykowski, 2006). Then, RecAEco forms a pre-synaptic filament that efficiently displaces SSBEco bound to other regions of ssDNA (Spies et al., 2005; Dillingham and Kowalczykowski, 2008). This species-specific RecA

loading mechanism might be conserved among bacteria of the Proteobacteria phylum. Indeed, it was necessary to co-express recBCD and recA genes of Serratia marcescens, Proteus mirabilis or Helicobacter pylori to suppress the DNA repair defect in recBCDEco cells (Rinken et al., 1991; Amundsen et al., 2009). However, the expression of only B. subtilis AddAB or the Streptomyces ambofaciens AdnAB complex was sufficient to partially restore the recombination deficiency of the distantly related E. coli ∆recB or recBCD strain (Kooistra et al., 1993, Zhang et al., 2014), suggesting that a species-specific RecA-loading mechanism was not required. Alternatively, the AddAB or AdnAB enzymes do not contribute to RecA loading onto naked ssDNA (in E. coli, RecBCD does so). Biochemical data consistent with the later hypothesis have shown that B. subtilis AddAB cannot load and activate its cognate RecA·ATP to polymerize onto SsbA-coated ssDNA or to catalyse DNA three strand exchange (Carrasco et al., 2015). This is consistent with the idea that (i) both end-resection pathways (AddAB or RecJ-RecQ/S-SsbA) require RecO for recombination (Carrasco et al., 2015); and (ii) AddAB translocation is not coupled to DNA unwinding in the absence of SsbA, because nascent ssDNA immediately re-anneals behind the moving enzyme (Yeeles et al., 2011). In E. coli the activated RecF pathway requires RecOR or the RecORF complex to load RecA onto SSB-coated SSGs, and RecA nucleoprotein filament growth displaces the SSB protein to catalyse DNA recombination (Menetski and Kowalczykowski, 1989, Umezu and Kolodner, 1994; Hobbs et al., 2007; Sakai and Cox, 2009; Handa et al., 2009; Morimatsu et al., 2012). RecAEco can catalyse recombination and only requires mediators to stimulate DNA strand exchange. In contrast, in the ATP bound form, RecA from bacteria that can develop natural competence (see Chapter 13), cannot nucleate or polymerize onto SsbA-coated ssDNA in vitro and they are unable to promote strand exchange to form heteroduplex DNA in vitro without additional proteins (Lovett and Roberts, 1985, Steffen and Bryant, 1999, Carrasco et al., 2015). Accessory factors can activate RecAmediated DNA strand exchange. These factors can be divided into two broad classes: those that act before homology search by promoting assembly of recombinase filaments (mediators), and those that

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act during homology search and strand exchange (modulators). Not all the RecA mediators might be conserved among free-living bacteria (Fig. 2.3 stage d). Indeed, > 99% of the > 820 genomes of free-living bacteria examined contained recA and ssbA/ssb genes, > 95% contained recO and > 85% contained recR. However, only  820 genomes examined contained the ruvB gene, and > 95% contained ruvA, ruvC/recU and recG genes (Garcia-Gonzalez et al., 2013), suggesting that branch migration and HJ resolution is a well-conserved mechanism to resolve HJ intermediates. Bacteria of the Firmicutes phylum contain homologues of RuvA, RuvB and RecU, but not of RuvC (Ayora et al., 2004). The RecU HJ-specific endonuclease, which is structurally and biochemically unrelated to RuvC, resolves HJs (Ayora et al., 2004; McGregor et al., 2005). A B. subtilis recU-, recG- or ruvAB-deficient strain is very sensitive to DNA damaging agents, and the ruvAB mutations are synthetically lethal with recU or recG deletions (Sanchez et al., 2005, 2007b). Cytologic experiments with a functional RecU– GFP fusion showed that RecU forms discrete foci, late in the process of DSB repair (~120 min after addition of MMC) and it co-localizes with RecN at the RC, which confirms that the repair of DSBs is a sequential process (Fig. 2.3 and Table 2.3) (Sanchez et al., 2005). The RuvAB branch migration translocase is required for such step because RecU foci are not observed in ∆ruvAB cells (Sanchez et al., 2005, Mascarenhas et al., 2006, Sanchez and Alonso, 2005, Carrasco et al., 2009). The absence of RuvAB does not completely disable HR, suggesting that RecG catalyses branch migration along with RecU cleaving these recombination intermediates (Cañas et al., 2014; Sanchez et al., 2007b). RecU structure shows certain degree of similarity with type II restriction endonucleases of the EcoRII/PvuII superfamily and HJ endonucleases (Hjc or Hje) from Archaea (McGregor et al., 2005). The overall RecU fold has been described as a ‘mushroom’ with a cap and a stalk region. The cap contains the catalytic residues important for

HJ cleavage and RuvB interaction (Cañas et al., 2008, 2014; Carrasco et al., 2009). The stalk region is important for HJ binding and interaction with RecA (Cañas et al., 2008, 2011). In vitro, RuvA binds HJs and recruits the RuvB translocase to catalyse branch migration (Cañas et al., 2014; Suzuki et al., 2014). Concomitantly, RecU bound to HJ DNA, by a direct protein–protein interaction, recruits RuvB onto the RecU-HJ DNA complex and ultimately cleaves the HJs (Ayora et al., 2004; Carrasco et al., 2005; Cañas et al., 2014; Suzuki et al., 2014). The RuvAB translocase moves the junction until a RecU cognate site is exposed and RecU promotes HJ cleavage (Cañas et al., 2014). Alternatively, as shown for E. coli proteins, RecG and RuvAB catalyse replication fork regression, but RuvC preferentially cleaves the HJs formed by RecG at stalled forks (Atkinson and McGlynn, 2009; Gupta et al., 2014). At present we cannot rule out that in the absence of RuvAB, RecU alone, or with the help of the branch migration translocase RecG, is loaded at the HJ to process them (Fig. 2.2) (Cañas et al., 2014). Processing of recombination intermediates During the repair of one-ended DSB the D-loop intermediate might be processed by a nuclease: perhaps MutS2 disrupts the D-loop (Pinto et al., 2005; Fukui et al., 2008; Damke et al., 2015), or an uncharacterized nicking enzyme (Fig. 2.4, stages e–g). Alternatively, RecG or RuvAB may catalyse the regression of the extended heteroduplex so that the D-loop becomes a HJ intermediate (Fig. 2.4, stages h–j) (Carrasco et al., 2012; Alonso et al., 2013; West, 2003). During the repair of two-ended DSB the D-loop recombination intermediate can be processed by three different mechanisms (Fig. 2.2, stages e–j). In one of these mechanisms, non-reciprocal transfer of DNA via SDSA takes place (Pâques and Haber, 1999; Fig. 2.2, stages h–j). The joint molecule might be disrupted by DNA unwinding (perhaps by PcrA). The invading and extended 3′-end anneals to a complementary sequence at the other side of the break, leading to NCO products via RecO in bacteria (Fig. 2.2 stages h–j) or via Rad52 in yeast (Pâques and Haber, 1999). In B. subtilis, there is no evidence that asymmetrically extended D-loop intermediates could proceed via a putative Mus81-Mms4 (Eme1)

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structure specific endonuclease, as it has been reported in eukaryotic cells (Osman et al., 2003; Meddows et al., 2004; Table 2.1). In the other two avenues, the extension of the strand exchange reaction leads to the conversion of a D-loop into a HJ and after second end capture directly by RecA (or by RecO), and DNA synthesis to a genuine double HJ (dHJ) (Fig. 2.2, stages e–g) (Holliday, 1967; Szostak et al., 1983; Lopez et al., 2005; Bachrati and Hickson, 2008; Wu et al., 1999; Fig. 2.2, stage g). As predicted by the canonical DSB repair model, the strand cleavage of opposite arms of the HJ by RecU in concert with RuvAB, at certain preferred structures, and concomitant strand ligation leads to CO or NCO products (Fig. 2.2, stage f) (Alonso et al., 2013). The orientation of resolution of the HJ(s) could result in monomeric, NCO, or dimeric, CO, chromosomes with a 50% ratio (Szostak et al., 1983). However, in wt cells, molecular mechanisms that introduce a strong bias for the direction of HJ processing towards NCOs have been uncovered (van Gool et al., 1999; Cromie and Leach, 2000; Carrasco et al., 2004). Dimeric chromosomes (CO products) formed may be decatenated by the action of Topo IV and converted to monomers by the action of the RipX-CodV tyrosine recombinase (counterpart of XerCDEco, reviewed by Sherratt, 2003), in concert with the SpoIIIE and/or SftA translocases upon recognition of their cognate sites (Kaimer et al., 2009, Ptacin et al., 2008). They act at the dif recombination sites, located close to the terminus region that is segregated last, just before cell division occurs, and thus when chromosome dimers need to be resolved. The role of the SpoIIIE translocase in the site-specific recombination reaction is to locate the two dif sites into close proximity (Ptacin et al., 2008). In the absence of RecU or any of the branch migration translocase, a chromosomal segregation defect was observed in 4–8% of total cells, suggesting that alternative avenues can be involved in the separation of entangled chromosomes (Carrasco et al., 2004). E. coli Topo III, which co-purifies with SSB and RecQ (Butland et al., 2005), in concert with RecQ and SSB can dissolve the HJs to render only NCOs products (Lopez et al., 2005, Bachrati and Hickson, 2008, Wu et al., 1999). It is likely that a B. subtilis Topoisomerase I (Topo I or Topo III) in concert with a RecQ-like helicase (RecQ or RecS) and perhaps SsbA disentangles the recombination

intermediates (Fig. 2.2, stage g). The recG ruvAB double mutant strain exhibits a synthetic growth defect (Sanchez et al., 2007b). It is likely that RecG or RuvAB catalyses the interconversion of replication forks and HJs, and that these intermediates can be either resolved by RuvAB-RecU, or dissolved by Topo I/III-RecQ(RecS)-SsbA, rather than disrupted by the action of a putative D-loop disruptase (MutS2) (Fukui et al., 2008), by a DNA helicase (PcrA, in SDSA pathway) or processed by a Mus81Mms4-like mechanism (Table 2.1). Genetic and biochemical evidences suggest that RecU itself and the human Rad51 paralogues associated with the human Gen1 HJ resolvase might share many features in common: (i) both RecU and the Rad51BCD–XRCC2 complex bind specifically to HJs and help RecA and Rad51, respectively, to initiate DNA strand exchange (Ayora et al., 2004; Carrasco et al., 2005; Lio et al., 2003); (ii) both RecU and Rad51C show synergy in their binding affinity with RecA and Rad51, respectively (Sigurdsson et al., 2001; Kurumizaka et al., 2001; Carrasco et al., 2005); and (iii) RecU and indirectly the Rad51C–XRCC3 complex are involved in the processing and resolution of HJs by the Gen1 resolvase (Liu et al., 2004; Ayora et al., 2004; Carrasco et al., 2005; Ip et al., 2008). However, the suggested role of human Rad51 paralogs in the positive and negative modulation of Rad51 function remains to be documented. Conclusions and perspectives Many of the functions and pathways involved in dedicated DSB repair share conserved features, as well as differences in Bacteria (Shuman and Glickman, 2007; Ayora et al., 2011; Carrasco et al., 2012; Lenhart et al., 2012). In B. subtilis, genetic, cytological and biochemical data allow us to postulate that during DSB repair, the RecN protein recognizes ssDNA tails on duplex DNA ends, and with the help of PNPase, specifically binds to the 3′-OH ends (Cárdenas et al., 2009, 2011; Sanchez and Alonso, 2005). After basal end resection, the AddAB complex, or RecJ, in concert with a RecQ-like helicase (RecQ or RecS) and SsbA, resects the 5´-ends, generating a proper substrate for RecA·ssDNA filament formation (Figs. 2.2–2.4) (Kidane et al., 2004; Kidane and Graumann, 2005a; Sanchez et al., 2006). However, RecA·ATP cannot be loaded onto

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SsbA-coated ssDNA (Manfredi et al., 2008, Yadav et al., 2012, 2013, 2014). The loading of RecA onto the 3′-tailed duplex or onto SsbA-coated ssDNA relies on RecO and RecR. We postulate that RecO, in concert with RecN and/or RecR, displaces SsbA from ssDNA and promotes RecA loading (Figs. 2.2 and 2.4) (Carrasco et al., 2008; Kidane et al., 2004; Manfredi et al., 2008; Sanchez et al., 2006; Shan et al., 1997). The role of RecX, RecF and PcrA in the modulation of RecA·ssDNA filament extension remains elusive (Cárdenas et al., 2012). RecU modulates RecA activities by promoting RecA-catalysed strand invasion and by inhibiting RecA-mediated branch migration (Ayora et al., 2004, Carrasco et al., 2005, Cañas et al., 2008, Kidane et al., 2009). RecA promotes partial disassembly of the RecN-induced large protein complex at the RC, and forms discrete threads or filaments that search for homology along the sister chromosome located in the other cell half (Kidane and Graumann, 2005a). Then, RecF and RecX may contribute to RecA filament assembly/ disassembly in an antagonistic manner (Ayora and Alonso, 1997; Lusetti et al., 2006), with RecX perhaps exerting a negative effect on the extension of RecA filaments (Cárdenas et al., 2012). Cytological studies have been used to demonstrate that at least the RecN, RecO, RecR, RecA, RecF or RecU proteins do not exist as a pre–assembled complex but rather assemble in an ordered fashion at the site of DSB (Kidane et al., 2004; Kidane and Graumann, 2005a; Sanchez et al., 2006). In particular, the dynamic movements of RecA protein have been monitored in living cells (Kidane and Graumann, 2005a; Lenhart et al., 2014). If the DSB occurs at the replication fork (oneended DSB), RecA-mediated strand exchange creates a D-loop that PriA-DnaD-DnaB-DnaI could exploit to assemble the DnaC helicase. The D-loop or the nicked HJ is resolved by RuvAB-RecU or RecG-RecU, dissociated by Topo I/III-RecQ-like helicase (RecQ or RecS)-SsbA or disrupted, perhaps by MutS2 (Fig. 2.4). Then, activated DnaC recruits DnaG and the chromosomal replicases (see Chapter 1). Preliminary data suggest that B. subtilis RecA can assemble on both the 3′- or 5′-ssDNA termini associated with the DSB (Carrasco et al., 2016). If the DSB occurs at any place, RecAmediated DNA strand invasion followed by DNA synthesis and end capture may form a genuine dHJ. The dHJ is branch migrated by RecG or RuvAB and

then resolved or dissolved, and later the replication fork is fully re-established (Fig. 2.2). Alternatively, the D-loop can be disrupted after DNA synthesis from the first processed DNA end, and this end can anneal with the other resected end of the break, leading to SDSA that can be the primary source of NCO (Fig. 2.2, stages h–j). In the absence of an homologous template or absence of end resection the NHEJ machinery reconnects the broken ends, but in the presence of end resection the poorly characterized SSA catalyses the joining of short patches of homology with very low accuracy during stationary phase (Weller et al., 2002) or dormant spores (Wang et al., 2006; Vlasic et al., 2014). Acknowledgements This work was supported by grants BFU201239879-C02-01, BFU2012-39879-C02-02 and BFU2015-67065-P from Ministerio Economia y Competividad-FEDER to SA and JCA, respectively. We are grateful to Carolina César, Cristina Cañas, Humberto Sánchez, and Tribhuwan Yadav for their contribution in early editions of this review, and to Candela Manfredi and Chiara Marchisone for the communication of unpublished results. We wish to offer our apologies to all researchers whose important work is not cited owing to space limitations. References Alonso, J.C., Cardenas, P.P., Sanchez, H., Hejna, J., Suzuki, Y., and Takeyasu, K. (2013). Early steps of double-strand break repair in Bacillus subtilis. DNA. Repair. 12, 162– 176. http://dx.doi.org/10.1016/j.dnarep.2012.12.005 Alonso, J.C., and Lüder, G. (1991). Characterization of recF suppressors in Bacillus subtilis. Biochimie 73, 277–280. Alonso, J.C., and Stiege, A.C. (1991). Molecular analysis of the Bacillus subtilis recF function. Mol. Gen. Genet. 228, 393–400. Alonso, J.C., Stiege, A.C., and Lüder, G. (1993). Genetic recombination in Bacillus subtilis 168: effect of recN, recF, recH and addAB mutations on DNA repair and recombination. Mol. Gen. Genet. 239, 129–136. Alonso, J.C., Tailor, R.H., and Lüder, G. (1988). Characterization of recombination-deficient mutants of Bacillus subtilis. J. Bacteriol. 170, 3001–3007. Amundsen, S.K., Fero, J., Salama, N.R., and Smith, G.R. (2009). Dual nuclease and helicase activities of Helicobacter pylori AddAB are required for DNA repair, recombination, and mouse infectivity. J. Biol. Chem. 284, 16759–16766. http://dx.doi.org/10.1074/jbc. M109.005587 Anand, S.P., Zheng, H., Bianco, P.R., Leuba, S.H., and Khan, S.A. (2007). DNA helicase activity of PcrA is not required for the displacement of RecA protein from DNA or inhibition of RecA-mediated strand exchange. J.

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Chromosome Arrangement and Segregation Peter L. Graumann

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Faculty of Chemistry and SYNMIKRO (Centre for Synthetic Microbiology), University of Marburg, Marburg, Germany. Correspondence: [email protected] https://doi.org/10.21775/9781910190579-03

Abstract After a bit more than two decades of the use of green fluorescent protein (GFP) – or immunofluorescence microscopy – to study bacterial chromosome segregation, it has become clear that this process is highly organized, temporally as well as spatially, and that a machinery exists that mediates an overall gradual separation of sister chromosomes. Several key factors in this process have been identified, and at least a rough overall picture can be drawn on how chromosomes are separated so highly rapidly and efficiently. Bacillus subtilis has a circular chromosome. Replication initiates at the origin of replication that is defined as 0°, and two replication forks proceed bidirectionally to converge at the terminus region, which is defined as 180°. All other regions on the chromosome are defined as the corresponding site on a circle (Fig. 3.1). DNA replication occurs in the cell centre, and duplicated regions are moved away from the cell centre towards opposite cell poles. How this process is energetically driven is still unknown, but entropic forces could play a major role. A dedicated protein complex called SMC forms two subcellular centres that organize newly duplicated chromosome regions within each cell half, setting up the spatial organization that characterizes bacterial chromosome segregation. Several proteins, including topoisomerases, DNA translocases and recombinases, ensure that entangled sister chromosomes or chromosome dimers can be completely separated into the future daughter cells shortly before cell division occurs at the middle of the cells.

Figure 3.1 Schematic drawing of the B. subtilis chromosome. Regions are defined as corresponding degrees on a circle. A plasmid carrying a tandem repeat of lacO sequences can be integrated at any position on the chromosome, which is visible in a fluorescence microscope due to binding of many LacI-GFP molecules to the compacted region.

Chromosome compaction The B. subtilis chromosome with its 4.2 million base pairs has a contour length (i.e. when the circular chromosome would be fully extended into a single circle) of about 1.5–2 mm, but it occupies only a space of about 1–2 µm within the cell, called the nucleoid (Fig. 3.2A). One level of compaction is achieved by HBsu protein, which belongs to the

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Figure 3.2  Arrangement of the B. subtilis chromosome. (A) Chromosomes are compacted into nucleoids that fill the centre of the cell. Late in the cell cycle, two new nucleoids arise within each cell half (indicated by grey triangles), that are completely separated from each other before cell division occurs in the cell middle (indicated by white triangles). Very rarely, nucleoids are not completely separated when septation occurs (indicated by white arrow). In this case, the SpoIIIE rescue system (Fig. 3.7 and text) pumps the entrapped sister chromosome completely into the two cell halves. (B) Cartoon showing the preferred arrangement of the B. subtilis chromosome. Replication origins are moved to the cell poles at an early time point during the cell cycle, while the terminus region remains close to the cell centre, until the duplicated termini are separated soon before cell division occurs. Regions between origin and terminus are located between the subcellular sites, such that the chromosome is compacted roughly according to its physical structure. (C) Schematic view of the orientation of chromosomes in B. subtilis, Escherichia coli, Vibrio cholerae and Caulobacter crescentus cells.

group of bacterial histone-like proteins. HBsu is a small, basic protein that bind to dsDNA in a nonspecific manner, but prefers DNA with an intrinsic bend. About 50000 copies of HBsu are present within growing cells (Ross and Setlow, 2000), which can cover a good portion of the chromosome, but by far not all chromosomal DNA. HBsu is essential for viability (Micka et al., 1991), and the depletion of HBsu leads to strong chromosome

decondensation (Köhler and Marahiel, 1997), showing that it is a bona fide DNA compaction factor. The homologous protein HBst was shown to form a dimer in the crystal structure, with two basically charged arms that appear to interact with the phosphate backbone of the DNA (Tanaka et al., 1984). Based on its similarity to IHF, one could speculated that like IHF (Rice et al., 1996), HBsu may bend DNA upon binding with its long

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symmetrical arms and thereby induce a kink in the DNA, or stabilize DNA with an intrinsic bend to remain in the more compacted, bent form. However, the exact mechanism of DNA compaction by HBsu is not clear. A second major factor for chromosome compaction is the SMC complex, which is described further below. Chromosome arrangement and sister chromosome segregation To address the question if bacterial chromosomes are a random coil of DNA, or have something of an ordered structure, it was crucial to visualize defined regions on the chromosome. This can be done in live cells, using a technique originally designed for yeast cells by the Murray and Belmont laboratories (Robinett et al., 1996). A tandem array of lactose operators (200–400 copies) integrated into a plasmid can be incorporated at any site on the chromosome, when a 1 kb fragment from a site on the chromosome is also present in the plasmid, which will be used for homologous recombination with and integration into the chromosome. The region carrying the lacO cassette is visible in cell expressing LacI-GFP (Fig. 3.1). Initial experiments using this system showed that exponentially growing cells that contain only one or two chromosome copies usually contain two origin signals, one within each cell half (and usually each one close to a cell pole), and one or rarely two signals for the terminus region, close to the cell centre (Webb et al., 1997). When a second site on the chromosome was visualized using immunofluorescence (using antibodies against Spo0J protein that binds to several sites surrounding the origin of replication), it became clear that the B. subtilis chromosome has a conserved and preferred arrangement within the cell, and is compacted according to its physical structure (Fig. 3.2B) (Teleman et al., 1998): during the largest part of the cell cycle, origin regions are closest to the cell poles, while the terminus region (or both termini at the end of the cell cycle) are located towards the cell centre. The arms of the chromosomes lie side by side along the length of the cell. In a cell with a single chromosome, the arrangement would be as depicted in Fig. 3.2C, with origin and terminus at opposite cell poles, and arms side by side. This arrangement is similar to that seen in Caulobacter crescentus or Vibrio cholerae, and different from that

of slow-growing Escherichia coli cells, which have the chromosome arms within opposite cell poles (Graumann, 2014). Further experiments showed that origin regions on the chromosome are replicated at an early time point during the cell cycle, and are separated soon after their duplication (Sharpe and Errington, 1998). Using time lapse microscopy, it was found that origin separation, as well as separation of 90°, 180° or 270° regions, occurs in a rapid manner (Teleman et al., 1998; Webb et al., 1998). Origin regions move towards opposite cell poles with an average speed of 0.17 µm/min, while 90° or 270° regions are separated at a later time point during the cell cycle, and 180° last (Fig. 3.3). Thus, chromosome segregation occurs concomitantly with replication, and duplicated regions are separated in a sequential manner, soon after they have been replicated. When cell wall growth was inhibited, origin regions still showed rapid segregation (Webb et al., 1998), revealing that an intracellular machinery or mechanism must exist that efficiently partitions chromosomes during ongoing replication. In a recent study, it was shown that even during slow growth, B. subtilis cells are partially diploid, that is, a majority of cells contains two origin regions and partially replicated chromosomes after cell division (Wang et al., 2014a). In other words, cells avoid having a single chromosome, and instead alternate between a predominant state, in which chromosome arms (left arm and right arm with oriC and ter in the middle; Fig. 3.1) lie next to each other along the long axis of the cell, as they are being duplicated (Fig. 3.4), or switch to an orientation in which they lie side by side, left and right of the replication fork (Wang et al., 2014a). Partial or full diploidity is a helpful strategy for the repair of DNA breaks, which can be most efficiently repaired using homologous replication (see Chapter 2), i.e. one broken chromosome uses the intact sister copy for faithful repair. Replication Replication of the chromosome is discussed in detail in Chapter 1. Here, it is briefly summarized why the spatial organization of replication is one of the central elements in chromosome segregation. The first key insight that helped understand how ordered chromosome arrangement and segregation

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Figure 3.3 Time-lapse microscopy showing rapid separation of origin or terminus regions on the chromosome towards opposite cell poles. The left two panels show origins of replication decorated by GFP-LacI, the right two panels terminus regions. The first and third panels show GFP channels, the second and fourth panel Nomarski DIC channels that reveal the outlines of the cells. Ends of cells and new division septa are indicated by white lines. Numbers indicate time in minutes for image capturing. Between minutes 20 and 24, origin regions separate within the small cell (and thus early during the cell cycle), while the terminus regions separate between minutes 40 and 44 within a large cell (and thus late in the cell cycle), just before a septum is formed at the cell middle.

are achieved within the cell was the finding that the replicative polymerase complex is a static factory in the cell centre (Fig. 3.4). In slow-growing cells, one replication centre can be observed after initiation of replication, and late in the cell cycle, two centres are present in most cells, one within each cell half (Lemon and Grossman, 1998) (see Fig. 1.1). The implication of these observations is that the DNA polymerase does not move along the chromosome, but that the chromosome must move through the polymerase complex. Indeed, Lemon and Grossman (2000) were able to show

that the chromosome moves through the central replication factory, which is feasible because DNA polymerase is a very strong motor. However, DNA is too flexible to be pushed over 0.5–1 µm from mid-cell towards the poles, so DNA polymerase can not be the motor that drives origin separation. The movement of the chromosome through the cell centre, where it is duplicated, and the ensuing separation of duplicated regions towards the poles can explain how chromosome regions are sequentially segregated during replication, a prerequisite for the establishment of the preferred arrangement of the chromosome (Fig. 3.2B). When exponentially growing cells are deprived of amino acids supplied in the medium, the stringent response kicks in and arrests replication at the STer sites, present on both sides less than 200 kbp away from the replication origin (Levine et al., 1995). In most stringently arrested cells, two replication origins are well separated and positioned close to the poles, showing that the segregation of origin regions must occur soon after initiation of replication. Therefore, there is little or no chromosome cohesion in Bacillus subtilis, as opposed to eukaryotic cells, in which sister chromosomes stay together from S-phase until Anaphase. Topoisomerases Owing to the helical nature of DNA, duplication of a DNA double helix will lead to the entanglement of sister chromosomes behind the replication fork (if the DNA polymerase turns around the DNA helix during DNA synthesis), or will lead to the formation of superhelical tension in front of the replication fork, caused through the action of DNA helicase. In other words, unwinding of the parent DNA strand to yield two ssDNA strands generates a torsion in the DNA ahead of the replication fork that would resist further unwinding after accumulation of too much tension. Topoisomerases can remove torsions in the DNA in several different ways. DNA gyrase and Topoisomerase IV can remove positive supercoils ahead of the DNA polymerase, and can also remove entanglements between sister chromosomes that would lead to complete chromosome segregation towards the end of the cell cycle (Espeli and Marians, 2004). Apparently, it is mostly DNA gyrase that works at the replication forks, because DNA gyrase forms discrete accumulations and foci

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Figure 3.4  Model for the B. subtilis cell cycle. Note that this model is oversimplified, because B. subtilis cells are usually partially diploid (i.e. enter the cell cycle with two origin regions and a single terminus region), even under slow growth conditions (Wang et al., 2014a). For simplicity, a ‘single chromosome’ cell cycle is depicted. Soon after initiation of replication, origin regions are moved towards opposite cell poles, possibly assisted through an MreB/RNA polymerase motor (indicated by dashed helices underneath the envelope, upper panel). At this point, the SMC complex is located at the cell centre, but soon thereafter moves towards opposite poles to two positions close to the origin regions (right panel). DNA gyrase accumulates at the central replication machinery, while Topo I forms accumulations within each cell half. After termination of replication, sister chromosomes need to be disentangled, which is mostly performed through Topo IV, and the SMC complex is required for the complete separation of chromosomes (lower panel). If a chromosome dimer has been formed between duplicated sister copies, the dimer is resolved into two chromosomes by the dif/RipX/SftA-SpoIIIE system, whose activity is enhanced by the RTP replication terminator protein (lower left panel). After complete separation of chromosomes, the division machinery can commence to synthesize the septum at mid cell, which will generate two daughter cells (left panel).

at the central replisome in growing cells (Tadesse and Graumann, 2006) (Fig. 3.4). DNA gyrase and Topo IV can move one DNA double strand through another by generating a double strand break, through with a DNA duplex is moved, and which is then resealed. This can also generate negative supercoiling (underwinding of DNA), which is essential for chromosome compaction. Contrarily, Topo I and III only generate a single strand break, through which the other strand of the duplex is moved. This can lead to the relaxation of negative supercoils generated by DNA gyrase, and indeed, in vivo, Topo I antagonizes the action of DNA gyrase. Mesophilic bacteria such as B. subtilis have an overall negative supercoiling (DNA is slightly underwound, facilitating replication and open complex formation through RNA polymerase), which is generated through the dynamic interplay of DNA gyrase and Topo I. Like most other bacteria, B. subtilis contains

four topoisomerases, three of which are essential (Topoisomerase I, IV and gyrase). Depletion of Topo I, IV or gyrase leads to the formation of large cells with a single nucleoid, and the formation of anucleate cells that arise through division within the DNA-free spaces away from the nucleoid (Kato et al., 1990; Steck, 1984; Tadesse et al., 2005; Zhu et al., 2001). Thus, these three topoisomerases are essential for chromosome segregation, and indeed localize to the nucleoids in growing cells. In contrast to gyrase, which forms foci at the replisome, Topo IV does not form foci on, but is uniformly dispersed throughout the nucleoids (Tadesse and Graumann, 2006). A major main function of Topo IV is the decatenation of sister chromosomes (removal of interlocking of chromosomes) at the end of the replication/segregation cycle (Fig. 3.4, lower panel). In the absence of Topo IV activity, sister chromosomes remain entangled and resist

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complete separation, blocking cell division. However, Topo IV also helps maintain overall negative supercoiling in B. subtilis cells (Tadesse et al., 2005). Like gyrase, Topo I also forms dynamic foci (which accumulate and dissipate in 1-min intervals, like those formed by gyrase), however, different from gyrase, foci are located away from the cell centre, and usually one within each cell half (Tadesse and Graumann, 2006). The bipolar localization pattern of Topo I is similar to the localization of RNA polymerase (RNAP) as bipolar transcription foci (TF; see Fig. 5.2A). Indeed, RNAP generates positive supercoils during transcription, and Topo I and RNAP have been shown to physically interact, suggesting that Topo I accumulates in TF to counterbalance RNAP action. Thus, areas with different degrees of supercoiling exist in the chromosome (more positive supercoils in the cell centre, and more negative supercoils at positions away from the cell centre). Interestingly, about 400 different superhelical domains exist in the chromosome (Postow et al., 2004; Winston and Pettijohn, 1988), that is, an increase or a decrease in superhelical density at one position doe not influence supercoiling at some distance away, upstream or downstream from that position. However, the on average 10,000 kb comprising domains are not fixed, rather, domain barriers appear to be rather fluid. The basis for these domains and the function of topoisomerases will be further described in the following section. Proteins mediating homologous recombination are involved in chromosome segregation Homologous recombination describes the process in which homologous DNA strands can be exchanged between different DNA molecules (see Chapter 2). Double strand breaks (DSBs) in DNA can be efficiently repaired via HR, because the genetic information of the non-broken sister chromosome is used to repair a DSB (see Fig. 2.2). Mutations in HR proteins frequently cause generation of tumours and genetic diseases in humans, and cells (e.g. bacteria) that become highly sensitive to agents causing DNA damage. HR also mediates insertion of foreign DNA into chromosomes, in case the foreign DNA bears sufficient homology to the host chromosome. This process is called transformation (see Chapter 13, ‘Competence’) and yields

genetically altered cells; it is important in terms of biotechnological manipulation of cells, and also in terms of health, because bacteria acquire resistance genes via HR, and thus become immune against antibiotics inside and outside of hospitals/clinics. A multitude of proteins are required for HR to occur efficiently. Central to HR is the RecA protein (bacteria) and its homologue Rad51 (eukaryotes) that promotes strand exchange and formation of crossovers (Fig. 2.2), a stage called synapsis. RecA/ Rad51 need to be loaded onto DNA, which is achieved by several proteins acting prior to RecA, a stage termed presynapsis. Crossovers need to be resolved to yield two parental or recombined DNA molecules, and again, several proteins and protein complexes act during this stage of postsynapsis (Cromie et al., 2001) (see Chapter 2). Several mutations in genes encoding for HR proteins cause defects in chromosome segregation. Deletions in recF (presynapsis), ruvA, recU and recG (postsynapsis; Fig. 2.2) cause formation of anucleate cells, between 1% and 3%, respectively (Carrasco et al., 2004; Pedersen and Setlow, 2000; Sciochetti et al., 2001a). Deletion of recU exacerbates the segregation defect in smc or in spo0J mutant cells (two proteins involved in chromosome segregation, see below), showing that RecU truly affects chromosome segregation, and that therefore, resolution of crossovers is a vital aspect in segregation during most cell cycles. Although only few or no anucleate cells have been observed in the absence of RecA, recA mutant cells contain highly abnormal nucleoids and show highly reduced viability compared with wild type cells. Therefore, formation of crossovers is important during the cell cycle in most cells. It is possible that RecA helps restart stalled or blocked replications forks (see also Chapter 2). The finding that depletion of PolC, one or the replicative DNA polymerases, prevents most cells from going through a whole cell cycle (Dervyn et al., 2001) indicates that replication forks frequently disintegrate, possibly because the polymerase needs to be exchanged for a ‘fresh’ enzyme. HR is one avenue to restart a collapsed replication fork, explaining why RecA may play such an important role during the cell cycle. On the other hand, a recA deletion reduces the formation of anucleate cells in recU mutant cells (Carrasco et al., 2004), supporting the idea that anucleate cells arise when crossovers can not be resolved. These data

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show that HR occurs within ever cell cycle, and that a defect in the process causes severe disturbance of proper chromosome segregation. SMC proteins and the SMC/ ScpAB complex organize the arrangement of duplicated chromosome arms The third important element of the chromosome segregation machinery (counting the replisome and topoisomerases as the first two components) was identified as Smc (structural maintenance of chromosomes) protein. SMC proteins are found in almost all organisms, exceptions are such illustrious bacteria as Spirochaetes or Deinocuccus radiodurans, which have multiple chromosome copies (note that most bacteria that have multiple chromosomes, e.g. Cyanobacteria, do have Smc proteins). SMC proteins are central components of several protein complexes in pro- and eukaryotes (Kleine Borgmann and Graumann, 2014). Condensin and cohesin complexes are essential for mitosis, and mediate proper chromosome compaction/structure, or cohesion of sister chromosomes, which is essential for entry into Prophase, or for alignment of chromosomes at the metaplane and transition to Anaphase, respectively (Nasmyth et al., 2000; Strunnikov and Jessberger, 1999). SMC proteins consist of an N-terminal region containing the nucleotide binding Walker A motif, one part of the ATP binding pocket, two long central coiled coil regions separated by a hinge domain, and a C-terminal region that contains a Walker B motif (the other part of the ATP binding pocket) and a ‘C’-motif required for ATP hydrolysis (Koshland and Strunnikov, 1996). The coiled coil regions fold back onto each other, forming a 50-nm-long coiled coil. The functional ATPase pocket is jointly formed by the N- and the C-terminal regions, which compose the head domain, as revealed in the crystal structures (Hopfner et al., 2000; Lowe et al., 2001). SMC proteins generally form dimers (Hirano et al., 1997; Melby et al., 1998), mediated by the hinge domain (Haering et al., 2002). In eukaryotes, Smc2 and Smc4 form the condensin SMC heterodimer, and Smc1 and Smc3 the cohesin dimer. Contrarily, bacterial SMC proteins generally form symmetrical homodimers, composed of a central hinge and two long arms with two head domains at their ends

(Fig. 3.5A). These molecules can adopt an open V-shape, or a closed structure, with closure apparently occurring at the head domains. Interestingly, when not-bound to DNA, B. subtilis Smc coiled coil arms are closed, but open up at the hinge domain when interacting with DNA, and stably encircle DNA when head domains engage and dimerize (Soh et al., 2015; Wilhelm et al., 2015) (see below for more details on the mode of DNA binding of SMC). In vitro, the condensin complex introduces positive writhe (a right-handed superhelix) into DNA (Kimura et al., 1999), which probably leads to the introduction of negative supercoiling in vivo. The cohesin complex and prokaryotic SMC have been shown to bind to DNA as a ring-like structure (Gruber et al., 2003; Hirano et al., 2001; Volkov et al., 2003), in which DNA is most likely bound by wrapping of the long coiled coil arms around the DNA, and by closing of the ring at the head domains. DNA is condensed in a highly cooperative and repetitive manner (Strick et al., 2004), but the actual mode of condensation is yet unknown. Hiraga and coworkers identified the fist SMClike protein, MukB, in E. coli as a central factor in chromosome segregation (Niki et al., 1991). The group hunted for genes whose mutation generates a high number of anucleate cells (lacking DNA), and identified the mukBEF operon via this approach. MukB was shown to form a complex with MukE and MukF, all of which are required for proper chromosome segregation (Yamazoe et al., 1999). Because MukB is a somewhat unusual SMC type protein, the wide distribution and important function for SMC proteins became only apparent with the finding that the smc gene in B. subtilis confers an important role in chromosome compaction and segregation (Britton et al., 1998; Graumann et al., 1998; Moriya et al., 1998). Interestingly, like its eukaryotic counterparts, B. subtilis Smc forms a complex with two other proteins, ScpA and ScpB, which were identified through sequence gazing (Mascarenhas et al., 2002; Soppa et al., 2002). ScpA and ScpB are widely conserved in prokaryotes, although some organisms exist that have an smc and an scpA gene, but no scpB, or vice versa. In vitro, ScpA forms a monomer, and to a minor extend dimers, while ScpB exclusively forms dimers (Volkov et al., 2003). An ScpA/2 ScpB complex binds asymmetrically to the Smc head domains, in which the C-terminus

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Figure 3.5  A) Architecture of the bacterial SMC complex, ScpA is shown as the blue monomer, ScpB dimers are in brown. B) 3D Structure of the Smc head domain/ScpAB interface of the SMC complex, Smc head domains and part of the coiled coils are in light blue and green, ScpA is dark blue, ScpB dimers are purple. C) Model for the function of the SMC complex. Inset shows Spo0J (ParB) molecules spreading on DNA and bridging distinct DNA strands; Spo0J facilitates the loading of Smc at sites surrounding origin regions on the chromosome. Smc molecules devoid of ScpAB can diffuse throughout the chromosome, while condensation centres contain the entire Smc/ScpAB complex that bridges chromosome arms and condenses DNA.

of ScpA interacts with the tip of one head domain (called the ‘cap’), while the N-terminus binds to the coiled coil region directly adjacent to the other head domain (called the ‘neck’) (Bürmann et al., 2013) (Fig. 3.5B). Bridging of Smc heads by ScpAB reduces Smc ATPase activity in vitro (Hirano and Hirano, 2004). Smc protein binds non specifically to double stranded DNA (Volkov et al., 2003), even in the absence of ATP, and an ATPase mutant form of Smc that is unable to bind ATP is fully proficient in DNA binding (TZ. Knust and PLG, unpublished results). Therefore, regulation of Smc ATPase activity may be an important function performed by ScpA and ScpB. The deletion of scpA or of scpB leads to an smc-like phenotype, the cells grow only below 23°C in rich medium, show very poor growth (growth is even slower in smc mutant cells than in scpA or in scpB mutant cells), form anucleate cells at high frequency, and have strongly decondensed

chromosomes (Mascarenhas et al., 2002). Deletion of smc and of scpA or of scpB does not acerbate the phenotype, showing that the genes are epistatic, i.e. that all three proteins act on a similar aspect of chromosome segregation and condensation, and are all required for proper function of the SMC complex. In vivo, the SMC complex appears to be in a dynamic equilibrium between free and ScpA/B-bound Smc, and between free ScpA and ScpB and a ScpA/B subcomplex (Mascarenhas et al., 2005b). In vitro, the ScpA/B subcomplex is recruited to form a complex with Smc, which most consists of an Smc dimer, an ScpA monomer and two ScpB molecules (Bürmann et al., 2013; Fuentes-Perez et al., 2012). It is possible that Smc and the ScpA/B subcomplex associate and dissociate at certain times in the cell cycle, or even at certain positions within the cells, but this issue has not yet been addressed experimentally.

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An important clue about the function of SMC proteins came from the findings that SMC protein shows connections with three topoisomerases at the genetic level, revealing that a key function of SMC lies in chromosome supercoiling. Firstly, a mukB deletion in E. coli can be suppressed by a mutation in topA, encoding for Topo I (Sawitzke and Austin, 2000), and the phenotype of an smc deletion in B. subtilis can be substantially rescued by a reduction in the level of Topo I (Lindow et al., 2002a). Secondly, the inhibition of gyrase severely acerbates the smc phenotype, the cells are hypersensitive to chemical reducing gyrase activity (Lindow et al., 2002a). These experiments show that SMC has an effect on supercoiling, working in the opposite direction of Topo I and in the same direction as gyrase. Because gyrase induces negative supercoils, which are relaxed by Topo I (see above), it is clear that the net effect of SMC action is the introduction of negative supercoils. Indeed, in vitro, condensin can introduce positive writhe into DNA (Kimura et al., 1999), which translates into negative supercoils. Thirdly, a genetic interaction also exists between SMC and Topo IV. Lower levels of Topo IV lead to chromosome decondensation, while overproduction of Topo IV induces nucleoid hypercompaction. Overproduction of Topo IV in smc mutant cells almost completely reverses the decondensation defect of nucleoids that now resemble those in wild type cells (Tadesse et al., 2005). However, there are still 7–8% anucleate cells in smc mutant cells during overproduction of Topo IV, and although the cells grow much faster than mutant cells with normal Topo IV levels, growth is still slower than in wild type cells (Tadesse et al., 2005). These experiments show that overproduction of Topo IV only partially compensates for the segregation and slow growth defect in the absence of SMC, but fully rescues the condensation defect, most likely through compensation of missing negative supercoils in smc mutant cells. Interestingly, global protein synthesis is strongly deregulated in smc mutant cells, but can be almost completely restored to the wild type pattern of expression through overproduction of Topo IV (Tadesse et al., 2005). These data support the idea that SMC has a strong influence on global supercoiling, which is known to affect transcription of a large fraction of all genes. Further important information on the mode of

action of the SMC complex came from the visualization of the proteins in live cells. Rather than localizing throughout the nucleoid like HBsu, the SMC complex localizes as discrete foci, in a cell cycle dependent manner (Fig. 3.6). Smc, ScpA and ScpB colocalize in defined foci that are present in the middle of small cells, and early in the cell cycle, two foci move towards opposite cell poles, in a manner similar to origins of replication. Later in the cell cycle, two or (rarely) four foci were present close to each cell pole (Lindow et al., 2002b; Mascarenhas et al., 2002; Volkov et al., 2003). Dual labelling confirmed that Smc, ScpA and ScpB colocalize. When Smc protein is overproduced in B. subtilis cells, nucleoids are strongly hypercondensed, although Smc protein is largely retained within the bipolar SMC centres (Volkov et al., 2003), showing that SMC has an effect on global chromosome condensation from the defined assemblies it forms. The dynamics of separation of Smc/Scps was monitored using time lapse microscopy, which showed that the separation is abrupt and its speed comparably fast to that of the movement of origin regions.

Figure 3.6 Formation of subcellular assemblies on the nucleoids of the SMC complex. Fluorescence microscopy of exponentially growing cells expressing ScpA-YFP, which localizes close to the cell centre in small cells (indicated by single triangle), and to two (or rarely four) foci, one (or two) within each cell half (indicated by two triangles per cell). SMC foci duplicate at the cell centre and move towards opposite cell poles with dynamic similar to those of replication origins, but after the origins have moved towards the poles.

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Interestingly, the movement of fluorescent foci was asymmetrical, with one focus moving 5–10 min before the other towards the poles, as observed in a majority of cells, which was supported by statistical analysis. Average movement of foci was 0.09 µm/ min over an average distance of separation of 1.3 µm (Volkov et al., 2003), compared with 0.17 µm/min of movement over 1.4 µm for origin regions of the chromosomes in cells grown in a similar medium (Webb et al., 1998). During the B. subtilis cell cycle, the SMC complex appears to act soon after the initial separation of chromosome origins. This is based on several observations: colocalization of the SMC complex and of origin regions showed that origin regions move out towards the cell poles before the SMC complex, which assumes a bipolar position soon after origin movement (Volkov et al., 2003). Secondly, origin regions on the chromosome were found to still move apart in a rapid manner in a smc deletion strain (Graumann, 2000), and, thirdly, cells devoid of SMC contain multiple origin regions but much fewer termini signals than expected. Indeed, when purified spores from a smc deletion strain were germinated synchronously (spores contain only one chromosome), origin regions were able to separate, but not the terminus, in contrast to spores from wild type cells (Graumann, 2000). However, during rapid growth in rich medium, cells are not even able to separate origin regions when Smc or ScpAB are degraded (Gruber et al., 2014; Wang et al., 2014b), showing that during growth with multiple replication forks per cell, an additional need arises to efficiently separate origin regions. Thus, the SMC complex is clearly required for complete separation of chromosomes during each cell cycle, and for the separation of origin regions under rapid growth conditions. Likely as a consequence of a failure for sister chromosome segregation, ordered chromosome arrangement is lost in the absence of SMC, showing that SMC is required for the maintenance of chromosome organization. How could the SMC complex achieve efficient segregation of duplicated chromosome regions? I will first present a model at the end of this section, then present findings on Smc and its complex partners, which are important to understand its probably mode of action. During the cell cycle, the SMC complex localizes close to origin regions soon after the bipolar

separation of the origins, but later moves away from the origin regions, suggesting that the complex interacts with several regions on the chromosome, rather than with any specific region(s). Indeed, Smc binds non specifically to a wide range of DNA sequences (Volkov et al., 2003), and CHIP experiments showed that it interacts with several distinct sites on the B. subtilis chromosome (Lindow et al., 2002b). Interestingly, detailed chromosome binding studies have shown that Smc is highly enriched at several sites surrounding the origin region of replication, and found at most other sites at much lower frequency (Gruber and Errington, 2009). Indeed, Spo0J, and ParB protein, facilitates the binding of Smc to parS DNA in vitro and in vivo; Spo0J binds to at least 10 parS sites, which are present on the chromosome in the vicinity of origin regions. Therefore, Spo0J is a loading factor for Smc, and increase binding of Smc close to origin regions is a likely cause that most of the SMC clusters are present near oriC. Interestingly, purified Smc forms striking highorder structures in vitro, as visualized using atomic force microscopy (Mascarenhas et al., 2005b). The structures appear to consist of rosette-like arrays of Smc dimers that might form through protein–protein interaction of the head domains [hinge domains form intramolecular dimers only (Volkov et al., 2003)]. Because Smc binds to DNA by embracing its substrate with the long coiled coil arms, it is possible that many DNA loops are connected through the Smc rosette arrays, which could explain how SMC condenses DNA, and which might be the basis for the formation of the discrete subcellular SMC structures observed in vivo. When replication is arrested, the SMC complex rapidly loses its bipolar localization, rather, single central assemblies are formed. Upon restart of replication, the SMC complex rapidly resumes its bipolar localization pattern, showing that it depends on active replication (Mascarenhas et al., 2005b). It has been speculated that DNA-bound Smc may associate with the rosette-like superstructures observed in vitro, which could connect ordered DNA loops at their basis. This would explain the existence of independent superhelical domains (about 400 of which exist in B. subtilis or in E. coli cells) that may be set up through the SMC superstructures. The superhelical domains are not static, but flexible, possibly because SMC embraces DNA

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loops, but does not appear to bind with a specific domain, which would allow for flexible binding and slipping of DNA through the Smc arms (Fig. 3.6C). However, less than 60 Smc dimers per chromosome equivalent (dependent on the growth rate) are present within a cell (Wilhelm et al., 2015), and therefore, not enough molecules are present to set up the domain structure of the chromosome. FRAP (fluorescence recovery after photobleaching) analyses have revealed that the SMC complex has a slow turnover within the condensation centres, with half time recovery of around 4 min (Kleine Borgmann et al., 2013a). Therefore, once formed, condensation centres/Smc/ScpAB clusters are rather stable. A mutation that stabilizes the dimerization of head domains (‘transition state mutant’) displays a dominant negative phenotype (it hinders the function of wild type Smc) (Kleine Borgmann et al., 2013a) and forms fewer (but brighter based on fluorescence) condensation centres per cell (Minnen et al., 2016). ATP-binding mutant Smc or Smc that is unable to dimerize head domains do not form condensation centres, and are non-functional (Mascarenhas et al., 2005a), showing that an ATP cycle must operate that mediates the setting up and turnover of condensation centres, and that is essential for the function of Smc. Another interesting aspect is the finding that Smc protein is subject to posttranslational control. As the cells enter stationary phase, SMC is gradually degraded, until it becomes undetectable in stationary phase cells, although transcription of the gene remains rather constant. Smc is degraded to a lesser extent in protease mutants, showing that the protein is subject to proteolysis as cell growth ceases (Mascarenhas et al., 2005b). Surprisingly, Smc becomes unstable in cell extracts from growing cells after removal of all DNA present in the extract. Again, the protein is a bit more stable in DNasetreated cell extracts from protease mutants, showing that Smc is prone to proteolysis, and is protected from degradation in the presence of DNA. It is possible that this constitutes a mechanism for release of the SMC complex from DNA: ATP hydrolysis at the Smc head domains may trigger release of Smc, which may be followed by rapid degradation of the protein, rather by rebinding. Single molecule microscopy allows the visualization of individual GFP-labelled proteins in real time, and has revealed that only 20% of Smc

proteins are statically positioned within the condensation centres, while 80% move throughout the chromosome. Static Smc molecules are bound to ScpA and ScpB, while the dynamic Smc dimers move throughout the entire chromosome without their complex partners (Kleine Borgmann et al., 2013b). Interestingly, diffusion of single purified Smc dimers along stretched out DNA in vitro is independent of ATP and in a similar range as that seen for the dynamic molecules in vivo (Kim and Loparo, 2016). Upon an increase in the concentration of Smc, clusters are formed on the DNA that induce condensation of the bound DNA strands; when Smc is preincubated with ScpAB, the number of clusters is reduced, but DNA is still being compacted (Kim and Loparo, 2016). Interestingly, only Smc by itself is able to bind to non-specific DNA, while the entire Smc/ScpAB complex is unable to do so (Kleine Borgmann et al., 2013b). These data suggest that Smc dimers move through the chromosome by Brownian motion and interact with DNA in a non-specific manner, and form multimers upon meeting each other, with ScpAB playing a stabilizing function of cluster formation. Because Smc is efficiently loaded onto DNA at parS sites surrounding the origin region, through the activity of Spo0J (see below), Smc/ScpAB clusters are most likely to form in the vicinity of origin regions, in agreement with localization data. In vivo DNA entrapment assays show that indeed, the Smc/ScpAB complex binds to DNA as a stable ring the DNA runs through (Wilhelm et al., 2015). About 20% of all Smc dimers were seen to be bound to DNA in this way, which correlates with the 20% static ScpAB bound Smc molecules seen by in vivo single molecule microscopy (Kleine Borgmann et al., 2013b). These data suggest that Smc may move through the chromosome as a rod, transiently interacting with DNA through opening of coiled coil arms at the hinge region, while binding to ScpAB leads to ring opening at the head domains and stable binding as a ring structure (Soh et al., 2015), within the condensation centres. Taken together, the following – speculative – model for the function of the SMC complex can be drawn: newly synthesized Smc binds to DNA adjacent to origin regions, mediated by Spo0J (ParB) protein, and may introduce stable loops into the DNA, by bridging distant DNA strands (Fig. 3.5C). Smc molecules slide along both arms

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of the chromosome, and upon binding of the ScpAB subcomplex may adopt a DNA binding mode in which DNA is entrapped within the ring structure, which is stabilized by ScpAB bridging of the head domains, plus by reducing ATPase activity. SMC condensation centres could bridge chromosome arms and thereby facilitate separation of origin regions towards the cell poles. We have recently observed the existence of more than one condensation centre per cell half using super resolution fluorescence microscopy (Schibany and Graumann, unpublished results), and determined the number of Smc molecules to about 50–60 per cell in slow-growing cells, in agreement with data from the Gruber laboratory (Wilhelm et al., 2015). Because only 20% of these are statically bound to DNA (corresponding to the condensation clusters), only 10 or so Smc molecules appear to set up the clusters. Therefore, only few Smc dimers or possibly dimers of dimers might connect many individual DNA loops, but can not be the basis for the 200–400 chromosome domain barriers existing in bacterial chromosomes (Postow et al., 2004). Slow ATP hydrolysis might open the ring, leading to dissociation and proteolysis of Smc, such that a new cycle of binding at origin regions, diffusion through the chromosome and condensation by cluster formation can start (Fig. 3.5C). Alternatively, or additionally, Smc may help solve catenanes (intertwining) of chromosomes during replication, As sister chromosomes are expelled from the replication forks, they are intertwined, because of the helical nature of DNA. Based on the interactions with topoisomerases, Smc rings could help decatenation of DNA strands, which will be especially important during rapid growth and multiple origins per cell. Soj and Spo0J protein Soj belongs to the ParA ATPase protein family, while Spo0J belongs to the ParB protein family, both of which are found in many other bacterial species. Spo0J has been described before as a loading factor for Smc. Being a ParB type protein, Spo0J induces ATPase activity of Soj, its ParA partner, and binds to a specific site, called parS. Spo0J is a negative regulator of Soj, which in turn is a negative regulator of sporulation functions (Quisel et al., 1999). The deletion of spo0J therefore leads to

a block in sporulation (which is relieved when soj is also deleted), wherefore it was first identified as spo gene. However, a major function for Spo0J can be found in growing cells, where its absence leads to the formation of about 1% anucleate cells (a 100 fold increase over the wild type) (Ireton et al., 1994). Spo0J binds to several specific binding sites (parS) present on a 800 kbp region (about 20% of the chromosome) surrounding the origin region on the chromosome, and thus defines a centromerelike region on the B. subtilis chromosome (Lewis and Errington, 1997; Lin and Grossman, 1998). Its true function is still not entirely clear, but experiments from the Grossman and Moriya laboratories have suggested a role in the control of replication initiation, possibly via an effect on Soj. The absence of Spo0J or an increase in the ration of Soj to Spo0J results in an increase in the number of replication origins per cell, indicating increased and asynchronous initiation events (Lee et al., 2003; Ogura et al., 2003). Interestingly, Soj has been shown to interact with DnaA protein, the key replication initiator (see Chapter 1). Mutations in Soj affect replication initiation (Murray and Errington, 2008), suggesting that Soj has an effect on replication via DnaA protein. Thus, by influencing the timing of replication and copy number of the chromosome, Soj and Spo0J most likely have an indirect effect on chromosome segregation, because a higher number of origins would interfere with proper segregation. Additionally, Spo0J facilitates binding of Smc protein to parS sites in vitro (Gruber and Errington, 2009; Sullivan et al., 2009). The absence of Spo0J alters the distribution of Smc around origin regions on the chromosome, suggesting that at least a considerable fraction of Smc is loaded onto the chromosome at parS sites guided by Spo0J protein. It is not yet clear how important loading this loading step is, because a spo0J deletion results in a very minor phenotype, while an smc deletion is highly deleterious for the cell. Hi-C (chromosome conformation capture) experiments have shown that during most of the cell cycle, the B. subtilis chromosome has three major features: (a) origin regions show the highest degree of interactions between DNA strands within this vicinity; (b) the left and right arm of the chromosome (or in fact duplicated regions of left and right arms) lie side by side along the long axis of the cell, and interact with each other along their length; and

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(c) most interactions are seen within chromosome arms, where about 20 so-called CIDs (chromosome interaction domains) exist, which are often flanked by highly expressed genes (Wang et al., 2015). The experiments suggest that the chromosome most likely adopts a 3D arrangement similar to that of a lamp brush, with individual superhelical domains emanating in a helical fashion. Recent experiments have shown that ParB is important form the connection between DNA regions within the origin region, and the SMC complex for the connections between the two chromosome arms (Wang et al., 2015). Possibly, the special tightly folded structure of replication origins (Fig. 3.5C) and the connection of chromosome arms behind the replication forks help to orient duplicated origin regions towards opposite cell poles, which appears to be a crucial event at the onset of chromosome segregation. Two DNA translocases couple late stages of segregation with cell division SMC mutant cells are unable to properly separate the chromosomes, but are still viable at low growth rates. They can only divide through non-separated nucleoids (Graumann, 2000), leading to so called ‘guilloutining’ of the chromosome, in that the septum simply comes down onto the chromosome. The reason that the cells can survive this situation is that the SpoIIIE protein can pump DNA through the closed septum and thus rescue the entrapped chromosomes (Burton et al., 2007). SpoIIIE is a DNA pump that moves DNA from the mother cell into the forespore after asymmetrical division during sporulation (see Fig. 11.5C), and is essential for sporulation (Wu and Errington, 1994). In exponentially growing cells, SpoIIIE is constitutively expressed and is dispensable for growth (Wu and Errington, 1997). The translocase localizes throughout the membrane, but assembles at the division septum when DNA becomes trapped within the membrane (Biller and Burkholder, 2009; Kaimer et al., 2009). The deletion of both, smc and spoIIIE is lethal, and the depletion of Smc in spoIIIE mutant cells leads to guilloutining, and to cessation of growth (Britton and Grossman, 1999). Thus, SpoIIIE acts as a rescue system that can move entrapped chromosomes into daughter cells (Fig.

3.7B), which most likely requires an extended amount of time. This may explain why smc mutant cells can only grow up to a certain (low) growth rate (up to 23°C in rich medium, up to 37°C in minimal medium), because at higher growth rate, the SpoIIIE system is overwhelmed. The essential nature of SpoIIIE in an smc background shows that in the absence of a rescue system, the activity of SMC is essential for chromosome segregation and thus for viability. Recently, it has become clear that B. subtilis has a second DNA translocase, SftA, which also belongs to the FtsK/SpoIIIE DNA translocase family (Biller and Burkholder, 2009; Kaimer et al., 2009). These proteins form hexamers with a central channel that encompasses dsDNA, and

Figure 3.7 Model for a two-step segregation machinery that couples late steps of segregation with cell division. (A) SftA forms hexamers and binds to the FtsZ ring during septation and moves DNA away from the central division plane. SpoIIIE is dispersed within the membrane and only assembles after closure of the septum and in case DNA is entrapped within the septum. (B) SpoIIIE forms hexameric membrane-bound rings that transport DNA through the membrane. Most likely, SpoIIIE forms a hexamer in each membrane (only one is shown), and thus a double hexamer for each membrane-entrapped DNA strand.

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share a conserved translocase domain at their C-terminus, which hydrolyses ATP to perform the motor function. Contrarily to FtsK and SpoIIIE, SftA is a soluble protein lacking any membrane spans. It is recruited to the FtsZ ring at an early time point during cell division (see Chapter 4), and its absence causes a very mild segregation defect at the end of the cell cycle. However, when DNA damage occurs, which leads to an arrest in the cell cycle, the action of SftA becomes much more important, in finishing segregation before the block in cell division is lifted. When SftA and SpoIIIE are both lacking, all phenotypes are exacerbated, showing that the translocases synergistically facilitate late stages of chromosome segregation. They do this in a two-step mechanism: SftA moves DNA while the septum closes, while SpoIIIE only becomes active when DNA is entrapped within the membrane (Fig. 3.7A). How do DNA translocases ‘know’ which of the sister chromosomes to pump into which direction? For SpoIIIE and FtsK, it has been shown that short polar sequences (8 bases) exist, which are highly enriched around the terminus region of the chromosome (the last region to be segregated). The polar sequences are oriented towards the terminus, and provide the directionality of DNA pumping of the translocases, which recognize the sequences via their very C-terminal part (g-domain). Most likely, SftA also recognizes an as yet unknown polar sequence, since its g-domain is highly similar to that of SpoIIIE and to other FtsK like proteins. Dimer resolution Recombination between newly replicated chromosome regions appears to occur frequently in growing cells, and can lead to the formation of a chromosome dimer, when an uneven number of crossovers occurs. This means that towards the end of chromosome segregation, when DNA replication has terminated, a single chromosome exists that can not be further segregated (Fig. 3.4, lower left panel). This occurs at a frequency of about 15% of cell cycles, that is, 15% of the cells in a growing culture show a cytological block in segregation in the absence of a dimer resolution system. A chromosome dimer can be resolved by a last round of recombination between sites close to the terminus, provided that recombination is resolved to yield recombinant rather than parental products. An intricate system exists that mediates efficient

resolution of chromosome dimers. A specific site for recombination, called dif, exists close to the terminus of replication on the E. coli and on the B. subtilis chromosome. Dif is recognized and used for dimer resolution by two site specific recombinases called XerC and XerD in E. coli, and CodV and RipX in B. subtilis (Cornet et al., 1997; Sciochetti et al., 2001b). The dif sites are moved close to each other by the DNA-pumps FtsK (E. coli) or synergistically by SftA and SpoIIIE (B. subtilis) (Kaimer et al., 2011; Lemon et al., 2001; Recchia et al., 1999; Steiner et al., 1999); SftA moves dif sites during septation, and SpoIIIE after septation (in case DNA is entrapped in the septum) (Biller and Burkholder, 2009; Kaimer et al., 2009) (Fig. 3.4, lower left panel). The translocases juxtapose dif sites guided by the polar sequences, and recombinases ensure that crossovers between the dif sites are resolved to yield recombinant resolution, such that two separate chromosome are generated. Interestingly, RipX and CodV are pre-assembled at dif throughout the cell cycle, such that resolution can be instantaneous as soon as the two sites meet at mid-cell (Kaimer et al., 2011). In B. subtilis, the SpoIIIE/dif system also interacts with the replication terminator protein, RTP, which arrests replication forks at specific sites in the terminus region on the chromosome, regulating that replication forks do not run into each other in an uncontrolled way. RTP also appears to facilitate dimer resolution, because a rtp spoIIIE double deletion mutant is more severe than the single mutants (Lemon et al., 2001). Thus, termination of replication appears to be connected to dimer resolution at the end of chromosome segregation. Coordination of chromosome segregation and cell division Chromosome segregation is independent of cell division, because depletion of FtsZ, the first cell division protein to assemble as a ring at the future division site (Fig. 4.3), leads to the formation of long filamentous cells containing nicely separated nucleoids (Beall and Lutkenhaus, 1991; Graumann et al., 1998). On the other hand, inhibition of segregation (e.g. through inhibition of replication, of topoisomerase IV, or depletion of SMC) leads to the formation of long cells, and thus to a block or delay of cell division. One component that appears to delay the onset of cell division until termination

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of segregation is NOC protein. NOC stands for ‘nucleoid occlusion’, and refers to the observation that cell division usually does not take place across non-divided nucleoids, i.e. that the nucleoid blocks cell division (Fig. 4.5). In the absence of NOC, FtsZ forms rings even at positions occupied by the nucleoid (Wu and Errington, 2004). NOC is homologous to Spo0J and localizes to the nucleoid, where it may interfere with the formation of the FtsZ ring. This would be a simple mechanism ensuring that FtsZ only polymerizes between segregated nucleoids, i.e. only after segregation has been completed. Indeed, NOC has recently been shown to bind to a consensus sequence that is present throughout the chromosome, but less often around the terminus region on the chromosome (Wu et al., 2009). This way, NOC efficiently blocks Z ring formation throughout the replication/segregation of most of the chromosome, but repression is relieved during the last stages of segregation. So the signal for FtsZ assembly appears to be the movement of most NOC molecules away from the cell centre, where only the last part of the chromosome still needs to be duplicated and segregated. A more detailed description of NOC and regulation of the localization of FtsZ is given in Chapter 4. The segregation motor From the above section, it is clear that the SMC complex is not the driving force to separate origin regions of the chromosome. It is also unlikely that Soj/Spo0J play a motor-like role like in, for example, ParA and ParB C. crescentus (Lim et al., 2014), because of their mild phenotypes when deleted. It has been speculated that entropy may be a primary source for the separation of sister chromosomes: the bacterial chromosome can be viewed as many beads on a string, with the beads corresponding to the several hundred topological domains that have been observed. The polymeric beads will self-avoid each other and thereby unmix, which in a rod-shaped cell will lead to a gradual separation into opposite cell halves ( Jun and Mulder, 2006). Centrally located replication forks in fact facilitate the ordered unmixing, and it has been shown by computer simulation that bacterial sister chromosome can largely separate from each other during the cell cycle. However, unmixing is inefficient early after initiation of replication, and can only be

simulated if a soft surface of the nucleoid is assumed along which duplicated chromosome regions can move towards the poles ( Jun and Mulder, 2006). I propose that the dynamic tethering of chromosome arms via the SMC complex rigidifies origin regions (in conjunction with Spo0J/ParB-mediated strand bridging) and renders the movement of earlyreplicated chromosome regions away from the cell centre efficient enough to guarantee rapid separation, whose major source of energy is simply entropic repulsion. Unmixing becomes highly inefficient towards the end of the process (Di Ventura et al., 2013). DNA translocases are good candidates to speed up the complete unmixing of chromosomes. Entropy being the motor for chromosome segregation has the charm that it harnesses basic physical forces and avoids the need to invent a machinery whose directionality and activity need to be tightly regulated, and would fit the idea that bacteria evolved as geared towards highly efficient growth yet simple structural composition. It has been proposed that bacterial actin-like proteins can facilitate chromosome segregation. These proteins are essential for the formation of proper cell morphology, which is discussed in detail in Chapter 8. Bacterial MreB proteins are closely related to actin according to their threedimensional structure and ability to form long ATP-dependent filamentous structures in vitro (van den Ent et al., 2001). In vivo, MreB forms helical structures underneath the cell membrane, and is essential for cell viability ( Jones et al., 2001) (see Figs. 8.2B and 8.3). The depletion of MreB in B. subtilis, or in Caulobacter crescentus, or the overproduction of a dominant negative mutant MreB allele in E. coli severely affect chromosome segregation at the stage of separation of origin regions (Defeu Soufo and Graumann, 2003; Gitai et al., 2005; Kruse et al., 2003). In C. crescentus, inhibition of the polymerization of MreB blocks origin movement, but not that of the termini. Moreover, the existence of MreB spirals correlates with the existence of DNA within B. subtilis cells [that is, MreB spirals do not form in mutant cells lacking a chromosome (Defeu Soufo and Graumann, 2004)], and C. crescentus MreB has been shown to physically interact with the origin regions, directly or indirectly, while this was not the case for the terminus region (Gitai et al., 2005). In B. subtilis, MreB and Mbl have been shown to perform dynamic movement in the rage

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of few minutes to few seconds, dependent on the technique used, suggesting that they could perform motor-like tasks (Carballido-Lopez and Errington, 2003; Defeu Soufo and Graumann, 2004). Time lapse microscopy has suggested that MreB can move (that is polymerize and extend on one end of the filament and be released from the other end) towards the cell poles with a speed of 0.24 µm/min (Defeu Soufo and Graumann, 2004), which would allow for the measured separation of 0.17 µm/min for origin regions (Webb et al., 1998). However, it is still controversial if B. subtilis MreB protein is indeed the motor for chromosome segregation (Fig. 3.4, upper panel), because an mreB deletion that is lethal under normal growth conditions can be generated under special media conditions (including addition of high magnesium and sucrose concentrations), where apparently, no segregation phenotype was detectable (Formstone and Errington, 2005). Experiments by Dworkin and Losick (2002) have shown that RNA polymerase plays an important role in the process. Inhibition of transcription blocks the separation of duplicated origin regions, which remain in the cell centre, until transcription resumes. On most bacterial chromosomes, the prevalent orientation of transcription is away from the origin region, such that both arms of the circle have the opposite orientation for the majority of genes. Additionally, highly transcribed genes tend to be located closer to the origins, while genes towards the terminus region are usually associated with lower transcription rates. It was proposed that the concerted movement of RNA polymerase complexes away from the origin regions can lead to the movement of the regions towards the cell poles, based on the fact the RNA polymerase has strong motor activity when moving along DNA. Owing to the high viscosity of the bacterial cytosol, DNA would rather move through the RNA polymerizes, rather than the reverse. Interestingly, when the orientation of one of the chromosome arms is inverted in E. coli cells, a severe segregation phenotype was observed, which was not due to abnormal termination of replication (Niki et al., 2000). This suggests that the orientation of transcription away from the origin region is important for efficient chromosome segregation. Interestingly, E. coli MreB has been shown to specifically interact with RNA polymerase (Kruse et al., 2006). This raises the intriguing

possibility that many RNA polymerase molecules could be anchored and fixed through MreB, and may direct movement of duplicated origin regions towards the cell poles. Such a model would incorporate the need for both, RNA polymerase and bacterial actin, to drive segregation of bacterial origin regions. However, the question of the motor still awaits for definitive experiments. Chromosome segregation during sporulation At the onset of sporulation, cells divide asymmetrically to yield two cells with dissimilar size, the larger mother cell and the small forespore that develops into the mature spore (see Fig. 11.1). Exponentially growing cells do not contain DNA close to the cell pole (Figs. 3.2A and 3.8A), and division close to the cell pole results in the formation of small anucleate cells (so called minicells, which arise in Min mutants; see Chapter 4). Evidently then, the organization of the chromosome must be changed at the onset of sporulation, such that the small forespore compartment contains some DNA, ensuring that a protein pump can translocate a complete chromosome into the forespore. Indeed, while replication origins are close to the cell poles but never at the poles in exponentially growing cells, they are recruited to the poles by a sporulation-induced protein, RacA (Ben-Yehuda et al., 2003b). RacA binds to several sites on the chromosomes, and predominantly close to the replication origins, where its binding sites are heavily enriched. RacA specifically interacts with DivIVa, a membrane protein that is exclusively present at both cell poles, such that each origin is attached to a cell pole (Fig. 3.8B). This switch in chromosome arrangement ensures entrapment of a chromosome in the forespore, since the pole that is used for the establishment of the sporulation septum is chosen in a stochastic fashion. Transcription of racA is induced through activated Spo0a protein and sigma H and thus early during sporulation. The special structure of the chromosome with both origins attached to the poles is called axial filament. Axial filament formation is highly efficient, because forespores devoid of an origin region can not be detected in more than 5000 sporulating cells (Graumann and Losick, 2001). In the absence of RacA, many forespores completely lack DNA and are thus lost. After polar

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segregation machinery exists in sporulating cells that has little in common with the apparatus segregating chromosomes in exponentially growing cells.

Figure 3.8 Model for chromosome segregation during sporulation. (A) origin regions are located at the outer edges of the nucleoids, but are almost never present at the cell poles. DivIVa localizes specifically to both cell poles in growing cells (A) and in cells that have initiated sporulation (B). To ensure entrapment of the chromosome in the future forespore compartment, the specifically induced RacA protein binds to many sites surrounding the origin regions and mediates the binding of origin regions to the DivIVa protein at the cell pole. After formation of the single polar septum (C), SpoIIIE pumps the remaining two-thirds of the forespore chromosome into the small compartment. Thus, RacA achieves the switch in chromosome segregation during sporulation, and together with SpoIIIE mediates efficient segregation during asymmetrical division.

division, the two-thirds of the forespore chromosome that are still located in the mother cell must be transported into the forespore, which is achieved by the SpoIIIE protein (Wu and Errington, 1994), and hexameric pore-forming genuine DNA pump (Figs. 3.7B and 3.8C). Interestingly, SpoIIIE does not assemble in the sporulation septum in DNA-less forespores, revealing that the DNA pump requires a septum-entrapped chromosome for its assembly (Ben-Yehuda et al., 2003a). Only SpoIIIE produced in the mother cell assembles properly (Fig. 11.5C), which dictates the direction of DNA transport, i.e. the translocation of the remaining two-thirds of the chromosome into the forespore, rather than the reverse. Thus, a specialized chromosome

Conclusions The known events during the B. subtilis cell cycle and steps during chromosome segregation are summarized in Fig. 3.4. Chromosome segregation commences soon after initiation of replication, and runs in parallel with replication. This is in marked contrast to eukaryotic cells, where chromosome segregation during M phase operates completely separately from replication in S phase. Therefore, there is little or no chromosome cohesion in B. subtilis cells, which has strong implications for DNA repair via homologous recombination (see Chapter 2). Chromosome segregation features (a) a centrally located replication machinery, (b) topoisomerases that maintain proper superhelicity and untangle interlocked sister chromosomes, (c) putative help from MreB proteins and RNA polymerase activity, (d) ordered 3D arrangement and tight interstrand connections at the origin regions set up by the ParABS (Soj/Spo0J) system and (e) the SMC complex that helps maintain proper chromosome arrangement and supercoiling, as well as separation of duplicated chromosome regions. The current model includes an extrusion/capture mechanism, in which duplicated region on the chromosome are pushed or pulled towards opposite cell poles, possibly through an MreB/RNA polymerase motor, or simply by physical forces of unmixing of macromolecules, and chromosome arms that are arranged by the SMC complex such that duplicated regions find their appropriate location(s) within each cell half. The components of the segregation machinery set up the preferred ordered arrangement of the chromosome and mediate/aid the rapid separation of duplicated chromosome regions towards opposite cell poles. How is the spatial arrangement of the central replicase and bipolar SMC centres set up? It is interesting to note that at an early time point during depletion of MreB, when cell shape is not yet affected, DNA polymerase loses its central localization (Defeu Soufo and Graumann, 2005), and the position of the SMC centres also becomes abnormal (Defeu Soufo and Graumann, 2003). Also, the two replication forks can visible separate

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from each other, to come back together (i.e. closer than 250 nm) within a time frame of few minutes (Migocki et al., 2004). These data suggest that the replication machinery is not anchored at the cell centre, but is held at this position by a dynamic process. Possibly, the physical process that pulls or pushed replicated DNA regions towards the poles sets up an equilibrium, in which the DNA that is moved away from the cell centre towards opposite directions positions the replication machineries to the cell centre, in an equilibrium reaction (Fig. 3.4). Additional factors such as dimer resolution and the SpoIIIE/SftA rescue systems resolve occasionally occurring chromosome dimers and rarely arising septum-entrapped chromosomes, and thus act at the terminal phase of chromosome segregation. Thus, it appears that the segregation machinery in B. subtilis (and in other bacteria) is composed of few elements that are central cellular components (DNA polymerase, cytoskeletal element MreB, RNA polymerase, SMC condensation complex, ParAB, topoisomerases), which are anyway required for the basic cell functions of replication, maintenance of cell shape, transcription and chromosome compaction. It is tempting to speculate that ancient bacteria may have segregated their chromosomes through a slow purely entropic process, and that the achievement of fast growth that requires fast and efficient chromosome segregation evolved through adaptation of existing cellular components without the need for evolution of specialized mitotic factors such as motor proteins. In fact, with the exception of membrane GTPase dynamin (Low and Lowe, 2006), no nucleotide hydrolysis-driven motor protein has so far been found in bacteria. Acknowledgments Work in my laboratory is supported by the Deutsche Forschungsgemeinschaft (DFG), the Bundesministerium für Bildung und Forschung (BMBF) and the LOEWE initiative for science in Hessen, funding the LOEWE Centre SYNMIKRO (Centre for Synthetic Microbiology). I would like to thank Luise Kleine Borgmann (University of Luxemberg), Stephan Gruber (University of Lausanne) and Byung-Ha Oh (Korea Advanced Institute of Science and Technology) for Fig. 3.5.

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Wilhelm, L., Burmann, F., Minnen, A., Shin, H.C., Toseland, C.P., Oh, B.H., and Gruber, S. (2015). SMC condensin entraps chromosomal DNA by an ATP hydrolysis dependent loading mechanism in Bacillus subtilis. Elife 4. Winston, S., and Pettijohn, D.E. (1988). Relaxation of DNA torsional tension in defined domains of bacterial chromosomes in vivo. Can. J. Microbiol. 34, 522–525. Wu, L.J., and Errington, J. (1994). Bacillus subtilis SpoIIIE protein required for DNA segregation during asymmetric cell division. Science 264, 572–575. Wu, L.J., and Errington, J. (1997). Septal localization of the SpoIIIE chromosome partitioning protein in Bacillus subtilis. EMBO J. 16, 2161–2169. http://dx.doi. org/10.1093/emboj/16.8.2161 Wu, L.J., and Errington, J. (2004). Coordination of cell division and chromosome segregation by a nucleoid occlusion protein in Bacillus subtilis. Cell 117, 915–925. http://dx.doi.org/10.1016/j.cell.2004.06.002

Wu, L.J., Ishikawa, S., Kawai, Y., Oshima, T., Ogasawara, N., and Errington, J. (2009). Noc protein binds to specific DNA sequences to coordinate cell division with chromosome segregation. EMBO J. 28, 1940–1952. http://dx.doi.org/10.1038/emboj.2009.144 Yamazoe, M., Onogi, T., Sunako, Y., Niki, H., Yamanaka, K., Ichimura, T., and Hiraga, S. (1999). Complex formation of MukB, MukE and MukF proteins involved in chromosome partitioning in Escherichia coli. EMBO J. 18, 5873–5884. http://dx.doi.org/10.1093/ emboj/18.21.5873 Zhu, Q., Pongpech, P., and DiGate, R.J. (2001). Type I topoisomerase activity is required for proper chromosomal segregation in Escherichia coli. Proc. Natl. Acad. Sci. USA 98, 9766–9771. http://dx.doi. org/10.1073/pnas.171579898

4

Cell Division Frederico Gueiros-Filho Departamento de Bioquímica, Instituto de Química, Universidade de São Paulo, São Paulo, Brazil. Correspondence: [email protected] https://doi.org/10.21775/9781910190579-04

Abstract Cell division in rod-shaped bacteria like Bacillus subtilis is carried out by a contractile protein ring, made up of about a dozen different polypeptides and known as the divisome. This sophisticated macromolecular machine, which is centered around the tubulin-like protein FtsZ, is capable of promoting the coordinated invagination of the cell membrane and cell wall to create the so-called division septum. The goal of this chapter is to provide an overview of the mechanism of septum formation in B. subtilis. Emphasis will be placed on describing the properties of the individual division proteins and how they assemble into the divisome complex, and on a discussion of the regulatory mechanisms that ensure that septum formation will happen with great spatial and temporal precision. In addition, the peculiar asymmetrical division that happens during B. subtilis sporulation will be described. Introduction Cell division is the process of generating two viable descendants from a progenitor cell. This involves two coordinated events: the replication and segregation of the bacterial chromosome and the splitting of the progenitor cell by cytokinesis, which in bacteria is also known as septum formation. This chapter focuses on the mechanism of septum formation. For a detailed description of chromosome replication and segregation see Chapters 1 and 3. Given the importance of efficient reproduction for evolutionary success, bacterial cells have developed a remarkably sophisticated protein machine capable of precisely splitting a progenitor

cell at the right place and time in every cell cycle. This machine, which is known as the divisome (Nanninga, 1998), is based on a contractile protein ring, as in the case of eukaryotes. In contrast to eukaryotic cells, however, which use actin and myosin in their contractile protein ring, the bacterial contractile machine is based on the tubulin-like protein FtsZ. Understanding bacterial cell division started more than 40 years ago when the first mutants impaired on division were isolated (Hirota et al., 1968; Nukushina and Ikeda, 1969). These mutants provided a first handle on the components of the division pathway. It was not until the pioneering work of Bi and Lutkenhaus in the early 1990s, which used electron microscopy to determine the distribution of FtsZ during the cell cycle and discovered the Z ring (Bi and Lutkenhaus, 1991), that the function of these components began to be understood. Since then, the field has blossomed, advanced by the powerful genetic tools available for Escherichia coli and B. subtilis, as well as the intensive use of cell biological approaches. In this regard, it is noteworthy that the study of cytokinesis, by popularizing the use of protein localization methods, has helped establishing a new view of cellular microbiology, centred on the idea that bacterial cells possess bona fide cytoskeletal elements and discretely localized proteins and thus are far more architecturally complex than previously anticipated. The goal of this chapter is to review the mechanism of cytokinesis in B. subtilis. This mechanism is generally conserved in bacteria and often information derived from other model organisms such as E. coli and Caulobacter crescentus will be used to fill

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gaps in the B. subtilis literature. Despite the general conservation of the process, however, there are specializations that are unique to B. subtilis and its closest relatives. Thus, whenever appropriate, such specializations will be highlighted. Anatomy of Bacillus subtilis and Escherichia coli envelopes: implications for cytokinesis Cytokinesis requires the coordinated remodelling of the different layers that make up the bacterial envelope. Even though the overall mechanism of cytokinesis is conserved in B. subtilis and E. coli, there are differences in the architecture of their envelopes and the way in which the septum is constructed that are worth highlighting. The envelope of B. subtilis, typical of Gram-positive bacteria, consists of the cytoplasmic membrane plus a thick cell wall made of peptidoglycan and associated anionic polymers such as teichoic acid (Fig. 4.1A; see also Chapter 10). The situation in E. coli is more complex, since it

has an LPS-containing outer membrane in addition to the cell wall and the cytoplasmic membrane (Fig. 4.1B). The presence of an outer membrane creates a need for specialized proteins involved in connecting the outer membrane with the cell wall during the invagination of the division septum (Gerding et al., 2007). Such proteins are not present in Grampositive bacteria and thus will not be discussed here. The cell wall in E. coli is also much thinner than in B. subtilis, probably a single peptidoglycan layer in thickness (Holtje, 1998). The difference in cell wall thickness creates a marked difference in the anatomy of the B. subtilis and E. coli developing division sites. In B. subtilis, division happens by the synthesis of a straight cross wall of peptidoglycan. Initially, there is no obvious invagination or constriction at the division site (Fig. 4.1A, B and E). Invagination and maturation of the septa into new hemispherical poles will eventually occur by the degradation of the inner part of the septal peptidoglycan by the action of specific hydrolases. In contrast, in E. coli synthesis of the division septum

Figure 4.1 Anatomy of septation in B. subtilis and E. coli. (A) Schematic representation of the B. subtilis envelope and (B) fluorescence microscopy image of a septating B. subtilis cell. (C) Schematic representation of the E. coli envelope and (D) fluorescence microscopy image of a septating E. coli cell. Images (B) and (D) represent live cells that were treated with a fluorescent membrane dye. Note the absence of constriction at the B. subtilis septum. (E) Electron micrograph of a growing septum in B. subtilis, highlighting the simultaneous invagination of the cell wall and plasma membrane. (F) Typical chain of cells in an exponentially growing culture of B. subtilis. Chain formation is due to the delay that exists between septation and cell separation. Membrane stains are often used to assess septation in B. subtilis cultures because septa that have not yet undergone constriction cannot be detected by phase contrast or DIC methods. CW, cell wall; PM, plasma membrane; OM, outer membrane.

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is concomitant with constriction (Fig. 4.1C and D). This is because the thinner peptidoglycan septal wall in E. coli is split in the middle as it is synthesized (Holtje, 1998). Another consequence of the thick septum of B. subtilis is that division will often happen faster than cell separation. This gives rise to ‘chains of cells’, such as the one depicted in Fig. 4.1F, which are a hallmark of fast growing B. subtilis cultures. Each cell in these chains is fully separated from the next by a complete septum but will not become individualized until the septa connecting it to its siblings mature into hemispherical poles. Overview of cytokinesis Cytokinesis in B. subtilis can be summarized in the following series of events. First, FtsZ, which is normally a cytoplasmic protein (Fig. 4.2A), assembles into a ring structure associated with the inner face of the cytoplasmic membrane (Fig. 4.2B). This structure, known as the Z ring, is a cytoskeletal element that serves as a scaffold onto which the divisome complex will assemble. Z ring formation is the target of careful regulatory mechanisms, described in detail below, that ensure that division will happen in coordination with other cell cycle events and at the proper place along the rod-shaped cell. Next, the Z ring nucleates the assembly of the divisome by attracting about a dozen other proteins to the division site. These are mostly integral membrane proteins and their recruitment to the Z ring occurs through both direct and indirect interactions with FtsZ (Fig. 4.2C). Once a mature divisome has formed, it will promote the synthesis of a transversal cell wall and the coordinated invagination of the cytoplasmic membrane (Fig. 4.2D), eventually producing a septum separating the progenitor cell into two independent daughter cells. Completion of the septum requires a membrane fission event to separate the membranes of the daughter cells (Fig. 4.2E). Septum formation is accompanied by the constriction and disassembly of the divisome into its elementary parts (Fig. 4.2D) such that division proteins will have been redistributed back to the cytoplasm and cell membrane at the end of cytokinesis. The last step of cytokinesis is separation of the daughter cells by the action of murein hydrolases that degrade the inner part of the septal peptidoglycan wall (Fig. 4.2F).

Figure 4.2 The steps of cytokinesis. (A) In a newborn cell, FtsZ (grey circles) and the other divisome proteins, here represented as a membrane associated complex (black ‘paddle’), are dispersed throughout the cell. (B) Upon the decision to divide, FtsZ self-associates into the Z ring at the cell centre. (C) The Z ring acts as a scaffold and recruits the other division proteins to the division site, forming the divisome. (D) The assembly of a complete divisome triggers the ingrowth of the peptidoglycan cell wall and the invagination of the plasma membrane. Septum formation is followed by constriction of the divisome and disassembly of its components. (E) The ingrowth of the cell wall and membrane will eventually produce a complete septum separating the progenitor cell into two daughter cells. Finishing of the septum requires a membrane fission event to separate the membranes of the daughter cells. This step has been omitted from the figure for simplicity. (F) Separation of the daughter cells is accomplished by the degradation of the inner part of the septal peptidoglycan through the action of murein hydrolases.

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Bacillus subtilis division proteins There are currently 24 proteins known to be associated with division in B. subtilis: ClpX, DivIB, DivIC, DivIVA, EzrA, FtsA, FtsL, FtsW, FtsZ, GpsB, MciZ, MinC, MinD, MinJ, Noc, PBP1, PBP2B, SepF, SftA, SpoIIE, SpoIIIE, UgtP, YneA and ZapA. These can be divided in two main groups: (a) proteins that make up the divisome and are directly involved in the construction of the septum (DivIB, DivIC, EzrA, FtsA, FtsL, FtsW, FtsZ, GpsB, PBP1, PBP2B, SepF and ZapA) and (b) proteins that regulate the assembly of the divisome (ClpX, DivIVA, MciZ, MinC, MinD, MinJ, Noc, UgtP and YneA). SpoIIIE and SftA, proteins whose function is to coordinate septum formation with chromosome segregation, represent a third functional group. A summary of the features of the B. subtilis division proteins is presented in Table 4.1. Most of the components of the B. subtilis divisome have counterparts in E. coli and many other bacteria. These proteins represent the core division machine that was probably already present in the last common ancestor of all bacteria (Robson F. Souza, Sandro de Souza and Frederico GueirosFilho, unpublished observations). The genes for many cell division proteins are often found next to each other and to some cell wall biosynthetic genes in a conserved chromosomal region that has been called the dcw (for ‘division and cell wall’) cluster. The remarkably conserved gene order of this region in diverse bacterial genomes suggests that there are functional consequences of this arrangement for division and for the maintenance of a rod shape (Mingorance et al., 2004). The dcw cluster of B. subtilis is located at around 135 degrees on the chromosome. ftsA, ftsZ, divIB, pbpB, ftsL and ftsW are some of the genes commonly present at the dcw cluster. In B. subtilis and several other Gram-positives, the dcw cluster extends further on one side to include also the genes for DivIVA and SepF, in addition to several conserved ‘ylm’ genes whose function is still unknown (Massidda et al., 1998; Fadda et al., 2003; Hamoen et al., 2006). The remaining genes are scattered throughout the genome. In the next sections, the current knowledge about the B. subtilis division proteins will be summarized, following the order in which they assemble in the divisome.

FtsZ and the Z ring FtsZ is the master coordinator of septum formation and the most widely conserved division protein, being present in essentially all bacterial genomes that have been sequenced to date. There are exceptions, however, such as the obligate intracellular Chlamydia, the Planctomycete Pirellula and the Mollicute Ureaplasma (Vaughan et al., 2004), which lack FtsZ and thus must employ a different mechanism to divide (Erickson and Osawa, 2010). Remarkably, recent studies showed that wall-less forms of B. subtilis (l-forms) are also capable of reproducing in the absence of FtsZ and do so by a process of membrane tubulation and blebbing (Leaver et al., 2009; Mercier et al., 2014). This FtsZindependent process may represent an ancient form of division that was substituted by the FtsZbased divisome in most bacteria but that may still be operational in some present day species. Despite their low sequence identity, FtsZ has been shown to be very similar to tubulin both at the structural and functional levels. FtsZ is a 40 kDa protein that folds into two independent globular domains (the N- and C- terminal domains) and has an unstructured tail of about 50 amino acids followed by a 15–17 conserved amino acid sequence at its extreme C-terminus (Lowe and Amos, 1998; Oliva et al., 2004) (Fig. 4.3A). This conserved terminal sequence is also known as the ‘C-terminal peptide’, or CTP. Self-assembly of FtsZ involves interactions between the C-terminal globular domain of one subunit with the N-terminal globular domain of another subunit (Oliva et al., 2004) (Fig. 4.3B). The CTP, on the other hand, is the binding site for several of the proteins that interact with FtsZ. Similarly to tubulin, FtsZ polymer formation is regulated by guanine nucleotides. Binding of GTP to a pocket located in the N-terminal domain induces the assembly of polymers (Erickson et al., 1996; Mukherjee and Lutkenhaus, 1998) (Fig. 4.3C). In contrast, when FtsZ is bound to GDP instead of GTP polymerization is inhibited. FtsZ polymers are highly dynamic both in vitro and in vivo (Chen and Erickson, 2005; Mukherjee and Lutkenhaus, 1998; Stricker et al., 2002), as would be expected of a cytoskeletal element that needs to undergo recurring assembly and disassembly at every cell cycle. The dynamic behaviour of FtsZ arises from its differential response to GTP and GDP and the fact that FtsZ polymerization is

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Table 4.1  B. subtilis division genes and their E. coli counterparts. Gene names that differ from the corresponding protein name are presented in parentheses B. subtilis E. coli orthologue orthologue Features and function ClpX

ClpX

46 kDa cytoplasmic protein. Substrate recognition subunit of the ClpXP protease complex. Negative modulator of Z-ring formation

DivIB

FtsQ

30 kDa membrane protein with extracytoplasmic domain. Part of multiprotein complex connecting the peptidoglycan synthesizing proteins to the early divisome.

DivIC

FtsB

15 kDa membrane protein with extracytoplasmic domain. Part of multiprotein complex connecting the peptidoglycan synthesizing proteins to the early divisome.

DivIVA

–

19 kDa membrane-associated protein. Topological determinant of Min system. Localizes MinCD to cell poles thus restricting division inhibition to this location. Analogous function to MinE in E. coli. Localizes to cell poles by recognizing curved membranes

EzrA

–

65 kDa membrane protein with cytoplasmic domain. Negative regulator of FtsZ polymerization. Prevents formation of multiple Z rings

FtsA

FtsA

48 kDa membrane associated protein. Structurally similar to actin. Stabilizes and tethers Z ring to membrane. Recruits downstream divisome proteins

FtsL

FtsL

13 kDa membrane protein with extracytoplasmic domain. Part of multiprotein complex connecting the peptidoglycan synthesizing proteins to the early divisome

FtsW

FtsW

44 kDa multipass transmembrane protein. Putative peptidoglycan precursor transporter

FtsZ

FtsZ

40 kDa cytoplasmic protein. Structurally and functionally related to tubulin. Self-assembles to forms cytoskeletal scaffold (Z ring) for divisome assembly

GpsB or YpsB

–

11 kDa cytoplasmic protein. Possibly involved in recruitment of PBP1 to divisome. Similar to DivIVA in N-terminus, may associate peripherally with cytoplasmic membrane

MciZ

–

40 amino acid peptide. Sporulation inhibitor of Z ring formation. Expressed in mother cell to prevent unwanted divisions during spore development

MinC

MinC

25 kDa cytoplasmic protein. Site-specific FtsZ inhibitor. Prevents polar Z-ring formation (with MinD)

MinD

MinD

29 kDa membrane-associated ATPase. Interacts with MinC and brings it to membrane. Prevents polar Z-ring formation

MinJ

–

44 kDa transmembrane protein with intracytoplasmic PDZ domain. Part of the Min system. Connects MinCD to DivIVA

Noc

?a

33 kDa DNA-associated FtsZ inhibitor. Effector of nucleoid occlusion. Analogous function to SlmA in E. coli

PBP1 (ponA)

PBP1A (mrcA)

99 kDa membrane protein. Bifunctional transglycosylase/transpeptidase involved in septal peptidoglycan synthesis. Recruited to divisome late in the cell cycle, probably with the help of GpsB.

PBP2B (pbpB)

FtsI or PBP3 (ftsI or pbpB)

79 kDa membrane protein with large extracytoplasmic domain. Transpeptidase involved in septal peptidoglycan biosynthesis. Likely to be associated with DivIB, DivIC and FtsL in a multiprotein complex comprising a subassembly of the divisome

SepF or YlmF

–

17 kDa cytoplasmic FtsZ-binding protein. Organizes FtsZ polymers in regular bundles. Absence reduces division frequency and affects septum structure. Overexpression can compensate for the absence of FtsA.

SftA

FtsK

107 kDa cytoplasmic protein. Second DNA pump of B. subtilis. Associates with divisome like FtsK and helps segregation before septation is complete. Works in tandem with SpoIIIE

SpoIIE

–

92 kDa transmembrane protein with cytoplasmic domain. Sporulation protein needed for efficient polar septum formation. Binds to FtsZ. Exhibits also phosphatase activity required for the activation of forespore-specific gene expression

SpoIIIE

FtsK

87 kDa protein with four transmembrane segments and a large cytoplasmic domain. DNA pump involved in chromosome segregation and dimer resolution. Dispensable for septum formation in B. subtilis but required in E. coli

UgtP

–

43 kDa cytoplasmic protein. UDP-glucose diacylglycerol glucosyltransferase that also functions as inhibitor of Z ring formation. Coordinates timing of Z ring formation with growth rate

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Table 4.1 Continued B. subtilis E. coli orthologue orthologue Features and function YneA

–

12 kDa membrane protein. SOS-induced septation inhibitor. Analogous function to SulA in E. coli

ZapA

ZapA

10 kDa cytoplasmic protein. Positive modulator of FtsZ. Binds to FtsZ and promotes polymer bundling in vitro. Stabilizes Z ring in vivo. Dispensable for septum formation

aNoc

is a member of the ParB family of DNA binding proteins that have several orthologues in E. coli. It is unclear which of the E. coli family members is the true orthologue of Noc.

Figure 4.3  FtsZ and the Z ring. (A) Crystal structure of FtsZ highlighting its main features. The protein can be divided by the helix H7 into two independently folding domains (boundaries marked by dashed line): the N- and C-terminal domains. The nucleotide binding pocket is in the N-terminal domain whereas the T7 loop involved in catalysing nucleotide hydrolysis is located in the C-terminal domain. The C-terminal peptide (CTP) is not seen in the crystal structure because it is unstructured. (B) Model of an FtsZ protofilament showing the contacts involved in the self-association of monomers. The spacing between subunits is 42 Å, very similar to the spacing between subunits in a tubulin protofilament (40 Å). Molecules of GDP are present in the binding interface between the monomers. Plus and minus signs indicate the polarity of the polymer. The plus end corresponds to the nucleotide binding surface of the protein. (C) Electron micrograph of a negatively stained sample of B. subtilis FtsZ polymerized with GTP. This sample exhibits predominantly straight protofilaments. (D) Fluorescence image of a Z ring in a dividing B. subtilis cell expressing GFP-tagged FtsZ. (E) 3D reconstruction of a Z ring carried out from optical sections of a cell expressing FtsZ-GFP like the one in (D). The dotted white line shows the boundaries of the imaged cell. Images in panels (A) and (B) adapted from Lowe et al., Annu. Rev. Biophys. Biomol. Struct. 2004;33:177–98 and reprinted with permission. Microscopy images in panels (D) and (E) adapted from Ben-Yehuda and Losick (2002) and reprinted with permission.

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coupled with activation of its GTPase activity. Thus, GTP-induced polymerization of FtsZ is quickly followed by the conversion of GTP to GDP within the polymer subunits. Because GDP-bound FtsZ has low affinity for itself, this then leads to polymer disassembly. Structurally, activation of GTP hydrolysis in FtsZ polymers is due to the positioning of a catalytic aspartate present in a C-terminal domain loop (the T7 loop) of one FtsZ monomer next to the GTP sitting in the binding pocket of another monomer. Another similarity between the biochemistry of FtsZ and tubulin is that FtsZ assembly is a cooperative process (Chen et al., 2005; Huecas and Andreu, 2003; Wang and Lutkenhaus, 1993). Structural studies of FtsZ polymerization using electron microscopy under different conditions have shown that FtsZ is capable of forming various types of polymers (Romberg and Levin, 2003). The simplest kind of polymer formed in the presence of GTP is the protofilament, a one-subunit thick head-to-tail string of FtsZ monomers (Fig. 4.3C). Protofilaments induced by GTP are straight polymers and can exist in isolation or associate into two dimensional sheets or 3D bundles through lateral interactions (Gonzalez et al., 2003). The tendency of FtsZ filaments to form lateral interactions is greatly increased by the presence of cations such as Ca2+ (Lowe and Amos, 1999; Mukherjee and Lutkenhaus, 1999), DEAE-dextran, or cationic phospholipids (Erickson et al., 1996) and by FtsZ binding proteins, such as ZapA and SepF, discussed below. Polymerization can be achieved also in the presence of GDP as long as stabilizing factors such as DEAE-dextran are present in the reaction. In contrast to GTP-FtsZ, polymers formed by GDPbound FtsZ are mostly curved. Helical tubes, mini rings and curved protofilaments have been detected in different preparations (Erickson et al., 1996; Lu et al., 1998). Much less is known about the structure of the Z ring in live cells. The Z ring has never been detected using standard transmission electron microscopy. This is likely because the polymer that makes up the ring is not bulky enough to be distinguished above the high density of the bacterial cytoplasm. If the ring were some form of protofilament sheet continuously encircling the cell it would exhibit a width of three to twelve protofilaments, based on estimates that there are 5000 to 20000 FtsZ molecules in a single B. subtilis or E. coli cell (Feucht et al., 2001;

Romberg and Levin, 2003). This width is likely significantly lower, however, if we consider that only one third of the total cellular FtsZ is present at the Z ring at any given time (Stricker et al., 2002). The recent application of cryoelectron tomography and super resolution fluorescence microscopy allowed a first glimpse on how FtsZ filaments are organized inside cells and revealed that the Z ring seems to be made up of relatively short protofilaments loosely associated via lateral and radial interactions. The ring, which spans a width of ~60–110 nm, is irregular and discontinuous, in particular in the case of Caulobacter crescentus, where partial rings are often observed at the site of cell division (Li et al., 2007; Holden et al. 2014). In E. coli and B. subtilis the ring tends to be continuous but is still heterogeneous, with high and low density regions (Strauss et al. 2012). In contrast, the Löwe group recently reported long continuous FtsZ filaments in cells and reconstituted liposome systems (Szwedziak et al., 2014). Further developments of these technologies, which still exhibit shortcomings (for example, cryoelectron tomography is limited to specimens