The Regulation of Dnaa in Caulobacter Crescentus

The Regulation of DnaA in Caulobacter crescentus

Richard Burns Wargachuk Department of Microbiology and Immunology McGill University, Montreal August 2012

A thesis submitted to McGill University in partial fulfillment of the requirements of the degree of Doctor of Philosophy © Richard Burns Wargachuk, 2012

Table of Contents Table of Contents ...... I Table of Figures ...... IV Abstract ...... VI Résumé ...... IX Acknowledgements ...... XII Contributions to original knowledge ...... XIII Chapter One ...... 1 Overview of the literature review ...... 2 Prokaryotic Chromosome Replication ...... 4 The E. coli model ...... 4 The E. coli origin of chromosome replication oriC ...... 5 The steps in the initiation of chromosome replication ...... 9 Diversity among bacterial origins of chromosome replication ...... 14 DnaA independent chromosome replication ...... 15 Bacteria with multiple chromosomes ...... 16 Replication of linear bacterial chromosomes ...... 18 Plasmid replication ...... 21 DnaA - Structure and function ...... 22 DnaA interacting proteins ...... 30 DnaA as a transcription factor ...... 32 AAA+ proteins -Structure and function...... 33 Mechanisms used to limit initiation of chromosome replication to once per cell cycle 35 Sequestration of the origin of replication ...... 36 Titration of DnaA away from the origin of replication ...... 37 Regulatory inactivation of DnaA (RIDA) ...... 38 Reactivation of DnaA ...... 45 DNA replication in eukaryotic cells ...... 47 Comparison of prokaryotic and eukaryotic chromosome replication ...... 47

Eukaryotic origins of replication ...... 47 Steps involved in eukaryotic chromosome replication ...... 48 Chromosome replication in the Archaea ...... 55 Caulobacter crescentus as a model for prokaryotic cell development ...... 56 Alphaproteobacteria ...... 56 Caulobacter crescentus cell cycle ...... 63 Master cell cycle regulators, CtrA, GcrA, DnaA, CcrM ...... 64 Chromosome replication ...... 68 Stationary Phase ...... 70 Proteolysis ...... 72 Rational and Objectives ...... 74 References ...... 77 Chapter Two ...... 91 Abstract ...... 92 Introduction ...... 94 Results ...... 98 Discussion ...... 143 Experimental Procedures ...... 148 References ...... 151 Chapter Three ...... 152 Abstract ...... 153 Introduction ...... 155 Results ...... 158 Experimental Procedures ...... 181 Discussion ...... 185 References ...... 188 Chapter Four ...... 189 Abstract ...... 190 Introduction ...... 192

Results ...... 193 Discussion ...... 219 Experimental Procedures ...... 223 References ...... 226 Chapter Five ...... 227 References ...... 233

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Table of Figures Chapter One Figure 1: Variation among the prokaryotic origins of chromosomal replication ...... 7 Figure 2: Steps involved in initiation of Chromosome replication in E. coli ...... 12 Figure 3: Replication of linear chromosomes in bacteria ...... 20 Figure 4: The domain structure of DnaA ...... 28 Figure 5: Mechanisms used to limit chromosome replication in E. coli ...... 41 Figure 6: Licensing of eukaryotic origins of replication ...... 51 Figure 7: Comparison of the initiation of chromosome replication in bacteria and eukaryotic cells ...... 53 Figure 8: The Caulobacter crescentus cell cycle ...... 60

Chapter Two Figure 1: A comparison of the domain structure of the DnaA protein with the HdaA protein ... 99 Figure 2: Creation of C. crescentus strain GM3700 that conditionally transcribes hdaA/cc1711 ...... 102 Figure 3: HdaA is required for the normal growth of C. crescentus ...... 105 Figure 4: HdaA is cleared from GM3700 cells when shifted to media lacking xylose ...... 108 Figure 5: HdaA is required for normal growth of C. crescentus ...... 110 Figure 6: Blocked expression of hdaA leads to over-replication of chromosomal DNA ...... 115 Figure 7: Depletion of HdaA results uncontrolled initiation of chromosome replication ...... 118 Figure 8: Clearing by proteolysis of DnaA during stationary phase requires HdaA ...... 121 Figure 9: HdaA promotes DnaA proteolysis in logarithmically growing cells ...... 124 Figure 10: Transcriptional activity of dnaA promoter in absence of HdaA ...... 129 Figure 11: Proteolysis of DnaA can be stimulated by starvation independent of HdaA ...... 131 Figure 12 Effect of depleted media on DnaA abundance in wild type and GM3700 cells ...... 136 Figure 13: SpoT is not required for the removal of DnaA upon entry into stationary phase .... 139 Figure 14: Proteolysis of DnaA during stationary phase is conserved among related fresh water Caulobacter species ...... 142

Chapter Three Figure 1: A comparison of the domain structure of the DnaA protein with the predicted HdaB protein ...... 159 Figure 2: Alignment of a highly conserved section of the DnaA binding domain from E. coli, C. crescentus with the HdaB ...... 161 Figure 3: List of bacteria among the alphaproteobacteria which contain a hdaB gene ...... 163 Figure 4: Replacement of the cc1058 locus with the omega antibiotic resistance cassette ..... 165 Figure 5: The hdaB gene is not required for the normal growth C. crescentus ...... 168 Figure 6: Environmental stress simulated by a sudden change in nutrients ...... 171 Figure 7: The relative abundance of DnaA in wild type C. crescentus (NA1000) and the Δ1058 strain ...... 174 Figure 8: Electrophoretic mobility shift assay ...... 176 Figure 9: DNase I protection assay ...... 179

Chapter Four Figure 1: Alignment of amino acid sequences of DnaA of E. coli and C. crescentus ...... 194 Figure 2: Expression and purification of C. crescentus DnaA proteins ...... 198 Figure 3: Intrinsic ATPase activity of C. crescentus DnaA ...... 200 Figure 4: The specificity of the new and the previously published DnaA antisera ...... 205 Figure 5: C. crescentus DnaA binds both ATP and ADP in vivo ...... 207 Figure 6: Abundance of DnaA in cells expressing mutant DnaA ...... 209 Figure 8: The stability of DnaA in cells expressing the DnaA ATPase deficient mutants ...... 212 Figure 8: Moderate overexpression of DnaAR357A and DnaAR357H results in a cell division defect ...... 216 Figure 9: The moderate overexpression of DnaAR357A or DnaAR357H results in an increase in chromosome replication ...... 217

Abstract All growing cells must ensure that their genetic material is faithfully replicated and divided equally to the newly formed daughter cells. Bacterial cells coordinate several pieces of information before beginning the task of chromosome replication, including cells mass and availability of resources to complete chromosome replication. Once chromosome replication has begun the cell must assure that it does not begin again until the next cell cycle. In nearly all bacteria chromosome replication begins at a single point on the chromosome, termed the origin of replication. In the bacterium Escherichia coli the activity of the initiator protein DnaA has been shown to be down regulated by the sequestration of newly replicated origins and titration of the initiator protein DnaA down-regulate replication. The third mechanism in E. coli is the regulatory inactivation of DnaA (RIDA). Shortly after initiation of replication the intrinsic

ATPase activity of DnaA is stimulated by a protein (Hda) with homology to the ATPase domain of DnaA, converting DnaA from the active ATP bound form to its inactive ADP bound form. The current models of chromosome replication have been developed using the model bacteria E. coli. DnaA from other bacteria differs from that of E. coli in structure and biochemical properties. A model based solely on E. coli may not satisfactory take into account the diversity found among bacteria.

The free-living Gram-negative bacterium Caulobacter crescentus is found in nutrient poor aqueous environments such as fresh water lakes and streams. Genomic analysis suggests that C. crescentus lacks canonical sequestration and titration mechanisms of DnaA regulation.

C. crescentus does however encode a protein with homology to Hda of E. coli. C. crescentus also makes use of novel regulatory mechanisms not found in E. coli to regulate DnaA. Unlike DnaA of

E. coli the DnaA protein of C. crescentus is unstable in growing cells, rapidly degraded during stationary phase and upon nutrient starvation. Intriguingly, C. crescentus also contains a second gene (hdaB) predicted to encode a protein with homology to the DNA binding domain of DnaA.

I performed the following experiments to determine if the RIDA mechanism, originally identified in E. coli, is functionally conserved in C. crescentus. I have identified a C. crescentus gene (hdaA) with homology to the E. coli hda gene. I have created a strain (GM3700) of C. crescentus with hdaA expressed under the control of a xylose dependent promoter. I demonstrated that C. crescentus depends upon a RIDA mechanism to prevent multiple initiations of chromosome replication in the same cell cycle, as blocking hdaA expression causes an increased frequency of chromosome replication as well as a blockage of cell division. I have also uncovered an unexpected role for HdaA in the stability control of DnaA. Removing HdaA from C. crescentus stabilizes the DnaA protein in growing cells and prevents DnaA protein removal from stationary phase cells. My experiments have identified a new role for HdaA in

DnaA proteolysis that presumably works in exponentially growing cells to aid the RIDA mechanism. I also demonstrate that HdaA unexpectedly participates in the programmed transition from exponentially growing cells to the stationary phase.

I have identified a second gene which displays homology to dnaA in C. crescentus. I called this second gene hdaB; it displays significant homology to the DNA binding domain IV of

DnaA. Sequence analysis of hdaB suggests that it is a paralogue of DnaA that retains key amino acids used by DnaA to make sequence-specific contacts with the DnaA R box. The hdaB gene does not appear to be essential for the normal growth of C. crescentus under laboratory conditions. A deletion strain with the entire open reading frame of hdaB replaced with an

VII antibiotic resistance cassette was indistinguishable from the wild type strain. Despite the obvious sequence conservation with the DNA binding domain of DnaA, experiments performed in vitro on purified HdaB protein were unable to detect HdaB -DNA interactions. These negative results do not indicate a role for hdaB in C. crescentus; the fact that homologs of HdaB are found in the genomes of other alphaproteobacteria suggests that, while not obvious under laboratory conditions, hdaB has an evolutionary conserved role in these bacteria.

The depletion of HdaA not only alters DnaA activity but DnaA stability as well. This suggests that C. crescentus employs both nucleotide binding/hydrolysis, as seen in E. coli, and a novel proteolytic mechanism to regulate DnaA. However, no previous studies have directly addressed the nucleotide binding and ATP hydrolysis of C. crescentus DnaA. To clarify the link between ATPase activity and protein stability I have created four independent single amino acid mutations in two conserved positions of C. crescentus DnaA, DnaAR300 and DnaAR357. I have shown that mutations in either arginine reduce DnaA ATPase activity in vitro. Mutations in the

C. crescentus DnaAR357 result in increased chromosome replication as well as an increase in

DnaA stability. Despite the effects of mutations in C. crescentus DnaAR300 observed in vitro, the C. crescentus DnaAR300A behaved similarly to wild type DnaA in vivo. These results indicate that the stability of DnaA in vivo is linked with its activity state and ties DnaA proteolysis with C. crescentus cell cycle progression.

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Résumé Toutes les cellules vivantes doivent s'assurer que leur matériel génétique soit fidèlement reproduit et divisé également parmi ses cellules filles. Chez les bactéries, le contrôle de la réplication des chromosomes se fait principalement au niveau de la fréquence de l'initiation. La réplication des chromosomes commence à un point prédéterminé sur le chromosome nommé l'origine de réplication. La fréquence d’initiation de réplication dans la bactérie Escherichia coli est contrôlée par trois mécanismes. Le premier mécanisme bloque l’accès aux origines de réplication à DnaA. Le deuxième redirige la protéine DnaA de l’origine. Le troisième se nomme le RIDA (regulatory inactivation of DnaA). Après l’initiation de la réplication, l'activité ATPase intrinsèque de DnaA est stimulée par une protéine ayant une homologie avec DnaA (Hda). Ceci stimule la conversion de la forme DnaA-ATP active en forme DnaA-ADP inactive. La protéine responsable de l'initiation de réplication des chromosomes est très bien conservée parmi presque toutes les bactéries. Plusieurs de leurs systèmes ont évolué pour régler l’activité de

DnaA qui répond aux besoins des bactéries vivant dans des milieux diversifiés.

La bactérie Caulobacter crescentus est une bactérie à Gram négative qui vit dans un environnement pauvre en aliments nutritifs. Seulement un des trois mécanismes identifiés chez

E. coli pour contrôler la réplication des chromosomes existe chez C. crescentus. Cette dernière encode une protéine (HdaA) avec homologie à la protéine Hda d’E. coli. C. crescentus utilise

également de nouveaux mécanismes de régulation qui ne se trouvent pas chez E. coli, afin de maîtriser l’activité de DnaA. Contrairement à DnaA chez E. coli, celle de C. crescentus est instable dans les cellules en croissance. Elle se dégrade rapidement pendant la phase stationnaire et lorsqu’il y a carence en éléments nutritifs. Curieusement, C. crescentus contient

IX un deuxième gène (hdaB) qui encode une protéine ayant une homologie avec le domaine de liaison de l’ADN de DnaA. Le rôle du gène hdaB n’est pas encore déterminé.

Afin de déterminer le rôle de HdaA, j'ai créé une souche (GM3700) de C. crescentus avec hdaA exprimée sous le contrôle d'un promoteur qui dépend de la présence de xylose. Je démontre que C. crescentus dépend du mécanisme RIDA afin d’empêcher les initiations multiples de la réplication des chromosomes dans un cycle cellulaire. Bloquer l'expression de hdaA provoque une augmentation de la fréquence d’initiation de la réplication des chromosomes ainsi qu'un blocage de la division cellulaire. J'ai également découvert un rôle inattendu pour HdaA en lien avec la stabilité de DnaA. En l’absence de la protéine HdaA, la protéine DnaA se stabilise dans les cellules en croissance et n'est pas dégradée en phase stationnaire. Mes expériences ont identifié un nouveau rôle pour HdaA dans la protéolyse de

DnaA. J'ai aussi découvert que HdaA participe à la transition programmée des cellules en croissance exponentielle à la phase stationnaire.

J'ai identifié un deuxième gène pour C. crescentus avec homologie à dnaA que j'ai nommé hdaB. Il affiche une homologie significative avec le domaine IV de la DnaA, celui qui est responsable des contacts avec l’ADN. L'analyse des séquences de hdaB suggère qu'il s'agit d'un paralogue de DnaA conservant les résidus principaux utilisés par DnaA pour faire des contacts de séquences spécifiques avec l’ADN. Le hdaB n’est pas essentiel à la croissance normale de C. crescentus en laboratoire car il peut être éliminé sans conséquence évidente pour la cellule.

Malgré la conservation évidente de la séquence avec le domaine de contact de l'ADN, des expériences avec des protéines purifiées in vitro démontrent qu’elles sont incapables de détecter les contacts entre HdaB et l’ADN. Ces résultats négatifs n'indiquent pas précisément

X un rôle du gène hdaB dans C. crescentus. Le fait que des homologues de hdaB se trouvent dans le génome d’autres alphaprotéobactéries suggère que hdaB donne un avantage aux bactéries qui n’apparaissent pas dans des conditions de laboratoire.

Pour mieux comprendre le lien entre RIDA et la dégradation de la protéine DnaA, J’ai créé quatre mutants de DnaA présentant des déficiences dans leurs activités ATPase. Dans la bactérie C. crescentus, l’expression de ces mutants donne un phénotype similaire aux cellules manquant d’HdaA: une plus haute fréquence d’initiation de la réplication des chromosomes ainsi que la stabilisation de DnaA. Ces résultats indiquent que la stabilité de DnaA est directement liée à son état d'activité et à la progression du cycle cellulaire.

Acknowledgements

I am extremely grateful for the opportunity to pursue my Ph.D. in the laboratory of Dr. Gregory

Marczynski. I have benefitted immensely from his guidance over the years and have learned many lessons that will continue to guide my future decisions both inside and outside of the laboratory.

I would also like to thank Dr. Benoit Cousineau and Dr. Maria Zannis-Hadjopoulos who have free given of their time to serve on my supervisory committee. I am appreciative of their support, encouragement and the constructive ideas they have provided during the course of my degree.

Many thanks go out to the past and present members of the Marczynski laboratory who have made going to work every day a true pleasure. Their scientific and technical expertise has been an invaluable resource to me. In particular I would like to thank Dr. James Taylor and Dr.

Gregory Marczynski for their critical reading of this thesis. I also thank Ms. Nadia Prud’homme for her assistance translating the abstract into French.

None of this would be possible without the constant love and support of my family. I am fortunate to have four great sisters Lise, Stasi, Mary, and Anna, two supportive parents, Yvonne and Michael, and a wonderful wife Nadia.

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Contributions to original knowledge

1. I employed genetic techniques to establish that the hdaA gene is essential in

Caulobacter crescentus.

a. I created a strain of C. crescentus with the only copy of the hdaA gene under the

control of a xylose inducible promoter

b. I have demonstrated that loss of HdaA leads to a loss of cell viability.

2. I demonstrated that C. crescentus utilizes the regulatory inactivation of DnaA (RIDA) to

limit chromosome replication to once per cell cycle.

3. I demonstrated the physiological and morphological consequences of blocked hdaA

expression, which include cell filamentation and the accumulation of unmethylated

chromosomal DNA.

4. I identified a link between RIDA and the stability of DnaA in C. crescentus. DnaA is

stabilized in the absence of the RIDA protein HdaA in log phase cells and is not cleared

from stationary phase cells.

5. I was able to stimulate the proteolysis of C. crescentus DnaA which had been stabilized

in stationary phase by the depletion of HdaA. This provides evidence that the

degradation of DnaA upon entry into stationary phase is not simply due to the depletion

of nutrients in the environment.

6. I demonstrated that the spoT gene which has previously been shown to be required for

the starvation induced degradation of DnaA is not required for the stationary phase

degradation of DnaA in C. crescentus.

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7. I created a strain of C. crescentus with a genetic deletion in the hdaB gene,

demonstrating that the gene is not essential

8. I identified homologs of the hdaB gene in other members of the alphaproteobacteria.

The conservation of the hdaB suggests that it may have a beneficial function.

9. I have provided the first direct evidence that C. crescentus DnaA binds ATP and ADP in

vivo.

10. I have shown that mutations in C. crescentus DnaAR300K and DnaAR357H both reduce

the intrinsic ATPase activity of DnaA in vitro.

11. I demonstrated the consequences of expressing the C. crescentus DnaAR357H, and

DnaAR357A mutants in vivo result in increased chromosome replication, cell

filamentation and improved stability of the DnaA protein.

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Chapter One Literature Review

Overview of the literature review The central theme of this thesis is the regulation of chromosome replication in Caulobacter crescentus. All living cells must replicate their genetic material and transmit their genes to their daughter cells. This essential process must also be precisely regulated in ways that have yet to be fully explored. During my Ph.D. I have explored the regulation of chromosome replication, using the model bacterium Caulobacter crescentus. In a growing cell there is a point during the cell cycle when the decision is made to begin chromosome replication; once replication has begun, the system needs to be reset to prevent the immediate re-initiation of chromosome replication. The main question that framed my studies is: How does C. crescentus limit chromosome replication to once per cell cycle?

The purpose of this literature review is to provide background material related to the topic of prokaryotic chromosome replication necessary to understand the significance of my work. Much of what is known about chromosome replication in prokaryotes was derived from work done on the model bacterium Escherichia coli. The first section summarises the E. coli model of chromosome replication. It will outline the steps involved in the initiation of chromosome replication. Particular emphasis will be placed upon DnaA, the highly conserved protein responsible for initiating chromosome replication, and the mechanisms that restrict chromosome replication to once per cell cycle. The following sections deal with variations on the theme of DNA replication, including structural differences found among bacteria with regard to the origin of replication. This includes bacteria that contain multiple chromosomes, and the replication of plasmid DNA in bacteria. This section ends with a brief examination of some of the exceptions to the otherwise universal requirement of DnaA for bacterial chromosome replication.

The next two main sections of this literature review deal with chromosome replication in the other two domains of life before returning to my model organism. I have compared the steps involved in chromosome replication between bacteria, archaea, and eukaryotes. The section entitled, “Caulobacter crescentus as a model for prokaryotic cell development” explains the choice of C. crescentus as my model organism for studying chromosome replication. I have described how C. crescentus relates to the other diverse members of the alphaproteobacteria, as well as their economic, scientific, environmental importance. This review finishes with a summary of the current state of knowledge on the regulation of the C. crescentus cell cycle.

This last section focuses on the master cell cycle regulators CtrA, GcrA, CcrM and DnaA, their regulation by proteolysis and the distinction between logarithmic growth and stationary phase.

These final topics are the most relevant to my studies.

Prokaryotic Chromosome Replication

The E. coli model

Research into the control of chromosome replication began with the early work on E. coli by

Jacob et al (1963). They proposed a replicon model consisting of two elements: i) a “replicator” site on the chromosome where replication initiates, and ii) the “initiator” that is the protein that begins the process of chromosome replication (Jacob and Brenner 1963). The replicator was later designated as the origin of chromosome replication (oriC) but originally it was inferred from indirect genetic criteria. For example, the replicator was identified in E. coli by its inability to permit the integration of F plasmids on the chromosome and thereby blocking the creation of Hfr strains. We now know that the chromosome does not tolerate a second origin of replication carried by the F plasmid. Therefore, the replicator was originally genetically designated as poh+ (permissive on Hfr) (Hiraga 1976). The confirmation of oriC as the replicator came from its ability to support the extra-chromosomal replication of DNA (Messer, Bergmans et al. 1978, von Meyenburg, Hansen et al. 1978, Messer, Meijer et al. 1979). The initiator was identified as the protein DnaA, which was shown to be essential for both in vivo and in vitro replication of DNA (Fuller, Kaguni et al. 1981, Tabata, Oka et al. 1983). The primary mechanism governing chromosome replication in E. coli derives from the sequence specific contacts between the initiator DnaA and the replicator oriC.

The E. coli origin of chromosome replication oriC The majority of our knowledge of chromosome replication comes from the model bacterium E. coli (Weigel and Messer 2001). E. coli chromosome replication initiates from a singular point on the chromosome, oriC, and replication then proceeds bi-directionally. It is argued that bacteria regulate chromosome replication by regulating the frequency of replication initiation from oriC.

Once chromosome replication has begun, the cell has committed itself to completely replicating the chromosome as a partially replicated chromosome would be fragile and potentially detrimental to the cell. Therefore, chromosome replication must be regulated to preclude multiple initiations within one cell cycle and oriC must be a key regulatory point.

The minimum essential length of E. coli oriC (Figure 1) is approximately 250 base pairs

(Fuller, Funnell et al. 1984). E. coli oriC can be divided into two functional domains: i) the DNA domain involved in binding and assembling DnaA proteins into a structured complex and ii) the

DNA domain termed the “unwinding element” (DUE) where double-stranded oriC first becomes single-stranded (Bramhill and Kornberg 1988). DNAase I footprinting was used to identify DnaA binding sites located in oriC (Matsui, Oka et al. 1985). These sites were called DnaA R boxes and consist of a 9 base pair sequence (5`-TTATNCACA-3`) which is recognized by DnaA. There are five DnaA R boxes found in oriC, R1, R2, R3, R4 and R5 (Figure 1). The sites R1, R2 and R4 are high-affinity binding sites for DnaA and interestingly, they are occupied throughout most of the cell cycle. In contrast, R3 and R5 have a lower affinity for DnaA (Schaper and Messer 1995,

Margulies and Kaguni 1996) and these low-affinity sites are only occupied by DnaA proteins during the initiation of chromosome replication (Miller, Grimwade et al. 2009). Other sites of

DnaA binding have also been identified in E. coli oriC. These sites bind DnaA with even lower affinity, and are termed I1, I2, and I3 as well as two τ sites, τ1 and τ2 (Figure 1). These lower

5 affinity sites are preferentially bound by DnaA-ATP in vitro and they are occupied only at the time of initiation in vivo (McGarry, Ryan et al. 2004, Kawakami, Keyamura et al. 2005). As explained further below, DnaA is bound by ATP (DnaA-ATP) or by ADP (DnaA-ADP). When the

DnaA-ATP form is present at a sufficiently high concentration, DnaA binds cooperatively to the lower affinity oriC sites, causing a melting of the DNA-duplex in the adjacent AT-rich DNA

Unwinding Element (DUE) (Kowalski and Eddy 1989, Erzberger, Mott et al. 2006). DnaA-ATP then stabilizes the open complex by binding the single stranded DNA (Speck and Messer 2001,

Duderstadt, Mott et al. 2010). At this time, DnaA-ATP also recruits proteins required for DNA replication to the oriC (Abe, Jo et al. 2007, Mott, Erzberger et al. 2008). The initiation of chromosome replication is a highly regulated process that requires specific steps.

Figure 1: Variation among the prokaryotic origins of chromosomal replication

Illustration of the organization of the origins of chromosome replication of Escherichia coli,

Caulobacter crescentus, Bacillus subtilis, Thermus thermophilus, and Streptomyces lividans. The positions of the AT-rich DNA unwinding element (DUE) is indicated by an orange box. The positions of the DnaA boxes are indicated by arrow heads. The E. coli strong DnaA boxes (R1 and R4), the Weak DnaA boxes (R2, R2, R3,I1, I2, I3, τ1, and , τ2) are indicated by blue and purple arrow heads respectively. The location of binding sites for IHF and FIS are indicated by yellow and green boxes. The details about these protein binding sites are described in the text.

The C. crescentus strong DnaA (G1, and, G2) are indicated by blue arrow heads and the weak

DnaA boxes are (w1, w2, w3, w4, w5, and, w6) are indicated with purple arrow heads. The binding site for IHF is indicated by a yellow bar. The binding sites for the master cell cycle regulator CtrA are indicated as red bars.

Figure 1: Variation among the prokaryotic origins of chromosomal replication

The steps in the initiation of chromosome replication A highly choreographed series of protein-DNA, as well as protein-protein interactions are required to initiate DNA replication from oriC. As described in Figure 2, the initiation of chromosome replication can be divided into a series of discrete steps i) binding of DnaA to high affinity DnaA binding sites, ii) cooperative binding of DnaA to low affinity DnaA binding sites, iii) the unwinding of the DNA in the AT-rich region by DnaA, iv) the recruitment of the replication machinery to the origin of replication.

As noted above, throughout most of the cell cycle, DnaA occupies the high-affinity R1,

R2 and R4 sites in oriC (Samitt, Hansen et al. 1989). These DnaA proteins seem to serve as markers for oriC. The E. coli oriC contains binding sites for other proteins which regulate chromosome replication (Figure 1). The protein FIS (factor for inversion stimulation) is also bound to oriC for the majority of the cell cycle (Cassler, Grimwade et al. 1995). The initiation of

DNA synthesis is concomitant with a reduction in FIS binding, an increase in IHF (integration host factor) protein binding and an increase in DnaA binding at oriC to include the R boxes R5 and R3 and the lower-affinity I sites (Cassler, Grimwade et al. 1995, Ryan, Grimwade et al.

2002). It is at this point that DnaA now associated with ATP forms a right handed helix which stimulates the unwinding of the DNA at the AT rich DUE (Hwang and Kornberg 1992, Erzberger,

Mott et al. 2006). The creation of single stranded DNA at oriC, is termed open complex formation and is necessary for the loading of the replication machinery in subsequent steps.

The open complex is stabilized by the binding of both DnaA (Speck and Messer 2001,

Duderstadt, Mott et al. 2010) and SSB (single stranded DNA-binding protein)(Krause and

Messer 1999).

DnaA next directs the entry and the loading of the helicase DnaB onto the single stranded DNA (Marszalek and Kaguni 1994, Abe, Jo et al. 2007). DnaB is loaded onto the single stranded DNA by contacts with DnaA and by the action of the ATP-dependent helicase loader protein DnaC (Mott, Erzberger et al. 2008). This action forms the “pre-priming oriC complex”

(Fang, Davey et al. 1999, Carr and Kaguni 2001). DnaB forms a homohexameric ring that stably complexes with six molecules of DnaC (Wahle, Lasken et al. 1989). The DnaBC complex requires DnaC to be bound to ATP (Wahle, Lasken et al. 1989). The ATPase activity of DnaC is stimulated by interaction with single stranded DNA and DnaB, resulting in DnaC-ADP which removes the inhibition of the helicase activity of DnaB (Learn, Um et al. 1997, Davey, Fang et al.

2002). The DnaB helicase is positioned at the apex of the replication fork expanding the unwound region (LeBowitz and McMacken 1986) and interacting with DnaG primase (Tougu and Marians 1996).

The term “replisome” describes the group of proteins required for replication fork progression. In addition to the DnaG primase, the assembly of the replisome includes the loading of a so called “sliding clamp”. The sliding clamp is made up of two DnaN proteins that form a complete ring around the unwound DNA strands. DnaN can bind and thereby “clamp” several proteins to the DNA (described further below) including the replication fork DNA polymerase III. The DNA polymerase III holoenzyme extends the primers synthesized by DnaG primase and replication proceeds outward from the origin. The DnaN sliding clamp is required for processivity of this polymerase. In the absence of the sliding clamp, the core DNA polymerase III is very slow, it incorporates only ~20 nucleotides per second and it is weakly processive, it extends only ~10 nucleotides before detaching from the template DNA (Maki and

Kornberg 1985). Binding to the DnaN sliding clamp greatly enhances the speed of DNA polymerase III to approximately 750 nucleotides per second as well as increasing its processivity to greater than 50 kilobases before detaching from the template (Stukenberg, Studwell-

Vaughan et al. 1991). Interestingly, the DnaN protein also binds and tethers other proteins to the DNA including the Hda protein that regulates DnaA activity and that will be described further below (Kato and Katayama 2001).

Figure 2: Steps involved in initiation of Chromosome replication in E. coli

DnaA bound to ADP (red balls) is found associated with the high affinity DnaA boxes (blue arrow heads) in the E. coli origin of replication throughout most of the cell cycle. When the concentration of DnaA bound to ATP (blue balls) reaches a critical level DnaA binds cooperatively to the low affinity DnaA binding sites which stimulates the unwinding of the DNA in the AT-rich DNA unwinding element, DUE (orange box) forming an open complex. The single stranded DNA is stabilized by DnaA. The DnaB helicase (green ovals) expands the open complex, recruiting the DnaG primase which synthesizes a RNA primer. The DnaN sliding clamp ensures processivity as the DNA polymerase III replicates the chromosome from the RNA primer.

Figure 2: Steps involved in initiation of Chromosome replication in E. coli

Diversity among bacterial origins of chromosome replication The origin of replication is defined as the location on the chromosome where replication is initiated. Unlike higher eukaryotic origins, bacterial origins are usually well defined at the DNA sequence level. There is a considerable variation in the size and organization of bacterial origins of chromosome replication (Figure 1). However, they possess a number of characteristic functional elements in common, including specific binding sites for the initiator protein (DnaA), binding sites for accessory replication proteins and regions with a high content of adenine and thymine residues (AT-rich).

The most abundant repeated sequences found in bacterial origins are the DnaA boxes.

Although DnaA is widely conserved, the number and arrangement of DnaA boxes varies greatly between bacteria (Zawilak-Pawlik, Kois et al. 2005). In nearly all bacteria the origin consists of a single cluster of DnaA boxes except for Bacillus subtilis and related bacteria. The B. subtilis origin is divided into two regions, 5’ and 3’ to the dnaA gene (Ogasawara, Moriya et al. 1985).

The initiation of B. subtilis chromosome replication requires both 5’ and 3’ DnaA box regions

(Moriya, Atlung et al. 1992). However, the dnaA gene may be displaced from the origin as long as the DnaA protein in complemented in trans (Moriya, Atlung et al. 1992). In contrast, the

Thermus thermophilus chromosome is replicated from a symmetrical origin which is located between the dnaA and dnaN genes. The origin of T. thermophilus contains 13 symmetrically arranged DnaA boxes (Schaper, Nardmann et al. 2000). As another example, the large linear chromosomes of Streptomyces species are replicated from a centrally positioned origin, which contains 19 DnaA boxes whose location and orientation is conserved (Jakimowicz, Majka et al.

1998). As will be described further below, not all bacterial replication origins have such conspicuous clusters of DnaA boxes.

DnaA independent chromosome replication In addition to the normal mode of chromosome replication mediated by DnaA binding to the oriC, E. coli possesses at least three alternate modes of chromosome replication, that are normally repressed, but which can be activated with the right stimulus. Unlike the normal mode of chromosome replication, theses alternate modes do not rely on DnaA to unwind the origin. These three alternate modes are called the inducible stable DNA replication

(iSDR)(Kogoma and Lark 1970), the constitutive stable DNA replication (cSDR), and the nutritional shiftup- activatable stable DNA replication (nSDR) (Hong, Cadwell et al. 1996). These three modes are differentiated by the manner by which they unwind the DNA at first step in chromosome replication. They also differ from the DnaA mode, because they initiate replication outside of oriC.

iSDR is a form of recombination-dependent DNA replication that is activated by SOS induction (Kogoma, Torrey et al. 1979). It is believed that iSDR initiates replication from a DNA structure termed a “D-loop” which is also a recombination intermediate created by the single strand DNA insertion into duplex DNA (Asai, Imai et al. 1994).

cSDR is activated by the loss of RNaseH activity (Horiuchi, Maki et al. 1984). E. coli possessing mutations in the rnhA gene can overcome the need for chromosome replication from oriC, resulting in cells that retain viability despite deletions in the dnaA gene or the disruption of the oriC locus (Kogoma and Vonmeyenburg 1983). It is believed that cSDR initiates replication from an RNA-DNA structure termed an “R-loop” which remains in the DNA duplex if RNA is not for example displaced by transcribing RNA polymerase.

nSDR is also a non SOS dependent stable replication that is observed in rapidly growing wild type cells upon entry into stationary phase (Hong, Cadwell et al. 1996). The inhibition of

15 chromosome replication by chloramphenicol is transiently overcome when E. coli is shifted from minimal medium to rich media (Luria broth). Like iSRD, nSDR absolutely requires recA, but replication does not require recB, or dnaA (Hong, Cadwell et al. 1996). These modes of chromosome replication are considered as emergency systems of last resort and they are not a part of the normal growth and cell cycle, as is the DnaA mode.

Bacteria with multiple chromosomes

Early work done by Jacobs et al. (1963) established the E. coli model of the bacterial genome consisting of a single circular chromosome, with a single “replicator” now termed the origin of replication (Jacob and Brenner 1963). Since then, many bacteria have been identified with multiple and/or some with linear chromosomes. The distinction between a mega-plasmid and a true chromosome is based upon the presence of essential genes. This distinction is fundamentally arbitrary, since it can result from the presence of a single essential open reading frame (orf). The distribution of genes is often not divided equally between the two chromosomes; the majority of essential genes are located on one dominant chromosome while the second chromosome often contains only a few essential genes. There are several mechanisms by which a bacterium could have acquired a secondary chromosome. The second chromosome could be the result of a duplication of the primary chromosome, the result of the primary chromosome being divided into two autonomously replicating units, or the second chromosome may have evolved from a plasmid which has acquired essential genes. The non- essential plasmid origin theory is supported by the relatively few essential genes usually found

16 on the second chromosome as well as the fact that many secondary plasmids contain plasmid- like origins of replication (Egan, Fogel et al. 2005).

However, no matter how they might have evolved, bacteria with multiple chromosomes must coordinate the replication of the individual chromosomes to ensure that all of the genes are faithfully replicated. Chromosome replication is linked with the cell cycle to ensure that chromosome replication occurs once per cell cycle, while autonomous plasmid replication may occur more than once per cell cycle. Small plasmid replicons are structured so as to maintain the plasmid copy-number and this does not need strict cell cycle coordination. This difference poses a challenge for bacteria with two chromosomes of unequal size. For example, Vibrio cholerae overcomes this challenge by somehow delaying replication of the smaller chromosome (Rasmussen, Jensen et al. 2007). Bacteria may have evolved a number of mechanisms of coordinating replication between multiple chromosomes. One possibility is that the two chromosomes may use completely different mechanisms to ensure that each is replicated and this ensures that the two origins of replication do not interfere with each other.

Alternatively, the two chromosomes may share a common regulatory mechanism, which ensures that initiation occurs synchronously for both chromosomes. It is also possible that the chromosomes use a combination of common and unique regulatory proteins but mechanistic details have yet to be understood.

Regarding our model, although C. crescentus has only one chromosome, several related alphaproteobacteria have multiple chromosomes, including Agrobacterium tumefaciens

(Goodner, Hinkle et al. 2001), and, Brucella suis (Paulsen, Seshadri et al. 2002). The larger chromosome of each of these bacteria contains an origin or replication similar to that of C.

17 crescentus (described below) while the smaller chromosome contains a repABC-like origin similar to those found on plasmids (also described below).

Replication of linear bacterial chromosomes The development of pulse-field gel electrophoresis allowed for the separation and visualization of very large fragments of DNA (Schwartz and Cantor 1984). The ability to separate whole bacterial chromosomes lead to the identification of a linear chromosome in Borrelia burgdorferi, the causative agent of Lyme disease (Ferdows and Barbour 1989). Since then, linear chromosomes have been identified in several unrelated bacteria, including, Streptomyces lividans (Lin, Kieser et al. 1993) and Agrobacterium tumefaciens (Goodner, Markelz et al. 1999).

The replication of linear chromosomes is similar to that of circular chromosome. Replication is initiated from a single origin of replication that is generally located at the center of the linear chromosome. In B. burgdorferi the origin of replication was mapped to a 240 bp region between dnaA and dnaN positioned in the center of the chromosome (Picardeau, Lobry et al.

1999). Similarly, quantitative DNA hybridization analysis was used to locate the origin of replication near the dnaA-gyrB region located at the centre of the linear chromosome of

Streptomyces coelicolor (Musialowski, Flett et al. 1994). These origins of replication resemble those of other bacteria, so linear chromosomes appear to initiate replication by the same DnaA mode as do their circular counterparts.

However, linear chromosomes present special problems. DNA synthesis requires priming and it only occurs in a polar 5’ to 3’ fashion. These constraints create gaps at the linear ends of chromosomes. Bacteria have evolved at least two mechanisms to completely replicate the ends of linear DNA. One mechanism to resolve this problem involves the formation of circular intermediates (Figure 3). B. burgdorferi replication of the linear DNA results in the

18 formation of circular dimer junctions (Casjens, Murphy et al. 1997). The replicated telomeres are recognized by the telomere “resolvase” (ResT), which resolves the dimer junctions into two closed hairpin telomeres (Chaconas, Stewart et al. 2001, Kobryn, Burgin et al. 2005). Therefore, this resolvase mechanism is an example of site-specific recombination mechanisms, e.g. phage integration/ excision reactions, that are very common among bacteria. A different mechanism evolved in the genus Streptomyces. It uses proteins which are covalently bound to the ends of linear chromosomes, as well as to their linear plasmids (Lin, Kieser et al. 1993). These

“telomere” associated proteins replicate the ends of chromosomes by terminal-protein-primed

DNA synthesis (Qin and Cohen 1998).

Figure 3: Replication of linear chromosomes in bacteria

One method used by bacteria to replicate linear chromosomes involves the use of a circular intermediate step. One example of this type of chromosome replication occurs in the bacterium Borrelia burgdorferi. The linear chromosome of B. burgdorferi ends are sealed with hairpin ends, replication begins at a single centrally located origin of replication. Chromosome replication results in the creation of a circular chromosome which is then resolved into two identical linear chromosomes.

Plasmid replication By definition plasmids do not carry genes essential for the survival of the cell under non- stressed conditions, but rather contain genes which confer a selective advantage such as antibiotic resistance or the ability to utilise a novel food source. Plasmids provide a means of genetic exchange between bacteria. A plasmid consists of a double-stranded circular or a linear

DNA molecule capable of autonomous replication (Leplae, Hebrant et al. 2004). Plasmid replication is regulated such that the plasmid is maintained at a characteristic copy number

(Kollek, Oertel et al. 1978), which can range from one copy to thousands of copies per cell. The minimal portion of the plasmid required to maintain the characteristic copy number is termed the replicon. The replicon contains the origin of replication (ori) as well as specific replication initiator genes and their regulators. Several families of E. coli plasmids have been identified which utilise DnaA as an initiator of replication. The participation of DnaA in the unwinding of the origin and initiation of replication varies depending upon the plasmid type. DnaA acts cooperatively with plasmid encoded initiator proteins. The organisation of the plasmid origin of replication varies between different plasmid types, however, they will generally contain one or more DnaA binding sites, an AT rich region where unwinding of the DNA occurs, and binding sites for plasmid encoded initiator proteins. repABC family of plasmids The presence of repABC operons is widespread among high molecular weight, low copy plasmids in the alphaproteobacteria. Examples include the tumour inducing (Ti) plasmid of

Agrobacterium (Tabata, Hooykaas et al. 1989) and the symbiotic plasmid (pSym) of Rhizobium

(Mazur, Majewska et al. 2011). The operon consists of three genes in the same order repA, repB, followed by repC. The region between repB and repC is highly conserved and it contains a

21 small antisense RNA (57-59 nucleotides). Mutations in the promoter region of the antisense

RNA abolished the plasmid incompatibility phenotype (Venkova-Canova, Soberon et al. 2004,

Chai and Winans 2005, MacLellan, Smallbone et al. 2005). The anti-sense RNA has also been shown to be a negative regulator of the repC gene (Chai and Winans 2005). The repA and repB genes encode homologs of the ParA and ParB plasmid partitioning proteins (Bignell and Thomas

2001), while RepC is the initiator of replication. The origin of replication maps to within the repC gene (Ramirez-Romero, Soberon et al. 2000). Disruption of either repA or repB (Ramirez-

Romero, Soberon et al. 2000) results in a change in copy number, while disruption of repC completely abolishes plasmid replication (Tabata, Hooykaas et al. 1989).

The repABC operon is not restricted to the replication of repABC plasmids. The repABC system is also used to replicate the second chromosome found in some of the alphaproteobacteria. For example, the second chromosome of Agrobacterium vitis S4 and

Agrobacterium tumefaciens C58 are replicated using a repABC origin of replication (Slater,

Goldman et al. 2009).

DnaA - Structure and function

Since the DnaA protein is responsible for initiating chromosome replication in the vast majority of bacteria, I will now describe its structure and biochemical properties. The crystal structures of DnaA domains, summarized in Figure 4, have greatly improved our understanding of how

DnaA functions both as an initiator of chromosomal replication and as a transcriptional regulator of genes involved in chromosome replication. DnaA is a member of an ATPase family termed ATPase associated with various cellular activities (AAA+) proteins (Neuwald, Aravind et

22 al. 1999). First based on DNA alignment and then based on functional studies, the DnaA protein can be regarded as having four functional domains (I, II, III, IV) (Sutton and Kaguni 1997).

DnaA domain I

Domain I (amino acids 1 to 90 in E. coli) of DnaA contains three α helices and three β sheets

(Abe, Jo et al. 2007) and is involved in protein-protein interactions, with multiple proteins including the helicase DnaB, a DnaA-binding protein DiaA (that promotes DnaA self- association), as well as other DnaA proteins. Oligomerization of DnaA at the origin of replication is necessary for the initiation of chromosome replication (Simmons, Felczak et al.

2003). The expression of a truncated DnaA protein consisting of only domain I (amino acids 1 to

86) had an inhibitory effect on chromosome replication and was sufficient to repress the over- initiation of chromosome replication of a dnaA219(Cos) mutant in vivo (Weigel, Schmidt et al.

1999).

Domain I of DnaA is sufficient and essential for the homo-dimerization of DnaA. A ‘one hybrid assay’ has been used to show that domain I is able to functionally replace the dimerization domain of coliphage lambda cI repressor (Weigel, Schmidt et al. 1999). Deletion of Domain I from DnaA renders it incapable of supporting replication from oriC. Domain I is not as conserved as domain III and IV among DnaA orthologs, only key amino acids such as Trp6 are conserved (Felczak, Simmons et al. 2005, Abe, Jo et al. 2007). The Trp6 residue has been identified as being required for domain I function (Abe, Jo et al. 2007). For example, Trp6 which is required for homo-dimerization of DnaA is also needed for loading of DnaA (Felczak,

Simmons et al. 2005, Abe, Jo et al. 2007).

DnaA-DnaA contacts in domain I also help to recruit the DnaB helicase. Both Domains I and III are believed to be necessary for loading of the DnaB helicase(Seitz, Weigel et al. 2000).

The Glu21 residue located in domain I of DnaA is also required for loading of DnaB (Abe, Jo et al. 2007). These observations argue that the loading of DnaB is dependent upon a stable formation of DnaA multimers created during oriC initiation (Felczak, Simmons et al. 2005).

DnaA domain II

Domain II (amino acids 90 to 130 in E. coli) is a flexible linker domain which allows domain I and

III to move independently of each other (Abe, Jo et al. 2007). In support of this interpretation, the sequence of domain II is the least conserved domain among DnaA homologs (Skarstad and

Boye 1994, Messer 2002). Sections of 30 to 36 consecutive amino acids can be deleted from various positions of domain II without loss of function of the DnaA protein. A minimum of 21 to

27 amino acids are required to keep domains I and III separated (Nozaki and Ogawa 2008).

Interestingly, a GFP (Green Fluorescence Protein) tag can be inserted into this domain without loss of viability (Boeneman, Fossum et al. 2009). This GFP-tagged DnaA has been used to study protein sub-cellular localization in E. coli and apparently DnaA multimerizes to form a helical structure.

DnaA domain III

Domain III (amino acids 131 to 346 in E. coli) is a highly conserved domain involved in ATP binding and DnaA oligomerization. The crystal structure of domain III of Aquifex aeolicus reveals structural similarities with both archaeal and eukaryotic initiators of chromosome replication

(Erzberger, Pirruccello et al. 2002). Domain III contains the motifs which group DnaA into the

AAA+ superfamily of proteins. The AAA+ superfamily is a diverse group of ring-shaped P-loop

NTPases. These proteins typically include the walker A and B motifs, a sensor 1 motif, Box VII arginine finger and Box VIII sensor 2 motifs (Neuwald, Aravind et al. 1999). AAA+ proteins exert energy in the form of conformational change to perform a wide range of cellular processes, including DNA replication, degradation of damaged or misfolded proteins, microtubule dynamics and membrane fusion.

Domain III of DnaA has high affinity for both ATP and ADP with a KD of 0.03μM and

0.1μM, respectively (Sekimizu, Bramhill et al. 1987). The structure derived from the bacterium

A. aeolicus predicts that DnaA can self-assemble into a right-handed helical filament when it is bound by ATP but not when DnaA is bound by ADP (Erzberger, Mott et al. 2006, Duderstadt,

Mott et al. 2010). These structures support earlier proposals based on biochemical studies that the nucleotide bound to DnaA is acting as a switch to alter the DnaA oligomerization state and in turn the assembly of DnaA on oriC DNA. The hydrolysis of ATP does not provide energy for the unwinding reaction because a non-hydrolysable ATP analog (ATP-gamma-S) is able to function in vitro to unwind the oriC in the absence of ATP hydrolysis (Sekimizu, Bramhill et al.

1987).

Several amino acid residues have been identified as important for ATP binding and hydrolysis. The binding of ATP and ADP requires Asp269 located in the sensor 1 motif

(Kawakami, Ozaki et al. 2006). Amino acids which are specifically important in the initiation activity of DnaA include the Box VII arginine finger Arg285 as well as the B/H motif residues.

While Box VIII sensor 2 Arg334 is not required for the initiating activity of DnaA, it is required for the ATPase activity of DnaA. The DnaAR334A mutant is unaltered in its ability to bind both

ATP and ADP, however it is inert to the hydrolysis of bound ATP (Nishida, Fujimitsu et al. 2002).

ATP/ADP binding and ATP hydrolysis have obviously important consequences for DnaA regulation which will be described below in the studies presented in this thesis.

DnaA domain IV

Domain IV (amino acids 347 to 467 in E. coli) is the domain which makes sequence specific contacts with DNA. As noted above, DnaA binds to DNA most strongly at a specific 9 base pair element called a DnaA R box (5`-TTATNCACA-3`) and located within the 260 base pairs of the E. coli oriC are 5 DnaA R boxes (Fuller, Funnell et al. 1984).

The protein-DNA co-crystal structure of DnaA protein with a DnaA R box has been solved. The helix-turn-helix motif of domain IV makes contacts along both the major and minor grooves of the DnaA R box (Fujikawa, Kurumizaka et al. 2003). Several residues important for making sequence specific contacts have been identified including DnaA-Arg339 which contacts the DNA minor groove to recognize the TTA sequence of the TTATNCACA R box consensus sequence. The TNCACA sequence of the R box is recognized by a helix inserted in the DNA major groove (Fujikawa, Kurumizaka et al. 2003). Mutations in residues DnaA-Asp433, His434 and Thr435, cause a defect in DnaA R box recognition without affecting non-specific DNA binding (Sutton and Kaguni 1997, Blaesing, Weigel et al. 2000). The binding of DnaA to the

DnaA box introduces a bend of approximately 40 degrees (Schaper and Messer 1995). While it is clear that Domain IV binds the R box, it not yet clear how DnaA makes the other DNA sequence-specific contacts during the initiation of chromosome replication.

The domain organization of DnaA will help us to understand how it functions to initiate chromosome replication and thereby suggest how initiation can be regulated. Interestingly,

26 regulators of DnaA themselves correspond to DnaA protein domains and these will be described and studied further below.

Figure 4: The domain structure of DnaA

DnaA is composed of four structural and functional domains. Panel A shows a linear diagram of

E. coli DnaA. Domain I (blue) is primarily responsible for protein-protein interactions, Domain II

(green) is the least conserved domain and is believed to be a flexible linker domain, Domain III

(orange) contains the AAA+ ATPase domain, and Domain IV (purple) contains the helix-turn- helix DNA binding domain. The amino acids (based upon E. coli) which begin and end each

Domain are indicated above the diagram. Amino acids with key regulatory functions are indicated below the diagram. Panel B shows the crystal structures below the corresponding linear domains. Domains I and II are represented by the crystal structure (2E0G) which was obtained from E. coli, domain II appears as a filament due to lack of secondary structure, the structure domain III (2Z4S) was determined from the DnaA of Thermotoga maritima (Ozaki,

Kawakami et al. 2008), the structure of the DNA binding domain ( 1J1V) of DnaA from E. coli domain IV is shown in association with a double stranded DnaA box (Fujikawa, Kurumizaka et al. 2003).

Figure 4: The domain structure of DnaA

DnaA interacting proteins The activity of DnaA protein is regulated by multiple protein-protein interactions, including

DnaA multimerization at the origin of replication, interaction with the proteins of the replication machinery and regulatory proteins which can either enhance or inhibit DnaA activity.

DiaA/HU/HobA In E. coli, the protein DiaA (DnaA initiator-associating factor) is required for timely initiation of chromosome replication during the cell cycle. Loss of diaA supresses the over-initiation phenotype caused by the dnaAcos mutant (Ishida, Akimitsu et al. 2004). The histone-like dimeric protein HU has also been shown to influence the activity of DnaA at oriC (Dixon and

Kornberg 1984). The HU protein is composed of 2 alpha or one alpha and one beta subunits during early to mid-log phase or 2 beta subunits during stationary phase (Claret and Rouviere-

Yaniv 1997). Both HU (alpha dimer or the alpha-beta heterodimer) and DiaA interact with the

N-terminus of DnaA to promote DnaA oligomerization at oriC (Chodavarapu, Felczak et al. 2008,

Keyamura, Abe et al. 2009).

A yeast two-hybrid screen looking for proteins which interact with DnaA in the human pathogen Helicobacter pylori, identified the protein HP1230 (HobA) which complexes with

DnaA (Rain, Selig et al. 2001). The interaction of HobA with DnaA was confirmed in vitro and later in vivo (Terradot, Durnell et al. 2004, Zawilak-Pawlik, Kois et al. 2007). HobA is essential for cell growth, since depletion of HobA leads to growth arrest and a failure to initiate chromosome replication (Zawilak-Pawlik, Kois et al. 2007). The crystal structure of HobA revealed that it is a structural homolog of DiaA from E. coli, despite lacking any obvious sequence homology (Natrajan, Hall et al. 2007).

YabA/Hda The activity of DnaA may also be negatively regulated through protein-protein interactions. The dominant mechanism in E. coli is the regulatory inactivation of DnaA (RIDA) (Camara, Breier et al. 2005). The intrinsic ATPase activity of DnaA is stimulated by the interaction of DnaA with the

Hda carried by DnaN (also the β-subunit of DNA polymerase III). The Hda protein displays homology to the AAA+ domain of DnaA. Hda was initially identified in a genetic screen for high copy suppressors of a dnaN36 mutant (Kato and Katayama 2001). The Hda mediated inactivation of DnaA is an important process preventing over-initiation since Hda deficient E. coli over-initiates replication from oriC (Camara, Breier et al. 2005). This thesis extends our knowledge of Hda in C. crescentus and Hda will be described further below.

Bacillus subtilis genome-wide two-hybrid screens predicted the interaction of the protein YabA with both DnaA and DnaN (Noirot-Gros, Dervyn et al. 2002). Like Hda of E. coli,

YabA is also a negative regulator of DnaA and YabA is involved in the timing of the initiation of chromosomal replication. Inactivation of YabA or mutations which alter its binding to DnaA or

DnaN result in extra initiation of chromosome replication (Hayashi, Ogura et al. 2005, Noirot-

Gros, Velten et al. 2006). While Hda and YabA proteins appear to be similar, since they both use DnaN to contact DnaA, they are in fact completely different proteins, that evolved independently and that use different mechanisms to inactivate DnaA. The Hda mechanism stimulates DnaA-ATP hydrolysis (described further below). In contrast, the YabA mechanism apparently disrupts DnaA oligomerization and assembly on the replication origin(Merrikh and

Grossman 2011).

DnaA as a transcription factor In addition to its role in initiating chromosome replication, DnaA also acts as a cell cycle transcriptional regulator. Several bacteria, including E. coli (Messer and Weigel 1997), C. crescentus (Hottes, Shapiro et al. 2005) and B. subtilis (Burkholder, Kurtser et al. 2001), use

DnaA as a transcription factor to coordinate chromosome replication with other events in the cell cycle, including the transcription of genes that support DNA synthesis and cell division.

DnaA can either activate or repress various genes including dnaA itself. For example, the E. coli dnaA gene is auto-regulated (Atlung, Clausen et al. 1985, Braun, O'Day et al. 1985, Kucherer,

Lother et al. 1986, Wang and Kaguni 1987). The dual nature of DnaA as both a transcription factor and the initiator of chromosome replication allows it to coordinate chromosome replication with the expression of genes needed for DNA replication. The E. coli genome carries more than 3,000 high affinity DnaA binding sites (Roth and Messer 1998). One important example of this regulation is the enzyme ribonucleotide reductase, which catalyzes the last step in the formation of deoxynucleotides (dNTPs) required for DNA replication (Nordlund and

Reichard 2006). DnaA will either activate or repress transcription from the nrdAB operon depending on its nucleotide-bound state (Olliver, Saggioro et al. 2010). The conversion of

DnaA-ATP to DnaA-ADP by Hda following the initiation of chromosome replication is believed to synchronize the expression of the nrdAB operon with the onset of chromosome replication

(Gon, Camara et al. 2006). Consequently, E. coli synthesizes more dNTPs when they are needed for chromosome replication.

AAA+ proteins -Structure and function As noted above, the largest domain III of DnaA belongs to the AAA+ family, so we can gain functional insights from related proteins. The AAA+ (ATPase associated with various cellular activities) family is a diverse group of proteins, which contain a 200 to 230 amino acid ATP binding domain (AAA domain) (Patel and Latterich 1998). The AAA+ proteins are macromolecular machines which use the energy of ATP hydrolysis to remodel target substrates in various ways. AAA+ proteins have been shown to be involved in disassembly of protein complexes, the unfolding and degradation of proteins, unwinding of DNA, or alteration of the state of DNA-protein complexes (Ogura and Wilkinson 2001). The AAA+ group of proteins includes proteins involved in DNA replication, including the prokaryotic initiator DnaA, the helicase DnaB, the helicase loader DnaC, the eukaryotic origin recognition complex (ORC) and the MCM DNA-licensing factors (Figure 6). Additional AAA+ proteins include the Bacillus sporulation protein SpoVJ, Mg2+, and Co2+ chelatases and dynein motor proteins. This multiplicity of functions is due to the variety of domains which are found associated with the

AAA domain.

Based upon sequence analysis and structural similarities the AAA family was expanded to become the AAA+ superfamily of proteins (Neuwald, Aravind et al. 1999). The AAA domain contains two defining features of the AAA+ class of proteins, a core αβα nucleotide-binding domain and a smaller α-helical domain consisting of two helical hairpins arranged in a left- handed super-helical structure. The AAA+ domains show a greater degree of conservation at the structural level than at the sequence level (Ogura and Wilkinson 2001, Ammelburg, Frickey et al. 2006).

The AAA+ domain contains the following distinct signature sequence motifs. These include the Walker A and Walker B motifs (Walker, Saraste et al. 1982). The Walker A motif has the consensus sequence GXXXXGK [S/T], where G is glycine, K is lysine, S is serine, T is threonine, and x is any residue . The Walker B motif has the consensus sequence hhhhDE, where h is any hydrophobic residue D is aspartate and E is glutamate (Hanson and Whiteheart

2005). A conserved region, positioned C terminal to the Walker B motif, is called the second region of homology (SRH). The SRH contains the sensor 1 motif as well as the Arginine fingers.

The role of the SRH is to coordinate nucleotide hydrolysis and conformational change between subunits (Ogura, Whiteheart et al. 2004). The SHR is not always apparent from inspecting the sequence alone but the structural features are conserved. The sensor 1 motif is present in all

AAA+ proteins and mutations result in impaired ATP hydrolysis (Steel, Harley et al. 2000).

Structural analysis of AAA+ proteins shows that the arginine fingers from one subunit contact the ATP binding site of the adjacent subunit. Mutation of the arginine finger residue alters ATP hydrolysis and/or the propagation of conformational changes (Ogura, Whiteheart et al. 2004).

The sensor 2 located near the C terminus contains a conserved arginine which participates in

ATP binding and hydrolysis (Joshi, Baker et al. 2003, Ogura, Whiteheart et al. 2004).

The majority of AAA+ proteins form closed ring structures (Ogura and Wilkinson 2001).

For example, the DnaB helicase forms a hexameric ring. Exceptions to this paradigm are the initiators of DNA replication, which form open helix structures (Erzberger, Mott et al. 2006).

Structural studies performed on the DnaA protein of Aquifex aeolicus shows that binding of

DnaA by ATP induces a conformational change which favors the formation of a right handed helix (Erzberger, Mott et al. 2006). This assembly mechanism may be conserved in all domains

34 of life since eukaryotic and archaea initiators also share similar structural motifs that promote open helix formation (Giraldo 2003, Clarey, Erzberger et al. 2006). A subset of the AAA+ proteins do not have an ATPase activity, while some do not have the ability to bind ATP at all.

These AAA+ family members however do form complexes with other family members that do have ATPase activity (Ogura and Wilkinson 2001).

Mechanisms used to limit initiation of chromosome replication to once per cell cycle

Electron microscopy has shown that approximately 20 monomers of DnaA are needed to form the pre-RC “pre-replication complex” of oriC (Crooke, Thresher et al. 1993). E. coli contains between 800 to 2100 molecules per cell of DnaA (Sekimizu, Yung et al. 1988). Over-expressing

DnaA in E. coli causes over-initiation of chromosome replication (Atlung, Lobner-Olesen et al.

1987, Skarstad, Lobner-Olesen et al. 1989), suggesting the amount of DnaA is rate limiting for the initiation of chromosome replication. However, the amount of DnaA in the cell greatly exceeds the amount required for the formation of the replication complex. Therefore, mechanisms must exist which limit the activity and/or accessibility of DnaA to oriC. In E. coli cells that were synchronized using a temperature sensitive dnaC mutant, the ratio of DnaA-ATP

/DnaA-ADP varied in a cell cycle dependent manner. The DnaA-ATP form peaks at about the time of initiation of replication (Kurokawa, Nishida et al. 1999). This indicates that the activity of DnaA is regulated during the normal E. coli cell cycle. E. coli possesses at least three mechanisms which prevent the over initiation of oriC replication, one which stimulates the intrinsic ATPase activity of DnaA and two which restrict DnaA binding to the origin of replication.

Sequestration of the origin of replication

Premature replication of the chromosome can be prevented by restricting access of DnaA to the oriC. Newly replicated origins are “sequestered” from DnaA and hence the whole replication machinery by binding the SeqA protein (Figure 5A). The sequestration of newly replicated origins requires specific DNA methylation. In E. coli the Dam methyltransferase methylates DNA at GATC sites. Prior to chromosome replication the GATC sites are methylated on both top and bottom DNA strands. However, the passage of a replication fork yields two

DNA molecules which are hemi-methylated until the Dam enzyme finds the un-methylated strands. The methylation of GATC sites is usually rapid (less than 1 min) at typical places on the chromosome, but at the origin of replication (oriC) and at the promoter of dnaA, the methylation of GATC sites is much slower (over 20 min). This delay is due to the sequestration of the hemi-methylated DNA by the SeqA protein. The SeqA protein preferentially binds to hemi-methylated DNA blocking access to the origin and to the promoter of dnaA (Taghbalout,

Landoulsi et al. 2000). Disruption of the seqA gene results in over-initiation of chromosome replication. Over expression of SeqA protein also supresses the over-replication phenotype associated with the DnaAcos mutant (that produces hyperactive DnaA-ATP).

Vibrio cholera encodes a seqA gene with homology (54% identity and 69% similarity to E. coli seqA). The SeqA protein is essential in V. cholera, unlike in E. coli. Excess SeqA in V. cholera leads to replication arrest and a cell division defect (Saint-Dic, Kehrl et al. 2008). This suggests the role SeqA in V. cholera may be different than in E. coli.

While homologs of SeqA have not been widely observed, DNA methylation nonetheless may be an important factor in cell cycle regulation. Several members of the

36 alphaproteobacteria subdivision of gram-negative bacteria including Rhizobium meliloti, Brucella abortus, Agrobacterium tumefaciens, and Rhodobacter capsulatus (Wright, Stephens et al.

1997), encode a cell cycle regulated DNA methyltransferase CcrM which has been shown to be essential in Agrobacterium tumefaciens (Kahng and Shapiro 2001), in Caulobacter crescentus

(Wright, Stephens et al. 1997) and in Brucella abortus (Robertson, Reisenauer et al. 2000).

Overexpression of CcrM results in defects in cell morphology and in the initiation of DNA replication in both C. crescentus and R. meliloti (Wright, Stephens et al. 1997), while overexpression of CcrM attenuates intracellular replication of B. abortus in murine macrophages (Robertson, Reisenauer et al. 2000). It remains unclear if these phenotypes are due to the CcrM methylation at the origin of replication or if they are due to CcrM methylation in the promoter regions of cell cycle transcriptional regulators such as CtrA (Reisenauer and

Shapiro 2002) and DnaA (Collier, McAdams et al. 2007), as described further below.

Titration of DnaA away from the origin of replication

This is another mechanism that restricts DnaA binding to oriC but instead of blocking access to oriC, in this case DnaA is diverted away from oriC by titration (Figure 5B). The datA locus has a very high affinity for DnaA, binding eight times more DnaA than the oriC (Kitagawa, Mitsuki et al. 1996). The datA locus is located near the origin of replication. Thus, early datA duplication serves to reduce the amount of free DnaA available for initiation.

A study which examined 120 completed sequences of bacterial genomes identified many with clusters of high-affinity DnaA boxes located near the inferred origins of replication

(Mackiewicz, Zakrzewska-Czerwinska et al. 2004). In Streptomyces coelicolor clusters of DnaA binding sites have been identified in the vicinity of the origin of chromosome replication.

Deletion of one of these clusters resulted in an increased rate of initiation of chromosome replication, while extra copies resulted in a slower colony growth, presumably due to a decrease in initiation of chromosome replication (Smulczyk-Krawczyszyn, Jakimowicz et al.

2006). Taken together, this suggests that titration of DnaA away from the origin of replication is a widespread mode of regulating DnaA activity.

Regulatory inactivation of DnaA (RIDA)

The dominant mechanism used by E. coli to limit chromosome replication is called the regulatory inactivation of DnaA (RIDA). Summarized in Figure 5C, RIDA involves the inactivation of DnaA by the stimulation of the ATPase activity of DnaA, converting the replication competent

DnaA-ATP form to a DnaA-ADP form which is unable to stimulate chromosome replication.

RIDA was originally identified biochemically to require the β-subunit of DNA polymerase III

(DnaN) and a partially purified protein IdaB (Katayama, Kubota et al. 1998). The IdaB protein was identified as Hda by combining this biochemical analysis with a genetic screen for a high copy suppressors of a DnaN mutant (Kato and Katayama 2001). The Hda protein displays homology to the ATPase AAA+ domain III of DnaA. Hda was later independently identified to be involved in the replication of the broad host range RK2 plasmid, which requires both DnaA and a plasmid replication factor TfrA for replication. Hda was found to repress the toxicity of a peptide derived from TfrA (Kim, Banack et al. 2003). The Hda mediated inactivation of DnaA is an important process preventing over initiation; Hda deficient E. coli over initiate replication from oriC (Camara, Breier et al. 2005).

The ability to bind and hydrolyze ATP to ADP is a characteristic of all the DnaA proteins identified so far. In addition to E. coli DnaA, ATPase activity has been demonstrated for DnaA

38 proteins of several other bacteria including Bacillus subtilis (Fukuoka, Moriya et al. 1990),

Streptomyces lividans (Majka, Messer et al. 1997), Thermus thermophilus (Schaper, Nardmann et al. 2000), Thermotoga maritima (Ozaki, Fujimitsu et al. 2006), and, Mycobacterium tuberculosis (Yamamoto, Muniruzzaman et al. 2002) and Caulobacter crescentus (this thesis chapter #4).

Unlike SeqA, RIDA is not limited to acting at oriC and RIDA acts to inactivate DnaA at the replication forks as DNA synthesis progresses. For example, temperature sensitive mutants with blocked DNA synthesis at non-permissive temperature accumulate DnaA-ATP. In contrast, temperature sensitive mutants of proteins involved in cell division have DnaA-ATP levels similar to wild type (Kurokawa, Nishida et al. 1999). RIDA mediated hydrolysis of the nucleotide bound to DnaA does not require sequence-specific elements at oriC, since rnhA deletion mutants of E. coli which initiate chromosome replication from an alternate origin also show RIDA activity

(Kurokawa, Nishida et al. 1999). A study done in vitro shows that DnaN loaded onto DNA is sufficient for Hda to stimulate the hydrolysis of ATP associated with DnaA in the absence of

DNA synthesis and this does not require sequence-specific elements. Hda makes contact with

DnaN with a specific peptide motif QL[SD]LF found near the N terminus of Hda (Dalrymple,

Kongsuwan et al. 2001, Kurz, Dalrymple et al. 2004), and this motif is found in other DnaN interaction proteins (Dalrymple, Kongsuwan et al. 2001).

The crystal structure of Hda of Shewanella amazonensis SB2B has been solved with a resolution of 1.75 Å (Xu, McMullan et al. 2009). The Hda monomer displays features typical of an AAA+ protein. The structure of Hda is divided into two subdomains (residues 1–174 and

175–241). The N-terminal domain I (residues 1–174) contains a RecA-like fold containing the

Walker A (the P-loop), Walker B, and sensor I motifs. An extended N-terminal loop contains a short strand (β1) required for binding the sliding clamp (DnaN) (Su'etsugu, Shimuta et al. 2005).

The C-terminal domain IV (residues 175–241) contains the sensor II motif and is comprised of a short four-helix bundle (α5–α8).

HdaA is believed to form a homo-dimer with a novel mode of AAA+ oligomerization as

AAA+ proteins often form hexameric rings. The dissociation of the Hda homo-dimer is required for activation of the Hda protein. Despite being an AAA+ protein, experiments show that Hda specifically binds to ADP but not ATP in vitro and in vivo (Su'etsugu, Nakamura et al. 2008). The binding of ADP by Hda promotes dissociation of the Hda homo-dimers to monomers and such activated Hda monomers function in the RIDA system in vitro (Su'etsugu, Nakamura et al.

2008). However, the biological significance of this reaction is not known.

Figure 5: Mechanisms used to limit chromosome replication in E. coli

E. coli possesses at least three mechanisms which prevent the over initiation from one origin of replication.

A) Newly replicated origins are sequestered from the replication machinery by the SeqA protein. The origin of replication contains multiple recognition sites (GATC) for the Dam methyltransferase (indicated by red bars). The chromosome begins the cell cycle fully methylated, i.e. on both strands. Replication of fully methylated DNA yields DNA which is hemi- methylated. The SeqA protein binds preferentially to hemi-methylated DNA blocking access to

DnaA to oriC.

B) DnaA is also shown regulated by titration. The datA locus has a very high affinity for DnaA, binding eight times more DnaA than the oriC. The datA locus is located near the origin of replication, thus its early duplication serves to reduce the amount of free DnaA available for initiation at oriC.

C) The activity of DnaA is controlled by the nucleotide ATP or ADP bound to it. The inactivation of DnaA-ATP by its conversion to DnaA-ADP is mediated by a process termed the regulatory inactivation of DnaA (RIDA). Once the replication machinery has been assembled at the origin, the intrinsic ATPase activity of DnaA is stimulated by the β-subunit of DNA polymerase III,

DnaN, together with a protein with homology to DnaA, Hda. DnaN binds Hda and brings it into contact with DnaA-ATP to stimulate hydrolysis and DnaA-ADP +Pi formation. Like the two preceding examples, this is also a negative feedback mechanism because replication creates the conditions, i.e. the assembly of DnaN around the DNA, which inhibit DnaA activity.

Figure 5 Mechanisms used to limit chromosome replication in E. coli (A) Sequestration of newly replicated origins

Figure 5 Mechanisms used to limit chromosome replication in E. coli (B) Titration of DnaA away from the origin

oriC datA

Chromosome Replication

Figure 5 Mechanisms used to limit chromosome replication in E. coli (C) Regulatory Inactivation of DnaA (RIDA)

Reactivation of DnaA The excess of ATP over ADP in the cell means that newly synthesized DnaA is formed in an active DnaA-ATP configuration. Chromosome replication and RIDA result in the hydrolysis of the bound ATP forming an inactive DnaA-ADP. Two mechanisms have been identified which can “rejuvenate” the inactive DnaA-ADP to an active DnaA-ATP form. Both mechanisms described below rely on releasing the tightly bound ADP so that the empty “apo-DnaA” protein becomes free to bind a new ATP molecule.

DARS (DnaA Reactivation Sequences) In E. coli, the amount of DnaA-ATP in the cell varies in a cell cycle dependent manner, peaking at the time of chromosome initiation (Kurokawa, Nishida et al. 1999). Newly synthesized DnaA is most likely to bind ATP because it is present at much higher concentrations in the cell than

ADP. DnaA-ATP can also be generated from DnaA-ADP by DNA sequence induced nucleotide exchange. These DNA sites are termed DARS (DnaA Reactivation Sequences). The DARS site consists of three DnaA boxes whose configuration favors the removal of the nucleotide from

DnaA. DARS promote a binding and release cycle that favors the release of the apo-DnaA form and the apo-DnaA is then free to associate with ATP (Fujimitsu and Katayama 2004). Two DARS sites have been identified on the E. coli chromosome (Fujimitsu, Senriuchi et al. 2009).

Additional copies of the DARS resulted in an increase in the DnaA-ATP level and increased initiation of chromosome replication. Deletion of the DARS sites resulted in a decrease in the

DnaA-ATP level in the cell and so inhibited the initiation of replication. This deletion of the

DARS sites also suppressed the over-initiation caused by the lack of seqA or datA (Fujimitsu,

Senriuchi et al. 2009).

DnaA membrane interactions Interactions with the cell membrane can also convert inactive DnaA-ADP to the replication competent DnaA-ATP form. Treatment of E. coli DnaA with phospholipids in a fluid phase caused the dissociation of tightly bound ADP or ATP from DnaA protein (Sekimizu and Kornberg

1988). Acidic phospholipids (cardiolipin, phosphatidylglycerol, and phosphatidic acid) together with DNA from oriC readily stimulate the exchange of ADP for ATP (Sekimizu and Kornberg

1988, Yung and Kornberg 1988, Castuma, Crooke et al. 1993).

In E. coli, the disruption of the pgsA gene renders the organism incapable of synthesizing phosphatidylglycerol or cardiolipin, resulting in cells that are depleted of acidic phospholipids for growth (Heacock and Dowhan 1987). A mutation in the rnhA gene, which encodes the

RNase H, allows for chromosome replication in the absence of DnaA by induction of constitutive stable DNA replication (described above) (Kogoma and von Meyenburg 1983). This rnhA mutation also relieves the growth arrest phenotype of the pgsA mutant (Xia and Dowhan

1995). As noted above, rnhA bypasses the need for DnaA and replication from oriC, arguing that DnaA is the direct target and cause of the pgsA growth arrest (Xia and Dowhan 1995). In summary, these results suggest a direct connection between the production of phospholipids,

DnaA-ATP formation and the DnaA-dependent initiation of chromosome replication.

DNA replication in eukaryotic cells Comparison of prokaryotic and eukaryotic chromosome replication In both prokaryotic and eukaryotic cells, the faithful duplication and transmission of genetic information requires the assembly of highly organized nucleoprotein complexes on the chromosome. There exist both functional and structural similarities in the pre-replicative (pre-

RC) complex of prokaryotic and eukaryotic cells. Unlike prokaryotic chromosomes which initiate chromosome replication from a single origin, eukaryotic chromosomes contain multiple origins of replication. The use of a single origin of replication effectively limits the amount of

DNA which can be replicated during a cell cycle. As described above, E. coli initiates replication from a single origin of replication (oriC) and replication proceeds bidirectionally at a rate of approximately 60 kb per minute. However in the much larger human genome (about 700 times the size of the E. coli genome), replication forks travel at the much slower rate of 2 to 3 kb per minute. Eukaryotic chromosome replication would be a prohibitively long process if only one origin was used per chromosome. Eukaryotic chromosomes initiate replication from multiple origins. Budding and fission yeast contain approximately 500 origins distributed over a genome of about 1.4 X107 base pairs (Wyrick, Aparicio et al. 2001, Heichinger, Penkett et al. 2006). Work with Chinese hamster cells has shown that mammalian cells initiate replication from between

30,000 and 50,000 origins per cell cycle (Huberman and Riggs 1966).

Eukaryotic origins of replication Yeast origins of replication The best studied eukaryotic origins of replication are from the budding yeast Saccharomyces cerevisiae. These origins of replication were originally identified as segments of the chromosome which conferred high frequency transformation due to the autonomous replication of the recombinant plasmids (Stinchcomb, Struhl et al. 1979). The sizes of these

“autonomously replicating sequences” termed ARS in S. cerevisiae, are short and often range from 100 to 150 base pairs. Further analysis of these ARSs showed that they contain a highly conserved 11 base pair sequence (Broach, Li et al. 1983) called the ‘A’ element as well as a less conserved ‘B’ element (Marahrens and Stillman 1992). The A element is recognized by the ORC proteins (Bell and Stillman 1992). The origins observed in S. cerevisiae follow the model of chromosome replication proposed for bacteria (Jacob and Brenner 1963), with a cis-acting replicator (the ARS) and a trans-acting initiator (the ORC).

The presence of well-defined sequence specific origins of replication has not been observed outside of S. cerevisiae. For example, the selection of origins of replication in South

African clawed toad Xenopus laevis does not require any sequence specific information, as practically any DNA injected into X. laevis egg will be replicated (Harland and Laskey 1980,

Mechali and Kearsey 1984). Most significantly, the replication of the injected DNA occurs in a controlled cell cycle manner, being replicated no more than once per cell cycle (Harland and

Laskey 1980, Mechali and Kearsey 1984). Therefore, although eukaryotes replicate their chromosomes precisely during the cell cycle, the choice of the replication origins need not be precisely made.

Steps involved in eukaryotic chromosome replication Initiation of chromosome replication in eukaryotes can be divided into three stages: the recognition of the origin sequences on the chromosome, the assembly of the pre-replicative complex (pre-RC) on the origin, and the activation of the pre-RC. The proteins which make up the pre-RC are the origin recognition complexes (ORC) and Cdc6. The first ORC was purified from the budding yeast S. cerevisiae, as a heterohexameric complex which binds to origins of

DNA replication in a sequence specific manner (Bell and Stillman 1992). The subunits were

48 named ORC1 through 6. Orc1, Orc4, and Orc5 are AAA+ proteins (Iyer, Leipe et al. 2004). The proteins Orc1 to Orc5 are highly conserved. Orthologs have been identified in a diverse range of eukaryotic organisms including plants (Arabidopsis thaliana) (Masuda, Ramos et al. 2004) , insects (Drosophila melanogaster) (Remus, Beall et al. 2004), and mammals (Homo sapiens)(Vashee, Cvetic et al. 2003).

The regulation of ORC by ATP The activity of the eukaryotic ORC is regulated by binding ATP. In a manner reminiscent of prokaryotic DnaA, ORC is able to bind and hydrolyse ATP (Bell and Stillman 1992, Austin, Orr-

Weaver et al. 1999). Research performed using S. cerevisiae and D. melanogaster, has revealed that the Orc1 subunit of ORC is responsible for binding and hydrolysing ATP (Klemm, Austin et al. 1997, Chesnokov, Remus et al. 2001). The binding of ATP by ORC is required for DNA binding; however, ATP hydrolysis is not required for DNA binding (Klemm, Austin et al. 1997).

The ATPase activity of S. cerevisiae ORC (ScORC) is inhibited by DNA in a sequence-specific manner. For example, ScORC mixed and bound with wild type origin DNA has a tenfold decrease in ATPase activity compared with ScORC mixed with origin DNA which contains mutations that inhibit ORC binding (Klemm, Austin et al. 1997). A less pronounced decrease was observed in the ATPase activity of DmORC (D. melanogaster ORC) when bound to DNA. The binding of ORC to the double-stranded DNA of the origin of replication stabilizes ORC in an ATP bound state. However, single-stranded DNA stimulates the ATPase activity of ORC (Lee,

Makhov et al. 2000). Since unwinding of double-stranded DNA creating single-stranded DNA is one of the early steps in chromosome replication, the binding and hydrolysis of ATP suggests a negative feedback that inactivates ORC in S-phase.

In a process termed “origin licencing” the ORC recruits other proteins to form a larger complex called the pre-replicative complex (pre-RC) (Diffley, Cocker et al. 1994)(Figure 6). The proteins which compose the pre-RC complex include Cdc6 (Liang, Weinreich et al. 1995), cdt1

(Maiorano, Moreau et al. 2000, Nishitani, Lygerou et al. 2000) and the mini chromosome maintenance proteins (Mcm2, Mcm3, Mcm4, Mcm5, Mcm6, and Mcm7) (Chong, Mahbubani et al. 1995, Kubota, Mimura et al. 1995, Madine, Khoo et al. 1995). Two replication factors have been shown to be important for loading the MCM complex onto the ORC: Cdc6 and Cdt1. The replication factor Cdc6 is a highly conserved protein with homology to the Orc1 protein (Bell,

Mitchell et al. 1995). Cdc6 was originally identified in S. cerevisiae (Hartwell 1976) and has since been identified in all eukaryotic cells. Orthologs of Cdc6 have also been identified in the

Archaea (Barry and Bell 2006). Cdc6 plays an essential role in the formation of the pre-RC, as a conditional knock out of cdc6 fails to produce the pre-RC (Cocker, Piatti et al. 1996). Cdc6 is responsible for loading the DNA helicase MCM protein complex onto the origin of replication

(Donovan, Harwood et al. 1997, Liang and Stillman 1997, Tanaka, Knapp et al. 1997). Cdt1 was originally found in the fission yeast Schizosaccharomyces pombe as an essential target of the

Cdc10 transcription factor (Hofmann and Beach 1994). In S. pombe, both Cdc18 (equivalent of

Cdc6 in S. cerevisiae) and Cdt1 are required to load the MCM complex onto the chromosome

(Nishitani, Lygerou et al. 2000).

It is interesting to note that Eukaryotes (and presumably the Archaea) recruit their MCM replicative helicases to the origins long before the initiation of chromosome replication. In contrast, E. coli (and presumably most bacteria) recruits the DnaB replicative helicase only during the initiation process (Figure 7).

Figure 6: Licensing of eukaryotic origins of replication

The origin recognition complex is made up of six subunits (Orc 1 to 6) and is found associated with the origin throughout the cell cycle. During G1 the origin becomes licensed for replication by the loading of Cdc6, Cdt1 and the MCM helicase complex. Once replication has been initiated, the origin is returned to an un-licensed state by a combination of ATP hydrolysis and protein phosphorylation that results in the dissociation of Cdc6, Cdt1 and the MCM complex.

Figure 6: Licensing of eukaryotic origins of replication

Figure 7: Comparison of the initiation of chromosome replication in bacteria and eukaryotic cells

In bacteria the protein used to recognise the origin of replication is DnaA, in eukaryotic cells the origin of replication is recognised by a complex of six proteins called the origin recognition complex (ORC). While both DnaA and ORC bind to DNA, the binding of DnaA to the origin is more dependent upon sequence specificity than the ORC of eukaryotic cells. DnaA is found associated with the high affinity binding sites at the origin throughout the cell cycle. Both DnaA and ORC are AAA+ proteins capable of binding ATP, which results in conformational changes which stimulate unwinding of the origin of replication and recruitment of the replication machinery. DnaA interacts directly with the DnaB helicase while ORC binds Cdc6 and Cdt1 which are required for helicase loading. Once chromosome has been initiated, DnaA is inactivated by hydrolysis of the bound ATP, while ORC is inactivated via a combination of hydrolysis of ATP, the disassociation of Cdc6 and Cdt1 and the phosphorylation of the ORC subunits. This figure is based upon the model presented in Scholefield et al. (2011).

Figure 7: Comparison of the initiation of chromosome replication in bacteria and eukaryotic cells

Bacterial Eukaryoti c

6 2 Initiator 5 3 4 DnaA 1 ORC proteins

Origin Recognition

Activation of Replication

Complex

Initiation of Replication

-P Inactivation of -P Chromosome

Replication -P Complex

Chromosome replication in the Archaea Chromosome replication in the Archaea more closely resembles that of the eukaryotes than that of bacteria (Edgell and Doolittle 1997). The replication machinery resembles that found in eukaryotic cells, as archaeal homologs of many eukaryotic replication proteins have been identified, including the origin recognition complex (ORC), as well as the MCM helicase. The number of origins of replication is variable among archaeal species, some containing a singular origin of replication resembling bacteria, such as Pyrococcus abyssi (Myllykallio, Lopez et al.

2000), while others resemble eukaryotic cells containing multiple origins of replication such as

Sulfolobus acidocaldarius and Sulfolobus solfataricus (Lundgren, Andersson et al. 2004).

The origin binding proteins of Archaeal cells are homologues of the Orc1 and Cdc6 of eukaryotic cells. Like their eukaryotic counterparts the Archaeal OrC1/Cdc6 proteins are members of the AAA+ family of proteins. With only a few exceptions, all the archaeal genomes sequenced contain at least one gene with homology to Orc1 and Cdc6(Barry and Bell 2006).

The replication origins of Archaeal species are defined by sequence specific elements similar to origins found in bacteria and S. cerevisiae. Analysis of the S. solfataricus origins of replication (oriC1, oriC2) has led to the identification of inverted repeats sequence elements which are recognized by Cdc6 at oriC1 and these elements are called origin recognition boxes

(ORB). The ORBs are conserved across many archaeal species. The oriC2 contains sequences which are homologous to the central elements of the ORBs and are called mini-ORBs (Robinson,

Dionne et al. 2004).

Following this background material on chromosome replication, I will next present material that frames my research goals. I will start by describing the alphaproteobacteria,

55 because this background will give a better appreciation for my choice of C. crescentus as a model organism.

Caulobacter crescentus as a model for prokaryotic cell development

Alphaproteobacteria Caulobacter crescentus is a model organism for the alphaproteobacteria, which comprise a diverse group of bacteria. Alphaproteobacteria frequently adopt either a parasitic or symbiotic intracellular lifestyle with eukaryotic host cells (Batut, Andersson et al. 2004). For example, the

Rickettsia genus is composed of obligate intracellular parasites, which include the human pathogens Rickettsia typhi and Rickettsia prowazekii. R. typhi is the causative agent of Murine typhus and is transmitted by fleas, while R. prowazekii is the causative agent of epidemic typhus and is transmitted by lice associated with poor sanitary conditions. The main vector of

R. prowazekii in North America is the flying squirrel (Glaucomys volans). R. prowazekii may develop into a latent infection but stress or a failing immune system may trigger reactivation of the latent infection referred to as Brill-Zinsser disease. The fact that R. prowazekii is stable in louse faeces and could be easily dispersed in aerosol has promoted its classification as a category B bioterrorism agent. Rickettsia species contain a highly streamlined genome which resembles that of the eukaryotic mitochondria.

Brucella is the causative agent of the zoonotic disease brucellosis. Several Brucella species have been identified which cause Brucellosis in cattle (B. abortus), sheep (B. melitensis), pigs (B. suis), and dogs (B. canis). Transmission to humans can occur via direct contact with an infected animal or through the consumption of unpasteurized milk. Brucella species which infect marine mammals, B. ceti (porpoises and dolphins) and B. pinnipedialis (seals) (Foster,

Osterman et al. 2007), have been shown to also infect humans (Whatmore, Dawson et al.

2008). Recently two novel species of Brucella have been identified in human infections. B. inopinata was isolated from an infected breast implant (Scholz, Nockler et al. 2010) and a related strain (BO2) was found in a person with chronic lung disease (Tiller, Gee et al. 2010).

The natural reservoirs of these bacteria have yet to be determined. Brucella species have also been categorized as a category B bioterrorism agent.

In contrast to these pathogens, the bacterium Sinorhizobium meliloti is a member of the alphaproteobacteria with significant environmental and agricultural importance. S. meliloti is a soil bacterium which is able to induce the formation of nodules in specific legumes including the forage crop alfalfa (Medicago sativa). These bacteria colonize the roots of the plant through infection threads and ultimately enter the cytoplasm of the plant cells where they differentiate into bacterioids. The S. meliloti bacteroids become specialized organelles that reduce inorganic atmospheric nitrogen to ammonia, a bioavailable form of nitrogen. The ammonia is directly assimilated by the plant and when the plant dies, the nitrogen is released to the soil and is thus available for other plants.

The plant pathogen Agrobacterium tumefaciens is the causative agent of crown gall disease. A. tumefaciens is a broad host range pathogen which is capable of inducing tumorous growths on most species of dicot as well as some monocots (De Cleene and De Ley 1976). The pathogenicity of A. tumefaciens is dependent upon the presence of a large (approximately 200 kb) tumor inducing (Ti) plasmid. A portion of the plasmid flanked by 23 base pair repeats is transferred into the plant cell where it is integrated into the nuclear DNA. The expression of this transferred DNA (T-DNA) causes changes in the plant hormones resulting in tumour formation, as well as the expression of genes for the synthesis of novel amino acid-sugar

57 conjugates, called opines. The formation of opines is advantageous for the colonizing bacteria since only they can utilize this uncommon carbon and nitrogen source. Unlike the plant symbiont Sinorhizobium meliloti, A. tumefaciens does not benefit the plant and it is detrimental to many commercial crops. (Review in (McCullen and Binns 2006). A. tumefaciens has become a valuable tool for creating genetically modified plants, since the transfer of DNA to the plant does not require the tumour inducing genes and any DNA can be integrated into the plant chromosome (Hooykaas and Schilperoort 1992).

The alphaproteobacteria also includes some of the most successful bacterial species on earth. The SAR11 clade of alphaproteobacteria is believed to be the most abundant cell in the ocean. On average one in three cells present in surface waters and one in five of the cells present in the mesopelagic zone belong to the SAR11 clade (Morris, Rappe et al. 2002).

C. crescentus is grouped among the alphaproteobacteria and it can therefore serve as a model for these and many other interesting bacteria that are often difficult to grow and to study. C. crescentus is a free living Gram-negative non-pathogenic bacterium which is found in fresh water lakes and streams. Caulobacter species have been isolated from both fresh and saline water environments (Anast and Smit 1988). These include waste water treatment plants which contain relatively high concentrations of nutrients (MacRae and Smit 1991) as well as ground water from highly polluted environments, e.g. polluted by organic chemicals (Mannisto,

Tiirola et al. 1999) or by heavy metals (North, Dollhopf et al. 2004). Caulobacter species have also been detected in extreme environments found in gold mines (Inagaki, Takai et al. 2003) and in deep-sea sediments (Marchesi, Weightman et al. 2001).

The original CB15 strain of Caulobacter crescentus was isolated in 1960. Since then, C. crescentus has become a model bacterium for studying many processes including the bacterial cell cycle, asymmetric cell division and bacterial aging. Like all Caulobacter species, C. crescentus produces two cell types, a motile “swarmer” cell, and an adherent non-motile

“stalked” cell (Figure 8). The swarmer cells must differentiate into stalked cells in order to grow and divide. Simple density-gradient centrifugation is sufficient to separate these two cell types, effectively creating a culture with synchronized swarmer cells that can be studied as they differentiate into growing and dividing stalked cells. This growth process has been termed the

“dimorphic” cell cycle (Poindexter 1964). The NA1000 strain of C. crescentus is a so called

“synchronizable” derivative strain of CB15 because its swarmer cells can be more efficiently isolated by density gradient centrifugation (Evinger and Agabian 1977). NA1000 has since become the main laboratory strain of C. crescentus. CB15 and NA1000 have several additional differences, including the loss of surface adhesive in the NA1000 strain. The exact genetic differences between these two strains have recently been identified (Marks, Castro-Rojas et al.

2010).

Figure 8: The C. crescentus cell cycle

C. crescentus is a free living fresh water Gram-negative bacterium, with a dimorphic life cycle.

The stalked cell is non-motile; in nature the stalked cell would be attached to a surface by the holdfast located at the end of the stalk. Chromosome replication (red oval) is restricted to the stalked cell type; the stalked cell elongates dividing asymmetrically to yield one stalked cell and one swarmer cell. The swarmer cell contains a single flagellum positioned on the pole opposite to the stalked cell. The swarmer cell is motile but not competent for chromosome replication

(blue oval) and must differentiate into a stalked cell by ejecting the flagellum and producing a stalk prior to chromosome replication

Figure 8: The Caulobacter crescentus cell cycle

Regulatory mechanisms first studied in C. crescentus have been found to be conserved among other members of the alphaproteobacteria. The four master cell cycle regulators of C. crescentus, CtrA, GcrA, CcrM, and, DnaA are widely conserved among alphaproteobacteria

(Brilli, Fondi et al. 2010). The master transcriptional regulator CtrA is essential for cell viability in C. crescentus and orthologs of CtrA have been identified in Agrobacterium tumefaciens

(Kahng and Shapiro 2001), Brucella abortus (Bellefontaine, Pierreux et al. 2002), Sinorhizobium meliloti (Barnett, Hung et al. 2001), and Rickettsia prowazekii (Brassinga, Siam et al. 2002).

However, CtrA is not essential in all alphaproteobacteria, as for example ctrA may be deleted from Rhodospirillum centenum (Bird and MacKrell 2011), Silicibacter sp. TM1040 (Miller and

Belas 2006), Rhodobacter capsulatus (Lang and Beatty 2002) and Magnetospirillum magneticum (Greene, Brilli et al. 2012) resulting only in a loss of cell motility but not in cell viability. This suggests that while the CtrA protein is highly conserved it may be adapted to severe different purposes in different species of bacteria.

The CcrM DNA methyltransferase is another highly conserved cell cycle regulator first discovered in C. crescentus. CcrM homologs have been identified in at least 20 members of the alphaproteobacteria (Stephens, Reisenauer et al. 1996). CcrM was shown to be essential for viability in bacteria with agricultural and economic importance including, Agrobacterium tumefaciens (Kahng and Shapiro 2001), Brucella abortus (Robertson, Reisenauer et al. 2000), and Sinorhizobium meliloti (Wright, Stephens et al. 1997).

Such examples argue that information gleaned from the study of C. crescentus could have widespread medical and economic implications, including wide ranging fields such as antibiotics, agriculture, and counter bioterrorism.

Caulobacter crescentus cell cycle

C. crescentus offers an opportunity to study the spatial and temporal mechanisms of bacterial cell development. Progression through the bacterial cell cycle requires a sequence of events including changes in cellular morphology, chromosome replication, chromosome segregation, and cell division. Unlike many prokaryotes which may have overlapping rounds of chromosome replication, C. crescentus initiates chromosome replication exactly once during the cell division cycle (Marczynski 1999). Furthermore the G1, S, and G2 phases of the cell cycle are tied to cell morphology making it possible to monitor the cell cycle using simple light microscopy.

C. crescentus has an obligate dimorphic life cycle (Figure 8), cell division results in two cell types: one sessile stalked cell and one motile swarmer cell. The swarmer cell possesses multiple pili and a single flagellum at one end as well as an intricate chemosensory network, enabling its response to chemo-attractants and movement towards potential food sources. The swarmer cell is not able to undergo chromosome replication and it must first differentiate into a stalked cell, by ejecting the flagellum, disassembling the pili and extending a stalk in the place of the flagellum. The stalk is a thin extension of the cell envelope which is filled with cytoplasm.

An adhesive organelle termed the holdfast is located at the end of the stalk. The holdfast is made up of a polysaccharide adhesin which is responsible for the strong attachment of C. crescentus to surfaces (Smith, Hinz et al. 2003). This adhesive is believed to be the strongest glue found in nature (Tsang, Li et al. 2006). The stalked cell begins chromosome replication almost immediately. The newly replicated parts of the chromosome are rapidly transported to the cell-pole opposite from the stalk (Viollier, Thanbichler et al. 2004). Also during chromosome replication, a new flagellum is synthesized and assembled at the pole opposite

63 from the stalk. The cell continues chromosome replication while elongating and assembling the flagellum. As replication is complete, the cell begins to constrict at the future site of cell division to form a pre-divisional cell (Jensen 2006). The pre-divisional cell takes on a characteristic crescent shape from which its name is derived. The cell will then divide asymmetrically yielding one swarmer cell and a stalked cell. The stalked cell may begin a new round of replication immediately, while the swarmer cell must first differentiate into a stalked cell to start chromosome replication and to complete the cell cycle. Interestingly, the stalked cell does not undergo an unlimited number of cell divisions, making C. crescentus also a model for bacterial aging (Ackermann, Stearns et al. 2003).

In summary, by analogy to the eukaryotic cell cycle, the swarmer cell is in the pre- synthetic G1 phase, the cell enters S phase shortly after differentiating to become a stalked cell, while the pre-divisional stalked cell is in the post-synthetic G2 phase.

Master cell cycle regulators, CtrA, GcrA, DnaA, CcrM

Temporal control of the cell cycle is maintained by synthesis and proteolysis of the components needed at each of the stages. The transcripts of 553 genes, approximately 20% of the predicted genes in the C. crescentus genome, have been shown to fluctuate in a cell cycle dependent manner (Laub, McAdams et al. 2000). The transcription of many of these genes including genes for DNA replication, chromosome segregation, cytokinesis and for biogenesis of the stalk, flagellum and pili coincide with the times the gene products are needed in the cell (Laub,

McAdams et al. 2000).

CtrA The transcriptional regulator CtrA has been show to regulate several cell cycle events including flagella and pilus biogenesis, cell division, DNA methylation, and DNA replication (Quon,

Marczynski et al. 1996, Kelly, Sackett et al. 1998, Quon, Yang et al. 1998, Skerker and Shapiro

2000, Laub, Chen et al. 2002). CtrA binds to a specific motif (consensus TTAA-N7-TTAAC)

(Quon, Marczynski et al. 1996) in the promoter regions of 55 operons, allowing it to affect transcription of 95 cell cycle regulated genes (Laub, Chen et al. 2002). CtrA auto-regulates its own transcription from two promoters. The P1 is a weak promoter which is repressed by phosphorylated CtrA. The P1 promoter is active in stalked cells and CtrA accumulates in early pre-divisional cells. The accumulation of CtrA in the cell then represses transcription from the

P1 promoter while it activates the stronger P2 promoter (Domian, Quon et al. 1997).

The activity of CtrA is controlled by its phosphorylation state. CtrA is phosphorylated by the CckA histidine kinase through the ChpT phosphotransferase (Jacobs, Domian et al. 1999,

Biondi, Reisinger et al. 2006). Phosphorylated CtrA (CtrA-P) is found in swarmer cells and in pre-divisional cells. CtrA is inactivated by dephosphorylation and is rapidly degraded by the

ClpXP protease at the swarmer to stalked cell transition and in the stalked compartment of late pre-divisional cells (Domian, Quon et al. 1997, Jenal and Fuchs 1998);(Spencer, Siam et al.

2009).

The C. crescentus origin of replication contains five binding sites for CtrA. It was previously believed that the role of CtrA binding to the origin was to repress chromosome replication in swarmer cells. However, under laboratory conditions it is possible to disrupt all five of these binding sites, without hindering the viability of the cell (Bastedo and Marczynski

2009). It is only when the cell is challenged by antibiotics, or by a dramatic change in nutrient levels that loss of CtrA binding become deleterious to the cell (Bastedo and Marczynski 2009).

These results indicate that CtrA processes environmental stress signals and that it integrates this information with control of chromosome replication.

GcrA GcrA has been shown to interact with promoter sequences in vivo (Holtzendorff, Hung et al.

2004), although this interaction may be indirect as GcrA does not have an obvious DNA binding domain. Like CtrA, GcrA is an essential cell cycle regulated protein, depletion of GcrA alters directly or indirectly the expression of 125 genes, 49 of which are cell cycle regulated genes.

GcrA protein abundance oscillates out of phase with the transcriptional regulator CtrA during the cell cycle. The 125 GcrA regulated genes are involved in multiple aspects of the cell cycle, including DNA replication and repair, cell motility, polar development and cell wall synthesis as well as other regulatory genes. These regulatory genes include the response regulators CtrA and DivK, the histidine kinase PleC as well as the sigma-54 factor activator TacA (Holtzendorff,

Hung et al. 2004).

Tightly regulated transcription and proteolysis work together and thereby segregate

CtrA and GcrA to the different cell types during the cell cycle. CtrA, but not GcrA, is present in swarmer cells, while GcrA but not CtrA is present in stalked cells. The promoter of gcrA is inhibited by CtrA, such that gcrA is only expressed once CtrA has been degraded at the swarmer to stalked cell transition. The accumulation of GcrA then triggers the expression of ctrA: GcrA stimulates transcription of ctrA from the weak P1 promoter; once CtrA-P has accumulated in the cell it then represses transcription of gcrA. GcrA accumulates in the stalked cell and the stalked compartment of the late pre-divisional cell (Holtzendorff, Hung et al. 2004).

CcrM CcrM (cell-cycle-regulated methyltransferase ) is an essential DNA methyltransferase which was originally identified in C. crescentus, and then it was found to be widespread among the alpha subdivision of proteobacteria (Zweiger, Marczynski et al. 1994, Stephens, Reisenauer et al.

1996), including Rhizobium meliloti, Brucella abortus, Agrobacterium tumefaciens, and

Rhodobacter capsulatus (Wright, Stephens et al. 1997). CcrM specifically methylates the adenine base of the sequence GANTC. CcrM is synthesized late in the cell cycle when it completely methylates the chromosomes. Then CcrM protein is completely degraded just prior to cell division (Stephens, Reisenauer et al. 1996). Expression of the ccrM gene is controlled in a strict cell cycle dependent manner, with maximal expression late in S phase. Expression of ccrM is controlled by CtrA (Reisenauer, Quon et al. 1999). Chromosome replication during S phase results in the creation of hemi-methylated DNA, and it is late in S phase when the approximately 30,000 GANTC CcrM sites of the newly replicated chromosomes are remethylated (Reisenauer, Kahng et al. 1999). CcrM is then removed from the cell by the Lon protease, limiting the activity of CcrM to late S phase (Wright, Stephens et al. 1996).

CcrM does not directly alter gene transcription but acts indirectly by methylating sites in the promoter sequences of genes and thereby affecting their transcription. CcrM has also been proposed to play a role in the cell cycle transcription of dnaA. CcrM plays a role in regulating its own transcription (Stephens, Zweiger et al. 1995). CcrM also helps regulates the transcription of ctrA. For example, the weak P1 promoter of ctrA contains a CcrM GANTC methylation site located in proximity to the -35 site (Reisenauer and Shapiro 2002). Transcription from this promoter is repressed when the promoter is fully methylated. Passage of a replication fork

67 during S phase results in the creation of hemi-methylated DNA and activation of the P1 promoter.

DnaA In the C. crescentus genome, DnaA R boxes are present upstream of many genes whose expression is DnaA-dependent (Hottes, Shapiro et al. 2005). The DnaA regulon includes components of the replisome, the master regulator GcrA, the cell division protein FtsZ, and nucleotide biosynthesis enzymes (Hottes, Shapiro et al. 2005). Intriguingly, while the nrdA and nrdB genes are not located in the same operon, they are still independently regulated by DnaA

(Hottes, Shapiro et al. 2005).

In the Gram positive bacterium Bacillus subtilis, DnaA regulates the transcription of more than 50 genes expressed from approximately 20 operons (Goranov, Katz et al. 2005).

DNA-protein crosslinking followed by DnaA immunoprecipitation showed that DnaA binds to at least 17 loci, 15 of which respond transcriptionally to blocked chromosome replication. A significant increase in DnaA binding was seen at 6 loci when DNA replication was interrupted.

DnaA binding was rapidly reduced when DNA replication resumed (Breier and Grossman 2009).

Chromosome replication

C. crescentus origin of replication (Cori) The Caulobacter crescentus origin of replication (Cori) was cloned (Marczynski and Shapiro

1992). Cori was mapped to a 1.6 Kb fragment spanning the intergenic region between the genes hemE and duf299. This placement is conserved among bacteria in the alphaproteobacteria subdivision (Brassinga and Marczynski 2001). Deletion analysis indicates that the minimum essential sequence for autonomous replication is somewhere between 430

68 and 720 base pairs (Marczynski and Shapiro 1992). Plasmid replication from Cori was observed only in stalked cells mirroring the replication of the chromosome (Marczynski and Shapiro

1992). Broad host range plasmids are able to replicate in albeit at a much reduced frequency in swarmer cells (Marczynski, Dingwall et al. 1990). This argues that the DNA replication machinery is present in both cell types and that regulatory elements located in Cori are responsible for determining proper cell cycle timing of replication.

The Caulobacter crescentus origin Cori contains several motifs which bind multiple regulatory proteins. Cori contains two classes of DnaA binding sites, two high-affinity DnaA binding sites termed G boxes (Marczynski and Shapiro 1992), and five low affinity DnaA binding sites termed W boxes (Taylor, Ouimet et al. 2011). Interestingly, these G boxes do not have the maximum affinity of the standard R boxes and, contrary to for example E. coli oriC, Cori evolved a relatively low affinity for DnaA.

Cori also contains five binding sites for the response regulator CtrA (Quon, Yang et al.

1998) designated a, b, c, d and e. These are among the strongest affinity CtrA binding sites in the whole genome. The exact function of each site is not known but in general, strong CtrA binding blocks Cori replication in the swarmer cell types. Cori also contains one binding site for the integration host factor (IHF) which overlaps with a CtrA binding site (site c) (Brassinga, Siam et al. 2002). Binding of IHF interferes with CtrA binding (Siam, Brassinga et al. 2003). Cori also contains a region of approximately 40 bases with a high (85%) AT content (Marczynski, Lentine et al. 1995). It is not known if this is the DUE or DNA unwinding element. However, this DNA is overlapped by 2 CtrA binding sites (sites a and b) and it overlaps the strong transcription promoter that points into the hemE gene. Transcription of the hemE gene adjacent to the Cori

(Brassinga and Marczynski 2001, Brassinga, Siam et al. 2001) may help regulate chromosome replication since it is required for autonomous Cori-plasmid replication (Marczynski, Lentine et al. 1995).

Stationary Phase Bacteria which are studied in laboratory environments have for the most part been selected for their ease of growth and manipulation. E. coli growing under ideal laboratory growth conditions will divide every 30 minutes. However, exponential growth is only seen for brief periods in laboratory settings, while conditions which permit sustained exponential growth of bacteria are rarely seen in nature. Furthermore, bacteria must deal with fluctuations in the levels of water, light, nutrient availability, temperature, and competition with other organisms.

When nutrients are plentiful, bacteria can grow rapidly but these same bacteria must be able to survive prolonged periods of nutritional starvation. Some bacteria including Bacillus subtilis

(Errington 1996), Clostridium difficile (Underwood, Guan et al. 2009), Myxococcus xanthus

(Sudo and Dworkin 1969) enter an alternate developmental pathway which leads to the development of spores which are dormant. These spores are then resistant to starvation, as well as extremes in temperature, desiccation, and ultraviolet radiation. During periods that are not amenable to bacterial growth, non-sporulating bacteria such as E. coli enter stationary phase. This is a form of cellular differentiation that renders bacteria resistant to multiple environmental stresses, including oxidative stress, heat shock, and acidic conditions (Jenkins,

Schultz et al. 1988, Small, Blankenhorn et al. 1994).

Bacteria growing in a culture will grow exponentially until the available nutrients have been exhausted and the growth rate decreases, indicating the onset of stationary phase.

Stationary phase is defined qualitatively as the point where the number of cells in the culture

70 does not increase. The onset of stationary phase could come from any point in the cell cycle due to sudden depletion of a key nutrient. However a more ordered entry into stationary phase is preferred as a sudden interruption in a key event such as DNA replication could have fatal consequences for the cell.

Stationary phase was previously viewed as a period of metabolic inactivity, during which the cell simply waits for conditions favorable to cell growth. It is now recognized that entry into stationary phase is characterized by a precise series of metabolic changes. This programmed genetic response allows the cell to survive changing nutrient levels and other environmental stresses. In E. coli and many of the gammaproteobacteria, gene expression during stationary phase is regulated by the alternate sigma factor Sigma S, also known as RpoS (Lange and

Hengge-Aronis 1991). The main role of the Sigma factor in the cell is to bind the core RNA polymerase and confer promoter specificity. Upon entry into stationary phase and during periods of stress, the Sigma S may partially replace the housekeeping Sigma factor (Sigma 70 in

E. coli). Genome-wide expression profiling has indicated that up to 10 percent of the E. coli genome is either directly or indirectly under the control of RpoS (Weber, Polen et al. 2005). In healthy growing cells, RpoS is produced but the adaptor protein SprE immediately targets it for degradation by the ClpXP protease (Pratt and Silhavy 1996). During periods of carbon starvation, there is a decrease in the energy storing molecule ATP. The reduction in the cellular level of ATP then inhibits the proteolysis of RpoS, resulting in its rapid accumulation in the cell

(Peterson, Levchenko et al. 2012). RpoS then competes with other sigma factors for binding to the core RNA polymerase, which in turn alters gene expression, increasing expression of genes which protect the cell against stress (Maeda, Fujita et al. 2000). Upon the return of carbon to

71 the environment, the normal cellular levels of ATP are re-established and RpoS is rapidly degraded and the RpoS controlled genes are no longer expressed (Peterson, Levchenko et al.

2012).

Although no homologs of the RpoS (Sigma S) have been identified in the alphaproteobacteria, bacteria such as C. crescentus enter a stationary phase characterized by reduced growth and increased resistance to environmental stresses. The induction of antioxidant molecules such as catalases and catalase-peroxidases are required for the removal of toxic reactive oxygen species. The katG gene of C. crescentus has been shown to be dramatically up regulated in stationary phase (Schnell and Steinman 1995). KatG has been shown to be essential for survival of C. crescentus in stationary phase, as the viability of a katG null mutant was reduced by more than 3 orders of magnitude after 24 h in stationary phase and more than 6 orders of magnitude after 48 h (Steinman, Fareed et al. 1997).

In addition to an early stationary phase response, when grown to saturation for extended periods, C. crescentus displays dramatic changes in morphology. After a long period of cell death, the surviving cells lose their characteristic crescent shape taking on an elongated coiled shape with no constrictions and an average length of 20 μm, which is 20 times longer than exponentially growing cells. When returned to fresh media, these cells revive and return to the normal C. crescentus size and morphology (Wortinger, Quardokus et al. 1998).

Proteolysis In bacteria, proteolysis plays a critical role in maintaining the concentration of regulatory proteins, ensuring that proteins are only present in the appropriate compartment of the cell, and in recycling proteins whose function is no longer required. Four classes of energy dependent proteases have been identified in E. coli. All four proteases consist of an ATPase

72 domain and a proteolytic domain, these domains may be found in different subunits as is the case for the ClpAP/XP and ClpYQ (also called HslUV) proteases, or they may be different domains of the same protein as is the case for the Lon and FtsH proteases.

Due to evolutionary pressure, the majority of bacterial cellular proteins are stable.

Nonetheless, the regulated proteolysis of proteins is an important mechanism which ensures that they are removed once they are no longer useful to the cell. A global analysis of the proteins present in C. crescentus was able to identify approximately one quarter (979) of the predicted proteins by two-dimensional gel electrophoresis. Of these, a total of 48 proteins were rapidly degraded during the course of one cell cycle and half of these proteins were expressed in a cell cycle-dependent manner (Grunenfelder, Rummel et al. 2001). There are several well characterized examples of proteolysis used to ensure that key cell cycle regulators are present at the correct time and place in the cell.

The master cell cycle regulators CtrA, GcrA, DnaA, and CcrM are all themselves regulated by proteolysis. Proteolysis limits the presence of the response regulator CtrA to the swarmer cell period of the cell cycle and it directs the sub-cellular CtrA distribution in the pre- divisional cells (Domian, Quon et al. 1997). The CcrM DNA methyltransferase is degraded in a cell cycle dependent manner (Wright, Stephens et al. 1996). The FtsZ ring which is used in cell division is rapidly degraded following cell division. FtsZ is degraded from the swarmer compartment but not the stalked compartment of pre-divisional cells (Kelly, Sackett et al.

1998). The DnaA initiator of chromosome replication is rapidly degraded upon entry into stationary phase and upon sudden nutrient starvation (Gorbatyuk and Marczynski 2005).

Rational and Objectives

In bacteria, chromosome replication is regulated by changing the frequency of initiations. This regulation is presumably mediated by the DnaA protein since at least in E. coli, DnaA unwinds the oriC and starts the replisome assembly. Several mechanisms have been demonstrated which limit chromosome replication in E. coli by affecting DnaA. While both E. coli and C. crescentus use DnaA to initiate chromosome replication, they are different in that quickly growing E. coli are able to divide in less time than it takes to replicate the chromosome. This is accomplished by beginning subsequent rounds of replication prior to cell division. In contrast C. crescentus has chromosome replication strictly limited to once per cell cycle. This may require novel modes of regulating DnaA. I therefore set out to determine if the mechanisms identified in E. coli were conserved in C. crescentus.

My first objective was to determine if the regulatory inactivation of DnaA (RIDA mechanism) is also present in C. crescentus. Therefore, Chapter 2 describes how I created a conditional Hda depletion strain. I demonstrated that C. crescentus Hda (here termed HdaA) is essential. I also showed that loss of HdaA leads to over-initiation of chromosome replication.

Unexpectedly, the depletion of HdaA from the cells resulted in an increase in DnaA protein stability. Unlike DnaA in wild type C. crescentus cells, DnaA in cells lacking transcription of the hdaA gene was not completely degraded when these cells entered stationary phase.

Furthermore, DnaA became stabilized in growing cells coincident with loss of the HdaA protein, as measured by western blot. The HdaA protein was not absolutely required for degradation of

DnaA, as I was able to stimulate DnaA degradation in the absence of HdaA by shifting the cells to media lacking either nitrogen or carbon. This suggested that DnaA proteolysis during the

74 entry into stationary phase is not simply due to the cell sensing the decreasing levels of nutrients in the media, but rather DnaA proteolysis is a part of a developmental program. This result was confirmed using a strain with a disruption in the spoT gene required for the starvation induced degradation of DnaA. I showed that DnaA was cleared with equal rates in wild type (NA1000) and in the spoT deletion strain as they entered stationary phase. These observations suggested two possible mechanisms, either HdaA is acting as a chaperone targeting DnaA for degradation or HdaA inactivates DnaA by stimulating the hydrolysis of the bound ATP to ADP, and the DnaA-ADP is less stable than DnaA-ATP. The creation of DnaA mutants with reduced ATPase activity (detailed in Chapter four) display an increased protein stability, favoring the hypothesis that the stability of DnaA is regulated at least in part by its bound nucleotide.

In the third chapter, I set out to determine the role of a second protein with homology to DnaA (hdaB). C. crescentus encodes two genes with homology to dnaA, one hdaA with homology to the ATPase domain, and a second hdaB with homology to the DNA binding domain of DnaA. To explore the role of HdaB, I created an hdaB knock out strain by replacing the hdaB open reading frame with an antibiotic resistance cassette. I demonstrate that the HdaB is not essential for normal growth under laboratory conditions. I also demonstrate that purified HdaB protein lacks conspicuous DNA binding activity inside Cori. Although the hdaB gene is conserved in alphaproteobacteria, the role of HdaB remains to be determined.

In the fourth chapter, I explore the role of arginine fingers in DnaA regulation. The position of the arginine finger of C. crescentus differs from those in E. coli. There is a single arginine in the DnaA of C. crescentus which lies between the two characterized arginine fingers

75 of E. coli (DnaAR281 and R285). There is also an arginine in the ATPase domain which is required for the ATPase activity of DnaA in E. coli (EcDnaAR334) which is conserved in C. crescentus (CcDnaAR357). I have made single amino acid point mutants corresponding to these arginines in C. crescentus. I have shown that mutations in either arginine reduce DnaA ATPase activity in vitro. Mutations in the C. crescentus DnaAR357 cause increased chromosome replication as well as an increase in DnaA protein stability. Despite the effects of mutations in

C. crescentus DnaAR300 observed in vitro, the C. crescentus DnaAR300A behaved similarly to wild type DnaA in vivo. The DnaA content and DnaA stability are both comparable to wild type

DnaA: this is likely due to the effect of the ATPase activity induced by RIDA. The comparable mutation in E. coli (DnaAR285A) has no ATPase activity in vitro, however it’s ATPase may be stimulated in vivo by RIDA (Kawakami, Keyamura et al. 2005). In summary, my results show that while C. crescentus clearly uses a RIDA-like mechanism, it is a more sophisticated mechanism that uses both ATP hydrolysis and proteolysis to regulate DnaA activity.

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Chapter Two Identification and functional analysis of HdaA, a protein with homology to the ATPase

domain of DnaA

Abstract The protein that initiates chromosome replication, DnaA, is widely conserved among bacteria.

DnaA has been most extensively studied in Escherichia coli where only the ATP associated form of DnaA is able to initiate chromosome replication from oriC. Regulatory inactivation of DnaA

(RIDA) is the predominant mechanism which prevents chromosome over-replication in E. coli.

Two proteins are necessary for RIDA activity, the β-subunit of DNA polymerase III, DnaN, and a homolog of DnaA, Hda. Shortly after initiation of replication, Hda stimulates the conversion of the active ATP-DnaA to its inactive form ADP-DnaA. In E. coli, disruption of the hda gene increases in the proportion of ATP-DnaA and this increase overstimulates the initiation of chromosome replication.

I have identified a C. crescentus gene (hdaA) with homology to the E. coli hda gene. I have created a strain (GM3700) of C. crescentus with hdaA expressed under the control of a xylose dependent promoter. The GM3700 strain cannot form colonies when grown on media lacking xylose, demonstrating that the hdaA gene is essential for the normal growth of C. crescentus. More significantly, I have shown that HdaA is necessary to prevent multiple initiations of chromosome replication in the same cell cycle. Blocking hdaA expression causes an increased frequency of chromosome replication as well as a blockage of cell division.

Therefore, despite their apparent differences, both E. coli and C. crescentus share the RIDA mechanism. However, an unexpected consequence of removing HdaA from C. crescentus cells is that DnaA protein becomes stabilized in growing cells and it is not cleared from stationary phase cells. My experiments demonstrate a new role for HdaA in DnaA proteolysis that presumably works in exponentially growing cells to aid the RIDA mechanism. I also demonstrate that HdaA unexpectedly participates in the programmed transition from

92 exponentially growing cells to the stationary phase. This programmed transition, which normally removes DnaA protein from non-growing cell, is evolutionarily conserved among diverse Caulobacter species.

Introduction

While nearly all bacteria use the protein DnaA to initiate chromosome replication, bacteria responding to different environmental pressures may have evolved different mechanisms of

DnaA regulation. The free-living bacterium Caulobacter crescentus is a Gram-negative bacterium found in nutrient poor aqueous environments such as fresh water lakes and streams

(Poindexter 1981). C. crescentus divides asymmetrically to produce a motile “swarmer cell” and a non-motile “stalked cell”. Chromosome replication is tied to this cell morphology and cell cycle progression, as only the stalked cell is capable of chromosome replication, the swarmer cell must first differentiate into a stalked cell before it can replicate its chromosome. This cell cycle is different from E. coli which divides symmetrically producing two identical cells, both capable of chromosome replication. Rapidly growing E. coli may also begin subsequent rounds of chromosome replication prior to cell division. In contrast, C. crescentus chromosome replication occurs once and only once per cell cycle (Marczynski 1999). These differences in the overall cell cycle and in the timing of chromosome replication suggest that the mechanisms which regulate chromosome replication might have evolved differently in these two bacteria.

For example, in contrast to the E. coli DnaA protein, the C. crescentus DnaA protein is unstable:

C. crescentus DnaA is rapidly turned-over during the exponential phase growth and it is completely absent during the stationary phase (Gorbatyuk and Marczynski 2005). This difference suggests that C. crescentus utilises novel proteolytic mechanisms and perhaps other novel mechanisms to regulate DnaA.

E. coli possesses at least three mechanisms which prevent the over-initiation from one origin of replication. As reviewed in Katayama (Katayama 2001), E. coli possesses one mechanism that stimulates DnaA ATPase activity and two mechanisms that restrict DnaA binding to the origin of replication. As described in Chapter 1, the newly replicated oriC DNA becomes hemi-methylated and becomes “sequestered” from the DnaA replication proteins by the SeqA protein (Taghbalout, Landoulsi et al. 2000). DnaA is also regulated by titration by the datA locus which has a very high affinity for DnaA (Kitagawa, Mitsuki et al. 1996). However, the dominant mechanism in E. coli is the regulatory inactivation of DnaA (RIDA) that stimulates

DnaA ATPase activity and thereby increases the proportion of the inactive ADP-DnaA form

(Camara, Breier et al. 2005). RIDA requires the β-subunit of DNA polymerase III, DnaN, and

Hda. The Hda protein displays homology to the AAA+ domain of DnaA and it was initially identified in a genetic screen for high copy suppressors of a DnaN mutant (Kato and Katayama

2001). Hda mediated inactivation of DnaA is an important cell cycle process since Hda deficient

E. coli over-initiate replication from oriC (Camara, Breier et al. 2005).

While methylation of the C. crescentus chromosome by the CcrM methyltransferase has been shown to be essential, analysis of the C. crescentus genome has not revealed the presence of homologs to the SeqA protein. Also, no high-affinity sites capable of titrating DnaA away from the origin of replication have been identified in the C. crescentus genome. The absence of a SeqA homolog and the absence of a locus equivalent to datA suggest that their functions may be performed by other mechanisms in C. crescentus. Analysis of the C. crescentus genome does show a gene with homology to the ATPase domain of DnaA. This observation suggests that C. crescentus may use a RIDA-like mechanism to inactivate DnaA.

The natural habitats of C. crescentus are nutrient-poor oligotrophic aquatic environments. It is in these environmental niches that C. crescentus has evolved an asymmetric cell cycle, with an obligatory motile phase before chromosome replication. This cell cycle serves to limit competition for scarce nutrients between the newly formed swarmer cell and the parental stalked cell. In scarce nutrients environments, it would be deleterious to initiate overlapping rounds chromosome replication if the food resources were not sufficient to completely replicate the chromosome. The limiting of chromosome replication to one cell type prohibits the multiple overlapping rounds of chromosome replication. This strict control of chromosome replication may require novel mechanisms not found in E. coli.

One novel mechanism involves the response regulator protein CtrA. As described in chapter 1, the simplest regulatory model proposes that CtrA blocks chromosome replication in the swarmer cells while DnaA promotes chromosome replication in the stalked cells. However, even when much of the CtrA binding to the replication origin is removed, the timing of C. crescentus chromosome replication remains very similar to that of the wild type (Bastedo

Marczynski 2009). Such observations emphasize the importance of regulating C. crescentus DnaA activity. In contrast to the E. coli protein, C. crescentus DnaA is unstable during exponential growth so apparently proteolysis is one means of adjusting DnaA activity. Also, C. crescentus

DnaA is absent in stationary phase cells and C. crescentus DnaA is rapidly degraded in response to nutrient starvation in a SpoT dependent manner. SpoT is the signalling protein that makes the starvation induced signal that promotes the degradation of DnaA. Consequently, C. crescentus DnaA is also removed from cells when chromosome replication is disadvantageous in starved and in stationary phase cells.

In this chapter, I show that the loss of HdaA causes uncontrolled chromosome replication in accordance with a RIDA-like mechanism operating in C. crescentus. Unexpectedly,

I also show that the loss of HdaA stabilizes the DnaA protein in both exponential growing and in stationary phase cells. The starvation induced proteolysis of DnaA does not require HdaA, as I was able to starve cells and stimulate DnaA degradation in the absence of HdaA. These results argue that the degradation of DnaA during entry into stationary phase is not primarily due to starvation signaling when nutrients are consumed. Supporting this interpretation, I show by genetic deletion that SpoT, the signalling protein required for the starvation induced degradation of DnaA, is not required for the stationary phase degradation of DnaA. Therefore, I propose that the proteolysis of DnaA is accomplished by two mechanisms: The first mechanism requires HdaA and it targets DnaA for proteolysis in exponentially growing cells and as these cells transition from exponential growth into stationary phase. The second proteolysis mechanism is independent of HdaA and it responds to sudden changes in nutrient levels. My experiments therefore provide new roles for the key regulator HdaA and a new perspective on how a model bacterium transitions from logarithmic growth into the stationary phase.

Results

Caulobacter crescentus encodes a gene with homology to the ATPase domain of DnaA. The regulatory inactivation of DnaA (RIDA) has been shown to be the predominant mechanism to limit initiation of chromosome replication to once per cell cycle. Hda was originally identified in E. coli by biochemical and genetic methods and later showed to be encoded by a gene with homology to the ATPase domain of DnaA. Homologs of the hda gene are widespread in the proteobacteria. While RIDA has been extensively studied in E. coli, it has not been demonstrated to operate in other bacteria. A scan of the C. crescentus genome reveals a gene

(cc1711) with homology to the ATPase domain of DnaA in C. crescentus (Figure 1). An alignment of the amino acid sequences of C. crescentus DnaA (CC0008) with the putative Hda

(CC1711) reveals that the two proteins have 28% identity and 48% similarity in the amino acids sequence. Based upon this sequence similarity, cc1711 gene was initially annotated as a dnaA related gene. The cc1711 gene also displays homology with the hda gene of E. coli, 44 percent of the nucleotides conserved between the two genes. The presence of a gene with homology to hda suggests that it may be used by C. crescentus for a RIDA-like mechanism to limit chromosome replication to once per cell cycle.

Figure 1: A comparison of the domain structure of the DnaA protein with the HdaA protein

(A) The DnaA protein contains four functional domains: domain I is involved in protein-protein interactions, domain II a flexible linker domain, domain III is capable of binding and hydrolyzing

ATP, domain IV contains the DNA binding domain. The HdaA protein contains one functional domain which aligns with the ATP binding / ATPase domain of DnaA.

(B) An amino acid alignment of the predicted amino acids coded by the dnaA gene (cc0008) and the hdaA gene (cc1711) using the BLOSUM62 exchange weights matrix. The degree of conservation between amino acids of each protein is color coded from blue being not conserved to red being completely conserved, there is a 29% identity and 48% similarity in the amino acid sequences between the two proteins.

Figure 1: A comparison of the domain structure of the DnaA protein with the HdaA protein

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HdaA is essential in Caulobacter crescentus In the E. coli literature, there is disagreement as to whether the hda gene is essential, i.e. whether it is advantageous or absolutely required for cell viability. To test if C. crescentus hdaA gene is essential and to further explore its functions, I created a strain GM3700 with the only copy of hdaA under the control of the xylose inducible promoter, PxylX.

I created GM3700, an hdaA conditional expression strain, by replacing the coding sequence of hdaA (cc1711) with an “omega-cassette” conferring spectinomycin and streptomycin antibiotic resistance and by integrating a complementing copy of hdaA at the xylX locus immediately downstream from the xylose dependent promoter (PxylX). Creating GM3700 required the following steps: First the flanking 5’ and 3’ regions of the cc1711 gene were separately placed into the pNPTS138 vector that allows for DNA exchange by integration then excision with sacB counter selection. Then the omega-cassette was placed in between the flanking 5’ and 3’ regions of the cc1711 on pNPTS138. This plasmid was stably integrated at the cc1711 locus and this was confirmed by Southern blot analysis (Figure 2). The subsequent intermediate steps for creating GM3700 were also verified by Southern blot analysis (Figure

2B). Next, a copy of the hdaA gene under the control of a xylose inducible promoter was integrated at the PxylX locus. The sacB gene is lethal when the cells attempt growth in the presence of sucrose, shifting these cells to media containing sucrose selects for cells which have excised the pNPTS138 plasmid by homologous recombination. If the first and second recombination events occur on opposite sides of cc1711, then this gene will be replaced by the

“omega-cassette” genes. In fact, the second recombination event to replace the hdaA/cc1711 gene with the “omega-cassette” required the PxylX::cc1711 gene in trans. This observation was the first indication that HdaA is essential for the normal growth of C. crescentus.

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Figure 2: Creation of C. crescentus strain GM3700 that conditionally transcribes hdaA (cc1711)

The replacement of the natural hdaA open reading frame with the omega antibiotics resistance cassette required the placement of a cloned hdaA open reading frame downstream of the xylose regulated PxylX transcription promoter.

(A) Restriction map showing the SalI and NotI sites in the region of the cc1711 gene on the C. crescentus chromosome. The hdaA open reading frame is shown as a green arrow. The numbers indicate predicted fragment sizes when cut with SalI and NotI in wild type, 5’ integration of the pNTPS138 1711Ω, plasmid, 3’ integration of the pNTPS138 1711Ω, and replacement of the cc1711 gene with the Ω cassette.

(B) Southern blot of C. crescentus chromosomal DNA from wild type (NA1000), wild type

(NA1000) with the sacB selection vector (pNTPS138 1711Ω) integrated at the cc1711 locus, and the GM3700 strain with the cc1711 gene replaced by the omega cassette. The Southern blot was hybridized to a 32-P labeled probe with sequence homology to the region 500 bp immediately upstream of cc1711 coding region. The wild type (NA1000) with the pNTPS138

1711Ω vector integrated at the cc1711 locus appears as two bands on the Southern blot due to the presence of a region with homology to the region upstream cc1711 integrated into the pNTPS138 1711Ω vector. The band on the Southern blot in the lane with the G3700 strain appears higher because the omega cassette is larger than the cc1711 gene it replaced.

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Figure 2: Creation of C. crescentus strain GM3700 that conditionally transcribes hdaA (cc1711)

NA1000

+ or pNPTS138-1711Ω

GM3700 and

Pxyl cc1711

B NA1000 + NA1000 pNPTS138-1711Ω GM3700

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Blocked PxlyX::cc1711/hdaA transcription rapidly removes the HdaA protein To create strain GM3700, I placed cc1171/hdaA under the control of the xylose inducible promoter (PxylX). Expression from the PxylX promoter is induced by and requires the presence of xylose. To create the PxylX::cc1711 integrating plasmid, the open reading frame of hdaA minus the stop codon was cloned into pXTCYC-6 which creates a C-terminal fusion protein adding a tetra-cysteine tag to the end of HdaA. This plasmid cannot replicate in C. crescentus but must integrate into the chromosome at the xylX locus by homologous recombination.

GM3700 required the presence of xylose in the media to express hdaA, as for example, no colonies could form without xylose in the PYE rich media plates (Figure 3). I directly confirmed the conditional expression of HdaA protein by immune blot analysis (Figure 4). Also, the kinetics of HdaA protein disappearance was determined by comparing a GM3700 culture split into rich media with and without xylose (Figure 4). Most HdaA protein disappeared after 1-2 hours and it was not detectable after 4-5 hours. Therefore, this experiment also established that GM3700 is an efficient and rapid shut-off strain that can allow us to draw meaningful comparisons between the HdaA shut-off and other physiological consequences.

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Figure 3: HdaA is required for the normal growth of C. crescentus

The strain C. crescentus GM3700 has only one copy of hdaA that is transcribed from the xylose inducible PxylX promoter. GM3700 cells are unable to form colonies when plated on media lacking xylose (PYE). GM3700 colonies were observed only when 0.2% xylose was added to the media (PYEX). The wild type (NA1000) strain grew equally well on both PYE and PYEX plates.

Also note that GM3700 forms smaller colonies than NA1000 on PYEX. This poorer growth probably reflects insufficient expression of hdaA from the artificial PxylX promoter.

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Figure 3: HdaA is required for the normal growth of C. crescentus

PYEX

Wild type GM3700 (NA1000)

PYE

Wild type GM3700 (NA1000)

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Turning off hdaA permits cell growth while inhibiting cell division Once I established that HdaA performs an essential function(s) in the cell, to better understand how and when these cells die, I performed the following time-course and shut-off experiment.

The bacteria were grown overnight in PYE media supplemented with glucose and xylose

(PYEGX) and then transferred to PYEGX or PYEG (without xylose). Shifting these cells to PYEG stops transcription from the PxylX promoter and rapidly reduces the amount of HdaA in the cells (Figure 4).

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Figure 4: HdaA is cleared from GM3700 cells when shifted to media lacking xylose

Expression of hdaA was monitored by splitting a growing culture of GM3700 into media with

and without xylose. The HdaA protein in the cells was monitored by immunoblot analysis,

anti-hdaA antibodies generously provided by Justine Collier. Samples of a volume equivalent to

1.0 ml at an OD660 of 0.1 were taken at one hour time intervals. Purified His-tagged HdaA

protein (10ng) was used to confirm the size of the band corresponding to HdaA. The amount of

HdaA present in cells decreased 1 hour after the removal of xylose from the media. The

difference in abundance of HdaA between GM3700 cells grown in the presence of xylose and

those grown without xylose increased with time.

hours 0 1 2 3 4 5 His-HdaA xylose + - + - + - + - + - + - HdaA

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Blocked hdaA transcription and the absence of HdaA protein also inhibited cell division while permitting cell growth (Figure 5). The rate of growth was determined by optical density which increases as the bacterial protoplasm increases. The number of viable cells in the culture was also measured by preparing serial dilutions and plating on PYEX. These colonies were counted to determine the colony forming units per ml (CFU/ml). When the PxylX::hdaA was turned off, by shifting the cells from PYEGX to PYEG, there was a steady increase in the optical density accompanied by a decrease in the number of viable cells (compare Figure 5A with 5B).

After 4 hours post-shift, most of the HdaA protein (Figure 4) is gone and most of the cells fail to recover and do not from colonies on xylose containing plates (Figure 5B). Therefore, adding back xylose and restoring HdaA protein does not rescue these cells and yet they remain metabolically active since they continue to grow (Figure 5A). Under the microscope, these cells grow long and filamentous, i.e. they increase in length but they failed to divide (Figure 5C).

Apparently, when C. crescentus cells are growing, the loss of HdaA for as little as 4 hours creates an irreversible or a dead-end state.

When grown in rich media the GM3700 cells were slightly elongated even in the presence of xylose (PYEGX media, Figure 5C), potentially signifying incomplete complementation by hdaA supplied by the artificial PxyX promoter. When these cells were grown in minimal media supplemented with xylose (M2GX), they were indistinguishable from wild type cells (data not shown). This observation suggests that proper expression of HdaA is more important in rapidly dividing cells which presumably have a greater need for precise chromosome replication control.

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Figure 5: HdaA is required for normal growth of C. crescentus

GM3700 cells were grown in PYEGX, the culture was then split and the cells resuspended in either PYEGX or PYEG which turns off transcription of hdaA, leading to a depletion of the HdaA protein (Figure 4).

(A) Cell growth was monitored by optical density and serial dilutions were plated on PYEX plates to obtain the number of colony forming units CFU/ml. The optical density of the cultures increased with (PYEGX) or without (PYEG) the presence of xylose in the media.

(B) The number of viable cells decreased once HdaA had been depleted for two hours. This is in agreement with our observation that HdaA is cleared from the cell after 2 hours (Fig. 4).

(PYEGX) and when transferred to media lacking xylose (PYEG), these cells became filamentous without any visible pinching of the cell wall where cell division might take place. This division defect was not observed when transferred to fresh media containing xylose.

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Figure 5: HdaA is required for normal growth of C. crescentus

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Figure 5: HdaA is required for normal growth of C. crescentus

C GM3700

PYEG 20 hours

GM3700

PYEGX 20 hours

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Loss of HdaA leads to over-initiation of chromosome replication As E. coli Hda (Kato and Katayama 2001) and the homologous protein from C. crescentus HdaA

(Collier and Shapiro 2009) were both implicated in suppressing chromosome over-replication, I also investigated this role for HdaA in our C. crescentus strain (GM3700) but I used a more sensitive assay for chromosome replication. In C. crescentus, over-replication produces unmethylated DNA because the chromosome is normally re-methylated towards the end of each cell cycle by the CcrM DNA methyltransferase (Zweiger, Marczynski et al. 1994). As explained in

Chapter 1, when the first replication fork passes through fully-methylated DNA, it produces two un-methylated strands and hence two hemi-methylated duplex DNAs. If a second replication fork passes before the methylation sites can be re-methylated, then this will produce one hemi- methylated duplex DNA and one un-methylated duplex DNA. Such un-methylated DNA can be differentiated from both fully-methylated and hemi-methylated DNA by Southern blot analysis, as only un-methylated DNA is susceptible to cutting by HinfI restriction endonuclease. The C. crescentus origin (Cori) of replication contains multiple recognition sites for HinfI (Figure 6A).

Such an assay is very sensitive and it focusses on replication just from Cori. So, unlike DNA- fluorescence assays, e.g. FACS assays that count chromosomes per cell, this assay does not require that DNA synthesis extends over long distances.

I applied this replication assay to the xylose regulated HdaA strain (GM3700) grown on rich media (Figure 6B) and on minimal media (Figure 6C) as follows: Exponentially growing

GM3700 cultures were supplemented with xylose and glucose and then shifted to media supplemented with only glucose or glucose and xylose. Samples were taken at the indicated times for Southern blot analysis. The chromosome DNA was cut with BamHI and HinfI and the

Southern blot membranes were hybridized with 32-P labeled DNA derived from the cloned Cori

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BamHI fragment (Figure 6A). This analysis reveals a 1.6 kb Cori BamHI band and smaller bands if Cori DNA becomes susceptible to cutting by HinFI. In rich media, such un-methylated Cori

DNA becomes detectable only after 6 hours post shift and after 8 hours the lower unmethylated DNA bands are clearly visible (Figure 6B). A similar result is seen in minimal media

(Figure 6C). Un-methylated Cori DNA was never observed in the wild type strain (NA1000)(data not shown) and it was never detected in GM3700 cultured with xylose (Figures 6BC). I conclude that loss of HdaA causes over-replication from Cori and that HdaA is required for the normal chromosome replication cycle.

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Figure 6: Blocked expression of hdaA leads to over-replication of chromosomal DNA

(A) Location of CcrM methylation/ HinfI target sites at the C. crescentus origin of replication

(Cori). Multiple recognition sites (GAnTC) for CcrM DNA methyltransferase are located in Cori.

These sites are methylated in a cell cycle dependent manner.

(B) C. crescentus depleted of HdaA display an increased frequency of chromosomal replication.

GM3700 cells were grown in (PYE) supplemented with xylose and the culture was then split and resuspended in media with or without xylose. Samples were taken at the indicated time points and total chromosomal DNA was extracted. The chromosomal DNA was digested with the restriction endonucleases BamHI and HinfI and used for Southern blot analysis. Southern blots were hybridized with [32-P] radio-labelled DNA prepared from a 1.7 kb BamHI Cori fragment

(shown above in 6A). The restriction endonuclease HinfI only cuts completely unmethylated

DNA produced by the passage of two replication forks during the same cell cycle. Unmethylated

DNA was observed only after 6 and 8 hours without xylose.

(C) When GM3700 cells were grown in minimal media with xylose (M2GX) unmethylated DNA was observed 8 hours after blocking hdaA transcription by removal of xylose (M2G) from the media. Unmethylated DNA was never observed in wild type (NA1000, not shown) or in GM3700 cells grown with xylose.

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Figure 6: Blocked expression of hdaA leads to over-replication of chromosomal DNA

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To support these results, I also demonstrated that loss of HdaA causes C. crescentus cells to accumulate extra chromosomes. I used flow cytometry (FACS analysis) to measure the total amount of DNA per cell after removal of xylose from the media. As expected, removal of xylose from GM3700 cultures substantially increased the amount chromosome DNA in these cells (Figure 7). Even with complementation (xylose), a sub population contains greater than two chromosomes equivalents of DNA. This is likely due to incomplete complementation. The proportion of cells with greater than two chromosome equivalents is increased two hours after xylose (hdaA transcription) was removed from the media. It is at this time that the lower levels of HdaA are observed (Figure 4).

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Figure 7: Depletion of HdaA results in uncontrolled initiation of chromosome replication

The GM3700 strain was grown over night in PYE supplemented with glucose and xylose, the cells were pelleted and resuspended in either PYE with glucose (G) or PYE with glucose and xylose (GX). The cultures were grown for 1, 2, 3 and 4 hours labelled T1, T2, T3, and T4 respectively. The DNA content of the conditional hdaA expression strain GM3700 was measured by flow cytometry. The cells were treated with rifampicin (to block initiation of replication) and cephalexin (to block cell division). overnight. The cells were fixed in 70% ethanol and the chromosomes were stained with propidium iodide. The quantity of DNA per cell was measured using a BD FACSCalibur (BD

Biosciences) and the Cell Quest Pro acquisition software (BD Biosciences). Analysis of the data was performed using the FlowJo software (TreeStar Inc.)

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Clearing of DnaA protein during stationary phase requires HdaA In addition to its role as the initiator of chromosomal replication, DnaA also acts as a transcription factor that modulates its own transcription in E. coli (Speck, Weigel et al. 1999). It had been previously shown that removal of Hda from E. coli shifts the activation state of DnaA from the predominantly inactive ADP-DnaA state to the predominantly active ATP-DnaA state

(Kato and Katayama 2001). In E. coli, this shift causes decreased dnaA transcription and less

DnaA synthesis. I therefore, examined if removing HdaA from C. crescentus strain GM3700 would affect the level of DnaA in the cell. I used an immunoblot assay to examine the abundance of DnaA in the GM3700 strain. I did not see any obvious change in the levels of

DnaA protein in growing GM3700 cells, with or without xylose (e.g. Figures 8, 10, 11 and data not shown). Therefore, blocking the expression the hdaA gene does not significantly influence

DnaA protein synthesis and presumably dnaA transcription is not dramatically affected.

However, during such experiments I discovered that blocking the expression of the hdaA gene does block the removal of DnaA protein when cells enter stationary phase. I used the immunoblot assay to examine the abundance of DnaA in wild type C. crescentus (NA1000) as well as in the GM3700 strain. Very unexpectedly, DnaA protein was not cleared from GM3700 cells during their entry into stationary phase when xylose was removed (Figure 8A, lane 8). As all of these cultures grew to the same optical cell densities, I conclude that DnaA protein is retained in these GM3700 cells because HdaA is not present to somehow promote DnaA proteolysis. This conclusion implies a new and unexpected role for HdaA.

To confirm that this retention of DnaA is due to loss of HdaA, I stimulated the expression of hdaA by adding a trace amount (0.02%) of xylose to the stationary phase GM3700 cells. I anticipated that if HdaA promotes proteolysis then restoring HdaA should remove the retained

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DnaA. As in the previous experiment, the immunoblot experiments in Figure 8B confirm that

GM3700 cells grew to saturation but retained DnaA (compare lane 2 without xylose to the control in lane 1). As anticipated, adding only 0.02% xylose to these stationary phase cells completely removed DnaA (Figure 8B, lane 4). This 0.02% xylose addition did not stimulate cell growth (data not shown), yet it is sufficient to activate the PxylX promoter. Therefore, HdaA can operate to remove DnaA in the absence of cell growth. This experiment also indicated that while these cells suffer from division defects they are still metabolically active, since DnaA proteolysis requires an ATP-dependent mechanism and a ClpP-dependent protease. Adding higher concentrations of xylose (0.2%) also reduced but did not completely remove all of the

DnaA protein (Figure 8B, lane 3). Since the 0.2% xylose did stimulate growth, some of this DnaA may come from new protein synthesis and not incomplete proteolysis. Adding glucose did not clear DnaA from stationary phase GM3700 cells (data not shown).

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Figure 8: Clearing by proteolysis of DnaA during stationary phase requires HdaA

The clearing of DnaA from stationary phase cells requires hdaA expression. (A) GM3700 and wild type (NA1000) cells were grown in rich media supplemented with xylose, the cells were transferred to media with or without xylose and samples were taken at 0 hours and 20 hours.

Samples were adjusted to the equivalent of 1 ml at an optical density (A660) of 0.1 and immunoblots were performed with anti-DnaA antiserum. The initial abundance of DnaA was similar between the GM3700 and wild type cells. In contrast to both wild type cells and

GM3700 grown in media containing xylose, the GM3700 cells grown in media without xylose had not cleared DnaA at the 20 hour time point, when the cells had reached stationary phase.

(B) GM3700 cells were grown to stationary phase in M2G minimal media with xylose (lane 1) or without xylose (lane2). The clearing of DnaA from stationary phase cells was stimulated by restarting hdaA transcription by adding xylose to the media (lanes 3 and 4). Adding only 0.02% xylose did not stimulate cell growth but it completely cleared the DnaA protein (lane 4).

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Figure 8: Clearing by proteolysis of DnaA during stationary phase requires HdaA

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HdaA also has a role in the logarithmic growth phase proteolysis of DnaA C. crescentus DnaA protein is also unstable and targeted for proteolysis during logarithmic growth. To determine if HdaA causes DnaA instability only during stationary phase (Figure 8) or if this role extends to growing cells, I measured DnaA instability with the following shut off experiment: First, a logarithmically growing GM3700 culture was split into with xylose (PYEGX) and without xylose (PYEG) media for 180 min. This split created parallel growing cultures with and without HdaA. Next, both cultures were treated with the antibiotic tetracycline to block translation, sampled at time intervals and the amount of DnaA present was measured by immunoblot analysis (Figure 9A). DnaA protein rapidly disappeared from the PYEGX (with xylose/HdaA) cells with an approximate half-life of 30 min. In contrast, DnaA protein slowly disappeared from the PYEG (without xylose/HdaA) cells with an approximate half-life of 120 min. Significant amounts of DnaA protein remained after 150 min in these PYEG cells compared to undetectable levels of DnaA protein after 150 min in PYEGX (Figure 9A).

Since I chose a fixed 180 min period to prepare the without xylose/HdaA cells, I next modified the experiment in Figure 9A by varying this shut off period for 0, 2 and 5 hours. Then tetracycline was added and cells were sampled for DnaA immunoblot analysis at T = 0 and at T

= 4 hours (Figure 9B). For all conditions, 4 hours is sufficient to degrade all DnaA protein, except when the xylose/HdaA shut off is 5 hours. In this case, about half the DnaA protein remains after 4 hours (Figure 9B, far right lanes). This experiment showed that a shut off period of 2 hours is too short while 3 hours (Figure 9A) is sufficient to block normal DnaA proteolysis.

In summary, HdaA clearly promotes DnaA proteolysis when cells are growing logarithmically.

Presumably, HdaA also uses proteolysis to adjust DnaA activity so as to initiate chromosome replication at the correct time in the stalked cell-type.

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Figure 9: HdaA promotes DnaA proteolysis in logarithmically growing cells

(A)GM3700 cells were grown in rich media supplemented with glucose and xylose (PYEGX) the cells were transferred to media with or without xylose and grown for 180 minutes. Tetracycline was added to halt translation, and thus production of new DnaA. To determine the stability of

DnaA at the time when tetracycline was added, aliquots of equivalent optical density were taken at 30 minute time points. Immunoblot analysis was used to determine DnaA abundance.

DnaA was not cleared from the cells that were depleted of xylose (HdaA) for 180 minutes or more.

(B) To determine how long after removal of xylose (hdaA transcription) DnaA was stabilized.

GM3700 cells were grown in rich media supplemented with glucose and xylose (PYEGX) the cells were transferred to media with or without xylose and grown for 0, 2, or 5 hours.

Tetracycline was added to halt translation and thus production of new DnaA. To determine the stability of DnaA at the time when tetracycline was added aliquots of equivalent optical density were taken at time 0 and after 4 hours. Immunoblot analysis was used to determine DnaA abundance. DnaA is cleared up until at least 2 hours after transcription of hdaA was blocked in

GM3700.

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Figure 9: HdaA promotes DnaA proteolysis in logarithmically growing cells (A) DnaA stability 3 hours after termination of hdaA transcription

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Figure 9: HdaA promotes DnaA proteolysis in logarithmically growing cells (B) DnaA stability 0, 2, and 5 hours after termination of hdaA transcription

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HdaA does not significantly affect dnaA transcription from log to stationary phase The cellular abundance of DnaA is determined by a balance between its synthesis and proteolysis. I examined the abundance of DnaA in the GM3700 strain but I did not see any significant change in the levels of DnaA protein in logarithmically growing GM3700 cells with or without xylose (e.g. Figure 8, 10, 11 and data not shown). Therefore, removing HdaA does not profoundly influence DnaA protein synthesis and presumably dnaA transcription is not significantly affected during logarithmic growth. This situation is very different than that of E. coli where Hda provides a negative feedback to down regulate DnaA transcription and synthesis. However, since GM3700 cells without xylose accumulate more DnaA protein when they exit logarithmic growth and enter stationary phase, it is formally possible that dnaA genetic transcription increases at this time and that both higher transcription and lower proteolysis account for DnaA protein accumulation.

I therefore tested the possibility that HdaA regulates transcription when cells transition from logarithmic to stationary phase with the following experiment: The 5’ dnaA promoter region was placed upstream of lacZ on the transcription reporter plasmid as described in the experimental procedures and this plasmid was introduced into GM3700 cells. During logarithmic grow, these cells contain about 525 Miller units of LacZ activity (Figure 10). This culture was split into with xylose M2GX and without xylose M2G media and both cultures were left to enter stationary phase. These cells displayed about 400 Miller units of β-galactosidase activity activity (Figure 10). The drop from 525 to 400 Miller units indicates that β-galactosidase activity does not keep pace with cell growth as the cells enter the stationary phase. So dnaA transcription is probably down-regulated at this time. However this regulation does not involve

HdaA since both M2GX and M2G cultures contain very similar β-galactosidase activity

127 measured in Miller units (Figure 10). Therefore, the presence of DnaA in stationary phase

GM3700 cells is due to a decrease in proteolysis alone and not an increase in dnaA transcription.

128

Figure 10: Transcriptional activity of dnaA promoter in absence of HdaA

Blocked expression of hdaA does not alter transcription of the dnaA gene as the cells transition from logarithmic growth into stationary phase. The level of dnaA transcription was monitored using a LacZ reporter construct. GM3700 was grown with a plasmid containing a lacZ reporter gene under the control of the dnaA promoter. The GM3700 strain was grown in M2GX. The culture was split and one half grown in M2G, and the other half in M2GX for 20 hours. The lacZ expression was measured in the original culture and in both the stationary phase cultures with hdaA expression (M2GX) and without hdaA expression (M2G). While a slight decrease in expression was observed no significant difference was seen between stationary phase cells with or without the presence of HdaA.

129

At least two mechanisms regulate DnaA proteolysis in logarithmically growing C. crescentus C. crescentus DnaA is degraded during the logarithmic growth phase and during periods of nutrient starvation. For example, when the nitrogen source ammonium is removed from the growth media, the degradation of DnaA is accelerated and it is quickly lost from these cells. I have shown that DnaA proteolysis is significantly decreased in logarithmically growing cells if

HdaA is removed (Figure 9). This retention of DnaA in growing cells was unexpected. HdaA somehow promotes DnaA degradation in rapidly growing cells replete with nutrients.

Therefore theoretically, HdaA might also accelerate DnaA degradation when nutrients are suddenly withdrawn and in cells that experience starvation.

To determine if C. crescentus HdaA is also required for starvation-accelerated DnaA proteolysis, I tested the degradation of DnaA by nutrient starvation as follows: Wild type and

GM3700 cells were grown in minimal media (M2G) with xylose then their cultures were split into the same media with and without xylose. These cells were grown for four hours to remove

HdaA from the GM3700 cells and the abundance of DnaA was measured by immunoblot analysis. As shown in the left panel of Figure 11A, all 4 cultures, wild type and GM3700, with and without xylose all had comparable amounts of DnaA protein. Next, cells from all 4 cultures were placed in starvation media, i.e. the same media but lacking the nitrogen source ammonium (M2G –N) for 2 more hours. The immunoblot in the right panel of Figure 11A shows that for all 4 cultures, the DnaA protein was efficiently removed by nitrogen starvation and barely detectable. Therefore, GM3700 cells lacking HdaA have efficient starvation- accelerated DnaA proteolysis indistinguishable from that of wild type cells. Apparently, this proteolysis mechanism, which involves SpoT to produce a starvation signal (Lesley and Shapiro

2008), does not use HdaA to degrade DnaA.

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Figure 11: Proteolysis of DnaA can be stimulated by starvation independent of HdaA

C. crescentus cells which have been depleted of HdaA retain the ability to degrade the DnaA protein in response to sudden starvation conditions.

(A) Wild type C. crescentus (NA1000) and GM3700 were grown in M2GX and transferred to either M2GX or M2G to block hdaA transcription. The cultures were grown for 4 hours to deplete HdaA from the cells. DnaA abundance was measured by immunoblot analysis. Both wild type and the GM3700 strain with or without xylose rapidly degraded DnaA when placed in a medium lacking nitrogen.

(B) The proteolysis of DnaA which had been stabilized by entry into stationary phase without expression of hdaA could be stimulated by starving the cells of nitrogen. The abundance of

DnaA was decreased in cells which had been resuspended in M2G-N, while the abundance of

DnaA increased when the cells were resuspended in fresh M2G, or M2GX.

131

Figure 11: Proteolysis of DnaA can be stimulated by starvation independent of HdaA

132

At least two mechanisms regulate DnaA proteolysis in stationary phase C. crescentus I have also shown that DnaA is retained in stationary phase cells if HdaA is removed (Figure 8).

This retention of DnaA was even more unexpected because our lab previously regarded the entry into stationary phase as a response to nutrient starvation and there was no obvious connection between nutrient starvation and the established roles for E. coli Hda. Furthermore, the preceding experiments in Figure 11A argue that HdaA is not involved in starvation induced degradation of DnaA. Therefore, if cells enter stationary phase solely because they are starved, then DnaA should have been completely removed from the stationary phase GM3700 cells without xylose.

To explain this paradox, I needed to re-evaluate how C. crescentus enters stationary phase. Since stationary phase is a substantially different metabolic state, I propose that HdaA coordinates DnaA proteolysis with the transition into the stationary state. I also propose that this transition must be gradual since C. crescentus needs time to adjust. This condition requires that C. crescentus cells commit to stationary phase long before the nutrients are exhausted and it is likely that some nutrients remain during the stationary phase at sufficiently high concentrations to maintain basal metabolism. This consideration predicts that a sudden and complete removal of an essential nutrient such as the sole nitrogen source will be treated very differently by C. crescentus cells.

I therefore determined if nutrients remain in C. crescentus stationary phase and if HdaA is required for starvation-induced DnaA proteolysis, as follows: GM3700 cells were grown in minimal media with xylose (M2GX), split into the same media with and without xylose (M2GX and M2G) and allowed to reach stationary phase. As shown by immunoblot analysis, the left two lanes in Figure 11B reproduce the expected result that stationary phase GM3700 cells

133 retain DnaA protein only without xylose (M2G), and therefore without HdaA. This stationary phase culture was then split 3-ways and its cells were placed into fresh media without ammonium as the sole nitrogen source (M2G-N), and into control fresh M2G and M2GX media.

These split cultures were incubated for 2 more hours and analyzed on the same immunoblot as shown in the right 3 lanes of Figure 11B. The M2G-N lane clearly shows that most of the DnaA protein, that was stable in the stationary phase cells, became unstable and rapidly degraded when all of the ammonium was removed. The 2 adjacent lanes show that these cells were viable when returned into fresh media M2GX and M2G. The extra DnaA protein seen in these 2 control lanes is due to revived growth and probably new DnaA synthesis. Therefore, stationary phase cells are not completely starved for nitrogen and when the HdaA-dependent mechanism is blocked, a separate starvation induced mechanism can still degrade the DnaA protein.

Nutritional cues that promote DnaA proteolysis are present in the exhausted growth media but they are not utilized by GM3700 cells lacking HdaA It might be argued that GM3700 cells retain DnaA in stationary phase not because they lack

HdaA but because they consume fewer nutrients and that they are therefore not starved as much as the wild type cells. This interpretation is unlikely because both wild type and GM3700 cells grow to the same optical cell densities in the same media (data not shown). To show that both wild type and GM3700 respond differently to exactly the same nutrients, I first prepared nutrient exhausted growth media as follows: Both wild type and GM3700 cells were grown to stationary phase in rich PYE media without xylose. The cells were removed from the exhausted media by centrifugation and the cleared supernatant was used to re-suspend either wild type or GM3700 cells that were growing in PYE with or without xylose. Following 2 hour incubations, the DnaA immunoblots in Figure 12 demonstrate that wild type and GM3700 cells

134 respond differently to the same nutrient exhausted media. While the GM3700 cells retain

DnaA protein, the wild type cells rapidly remove all DnaA protein. Note that exhausted media from either wild type or from GM3700 cultures is sufficiently depleted of nutrients to cause wild type cells to remove all DnaA. Yet the same exhausted media does not cause the GM3700 cells to do the same and DnaA protein is retained. This difference is due to lack of HdaA since

GM3700 cells grown without xylose retain more DnaA (Figure 12 and data not shown). That

GM3700 cells grown on xylose also retain DnaA under these circumstances is probably due to the insufficient expression of hdaA from the relatively weak PxylX promoter, as noted above

(Figure 3).

135

Figure 12: Effect of depleted media on DnaA abundance in wild type and GM3700 cells

Extracellular cues that trigger proteolysis of DnaA from stationary phase cultures are present but not utilized in cells lacking HdaA. Growing wild type cells (NA1000) or GM3700 cells were resuspended in media that had been previously used to grow wild type C. crescentus (NA1000) or GM3700 cells to saturation. The depleted media was cleared of cells by high speed centrifugation. The abundance of DnaA was measured by immunoblot analysis. The depleted media was able to stimulate the degradation of DnaA from wild type cells but not from the

GM3700 cells.

136

Figure 12: Effect of depleted media on DnaA abundance in wild type and GM3700 cells

137

SpoT is not required for the stationary phase degradation of DnaA My results suggested that DnaA degradation involves two mechanisms one starvation induced and a second one linked by HdaA to the transition into stationary phase. It has been previously reported that SpoT produces the messenger molecule ppGpp that is somehow necessary for the degradation of DnaA due to carbon starvation (Lesley and Shapiro 2008). To determine the role of nutrient starvation in the stationary phase degradation of DnaA, I determined the abundance of DnaA in growing and stationary phase cultures of C. crescentus lacking SpoT. To confirm the published effect, i.e. that the spoT C. crescentus strain does not stimulate DnaA degradation when suddenly starved, these cells growing in M2G were re-suspended in media lacking glucose (the sole carbon source). As expected, I observed that DnaA was rapidly cleared from wild type cells but was cleared much more slowly from the spoT knock out strain (data not shown), as previously reported (Lesley and Shapiro 2008).

Therefore, to determine if SpoT is required for DnaA degradation as the cells transition from logarithmic growth to the stationary phase, the abundance of DnaA was measured by immunoblot from parallel wild type and spoT null cultures. Time points were taken at two hour intervals as the cells entered stationary phase. Note that DnaA was cleared at the same rate from both cultures (Figure 13). This result clearly indicates that SpoT, and therefore the starvation signal ppGpp, does not significantly contribute to DnaA degradation during this transition period.

138

Figure 13: SpoT is not required for the removal of DnaA upon entry into stationary phase

(A) Immunoblot analysis to determine the abundance of DnaA in wild type and ΔspoT as they transitioned from growing cells to stationary phase cells. Samples of equivalent optical density were taken at two hour intervals and DnaA abundance was determined by immunoblot analysis. The abundance of DnaA decreased in both wild type and the ΔspoT strain.

(B)The graph shows the optical densities of the cultures at the time points in panel A. The rate of growth of the wild type (NA1000) and ΔspoT was monitored by the optical density of the cultures. Both cultures reached stationary phase at the same rate.

139

Figure 13: SpoT is not required for the removal of DnaA upon entry into stationary phase

140

Proteolysis of DnaA is conserved among freshwater Caulobacters C. crescentus DnaA is degraded during entry into stationary phase. To determine if this regulated DnaA proteolysis is unique to C. crescentus or if it is more generally used by other bacteria, I assayed for DnaA protein in extracts from three additional fresh water Caulobacter species under both exponential growth and stationary phase conditions. The serum raised against C. crescentus DnaA detects a single protein band of comparable size in the related

Caulobacter species (Figure 14). Interestingly, all of the freshwater Caulobacter species in

Figure 14 show significantly reduced levels of DnaA in stationary phase (S) compared to logarithmic phase growth (L). These results argue that proteolysis of DnaA is not incidental to our standard laboratory strains of C. crescentus. Instead, proteolysis of DnaA appears to be a generally conserved mechanism for regulating this essential replication protein.

141

Figure 14: Proteolysis of DnaA during stationary phase is conserved among related fresh water Caulobacter species.

DnaA is cleared from C. crescentus (WT, NA1000) and from related fresh water Caulobacter species (FWC) during stationary phase. DnaA abundance was measured by immunoblot analysis. Samples were taken from log phase growing cells (L) and after the same culture reached stationary phase (S), approximately 20 hours later. All samples were adjusted to a volume equivalent of 1 ml at an optical density of 0.1. Lanes 3-10 show that each of these 4

Caulobacters decreases DnaA abundance in stationary phase cells relative to log phase cells.

Control lanes 1-2 show that C. crescentus strain GM2471, with dnaA under the control of the xylose inducible promoter, produces DnaA with xylose (X) but not with glucose (G).

142

Discussion The control of chromosome replication in bacteria occurs at the level of initiation. The protein

DnaA is responsible for initiating chromosome replication in nearly all bacteria, including all human pathogens. There are currently no antibiotics which target DnaA. While regulation of

DnaA has been extensively studied in the bacterium E. coli, it has not been established that these mechanisms are conserved among other bacteria. Other bacteria may have evolved mechanisms to regulate DnaA and chromosome replication which are not seen in E. coli.

I have identified a gene in C. crescentus, hdaA (cc1711) , which is similar to hda of E. coli.

Analysis of the predicted amino acid sequence reveals homology to the ATPase domain of

DnaA. I have created a strain of C. crescentus which requires xylose for the expression of hdaA.

I have demonstrated that C. crescentus HdaA is essential and that blocking hdaA expression leads to greater than once per cell cycle initiation of chromosome replication, inhibition of cell division, accumulation of multiple chromosomes per cell and loss of viability. Intriguingly loss of

HdaA not only affects the activity of DnaA but also affects the stability of the DnaA protein.

Southern blot analysis of the C. crescentus chromosome reveals the accumulation of unmethylated DNA at the origin of replication (Cori). This is a strong indication that the DNA has been replicated at least twice in the same cell cycle. Interestingly Southern blot analysis does not indicate the presence of unmethylated DNA when probed at a point further from the origin (data not shown). This is consistent with what was observed in E. coli Hda deficient strains. Using flow cytometry to measure the DNA content of cells, I showed that removal of xylose from the media leads to an increase in the number of chromosome equivalents in the cells when HdaA is depleted in the strain where the only copy of hdaA is under the control of

143

Pxyl. Microscopic analysis of GM3700 cells with blocked hdaA expression reveals that the cells become filamentous.

The effect of HdaA on DnaA abundance was measured by immunoblot analysis with poly clonal DnaA antibodies. There was no difference observed in relative abundance of DnaA between growing wild type cells, or GM3700 cells with or without the expression of HdaA.

However GM3700 cell with blocked expression of hdaA failed to clear DnaA from the cell upon growth to stationary phase. Several possibilities for this phenotype were considered. Since the loss of HdaA negatively affects viability and the proteolysis of DnaA requires energy in the form of ATP, the DnaA detected in the cells lacking HdaA could be from dead or metabolically inactive cells. I rejected this possibility because I could stimulate the proteolysis of DnaA by the addition of xylose to the media, indicating that the DnaA protein was present inside metabolically active cells capable of responding to xylose and presumably synthesizing more

HdaA which in turn must have mobilized a mechanism of selective proteolysis. Blocked expression of hdaA in GM3700 cells alters DnaA stability in logarithmic phase and in stationary phase. DnaA stability increases as early as within three hours after the xylose was removed from the media containing GM3700 cells. This is concurrent with the decrease of HdaA abundance in the cell as measured by immunoblot assay.

In C. crescentus, DnaA is actively degraded upon entry into stationary phase as well as during nutrient starvation. I have shown that HdaA is not required for the nutrient starvation induced proteolysis of DnaA. The starvation of GM3700 cells with blocked hdaA expression was able to stimulate the proteolysis of DnaA. The fact that GM3700 cells with blocked expression of hdaA in stationary phase were able to degrade DnaA when starved for nitrogen indicates

144 that these cells might not be experiencing starvation conditions even during stationary phase.

Therefore, I examined the properties of the media; I could stimulate the proteolysis of DnaA from wild type cells suspended in the media collected from stationary phase GM3700 cells with blocked hdaA expression. However, transferring GM3700 cells to media obtained from wild type cells in stationary phase did not result in the degradation of DnaA. This suggests that factors other than a simple lack of nutrients in the media cause DnaA proteolysis in an HdaA dependent manner.

The acceleration of DnaA proteolysis upon nutrient starvation requires SpoT. I have shown that spoT is not required for the degradation of DnaA upon entry into stationary phase.

This is consistent with our previous results indicating that at least two distinct mechanisms govern DnaA stability: one HdaA dependent mechanism which is required upon entry into stationary phase and one SpoT dependent mechanism required for the starvation induced proteolysis of DnaA.

The regulation of DnaA by proteolysis is not limited to C. crescentus. I examined three other fresh water Caulobacter species. Immunoblot analysis with an antibody raised against C. crescentus DnaA revealed a cross reacting band of approximately the same size as DnaA. This band was present in growing cells but dramatically reduced in stationary phase cells.

While this work was in progress Collier and Shapiro (2009) independently created a conditional expression strain for the cc1711 gene that encodes HdaA. They concluded that the hdaA gene is essential for the normal growth of C. crescentus as they were unable to obtain a disruption of the cc1711 locus without the presence of a complementing copy of the gene on a plasmid. They showed by flow cytometry that C. crescentus that has been depleted of HdaA

145 accumulates extra chromosomes and by Southern blot analysis that there is an increase in the frequency of chromosome initiation. A fluorescently labelled HdaA was used to show that HdaA associates with the replisome during the cell cycle. These results confirmed work I had previously done. We both showed that HdaA was essential due to the inability to disrupt the cc1711 locus without complementation. Additionally, I have shown that depleting HdaA results in cells which lose their ability to proceed normally through the cell cycle. Collier et al. did not observe a defect in viability when their conditional cc1711 expression strain was shifted to media lacking the inducer (xylose) to express the cc1711 gene (Collier, personal communication). This discrepancy is likely due to the presence of a tetracysteine tag on the

HdaA protein produced in GM3700, which may have made the protein less stable so that it could be depleted from the cell more quickly allowing for the observation of the phenotype. In addition to flow cytometry, I employed a Southern blot based assay to identify chromosomal

DNA which had been replicated more than once in the same cell cycle. This allowed me to determine the time interval between the removal of the inducer of hdaA transcription and its effect on chromosome replication. Collier et al. also employed a technique based on the

Southern blot to quantify the numbers of copies of the DNA at the origin with the copy number of the DNA at the terminus. The results of this type of quantification can be difficult to interpret as a growing culture would normally have an increased content of DNA from the region surrounding the origin, which is replicated early compared with DNA replicated later on from the terminus.

In addition to the role of HdaA in regulating DnaA activity, presumably through a RIDA like mechanism, I have uncovered a novel role for HdaA in regulating the stability of DnaA in

146 both log cell and stationary phase cells, which is different than the starvation induced proteolysis of DnaA. I propose that DnaA stability is linked to the state of the nucleotide (ATP or ADP) bound to DnaA. Evidence for this hypothesis will be presented in chapter four of this thesis.

147

Experimental Procedures Bacterial strains and growth conditions The strains and plasmids used in this study are listed in Table 1. C. crescentus strains were grown in peptone-yeast extract (PYE) rich media or M2G minimal glucose media. Media used for the growth strains requiring the presence of xylose was supplemented with 0.2% xylose and

0.2% glucose (PYEGX, or M2GX). Xylose wash out experiments were performed by washing the cells twice in PYE supplemented with 2% glucose.

Table 1: Strains and Plasmids

Strain/Plasmids Description Source E. coli Strains DH5α Cloning strain Invitrogen BL21(DE3) Protein expression strain

C. crescentus NA1000 wild type C. crescentus Evinger and Agabian (1977) synchronizable strain GM3700 C. crescentus with xylose This work dependent expression of hdaA ΔspoT C. crescentus with a genetic Lesley and Shapiro (2008) deletion of the spoT gene GM2471 Conditional dnaA expression Gorbatyuk and Marczynski strain (2005) FWC14 Independent isolate of fresh John Smit (UBC) water Caulobacter species FWC21 Independent isolate of fresh John Smit (UBC) water Caulobacter species FCW38 Independent isolate of fresh John Smit (UBC) water Caulobacter species

Plasmids pGM2727 dnaA promoter reporter plasmid pGM2876 pET19B-dnaA C. crescentus his- Shaheen et al.(2009) dnaA expression vector

148

Creation of the hdaA conditional expression strain I created a conditional hdaA expression strain by replacing the coding sequence of cc1711 with an omega (spectinomycin and streptomycin antibiotic resistance) cassette. Regions of homology extending 1Kb upstream (5’GATATCTCCTCAGGGTCCGGCGTGGTGAACGTC, 5’

GGATCCGCGAGGTTTTGCTTA TGTTGACCACGAGCG) and downstream

(5’GATATCGCCGTGACGCTCCTTCTCCCCTTGCG, 5’AAGCTTCGGCCTCTTCCTCGATCTCGAG) of the cc1711 gene were PCR amplified and cloned into the non-replicating vector pNPTS138. The omega cassette was then inserted between the two regions of homology. A stable integration of this plasmid into the cc1711 locus was confirmed by Southern blot analysis. The disruption of the cc1711 locus required the presence of a hdaA gene complementing in trans. A copy of the hdaA gene under the control of a xylose inducible promoter was integrated at the xylX locus.

The predicted hdaA open reading frame was amplified by PCR (5’

GGTACCTTGTCCACCCAGTTCAAACTGCCGC , 5’ GAGCTCCTACCCCTCATCCCCCTCGAACC) and cloned into the pXTCYC-6 vector, creating a xylose inducible hdaA marked with an tetracysteine tag. The HdaA depletion strain was only able to grow in media supplemented with xylose.

Nutrient starvation of C. crescentus C. crescentus cells were starved for nutrients by growing the cell in minimal media (M2G, or

M2GX) then splitting the culture and washing and re-suspending in minimal media prepared without nitrogen (M2G-N, or M2GX-N) or minimal media prepared without a carbon source

(M2).

Flow cytometry The chromosomal content of the cells were determined using flow cytometry. Cells were treated with 60 μg ml−1 rifampicin to block new RNA synthesis preventing new initiations of

149 chromosomal replication and 35 μg ml−1 cephalexin to prevent cell division, for three hours to let any chromosomes that had begun replication to proceed to completion. The number of chromosome peaks observed would then be equivalent to the number of origins of replication present when the antibiotics were added. Cells were then washed in staining buffer (50 mM sodium citrate, 10 mM Tris-HCl, 1 mM EDTA, pH = 7.2), and permeabilized by adding ethanol to a final concentration of 70%. The RNA was removed from the cells by the addition of RnaseA and the DNA stained with propidium iodide (20 μg ml−1). The data was acquired using the FL2 detector of a BD FACSCalibur (BD Biosciences) and the Cell Quest Pro acquisition software (BD

Biosciences). Analysis of the data was performed using the FlowJo software (TreeStar Inc.)

Antibody production

Polyhistidine tagged C. crescentus DnaA protein was expressed in E. coli BL21(DE3) from the pGM2876 plasmid. The tagged protein was purified as previously described (Li and Crooke

1999) and used to immunize two rabbits for the production of polyclonal antibodies (McGill

Animal Resources Centre). Antibody specificity was determined by western blot analysis and by

35-S immunoprecipitation, using a DnaA conditional expression strain (GM2471).

Immunoblot analysis Immunoblot analysis was used to determine the cellular levels of DnaA. Whole cell lysate were resolved using 8% SDS-PAGE. Immunoblots were performed using DnaA anti-serum diluted

1:10,000 and anti-rabbit immunoglobulin G conjugated to horseradish peroxidase at a 1:10,000 dilution. Blots were visualised with Chemiluminescent ECL (Perkin Elmer).

150

References Camara, J. E., A. M. Breier, T. Brendler, S. Austin, N. R. Cozzarelli and E. Crooke (2005). "Hda inactivation of DnaA is the predominant mechanism preventing hyperinitiation of Escherichia coli DNA replication." EMBO Rep 6(8): 736-741. Collier, J. and L. Shapiro (2009). "Feedback control of DnaA-mediated replication initiation by replisome- associated HdaA protein in Caulobacter." J Bacteriol 191(18): 5706-5716. Evinger, M. and N. Agabian (1977). "Envelope-associated nucleoid from Caulobacter crescentus stalked and swarmer cells." J Bacteriol 132(1): 294-301. Gorbatyuk, B. and G. T. Marczynski (2005). "Regulated degradation of chromosome replication proteins DnaA and CtrA in Caulobacter crescentus." Mol Microbiol 55(4): 1233-1245. Katayama, T. (2001). "Feedback controls restrain the initiation of Escherichia coli chromosomal replication." Mol Microbiol 41(1): 9-17. Kato, J. and T. Katayama (2001). "Hda, a novel DnaA-related protein, regulates the replication cycle in Escherichia coli." EMBO J 20(15): 4253-4262. Kitagawa, R., H. Mitsuki, T. Okazaki and T. Ogawa (1996). "A novel DnaA protein-binding site at 94.7 min on the Escherichia coli chromosome." Mol Microbiol 19(5): 1137-1147. Lesley, J. A. and L. Shapiro (2008). "SpoT regulates DnaA stability and initiation of DNA replication in carbon-starved Caulobacter crescentus." J Bacteriol 190(20): 6867-6880. Li, Z. and E. Crooke (1999). "Functional analysis of affinity-purified polyhistidine-tagged DnaA protein." Protein Expr Purif 17(1): 41-48. Marczynski, G. T. (1999). "Chromosome methylation and measurement of faithful, once and only once per cell cycle chromosome replication in Caulobacter crescentus." J Bacteriol 181(7): 1984-1993. Poindexter, J. S. (1981). "The caulobacters: ubiquitous unusual bacteria." Microbiol Rev 45(1): 123-179. Shaheen, S. M., M. C. Ouimet and G. T. Marczynski (2009). "Comparative analysis of Caulobacter chromosome replication origins." Microbiology 155(Pt 4): 1215-1225. Speck, C., C. Weigel and W. Messer (1999). "ATP- and ADP-dnaA protein, a molecular switch in gene regulation." EMBO J 18(21): 6169-6176. Taghbalout, A., A. Landoulsi, R. Kern, M. Yamazoe, S. Hiraga, B. Holland, M. Kohiyama and A. Malki (2000). "Competition between the replication initiator DnaA and the sequestration factor SeqA for binding to the hemimethylated chromosomal origin of E. coli in vitro." Genes Cells 5(11): 873-884. Zweiger, G., G. Marczynski and L. Shapiro (1994). "A Caulobacter DNA methyltransferase that functions only in the predivisional cell." J Mol Biol 235(2): 472-485.

151

Chapter Three

Identification and analysis of HdaB, a novel protein with homology to the DNA-binding

domain of DnaA

152

Abstract

The correct timing of chromosome replication in C. crescentus requires a homolog of DnaA called HdaA that displays homology to the AAA+ (ATP/ADP binding) domain III of DnaA.

Intriguingly, a second gene with homology to DnaA is present in C. crescentus. I called this second gene hdaB since it displays significant homology to the DNA binding domain IV of DnaA.

Sequence analysis of hdaB suggests that it is a paralogue of DnaA that retains key amino acids used by DnaA to make sequence-specific contacts with the DnaA R box. Homologs of HdaB are found in the genomes of other alphaproteobacteria, suggesting that hdaB has an evolutionary conserved role in these bacteria.

To address the function of hdaB in C. crescentus, I created a strain called ∆1058 where the entire hdaB coding sequence was deleted from the chromosome. Light microscopy showed that ∆1058 cells are indistinguishable from wild type C. crescentus cells. Similar side by side comparisons showed that both ∆1058 and wild type C. crescentus grow with very comparable rates and they both retained comparable cell viabilities when shifted from rich medium to poorer growth medium. Since HdaA participates in the proteolysis of DnaA, I similarly tested how ∆1058 and wild type C. crescentus cultures degrade DnaA protein. However, DnaA immunoblots showed that both cultures responded indistinguishably to sudden carbon starvation and to entry into stationary phase.

To test the DNA-binding potential of HdaB, I affinity-purified a GST-HdaB fusion protein from an engineered E. coli strain. The binding affinity of the established DNA-binding protein

CtrA was compared with that of HdaB by the electrophoretic mobility shift assay (EMSA) and by the DNAase I protection footprint assay. Both EMSA and footprint assays used specific DNA

153 targets from the C. crescentus chromosome origin of replication Cori. These DNA binding targets contained established CtrA and DnaA binding sites. Affinity purified a GST-CtrA protein produced conspicuous EMSA band-shifts while comparable concentrations of GST-HdaB protein did not, even though these DNA targets also contained the strongest Cori DnaA binding sites.

Similarly, GST-CtrA protein produced conspicuous footprints on 32-P end labeled Cori DNA while comparable concentrations of GST-HdaB protein and HdaB (with the GST tag removed by thrombin cleavage) did not produce any detectable footprints.

Therefore, considering the negative results of these in vivo and in vitro experiments, the role of the hdaB gene remains to be determined. Considering the homology and conservation of hdaB, my hypothesis is that HdaB is involved in chromosome replication and/or genetic transcription. C. crescentus and other alphaproteobacteria may have evolved more subtle mechanisms to help them adapt rapidly to changes in the environment but these were not reproduced in the laboratory or went undetected by the assays performed.

154

Introduction

DnaA binds DNA at 9 base pair elements called “DnaA boxes” (5` TTATNCACA). The E. coli origin of replication (oriC) contains 5 DnaA boxes (Fuller, Funnell et al. 1984). In addition to the DnaA boxes, oriC also contains lower affinity DnaA binding sites, termed I sites (McGarry, Ryan et al.

2004). The high affinity binding sites for DnaA are occupied throughout most of the cell cycle, while the lower affinity sites and I sites are occupied only at the time of initiation (McGarry,

Ryan et al. 2004). When in the ATP bound state, DnaA binds cooperatively to the lower affinity sites, causing a melting of the DNA in the adjacent AT-rich DNA-unwinding element (DUE)

(Kowalski and Eddy 1989). In addition to its role as the initiator, DnaA also acts as a transcription factor (Messer and Weigel 1997). DnaA boxes are present in the promoter regions of many genes where they can mediate repression or transcriptional activation.

Caulobacter crescentus similarly employs DnaA as the initiator of chromosomal replication and as a transcription factor which regulates several genes involved in chromosome replication. DnaA initiates chromosomal replication by binding to several sites located within the origin of replication (Cori). Encoded within Cori are multiple binding sites for DnaA, two sites with high affinity for DnaA termed G-boxes (Marczynski and Shapiro 1992) and five lower affinity DnaA binding termed W-boxes (Taylor, Ouimet et al. 2011).

DnaA is composed of four functional domains (Sutton and Kaguni 1997). Domain III contains the ATP-binding region homologous to the AAA+ ATPase proteins. DnaA is found in two states, an ATP bound state and an ADP bound state. Only the ATP bound form of DnaA is

“active” or capable of initiating oriC replication in vitro. In E. coli cells that were synchronized using a temperature sensitive DnaC mutant, the ratio of ATP-DnaA / ADP-DnaA varied in a cell

155 cycle dependant manner (Kurokawa, Nishida et al. 1999). This variation allows DnaA to act as a cell cycle regulator. Also as described in the previous 2 chapters, DnaA is primarily regulated by a homolog of DnaA domain III called Hda in E. coli and HdaA in C. crescentus. This precedent suggests that bacteria can use protein domains as modules for novel regulatory pathways.

DnaA domain IV is responsible for protein-DNA interactions at the DnaA boxes (Roth and Messer 1995). Domain IV contains a helix-turn-helix motif which makes contacts along both the major and minor grooves of the DnaA box (Fujikawa, Kurumizaka et al. 2003) . Several residues have been identified as being important for sequence specific contacts, including

DnaA- Arg339 which contacts the DNA minor groove to recognize the TTA sequence of the

TTATNCACA 9-mer consensus sequence. The TNCACA sequence is recognized by a helix inserted in the DNA major groove (Fujikawa, Kurumizaka et al. 2003). Mutations in DnaA positions

Asp433, His434 and Thr435, cause defective DnaA box recognition without affecting non- specific DNA binding (Sutton and Kaguni 1997, Blaesing, Weigel et al. 2000). These amino acids which are believed to be responsible for making sequence-specific contacts are conserved in the DnaA protein of C. crescentus where they correspond to DnaA-Asp456, His457 and Thr458.

I will show an alignment of a novel protein termed HdaB with DnaA, where these amino acids are conserved and therefore imply a role in DNA-binding for HdaB as well.

The C. crescentus genome encodes not one but two proteins with homology to DnaA.

The HdaA protein displays homology with the ATPase domain III of DnaA (Collier and Shapiro

2009). It has been shown that HdaA is essential in preventing multiple initiations of chromosome replication in the same cell cycle (Collier and Shapiro 2009)(Chapter two, of this thesis). Interestingly, C. crescentus has a second gene (cc1058) of unknown function encoding a

156 protein with homology to DnaA. Here termed HdaB, this novel protein displays homology to the DNA binding domain IV of DnaA. While E. coli contains no obvious homolog of cc1058, I will show that cc1058/hdaB homologs are found throughout the alphaproteobacteria. Chapter 3 presents my systematic attempts to find a function for hdaB.

157

Results

Caulobacter crescentus encodes a protein with homology to the DNA binding domain of DnaA. Three amino acids of DnaA have been shown by mutational analysis to be required for DNA sequence-specific binding (E. coli DnaA positions Asp433, His434 and Thr435). In E. coli, mutations in any one of these amino acids cause a defect in DnaA box recognition without affecting non-specific DNA binding (Sutton and Kaguni 1997, Blaesing, Weigel et al. 2000).

These amino acids which are believed to be responsible for making sequence-specific contacts are conserved in the DnaA protein of C. crescentus and correspond to Asp456, His457 and

Thr458. Presumably, this conservation explains how C. crescentus DnaA binds the very similar

G boxes in Cori and how it has an even higher affinity for the E. coli-like R boxes (Taylor,

Ouimet, et al. 2011).

An analysis of the C. crescentus genome reveals the presence of a novel gene cc1058 with homology to the DNA binding domain of DnaA, here called hdaB (Figure 1). The amino acid sequence has a 23 percent identity with the amino acid sequence of the last 70 amino acids of DnaA. An alignment of HdaB with DnaA shows that two of the three amino acids are conserved HdaB -Arg80 and -Thr82 the third amino acid HdaB –Arg82 is similar to that of DnaA, changing from a histidine to an arginine (Figure 2). These results predict that HdaB should act as a DNA binding protein, recognizing a similar sequence motif as the DnaA protein.

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Figure 1: A comparison of the domain structure of the DnaA protein with the predicted HdaB protein (A) The DnaA protein contains four functional domains: domain I is involved in protein-protein interactions, domain II is a flexible linker domain, domain III is capable of binding and hydrolyzing ATP, domain IV contains the DNA binding domain. The HdaB protein contains one functional domain which aligns with the DNA binding of DnaA.

(B) An amino acid alignment of the predicted amino acids coded by the dnaA gene (cc0008) and the hdaB gene (cc1058). The degree of conservation between amino acids of each protein is color coded from blue being not conserved to red being completely conserved.

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Figure 1: A comparison of the domain structure of the DnaA protein with the predicted HdaB protein

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Figure 2: Alignment of a highly conserved section of the DnaA binding domain from E. coli and C. crescentus with the HdaB protein. Alignment of a highly conserved section of the DnaA binding domain from E. coli and C. crescentus with the HdaB protein. Amino acid that are believed to interact with DnaA binding sites on the Chromosome are indicated with arrows. The Domain IV of E. coli DnaA is shown bound to DNA with the DnaA box TTAATCCACA. The atomic structures and the protein-DNA contact information are from Fujikawa, Kurumizaka, et al. (2003).

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Phylogenetic analysis of hdaB distribution Phylogenetic analysis reveals that HdaB homologs are present in most of the alphaproteobacteria. The presence of an hdaB homolog was detected in 36 members of the alpha proteobacteria representing the orders Rhizobiales, Caulobacterales, Rhodobacterales,

Parvularcula. The presence of hdaB in other members of the alphaproteobacteria is suggestive that it provides a selective advantage.

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Figure 3: List of bacteria among the alphaproteobacteria which contain a hdaB gene A blast (Max E value 0.1 and min percentage identity 10) search was used to identify species among the alphaproteobacteria which carry a gene that encodes a protein that is homologous to the DNA binding domain of DnaA. The results were sorted alphabetically.

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Construction of the hdaB deletion strain The C. crescentus strain ∆1058 (GM3705) was created by replacing the coding sequence of hdaB

(cc1058) with an “omega cassette” conferring spectinomycin/streptomycin antibiotic resistance. To create this deletion, the flanking 5’ and 3’ regions of the cc1058 gene were cloned into the pNPTS138 sacB counter selection vector. The pNPTS138 vector is not capable of autonomous replication in C. crescentus. A stable integration of this plasmid was selected based upon the ability to grow in media containing kanamycin. The sacB gene is lethal when the cells are grown in the presence of sucrose. When plated on media containing sucrose a second recombination event occurred either replacing the hdaB/cc1058 gene with the “omega cassette” or reverting to the wild type locus. Sucrose resistant colonies, which had excised the pNPTS138 plasmid, were then screened for resistance to kanamycin and to spectinomycin / streptomycin. Colonies which were sucrose resistant, kanamycin sensitive and spectinomycin / streptomycin resistant were presumed to have replaced the cc1058 locus with the omega antibiotic resistance cassette (Figure 4). Colony screening by PCR was used to test for the presence or absence of the cc1058 locus. If HdaB did not serve an important function, then on average the cc1058 locus should have been deleted and replaced by the “omega cassette” 50% of the time. However, contrary to expectations, I observed that less than 5% of the sucrose resistant colonies gave the expected antibiotic resistance (Figure 4) and PCR amplification results (data not shown). The strain with the confirmed replacement of the cc1058 locus was designated ∆1058 (GM3705) and it was used for further studies.

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Figure 4: Creation of an hdaB (cc1058) knock out strain Regions of homology flanking hdaB (cc1058) were cloned into the pNPTS138 plasmid, the omega cassette which confers spectinomycin and streptomycin resistance was cloned between the two regions of homology. The plasmid was then electroporated into NA1000 and transformed cells were grown overnight in PYE with spectinomycin and streptomycin and then plated onto PYE plates supplemented with 2% sucrose. Sucrose resistant colonies were then screened for the replacement of the cc1058 locus. Successful replacement of the cc1058 locus would result in cells which are resistant to sucrose (PYE- Sucrose), resistant to spectinomycin and streptomycin (PYE-Spec/Strep) but sensitive to kanamycin (PYE- Kan). The strains were plated on PYE plates with no selection as a positive control.

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Figure 4: Replacement of the cc1058 locus with the omega antibiotic resistance cassette

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Growth of the wild type and the hdaB deletion strain ∆1058 Since cells with a genetic deletion of hdaB are viable, but obtained a lower than expected frequency, I attempted to identify a phenotype in the deletion strain that would give a selective advantage to the wild type strain. One possible explanation for the increased retention of the wild type cc1058 locus (after the sucrose selection step) is that this locus provides a growth advantage, so that wild type cells will proliferate faster than the ∆1058 cells. Therefore, the growth rates were measured. Wild type and Δ1058 cells were grown in rich media (PYE) overnight and diluted back to an OD660 of 0.1. The optical density was taken each hour as a measure of cell growth. The disruption of the hdaB gene did not alter the growth rate of the cells and the wild type and Δ1058 growth curves were almost identical (Figure 5). The doubling time for both wild type C. crescentus (NA1000) and the cc1058 deletion strain were both approximately 137 minutes. The increase in optical density is a measure of the culture’s ability to grow and assimilate nutrients to increase the cell mass. Since I was unable to detect a difference in the growth between wild type cells and those lacking the hdaB gene, HdaB does not seem to be essential for normal cell growth under laboratory conditions. Light microscopy observations also showed that the cells from both cultures had the sizes and shapes typical of wild type C. crescentus cells (Figure 5). These results do not preclude the requirement for hdaB under other conditions such as for example extremely slow growth that occurs in the natural environment.

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Figure 5: The hdaB gene is not required for the normal growth C. crescentus (A) The growth rate of wild type (NA1000) and Δ1058 strains was determined by optical density.

Both wild type and Δ1058 cultures were grown in PYE media overnight and diluted back to an

OD660 of 0.1. The optical density was taken each hour as a measure of the increase in cell bulk.

(B) The overall morphology of wild type (NA1000) and Δ1058 strain was determined by microscopic analysis. The morphology of both wild type (NA1000) cells and Δ1058 cells reveals no obvious changes in the appearance of the cells.

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Figure 5: The hdaB gene is not required for the normal growth C. crescentus.

Δ1058

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Effect of nutritional stress on deletion strain ∆1058 The hdaB gene is not required for normal growth under laboratory conditions; however it may provide a selective advantage under certain growth conditions. In nature, bacteria are often in stressful conditions due to rapid changes in the environment. DnaA protein is itself degraded when nutrients are removed. To simulate a natural stress in the laboratory, I grew both wild type C. crescentus (NA1000) and the ∆1058 deletion strain in rich media (PYE) and then spotted agar plates made with minimal media (M2G). This treatment suddenly exposes cells to lower nutrients. However, as shown in Figure 6, no differences in viability or in colony sizes were observed between the wild type and the ∆1058 deletion strain. Shifting the cells from minimal media to rich media gave similar null results (data not shown). These results indicate that hdaB does not appear to be required to help C. crescentus adapt to rapid changes in nutrient levels.

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Figure 6: Environmental stress simulated by a sudden change in nutrients In nature bacteria must deal with sudden changes in their environment and these stresses are mitigated in the carefully controlled growing environment of a laboratory. As a result, genes that are essential in nature to respond to these changes may be dispensable in a laboratory setting. To evaluate a potential role for the hdaB gene in responding to a sudden nutrient stress, both wild type C. crescentus (NA1000) and the Δ1058 strain cells were grown in rich media (PYE), serial dilutions were then placed onto either rich media or minimal media plates.

We did not observe a difference in the survival of cells plated on minimal media plates or rich media plates.

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Figure 6: Environmental stress simulated by a sudden change in nutrients

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Effect of ∆1058 on proteolysis of DnaA The DnaA protein is regulated by proteolysis during stationary phase and during sudden nutrient starvation. The promoter region of the dnaA gene contains a DnaA binding site that might theoretically bind HdaB. One potential role for HdaB is to fine tune the expression of dnaA during nutrient starvation or upon entry into stationary phase. To determine if HdaB has a role in the expression or in the stability of the DnaA protein, the relative abundance of DnaA in wild type C. crescentus (NA1000) and the Δ1058 strain was determined by immunoblot, for growing cells, for cells which have been starved for carbon, and for stationary phase cells. Wild type C. crescentus (NA1000) and the Δ1058 strain were grown in minimal media (M2G) overnight and diluted back to an optical density (660nm) of 0.1. The cells were then allowed to grow for two doubling times and an aliquot was transferred to minimal media prepared without a carbon source (M2 without glucose) for two hours. The remainder of the cells were left to grow and enter stationary phase overnight. The volume of the sample taken for immunoblot analysis was adjusted to be the equivalent of 1ml at an optical density (660nm) of 0.1. As shown in Figure 7, DnaA was cleared indistinguishably from the wild type (NA1000) and the

Δ1058 strain in stationary phase and after carbon starvation. Therefore, there are no significant differences between the wild type (NA1000) and the Δ1058 strains in either the synthesis or the degradation of DnaA protein.

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Figure 7: Effect of hdaB (cc1058) on DnaA proteolysis The relative abundance of DnaA in wild type C. crescentus (NA1000) and the Δ1058 strain was determined by immunoblot, for growing cells, cells which have been starved for carbon, as well as stationary phase cells. Wild type C. crescentus (NA1000) and the Δ1058 strain were grown in minimal media (M2G) overnight and diluted back to an optical density (660nm) of 0.1. The cells were then grown for two doubling times and an aliquot was transferred to minimal media prepared without a carbon source (M2 without glucose) for two hours. The remainder of the cells were left to grow overnight and reach stationary phase. The volume of the sample taken for immunoblot analysis was adjusted to be the equivalent of 1ml at an optical density (660nm) of 0.1.

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Electrophoretic Mobility Shift Assay (EMSA) analysis of HdaB binding potential Sequence analysis of hdaB suggests that it encodes a protein consisting almost entirely of the

DNA binding domain (IV) of DnaA (Figure 1). An artificial protein based only on E. coli domain IV selectively binds the 9-mer DnaA box and this accounts for the full length DnaA protein binding to the E. coli oriC (Fujikawa, Kurumizaka et al. 2003). The DnaA protein of C. crescentus has been shown to bind the origin of replication of C. crescentus at multiple high and low affinity binding sites (Taylor, Ouimet, et al. 2011).

To determine if the HdaB protein is capable of interacting with DnaA binding sites, I performed an electrophoretic mobility assay (EMSA). The purified GST-HdaB protein was incubated with annealed oligonucleotides derived from Cori (termed C and E) which contain both DnaA binding sites and CtrA binding sites. The protein DNA mixture was run on a polyacrylamide gel and stained with ethidium bromide. As shown in Figure 8, the positive control protein CtrA bound DNA as expected and retarded its progress through the gel. The purified HdaB protein did not interact with the oligonucleotides.

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Figure 8: Electrophoretic mobility shift assay Electrophoretic mobility shift assay (EMSA) was used to test binding of HdaB to DnaA binding sites on a double stranded oligonucleotide. The EMSA was carried out using the following oligonucleotide pairs which were PAGE-purified and annealed at 1 µM in 10 mM Tris (pH

7.9).The oligonucleotide pairs C (5’-CTTCCCACATGGGGTTAACGCTCTGTTAATCATGGGGATG-3’) and E (5’-CAGGGCAATGGTTAAGCAACCGTTAACGGATGATGATCCACAGGAGAGTCTG-3’) contained both a CtrA binding site (underlined) as well as a binding site for DnaA (in bold). The third oligonucleotide pair, R (5’-GGTTCGCAGCATCTAGCCTCTGGTGAAGGCAGG-3’) was made up of a randomized nucleotide sequence. The oligonucleotide pairs were incubated with either HdaB or

CtrA for 30 min and loaded onto a native 8% polyacrylamide gel which had been pre-run in 1 ×

TBE to resolve bound and free oligonucleotides. The gel was stained with ethidium bromide and the position of the oligonucleotides visualized under ultra violet light. The positive control protein CtrA bound DNA as expected and retarded it progress through the gel. The purified

HdaB protein did not interact with the oligonucleotides.

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Figure 8: Electrophoretic mobility shift assay

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HdaB binding potential tested by DNase I protection footprint assays The DnaA protein of C. crescentus has been shown to bind the origin of replication of C. crescentus at multiple high and low affinity binding sites (Taylor, Ouimet, et al. 2011). I also assessed the ability of HdaB to interact within Cori by means of a DNase I footprint assay. This assay is more sensitive than the previous EMSA and it can test binding to more DNA sequences than those chosen for the EMSA oligonucleotides. As shown in Figure 9, the positive control protein CtrA bound DNA as expected. Zones of protection can be clearly seen at the five well characterized CtrA binding sites. The HdaB protein was purified with a GST tag. The GST tag was cleaved using the thrombin enzyme. Neither the GST-HdaB nor HdaB cleaved of the GST tag showed any protection of the Cori DNA. There are several possible explanations for these negative results. HdaB may bind to a DNA sequence not present in Cori (something other than a DnaA box), HdaB may bind DnaA boxes with the aid of accessory proteins, or the purified protein may have become denatured and non-functional.

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Figure 9: DNase I protection footprint assay The potential binding of HdaB to sequences in the origin of replication of C. crescentus (Cori) was assessed by a DNase I protection footprint assay. A fragment of Cori fragment labeled with

32P at the 5′ end containing all five CtrA binding sites (labeled by red bars) as well as two high- affinity DnaA and the five low affinity DnaA binding sites. The HdaB protein was purified with a

GST tag. The GST tag was cleaved using the thrombin enzyme. The 0.1 μM of GST-CtrA or a 10 fold dilution series of 0.1 μM HdaB protein was incubated a room temperature for 30 minutes prior to digestion with DNase I. The DNA was then purified and analyzed by 5% sequencing gel.

The gel was then exposed to x-ray film (Kodak). Neither the GST-HdaB nor HdaB cleaved of the

GST tag showed any protection of the Cori DNA.

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Figure 9: DNase I protection footprint assay

180

Experimental Procedures Bacterial strains and growth conditions

The strains and plasmids used in this study are listed in Table 1. C. crescentus strains were grown in peptone-yeast extract (PYE) rich media or M2G minimal glucose media. Media used for the growth strains requiring the presence of xylose was supplemented with 0.2% xylose and

0.2% glucose (PYEGX, or M2GX). Xylose wash out experiments were performed by washing the cells twice in PYE supplemented with 2% glucose.

Table 1: Strains and Plasmids Strain/Plasmids Description Source

E. coli Strains DH5α Cloning strain Invitrogen BL21(DE3) Protein expression strain

C. crescentus NA1000 wild type C. crescentus Evinger and Agabian (1977) synchronizable strain GM3705 C. crescentus with a genetic This work deletion of the hdaB gene

Plasmids pGM2589 pGEX::CtrA C. crescentus gst-ctrA Spencer et al.(2009) expression vector pGM2876 pET19B::dnaA C. crescentus his- Shaheen et al.(2009) dnaA expression vector pGM2904 pGEX::HdaB C. crescentus gst- hdaB expression vector

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Creation of a hdaB knock out strain

I have created a conditional hdaB knock out strain by replacing the coding sequence of cc1058 with an omega (spectinomycin/streptomycin antibiotic resistance) cassette as previously described. Regions of homology extending 1Kb upstream

(5’GGATCCGCCGGGCCTCTACGCCATCG, 5’GATATCGGCGTCCTCTCCTCCAGGCC) and downstream

(5’GATATCTGACCGGCGGGCTTCAGATCC, 5’AAGCTTTGTGGCGGGGCGAATGCTTCGTC) of the cc1058 gene were PCR amplified and cloned into the non-replicating vector pNPTS138. The omega cassette was then cloned between the two regions of homology. The pNPTS138 plasmid with the regions of homology and the omega antibiotic resistance cassette were transformed into wild type (NA1000) C. crescentus cells. Cells with the plasmid integrated into the chromosome were selected for based upon their ability to grow in the presence of kanamycin. I was able to disrupt the cc1058 locus without the need to complement the gene in trans. The presence of a complementing copy of the hdaB gene did not alter the frequency at which I was able to replace the cc1058 locus with the omega antibiotic resistance cassette.

Immunoblot analysis

Immunoblot analysis was used to determine the cellular levels of DnaA. Whole cell lysate were resolved using 8% SDS-PAGE. Immunoblots were performed using DnaA anti-serum was diluted

1:10,000 anti-rabbit immunoglobulin G conjugated to horseradish peroxidase 1:10,000 dilution and visualised with Chemiluminescent ECL (Perkin Elmer).

Electrophoretic mobility shift assay (EMSA)

I used the electrophoretic mobility shift assay (EMSA) to identify binding of HdaB with DnaA binding sites on a double stranded oligonucleotide. The EMSA were carried out using the

182 following oligonucleotide pairs which were PAGE-purified and annealed at 1 µM in 10 mM Tris

(pH 7.9).Two oligonucleotide pairs contained both a CtrA binding site as well as a binding site for DnaA. The third oligonucleotide pair was made up of a randomized nucleotide sequence.

The oligonucleotides pairs were incubated with either HdaB or CtrA for 30 min and loaded onto a native 8% polyacrylamide gel which had been pre-run in 1 × TBE to resolve bound and free oligonucleotides. The gel was stained with ethidium bromide and the position of the oligonucleotides visualized under ultra violet light.

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DNase I protection footprint analysis

DNase I footprinting was performed as previously described (Kawakami, Keyamura et al. 2005).

A fragment of Cori fragment labeled with 32P at the 5′ end containing all five CtrA binding sites as well as two high-affinity DnaA and the five low affinity DnaA binding sites. The Dnase I assay was performed using 0.1 μM GST-CtrA, 0.1 μM GST-HdaB and a 10 fold dilution series of 0.1

μM HdaB (GST-HdaB with the GST tag cleaved by thrombin digestion). The proteins were incubated a room temperature for 30 minutes prior to digestion with DNase I. The DNA was then purified and analyzed by 5% sequencing gel. The gel was then exposed to x-ray film

(Kodak).

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Discussion

The mechanisms controlling chromosome replication have been most extensively studied in E. coli. However, the E. coli model of chromosome replication may not be sufficient to account for the variability in life styles found among bacteria. A broader understanding of the dynamics of chromosome replication in bacteria could lead to the development of novel antibiotics against pathogenic bacteria as well a better understanding of the role of bacteria in the environment.

C. crescentus is a free living Gram negative bacteria which undergoes chromosome replication once and only once per cell cycle (Marczynski 1999). There are a number of similarities and differences between C. crescentus and E. coli in the manner in which they regulate chromosome replication. For example, both C. crescentus and E. coli require the protein DnaA for the initiation of chromosome replication (Fuller, Funnell et al. 1984, Gorbatyuk and

Marczynski 2001). However, the mechanisms which control chromosome replication in E. coli are not found in all bacteria. C. crescentus does not encode an obvious homolog of the SeqA protein nor has it an equivalent of the datA locus. C. crescentus does depend on a RIDA like mechanism to limit chromosome replication to once per cell cycle. C. crescentus actually encode two proteins with homology to DnaA, a homolog of Hda called HdaA, which displays homology to the ATPase domain of DnaA, as well as a second gene called hdaB which is predicted to encode a protein with homology to the DNA binding domain of DnaA. Sequence analysis of the hdaB gene predicts that it encodes a protein of 13.76 KiloDaltons, protein that is homologous to the DNA binding domain of DnaA. The role of the hdaB gene has yet to be elucidated.

185

Homologs of the hdaB gene are conserved among other members of the alphaproteobacteria. I found hdaB in the sequenced genomes of the Rhizobiales, Caulobacterales,

Rhodobacterales, Parvularcula. However, the hdaB gene is not conserved in all of the alphaproteobacteria. HdaB homologs are absent from Sphingomonadlaes, Rodospirillales and the Rickettsiales orders.

The purified GST tagged HdaB protein does not interact with Cori / DnaA boxes in vitro.

Based upon sequence homology, I predicted that HdaB would interact with the same binding sites as DnaA. I used EMSA with double stranded oligonucleotides which contained DnaA binding sites as well as binding sites for CtrA. The response regulator CtrA was used as a positive control. CtrA-DNA binding was clearly observed as a reduced rate of migration of the oligonucleotides in an ethidium stained native polyacrylamide gel. The oligonucleotides incubated with HdaB migrated at the same rate as the oligonucleotide without the protein. No non-specific binding for either CtrA or HdaB was observed to the randomised pair of oligonucleotides. There are several possible explanations for this negative result. The purified

HdaB protein was not functional, or had become denatured when the tag was cleaved, the

HdaB protein may require one or more other proteins and bind DNA as a complex. Another possibility is that despite the sequence homology to DnaA, HdaB may recognise an alternate sequence and not bind DnaA binding sites at all. Since HdaB might bind sequences other than the predicted DnaA box. I then used a less sequence specific method to look for protein-DNA interactions. I performed a DNase I footprint assay to look for HdaB binding anywhere in the

Cori. Since CtrA has well characterized binding sites in Cori, it was again chosen as the positive control. I observed CtrA binding to all five established CtrA binding sites. However, no zones of

186 protection from the DNase I enzyme were seen with either the GST tagged HdaB or the HdaB protein with the tag cleaved. While the results of the in vitro experiments did not demonstrate

HdaB DNA interactions, this negative result did not exclude the possibility that HdaB does bind the origin in vivo. The binding of HdaB may require the presence of other proteins or some other set of yet unidentified factors.

To help determine the biological role of the hdaB gene I created an hdaB deletion strain, where the hdaB (cc1058) open reading frame was replaced with the omega streptomycin/spectinomycin antibiotics resistance cassette. The replacement of the hdaB open reading frame was achieved at a reduced rate approximately 10 percent, of the kanamycin sensitive screened colonies contained the omega cassette. This is significantly lower that the theoretical value of 50 percent. The replacement of the hdaB locus was confirmed by colony

PCR. The reduced rate at which I observed for the replacement of the hdaB locus could indicate the gene while not essential is conferring a selective advantage to the cell. The presence of a complementing copy of hdaB integrated at the PxylX locus did not alter the results. This result suggests that the integration of the omega cassette may affect the expression of neighboring genes. The overall cell morphology of the hdaB deletion strain was indistinguishable for that of the wild type (NA100) C. crescentus strain when grown in rich medium (PYE) or minimal medium (M2G). The growth rate of the hdaB deletion strain was identical to that of the wild type (NA1000) strain. Taken together, the negative results in this chapter indicate that the hdaB does not play a conspicuous role in C. crescentus growth and differentiation under normal laboratory conditions.

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References

Blaesing, F., C. Weigel, M. Welzeck and W. Messer (2000). "Analysis of the DNA-binding domain of Escherichia coli DnaA protein." Mol Microbiol 36(3): 557-569. Collier, J. and L. Shapiro (2009). "Feedback control of DnaA-mediated replication initiation by replisome- associated HdaA protein in Caulobacter." J Bacteriol 191(18): 5706-5716. Evinger, M. and N. Agabian (1977). "Envelope-associated nucleoid from Caulobacter crescentus stalked and swarmer cells." J Bacteriol 132(1): 294-301. Fujikawa, N., H. Kurumizaka, O. Nureki, T. Terada, M. Shirouzu, T. Katayama and S. Yokoyama (2003). "Structural basis of replication origin recognition by the DnaA protein." Nucleic Acids Res 31(8): 2077- 2086. Fuller, R. S., B. E. Funnell and A. Kornberg (1984). "The dnaA protein complex with the E. coli chromosomal replication origin (oriC) and other DNA sites." Cell 38(3): 889-900. Gorbatyuk, B. and G. T. Marczynski (2001). "Physiological consequences of blocked Caulobacter crescentus dnaA expression, an essential DNA replication gene." Molecular Microbiology 40(2): 485-497. Kawakami, H., K. Keyamura and T. Katayama (2005). "Formation of an ATP-DnaA-specific initiation complex requires DnaA Arginine 285, a conserved motif in the AAA+ protein family." J Biol Chem 280(29): 27420-27430. Kowalski, D. and M. J. Eddy (1989). "The DNA unwinding element: a novel, cis-acting component that facilitates opening of the Escherichia coli replication origin." Embo J 8(13): 4335-4344. Kurokawa, K., S. Nishida, A. Emoto, K. Sekimizu and T. Katayama (1999). "Replication cycle-coordinated change of the adenine nucleotide-bound forms of DnaA protein in Escherichia coli." EMBO J 18(23): 6642-6652. Marczynski, G. T. (1999). "Chromosome methylation and measurement of faithful, once and only once per cell cycle chromosome replication in Caulobacter crescentus." J Bacteriol 181(7): 1984-1993. Marczynski, G. T. and L. Shapiro (1992). "Cell-cycle control of a cloned chromosomal origin of replication from Caulobacter crescentus." J Mol Biol 226(4): 959-977. McGarry, K. C., V. T. Ryan, J. E. Grimwade and A. C. Leonard (2004). "Two discriminatory binding sites in the Escherichia coli replication origin are required for DNA strand opening by initiator DnaA-ATP." PNAS 101(9): 2811-2816. Messer, W. and C. Weigel (1997). "DnaA initiator--also a transcription factor." Mol Microbiol 24(1): 1-6. Shaheen, S. M., M. C. Ouimet and G. T. Marczynski (2009). "Comparative analysis of Caulobacter chromosome replication origins." Microbiology 155(Pt 4): 1215-1225. Spencer, W., R. Siam, M. C. Ouimet, D. P. Bastedo and G. T. Marczynski (2009). "CtrA, a global response regulator, uses a distinct second category of weak DNA binding sites for cell cycle transcription control in Caulobacter crescentus." J Bacteriol 191(17): 5458-5470. Sutton, M. D. and J. M. Kaguni (1997). "The Escherichia coli dnaA gene: four functional domains." J Mol Biol 274(4): 546-561. Sutton, M. D. and J. M. Kaguni (1997). "Threonine 435 of Escherichia coli DnaA protein confers sequence-specific DNA binding activity." J Biol Chem 272(37): 23017-23024. Taylor, J. A., M. C. Ouimet, R. Wargachuk and G. T. Marczynski (2011). "The Caulobacter crescentus chromosome replication origin evolved two classes of weak DnaA binding sites." Mol Microbiol 82(2): 312-326.

188

Chapter Four

The role of key arginine motifs in C. crescentus DnaA regulation

189

Abstract The activity of DnaA in E. coli is regulated by a process called the regulatory inactivation of

DnaA (RIDA) which requires the presence of a protein with Homology to DnaA (Hda). Hda stimulates the intrinsic ATPase of DnaA converting it from active ATP-DnaA to an inactive ADP-

DnaA. Presumably, the inactivation of C. crescentus DnaA also occurs through RIDA or a RIDA- like mechanism mediated by homologue of Hda, called HdaA. However, in contrast to the E. coli protein, C. crescentus DnaA is unstable during both exponential growth and stationary phases. This suggests that C. crescentus employs both nucleoside binding/hydrolysis, as seen in

E. coli, and a novel proteolytic mechanism to regulate DnaA. However, no previous studies have directly addressed the nucleotide binding and ATP hydrolysis of C. crescentus DnaA.

I created four independent single amino acid mutations in two conserved positions of C. crescentus DnaA, DnaAR300 and DnaAR357. The position of C. crescentus DnaAR300 corresponds to the E. coli DnaAR281 and/or DnaAR285 which are primarily required for E. coli

DnaA protein multimerization. The position of C. crescentus DnaAR357 corresponds to the E. coli DnaAR334 which is directly required for ATP hydrolysis. I created two versions of each mutant: severe arginine to alanine mutations (DnaAR300A, and DnaAR357A) and conservative arginine to histidine and lysine mutations (DnaAR300K, and DnaAR357H).

I purified histidine tagged mutant C. crescentus DnaAR300K, and DnaAR357H proteins and demonstrated that they were deficient in ATPase activity. When expressed in C. crescentus from a low copy plasmid, the DnaAR357A and DnaAR357H mutants increased the amount of

DNA per cell, presumably due to increased activity of the DnaA and an over-initiation of chromosome replication. As described in chapter 2, I showed that disruption of RIDA induced

ATPase activity caused by depleting the cell of HdaA leads to both an increase in the frequency

190 of chromosome replication and a stabilization of DnaA. Here, I assessed the stability of DnaA protein in wild type (NA1000) cells expressing the DnaA wt, DnaAR300A, DnaAR357H, or

DnaAR357A from a xylose inducible promoter on low copy plasmid. Both DnaAR357A and

DnaAR357H displayed increased stability. The DnaAR300A mutant did not show an increase in

DnaA protein stability when expressed in C. crescentus. This is likely due to the presence of

RIDA induced ATPase, as mutations made in the equivalent amino acid of E. coli DnaA have been shown to retain their RIDA induced ATPase activity. Taken together, these results agree with the hypothesis that the active form of DnaA is ATP-DnaA and that it is more stable than the inactivated ADP-DnaA form. This is a novel modification of the E. coli RIDA mechanism which ties DnaA proteolysis with C. crescentus cell cycle progression.

191

Introduction DnaA is a member of an ATPase family termed ATPase associated with various cellular activities

(AAA+) proteins (Neuwald, Aravind et al. 1999). The E. coli DnaA has been biochemically shown to have high affinity for both ATP and ADP, as well as a weak intrinsic ATPase activity (Sekimizu,

Bramhill et al. 1987). The binding of DnaA-ATP to the origin of replication is required for initiation of chromosome replication; however, the ATPase activity of DnaA appears to be dispensable for initiation, as the ATP may be replaced with a non-hydrolysable analog of ATP,

ATPγS (Sekimizu, Bramhill et al. 1987). ATP binding to DnaA induces a conformational change in the protein that stimulates oligomerization of DnaA and open complex formation at the origin of replication (Erzberger, Mott et al. 2006). The intrinsic ATPase activity of DnaA is stimulated by the interaction of DnaA with Hda and the sliding clamp loaded on to the DNA (Kato and

Katayama 2001).

The regulation of E. coli DnaA by a homolog of Hda, and the similar regulation of C. crescentus DnaA by HdaA (Chapter 2) suggest that C. crescentus DnaA activity is similarly governed by the ATP/ADP nucleotide state. However, it has not been shown that C. crescentus

DnaA is more active when bound to ATP or even that DnaA binds ATP/ADP. In this chapter, I address this essential issue. I clearly show that C. crescentus DnaA binds ATP/ADP in vitro and in vivo. I also show that mutations that block intrinsic DnaA ATPase also increase DnaA protein stability and stimulate chromosome replication. My results support the original hypothesis from

Chapter 2 that the active form is ATP-DnaA. These results also suggest at least one mechanism for connecting DnaA proteolysis with C. crescentus cell cycle progression. Apparently, the inactive form ADP-DnaA is marked, presumably by a conformational change, for proteolysis.

192

Results Caulobacter DnaA has an unusual Arg-finger and a conserved ATPase motif

Arg fingers are the AAA+ motifs that reach from one protein subunit into the next protein subunit and thereby complete the ATP binding and hydrolysis sites of these interacting proteins. The role of the DnaA Arginine fingers has been studied in E. coli. Two key arginines have been identified, E. coli DnaAR281 which has been implicated in the stabilization of DnaA multimers and DnaAR285 believed to play a direct role in ATP recognition, hydrolysis and the conformational changes of DnaA. Interestingly, C. crescentus (see Figure 1) and related

Caulobacters (data not shown) lack the arginines that directly correspond to E. coli R281 or

R285. The closest corresponding arginine is the C. crescentus R300 and I will assume therefore that this is the arginine finger of C. crescentus DnaA.

Another key arginine required for ATP binding and hydrolysis is the E. coli DnaA R334 in the so called sensor 2 motif. This is a very well conserved motif and the C. crescentus DnaA

R357 corresponds directly to E. coli DnaA R334 (Figure 1). Experiments presented below will show that mutations at this position produce the strongest consequences for C. crescentus

DnaA.

193

Figure 1: Alignment of amino acid sequences of DnaA of E. coli and C. crescentus The amino acid sequence of E. coli DnaA displays a high degree of conservation with the amino acid sequence of C. crescentus DnaA. The degree of conservation between amino acids of each protein is color coded from blue being not conserved to red being completely conserved. The positions of arginines C. crescentus (Ccr) DnaAR300, CcrDnaAR357 studied in this chapter are indicated by circles.

194

Figure 1: Alignment of amino acid sequences of DnaA of E. coli and C. crescentus

195

To determine the role of key arginines in the regulation of DnaA in C. crescentus, I have set out to create point mutations in DnaA, replacing the arginine present with an alanine. I have also created conservative mutations where the arginine present was replaced with either a lysine or histidine. All of the DnaA mutants were created in the pET19b histidine vector, so that the mutant DnaA could be his-tagged for ease of purification.

Intrinsic ATPase activity of DnaA in C. crescentus

The E. coli DnaA has been biochemically shown to have high affinity for both ATP and ADP and to have a weak intrinsic ATPase activity (Sekimizu, Bramhill et al. 1987). The E. coli origin of replication contains two types of DnaA binding sites, R-boxes which bind equally to both ATP-

DnaA and ADP-DnaA (Schaper and Messer 1995) and I sites which preferentially bind ATP-DnaA

(McGarry, Ryan et al. 2004). The C. crescentus origin of replication does not contain DnaA binding sites with preference for either the ATP or ADP associated form of DnaA (Taylor,

Ouimet et al. 2011). The significance of this unexpected observation is not yet clear but it does suggest that the C. crescentus origin of replication may not respond to ATP-DnaA as does the E. coli replication origin. This suggestion contradicts the main conclusion of Chapter 2 that C. crescentus has a RIDA mechanism similar to that of E. coli. However, up to this time there was no direct evidence that C. crescentus DnaA in fact binds ATP or that it has an ATPase activity.

Therefore, I tested C. crescentus DnaA for ATPase activity in vitro. C. crescentus DnaA with a polyhistidine tag was expressed and purified from E. coli cells (Figure 2). The protein was incubated at room temperature with ATP labelled at the alpha position with 32P. Samples were taken at various time points and filtered to remove excess ATP away from the solution. The

DnaA protein was bound to the nitrocellulose filters. The nucleotides associated with DnaA

196 were then separated by thin layer chromatography (TLC). The TLC plates were exposed to a phosphorimager screen to measure the relative amounts of 32P ATP and 32P ADP. To control for spontaneous degradation of the ATP, a no protein control was performed. The purified DnaA was able to convert the majority ATP to ADP in less than 30 minutes and there was no ADP produced in the no protein control (Figure 3A). This result indicates that C. crescentus may also use the hydrolysis of ATP to regulate DnaA activity in a manner similar that of E. coli.

197

Figure 2: Expression and purification of C. crescentus DnaA proteins Proteins suitable for in vitro analysis were obtained by expressing DnaA (wild type), DnaAR300K and DnaAR357H from the pET19b vector containing an N-terminal poly-histidine tag. The histidine tagged proteins were expressed in BL21 (DE3) E. coli cells. The size and purity of the proteins was confirmed by polyacrylamide gel electrophoresis and staining with Pageblue protein stain (Fermentas). E. coli DnaA (Ec DnaA) was used as a reference; it migrates faster as it is smaller than DnaA from C. crescentus (Ccr DnaA).

198

Arginine mutations decrease C. crescentus DnaA ATPase activity in vitro

In order to confirm the inferred role of the arginine fingers which have been previously characterized in E. coli, mutant C. crescentus DnaA proteins were purified and their ATPase activity was measured and compared with a wild type DnaA protein which had been prepared in a similar manner. C. crescentus DnaAwt, DnaR300K and DnaAR357H were purified using a histidine tag (Figure 2). The DnaA proteins were incubated with alpha 32P ATP, the excess ATP removed and the hydrolysis reaction was performed at room temperature with samples taken at various time points. The nucleotides associated with DnaA were then separated by thin layer chromatography (TLC) and the TLC plate was exposed to a phosphorimager screen. The relative amounts of ATP and ADP were determined. Both DnaAR300K and DnaAR357H showed decreased rates of ATP hydrolysis compared with the wild type DnaA protein (Figures 3C,D).

Both previous mutations, DnaAR300K and DnaAR357H were conservative changes that replaced a basic arginine with another basic amino acid. Consequently, I observe that both of these mutations retain at least some intrinsic ATPase activity (Figures 3CD). To obtain DnaA with even lower ATPase activity, I created C. crescentus DnaA proteins with the corresponding changes to alanine, DnaAR300A and DnaAR357A.

199

Figure 3: Intrinsic ATPase activity of C. crescentus DnaA (A) Hydrolysis of ATP bound to wild type C. crescentus DnaA protein was monitored on thin layer PEI-cellulose plates. A no protein control was used to assess the spontaneous hydrolysis of

ATP. Poly-histidine tagged DnaA protein was incubated with alpha 32P ATP on ice for 15 minutes to allow for ATP binding to the protein. The DnaA protein samples were then incubated at room temperature for the times indicated. Samples were filtered through nitrocellulose membranes.

The protein attached to the membrane was extracted with 1M formic acid and spotted onto a thin layer PEI Cellulose plates. The labeled ADP and ATP were separated by thin layer chromatography and quantified using a phosphorimager. The ADP and ATP positions were confirmed by UV fluorescence of unlabeled samples (not shown). The autoradiograms were scanned by densitometry and the percentage of ADP was calculated as %ADP =

(ADP/(ATP+ADP))X100.

(B) Time course of hydrolysis of ATP bound to wild type C. crescentus DnaA protein. Hydrolysis of ATP bound to wild type DnaA protein was monitored on thin layer PEI-cellulose plates, as described in A, except that shorter time points were taken before the exhaustion of ATP. The autoradiograms were scanned by densitometry and the percentage ADP was determined.

(C) Time course of hydrolysis of ATP bound to the DnaAR300K protein. Hydrolysis of ATP bound to DnaAR300K or wild-type DnaA protein was monitored on thin layer PEI-cellulose plates, as described above.

(D) Time course of hydrolysis of ATP bound to the DnaAR357H protein. Hydrolysis of ATP bound to DnaAR357H or wild-type DnaA proteins was monitored on thin layer PEI-cellulose plates, as described in above.

200

Figure 3: Intrinsic ATPase activity of C. crescentus DnaA (A) ATPase activity of wild type C. crescentus DnaA vs no protein control

201

Figure 3: Intrinsic ATPase activity of C. crescentus DnaA (B) Intrinsic ATPase activity of wild type C. crescentus DnaA

202

Figure 3: Intrinsic ATPase activity of C. crescentus DnaA

(D) Intrinsic ATPase activity of wild type C. crescentus DnaA and DnaAR357H

203

C. crescentus DnaA binds both ATP and ADP in vivo The E. coli RIDA mechanism requires that DnaA binds ATP, hydrolyzes ATP and then holds ADP.

Therefore, to support the hypothesis that C. crescentus also uses a similar RIDA mechanism, I immune-precipitated DnaA protein from rapidly lysed C. crescentus cells and demonstrated that both ATP and ADP are bound as predicted by the RIDA hypothesis (Figure 5). The validity of this apparently simple experiment requires an antiserum that is specific for the native form of C. crescentus DnaA. This reagent was not available when I started my studies and so I produced a new antiserum for this purpose (and in fact for many of the experiments in this thesis). The previously available serum contained antibodies to DnaA as well as other proteins including the

GroEL heat shock protein which also binds ATP/ADP. The experiments described in Figure 4 demonstrate that my new serum reacts only with C. crescentus DnaA in both immune- precipitation (native protein conditions, Figure 4A) and in immunoblot (denatured protein conditions, Figure 4BC).

Therefore, I used this new serum to demonstrate that both ATP and ADP are bound to C. crescentus DnaA in vivo. Logarithmically growing cells were incubated with alpha-32P ATP for 15 minutes and then rapidly lysed under native conditions. The released proteins were then treated with the antiserum; the C. crescentus DnaA was immune-precipitated, the nucleotides were analyzed by TLC and visualized by autoradiography. The results in Figure 5 clearly show that both ATP and ADP associate with DnaA. The approximately equal amounts of ATP and ADP imply that a mixed population of swarmer, stalked and dividing cells have on average approximately equal amounts of ATP-DnaA and ADP-DnaA.

204

Figure 4: The specificity of the new and the previously published DnaA antisera (A) The proteins of wild type C. crescentus (NA1000) were labelled by growing the cells in the presence of 35-S labelled methionine. The cells were lysed and an immunoprecipitation was performed using the previously published antiserum against DnaA from the Shapiro lab (lane 1), the pre-immune serum (lane 2), or the newly developed antiserum raised against C. crescentus

DnaA (lane 3). The previously published antibody cross-reacts with a heat shock protein GroEL.

The per-immune serum showed no detectable cross reactivity with the C. crescentus proteins.

The new anti-DnaA antiserum specifically immune-precipitated DnaA.

(B) “Western” immunoblots comparing the specificity of the new DnaA antiserum with the currently widely available anti-DnaA antibody. Western blots were performed using wild type C. crescentus (NA1000) and a strain (GM2472) of C. crescentus with the sole copy of dnaA under the control of a xylose inducible promoter, grown in media without xylose (M2G) and media with xylose (M2GX). The western blot performed with the new anti-DnaA antibody produced a single band from the wild type and GM2472/M2GX cells (lanes 1, and, 3), and no signal from the GM2427 /M2G not expressing dnaA cells (lane 2)

(C) “Western” immunoblots using the previously available anti-DnaA antiserum detect a major band corresponding to GroEL (Gorbatyuk and Marczynski 2005). Otherwise these lanes 4-6 correspond to lanes 1-3 in B. The DnaA band, identified by strains GM2472, is barely detected and GroEL dominates.

205

Figure 4: The specificity of the new and the previously published DnaA antisera

206

Figure 5: C. crescentus DnaA binds both ATP and ADP in vivo Wild type C. crescentus cells were incubated with alpha-32P ATP for 15 minutes with shaking.

The cells were first washed to remove excess ATP. Then these cells were lysed and the DnaA protein was immunoprecipitated with antiserum to C. crescentus DnaA (DnaA antiserum 4300).

The nucleotide bound to the DnaA protein was determined by thin layer chromatography on

PEI-cellulose plates. The plus and minus TCL runs indicate the presence of the anti-DnaA antibody and the identical mock control experiment performed using the pre-bleed serum.

207

C. crescentus DnaA mutants with decreased ATPase activity show increased protein stability

In Chapter 2, I showed that lack of HdaA, a protein predicted to stimulate DnaA ATPase, increases DnaA protein stability. To determine if the intrinsic DnaA ATPase activity influences

DnaA protein stability, I examined the abundance of DnaA in cells expressing either DnaA(wt),

DnaAR300A, DnaAR357H, or DnaAR357A under the control of a xylose inducible promoter on a low copy plasmid. Such cells express predominantly the wild type DnaA from the chromosome in PYEG (glucose) media and they also express DnaA from the plasmid in PYEX (xylose) media.

The abundance of DnaA in wild type C. crescentus (NA1000) cells expressing mutant

DnaAR357A was most greatly increased compared with cells expressing wild type DnaA (Figure

6). These differences were observed despite the fact that all proteins were expressed from the same plasmid and promoter. This result implies that differences in the abundances of these proteins are due to different rates of proteolysis, because their rates of synthesis should be the same.

208

Figure 6: Abundance of DnaA in cells expressing mutant DnaA Wild type C. crescentus (NA1000) cells were transformed with a plasmid that can express either

DnaA (wt), DnaAR300A, DnaAR357H, or DnaAR357A from xylose inducible promoter. The strains were grown for 6 hours in either PYEG, to repress the xylose inducible promoter or PYEX to activate the xylose inducible promoter. A volume of cells equivalent to 1ml at an optical density (660nm) of 0.1 was sampled for western blot analysis. The density of the band corresponding to DnaA was quantified and normalized to the strain expressing the wild type

DnaA. The strains expressing DnaAR300A, DnaAR357H, and DnaAR357A all have increased levels of DnaA despite being expressed from the same promoter.

209

Figure 6: Abundance of DnaA in cells expressing mutant DnaA

210

The effect of altered ATPase activity on DnaA stability was confirmed by translational shut off experiments. Wild type C. crescentus (NA1000) containing a low copy plasmid with

DnaA (wt), DnaAR300K, DnaAR357H, or DnaAR357A under the control of a xylose inducible promoter were grown in rich media (PYE) with antibiotic selection for the plasmid

(chloramphenicol) and with or without xylose for 6 hours to allow plasmid-borne DnaA to be expressed. The cultures were then treated with tetracycline to block protein translation. The abundance of DnaA in the cell was monitored by immunoblot analysis, using an antibody against DnaA. The results in Figure 7 show that both the DnaAR357H and DnaAR357A mutant proteins had increased DnaA protein stability as compared with the wild type DnaA and the

DnaAR300K proteins.

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Figure 7: The stability of DnaA in cells expressing the DnaA ATPase deficient mutants Wild type C. crescentus (NA1000) expressing either DnaA(wt), DnaAR300A, DnaAR357H, or

DnaAR357A from xylose inducible promoter. The strains were grown for 6 hours in either

PYEG, to repress the xylose inducible promoter or PYEX to activate the xylose inducible promoter. Tetracycline was added to halt translation, thus production of new DnaA. To determine the stability of the DnaA at the time when tetracycline was added aliquots of equivalent optical density were taken at 30 minute time points. Immunoblot analysis was used to determine DnaA abundance. DnaA was stabilized in the cells expressing the ATPase deficient mutant DnaA proteins DnaAR357A, and DnaAR557H. The stability of DnaA was not altered in cells expressing either wild type DnaA or the DnaAR300A mutant.

212

Figure 7: The stability of DnaA in cells expressing the DnaA ATPase deficient mutants

213

C. crescentus DnaA mutants with decreased ATPase activity over-replicate their chromosomes

I have previously shown that the RIDA related protein HdaA is required to limit chromosome replication to once per cell cycle (Chapter 2, Figures 6-7), presumably through stimulation of the ATPase activity of DnaA. To assess the role of intrinsic ATP hydrolysis, I expressed the

DnaAR357A and DnaAR357H mutants in wild type (NA1000) C. crescentus. The DnaAR357A and

DnaAR357H mutants were placed under the control of a xylose inducible promoter on a low copy plasmid. When grown in the presence of the inducer (0.2% xylose), cells expressing either

DnaAR357A or DnaAR357H became very elongated (Figure 8). This appearance was reminiscent of the elongated HdaA depleted cells that I observed (Chapter 2, Figure 5C). Flow cytometry showed that DnaAR357A and to a lesser extent DnaAR357H over-initiated chromosome replication (Figure 9). This over-replication is shown by the extra trail of DNA fluorescence, seen after the 2-chromsome peaks, in the +xylose DnaAR357A and +xylose DnaAR357H cells.

This trail of DNA fluorescence is not seen in the corresponding +glucose cells or in and of the cell with wild type of the DnaAR300A expressing plasmids (Figure 9). Therefore, the expression of an ATPase defective DnaA protein phenocopies the GM3700 strain which had been depleted of HdaA (Chapter 2).

Wild type C. crescentus moderately over expressing DnaAR300A did not show a dramatic increase in chromosome content when compared with cells moderately over- expressing wild type DnaA (Figure 9). Despite the fact that a point mutation (DnaAR300K) caused a decrease in ATPase activity, the stability of DnaA in the cells was not significantly increased in cells moderately over expressing DnaAR300A (Figure 7), as was seen with cells moderately over expressing the DnaAR357A, or DnaAR357H mutant proteins. In E. coli, the

214 corresponding arginines (E. coli R281 or R285 correspond to C. crescentus R300, Figure 1) are not directly involved in ATPase activity but in DnaA multimerization and the recognition of ATP.

It is possible that the DnaAR300A mutants are less able to participate in multimerization and have less of an effect on the initiation of chromosome replication than mutants which are inert with respect to ATPase activity in vivo or because they retain RIDA induces ATPase activity.

215

Figure 8: Moderate overexpression of DnaAR357A and DnaAR357H results in a cell division defect Representative images from wild type C. crescentus (NA1000) containing the pRXMCS6-

DnaAR300A plasmid, the pRXMCS6-DnaAR357A plasmid, and the pRXMCS6-DnaAR357H plasmid. The cultures were grown to exponential phase in PYE, then 0.2% glucose (PYEG) was added to half the culture and 0.2% xylose (PYEX) was added to the other half. The cultures were grown for an additional 20 hours. The expression of DnaAR357A, and DnaAR357H resulted in cells with blocked cell division, while the expression of DnaAR300A only had a mild effect on cell morphology.

216

Figure 9: The moderate overexpression of DnaAR357A or DnaAR357H results in an increase in chromosome replication Flow cytometry performed on wild type (NA1000) cells with either DnaA wt, DnaAR300A,

DnaAR357A, DnaAR357H under the control of a xylose inducible promoter on a low copy plasmid (pRXMCS-6). The cells were grown 6 hours in PYE-chloramphenicol with glucose (0.2%), or xylose (0.2% each). The cells were treated with rifampicin and cephalexin overnight. The chromosomes were stained with propidium iodide. The quantity of DNA per cell was measured using a BD FACSCalibur (BD Biosciences) and the Cell Quest Pro acquisition software (BD

Biosciences). Analysis of the data was performed using the FlowJo software (TreeStar Inc.)

The moderate overexpression of DnaAR357A, and DnaAR357H resulted in cells with an increased DNA content per cell indicating over replication of the chromosome, while the moderate overexpression of wild type DnaA or the mutant DnaAR300A protein did not alter the

DNA content of the cell.

217

Figure 9: The moderate overexpression of DnaAR357A or DnaAR357H results in an increase in chromosome replication

218

Discussion Based upon sequence alignments between DnaA of C. crescentus and E. coli, two arginines were selected for mutagenesis. I created independent point mutations in C. crescentus DnaA arginine

300 and arginine 357. The DnaA arginine 357 of C. crescentus is equivalent to the highly conserved arginine 334 residue within the Box VIII sensor 2 motif of the AAA+ domain of E. coli

DnaA. The E. coli DnaAR334A mutant is inert with respect to ATPase activity and therefore does not hydrolyze ATP, despite retaining binding affinities for both ATP and ADP comparable to that of wild type DnaA (Nishida, Fujimitsu et al. 2002). Experiments performed in vitro show that when bound to ATP, DnaAR334A initiates minichromosomal replication at a level similar to that seen for wild-type DnaA. When expressed at moderate levels in vivo, DnaA R334A is predominantly in the ATP-bound form resulting in over initiation of chromosome replication, unlike the wild-type DnaA (Nishida, Fujimitsu et al. 2002). Results presented in Chapter 2 indicate that in C. crescentus, there is a link between the ATPase induced inactivation of DnaA and DnaA proteolysis. The ATPase activity of the DnaAR334A mutant is not stimulated by RIDA.

Therefore, I chose to create a C. crescentus DnaAR357A and a DnaAR357H mutant to explore the role of ATPase activity on DnaA activity and stability.

C. crescentus DnaA contains a single arginine (C. crescentus DnaAR300) in the same relative position as the two arginines of E. coli DnaAR281 and DnaAR285. It is believed that E. coli DnaAR285 acts as an arginine finger which recognizes ATP bound to the adjacent subunit in a multimeric complex (Kawakami, Keyamura et al. 2005). The E. coli DnaAR285A mutant has been shown to be inactive both in vitro and in vivo for chromosome replication due to defects in unwinding of the origin of replication (Kawakami, Keyamura et al. 2005). The DnaAR281 mutant is able to bind ATP and ADP but is inactive for replication from oriC. This is believed to

219 be due to an inability to form the multimeric structures on oriC needed to recruit the DnaB helicase to the origin of replication (Felczak and Kaguni 2004). Mutations in these two arginines have different effects on the ATPase activity of DnaA. The DnaAR285A mutant is inactive for intrinsic ATPase activity, while the DnaAR281A shows a fivefold increase in intrinsic ATPase activity compared with wild type DnaA protein (Kawakami, Keyamura et al. 2005). However, both the DnaAR281A and DnaAR285A mutants hydrolyze ATP in a RIDA dependent manner at rates comparable to that of wild type DnaA. Therefore, I chose to create a C. crescentus

DnaAR300A mutant, and a DnaAR300K mutant to determine the role of multimerization on

DnaA activity and protein stability.

The experiments detailed in this chapter demonstrate a link between the state of the nucleotide bound to DnaA, the activity of DnaA in chromosome replication, and the stability of

DnaA. Apparently, in C. crescentus the activity of the DnaA is regulated by both nucleotide state and by proteolysis. Mutations in DnaAR357 result in a decrease in ATPase activity in vitro.

When expressed at moderate levels in vivo DnaA proteins with mutations in arginine 357

(DnaAR357H or DnaA357A) result in increased chromosome replication. The stability of DnaA is also increased in cells expressing either DnaAR357A or DnaAR357H and these cells also showed cell division defects consistent with over-replication. These observations suggest that DnaA protein stability and activation are functionally connected. These results are also similar to those I obtained from cells which have been depleted of the HdaA protein. HdaA is a negative regulator of DnaA in C. crescentus and is required for the regulatory inactivation of DnaA

(RIDA). One interpretation of these results is that loss of ATPase activity stabilizes and locks the

DnaA in an active form which results in an increased frequency of initiation of chromosome

220 replication. Once DnaA is inactivated by the hydrolysis of the attached ATP it is removed from the cell by proteolysis.

To identify the role of the arginine finger in C. crescentus, we created two mutants of

DnaA, a DnaA arginine to alanine mutant (DnaAR257A) and a less severe arginine to lysine mutant (DnaAR300K). Mutations of the comparable arginine fingers of E. coli gave two different results; the E. coli DnaAR281A displayed an increased intrinsic ATPase activity while the E. coli

DnaAR285A mutation abolished the intrinsic ATPase activity. The DnaAR300K mutation we created had a reduced ATPase activity. When expressed in wild type C. crescentus, the

DnaAR300A mutant showed only moderate morphological defect. We used flow cytometry to monitor chromosome content; we did not observe increased DNA content per cell in cells expressing the mutant DnaAR300A protein. The stability of DnaA was unchanged in cells expressing the DnaAR300A compared with cells expressing wild type DnaA. Mutations in arginine 300 of DnaA in C. crescentus display an effect on the intrinsic ATPase activity in vitro but have only a mild effect on cellular physiology in vivo. This is likely due the ATPase activity of

DnaA which is induced by RIDA. The E. coli DnaAR281A and DnaAR285A mutants have RIDA induced ATPase activities similar to wild type DnaA. If the C. crescentus DnaAR300A mutant is also responsive to RIDA induced inactivation we would not expect to see an effect in vivo.

Despite the biochemical differences between DnaA of E. coli and C. crescentus the core mechanism of DnaA being activated when associated with ATP and inactivated by the hydrolysis of ATP to ADP is conserved. This conserved mechanism has been modified by C. crescentus to allow it to strictly control chromosome replication in nutrient poor environments.

The proteolytic regulation of DnaA in C. crescentus which I have shown is tied to RIDA (Chapter

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2) appears to be regulated by the nucleotide bound state of DnaA, providing a direct link between the proteolysis of DnaA and cell cycle progression.

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Experimental Procedures Bacterial strains and growth conditions

The strains and plasmids used in this study are listed in Table 1. C. crescentus strains were grown in peptone-yeast extract (PYE) rich media or M2G minimal glucose media.

Strain/Plasmids Description Source

E. coli Strains DH5α Cloning strain Invitrogen BL21(DE3) Protein expression strain

C. crescentus NA1000 wild type C. crescentus Evinger and Agabian (1977) synchronizable strain Plasmids pGM2876 pET19B::dnaA C. crescentus his- Shaheen et al.(2009) dnaA expression vector pGM3769 pRXMCS6::danA xylose dependent This work expression of C. crescentus DnaA pGM3770 pRXMCS6::danAR300A xylose This work dependent expression of C. crescentus DnaAR300A pGM3771 pRXMCS6::danAR300K xylose This work dependent expression of C. crescentus DnaAR300K pGM3769 pRXMCS6::danAR357Axylose This work dependent expression of C. crescentus DnaAR357A pGM3769 pRXMCS6::danAR367H xylose This work dependent expression of C. crescentus DnaAR357H

Purification of His tagged DnaA

Histidine – tagged wild type, DnaAR300K, DnaAR357H were expressed pET19bDnaA, pET19bDnaAR300K, and pET19bDnaAR357H plasmids respectively. The histidine tagged

223 proteins were expressed in BL21 E. coli cells and the purification was performed as described in

(Li and Crooke 1999).

DnaA intrinsic ATPase activity

Histidine tagged DnaA protein was incubated with alpha 32P ATP on ice for 15 minutes to allow for ATP binding to the protein. The samples were then incubated at room temperature for various times. Samples were filtered through nitrocellulose membranes. The protein attached to the membrane was extracted with 1M formic acid and spotted onto a thin layer PEI Cellulose plates. The labeled ADP and ATP were separated by thin layer chromatography and quantified using a phosphorimager. The rate of production of percentage of ADP was calculated as %ADP =

((ADP/ATP+ADP)X100).

Flow cytometry

The chromosomal content of the cells was determined using flow cytometry. Cells were treated with 60 μg ml−1 rifampicin to block RNA synthesis, preventing new initiations of chromosomal replication and 35 μg ml−1 cephalexin to prevent cell division, for three hours to let any chromosome that had begun replication to proceed to completion. The number of chromosome peaks observed would then be equivalent to the number of origins of replication present when the antibiotics were added. Cells were then washed in staining buffer (50 mM sodium citrate, 10 mM Tris-HCl, 1 mM EDTA, pH = 7.2), and permeabilized by adding ethanol to a final concentration of 70%. The RNA was removed from the cells by the addition of RnaseA and the DNA stained with propidium iodide (20 μg ml−1). The data was acquired using the FL2 detector of a BD FACSCalibur (BD Biosciences) and the Cell Quest Pro acquisition software (BD

Biosciences). Analysis of the data was performed using the FlowJo software (TreeStar Inc.)

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Immunoblot analysis

Immunoblot analysis was used to determine the cellular levels of DnaA. Whole cell lysates were resolved using 8% SDS-PAGE. Immunoblots were performed using DnaA anti-serum diluted 1:10,000 anti-rabbit immunoglobulin G conjugated to horseradish peroxidase 1:10,000 dilution and visualised with Chemiluminescent ECL (Perkin Elmer).

Antibody production

Polyhistidine tagged C. crescentus DnaA protein was expressed in E. coli BL21(DE3) from the pGM2876 plasmid. The tagged protein was purified as previously described (Li and Crooke

1999) and used to immunize two rabbits for the production of polyclonal antibodies (McGill

Animal Resources Centre). Antibody specificity was determined by western blot analysis and by

35-S immunoprecipitation, using a DnaA conditional expression strain (GM2471).

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References Erzberger, J. P., M. L. Mott and J. M. Berger (2006). "Structural basis for ATP-dependent DnaA assembly and replication-origin remodeling." Nat Struct Mol Biol 13(8): 676-683. Evinger, M. and N. Agabian (1977). "Envelope-associated nucleoid from Caulobacter crescentus stalked and swarmer cells." J Bacteriol 132(1): 294-301. Felczak, M. M. and J. M. Kaguni (2004). "The box VII motif of Escherichia coli DnaA protein is required for DnaA oligomerization at the E. coli replication origin." J Biol Chem 279(49): 51156-51162. Kato, J. and T. Katayama (2001). "Hda, a novel DnaA-related protein, regulates the replication cycle in Escherichia coli." EMBO J 20(15): 4253-4262. Kawakami, H., K. Keyamura and T. Katayama (2005). "Formation of an ATP-DnaA-specific initiation complex requires DnaA Arginine 285, a conserved motif in the AAA+ protein family." J Biol Chem 280(29): 27420-27430. Li, Z. and E. Crooke (1999). "Functional analysis of affinity-purified polyhistidine-tagged DnaA protein." Protein Expr Purif 17(1): 41-48. McGarry, K. C., V. T. Ryan, J. E. Grimwade and A. C. Leonard (2004). "Two discriminatory binding sites in the Escherichia coli replication origin are required for DNA strand opening by initiator DnaA-ATP." Proc Natl Acad Sci U S A 101(9): 2811-2816. Neuwald, A. F., L. Aravind, J. L. Spouge and E. V. Koonin (1999). "AAA+: A class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes." Genome Res 9(1): 27- 43. Nishida, S., K. Fujimitsu, K. Sekimizu, T. Ohmura, T. Ueda and T. Katayama (2002). "A nucleotide switch in the Escherichia coli DnaA protein initiates chromosomal replication: evidnece from a mutant DnaA protein defective in regulatory ATP hydrolysis in vitro and in vivo." J Biol Chem 277(17): 14986-14995. Schaper, S. and W. Messer (1995). "Interaction of the initiator protein DnaA of Escherichia coli with its DNA target." J Biol Chem 270(29): 17622-17626. Sekimizu, K., D. Bramhill and A. Kornberg (1987). "ATP activates dnaA protein in initiating replication of plasmids bearing the origin of the E. coli chromosome." Cell 50(2): 259-265. Shaheen, S. M., M. C. Ouimet and G. T. Marczynski (2009). "Comparative analysis of Caulobacter chromosome replication origins." Microbiology 155(Pt 4): 1215-1225. Taylor, J. A., M. C. Ouimet, R. Wargachuk and G. T. Marczynski (2011). "The Caulobacter crescentus chromosome replication origin evolved two classes of weak DnaA binding sites." Mol Microbiol 82(2): 312-326.

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Chapter Five Conclusion and Future Directions

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The central theme of this thesis is the regulation of the bacterial initiator of chromosome replication, DnaA, in Caulobacter crescentus. Much of our current understanding of the regulation of chromosome replication is based upon studies performed using the model bacteria Escherichia coli. The protein DnaA is essential for the initiation of chromosome replication in nearly all bacteria. To adapt to the wide range of environments in which bacteria are found, individual bacteria have adapted the regulatory mechanism and biochemical properties of DnaA to allow them to better exploit their environment. A model of chromosome replication based solely on E. coli may not satisfactory to account the diversity found among bacteria.

The work presented in this thesis demonstrates that C. crescentus utilizes a homolog of the E. coli Hda protein (HdaA) and presumably the regulatory inactivation of DnaA (RIDA) to limit chromosome replication to once per cell cycle (Chapter 2). Although homologs of Hda have been identified in a wide range of bacteria in the phylum proteobacteria (Chapter 2), the

RIDA has only previously been reported in E. coli. The identification of a RIDA mechanism does not preclude the existence of other complementary mechanisms that govern chromosome replication. Three separate mechanisms have been identified in E. coli which limit chromosome replication to once per cell cycle and disabling any one of these mechanisms results in an increased frequency of initiation of chromosome replication.

In C. crescentus, chromosome replication is strictly limited to once per cell cycle and the initiation of chromosome replication occurs only in the stalked cell type. The disruption of the

RIDA mechanism would presumably lead to an accumulation of DnaA locked in an active form both in the swarmer and stalked cell types. Future studies should aim to determine if the over-

228 replication phenotype occurs only in the stalked cell type, indicating the existence of independent mechanisms preventing chromosome replication in swarmer type cells. It has been previously speculated that binding of the response regulator CtrA to the origin of replication in swarmer cells rendered them incapable of supporting chromosome replication, however recent work by Bastedo and Marczynski (2009) show that all five binding sites for CtrA can be disrupted without an increase in the frequency of chromosome replication. This result may be due to the presence of redundant mechanisms such as RIDA which limit chromosome replication. It would be fascinating to determine if the mutations in the CtrA binding sites were synergistic with the HdaA depletion phenotype. The absence of CtrA at the origin and presence of DnaA in an active ATP bound state could allow for the ectopic initiation of chromosome replication in swarmer cells.

The work presented in this thesis indicates a novel connection between the activity

(nucleotide) states of DnaA with its stability. The depletion of the negative regulator of DnaA

HdaA resulted in the stabilization of the DnaA protein. Single amino acid substitutions which impair the ATPase activity of DnaA also stabilize the DnaA protein. Taken together these results indicate that DnaA is stabilized when found in an active state associated with ATP. The DnaA which had been stabilised by the depletion of HdaA was still subject to starvation induced proteolysis, suggesting that the proteolysis of DnaA is governed by two distinct pathways. This result was confirmed by analysis of a strain of C. crescentus carrying a genetic deletion of spoT

(Lesley and Shapiro 2008). The SpoT signalling protein, required for the starvation induced degradation of DnaA, is not required for the stationary phase degradation of DnaA (Chapter 2). This result was surprising as it was previously believed that DnaA proteolysis during stationary phase

229 was caused by the cell sensing the depleting nutrients in the environment. The mechanism which governs this selective degradation of the inactive ADP associated form of DnaA could be an interesting area for future studies.

Chapter three of this thesis deals with a conserved gene that is predicted to encode a protein with homology to the DNA binding domain of DnaA. Sequence analysis of this gene suggests that it should recognise the same sequences as the DnaA protein. The role of this gene remains unknown as I was able to replace the complete open reading frame with an antibiotics cassette without any obvious physiological consequences to the cell. This result was confirmed by a supersaturated transposon mutagenesis screen that identified all of the essential genes in

C. crescentus, the hdaB gene was identified as non-essential by this screen. The purified protein did not bind to known DnaA binding sites in vitro (chapter 3). While these negative results fail to reveal the role of the hdaB gene in C. crescentus, its conservation among alphaproteobacteria suggests that it has been retained because it delivers a selective advantage. It has still to be determined if the HdaB protein is capable of binding DNA. This could be determined by the creation of a chimeric protein of DnaA with the DNA binding domain

(domain IV) replaced by HdaB. If this chimeric DnaA-HdaB protein is able to replace the native copy of DnaA, this would be strong evidence that the HdaB protein capable of binding DNA in a sequence specific manner. The creation of an antibody against HdaB would allow for immunoprecipitation experiments such as ChIP-SEQ to identify HdaB binding sites on the chromosome. The hdaB gene may be dispensable due to the presence of a second gene which serves a redundant purpose. This could be assayed for by a screen for genes which are synthetically lethal with HdaB.

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In chapter 2 of this thesis, I identify a novel role for the RIDA protein HdaA in DnaA stability. The stabilization of DnaA observed when HdaA was depleted from C. crescentus could be explained by two possible mechanisms. Either the ADP associated form of DnaA may be preferentially targeted for proteolysis or HdaA may itself act as a chaperon protein that brings

DnaA to the protease for degradation. In order to differentiate between these two possibilities

I have created four independent ATPase deficient mutants of DnaA. In chapter 4 I describe the effects of expressing these mutants in C. crescentus. The stabilization of DnaA in C. crescentus

(containing a wild type hdaA gene) expressing the ATPase deficient mutants (DnaAR357A, or

DnaAR357H) suggests that the stabilization of DnaA seen in the HdaA depleted cells is due to the effect of HdaA on the nucleotide state. The regulation studies of DnaA in bacteria other than E. coli are just beginning. It would be interesting to characterize the ATP and ADP bound states of DnaA as a function of the cell cycle as well as how these two forms of DnaA are organized at the sub-cellular level.

In addition to its role as the initiator of chromosome replication, DnaA also functions as a transcription factor. The conditional hdaA expression described in Chapter 2 and the DnaA

ATPase mutants described in Chapter 4 would be a valuable resource for further studies of the role of the nucleotide state in regulating the transcriptional activity DnaA has yet to be fully explored.

Bacteria are found nearly everywhere on our planet. While they have adapted to inhabit a diverse set of ecological niches, they share a common set of challenges since they must all consume nutrients, grow, replicate their genetic material and divide. It is amazing that nearly all bacteria utilize the protein DnaA to initiate chromosome replication. This conservation of

231 function across an entire domain of life indicates that DnaA-based replication initiation is both robust enough to be maintained throughout bacterial evolution, and yet malleable enough to be regulated in a variety of ways to meet the needs of bacteria with very different lifestyles.

The regulation of DnaA has been adapted by individual bacteria to allow them to grow and divide at a rate best suited to their particular environment. A thorough understanding of how

DnaA functions in a wide variety of bacteria would be of great scientific, economical, and medical importance.

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References Bastedo, D. P. and G. T. Marczynski (2009). "CtrA response regulator binding to the Caulobacter chromosome replication origin is required during nutrient and antibiotic stress as well as during cell cycle progression." Mol Microbiol 72(1): 139-154. Lesley, J. A. and L. Shapiro (2008). "SpoT regulates DnaA stability and initiation of DNA replication in carbon-starved Caulobacter crescentus." J Bacteriol 190(20): 6867-6880.

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