Initiation of DNA Replication?
Module
4.4.1
ALAN C. LEONARD* AND JULIA E. GRIMWADE
[SECTION EDITOR: SUE LOVETT]
Posted January 05, 2010 Nikhil
Florida Institute of Technology, Department of Biological Sciences, Melbourne, FL 32901–6975
*Corresponding author. Mailing address: Florida Institute of Technology, Department of Biological Sciences, Olin Life Science Building, Rm 234, 150 West University Blvd., Melbourne, FL 32901–6975. Phone: (321) 674–8577, Fax: (321) 674–7990, E-mail:
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†We dedicate this review to the memory of Walter Messer, not only to honor his considerable scientific contributions but also to recognize the great kindness and support he bestowed on all who joined him in the quest to understand the regulation of bacterial cell growth.
Before dividing into two daughter cells, Escherichia ddddd coli and Salmonella enterica must duplicate their circular genomes, each encompassing about 4.7 Mbp of double-stranded DNA (25, 164). Successful completion of this critical step of cellular reproduction demands precise coordination between the nutritional state of the cell and the mechanism that triggers the start of new DNA polymerization. In recent years it has become clear that complex regulatory circuits control the initiation step of DNA replication by directing the assembly of a multicomponent molecular machine (the orisome) that separates DNA strands and loads replicative helicase at oriC, the unique chromosomal origin of replication. After assembly of the replisome and passage of new replication forks, the orisome must then be disassembled, and its components must be inactivated, to ensure that, during a cell division cycle, only one new round of DNA synthesis is triggered from every replication origin.
Although many details of the process still remain unclear, in this chapter we will discuss recent efforts to understand the regulated protein-DNA interactions that are responsible for properly timed initiation of chromosome replication. Information about newly identified nucleotide sequence features within E. coli oriC and new structural and biochemical attributes of the bacterial initiator protein DnaA will be reviewed. We will also discuss the coordinated mechanisms that prevent improperly timed DNA replication.
A number of recent and noteworthy reviews cover several topics presented here in greater detail (116, 122, 176, 180, 197, 267), and the reader is also referred to two new EcoSal chapters covering DNA replication machinery (chapter 4.4.2) and replication, segregation, and cell division (chapter 5.2.1), as well as a previous chapter on timing of synthetic activities in the cell cycle (101) in the last print edition of Escherichia coli and Salmonella: Cellular and Molecular Biology. It is also important to note that two chapters on initiation of DNA synthesis were written for the last print editions of Escherichia coli and Salmonella: Cellular and Molecular Biology, by Kaspar von Meyenburg and Flemming Hansen (249) and by Walter Messer and Christoph Weigel (169). The reader is referred to both of these excellent chapters for details that we are unable to include in this version.
Bacteria contain more DNA when growing rapidly and less when growing slowly, with the amount of DNA per cell varying continuously with growth rate. This observation, presented in 1958 by Schaechter et al. (214), raised questions about how growth-rate-regulated expansion and contraction of DNA content was achieved. Although it is reasonable to expect that replication forks simply move more rapidly at ever faster growth rates, this is not the case. Studies done by Helmstetter and Cooper (47, 100), using synchronously dividing E. coli B/r cultures, demonstrated that the average rate of DNA chain elongation is the same in cells with doubling times between 20 and 100 min, reviewed in reference 101. Over this range, approximately 40 min is required to duplicate the chromosome (termed the C period), and an additional 20 min (D period) is required for the cell to complete septum formation and divide (see Fig. 1) (47, 100), although the duration of C and D periods may be different in other strains (see, for example, reference 102). Exponentially growing E. coli B/r will always divide 60 min after the onset of each round of chromosomal DNA synthesis. However, the time required to prepare to initiate a round of chromosome replication (I period) is not constant, but depends strictly on cellular growth rate. Based on this relationship of I and C+D, new rounds of DNA synthesis will initiate in the early, middle, or late portion of the cell cycle, independent of the status of the cell's ongoing chromosome replication and septum formation (101) (see Fig. 1).
One caveat of the constancy of C + D is that during rapid growth, at doubling times less than 60 min, there is insufficient time for cells to complete a round of chromosome replication and divide before new rounds of DNA synthesis must be triggered (Fig. 1). Since replication forks progress bidirectionally from fixed replication origins on Escherichia and Salmonella chromosomes (78, 161, 162), newly divided daughter cells will inherit dichotomously branched chromosomes (termed theta structures, because they resemble the Greek letter θ). Chromosome configurations in E. coli B/r cells growing with generation times of 20, 40, 50, and 80 min are shown in Fig. 1. The reader is also referred to http://simon.bio.uva.nl/Object-Image/CellCycle/index.html (268) for an animated simulation of chromosome configurations at a variety of growth rates as well as different C + D periods. It is also worth noting that, because of these overlapping rounds of DNA replication, cells must contain more replication origin copies at faster growth rates (Fig. 1). However, all copies initiate replication synchronously during the cell division cycle (225).
The I + C + D rule provides an explanation for the increased DNA content in rapidly growing bacteria and focuses attention on the mechanism that triggers initiation of DNA synthesis as a key regulatory step in the bacterial cell cycle.
What is the nature of a regulatory mechanism that triggers new rounds of DNA synthesis at the correct time during the cell cycle? A simple regulatory circuit to control the initiation step, referred to as the replicon model, was described in 1963 by Jacob, Brenner, and Cuzin (114) and is considered to be the first formal conceptualization of a defined genetic site from which DNA synthesis begins (originally called the replicator, and now referred to as the replication origin), and replicon-encoded, origin-activating factors, called initiators. The concept that the initiator was a protein in E. coli was supported by early studies demonstrating that new DNA synthesis is sensitive to chloramphenicol because ongoing protein synthesis is required for initiation (142, 159, 213). Additional studies demonstrated that cells always initiate DNA replication at a constant mass per chromosomal origin (57), and led to the conclusion that a threshold level of one or more initiator proteins triggers new rounds of DNA synthesis.
Identification of the genes that encoded the initiators came from studies on temperature-sensitive, conditional-lethal mutants of E. coli, in which two DNA replication-defective phenotypes were identified (132, 133). One class, “immediate stop” mutants, halted DNA synthesis immediately upon shift to nonpermissive temperature, indicating that the mutated gene products were required for the elongation phase of DNA replication. The “delayed stop” mutants, in contrast, continued to make DNA after a shift to nonpermissive temperature, gradually stopping after a reproducible period of time (132, 133, 169). The kinetics of the delayed stop mutants suggested that the defective gene products were required specifically for the initiation step of DNA synthesis, and subsequently, two genes, dnaA and dnaC, were identified (reviewed in references 169 and 249). As described in more detail below, the DnaA protein is the bacterial initiator, and in E. coli, the DnaC protein is required to load replicative helicase.
The first step toward identifying the replicator came from studies in which the position of the unique E. coli replication origin was mapped by marker frequency analysis to a chromosomal site near ilv (22, 23). Hiraga then demonstrated that this region of chromosomal DNA, when carried on F′ plasmids, was sufficient to suppress the incompatibility of F plasmids to replicate in Hfr strains (105). He first called this region poh+ (permissive on Hfr), and then named the genetic locus “oriC” to designate the origin of chromosome replication (105). oriC was more precisely mapped to be in, or near a 1.3-kb HindIII fragment located at 84.3 minutes on the E. coli chromosome (161). A 9-kbp EcoRI fragment from this region of the chromosome was ultimately found to be capable of directing autonomous extrachromosomal replication when circularized into a plasmid (250, 262).
Several modifications to the replicon model were made to account for constancy of initiation mass (202, 230), and these models enforced the importance of having both positive and negative regulatory features to ensure both proper timing and only once-per-cell cycle initiations. As described in detail in the following sections, the present view is that, in E. coli and S. enterica, DnaA protein is the positive regulator whose threshold level triggers new rounds of DNA replication and determines initiation mass (116, 154), with a variety of postinitiation regulators quickly repressing the potential to reinitiate. It is also clear that to ensure that initiations are precisely timed, it is necessary to assemble multicomponent complexes to activate replication origins (61); these complexes have been termed “orisomes” (49, 146, 269). Regulation of DnaA accessibility to oriC, the ordered assembly and disassembly of a multi-DnaA complex at oriC, and the means by which DnaA unwinds oriC remain important questions to be answered and we will discuss the current state of knowledge on these topics in the following sections.
Before assembly of bidirectional replication forks, a replication origin is unwound and two molecules of replicative DNA helicase are loaded onto available single strands (136). In Escherichia and Salmonella, three proteins, DnaA, DnaB, and DnaC, are sufficient to perform these activities. DnaA is responsible for site-specific unwinding of oriC and assists DnaC, a dedicated helicase-loading protein, in positioning DnaB, the replicative DNA helicase, onto available single-stranded DNA (136). All three components are members of the AAA+ family (ATPases associated with various cellular activities) (reviewed in reference 244), whose activity is regulated by the binding and hydrolysis of ATP.
DnaA is the initiator protein.
DnaA is a highly conserved 52-kDa protein that has been identified as the primary initiator protein in eubacteria (154, 176, 272). DnaA levels are the critical factor in determining the cellular mass at initiation (154), and overproduction of wild-type DnaA results in increased initiations from oriC in vivo (11) and abolition of cell cycle-specific initiation timing (199). These extra initiations do not necessarily increase intracellular DNA content due to replication fork collapse near oriC (10, 222, 226).
DnaA binds to DNA at a 9-mer sequence with the consensus 5′-TTA/TTNCACA-3′ (24, 215), although lower-affinity binding sites that deviate from the consensus in one or more base pairs have been identified (87, 95, 124). As described in more detail in the following sections, both high- and low-affinity DnaA recognition sites exist in oriC and become occupied prior to DnaA-catalyzed unwinding (146). Approximately 300 additional consensus recognition sites for DnaA are found distributed throughout the chromosome (207).
DnaA has a high affinity for ATP (KD = 30 nM) and ADP (KD = 100 nM) (218). Since the ATP levels in the cell are higher than ADP, newly synthesized DnaA is commonly believed to be in the ATP form. DnaA-ATP is the active form of the protein, and, in vitro, DnaA-ATP is necessary for both oriC unwinding and initiation of DNA replication (218). The activity conferred by ATP is allosteric, since DnaA bound to a poorly hydrolyzable analog of ATP is also active for unwinding and replication in vitro (218). An intrinsic ATPase activity converts DnaA-ATP to inactive DnaA-ADP, but this activity is too slow to be effective in cells, and so must be stimulated in a postinitiation, DNA replication-coupled mechanism called RIDA (Regulatory Inactivation of DnaA) (117) (see “Hda: post-initiation inactivator of DnaA,” below). Hydrolysis of DnaA-ATP is critical, since overinitiation, fork collapse, and cell death are observed in cells carrying mutations, for example, dnaAcos, that cannot downregulate DnaA by hydrolysis (36, 123, 127, 182, 222).
Structural properties of DnaA.
DnaA is divided into four functional domains (238) (Fig. 2). Domain I (N-terminal amino acids 1 to 90) interacts at separate locations with DnaB (217, 237) and other regulatory proteins, such as DiaA (113), Hda (236), HU (44), and Dps (45) (see below). Domain I also plays a role in DnaA self-oligomerization (68, 256). Several laboratories have performed deletion and site-directed mutagenesis of DnaA domain I, and these mutants reveal critical regions needed for protein-protein interactions (1, 68, 223, 256).
Domain II (amino acids 90 to 130) is a flexible linker region without a clearly defined role in initiation of DNA replication. Domain II is the least conserved domain among eubacterial DnaAs, but the length of the linker appears to be important for function (184), since only some deletions in this region are tolerated, and one viable deletion mutant is reported to have an underinitiating phenotype (172). Domain II appears to be the only region in which large insertions can be made in DnaA, a feature that has been used to construct a functional green fluorescent protein-tagged DnaA (27).
Domain III (amino acids 130 to 347) is the region that establishes DnaA as a member of the AAA+ family of ATPases (60, 118). Domain III can be further divided into smaller subdomains that are common among AAA+ proteins (Fig. 2). Most of the N-terminal and central region of domain III (termed the Base), is an αβα-nucleotide binding fold that carries highly conserved Walker A and B motifs involved in adenine nucleotide binding and magnesium ion interaction (reviewed in reference 116, 118, and 167). This region also contains sensor I (86) and box VII motifs, which site-directed mutational analyses have shown to be important for γ-phosphate hydrolysis and DnaA oligmerization (67, 124). Box VII contains a conserved arginine (Arg 285) that is required for DnaA function (67, 124). The remaining C-terminal region of domain III is an α-helical bundle (termed the Lid) that carries the conserved AAA+ motif sensor II, also involved in monitoring nucleotide binding (118). An invariant arginine critical for DnaA function is also found at the tip of a helix within the sensor II motif (182). Properly folded DnaA produces a nucleotide binding pocket at the juncture between Base and Lid that is accessible to all of the aforementioned motifs simultaneously.
Domain IV (amino acids 347 to 467) is the DNA binding domain that interacts with 9-mer recognition sites (74, 206, 239), and also contains amino acids that are responsible for membrane interaction (82).
Structural studies of DnaA domain III and IV in the thermophilic bacterium Aquifex aeolicus (64, 65) reveal a novel basis for ATP-dependent DnaA assembly into an oligomer. DnaA-DnaA interactions are modeled within a bipartite nucleotide binding pocket comprising sensor II residues of one protomer and the box VII residues of its neighbor. An adjacent box VII arginine finger becomes accessible to its ATP-bound neighbor because of an open configuration of the Base and Lid that does not exist in DnaA-ADP (64) (Fig. 2). Through the binding and hydrolysis of ATP, DnaA switches back and forth between oligomer-promoting and oligomer-inhibiting states. The assembled DnaA oligomer exists as an extended right-handed superhelical filament rather than the ring shape often observed for AAA+ proteins, due to amino acid residues that form a V-shaped steric wedge protruding from domain III in such a way as to prevent a flat ring assembly (64).
Structural studies have also revealed two modes for DnaA interaction with DNA. The axial channel produced by the assembly of the DnaA-ATP helical filament has been proposed to directly engage unwound single-stranded DNA in the origin (64, 196) and provide a structure to assist in the delivery of DnaB-C complex (177). Domain IV contains a helix-turn-helix (HTH) motif for double-stranded DNA binding termed the DnaA signature sequence (KDHTTVI), as well as a proximal region termed the Basic loop (65). Both major and minor grooves of an oriC R box interact with DnaA (74). One helix and one loop of the HTH insert into the major groove, and an arginine in the Basic loop is recognized by bases in the minor groove (65, 74). DNA is bent by 28 degrees when bound to DnaA (74).
dnaA mutants.
DnaA is an essential protein, but a variety of conditional lethal (temperature-sensitive) mutants have been isolated; for example, see references 39 and 132. DnaA(ts) mutants include alleles with defects in domain I (dnaA508), domain III (dnaA5, dnaA46, dnaA601/602, and dnaA604/605), and domain IV (dnaA203/204) (97), although most also contain an additional amino substitution (97). Excellent reviews of these dnaA(ts) alleles are available (169, 224), and the reader is referred to them for more details. It should be noted that the domain III mutants all share the same amino acid substitution in the ATP binding cleft, which helped identify the importance of this region of DnaA for nucleotide binding and activity. More recently, mutagenesis of DnaA has been driven by structural studies to identify amino acid residues critical for domain-specific functions; for examples, see references 68, 124, 125, and 239.
Regulation of dnaA expression.
The dnaA gene, located 42 kbp counterclockwise to oriC (25), is the first gene in an operon that also contains dnaN and recF (92, 96, 192). The dnaN gene encodes the β-clamp of DNA polymerase III holoenzyme and the recF gene encodes a DNA recombination protein. The entire operon is transcribed from two promoters upstream of the dnaA gene, termed dnaA1p and dnaA2p, although dnaN and recF also appear to have separate promoters (4). The two dnaA promoters are separated by approximately 80 bp, and this region of DNA contains a consensus DnaA box and one box deviating from consensus in one position (96), separated by a putative third box, deviating from consensus at two positions (95). There are also three ATP-DnaA boxes (Speck-Messer sites) flanking the DnaA boxes (232). Both dnaA promoters are negatively regulated by the DnaA protein (9, 33, 138, 169), and it was proposed that a DnaA oligomer formed around the consensus DnaA box, and blocked RNA polymerase from binding to either dnaA1p or dnaA2p (144). Autorepression appears to be largely mediated by DnaA-ATP binding to the promoter region, with DnaA-ADP having less of an effect on transcription (232). Other negative regulators, including the DNA bending protein Fis, and the stringent response nucleotide ppGpp, also have been reported to bind to the promoter region, causing growth-rate regulation of transcription (43, 73, 201), although there are conflicting data as to whether or not the cellular levels of DnaA protein are also growth rate regulated (43, 93). The promoter region also contains several GATC sequences that, as described below, are involved in binding the SeqA protein immediately after the gene is duplicated (38, 138, 158); this binding represses transcription (38).
Transcription of the dnaA gene fluctuates during the cell cycle, primarily due to SeqA blocking the hemimethylated promoter region (38, 241) (described in more detail below). It seems less likely that cell-cycle-specific dnaA expression results from autorepression, since cellular DnaA-ATP levels are highest immediately before initiation (140), and transcription of the dnaA gene is not decreased until after initiation of new DNA synthesis (28). Rather, it appears that autoregulation may be more important in allowing the expression level of dnaA to increase when additional copies of oriC, or other DnaA titrating sites, are present in the cell. Addition of DnaA titrating sites to E. coli causes an increase in expression from the dnaA gene, proportional to the number of DnaA molecules bound by the titrating site (98, 174). This derepression would allow rapidly growing cells to accommodate additional copies of oriC without changing the cellular age at initiation.
DnaB and DnaC.
DnaB helicase (50 kDa) oligomerizes into a hexomeric toroidal ring with a central channel (15, 19, 58, 143) that encircles single-stranded DNA and unwinds the helix in advance of the replication fork. To load onto single-stranded DNA, the DnaB ring must be broken by DnaC-ATP (29 kDa) (130, 251, 252), a structural paralog of DnaA (177). After loading DnaB onto oriC, DnaC dissociates from the complex (66). DnaB-C loading also requires DnaA since mutations in the N-terminal region of DnaA disrupt the loading (237), although details regarding DnaA's role in the process are lacking. A recent model (177) suggests a novel mode of interaction among DnaA, B, and C during helicase loading and is discussed below.
DnaC is essential for initiation, and temperature-sensitive, conditional lethal dnaC mutants have been used extensively to align initiation of chromosome replication in populations of growing E. coli cells (39). Cultures of a dnaC(ts) strain that are shifted to 42°C for 1 h complete all previously initiated rounds of DNA synthesis, but are unable to start new rounds. During the time at nonpermissive temperature, all other components necessary to intiate new rounds are synthesiszed in excess so that, on shifting back to permissive temperature (25°C), all oriC copies intiate DNA synthesis simultaneously (91). Initiation potential for two rounds of DNA synthesis accumulates at the nonpermissive temperature, and a second burst of initiation follows about 10 min after the first. The time interval between these successive initiations is termed the eclipse period (245) and is considered to be the minimum time during which a cell is capable of successive initiations from the same origin. It is not clear what normally determines the length of the eclipse period, but the topological state of oriC and sequestration (see below) may contribute (116).
Analysis of A. aeolicus DnaC structure reveals that it shares DnaA's ability to form a right-handed helical filament (177). This structural similarity is sufficent to allow placement of DnaC AAA+ domains onto the end of a DnaA oligomer in an ATP-dependent fashion. It has been proposed that DnaC-ATP associates with the DnaA bound to the 13-mer region to specifically position DnaB on top and bottom strands within the unwound region of oriC (177).
E. coli and S. enterica each have a single, fixed origin of chromosome replication termed oriC, which in E. coli K-12, comprises nucleotides 3,923,767 to 3,924,025 on the E. coli K-12 genomic sequence (25). In E. coli and S. enterica, the oriC region does not encode any proteins involved in DNA replication (Fig. 3). This is not true for all eubacteria, because many harbor dnaA within 2 to 3 kb of oriC. In E. coli and S. enterica, DnaA remains the closest replication gene to oriC, but the two loci are separated by 42 kb (186). The genes directly flanking oriC are mioC (clockwise) and gidA (counterclockwise) (Fig. 3). mioC encodes a flavodoxin implicated in biotin synthesis (21), and gidA encodes a flavin adenine dinucleotide binding protein that modifies tRNA (170, 263). The protein products of both genes are dispensable, but transcriptional activity from both gidA and mioC promoters is implicated in oriC regulation (see below). Immediately beyond mioC and gidA lie genes encoding asparagine synthetase (asnC and asnA) and ATP synthetase, respectively.
Cloned versions of oriC form minichromosomes.
Although the location of oriC was precisely mapped on the E coli chromosome, the size of the chromosome was prohibitively large for detailed functional analysis. This problem was solved by constructing oriC plasmids (termed minichromosomes), selected for autonomous replication in the absence of any other replication origin (250, 262). Minichromosomes share regulatory attributes with the chromosome including bidirectional replication (166), dependence on the same DNA replication factors (250), and synchronous, cell cycle-specific, one round per cell cycle initiation (103, 134, 148). Although minichromosomes replicate only once per cell cycle, they lack an equipartition mechanism and tend to segregate in clumps, causing recipient cells to harbor more than one copy per cell (52). Synchronous cell-cycle replication of up to 30 additional oriC copies is permitted, demonstrating that the levels of diffusible factors are not generally limiting (151). Cell cycle-specific minichromosome replication does not depend on chromosomal oriC function, and minichromosomes are retained in cells with randomly replicating chromosomes, in which chromosomal oriC is replaced by a plasmid replication origin (62). Combined, these studies demonstrated that the wild-type oriC sequence contains all of the information needed to direct cell cycle-specific timing of initiation.
Sequence features of oriC.
Functional oriC can be dissected into two distinct regions that play different, but interrelated roles in initiating DNA replication. The right/central region (bases 80 to 270; Fig. 3) contains a set of protein recognition sites that are separated by fixed numbers of base pairs (5), forming a platform for ordered assembly of multiple DnaA molecules (discussed in more detail in a later section). The leftmost region of oriC (bases 10 to 66 of Fig. 3) contains the DNA Unwinding Element (DUE) (137); an A-T-rich region with three tandem 13-mer repeats of 5′-GATCTNTTNA/TA/TA/TG/T-3′, where the DNA helix unwinds in response to assembly of a higher-order DnaA complex (32). In the single-stranded state, the 13-mers in the DUE are reported to interact directly with DnaA-ATP, at several 6-mer A/TGATCT motifs (referred to in Fig. 3 as Speck-Messer, or S-M, sites) (231), similar to the ATP-DnaA boxes observed in the dnaA promoter (232). Interaction of DnaA with these sites is proposed to stabilize origin unwinding (196, 231). An additional AT-rich sequence is found between bases 10 and 20 at the left-most end of oriC, and although it is required for proper oriC function, its specific role remains to be determined (7).
The DUE is separated from the right/central DnaA-loading platform by a precise distance of 13 bp (encompassing bases 67 to 79); adding or deleting nucleotides in the spacer sequence is not tolerated (109). The right/central region contains at least ten DnaA 9-mer recognition sites whose occupation is detected by DNA footprinting (79, 124, 146, 163). The nucleotide sequence and positions of all DnaA binding sites in oriC are highly conserved among all members of the Enterobacteriacae (99, 271).
Each DnaA recognition site in oriC may be classified based on two criteria: affinity for DnaA and preference for DnaA nucleotide form. There are three high-affinity (KD = 4 to 20 nM) (215) consensus recognition sites: R1, R2, and R4 (Fig. 3). These sites bind DnaA-ATP and DnaA-ADP equally well in vitro (79, 215). An additional seven low-affinity (KD > 200 nM) (215) sites have been identified in oriC; these deviate from the consensus sequence at one or more bases. Of the lower-affinity sites, I sites (I1, I2, and I3) (87) and τ-sites (τ1 and τ2) (124) clearly discriminate between nucleotide forms of DnaA, showing a 4-fold preference for DnaA-ATP (124, 165). There are conflicting data regarding binding of DnaA-ADP to R5M and R3; DMS footprinting studies indicated that these two sites have no preference for a particular DnaA nucleotide form (88, 165), while DNase I footprinting experiments suggested that all low-affinity sites, as well as R2, preferred to bind DnaA-ATP (124). The reason for these differing results is not clear, but may reflect a difference in the sensitivity of the methods used to examine binding. DnaA binding to its high- and low-affinity recognition sites is discussed in greater detail in later sections.
In addition to DnaA recognition sites, oriC contains specific binding sites for two DNA bending proteins, Factor for inversion stimulation (Fis) and Integration Host Factor (IHF) (70, 200, 208). The Fis binding site lies in the right half of oriC between R2 and R3, and IHF binds between R1 and R5M (Fig. 3). Both proteins modulate DnaA interactions, as discussed below.
The entire oriC sequence contains an unexpected number of GATC palindromes (Fig. 3), with four located in the 13-mer region and seven found between bases 80 and 250. No GATC is found within any high-affinity DnaA recognition site, but four low-affinity recognition sites (τ1, τ2, I2, and I3; Fig. 3) contain an internal GATC, and R5M overlaps a GATC. GATC is the recognition sequence for both deoxyadenosine methyltransferase (dam methylase) and the hemimethylated DNA binding protein, SeqA. The DNA replication-dependent conversion of fully methylated GATC to the hemimethylated state plays an important regulatory role in limiting initiations from oriC and resetting the origin for the next cell division cycle (described in more detail below).
Several transcriptional promoters were also previously mapped within oriC (115, 216), but a specific role during initiation for these transcripts remains unclear.
Mutational analysis of oriC.
Mutations have been placed at specific locations within oriC to evaluate the roles of individual protein binding sites, as well to examine the role of the intervening nucleotides between DnaA recognition sites (108, 141, 165, 194, 195, 208, 254, 257). In general, mutations are introduced into oriC on plasmids that carry an additional origin of replication, such as ori pBR322. This chimeric configuration (149, 255) allows recovery of defective versions of oriC, and the extent of the defect can be assayed in strains lacking DNA polymerase I (required for initiation from ori pBR322, but not oriC).
While some single-base-pair changes in DnaA recognition sites do not affect either DnaA binding or in vivo replication (108), mutations that reduce DnaA binding resulted in decreased minichromosomal oriC function, as do mutations that alter helical phasing between recognition sites (141, 165). The loss of minichromosome replication in vivo indicates that these mutations cripple oriC so that it is unable to compete effectively with the wild-type chromosomal copy (108, 151). Combined, the results of these studies support a role for all DnaA recognition sites and spacer regions during normal assembly of higher-order complexes. However, with the exception of mutations that eliminate binding to R1, reducing the activity of a single recognition site does not completely inactivate oriC when it is functioning as the sole origin on the chromosome, although many mutations alter initiation timing and/or perturb initiation synchrony (20, 203, 254). This resilience of function suggests that there are likely to be multiple ways for DnaA complexes to assemble, unwind oriC, and load helicase. However, the failure of mutant origins to function in the presence of wild-type competition suggests that none of the potential alternative assembly mechanisms is as efficient as the normal pathway. A model for the precise assembly of prereplication complexes based on binding to all high- and low-affinity recognition sites is discussed in a later section.
Mutations have also been made in the binding sites for Fis and IHF (50, 208), as well as in the GATC sequences in the left part of oriC (13). These mutations are discussed below.
Although DnaA, DnaB, and DnaC are essential genes, there are other regulators of initiation that can be mutated without loss of viability, but whose inactivation results in abnormal initiation timing during the cell cycle. These regulators generally work by: (i) controlling availability of active initiator DnaA (Fig. 4A), (ii) enhancing or repressing the ordered assembly of DnaA complexes at oriC (Fig. 4B), or (iii) conditionally blocking initiation in response to unfavorable conditions (Fig. 4B). It is sometimes difficult to evaluate the function of these regulators by examining the phenotype of mutant strains with severe growth perturbations, since compensatory mutations that improve cell survival may arise spontaneously in strains with severe initiation defects (205).
Architectural proteins IHF, HU, and Fis: DNA bending proteins that modulate assembly of DnaA complexes at oriC.
Fis (71), IHF (63), and HU (59) are small, abundant, nucleoid-associated proteins that bend DNA and regulate many aspects of chromosome biology, including specific steps in orisome assembly. IHF places a severe bend (120°) in oriC between R1 and R5M, while the Fis-induced bend, between R2 and R3, is less acute, at 55° (85, 200, 201). HU lacks a specific recognition site in oriC, but binding of several HU molecules is reported to introduce a curve into DNA (30, 107). Minichromosomes cannot be stably maintained in cells lacking Fis or HU (70, 85, 189), while IHF mutants are able to maintain minichromosomes unless polA is also absent (69). The reason for the polA requirement is not clear, but it has been proposed that these cells use an alternative mode of DNA replication (69).
Both IHF and HU are enhancers of DnaA-catalyzed unwinding of oriC (56, 110), but two different mechanisms are involved (210). IHF binding promotes DNA unwinding by increasing DnaA occupation of low-affinity recognition sites (87). IHF mutants initiate DNA replication later in the cell cycle than wild-type cells, and they have asynchronous, rifampin (rifampicin)-resistant initiations (31, 248), a phenotype that is consistent with a requirement for a higher cellular level of DnaA to initiate chromosome replication. Scrambling the IHF binding site on oriC eliminates minichromosomal oriC function, and causes late and asynchronous initiations on the chromosome (208, 254), providing supporting evidence for IHF's role as a positive regulator of initiation.
The mechanism used by HU to assist oriC unwinding is not yet clear. HU does not appreciably alter DnaA binding to oriC (211), although it might affect IHF binding under some conditions (30). It is possible that, by placing curves in the oriC DNA, HU increases torsional stress and stabilizes unwinding. HU was also recently shown to interact directly with DnaA (44), which could be a mechanism to bring HU to specific positions within the orisome. HU mutants, in particular, those with defects in the α-subunit of HU, display an asynchronous phenotype (14), indicating that HU and IHF are not interchangeable in vivo.
Fis represses oriC unwinding (210) and in vitro replication of oriC plasmids (104, 258) by blocking DnaA binding to low-affinity sites (210). Fis also inhibits IHF binding to oriC (210). In vivo, cells lacking Fis initiate asynchronously (31). Mutant versions of oriC with a defective Fis binding site are not functional as minichromosomes (50, 85), and replicate with altered timing (earlier) when operating as the chromosomal origin (203). Fis occupies its primary site in oriC throughout the cell cycle in rapidly growing E. coli, but DnaA-catalyzed displacement of Fis near the time of initiation (40) promotes binding of IHF (210). Fis is the only bending protein in this group of initiation regulators whose synthesis is growth-rate regulated (18), and its presence at oriC is likely to also be growth-rate dependent.
dam methylation and SeqA protein.
dam methylation of GATC (83) plays a role in many different DNA-based activities in E. coli including gene transcription, DNA mismatch repair, initiation of chromosome replication, and maintenance of nucleoid structure (155) (for more details, the reader is referred to chapter 4.4.5). As mentioned above, oriC contains an unusually high number of GATC motifs (Fig. 3), and GATC is also found in the promoter region of dnaA, allowing methylation to play a regulatory role in both DnaA synthesis and DnaA binding to oriC (34, 181).
After the passage of a replication fork, fully methylated GATC on duplex DNA becomes transiently hemimethylated, but in most regions of the genome, remethylation by dam methyltransferase is rapid. However, both oriC and the DnaA promoter region remain hemimethylated for about one-third of the cell cycle, termed the sequestration period (38). During sequestration, hemimethylated oriC is inactive, and for this reason fully methylated minichromosomes are unable to replicate in Dam− strains (209, 229). In addition, transcription from both dnaA1p and dnaA2p is blocked during the sequestration period (38).
Sequestration is mediated by the protein SeqA (21 kDa), which preferentially binds hemimethylated GATC sequences (158, 228). E. coli mutant strains lacking SeqA activity have asynchronous initiations, and suffer from overinitiation of DNA replication (158). In oriC, SeqA binding to hemimethylated GATCs blocks rebinding of DnaA to low-affinity sites after initiation (181) (discussed in a later section). Presumably, SeqA also blocks RNA polymerase from binding to the dnaA promoter region. The orientation of the 11 GATCs within oriC is sufficient to allow prolonged binding of SeqA and slow remethylation, since this pattern of sites also shows delayed remethylation if inserted at non-oriC regions of the chromosome (12). Further, mutation of GATC sequences in the left region of oriC resulted in loss of sequestration and overinitiation (13). Hemimethylated oriC is also associated with E. coli's membrane fraction (51, 191) and this interaction is reported to require SeqA (228).
SeqA forms homotetramers that bind to two hemimethylated GATCs that are separated by no more than 31 bp (35). Based on structural analysis, SeqA forms higher-order oligomeric filaments (89) similar to DnaA (64). With an intracellular concentration of about 1,000 molecules (228), there is also sufficient SeqA to perform other duties in the cell including formation of aggregates that travel with the replication forks (106, 261) and a role in chromosome organization (243). While these attributes of SeqA are beyond the scope of this chapter, interested readers are referred to an excellent recent review (253).
DiaA: a DnaA binding protein that enhances DnaA oligomerization.
DiaA is a recently discovered positive regulator of initiation, identified in a screen for mutations that suppress the overinitiation phenotype of dnaAcos (113). DiaA-deficient strains are viable, but show perturbed initiation timing, with delayed, asynchronous initiations in rich media (113). DiaA stimulates initiation by enhancing oligomeric DnaA binding to oriC, so that less DnaA is required for unwinding of the 13-mer region (77, 128) (Fig. 4B). The active form of DiaA (50 to 60 kDa) is a homotetramer that interacts specifically with the N terminus of DnaA (128). A single homotetramer can bind to multiple DnaA molecules simultaneously and enhance assembly of both DnaA-ATP and DnaA-ADP complexes at oriC (128). About three DiaA tetramers are associated with a complete DnaA complex at oriC (113, 128). DiaA also contains a putative phosphosugar binding domain that may regulate DiaA activity.
Dps and IciA: conditional repressors of initiation.
Dps (178) and IciA (242) are negative regulators that act in response to stress or starvation conditions to decrease initiation from oriC (Fig. 4B). Dps is a DNA binding protein that is abundant in stationary phase cells and in starved cells (2), where it appears to play a role in protecting the bacterial genome from environmental stress. Dps also is responsible for increased chromosome condensation in stationary phase (193). Recently, Dps was shown to interact directly with the N terminus of DnaA and to impede in vitro replication by interfering with oriC unwinding (45). This repression of initiation may allow stress-induced DNA damage to be repaired in the absence of new replication forks.
IciA is a 33-kDa protein that blocks oriC unwinding by interacting directly with oriC within the A-T-rich 13-mer region (111, 112). IciA is a member of the LysR family of transcription regulators, and was shown to be identical to ArgP, a regulator of arginine transport (42). IciA/ArgP levels increase in response to phosphate starvation (90), and presumably block initiations under conditions where successful elongation is unlikely. IciA also binds to the dnaA promoter region, simultaneously enhancing transcription and blocking DnaA autoregulation (145). Thus, during phosphate starvation, while oriC is inactivated, levels of DnaA-ATP would be expected to increase and remain high enough to trigger initiation once phosphate was no longer limiting. When conditions are appropriate for initiation, IciA levels are reduced by specific proteolysis (264).
Hda: postinitiation inactivator of DnaA.
As discussed above (section on DnaA), DnaA-ATP is rapidly converted into the inactive ADP form during RIDA (120) (Fig. 4A). Initially, the factors involved in RIDA were termed IdaA and IdaB (119, 121, 139); IdaA was subsequently found to be the sliding clamp of the DNA polymerase holoenzyme (121), and IdaB was isolated and renamed Hda (123), since the protein is homologous to DnaA. Hda forms a stable complex with the sliding clamp, where its primary function is to stimulate the intrinsic ATPase activity of DnaA (121, 236). The mechanism used by Hda to do this is not fully characterized, but it appears that active Hda must be bound to a DNA-loaded clamp (126). The interaction of Hda with DnaA also requires amino acid residues in DnaA's N-terminal region (236). Hda is a member of the AAA+ family of ATPases (123), but there is no evidence that it binds ATP (235). Rather, a conserved arginine (arg-168) in the box VII motif (arginine finger) is required for the stimulation of hydrolysis, and a recent structural analysis of Hda suggests that the box VII region interacts directly with the ATPase region of DnaA (260). ADP binding to Hda may also be important for in vivo activity (235).
Because Hda activity depends on association with DNA-loaded sliding clamp, RIDA is coordinated with movement of replication forks. It was first proposed that, as the newly formed replication forks proceed from oriC, the associated clamp/Hda interacts with DnaA-ATP, stimulating ATP hydrolysis (236). In this scenario, DnaA would have to be present at the fork. It is also possible that some clamps are left behind on the DNA after lagging strand synthesis, and these “leftover” clamp/Hda complexes might also stimulate DnaA-ATP hydrolysis. In either case, DnaA-ATP inactivation would not be expected during cell cycle periods devoid of ongoing DNA synthesis, which occur in slowly growing cells (see Fig. 1).
There is controversy in the literature regarding the phenotype of hda mutants. Loss of Hda activity is reported to produce effects of varying severity, ranging from lethality (123) to significant overinitiation (36) or only modest overinitiation and asynchrony (37, 205). However, several compensatory mutations were identified in Hda-defective strains (205), and much of the variation in phenotype could be explained by differences in the compensatory mutations generated in the strains used in the studies.
datA and other chromosomal DnaA titration sites.
There are approximately 1,000 molecules of DnaA per E. coli cell (93, 221). Despite the need to activate multiple oriC copies in rapidly growing cells, this level of DnaA is unexpectedly high, and it is clear that there must be negative regulators of DnaA activity to ensure that excess free DnaA-ATP is not available to bind oriC and promote unscheduled initiation events. There are several ways this can be accomplished, two of which (RIDA and sequestration) were discussed above. An additional mechanism is to titrate DnaA away from oriC by providing competing initiator recognition sites on the chromosome. As originally formalized in the initiator titration model (94), ongoing DNA replication causes the duplication of DnaA recognition sites dispersed along the chromosome that bind DnaA and lower its availability. Orisome assembly is then prevented until the number of newly synthesized DnaA molecules is greater than the number of titration sites. For initiator titration to be effective, the relative affinities of DnaA recognition sites within oriC must be lower than the titration sites dispersed on the chromosome. This requirement raises interesting issues for mechanisms that precisely order orisome assembly (see below).
Although some DnaA recognition sites on the E. coli chromosome are in gene promoter regions, where DnaA acts as a transcriptional regulator (168), there are other chromosomal DnaA recognition sites that appear to serve no role other than to titrate available DnaA away from oriC. The primary DnaA titration activity on the E. coli chromosome is localized within a 950-bp region, termed datA, that carries five DnaA recognition sites (190). datA is reported be able to bind large amounts of DnaA (up to 300 molecules) (129). Such high titration capacity from only a few recognition sites must require cooperative protein-protein interactions and a higher-order structure among assembled DnaA molecules. Although loss of datA is not lethal, initiations from oriC become asynchronous and an overinitiation phenotype was observed by flow cytometry (129). More recently, datA mutants were shown to continue to initiate in the presence of rifampin (175), resulting in the apparent asynchronous phenotype. Rifampin-resistant initiations do not occur in normal cells, and generally indicate that there is an increased availability of DnaA in the cells. Excess datA delays initiations from oriC (173, 174), as would be expected if it were titrating DnaA away from oriC.
Membrane phospholipids and DARS: two DnaA-ADP recharging mechanisms.
DnaA-ADP produced by RIDA does not spontaneously reactivate by exchanging bound ADP for ATP (218). Rather, two different recycling pathways exist to regenerate DnaA-ATP, and these pathways may be required, at least under some growth conditions, to supplement the amount of DnaA-ATP generated by new synthesis (76, 147, 259). The first pathway depends on acidic phospholipids (26). DnaA interacts directly with phospholipids via a short stretch of amino acids between domains III and IV (26, 65) (Fig. 2), and in vitro, acidic phospholipids can catalyze exchange of DNA-bound DnaA-ADP to DnaA-ATP (48, 220). In cells, DnaA associates with phospholipid membranes (27, 179), and about one-half of the intracellular DnaA appears to be retained in the insoluble lipid fraction after cell lysis (221). E. coli cells that are unable to make the acidic phospholipids phosphatidylglycerol and cardiolipin are growth arrested at the initiation stage (49, 259), and this growth arrest can be suppressed by mutations in the DnaA membrane binding domain (e.g., DnaAL366K) (270), although the mechanism for the suppression is not yet clear. It is also suggested that a high local DnaA concentration in the membrane may be necessary to regulate ADP to ATP exchange (3).
A second recharging mechanism requires specialized chromosomal DNA sequences termed DARS (DnaA Reactivation Sequences) (75, 76). Although originally identified in the ColE1 replication origin, DARS1 and DARS2 are found on the E. coli chromosome near bioD and mutH, respectively (76). While the exact recharging mechanism remains to be elucidated, DARS share two closely spaced DnaA recognition sites that are oriented in back-to-back positions, and this configuration is required to stimulate the conversion of bound DnaA-ADP into the apo-DnaA (no-nucleotide) form. Once the nucleotide is removed, DnaA quickly recharges back to DnaA-ATP spontaneously because of the higher cellular ATP concentration. Loss of DARS delays initiation during the cell cycle (76), suggesting these sequences normally contribute to the threshold level of DnaA-ATP required for initiation. A yet to be identified soluble factor has also been implicated in the regulation of DARS2 activity (76).
Topoisomerases and RNA Polymerase.
Initiation of E. coli chromosomal DNA replication requires supercoiled oriC DNA (32), and any changes in the level of DNA supercoiling or changes in transcriptional activity from promoters adjacent to oriC have the potential to alter origin function. For this reason, both DNA topoisomerases and RNA polymerases must be considered regulators of oriC activity.
DNA supercoiling is altered in strains carrying mutations in topoisomerase genes (84). Increased supercoiling, as is seen in strains with mutations in topA encoding topoisomerase I, suppresses the temperature sensitivity of the dnaA46(ts) allele, which normally underinitiates (157). Decreased supercoiling also causes asynchronous initiations from oriC (246, 247), and it adversely affects minichromosome maintenance (150). These findings suggest a direct relationship between DNA supercoiling and initiation efficiency, most likely by modulation of the amount of DnaA required to unwind oriC.
Transcription near oriC generally promotes initiations under conditions where DnaA levels are deficient (17). In vivo, the gidA promoter, located to the left (counterclockwise) of oriC, is the most likely candidate for a transcriptional activator of initiation, since the direction of gidA transcription should generate negative supercoiling within oriC (see Fig. 3). In support of this idea, gidA transcription is reported to be required for normal origin function (7), and the gidA promoter is shut down during oriC sequestration (28), as would be expected for a positive modulator of initiation. The mioC gene on the right (clockwise) side of oriC was also originally thought to play a role in initiation timing (mioC stands for modulator of initiation at oriC). mioC transcripts enter and pass through oriC (28, 29, 115, 156, 216). Normally, mioC transcription is shut down by DnaA binding to the mioC promoter, shortly before initiation (29, 187, 234). Continuous mioC transcription is reported to be deleterious to oriC function (183, 234), but does not completely repress new rounds of DNA synthesis (29). However, despite its intriguing cell cycle-specific regulation, mioC is not necessary for proper timing of initiation, since its deletion does not affect replication timing of either minichromosomes (148) or the chromosome (152).
Direct interaction of DnaA and RNA polymerase was recently detected by immunoprecipitation (72), and this observation is consistent with previous findings that several mutations in rpoB (encoding RNA polymerase β-subunit) are able to suppress temperature-sensitive dnaA initiation mutants (8). Further examination of this interaction may reveal new roles for both proteins at specific stages of initiation.
The development of in vitro systems to study E. coli DNA replication revolutionized our understanding of the steps used to assemble the replication machinery at oriC (135, 219). By characterizing the complexes generated by combining purified replication proteins and supercoiled oriC plasmids (16, 79, 80, 219), the Kornberg laboratory defined a series of stages required to initiate DNA synthesis in vitro from oriC (described below).
Initial complex.
The initial complex was originally defined as the association of DnaA with R boxes on oriC templates (219). Neither supercoiled oriC nor ATP is required to assemble this complex. When examined by electron microscopy, the initial complex was classified further based on correlation of a specific structure with in vitro replication activity (50). The active initial complex was a compact ellipsoid containing approximately 200 bp of oriC wrapped around approximately 20 molecules of DnaA-ATP, with the DNA strands crossing as they emerge from the ellipse (50, 80). At lower DnaA concentrations, smaller complexes were observed, suggesting that initial complexes are built from distinct subcomplexes (50). The observation that small complexes transitioned into a reproducible and uniquely shaped active initial complex was one of the first indications that DnaA interaction with oriC is ordered (discussed in greater detail below). A complex that is similar in appearance to the compact ellipse can be formed with DnaA-ADP, but the DnaA-ADP complex is inactive in the in vitro replication assay (50).
Open complex.
The second distinctive stage, the open complex, is characterized by DnaA-mediated unwinding of supercoiled oriC templates in the absence of DNA helicase (219). The location of the single-stranded bubble has been mapped by its sensitivity to P1 endonuclease, and is found within a 26-bp region between the middle and rightward 13-mer repeat in the DUE (32, 85). Patterns of cutting are different for the top and bottom strands (32) and may result from one strand interacting with DnaA more extensively than the other strand (196, 231). DnaA-ATP is sufficient to unwind oriC in vitro at 37°C, but HU or IHF must be included for any unwinding to take place at lower temperatures (110). The presence of HU or IHF also reduces the amount of DnaA that is required for unwinding at 37°C (56, 87, 110). In addition, while it is clear that some of the DnaA used to make the open complex must be in the ATP-bound form, a combination of DnaA-ADP and DnaA-ATP can support in vitro replication of oriC plasmids (265). The DnaA-ATP requirement for initiation is discussed in more detail below.
Conversion of the initial complex, made with either all DnaA-ATP or a combination of DnaA-ATP and DnaA-ADP, to the open complex requires the additional presence of high ATP concentrations (5 mM) (265), even though the KD for ATP binding to DnaA is much lower (30 nM) (218). The reason for this requirement remains unclear, since there is no evidence suggesting that there is more than one ATP binding site in DnaA.
Prepriming complexes I and II are stages of helicase loading that follow open complex formation. Prepriming complex I is defined by two DnaB-C complexes (251, 252) loaded sequentially at unwound oriC (top and bottom single strands). In prepriming complex II, the loaded helicases are translocated within oriC, expanding the unwound region sufficiently to assemble the replisome (16, 80).
Prepriming complex I requires addition of DnaA-ATP, DnaB, and DnaC to the supercoiled oriC template, with HU, if present, stimulating the reaction (80, 218). DnaA-ATP is necessary to produce both the open complex and to stably retain of DnaB in the prepriming complex (237). The latter function is mediated through direct interaction of the N-terminal domain of DnaA with DnaB (237).
DnaC loads replicative helicase by interacting with a 1:1 stoichiometry on one face of the DnaB hexamer, creating a gap in the ring through which single-stranded DNA can pass (54). DnaC binds both ATP and ADP, and the dual ATP/ADP switch activity is necessary to form the prepriming complexes. DnaC-ATP is proposed to first interact with single-stranded DNA, expanding the unwound bubble at oriC as it delivers DnaB. DnaC-ATP also inhibits helicase activity, preventing the DnaB oligomer from translocating until it is properly loaded (53). DnaC is also modeled to interact directly with DnaA-ATP oligomeric filaments, directing helicase loading to the correct location on the top strand of the unwound 13-mer region (177). In the presence of DnaB and single-stranded DNA, DnaC-ATP is hydrolyzed to DnaC-ADP, activating helicase and allowing its translocation along the DNA (53), forming prepriming complex II. The association of DnaC with oriC is transient, and DnaC cannot be detected in the prepriming complex after it is assembled (80).
Helicase hexamers are loaded face to face (one on each strand) and pass one another as they move along the DNA (66). By passing one another in this way, the unwound region remains melted and can be coated with single-strand binding protein prior to priming and replisome assembly.
Priming complex and replisome assembly stages do not require DnaA; for details, the reader is referred to chapter 4.4.2. Limited helicase unwinding (65 bp or more) is required prior to the interaction of DnaG primase. Primase interacts with the moving helicase and, for this reason, RNA/DNA junction positions are not fixed (66). Predominant priming sites fall within oriC (between the rightmost 13-mer and R1) for counterclockwise synthesis, but outside oriC (approximately 60 to 70 bp left of the left boundary) for clockwise synthesis (131). The primases also appear to pass one another during bidirectional replication. Following the synthesis of two 10- to 12-bp primer RNAs per oriC, two β-clamps assemble on the DNA followed by two DNA polymerase III complexes (66).
Although characterization of in vitro complexes provided invaluable information regarding the biochemical processes needed to begin DNA replication, these studies did not address how the orisome was assembled in growing cells at the correct time during the cell cycle. To answer this critical question, high-resolution in vivo footprinting methods were developed, making it possible to follow the occupation of DnaA recognition sites in oriC as a function of cell cycle timing (40, 181, 211, 212). These studies revealed that the three highest-affinity sites, R1, R2, and R4, are the only sites that become occupied immediately after initiation during the sequestration period (181). Further, these three sites remain the only oriC sites bound by DnaA throughout the majority of the cell cycle (40, 181, 211, 212). This persistent DnaA complex is temporally similar to the nucleoprotein structure observed at replication origins in budding yeast, in which the Origin Recognition Complex (ORC) proteins remain associated with replication origins throughout the yeast cell cycle (233). For this reason, the DnaA complex bound to R1, R2, and R4 is defined as the bacterial ORC (181). X-ray crystallographic analysis of DnaA from A. aeolicus supports this definition by revealing that DnaA has remarkable structural similarity to components of the eukaryotic ORC and to archaeal initiators (60, 65). In addition to regulation by ATP, bacterial and eukaryotic initiator proteins appear to share the ability to form right-handed helical filaments (46, 64, 185), and the structural similarites are sufficiently strong that it is possible to dock a helical pentamer of DnaA into the core of Drosophila ORC (46). Thus, it seems that all domains of life may use similar mechanisms to regulate initiation of DNA synthesis.
In E. coli, at the time of initiation of DNA replication, in vivo footprinting studies revealed that additional DnaA is bound to oriC, where it occupies the lower-affinity sites R5M, R3, I1, I2, I3, and τ2 (40, 181, 211) and mediates localized strand separation in the DUE (87, 165). This distinctive fully assembled DnaA/oriC complex with an unwound DUE is reasonably defined as the bacterial prereplication complex, pre-RC (181). In yeast, before entering S phase, a similar increase in protein binding to replication origins is observed, where additional proteins (Cdc6, Cdt1, and MCM2–7) associate with ORC to form yeast pre-RC before loading DNA polymerase (55). The similarities between eukaryotic and E. coli initiation can be extended further to include mechanisms used to load replicative helicase, with DnaA, DnaB, and DnaC being the functional equivalents of ORC/Cdc6, replicative helicase MCM 2–7, and Cdt1, respectively. However, the distinct DNA strand melting observed in the E. coli pre-RC before helicase loading has not been observed at eukaryotic origins.
In vitro, the differing affinities of the DnaA recognition sites in oriC causes DnaA loading to take place in the order: R1 = R4 > R2 > R5M = I2 = I3 > I1 = τ2 = R3 (87, 160, 165). In vivo, only two stages of DnaA occupation of oriC are easily observed: filling of R1, R2, and R4 (forming the bacterial ORC), and the filling of the remaining, lower-affinity sites to form the fully occupied pre-RC (181, 211). Conversion of the E. coli ORC to the pre-RC must proceed identically during every cell division cycle to ensure correct initiation timing. Since DnaA binding to high-affinity sites persists throughout the majority of the cell cycle, their occupation cannot be part of the timing mechanism, nor is the formation of bacterial ORC likely to be the target of mechanisms that regulate initiation timing. Rather, it is the filling of the lower-affinity sites by DnaA that is the rate-limiting and regulated process. Although in vivo, low-affinity sites appear to fill abruptly at the time of initiation, it seems likely that the ORC to pre-RC transition takes place in separate and successive stages, since this type of assembly would provide more opportunities for regulation.
Evidence for the existence of separate stages of pre-RC assembly comes from analysis of the binding patterns of accessory bending proteins Fis and IHF to oriC during the cell cycle, and a model for staged pre-RC assembly is shown in Fig. 5A. During rapid growth, Fis is associated with the bacterial ORC, occupying its primary recognition site immediately rightward of R2 throughout most of the cell cycle (40). In vitro analysis of the effect of Fis on DnaA binding to oriC revealed that the occupation of low-affinity recognition sites R5M, I2, and I3 requires higher DnaA levels than are needed in the absence of Fis (210).
The next observable stage of pre-RC assembly is the displacement of Fis. As DnaA levels are raised in vitro, and at the time of initiation in vivo, Fis is lost from its primary site in oriC (40, 210). Although the mechanism responsible for the displacement of Fis is unknown, one attractive possibility is that Fis is displaced by DnaA oligomers that extend between R2 and R4.
Immediately after Fis displacement, IHF is detected at its primary recognition site between R1 and R2 (40, 210). In vitro footprinting analysis demonstrated that Fis represses the ability of IHF to bind to oriC (210), although the mechanism for this repression is unclear. Once bound, IHF promotes DnaA occupation at low-affinity sites R5M, I1, I2, and I3, and it is this IHF-mediated stimulation that led to the naming of I sites (for IHF-stimulated sites) (87). The τ-sites, located to the left and right of R5M, are also reported to become occupied coincidently with the other lower-affinity sites in vitro and in vivo (124, 165, 181). It is possible that the bending of oriC DNA by IHF promotes interaction of the DnaA molecules bound to R1 and R5M, providing a stable anchor site from which to extend an oligomeric filament toward R2. This would help explain the rapid filling of the lower-affinity sites in this region, but again, such structures bound to oriC have yet to be observed.
The transition from Fis-bound ORC to IHF-bound pre-RC appears to be a switch-like mechanism for rapid conversion of ORC to pre-RC in a DnaA-dependent fashion. Because of the dynamic interplay among DnaA, Fis, and IHF (210), cell cycle-specific DNA bending by Fis and IHF would be an efficient mechanism to rapidly change positional relationships of DnaA binding sites during the cell cycle to either promote or inhibit the occupation of lower-affinity DnaA recognition sites.
The binding of IHF and the IHF-stimulated filling of low-affinity sites leads to localized unwinding of the DNA in the DUE (32, 87, 110, 210). After the 13-mer DNA becomes single stranded, there is evidence that DnaA-ATP associates with, and stabilizes the unwound region (196, 231, 266). Although this interaction has been proposed to be directed to single-stranded ATP-DnaA boxes (AGatct; labeled S-M sites in Fig. 3) found in each 13-mer repeat (231, 232), it seems likely that DnaA-ATP does not use domain IV for binding of single-stranded DNA. Instead, both structural and mutational analyses suggest that the central region of a DnaA-ATP filament associates with the single-stranded DUE (64, 196).
The mechanism by which the DnaA that is bound to high- and low-affinity sites in oriC mediates strand separation in oriC and directs helicase loading is among the least understood aspects of initiation in E. coli. Two models for unwinding, which are not mutually exclusive, have been proposed. In one, oriC DNA wraps around a right-handed oligomeric filament of DnaA-ATP. This wrapping would generate positively supercoiled DNA, but could simultaneously produce localized negative supercoiling in the adjacent DUE (64). In another model, the DnaA complex bound to oriC would prevent the DNA helix outside of the DUE from unwinding in the presence of topological change directed by transcriptional activity, presumably from gidA (6, 188). This inhibition of helical twist would thereby direct local unwinding to AT-rich regions within the DUE. Models for DNA helicase loading currently focus on the putative DnaA-ATP filament that extends into, but not through, the unwound 13-mer region. The availability of an oligomeric DnaA structure fixed in location is an attractive target for the interaction of DnaB-DnaC and the delivery of helicase to a specific location on both strands (177).
The role of DnaA-ATP in forming the pre-RC.
Although it is clear that DnaA-ATP is the active form of the initiator, the reason for this requirement is not immediately obvious, since R boxes bind DnaA-ATP and DnaA-ADP with equal affinities (165, 215, 218). Two compatible scenarios seem likely; first, that oriC contains binding sites with a preference for DnaA-ATP, and second, that DnaA-ATP is uniquely capable of forming a complex with the capability of unwinding the 13-mer region. Current data provide supporting evidence for both cases. Sites that preferentially bind DnaA-ATP have been identified, and they fall into two major classes. The DnaA-ATP-specific S-M sites are in the 13-mer region and preferentially bind DnaA as single-stranded DNA (231). I sites and τ-sites are located in the body of oriC between high-affinity sites (87, 124), and must be filled before strand separation (87, 165). I sites and τ-sites have a 3- to 4-fold preference for DnaA-ATP (124, 165). The placement of these sites may be critical for pre-RC formation, to ensure that DnaA-ATP is correctly positioned to form a specific structure, such as the previously mentioned right-handed helical filament (64).
Questions remain about the role of DnaA-ADP in the assembly of pre-RC. In vitro, only a fraction of the DnaA needed to support DNA replication needs to be DnaA-ATP (265), and our unpublished results indicate that open complexes can be made when only 30% of the DnaA in the reaction is in the ATP form. There is also in vivo evidence that DnaA-ADP is used during pre-RC formation, since increasing the requirement for DnaA-ATP by converting R5M to I2 makes the mutated oriC less efficient (88). Since structural studies indicate that only DnaA-ATP can form a helical filament (64), it remains to be determined how DnaA-ADP is accommodated in the pre-RC. It also remains uncertain to what degree oligomer formation in vivo requires assistance from other cellular factors. For example, DiaA stimulates formation of both DnaA-ATP and DnaA-ADP oligomers and may play a role in promoting specific interactions between DnaA molecules in growing complexes.
Synchronous initiation from multiple copies of oriC: the initiation cascade model.
It is clear that every oriC copy must build a complete pre-RC to unwind the DUE. However, potential problems arise in rapidly growing cells or cells containing minichromosomes. These cells, containing multiple copies of oriC (see Fig. 1) (101), initiate chromosome replication at the same mass per chromosomal origin (57), and all origins initiate synchronously during the cell cycle (148, 225). To account for the fact that the cell does not need to make more DnaA to initiate more copies of oriC, the “initiation cascade” model was proposed, in which newly triggered origins are able to release active DnaA that becomes available for rapid pre-RC assembly at remaining “uninitiated” oriC copies (94, 153). For an initiation cascade to function properly, there must be mechanisms to prevent pre-RC from being reassembled at origins that have recently initiated. These mechanisms are discussed in the following sections.
To trigger DNA synthesis only once from each oriC copy per cell cycle, DnaA interacting with each newly initiated oriC must be removed and prevented from reassembling pre-RC for one cell generation. Once all oriC copies initiate, any remaining free DnaA-ATP must become unavailable and ultimately inactivated to ensure that new synthesis of active initiator is required for the next round, so that initiation frequency remains coupled to growth rate. Additionally, the mechanisms that disassemble and inactivate the orisome must also allow each oriC copy to be reset by re-forming the bacterial ORC, so that the next round of pre-RC assembly can proceed in the correct order. All of these activities are accomplished by complementary and cell cycle-specific mechanisms, described below and shown in Fig. 5B.
Removal of oriC-bound DnaA.
In vivo DMS footprinting studies have demonstrated that the pre-RC is a transient complex, with oriC being fully occupied by DnaA-ATP for only a small fraction of the cell cycle (181). However, the mechanism by which DnaA is removed from oriC is not known. It is likely that either translocation of DNA helicase through oriC, or the assembly and movement of the replisome would generate enough force to displace DnaA. If displaced by DNA helicase, then DnaA-ATP may leave oriC while remaining in the active form for use at other unfired origins in the initiation cascade. If the replisome were to remove the DnaA, then it is possible that the Hda bound to the sliding clamp could stimulate hydrolysis of DnaA-ATP, which might preclude its reuse at another origin. However, the rapid and multiple reinitiation events that take place in SeqA mutants (158) suggest that, if the replisome is the factor that removes DnaA from oriC, RIDA activity may not be sufficient to inactivate enough DnaA-ATP to prevent re-formation of the pre-RC.
oriC sequestration and resetting of ORC.
Replication of oriC DNA results in hemimethylation of the 11 GATC residues located in the origin, followed by binding of SeqA to the hemimethylated sites (181, 227, 228). These SeqA-oriC interactions mark the start of the sequestration period, where oriC remains refractory to reinitiation and blocked from remethylation by Dam methyltransferase for approximately one-third of the cell cycle (38, 158). During sequestration, bound SeqA prevents reassembly of pre-RC at newly replicated origins by blocking DnaA binding sites that contain or overlap a GATC (181, 240). These sites include the low-affinity sites in the body of oriC (R5M, τ2, I2, and I3), as well as the ATP-DnaA/S-M sites in the 13-mer region (see Fig. 3). In mutants lacking SeqA, loss of this blocking activity allows rapid rebinding of DnaA to the lower-affinity sites and re-formation of the pre-RC, (181), leading to inappropriate reinitiation and overreplication (158).
The high-affinity sites R1, R2, and R4 do not contain or overlap a GATC sequence, and SeqA does not block DnaA from binding to these sites (181). Therefore, early in the sequestration period, DnaA rebinds the high-affinity sites in oriC, resetting the bacterial ORC so that the origin is ready to begin the process of pre-RC assembly to ensure that chromosome replication will start at the correct time in the next cell cycle.
Does the bacterial ORC need to be recharged to the ATP form?
There is no known mechanism that would ensure that the DnaA that rebinds to high-affinity sites during sequestration is in the ATP form, since R1, R2, and R4 have equal affinity for both DnaA-ATP and DnaA-ADP, and both forms are likely to be available immediately after initiation. However, if all origins are to build orisomes identically each and every cell cycle, it seems logical to propose that they should all start from the same ground state, although there is little evidence to suggest or refute that this is, indeed, the case. If the transition from ORC to pre-RC requires that the ORC contain only DnaA-ATP, then special mechanisms would be needed to exchange any DnaA-ADP or replace bound DnaA-ADP with DnaA-ATP. Since hemimethylated oriC DNA interacts with the membrane fraction (191), there is an opportunity during the sequestration period for any oriC-bound DnaA-ADP to interact with acidic phospholipids that stimulate the nucleotide exchange to recharge DnaA-ATP (41, 49). Alternatively, DnaA is detected near the membrane surface (27, 179), and it is possible that the membrane provides a localized reservoir of DnaA-ATP that could replace any DnaA-ADP bound to newly replicated oriC.
Decreasing the cellular levels of available DnaA-ATP.
SeqA blocks re-formation of the pre-RC for approximately one-third of the cell cycle (38). During this time, several other mechanisms are active, working to reduce the cellular levels of available DnaA-ATP, so that at the end of sequestration, the pre-RC does not re-form prematurely. During sequestration SeqA binds and sequesters the dnaA gene promoter, shutting down transcription (38, 241), and causing a rapid decrease of cellular dnaA mRNA levels, and a halt to new DnaA-ATP synthesis for the duration of the sequestration period. In addition, as each new round of DNA synthesis begins, RIDA becomes active and Hda associated with the replication forks stimulates the intrinsic ATPase activity of bound DnaA-ATP (123). Thus, RIDA, coupled with the lack of new DnaA-ATP synthesis, results in a rapid decrease in cellular DnaA-ATP levels after initiation (140). Mutational studies have demonstrated that both timely dnaA promoter sequestration and Hda/RIDA are necessary to prevent reinitiation (123, 204). Moving the dnaA gene to a location more distant from oriC, thus delaying its shutdown by sequestration, results in asynchronous initiations (204). While this latter study indicates that turning off dnaA expression at the wrong time in the cell cycle perturbs initiation regulation, the result of keeping the dnaA gene on throughout the entire cell cycle is less clear. In studies where DnaA, driven by the inducible plac, was constitutively expressed from a plasmid, changes in dnaA expression levels altered the mass at initiation, but did not change once-per-cycle regulation, unless the DnaA levels were raised to such an extent as to cause overinitiation (10, 154). There is no clear explanation for these conflicting results, but they may indicate that there is a very strict balance between DnaA levels, RIDA, and sequestration in preventing reinitiation events.
As replication forks proceed, DnaA recognition sites on the chromosome are duplicated and become available to titrate any freely available DnaA in the cell (94), including DnaA displaced from oriC that is precluded from rebinding by sequestration. The primary DnaA titration locus, datA is located 450,000 bp rightward of oriC and, based on the fixed rate of replication fork movement, becomes duplicated approximately 8 min after initiation, producing a cell cycle-specific titration activity during the sequestration period. However, it is not clear whether this location is critical, since moving datA further away from oriC has little effect until relocation places it near the terminus region (129).
Regulating the availability of DnaA-ATP during the cell cycle: a proper balance of synthesis, inactivation, and recharging.
Although newly synthesized as DnaA-ATP (218), in exponentially growing E. coli, DnaA-ADP predominates, making up approximately 70% of total DnaA (140). In synchronized cells, the relative levels of DnaA-ATP and DnaA-ADP fluctuate. Near the time of initiation of DNA synthesis, DnaA-ATP comprises 80% of total DnaA. After initiation, there is a rapid decrease in DnaA-ATP levels (to approximately 30% of total DnaA), and then, after sequestration ends and approximately half of the chromosome is replicated, the DnaA-ATP levels begin to rise again as cells approach the time to start the next round of DNA replication (140).
Regulating this dramatic cell cycle-specific fluctuation in DnaA-ATP levels over a wide variety of growth rates demands careful orchestration among different regulatory pathways. As described above, decreased DnaA activity immediately following initiation can be explained by the combination of RIDA and sequestration of the dnaA promoter. After sequestration is completed, new DnaA-ATP synthesis will conflict with RIDA in establishing DnaA-ATP levels, and this conflict will continue throughout the cell cycle, since there are always active replication forks in rapidly growing cells (Fig. 1). However, for a cell to begin the next new round of DNA synthesis, at some cell cycle time the production of DnaA-ATP must outpace inactivation by Hda so that DnaA-ATP levels can reach the threshold needed to trigger the initiation event. Resumption of DnaA-ATP synthesis alone, at least initially, does not appear to be able to raise DnaA-ATP levels in the face of ongoing RIDA, since there is a gap in time between the end of sequestration and the rise in DnaA-ATP levels (140). It has been proposed that reactivation of DnaA-ADP is required to supplement the DnaA-ATP levels during the “gap” (76, 259) using the two independent DnaA-recharging mechanisms described in a previous section. It remains to be determined whether or not the recharging mechanisms operate over specific fractions of time during the cell cycle. For example, the phospholipid-based exchange mechanism could be restricted to a particular time since phospholipid synthesis and assembly of membrane domains is cell cycle specific (171, 198). Sequestration may also provide a specific time period in which DnaA-ADP bound to strong sites in hemimethylated oriC associates with the membrane (191) and helps mediate phospholipid-stimulated nucleotide exchange.
The two DARS loci are mapped approximately equidistant from oriC, located on opposite sides of the replicore (76). Both are replicated approximately three quarters of the way through the C period, and since their duplication is later than the time that DnaA-ATP levels begin to rise, it seems improbable that the increase in DARS copy number plays a significant timing role. DARS could, however, have a role in cell cycle-specific recharging if the sites were supplied with DnaA-ADP substrate at a specific time during the cell cycle. datA is an obvious “sink” for DnaA-ATP, with the ability to produce a cell cycle-specific increase in DnaA-ADP available to DARS as replication forks move through the titration locus and Hda hydrolyzes the bound DnaA-ATP. It should be noted, however, that RIDA appears to be functional in datA mutant strains (120), suggesting that other chromosomal recognition sites also contribute DnaA-ATP for inactivation and supply DnaA-ADP for DARS-mediated recharging.
It also remains possible that DnaA-ADP is selectively removed from the pool of available free DnaA causing an increase in DnaA-ATP relative to DnaA-ADP. The membrane is the most likely destination for DnaA-ADP; since much of the DnaA in the cells has been shown to be localized near the surface (27, 179) and in cell lysates, approximately half of the cellular DnaA is found in an insoluble fraction associated with phospholipids (221). However, it is not yet known whether any particular nucleotide form of DnaA predominates in the membrane-bound fraction.
The conservation of DnaA suggests that efforts to dissect the mechanisms that control initiation of replication in E. coli will assist our understanding of the process in other eubacteria. However, there are surprisingly large differences in the nucleotide sequences that constitute eubacterial replication origins, with a wide range of numbers and placements of DnaA recognition sites (81). These differences suggest that eubacterial ORC and pre-RC are likely to be assembled and regulated by mechanisms that are most appropriate for the lifestyle of the bacteria. For example, large numbers of DnaA recognition sites within oriC may be an advantage for slow-growing bacteria (for examples, see reference 269), that initiate exclusively from one copy of oriC and have extended periods of the cell cycle devoid of ongoing DNA replication. Adding oriC recognition sites to titrate DnaA may enhance the sensitivity of the initiation timing mechanism during slow growth, in particular, if extremely low rates of DnaA synthesis are difficult to achieve. In the case of enteric bacteria like E. coli and Salmonella, relatively low numbers of DnaA recognition sites within oriC may allow for additional modulators to extend the dynamic range of initiator complex assembly to accommodate both fast (multifork replication) and slow growth.
We thank our many colleagues for helpful discussions in the preparation of this review. Work from our laboratories (A.C.L. and J.E.G.) was supported by Public Health Service Grant GM054042.
References
1. Abe, Y., T. Jo, Y. Matsuda, C. Matsunaga, T. Katayama, and T. Ueda. 2007. Structure and function of DnaA N-terminal domains: specific sites and mechanisms in inter-DnaA interaction and in DnaB helicase loading on oriC. J. Biol. Chem.282:17816–17827.[PubMed] [CrossRef]
2. Almiron, M., A. J. Link, D. Furlong, and R. Kolter. 1992. A novel DNA-binding protein with regulatory and protective roles in starved Escherichia coli. Genes Dev. 6:2646–2654.[PubMed] [CrossRef]
3. Aranovich, A., A. H. Parola, and I. Fishov. 2007. The reactivation of DnaA(L366K) requires less acidic phospholipids supporting their role in the initiation of chromosome replication in Escherichia coli. FEBS Lett. 581:4439–4442.[PubMed] [CrossRef]
4. Armengod, M. E., M. Garcia-Sogo, and E. Lambies. 1988. Transcriptional organization of the dnaN and recF genes of Escherichia coli K-12. J. Biol. Chem.263:12109–12114.[PubMed]
5. Asada, K., K. Sugimoto, A. Oka, M. Takanami, and Y. Hirota. 1982. Structure of replication origin of the Escherichia coli K-12 chromosome: the presence of spacer sequences in the ori region carrying information for autonomous replication. Nucleic Acids Res.10:3745–3754.[PubMed] [CrossRef]
6. Asai, T., C. P. Chen, T. Nagata, M. Takanami, and M. Imai. 1992. Transcription in vivo within the replication origin of the Escherichia coli chromosome: a mechanism for activating initiation of replication. Mol. Gen. Genet.231:169–178.[PubMed]
7. Asai, T., M. Takanami, and M. Imai. 1990. The AT richness and gid transcription determine the left border of the replication origin of the E. coli chromosome. EMBO J.9:4065–4072.[PubMed]
8. Atlung, T. 1984. Allele-specific suppression of dnaA(Ts) mutations by rpoB mutations in Escherichia coli. Mol. Gen. Genet.197:125–128.[PubMed] [CrossRef]
9. Atlung, T., E. S. Clausen, and F. G. Hansen. 1985. Autoregulation of the dnaA gene of Escherichia coli K12. Mol. Gen. Genet.200:442–450.[PubMed] [CrossRef]
10. Atlung, T., and F. G. Hansen. 1993. Three distinct chromosome replication states are induced by increasing concentrations of DnaA protein in Escherichia coli. J. Bacteriol.175:6537–6545.[PubMed]
11. Atlung, T., A. Lobner-Olesen, and F. G. Hansen. 1987. Overproduction of DnaA protein stimulates initiation of chromosome and minichromosome replication in Escherichia coli. Mol. Gen. Genet.206:51–59.[PubMed] [CrossRef]
12. Bach, T., and K. Skarstad. 2005. An oriC-like distribution of GATC sites mediates full sequestration of non-origin sequences in Escherichia coli. J. Mol. Biol.350:7–11.[PubMed] [CrossRef]
13. Bach, T., and K. Skarstad. 2004. Re-replication from non-sequesterable origins generates three-nucleoid cells which divide asymmetrically. Mol. Microbiol.51:1589–1600.[PubMed] [CrossRef]
14. Bahloul, A., F. Boubrik, and J. Rouviere-Yaniv. 2001. Roles of Escherichia coli histone-like protein HU in DNA replication: HU-beta suppresses the thermosensitivity of dnaA46ts. Biochimie83:219–229.[PubMed] [CrossRef]
15. Bailey, S., W. K. Eliason, and T. A. Steitz. 2007. Structure of hexameric DnaB helicase and its complex with a domain of DnaG primase. Science318:459–463.[PubMed] [CrossRef]
16. Baker, T. A., B. E. Funnell, and A. Kornberg. 1987. Helicase action of dnaB protein during replication from the Escherichia coli chromosomal origin in vitro. J. Biol. Chem.262:6877–6885.[PubMed]
17. Baker, T. A., and A. Kornberg. 1988. Transcriptional activation of initiation of replication from the E. coli chromosomal origin: an RNA-DNA hybrid near oriC. Cell55:113–123.[PubMed] [CrossRef]
18. Ball, C. A., R. Osuna, K. C. Ferguson, and R. C. Johnson. 1992. Dramatic changes in Fis levels upon nutrient upshift in Escherichia coli. J. Bacteriol.174:8043–8056.[PubMed]
19. Barcena, M., T. Ruiz, L. E. Donate, S. E. Brown, N. E. Dixon, M. Radermacher, and J. M. Carazo. 2001. The DnaB.DnaC complex: a structure based on dimers assembled around an occluded channel. EMBO J.20:1462–1468.[PubMed] [CrossRef]
20. Bates, D. B., T. Asai, Y. Cao, M. W. Chambers, G. W. Cadwell, E. Boye, and T. Kogoma. 1995. The DnaA box R4 in the minimal oriC is dispensable for initiation of Escherichia coli chromosome replication. Nucleic Acids Res. 23:3119–3125.[PubMed] [CrossRef]
21. Birch, O. M., K. S. Hewitson, M. Fuhrmann, K. Burgdorf, J. E. Baldwin, P. L. Roach, and N. M. Shaw. 2000. MioC is an FMN-binding protein that is essential for Escherichia coli biotin synthase activity in vitro. J. Biol. Chem.275:32277–32280.[PubMed] [CrossRef]
22. Bird, R., and K. G. Lark. 1968. Initiation and termination of DNA replication after amino acid starvation of E. coli 15T. Cold Spring Harbor Symp. Quant. Biol.33:799–808.[PubMed]
23. Bird, R. E., J. Louarn, J. Martuscelli, and L. Caro. 1972. Origin and sequence of chromosome replication in Escherichia coli. J. Mol. Biol.70:549–566.[PubMed] [CrossRef]
24. Blaesing, F., C. Weigel, M. Welzeck, and W. Messer. 2000. Analysis of the DNA-binding domain of Escherichia coli DnaA protein. Mol. Microbiol.36:557–569.[PubMed] [CrossRef]
25. Blattner, F. R., G. Plunkett III, C. A. Bloch, N. T. Perna, V. Burland, M. Riley, J. Collado-Vides, J. D. Glasner, C. K. Rode, G. F. Mayhew, J. Gregor, N. W. Davis, H. A. Kirkpatrick, M. A. Goeden, D. J. Rose, B. Mau, and Y. Shao. 1997. The complete genome sequence of Escherichia coli K-12. Science277:1453–1474.[PubMed] [CrossRef]
26. Boeneman, K., and E. Crooke. 2005. Chromosomal replication and the cell membrane. Curr. Opin. Microbiol.8:143–148.[PubMed] [CrossRef]
27. Boeneman, K., S. Fossum, Y. Yang, N. Fingland, K. Skarstad, and E. Crooke. 2009. Escherichia coli DnaA forms helical structures along the longitudinal cell axis distinct from MreB filaments. Mol. Microbiol.72:645–657. [CrossRef]
28. Bogan, J. A., and C. E. Helmstetter. 1997. DNA sequestration and transcription in the oriC region of Escherichia coli. Mol. Microbiol.26:889–896.[PubMed] [CrossRef]
29. Bogan, J. A., and C. E. Helmstetter. 1996. mioC transcription, initiation of replication, and the eclipse in Escherichia coli. J. Bacteriol.178:3201–3206.[PubMed]
30. Bonnefoy, E., and J. Rouviere-Yaniv. 1992. HU, the major histone-like protein of E. coli, modulates the binding of IHF to oriC. EMBO J.11:4489–4496.[PubMed]
31. Boye, E., A. Lyngstadaas, A. Lobner-Olesen, K. Skarstad, and S. Wold. 1993. Regulation of DNA replication in Escherichia coli, p. 15–26. In E. Fanning, R. Knippers, and E.-L. Winnacker (ed.), DNA Replication and the Cell Cycle. Springer, Berlin, Germany.
32. Bramhill, D., and A. Kornberg. 1988. Duplex opening by dnaA protein at novel sequences in initiation of replication at the origin of the E. coli chromosome. Cell52:743–755.[PubMed] [CrossRef]
33. Braun, R. E., K. O'Day, and A. Wright. 1985. Autoregulation of the DNA replication gene dnaA in E. coli K-12. Cell40:159–169.[PubMed] [CrossRef]
34. Braun, R. E., and A. Wright. 1986. DNA methylation differentially enhances the expression of one of the two E. coli dnaA promoters in vivo and in vitro. Mol. Gen. Genet. 202:246–250.[PubMed] [CrossRef]
35. Brendler, T., and S. Austin. 1999. Binding of SeqA protein to DNA requires interaction between two or more complexes bound to separate hemimethylated GATC sequences. EMBO J.18:2304–2310.[PubMed] [CrossRef]
36. 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.6:736–741.[PubMed] [CrossRef]
37. Camara, J. E., K. Skarstad, and E. Crooke. 2003. Controlled initiation of chromosomal replication in Escherichia coli requires functional Hda protein. J. Bacteriol.185:3244–3248.[PubMed] [CrossRef]
38. Campbell, J. L., and N. Kleckner. 1990. E. coli oriC and the dnaA gene promoter are sequestered from dam methyltransferase following the passage of the chromosomal replication fork. Cell62:967–979.[PubMed] [CrossRef]
39. Carl, P. L. 1970. Escherichia coli mutants with temperature-sensitive synthesis of DNA. Mol. Gen. Genet.109:107–122.[PubMed] [CrossRef]
40. Cassler, M. R., J. E. Grimwade, and A. C. Leonard. 1995. Cell cycle-specific changes in nucleoprotein complexes at a chromosomal replication origin. EMBO J.14:5833–5841.[PubMed]
41. Castuma, C. E., E. Crooke, and A. Kornberg. 1993. Fluid membranes with acidic domains activate DnaA, the initiator protein of replication in Escherichia coli. J. Biol. Chem.268:24665–24668.[PubMed]
42. Celis, R. T. 1999. Repression and activation of arginine transport genes in Escherichia coli K 12 by the ArgP protein. J. Mol. Biol.294:1087–1095.[PubMed] [CrossRef]
43. Chiaramello, A. E., and J. W. Zyskind. 1989. Expression of Escherichia coli dnaA and mioC genes as a function of growth rate. J. Bacteriol.171:4272–4280.[PubMed]
44. Chodavarapu, S., M. M. Felczak, J. R. Yaniv, and J. M. Kaguni. 2008. Escherichia coli DnaA interacts with HU in initiation at the E. coli replication origin. Mol. Microbiol.67:781–792.[PubMed] [CrossRef]
45. Chodavarapu, S., R. Gomez, M. Vicente, and J. M. Kaguni. 2008. Escherichia coli Dps interacts with DnaA protein to impede initiation: a model of adaptive mutation. Mol. Microbiol.67:1331–1346.[PubMed] [CrossRef]
46. Clarey, M. G., J. P. Erzberger, P. Grob, A. E. Leschziner, J. M. Berger, E. Nogales, and M. Botchan. 2006. Nucleotide-dependent conformational changes in the DnaA-like core of the origin recognition complex. Nat. Struct. Mol. Biol. 13:684–690.[PubMed] [CrossRef]
47. Cooper, S., and C. E. Helmstetter. 1968. Chromosome replication and the division cycle of Escherichia coli B/r. J. Mol. Biol.31:519–540.[PubMed] [CrossRef]
48. Crooke, E. 2001. Escherichia coli DnaA protein—phospholipid interactions: in vitro and in vivo. Biochimie83:19–23.[PubMed] [CrossRef]
49. Crooke, E., C. E. Castuma, and A. Kornberg. 1992. The chromosome origin of Escherichia coli stabilizes DnaA protein during rejuvenation by phospholipids. J. Biol. Chem.267:16779–16782.[PubMed]
50. Crooke, E., R. Thresher, D. S. Hwang, J. Griffith, and A. Kornberg. 1993. Replicatively active complexes of DnaA protein and the Escherichia coli chromosomal origin observed in the electron microscope. J. Mol. Biol.233:16–24.[PubMed] [CrossRef]
51. d'Alencon, E., A. Taghbalout, R. Kern, and M. Kohiyama. 1999. Replication cycle dependent association of SeqA to the outer membrane fraction of E. coli. Biochimie81:841–846.[PubMed] [CrossRef]
52. Dasgupta, S., and A. Lobner-Olesen. 2004. Host controlled plasmid replication: Escherichia coli minichromosomes. Plasmid52:151–168.[PubMed] [CrossRef]
53. Davey, M. J., L. Fang, P. McInerney, R. E. Georgescu, and M. O'Donnell. 2002. The DnaC helicase loader is a dual ATP/ADP switch protein. EMBO J.21:3148–3159.[PubMed] [CrossRef]
54. Davey, M. J., and M. O'Donnell. 2003. Replicative helicase loaders: ring breakers and ring makers. Curr. Biol. 13:R594–R596.[PubMed] [CrossRef]
55. Diffley, J. F. 2004. Regulation of early events in chromosome replication. Curr. Biol.14:R778–R786.[PubMed] [CrossRef]
56. Dixon, N. E., and A. Kornberg. 1984. Protein HU in the enzymatic replication of the chromosomal origin of Escherichia coli. Proc. Natl. Acad. Sci. USA81:424–428.[PubMed] [CrossRef]
57. Donachie, W. D. 1968. Relationship between cell size and time of initiation of DNA replication. Nature219:1077–1079.[PubMed] [CrossRef]
58. Donate, L. E., O. Llorca, M. Barcena, S. E. Brown, N. E. Dixon, and J. M. Carazo. 2000. pH-controlled quaternary states of hexameric DnaB helicase. J. Mol. Biol.303:383–393.[PubMed] [CrossRef]
59. Drlica, K., and J. Rouviere-Yaniv. 1987. Histonelike proteins of bacteria. Microbiol. Rev.51:301–319.[PubMed]
60. Duderstadt, K. E., and J. M. Berger. 2008. AAA+ ATPases in the initiation of DNA replication. Crit. Rev. Biochem. Mol. Biol.43:163–187.[PubMed] [CrossRef]
61. Echols, H. 1986. Multiple DNA-protein interactions governing high-precision DNA transactions. Science233:1050–1056.[PubMed] [CrossRef]
62. Eliasson, A., and K. Nordstrom. 1997. Replication of minichromosomes in a host in which chromosome replication is random. Mol. Microbiol.23:1215–1220.[PubMed] [CrossRef]
63. Ellenberger, T., and A. Landy. 1997. A good turn for DNA: the structure of integration host factor bound to DNA. Structure5:153–157.[PubMed] [CrossRef]
64. 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.13:676–683.[PubMed] [CrossRef]
65. Erzberger, J. P., M. M. Pirruccello, and J. M. Berger. 2002. The structure of bacterial DnaA: implications for general mechanisms underlying DNA replication initiation. EMBO J.21:4763–4773.[PubMed] [CrossRef]
66. Fang, L., M. J. Davey, and M. O'Donnell. 1999. Replisome assembly at oriC, the replication origin of E. coli, reveals an explanation for initiation sites outside an origin. Mol. Cell4:541–553.[PubMed] [CrossRef]
67. 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. 279:51156–51162.[PubMed] [CrossRef]
68. Felczak, M. M., L. A. Simmons, and J. M. Kaguni. 2005. An essential tryptophan of Escherichia coli DnaA protein functions in oligomerization at the E. coli replication origin. J. Biol. Chem.280:24627–24633.[PubMed] [CrossRef]
69. Filutowicz, M., and J. Roll. 1990. The requirement of IHF protein for extrachromosomal replication of the Escherichia coli oriC in a mutant deficient in DNA polymerase I activity. New Biol.2:818–827.[PubMed]
70. Filutowicz, M., W. Ross, J. Wild, and R. L. Gourse. 1992. Involvement of Fis protein in replication of the Escherichia coli chromosome. J. Bacteriol.174:398–407.[PubMed]
71. Finkel, S. E., and R. C. Johnson. 1992. The Fis protein: it's not just for DNA inversion anymore. Mol. Microbiol.6:3257–3265.[PubMed] [CrossRef]
72. Flatten, I., and K. Skarstad. 2009. DnaA protein interacts with RNA polymerase and partially protects it from the effect of rifampicin. Mol. Microbiol.71:1018–1030.[PubMed] [CrossRef]
73. Froelich, J. M., T. K. Phuong, and J. W. Zyskind. 1996. Fis binding in the dnaA operon promoter region. J. Bacteriol.178:6006–6012.[PubMed]
74. 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.31:2077–2086.[PubMed] [CrossRef]
75. Fujimitsu, K., and T. Katayama. 2004. Reactivation of DnaA by DNA sequence-specific nucleotide exchange in vitro. Biochem. Biophys. Res. Commun.322:411–419.[PubMed] [CrossRef]
76. Fujimitsu, K., T. Senriuchi, and T. Katayama. 2009. Specific genomic sequences of E. coli promote replicational initiation by directly reactivating ADP-DnaA. Genes Dev.23:1221–1233.[PubMed] [CrossRef]
77. Fujimitsu, K., M. Su'etsugu, Y. Yamaguchi, K. Mazda, N. Fu, H. Kawakami, and T. Katayama. 2008. Modes of overinitiation, dnaA gene expression, and inhibition of cell division in a novel cold-sensitive hda mutant of Escherichia coli. J. Bacteriol.190:5368–5381.[PubMed] [CrossRef]
78. Fujisawa, T., and A. Eisenstark. 1973. Bi-directional chromosomal replication in Salmonella typhimurium. J. Bacteriol.115:168–176.[PubMed]
79. 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. Cell38:889–900.[PubMed] [CrossRef]
80. Funnell, B. E., T. A. Baker, and A. Kornberg. 1987. In vitro assembly of a prepriming complex at the origin of the Escherichia coli chromosome. J. Biol. Chem.262:10327–10334.[PubMed]
81. Gao, F., and C. T. Zhang. 2007. DoriC: a database of oriC regions in bacterial genomes. Bioinformatics23:1866–1867.[PubMed]
82. Garner, J., and E. Crooke. 1996. Membrane regulation of the chromosomal replication activity of E. coli DnaA requires a discrete site on the protein. EMBO J.15:3477–3485.[PubMed]
83. Geier, G. E., and P. Modrich. 1979. Recognition sequence of the dam methylase of Escherichia coli K12 and mode of cleavage of Dpn I endonuclease. J. Biol. Chem.254:1408–1413.[PubMed]
84. Giaever, G. N., L. Snyder, and J. C. Wang. 1988. DNA supercoiling in vivo. Biophys. Chem.29:7–15.[PubMed] [CrossRef]
85. Gille, H., J. B. Egan, A. Roth, and W. Messer. 1991. The FIS protein binds and bends the origin of chromosomal DNA replication, oriC, of Escherichia coli. Nucleic Acids Res.19:4167–4172.[PubMed] [CrossRef]
86. Giraldo, R. 2003. Common domains in the initiators of DNA replication in Bacteria, Archaea and Eukarya: combined structural, functional and phylogenetic perspectives. FEMS Microbiol. Rev.26:533–554.[PubMed] [CrossRef]
87. Grimwade, J. E., V. T. Ryan, and A. C. Leonard. 2000. IHF redistributes bound initiator protein, DnaA, on supercoiled oriC of Escherichia coli. Mol. Microbiol.35:835–844.[PubMed] [CrossRef]
88. Grimwade, J. E., J. J. Torgue, K. C. McGarry, T. Rozgaja, S. T. Enloe, and A. C. Leonard. 2007. Mutational analysis reveals Escherichia coli oriC interacts with both DnaA-ATP and DnaA-ADP during pre-RC assembly. Mol. Microbiol.66:428–439.[PubMed] [CrossRef]
89. Guarne, A., T. Brendler, Q. Zhao, R. Ghirlando, S. Austin, and W. Yang. 2005. Crystal structure of a SeqA-N filament: implications for DNA replication and chromosome organization. EMBO J.24:1502–1511.[PubMed] [CrossRef]
90. Han, J. S., J. Y. Park, Y. S. Lee, B. Thony, and D. S. Hwang. 1999. PhoB-dependent transcriptional activation of the iciA gene during starvation for phosphate in Escherichia coli. Mol. Gen. Genet.262:448–452.[PubMed] [CrossRef]
91. Hanna, M. H., and P. L. Carl. 1975. Reinitiation of deoxyribonucleic acid synthesis by deoxyribonucleic acid initiation mutants of Escherichia coli: role of ribonucleic acid synthesis, protein synthesis, and cell division. J. Bacteriol.121:219–226.[PubMed]
92. Hansen, E. B., F. G. Hansen, and K. von Meyenburg. 1982. The nucleotide sequence of the dnaA gene and the first part of the dnaN gene of Escherichia coli K-12. Nucleic Acids Res.10:7373–7385.[PubMed] [CrossRef]
93. Hansen, F. G., T. Atlung, R. E. Braun, A. Wright, P. Hughes, and M. Kohiyama. 1991. Initiator (DnaA) protein concentration as a function of growth rate in Escherichia coli and Salmonella typhimurium. J. Bacteriol.173:5194–5199.[PubMed]
94. Hansen, F. G., B. B. Christensen, and T. Atlung. 1991. The initiator titration model: computer simulation of chromosome and minichromosome control. Res. Microbiol.142:161–167.[PubMed] [CrossRef]
95. Hansen, F. G., B. B. Christensen, and T. Atlung. 2007. Sequence characteristics required for cooperative binding and efficient in vivo titration of the replication initiator protein DnaA in E. coli. J. Mol. Biol.367:942–952.[PubMed] [CrossRef]
96. Hansen, F. G., E. B. Hansen, and T. Atlung. 1982. The nucleotide sequence of the dnaA gene promoter and of the adjacent rpmH gene, coding for the ribosomal protein L34, of Escherichia coli. EMBO J.1:1043–1048.[PubMed]
97. Hansen, F. G., S. Koefoed, and T. Atlung. 1992. Cloning and nucleotide sequence determination of twelve mutant dnaA genes of Escherichia coli. Mol. Gen. Genet.234:14–21.[PubMed]
98. Hansen, F. G., S. Koefoed, L. Sorensen, and T. Atlung. 1987. Titration of DnaA protein by oriC DnaA-boxes increases dnaA gene expression in Escherichia coli. EMBO J.6:255–258.[PubMed]
99. Harding, N. E., J. M. Cleary, D. W. Smith, J. J. Michon, W. S. Brusilow, and J. W. Zyskind. 1982. Chromosomal replication origins (oriC) of Enterobacter aerogenes and Klebsiella pneumoniae are functional in Escherichia coli. J. Bacteriol.152:983–993.[PubMed]
100. Helmstetter, C., S. Cooper, O. Pierucci, and E. Revelas. 1968. On the bacterial life sequence. Cold Spring Harbor Symp. Quant. Biol.33:809–822.[PubMed]
101. Helmstetter, C. E. 1996. Timing of synthetic activities in the cell cycle, p. 1627–1639. In F. C. Neidhardt, R. Curtis, III, J. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: Cellular and Molecular Biology. ASM Press, Washington, DC.
102. Helmstetter, C. E., C. Eenhuis, P. Theisen, J. Grimwade, and A. C. Leonard. 1992. Improved bacterial baby machine: application to Escherichia coli K-12. J. Bacteriol.174:3445–3449.[PubMed]
103. Helmstetter, C. E., and A. C. Leonard. 1987. Coordinate initiation of chromosome and minichromosome replication in Escherichia coli. J. Bacteriol.169:3489–3494.[PubMed]
104. Hiasa, H., and K. J. Marians. 1994. Fis cannot support oriC DNA replication in vitro. J. Biol. Chem.269:24999–25003.[PubMed]
105. Hiraga, S. 1976. Novel F prime factors able to replicate in Escherichia coli Hfr strains. Proc. Natl. Acad. Sci. USA73:198–202.[PubMed] [CrossRef]
106. Hiraga, S., C. Ichinose, H. Niki, and M. Yamazoe. 1998. Cell cycle-dependent duplication and bidirectional migration of SeqA-associated DNA-protein complexes in E. coli. Mol. Cell1:381–387.[PubMed] [CrossRef]
107. Hodges-Garcia, Y., P. J. Hagerman, and D. E. Pettijohn. 1989. DNA ring closure mediated by protein HU. J. Biol. Chem.264:14621–14623.[PubMed]
108. Holz, A., C. Schaefer, H. Gille, W. R. Jueterbock, and W. Messer. 1992. Mutations in the DnaA binding sites of the replication origin of Escherichia coli. Mol. Gen. Genet.233:81–88.[PubMed] [CrossRef]
109. Hsu, J., D. Bramhill, and C. M. Thompson. 1994. Open complex formation by DnaA initiation protein at the Escherichia coli chromosomal origin requires the 13-mers precisely spaced relative to the 9-mers. Mol. Microbiol.11:903–911.[PubMed] [CrossRef]
110. Hwang, D. S., and A. Kornberg. 1992. Opening of the replication origin of Escherichia coli by DnaA protein with protein HU or IHF. J. Biol. Chem.267:23083–23086.[PubMed]
111. Hwang, D. S., and A. Kornberg. 1992. Opposed actions of regulatory proteins, DnaA and IciA, in opening the replication origin of Escherichia coli. J. Biol. Chem.267:23087–23091.[PubMed]
112. Hwang, D. S., B. Thony, and A. Kornberg. 1992. IciA protein, a specific inhibitor of initiation of Escherichia coli chromosomal replication. J. Biol. Chem.267:2209–2213.[PubMed]
113. Ishida, T., N. Akimitsu, T. Kashioka, M. Hatano, T. Kubota, Y. Ogata, K. Sekimizu, and T. Katayama. 2004. DiaA, a novel DnaA-binding protein, ensures the timely initiation of Escherichia coli chromosome replication. J. Biol. Chem.279:45546–45555.[PubMed] [CrossRef]
114. Jacob, F., S. Brenner, and F. Cuzin. 1963. On the regulation of DNA replication in bacteria. Cold Spring Harbor Symp. Quant. Biol.28:239–348.
115. Junker, D. E., Jr., L. A. Rokeach, D. Ganea, A. Chiaramello, and J. W. Zyskind. 1986. Transcription termination within the Escherichia coli origin of DNA replication, oriC. Mol. Gen. Genet.203:101–109.[PubMed] [CrossRef]
116. Kaguni, J. M. 2006. DnaA: controlling the initiation of bacterial DNA replication and more. Annu. Rev. Microbiol.60:351–375.[PubMed] [CrossRef]
117. Katayama, T. 2001. Feedback controls restrain the initiation of Escherichia coli chromosomal replication. Mol. Microbiol.41:9–17.[PubMed] [CrossRef]
118. Katayama, T. 2008. Roles for the AAA+ motifs of DnaA in the initiation of DNA replication. Biochem. Soc. Trans. 36:78–82.[PubMed] [CrossRef]
119. Katayama, T., and E. Crooke. 1995. DnaA protein is sensitive to a soluble factor and is specifically inactivated for initiation of in vitro replication of the Escherichia coli minichromosome. J. Biol. Chem.270:9265–9271.[PubMed] [CrossRef]
120. Katayama, T., K. Fujimitsu, and T. Ogawa. 2001. Multiple pathways regulating DnaA function in Escherichia coli: distinct roles for DnaA titration by the datA locus and the regulatory inactivation of DnaA. Biochimie83:13–17.[PubMed] [CrossRef]
121. Katayama, T., T. Kubota, K. Kurokawa, E. Crooke, and K. Sekimizu. 1998. The initiator function of DnaA protein is negatively regulated by the sliding clamp of the E. coli chromosomal replicase. Cell94:61–71.[PubMed] [CrossRef]
122. Kato, J. 2005. Regulatory network of the initiation of chromosomal replication in Escherichia coli. Crit. Rev. Biochem. Mol. Biol.40:331–342.[PubMed] [CrossRef]
123. Kato, J., and T. Katayama. 2001. Hda, a novel DnaA-related protein, regulates the replication cycle in Escherichia coli. EMBO J.20:4253–4262.[PubMed] [CrossRef]
124. 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.280:27420–27430.[PubMed] [CrossRef]
125. Kawakami, H., S. Ozaki, S. Suzuki, K. Nakamura, T. Senriuchi, M. Su'etsugu, K. Fujimitsu, and T. Katayama. 2006. The exceptionally tight affinity of DnaA for ATP/ADP requires a unique aspartic acid residue in the AAA+ sensor 1 motif. Mol. Microbiol.62:1310–1324.[PubMed] [CrossRef]
126. Kawakami, H., M. Su'etsugu, and T. Katayama. 2006. An isolated Hda-clamp complex is functional in the regulatory inactivation of DnaA and DNA replication. J. Struct. Biol.156:220–229.[PubMed]
127. Kellenberger-Gujer, G., A. J. Podhajska, and L. Caro. 1978. A cold sensitive dnaA mutant of E. coli which overinitiates chromosome replication at low temperature. Mol. Gen. Genet.162:9–16.[PubMed] [CrossRef]
128. Keyamura, K., N. Fujikawa, T. Ishida, S. Ozaki, M. Su'etsugu, K. Fujimitsu, W. Kagawa, S. Yokoyama, H. Kurumizaka, and T. Katayama. 2007. The interaction of DiaA and DnaA regulates the replication cycle in E. coli by directly promoting ATP DnaA-specific initiation complexes. Genes Dev.21:2083–2099.[PubMed] [CrossRef]
129. Kitagawa, R., T. Ozaki, S. Moriya, and T. Ogawa. 1998. Negative control of replication initiation by a novel chromosomal locus exhibiting exceptional affinity for Escherichia coli DnaA protein. Genes Dev.12:3032–3043.[PubMed] [CrossRef]
130. Kobori, J. A., and A. Kornberg. 1982. The Escherichia coli dnaC gene product. II. Purification, physical properties, and role in replication. J. Biol. Chem.257:13763–13769.[PubMed]
131. Kohara, Y., N. Tohdoh, X. W. Jiang, and T. Okazaki. 1985. The distribution and properties of RNA primed initiation sites of DNA synthesis at the replication origin of Escherichia coli chromosome. Nucleic Acids Res.13:6847–6866.[PubMed] [CrossRef]
132. Kohiyama, M. 1968. DNA synthesis in temperature sensitive mutants of Escherichia coli. Cold Spring Harbor Symp. Quant. Biol.33:317–324.[PubMed]
133. Kohiyama, M., D. Cousin, A. Ryter, and F. Jacob. 1966. [Thermosensitive mutants of Escherichia coli K 12. I. Isolation and rapid characterization]. Ann. Inst. Pasteur (Paris)110:465–486.[PubMed]
134. Koppes, L. J., and K. von Meyenburg. 1987. Nonrandom minichromosome replication in Escherichia coli K-12. J. Bacteriol.169:430–433.[PubMed]
135. Kornberg, A. 1987. Enzyme systems initiating replication at the origin of the Escherichia coli chromosome. J. Cell Sci. Suppl.7:1–13.[PubMed]
136. Kornberg, A., and T. A. Baker. 1992. DNA Replication. W.H. Freeman and Company, New York, NY.
137. 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.8:4335–4344.[PubMed]
138. Kucherer, C., H. Lother, R. Kolling, M. A. Schauzu, and W. Messer. 1986. Regulation of transcription of the chromosomal dnaA gene of Escherichia coli. Mol. Gen. Genet.205:115–121.[PubMed] [CrossRef]
139. Kurokawa, K., T. Mizushima, T. Kubota, T. Tsuchiya, T. Katayama, and K. Sekimizu. 1998. A stimulation factor for hydrolysis of ATP bound to DnaA protein, the initiator of chromosomal DNA replication in Escherichia coli. Biochem. Biophys. Res. Commun.243:90–95.[PubMed] [CrossRef]
140. 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.18:6642–6652.[PubMed] [CrossRef]
141. Langer, U., S. Richter, A. Roth, C. Weigel, and W. Messer. 1996. A comprehensive set of DnaA-box mutations in the replication origin, oriC, of Escherichia coli. Mol. Microbiol.21:301–311.[PubMed] [CrossRef]
142. Lark, K. G., and H. Renger. 1969. Initiation of DNA replication in Escherichia coli 15T-: chronological dissection of three physiological processes required for initiation. J. Mol. Biol.42:221–235.[PubMed] [CrossRef]
143. LeBowitz, J. H., and R. McMacken. 1986. The Escherichia coli dnaB replication protein is a DNA helicase. J. Biol. Chem.261:4738–4748.[PubMed]
144. Lee, Y. S., and D. S. Hwang. 1997. Occlusion of RNA polymerase by oligomerization of DnaA protein over the dnaA promoter of Escherichia coli. J. Biol. Chem.272:83–88.[PubMed]
145. Lee, Y. S., H. Kim, and D. S. Hwang. 1996. Transcriptional activation of the dnaA gene encoding the initiator for oriC replication by IciA protein, an inhibitor of in vitro oriC replication in Escherichia coli. Mol. Microbiol.19:389–396.[PubMed] [CrossRef]
146. Leonard, A. C., and J. E. Grimwade. 2005. Building a bacterial orisome: emergence of new regulatory features for replication origin unwinding. Mol. Microbiol.55:978–985.[PubMed] [CrossRef]
147. Leonard, A. C., and J. E. Grimwade. 2009. Initiating chromosome replication in E. coli: it makes sense to recycle. Genes Dev.23:1145–1150.[PubMed] [CrossRef]
148. Leonard, A. C., and C. E. Helmstetter. 1986. Cell cycle-specific replication of Escherichia coli minichromosomes. Proc. Natl. Acad. Sci. USA83:5101–5105.[PubMed] [CrossRef]
149. Leonard, A. C., P. W. Theisen, and C. E. Helmstetter. 1990. Replication timing and copy number control of oriC plasmids, p. 279–286. In K. Dirlica and M. Riley (ed.), The Bacterial Chromosome. American Society for Microbiology, Washington, DC.
150. Leonard, A. C., W. G. Whitford, and C. E. Helmstetter. 1985. Involvement of DNA superhelicity in minichromosome maintenance in Escherichia coli. J. Bacteriol.161:687–695.[PubMed]
151. Lobner-Olesen, A. 1999. Distribution of minichromosomes in individual Escherichia coli cells: implications for replication control. EMBO J.18:1712–1721.[PubMed] [CrossRef]
152. Lobner-Olesen, A., and E. Boye. 1992. Different effects of mioC transcription on initiation of chromosomal and minichromosomal replication in Escherichia coli. Nucleic Acids Res.20:3029–3036.[PubMed] [CrossRef]
153. Lobner-Olesen, A., F. G. Hansen, K. V. Rasmussen, B. Martin, and P. L. Kuempel. 1994. The initiation cascade for chromosome replication in wild-type and Dam methyltransferase deficient Escherichia coli cells. EMBO J.13:1856–1862.[PubMed]
154. Lobner-Olesen, A., K. Skarstad, F. G. Hansen, K. von Meyenburg, and E. Boye. 1989. The DnaA protein determines the initiation mass of Escherichia coli K-12. Cell57:881–889.[PubMed] [CrossRef]
155. Lobner-Olesen, A., O. Skovgaard, and M. G. Marinus. 2005. Dam methylation: coordinating cellular processes. Curr. Opin. Microbiol.8:154–160.[PubMed] [CrossRef]
156. Lother, H., R. Kolling, C. Kucherer, and M. Schauzu. 1985. dnaA protein-regulated transcription: effects on the in vitro replication of Escherichia coli minichromosomes. EMBO J.4:555–560.[PubMed]
157. Louarn, J., J. P. Bouche, J. Patte, and J. M. Louarn. 1984. Genetic inactivation of topoisomerase I suppresses a defect in initiation of chromosome replication in Escherichia coli. Mol. Gen. Genet.195:170–174.[PubMed] [CrossRef]
158. Lu, M., J. L. Campbell, E. Boye, and N. Kleckner. 1994. SeqA: a negative modulator of replication initiation in E. coli. Cell77:413–426.[PubMed] [CrossRef]
159. Maaloe, O., and P. C. Hanawalt. 1961. Thymine deficiency and the normal DNA replication cycle. I. J. Mol. Biol.3:144–155.[PubMed] [CrossRef]
160. Margulies, C., and J. M. Kaguni. 1996. Ordered and sequential binding of DnaA protein to oriC, the chromosomal origin of Escherichia coli. J. Biol. Chem.271:17035–17040.[PubMed] [CrossRef]
161. Marsh, R. C., and A. Worcel. 1977. A DNA fragment containing the origin of replication of the Escherichia coli chromosome. Proc. Natl. Acad. Sci. USA74:2720–2724.[PubMed] [CrossRef]
162. Masters, M., and P. Broda. 1971. Evidence for the bidirectional replications of the Escherichia coli chromosome. Nat. New Biol.232:137–140.[PubMed] [CrossRef]
163. Matsui, M., A. Oka, M. Takanami, S. Yasuda, and Y. Hirota. 1985. Sites of dnaA protein-binding in the replication origin of the Escherichia coli K-12 chromosome. J. Mol. Biol.184:529–533.[PubMed] [CrossRef]
164. McClelland, M., K. E. Sanderson, J. Spieth, S. W. Clifton, P. Latreille, L. Courtney, S. Porwollik, J. Ali, M. Dante, F. Du, S. Hou, D. Layman, S. Leonard, C. Nguyen, K. Scott, A. Holmes, N. Grewal, E. Mulvaney, E. Ryan, H. Sun, L. Florea, W. Miller, T. Stoneking, M. Nhan, R. Waterston, and R. K. Wilson. 2001. Complete genome sequence of Salmonella enterica serovar Typhimurium LT2. Nature413:852–856.[PubMed] [CrossRef]
165. 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. USA101:2811–2816.[PubMed] [CrossRef]
166. Meijer, M., and W. Messer. 1980. Functional analysis of minichromosome replication: bidirectional and unidirectional replication from the Escherichia coli replication origin, oriC. J. Bacteriol.143:1049–1053.[PubMed]
167. Messer, W. 2002. The bacterial replication initiator DnaA. DnaA and oriC, the bacterial mode to initiate DNA replication. FEMS Microbiol. Rev.26:355–374.[PubMed] [CrossRef]
168. Messer, W., and C. Weigel. 2003. DnaA as a transcription regulator. Methods Enzymol.370:338–349.[PubMed] [CrossRef]
169. Messer, W., and C. Weigel. 1996. Initiation of chromosome replication, p. 1579–1601. In F. C. Neidhardt, R. Curtis, III, J. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: Cellular and Molecular Biology. ASM Press, Washington, DC.
170. Meyer, S., A. Scrima, W. Versees, and A. Wittinghofer. 2008. Crystal structures of the conserved tRNA-modifying enzyme GidA: implications for its interaction with MnmE and substrate. J. Mol. Biol.380:532–547.[PubMed] [CrossRef]
171. Mileykovskaya, E., and W. Dowhan. 2000. Visualization of phospholipid domains in Escherichia coli by using the cardiolipin-specific fluorescent dye 10-N-nonyl acridine orange. J. Bacteriol. 182:1172–1175.[PubMed] [CrossRef]
172. Molt, K. L., V. A. Sutera, Jr., K. K. Moore, and S. T. Lovett. 2009. A role for nonessential domain II of initiator protein, DnaA, in replication control. Genetics183:39–49.[PubMed] [CrossRef]
173. Morigen, E. Boye, K. Skarstad, and A. Lobner-Olesen. 2001. Regulation of chromosomal replication by DnaA protein availability in Escherichia coli: effects of the datA region. Biochim. Biophys. Acta1521:73–80.[PubMed]
174. Morigen, A. Lobner-Olesen, and K. Skarstad. 2003. Titration of the Escherichia coli DnaA protein to excess datA sites causes destabilization of replication forks, delayed replication initiation and delayed cell division. Mol. Microbiol.50:349–362.[PubMed] [CrossRef]
175. Morigen, F. Molina, and K. Skarstad. 2005. Deletion of the datA site does not affect once-per-cell-cycle timing but induces rifampin-resistant replication. J. Bacteriol.187:3913–3920.[PubMed] [CrossRef]
176. Mott, M. L., and J. M. Berger. 2007. DNA replication initiation: mechanisms and regulation in bacteria. Nat. Rev. Microbiol.5:343–354.[PubMed] [CrossRef]
177. Mott, M. L., J. P. Erzberger, M. M. Coons, and J. M. Berger. 2008. Structural synergy and molecular crosstalk between bacterial helicase loaders and replication initiators. Cell135:623–634.[PubMed] [CrossRef]
178. Nair, S., and S. E. Finkel. 2004. Dps protects cells against multiple stresses during stationary phase. J. Bacteriol.186:4192–4198.[PubMed] [CrossRef]
179. Newman, G., and E. Crooke. 2000. DnaA, the initiator of Escherichia coli chromosomal replication, is located at the cell membrane. J. Bacteriol.182:2604–2610.[PubMed] [CrossRef]
180. Nielsen, O., and A. Lobner-Olesen. 2008. Once in a lifetime: strategies for preventing re-replication in prokaryotic and eukaryotic cells. EMBO Rep.9:151–156.[PubMed] [CrossRef]
181. Nievera, C., J. J. Torgue, J. E. Grimwade, and A. C. Leonard. 2006. SeqA blocking of DnaA-oriC interactions ensures staged assembly of the E. coli pre-RC. Mol. Cell24:581–592.[PubMed] [CrossRef]
182. 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: evidence from a mutant DnaA protein defective in regulatory ATP hydrolysis in vitro and in vivo. J. Biol. Chem.277:14986–14995.[PubMed] [CrossRef]
183. Nozaki, N., T. Okazaki, and T. Ogawa. 1988. In vitro transcription of the origin region of replication of the Escherichia coli chromosome. J. Biol. Chem.263:14176–14183.[PubMed]
184. Nozaki, S., and T. Ogawa. 2008. Determination of the minimum domain II size of Escherichia coli DnaA protein essential for cell viability. Microbiology154:3379–3384.[PubMed] [CrossRef]
185. O'Donnell, M., and D. Jeruzalmi. 2006. Helical proteins initiate replication of DNA helices. Nat. Struct. Mol. Biol.13:665–667.[PubMed] [CrossRef]
186. Ogasawara, N., and H. Yoshikawa. 1992. Genes and their organization in the replication origin region of the bacterial chromosome. Mol. Microbiol.6:629–634.[PubMed] [CrossRef]
187. Ogawa, T., and T. Okazaki. 1994. Cell cycle-dependent transcription from the gid and mioC promoters of Escherichia coli. J. Bacteriol.176:1609–1615.[PubMed]
188. Ogawa, T., and T. Okazaki. 1991. Concurrent transcription from the gid and mioC promoters activates replication of an Escherichia coli minichromosome. Mol. Gen. Genet.230:193–200.[PubMed] [CrossRef]
189. Ogawa, T., M. Wada, Y. Kano, F. Imamoto, and T. Okazaki. 1989. DNA replication in Escherichia coli mutants that lack protein HU. J. Bacteriol.171:5672–5679.[PubMed]
190. Ogawa, T., Y. Yamada, T. Kuroda, T. Kishi, and S. Moriya. 2002. The datA locus predominantly contributes to the initiator titration mechanism in the control of replication initiation in Escherichia coli. Mol. Microbiol.44:1367–1375.[PubMed] [CrossRef]
191. Ogden, G. B., M. J. Pratt, and M. Schaechter. 1988. The replicative origin of the E. coli chromosome binds to cell membranes only when hemimethylated. Cell54:127–135.[PubMed] [CrossRef]
192. Ohmori, H., M. Kimura, T. Nagata, and Y. Sakakibara. 1984. Structural analysis of the dnaA and dnaN genes of Escherichia coli. Gene28:159–170.[PubMed] [CrossRef]
193. Ohniwa, R. L., K. Morikawa, J. Kim, T. Ohta, A. Ishihama, C. Wada, and K. Takeyasu. 2006. Dynamic state of DNA topology is essential for genome condensation in bacteria. EMBO J.25:5591–5602.[PubMed] [CrossRef]
194. Oka, A., H. Sasaki, K. Sugimoto, and M. Takanami. 1984. Sequence organization of replication origin of the Escherichia coli K-12 chromosome. J. Mol. Biol.176:443–458.[PubMed] [CrossRef]
195. Oka, A., K. Sugimoto, H. Sasaki, and M. Takanami. 1982. An in vitro method generating base substitutions in preselected regions of plasmid DNA: application to structural analysis of the replication origin of the Escherichia coli K-12 chromosome. Gene19:59–69.[PubMed] [CrossRef]
196. Ozaki, S., H. Kawakami, K. Nakamura, N. Fujikawa, W. Kagawa, S. Y. Park, S. Yokoyama, H. Kurumizaka, and T. Katayama. 2008. A common mechanism for the ATP-DnaA-dependent formation of open complexes at the replication origin. J. Biol. Chem.283:8351–8362.[PubMed] [CrossRef]
197. Paulsson, J., and D. K. Chattoraj. 2006. Origin inactivation in bacterial DNA replication control. Mol. Microbiol.61:9–15.[PubMed] [CrossRef]
198. Pierucci, O. 1979. Phospholipid synthesis during the cell division cycle of Escherichia coli. J. Bacteriol.138:453–460.[PubMed]
199. Pierucci, O., M. Rickert, and C. E. Helmstetter. 1989. DnaA protein overproduction abolishes cell cycle specificity of DNA replication from oriC in Escherichia coli. J. Bacteriol.171:3760–3766.[PubMed]
200. Polaczek, P., K. Kwan, D. A. Liberies, and J. L. Campbell. 1997. Role of architectural elements in combinatorial regulation of initiation of DNA replication in Escherichia coli. Mol. Microbiol.26:261–275.[PubMed] [CrossRef]
201. Polaczek, P., and A. Wright. 1990. Regulation of expression of the dnaA gene in Escherichia coli: role of the two promoters and the DnaA box. New Biol.2:574–582.[PubMed]
202. Pritchard, R. H., P. T. Barth, and J. Collins. 1969. Control of DNA synthesis in bacteria. Symp. Soc. Gen. Microbiol.19:263–297.
203. Riber, L., K. Fujimitsu, T. Katayama, and A. Lobner-Olesen. 2008. Loss of Hda activity stimulates replication initiation from I-box, but not R4 mutant origins in Escherichia coli. Mol. Microbiol.71:107–122.[PubMed] [CrossRef]
204. Riber, L., and A. Lobner-Olesen. 2005. Coordinated replication and sequestration of oriC and dnaA are required for maintaining controlled once-per-cell-cycle initiation in Escherichia coli. J. Bacteriol.187:5605–5613.[PubMed] [CrossRef]
205. Riber, L., J. A. Olsson, R. B. Jensen, O. Skovgaard, S. Dasgupta, M. G. Marinus, and A. Lobner-Olesen. 2006. Hda-mediated inactivation of the DnaA protein and dnaA gene autoregulation act in concert to ensure homeostatic maintenance of the Escherichia coli chromosome. Genes Dev.20:2121–2134.[PubMed] [CrossRef]
206. Roth, A., and W. Messer. 1995. The DNA binding domain of the initiator protein DnaA. EMBO J.14:2106–2111.[PubMed]
207. Roth, A., and W. Messer. 1998. High-affinity binding sites for the initiator protein DnaA on the chromosome of Escherichia coli. Mol. Microbiol.28:395–401.[PubMed] [CrossRef]
208. Roth, A., B. Urmoneit, and W. Messer. 1994. Functions of histone-like proteins in the initiation of DNA replication at oriC of Escherichia coli. Biochimie76:917–923.[PubMed] [CrossRef]
209. Russell, D. W., and N. D. Zinder. 1987. Hemimethylation prevents DNA replication in E. coli. Cell50:1071–1079.[PubMed] [CrossRef]
210. Ryan, V. T., J. E. Grimwade, J. E. Camara, E. Crooke, and A. C. Leonard. 2004. Escherichia coli prereplication complex assembly is regulated by dynamic interplay among Fis, IHF and DnaA. Mol. Microbiol.51:1347–1359.[PubMed] [CrossRef]
211. Ryan, V. T., J. E. Grimwade, C. J. Nievera, and A. C. Leonard. 2002. IHF and HU stimulate assembly of pre-replication complexes at Escherichia coli oriC by two different mechanisms. Mol. Microbiol.46:113–124.[PubMed] [CrossRef]
212. Samitt, C. E., F. G. Hansen, J. F. Miller, and M. Schaechter. 1989. In vivo studies of DnaA binding to the origin of replication of Escherichia coli. EMBO J.8:989–993.[PubMed]
213. Schaechter, M. 1961. Patterns of cellular control during unbalanced growth. Cold Spring Harbor Symp. Quant. Biol.26:53–62.[PubMed]
214. Schaechter, M., O. Maaloe, and N. O. Kjeldgaard. 1958. Dependency on medium and temperature of cell size and chemical composition during balanced grown of Salmonella typhimurium. J. Gen. Microbiol.19:592–606.[PubMed]
215. Schaper, S., and W. Messer. 1995. Interaction of the initiator protein DnaA of Escherichia coli with its DNA target. J. Biol. Chem.270:17622–17626.[PubMed] [CrossRef]
216. Schauzu, M. A., C. Kucherer, R. Kolling, W. Messer, and H. Lother. 1987. Transcripts within the replication origin, oriC, of Escherichia coli. Nucleic Acids Res. 15:2479–2497.[PubMed] [CrossRef]
217. Seitz, H., C. Weigel, and W. Messer. 2000. The interaction domains of the DnaA and DnaB replication proteins of Escherichia coli. Mol. Microbiol.37:1270–1279.[PubMed] [CrossRef]
218. 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. Cell50:259–265.[PubMed] [CrossRef]
219. Sekimizu, K., D. Bramhill, and A. Kornberg. 1988. Sequential early stages in the in vitro initiation of replication at the origin of the Escherichia coli chromosome. J. Biol. Chem.263:7124–7130.[PubMed]
220. Sekimizu, K., and A. Kornberg. 1988. Cardiolipin activation of dnaA protein, the initiation protein of replication in Escherichia coli. J. Biol. Chem.263:7131–7135.[PubMed]
221. Sekimizu, K., B. Y. Yung, and A. Kornberg. 1988. The dnaA protein of Escherichia coli. Abundance, improved purification, and membrane binding. J. Biol. Chem.263:7136–7140.[PubMed]
222. Simmons, L. A., A. M. Breier, N. R. Cozzarelli, and J. M. Kaguni. 2004. Hyperinitiation of DNA replication in Escherichia coli leads to replication fork collapse and inviability. Mol. Microbiol.51:349–358.[PubMed] [CrossRef]
223. Simmons, L. A., M. Felczak, and J. M. Kaguni. 2003. DnaA protein of Escherichia coli: oligomerization at the E. coli chromosomal origin is required for initiation and involves specific N-terminal amino acids. Mol. Microbiol.49:849–858.[PubMed] [CrossRef]
224. Skarstad, K., and E. Boye. 1994. The initiator protein DnaA: evolution, properties and function. Biochim. Biophys. Acta1217:111–130.[PubMed]
225. Skarstad, K., E. Boye, and H. B. Steen. 1986. Timing of initiation of chromosome replication in individual Escherichia coli cells. EMBO J.5:1711–1717.[PubMed]
226. Skarstad, K., A. Løbner-Olesen, T. Atlung, K. von Meyenburg, and E. Boye. 1989. Initiation of DNA replication in Escherichia coli after overproduction of the DnaA protein. Mol. Gen. Genet.218:50–56.[PubMed] [CrossRef]
227. Skarstad, K., G. Lueder, R. Lurz, C. Speck, and W. Messer. 2000. The Escherichia coli SeqA protein binds specifically and co-operatively to two sites in hemimethylated and fully methylated oriC. Mol. Microbiol.36:1319–1326.[PubMed] [CrossRef]
228. Slater, S., S. Wold, M. Lu, E. Boye, K. Skarstad, and N. Kleckner. 1995. E. coli SeqA protein binds oriC in two different methyl-modulated reactions appropriate to its roles in DNA replication initiation and origin sequestration. Cell82:927–936.[PubMed] [CrossRef]
229. Smith, D. W., A. M. Garland, G. Herman, R. E. Enns, T. A. Baker, and J. W. Zyskind. 1985. Importance of state of methylation of oriC GATC sites in initiation of DNA replication in Escherichia coli. EMBO J.4:1319–1326.[PubMed]
230. Sompayrac, L., and O. Maaloe. 1973. Autorepressor model for control of DNA replication. Nat. New Biol.241:133–135.[PubMed] [CrossRef]
231. Speck, C., and W. Messer. 2001. Mechanism of origin unwinding: sequential binding of DnaA to double- and single-stranded DNA. EMBO J. 20:1469–1476.[PubMed] [CrossRef]
232. Speck, C., C. Weigel, and W. Messer. 1999. ATP- and ADP-dnaA protein, a molecular switch in gene regulation. EMBO J.18:6169–6176.[PubMed] [CrossRef]
233. Stillman, B. 2005. Origin recognition and the chromosome cycle. FEBS Lett.579:877–884.[PubMed] [CrossRef]
234. Su'etsugu, M., A. Emoto, K. Fujimitsu, K. Keyamura, and T. Katayama. 2003. Transcriptional control for initiation of chromosomal replication in Escherichia coli: fluctuation of the level of origin transcription ensures timely initiation. Genes Cells8:731–745.[PubMed] [CrossRef]
235. Su'etsugu, M., K. Nakamura, K. Keyamura, Y. Kudo, and T. Katayama. 2008. Hda monomerization by ADP binding promotes replicase clamp-mediated DnaA-ATP hydrolysis. J. Biol. Chem.283:36118–36131.[PubMed] [CrossRef]
236. Su'etsugu, M., T. R. Shimuta, T. Ishida, H. Kawakami, and T. Katayama. 2005. Protein associations in DnaA-ATP hydrolysis mediated by the Hda-replicase clamp complex. J. Biol. Chem.280:6528–6536.[PubMed] [CrossRef]
237. Sutton, M. D., K. M. Carr, M. Vicente, and J. M. Kaguni. 1998. Escherichia coli DnaA protein. The N-terminal domain and loading of DnaB helicase at the E. coli chromosomal origin. J. Biol. Chem.273:34255–34262.[PubMed] [CrossRef]
238. Sutton, M. D., and J. M. Kaguni. 1997. The Escherichia coli dnaA gene: four functional domains. J. Mol. Biol.274:546–561.[PubMed] [CrossRef]
239. Sutton, M. D., and J. M. Kaguni. 1997. Threonine 435 of Escherichia coli DnaA protein confers sequence-specific DNA binding activity. J. Biol. Chem.272:23017–23024.[PubMed] [CrossRef]
240. 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 Cells5:873–884.[PubMed] [CrossRef]
241. Theisen, P. W., J. E. Grimwade, A. C. Leonard, J. A. Bogan, and C. E. Helmstetter. 1993. Correlation of gene transcription with the time of initiation of chromosome replication in Escherichia coli. Mol. Microbiol.10:575–584.[PubMed] [CrossRef]
242. Thony, B., D. S. Hwang, L. Fradkin, and A. Kornberg. 1991. iciA, an Escherichia coli gene encoding a specific inhibitor of chromosomal initiation of replication in vitro. Proc. Natl. Acad. Sci. USA88:4066–4070.[PubMed] [CrossRef]
243. Torheim, N. K., and K. Skarstad. 1999. Escherichia coli SeqA protein affects DNA topology and inhibits open complex formation at oriC. EMBO J.18:4882–4888.[PubMed] [CrossRef]
244. Tucker, P. A., and L. Sallai. 2007. The AAA+ superfamily—a myriad of motions. Curr. Opin. Struct. Biol.17:641–652.[PubMed] [CrossRef]
245. von Freiesleben, U., M. A. Krekling, F. G. Hansen, and A. Løbner-Olesen. 2000. The eclipse period of Escherichia coli. EMBO J.19:6240–6248.[PubMed] [CrossRef]
246. von Freiesleben, U., and K. V. Rasmussen. 1991. DNA replication in Escherichia coli gyrB(Ts) mutants analysed by flow cytometry. Res. Microbiol.142:223–227.[PubMed] [CrossRef]
247. von Freiesleben, U., and K. V. Rasmussen. 1992. The level of supercoiling affects the regulation of DNA replication in Escherichia coli. Res. Microbiol.143:655–663.[PubMed] [CrossRef]
248. Von Freiesleben, U., K. V. Rasmussen, T. Atlung, and F. G. Hansen. 2000. Rifampicin-resistant initiation of chromosome replication from oriC in ihf mutants. Mol. Microbiol.37:1087–1093.[PubMed] [CrossRef]
249. von Meyenburg, K., and F. G. Hansen. 1987. Regulation of chromosome replication, p. 1555–1577. In F. C. Neidhardt, J. Ingraham, K. B. Low, B. Magasanik, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: Cellular and Molecular Biology. ASM Press, Washington, DC.
250. von Meyenburg, K., F. G. Hansen, E. Riise, H. E. Bergmans, M. Meijer, and W. Messer. 1979. Origin of replication, oriC, of the Escherichia coli K12 chromosome: genetic mapping and minichromosome replication. Cold Spring Harbor Symp. Quant. Biol.43(Pt 1):121–128.[PubMed]
251. Wahle, E., R. S. Lasken, and A. Kornberg. 1989. The dnaB-dnaC replication protein complex of Escherichia coli. I. Formation and properties. J. Biol. Chem.264:2463–2468.[PubMed]
252. Wahle, E., R. S. Lasken, and A. Kornberg. 1989. The dnaB-dnaC replication protein complex of Escherichia coli. II. Role of the complex in mobilizing dnaB functions. J. Biol. Chem.264:2469–2475.[PubMed]
253. Waldminghaus, T., and K. Skarstad. 2009. The Escherichia coli SeqA protein. Plasmid61:141–150.[PubMed] [CrossRef]
254. Weigel, C., W. Messer, S. Preiss, M. Welzeck, Morigen, and E. Boye. 2001. The sequence requirements for a functional Escherichia coli replication origin are different for the chromosome and a minichromosome. Mol. Microbiol.40:498–507.[PubMed] [CrossRef]
255. Weigel, C., A. Schmidt, B. Ruckert, R. Lurz, and W. Messer. 1997. DnaA protein binding to individual DnaA boxes in the Escherichia coli replication origin, oriC. EMBO J.16:6574–6583.[PubMed] [CrossRef]
256. Weigel, C., A. Schmidt, H. Seitz, D. Tungler, M. Welzeck, and W. Messer. 1999. The N-terminus promotes oligomerization of the Escherichia coli initiator protein DnaA. Mol. Microbiol.34:53–66.[PubMed] [CrossRef]
257. Woelker, B., and W. Messer. 1993. The structure of the initiation complex at the replication origin, oriC, of Escherichia coli. Nucleic Acids Res.21:5025–5033.[PubMed] [CrossRef]
258. Wold, S., E. Crooke, and K. Skarstad. 1996. The Escherichia coli Fis protein prevents initiation of DNA replication from oriC in vitro. Nucleic Acids Res.24:3527–3532.[PubMed] [CrossRef]
259. Xia, W., and W. Dowhan. 1995. In vivo evidence for the involvement of anionic phospholipids in initiation of DNA replication in Escherichia coli. Proc. Natl. Acad. Sci. USA92:783–787.[PubMed] [CrossRef]
260. Xu, Q., D. McMullan, P. Abdubek, T. Astakhova, D. Carlton, C. Chen, H. J. Chiu, T. Clayton, D. Das, M. C. Deller, L. Duan, M. A. Elsliger, J. Feuerhelm, J. Hale, G. W. Han, L. Jaroszewski, K. K. Jin, H. A. Johnson, H. E. Klock, M. W. Knuth, P. Kozbial, S. Sri Krishna, A. Kumar, D. Marciano, M. D. Miller, A. T. Morse, E. Nigoghossian, A. Nopakun, L. Okach, S. Oommachen, J. Paulsen, C. Puckett, R. Reyes, C. L. Rife, N. Sefcovic, C. Trame, H. van den Bedem, D. Weekes, K. O. Hodgson, J. Wooley, A. M. Deacon, A. Godzik, S. A. Lesley, and I. A. Wilson. 2009. A structural basis for the regulatory inactivation of DnaA. J. Mol. Biol.385:368–380.[PubMed] [CrossRef]
261. Yamazoe, M., S. Adachi, S. Kanaya, K. Ohsumi, and S. Hiraga. 2005. Sequential binding of SeqA protein to nascent DNA segments at replication forks in synchronized cultures of Escherichia coli. Mol. Microbiol.55:289–298.[PubMed] [CrossRef]
262. Yasuda, S., and Y. Hirota. 1977. Cloning and mapping of the replication origin of Escherichia coli. Proc. Natl. Acad. Sci. USA74:5458–5462.[PubMed] [CrossRef]
263. Yim, L., I. Moukadiri, G. R. Bjork, and M. E. Armengod. 2006. Further insights into the tRNA modification process controlled by proteins MnmE and GidA of Escherichia coli. Nucleic Acids Res.34:5892–5905.[PubMed] [CrossRef]
264. Yoo, S. J., J. H. Seol, S. K. Woo, S. W. Suh, D. S. Hwang, D. B. Ha, and C. H. Chung. 1993. Hydrolysis of the IciA protein, an inhibitor of DNA replication initiation, by protease Do in Escherichia coli. FEBS Lett. 327:17–20.[PubMed] [CrossRef]
265. Yung, B. Y., E. Crooke, and A. Kornberg. 1990. Fate of the DnaA initiator protein in replication at the origin of the Escherichia coli chromosome in vitro. J. Biol. Chem.265:1282–1285.[PubMed]
266. Yung, B. Y., and A. Kornberg. 1989. The dnaA initiator protein binds separate domains in the replication origin of Escherichia coli. J. Biol. Chem.264:6146–6150.[PubMed]
267. Zakrzewska-Czerwinska, J., D. Jakimowicz, A. Zawilak-Pawlik, and W. Messer. 2007. Regulation of the initiation of chromosomal replication in bacteria. FEMS Microbiol. Rev.31:378–387.[PubMed] [CrossRef]
268. Zaritsky, A., N. Vischer, and A. Rabinovitch. 2007. Changes of initiation mass and cell dimensions by the ‘eclipse.’ Mol. Microbiol.63:15–21.[PubMed] [CrossRef]
269. Zawilak-Pawlik, A., A. Kois, J. Majka, D. Jakimowicz, A. Smulczyk-Krawczyszyn, W. Messer, and J. Zakrzewska-Czerwinska. 2005. Architecture of bacterial replication initiation complexes: orisomes from four unrelated bacteria. Biochem. J.389:471–481.[PubMed] [CrossRef]
270. Zheng, W., Z. Li, K. Skarstad, and E. Crooke. 2001. Mutations in DnaA protein suppress the growth arrest of acidic phospholipid-deficient Escherichia coli cells. EMBO J.20:1164–1172.[PubMed] [CrossRef]
271. Zyskind, J. W., J. M. Cleary, W. S. Brusilow, N. E. Harding, and D. W. Smith. 1983. Chromosomal replication origin from the marine bacterium Vibrio harveyi functions in Escherichia coli: oriC consensus sequence. Proc. Natl. Acad. Sci. USA80:1164–1168.[PubMed] [CrossRef]
272. Zyskind, J. W., L. T. Deen, and D. W. Smith. 1977. Temporal sequence of events during the initiation process in Escherichia coli deoxyribonucleic acid replication: roles of the dnaA and dnaC gene products and ribonucleic acid polymerase. J. Bacteriol.129:1466–1475.[PubMed]
Module4.4.1
