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9 Post-DNA Replication Nucleoid Formation; Slow and Fast Doubling Times

9 Post-DNA Replication Nucleoid Formation; Slow and Fast Doubling Times

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7  Molecular Structure and Dynamics of Bacterial Nucleoids



133



7.9.1 Nucleoid Formation in Cells Growing on Defined Medium

At slow growth conditions, synchronous cultures of E. coli cells made using a

“baby machine” method (Bates et al. 2005) with a doubling time of 125 min exhibit

eight important time signatures (Fig.  7.5). (1) DNA replication begins at 17 min

after elution from the column and DNA synthesis continues for 50–60 min. (2) For

10–14 min period after replication begins, the duplicate copies of oriC remain

together (Bates and Kleckner 2005). (3) At 27–32 min, the two oriC regions move

to the ¼ and ¾ positions of the cell, which is the position that will become cell

centers in the new daughter cells. (4) Nucleoids can be visualized in a special

medium where the refractive index is carefully controlled, or by staining with

DAPI. In baby machine conditions, nucleoids split at about 60 min. (5) Replication

is complete by 67–77 min. (6) Separation of fluorescent markers near the terminus

occurs at about 80 min. (7) At 100 min closure of the septum begins. (8) Finally



DNA

events



27-31”

17”

Initiate Two oriC

foci

DNA

Repliction



63-70”

Terminate

DNA

Replication



80-90”

Two Ter

foci



Cell

events

0“

New

Cell

1 oriC focus



28-32”

oriC’s

Move to

1/4 & 3/4

Position



60”

Nucleoid

Splitting



100”

Septation

Begins



125”

Cell

Doubling



15-20”

Two Ter

foci



25“

0“

Cell

New

Doubling

Cell

3 or more

oriC foci

1 Ter focus



Fig.  7.5  Time signature of discernable events in an “average” cell growing exponentially in

minimal medium with a 125 min doubling time (top) or in rich LB medium with a 20 min doubling time (bottom). Data abstracted from (Bates and Kleckner 2005; Nielsen et al. 2006; Wang

et al. 2006; Wang et al. 2008)



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cell division is completed 40 min later at 120 min. A diagram of nucleoid division

patterns is shown in Fig. 7.6. In newborn cells there is a single MukBEF focus for

slow growing cells. During the synthetic period, the focus doubles and each moves



O

oriC

Replichore

II



Replichore

I



Slow Growth

Doubling@ 125 min.



dif



DichotomousGrowth

Doubling@ 20 min.



MukBEF



Division Plane



Division Plane



New Nucleoids



Replichore

II



Old Nucleoid



1/4



Mid-Cell



dif



Replichore

I



3/4



1/4



Mid-Cell



3/4



dif



Fig.  7.6  Chromosome organization and segregation patterns during slow growth (left) and

dichotomous growth (right). The DNA replication pattern from oriC to the terminus near dif is

shown at top with replichore I blue and replichore II red. Replication of the continuously synthesized and discontinuously synthesized strands from each replichore are shown as solid and broken

lines, respectively. MukBEF condensin foci are indicated as green ovals near the quarter positions

of the long cell axis. The cell septum is marked by a purple line at mid-cell



7  Molecular Structure and Dynamics of Bacterial Nucleoids



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to the quarter positions of the cell, i.e. between midcell and cell poles on the long

cell axis.

The structure and binding force(s) that stabilize oriC near the quarter positions

are not understood, although the MukBEF condensin plays a part. Estimates on the

copy number of MukBEF molecules differ from 100–500 per cell. MukBEF forms

a focus at the quarter positions (shown as green ovals in Fig.  7.5) (Niki et  al.

2000), and this complex helps condense nucleoids into structures that segregate

efficiently (Sawitzke and Austin 2000). A mukB null mutant fails to position oriC

at the quarter cell locations, which results in the formation of nucleoids with a

more random organization than WT nucleoids (Danilova et al. 2007).

Nucleoid development has been followed in a number of different labs using

E. coli strains tagged with Lac and Tet operator arrays decorated with two colored

fluorescent repressors or parC sites decorated with fluorescent ParB protein. The

following summary of events represents a hypothetical average cell in an asynchronous population (Fig. 7.6). The continuous strand from one replichore (here arbitrarily labeled replichore I) is assembled into chromatin and deposited at the inner

margin of the left nucleoid. The discontinuously synthesized strand of replichore I,

shown as broken blue line, is deposited at the outer margin of the right nucleoid.

The continuous and discontinuous strands from replichore II move to the inner and

outer edges of the right and left nucleoids respectively. E. coli generates nucleoids

that contain genes from different replichores lying bilaterally on different sides of

the origin. Genes closest to the origin are followed by genes positioned toward the

terminus. This leads to a direct replichore repeat pattern in most cells (63%) with a

minor fraction of cells (30%) becoming mirror image replichore nucleoids (Wang

et al. 2006; Wang et al. 2005). This pattern is distinctly different from Caulobacter

cresentus, which generates exclusively mirror image nucleoids with oriC at one

pole and dif at the pole that has either a stalk or a flagellum (Viollier et al. 2004).

Both early and late stages of segregation appear to take extra time in slow

growth conditions. The delay between initiation and movement of the oriC to relatively fixed positions has been called cohesion, and the mechanism may be complex (Bates and Kleckner 2005; Sunako et  al. 2001). One possibility is that

pre-catenane links between the sisters (Fig.  1.2) requires time to organize and

untangle this region to allow freedom to move to quarter positions (Grainge et al.

2007). There is also a significant delay between the completion of DNA replication and separation of fluorescent foci near the terminus, which may be linked to

the MatP system. MatP binds specific sites and organizes a domain near the terminus (Mercier et al. 2008). Complete decatenation of links between sister chromatids, is necessary for complete segregation, and this process is catalyzed most

efficiently by Topo IV, which uses the DNA binding and motor domain of FtsK to

do the job efficiently.

About 15% of the cells undergo RecA-dependent homologous recombination

that generates a dimer chromosome. Chromosome dimers are resolved by the combined action of XerC, XerD, and FtsK working at the dif site. Complete decatenation is carried out by the combined action of gyrase, Topo IV, FtsK, XerC, XerD

and polarized G-rich Kops sequences that are polarized about the dif site.



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7.9.2 Nucleoid Formation During Dichotomous Replication

Some bacteria have evolved an adaptive ability to compress cell division stages to

achieve a very rapid doubling time. Rapid growth is a strong selective advantage

during the competition for a niche in the mammalian gut (Freter et  al. 1983).

Dichotomous replication drastically constricts the time period for carrying out all

the functions that are carried out in sequential order during slow growth (Fig. 7.5,

lower). Dichotomous growth depends on having a large biosynthetic capacity so

that cells can synthesize proteins, nucleic acids, and all cellular components at

break-neck speeds needed to produce viable cells every 20 min. Organisms that

carry out dichotomous growth exhibit a signature of multiple copies of ribosomal

RNA operons. Sueoka described dichotomous growth first in B. subtilis (Sueoka

1971), which has 12 rrn (ribosomal RNA) operons. Both E. coli and Salmonella

grow dichotomously and have 7 rrn operons (Fig. 7.4). By contrast, Caulobacter

cresentus, which doubles slowly in over an hour, has only 2 rrn operons to satisfy

its protein requirements, and the very slow growing M. tuberculosis with a 12 h

doubling time requires only one rrn operon.

Chromosome segregation becomes more complicated under dichotomous

growth conditions. First, each newborn cell has two MukBEF foci, and during the

division cycle 3–4 foci are seen. When a critical mass to volume is reached in

E. coli, and replisomes have moved a significant distance toward the terminus, cells

synchronously reinitiate new forks using both replicated oriC sequences. This adds

four additional nucleoids to the picture (Fig. 7.5 right side.) The segregation pattern

is now complicated and the timing of each step is constricted and difficult to sort

out using marked chromosomes in live cells. Nonetheless, at the time of cell division, each daughter cells must untangle one complete sister chromosome and 2 (or

even 4) partially replicated arcs with bi-directional replicating oriC elements.



7.10 Dichotomous Chaos at dif

All barriers to the replicative DnaB helicase must be eliminated to allow unimpeded fork

movement at the normal rate of >500 bp/s. DNA replication in cells carrying the gyrB652

mutation bogs down due to the sluggish supercoiling action of a hypomorphic gyrase

(Pang et al. 2005). This impediment mimics the effect of gyrase inhibited by quinolones,

which impedes RNA polymerase (Willmott et  al. 1994) and causes fork arrest when

DNA polymerase encounters the gyrase-quinolone-DNA complex (Hiasa and Marians

1996). Once a fork becomes arrested, there is a critical time for rescue before it is overtaken by the next round of synthesis. A second fork running into a stalled fork causes

chromosome breaks (Bidnenko et al. 2002). The experimental signature of dichotomous

chaos in Salmonella is a dramatic (30-fold or more) loss of gd-catalyzed resolution near

the dif site (Pang et al. 2005). This loss in recombination potential is likely caused by

unresolved catenane links that make synapsis inefficient, and possibly by a loss of −SC

around the dif site caused by replication fork convergence.



7  Molecular Structure and Dynamics of Bacterial Nucleoids



137



In addition to death caused by a sluggish gyrase, a similar phenotype is triggered

in null mutants of MukB (Niki et al. 1991) or by hyper-initiation. Grigorian et al.

showed that over-production of DnaA leads to over-initiation at oriC and fork collapse (Grigorian et al. 2003). More extensive analysis by Simmons et al. demonstrated that hyper-initiation of DNA replication in E. coli leads to breakage of

chromosomal DNA (Simmons et  al. 2004). They analyzed fork movement with

microarrays but found no specific sites of fork arrest. They suggested that merging

forks could cause breaks under hyper-initiation conditions. This hypothesis also

explains the sickness of seqA mutations. These strains are only healthy in minimal

medium at low temperature because they lack the sequestration mechanism in

which SeqA binding inhibits premature reinitiation by blocking the DnaA protein’s

access to oriC (Campbell and Kleckner 1990; Skarstad et al. 2001).



7.11 Differences in Species-Specific Supercoil Set Points

Supercoiling is a critical factor for nucleoid formation and segregation. The multiple functions of negative supercoiling include: (1) Gyrase-dependent compaction

of DNA into an interwound and branched plectoneme. (2) Segregation of sister

chromosomes requires unknotting and decatenation of newly synthesized double

strands. All simplifications in DNA topology are dramatically stimulated by negative supercoiling, which brings the interwound strands of DNA close together and

makes inter-chromosomal links more apparent to the decatenase, Topo IV

(Rybenkov et al. 1997). Single molecule experiments show that Topo IV acts preferentially on positive nodes rather than on the negative nodes of −SC DNA (Stone

et al. 2003). (3) Over 300 genes, including the promoters for gyrA, gyrB, and topA,

are regulated by supercoiling stress (Peter et  al. 2004). Considering the central

importance of gyrase in nucleoid formation, it was surprising to discover that WT

strains of E. coli and S. Typhimurium generate substantially different levels of

in vivo torsional strain. Nonetheless, plasmid pBR322 extracted from E. coli has a

median s value of −0.70 whereas the same plasmid from S. Typhimurium has a

value of −0.06 (Fig. 7.7) (Champion and Higgins 2007). As one illustration of the

significance of this difference, WT E. coli is not viable when grown at the

Salmonella level of s.



7.12 The Paradox of Supercoil Dynamics in E. coli

and Salmonella

Why have E. coli and Salmonella evolved to maintain different levels of s? How

do two organisms regulate >300 supercoil-responsive genes at different levels of

torsional strain? How does nucleoid formation differ between two species with such

large differences in average supercoiling? Answers to both questions are not



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E. coli - High Strung



Salmonella - Low Strung



σ = -0.69



σ = -0.59



TopA lethal



TopA tolerated



GyrB652 lethal



GyrB652 tolerated



SeqA sick in LB



SeqA healthy in LB



MukB viable in LB



MukB lethal in LB



H-NS null healthy



H-NS null sick



Fig.  7.7  Comparison of supercoil density and nucleoid related phenotypes of E. coli and

Salmonella typhimurium. Red = severe to lethal phenotype, Black = mild to non-detectable phenotype



entirely solved, but it is clear that these organisms have many significant phenotypic

differences for identical mutations in topologically sensitive genes. The list includes

gyrB, topA, seqA, mukB, and hns see Fig. 7.7 (Champion and Higgins 2007; Falconi

et al. 1991; Navarre et al. 2006). Although there could be complicated explanations

that cannot be excluded in each case, supercoiling provides a simple unifying

explanation for all of the observed phenotypes.

Amber mutations in topA (w protein) have long been known to be lethal in E. coli

but are healthy in S. Typhimurium or Shigella flexneri (Ní Bhriain and Dorman 1993;

DiNardo et al. 1982; Margolin et al. 1985). In E. coli, high torsional strain caused by

elimination of topA results in transcription-driven inter-molecular RNA triplex formation, or R-loops (Drolet et  al. 2003; Hraiky et  al. 2000). R-loops block further

transcription and stall replication forks (Higgins and Vologodskii 2004). Because

Salmonella generates 15% lower s, −SC is already lower than E. coli strains carrying

permissive gyrB compensatory mutations that allow introduction of topA amber

mutations (DiNardo et al. 1982; Jaworski et al. 1991). R-loop formation would be

expected to not be as severe in WT Salmonella as it is in E. coli.

Many gyrB mutations represented by a single nucleotide substitution in

Salmonella are lethal when introduced into WT E. coli. This includes the R436S

arginine to serine substitution that gives a TS phenotype at 42° in Salmonella but



7  Molecular Structure and Dynamics of Bacterial Nucleoids



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grows normally at 30° (Pang et al. 2005). Recombination experiments that would

result in exchange of the WT Salmonella gyrB sequence for the WT E. coli gene

have proven to be lethal, and more surprisingly, even low level expression of either

mutant or WT S. Typhimurium GyrB protein in WT E. coli is toxic (Champion and

Higgins 2007). The reciprocal procedure of exchanging either WT or mutant

Salmonella gyrB alleles with the WT E. coli gene is easily done, and expressing

E. coli GyrB in Salmonella is non-toxic except at extremely high expression levels.

The simplest hypothesis to explain these results is that the GyrB protein is an

important determinant of maintaining chromosomal s, and that the Salmonella

GyrB protein does not generate sufficient torsional strain to meet minimum requirements for E. coli growth in rich medium.

seqA null mutants in E. coli exhibit growth-rate toxicity in rich LB medium

while Salmonella does not (Fig.  7.7). In E. coli SeqA helps quell lethal consequences of over-initiation by sequestering the hemi-methylated segments of oriC

region for 10 min or more after initiation. This inhibits DnaA-dependent replisome

reassembly (Camara and Crooke 2005) and toxic problems associated with re-initiation of replication at oriC leads to filamentation and cell death due replication

fork failure (Grigorian et al. 2003; Simmons et al. 2004). One critical step in the

oriC initiation is unwinding of A/T rich sequences adjacent to DnaA binding sites

at the origin (Baker et al. 1986; Funnell et al. 1986; 1987). The 15% lower supercoil

density that Salmonella attains during dichotomous replication may decrease oriC

initiation to acceptable levels in rich media without the contribution of SeqA

sequestration.

An exception that proves the rule is the mukB mutation. Null mutants in

Salmonella have a much more severe phenotype than E. coli (Fig. 7.7). An E. coli

mukB mutant shows toxicity at 40° on LB, but grows well on LB medium at 30°.

A Salmonella mukB mutant is viable only on minimal medium and plates on LB

medium six orders of magnitude less efficiently than E. coli. In E. coli, supercoiling

influences the phenotype of mukB mutants. Sawitzke and Austin demonstrated that

growth in the presence of low levels of the GyrB inhibitor novobiocin increased the

E. coli reliance on MukB, whereas increasing the median supercoil density by the

introducing a topA mutation made it possible for E. coli mukB mutants to plate on

LB up to 42° (Holmes and Cozzarelli 2000; Sawitzke and Austin 2000). Salmonella

fits this pattern. The lower level of s correlates with growth of E. coli on novobiocin and a consequential heavier reliance on MukB for condensation and segregation

in rich medium (Fig. 7.7).

The fifth example is the hns gene. Like MukB, deletion of H-NS results in a more

severe phenotype in Salmonella than it does in E. coli, where growth rates and general

physiology of WT and hns null mutations are nearly indistinguishable (Falconi et al.

1991; Owen-Hughes et al. 1992). H-NS participates in chromosome condensation,

regulation of gene expression, and the targeting of transposons (Johnson et al. 2005).

In E. coli H-NS influences expression directly or indirectly of about 200 genes

(Hommais et al. 2001) while experiments in Salmonella indicate that expression of

over 400 genes are altered in an hns null mutant (Navarre et al. 2006). Most of the

genes in both organisms are derepressed when H-NS is eliminated. In addition to



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regulating genes, H-NS is one of the proteins that may organize loops in the bacterial chromosome (Deng et al. 2005). H-NS-induced looping in vitro has been demonstrated by atomic force microscopy (Dame et  al. 2000) as well as by single

molecule manipulation using optical tweezers (Dame et al. 2006). It has been proposed, although it remains to be proven, that the apparently stochastic distribution

of regions of H-NS binding along the genome reflects the organization of DNA into

supercoiled domains. (Noom et al. 2007). When over-expressed, H-NS can cause a

dramatic condensation of the nucleoid which leads to inhibition of global transcription and an artificial stationary phase (McGovern et al. 1994). The lower natural

supercoiling level of Salmonella may contribute to the increased dependence of this

organism on H-NS for maintaining nucleoid structure. Many of the genes that are

bound by H-NS in Salmonella appear to be A/T rich and transferred horizontally

from distantly related organisms (Navarre et al. 2006).



7.13 Major Unanswered Questions

Whereas recent experimental work has made great strides in shaping a new

understanding of what happens in nucleoid development, there are new questions

that are being uncovered as well. For example, what machinery channels the

nascent strands to form two separate nucleoids? What forces cause the oriC

region to be more restricted than other parts of the genome? And what about

loops and evolution?



7.13.1 Why Did E. coli and Salmonella Develop Different

Supercoil Set Points?

One selective pressure for reducing the value of s may come from prophages in

the Salmonella genome (Fig.  7.4). E. coli K12 harbors no prophage that can

make an infectious virus in a lytic cycle whereas S. Typhimurium has 4 lysiscompetent prophage; they are Gifsy-1, Gifsy-2, Fels-1, and Fels-2 (McClelland

et  al. 2001). How long Salmonella Typhimurium has harbored 4 prophages is

uncertain, but both Gifsy-1 and Gifsy-2 contain genes that contribute to pathogenicity in infected mice (Figueroa-Bossi and Bossi 1999). Another way to pose

the question is to compare the behavior of the same bacteriophage in both organisms. One ideal phage for this is bacteriophage Mu, which can infect both organisms.

In Salmonella, bacteriophage Mu is much less prone to respond to induction

stimuli and the fraction of cells surviving induction is orders of magnitude

higher than in E. coli (Champion and Higgins 2007). Thus, reduced supercoiling

might have been a mechanism to reduce the toxicity of prophage in the

Salmonella genome.



7  Molecular Structure and Dynamics of Bacterial Nucleoids



141



7.13.2 How Does One Prove/Disprove Loops in DNA?

The most popular theory about domain structure is that the proteins involved in

creating a domain form DNA loops in vivo Fig. 7.1 (Cook 2003; Luijsterburg et al.

2006). Electron micrographs of nucleoids from the 1970s are one reason loops

originally seemed reasonable. These images illustrated a rosette of interwound

supercoiled loops with a rather amorphous central region. However, alternative

explanations to looping have been discussed in a recent review (Higgins et  al.

2005). Biochemical studies show that two of the NAPs, H-NS and FIS can loop

DNA under some circumstances (Dame et  al. 2006; Skoko et  al. 2006). The

MukBEF complex compacts DNA with a mechanism that appears to stimulate

formation of positive chiral nodes (Petrushenko et  al. 2006) and MukBEF has a

non-specific DNA binding affinity that could stabilize 100–500 loops. However,

genetic experiments also show that the structure of the chromosome remains intact,

even when many of the potential looping elements are eliminated by mutation.

Must all domainins organize DNA loops that are cross-linked by proteins at the

loop base (Fig. 7.1)?

Currently, no solid evidence exists for long range (10 kb) DNA loops in vivo.

There are two problems that make testing loops experimentally difficult. First, stochastic boundary elements are very likely created by multiple mechanisms, which

makes identifying specific candidates difficult and proving a looping mechanism

in  vivo extremely difficult. One could carry out experiments comparable to the

chromosome conformation capture technique that shows eukaryotic chromatin

loops (Dekker 2006), but what are the control predictions of a stochastic pattern?

MukBEF, H-NS and FIS are three prime domainin candidates, but mutations disrupting H-NS and FIS show a only modest nucleoid phenotype in tests of supercoil

structure in vivo (Hardy and Cozzarelli 2005), and cells still form nucleoids in the

absence of MukBEF (Danilova et al. 2007).

Second, non-looping mechanisms clearly influence supercoil dynamics. Lynch

and Wang found that two group of genes become tethered or handcuffed to the membrane by transcription coupled translation. This process restricts rotation of the DNA

and can lead to hypersupercoiling of plasmids (Lynch and Wang 1993). One class of

genes includes tetA, lacY, and phoA, which are integral membrane proteins that are

inserted into the membrane via the SecA pathway. A second class of genes that

become membrane attached include tolC and ampC, which are proteins exported

through the cytoplasmic membrane to the outer membrane. These genes are not a

minor genome fraction. Of all protein-encoding genes in E. coli and Salmonella,

25% are membrane proteins and 10% are proteins that become exported through

the cytoplasmic membrane to the periplasm and outer membrane (Baars et al. 2008;

Daley et  al. 2005; Rey et  al. 2005). Each may be a transient or stable barrier to

supercoil diffusion depending on transcription/translation kinetics. Although many

of these are not highly transcribed, some are and in toto they represent 1,500 sites

in distributed throughout the genome that can restrict supercoil diffusion as a tether

between DNA and the membrane.



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7.13.3 Are Other Nucleoid Organizing Proteins Barriers

to Horizontal Gene Transfer?

GyrB is a strong barrier to horizontal gene transfer from Salmonella to E. coli. Even

relatively minor expression of the Salmonella GyrB in E. coli, which results in less

than 10% of normal GyrB expression, is under strong negative selection (Champion

and Higgins 2007). Moreover, introducing a single nucleotide substitution encoded

by the Salmonella gyrB652 mutation, or swapping the entire E. coli gyrB allele with

the WT sequence from Salmonella is lethal. Thus, gyrB and all linked genes

(including the oriC initiation region) should be excluded from recombination with

E. coli genome, even when nucleotide restriction barriers are suppressed. Oddly,

the reciprocal experiment of replacing the Salmonella gyrB locus with the E. coli

sequence is easy to carry out. Recent evidence from gaps in sequence assembly of

shotgun bacterial plasmid cloning projects shows that a surprising group of genes

can act as barriers to horizontal transfer. The genes include the replication initiators,

some NAPs, tRNA synthetases, outer membrane proteins, and many ribosomal

proteins from close relatives of E. coli (Sorek et  al. 2007). Some, but not all, of

these inhibitory effects are caused by multi-copy expression problems. However,

the only full-length genes tested for creating problems in the sequencing projects

are small due to the size restrictions of 1 kb for shotgun cloning strategies. How

many of the large proteins are intra-species restricted at low copy is a new biochemical and evolutionary question to think about.

Finally the comparison of segregation mechanics of E. coli and Salmonella

demonstrates that intricate cooperation between different types of enzymes is necessary to complete DNA untangling of converging replication forks. DNA gyrase,

Topo IV, FtsK, MukBEF, XerCD, and the dif site and KOPS sites are elements we

know about so far. The lessons from bacteria will surely apply to the eukaryotic

world because untangling DNA strands at points of replication fork convergence is

every chromosome’s problem. It seems improbable that a eukaryotic untangling

process, which must occur at thousands of sites around the genome, will involve

fewer coordinating components than are used to untangle the single chromosomes

of most bacteria.

Acknowledgement  Work in the Higgins Laboratory is supported by grants from the NSF –

MCB9122048 and NIH – RO1GM33143.



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9 Post-DNA Replication Nucleoid Formation; Slow and Fast Doubling Times

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