Organization of supercoil domains and their reorganization by transcription

doi:10.1111/j.1365-2958.2005.04796.x

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Mol Microbiol. Author manuscript; available in PMC 2006 February 24.

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Mol Microbiol. 2005 September; 57(6): 1511–1521.

doi: 10.1111/j.1365-2958.2005.04796.x.

PMCID: PMC1382059

NIHMSID: NIHMS5073

Organization of supercoil domains and their reorganization by transcription

Shuang Deng, Richard A. Stein, and N. Patrick Higgins^*

Department of Biochemistry and Molecular Genetics, University of Alabama at Birmingham, Birmingham, AL-35294, USA.

*For correspondence. E-mailnphiggin/at/uab.edu; Tel. (+1) 205 934 3299; Fax (+1) 205 975 5955.

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Summary

During a normal cell cycle, chromosomes are exposed to many biochemical reactions that require specific types of DNA movement. Separation forces move replicated chromosomes into separate sister cell compartments during cell division, and the contemporaneous acts of DNA replication, RNA transcription and cotranscriptional translation of membrane proteins cause specific regions of DNA to twist, writhe and expand or contract. Recent experiments indicate that a dynamic and stochastic mechanism creates supercoil DNA domains soon after DNA replication. Domain structure is subsequently reorganized by RNA transcription. Examples of transcription-dependent chromosome remodelling are also emerging from eukaryotic cell systems.

Introduction

A working model of the bacterial chromosome has been a long-standing goal of molecular biology. Once the outlines of replication and transcription were revealed in the late 1960s, the physical implications of Watson and Crick’s structure raised basic questions about chromosome mechanics (Maaloe et al., 1966). Resolving many of the most basic questions about chromosome behaviour proved to be a daunting challenge [see the review by Higgins et al. (2005)]. Recent discoveries in Escherichia coli (Postow et al., 2004), Salmonella typhimurium (Deng et al., 2004; Stein et al., 2005), Caulobacter crescentus (Viollier et al., 2004; Gitai et al., 2005) and Bacillus subtilis (Britton et al., 1998; Lemon and Grossman, 1998; Dworkin and Losick, 2002; Lindow et al., 2002; Wu and Errington, 2003; 2004) move us closer to a composite view of dynamic chromosome behaviour, and make it easier to test and understand the 270-odd sequenced bacterial genomes in the current NCBI database.

If fully extended, the 4.6 million base pair E. coli chromosome would stretch over 1 mm. How does a 2-μm-long bacterium condense DNA 1000-fold and still allow the dynamic DNA movement necessary for RNA transcription and DNA replication to proceed without everything getting snarled (Holmes and Cozzarelli, 2000)? A partial solution to the packing problem is negative supercoiling, which causes the double helix to adopt a branched and plectonemic or interwound structure. Cells treated with lysozyme and mild ionic detergent release non-viscous ‘nucleoids’¹ that can be analysed by sedimentation through sucrose gradients and by electron microscopy (Worcel and Burgi, 1972; Giorno et al., 1975; Kavenoff and Ryder, 1976). The unfolded structure reveals an interwound DNA conformation. Sensitive and non-disruptive topological tests of plasmid DNA structure that are produced by recombination with the lambda integrase confirmed the presence of interwound DNA in vivo (Bliska and Cozzarelli, 1987).

The chromosome superhelix density for mid-log phase cultures of E. coli is about σ= −0.06, where σ= ΔLk/Lk₀ (Bauer et al., 1980; Drlica and Rouviere-Yaniv, 1987; Pettijohn, 1996). This value results from a dynamic equilibrium established by competing activities of DNA gyrase, which introduces negative supercoils, and two enzymes that remove negative supercoils – TopoI (Menzel and Gellert, 1983) and Topo IV (Zechiedrich et al., 2000). The supercoil density of chromosomal DNA must exist in a window of ±20% of the normal mean value for cell growth (Drlica, 1992). Hypo-supercoiling creates problems in segregation (Steck and Drlica, 1984; Hiraga et al., 1989; Holmes and Cozzarelli, 2000; Sawitzke and Austin, 2000) and leads to poor chromosome function that is either toxic or lethal (Zechiedrich et al., 1997). Hyper-supercoiling promotes the formation of Z-DNA at positions with long alternating GC or AT base pairs, causes the extrusion of cruciforms at inverted repeats, and stabilizes other unusual structures such as intramolecular triplexes (H-DNA), intermolecular triplexes and R-loops. These unusual structures impede RNA polymerase, slow or stall DNA replication forks, and create substrates for endonucleases and Holliday junction resolving enzymes (Higgins and Vologodskii, 2004).

In addition to compacting DNA, negative supercoils are dynamic. The slithering and branching of the interwound strands allow DNA to act like a chaperone, promoting the long-range assembly and disassembly of protein–DNA complexes (Higgins et al., 2005). Furthermore, negative supercoils facilitate the transition from duplex- to single-stranded conformation, a state through which DNA replication, transcription and recombination all proceed (Funnell et al., 1986; Artsimovitch, 2005; Kuzminov and Stahl, 2005).

Domain numerology

A supercoil domain can be defined as the segment of DNA relaxed by the introduction of a single- or double-strand break (Pettijohn, 1996; Postow et al., 2004). In vivo, single-strand breaks occur through a variety of mechanisms (Strauss, 2005). These include chemical damage from oxygen radicals, alkylation and depurination by chemicals generated by endogenous and exogenous mechanisms, attack from cellular endonucleases and the process of semi-discontinuous DNA replication. Because supercoiled DNA strands rotate rapidly (Oram et al., 1994), large chromosomes are partitioned into small units to shield the genome from catastrophic negative superhelical loss. Early DNase I nicking studies of E. coli nucleoids by Worcel and Burgi showed that multiple breaks were required to relax a chromosome fully in vitro; the estimated number of domains was 4–30 (Worcel and Burgi, 1972). Using ³H-tri-methylpsoralen binding as a probe of supercoil structure in vivo, Sinden and Pettijohn treated cells with X-rays to introduce breaks and estimated that E. coli contained 40 domains per genome equivalent (Sinden and Pettijohn, 1981). Recent studies indicate the presence of 400 domains [for an explanation of why the numbers differ, see the review by Higgins et al. (2005)].

A method developed in 1996 permitted the examination of domains in defined segments of the large bacterial chromosome using site-specific recombination as an assay. This test involved less structural perturbation than either physical extraction of DNA or X-irradiation (Higgins et al., 1996). S. typhimurium chromosomes marked with pairs of directly repeated res sites spaced at different intervals were exposed to the controlled induction of Tn3 or γδ resolvases. Unlike recombination catalysed by λ Int, phage P1 Cre, or yeast 2μ Flp recombinases, which pair sites by random DNA collisions, the resolvase system relies on negative supercoiling to precisely align two 140 bp res sites (Stark and Boocock, 1995). Two res sites can recombine and mark a chromosome with a deletion of non-essential genes if and only if they reside in the same supercoil domain (Fig. 1A

). Three conclusions from this study challenged the conventional notions of chromosome structure. First, a distance penalty for recombination efficiency approximated first order decay. This suggested cell-to-cell variation in domain location. Second, the slope of the first order decay revealed that resolution efficiency decreased by 50% for each 15–17 kb distance separating a pair of res sites (a term called the 1/2D). Assuming that two points in a chromosome form a loop (see below) and that barrier placement has a Poisson distribution, the mean domain size was estimated to be 20 kb, which leads to an estimate three to four times larger than the 40 domain structure predicted by previous methods (Higgins et al., 1996). Third, domains appeared and disappeared over time. Surprisingly, chromosome movement was more dynamic in stationary-phase cells than in exponential log phase cells (Higgins et al., 1996; Staczek and Higgins, 1998; Higgins, 1999).

Fig. 1

Assays measuring supercoil domain structure in Salmonella and E. coli

Recently, Cozzarelli’s laboratory devised a new method for enumerating domains in E. coli (Postow et al., 2004). Negative supercoiling is a product of transcription as well as a factor that can modulate increased or decreased output from specific promoters (McGovern et al., 1994; Willenbrock and Ussery, 2004; Travers and Muskhelishvili, 2005). The Postow method takes advantage of 306 genes that are widely distributed in the genome and which respond rapidly and reliably to supercoil relaxation. After DNA relaxation, a third of the supercoil-sensitive genes (SSGs) become induced for transcription and two-thirds respond with transcriptional repression (Peter et al., 2004). By controlling the in vivo expression of type II restriction enzymes, Postow made double-strand breaks at precise chromosomal locations. The distance from a SwaI site to the promoter of an SSG was combined with microarray expression patterns before and after chromosomal cleavage. The best-fit model of the E. coli chromosome using SSG transcription data consisted of variable loops, random barrier positions and a 10 kb mean domain size. This model predicts ~450 domains per genome equivalent (Postow et al., 2004).

Support for a 400-domain chromosome came from two sources. First, tracings of EM images of gently lysed nucleoids showed that loops ranged from 2 to 66 kb and the best-fit model supported variable loops with an 11 kb mean (Postow et al., 2004).

Second, Postow’s domain estimates in E. coli agreed with new results in S. typhimurium, which developed from studies aimed at understanding how domains change over time (Scheirer and Higgins, 2001; Deng et al., 2004). Stein et al. discovered that once the WT Tn3 resolvase is produced, it remains active inside cells for more than 3 h. Because short-lived domains would be invisible to such a long lasting enzyme, Stein worked out a method to modulate the time-span of cell exposure to γδ resolvase. An 11-amino-acid extension was added to the C-terminus of Res protein (Res-SsrA). The extension corresponds to a sequence that is normally appended to proteins by the SsrA or Tm RNA trans-translation system (Stein et al., 2005). The tag had no effect on the enzyme’s catalytic efficiency or mechanism, but it made the resolvase an efficient substrate for ClpXP degradation (Keiler et al., 1996; Hayes et al., 2002a,b). By modifying selected residues in the tag (Flynn et al., 2001), Stein generated five resolvases that persist for periods varying from 5 min to several hours. Over a 100 kb chromosomal segment, domain structure changed over a 3 h window. Assayed for a 10 min period in exponential growth, the mean domain size was 11–13 kb. Thus, like E. coli, Salmonella has ~400 domains per genome equivalent, and the agreement between three different techniques measuring domain sizes in two species of bacteria is striking (Fig. 1C

When do supercoil domains form? The process of DNA replication requires all DNA binding proteins and all topological impediments to unwinding of the helix by the DnaB helicase to be removed to allow efficient DNA synthesis. DNA replication proceeds at the rate of 800 bp per second per fork, and supercoils behind the fork are lost because of discontinuities in strand synthesis (Hingorani and O’Donnell, 2005). Peter showed that domains must reform rapidly after DNA replication. He used the 306 super-coil reporter SSGs mentioned above to measure how fast domains form after replication (Peter, 2000). Because there was no significant transcription change in the SSGs following replication, Peter concluded that domains reformed so rapidly that RNA polymerase had no time to synthesize significant levels of RNA before supercoil density was restored. The post-replication period is undoubtedly the time when most of the 400 stochastic supercoil domains are established.

Domains remodelled by transcription

In 2001, Scheirer reported that transcription from a phage promoter caused the appearance of a supercoil diffusion barrier in a Mu prophage (Scheirer and Higgins, 2001). To follow up, Deng created test intervals at several locations in the Salmonella chromosome (Deng et al., 2004). The impact of transcription was tested using a module derived from Tn10. Constitutive transcription of tetA, the gene for an integral plasma membrane tetracycline efflux pump, created region-specific barriers to γδ recombination and reduced resolution by 20-fold in a 14 kb interval at several chromosome locations. In plasmid studies, transcription anchors DNA to the cell membrane by cotranscriptional translation, and this attachment enhanced topological effects on DNA (Lynch and Wang, 1993). However, in the Salmonella chromosome, two cytoplasmic proteins, β-galactosidase (β-gal) and aminoglycoside-3′-O-phospho-transferase, the product of the Tn5 kan gene, reduced resolution as much as transcription of tetA. Thus, membrane attachment was not required to generate supercoil barriers in a wild-type (WT) Salmonella chromosome.

How does transcription remodel chromosome structure? X-ray crystal structures of RNA polymerase show that DNA enters RNA polymerase through one channel and RNA emerges from another (Perederina et al., 2004; Geszvain and Landick, 2005). Thus, transcription would either remove any previously established DNA attachments or move them along the DNA and out of the path of RNA polymerase. This would generate an impediment free zone for RNA synthesis and possibly pile up barriers near the transcription terminus. What does a transcription-generated domain look like? Two rules suggest a loop. First, any gene positioned between a pair of res sites always inhibits resolution once the gene is highly transcribed. Second, when a pair of res sites lies downstream of a highly transcribed gene, resolution is inhibited only when one res site is within 1 kb of the transcription terminator sequence of the highly transcribed gene (S. Deng and N.P. Higgins, unpubl. data). A loop that includes the transcribed sequence plus a little extra DNA (average about 500 bp) is the simplest model that explains this pattern.

A bonus in the Salmonella transcription experiments came from discovering the time-course for the formation and dissolution of a transcription domain. Using a short-lived resolvase discussed above, cells were induced to transcribe tetA by adding chlorotetracycline (CLT). The maximum impact of the transcription on inhibiting resolution required 20 min of cell growth. Conversely, when maximally induced cultures were washed free of CLT, barriers to resolution disappeared over a 20 min interval. This shows that domain formation and dissolution does not coincide with transcription; transcription domains persist for a length of time roughly equal to the time for a new wave of replication to reach dichotomously replicating chromosomes. However, once high transcription established a domain, it persisted, representing a type of topological memory.

Is transcription memory functionally useful?

New methods allow the analysis in variation of expression at the single cell level (Rosenfeld et al., 2005). The mean lac operon expression profile of growing cultures spans a several hundred-fold range. However, all cells do not respond alike, and in most individual cells (Fig. 2A

), lac expression behaves like a switch that modulates between low and high expression states (Novick and Weiner, 1957). Ozbudak found cell populations to behave hysteretically over a broad range of inducer levels (Ozbudak et al., 2004). With E. coli cultures started from cells that were un-induced for lac expression (Fig. 2B

, bottom panel), growth for 20 h in different inducer concentrations yielded reproducible bi-stable cell populations with high or low GFP expression. With cell cultures initiated from a maximally induced cell population (Fig. 2B

, top panel), the same inducer concentrations resulted in higher mean cellular expression levels and bi-stable populations containing much higher frequencies of high expression cells. Hysteresis shows that bacterial cultures ‘remember’ the initial growth conditions. One mechanism contributing to hysterisis involves the LacY permease, which concentrates inducer by actively transporting it into cells. However, the formation of a stable free-running transcription domain may also contribute to efficient transcription persistence in cells that achieve induction.

Fig. 2

Hysteresis and bistability in single cell transcription profiles

Transcription memory may contribute to the expression of operons that lack a repressor control system. The bgl operon has no repressor but changes expression in response to DNA structure and TopoI and gyrase activity (DiNardo et al., 1982; Manna et al., 2001). There are also interesting genes like hipA (Moyed and Bertrand, 1983; Balaban et al., 2004), which cause a variable fraction of cells to stop growth and thereby survive exposure to antibiotics. Transcription memory could provide the epistatic mechanism to generate lineages with different expression states as minor cellular subpopulations.

Eukaryotic cells also move chromosome proteins with RNA polymerase and mark genes with transcription memory signals. One example in budding yeast is cohesin. Following DNA replication, chromosomes are decorated with cohesin, which has three important roles in cell division. In mitotic yeast cells, cohesin maintains a contact between sister chromatids after DNA synthesis, and it promotes attachment of chromosomes to the spindle. In meiosis, cohesin produces cohesion between sister chromatids that is necessary for efficient homologous recombination (Ross and Cohen-Fix, 2004). New studies using ChIP-on-chip technology showed that cohesins are dispersed by RNA polymerase from specific loading sites to positions near the 3′ end of transcribed genes (Glynn et al., 2004; Lengronne et al., 2004; Ross and Cohen-Fix, 2004). Another example is transcription-mediated nucleosome replacement. Histones deposited during DNA replication are naturally stable. However, if a Drosophila gene is expressed after replication, nucleosomes containing the histone H3 subunit disappear and are replaced with nucleosomes containing the histone variant H3.3 (Schwartz and Ahman, 2005). Third, the condensin proteins, closely related in structure to cohesins, mediate dosage compensation of sex chromosomes and serve as bookmarks for the inducible expression of heatshock genes (Xing et al., 2005).

Transcription neighbourhoods

How many transcription domains exist in the average bacterial cell? Deng tested the relationship between transcription activity and resolution efficiency in Salmonella (Deng et al., 2004). Using a strain with lacZ regulated from the tetA promoter with a WT TetR repressor, un-induced cultures gave 15 units of β-galactosidase activity and the recombination efficiency (1/2D) was comparable to previously analysed stochastic neighbourhoods with a mean 12 kb domain. Cells incubated with 5 μg ml⁻¹ CLT generated >600 units of β-gal and the resolution efficiency of this region fell 10-fold. The measured RNA:DNA ratio of lacZ message for cells expressing 15 and 600 units of β-gal was 0.45 and 15 respectively (Wei et al., 2001). With the microarray scale it was possible to predict transcription effects genome-wide using RNA:DNA ratios for genes in Salmonella or E. coli.

Most (~70%) of the >4200 genes with measurable expression in E. coli or Salmonella give an RNA/DNA signal of less than one (Wei et al., 2001; Bernstein et al., 2002). In fact, all organisms for which RNA abundance has been studied including yeast, mouse and humans show the same genome-wide transcript structure; >70% of all eukaryotic protein genes contain steady-state RNA levels of less than one molecule per cell (Kuznetsov et al., 2002; Blewitt et al., 2004). In Gram-negative bacteria, most mRNAs turn over with a half-life of 3–4 min, so the transcription rate of 70% of the chromosome must be less than one message per 3 min, including 30% of the known essential genes. For 27% of E. coli genes, the steady-state RNA:DNA ratio is less than four, which would hinder resolution by less than 30%. Thus, 97% of bacterial genes have little or no transcriptional impact on domain structure. This confirms results obtained by French using electron microscopy to locate transcribing RNA polymerase molecules in E. coli DNA. The images from this study showed long stretches of DNA with an occasional single polymerase and very rare zones with ‘Christmas trees’ of multiple polymerases with short branches at the operon start and long branches near the operon 3′ end (French and Miller, 1989).

Only about 110–130 (2.5%) of E. coli genes located at 50 positions would inhibit resolution of a 14 kb interval by twofold or more, and only 25 protein encoding genes at about 15 locations have mRNA:DNA ratios of >10, which inhibits resolution by fivefold or more. High transcription zones in bacterial chromosomes are spaced at intervals (Allen et al., 2003; Lobner-Olesen et al., 2003; Jeong et al., 2004; Manna et al., 2004) and if the stable RNA genes, which include seven ribosomal operons and several tRNA gene clusters, are included, only ≈30 sites would show region-specific domains with fivefold or greater effects on resolution. Specific regions in Salmonella with highly transcribed non-essential genes were tested for agreement between the steady-state microarray RNA:DNA ratio and the resulting impact on resolution. The results validated microarray expression data for predicting where region-specific domains can be found [for specific examples see supplementary data in the study by Deng et al. (2004)].

Domain barrier elements

With a 400-domain chromosome and transcription causing formation of region-specific domains, two questions arise. First, what makes a domain? One hypothesis is that barriers are simple knots and tangles in DNA (Higgins et al., 1996), but the stability of domains made by transcription suggests that proteins are involved. A set of small structural proteins plus a handful of nucleotide cofactor-driven enzymes play key roles in chromosome behaviour. Small abundant ‘nucleoid-associated proteins’ (NAPs) act as architectural components that shape DNA. These proteins are often present at concentrations exceeding the level needed to saturate their known high-affinity sites, and they contribute to global chromosome structure by binding DNA ‘non-specifically’ (Crozat et al., 2005; Johnson et al., 2005). These proteins include HU, IHF, H-NS, StpA, FIS, LRP, SeqA and DPS. Each protein was originally discovered through its ability to condense DNA or to stimulate complex DNA transactions like transcription, transposition and site-specific recombination. Johnson et al. recently reviewed the biochemical properties as well as structural models of X-ray DNA-cocrystals for most of these proteins (Johnson et al., 2005).

Other than RNA and DNA polymerases and the type I and type II topoisomerases mentioned above, three catalytic enzymes channel chromosome motion in living cells. A three-protein complex of MukB, MukE and MukF (Muk-BEF) (Yamazoe et al., 1999) is a member of a highly conserved structural maintenance of chromosome (SMC) proteins. This family includes condensins and cohesins. MreB is an actin-like protein involved in moving chromosomal DNA after replication (van den Ent et al., 2001), and FtsK is an ATP-driven DNA conveyer (Aussel et al., 2002; Pease et al., 2005).

What are the most likely proteins for controlling formation of loop domains? Four proteins warrant special mention: DNA gyrase, MukBEF, FIS and HNS. DNA gyrase re-establishes a proper value of σ and it also controls the mean domain number. Certain gyrase hypomorphs have more than twice the WT number of domains (Staczek and Higgins, 1998). Stein confirmed this result with short-lived resolvases and showed that some gyrase hypomorphs limit resolution to segments much shorter than 12 kb (Stein et al., 2005). The topological impact of severe gyrase hypomorphs is most dramatic at the terminus of DNA replication. This can be seen as a large effect on Tn3 resolution in the region that lies adjacent to the dif site (Z. Pang, R. Chen and N.P. Higgins, submitted). Microarray studies also show strong effects of gyrase mutants on transcription profiles near the terminus (Jeong et al., 2004).

The E. coli and Salmonella MukBEF complex is a prime candidate for stabilizing loop domains in chromosomes. SMC proteins are conserved across the bacteria, archaea and eukaryotic kingdoms and this family includes cohesins and condensins. SMC proteins have globular amino- and carboxyl-terminal domains separated by long coiled-coil regions with a central flexible hinge (Jessberger and Gasser, 1998). SMC proteins form antiparallel, interwound dimers that hydrolyse ATP and bind DNA at each end (Hirano and Hirano, 1998; Hirano et al., 2001). Although not enough MukB is present in E. coli cells to account for 400 domains, this protein could account for the transcription-dependent domains. The movement of RNA polymerase would reduce negative supercoiling ahead of RNA polymerase, thereby creating a high affinity site for gyrase (Higgins and Cozzarelli, 1982; Moulin et al., 2005). A complex involving both MukBEF and gyrase could stabilize a loop and facilitate transcription by relieving the supercoil flux generated by high level transcription. In other organisms, SMC complexes like MukBEF are involved in chromosome condensation, dosage compensation of transcription, DNA repair and recombination (Cobbe and Heck, 2000; Holmes and Cozzarelli, 2000; Hirano, 2002).

Two NAPs make DNA loops in vitro. Electron microscopy studies show that H-NS and FIS both stabilize branched loops through cooperative interactions when mixed with plasmid DNA at appropriate concentrations (Dame et al., 2000; Schneider et al., 2001). Interestingly, both proteins have been implicated in global chromosome structure in several assays. H-NS influences both the frequency (Falconi et al., 1991) and the location (Swingle et al., 2004) of transposon insertions in the bacterial chromosome. H-NS is present at 10 000–20 000 copies per cell (Azam et al., 1999), and deletions of hns alter expression of about 5% of E. coli proteins (Hommais et al., 2001). FIS also has intriguing properties. In early log phase, FIS is abundant (30 000 molecules per cell), and it is the only NAP that disappears when cells enter stationary phase (<1000 molecules per cell). As domain barriers decrease in stationary phase, this is an intriguing correlation. FIS has also been implicated in the regulation of both the gyrA and the gyrB genes (Schneider et al., 1997, 1999; Keane and Dorman, 2003)

Finally, Hardy devised a genetic screen to find chromosomal domain barrier genes (Hardy and Cozzarelli, 2005). Modules with supercoil-responsive promoters were introduced into the E. coli chromosome at two locations. One promoter was linked to β-lactamase and the other controlled β-gal expression. Chromosome supercoiling relies on domains to maintain the mean chromosomal value of σ, but plasmids are single molecules without domain boundaries. Mutations were sought that alter chromosome supercoiling while leaving the plasmid σ-values at the normal mean value. Transposon mutagenesis uncovered five non-essential genes that appear to contribute to chromosome domain structure. The screen/selection uncovered H-NS and Fis mutations, which are in the category of expected genes (see above), plus three unexpected genes. These ‘new’ domain candidates are tktA, a transketolase, pgm, phosphoglucomutase and dskA, a multicopy suppressor of dnaK and mukB knockouts. Deletions of each of these genes showed reproducible effects on the supercoil-responsive reporters in the chromosome but did not change the superhelical density of pBR322-derived plasmids in the same cell. How these new genes alter chromosomal DNA structure has yet to be explained.

Conclusions

Some scientists have been reluctant to accept the notion that variable domain organization is good design for chromosomal DNA. Nonetheless, stochastic events in cell development are not rare (Rosenfeld et al., 2005), and variation is an inherent aspect of regulated gene expression in all tissues and organisms (Blewitt et al., 2004). The key property of bacterial chromosome compaction is pliability. Replication takes place within a DNA ‘factory’ and temporal rules for initiation are predictable for different growth rates. Transcription changes in response to internal and external environment, and during cotranscriptional translation, insertion of a protein into the membrane or periplasm will tether its gene to the cell membrane. As most of the DNA is rarely transcribed, workable units with variable divisions are acceptable most of the time. For the 3% of DNA with very active transcription, what is made during replication can be reworked to accommodate efficient gene transcription, even though life in the fast lane requires both RNA and DNA synthesis to occur simultaneously at different locations and at 10-fold different rates. Thus, a bacterial chromosome requires a statistical view in lieu of the highly ordered X-ray crystal structures of proteins that do the biochemical work on a chromosome.

Above the stochastic 10 kb domain level, long range order is beginning to come into focus. Using fluorescent in situ hybridization or fluorescent DNA binding proteins with specific DNA binding sites, specific genetic regions have been located and tracked in growing cell populations (Glaser et al., 1997; Gordon et al., 1997; Niki and Hiraga, 1998; Lemon and Grossman, 2000; Niki et al., 2000; Roos et al., 2001; Li et al., 2002; 2003; Lau et al., 2003) and by using random collision site-specific reactions catalysed by the phage λ Int and Xis proteins with attR and attL sites placed in the genome as unique locations, Garcia-Russell showed that some segments of chromosome do not often make intimate contact with each other (Garcia-Russell et al., 2004). In a similar study of E. coli with many endpoints, Valens et al. found a distance rule for lambda attL X attR recombination that indicated colocalization of very large regions (Valens et al., 2004). Furthermore, an amazing structure was revealed in C. crescentus using ‘tagged’ chromosomes with tet- and lac-operator modules bound with TetR-cyan fluorescent protein and LacI-yellow fluorescent protein (Lau et al., 2003). Foci were resolved in 112 different strains, using thousands of images for each strain (Viollier et al., 2004). Every tagged region moved to a replication factory and then to a post-replication position in sister cells that reflected genetic linkage to the origin or terminus of the chromosome. More recently, using chromosomes marked at two sites close to the dif site, Wang et al. demonstrated an intriguing genome organization in E. coli (Wang et al., 2005). For slow growing cells, sites separated by 150 kb move independently to the centrally located replication factory, and then to a unique location of the replicated nucleoids. Moreover, sites near the terminus in different replichores moved to opposite edges the nucleoid. These studies show that the left and right replichores, which have either near-continuous or discontinuous synthetic patterns, also have different traffic patterns in living cells. High fidelity chromosome positioning has been equated to geologic stratification or stabilization after deposition. For the chromosome, stability may be underpinned by numerous 10 kb domains (Breier and Cozzarelli, 2004). Whatever accounts for 400 domains and the longer range order in bacterial genomes, combinatorial mutation schemes will be required to pin down domain size phenotypes and more unpredictable players like tktA and pgm seem likely to emerge.

Acknowledgments

We thank Nick Cozzarelli and Christine Hardy for sharing data before publication. Lisa Postow generated the graph in Fig. 1C

, and David Sherratt provided prepublication data on nucleoid structure in E. coli. We thank members of the Higgins lab for comments and criticism of the manuscript. Work in the NPH lab was supported by grants from the National Institutes of Health – GM33143 and the National Science Foundation – MCB9122048.

Footnotes

¹The term nucleoid is also used in reference to a subcellular region of a bacterium that stains with DNA-binding drugs like DAPI (Bohrmann et al., 1991; Cunha et al., 2001).

Note added in proof

In the brief time since this manuscript was submitted, two papers appeared with a focus on the long-range organization of the E. coli chromosome. 1. Bates, D., and Kleckner, N. (2005) Chromosome and replisome dynamics in E. coli: loss of sister cohesion triggers global chromosome movement and mediates chromosome segregation. Cell 121: 899–911. 2. Elmore, S., Muller, M., Vischer, N., Odijk, T., and Woldringh, C. (2005) Single-particle tracking of oriC-GFP fluorescent spots during chromosome segregation in Escherichia coli. J Struct Biol 136: 53–66.

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