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Genes to Cells (2007) 12, 1141-1152. doi:10.1111/j.1365-2443.2007.01125.x
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Transcription-coupled nucleoid architecture in bacteria

 Ryosuke L. Ohniwa1,*, Kazuya Morikawa2, Sayaka L. Takeshita2, Joongbaek Kim1, Toshiko Ohta2, Chieko Wada1,3 and Kunio Takeyasu1

1 Laboratory of Plasma Membrane and Nuclear Signaling, Kyoto University, Graduate School of Biostudies, Yoshidahonmachi, Sakyo-ku, Kyoto 606-8501, Japan
2 Institute of Basic Medical Sciences, Graduate School of Comprehensive Human Sciences, University of Tsukuba, Tennoh-dai, Tsukuba 305-8575, Japan
3 Yoshida Biological Laboratory, Yamashina, Kyoto 606-8081, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
The circular bacterial genome DNA exists in cells in the form of nucleoids. In the present study, using genetic, molecular and structural biology techniques, we show that nascent single-stranded RNAs are involved in the step-wise folding of nucleoid fibers. In Escherichia coli, RNase A degraded thicker fibers (30 and 80 nm wide) into thinner fibers (10 nm wide), while RNase III and RNase H degraded 80-nm fibers into 30-nm (but not 10-nm) fibers. Similarly in Staphylococcus aureus, RNase A treatment resulted in 10-nm fibers. Treatment with the transcription inhibitor, rifampicin, in the absence of RNase A changed most nucleoid fibers to 10-nm fibers. Proteinase-K treatment of nucleoids exposed DNA. Thus, the smallest structural unit is an RNase A-resistant 10-nm fiber composed of DNA and proteins, and the hierarchical structure of the bacterial chromosome is controlled by transcription itself. In addition, the formation of 80-nm fibers from 30-nm fibers requires double-stranded RNA and RNA–DNA hetero duplex. RNA is evident in the architecture of log-phase uncondensed and stationary-phase condensed nucleoids.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
The genomes of all living organisms are flexible higher-order structures and the transition can occur between functionally active and inactive states of gene replication and expression. In bacteria, genome DNA is packed into "nucleoids" (Robinow & Kellenberger 1994; Poplawski & Bernander 1997). Nucleoids contain a set of distinct DNA binding proteins, such as Hu and RNAs that (in conjunction with the physical and chemical properties of the nucleoid and cytosol) help organize bacterial DNA into higher-order structures (Hecht & Pettijohn 1976; Zimmerman & Murphy 1996; Murphy & Zimmerman 1997; Azam & Ishihama 1999).

Electron microscopy (EM) has revealed that the circular fibrous nucleoids in bacteria form the core of a rosette-like structure with interwoven loops extending radially (Kavenoff & Bowen 1976; Kavenoff & Ryder 1976; Sloof et al. 1983). Recently, we demonstrated that the fiber is not naked DNA (Kim et al. 2004; Takeyasu et al. 2004). When Escherichia coli, Staphylococcus aureus and Clostridium perfringens were lysed in physiological ionic strength buffer, fibers (~ 40 and ~ 80 nm wide), but not naked DNA, could be observed by atomic force microscopy (AFM). In E. coli, the loop structure composed of 80-nm fibers also appeared under the same lysis conditions. These results suggest the occurrence of a step-wise folding of the genome, in which the 40-nm fibers of each loop hanging from the nucleoid core become 80-nm fibers.

Biochemical studies showing reduced sedimentation rate of the isolated nucleoid after RNase treatment have suggested that RNA is situated at the boundaries of nucleoid folding in the core region (Worcel & Burgi 1972; Pettijohn & Hecht 1974). EM studies have shown that the nucleoid shrinks in response to rifampicin treatment, suggesting that RNA anchors the nucleoid to the cell membrane (Dworsky & Schaechter 1973; Kleppe et al. 1979; Vos-Scheperkeuter & Witholt 1982). In apparent contradiction, dispersed nucleoids can be observed when cultured cells are stained by DAPI (4',6-diamino-2-phenylindole) after addition of rifampicin (Sun & Margolin 2004; Kruse et al. 2006). Taken together, these results suggest that nascent RNA stabilizes nucleoid structure (Zimmerman & Murphy 1996).

The involvement of RNA in the architecture of 40–80 nm fibers is suggested by the thinning of E. coli nucleoid fibers after RNase treatment (Ohniwa et al. 2007). In this study, we show that the RNA in the nucleoid fibers is nascent single-stranded RNA. Treatment of the E. coli and S. aureus nucleoids with RNase A (but not RNase III or RNase H) exposed the 10-nm fibers. These 10-nm fibers also appeared when E. coli was treated with rifampicin. The RNA in condensed nucleoids during stationary phase was single-stranded, suggesting that transcription-coupled mechanisms mediate the thickening of 10-nm fibers.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Nucleoid fibers thinner than 80 nm

It is critical to compare the sizes of nucleoid fibers in bacteria and eukaryotes to elucidate genome structures in general. We previously applied the "half maximum height (FWHM; full width at half-maximum)" method (Schneider et al. 1998) to estimate the dimensions of nucleoid fibers, and found nucleoid fibers with width of ~40 and ~80 nm in E. coli and S. aureus (Takeyasu et al. 2004). However, the considerations used to develop the FWHM method for analyzing AFM images were empirically and not theoretically based. Since the dimensions of molecules apparent by AFM depend on edge curvature and tilt angle of the cantilever, these must be taken into consideration to eliminate the "tip-effect" before the real dimensions can be determined. Here, using the EM image of the cantilever, we developed the "circular cone model" to correct for the tip-effect (see Experimental procedures and Fig. 1), and re-evaluated the width of nucleoid fibers. As a result, the diameters of 40- and 80-nm nucleoid fibers in both E. coli and S. aureus were re-estimated to be 30 and 80 nm, respectively (Fig. 2). We applied this method to thinner fibers to obtain their diameters.


Figure 1
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  		Figure 1  Circular cone model for the estimation of tip effect. (A) Relation between a cantilever tip and a sample when the sample size is smaller than the tip. (B) Relation between a cantilever tip and a sample when the sample size is larger than the tip. (C) Relation between a cantilever tip and a sample when the edge of tip is an ideal incircle. (D) AFM images showing the apparent widths of reconstituted 30 nm chromatin and nucleosomes (Hizume et al. 2004, 2005). Solid lines are obtained by Gaussian fitting, and the mean apparent widths are as follows: (i) for 30 nm chromatin, 58.0 ± 6.1 (mean ± SD) (the total number of observations, n = 30); and (ii) for nucleosomes, 33.6 ± 2.9 nm (n = 30). (E) AFM images of 10, 30 and 80 nm gold particles, and the relation between the measured (X-axis) and real (Y-axis) dimensions of the particles. Standard error bars are shown in the graph for a total number of the observations: n = 36 for 10 nm, n = 20 for 30 nm and n = 14 for 80 nm particles. A red dot on the graph (E) indicates the measured width of naked DNA (pRSFDuet-1, Novagen) (see Fig. 3G,H). Scale bars in the AFM images represent 500 nm.

 

Figure 2
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  		Figure 2  Comparison of the nucleoid fibers in E. coli and S. aureus. AFM images of lysed, log-phase (A) E. coli and (B) S. aureus cells. The distribution of nucleoid fiber widths using the "circular cone model" in (C) E. coli and (D) S. aureus. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (C) 31.9 ± 7.3 (mean ± SD) and 70.5 ± 12.3 nm (the total number of observations, n = 174); and (D) 34.0 ± 11.4 and 71.2 ± 13.3 nm (n = 188). Scale bars in the AFM images represent 500 nm.

 
RNA contribution to nucleoid architecture in bacteria

An "on-substrate lysis" procedure has been developed to visualize nucleoids in bacterial species by AFM (Kim et al. 2004; Morikawa et al. 2006). After the lysis of E. coli K-12 W3110 cells in log phase, their nucleoids were treated with 5 µg/mL RNase A for 5 min and observed by AFM. RNase A plus lysis treatment spread out more fibers around the cells than lysis treatment alone (Fig. 3A). RNase treatment of the isolated nucleoids decreased their sedimentation rate (Worcel & Burgi 1972; Pettijohn & Hecht 1974; Murphy & Zimmerman 2000), and RNA was localized at the site of nucleoid folding in the core (Worcel & Burgi 1972; Pettijohn & Hecht 1974). Therefore, the lack of RNA-clamps or placeholders in the nucleoid likely allowed the nucleoid fibers to unbind, spread and be easily released from the cell.


Figure 3
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  		Figure 3  RNase A treatment has a marked effect on the hierarchy of E. coli nucleoid organization. Lysed E. coli cells in log phase were treated with RNase A for 5 min (A, E), 30 min (B, F) and 120 min (C, G) at room temperature. (A, B, C) AFM images and (E, F, G) the widths of released fibers. (D) AFM image of naked plasmid DNA (pRSFDuet-1, 3829 bp, Novagen) and (H) the width of DNA. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (E) 7.9 ± 4.7 (mean ± SD) and 27.2 ± 2.5 nm (the total number of observations, n = 171); (F) 11.4 ± 3.9 and 26.5 ± 2.6 nm (n = 160); (G) 10.6 ± 4.6 and 25.7 ± 2.8 nm (n = 152); and (H) 1.3 ± 2.7 nm (n = 30). Scale bars in the AFM images represent 500 nm.

 
AFM images detected the 10-nm wide fibers that resulted from RNase A treatment (Fig. 3E). The 10-nm fibers were not naked DNA [diameter 1.4 ± 3.1 nm (mean ± SD), using the current "circular cone model" (Fig. 3D,H)]. Longer treatment with RNase A did not convert 10-nm fibers into naked DNA, and some 30-nm fibers persisted (Fig. 3B,C,F,G). These results suggest that RNA is a component of some thick fibers, and that the RNase-resistant 10-nm fibers without RNA appear to be the thinnest structural units of nucleoids in E. coli. Interestingly, no eukaryotic nucleosome-like "beads-on-a-string" structure was identified, indicating E. coli nucleoids do not seem to possess this structure.

Escherichia coli belongs to the Proteobacteria, and more than ten major nucleoid proteins of E. coli have been identified (Azam & Ishihama 1999). These proteins are mostly Proteobacteria-specific (Kim et al. 2004). Other bacterial species have much simpler nucleoid components. To investigate the general contribution of RNA to the bacterial nucleoid architecture, log phase nucleoids of S. aureus N315, which is a member of the Firmicutes and lacks various E. coli-type nucleoid proteins like IHF, Fis and H-NS (Kim et al. 2004), was also treated with RNase A. AFM of this nucleoid showed 10 and 30-nm wide fibers, but not the "beads-on-a-string" structure or 80-nm wide fibers (Fig. 4). In bacteria, since the thinnest observable fiber before RNase A treatment is 30 nm wide, RNA was expected to be essential to the assembly of higher-order fibers thicker than 10 nm.


Figure 4
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  		Figure 4  Effect of RNase A treatment on the hierarchy of S. aureus nucleoid organization. Lysed S. aureus cells in log phase were treated with RNase A for 30 min at room temperature. (A) AFM image and (B) the widths of released fibers. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (B) 10.0 ± 2.7 (mean ± SD) and 27.8 ± 7.1 nm (the total number of observations, n = 103). Scale bars in the AFM images represent 500 nm.

 
Structural dependence on nascent single-stranded RNA and nucleoid protein

RNase A digests mainly single-stranded RNA (Beintema & Kleineidam 1998). RNase III and RNase H specifically break down double-stranded RNA and RNA–DNA hetero duplexes, respectively. Even after 120-min RNase III and RNase H treatment at 37 °C, the 30-nm fibers remained intact (Fig. 5). Therefore, we concluded that the RNA species is single-stranded. However, since the 80-nm fibers disappeared with RNase III and RNase H treatments, we concluded that both double-stranded RNA and RNA–DNA hetero duplex are involved in the assembly of 80-nm fibers from 30-nm fibers.


Figure 5
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  		Figure 5  RNase III- and RNase H treatments do not influence 30-nm fiber units. Lysed E. coli cells in log phase were treated with RNase III (A, C) or RNase H (B, D) for 120 min at 37 °C. (A, B) AFM image and (C, D) the widths of released fibers. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (C) 25.3 ± 5.2 nm (mean ± SD) (the total number of observations, n = 101) and (D) 27.2 ± 8.9 nm (n = 160). Scale bars in the AFM images represent 500 nm.

 
In contrast, when log-phase cells were treated with rifampicin (100 µg/mL), which binds to RNA polymerase and inhibits the transcription, the 10-nm fibers appeared upon lysis (Fig. 6). Under our experimental conditions, transcription level was indeed reduced after the addition of rifampicin to the culture medium (Fig. 6A), and the number of 10-nm fibers increased (Fig. 6C–D). Thus, maintenance of the nucleoid structures containing fibers thicker than 10 nm is likely to be coupled to transcriptional activity. In particular, the lack of rapid turnover proteins, whose transcription and translation are inhibited by treatment with rifampicin, may affect nucleoid formation.


Figure 6
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  		Figure 6  Transcriptional activity of the genome correlates with nucleoid structure. (A) The transcriptional activity (in cpm) of rifampicin-treated cells in log phase was measured and normalized to the amount of cells used in the experiment (OD600). The transcriptional activity of treated cells relative to that of untreated cells is shown. Error bars represent standard deviation from three independent experiments. (B) AFM image of lysed E. coli treated with rifampicin for 30 min at 37 °C. Scale bars in the AFM images represent 500 nm. (C–E) The widths of the released fiber from the lysed cells cultured with rifampicin for (C) 5 min, (D) 30 min and (E) 60 min. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (C) 11.7 ± 3.1 (mean ± SD) and 26.0 ± 4.6 nm (the total number of observations, n = 101); (D) 12.3 ± 4.2 and 28.5 ± 3.7 nm (n = 101); and (E) 10.0 ± 4.5 and 25.7 ± 3.7 nm (n = 101).

 
When lysed log-phase cells were treated with proteinase K for 30 min, naked DNA-like fibers were released (Fig. 7A,C), suggesting that the 10-nm fibers are comprised of proteins and naked DNA without RNA. Among the nucleoid proteins, Hu is a strong candidate component of 10-nm fibers, because it is abundant in the log phase and present both in E. coli and S. aureus. Longer incubation with proteinase K did not change the distribution of fiber widths (Fig. 7B,D).


Figure 7
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  		Figure 7  The 10 nm-fiber consists of DNA and proteins. Lysed E. coli cells in log phase were treated with proteinase K for 30 min (A, C) or 120 min at 37 °C (B, D). (A, B) AFM images and (C, D) the widths of released fibers. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (C) 0.9 ± 1.4 (mean ± SD), 10.0 ± 2.9 nm and 30.6 ± 7.4 nm (the total number of observations, n = 101); and (D) 3.1 ± 3.3, 11.7 ± 0.6 nm and 27.4 ± 11.1 nm (n = 103). Scale bars in the AFM images represent 500 nm.

 
Nucleoid architecture in the stationary phase

In the stationary phase, the nucleoids in E. coli were condensed and their fibers unable to escape from the cell (Fig. 8A). To evaluate the RNA contribution to nucleoid architecture during stationary phase, stationary-phase E. coli cells were lysed and treated with 5 µg/mL RNase A for 5 min. AFM revealed decondensation of the condensed nucleoid and release of nucleoid fibers (Fig. 8B). Since no other kinds of nucleoid fibers were detected under our experimental conditions without RNase A treatment, we concluded that RNA was critical to the maintenance of condensed nucleoids in the stationary phase.


Figure 8
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  		Figure 8  Effect of RNase A treatment on stationary-phase E. coli nucleoids. (A) AFM image of the lysed cells in the stationary phase without the treatment with RNase A. (B) AFM image of the RNase A-treated lysed cells. (C) The fiber widths of RNase A-treated nucleoids. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (B) 2.6 ± 1.9 (mean ± SD), 12.1 ± 2.6 and 29.7 ± 6.6 nm (the total number of observations, n = 103). Scale bars in the AFM images represent 500 nm.

 
Notably, the degree of fiber release from RNase A-treated nucleoids was lower in the stationary phase than in the log phase (Table 1), and many fibers released from stationary-phase cells were bundled. Section analysis found 10 and 30-nm single fibers in the stationary phase (Fig. 8C). These results suggest that in E. coli, the hierarchical structures are the same in both the stationary and log phases. Interestingly, RNase A treatment in the stationary phase (unlike the log phase) released a significant amount of naked DNA, probably because the protein composition in the stationary phase differs from that in the log phase (Azam & Ishihama 1999).


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  		Table 1  Efficiency of fiber release upon RNase A treatment
 
The induction of Dps (DNA-binding protein from starved cells), which is the most abundant nucleoid protein in stationary-phase E. coli (Azam & Ishihama 1999), causes nucleoid condensation (Almiron et al. 1992; Wolf et al. 1999; Frenkiel-Krispin et al. 2001; Kim et al. 2004; Ohniwa et al. 2006). To investigate the possible involvement of Dps in sustaining the hierarchical fibrous structure and bundling revealed by RNase A treatment, RNase-treated nucleoids of the dps-deficient strain ({Delta}dps) were subjected to AFM analysis. The RNase A treatment released fibers with diameters similar to those of wild-type cells, indicating that Dps is not required to sustain the hierarchical fibrous units (10- and 30-nm fibers) during stationary phase. In contrast, there were fewer bundled fibers in the {Delta}dps than in the wild-type (Fig. 9), and nucleoid fibers were efficiently released (Table 1). Therefore, the fiber-bundling observed after RNase A treatment seems to depend on Dps-induced nucleoid condensation.


Figure 9
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  		Figure 9  Dps does not affect the 10 and 30-nm fiber structures. Effect of RNase A treatment on the {Delta}dps nucleoid was examined under AFM in (A) the stationary and (B) late stationary phases. The distributions of fiber widths are shown for the (C) stationary and (D) late stationary phases. Solid lines are obtained by Gaussian fitting, and the mean estimated widths are as follows: (C) 10.8 ± 4.5 (mean ± SD) and 28.1 ± 3.9 nm (the total number of observations, n = 101); and (D) 9.6 ± 5.4 and 25.7 ± 10.8 nm (n = 101). Scale bars represent 500 nm.

 

   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
We, in this report, evaluated the hierarchical architecture of bacterial nucleoids using a new model, the "circular cone model" for estimation of tip effects, and identified hierarchical 30- and 80-nm wide fibers in E. coli and S. aureus. RNase A treatment of these nucleoids released 10-nm fibers without nucleosome-like "beads-on-a-string" structure. The 30-nm fibers were resistant to RNase H and RNase III, but 80-nm fibers were disrupted by these treatments. When log-phase bacteria were treated with a transcription inhibitor, rifampicin, 10-nm fibers were also released upon lysis without RNase A treatment. These results indicate that transcription products contribute to bacterial nucleoid architecture; nascent single-stranded RNAs are essential to the assembly of 30-nm fibers from 10-nm fibers, and double-stranded RNAs/RNA–DNA hetero duplexes are required for the formation of 80-nm fibers from 30-nm fibers (Fig. 10).


Figure 10
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  		Figure 10  (A) Schematic representation of the nucleoid response to lysis and/or RNase A treatment, and (B) a model of the hierarchical structure of bacterial nucleoids. (A) Nucleoids in cells release 30 and 80-nm fibers upon treatment with weak detergent in physiological salt. After RNase A treatment of the lysed cells, most of the structural units released from nucleoids are 10-nm fibers. Nucleoids in cells cultured with rifampicin are destabilized and release 10-nm fibers after cell lysis. (B) The 10-nm fiber is composed of DNA and nucleoid proteins, and further folds up into the 30-nm fiber with the help of RNA molecules. The 30-nm fiber folds or coils into a super-solenoidal 80-nm fiber, which then forms a loop structure. The expression of Dps induces a highly condensed structure.

 
Nucleoid architecture coupled with transcription

RNA hybridization analysis revealed that the RNA molecules present in nucleoids isolated from E. coli were not nucleoid-specific but rather nascent rRNA and mRNA (Hecht & Pettijohn 1976). Although various non-coding small RNA species, some of which are fragments of mRNA or rRNA, have been identified in E. coli (Kawano et al. 2005), a unique class of chromosome-stabilizing RNA has not been found yet. One of the major proteins in isolated nucleoids is RNA polymerase (Murphy & Zimmerman 1997), suggesting that transcriptional activity must affect nucleoid architecture.

In addition, 5–10 major DNA-binding proteins and about 100 species of transcription factor are associated with nucleoids (Kundu et al. 1997). Specifically, HU (heat-unstable nucleoid protein), IHF (integration host factor protein), H-NS (histone-like nucleoid structuring protein), Fis (factor for inversion stimulation), Hfq (host factor for phage Qb replication), StpA (suppressor of td mutant phenotype A) and Dps are major nucleoid components in E. coli (Rouviere-Yaniv et al. 1979; Drlica & Rouviere-Yaniv 1987; Azam & Ishihama 1999). Interestingly, many of these proteins preferentially bind to single-stranded RNA rather than double-stranded DNA. Hu and H-NS recognize the 5'-UTR secondary structure of mRNA (such as rpoS mRNA) and non-coding small RNA (such as DsrA RNA), and regulate translational activities (Yamashino et al. 1995; Nakamura et al. 1999; Balandina et al. 2001, 2002; Brescia et al. 2004). Hfq has only RNA and not DNA binding activity and, in general, works as an RNA chaperone to bridge mRNA and non-coding small RNA to control their translation (Brescia et al. 2003; Moll et al. 2003; Zhang et al. 2003; Morita et al. 2006; Arluison et al. 2007). These results suggest the presence of complexes composed of non-coding small RNA, mRNA and proteins. Recently, the distribution of the RNA polymerase and three nucleoid proteins (H-HS, Fis and IHF) in whole genome DNA has been determined in E. coli using ChIP-chip (chromatin immunoprecipitation and high-density microarrays) (Grainger et al. 2005, 2006). The positions of RNA polymerase on the genome overlapped with those of H-NS and Fis. Interestingly, the amounts of H-NS and Fis together with RNA polymerase in the open reading frame (ORF) regions correspond to the degree of transcription activation or repression. Thus, transcription and translation occur concurrently on bacterial chromosome, nucleoid fiber architecture (involving these proteins, DNA and RNAs) must reflect the dynamic state needed for gene expression.

In eukaryotes, RNase A, but not RNase H, disrupts heterochromatin structure (Maison et al. 2002). AFM revealed 30-nm chromatin fibers in the RNase A-treated HeLa cell nucleus (Ohniwa et al. 2007), but not in the nuclei of untreated cells (Yoshimura et al. 2003). Thus, it is likely that these single-stranded RNAs maintain the integrity of chromatin fibers thicker than 30 nm and have transcription activity. Small RNAs may play a crucial role in heterochromatin formation control via RNA interference (Grewal & Elgin 2007). Similar small RNA mechanisms to control the assembly and maintenance of higher order chromosome structures might be present in both prokaryotes and eukaryotes.

Nucleoid architecture inside the cell

Rifampicin affects nucleoid condensation (Morgan et al. 1967; Dworsky & Schaechter 1973; Kleppe et al. 1979; Vos-Scheperkeuter & Witholt 1982; Sun & Margolin 2004; Kruse et al. 2006). Many reports have described the indirect attachment of DNA to the plasma membrane via coupled transcription, translation and translocation of membrane proteins (Kleppe et al. 1979; Vos-Scheperkeuter & Witholt 1982). Therefore, it has been proposed that, without RNA linkers between the strands of genomic DNA and the membrane, the force of macromolecular crowding due to non-bonding polymers and cations (like spermidine) compresses stable nucleoids causing their collapse (Lerman 1971; Zimmerman & Murphy 1996; Cunha et al. 2001). These facts and the proposed model are consistent with our observation that rifampicin-treated, lysed cells release 10-nm fibers, and have fewer nucleoids with compact, higher-order structure (Fig. 6). The nucleoids in rifampicin-treated cells are an assembly of less-organized nucleoid fibers that can be easily released by cell lysis (Fig. 10).

The in vitro structure of nucleoid fibers probably reflects their in vivo structure since the same proteins, RNAs and DNA involved in transcription are also involved in maintenance of nucleoid structure. It is important to realize that, upon cell lysis, these structures could be solidified and assemble into matrices as occurs for eukaryotes, forming a eukaryote-like nuclear matrix (or scaffold). It is possible that the 30–80 nm fibers seen in vitro do not exist in vivo and that more soluble forms persist instead.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Bacterial cultures

The wild-type (W3110) and dps deletion mutant [W3110 ({Delta}dps::Km)] are E. coli K-12 derived strains (Kim et al. 2004). Glycerol stocks of W3110 strains were inoculated into LB medium and cultured at 37 °C with constant shaking (180 rpm, Bioshaker BR-15; TAITEC, Tokyo, Japan) for 24 h. A 10-µL sample of the saturated culture was inoculated into 5 mL of fresh LB medium and cultured at 37 °C with constant shaking (180 rpm, Bioshaker BR-15) to an appropriate cell density. The cell density was determined by measuring absorbance at 600 nm using a UV-160A spectrophotometer (Shimadzu, Tokyo, Japan). Each strain was grown in LB medium until the OD600 reached 0.5 (log phase) and 2.0 (stationary phase). {Delta}dps cells in late stationary phase were collected after 24 h from the initiation of the culture. Rifampicin (dissolved in ethanol at 50 mg/mL) was added to the log phase culture to stop the transcription (final concentration, 100 µg/mL), and the cultures were incubated for 30 min or 60 min at 37 °C. Glycerol stocks of S. aureus (N315) (Kuroda et al. 2001) were inoculated into Brain Heart Infusion (BHI) medium (Difco, Detroit, MI) and cultured at 37 °C with constant shaking (180 rpm, Bioshaker BR-15) for 24 h. A 3-mL of the saturated culture was inoculated into 400 mL of fresh BHI and cultured at 37 °C, with constant shaking, to an appropriate cell density.

Lysis procedures

Bacterial cells were harvested from 100 µL of culture by centrifugation (13 000 g, 1 min at 4 °C) and washed once with 1 mL of phosphate buffered saline (PBS; pH 7.2). The cells were resuspended in 250 µL of PBS and a 50-µL aliquot was placed on a round cover glass (15 mm in diameter). The extra liquid was removed by evaporation with nitrogen gas. Escherichia coli cells were immersed in 2 mL of a buffer containing 10 mM Tris–HCl (pH 8.2), 1 mM NaN3 and 0.1 M NaCl for 5 min, followed by an addition of 25 µg/mL lysozyme. Staphylococcus aureus cells were immersed in 100 µL of buffer containing 10 mM Tris–HCl (pH 8.2) and 0.1 M NaCl for 5 min, and treated sequentially with 10 µL of lysostaphin (1 mg/mL) (Wako, Osaka, Japan) and 10 µL of 2 mg/mL N-acetylmuramidase SG (Seikagaku Corporation, Tokyo, Japan). After 2 min incubation at 25 °C, Brij 58 (polyoxyethylene hexadecyl ether) and sodium deoxycholate were added to the final concentrations of 0.25 and 0.1 mg/mL, respectively. After 10 min, the cover glass was dried under nitrogen gas. The surface of the cover glass was gently washed with distilled water and dried again for AFM analyses. In some experiments, the lysed cells on the cover glass were treated with 5 µg/mL DNase-free RNase isolated from bovine pancreas (RNase A, Roche, Welwyn, Garden City, UK), 10 U of RNase III (New England Biolabs, Ipswich, MA), 10 U of RNase H (Promega, Madison, WI) or 1 mg proteinase K (DNase- and RNase-free, Invitrogen, Carlsbad, CA) for 5, 30 or 120 min before AFM observation.

Transcription assay

Escherichia coli W3110 was grown at 37 °C with shaking in LB medium. When the OD600 reached 0.5, rifampicin or ethanol (solvent negative control) was added to the medium (final dilution, 1 : 500). After incubation at 37 °C for 5, 30 or 60 min, 1-mL aliquots of culture were harvested and chilled on ice. The cells were harvested by centrifugation, washed once with 0.05 M Tris–HCl (pH 7.5) (nakarai tesque, Tokyo, Japan) and resuspended in 10 µL of 0.05 M Tris–HCl (pH 7.5). An aliquot of 3.75 µL of the cell suspension was mixed with 21.25 µL of assay solution. The final reaction mixture (25 µL) contained 75 mM Tris–HCl (pH 7.5), 12 mM MgCl2, 60 mM KCl, 1 mM dithiothreitol (DTT), 240 U of RNase inhibitor, 1 mM each of ATP, CTP and GTP, 0.1 mM UTP and 12.5 µCi of [{alpha}-32P] UTP (3000 Ci/mmole; MP Biomedicals, Irvine, CA). After the incubation at 37 °C for 10 min, the reaction mixtures were put on ice. A 5-µL sample of each reaction mixture was mixed with 5 µL of 20 mg/mL lysozyme, Brij 58 (final concentration, 0.25 mg/mL) and sodium deoxycholate (final concentration, 0.1 mg/mL). Each mixture was incubated at room temperature for 2 min, followed by on-ice incubation for 10 min. Then 2 µL of each mixture was put onto a DE81 filter (ion exchange cellulose papers: Whatman Inc., Clifton, NJ). The filters were dried, and washed 5 times with 5% Na2HPO4 for 5 min, 2 times with water for 2 min and 2 times with ethanol for 1 min. After drying, the retained radioactivity was measured in a liquid scintillation counter.

Microscopy

The atomic force microscope (Nanoscope IIIa or IV) from Digital Instruments (Santa Barbara, CA) was used for the imaging of E. coli nucleoid structures in air at room temperature. The system was operated in tapping mode with a 100-µm scanner. Probes made of a single silicon crystal with cantilever length of 129 µm and spring constant of 33–62 N/m (OMCL-AC160TS-W2, Olympus, Tokyo, Japan) were used for imaging. Data were collected in the height mode using a scanning rate of 0.5–1.0 Hz and driving amplitude of 40–80 mV. The images were captured in a 512 x 512 pixels format and the captured images were flattened and plane-fitted before analysis. The image analyses were performed using software provided with the imaging module.

Correction of tip-effect: circular cone model

The apparent horizontal dimensions of the samples measured by AFM are generally much larger than the real dimensions owing to the effects of the edge curvature and point angle of the cantilever. From the EM images of the tip (OMCL-AC160TS-W2, Olympus) <Catalog: http://www.olympus.co.jp/en/insg/probe/>, the geometry of the cantilever and the globular sample can be drawn as in Fig. 1; the cantilever is circular cone shaped and curved on the edge. In practice, two situations are possible: (i) the sample is small but there is enough to be in contact with the curved surface of the tip (Y > H; Y is the distance between the surface and the end of the curvature, and H is the distance between the surface and the contact point between the sample and the tip, Fig. 1A); and (ii) the sample is big and in contact with a side face of the tip (Y < H, Fig. 1B). The relationship between the apparent width of the sample in the image (W), the radius of curvature of the tip (Rc), the real radius of the sample (Rm) and the point angle of the tip (2 x {theta}) can be given as follows:



Formula 1

(1)



Formula 2

(2)



Formula 3

(3)

N in Eq. (2) is the distance that depends on the shape of the tip edge (Fig. 1B). Equation (3) provides the value used to judge which equation, (1) or (2), should be applied to estimate W and Rm. In the case of naked DNA, since the Rm of DNA is 1 nm and the average {theta} of the tip is 12.5° (from the catalog), H is 1.2 nm. If the edge of the tip is an ideal incircle of the circular cone (Fig. 1C), Y is Rc x (1 – sin {theta}). In this case, Y is 5.9 nm for Rc = 7.5 nm and {theta} = 12.5° (from the catalog). Therefore, small samples like DNA should be evaluated by Eq. (1). In contrast, the H of a nucleosome in eukaryotic chromosomes is 6.7 using Rm = 5.5 nm from the X-ray crystal structure of the nucleosome (Luger et al. 1997) and {theta} = 12.5°, and, therefore, samples bigger than the nucleosome should be calculated using Eq. (2).

To clarify whether Eq. (2) is valid and practical, we applied this equation to samples of 30 nm chromatin and nucleosomes of eukaryotic chromosomes. Since the Rm and W of the nucleosome are, respectively, 5.5 and 33.8 nm (Fig. 1D), N based on the nucleosome was 10.2 nm ({theta} = 12.5°). Application of this N-value for 30 nm chromatin (W = 58.1 nm, Fig. 1D) gave 15.1 nm as the value of Rm for this chromatin. This value is close to the real radius of the 30 nm chromatin fiber (15.0 nm). Our estimates of N (based on the Rm and W of 30 nm chromatin fibers) and Rm of the nucleosomes were 10.7 and 5.0 nm, respectively. This Rm value also approximated the real radius of nucleosomes (5.5 nm), and, thus, Eq. (2) is useable for estimating the real dimensions of the samples.

Since the Eq. (2) is a linear function and the real width of the sample (S) is 2 x Rm, the Eq. (2) can be given as follows:



Formula

(4)

A and B are constants determined by the tip characters (N and {theta}). To obtain the values of A and B, in this study, we measured gold particles (with diameters 10, 30 and 80 nm; BBInternational, Cardiff, UK) as the standards, performed line fitting (Fig. 1E), and obtained the values 0.75 and –16.14 for A and B, respectively. These values are tip-specific and remain unchanged during use of the same tip. In this article, Eq. (4) was used to estimate the real dimensions of samples.


   Acknowledgements
 
This study was supported by the Special Co-ordination Fund, and a COE Research Grant and Basic Research Grant (B) from the Ministry of Education, Culture, Sports, Science and Technology of Japan. We thank the Japan Science Society (the Sasakawa Scientific Research Grant) and the Sumitomo Foundation for their strong support of this work. We also thank Dr Koji Hizume for providing us with AFM images of nucleosomes and 30 nm chromatin, and Ms. Shizuka Iwasaka for her technical support in molecular biology.


   Footnotes
 
Communicated by: Hiroji Aiba

* Correspondence: E-mail: ohniwa{at}lif.kyoto-u.ac.jp


   References
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 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
 References
 
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Received: 21 May 2007
Accepted: 5 July 2007





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