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Genes to Cells (2006) 11, 409-423. doi:10.1111/j.1365-2443.2006.00949.x
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Bacterial nucleoid dynamics: oxidative stress response in Staphylococcus aureus

Kazuya Morikawa1,*, Ryosuke L. Ohniwa2, Joongbaek Kim2, Atsushi Maruyama1, Toshiko Ohta1 and Kunio Takeyasu2

1 Institute of Basic Medical Sciences, Graduate School of Comprehensive Human Sciences, University of Tsukuba, Tennoh-dai, Tsukuba 305-8575, Japan
2 Laboratory of Plasma Membrane and Nuclear Signaling, Graduate School of Biostudies, Kyoto University, Sakyo-ku, Kyoto 606-8502, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
A single-molecule-imaging technique, atomic force microscopy (AFM) was applied to the analyses of the genome architecture of Staphylococcus aureus. The staphylococcal cells on a cover glass were subjected to a mild lysis procedure that had maintained the fundamental structural units in Escherichia coli. The nucleoids were found to consist of fibrous structures with diameters of 80 and 40 nm. This feature was shared with the E. coli nucleoid. However, whereas the E. coli nucleoid dynamically changed its structure to a highly compacted one towards the stationary phase, the S. aureus nucleoid never underwent such a tight compaction under a normal growth condition. Bioinformatic analysis suggested that this was attributable to the lack of IHF that regulate the expression of a nucleoid protein, Dps, required for nucleoid compaction in E. coli. On the other hand, under oxidative conditions, MrgA (a staphylococcal Dps homolog) was over-expressed and a drastic compaction of the nucleoid was detected. A knock-out mutant of the gene encoding the transcription factor (perR) constitutively expressed mrgA, and its nucleoid was compacted without the oxidative stresses. The regulatory mechanisms of Dps/MrgA expression and their biological significance were postulated in relation to the nucleoid compaction.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
In bacteria, the genomic DNA is packed in a cell as a form of ‘nucleoid’ (Robinow & Kellenberger 1994; Poplawski & Bernander 1997; Azam et al. 2000), whereas, in eukaryotic cells, the genomic DNA exists in a form of chromatin and is packed in a nucleus (Wolffe 1995). In either case, to organize the DNA into higher-order structures, a set of distinct structural DNA binding proteins, such as histones in eukaryotic cells, constitutively play major roles by utilizing the physical/chemical properties of DNA-protein interactions. A number of additional proteins also play crucial roles in the maintenance of well organized higher order structures of chromosome (Hayat & Mancarella 1995; Swedlow & Hirano 2003). These compacted genomes of prokaryotes and eukaryotes have been known to undergo dynamic structural changes depending upon their activities during the cell cycle. Therefore, it is critical to elucidate the transition mechanisms between the distinct hierarchies of the chromosome organization.

Atomic force microscope (AFM) scans a sample surface with a very sharp probe and reveals the topography of the sample surface (Hansma et al. 1988; Hansma & Hoh 1994; Bustamante et al. 1997). We have developed a series of procedures for the biological application of AFM (Nettikadan et al. 1996; Ohta et al. 1996; Sato et al. 1999; Yoshimura et al. 2000a,b). In our previous study on the Escherichia coli nucleoid (Kim et al. 2004), a hierarchy of 40 nm, 80 nm, and thicker fibers was found to be a key feature for the architecture of the genome organization, and that this hierarchy undergoes a series of changes during the growth. On the basis of these results, a structural model has been proposed for the nucleoid organization (Fig. 1), in which the 80 nm fibers play a role as a fundamental structural unit. The 80 nm fiber builds up a 300 nm loop in cells and a coral reef structure is formed during the progression of cell growth phases. In the previously proposed model (Trun & Marko 1998), a random coil structure of DNA is turned into a loop domain every 50 kbp. This model could explain the structural relationship between the 300 nm loop and the coral reef structure. The coral reef structure might be composed of a number of 300 nm loops, and undergoes further compaction towards the stationary phase. Mutant analyses revealed that a nucleoid protein, Dps, plays a key role for the nucleoid compaction towards the stationary phase (Kim et al. 2004).


Figure 1
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  		Figure 1  A model of the architectural hierarchy in E. coli nucleoid (Kim et al. 2004). The DNA and nucleoid proteins form a 40 nm fiber structure. The 40 nm fiber is super-solenoided into an 80 nm fiber (their AFM image is shown in left panel). These fiber structures are commonly found in the bacterial nucleoid. The 80 nm fiber, then, forms a loop structure (middle panel, arrows). The nucleoid undergoes a series of drastic structural changes into a tightly compacted state (right panel). The nucleoid compaction is mainly mediated via a nucleoid protein, Dps. In the present study, the loop and coral reef structures were not detected in S. aureus. Scale bars, 500 nm.

 
The Dps protein, which is encoded by a gene, dps, is a member of the Fe-binding protein family that forms multimers in cells (Grant et al. 1998; Ren et al. 2003). The Dps protein and its homologs have DNA binding activity (Chen & Helmann 1995; Martinez & Kolter 1997), and E. coli Dps is known to be the dominant nucleoid protein in the stationary phase cells (Talukder et al. 1999). The dps gene expression in E. coli is up-regulated by a transcription factor, IHF, and the {sigma}s-factor towards stationary phase (Altuvia et al. 1994; Lomovskaya et al. 1994). Bioinformatic analysis has revealed that not all bacterial species possess the dps gene and its regulators; e.g. Clostridium perfringens lacks them (Takeyasu et al. 2004). Our preliminary analysis has suggested that, although Staphylococcus aureus has a dps homolog gene, mrgA, the regulatory mechanisms of the nucleoid architecture are quite different from those in E. coli, because 40 and 80 nm fibers were still observed at the stationary phase in S. aureus (Takeyasu et al. 2004).

In this study, the nucleoid architectures of S. aureus were investigated in detail by AFM. The results obtained showed that the S. aureus nucleoid always exhibited 40 and 80 nm fibers and was never compacted during a normal growth condition. However, it did undergo drastic conformational changes into a compacted clump under an oxidative stress condition. Bioinformatics on the transcriptional regulators for the dps gene and its homologs suggested that the oxidative stress, which is directly sensed by evolutionary distinct regulators, is a common induction signal among bacterial phyla, whereas the stationary phase-specific regulation is rather restricted to some species.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Staphylococcal nucleoids do not undergo a stationary-phase compaction

A clinical isolate designated N315 was selected as a representative of S. aureus, because its defined genomic sequences certify the intactness of the important genes focused in this study (Kuroda et al. 2001).

The S. aureus N315v cells, control strain cells, were collected at intervals during their growths, lysed as described in Experimental procedures, and subjected to AFM analyses. The nucleoids from the log- and the late log-phase cells were composed of 40 and 80 nm fibers (Fig. 2C,D). The nucleoids from the cells that had been incubated for 2 days still released the 40 and 80 nm fibers upon lysis (Fig. 2E). This finding suggests that the fundamental fiber-units required to build up the higher-order architectures in S. aureus are essentially the same as those in E. coli. However, the nucleoids of both species evidently behave differently towards the stationary phase; i.e. the E. coli nucleoid undergoes a compaction under a normal growth condition (Kim et al. 2004), but the S. aureus nucleoid does not.


Figure 2
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  		Figure 2  (A–E) Nucleoid structures of S. aureus in a normal growth condition. Cells were collected in the mid-log phase (C), late-log phase (D), and stationary phase (E), lysed as described in Experimental procedures and subjected to AFM analyses. The sampling points were marked by arrowheads on the growth curve (A). Close observations of the stationary phase nucleoid showed two different fiber structures (40 and 80 nm). The dotted square areas (C–E) were directly rescanned at higher resolutions and shown as inlets. A curve obtained by a Gaussian fitting was overlaid in the distribution histogram of the fiber widths (bin size, 10 nm) (B). Scale bars, 500 nm. (F) Northern blot analysis of the mrgA transcripts. N315 cells were harvested at various growth phases. Lanes 1 through 3: O.D.600 = 0.5, 1.0, 1.5, respectively. Lanes 4 through 10: 6, 7.5, 9, 10.5, 12, 26 and 48 h after inoculation, respectively. The expression of mrgA was constitutive throughout the growth. Note that lane 1 is identical to lane 1 in Figure 4A. Bottom panel: Ethidium bromide staining of rRNAs.

 
MrgA is the key factor in the nucleoid compaction

We previously demonstrated that the key factor for the nucleoid compaction is the Dps protein in E. coli (Kim et al. 2004). In S. aureus and other Gram-positive bacteria such as Bacillus subtilis, the Dps homolog is designated as MrgA (Metallo regulated genes A), which has a 40% amino acid sequence similarity (23.8% identity) to E. coli Dps. Northern blot analyses were used to define the timing when the endogeneous mrgA gene can be expressed. In the BHI medium, the mrgA gene was expressed at a certain level in the log phase, and this level was rather sustained throughout the growth (Fig. 2F), which is quite different from the case of the E. coli dps gene whose expression is drastically up-regulated towards the stationary phase (Azam et al. 1999).

To test whether staphylococcal MrgA is functionally equivalent to E. coli Dps in the nucleoid compaction, we examined the effect of MrgA over-expression on the nucleoid structure. The MrgA-over-expressing strain, N315M, was grown in the BHI medium and the nucleoids from the log- and stationary-phase cells were subjected to the AFM analyses (Fig. 3). The fibers released from the cells were hardly detectable. Instead, N315M exhibited a highly clumped structure (Fig. 3A,B); 96.6% (n = 59) of N315M and 0% (n = ~31) of N315v cells exhibited the clumped structure) which contained DNA as proved by DAPI staining (compare Fig. 3C). These results indicate that an over-expression of MrgA promotes the formation of the clumped structure, and suggest that MrgA is the functional counterpart of Dps.


Figure 3
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  		Figure 3  Nucleoid compaction induced by MrgA over-expression. N315M cells harboring the mrgA-expression plasmids were collected in (A) the mid-log phase and (B) the stationary phase, lysed as described in Experimental procedures and subjected to AFM analyses. The dotted square areas were directly rescanned at higher resolutions and shown as inlets. Scale bars, 500 nm. Panels in C are the DAPI-stained images of the lysed cells of N315M and N315v. Scale bars, 20 µm.

 
Oxidative stress induces the expression of mrgA and nucleoid compaction

We searched for certain environmental conditions or stresses that could induce the mrgA expression. The S. aureus cells grown in the BHI medium until mid-log phase were exposed to different growth conditions or stresses and subjected to a Northern blot analysis (Fig. 4A). A shift to the oxygen-limiting condition, where S. aureus could grow anaerobically, had no positive effect on the mrgA expression. Similarly, other stresses such as a hypotonic shock, a heat shock, a ß-lactam antibiotic, and a different culture medium (RPMI1640), had no inducible effect on the mrgA expression. In contrast, the oxidative stresses by phenanthrenquinone (PQ), menadione (MD), and H2O2 drastically induced the mrgA gene expression. This indicates that the reactive oxygen species are effective inducers for the mrgA expression. This is expected to promote the nucleoid compaction.


Figure 4
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  		Figure 4  (A) Drastic induction of the mrgA expression by oxidative stresses. N315 cells were grown in BHI media until the mid-log phase (lane 1) and exposed to stresses: anaerobic condition (lane 2), heat shock (46 °C, lane 3), osmotic stress by 1.5 M NaCl (lane 4), oxidative stresses by 80 µM of MD (lane 5), by 20 µM PQ (lane 6), by 500 µM H2O2 (lane 7), by 50 mM H2O2 (lane 10), 16 µg/mL (sub-MIC) of oxacillin (ß-lactam antibiotics, lane 8), different medium (RPMI 1640, lane 9) for 30 min. Four microgram of the total RNAs were subjected to the Northern blot analysis with a mrgA specific probe (see Experimental procedures). Ethidium bromide staining of rRNAs shows equal loading of RNA. (B–D) Oxidative stress-induced nucleoid compaction. Cells were grown in BHI media as in Figure 2, but exposed to the oxidative stress by adding 20 µM of PQ (B,C), or 50 mM H2O2 (D) when O.D.600 reached 0.5. The cells were harvested after 30 min (B,D) or 2 days (C), and, then, their nucleoids were subjected to the AFM analysis.

 
To examine the expected conformational dynamics, the nucleoids from the S. aureus cells exposed to the oxidative stresses were subjected to the structural analyses under AFM. During the incubation for 30 min in the presence of the redox-cycling reagent, PQ, the nucleoids underwent a drastic structural change into the clumped complex (Fig. 4B); 100% (n = 32) of the cells treated with PQ exhibited the compacted structure), and the 40 and 80 nm fibers were rarely detected. When the cells were cultured for 2 days under the PQ exposure (Fig. 4C), we could never find the 40 and 80 nm fibrous structures released from the cell.

Other oxidative stresses by MD (data not shown) and H2O2 (Fig. 4D) also induced the nucleoid compaction (percentages of the cells exhibiting the compacted structure is 59.2 (n = 152) and 38.8 (n = 49) for the MD and 50 mM H2O2 treatments, respectively.) In contrast, it should be noted that, without the oxidative stress, the nucleoids never underwent such structural changes (Fig. 2). These results indicate that the oxidative stress is a key signal that triggers the MrgA-dependent nucleoid compaction.

The mrgA mutant was generated as described in Experimental procedures. The growth rate and colony morphology were not affected by the mrgA mutation, indicating that mrgA is dispensable in the normal growth condition. However, the mrgA mutant could not exert the nucleoid compaction response when exposed to the oxidative stress (Fig. 5A,B). This indicates that mrgA gene is essential to construct the compacted nucleoid structure under the oxidative stress. We also confirmed that the mrgA mutant exhibited increased susceptibility to H2O2 (Fig. 5C) as well as to UV irradiation (Fig. 5D) that can cause direct DNA damage and possibly indirect damage via the production of reactive oxygen species (ROS) (Cadet et al. 2005). Theses results are in line with the previous reports in other species (Nair & Finkel 2004).


Figure 5
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  		Figure 5  (A,B) Nucleoid images of the mrgA mutant under oxidative stress. The mrgA null mutant was exposed to the oxidative stress by adding 20 µM of PQ when O.D.600 reached 0.5. The cells were harvested after 30 min, and their nucleoids were subjected to the AFM analysis (A) or the fluorescent microscopy by DAPI-staining (B). Scale bars, 500 nm in (A) and 20 µm in (B). (C,D) Increased susceptibilities to oxidative stress and UV irradiation in mrgA mutant. (C) Viabilities of cells after the 200 mM or 400 mM H2O2 treatment. N315: open column, mrgA mutant: filled column. (D) UV tolerance. Cells were evenly spread on the BHI-agar plate, and UV was irradiated for the indicated periods. The plate was incubated at 37 °C for overnight to detect colonies from survived cells. The mrgA mutant (right) was more susceptible to UV than N315 (left).

 
OxyR and PerR, regulators of dps/mrgA, spread widely in the bacterial kingdom

OxyR and PerR are evolutionarily distinct transcription factors that are known to regulate the expression of dps (Altuvia et al. 1994) and mrgA (Horsburgh et al. 2001a), respectively, in response to oxidative stresses (see Discussion). OxyR acts as an activator and belongs to the large LysR-family of transcriptional regulators (Schellhorn 1995), and PerR acts as a repressor and is homologous to Fur and Zur that are different from the LysR-family (Horsburgh et al. 2001a). Phylogenetic analysis of the LysR-family and the Fur homologs revealed that OxyR and PerR form distinct clades in their phylogenetic trees (Fig. 6A,B). S. aureus possess SA2330 and SA2123 as the LysR-family proteins, but they were excluded from the OxyR clade, implying that they may not share the functions with OxyR. When the occurrences of Dps/MrgA (Takeyasu et al. 2004) were compared with those of OxyR and PerR, 85.3% of the 66 strains containing Dps/MrgA possessed at least one of them, and, interestingly, most species in Bacillales and Lactobacillales carried PerR, but not OxyR (Table 1). Another transcription factor, IHF, which is known to regulate the E. coli dps gene towards the stationary phase (Altuvia et al. 1994), was restricted mostly in Proteobacteria. The PerR was expected as the key regulator in the nucleoid compaction in S. aureus.


Figure 6
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  		Figure 6  Phylogenetic analyses of the LysR family and the Fur homologs. (A) 879 OxyR homologs. The OxyR clade (red) was clearly separated from other LysR family proteins. The essential cystein residues (199th and 208th from N-terminal) required to form the sensor disulfide bridge (Choi et al. 2001) were conserved in the defined OxyR clade (BRA0709 in Brucella meltensis and BMEII0576 in Brucella suis lack 208th Cys), and never found in other LysR family proteins including SA2330 and SA2123. (B) 219 Fur homologs. The genes identified are colored red in E. coli, blue in B. subtilis and green in S. aureus.

 

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  		Table 1 Prevalence of OxyR and PerR homologs. The OxyR and PerR homologs were defined as the members in the OxyR clade in Fig. 6A and the PerR clade in Fig. 6B, respectively. The Dps distribution was derived from our previous analyses (Kim et al. 2004; Takeyasu et al. 2004)
 
Based on the above considerations, we examined the effect of the perR mutation on the nucleoid structure in S. aureus. Our perR mutant generated from N315 constitutively expressed the mrgA gene in the absence of the oxidative stress (Fig. 7A), as expected. This is consistent with the previous study (Horsburgh et al. 2001a; Morrissey et al. 2004). The growth rate was not altered by the perR mutation in the employed growth condition. The nucleoid of the perR mutant exhibited the compacted structure in the absence of the oxidative stress, and no fibrous structure was detected (Fig. 7B).


Figure 7
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  		Figure 7  Effect of perR mutation on mrgA expression and nucleoid structure. (A) The perR mutant expresses mrgA without the oxidative stress. N315 (lanes 1 and 2) and the perR mutant cells (lanes 3 and 4) were grown in BHI until mid-log phase (lanes 1 and 3), and exposed to 20 µM PQ for 30 min (lanes 2 and 4). Four microgram of the total RNAs were subjected to the Northern blot analysis with the mrgA specific probe. Lower panel: Ethidium bromide staining of rRNAs. (B) Nucleoid image of the perR mutant under normal growth condition. Dotted square area was directly rescanned at the higher resolution, and shown as inlet. Scale bars, 500 nm.

 

   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
AFM combined with the ‘on-substrate lysis’ method revealed that the 80 nm and 40 nm fibers dominated when the nucleoid was released from the S. aureus cells at any growth phases (Fig. 2). These fibers were stable, never unfolded further in the present experimental condition, and seem to be the common structural units of bacterial nucleoids, since they have been found in the log phase cells of S. aureus, C. perfringens and E. coli (Takeyasu et al. 2004). These fibers in S. aureus underwent a tight compaction under certain conditions as seen in E. coli (Fig. 4). Possible genetic switches for the nucleoid compaction can be proposed where the dps/mrgA genes and their products (Dps/MrgA) play a key role under different growth phases and oxidative stresses (Figs 1 and 3). In S. aureus, the nucleoid compaction is thought to hardly hinder the cellular processes required for the normal growth in the employed growth condition; N315M and perR mutant cells could grow normally. In addition, loss of the mrgA gene did not influence the progression into the stationary phase. This is also true in E. coli; the dps mutant, which is deficient in constructing higher order nucleoid structures, can enter into the stationary phase (Kim et al. 2004; Nair & Finkel 2004). Thus, the dps/mrgA dependent nucleoid compaction seems to be dispensable in the control of the log-phase to stationary-phase transition itself.

Roles of Dps/MrgA in the oxidative stress resistance

ROS are highly reactive molecules, which are generated during the oxidative respiration of bacteria themselves, leading to damages of various macromolecules including the genomic DNA (Nunoshiba et al. 1999). In our defense system, phagocytes such as neutrophils and macrophages produce ROS to kill the infectious bacteria (Clements & Foster 1999; El-Benna et al. 2005). On the other hand, bacteria are often equipped with several surviving mechanisms against the oxidative stress. For example, bacterial enzymes such as super oxide dismutase and catalase convert the ROS into safe molecules (Clements et al. 1999; Sanz et al. 2000; Valderas & Hart 2001). The Dps/MrgA family proteins also constitute the oxidative stress-responsive systems (Almiron et al. 1992; Martinez & Kolter 1997). The Dps protein is a stress-induced protein with a molecular weight of 19 kDa and is known to be a member of the Fe-binding protein family that forms multimers in cells (Grant et al. 1998). A dps homolog, mrgA in B. subtilis was found as the metal ion responsive gene (Chen et al. 1993). Its product, MrgA, can also form a multimer complex in vitro and possibly can bind DNA (Chen & Helmann 1995). The protective mechanism by Dps/MrgA has been attributed to their ability to reduce the intracellular level of iron (Fe2+) that generates the super oxide anion through the Fenton reaction (Zhao et al. 2002). Alternatively, it has been suggested that their DNA binding might protect the genome against the ROS (Wolf et al. 1999). An additional possible resistance mechanism may be that a reduction of the nucleoid ‘surface’ by Dps/MrgA-dependent compaction (Fig. 3) reduces the challenges of ROS, although we have no direct evidence for the relationship of the nucleoid compaction and the resistance.

Additional factors regulating the nucleoid compaction

It is surprising that an over-expression of MrgA in S. aureus could induce a nucleoid compaction in log-phase cells (Fig. 3A), because an over-expression of Dps in E. coli cannot induce such a compaction in the log-phase (Kim et al. 2004). It may be that certain factor(s) that cooperatively functions with MrgA is constitutively active in S. aureus. Alternatively, certain inhibitory factor(s) that inhibits the nucleoid compaction may exist in the log-phase E. coli cells, but not in S. aureus. For the latter case, the candidates may include the E. coli-specific components of the nucleoid, such as Fis and H-NS. It is interesting to note that the expression level of Fis declines toward the stationary phase (Ball et al. 1992; Talukder et al. 1999).

It can be speculated that the nucleoid compaction requires a modulation of the DNA superhelical density. In fact, the previous biochemical studies have revealed that oxidative stresses affect the DNA topology (Weinstein-Fischer et al. 2000). Two enzymes, topoisomerase and gyrase, can modulate the superhelical density of DNA in response to the oxidative stresses, and the Fis protein reduces the activity of DNA gyrase and counteracts the increase of the overall superhelicity of DNA during early exponential growth phase (Muskhelishvili & Travers 2003). It is likely that Fis acts as an inhibitory factor in the nucleoid compaction in E. coli, and its counterpart is missing in S. aureus.

Regulatory mechanism of the dps/mrgA genes

In E. coli, IHF together with {sigma}s-factor up-regulates the dps gene expression towards the stationary phase (Fig. 8) (Altuvia et al. 1994), and the amount of its product, Dps, reaches up to 180 000 molecules/cell (~25-fold induction) (Talukder et al. 1999). In contrast, the staphylococcal mrgA gene was not induced toward the stationary phase (Fig. 2F); observation attributable to the lack of the IHF/{sigma}s system (Takeyasu et al. 2004; Fig. 8). The differences of such regulatory modes are well correlated with the nucleoid compaction status in normal growth condition.


Figure 8
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  		Figure 8  Schematic representation of the regulatory mechanisms of the dps/mrgA genes in E. coli and S. aureus. In E. coli, IHF and {sigma}s-factor regulate the dps gene expression towards the stationary phase, and OxyR and {sigma}70 stimulate the dps gene expression under the H2O2 stress. In S. aureus, only the PerR/{sigma}A pathway exists due to the lack of the genes for IHF/{sigma}s in the genome. Therefore, the staphylococcal mrgA gene cannot be induced toward the stationary phase, although it is expressed under oxidative stresses.

 
A phylogenetic analysis of LysR family defined the OxyR group as a tight clade (Fig. 6A). The two cystein residues that are essential to form a disulfide bridge within the sensor domain in E. coli OxyR (Linke & Jakob 2003) were conserved in the defined OxyR group proteins, but never found in the other LysR-family proteins. Therefore, it is unlikely that SA2330 and SA2123 are functionally equivalent to OxyR. This is consistent with the fact that the perR mutation is sufficient to abolish the oxidative stress-response of the mrgA gene expression (compare lanes 3 and 4 in Fig. 7A).

Under oxidative stress, PerR derepresses the mrgA gene expression in the presence of iron but not manganese (Horsburgh et al. 2001a, 2002; Morrissey et al. 2004). Fur and Zur are the other members of Fur homologs, and dependent on iron and zinc, respectively (Gaballa & Helmann 1998; Patzer & Hantke 1998; Horsburgh et al. 2001b; Andrews et al. 2003). The phylogenetic tree of Fur homologs (Fig. 6B) revealed that E. coli lacks PerR but possesses Fur (b0683) and Zur (b4046). However, b0683 and b4046 cannot compensate for the OxyR function in the dps gene regulation, i.e. dps was not induced under oxidative stress in {Delta}oxyR mutant of E. coli in which the b0683 and b4046 genes were intact (Zheng et al. 2001).

S. aureus and E. coli exclusively utilizes PerR and OxyR, respectively, to regulate the mrgA/dps expression as an oxidative stress response (Fig. 8). These systems seem to have evolved independently in Proteobacteria and Firmicutes (Table 1).

Bacillus subtilis, a close relative of S. aureus, also lacks the ihf gene but possesses two mrgA genes. One is under a control of PerR (Chen & Helmann 1995; Bsat et al. 1998; Fuangthong & Helmann 2003), and the other seems to be regulated by SigB in a growth phase-dependent manner (Antelmann et al. 1997). Although SigB is conserved in Bacillales, the mrgA genes of S. aureus, Staphylococcus epidermidis, Oceanobacillus iheyensis, Bacillus anthracis, Bacillus thuringiensis and Bacillus cereus harbor no potential sequence that could be recognized by SigB. In fact, staphylococcal mrgA was not induced toward the stationary phase (Fig. 2F) in spite of the activation of SigB (Bischoff et al. 2001; Giachino et al. 2001).

In summary, although Proteobacteria and Firmicutes have different regulatory systems for the dps/mrgA gene expression, they seem to respond to the oxidative stress in a similar fashion (i.e. condensing their nucleoids) via the OxyR and PerR systems, respectively. We suspect that bacteria generally respond to the oxidative stress by constructing highly organized nucleoid architectures, and that the certain species have acquired the ability to respond to the starvation or stationary phase signals. Extra-ordinal regulatory systems of the nucleoid conformation may be found in species such as Streptococcus pneumoniae and Treponema pallidum (Table 1) carrying the dps genes that are not regulated by OxyR and PerR.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Construction of mrgA over-expressing strain

The mrgA gene (SA1941, dps homolog) of S. aureus N315 (a clinical isolate in 1982, pre-MRSA; Kuwahara-Arai et al. 1996; Kuroda et al. 2001) was amplified by PCR with a set of primers, SA1941-f(Eco): 5'-GGAATTCGACAATTTTAAFormula T-3', and SA1941-r(Sal): 5'-CGCGTCGACTGTAGATTAGCTTAAGTAAGA-3'. The SA1941-f(Eco) primer was designed to include the Shine-Dalgarno sequence (double underlined). The amplified 468 bp fragment was digested with EcoR I and Sal I, and cloned into the EcoR I-Sal I sites of pRIT5H (Morikawa et al. 2003). The resulting plasmid, pRIT-mrg, can over-express MrgA under a control of the proteinA promoter. The pRIT-mrg was introduced into the N315 strain to generate a strain, N315M. To generate a control strain, N315v, pRIT5H was introduced into N315. The plasmids were stably maintained in the presence of 12.5 µg/mL chloramphenicol.

Growth condition

Glycerol stocks of S. aureus strains (Table 2) were inoculated into Brain-Heart Infusion (BHI) media containing 12.5 µg/mL chloramphenicol and cultured at 37 °C with constant shaking (180 r.p.m) for 24 h. Three millilitre of the saturated culture was inoculated into 400 mL of fresh BHI with chloramphenicol and cultured at 37 °C with constant shaking to an appropriate cell density. The cell density was determined by measuring the absorbance at 600 nm.


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  		Table 2 S. aureus strains used in this study
 
RNA isolation and Northern hybridization analyses

S. aureus strains were grown in drug-free BHI medium until the O.D.600 reached at 0.5 (log phase), and were exposed to the appropriate stresses for 30 min. Anaerobic condition was made by using AnaeroPack (Mitsubishi Gas Chemical). Oxidative stresses were given by the addition of 20 µM 9,10-phenanthrenquinone (PQ), 80 µM 2-methyl-1,4-naphthoquinone (menadione; MD), and 500 µM or 50 mM of H2O2. After 30 min incubation, the cells were harvested by a centrifugation at 13 000 x g for 30 s at 4 °C, suspended in a lysis buffer containing 10 mM Tris-HCl (pH 8.0), 10 mM EDTA, and 100 µg/mL of lysostaphin, and, then, incubated at 37 °C for 2 min followed by the on-ice incubation for 30 min. The total RNA was extracted from the lysate by the SV total RNA isolation system (Promega). Four microgram of the total RNAs was separated on a 1% agarose-formamide denaturing gel and transferred on to the Hybond N+ membrane (Amersham Biosciences). The DNA fragments of the mrgA gene prepared by PCR from the N315 genomic DNA were labeled by a random priming method using the Ready-to-Go DNA labeling Beads (Amersham Biosciences), and used as the probes. The hybridization was conducted at 60 °C in the hybridization solution containing 5 x SSPE, 5 x Denhardt's solution, 0.5% SDS, and 20 µg/mL of salmon sperm DNA for 16 h, and the final washing was done at 60 °C in 0.1 x SSC and 0.1% SDS for 30 min

Bacterial cell lyses procedures

S. aureus cells were harvested from a 500 µL culture by centrifugation (13 000 x g, 1min at 4 °C) and washed once with 1 mL PBS (pH 7.2). The cells were resuspended in 0.25 mL or 2 mL of PBS and a 5 µL aliquot was placed on to a round-shape cover glass, 18 mm in diameter. The extra liquid was removed by nitrogen gas blow. The sample was immersed in 100 µL of a buffer containing 10 mM Tris-HCl (pH 8.2), and 0.1 M NaCl for 5 min, followed by a sequential addition of 10 µL Lysostaphin (1 mg/mL) (Wako, Japan) and 10 µL of 2 mg/mL N-Acetylmuramidase SG (Seikagaku Corporation, Japan). After 2 min incubation at 25 °C, Brij 58 (polyoxyethylene hexadecyl ether) and sodium deoxycholate were added for 10 min to the final concentrations of 0.25 mg/mL and 0.1 mg/mL, respectively. After the specimen was gently washed with distilled water, the excess water on the specimen was removed under nitrogen gas for microscopic analyses.

Microscopy

The atomic force microscope (SPI3800N-SPA400) from Seiko Instrument Inc. was used for the imaging of S. aureus nucleoid structures in air at room temperature under a dynamic force mode with a 150 µm scanner. Probes made of a single silicon crystal with the cantilever length of 129 µm and the spring constant of 33-62 N/m (OMCL-AC160TS-W2, Olympus) were used for imaging. Data was collected in the height mode with a scanning rate of 0.2-0.5 Hz and the driving amplitude of 40-80 mV. The images were captured in a 512 x 512-pixel format and the captured images were flattened and plane-fitted before analysis. The image analyses were performed with the software accompanying with the imaging module (Seiko Instrument Inc., Japan).

All of the AFM images contain ‘tip effect.’ The sizes of the objects in the images were estimated at the half maximum height (FWHM; full width at half-maximum) for correction of the tip effect (Schneider et al. 1998). In the present study, we have found that the AFM cantilevers purchased from Olympus had a constant tip angle (~35 degrees) and tip radius (20 nm+/- 2 nm) by measuring the apparent width of double stranded DNA in the AFM images.

For the fluorescence microscopy, the sample was incubated in 4',6-diamino-2-phenylindole (DAPI, Sigma) solution (1 µg/mL), and observed under a fluorescence microscope (DM RBE, Leica, Germany), and the images were captured by a chilled CCD camera (C5985, Hamamatsu Photonics, Japan).

Construction of mrgA and perR mutants

The mrgA targeting vector, pKILts-cat 1941, was constructed as follows.

The chloramphenicol acetyl transferase gene (cat) including its promoter region was amplified from pRIT5H by PCR with a set of primers CAT(+ P)F: 5'-CGAAAATTGGGTACCGTGGGATATTTT-3' and CAT(+ T)R: 5'-CAACTAACGGGGCATATGAGTGACATT-3'. The fragment was blunt-ended and cloned between BamH I and Hind III sites of pKILts (Morikawa et al. 2001), which has tetracycline resistance marker and the temperature-sensitive replication origin for S. aureus. The resulting plasmid was designated pKILts-cat.

The upstream and downstream regions of the mrgA gene were amplified by PCR with the primers: UF1941-1: 5'-CCGGATCCTATCGTGAAGGGTTTATTAC-3' and UF1941-2: 5'-GCGGATCCTAATCTAAATGTAAGGTG-3' (for the upstream 1.25 kbp fragment); DF1941-1: 5'-GGCAAGCTTAAGCTAATCTACAGATAAGT-3' and DF1941-2: 5'-GGCAAGCTTTCGCTTCATCGATGATTTG-3' (for the downstream 1.3 kbp fragment). The PCR fragments obtained were sequentially cloned into the BamH I-site and the Hind III-site of pKILts-cat. In the resulting plasmid, pKILts-cat 1941, the cat gene lies between the two fragments.

The perR targeting vector was constructed as follows. The 2.2 kbp region encompassing the perR gene was amplified by PCR with a set of primers: UF-perR-1: 5'-GCGGATCCTGAGAGTGACTT-3' and DF-perR-2: 5'-GGCAAGCTTCTCGTCCGTTC-3'. The resultant fragment was blunt-ended and ligated into the PvuII site of pSP72. The cat gene was amplified by PCR with the primers, CAT(+ P)F and CAT(+ T)R (see above), and inserted into the internal PvuII site within the perR coding sequence (+ 136 from the translation initiation site). The fragment encompassing the disrupted perR gene was excised from the plasmid by BamH I and HindIII, blunt-ended, and ligated into pKILts to generate the perR targeting vector, pKILts-cat-per. This plasmid was passed through a S. aureus strain, RN4220, prior to the introduction into N315.

Each of pKILts-cat 1941 and pKILts-cat-per was introduced into S. aureus N315 by electroporation. The N315 cells carrying the plasmids were grown in BHI containing 10 µg/mL of tetracycline at 30 °C. This growing temperature allows the replication of these plasmids. Then the cells were plated on to a BHI-agar plate containing 12.5 µg/mL of chloramphenicol and incubated at 43 °C in order to select the transformants that had the cat gene on its genome. The chloramphenicol resistant colonies were further selected for the absence of the tetracycline resistance by a replica method. These procedures generated the mrgA and perR mutants, where the original mrgA or perR gene was replaced with the cat gene by a double-cross homologous recombination event. The absence of the mrgA or perR gene in the mutant was confirmed by PCR.

Oxidative-stress assay

N315 and the mrgA mutant cells were washed with PBS. The cells were incubated at 25 °C for 3 min in PBS with or without H2O2, and then serially diluted with ice-cold saline. The viable cells were counted as colony forming unit (cfu) on BHI agar plates. The assay was independently performed twice, and the representative result was shown in Fig. 5C.

UV irradiation

N315 and the mrgA mutant cells were serially diluted with ice-cold saline, and plated on BHI agar. The plates were placed under a germicidal lamp (254 nm, National GL-15), and exposed for 0–60 s. The assay was independently performed twice, and the representative result was shown in Fig. 5D.

Database search

For the 97 bacteria whose genome projects had been completed, the genes encoding OxyR, and PerR were searched within the SSDB (Kanehisa & Goto 2000). When the genes were not found in certain species, FASTA search of the genes and genomes was conducted on the KEGG database (Kanehisa & Goto 2000) using the amino acid sequences for OxyR of E. coli, and PerR of S. aureus and Synechocystis sp. as queries.

The amino acid sequences of the 879 retrieved proteins homologous to OxyR (e-value is under e-10) and 219 retrieved proteins homologous to PerR (e-value is under 0.0001) were aligned with the ClustalX program (Jeanmougin et al. 1998). The alignment was used for the phylogenetic analysis with the PROTDIST and NEIGHBOR programs of the PHYLIP 3.6 package (Felsenstein 1989). The phylogenetic tree was inferred by the neighbor-joining method (Saitou & Nei 1987) and tested by 1000 replications of the bootstrap analysis which was carried out with the SEQBOOT and CONSENSE programs in the same package, and, then, visualized using the TREEVIEW program (Page 1996).


   Acknowledgements
 
This study was supported by the Special Co-ordination Funds, the COE Research Grant and the 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 for this work.


   Footnotes
 
Communicated by: Masayuki M. Yamamoto

* Correspondence: E-mail: morikawa{at}sakura.cc.tsukuba.ac.jp


   References
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 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
 References
 
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Received: 16 November 2005
Accepted: 3 January 2006




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