Multiple Regulatory Systems Coordinate DNA Replication with Cell Growth in

Summary

DNA replication must be coordinated with cellular physiology to ensure proper genome inheritance. Model bacteria such as the soil-dwelling Bacillus subtilis can achieve a wide range of growth rates in response to nutritional and chemical signals. In order to match the rate of DNA synthesis to the rate of nutrient-mediated cell growth, bacteria regulate the initiation frequency of DNA replication. This control of bacterial DNA replication initiation was first observed over forty years ago, however the molecular basis for this regulation has remained hotly debated. In this paper we test one of the leading models for nutrient-mediated growth rate regulation in bacteria, namely that the abundance of the master DNA replication initiation protein DnaA dictates the frequency of DNA replication events. Critically, our results show that changes in DnaA protein level are not sufficient to account for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis. We then go on to show that there are strong connections between DNA replication and several essential cellular activities, which unexpectedly indicates that there is likely more than one single regulatory pathway involved in coordinating DNA replication with cell physiology. We believe that our work changes thinking regarding this long-standing biological question and reinvigorates the search for the molecular basis of these critical regulatory systems.

Introduction

DNA replication must be coordinated with the cell cycle to ensure proper genome inheritance. For many bacteria cellular physiology dictates the rate of growth and division. In nutrient-rich media that support rapid growth rates, bacteria synthesize DNA more rapidly by increasing the frequency of DNA replication initiation [1]–[3]. This control system is termed nutrient-mediated growth rate regulation and although it has been appreciated for decades, the molecular mechanisms that connect cell physiology with DNA replication initiation remain debatable.

Historically it has been thought that there is a constant cell mass or cell size at the time of bacterial DNA replication initiation and it has been proposed that a positive regulator would accumulate in a growth-dependent manner to trigger DNA replication initiation when cells attained a critical size [4]. However, modern quantitative analysis of single bacterial cells within steady-state populations has shown that the relationship between DNA replication initiation and cell mass is variable, indicating that the control for timing of DNA replication initiation is not governed by a direct connection with mass accumulation [5].

DnaA is the master bacterial DNA replication initiator protein and is a candidate factor to connect cell physiology with DNA synthesis. DnaA is a member of the AAA+ family of ATPases and shares homology with archaeal and eukaryotic initiator proteins. DnaA directly stimulates DNA replication initiation from a single defined origin of replication (oriC) once per cell cycle. Multiple ATP-bound DnaA molecules bind to an array of recognition sequences (DnaA-box 5′-TTATCCACA-3′) within oriC where they assemble into a helical filament that promotes duplex DNA unwinding [6], [7].

Studies in Escherichia coli have suggested that the amount of ATP-bound DnaA dictates the rate of DNA replication initiation. Artificial overexpression of DnaA increases the frequency of DNA replication initiation [8], [9]. Conversely, decreasing the amount of DnaA per cell by synthetically promoting early cell division delays DNA replication initiation and modest increases in DnaA levels alleviate this delay, supporting the view that growth-dependent accumulation of DnaA is the trigger for replication initiation in E. coli [10]. However, it remains uncertain whether the amount of ATP-bound DnaA is the primary regulator that coordinates DNA replication initiation with cell growth in wild-type E. coli cells [11].

In contrast to E. coli, studies in Bacillus subtilis have suggested that the amount of DnaA may not dictate the rate of DNA replication initiation. Artificially decreasing cell size by stimulating cell division (thereby lowering the amount DnaA per cell to ∼70% of wild-type) did not affect DNA replication initiation [10]. Moreover, results from overexpression of DnaA in B. subtilis are not clear. Increased expression of DnaA alone causes cell elongation, cell growth inhibition, and induction of the SOS DNA damage response due to depletion of DnaN because of autoregulation of the dnaA-dnaN operon by DnaA [12]. To circumvent this problem DnaA was co-overexpressed with DnaN, and under this condition DNA replication initiation does increase [12]. However, subsequent experiments demonstrated that overexpression of DnaN alone increases DNA replication initiation by repressing the activity of the regulatory protein YabA (an inhibitor of DnaA)[13], suggesting that this could account for the effect on DNA replication initiation when DnaA and DnaN were co-overexpressed.

In this study we have investigated nutrient-mediated growth rate control of DNA replication initiation in B. subtilis. We find that changes in DnaA protein level are not sufficient to account for nutrient-mediated growth rate regulation of DNA replication initiation, although this regulation does require both DnaA and oriC. We then present evidence suggesting that multiple regulatory systems are involved in coordinating DNA synthesis with cell physiology, and that depending on the nature of the growth limitation, control of DNA replication acts through either oriC-dependent or oriC-independent mechanisms.

Results

Changes in DnaA levels cannot account for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis

Steady-state bacterial growth rates can be manipulated by culturing cells in media that contain differing amounts of nutrients, with rich media supporting faster growth because resources are not required to synthesize cellular building blocks de novo. In response to different nutrient-mediated steady-state growth rates, bacteria control DNA synthesis by varying the frequency of DNA replication initiation while maintaining a constant speed of elongation [1]–[3], [14]. The rate of DNA replication initiation can be determined by marker frequency analysis (i.e. - measuring the ratio of DNA at the replication origin (ori) versus the replication terminus (ter) using quantitative PCR), and Figure 1A shows the positive correlation between DNA replication initiation and nutrient-mediated growth rates (cell doublings per hour measured using spectrophotometry). It is important to state that experimental approaches which change bacterial growth rates without altering the chemical composition of the cell (e.g. – varying temperature) do not influence the rate of DNA replication initiation (Figures 1B, S1; [15], [16]). Thus, varying nutrient availability modulates bacterial physiology, in turn affecting cell growth and DNA replication initiation [3].

It has been reported that DnaA protein level determines the frequency of DNA replication initiation in E. coli [8], [9], therefore we wondered whether the amount of DnaA could account for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis [14]. Western blot analysis shows that DnaA concentration increases with faster steady-state growth rates (Figure 1C; the tubulin homolog FtsZ was used as a loading control because its concentration is growth-rate independent [17],[18]). Since B. subtilis cell size increases as a function of growth rate, the number of DnaA molecules would also be greater in larger cells formed during fast growth (Figure S2)[14]. This conclusion is in agreement with absolute quantification of DnaA proteins per cell determined at different growth rates using mass spectrometry (163-337 molecules at 0.5 doublings/hr; 875-1791 molecules at 1.0 doublings/hr)[18]. These results indicate that the amount of DnaA protein could account for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis.

To directly test whether the amount of DnaA protein determines the rate of DNA replication initiation, the endogenous dnaA gene was placed under the control of an IPTG-inducible promoter (this also alleviated autoregulation of the dnaA-dnaN operon [12]). At near wild-type DnaA levels growth rates were normal, ori:ter ratios were unchanged, and the distribution of origin regions per cell visualized using a TetR-YFP/tetO reporter system was equivalent to wild-type (Figures 2A-C, S4A). In contrast, when the amount of DnaA fell significantly (between ∼50–30% of wild-type, depending upon the media), growth rates slowed, ori:ter ratios dropped, DNA replication was inhibited as judged by origin region localization, and cells became elongated (Figures 2A–C, S4A). As noted above dnaA is located in an operon upstream of dnaN (encoding the sliding clamp component of the replisome) in B. subtilis, and Western blot analysis confirmed that the level of DnaN correlated with the level of DnaA (Figure S3B). Depletion of DnaN can cause replication fork stalling and induction of the SOS DNA damage response, which likely contributes to the slow growth and cell elongation phenotypes observed at low IPTG concentrations [12]. However, replication fork stalling would also be expected to cause an increase in the ori:ter ratio, suggesting that the observed decreases may be an overestimate of the true initiation frequency. We conclude that wild-type DnaA levels are necessary to achieve the proper frequency of DNA replication initiation at both slow and fast steady-state growth rates.

Only modest overexpression of DnaA could be achieved using the IPTG-inducible promoter (Figure 2A), much lower than the changes in DnaA concentration observed at different nutrient-mediated growth rates (Figure 1C). Therefore, to further increase DnaA protein levels a second copy of the dnaA gene was integrated at an ectopic locus under the control of a xylose-inducible promoter (again the endogenous dnaA-dnaN operon was expressed using an IPTG-inducible promoter to avoid autorepression). This strain was grown in media that supported a slow growth rate and varying amounts of xylose were added to induce DnaA (>10 fold overexpression was achieved, which was in the range observed for different nutrient-mediated growth rates; Figures 2E, 4D). When DnaA levels were elevated ∼2–4 fold a modest increase in the ori:ter ratios was observed, although critically the resulting initiation frequencies remained well below the rate generated in rich media (Figures 2D, S4B). These results indicate that changes in DnaA protein levels are not sufficient to account for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis.

Surprisingly, further overexpression of DnaA lead to a dramatic decrease in the ori:ter ratios. To determine whether this inhibition was specific, DnaA was overexpressed in a strain where oriC was inactivated by partial deletion (ΔoriC), the endogenous dnaA-dnaN operon was expressed using a constitutive promoter to avoid autorepression, and genome replication was driven by a plasmid-derived replication origin (oriN; integrated ∼1 kb to the left of oriC) that is recognized and activated by its cognate initiator protein (RepN). It is important to note that while initiation at oriN does not require either oriC or DnaA, the downstream B. subtilis initiation proteins DnaD, DnaB and DnaC (helicase) are necessary for oriN activity [19]. Therefore, if overexpression of DnaA was either inhibiting the expression of genes required for DNA replication (e.g. – nucleotide biosynthesis [20]) or sequestering essential replication factors, then DNA replication initiation from oriN would be expected to decrease. However, overexpression of DnaA in the ΔoriC oriN⁺ background did not alter ori:ter ratios, showing that high overexpression of DnaA specifically inhibits DNA replication initiation at oriC (Figure S4C).

Neither Soj nor YabA nor (p)ppGpp are required for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis

We hypothesized that nutrient-mediated growth rate control of DNA replication initiation could act via regulation of DnaA activity rather than protein abundance. There are two known trans-acting regulators of B. subtilis DnaA during steady-state growth, Soj and YabA. Soj is a dynamic protein that can act as either a negative or a positive regulator of DnaA, depending upon its quaternary state [21]–[23]. YabA is a negative regulator of DnaA that forms a protein bridge between the initiator DnaA and the DNA polymerase sliding clamp processivity factor, DnaN, and is thought to inhibit DNA replication by spatially sequestering DnaA away from the replication origin and by inhibiting DnaA oligomerization [24]–[27]. Interestingly, the number of both proteins per cell was found to positively correlate with growth rate [18].

To determine whether either of these regulatory proteins is required for nutrient-mediated growth rate regulation of DNA replication initiation, single knockout mutants were cultured in a range of media and analyzed using marker frequency analysis. It was found that both of the mutant strains retained the ability to coordinate DNA replication initiation with nutrient-mediated changes in growth rate (Figures 3A, S5A). To test whether Soj and YabA acted redundantly to control the nutrient-mediated activity of DnaA, the double mutant was constructed and analysed by marker frequency analysis. Again proper regulation of DNA replication initiation was maintained in the Δsoj ΔyabA mutant, indicating that neither regulatory protein is required (Figures 3A, S5B). Interestingly, both the single and double mutants displayed a reduced growth rate in rich media, suggesting that the burden of overactive DNA replication initiation may be exacerbated during multifork replication. In the case of the soj mutant it is also possible that the slow growth phenotype is related to its role in chromosome origin segregation [28]–[30].

We also determined whether the alarmone (p)ppGpp, a small molecule induced during nutrient limitation, is required for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis. Marker frequency analysis was performed on a strain lacking the three known (p)ppGpp synthases (RelA, YwaC, YjbM). It was found that regulation of DNA replication initiation was unaffected by the absence of (p)ppGpp, suggesting that (p)ppGpp is not involved in the regulatory mechanisms coordinating DNA replication with nutrient availability during steady-state cell growth (Figures 3B, S5C).

DnaA and oriC are necessary for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis

Since both overexpression of DnaA and deletion of DnaA regulatory proteins did not alter nutrient-mediated growth rate regulation of DNA replication initiation, it was unclear whether DnaA was actually a component of this system. To determine whether DnaA activity at oriC is required, marker frequency analysis was performed in a strain where oriC was inactivated (ΔoriC) and DNA replication initiates from the plasmid-derived oriN. It was found that ori:ter ratios remain constant over a wide range of growth rates in the ΔoriC mutant, indicating that nutrient-mediated growth rate regulation of DNA replication was lost (Figures 4A, S6A). Critically, DNA replication initiation from oriC is unaffected by the addition of oriN (Figures 4B, S6B). This shows that it is the absence of DnaA activity at oriC, rather than the presence of oriN, which accounts for the loss of nutrient-mediated growth rate regulation in the ΔoriC mutant.

However, it could not be concluded whether the ΔoriC mutation acted by removing replication origin function or by deleting a site that is required for the nutrient-mediated growth rate regulation. Therefore, a mutation was introduced into dnaA that alters the critical “arginine finger” residue (Arg²⁶⁴→Ala), thereby disabling DnaA filament assembly and initiation activity (note that a DnaA arginine finger mutant remains competent for DNA binding and ATP binding) [22], [31]. Again the dnaA^R264A mutant strain contains oriN in order to maintain viability. Like the ΔoriC mutant the DnaA^R264A variant also lost growth rate regulation in response to nutrient availability, indicating that DnaA activity at oriC is necessary for growth rate regulation in B. subtilis (Figures 4C, S6C). Moreover, since the ori:ter ratios of the ΔoriC and dnaA^R264A mutants remains constant during nutrient-mediated growth rate changes and since DNA replication elongation speed is independent of the nutrient-mediated growth rate [2], [14], the results suggest that within a population of cells the average frequency of DNA replication initiation at oriN is independent of the nutrient-mediated growth rate.

Western blot analysis showed that in rich media there was less DnaA and DnaN in the ΔoriC strain, whereas conversely there was more DnaA and DnaN in the dnaA^R264A mutant (Figure S3C). The latter result suggests that autoregulation of the dnaA promoter requires ATP-dependent filament formation by DnaA, however, the former result was more puzzling. To investigate this further the amount of DnaA in the ΔoriC strain was determined over a range of nutrient-mediated growth rates. While the concentration of DnaA was observed to increase as a function of growth rate in the wild-type strain, DnaA levels did not display the same correlation with growth rate in the ΔoriC mutant (Figure 4D). This suggests that in the ΔoriC mutant nutrient-mediated growth rate-dependent expression of DnaA is lost either because DNA replication initiation from oriN is constitutive or because the deletion within oriC affects dnaA expression (although this region is downstream of the dnaA gene).

The apparent decrease in growth rates observed for strains initiating DNA replication solely through oriN, particularly in rich media (Figures 4A, 4C, S6A, S6C), is likely due to the formation of cells lacking DNA as a direct consequence of decoupling DNA replication initiation from growth rate (Figure 4E). This result underscores the importance of growth rate regulation of DNA replication initiation to ensure bacterial fitness.

Slowing growth rate by limiting essential cellular activities inhibits DNA replication

Since nutrient-mediated growth rate regulation of DNA replication initiation did not appear to act through either DnaA protein accumulation or known DnaA regulators, we considered alternative possibilities for how DNA replication could be connected to cell physiology. One hypothesis was that this regulation could be linked to metabolism, either through the amount of a metabolic intermediate or the activity of a critical enzyme. Genetic evidence has suggested a relationship between several glycolytic enzymes and DNA replication in B. subtilis and E. coli [32], [33], but it has not been established whether these connections act directly at the level of DnaA-dependent initiation. ATP would be another rational candidate since DnaA is an ATP-dependent protein, but it has been found that the concentration of ATP in B. subtilis (as well as in E. coli) is invariant over a wide range of growth rates (L. Krasny and R. Gourse, personal communication; [34], [35]). Another hypothesis was that this regulation could be linked to the synthesis of an essential cellular complex, such as the ribosome or the cell membrane [36], [37]. In this way a bacterial cell would integrate nutritional information based on the availability of multiple substrates required to construct such macromolecules.

In order to identify possible routes through which nutrient availability could impact DNA replication we analyzed a range of genetically altered strains, targeting respiration, central carbon metabolism, protein synthesis, fatty acid synthesis, and phospholipid synthesis (Table 1), that all manifest decreased steady-state growth rates in rich complex media. Genes were either disrupted by antibiotic cassettes or depleted using regulated expression systems; importantly, depletion of essential genes was not lethal under the experimental conditions used. Knock-out strains were compared to wild-type while depletion strains were analyzed without and with inducer (indicated in Figures 5, 6, S7, S8 with “-” and “+”, respectively). DNA replication was measured using marker frequency analysis. Strikingly, in all of the strains examined the ori:ter ratio decreased to match the slower growth rates caused by gene disruption/depletion (Figures 5–6, S7–S8; black symbols).

Evidence for an oriC-independent response to changes in growth rate

The uniform response of DNA replication in slow growing mutants suggested that a single mechanism might account for this regulation, in accord with nutrient-mediated regulation of DNA replication initiation (Figures 4, S6). To examine this hypothesis the deletion and depletion strains were crossed into the ΔoriC strain that initiates DNA replication using oriN. For several mutants (ndh, gapA, pdhB, fabHA, plsC and ltaS) the ori:ter ratio was not significantly affected (≤1/10 of the percentage decrease observed for oriC⁺), suggesting that the regulatory signal specifically targeted DNA replication initiation at oriC (Figures 5, S7; red symbols). However, there were a number of mutants (pykA, pgsA, and multiple ribosomal protein genes) that produced a marked decrease in the ori:ter ratio of the ΔoriC strain (≥1/2 of the percentage decrease observed for oriC⁺), suggesting that in these cases DNA replication was being regulated through an oriC-independent mechanism (Figures 6A–C, S8A-C; red symbols). Interestingly, in some cases manipulation of different genes within a single biological pathway (e.g. – carbon metabolism or phospholipid synthesis) resulted in the regulation of DNA replication through the different regulatory systems.

Evidence for a DnaA-independent response to changes in growth rate

Since the ΔoriC oriN⁺ strain does not require DnaA activity to initiate DNA replication, it suggested that the observed oriC-independent growth rate regulation might be DnaA-independent. To address this possibility the oriC-independent mutants (Figures 6A–C, S8A–C) were crossed into a ΔdnaA strain that initiates DNA replication using oriN (Figure S3C). When PykA was depleted in the ΔdnaA mutant the ori:ter ratio no longer decreased, indicating that DnaA was required for this response, although apparently not for its role in origin recognition and DNA unwinding (Figures 6D, S8D; green symbols). In contrast, when either ribosomal genes were deleted or PgsA was depleted in the ΔdnaA mutant the ori:ter ratios did decrease, suggesting that DnaA-independent mechanisms act under these conditions (Figures 6E–F, S8E–F; green symbols). Taken together, the genetic analysis reveals that in B. subtilis there is likely more than one regulatory system linking DNA replication with cell growth.

Slowing growth rate by targeting essential cellular activities with small molecules inhibits DNA replication initiation

The importance of growth rate regulation of DNA replication in response to nutrient availability is self-evident, but the biological relevance of growth rate regulation of DNA replication in response to genetic manipulations is less clear. To address this issue we evaluated the response of DNA replication to sublethal concentrations of antibiotics that produced slow steady-state growth rates. We chose small molecules that inhibit either fatty acid synthesis (cerulenin) or protein synthesis (chloramphenicol) because our genetic analyses indicated that the former regulated DNA replication through oriC while the latter acted independently of both oriC and DnaA. Incubation with either antibiotic caused a decrease in the ori:ter ratios in the wild-type strain, showing that growth rate regulation of DNA replication in response to genetic perturbations of essential cellular activities is physiologically relevant (Figures 7A–B, S9A–B; black symbols).

To further assess whether changes in DNA replication caused by these small molecules reflected the results using genetic approaches, the ΔoriC oriN⁺ strain was analyzed. Only chloramphenicol elicited a significant decrease in the ori:ter ratios in the ΔoriC strain (Figures 7A–B, S9A–B; red symbols). Finally, the ΔdnaA oriN⁺ strain was analyzed in the presence of chloramphenicol and again the ori:ter ratio decreased (Figures 7B, S9B). This result is fully consistent with the data from ribosomal gene deletions and indicates that regulation of DNA replication in response to perturbed ribosome activity is DnaA-independent.

Discussion

We have found that nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis requires both DnaA and oriC. To our knowledge this is the first time that a specific DNA replication initiation protein has been shown to play an essential role in this regulatory system, and because DnaA is the earliest acting initiation factor we propose that DnaA is the target for the nutrient-mediated growth rate regulatory system. Critically however, we show that changes in DnaA protein level are not sufficient to account for nutrient-mediated growth rate regulation of DNA replication initiation in B. subtilis. This is in contrast to the generally accepted mechanism for control of bacterial DNA replication initiation based on work using E. coli [8], [9].

B. subtilis contains a bipartite origin that flanks the dnaA gene (incA and incB regions containing the dnaA promoter lie 1.3 kb upstream of the incC region which contains the DNA unwinding element) [38]. When the expression of the dnaA-dnaN operon was placed under the control of the inducible P_spac promoter in order to test the effect of DnaA overexpression on DNA replication initiation, a ∼9kb plasmid was recombined upstream of dnaA by single cross-over. Therefore, integration of this vector resulted in the considerable displacement of the two origin regions from one another without any significant consequence. It will be extremely interesting to determine the maximum and minimum distances that the inc regions can be moved, as well as ascertaining the role of the upstream region in DNA replication initiation.

We have shown that neither of the known DnaA regulatory proteins present during vegetative growth, Soj and YabA, are required for nutrient-mediated growth rate regulation of DNA replication initiation. We have also determined that the alarmone (p)ppGpp is not required for this regulation, consistent with a previous report that induction of the stringent response inhibits DNA replication elongation but not initiation in B. subtilis [39], [40]. This result marks an apparent distinction between the role of (p)ppGpp in B. subtilis and in proteobacteria such as E. coli and Caulobacter crescentus where (p)ppGpp has been shown to regulate DNA replication initiation [41], [42].

The use of genetic manipulations and small molecule inhibitors presented here reinforce and extend previously observed connections for bacterial DNA replication with central carbon metabolism [32], [33] and with phospholipid synthesis [43], [44]. In addition our work identifies new links for B. subtilis DNA replication with respiration, fatty acid synthesis, lipoteichoic acid synthesis, and ribosome biosynthesis. The results indicate that growth rate regulation of DNA replication in B. subtilis can be controlled through either oriC-dependent, oriC-independent/DnaA-dependent, or oriC-independent/DnaA-independent mechanisms (summarized in Figure 7C). Based on these novel findings we propose that multiple systems coordinate DNA replication with bacterial cell growth, with distinct regulators responding to diverse physiological and chemical changes. This model deviates from the long-standing concept of a single universal cellular property utilized to link bacterial DNA replication with cell growth [4].

Since nutrient-mediated growth rate regulation of DNA replication initiation requires DnaA activity at oriC, this suggests that factors affecting DNA synthesis through an oriC-independent mechanism (PykA, PgsA, and ribosomal proteins) are unlikely to be responsible for the nutrient sensing system. We suspect that nutrient-mediated regulation may be influenced by more than one control system, thereby forming a robust network capable of integrating information from multiple metabolic and cellular sources.

We note that for deletion/depletion mutants regulating DNA replication through oriC-dependent and oriC-independent/DnaA-dependent mechanisms, ori:ter ratios either remained constant or slightly increased in the ΔoriC and ΔdnaA strains, respectively (Figures 5, 6D, S7, S8D). Because the average initiation frequency of oriN appears to be growth rate independent (Figures 4A, 4C, S6A, S6C), the measured ori:ter ratios of these strains indicates that replication elongation speeds are either not being affected or are slightly decreasing. Therefore, for both oriC-dependent and oriC-independent/DnaA-dependent regulatory mechanisms, the observed decrease in ori:ter ratios in the oriC⁺ strain most likely reflects inhibition of DNA replication initiation. In contrast, for the oriC-independent/DnaA-independent mutants where the ori:ter ratio was decreased when DNA replication initiated from oriN, this difference could be due to a change in DNA replication elongation (although this would mean that the elongation speed was increased).

Our current aim is to determine the molecular mechanisms underlying each system that coordinates DNA replication with cell growth. We hypothesize that the oriC-dependent regulatory system targets DnaA activity at oriC. We speculate that the oriC-independent/DnaA-dependent regulatory system could influence DNA replication initiation by affecting the abundance or activity of a downstream replication initiation factor. For example, DnaA is also a transcription factor that is thought to directly regulate the expression of>50 genes, including dnaB [20], [45]. Alternatively, DnaA could act by titrating initiation factors away from oriN. Lastly, in the case of the oriC-independent/DnaA-independent regulatory system it needs to be established whether DNA replication is affected at the step of initiation or elongation, after which the role of appropriate candidate proteins can be investigated.

Materials and Methods

Strains and plasmids

Strains are listed in Table S1. Plasmids are listed in Table S2 and Table S3.

Media and chemicals

Nutrient agar (NA; Oxoid) was used for routine selection and maintenance of both B. subtilis and E. coli strains. For experiments in B. subtilis cells were grown using a range of media (using the following concentrations unless otherwise noted): Luria-Bertani (LB) medium, Antibiotic 3 (PAB) medium, Brain-Heart Infusion (Bacto), or defined minimal medium base (Spizizen minimal salts supplemented with Fe-NH₄-citrate (1 µg/ml), MgSO₄ (6 mM), CaCl₂ (100 µM), MnSO₄ (130 µM), ZnCl₂ (1 µM), thiamine (2 µM)) supplemented with casein hydrolysate (200 µg/ml) and/or various carbon sources (succinate (1%), glycerol (0.5%), glucose (0.5%)). Supplements were added as required: tryptophan (20 µg/ml), phenylalanine (40 µg/ml), chloramphenicol (5 µg/ml), erythromycin (1 µg/ml), kanamycin (2 µg/ml), spectinomycin (50 µg/ml), tetracycline (10 µg/ml), zeocin (10 µg/ml). Unless otherwise stated all chemicals and reagents were obtained from Sigma-Aldrich.

Marker frequency analysis

Sodium azide (0.5%; Sigma) was added to exponentially growing cells to prevent further metabolism. Chromosomal DNA was isolated using a DNeasy Blood and Tissue Kit (Qiagen). The DNA replication origin (oriC) region was amplified using primers 5′-GAATTCCTTCAGGCCATTGA-3′ and 5′-GATTTCTGGCGAATTGGAAG-3′; the DNA replication terminus (ter) region was amplified using primers 5′-TCCATATCCTCGCTCCTACG-3′ and 5′-ATTCTGCTGATGTGCAATGG-3′. Either Rotor-Gene SYBR Green (Qiagen) or GoTaq (Promega) qPCR mix was used for PCR reactions. Q-PCR was performed in a Rotor-Gene Q Instrument (Qiagen). By use of crossing points (C_T) and PCR efficiency a relative quantification analysis (ΔΔC_T) was performed using Rotor-Gene Software version 2.0.2 (Qiagen) to determine the origin:terminus (ori:ter) ratio of each sample. These results were normalized to the ori:ter ratio of a DNA sample from B. subtilis spores which only contain one chromosome and thus have an ori/ter ratio of 1.

Microscopy

To visualize cells during exponential growth starter cultures were grown overnight and then diluted 1∶100 into fresh medium and allowed to achieve at least three doublings before observation. Cells were mounted on ∼1.2% agar pads (0.25× minimal medium base) and a 0.13–0.17 mm glass coverslip (VWR) was placed on top. To visualize individual cells the cell membrane was stained with either 2 µg/ml Nile Red (Sigma) or 0.4 µg/ml FM5-95 (Molecular Probes). To visualize nucleoids DNA was stained with 2 µg/ml 4′-6-diamidino-2-phenylindole (DAPI) (Sigma). Microscopy was performed on an inverted epifluorescence microscope (Nikon Ti) fitted with a Plan-Apochromat objective (Nikon DM 100x/1.40 Oil Ph3). Light was transmitted from a 300 Watt xenon arc-lamp through a liquid light guide (Sutter Instruments) and images were collected using a CoolSnap HQ² cooled CCD camera (Photometrics). All filters were Modified Magnetron ET Sets from Chroma and details are available upon request. Digital images were acquired and analysed using METAMORPH software (version V.6.2r6). Analysis was performed using ImageJ software: foci counting utilized the particle analysis plugin; cell lengths and widths were measured using the ObjectJ plugin.

Western blot analysis

Proteins were separated by electrophoresis using a NuPAGE 4-12% Bis-Tris gradient gel run in MES buffer (Life Technologies) and transferred to a Hybond-P PVDF membrane (GE Healthcare) using a semi-dry apparatus (Hoefer Scientific Instruments). Proteins of interest were probed with polyclonal primary antibodies and then detected with an anti-rabbit horseradish peroxidase-linked secondary antibody using an ImageQuant LAS 4000 mini digital imaging system (GE Healthcare). Quantification was determined by densitometry using Image J software. Figure S3A shows that detection of DnaA, FtsZ and DnaN was within a linear range.

Supporting Information

Attachment 1

Attachment 2

Attachment 3

Attachment 4

Attachment 5

Attachment 6

Attachment 7

Attachment 8

Attachment 9

Attachment 10

Attachment 11

Attachment 12

Attachment 13