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Journal of Bacteriology, July 2004, p. 4655-4664, Vol. 186, No. 14
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.14.4655-4664.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Response of Bacillus subtilis to Nitric Oxide and the Nitrosating Agent Sodium Nitroprusside{dagger}

Charles M. Moore,1 Michiko M. Nakano,2 Tao Wang,3 Rick W. Ye,3 and John D. Helmann1*

Department of Microbiology, Cornell University, Ithaca, New York 14853,1 Department of Environmental and Biomolecular Systems, Oregon Graduate Institute School of Science and Engineering, Oregon Health & Science University, Beaverton, Oregon 97006,2 Experimental Station E328/148B, DuPont Central Research and Development, Wilmington, Delaware 198803

Received 23 February 2004/ Accepted 6 April 2004


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ABSTRACT
 
We examined the effects of nitric oxide (NO) and sodium nitroprusside (SNP) on Bacillus subtilis physiology and gene expression. In aerobically growing cultures, cell death was most pronounced when NO gas was added incrementally rather than as a single bolus, suggesting that the length of exposure was important in determining cell survival. DNA microarrays, Northern hybridizations, and RNA slot blot analyses were employed to characterize the global transcriptional response of B. subtilis to NO and SNP. Under both aerobic and anaerobic conditions the gene most highly induced by NO was hmp, a flavohemoglobin known to protect bacteria from NO stress. Anaerobically, NO also induced genes repressed by the Fe(II)-containing metalloregulators, Fur and PerR, consistent with the known ability of NO to nitrosylate the Fe(II) center in Fur. In support of this model, we demonstrate that NO fails to induce PerR-regulated genes under growth conditions that favor the formation of PerR:Mn(II) rather than PerR:Fe(II). Aerobically, NO gas induced hmp, the {sigma}B general stress regulon, and, to a lesser extent, the Fur and PerR regulons. Surprisingly, NO gas induced the {sigma}B regulon via the energy branch of the {sigma}B regulatory cascade while induction by SNP was mediated by the environmental stress branch. This emphasizes that NO and SNP elicit genetically distinct stress responses.


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INTRODUCTION
 
Nitric oxide (NO) is a lipophilic, freely diffusible radical that can inhibit enzymes, damage DNA, initiate lipid peroxidation, and exacerbate peroxide-induced damage (49, 53, 66). NO chemistry can be divided into those reactions that occur between NO and biomolecules (direct effects) and those reactions that can only occur subsequent to NO reacting with oxygen or superoxide to form reactive nitrogen oxide species (RNOS) (indirect effects). Direct effects of NO include the formation of metal-nitrosyl complexes (63) and reactions with lipid-derived (50) and other high-energy radicals (34). For example, NO coordinates free or enzyme-bound Fe(II) to form Fe-NO as described for cytochrome P450 (36, 43, 64). Fe nitrosylation leads to altered activity of at least two bacterial metalloregulatory proteins: Fur and Fnr (13, 14). Indirect effects occur after NO reacts with either oxygen to generate N2O3 or superoxide to generate the highly reactive oxidant peroxynitrite (OONO–). Peroxynitrite rapidly decomposes to form nitrate (NO3–), hydroxyl radical ( · OH), and nitrogen dioxide radical ( · NO2). Some RNOS have the propensity to react with thiol groups and amines to form S-nitrosothiols and nitrosamines.

Bacteria encounter NO from a variety of sources. Macrophages of the mammalian immune system generate NO with an inducible NO synthase as part of their arsenal employed against microbial pathogens (38, 57). NO synthases have also been identified in some bacteria, including Bacillus subtilis (1, 2), although their function in many cases remains elusive. Denitrifying bacteria produce NO as an intermediate in the reduction pathway from nitrate to dinitrogen (33). B. subtilis is not capable of denitrification, yet it coexists with denitrifying bacteria in subsurface environments. It is therefore likely that B. subtilis has developed a targeted response to exogenously and perhaps endogenously produced NO.

Several bacterial enzymes can alleviate NO stress. The Escherichia coli flavohemoglobin Hmp has NO reductase activity under anaerobic conditions (31) and NO dioxygenase (22) or denitrosylase activity (25) under aerobic conditions; however, only the aerobic Hmp activities appear to confer NO stress resistance (20). In addition, both E. coli and Salmonella enterica hmp mutants are hypersensitive to NO (41, 56, 57). Nakano showed that B. subtilis hmp is regulated by ResDE, a two-component system that regulates several genes required for anaerobic growth (46). Another hemoglobin, HbN from Mycobacterium tuberculosis, displays NO dioxygenase activity and protects heterologous hosts (E. coli and Mycobacterium smegmatis) from NO challenge (51). In response to NO, E. coli expresses a second NO reductase (norVW) which is under positive control by the regulator NorR (21, 45). Inactivation of norVW or norR results in an NO-sensitive mutant (30). In addition, alkyl hydroperoxide reductase (AhpC) of S. enterica, Helicobacter pylori, and M. tuberculosis protects cells from peroxynitrite insult (7, 12, 40).

Here we investigate the physiological and genetic responses of B. subtilis to NO (gas) and sodium nitroprusside (SNP). We demonstrate that the NO stimulon is largely defined by hmp and the Fur, PerR, and {sigma}B regulons. The pattern of induction is significantly altered by the presence of oxygen and by changes in the metal ion content of the growth medium. Furthermore, we show that the pathway of induction for the {sigma}B regulon for NO (gas) is different from that for SNP.


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MATERIALS AND METHODS
 
Media and growth conditions. B. subtilis strains were maintained on Luria-Bertani (LB) agar, and all experiments (except Fig. 7; see below) were conducted at 37°C in fermentation broth consisting of 2x yeast tryptone broth supplemented with 0.5% glucose and 0.5% pyruvate. Kanamycin (40 µg ml–1) or spectinomycin (100 µg ml–1) was used for selection of various B. subtilis strains. All inoculations were made at 1% volume from an overnight culture grown in LB or anaerobic minimal medium to an optical density at 600 nm (OD600) of approximately 0.75 or 0.40, respectively.



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  		FIG. 7. A fur mutant is sensitive to NO and SNP. Anaerobic growth of B. subtilis wild type (squares) and fur mutant (circles) in fermentation broth either unamended (black), with 2 mM SNP (gray), or with 50 µM NO (white). SNP and NO was added at time zero after cultures had been purged for 30 min with N2.

To influence the metallation of PerR (see Fig. 8), we used anaerobic minimal medium consisting of 40 mM potassium morpholinepropanesulfonic acid (MOPS) (adjusted to pH 7.4 with KOH), 2 mM potassium phosphate buffer (pH 7.0), 2% (wt/vol) glucose, (NH4)2SO4 (2 g/liter), MgSO4 · 7H2O (0.2 g/liter), trisodium citrate · H2O (1 g/liter), potassium glutamate (1 g/liter), tryptophan (10 mg/liter), 3 nM (NH4)6Mo7O24, 400 nM H3BO3, 30 nM CoCl2, 10 nM CuSO4, 10 nM ZnSO4. MnCl2 and FeSO4 were added at various concentrations as noted.



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  		FIG. 8. PerR:Fe responds to NO and SNP while PerR:Mn does not. Northern hybridization of total RNA extracted from wild-type cells either induced by 2 mM sodium nitroprusside (+) or uninduced (–) and grown in minimal medium supplemented with 10 µM Fe(II) (Fe), 10 µM Mn(II) (Mn), or both metal concentrations simultaneously (Fe Mn). Membranes were hybridized with DNA probes for hmp (A), katA (B), or mrgA (C). Note that when 10 µM Fe(II) is present in the growth medium, the PerR-regulated genes katA and mrgA are induced by SNP. When Fe(II) is not added to the growth medium and Mn(II) is added at a concentration of 10 µM, these genes are no longer induced. Induction of hmp occurs at all metal concentrations tested.

NO-saturated water solution was prepared by sparging 50 ml of water with N2 for 30 min to remove O2 and NO gas for 30 min in 100-ml gas-tight crimp seal vials. The resulting solution was assumed to be 1.6 mM (at 37°C) with respect to NO (62).

Bacterial killing by NO. Cultures were grown aerobically to an OD600 of approximately 0.3, and NO-saturated water was added as an initial bolus or at even time intervals over a 20-min period. Twenty minutes after the initial addition of NO, cells were diluted in series and 10 µl was applied to LB agar and incubated overnight prior to viability assessment. All experiments were performed in duplicate.

Microarray analyses: RNA isolation, cDNA synthesis, and slide hybridization. Anaerobic growth for microarray analysis was performed in 50-ml serum bottles (Bellco Glass) sealed with black butyl rubber crimp sealed stoppers as previously described (47). Bottles were filled with 50 ml of fermentation broth and were inoculated, and the headspace was sparged with N2 and was sealed. Bottles were rotated at 37°C. At an OD600 of approximately 0.3, NO was added to a final concentration of 50 µM from gas-saturated water. After 15 min of incubation, samples were centrifuged at 9,000 rpm in a Sorval RC-5B centrifuge (when the speed attained 9,000 rpm, centrifugation was stopped). The supernatant fraction was discarded, and the pellet was frozen and stored at –80°C. Total RNA was extracted using an RNeasy minikit (QIAGEN). Contaminating DNA was removed with a DNA-free kit (Ambion), and RNA was stored at –80°C for future use. Experiments performed under aerobic conditions were identical to those performed under anaerobic conditions, with the following exceptions. Aerobic growth experiments were performed in 250-ml flasks containing 50 ml of fermentation broth shaken at 200 rpm at 37°C. At an OD600 of approximately 0.3, 50 µM NO was added to the aerobic cultures 5 min prior to centrifugation. In some experiments, 25 µM NO was added twice to aerobic cultures with 2 min between additions and samples were harvested 5 min after the initial NO addition.

The effects of NO were analyzed by competitive hybridization of fluorescently labeled cDNA samples to DNA microarrays that contained 4,020 PCR products spotted in duplicate on each glass slide. Slide construction and labeling (using random hexamers) was performed as previously described (67, 68). RNA preparations were used to generate both Cy3- and Cy5-labeled cDNAs, and all competitive hybridizations were performed twice to control for any differences in labeling with the two fluorophores. Because all PCR products are spotted twice on each slide, all signal intensities and calculated ratios are the averages of four values.

Microarray data analysis. Signal intensities were detected and quantified with ArrayVision software (Molecular Dynamics) and were assembled into Excel spreadsheets (Microsoft). Mean fluorescence intensity was set to 1.0, with a value of 0.1 corresponding to background. The mean expression ratio and standard deviation were calculated as wild-type NO-treated/wild type for each experiment. Data were filtered to remove genes with high variability (standard deviation equal to or greater than the mean).

Northern hybridization. Anaerobic growth for Northern hybridization was performed in 16-ml Hungate tubes (Bellco Glass Co.) sealed with black butyl rubber stoppers as previously described (15). Tubes were filled with 15 ml of fermentation broth or anaerobic minimal medium, the headspace was sparged with N2, and the tubes were sealed. Tubes were gently rocked at 37°C on their sides to prevent cell aggregation. At an OD600 of approximately 0.4, SNP (Sigma) was added to a final concentration of 2 mM (from a 200 mM stock solution made fresh daily). Cells were grown in the presence of SNP for 2 h, and the cells were harvested. Aerobic growth experiments were performed in 250-ml flasks containing 50 ml of fermentation broth shaken (200 rpm) at 37°C. All other aerobic procedures were identical to anaerobic experiments.

To isolate RNA, 10 ml of bacterial culture was added to 2 ml of 95% ethanol-5% phenol (pH 4.5) on ice and centrifuged for 5 min at 5,000 rpm in a Sorval RC-5B centrifuge (4°C). The supernatant was discarded and the pellet was snap frozen and stored at –80°C. Total RNA was extracted by using an RNeasy mini kit (QIAGEN) according to the manufacturer's instructions. RNA was quantified by A260 with a Lambda 25 UV/VIS Spectrophotometer (Perkin Elmer). DNA probes were constructed via PCR using HotStarTaq Master Mix kit (QIAGEN) according to the manufacturer's instructions, using 100 pM primers with 50 ng of CU1065 chromosomal DNA template. The temperature profile was as follows: 15 min at 95°C (1 cycle); 30 s at 94°C; 1 min at 50 to 60°C (primer dependent); 1 min at 72°C (35 cycles); and 3 min at 72°C (1 cycle) in a Master Cycle Gradient Thermocycler (Eppendorf). Primers (sequences available on request) contained either a HindIII or EcoRI restriction site for subsequent use in the probe labeling reaction. HindIII- and EcoRI (New England Biolabs)-restricted DNA probes were purified with a PCR cleanup kit (QIAGEN) and were labeled with [{alpha}-32P]dATP via Klenow fragment of DNA polymerase I (Klenow exo–; New England Biolabs) fill-in of single-stranded overhangs (2 h at 37°C). Unincorporated [{alpha}-32P]dATP was removed by using Nuc Away Spin Columns (Ambion) according to manufacturer's instructions. Northern blot hybridizations were performed using a Northern Max Northern Hybridization kit (Ambion) with Zeta-Probe Blotting Membrane (Bio-Rad) according to the manufacturers' instructions, except that hybridizations were performed overnight at 50°C (42°C for the mrgA probe) and membranes were washed two times at room temperature in 2x SSPE (Ambion) (1x SSPE is 0.18 M NaCl, 10 mM NaH2PO4, and 1 mM EDTA [pH 7.7]) followed by two washes at 50°C (42°C for mrgA probe) in 0.1x SSPE. Northern hybridization membranes were developed on a Storm 840 PhosphorImage scanner (Molecular Dynamics) after a 12- to 24-h exposure of a PhosphorImage screen.

RNA slot blot hybridization. Aerobic cultures were grown in 50 ml of fermentation broth and were shaken at 200 rpm at 37°C in 250-ml flasks. NO (25 µM) was added at an OD600 of approximately 0.3 and again 2 min after initial addition. Anaerobic experiments were conducted in Hungate tubes as described above and received 50 µM (final) NO at an OD600 of approximately 0.3. Ten milliliters of culture was removed 5, 10, 15, and 25 min after initial NO addition. RNA was extracted from two independent samples for each condition and was quantified as described above. RNA samples containing 1, 0.1, 0.01, and 0.001 µg were loaded onto nitrocellulose membrane (a 0.1-µg signal was used to generate Fig. 5). Probe construction, membrane hybridization, washing, and development were as described for Northern hybridization. PhosphorImage signals were quantified by using ImageQuaNT software version 4.2 (Molecular Dynamics).



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  		FIG. 5. Time course induction of hmp, ykuO (Fur regulated), mrgA (PerR regulated), and csbC ({sigma}B regulated) by 50 µM NO under aerobic conditions (circles) and anaerobic conditions (squares). A concentration of 50 µM NO was added as two 25 µM additions separated by 2 min for aerobic cultures. Time zero is the initial addition of NO at an OD600 of approximately 0.3. Data points represent fold induction (n = 2). Transcript levels were measured by RNA slot blot analysis.

Aerobic respiration assay. Oxygen utilization was monitored by using a Digital Model 10 Controller Clark type O2 electrode (Rank Brothers, Ltd.). Cells were grown in 50 ml of fermentation broth to an OD600 of approximately 0.3, at which point 10 ml of culture was centrifuged at 10,000 rpm in a Sorval RC-5B centrifuge and resuspended in 100 µl of a buffer solution containing 50 µM MOPS (pH 7.4) and 50 µM NaCl. The cell resuspensions were added to 2.9 ml of the same buffer solution with 30 µM glucose in an incubation chamber at 37°C. NO was added into the reaction vessel syringe port when 75% of the initial [O2] remained. Sodium dithionite was added to cells at the end of each experiment to measure residual O2.

ß-Galactosidase assays. Overnight cultures were grown in LB to late logarithmic phase and were used to inoculate 50 ml of fermentation broth in 250-ml flasks (aerobic) or 15 ml of fermentation broth in Hungate tubes (anaerobic). Aerobic cultures were shaken at 200 rpm at 37°C and were grown to an OD600 of approximately 0.2, at which time the cultures were amended with SNP to a final concentration of 2 mM or NO to a final concentration of 50 µM (two additions of 25 µM with 2 min between additions). Anaerobic cultures received either 2 mM SNP or 50 µM NO (as indicated). Cells were harvested in 1-ml volumes after 15 min of incubation and were centrifuged at 13,000 rpm in an Eppendorf 5415 D centrifuge for 30 s at 4°C prior to overnight storage at –80°C. Cells were assayed for ß-galactosidase activity as previously described (42).

Supplementary material. Complete microarray datasets are available in Microsoft Excel and Tab-delimited format at http://www.micro.cornell.edu/faculty.Jhelmann.html. This site also contains Fig. S1.


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RESULTS AND DISCUSSION
 
Physiological effects of NO during aerobic growth. We have measured the physiological effects of NO (gas) on B. subtilis by following cell viability, growth rate, and respiration. To measure effects on cell viability, NO was added to aerobically growing B. subtilis cultures to 50 or 200 µM, either as a single bolus or divided into a series of smaller additions spread out over 20 min. Single additions of either 50 or 200 µM NO were tolerated with no significant loss in viability (Fig. 1A). Interestingly, addition of 25 µM NO two times (2 min between additions) led to little or no loss in viability, whereas addition of 10 µM NO five times (4 min between additions) led to a nearly 100-fold reduction in viability. Similarly, 200 µM NO was well tolerated, whereas addition of 50 µM NO four times (5 min between each addition) resulted in a 104-fold loss of viability (Fig. 1A). These results demonstrate that NO is a much more effective cell stress agent when applied to cells over time than as a single large addition. This effect may be due to the rapid loss of NO that occurs via degassing in open systems (such as those used during aerobic culture) or could reflect a process of sensitization to the killing action of NO.



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  		FIG. 1. NO effects on B. subtilis physiology. (A) NO effect on B. subtilis viability. B. subtilis serial dilutions were performed 20 min after initial NO addition, and 10 µl was applied to an LB agar plate and allowed to grow at 37°C overnight. Rows designated 50 µM bolus and 200 µM bolus were both single initial additions. (25 µM NO) x 2, two additions separated by 2 min; (10 µM NO) x 5, five additions separated by 4 min; (50 µM NO) x 4, four additions separated by 5 min. (B) NO effect on B. subtilis aerobic growth. Squares, untreated; diamonds, 50 µM NO bolus; triangles, 50 µM NO added as two 25 µM additions with 2 min between additions; circles, 200 µM NO added as four 50 µM additions with 5 min between additions. (C) NO effect on B. subtilis anaerobic growth. Squares, untreated; diamonds, 50 µM NO bolus; triangles, 100 µM NO bolus; circles, 200 µM bolus.

To measure the effects of NO on growth rate, we monitored cell density after addition of sublethal concentrations of NO to mid-logarithmic-phase cells. NO temporarily suspended growth of aerobic cultures and resulted in reduced growth rates at high concentrations. Addition of 50 µM NO as a bolus or as two 25 µM additions with 2 min between additions slowed growth for approximately 15 min before cultures resumed growth at rates similar to that of an untreated control (Fig. 1B). These conditions produced minimal killing in the viability experiments, and therefore the pause observed in growth rate may be the result of reversible respiratory chain inhibition by NO (see below). In contrast, 50 µM NO added four times (5 min between additions) significantly reduced the growth rate of the culture for the duration of the experiment. NO delivery in this manner resulted in significant killing (Fig. 1A) and may have generated culture conditions (e.g., nitrosylation of media constituents, lysis of cells) that resulted in poor growth for those cells that survived the NO stress.

NO inhibits aerobic respiration by reversibly binding to cytochromes (5, 6, 63). To determine if the transient inhibition of growth observed upon NO addition correlated with inhibition of aerobic respiration, oxygen consumption was monitored with a Clark type O2 electrode. The addition of 25 µM NO to actively respiring cells resulted in a temporary inhibition of respiration (7 min in duration), after which respiration resumed at pretreatment rates (Fig. 2). Interestingly, 50 µM NO inhibited respiration for approximately the same time period (7 min). After respiration resumed, a steady decrease in rate was observed with ~40 µM O2 remaining unrespired. This suggests that 50 µM NO addition caused some irreversible damage to the cells, possibly from RNOS formed during the experiment, which damaged the cells in a time-dependent manner. Alternatively, the high-affinity aerobic respiratory system, characterized by two bd-type cytochrome oxidases (61, 65), may have been irreversibly damaged while inhibition of the lower affinity systems, characterized by two heme-copper-type cytochrome oxidases (cytochromes aa3 and caa3) (59, 61), was temporary. Indeed, NO-induced damage may account for the aerobic induction by NO of two genes (cydC and cydD) involved with the synthesis of one of the bd-type cytochrome oxidases (65) (Table 1).



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  		FIG. 2. NO inhibits aerobic respiration of B. subtilis in a reversible manner. Oxygen depletion in 3 ml of B. subtilis culture was measured with a Clark type O2 electrode with 25 or 50 µM NO added at 75% O2 saturation (indicated by arrows). Addition of sodium dithionite crystals to reduce remaining O2 is indicated by an asterisk.


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  		TABLE 1. Miscellaneous genes induced more than fivefold by 50 µM NO under either aerobic or anaerobic growth conditions

These results suggest that transient growth arrest by NO could be due to inhibition of respiration. However, it is important to note that the respiration inhibition experiments were performed in a closed vessel without continual aeration. Therefore, degassing of NO was prevented and all added NO was available to react with O2 and cellular material contained in the system. In contrast, cell viability and aerobic growth inhibition experiments were conducted in an open system. This difference in experimental conditions undoubtedly has an effect on the kinetics of RNOS formation, and therefore caution is warranted when comparing these results.

Physiological effects of NO: anaerobic growth. Under anaerobic (fermentative) conditions, NO impaired growth in direct correlation to the amount of NO added (Fig. 1C). NO (50 µM) had little effect on anaerobic growth, whereas 100 and 200 µM NO significantly decreased both growth rate and final cell yield. Note that under the closed culture conditions of anaerobic growth, NO is unable to diffuse out of the system or to react with O2. While it is likely that NO inhibition of cytochromes accounts for some, if not all, of aerobic growth inhibition, during anaerobic fermentative growth respiratory chain cytochromes are not utilized. The absence of O2 further eliminates RNOS as a possible source of toxicity. It is therefore likely that direct effects of NO are responsible for anaerobic growth inhibition, perhaps via nitrosylation of thiol groups or Fe cofactors in essential enzymes.

Overview of the nitric oxide stimulons of B. subtilis. We employed DNA microarray analysis to investigate the transcriptional response of B. subtilis to NO under both aerobic and anaerobic conditions. Addition of 50 µM NO was chosen for the aerobic cultures, because this concentration leads to very little effect on cell viability (Fig. 1A) or growth rate (Fig. 1B), although it does lead to a transient inhibition of respiration (Fig. 2) and a transient growth lag (Fig. 1B). Similarly, 50 µM NO is the highest amount tested with the anaerobic cultures that did not significantly impair growth rate and final cell yield (Fig. 1C). Because stress responses are often transient in nature, we isolated RNA after 5 min for aerobic cultures and after 15 min for the more slowly growing anaerobic cultures. A second set of three independent microarray experiments was performed under aerobic conditions, with NO added as two 25 µM additions separated by 2 min (RNA was extracted 5 min after the initial addition) (Fig. 1S, supplemental data). The results from all aerobic microarray experiments were in general agreement with each other.

To identify genes that are strongly regulated in response to NO exposure, we generated scatter plots to compare the observed fluorescence intensity (after normalization) corresponding to each gene in the presence and absence of NO. The vast majority of genes cluster along a diagonal line with a slope of 1.0, which indicates that the levels of most mRNAs were unaffected by NO under these conditions (Fig. 3). However, in each case a significant subset of genes were clearly induced by NO. Aerobically, the most strongly induced genes were hmp and members of the {sigma}B and Fur regulons, with less induction of the PerR regulon (Fig. 3A). Under the conditions of the anaerobic experiment, the most strongly induced genes were hmp and members of the PerR and Fur regulons, whereas {sigma}B-regulated genes were induced weakly if at all (Fig. 3B). To compare the NO stimulons in a more direct manner, the effects of NO on gene expression (induced versus uninduced) were compared for the aerobic and anaerobic culture conditions (Fig. 4). This further emphasizes that induction of the Fur and PerR regulons is most dramatic in the anaerobic experiment, whereas induction of the {sigma}B regulon is most pronounced aerobically.



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  		FIG. 3. Gene expression of wild-type B. subtilis with 50 µM NO compared to the uninduced wild-type strain grown under aerobic growth conditions (A) or anaerobic growth conditions (B). Relative hybridization intensities (mean normalized hybridization quanta) are plotted for genes as measured for uninduced (x axis) and NO induced (y axis). Symbols correspond to known regulons induced by NO (black square, hmp; white square, Fur regulon; gray triangle, PerR regulon; cross, {sigma}B regulon). Under aerobic conditions hmp, the {sigma}B regulon, and the Fur regulon are induced by NO. Under anaerobic conditions hmp, the Fur regulon, and the PerR regulon are induced by NO.



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  		FIG. 4. The NO stimulon under aerobic and anaerobic growth conditions. The ratio of gene expression after NO addition (fold induction) is plotted for anaerobic (x axis) versus aerobic (y axis) growth. Genes induced equally by NO under anaerobic and aerobic conditions cluster in the upper-right quadrant along the diagonal dashed line (slope, 1.0). Genes more strongly induced under aerobic conditions cluster nearer to the y axis, while genes more strongly induced under anaerobic conditions cluster nearer to the x axis. Symbols: black square, hmp; white square, Fur regulon; gray triangle, PerR regulon; cross, {sigma}B regulon. WT, uninduced wild type.

We confirmed and extended these microarray results by slot blot hybridization of RNA from cells exposed to NO for 5 to 25 min (Fig. 5A to D). The results confirm that hmp and the PerR-regulated gene mrgA are rapidly induced by NO stress in anaerobic cultures, with somewhat slower induction noted for the Fur-regulated ykuO gene. As also noted in the microarray study, induction of the {sigma}B-regulated csbC gene is most notable under aerobic conditions.

Several other genes were induced by NO and fell outside the context of these regulons (Table 1). The regulation of most of these genes is unclear, but we do note the appearance of one member of the Spx regulon known to be induced by the thiol-specific oxidant diamide (37, 48). Because NO can modify reactive thiols, we tabulated the effects of NO on genes under the control of regulators that utilize reactive cysteine residues (Table 2). For example, at least three Spx-regulated genes (48) are significantly induced by NO under anaerobic conditions. OhrR is a regulator that responds to organic peroxide stress via oxidation of a cysteine residue and represses ohrA (16). The ohrA gene was induced by NO aerobically. Finally, the class III heat shock response, known to be induced by diamide (37), was induced by NO under aerobic conditions. These genes are regulated in part by McsA, a protein that is postulated to sense changes in redox and temperature via the oxidation of reactive cysteine residues (32, 37). Collectively, these results suggest that the reaction of NO (and RNOS under aerobic conditions) with reactive cysteine residues contributes to the observed NO stimulons.


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  		TABLE 2. Genes induced by regulatory proteins that utilize reactive cysteine residues to sense redox shiftsb

SNP induces many of the same genes as NO. SNP is a nitrosating agent and contains an NO+ (nitrosyl) group that can be donated to nucleophilic thiols and amines (53). NO can be released by SNP indirectly following nitrosation of a thiolate group and the subsequent degradation of S-nitrosothiol (53).

To determine whether genes induced by NO are also induced by SNP, we performed Northern hybridizations on RNA extracted from SNP-challenged cells. Under aerobic conditions, hmp, csbC ({sigma}B regulon), and ykuO (Fur regulon) were induced, while under anaerobic conditions hmp, two Fur-regulated genes (ykuO and dhbA), and two PerR-regulated genes (mrgA and katA) were induced (Fig. 6A to H). To document the selectivity of these responses, we monitored transcripts for mntH (MntR regulon [24]) and yciC (Zur regulon [19]). Neither of these genes was induced by SNP (Fig. 6I and J), demonstrating that the response of the Fur and PerR regulons is not shared by other metal-dependent repressor systems. In general, the transcriptional response to SNP is considerably slower than that observed for NO and required 20 to 60 min before expression was observed. Furthermore, millimolar concentrations of SNP (compared to 50 µM for NO) were required to elicit a measurable transcriptional response from B. subtilis. It is likely that SNP penetrates the cell poorly relative to NO. This may account for the high concentration requirement and slow response of B. subtilis to SNP. Despite these notable differences in the dose and kinetics of the response, the overall pattern of gene induction by SNP was in general agreement with that observed for NO under aerobic and anaerobic conditions.



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  		FIG. 6. SNP induces many of the same genes as NO. Northern hybridization of total RNA extracted from uninduced wild type (–) or wild type induced by 2 mM SNP (+). As controls for full derepression, RNA was also isolated from untreated culture of the fur mutant, perR mutant, mntR mutant, or zur mutant. Probes employed under aerobic conditions included (corresponding regulon is in parentheses): hmp (ResDE) (A); cscC ({sigma}B) (B); and dhbA (Fur) (C). Probes employed under anaerobic conditions included dhbA (Fur) (D), ykuO (Fur) (E), katA (PerR) (F), mrgA (PerR) (G), hmp (ResDE) (H), mntH (MntR) (I), and yciC (Zur) (J). wt, wild type.

Induction of hmp by NO. The gene most highly induced by NO (gas) under both aerobic and anaerobic conditions was hmp, although its expression was almost 10-fold higher under anaerobic versus aerobic conditions (Fig. 3, 4, 5A). NO-mediated induction of hmp is expected given its documented cytoprotective role against NO challenge in E. coli and S. enterica (see Introduction). Induction of hmp requires the two-component system ResDE (46). Although ResDE was initially believed to sense anaerobiosis (35), subsequent studies have revealed that NO is also required (46). In addition to hmp, the ResDE two-component system regulates several other genes, one of which is nasD, an assimilatory nitrite reductase subunit. Under aerobic conditions, NO induced nasD 13.7-fold while under anaerobic conditions it was induced 8.8-fold (Table 1). It is noteworthy that hmp and nasD are among the ResDE-regulated genes that are most highly induced under nitrate and nitrite respiration conditions (46).

Induction of the Fur regulon by NO. The B. subtilis ferric uptake regulator (Fur) regulates iron homeostasis by repressing genes involved in siderophore biosynthesis and uptake under conditions of iron sufficiency (4, 8, 9, 28). The Fur regulon is induced by NO under anaerobic conditions and, to a lesser degree, under aerobic conditions (Fig. 3, 4, 5B). Under aerobic conditions, ykuO achieves a maximum induction of 7-fold at 15 min, while anaerobically ykuO is induced 30-fold after 25 min (Fig. 5B).

Fur is a dimeric DNA-binding protein that requires bound Fe(II) to bind DNA (8). Insight into the possible mechanism of NO induction of the Fur regulon has been provided by D'Autreaux et al. (14), who demonstrated that direct nitrosylation of the Fe corepressor of E. coli Fur was responsible for NO-mediated inhibition of Fur DNA binding. It is likely that the same mechanism accounts for NO-mediated Fur regulon induction in B. subtilis.

Effect of NO and SNP on fur mutants. The observation that NO and SNP induce the Fur and PerR regulons suggests that these regulons may provide a defense against NO stress. Indeed, the PerR regulon provides defense against another redox-active agent, peroxide, and includes AhpC, an enzyme with demonstrated RNOS cytoprotective properties (7, 12, 40). Under our experimental conditions, SNP induced genes under the control of both of these metalloregulators, albeit at levels significantly lower than those observed in null mutant strains (Fig. 6C to G).

To determine whether derepression of the Fur or PerR regulons results in resistance to NO and SNP, we monitored growth of fur and perR null mutants in the presence of NO and SNP under aerobic and anaerobic conditions. The perR mutation did not significantly affect growth in the presence of either NO (gas) or SNP (data not shown). Surprisingly, the fur mutant was impaired in growth in the presence of either NO or SNP (Fig. 7). Because a fur mutant contains elevated levels of intracellular iron (23), it is possible that the Fe-nitrosyl complexes formed in the presence of NO and SNP are themselves toxic to the cell. If correct, induction of the Fur regulon by NO and SNP could contribute to the impaired growth noted in the presence of these agents. Alternatively, Fur itself may act as a sink (in its metallated form) for NO. Indeed, it has been speculated that Fur may be sufficiently abundant in E. coli to act as a Fe storage protein in addition to its well-defined role as a metalloregulator (69).

The response of PerR to NO is suppressed by Mn. PerR is a Fur homolog that negatively regulates the major vegetative catalase (katA), a Dps homolog (mrgA), an operon involved with heme biosynthesis (hemAX-CDBL), alkyl hydroperoxide reductase (ahpCF), a Zn(II) uptake system (zosA), fur, and perR (17, 18, 28). Some, but not all, PerR-regulated genes (katA, mrgA, ahpCF, zosA) are induced by the addition of peroxide (27) or diamide (37). In general, the subset of PerR-regulated genes that are strongly induced by oxidative stress are those that are induced by NO.

Induction patterns for mrgA were similar to those observed for the Fur-regulated gene ykuO. Under aerobic conditions, mrgA expression peaked at a value of 10-fold 15 min after NO addition, while under anaerobic conditions 40-fold induction was observed 25 min after NO addition (Fig. 5C). Indeed, only the anaerobic induction of mrgA, katA, and zosA by NO exceeded fivefold as observed by microarray analysis, while other PerR regulon members were relatively unaffected (Table 3).


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  		TABLE 3. PerR-regulated gene induction by NO under either aerobic or anaerobic growth conditions

In contrast to Fur, PerR can use either Fe(II) or Mn(II) in its sensing site (17). Furthermore, the selectivity of metal repression varies among PerR members, with some genes repressed by either Mn(II) or Fe(II) while others are repressed only by Mn(II) (17). We hypothesized that NO inactivates PerR by nitrosylation of the bound Fe(II) corepressor, as previously documented for E. coli Fur both in vitro and in vivo (14).

To test this model in vivo we examined the induction of two PerR-regulated genes (katA and mrgA) by NO or SNP in defined anaerobic medium with 10 µM Fe(II) or 10 µM Mn(II) to favor either the PerR:Fe or PerR:Mn form (9, 17). Significantly, katA and mrgA were induced by NO and SNP in the medium favoring the PerR:Fe form (Fig. 8C to F) but not when PerR:Mn was the dominant form. Furthermore, when Fe(II) and Mn(II) were both present, katA and mrgA were induced at levels similar to those observed with Fe(II) alone. This corroborates results obtained using H2O2 as an inducer, which suggested that PerR assumes the Fe(II) form when both metals are present (17). As a control, we demonstrated that under identical culture conditions hmp was induced by NO and SNP regardless of the Fe(II) and Mn(II) levels (Fig. 8A and B). These results provide strong evidence to support the idea that PerR exists in vivo in either a PerR:Fe or a PerR:Mn form and indicates that these two forms differ greatly in their ability to respond to NO. This extends a previously presented model proposing that the PerR:Fe form is primarily responsible for sensing H2O2 (44).

The {sigma}B regulon is activated by NO and SNP via different pathways. The {sigma}B regulon includes an estimated 200 genes that are induced by entry into early stationary phase (energy stress) or by a variety of environmental stresses (reviewed in references 26 and 54). Although only a subset of the {sigma}B regulon was induced by NO under aerobic conditions, partial induction of this regulon is commonly observed for other {sigma}B regulon-activating conditions (26, 52, 54). Although it is unclear which of these genes (if any) provide cytoprotective benefits to the cell against the effects of NO, the {sigma}B general stress response includes dps, a ferritin-like protein with a demonstrated role in defending against oxidative stress (39), three catalase homologues (katB, katX and ydbD), and other factors known to protect against redox-active agents (54).

The activity of {sigma}B is directly regulated by the anti-{sigma} RsbW. In response to stress, the anti-{sigma} antagonist RsbV interacts with RsbW, thereby releasing {sigma}B. RsbV, in turn, is regulated by reversible phosphorylation. Dephosphorylation of RsbV requires either of two PP2C phosphatases (RsbU and RsbP) and results in activation of {sigma}B (54). It has been shown that the phosphatase activity of RsbU is responsive to environmental stress while the phosphatase activity of RsbP is induced by cellular energy stress.

To determine whether NO and SNP activation occurs by the environmental (RsbU) or the energy stress (RsbP) pathway, the response of a ctc-lacZ reporter was analyzed in rsbP and rsbU mutant backgrounds (Fig. 9). The ctc gene encodes a {sigma}B-induced ribosomal protein (55) and has been used extensively to monitor the {sigma}B general stress response. Under aerobic conditions the NO-dependent induction of ctc-lacZ expression was greatly reduced in the rsbP mutant, suggesting that activation is primarily dependent on the energy stress pathway. Under anaerobic conditions NO did not induce ctc expression. Because NO inhibits aerobic respiration under similar conditions (Fig. 2), we suggest that depressed ATP levels may trigger the {sigma}B response via the energy stress branch (10, 29).



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  		FIG. 9. The {sigma}B regulon is activated by NO and SNP. All B. subtilis strains contain a ctc-lacZ fusion at the amyE locus. Shown are B. subtilis PB198 (wild type), PB567 rsbP::spc (rsbP), PB495 rsbU::{Delta}2 (rsbU), HB2617 rsbP::spc, and rsbU::{Delta}2 (rsbP rsbU). Error bars represent standard errors.

Unexpectedly, SNP activated {sigma}B induction through the environmental rather than the energy stress pathway and induction was noted both aerobically and, to a lesser extent, anaerobically (Fig. 9). SNP induction of ctc was essentially eliminated in an rsbU mutant. Thus, NO and SNP activate the {sigma}B general stress response through different mechanisms. It is unclear where SNP acts within the complex environmental signal transduction cascade. Several upstream regulators of the RsbU phosphatase have been identified that may monitor the various environmental stresses that induce the {sigma}B general stress response (3, 11). One of these regulators, YtvA, possesses a PAS sensory domain (3) thought to respond to redox-reactive signals, such as O2 tension and light (58), and this regulator could also be involved in sensing SNP-induced stress. Similarly, energy sensing by RsbP may also involve a PAS domain (60). These results highlight the fact that NO and SNP affect cellular physiology in different manners.

The much stronger activation of the {sigma}B response in aerobic conditions suggests that the active agent might be RNOS derived from the reaction of NO with O2 and superoxide. Alternatively, NO may preferentially activate {sigma}B when the cell is expressing genes required for aerobic metabolism. In an attempt to address this question, NO-saturated water and fermentation broth were allowed to react with O2 for 5 and 15 min and were added to anaerobically grown cells in concentrations up to 200 µM (initial NO concentration before aeration). This did not elicit a {sigma}B response as judged by RNA slot blot analysis (data not shown). While this suggests that RNOS are not sufficient to cause a {sigma}B response in anaerobically grown cells, the transient nature of RNOS and the fact that they were generated in a cell-free system complicates the interpretation.

Concluding remarks. We have explored the physiological and genetic consequences of NO stress by using B. subtilis as a model system. We have demonstrated that aerobic cells are most sensitive to NO when added repeatedly over several minutes rather than as a single bolus. This may indicate that NO-treated cells, which experience a transient inhibition of respiration, become sensitized to the killing action of subsequently added NO. Under conditions that lead to minimal loss of viability, we demonstrated that NO (gas) strongly induces hmp and the PerR, Fur, and {sigma}B regulons. The precise set of genes induced is dependent on culture conditions, with induction of the PerR and Fur regulons being most pronounced anaerobically while the {sigma}B regulon is most strongly induced aerobically. Induction of the PerR regulon can be blocked by manganous ion, supporting a model in which induction of the Fur and PerR regulons results from Fe(II) nitrosylation. In general, the nitrosating agent SNP induces many of these same regulons, but the pathways of signal transduction responsible for NO and SNP induction of the {sigma}B regulon are distinct.


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ACKNOWLEDGMENTS
 
We thank Chester Price for generously providing us with ctc-lacZ fusion strains used for {sigma}B analysis. We further thank Jim P. Shapleigh and Peter S. Choi for their invaluable advice and discussion.

This work was supported by grants from the National Science Foundation to J.H. (MCB0235255) and to M.N. (MCB0110513).


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Microbiology, Wing Hall, Cornell University, Ithaca, NY 14853. Phone: (607) 255-6570. Fax: (607) 255-3904. E-mail: jdh9{at}cornell.edu. Back

{dagger} Supplemental material for this article may be found at http://jb.asm.org/. Back


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Journal of Bacteriology, July 2004, p. 4655-4664, Vol. 186, No. 14
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.14.4655-4664.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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