The nitrosative stress response of Staphylococcus aureus is required for resistance to innate immunity

Authors


*E-mail fcfang@u.washington.edu; Tel. (+1) 206 221 6770; Fax (+1) 206 616 1575.

Summary

Staphylococcus aureus is a highly virulent human pathogen with an extensive array of strategies to subvert the innate immune response. An important aspect of innate immunity is the production of the nitrogen monoxide radical (Nitric Oxide, NO·). Here we describe an adaptive response to nitrosative stress that allows S. aureus to replicate at high concentrations of NO·. Microarray analysis revealed 84 staphylococcal genes with significantly altered expression following NO· exposure. Of these, 30 are involved with iron-homeostasis, potentially under the control of the Fur regulator. Another seven induced genes are involved in hypoxic/fermentative metabolism, including the flavohaemoprotein, Hmp. The SrrAB two-component system has been shown to regulate the expression of many of the NO·-induced metabolic genes. Indeed, inactivation of hmp, srrAB and fur resulted in heightened NO· sensitivity. Hmp was responsible for c. 90% of measurable staphylococcal NO· consumption and therefore critical for efficient NO· detoxification. While SrrAB was required for maximal hmp expression, srrAB mutants still exhibited significant NO· scavenging and NO·-dependent induction of hmp. Yet S. aureus lacking SrrAB were more sensitive to nitrosative stress than hmp mutants, indicating that the contribution of SrrAB to NO· resistance extends beyond the regulation of hmp expression. Both Hmp and SrrAB were required for full virulence in a murine sepsis model, however, only the attenuation of the hmp mutant was restored by the abrogation of host NO· production. Thus, the S. aureus Hmp protein has evolved to serve as an iNOS-dependent virulence determinant.

Introduction

Infections caused by the Gram-positive bacterium Staphylococcus aureus are notoriously difficult to prevent and treat. S. aureus has been shown to interfere with virtually every facet of the host innate immune response, ranging from leucocyte chemotaxis to phagocytosis and intracellular killing (Foster, 2005). Factors such as CHIPS (CHemotaxis Inhibitory Protein of S. aureus) and Eap (Extracellular Adherence Protein) can interfere with the influx of neutrophils, the host's primary line of defence against invading S. aureus (Chavakis et al., 2002; Haas et al., 2004). Furthermore, the staphylococcal capsule, protein A, fibrinogen binding proteins and staphylokinase confer resistance to opsonin-mediated phagocytosis (Wilkinson et al., 1979; Palmqvist et al., 2004; Rooijakkers et al., 2005). When phagocytes are able to internalize staphylococci, engulfed cells resist killing by host-derived reactive oxygen species through the elaboration of antioxidant defences such as carotenoid pigments, two superoxide dismutases, manganese homeostasis and catalase (Mandell, 1975; Horsburgh et al., 2002; Karavolos et al., 2003; Liu et al., 2005). Thus, the subversion of innate immunity is a major reason for the remarkable success of S. aureus as a pathogen.

Nitric Oxide (NO·) is an important effector of host innate immunity attributed both to its antimicrobial activity and its immunomodulatory roles. Inflammatory NO· has been shown to be indispensable for normal clearance of diverse pathogens including viral, fungal, bacterial and parasitic microorganisms (De Groote, 1999). Activated leucocytes can generate local concentrations of NO· in the micromolar range through the action of an inducible Nitric Oxide Synthase (iNOS)(Lewis et al., 1995; Nalwaya and Deen, 2005). NO· can react directly with invading organisms and surrounding host tissues, or can be further oxidized into potent reactive nitrogen species (RNS). Moderate levels of RNS are responsible for the nitrosation of protein thiols (RSNO) and the nitrosylation of metal centres, particularly iron (Wink and Mitchell, 1998). At high concentrations, RNS can mediate nitration of protein tyrosine residues (Y-NO2), peroxidation of lipids and deamination of DNA bases (Radi et al., 1991; Wink et al., 1991; Schopfer et al., 2003).

Accordingly, bacterial cells exposed to NO· must respond to a diverse array of biochemical insults. Thisnitrosative stress response often includes the maintenance and replenishment of cytosolic thiol pools, altered metal homeostasis, activation of specific DNA repair processes, and increased enzymatic NO· detoxification (Ohno et al., 2003; Firoved et al., 2004; Moore et al., 2004; Mukhopadhyay et al., 2004; Flatley et al., 2005; Hromatka et al., 2005; Justino et al., 2005). For instance, the depletion of thiol pools by RNS induces the cysteine and homocysteine biosynthetic operons in Escherichia coli and Salmonella Typhimurium, and glutathione biosynthetic genes in Candida albicans (De Groote et al., 1996; Mukhopadhyay et al., 2004; Hromatka et al., 2005). Induction of iron-regulated genes by NO· has been demonstrated in E. coli and attributed to nitrosylation of the ferrous iron/Fur regulatory complex (D'Autreaux et al., 2002; Mukhopadhyay et al., 2004). Due to the reactivity of RNS for DNA, the SOS response is induced under nitrosative stress in E. coli and S. Typhimurium (Schapiro et al., 2003; Mukhopadhyay et al., 2004). Finally, NO· exposure often leads to the increased expression of enzymes evolved explicitly to detoxify this radical. Bacterial flavohaemoglobins, the NO· reductases of denitrifying bacteria, cytochrome c nitrite reductases, and flavorubredoxins are potent scavengers of both endogenously generated and exogenously supplied NO· (Braun and Zumft, 1991; Gardner et al., 1998; 2002; Watmough et al., 1999; Poock et al., 2002). Homologues of a flavohaemoglobin, Hmp, are known to be protective in cells exposed to NO·in vitro for a wide variety of microorganisms including S. aureus (Ullmann et al., 2004; Poole, 2005; Goncalves et al., 2006). Indeed, the flavohaemoglobin of S. Typhimurium is critical for intraphagosomal survival in cultured human macrophages (Stevanin et al., 2002). The regulation of these proteins is not generally understood, but the signals leading to full induction often include microaerobiosis and the presence of NO·, or nitrite/nitrate (LaCelle et al., 1996; Hu et al., 1999; Cruz-Ramos et al., 2002). These NO·-detoxifying enzymes may have evolved to protect cells from endogenously produced RNS during anaerobic respiration using nitrate/nitrite as terminal electron acceptors.

The ability to produce NO· via iNOS has been shown to protect murine hosts from infections caused by S. aureus (Sakiniene et al., 1997; McInnes et al., 1998; Sasaki et al., 1998). Indeed, iNOS-dependent NO· production can be detected in experimentally infected wounds or in cultured cells exposed to staphylococcal lipoteichoic acid (Mahoney et al., 2002; Chang et al., 2005). Furthermore, NO· was demonstrated to increase killing of ingested S. aureus by PMN-like cytokineplasts (Malawista et al., 1992). Recent advances in NO· releasing sol-gel materials have shown promise in controlling staphylococcal colonization of implanted medical devices (Nablo et al., 2005), and preliminary studies have suggested that NO· exposure can reduce staphylococcal viability in vitro (Kaplan et al., 1996). Thus, while host-derived NO· is a valuable weapon against invading staphylococci, the mechanisms by which NO· exerts anti-staphylococcal activity have yet to be systematically examined. Nor has any investigations into the adaptive response of S. aureus to nitrosative stress been undertaken. This study represents an initial examination of the role of the nitrosative stress response in staphylococcal pathogenesis.

Results

Staphylococcus aureus is relatively resistant to growth inhibition by NO·

Published studies using the ‘NO· donor’S-nitroso N-acetyl dl-penicillamine (SNAP) have suggested that NO· is bactericidal for S. aureus (Kaplan et al., 1996). However, in these experiments, the loss of cell viability did not occur rapidly, and bacterial replication was hindered by the lack of essential nutrients in KRPG (Krebs–Ringer Phosphate buffer supplemented with Glucose) (see Experimental procedures). While these findings were faithfully replicated in our laboratory, conducting experiments in PN growth medium (see Experimental procedures) yielded strikingly different results (Fig. 1A). While the addition of 1 mM SNAP had minimal effect on the growth of S. aureus strains COL and Newman (not shown) in PN medium, this concentration was lethal in KRPG buffer. S-nitrosothiols such as SNAP release finite doses of NO· that are only transiently sustainable (NO· can undergo auto-oxidation with dissolved oxygen, react with medium components, or dissipate out of the reaction vessel). Thus, the inability of SNAP to inhibit staphylococcal growth in PN medium as compared with KRPG buffer might be attributed to differences in basal NO· scavenging of the solutions. However, the loss of authentic NO· in 8 ml of either solution occurred at comparable rates (initial rate of 1.2 × 10−9 mol min−1 for KRPG and 1.4 × 10−9 mol min−1 for PN medium); therefore, differential NO· scavenging cannot account for staphylococcal resistance to SNAP in PN medium.

Figure 1.

S. aureus is resistant to high concentrations of NO· in growth medium.
A. Viable cfu ml−1 of cultures supplemented with either 1 mM of the NO· donor compound SNAP or 1 mM of the parent compound NAP in PN medium versus KRPG buffer.
B. Growth rates (doublings h−1) of various strains of S. aureus, B. subtilis and E. coli exposed to c. 2 mM authentic NO· during mid-logarithmic growth at an approximate cell density of 108 cfu ml−1 (see Experimental procedures).

While SNAP can release authentic NO· in the presence of light or metal ions, the rate of NO· release is relatively slow, and trans-nitrosation reactions (donation of NO+) are generally favoured. Therefore, exposure of mid-logarithmic S. aureus cultures to authentic NO· via the addition of diazeniumdiolate donors (NOC-12 and DEA/NO at a 9:1 molar ratio) was examined (see Experimental procedures). However, as with SNAP, authentic NO· failed to significantly inhibit staphylococcal growth. In addition, S. aureus strains COL and Newman grown to mid logarithmic phase displayed only a transient growth arrest after exposure to millimolar concentrations of NO· (Fig. 1B, Fig. S3). The growth rate of NO·-exposed S. aureus quickly returned to approximately 50% of that of unexposed cells within minutes, whereas E. coli (MG1655) and Bacillus subtilis (168) resumed growth only after a 4 h period of complete growth inhibition (Fig. 1B). Thus, in contrast to other bacteria, S. aureus appears to escape the cytostatic actions of NO· when provided appropriate nutrients.

One explanation for the ability of S. aureus to divide despite the presence of relatively high levels of NO might be the efficiency of staphylococcal NO· scavenging. Indeed, cell suspensions of S. aureus COL at c. 1 × 108 cfu ml−1 exhibit a marked increase in NO· consumption compared with that of medium alone (Fig. 2A). However, this NO· scavenging rate is not kinetically different from that of E. coli at comparable cell densities (Fig. 2A). While in the absence of induction, B. subtilis (168) did not display significant NO· scavenging (Fig. 2A), pre-exposure to 1 mM SNAP induced high rates of cellular NO· scavenging in B. subtilis, E. coli and S. aureus (Table 1). Therefore, superior NO·-scavenging cannot account for the relative resistance of S. aureus to NO· as compared with other organisms.

Figure 2.

NO· metabolism of bacterial cell suspensions.
A. Cell suspensions (108 cfu ml−1) in PBS of mid-logarithmic cultures of E. coli (MG1655), B. subtilis (168) and S. aureus (COL) were assayed for NO· consumption at 37°C. Measurements were obtained using an ISO-MARK NO· probe after addition of 1 μM ProliNO. Results are expressed as fraction of the initial 2 μM NO· remaining over time.
B. Same as in (A), but a comparison of WT S. aureus COL with isogenic hmp and srrAB mutants.

Table 1.  Initial NO·-consumption rates of bacterial cell suspensions.
 UninducedaInduceda
  • a. 

    Uninduced cells were exposed to 1 mM NAP for 25 min; induced cells were exposed to 1 mM SNAP for 25 min prior to measurement.

  • Rates are expressed as nmol min−1·108 cfu−1 or nmol min−1 for PN Medium.

PN Medium1.6N/A
B. subtilis 1682.2128
E. coli MG165519.4148
S. aureus COL16.247.5
COL hmp2.75.3
COL srrAB7.317.2

Transcriptional microarray analysis of NO·-exposed S. aureus COL cells

Global levels of transcript abundance from mid-logarithmic cells grown in PN medium exposed to either 1 mM SNAP or the native thiol NAP (N-acetyl-d,l-penicillamine) for 25 min were compared by Affymetrix microarray analysis. A total of 84 genes were differentially expressed during nitrosative stress from SNAP exposure compared with NAP alone (Fig. 3A, Table S1). Of these, 57 (68%) genes were proposed to be within 16 operons (Wang et al., 2004). Five operons containing 14 open reading frames (ORFs) were repressed by NO·, along with an additional four monocistronic ORFs, comprising 18 NO·-repressed genes in total. Eleven operons encompassing 43 genes were induced by SNAP exposure, with another 23 monocistronic genes also displaying significant NO· induction (66 NO·-induced genes in total).

Figure 3.

Microarray analysis of the nitrosative stress response in S. aureus.
A. Y-axis: Z-values (see Experimental procedures) for each of the 4984 probe sets with significant signal; x-axis: probe sets ordered as on the Affymetrix GeneChip. Dotted lines demark the Z = ±3 thresholds. Red symbols indicate genes with putative iron homeostasis function, green symbols represent genes involved with hypoxic/fermentative metabolism, and blue symbols represent PerR repressed genes. Numbers correspond to 13 genes selected for further verification by Q-RT PCR.
B. Q-RT PCR results of RNA from SNAP versus NAP-exposed cells for 13 genes. Signals were normalized to rpoD levels.
C. Q-RT PCR analysis of hmp expression in NAP-exposed (open bars) or SNAP-exposed (grey bars) cells in both WT and srrAB backgrounds. Signals were normalized to rpoD levels.

Of the 66 genes induced by NO·, 30 (45%) are most likely involved in iron homeostasis (Fig. 3A, red symbols), many of which are repressed by the Ferric Uptake Regulator (Fur). Of the 14 repressed genes, five (ahpCF, katA, ftnA and mrgA) are known to be repressed by PerR, a peroxide responsive Fur homologue (Fig. 3A, blue symbols) (Horsburgh et al., 2001; Morrissey et al., 2004). However, homologues of some of these genes (acnA, ftnA and sdhB) are known to be Fur-activated in other organisms (Masse and Gottesman, 2002), so the exact mechanism by which NO· represses the expression of some of these genes remains to be elucidated.

Another prominent feature of the staphylococcal nitrosative stress response was the subset of genes involved in hypoxic/fermentative metabolism that displayed altered expression (Fig. 4A, green symbols). NO· induced ldh, hmp, fdaB, nrdDG and cydAB transcription and repressed the expression of the pyrimidine biosynthetic operon, pyrBCcarABpyrF. NO· stress also triggered the expression of members of the Clp ATPase family proteins involved in stress-induced protein turnover. SNAP exposure leads to increased expression of clpB and clpL, both of which have been shown to be induced in heat shocked cells (Frees et al., 2004). Furthermore, the ctsR gene, a regulator of clpB, was also induced in NO·-exposed cells. While CtsR represses the expression of clpB, both are known to accumulate in heat stressed cells (Frees et al., 2004).

Figure 4.

Relative resistance of S. aureus COL and mutant derivatives to NO· in TSB. Cells were inoculated at a density of c. 106 cfu ml−1 and exposed to NO· by addition of 5 mM NOC-12.

Microarray results were validated by quantitative real time reverse transcriptase-polymerase chain reaction (Q-RT PCR) analysis of SNAP-induced transcript levels. Thirteen of the SNAP-induced genes showed > twofold increases in expression by Q-RT PCR (Fig. 3B, Table S1). Five negative control genes (SA2618, SA0119, gyrA, rpsL, rpoD) that were not determined to be differentially regulated by microarray analysis did not demonstrate differential transcript titers.

SrrAB and Fur regulons are essential for NO· resistance

Given the extent of the staphylococcal NO· response that can be attributed to hypoxic metabolism and iron homeostasis, we determined whether mutations in srrAB (Staphylococcal Respiratory Regulator) or fur (Ferric iron Uptake Regulator) affect NO· resistance. SrrAB is known to be important for the expression of many genes involved in anaerobic metabolism and Fur represses many of the iron acquisition genes in S. aureus (Horsburgh et al., 2001; Throup et al., 2001). Inactivation of srrAB or fur significantly enhanced staphylococcal NO· susceptibility (Fig. 4).

ResDE, a two-component regulatory system homologous to SrrAB activates expression of the flavohaemoglobin Hmp in B. subtilis (Nakano, 2002). In enteric bacteria, Hmp is known to reduce NO· to the nitroxyl ion (NO) which can then yield nitrate (NO3) in the presence of molecular oxygen or nitrous oxide (N2O) under anaerobiosis (Hausladen et al., 2001). As the staphylococcal hmp gene was induced by NO· in the microarray analyses, we investigated whether the heightened NO· sensitivity of an srrAB mutant results from an inability to activate hmp, a critical NO· detoxification system in S. aureus (Goncalves et al., 2006). While inactivation of hmp did result in increased NO· sensitivity, the NO·-sensitive phenotype was not as severe as in the srrAB mutant (Fig. 4, Fig. S4). Moreover, srrAB mutant bacteria displayed significantly more NO· scavenging capability than hmp mutant cells (Fig. 2B). Analysis of hmp transcription in both wild-type (WT) and srrAB cells demonstrated that SrrAB is only required for maximal expression, and that NO· still induced hmp expression sixfold in srrAB staphylococci (Fig. 3C). Therefore, the critical contribution of SrrAB to NO·-resistance in S. aureus is only partially attributed to the regulation of Hmp and NO· scavenging.

Inactivation of fur significantly reduced the growth rate of S. aureus in PN defined medium (data not shown). However, growth in complex medium such as trypticase soy broth (TSB) alleviated most of the fur-associated growth defect. Inactivation of fur resulted in a more marked increase in NO· sensitivity in TSB than could be attributed simply to a reduced growth rate (Fig. 4).

Hmp expression is required for full virulence in NO·-proficient mice

Intravenous inoculation of C57BL/6 mice with 5 × 107 cfu of S. aureus strain Newman resulted in the onset of arthritic symptoms within 3 days followed by increasing mortality until day 10 (Fig. 5A). However, mice infected with isogenic hmp and srrAB mutant bacteria showed little or no indications of septic arthritis, and significantly less mortality than mice infected with WT bacteria (Fig. 5A). Additionally, kidney tissue sampled from mice infected with 2 × 107 cfu of hmp or srrAB Newman contained 8- and 58-fold fewer viable cfu per gram tissue, respectively, than WT Newman at day 5 (data not shown). C57BL/6 iNOS–/– mice lack the ability to generate high levels of NO· in response to infection due to inactivation of the NOS2 gene. These mice are more susceptible to infection with S. aureus as seen by the increased mortality between days 5 and 8 (Fig. 5B). Additionally, iNOS–/– mice infected with hmp mutant bacteria succumb to infection with similar kinetics as iNOS–/– mice inoculated with WT Newman (Fig. 5B). Furthermore, the numbers of viable cfu per gram kidney tissue from iNOS–/– mice infected with WT versus hmp Newman were indistinguishable (3.2 × 108 versus 2.6 × 108 respectively). Thus the role of hmp as a virulence determinant is solely dependent on the ability of the host to produce NO· via iNOS. While avirulent srrAB mutant S. aureus displayed in vitro susceptibility to NO· (Fig. 4), virulence with this mutant could not be restored in the iNOS–/– mouse indicating additional roles for members of this regulon in vivo (Fig. 5B).

Figure 5.

Kaplan–Meier survival plots of mice infected with S. aureus Newman and mutant derivatives. WT C57BL/6 mice (A) or iNOS–/– C57BL/6 mice (B) were infected i.v. with 5 × 107 cfu of WT S. aureus Newman (circles), isogenic hmp mutant (squares), or srrAB mutant cells (triangles).

Discussion

In order for S. aureus to survive within the host during infection, the bacterium must initially subvert a formidable innate immune defence. This report describes an immune evasion strategy employed by S. aureus to avoid clearance by host-derived NO·. Several studies report the importance of NO· in protecting mammalian hosts from a variety of bacterial pathogens, in that the lack of high-level NO· production severely impairs the host's ability to control infections (De Groote, 1999). Indeed, mice lacking iNOS show a modest defect in resistance to S. aureus following intravenous inoculation (Fig. 5). Thus it seems that NO· plays only a minor protective role in murine hosts against this pathogen. However, further inspection reveals that S. aureus mutants lacking components of the nitrosative stress response (i.e. hmp) are avirulent in WT mice, but readily causes lethal infections in iNOS–/– mice. This suggests that invading S. aureus are exposed to potentially growth-limiting concentrations of NO·, but the production of Hmp as an adaptive response to nitrosative stress effectively neutralizes host-derived NO·.

While the role of staphylococcal Hmp in NO·-resistance has been clearly demonstrated, the mechanism of NO·-dependent hmp induction in S. aureus is not yet clear. Previous work has shown that microaerobiosis and nitrosative stress appear to induce hmp expression in S. aureus (Goncalves et al., 2006). The regulation of hmp expression is known to involve the ResDE two-component system in B. subtilis (Nakano, 2002). However, the staphylococcal ResDE counterpart, SrrAB, is not solely responsible for NO·-dependent activation of hmp (Fig. 3C). In E. coli, the FNR protein has been implicated in hmp regulation; however, S. aureus lacks any recognizable FNR homologue (Poole et al., 1996). Another iron-sulphur containing protein, NsrR, has been predicted to control hmp expression in many species of proteobacteria including E. coli, and Gram-positive species such as B. subtilis (Rodionov et al., 2005). Indeed, this gene has been demonstrated to control the NO·-dependent induction of Hmp in E. coli (Bodenmiller and Spiro, 2006). Work is ongoing to determine whether the staphylococcal homologue of NsrR (SA-COL1681) controls hmp expression in S. aureus.

Direct NO· detoxification is, however, only part of the Staphylococcal RNS response. E. coli and B. subtilis both contain Hmp homologues and efficiently scavenge NO· (Table 1), yet these species are unable to replicate in the presence of high concentrations of NO· (Fig. 1B). In contrast, S. aureus resumes dividing at approximately 50% of its normal growth rate just minutes after the initiation of NO· exposure (Fig. 1B). This suggests that S. aureus possesses other mechanisms to escape NO·-mediated cytostasis. The cDNA microarray analysis of NO·-exposed S. aureus was specifically conducted to identify genes with altered expression during this adaptation period. The staphylococcal Fur regulon appears to be globally derepressed during nitrosative stress, as has been previously demonstrated in E. coli and B. subtilis (Moore et al., 2004; Mukhopadhyay et al., 2004). It is known that NO· can readily react with the Fur-bound ferrous ion creating dinitrosyl iron complexes (DNICs) and rendering the Fur-Fe2+ dimer inactive, thus derepressing the regulon (D'Autreaux et al., 2002). Interestingly, inactivation of fur in S. aureus resulted in cells extremely sensitive to NO· (Fig. 4). This was also shown in E. coli (Mukhopadhyay et al., 2004) and suggests that inactivation of Fur by NO· may represent a pathological consequence of nitrosative stress rather than an adaptive response. Alternatively, Fur is a highly abundant protein (Watnick et al., 1997; Zheng et al., 1999) and may possess NO·-resistance functions outside the role of an iron-responsive transcriptional regulator.

While deprepression of the staphylococcal Fur regulon by NO· is consistent with the NO·-responses of several species, many key aspects of the staphylococcal nitrosative stress response differ significantly from that of its Gram-positive counterpart B. subtilis. The most striking is that NO· induced the expression of several SigB controlled genes in B. subtilis (Moore et al., 2004), while the clpL gene represents the sole member of the staphylococcal SigB regulon to be induced by NO· exposure (Fig. 3B). Thus, in contrast to B. subtilis, the extensive SigB regulon was virtually unaffected by NO· in S. aureus. Additionally, the staphylococcal PerR regulon was further repressed by NO· exposure (Fig. 3A), a trend not apparent in NO·-exposed B. subtilis. The biological basis for the divergence in NO·-responses between these two species is unknown, but it may reflect the significant difference in NO· sensitivity between S. aureus and B. subtilis (Fig. 1B).

Several hypoxic/fermentative metabolic genes including hmp, the anaerobic nucleotide reductase genes, nrdGD, the alternative bd-type cytochrome cydAB, a class one 1,6-fructosebisphosphate aldolase fdaB, and one of the staphylococcal l-lactate dehydrogenases ldh were significantly induced in NO·-treated cells. Similarly, the transcriptional response of Mycobacterium tuberculosis to NO·-exposure overlapped significantly with that of cells grown under hypoxic conditions (Ohno et al., 2003). Interestingly some NO·-stimulated hypoxic/fermentative genes (cydAB, nrdDG and ldh) were also induced in S. aureus upon H2O2 exposure as well as in mutant staphylococci devoid of haem biosynthesis (hemB mutants: ldh, fdaB, hmp and clpL) (Kohler et al., 2003; Chang et al., 2006). Staphylococci grown under nitrosative stress, oxidative stress or with impaired haem biosynthesis would share a common defect in basic respiratory function and, in turn, induce the expression of hypoxic/fermentative genes. The mechanism by which this happens is ill-defined, but the two-component system SrrAB most likely plays a key role in the regulation of these genes. S. aureus lacking srrAB grow very poorly under anaerobiosis (Throup et al., 2001). Furthermore, proteomic analyses indicate that this regulon affects the expression of adh, ldh, arcAB, scdA, acnA, fumA and others (Throup et al., 2001). Many of these genes showed altered expression in cells exposed to NO· (Table S1).

Insight into the true complement of SrrAB regulated genes may shed light on the NO· sensitivity as well as the avirulence of staphylococcal srrAB mutants. Indeed, in addition to NO· sensitivity, srrAB cells may be attenuated simply because hypoxic environments are often encountered in the host and thus the ability to grow in these niches requires SrrAB. Alternatively, the SrrAB regulon may be required for virulence because it includes essential staphylococcal virulence factors (Yarwood et al., 2001; Pragman et al., 2004). Indeed, haemolysin D (hld) encoded on the RNAIII molecule was significantly repressed by NO· exposure (Table S1). Thus, the role of SrrAB in NO· resistance in vivo may add to its proposed regulation of staphylococcal virulence determinants and/or its requirement for growth in hypoxic niches of the host.

In summary, this work demonstrates that the nitrosative stress response of S. aureus is required for NO· resistance both in vitro and in vivo. In particular, the staphylococcal Hmp protein provides protection by scavenging host-derived NO·. The SrrAB two-component regulatory system is also required for resistance to NO·, in part due to its activation of hmp and possibly due to its regulation of fermentative metabolic genes. The ability to replicate despite the presence of high concentrations of NO· appears to distinguish S. aureus from other Hmp-producing bacteria including E. coli and B. subtilis. Additional work will be required to determine the mechanism by which S. aureus escapes from NO·-mediated cytostasis. Detoxification and evasion of host-derived NO· can be added to the strategies by which this extraordinarily successful human pathogen is able to subvert the host innate immune system.

Experimental procedures

Chemicals and O2/NO· detection

Nitric oxide donating compounds used in this study were obtained from Alexis Biochemicals (San Diego, CA). Diethylamine NONO-ate (DEA/NO, t1/2 = 2 min), Proline NONO-ate (ProliNO, t1/2 = 1.8 s), and Diethylethylenediamine NONO-ate (NOC-12, t1/2 = 100 min) were solubilized to 500 mM final concentration in 0.01 N NaOH and stored at −80°C. SNAP was resuspended to a final concentration of 500 mM in DMSO and aliquots were stored at −80°C. The conjugate donor compounds diethylamine (Sigma D-0806), proline (Sigma P-0380), diethylethylenediamine (Aldrich 126942), and NAP (Aldrich A19008) were diluted to same concentrations as cognate NONO-ates in like solvents and stored at −80°C. Given an empirically determined NO· mass transfer coefficient of 4.32 × 10−4 cm · s−1 in anoxic phosphate-buffered saline (PBS) at 37°C, and a surface area to volume ratio of 1.3:1, the combination of 1.1 mM DEA/NO and 10 mM NOC-12 resulted in a bolus of NO· that peaked at 2 mM in 15–20 min and dissipated to 1 mM by 4 h [calculated as previously described (Lewis and Deen, 1994)]. These determinations were verified empirically and do not account for rates of autooxidation nor consumption by suspended cells. NO· detection was performed using an ISO-NOPMC Mark II electrode (WPI Instruments, Sarasota, FL) and standard curves generate as per manufacturer instruction. Dissolved oxygen was measured using a Clark-type electrode MLT1120 (ADI Instruments, Milford, MA) run through an Analog Adapter MLT1122 (ADI Instruments).

Bacterial strains and growth conditions

Staphylococcus aureus strains COL, Newman, and RN4220 were generous gifts from W. Shafer (Emory University) as was the E. coli/S. aureus shuttle vector, pBT2 (Bruckner, 1997). pABG5, harbouring the aph-A3 kanamycin resistance determinant, was obtained from C. Collins (University of Washington), and pTA-Erm harbouring an erm cassette from Tn1545 was from I. Stojiljkovic (Emory University). S. aureus strains were either grown in TSB/agar (Difco, Detroit, MI) or PN medium (see below) when appropriate. Chemically defined PN medium was prepared as previously described (Pattee and Neveln, 1975), and consists of a phosphate buffer supplemented with a carbon source (glucose), nitrogen and sulphur sources [(NH4)2SO4 and MgSO4], amino acids, nucleic acid bases, and vitamins (thiamine, niacin, biotin, and pantothenic acid). KRPG was prepared as in Kaplan et al. (1996). Cultures were shaken at 250 rpm at 30°C, 37°C or 43°C when appropriate. For antibiotic selection the following concentrations were used in S. aureus (E. coli): chloramphenicol 10 μg ml−1 (10 μg ml−1), erythromycin 5 μg ml−1 (300 μg ml−1), kanamycin 50 μg ml−1 (50 μg ml−1), and penicillin G (250 μg ml−1). Inactivation of hmp, srrAB and fur was carried out in S. aureus RN4220 using previously described methods (Bruckner, 1997). The hmp, srrAB and fur loci were first PCR amplified using primer sets indicated in Table 1 and cloned into the pCR2.1-Topo Vector (Invitrogen, Carlsbad, CA). An erm cassette was inserted into the unique ClaI site of hmp and the hmp::ErR fragment was cloned into the EcoRI site of pBT2ts to yield pTR40. Likewise, replacing the internal 1.3 kb HincII fragment of srrAB with an erm cassette and moving the srrAB::ErR allele into the EcoRI site of pBT2ts produced pTR43. Finally, pTR46 (fur::KmR) was constructed by inserting the aph-A3 cassette from pABG5 into the unique fur SalI site followed by subcloning the fur::KmR fragment into the EcoRI site of pBT2ts. Mutations were transduced via Φ-11 into S. aureus COL and Newman by methods described in Novick (1991). PCR analysis confirmed insertion of antibiotic resistance cassettes in all backgrounds (see Table 2).

Table 2.  Bacterial strains, constructed plasmids, and primer sets used in this study.
Strain IDGenotypeSource
AR74S. aureus RN4220 hmp::ErRThis study
AR77S. aureus RN4220 srrAB::ErRThis study
AR85S. aureus RN4220 fur::KmRThis study
AR87S. aureus COL hmp::ErRThis study
AR90S. aureus COL srrAB::ErRThis study
AR92S. aureus COL fur::KmRThis study
AR94S. aureus Newman hmp::ErRThis study
AR97S. aureus Newman srrAB::ErRThis study
AR99S. aureus Newman fur::KmRThis study
Plasmid IDDescriptionSource
pABG5aphA3 cassette flanked by SalI sitesC. Collins
pTA-ErErm cassette of Tn1545 flanked by EcoRI and BamHI sitesI. Stojiljkovic
pTR40hmp::ErR in pBT2ts (ErR in internal SspI site in hmp)This study
pTR43srrAB::ErR in pBT2ts (ErR in internal HincII sites in srrAB)This study
pTR46fur::KmR in pBT2ts (KmR in internal SalI site in fur)This study
Primer IDSequence
hmp.1a5′-agaaagacattatcaaacaaacgg-3′
hmp.1b5′-taatgcaaatacttttacgtaacg-3′
srrAB.1a5′-aataactgtatgcgctttcctgtg-3′
srrAB.1b5′-agagtctctaatcaataacatgcg-3′
fur.1a5′-agactatcattgcattgcaacacc-3′
fur.1b5′-accaattgtgttagaacttagtcc-3′
hmpRT.1a5′-tgactttagtgaatttacaccagg-3′
hmpRT.2b5′-cgtttaacgccaaaagttaaatgg-3′
ldh1RT.1a5′-aaaacatgccacaccatattctcc-3′
ldh1RT.1b5′-tactaaatctaaacgtgtttctcc-3′
rpoDRT.1a5′-aactgaatccaagtgatcttagtg-3′
rpoDRT.1b5′-tcatcaccttgttcaatacgtttg-3′
fdaBRT.1a5′-cttagctgataaaggtgttgttcc-3′
fdaBRT.1b5′-aacaacgtctttgataccttgctc-3′
nrdGRT.1a5′-cagtgtttatgtatcaggatgtcc-3′
nrdGRT.1b5′-gttcgccacctaatagacttagcc-3′
ahpFRT.1a5′-accaagtttctcagtcaatcgtcc-3′
ahpFRT.1b5′-ctgtttttctttaggtgcacgacc-3′
acnART.1a5′-gagcaaggtattactaaagtttcc-3′
acnART.1b5′-ctcgccttcatttccatcttttcc-3′
sbnDRT.1a5′-gacagtacttgtgccattattgcc-3′
sbnDRT.1b5′-gagcagcaatcgctataccactcc-3′
cydART.1a5′-catttcgatacatcttcccatgcc-3′
cydART.1b5′-atctgctaacaaactcaatagtcc-3′
trkART.1a5′-acatggatgtaatggccatcgacc-3′
trkART.1b5′-gacatgatcaaagttacggatacc-3′
glnART.1a5′-gcttcgatattgaagctagtcacc-3′
glnART.1b5′-gtgcataccgctaccattcacacc-3′
clpLRT.1a5′-aagcacgtgacggtttattagatcc-3′
clpLRT.1b5′-ttcaacgattgcctgtgctaaacc-3′
2368RT.1a5′-aaggattaacgcaggtatgagtcc-3′
2368RT.1b5′-atcgccaatcactctacccatacc-3′
2535RT.1a5′-gctacaattacagcttatgacgcc-3′
2535RT.1b5′-tgctgcgttaactaagattgcacc-3′

Determination of NO· consumption rate

Five millilitre cultures in PN medium were grown shaking at 37°C to an OD660 ≈ 0.4. Cells were supplemented with 1 mM SNAP or its parent compound NAP, and allowed 25 min continued incubation in ambient fluorescent light. Cells were then resuspended to 1 × 108 cfu ml−1 in 8 ml final volume. A two-hole rubber stopper sealed with parafilm enclosed the cell suspension in an 8 ml glass vial with no gaseous headspace. Cells were stirred vigorously at 37°C as ProliNO was added through one open port to 1 μM. The resulting immediate release of c. 2 μM NO· followed by the gradual decay of detectible signal was recorded and normalized to the fraction of initial [NO·]. Measurements were performed in triplicate for each strain tested.

Virulence determination

Six-week-old C57BL/6 mice (Jackson Laboratories, Bar Harbor, ME) or C57BL/6 iNOS–/– mutant mice were housed in Modified Specific Pathogen Free facilities with access to food and water ad libitum. Overnight cultures of S. aureus Newman and derivatives grown in TSB were washed twice in sterile PBS. Mice were injected via tail vein with 5 × 107 cfu of S. aureus Newman in a 100 μl volume. Survival was monitored over 2 weeks, and moribund animals were euthanized per IACUC-approved protocols. Alternatively, mice were injected intravenously with 2 × 107 cfu of S. aureus Newman and derivatives and sacrificed via CO2 asphyxiation 5 days post infection. Kidneys were homogenized in sterile PBS and viable cfu per gram was determined by plating serial dilutions of tissue homogenate.

RNA isolation

RNA was isolated from mid-log (OD660 ≈ 0.5) cultures of S. aureus COL and derivatives using Qiagen RNA-Midi/-Mini Kits (Valencia, CA) with modifications for cell disruption. For microarray/Q-RT PCR analysis, 75 ml/5 ml of bacterial culture was harvested at desired times and immediately diluted 1:1 in ice-cold Ethanol:Acetate then incubated at −80°C for ≥ 20 min. Cells were pelleted, resuspended in 1 ml RNase-free TE [or 700 μl RTL (Qiagen) for scaled down Q-RT PCR preps] then added to tubes containing lysing matrix B (Q-biogene, Irvine, CA) for mechanical disruption. Lysing matrix suspension was centrifuged at 10 000 g for 10 min at 4°C and the supernatants decanted for further preparation using Qiagen Midi/Mini Spin-Prep procedures. An on-the-column DNase treatment (25 min at room temperature) preceded the final washes and elution of RNA in RNase-free ddH2O.

cDNA microarray analysis

The Affymetrix S. aureus GeneChips (Santa Clara, CA) used in this study represented genomic sequences, including intergenic regions, from S. aureus strains NCTC 8325, COL, N315 and Mu50. Total RNA from three independent experiments was labelled, pooled and hybridized in duplicate, as previously described (Roberts et al., 2006). Each qualifier signal intensity value from a given GeneChip was normalized to the median signal value of that GeneChip using GenesSpring 6.2 Software (Silicon Genetics, Redwood City, CA). Normalized signal intensities from GeneChips hybridized with SNAP-exposed versus NAP-exposed total RNA were then used to determine intensity ratios for each probe set and averaged over the two technical replicates (Table S2). Z-values were calculated as follows (Larsson et al., 2005):

Z(i) = [x(i) − μ(i)]/SD

Where x(i) is the average log2 induction ratio of the ith probe set, μ(i) represents the mean log2 induction ratio for all probe sets (0.033), and SD is the standard deviation about μ(i) (0.185). Thus Z(i) is a measure of the number of standard deviations that the intensity ratio of the ith probe set differs from that of the mean intensity ratio for the array. An arbitrary cut-off of −3 ≥ Z(i) ≥ 3 was chosen as probe sets that represent differentially expressed genes within a 99.73% confidence interval. A total of 84 genes were either induced ≥ 1.50-fold or repressed ≤ 1.43-fold and thus met these requirements (Table S1 and Fig. S1). As controls, technical replicates of the two SNAP-exposed RNA pools as well as the two NAP-exposed pools were analysed and only eight of the 4984 total probe sets met the above 3 ≤ Z(i) ≤ −3 criteria (Fig. S2).

For verification of cDNA microarray analysis, quantitative real-time PCR was performed on RNA extracts from SNAP- and NAP-exposed cultures. RNA was spectrophotometrically quantified and 50 ng of total RNA was analysed per reaction using the QuantiTect™ SYBR® Green RT-PCR kit (Qiagen). Reaction conditions were as specified by Qiagen and reactions were performed and analysed using a Rotor Gene™ 2000 Real Time Cycler (Corbett Research, Sydney, Australia).

Acknowledgements

This work was supported by the National Institutes for Health (AI039557 to F.C.F and AI055396 to A.R.R.).

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