The msaABCR Operon Regulates the Response to Oxidative Stress in Staphylococcus aureus

This study shows the involvement of the msaABCR operon in resisting oxidative stress by Staphylococcus aureus generated under in vitro and ex vivo conditions. We show that MsaB regulates the expression and production of a carotenoid pigment, staphyloxanthin, which is a potent antioxidant in S. aureus. We also demonstrate that MsaB regulates the ohr gene, which is involved in defending against oxidative stress generated by organic hydroperoxides. This study highlights the importance of msaABCR in the survival of S. aureus in the presence of various environmental stimuli that mainly exert oxidative stress. The findings from this study indicate the possibility that msaABCR is involved in the persistence of staphylococcal infections and therefore could be a potential antimicrobial target to overcome recalcitrant staphylococcal infections.

ABSTRACT

Staphylococcus aureus has evolved a complex regulatory network that controls a multitude of defense mechanisms against the deleterious effects of oxidative stress stimuli, subsequently leading to the pathogen’s survival and persistence in the hosts. Previously, we characterized the msaABCR operon as a regulator of virulence, antibiotic resistance, and the formation of persister cells in S. aureus. Deletion of the msaABCR operon resulted in the downregulation of several genes involved in resistance against oxidative stress. Notably, those included carotenoid biosynthetic genes and the ohr gene, which is involved in resistance against organic hydroperoxides. These findings led us to hypothesize that the msaABCR operon is involved in resisting oxidative stress generated in the presence of both H₂O₂ and organic hydroperoxides. Here, we report that a protein product of the msaABCR operon (MsaB) transcriptionally regulates the expression of the crtOPQMN operon and the ohr gene to resist in vitro oxidative stresses. In addition to its direct regulation of the crtOPQMN operon and ohr gene, we also show that MsaB is the transcriptional repressor of sarZ (repressor of ohr). Taken together, these results suggest that the msaABCR operon regulates an oxidative stress defense mechanism, which is required to facilitate persistent and recurrent staphylococcal infections. Moving forward, we plan to investigate the role of msaABCR in the persistence of S. aureus under in vivo conditions.

IMPORTANCE This study shows the involvement of the msaABCR operon in resisting oxidative stress by Staphylococcus aureus generated under in vitro and ex vivo conditions. We show that MsaB regulates the expression and production of a carotenoid pigment, staphyloxanthin, which is a potent antioxidant in S. aureus. We also demonstrate that MsaB regulates the ohr gene, which is involved in defending against oxidative stress generated by organic hydroperoxides. This study highlights the importance of msaABCR in the survival of S. aureus in the presence of various environmental stimuli that mainly exert oxidative stress. The findings from this study indicate the possibility that msaABCR is involved in the persistence of staphylococcal infections and therefore could be a potential antimicrobial target to overcome recalcitrant staphylococcal infections.

INTRODUCTION

Staphylococcus aureus is an important human pathogen that causes a range of pathologies, from minor soft tissue infections to chronic and often life-threatening conditions, such as septicemia, endocarditis, and pneumonia (1, 2). The acquisition of antibiotic resistance and/or tolerance and its numerous virulence factors contribute to this pathogen’s ability to cause such a variety of infections (3–6). In addition, this pathogen has acquired the ability to evade or invade the host defense system to adapt and proliferate inside host immune cells during infectious processes, leading to persistent infections (7–9). Host immune cells, such as neutrophils, monocytes, and macrophages, express NADPH oxidase, which is responsible for generation of the superoxide anion (O₂^·−) during oxidative bursts (10). The O₂^·− is presumed to contribute to bacterial killing and is involved in an array of chemical reactions that generate reactive oxygen species (ROS), including hydroxyl radicals (OH·), peroxynitrite (ONOO⁻), and H₂O₂, which are all highly lethal to bacterial cells (11). Killing by the host immune cells depends primarily on damaging the pathogen’s lipids, proteins, and DNA. To combat these lethal effects, the pathogen has evolved complex regulatory systems. These systems include the characteristic golden pigment carotenoid (crtMN) (12, 13), catalase (katA) (14, 15), and superoxide dismutases (sodA-M) (16–18). In addition, many genes, including those of the PerR regulon (katA, mrgA, zosA, and ohrA) and the σ^B regulon, have been shown to be involved in the response to oxidative stress (19, 20).

S. aureus also possesses several peroxiredoxins, which are induced on exposure to H₂O₂ (21, 22). These include two enzymatic groups, the alkyl hydroperoxide reductase (ahpC) family and the osmotically inducible protein/organic hydroperoxide reductase superfamily, both associated with detoxification of alkyl and organic hydroperoxides (23). Using NADH or NADPH as reducing equivalents, peroxiredoxins detoxify alkyl hydroperoxides by reducing them to their alcohols. Moreover, S. aureus AhpC also confers resistance to a wide spectrum of ROS (15). Since oxidative stress imparts deleterious effects to all aspects of physiology, including the enzymatic activity of bacterial pathogens, the regulation of defensive systems against these stimuli is complex (13). Alteration of enzymatic activity in turn changes metabolite concentrations and redox poise. These changes ultimately alter the activity of redox or metabolite-responsive regulators such as CcpA or CodY (24, 25). The global regulators MgrA, SarZ, and PerR are important S. aureus proteins that are involved in sensing oxidative stresses (22, 26). Possession of all these oxidative stress-responsive systems contributes to the success of S. aureus as a human pathogen capable of causing persistent infections (27).

Indeed, S. aureus has been reported to be a major intracellular persisting pathogen in chronic and recurrent infections (6, 28). Richards et al. (29) reported the existence of S. aureus clinical isolates with unique immune evasion ability and increased levels of stress-response proteins, including cold shock protein CspA, in persistent bacteremia cases (29). In addition to this study, several other studies also reported increased levels of MsaB (also known as CspA) and oxidative stress-response proteins, including thioredoxin (Trx), in staphylococcal infections (29–31), highlighting their importance in stress response and their role in causing persistent S. aureus infections in the host. In our previous studies, we reported a regulatory role for the msaABCR operon (MsaB) in capsule production (32, 33), biofilm development (34), antibiotic resistance (35), and the formation of persister cells in S. aureus (36). On transcriptomic analysis, deletion of the msaABCR operon in the USA300 LAC strain induced differential expression of more than 10 genes that are involved in resisting oxidative stress (36). Among the downregulated genes, notably, were the carotenoid biosynthetic genes crtM and crtN, ohr (SAUSA300_0786), and katA. The carotenoid pigment is the membrane-bound orange-red C₃₀ triterpenoid staphyloxanthin (STX), the biosynthesis of which is encoded by the genes within the operon containing crtOPQMN and aldH (37, 38). In this study, we observed an isogenic msaABCR deletion mutant in both USA300 LAC and UAMS strains with reduced expression of crt genes and production of STX, which is a potent staphylococcal antioxidant. Therefore, we hypothesized that MsaB is a positive transcriptional regulator of crtOPQMN, an STX biosynthetic operon. In support of this hypothesis, we here demonstrate modulation of STX production by MsaB via direct regulation of the crtOPQMN operon. Furthermore, we also show differential expression of its transcripts with sigB inactivation, a positive transcriptional regulator of the crtOPQMN operon. We show the importance of the ohr gene and its transcriptional regulation by MsaB for resistance against organic hydroperoxide. In addition, we also demonstrate the transcriptional regulation of sarZ, which negatively regulates the ohr gene.

Understanding the S. aureus response under defined in vitro stressed conditions is a prerequisite for obtaining a more comprehensive explanation of persistence in host cells, which certainly requires further investigation. This study will add new insights into the regulatory role of a transcriptional regulator, MsaB, in resisting oxidative stress, which aids S. aureus in causing persistent infections.

RESULTS

msaABCR deletion mutants are more susceptible to oxidative stress in vitro.

To study the role of the msaABCR operon in response to oxidative stress in S. aureus, in vitro survival assays in the presence of inorganic hydrogen peroxide (H₂O₂), organic hydroperoxides such as tert-butyl hydroperoxide (tBOOH) and cumene hydroperoxide (CHP), and paraquat were performed. To test the effect of msaABCR mutants on H₂O₂ resistance, we employed an oxidative stress assay as previously described (39, 40). Briefly, postexponentially grown cells of wild-type USA300 LAC and UAMS-1 strains, an isogenic msaABCR deletion mutant, and their complementation cells normalized to and optical density at 600 nm (OD₆₀₀) of 0.1 in phosphate-buffered saline (PBS) were exposed to a final concentration of 34 mM H₂O₂ for 1 h at 37°C. The survival percentage measured after the incubation showed increased susceptibility of msaABCR mutant cells (1-log-fold reduction compared to that of the wild type), with 10% to 12% survival in wild-type and 8% to 10% in complementation strains, while survival was ∼0.5% for msaABCR mutant cells in both strain backgrounds (Fig. 1). We also tested the growth pattern of the msaABCR mutant in the presence of 25 mM H₂O₂ in the nutrient-rich tryptic soy broth (TSB) medium, as previously described (41). We observed that the growth of USA300 LAC and complementation cells was halted for 5 h. After this tolerance phase, the wild-type and complementation cells resumed growth to similar levels as those under unstressed conditions. However, growth of the msaABCR mutant was reduced gradually to below the limit of detection (100 CFU ml⁻¹), resulting in a clear culture (Fig. 2A and C). On the other hand, the growth of UAMS-1 and msaABCR mutant cells was reduced in the presence of 25 mM H₂O₂ compared to that of the unstressed cells after an overnight incubation, while such growth delay was not observed in complementation cells (Fig. 2B and D). To confirm whether the killing was by ROS production, we added 50 mM thiourea, an ROS scavenger, to the msaABCR mutant cells after 10 min of H₂O₂ treatment. In the presence of thiourea, they resumed growth similar to that under the H₂O₂-unstressed condition (Fig. 2E). These observations suggest that the msaABCR mutant is more susceptible to ROS generated from H₂O₂. Likewise, to test susceptibility to organic hydroperoxide, we employed plate assays as previously described by Chen et al. (22) using tryptic soy agar (TSA) plates containing 5 mM tBOOH and 4 mM CHP. In the presence of both hydroperoxides, after 24 h of incubation, we observed a significantly decreased survival percentage of msaABCR mutant cells compared with those of wild-type and complementation cells in both USA300 LAC and UAMS-1 strains (Fig. 3).

FIG 1.

Oxidative stress survival assay in the presence of H₂O₂. S. aureus cells grown for 24 h were washed, normalized to an OD₆₀₀ of 0.1 in 1 ml phosphate-buffered saline (PBS) containing 34 mM H₂O₂, and incubated for 1 h at 37°C. After incubation, catalase (2,000 U ml⁻¹) was added, and the mixture was incubated for 1 min at room temperature to decompose H₂O₂. Cells were diluted and plated for CFU count. The survival percentages represent the percentage of initial CFU that survived the H₂O₂ exposure in 1 h. The data present the standard errors of the means (SEMs) of the results from at least three independent experiments. Student’s unpaired t tests were used to compare the results from wild types and isogenic mutants. ***, P ≤ 0.001; FSmut-msaB, msaABCR complementation with frameshift-mutated msaB.

FIG 2.

Growth of msaABCR mutant cells is severely disrupted in the presence of 25 mM H₂O₂. S. aureus cells grown for 24 h were normalized to an OD₆₀₀ of 0.1 in 5 ml of TSB with 25 mM H₂O₂. (A and B) Growth was monitored by CFU counts at different time points as indicated. (C and D) Observation of growth of cultures with and without 25 mM H₂O₂ incubated overnight. (E) Addition of 50 mM thiourea to the H₂O₂-stressed cells. The data present the means ± SEMs of the results, which were repeated at least three times.

FIG 3.

Oxidative stress assay in the presence of the organic hydroperoxides tert-butyl hydroperoxide (tBOOH) and cumene hydroperoxide (CHP). S. aureus cells grown for 16 h were harvested, washed, and normalized in PBS to an OD₆₀₀ of 6.0 for all strains tested. Then, 10 μl of serially diluted cells (10⁻¹ to 10⁻⁸) was dropped onto the TSA plates containing 5 mM tBOOH (A and B) and 4 mM CHP (C and D). Plates were incubated 24 h for CFU counts. Graphs represent the percentages of survived CFU. Error bars represent SEMs of the results repeated three times. Student’s unpaired t tests were used to compare the results from wild types and isogenic mutants. *, P ≤ 0.05; **, P ≤ 0.005.

The upregulated expression of superoxide dismutases (SODs) in msaABCR mutants led us to test the susceptibility to paraquat, which generates O₂^·−. The strains, including sodM transposon mutants, were monitored in the presence of 40 mM paraquat. While the addition of paraquat resulted in growth arrest, it did not show a killing effect in any strain for an extended period (Fig. 4). The SODs convert O₂^·− to H₂O₂, which in turn is degraded to H₂O and O₂ by catalases, mainly encoded by katA (23). A decreased transcript level for the katA gene in the msaABCR mutant (Table 1) led us to examine whether this mutant has decreased catalase activity. Within 15 min, 91.17 ± 1.31 μM and 92.22 ± 1.91 μM of exogenously added 100 μM H₂O₂ was degraded in wild-type USA300 and msaABCR mutant cells, respectively. Similarly, in UAMS-1, 91.2 ± 2.09 μM and 92.38 ± 1.29 μM of H₂O₂ were degraded with ∼99% degradation in wild-type and mutant strains, respectively, within 1 h (Fig. 5A and B). The control katA transposon mutant (USA300 LAC kat::Tn) and TSB medium without bacteria showed no catalase activity, while the addition of catalase completely detoxified H₂O₂ within 15 min (Fig. 5C). Therefore, the lack of sensitivity to paraquat appears also to depend on a similar ability to detoxify the H₂O₂ produced by SODs in the presence of paraquat. Taken together, these results indicate that protection of S. aureus by msaABCR against oxidative stress, although specific for H₂O₂, does not appear to depend on detoxification ability, at least in the presence of low concentrations of H₂O₂.

FIG 4.

Oxidative stress assay in the presence of paraquat in USA300 LAC (A) and UAMS-1 (B) strains. Cells grown for 24 h were washed and normalized to an OD₆₀₀ of 0.1 in 1 ml PBS. The cells (100 μl) were dispensed into 96-well plates supplemented with and without 40 mM paraquat (PQ) and incubated at 37°C with continuous shaking. Growth was monitored by taking OD₆₀₀ measurements at designated time points. The sodM mutants (sodM::Tn) for each strain were also included in the experiment as controls. The data represent the SEMs of the results from three independent experiments.

TABLE 1.

Expression of oxidative stress response genes in the msaABCR deletion mutant relative to that in wild-type strains

Gene (locus IDa )	Functional name	Fold change in expressionb
		USA300 LAC	UAMS-1
crtM (SAUSA300_2499)	Squalene desaturase	−3.58 ± 0.24	−3.25 ± 0.05
crtN (SAUSA300_2498)	Squalene synthase	−2.99 ± 0.74	−2.25 ± 0.15
ohr family protein (SAUSA300_0786)	OsmC/Ohr family protein	−3.8 ± 0.58	−2.18 ± 0.11
katA (SAUSA300_1232)	Catalase	−2.38 ± 0.48	−3.7 ± 0.10
sodA (SAUSA300_0135)	Superoxide dismutase	+2.98 ± 0.25	+3.44 ± 0.74
sodM (SAUSA300_1531)	Superoxide dismutase	+3.24 ± 0.14	+2.67 ± 0.54
Membrane protein gene (SAUSA300_0374)	Putative membrane protein	−3.2 ± 0.15	−7.4 ± 1.83
ahpC (SAUSA300_0380)	Alkyl hydroperoxide reductase	−1.01 ± 0.28	−4.49 ± 0.83
trx (SAUSA300_0747)	Thioredoxin	−1.05 ± 0.19	+1.8 ± 0.70
dps (SAUSA300_2092)	General stress protein 20U	−1.26 ± 0.12	−1.28 ± 0.14
Glutathione peroxidase gene (SAUSA300_1197)	Glutathione peroxidase	+1.28 ± 0.41	−2.65 ± 0.44
uvrA (SAUSA300_0742)	Excinuclease ABC subunit A	−1.02 ± 0.78	+2.43 ± 0.37

^aID, identifier.

^bValues are means ± standard errors of the means. Results are representative of at least three independent experiments for each sample set.

FIG 5.

Hydrogen peroxide detoxification assay in wild-type, msaABCR deletion mutant, and complementation cells in USA300 LAC (A) and UAMS-1 (B) strains. Cells grown for 16 h were prewashed with PBS and normalized to 2 × 10⁵ cells in 5 ml TSB containing 100 μM H₂O₂. The cells were incubated for 1 h at 37°C, and 1-ml culture aliquots were taken every 15 min to measure remaining H₂O₂ using an H₂O₂ detoxification kit (Cell Biolabs). (C) Control H₂O₂ detoxification assay using katA::Tn mutant, katA::Tn plus 10 mM catalase, and in the absence of bacteria. The graph represents the amount of H₂O₂ degraded at designated time points. The data represent the SEMs of the results from three independent experiments.

msaABCR mutant cells have low NADPH levels and low total antioxidant capacity compared with those of the wild type.

The elimination of pathogens by phagocytic cells depends largely on the ROS generated by NADPH oxidase (42). In Gram-negative bacteria, the cellular NADPH level has also been shown to confer resistance against oxidative stress (43). In this study, we sought to examine whether deletion of the msaABCR operon results in changes in the NADPH level in S. aureus. We observed reduced levels of NADP⁺ and NADPH in the msaABCR mutant relative to those in the wild type, while in the complementation strain, an increased NADPH level was observed, especially in the case of UAMS-1 (Fig. 6). The decreased level of NADPH suggests a reduced redox capacity of the msaABCR mutant cells relative to that of the parental strains. We therefore measured the total antioxidant capacity (TAC), which is proportional to copper-reducing equivalents, using the OxiSelect total antioxidant capacity assay kit. We observed reduced levels of TAC in msaABCR mutant cells relative to that in the isogenic wild-type and complementation cells, indicating that the deletion of the msaABCR operon renders S. aureus cells with reduced redox capacity (Fig. 7).

FIG 6.

Concentrations of NADP⁺ and NADPH in USA300 LAC (A) and UAMS-1 (B) strains. Overnight cultures were diluted to an OD₆₀₀ of 0.05 and grown until late exponential phase (OD₆₀₀ of 4.0). Cells were harvested and washed with ice-cold PBS, and NADPH levels were measured according to the manufacturer’s instructions (Biovision Inc.). Values represent the absolute concentrations of NADP⁺ and NADPH per microgram of protein. Error bars represent the SEMs from three independent experiments. Student’s unpaired t tests were used to compare the results from wild types and isogenic mutants. *, P ≤ 0.05.

FIG 7.

Measurement of total antioxidant capacity (TAC). Cells grown to an OD₆₀₀ of ∼0.7 were washed thrice and resuspended in 600 μl of ice-cold PBS. Cells were then freeze-thawed twice before mechanical lysis, and the protein was collected by centrifugation at 5,000 × g for 10 min at 4°C. The protein concentrations of the samples were determined using the BCA protein assay kit (Pierce), and the remaining portions of the samples were assayed using the OxiSelect total antioxidant capacity assay kit (Cell Biolabs). Assays were performed in duplicate and repeated once, and values represent relative levels compared with wild-type strains, which were set at 100%. Error bars represent the SEMs. Student’s unpaired t tests were used to compare the results from wild types and isogenic mutants. *, P ≤ 0.05.

We next sought to examine whether reduced survival due to in vitro oxidative stresses along with reduced NADPH and total antioxidant capacity in the msaABCR mutant is translated into decreased bacterial resistance against innate immune clearance in ex vivo systems, including macrophages, neutrophils, and whole-blood cells that additionally contain complementing proteins. Postexponentially grown cultures of bacterial cells were incubated with macrophages (RAW 264.7 cells), freshly collected neutrophils, and heparinized whole-blood cells from volunteers. We observed that msaABCR mutant cells were significantly defective in intracellular survival within macrophages, human neutrophils, and whole-blood cells. Complementation of the mutants with a functional copy of the msaABCR operon restored their survival to a level comparable with that of the wild type in neutrophils and human blood and to lesser extent (∼70% survival compared to wild types) in macrophages, indicating that msaABCR is required for S. aureus to evade host immune systems, which produce ROS that kill pathogens (Fig. 8).

FIG 8.

Ex vivo survival assays with macrophages, neutrophils, and whole-blood cells. (A) S. aureus cells with a multiplicity of infection (MOI) of 10 were incubated with macrophages (RAW 264.7 cells) at 37°C in the presence of CO₂. (B) S. aureus cells were incubated with heparinized human whole blood and freshly isolated human neutrophils to measure their intracellular survival rates. Results with human blood and neutrophils for the UAMS-1 strain were reported in a previous study (32). The data represent the means from three independent experiments. Error bars represent the standard errors (SEs). Student’s unpaired t tests were used to compare the results from wild types and their isogenic msaABCR mutants. **, P ≤ 0.005.

The msaABCR operon regulates the genes involved in responding to oxidative stress.

Previously, our RNA transcriptomic data showed that more than 10 genes involved in the oxidative stress response were differentially expressed in the msaABCR deletion mutant in the USA300 LAC strain (36). Our transcriptomic results showed that the genes encoding dehydrosqualene synthase (crtM) and dehydrosqualene desaturase (crtN), which are involved in the staphyloxanthin biosynthetic pathway, were downregulated by 9.8- and 8.6-fold, respectively. In addition, the organic hydroperoxide reductase gene (ohr) was also downregulated by 13.8-fold in the msaABCR mutant (36). In this study, we further confirmed the differential expression of oxidative stress-resistant genes by quantitative real-time PCR (qRT-PCR) in the msaABCR mutant relative to that in the isogenic parental USA300 LAC and UAMS-1 strain backgrounds at the stationary growth phase (Table 1). We observed downregulated expression of crtM, crtN, ohr, katA, and putative membrane protein genes in the msaABCR operon mutant in both strain backgrounds. However, the alkyl hydroperoxide reductase gene (ahpC) was downregulated only in the UAMS-1 msaABCR mutant strain, with no apparent change in the USA300 LAC msaABCR mutant strain. Interestingly, however, the ROS-detoxifying superoxide dismutase genes sodA and sodM were upregulated in the operon mutants of both strains (Table 1). These results show that deletion of the msaABCR operon in S. aureus leads to the differential expression of genes involved in resisting oxidative stress.

MsaB is a positive transcriptional regulator of the crtOPQMN operon.

Previously, we showed that deletion of the msaABCR operon significantly reduced pigmentation in S. aureus (44). On trans complementation of the msaABCR mutant with the native msaABCR operon, the STX level was completely restored. However, trans complementation with the frameshift mutant (FSmut)-msaB msaABCR operon did not restore STX production in the msaABCR mutant. In fact, the FSmut-msaB msaABCR operon showed a similar level of STX defect as the msaABCR operon mutant (Fig. 9). Furthermore, the trans-complemented FSmut-msaB msaABCR operon cells also showed similar susceptibility to H₂O₂ killing as msaABCR cells (Fig. 1). Taken together, these observations indicate that the msaB gene and/or protein is responsible for the STX production that corresponds to H₂O₂ susceptibility in S. aureus. In addition to significant downregulation of the crtM and crtN genes, our proteomics results also revealed a decreased level of carotenoid protein in the USA300 LAC msaABCR mutant. To further verify the importance of STX in ROS resistance in our strain background, the crtM mutant, which is devoid of pigment production, was exposed to H₂O₂ stress. We observed crtM cells with comparable susceptibility to H₂O₂ as that of msaABCR mutant cells. All these findings indicate the importance of functional MsaB in resisting H₂O₂ killing, which is likely to be mediated through STX production.

FIG 9.

Measurement of staphyloxanthin (STX) production. (A) Standardized S. aureus cells in 10 ml TSB were grown for 16 h at 37°C with continuous shaking. Cells were harvested, washed with deionized water, and normalized to an OD₆₀₀ of 10 in 1 ml. Then, STX was extracted using the methanol extraction method. (B) The corresponding pictorial representation of pigmentation. The values are presented relative to the wild type normalized to 1. The data represent the SEMs of the results from four independent experiments. Student’s unpaired t tests were used to compare the results from wild types and mutants. *, P ≤ 0.05.

To investigate whether the MsaB regulation of the crtOPQMN operon is direct, we performed a chromatin immunoprecipitation (ChIP) assay using an MsaB antibody at the stationary growth phase, in which STX production is visible. The cross-linking of MsaB to its target DNA promoters was facilitated by treating stationary-phase cells with sodium phosphate and formaldehyde. The MsaB-DNA complexes were then pulled down using an MsaB antibody. To confirm specificity, the msaABCR mutant cells were included as a negative control in each assay. In our previous study, we demonstrated the existence of a consensus sequence including the TTGTTTAAA region upstream of the cap promoter for MsaB binding (32). The primers specific for the crtOPQMN operon promoter region comprising the consensus sequence of MsaB were designed and used to test the amplification. The binding activity of MsaB revealed via ChIP assay indeed confirms that MsaB is a direct regulator of the crtOPQMN operon (Fig. 10A). Furthermore, we also employed an electrophoretic mobility shift assay (EMSA) to observe the binding of MsaB to the crtOPQMN operon. We prepared purified MsaB protein and examined the gel shift of the protein-DNA complex with an amplified 5′-biotinylated crtOPQMN operon promoter region. The presence of a shifted band as a result of MsaB binding to the 5′-biotinylated crtOPQMN operon promoter thus confirmed that MsaB directly regulates the crtOPQMN operon (Fig. 11A).

FIG 10.

Chromatin immunoprecipitation (ChIP) assay. ChIP using an anti-MsaB antibody was performed to determine whether MsaB binds to the promoter region of the crtOPQMN operon (A), the ohr promoter (B), and the sarZ promoter (C). Primers specific for the corresponding promoter regions were used to amplify DNA after immunoprecipitation of wild-type USA300 LAC cells. msaABCR, the negative control, which is a whole-cell extract from the msaABCR deletion mutant with anti-MsaB antibody; + Control, PCR amplification from genomic DNA of the wild-type strain; LAC::Tn SigB, immunoprecipitated DNA isolated from a transposon mutant of sigB in the USA300 LAC strain; P, promoter region of corresponding gene.

FIG 11.

Electrophoretic mobility shift assay (EMSA). (A) The shifted DNA plus protein complex is shown when an increased concentration of MsaB-His was incubated with a biotin-labeled crt promoter target (labeled probe). The gel shift was reversed when 100 M excess concentration (50 pmol) of nonlabeled specific crt promoter probe (unlabeled probe) was added to the reaction mixture. *, nonlabeled nonspecific gene (unlabeled probe) was added to the reaction to rule out nonspecificity (that the addition did not affect the binding). (B) The shifted DNA plus protein complex is shown when an increased concentration of MsaB-His was incubated with the biotin-labeled ohr promoter target (labeled probe). **, mutated ohr probe. Binding was not observed in the mutated probe (lane 7). The assay was performed in a total reaction volume of 20 μl, and the assay mixture was incubated at room temperature for 20 min according to the manufacturer’s instructions (LightShift chemiluminescent EMSA kit).

Previous studies have shown sigB to be a positive transcriptional regulator of the crtMN operon (39, 45). In this study too, we further verified decreased crtM expression (4.07-fold) by qRT-PCR and also observed reduced pigmentation in the sigB mutant of the USA300 LAC strain. Deletion of msaABCR downregulated the transcription of sigB by 3.2- and 2.31-fold in the USA300 LAC and UAMS-1 strain backgrounds, respectively (Table 2). To examine whether SigB affects the binding of MsaB to the crtOPQMN operon promoter, we employed a ChIP assay with a sigB transposon mutant of the USA300 LAC strain. Our result showed that MsaB also binds to the crtOPQMN promoter in the absence of sigB (Fig. 10A). However, MsaB binding to the crtOPQMN operon alone is not sufficient to activate crtMN expression, since the sigB mutant also displayed reduced crtM and/or STX production. These results suggest that both MsaB and SigB are required for crtOPQMN operon expression. We were then interested to see how msaB transcription was affected by sigB. The sigB mutation resulted in upregulated msaB transcription by 9-fold, indicating that sigB is a repressor of msaB expression (Table 3). These results indicate the interaction between MsaB and SigB and the mechanism of regulation of the crtOPQMN operon are indeed complex and need further investigation for complete understanding.

TABLE 2.

Expression of genes in the msaABCR deletion mutant relative to that in wild-type strains

Gene (locus ID)	Fold change in expressiona
	USA300 LAC	UAMS-1
sigB (SAUSA300_2025)	–3.29 ± 0.12	–2.31 ± 0.14
ssrA (SAUSA300_ 0766)	+2.21 ± 0.08	+10.83 ± 0.19
sarZ (SAUSA300_2331)	+2.65 ± 0.05	+3.39 ± 0.18

^aValues are means ± standard errors of the means. Results represent at least 3 independent experiments in each sample set.

TABLE 3.

Expression of msaB in sigB and sarZ mutants in the USA300 LAC strain

Mutant type	msaB expression relative to that in the wild type (mean ± SEM)a
sigB	+9.4 ± 1.04
sarZ	+5.7 ± 0.32

^aValues are fold changes. Results represent at least 3 independent experiments in each sample set.

MsaB positively regulates the ohr gene to resist oxidative stress generated in the presence of organic hydroperoxides.

The ohr gene encodes a protein that confers resistance to organic hydroperoxide in different bacterial species, including Escherichia coli, Pseudomonas aeruginosa, Xanthomonas campestris, and S. aureus (46–50). Deletion of msaABCR leads to the downregulation of ohr transcripts (Table 1). We observed that ohr mutant cells have increased susceptibility to tBOOH- and CHP-induced oxidative stress that was comparable to that of msaABCR mutant cells (Fig. 3). To examine whether the altered sensitivity to organic peroxides in msaABCR mutant cells was due to repression of ohr, a plasmid for inducible expression of the ohr gene (msaABCR::POhr) was constructed and then transformed into msaABCR mutant cells. Overexpression of the ohr gene in the msaABCR mutant cells increased resistance against tBOOH and CHP to a level comparable to that of the wild type. These results suggest that the response of MsaB to organic hydroperoxide stress is likely to be dependent on the ohr gene in S. aureus (Fig. 3). To observe the direct regulation of ohr transcripts by MsaB, we employed ChIP and EMSA using the previously published ohr promoter region sequence (see Table 5) (22). In our previous study, we demonstrated existence of a consensus sequence including the TTGTTTAAA region upstream of the cap promoter for MsaB binding. When this region was mutated, binding was affected, indicating specific MsaB binding activity in this promoter region (32, 33). In this study, we mutated this region TTTAAA to TTGGAA in the promoter region of ohr gene. As expected, we found that this mutation abolished binding by MsaB. Therefore, these specific binding results confirmed that MsaB directly regulates the ohr gene (Fig. 10B and 11B). A previous study by Chen et al. (22) showed that SarZ is a multigene regulator (Mgr) homologue that negatively regulates the ohr gene in S. aureus strain Newman (22). Under stationary growth conditions, we observed that sarZ expression was upregulated by 2.65- and 3.39-fold in the msaABCR mutants of the USA300 LAC and UAMS-1 strains, respectively (Table 2). We then sought to determine whether MsaB also regulates sarZ expression. The positive binding activity observed in the ChIP assay indeed demonstrated that MsaB also regulates sarZ (Fig. 10C). The findings from this study thus led us to conclude that MsaB directly regulates the ohr gene as a transcriptional activator and sarZ as a transcriptional repressor.

TABLE 5.

List of primers used in this study

Function and primer name	Sequence (5′→3′)a
For qRT-PCR
gyrA F	GCCGTCAGTCTTACCTGCTC
gyrA R	TTTGCATCCTTACGCACATC
gyrB F	GGTGCTGGGCAAATACAAGT
gyrB R	TCCCACACTAAATGGTGCAA
crtM F	AAGTATTTAGTATTGAAGCACAAC
crtM R	ACGTTCATGTAATGTATAGTTAGC
crtN F	TACTAGGATTTTTAGCGTCGAAAC
crtN R	AAGAAGTTTTTGATGAAGCGATAG
ohr F	CGTAAAGGACATGTTTATACTGATGA
ohr R	TGTAAATATTTTTCAGCTTCTTCTTG
katA F	TTTAGTAGGGAATAACACACCAGT
katA R	CTCATATTTGTTCTAGGATCTCGT
sodA F	ACTAGCGGATATGATTGCTAACTT
sodA R	ATTTCCCAGAATAATGAATGGTTA
sodM F	CTGTACCTTCTACTGCAGCATTTA
sodM R	TTAGAACCACATTTTGACAAAGAA
Membrane protein F	ATGTTTGGATTTATTGGAATGTTAAT
Membrane protein R	AAAATACCACCTGGGATATCTTTAC
ahpC F	TTAGGCGTAAATGTATTCTCAGTA
ahpC R	CACCAATCATAGTGTAAGTGATTT
trx F	CTATGAAGGTAAAGCTGACATTTT
trx R	TTTAAAGACGATTAATGTTGGAAT
dps F	AAAAGAATTGAATCAACAAGTAGC
dps R	AACGTGTAATGAGAAGAAGTTAGG
Glutathione peroxide F	CGTTATGCTAATTGTTAATACAGC
Glutathione peroxide R	ATTACAAACCCTTGATCTTTGTAT
uvrA F	ACACGTGAAATGATGAGTAAAT
uvrA R	CCACCTACATAAACAGATAACG
sarZ F	GTTCTTAGATTCTGGAACACTGAC
sarZ R	TTGTAGGTTTCTTTCATCTTTCTC
sigB F	GAAGCTAAGTCTATCTCTTTATCGTGAA
sigB R	CAAGAAATCGTTAAAGGCTTTGGTTATA
ssrA F	ATCAAACAATAATTTCGCAGTA
ssrA R	GCATATCCTATTAAGGTTGAATC
For frameshift mutant msaB (FSmut-msaB)
Fsmut-msaB F1	ACTTGTAAATGGATCCGTCTCATTTTTACCACCTCA
Fsmut-msaB R1	TCCTTTTTCAGCGTTAAACCATTTAAACTGTACCTTGTTTCAT
Fsmut-msaB F2	ATGAAACAAGGTACAGTTTAAATGGTTTAACGCTGAAAAAGGA
Fsmut-msaB R2	AGTTATAAAGCGTACATCGTTAAGACAACTCATTA
For MsaB protein expression
UpA	AGATGGATCCTTGAACGCGTCCGAATACTTGTAA
UpB	TTAATGGTGATGATGGTGATGTAGTTTAACAACGTTTGCAGCTTGT
DnC	TACATCACCATCATCACCATTAATTCTTAGATTTGAATCATTGATTTTA
DnD	AGATGAATTCGAAGTTATAAAGCGACAATCGTTAAGACAACT
For complement and overexpression constructs
ohr-Comp-F	AGTCTGGATCCGTTTCTTCGATAGATAATTGTGGC
ohr-Comp-R	ACGTTGAATTCTCTCTTTAAGTTAATGCTAATCTA
msaABCR::POhr-F	ATGTATTTTAAATTAGGATCCTATAAGTATGGCA
msaABCR::POhr-R	ACGTTGAATTCTCTCTTTAAGTTAATGCTAATCTA
For ChIP and EMSAb
crtOPQMN-ChIP-F	CTAATGGTTATGCATCAGGAAGTAAC
crtOPQMN-ChIP-R	CTAAATTGAATCACTCTCAATCATACTGAC
crtOPQMN-EMSA-F	Biotin-CTAATGGTTATGCATCAGGAAGTAAC
crtOPQMN-EMSA-R	Biotin-CTAAATTGAATCACTCTCAATCATACTGAC
ohr-ChIP-F	GAATCCCTCATTTCAGATACTC
ohr-ChIP-R	GCCATACTTATAACCTCCTAATT
ohr-EMSA probe	Biotin-duplex-TTTTCGAATGGGTAAAGCATAAATGTATTTTAAATTAGGAGGTTATAAGT
sarZ-ChIP-F	AACAATTAAGGATGATACAATTACA
sarZ-ChIP-R	CAATCACTCCTTGTTAAAATAAA

^aRestriction sites are underlined.

^bChIP, chromatin immunoprecipitation; EMSA, electrophoretic mobility shift assay.

DISCUSSION

In the versatile pathogen S. aureus, oxidative stress acts as a trigger for the expression of a multitude of responses (51, 52) that have evolved as complex regulatory systems in the pathogen to combat deleterious effects of environmental stimuli (13, 53). We characterized the msaABCR operon, which regulates virulence, antibiotic resistance, and in vitro persister cell formation in S. aureus (34–36, 44). In this study, we demonstrate the importance of the msaABCR operon in resistance against oxidative stress via direct regulation of the crtOPQMN operon and the ohr gene. In addition to this, we also show msaABCR (MsaB) as a transcriptional repressor of sarZ (repressor of ohr). The regulatory relationship between several regulators of the oxidative stress defense system is depicted in Fig. 12.

FIG 12.

Regulatory map of msaABCR (MsaB), sigB, and sarZ in the oxidative stress defense system. MsaB positively activates transcription of the STX biosynthetic operon crtOPQMN and ohr gene. In addition, MsaB also activates the expression of sigB, which is a positive transcriptional regulator of crtOPQMN expression, and the regulation of the two genes appears to be mutually epistatic. On the other hand, MsaB represses sarZ, which in turn also represses the ohr gene. The oxidative sensing pathway by MsaB and the coordinated expression of genes remain to be elucidated. At this time, although we do not know whether SigB and SarZ directly regulate MsaB, the increased transcription of msaB in the sarZ and sigB mutants indicates the possibility of interplay between these regulators and MsaB.

Transcription of the crtOPQMN operon is positively regulated by the RsbUVW-sigmaB system (39, 54–57) and MsaB (39) and is negatively regulated by small stable RNA A (ssrA) (58). In addition, the production of STX is also regulated by metabolic activity (59). Recently, a two-component system, AirSR, was shown to transcriptomically regulate the crtOPQMN operon in the S. aureus strain WCUH29 (45). The carotenoid pigment STX is known to quench toxic singlet oxygen due to the presence of numerous conjugated double bonds (60–62). Studies have shown that mutants deficient in carotenoid production have increased susceptibility to in vitro oxidative stress, are more readily cleared by host immune cells (12, 53, 63, 64), and consequently demonstrate reduced virulence and survival in a murine model (64–66). Previous studies showed that inactivation of msaB led to downregulation of crtM and reduced production of STX, resulting in increased susceptibility toward H₂O₂ under ex vivo conditions (40, 64). Katzif et al. (39) demonstrated reduced crtM transcription in the msaB mutant. Although the putative DNA-binding sites for MsaB were predicted upstream of the promoter of the crtMN transcript (39), studies remained inconclusive whether MsaB mediates direct regulation of pigment production (38–40, 54). In this study, we found that MsaB indeed directly binds the crtOPQMN operon. A shift of the biotinylated crtOPQMN promoter-3′-MsaB_His complex, which was reversed by adding a 50-fold excess of specific but unlabeled DNA in EMSA, confirmed that MsaB directly regulates transcription of the crtOPQMN operon. The RsbUVW-sigmaB system also positively regulates transcription of the crtOPQMN operon (39, 54–57). Importantly, in the sigB mutant, the binding of MsaB to the crtOPQMN promoter was not regulatory, since the binding by MsaB does not appear to increase STX production in the absence of sigB. These observations indicate that transcriptional regulation of the crtOPQMN operon by the two regulators MsaB and SigB is rather complex, and we further plan to study their interaction. Moreover, a recent study by Caballero et al. (40) showed MsaB binding to rsbVWsigB mRNA as a RNA chaperone, indicating the role of MsaB in sigB regulation (40).

However, consistent with the previous study, we observed a lower level of STX production in the sigB mutant than in the msaABCR mutant (39). Interestingly, in agreement with a previous report (54), the sigB mutant demonstrated no sensitivity toward H₂O₂ compared with that of the isogenic wild-type strain. These observations indicate that although SigB and MsaB are both required for STX production, the resistance to oxidative stress that is mediated through pigmentation is dependent on msaABCR operon regulation. Liu et al. (58) showed the involvement of ssrA RNA as a negative regulator for STX production via base pairing with the ribosomal binding site of crtM mRNA in S. aureus (58). In msaABCR deletion mutants, transcripts of ssrA are upregulated. However, it is unclear whether or how this effect is linked to crtOPQMN regulation by MsaB.

We observed downregulation of katA and ahpC transcripts in msaABCR mutants in both the USA300 LAC and UAMS-1 strains. Of note, the katA and ahpC genes are required for survival, nasal colonization, and persistence of S. aureus via peroxide resistance (15). Reduced transcription of katA and ahpC in the msaABCR mutant indicates the possibility that the msaABCR mutant could also be less persistent under in vivo conditions. This speculation is corroborated by our findings showing that msaABCR mutants are highly susceptible to clearance by host immune cells.

OsmC and Ohr proteins reduce alkyl and organic hydroperoxides in different bacterial pathogens (67–72). Inactivation of these genes also results in increased susceptibility toward intracellular killing by macrophages, which clear pathogens mainly by ROS-mediated killing (11, 73). The inactivation of these proteins in bacterial species was shown to be associated with increased susceptibility toward organic hydroperoxides, with little to no effect on H₂O₂ susceptibility (50, 74–77). Previous studies have shown that the expression of Ohr is regulated by the organic hydroperoxide stress resistance regulator (OhrR) in different bacterial species (48, 49). Furthermore, Ohr was also shown to be involved in increased survival inside macrophages (48). In S. aureus, MgrA and SarZ have been considered paralogs of OhrR, functioning as a transcriptional regulator of Ohr (22, 52), which is mainly involved in sensing and resisting organic peroxide stresses. Chen et al. (22) identified SarZ as a functional homologue of MgrA, which acts as a global regulator for the expression of ∼87 genes in S. aureus. The thiol-based oxidative sensing pathway (via Cys13) in SarZ was demonstrated to play a global regulatory role over metabolic pathways, antibiotic and oxidative stress resistance, and virulence in S. aureus. The sarZ mutant was shown to have higher resistance activity against CHP, with a lesser impact toward H₂O₂. In addition, the authors found SarZ to be a repressor of Ohr, which was shown to be involved in resisting peroxide stress in S. aureus strain Newman (22). A recent study by Caballero et al. (40) showed MsaB stabilizes sarZ mRNA (40). SarZ represses not only ohr transcription but also msaABCR. Conversely, msaABCR activates both ohr and sarZ transcription. Recently, MsaB was shown to bind mgrA mRNA as an RNA chaperone (40), suggesting its involvement in the regulation of mgrA. Of note, MgrA transcriptionally activates sarZ (78). All these results indicate the complex regulatory interplay among these regulators in sensing and responding to oxidative stress. Consistent with previous studies (50, 74–77), we found that inactivation of ohr resulted in hypersusceptibility toward both of the organic hydroperoxides tBOOH and CHP, with little to no sensitivity toward H₂O₂. Furthermore, overexpression of the ohr gene in the msaABCR mutant increased resistance against tBOOH and CHP, suggesting that MsaB responds to organic hydroperoxide stress that is likely to depend on the ohr gene. However, we need further investigation into how MsaB senses peroxide stress and coordinates the expression of genes in response to oxidative stress stimuli.

Oxidative stress triggers a multitude of responses in bacterial pathogens. While these protective strategies are crucial for bacterial survival and persistence in response to various environmental stimuli, the inability to cope with these stresses leads to the rapid clearance of pathogens by host immune cells. Likewise, antibiotic-generated ROS have been discovered to be crucial molecules that enhance the bactericidal activity of drugs against antibiotic-tolerant persister cells (79), whereas tolerance to antibiotics has been suggested as depending on the cell’s protective strategies against lethal ROS (80, 81). Of note, msaABCR mutants have reduced ability to tolerate the stress of several antibiotics (36). All these observations show that the msaABCR operon is highly important to S. aureus, contributing to its ability to survive in diverse environments, an ability that facilitates the pathogen in causing persistent infections.

MATERIALS AND METHODS

Bacterial strains and culture conditions.

S. aureus strains USA300 LAC and UAMS-1, their isogenic msaABCR operon mutants (msaABCR), complemented strains (complementation), and their isogenic transposon mutants were used (Table 4). Cells were grown in tryptic soy broth (TSB) unless otherwise stated. The DH5α strain of Escherichia coli and the restriction-deficient laboratory strain of S. aureus, RN4220, were used to move plasmid constructs into the strains of choice through transformation and phage transduction, respectively, as described elsewhere (35, 44, 82). When required, either erythromycin (10 μg ml⁻¹) or chloramphenicol (10 μg ml⁻¹) was added to the medium for selection during the plasmid construction and transformation processes. The E. coli strain DH5α was grown in Luria-Bertani (LB) medium, with ampicillin (100 μg ml⁻¹) used for selection when required.

TABLE 4.

Strains and plasmids used in this study

Strain or plasmid	Relevant feature(s)a	References or source
Strains
E. coli DH5α	F− ϕ80lacZΔM15 recA1	Life Technologies
S. aureus
RN4220	Restriction-deficient laboratory strain	NARSAb
USA300 LAC	Methicillin resistant
UAMS-1	Methicillin sensitive
msaABCR	msaABCR operon knockout in USA300 LAC and UAMS-1 strains	32, 33, 44
Complementation	1.7-kb PCR fragment containing msaABCR operon cloned into pCN34 (Cmr)	32, 33, 44
FSmut-msaB	Frameshift mutation on msaB transduced into msaABCR mutant	This study
ohr-comp	ohr complementation	This study
msaABCR::POhr	Pohr::pCN51 transduced into msaABCR mutant	This study
USA300 JE2	Transposon mutants in JE2 strain	NARSA
ohr::Tn	ohr transposon mutant in USA300 LAC and UAMS-1	This study
sodM::Tn	sodM transposon mutant in USA300 LAC and UAMS-1	This study
katA::Tn	katA transposon mutant in USA300 LAC	This study
crtM::Tn	crtM transposon mutant in USA300 LAC	This study
sigB::Tn	sigB transposon mutant in USA300 LAC	This study
Plasmids
pCN34	pCN34 antibiotic marker Ermr	32, 44
pCN34 (Cmr)	pCN34 antibiotic marker Ermr changed to Cmr	32, 35
pCN51	Cad-inducible shuttle vector (Ampr Ermr)	NARSA

^aAmp^r, ampicillin resistant; Cm^r, chloramphenicol resistant; Emr^r, erythromycin resistant.

^bNARSA, Network on Antimicrobial Resistance in Staphylococcus aureus.

In vitro oxidative stress assays.

Frozen S. aureus cells inoculated on TSB were grown for ∼3 h to prepare precultures with an optical density at 600 nm (OD₆₀₀) of 0.05 in 5 ml TSB. The cultures were then incubated for 24 h at 37°C with continuous shaking (225 rpm). An oxidative stress assay with H₂O₂ was performed as previously described (39, 40) with slight modifications. Briefly, the cells were washed with phosphate-buffered saline (PBS) and diluted to an OD₆₀₀ of 0.1 in 1 ml PBS containing 34 mM H₂O₂ for 1 h. After incubation, catalase (2,000 U ml⁻¹) was added to each tube, which was incubated for 1 min at room temperature to decompose the remaining H₂O₂. The cells were diluted and plated to obtain the CFU counts. The survival percentage represents the percentage of initial CFU that survived the H₂O₂ treatment after 1 h. Similarly, the cells prepared as described above were stressed with 25 mM H₂O₂ in 5 ml of TSB, and growth was monitored by CFU counts at designated time points. A detection limit of 100 CFU/ml was established for the results that showed ≤10²/ml in undiluted culture treated with H₂O₂. For the paraquat assay, the cells were grown for 24 h, washed, and normalized to an OD₆₀₀ of 0.1 in 1 ml of PBS. Aliquots of 100 μl of bacterial suspension were dispensed into 96-well plates supplemented with and without 40 mM paraquat and incubated at 37°C with continuous shaking. Growth was monitored by taking an OD₆₀₀ measurement at designated time points. For organic hydroperoxide assays, cells were prepared as described above, washed with PBS, and diluted to an OD₆₀₀ of 6.0 for all strains tested. The cells were then serially diluted (up to 10⁻⁸), and 10 μl of each dilution was pipetted onto tryptic soy agar (TSA) plates containing 5 mM tert-butyl hydrogen peroxide (tBOOH) and 4 mM cumene hydrogen peroxide (CHP) to obtain the CFU counts after 24 h of incubation.

Hydrogen peroxide detoxification assay.

S. aureus cells grown for 16 h as described above were washed with PBS, diluted to ∼2 × 10⁵ CFU in 5 ml of TSB containing a 100 μM final concentration of H₂O₂, and incubated for 1 h at 37°C with shaking. Every 15 min, 1 ml of culture was withdrawn and centrifuged, and the supernatant was filtered to remove any bacteria. The amount of H₂O₂ in the supernatant was measured using an H₂O₂ detoxification kit (Cell Biolabs). The amount of H₂O₂ degraded at designated time points was represented graphically.

NADP⁺/NADPH measurement.

Quantification of NADP⁺/NADPH was conducted using an NADPH/NADP kit from Biovision Inc., as described elsewhere (41). Briefly, overnight cultures were diluted to an OD₆₀₀ of 0.05 in 5 ml of TSB and grown for 4 h at 37°C. After incubating, the cells were pelleted by centrifuging at 5,000 × g at 4°C, washed with cold sterile PBS, resuspended in 400 μl of NADP⁺/NADPH extraction buffer (Biovision Inc.) within 10 min, and lysed with 0.1 mM silica beads in a bead beater (MP Biomedical). The sample obtained after centrifuging lysed samples at 16,000 × g for 10 min at 4°C was treated with protein denaturation reagents (Amicon) to remove enzymes that utilize NADPH. Next, prepared samples (50 μl) were transferred in triplicates to a 96-well plate, 100 μl of NADP cycling mix was added, and the samples were incubated in the dark at room temperature for 5 min. Next, 10 μl of NADPH developer was added to each of the wells, which were incubated at room temperature for 2 h. The OD₄₅₀ was taken every hour to measure color development, and the data represent the values measured at 1 h. The relative protein concentrations were calculated for each strain using a bicinchoninic acid (BCA) protein assay kit (Pierce).

Total antioxidant capacity.

Total antioxidant capacity was measured using the OxiSelect total antioxidant capacity assay kit (Cell Biolabs) as previously described (41). Briefly, precultures prepared as described above were further grown to an OD₆₀₀ of 0.5 to 0.6, and cells were harvested by pelleting. The cells were then washed twice, resuspended in 500 μl of ice-cold PBS, kept frozen at −80°C, thawed twice, and then mechanically disrupted using a bead beater. The protein was then harvested by centrifugation at 5,000 × g for 10 min at 4°C. The protein concentrations were determined using a BCA protein assay kit (Pierce), and the remaining portions were assayed for total antioxidant capacity.

In vitro whole-blood and neutrophil survival assay.

The survival of S. aureus in whole human blood was measured in vitro as described previously (32, 83). In brief, heparinized venous blood samples were collected from healthy donors according to a protocol approved by the Institutional Review Board for Human Subjects at the University of Southern Mississippi. S. aureus cells were harvested in the postexponential phase of growth. The blood samples (3 ml) were inoculated with 3 × 10⁵ S. aureus cells in a 14 -ml culture tube and incubated at 37°C for 2 h with end-over-end rotation at 20 rpm. The samples were then diluted with sterile deionized water to lyse the blood cells, and the number of CFU was determined after plating on TSA plates. The neutrophil survival of S. aureus was measured using freshly isolated human polymorphic neutrophils (PMNs) as previously described (7), with minor modifications (32). In brief, PMNs (1 × 10⁶) were combined with S. aureus (1 × 10⁷) in a 24-well tissue culture plate and centrifuged at 380 × g for 8 min. The CFUs were then incubated for 2 h at 37°C with 5 % CO₂. After incubating, the neutrophils were lysed using deionized water, plated on TSA plates, and incubated overnight at 37°C. The CFU was counted, and the test samples were compared with controls, which contained deionized water instead of the neutrophil suspension. This assay measures the total number of viable bacteria.

Macrophage survival assay.

Macrophages (RAW 264.7) were prepared by growing in Cyto 6-well plates in sterile RPMI 1640 medium at 37°C supplemented with CO₂. Stationary-phase S. aureus cells (24 h) prepared as described above were harvested, washed, and diluted in PBS. The bacterial cells (2 × 10⁵ cells) with a multiplicity of infection (MOI) of 10 were incubated with macrophages at 37°C supplemented with CO₂. At a designated time point, gentamicin 200 μg/ml was added to each well for 20 min to kill extracellular bacterial cells. The content of each well was gently removed and washed 3 times with PBS, and finally, 1 ml ice-cold deionized water was added to rupture the macrophages. The released bacterial cells were serially diluted and plated on Columbia agar to determine the intracellularly surviving CFU counts after incubation at 37°C for 24 h.

Extraction of staphyloxanthin.

Staphyloxanthin (STX) was extracted using the methanol extraction method as described previously (44). Briefly, cells grown overnight were normalized to an OD₆₀₀ of 0.05 in 10 ml of TSB and cultured for 16 h at 37°C with continuous shaking. Cells were harvested, washed twice with deionized water, and pelleted after normalizing all the strains to an OD₆₀₀ of 10.0 in 1 ml. Then, the cells were mixed with 1 ml of methanol and incubated in a 58°C water bath for 10 min with occasional vortexing. The supernatant was harvested by centrifugation at 16,000 × g for 1 min, and the absorbance of STX was measured at OD₄₆₅, with methanol as a blank.

RNA extraction, reverse transcription, and qRT-PCR.

The expression of genes was measured with quantitative real-time PCR (qRT-PCR) in the wild type and isogenic mutants. Briefly, 5 ml of a 16-h culture was prepared as described above with continuous shaking at 37°C. Aliquots of 500-μl cultures were treated with equal volumes of RNAprotect bacterial reagent (Qiagen, Valencia, CA) in a 1.5-ml tube for 5 min at room temperature. The cells were pelleted by centrifugation at 10,000 × g for 5 min and stored at −80°C until analysis. The samples were thawed on ice, and total RNA was extracted as previously described using the Qiagen RNeasy kit (32, 33, 35, 44). RNA concentration and quality were analyzed by measuring absorbance at 260/280 nm using a NanoDrop spectrophotometer (Thermo Scientific). Reverse transcription was performed with the iScript cDNA synthesis kit (Bio-Rad Laboratories, Hercules, CA), using 1 μg of the total RNA isolated according to the manufacturer’s instruction. qRT-PCR was performed using the primers in Table 5, and the relative fold change in gene expression was calculated using gyrA and gyrB as endogenous control genes. The results were consistent between the control genes. The data represent the results from three independent experiments analyzed using gyrB as an internal control.

Generation of transposon mutants and complementation.

The allelic replacement method was used to generate msaABCR deletions in the strains USA300 LAC and UAMS-1, as previously described (32, 44). For trans complementation, the msaABCR region was cloned into the pCN34 low-copy-number vector with the modification of changing the kanamycin selectable marker to a chloramphenicol resistance marker, as described elsewhere (35, 84). The mutants of other genes in this study (Table 4) were generated by the insertion of a transposon into their open reading frame (ORF), as described previously (32, 33). Briefly, strain JE2 was obtained from the Network on Antimicrobial Resistance in S. aureus (NARSA) collection (Bei Resources). This strain contains the Bursa aurealis mariner-based erythromycin resistance expression transposon within the coding region of the respective genes. The mutation was moved by generalized transduction using bacteriophage ϕ11 (32, 33, 35, 85). Introduction of the transposon mutation into the recipient strains was verified by PCR, followed by sequencing. The transposon mutants and primers used in this study are listed in Tables 4 and 5. For trans complementation, the corresponding gene’s open reading frame (ORF) was cloned into the pCN34 low-copy-number vector with the modification of changing the kanamycin selectable marker to a chloramphenicol resistance marker as described previously (32, 35).

Construction of FSmut-msaB.

We used an overlap extension PCR cloning technique to generate a frameshift mutation in msaB ORF as previously described by Bryksin et al. (86). The upper 706-bp fragment of msaABCR operon was PCR amplified by using primer set Fsmut-msaB F1 and Fsmut-msaB R1, and the lower 1,090-bp fragment of msaABCR operon region was amplified using primer set Fsmut-msaB F2 and Fsmut-msaB R2. Primers Fsmut-msaB R1 and Fsmut-msaB F2 overlap, such that an insertion of one nucleotide causes a frameshift mutation in msaB. Both the PCR fragments were PCR purified using a Promega DNA cleanup kit, and then 50 ng of each fragment was used in the PCR ligation, which contained all the ingredients of the PCR mix except the terminal primers. The normal PCR cycle was carried out for 15 cycles, and finally, the terminal primers (Fsmut-msaB F1 and Fsmut-msaB R2) were added to the reaction mixture and 20 additional cycles were performed. The final amplified PCR product was ligated with the pCN34 low-copy-number plasmid vector and transduced into the msaABCR operon deletion mutant. The FSmut-msaB construct was verified by sequencing.

Expression and purification of MsaB.

The MsaB protein was expressed in msaABCR mutant of USA300 LAC strain. The primers UpA, UpB, DnC, and DnD listed in Table 5 were used to label MsaB protein with 6×His at the C-terminal end (MsaB-His). To generate the expression construct, the ORF of msaB was PCR amplified using primers listed in Table 4 and cloned into the inducible plasmid pCN51 using the BamHI and EcoRI restriction enzymes. The plasmid construct was then transformed into E. coli strain DH5α. The plasmid isolated from transformed DH5α cells was used to transduce the restriction-less S. aureus RN4220 competent cells. The plasmid was then moved to USA300 LAC msaABCR mutant through generalized transduction using bacteriophage ϕ11, as described previously (35, 44, 82). Protein expression was induced by adding 10 μM cadmium chloride (CdCl₂) at exponential phase and incubating further with shaking. The cells were pelleted after 4 h of induction, resuspended in PBS (pH 7.4) with a protease inhibitor cocktail, and then lysed by bead beating followed by sonication. The cell lysate was centrifuged at 10,000 × g for 30 min to remove the cell debris. The 6×His-MsaB fusion protein was purified from the clear lysate with a nickel column (HisPur Ni-nitrilotriacetic acid [Ni-NTA] resin; Thermo Scientific).

Overexpression of the ohr gene.

Escherichia coli strain DH5α and the restriction-deficient S. aureus laboratory strain RN4220 were used to move plasmid constructs into the strains of choice through transformation and phage transduction as described previously. To generate the overexpression construct, the ohr gene was PCR amplified using primers listed in Table 4 and cloned into the inducible plasmid pCN51 using the BamHI and EcoRI restriction enzymes. The plasmid construct was then transformed into Escherichia coli strain DH5α. The plasmid was then isolated from transformed DH5α cells and used to transduce the restriction-less Staphylococcus aureus RN4220 competent cells. The plasmid was moved to the msaABCR mutants (msaABCR::pOhr) through generalized transduction using bacteriophage ϕ11, as described previously (35, 44, 82). The overexpression of the gene was analyzed with qRT-PCR by adding CdCl₂ (10 nM to 1 μM) in the exponential growth phase. Induction with the 100 nM CdCl₂ concentration overexpressed the gene by 5- to 10-fold; below and above this concentration, the expression level was lowered. Therefore, we chose this concentration for the induction. When preparing msaABCR::pOhr cells for the oxidative stress assay, cells were induced by adding 100 nM CdCl₂ in the early exponential (2 h) and late and postexponential growth phases (4 h and 12 h) of culture.

Chromatic immunoprecipitation assay.

The ChIP assay was performed as described by Sengupta et al. (87), with slight modifications adopted in our lab (32). Briefly, S. aureus cells in stationary growth phase were treated with 1% formaldehyde and 10 mM sodium phosphate to facilitate the cross-linking of MsaB to its targets. After 20 min, the cross-linking reaction was quenched by the addition of 0.1 volumes of 3 M glycine. The cultures were then pelleted and washed with equal volumes of 0.1 M phosphate buffer to remove excess formaldehyde. The phosphate buffer was removed by centrifugation, and the cells were resuspended in 600 μl IP buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride [PMSF], 5% [vol/vol] glycerol, 1% Triton X-100) and lysed with a bead beater. The 400-μl cell lysates were then sheared by sonication to obtain fragment sizes of ∼500 bp. The clear supernatant obtained after brief centrifugation was diluted with 1 ml IP buffer, an anti-MsaB antibody (diluted 1:1,000) was added, and the mixture was incubated at room temperature for 2 h with gentle shaking. This antigen-antibody mixture was then added to prewashed protein G-coupled magnetic beads (Thermo Scientific) and incubated further for 1 h as described above. The antigen-antibody-bead complex was collected with a magnetic tube holder, washed five times with wash buffer (Tris-buffered saline, 0.05% Tween 20, 0.5 M NaCl), followed by washing once with ultrapure water. The beads were then resuspended in 100 μl elution buffer (0.1 mM glycine, pH 2.0) and neutralized with neutralization buffer (Tris-EDTA, pH 8.0), followed by an incubation at 65°C overnight to reverse the cross-links (decoupling). The DNA extracted with the phenol-chloroform extraction method was used as the template to detect the MsaB-bound promoter sequences with PCR amplification using promoter-specific primers. The msaABCR mutant was used as an internal negative control to demonstrate that the MsaB antibody is specifically enriched in the promoter regions.

Electrophoretic mobility shift assay.

To determine the binding activity of MsaB on the crtOPQMN operon promoter, we used amplified DNA from the region upstream of the crtOPQMN operon, as described previously (45). Briefly, the 5′-biotinylated Pcrt-EMSA forward/reverse primers were used to amplify the promoter region. A total of 250 μl of the PCR product was loaded into a 2% agarose gel. The DNA band of the corresponding Pcrt fragment size was excised and cleaned using the Wizard SV gel and PCR cleanup system (Promega). The purified product was analyzed for DNA concentration and quality using a NanoDrop spectrophotometer (Thermo Scientific). The purified probe was again electrophoresed to verify the size and the purity of the labeled Pcrt probe. To determine the binding activity of MsaB on the ohr promoter, we synthesized 5′-biotinylated duplex fragments of the same oligonucleotide sequence in the promoter region of ohr, which was published in a previous study (22). Electrophoretic mobility shift assays were performed using the LightShift Chemiluminescent EMSA kit (Pierce), according to the manufacturer’s protocol. Briefly, the binding reaction mixture was prepared with ultrapure water, 1× binding buffer, 50 ng μl^− 1 poly(dI-dC), 2.5% (vol/vol) glycerol, 0.05 % NP-40, 5 mM MgCl₂, and the purified 5′-biotin-labeled DNA probe. Increasing concentrations of MsaB-His protein and unlabeled specific probe, when required, were added. Various amounts of RNase/DNase-free water were added to the tubes to adjust the reaction volume to 20 μl. The reaction mixture was prepared by adding the individual components in the order instructed by the manufacturer. The mix was then incubated at room temperature for 20 min and subjected to electrophoresis at 100 V for 1 h in a prerun 5% Tris-borate-EDTA (TBE) gel. The samples were then transferred to a nylon membrane (1 h in the cold), cross-linked, and processed for the detection of samples. The protein-DNA complexes in the gel were then visualized using the detection module supplied with the kit, according to the manufacturer’s protocol, and imaged with the ChemiDoc system (Bio-Rad).