Strains and plasmids used in this study are listed in Table 1. GAS was grown under static conditions at 37°C in Todd–Hewitt broth with 0.2% (wt/vol) yeast extract (THY broth, Difco Laboratories) or in C-medium consisting of 0.5% Proteose Peptone #3 (BD), 1.5% yeast extract (BD), 10 mM K₂HPO₄, 0.4 mM MgSO₄, 17 mM NaCl, adjusted to pH 7.5 (Lyon et al., 1998). In some experiments, hemin chloride (Sigma) from stock solutions prepared in DMSO was added to the growth media at different concentrations. E. coli cells were used for cloning and protein expression purposes. E. coli was grown aerobically in a Luria–Bertani medium (pH 7.0) supplemented with kanamycin (30 μg/ml) at 37°C.

GAS culture samples with equal cell numbers were harvested 30, 60, and 90 min following treatment with 4 μM heme (in 0.035% DMSO) or with mock (0.035% DMSO, control) and washed twice with phosphate buffer saline, pH 7.4 (PBS). The resulting cell pellets were resuspended in 5 ml of PBS with lysozyme (1 mg/ml, Sigma) and 400 U of mutanolysin (sigma) and incubated at 37°C for 30 min. The cells were then subjected to sonication (5 s, 10% amplitude). TBARS formation in the membrane samples was quantified using the TBARS assay kit (ZeptoMetrix Corporation) according to the manufacture's recommendations. Briefly, 100 μl cell lysate samples were treated with 100 μl of SDS and 2.5 ml of the TBA reagent and incubated at 95°C for 60 min. The sample supernatant was collected following 10 min incubation on ice by centrifugation (15 min at 835 × g). The absorbance at 532 nm (indicative of TBARS) was measured using the DU 730 Life Science UV/Vis spectrophotometer. The amount of TBARS in the experimental samples was derived from a standard curve generated using the malondialdehyde (MDA) reagent supplied with the kit.

The TBARS assay was followed as above except the cell pellets were washed twice with TSM buffer (100 mM Tris, 500 mM sucrose, 10 mM MgCl₂, pH 7.0), resuspended in 0.5 ml of TSM with 400 U of Mutanolysin (Sigma) and incubated at 37°C for 30 min. The protoplasts were centrifuged at 20,000 × g for 5 min at 4°C, suspended in 0.2 ml of lysis buffer (50 mM Tris-HCl, 60 mM KCl, 10 mM MgCl₂, pH 7.0, 2% β-mercaptoethanol) and subjected to sonication (10 s, 10% amplitude). The membrane components were collected by centrifugation at 100,000 × g for 30 min at 4°C. The supernatant was retained as a cytoplasmic fraction and pellets were resuspended in 0.2 ml of lysis buffer and were considered as the membrane fractions. Protein damage was detected using the OxyBlot™ protein oxidation detection kit (Millipore) according to the manufacturer's instructions. In this assay, oxidized proteins are derivatized with 2,4-dinitrophenylhydrazine (DNP), which is then detected with primary antibodies specific for DNP and secondary antibody conjugated to horseradish peroxidase (HRP) using a chemiluminescence protocol. The intensity of the signal in individual lanes was quantified using ImageQuant LAS 4000 and the ImageQuant TL software (GE).

Cell samples were harvested 30, 60, and 90 min following heme or mock treatment by centrifugation (5000 × g for 20 min) at 4°C. RNA was extracted from the cell pellet using the RiboPure™ RNA purification kit (Ambion) followed by DNaseI digestion (Ambion) performed according to manufacturer's instructions. Microarray analysis was performed with total RNA extracted from 90 min post treatment cell samples. Real-time RT-PCR transcript analysis for selected genes was performed using total RNA extracted from all three-time points post treatments.

The GAS microarray used in this study (Ribardo and McIver, 2006) consists of 2328 70-mer oligonucleotide probes targeting unique non-repetitive ORFs from the sequenced genomes of serotypes M1 (SF370), M3 (MGAS315), and M18 (MGAS8232). Probes were synthesized by Qiagen Operon with a melting temperature of 76 ± 5°C, ≤70% cross-hybridization identity to another gene within the same strain, ≤20 contiguous bases in common with another gene, and probe location within 3′ end of ORF. Microarrays were printed at Microarrays Inc. (http://www.microarrays.com), with 10 pl of each oligonucleotide probe spotted onto slides (UltraGAPS2; Corning) using a 12-pin contact printer. The microarray hybridization was performed as previously described (Jiang et al., 2008). Briefly, 10 μg of DNase I-treated total RNAs to be compared were used for reverse transcription into single-stranded cDNA using 200 U Superscript II reverse transcriptase (Life Technologies), 6 μg random hexamers, 1X first strand buffer, 10 mM dithiothreitol (DTT), 0.5 mM dATP, 0.5 mM dCTP, 0.5 mM dGTP, 0.3 mM dTTP, and 0.2 mM of amino-allyl dUTP. The mixture was incubated at 42°C for 2 h and the reaction stopped by addition of 10 μl 0.5 M EDTA and 1 M NaOH. Amine-modified cDNA was purified by ethanol precipitation followed by chemical labeling with Cy3- or Cy5-NHS-ester fluorescent dyes (GE Healthcare) in a final step. Yield and incorporation of dye was determined using a Nanodrop ND-1000 (Thermo Scientific). Slides were pre-hybridized in a 50 ml solution of 5X SSC, 0.1% SDS and 1% BSA for 30 min at 42°C, washed 4X in water and once in isopropanol, then dried by brief centrifugation. Labeled probes were resuspended in hybridization buffer (30% formamide, 5X SSC, 0.1% SDS, 0.6 μg/μL salmon sperm DNA) and hybridized to the microarray slides in a 42°C water bath for 16–20 h. Slides were washed twice in a low stringency buffer (2X SSC, 0.1% SDS) at 55°C for 5 min, twice in a medium stringency buffer (0.1X SSC, 0.1% SDS) at room temperature for 5 min and finally twice in a high stringency buffer (0.1X SSC) at room temperature for 5 min, and then dried by brief centrifugation. Synthesized cDNA from each RNA sample from three independent cell cultures was hybridized on three separate microarray slides (biological replicates), and independently synthesized cDNA from each of these same RNA samples was hybridized in a repeat dye-swap experiment (technical replicates) to test technical reproducibility. Slides were scanned using an Axon 4100A personal array scanner and GenePixPro 6.0 software (Molecular Devices). Data obtained from MGAS5005 cells incubated in the presence of DMSO or 4 μM of heme were compared for twofold changes in expression, ≥2.0 or ≤0.50 with Acuity 4.0 software (Molecular Devices). Using a ratio-based normalization, data were normalized by the ratio of the means (635/532) and samples were removed when four out of the six experiments did not show significance. Data was validated on 11 independent genes by real-time RT-PCR as described below. Correlation coefficients for the arrays were determined by plotting the log value of the array on the x-axis to the log value of the real-time RT-PCR on the y-axis. An equation determining the line of best fit was determined, and the resulting R² value was calculated to be 0.889. Array data was submitted to the Gene Expression Omnibus (GEO) at the National Center for Biotechnology Information under the accession number GSE61415.

For microarray data validation, real-time RT-PCR analysis was carried out using primers in Table 2 as follows: 25 ng of DNaseI-treated total RNAs were added to SYBR green master mix (AB) containing 200 nM of each specific real-time primer for the one-step protocol. The real-time RT-PCR experiments were completed using a LightCycler 480 instrument (Roche), and levels represent the ratio of non-treated to treated experimental values relative to the level of expression of gyrA transcript as an endogenous control. For expression of the pefRC operon, a SYBR Green based quantitative PCR reactions were performed using the Power SYBR® Green RNA-to-Ct™ 1-Step Kit (AB) on 7500 Fast Real-Time PCR machine (AB) according to the manufacturer's specifications. Briefly, the reaction mixture (20 μl) contained, 10 μl of RT-PCR Mix (final 1X), 200 nM of each forward and reverse primers, 25 ng of total RNA, and RT enzyme Mix in 1X final concentrations. The relative quantification with comparative ΔΔCT method was employed to calculate differences in pefRC expression levels at different times after heme exposure. The relative expression levels of pefR and pefC genes were normalized to the level of rpsL transcript as an endogenous control. The levels of transcripts in heme treated samples were compared to the control samples.

The PefR binding motif was identified within the putative promoter region of MGAS5005 spy_0195 (pefR) using Multiple Em for Motif Elicitation (MEME) suit hosted by the National Biomedical Computation Resource (Bailey et al., 2006). The outcome was further analyzed by the MAST application of MEME for its genome wide occurrence within the upstream sequences of GAS under both the stringent (E-value ≤ 0.01) and relaxed (E-value ≤ 10) parameters. All of the sequences were acquired from Kyoto Encyclopedia of Genes and Genomes (KEGG) database (Kanehisa, 2002) for comparative sequence analysis; sequences were aligned using ClustalW (Thompson et al., 2002).

The spy_0195 ORF was amplified from GAS MGAS5005 genomic DNA by PCR using primers ZE515 and ZE516 (Table 2). To generate pAJS11 plasmid (Table 1), the PCR fragment with pefR ORF was cloned into the pRham™ expression vector (Lucigen) by Expressioneering™ technology and introduced into E. cloni® 10G competent cells. The cloning was confirmed by sequence analysis. For PefR expression, cells harboring pAJS11 were induced at OD_{600 nm} = 0.8 with 0.2% rhamnose. The cells were harvested (8000 × g for 5 min at 4°C) following 16 h incubation at 28°C. The resulting pellets were resuspended in extraction buffer (20 mM Tris pH 8, 100 mM NaCl, 0.1% Triton X-100) containing 0.5 mg/mL Complete, mini-EDTA-free protease inhibitor cocktail (Roche), sonicated and the cell debris were removed by centrifugation. The resulting lysate was purified over 5 ml HisTrap™ HP (GE) nickel affinity column. Protein fractions were dialyzed overnight in sodium phosphate buffer (SPB: 20 mM sodium phosphate, 500 mM NaCl pH 7.4). Expression of the recombinant PefR was evaluated by SDS-PAGE and western blot analysis with mouse anti-His antibodies (Sigma).

Heme binding by PefR was assessed spectroscopically as described in Puri and O'brian (2006) and Ouattara et al. (2010) with minor modifications. Briefly, an increasing concentration of hemin chloride (2–30 μM) was added to both the test cuvette containing 10 μM of PefR protein in SPB and the reference cuvette (containing SPB alone) and the changes in absorbance across the wavelength of 250–700 nm region were recorded every 6 min. The PefR to heme stoichiometry and dissociation constant (K_d) were determined by plotting the absorbance at 435 nm as a function of the heme concentrations. The extinction coefficient (ε_max) for PefR was calculated from the hemocromogen method described in Asakura et al. (1964). All of the spectroscopic measurements were made using the DU730 Life Science UV/Vis spectrophotometer (Beckman Coulter). PPIX binding was tested by titrating PefR (5 μM) with 0–6 μM PPIX (in 0.5 μM increments) prepared in acidified methanol:dimethylformamide (1:1). The changes in absorbance were recorded.

Host heme is the immediate source of iron during infection for invading pathogens such as GAS. Recent studies demonstrated that bacteria must maintain the intracellular levels of heme at equilibrium in order to benefit from its nutritional value, while eluding the toxicity that is associated with heme overload (Torres et al., 2007; Fernandez et al., 2010; Mike et al., 2014). In this work, we began evaluating the impact of heme on GAS physiology. We found that the addition of a disc saturated with 10 mM heme onto agar plates seeded with a confluent lawn of GAS resulted in a zone of clearance similar to those observed with antibiotic-impregnate discs. This observation indicates that heme can have a bacteriostatic effect on GAS (Table 3). To compare the sensitivity to heme among GAS isolates we determined the MIC values of heme using two common methods (Table 3). The agar dilution assay employs solid C-medium containing varying concentrations of heme. We determined that the heme MIC in this method is between 10 and 20 μM with the nephritogenic M49 GAS skin isolate, NZ131 (Simon and Ferretti, 1991; Mcshan et al., 2008), exhibiting the highest sensitivity of all GAS strains tested. The highly pathogenic M1T1 MGAS5005 strain (Sumby et al., 2005) as well as clinical isolates from patients with invasive diseases (Table 1) demonstrated higher heme tolerance. In the broth dilution assay, we measured bacterial growth in THYB supplemented with varying heme concentrations. This method resulted in higher values of heme MIC (30–50 μM) for MGAS5005 and the clinical isolates. However, both methods indicated that the invasive strains were more resistant to heme than NZ131.

To learn why heme is detrimental for GAS growth, we investigated its impact on GAS macromolecules. Heme is reported to damage directly or through the generation of reactive oxidative species the integrity of membrane lipids (Chang et al., 2003; Chiabrando et al., 2014). Therefore, we evaluated the extent of lipid peroxidation in GAS following heme exposure. Using low heme concentration that could still support cell growth when applied at the early logarithmic phase allowed us to examine the kinetics of damage production and later the bacterial transcriptome response to heme. As seen in Figure 1A, the addition of 4 μM heme to MGAS5005 at the early logarithmic phase of growth did not result in a significant change in the bacterial growth profile. To detect lipid peroxidation, we used an established method that relies on the tendency of oxidized lipids to react with a thiobarbituric acid (TBA) reagent to form adducts (named TBARS) that absorb at 532 nm (Hong et al., 2012). The time course of TBARS production in GAS cells exposed to heme (or in control samples) was quantified using a standard curve (Figure 1B). Significantly higher occurrence of lipid peroxidation was observed in all of the heme-treated samples compared with control samples (Figure 1C). Lipid peroxidation was the highest (22.5 fold over background) 30 min after heme treatment; while lower levels (~5 fold over background) were found in the samples collected 60 and 90 min post-heme treatment (Figure 1C), indicating possible repair mechanism. In eukaryotes, membrane repair involves enzymes such as phospholipase A2 that hydrolyse peroxidized fatty acid esters and glutathione peroxidase (van Kuijk et al., 1987; Thomas et al., 1990).

We next evaluated the effect of heme on GAS proteins. Oxidation introduces carbonyl groups into protein side chains that can then react with 2,4-dinitrophenylhydrazine (DNPH) to generate 2,4-dinitrophenylhydrazone (DNP) derivatives (Dalle-Donne et al., 2003). To monitor protein oxidation in GAS, membrane and cytoplasmic proteins were extracted from culture samples collected at 30, 60, and 90 min post treatment and allowed to react with DNPH. Oxidation was then visualized by immunoblot with antibodies specific for DNP and the damage was quantified by densitometry. Analysis of the membrane fractions revealed that the anti-DNP antibody strongly reacted with all of the samples from heme exposed cells (Figure 2A, upper panel). While general staining confirmed the presence of proteins in all samples (Figure 2A, bottom panel), protein oxidation was only observed in the heme-treated cells in the early sample. At the later time points (60 and 90 min) some background oxidation also occurred in the control samples. In all cases though, a significantly higher amount of protein damage was observed in the heme-exposed cells compared to the controls (Figure 2C). In addition, the degree of protein oxidation was the highest in the 90 min samples. Similar analysis of cytoplasmic proteins did not reveal detectable damage (Figure 2B); therefore, protein oxidation following heme exposure is primarily linked to the membrane fraction under our experimental conditions.

When overnight cultures of NZ131 grown in heme-free medium were inoculated into medium containing 1 μM heme, which is below the growth inhibiting concentrations at the early logarithmic phase, an extended lag phase that lasted for more than 8 h was observed (Figure 3). Although exposure to heme inhibited growth initially, the cultures reached after overnight incubation the cell density observed in cultures that grew in heme-free medium (data not shown). To test for possible adaptation to environmental heme, GAS cells were grown overnight in the presence of 0.1 μM heme before they were inoculated into medium with 1 μM heme. Indeed, pre-exposure to low heme level eliminated the extended lag period (Figure 3). These observations suggest that the treatment with low level of heme-triggered adaptation to environmental heme.

In light of recent evidence that bacteria can sense and respond to heme (Fernandez et al., 2010; Lechardeur et al., 2012; Mike et al., 2014; Wakeman et al., 2014) and with evidence for a similar response in GAS, we wanted to gain insight into the pathogen's mechanisms for heme sensing and tolerance. We examined the differential global gene expression pattern between GAS cells subjected to heme stress (4 μM) and control treatment. Total RNA was isolated from cells 90 min post treatment and analyzed using a 70-mer oligonucleotide microarray (representing GAS genomes M1, M3, and M18) (Ribardo and McIver, 2006). Our data showed a significant shift in the transcript profile of different genes and operons in response to heme stimulus; referred to as the “GAS heme stimulon.” This stimulon involved changes in 163 total genes, including 79 genes that were up regulated and 84 genes that were down regulated (Figure 4 and Table S1). The array analysis was validated by quantitative RT-PCR (qPCR) on 11 differentially regulated genes (Table S1), showing a strong correlation, with an R² value of 0.889 (Figure S1).

Genes that were down regulated significantly in response to heme are found in the categories of metabolic pathways, sugar and metabolite transport, and two-component system (TCS, Figure 4A). Of particular interest is the down regulation of the TCS, trxSR. The response regulator, TrxR activates the Mga virulence regulon in GAS, and a trxR mutant is attenuated for virulence in a murine infection model (Leday et al., 2008; Baruch et al., 2014). Therefore, environmental heme may impact the expression of key virulence factors in GAS via the TrxSR/Mga pathway. Since the TrxSR system is implicated in asparagine sensing, our observations also imply that the damage to the cell envelope introduced by heme is associated with increased levels of asparagine (or other amino acids) (Baruch et al., 2014). In contrast, heme-activated genes and highly expressed transcripts encode for protein folding and degradation (such as groEL/ES ~8 fold, DnaJ/K 3-4 fold, and clpE/L ~4 fold) and components of the Clp protease machinery (clpCP- 2.5 fold) (Lund, 2001; Alexopoulos et al., 2012) (Figure 4B). In addition, the GAS heme-activated stimulon includes genes whose functions are linked to regulation of redox stress management, including spxA2 (5 fold) and ctsR (2.5 fold) (Elsholz et al., 2010; Antelmann and Helmann, 2011). Accordingly, members of the downstream genes of these regulators were also activated by heme such as thioredoxin (trx ~2.5 fold) and oxidoreductases (3 fold). Finally, the expression of two TCSs was also up regulated in response to heme exposure, including the pneumococcal-like TCS homologous system, ciaHR (~2.3 fold) (Ahn et al., 2006) and the ihk/irr TCS (2.5 fold) involved in GAS response to oxidative stress and survival within phagocytes (Han et al., 2012).

The transcriptomic analysis also revealed that several putative efflux proteins also showed an increase in transcriptional activity in response to heme stress. These included a 3-gene cluster (MGAS5005 spy_0195, 0196, and 0197) with homology to the pefRCD genes from GBS (Fernandez et al., 2010). Since the pefRCD genes are important for the management of heme stress in GBS, we focused our attention on their heme-induced homologs in GAS. Comparative sequence analysis demonstrated that the putative GAS genes, spy_0195 (84% similarity; 56% identity), spy_0196 (76% similarity; 48% identity), and spy_0197 (76% similarity; 51.3% identity) show significant sequence homology to the GBS pefRCD genes, respectively (Figure 5). In addition to the high sequence homology demonstrated by each of these GAS genes, the genomic arrangement of the GBS pefRCD locus and its immediate chromosomal location are also conserved in all of the published GAS genomes (Figure 6A). Moreover, using the MEME algorithm, we identified a 17-bp inverted repeat region within the putative promoter of spy_0195 that shared 76% identity with the PefR binding site from GBS (Figure 6A). Based on this sequence conservation and the observed induction by heme, we hypothesized that like GBS, the above-mentioned GAS genes encode a heme dependent repressor (spy_0195) and an ABC heme exporter (spy_0196 and spy_0197) and named them pefRCD, respectively.

We examined the time course of pefRC expression in response to heme by qPCR (Figure 6B). As seen in the microarray analysis, the expression of pefR and C was comparable to one another in all of the time points, supporting an operon structure. Heme exposure resulted in an increase of pefR and pefC expression over time, reaching 2.5 fold over background at 30 min, 4.5 fold at 60 min and remained high at 90 min post treatment. This temporal increase in pef expression in response to heme insinuates an active participation of these genes in heme tolerance.

In GBS, PefR is a MarR-like repressor that binds to DNA and blocks the transcription of both pefRCD and pefAB operons. In this system, heme acts as an inducer that binds to PefR protein to alleviate repression. The GAS protein shares 43% identity and 74% similarity with PefR from GBS (Figure 5), implying the two proteins may function in a similar manner. We cloned the pefR gene from GAS and expressed it as an N-terminal hexahistidine (6x-His) fusion protein. SDS-PAGE and western blot analysis of the recombinant protein following purification from E. coli confirmed the presence of a single protein at the expected size that reacts with anti-his tag antibodies (Figure 7A and data not shown). Interestingly, the purified PefR had a light brown color (Figure 7B, left inset) and exhibited an absorption in the Soret region (435 nm). Upon titration with free heme, PefR displayed a growing Soret band and concomitant increase in absorption at 530 and 560 nm (β and α bands of heme). These spectral features are characteristics of heme bound protein and are easily separated from the absorption displayed by heme that is free in solution (Zhu et al., 2000). The holo-PefR maintained a bright red color after the removal of heme excess by gel filtration and dialysis (Figure 7B, right inset). Therefore, PefR was purified from E. coli with some heme and readily bound heme in vitro. The plot of changes in absorbance at 435 nm as a function of heme concentrations indicated a binding stoichiometry of 1:2 (PefR:heme, Figure 7C). We determined the dissociation constant (Kd) for PefR by fitting the plot of changes at 435 nm in the modified Hill's equation (Goutelle et al., 2008). We found the Kd to be 10 μM and Hill's coefficient (nH) to be 1.3, greater than 1 (Yifrach, 2004). The molar extinction co-efficient at 435 nm of holo-PefR was determined using the pyridine hemocromogen method (Asakura et al., 1964) and resulted in ε₄₃₅ = 30,407 M⁻¹cm⁻¹. Similarly, titration of PefR with PPIX indicated that it could also bind PPIX in 1:1 stoichiometry with Kd of 9 μM (Figure 8).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.