Recently, the emergence of antibiotic-resistant bacteria has resulted in serious problems for chemotherapy. In addition, many antibacterial agents, such as disinfectants and food additives, are widely used. Therefore, there is a possibility that bacteria are becoming resistant to some antibacterial agents. In this study, we investigated whether Staphylococcus aureus can become resistant to nisin A, one of the bacteriocins applied as a food additive. We isolated a highly nisin A-resistant strain designated SAN2 that displayed increased expression of Pmt proteins, which are involved in the secretion of virulence factors called phenol-soluble modulins (PSMs). This strain also showed decreased susceptibility to human antimicrobial peptides and increased hemolytic activity. In addition, SAN2 showed increased lethal activity in a mouse bacteremia model. Our study provides new insights into the possibility that the acquisition of resistance against food preservatives may modulate virulence in S. aureus, suggesting that we need to pay more attention to the use of food preservatives together with antibiotics.
KEYWORDS: ABC transporters, Staphylococcus aureus, bacteriocins, two-component regulatory systems
ABSTRACT
Nisin A is a bacteriocin produced by Lactococcus lactis and is widely used as a food preservative. Staphylococcus aureus has the BraRS-VraDE system that provides resistance against low concentrations of nisin A. BraRS is a two-component system that induces the expression of the ABC transporter VraDE. Previously, we isolated a highly nisin A-resistant strain with increased VraDE expression due to a mutation in braRS. In this study, we isolated S. aureus MW2 mutants with BraRS-VraDE-independent nisin A resistance. These mutants, designated SAN2 (S. aureus nisin resistant) and SAN469, had a mutation in pmtR, which encodes a transcriptional regulator responsible for the expression of the pmtABCD operon. As a result, these mutants exhibited increased expression of PmtABCD, a transporter responsible for the export of phenol-soluble modulin (PSM). Characterization of the mutants revealed that they have decreased susceptibility to human β-defensin-3 (hBD3) and LL37, which are innate immune factors. Additionally, these mutants showed higher hemolytic activity than the original MW2 strain. Furthermore, in a mouse bacteremia model, the SAN2 strain exhibited a lower survival rate than the original MW2 strain. These results indicate that the increased expression of pmtABCD due to a pmtR mutation is an alternative nisin A resistance mechanism that also affects virulence in S. aureus.
IMPORTANCE Recently, the emergence of antibiotic-resistant bacteria has resulted in serious problems for chemotherapy. In addition, many antibacterial agents, such as disinfectants and food additives, are widely used. Therefore, there is a possibility that bacteria are becoming resistant to some antibacterial agents. In this study, we investigated whether Staphylococcus aureus can become resistant to nisin A, one of the bacteriocins applied as a food additive. We isolated a highly nisin A-resistant strain designated SAN2 that displayed increased expression of Pmt proteins, which are involved in the secretion of virulence factors called phenol-soluble modulins (PSMs). This strain also showed decreased susceptibility to human antimicrobial peptides and increased hemolytic activity. In addition, SAN2 showed increased lethal activity in a mouse bacteremia model. Our study provides new insights into the possibility that the acquisition of resistance against food preservatives may modulate virulence in S. aureus, suggesting that we need to pay more attention to the use of food preservatives together with antibiotics.
INTRODUCTION
Staphylococcus aureus is a commensal bacterium in humans; it generally localizes in the nasal cavity, skin, and intestine and sometimes causes opportunistic infections, such as suppurative diseases, pneumonia, and sepsis (1–3). Additionally, S. aureus causes foodborne illness because this organism produces several heat-stable enterotoxins (2). To cure S. aureus infections, antibacterial agents are generally administered. However, the emergence of methicillin-resistant S. aureus (MRSA) sometimes causes difficulties in chemotherapeutic treatment. Glycopeptides such as vancomycin and teicoplanin are now used for MRSA infection, although vancomycin-intermediate and/or vancomycin-resistant bacteria have been reported (4–6). In addition to antibiotics, other antibacterial agents, such as disinfectants and food preservatives, are widely used. qac genes in S. aureus were identified as resistance factors for quaternary ammonium compounds, such as chlorhexidine and mupirocin (7, 8). Therefore, S. aureus shows resistance not only to several antibiotics but also to other antibacterial agents.
Nisin A is a bacteriocin produced by Lactococcus lactis (9). Nisin A belongs to the lantibiotics, which are antimicrobial peptides containing unusual amino acids, lanthionines (10, 11). The target of nisin A is lipid II, which is involved in cell wall biosynthesis (12, 13). The binding of nisin A to lipid II inhibits cell wall biosynthesis and eventually causes the formation of pores or disturbances in bacterial membranes. Nisin A has a broad range of antibacterial activities against mainly Gram-positive bacteria, including species of Streptococcus, Staphylococcus, and Clostridium (14–19). We and other groups have demonstrated that a two-component system (TCS) named BraRS is responsible for resistance to nisin A (20, 21). When S. aureus cells are exposed to nisin A, BraS senses nisin A, and phosphorylation of BraR then occurs. Phosphorylated BraR binds upstream of vraDE, which encodes an ABC transporter responsible for nisin A resistance. Inactivation of braRS resulted in increased susceptibility to nisin A. However, a high concentration of nisin A is still effective against S. aureus cells (18). Due to its broad-spectrum activity, nisin A is widely used as a food additive worldwide for the prevention of foodborne illness (14, 22, 23). Additionally, the use of bacteriocins, including nisin A, as clinical antibacterial agents has been investigated (14, 17, 24).
Previously, to determine whether nisin A treatment induces increased nisin A resistance, we isolated mutants that showed increased levels of nisin A resistance by incubation with a sub-MIC amount of nisin A and ultimately obtained three mutants that constitutively expressed high levels of VraDE (25). Then, we found point mutations in the promoter region of braRS in the braR or braS coding region in each mutant.
In this study, to explore potential nisin A resistance mechanisms independent of the BraRS-VraDE system, we isolated S. aureus strains that are highly resistant to nisin A by exposing S. aureus to nisin A and eventually obtained mutants with BraRS-VraDE-independent nisin A resistance. Our analysis of these mutants demonstrated that they utilize an alternative nisin A resistance mechanism and that the mutations also affect the virulence of S. aureus.
RESULTS
Isolation of S. aureus strains highly resistant to nisin A.
Previously, we obtained three types of S. aureus strains highly resistant to nisin A, which showed increased levels of vraDE expression (25). In this study, we isolate additional strains highly resistant to nisin A. From two independent experiments, we obtained two mutants that showed no increased expression of vraDE. We designated these mutants SAN2 and SAN469. The nisin A MICs for SAN2 and SAN469 were 2,048 μg/ml, while the MIC for the original MW2 strain was 512 μg/ml (Table 1).
TABLE 1.
Susceptibility of nisin A, bacitracin, gallidermin, and nukacin ISK-1 against S. aureus strains
| Strain | MIC (μg/ml) for: |
|||
|---|---|---|---|---|
| Nisin A | Bacitracin | Gallidermin | Nukacin ISK-1 | |
| MW2 | 512 | 64 | 8 | 16 |
| SAN2 | 2,048 | 64 | 8 | 16 |
| SAN469 | 2,048 | 64 | 8 | 16 |
| COL | 1,024 | 64 | 8 | 16 |
| SAN233 | 2,048 | 64 | 8 | 16 |
| TY34 | 512 | 64 | 8 | 16 |
| SAN455 | 2,048 | 64 | 8 | 16 |
We also evaluated the MICs of these mutants against bacitracin, gallidermin, and nukacin ISK-1, which also inhibit the lipid cycle for cell wall biosynthesis (Table 1). There were no differences in the MICs for these three agents between the mutants and the original MW2 strain.
Analysis of gene expression in MW2 and SAN2 by microarray analysis.
To identify the factors responsible for high resistance to nisin A, we investigated the expression of all genes located on the chromosome of MW2. As shown in Table 2, the expression of genes MW1875 to MW1871 was significantly increased in the SAN2 strain, with more than 30-fold-higher expression levels than those of the original MW2 strain in the absence of nisin A. MW1875 to MW1871 were previously associated with phenol-soluble modulin (PSM) transport (PmtABCD; MW1874 to MW1871) and a transcriptional regulator (PmtR; MW1875) (26). In the strain SAN2, the expression levels of several other genes were also increased, but to lesser extents (2- to 3-fold greater than the MW2 levels).
TABLE 2.
Genes upregulated in S. aureus SAN2
| Gene IDa | Gene name | Function | Fold differenceb | SD | P valuec |
|---|---|---|---|---|---|
| MW1873 | pmtB | ABC transporter, membrane domain | 62.2 | 13.7 | 0.01 |
| MW1874 | pmtA | ABC transporter, ATPase domain | 54.7 | 2.8 | <0.001 |
| MW1875 | pmtR | Transcription regulator, GntR family | 49.0 | 3.3 | 0.001 |
| MW1872 | pmtC | ABC transporter, ATPase domain | 47.7 | 3.2 | 0.002 |
| MW1871 | pmtD | ABC transporter, membrane domain | 38.1 | 6.2 | 0.007 |
| MW1869 | Hypothetical protein | 2.6 | 0.3 | 0.008 | |
| MW1868 | Hypothetical protein | 2.4 | 0.9 | 0.004 | |
| MW1867 | Hypothetical protein | 2.1 | 0.3 | 0.02 |
Gene identification numbers (IDs) are from the Gene Expression Omnibus (GEO) of the NCBI Database (https://www.ncbi.nlm.nih.gov/geo/).
Expression level of SAN2 relative to that of MW2.
The P values were analyzed for statistically significant differences using Student’s t test.
Effects of MW1875 or MW1874 inactivation on nisin A susceptibility.
From microarray analysis, we thought MW1875 to MW1871 were associated with high resistance to nisin A. Therefore, we constructed corresponding inactivation mutants to examine whether these genes truly contributed to high nisin A resistance. Since pmtR and pmtABCD (MW1875 to MW1871) formed an operon, we constructed two insertional mutants (MM2278 and MM2153) in which pYT1 was integrated in MW1875 or MW1874 of strain SAN2 and one insertional mutant (MM2202) with an insertion in MW1875 of strain MW2 (Table 3). The two SAN2 mutants showed the same nisin A MIC as MW2. Similarly, the MW2 mutant (MM2202) retained the same MIC as MW2 (Table 3). A complemented strain, MM2279, in which the pmtRABCD operon (MW1875 to MW1871) was expressed by pCL15 in MM2278, also showed an MIC similar to that of SAN2, indicating that the increased expression of pmtRABCD was involved in high nisin A resistance in SAN2. We also constructed one mutant (MM2259) in which only MW1875 was inactivated but in which MW1874 to MW1871 were constitutively expressed and found that MM2259 showed the same MIC as the SAN2 strain.
TABLE 3.
Strains used in this study
| Strain | Characteristics | MIC of nisin A (μg/ml) | Reference or source |
|---|---|---|---|
| S. aureus | |||
| MW2 | Clinical strain, methicillin-resistant (mecA+) | 512 | 45 |
| COL | Clinical strain, methicillin-resistant (mecA+) | 1,024 | 46 |
| TY34 | Clinical strain, methicillin-resistant (mecA+) | 512 | 47 |
| MM2202 | pYT1 insertion in MW1875 (MW1875 to MW1871 inactivation) in MW2, TETr | 512 | This study |
| MM2280 | Overexpression of MW1875 to MW1871 (pCL8) in MW2, CHLr | 2,048 | This study |
| MM2259 | pYT1 insertion in MW1875 (MW1874 to MW1871 overexpression) in MW2, TETr | 2,048 | This study |
| SAN2 | Nisin A-resistant mutant from MW2 | 2,048 | This study |
| MM2278 | pYT1 insertion in MW1875 (MW1875 to MW1871 inactivation) in SAN2, TETr | 512 | This study |
| MM2153 | pYT1 insertion in MW1874 (MW1874 to MW1871 inactivation) in SAN2, TETr | 512 | This study |
| MM2279 | MW1875 to MW1871 (SAN2) complementation in MM2278, TETr, CHLr | 2,048 | This study |
| SAN469 | Nisin A-resistant mutant from MW2 | 2,048 | This study |
| SAN233 | Nisin A-resistant mutant from COL | 2,048 | This study |
| SAN455 | Nisin A-resistant mutant from TY34 | 2,048 | This study |
| E. coli | |||
| XL-II | endA1 supE44 thi-1 hsdR17 recA1 gyrA96 relA1 lac [F′ proAB lacIqZ∆M15 Tn10 (TETr) Amy CHLr] | Stratagene | |
| MM1127 | His tag-fused MW1875 (MW2) gene in XL-II, AMPr | This study | |
| MM1128 | His tag-fused MW1875 (SAN2) gene in XL-II, AMPr | This study |
DNA sequences of the MW1875 to MW1871 (pmtR and pmtA to pmtD) region.
The DNA sequences of the MW1875 to MW1871 region in MW2, SAN2, and SAN469 were determined. In SAN2, only one mutation was detected in MW1875 (Fig. 1). This mutation induced the replacement of alanine (Ala) at the 43rd amino acid of MW1875 with aspartic acid (Asp) in the SAN2 strain. In SAN469, the fifth amino acid position in MW1875 was mutated to a stop codon.
FIG 1.
Open reading frame (ORF) map of the pmt region (MW1875 to MW1871) and mutation sites in pmtR of isolated mutants. The mutation sites in the pmtR region are indicated by white arrows.
Isolation of highly nisin A-resistant mutants derived from other S. aureus strains.
To determine whether similar pmtR mutants were able to be obtained from other strains after exposure to nisin A, we isolated nisin A-resistant mutants of S. aureus COL and TY34. From two independent experiments with each strain, we obtained two mutants with increased expression of pmtRABCD, one from each strain (Table 3). We designated the mutants SAN233 (from COL) and SAN455 (from TY34). DNA sequencing analysis identified point mutations at the 16th and 93rd amino acids of MW1875 (PmtR) in SAN233 and SAN455, respectively (Fig. 1), while there were no mutations in the pmtABCD genes. These two mutations introduced stop codons within the pmtR gene. Therefore, SAN233 and SAN455 did not express full-length PmtR.
Expression of pmtR (MW1875), pmtA (MW1874), and vraD in MW2 and its mutants.
We investigated the expression of pmtR, pmtA, and vraD by quantitative PCR. In SAN2, the expression of pmtR and pmtA (a more than 70-fold increase) was significantly increased compared to that in MW2 in the absence of nisin A (Fig. 2A). We observed the same result using immunoblot analysis (Fig. 2B). We also confirmed the increased expression of pmtR and pmtA in SAN469 using quantitative PCR (see Fig. S1 in the supplemental material). In addition, the PmtR and PmtA expression patterns of MW2 and SAN2 in the absence of nisin A were quite similar to those in the presence of nisin A because the expression of pmtR and pmtA was not induced by nisin A (Fig. S2). Additionally, we investigated the expression of vraD and found that SAN2 showed no increase in vraD expression at a low concentration of nisin A (32 μg/ml), while MW2 showed increased expression of vraD (Fig. 2C). However, the inactivation of pmtRABCD in SAN2 resulted in increased vraD expression in the presence of nisin A.
FIG 2.
Expression of pmtR, pmtA, and vraD in the mutants. (A) Expression of pmtR and pmtA in MW2, SAN2, MM2278 (SAN2 with inactivation of pmtR-pmtD), and MM2279 (the complemented strain in MM2278) was evaluated by quantitative PCR using specific primers. *, P < 0.001; determined with Dunnett’s post hoc test compared to untreated MW2. (B) Expression levels of PmtR and PmtA in S. aureus strains were evaluated by immunoblotting analysis using respective specific antibodies. (C) Expression of vraD in S. aureus strains in the presence or absence of 1/16 of the MIC of nisin A (32, 128, 32, and 512 μg/ml in MW2, SAN2, MM2278, and MM2279, respectively) was evaluated by quantitative PCR using specific primers. *, P < 0.005; determined with Dunnett’s post hoc test compared to untreated MW2.
Then, we investigated vraD expression in the presence of various concentrations of nisin A (Fig. 3A). In MW2, the expression of vraD was induced at concentrations above 1/32 of the MIC for nisin A (16 μg/ml), while in SAN2, it was induced by concentrations above half of the MIC for nisin A (1,024 μg/ml). We further investigated the induction of vraD expression by bacitracin (1 MIC = 64 μg/ml). Since the MIC of bacitracin was the same in the MW2 and SAN2 strains (Table 1), we analyzed the effect of bacitracin at a range from 1/64 MIC to 1 MIC. The vraD expression in both strains was similarly induced by bacitracin at each concentration (Fig. 3B).
FIG 3.
Expression of vraD in MW2 and SAN2 with various concentrations of nisin A and bacitracin. The induction of vraD expression in S. aureus MW2 and SAN2 by the addition of various concentrations of nisin A (A) or bacitracin (B) was evaluated by quantitative PCR using specific primers. *, P < 0.05; **, P < 0.01; determined with Dunnett’s post hoc test compared to untreated MW2 or SAN2.
Binding of the wild-type and mutated PmtR protein to the upstream region of MW1875.
Fig. 4A shows the previously reported promoter region of pmtR, the transcriptional start site, and the binding region of PmtR (27). The results of our electrophoretic mobility shift assay (EMSA) revealed that MW2-rPmtR bound the upstream region of pmtR (pmtR-F), while SAN2-rPmtR did not bind its region (Fig. 4B). This binding of MW2-rPmtR was inhibited by the addition of an excess amount of unlabeled DNA fragments.
FIG 4.
EMSAs of PmtR. (A) The nucleotide sequence of the pmtR promoter region. Squares, –35, –10 box; *, pmtR transcriptional start site; bold, PmtR-binding region. (B) EMSAs of PmtR with a DNA fragment labeled with DIG were performed with the method described in Materials and Methods.
Hemolytic activity of MW2, SAN2, MM2278, and MM2279.
Since PSM transported by PmtABCD has been demonstrated to be involved in hemolytic activity (28), we hypothesized that the increased expression of PmtABCD would affect hemolytic activity, and we next analyzed the hemolytic activities of the strains MW2 and SAN2 on sheep blood agar. Compared to wild-type strain MW2, SAN2 produced a larger hemolytic zone (Fig. 5A). When pmtR and pmtABCD (pmtR-D, MW1875 to MW1871) were inactivated in SAN2 (MM2278), the hemolytic zone became smaller than that of SAN2. The hemolytic zone produced by the complemented strain MM2279 was similar in size to that of SAN2.
FIG 5.
Hemolytic assay of S. aureus mutants. (A) Three microliters each of 10-fold-diluted overnight cultures of MW2, SAN2, MM2278 (SAN2 with inactivation of pmtR-pmtD), and MM2279 (the complemented strain in MM2278) was spotted individually on a sheep blood agar plate and incubated at 37°C for 2 days. The percentage of hemolysis area was evaluated using ImageJ. *, P < 0.01; **, P < 0.005; determined with Dunnett’s post hoc tests compared to MW2. (B) Three microliters of a 10-fold-diluted overnight culture of RN4220 (a β-hemolysin-producing strain) was spotted in the center of a sheep blood agar plate, and equal volumes of MW2, SAN2, MM2278, MM2279, and SAN469 cultures were spotted around RN4220 (distance from the center of the RN4220 spot was 17 mm). The agar plate was incubated at 37°C for 20 h and then kept at 4°C for 2 days. White arrows indicate a synergistic effect. The percentage of hemolysis area was evaluated using ImageJ. *, P < 0.01; **, P < 0.005; determined with Dunnett’s post hoc tests compared to MW2.
It was reported that PSM or δ-hemolysin enhanced the activity of β-hemolysin (28). Since MW2 (and SAN2) does not produce β-hemolysin, we investigated the synergistic hemolytic activity of MW2 and its derivatives with RN4220 (which produces β-hemolysin). An enhanced hemolytic zone with RN4220 was produced (white arrows in Fig. 5B) by SAN2, MM2279, and SAN469, while MW2 and MM2278 did not have an enhanced hemolytic effect with RN4220 (Fig. 5B). We also investigated the hemolytic activity of COL, SAN233, TY34, and SAN455 and found similar results (Fig. S3).
Susceptibility to hBD3 and LL37 in MW2, SAN2, MM2278, and MM2279.
Since it was reported in a previous study that pmtR-D inactivation increased the susceptibility to hBD3 and LL37 (29), we investigated the effect of the increased expression of pmtABCD in SAN2 on susceptibility to hBD3 (Fig. 6) and LL37 (Fig. S4). As shown in Fig. 6, compared to the original MW2 strain, SAN2 showed a decrease in susceptibility to hBD3, while the inactivation of pmtRABCD increased susceptibility to the peptides, resulting in the same susceptibility as that of MW2. The complemented strain (MM2279) showed susceptibility similar to that of SAN2.
FIG 6.
Susceptibility of S. aureus mutants to hBD3. An antibacterial assay for hBD3 was performed as described in Materials and Methods. S. aureus was incubated with various concentrations of hBD3 for 10 min (A) and 2 h (B). *, P < 0.05, determined by Dunnett’s post hoc test compared to untreated MW2.
Mouse survival rate after injection of MW2, SAN2, MM2278, and MM2279.
We performed a mouse survival experiment using a bacteremia model. Injection of the MW2 strain killed only 1 out of 8 mice by 3 days after injection, while 5 of 8 mice were killed 1 to 5 days after the injection of SAN2 (Fig. 7A; P = 0.010, log-rank test). No mouse was killed by injection of the SAN2 pmtR-D inactivation strain (MM2278), while 4 of 5 mice were killed 1 to 3 days after the injection of the complemented strain (MM2279) (Fig. 7B, P = 0.002; Fig. 7C, P = 0.604, log-rank test).
FIG 7.
Mouse survival experiment. Survival percentage of Slc:ddY mice after being challenged with intravenous injection of 1.5 × 108 CFU of S. aureus. (A) Survival comparison between MW2- and SAN2-inoculated mice. (B) Survival comparison between wild-type SAN2 and MM2278 (SAN2 with inactivation of pmtR-pmtD). (C) Survival comparison between wild-type SAN2 and MM2279 (the complemented strain in MM2278). Significant differences between the two strains are shown. Survival statistics were calculated with the log-rank test (Mantel-Cox).
DISCUSSION
In this study, we demonstrated a novel high nisin A resistance mechanism that functions independently of the BraRS-VraDE system. We obtained 4 mutant strains with increased expression of PmtABCD from MW2, COL, and TY34 that acquired high nisin A resistance. All these mutants had a point mutation in the pmtR gene, yielding a mutant PmtR with an Ala43Asp substitution (SAN2 from MW2) or truncated PmtR (SAN469 from MW2, SAN233 from COL, and SAN455 from TY34). EMSAs showed that the SAN2-derived PmtR protein (SAN2-rPmtR) did not bind the DNA region upstream of pmtR-D. Since PmtR is a negative transcriptional regulator of the pmtRABCD operon (27), SAN2-PmtR and the three truncated PmtRs could not suppress the expression of the pmtR-D operon, resulting in the increased expression of pmtR-D. The mutation site of PmtR in SAN2 is within the helix-turn-helix DNA-binding region. Based on our EMSA, a mutated PmtR in SAN2 lost the ability to bind to the target DNA region. This result suggests two possibilities. One possibility is that the mutation site is critical for DNA binding. The other possibility is that the mutation causes a structural change in PmtR, leading to the loss of DNA binding.
PmtABCD forms an ABC transporter from two membrane proteins (PmtA and C) and two ATPases (PmtB and D) (26). PmtABCD is involved in the transport of PSMs and δ-hemolysin (Hld) from the cytoplasm to the extracellular space (26, 28). PSMs have broad virulence activities, such as the surface spreading activity responsible for epithelial colonization, biofilm formation, proinflammatory activity, cytolytic activity, and antimicrobial activity (30–32). In addition, Cheung et al. recently reported that the Pmt transporter is also associated with human-derived antimicrobial peptides, such as hBD3 and LL37 (29). We also found that high expression of Pmt transporters in SAN2 resulted in high resistance against hBD3 and LL37. These results suggest the association of Pmt transporters with the susceptibility of human antimicrobial peptides. Although there are no clear structural similarities among PSMs, δ-hemolysin, hBD3, and LL37, we speculate that PmtABCD may be associated with the export of these peptides with membrane insertional activity (Fig. 8). However, we also evaluated the susceptibility to bacitracin, gallidermin, and nukacin ISK-1 and found no difference between the MW2 and SAN2 strains. Nukacin ISK-1 is functionally similar to nisin A because they are both lantibiotics with the same target, lipid II (12, 33). Furthermore, we found a different vraD expression response; bacitracin induced vraD expression in both the MW2 and SAN2 strains with the same low concentration of nisin A, while only a high concentration of nisin A than that for MW2 induced the expression of vraD (Fig. 3A and B). These results indicate that the Pmt system recognizes the limited structure of antimicrobial peptides, such as nisin A and LL37, with membrane insertional activity.
FIG 8.
Proposed high nisin A resistance mechanism mediated by the Pmt system. The MW2 strain shows resistance to low concentrations of nisin A by the BraRS system. In the SAN2 strain, the Pmt system contributes to high nisin A resistance due to the mutation in PmtR.
Joo et al. demonstrated that PSMα1-3 binds PmtR to release it from its target DNA region, which is followed by induction of the expression of the pmt operon (27). PSM expression is upregulated by the agr system (28, 31). Since agr expression increases in the late exponential phase (34, 35), the expression levels of PSM and PmtABCD are increased at the late exponential phase. In this study, we collected bacterial cells for the evaluation of gene expression at the mid-exponential phase (optical density [OD] at 660 nm, 0.5) and found strong expression of pmtA in SAN2 but not MW2 (Fig. 2A and S1), indicating that in the SAN2 mutant, PmtABCD expression is independent of the growth phase due to the lack of functional PmtR. Therefore, the expression of PmtABCD is constitutively increased; thus, PSMs are constantly exported from the cell during SAN2 growth. As mentioned above, PSMs have broad virulence activities, so the increased PSM transport may modulate the virulence of S. aureus. We found that the hemolytic activity of SAN2 was greater than that of the wild-type MW2, likely because PSMα has an enhanced effect on hemolytic activity (28). Additionally, susceptibility to human antimicrobial peptides was decreased in the mutants. Finally, in a mouse experiment, we found that compared to the wild-type MW2, SAN2 showed an increase in fatality rate due to its increased virulence. The Pmt-dependent nisin A resistance identified in this study is different from the previously identified BraRS-VraDE system in terms of the modulation of virulence in S. aureus.
Previously, we isolated strains that were highly resistant to nisin A and that showed high constitutive expression of VraDE (25). VraDE, which is regulated by BraRS, is an intrinsic resistance factor with activity against nisin A, bacitracin, gallidermin, and nukacin ISK-1. In MW2, sub-MIC (1/16 MIC) amounts of nisin A induced the expression of VraDE, while sub-MIC amounts of nisin A did not induce its expression in SAN2. However, pmtABCD inactivation in SAN2 caused inducible expression of VraDE by sub-MIC amounts of nisin A. Additionally, induction of vraD expression occurred at high concentrations of nisin A in SAN2, suggesting that the Pmt system is a dominant system contributing to nisin A resistance in SAN2 (Fig. 8).
In this study, we used nisin A obtained from Sigma-Aldrich with a 2.5% formulation. Since the MIC of nisin A in MW2 was measured as 512 μg/ml in this study, the MIC of purified nisin A is estimated to be 12.8 μg/ml. There are several reports regarding the concentration of nisin A in several foods, such as cheese, sausage, and milk (36, 37). Although the nisin A concentration is varied among foods, its concentration is mostly above 10 μg/g. Therefore, the concentration of nisin A for the isolation of the resistant mutant in this study is close to that for food preservatives, suggesting the possibility that nisin A resistance in S. aureus might be generated by food preservatives.
In conclusion, we found a new mechanism for high nisin A resistance that is mediated by the increased expression of Pmt proteins in S. aureus. It is important to note that the increased expression of Pmt proteins causes high nisin A resistance and enhances virulence. Since nisin A is widely used as a food additive, it is important to use it cautiously.
MATERIALS AND METHODS
Bacterial strains and growth conditions.
The bacterial strains used in this study are shown in Table 3. S. aureus and Escherichia coli XL-II were grown in Trypticase soy broth (TSB) (Becton, Dickinson and Company, Franklin Lakes, NJ, USA) and Luria-Bertani (LB) broth, respectively. Tetracycline (Wako Chemicals, Osaka, Japan) (TET, 5 μg/ml) and chloramphenicol (Sigma-Aldrich, St. Louis, MO, USA) (CHL, 5 μg/ml) were used for S. aureus, and ampicillin (Wako Chemicals, Osaka, Japan) (AMP, 100 μg/ml) was used for E. coli when necessary.
MIC determination.
The MICs of several antibacterial agents were determined by microdilution with liquid culture (TSB), as described previously (38). Nukacin ISK-1 was purified as described previously (20). Nisin A with a 2.5% formulation (Sigma-Aldrich, St. Louis, MO, USA) was used in this study. Bacitracin and gallidermin were obtained from Wako Chemicals, Osaka, Japan, and Santa Cruz Biotechnology, TX, USA, respectively. S. aureus strains (105 cells) were inoculated into 100 μl of TSB containing various concentrations of nisin A (4 to 4,096 μg/ml), nukacin ISK-1 (0.125 to 128 μg/ml), gallidermin (0.125 to 128 μg/ml), and bacitracin (0.25 to 256 to μg/ml). MICs were determined after incubation for 24 h at 37°C. MIC determination was repeated independently three times.
Isolation of mutants highly resistant to nisin A.
S. aureus mutants highly resistant to nisin A were obtained using a method described elsewhere (25). Briefly, a microdilution method that is generally used to evaluate the MICs of antibacterial agents was used for isolation of the mutants. An overnight culture of S. aureus MW2 was diluted to 107 cells/ml, and 10 μl of the diluted culture was applied to each well of a plate (100 μl), which contained a 2-fold dilution series of nisin A (16 to 16,384 μg/ml). After incubation at 37°C overnight, bacterial cells that grew in 1/2 of the MIC of nisin A were collected and diluted 100-fold. Then, 10 μl of this diluted sample was applied to each well (100 μl), which contained serial 2-fold dilutions of nisin A (16 to 16,384 μg/ml). This procedure was repeated three times. Ultimately, bacterial cells grown in the presence of 1/2 of the MIC of nisin A were appropriately diluted and plated on tryptic soy agar (TSA). After overnight incubation, 14 colonies were picked and incubated in 5 ml of TSB. Overnight culture was used for the determination of the MIC of nisin A. This experiment was independently performed 3 times. Then, we investigated the vraD expression of the strains with high nisin A resistance using quantitative PCR and found no increase in vraD expression.
We also tried to isolate strains highly resistant to nisin A from S. aureus COL and TY34 using the method described above.
Microarray analysis.
Overnight cultures of S. aureus (108 cells) were inoculated into 30 ml of fresh TSB and cultured at 37°C with shaking. When the OD at 660 nm (OD660) reached 0.4, the bacterial cells were collected by centrifugation at 5,000 × g for 5 min at 4°C. Total RNA was extracted using a FastRNA Pro Blue kit (MP Biomedicals, Cleveland, OH, USA) according to the manufacturer’s protocol. Then, cDNA from each sample was synthesized from 10 μg of total RNA using a FairPlay III microarray labeling kit (Agilent Technologies, Santa Clara, CA, USA) according to the manufacturer’s instructions. The Agilent eArray platform (Agilent Technologies) was used to design the microarray; 13,939 probes (60-mers) were designed to target the 2,628 protein-coding genes of S. aureus MW2 (up to 5 probes per gene). Microarray analysis was performed using a method described elsewhere (39). The experiments were performed using three biological replicates (three technical replicates for each set of conditions), and the expression data were deposited into the Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/) under accession number GSE131352. Statistical significance was determined using Student’s t test. The P values are shown in Table 2.
DNA sequences of the MW1875 to MW1871 region.
Since we found the increased expression of genes corresponding to MW1875 to MW1871 in the SAN2 strain, we determined the DNA sequence of the MW1875 to MW1871 region. Primers were constructed to amplify MW1875 to MW1871 with their corresponding flanking regions, including the promoter regions of MW1875 (Table 4). To prepare chromosomal DNA from the original MW2 strain and the mutant strains, the cells from 1 ml of overnight culture were collected. The cells were suspended in 100 μl of 10-mM Tris-HCl (pH 6.8) containing 10 μg of lysostaphin (Sigma-Aldrich), incubated at 37°C for 20 min, and then incubated at 95°C for 15 min. After centrifugation, cell lysates were used as the template DNA for PCR. PCR was performed using the TaKaRa Ex Taq system, and the amplicons were purified using a QIAquick kit (Qiagen, Hilden, Germany). The nucleotide sequence of each DNA fragment was determined using specific primers. The primers used to amplify DNA sequences are listed in Table 4.
TABLE 4.
Primers used in this studya
| Target gene ID | Primer-forward | Primer-reverse |
|---|---|---|
| Construction of gene-inactivated mutants | ||
| pmtR + pmtA-D | 5′-TTGGATCCACAATAGTGATTTTCCGATT-3′ | 5′-CAAAGCTTTGGCTTGCGCTTCATTAA-3′ |
| pmtA-D | 5′-AGGGATCCATATGGCTCTCAATCCG-3′ | 5′-AAAAGCTTCACTCACAACTTGATATC-3′ |
| Construction of the plasmid for gene complementation | ||
| pmtR + pmtA-D-pCL15 | 5′-ATAAGCTTATACAGAAAGTGATAGGG-3′ | 5′-TTGGATCCAACTGATCACTTGAATAATT-3′ |
| pmtA-D-pCL15 | 5′-ATAAGCTTTTTTAAGCTTCATTTATGAG-3′ | 5′-TTGGATCCAACTGATCACTTGAATAATT-3′ |
| pmtR + pmtA-D-pCL8 | 5′-CTAAGCTTTTGAAGTAGACAATGCAAG-3′ | 5′-ATCCCGGGTTCCCAACCTCAAAATTAT-3′ |
| Pmt promoter + pmtA-D-pYT1 | 5′-CAAGATCTAATGGTAGTGTCATTTCATT-3′ | 5′-CAGGATCCCAACGTCCCCCTATCAC-3′ |
| 5′-TTGGATCCACAATAGTGATTTTCCGATT-3′ | 5′-CAAAGCTTTGGCTTGCGCTTCATTAA-3′ | |
| Amplification of DNA fragments used in gel shift assay | ||
| pmtR-F1 | 5′-AATGGTAGTGTCATTTCATTT-3′ | 5′-CAACGTCCCCCTATCAC-3′ |
| Construction of the plasmid for recombinant protein | ||
| rPmtR | 5′-CCGGATCCATGAAAATAATTTTAAAAAACAAT-3′ | 5′-TTAACGTTTCATGATGATTCCTCCTCA-3′ |
| rPmtA | 5′-CCGGATCCATGAATGCCATAGAATTAAG-3’ | 5′-TTAAGCTTTTAAAAACCTTCTTCCATCA-3′ |
| Primers for quantitative PCR | ||
| pmtR | 5′-AATTGGTTAATGAAGCGCAAG-3′ | 5′-GATTCCTCCTCATAAATGAACG-3′ |
| pmtA | 5′-TAAAGCTTCGTTCATTTATGAGGAGG-3′ | 5′-ACGATAAAAAGGGGCAATCA-3′ |
| vraD | 5′-CACTTGCCAAATTCCGTA-3′ | 5′-AATACCTAATGCTGTCGTGA-3′ |
| gyrB | 5′-AGGTCTTGGAGAAATGAATG-3′ | 5′-CAAATGTTTGGTCCGGTT-3′ |
| Primers used for the DNA sequence | ||
| pmtR, A-D-seq-F-500 | 5′-TGAAATTCAATAACTTATTAAA-3′ | |
| pmtR, A-D-F-26 | 5′-ATACAGAAAGTGATAGGG-3′ | |
| pmtR, A-D-F351 | 5′-AACGTTCATTTATGAGGAGG-3′ | |
| pmtR, A-D-F967 | 5′-ATTATATCATTCACTTAAGTG-3′ | |
| pmtR, A-D-F1429 | 5′-TTTGGATTTTAGATGCTGGTCA-3′ | |
| pmtR, A-D-F1967 | 5′-ATTACAAAAAAATACGGCTC-3′ | |
| pmtR, A-D-F2409 | 5′-ATGGCTCAATTGATGTGCTG-3′ | |
| pmtR, A-D-F3047 | 5′-CGCGTGATTTTTCACAAGGT-3′ | |
| pmtR, A-D-seq-R | 5′-TTTAAAATTCCCAACCTCA-3′ |
Underlines indicate the recognition site of the respective restriction enzyme.
Inactivation of MW1875 and MW1874 in S. aureus and their complementation.
The strains used in this study are listed in Table 3. Specific gene inactivation by the insertion of the thermosensitive plasmid pYT1 was performed using a method described elsewhere (40). An internal DNA fragment of MW1875 or MW1874 was amplified by PCR using specific primers and then cloned into the pYT1 vector. The plasmid was electroporated into S. aureus RN4220, and then the plasmid in RN4220 was transferred into respective strains by transduction using phage 80α. The obtained strains were grown overnight at 30°C. Appropriate dilutions of the culture were spread on TSA plates containing TET (10 μg/ml) and then incubated at 42°C overnight. Colonies were picked and replated on TSA containing TET. Disruption of the target gene was checked by PCR. For gene complementation, the vector pCL8, which is a shuttle vector for E. coli and S. aureus, was used (41). A DNA fragment for complementation was PCR amplified using chromosomal DNA from the MW2 or mutant strains. The DNA fragment was cloned into pCL8 using E. coli XL-II competent cells. The obtained plasmid was electroporated into S. aureus RN4220 and was subsequently transduced into the appropriate strain using phage 80α.
To obtain a mutant that constitutively expresses MW1874 to MW1871 by its own promoter upstream of MW1875 without MW1875 expression, we constructed a plasmid using PCR cloning of a DNA fragment containing the internal region of MW1875 (295 bp) and the promoter region of MW1875 to MW1871 (149 bp). Two PCR fragments were obtained and then cloned into the pYT1 vector by using restriction enzymes. The plasmid was finally transduced into the MW2 strain. The plasmid integration was performed using the method described above. Two types of mutants were obtained (Fig. S5). Finally, the strain that contained inactivated MW1875 and expressed MW1874 to MW1871 was verified by PCR and DNA sequencing.
Quantitative PCR and immunoblotting analysis.
Quantitative PCR was performed to investigate the expression of MW1875 (pmtR), MW1874 (pmtA), and vraD. A small portion of overnight culture (108 cells) was inoculated into 5 ml of fresh TSB and then grown at 37°C with shaking. When the optical density at 660 nm reached 0.5, nisin A (16 to 2,048 μg/ml) or bacitracin (1 to 64 μg/ml) was added to the bacterial culture. After incubating for 15 min (for quantitative PCR) and 2 h (for immunoblotting), the bacterial cells were collected. For quantitative PCR, RNA extraction, cDNA synthesis, and PCR were performed as described previously (25). Statistical significance was determined (*, P < 0.01; **, P < 0.005) using Dunnett’s post hoc tests compared to the control in each experiment.
For immunoblotting, antiserum against MW1875 and MW1874 was obtained by immunizing a rabbit with the recombinant protein, as described previously (42). Briefly, the coding region of MW1875 or MW1874 amplified by PCR using specific primers was cloned into pQE30 (Qiagen, Tokyo, Japan), which is used for the construction of histidine-tagged recombinant proteins. The obtained plasmid was transformed into E. coli M15(pREP4). The resulting recombinant protein was purified according to the manufacturer’s instructions. Antiserum was obtained using the recombinant protein. We also immunoblotted VraD using anti-rVraD antibodies obtained previously (25).
The collected bacterial cells were resuspended in 200 μl of Tris-HCl (pH 6.8) containing 10 μg of lysostaphin and incubated for 20 min at 37°C. The cells were then heated at 95°C for 15 min. After centrifugation, the supernatant was obtained as a whole-cell lysate. Lysate proteins mixed with an equal volume of sample loading buffer were resolved by 15% SDS-polyacrylamide gel electrophoresis (PAGE). Then, the proteins were transferred to a nitrocellulose membrane. After blocking with 2% skim milk in Tris-buffered saline (TBS; 20 mM Tris, 137 mM NaCl [pH 8.0]) containing 0.05% Tween 20 (TBS-T), the membrane was incubated with specific antiserum (diluted 1:1,000 in 1% skim milk in TBS-T) for 1 h at 37°C. The membrane was then washed with TBS-T and incubated with horseradish peroxidase-conjugated anti-mouse IgG (diluted 1:1,000 in TBS-T) (Promega, Madison, WI, USA) for 1 h at 37°C. The membrane was then washed 5 times with TBS-T, and the protein band reacting with the antiserum was detected using a chemiluminescence detection system (PerkinElmer, Waltham, MA, USA).
Hemolysis assay.
Hemolysis assays were performed using TSB agar plates containing 5% sheep blood (Becton, Dickinson and Company). To evaluate the hemolysis activity of individual strains, 3 μl each of 10-fold-diluted overnight cultures of MW2, SAN2, MM2278 (pmtR, pmtABCD inactivated strain), and MM2279 (complemented strain of MM2278) were spotted individually on a sheep blood agar plate and incubated at 37°C for 2 days. The hemolysis area for each strain was calculated using ImageJ analysis. This experiment was performed independently 3 times. To evaluate the synergistic effect on β-hemolysin of individual strains, 3 μl of a 10-fold-diluted overnight culture of the RN4220 strain (a β-hemolysin-producing strain) was spotted. Additionally, 3 μl each of 10-fold-diluted overnight cultures of MW2, SAN2, MM2278, and MM2279 was spotted around RN4220 (distance from the center of the RN4220 spot was 17 mm). The agar plate was incubated at 37°C for 20 h and then kept at 4°C for 2 days. This experiment was performed independently 3 times to confirm the results. We quantified hemolytic activity by measuring the hemolysis zone for each strain using ImageJ analysis. Briefly, we excised the area covering the hemolysis zone (1.5 cm × 1.5 cm). Then, the percentage of hemolysis area (%) for MW2, SAN2, MM2278, MM2279, and SAN469 was evaluated using ImageJ. Statistical significance was determined (*, P < 0.01; **, P < 0.005) using Dunnett’s post hoc tests compared to MW2.
Susceptibility to hBD3 and LL37.
An antibacterial assay was performed as described elsewhere (43). Briefly, overnight cultures of S. aureus strains were collected and washed with 10 mM sodium phosphate buffer (PB). The bacterial suspension was diluted to 107 cells/ml with PB, and 10 μl of the bacterial suspension (105 cells) was inoculated into 500 μl of PB with or without human antimicrobial peptides (β-defensin-3 [Peptide Institute, Inc., Osaka, Japan] or LL37; 0.5, 0.4, 0.2, 0.1, and 0.05 μM) and incubated aerobically for 2 h and 10 min at 37°C. Dilutions of the reaction mixture (100 μl) were plated on agar medium and incubated at 37°C overnight. The CFU were determined as the total number of colonies on each plate. The antibacterial effect was calculated as the ratio of the number of surviving cells (survival rate [%]) to the total number of bacteria incubated in a control PB solution after exposure to antimicrobial peptides. Statistical significance was determined (*, P < 0.05) using Dunnett’s post hoc test compared to untreated MW2.
Electrophoretic mobility shift assay.
For an electrophoretic mobility shift assay (EMSA), two 6-histidine-tagged recombinant PmtR (rPmtR) proteins each from the original MW2 strain and SAN2 were used. Construction of the rPmtRs was performed as described above. Briefly, a DNA fragment encoding pmtR (MW1875) was amplified with specific primers using chromosomal DNA of MW2 and SAN2. Then, DNA fragments were cloned into pQE30. The plasmid was then transformed into E. coli M15(pREP4). The recombinant protein was purified according to the manufacturer’s instructions. To assess the binding of rPmtR to a region upstream of pmtR, an EMSA was performed as described previously (25). A DNA fragment encompassing the region upstream of pmtR, pmtABCD, was amplified with the specific primers listed in Table 1. The DNA fragments were labeled at the 3′ end with digoxigenin (DIG) using a DIG gel shift kit, 2nd generation (Roche, Mannheim, Germany). The DIG-labeled fragments (5 ng) were reacted with rPmtR protein (50 mM) in the labeling buffer provided with the kit. When necessary, nonlabeled DNA fragments (10 ng) were added to the reaction mixture. After native PAGE on a 6% polyacrylamide gel, the DNA fragments were transferred to a positively charged nylon membrane (Roche, Mannheim, Germany) and visualized according to the manufacturer’s protocol.
Mouse bacteremia experiment.
Since we found the alteration of virulence in the SAN2 strain, we then performed the animal experiment to see whether the SAN2 strain showed strong virulence in vivo. A mouse bacteremia experiment was performed as described previously (44). Six-week-old female Slc:ddY mice were purchased from SLC (Shizuoka, Japan). Small portions of overnight cultures of S. aureus MW2, SAN2, SAN2 with MW1875 to MW1871 inactivation (MM2278), and MM2278 complemented with MW1875 to MW1871 (MM2279) were inoculated into 5 ml of fresh TSB and incubated at 37°C with shaking. When the OD660 reached 0.8, bacterial cells were collected and washed with PBS. Then, the cells were resuspended in PBS at a concentration of 1.0 × 109 CFU/ml. An aliquot of 100 μl (1.0 × 108 CFU) was injected into the tail vein of the mice. We divided the mice into the following 3 groups for this experiment: (A) survival comparison between MW2 and SAN2, (B) the comparison between SAN2 and MM2278, and (C) the comparison between SAN2 and MM2279. The number of trials was as follows: (A) 8 MW2, 8 SAN2; (B) 8 SAN2, 6 MM2278; and (C) 6 SAN2, 5 MM2279. Mouse survival was monitored for 6 days. Survival statistics were calculated using the log-rank test (Mantel-Cox). The animal experimentation performed in this study was conducted according to a protocol approved by the president of Kagoshima University after review by the Institutional Animal Care and Use Committee (ethical number D18015).
Data availability.
Newly determined data were deposited in the Gene Expression Omnibus under accession number GSE131352.
Supplementary Material
ACKNOWLEDGMENTS
This study was supported in part by a Grant-in-Aid for Scientific Research (C) (grant no. 18K09553) from the Ministry of Education, Culture, Sports, Sciences, and Technology of Japan. This work was supported by the Facility of Laboratory Animal Science, Research Support Center, Institute for Research Promotion, Kagoshima University.
We declare no competing financial interests in relation to the work described.
Footnotes
Supplemental material is available online only.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
Data Availability Statement
Newly determined data were deposited in the Gene Expression Omnibus under accession number GSE131352.