Skip to main content
NCBI home page
Antimicrobial Agents and Chemotherapy logoLink to Antimicrobial Agents and Chemotherapy
. 2016 Apr 22;60(5):2684–2691. doi: 10.1128/AAC.03022-15

Daptomycin Tolerance in the Staphylococcus aureus pitA6 Mutant Is Due to Upregulation of the dlt Operon

Lukas Mechler a, Eve-Julie Bonetti b, Sebastian Reichert a, Matthias Flötenmeyer c, Jacques Schrenzel b, Ralph Bertram a,d, Patrice François b, Friedrich Götz a,✉
PMCID: PMC4862447  PMID: 26883712

Abstract

Understanding the mechanisms of how bacteria become tolerant toward antibiotics during clinical therapy is a very important object. In a previous study, we showed that increased daptomycin (DAP) tolerance of Staphylococcus aureus was due to a point mutation in pitA (inorganic phosphate transporter) that led to intracellular accumulation of both inorganic phosphate (Pi) and polyphosphate (polyP). DAP tolerance in the pitA6 mutant differs from classical resistance mechanisms since there is no increase in the MIC. In this follow-up study, we demonstrate that DAP tolerance in the pitA6 mutant is not triggered by the accumulation of polyP. Transcriptome analysis revealed that 234 genes were at least 2.0-fold differentially expressed in the mutant. Particularly, genes involved in protein biosynthesis, carbohydrate and lipid metabolism, and replication and maintenance of DNA were downregulated. However, the most important change was the upregulation of the dlt operon, which is induced by the accumulation of intracellular Pi. The GraXRS system, known as an activator of the dlt operon (d-alanylation of teichoic acids) and of the mprF gene (multiple peptide resistance factor), is not involved in DAP tolerance of the pitA6 mutant. In conclusion, DAP tolerance of the pitA6 mutant is due to an upregulation of the dlt operon, triggered directly or indirectly by the accumulation of Pi.

INTRODUCTION

Daptomycin (DAP) is a cyclic lipopeptide antibiotic that was approved by the Food and Drug Administration (FDA) for treatment of complicated skin infections and bacteremia, as well as endocarditis caused by Gram-positive bacteria (1). DAP is highly efficient not only against growing cells but also against stationary-phase cultures of Staphylococcus aureus, including methicillin-resistant S. aureus (MRSA) strains, and is used as a last-resort antibiotic (2). Interestingly, a MIC creep for DAP has not been observed since its approval in 2003 (3); however, the number of cases with DAP therapy failure due to DAP-resistant (DAP-R) S. aureus strains has increased (1, 4).

Daptomycin is known to bind to the bacterial membranes of Gram-positive bacteria and kills the cell by disrupting the proton motive force (1). It is known that most of the adaptations in DAP-R strains concern the bacterial cell membrane and cell wall (5); however, the precise mode of action of DAP is still enigmatic.

In addition to antibiotic resistance, antibiotic tolerance can also be responsible for the failure of antibiotic therapy (4, 6). In contrast to resistant cells, tolerant strains are unable to grow in the presence of DAP; however, they survive the bactericidal action of DAP at least for a restricted period of time (7).

Antibiotic tolerance can be categorized into phenotypic and genetic tolerance. Phenotypic tolerance is independent from genetic alterations and becomes apparent, for instance, in bacterial persister cells, representing a tolerant subpopulation within isogenic bacterial populations (8, 9). Persisters are dormant or nongrowing, and their tolerance extends to different kind of antibiotics (6). Drug indifference is another example of phenotypic tolerance which is distributed, however, over the entire population and is caused by the inhibition of proliferation due to nutrient starvation (9, 10). Nonproliferating stationary-phase cells that are less sensitive toward treatment with most antibiotics can be named as a well-known example of drug indifference. In comparison, genetic tolerance is caused by genetically manifested and vertically transmitted mutations (8). All types of tolerance share the fact that an increase in the MIC does not occur. Although genetic antibiotic tolerance has been known for several decades, little is known about its generation (11).

In a recent study, we selected a mutant of S. aureus strain HG003 with a 10,000-fold-increased DAP tolerance (7). The tolerance of the corresponding mutant HG003pitA6 differs from that of the wild-type strain as it fulfills all criteria of genetic tolerance with a rather slow but linear decrease of viability during DAP treatment (8, 12). In contrast, the wild-type strain shows a biphasic killing behavior after the addition of DAP, which is typical of phenotypic tolerance by persister cells (6, 12). The DAP tolerance of HG003pitA6 was shown to be restricted to the stationary growth phase and associated with an increased uptake of inorganic phosphate (Pi), which was stored as polyphosphate (polyP) (7). We showed that the genetic basis of this increased DAP tolerance is represented by an adaptive single point mutation (pitA6) in the Pi transporter-encoding gene, pitA, which is part of the pitRA operon.

In this follow-up study, we found that heterologous expression of exopolyphosphatase (PPX) in the tolerant strain HG003pitA6 diminishes polyP to a wild-type level; however, the intracellular Pi level and growth phase-dependent DAP tolerance were not affected. Transcriptome analysis followed by quantitative real-time PCR (qRT-PCR) revealed that the dlt operon is upregulated in the pitA6 mutant, which is most likely responsible for the DAP tolerance. The responsibility of the dlt operon in the pitA6 DAP tolerance phenotype was confirmed as the deletion of dltA completely eliminated DAP tolerance caused by the pitA6 allele.

MATERIALS AND METHODS

Bacterial strains, plasmids, and culture conditions.

All bacterial strains used in this study are listed in Table S2 in the supplemental material. Escherichia coli DC10B (13) was used for molecular cloning of plasmids pMAD (14) and pRAB11 (15). E. coli cells bearing pMAD, pRAB11, or their derivatives were grown at 37°C in basic medium (BM; 1% [wt/vol] soy peptone, 0.5% [wt/vol] yeast extract, 0.5% [wt/vol] NaCl, 0.1% [wt/vol] K2HPO4, 0.1% [wt/vol] glucose), supplemented with 100 μg/ml ampicillin (Carl Roth). S. aureus cells with episomal pMAD plasmids were incubated at 30°C in BM or tryptic soy broth (TSB; 1.7% [wt/vol] peptone from casein, pancreatic digest, 0.3% [wt/vol] soy peptone, 0.5% [wt/vol] NaCl, 0.25% [wt/vol] K2HPO4, 0.25% [wt/vol] glucose, pH 7.3 ± 0.2) supplemented with 2.5 μg/ml erythromycin, and those carrying pRAB11 were incubated at 37°C in medium supplemented with 10 μg/ml chloramphenicol (Carl Roth).

Molecular cloning and DNA isolation.

Genomic DNA from S. aureus was isolated as described previously (16). Qiagen kits were used for isolation of bacterial plasmids. Successful cloning of plasmid constructs was verified by colony PCR and sequencing (GATC Biotech). RbCl was used for preparation of chemically competent E. coli cells (17), and electrocompetent cells of S. aureus were prepared as described previously by Mechler et al. (7).

Allelic exchange constructs and homologous recombination.

We used Gibson assembly (18) for cloning of the constructs for chromosomal deletion of genes dltA, mprF, graR, and graS in S. aureus HG003 wild type and HG003pitA6. DNA from S. aureus HG003 was used as a template. The primer pairs LM_graR_up_f /LM_graR_up_r and LM_graR_down_f /LM_graR_down_r (see Table S3 in the supplemental material for oligonucleotides) were used to amplify the upstream and downstream regions of graR in S. aureus HG003. The deletion region for graR includes 9 bp upstream from the start codon of graR, including parts of the putative ribosomal binding site. graXS fulfills all requirements for functionality in the corresponding strains. The primer pairs LM_graSup1/LM_graSup2 and LM_graSdown1/LM_graSdown1 were used for the deletion construct of graS. Regions consisting of 12 bp encoding the N-terminal part and 21 bp encoding the C terminus remained, resulting in functional graR and graX. For dltA deletion, the primer pairs LM_dlt_up1/LM_dlt_up2 and LM_dlt_down1/LM_dlt_down2 were used for amplification of regions bracketing dltA, leaving dltX, dltB, dltC, and dltD intact. The deletion part contains 10 bp upstream of dltA, disrupting the putative Shine-Dalgarno (SD) sequence, but retains 21 bp encoding the N terminus and encompassing the putative SD sequence of dltB. For deletion of mprF, the primer pairs LM_mprFup1/LM_mprFup2 and LM_mprFdown1/LM_mprFdown2 were used. In this construct, 6 bp encoding the N terminus and 16 bp encoding the C terminus remain, resulting in a frameshift. For cloning of all deletion constructs, the vector pMAD was digested with the enzymes BglII and SalI, and PCR products were incubated in a 2.5-fold excess of the plasmid backbone for 60 min at 50°C, in a final volume of 20 μl of reaction mixture (0.08 U of T5 exonuclease, 0.5 U of Phusion DNA polymerase, 80 U of Taq DNA ligase [all from New England BioLabs]) and 4 μl of 5× isothermal reaction buffer (25% [wt/vol] polyethylene glycol [PEG] 8000, 500 mM Tris-HCl [pH 7.5], 50 mM MgCl2, 50 mM dithiothreitol [DTT], 1 mM concentrations of each of the four deoxynucleoside triphosphates [dNTPs], and 5 mM NAD). The procedure of homologous recombination was performed as described previously by Mechler et al. (7).

Inducible expression using plasmid pRAB11.

The exopolyphosphatase (ppx) gene, A284_02120, including its native ribosome-binding sequence, was PCR amplified from genomic DNA of Staphylococcus warneri (19), using the primer pair ppx(warneri)fw and ppx(warneri)rev. Plasmid pRAB11 was digested with BglII. Again, Gibson assembly gave rise to pRAB11-ppx, harboring ppx downstream of the anhydrotetracycline (ATc)-inducible promoter Pxyl-tet. pRAB11-ppx was finally used to transform HG003, as well as HG003pitA6. For heterologous expression and downstream experiments, strains carrying pRAB11-ppx were inoculated to an optical density at 578 nm (OD578) of 0.07 and were grown for 2 h in TSB at 37°C with agitation before the addition of ATc (0.4 μM). Incubation was continued for a total of 16 h, and then polyP levels, intracellular Pi levels, and DAP tolerance were determined.

Determination of MIC and antibiotic tolerance.

MIC values were determined as described previously (20). The MIC of DAP is 1 μg/ml for both HG003 and HG003pitA6. For antibiotic tolerance assays, cells were grown for 16 h at 37°C in TSB with agitation. Afterwards, 2.24 ml of the overnight culture was transferred to 14-ml round-bottom tubes. DAP and CaCl2 were added to final volumes of 100 μg/ml and 50 μg/ml, respectively. The number of CFU/milliliter was determined at time points indicated in the figures.

Intracellular inorganic phosphate (Pi) and polyphosphate (polyP) determination.

Intracellular Pi was quantified as described previously (7) using a commercially available colorimetric approach (ab65622; Abcam). In brief, cells were grown at 37°C in TSB with agitation, harvested at time points indicated in the figures, washed with double-distilled H2O (ddH2O), and disrupted using a FastPrep (MP Biomedicals) instrument. Cell debris was pelleted, and supernatant was used to quantify Pi according to the manual of the colorimetric approach (ab65622; Abcam). Intracellular polyP was determined using 4′-6-diamidino-2-phenylindole (DAPI; Sigma) as described previously (7, 21, 22). Again, cells were grown in TSB at 37°C with agitation before being harvested and washed twice in Tris-HCl buffer (100 mM Tris; pH 7.5). Samples were adjusted to an OD578 of 0.5, and DAPI was added to a final concentration of 20 μM. The polyP fluorescence signal was determined using a microplate reader (TECAN Infinite 200M) with excitation at 415 nm and an emission wavelength of 550 nm.

RNA isolation for microarray and qRT-PCR.

RNA for microarray and qRT-PCR was isolated as described previously (7). In brief, cells were adjusted to an OD578 of 0.07 in TSB and cultured at 37°C. At the time points indicated in the figures, cells were harvested and adjusted to an OD578 of 70 before lysis with 1 ml Trizol (Life Technologies). DNA was removed by 2-fold DNase I digestion as described previously (23). A pool of 3 μg of total RNA for each condition was reverse transcribed using SuperScript II (Invitrogen, Basel, Switzerland). A volume of 5 μl of a 1:100-fold dilution was used for quantitative PCRs (qPCRs). Gene-specific primers and 6-carboxyfluorescein (FAM)-coupled probes for hu and mprF (see Table S4 in the supplemental material) were designed using Primer Express, version 3.0 (Applied Biosystems), and mixed in qPCR reagents (ABgene). Other genes were quantified by using a Brilliant SYBR Green master mix (Agilent) with primers described in Table S4 in the supplemental material. Reactions were performed in a Bio-Rad CFX96 and normalized using intensity levels recorded for the hu gene as described previously (24).

Microarray manufacturing and microarray design.

The microarray was manufactured by in situ synthesis of 60-base-long oligonucleotide probes (Agilent), selected as previously described (25). The microarray consists of a glass slide with 15,000 printed oligonucleotides, covering >95% of all open reading frames (ORFs) annotated in strains NCTC 8325 (26), UAMS-1 (27), and SA564 (28) as well as Newman (29), including their respective plasmids.

Preparation of labeled nucleic acids for expression microarrays.

Total RNA was purified from two independent cultures from strain HG003 and its pitA6 mutant. After additional DNase treatment, the absence of DNA traces was confirmed by quantitative PCR (SDS 7700; Applied Biosystems) with assays specific for 16S rRNA (30). Batches of 5 μg of total S. aureus RNA were labeled by Cy3-dCTP, using SuperScript II (Invitrogen) according to the manufacturer's instructions. Labeled products were then purified onto QiaQuick columns (Qiagen). Purified genomic DNA was extracted (DNeasy; Qiagen) from the different sequenced strains used for the design of the microarray, labeled with Cy5-dCTP using the Klenow fragment of DNA polymerase I (BioPrime), and used for the normalization process (31). Cy5-labeled DNA (500 ng) and a Cy3-labeled cDNA mixture were diluted in 50 μl of Agilent hybridization buffer and hybridized at a temperature of 60°C for 17 h in a hybridization oven (Robbins Scientific). Slides were washed, dried under nitrogen flow, and scanned (Agilent) using 100% photon multiplier tube power for both wavelengths.

Microarray analysis.

Fluorescence intensities were extracted using Feature Extraction software (version 9; Agilent). Local background-subtracted signals were corrected for unequal dye incorporation or unequal loading of the labeled product. The algorithm consisted of a rank consistency filter and a curve fit using the default LOWESS (locally weighted linear regression) method. Data consisting of three independent biological experiments were expressed as log10 ratios and analyzed using GeneSpring, version 8.0 (Silicon Genetics). A filter was applied to select oligonucleotides mapping ORFs in the Newman genome, yielding approximately 95% coverage. Statistical significance of differentially expressed genes was calculated by analysis of variance (32) using GeneSpring, including the Benjamini-Hochberg false discovery rate correction of 5% (P value cutoff, 0.05) and an arbitrary cutoff of 2-fold for expression ratios.

FITC–poly-l-lysine binding assay.

To determine the relative surface charge, overnight cultures were washed twice with HEPES buffer (20 mM, pH 7.25) and resuspended to an OD578 of 0.1. fluorescein isothiocyanate (FITC)–poly-l-lysine (FPLL) was added to a final concentration of 5 μg/ml, and the percentage of bound FPLL was determined as described previously (33, 34).

Microarray data accession numbers.

The complete microarray data set has been deposited in the Gene Expression Omnibus database (http://www.ncbi.nlm.nih.gov/geo/) under accession numbers GSE77069 and GPL10537 (for platform design).

RESULTS

DAP tolerance of HG003pitA6 is independent from polyP.

Previously, we selected a mutant of S. aureus HG003 which was extremely tolerant against DAP. The corresponding strain HG003pitA6 further showed an increased level of intracellular inorganic phosphate (Pi) and inorganic polyphosphate (polyP) (Fig. 1) (7). We heterologously expressed the exopolyphosphatase gene (ppx) from Staphylococcus warneri using the inducible plasmid pRAB11 to artificially hydrolyze the polyP level in both the HG003 wild type and mutant HG003pitA6 (Fig. 1A). Overexpression of ppx (Fig. 1, +ppx) in HG003pitA6 decreased polyP to a basal wild-type level (Fig. 1A). The Pi level and DAP tolerance, however, remained high in HG003pitA6 irrespective of ppx expression (Fig. 1B and C). Heterologous expression of ppx in HG003 wild type did not influence the polyP or intracellular Pi level or DAP tolerance (Fig. 1A to C).

FIG 1.

FIG 1

Open in a new tab

DAP tolerance of HG003pitA6 is independent of polyP. Graphs represent the following: quantification of polyP levels in DAPI-stained cells with excitation at 415 nm and emission at 550 nm (A), intracellular Pi concentration (B), and killing kinetics of stationary-phase cultures treated with 100 μg/ml DAP (C). Strains harboring the empty plasmid are indicated by −ppx, whereas those expressing ppx are indicated by +ppx. All values are averages from at least three independent experiments. Error bars indicate the standard deviations, and significance compared to results with HG003 with the empty plasmid pRAB11 was determined using Student's t test. Asterisks indicate P values of <0.05. wt, wild type; RFU, relative fluorescence units.

Transcriptome analysis disclosed predominant downregulation of genes from several functional categories.

We performed transcriptome analysis using the microarray technique to better understand molecular processes affecting the increase of DAP tolerance in HG003pitA6.

We showed previously that DAP tolerance and increased Pi level were restricted to the stationary growth phase (7). We therefore compared relative gene expression of HG003pitA6 with that of the wild type after 6 h (early stationary phase) and 8 h (advanced stationary phase) of growth. The complete results are summarized in Table S1 in the supplemental material. A total of 234 genes were found to be at least 2.0-fold differentially expressed in HG003pitA6 compared to expression in the HG003 wild-type strain after 6 h and/or 8 h of growth.

The majority of 170 genes were downregulated, in contrast to 65 genes that were upregulated (Fig. 2A). In general, differences in gene expression between the two strains were mostly moderate, in a range of 2- to 5-fold (see Table S1 in the supplemental material); however, downregulation was common in most functional categories (Fig. 2B). Predominantly genes involved in translation (100%), transcription (90%), and amino acid (65%) and nucleotide (67%) metabolism, as well as posttranslational modification (82%), were repressed in HG003pitA6 (Fig. 2B; see also Table S1). The percentage reflects the portion of downregulated genes in the corresponding functional category. Furthermore, a high number of repressed genes are involved in carbohydrate metabolism (77%), energy production (100%), or lipid transport (100%).

FIG 2.

FIG 2

Open in a new tab

Transcriptome analysis of genes differently expressed in HG003pitA6 compared to expression in the HG003 wild-type strain. (A) Venn diagrams showing numbers of genes that are at least 2.0-fold up- or downregulated after 6 h and/or 8 h of growth in TSB. (B) COG (clusters of orthologous groups of proteins) functional categories with the number of genes being at least 2.0-fold up- or downregulated.

Interestingly, many genes associated with the synthesis of the cell envelope, which was shown before to be affected in DAP-nonsusceptible strains (35–38), are dysregulated, with no clear tendency to up- or downregulation (Fig. 2B). Among them are the murC, murD, and murG genes that are 2- to 4-fold upregulated in HG003pitA6 (see Table S1 in the supplemental material). These genes are involved in the synthesis of peptidoglycan (39). However, murA, which represents the first step in peptidoglycan synthesis (39), was found to be 2-fold downregulated in our approach (see Table S1).

The dlt operon is upregulated in the HG003pitA6 mutant.

The mprF gene and the dlt operon are well-known hot spots with increased activity in DAP-nonsusceptible strains (35, 36, 40). Table 1 shows the relative expression profiles of mprF and the dlt operon during log and stationary phases. Neither mprF nor the dlt operon was upregulated during exponential growth according to quantitative real-time PCR (qRT-PCR) data. In contrast, the dlt operon was moderately downregulated. Both qRT-PCR and microarray analyses showed an upregulation of the dlt operon in stationary phase, in contrast to mprF, which was mildly downregulated.

TABLE 1.

Expression profiles of mprF and dltABCD in HG003pitA6 compared to those in HG003

Method of analysisa Relative expression (HG003pitA6 vs HG003)
dltA dltB dltC dltD mprF
qRT-PCR (4 h) −1.40 −1.27 −1.41 −1.05
qRT-PCR (8 h) 1.20 3.10 2.70 2.70 −1.84
Microarray (8 h) 2.40 −1.52
Open in a new tab
a

Times in parentheses refer to growth periods.

Deletion of dltA removes DAP tolerance in HG003pitA6.

As shown in Fig. 3A, markerless deletion of mprF from the HG003 wild-type strain and the HG003pitA6 mutant had only minor effects on DAP susceptibility of both strains in the time-kill curve. The additional deletion of dltA, however, increased the susceptibility of both wild-type HG003 and mutant HG003pitA6. This is reflected by the complete sterilization of the HG003pitA6 culture after 8 to 24 h of DAP treatment (Fig. 3B) and by an even faster eradication of the wild-type culture within 5 h of exposure to the drug. The DAP MIC was decreased in all deletion mutants (0.125 μg/ml) compared to the MICs of the HG003 wild-type strain and HG003pitA6 with intact mprF and dltA alleles (1 μg/ml).

FIG 3.

FIG 3

Open in a new tab

Time-dependent killing of ΔmprF and ΔmprF ΔdltA strains by DAP. CFU counts were determined for stationary-growth-phase cultures of the HG003 ΔmprF and HG003 ΔmprF ΔdltA strains (A) and for the HG003pitA6 ΔmprF and HG003pitA6 ΔmprF ΔdltA strains (B) upon challenge with 100 μg/ml DAP. Values represent the arithmetic means from three independent experiments. Error bars represent standard deviations.

Surface charge is unaffected in HG003pitA6.

An increase in the surface charge was shown earlier to be a possible phenotype for the upregulation of the dlt operon (33); however, no differences in the relative surface charges of the HG003 wild-type and HG003pitA6 strains could be detected when the FITC-poly-l-lysine (FPLL) approach was used (Fig. 4).

FIG 4.

FIG 4

Open in a new tab

Relative surface charge of HG003 and HG003pitA6. The percentage of FPLL bound to HG003 and HG003pitA6 grown for 16 h in TSB and incubation of cells with FPLL for 10 min at room temperature are depicted. All values are averages from at least three independent experiments. Error bars indicate the standard deviations, and significance was determined using Student's t test (P < 0.05). n.s., not significant.

Upregulation of the dlt operon is independent from GraXRS.

The GraXRS-VraFG five-component system is known to increase the expression of both mprF and the dlt operon (41). We found, however, that neither the markerless deletion of graR nor that of graS in either wild-type HG003 or HG003pitA6 was able to alter DAP tolerance (Fig. 5).

FIG 5.

FIG 5

Open in a new tab

Time-dependent killing of S. aureus strains with markerless deletion of graR or graS by DAP. Stationary-growth-phase cultures of the HG003 ΔgraS, HG003 ΔgraR, HG003pitA6 ΔgraS, and HG003pitA6 ΔgraR strains were challenged with 100-fold the MIC of DAP, and CFU counts were recorded. Values are the averages from three independent experiments. Error bars represent standard deviations.

DISCUSSION

A fundamental understanding of the mechanisms by which bacteria are able to escape bactericidal activity of antibiotics is essential for the development of new and possibly better antimicrobial therapies. Antibiotic tolerance is one aspect explaining the recalcitrance of relapsing and chronic infection by genetically susceptible bacteria (6). In an earlier study, we developed a strain showing decreased levels of DAP susceptibility in stationary phase (7). This phenotype differed from classical resistance since there was no increase in the MIC. The only adaptation required to decrease DAP susceptibility was a nonsynonymous mutation within the inorganic phosphate transporter gene pitA. Additionally, we showed that this kind of DAP tolerance in the mutant S. aureus HG003pitA6 was restricted to the stationary growth phase and correlated with an increased level of intracellular Pi and polyP. Previous studies by Gerdes and Maisonneuve demonstrated that the accumulation of polyP increased antibiotic tolerance by persister cells in E. coli (42). As polyP levels were also increased in the mutant HG003pitA6 (Fig. 1A), we wanted to elucidate a potential role of increased amounts of polyP in the DAP tolerance phenotype. PolyP is synthesized by polyphosphate kinases (PPKs) and hydrolyzed by exopolyphosphatases (PPXs) (43). S. aureus HG003 contains a putative exopolyphosphatase gene (SAOUHSC_01812) and a putative inorganic polyphosphate/ATP-NAD kinase gene (SAOUHSC_00943), but they are poorly characterized and different from the well-known ppk-ppx operon of E. coli (44). For heterologous expression, we decided to use the S. warneri ppx gene (A284_02120), which is a homologue of ppx from E. coli but closely related to the S. aureus gene (45). According to expectations (43), inducible overexpression of ppx reduced the amount of polyP in HG003pitA6/pRAB11-ppx to a basal wild-type level (Fig. 1A). Concentrations of intracellular Pi and the increase in DAP tolerance, however, remained unaffected (Fig. 1B and C), a fact clearly indicating that DAP tolerance mediated by the pitA6 allele is independent from polyP.

Transcriptome analysis using a microarray technique was applied to unravel the cellular basis of the HG003pitA6 phenotype (Fig. 2A and B; see also Table S1 in the supplemental material). As the increase in DAP tolerance is restricted to the stationary phase (7), we checked the expression of HG003pitA6 relative to that of the HG003 wild-type strain after 6 h (early stationary phase) and 8 h (advanced stationary phase) of growth.

Among a total of 234 genes that were at least 2.0-fold differentially expressed after 6 h and/or 8 h (Fig. 2A; see also Table S1 in the supplemental material), 65 were significantly upregulated, and 170 were repressed, in most cases with moderate changes (2- to 5-fold). A broad range of differentially expressed genes from central metabolic function was observed, with a predominant downregulation in most functional categories (Fig. 2B). A prevalent downshift of expression profiles for genes being involved in translation, transcription, amino acid metabolism, nucleotide metabolism, or posttranslational modification (Fig. 2B; see also Table S1) indicates that protein biosynthesis is low in HG003pitA6. Likewise, downregulation of genes implicated in carbohydrate or lipid metabolism and especially those responsible for energy generation suggests that a metabolic downshift prevails in the mutant HG003pitA6 at stationary growth phase. This is endorsed by the fact that HG003pitA6 is impaired in growth in comparison to HG003 wild type as soon as it reaches stationary phase (7). An increase in the cell wall thickness is assumed to be one reason for decreased susceptibility to DAP (36−38, 46), and indeed we found several genes involved in cell wall, membrane, and envelope maintenance or synthesis to be significantly upregulated. Among them were murC, murD, and murG (Fig. 2B; see also Table S1), which are responsible for the synthesis of peptidoglycan, a major component of the bacterial cell wall (39). However, MurA executes the first step in peptidoglycan synthesis (39) and is 2-fold downregulated in our data set, which contradicts an increased cell wall thickness phenotype. Accordingly, differences in cell wall thickness of the HG003 wild-type and HG003pitA6 strains were not detected by transmission electron microscopy (see Fig. S1 in the supplemental material). Furthermore, the content of wall teichoic acids (WTA) does not differ significantly in these strains (see Fig. S2), indicating that DAP tolerance is independent from alteration of cell wall thickness.

Beside an increase in thickness, alteration in composition of the bacterial cell wall as well as the membrane is a common adaptation in DAP-R mutants (1, 5). Genes involved in these processes and well-known hot spots for modification in DAP-nonsusceptible strains are mprF and the dltXABCD operon (1, 35, 36).

The multipeptide resistance factor gene mprF transfers lysyl from an l-lysyl-tRNA to membrane-bound phosphatidylglycerol, forming lysyl-phosphatydylglycerol (LPG) (47). The dlt operon is known to link positively charged d-alanine to both lipid and wall teichoic acids (40). Enhanced expression or activity of either or both is known to cause DAP nonsusceptibility (35, 36, 40, 48).

We found a time-dependent upregulation of the dlt operon but not the mprF gene in HG003pitA6 compared to the level in the wild type that could be linked to an increased level of intracellular Pi and DAP tolerance (Table 1 and Fig. 1B and C) (7). In accordance with previous studies (49), we found that the markerless deletion of mprF in both wild-type HG003 and mutant HG003pitA6 increased the general susceptibility for these strains, with a MIC of DAP of 0.125 μg/ml in contrast to a MIC of 1 μg/ml for HG003 and HG003pitA6 with an intact mprF gene. However, no dramatic increase in DAP susceptibility was detectable in time-kill studies in any of the strains (Fig. 3A). Interestingly, the subsequent deletion of dltA did not further decrease the MIC of DAP but completely reversed tolerance for both the HG003 ΔmprF ΔdltA and HG003pitA6 ΔmprF ΔdltA strains, with a complete sterilization within 24 h of DAP treatment (Fig. 3A and B).

According to our results, the dlt operon seems to be necessary for both the basal tolerance of the wild-type strain and the increased DAP tolerance of HG003pitA6. The growth phase-dependent upregulation of the dlt operon in the mutant HG003pitA6 is therefore the most plausible explanation for the increase in DAP tolerance in the HG003pitA6 strain. Nevertheless, it should be mentioned that many more genes were dysregulated in HG003pitA6 according to our microarray results, and their possible involvement in DAP tolerance was not further investigated and thus cannot be excluded entirely.

It further remains an open question as to how the upregulation of dlt is mediated. The five-component system GraXRS-VraFG is known to increase the expression of both mprF and the dlt operon (41, 50). We found, however, that the deletion of graR or graS, which will disrupt the function of the system (50), had no effect on DAP tolerance in the HG003 wild-type strain or HG003pitA6 (Fig. 5), an observation indicating that the regulation is mediated by a different mechanism.

The prevailing hypothesis of how dlt decreases DAP susceptibility is that there is an increase in the relative surface charge due to an increased d-alanylation of teichoic acids, resulting in repulsion of cationic peptides like DAP (48). Previous studies suggested that increased dlt expression would possibly, but not necessarily, increase the surface charge (36). Our experiments using HG003pitA6 support these findings as higher expression in HG003pitA6 of dlt did not produce an increase in the net charge (Fig. 4), clearly indicating that dlt contributes to a DAP-nonsusceptible phenotype by a means other than an increase in the cell surface charge.

According to our results, we propose a graphical summary of the DAP tolerance associated with pitA6 in Fig. 6. We conclude that the growth phase-dependent increase of intracellular Pi and DAP tolerance are linked to the upregulation of the dlt operon. As this was shown before to be causative for DAP nonsusceptibility (35, 36, 48), it is reasonable to argue that dlt overexpression is causative for DAP tolerance of HG003pitA6, which once more emphasizes the potential role of this operon in DAP nonsusceptibility.

FIG 6.

FIG 6

Open in a new tab

Putative model for DAP tolerance due to pitA6. A mutation in pitA generating PitA6 increases intracellular Pi levels and polyP levels. Elevated concentrations of intracellular Pi directly or indirectly promote the upregulation of the dlt operon, which is responsible for DAP tolerance. PitR is a PhoU-like protein that interacts with itself to most likely form dimers. The precise function of PitR is not known; however, it seem to act as a helper protein for phosphate transport and is mandatory for increased intracellular amounts of Pi and DAP tolerance in HG003pitA6.

Supplementary Material

Supplemental material
supp_60_5_2684__index.html (815B, html)

ACKNOWLEDGMENTS

We thank Panagiotis Papadopoulos for the excellent technical assistance.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Footnotes

Supplemental material for this article may be found at http://dx.doi.org/10.1128/AAC.03022-15.

REFERENCES

  • 1.Humphries RM, Pollett S, Sakoulas G. 2013. A current perspective on daptomycin for the clinical microbiologist. Clin Microbiol Rev 26:759–780. doi: 10.1128/CMR.00030-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Mascio CT, Alder JD, Silverman JA. 2007. Bactericidal action of daptomycin against stationary-phase and nondividing Staphylococcus aureus cells. Antimicrob Agents Chemother 51:4255–4260. doi: 10.1128/AAC.00824-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Sancak B, Yagci S, Mirza HC, Hascelik G. 2013. Evaluation of vancomycin and daptomycin MIC trends for methicillin-resistant Staphylococcus aureus blood isolates over an 11 year period. J Antimicrob Chemother 68:2689–2691. doi: 10.1093/jac/dkt247. [DOI] [PubMed] [Google Scholar]
  • 4.Mulcahy LR, Burns JL, Lory S, Lewis K. 2010. Emergence of Pseudomonas aeruginosa strains producing high levels of persister cells in patients with cystic fibrosis. J Bacteriol 192:6191–6199. doi: 10.1128/JB.01651-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Bayer AS, Schneider T, Sahl HG. 2013. Mechanisms of daptomycin resistance in Staphylococcus aureus: role of the cell membrane and cell wall. Ann N Y Acad Sci 1277:139–158. doi: 10.1111/j.1749-6632.2012.06819.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Lewis K. 2010. Persister cells. Annu Rev Microbiol 64:357–372. doi: 10.1146/annurev.micro.112408.134306. [DOI] [PubMed] [Google Scholar]
  • 7.Mechler L, Herbig A, Paprotka K, Fraunholz M, Nieselt K, Bertram R. 22 June 2015. A novel point mutation promotes growth phase-dependent daptomycin tolerance in Staphylococcus aureus. Antimicrob Agents Chemother doi: 10.1128/AAC.00643-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Tuomanen E, Durack DT, Tomasz A. 1986. Antibiotic tolerance among clinical isolates of bacteria. Antimicrob Agents Chemother 30:521–527. doi: 10.1128/AAC.30.4.521. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Levin BR, Rozen DE. 2006. Non-inherited antibiotic resistance. Nat Rev Microbiol 4:556–562. doi: 10.1038/nrmicro1445. [DOI] [PubMed] [Google Scholar]
  • 10.Corona F, Martinez JL. 2013. Phenotypic resistance to antibiotics. Antibiotics 2:237–255. doi: 10.3390/antibiotics2020237. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Tomasz A, Albino A, Zanati E. 1970. Multiple antibiotic resistance in a bacterium with suppressed autolytic system. Nature 227:138–140. doi: 10.1038/227138a0. [DOI] [PubMed] [Google Scholar]
  • 12.NCCLS. 1999. Methods for determining bactericidal activity of antimicrobial agents: approved guideline. National Committee for Clinical Laboratory Standards, Wayne, PA. [Google Scholar]
  • 13.Monk IR, Shah IM, Xu M, Tan MW, Foster TJ. 2012. Transforming the untransformable: application of direct transformation to manipulate genetically Staphylococcus aureus and Staphylococcus epidermidis. mBio 3:e00277-11. doi: 10.1128/mBio.00277-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Arnaud M, Chastanet A, Debarbouille M. 2004. New vector for efficient allelic replacement in naturally nontransformable, low-GC-content, gram-positive bacteria. Appl Environ Microbiol 70:6887–6891. doi: 10.1128/AEM.70.11.6887-6891.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Helle L, Kull M, Mayer S, Marincola G, Zelder ME, Goerke C, Wolz C, Bertram R. 2011. Vectors for improved Tet repressor-dependent gradual gene induction or silencing in Staphylococcus aureus. Microbiology 157:3314–3323. doi: 10.1099/mic.0.052548-0. [DOI] [PubMed] [Google Scholar]
  • 16.Nolle N, Schuster CF, Bertram R. 2013. Two paralogous yefM-yoeB loci from Staphylococcus equorum encode functional toxin-antitoxin systems. Microbiology 159:1575–1585. doi: 10.1099/mic.0.068049-0. [DOI] [PubMed] [Google Scholar]
  • 17.Hanahan D. 1983. Studies on transformation of Escherichia coli with plasmids. J Mol Biol 166:557–580. doi: 10.1016/S0022-2836(83)80284-8. [DOI] [PubMed] [Google Scholar]
  • 18.Gibson DG, Young L, Chuang RY, Venter JC, Hutchison CA III, Smith HO. 2009. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 6:343–345. doi: 10.1038/nmeth.1318. [DOI] [PubMed] [Google Scholar]
  • 19.Wood CA. 1992. Significant infection caused by Staphylococcus warneri. J Clin Microbiol 30:2216–2217. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Lechner S, Lewis K, Bertram R. 2012. Staphylococcus aureus persisters tolerant to bactericidal antibiotics. J Mol Microbiol Biotechnol 22:235–244. doi: 10.1159/000342449. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Schurig-Briccio LA, Farias RN, Rintoul MR, Rapisarda VA. 2009. Phosphate-enhanced stationary-phase fitness of Escherichia coli is related to inorganic polyphosphate level. J Bacteriol 191:4478–4481. doi: 10.1128/JB.00082-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Aschar-Sobbi R, Abramov AY, Diao C, Kargacin ME, Kargacin GJ, French RJ, Pavlov E. 2008. High sensitivity, quantitative measurements of polyphosphate using a new DAPI-based approach. J Fluoresc 18:859–866. doi: 10.1007/s10895-008-0315-4. [DOI] [PubMed] [Google Scholar]
  • 23.Schuster CF, Bertram R. 2014. fluorescence based primer extension technique to determine transcriptional starting points and cleavage sites of RNases in vivo. J Vis Exp 92:e52134. doi: 10.3791/52134. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Garzoni C, Francois P, Huyghe A, Couzinet S, Tapparel C, Charbonnier Y, Renzoni A, Lucchini S, Lew DP, Vaudaux P, Kelley WL, Schrenzel J. 2007. A global view of Staphylococcus aureus whole genome expression upon internalization in human epithelial cells. BMC Genomics 8:171. doi: 10.1186/1471-2164-8-171. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Charbonnier Y, Gettler B, Francois P, Bento M, Renzoni A, Vaudaux P, Schlegel W, Schrenzel J. 2005. A generic approach for the design of whole-genome oligoarrays, validated for genomotyping, deletion mapping and gene expression analysis on Staphylococcus aureus. BMC Genomics 6:95. doi: 10.1186/1471-2164-6-95. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Gillaspy A, Worrell V, Orvis J, Roe B, Dyer D, Iandolo J. 2006. The Staphylococcus aureus NCTC 8325 genome, p 381–212. In Fischetti V, Novick R, Ferretti J, Portnoy D, Rood J (ed), Gram-positive pathogens. ASM Press, Washington, DC. [Google Scholar]
  • 27.Sassi M, Sharma D, Brinsmade SR, Felden B, Augagneur Y. 2015. Genome sequence of the clinical isolate Staphylococcus aureus subsp. aureus strain UAMS-1. Genome Announc 3:e01584-14. doi: 10.1128/genomeA.01584-14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Giraud C, Hausmann S, Lemeille S, Prados J, Redder P, Linder P. 2015. The C-terminal region of the RNA helicase CshA is required for the interaction with the degradosome and turnover of bulk RNA in the opportunistic pathogen Staphylococcus aureus. RNA Biol 12:658–674. doi: 10.1080/15476286.2015.1035505. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Baba T, Bae T, Schneewind O, Takeuchi F, Hiramatsu K. 2008. Genome sequence of Staphylococcus aureus strain Newman and comparative analysis of staphylococcal genomes: polymorphism and evolution of two major pathogenicity islands. J Bacteriol 190:300–310. doi: 10.1128/JB.01000-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Pohl K, Francois P, Stenz L, Schlink F, Geiger T, Herbert S, Goerke C, Schrenzel J, Wolz C. 2009. CodY in Staphylococcus aureus: a regulatory link between metabolism and virulence gene expression. J Bacteriol 191:2953–2963. doi: 10.1128/JB.01492-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Talaat AM, Howard ST, Hale Wt Lyons R, Garner H, Johnston SA. 2002. Genomic DNA standards for gene expression profiling in Mycobacterium tuberculosis. Nucleic Acids Res 30:e104. doi: 10.1093/nar/gnf103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Churchill GA. 2004. Using ANOVA to analyze microarray data. Biotechniques 37:173–175, 177. [DOI] [PubMed] [Google Scholar]
  • 33.Taneja NK, Ganguly T, Bakaletz LO, Nelson KJ, Dubey P, Poole LB, Deora R. 2013. d-Alanine modification of a protease-susceptible outer membrane component by the Bordetella pertussis dra locus promotes resistance to antimicrobial peptides and polymorphonuclear leukocyte-mediated killing. J Bacteriol 195:5102–5111. doi: 10.1128/JB.00510-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Mukhopadhyay K, Whitmire W, Xiong YQ, Molden J, Jones T, Peschel A, Staubitz P, Adler-Moore J, McNamara PJ, Proctor RA, Yeaman MR, Bayer AS. 2007. In vitro susceptibility of Staphylococcus aureus to thrombin-induced platelet microbicidal protein-1 (tPMP-1) is influenced by cell membrane phospholipid composition and asymmetry. Microbiology 153:1187–1197. doi: 10.1099/mic.0.2006/003111-0. [DOI] [PubMed] [Google Scholar]
  • 35.Cafiso V, Bertuccio T, Purrello S, Campanile F, Mammina C, Sartor A, Raglio A, Stefani S. 2014. dltA overexpression: a strain-independent keystone of daptomycin resistance in methicillin-resistant Staphylococcus aureus. Int J Antimicrob Agents 43:26–31. doi: 10.1016/j.ijantimicag.2013.10.001. [DOI] [PubMed] [Google Scholar]
  • 36.Mishra NN, Bayer AS, Weidenmaier C, Grau T, Wanner S, Stefani S, Cafiso V, Bertuccio T, Yeaman MR, Nast CC, Yang SJ. 2014. Phenotypic and genotypic characterization of daptomycin-resistant methicillin-resistant Staphylococcus aureus strains: relative roles of mprF and dlt operons. PLoS One 9:e107426. doi: 10.1371/journal.pone.0107426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Bertsche U, Yang SJ, Kuehner D, Wanner S, Mishra NN, Roth T, Nega M, Schneider A, Mayer C, Grau T, Bayer AS, Weidenmaier C. 2013. Increased cell wall teichoic acid production and d-alanylation are common phenotypes among daptomycin-resistant methicillin-resistant Staphylococcus aureus (MRSA) clinical isolates. PLoS One 8:e67398. doi: 10.1371/journal.pone.0067398. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Bertsche U, Weidenmaier C, Kuehner D, Yang SJ, Baur S, Wanner S, Francois P, Schrenzel J, Yeaman MR, Bayer AS. 2011. Correlation of daptomycin resistance in a clinical Staphylococcus aureus strain with increased cell wall teichoic acid production and d-alanylation. Antimicrob Agents Chemother 55:3922–3928. doi: 10.1128/AAC.01226-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Typas A, Banzhaf M, Gross CA, Vollmer W. 2012. From the regulation of peptidoglycan synthesis to bacterial growth and morphology. Nat Rev Microbiol 10:123–136. doi: 10.1038/nrmicro2677. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Weidenmaier C, Peschel A, Kempf VA, Lucindo N, Yeaman MR, Bayer AS. 2005. DltABCD- and MprF-mediated cell envelope modifications of Staphylococcus aureus confer resistance to platelet microbicidal proteins and contribute to virulence in a rabbit endocarditis model. Infect Immun 73:8033–8038. doi: 10.1128/IAI.73.12.8033-8038.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Falord M, Karimova G, Hiron A, Msadek T. 2012. GraXSR proteins interact with the VraFG ABC transporter to form a five-component system required for cationic antimicrobial peptide sensing and resistance in Staphylococcus aureus. Antimicrob Agents Chemother 56:1047–1058. doi: 10.1128/AAC.05054-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Gerdes K, Maisonneuve E. 2012. Bacterial persistence and toxin-antitoxin loci. Annu Rev Microbiol 66:103–123. doi: 10.1146/annurev-micro-092611-150159. [DOI] [PubMed] [Google Scholar]
  • 43.Rao NN, Gomez-Garcia MR, Kornberg A. 2009. Inorganic polyphosphate: essential for growth and survival. Annu Rev Biochem 78:605–647. doi: 10.1146/annurev.biochem.77.083007.093039. [DOI] [PubMed] [Google Scholar]
  • 44.Akiyama M, Crooke E, Kornberg A. 1993. An exopolyphosphatase of Escherichia coli. The enzyme and its ppx gene in a polyphosphate operon. J Biol Chem 268:633–639. [PubMed] [Google Scholar]
  • 45.Lamers RP, Muthukrishnan G, Castoe TA, Tafur S, Cole AM, Parkinson CL. 2012. Phylogenetic relationships among Staphylococcus species and refinement of cluster groups based on multilocus data. BMC Evol Biol 12:171. doi: 10.1186/1471-2148-12-171. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Yang SJ, Nast CC, Mishra NN, Yeaman MR, Fey PD, Bayer AS. 2010. Cell wall thickening is not a universal accompaniment of the daptomycin nonsusceptibility phenotype in Staphylococcus aureus: evidence for multiple resistance mechanisms. Antimicrob Agents Chemother 54:3079–3085. doi: 10.1128/AAC.00122-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Ernst CM, Peschel A. 2011. Broad-spectrum antimicrobial peptide resistance by MprF-mediated aminoacylation and flipping of phospholipids. Mol Microbiol 80:290–299. doi: 10.1111/j.1365-2958.2011.07576.x. [DOI] [PubMed] [Google Scholar]
  • 48.Yang SJ, Kreiswirth BN, Sakoulas G, Yeaman MR, Xiong YQ, Sawa A, Bayer AS. 2009. Enhanced expression of dltABCD is associated with the development of daptomycin nonsusceptibility in a clinical endocarditis isolate of Staphylococcus aureus. J Infect Dis 200:1916–1920. doi: 10.1086/648473. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Cameron DR, Mortin LI, Rubio A, Mylonakis E, Moellering RC Jr, Eliopoulos GM, Peleg AY. 2015. Impact of daptomycin resistance on Staphylococcus aureus virulence. Virulence 6:127–131. doi: 10.1080/21505594.2015.1011532. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Yang SJ, Bayer AS, Mishra NN, Meehl M, Ledala N, Yeaman MR, Xiong YQ, Cheung AL. 2012. The Staphylococcus aureus two-component regulatory system, GraRS, senses and confers resistance to selected cationic antimicrobial peptides. Infect Immun 80:74–81. doi: 10.1128/IAI.05669-11. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental material
supp_60_5_2684__index.html (815B, html)
AAC.03022-15_zac005165098so1.pdf (376.3KB, pdf)

Articles from Antimicrobial Agents and Chemotherapy are provided here courtesy of American Society for Microbiology (ASM)

RESOURCES