ABSTRACT
Methicillin-resistant Staphylococcus aureus (MRSA) resistant to decolonization agents such as mupirocin and chlorhexidine increases the need for development of alternative decolonization molecules. The absence of reported severe adverse reactions and bacterial resistance to polyhexanide makes it an excellent choice as a topical antiseptic. In the present study, we evaluated the in vitro and in vivo capacity to generate strains with reduced polyhexanide susceptibility and cross-resistance with chlorhexidine and/or antibiotics currently used in clinic. Here we report the in vitro emergence of reduced susceptibility to polyhexanide by prolonged stepwise exposure to low concentrations in broth culture. Reduced susceptibility to polyhexanide was associated with genomic changes in the mprF and purR genes and with concomitant decreased susceptibility to daptomycin and other cell wall-active antibiotics. However, the in vitro emergence of reduced susceptibility to polyhexanide did not result in cross-resistance to chlorhexidine. During in vivo polyhexanide clinical decolonization treatment, neither reduced polyhexanide susceptibility nor chlorhexidine cross-resistance was observed. Together, these observations suggest that polyhexanide could be used safely for decolonization of carriers of chlorhexidine-resistant S. aureus strains; they also highlight the need for careful use of polyhexanide at low antiseptic concentrations.
KEYWORDS: methicillin-resistant Staphylococcus aureus, decolonization, MIC, antibiotic resistance, daptomycin, MRSA, polyhexanide
INTRODUCTION
Prevention of health care-associated infections includes the use of antiseptic agents. Chlorhexidine antiseptic solution has been one of the most widely used antiseptics since the 1950s and is administered for hand and skin disinfection prior to surgical intervention, bathing patients in intensive care units, decolonization of carriers of methicillin-resistant Staphylococcus aureus (MRSA), and prevention of vascular catheter infections (1). Broad range and long residual activity, safety and good tolerance are the key advantages of this antiseptic agent. However, reduced chlorhexidine susceptibility associated with biocide efflux pumps (1, 2) has been shown to impact clinical outcomes (3, 4).
An increase in reduced chlorhexidine susceptibility due to its intensive clinical use (5) has led to the development of new antiseptics such as polyhexanide (polyhexamethylene biguanide). This antiseptic was originally developed as a surface disinfectant, but in the early 1990s it was introduced into medicine for local antiseptic treatment (6) and is currently used in the United States for wound disinfection. Polyhexanide shows good safety, tissue compatibility, and reduction of bacterial load and infection rate of chronic and burn wounds and is proposed as an alternative to topical antibiotic treatment (7, 8).
Polyhexanide is a cationic polymer attaching primarily to negatively charged membrane phospholipids, interfering with their stability and leading to membrane permeability. Lipopolysaccharides and teichoic acids from Gram-negative and Gram-positive bacteria, respectively, and peptidoglycan components of the cell wall were also identified as polyhexanide targets (9–11). Accordingly, polyhexanide was shown to have potent antimicrobial activity against both Gram-positive and Gram-negative bacteria (8, 12). In contrast to the situation with other antiseptics, reduced susceptibility to polyhexanide and its association with antibiotic resistance has not yet been detected (12–16). Another exceptional characteristic recently identified for polyhexanide is its intracellular bactericidal activity, recognized as an important property for potentially treating skin infections caused by intracellular bacteria (17).
Polyhexanide lacks reported severe adverse reactions, selection of bacterial resistance, and antagonisms with antibiotic activities and has potential use as an intracellular bactericidal agent, making it an excellent choice as a topical antiseptic to prevent and treat bacterial infections. To test polyhexanide, we previously assessed its efficacy in eradicating MRSA carriage in vivo by a randomized placebo-controlled clinical trial (18), which showed that a single polyhexanide decolonization course was not sufficient to significantly eradicate MRSA carriage (18).
In the current microbiological study we analyzed the potential reasons that could explain low polyhexanide decolonization rates in vivo. We tested the in vitro emergence of polyhexanide resistance and the potential cross-resistance with chlorhexidine antiseptic and identified the genetic mutations potentially leading to reduced susceptibility to polyhexanide.
RESULTS
In vitro emergence of reduced polyhexanide susceptibility and cross-resistance with chlorhexidine.
To analyze the potential emergence of reduced polyhexanide susceptibility in vitro, we selected three different clinical MRSA strains (COL, 134947, and 128822) that were subjected to a stepwise training method in polyhexanide broth cultures (see Materials and Methods). As shown in Table 1, after several passages of 2 days each on increasing concentrations of polyhexanide, two out of three MRSA strains (with initial polyhexanide MICs of 0.5 and 1 μg/ml) ultimately grew at polyhexanide concentrations of 2, 4, and 8 μg/ml. Prolonged exposure did not increase further the levels of polyhexanide MICs.
TABLE 1.
Polyhexanide and chlorhexidine resistance profiles of in vitro selected antiseptic mutants
| Strain | Antiseptic selection | No. of passages | MIC (μg/ml) fora: |
|
|---|---|---|---|---|
| Polyhexanide | Chlorhexidine | |||
| COL | None | 0 | 0.5 | 2 |
| COL P3 | Polyhexanide | 3 | 2 | 2 |
| COL P5 | Polyhexanide | 5 | 4 | 2 |
| 134947 | None | 0 | 1 | 4 |
| 134947 P6 | Polyhexanide | 6 | 8 | 4 |
| 128822 | None | 0 | 1 | 8 |
| 128822 P6b | Polyhexanide | 6 | 1 | 8 |
| COL | None | 0 | 0.5 | 2 |
| COL P10 | Chlorhexidine | 10 | 0.5 | 8 |
| COL P7 | Chlorhexidine | 7 | 0.5 | 8 |
| 134947 | None | 0 | 1 | 4 |
| 134947 P10 | Chlorhexidine | 10 | 1 | 8 |
| 128822 | None | 0 | 1 | 8 |
| 128822 P10 | Chlorhexidine | 10 | 1 | 8 |
Polyhexanide and chlorhexidine MICs were measured by the macrodilution method.
Selection of polyhexanide stepwise mutants was impossible with this strain.
To further analyze cross-resistance development, we assessed the emergence of reduced chlorhexidine susceptibility in vitro, using the same methodology as described above for polyhexanide. After several passages of 2 days each on chlorhexidine, again two out of three MRSA strains (with initial chlorhexidine MICs of 2 and 4 μg/ml) ultimately grew at a chlorhexidine concentration of 8 μg/ml (Table 1). Emergence of reduced chlorhexidine susceptibility was not accompanied by changes in polyhexanide MICs (Table 1). Similarly, emergence of reduced polyhexanide susceptibility was not accompanied by changes in chlorhexidine MICs, suggesting an absence of cross-resistance between these antiseptics.
Antibiotic susceptibility profiles and genomic sequencing of MRSA strains with reduced susceptibility to antiseptics.
Previous studies have shown an association of chlorhexidine resistance with resistance to antibiotics (19). To determine whether in vitro emergence of reduced susceptibility to polyhexanide or chlorhexidine in our strains was associated with emergence of antibiotic resistance, we analyzed the antibiotic susceptibility pattern of each parental isolate and its cognate in vitro-derived polyhexanide- or chlorhexidine-exposed derivative showing altered susceptibility (COL/COL P3/COL P5, 134947/134947 P6, 128822/128822 P6, COL/COL P7/COL P10, 134947/134947 P10, 128822/128822 P10). Antibiotic disc diffusion and Etest assays showed that reduced susceptibility to polyhexanide was accompanied by changes in antibiotic susceptibility profiles (vancomycin, teicoplanin, daptomycin) compared to the parental strains (Table 2). Interestingly, strains showing reduced susceptibility to polyhexanide showed reduced susceptibility to daptomycin with or without concomitant alteration of susceptibility to vancomycin or teicoplanin. In contrast to the polyhexanide susceptibility profile, no consistent association between reduced susceptibility to chlorhexidine and antibiotic resistance was observed; of three strains with reduced chlorhexidine susceptibility, only one showed an association with reduced ciprofloxacin susceptibility, in agreement with previous observations (20). In a single background (MRSA strain 128822) showing a high initial MIC level against chlorhexidine (MIC = 8 μg/ml), we did not observe reduced susceptibility to daptomycin (Table 2).
TABLE 2.
Antibiotic resistance profiles and single nucleotide polymorphisms present in MRSA antiseptic-selected mutants compared to their corresponding wild-type strains
| Strain | Antiseptic selectiona | MIC (μg/ml) |
Antibiotic susceptibility pattern (MIC [μg/ml])b | ST | Gene name | SNPc | Amino acid change | Gene function | |
|---|---|---|---|---|---|---|---|---|---|
| Polyhex | Chlorhex | ||||||||
| COL | 0.5 | 2 | Penr, Oxar, ACr, Cefr, Vans (1.5), Tei (1.5), Dap (0.5) | 250 | |||||
| COL P3 | Polyhex | 2 | 2 | Penr, Oxar, ACr, Cefr, Vanr (3), Teir (3), Dapr (3) | 250 | mprF | C1010T | L337S | Phosphatidylglycerol lysyltransferase |
| COL P5 | Polyhex | 4 | 2 | Penr, Oxar, ACr, Cefr, Teir (4), Dapr (4) | 250 | mprF | C1010T | L337S | Phosphatidylglycerol lysyltransferase |
| 134947 | 1 | 4 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr | 228 | |||||
| 134947 P6 | Polyhex | 8 | 4 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr, Dapr (1.5–2) | 228 | purR | T686G | V229G | Pur operon repressor |
| mprF | C884T | S295L | Phosphatidylglycerol lysyltransferase | ||||||
| 128822 | 1 | 8 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr, Genr | 228 | |||||
| 128822 P6 | Polyhex | 1 | 8 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr, Genr | 228 | NDd | |||
| COL | 0.5 | 2 | Penr, Oxar, ACr, Cefr | 250 | |||||
| COL P10 | Chlorhex | 0.5 | 8 | Penr, Oxar, ACr, Cefr, Cipr (D17) | 250 | T to A | Intergenic region (position 775770) between SACOLSACOL_RS03870 and NorA | ||
| G to A | Intergenic region (position 814686) between SACOL_RS04065 and NrdL | ||||||||
| T to A | Intergenic region (position 1227842) upstream SACOL_RS06240 | ||||||||
| G to A | Intergenic region (position 1227910) upstream SACOL_RS06245SACOL_RS06245 | ||||||||
| COL P7 | Chlorhex | 0.5 | 8 | Penr, Oxar, ACr, Cefr | 250 | mepA | C127T | T376I | Multidrug export protein MepA |
| purR | G403T | V135F | Pur operon repressor | ||||||
| 134947 | 1 | 4 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr | 228 | |||||
| 134947 P10 | Chlorhex | 1 | 8 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr | 228 | ND | |||
| 128822 | 1 | 8 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr, Genr | 228 | |||||
| 128822 P10 | Chlorhex | 1 | 8 | Penr, Oxar, ACr, Cefr, Cipr, Clir, Eryr, Genr, Dapr (2) | 228 | pldB | G89T | G30V | Lysophospholipase L2 |
| glpD | C293A | T98K | Aerobic glycerol-3-phosphate dehydrogenase | ||||||
| mprF | C1001T | P334L | Phosphatidylglycerol lysyltransferase | ||||||
Polyhex, polyhexanide; Chlorhex, chlorhexidine.
Antibiotic susceptibility was measured by disc diffusion assays for all antibiotics except vancomycin, teicoplanin, and daptomycin (D corresponds to diameter size measurement; EUCAST ciprofloxacin susceptible diameter = 19–20). For vancomycin, teicoplanin, and daptomycin, MIC values were determined by Etest assays. EUCAST susceptibility breakpoints for are ≤2 μg/ml for vancomycin, ≤2 μg/ml for teicoplanin, and ≤1 μg/ml for daptomycin. Changes in antibiotic resistance patterns compared to those in wild-type strains are indicated by bold type. Pen, penicillin; Oxa, oxacillin; AC, amoxicillin-clavulanate; Cef, cefoxitine; Cip, ciprofloxacin; Cli, clindamycin; Ery, erythromycin; Gen, gentamicin; Van, vancomycin; Tei, teicoplanin; Dap, daptomycin; r, resistant; s, susceptible.
SNP, single-nucleotide polymorphism.
ND, not determined.
To identify genomic changes associated with reduced antiseptic/antibiotic susceptibility, we performed de novo whole-genome sequencing (see Materials and Methods). Using Illumina-Solexa technology, we obtained between 4,107,708 and 3,452,730 of 300-bp paired-end reads for all strains leading to between 140× and 287× of coverage depth after quality filtering. Genome assembly resulted in 2,821,361 bp for the S. aureus COL strain, 2,794,034 bp for S. aureus SA134947, and 2,922,225 bp for MRSA128822. After quality assessment, filtering, and genome assembly, single nucleotide polymorphisms, insertions, and deletions were identified between antiseptic-selected mutants and their cognate parents. As shown in Table 2, antiseptic-selected mutants showing changes in polyhexanide or chlorhexidine MICs possessed mutations in the mprF, purR, mepA, pldB, and glpD genes and in some intergenic regions near norA, ndrL, or other hypothetical genes. Interestingly, these genes affect lipid metabolism (mprF, pldB, and glpD) or are already known to affect resistance to chlorhexidine (MepA efflux pump) (20) or to daptomycin (21) and nisin, cationic antimicrobial peptide antibiotics (22) (MprF protein and PurR transcriptional activator, respectively). To establish the contribution of the observed purR mutation to polyhexanide resistance, the MIC of polyhexanide was determined against S. aureus SH1000:purR T686G, the construction of which is described elsewhere (22). No change in polyhexanide susceptibility was observed compared with that of either parental strain, SH1000 or COL, suggesting that this mutation does not contribute to the observed resistance phenotype. Given that nonsynonymous mprF mutations were identified (Table 2) in all strains displaying polyhexanide resistance, it seems likely that these mutations are responsible for resistance. MIC determination was also assessed in strains harboring mprFC884T identified in a different experimental context (23). This mutation leading to daptomycin resistance was responsible for a 2-fold increase in polyhexanide MIC (from 4 to 8 μg/ml). Note that we tried several times but failed to transfer an individual mutation into the parental strain COL by transduction using several staphylococcal phages.
Polyhexanide and chlorhexidine susceptibility profiles of MRSA isolates before and after polyhexanide decolonization.
Our previously published study suggested a limited efficacy of a single polyhexanide decolonization course in eradicating MRSA carriage (18). Despite several possible limitations of our study, one possible explanation was the emergence of resistance to polyhexanide or cross-resistance between chlorhexidine and polyhexanide antiseptics. Indeed, we previously reported that resistance to chlorhexidine in our hospital was associated with the dominant clone, the South German staphylococcal cassette chromosome mec element (SCCmec) I sequence type (ST) 228 MRSA (3). To monitor potential reduced polyhexanide and chlorhexidine susceptibility in our strain collection, we selected nasal MRSA strains isolated before (D0) and after active polyhexanide decolonization treatment (D28) (Table 3). Multilocus variable-number tandem-repeat analysis (MLVA) analysis was performed to confirm the clonal relationship between D0 and D28 bacterial strains isolated from the same patient and to deduce the ST of our strain collection (not shown). All selected pairs of strains isolated from the same patients were indeed clonally related and showed ST228 (n = 20), ST5 (n = 2), ST8 (n = 2), ST105 (n = 2), or ST22 (n = 1).
TABLE 3.
Chlorhexidine and polyhexanide MIC values of 54 MRSA clinical strains isolated before and after polyhexanide patient decolonizationa
| Strain no. (day 0)b | MIC (μg/ml) for: |
Strain no. (day 28)c | MIC (μg/ml) for: |
||
|---|---|---|---|---|---|
| Chlorhexidine | Polyhexanide | Chlorhexidine | Polyhexanide | ||
| 1 | 4 | 0.5 | 1 | 4 | 0.5 |
| 2 | <0.5 | 0.5 | 2 | 1/0.5 | 1 |
| 3 | 4 | 0.5 | 3 | 4 | 1 |
| 4 | <0.5 | 0.5 | 4 | <0.5 | 0.5 |
| 5 | 4 | 0.25 | 5 | 2/4 | 0.5 |
| 6 | 4 | 0.5 | 6 | 2 | 0.5 |
| 7 | 4 | 0.5 | 7 | 4 | 0.5 |
| 8 | 1 | 0.5 | 8 | 1 | 0.5 |
| 9 | <0.5 | 0.5 | 9 | <0.5 | 0.5 |
| 10 | 4 | 0.5 | 10 | 4 | 0.5 |
| 11 | 4 | 0.5 | 11 | 1 | 0.5 |
| 12 | 4 | 0.25 | 12 | 4 | 0.5 |
| 13 | 4 | 1 | 13 | <0.5 | 0.5 |
| 14 | 4 | 0.5 | 14 | 2 | 1 |
| 15 | 1 | 1 | 15 | 4 | 0.5 |
| 16 | 2 | 0.5–1 | 16 | 4 | 0.5 |
| 17 | <0.5 | 1 | 17 | <0.5 | 0.5 |
| 18 | 4 | 0.5 | 18 | 4 | 0.25 |
| 19 | 4 | 0.5 | 19 | 4 | 0.5 |
| 20 | 2 | 0.5 | 20 | 2 | 0.5 |
| 21 | 2 | 0.25 | 21 | 2 | 0.25 |
| 22 | <0.5 | 0.5 | 22 | 1 | 1 |
| 23 | 2 | 1 | 23 | 1 | 1 |
| 24 | 4 | 0.5 | 24 | 4 | 0.5 |
| 25 | 4 | 0.5 | 25 | 1 | 0.5 |
| 26 | 4 | 0.5 | 26 | 4 | 0.5 |
| 27 | 2 | 1 | 27 | 4 | 0.25 |
Bold type is used for pairs of strains showing the most important changes.
MRSA clinical strains isolated prior to polyhexanide decolonization (day 0) or after 28 days of polyhexanide treatment (day 28).
MRSA day 28 bacteria were clonally related to day 0 bacteria, isolated from the same patient.
Reduced susceptibility to polyhexanide and chlorhexidine was further measured using macrodilution MIC method. Our strain collection shows polyhexanide and chlorhexidine MICs ranging from 0.25 to 1 μg/ml and 0.5 to 4 μg/ml, respectively, with a modal polyhexanide MIC of 0.5 μg/ml and a modal chlorhexidine MIC of 4 μg/ml (Table 3). According to the epidemiological cutoff value proposed by Fabry et al., our S. aureus collection is considered susceptible to polyhexanide and 50% resistant to chlorhexidine (15). However, no correlation between chlorhexidine and polyhexanide susceptiblity profiles or cross-resistance was observed. Moreover, the majority of our isolates at D28 showed neither polyhexanide nor chlorhexidine MIC changes compared to isolates at D0. Altogether, our results suggest that the limited MRSA decolonization rate previously observed (17, 18) is not related to the presence or selection of strains with reduced susceptibility to polyhexanide or with cross-resistance toward chlorhexidine.
DISCUSSION
This study focused on the development of reduced polyhexanide susceptibility and the emergence of cross-resistance with other antiseptics or antibiotics in various MRSA strains. We previously found that a single polyhexanide decolonization course was not effective in eradicating MRSA carriage (18). This study, performed in a population composed mainly of MRSA strains harboring qac genes, excludes the possibility that the moderate decolonization rate of MRSA relies on the emergence of isolates showing reduced polyhexanide susceptibility or on potential cross-resistance between polyhexanide and chlorhexidine in MRSA.
In the present study, we provide evidence that prolonged in vitro exposure to low levels of polyhexanide results in the emergence of reduced polyhexanide susceptibility in MRSA strains without cross-resistance to chlorhexidine. Moreover, we repeatedly observed concomitant changes in the resistance profiles of daptomycin and glycopeptides, antibiotics used for S. aureus clinical treatment. This observation should encourage further in vivo studies, as various local and low disinfectant concentrations can potentially be found after topical administration of this substance or at residual levels on surfaces (15, 24).
In vitro, we detected reduced polyhexanide susceptibility (MIC changes from 0.5 to 4 μg/ml) following stepwise and prolonged (2 days) passages in low concentrations of polyhexanide (<2 μg/ml of polyhexanide). The occurrence of reduced polyhexanide susceptibility under low concentrations in vitro does not argue against the general use of polyhexanide for decolonization, because the high therapeutic concentration used greatly exceeds the low concentrations that permit resistance development and rapidly eradicates bacteria. However, it suggests careful follow-up of resistance profiles during topical administration.
To understand the molecular pathways leading to reduced polyhexanide susceptibility, we performed whole-genome sequencing and identified genetic changes in strains selected in vitro under polyhexanide exposition compared to wild-type strains. Mutations were found in mprF genes that can be correlated with reduced polyhexanide susceptibility. Indeed, polyhexanide is a cationic polymer attaching to negatively charged molecules and acting on bacterial membrane phospholipids, lipopolysaccharides, teichoic acids, and peptidoglycan components of the cell wall (9, 10). The integral membrane protein MprF lysinylates membrane lipid phosphatidyl glycerol (PG) and subsequently flips lysyl phosphatidyl glycerol (L-PG) to the outer leaflet of the plasma membrane (21). The mprF mutations detected in our strains may potentially increase L-PG synthesis and flipping, leading to an increase of membrane positive surface charge and consequently charge repulsion for cationic molecules, such as polyhexanide. Interestingly, our mutants showing mprF mutations and reduced susceptibility to polyhexanide also showed resistance to the cationic antibiotic daptomycin. The identified mprF mutation L337S is located in the so-called bifunctional domain of mprF known to be a hot spot for mprF mutations leading to daptomycin resistance (25). Further studies are under way to highlight the association of mprF mutations with antiseptic resistance, a mechanism that to our knowledge has not been previously identified. Regarding purR mutations, further studies are needed to understand the mechanistic link between purR mutations and reduced susceptibility to polyhexanide. In addition to containing the mprF mutation described above, a polyhexanide-resistant mutant of 134947 (P6) was found to contain a nonsynonymous mutation in purR. This mutation has been encountered elsewhere during selection for resistance to the lantibiotic nisin (22), and other purR mutations were discovered in mutants displaying resistance to vancomycin (26). However, we found no effect of purR mutation on polyhexanide resistance, as a strain containing PurR(V229G) was no more resistant to polyhexanide than its parent. It is not clear why mutations in purR emerge during selection for resistance to antibiotics or antiseptics; however, it is apparent that they are not required to confer resistance to these agents.
An important observation of this study is the potential emergence of cross-resistance between antiseptics and antibiotics used in clinical routine. This has been observed for antiseptics such as chlorhexidine or triclosan in other bacterial species (27–29). In S. aureus, cross-resistance between antiseptics and antibiotics was previously observed after chlorhexidine exposure selecting for resistance to several β-lactam antibiotics (30). To date, a single study assessed and found no correlation between polyhexanide and antibiotic resistance profiles. However, the analyzed collection lacks isolates with reduced polyhexanide susceptibility, which prevents any conclusion on cross-resistance between these molecules (15). Our results showed that development of reduced susceptibility to polyhexanide can be accompanied by changes in resistance not only to daptomycin but also to cell wall-active antibiotics such as vancomycin and teicoplanin. This can be expected by taking into account the mode of action of polyhexanide. Any molecular change leading to alteration in the cell wall could potentially affect the net charge of the cell wall and indirectly affect polyhexanide binding. This link was reliably observed in independent experiments and in different bacterial genetic backgrounds. However, we did not observe the development of reduced polyhexanide susceptibility accompanied always by an identical antibiotic resistance pattern, even though identical genetic mutations were identified. Studies related to whole transcriptomics would probably contribute to clarify the mechanisms leading to alteration of susceptibility.
Our experiments were performed in vitro, which appears to be their main limitation. In a recent clinical trial dedicated to assessing the decolonization efficacy of polyhexanide (18), we were able to collect 27 pairs of MRSA isolates resulting from cases of decolonization failure. No significant MIC alterations for antibiotics were observed between pairs of isolates in this small collection following polyhexanide exposition. Note, however, that in vivo, bacteria are probably exposed to the concentrations used in our report at specific body sites and that our design mimicking potential prolonged or repeated exposition to antimicrobial solutions may reflect the in vivo situation. Under these conditions, we reliably obtained alteration of susceptibility to polyhexanide as well as alteration in MRSA antibiotic susceptibility profiles, which is a potential risk of emergence of antibiotic resistance, particularly in areas showing generalized and extensive utilization of antiseptic solutions.
MATERIALS AND METHODS
Bacterial strains.
Bacterial strains used in this study are listed in Table 3. Strains COL, 134947, and 128822 are MRSA strains belonging to different ST types and used to analyze phenotypic and genetic alterations following exposure to antiseptic solutions. The other MRSA isolates were collected during a previously published randomized, placebo-controlled trial, assessing the clinical efficacy of polyhexanide in eradicating MRSA carriage at day 28 (D28) after decolonization (18). Briefly, selected MRSA-colonized patients fulfilling inclusion criteria were randomized to receive either active treatment or placebo for 10 days. Active treatment (Prontoderm solution [B. Braun Medical AG, Sempach, Switzerland], a mix of polyhexanide, allantoin, a cationic component, surfactants, and purified water, and Prontoderm Gel Light [B. Braun Medical AG, Sempach, Switzerland], a mix of polyhexanide, glycerol, cellulose polymer, and purified water) was applied to the skin (once daily) and the anterior nares (thrice daily). Placebos were similar in appearance but did not contain polyhexanide. After 28 days of treatment, swabs were taken from the inguinal/perineal region (one swab) and both nares (one swab) and identification of MRSA was performed as previously described (18). MRSA strains before (D0) and after treatment (D28) were saved frozen in skimmed milk for further determinations.
Molecular MRSA typing.
MRSA isolates were subjected to a rapid genotyping assay using multiple-locus variable number of tandem repeats analysis (MLVA). Briefly, this assay is based on a multiplex PCRs using 10 primer pairs targeting 9 genes showing variable numbers of tandem repeats and an additional pair of primers allowing amplification of the mecA gene as an internal control. This method shows discriminatory power that is at least similar to that of pulsed-field gel electrophoresis (31). The analysis was performed on isolated colonies grown on Mueller–Hinton agar disrupted by vortex agitation. PCR amplification was then evaluated using a microcapillary electrophoresis system (2100 Bioanalyzer, Agilent Technologies) and automatically analyzed using specifically developed software (31). The genotype of each strain was deduced by comparison with profiles obtained with characterized control isolates (31).
Polyhexanide susceptibility testing.
Polyhexanide 20% solution was obtained and prepared as recommended by the manufacturer. The stock solution was diluted in the test broth to final polyhexanide concentrations of 0.25, 0.5, 1, 2, and 4 μg/ml. Polyhexanide MICs were determined as previously described (8) but using a macrodilution method. Briefly, one bacterial colony growing on Mueller-Hinton agar was used to inoculate 1 ml of Mueller-Hinton broth (MHB). Overnight culture at 37°C was diluted to deliver the final inoculum of 1.5 × 106 CFU/ml into each tube containing different polyhexanide concentrations (MHB containing 0, 0.25, 0.5, 1, 2, and 4 μg/ml of polyhexanide). After incubation for 24 to 48 h at 37°C, MIC was defined as the lowest concentration allowing bacterial growth. Three independent MIC determinations were performed for each isolate. Modal MICs for each isolate are represented. Chlorhexidine MICs were determined as described above but using chlorhexidine diluted to final concentrations of 0.12 to 16 μg/ml.
In vitro selection of polyhexanide and chlorhexidine mutants with reduced susceptibility.
The selected MRSA strains COL, MRSA134947 and MRSA128822 showing initial polyhexanide MICs of 0.5, 1, and 1 μg/ml, respectively, were serially passaged onto increasing concentrations of polyhexanide. Briefly, 100 ml of Mueller-Hinton broth (CAMHB) containing polyhexanide (MIC 0.5 or 1 μg/ml) was inoculated overnight with bacteria at a concentration of 1 × 109 bacteria/ml. After incubation for 2 days at 37°C, bacteria growing at a polyhexanide concentration of 0.5 or 1 μg/ml, were used for a second step passage in increased polyhexanide concentrations. Further stepwise passages were done when indicated. After passages bacteria were collected and macrodilution MICs were determined. An identical methodology was used for selection of reduced-chlorhexidine-susceptible mutants of strains COL, MRSA134947, and MRSA128822, showing initial chlorhexidine MICs of 2, 4, and 8 μg/ml, respectively.
Antimicrobial susceptibility testing.
The bacterial inoculum suspension was prepared by selecting several colonies from overnight growth (16 to 24 h of incubation) on Columbia agar plates with a cotton swab and suspending the colonies in sterile saline (0.85% NaCl, wt/vol, in water) to the density of a 0.5 McFarland standard, corresponding to 3 to 4 × 108 CFU/ml. The inoculum was spread over the entire surface of the Mueller–Hinton agar plate by swabbing in three directions, and the plates were incubated in a humid atmosphere at 35 ± 1°C for 18 ± 2 h. Antibiotic resistance profiles were tested using disc diffusion assays according to EUCAST methods.
Genome sequencing.
High-throughput sequencing was used to sequence the genomes of all isolates. Genomic DNA from each isolate was purified by using DNeasy columns (Qiagen), and then sequenced on the Illumina HiSeq 2500 (Illumina, San Diego, CA) using 100 bases, paired-ends, and barcodes according to the Nextera XT kit (Illumina). Read sequence quality was assessed with the FastQC program (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/) and filtered using the fastq-mcf program (https://github.com/ExpressionAnalysis/ea-utils/blob/wiki/FastqMcf.md). Genome assembly was performed using the Edena v3 assembler (32). Assembled genomes were annotated using the Prokka v1.10 program (33). The phylogenetic relationships of isolates were investigated by genomic single-nucleotide polymorphism (SNP)-based analysis using the Parsnp v1.0 program (34). The proteome comparison of all isolates was performed using the CGView Comparison Tool program (35). The blastp analysis was used to detect nonsynonymous mutations.
ACKNOWLEDGMENTS
We thank all investigators and infection control staff members who contributed to and helped to conduct the clinical trial.
S.H. has received a peer-reviewed research grant funded by Pfizer and is also a member of the advisory boards of DNA Electronics, GlaxoSmithKline, Novartis, and Bayer. J. Schrenzel participated in advisory boards and received honoraria from bioMérieux. All other authors have no conflicts of interest to declare.
This work was funded in part by B. Braun (Sempach, Switzerland), which had no influence on data analysis and reporting. This study was partially funded by Swiss National Science Foundation grants 310030-149762 (to A.R.) and 31003A_153474 (to P.F). For E.V.D., C.L., and S.H., the research leading to this paper received support from the Innovative Medicines Initiative Joint Undertaking under grant agreement numbers 115523, 115620, and 115737 (Combatting Bacterial Resistance in Europe projects [COMBACTE]), resources of which are composed of financial contribution from the European Union's 7th Framework Programme (FP7/2007 ± 2013) and the European Federation of Pharmaceutical Industries and Associations (EFPIA) companies' in-kind contribution.
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