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Antimicrobial Agents and Chemotherapy, May 2009, p. 2059-2065, Vol. 53, No. 5
0066-4804/09/$08.00+0     doi:10.1128/AAC.00871-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Genetic Determinants of Resistance to Fusidic Acid among Clinical Bacteremia Isolates of Staphylococcus aureus

Jonas Lannergård, Tobias Norström, and Diarmaid Hughes^*

Microbiology Programme, Department of Cell and Molecular Biology, Box 596, Biomedical Center, Uppsala University, S-75124 Uppsala, Sweden

Received 1 July 2008/ Returned for modification 23 October 2008/ Accepted 5 March 2009

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Resistance to fusidic acid in Staphylococcus aureus is caused by mutation of the elongation factor G (EF-G) drug target (FusA class) or by expression of a protein that protects the drug target (FusB and FusC classes). Recently, two novel genetic classes of small-colony variants (SCVs) were identified among fusidic acid-resistant mutants selected in vitro (FusA-SCV and FusE classes). We analyzed a phylogenetically diverse collection of fusidic acid-resistant bacteremia isolates to determine which resistance classes were prevalent and whether these were associated with particular phylogenetic lineages. Each isolate was shown by DNA sequencing and plasmid curing to carry only one determinant of fusidic acid resistance, with approximately equal frequencies of the FusA, FusB, and FusC genetic classes. The FusA class (mutations in fusA) were distributed among different phylogenetic types. Two distinct variants of the FusC class (chromosomal fusC gene) were identified, and FusC was also distributed among different phylogenetic types. In contrast, the FusB class (carrying fusB on a plasmid) was found in closely related types. No FusE-class mutants (carrying mutations in rplF) were found. However, one FusA-class isolate had multiple mutations in the fusA gene, including one altering a codon associated with the FusA-SCV class. SCVs are frequently unstable and may undergo compensatory evolution to a normal growth phenotype after their initial occurrence. Accordingly, this normal-growth isolate might have evolved from a fusidic acid-resistant SCV. We conclude that at least three different resistance classes are prevalent among fusidic acid-resistant bacteremia isolates of S. aureus.

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Staphylococcus aureus is an important hospital and community pathogen, and fusidic acid is one of several antibiotics used in its management (1, 9, 29, 36, 37, 42, 43). Fusidic acid interacts with translation elongation factor G (EF-G) on the ribosome, where it inhibits protein synthesis by preventing the release of EF-G·GDP (7, 44). There are two mechanisms known to cause resistance to fusidic acid in S. aureus: (i) alteration of the drug target (EF-G in complex with the ribosome) by mutation (5, 22) and (ii) protection of the drug target by FusB-family proteins (25). Each of these mechanisms has multiple genetic causes, some of which have only recently been discovered (23, 28).

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The first mechanism of fusidic acid resistance to be identified was target alteration caused by mutations in fusA, the chromosomal gene encoding EF-G (8). More than 30 different amino acid substitution mutations causing fusidic acid resistance in S. aureus have now been identified in EF-G (5, 22, 23). The resistance mutations occur mostly in structural domain III of EF-G, but some also occur in domains I and V (17). In a few clinical isolates, multiple resistance-associated mutations have been found in fusA, suggestive of multiple rounds of selection (6, 22). Data from in vitro translation experiments (12, 32) show that fusA mutations reduce the affinity of fusidic acid for the EF-G-ribosome complex. We refer to resistance due to fusA mutations as FusA-class resistance. A subset of the mutations in fusA resulting in resistance to fusidic acid, most of which are located in domain V of EF-G, cause the small-colony variant (SCV) phenotype. In addition to being resistant to fusidic acid, these mutants have reduced susceptibility to aminoglycosides and are auxotrophic for hemin. We refer to this group of mutants as the FusA-SCV class (23). Another group of fusidic acid-resistant mutants that display an SCV phenotype are referred to as FusE and carry mutations in rplF, the gene encoding ribosomal protein L6 (23). Protein L6 maps in the part of the ribosome where EF-G interacts (3, 35).

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The FusB protein prevents fusidic acid from interacting with EF-G, thus protecting the translation apparatus from inhibition by the antibiotic (25). Several decades ago it was recognized that resistance to fusidic acid could be associated with the presence of a penicillinase plasmid in S. aureus (15, 16). An example of this plasmid, pUB101, was sequenced and shown to carry genes coding for cadmium resistance and a gene designated fusB (also known as far), responsible for fusidic acid resistance mediated by target protection (24-26). The fusB gene has also been found to be integrated into the chromosome of S. aureus in an epidemic fusidic acid-resistant impetigo clone (27). In addition, chromosomal genes encoding proteins with 45% amino acid similarity to FusB have been identified (28). These genes, fusC (found in S. aureus and Staphylococcus intermedius) and fusD (found in Staphylococcus saprophyticus), confer resistance to fusidic acid on S. aureus and are presumed to encode proteins that, like FusB, protect EF-G from the antibiotic (28). The fusB and fusC genes have also been identified in fusidic acid-resistant isolates of Staphylococcus epidermidis (20).

Previous studies of the genetic basis of fusidic acid resistance in clinical isolates of S. aureus have been limited to an examination of strains recovered from patients with impetigo (26-28). In addition, previous studies could not have looked for evidence of the FusE genotype (mutation in rplF), as it has only recently been described and is associated with an SCV phenotype (23). SCVs are of special interest in S. aureus infections because they have reduced susceptibility to aminoglycosides and have been isolated from patients with chronic, persistent, and/or recurrent infections (13, 30, 31, 38, 39). This distribution of SCVs is probably a consequence of their ability to persist intracellularly and thus to be shielded from the host immune response (2, 38, 40). The SCV phenotype in S. aureus is often unstable, and many strains revert or evolve at a high frequency to a normal colony phenotype (4, 41). For practical reasons (very slow growth, phenotypic instability), SCVs are only rarely identified in clinical laboratory isolates unless special efforts are made to look specifically for them. However, if the phenotypic reversion of the SCV to a normal growth phenotype involves secondary compensatory mutations, then genetic traces of the original SCV mutation could remain in the genotypes of resistant isolates with a normal growth phenotype. It is therefore of interest to determine which fusidic acid resistance mechanisms are associated with invasive S. aureus infections. Here, we examined the genetic basis of fusidic acid resistance in a phylogenetically diverse set of clinical bacteremia isolates of S. aureus.

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Bacterial strains. Twenty fusidic acid-resistant S. aureus bacteremia isolates, originally from the Statens Serum Institut (Copenhagen, Denmark), were described previously in terms of fusidic acid MICs (22). The isolates were collected from patients in 18 Danish hospitals during 1996. In the two cases where two isolates were collected from the same hospital, both spa typing and fusA analysis showed that the isolates were not clonal (see Results and Fig. 1).

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FIG. 1. Phylogenetic tree based on the spa sequences of the 20 bacteremia isolates studied here, shown as a UPGMA dendrogram with the sum of branch lengths being 1.49. The tree is drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree. Clustering was performed using the BURP algorithm (21), with the Ridom StaphType 1.5 program (Ridom GmbH, Würzburg, Germany). Isolate identification numbers (INXXX), each with its Fus resistance class, follow the spa type. C and C* indicate the two different genome contexts of the fusC gene in the chromosome.

Media and growth conditions. Standard liquid growth medium was LB, and solid growth medium was LA (LB plus 1% agar; Merck, Darmstadt, Germany). B2 broth (1% casein hydrolysate [Merck, Darmstadt, Germany], 2.5% yeast extract [Oxoid, Hampshire, England], 0.1% K₂HPO₄, 0.5% glucose, 2,5% NaCl) was used when S. aureus cells were made electrocompetent and as an outgrowth medium after electroporation. NYE agar plates (1% casein hydrolysate, 0.5% yeast extract, 0.5% NaCl, 1.5% agar) were used for plating electroporated cells. Difco Mueller-Hinton agar (Becton, Dickinson and Company, Sparks, MD) was used for all MIC tests. Liquid cultures were grown at 37°C in flasks or tubes on a shaker set at 200 rpm.

Antimicrobials and MIC tests. The antibiotics ampicillin (Sigma Aldrich, Stockholm, Sweden) and fusidic acid (Leo Pharma, Ballerup, Denmark) were used for selection and screening in solid medium, each at 1 µg/ml. Cadmium acetate (Sigma Aldrich) was used at 10^–4 M (final concentration) in solid medium. MICs of antibiotics were measured on Mueller-Hinton agar using an Etest (AB Biodisk, Solna, Sweden) according to the manufacturer's instructions.

Plasmid DNA preparation. We examined the clinical S. aureus strains for the presence of an 21-kb pUB101-like plasmid (24) coding for ampicillin resistance, cadmium resistance, and fusidic acid resistance. Clinical isolates were grown overnight without shaking and without antibiotic selection in LB supplemented with 0.5% glycine (Sigma-Aldrich, Stockholm, Sweden). Cells from 15 ml of overnight culture were pelleted by centrifugation at 3,000 x g for 15 min. The cell pellet was resuspended in 100 µl TE buffer (10 mM Tris-Cl [pH 8.0], 1 mM EDTA [pH 8.0]), to which was added 100 µl of lysostaphin (L7386; Sigma-Aldrich; 300 µg/ml), 100 µl lysozyme (Sigma-Aldrich; 100 mg/ml) and 45 µl proteinase K (Sigma-Aldrich; 20 mg/ml). This mixture was incubated at 37°C for 30 to 60 min, after which 300 µl of 0.2 M NaOH, 1% sodium dodecyl sulfate was added and incubation continued at room temperature (18°C) for a further 5 to 10 min. Potassium acetate (300 µl, 3 M) was added, and the tube was inverted several times and then kept on ice for 15 min. The solution was centrifuged at 14,000 x g for 30 min at 4°C. The clear supernatant was removed, extracted with phenol-chloroform followed by chloroform, loaded onto a GenElute plasmid miniprep kit (PLN-70; Sigma-Aldrich), washed, and eluted according to the manufacturer's instructions. The eluate was mixed with 0.7 volume isopropanol, mixed, and centrifuged at 14,000 x g in a microcentrifuge at 4°C. The plasmid DNA pellet was washed once with 500 µl 70% ice-cold ethanol, dissolved in 50 µl distilled water, and stored at –20°C. Plasmid DNA was visualized on agarose gels, and the presence of cadmium resistance (cadXD) and fusidic acid resistance (fusB) sequences was assayed by PCR and DNA sequencing. Plasmids from clinical isolates were transformed into the restriction-negative S. aureus strain RN4220 by electroporation.

Electrocompetent cells. S. aureus RN4220 (generously supplied by Molly Schmid and Bret M. Benton, formerly of Microcide Pharmaceuticals, Mountain View, CA) was grown overnight in B2 broth at 37°C, diluted 25 times in fresh B2 in an E-flask (total volume, 25 ml), and grown at 37°C to an optical density at 600 nm of 0.5. Cells were washed three times in 1 volume of filter-sterilized deionized water, followed by two washes with 10% glycerol in 1/2 and 1/5 volumes, respectively. Washing was by centrifugation in SS-34 tubes at 5,000 x g. Before the last centrifugation, the cells were left at room temperature for 15 min. All steps were carried out at room temperature. The final cell pellet was dispersed in 800 µl 10% glycerol and if not used immediately was stored at –80°C in 80-µl aliquots for up to 1 month.

Electroporation. Plasmids purified from clinical S. aureus were transformed into RN4220 by electroporation, essentially as described previously (33). All steps were at room temperature. Seventy microliters of electrocompetent cells was mixed with 0.5 to 1 µg of plasmid DNA (50 to 100 ng/µl). Sixty microliters of this mixture was transferred to a 1-mm electroporation cuvette (ECU101; EquiBio, Ashford, Middlesex, United Kingdom) and electroporated using a Bio-Rad gene pulser (Bio-Rad Laboratories AB, Sundbyberg, Sweden). The gene pulser was set at 100 resistance, 25 µF capacitance (2.5 ms optimal time constant), and 2.3 kV. After electroporation, cells were resuspended in 400 µl B2 broth, transferred to a microcentrifuge tube and incubated at 37°C for 24 h to allow recovery. Cells were spread on NYE agar containing ampicillin at 1 µg/ml and incubated at 37°C for 24 to 48 h. Typically, 200 transformants were recovered.

PCR. To extract template DNA for PCR, a fresh bacterial colony was suspended in 100 µl sterile water to which was added 100 µl acid-washed glass beads (catalog no. G1277; Sigma-Aldrich, Stockholm, Sweden). The bacterium-bead mixture was vortexed at maximum speed on a bench vortex device for 45 to 60 s to disrupt the cells. A 0.5-µl portion of this solution was used as the DNA template to initiate a PCR. PCRs were carried out using a Ready-2-Go PCR bead kit (GE Health, Uppsala, Sweden) in a PTC-200 Peltier thermal cycler (SDS Diagnostics, Falkenberg, Sweden) with oligonucleotides from Sigma (Sigma Genosys Ltd, Sigma-Aldrich House, Haverhill, United Kingdom). PCRs were initiated with 5 min denaturation at 95°C, followed by 30 cycles of 20 s at 95°C, 20 s at 50°C, and either 1 min (fusB, fusC, rplF, and cadXD) or 2.5 min (fusA) at 72°C. PCR products were visualized by agarose gel electrophoresis, purified prior to sequencing using a QIAquick PCR purification kit (Qiagen, VWR International AB, Stockholm, Sweden) according to the manufacturer's instructions, and quantified after purification using an NO-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE). The oligonucleotides used for PCR and DNA sequencing are listed in Table 1.

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TABLE 1. Oligonucleotides used for PCR and DNA sequencinga

Single-primer semirandom PCR. The genome location of fusC in the different strains was determined using a single-primer semirandom PCR method (10). The PCR amplification was performed with one primer per reaction (Fus3F for the downstream region and FusC3R for the upstream region), with the annealing temperature set to 45°C to allow for unspecific reverse priming outside of fusC. The resulting mixture of PCR products was sequenced with the nested primers FusC4F (downstream) and FusC1R (upstream).

DNA sequencing. DNA sequencing was performed at Macrogen Inc., Seoul, South Korea, and at the DNA Sequencing Facility (Rudbeck Laboratory, Uppsala University) on an ABI Prism 3700 capillary sequencer using a BigDye terminator version 3.1 cycle sequencing kit (Applied Biosystems, Foster City, CA). Each sequencing reaction mixture contained 8 ± 4 ng purified PCR product, 1.6 pmol sequencing primer, 4 µl Terminator Ready Reaction mix, and water to bring the total volume to 10 µl.

Plasmid curing. Clinical isolates carrying pUB101-like plasmids were cured of the plasmid by inoculating 100 cells into 100 ml LB and growing overnight at 43°C with shaking for 30 generations and then screening for loss of cadmium resistance by replica plating at 37°C (15). Cured strains were obtained at a frequency of 0.1% and purified on LA plates. Loss of the plasmid was verified by loss of unselected phenotypic traits (ampicillin resistance and fusidic acid resistance) and by PCR of cadXD.

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Phylogenetic diversity of the clinical isolates. The spa typing method (14, 34) was used to assess the phylogenetic diversity of the 20 Fus^r bacteremia isolates examined here. The spa sequences were analyzed using the RIDOM StaphType 1.5 program (Ridom, GmbH, Würzburg, Germany; http://spaserver.ridom.de/). The 20 isolates belong to 13 different spa groups. No more than three isolates belonged to any one spa group. A phylogenetic tree based on these spa sequences (Fig. 1) was built using the BURP (Based Upon Repeat Patterns) algorithm (21). To validate the strain diversity apparent from the spa typing, we also made a phylogenetic tree based on the fusA sequences of the 20 isolates. We aligned and compared the fusA sequences from 14 S. aureus strains for which complete genome sequences were available (www.ncbi.nlm.nih.gov) and from the 20 Fus^r bacteremia isolates examined here. We found a total of 24 base pair differences and used these to classify the fusA genes into 11 phylogenetic groups. The phylogenetic grouping based on fusA is in good agreement with the spa typing (data not shown) and with the grouping previously made by multilocus sequence typing for a subset of these genomes (18). The phylogenetic information can be used to evaluate the likelihood of clonal relationships between different clinical isolates and thus the significance of the distribution of different resistance classes. We concluded that this set of 20 bacteremia isolates is genetically diverse.

FusA class resistance. In a previous study, we showed that a number of these clinical isolates had mutations in fusA causing fusidic acid resistance (22), and here we found fusA mutations in 6 of the 20 clinical strains. The phylogenetic classification made using spa typing (Fig. 1) shows that none of these six fusA genes belong to the same group (Table 2). The isolates IN441 and IN448 carry an identical resistance mutation in fusA (L461S), but because the isolates belong to different phylogenetic groups (t167 and t230, respectively), the mutations must have arisen independently. Interestingly, one isolate, IN442, carried four amino acid substitution mutations in fusA, including one in codon 655. Mutations in this region of EF-G, codons 655 to 666, are closely associated with the FusA-SCV phenotype (23), suggesting that this isolate might have evolved by compensatory evolution from an SCV.

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TABLE 2. Classification of resistance type in clinical Fusr isolates

FusB class resistance. A second source of fusidic acid resistance is the fusB gene, frequently borne on the 21-kb plasmid pUB101 (24). We assayed the 20 clinical isolates for the presence of plasmids and found that 17 carried an 21-kb plasmid, 1 carried a plasmid of 3 to 4 kb (IN439), and 2 had no detectable plasmids (Table 2). The complete sequences of 50 different plasmids from strains of S. aureus are available in public genome databases (www.ncbi.nlm.nih.gov). Three of these plasmids, pUB101, pMW2, and p21, are 21 kb in size, while most others are either much larger (>38 kb) or smaller (8 kb). Each of the sequenced 21-kb plasmids carries a cadDX operon, encoding cadmium resistance, and a bla operon, encoding ß-lactamase activity, but only pUB101 carries fusB, encoding resistance to fusidic acid. Each of the 17 clinical isolates carrying an 21-kb plasmid was resistant to cadmium and to ampicillin, whereas the remaining three isolates were susceptible to both. PCR was used to assay the 20 isolates for the presence of cadXD and fusB, using both purified plasmid preparations and total cell lysates. The cadXD sequence was found in all 17 clinical isolates that carry an 21-kb plasmid but not in any of the remaining three isolates. In contrast, fusB could be amplified from only 6 of the 20 isolates (Table 2). The fusB sequence was determined from each of these six plasmids and was identical to the published sequence on pUB101 (24). None of these six isolates carried a fusidic acid resistance mutation in fusA. The six FusB-class mutants belong to four closely related spa types, t065, t715, t3000, and t362 (Table 2 and Fig. 1). Three of these were spa type t065, suggesting a possible clonal relationship between these isolates.

We assigned the 17 21-kb plasmids to three groups based on the sequence of cadX and its surrounding sequences. One group, consisting of the six plasmids with fusB, is identical in sequence to pUB101. A second group, consisting of ten plasmids, is identical in sequence to pMW2. The third group, consisting of two plasmids designated pMW2(), is identical to pMW2 in the cadX coding sequence and for at least 60 nucleotides downstream, except for a deletion of 9 bp downstream of the cadX termination codon (TGAAACGAGTGAAACGAGTTT TGAAACGAGTTT; the termination codon is underlined). An identical 9-bp deletion is found in p21. The deletion occurs where there is a direct repeat of nine nucleotides, suggesting that it is a result of slippage and mispairing (19) during replication. Because p21 differs from the pMW2() plasmids at a large number of other nucleotide positions, including some within and downstream of cadX, we suggest that pMW2() is a derivative of pMW2 that has independently acquired a deletion identical to that found in p21.

FusB plasmid curing and transfer. The MIC of fusidic acid for each of the six isolates carrying pUB101 (fusB) was measured before and after curing of the plasmid (Table 3). In each case, the original isolates had similar MICs (within one step), and each became susceptible to fusidic acid after curing, showing that the only fusidic acid resistance determinant in these six isolates was carried on the plasmid. Plasmids purified from each of the six isolates were also transformed into RN4220, and the fusidic acid MIC was measured again. The MICs for the transformed RN4220 were similar with all six plasmids and were within one step of the MICs measured in the clinical strains. On this basis, we conclude that these clinical isolates do not have any fusidic acid resistance determinants other than the plasmid-borne fusB gene.

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TABLE 3. FusB class resistance is plasmid borne

FusC class resistance. The remaining eight isolates that lack a fusA mutation and have no fusB gene were each shown by PCR and DNA sequencing to carry the fusC gene on the chromosome and were thus classified as the FusC resistance class (Table 2). In four out of eight FusC isolates, a synonymous mutation (TCC to TCT) was present at nucleotide position 90 in fusC, and since these strains also belong to two closely related spa types, t630 and t015, it is likely that these four are clonally related. However, the remaining four FusC isolates belong to three different spa types (t015, t127, and t008). Thus, the eight fusC isolates are distributed among four spa types. The fusC gene was originally identified in the genome sequence of S. aureus MSSA476, where it was found inside the staphylococcal chromosome cassette SCC₄₇₆ (11). Using single-primer semirandom PCR, we amplified and sequenced the up- and downstream regions adjacent to fusC in the eight strains carrying this gene. An alignment of the obtained sequences upstream of fusC showed 99 to 100% identity to S. aureus MSSA476 in all eight isolates, indicating that the fusC gene has been incorporated in the genome of these strains as part of an SCC element, independently or in a common ancestor. Strains IN439, IN456, and IN449 (spa types t008 and t127) also displayed complete sequence identity to MSSA476 in the noncoding region downstream to fusC, in contrast to the remaining five strains (belonging to spa types t630 and t015), where the sequence similarity ends 70 bases downstream from the stop codon. We conclude that there are at least two variants of the fusC region present in clinical S. aureus isolates.

FusE class resistance. No FusE-class mutations were found in any of the 20 strains, as DNA sequencing showed that they all carried wild-type rplF genes.

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Resistance to fusidic acid is associated with fitness costs for S. aureus both in vitro and in vivo (6, 22). It would not be surprising if the magnitude or nature of these resistance-associated fitness costs was influenced by genomic context, resulting in a skewed distribution of resistance determinants as a function of genotype. Indeed, of recent S. aureus impetigo isolates in Europe, the great majority belong to the same phylogenetic group and carry the same resistance determinant (26, 27), suggesting that the particular combination may be associated with a high fitness.

A priori, one would expect fusidic acid resistance by mutation (FusA class) to be phylogenetically widespread unless it was counterselected in some genotypes. Regarding plasmid-borne fusidic acid resistance (FusB class), its phylogenetic distribution is expected to reflect the complex dynamics of plasmid transmission between strains and the clonal selection of resistant strains. Finally, the distribution of intrinsic fusidic acid resistance associated with the presence of a resistance gene on the chromosome (FusC class) might be expected to be phylogenetically restricted, depending on the probability of independent acquisition of the element into one or more lineages and the time for subsequent genetic diversification of a lineage after acquisition.

Here, we addressed the genetic causes of fusidic acid resistance in a phylogenetically diverse collection of S. aureus bacteremia isolates. The resistance determinants we found in these isolates belonged to the FusA class (mutations in fusA), the FusB class (presence of pUB101 carrying the fusB gene), and the FusC class (fusC gene inserted in the chromosome as part of an SCC island). No representatives of the FusE SCV class (mutation in rplF) were found. Each isolate carried a single determinant for resistance, and there were approximately equal distributions of the FusA, FusB, and FusC classes among the 20 isolates studied (6:6:8, respectively). Based on phylogenetic relationships (spa typing), we could conclude that fusidic acid resistance in all six of the FusA-class isolates most likely arose independently. This is not unexpected for a resistance that arises by mutation, but it does indicate that this resistance class is not closely associated with a particular genetic background (Fig. 1). In contrast, the FusB-class isolates are closely related and possibly clonal based on their spa types and their fusA and cadXD sequences. Likewise, four of eight FusC strains are closely related and possibly clonal based on their identical fusA and fusC sequences and the identical genomic context of the fusC gene, while the remaining four strains (IN439, IN456, IN447, and IN449) are genetically dissimilar. In the genome sequence of S. aureus MSSA476, fusC is located inside the staphylococcal chromosome cassette SCC₄₇₆ (11). Even though all FusC strains have the fusC gene inserted at the same site as MSSA476, the sequence downstream of fusC shows genetic rearrangements that divides the strains into two groups. This could mean either that the FusC strains belong to lineages that acquired different SCC elements in at least two independent events or that the element is inherited from an ancestor common to all strains and has since then gone through genetic rearrangements.

No fusidic acid-resistant isolates of the FusE class could be identified among the strains studied, and as this class of mutants has so far been identified only in vitro, it could be taken as evidence that mutants of this genotype are unlikely to thrive in the clinical setting. In a set of bacteremia isolates, however, this result is expected, as strains with a small-colony phenotype are unlikely to be collected from patients with blood infections.

Interestingly, the FusA isolate IN442 carries a mutation in codon 655, where a different mutation has previously been associated with the FusA-SCV class (23), in addition to two other substitutions (Leu461Phe and Asp463Gly) associated with the FusA class and one substitution associated with growth fitness compensation (Ala376Val) (22). This raises the possibility that IN442 might have initially been a FusA-SCV isolate and evolved to a normal growth phenotype by acquiring intragenic compensatory mutations in fusA. Further studies of clinical isolates are required to determine the extent to which SCVs are involved in the development of fusidic acid resistance. Our overall conclusion from this study is that three different genetic determinants of fusidic acid resistance (FusA, FusB, and FusC) are common among S. aureus bacteremia isolates. Furthermore, FusA and FusC resistance is widely distributed among different phylogenetic groups of S. aureus.

 
The work was supported by grants from the Swedish Natural Science Research Council (Vetenskapsrådet) and a European Union 6th Framework grant (EAR project) to D.H. T.N. acknowledges support from Leo Pharma, Ballerup, Denmark, while J.L. was supported in part by a postdoctoral fellowship from the Carl Trygger Foundation.

We gratefully acknowledge Ann-Cathrine Petersson and Bo Nilsson (Lund University, Sweden) for their assistance with the spa analysis and tree building.

 
* Corresponding author. Mailing address: Microbiology Programme, Department of Cell and Molecular Biology, Box 596, Biomedical Center, Uppsala University, S-75124 Uppsala, Sweden. Phone: 46-18-4714354. Fax:. 46-18-530396. E-mail: diarmaid.hughes{at}icm.uu.se

Published ahead of print on 16 March 2009.

These authors contributed equally to the manuscript.

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Antimicrobial Agents and Chemotherapy, May 2009, p. 2059-2065, Vol. 53, No. 5
0066-4804/09/$08.00+0     doi:10.1128/AAC.00871-08
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