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Journal of Clinical Microbiology, April 2009, p. 1172-1180, Vol. 47, No. 4
0095-1137/09/$08.00+0     doi:10.1128/JCM.01891-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Biofilm Formation by Staphylococcus haemolyticus

Elizabeth Gladys Aarag Fredheim,¹ Claus Klingenberg,^1,2 Holger Rohde,³ Stephanie Frankenberger,³ Peter Gaustad,⁴ Trond Flægstad,^1,2 and Johanna Ericson Sollid^5*

Department of Pediatrics, Institute of Clinical Medicine, University of Tromsø, Tromsø, Norway,¹ Department of Pediatrics, University Hospital of North Norway, Tromsø, Norway,² Institut für Medizinische Mikrobiologie, Virologie und Hygiene, Universitätsklinikum Hamburg-Eppendorf, Hamburg, Germany,³ Institute of Microbiology, Rikshospitalet, and University of Oslo, Oslo, Norway,⁴ Department of Microbiology and Virology, Institute of Medical Biology, University of Tromsø, Tromsø, Norway⁵

Received 30 September 2008/ Returned for modification 8 November 2008/ Accepted 7 January 2009

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Infections due to coagulase-negative staphylococci (CoNS) most frequently occur after the implantation of medical devices and are attributed to the biofilm-forming potential of CoNS. Staphylococcus haemolyticus is the second most frequently isolated CoNS from patients with hospital-acquired infections. There is only limited knowledge of the nature of S. haemolyticus biofilms. The aim of this study was to characterize S. haemolyticus biofilm formation. We analyzed the biofilm-forming capacities of 72 clinical S. haemolyticus isolates. A detachment assay with NaIO₄, proteinase K, or DNase was used to determine the main biofilm components. Biofilm-associated genes, including the ica operon, were analyzed by PCR, and the gene products were sequenced. Confocal laser scanning microscopy (CLSM) was used to elucidate the biofilm structure. Fifty-three isolates (74%) produced biofilms after growth in Trypticase soy broth (TSB) with glucose, but only 22 (31%) produced biofilms after growth in TSB with NaCl. It was necessary to dissolve the biofilm in ethanol-acetone to measure the optical density of the full biofilm mass. DNase, proteinase K, and NaIO₄ caused biofilm detachment for 100%, 98%, and 38% of the isolates, respectively. icaRADBC and polysaccharide intercellular adhesin (PIA) production were found in only two isolates. CLSM indicated that the biofilm structure of S. haemolyticus clearly differs from that of S. epidermidis. We conclude that biofilm formation is a common phenotype in clinical S. haemolyticus isolates. In contrast to S. epidermidis, proteins and extracellular DNA are of functional relevance for biofilm accumulation, whereas PIA plays only a minor role. The induction of biofilm formation and determination of the biofilm mass also needed to be optimized for S. haemolyticus.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Coagulase-negative staphylococci (CoNS) are major nosocomial pathogens. Among the CoNS, Staphylococcus haemolyticus is second only to S. epidermidis in its frequency of isolation from human blood cultures (3, 14). S. haemolyticus plays an important role in hospital-acquired opportunistic infections related to implanted medical devices (29, 31, 34). Furthermore, S. haemolyticus has the highest level of antimicrobial resistance among all CoNS species, and heteroresistance to glycopeptides is common (8, 19). This limits the therapeutic options available and makes an S. haemolyticus infection a serious threat. Therefore, a deeper understanding of how S. haemolyticus turns into a pathogen at the molecular level is needed in order to develop new approaches for prevention and therapy.

The ability to form a biofilm is considered the most important virulence factor in CoNS foreign material-associated infections (29). Biofilm formation is a two-step process that has been described in detail for S. epidermidis. The initial attachment is mediated by a number of factors like cell wall-anchored surface proteins (i.e., Fbe and Bhp) and the cell wall lytic enzyme autolysin E (AtlE) (21). Fbe (12) and AtlE both have adhesive properties and bind to the host factors fibrinogen and vitronectin, respectively. Bhp has been suggested to contribute to primary attachment as well as intercellular adhesion (11).

In S. epidermidis, the accumulative phase is linked to the production of polysaccharide intercellular adhesin (PIA), which is synthesized by icaADBC-encoded proteins (22, 31). However, it has become clear that PIA is neither an essential nor necessarily the major component of CoNS biofilms (18, 28). Intercellular adhesion mediated by the accumulation-associated protein (Aap), independently or in cooperation with the ica operon, is well described (24, 26, 42). Additional components, such as proteins other than Aap, DNA, and RNA and polysaccharides other than PIA (24, 40, 43, 47), have also been suggested to be important in CoNS biofilms.

In contrast to S. epidermidis, the molecular basis of the virulence of S. haemolyticus in general and in the context of foreign material-associated infections is largely unknown. Complete genome sequencing of S. haemolyticus JCSC1435 uncovered a wide range of open reading frames encoding putative virulence factors (45). Furthermore, passage in drug-free medium causes frequent genomic rearrangements (45). Both antimicrobial resistance and genetic rearrangements are probably mediated by insertion elements, which are abundant in S. haemolyticus (49). Recently, a capsular polysaccharide was proposed to be an important virulence factor in S. haemolyticus (16). S. haemolyticus biofilm formation in vitro has been reported, but the molecular mechanisms involved were not elucidated. The presence of an ica operon in S. haemolyticus has been reported (17), but to date its contribution to biofilm formation is unclear.

The aim of this study was to characterize S. haemolyticus biofilm formation by a collection of 72 clinical isolates. In addition, the prevalence of genes associated with biofilm formation in CoNS was investigated.

(This work was partly presented as a poster at the 43rd ESCMID International Symposium, BAMBHAR, Palma de Mallorca, Spain, November 2007, and at the 48th Interscience Conference on Antimicrobial Agents/IDSA 46th Annual Meeting, Washington, DC, October 2008.)

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Patients and isolates. A total of 72 clinical isolates were included in this study. Fifty-one blood culture isolates were obtained from the Rikshospitalet (Oslo, Norway) over the period from 1992 to 2005. Twenty-one isolates were collected from the University Hospital of North Norway (Tromsø, Norway) over the period from 1989 to 2005; 14 isolates were from blood cultures and 7 isolates were from other clinical sources. The clinical isolates were obtained from neonates admitted to a neonatal intensive care unit (n = 32), immunosuppressed adult patients with cancer (n = 22), and adult patients treated in other hospital units (n = 18). The regional committee for medical research ethics approved the collection and analysis of patient data.

The control isolates used for the PCRs and the biofilm assays were S. epidermidis ATCC 35984 (RP62A), S. epidermidis ATCC 12228, S. epidermidis ATCC 27626, S. epidermidis 1457 (32), S. epidermidis 1457-M10 (33) Enterococcus faecium ATCC 51559, and S. aureus NCTC 8325.

Species identification and antibiotic susceptibility testing. All clinical isolates were identified as S. haemolyticus by using the ID32 Staph gallery (bioMèrieux, Marcy l'Etoile, France), according to the manufacturer's instructions. Species identification of all isolates was confirmed by 16S rRNA gene sequencing (37). For selected isolates, a 415-bp region of the sodA gene (38) or additional parts of the 16S rRNA gene were sequenced. The primers and amplicons used in this study are listed in Table 1. Vancomycin resistance was determined by Etest (AB Biodisk, Solna, Sweden). Susceptibility was determined according to Norwegian guidelines (4).

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TABLE 1. Oligonucleotide primers and PCR amplicons used in this studya

Analyses of genomic DNA. Total DNA was extracted in agarose plugs for pulsed-field gel electrophoresis (PFGE), as described previously (20). Chromosomal DNA was analyzed after SmaI digestion, and plasmid DNA was analyzed after S1 nuclease digestion, as described by Barton et al. (5). To differentiate between the chromosomal and the plasmid location of the ica operon, PFGE of S1- and SmaI-digested total DNA was performed. By this approach, S1 nuclease-digested plasmids are linearized to allow separation by PFGE, while the chromosomal DNA remains in the agarose plug.

Capillary blotting was performed with a Turboblotter system, according to the manufacturer's recommendations (Whatman Schleicher & Schuell, Kent, United Kingdom) by using 1 M Tris base-1.5 M NaCl as the transfer buffer, after the gel had first been denatured for 15 min in 0.25 M HCl, depurinated for 15 min in 1.5 M NaCl-0.5 M NaOH, and equilibrated twice for 15 min each time in 1 M Tris base-1.5 M NaCl. Hybridization and detection under high stringency conditions were performed as described previously, with minor modifications (20); i.e., we used an EZNA cycle pure kit (D6492-02; Omega Biotek Inc.) for purification of the PCR product used as the template for probe synthesis.

PCR. Template DNA for PCR amplification was prepared as described previously (20). The reaction mixture was based on Reddymix (catalog no. AB-0815; ABgene, Surrey, United Kingdom) to which MgCl₂ was added at a concentration of 4.5 mM and oligonucleotide primers were added at 0.4 pmol/sample. The primers specific for biofilm-associated genes (aap, atlE, bhp, icaD, ica, and fbe) and antibiotic resistance determinants are listed in Table 1.

DNA sequencing. The PCR products were purified by using an Exo/Sap PCR product presequencing kit (USB Corp.). Heterogeneity in the ica operon, atl, fbe, and the 16S rRNA gene was identified by cycle sequencing of both strands with a BigDye Terminator (version 3.1) cycle sequencing kit (Applied Biosystems, Warrington, United Kingdom) on an ABI Prism 377 sequence analyzer.

The sequences were edited by using the Chromas lite (version 2.01) program (Technylesium Pty. Ltd.) and were aligned by using the BioEdit sequence alignment editor (version 7.0.5.3) (T. Hall) and/or the SeqManII (version 5.50) program (DNAStar Inc., Madison, WI).

Phylogenetic analyses. To determine the overall genetic relationship between the isolates, the GelCompar (version 2.5) program (Applied Maths, Kortrijk, Belgium) was used to analyze the SmaI PFGE banding patterns. The Dice band-based similarity coefficient was calculated with a band position tolerance of 2%. PFGE dendrograms were constructed by the unweighted pair group method with arithmetic means. We defined clusters as three or more isolates with 80% similarity.

The phylogenetic relationship between the ica sequences was analyzed by constructing phylogenetic dendrograms from aligned sequences of a 372-bp fragment of the icaA gene and the ica operon from the five staphylococcal species whose operons had been fully sequenced. The accession numbers of the gene fragments or genes used for the alignment of icaA are given in the legend to Fig. 3. MEGA (version 4.0) software and maximum-parsimony and distance methods were used to generate the dendrograms. The topology was evaluated by bootstrap analyses with 1,000 replicates to give the confidence interval for each node in the dendrograms.

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FIG. 3. Phylogenetic tree based on 372-bp icaA sequences from 10 staphylococcal species. The tree was generated by using the neighbor-joining algorithm. Bootstrap values for internal nodes are given. The GenBank accession numbers for the sequences are as follows: S. epidermidis, NC_002976.3; S. saprophyticus, AF500270; S. cohnii, AF500268; S. condimenti, AF500266; S. simulans, AF500263; S. caprae, AF246926; S. capitis, AF500269; S. lugdunensis, EF546621; and S. haemolyticus, FJ472951.

Semiquantitative determination of biofilm formation. Semiquantitative determination of biofilm formation was performed as described previously (9, 25). All isolates were tested in eight wells in three parallel runs. For each parallel run, the highest and the lowest optical density (OD) values were removed to exclude outliers, and the remaining values were averaged. We tested the biofilm-forming capacities of all isolates under two different growth conditions: in Trypticase soy broth (TSB) supplemented with 1% glucose (TSB_glu) and in TSB supplemented with 3% NaCl (TSB_NaCl). A modified assay for the determination of biofilm production was performed by dissolving crystal violet from the biofilm with an ethanol-acetone mixture (70:30). Isolates were considered biofilm positive if they had an OD of 0.12 in the standard assay or an OD of 0.25 in the modified assay. Cutoff values were chosen to distinguish between isolates that produced significant amounts of biofilm and those that did not, taking into account the OD values for the negative controls included in each experiment.

S. epidermidis RP62A was used as a positive control (51). S. epidermidis ATCC 12228 (51), S. haemolyticus 51-03, and S. haemolyticus 8-79 were used as negative controls. The S. haemolyticus isolates were chosen from the material because they gave approximately the same ODs in the biofilm assay after growth in TSB_glu and/or TSB_NaCl as ATCC 12228 did after growth in TSB_NaCl.

Detachment assay. The major components of S. haemolyticus biofilms was analyzed by using (i) sodium meta-periodate (NaIO₄) to degrade β-1,6-linked polysaccharides, (ii) proteinase K to degrade proteins, and (iii) DNase I to degrade DNA (28, 40, 48). Briefly, biofilm formation and detection were performed as described above for the modified biofilm assay. Mature biofilms cultivated in TSB_glu and/or TSB_NaCl were washed three times with phosphate-buffered saline (PBS) and were treated with (i) 40 mM NaIO₄ in double-distilled H₂O, (ii) 0.1 mg/ml proteinase K (70663; Novagen) in 20 mM Tris-HCl (pH 7.5) with 100 mM NaCl, or (iii) 0.5 mg/ml DNase I (DN25; Sigma) in 5 mM MgCl₂ for 24 h at 37°C. S. epidermidis RP62A, for which the PIA is the most abundant matrix molecule, was included as a control (43).

Detachment assays were performed in quadruplicate wells in three parallel runs. For each parallel run, the highest and the lowest OD values were removed to exclude outliers, and the remaining values were averaged. Percent detachment was calculated on the basis of the average difference between the treated wells and the control wells. We divided the detachment results into three categories: no detachment (<10%), intermediate detachment (10 to 50%), and strong detachment (>50%).

Stability assay. The icaD-positive isolates were subjected to serial passages on blood plates (Oxoid blood agar base Cm0271, 5 M NaOH, 7% human blood) by transfer of a single colony to a new plate every day and successive incubation for 24 h at 37°C before a colony was transferred to a new plate. Colonies were harvested each day for DNA isolation, icaD PCR, and species identification by use of the ID32 Staph system. PCR of the 16S rRNA gene confirmed that all DNA extracts contained DNA.

PIA detection. PIA synthesis by a selection of isolates was tested by using a semiquantitative dot blot assay (1). In brief, isolates were grown overnight in TSB (Becton Dickinson, Cockeysville, MD) at 37°C before they were diluted 1:100 in fresh TSB_NaCl and grown overnight in tissue culture plates (Nunc, Roskilde, Denmark). Cells were harvested and washed once with PBS, and the cell suspensions were adjusted to an A₆₀₀ of 1. Cell surface-associated structures were then removed by the use of ultrasound, and the extracts were centrifuged. Serial dilutions of the supernatants were spotted onto polyvinylidene fluoride membranes that were blocked overnight with bovine serum albumin (10% [wt/vol] in PBS). After the membrane was rinsed with PBS, PIA was detected with a PIA-specific antiserum (30) preabsorbed with PIA-negative mutant 1457-M10 (33). Bound antibodies were made visible by using an enhanced chemiluminescence Western blotting detection kit (Amersham Biosciences), according to the manufacturer's instructions. S. epidermidis 1457 was used as a positive control.

CLSM. We investigated the structural differences between S. haemolyticus 51-10 and S. epidermidis RP62A using confocal laser scanning microscopy (CLSM). S. haemolyticus 51-10 is a strong biofilm producer (OD, 0.7) and PIA negative and showed a high level of detachment (66%) with proteinase K treatment.

One-milliliter aliquots of TSB_glu-diluted overnight cultures were used to grow biofilms on plastic cover slides for 24 h (Thermanox; Nunc) in 24-well microtiter plates (Falcon 3047; Becton Dickinson) at 37°C. After fixation for 1 min in 4% PBS-buffered paraformaldehyde solution (1x PBS with 4% paraformaldehyde) the cover slides were stained with wheat germ agglutinin (WGA; W11261; Invitrogen, Paisley, United Kingdom), which stains N-acetyl glucosaminyl residues, and 4',6-diamidino-2-phenylindole (DAPI; 32670; Sigma-Aldrich), which stains DNA.

Images were obtained on a TCS SP5 confocal laser scanning microscope (Leica Microsystems CMS GmbH, Mannheim, Germany) with a x63 1.2-numerical-aperture HCX PL APO water-immersion lens. For the detection of WGA (green), we used the 488-nm line of the argon laser and a detection bandwidth of 500 to 580 nm. For DAPI (blue), we used the 405-nm line and a detection bandwidth of 420 to 530 nm. The two fluorescent signals were collected sequentially at 400 Hz. Z stacks were exported as .tif files from Leica LAS AF (version 1.8.2) software and imported into the ImageJ (version 1.41) program (U.S. National Institutes of Health, Bethesda, MD). Orthogonal projections were generated for each channel by using the OrtView plug-in. The channels were merged, and image contrast and brightness were adjusted with the Photoshop CS3 program (Adobe Systems Incorporated).

Nucleotide sequence accession numbers. The sequences determined in this study were submitted to the EMBL/GenBank database and were assigned the following accession numbers: putative atlE, FJ472949; putative fbe, FJ472950; putative ica operon, FJ472951.

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Population structure. On the basis of SmaI macrorestriction enzyme analysis, the clonal relationship of 72 S. haemolyticus isolates was analyzed. Figure 1 shows the genetic diversity between the S. haemolyticus isolates. One large cluster consisting of 16 isolates and six smaller clusters consisting of 3 to 8 isolates each were identified. The largest cluster consisted of isolates from neonates admitted to the neonatal unit of Rikshospitalet. We found no association between the genetic relationship of the isolates and their biofilm-forming capacity.

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FIG. 1. Dendrogram generated from the PFGE profiles of 72 Staphylococcus haemolyticus by the unweighted pair group method with arithmetic means. The Dice band-based similarity coefficient was calculated with a band position tolerance of 2%. Clusters were designated when the branching points exceeded 80% similarity (7) and contained three or more isolates. The scale bar at the top of the dendrogram represents similarity.

Antibiotic resistance. The mecA gene, which codes for resistance to all beta-lactam antibiotics, was detected in 64/72 (89%) of the isolates. The most important gene coding for aminoglycoside resistance [aac(6')-Ie-aph(2''')] was detected in 61/72 (85%) of the isolates. Eighty-two percent of the isolates harbored both of these resistance determinants. No isolates were resistant to vancomycin. However, 10 (14%) isolates had MICs of 4 mg/ml, the breakpoint value between susceptible and intermediate resistance.

Phenotypic biofilm assay and biofilm induction. We observed that in polystyrene microtiter plates, the S. haemolyticus isolates commonly formed biofilms at the air-liquid interface. In the modified assay, we aimed to include all the crystal violet in the measurements. Thus, the proportion of isolates considered biofilm positive increased substantially by the modified assay. The growth medium had a strong effect on the proportion of isolates producing biofilms, and all isolates that were biofilm positive when they were cultivated in TSB_NaCl were also positive when they were cultivated in TSB_glu. By the modified assay, 53 (74%) isolates produced biofilms in TSB_glu, whereas only 22 (31%) isolates produced biofilms in TSB_NaCl. In contrast, by the original method (the standard crystal violet assay), 18 (25%) isolates produced biofilms in TSB_glu and 2 (3%) isolates produced biofilms in TSB_NaCl.

Two of the 72 isolates diverged from the rest of the collection in their biofilm-forming ability. Both produced more biofilm in TSB_NaCl than in TSB_glu, and both were ica positive. In the cluster analysis, these strains did not cluster together.

Prevalence and sequencing of biofilm-associated genes. Only three isolates were positive for icaD. Two of these were also positive for the additional biofilm-associated genes tested. S. haemolyticus 25-59 contained atlE, fbe, and icaD; S. haemolyticus 6-49 contained atlE and icaD; and S. haemolyticus 51-51 contained only icaD. All amplicons obtained from the icaD, fbe, and atlE PCRs were sequenced.

The sequences of three icaD amplicons were identical to the sequence of the corresponding region in S. epidermidis RP62A (GenBank accession no. CP000029). Primer sets Ica1 to Ica6 and Ica10 to Ica13 were designed for determination of the complete sequence of the ica operon in the three S. haemolyticus isolates. The complete ica operon was assembled, and all three sequences were identical. However, the last 100 bp of the icaR sequence of isolate 51-51 gave somewhat ambiguous results, which might be explained by the instability observed for this operon (see below).

Figure 2 presents the percent identity calculated for the full-length ica operon and the individual coding sequences to the different staphylococcal sequences available. The sequence of the S. haemolyticus ica operon showed an overall 99% similarity to the sequence of the strain RP62A ica operon (Fig. 2). The differences were located within the icaR and icaA genes. The S. haemolyticus icaA sequence was 98% identical to the corresponding S. epidermidis sequence, whereas the icaR sequence showed 99% identity.

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FIG. 2. Organization of the ica operon, as described for S. epidermidis RP62A at the top. The percent identities between the individual ica genes or the complete ica operon from S. haemolyticus and the different published ica sequences from four staphylococcal species are shown below the organizational diagram. S. lugdunensis does not have the icaR gene. The GenBank accession numbers for the sequences are as follows: S. epidermidis, NC_002976.3; S. caprae, AF246926; S. lugdunensis, EF546621; S. haemolyticus, FJ472951. NA, not available.

Figure 3 displays a phylogenetic dendrogram based on the 372-nucleotide region of the icaA sequence available in GenBank from different staphylococcal species. The dendrogram illustrates a close phylogenetic relationship between the icaA genes of S. epidermidis and S. haemolyticus. A similar phylogenetic relationship was found in an analysis based upon the five fully sequenced ica operons (data not shown).

The two 340-bp atlE sequences obtained were identical to each other and to the corresponding region in S. epidermidis RP62A (GenBank accession no. CP000029). The 505-bp sequence obtained from the fbe amplicon showed 1 nucleotide difference compared to the sequence of the corresponding region in RP62A (GenBank accession no. CP000029).

Southern blot hybridization-stability assay. Isolates positive for the ica operon by PCR were further investigated by Southern blot hybridization. A probe specific for a region of the ica operon (amplified by the Ica4 primer pair) was used. The hybridization confirmed the occurrence of the ica operon in two of the three isolates but not in S. haemolyticus 51-51. The hybridization pattern was consistent with a chromosomal location of the operon (data not shown). However, the ica operon in S. haemolyticus seemed to be unstable. All three ica-positive isolates showed an ica-negative genotype by PCR within four serial passages on nonselective blood agar. No changes in their phenotypic profiles, as determined with the ID32 Staph system, were detected. These results were reproduced twice.

Biofilm detachment assay. We used proteinase K, DNase, and NaIO₄ to investigate the relative contributions of proteins, DNA, and polysaccharides in the S. haemolyticus biofilm matrix, respectively. The effects of proteinase K and NaIO₄ were tested in biofilms induced with both TSB_glu and TSB_NaCl, whereas DNase was tested only after induction with TSB_glu. However, the results from the detachment assays followed the same pattern, independent of the cultivation conditions. Thus, only the detachment results after cultivation in TSB_glu are presented in Fig. 4.

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FIG. 4. Percent biofilm detachment of 53 biofilm-producing S. haemolyticus isolates after treatment with NaIO4 (light checks), proteinase K (dark checks), or DNase (stripes).

The positive control isolate, S. epidermidis RP62A, displayed a level of detachment of 90% when it was treated with NaIO₄ but displayed levels of detachment of only 10% and 20% after treatment with proteinase K and DNase, respectively. Among the 53 biofilm-positive S. haemolyticus isolates, DNase caused detachment in all (100%) isolates and proteinase K caused detachment in 52/53 (98%) isolates, while NaIO₄ caused detachment in only 20/53 (38%) isolates. The majority of the isolates showed intermediate detachment, and a few isolates showed strong detachment (Fig. 4). All isolates showing strong detachment were also rather strong biofilm producers (ODs, >0.5). None of the three ica operon-containing isolates showed detachment when they were treated with NaIO₄ under any of the conditions tested.

Dot blot assay for PIA synthesis. All 20 isolates showing intermediate or strong detachment after NaIO₄ treatment, as well as the 3 ica-positive isolates, were investigated for PIA synthesis in a dot blot assay. Two of the ica-positive isolates produced material that specifically reacted with a PIA-specific antiserum. However, the amount of PIA produced was less than that produced by S. epidermidis 1457 (data not shown). We did not find evidence of PIA synthesis in any of the other isolates investigated.

CLSM. Figure 5 exemplifies the morphological differences between the PIA-positive isolate S. epidermidis RP62A and biofilm-positive but PIA-negative isolate S. haemolyticus 51-10. S. epidermidis RP62A created a thick and complex biofilm structure with a large amount of extracellular polysaccharides, which stained green in our assay. In contrast, the protein-rich S. haemolyticus biofilm was thinner and showed a simpler three-dimensional structure with only small amounts of polysaccharides.

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FIG. 5. CLSM images of 24-h biofilms of S. epidermidis RP62A (A) and S. haemolyticus 51-10 (B) stained with DAPI (blue) and WGA (green). The S. epidermidis RP62A biofilm shows a complex three-dimensional structure, and polysaccharides dominate the extracellular matrix component. In comparison, the protein-rich biofilm of S. haemolyticus 51-10 is more condensed, and only small amounts of polysaccharides can be observed. Also, the RP62A biofilm is more than twice as thick as the S. haemolyticus biofilm.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
To our knowledge, the clinical S. haemolyticus isolates evaluated in the present study are the largest collection of such isolates analyzed for their biofilm-forming capacities. Our data confirm that S. haemolyticus is a significant pathogen in selected high-risk patient groups, such as neonates and immunosuppressed patients with cancer. We found a very high prevalence of genes encoding resistance to beta-lactam antibiotics and aminoglycosides, as well as a significant proportion of isolates with vancomycin MICs close to the defined breakpoint for resistance. This underlines the importance of S. haemolyticus as a multidrug-resistant pathogen that is difficult to treat with conventional antibiotics (6, 19).

A broad range of model systems for the in vitro study of biofilm formation have been described (36). The microtiter assay with crystal violet staining of biofilm formation at the bottom of the wells has been a standard method for the evaluation of S. epidermidis biofilm formation for many years (9). The visual investigation of S. haemolyticus biofilms showed that they frequently form biofilms at the liquid-air interface. We therefore recommend the use of a modified assay for the quantification of an S. haemolyticus biofilm. This allows determination of the total biomass in each well, as also described by Stepanovic et al. (44).

According to our results, biofilm formation by S. haemolyticus is induced when the organism is cultivated in TSB_glu. In contrast, a high NaCl concentration greatly reduced the level of biofilm production by ica-negative isolates. This is consistent with the findings of a previous study of an S. epidermidis ica-knockout mutant (23). In that study, the biofilm-forming phenotype was maintained by the mutant as a proteinaceous biofilm mainly consisting of Aap, but the phenotype was abolished by cultivation in growth medium containing high NaCl concentrations.

The low prevalence of the ica operon in our strain collection indicates that S. haemolyticus mainly forms PIA-independent biofilms. Both ica-positive strains exhibited only weak PIA synthesis. Furthermore, the detachment results indicate that ß-1,6-linked polysaccharides are probably not the major matrix component of PIA-independent biofilms. PIA-independent biofilm formation has also been reported in S. epidermidis, S. aureus, and S. lugdunensis (15, 18, 41, 43, 46). Only two isolates in our study showed >75% detachment when they were treated with NaIO₄, which is comparable to the level of detachment observed for S. epidermidis RP62A (43). However, both these isolates were ica negative, and they also showed >50% detachment when they were treated with proteinase K, indicating that proteins and polysaccharides play equal roles in their biofilms. Still, NaIO₄ induced biofilm disruption in 38% of the isolates, which may reflect the presence of hitherto unknown β-1,6-linked polysaccharides that are part of the biofilm matrix in S. haemolyticus. Further studies will have to identify and characterize the polysaccharide structures involved.

There is evidence that extracellular DNA may function as cell-to-surface and/or cell-to-cell adhesins in the initial phase of biofilm formation (50). Others have found that DNase treatment disrupted the biofilm if the DNase was introduced at the early stages of biofilm formation but that the DNase had no effect on a mature S. epidermidis biofilm (40). In our study, DNase disrupted the 24-h mature biofilm in all isolates. DNase treatment of biofilms has not been extensively studied. Furthermore, different protocols, DNase products, and DNase concentrations have been used. The DNase concentration used in our study caused only 20% detachment of the strong biofilm producer S. epidermidis RP62A. However, 33 of 53 S. haemolyticus isolates showed more than 25% detachment of the 24-h biofilm after DNase treatment. This potential important role of DNA in the mature biofilm matrix of S. haemolyticus is novel among CoNS.

Significant proteinase K-induced biofilm disruption was found in all but one isolate. Different proteins have been reported in CoNS biofilms. The best studied is Aap, which in S. epidermidis mediates intercellular adhesion and biofilm formation, independently or cooperatively, with the ica operon (26, 42). We did not detect aap or bhp in any isolate by PCR. Proteinaceous biofilms formed with hitherto unknown proteins have also been reported for S. epidermidis. Qin et al. investigated two ica-negative, biofilm-positive isolates that formed a proteinaceous biofilm (41). Both these isolates were negative for bhp and aap and therefore lacked the genotype for the well-known protein components in S. epidermidis biofilms. However, as the primers were designed from the published S. epidermidis gene sequences, we cannot exclude the possibility of primer failure in our assay. Further studies are needed to identify and purify the proteins involved in S. haemolyticus biofilm formation.

The images obtained by CLSM indicate that the three-dimensional biofilm structure of a protein-rich S. haemolyticus biofilm is quite different from that of the characteristic S. epidermidis PIA-positive biofilm.

In staphylococcal species like S. epidermidis, S. aureus (10, 52), and S. lugdunensis (18), alternative mechanisms of biofilm development seem to be a secondary choice or a response to more extreme environmental conditions. In our study, the biofilm-forming phenotype did not follow any specific clonal lineage but was spread throughout the collection of genetically diverse isolates. In S. haemolyticus, no pattern that indicates a predominant matrix component has arisen, and our results also indicate that biofilm formation may rely on the simultaneous contributions of several factors that function together.

icaADBC has been reported in several other CoNS species and has been sequenced (2, 10, 18). We confirmed by sequencing that the complete icaADBC operon was present in 3 of 72 S. haemolyticus isolates; thus, it was present at a low prevalence compared with the prevalence in S. epidermidis. In concurrence with previously reported sequences, we found a high degree of similarity between the ica operons in different staphylococcal species. Phylogenetic analyses show that the ica operon of S. haemolyticus is closely related to that of S. epidermidis. Thus, the ica operon may have been subject to recent genetic exchange between these two species. A high degree of instability of the S. haemolyticus genome, in which DNA fragments of up to 420 kb are deleted during serial passage, has been reported (45, 49). In our study, we discovered that the ica operon was very unstable. In the attempt to confirm the ica-positive PCR results by Southern blotting, only two of the three strains hybridized to the probe. However, this could be explained by the sensitivity of the PCR compared with that of Southern blotting.

The instability of the ica operon may be explained by several models: genomic rearrangements mediated by insertion elements have been observed in S. haemolyticus, as mentioned above (49). Furthermore, Nuryastuti et al. (35) showed that deletion of the ica operon from S. epidermidis can be caused by the deregulation of RecA, which is involved in genetic deletions and rearrangements.

We have investigated the biofilm formation phenotypes and genotypes of a large collection of clinical S. haemolyticus isolates. The S. haemolyticus biofilms showed high degrees of diversity in their biochemical profiles. The genetic background for biofilm formation is clearly different from what is commonly found in S. epidermidis. Further studies are needed to define the roles of the different components of S. haemolyticus biofilms and how they are regulated.

 
This study was supported by grants from the Northern Norway Regional Health Authority, Fredriksens legat, and the Deutsche Forschungsgemeinschaft (SFB470/C10 to H.R.).

We declare that none of the authors have any commercial relationship or other association that might pose a conflict of interest.

We thank Jorunn Pauline Cavanaugh, Merethe Sletteng, and the Bioimaging Core Facility at the University of Tromsø for excellent technical assistance.

 
* Corresponding author. Mailing address: Department of Microbiology and Virology, Institute of Medical Biology, MH Building, University of Tromsø, Tromsø, Norway. Phone: 47 77 64 46 63. Fax: 47 77 64 53 50. E-mail: johanna.e.sollid{at}uit.no

Published ahead of print on 14 January 2009.

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Journal of Clinical Microbiology, April 2009, p. 1172-1180, Vol. 47, No. 4
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