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Methods

Since 1999, nosocomial infections due to and colonization by MRSA have been surveyed monthly at Kagoshima University Hospital, a 700‐bed tertiary‐care hospital.³ We defined cases of colonization as those involving patients for whom culture results were positive for MRSA but for whom there was no evidence of tissue invasion. The diagnosis of nosocomial infection was based on the criteria proposed by the US Centers for Disease Control and Prevention.⁷ Nasal swab specimens were collected routinely for surveillance culture from patients undergoing invasive treatments, such as surgery or tracheal intubation. Culture specimens from the other sites were collected as usual for routine clinical culture. S. aureus was identified by colony morphological analysis and by coagulase testing. Resistance to methicillin was determined by subculturing the isolate on MRSA screening plates (Becton Dickinson) at 35°C for 24 hours. During a 3‐year period from 2000 through 2002, we found 121 patients who were nosocomially infected with MRSA and 287 patients who were colonized in the nose and pharynx. Because only 1 MRSA strain was selected from each patient, 121 isolates from nosocomial infection sites (the infection group) and 287 nasal or pharyngeal strains (the carrier group) were examined. The sources of the specimens in the infection group were as follows: surgical site, 49 (40.5%); sputum, 42 (34.7%); catheter, 18 (14.9%); pleural effusion, 10 (8.3%); and ascites, 2 (1.7%). The sources of the specimens in the carrier group were as follows: nasal cavity, 239 (83.3%) and pharynx, 48 (16.7%).

Biofilm formation by MRSA strains on polystyrene was quantified using the microtiter plate assay first described by Christensen et al.⁸ Briefly, strains were grown overnight in trypticase soy broth with 0.25% glucose. The next day, the strains were diluted 1:40 in trypticase soy broth with 0.25% glucose, and 200 μL of this suspension was inoculated in duplicate in a 96‐well, flat‐bottom, polystyrene microtiter plate and was incubated for 18 hours at 37°C without shaking. Next, the cells were decanted, and the wells were washed gently 4 times with tap water. The cells that remained in the wells were stained with 0.1% crystal violet for 5 min. The wells were washed 4 times with tap water. The stained cells were resolved by the addition of 200 μL of 95% ethanol. Absorbance was measured at 595 nm by use of an ELISA plate reader. The biofilm index was defined as the optical density at 595 nm (OD₅₉₅). To evaluate the reproducibility of this method, biofilm indexes of all the strains were measured twice independently by K.M. and J.N. Both authors' results correlated well with each other (correlation coefficient, 0.938; ; ).

The genotyping methods included spa typing, agr typing, pulsed‐field gel electrophoresis, mec‐hypervariable region polymerase chain reaction or sequence typing, and toxin genotyping, as described elsewhere.³ Four agr types based on the variability of agrD or agrC sequences were determined by use of the agr typing method using multiplex polymerase chain reaction, as described elsewhere.⁹ One universal forward primer and 4 type‐specific reverse primers were used. DNA was prepared for polymerase chain reaction by standard biochemical methods. Amplification reactions were performed in a Thermal Cycler PC 707 (ASTEC). The polymerase chain reaction products were electrophoresed on 1.2% agarose gels, and the agr type (1, 2, 3, or 4) was determined by the expected product size.

Statistical testing was performed using the Mann‐Whitney U test and Fisher’s exact test. was considered to be statistically significant.

Results

The examined isolates were classified into 297 genotypes: 91 genotypes in the infection group and 225 genotypes in the carrier group. Nineteen genotypes were observed in both groups. The members of the same genotype in each group were eliminated, except for one representative strain, so as not to count members of the same clone repeatedly as different test samples. Finally, 91 isolates in the infection group and 225 isolates in the carrier group were evaluated for biofilm formation. The sources of the specimens in the infection group were as follows: surgical site, 36 (39.6%); sputum, 32 (35.2%); catheter, 16 (17.6%); pleural effusion 6 (6.6%), and ascites 1 (1.1%). The sources of the specimens in the carrier group were as follows: nasal cavity, 191 (84.9%) and pharynx, 34 (15.1%).

A comparison of quantitative biofilm analysis in each group is shown in panel A of the Figure. The average biofilm index in the infection group (mean ± SD, OD₅₉₅; range, 0.020‐3.251 OD₅₉₅) was significantly higher than that in the carrier group (mean ± SD, OD₅₉₅; range, 0.001‐2.084 OD₅₉₅) ( ). We defined a strongly biofilm‐producing strain as one with a biofilm index higher than 0.304 OD₅₉₅, which was the 75th percentile in the carrier group. The incidence of strongly biofilm‐producing strains in the infection group (35/91 [38.5%]) was significantly higher than that in the carrier group (56/225 [24.9%]) ( ).

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Figure.  A, Quantitative analysis of biofilm formation by 91 methicillin‐resistant Staphylococcus aureus strains recovered from nosocomial infection sites (the infection group) and 225 nasal or pharyngeal strains recovered from asymptomatic carriers (the carrier group). B, Comparison of biofilm formation among the agr types: agr‐1, ; agr‐2, ; and agr‐3, . Box plots show median and the 10th, 25th, 75th, and 90th percentiles. OD₅₉₅, optical density at 595 nm.

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The average biofilm index of each specimen site was as follows: surgical site, OD₅₉₅ (range, 0.020‐3.251 OD₅₉₅); catheter, OD₅₉₅ (range, 0.036‐1.877 OD₅₉₅); sputum, OD₅₉₅ (range, 0.034‐2.372 OD₅₉₅); pleural effusion and ascites, OD₅₉₅ (range, 0.072‐0.868 OD₅₉₅); nasal cavity, OD₅₉₅ (range, 0.001‐2.084 OD₅₉₅); and pharynx, OD₅₉₅ (range, 0.003‐1.846 OD₅₉₅). There was no significant difference among the specimen sites within either group.

The prevalences of agr types in both groups are shown in the Table. The prevalence of agr‐1 strains in the infection group was significantly lower than that in the carrier group ( ), whereas that of agr‐2 strains in the infection group was significantly higher than that in the carrier group ( ). No agr‐4 strains were found in this study. A comparison of biofilm index by each agr type is shown in panel B of the Figure. The biofilm index of the agr‐1 strains (mean ± SD, OD₅₉₅; range, 0.001‐0.928 OD₅₉₅) was significantly lower than those of the agr‐2 strains (mean ± SD, OD₅₉₅; range, 0.012‐3.251 OD₅₉₅) ( ) and the agr‐3 strains (mean ± SD, OD₅₉₅; range, 0.011‐0.820 OD₅₉₅) ( ).

Table.  The Prevalence of Methicillin‐Resistant Staphylococcus aureus agr Types in the Carrier and Infection Groups.

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Discussion

In the present study, we found that MRSA strains recovered from patients with nosocomial infection sites were more likely to produce strong biofilm adhesions to polystyrene than were strains recovered from the nasal cavity or pharynx of asymptomatic carriers. This finding suggests that strongly biofilm‐producing MRSA strains are associated with nosocomial infection, such as surgical site infection and pneumonia. Using a tissue‐culture plate assay, Knobloch et al.¹⁰ compared the formation of biofilm by clinical S. aureus isolates recovered from patients with that by nasal S. aureus isolates recovered from healthy students. Consistent with our results, they found that significantly more clinical isolates than nasal isolates were able to form biofilms. To our knowledge, however, ours is the first report to study biofilm formation in an epidemiological study of MRSA in a hospital. Our study has the limitation of cross‐sectional approach. To investigate whether strongly biofilm‐producing MRSA strains preferentially induce nosocomial infections, further analysis, including a longitudinal study, will be needed.

Some strains in the carrier group produced strong biofilms. Previously, we reported that the prevalent genotypes in nonnasal strains causing nosocomial infections were also found in the nasal strains of asymptomatic carriers in our hospital.³ Strongly biofilm‐producing strains present in the nasal cavity may possibly cause nosocomial MRSA infections, if the carriers undergo invasive treatments. Thus, investigation of the biofilm formation of MRSA strains carried by patients preparing to undergo invasive treatments may be useful for the prevention of nosocomial infection.

The agr gene is a factor of the quorum‐sensing system in staphylococci that senses the density of bacteria.⁴ In our study, most of the agr‐1 strains showed poor biofilm formation, compared with agr‐2 and agr‐3 strains. We also found a lower prevalence of agr‐1 strains and a higher prevalence of agr‐2 strains in the infection group. Goerke et al.⁶ reported that the majority of S. aureus strains recovered from patients undergoing intubation and from patients with chronic wounds were type agr‐2, in contrast to those recovered from healthy control subjects. Our results are consistent with their finding and suggest that the agr type is associated with biofilm formation and nosocomial MRSA infection.