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Infection and Immunity, October 2004, p. 6132-6138, Vol. 72, No. 10
0019-9567/04/$08.00+0     DOI: 10.1128/IAI.72.10.6132-6138.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Biofilm Formation by Neisseria meningitidis

Kyungcheol Yi,^1* Andrew W. Rasmussen,¹ Seshu K. Gudlavalleti,¹ David S. Stephens,^1,2 and Igor Stojiljkovic¹

Department of Microbiology and Immunology,¹ Division of Infectious Diseases, Department of Medicine, Emory University School of Medicine, and Laboratories of Microbial Pathogenesis, Veterans Affairs Medical Center, Atlanta, Georgia²

Received 12 April 2004/ Returned for modification 7 May 2004/ Accepted 26 June 2004

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Biofilm formation by the human pathogen Neisseria meningitidis was analyzed. Biofilm-forming meningococcal strains were identified and quantitated by crystal violet staining. Laser scanning confocal microscopy of the meningococcal biofilm revealed variable layers up to 90 µm in thickness. A total of 39 meningococcal isolates were studied; 23 were nasopharyngeal-carriage isolates, and 16 were invasive-disease isolates. Thirty percent of carriage isolates and 12.5% of invasive-disease isolates formed biofilms proficiently on a polystyrene surface. Generally, the strains that formed biofilms showed high-level cell surface hydrophobicity, characteristic of strains lacking a capsule. The inhibitory role of capsule in biofilm formation was further confirmed by comparing the biofilm-forming capabilities of a serogroup B wild-type strain of a disease-associated isolate to those of its capsule-deficient mutant (ctrA). Some strains of meningococci form biofilms, and this process is likely important in menigococcal colonization.

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Bacterial biofilms are sessile bacterial communities that adhere to each other and solid surfaces and are enclosed in an exopolysaccharide matrix (6). Biofilms are the predominant communities of many bacterial species in numerous ecosystems. Formation of biofilms involves participation of the extracellular-matrix and cellular-surface molecules, including membrane proteins. Biofilm formation also requires considerable bacterial energy and resources. The formation of biofilms begins with the attachment of the planktonic cells to a suitable surface, followed by replication and spreading. Eventually, the biofilms mature to differentiated forms. Exopolysaccharides play a key role in the establishment of biofilm architecture (6).

In clinical settings, bacteria in biofilms are less susceptible to antimicrobial agents and host immune responses, thereby becoming persistent colonizers or sources of chronic infections (8). Bacteria are released from biofilms as individual planktonic cells or as a result of the sloughing of the biofilms. While many biofilms form on abiotic surfaces such as medical devices, some also develop on living tissues, as in the case of endocarditis or cystic fibrosis (8).

Studies of biofilm formation by the Neisseria species are very limited, and most of those species examined have been oral commensals (4, 20, 24, 26, 38). Biofilm formation by Neisseria meningitidis, an etiologic agent of epidemic sepsis and bacterial meningitis, has not been documented. Meningococci are isolated from 5 to 10% of the normal population, and the colonization of the human nasopharyngeal mucosal surface by meningococci is the first step of the host-parasite interaction. Successful meningococcal colonization requires initial attachment facilitated by pili and subsequent interaction of other secondary-surface molecules with the host mucosal surface (12, 31, 36, 43).

In this study, the formation of the biofilms by N. meningitidis was assessed. In addition, the roles of the bacterial-surface molecules (pilus, capsule, and lipooligosaccharide [LOS]) in the biofilm formation were also determined. The meningococcal strains used in this study are listed in Table 1. Meningococci were grown on GC medium base (GCB) (Difco) agar containing Kellog's supplements and incubated at 37°C and 5% CO₂ tension or in liquid cultures in GC broth (1.5% proteose peptone no. 3 [Difco], 0.4% K₂HPO₄ [Sigma], 0.1% KH₂PO₄ [Sigma], and 0.5% NaCl [Sigma] plus Kellog's supplements I and II) at 37°C and 5% CO₂ with agitation. When necessary, streptomycin (750 µg/ml) was added to the medium.

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TABLE 1. Genetically modified meningococcal strains used in the study

One of the biofilm-forming strains was selected for the microscopic examination by confocal laser scanning microscopy. A 1:100 dilution of an overnight culture in GCB of strain IR3501 (Table 2), a serogroup B meningococcal-carriage isolate, was inoculated into 10 ml of fresh GCB in a sterile polystyrene 100- by 15-mm petri dish containing sterile borosilicate glass coverslips. The coverslips were removed after overnight incubation at 37°C in a 5% CO₂ atmosphere without agitation and carefully rinsed with fresh GCB. The coverslips were then stained with 30 µg of acridine orange (Sigma)/ml, rinsed again with GCB, and mounted onto a microscope slide as follows: a circle 10 mm in diameter was cut out of the middle of a 25-mm² piece of parafilm, greased on both sides with stopcock grease, and laid on the slide. Fresh GCB was applied to the center well in the parafilm, and the coverslip was fitted over the top of the parafilm gasket. The slide was then observed microscopically. All confocal microscopy experiments were performed on an Axiovert 135 inverted microscope equipped with an LSM-410 inverted confocal laser scanning microscope (Carl Zeiss, Jena, Germany). A x40 1.2-numerical-aperture C-Apochromat objective lens was used for observing the biofilms. All images were captured with a charge-coupled-device camera. Cross-sections were captured in 2-µm sections through the 100-µm sample depth of the N. meningitidis biofilm unless otherwise indicated. Excitation and emission were 488 and 510 to 525 nm, respectively. The examination showed a thick layer of meningococci and matrix material that ranged from less than 30 µm to above 60 µm (Fig. 1A). A sagittal view of the biofilm additionally shows the thick but variable layer of meningococci and matrix (Fig. 1B). Potential pillars of the biofilms are indicated.

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TABLE 2. Meningococcal nasopharyngeal-carriage and invasive-disease isolates

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FIG. 1. Confocal laser scanning microscopy of biofilms formed by meningococcal strain IR3501, a nasopharyngeal-carriage isolate. (A) Topographical illustration of the biofilms formed by IR3501. The composite images were computer generated after the biofilms on a coverslip were scanned horizontally by a confocal laser scanning microscope. Virtual color changes represent the height of the biofilm above the coverslip. A color-coded scale of biofilm depth is located in the upper left corner. (B) Sagittal view of a section of the biofilm. Various sizes of potential biofilm pillars are indicated by the letter P.

Formation of biofilms by strain IR3501 over a 12-h time course was assessed by Congo red staining and light microscopy (Fig. 2). Congo red stains starch, amylose, and polysaccharides containing contiguous ß-(14)-linked D-glucopyranosyl units or ß-(13)-D-glucans and has been used to detect exopolysaccharides constituting the extracellular materials of biofilms (28, 41, 45). At various time points, bacterial exopolysaccharides were visualized according to a modification of the staining method of Harrison-Balestra et al. (18). Ten microliters of an overnight culture was inoculated into 10 ml of fresh GCB in a sterile polystyrene 100- by 15-mm petri dish (1:1,000 dilution) containing sterile borosilicate glass coverslips. At various times, the coverslips were removed, and cetylpyridinium chloride (Sigma) (10 mM) was applied for 30 s and then discarded. The coverslips were air dried for 20 to 30 min. The specimens were stained with a 2:1 (vol/vol) mixture of saturated Congo red (Sigma) solution and 10% Tween 80 (Sigma) for 15 min after gentle heat fixation. Subsequently, the meningococci were stained with 10% Carbol Fuschin for 6 min. After air drying, the coverslips were mounted on slides and observed by light microscopy as noted above. After 5 h of inoculation, microcolonies were observed (Fig. 2a). The microcolonies expanded to form structures of biofilms and exopolysaccharides after 7 and 9 h of incubation (Fig. 2b and c). This coincides with previous observations that exopolysaccharides are not visible after 5 h or later (10, 17, 44). The biofilms organized after 12 h, and exopolysaccharides were more visible than at earlier time points (Fig. 2d).

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FIG. 2. Microscopic views of the meningococcal biofilm formation of strain 3501 (x1,000 magnification, light microscopy). The process of biofilm formation was visualized by double staining meningococci grown on coverslips with Congo red and Carbol Fuschin at 5 h (a), 7 h (b), 9 h (c), and 12 h (d). Congo red stains the exopolysaccharides pink (indicated by arrows). Meningococci were stained purple.

In contrast to the results seen with strain IR3501, serogroup B meningococcal strain IR4127 did not form a biofilm (Table 1 and Fig. 3). To examine the effect of surface-exposed molecules on the formation of biofilms, isogenic mutants of IR4127 were examined. These included mutants that had defects in capsule (ctrA mutant; strain IR5390) or that produced truncated LOS (rfaC mutant; strain IR5389) (Table 1). To visualize biofilms, bacteria were inoculated at a 1:100 dilution from the overnight culture and grown as described previously for 24 h in 24-well polystyrene plates containing 500 µl of GCB. The wells containing bacterial culture were stained with 300 µl of 0.3% crystal violet (CV; Difco) per well for 2 min after two washes with distilled water. The stained wells were subsequently washed with distilled water twice to remove residual CV. The stained biofilms were dissolved in 33% acetic acid and quantitated by measuring optical density at 630 nm (OD₆₃₀) (19). The proficiency of biofilms was quantitated by measuring the OD of dissolved CV in acetic acid. The OD values from the wells that had not been inoculated with bacteria were used as the negative control. The cutoff value for determining a biofilm producer was set as two times the negative-control value. The capsule-deficient mutant formed biofilms which were visualized by CV staining (Fig. 3B). The rfaC mutant with a KDO₂-lipid A LOS molecule also formed a biofilm at levels lower than those seen with the capsule mutant strain (Fig. 3A). These results indicated that the capsule and LOS oligosaccharide - and ß-chain structures play inhibitory roles in the biofilm formation and that meningococci can develop biofilm most effectively when capsule is absent or LOS truncated. The absence of the negatively charged capsule, in particular, suggests that hydrophobic interactions may play a significant role in biofilm formation.

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FIG. 3. Crystal violet staining of biofilms and quantitation. Biofilms formed by meningococcal mutants of rfaC (strain IR5389) and ctrA (strain IR5390) were compared with those formed by the parent strain (IR4127 [strain NMB, an invasive-disease isolate]) by visualization with crystal violet staining on a polystyrene surface (B). The amount of biofilm was quantitatively measured as readings at OD630 (x axis) (A).

To better understand the role of hydrophobicity in meningococcal biofilm formation, cell surface hydrophobicity was measured for selected strains. The surface hydrophobicity of meningococcal strains was measured using a modification of a previous protocol (21). Disposable plastic columns packed with octyl Sepharose CL-4B (Sigma) to a height of 2 cm were washed with 10 ml of buffer A (0.2 M ammonium sulfate in 10 mM sodium phosphate buffer; pH 6.8). Meningococci collected from overnight plate cultures were suspended in phosphate-buffered saline to an OD of 10, and a 100-ml aliquot was gently pipetted onto the surface of the column and eluted with 5 ml of buffer A. A 100-µl cell suspension diluted directly into 5 ml of buffer A was also prepared as a control. The OD₆₀₀ of both the column flowthrough and control samples was determined. Results were calculated as the OD₆₀₀ of the flowthrough divided by that of the control and expressed as the percentage of menigococci adsorbed to the column. The amount of biofilm determined by CV staining was compared with the cell surface hydrophobicity (Fig. 4). The plot indicates that the proficiency of biofilm formation correlates with the surface hydrophobicity.

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FIG. 4. Correlation between cell surface hydrophobicity and biofilm formation. Four biofilm-negative strains and eight biofilm-forming strains were assessed for their percent cell surface hydrophobicity (CSH) (y axis) versus biofilm formation as determined by crystal violet quantification in OD630 units (x axis). CSH is presented as the percentage of retained bacteria in the octyl-Sepharose column. The trend is shown as a straight line, with correlation coefficient r2 equal to 0.8056.

A pilQ mutant (IR5545) of the biofilm-forming strain (IR3501) was examined to evaluate the role of pili in biofilm formation. PilQ is part of the pilus secretion system (16) and is required for pilus formation. The pilQ mutant was nonpiliated and developed biofilms on the polystyrene surface that were indistinguishable in the way that they stained with CV from the biofilms of the parent strain. However, further analysis of the pilQ mutant biofilm by confocal laser-scanning microscopy revealed that the mutant formed biofilms that ranged from 20 to 30 µm in thickness (data not shown). This indicates that the meningococcal pilus may affect the architecture of the biofilm but not the quantity of the biofilm.

A total of 16 disease strains and 23 carriage strains were screened for biofilm formation on the polystyrene surface by crystal violet staining (Table 2). Of these, 30% (7 of 23) of the carriage strains formed biofilms whereas 12% (2 of 16) of invasive-disease isolates formed biofilms, suggesting that biofilm-forming strains are found more often in carriage strains.

Conclusions. Bacterial biofilms are found on a range of biotic and abiotic surfaces (2, 8, 23). Biofilms can consist of a single species or multiple species that show commensalism and competitive behavior within the biofilm milieu (5). While mixed-species biofilms predominate in nature, single-species biofilms can be found in clinical settings, including medical implants (1, 3, 13). The present study focused on a single-species biofilm formed by the human pathogen N. meningitidis. Typical architectural structures of the biofilms reveal water channels between pillars, which are used to deliver the nutrients necessary for bacterial survival (7). Meningococci formed pillars of cells that are approximately 50 to 60 µm in height. Cell surface molecules, including pili, flagella, lipopolysaccharides (LPS), and outer membrane proteins as well as secreted materials such as exopolysaccharides, are involved in the formation of the biofilms in Pseudomonas aeruginosa and Escherichia coli (9, 14, 25, 27, 32, 33). The identity of the meningococcal exopolysaccharides is of interest. No homologous genes were identified in a search of the meningococcal genome for the genes encoding colanic acid of E. coli and alginate of P. aeruginosa.

The role of pili in the formation of biofims in P. aeruginosa and E. coli has been extensively studied (14, 33, 34). Type IV pilus mutants of P. aeruginosa do not form biofilms, suggesting that pili are required for biofilm formation (33). However, growth on citrate minimal medium eliminates the need for pili on initial attachment or microcolony formation on surfaces and pilus-deficient mutants can form flat biofilms (25). This study also showed that twitching motility mediated by type IV pili was responsible for the migration of the microcolonies. The meningococcal pilQ mutant used in this study formed biofilms with a structure thinner than that of the biofilms formed by the wild-type strain. Altered biofilm structure formed by the pilQ mutant indicates a potential role for pili in meningcoccal biofilm formation.

LPS also seems to play an inhibitory role in the biofilm formation. Mutants of Salmonella enterica that produce truncated LPS structures form more proficient biofilms than the wild-type bacteria that produce elongated LPS structures (29). This agrees with our observation that the meningococcal LOS mutant (rfaC) is more proficient in biofilm formation than its parental strain. The mutant has a truncated LOS structure (KDO₂-lipid A) devoid of heptose and - and ß-chain oligosaccharides, including the -chain-terminal sialic acid. The absence of the structures may attenuate the steric hindrance and negative charges on the meningococcal surfaces, thereby allowing increased intimate contact between bacterial cells. This hypothesis was further supported by our observation that the LOS mutant autoaggregates more rapidly than the parental strain.

Our study indicates that prevalence of biofilm formation among carriage isolates is greater than that of disease isolates. Not surprisingly, this may be due to capsular expression. The serotyping and surface hydrophobicity profiles suggest that the majority of the non-biofilm-forming strains are encapsulated. Further, in contrast to the encapsulated parent a mutant defective in capsule expression gained the ability to form a biofilm. This suggests that all meningococci have the necessary cellular machinery to form biofilms, for example, when phase variation mechanisms (e.g., slipped strand mispairing) turn off capsule expression. The maintenance of biofilm-forming capabilities despite a metabolic burden suggests a selective advantage. Several lines of evidence suggest that the absence or down regulation of capsule promotes meningococcal colonization, possibly aided by biofilm formation. In natural settings, approximately 30% of the carriage strains are nonserogroupable (15). Following a community-based intervention program, a higher percentage of ctrA-negative isolates was recovered from a school population under study, suggesting that unencapsulated strains recolonize more rapidly than capsulated strains (37). At the molecular level, genes for capsule biosynthesis are down regulated upon contact with the host epithelium, facilitating meningococcal adherence (11). Additionally, gene transfer occurs more efficiently in the biofilms (30); this could also be a selective advantage for meningococci in the nasopharynx.

 
We thank Tony Romeo and William Shafer for critical review of the manuscript. We also thank Konstantin Agladze for help provided with confocal microscopy.

This work was supported by public service grant R01-AI42870-01A1 (to I.S.).

 
* Corresponding author. Mailing address: Department of Microbiology and Immunology, Emory University School of Medicine, 1510 Clifton Rd. NE, Atlanta, GA 30322. Phone: (404) 727-5968. Fax: (404) 727-3659. E-mail: kyi{at}emory.edu.

Editor: J. T. Barbieri

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Infection and Immunity, October 2004, p. 6132-6138, Vol. 72, No. 10
0019-9567/04/$08.00+0     DOI: 10.1128/IAI.72.10.6132-6138.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

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