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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

ABSTRACT
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.

TEXT
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.
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%
CO
2 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-mm
2 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.
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-(1

4)-linked
D-glucopyranosyl units or
ß-(1

3)-
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).
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 (OD
630) (
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 KDO
2-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.
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 OD
600 of both the column flowthrough
and control samples was determined. Results were calculated
as the OD
600 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.
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.

ACKNOWLEDGMENTS
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.).

FOOTNOTES
* 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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