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Applied and Environmental Microbiology, October 1998, p. 3648-3655, Vol. 64, No. 10
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Vibrio cholerae O1 Strain TSI-4 Produces the
Exopolysaccharide Materials That Determine Colony Morphology,
Stress Resistance, and Biofilm Formation
Sun Nyunt
Wai,1,*
Yoshimitsu
Mizunoe,1
Akemi
Takade,1
Shun-Ichiro
Kawabata,2 and
Shin-Ichi
Yoshida1
Department of Bacteriology, Faculty of
Medicine,1 and
Department of
Biology, Faculty of Science,2 Kyushu
University, Fukuoka 812-8582, Japan
Received 12 May 1998/Accepted 5 August 1998
 |
ABSTRACT |
Vibrio cholerae O1 strain TSI-4 (El Tor, Ogawa) can
shift to a rugose colony morphology from its normal translucent colony morphology in response to nutrient starvation. We have investigated differences between the rugose and translucent forms of V. cholerae O1 strain TSI-4. Electron microscopic examination of the
rugose form of TSI-4 (TSI-4/R) revealed thick, electron-dense
exopolysaccharide materials surrounding polycationic ferritin-stained
cells, while the ferritin-stained material was absent around the
translucent form of TSI-4 (TSI-4/T). The exopolysaccharide produced by
V. cholerae TSI-4/R was found to have a composition of
N-acetyl-D-glucosamine, D-mannose, 6-deoxy-D-galactose, and
D-galactose (7.4:10.2:2.4:3.0). The expression of an
amorphous exopolysaccharide promotes biofilm development under static
culture conditions. Biofilm formation by the rugose strain was
determined by scanning electron microscopy, and most of the surface of
the film was colonized by actively dividing rod cells. The
corresponding rugose and translucent strains were compared for stress
resistance. By having exopolysaccharide materials, the rugose strains
acquired resistance to osmotic and oxidative stress. Our data indicated
that an exopolysaccharide material on the surface of the rugose strain
promoted biofilm formation and resistance to the effects of two
stressing agents.
| |
INTRODUCTION |
Cholera is a serious epidemic
disease that has killed millions of people and continues to be a major
health problem worldwide. Vibrio cholerae, the bacterium
that causes cholera, is a motile, gram-negative, curved rod with a
single polar flagellum. The hypothesis that V. cholerae
occupies an ecological niche in the estuarine environment requires that
this organism be able to survive the dynamics of various physiochemical
changes, including variations in nutrient concentrations. As a response
to nutrient depletion, copiotrophic (31, 42), heterotrophic
bacteria may undergo considerable morphological, physiological, and
chemical changes (13, 22, 23, 26-28). In fact, to survive
energy- and nutrient-deprived conditions, non-spore-forming,
heterotrophic bacteria are known to undergo an active adaptation
program (28). Brown and Williams have provided detailed
experimental evidence that the molecular composition of the bacterial
cell walls is essentially plastic and is remarkably responsive to the
cell's growth environment (5). Rice et al. (33)
discovered that V. cholerae O1 from the Peru epidemic
was able to shift to a phenotype having a wrinkled or rugose colony
morphology. They also suggested that the V. cholerae rugose phenotype represents a fully virulent survival form of the
organism that can persist in the presence of free chlorine. Morris et
al. (29) reported that V. cholerae can shift
to a rugose colony morphology associated with the expression of an amorphous exopolysaccharide (EPS) that promotes cell aggregation, and
they also confirmed that rugose strains displayed resistance to killing
by chlorine and complement-mediated serum bactericidal activity. They
also indicated that these rugose strains cause human disease. However,
the phenotypic characteristics associated with rugose morphology,
relationships between these characteristics, and their relative
importance in pathogenicity still remained to be identified.
A large variety of EPSs are synthesized by gram-negative bacteria.
While some have been implicated in the pathogenicity of plant and
mammalian hosts, others have not been assigned a function, but many
serve a structural role, benefiting the bacterium by enabling
attachment to surfaces, improving nutrient acquisition, or providing
protection from environmental stresses and host defenses (36). The EPSs cover the surfaces of many gram-negative and gram-positive bacteria. They may form a capsule composed of a high-molecular-weight polysaccharide attached to the cell surface, or
they may produce slime either loosely attached to the cell surface or
released to the culture fluid. Bacterial cells initiate the process of
irreversible adhesion by binding to the surface by using EPS glycocalyx
polymers and the development of microcolonies. The eventual production
of a continuous biofilm on the colonized surface is a function of cell
division within microcolonies and recruitment of bacteria from the
planktonic phase. The biofilm concept has drawn attention to the
bacterium's ecological and biotechnological importance
(8-11). We must now accept the unequivocal evidence that
bacteria respond to changes in their environment by profound phenotypic
variations in enzymatic activity, cell wall composition
(34), and surface structure (2).
In this study, we have isolated the rugose variants of V. cholerae O1 strain TSI-4 from starvation medium and determined EPS expression on the cell surface of the rugose strain by polycationic ferritin-labeled thin-section electron microscopy. While examining the
morphological characteristics of these rugose strains, we found that
they produced a continuous biofilm on the colonized surface and culture
tube walls. Directly sampled, intact biofilms were subjected to
electron microscopic analysis. We have also studied the role of the
slime polysaccharide of V. cholerae TSI-4 in the
bacterium's resistance to osmotic and oxidative stress.
| |
MATERIALS AND METHODS |
Organism and microcosm conditions.
V. cholerae O1
strain TSI-4 (El Tor Ogawa) was used in this study. Frozen stocks were
maintained at 
80°C in L broth (25) containing 50%
glycerol. The original isolate of strain TSI-4 had a translucent colony
morphology. Cells of TSI-4 were routinely grown at 37°C on a rotary
shaker in L broth. The culture was incubated to mid-log phase, which
corresponded to an A600 of 0.4. The cells were
then harvested by centrifugation (13,000 × g for 10 min), washed three times with cold M9 salts (37),
resuspended in starvation medium (M9 salts) to give a final
concentration of approximately 5 × 107 cells/ml, and
incubated at 16°C without shaking. Strain TSI-4 exhibits a shift of
colony morphology to the rugose form under starvation conditions at 2 months after inoculation. The rate of phase variation from the rugose
form to the translucent form was assessed by inoculating an isolated
rugose colony into L broth and incubating it overnight with shaking at
37°C and then plating serial dilutions of the bacteria onto L agar
incubated overnight at 37°C.
Polycationic ferritin labeling and electron microscopy.
Bacteria were grown on L agar overnight at 37°C, harvested, and
washed twice with cacodylate buffer (0.1 M, pH 7.0). Bacterial cells
were fixed for 2 h at 20°C in cacodylate buffer containing 5%
glutaraldehyde. Fixed bacteria were washed and resuspended in
cacodylate buffer and allowed to react with polycationic ferritin (Sigma Chemical Co., St. Louis, Mo.) (final concentration, 1.0 mg/ml)
for 30 min at 20°C (12, 15, 19). The samples were diluted
1:10 with the same buffer, and the bacterial cells were centrifuged and
washed three times in cacodylate buffer. Bacterial cells were then
immobilized in 4% agar, postfixed for 2 h with 2% osmium
tetroxide, and then washed three times. The samples were dehydrated in
a graded series of acetone washes, washed twice in propylene oxide, and
then embedded in Epon by a rapid embedding technique (30).
Thin sections were poststained with uranyl acetate and lead citrate and
examined with a JEM 2000EX electron microscope (JEOL, Ltd., Tokyo,
Japan) at 100 kV.
Antiserum.
An anti-EPS antibody was prepared by the
specific-adsorption method from the serum of a rabbit immunized with
rugose variant strain TSI-4/R as follows. A rabbit was immunized with a
bacterial suspension (108 CFU/ml) of strain TSI-4/R by
three injections at 5-day intervals, and 7 days after the last
injection, blood was collected from the ear vein. Antibodies other than
the anti-EPS antibody were removed from the serum by the adsorption
technique. About 2 ml of the serum and about 109 CFU of
heat-inactivated strain TSI-4/T (translucent variant) per ml were
mixed, and the mixture was stirred during overnight incubation at
4°C. The bacteria were then removed by centrifugation at 5,000 × g for 10 min, and the supernatant was again mixed with the same strain. This adsorption process was repeated three times, and
the final supernatant was used as anti-EPS serum. The final titer of
the antiserum was measured by tube agglutination. Strain TSI-4/R was
used as the antigen to determine the titer of the anti-EPS antibody.
The agglutination titer of anti-EPS serum against TSI-4/R was 1:256,
and it did not agglutinate strain TSI-4/T.
Immunoelectron microscopy.
For immunoelectron microscopy, a
colloidal gold probe (Wako Pure Chemical Industries Ltd., Osaka, Japan)
was used to label the specific reaction sites of anti-EPS serum and an
anti-O1 monoclonal antibody (mouse immunoglobulin G anti-V.
cholerae monoclonal antibody reacting with lipopolysaccharide
[LPS] epitopes present in the bacterial cell wall). The
anti-V. cholerae monoclonal antibody was purchased from
Cosmo Bio Co., Ltd. (6). To label the specimens, 1 ml of a
bacterial suspension of about 108 CFU/ml was treated with
antiserum appropriately diluted with phosphate-buffered saline (PBS)
for 30 min at 37°C. The bacteria then were separated from the serum
by centrifugation at 12,000 × g for 10 min. After
being washed three times with PBS, the bacteria were mixed with a
suspension of the colloidal gold probe, and the mixture was kept at
room temperature for 30 min. After being washed with PBS to remove
nonreacted gold particles, the bacteria were processed for thin-section
electron microscopy (1).
EPS isolation and purification.
Cells were harvested from a
24-h culture on an L agar plate and resuspended in physiological saline
(0.85% sodium chloride). Samples were centrifuged at 1,600 × g for 20 min, and supernatants were dialyzed with multiple
changes of distilled water. The specimens were then ultracentrifuged at
154,000 × g for 15 h at 20°C, and the
supernatants were removed and subjected to enzymatic digestion with
RNase (100 µg/ml), DNase I (50 µg/ml plus 1 mM MgCl2),
and pronase (250 µg/ml), followed by sequential phenol-chloroform extraction (32). Purity was assessed by lack of detectable
protein on silver-stained sodium dodecyl sulfate-polyacrylamide gel and by wavelength scanning spectrophotometric analysis (Milton Roy Spectronic Genesys 5 spectrophotometer; Milton Co. Ltd., New York, N.Y.). The total neutral sugar content was determined by the
phenol-sulfuric acid method (14). Qualitative and
quantitative sugar analyses were carried out after pyridylamination by
using a Palstation pyridylamination reagent kit (Takara Biomedicals,
Kyoto, Japan) (35).
Outer membrane and LPS preparation.
The outer membrane was
prepared from a broth culture of either strain TSI-4/T or strain
TSI-4/R by the method of Filip et al. (16). LPS was prepared
from 1 ml of an overnight culture (18). LPS and outer
membrane samples were electrophoresed and detected by silver staining
as previously described (24).
Preparation of DNA.
The high-molecular-weight chromosomal
DNAs of V. cholerae TSI-4/T and TSI-4/R were prepared
essentially by the method of Berns and Thomas (4). Plasmid
DNA was isolated by the alkaline method (25).
HindIII, BglII, and PstI were used
for chromosomal DNA and plasmid fingerprinting (40).
Scanning electron microscopy.
V. cholerae TSI-4
was cultured in L broth at 37°C without shaking for 3 to 5 days. The
biofilms growing on the upper surface of the L broth and on the wall of
a culture tube were sampled for scanning electron microscopy. The
specimens were fixed with 2% glutaraldehyde in PBS for 1 h
followed by 1% osmium tetroxide overnight at 4°C, dehydrated with a
series of acetone concentrations ranging from 50 to 100%, dried by the
critical-point drying method, coated with gold-palladium for surface
conductivity, and examined with a scanning image observing device
(ASID) equipped with a JEOL JEM 2000EX electron microscope.
Stress resistance assay.
Stress resistance was assessed by
using a 1:1,000 dilution of an overnight culture of either TSI-4/T or
TSI-4/R in 3.0 ml of minimal glucose medium. The diluted cultures were
grown to approximately 2 × 109 cells per ml, which
corresponded to an A600 of 0.4, and cells were
harvested by centrifugation and resuspended in minimal glucose medium
(pH 7.0) which contained either 20 mM H2O2
(oxidative stress) or 2.5 M NaCl (osmotic stress). Survival was
measured at various times, depending upon the type of stress applied.
Percent survival was calculated by dividing the number of CFUs at a
given time by the number of CFUs at time zero and multiplying the
result by 100. One hundred percent viability was taken to be
approximately 2 × 109 cells per ml and represents the
viable counts obtained immediately before challenge. Percent survival
of TSI-4/T and that of TSI-4/R were compared (see Fig. 6).
Statistical analysis.
Data are expressed as means ± standard errors. Differences between experimental groups were analyzed
by Student's t test (unpaired), and a P of
<0.05 was accepted as statistically significant.
| |
RESULTS |
Isolation of the rugose strain and phase variation.
We have
found that V. cholerae O1 strain TSI-4 is able to shift
to a phenotype having a rugose colony morphology under starvation conditions. To isolate spontaneous rugose variants, bacteria from smooth colonies were starved in M9 salts for 2 months at 16°C and
then plated on L agar at 37°C. TSI-4 underwent phase variation, converting from translucent to rugose in M9 salts and back again at a
low rate in L broth. Rugose TSI-4/R colonies inoculated into L broth
and subcultured on L agar produced translucent TSI-4/T colonies at a
frequency of 1.5 × 10
5. The two distinct colony
morphologies are shown in Fig. 1.
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FIG. 1.
Photomicrograph of V. cholerae O1 rugose
TSI-4/R (arrow) and translucent TSI-4/T (arrowhead) colonies.
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Thin-section electron microscopy.
To determine the nature of
the colony morphology differences, bacterial pellets were stained with
polycationic ferritin and thin sections were observed by electron
microscopy. Representative profiles are shown in Fig.
2. EPS materials of TSI-4/R were
recognized as a heavy, fibrous, electron-dense, ferritin-stained layer
completely surrounding the cell (Fig. 2A), but TSI-4/T did not appear
to have this external layer surrounding its cells (Fig. 2B). The staining of the extracellular fibrous layer by polycationic ferritin strongly suggests the presence of acidic polysaccharide, and it shows
the electrostatic binding of a derivative of ferritin to anionic sites
on the cell surface (12, 20).
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FIG. 2.
Thin sections of V. cholerae stained
with polycationic ferritin showing a thick, electron-dense EPS layer
completely surrounding TSI-4/R cells (A) and the absence of this layer
surrounding a cell of TSI-4/T (B). Bars, 0.5 µm.
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Immunoelectron microscopy.
The reactivity of the
anti-V. cholerae O1 monoclonal antibody with the
cell surfaces was tested with both strains TSI-4/R and TSI-4/T. The
reaction of anti-O1 serum with both strains is shown in Fig.
3. The gold particles were specifically
found on the outer membrane surfaces and were believed to be the
antibody reacting with the surface O antigens of both strains. The
anti-EPS serum prepared by specific adsorption of anti-TSI-4/R serum
with strain TSI-4/T was reactive only with TSI-4/R and not with TSI-4/T (Fig. 4A and B). The gold particles were
specifically bound to the EPS material on the surface of strain TSI-4/R
and at the intercellular spaces (Fig. 4A).
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FIG. 3.
Immunoelectron micrographs of the surface labeling of
strains TSI-4/R (A) and TSI-4/S (B) with anti-V.
cholerae O1 monoclonal antibody. Bars, 0.5 µm.
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FIG. 4.
Strains TSI-4/R (A) and TSI-4/T (B) treated with
anti-EPS serum and labeled with a gold probe. Bars, 0.5 µm.
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LPS and outer membrane profiles.
Additional testing of the two
morphotypes was undertaken to identify differences which might
contribute to different colony morphologies, outer membrane proteins
and LPS were prepared from both cell types of TSI-4, electrophoresed,
and detected. No outer membrane protein or LPS differences between cell
types were detected (data not shown). To further ensure that the rugose
strain was a phenotypic variant of V. cholerae O1
strain TSI-4, we also performed plasmid and DNA fingerprint analyses.
The plasmid and DNA patterns of the two strains were the same (data not
shown).
EPS extraction.
The purified EPS from TSI-4/R was quantified
by the phenol-sulfuric acid method of Dubois et al. (14).
The reaction of extracted EPS material with phenol and sulfuric acid
confirms the presence of sugars and suggests that these sugars are
hexoses and methylhexoses because of a characteristic absorbance
peak at 490 nm. Spectrophotometric scanning of the purified
EPS also showed a lack of absorbance at 260 or 280 nm, indicating an
absence of contaminating protein or nucleic acids. Qualitative and
quantitative sugar analyses showed that the extracellular
polysaccharide of V. cholerae TSI-4/R contains
N-acetyl-D-glucosamine, D-mannose,
6-deoxy-D-galactose, and D-galactose at a
molar ratio of 7.4:10.2:2.4:3.0.
Biofilm growth of V. cholerae TSI-4/R and scanning
electron microscopy.
The biofilms of V. cholerae
TSI-4/R were clearly visible on the upper surface of L broth and
culture tube walls after 5 days of static incubation at 37°C, whereas
TSI-4/T did not have the biofilm-forming property and produced a smooth
suspension of bacteria. Figure 5 shows a
biofilm under scanning electron microscopy; the surface of the
film was completely covered with a layer of contiguous bacterial
cells embedded within a polymeric matrix. Throughout the
biofilm, cells were interconnected by a finger-like glycocalyx matrix that extended from the substratum to the outer boundaries of the
biofilm.
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FIG. 5.
Scanning electron micrographs of biofilm formation by
V. cholerae O1 strain TSI-4/R. (A) Most of the surface
has been colonized with actively dividing rod cells, and finger-like
projections of extracellular polymeric material are present. Bar, 5 µm. (B) High magnification indicates the presence of extracellular
polymeric materials on the surfaces of bacterial cells. Bar, 1 µm.
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Rugose TSI-4 exhibits elevated resistance to osmotic and oxidative
stress.
Cells of both TSI-4/T and TSI-4/R were collected at
mid-exponential phase and tested as described in Materials and
Methods. V. cholerae O1 strain TSI-4/R was much more
resistant to oxidative and osmotic stress than was strain TSI-4/T,
showing viability more than 10 times greater than that of strain
TSI-4/T (Fig. 6).
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FIG. 6.
Increased resistance of strain TSI-4/R to oxidative
stress (A) and osmotic stress (B). Strains TSI-4/R and TSI-4/T were
collected at mid-exponential phase and tested as described in Materials
and Methods. The time points for oxidative challenge
(H2O2) were 0, 5, 10, and 15 min, and the time
points for osmotic challenge (NaCl) were 0, 15, 30, 45, and 60 min.
Survival is expressed as the percentage of the initial cell input that
survived the treatment. Bars show standard errors (each point is the
average of three separate experiments). The P values
(t test) for the 10-min time point in the oxidative stress
resistance test and the 30-min time point in the osmotic stress
resistance test are <0.001 and <0.01, respectively.
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| |
DISCUSSION |
When bacteria are transported from one environment to another, the
environmental changes with which they are confronted include changes in
temperature, nutrient concentration, salinity, osmotic pressure, pH,
and many other factors. However, bacterial cells dynamically adapt to
shifts in environmental parameters by employing a variety of
genotypic and phenotypic mechanisms (7). Starvation-induced changes in bacterial surfaces have been reported for several
strains of marine bacteria by Kjelleberg and Hermansson
(21). The cells of V. cholerae demonstrate a
predilection for association with chitinaceous surfaces and the
mucilaginous sheath of algae that can be interpreted as an ecological
advantage (3).
In this paper, we report that the growth of V. cholerae
O1 strain TSI-4 (El Tor, Ogawa) in starvation medium resulted in a change to a wrinkled or rugose colony morphology from the normal translucent colony morphology and, at the same time, significant production of EPSs, some of which are loosely attached to the cell
surface and some of which are released into the intercellular spaces.
Analogies to rugosity can be found in a number of other bacterial
species, including the expression of alginate by mucoid strains of
Pseudomonas aeruginosa and the expression of an adhesive EPS
by members of the marine genus Hyphomonas. Wrangstadh et al. (41) demonstrated that energy and nutrient starvation of
marine Pseudomonas sp. strain S9 induced the production and
release of an EPS with resulting pronounced effects on the adhesion and
aggregation of the bacterial cells. Our data indicate that cell surface
EPS materials confer a rugose colony morphology, biofilm formation ability, and resistance to osmotic and oxidative stress. We suggest that the spontaneous and reversible variation in cell-associated and
cell-free EPS production represents an optimal adaptive mechanism that
facilitates survival in stressful environments.
The rugose form of V. cholerae was first described in
1938 by Bruce White, who recognized that it might serve as a survival form of the organism (39). Rice et al. (33)
suggested that the V. cholerae rugose phenotype
represents a fully virulent survival form of the organism that can
persist in the presence of free chlorine and that this phenotype may
limit the usefulness of chlorination in blocking the endemic and
epidemic spread of cholera. Morris et al. (29) have
supported and confirmed his finding that rugose strains appear to
produce an EPS that promotes cell aggregation and causes human disease.
This aggregation may shield individual cells from killing by
disinfectants, such as chlorine, or lysis by complement. He also
suggested that the EPS produced by V. cholerae plays a
role in marine biofilm formation; this, in turn, may contribute to
attachment of bacteria to marine organisms, such as plankton. Our
findings confirm and support their suggestions that EPS-producing rugose vibrios promote cell aggregation in a shaking culture at stationary phase and that this adhesive EPS and rugose vibrio can form
a biofilm in a static culture. We identified the biofilm of TSI-4/R
macroscopically and electron microscopically. EPS materials were also
identified by ferritin-labeled thin-section electron microscopy.
Polycationic ferritin has previously been used to visualize the
capsular material of many organisms, such as Klebsiella sp.
(38), Neisseria gonorrhoeae (17), and
non-O1 V. cholerae (20).
Routine practices in clinical laboratories and researchers working with
V. cholerae have routinely selected smooth colonies from culture plates for investigation. A rugose colony is easily confused with a contaminated colony because of its unusual morphology and is likely to be dismissed as an avirulent rough variant or a
contaminant. This rugose form can survive in the presence of the
first-line disinfectant chlorine and remain fully virulent (29,
33). Our studies extend previous findings by indicating that the
rugose strain acquired high resistance to stress conditions and biofilm
formation ability. EPS materials of this rugose variant are considered
to be important for shielding of the organism from adverse
environmental conditions. The rugose form might serve as a survival
form of the organism. The ability of V. cholerae O1 to
assume this phenotype and survive environmental stress is very
important in the ecology of this organism.
| |
ACKNOWLEDGMENTS |
We thank H. Nakayama for the scientific discussion.
This work was supported by a grant from the Japan Society for the
Promotion of Science.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Bacteriology, Faculty of Medicine, Kyushu University, Fukuoka 812-8582, Japan. Phone: 81-92-642-6130. Fax: 81-92-642-6133. E-mail:
sunwai{at}bact.med.kyushu-u.ac.jp.
| |
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0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
