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Previous Article | Next Article 
Journal of Bacteriology, October 2001, p. 5848-5854, Vol. 183, No. 20
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.20.5848-5854.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Salmonella enterica Serovar Typhimurium Swarming
Mutants with Altered Biofilm-Forming Abilities: Surfactin Inhibits
Biofilm Formation
Joe Robert
Mireles II,
Adam
Toguchi, and
Rasika M.
Harshey*
Section of Molecular Genetics and
Microbiology and Institute of Cellular and Molecular Biology,
University of Texas at Austin, Austin, Texas 78712
Received 8 March 2001/Accepted 16 July 2001
 |
ABSTRACT |
Swarming motility plays an important role in surface colonization
by several flagellated bacteria. Swarmer cells are specially adapted to
rapidly translocate over agar surfaces by virtue of their more
numerous flagella, longer cell length, and encasement of slime.
The external slime provides the milieu for motility and likely harbors
swarming signals. We recently reported the isolation of
swarming-defective transposon mutants of Salmonella enterica serovar Typhimurium, a large majority of which were
defective in lipopolysaccharide (LPS) synthesis. Here, we have examined the biofilm-forming abilities of the swarming mutants using a microtiter plate assay. A whole spectrum of efficiencies were observed,
with LPS mutants being generally more proficient than wild-type
organisms in biofilm formation. Since we have postulated that O-antigen
may serve a surfactant function during swarming, we tested the effect
of the biosurfactant surfactin on biofilm formation. We report that
surfactin inhibits biofilm formation of wild-type S.
enterica grown either in polyvinyl chloride microtiter wells or
in urethral catheters. Other bio- and chemical surfactants tested had
similar effects.
| |
INTRODUCTION |
Swarming bacteria are prime
examples of how prokaryotes interact with and adapt to surface
environments (5, 6, 8, 11, 12). Upon propagation on a
surface, many flagellated bacteria turn on a new genetic program that
leads to formation of swarmer cells, which are typically longer and
have more flagella than cells of the same species grown in
aqueous conditions (swimmer cells). These morphological changes appear
to be necessary to overcome frictional forces presented by a surface,
allowing colony expansion. Another important feature of swarming
colonies is their encapsulation in slime (a generic term for a mixture
of carbohydrates, proteins, peptides, surfactants, etc.), which not
only provides an essential milieu for swarming motility, but likely
harbors signals for swarmer cell differentiation (18).
We recently reported the isolation of a large number of transposon
mutants of Salmonella enterica serovar Typhimurium that affect swarming motility (18). A majority of these mutants
were defective in lipopolysaccharide (LPS) synthesis, many were
defective in chemotaxis, and some had defects in putative two-component signaling components. While the latter two classes were defective in
swarmer cell differentiation, representative LPS mutants were not and
could be rescued for swarming by external addition of a biosurfactant.
We have suggested that the O-antigen improves surface wettability,
required for swarm colony expansion, and that multiple two-component
systems cooperate to promote swarmer cell differentiation.
In contrast to the expansionist behavior of swarming colonies, there is
a prevalence in nature of microbial colonies that remain attached to a
surface in associations called biofilms (2). Biofilms are
composed of exopolysaccharides or slime secreted by the adherent
bacteria (20). Genetic studies have shown that, like the
swarming bacteria, single-species biofilms form in multiple steps
(21), require intercellular signaling (4),
and show a pattern of gene expression that is distinct from that of
planktonic cells (16). Bacteria that have formed adherent
biofilms exist not as a tightly packed unit but rather as columns of
loosely associated cells, some fixed, others motile. Water channels
between pillars of cells in such biofilms allow nutrients to disperse. Biofilms are medically and industrially important because they can
accumulate on a wide variety of substrates and are resistant to
antimicrobial agents and detergents. What feature of a biofilm allows
adherence in one case and expansion in another?
It is clear that swarming and biofilm formation have common features,
such as generation of a large bacterial mass within which cells
interact closely and generation of slime, which promotes association or
colonization of the growth surface. In order to investigate whether our
swarming mutants might shed light on the expansionist versus adherent
nature of the two types of surface colonies, we tested these mutants
for their ability to form biofilms in polyvinyl chloride (PVC)
microtiter wells. We find an inverse relationship between the presence
of a wettability factor (either LPS or surfactin) and biofilm
formation; thus, the presence of the wettability factor favors swarming
but inhibits biofilm formation, and vice versa.
| |
MATERIALS AND METHODS |
Bacterial strains, media, and chemicals.
S.
enterica serovar Typhimurium SJW1103 (a fliC stable
derivative of serovar Typhimurium LT2) and its transposon mutants have been described (18). The bacteria were grown in
Luria-Bertani (LB) medium. Surfactin, sodium dodecyl sulfate (SDS), and
Tween 80 were purchased from Sigma Chemical Co. Rhamnolipid (extracted from Pseudomonas aeruginosa) was a gift from Jeneil
Biosurfactant Co. Inc. PVC 96-well microtiter plates were from Costar
(2596; nontreated). Clear vinyl urethral catheters (14Fr) were from
Kendall Co., Mansfield, Mass.
Quantification of biofilm formation in microtiter wells.
The
quantification assay was performed as previously described
(14). Typically, 10 µl of an overnight culture was used
to inoculate PVC microtiter wells containing 90 µl of LB without NaCl
but with 2% glucose. The covered microtiter dish was sealed with
Parafilm during incubation at 30°C. Cultures were removed to
determine the optical density at 630 nm (OD630),
and the wells were rinsed with distilled water. After drying at room
temperature for 15 min, 200 µl of crystal violet (1%) was added to
the wells for 20 min. The stained biofilms were rinsed several times
with distilled water, allowed to dry at room temperature for 15 min, and extracted twice with 200 µl of 95% ethanol. The
OD550 was estimated using a Beckman DU-640B
spectrophotometer after adjusting the volume to 1 ml with distilled water.
Visualizing biofilm formation in catheters.
An overnight
culture of S. enterica (10 µl) was inoculated into 500 µl of medium and injected into clear vinyl urethral catheters. The
catheters were capped at both ends and incubated at 30°C overnight. Media and growth conditions were as described above for PVC wells. Cultures were removed to determine the OD630, and
the catheters were rinsed with distilled water. After drying at room
temperature for 15 min, 700 µl of crystal violet (1%) was added to
the catheters for 20 min. The stained biofilms were rinsed several
times with distilled water and allowed to dry at room temperature for
15 min before examination.
| |
RESULTS AND DISCUSSION |
Analysis of biofilm formation by wild-type S.
enterica in microtiter wells.
The biofilm assay used in
this study monitors the ability of S. enterica to attach to
the wells of microtiter dishes (14). The biofilm forms at
the interface between the air and the liquid medium and is quantitated
by staining with crystal violet as described in Materials and Methods.
Initial experiments with different abiotic materials (PVC,
polystyrene, and borosilicate glass) showed that the wild-type
strain SJW1103 forms the best biofilms on PVC, in LB broth without
NaCl but with 0.2% glucose, and at 30°C; these conditions were
used throughout. SJW1103 did not form a detectable biofilm in M9
minimal medium, consistent with the results reported for
Escherichia coli (15). Figure
1 shows the kinetics of biofilm formation
by wild-type S. enterica. The exponential phase of biofilm formation coincided with that of cell growth. Biofilm formation began
to slow around 13 h, decreased up to 17 h, and then leveled off, coincident with entry of the culture into stationary phase.
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FIG. 1.
Kinetics of biofilm formation by wild-type S.
enterica. SJW1103 was cultured at 30°C in PVC 96-well plates
containing LB without NaCl plus 0.2% glucose. Samples were harvested
at designated time points to determine growth (OD630) and
biofilm formation after crystal violet staining (OD550) as
described in the text.  , OD630;  ,
OD550.
|
|
Analysis of biofilm formation by swarming mutants.
Fifty-nine
S. enterica swarming mutants (18), all
completely defective in swarming, were tested for their ability to form biofilms. A subset of these is shown in Fig.
2. Also included in
this analysis were four mutants with mutations that did not affect
swarming (luxS [18]; 1, 2, and 3 are random
Tn10dCm insertions in SJW1103). Biofilms were harvested when
cultures reached an OD630 of approximately 0.2. The data (arranged in increasing biofilm formation efficiency) are
shown in Fig. 2. Overall, a continuous spectrum of biofilm formation
was observed in the mutants, from 10 to 156% of the wild-type level.
Except for the waaP and waaL mutants, all the LPS
mutants clustered at the far end of this spectrum. Mutants not affected
in swarming showed biofilm formation similar to that of wild-type cells
(Fig. 2, arrowheads and arrow, respectively). Specific mutants are
discussed below.
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FIG. 2.
Biofilm formation by S. enterica swarming
mutants. Growth conditions were as described in the legend to Fig. 1.
The OD630 of all samples was approximately 0.2. Arrow
points to biofilm levels (OD550) of wild-type SJW1103, and
arrowheads point to those of mutants not affected in swarming. The data
are means of triplicate samples monitored on the same day. There was a
significant variation in the absolute biofilm values of samples
processed on different days, but the relative values with respect to
the wild-type value were very reproducible.
|
|
As shown previously with E. coli (15), both
motA and motB mutants were significantly
defective in biofilm formation, reaching only about 10% of the
wild-type level, while the che mutants were less defective,
attaining between 38 and 62% of wild-type levels. Biofilm formation by
mutants not defective in swarming (Fig. 2, arrowheads) was within 10 to
20% of the wild-type level. Among the swarming mutants, a whole range
of biofilm formation levels was observed. We note that the crystal
violet staining technique is only a measure of adherence and that this
analysis only measures gross biofilm formation. For example, in the
lasI mutant of P. aeruginosa, total biofilm
formation was unaffected but the structure was physically different
and, in contrast to the wild-type parent, could be dislodged with 0.2%
SDS (4). Similarly, a colanic acid mutant of E. coli showed normal crystal violet staining, but when examined
under the microscope, the mutant did not display the complex pillars of
cells typical of the wild-type strain (3). Thus, it is
possible that mutants that appear to be unaffected by this assay could
have different structures to their biofilms. If we consider only the
very low or very high end of the biofilm formation spectrum, then of
the uncharacterized two-component signaling mutations, baeS
is at the low end and yfhk and yojN are towards
the middle. At the high end there are a cluster of LPS mutations,
waaK, waaC, wbaP, wzx, and
ddhC. The kinetics of biofilm formation for one of these
mutants (ddhC) was determined to be similar to the wild-type
kinetics in the initial phase of growth; however, the mutant continued
to accumulate more biofilm after the wild type had begun to slow (data
not shown). None of the LPS mutants (or the wild type) formed biofilm
on a hydrophilic surface such as glass under these growth conditions
(data not shown). Attempts to measure cell surface hydrophobicity of
the wild type and the mutants by measuring adherence to hydrocarbons (17) were not successful.
Effect of a surfactant on biofilm formation.
We have recently
shown that the swarming defect of the LPS mutants could be rescued by
addition of the biosurfactant surfactin isolated from Bacillus
subtilis (18). To analyze the effect of surfactin on
biofilm formation, either the PVC wells were coated with surfactin or
surfactin was included in the growth medium. Figure
3 shows that increasing amounts of
surfactin led to a decrease in the amount of biofilm formed by the
wild-type strain and that 5 µg of surfactin was more than sufficient
to completely abolish biofilm formation. Bacterial growth was
unaffected under all surfactin concentrations tested (data not shown).
Surfactin had a similar inhibitory effect on biofilm formation by the
ddhC mutant (data not shown).
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FIG. 3.
Surfactin inhibits biofilm formation by wild-type
S. enterica. PVC wells were coated with the indicated
amounts of surfactin before inoculation with S.
enterica. After overnight incubation at 30°C, the wells were
rinsed out and stained with crystal violet. The biofilm is concentrated
at the interface between the air and the liquid medium (indicated by
arrows).
|
|
To test if surfactin would dislodge a preformed biofilm, surfactin was
added to the PVC wells after the culture had reached an
OD630 of 
0.15 to 0.2 (Fig.
4A). The OD550 of
the surfactin-treated sample decreased at a faster rate than that of
the untreated sample for the initial sloughing phase of biofilm
formation, resulting in an approximately 85% decrease in total biofilm
by the end of the experiment at 22 h (Fig. 4B). To determine
if other surfactants would have similar effects on preformed biofilms,
we tested SDS (ionic surfactant), Tween 80 (anionic surfactant), and
rhamnolipid. While all the chemicals tested dispersed the preformed
biofilm, SDS and rhamnolipid decreased the OD630
of the culture as well (Fig. 4A). (Surfactin concentrations in this and
the rest of the experiments were maintained at 100 µg in order to
compare its activity to that of the biosurfactant rhamnolipid, which
affected biofilm formation at higher concentrations.)
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FIG. 4.
Addition of surfactin and other surfactants to a
preformed biofilm accelerates biofilm dispersal. After S.
enterica had reached an OD630 of 0.15 to 0.2, the indicated surfactants were gently mixed into the cultures in
microtiter wells. Samples were harvested, and either growth (A),
as determined by OD630, or biofilm levels (B), as measured
by OD550 of crystal violet-stained material, were analyzed
at the indicated time points.  , no treatment;  , 100 µg of
surfactin; ×, 0.25% Tween 80; , 100 µg of rhamnolipid;  ,
0.2% SDS. The data are means of triplicate samples monitored on the
same day.
|
|
To investigate the biofilm-forming ability of bacteria known to
produce surfactants, we tested Serratia marcescens
and B. subtilis as well as mutants defective in
surfactant production. S. marcescens mutants
defective in production of the surfactant serrawettin are unable to
swarm (9, 13), as are surfactin mutants of B. subtilis (obtained from P. Zuber, Oregon Graduate Institute of
Science and Technology; our unpublished data). Mutants of S. marcescens that were defective in serrawettin synthesis made approximately threefold more biofilm that their wild-type parent
(Fig. 5). These results are
consistent with the notion that absence of the biosurfactant promotes
biofilm formation. To test if surfactin would inhibit biofilm formation
by the serrawettin mutants, surfactin was included in parallel wells
(Fig. 5). A significant reduction in biofilm formation was observed for
all the strains in the presence of surfactin. Purified serrawettin (gift from T. Matsuyama, University of Niigata, Niigata, Japan), had an inhibitory effect on biofilm formation but was not as effective as surfactin (data not shown). We were unable to observe biofilm formation by wild-type B. subtilis or its surfactin mutants
under these conditions (data not shown).
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FIG. 5.
Biofilm formation by S.
marcescens and its mutants. Wild-type S.
marcescens 274 and its serrawettin mutants (SMu1a,
SMu4e, SMu13a, and SMu4853b [9]) were analyzed for
biofilm formation in the absence and presence of surfactin as described
in the legend to Fig. 3. The OD630 of all the cultures was
0.15 (not shown). Open bars, no surfactin; shaded bars, 100 µg of
surfactin.
|
|
Surfactin prevents biofilm formation on urethral catheters.
To
test the effect of surfactin on medically relevant objects, we grew
S. enterica in clear vinyl urethral catheters. The stained
catheters are shown in Fig. 6. The
biofilm formed by S. enterica was dispersed all along the
growth surface. Surfactin eliminated formation of the biofilm.
Precoating the catheters by running the surfactin solution through them
prior to inoculation with medium was just as effective as including
surfactin in the growth medium (data not shown). Among other
surfactants tested on S. enterica, Tween 80 (0.25%) was as
effective as surfactin, while rhamnolipid seemed to be only half as
effective (data not shown). Given the importance of opportunistic
infections with Salmonella species, including urinary tract
infections of AIDS patients (7), these results have the
potential for practical applications.
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FIG. 6.
Surfactin inhibits biofilm formation on urethral
catheters. The indicated organisms were grown overnight at 30°C in
sterile urethral catheters containing medium with and without 100 µg
of surfactin as described in the text. Biofilms were analyzed by
staining with crystal violet.
|
|
To test the effect of surfactin on biofilm formation by other medically
relevant organisms, we grew E. coli, Proteus
mirabilis, and Pseudomonas aeruginosa in urethral
catheters. E. coli and Proteus mirabilis formed a
biofilm mainly at the air-liquid interface, while the biofilm formed by
P. aeruginosa, like that formed by S. enterica,
was dispersed all along the catheter (Fig. 6). Surfactin inhibited
biofilm formation (but not growth) by all organisms except P. aeruginosa. Similar inhibition profiles were obtained with Tween
80 (data not shown). We note that there are earlier reports on
inhibition of biofilms formed by uropathogens and yeasts on silicone
rubber by biosurfactants produced by Streptococcus thermophilus and lactobacilli (1, 19).
Summary.
The main conclusions from this work can be summarized
as follows. At least one difference between adherent and moving
biofilms may lie in the surfactant composition of the slime, since the absence of surfactants inhibits swarming but promotes biofilm formation
and vice versa. O-antigen mutants of S. enterica, which are
defective in swarming (18), were generally more proficient in biofilm formation than the wild type (Fig. 2). O-antigen has been
postulated to provide a surfactant or wettability function that allows
spreading of the swarmer colony (18). Surfactin, a
cyclic lipopeptide from B. subtilis which was
shown to rescue the swarming defect of the LPS mutants
(18), inhibited biofilm formation (Fig. 3). Mutants of
S. marcescens defective in serrawettin production
(serrawettin is an analog of surfactin [10]) are defective in swarming (9, 13) and make better biofilms
(Fig. 5). Surfactin (as well as the chemical surfactant Tween 80)
dispersed preformed biofilms without affecting cell growth (Fig. 4) and also prevented biofilm formation in urethral catheters by organisms such as S. enterica, E. coli, and P. mirabilis (Fig. 6). The nonbactericidal nature of these compounds
would be advantageous in preventing the selection of resistant
organisms if used for inhibiting biofilm formation. Biofilms formed by
P. aeruginosa were not affected by the surfactants tested in
this study (Fig. 6). Our results suggest the need for additional
research into the use of this genre of compounds for biofilm
containment. Since P. aeruginosa did not form
biofilms on a hydrophilic glass surface under our growth conditions,
manipulating the hydrophilicity of the growth surface might provide
another avenue to the control of this class of biofilms.
| |
ACKNOWLEDGMENT |
This work was supported by a grant from NIH (GM57400).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Section of
Molecular Genetics and Microbiology and Institute of Cellular and
Molecular Biology, University of Texas at Austin, Austin, TX 78712. Phone: (512) 471-6881. Fax: (512) 471-7088. E-mail:
rasika{at}uts.cc.utexas.edu.
| |
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Journal of Bacteriology, October 2001, p. 5848-5854, Vol. 183, No. 20
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.20.5848-5854.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
