Pellicle formation in Shewanella oneidensis

School of Minerals processing and Bioengineering, Central south University, Changsha, PR China.
BMC Microbiology (Impact Factor: 2.73). 11/2010; 10(1):291. DOI: 10.1186/1471-2180-10-291
Source: PubMed
ABSTRACT
Although solid surface-associated biofilm development of S. oneidensis has been extensively studied in recent years, pellicles formed at the air-liquid interface are largely overlooked. The goal of this work was to understand basic requirements and mechanism of pellicle formation in S. oneidensis.
We demonstrated that pellicle formation can be completed when oxygen and certain cations were present. Ca(II), Mn(II), Cu(II), and Zn(II) were essential for the process evidenced by fully rescuing pellicle formation of S. oneidensis from the EDTA treatment while Mg (II), Fe(II), and Fe(III) were much less effective. Proteins rather than DNA were crucial in pellicle formation and the major exopolysaccharides may be rich in mannose. Mutational analysis revealed that flagella were not required for pellicle formation but flagellum-less mutants delayed pellicle development substantially, likely due to reduced growth in static media. The analysis also demonstrated that AggA type I secretion system was essential in formation of pellicles but not of solid surface-associated biofilms in S. oneidensis.
This systematic characterization of pellicle formation shed lights on our understanding of biofilm formation in S. oneidensis and indicated that the pellicle may serve as a good research model for studying bacterial communities.

Full-text

Available from: Haichun Gao
RESEARC H ARTIC L E Open Access
Pellicle formation in Shewanella oneidensis
Yili Liang
1,2
, Haichun Gao
2,3*
, Jingrong Chen
2
, Yangyang Dong
3
, Lin Wu
3
, Zhili He
1
, Xueduan Liu
1
, Guanzhou Qiu
1
,
Jizhong Zhou
1,2*
Abstract
Background: Although solid surface- associated biofilm development of S. oneidensis has been extensively studied
in recent years, pellicles formed at the air-liquid interface are largely overlooked. The goal of this work was to
understand basic requirements and mechanism of pellicle formation in S. oneidensis.
Results: We demonstrated that pellicle formation can be completed when oxygen and certain cations were
present. Ca(II), Mn(II), Cu(II), and Zn(II) were essential for the process evidenced by fully rescuing pellicle formation
of S. oneidensis from the EDTA treatment while Mg (II), Fe(II), and Fe(III) were much less effective. Proteins rather
than DNA were crucial in pellicle formation and the major exopolysaccharides may be rich in mannose. Mutational
analysis revealed that flagella were not required for pellicle format ion but flagellum-less mutants delayed pellicle
development substantially, likely due to reduced growth in static media. The analysis also demonstrated that AggA
type I secretion system was essential in formation of pellicles but not of solid surface-associated biofilms in S.
oneidensis.
Conclusion: This systematic cha racterization of pellicle formation shed lights on our understanding of biofilm
formation in S. oneidensis and indicated that the pellicle may serve as a good research model for studying bacterial
communities.
Background
Most microbes in natural ecosystems exist in highly
organized and functional interactive communities, which
are composed of cells attached to surfaces and/or to
each other either from a single species or multiple spe-
cies [1-7]. Microbial communities confer a number of
advantages for survival, such as nutrient availability with
metabolic cooperati on, acquis ition of new ge netic trait s,
and protection from the environment [4,8]. The most
common microbial communiti es are biofilms, which
refer to assemblages of cell on solid biotic or abiotic
surfaces. In recent years, the subject of microbial bio-
films has drawn a lot of attenti on and numerous studies
have provided important insights into the genetic basis
of biofilm development [5,7].
Pellicles, arising from the interface between air and
liquid and therefore frequently called air-liquid (A-L)
biofilms [9], have been well studied i n an array of
bacteria, such as Bacillus subtilis, Pseudomonas aerugi-
nosa,andVibrio parahaemolyticus [7,10-12]. Pellicle
formation consists of at least three distinctive steps: (i)
initial attachment of bacteria to the solid surface (wall
of culture device) at the interface between air and liquid,
(ii) development of the monolaye r pellicle initiated from
the a ttached cells, and (iii) maturation of pellicles with
characteristic three-dimensional a rchitecture [1,11]. In
addition to cells, a variety of compone nts, mainly extra-
cellular polymeric substances (EPS), are nee ded for
developing and maintaining the pellicle matrix. The
most extensively studied EPS include exopolysacchar-
ides, proteins, and extracellular DNA although contribu-
tions of these agents to the integrity of the pellicle
matrix may vary [11]. While the pellicle is ge nerally
taken into account as a special form of biofilms [5,7,13],
its distinguishing characteristics justify that this type of
biofilm may serve as an independent researc h model
[12-14].
Many factors, including ext racellular organelles such
as flagella and type IV pili, secreted proteins, and chemi-
cal agents supplemented in media such as iron and
phosphate, have been shown to play important roles in
* Correspondence: haichung@zju.edu.cn; jzhou@ou.edu
1
School of Minerals processing and Bioengineering, Central south University,
Changsha, 410083, PR China
2
Institute for Environmental Genomics and Department of Botany and
Microbiology, University of Oklahoma, Norman, 73019, USA
Full list of author information is available at the end of the article
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
© 2010 Liang et al; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons
Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in
any medium, provided the or iginal work is properly cited.
Page 1
biofilm formation [5]. However, effects of these factors
on the biofilm formation process depend on the bacter-
ium under study. For example, flagella facilitate surf ace
adhesion fo r many species but it has been also observed
in other species that mutations resulting in aflagellate
and paralyzed nonmotile cells promote formation of a
multilayer biofilm [7]. In the case of iron, results are
even more inconsistent. In P. aeruginosa and Vibrio cho-
lerae, i ron limit ation hinders biofilm formation whereas
it facilitates the process in Actinomyces naeslundii and
Staphylococcus epidermidis [15,16]. It has been sug-
gested that variation in effects of these factors on bio-
film formation by particular species of bacteria may be
reflection of the different environmental niches where
they live [14,17-19].
Shewanella oneidensis MR-1, a facultative Gram- nega-
tive anaerobe with a remarka ble respiratory versatility,
has been extensively studied for its biofilm de velopment
[20-26]. However, little progress has been made to
understand biological mechanisms of pellicle formation.
This work represents the initial steps in cha racterizing
the process in S. oneidensis. We showed th at successful
pellicle formation required the availability of oxygen and
the presence of ce rtain metal cations. A further analysis
on metal cations revealed that Fe(II) and Fe(III) were
not as essential as Ca(II), Cu(II), Mn(II), and Zn(II) for
pellicle formation. In addition, results presented demon-
strated that a type I secretion pathway of S. oneidensis is
required for the pellicle development but not for attach-
ment to abiotic surface.
Results
Characteristics of S. oneidensis growth in still media under
aerobic conditions
The S. oneidensis MR-1 cells grew rapidly in LB i n a
flask when aeration of the media wa s provided by vigor-
ously shaking, with a doubling time of approximately 40
min at the room temp erature (Figure 1A). Such growth
eventually led to formation of the solid surface-asso-
ciated (SSA) biofilms on the flask wall, especially around
the A-L interface. Cells in static media accessible to
ambient air, however, displayed a different growth pat-
tern. Before pel licles were formed, cell s lived in the
planktonic form with a much longer doubling time,
approximately 2.6 h (Figure 1A). Once pellicle formation
initiated, some of the planktonic cells started to form
pellicles wh ile the rest remained in the planktonic form.
During the development of pellicles, the planktonic cells
grew at a much lower rate with a doubling time of
approximately 6 h (Figure 1A). In this study, initiation
of pellicle formation was determined by the time point
where the growth rate of the planktonic cells changed
although pellicles visible to naked eyes appeared much
later, ab out 12 hours after inoculation at the room
temperature. Both complex and defined media sup-
ported pellicle formation of S. oneidensis. Howev er, pel-
licles from LB were thick and fairly uniform compared
to thin and porous ones from the defined medium, indi-
cating an impact of nutrition on pellicle formation
(Figure1B).WethereforechoseLBthroughtherestof
this study unless otherwise noted.
Oxygen is required for pellicle formation in S. oneidensis
As demon strated above, S. oneidensis initiated the pelli-
cle formation process under aerobic conditions. We
then asked whether oxygen is an essential fac tor for pel-
licle formatio n of this microorganism. The pellicle for-
mation assay was carried out under anaerobic
conditions with lactate as the elect ron donor and one of
following agents as the electron acceptors: fumarate
(20 mM), nitrate (5 mM), DMSO (20 mM), TMAO (20
mM), or ferrous citrate (10 mM). In a ll cases, the capa-
city of S. oneidensis cells to form pellicles was abolished
(data not shown), indicating that oxygen is required for
the process. This is in agreement with the findings that
the lack of oxygen also resulted in a defect in SSA bio-
film formation and a sudden decrease in oxygen concen-
tration led to rapid detachment of SSA biofilms [25,27].
To further elucidate the role of oxygen i n pellicle for-
mation, dissolved oxygen concentrations (DOC) at four
different depths be low the s urface in the unshaken cul-
tures were measured in a time-course manner. Results
revealed that DOC at 0.5, 1, and 2 cm below the surface
in the unshake n culture s displayed a s imilar declining
pattern with time, decreased rapidly from approximately
8to0.04mg/Lduringthefirsttwoandhalfhours,and
then remained stable at 0.04 mg/L (Figure 1C). However,
DOC at the depth immediately below the surface (0.1 cm
but the detector immersed in the liquid) reduced in a
much slow er rate and reached the lowest level of 0.04
mg/L only after the pellicle formed. These data indicate
that the majority of dissolved oxygen is like ly consumed
by the cells close to the surface and the cells below the
surface were grown under microaerobic/anaerobic condi-
tions even before the pellicle was formed.
Proteins are essential in pellicle formation of S. oneidensis
Since EPS, i nclud ing proteins, polysaccharides, extracel-
lular DNA, humic acid, and sugar, are important in SSA
biofilm and pellicle formation of various bacteria, we
speculated that these biopolymers may play a role in
pellicle formation of S. oneidensis. To this end, effects of
proteinase K and DNase I on pellicle formation and
developed pellicles were ass essed. The pellicles were
prevented from formation in the presence of 100 μg/ml
proteinase K (Figure 2A). Consistently, 100 μg/ml of the
proteinase K was able to degrade the developed pellicles
in 24 h, resulting in the semi-tr ansparent membrane-
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 2 of 11
Page 2
like complexes (Figure 2A). In the control experimen t,
proteinase K at concentrations up to 300 μg/ml did not
show a noticeable inhibitory influence on growth of
S. oneidensis under agitated conditions. On the contrary,
DNase I (up to 1000 U/ml) was not effective to inhibit
pellicle formation or to degrade of the developed
pellicles (data not shown), suggesting that DNA plays a
negligible role in the process. Since proteinase K unspe-
cifically removes polypeptides in the extracellular space
and in the outer-membrane exposed to environments,
the results could not conclude whether specific extracel-
lular proteins are required for the process.
Figure 1 Pellicle formation of S. oneidensis in LB under aerobic conditions. (A) Growth of S. oneidensis in static liquid LB under aerobic
conditions. Cell density of all cells (planktonic and pellicle cells combined) (brown square), pellicle cells (yellow triangle), planktonic cells (blue
circle), and the ΔflgA mutant (green cross) was shown. Growth of agitated cultures (black diamond) is included for comparison. Presented are
averages of four replicates with the standard deviation indicated by error bars. (B) Pellicle formation of MR-1 in static liquid LB under aerobic
conditions. The pellicles started to form about 12 h after inoculation based on the altered growth rate of planktonic cells at the room
temperature. (C) Dissolved oxygen concentrations at 1 cm below the surface in the static MR-1 cultures.
Figure 2 EPS analysis. (A) Effects of proteinase K on pellicle formation and developed pellicles. Upper-panel, pellicle formation of the WT in
static LB, in which the proteinase K was added at inoculation to 100 mg/ml (final concentration). Lower panel, developed pellicles of the WT
(48 h after inoculation) were treated with 100 mg/ml (final concentration). (B) TLC analysis of monosaccharide in pellicles and supernatants.
P and S represent pellicle and supernatant, respectively. Man, gal, and glu represent mannose, galactose, and glucose, respectively. Supernatants
of the aggA mutant culture were included in the analysis.
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 3 of 11
Page 3
Attempts were made to solve the major polysaccharide
components of S. oneidensis pellicles by the thin layer
chromatography (TLC) analysis. Culture supernatants
and pellicles were collected independently after 36 h of
growth and pellicles were then treated with 100 μg/ml
proteinase K to removed cells. Polysaccharides were
extracted and subjected to TLC analysis as described in
Methods. A preliminary experiment was performed with
six monosaccharides as standards, including ribose,
mannose, glucose, galactose, rhamnose, and N-acetyl-
glucosamine. The monosaccharides visualized on the
TLC plates were close to mannose, glucose, and galac-
tose (data not shown). To further confirm the observa-
tion, the experiment was conducted again with these
three monosaccharide standards only. As shown in Fig-
ure 2B th e major monosaccha rides identified were most
likely to be mannose in both supernatants and pellicles.
To va lidate this res ult, the aggA mutant, a pellicle-less
strain was included in the analysis and the same result
was obtaine d. These data suggest that the mannose-rich
polysaccharides identified in pellicles are not pellicle
specific.
Certain metal cations are required for pellicle formation
in S. oneidensis
On the b asis that metal cations are of general impo r-
tance in biofilm formation, we examined the effects of
certain metal cations on pellicle formation of S. oneiden-
sis. The metal chelator ethylenediaminetetraacetate
(EDT A) has been shown to have an activity against bio-
films of various bacteria by removing metal cations
[28,29]. As shown in Figure 3A, 0.3 mM EDTA comple-
tely blocked pellicle formation of S. oneiden sis.Asevere
inhibitory effect was also observed in the presence of 0.1
and 0.2 mM of EDTA, reducing the pellicles to approxi-
mately 50 and 70% (by OD
600
readings), respectively
(Figure 3 B). In addition, the pellicle development was
much slower than the non-EDTA control. To rule out
that the observation was due to toxicity of EDTA to S.
oneidensis, the same experiment was conducted again
under agitated conditions. No noticeable difference in
growth between samples containing 0.3 m M EDTA and
the non-EDTA control. All these results indicate that
EDTA at the tested concentration has a detrimental
effect on pellicle formation of S. oneidensis.
We reasoned that the inhibitory effect of EDTA on
pellicle formation of S. oneidensis was due to the
absence of free metal cations in the c ultures. Theref ore,
the role of a specific cation in the process can be
assessed by the addition of this cation to the cultures
containing EDTA. Given that 0.3 mM EDTA appears to
be close to the minimal EDTA concentration for com-
plete inhibition of pellicle formation, we chose the con-
centration for this analysis to determine the importance
of a variety of metal cations in pellicle formation. An
array of metal cations with different stability constants
[log(K
c
)] were tested, including Cu(II) [K
c
= 5.77], Mg
(II) [K
c
= 8.83], Ca(II) [K
c
= 10. 61], Mn(II) [K
c
= 15. 6],
Zn(II) [K
c
= 17.5], Fe(II) [K
c
= 25.0], and Fe(III) [K
c
=
27.2]. To saturate 0.3 mM EDTA, the concentration for
each metal cation used was 0.3 mM as well.
The addition of Ca(II), Mn(II), Cu(II), or Zn(II) fully
rescued the initiation of pellicle formation at the cell
density threshold and subsequent development (Figure
3A (only Ca(II) was shown), 3C). On the contrary, the
inhibitory effect of EDTA w as noticeably lessened but
not fully removed when Mg(II) was added ( Figure 3A).
In the case of Fe(II) and Fe(III), the addition of either
agent partially rescued ( ~40%) the pellicle formation
defect caused by EDTA (Figure 3A). In addition, unlike
pellicles formed in the non-EDTA control or in the pre-
sence of Ca (II), Mn(II), Cu(II), or Zn(II), the Fe-enabled
pellicles were weakly attached to the container wall and
fragile. As a result, the pellicles can be detached from
the wall and broken into pieces with a slight shake. The
same results were observed with even higher levels of Fe
(II) or Fe(III) (up to 0.9 mM). In solution, the addition
of an extra amount o f certain metal cation may release
other cations with lower stability constants from EDTA.
However, this is unlikely to be the underlying reason for
the observed results because the inhibitory effects of
these tested cations on pellicle formation are not corre-
lated to the stability constants of the tested metal
cations.
Progression of pellicle formation was delayed but not
prevented in flagella-less S. oneidensis
Flagella-less and paralyzed flagellar mutants of many
motile bacteria are defective in SSA b iofilm and pellicle
formation because initial surface attachment depends on
flagella-mediat ed motility [30,31]. Ho wever, reports that
biofilm and pellicle formation is not affected or even
promoted by mutation resulting in impaired flagella in
some othe r bacteria are not scarce [1,32,33]. To assess
theroleofflagellainpellicleformationofS. oneidensis,
we tested a flagellum-less strain derived from MR-1 in
which flgA(so3253) was knocked out. FlgA is a molecu-
lar chaperone required for P ring assembly in the peri-
plasmic space [34]. The mutant was unable to swarm or
swim, indicating that the mutati on resulted in function-
ally impaired flagella (Figure 4A). In addition, the fla-
gella were not found on the mutant under an electron
microscope (Figure 4A). To confirm this observation,
the intact flgA was cloned into plasmid pBBRMCS-5 for
complementation. The ability of the mutant to swarm
and swim was restored by the corresponding DNA frag-
ment, indicating that the nonmotile phenotype was due
to mutation in the gene (Figure 4A).
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 4 of 11
Page 4
Compared to MR-1, mutation in flgA failed to elicit
any significant difference in growth under agitated con-
ditions and SSA biofilm formation (data not shown).
However, the mutant displayed a growth defect in the
still media and the pellicle formation was drastically
delayed. As presented in (Fig ure 4B), mutation in flgA
resulted in slow growth with a doubling t ime of ~7 h,
approximately 3 times longer than that of the wild type
before pellicles were formed (Figure 1A). Once pellicle
formation i nitiated, that did not occur until 30 h after
inoculation, the mutant grew at the rate comparable to
the wild type. Interestingly, the development of pellicles
in mutants appeared to b e normal. As a result, the
mutants manag ed to catch up the wild-type in pellicle
production (10 days) (Figure 4B). All of these results
suggest that the delayed initiation o f pellicle formation
of the flgA mutant was po ssibly due to the slow growth
of the mutant cells in the unshaken media and flagella
were unlikely to play a significant role in the attachment
of S. oneidensis cells to the wall or pellicle maturation.
AggA type I secretion pathway is essential in pellicle
formation of S. oneidensis
Previously, a type I secretion system (TISS) consisting of
an ATP-binding protein in the inner membrane RtxB
(SO4318), an HlyD-family membrane-fusion protein
SO4319, and an agglutination protein AggA (SO4320)
was suggested to be important in SSA biofilm formation
of S. oneidensis [21,22,35]. A following mutational analy-
sis revealed that AggA was critical to hyper-aggregatio n
of th e COAG strain, a spontane ous mutant from MR-1
[22]. In the case of SSA biofilm formation, the impact of
mutation in aggA was rather mild, reducing the robust
biofilm-forming capacity of the COAG strain to t he
level of the wild-type.
Given the importance of AggA in biofilm formation
sugge sted by abov e-mentioned studies, it is necessary to
assess its role in biofilm formation of S. oneidensi s with
a wild-type genetic background. To this end, we c on-
structed an aggA in-frame deletion mutant with MR-1
as the parental strain. The physiological characterization
revealed that the mutant grew at the rate comparabl e to
that of the parental strain either in the shaking or static
conditions. However, the aggA mutant was unable to
formed pellicles in 5 days (Figure 5A). Introduction of
aggA on plasmid pBBR-AG GA into the mutant restored
its ability to form pellicles, verifying that the phenotype
of the aggA mutant was specific to the mutation in the
aggA gene (Figure 5A). As a result, the aggA strain dis-
played a growth pattern different from the wild type
strain in the static media by the lack of the growth rate
change which signaled the initiation of pellicle
Figure 3 Treatment of S. oneidensis pellicles with EDTA and divalent cations. (A) Pellicle formation of the WT after 48 h in static LB in the
presence of 0.3 mM EDTA and certain divalent cation (0.3 mM) under aerobic conditions. (B) Cells in pellicles formed in the presence of 0 (light
blue), 0.1 (dark red), 0.2 (light yellow), and 0.3 mM (dark blue) EDTA at the different time points. Presented are averages of four replicates with
the standard deviation indicated by error bars. (C) Effects of divalent cations on the inhibition of pellicle formation by EDTA. Pellicle formation of
the WT after 48 h in static LB in the presence of 0.3 mM EDTA and one of indicated divalent cations (0.3 mM) under aerobic conditions was
shown. The WT in static LB without EDTA was used as the control. The relative pellicle formation ((EDTA and indicated cation)/EDTA-absence
control) was presented in the figure. EDTA only (No cation was used as the negative control. Presented are averages of four replicates with the
standard deviation indicated by error bars.
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 5 of 11
Page 5
formation (Figure 1A). However, the mutant was able to
attach to the glass wall at the air-liquid i nterface, sug-
gesting that AggA is not essential for this step of biofilm
formation (Figure 5A). This proposal gained support
from the SSA biofilm formation of the mutant, which
differed from that of the wild type strain insignificantly
(Figure 5B). All these data implicate that AggA TISS is
require d for pel licle formation, most likely at t he mono-
layer pellicle formation stage, which appears to be differ-
ent from that in SSA biofilm formation.
Discussion and Conclusions
In the microbial world, existence within surface-asso-
ciated structured multicellular communities is the pre-
vailing lifestyle [36,37]. The pellicles of facultative
bacteria formed at the liquid-air interface can be selec-
tively advantageous given that respiration with oxygen
as the terminal electron acceptor is the most productive.
In S. oneidensis, the growth rate was promoted by better
access to oxygen evidenced by that the c ells grew much
faster in shaking than in static cultures. Along with the
observation that SSA biofilm formation of S. oneidensis
was inhibited under a naerobic conditions, the require-
ment of oxygen for pellicle formation may mainly come
from its facilitation of aggregation and attachment of
cells to the solid surfaces. This is consistent with pre-
vious findings that oxygen promotes autoaggregation of
and sudden depletion of molecular oxygen was shown
to act as the predominant trigger for initiating detach-
ment of individual cells from biofilms [26,38]. We
Figure 4 The ΔflgA mutant displayed slow pellicle format ion. (A) Swimming and swarming motility assays of the ΔflgA mutant. In both
panels, the ΔflgA mutant (Upper) was compared to the WT (Lower). The ΔflgA* strain refers to the ΔflgA mutant containing pBBR-FLGA. (B)
Electron micrographs of WT and the ΔflgA mutant. No flagellum was observed on the mutant. (C) Left panel, pellicle formation of the ΔflgA
mutant. Right panel, the cell densities of cells in pellicles of the WT and the ΔflgA mutant. The WT, dark red; the ΔflgA mutant, light blue. E
represents the time at which the cell density of ΔflgA mutant catches up (10 days after inoculation in the experiment). Presented are averages of
four replicates with the standard deviation indicated by error bars.
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 6 of 11
Page 6
therefore propose that an oxygen gradient established in
static cultures with the highest oxygen concentration at
the surface resulted in a larger number of cells at the A-
L interface to form pellicles, which eventually induce
attachment of individual cells to the abiotic surface.
To form pellicles, S. oneidensis cultures require certain
divalent ions. Involv ement of metals in biofilm forma-
tion either as a facilitator or an inhibitor has been well
documented. In recent years, many elegant studies
about the susceptibility of biofilms to metals (as an inhi-
bitor) have been published [39-41]. Although metals as
a biofilm formation facilitator have been studied for
more than two decades, only a few metals (Ba(II), Mg
(II), Ca(II), Fe(III), and Fe( III)) have been investigated
[34,42,43]. In P. aeruginosa, all these metals but Ba(II)
are able to protect P. aeruginosa biofilms against EDTA
treatment, presumably by stabilizing the biofilm matrix.
In addition, it has been shown that there is a positive
correlation between calcium concentration and amount
of biofilm accumulation [44]. While our data support
previous conclusions that calcium plays an impo rtant
role in stabilizing biofilms of bacteria [34,43,44] , most of
other findings are either new or surprising. Among
tested metal cations , Cu(II), Ca(II ), Mn(II), and Zn(II)
Figure 5 Biofilm assay of MR-1 and aggA mutant. (A) Pellicle formation of MR-1, ΔaggA, ΔaggA* (aggA in-frame deletion mutant containing
pBBR-AGGA). (B) SSA Biofilm was assessed for the strains indicated after 16 and 24 h, respectively. Cultures were prepared as described in
Methods. The averaged OD readings of four independent culture tubes were given with images of representative CV-stained tubes.
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 7 of 11
Page 7
belong to the same class, which are capable of restoring
the a bility of S. oneidensis to form pellicles in the pre-
sence of EDTA completely. In contrast, Mg(II) shows
mild effects on relieving EDTA inhibition whereas Fe(II)
and Fe(III) counteracted EDTA in a way different from
other tested cations evidenced by the fragile pellicles. In
combination, these data suggest that the relative stability
constants of metal cations (Cu(II) [5.77], Mg(II) [8.83],
Ca(II) [10.61], Mn(II) [15.6], Zn(II) [17.5], Fe(II) [25.0],
and Fe(III) [27.2]) and their af fect on EDTA inhibition
are not correlated.
It is particularly worth discussing roles of Fe(II) and Fe
(III) in pellicle formation of S. oneidensis. In recent years,
many reports have demonstrated that the iron cations
are important, if not essential, in bacterial biofilm forma-
tion [34,45-47]. In P. aeruginosa, influence of Fe( II) and
Fe(III) on the process was equivalent to that of Ca(II)
[34]. In S. oneidensis, irons in forms of Fe(II) and Fe( III)
were not only unable to neutralize the inhibitory effect of
EDTA on pellicle formation completely but also resulted
in structurally impaired pellicles although these agents
indeed play a role in pellicle formation. Thi s observation
indicates that irons are not so crucial as Cu(II), Ca(II),
Mn(II), and Zn(II) in pellicle formation of S. oneidensis.
In fact, this may not be su rprisi ng. In Acinetobacter bau-
mannii and Staphylococcus aureus, iron limitation
improved bio film formation [48,49]. Therefor e, it is pos-
sible that different bacteria respond to irons in a different
way with respect to biofilm formation.
Like SSA biofilms, pellicles require EPS to form a
matrix to support embedded cells. Although EPS are
now w idely recognized as the essential compone nts for
biofilm formation and development in all biofilm-form-
ing microorganisms studied so far, diversity in their
individual composition and relative abundance of certain
elements is substantial [50]. For example, extracellular
nucleic acids, which are not important in most biofilm-
forming microorganisms, are required for SSA biofi lm
formation in a variety o f bacteria [11,36,37,51,52]. In S.
oneidensis, proteins not extracellular DNA s are required
to pellicle formation. While essential extracellular pro-
teins for S. oneiden sis pellicle formation are largely
unknown, results from this study demonst rated that the
AggA TISS is crucial in the process, likely at the devel-
opment of the monolay er. One of substrates of this
transporter is predicted to be SO4317, a large putative
RTX toxin [35], implicating that the protein m ay be
involved in pellicle formation. In the case of polysac-
charides, mannose dominates not only in pellicles but
also in supernatants, implicating that mannose-based
polysaccharides may have a more general role in the
bacterial physiology.
Like in B. subtilis, mutations in S. oneidensis flagellar
genes resulting i n the nonmotile phenotype significantly
delayed the initiation and development of pellicle forma-
tion [17]. Here we further illustrated that neither SSA
biofilm formation nor the maturization of pellicle was
impaired by the mutations. In agreement with findings
on biofilm formation of Bacillus c ereus [13], this obser-
vation suggests that motility not only promotes cells to
move to surfaces where the pellicle forms but also facili-
tate planktonic cells entrance into the pellicle.
Overall, the results presented here provided the first
insights into pellicle formation of S. oneidensis,making
pellicle formation of S. oneidensis a simple research
mode l for biofilm formation in general. The study high-
lights p arallels and significant differences between this
process and well-documented paradigms, raising some
key questions demanding i mmediate investigations.
These include what the major polysaccharides in S. onei-
densis pellicles are, why irons result in fragile pellicles in
the presence of EDTA, and w hich proteins a nd their
secretion pathway(s) a re directly related to pellicle
formation.
Methods
Bacterial strains, plasmids, and culture conditions
Bacterial strains and plasmids used in this study are
listed in Table 1 [53]. Escherichia coli and S. oneidensis
strains were routinely grown in LB broth or on LB
plates at 37°C and the room temperature for gene tic
manipulation, respectively. When needed, antibiotics
were used at the following concentrations: ampicillin at
50 μg/ml and gentamycin at 15 μg/ml.
Pellicle formation, measurement of growth, and
quantification of pellicles
A fresh colony grown overnight on a LB plate was used
to inoculate 5 0 ml LB and incubated in a shaker (200
rpm) to an OD
600
of 0.8 at the room temperature. This
culture was then diluted 500-fold with fresh LB, result-
ing in the starting cultures. Throughout the study, all
starting cultures of S. oneidensis strains were prepared
this way. Aliquots of 30 ml starting cultures were trans-
ferred to 50 ml Pyrex beakers. The beakers were kept
still for pellicle formation at the room temperature and
dissolved oxygen (DO) of the cultures was recorded
every hour with an Accumet X L40 meter (Fisher Scien-
tific). M1 defined medium containing 0.02% (w/v) of
vitamin-free Casamino Acids and 15 mM lac tate with
one of electron acceptors including fumarate (20 mM),
nitrate (5 mM), trimethylamine N-ox ide (TMAO) (20
mM), dimethyl sulfox ide (DMSO) (20 mM) and ferrous
citrate (10 mM), was used to test pellicle formation in
the defined medium [54]. To separate cells in pellicle
and underneath, cultures were withdrawn care fully for
collecting planktonic cells and the left pellicles. For
growth measurement, 27 parallel starting cultures were
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 8 of 11
Page 8
used and 3 were collected at each time point and the
rest remained undisturbed. The cell density (OD
600
)of
cultures containing planktonic cells was measured first
as the planktonic cell density and measured again as the
overall cell density after cells from pellicles were added
and extensively vortexed. To quantify the pellicles
formed by the S. oneidensis wild-type and mutant
strains, cells from pellicles were coll ected, suspended in
30 ml fresh LB, violently vortexed, and applied to the
spectrometer at 600 nm.
Proteinase K and DNase I treatment of S. oneidensis
pellicles
S. oneidensis was statically cultured in LB broth with the
addition of proteinase K (0 μg/mL, 100 μg/mL, and 500
μg/mL) or DNase I (Qiagen, 0U/mL, 100U/mL, 500U/
mL and 1000U/mL) for 3 days [55]. We also investi-
gated whether these 3 enzym es could dissolve estab-
lished pellicles. 2-day old pellicles were rinsed with 20
mM Tris-HCl (pH = 8.0) and incubated in the same
buffer supplemented with proteinase K at 37°C for 2
days. Similarly, 2-day old pellicles were incubated with
DNase I to examine the DNA content at room tempera-
ture for 2 days.
Mutagenesis, physiological characterization and
complementation of the resulting mutants
Deletion mutation strains were constructed using the
fusion PCR method illustrated previously [56]. Primers
used for mutagenesis were listed in Additional fil e 1. In
brief, two DNA fragments flanking the target gene were
generated from S. oneidensis genomic DNA by PCR
with primers 5F/5R and 3F/3R, respectively. Fusion PCR
was then performed to join these two DNA fragments
with primers 5F/3R. The resulting single fragment was
digested with SacI and ligated into the SacI-digested
and phosphatase-treated suicide vector pDS3.0. The
resultant vectors were electroporated into the donor
strain, E. coli WM3064 and then moved to S. oneidensis
by conjugation. Integration of the mutagenesis construct
into the chromosome and resolution were performed to
generate the final deletion strains. The deletion was ver-
ified by PCR and DNA sequencing.
For complementation, DNA fragments containing
aggA or flgA were generated by PCR amplification with
MR-1 genomic DNA as the template using primers
SO4320- COM-F/SO3988-COM-R and SO3253-COM-F/
SO32 53-COM-R, respectively as listed in A dditional file
1. These fragments were digested with SacI and ligated
to SacI-digested pBBR1MCS-5 to form pBBR-AGGA
and pBBR-FLGA, which was electroporated into
WM3064. Introduction of pBBR-AGGA or pBBR-FLGA
into the corresponding mutant was done by conjugation,
and gentamycin-resistant colonies were selected. The
presence of pBBR-AGGA or pBBR-FLGA i n t he corre-
sponding mutant was confirmed by plasmid purification
and restriction enzyme digestion.
Swarm and swimming motility assay
A fresh colony of tested strains w as grown to an OD
600
of 0.8 i n L B media. The cultures (1 ml) w ere spotted
onto a swarm LB plate (0.5% agar) or stabbed into a
swimming LB plate (0.2% agar). All plates were incu-
bated at the room temperature for 48 h. Images were
acquired using Alpha Innotech s Fluorchem imaging
system.
SSA biofilm assay
The SSA biofilm formation assay used is b ased on the
method previously reported [57]. In brief, 3 ml of f resh
LB in 15 ml glass tubes were inoculated with S. oneiden-
sis strains from an overnight culture in LB at 200 rpm.
Table 1 Strains and plasmids used in this study
Strain or plasmid Relevant genotype Reference or source
E. coli
WM3064 Donor strain for conjugation; ΔdapA [53]
S. oneidensis
MR-1 Wild-type ATCC 700550
JZ3253 flgA deletion mutant derived from MR-1; Δ flgA This study
JZ4320 aggA deletion mutant derived from MR-1; ΔaggA This study
Plasmid
pDS3.0 Ap
r
,Gm
r
, derivative from suicide vector pCVD442 Lab stock
pBBR1MCS-5 Gm
r
vector used for complementation Lab Stock
pDS-AGGA aggA deletion construct in pDS3.0 This study
pDS-FLGA flgA deletion construct in pDS3.0 This study
pBBR-AGGA pBBR1MCS-5 containing aggA of S. oneidensis This study
pBBR-FLGA pBBR1MCS-5 containing flgA of S. oneidensis This study
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 9 of 11
Page 9
After 16, 24, 32, or 40 h of incubation at 200 rpm at
room temperature, 500 μl of 1% (wt/vol) crystal violet
(CV) solution was added to each tube and incubated for
15 min. Tubes were rinsed three times with 5 ml of dis-
tilled H
2
O and air dried. Biofilm formation was quanti-
fied by measuring the absorbance at 575 nm. Each assay
was performed four times.
Thin layer chromatography (TLC) analysis
Supernatants and pellicles were collected after 36 h of
growth in st atic LB media. Pellicles were treated with
100 μg/mL proteinase K for removal of cells. Cell-less
pellicles and supernatants were subjected to exopolysac-
charide extraction and hydrolysis with trifluoroacetic
acid as described previously [58]. The resulting mono-
saccharides were dissolved in ddH
2
O in the co ncentra -
tion of 10 mg/ml, and 2 μl of the sample was spotted
onto TLC plates (silica gel 60 F
254
; Merck). After devel-
opment in butan-1-ol-acetone-water (4:5:1), the TLC
plates were dipped in the reagent aniline-diphenylamine
in acetone and incubated for 2 to 5 min at 100°C.
Additional material
Additional file 1: Primers used in this study. File contains all primers
used in this study
Acknowledgements
This research was supported by Major State Basic Research Development
Program (973 Program: 2010CB833803) and National Natural Science
Foundation of China (30870032) to HG. This research was also supported by
Chinese Science Foundation for Distinguished Group (No.50321402) to YL.
This research was also supported by The U.S. Department of Energy under
the Genomics: GTL Program through Shewanella Federation, Office of
Biological and Environmental Research, Office of Science.
Author details
1
School of Minerals processing and Bioengineering, Central south University,
Changsha, 410083, PR China.
2
Institute for Environmental Genomics and
Department of Botany and Microbiology, University of Oklahoma, Norman,
73019, USA.
3
Institute of Microbiology and College of Life Sciences, Zhejiang
University, Hangzhou, Zhejiang 310058, PR China.
Authors contributions
YL carried out pellicle formation and characterization experiments and
drafted the manuscript. HG conceived of the study, and participated in its
design, and directed all experiments and coordination and drafted the
manuscript. JC carried out the mutagenesis experiments. YD and LW carried
out SSA biofilm and TLC assays. ZH participated in design of the study and
helped to draft the manuscript. XL and GQ participated in the design of the
study and helped to draft the manuscript. JZ conceived of the study, and
participated in its design and coordination and helped to draft the
manuscript. All authors read and approved the final manuscript.
Received: 29 May 2010 Accepted: 16 November 2010
Published: 16 November 2010
References
1. OToole G, Kaplan HB, Kolter R: Biofilm formation as microbial
development. Ann Rev Microbiol 2000, 54:49-79.
2. Watnick P, Kolter R: Biofilm, city of microbes. J Bacteriol 2000,
182:2675-2679.
3. Stoodley P, Sauer K, Davies DG, Costerton JW: Biofilms as complex
differentiated communities. Ann Rev Microbiol 2002, 56:187-209.
4. Kolter R, Greenberg EP: Microbial sciences-The superficial life of microbes.
Nature 2006, 441:300-302.
5. Goller CC, Romeo T: Environmental Influences on Biofilm Development.
In Bacterial Biofilms 2008, 37-66.
6. Spormann AM: Physiology of microbes in biofilms. In Bacterial Biofilms
2008, 17-36.
7. Karatan E, Watnick P: Signals, Regulatory Networks, and Materials That
Build and Break Bacterial Biofilms. Microbiol Mol Biol Rev 2009, 73:310-347.
8. Liu M, Alice AF, Naka H, Crosa JH: HlyU protein is a positive regulator of
rtxA1, a gene responsible for cytotoxicity and virulence in the human
pathogen Vibrio vulnificus. Infect Immun 2007, 75:3282-3289.
9. Rainey PB, Travisano M: Adaptive radiation in a heterogeneous
environment. Nature 1998, 394:69-72.
10. Ude S, Arnold DL, Moon CD, Timms-Wilson T, Spiers AJ: Biofilm formation
and cellulose expression among diverse environmental Pseudomonas
isolates. Environ Microbiol 2006, 8:1997-2011.
11. Lemon KP, Earl AM, Vlamakis HC, Aguilar C, Kolter R: Biofilm development
with an emphasis on Bacillus subtilis. In Bacterial Biofilms 2008, 1-16.
12. Enos-Berlage JL, Guvener ZT, Keenan CE, McCarter LL: Genetic
determinants of biofilm development of opaque and translucent Vibrio
parahaemolyticus. Mol Microbiol 2005, 55:1160-1182.
13. Joshua GWP, Guthrie-Irons C, Karlyshev AV, Wren BW: Biofilm formation in
Campylobacter jejuni. Microbiology 2006, 152:387-396.
14. Houry A, Briandet R, Aymerich S, Gohar M: Involvement of motility and
flagella in Bacillus cereus biofilm formation. Microbiology 2010,
156
:1009-1018.
15. Deighton M, Borland R: Regulation of slime production in Staphylococcus
epidermidis by iron limitation. Infect Immun 1993, 61:4473-4479.
16. Moelling C, Oberschlacke R, Ward P, Karijolich J, Borisova K, Bjelos N,
Bergeron B: Metal-dependent repression of siderophore and biofilm
formation in Actinomyces naeslundii. FEMS Microbiol Lett 2007,
275:214-220.
17. Kobayashi K: Bacillus subtilis pellicle formation proceeds through
genetically defined morphological changes. J Bacteriol 2007,
189:4920-4931.
18. Solano C, Garcia B, Valle J, Berasain C, Ghigo JM, Gamazo C, Lasa I: Genetic
analysis of Salmonella enteritidis biofilm formation: critical role of
cellulose. Mol Microbiol 2002, 43:793-808.
19. Spiers AJ, Bohannon J, Gehrig SM, Rainey PB: Biofilm formation at the
air-liquid interface by the Pseudomonas fluorescens SBW25 wrinkly
spreader requires an acetylated form of cellulose. Mol Microbiol 2003,
50:15-27.
20. Bagge D, Hjelm M, Johansen C, Huber I, Grami L: Shewanella putrefaciens
adhesion and biofilm formation on food processing surfaces. Appl
Environ Microbiol 2001, 67:2319-2325.
21. De Vriendt K, Theunissen S, Carpentier W, De Smet L, Devreese B, Van
Beeumen J: Proteomics of Shewanella oneidensis MR-1 biofilm reveals
differentially expressed proteins, including AggA and RibB. Proteomics
2005, 5:1308-1316.
22. De Windt W, Gao H, Kromer W, Van Damme P, Dick J, Mast J, Boon N,
Zhou J, Verstraete W: AggA is required for aggregation and increased
biofilm formation of a hyper-aggregating mutant of Shewanella
oneidensis MR-1. Microbiology 2006, 152:721-729.
23. Teal TK, Lies DP, Wold BJ, Newman DK: Spatiometabolic stratification of
Shewanella oneidensis biofilms. Appl Environ Microbiol 2006, 72:7324-7330.
24. Thormann KM, Saville RM, Shukla S, Pelletier DA, Spormann AM: Initial
phases of biofilm formation in Shewanella oneidensis MR-1. J Bacteriol
2004, 186:8096-8104.
25. Thormann KM, Saville RM, Shukla S, Spormann AM: Induction of rapid
detachment in Shewanella oneidensis MR-1 biofilms. J Bacteriol 2005,
187:1014-1021.
26. Thormann KM, Duttler S, Saville RM, Hyodo M, Shukla S, Hayakawa Y,
Spormann AM: Control of formation and cellular detachment from
Shewanella oneidensis MR-1 biofilms by cyclic di-GMP. J Bacteriol 2006,
188:2681-2691.
27. Walters MC, Roe F, Bugnicourt A, Franklin MJ, Stewart PS: Contributions of
Antibiotic penetration, oxygen limitation, and low metabolic activity to
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 10 of 11
Page 10
tolerance of Pseudomonas aeruginosa biofilms to ciprofloxacin and
tobramycin. Antimicrob Agents Chemother 2003, 47:317-323.
28. Kite P, Eastwood K, Sugden S, Percival SL: Use of In Vivo-generated
biofilms from hemodialysis catheters to test the efficacy of a novel
antimicrobial catheter lock for biofilm eradication In Vitro. J Clin Microbio
2004, 42:3073-3076.
29. Banin E, Brady KM, Greenberg EP: Chelator-induced dispersal and killing
of Pseudomonas aeruginosa cells in a biofilm. Appl Environ Microbiol 2006,
72:2064-2069.
30. Pratt LA, Kolter R: Genetic analysis of Escherichia coli biofilm formation:
roles of flagella, motility, chemotaxis and type I pili. Mol Microbiol 1998,
30:285-293.
31. Lemon KP, Higgins DE, Kolter R: Flagellar motility is critical for Listeria
monocytogenes biofilm formation. J Bacteriol 2007, 189:4418-4424.
32. Merritt PM, Danhorn T, Fuqua C: Motility and chemotaxis in
Agrobacterium tumefaciens surface attachment and biofilm formation. J
Bacteriol 2007, 189:8005-8014.
33. Parsek MR, Tolker-Nielsen T: Pattern formation in Pseudomonas aeruginosa
biofilms. Curr Opin Microbiol 2008, 11:560-566.
34. Nambu T, Kutsukake K: The Salmonella FlgA protein, a putative
periplasmic chaperone essential for flagellar P ring formation.
Microbiology 2000, 146:1171-1178.
35. Theunissen S, Vergauwen B, De Smet L, Van Beeumen J, Van Gelder P,
Savvides SN: The agglutination protein AggA from Shewanella oneidensis
MR-1 is a TolC-like protein and forms active channels in vitro. Biochem
Biophys Res Commun 2009, 386:380-385.
36. Whitchurch CB, Tolker-Nielsen T, Ragas PC, Mattick JS: Extracellular DNA
required for bacterial biofilm formation. Science 2002, 295:1487-1487.
37. Branda SS, Vik A, Friedman L, Kolter R: Biofilms: the matrix revisited. Trends
Microbiol 2005, 13:20-26.
38. McLean JS, Pinchuk GE, Geydebrekht OV, Bilskis CL, Zakrajsek BA, Hill EA,
Saffarini DA, Romine MF, Gorby YA, Fredrickson JK, Beliaev AS: Oxygen-
dependent autoaggregation in
Shewanella oneidensis MR-1. Environ
Microbiol 2008, 10:1861-1876.
39. Teitzel GM, Parsek MR: Heavy metal resistance of biofilm and planktonic
Pseudomonas aeruginosa. Appl Environ Microbiol 2003, 69:2313-2320.
40. Priester JH, Olson SG, Webb SM, Neu MP, Hersman LE, Holden PA:
Enhanced exopolymer production and chromium stabilization in
Pseudomonas putida unsaturated biofilms. Appl Environ Microbiol 2006,
72:1988-1996.
41. Harrison JJ, Ceri H, Turner RJ: Multimetal resistance and tolerance in
microbial biofilms. Nature Rev Microbiol 2007, 5:928-938.
42. Turakhia MH, Characklis WG: Activity of Pseudomonas aeruginosa in
biofilms-effect of calcium. Biotechnol Bioeng 1989, 33:406-414.
43. Huang J, Pinder KL: Effects of calcium on development of anaerobic
acidogenic biofilms. Biotechnol Bioeng 1995, 45:212-218.
44. Kierek K, Watnick PI: The Vibrio cholerae O139O-antigen polysaccharide is
essential for Ca2+-dependent biofilm development in sea water. Proc
Natl Acad Sci USA 2003, 100:14357-14362.
45. Singh PK, Parsek MR, Greenberg EP, Welsh MJ: A component of innate
immunity prevents bacterial biofilm development. Nature 2002,
417:552-555.
46. Chen X, Stewart PS: Role of electrostatic interactions in cohesion of
bacterial biofilms. Appl Microbiol Biotechnol 2002, 59:718-720.
47. Berlutti F, Morea C, Battistoni A, Sarli S, Cipriani P, Superti F,
Ammendolia MG, Valenti P: Iron availability influences aggregation,
biofilm, adhesion and invasion of Pseudomonas aeruginosa and
Burkholderia cenocepacia. Inter Journal Immunopath Ph 2005, 18:661-670.
48. Tomaras AP, Dorsey CW, Edelmann RE, Actis LA: Attachment to and
biofilm formation on abiotic surfaces by Acinetobacter baumannii:
involvement of a novel chaperone-usher pili assembly system.
Microbiology 2003, 149:3473-3484.
49. Johnson M, Cockayne A, Williams PH, Morrissey JA: Iron-responsive
regulation of biofilm formation in Staphylococcus aureus involves fur-
dependent and fur-independent mechanisms. J Bacteriol 2005,
187:8211-8215.
50. Tart AH, Wozniak DJ: Shifting paradigms in Pseudomonas aeruginosa
biofilm research. In Bacterial Biofilms 2008, 193-206.
51. Spoering AL, Gilmore MS: Quorum sensing and DNA release in bacterial
biofilms. Curr Opin Microbiol 2006, 9:133-137.
52. Lappann M, Claus H, Van Alen T, Harmsen M, Elias J, Molin S, Vogel U: A
dual role of extracellular DNA during biofilm formation of Neisseria
meningitidis. Mol Microbiol 2010, 75:1355-1371.
53. Saltikov CW, Newman DK: Genetic identification of a respiratory arsenate
reductase. Proc Natl Acad Sci USA 2003, 100:10983-10988.
54. Gao H, Wang XH, Yang ZK, Palzkill T, Zhou JZ: Probing regulon of ArcA in
Shewanella oneidensis MR-I by integrated genomic analyses. BMC
Genomics 2008, 9:42.
55. Yap MN, Rojas CM, Yang CH, Charkowski AO: Harpin mediates cell
aggregation in Erwinia chrysanthemi 3937. J Bacteriol 2006, 188:2280-2284.
56. Gao WM, Liu YQ, Giometti CS, Tollaksen SL, Khare T, Wu LY, Klingeman DM,
Fields MW, Zhou J: Knock-out of SO1377 gene, which encodes the
member of a conserved hypothetical bacterial protein family COG2268,
results in alteration of iron metabolism, increased spontaneous
mutation and hydrogen peroxide sensitivity in Shewanella oneidensis
MR-1. BMC Genomics 2006, 7:76.
57. OToole GA, Kilter R: Flagellar and twitching motility are necessary for
Pseudomonas aeruginosa biofilm development. Mol Microbiol 1998,
30:295-304.
58. Wall P: Thin layer Chromatography: A modern practical approach RSC
publishing; 2005.
doi:10.1186/1471-2180-10-291
Cite this article as: Liang et al.: Pellicle formation in Shewanella
oneidensis. BMC Microbiology 2010 10:291.
Submit your next manuscript to BioMed Central
and take full advantage of:
Convenient online submission
Thorough peer review
No space constraints or color figure charges
Immediate publication on acceptance
Inclusion in PubMed, CAS, Scopus and Google Scholar
Research which is freely available for redistribution
Submit your manuscript at
www.biomedcentral.com/submit
Liang et al . BMC Microbiology 2010, 10:291
http://www.biomedcentral.com/1471-2180/10/291
Page 11 of 11
Page 11
  • Source
    • "In Pseudomonas fluorescens, Mg2+ cation increase cell attachment and condition the structure and further development of the biofilms [46]. Cations such as Ca2+, Mn2+, Cu2+ or Zn2+ have also been found to be essential for the formation of air-liquid interface biofilms in Shewanella oneidensis[47]. In fact, when MH2 is supplemented with 20 mg/L Ca2+ and 10 mg/L Mn2+ (CAMH2 medium), a shift in biofilm production is observed (Figure 2B). "
    [Show abstract] [Hide abstract] ABSTRACT: A variety of conditions (culture media, inocula, incubation temperatures) are employed in antifouling tests with marine bacteria. Shewanella algae was selected as model organism to evaluate the effect of these parameters on: bacterial growth, biofilm formation, the activity of model antifoulants, and the development and nanomechanical properties of biofilms.The main objectives were:1) To highlight and quantify the effect of these conditions on relevant parameters for antifouling studies: biofilm morphology, thickness, roughness, surface coverage, elasticity and adhesion forces.2) To establish and characterise in detail a biofilm model with a relevant marine strain. Both the medium and the temperature significantly influenced the total cell densities and biofilm biomasses in 24-hour cultures. Likewise, the IC50 of three antifouling standards (TBTO, tralopyril and zinc pyrithione) was significantly affected by the medium and the initial cell density. Four media (Marine Broth, MB; 2% NaCl Mueller-Hinton Broth, MH2; Luria Marine Broth, LMB; and Supplemented Artificial Seawater, SASW) were selected to explore their effect on the morphological and nanomechanical properties of 24-h biofilms. Two biofilm growth patterns were observed: a clear trend to vertical development, with varying thickness and surface coverage in MB, LMB and SASW, and a horizontal, relatively thin film in MH2. The Atomic Force Microscopy analysis showed the lowest Young modulii for MB (0.16 +/- 0.10 MPa), followed by SASW (0.19 +/- 0.09 MPa), LMB (0.22 +/- 0.13 MPa) and MH2 (0.34 +/- 0.16 MPa). Adhesion forces followed an inverted trend, being higher in MB (1.33 +/- 0.38 nN) and lower in MH2 (0.73 +/- 0.29 nN). All the parameters significantly affected the ability of S. algae to grow and form biofilms, as well as the activity of antifouling molecules. A detailed study has been carried out in order to establish a biofilm model for further assays. The morphology and nanomechanics of S. algae biofilms were markedly influenced by the nutritional environments in which they were developed. As strategies for biofilm formation inhibition and biofilm detachment are of particular interest in antifouling research, the present findings also highlight the need for a careful selection of the assay conditions.
    Full-text · Article · Apr 2014 · BMC Microbiology
  • Source
    • "It is worth noting that Shewanella species thrive in oxic-anoxic interfaces, where redox conditions change rapidly with frequent shifts in the main electron acceptors11. It has been demonstrated that oxygen can affect the formation and three-dimensional structure of a community formed by Shewanella species, such as pellicles at air-liquid interfaces, aggregates in aerobic chemostat cultures and biofilms in hydrodynamic flow cells121314. Biofilms formed under aerobic conditions show a hollow and seeding dispersal structure, while a round and densely-packed structure under anaerobic conditions15. "
    [Show abstract] [Hide abstract] ABSTRACT: Although oxygen has been reported to regulate biofilm formation by several Shewanella species, the exact regulatory mechanism mostly remains unclear. Here, we identify a direct oxygen-sensing diguanylate cyclase (DosD) and reveal its regulatory role in biofilm formation by Shewanella putrefaciens CN32 under aerobic conditions. In vitro and in vivo analyses revealed that the activity of DosD culminates to synthesis of cyclic diguanylate (c-di-GMP) in the presence of oxygen. DosD regulates the transcription of bpfA operon which encodes seven proteins including a large repetitive adhesin BpfA and its cognate type I secretion system (TISS). Regulation of DosD in aerobic biofilms is heavily dependent on an adhesin BpfA and the TISS. This study offers an insight into the molecular mechanism of oxygen-stimulated biofilm formation by S. putrefaciens CN32.
    Full-text · Article · Jun 2013 · Scientific Reports
  • Source
    [Show abstract] [Hide abstract] ABSTRACT: Background: Although solid surface-associated biofilm development of S. oneidensis has been extensively studied in recent years, pellicles formed at the air-liquid interface are largely overlooked. The goal of this work was to understand basic requirements and mechanism of pellicle formation in S. oneidensis.|Results: We demonstrated that pellicle formation can be completed when oxygen and certain cations were present. Ca(II), Mn(II), Cu(II), and Zn(II) were essential for the process evidenced by fully rescuing pellicle formation of S. oneidensis from the EDTA treatment while Mg (II), Fe(II), and Fe(III) were much less effective. Proteins rather than DNA were crucial in pellicle formation and the major exopolysaccharides may be rich in mannose. Mutational analysis revealed that flagella were not required for pellicle formation but flagellum-less mutants delayed pellicle development substantially, likely due to reduced growth in static media. The analysis also demonstrated that AggA type I secretion system was essential in formation of pellicles but not of solid surface-associated biofilms in S. oneidensis.|Conclusion: This systematic characterization of pellicle formation shed lights on our understanding of biofilm formation in S. oneidensis and indicated that the pellicle may serve as a good research model for studying bacterial communities.
    Full-text · Article · Jan 2010 · BMC Microbiology
Show more