APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Aug. 2011, p. 5342–5351
Copyright © 2011, American Society for Microbiology. All Rights Reserved.
Vol. 77, No. 15
Roles of Two Shewanella oneidensis MR-1 Extracellular Endonucleases?†
Julia Go ¨deke,‡ Magnus Heun,‡ Sebastian Bubendorfer, Kristina Paul, and Kai M. Thormann*
Department of Ecophysiology, Max-Planck-Institut fu ¨r terrestrische Mikrobiologie, Marburg, Germany
Received 22 March 2011/Accepted 16 May 2011
The dissimilatory iron-reducing bacterium Shewanella oneidensis MR-1 is capable of using extracellular DNA
(eDNA) as the sole source of carbon, phosphorus, and nitrogen. In addition, we recently demonstrated that S.
oneidensis MR-1 requires eDNA as a structural component during all stages of biofilm formation. In this study,
we characterize the roles of two Shewanella extracellular endonucleases, ExeS and ExeM. While ExeS is likely
secreted into the medium, ExeM is predicted to remain associated with the cell envelope. Both exeM and exeS
are highly expressed under phosphate-limited conditions. Mutants lacking exeS and/or exeM exhibit decreased
eDNA degradation; however, the capability of S. oneidensis MR-1 to use DNA as the sole source of phosphorus
is only affected in mutants lacking exeM. Neither of the two endonucleases alleviates toxic effects of increased
eDNA concentrations. The deletion of exeM and/or exeS significantly affects biofilm formation of S. oneidensis
MR-1 under static conditions, and expression of exeM and exeS drastically increases during static biofilm
formation. Under hydrodynamic conditions, a deletion of exeM leads to altered biofilms that consist of densely
packed structures which are covered by a thick layer of eDNA. Based on these results, we hypothesize that a
major role of ExeS and, in particular, ExeM of S. oneidensis MR-1, is to degrade eDNA as a matrix component
during biofilm formation to improve nutrient supply and to enable detachment.
Extracellular DNA (eDNA) occurs in significant amounts in
aquatic and terrestrial ecosystems (68). Concentrations vary
among different environments, ranging from 2 ?g g?1in dry
soil (49) to up to 20 mg g?1organic matter in activated sludge
from wastewater treatment plants (51). The pool of eDNA
provides a reservoir for horizontal gene transfer (45, 68) and a
valuable source of carbon, nitrogen, and, in particular, phos-
phorus (11, 12, 48, 53). Accordingly, it has been demonstrated
that several bacteria have the capacity to use DNA as the sole
nutrient source (17, 48, 53).
In biofilms, the predominant bacterial lifestyle in nature, the
cells are embedded in a self-produced matrix, forming highly
structured aggregates that are commonly attached to surfaces
(60). This matrix, which in many cases comprises the majority
of the communities’ biomass, consists of different biopolymers.
It has previously been reported that eDNA is abundant in
bacterial biofilms (19, 51); in addition, polysaccharides and
proteins are common components (18). Studies of Pseudomo-
nas aeruginosa demonstrated that eDNA in the matrix is re-
quired for structural integrity during the early stages of biofilm
formation (70). Since then, eDNA has been recognized as an
important factor for cell adhesion and as a structural compo-
nent of the biofilm matrix for a growing number of Gram-
positive and Gram-negative species and mixed communities
(15, 59). However, its exact role in biofilm formation remains
to be elucidated. In addition to a structural and/or adhesive
role in microbial biofilms, eDNA was demonstrated to have
antimicrobial activity through chelation of ions required for
stabilizing lipopolysaccharides and/or outer membranes (47).
The release of DNA is thought to be mediated by cell lysis (37,
41, 55, 56, 63), active transport (25), or vesiculation (1, 70). In
contrast, given the significance of eDNA as a structural com-
ponent, potential nutrient, antimicrobial agent, and reservoir
for gene transfer, surprisingly little is known about the role of
eDNA degradation in biofilms. Recently, it was demonstrated
that nuclease-mediated degradation of eDNA is required for
normal biofilm formation. It is thought that the nuclease ac-
tivity enables detachment of cells from Staphylococcus aureus
and Neisseria gonorrhoeae biofilms (41, 58).
The dissimilatory iron-reducing gammaproteobacterium
Shewanella oneidensis MR-1 adheres to surfaces and forms
complex and dynamic surface-associated biofilms (2, 42, 62,
64–66). Biofilm formation is thought to positively impact the
ability of Shewanella species to access insoluble alternative
electron acceptors, such as ferric or manganese minerals (10,
24, 43). We have recently demonstrated that biofilm formation
of S. oneidensis MR-1 strongly depends on eDNA as a struc-
tural component. The DNA is predominantly released by pro-
phage-mediated lysis of a subpopulation of the cells. In the
absence of eDNA, S. oneidensis MR-1 is severely reduced in
surface adhesion and subsequent formation of three-dimen-
sional structures. DNase I treatments of S. oneidensis MR-1
biofilms released large amounts of biomass (22). In addition, a
recent study showed that Shewanella species can use eDNA as
the sole source of phosphorus, carbon, and energy. Accord-
ingly, nuclease activity was determined in S. oneidensis MR-1
cultures, and two putative extracellular nucleases were identi-
fied (27, 53). The activities and functions of both proteins are
not yet known. However, with respect to the important role of
eDNA as structural component, we hypothesized that these
nucleases might also affect S. oneidensis MR-1 biofilm forma-
tion. In this study, we characterized whether the two predicted
extracellular endonucleases are involved in degradation of
eDNA by S. oneidensis MR-1. Furthermore, we determined
* Corresponding author. Mailing address: MPI fu ¨r terrestrische
Mikrobiologie, Karl-von-Frisch-Strasse 10, 35043 Marburg, Germany.
Phone: 49 6421 178301. Fax: 49 6421 178309. E-mail: thormann
‡ These authors contributed equally to this study.
† Supplemental material for this article may be found at http://aem
?Published ahead of print on 24 June 2011.
whether both proteins affect the ability of this species to exploit
eDNA as a nutrient, to tolerate large amounts of eDNA, or to
MATERIALS AND METHODS
Bacterial strains and growth conditions. The bacterial strains used in this
study are listed in Table 1. Escherichia coli and S. oneidensis strains were rou-
tinely grown in LB medium at 37°C and 30°C, respectively. For plates, agar was
added to a final concentration of 1.5% (wt/vol). For the conjugation strain E. coli
WM3064, 2,6-diaminopimelic acid (DAP) was added to the medium to a final
concentration of 300 ?M.
Anaerobic growth was assayed in LB medium adjusted to pH 7.5, containing
40 mM lactate and 20 mM fumarate as described previously (38). For growth
assays, S. oneidensis strains were cultivated in modified M1 minimal medium
(53), containing 30 mM piperazine-N,N?-bis(2-ethanesulfonic acid) (PIPES), 30
mM NaCl, 3 mM MgCl2, 1.34 mM KCl, 6.8 ?M CaCl2, and 10 ml of a 100?
mineral solution used for 4M medium, containing 6.72 mM Na2EDTA, 5.66 mM
H3BO3, 0.54 mM FeCl2, 0.5 mM CoCl2, 0.5 mM NiCl2, 0.39 mM Na2MO4, 0.15
mM Na2SeO4, 0.13 mM MnSO4, 0.1 mM ZnSO4, and 0.02 mM CuSO4(20).
Either 40 mM lactate, 0.86 mM NaH2PO4, 28 mM NH4Cl, or DNA was used as
the source of carbon, phosphorus, and nitrogen, respectively. All growth exper-
iments were performed using DNA from salmon sperm (Sigma, Deisenhofen,
If necessary, media were supplemented with 6 ?g ml?1chloramphenicol
and/or 30 ?g ml?1kanamycin. Biofilms of S. oneidensis were cultivated in LM
medium (52) without antibiotics containing 0.5 mM lactate. DNase I (Serva
Electrophoresis GmbH, Heidelberg, Germany) was used at a concentration of 30
?g ml?1in medium supplemented with 5 mM MgCl2. DDAO [7-hydroxy-9H-
(1,3-dichloro-9,9-dimethylacridin-2-one)] was used at a concentration of 4 ?M to
stain extracellular DNA in biofilms grown under hydrodynamic conditions.
Vector and strain construction. Molecular methods were carried out accord-
ing to standard protocols (57) or according to the manufacturer’s instructions.
Kits for the isolation of plasmids and the purification of PCR products were
purchased from HISS Diagnostics GmbH (Freiburg, Germany). Enzymes were
purchased from New England BioLabs (Frankfurt, Germany) and Fermentas
(St. Leon-Rot, Germany). The strains and plasmids used in this study are sum-
marized in Table 1.
Markerless in-frame deletion mutants of S. oneidensis MR-1 were constructed
essentially as previously reported (38) using the suicide vector pNTPS-138-R6K
and appropriate primer pairs, as summarized in Table S1 in the supplemental
material. Complementation of the mutants was carried out by reintegration of
the deleted fragment into the appropriate deletion strain using the same protocol
as for the deletion. After complementation, all strains retained the wild-type
For biofilm studies, S. oneidensis MR-1 strains constitutively expressing the
green fluorescent protein (GFP) gene gfp were constructed by using a modified
Tn7 delivery system as reported earlier (22). Briefly, the plasmid pUC18-R6KT-
miniTn7T-egfp coding for enhanced GFP (EGFP) was used for tagging S. one-
idensis strains by three-parental mating from the DAP auxotroph E. coli
WM3064 and E. coli strain WM3064 harboring the helper plasmid pTNS2.
Comparative growth and biofilm experiments ensured that no detrimental effects
based on fluorescent protein expression occurred.
For the construction of constitutively luminescent S. oneidensis MR-1 strains,
the luxCDABE operon of Photorhabdus luminescens and a transcriptional termi-
nator cassette were amplified from pUC18-mini-Tn7T-Gm-lux and cloned into
the broad-host-range vector pBBR1-MCS5, using KpnI and SacI/XhoI, respec-
tively, yielding vector pBBR1-MCS5-TT-RBS-lux. Lux expression was conferred
TABLE 1. Bacterial strains and plasmids used in this study
Strain or plasmid Relevant genotypea
Source or reference
DH5? ? pir
?80dlacZ?M15 ?(lacZYA-argF)U169 recA1 hsdR17 deoR thi-l supE44 gyrA96
thrB1004 pro thi rpsL hsdS lacZ?M15 RP4-1360 ?(araBAD)567
?dapA1341::?erm pir (wt)?
WM3064W. Metcalf, University
of Illinois, Urbana
S. oneidensis MR-1 wild type
MR-1, tagged with EGFP in a mini-Tn7 construct; Cmr
MR-1 ?exeM; deletion of gene SO_1066
MR-1 ?exeM, tagged with EGFP in a mini-Tn7 construct; Cmr
Wild type (reintegration of ?exeM)
MR-1 ?exeS; deletion of gene SO_1844
MR-1 ?exeS, tagged with EGFP in a mini-Tn7 construct; Cmr
Wild type (reintegration of ?exeS)
MR-1 ?exeM ?exeS
MR-1 ?exeM ?exeS, tagged with EGFP in a mini-Tn7 construct; Cmr
oriT mobRK2 oriR lacZ?, cloning vector; Gmr
luxCDABE and terminators lambda T0 rrnB1 T1 cloned into pBBR1-MCS5
for plasmid-based transcriptional fusions; Gmr
motAB promoter of S. oneidensis MR-1 cloned into pBBR1-MCS5-TT-RBS-
oriT ori-R6K sacB, suicide plasmid for generating in-frame deletions; Kmr
Fragment for in-frame deletion of SO_1066 in pNTPS-R6K; Kmr
exeM in pNTPS-R6K for reintegration; Kmr
Fragment for in-frame deletion of SO_1844 in pNTPS-R6K; Kmr
exeS in pNTPS-R6K for reintegration; Kmr
ori-R6K; encodes the TnsABC ? D-specific transposition pathway; Apr
ColE1 replicon, mini-Tn7; luxCDABE AprGmr
NotI-egfp-Cmr-NotI fragment from pBK-miniTn7-gfp3 in pUC18-R6KT-
aApr, ampicillin resistance; Cmr, chloramphenicol resistance; Gmr, gentamicin resistance; Kmr, kanamycin resistance.
VOL. 77, 2011NUCLEASES AFFECT SHEWANELLA BIOFILM FORMATION5343
by cloning the constitutively transcribed S. oneidensis MR-1 motAB promoter
upstream of the lux operon, using PstI and BamHI. The resulting vector, pBBR1-
MCS5-TT-Pmot-RBS-lux, was introduced into the appropriate strains by elec-
troporation or conjugation.
Cultivation of S. oneidensis MR-1 biofilms. (i) Static conditions. Biofilm cul-
tivation in polystyrene microtiter plates (Sarstedt, Newton, NC) was carried out
essentially as previously described (65). Briefly, freshly diluted overnight cultures
of S. oneidensis MR-1 strains (1:35 in LM medium) were transferred to wells of
polystyrene microtiter plates (170 ?l per well) and incubated for the indicated
times at 30°C. Prior to processing, the density of the planktonic population in the
wells was determined at 600 nm. Subsequently, 10 ?l crystal violet (0.5% [wt/vol])
was added to the wells followed by 10 min of incubation. The wells were washed
with 200 ?l distilled water, and the remaining surface-attached biomass was
quantified indirectly by the solubilization of retained crystal violet by the addition
of 200 ?l ethanol (96% [wt/vol]) followed by measurements of the absorbance at
570 nm using an Infinite M200 plate reader (Tecan, Switzerland). The relative
degree of surface attachment was normalized to that of the wild type. The assay
was repeated in at least three independent experiments.
(ii) Hydrodynamic conditions. Biofilms were cultivated at room temperature
in LM medium in custom-made three-channel flow cells with individual channel
dimensions of 1 by 4 by 40 mm. Microscope coverslips (Roth, Germany) were
used as a colonization surface and glued onto the channels with silicone (Sista-
Henkel, Germany). Assembly, sterilization, and inoculation of the flow system
were performed essentially as previously described (65). Analyses were carried
out in triplicate in at least two independent experiments. For DDAO staining,
the flow was briefly arrested and the dye was added to the medium in the bubble
trap. This process took no longer than 1 min, and control channels ensured that
the short arrest did not affect biofilm development. The biofilm cells were
incubated with the DDAO for 1 h. Microscopic visualization was performed at
defined spots close to the inflow before and after the treatment.
Microscopy and image acquisition. Microscopic visualization of biofilms and
image acquisition were conducted using an inverted Leica TCS SP5 confocal
laser scanning microscope (Leica Microsystems, Wetzlar, Germany) equipped
with ?10/0.3 Plan-Neofluar and ?63/1.2 W C-Apochromate objectives. The flow
chambers were mounted on the microscopic stage without interrupting the flow.
Live imaging of the cells was enabled by using strains that were constitutively
expressing gfp; however, biofilms were not further incubated after DDAO stain-
ing. To display biofilm images, confocal laser scanning microscopy (CLSM)
image stacks were processed using the IMARIS software package (Bitplane AG,
Zu ¨rich, Switzerland) and Adobe Photoshop. For the quantification of the surface
coverage, the image of the confocal plane displaying the cell layer attached to
surface was selected. The amount of surface-attached biomass was determined
by the amount of green pixels (cells) in relation to that of the background (black)
using Adobe Photoshop CS2. For each data point, at least four individual images
from at least two independent experiments were analyzed.
Quantification of extracellular DNA. The quantification of extracellular DNA
in the biofilm supernatant was carried out as previously described (22). Briefly,
the supernatants of statically grown S. oneidensis MR-1 biofilms were collected
after 1, 4, and 24 h of incubation and filter sterilized. One hundred microliters of
a 1:200 dilution of PicoGreen fluorescent dye (Invitrogen/Molecular Probes,
Darmstadt, Germany) was added to biofilm supernatant, and the DNA release
was immediately measured fluorometrically at an excitation wavelength of 485
nm and an emission wavelength of 535 nm using a Tecan Infinite M200 reader
(Tecan, Switzerland). The concentration of extracellular DNA was then calcu-
lated using DNA reference standards prepared in culture medium.
DNA degradation assay. Nuclease activity was determined in a DNA degra-
dation assay essentially as described earlier (53). Overnight cultures of S. one-
idensis MR-1 strains grown in LB medium were diluted to an optical density at
600 nm (OD600) of 0.05 and incubated in LB or 4M medium until cells reached
an OD600of 1.9. Afterwards, 230-?l aliquots of washed cell culture and filter-
sterilized supernatants, respectively, were mixed with 1,006 bp or 3,140 bp of
PCR-amplified DNA fragments at a final concentration of 5 ?g ml?1. The
samples were incubated at 30°C. Every 15 min, 25-?l aliquots were removed and
analyzed by agarose gel electrophoresis. The absence of DNA on the gel was
assumed to be an indicator for complete DNA digestion. The assay was repeated
in at least two independent experiments.
Extraction of total RNA from S. oneidensis MR-1. RNA was extracted from
static biofilm cultures incubated in petri dishes. To this end, overnight cultures of
S. oneidensis MR-1 strains grown in LM medium were diluted to an OD600of
0.01 and transferred into petri dishes for static biofilm formation. At appropriate
time points, the medium supernatant was collected and the surface-associated
cells were collected in 1:10 stop solution (5% [vol/vol] phenol, 95% [vol/vol]
ethanol [pH 7.4]) via scraping prior to RNA extraction. Cells from biofilm
supernatants and planktonic cultures were harvested by centrifugation (15 min at
4,600 ? g and 4°C). Anaerobic cultures for RNA extraction were grown up to an
OD600of 0.5 before harvesting.
Harvested cells were washed with 2 ml AE buffer (20 mM sodium acetate, 1
mM EDTA) and resuspended in 600 ?l AE buffer. Subsequently, 900 ?l of hot
phenol (preheated to 60°C in a water bath) and 10 ?l of 25% (wt/vol) SDS were
added to the solution, which was then incubated at 60°C for 10 min with occa-
sional inversion and finally cooled on ice. After centrifugation (10 min, 13,000 ?
g), the aqueous phase was transferred to a Phase Lock gel tube (light, 2 ml; 5
PRIME GmbH, Hamburg, Germany) and further supplemented with 900 ?l
60°C hot phenol and 62.5 ?l 2 M sodium acetate solution (pH 5.2) followed by
another 10 min of centrifugation (13,000 ? g). These steps were repeated with-
out adding sodium acetate until no interphase was visible. A total of 2.5 volumes
of ice-cold ethanol (96% [vol/vol]) was used to precipitate the RNA in the
samples by incubation at ?80°C for at least 2 h. After centrifugation at 4°C for
at least 30 min, the RNA was washed with ice-cold ethanol (70% [vol/vol]). After
removal of the supernatant, the RNA precipitate was dried at room temperature
for approximately 1 h and then dissolved in 100 ?l diethyl pyrocarbonate
(DEPC)-treated water. Contaminating DNA was removed using the Turbo
DNA-free kit (Applied Biosystems, Darmstadt, Germany). RNA quality was
determined by agarose gel electrophoresis.
Quantitative RT-PCR (qRT-PCR). For quantitative reverse transcription-PCR
(qRT-PCR), extracted total RNA was applied as a template for random-primed
first-strand cDNA synthesis by using Bioscript reverse transcriptase (Bioline,
Luckenwalde, Germany) according to the manufacturer’s instructions. The
cDNA was used as a template for quantitative PCR (real-time 7300 PCR ma-
chine; Applied Biosystems, Darmstadt, Germany) by using the Sybr green de-
tection system (Applied Biosystems, Darmstadt, Germany). The signals were
standardized to recA, with the cycle threshold (CT) determined automatically by
the Real Time 7300 PCR software (Applied Biosystems), and the total number
of cycles was set to 40. Samples were assayed in duplicate. The efficiency of each
primer pair was determined using four different concentrations of S. oneidensis
MR-1 chromosomal DNA (10, 1.0, 0.1, and 0.01 ng liter?1) as a template in
Growth inhibition and killing assays. Killing assays were performed as previ-
ously described (29, 47). Briefly, overnight cultures of wild-type and mutant
strains carrying pBBR1-MCS5-TT-Pmot-lux were diluted to an OD600of 0.05 in
LB medium and were grown to an OD600of 1.9. Luminescence of 180 ?l cells
was measured using a Tecan Infinite M200 plate reader (Tecan, Switzerland).
Subsequently, 20 ?l herring sperm DNA dissolved in LB medium was directly
added to the wells to yield the appropriate end concentration and mixed well.
Subsequently, the luminescence of the cells was monitored over time as a mea-
sure of viability. Each measurement was performed in duplicate, and the exper-
iments were repeated at least two times. In addition, cells were removed 30 min
after addition of eDNA and diluted in 1:10 steps, and 10 ?l of each deletion was
spotted onto an LB plate.
Identification of two extracellular endonucleases in S. one-
idensis MR-1. Previous bioinformatic analysis of S. oneidensis
MR-1 revealed two genes, SO_1066 and SO_1844, which en-
code putative extracellular endonucleases. Both genes are fol-
lowed by a Rho-independent transcriptional terminator struc-
ture and are likely to be transcribed monocistronically.
SO_1844 is 2,847 bp in length and encodes a protein of 948
amino acids with a predicted molecular mass of 101 kDa. The
protein has a predicted N-terminal signal sequence and is
assumed to be transported in a Sec-dependent manner. Con-
sistently, SO_1844 has so far only been identified in S. one-
idensis MR-1 culture supernatants (53). SO_1066 is 2,616 bp in
length, and the deduced protein of 871 amino acids has a
predicted molecular mass of 93.7 kDa. SO_1066 has an N-ter-
minal signal sequence and, in addition, a putative transmem-
brane domain at the C terminus which could function as a
membrane anchor. In accordance with the predicted localiza-
tion, SO_1066 was found to be associated with the cell enve-
lope in S. oneidensis MR-1 (6, 61). Thus, we will refer to the
5344GO ¨DEKE ET AL.APPL. ENVIRON. MICROBIOL.
proteins encoded by SO_1066 and SO_1844 as ExeM (extra-
cellular endonuclease, membrane associated) and ExeS (extra-
cellular endonuclease, secreted), respectively.
To determine whether ExeM and ExeS are active nucleases,
we performed DNA degradation assays with S. oneidensis
strains lacking one or both enzymes. To this end, we intro-
duced in-frame deletions in the corresponding genes resulting
in single mutants (?exeM and ?exeS) and a double mutant
(?exeM ?exeS). We then determined the nuclease activity of
filter-sterilized supernatants derived from exponentially grow-
ing cultures of the appropriate strains in LB medium (Fig. 1).
In the supernatant of a wild-type culture, degradation of a
defined PCR-derived DNA fragment of 1,006 bp was observed
after 15 min, and degradation was completed after 45 min,
while no nuclease activity was observed in plain LB medium. In
contrast to the wild type, nuclease activity was significantly
decreased in the supernatant of the mutant strain lacking
ExeS. DNA degradation was visible after 30 min and was
completed after 75 min. A mutant lacking ExeM exhibited
even less nuclease activity, and nondegraded DNA fragments
of the original size were still present in the supernatant after 75
min of incubation. The DNA degradation pattern of the dou-
ble mutant equaled that of the ?exeM mutant. Degradation of
a larger 3,140-bp DNA fragment occurred significantly faster
(Fig. 1); however, the relative phenotype of the mutants re-
mained consistent. When a DNA fragment was directly added
to cultures of the wild-type and mutant strains, a degradation
pattern similar to that of the corresponding supernatants oc-
curred (see Fig. S1 in the supplemental material). In a parallel
analysis, we also determined the nuclease activity of the wild
type and the nuclease mutants in mineral medium. The DNA
degradation patterns equaled those obtained in LB medium
(data not shown). Together, the results indicated that both
ExeM and ExeS are active nucleases and that ExeM, which is
predicted to be membrane associated, is also present in cell-
free culture supernatants. Since significant DNA degradation
occurs in the absence of ExeS and ExeM, S. oneidensis MR-1
likely possesses one or more additional proteins with nucleo-
ExeM contributes to utilization of eDNA as the sole source
of phosphorus. Having established that ExeM and ExeS are
active nucleases, we determined whether both enzymes are
required to utilize DNA as the source of phosphorus. To this
end, we tested growth of the wild type and the corresponding
mutants (?exeM, ?exeS, and ?exeM ?exeS) in mineral medium
supplemented with 0.5 g liter?1(0.05%) DNA in the absence
of alternative phosphorus sources (Fig. 2). By quantitative
RT-PCR (qRT-PCR), we determined that both exeM and exeS
are highly upregulated under these conditions (see Fig. 5A).
FIG. 1. DNA degradation by ExeM and ExeS in medium superna-
tants. Filter-sterilized supernatants of the S. oneidensis MR-1 wild-type
strain and nuclease mutants were mixed with PCR-amplified She-
wanella 1,006-bp (A) and 3,140-bp (B) DNA fragments. At the indi-
cated times, aliquots of the samples were removed and analyzed by
agarose gel electrophoresis. In the control lane, the PCR fragment was
added to plain LB medium.
FIG. 2. Aerobic growth of S. oneidensis MR-1 strains with DNA as
the sole source of phosphorus. Growth of the wild-type (WT; circles)
strain, the ?exeM (triangles) and ?exeS (squares) single mutant strains,
and the ?exeM ?exeS (diamonds) double-mutant strain was followed
for 60 h in mineral medium supplemented with either 0.86 mM
NaH2PO4(dashed lines), salmon sperm DNA (0.5 g liter?1; solid
lines), or no source of phosphorus (dotted lines). The error bars
represent the standard deviation.
VOL. 77, 2011NUCLEASES AFFECT SHEWANELLA BIOFILM FORMATION5345
The wild type grew with DNA as the sole source of phospho-
rus; however, a significantly prolonged lag phase was observed
(?24 h). Notably, the exeS mutant grew comparably well with
DNA, like the wild type, strongly indicating that ExeS is not
required for utilization of eDNA. In contrast, the ?exeM mu-
tant had a significantly reduced growth rate with DNA as the
phosphate source, and accordingly, the ?exeM ?exeS double
deletion mutant exhibited a ?exeM growth phenotype. Growth
on DNA as the phosphate source was not totally abolished in
these mutants. In contrast to phosphate limitation, almost no
growth occurred with wild-type and mutant strains when
eDNA was the sole source of nitrogen or carbon (data not
shown). Based on these results, we concluded that ExeM con-
tributes to utilizing eDNA as source of phosphate. As it was
previously established that ExeM and ExeS are not the only
proteins with nucleolytic activity in the supernatant, another
nuclease might be involved in this process. In contrast, ExeS
appears not to be required to use DNA as a nutrient in growing
cultures and probably has a different role in S. oneidensis
ExeM and ExeS do not significantly contribute to the toler-
ance toward elevated eDNA concentrations under planktonic
conditions. Since the matrix of S. oneidensis MR-1 biofilms was
demonstrated to contain significant amounts of eDNA, we
determined whether ExeM and ExeS could alleviate potential
toxic effects of DNA. To this end, we constructed wild-type and
mutant strains constitutively expressing the lux operon of Pho-
torabdus luminescens and recorded the loss of luminescence by
the S. oneidensis wild type and nuclease mutants upon addition
of DNA to exponentially growing cultures. Under these con-
ditions, significant inactivation of the wild type was observed
upon addition of ?1% DNA to the growth medium. Almost
instant cell death occurred upon addition of 2% DNA (Fig.
3A), and the complete loss of viability due to cell lysis was
confirmed by plating and microscopic observation (data not
shown). Notably, we never observed a significant difference in
the detrimental effect of DNA addition to cultures of the wild
type or strains lacking exeM, exeS, or both at any of the con-
centrations tested (Fig. 3B). When sublethal concentrations of
DNA were added to growing cultures, the growth rate was
significantly decreased for a period of time before normal
growth resumed. However, also under those conditions, no
difference in growth rates occurred between the wild-type and
mutant strains lacking ExeM, ExeS, or both (Fig. 3C). The
analysis of exeM and exeS transcriptional levels by qRT-PCR 30
min after addition of eDNA revealed that expression of neither
gene was significantly induced by the presence of DNA (data
not shown). These results demonstrated that S. oneidensis
MR-1 is susceptible to elevated levels of eDNA; however,
under planktonic conditions, ExeM and ExeS do not protect
the cells from these detrimental effects.
ExeM and ExeS affect biofilm formation. Since eDNA is a
major factor in mediating cell-cell and cell-surface interactions
in S. oneidensis MR-1, we next analyzed whether ExeM and/or
ExeS contribute to biofilm formation of this species. Biofilm
formation of the wild-type and mutant strains was character-
ized in static microtiter plate assays and in a hydrodynamic
flow chamber system. When grown in microtiter plates (Fig.
4A), a ?exeS mutant accumulated substantially more (164%)
surface-attached biomass than the wild type. In contrast, the
?exeM mutant displayed decreased biofilm formation (28%
compared to that of the wild type). The ?exeM ?exeS double
mutant formed less (48%) biofilm than wild-type cells. We also
determined whether the absence of ExeM and ExeS results in
a lower concentration of eDNA in the supernatant of static
biofilm cultures (Fig. 4). After 4, 24, and 48 h of incubation, the
amount of eDNA in the supernatants of the exeM mutant and
FIG. 3. Detrimental effect of eDNA. (A) Exponentially growing
LB cultures of S. oneidensis MR-1 harboring pBBR1-MCS5-TT-Pmot-
RBS-lux were supplemented with DNA in the concentrations indi-
cated, and the luminescence (counts per second [cps]) was measured
over time. The data presented display luminescence relative to the
untreated control. (B) Exponentially growing LB cultures of the S.
oneidensis MR-1 wild-type strain and nuclease mutants harboring
pBBR1-MCS5-TT-Pmot-RBS-lux were supplemented with eDNA at a
concentration of 1.5%, and luminescence was measured over time.
(C) Growth of S. oneidensis MR-1 wild type and ?exeM ?exeS mutant
cultures in LB medium in the presence and absence of 0.25% DNA.
All error bars display the standard deviation.
5346GO ¨DEKE ET AL.APPL. ENVIRON. MICROBIOL.
the double mutant was significantly higher (?15 to 30%), re-
spectively, than that in supernatants obtained from the wild
type and ?exeS mutants, indicating the nucleolytic activity of
ExeM. After 48 h of incubation, the concentration of eDNA in
the supernatant reached up to 10 ng ?l?1.
Due to the pronounced biofilm phenotype of exeM and exeS
mutants in static biofilm cultures, we hypothesized that expres-
sion of exeM and/or exeS might be induced under these condi-
tions. To analyze the regulation patterns of the two genes
during biofilm formation under static conditions, we harvested
cells from the supernatant and surface-associated cells at dif-
ferent time points during biofilm formation. RNA was pre-
pared from the cells, and the exeM and exeS transcript levels
were quantified by qRT-PCR. Both genes displayed highly
increased transcript levels in surface-associated cells compared
to cells from the supernatant already early during biofilm for-
mation. One hour after surface attachment, exeM was upregu-
lated by a factor of 7.9 and exeS by a factor of ?20. Similar
values were determined after 24 h of biofilm development (Fig.
5B). Thus, growth under static biofilm conditions strongly in-
duces expression of exeM and exeS and might indicate that the
cells undergo phosphate starvation under such conditions. Pre-
vious studies of S. oneidensis MR-1 biofilms have identified
molecular oxygen levels as a major signal involved in biofilm
formation (66). Based on these studies, we hypothesized that,
in addition to phosphate limitation, expression of exeM and
exeS might also be directly or indirectly controlled by oxygen
levels. During exponential anaerobic growth, exeM was upregu-
lated by a factor of about 2.9 (Fig. 5C). In contrast, expression
of exeS decreased by a factor of 4.4 during anaerobic growth.
Thus, oxygen levels might contribute to the upregulation of
exeM during biofilm formation, while the drastically increased
expression levels of exeS are independent from oxygen levels.
To analyze the biofilm formation in the flow chamber system
via confocal laser scanning microscopy (CLSM), we con-
structed corresponding mutant strains constitutively expressing
gfp. Under hydrodynamic conditions in the flow chamber, no
visible phenotypic change occurred in the ?exeS mutant (Fig.
6). Similar to the wild type, the mutant covered the surface
after 24 h (94.73% ? 2.0% and 91.72% ? 7.0% surface cov-
erage for the wild type and ?exeS mutant, respectively), and
distinct three-dimensional structures were formed after 48 h.
However, under these conditions, the ?exeM mutant had a
distinct phenotype in biofilm formation. While the initial at-
tachment was unaffected, ?exeM mutants were not able to
cover the surface after 24 h (44.95% ? 6.33% coverage versus
94.73% ? 2.0% for the wild type), but instead formed smaller,
densely packed microcolonies. After 48 h, towering three-di-
mensional structures were formed. Notably, few cells of the
?exeM mutant were observed that were not associated with the
biofilm. As in the static microtiter plate system, the biofilm
phenotype of the ?exeM ?exeS double mutant resembled that
of the ?exeM single mutant. To visualize the amount of eDNA
in biofilms grown under hydrodynamic conditions, we applied
DDAO staining (Fig. 6). eDNA was present in both wild-type
and ?exeS mutant biofilms in similar amounts to those previ-
ously described (22). However, a striking difference occurred
in ?exeM and ?exeM ?exeS mutant biofilms, which were cov-
ered by a thick layer of eDNA. Together, the results demon-
strated that ExeM and ExeS affect biofilm formation. While a
significant phenotype of a ?exeS mutant only occurred under
static conditions, the loss of ExeM results in altered biofilm
formation under both static and hydrodynamic conditions, and
large amounts of eDNA accumulated in the matrix.
Earlier studies have identified substantial concentrations of
eDNA in marine sediments (13, 14), an environment from
which numerous Shewanella species have been isolated (26).
The eDNA is assumed to provide a major source of phospho-
rus in marine sediments, and significant nuclease activities
have been determined in these environments (11, 12). Accord-
ingly, a recent study demonstrated that Shewanella species are
capable of eDNA degradation and that eDNA is a particularly
valuable source of phosphorus (53). Significant nuclease activ-
ity occurred in Shewanella cultures, indicating a role for extra-
FIG. 4. Role of ExeM and ExeS in S. oneidensis MR-1 static biofilm
formation. (A) Biofilm formation of the S. oneidensis MR-1 wild-type
strain and ?exeM, ?exeS, and ?exeM ?exeS nuclease mutant strains, as
well as the corresponding complemented mutants (KI-) in microtiter
plates. The strains were incubated in LM medium for 24 h. The
surface-associated biomass was quantified using a crystal violet assay.
The values are means of three replicates, and the standard deviations
are displayed by error bars. (B) Amount of eDNA in the supernatant
of static biofilm cultures of the wild type and the indicated nuclease
mutants 1, 4, 24, and 48 h after surface attachment. The error bars
display the standard deviation.
VOL. 77, 2011NUCLEASES AFFECT SHEWANELLA BIOFILM FORMATION 5347
cellular nucleases in concert with phosphatases in exploiting
DNA as a nutrient, as has recently been demonstrated for
Pseudomonas aeruginosa (48). Two putative extracellular en-
donucleases were previously identified in Shewanella oneiden-
sis MR-1 (53), and both are present in all Shewanella species
sequenced so far. Our study provides evidence that these two
nucleases, now designated ExeM and ExeS, are contributing to
eDNA degradation. Both are highly upregulated under phos-
FIG. 5. Expression of exeM and exeS genes in S. oneidensis MR-1. (A) Transcriptional levels of exeM and exeS during exponential growth on
DNA as the sole source of phosphate as determined by qRT-PCR. The bars display exeM and exeS expression during growth on DNA relative to
expression levels during growth on NaH2PO4. (B) Differences in exeM and exeS transcriptional levels of surface-attached cells compared to those
of planktonic cells during static biofilm formation. The transcription levels of exeM and exeS were determined by qRT-PCR 0.25, 1, and 24 h after
attachment. Displayed are the transcription levels of the nuclease genes in surface-attached cells relative to transcription levels of cells in the
supernatant. (C) Transcriptional levels of exeM and exeS under anaerobic conditions relative to aerobic conditions in planktonic cultures as
quantified by qRT-PCR. All values are means of three replicates. Error bars display the standard deviations.
FIG. 6. Role of ExeM and ExeS in S. oneidensis MR-1 biofilm formation under hydrodynamic conditions. Biofilm formation of GFP-tagged
wild-type and nuclease mutant strains under hydrodynamic conditions was monitored by CLSM after 1, 4, 24, and 48 h of attachment. After 48 h
of incubation, eDNA was visualized by DDAO staining (red). The lateral edge of each micrograph equals 250 ?m.
5348 GO ¨DEKE ET AL.APPL. ENVIRON. MICROBIOL.
phate-limited conditions, as has been observed for other spe-
cies, such as Corynebacterium glutamicum (31), Bacillus licheni-
formis (30, 69), and P. aeruginosa (48). However, the ability of
S. oneidensis to utilize DNA as a source of phosphate was
not abolished in the absence fof ExeM and not affected after
loss of ExeS under all conditions tested, which is likely due
to the fact that S. oneidensis produces one or more addi-
tional nucleases. This remarkable abundance of nucleolytic
activity underlines the significance of eDNA for Shewanella
species. In addition, our study strongly suggests that ExeS
and, in particular, ExeM have functions beyond exploiting
DNA as a nutrient.
ExeS is assumed to be transported via the cytoplasmic mem-
brane, and the protein has so far only been identified in cell-
free supernatants (53). In contrast, two studies addressing the
membrane proteome of S. oneidensis MR-1 have provided
evidence that ExeM is associated with the cell envelope. How-
ever, it is not yet clear whether ExeM is an outer membrane
protein (61) or whether the protein is localized to the cyto-
plasmic membrane (6). The latter would be rather inconsistent
with a role in degradation of an extracellular compound such
as eDNA unless it is first transported through the outer mem-
brane. Although ExeM has been proposed to be associated
with the cell envelope, our study indicates that ExeM-depen-
dent nuclease activity occurs in the cell-free supernatant. If
ExeM is located on the outside of the outer membrane, the
protein might be occasionally released into the supernatant. In
addition, we have shown that S. oneidensis MR-1 undergoes
cell lysis (22), and ExeM might also be transported by or
attached to vesicles produced by Shewanella (23). Notably,
DNA uptake systems involved in natural transformation are
thought to involve membrane-associated nuclease activity (7,
8). In addition, a recent study provides evidence that an extra-
cellular nuclease, Dns, affects natural transformation in Vibrio
cholerae (3). This nuclease is expressed in a cell-density-depen-
dent fashion and prevents transformation by degradation of
eDNA at low cell densities. ExeM and/or ExeS may have a
similar role in S. oneidensis MR-1; however, it remains to be
demonstrated whether this species is naturally competent and
whether the two nucleases are involved in or affect DNA up-
take. Bioinformatic analysis readily identifies numerous genes/
gene clusters that have most likely been acquired by S. one-
idensis MR-1 through lateral gene transfer, such as a second
set of flagellar stators, motAB (52). Moreover, this species
harbors an integron integrase system (16), indicating active
uptake of eDNA in S. oneidensis MR-1.
While ExeM is involved in utilizing eDNA as a source of
phosphate, our study provided evidence that both ExeM and
ExeS contribute to biofilm formation of S. oneidensis MR-1.
The pronounced biofilm phenotype and the significant upregu-
lation of the nucleases during surface-associated growth
strongly suggest functional roles of ExeM and ExeS in the
degradation of eDNA in the biofilm matrix. A recent study of
Pseudomonas aeruginosa and other species has demonstrated
that high concentrations of eDNA are lethal to bacterial cells
(47). This effect is thought to be due to the chelating properties
of DNA that result in a cation-limited environment leading to
the perturbation of inner and outer membranes. Here, we
provide evidence that S. oneidensis is susceptible to detrimen-
tal effects of eDNA in a similar fashion. Our previous studies
indicated that, under hydrodynamic conditions, eDNA is not
evenly distributed but is particularly abundant in the densely
packed three-dimensional structures occurring after 24 h (22).
We have shown that, under planktonic conditions, ExeM and
ExeS do not provide short-term protection from toxic effects of
eDNA. This is not surprising since smaller fragments of DNA
probably can also provide chelating functions until being com-
pletely degraded. However, it is conceivable that ExeM and
ExeS might help to counteract a gradual accumulation of
eDNA in the matrix before it reaches inhibitory levels.
An important but not well understood stage occurring dur-
ing biofilm formation is the detachment of cells from the com-
munity, which enables bacteria to leave under unfavorable
conditions and contributes to biological dispersal and survival
of the cells (32, 34). Previous studies on S. oneidensis MR-1
have shown that cells constantly detach from biofilms and that
rapid detachment can be induced by a rapid drop in the mo-
lecular oxygen level (64, 66). To actively leave the biofilm, cells
are required to modify the matrix that keeps them in the
community. A number of matrix-degrading enzymes have been
described for various species that contribute to detachment.
Some target protein compounds of the matrix (4, 21, 39), while
others degrade the polysaccharide components (5, 33). A re-
cent study has shown that the staphylococcal thermonuclease
(nuc) affects biofilm formation of Staphylococcus aureus (41).
In the absence of nuc, S. aureus forms significantly thicker
biofilms, indicating that this nuclease promotes biofilm disper-
sal. Similar observations were made for Neisseria gonorrhoeae
(58). Similarly, with respect to the important role of eDNA as
a structural matrix component in biofilms of S. oneidensis (22),
we hypothesize that ExeM and ExeS are involved in detach-
ment and dispersal of Shewanella cells from biofilms. ExeS is
predicted to be released into the supernatant to degrade
eDNA within the surface-associated community. Thus, a loss
of ExeS would be expected to result in a more resistant biofilm
matrix containing more biomass, as observed in the static mi-
crotiter plate assay. However, under hydrodynamic conditions,
ExeS is probably removed by the medium flow before a con-
centration is reached that affects matrix formation. Accord-
ingly, we did not observe a phenotype of exeS mutants in flow
chamber-grown biofilms of S. oneidensis MR-1. In contrast to
ExeS, ExeM is thought to be associated with the cell envelope.
Hence, this nuclease would not be removed under hydrody-
namic conditions and might degrade DNA in close proximity
to the cell. Biofilms formed by exeM mutants in the hydrody-
namic flow chamber exhibit tight cell-cell interactions and are
covered by a thick layer of eDNA. In contrast, in the static
microtiter plate assay this mutant displays less surface-associ-
ated biomass. A previous study has demonstrated that tight
cell-cell interactions due to hyperpiliation cause a similar phe-
notype of S. oneidensis MR-1 (65). This finding indicates that
detachment is an important prerequisite for normal biofilm
formation of this species and suggests that a role of ExeM is to
degrade eDNA to prevent the formation of too tight cell-cell
interactions that would prevent detachment.
exeM and exeS were upregulated upon phosphate starvation
in S. oneidensis MR-1. Interestingly, there are reports from
other species that directly link the availability of phosphorus to
biofilm formation. In Pseudomonas fluorescens Pf0-1, a low
level of phosphate leads to activation of the Pho regulon and
VOL. 77, 2011NUCLEASES AFFECT SHEWANELLA BIOFILM FORMATION 5349
increased production of RapA (46). RapA is a diesterase that
degrades the signaling molecule c-di-GMP. Generally, a low
level of c-di-GMP is associated with a transition from a sessile
to planktonic lifestyle in bacteria (28). More specifically, the
degradation of c-di-GMP by RapA is thought to inhibit the
secretion of a large adhesin, LapA, which is required for P.
fluorescens biofilm formation (46). Similarly, activation of the
Pho system positively regulates motility and decreases biofilm
formation in Vibrio cholerae (54). Biofilm formation is also
severely impaired in Proteus mirabilis mutants lacking the high-
affinity transporter Pst (50). Thus, for a number of species,
phosphate limitation appears to favor the transition from the
biofilm to the planktonic lifestyle. Accordingly, production of a
nuclease that degrades the matrix component eDNA under
those conditions not only might improve the acquisition of
nutrients but also would facilitate detachment from the com-
munity. It will be interesting to determine whether Shewanella
biofilms also depend on phosphate in a similar fashion and
whether growth in biofilms might lead to rapid depletion in
phosphate. Thus, further studies will address how regulation of
ExeM and ExeS occurs and whether they are specifically pro-
duced or activated, e.g., during induced biofilm detachment.
Taken together, we have demonstrated here that ExeM and
ExeS are required for normal biofilm formation of S. oneiden-
sis MR-1. We hypothesize that this effect is due to degradation
of eDNA as a matrix component, enabling detachment from
biofilms. Since Shewanella species are not confined to extreme
habitats, it is likely that they commonly occur in mixed-species
rather than in monospecies communities (35, 40). In contrast
to many other extracellular polymeric substance (EPS) com-
pounds, eDNA represents a universal matrix component and is
required for interaction of mixed-species biofilms (15). Thus,
the production of extracellular nucleases might be an effective
means to enable detachment from such multispecies biofilms.
Given the wide distribution of eDNA as a structural compo-
nent of bacterial biofilms, we expect that similar systems are
required for biofilm formation and dispersal of other species.
We are grateful to Penelope Higgs, Martin Thanbichler, and Chris
van der Does for critical reading of the manuscript.
The study was supported by a grant from the Deutsche Forschungs-
gemeinschaft (DFG; TH831/3-1) and the Max-Planck-Gesellschaft.
1. Allesen-Holm, M., et al. 2006. A characterization of DNA release in Pseu-
domonas aeruginosa cultures and biofilms. Mol. Microbiol. 59:1114–1128.
2. Bagge, D., M. Hjelm, C. Johansen, I. Huber, and L. Gram. 2001. Shewanella
putrefaciens adhesion and biofilm formation on food processing surfaces.
Appl. Environ. Microbiol. 67:2319–2325.
3. Blokesch, M., and G. K. Schoolnik. 2008. The extracellular nuclease Dns and
its role in natural transformation of Vibrio cholerae. J. Bacteriol. 190:7232–
4. Boles, B. R., and A. R. Horswill. 2008. Agr-mediated dispersal of Staphylo-
coccus aureus biofilms. PLoS Pathog. 4:e1000052.
5. Boyd, A., and A. M. Chakrabarty. 1994. Role of alginate lyase in cell de-
tachment of Pseudomonas aeruginosa. Appl. Environ. Microbiol. 60:2355–
6. Brown, R. N., M. F. Romine, A. A. Schepmoes, R. D. Smith, and M. S.
Lipton. 2010. Mapping the subcellular proteome of Shewanella oneidensis
MR-1 using sarkosyl-based fractionation and LC-MS/MS protein identifica-
tion. J. Proteome Res. 9:4454–4463.
7. Burton, B., and D. Dubnau. 2010. Membrane-associated DNA transport
machines. Cold Spring Harbor Perspect. Biol. 2:a000406.
8. Chen, I., P. J. Christie, and D. Dubnau. 2005. The ins and outs of DNA
transfer in bacteria. Science 310:1456–1460.
9. Choi, K. H., et al. 2005. A Tn7-based broad-range bacterial cloning and
expression system. Nat. Methods 2:443–448.
10. Das, A., and F. Caccavo, Jr. 2000. Dissimilatory Fe(III) oxide reduction by
Shewanella alga BrY requires adhesion. Curr. Microbiol. 40:344–347.
11. Dell’Anno, A., and C. Corinaldesi. 2004. Degradation and turnover of ex-
tracellular DNA in marine sediments: ecological and methodological con-
siderations. Appl. Environ. Microbiol. 70:4384–4386.
12. Dell’Anno, A., and R. Danovaro. 2005. Extracellular DNA plays a key role in
deep-sea ecosystem functioning. Science 309:2179.
13. Dell’Anno, A., M. Fabiano, G. C. A. Duineveld, A. Kok, and R. Danovaro.
1998. Nucleic acid (DNA, RNA) quantification and RNA/DNA ratio deter-
mination in marine sediments: comparison of spectrophotometric, fluoro-
metric, and high-performance liquid chromatography methods and estima-
tion of detrital DNA. Appl. Environ. Microbiol. 64:3238–3245.
14. Dell’Anno, A., M. Fabiano, M. L. Mei, and R. Danovaro. 1999. Pelagic-
benthic coupling of nucleic acids in an abyssal location of the northeastern
Atlantic Ocean. Appl. Environ. Microbiol. 65:4451–4457.
15. Dominiak, D. M., J. L. Nielsen, and P. H. Nielsen. 2011. Extracellular DNA
is abundant and important for microcolony strength in mixed microbial
biofilms. Environ. Microbiol. 13:710–721.
16. Drouin, F., J. Melancon, and P. H. Roy. 2002. The IntI-like tyrosine recom-
binase of Shewanella oneidensis is active as an integron integrase. J. Bacte-
17. Finkel, S. E., and R. Kolter. 2001. DNA as a nutrient: novel role for bacterial
competence gene homologs. J. Bacteriol. 183:6288–6293.
18. Flemming, H. C., and J. Wingender. 2010. The biofilm matrix. Nat. Rev.
19. Frølund, B., R. Palmgren, K. Keiding, and P. H. Nielsen. 1996. Extraction of
extracellular polymers from activated sludge using a cation exchange resin.
Water Res. 30:1749–1758.
20. Gescher, J. S., C. D. Cordova, and A. M. Spormann. 2008. Dissimilatory iron
reduction in Escherichia coli: identification of CymA of Shewanella oneidensis
and NapC of Escherichia coli as ferric reductases. Mol. Microbiol. 68:706–
21. Gjermansen, M., M. Nilsson, L. Yang, and T. Tolker-Nielsen. 2010. Char-
acterization of starvation-induced dispersion in Pseudomonas putida bio-
films: genetic elements and molecular mechanisms. Mol. Microbiol. 75:815–
22. Go ¨deke, J., K. Paul, J. Lassak, and K. M. Thormann. 2011. Phage-induced
lysis enhances biofilm formation in Shewanella oneidensis MR-1. ISME J.
23. Gorby, Y., et al. 2008. Redox-reactive membrane vesicles produced by She-
wanella. Geobiology 6:232–421.
24. Gorby, Y. A., et al. 2006. Electrically conductive bacterial nanowires pro-
duced by Shewanella oneidensis strain MR-1 and other microorganisms.
Proc. Natl. Acad. Sci. U. S. A. 103:11358–11363.
25. Hamilton, H. L., N. M. Dominguez, K. J. Schwartz, K. T. Hackett, and J. P.
Dillard. 2005. Neisseria gonorrhoeae secretes chromosomal DNA via a novel
type IV secretion system. Mol. Microbiol. 55:1704–1721.
26. Hau, H. H., and J. A. Gralnick. 2007. Ecology and biotechnology of the
genus Shewanella. Annu. Rev. Microbiol. 61:237–258.
27. Heidelberg, J. F., et al. 2002. Genome sequence of the dissimilatory metal
ion-reducing bacterium Shewanella oneidensis. Nat. Biotechnol. 20:1118–
28. Hengge, R. 2009. Principles of c-di-GMP signalling in bacteria. Nat. Rev.
29. Hilpert, K., and R. E. Hancock. 2007. Use of luminescent bacteria for rapid
screening and characterization of short cationic antimicrobial peptides syn-
thesized on cellulose using peptide array technology. Nat. Protoc. 2:1652–
30. Hoi le, T., et al. 2006. The phosphate-starvation response of Bacillus licheni-
formis. Proteomics 6:3582–3601.
31. Ishige, T., M. Krause, M. Bott, V. F. Wendisch, and H. Sahm. 2003. The
phosphate starvation stimulon of Corynebacterium glutamicum determined
by DNA microarray analyses. J. Bacteriol. 185:4519–4529.
32. Kaplan, J. B. 2010. Biofilm dispersal: mechanisms, clinical implications, and
potential therapeutic uses. J. Dent. Res. 89:205–218.
33. Kaplan, J. B., C. Ragunath, N. Ramasubbu, and D. H. Fine. 2003. Detach-
ment of Actinobacillus actinomycetemcomitans biofilm cells by an endoge-
nous beta-hexosaminidase activity. J. Bacteriol. 185:4693–4698.
34. Karatan, E., and P. Watnick. 2009. Signals, regulatory networks, and mate-
rials that build and break bacterial biofilms. Microbiol. Mol. Biol. Rev.
35. Kobayashi, T., et al. 2008. Phylogenetic and enzymatic diversity of deep
subseafloor aerobic microorganisms in organics- and methane-rich sedi-
ments off Shimokita Peninsula. Extremophiles 12:519–527.
36. Kovach, M. E., et al. 1995. Four new derivatives of the broad-host-range
cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes.
37. Lappann, M., et al. 2010. A dual role of extracellular DNA during biofilm
formation of Neisseria meningitidis. Mol. Microbiol. 75:1355–1371.
38. Lassak, J., A. L. Henche, L. Binnenkade, and K. M. Thormann. 2010. ArcS,
5350GO ¨DEKE ET AL.APPL. ENVIRON. MICROBIOL.
the cognate sensor kinase in an atypical Arc system of Shewanella oneidensis
MR-1. Appl. Environ. Microbiol. 76:3263–3274.
39. Lee, S. F., Y. H. Li, and G. H. Bowden. 1996. Detachment of Streptococcus
mutans biofilm cells by an endogenous enzymatic activity. Infect. Immun.
40. Liu, W., et al. 2010. Geochip-based functional gene analysis of anodophilic
communities in microbial electrolysis cells under different operational
modes. Environ. Sci. Technol. 44:7729–7735.
41. Mann, E. E., et al. 2009. Modulation of eDNA release and degradation
affects Staphylococcus aureus biofilm maturation. PLoS One 4:e5822.
42. McLean, J. S., et al. 2008. Investigations of structure and metabolism within
Shewanella oneidensis MR-1 biofilms. J. Microbiol. Methods 74:47–56.
43. McLean, J. S., et al. 2010. Quantification of electron transfer rates to a solid
phase electron acceptor through the stages of biofilm formation from single
cells to multicellular communities. Environ. Sci. Technol. 44:2721–2727.
44. Miller, V. L., and J. J. Mekalanos. 1988. A novel suicide vector and its use
in construction of insertion mutations: osmoregulation of outer membrane
proteins and virulence determinants in Vibrio cholerae requires toxR. J.
45. Molin, S., and T. Tolker-Nielsen. 2003. Gene transfer occurs with enhanced
efficiency in biofilms and induces enhanced stabilisation of the biofilm struc-
ture. Curr. Opin. Biotechnol. 14:255–261.
46. Monds, R. D., P. D. Newell, R. H. Gross, and G. A. O’Toole. 2007. Phosphate-
dependent modulation of c-di-GMP levels regulates Pseudomonas fluo-
rescens Pf0-1 biofilm formation by controlling secretion of the adhesin LapA.
Mol. Microbiol. 63:656–679.
47. Mulcahy, H., L. Charron-Mazenod, and S. Lewenza. 2008. Extracellular
DNA chelates cations and induces antibiotic resistance in Pseudomonas
aeruginosa biofilms. PLoS Pathog. 4:e1000213.
48. Mulcahy, H., L. Charron-Mazenod, and S. Lewenza. 2010. Pseudomonas
aeruginosa produces an extracellular DNase that is required for utilization of
DNA as a nutrient source. Environ. Microbiol. 12:1621–1629.
49. Niemeyer, J., and F. Gessler. 2002. Determination of free DNA in soils. J.
Plant Nutr. Soil Sci. 165:121–124.
50. O’May, G. A., et al. 2009. The high-affinity phosphate transporter Pst in
Proteus mirabilis HI4320 and its importance in biofilm formation. Microbi-
51. Palmgren, R., and P. H. Nielsen. 1996. Accumulation of DNA in the exo-
polymeric matrix of activated sludge and bacterial cultures. Water Sci. Tech-
52. Paulick, A., et al. 2009. Two different stator systems drive a single polar
flagellum in Shewanella oneidensis MR-1. Mol. Microbiol. 71:836–850.
53. Pinchuk, G. E., et al. 2008. Utilization of DNA as a sole source of phospho-
rus, carbon, and energy by Shewanella spp.: ecological and physiological
implications for dissimilatory metal reduction. Appl. Environ. Microbiol.
54. Pratt, J. T., E. McDonough, and A. Camilli. 2009. PhoB regulates motility,
biofilms, and cyclic di-GMP in Vibrio cholerae. J. Bacteriol. 191:6632–6642.
55. Qin, Z., et al. 2007. Role of autolysin-mediated DNA release in biofilm
formation of Staphylococcus epidermidis. Microbiology 153:2083–2092.
56. Rice, K. C., et al. 2007. The cidA murein hydrolase regulator contributes to
DNA release and biofilm development in Staphylococcus aureus. Proc. Natl.
Acad. Sci. U. S. A. 104:8113–8118.
57. Sambrook, K., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a
laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold
Spring Harbor, NY.
58. Steichen, C. T., C. Cho, J. Q. Shao, and M. A. Apicella. 2011. The Neisseria
gonorrhoeae biofilm matrix contains DNA and an endogenous nuclease con-
trols its incorporation. Infect. Immun. 79:1504–1511.
59. Steinberger, R. E., and P. A. Holden. 2005. Extracellular DNA in single- and
multiple-species unsaturated biofilms. Appl. Environ. Microbiol. 71:5404–
60. Stoodley, P., K. Sauer, D. G. Davies, and J. W. Costerton. 2002. Biofilms as
complex differentiated communities. Annu. Rev. Microbiol. 56:187–209.
61. Tang, X., et al. 2007. Profiling the membrane proteome of Shewanella one-
idensis MR-1 with new affinity labeling probes. J. Proteome Res. 6:724–734.
62. Teal, T. K., D. P. Lies, B. J. Wold, and D. K. Newman. 2006. Spatiometabolic
stratification of Shewanella oneidensis biofilms. Appl. Environ. Microbiol.
63. Thomas, V. C., L. R. Thurlow, D. Boyle, and L. E. Hancock. 2008. Regulation
of autolysis-dependent extracellular DNA release by Enterococcus faecalis
extracellular proteases influences biofilm development. J. Bacteriol. 190:
64. Thormann, K. M., et al. 2006. Control of formation and cellular detachment
from Shewanella oneidensis MR-1 biofilms by cyclic di-GMP. J. Bacteriol.
65. Thormann, K. M., R. M. Saville, S. Shukla, D. A. Pelletier, and A. M.
Spormann. 2004. Initial phases of biofilm formation in Shewanella oneidensis
MR-1. J. Bacteriol. 186:8096–8104.
66. Thormann, K. M., R. M. Saville, S. Shukla, and A. M. Spormann. 2005.
Induction of rapid detachment in Shewanella oneidensis MR-1 biofilms. J.
67. Venkateswaran, K., et al. 1999. Polyphasic taxonomy of the genus Shewanella
and description of Shewanella oneidensis sp. nov. Int. J. Syst. Bacteriol.
68. Vlassov, V. V., P. P. Laktionov, and E. Y. Rykova. 2007. Extracellular nucleic
acids. Bioessays 29:654–667.
69. Voigt, B., et al. 2006. The extracellular proteome of Bacillus licheniformis
grown in different media and under different nutrient starvation conditions.
70. Whitchurch, C. B., T. Tolker-Nielsen, P. C. Ragas, and J. S. Mattick. 2002.
Extracellular DNA required for bacterial biofilm formation. Science 295:22.
VOL. 77, 2011NUCLEASES AFFECT SHEWANELLA BIOFILM FORMATION5351