TOL plasmid carriage enhances biofilm formation and increases extracellular DNA content in...

R E S E A R C H L E T T E R

TOLplasmid carriage enhancesbio¢lmformationand increasesextracellularDNAcontent inPseudomonasputidaKT2440Paul W. D’Alvise1, Ole R. Sjøholm2, Tatiana Yankelevich1, Yujie Jin3, Stefan Wuertz3 & Barth F. Smets1

1Department of Environmental Engineering, Technical University of Denmark, Kongens Lyngby, Denmark; 2Department of Agriculture and Ecology,

Faculty of Life Sciences, University of Copenhagen, Frederiskberg, Denmark; and 3Department of Civil and Environmental Engineering, University of

California, Davis, CA, USA

Correspondence: Barth F. Smets,

Department of Environmental Engineering,

Technical University of Denmark, Miljøvej, DK-

2800 Kongens Lyngby, Denmark. Tel.: 145 45

252 230; fax: 145 24 217 034; e-mail:

[email protected]

Received 19 March 2010; revised 16 July 2010;

accepted 16 August 2010.

Final version published online 15 September

2010.

DOI:10.1111/j.1574-6968.2010.02105.x

Editor: Steve Diggle

Keywords

TOL plasmid; biofilm; extracellular DNA;

air–liquid interface.

Abstract

Adherent growth of Pseudomonas putida KT2440 with and without the TOL

plasmid (pWWO) at the solid–liquid and air–liquid interface was examined. We

compared biofilm formation on glass in flow cells, and assayed pellicle (air–liquid

interface biofilm) formation in stagnant liquid cultures by confocal laser scanning

microscopy. The TOL-carrying strains formed pellicles and thick biofilms, whereas

the same strains without the plasmid displayed little adherent growth. Microscopy

using fluorescent nucleic acid-specific stains revealed differences in the production

of extracellular polymeric substances: TOL carriage leads to more extracellular

DNA (eDNA) in pellicles and biofilms. Pellicles were dissolved by DNase I

treatment. Enhanced cell lysis due to plasmid carriage was ruled out as the

mechanism for eDNA release. We report, for the first time, that carriage of a

conjugative plasmid leads to increased biofilm formation by production of eDNA.

Introduction

The ubiquitous soil bacterium Pseudomonas putida can meta-

bolize a wide range of natural and synthetic organic com-

pounds and may play a central role in the natural degradation

of soil pollutants. Pseudomonas putida’s natural niche is in

soils, where it colonizes and forms biofilms on roots or soil

particles. When examined in laboratory flow cells, P. putida

strains are often poor biofilm formers, yielding patchy, thin,

discontinuous, and weak biofilms (Tolker-Nielsen et al., 2000;

Gjermansen et al., 2005; Chang et al., 2007; Rochex &

Lebeault, 2007). The ability to form biofilms is, however, a

prerequisite for a number of industrial and environmental

applications, driving the interest in understanding the mole-

cular and environmental factors that govern biofilm forma-

tion in P. putida. The consensus is that biofilm formation can

be described as a sequential process that involves (1) transport

of cells to a surface, (2) initial reversible attachment, (3)

formation of microcolonies, and (4) the further expansion

and maturation of the biofilm (O’Toole et al., 2000).

Working with various model organisms, it has been

demonstrated that several physiological events are impor-

tant or essential in the initial development and maturation

of biofilms, such as cellular motility, synthesis of exopoly-

meric substances, synthesis of adhesins, and cell-to-cell

signalling (O’Toole et al., 2000; Monds & O’Toole, 2009).

The genetic elements underlying these processes, as well as

the environmental cues controlling their expression, have

been increasingly documented. Nevertheless, it appears that

multiple pathways might exist for biofilm development,

even within a single species, and that environmental condi-

tions may play a significant role (Karatan & Watnick, 2009).

In a seminal paper, Ghigo (2001) demonstrated that

derepressed conjugal plasmids have a stimulatory effect on

Escherichia coli K-12 biofilm formation: using the F plasmid,

the presence and expression of the traA gene was shown to

be sufficient and necessary to observe the stimulatory effect,

and the direct involvement of conjugal pili was inferred. It

was later documented that expression of conjugal pili alone

obviates the need for expression of other cellular factors

typically assumed to be necessary for biofilm formation (e.g.

flagella, fimbriae, curli) in E. coli (Reisner et al., 2003).

The observation that carriage of conjugal plasmids can

enhance biofilm formation, then, suggests a simple way by

FEMS Microbiol Lett 312 (2010) 84–92c� 2010 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

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which an organism can be engineered into a stronger biofilm

former. This is especially interesting for strains such as P.

putida, which can be host to a variety of catabolic conjugal

plasmids (Sevastsyanovich et al., 2008), expanding their

intrinsic metabolic range. The TOL plasmid, originally

isolated from P. putida strain mt-2, is one of the best-studied

catabolic plasmids belonging to the IncP-9 group, encoding

biodegradation pathways for toluenes and xylenes (Williams

& Murray, 1974). The TOL plasmid can be transferred to

other pseudomonads and has earlier been reported as

derepressed for transfer (Benson & Shapiro, 1978; Bradley

& Willams, 1982; Ramos-Gonzalez et al., 1991).

During earlier studies, we had observed a stimulatory effect

of TOL plasmid carriage on biofilm formation in P. putida

KT2440 at the air–water interface (Arango Pinedo et al.,

2003). Here, we provide quantitative support for this biofilm

enhancement at both the air–liquid and the liquid–solid

interface and show that extracellular DNA (eDNA) may be

responsible for the plasmid-stimulated interfacial growth.

Materials and methods

Pseudomonas putida KT2440 is a plasmid-free, restriction-

deficient derivative of P. putida mt-2 (Bagdasarian et al.,

1981). TOL is the archetypical catabolic plasmid pWWO

(Williams & Murray, 1974). The TOL-free strain was chromo-

somally tagged with a miniTn5-Plac-gfpmut3b-kanR cassette

(Normander et al., 1998), the TOL plasmid was tagged with a

miniTn5-Plac-gfpmut3b-tetR cassette as per Christensen et al.

(1996), and carried in a wild-type KT2440 host. Strains were

cultured in AB medium [15.1 mM (NH4)2SO4, 42.2 mM

Na2HPO4, 22 mM KH2PO4, 51.3 mM NaCl, 100mM MgCl2,

10mM CaCl2, 1mM FeCl3, Clark & Maaløe, 1967] supplemen-

ted with 1 mM (flow cells) or 40 mM (static cultures) sodium

citrate. Solid media were prepared by adding 20 g L�1 agar to

AB medium. All cultivations were at 25 1C, unless observed

otherwise. Antibiotics were added to all precultures to a final

concentration of 50mg mL�1.

Static cultures were performed in replicate 500-mL Er-

lenmeyer flasks (70 mL medium) for bulk measurements,

EPS extractions, and viscosity measurements or in 20 mL

test tubes (5 mL medium) for microscopy, flow cytometry,

and b-glucosidase assays.

To test DNase effect, duplicate cultures were supplemented

with DNaseI (Qiagen, 20 U mL�1), magnesium chloride

(250mM), and calcium chloride (4mM). Pellicles were sampled

with a 10-mL inoculation loop, or tweezers for very eDNA-rich

pellicles, applied to a microscope slide, and stained with PI

(15mL, 10mg mL�1) or Cytox Orange before viewing.

Flow cell biofilms were established in three-channel flow

cell setups, as described before (Møller et al., 1996). Flow cells

were inoculated by adjusting the OD600 nm of precultures to

1.0, washing and resuspending the cells in 0.9% NaCl, and

injecting 300mL of this suspension into each channel with an

insulin syringe. AB medium was continuously supplied by a

peristaltic pump (Watson & Marlow 205S) to each channel at

a rate of 2.7 mL h�1. After 2 and 7 days of incubation, three-

dimensional image stacks of the biofilms (3 z-stacks from

three replicate channels in duplicate flow cells for each strain)

were recorded by confocal laser scanning microscopy (CLSM).

To study the distribution of dead cells and eDNA in situ, PI

(300mL, 10mg mL�1) was injected into each channel of

another set of 7-day-old flow cell biofilms, after stopping

medium flow, before CLSM.

For CLSM, a Leica TCS SP5 system, equipped with a � 63

apochromatic objective (NA = 1.4) was used. Both green

fluorescent protein (GFP) and PI were excited at 488 nm using

an argon laser. GFP fluorescence signal was collected between

500 and 540 nm, and PI between 610 and 660 nm. Cytox

Orange was excited at 543 nm using a helium–neon laser, and

its emission light was collected between 545 and 615 nm.

Image stacks were analyzed using the computer program

COMSTAT (Heydorn et al., 2002) and values for biovolume and

average biofilm thickness were recorded. Optical sections were

created using the IMARIS image processing software (Bitplane,

Zurich, Switzerland).

To obtain eDNA, culture samples were treated with

10 U mL�1 cellulase at 37 1C for 1 h, followed by treatment

with 10 U mL�1 proteinase K for another 1 h (Wu & Xi,

2009). Treated samples were centrifuged at 10 000 g for

10 min and the resulting supernatant was amended with

0.25 M NaCl, followed by precipitation in 2� 95–100%

ethanol. The precipitate was collected by centrifuging at

10 000 g for 10 min and then washed twice with 95–100%

ethanol. The purified precipitate was dissolved in TE buffer.

Cellular DNA was extracted by first placing the samples in

boiling water for 10 min and then at � 80 1C for 10 min. The

process was repeated and then the sample was centrifuged at

10 000 g for 10 min, and the supernatant was collected.

RAPD analysis was performed as described previously

(Verma et al., 2007) using two different oligonucleotide

primers (OPB07, 50-GGGTAACGCC and OPA09, 50-

GGTGACGCAG). Each 25-mL reaction contained 45 ng

template DNA, 40 pmol of oligonucleotide primers, 1 U

Taq DNA polymerase, 1� PCR buffer, 200mM each dNTP,

and 2.5 mM MgCl2. Amplification was performed by dena-

turation at 94 1C for 3 min, followed by 40 cycles at 94 1C for

1 min, 37 1C for 1 min, 72 1C for 2 min, and a final extension

at 72 1C for 10 min. The RAPD products were analyzed by

gel electrophoresis in a 2% agarose gel. Fragment sizes were

determined by comparison with a standard curve obtained

by plotting known ladder fragment size against the distance

from the loading well to the center of each band, where log

(fragment size) =� 0.0258� distance14.1714, R2 = 0.9385).

Particulate protein contents of the cultures were mea-

sured using the QuantiProTM BCA Assay Kit (Sigma).

FEMS Microbiol Lett 312 (2010) 84–92 c� 2010 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

85TOL plasmid enhances P. putida biofims

Cultures were subject to EPS extraction after Frølund’s

method (Frølund et al., 1996), by adding 10 g of cation-

exchange resin (AB-washed Dowex Marathon, Sigma 91973)

to each culture, intense stirring (300 r.p.m.) overnight at 4 1C,

and centrifugation at 5000 g for 20 min. The supernatants

were stored at 4 1C before further analysis.

Carbohydrates were quantified by the phenol–sulfuric acid

method (Dubois et al., 1956): Homogenized liquid cultures or

extracts were diluted fivefold in 0.2mm filtered distilled water.

Also, 125mL of 5% (w/v) phenol and 625mL of�95% sulfuric

acid were added simultaneously to the diluted samples in

semi-micro photometer cuvettes (Plastibrand 759015). After

10 min incubation at room temperature and 15 min incuba-

tion at 30 1C, absorbance was measured at 490 nm against a

glucose standard (0–100mg mL�1).

For bulk DNA measurements, 1 mL aliquots of the homo-

genized biofilm-containing liquid cultures were lyzed by three

freeze/thaw cycles (5 min at � 20 1C and 5 min at 60 1C).

DNA contents of the lyzate, and also DNA contents of the EPS

extracts, were measured in 200mL dilution series in black 96-

well optical bottom microtiter plates (Nunc 265301). PI was

added to a final concentration of 50mM. Total fluorescence

was measured on a Beckman–Coulter DTX880 multimode

detector using a long-pass emission filter and 535 nm excita-

tion light. Solutions of 0–100mg mL�1 fish sperm DNA (Sigma

74782) were used as a standard.

Nonpurgeable organic carbon contents were determined

with a Total Organic Carbon Analyzer (TOC-Vwp; Shimad-

zu, Columbia, MD) using a standard protocol. The sensitiv-

ity range of the method was 0.5–3500 mg L�1.

Dynamic viscosities of cultures and extracts were deter-

mined by measuring elution times for draining 50 mL

aliquots of the cultures or culture extracts through a glass

burette (i.d. = 2 mm). Using deionized water as a reference

(Z20�C = 1.003 cSt), dynamic viscosities of the suspensions

were determined with Z20�C = tC, with t as elution time and

C as instrument constant.

Exocellular b-glucosidase activity was measured according to

Rath & Herndl, 1994. Replicate static cultures were mixed by

vortexing, sampled, and supernatants were collected by centri-

fugation (10 000 g, 1 min) and microfiltration (0.2mm). 4-

Methylumbelliferyl b-D-glucopyranoside was added (2.5mM)

and fluorescence was read after 30 min (265 nm excitation,

445-nm band pass emission; Cary Eclipse, Varian) against

nonspiked control samples. Microfiltered sonication-lyzed

P. putida cells were used to check method response.

Live/dead staining was performed as per an established

protocol (Nielsen et al., 2009). In brief, replicate static cultures

were mixed and two subsamples of 100mL were removed; each

of these was supplemented with 10mL Sybr Green (Sybr Green

I; Molecular Probes, Eugene, OR) from a 100� stock solution,

10mL PI (Invitrogen, Carlsbad, CA) from a 0.5 mg mL�1 stock

solution, and 10mL yellow–green fluospheres–carboxylate

microspheres (F-8827; Molecular Probes) in a concentration

of 2� 107 mL�1 as internal standards for flow control. The

samples were then supplemented with 870mL MilliQ water

containing 5 mM EDTA for outer-membrane permeabilization

(Nebe-von-Caron et al., 2000). Samples were then vortexed,

incubated in the dark for 15 min, and briefly vortexed again

before being analyzed. Flow cytometric analysis was performed

with a Facs Calibur instrument (Becton Dickinson, CA)

equipped with 488-nm excitation from an argon-ion laser at

15 mW. The channels forward scatter (FSC), side scatter and

fluorescent channels FL1 (530/30 BP) and FL2 (661/16 BP) were

used for detection. Threshold was set for SSC and compensation

was not used. The carrier liquid used was 0.22mm filtered

MilliQ water. Samples were measured for 30 s at low flow speed

(12� 3mL min�1) with event counts below 3000 s�1.

Results and discussion

Increased biofilm formation by TOL-containingstrains

In our hands, the plasmid-free P. putida KT2440 wild-type

strain is a rather weak biofilm former in minimal medium

citrate-fed flow cell experiments, whereas earlier reports

indicated stronger biofilm formation (Tolker-Nielsen et al.,

2000), especially with different carbon sources or under

coculture conditions (Hansen et al., 2007). After 2 days,

small microcolonies were found, but after 7 days, these had

either died or detached, and hardly any adherent biomass

was found (Fig. 1 and Table 1). Carriage of the TOL plasmid

considerably enhanced biofilm formation: all TOL biofilms

consisted of multiple cell layers after 2 days, with single

microcolonies measuring up to 50mm in height and 25 mm

in diameter. After 7 days, some microcolonies measured up

to 100 mm in height, suggesting that detachment had not

affected KT2240 (TOL) biofilms (Fig. 1 and Table 1). All

differences in biovolume and average thickness between

plasmidless and TOL-carrying strains were significant

(Po 0.0001).

In addition, P. putida forms poor air–liquid interface

biofilms (Ude et al., 2006). Here, again, the TOL-carrying

strain formed slimy, coherent pellicles at the air–liquid

interface of liquid cultures, even with shaking at moderate

speed, whereas the plasmid-free strain did not form coher-

ent pellicles (Supporting Information, Fig S1). After pro-

longed incubation, the liquid cultures of the TOL strain

became increasingly viscous [KT2440: 1.6 centistoke (cSt)

(cSt = mm2 s�1) vs. KT2440 (TOL) 6.6 cSt], suggesting that

extracellular polymeric substances (EPS) were produced.

We dismiss the possibility that enhanced biofilm and

pellicle formation is due to a growth enhancement asso-

ciated with TOL plasmid carriage per se. First, plasmid

carriage, under nonselective conditions – as used here –


86 P. W. D’Alvise et al.

typically results in growth impairment, rather than in

enhancement, and we have – specifically for these two

strains –documented a slight reduction in intrinsic growth

kinetics due to plasmid carriage (Seoane et al., 2010).

Second, detailed monitoring of total cell densities in both

static (Table 2) and shaken (data not shown) cultures

indicates very similar profiles and final cell densities of

approximately 108 after 1 day and 109 from day 3 onward.

Only with genetic modification (e.g. by loss of a genomic

EAL domain-encoding gene, or expression of a heterologous

GGDEF domain-encoding gene) does P. putida form persis-

tent biofilms or perceivable pellicles (Gjermansen et al.,

2006; Ude et al., 2006). Here, we show that carriage of the

TOL plasmid significantly enhances flow cell biofilm forma-

tion and results in enhanced pellicle formation.

EPS analysis

Biofilm EPS can contain polysaccharides, proteins, nucleic

acids (Flemming & Wingender, 2002), but few specific

reports exist on the EPS matrix of P. putida biofilms. As a

first approach to address the increased biofilm formation by

the TOL-carrying strain, we attempted to extract EPS from

biofilm-containing stagnant liquid cultures by a standard

protocol using a cation-exchange resin (Frølund et al.,

1996). However, it was not possible to extract the EPS

component that caused the higher viscosity: both extracts

had similar low viscosities [KT2440: 1.3 cSt vs. KT2440

(TOL) 1.8 cSt], and extractable carbohydrate and nucleic

acid contents were similar for both strains (Table 3). This

suggested that a major part of the EPS was not extractable

and remained cell associated. Further attempts to increase

extraction intensities rapidly caused cell lysis, as observed by

rRNA release in extracts (results not shown).

Consequently, we attempted to analyze EPS components

directly in the biofilm-containing liquid cultures. Total

carbohydrate contents were about double as high as in the

extracts, but no striking differences between the two strains

were found. Total DNA contents of the two cultures were,

however, clearly different: The TOL-carrying strain con-

tained twice the amount of total DNA, as compared with

the respective plasmid-free cultures, as similar cellular

biomass contents measured as particulate protein.

eDNA in flow cell biofilms

The next approach involved direct visualization by micro-

scopy. To check for the differential presence of eDNA in

liquid–solid interface biofilms, 7-day-old flow cell biofilms of

the TOL-free or TOL-carrying KT2440 were stained with

propidium iodide (PI). Diffuse PI fluorescence was colocalized

with the larger (presumably older) microcolonies formed by

the TOL-carrying strain, indicating that eDNA was a major

constituent (Fig. 1). The plasmid-free strains, again, did not

form thick biofilms, and eDNA was not observed.

Using similar approaches, eDNA has been observed in

various pure-culture biofilms (e.g. Pseudomonas aeruginosa,

Bacillus subtilis, Enterococcus faecalis, environmental isolate)

(Whitchurch et al., 2002; Bockelmann et al., 2006; Thomas

et al., 2009; Vilain et al., 2009), and eDNA is, therefore,

Table 1. Biovolume and thickness of Pseudomonas putida biofilms after

2 and 7 days formed in flow cells

Days

2 7

KT2440

Biovolume (mm3mm�2) 0.32�0.42 0.04� 0.05

Thickness (mm) 1.4�1.7 4.8� 5.1

KT2440 (TOL)

Biovolume (mm3mm�2) 8.15�4.75 32.56� 19.79

Thickness (mm) 30.4�8.9 15.9� 13.2

Data are mean values (n = 6) and SD is included as � .

Fig. 1. Optical sections of 7-day-old flow cell biofilms of Pseudomonas putida KT2440 (left), P. putida KT2440 TOL (middle), and P. putida KT2440 (TOL)

after staining with PI (right). The main frames are x–y sections, the side frames are z–x and z–y sections. eDNA and dead cells (red) constitute a major

part of the older colonies (right).



increasingly being considered a potential core element of the

biofilm matrix. eDNA has similarly been observed in flocs and

unsaturated biofilms of environmental Pseudomonas (Palmg-

ren & Nielsen, 1996; Steinberger & Holden, 2005). In contrast

to our observations, this eDNA remained easily extractable by

chemical or thermal treatment methods isolates (Palmgren &

Nielsen, 1996; Steinberger & Holden, 2005).

Random amplification of polymorphic DNA(RAPD) analysis of eDNA and cellular DNA

Ultimately, eDNA was successfully extracted from TOL-free

and TOL-carrying KT2440 cultures by enzymatic treatment

using cellulase and proteinase K followed by centrifugation

(Wu & Xi, 2009), without concurrent increase in cell lysis as

ascertained using live/dead cell staining. RAPD analysis was

performed with two oligonucleotide primers (OPB07 and

OPA09) on eDNA as well as cellular DNA extracted from

both strains (Fig. 2). For primer OPB07, each strain yielded

identical eDNA and cellular DNA band patterns (Fig. 2a),

although the patterns were distinct between strains: 11

bands, ranging from 200 bp to 12 kb, were observed for the

wild type, and six bands, ranging from 400 bp to 3 kb, for the

TOL-carrying strain. None of the bands were identical. For

primer OPA09, eDNA and cellular DNA RAPD band

patterns were slightly different after RAPD analysis (Fig.

2b). Cellular DNA from the wild-type strain (yielding

approximately 12 bands) revealed a 4390 bp amplicon

(named B1 in Fig. 2b), which was not found in eDNA

extracts. eDNA yielded approximately 13 bands, of which

two – B3 at 310 bp and B5 at 12 kb – were not visible in

cellular DNA extracts. For the strain carrying TOL, two of

the eight bands in eDNA – B2 at approximately 2150 bp and

B4 at 250 bp – were not identical in size to any of the bands

found in cellular DNA. Overall, eDNA and cellular DNA

RAPD profiles are very similar, consistent with previous

work done on P. aeruginosa strains PG201 and PAO1

(Steinberger & Holden, 2005; Allesen-Holm et al., 2006).

Because eDNA is either released after cell lysis (Lorenz et al.,

1991) or by an active release mechanism (Kreth et al., 2009),

Table 2. Viability, size, and exocellular enzyme activity of KT2440 and

KT2440 (TOL) static cultures

Days

1 2 3 7

KT2440

%Live 45� 7 60� 16 49� 6 30� 7

FSC 1.0� 0.0 1.8� 0.5 1.2� 0.4 1.8� 0.5

%Dead 55� 10 40� 3 51� 2 70� 23

FSC 2.3� 0.1 2.4� 0.5 2� 0.7 2.5� 0.6

Total mL�1 8.2E7� 1.4E7 2.4E8� 3.5E7 5.4E8� 2.1E7 9.9E8� 2.9E8

b-

Glucosidase

0.09� 0.04 0.22� 0.00 0.29� 0.09 1.51� 0.07

KT2440 (TOL)

%Live 93� 14 78� 13 70� 29 58� 7

FSC 0.4� 0.1 1.1� 0.1 1.1� 0.6 2.6� 0.1

%Dead 7� 1 22� 3 30� 16 42� 1

FSC 1.5� 0.2 1.9� 0.1 2� 0.8 3.4� 0.2

Total mL�1 1.3E7� 1.8E6 3.2E8� 4.3E7 7.9E8� 3.5E8 1.3E9� 8.7E7

b-

Glucosidase

0.06� 0.01 0.14� 0.02 0.22� 0.03 2.02� 0.16

Data are mean values (n = 4 for FCM, and n = 2 for b-glucosidase assay)

and SD is included as � .

FSC values are expressed relative to strain KT2440 at day 1; b-glucosi-

dase is expressed in arbitrary fluorescence units; bold entries mark

significant differences between KT2440 and KT2440 (TOL) based on

pair-wise t-tests (P � 0.05).

Table 3. Macromolecular composition of 5-day-old static Pseudomonas

putida cultures and derived extracts

Protein

(mg L�1)

Carbohydrate

(mg L�1)

DNA

(mg L�1)

Organic C

(mg L�1)

KT2440

Culture 604� 16 512� 33 679� 11 ND

Extract ND 270� 36 206� 44 249� 21

KT2440 (TOL)

Culture 581� 103 393� 118 1430� 41 ND

Extract ND 214� 30 144� 26 366� 131

Data are mean values (n = 4) and SD is included as � .

Fig. 2. Comparative RAPD analysis of eDNA and cellular DNA from

Pseudomonas putida 2440 and P. putida 2440 TOL. (a) RAPD of eDNA

and cellular DNA of P. putida 2440 (lanes 2 and 3) and P. putida 2440

carrying TOL plasmid (lanes 4 and 5) with primer OPB07; (b) RAPD of

eDNA and cellular DNA of P. putida 2440 (lanes 2 and 3) and P. putida

2440 carrying TOL plasmid (lanes 4 and 5) with primer OPA09; lane 1

(both panels): -ladder DNA with known fragment sizes.



cellular DNA should be the main source of eDNA. The

difference in RAPD patterns is likely due to partial eDNA

degradation in the extracellular environment. The presence of

the TOL plasmid altered the RAPD band pattern in both

eDNA and cellular DNA, which has not been reported before.

Effect of DNase I on air--liquid interface biofilms

Pellicles (air–liquid interface biofilms) stained with PI or

Cytox Orange, similarly, revealed large amounts of dead cells

and eDNA in the coherent, viscous pellicles of the TOL-

carrying strain (Fig. 3, Fig S2). eDNA was so abundantly

present that eDNA bundles could be directly observed as large

fibrous structures (Fig. 3), which might form as a result of the

sample preparation procedure. The non-TOL-carrying strain

formed loose, noncoherent air–liquid interface biofilms con-

taining fewer dead cells and no visible eDNA. Calcofluor

staining (specific for b1! 4 polysaccharidic bonds) did not

reveal obvious differences between the strains (not shown),

suggesting that cellulose production, observed in some pseu-

domonad biofilms and pellicles (Ude et al., 2006), is not

responsible for the enhanced biofilm phenotype.

To investigate the structural role of eDNA in the pellicles,

a duplicate set of static cultures was grown in the presence of

DNase I (20 U mL�1). The macro- and microscopic appear-

ance and consistency of the pellicles formed by the TOL

strain were markedly altered by incubation with DNase I.

Accumulation of eDNA in the pellicles was prevented,

resulting in strongly reduced cohesiveness and in a smaller

fraction of dead cells. In the cultures of the plasmid-free

strain, DNase I did not change the macroscopic properties of

interfacial growth, but here also, a reduction in the number

of dead cells could be observed (Fig. 3, Fig S2). Similar

strong effects of DNase treatment on biofilm integrity has

been observed for P. aeruginosa, Streptococcus mutans, and

Streptococcus intermedius (Whitchurch et al., 2002; Petersen

et al., 2005). Hence, eDNA may be responsible for the

development or stabilization of the air–liquid interface

biofilm formed by KT2440 TOL. Its removal by DNase

treatment reduces the cohesiveness of the pellicle and

probably results in a higher turnover of the pellicle.

Population-averaged observations in staticcultures

eDNA release in biofilms (P. aeruginosa, E. faecalis) is often

caused by cell lysis under control of density-dependent

mechanisms (Allesen-Holm et al., 2006; Qin et al., 2007;

Thomas et al., 2009), while in other cases, the mechanisms

of its excretion are not clear (Bockelmann et al., 2006; Vilain

Fig. 3. Micrographs of 7-day-old Pseudomonas

putida KT2440 pellicles, with and without TOL

plasmid, grown in the presence or absence of

DNaseI. Live cells appear green (GFP), dead cell

DNA and eDNA appears red (Cytox Orange). The

untreated pellicle of the TOL-carrying strain con-

tains ample amounts of eDNA, which is concen-

trated as thick bundles during sampling with

tweezers.



et al., 2009). Hence, we examined differential culture

viability in the static cultures. Using a live/dead staining

procedure and flow cytometric quantification of cells, three

core observations were made (Table 2). First, TOL carriage

delayed initial increase in culture densities, but final densities

of both cultures were similar. Second, the fraction of dead cells

increased at the end of incubation, but was not affected by

plasmid carriage. Third, cell sizes increased slightly with

culture age, and this effect was strongest for the TOL-carrying

strain (Table 2). Exocellular b-glucosidase activity increased in

both cultures with time, and sharply after 7 days, but with

little relation to TOL carriage. Therefore, we could not obtain

proof for plasmid-carriage-dependent cell lysis as the reason

for increased eDNA concentrations. Similar cell counts and

live/dead fractions were observed in static cultures of both

strains irrespective of plasmid carriage, and measures of

released cellular enzymatic activity were similar.

The stimulatory role of plasmid carriage on biofilm

formation was first documented and examined with E. coli

K-12. The effect was restricted to derepressed plasmids, and

pointed to the need for traA-like gene expression, suggesting

a direct involvement of conjugal pili as adhesion factors

(Ghigo, 2001; Reisner et al., 2003). Observations with a

range of E. coli isolates confirmed that biofilm stimulation

was contingent on active conjugal plasmid transfer (Reisner

et al., 2006). Although some direct proof of IncF-mating

pili involvement in initial biofilm establishment has been

provided (May & Okabe, 2008), the exact mechanisms

responsible for plasmid-mediated biofilm enhancement re-

main unresolved. Yang et al. (2008) have shown that en-

hanced biofilm formation caused by the presence of R1drd19

in E. coli is contingent on the envelope stress response

system, speculating that pili synthesis imposes stress on

membranes. The virulence plasmid pO157 enhances biofilm

formation in E. coli 0157:H7 due to increased exopolysac-

charide production (Lim et al., 2010); carbohydrate mea-

surement did not support this mechanism in our study.

Here, we demonstrate a new link between plasmid

carriage, biofilm formation, and eDNA for P. putida

KT2440. The potential universality and molecular mechan-

ism by which TOL carriage results in excess eDNA remains,

so far, unresolved but do not appear to be related to

enhanced cell lysis, and suggest secretion. Additional studies

will be required to examine the exact mechanism of eDNA

release and the nature of the released eDNA associated with

TOL carriage in P. putida KT22440.

Acknowledgements

This study was supported by an EC-FP6 Marie Curie Excel-

lence Grant (MEXT-CT-2005-024004) to B.F.S. and the Vil-

lum Kan Rasmussen Center for Environmental and

Agricultural Microbiology. We thank N. Kroer and S. Molin

for providing strains and plasmids, B.S. Lauritzen for plasmid

tagging, and N. El Azhari for initial flow cell observations.

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Supporting Information

Additional supporting information may be found in the

online version of this article:

Fig. S1. Observation of little and abundant pellicle forma-

tion in 5-day-old static cultures of Pseudomonas putida

KT2440 and KT2440.

Fig. S2. Micrographs of 1–7-day-old Pseudomonas putida

KT2440 pellicles, with and without TOL plasmid, grown in

presence or absence of DNase I.

Please note: Wiley-Blackwell is not responsible for the

content or functionality of any supporting materials supplied

by the authors. Any queries (other than missing material)

should be directed to the corresponding author for the article.



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