Environmental bacteria produce abundant and diverse antibiofilm compounds

ORIGINAL ARTICLE

Environmental bacteria produce abundant and diverseantibiofilm compoundsJ.T. Farmer1, A.V. Shimkevitch1, P.S. Reilly1, K.D. Mlynek1, K.S. Jensen2,*, M.T. Callahan1,†,K.L. Bushaw-Newton2,‡ and J.B. Kaplan1

1 Department of Biology, American University, Washington, DC, USA

2 Department of Environmental Science, American University, Washington, DC, USA

Keywords

antibiofilm, biofilm, cave, environmental

bacteria, river, screen, soil, Staphylococcus

aureus.

Correspondence

Jeffrey B. Kaplan, Department of Biology,

American University, 4400 Massachusetts

Ave. N.W., Washington, DC 20016, USA.

E-mail: [email protected]

*Present address: Department of Cell Biology

and Molecular Genetics, University of

Maryland, Rockville, MD 20850, USA†Present address: USDA-ARS, Environmental

Microbial and Food Safety Laboratory,

Beltsville, MD 20705, USA‡Present address: Department of Biology,

Northern Virginia Community College,

Annandale, VA 22003, USA

2014/1116: received 29 May 2014, revised

23 July 2014 and accepted 22 August 2014

doi:10.1111/jam.12639

Abstract

Aims: The aim of this study was to isolate novel antibiofilm compounds

produced by environmental bacteria.

Methods and Results: Cell-free extracts were prepared from lawns of bacteria

cultured on agar. A total of 126 bacteria isolated from soil, cave and river

habitats were employed. Extracts were tested for their ability to inhibit

Staphylococcus aureus biofilm in a 96-well microtitre plate assay. A total of 55/

126 extracts (44%) significantly inhibited Staph. aureus biofilm. Seven extracts

were selected for further analysis. The antibiofilm activities in all seven extracts

exhibited unique patterns of molecular mass, chemical polarity, heat stability

and spectrum of activity against Staph. aureus, Staphylococcus epidermidis and

Pseudomonas fluorescens, suggesting that these seven antibiofilm activities were

mediated by unique chemical compounds with different mechanisms of action.

Conclusions: Environmental bacteria produce abundant and diverse

antibiofilm compounds.

Significance and Impact of the Study: Screening cell-free extracts is a useful

method for identifying secreted compounds that regulate biofilm formation.

Such compounds may represent a novel source of antibiofilm agents for

technological development.

Introduction

Biofilms are communities of bacteria, encased in a self-

synthesized polymeric matrix, growing attached to a bio-

tic or abiotic surface (Hall-Stoodley et al. 2004). Biofilms

protect bacteria from cell stressors such as predators, des-

iccation and antibiotics, and they contribute to numerous

problems in both industrial and clinical settings (Kumar

and Anand 1998; Parsek and Singh 2003; Flemming et al.

2013). New methods for preventing and dispersing bio-

films are needed.

The major adhesive components of most biofilms are

extracellular polymeric substances such as proteins,

polysaccharides and DNA (Flemming and Wingender

2010). In addition, secreted molecules such as quorum-

sensing signals, surfactants, matrix-degrading enzymes

and antibiofilm polysaccharides function to regulate bio-

film architecture and mediate the release of cells from

biofilms during the dispersal stage of the biofilm life cycle

(Kaplan et al. 2003; Karatan and Watnick 2009; Kaplan

2010; Worthington et al. 2012; Solano et al. 2014). These

agents sometimes exhibit broad spectrum antibiofilm

activity when tested against biofilms cultured in vitro

(Kaplan et al. 2004; Valle et al. 2006; Bendaoud et al.

2011; Jiang et al. 2011; Rendueles et al. 2011; Karwacki

et al. 2013). Antibiofilm compounds also increase the

Journal of Applied Microbiology 117, 1663--1673 © 2014 The Society for Applied Microbiology 1663

Journal of Applied Microbiology ISSN 1364-5072

effectiveness of conventional antibiotics against biofilms,

and they are nonbiocidal which reduces selective pressure

for the evolution of resistance (Yang et al. 2012; Blackl-

edge et al. 2013; Kostakioti et al. 2013). Such compounds

may represent a novel source of antibiofilm compounds

for technological development.

Previous studies showed that compounds secreted into

the biofilm matrix can readily be isolated from lawns of

bacteria cultured on agar (Bendaoud et al. 2011; Kar-

wacki et al. 2013). Bacterial lawns, sometimes referred to

as colony biofilms (Merritt et al. 2005), exhibit many

properties characteristic of biofilms cultured in broth,

including high cell density, extracellular matrix, spatially

dependent growth, chemical gradients and increased anti-

biotic resistance (Auerbach et al. 2000; Dietrich et al.

2008; Kim et al. 2009). Cell-free extracts isolated from

lawns of bacteria, or from biofilms cultured in a continu-

ous-flow fermentor that produces a similarly high

amount of biofilm biomass, have been shown to be

enriched for soluble molecules produced within the bio-

film matrix (Bendaoud et al. 2011; Rendueles et al. 2011,

2013).

In the present study, we screened a panel of 126 cell-

free extracts prepared from environmental bacteria iso-

lated from soil, cave and river habitats. Using a 96-well

microtitre plate assay, we measured the ability of each

extract to inhibit biofilm formation by Staphylococcus

aureus, an important human pathogen. We found that

nearly half of the extracts inhibited Staph. aureus biofilm,

most without inhibiting growth. Here we present evi-

dence that these extracts contain a variety of chemical

compounds that inhibit biofilm by a variety of mecha-

nisms.

Materials and methods

Isolation and characterization of environmental bacteria

A total of 126 bacterial strains were isolated from cave,

river and soil samples collected at four locations in and

around Washington, DC (Table 1). Thirty-four strains

were isolated from mud and pool sediment collected in

Rogers Belmont Cave, Warren County, Virginia. Samples

were collected in the dark zone of the cave. Thirty-four

strains were isolated from water samples taken from the

Anacostia River in Washington, DC, and the Chester

River in Queen Anne’s County, Maryland. River samples

were collected from sites near run-off or drainage points,

bridge run-off points and sewage discharge points. In

addition, 58 strains were isolated from soil samples col-

lected at 16 different sites on the American university

campus in Washington, DC. River samples were collected

in 2006, and other samples were collected in 2013.

Mud, sediment and soil samples were resuspended in

sterile water, disrupted by vortex agitation for 30 s and

then diluted and plated on LB agar. Plates were incubated

at room temperature for 1–3 days. Individual colonies

were streaked to purity and stored at �80°C in 8% DMSO.

Strains that exhibited diverse colony morphologies on agar

were selected to maximize the number of different taxa in

the screen. River water samples were diluted and plated on

LB agar supplemented with 10 mg l�1 tetracycline, and

then incubated at 28°C for 4 days. Individual colonies were

streaked to purity and stored at �80°C.All strains were Gram-stained and analysed by bright-

field oil immersion microscopy to confirm a bacterial cell

morphology. A total of 29 strains were further analysed

by PCR amplification and partial DNA sequence analysis

of the 16S rRNA gene using PCR primers 27F and 534R

(Muyzer et al. 1993). Genera were assigned using the

RDP Classifier program (Wang et al. 2007).

Preparation of cell-free extracts

A loopful of cells from a fresh agar plate was resuspended

in 100 ll of LB broth and spread onto the surface of a

100-mm-diameter LB agar plate using a sterile glass sprea-

der. The plate was incubated at room temperature for 48–96 h until a robust lawn of microbial growth developed.

The cell paste was then scraped from the surface of the

agar using a plastic inoculating loop, and the cells were

transferred to a microcentrifuge tube containing 750 ll ofphosphate-buffered saline (PBS). The tube was mixed by

vortex agitation for 10 min, and the cells were then pel-

leted by centrifugation. The supernatant was sterilized by

passage through a 0�22-lm pore size centrifugal filter. The

resulting cell-free extracts were stored at 4°C.

Biofilm assay

The biofilm-forming test strains used in this study were

Staph. aureus JE2 (Fey et al. 2013), Staph. aureus SH1000

(Horsburgh et al. 2002), Staphylococcus epidermidis

RP62A (ATCC 35984; American Type Culture Collection,

Manassas, VA) and Pseudomonas fluorescens WCS365

(O’Toole and Kolter 1998). Staphylococci were cultured

at 37°C in (TSB) supplemented with 6 g l�1 yeast extract

and 8 g l�1 glucose. Staphylococcus aureus JE2 cultures

were further supplemented with 0�2 mg l�1 amoxicillin

to induce biofilm formation (Kaplan et al. 2012). Pseudo-

monas fluorescens was cultured at 30°C in LB broth. Inoc-

ula were prepared from 18-h-old agar colonies as

previously described (Izano et al. 2008). A volume of

180 ll of inoculum (c. 104–105 CFU ml�1) was trans-

ferred to the well of a tissue-culture-treated polystyrene

microtitre plate (Falcon no. 353047). A total of 20 ll of

Journal of Applied Microbiology 117, 1663--1673 © 2014 The Society for Applied Microbiology1664

Screen for antibiofilm compounds J.T. Farmer et al.

Table 1 Bacterial strains

Source Strain† Genus‡ DNase activity§

Per cent biofilm inhibition¶

Staphylococcus aureus JE2 Staphylococcus aureus SH1000

Cave C101 – – 30 � 3 –

C102 – – 0 –

C103 – – 0 –

C104 Bacillus No 92 � 2* 71 � 6*

C105 – – 17 � 7 –

C106 – – 22 � 1 –

C107 – – 0 –

C108 – – 0 –

C109 – – 0 –

C201 Bacillus – 95 � 5* 61 � 18

C202 – – 0 –

C203 – – 32 � 12 –

C204 – – 0 –

C205 – – 0 –

C206 – – 23 � 1 –

C207 – – 15 � 2 –

C208 – – 0 –

P101 – – 45 � 12 –

P102 – – 0 –

P103 – – 0 –

P104 Bacillus – 90 � 5* 0

P105 Bacillus – 90 � 3* 90 � 4*

P106 – – 54 � 14 –

P107 – – 0 –

P108 – – 0 –

P201 – – 42 � 16 –

P202 – – 0 –

P203 – – 0 –

P204 Bacillus Yes 67 � 5* 0

P205 – – 0 –

P206 Bacillus – 85 � 4* 17 � 6

P207 – – 65 � 6* –

P208 – – 0 –

P209 Rhodococcus – 89 � 5* 0

River AR17 – – 61 � 6* –

AR18 Serratia No 84 � 4* 94 � 5*

AR19 – – 75 � 5* –

AR20 – – 75 � 5* –

AR21 – – 36 � 10 –

AR22 – No 90 � 3* 75 � 0*

AR23 – – 35 � 4 –

AR24 – – 75 � 3* –

AR25 Serratia – 82 � 4* 91 � 5*

AR26 – – 68 � 11* –

AR27 – – 69 � 11* –

AR28 Serratia – 86 � 3* 83 � 3*

AR29 – – 45 � 6 –

AR30 Enterobacter – 86 � 6* 92 � 3*

AR31 – – 43 � 0 –

AR32 Enterobacter – 85 � 5* 0

AR33 – – 93 � 0* 96 � 1*

AR34 Serratia Yes 88 � 3* 93 � 4*

(Continued)


J.T. Farmer et al. Screen for antibiofilm compounds

Table 1 (Continued )




CR1 – – 71 � 5* –

CR2 – – 68 � 5* –

CR3 – – 19 � 10 –

CR4 – – 75 � 0* –

CR5 – – 21 � 3 –

CR6 Serratia Yes 90 � 5* 95 � 0*

CR7 – – 84 � 5* 96 � 1*

CR8 – – 55 � 5* –

CR9 – – 46 � 5 –

CR10 – – 39 � 8 –

CR11 – – 50 � 10 –

CR12 Serratia No 85 � 5* 95 � 1*

CR13 – – 31 � 10 –

CR14 Bacillus – 92 � 2* 85 � 9*

CR15 Serratia No 76 � 4* 94 � 2*

CR16 Serratia Yes 72 � 2* 95 � 1*

Soil AB1 – Yes 72 � 10* –

AB2 – – 96 � 0* –

AM1 – Yes 96 � 0* –

CB2 – Yes 96 � 0* –

CB3 – – 0 –

FA1 – No 0 –

FA2 – – 40 � 14 –

GF1 Bacillus – 67 � 7* 0

JF1 – – 23 � 2 –

JF2 – – 0 –

JK1 – No 96 � 0* –

JK2 – – 0 –

JS1 – – 54 � 9 –

JS2 – Yes 96 � 0* –

KA1 – – 23 � 1 –

KS1 – – 0 –

KS2 – – 0 –

MC1 – No 97 � 0* –

MC2 – – 0 –

MC5 – – 50 � 6 –

MC6 – – 0 –

MC7 – – 0 –

MC8 Stenotrophomonas No 95 � 0* 95 � 1*

MC9 – No 0 –

MC10 – – 0 –

MC11 – – 0 –

MC12 Pseudomonas No 94 � 0* 48 � 4

MC14 Stenotrophomonas No 96 � 0* 79 � 9*

PR1 – – 60 � 5* –

PR2 – Yes 94 � 0* –

SG1 – No 0 –

SG2 – – 10 � 2 –

SG3 – No 95 � 0* –

SG4 – – 32 � 5 –

SG5 Bacillus – 96 � 0* 95 � 2*

SP1 – – 30 � 6 –

SP2 – Yes 90 � 2* –

(Continued)



cell-free extract, or 20 ll of PBS as a control, was mixed

with the inoculum, and the plate was incubated statically

at 37°C (or 30°C for Ps. fluorescens) for 18 h. To measure

bacterial growth, the absorbance of the broth was mea-

sured in a microplate reader set to 450 nm. Control wells

containing sterile broth were used to measure back-

ground absorbance values. To measure biofilm formation,

wells were rinsed with water and stained for 1 min with

200 ll of Gram’s crystal violet. Wells were then rinsed

with water and dried. The amount of crystal violet bind-

ing was quantified by destaining the wells for 10 min

with 200 ll of 33% acetic acid, and then measuring the

absorbance of the crystal violet solution in a microplate

reader set to 595 or 620 nm.

Physical and chemical analyses of cell-free extracts

DNase activity was measured using DNase test agar, or

by incubating 1 ll of cell-free extract with 1 lg of MspI-

digested pBR322 DNA (New England BioLabs, Ipswich,

MA) in 10 mmol l�1 Tris-HCl (pH 7�5), 10 mmol l�1

MgCl2, and then assessing DNA degradation by agarose

gel electrophoresis. Size-exclusion filtration was carried

out using Microcon centrifugal concentrators (Millipore,

Darmstadt, Germany) with 10-kDa-molecular-weight cut-

off filters. Filtrates were tested in the biofilm assay, along

with crude extract as a positive control and PBS as a neg-

ative control. Heat stability was measured by incubating

cell-free extracts at 100°C for 15 min and cooling to

room temperature prior to adding them to the micro-

plate wells. Unheated cell-free extracts served as positive

controls and PBS as a negative control. Chemical polarity

was measured by adding an equal volume of hexane to

the extract, mixing by vortex agitation and separating the

phases by brief centrifugation. The aqueous phase was

then tested in the biofilm assay, along with nonextracted

cell-free extract as a positive control, and PBS and hex-

ane-extracted PBS as negative controls.

Statistics and reproducibility of results

All microtitre plate assays were performed in duplicate or

triplicate wells, which exhibited an average variation of

<10%. The significance of differences between mean

Table 1 (Continued )




SS2 Janthinobacterium No 96 � 0* –

SS4 – – 0 –

SS3 – – 0 –

SS5 Pseudomonas No 79 � 6* 49 � 13

SS6 Flavobacterium No 71 � 13 0

SS7 – – 0 –

SS8 – – 0 –

TA1 Pseudomonas Yes 85 � 3* 70 � 19

TA2 – – 36 � 2 –

TA3 – – 45 � 7 –

TA4 – – 16 � 2 –

TA5 – – 0 –

TA7 – – 19 � 9 –

TA9 – – 13 � 2 –

TA10 – – 0 –

TA11 – Yes 88 � 1* –

TA12 – – 80 � 2* –

TA13 Pedobacter – 30 � 3 0

TA14 Bacillus No 96 � 0* 85 � 2*

TA15 – – 54 � 7 –

TA16 – No 96 � 0* –

†AR, Anacostia River; CR, Chester River.

‡Based on 16S rRNA sequence. –, not tested.

§–, not tested.

¶Values show mean per cent biofilm inhibition (�SD) from 2 to 5 experiments. Per cent biofilm inhibition was calculated as 1 � (A[10% cell-free

extract]/A[10% PBS]) 9 100. Asterisks indicate values significantly different from phosphate-buffered saline (PBS) control (P < 0�05). 0, per cent bio-film inhibition <10; –, not tested.



absorbance values was calculated using a two-tailed Stu-

dent’s t-test. A P value of <0�05 was considered signifi-

cant. All assays were performed 2–5 times with similarly

significant differences in absorbance values.

Results

Cell-free extracts of environmental bacteria inhibit

Staphylococcus aureus biofilm

We isolated 126 bacterial strains from river, cave and soil

habitats in and around Washington, DC (Table 1). Gram

stain analyses revealed rods and cocci, some linked in

chains or clusters, 1–5 lm in size, which is indicative of

Bacteria. Several strains isolated from soil produced en-

dospores. Approximately 40% of the strains stained Gram

positive.

Cell-free extracts prepared from all 126 strains were

screened for their ability to inhibit biofilm formation by

methicillin-resistant Staph. aureus (MRSA) strain JE2 in a

96-well microtitre plate assay (Table 1). A total of 55/126

extracts (44%) significantly inhibited JE2 biofilm, includ-

ing 8 of 34 cave extracts (24%), 23 of 58 soil extracts

(40%) and 24 of 34 river extracts (71%).

Mechanisms of action

More than 90% of the 126 cell-free extracts that we

screened had no significant effect on Staph. aureus JE2

growth, and several extracts actually increased growth

(data not shown). Four extracts significantly inhibited

JE2 growth. These included extracts CR12 and SS2, which

also inhibited growth of methicillin-sensitive

Staph. aureus (MSSA) strain SH1000 (Fig. 1). Extracts

CR6 and CR14 also inhibited growth of strains JE2 and

SH1000 (data not shown). The antimicrobial activity in

these extracts probably accounts for their biofilm-inhibit-

ing activity.

As Staph. aureus biofilm formation is inhibited by

DNase (Izano et al. 2008), we tested 31 extracts that

inhibited Staph. aureus JE2 biofilm for DNase activity

(Table 1). These 31 extracts were selected because they

were obtainable in high yields and they exhibited stable

antibiofilm activity. A total of 12/31 extracts (39%)

exhibited DNase activity. DNase may account for the

observed antibiofilm activity in these extracts. Three

extracts that did not inhibit Staph. aureus JE2 biofilm

(FA1, MC9 and SG1) did not exhibit DNase activity

(Table 1).

SS2 extract CR12 extract

00 2 4 6 8 10

0 2 4 6 8 10

0 2 4 6 8 10

0 2 4 6 8 10

0·1

0·2

0·3

0·4

0·5

0·6

0

0·1

0·2

0·3

0·4

0·5

0·6

0·7

0

0·1

0·2

0·3

0·4

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0·6

0·7

0

0·1

0·2

0·3

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0·5

0·6

0·7

0·8

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(450

nm

)A

bsor

banc

e (4

50 n

m)

Abs

orba

nce

(450

nm

)

Time (h) Time (h)

Time (h)Time (h)

Str

ain

JE2

Str

ain

SH

1000

Figure 1 Growth of Staphylococcus aureus

strain JE2 (top graphs) and strain SH1000

(bottom graphs) in the presence of cell-free

extracts prepared from environmental

bacteria SS2 (left graphs) and CR12 (right

graphs). Open circles show growth in the

presence of 10% (by vol) of the indicated

extract. Filled circles show growth in the

presence of 10% phosphate-buffered saline

(PBS) as a control. Values show means from

duplicate wells. Error bars were omitted for

clarity.



We also tested 31 cell-free extracts that inhibited

Staph. aureus JE2 biofilm for their ability to inhibit

Staph. aureus SH1000 biofilm (Table 1). Both JE2 and

SH1000 produce biofilms dependent on proteinaceous

adhesins and extracellular DNA, but JE2 forms biofilm

only in the presence of low-level b-lactam antibiotics

(Kaplan et al. 2012). A total of 25/31 extracts (81%) sig-

nificantly inhibited biofilm by both JE2 and SH1000.

However, six extracts (P104, P204, P209, AR32, GF1 and

SS6) inhibited biofilm by JE2, but not by SH1000. Fig-

ure 2 shows growth and biofilm of JE2 and SH1000 in

the presence of increasing concentrations of the JE2-

inhibitory extract P104.

More than two-thirds of the cell-free extracts that

inhibited both JE2 and SH1000 biofilm exhibited biofilm

inhibition for up to 10 h with little or no effect on bacte-

rial growth. Examples of extracts that exhibited this pat-

tern of antibiofilm activity are shown in Fig. 3.

We selected seven cell-free extracts for further analysis

(Table 2). These seven extracts were chosen because they

significantly inhibited biofilm by both Staph. aureus JE2

and SH1000 without inhibiting growth, and their antibio-

film activities were potent and stable. Two of these extracts

(CR16 and TA1) exhibited DNase activity (Table 1).

The spectrum of activity of the seven selected extracts

was investigated by measuring their ability to inhibit

biofilm formation by Staph. epidermidis strain RP62A

and Ps. fluorescens strain WSC365 (Table 2). The

Staph. epidermidis biofilm matrix is composed primarily

of polysaccharides (Kaplan et al. 2004), whereas the

Ps. fluorescens matrix contains both polysaccharides and

proteinaceous adhesins (O’Toole and Kolter 1998; Itoh

et al. 2005). Two extracts (AR18 and TA1) significantly

inhibited biofilm by Staph. epidermidis, and four extracts

(AR18, C104, MC8 and SS5) significantly inhibited bio-

film by Ps. fluorescens (Table 2). Only one extract (AR18)

significantly inhibited biofilm by both species. Extract

TA1 inhibited Ps. fluorescens growth, but not Staph. epi-

dermidis growth, while none of the other six extracts

inhibited Staph. epidermidis or Ps. fluorescens growth

(data not shown).

Physical and chemical properties

To investigate the physical and chemical nature of the

biofilm-inhibiting activities in the seven selected extracts,

we measured their heat stability, molecular mass and

chemical polarity (Table 2). The antibiofilm activities in

two extracts (MC8 and MC14) were heat labile, the anti-

biofilm activities in three extracts (AR18, CR16 and

MC8) were <10 kDa in mass, and the antibiofilm activi-

ties in two extracts (CR16 and TA1) were abolished after

Growth Biofilm0·8

0·6

0·4

0·2

0

0·8

0·6

0·4

0·2

0

0·8

0·6

0·4

0·2

0

0 2 6 10

0 2 6 10 00

2

2

6 10

0 2 6 10

4

3

1

1

*

* *

Extract concentration (%) Extract concentration (%)

Abs

orba

nce

(450

nm

)A

bsor

banc

e (4

50 n

m)

Abs

orba

nce

(620

nm

)A

bsor

banc

e (6

20 n

m)

Extract concentration (%)Extract concentration (%)

Str

ain

SH

1000

Str

ain

JE2

Figure 2 Growth (left graphs) and biofilm

(right graphs) of Staphylococcus aureus strain

JE2 (top graphs) and strain SH1000 (bottom

graphs) in the presence of 0, 2, 6 and 10%

(by vol) of cell-free extract P204. Cultures

were incubated for 18 h. Values show means

from duplicate wells. Error bars indicate

range. Asterisks denote values significantly

different from no extract control (P < 0�05).



hexane extraction, indicating that nonpolar compounds

are active components in these extracts (Table 2). Taken

together, these findings suggest that the biofilm-inhibiting

activities in the seven selected extracts are mediated by

unique chemical compounds.

Discussion

In the present study, we screened 126 cell-free extracts

isolated from soil, cave and river bacteria for their ability

to inhibit Staph. aureus biofilm in a 96-well microtitre

plate assay. We found that 55/126 extracts (44%) signifi-

cantly inhibited Staph. aureus biofilm. Results from stud-

ies on the spectrum of activity and physicochemical

properties of these extracts suggested that biofilm inhibi-

tion was mediated by a variety of chemical compounds

that inhibited biofilm by a variety of mechanisms.

Bacteria that produced cell-free extracts with antibiofilm

activity included members of the Firmicutes (Bacillus),

Actinobacteria (Rhodococcus), Bacteroidetes (Flavobacteri-

um, Pedobacter) and Proteobacteria (Serratia, Pseudomo-

nas, Enterobacter, Janthinobacterium, Stenotrophomonas).

Growth Growth

Growth Growth

Biofilm Biofilm

Biofilm Biofilm

0·80·70·60·50·40·30·20·1

0

0·80·70·60·50·40·30·20·1

0

0·80·70·60·50·40·30·20·1

0

0·80·70·60·50·40·30·20·1

0

0·70·60·50·40·30·20·1

00 2 4 6 8

0 2 4 6 8 0 2 4 6 8

10 0 2 4 6 8 10

0 2 4 6 8 10 0 2 4 6 8 10

0 2 4 6 8 10 0 2 4 6 8 10

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(450

nm

)

Abs

orba

nce

(620

nm

)

Abs

orba

nce

(595

nm

)

Time (h) Time (h) Time (h) Time (h)

Time (h)Time (h)Time (h)Time (h)

0

1

2

3

4

0

1

2

3

4

0

1

2

3

4

(a) (b)

(d)(c)

Figure 3 Growth (left panels) and biofilm (right panels) of Staphylococcus aureus strain SH1000 in the presence of cell-free extracts isolated from

environmental bacteria AR34 (a), CR7 (b), CR15 (c) and C104 (d). Open circles show growth and biofilm in the presence of 10% (by vol) of the

indicated extract. Filled circles show growth and biofilm in the presence of 10% phosphate-buffered saline (PBS) as a control. Values show means

from duplicate wells. Error bars were omitted for clarity.

Table 2 Spectrum of activity and physicochemical properties of cell-free extracts

Cell-free extract

Spectrum of activity* Physicochemical properties†

Staphylococcus

epidermidis

RP62A

Pseudomonas

fluorescens

WCS365

100°C,

15 min

10-kDa

filter

Hexane

extraction

Serratia AR18 + + + + +

Bacillus C104 + + +

Serratia CR16 + +

Stenotrophomonas MC8 + + +

Stenotrophomonas MC14 +

Pseudomonas SS5 + + +

Pseudomonas TA1 + +

*+, cell-free extract exhibited significant biofilm-inhibiting activity against the indicated strain.

†+, cell-free extract exhibited significant biofilm-inhibiting activity against Staphylococcus aureus SH1000 after the indicated treatment.



Bacillus spp. dominated cave samples (6/7 strains) and was

the only genus found in three habitats (soil, cave, Chester

River). As expected, soil samples contained several mem-

bers of the common soil genera Bacillus and Pseudomonas.

The river bacteria that we analysed comprised a conve-

nience sample isolated during a 2006 study on bacterial

communities in river systems. These strains had been

selected on LB agar supplemented with 10 mg l�1 tetracy-

cline. Serratia spp. were the most common bacteria isolated

from river water, and tetracycline resistance has been

reported to be widespread among Serratia (Carbonell et al.

2000; Stock et al. 2003; Thompson et al. 2007). A total

of 5/8 Serratia strains were nonpigmented, which is a

common phenotype of human clinical strains (Hejazi and

Falkiner 1997). This suggests that these strains may be of

human origin.

Four of the 126 cell-free extracts (3%) significantly

inhibited Staph. aureus growth. These include extracts

isolated from Serratia spp. (CR12 and CR6), Janthino-

bacterium sp. (SS2) and Bacillus sp. (CR14). All three

of these genera have been shown to produce antimicro-

bial compounds that are active against Staph. aureus

(O’Sullivan et al. 1990; Romero-Tabarez et al. 2006; Ka-

douri and Shanks 2013). These findings suggest that

screening cell-free extracts may be a useful method for

isolating antimicrobial compounds from environmental

bacteria.

About 40% of the cell-free extracts that we tested had

detectable DNase activity, and all extracts that exhibited

DNase activity inhibited biofilm by both Staph. aureus

strains JE2 and SH1000 (Table 1). As Staph. aureus JE2

and SH1000 biofilms have been shown to be inhibited by

DNase (Izano et al. 2008; Kaplan et al. 2012), these find-

ings suggest that the antibiofilm activity in these extracts

is mediated by DNase. However, 19/31 extracts that did

not exhibit detectable DNase activity also inhibited

Staph. aureus JE2 biofilm, and 11/12 that were tested also

inhibited SH1000 biofilm (Table 1). Therefore, com-

pounds other than DNase probably contribute to the an-

tibiofilm activity in these extracts.

One interesting pattern of antibiofilm activity that we

observed was exhibited by extracts of Bacillus spp. P104,

P204 and GF1; Rhodococcus sp. P209; Enterobacter sp.

AR32; and Flavobacterium sp. SS6. These six extracts

inhibited biofilm by Staph. aureus JE2, but not by

Staph. aureus SH1000. As strain JE2 requires low-level b-lactam antibiotics to form biofilm, these six extracts may

contain b-lactamases or compounds that inhibit signal-

ling pathways regulating Staph. aureus antibiotic-induced

biofilm (Kaplan 2011).

Among the seven cell-free extracts that we tested

against Staph. epidermidis and Ps. fluorescens biofilms

(Table 2), one inhibited biofilm by both species (AR18),

three inhibited only Ps. fluorescens biofilm (C104, MC8

and SS5), one inhibited only Staph. epidermidis biofilm

(TA1) and two inhibited neither biofilm (CR16 and

MC14). These unique patterns of biofilm inhibition

against bacteria with different mechanisms of biofilm for-

mation support the notion that biofilm inhibition is

mediated by agents with multiple mechanisms of action.

In summary, a screen of 126 diverse cell-free extracts

yielded 55 extracts that exhibited antibiofilm activity

against MRSA in a microplate assay. Among these, we

found that 12 contained DNase, six may contain b-lac-tamase or compounds that inhibit b-lactam-induced bio-

film, five contained active compounds that were

thermostable, four contained antimicrobial compounds,

three contained active compounds <10 kDa in mass, two

contained hydrophobic active compounds and one exhib-

ited broadspectrum antibiofilm activity against

Staph. aureus, Staph. epidermidis and Ps. fluorescens.

Although further experiments are needed to characterize

the structures and mechanisms of action of these com-

pounds, our preliminary findings suggest that environ-

mental bacteria constitute an untapped source of natural

bioactive molecules antagonizing adhesion or biofilm

formation of other bacteria.

Acknowledgements

We thank the students in BIO-440 (Keide Akinola, Fariha

Alam, Cassandra Baker, Anne Ballard, Alexis Dobbs,

George Fountain, Sarah Goodheart, Benjamin Kussin-

Shoptaw, Alfred Mabika, Scott Piraino, Anusha Pundu,

Priscila Riccardi, Swati Samtani and Julie Shelton) for

isolating bacteria from American University soil; Daniel

W. Fong for providing cave samples; and Sophie West,

Don Campbell, James McRedmond, Kristen Bert and

Lauren Jesnig for technical assistance. This study was

funded by NIH Grant AI097182 and by American

University.

Conflict of Interest

The authors have no conflicts of interest.

References

Auerbach, I.D., Sorensen, C., Hansma, H.G. and Holden, P.A.

(2000) Physical morphology and surface properties of

unsaturated Pseudomonas putida biofilms. J Bacteriol 182,

3809–3815.

Bendaoud, M., Vinogradov, E., Balashova, N.V., Kadouri,

D.E., Kachlany, S.C. and Kaplan, J.B. (2011) Broad-

spectrum biofilm inhibition by Kingella kingae

exopolysaccharide. J Bacteriol 193, 3879–3886.



Blackledge, M.S., Worthington, R.J. and Melander, C. (2013)

Biologically inspired strategies for combating bacterial

biofilms. Curr Opin Pharmacol 13, 699–706.

Carbonell, G.V., Della Colleta, H.H., Yano, T., Darini, A.L.,

Levy, C.E. and Fonseca, B.A. (2000) Clinical relevance and

virulence factors of pigmented Serratia marcescens. FEMS

Immunol Med Microbiol 28, 143–149.

Dietrich, L.E., Teal, T.K., Price-Whelan, A. and Newman, D.K.

(2008) Redox-active antibiotics control gene expression

and community behavior in divergent bacteria. Science

321, 1203–1206.

Fey, P.D., Endres, J.L., Yajjala, V.K., Widhelm, T.J., Boissy,

R.J., Bose, J.L. and Bayles, K.W. (2013) A genetic resource

for rapid and comprehensive phenotype screening of

nonessential Staphylococcus aureus genes. mBio 4, e00537–

12.

Flemming, H.C. and Wingender, J. (2010) The biofilm matrix.

Nat Rev Microbiol 8, 623–633.

Flemming, H.C., Meier, M. and Schild, T. (2013) Mini-review:

microbial problems in paper production. Biofouling 29,

683–696.

Hall-Stoodley, L., Costerton, J.W. and Stoodley, P. (2004)

Bacterial biofilms: from the natural environment to

infectious diseases. Nat Rev Microbiol 2, 95–108.

Hejazi, A. and Falkiner, F.R. (1997) Serratia marcescens. J Med

Microbiol 46, 903–912.

Horsburgh, M.J., Aish, J.L., White, I.J., Shaw, L., Lithgow, J.K.

and Foster, S.J. (2002) rB modulates virulence

determinant expression and stress resistance:

characterization of a functional rsbU strain derived from

Staphylococcus aureus 8325-4. J Bacteriol 184, 5457–5467.

Itoh, Y., Wang, X., Hinnebusch, B.J., Preston, J.F. III and

Romeo, T. (2005) Depolymerization of beta-1,6-N-acetyl-

D-glucosamine disrupts the integrity of diverse bacterial

biofilms. J Bacteriol 187, 382–387.

Izano, E.A., Amarante, M.A., Kher, W.B. and Kaplan, J.B.

(2008) Differential roles of poly-N-acetylglucosamine

surface polysaccharide and extracellular DNA in

Staphylococcus aureus and Staphylococcus epidermidis

biofilms. Appl Environ Microbiol 74, 470–476.

Jiang, P., Li, J., Han, F., Duan, G., Lu, X., Gu, Y. and Yu,

W. (2011) Antibiofilm activity of an exopolysaccharide

from marine bacterium Vibrio sp. QY101. PLoS One 6,

e18514.

Kadouri, D.E. and Shanks, R.M. (2013) Identification of a

methicillin-resistant Staphylococcus aureus inhibitory

compound isolated from Serratia marcescens. Res Microbiol

164, 821–826.

Kaplan, J.B. (2010) Biofilm dispersal: mechanisms, clinical

implications, and potential therapeutic uses. J Dent Res 89,

205–218.

Kaplan, J.B. (2011) Antibiotic-induced biofilm formation. Int J

Artif Organs 34, 737–751.

Kaplan, J.B., Ragunath, C., Ramasubbu, N. and Fine, D.H.

(2003) Detachment of Actinobacillus

actinomycetemcomitans biofilm cells by an endogenous

beta-hexosaminidase activity. J Bacteriol 185, 4693–4698.

Kaplan, J.B., Ragunath, C., Velliyagounder, K., Fine, D.H. and

Ramasubbu, N. (2004) Enzymatic detachment of

Staphylococcus epidermidis biofilms. Antimicrob Agents

Chemother 48, 2633–2636.

Kaplan, J.B., Izano, E.A., Gopal, P., Karwacki, M.T., Kim, S.,

Bose, J.L., Bayles, K.W. and Horswill, A.R. (2012) Low

levels of b-lactam antibiotics induce extracellular DNA

release and biofilm formation in Staphylococcus aureus.

mBio 3, e00198–12.

Karatan, E. and Watnick, P. (2009) Signals, regulatory

networks, and materials that build and break bacterial

biofilms. Microbiol Mol Biol Rev 73, 310–347.

Karwacki, M.T., Kadouri, D.E., Bendaoud, M., Izano, E.A.,

Sampathkumar, V., Inzana, T.J. and Kaplan, J.B. (2013)

Antibiofilm activity of Actinobacillus pleuropneumoniae

serotype 5 capsular polysaccharide. PLoS One 8, e63844.

Kim, J., Park, H.J., Lee, J.H., Hahn, J.S., Gu, M.B. and Yoon,

J. (2009) Differential effect of chlorine on the oxidative

stress generation in dormant and active cells within colony

biofilm. Water Res 43, 5252–5259.

Kostakioti, M., Hadjifrangiskou, M. and Hultgren, S.J. (2013)

Bacterial biofilms: development, dispersal, and therapeutic

strategies in the dawn of the postantibiotic era. Cold

Spring Harb Perspect Med 3, a010306.

Kumar, C.G. and Anand, S.K. (1998) Significance of microbial

biofilms in food industry: a review. Int J Food Microbiol

42, 9–27.

Merritt, J.H., Kadouri, D.E. and O’Toole, G.A. (2005)

Growing and analyzing static biofilms. Curr Protoc

Microbiol Chapter 1, Unit 1B 1.

Muyzer, G., de Waal, E.C. and Uitterlinden, A.G. (1993)

Profiling of complex microbial populations by denaturing

gradient gel electrophoresis analysis of polymerase chain

reaction-amplified genes coding for 16S rRNA. Appl

Environ Microbiol 59, 695–700.

O’Sullivan, J., McCullough, J., Johnson, J.H., Bonner, D.P.,

Clark, J.C., Dean, L. and Trejo, W.H. (1990) Janthinocins

A, B and C, novel peptide lactone antibiotics produced by

Janthinobacterium lividum. I. Taxonomy, fermentation,

isolation, physico-chemical and biological characterization.

J Antibiot (Tokyo) 43, 913–919.

O’Toole, G.A. and Kolter, R. (1998) Initiation of biofilm

formation in Pseudomonas fluorescens WCS365 proceeds

via multiple, convergent signalling pathways: a genetic

analysis. Mol Microbiol 28, 449–461.

Parsek, M.R. and Singh, P.K. (2003) Bacterial biofilms: an

emerging link to disease pathogenesis. Annu Rev Microbiol

57, 677–701.

Rendueles, O., Traviera, L., Latour-Lamberta, P., Fontainec, T.,

Magnusa, J., Denamurd, E. and Ghigo, J.-M. (2011)

Screening of Escherichia coli species biodiversity reveals

new biofilm-associated antiadhesion polysaccharides. mBio

2, e00043–11.



Rendueles, O., Kaplan, J.B. and Ghigo, J.-M. (2013)

Antibiofilm polysaccharides. Environ Microbiol 15, 334–

346.

Romero-Tabarez, M., Jansen, R., Sylla, M., L€unsdorf, H.,

H€aussler, S., Santosa, D.A., Timmis, K.N. and Molinari, G.

(2006) 7-O-malonyl macrolactin A, a new macrolactin

antibiotic from Bacillus subtilis active against

methicillin-resistant Staphylococcus aureus, vancomycin-

resistant enterococci, and a small-colony variant of

Burkholderia cepacia. Antimicrob Agents Chemother 50,

1701–1709.

Solano, C., Echeverz, M. and Lasa, I. (2014) Biofilm dispersion

and quorum sensing. Curr Opin Microbiol 18C, 96–104.

Stock, I., Grueger, T. and Wiedemann, B. (2003) Natural

antibiotic susceptibility of strains of Serratia marcescens

and the S. liquefaciens complex: S. liquefaciens sensu

stricto, S. proteamaculans and S. grimesii. Int J Antimicrob

Agents 22, 35–47.

Thompson, S.A., Maani, E.V., Lindell, A.H., King, C.J. and

McArthur, J.V. (2007) Novel tetracycline resistance

determinant isolated from an environmental strain of

Serratia marcescens. Appl Environ Microbiol 73, 2199–2206.

Valle, J., Da Re, S., Henry, N., Fontaine, T., Balestrino, D.,

Latour-Lambert, P. and Ghigo, J.-M. (2006)

Broadspectrum biofilm inhibition by a secreted bacterial

polysaccharide. Proc Natl Acad Sci USA 103, 12558–12563.

Wang, Q., Garrity, G.M., Tiedje, J.M. and Cole, J.R. (2007)

Na€ıve Bayesian classifier for rapid assignment of rRNA

sequences into the new bacterial taxonomy. Appl Environ

Microbiol 73, 5261–5267.

Worthington, R.J., Richards, J.J. and Melander, C. (2012)

Small molecule control of bacterial biofilms. Org Biomol

Chem 10, 7457–7474.

Yang, L., Liu, Y., Wu, H., Song, Z., Høiby, N., Molin, S. and

Givskov, M. (2012) Combating biofilms. FEMS Immunol

Med Microbiol 65, 146–157.



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Environmental bacteria produce abundant and diverse antibiofilm compounds