- Research article
- Open Access
Marine bacteria from the French Atlantic coast displaying high forming-biofilm abilities and different biofilm 3D architectures
- Ibtissem Doghri1,
- Sophie Rodrigues2,
- Alexis Bazire2,
- Alain Dufour2,
- David Akbar1,
- Valérie Sopena1,
- Sophie Sablé†1Email author and
- Isabelle Lanneluc†1
© Doghri et al. 2015
Received: 24 March 2015
Accepted: 15 October 2015
Published: 24 October 2015
Few studies have reported the species composition of bacterial communities in marine biofilms formed on natural or on man-made existing structures. In particular, the roles and surface specificities of primary colonizers are largely unknown for most surface types. The aim of this study was to obtain potentially pioneering bacterial strains with high forming-biofilm abilities from two kinds of marine biofilms, collected from two different surfaces of the French Atlantic coast: an intertidal mudflat which plays a central role in aquaculture and a carbon steel structure of a harbour, where biofilms may cause important damages.
A collection of 156 marine heterotrophic aerobic bacteria isolated from both biofilms was screened for their ability to form biofilms on polystyrene 96-well microtiter plates. Out of 25 strains able to build a biofilm in these conditions, only four bacteria also formed a thick and stable biofilm on glass surfaces under dynamic conditions. These strains developed biofilms with four different three - dimensional architectures when observed by confocal laser scanning microscopy: Flavobacterium sp. II2003 biofilms harboured mushroom-like structures, Roseobacter sp. IV3009 biofilms were quite homogeneous, Shewanella sp. IV3014 displayed hairy biofilms with horizontal fibres, whereas Roseovarius sp. VA014 developed heterogeneous and tousled biofilms.
This work led for the first time to the obtaining of four marine bacterial strains, potentially pioneering bacteria in marine biofilms, able to adhere to at least two different surfaces (polystyrene and glass) and to build specific 3D biofilms. The four selected strains are appropriate models for a better understanding of the colonization of a surface as well as the interactions that can occur between bacteria in a marine biofilm, which are crucial events for the initiation of biofouling.
Biofilms are generally considered as surface-associated microorganism communities encased in a hydrated polymeric matrix . In marine environment, most of the solid man-made structures as well as natural surfaces are covered by microbial biofilms. Together with diatoms, bacteria constitute the major components of biofilms occurring in the marine environment . Furthermore, all types of bacteria can form biofilms, making this sedentary lifestyle their favourite mode of existence in nature . This mechanism is described as considerably important for survival of marine bacteria by providing a favourable environment . This lifestyle, compared to the planktonic one, indeed improves access to nutrients and protects against stress, antibiotics and predators [1, 5].
Quickly after immersion of a clean surface in the sea, microorganisms colonize it and subsequently develop biofilms with highly diverse three-dimensional (3D) structures which can include channels allowing the flow of liquids, nutrients and wastes [6–8]. The early stages of biofilm formation are based on the interactions of free-living bacteria with the surface which generates an initial layer of microorganisms and polymers . Thereafter, growth of the primary colonizing bacteria changes the surface characteristics of the substratum, rendering it suitable for subsequent colonization by other microorganisms. Finally, the mature biofilm community is formed through synergistic and/or competitive interactions .
Few studies have reported the species composition of bacterial communities in marine biofilms. In particular, the roles and surface specificities of primary colonizers are largely unknown for most surface types . The early-stage biofilms were dominated by the same major classes of bacteria that were most abundant in planktonic communities, with the latter demonstrating a higher diversity when compared with that of biofilm bacteria [9, 10]. The Alphaproteobacteria and Gammaproteobacteria were recognized as the pioneering microorganisms in marine biofilm formation [2, 5, 9–11]. Then, Acidobacteria, Actinobacteria, Bacteroidetes, Chloroflexi, Cyanobacteria, Firmicutes, Planctomycetes, Verrucomicrobia and Beta, Delta and Epsilon groups of Proteobacteria were identified as minor phyla also belonging to these biofilms [5, 8–10, 12–14]. However, the bacterial composition of early-stage biofilms may be affected by the physicochemical properties of the solid surface [5, 10] and by the variation of environmental conditions due for example to the seasons or the characteristics of the immersion sites .
In the present work, we studied bacteria of two original types of marine biofilms from the French Atlantic coast, displaying different characteristics. The first one is a non-permanent benthic biofilm sampled from an intertidal mudflat, which plays a central role in the production of oysters. Indeed, oyster larvae can directly digest and assimilate bacterial carbon . Moreover, this biofilm indirectly feeds the planktonic trophic network through re-suspension of the biofilm in the water column. However, current knowledge on the structure as well as the functioning of this biofilm is largely conceptual and theoretical. The second one is a permanent biofilm formed on carbon steel structures immersed in a French Atlantic harbour and involved in the early stages of the biocorrosion phenomenon . In seawater, complex biofilms closely linked with corrosion products quickly develop on metallic structures. This can influence the deterioration of metal by microorganism activity, thus causing great damages to harbour infrastructures, resulting in economic losses. To date, the initiation of the microbiologically-influenced corrosion processes remains unclear.
Understanding the initial stage of marine biofilm formation is highly important to explain the biofilm formation phenomenon. Unlike the studies in which the structure of pioneering communities, developed on immersed artificial surfaces in seawater, is directly investigated [9, 14], our approach was first to build a collection of culturable marine bacteria isolated from two kinds of biofilms and then to screen for the ability of each isolate to adhere to artificial surfaces and to form biofilms under the same controlled conditions. The objective of this work was to obtain model strains with high forming-biofilm abilities, suitable for further experiments which would allow a better understanding of the colonization of a surface as well as the interactions that can occur between bacteria in a marine biofilm.
Screening of bacteria for their ability to adhere and to form biofilms on polystyrene surfaces
Concerning the bacteria isolated from biofilms developed on corroded carbon steel immersed in sea water, 10 strains among the 70 isolates were able to form biofilms after 24 h (with a ratio of cells grown in biofilm/planktonic cells higher than 2) (Fig. 1). These strains were affiliated to the same taxonomic groups as the benthic bacteria, but the proportion of bacteria from each class varied: 50 % Alphaproteobacteria, 20 % Flavobacteriia, 10 % Gammaproteobacteria, 10 % Bacilli and 10 % Actinobacteria. Under our experimental conditions, the ratios of cel1s grown in biofilm/planktonic cells obtained for these bacteria were lower than for the benthic bacteria. The ratio of 9, for the Roseovarius sp. VA014 strain, was the highest value obtained for bacteria isolated from corroded structures (Fig. 1). Erythrobacter sp. IVA009 was also interesting with a ratio higher than 8 (Fig. 1). The results presented in Fig. 2b show that bacteria able to form a biofilm on polystyrene were found in all samples, but the highest number was isolated from the steel immersed for 2 weeks.
In conclusion, this first screening allowed us to detect 15 benthic bacteria and 10 bacteria from corroded structures able to develop a biofilm in 96-well polystyrene microplates.
Ability of the selected strains to adhere and to form biofilms under static conditions on glass surfaces
Study of the bacterial biofilm structures under dynamic conditions in flow cells
Through the previous steps, seven bacteria have been selected for their capability to form a thick biofilm under static conditions on polystyrene as well as glass surfaces. These bacteria were then studied under dynamic conditions, to further investigate strains able to develop stable biofilms. Thus, bacterial biofilms were grown on glass slides in three-channel flow cells and observed by confocal laser scanning microscopy after staining with the Syto 61 fluorescent dye.
In conclusion, only four strains, Flavobacterium sp. II2003, Roseobacter sp. IV3009, Shewanella sp. IV3014 from the mudflat and Roseovarius sp. VA014 from the steel structure, were able to develop a stable single-species biofilm under dynamic conditions. Each biofilm had a specific structure. Interestingly, these strains were the four bacteria that displayed the thickest biofilms (more than 30 μm) on glass surfaces under static conditions (Fig. 3b).
In this work, we studied bacteria that inhabited two types of marine biofilms. Out of 156 isolates of our marine bacterial collection, only 15 strains from the mudflat biofilms and 10 strains from the corroded metallic structures were able to form single-species biofilms on polystyrene surfaces. This low number of biofilm forming bacteria could be explained by the experimental conditions. For instance, monospecies biofilms were performed, whereas in natural environments, the presence of different bacteria may help to build a biofilm. Moreover, the substratum we used differed from that of the original ecosystem, and the time we let for the bacteria to attach to their support was 2 h only. This step was particularly relevant for pioneer bacteria known to settle in few hours in marine biofilms . Cell density, medium, temperature… may also influence the biofilm formation.
Among our marine cultured strains collection, the most important bacterial classes able to form a biofilm in microtiter polystyrene plates were Alphaproteobacteria (5 from the steel and 3 from the mudflat, of which 6 were Rhodobacterales), Flavobacteriia (4 from the mudflat and 2 from the steel, all being Flavobacteriales), Gammaproteobacteria (4 from the mudflat and 1 from the steel, of which 4 Alteromonadales) and Bacilli (3 from the mudflat and 1 from the steel, all being Bacillales). Only 1 strain from each kind of biofilm belonged to the Actinobacteria class. This is consistent with Dang et al. , who identified members of Alphaproteobacteria (mainly Rhodobacterales), Gammaproteobacteria (mainly Alteromonadales and Oceanospirillales), and Bacteroidetes (mainly Flavobacteriales) groups, as the most common and dominant surface colonizers in their study.
Out of the 8 Alphaproteobacteria able to build a biofilm on microtiter plates, we finally selected two strains belonging to the Roseobacter clade, Roseobacter sp. IV3009 and Roseovarius sp. VA014, for their capacity to also form a biofilm on glass surface under dynamic conditions. It is well known that the Roseobacter clade members are the dominant and ubiquitous primary surface colonizers whatever the type of surfaces, in temperate coastal waters (Pacific and Atlantic coasts) [5, 10, 11, 19]. Moreover, Dang et al.  suggested that Roseobacter were early steel surface colonizers, but also participated to the process of biofilm growth, while Roseovarius would only be pioneer surface colonizers. Roseobacter sp. IV3009 and Roseovarius sp. VA014 are therefore two interesting models of potential pioneering bacteria in marine biofilms.
Among the 5 Gammaproteobacteria able to form a biofilm on polystyrene, 4 were Alteromonadales, with 3 Shewanella and 1 Alteromonas. When Lee et al.  studied the succession of bacterial communities during the first 36 h of biofilm formation on acryl, glass and steel substratum in seawater, they observed that some species of Gammaproteobacteria, such as Alteromonas, were predominant during the first 9 h. Shewanella sp. was until now not described as a predominant bacterium in Atlantic marine biofilms, but was recently observed in early biofilms from Mediterranean Sea . Shewanella sp. IV3014 was selected in this work for its ability to develop an original hairy biofilm with horizontal fibres under dynamic conditions.
We observed a very high diversity among the 6 Flavobacteriia strains able to adhere on polystyrene: they belong to 6 different genera. Previous works showed that bacteria of the Bacteroidetes phylum (containing the Flabovacteriia class) constituted a dominant and diverse bacterial group on carbon steel coupons, at all early immersion stages [16, 19], and it was suggested that different strains might be involved at different stages of the surface colonizing and development microbiota . In our work, only one Flavobacteriia strain, Flavobacterium sp. II 2003, was finally able to form a biofilm in dynamic conditions. Both in static and dynamic conditions, Flavobacterium sp. II2003 displayed a very thick biofilm. It is known that pathogenic Flavobacterium species are responsible for great losses of fish in aquaculture farming worldwide [20, 21]. Aquaculture surfaces are easily colonized and persisting Flavobacterium sp. inhabiting biofilms might serve as a source of infection or reinfection . Although Flavobacterium sp. are important pathogens in the aquaculture setting and have been detected in industrial, domestic, and medical environment biofilms, the manner in which they form biofilms has not been elucidated . Therefore, the Flavobacterium sp. II2003 strain constitutes a very interesting model.
We detected 4 bacteria affiliated to Firmicutes (Bacillus) able to form biofilms on polystyrene. However, Firmicutes were identified as minor phyla found in biofilms formed on acryl, glass and steel substratum in seawater [9, 10, 22]. In our experiments, the Firmicutes then represent a high proportion compared to what occurs in natural environments. However, none of these strains was able to adhere to glass surfaces and they could not be retained as models. The same phenomenon was observed for the two Actinobacteria strains.
In conclusion, this work allowed us to finally select four bacteria able to form a thick biofilm on polystyrene as well as glass surface under dynamic conditions: Flavobacterium sp. II2003, Roseobacter sp. IV3009, Shewanella sp. IV3014 from the mudflat biofilm, and Roseovarius sp. VA014 from the corrosion product-microorganism composite biofilm. Moreover, each of the four strains was able to develop a biofilm with a specific 3D structure. It will then be possible to accurately study potential pioneering bacteria in marine biofilms. Primary colonizers are known to be responsible for the initiation of biofouling and may cause various damages in maritime activities and industries. The paramount importance of the bacterial primary colonizers in surface community formation, dynamics, and function needs to be explored. In future studies, we will investigate the interactions between these high forming-biofilm bacterial models and other marine bacteria from the same ecosystems in order to better understand the initial stage of marine biofilm formation.
Bacterial strains isolation and culture media
A wide range of heterotrophic aerobic bacteria (156) was isolated from two marine biofilms. Benthic bacteria were collected from the intertidal temperate mudflat biofilm of Marennes-Oléron Bay (45°55’N, 01°06’W, Atlantic Coast of France), during three days at low tide in February and July 2008, at 2 h, 3 h and 4 h after emersion. Mudflat samples were collected using core diameter of 20 cm, and the top 2–3 mm was taken. After sampling, mudflat samples were carried to the laboratory at 4° C and immediately processed. The second source of bacteria was the biofilm associated with the corrosion products formed on carbon steel structures immersed in seawater . Briefly, carbon steel coupons (70 × 70 × 6 mm) were immersed in a harbour of La Rochelle (Atlantic coast, France) for 1 week to 2 months at a constant depth of 1 m. The steel composition (98.2 % Fe, 0.122 % C, 0.206 % Si, 0.641 % Mn, 0.016 % P, 0.031 % S, 0.118 % Cr, 0.02 % Mo, 0.105 % Ni and 0.451 % Cu) was the same as that of the harbour metallic structures. At the end of the experiment, the corroded coupons were carried to the laboratory in sealed bags filled with seawater and immediately processed. The mudflat samples and biofilms scraped from the corroded coupons were resuspended in sterile artificial seawater (sea salts Sigma 35 g l-1) and inoculated on Marine Agar (Difco) supplemented with cycloheximide (Sigma 100 μg ml-1) to prevent eukaryotic growth. Bacterial isolates were obtained from the plates after incubation at 20° C in aerobic conditions. Strains were conserved as frozen stocks with 25 % glycerol at -80°C until further processing. For all subsequent tests, the strains were grown in Zobell broth (pastone Bio-Rad 4g l-1; yeast extract Bio-Rad 1g l-1; sea salts Sigma 30g l-1) at 22° C with shaking (150 rpm).
DNA extraction and 16S rRNA gene sequencing
The isolated bacterial strains were identified by 16S rRNA gene sequencing. The genomic DNA of bacteria was extracted with the Genomic DNA from Tissue Kit (Macherey Nagel) from 1 to 5 ml of overnight culture. Amplification of about 1400 bp of the 16S rRNA gene was carried out using 50 ng of genomic DNA in a total volume of 50 μl. The reaction mixtures contained 0.2 μmol l-1 16SUnivF (5’AGAGTTTGATCCTGGCTCA3’) and 16SUnivR (5’GGCTACCTTGTTACGACTT3’) primers, 3 mmol l-1 MgCl2, 320 μmol l-1of each dNTP, and 0.04 Taq polymerase Units (Fermentas), in the corresponding 1× buffer. Denaturation at 95° C for 2 min was followed by 30 cycles of amplification (92° C for 30 s, 54° C for 30 s, 72° C for 1 min 30). About 300 ng of each amplified DNA were sent to GenoScreen (Lille, France) for sequencing. The 16S rDNA sequences were compared with those in GenBank using the Blast software (National Institutes of Health, USA).
Growth of biofilm on polystyrene surfaces (microtiter plates) under static conditions, crystal violet staining
The ability of the bacterial strains to form biofilms onto polystyrene was tested individually by cultivating each of them in 96-well microtiter plates (MICROTEST™ 96, Falcon) under static conditions and by crystal violet staining. The protocol used was a modified version of that described by O’Toole and Kolter . Cells of an overnight bacterial culture were resuspended after 10 min of centrifugation at 7000 g in artificial seawater to a final optical density at 600 nm (OD600) of 0.25 and 150 μl of the resulting bacterial suspensions were loaded per well of the microtiter plates. After incubation for 2 h at 22° C, the wells were gently washed three times with artificial seawater. Thereafter, 150 μl of Zobell medium were transferred in each well and the plate was incubated at 22° C for 24 h. The planktonic fractions were transferred into a new microtiter plate and the absorbance was measured at 600 nm. The plates with biofilms were washed three times with artificial seawater. The biofilms were then stained with a 0.8 % crystal violet solution for 20 min. The wells were then rinsed with ultra-pure water until the wash-liquid was clear (10 times on average) and 150 μl of 96 % ethanol was added to solubilize the attached crystal violet from biofilms. Quantification was carried out by measuring the OD595. To be able to compare the results obtained with strains showing different growth speeds, the biofilm formation was expressed as the ratio of OD595 (cells grown in biofilm)/OD600 (planktonic cells). Assays were performed in triplicate for each strain.
Growth of biofilm on glass surfaces under static conditions, fluorescence microscopy and image analyses
For each tested strain, cells of an overnight bacterial culture were resuspended after 10 min of centrifugation at 7000 g in artificial seawater to a final OD600 of 0.25. One ml of the resulting bacterial suspension was loaded in a compartment of Petri dishes (CellView diameter 35 mm, Greiner Bio-one) containing four compartments and a glass bottom. After incubation for 2 h at 22° C, the compartments were gently washed with artificial seawater and 1ml of Zobell medium was poured in each compartment. Biofilms were then grown for 24 h at 22° C. The surfaces were then rinsed with artificial seawater and biofilms were stained with 4 μg l-1 4–6-diamidino-2-phenylindole (DAPI) in the dark for 20 min. After rinsing with ultra-pure water and drying, samples were analysed using a fluorescence microscope DMI6000B system (magnification 1000×, Leica Microsystems, Germany), over an average of 10 fields. The three dimensional (3D) structures were reconstituted by using IMARIS software. The percentages of colonized surface (%) were calculated using the NIH ImageJ software . All experiments were performed in triplicate.
Growth of biofilm on glass surface under dynamic conditions in three channel flow cell, confocal laser scanning microscopy and image analyses
Bacterial biofilms were grown on glass slides in three-channel flow cells (channel dimensions 1 by 4 by 40 mm, Technical University of Denmark Systems Biology, Denmark) . The flow system was assembled, prepared and sterilized as described by Tolker-Nielsen and Sternberg . The substratum consisted in a microscope glass coverslip (24 × 50 st1, KnittelGlasser, Germany). Flow cells were inoculated with overnight bacterial cultures diluted in artificial seawater to a final OD600 of 0.1. Bacteria were allowed to attach during 2 h at 22° C without medium flow. The channels were then washed by applying a flow of artificial seawater for 15 min at a rate of 2 ml h-1 to remove planktonic cells. Biofilm growth was then performed under a constant flow of Zobell (2 ml h-1) for 24 h at 22° C. Microscopic observations were performed by confocal laser scanning microscopy using a TCS-SP2 system (Leica Microsystems, Germany). The 2-h attached cells and the biofilms formed after 24 h on the glass surface were observed by staining bacteria with 5 μmol l-1 Syto 61 red. Images were obtained using the Leica confocal software. The surface coverages after the 2 h adhesion step were evaluated using the ImageJ software. The biofilm stacks were analysed with the COMSTAT software (developed in MATLAB, ) to estimate the maximal and average thicknesses (μm) and the biovolume (μm3 μm-2). The values were calculated from three independent experiments from which a total of 15 image stacks were obtained.
The standard deviations were calculated using Matlab software (Mathworks Inc., Natick, USA). The statistical analyses were determined by the Student t-test and considered as significant if p values are < 0.05.
The authors wish to thank the Conseil Général de la Charente Maritime (France) for the PhD grant of Ibtissem Doghri. This work was financially supported by the CNRS “Ecosphère Continentale et Côtière” program 2013–14 (France). The LIENSs and LBCM are supported by the Région Poitou-Charentes (France), the Région Bretagne (France) and the European FEDER funds.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science. 1999;284:1318–22.View ArticlePubMedGoogle Scholar
- Salta M, Wharton JA, Blache Y, Stokes KR, Briand JF. Marine biofilms on artificial surfaces: structure and dynamics. Environ Microbiol. 2013;15:2879–93.PubMedGoogle Scholar
- Karatan E, Watnick P. Signals regulatory network and materials that build and break bacterial biofilms. Microbiol Mol Biol Rev. 2009;73:310–47.PubMed CentralView ArticlePubMedGoogle Scholar
- Jefferson KK. What drives bacteria to produce a biofilm? FEMS Microbiol Lett. 2004;236:163–73.View ArticlePubMedGoogle Scholar
- Dang H, Lovell CR. Bacterial primary colonization and early succession on surfaces in marine waters as determined by amplified rRNA gene restriction analysis and sequence analysis of 16S rRNA genes. Appl Environ Microbiol. 2000;66:467–75.PubMed CentralView ArticlePubMedGoogle Scholar
- Davey ME, O’Toole GA. Microbial biofilms: from ecology to molecular genetics. Microbiol Mol Biol Rev. 2000;64:847–67.PubMed CentralView ArticlePubMedGoogle Scholar
- Stoodley P, Sauer K, Davies DG, Costerton JW. Biofilms as complex differentiated communities. Annu Rev Microbiol. 2002;56:187–209.View ArticlePubMedGoogle Scholar
- Huggett M, Nedved B, Hadfield M. Effects of initial surface wettability on biofilm formation and subsequent settlement of Hydroides elegans. Biofouling. 2009;25:387–99.View ArticlePubMedGoogle Scholar
- Lee JW, Nam JH, Kim YH, Lee KH, Lee DH. Bacterial communities in the initial stage of marine biofilm formation on artificial surfaces. J Microbiol. 2008;46:174–82.View ArticlePubMedGoogle Scholar
- Jones PR, Cottrell M, Kirchman DL, Dexter SC. Bacterial community structure of biofilms on artificial surfaces in an estuary. Microb Ecol. 2007;53:153–62.View ArticlePubMedGoogle Scholar
- Dang H, Li T, Chen M, Huang G. Crossocean distribution of Rhodobacterales bacteria as primary surface colonizers in temperate coastal marine waters. Appl Environ Microbiol. 2008;74:52–60.PubMed CentralView ArticlePubMedGoogle Scholar
- Dang H, Lovell CR. Numerical dominance and phylotype diversity of marine Rhodobacter species during early colonization of submerged surfaces in coastal marine waters as determined by 16S ribosomal DNA sequence analysis and fluorescence in situ hybridization. Appl Environ Microbiol. 2002;68:496–504.PubMed CentralView ArticlePubMedGoogle Scholar
- Webster NS, Smith LD, Heyward AJ, Watts JEM, Webb RI, Blackall LL, et al. Metamorphosis of a scleractinian coral in response to microbial biofilms. Appl Environ Microbiol. 2004;70:1213–21.PubMed CentralView ArticlePubMedGoogle Scholar
- Brian-Jaisson F, Ortalo-Magné A, Guentas-Dombrowsky L, Armougom F, Blache Y, Molmeret M. Identification of bacterial strains isolated from the Mediterranean sea exhibiting different abilities of biofilm formation. Microb Ecol. 2014;68:94–110.View ArticlePubMedGoogle Scholar
- Douillet P. Bacterivory in Pacific oyster Crassostrea gigas larvae. Mar Ecol Prog Ser. 1993;98:123–34.View ArticleGoogle Scholar
- Lanneluc I, Langumier M, Sabot R, Jeannin M, Refait P, Sablé S. On the bacterial communities associated with the corrosion product layer during the early stages of marine corrosion of carbon steel. Int Biodeterior Biodegrad. 2015;99:55–65.View ArticleGoogle Scholar
- Klausen M, Gjermansen M, Kreft J-U, Tolker-Nielsen T. Dynamics of development and dispersal in sessile microbial communities: examples from Pseudomonas aeruginosa and Pseudomonas putida biofilms. FEMS Microbiol Lett. 2006;261:1–11.View ArticlePubMedGoogle Scholar
- Barken KB, Pamp SJ, Yang L, Gjermansen M, Bertrand JJ, Klausen M, et al. Roles of type IV pili, flagellum-mediated motility and extracellular DNA in the formation of mature multicellular structures in Pseudomonas aeruginosa biofilms. Environ Microbiol. 2008;10:2331–43.View ArticlePubMedGoogle Scholar
- Dang H, Chen R, Wang L, Shao S, Dai L, Ye Y, et al. Molecular characterization of putative biocorroding microbiota with a novel niche detection of Epsilon- and Zetaproteobacteria in Pacific Ocean coastal seawaters. Environ Microbiol. 2011;13:3059–74.View ArticlePubMedGoogle Scholar
- Basson A, Flemming LA, Chenia HY. Evaluation of Adherence, Hydrophobicity, Aggregation, and Biofilm Development of Flavobacterium johnsoniae-Like Isolates. Microb Ecol. 2008;55:1–14.View ArticlePubMedGoogle Scholar
- Starliper EC. Bacterial coldwater disease of fishes caused by Flavobacterium psychrophilum. J Adv Res. 2011;2:97–108.View ArticleGoogle Scholar
- Chung H, Lee O, Huang Y, Mok S, Kolter R, Qian P. Bacterial community succession and chemical profiles of subtidal biofilms in relation to larval settlement of the polychaete Hydroides elegans. ISME J. 2010;4:817–28.View ArticlePubMedGoogle Scholar
- O'Toole GA, Kolter R. Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a genetic analysis. Mol Microbiol. 1998;28:449–61.View ArticlePubMedGoogle Scholar
- Rasband WS. ImageJ, U.S. National Institutes of Health, Bethesda, Maryland, USA, imagej.nih.gov/ij/, 1997–2012Google Scholar
- Pamp SJ, Sternberg C, Tolker-Nielsen T. Insight into the microbial multicellular lifestyle via flow-cell technology and confocal microscopy. Cytometry A. 2009;75:90–103.View ArticlePubMedGoogle Scholar
- Tolker-Nielsen T, Sternberg C. Growing and analyzing biofilms in flow chambers. Curr Protoc Microbiol. 2011;2:1–17.Google Scholar
- Heydorn A, Nielsen AT, Hentzer M, Sternberg C, Givskov M, Ersbøll BK, et al. Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology. 2000;146:2395–407.View ArticlePubMedGoogle Scholar