Microfluidic Assay to Quantify the Adhesion of Marine Bacteria
© The Author(s) 2012
Received: 2 December 2011
Accepted: 28 February 2012
Published: 21 March 2012
For both, environmental and medical applications, the quantification of bacterial adhesion is of major importance to understand and support the development of new materials. For marine applications, the demand is driven by the quest for improved fouling-release coatings. To determine the attachment strength of bacteria to coatings, a microfluidic adhesion assay has been developed which allows probing at which critical wall shear stress bacteria are removed from the surface. Besides the experimental setup and the optimization of the assay, we measured adhesion of the marine bacterium Cobetia marina on a series of differently terminated self-assembled monolayers. The results showed that the adhesion strength of C. marina changes with surface chemistry. The difference in critical shear stress needed to remove bacteria can vary by more than one order of magnitude if a hydrophobic material is compared to an inert chemistry such as polyethylene glycol.
Biofouling, the colonization of submerged artificial or natural surfaces by undesired biological organisms, is a major problem for many marine industries resulting in both, environmental and economic penalties [1, 2]. As application of biocidal antifouling (AF) paints is increasingly being restricted, fouling-release (FR) coatings are currently considered as alternative. Such non-toxic alternatives appear attractive, as they seem to reduce fuel consumption compared to conventional ablative AF coatings [3–5]. Bacteria are among the first microorganisms to colonize submersed interfaces to form biofilms . Both, bacteria and microalgae produce extracellular polymeric substances (EPS), which contain polysaccharides, lipopolysaccharides, proteins and nucleic acids . Such substances mediate the initial adhesion to surface and constitute the matrix of the biofilms . In some cases, marine bacteria influence subsequent colonization by invertebrates, algae  and tubeworms [9–11]. Understanding bacterial adhesion and optimization of coatings so that they can easily be cleaned are important to improve commercial fouling-release technologies.
In the past different techniques were used to quantify adhesion of biological material to surfaces: Atomic force microscopy (AFM) , spinning disk , hydrodynamic shear force assays such as a water jet apparatus , flow channels [14–16] or microfluidic channels [17, 18]. Most of these techniques are conventionally applied as laboratory assays. In field experiments, water jetting and grooming tools are used as techniques to assess cleanability of fouling-release coatings in real, mixed species environments [19–21]. Especially for laboratory tests, microfluidic assays have a number of advantages: they allow quantifying adhesion strength on relatively small sample areas and require only small amounts of bacteria. Experiments mostly only take some hours and the experiment can easily be parallelized. The main advantage of a microfluidic assay lies in the fact that typically ca. 400 cells can simultaneously be investigated and the variation of the shear force across several orders of magnitude allows to record detachment of each single cell [12, 18, 22]. In contrast to many other approaches this means that the cell density is accurately known from the beginning, and as the field of view remains unchanged, the same initial seeding density is valid for the entire experiment. We recently described a microfluidic device which allows the measurement of cell-surface interaction . Cells can be incubated in the channel for several hours after which they are removed by a stepwise increased flow. Using self assembled monolayers with different abilities to bind water we were able to detect that subtle changes in hydration strongly influence the adhesion strength of fibroblasts . Furthermore this new assay revealed that cell removal from directed nanostructures depends on the flow direction  and that CD44+ leukemic cells attach to hyaluronans by a catch bond activated binding .
In this work we apply the microfluidic shear force assay to quantify the adhesion strength of the marine bacterium Cobetia marina on chemically different model surfaces. This bacterium is used as a model system for marine biofouling because it is frequently found in biofilms and influences secondary colonization by invertebrates and algae . To demonstrate the applicability of the microfluidic assay, we used self-assembled monolayers as well-defined model surfaces. Self-assembled monolayers [25, 26] are highly useful tools to reproducibly prepare coatings and frequently applied to study response of marine biofouling organisms [14, 16, 27–32]. One major advantage is that the mechanical properties are determined by the substrate while physicochemical properties, such as wetting and hydration are determined by the thin organic film. The accumulation of C. marina on chemically differently terminated self-assembled monolayers (SAMs) revealed that surface properties change the amount of accumulated biomass [14, 29]. In this article we describe the effect of undecanethiol SAMs with –CH3, –NH2, –OC7F10CF3 termination and polyethylene glycol (PEG) terminated SAMs on the adhesion strength of the marine bacteria C. marina. We chose these surfaces as they cover a large range of wettabilities with different inert properties as numerous recent studies revealed [14, 27, 29, 31, 33–38].
2.1 Preparation and Characterization of SAMs
Ethanol (p.a.) was purchased from Sigma-Aldrich (Munich, Germany). Deionized water was purified with a Milli-Q plus system (Millipore, Schwalbach, Germany), the final resistivity was ≥18 MΩcm. Nexterion® B glass slides (Schott, Mainz, Germany) were used as substrates for adhesion experiments and as substrates for deposition of gold films. Thin films of polycrystalline gold were prepared by thermal vapor deposition of 30 nm gold (99.99 % purity) onto Nexterion® glass slides predeposited with a 5 nm titanium adhesion layer. Evaporation was performed at a pressure of 2 × 10−7 mbar and a deposition rate of 0.5 nm s−1, leading to a root-mean-square (rms) roughness of about 1 nm. The chemicals used for self-assembly were dodecanethiol (DDT, HS–(CH2)11–CH3) and 11-amino-undecanethiol (AUDT, HS–(CH2)11–NH2), purchased from Sigma-Aldrich. 11-(tridecafluorooctyloxy) undecanethiol (FUDT, HS–(CH2)11–O–(CH2)2–(CF2)5–CF3) were retrieved from Prochimia, and Hydroxy-PEG2000-thiol (PEG, HS–(CH2)2(OCH2CH2)44OH), was purchased from Rapp Polymere GmbH (Tuebingen, Germany). All chemicals were used as received without further purification. For the SAM formation the gold slides were first cleaned in an UV reactor for 2 h and then immersed into the corresponding 1 mM thiol solution in ethanol p.a. for 24 h, except for PEG where 48 h were required. Before and after immersion the samples were rinsed and sonicated for 3 min in ethanol p.a., and finally dried in a flow of nitrogen. The samples were stored under argon.
2.2 Surface Analysis
Successful assembly of the SAMs was verified by contact angle goniometry, spectral ellipsometry, and X-ray photoelectron spectroscopy (XPS). Sessile drop water contact angles were measured with a custom built goniometer under ambient conditions. Using digital images of the sessile droplet, the drop shape is modeled by the Young–Laplace equation and the contact angle at the interface is calculated. The contact angle was determined three times on each sample and the average is reported. SAM thickness measurements were performed with a M-44 multiple wavelength ellipsometer from J. A. Woollam Co., Inc. (Lincoln, NE). The organic film was modeled as a single Cauchy layer using the software WVASE from J. A. Woollam Co. The reported values are the average of three measurements. Film purity, composition, and thickness were analyzed by XPS using a Leybold-Heraeus MAX 200 X-ray photoelectron spectrometer with a magnesium anode as the X-ray source (Kα, 1253.6 eV).
2.3 Bacteria Culture
Most assays described in literature prefer to work with bacteria in the log phase for adhesion experiments  as the results are most reliable. To bring bacteria into the log phase for our microfluidics experiments, the overnight culture was diluted 1:100 in MB and held in liquid culture for approximately 3 h. After this, the OD was frequently measured until the desired OD600 of 0.1 was reached. This suspension was harvested by centrifugation (Hettrich, Mikro 22 R at 10,000 rpm for 2 min), washed in sterile (0.45 μm filtered) ASW to remove any residual marine broth, and resuspended in ASW. Prior to use in the microfluidic experiment, the suspension was filtered through a 5 μm filter to remove larger bacterial aggregates. The number of bacteria in the suspension with an OD of 0.1 was 107 cells mL−1as we determined by analysis of the number of colony forming units (CFU).
2.4 Microfluidic Bacteria Detachment Assay
Q is the volumetric flow rate, μ the viscosity of the medium (for sea water ~1 × 10−3 kg m−1 s−1 at 20 °C ), and the channel’s dimensions height h and width w. This model agrees well with more elaborate calculations .
3 Results and Discussion
3.1 Influence of Medium and Incubation Time on the Adhesion Strength of Bacteria
3.2 Adhesion of Cobetia marina to SAMs with Different Chemical Termination
Properties of the different self assembled monolayers: water contact angle and film thickness as determined by spectral ellipsometry
113 ± 2
16 ± 1
106 ± 1
13 ± 1
54 ± 1
16 ± 1
30 ± 1
30 ± 2
Especially the major difference in attachment strength of bacteria on the hydrophobic surfaces (≈3,500–4,000 dyn/cm2) and on PEG coated surfaces (≈200 dyn/cm2) by more than one order of magnitude showed that the microfluidic detachment assay was capable of discriminating the adhesion strength of bacteria to surfaces and thus to correlate surface properties with their ability to reduce bacterial attachment strength. In the future, we intend to apply this technique to test different coatings in order to find optimized surface compositions and properties, which are able to minimize bacterial adhesion strength.
We established a microfuidic assay to quantify adhesion strength of bacteria. The total duration of an experiment using four channels was less than 3 h, which allowed multiple experiments per day and thus a high sample throughout. Also the assay only required small glass chips (dimensions of 25 × 25 mm2). As the fluidic environment was well controlled, quantitative data on the attachment strength of ≈400 cells could be probed simultaneously. Most importantly, detachment of single, individual cells was observed and thus the shear stress needed for their removal was obtained for each single bacterium in the field of view (256 by 192 μm2). The assay covers six orders of magnitude of wall shear stresses and the situation in the microchannels was correlated with the turbulent hydrodynamic shear acting on the hull of a vessel cruising through the ocean. From detachment curves, both the adherent fraction and the critical shear stress for removal of 50 % of the adherent cells can be obtained. The decisive experimental parameters such as incubation time and medium were optimized for the biofouling marine bacterium C. marina. SAMs with different chemical termination were investigated towards their influence on both, fraction of attached cells and adhesion strength. The assay discriminated well between bacteria which adhere on hydrophobic SAMs and resistant PEG coatings and showed that the critical shear stress needed for bacterial removal differed by more than one order of magnitude. Thus the assay is a sensitive tool for the quantification of bacteria-surface interaction and capable to accurately discriminate the fouling-release potential of surfaces.
The article was written following the award of the Biointerfaces division poster price to M.P.A.S. sponsored by Biointerphases at the 58th AVS International Conference and Exhibition held in Nashville, TN. The authors thank the Office of Naval Research (Grant number N00014-08-1-1116) and the Helmholtz program Biointerfaces for support of this work. The stimulating discussions with M. Grunze are kindly acknowledged. We thank G. Albert for preparation of the gold coated glass slides. We are gratefully indebted to L. Ista and G. Lopez (University of New Mexico, NM, USA) for their kind introduction into bacteria culture. We also acknowledge helpful advices from John Finlay and Maureen Callow (University of Birmingham, UK).
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