Research Article Cite This: ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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Study of Biofilm Growth on Slippery Liquid-Infused Porous Surfaces Made from Fluoropor Nico Keller,† Julia Bruchmann,‡ Thomas Sollich,‡ Christiane Richter,† Richard Thelen,† Frederik Kotz,† Thomas Schwartz,‡ Dorothea Helmer,† and Bastian E. Rapp*,† †
Institute of Microstructure Technology and ‡Institute of Functional Interfaces, Karlsruhe Institute of Technology, 76344 Eggenstein-Leopoldshafen, Germany
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S Supporting Information *
ABSTRACT: Undesired growth of biofilms represents a fundamental problem for all surfaces in long-term contact with aqueous media. Mature biofilms resist most biocide treatments and often are a pathogenic threat. One way to prevent biofilm growth on surfaces is by using slippery liquid-infused porous surfaces (SLIPS). SLIPS consist of a porous substrate which is infused with a lubricant immiscible with the aqueous medium in which the bacteria are suspended. Because of the lubricant, bacteria cannot attach to the substrate surface and thus formation of the biofilm is prevented. For this purpose, we manufactured substrates with different porosity and surface roughness values via UVinitiated free-radical polymerization in Fluoropor. Fluoropor is a class of highly fluorinated bulk-porous polymers with tunable porosity, which we recently introduced. We investigated the growth of the biofilm on the substrates, showing that a reduced surface roughness is beneficial for the reduction of biofilm growth. Samples of low roughness effectively reduced Pseudomonas aeruginosa biofilm growth for 7 days in a flow chamber experiment. The lowroughness samples also become transparent when infused with the lubricant, making such surfaces ideal for real-time observation of biofilm growth by optical examination. KEYWORDS: slippery lubricant-infused porous surfaces, porosity, roughness, biofouling, long-term stability fluoropolymers, Fluoropor
1. INTRODUCTION Undesired growth of bacteria on technical substrates is among the most relevant challenges for medical implants,1 marine hulls,2 drinking water systems,3 oil pipelines,4 and medical devices such as contact lenses.5 When microorganisms grow on a surface, they can form a highly resistant layer of biomaterial. In this so-called “biofilm”, the microorganism is embedded in an extracellular polymeric substance which consists of polysaccharides and proteins as well as nucleic acids.6 Biofilms have an increased resistance to antibiotics and other antimicrobial treatments when compared to the planktonic cultures of the same cells.7,8 Biofilm formation on medical devices is the reason for many hospital-acquired infections.9 Growth of biofilms in pipelines can lead to clogging and therefore to failure of the technical system.10 In case of drinking water systems, biofilms represent a serious threat to human health.3 Preventing bacterial growth and biofilm formation on technical substrates is therefore of high interest, and several approaches for preventing biofilm growth have been reported either relying on the destruction of the adhering bacteria or on the prevention of bacterial adhesion. For the former approach, the surfaces are coated with matrices containing antibiotics such as gentamycin11 or antibacterial metallic ions such as silver(I).12 However, this strategy has several limitations. Over time, bacteria will develop resistances both against antibiotics13 as well as metallic coatings,12 which © XXXX American Chemical Society
renders the surfaces ineffective. Additionally, the leaching of antibiotic agents into the environment is problematic; therefore, widespread use of several of these coatings is now prohibited for many applications.13 However, the most limiting disadvantage of these strategies is the fact that antibacterial coatings only kill the first adhering layer of bacteria. This first layer of dead bacteria represents an ideal basis for attachment and growth of the next layer of bacteria. The latter will grow almost uninhibited as the antimicrobial coating is effectively passivated. Therefore, approaches which prevent the adhesion of bacteria are significantly more promising. Common methods to design such antifouling surfaces are coatings based on hydrophilic polymers such as poly(ethylene glycol)14 or poly(2-hydroxyethyl methacrylate),15 which limit the protein interaction with the technical substrate by formation of hydrate layers. Low surface energy polymers such as polytetrafluoroethylene have been used to prevent nonspecific bioadhesion by reducing molecular adhesive forces between the coating and the bacteria.16 However, none of the above methods have yet proven ideal for long-term antifouling. Received: July 24, 2018 Accepted: December 17, 2018
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DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
Research Article
ACS Applied Materials & Interfaces
blended with 1 wt % (with respect to the monomer) DMPAP in acetone (1 mg/μL). Different porosities of the Fluoropor substrates are generated by adjusting the concentration of cyclohexanol between 10 and 30 wt % (with respect to the reaction mixture mass) and the concentration of 1H,1H,2H,2H-perfluorooctanol between 40 and 20 wt % (with respect to the reaction mixture mass). For nonporous Fluoropor substrates, Fluorolink MD 700 is blended with 1 wt % (with respect to the monomer) DMPAP in acetone (1 mg/μL). Commercially available polycarbonate HybriWell chambers were placed on the functionalized glass slides. After filling the HybriWell chamber with the mixture, the chamber was irradiated via the glass slide with UV light (370 nm, Lumatec Superlite SUV mercury arc lamp, Lumatech, Germany) for 5 min. Subsequently, the chamber was removed and the polymer was rinsed with 2-propanol and placed in 2-propanol overnight. Afterward, the polymer was dried at room temperature for 4 h. After drying, the Fluoropor substrate was wiped with a tissue to remove the unstructured top polymer layer formed during polymerization in the closed chamber. 2.4. Preparation of Fluoropor SLIPS. 750 μL of Aflunox 100 was pipetted onto the porous Fluoropor substrate and left to wick the porous structure overnight. Excess lubricant was removed by tilting the surface about 45° for 1 h. 2.5. Scanning Electron Microscopy for Surface Characterization. A SUPRA 60 VP (Zeiss, Germany) was used for all scanning electron microscopy (SEM) measurements. Prior to analysis, the samples were sputtered with a 25 nm thick silver layer via physical vapor deposition. 2.6. Mercury Intrusion Porosimetry. The total porosity of the different Fluoropor substrates was measured using a Pascal 140/440 mercury porosimetry system (Porotec, Germany). For mercury, a contact angle of 140° and a surface tension of 480 mN/m were assumed. The mercury intrusion porosimetry was carried out up to a final pressure of 400 MPa. For each porosity assessed, a total of three samples were characterized. For sample preparation, 400 μL reaction mixture were placed in a mold (diameter: 1.2 mm) and cured with UV-light for 5 min. The samples were then placed in acetone overnight and dried afterward. The surface of the sample was opened with 2000 grain sandpaper. 2.7. Optical Transmission Measurement. The optical transmission of the Fluoropor and Fluoropor-SLIPS samples was determined using a UV/vis spectrometer (Evolution 201, Thermo Scientific, Germany). The samples were measured against a Nexterion clean room slide blank. 2.8. Bacteria Cultivation. Pseudomonas aeruginosa strain PA14 were cultured in Basal Medium 2 (BM2, 62 × 10−3 M potassium phosphate, 7 × 10−3 M (NH4)2SO4, 2 × 10−3 M MgSO4, 10 × 10−6 M FeSO4, 0.4% glucose).33 The samples were placed on a shaker and incubated overnight at 37 °C and 50 rpm. 2.9. Microfluidic Flow Cell Setup. The antifouling properties and the hydrodynamic stability of the Fluoropor-SLIPS were assessed using a microfluidic flow test. The flow cell was assembled by pressing a microfluidic channel made from polydimethylsiloxane (PDMS, width: 4 mm, height: 0.5 mm, length: 24 mm) onto glass slides onto which the different Fluoropor-SLIPS were created. The flow cell was generated via soft lithography using a master structure generated via milling. The two-component PDMS prepolymer (Wacker Elastosil RT 601) was mixed in 9:1 mass ratio and left to cure for 4 h. The flow cell was connected to a peristaltic pump (IPC 12/ISM 932, Ismatec, Germany) via Teflon tubing. 2.10. Biofilm Growth. Biofilm was grown on the FluoroporSLIPS substrate inside the microfluidic flow cell. The flow cell was first sterilized by pumping a mixture of 70 vol % ethanol in water through the cell at 0.1 mL/min for several minutes. Overnight cultures of P. aeruginosa strain PA14 were diluted to an optical density of 0.1 (∼108 CFU mL−1) in BM2, and the bacterial suspension was pumped through the microfluidic flow cell at 0.1 mL/min for 3 h to induce biofilm formation. Afterward, the bacterial suspension was exchanged for fresh BM2 medium to cultivate the growing biofilm for seven days with a flow rate of 0.1 mL/min.
Recently, approaches which shield the solid substrate with a liquid thus preventing direct contact of the bacteria with the technical surface have gained significant attention. These socalled slippery liquid-infused porous surfaces (SLIPS) are inspired by the pitcher plant Nepenthes.17 SLIPS consist of a lubricant which is spread on nano-/microstructured surfaces. The lubricant is immiscible with the aqueous media containing the bacteria, thus a slippery liquid/liquid interface forms which prevents bacterial adhesion. SLIPS are not only suitable for reducing biofilm formation18−20 but also can reduce biofouling by blood,21 algae,22,23 and mussels.24 In the earlier fabrication methods of SLIPS porous Teflon membrane and lithographically structured epoxy resin was used to hold the lubricant.17 Other approaches uses colloidal templating,25 porous polymer layers,19,20,22 layer-by-layer deposition, electrodeposition,23 anodization,26 breath figure patterns27,28 or ice templating29−31 to form the nano-/ microstructured surfaces. In this study, we use “Fluoropor” which are highly fluorinated polymer foams with adjustable porosity made via radical emulsion polymerization, which we have recently reported.32 Fluoropor has a very low surface energy of only 7.2 mN/m32 and does not require additional treatments for further reduction of the surface energy. SLIPS based on Fluoropor can thus be generated in a simple two-step protocol: (1) in situ emulsion polymerization and washing followed by (2) impregnation with a fluorinated lubricant. Fluoropor allows generating substrates with different roughness, which allows studying the influence of the surface topography on the formation of a biofilm on SLIPS generated from such rough Fluoropor substrates. We show that the surface roughness influences the formation and attachment of the biofilm. The roughness also influences the transparency of the generated SLIPS which, for low roughness substrates, can reach over 80% transmission effectively opening up applications where substrate transparency is crucial.
2. EXPERIMENTAL SECTION 2.1. Materials and Methods. Fluorolink MD700 was purchased from Acota (United Kingdom). 2,2-Dimethoxy-2-phenylacetophenone (DMPAP) was purchased from Sigma-Aldrich (Germany). Acetone, ammonium sulfate, cyclohexanol, ethanol, glucose, hydrochloric acid (37%), iron(II) sulfate, magnesium acetate, magnesium sulfate, 3-(methacryloyloxypropyl)dimethylchlorosilane, methanol, potassium phosphate, 2-propanol, toluene, and Tris-base were purchased from Merck (Germany). 1H,1H,2H,2H-Perfluorooctanol was purchased from Apollo Scientific (United Kingdom). Nexterion Slide Glass B clean room slides were purchased from Schott (Germany). HybriWell incubation chambers were purchased from BioCat (Germany). Aflunox 100 was a kind gift from Klüber Lubrication (Germany). ELASTOSIL RT 601 was purchased from Wacker (Germany). SYTO 17 red fluorescent nucleic acid stain was purchased from ThermoFisher Scientific (Germany). 2.2. Functionalization of Glass Slides. Clean room glass slides were placed in a mixture of concentrated hydrochloric acid and methanol (50:50 vol %) for 30 min. Thereafter, the glass slides were rinsed with 2-propanol and deionized water and dried with compressed air. Subsequently, the glass slides were placed in a 25 mM solution of (3-methacryloyloxypropyl)dimethylchlorosilane in toluene for 1 h. Afterward, the slides were rinsed with 2-propanol and deionized water and dried with compressed air. 2.3. Synthesis of Fluoropor. The process for generating porous Fluoropor substrates was described previously.32 Briefly, 50 wt % Fluorolink MD700 (Lot-Nr BL0916) was mixed with 50 wt % of a mixture of cyclohexanol and 1H,1H,2H,2H-perfluorooctanol and B
DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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ACS Applied Materials & Interfaces
Figure 1. Porosity of Fluoropor substrates made with different concentrations of the porogen. (a) SEM images of Fluoropor 10, Fluoropor 15, Fluoropor 20, Fluoropor 25 and Fluoropor 30. The numbers refer to the weight percentage of the porogen (cyclohexanol) in the mixture (scale bar: 1 μm). (b) SEM images of the same samples with a greater magnification (scale bar: 500 nm). (c) Porosity measured for the given samples by mercury intrusion porosimetry. (d) Roughness of the Fluoropor samples as given by the Rq values determined by white light interferometry. 2.11. Fluorescence Staining and Quantification of Bacterial Adhesion. SYTO 17 nucleic acid staining was used to visualize the biofilm. Samples were carefully washed in sterile cell wash buffer (5 × 10−3 M magnesium acetate, 10 × 10−3 M Tris-base, pH 8) and stained with 5 μM SYTO 17 (stock diluted 1:1000 in cell wash buffer) for 10 min. Afterward, samples were carefully washed in sterile cell wash buffer. Biofilm formation was assessed using fluorescence microscopy (Axio Imager.M2, Zeiss, Germany). The rhodamine channel was used for fluorescence imaging at 100-fold and 200-fold magnification. The surface coverage of the biofilm was assessed using the software ImageJ (version: 1.51 s). Images were converted to 8-bit and a constant threshold value (39) was employed to produce binary pictures. All pixels above the threshold intensity appear white and represent the biofilm-covered surface. All pixels under the threshold appear black and represents biofilm free surface. Biofilm coverage was calculated in percent of the total surface. The arithmetic mean and the standard deviation were calculated from these values. At least four fluorescence images were analyzed per sample. A two-sided Student’s t-test was used for statistical data analyses. 2.12. Shear Force Shaking Experiment. We tested the hydrodynamic stability of the Fluoropor-SLIPS by using a shear force shaking experiment. The Fluoropor-SLIPS samples were placed into a Petri dish and 30 mL of deionized water was filled. The samples were placed on a shaker and shaken for seven days at 50 rpm. 2.13. White Light Interferometry. Surface roughness was measured using white light interferometry of type Contour GT-K (Bruker, USA). For a better contrast, porous polymer samples were sputtered with a 25 nm thick silver layer. 2.14. Measurement of the Layer Thickness. The thicknesses of the polymer coatings on glass were determined using a length gauge MT 60M (Heidenhain, Germany). 2.15. Comparison of Background Staining of Fluoropor and Fluoropor-SLIPS. Fluoropor 20 and Fluoropor-SLIPS 20 samples were thoroughly washed with 70 vol % ethanol in water and dried under a clean bench. Afterward, the samples were placed into a Petri dish. Overnight cultures of P. aeruginosa strain PA14 were diluted to an optical density of 0.1 (∼108 CFU mL−1) in BM2. The samples were incubated for 3 days at 37 °C and 50 rpm. The medium was
changed every day. The staining was carried out as described in the Experimental Section.
3. RESULTS AND DISCUSSION Fluoropor substrates with varying pore sizes were created by adapting our previously described protocol via light-induced radical polymerization on glass slides functionalized with (3methacryloyloxypropyl)dimethylchlorosilane.32 Porosity is controlled by adding cyclohexane as immiscible nonsolvent porogen and 1H,1H,2H,2H-perfluorooctanol as the emulsifier. Substrates with varying porosity were generated using mixtures with 10, 15, 20, 25 and 30 wt % of cyclohexanol, respectively. The amount of emulsifier was adjusted accordingly in the mixtures (Materials and Methods section for details). In the following, we refer to these samples as Fluoropor 10, Fluoropor 15, Fluoropor 20, Fluoropor 25, and Fluoropor 30 based on the cyclohexanol content in the mixture. Mixtures with a cyclohexanol concentration higher than 30 wt %, that is, Fluoropor 35 and higher, could not be fabricated because of separation of the monomer/porogen phase. Figure 1a,b shows SEM images of each substrate and confirms the gradual increase in the size of microglobules and pores as the cyclohexanol content of the polymerization mixture is increased. The SEM images of Figure 1a,b suggest an increase in porosity for all samples. However, the porosimetry data in Figure 1c suggest the same porosity for Fluoropor 10, Fluoropor 15, and Fluoropor 20. The average pore sizes measured by mercury intrusion porosimetry assume cylindrical pores and depend on the largest connection between the pores within the network. Therefore, the pore sizes detected by porosimetry are always below the pore sizes observed by optical methods such as SEM.34 Because of the Washburn equation35 the theoretical smallest pore radius detectable at 400 MPa (the limit of the instrument used) are pores of 3.7 C
DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
Research Article
ACS Applied Materials & Interfaces
Figure 2. Optical transmission of the porous Fluoropor substrates and the lubricant-infused Fluoropor-SLIPS. (a) Images of noninfused Fluoropor and (b) corresponding infused Fluoropor-SLIPS coated on glass (Scale bar: 1 cm). (c) UV/vis spectra of the noninfused Fluoropor substrates are shown. The large increase in porosity for Fluoropor-SLIPS 25 and Fluoropor-SLIPS 30 results in nontransparent substrates. (d) Because of the optical index match of the lubricant Aflunox 100 and the Fluoropor samples, the optical transmission of the substrates increases significantly upon infusion. All samples were prepared on glass slides with a thickness of 118.4 ± 5.0 μm. Transmissions were measured against a glass slide blank.
Fluoropor-SLIPS are generated by infusing the Fluoropor substrates with a lubricant; we refer to the infused samples as Fluoropor-SLIPS 10, Fluoropor-SLIPS 15, Fluoropor-SLIPS 20, Fluoropor-SLIPS 25, and Fluoropor-SLIPS 30 (see Figure 2b). After infusion with the lubricant, the optical transmission of the samples increases drastically (see Figure 2d). The optical indices of refraction between bulk Fluoropor and the fluorinated lubricant are similar (nonporous Fluoropor bulk: nd = 1.3438, Aflunox 100: nd = 1.3039), thus reducing the effect of refraction at the pore/material interface. Fluoropor-SLIPS surfaces made by the infusion are transparent (FluoroporSLIPS 10, Fluoropor-SLIPS 15, Fluoropor-SLIPS 20) or translucent (Fluoropor-SLIPS 25, Fluoropor-SLIPS 30). The influence of porosity and surface roughness on the formation of a biofilm was investigated. We tested the longterm stability and the antifouling properties of FluoroporSLIPS by characterizing long-term biofilm growth using P. aeruginosa as the model bacteria. The experiments were conducted within a microfluidic flow cell using the FluoroporSLIPS substrates as the bottom of the flow cell. As a reference, glass slides were used to determine biofilm formation on noncoated substrates. After an incubation period of 7 days, the biofilm was assessed by fluorescence staining. The percentage coverage of the biofilm was evaluated by image analysis. For the statistical data evaluation, we used a two-sided Student’s ttest, and the results show that Fluoropor-SLIPS 10, FluoroporSLIPS 15, and Fluoropor-SLIPS 20 have a significant reduction of biofilm growing on the surface (see Figure 3). The data does not include biofilm growth on rough, noninfused Fluoropor surfaces. This is because it was not possible to analyze the biofilm on noninfused Fluoropor samples. Because of the porosity of the noninfused Fluoropor samples, the dye solution
nm. The average pore radii of samples Fluoropor 10, Fluoropor 15, and Fluoropor 20 are very close to that instrumental limit. It is likely that small variances in these values cannot be effectively resolved by the instrument. To further characterize the properties and differences of the samples, the surface roughness was evaluated by white light interferometry measurement (see Figure 1d). Examination of Rq values reveal that for increasing cyclohexanol content, the roughness increases until it reaches saturation for Fluoropor 30. The porosity and roughness of the samples influences the optical transmission of the samples. Optical transmissions decrease with increasing cyclohexanol content in the polymerization mixture (see Figure 2a). This is mainly due to the increase in pore size and diameters of the polymer globes, which result in light scattering. While Fluoropor 10 has a similar transmission as nonporous Fluoropor, the transmission decreases significantly for Fluoropor 15 and Fluoropor 20. Fluoropor 25 and Fluoropor 30 are nontransparent (see Figure 2c). The dramatic decrease in transmittance between samples Fluoropor 10, Fluoropor 15, and Fluoropor 20 can be explained by the large influence of the surface roughness on transmittance measurements. We have previously shown32 that scattering effects at the rough surface have a major impact on transmittance. After production, Fluoropor samples are smooth because of the wetting of the polymer foil covering the liquid samples during polymerization. After removal of the smooth top layer, transmittance decreases significantly, for example, the transmittance of a sample equivalent to Fluoropor 15 is decreased by 15%. For comparison, an image of nonporous Fluoropor is provided in the Supporting Information (see Figure S1). D
DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
Research Article
ACS Applied Materials & Interfaces
entering the pores so that an undeterminable amount of bacteria remains in the sample after washing. For realistic applications, SLIPS with long-term stability of the slip effect are required. Given our data, we reasoned that the roughness of the substrates will be a major influence on the long-term stability of the lubricant layer. White light interferometry data show that infusing the porous polymers (see Figure 4a) with a lubricant produces the desired perfectly smooth Fluoropor-SLIP surfaces (see Figure 4b). After seven days under shear stress in a flow chamber, the surfaces have lost most of the lubricant, exposing the underlying roughness of the polymer (see Figure 4c). In Figure 4e, the measured white light interferometry data Rq are shown. Fluoropor-SLIPS 10 and Fluoropor-SLIPS 15 still display lower roughness than the original noninfused samples, which is in good accordance with the observation that biofilm growth was significantly reduced after seven days of the flow experiment. For Fluoropor-SLIPS 20, Fluoropor-SLIPS 25 and FluoroporSLIPS 30 Rq values are increased after the flow experiment. More interestingly, the acquirable data point coverage in white light interferometry is significantly decreased, resulting in an image with a significant amount of black pixels. Black pixels represent regions of the substrate which do not backscatter light and are therefore not accessible to interferometric measurement. Compared to the noninfused Fluoropor samples which are strongly light scattering and thus yield a very good contrast in white light interferometry the index match between the lubricant of the porous polymers result in significantly more unidirectional reflection and scattering of the light which is not picked up by the low numeric aperture of the objective. This results in loss of information and thus a higher overall
Figure 3. Biofilm growth on Fluoropor-SLIPS evaluated by image analysis surface coverage. Biofilm growth was significantly reduced for Fluoropor-SLIPS 10, Fluoropor-SLIPS 15, and Fluoropor-SLIPS 20 coatings on glass slides compared to a noncoated glass slide. Fluoropor-SLIPS 25 and Fluoropor-SLIPS 30 did not show any significant reduction of biofilm growth on the surface (mean ± standard deviation, *t-test: p ≤ 0.05, **t-test: p ≤ 0.001).
enters the material and cannot be removed by washing. This results in a very bright background signal and prevents the detection of the biofilm. Fluorescence images of a Fluoropor 20 and Fluoropor-SLIPS 20 sample after biofilm formation is provided in the Supporting Information to emphasize the problem of high background staining signal (see Figure S2). It was tried to quantify the biofilm by washing it off and measuring the optical density of the supernatant. However, no reliable data could be achieved. This is likely due to the biofilm
Figure 4. (a) White light interferometry analysis of Fluoropor, (b) freshly prepared liquid-infused Fluoropor-SLIPS, (c) same Fluoropor-SLIPS used in a high shear-force flow experiment (duration: seven days), and (d) a low shear-force shaking experiment (duration: seven days). All white light interferometry images have the same scale bars. (e) Results of the measured white light interferometry data Rq and corresponding data points are listed in the table. Rq values cannot be reliably determined in images with data coverage below 90% and have therefore been excluded. E
DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
Research Article
ACS Applied Materials & Interfaces
Figure 5. Fluorescence images and the corresponding threshold-adjusted 8-bit images of biofilm formation on Fluoropor-SLIPS to enhance visibility. Compared to the glass slide, the formation of the biofilm is observed only on single spots is clearly visible. As expected, biofilm formation initiates via bacterial attachment to singular spots of the underlying substrate topography, which penetrates the lubricant layer. For FluoroporSLIPS 30, the loss of lubricant along the flow lines is visible. It is along these exposed areas of the substrate that bacterial attachment and thus biofilm formation initiates. Scale bars: 150 μm.
submerged in water (see Figure 4d). It is clearly visible that even upon low shear forces, the surface begins to expose its highest elevated bumps, which represent ideal attachment spots for the early stage of bacterial adhesion, ultimately resulting in the formation of biofilms. To elucidate the attachment phase of the biofilm on Fluoropor-SLIPS, we investigated the biofilms formed after seven days. When the stained biofilms on Fluoropor-SLIPS are observed under the microscope (see Figure 5), the attachment of bacteria to isolated spots on the surface is observed compared to a widespread biofilm formation on the glass reference. In Fluoropor-SLIPS 30, the flux lines of the lubricant caused by the shear flow are clearly visible by the attachment pattern of bacteria. Thus, the surface roughness as well as the porosity of the substrate plays an important role in biofilm formation on Fluoropor-SLIPS. We studied the degree of degradation in a low shear force experiment by evaluation of the weight of the samples before and after shaking in water for 7 days. We found the loss of lubricant to be 3.4, 2.9, 3.7, 3.2, and 1.4 wt % (relative to the amount of oil infused into the sample). At high shear force, the lubricant layer is largely removed after eight days, evaluated by staining of the sample (see Figure S4).
amount of black pixels. Pore size of all samples were determined by mercury intrusion porosimetry. Median pore sizes for Fluoropor 10−30 are: 8, 7, 6, 304, and 520 nm. Mean pore sizes for Fluoropor 10−30 are: 7, 6, 6, 35, and 43 nm. This indicates that samples Fluoropor 10, 15, and 20 possess very small pore sizes with a lean distribution of sizes, while Fluoropor 25 and Fluoropor 30 possess a broad distribution of pore sizes with a lean distribution of small pores and a broad distribution of large pores. The increasing amount of oil-filled small pores from Fluoropor-SLIPS 10 to Fluoropor-SLIPS 20 lead to an increase in missing data points and therefore an increase in black pixels. The oil from the larger pores in samples Fluoropor-SLIPS 25 and Fluoropor-SLIPS 30 is likely removed by high shear forces due to a lower capillary pressure and a reduced curvature of the meniscus in larger pores compared to smaller pores. Thus, the underlying distribution of smaller pores in samples Fluoropor-SLIPS 25 and Fluoropor-SLIPS 30 is exposed in the white light interferometry measurements. With the number and size of pores increasing between these two samples, there is an increased loss of data in Fluoropor-SLIPS 30, and the image therefore appears darker. The elevated structures in Fluoropor-SLIPS 10 and Fluoropor-SLIPS 20 after the low shear force experiments are a result of small Fluoropor particles on the surface (see Supporting Information S3). Because of the flexibility of Fluoropor, the low Vickers hardness (1.19 HV at 100 mN/20 s) and the low coefficient of friction (μ = 0.2), which have been previously reported,32 the particles are likely a result of opening the Fluoropor surface by abrasion with sandpaper. The surface charge of the material makes these particles very sticky, and they may only be removed by the hydrophobic oil at high shear force. The increase in surface roughness correlates well with the increase in biofilm surface coverage, indicating that a smooth surface can more effectively reduce the growth of a biofilm. Because biofilm formation commences with the attachment of single bacteria,36,37 it is likely that the early colony formation takes place before extensive loss of the lubricant. Thus, to simulate the early stage of lubricant removal, Fluoropor-SLIPS were subjected to a low shear force experiment, where the surfaces were gently shaken while
4. CONCLUSIONS The formation of Fluoropor-SLIPS offers the unique advantage of a universally applicable substrate coating which can be turned into a SLIPS by infusion. It is thus both highly suitable for coating applications in a wide variety of applications as well as for studying the underlying mechanics of bioadhesion as it allows convenient access to different physical substrate structures. In this study, we investigated the formation of biofilm of Fluoropor-SLIPS. We showed that the porosity and roughness increase with increasing content of porogen in Fluoropor mixture. Fluoropor-SLIPS are suitable for generating transparent SLIPS and thus enable applications where transparent substrates are required. As the Fluoropor technology is applicable for a large selection of substrate materials, the Fluoropor-SLIPS technology is equally broad in the application range. We developed Fluoropor-SLIPS F
DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
Research Article
ACS Applied Materials & Interfaces
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substrates with different porosities and characterized their long-term stability against biofouling. Fluoropor-SLIPS can significantly reduce the formation of a biofilm on the substrates over a time period of seven days. It was shown that a lower surface roughness of Fluoropor resulted in a more effective reduction of biofilm attachment on Fluoropor-SLIPS. This can be attributed to the attachment of biofilm on topographic features which penetrate the lubricant layer, thus forming an ideal spot for bacterial adhesion.
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.8b12542. SEM analysis of the nonporous Fluoropor. Fluoropor particles on the surface of Fluorpor-SLIPS. Comparison of background staining of Fluoropor and FluoroporSLIPS. Degree of degradation of the lubricant layer (PDF)
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. ORCID
Bastian E. Rapp: 0000-0002-3955-0291 Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was funded by the German Ministry of Education and Research (BMBF), funding code 03X5527 “Fluoropor”. REFERENCES
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DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
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DOI: 10.1021/acsami.8b12542 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX
