Studying the Influence of Surface Topography on Bacterial Adhesion

Article pubs.acs.org/Langmuir

Studying the Influence of Surface Topography on Bacterial Adhesion using Spatially Organized Microtopographic Surface Patterns David Perera-Costa,† José Morales Bruque,†,§ María Luisa González-Martín,†,§ Antonio Cándido Gómez-García,‡,§ and Virginia Vadillo-Rodríguez*,†,§ †

Department of Applied Physics, ‡Department of Biomedical Sciences and §Biomedical Research Networking Center in Bioengineering, Biomaterials and Nanomedicine (CIBER-BBN), University of Extremadura, Avda de Elvas s/n, 06006 Badajoz, Spain

ABSTRACT: The influence of surface topography on bacterial adhesion has been investigated using a range of spatially organized microtopographic surface patterns generated on polydimethylsiloxane (PDMS) and three unrelated bacterial strains. The results presented indicate that bacterial cells actively choose their position to settle, differentiating upper and lower areas in all the surface patterns evaluated. Such selective adhesion depends on the cells’ size and shape relative to the dimensions of the surface topographical features and surface hydrophobicity/hydrophilicity. Moreover, it was found that all the topographies investigated provoke a significant reduction in bacterial adhesion (30−45%) relative to the smooth control samples regardless of surface hydrophobicity/hydrophilicity. This remarkable finding constitutes a general phenomenon, occurring in both Grampositive and Gram-negative cells with spherical or rod shape, dictated by only surface topography. Collectively, the results presented in this study demonstrate that spatially organized microtopographic surface patterns represent a promising approach to controlling/inhibiting bacterial adhesion and biofilm formation.

1. INTRODUCTION In their natural state, bacterial cells tend to form self-organized multicellular communities called biofilms that typically develop at the solid−liquid interface of a large variety of living and inanimate substrata.1 The formation of these communities poses a serious problem in many industrial, environmental, and medical applications. Biofilms on ship hulls, for instance, which are usually followed by a progressive accumulation of larger marine organisms and seaweed, noticeably increase hydrodynamic drag and fuel consumption.2 Similarly, biofilm formation on heat exchangers, filters, pipelines, or separation membranes opposes heat and mass transfer and increases frictional resistance.3 These consequences clearly result in decreased production rates and increased costs in many industrial settings. In the medical field, the association of biofilms with synthetic biomaterials used for, for example, artificial organs, joint replacements, or voice and vascular prostheses, as well as for extracellular devices such as catheters or blood bags, is at the basis of severe and recurrent infections. Infected extracorporeal devices are routinely replaced worldwide and patients with, for instance, prosthetic infections, suffer from recurrent surgical interventions that come at a high cost for the patients, their families, and the healthcare systems.4,5 It has long been recognized that an effective strategy to deal with surface associated biofilms is to develop materials with antimicrobial surface properties. Accordingly, scientists have traditionally relied on surface modification methods to © 2014 American Chemical Society

chemically alter the surface properties of the materials. Hydrophilic poly(ethylene glycol) brush-like surface coatings, for instance, have been shown to successfully prevent the adhesion of a wide range of bacterial species; their brush-like structure provides a steric barrier, i.e., a physical separation between the cells and the underlying substratum, that effectively hinders adhesion.6,7 Natural and synthetic antimicrobial molecules such as cationic peptides, silver ions, quaternary ammonium compounds, or antibiotics have also been successfully incorporated in a large variety of engineering polymers and other materials surfaces.8−10 These compounds, able to penetrate into the cells and interfere with vital intracellular processes, are typically released into the surrounding aqueous environment using physically or chemically controlled release systems. These strategies, however, are transient: the nonspecific adsorption to the material surface of proteins and many other molecules secreted by the cells eventually masks the underlying chemical functionality and, on the other hand, the number of surface incorporated antimicrobial molecules is normally finite and subject to depletion. In addition, the manifestation of more and more multidrug-resistant bacterial strains highly compromises the effectiveness and use of antimicrobial and biocidal comReceived: January 9, 2014 Revised: April 3, 2014 Published: April 3, 2014 4633

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pounds.11−13 Therefore, materials that persistently withstand bacterial adhesion seem to be difficult to achieve by chemical surface modifications alone. Surface physical attributes may provide a more persistent form of inhibiting bacterial adhesion. In Nature, a variety of examples can be found in which a combination of physical and chemical surface characteristics effectively hampers biofilm formation. Marine organisms such as mussels, sharks, crabs, and skates possess at their most outer surface microscopic ridges and grooves that, combined with proteins and polysaccharides secreted by its constituent cells, confer to these natural surfaces antifouling properties.14,15 The inner surface of blood vessels constitutes another example: the distinct cobblestone-like topography of this layer, together with the bioactive compounds expressed by the cells that composed the endothelium, create an ideal anti-thrombogenic surface that genuinely resists the constant presence of fouling proteins and cells.16 Numerous experimental studies have also demonstrated that eukaryotic cells actively respond to surface topography, and that cellular processes such as spreading, proliferation, differentiation, and apoptosis highly depend on the spatial confinement of the cells.17−19 It has been shown, for instance, that cells seeded on parallel ridges align along the direction of the ridges during growth as long as the periodicity of the surface structures is above a critical value that is cell-type dependent.20,21 In addition, regularly spaced microfibers in contact with blood have been found to promote the formation of a stable pseudo-neointima, on which subsequent cellular migration and tissue healing was significantly enhanced.22,23 Moreover, patterned surfaces with square pillars of submicrometer dimensions have been shown to markedly reduce the adhesion of platelet from plasma at low shear stresses.24 The influence of surface topographical features on bacterial adhesion, however, remains poorly understood. Surface roughness (Ra), the most commonly used parameter to define surface topography in microbiological publications, does not appear to impact bacterial adhesion consistently. Taylor et al., for example, have reported that, while a slight increase in surface roughness (0.04 < Ra < 1.24 μm) promotes bacterial retention, a further increase (1.86 < Ra < 7.89 μm) results in lower retention values.25 Similarly, plate formation and maturation have been found to be encouraged on rough compared to similar polished surfaces.26,27 However, a few other studies working on a similar range of Ra values have shown no apparent correlation between surface roughness and bacterial adhesion.28,29 It is important to consider that the aforementioned studies involved different material types whose randomly roughened surfaces were mostly undefined in topographical terms. In particular, the morphological characteristics of the surfaces examined were solely described by the parameter Ra, which is typically defined as the arithmetic mean distance between surface peaks and valleys measured along a center line.30 For stainless steel, for instance, it has been shown that different finishes produce surfaces with different topographies while retaining similar values of Ra.31 Therefore, to truly assess how surface topography and specific surface features affect bacterial adhesion, surfaces with regular, well-defined topographical patterns and constant chemistry are essentially necessary. The purpose of this study is to investigate the influence of surface topography on bacterial adhesion by means of surfaces that contain spatially organized patterns of distinct microscopic topographical features. These surfaces were fabricated using

soft lithography and polydimethylsiloxane (PDMS), an elastomer widely used in pharmaceutical and medical applications.32,33 The experiments were performed with Staphylococcus epidermidis, one of the leading pathogens of nosocomial infections,34,35 and Escherichia coli and Bacillus subtilis. The last two strains, naturally found in the colon of virtually all warm-blood mammals and in soil and vegetation, respectively,36,37 were included to test whether the results obtained were limited to specific strains, cell shapes, or Gramstain type.

2. MATERIALS AND METHODS 2.1. Sample Preparation. PDMS surfaces with spatially organized microtopographic patterns were fabricated via soft lithography using two commercially available silicon masters purchased from Budget Sensors (Bulgaria), i.e., HS-20MG and HS-100MG. The masters feature silicon dioxide structure arrays on a 5 × 5 mm2 silicon chip fixed onto a 12 mm metal disc. Particularly, in the center of the chips, a 1 × 1 mm2 surface area comprises protruding and receding square and circular features of 10 and 5 μm pitch, respectively, and parallel channels with a 5 μm pitch. The configuration of the patterns, similar in both masters, is schematically depicted in Figure 1. The depth/

Figure 1. Schematic representation of the distinct topographies that comprise the silicon masters employed. On each master, protruding and receding square and circular features of a 10 and 5 μm pitch, respectively, and parallel channels with a 5 μm pitch, are found deposited on a 1 × 1 mm2 surface area as shown in this figure. height of all features was 21.1 nm (HS-20MG) and 117 nm (HS100MG), according to manufacturer’s specifications. Prior to use, the silicon masters were cleaned by sonicating for 5 min in 2% DSF surfactant solution in water (Derquim DSF 11, Panreac Quimica, S.A., Spain), rinsed thoroughly with demineralized water, again sonicated in ethanol for a period of 5 min, and finally, dried with nitrogen. The replication process, previously described elsewhere,38 consisted of mixing the elastomer base and curing agent (Sylgard 184 PDMS, Dow Corning Corporation, Ellsworth Adhesives, Spain) at a weight ratio of 10:1. After extensive mixing (∼20 min), the mixture was degassed in a vacuum desiccator for 30 min and carefully poured on the silicon masters, which had been previously fixed to the bottom half of a small Petri dish, to produce 4-mm-thick samples. The elastomer was then thermally cured for 1 h at 60 °C. After cooling, the PDMS molds were carefully peeled off, punched out with a circular die cutter of 12 mm in diameter, and stored at room temperature in a desiccator until use. A set of these samples were exposed to UV/O3 for a period of 4 h using a commercial 50 W UV/ozone cleaning system (PSD-UV Ozone System, Novascan Technologies, Inc., USA). This period was sufficient to guarantee a complete hydrophilization of the PDMS surfaces. In particular, the samples were positioned 2 cm from a light source predominantly emitting at a wavelength of 254 nm, receiving an estimated intensity of about 2.5 kW/m2. After the 4 h period, the 4634

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Figure 2. Optical representative images of the distinct topographical patterns transferred to PDMS from the HS-100MG and HS-20MG silicon masters. (A) and (B) illustrate protruding and receding square features, (C) and (D) protruding and receding circular features, and (E) parallel ridges and channels. As it is shown, the square and circular features are disposed in a two-dimensional square array. Optical view: ∼12 144 μm2. 2.4. Bacterial Strains and Adhesion Experiments. Three different bacterial strains were used in this study: Staphylococcus epidermidis ATCC12228 and Bacillus subtilis 168, both Gram-positive cells with an overall spherical and rod-shaped shape, respectively, and Escherichia coli K12, a Gram-negative rod-shaped cell. B. subtilis and E. coli were kindly supplied by Dr. John R. Dutcher, University of Guelph, Toronto, Canada. The three strains were maintained in blood agar plates and cultured in Trypticase Soy Broth (Becton Dickinson, and Company, USA) at 37 °C on a rotary shaker (150 rpm). S. epidermidis was cultured for 10 h, after which this culture was used to inoculate a second culture that was grown for 14 h to early stationary growth phase. E. coli and B. subtilis were only cultured for 16 h to reach the early stationary growth phase. Bacteria were harvested by centrifugation (5 min at 1000 g), washed three times with PBS, and resuspended in PBS at a concentration of 3 × 108 bacteria per milliliter. No sign of spore germination was detected after subjecting B. subtilis to the described growth conditions, as confirmed by the WirtzConklin stain. Prior to adhesion experiments, the PDMS samples were sterilized with ethanol, dried with nitrogen, and fixed into the wells of a sterile six-well culture plate. The wells were then filled with 10 mL of bacterial suspension and placed in an environmental chamber at 37 °C while being subjected to slight orbital shaking (20 rpm). After 30 min, the samples were carefully removed from the wells and dipped twice with PBS to remove loosely bound bacteria. It has been reported that the passage of an air−liquid interface results in an applied detachment force that amounts to ∼10−7 N.40 Therefore, solely the cells adhering to the surfaces with an adhesion force larger than 10−7 N are likely being accounting for in these adhesion experiments. Finally, the samples were air-dried and transferred to the optical microscope (Olympus BX41). For each sample, three images (field of view: 126.5 × 96 μm2) were recorded at randomly selected locations on each topographical region. Attached cells were quantified manually, distinguishing between the number of bacteria adhering to the top, bottom, and edge of the distinct topographical features with the help of regular square grids, grids of circles, and parallel channels drawn over the images. All experiments were performed in triplicate with separately grown bacterial suspensions and substrata surfaces. 2.5. Statistical Analysis. The statistical analysis was performed using two-tailed unpaired t tests and one-way ANOVA (GraphPad InStat v 3.05, GraphPad Software, Inc., USA). All data reported are mean ± SD. The confidential range selected was 95%, which gives statistical differences when p-value < 0.05.

samples were immediately immersed in demineralized water to effectively maintain the hydrophilicity of their surfaces until use ( 0.05) are detected between the density of cells adhering to the top or depressed areas (light areas in Figure 3A) as a function of the size and shape of the features, with the only exception being the lower values associated with the rodshaped cells adhering to the ridge topography (p < 0.05). The channel width on this last topography is larger than the cocci diameter (∼1 μm), but comparable to the longitudinal dimension of the rod-shaped cells (∼1 × 3 μm). It is thus expected that while cocci may easily settle within these surface features, rod-shaped bacteria may not, depending on cell alignment. A similar spatial restriction is initially anticipated for the topography with circular pillars whose interstitial space is of 2 μm, but results show otherwise. This fact likely relates to the degree of freedom of cell movement on each particular topography: while cell movement is only allowed on one coordinate direction in the channels between ridges, cells are permitted to move in two coordinate directions in the continuous and intersecting depressed grid formed between circular pillars. Moreover, the cells’ preference for attachment to the square and circular pits (on which cells are not allowed to move) rather than on the top area of the surfaces that contain these features could be associated with the larger 4637

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Figure 6. Total density of cells adhering to PDMS surfaces containing topographical features of (A) 115.6 nm height/depth and (B) 19.9 nm height/ depth. □ ↑ and □ ↓ represent the surfaces that comprise protruding and recessing square features, respectively, ○ ↑ and ○ ↓ protruding and recessing circular features, respectively, and ∥ ↑↓ ridges and channels. CONTROL refers to the smooth PDMS samples. For each topographical region, data (n = 12) were obtained from 3 to 5 independent samples. The data reported are mean ± SD.

Figure 7. Density of S. epidermidis cells adhering to the upper and lower area and total density of adhering cells (total area) on hydrophilic PDMS surfaces containing surface features of (A,B) 115.6 nm height/depth and (C,D) 19.9 nm height/depth. □ ↑ and □ ↓ represent the surfaces that comprise protruding and recessing square features, respectively; ○ ↑ and ○ ↓ protruding and recessing circular features, respectively; and ∥ ↑↓ ridges and channels. CONTROL refers to the smooth PDMS samples. For each topographical region, data (n = 12) were obtained from 3 to 5 independent samples. The data reported are mean ± SD.

percentage measured in this latter case is attributed to unfavorable interactions between the surface and cells oriented other than parallel to the channels, a behavior that, interestingly, is not observed when the height/depth of the ridges/channels is as low as 19.9 nm (Figure 6B). Examples of limiting adhesion through surface topography have previously been reported in the literature using other textured surface patterns. Inspired by the skin of sharks, for instance, Schumacher et al.45 developed through photolithographic techniques a particular topography composed of microsized ridges of various lengths that was found to markedly reduce the settlement of zoospores of the marine alga Ulva. Interestingly, the reduced number of attached Ulva spores and other marine cells to this particular and several other textured surfaces was found to inversely scale with the engineered roughness index (ERI), a dimensionless parameter accounting for the size, geometry, and spatial arrangement of the surface topographical

These results indicate that bacterial cells actively chose their position to settle depending on their size and shape relative to the dimensions and height/depth of the topographical features. Moreover, the described spatial disposition of the cells seems to be a general phenomenon dictated by topography, occurring in examples of both Gram-positive and Gram-negative bacteria, with a spherical or rod shape. 3.2.2. Total Number of Adhering Cells. Enumeration of the total number of adhering cells to the different topographical regions (Figure 6) illustrates that all surface patterns provoke a significant reduction (p < 0.05) in bacterial adhesion relative to smooth control surface. Interestingly, for all three strains and topographies investigated settlement density is reduced by a similar percentage that amounts to an average value of 28 ± 2%, with the only exception of the rod-shaped cells adhering to the 115.6 nm height/depth ridge topography on which the reduced overall adhesion is of 45 ± 3% (Figure 6A). The higher 4638

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features.46,47 The results here presented, however, did not show any direct correlation with this topographical index. The observed lack of trend likely relates to the fact that, unlike in the mentioned studies, the interstitial space between the examined features was not kept constant in all the topographies investigated and/or small enough to prevent cells from completely penetrating the recessed areas. Similarly, the attachment of barnacle larvae has been shown to be diminished by surface microtextures in the range of 50−100 μm that were achieved by molding PDMS against various mesh structures.49 Figure 6 also shows that S. epidermidis is the most adhesive (∼106 cells/cm2) of the three strains evaluated, whereas the rod-shaped cells, B. subtilis and E. coli, adhere in lower and similar numbers (∼10 5 cells/cm 2 ). This difference in adhesiveness is probably related to the distinct mechanisms (i.e., molecular physicochemical and ligand−receptor interactions) by which different bacterial species adhere to surfaces, and not to the particular shape and size of the cells investigated. Nevertheless, further experiments would be needed to corroborate this finding including a larger number of spherical and rod-shaped bacterial strains. Finally, the results presented also indicate that, for each particular strain and topography studied, cells adhere in similar numbers independently of the height/depth of the surface topographical features (Figure 6A vs B). This observation clearly demonstrates that the surface roughness parameter Ra (Table 1) is insufficient to adequately characterize the influence of surface topography on bacterial adhesion. 3.2.3. Role of Surface Hydrophobicity on the Spatial Distribution and Total Number of Adhering Cells. Surface wettability has been generally acknowledged to be a contributing factor in bacterial adhesion processes. In order to evaluate the influence of this surface parameter on the spatial distribution and overall adhesion of the cells, a set of the different patterned surfaces investigated was subjected to a previously reported hydrophilization process.39 Prior to adhesion experiments, hydrophobic (untreated) and hydrophilic (UV/ozone-treated) samples displayed an average water contact angle of 108 ± 7° and ∼0°, respectively, regardless of the presence or absence of the topographical surface patterns. Similarly, the determination of Wenzel’s roughness factor (r), which quantifies the effect of surface roughness in wettability,48 showed no differences in water contact angles between unpatterned and patterned surfaces for each of the topographical regions investigated (i.e., r ≅ 1). Interestingly, the results obtained for S. epidermidis show that the tendencies reported as a function of topography on the hydrophobic surfaces (described above) are qualitatively maintained on the hydrophilic ones (Figure 7). In quantitative terms, significant differences are detected in the ratio between the number of cells adhering to the lower and upper area associated with the distinct surface patterns. In particular, this ratio increases to 79 ± 11% for the topographies that contain topographical features of 115.6 nm height/depth (Figure 7A) and to 64 ± 3% for those that contain recessed features and channels of 19.9 nm depth (Figure 7C). Moreover, the percentage of cells adhering to the top of the protruded topographical features of 19.9 nm height was found to decrease to 18 ± 4% (Figure 7C). Regarding the total number of adhering cells, it is observed that surface hydrophilicity causes an significant decrease in cell adhesion (Figure 7B and D vs Figure 6A and B), in agreement with previous studies that have repeatedly shown that hydrophilic materials are more resistant to bacterial adhesion

than hydrophobic ones.50 Specifically, it is found that the overall reduction percentage, which amounts to 80 ± 5%, is independent of the topography (including the smooth samples) investigated, and thus, the influence of surface topography on the total adhesion relative the smooth control samples remains the same on hydrophobic (28 ± 2% reduction) and hydrophilic (29 ± 3% reduction) samples. Therefore, these results demonstrated that while surface hydrophobicity plays a significant (quantitative) role in the spatial disposition and total number of adhering cells, the overall effect of surface topography on bacterial adhesion is independent of surface hydrophobicity/hydrophilicity. Preliminary experiments with B. subtilis and E. coli reveal similar behavior in terms of this surface parameter. The data obtained were however disregarded, as they did not result in statistically significant differences, an observation that can be attributed to the low adhesiveness (markedly enhanced on hydrophilic surfaces) that characterize these strains.

4. CONCLUSIONS The results presented in this study suggest that bacterial cells are able to differentiate upper and lower areas in spatially organized microtopographic surface patterns. This selective adhesion depends on the size and shape of the cells relative to the dimensions and height/depth of the topographical features and on surface hydrophobicity/hydrophilicity. It is important to consider that tuning such parameters to control cell pattering on desired areas would, for instance, be advantageous to properly develop biosensors, bioelectronic devices, and cell based microsystem. Additionally, disrupting the natural packing arrangement of cells on a surface could potentially interfere with their associated cooperative behavior, mediated by cell− cell communications and other factors, and be used to control and/or discourage biofilm formation. The results presented also reveal that all the microtopographies investigated provoke a significant reduction (∼30−45%) in bacterial adhesion relative to the smooth control samples regardless of surface hydrophobicity/hydrophilicity. Importantly, this observation emerges as a consequence of a strictly structural property, and thus, it is likely to be persistent (i.e., not susceptible to masking or depletion). Collectively, the results presented in this study demonstrate that spatially organized micro/nanotopographic surface patterns appear to be a novel and successful approach to inhibiting/controlling bacterial adhesion and biofilm formation.

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*Tel.: (+34) 924 289300 ext. 86173. Fax: (+34) 924 289651. E-mail address: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors gratefully acknowledge financial support from the “Ministerio de Ciencia e Innovación” (Grant No. MAT201237736-C05-03) and the “Gobierno de Extremadura-FEDER: European Regional Development Fund” (Grant No. GR10149 and GR10031). V.V.R. thanks the “Ministerio de Ciencia e Innovación” for the Ramón y Cajal fellowship (RYC-200803482). 4639

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