Influence of the Available Surface Area and Cell Elasticity on Bacterial Adhesion Forces on Highly Ordered Silicon Nanopillars

Abstract

Initial bacterial adhesion to solid surfaces is influenced by a multitude of different factors, e.g., roughness and stiffness, topography on the micro- and nanolevel, as well as chemical composition and wettability. Understanding the specific influences and possible interactive effects of all of these factors individually could lead to guidance on bacterial adhesion and prevention of unfavorable consequences like medically relevant biofilm formation. On this way, the aim of the present study was to identify the specific influence of the available surface area on the adhesion of clinically relevant bacterial strains with different membrane properties: Gram-positive Staphylococcus aureus and Gram-negative Aggregatibacter actinomycetemcomitans. As model surfaces, silicon nanopillar specimens with different spacings were fabricated using electron beam lithography and cryo-based reactive ion etching techniques. Characterization by scanning electron microscopy and contact angle measurement revealed almost defect-free highly ordered nanotopographies only varying in the available surface area. Bacterial adhesion forces to these specimens were quantified by means of single-cell force spectroscopy exploiting an atomic force microscope connected to a microfluidic setup (FluidFM). The nanotopographical features reduced bacterial adhesion strength by reducing the available surface area. In addition, the strain-specific interaction in detail depended on the bacterial cell's elasticity and deformability as well. Analyzed by confocal laser scanning microscopy, the obtained results on bacterial adhesion forces could be linked to the subsequent biofilm formation on the different topographies. By combining two cutting-edge technologies, it could be demonstrated that the overall bacterial adhesion strength is influenced by both the simple physical interaction with the underlying nanotopography and its available surface area as well as the deformability of the cell.

Figure 1

Results of sample characterization. (a,b) Scanning electron micrographs of fabricated nanopillar arrays with their geometrical configuration. A0, blank silicon surface; A1, hexagonal grid size of 200 nm; A2, 300 nm; A3, 400 nm. All pillar structures have nominal diameters of 100 nm and heights of 500 nm. (c) Resulting normalized available surface area on top of the structures. (d) Measured contact angles showing hydrophilic behavior on blank silicon (A0), while more hydrophobic behavior occurs on the structured samples (A1–A3). (e) Droplet placed on the transitional area of the structured and unstructured areas to demonstrate the resulting meniscus in the three-phase interface indicating movement due to nanocapillary forces. (f) Light microscopic image of a droplet taking a hexagonal shape, induced by the underlying nanotopography.

Figure 2

Bacterial single-cell adhesion forces on different nanotopographies. (a) Representative force–distance curves of single bacterial cells of indicated strains on different nanotopographies after a 5 s adhesion time. From force–distance curves after 5 and 10 s adhesion times, the deepest peak was quantified as the maximum adhesion force, the number of peaks as attachment points, and the distance until the curve returns to the baseline as the detachment distance. The results are given as Tukey boxplots for S. aureus (b) and A. ac (c). Asterisks (*) indicate statistically significant differences with p ≤ 0.05 between groups (black brackets) and between time points (gray brackets).

Figure 3

Interaction of bacteria with the nanostructured surfaces. SEM images of A3 structures demonstrate (a) rigid S. aureus sitting on top without any sign of deformation and (b) A. ac with a large deformation, partly sunken into the nanotopography. (c) Young’s modulus for S. aureus and A. ac obtained from the approach force–distance curves reflecting the bacterial stiffness. The asterisk (*) indicates a statistically significant difference with p ≤ 0.05.

Figure 4

Initial bacterial attachment and viability after 5 h of incubation on different nanotopographies. Results are given as Tukey boxplots of attached colonies and the mean ± standard deviation of bacterial live/dead distribution for (a) S. aureus and (b) A. ac. Asterisks (*) indicate statistically significant differences with p ≤ 0.05. In (c), representative microscopic images of initial attached bacterial cells are shown. Living bacteria are stained in green, whereas dead bacteria are stained in orange/red. Scale bars = 50 μm.

Figure 5

Bacterial biofilm formation and viability after 24 h of incubation on different nanotopographies. Results are given as Tukey boxplots of the colonized area by the biofilm and the biofilm volume and the mean ± standard deviation of bacterial live/dead distribution for (a) S. aureus and (b) A. ac. Asterisks (*) indicate statistically significant differences with p ≤ 0.05. In (c), representative microscopic images of bacterial biofilms are shown. Living bacteria are stained in green, whereas dead bacteria are stained in orange/red. Scale bars = 50 μm.

Figure 6

Schematic drawing of the interaction mechanisms of bacteria on nanotopographies. (a) Rigid and less deformable bacteria like Gram-positive S. aureus show less contact to the nanostructured surface. (b) A deformable cell like Gram-negative A. ac increases its surface interaction by deformation and partly adapting to the nanotopography. Yellow dots indicate adhesion points.

Figure 7

Schematic illustration of parameters quantified from force–distance curves of bacterial adhesion force spectroscopy.