Nanotopography impedes Staphylococcus aureus biofilm without deforming the cell wall

Introduction:: There is an alarming rise in untreatable medical device-associated infections and in the emergence of antibiotic-resistant bacterial strains. These, coupled together, have made control of implant infection progressively challenging and costly. Microbial infection on medical devices starts with microbial adhesion and progresses through biofilm formation. Anti-biofilm strategies based on chemical coatings that affect initial adhesion, kill microbes, or inhibit their growth have shown limited long-term efficacy. Non-toxic biophysical approaches based on nanoscale topographies that hinder attachment or microscale topographies that restrict biofilm development function over long timescales. It is currently thought that the early adhesion of fungal and Gram-negative bacterial pathogens is guided by a balance between energy gain due to interface formation (adhesion) and expenditure due to cell wall deformation (stretching)^{[1, 2]}. Surface nanostructures that maximize the change in the total free energy of adherent cells are the least favorable for adhesion and show the least cell retention^[3]. However, the effect of nanotopography on the adhesion of rigid and non-deforming Gram-positive bacteria is less understood. We hypothesize that in absence of microscopically detectable cell deformation, nanotopography-induced mechanical strain of bacterial cell walls hinders microbial attachment through mechanobiological regulation of the adherent phenotype.


Materials and Methods:: Inspired by the antifouling properties of the Mediterranean mussel's surface, we used our non-electrospinning Spinneret Tunable Engineered Parameter (STEP) to develop aligned and regularly-spaced nanotopographies on surfaces (Fig 1A). In contrast to other surface coating methods, our technique enables independent control of the fiber diameter and spacing and the scalable deposition of the highly aligned nanofiber coating on 3D objects. To test our hypothesis, we deposited polystyrene STEP nanofibers (diameter: 300 and 750 nm; spacing: 900 nm) on flat polystyrene-coated glass surfaces to obtain specific nanotopography. Polystyrene-coated glass substrates (without nanofibers) were used as control. We integrated each substrate type in a polydimethylsiloxane (PDMS) based microfluidic assay device to discern the effect of surface nanotopography on S. aureus under static and physiologically relevant shear flow conditions (0 - 6.9 x 10^-2 dyne/cm², Fig 1A). To study biofilm growth dynamics over an extended timescale (~24 h), we opted for minimal media (M9 supplemented with CaCl₂, MgSO₄, glucose, and peptone). We selected S. aureus as the model microorganism in this study, as it is the most common etiological agent for healthcare-acquired infections (HCAIs), with the highest mortality rate in the US since 2007^[4]. We studied the effect of shear flow on initial attachment density, biofilm growth, and antibiotic susceptibility. Fluorescence and confocal microscopy are used to study the biofilm growth and morphological features. The biofilm is quantified by ImageJ software. The computational modeling of S. aureus on nanofiber-coated surfaces was performed using our previously developed model^[1].

Results, Conclusions, and Discussions:: Our previously developed experimentally-validated computational model^[1] predicts minimal differences in the binding energy for S. aureus on different nanotopographies (Fig 1B). Moreover, according to the attachment point theory^[3], the strength of cell adhesion to textured surfaces is regulated by the number of attachment points. Thus, we did not expect surface topography dependency in S. aureus attachment. Indeed, our experiments showed no significant difference in bacteria's initial adhesion (for 2h) on uncoated and fiber-coated surfaces under static and shear flow (Fig 1C.i). However, at 6h, topography-dependent differences emerged, wherein less growth was observed on fiber-coated surfaces than on uncoated ones under static and shear flow (Fig 1C.ii). Thus, we hypothesized that activation of mechano-sensitive channels, in the absence of cell wall deformation, regulates the binding of S. aureus bacteria on nano-textured surfaces. We are currently testing our hypothesis using wild-type and ΔMscL mutants. Overall, the nanotextured surface not only slows down the growth of biofilm (Fig 1D.i-ii) but also affects the 3D morphological makeup of the biofilms, as shown by the confocal microscopic images (Fig 1E and 1F). Our findings emphasize the role of mechanical forces on the nano-textured surfaces in biofilm formation and eradication. We are currently evaluating how altered biofilm architecture is susceptible to the commonly employed antibiotics for orthopedic implant infections (OII) infections. These observations suggest that there is a need to look beyond the attachment point theory and develop an understanding of how mechanobiological effects regulate microbial adhesion and biofilm formation. Such advancement in knowledge will help design medical device surface with tailored nanotopography to counter the challenges of biofilm-associated HCAIs.

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References (Optional): : [1] Z. Ye, A. Kim, C. Y. Mottley, M. W. Ellis, C. Wall, A. R. Esker, A. S. Nain, and B. Behkam, "Design of Nanofiber Coatings for Mitigation of Microbial Adhesion: Modeling and Application to Medical Catheters," ACS Applied Materials and Interfaces, vol. 10, no. 18, pp. 15477–15486, 2018, doi: 10.1021/acsami.8b02907.

[2] M. Kargar, J. Wang, A. S. Nain, and B. Behkam, "Controlling bacterial adhesion to surfaces using topographical cues: a study of the interaction of Pseudomonas aeruginosa with nanofiber-textured surfaces," Soft Matter, vol. 8, no. 40, pp. 10254–10259, 2012, doi: 10.1039/c2sm26368h.

[3] H. E. Scardino. A.J. de Nys. R., "Testing attachment point theory: diatom attachment on microtextured polyimide biomimics," Biofouling, vol. 22, pp. 55–60, 2006. doi: 10.1080/08927010500506094

[4] K. W. Lobdell, S. Stamou, and J. A. Sanchez, "Hospital-Acquired Infections," Surgical Clinics of North America, vol. 92, no. 1, pp. 65–77, 2012, doi: 10.1016/j.suc.2011.11.003.