University of Groningen
Bacterial interactions with nanostructured surfaces
Hizal, Ferdi
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Publication date: 2017
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Hizal, F. (2017). Bacterial interactions with nanostructured surfaces. Rijksuniversiteit Groningen.
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CHAPTER
3
Staphylococcal Adhesion, Detachment
and Transmission on Nanopillared
Si Surfaces
44
A B S T R A C T
Nanostructured surfaces are extensively considered with respect to their potential impact on bacterial adhesion from aqueous suspensions or air, but in real-life bacteria are often transmitted between surfaces. Mechanistically, transmission involves detachment of adhering bacteria from a donor and adhesion to a receiver surface, controlled by the relative values of the adhesion forces exerted by both surfaces. We here relate staphylococcal adhesion, detachment and transmission to, from, and between smooth and nanopillared-Si surfaces with staphylococcal adhesion forces. Nanopillared-Si surfaces were prepared with pillar-to-pillar distances of 200, 400 and 800 nm. On smooth surfaces, staphylococcal adhesion forces, measured using bacterial-probe Atomic-Force-Microscopy, amounted to 4.4–6.8 and 1.8–2.1 nN (depending on the AFM-loading force) for Extracellular-Polymeric-Substances producing and non-EPS producing strains, respectively. Accordingly the EPS producing strain adhered in higher numbers than the non-EPS producing strain. Fractional adhesion forces on nanopillared-Si surfaces relative to the smooth surface ranged from 0.30-0.95, depending on AFM-loading force, strain and pillar-to-pillar distance. However, for each strain, the number of adhering bacteria remained similar on all nanopillared surfaces. Detachment of adhering staphylococci decreased significantly with increasing adhesion forces, while staphylococcal transmission to a receiver surface also decreased with increasing adhesion force exerted by the donor. In addition, the strain with ability to produce EPS was killed in high percentages and induced to produce EPS during transmission on nanopillared-Si surfaces, presumably by high local cell-wall stresses. This must be accounted for in applications of nanostructured surfaces: whereas killing may be favorable, EPS production may reduce antimicrobial efficacy.
45
I N T R O D U C T I O N
Biofilm formation occurs onto almost any surface, regardless of environment and whether of natural or synthetic origin. Once bacteria adhere to a surface and start to grow, they embed themselves in a matrix of extracellular polymeric substances (EPS), providing shelter against environmental challenges, such as detachment forces, antimicrobials and when in a living host, against the host immune system. In many environments, biofilm formation is desirable such as in water purification or soil remediation plants, and also the gastrointestinal tract including the oral cavity, is colonized by selected bacterial strains and species indispensable to human health.1 Nevertheless biofilms are detrimental in food industry, water purification plants, on ship hulls and other surfaces exposed to the marine environment. Although indispensable for health in some niches of the human body, bacteria are also causative to many infectious diseases that become harder to control with the growing number of antibiotic resistance strains. Moreover, biofilms on biomedical equipment and biomaterials implants and devices can yield high morbidity and even mortality to implant recipients.2,3 Prosthetic-joint infections for instance, place patients at risk of mortality exceeding the one of many cancers.4
Traditionally, biofilm formation is separated into four different steps:5 1) transport of the
bacteria towards a surface, 2) initial, reversible adhesion of bacteria to a surface, 3) bond-maturation towards more irreversible adhesion and 4) growth of adhering bacteria into
a biofilm. This separation of biofilm formation in sequential steps has focused research mainly on the prevention of adhesion of bacteria from a transport medium, such as an aqueous suspension fluid or air. However, in many environments, bacteria are transmitted from one surface to another,6–12 examples being transmission of bacteria from a contact lens to the cornea,6 from the hands of health care workers to biomedical equipment and biomaterial implants and devices,7 or amongst different surfaces involved in food packaging.8,9 Different bacterial strains and species can survive on skin, gloves, sponges, cloths, surgical tools, coins and other surfaces for prolonged periods of time (several days to months) after initial adhesion and without metabolic activity or growth.11–14 Transmission of pathogens from kitchen sponges to stainless steel surfaces8 ongoing to food was found to be between 20% to 100% depending on the bacterial species, the contact pressure, the period of transmission and environmental humidity.
The study of bacterial transmission between surfaces requires a totally different approach than the study of bacterial adhesion from an aqueous suspension fluid or air, because it includes a detachment step of adhering bacteria from the donor surface followed by an adhesion step to the receiver surface. This implies that whether or not adhering bacteria are transmitted from a donor to a receiver surface will depend on the magnitude of the adhesion forces exerted by the receiver relative to the forces exerted on the adhering bacteria by the donor surface. Weibull analysis of bacterial adhesion forces measured using bacterial probe atomic force microscopy (AFM)15 to different surfaces has proven predictive of
Staphylococcus aureus transmission between contact lenses and storage cases.16 In an analogous way, surface thermodynamics of the preference of Pseudomonas aeruginosa,
46
Staphylococci and Serratia for adhesion to a donor or receiver surface is also predictive of whether bacterial transmission will occur or not.16
Micro- and nanostructured surfaces are extensively considered with respect to their potential impact on bacterial adhesion.17–19 Staphylococcal adhesion was significantly reduced on pillar-patterned poly(ethylene glycol) hydrogels with interpillar spacings below 1.5 µm.20 Nanostructured (pitted) titaniumoxide-coated silicon (Si) surfaces showed a reduction in bacterial adhesion by 15% to 85%, depending on the bacterial species,21 while gold-coated nanorough surfaces demonstrated significant variations in the morphological, genetic and proteomic expressions of adhering Escherichia coli.22 Importantly, random and ordered nano/microstructures have a different impact on bacterial adhesion of Pseudomonas fluorescens and their alignment was hindered on well-defined surface structures but not on random structures.23
In this work, we relate staphylococcal adhesion and detachment to and from smooth and highly-ordered nanostructured Si surfaces as well as the transmission from a smooth donor surface to a nanostructured receiver surface and vice versa with staphylococcal adhesion forces to the different surfaces. To this end, nanopillared Si surfaces were prepared with different center-to-center distances between pillars and the same surface chemistry. Bacterial adhesion forces were measured using bacterial probe AFM. Staphylococci present an important group of bacteria that can be found as commensals of the human skin and through transmission on virtually all environmental surfaces. Despite being commensals of the skin, especially strains with an extensive ability to produce EPS can also be causative to many infectious diseases, including biomaterial implant and device associated ones.
M A T E R I A L S A N D M E T H O D S
Si Nanopillar Fabrication. The Si nanopillar fabrication process is schematically summarized
in Figure S1. Briefly, polished 4-inch (10-cm) silicon wafers were cleaned by acetone and de-ionized water and dried by N2 gas. Next, a negative-tone photoresist (NR-250P, Futurrex, Inc., Franklin, NJ, USA) layer was spin-coated on the Si wafers (Figure S1a). Before laser exposure to define pore patterns in the photoresist layer, spin-coated Si wafers were baked at 150oC for 1 min. The mirror and substrate angles of a Lloyd-mirror laser interference lithography system with two degrees-of-freedom24 were regulated in such a way that pattern periodicities were ranging from 200 to 800 nm. The samples were then exposed to linearly polarized He-Cd laser with a wavelength of 325 nm (Kimmon, Tokyo, Japan) by using the two degrees-of-freedom laser interference lithography system24 (Figure S1b) and the samples were baked on a hot plate at 100oC for 1 min. The photoresist layers were subsequently developed using resist developer, RD 6 (Futurrex Inc., Franklin, NJ, USA), diluted (3:1) with de-ionized water, for different time periods (15 – 25 s) to achieve the desired pore diameters, followed by rinsing with de-ionized water (Figure S1c). Finally, a chrome metal layer was deposited through the nanopatterned photoresist layer onto the Si wafer by e-beam evaporation (PVD75, Kurt J Lesker, Jefferson Hills, PA, USA) (Figure S1d). The deposition rate was 0.2 nm s−1, which resulted in uniform metal film
47 thickness (~50 nm). Then, the photoresist layer was removed in piranha solution (a mixture of H2SO4 (98 %) and H2O2 (95 %) with a ratio 3:1), leaving the deposited chrome metals on the Si surface serving as a mask in order to etch the Si wafers to obtain high-aspect-ratio nanopillars (Figure S1e).
For the deep reactive ion etching of Si with the chrome metals on top, a cryogenic etch mode with O2 and SF6 was used at –100°C (Plasmalab 100, Oxford Instruments, Abington, UK) (Figure S1f). The number of etching cycles in the cryogenic process was varied according to the etch depth desired. Finally, the chrome masks were removed in chrome etchant (MicroChemicals Inc., Ulm, Germany) at 30°C for 1 min, followed by water rinsing and N
2 drying (Figure S1g), yielding a fully water wettable surface (i.e. a zero degrees water contact angle). The specimens were then imaged with a SEM (FEI-SEM Quanta FEG450, FEI, Hillsboro, Oregon 97124 USA) to examine the surface topographies and uniformity of the nanostructures over the sample area. Finally, 1 cm × 1 cm samples were cut out of the fabricated nanopillared Si wafers and used in the assays.
Bacterial Culturing and Harvesting. S. epidermidis ATCC12228 (non-EPS producer) and
S. aureus ATCC12600 (EPS producer) were cultured from a frozen stock, stored in 7% DMSO at -80°C. Bacteria were inoculated on blood agar plates and grown overnight aerobically at 37°C and after growth maintained for a maximum period of 12 days in the refrigerator. Single
colonies from the agar plates were inoculated in 10 ml tryptone soya broth (TSB, OXOID, Basingstoke, UK) and cultured for 24 h. This pre-culture was then used to
inoculate a 200 ml main culture in TSB, which was grown for 16 h before harvesting. Bacteria were harvested by centrifugation at 5000 × g for 5 min at 10°C, and washed twice with potassium phosphate buffered saline (PBS, 10 mM potassium phosphate, 0.15 M NaCl, pH 7.0). Staphylococcal aggregates were separated by sonication on ice, three times for 10 s at 30 W (Vibra Cell model 375, Sonics and Materials Inc., Danbury, Connecticut, USA). Finally, staphylococci were suspended in PBS to a density of 3 × 108 bacteria per ml for adhesion experiments and 2 × 109 bacteria per ml for transmission experiments, as determined using a Bürker-Türk counting chamber.
Bacterial Adhesion Force Measurements. Bacterial adhesion forces on smooth and
nanopillared Si surfaces were recorded using AFM (BioScope Catalyst atomic force microscope with ScanAsyst [Veeco Instruments Inc., Camarillo, CA]). Before each measurement, tipless cantilevers (NP-O10, Bruker AFM Probes, Camarillo, CA) were calibrated by the thermal tuning method,44 yielding an overall average spring constant of 0.047 ± 0.004 Nm-1. Bacterial probes were prepared by immobilizing single bacteria on a cantilever using electrostatic attraction.45 All adhesion force measurements were performed in PBS at room temperature with z-scan rates of 1.0 Hz under a loading force of 2, 5 and 10 nN at 0, 10 and 30 s surface delays. Ten force curves per spot and five randomly chosen spots on the surfaces were measured per probe from which the maximal adhesion force upon retraction were recorded. To confirm that the
48
bacterial probe recorded single bacterial contact with a substratum (see also Younes et al.,46 for confirmation of the presence of single bacteria on thus prepared probes), images taken with a bacterial probe on nanopillared Si surfaces were always checked for double contour lines, indicative of multiple contact points. Double contour lines were never observed. To confirm that a bacterial probe was not damaged during previous measurements, a force curve with 0 s surface delay was measured on bare glass after completion of each sample measurement and compared to initial force curves recorded on glass prior to further measurements. For each of the triplicated measurements, a different bacterial culture was used and a new bacterial probe was prepared.
Bacterial Adhesion and Detachment in a Parallel Plate Flow Chamber.Bacterial adhesion on smooth and nanopillared Si surfaces was carried out in a parallel plate flow chamber (7.6 cm × 3.8 cm × 0.058 cm). The top glass and bottom polymethylmethacrylate (PMMA) plates of the chamber were placed in the middle of a stainless steel frame and separated by two Delrin spacers creating a smooth channel with gradually diverging and converging (62 degrees) inlet and outlet regions. The chamber plates were sonicated for 3 min in 2% RBS35 (Omnilabo International BV, Breda, The Netherlands) followed by rinsing with tap water, demineralized water, methanol, tap water and finally demineralized water. Prior to use, the flow chamber was washed with 2% Extran (Merck, Darmstadt, Germany) and rinsed thoroughly with tap water and demineralized water.
Inserts were made in the bottom PMMA plate of the flow chamber, to allow placement of four different Si surfaces in a direction perpendicular to the flow. The locations of the smooth, 200, 400 and 800 nm nanopillared Si surfaces were randomly interchanged at each of triplicated adhesion experiments to compensate for possible differences in conditions at different locations on the bottom plate. Before each experiment, the flow chamber and tubes
were first filled with PBS and all air bubbles were removed from the system. Next, the staphylococcal suspension (3 × 108 per ml) was perfused through the flow chamber for
30 min under hydrostatic pressure at a laminar flow rate of 1 ml/min corresponding with a shear rate of 6 s-1, while recirculating the suspension using a roller pump. After 30 min, flow was switched to buffer at the same flow rate to remove non-adhering staphylococci from the system. For enumeration, the adhering staphylococci were stained in the flow chamber with live/dead stain (BacLight, Invitrogen, Breda, The Netherlands) for 15 min in the dark. The stock staining solution was prepared in a mixture of 3.34 mM SYTO®9 nucleic acid stain and 20 mM propidium iodide (1:1 in volume) for live/dead (Green/Red) viability. Five fluorescent images at different spots were taken from each sample using fluorescence microscopy (Leica DM4000B, Leica Microsystems GmbH, Heidelberg, Germany). Finally, the total number of adhering live and dead bacteria were counted using ImageJ software.
For bacterial detachment experiments, after staphylococcal adhesion, the flow was switched to buffer at a higher shear of 10 ml/min which corresponds to a shear rate of 60 s-1 to
49 stimulate detachment of adhering bacteria. Flow was stopped after 1 h and bacteria still adhering to the surface were counted using fluorescence microscopy after staining.
Bacterial Transmission. Donor surfaces were placed into six-well plates and 2 ml of 2 × 109 of staphylococci per ml was added and after 5 min the bacterial solutions were removed and the samples were gently rinsed 10 times in the well plates with PBS and left in 4 ml PBS. Surfaces were kept wet at all times to avoid dehydration of bacteria. Then, a receiver surface which was placed with double sided sticky tape on the bottom surface of a 500 g weight, was carefully placed on the donor surface. After 5 min of contact between the donor and receiver surface, the receiver surface was removed and immediately submerged into PBS. The staphylococci on both donor and receiver surfaces were stained with live/dead stain for 15 min in the dark and the transmitted bacteria on the receiver surface and the remaining bacteria on the donor
surface were enumerated by fluorescence microscopy. In addition, the initial number of staphylococci on the donor surface was also determined by live/dead stain and fluorescence
microscopy.
SEM Imaging of Staphylococcal Transmission. Smooth and nanopillared Si surfaces with
and without adhering staphylococci were prepared for SEM prior to and immediately after transmission. After removal of PBS from the well plate in which transmission was allowed to occur, a fixation solution (1% paraformaldehyde and 2% glutaraldehyde in 0.1 M cacodylate buffer) was introduced for 1 h at room temperature after which surfaces were washed with 0.1 M cacodylate buffer, followed by 1 h of incubation at room temperature with 1% OsO4 in 0.1 M cacodylate buffer. Subsequently, samples were washed with demineralized water and dehydrated with 30, 50, 70% ethanol for 15 min each and three times with absolute ethanol for 30 min at 4oC. Finally, the samples were incubated in ethanol (100%) and tetramethylsilane (1:1) for 10 min, followed by 15 min incubation in pure tetramethylsilane and dried in air. Samples were sputter-coated with electrically-conducting metal (Au/Pd) to prevent charging
of the non-conducting section on the surface (i.e. bacteria) and analyzed in the SEM (FEI-SEM Quanta FEG450), using 5, 10 and 20 kV accelerating voltage under high vacuum.
R E S U L T S
Interference lithography is an optical lithography technique used to fabricate periodic nanoscale patterns with high throughput over a relatively large surface area and with high accuracy,24–26 as can be seen in Figure 1. For the current study, three different kinds of nanopillared Si surfaces were made with different center-to-center distances between pillars, ranging from 200, 400 to 800 nm. All pillars were 500 nm in height regardless of the distance between pillars, while concurrent with center-to-center distance, also the pillar diameters at the tip varied from 90 to 75 and 300 nm, respectively. The variance in pillar diameters was unavoidably connected with the variance in pillar-to-pillar distance due to fabrication process settings.
50
Figure 1. SEM micrographs of lithographically prepared nanopillars on Si surfaces with different
center-to-center distances between pillars. (a) 200 nm distance, (b) 400 nm distance, (c) 800 nm distance. Scale bars equal 500 nm.
Adhesion forces between the staphylococci and the smooth or nanopillared Si surfaces were determined by bacterial probe AFM (see Figure 2a) applying different surface delay times and loading forces before the bacterial probe and substratum surface were retracted to yield
the adhesion force (see Figure 2b for an example of a force-distance diagram). In Figures 2c and 2d it can be seen that in general 30 s sufficed to obtain stable adhesion forces
which is the reason why we only present adhesion forces obtained after 30 s surface delay (Figure 2e and Figure 2f).
Staphylococcal adhesion forces for the EPS producing strain (Figure 2e) are statistically weaker (p < 0.05) on nanopillared Si surfaces than on the smooth Si surface, regardless of the loading force applied. However, for the non-EPS producing strain only the adhesion force of the 200 nm pillared surface is statistically different from the smooth surface (Figure 2f). Stronger adhesion forces (p < 0.05) were measured for the EPS producing staphylococcal strain than for the non-EPS producing strain on all surfaces, that amounted to 4.4, 6.9, and 6.8 nN on the smooth surfaces for EPS producing S. aureus ATCC12600 and 1.8, 1.6, and 2.1 nN for non-EPS producing S. epidermidis ATCC12228 at the respective loading forces of 2, 5, and 10 nN. Table 1 summarizes the fractional adhesion forces for all loading forces and different nanopillared Si surfaces relative to the force measured on the corresponding smooth surface. For the EPS producing staphylococcal strain, fractional adhesion forces range from 0.30 to 0.67 and regardless of the pillar-to-pillar distance, are smallest under a loading force of 5 nN. Fractional adhesion forces for the non-EPS producing strain vary over a wider range from 0.39 to 0.95, but moreover are the largest under a loading force of 5 nN.
Staphylococcal deposition and adhesion on the smooth and nanopillared Si surfaces were
measured in a parallel plate flow chamber (Figure 3a), under the influence of convective-diffusional mass transport (Figure 3b). Detachment was stimulated by increasing
the fluid flow therewith increasing the wall shear rate. In line with its higher adhesion force, EPS producing S. aureus ATCC12600 adhered in higher numbers than non-EPS producing
S. epidermidis ATCC12228 (Figure 3c, 3e, 3g), regardless of the loading force under which adhesion forces were measured. However, in neither strains did the numbers of adhering
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Figure 2. Bacterial probe AFM and influence of surface delay time and loading force on measured
staphylococcal adhesion forces on nanopillared Si surfaces. (a) Schematics of bacterial probe AFM.
(b) Example of a force-distance curve for S. aureus ATCC12600 and nanopillared Si surface (400 nm pillar-to-pillar distance) under 10 nN loading force and after 30 s surface delay. (c) Influence of
surface delay on the adhesion forces between EPS producing S. aureus ATCC12600 under different loading forces on a nanopillared Si surface with 200 nm center-to-center distance. (d) Same as in panel c but for non-EPS producing S. epidermidis ATCC12228 and a nanopillared Si surface with 400 nm center-to-center distance. (e) Adhesion forces after 30 s surface delay for EPS producing S. aureus ATCC12600 under different loading forces on smooth and nanopillared Si surfaces with different center-to-center distances. (f) Same as in panel e but for non-EPS producing S. epidermidis ATCC12228. Error bars represent the standard deviations over three bacterial probes with separately grown bacteria and ten force-distance curves each taken at five different spots on 1 cm × 1 cm sample yielding a total of 150 measurements for each data point. Asterisks indicate statistical significance (p < 0.05) between indicated groups.
52
staphylococci change with fractional adhesion force, that is, the presence of nanopillars. Opposite to staphylococcal adhesion, fluid shear induced detachment of adhering bacteria from the smooth and nanopillared Si surfaces was affected by fractional adhesion forces. Detachment significantly decreased in a near-linear fashion with increasing fractional adhesion forces, for both the EPS producing and the non-EPS producing staphylococcal strain following the same relation (Figure 3d, 3f, 3h).
Table 1. Fractional adhesion forces exerted on an EPS producing and a non-EPS producing staphylococcal
strain by nanopillared Si surfaces with different pillar-to-pillar distances relative to the forces measured on a smooth Si surface using bacterial probe AFM with a 30 s surface delay and at different loading forces.
AFM-loading force EPS producing S. aureus ATCC12600 Non-EPS producing S. epidermidis ATCC12228 200 nm 400 nm 800 nm 200 nm 400 nm 800 nm 2 nN 0.67 0.49 0.54 0.45 0.58 0.62 5 nN 0.36 0.39 0.30 0.60 0.95 0.97 10 nN 0.49 0.45 0.44 0.39 0.81 0.59
Transmission between smooth and nanopillared Si surfaces was investigated by pressing a bacteria-free receiver surface with a pressure of about 50 kPa for 5 min to a donor surface with 6.5 × 106 adhering staphylococci per cm2 (Figure 4a) and expressing transmission as the percentage number of bacteria left on the donor surface (Figure 4b). The percentage of remaining bacteria on the smooth donor surface after transmission to different Si receiver surfaces either slightly decreased with fractional adhesion forces exerted by the nanopillared
Si receiver surfaces (Figure 4c and 4f), or remained unaffected under a loading force of 5 nN (Figure 4e). Generally, a slightly higher percentage number of EPS producing
staphylococci was transmitted as compared to non-EPS producing staphylococci. Alternatively, the percentage of remaining bacteria on different Si donor surfaces after transmission to smooth receiver surfaces increased slightly with fractional adhesion forces exerted by the nanopillared Si donor surfaces (Figure 4d and 4h), the smallest increase occurring at an AFM-loading force of 5 nN (Figure 4f).
The morphology of the staphylococci prior to and after transmission differed considerably (Figure 5). Note that staphylococcal morphologies after transmission were similar regardless of whether a surface was used as a donor or a receiver surface. The morphology of EPS producing S. aureus ATCC12600 on the smooth surface shows no traces of EPS before or after transmission. However, on the nanopillared Si surfaces after transmission there is clear EPS production that appears induced by high local pressures in the region between pillars and an adhering staphylococcus (compare Figure 5a-5d with 5e-5h). Most EPS production was observed on the 200 nm nanopillared Si surfaces (Figure 5f) and only minor EPS production on the 800 nm pillared surface (Figure 5h). SEM images of S. epidermidis ATCC12228 did not show any indication of nanoscopic pressure-induced EPS production before or after transmission
53
Figure 3. Adhesion and detachment of EPS producing (S. aureus ATCC12600) and non-EPS producing
(S. epidermidis ATCC12228) staphylococcal strains to and from nanopillared Si surfaces as a function of the fractional adhesion force obtained under different loading forces in AFM. Fractional adhesion forces were calculated with respect to the force measured on the smooth Si surface (see Table 1). Staphylococcal detachment was stimulated by increasing the shear rate tenfold during fluid flow in the parallel plate flow chamber from 6 to 60 s-1. (a) Schematics of the parallel plate flow chamber. (b) Convective-diffusional mass
transport in the parallel plate flow chamber in which diffusion brings bacteria in a flowing suspension
close to the surface under the influence of adhesion forces arising from the substratum surface. (c, d) Numbers of adhering staphylococci and percentage detachment as a function of the fractional
adhesion force measured under a loading force of 2 nN. (e, f) Same as in panels c and d but for 5 nN AFM-loading force. (g, h) Same as in panels c and d but for 10 nN AFM-loading force. Error bars denote the standard deviations over triplicate experiments, carried out with different bacterial cultures and separately prepared samples. Drawn lines represent a linear fit through the data points with 95% confidence levels indicated by dashed lines. Note that for the percentage detachment, data for the EPS and non-EPS producing strains have been included in one fit.
a b
c d
e f
g h
54
(Figure 5i-5p). Instead, whereas before transmission staphylococci adhered on the tip of the pillars, after transmission they appeared pressed in between pillars (especially when the pillar-to-pillar distance was 800 nm, see Figure 5p) and at the contact points between the bacterial cell surfaces and nanopillars, the cell wall appeared to be slightly deformed (compare Figure 5i-l and 5m-p). Interestingly, about two-fold more EPS producing bacteria were killed during transmission on a nanopillared Si surface than on a smooth surface, but this effect was only minor for the non-EPS producing strain (Table 2).
D I S C U S S I O N
In this work, we relate staphylococcal adhesion to and detachment from smooth and highly-ordered nanostructured Si surfaces as well as the transmission from a smooth donor surface to a nanostructured receiver surface and vice versa with staphylococcal adhesion forces to the different surfaces. This study was conducted on lithographically-prepared, highly-ordered, nanopillared Si surfaces with different center-to-center distances between pillars. This is a unique approach compared to many other studies17,23,27,28 on bacterial adhesion to nanostructured surfaces that are often carried out on randomly structured surfaces, unavoidably possessing an associated surface heterogeneity.29 However, our nanopillared Si surfaces are fragile which makes them unsuitable for most practical applications, but highly suitable for a model study on the influence of nanostructuring on bacterial interactions with the surface.
In general, nanostructured surfaces show reduced bacterial adhesion due to the fact that nanostructuring decreases the contact area between a substratum surface and adhering bacteria.30 The contact area fraction for each of our nanopillared surfaces is 0.16, 0.02, and 0. 11 for 200, 400, and 800 nm pillar-to-pillar distances, respectively. Although the adhesion force on nanopillared surfaces is significantly lower than that on a smooth surface, we did not see significant difference in the adhesion force between nanopillared surfaces (compare Table 1), despite their significant difference in the contact area fractions. This suggests that the spherical morphology of the bacteria yields limited, pointed contacts with reduced bacterial adhesion forces for two strains of staphylococci on nanopillared Si surfaces compared to a smooth surface. However, contrary to the literature,27 this does not result in reduced adhesion. This contradiction is a result of the use of a parallel plate flow chamber with in situ observation implying that adhering bacteria are enumerated without ever having been “slightly rinsed or dipped to remove loosely adhering bacteria”. Slight rinsing or dipping to remove loosely adhering bacteria is one of the most awful procedures in bacterial adhesion research, especially since the detachment force accompanying slight rinsing or dipping is never specified. However, it is easily envisaged that the number of remaining bacteria on a substratum surface will critically depend on the detachment forces accompanying slight rinsing or dipping.31,32 Therefore, we have always advocated that bacterial adhesion should be enumerated under the shear conditions during which adhesion took place.31,33,34 Considering this, we conclude that under convective-diffusional fluid flow even minor adhesion forces
55
Figure 4. Percentage bacteria left on a donor surface for EPS producing S. aureus ATCC12600 and
non-EPS producing S. epidermidis ATCC12228 after transmission between nanopillared Si surfaces as a function of the fractional adhesion force exerted by the nanopillared Si surface obtained under different loading forces in AFM. Fractional adhesion forces were calculated with respect to the force measured on the smooth surface. (a) schematics of transmission from a nanopillared donor to a smooth receiver. (b) Bacteria on the smooth donor and nanopillared receiver after transmission and separation of both surfaces (see also panel a). (c) Percentage staphylococcal transmission from smooth donor to nanopillared receiver surfaces as a function of the fractional adhesion force (AFM-loading force equals 2 nN) on the nanopillared Si receiver surface, d) percentage staphylococcal transmission from nanopillared donor to smooth receiver surfaces as a function of the fractional adhesion force (AFM-loading force equals 2 nN) on the nanopillared Si donor surface. (e, f) same as in panels c and d but for 5 nN AFM-loading force. (g, h) Same as in panels c and d but for 10 nN AFM-loading force. Error bars denote the standard deviations over triplicate experiments, carried out with different bacterial cultures and separately prepared samples. Drawn lines represent a linear fit through the data points with 95% confidence levels indicated by dashed lines. a b c d e f g h
3
56
Figure 5. Morphology of EPS producing S. aureus ATCC12600 and non-EPS producing S. epidermidis
ATCC12228 before and after transmission. (a – h) SEM micrographs of EPS producing S. aureus ATCC12600 on smooth and nanopillared Si donor surfaces before (a – d) and after (e – h) transmission to a smooth Si surface. Center-to-center distances between pillars increase from smooth (left) to 200, 400, and 800 nm on the right. Arrows indicate pressure-induced EPS production. (i - p) Same as panels a – h but for non-EPS producing S. epidermidis ATCC12228. Note absence of pressure-induced EPS production. Arrows indicate cell wall deformation at the contact points. Scale bars equal 500 nm.
arising from nanopillared Si surfaces (due to their limited contact area with bacteria flowing in suspension), are adequate to cause bacteria to adhere to a surface. This coincides with the observation that bacteria also adhere to polymer brush coated surfaces that exert very weak adhesion forces under mild convective-fluid flow, provided observed without slight rinsing or dipping.30,35,36 This conclusion at the same time, implies that most literature studies concluding that bacterial adhesion to nanostructured surfaces is reduced after slight rinsing or dipping,27,37–40 actually have demonstrated that bacterial detachment under the influence of
b c d
EPS producing S. aureus ATCC12600 before (a-d) after (e-h) transmission
p o n e a m
Non-EPS producing S. epidermidis ATCC12228 before (i-l) after (m-p) transmission
f g l k j i h
57 an externally applied detachment force increases upon nanostructuring of a surface. Thus, instead of demonstrating reduced bacterial adhesion on nanostructured surfaces, they demonstrate higher detachment. This now, fully coincides with the results of the present study demonstrating that adhering staphylococci are more easily detached from nanopillared Si surfaces with reduced adhesion forces than from smooth surfaces (Figure 3d, 3f and 3h).
Table 2. Percentages of dead bacteria on smooth donor surfaces and smooth and nanopillared receiver
surfaces after transmission for the EPS and non-EPS producing staphylococcal strains involved in this study.
Since transmission involves detachment from a donor surface followed by adhesion to a receiver surface, it follows logically from our observations, that bacterial transmission will decrease with increasing adhesion forces exerted by a donor surface relative to those exerted by the receiver surface (Figure 4). SEM micrographs however, show a new and unexpected phenomenon that occurs under influence of the contact pressure during transmission and that interferes with the influence of adhesion forces as presented in Figure 5. When the non-EPS producing strain was pressed against (200 nm pillar-to-pillar distance) or between pillars (800 nm distance) during transmission under pressure, this resulted in highly local, minor cell wall deformations (see Figure 5n-5p), caused by a limited number of pointed contacts, as speculated upon above. However, when the EPS producing strain was under pressure during transmission, high local stresses exerted by the pillars on the cell wall caused the organism to produce excessive amounts of EPS during transmission that are clearly visible in SEM micrographs (see Figure 5f-5h). EPS production by adhering staphylococci is known to depend on the adhesion forces by which they adhere to a substratum surface,41 causing cell wall deformation42 as a stimulus for EPS production. EPS production represents a major challenge for bacteria, because it must be assembled and exported in a process spanning the envelope, without compromising the essential barrier properties of the cell wall and are known to do so through the opening of outer membrane efflux channels43 that strains not possessing the ability to produce EPS may lack. Speculatively, EPS producing staphylococci excessively open their outer membrane efflux channels to excrete EPS to relieve the pressure brought about
EPS producing S. aureus ATCC12600
smooth to smooth (%) smooth to 200 (%) smooth to 400 (%) smooth to 800 (%) dead on donor 15 30 25 15 dead on receiver 25 60 65 40
Non-EPS producing S. epidermidis ATCC12228
smooth to smooth (%) smooth to 200 (%) smooth to 400 (%) smooth to 800 (%) dead on donor 10 30 40 30 dead on receiver 20 40 50 30
3
58
during transmission but do so in an uncontrolled way compromising the membrane barrier function therewith causing cell death. Clearly, non-EPS producing strains may not have the ability to open membrane efflux channels and consequently stay alive to a greater extent during transmission involving pillared surfaces than EPS producing strains (Table 2).
Pressure-induced EPS production and associated cell death is a new phenomenon that has never been reported upon before and should be accounted for in future research on the influence of nanostructured surfaces on bacterial adhesion. Very often, it is not of major importance as to how many bacteria adhere to a surface since once adhering bacteria start growing with generation times of 20-30 min. Statistically significant differences by factors less than ten then rapidly disappear and only log-scale differences matter. More importantly in many practical situations where control of bacterial adhesion relies on the use of antimicrobials, it is important whether or not adhesion stimulates the adhering bacteria to produce their protective EPS matrix. Here we demonstrate that the likelihood that adhering bacteria after transmission from a donor surfaces produce their EPS matrix is enhanced when nanostructured surfaces are involved. On the other hand, however, bacteria can be killed by the high local pressures exerted by nanostructures to which they adhere.
C O N C L U S I O N S
Staphylococci experience weaker adhesion forces from nanopillared Si surfaces than from smooth Si surfaces. This, however, does not result in reduced numbers of staphylococci adhering to either surface under convective-diffusional fluid flow in a parallel plate flow chamber. Alternatively, detachment was much more stimulated by increasing the fluid flow when bacteria adhered to nanopillared Si surfaces than to a smooth Si surface. Transmission
between surfaces involves detachment from a donor surface followed by adhesion to a receiver surface. Hence transmission involving nanostructured surfaces was impacted by the
presence of nanopillars, the direction of impact depending on whether the donor was smooth or nanostructured. High local contact pressures during transmission exerted by nanostructures on the bacterial cell wall can induce EPS production through the opening of membrane efflux pumps therewith compromising the membrane barrier function and causing cell death. These events need to be accounted for in practical applications of nanostructured surfaces as the protective EPS matrix can strongly reduce the efficacy of antimicrobial treatments of biofilms. However, on the other hand, bacteria may be killed by high local pressures as well, arising from adhesion to nanostructures.
59
Supporting Information
Figure S1. Fabrication of nanopillared Si surfaces. (a) Spin coating of photoresist (PR) on to Si substrate.
(b) Laser interference lithography to define a nano-patterned PR layer. (c) Forming pore pattern via developing PR. (d) Physical vapor deposition of Cr through PR pores. (e) Removal of photoresist layer in piranha solution. (f) Deep reactive ion etching of Si with Cr dots serving as mask. (g) Removal of Cr dots in chrome etchant.
60
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