Article pubs.acs.org/Langmuir
Hot Embossed Microtopographic Gradients Reveal Morphological Cues That Guide the Settlement of Zoospores Linlin Xiao,*,†,∥ Stephanie E. M. Thompson,‡ Michael Röhrig,§,# Maureen E. Callow,‡ James A. Callow,‡ Michael Grunze,†,∥ and Axel Rosenhahn†,∥,⊥ †
Applied Physical Chemistry, University of Heidelberg, 69120 Heidelberg, Germany School of Biosciences, University of Birmingham, B15 2TT Birmingham, United Kingdom § Institute of Microstructure Technology, Karlsruhe Institute of Technology, 76021 Karlsruhe, Germany ∥ Institute of Functional Interfaces, Karlsruhe Institute of Technology, 76021 Karlsruhe, Germany ⊥ Analytical Chemistry - Biointerfaces, Ruhr-University Bochum, 44801 Bochum, Germany # Karlsruhe Nano Micro Facility, Helmholtz Research Infrastructure at Karlsruhe Institute of Technology, 76021 Karlsruhe, Germany ‡
ABSTRACT: Among different surface cues, the settlement of cells and larvae of marine macrofouling organisms has been found to be strongly influenced by surface microtopographies. In this article, the settlement of zoospores of the green alga Ulva linza on a surface topographic gradient has been investigated. “Honeycomb” gradient structures with feature sizes ranging from 1 to 10 μm were prepared by hot embossing, and the effect on the density of spores that attached in settlement assays was quantified. The highest density of spores was found when the size of the microstructures was similar to or larger than the size of the spores. With decreasing size of the structures, spore settlement density decreased. Interestingly, spore settlement density correlated with the Wenzel roughness of the surfaces. “Kink sites” on the surface played an important role and resembled preferred attachment positions. Furthermore, the gradients allowed the minimum pit size that the spores were able to squeeze into to be determined.
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INTRODUCTION
Ulva spp. are some of the most frequently observed softfouling organisms, being especially common around the water line. The life cycle of this macroalgal genus is characterized by the colonization of various substrates by zoospores, involving a temporary exploration phase followed by permanent settlement (adhesion).16,17 The quadriflagellate motile spores are pyriform “naked” cells; thus, without the support of cell walls they are easily deformed. When the spores commit to permanent settlement, they “round up” as they secrete adhesive, flatten the anterior region against the substratum, and retract the flagellar axonemes inside the cell.16,18 The settlement of the motile spores is modulated by different surfaces cues, including chemistry,17,19 morphology,20,21 surface nanoforce,22 and surface charge.23 Among the different surface properties, topography has been proven to have both deterrent and attractive effects on spore settlement. Settlement of spores of Ulva linza is highly dependent on topographic features including size, spacing, aspect ratio, and roughness.21,24,25 Generally, microtopographies slightly larger than the cell/larva of the fouling organisms facilitate settlement, in which case the surface
It is known that surface topographical cues are among the three key physiochemical properties, which influence and modulate adhesion on surfaces.1 Studies related to protein adsorption, cell differentiation, and microbial response on different surface microtopographies have been resported.2−4 For example, the microtopography of honeycomb-patterned porous films has been found to strongly affect the morphology and adhesion of cells.5 Besides the studies in the biomedical field, the role of surface micro- and nanotopography is intensively investigated and discussed in the area of marine antifouling. Marine biofouling, associated with the undesired accumulation of micro- and macrocolonizers on natural or manufactured submerged surfaces,6,7 has been a notable long-standing problem for marine-related industries (e.g., shipping, ocean energy, and aquaculture).8−10 The consequences of fouling raise significant problems in both economic and ecological aspects.11 Biocidal antifouling (AF) coatings have been the main technologies to control fouling organisms for the past decades.12,13 Increasingly restricted regulations on biocides have motivated and brought scientists from different disciplines together in order to research and develop novel nontoxic solutions to control biofouling.14,15 © 2012 American Chemical Society
Received: September 24, 2012 Revised: December 25, 2012 Published: December 28, 2012 1093
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Figure 1. (a) Diagram of “honeycomb” gradient microstructures. The gradient is based on the honeycomb motif and consists of hexagonal pits. To the left pits are getting smaller until they reach a minimal size of 1 μm. As the vertical rims have a smaller width, the structure converts into a zigzag structure when the size of the pits exceeds a critical diameter (right-hand side). The small circles on the left side of the structure resemble the symmetry of the centers of the hexagonal pits and their corresponding characteristic distances. (b) Diagram of hot-embossed PMMA “honeycomb” gradient replicas. (The gray structured areas are 1.512 cm × 1 cm, and the full dimensions of the samples are 4.5 cm × 4.5 cm.)
provides sufficient attachment sites to protect adherent biofoulers against hydrodynamic forces,20,26−29 while substantially reduced settlement and attachment strength of the settling propagules could be found on natural or man-made surfaces with features smaller than their body sizes.24,29 During settlement, spores also show their preference for certain attachment positions. Normally areas with higher dissimilarity are preferentially chosen by spores (i.e., recessed areas).22,30,31 For the correlation between surface topographic features and spore behaviors, different indices and models have been set up to help understand the mechanism and predict surface performance, which could finally lead to an optimal antifouling design. The Engineered Roughness Index (ERI), which considers the combined effect of surface roughness, freedom of movement, depressed area fraction, and amount of distinct features, has been defined24 and refined32,33 in order to interpret and predict zoospore settlement in relation to surface topography. The predictive regression model developed by Long et al.32 exhibits high correlation between several independent data sets for zoospore attachment. To identify critical surface features or favorable surface properties relevant for antifouling applications, gradients are highly useful.34,35 For example, with position-bound and gradually changing properties on the same surface, highthroughput and cost-effective analysis of microorganism behaviors can be quantified in a single experiment, which can further facilitate new surface designs. Since the first gradient surface was described in 1960s,36 many studies have focused on this field in order to produce gradients of surface wettability, chemistry, and topography. 37−39 However, besides the settlement of spores of U. linza on wettability gradients40 and hierarchically wrinkled coatings,41 there have been few applications in relation to marine fouling. In the present study, we aimed to develop an understanding about the response of spores of U. linza to a range of continuously changing microtopographic properties using a morphological gradient. In the work presented here, “honeycomb” microtopographic gradient samples were fabricated by hot embossing, and spore settlement on the topographic gradient was quantified in laboratory assays. The questions addressed were how spore settlement density changed with changing feature size and spacing and what was the critical pit size that spores would “choose” and be able to fit into. Furthermore, the
importance of local binding geometry (i.e., the attachment area or location) which spores “preferred” was analyzed.
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EXPERIMENTAL SECTION
Surface Preparation. Layout Design. Figure 1 shows the “honeycomb” gradient layout. The hexagonal microstructures appeared as pits on the poly(methyl methacrylate) (PMMA, PLEXIGLAS Folie Farblos 99530 from Röhm GmbH Chemische Fabrik, Darmstadt, Germany) replicas (Figure 1a). The pit size (the width of the hexagon in x direction in Figure 1a) changed continuously in micrometer step from 1 to 10 μm along the gradient, while the center distance between adjacent hexagons remained the same (10 μm in x direction in Figure 1a). On each PMMA sample, there were six structured areas (Figure 1b). Between the structured areas, there was smooth PMMA, which was used as the control for the bioassay. Hot Embossing. “Honeycomb” gradient structures were fabricated by micro hot embossing (Figure 2), a plastic moulding process for
Figure 2. Schematic view of the hot embossing process (adapted from ref 42). producing large-area surfaces with micro- and/or nanostructures.43 In order to avoid incomplete filling of the microstructures, the embossing parameters were optimized before producing the large number of replicas required for the biological tests. The embossing machine consists of the moulding platform and the demoulding plate. The structured nickel mould insert was fixed on the moulding platform and an unstructured PMMA sheet was positioned between the two-part moulding tool. After closing the embossing tool, the PMMA sheet was heated above the glass transition temperature (95 °C). When the moulding temperature (155 °C) was reached, the achieved embossing force (40 kN) pressed the softened PMMA into the cavities of the mould insert. While keeping the applied force constant, the polymer was cooled below the glass transition temperature. Finally, the 1094
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Figure 3. Top-down phase contrast microscopic images of the “honeycomb” gradient topography fabricated on the surface of PMMA. microstructured part was demoulded by opening the moulding tool, and the sample was peeled off the demoulding plate. Surface Characterization. Microscopy. The morphology of the “honeycomb” gradient was checked under an automated microscope (Nikon Eclipse 90i) in phase contrast mode with the objective magnification ×100. Images at different sites across the gradient were taken to show the change in surface microstructures. Contact Angle. Contact angle measurement was applied to determine the change in surface wettability across the gradient. With a self-build contact angle goniometer, the sessile water contact angles (CAs, droplet of Milli-Q water around 2 μL) were measured under ambient conditions with the tip withdrawn from the droplet. The presented values were the average of five measurements. Atomic Force Microscopy (AFM). The depth of the hexagonal pits was measured by a MFP-3D BioAFM (Asylum, Mannheim) in AC mode using a commercial Si3N4 cantilever with a spring constant of 7.5 N/m (μMasch) in air. The AFM was used in a closed loop operation mode on all three axes with the tip scanning back and forth at 0° along the horizontal line in the range of 80 μm. Obtained topographic images were evaluated with the corresponding software IGOR. Scanning Electron Microscopy (SEM). For classifying spores at different positions and measuring the displacement of spores from the centers of pits, samples were examined by SEM. Surfaces with settled spores were prepared as described. Before imaging, the samples were coated with a very thin layer of gold and graphite mixture to make the surface electrically conductive. Images were recorded in a LEO1530 Gemini electron microscope operating under high vacuum and with the EHT of 1 kV. Zoospore Settlement Assay. Plants of Ulva linza were collected from Llantwit Major, Glamorgan, Wales (52° 23′ N; 3° 30′ W) 2−5 days before the spring tide. Zoospores were released from the reproductive tips into filtered (0.22 μm) artificial seawater (ASW, Tropic Marin) and prepared for the assay as described by Cooper et al.31 In brief, after filtering through three layers of nylon mesh (100, 50, and 20 μm) to remove debris, the beaker containing the spore suspension was plunged into ice; spores swam rapidly toward the bottom. The concentrated spore suspension was removed with a pipet and refiltered through two layers of 20 μm nylon mesh. The spore suspension was diluted with filtered ASW to an absorbance of 0.15 at 660 nm, corresponding to a concentration of 1 × 106 spores mL−1. The spore suspension was kept on a magnetic stirrer and used in assays within 30 min of release.
Gradient samples were preincubated in Petri dishes containing filtered ASW on a vibrating platform for 1 h prior to the assay to ensure the surfaces were fully wetted and then rapidly transferred to the assay dishes to minimize the possibility of dewetting. 30 mL of spore suspension was added to 9 cm diameter Petri dishes, each containing a PMMA wafer. The dishes were incubated in the dark at room temperature (ca. 20−22 °C) for 45 min. The samples were then washed in filtered ASW to remove motile spores (i.e., spores that had not settled and undergone permanent attachment), by passing the samples 10 times through a beaker of ASW. The samples were then fixed in 2.5% glutaraldehyde in ASW for 20 min, followed by washing sequentially in ASW, 50% ASW, and deionized water and subsequently air-dried. Three separate experiments using different batches of spores were performed to check reproducibility of the data. It was not possible to count settled spores using automated image analysis as the “honeycomb” pattern was autofluorescent, leading to exaggeration of cell number. Therefore, spores were viewed in transmission light microscopy and counted manually. Three transects, corresponding to 31 fields of view, along each gradient were counted. Counts were recorded in an area of 0.038 mm2 at 0.5 mm intervals along the long axis of the gradient. The three cell counts taken at each of the 31 points along the gradient (top, middle, and bottom) were averaged to give a mean value (n = 3) for each patch. For the background (smooth control), 31 counts were made along transects of 15 mm length in three different areas of the wafer. The result is represented as cells mm−2 with respect to the size of microhexagonal pits.
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RESULTS AND DISCUSSION For quality control of the hot embossing samples, phase contrast microscopy was used (Figure 3). Across the gradient, the size of the microstructures increased. As the rims in the ydirection (Figure 1a) were slightly thinner compared to the other two directions (y′ and y″), they became smaller and finally disappeared with increasing diameter of the hexagons, and the surface pattern transformed into zigzag structures. The contact angle for the smooth area between the microstructures was ≈77°. Across the gradient the contact angle changed from 77° to 109° with increasing size of the hexagons. This increase was expected, as larger structures normally have a higher roughness which causes larger CAs. The depth of the hexagonal 1095
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pits measured by AFM was 1.57 ± 0.08 μm across the whole gradient. To correlate surface topography with spore settlement, the Wenzel roughness along the PMMA “honeycomb” gradients was calculated using the variables introduced in Figure 4, where
Figure 4. (a) Schematic representation of the repeating unit of the hexagonal structures. The distances between the centers of the hexagonal pits (x/3 and y, respectively) are fixed across the gradient with x/3 = 10 μm and y = 11.58 μm, while m and a change according to the size of the features and are connected by (a = (√3/3)m). (b) The roughness factor increases with the height of the structures h due to the available area of the side walls which are formed during the embossing process.
Figure 5. Correlation of Wenzel roughness and settlement density of zoospores of U. linza. Mean cell counts for five individual patches on one wafer; each point is the mean of three transects; counts were taken at 0.5 mm intervals along each gradient. The background value is the mean of 93 counts.
x and y represent respectively the length and width of the extracted unit and m describes the width of the hexagonal pit, while a is the side length of the hexagon. In Figure 4b, h stands for the height of the microstructures. The Wenzel roughness is defined as the ratio between the actual surface area and the geometric surface area.44 The actual surface area is the sum of projected surface area and area of the side walls. When the microstructures changed from hexagons to zigzag walls, the number of walls belonging to each hexagonal unit decreased from six to four. So for these two different parts of the gradient (i.e., the hexagonal part and the zigzag part), two sets of formulas were used to calculate the Wenzel roughness factor. For hexagonal structures (images a−e in Figure 3) R Wenzel = =1+
Sactual Sgeometric
=
Sgeometric + Swall Sgeometric
=
structures rather than in the middle. The gray area in Figure 5 represents the change in Wenzel roughness factor across the gradient, which showed a close correlation with spore settlement up to the point where hexagons became zigzag structures. Since the Wenzel roughness factor was introduced,44 it has been applied to a range of different studies. Developed by Brennan’s research group,24,32,33 the dimensionless Engineered Roughness Index (ERI) was used to correlate surface topographic properties and biofouling. For the fixed microstructure dimensions (feature spacing = 2 μm and feature depth = 3 μm), the spore settlement is inversely linear to the ERII. In both ERI equations (ERII and ERIII), the Wenzel roughness ratio (r) is proportional to ERI. Both ERII and ERIII also take into account the depressed surface area fraction (1 − φs). The degree of freedom (df) to move across the surface is considered in ERII, while for ERIII df is replaced by the number of distinct features (n).
xy + 18ah xy
10 3 mh 579
(1)
while for zigzag structures (image f in Figure 3) R Wenzel =
Sactual Sgeometric
20 3 =1+ mh 1737
=
Sgeometric + Swall Sgeometric
=
xy + 12ah xy (2)
The data in Figure 5 show the number of spores on different sections of the morphological gradient. The five curves were measured on five individual replicates. Reproducibility was verified for two additional batches of spores which yielded agreeing trends. As shown in Figure 5, spores attached at a higher density on the structured area compared to the smooth background. The number of settled spores was found to be gradually reduced with decreasing size of the pits. The reduction started to become apparent when the microstructures were smaller than the size of motile spores (pyriform spores have a diameter of approximately 4−5 μm at their widest part). As the size of the pits exceeded a diameter of 8.5 μm, the structures changed from hexagons to zigzag walls. Along with the structural change, a dramatic decrease in spore settlement was found. A closer inspection of the microscopic images revealed that the spores preferred to settle against edges in the
ERII =
r df 1 − φs
(3)
ERIII =
rn 1 − φs
(4)
Interestingly, the present results on the hexagonally structured morphology gradients reveal a trend of increasing settlement with increased Wenzel roughness (Figure 5). This is in contrast to the general ERI model which predicts lower settlement for increased Wenzel roughness, although only for topographies based upon feature size/spacing of 2 μm.24 In the present study even topographies based upon 2 μm stimulated settlement. On surfaces with a size gradient it is difficult to determine the number of “unique features” at each point. Moreover, the Wenzel roughness factor and depressed surface area fraction also change simultaneously across the gradient. Thus, it is not possible to describe the surface topographic features of the present study by the ERI model and the mechanistic basis of the difference in the response of spores to topographies in these different studies is not known. It is 1096
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Figure 6. (a) SEM image showing zoospores settled against a “kink site” and a “non-kink site”. (b) Sketch of local binding geometry of settled zoospore at a “kink site” and a “non-kink site”. (c) Mean cell counts for spores at different binding geometries during the transition of the gradient from hexagon to zigzag structures; each point is the mean of three counts taken at 0.5 mm intervals along each gradient with the error bar showing the standard error.
discharge of membrane-bound adhesive vesicles with the plasma membrane of the spore to contact the surface.45 The area available to be spread by the adhesive determines the effective contact area, which obviously is maximized at a kink position. This situation is qualitatively comparable with the “half-crystal position” or “terrase ledge kink” site in crystal growth on solid interfaces.46 By analyzing the SEM images in greater detail, the smallest pit size that spores could enter and settle in can be determined. Visual analysis of the images showed that if spores did not fit in the pits, they tended to settle asymmetrically and slightly displaced from the center of the pits. The same was observed if the pits were too large as spores tended to populate “kink sites” in the pits. This observation was quantified by SEM images (Figure 7) by measuring the distance between the center of the settled spore and the center of the pit. A gradual decrease in displacement was found when the size of the pits increased up to ∼2.0 μm. A steep drop in displacement to nearly zero was
possible that the presence of the gradient modifies responses of spores since the previous studies on gradients (albeit gradients of wettability rather than topography) generated some unexpected results.40 The settlement of spores of Ulva on surfaces with homogeneous wettability was substantially different to settlement on equivalent wettabilities when presented as part of a gradient. A steep drop in spore settlement density was found when the structure changed from hexagon to zigzag (size of hexagonal pits ≈8.5 μm). As pointed out above, scanning electron microscopy revealed that spores preferentially settled in the “kink sites”. The number of “kink sites” in the hexagonal area was 3 times larger compared with the zigzag area. Interestingly, the spore settlement density was also reduced to approximately one-third in the zigzag area. As shown in Figure 5, the change in spore settlement density was larger than that expected from the change in Wenzel roughness. Thus, a more detailed SEM investigation was carried out in which the spores were classified according to settlement positions. Figure 6c shows the change in the number of all attached spores, spores at “kink sites”, and “non-kink sites”. The difference between a “kink site” and a “non-kink site” or steep edge position is that three instead of two side walls are available for binding (Figure 6a,b). Figure 6c shows that during the transition from the hexagonal structure to the zigzag structure the decrease in the number of spores settled was mainly caused by the reduction of spores at “kink sites”, while the number of spores at “non-kink sites” remained almost the same. Thus, SEM provided substantial proof of the “kink site” effect. This gave rise to the interpretation that spores were able to detect sheltered pits and kinks on the surface and these sites facilitated settlement. A similar preference has also been observed for barnacle larvae, which showed strongly enhanced settlement on microstructures larger than the size of the cyprids providing maximum anchoring sites.28 The transition to the zigzag pattern changed the available number of “kink positions” which was reduced to one-third. Spores of U. linza are pear-shaped when swimming and then assume a spherical shape after they commit to settlement. “Kink sites” provide more attachment points, which facilitate the firm adhesion of spores against hydrodynamic forces (Figure 6b). This adhesion is facilitated by the endocytotic fusion and
Figure 7. Distance between the center of the pits and the center of the settled spores at different positions along the gradient. 1097
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found at a pit size of 2.6 μm, where spores were able to squeeze themselves into the pits. Spores remained in the center of the pits until the size of the pits increased to around 5 μm. Spores could be found sitting snugly when the size of the pit became closer to the dimension of the middle region of the swimming spores. When the size of the hexagons increased, spores tended to attach to the corner of the pits rather than sit in the middle and the displacement again increased. Thus, the minimum size of the pits across the gradient in which spores could fit was about 2.6 μm. The value of 2.6 μm obtained from SEM analysis was slightly smaller than the mean diameter of swimming spores (ca. 4−5 μm). Spores could fit into the 2.6 μm diameter pit because they were pear-shaped at the time they committed to settlement. After attaching, the spores became round and more compact in the pits.
(dimethylsiloxane) on protein adsorption, platelet and cell adhesion. Colloids Surf., B 2009, 71 (2), 275−281. (3) Hayes, J. S.; Khan, I. M.; Archer, C. W.; Richards, R. G. The role of surface microtopography in the modulation of osteoblast differentiation. Eur. Cells Mater. 2010, 20, 98−108. (4) Edwards, K. J.; Rutenberg, A. D. Microbial response to surface microtopography: the role of metabolism in localized mineral dissolution. Chem. Geol. 2001, 180 (1−4), 19−32. (5) Tanaka, M.; Takayama, A.; Ito, E.; Sunami, H.; Yamamoto, S.; Shimomura, M. Effect of pore size of self-organized honeycombpatterned polymer films on spreading, focal adhesion, proliferation, and function of endothelial cells. J. Nanosci. Nanotechnol. 2007, 7 (3), 763−772. (6) Wahl, M. Marine epibiosis. I. Fouling and antifouling: some basic aspects. Mar. Ecol.: Prog. Ser. 1989, 58 (1−2), 175−189. (7) Callow, M. E.; Callow, J. E. Marine biofouling: a sticky problem. Biologist 2002, 49 (1), 10−14. (8) Edyvean, R. Consequences of fouling on shipping. Biofouling 2010, DOI: 10.1002/9781444315462.ch15. (9) Zvyagintsev, A. Y.; Ivin, V. V. Study of biofouling of the submerged structural surfaces of offshore oil and gas-production platforms. Mar. Technol. Soc. J. 1995, 29 (2), 59−62. (10) Adams, C. M.; Shumway, S. E.; Whitlatch, R. B.; Getchis, T. Biofouling in marine molluscan shellfish aquaculture: a survey assessing the business and economic implications of mitigation. J. World Aquacult. Soc. 2011, 42 (2), 242−252. (11) Schultz, M. P.; Bendick, J. A.; Holm, E. R.; Hertel, W. M. Economic impact of biofouling on a naval surface ship. Biofouling 2011, 27 (1), 87−98. (12) Finnie, A. A.; Williams, D. N. Paint and coatings technology for the control of marine fouling. Biofouling 2010, DOI: 10.1002/ 9781444315462.ch13. (13) Thomas, K. V.; Brooks, S. The environmental fate and effects of antifouling paint biocides. Biofouling 2010, 26 (1), 73−88. (14) Dafforn, K. A.; Lewis, J. A.; Johnston, E. L. Antifouling strategies: history and regulation, ecological impacts and mitigation. Mar. Pollut. Bull. 2011, 62 (3), 453−465. (15) Callow, J. A.; Callow, M. E. Trends in the development of environmentally friendly fouling-resistant marine coatings. Nat. Commun. 2011, 2, 244. (16) Callow, M. E.; Callow, J. A.; Pickett-Heaps, J. D.; Wetherbee, R. Primary adhesion of Enteromorpha (Chlorophyta, Ulvales) propagules: quantitative settlement studies and video microscopy. J. Phycol. 1997, 33 (6), 938−947. (17) Heydt, M.; Pettitt, M. E.; Cao, X.; Callow, M. E.; Callow, J. A.; Grunze, M.; Rosenhahn, A. Settlement behavior of zoospores of Ulva linza during surface selection studied by digital holographic microscopy. Biointerphases 2012, 7 (1−4), 33. (18) Callow, M. E.; Callow, J. A. The Ulva spore adhesive system. Biol. Adhes. 2006, DOI: 10.1007/978-3-540-31049-5-4. (19) Schilp, S.; Rosenhahn, A.; Pettitt, M. E.; Bowen, J.; Callow, M. E.; Callow, J. A.; Grunze, M. Physicochemical properties of (ethylene glycol)-containing self-assembled monolayers relevant for protein and algal cell resistance. Langmuir 2009, 25 (17), 10077−10082. (20) Callow, M. E.; Jennings, A. R.; Brennan, A. B.; Seegert, C. E.; Gibson, A.; Wilson, L.; Feinberg, A.; Baney, R.; Callow, J. A. Microtopographic cues for settlement of zoospores of the green fouling alga Enteromorpha. Biofouling 2002, 18 (3), 237−245. (21) Cao, X.; Pettitt, M. E.; Wode, F.; Sancet, M. P. A.; Fu, J.; Ji, J.; Callow, M. E.; Callow, J. A.; Rosenhahn, A.; Grunze, M. Interaction of zoospores of the green alga Ulva with bioinspired micro- and nanostructured surfaces prepared by polyelectrolyte layer-by-layer selfassembly. Adv. Funct. Mater. 2010, 20 (12), 1984−1993. (22) Schumacher, J. F.; Long, C. J.; Callow, M. E.; Finlay, J. A.; Callow, J. A.; Brennan, A. B. Engineered nanoforce gradients for inhibition of settlement (attachment) of swimming algal spores. Langmuir 2008, 24 (9), 4931−4937. (23) Rosenhahn, A.; Finlay, J. A.; Pettit, M. E.; Ward, A.; Wirges, W.; Gerhard, R.; Callow, M. E.; Grunze, M.; Callow, J. A. Zeta potential of
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CONCLUSIONS Summarizing, morphological gradients allowed the influence of topographic cues on the settlement of spores to be studied systematically. We observed a clear correlation between spore settlement density and Wenzel roughness r. The specific role of the “kink site” revealed the importance of the local binding geometry as spores selectively populated these sites. And in agreement with previous notion, our studies also showed that spores preferred topographies of a size similar to or slightly bigger than their body size.20,47 Moreover, the smallest pit size occupied by spores was analyzed and determined to be 2.6 μm. This value was in line with previous studies showing that markedly reduced settlement was found on smaller microstructures (e.g., 2 μm).21
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AUTHOR INFORMATION
Corresponding Author
*E-mail
[email protected]. 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 The financial support from the European Community’s Seventh Framework Programme FP7/2007-2013 under Grant Agreement 237997 (SEACOAT) is thanked as well as the Biointerfaces programme, Helmholtz Society. We also acknowledge the kind help of Tatjana Ladnorg (IFG, KIT) with AFM measurements and Dr. Frank Friedrich (IFG, KIT) with SEM measurements. We thank Dr. Matthias Worgull, Dr. Hendrik Hölscher, and Prof. Volker Saile (IMT, KIT) for stimulating discussions as well as the Karlsruhe Nano Micro Facility (KNMF) for kind support. We acknowledge fabrication of the mold insert by A. Bacher, P.-J. Jakobs, and B. Matthis (IMT, KIT).
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REFERENCES
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dx.doi.org/10.1021/la303832u | Langmuir 2013, 29, 1093−1099
