ARTICLE pubs.acs.org/Langmuir
Charge Specific Protein Placement at Submicrometer and Nanometer Scale by Direct Modification of Surface Potential by Electron Beam Sylvain Robin,†,^ Abbasi A. Gandhi,‡,^ Maros Gregor,§ Fathima R. Laffir,‡ Tomas Plecenik,§ Andrej Plecenik,§ Tewfik Soulimane,† and Syed A. M. Tofail*,‡ †
Chemical and Environmental Science Department and Materials and Surface Science Institute, University of Limerick, Limerick, Ireland Materials & Surface Science Institute, University of Limerick, Limerick, Ireland § Department of Experimental Physics, Faculty of Mathematics, Physics and Informatics, Comenius University, 84248 Bratislava, Slovakia ‡
ABSTRACT: The understanding and the precise control of protein adsorption is extremely important for the development and optimization of biomaterials. The challenge resides in controlling the different surface properties, such as surface chemistry, roughness, wettability, or surface charge, independently, as modification of one property generally affects the other. We demonstrate the creation of electrically modified patterns on hydroxyapatite by using scanning electron beam to tailor the spatial regulation of protein adsorption via electrostatic interactions without affecting other surface properties of the material. We show that domains, presenting modulated surface potential, can be created to precisely promote or reduce protein adsorption.
’ INTRODUCTION Protein adsorption on biomaterials is of great importance as it controls many physiological responses and ultimately the biocompatibility of the material and the success of its implantation.1 The understanding and the precise control of protein deposition are therefore extremely important for the development and optimization of better biomaterials.2,3 Hydroxyapatite (HAp) is a bioceramic presenting an excellent biocompatibility, and for this reason the use of this material is extremely developed in the fields of bone or teeth replacement.4 Protein adsorption onto HAp plays an important role in this as it governs subsequent mechanisms such as cell attachment, proliferation, and migration that ensure the performance of this biomaterial. Protein adsorption is influenced by the physical and chemical properties of the surface such as roughness,5,6 chemical surface composition and structure, wettability,7 and surface charge.5,8 The challenge associated with a better understanding of protein adsorption mechanisms resides in controlling those different surface properties independently, as modification of one property generally affects the others.5,9 The surface charge of a material can lead to the attraction or repulsion of proteins depending on their isoelectric point and charge at a given pH.5 In the case of HAp, protein adsorption is mainly driven by the presence of P-sites and C-sites on a face (or b) and c face, respectively.10,11 The P-sites present an electronegative potential due to the exposition of phosphate groups, while C-sites are electropositive due to the presence of Ca2+ ions.11 It has been shown that positively charged proteins exhibits a higher adsorption on P-sites, while negatively charged proteins present a higher r 2011 American Chemical Society
affinity for C-sites.11 In the latter case, the electronegatively charged bovine serum albumin (BSA) adsorbed preferentially to C-sites through specific interactions with the bivalent calcium ions.12 While these studies have been conducted mainly on powder materials, the knowledge of such selective attachments of protein can be extended to solid continuous surfaces on a multiple length scales and is the focus of the present study. High resolution micro- and nanopatterning of proteins on solid continuum is of great practical interest for the development of many applications, from biosensors to templates for tissue engineering. The creation of these patterns by chemical selfassembly, lithographic, or morphological patterning have been demonstrated in the past. One approach consists in modifying the surface potential in order to tailor protein adsorption through electrostatic forces. To alter the surface potential in a spatially controlled manner, different strategies have been demonstrated. Proteins have been successfully patterned using chemical modification of the material surface.13 With such an approach Kumagai et al. achieved high-resolution single protein placement by patterning nanometric patches of positively charged 3-aminopropyltriethoxysilane (APTES) on negatively charged SiO2 substrate.14 Physical modifications can be used in order to affect only the surface potential without affecting other properties of a chosen material. Jacobs and coauthors have demonstrated the use of contact poling in order to create patterns with modified Received: September 12, 2011 Revised: October 11, 2011 Published: November 07, 2011 14968
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Langmuir surface potential.15 The drawback of the latter approach is that the technology is cumbersome. Recently, Aronov et al.16 have shown that the wettability of dense pellets of hydroxyapatite pellets can be easily modified to obtain selective protein adsorption on a macroscopic scale, but the extent of adsorption due to electrostatic attraction/repulsion is not known. On the other hand, our group has recently shown that a focused electron beam available as an EDX probe in a laboratory SEM can be used to create localized microdomains of electrostatic charge.17 In the present work, we use a rastered electron beam (e-beam) typically used in SEM imaging to create electrically modified patterns on hydroxyapatite that tailors protein adsorption without affecting the wettability, the chemistry or the roughness of the material. Lysozyme (LSZ), conjugated with fluorescein, was chosen as a model protein to probe the surface potential after irradiation. We show that the surface potential intensity can be modulated to such an extent that our approach offers more flexibility to control attraction or repulsion of biomolecules independently of chemistry, roughness, and wettability. While we demonstrate such patterning to the resolution limit of a laser scanning microscope (∼200 nm), we elaborate the methodology to obtain such selective placement of protein at and below the 150 nm scale.
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Scheme 1. Process Scheme for the Formation of Arrays of Electrically Modified Domains and the Specific Adsorption of FITCLysozyme via Electrostatic Interactionsa
(1) The grounded HAp film is irradiated using 30 keV e-beam through a nickel mask. (2) The electrically modified patterns are imaged by KPFM. (3) A solution containing FITClysozyme is deposited on the irradiated samples. The surface potential of the protein is represented: the blue, white, and red color indicates positive, neutral, and negative potential, respectively. (4) After washing and drying, the protein distribution is assessed by laser scanning confocal microscopy.
a
’ EXPERIMENTAL SECTION HAp Film Fabrication. HAp film on silicon substrate was deposited using the solgel method and spin-coating. A stoichiometric amount of calcium nitrate tetrahydrate (Ca(NO3)2 3 4H2O) and phosphorus pentoxide (P2O5) with absolute ethanol was mixed separately and filtered using a 45 μm filter. They are mixed then together for 10 min at room temperature to form an initial solgel precursor solution and for 2 h at 60 C in a closed reflux system. Once the solution is gelated, it is added dropwise onto the silicon wafer. This is then spin-coated at a 4000 speed with 500 rpm/s speed rate for 50 s. Once the sample has been coated, it is immediately dried on a hot plate at 150 C, and the sample is then annealed in the preheated oven at a 700 C for 1 h. LSZ Labeling and Protein Adsorption. Lysozyme (LSZ) (from hen egg, 95% purity) was purchased from Sigma-Aldrich (St. Louis, MO) and used as received. To allow detection using confocal microscopy, the LSZ was labeled using fluorescein isothiocyanate (FITC) in carbonate buffer (pH 9.0) for 45 min at 20 C. The reaction was stopped by adding a large excess of Tris-HCl buffer (pH 8.0). The unbound dye was removed by gel filtration followed by extensive dialysis against 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer (pH 7.3). The ratio fluorescein/protein was confirmed to be e1. For the protein adsorption on surfaces, the concentration of 0.5 g/mL FITCLSZ was used. The labeled protein was prepared in 1 mM HEPES buffer (pH 7.3). The protein solution was deposited for 1 min on the freshly e-beam irradiated HAp film room temperature. The HAp film was then rinsed with the same buffer, water, and air-dried. The protein distribution was determined by using a laser scanning confocal microscope (CarlZeiss Meta710). X-ray Photoelectron Spectroscopy (XPS). XPS was performed in a Kratos AXIS 165 spectrometer using monochromatic Al Kα radiation of energy 1486.6 eV. High-resolution spectra were taken at fixed pass energy of 20 eV, 0.05 eV step size, and 100 ms dwell time per step, resulting in an average instrumental resolution of 0.7 eV (Au 4f7/2). Surface charge was efficiently neutralized by flooding the sample surface with low-energy electrons. Core level binding energies were corrected using C 1s at 284.8 eV as charge reference. For construction and fitting of synthetic peaks of high-resolution spectra, a mixed GaussianLorenzian
function with a Shirley type background subtraction were used. Data were collected at a takeoff angle of 90 from the surface and analyzed using the CASA XPS Analysis program (Neil Farley, Casa XPS). SEM. HAp films on silicon substrates were masked with a patterned transmission electron microscopy (TEM) metal mask (Agar Scientific, Nickel 3.05 mm thick and 600 mesh) to create approximately 65 65 μm electrically modified areas. The electron irradiation was performed in a vacuum at room temperature, using SEM (FEG-SEM, Model: Joel JSM840) set to standard scanning mode with higher probe current range and larger spot size in which the electron beam raster scans 512 lines per area scanned and irradiates 512 discrete points per lines. The HAp films were scanned using different magnifications from 30 to 200, and the energy of the incident electrons was kept constant at 30 keV in vacuum (2 107 Pa) with a absorbed beam current of 30 nA, as measured by Faraday cup, for 20 min exposure time (2.34 s/line). The dose varied, ranging from Q = 1.8 to 78 mC/cm2. It should be noted that vacuum was achieved using diaphragm and turbomolecular pumps with zero oil contaminations.
Kelvin Probe Force Microscopy (KPFM)/Atomic Force Microscopy. In order to confirm the electrical modification by irradiation, the SP was measured by Kelvin probe force microscopy (KPFM) (Agilent Technology 5500), and the topography was monitored by tapping mode AFM, using the same apparatus. A TiPt-coated Si probe (MikroMasch ultrasharp cantilever, NSC14/TiPt) with a high electrical conductivity and spring constant (k) of ∼5 N/m was used for all tapping AFM and KPFM mode experiments. The surface potential polarity difference was confirmed by using bimetal (Au/Al) standard reference sample for Kelvin probe measurement. Contact Angle Measurements. To create uniformly irradiated HAp films for contact angle measurements, sample were directly e-beam irradiated without having been masked. Sessile drop measurements were made using a CAM200 contact angle meter (KSV Instruments) with a 14969
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Figure 1. XRD pattern of hydroxyapatite. Inset left: cross section of the HAp thin film. Inset right: nanocrystallinity of hydroxyapatite revealed by AFM. CCD camera that captures drop images digitally. A 2 μL drop of deionized Milli-Q water was dispensed on the surface at constant temperature and relative humidity (22 ( 1 C and 50 ( 5%, respectively). A minimum of three samples were tested for each measurement. The contact angles were determined 30 s after deposition of the drop from images using CAM Optical contact angle software (KSV Instruments). XRD. The phase identification of the produced films has been examined by glancing angle X-ray diffraction (XRD, Philips X’Pert PRO MPD) using a graphite-monochromatized Cu Kα radiation at 40 kV and 40 mA.
’ RESULTS AND DISCUSSION The principles and potentials of e-beam writing are wellknown.18 Scheme 1 shows a schematic outline of e-beam irradiation combined with a mask to pattern HAp films with discrete areas presenting modified surface potential. The HAp thin films were deposited on silicon substrate as described in the Experimental Section. The X-ray diffraction (XRD) pattern in Figure 1 shows characteristic peaks of HAp, matching with the standard hydroxyapatite peaks (JCPDS 9-432), and reveals that the film is nanocrystalline and phase pure. The HAp film thickness has been determined to be ∼500 nm by a field-emission gun (FEG) scanning electron microscope (SEM) (Figure 1, left inset), and the atomic force microscope (AFM) surface topography (measured in tapping mode) reveals the nanocrystalline nature of HAp film with crystal size of about 100 nm (Figure 1, right inset). Finally, the XRD data demonstrate that there is out-of-plane c-axis orientation in the HAp film as the peak ratio of (300)/(002) plane peak is around 1 on an average (Figure 1). Typically, a grounded HAp film was masked to allow the creation of approximately 65 65 μm arrays of electrically modified surface potential (SP) one-dimensional (1D) line-type patterned by using e-beam irradiation and the raster scan writing capabilities of a standard scanning electron microscope (SEM). The e-beam dose, relative surface potential on samples, width of the raster lines as well as the spacing between raster lines can be controlled by modifying the SEM magnification used during writing. Figure 2a shows the effect of this SEM magnification factor (from 30 to 200) on the estimated e-beam dose
Figure 2. (a) Dependence of the magnification factor, the calculated electron beam dose (black), and average relative surface potential (blue). The relative surface potential varies from 275 ( 4.9 mV to +12.5 ( 5.2 mV. (b) Dependence of the magnification factor, the measured raster lines width (black), and spacing between raster lines (blue). Inset: number of raster lines with magnification factor.
applied to the samples (from 1.8 to 78 mC/cm2) and the consequent surface potential relative to the unexposed film. For this range of magnification factor, the resulting SP difference can be modulated from a negative (275 ( 4.9 mV) to a positive (+12.5 ( 5.2 mV) SP. Both relations follow an approximately third-order polynomial fit. Interestingly, the transition from negative to positive SP relative to the unexposed area has been observed to occur between electron beam dose of 45 and 65 mC/cm2, corresponding to the magnification factor window of 160 to 180 (Figure 2a). X-ray photoelectron spectroscopy was performed to determine the surface chemistry of the untreated and irradiated HAp films, and compositions of elements identified C, O, Ca, and P are presented in Table 1. Differences in the surface compositions of the films indicate not significant chemical modification of the surface after irradiation, and any change is restricted to the adsorbed carbon overlayer. The results of the current study are contrary to the findings on e-beam poling of HAp pellets where contaminant carbon deposits were observed after irradiation.16,17 It may perhaps be the better cleanliness of the SEM chamber and the use of a FEG source than a typical filament in the low-energy electron gun in this study that eliminates carbon contamination. Finally, the 14970
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analysis of the irradiated areas using AFM shows in Figure 3a that the e-beam writing has no effect on the topography of the film. To demonstrate the electrical modification by irradiation using a magnification factor of 30 and a resulting electron dose of 1.8 mC/cm2 (Figure 2a), the SP of HAp films was measured by using Kelvin probe force microscopy (KPFM). Figures 3b and 3e show the creation, on the irradiated film, of arrays of 1D line patterns, presenting a width of ∼1 μm (Figure 3f) and exhibiting more electronegative SP. The surface potential of those patterns shows to be ∼70 mV lower than the SP of the space between lines (Figure 3c), while the SP difference between the lines and the unexposed HAp is around 275 mV. Table 1. XPS Surface Chemical Analysis of the HAp Thin Films Untreated and Irradiated by Electron Beam (30 keV) with an Electron Dose of 1.8 mC/cm2 elements C
O
bonding
untreated
e-beam irradiated
type
(atomic %)
(atomic %)
13.3
10.2
COC, COH
CC, CH
4.0
4.5
OdCO
1.1
1.4
CO3
1.4
1.4
50.3
50.6
Ca
18.3
19.3
P Ca/P ratio
11.7 1.56
12.6 1.53
2.75
2.62
O/Ca ratio in HAp
For selective protein placements on these electrostatic domains, we employed FITC-labeled lysozyme as a model protein. The LSZ is a small “hard” protein19 of 14 kDa, as its fold is reinforced by the presence of 4 disulfide bridges, providing a strong internal coherence. The pI of LSZ is 11, and at pH < 11, the polypeptide presents an overall positive surface charge (from 8 to 17 depending on pH)20 (Scheme 1). A solution containing this protein was added to HAp films on which electrostatic domains were created, and after washing the distribution of the protein on the HAp surface was assessed by laser scanning confocal microscopy. Figure 3h clearly shows that the adsorption of LSZ is higher on the electrically modified patterned arrays as they exhibit a stronger fluorescent signal. The fluorescence intensity profile (Figure 3i) reveals a high spatial correlation between the surface potential (in Figure 3b) and the enhanced protein adsorption. A Gaussian Lorentzian shape of the x-section of the line domains is also visible in both the SP profile (Figure 3c) and the fluorescent intensity profile (Figure 3i). This shape is most probably originating from the geometry of the electron beam that can be significantly defocused when the sample surface is charged.17 Additionally, as observed by Molina et al.21 and He et al.,22 migration of charge carriers inside the material could result from the local electric fields induced by the irradiation. In order to assess the stability of the electrostatic patterns, HAp samples were, after e-beam exposure, incubated in water for 1 min before addition of the LSZ containing solution. Similar results to the patterns shown in Figure 3h have been observed. This indicates that the charge carriers are injected into
Figure 3. (a) Topography and (b) surface potential distribution on HAp film of 100 100 μm area exposed to the e-beam with an electron dose of 1.8 mC/cm2 at a magnification factor of 30. (c) SP section profile corresponding to the black line marked in (b). (d) Topography and (e) surface potential distribution on HAp film of 20 20 μm area irradiated in the same conditions as in (a) and (b). (f) SP section profile corresponding to the black line marked in (e). (g) Schematic representation of the protein interaction with electrically modified HAp. (h) Fluorescence image of the FITC-LSZ adsorbed on the 1D line-type patterned arrays. (i) Intensity scans corresponding to the white line marked in (h). 14971
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Figure 4. Fluorescence images of the FITC-LSZ adsorbed on the area exposed to the e-beam at applied SEM magnification factor of (a) 30, (b) 30 with grid patterning, (c) 60, and (d) 100.
the material sufficiently deep to prevent immediate neutralization at the surface. As the LSZ is a “hard” protein, its adsorption is enthalpydriven and does not involve any structural rearrangement, in contrast to proteins with lower structural stability, such as BSA.23 For this reason, the adsorption of LSZ is only driven by its surface properties. The protein adsorbs onto the surface and presumably accumulates, forming a loosely packed monolayer24,25 to neutralize the charges created during patterning25 (Figure 3g, top). This was confirmed by successfully washing away the bound protein from the patterned surface by using a buffer presenting a higher ionic strength (1 M NaCl) (data not shown). Hydrophobicity is known to play a critical role in protein adsorption.26 In fact, e-beam exposure of HAp pellets has been reported to increase the hydrophobicity, which can be reversed by subsequent thermal annealing.16 The effect of e-beam irradiation on the wettability of HAp films in our case was nominal as the typical contact angle increases only slightly after irradiation to 97.9 ( 2.33 from a 91.8 ( 1.1 for unirradiated film. This shows that the increased adsorption of the model protein on localized charged areas is not driven by a decrease in wettability of the HAp film. This is in contrast to the findings of Aronov and coauthors that low-energy electron beam (100 eV) irradiation dramatically decreases the wettability of HAp pellets with the e-beam dosage.9 However, these authors have also noted that the wettability reduction is less dramatic when the excitation energy increases from 100 V to 1 keV. Higher excitation voltage usually results in deeper penetration depth of electrons, and a lower variation of the wettability is expected9 as it is for using a 30 keV excitation voltage in our study. The adsorption of LSZ protein onto the charged domains directly written by e-beam thus has been successfully tailored by modifying the surface potential of HAp film and independently of modifying its surface chemistry, hydrophobicity, or roughness. To demonstrate that our method can be used as a template to create directly more complex patterned arrays, we have also prepared cross patterns of electrostatic domains on HAp films. These cross-patterns have been created by irradiating HAp film by SEM raster similar to what has been used to create a pattern as
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in Figure 4a. The only difference is that after creating an initial array of 1D lines the film was rotated by 90, and the same process of 1D line creation by e-beam raster was repeated. Figure 4b demonstrates the creation of this cross patterns and the specific deposition of FITC-LSZ. Electrically modified patterned arrays with domains of smaller widths can be created by using the magnification power offered by the SEM. As shown in Figure 2b, the raster lines width and the spacing between raster lines decrease exponentially as the magnification factor increases. Theoretically from this relation, 1D line-type arrays presenting a width lower than 150 nm could be achieved by using higher magnification factor, which is ∼500. The density of electrically modified patterns will also increase as shown in Figure 2b (inset). Unfortunately, the spacing between domains also decreases with magnification factors, and if such spacing is too small, it will be very difficult to discern the domains from each other by using a KPFM. To demonstrate the effect of higher SEM magnification on the protein adsorption of e-beam exposed HAp, we have applied SEM magnification factors of 60 and 100 for e-beam poling, while keeping other parameters identical. This gradually increased the density of patterns on the exposed film as visible in the fluorescence images of HAp films, when LSZ proteins were placed on such domains (Figure 4c,d). Figure 4a,c,d clearly shows that the protein absorption is still higher on negative SP of the 1D line-type arrays for all three different magnification factors used for e-beam exposure. As expected, the increased magnification factor resulted in the creation of smaller features and in an increased density of charged patterns. If only 9 lines could be written at 30 per square feature, 18 and 28 lines could be created with a magnification factor of 60 and 100, respectively. Contact angle measurements confirm that the wettability of the exposed HAp is marginally affected by the irradiation (90.04 ( 2.9 at 60) and that the protein adsorption is still mainly driven by attractive electrostatic interactions between negatively charged HAp and positively charged LSZ.27 As depicted in Figure 2a, with increasing magnification factor the area of scanning decreases and as result the electron dose per raster line increases due to longer irradiation time at a given scanning speed. When a thin polymer or electret film, with thickness R, is irradiated by an e-beam of higher than a certain excitation energy (generally E > 2 keV), a negative surface charge (σT) is created at the average penetration depth of the film (r0 ). During irradiation, if one side of the metalized sample is grounded (at the ground potential), then the measured surface potential Us is given by Us ¼
σ T ðR r 0 Þ εε0
ð1Þ
where ε0 is the vacuum permittivity and ε the dielectric permittivity of the film.28 A higher electron dose will result in a subsequent higher penetration depth of electrons (r0 ). According to eq 1, the latter will result in decreased value of (R r0 ), consequently leading to an increase of the surface potential (Us). Therefore, it is, in theory, possible not only to modulate the geometry of the patterned arrays (e.g., to create nanolines) but also to modulate the local surface potential. This suggests that for a certain electron dosage the electron beam should penetrate the film layer and the surface potential (Us) will diminish. In other words, it will create relatively less negative SP on a HAp surface which initially possessed a negative SP. 14972
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Figure 5. (a) Topography and (c) surface potential distribution on HAp film of 90 90 μm area exposed to the e-beam at applied SEM magnification factor 200 during charging with an electron dose of 78 mC/cm2. (d) SP section profile corresponding to the black line marked in (c). (b) Wide-view fluorescence image of the FITC-LSZ adsorbed on the HAp film patterned as described above. (e) Fluorescence image of one array unit comprising 60 1D line-type patterns; Inset: patterns exhibiting a width of 175 nm. (f) Intensity scan profile corresponding to the white line marked in (b).
This phenomenon was observed when the HAp film was irradiated using SEM at an electron dose 78 mC/cm2 with a corresponding magnification factor of 200, as shown by KPFM imaging (Figure 5c). The individual 1D lines could not be resolved by KPFM due to their closer spacing; however, the corresponding SP section profile (see Figure 5d) clearly shows that the arrays present an overall 12.5 ( 5.2 mV higher SP than unexposed areas. The LSZ was further used to probe the newly created positive SP patterns. The analysis of the fluorescent signal by confocal microscopy shows (Figure 5b,e,f) that each 65 65 μm pattern exhibits a much lower fluorescent signal due to a lower protein adsorption. The change in protein behavior cannot be attributed to a change in HAp surface roughness as AFM analysis (Figure 5a) shows that the irradiation at an electron dose of 78 mC/cm2 has no effect on the topography of the film, similar to the observation made on films exposed to lower dosage of e-beam. The inset in Figure 5e shows that within a 65 65 μm area a total of 60 1D lines of adsorbed proteins have been created.
A lower adsorption of the LSZ on areas with modified SP thus means that repulsive interactions prevail (Figure 3g, bottom). Previously, it has been shown that the wettability of HAp can be tailored by e-beam irradiation at low energy.16,17 In clear contrast to these studies, it is unlikely that the change in wettability can explain the behavior of the protein in our present investigation as in our case the change in hydrophobicity is not significant. More importantly, it has been shown that the adsorption of LSZ increases with the hydrophobicity.29 As at pH 7.3, the LSZ is positively charged, and at low ionic strength (1 103 mol/L) used in the present study, the electrostatic interactions are expected to play a major role in the binding of this protein to HAp. Our results clearly thus show that the decrease in protein adsorption can mainly be explained by repulsive electrostatic interactions. Additionally, since hydroxyapatite has a lower dielectric constant than water, the image charge produced by a positive protein will also be positive, due to the ClausiusMossotti factor, and may participate in preventing protein adsorption. 14973
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Langmuir Of phenomenal importance is the observation of 1D lines of electrostatic domains with a width as small as 175 nm. Figure 2b shows that through a control of the magnification factor the width of the raster line can be reduced to below 150 nm. An associated problem to this approach is that the domain spacing will also decrease, thus making discerning nanodomains by KPFM or fluorescent microscopy extremely difficult. However, the principles of e-beam modification of surface potential have been clearly outlined and can be used to achieve domains of nanoscale dimensions by using an electron beam probe used in energy dispersive X-ray spectroscopy. The typical probe size of such an EDX probe with a FEG source is ∼10 nm. This can be used in conjunctions with the information disclosed in the current article to create nanodomains that will selectively interact with proteins through electrostatic interactions. This may provide a much more convenient methodology compared to current approaches nanopositioning of proteins via electrostatic interactions to study single protein interactions.14 The approach elucidated in the present study can also be applied to protein delivery, microfluidic devices, protein separation at nanoscale, and nanobiosensors.
’ CONCLUSION One-dimensional electrostatic domains of micrometer and submicrometer width have been created by exposing HAp films to e-beam available from a typical SEM raster. These domains have shown to tailor the spatial regulation of protein adsorption through electrostatic interactions independently of effects from changes in topography, wettability, or surface chemistry. By varying the electron dose, the surface potential can not only be modified to create highly negatively charged domains but that more positively charged patterns can be created. The physical modification of HAp can be used to modify its impact on different cellular activities and function via precisely immobilized proteins. The degree of control that this approach enables will be beneficial for better understanding of protein adsorption on HAp and other biomaterials and the implications of such adsorption in human tissue/implant applications. ’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected]. Author Contributions ^
Equal contribution and co-first author.
’ ACKNOWLEDGMENT This project has been funded with support from the European Commission (EC NMP4-SL-2008-212533 - BioElectricSurface). This publication reflects the views only of the authors, and the Commission cannot be held responsible for any use which may be made of the information contained therein. The AFM facility is enabled under the framework of the INSPIRE program, funded by the irish government’s Programme for Research in Third Level Institutions, Cycle 4, National Development Plan 2007-2013.
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(2) Andrade, J.; Hlady, V. Protein adsorption and materials biocompatibility: A tutorial review and suggested hypotheses. In Biopolymers/ Non-Exclusion HPLC; Springer: Berlin, 1986; Vol. 79, pp 163. (3) Keselowsky, B. G.; Collard, D. M.; García, A. J. J. Biomed. Mater. Res., Part A 2003, 66A (2), 247–259. (4) Ambard, A. J.; Mueninghoff, L. J. Prosthodontics 2006, 15 (5), 321–328. (5) Blanco, E. M.; Horton, M. A.; Mesquida, P. Langmuir 2008, 24 (6), 2284–7. (6) Rechendorff, K.; Hovgaard, M. B.; Foss, M.; Zhdanov, V. P.; Besenbacher, F. Langmuir 2006, 22 (26), 10885–10888. (7) Xu, L.-C.; Siedlecki, C. A. Biomaterials 2007, 28 (22), 3273–3283. (8) Lubarsky, G. V.; Browne, M. M.; Mitchell, S. A.; Davidson, M. R.; Bradley, R. H. Colloids Surf., B 2005, 44 (1), 56–63. (9) Aronov, D.; Rosen, R.; Ron, E. Z.; Rosenman, G. Process Biochem. 2006, 41 (12), 2367–2372. (10) Kandori, K.; Horigami, N.; Kobayashi, H.; Yasukawa, A.; Ishikawa, T. J. Colloid Interface Sci. 1997, 191 (2), 498–502. (11) Kawachi, G.; Watanabe, T.; Ogata, S.-i.; Kamitakahara, M.; Ohtsuki, C. J. Ceram. Soc. Jpn. 2009, 117 (1367), 847–850. (12) Kandori, K.; Masunari, A.; Ishikawa, T. Calcif. Tissue Int. 2005, 76 (3), 194–206. (13) Sivagnanam, V.; Sayah, A.; Vandevyver, C.; Gijs, M. A. M. Sens. Actuators, B 2008, 132 (2), 361–367. (14) Kumagai, S.; Yoshii, S.; Yamada, K.; Matsukawa, N.; Fujiwara, I.; Iwahori, K.; Yamashita, I. Appl. Phys. Lett. 2006, 88 (15), 153103–3. (15) Jacobs, H. O.; Whitesides, G. M. Science 2001, 291 (5509), 1763–1766. (16) Aronov, D.; Rosenman, G.; Karlov, A.; Shashkin, A. Appl. Phys. Lett. 2006, 88 (16), 163902–3. (17) Plecenik, T.; Tofail, S. A. M.; Gregor, M.; Zahoran, M.; Truchly, M.; Laffir, F.; Roch, T.; Durina, P.; Vargova, M.; Plesch, G.; Kus, P.; Plecenik, A. Appl. Phys. Lett. 2011, 98 (11), 113701–3. (18) Fontana, R. E.; Katine, J.; Rooks, M.; Viswanathan, R.; Lille, J.; MacDonald, S.; Kratschmer, E.; Tsang, C.; Nguyen, S.; Robertson, N.; Kasiraj, P. IEEE Trans. Magn. 2002, 38 (1), 95–100. (19) Arai, T.; Norde, W. Colloids Surf. 1990, 51, 1–15. (20) Xie, Y.; Zhou, J.; Jiang, S. J. Chem. Phys. 2010, 132 (6), 065101. (21) Molina, P.; Ramírez, M. O.; García-Sole, J.; Bausa, L. E. Opt. Mater. 2009, 31 (12), 1777–1780. (22) He, J.; Tang, S. H.; Qin, Y. Q.; Dong, P.; Zhang, H. Z.; Kang, C. H.; Sun, W. X.; Shen, Z. X. J. Appl. Phys. 2003, 93 (12), 9943–9946. (23) Kandori, K.; Takaragi, S.-i.; Murata, K.; Ishikawa, T. Adsorpt. Sci. Technol. 2005, 23 (10), 791–800. (24) Kim, D. T.; Blanch, H. W.; Radke, C. J. Langmuir 2002, 18 (15), 5841–5850. (25) Barroug, J.; Lemaitre, A.; Rouxhet, P. G. Colloids Surf. 1989, 37, 339–355. (26) Rabe, M.; Verdes, D.; Seeger, S. Adv. Colloid Interface Sci. 2011, 162 (12), 87–106. (27) Kawasaki, K.; Kambara, M.; Matsumura, H.; Norde, W. Colloids and Surfaces B. Biointerfaces 2003, 32 (4), 321–334. (28) Sessler, G. M.; Breazeale, M. A. J. Acoust. Soc. Am. 1988, 84 (6), 2298–2299. (29) Kandori, K.; Mukai, M.; Fujiwara, A.; Yasukawa, A.; Ishikawa, T. J. Colloid Interface Sci. 1999, 212 (2), 600–603.
’ REFERENCES (1) Allen, L. T.; Tosetto, M.; Miller, I. S.; O’Connor, D. P.; Penney, S. C.; Lynch, I.; Keenan, A. K.; Pennington, S. R.; Dawson, K. A.; Gallagher, W. M. Biomaterials 2006, 27 (16), 3096–3108. 14974
dx.doi.org/10.1021/la203491q |Langmuir 2011, 27, 14968–14974
