Elaboration of Nanostructured Biointerfaces with Tunable Degree of

Sep 22, 2011 - This study shows that electrophoretic deposition (EPD) is a fast and efficient technique for producing protein nanotube-based biointerf...
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Elaboration of Nanostructured Biointerfaces with Tunable Degree of Coverage by Protein Nanotubes Using Electrophoretic Deposition Deepak M. Kalaskar, Claude Poleunis, Christine Dupont-Gillain, and Sophie Demoustier-Champagne* Institute of Condensed Matter and Nanosciences - Bio & Soft Matter (IMCN/BSMA), Université catholique de Louvain, Croix du Sud, 1. B-1348 Louvain-la-Neuve, Belgium S Supporting Information *

ABSTRACT: This study shows that electrophoretic deposition (EPD) is a fast and efficient technique for producing protein nanotube-based biointerfaces. Well-shaped collagen-based nanotubes of controlled dimensions are synthesized by a template method combined with the layer-by-layer (LbL) assembly technique. Separation of nanotubes from the template material and collection of nanotubes on ITO glass carried out by EPD leads to a fairly homogeneous distribution of protein nanotubes at the support surface. Biointerfaces with different and tunable densities of protein nanotubes are obtained by changing either the applied voltage, solution concentration of nanotubes, or deposition time. Moreover, it is proved that the collected nanotubes are templatefree and keep their biofunctional outermost layer after EPD. A preliminary study of the behavior of preosteoblasts cells with the elaborated biointerfaces indicates a specific interaction of cells with the nanotubes through filopodia. This contribution paves the way to the easy preparation of a large variety of useful nanostructured collagen and other protein-based interfaces for controlling cell−surface interactions in diverse biomaterials applications.

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Collagen is the main structural protein found in ECM and is broadly used as a substrate or scaffold for cell attachment, proliferation, and differentiation.11 Very recently, we reported a simple, versatile, and cost-effective approach for synthesizing collagen-based nanotubes with controlled and tunable dimensions.12,13 The strategy is based on the nanotube template synthesis through layer-by-layer (LbL) assembly. An important feature of the template synthesis method, using track-etched polycarbonate membranes as templates, is the ability to control the dimensions of the resulting nanotubes over a wide range. The outside diameter of the nanotubes, determined by the diameter of the template pores, can indeed be varied from 30 nm up to a few micrometers, and the length of the nanotubes, determined by the thickness of the template, can range from 5 to 50 μm.14,15 The LbL technique, based on the alternate adsorption of oppositely charged species, has attracted much interest for biomaterials applications as it is a versatile technique, allowing the control of the multilayers properties (composition, thickness, and function).16,17 Combining this LbL process with the template method allows the wall thickness to be controlled, and correspondingly the inside diameter of template-synthesized nanotubes, by adjusting the number of layers of the material deposited along the pore walls.

INTRODUCTION

The past decades of scientific research in the field of biomaterials have revealed that in order to achieve predictable cellular responses a fine balance between chemistry, topography, and mechanical properties of materials is required.1,2 Engineering surfaces that closely mimic the complexity and functionality of the extracellular matrix (ECM) is thus currently one of the challenging aims in biomaterials science. Indeed, though cells have micrometer dimensions, they evolve in vivo in close contact with the ECM, a matrix with topographical and structural features of nanometer size, made of nanofibers of proteins such as collagen, elastin, and keratin that provide biological and physical support for cell attachment, proliferation, migration, differentiation, and ultimately cell fate. Artificial nanofibers, whose dimensions, strength, high surface area/volume ratio, and biocompatibility resemble those of natural ECM fibers, could play the same critical role in tissue regeneration process. Consequently, modification of biointerfaces using proteinbased nanocomponents has attracted an increasing interest in recent years. This approach benefits from progress made in biomacromolecules self-assembly and in the ability to prepare nanostructured materials with tailored properties, in particular, nanotubes and nanowires. Several recent papers describe the fabrication of nanotubes made from common globular proteins such as glucose oxidase, hemoglobin, cytochrome c, and serum albumin.3−10 © 2011 American Chemical Society

Received: August 19, 2011 Revised: September 21, 2011 Published: September 22, 2011 4104

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preosteoblast cells (product number CRL-2594), was purchased from the American Collection of Cell Culture (ATCC-LGC Standards S.a.r.l., France). Preparation of Polyelectrolyes and Collagen Solution. Polyelectrolyte solutions of PSS, PAH, and Flu-PAH were prepared in 100 mM acetate buffer (pH 4.7) at a concentration of 1 mg/mL. Collagen solution was also prepared by dissolution in acetate buffer solution at a concentration of 500 μg/mL. All solutions were freshly prepared before use. QCM-D. The buildup of (COL/PSS)n multilayers on flat substrate was monitored in situ by quartz crystal microbalance. Measurements were performed with a Q-Sense E4 System (Gothenborg, Sweden) at a temperature of 20.0 ± 0.1 °C. The crystal used is a thin AT-cut quartz coated with a thin SiO2 film (thickness ∼50 nm). Oscillations of the crystal at the resonant frequency (5 MHz) or at one of its overtones (15, 25, 35, 45, 55, 65 MHz) were obtained when applying ac voltage. The variation of the resonance frequency (Δf) was monitored upon adsorption of the polyelectrolytes. Solutions were injected into the measurement cell using a peristaltic pump (Ismatec IPC-N 4) at a flow rate of 50 μL/min. Prior to the multilayers buildup, acetate buffer solution was injected to establish the baseline. The construction of the (COL/PSS)n multilayers was performed as follows: First, COL solution was brought into the measurement cell for 30 min, allowing the establishment of the adsorption equilibrium at the crystal surface. Subsequently, rinsing was performed for 10 min at a flow rate of 50 μL/min using acetate buffer solution. PSS and collagen were then alternately injected according to the same procedure, except that the adsorption time for PSS was 15 min. After the buildup of 3.5 bilayers, the response of the sensor could not be longer monitored for all harmonics, probably because the film became too thick. Synthesis of Nanotubes. Synthesis of nanotubes was carried out by using classical layer-by-layer (LbL) method using polycarbonate track-etched templates as reported previously.12,13 Briefly, LbL assembly was performed by successive adsorption steps of COL (or PAH) and PSS in polycarbonate track-etched membranes with pore size of 500 nm. Throughout the experiments, acetate buffer at pH 4.7 was used. Membranes of 13 mm in diameter were immersed in cationic solution of COL for 2 h or PAH for 30 min, followed by rising in acetate buffer for 4 min. Alternative anionic layer was built by immersing the membranes for 30 min in a PSS solution. When the required number of bilayers was reached, samples were cleaned on both faces by using cotton bud with alkaline solution of aqueous NaOH (pH 12). Samples were dried overnight at room temperature. Fluorescently labeled nanotubes were also synthesized by incorporating a layer of poly(fluorescein isothiocyanate allylamine hydrochloride) (Flu-PAH) during LbL assembly along with PSS. By using this procedure, two different types of nanotubes were synthesized, namely (COL/PSS)3(Flu-PAH/ PSS)3 with collagen as outer layer and (Flu-PAH/PSS)3(COL/PSS)3 with Flu-PAH as outer later. Collection of Nanotubes by EPD. Nanotubes were collected on conducting solid surfaces. The nanotubes, synthesized into pores of polycarbonate membranes, were first liberated by dissolution of the template in DMF. 10 mm2 sample was cut into small pieces and dissolved in 4−5 mL of DMF and sonicated 2 or 3 times for 30 s until the membrane was completely dissolved and no particles in suspension were visible to naked eyes. This solution was filled in the EPD cell for collection of nanotubes. All electrochemical experiments were performed with a CHI660B electrochemical workstation (CH Inc.) in a specially design EPD cell (Figure 1a) at room temperature. Applied voltage and time of EPD were controlled by using chronoamperometry function of CHI electrochemical workstation software (version 5.19). During EPD, ITO-coated conducting glass coverslips were used as both working and counter electrodes, whereas a platinum wire was used as a pseudoreference electrode, as shown in Figure 1b. The distance between working and counter electrodes was maintained at 5 mm by using PDMS casing. The effect of applied voltage, solution concentration of nanotubes, and time of EPD on the density of collected nanotubes was studied using both (COL/PSS) 3(Flu-PAH/PSS)3 and (Flu-PAH/PSS)3(COL/PSS)3 nanotubes. When EPD was completed, the working electrode was rinsed 4−5 times with

In order to investigate the interaction of these protein nanotubes with cells, a remaining key issue is to find an efficient way for the collection and immobilization of the nanotubes on a surface. Indeed, one of the most important challenges for the practical use of template-synthesized nanotubes is the separation of the nanotubes from the template material (i.e., polycarbonate). Techniques such as filtration or centrifugation12,18 are often used for that purpose, but they are not very efficient in terms of extraction and collection of nano-objects. The major problems with these processes are that they usually lead to a significant fraction of broken nanotubes, retention of some polycarbonate on the outer surface of the nanotubes, and the formation of numerous aggregates of nanotubes. To overcome these shortcomings, we developed a new method based on electrophoretic deposition (EPD) for the collection of protein nanotubes and the elaboration of nanostructured biointerfaces. EPD of carbon nanotubes on conducting surfaces has already been widely reported.19,20 EPD of ceramic nanoparticles (less than 100 nm) has also been used for the production of a variety of materials including homogeneous coating on biomaterials. A complete review on this topic is reported elsewhere by Corni et al.21 Apart form carbon nanotubes and conventional ceramic materials, such as hydroxyapatite, numbers of other materials from micro- to nanoscale dimensions were also used for creating new biofunctional surfaces by EPD.22 In particular, to improve biocompatibilty of materials, thin coatings of various polysaccharides, such as chitosan, alginate, and hyaluronic acid, were studied by the EPD process. Finally, recent work reported by Baker et al. showed the preparation of self-supported collagen films by applying an electric field to an aqueous protein solution.23 In this contribution, we report the collection of collagenbased nanotubes by EPD. We evaluate the impact of the main determining experimental parameters (applied voltage, concentration of nanoparticles in the suspension, and time of electrophoretic deposition) on the density of collected nanotubes. To check that nanotubes deposited by EPD do not lose their structural integrity, are not aggregated on the collecting surface, and are free of polycarbonate, the various elaborated nanostructured biointerfaces were characterized by scanning electron microscopy (SEM), fluorescence microscopy, and time-of-flight secondary ion mass spectrometry (ToF-SIMS).

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EXPERIMENTAL SECTION

Materials. Poly(sodium-p-styrenesulfonate) (PSS, Mw ∼70 kDa) was purchased from Acros Organics. Poly(allylamine hydrochloride) (PAH, Mw ∼15 kDa), poly(fluorescein isothiocyanate allylamine hydrochloride) (Flu-PAH, with fluorescein:PAH (Mw ∼ 15 kDa) = 50:1), sodium acetate, acetic acid, dichloromethane (CH 2Cl2), dimethylformamide (DMF), and isopropanol were purchased from Sigma-Aldrich. Type I Collagen from calf skin (4 mg/mL at pH 3.0) was purchased from AutogenBioclear (UK). Type I collagen was chosen as it is recognized by many different types of cells and is therefore of great interest for studying cell−material interactions. Polycarbonate tracketched membranes with a thickness of 21 μm, a pore density of 4 × 107 pores cm−2, and an average pore size of 500 nm, supplied by it4ip, were used as nanoporous templates. Indium−tin oxide (ITO)-coated borosilicate glass coverslips of 20 mm × 20 mm (8−12 ohm resistivity) were purchased from SPI Supplies/Structure Probe, Inc., West Chester, PA. Fetal bovine serum was purchased from Harlan, Loughborough, UK. MEM α-medium with low glucose (1 g L−1) containing deoxyribonucleosides, L-glutamine, ribonucleosides, penicillin−streptomycine, and Alexa Fluor 594 phalloidin were bought from Invitrogen, Belgium. Sodium pyruvate solution was purchased from Sigma-Aldrich, Belgium. Vectashield mounting media with DAPI was purchased from Vectors Laboratories, Burlingame, CA. MC3T3-E1 Subclone 14, 4105

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Cell Culture. Nanotube-coated ITO glass slides were tested for cell culture using MC3T3 (subclone 14) preosteoblast cells. Cells were cultured in α-MEM medium supplemented with 10% of fetal bovine serum, 1% sodium pruvate, and 1% penicillin/streptomycine, referred to as cell culture media. Cells were passaged using trypsin−EDTA every 3 days and maintained at 5% CO2 in an incubator at 37 °C. The prepared biointerfaces were sterilized using ethanol for 30 min prior to cell seeding. Cells were seeded at a density of 20 000 cells/cm 2 for a period of 24 h on nanotube-coated surfaces to investigate cell− nanotube interactions. Cell shape and morphology were investigated by F-actin staining, while nuclei were stained using DAPI. For fluorescence microscopy, a Leica DM RA 2 microscope equipped with a Leica DFC 300Fx camera was used. All images were processed and scaled using Leica Q Win software (Version 3.0).

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RESULTS AND DISCUSSION LbL Assembly on Flat Substrate. The isoelectric point (iep) for collagen molecule reported in the literature varies from 4.8 to 9.24−26 The absence of a specific value of iep for collagen can be explained by the variable charge distribution along its fibrillar structure. Our recent work investigating the mechanism of interaction of collagen (COL) within polyelectrolyte multilayers showed that collagen can be used efficiently as a polycation.12,13 In these previous studies, we used an anchoring layer of poly(allylamine) (PAH) to initiate the LbL assembly of COL and poly(styrenesulfonate) (PSS). By doing so, when removing the polycarbonate template, the outermost layer of the nanotubes will be PAH and not collagen. In the present study, we therefore investigated the LbL assembly on flat surfaces of COL (used as polycation) and PSS (used as polyanion) without anchoring PAH layer, using in situ QCM-D measurements. The shifts in resonant frequency (Δf) and dissipation values (ΔD) after adsorption of COL followed by PSS were recorded (Supporting Information, Figure S1). The appreciably higher values of Δf and ΔD for COL compared to PSS indicate that collagen forms a thicker but softer and more viscoelastic layer after its adsorption than PSS. The observed decrease in dissipation when PSS is added suggests a contraction of the film. These results are consistent with our previous study12 where PAH was used as anchoring layer and show that the anchoring layer has no significant impact on the buildup of COL and PSS multilayers. Synthesis and Characterization of Collagen-Based Nanotubes. Based on the results obtained on flat surfaces, different COL/PSS-based nanotubes (NTs) with either COL or PAH as outer layer were built up by LbL deposition in nanoporous polycarbonate membranes. To facilitate the characterization, fluorescent nanotubes were also synthesized by incorporating fluorescein-labeled PAH (Flu-PAH) during the LbL assembly. Two types of nanotubes made of six bilayers were prepared, namely (COL/PSS)3(Flu-PAH/PSS)3 with collagen as outer layer and (Flu-PAH/PSS) 3(COL/PSS)3 with PAH as outer layer. SEM pictures of 500 nm diameter (COL/PSS)3(Flu-PAH/PSS)3 nanotubes collected by dissolution of the polycarbonate membrane and filtration on silver membranes show the presence of well-shaped protein nanotubes but also of a significant number of fragmented or aggregated nanotubes (Figure S2, Supporting Information). Moreover, by using this dissolution/filtration procedure, some residual polycarbonate was always present on the outer part of the nanotubes, as well as on the filter, even after several rinsings with the solvent. With the aim of overcoming all these drawbacks, we therefore developed and optimized a new method, based on EPD, for the collection of nanotubes and preparation of nanostructured biointerfaces. Though electrophoretic deposition of

Figure 1. Schematic (a) and picture (b) representation of the device used for collagen nanotubes collection using electrophoretic deposition (EPD). DMF to ensure that any loosely attached nanotubes were removed from the ITO surface. Characterization of Nanotubes by SEM and Fluorescence Microscopy. For scanning electron microscopy (SEM) analysis, the track-etched PC membranes were deposited on a silver membrane (average pore diameter of 0.45 mm, SPI Supplies) and then dissolved and rinsed with dichloromethane to entrap the liberated nanotubes. The samples were then imaged using a field effect gun digital scanning electron microscope (FE-SEM, DSM 982 Gemini from LEO) operating at 1 kV. The characterization of nanotubes deposited by EPD on ITOcoated glass coverslips was carried out by using both SEM and fluorescence microscopy. No further sample preparation was necessary for SEM microscopy due to conducting properties of ITO glass coverslips. For fluorescence microscopy, a Leica DM RA 2 microscope equipped with a Leica DFC 300Fx camera was used. All images were processed and scaled using Leica Q Win software (Version 3.0). Quantification of nanotubes density after electrophoretic deposition on ITO-coated glass was performed by using fluorescence microscopy. Ten random fields of views were captured, and nanotubes were manually counted using cell counter by using Image J (version 1.43u) analysis software. Time-of-Flight Secondary Ion Mass Spectrometry (ToF-SIMS). Chemical characterization and imaging of nanotubes were carried out by using ToF-SIMS. An Ion-TOF ToF-SIMS V instrument (IONTOF, GmbH, Münster, Germany) was used. ITO-coated glass coverslips which served as working and counter electrodes in the EPD process were mounted on sample holders. A Bi3+ metal ion source was used to produce a primary beam using an acceleration voltage of 60 kV. An ac target current of 0.002 pA with a bunched pulse width of less than 1.4 ns was used. Both positive and negative secondary ion species were analyzed. For imaging, a raster of 256 × 256 data points over an area of 100 × 100 μm2 was used. The total primary ion beam dose for each analyzed area was always kept below 1012 ions cm−2, ensuring static conditions. Lateral resolution of ∼0.3 μm and mass resolution m/Δm > 5000 at 29 m/z were maintained for acquisition of both images and corresponding spectra for positive and negative ions. All data analysis was carried out using the software supplied by the instrument manufacturer, SurfaceLab (version 6.1). 4106

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inorganic nanotubes, such as carbon nanotubes27 or tungsten oxide nanorods,28 on ITO-coated glass were previously reported, this is, to our knowlegde, the first time that EPD is used for the separation and collection of organic nanotubes, such as protein/polymer-based nanotubes. In this technique, separation and deposition of nanoparticles is achieved based on their surface charge. Once the nanotubes are formed into polycarbonate template, the template is dissolved in dimethylformamide (DMF), leading to a suspension of free nanotubes and polycarbonate. This suspension is filled between two conducting electrodes made of ITO-coated glass, separated by a PDMS casing (Figure 1). Nanotubes that contain cationic polymers (COL or PAH) in their outermost layer are expected to migrate toward the cathode when subjected to an external electric field, while polycarbonate should not. This could lead to an efficient separation of the nanotubes from the polycarbonate template and, at the same time, to the creation of a surface coated with protein-based nanotubes. Figure 2 shows SEM and fluorescence images of (COL/PSS)3(Flu-PAH/PSS)3 nanotubes collected by EPD at 7.5 V on

polycarbonate. Second, DMF is nonvolatile and has a high dielectric constant of ∼37.4 at 25 °C.33 These DMF properties ensure that the current passing through the system during EPD will mainly come from particles motion and not from solvent decomposition under the applied electric field. The influence of the main physical parameters of the EPD, including the applied voltage, the concentration of nanotubes in suspension, and the deposition time, on the deposition of two different types of protein-based nanotubes was then investigated. In order to systematically evaluate the impact of each parameter, the distance between the two electrodes was maintained constant at 5 mm with help of a PDMS casing (Figure 1). The design of the EPD cell was slighly adapted from EPD cells described in the literature.34,35 At first, to study the influence of nanotubes concentration in the suspension, the EPD cell was filled with a DMF suspension containing an increasing number of collagen-based nanotubes (ranging from 3 × 105 to 6 × 106 NTs/mL of DMF), and a constant voltage of 7.5 V was applied for 1000 s. As shown in Figure S3 (Supporting Information), the current density is increasing with the concentration of nanotubes in suspension. This effect was coupled with an increasing density of deposited nanotubes on the cathode (working electrode), while no nanotubes deposition was observed on the anode (counter electrode). Migration and deposition of collagen nanotubes exclusively on the cathode indicate that nanotubes have a net positive charge. By increasing the nanotubes concentration in the suspension up to 3 × 106 NTs/mL, a linear increase of the density of deposited nanotubes is achieved for both types of investigated nanotubes: (COL/PSS) 3(Flu-PAH/PSS)3 or (Flu-PAH/PSS)3(COL/PSS)3 (Figure 3a). The higher density of deposition obtained with the nanotubes containing PAH in their outer layer may result from a higher positive surface charge density on this type of nanotubes than on those with COL as outermost layer. Figure 3b−d shows fluorescence images of (COL/PSS)3(Flu-PAH/PSS)3 nanotubes collected by EPD at 7.5 V from suspensions at 3.75 × 105, 3 × 106, and 6 × 106 NTs/mL. On these pictures, a significant increase of the density of nanotubes present on the ITO glass surface (from ∼5000 to ∼50 000 NTs/cm2) is clearly observed with increasing concentration of nanotubes in the suspension used for EPD. Second, the effect of applied voltage on the collection of nanotubes was investigated. For this study, the concentration of nanotubes in suspension (1.2 × 106 NTs/mL in DMF) and time of deposition (1000 s) were maintained constant for all the experiments. As shown in Figure 4, when increasing the applied voltage, a progressive increase in density of collected nanotubes is observed up to 8 V for both types of nanotubes. However, further increase of the potential does not lead to a further increase of the nanotubes density on the collecting surface. Therefore, in order to investigate the effect of the deposition time on nanotubes collection, a constant voltage of 7.5 V was applied, and the concentration of nanotubes in suspension was fixed at 1.5 × 106 NTs/mL in DMF. Figure 5 shows the evolution of the amount of collected nanotubes with the deposition time. In the time range investigated (0−1000 s), a linear increase in density of collected nanotubes was observed with increasing time of EPD for both types of nanotubes. Whatever the studied deposition time, a higher density was obtained with the nanotubes containing PAH in their outermost layer compared to those containing COL as outermost layer. As previously mentioned, this can be

Figure 2. (a) SEM image of (COL/PSS)3(Flu-PAH/PSS)3 nanotubes deposited on ITO-coated glass by EPD at 7.5 V. (b) Corresponding fluorescence image of the nanotubes. Scale bars: 50 μm.

ITO-coated glass. These pictures show that nanotubes of uniform dimensions are collected by EPD and that they are randomly and homogeneously dispersed on the collecting surface. Factors Affecting Collection of Nanotubes Using EPD. EPD is based on the presence of small charged particles suspended in a suitable liquid which, on the application of a dc electric field, are forced to move toward the oppositely charged electrode where they can form a homogeneous coating.20 Despite ever increasing use of EPD for a large number of applications, the mechanism of EPD remains largely unexplained.21 The most fundamental mechanism of EPD described in the literature is based on the Derjaguin−Landau−Verwey−Overbeek (DLVO) theory.29 However, other theories have been proposed to explain particle−electrode interactions and kinetics of deposition, which include flocculation by particle accumulation, particle charge neutralization, electrochemical particle coagulation, electrical double layer distortion, and thinning mechanism. 21,29−32 Although, it is generally admitted that two groups of parameters determine the characteristics of the EPD process: those related to the suspension of particles to be collected and those related to physical parameters of the electrical process. For the efficient collection of particles by EPD, the first critical point is to produce a stable suspension of charged particles in an appropriate solvent, so that particles can be free to move when an electric field is applied. EPD was already carried out from aqueous or organic solvents or even from a mixture of both.20,21 In this work, DMF was chosen as solvent for two main reasons. First, polycarbonate readily dissolves in DMF,9 and therefore a good suspension is produced, containing both nanotubes and 4107

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Figure 3. (a) Effect of concentration of nanotubes suspension on the density of nanotubes collected at 7.5 V for 1000 s. Error bars are standard error from the mean (n = 10). Fluorescence images of (COL/PSS)3(Flu-PAH/PSS)3 nanotubes deposited by EPD on ITO glass from suspensions at a concentration of (b) 3.75 × 105, (c) 3 × 106, and (d) 6 × 106 NTs/mL. Scale bars = 100 μm.

Figure 4. Effect of the applied voltage on the density of nanotubes collected by EPD from a suspension containing a fixed concentration of nanotubes (1.2 × 106 NTs/mL in DMF) for 1000 s. Error bars are standard error of the mean (n = 10).

Figure 5. Effect of time of deposition on collection of nanotubes from a fixed concentration of nanotubes (1.5 × 106 NTs/mL in DMF) at a constant applied voltage of 7.5 V. Error bars are standard error of the mean (n = 10).

attributed to a higher positive surface charge density on the (Flu-PAH/PSS)3(COL/PSS)3 compared to the (COL/PSS)3(Flu-PAH/PSS)3 nanotubes. The density of deposited nanotubes was found to be directly related to the current density recorded during the EPD process (Figure S3). It is therefore possible to tune this density by adjusting the current density passing through the system, and this parameter can simply be adjusted by varying either the concentration of nanotubes in the suspension, the applied voltage, or the time of EPD. The EPD process is thus perfectly adapted for creating surfaces coated with the desired density of polymer- or protein-based nanotubes.

ToF-SIMS Imaging of Nanotubes. The presence of a functionally active outer layer on the nanotubes is of prime importance for most of the aimed applications. In particular, for biomaterial applications of protein-based nanotubes, where one intends to modulate the cell response through both protein function and nanotopography, it is absolutely critical to have the proteins in the outermost layer of the nanotubes. ToF-SIMS analysis, including ToF-SIMS imaging, was therefore used to determine the nature of the outermost layer (COL or PAH) in the different types of nanotubes. Both positive and negative ion images of the outermost layer of nanotubes (COL/PSS) 3(Flu-PAH/PSS)3 and (Flu-PAH/PSS)3(COL/PSS)3) obtained 4108

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Figure 6. ToF-SIMS images (field of view 100 μm2) of ITO glass coverslips used as both working and counter electrodes for nanotubes collection by EPD. (A) (COL/PSS)3(Flu-PAH/PSS)3 nanotubes on the working electrode; (B) (Flu-PAH/PSS)3(COL/PSS)3 nanotubes on the working electrode; (C) counter electrode. Sections (a) and (b) present images from positive and negative ions, respectively. The scale to the right of the image is from dark (low counts) to bright (high counts), with MC = maximum intensity counts. *Combined image of positive ion peaks of amino acids [C2H6N+, C4H8N+,C4H8NO+]. **Combined image of negative ion peaks of polycarbonate (PC)[C 8H5O−, C9H9O−,C14H11O2−].

from high-resolution ToF-SIMS spectra are shown in Figure 6. Both working (Figure 6A,B) and counter electrodes (Figure 6C) were analyzed. More details are given in Figures S4, S5, and S6 (see Supporting Information), where Figure S4 gives summary of positive and negative ion fragments used to generate ToFSIMS images. On the working electrode, the positive ion image of indium (In+) clearly shows the presence of nanotubes on the ITO-glass surface for both types of investigated nanotubes (Figure 6a, rows A and B). Nanotubes that were deposited on the working electrode show specific signature peaks which correspond to the outer layer of nanotubes. Figure 6a presents images of positive ions fragment CH4N+ (30.03 m/z) which is used to monitor PAH, and amino acid fragments (44.05, 70.06, and 86.06 m/z) were used to identify the presence of collagen. Total amino acid images were obtained after addition of amino acids fragments corresponding to alanine (44.05 m/z), proline (70.06 m/z), and leucine (86.06 m/z) where proline is a one of the major component of collagen (see Supporting Information Figure S4 for ToF-SIMS image of proline at 70.06 m/z).36,37 For (COL/PSS)3(Flu-PAH/PSS)3 nanotubes, only a weak signal at 30.03 m/z is visible while a much stronger total amino acids signal located along the nanotubes is observed, proving the presence of collagen in the outermost layer of this type of nanotubes. In the negative ion images (Figure 6b), the presence of sulfonate ion (SO3−) characteristic of fragments coming from PSS at 79.95 m/z indicates, as expected, that this polyelectrolyte is also present in the outermost layer of these nano-

tubes. But one more important observation to point out is the absence of any polycarbonate signals (shown here as total PC signal obtained by the sum of fragments at C8H5O− (117.03 m/z), C9H9O− (133.06 m/z), and C14H11O2− (211.07 m/z) on the working electrode coated with nanotubes. This confirms the efficiency of the EPD process to get nanotubes completely free of template material. For (Flu-PAH/PSS)3(COL/PSS)3 nanotubes, the presence of a strong and localized signal at 30.03 m/z along with the absence of signal from amino acids indicate that these nanotubes retain PAH in their outermost layer after EPD. Corresponding negative ion images of these nanotubes also show a strong and localized signal of sulfonate ions (SO 3− at 79.95 m/z), confirming the presence of PSS in the outermost layer of these nanotubes. Again, the absence of any localization of polycarbonate signal on this working electrode coated with nanotubes confirms that the outerlayer of the nanotubes is free from template material after deposition on ITO glass by EPD. On the counter electrode (Figure 6, row C), no signal was observed from the In+ positive ion image, indicating that the ITO surface of this electrode is covered after EPD. The absence of both signals corresponding to PAH (30.03 m/z) and collagen (total amino acids) indicates that no nanotubes migrated and deposited on the counter electrode. This was also proved by optical and fluorescence microscopy, where no nanotubes were oberved on the counter electrode (data not shown here). Similarly, the absence of sulfonate signal from PSS at 79.95 m/z in the negative ions images further confirms the absence of 4109

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nanotubes on the counter electrode. However, a significant signal corresponding to negative ions characteristic of polycarbonate was recorded on the counter electrode. This observation, combined with the absence of In+ ion signal, suggests that the counter electrode is progressively covered by some polycarbonate during the EPD process. Preliminary Study of the Interaction of Cells with Surfaces Modified by Collagen-Based Nanotubes. Collagen nanotube-based biointerfaces with tunable degree of NTs density prepared by EPD provide new biomaterial interfaces of great interest for both fundamental studies and potential applications. From a fundamental point of view, we are interested in studying the behavior of cells on this type of nanostructured protein-based surfaces. Use of ITO-coated glass for cell culture is not new, it was successfully used before for culture of various cell types including mouse embryonic fibroblasts and 3T3 fibroblasts.38,39 Cytotoxicity studies of ITO glass on 3T3 fibroblasts showed no apparent toxic effect.38 Indeed, 3T3 fibroblasts were shown to behave on ITO glass similarly as on uncoated glass or 316 stainless steel.38 Because of the transparent and conducting nature of ITO glass, cell analysis is possible without further sample preparation using techniques such as optical, fluoresence, and scanning electron microscopies.39 A first illustration of the interaction of cells (MC3T3 preosteoblast cells) with collagen nanotubes deposited on ITO glass is presented in Figure 7. In this figure, one can observe that cells are well

prepared. Interestingly, characterization of the extreme surface of the nanotubes after EPD by ToF-SIMS shows that nanotubes are template-free and that nanotubes with collagen as the outermost layer can be obtained. This is further confirmed by the ability of MC3T3 cells to interact directly with the nanotubes through filopodia. The availability of the outer protein layer is of special interest for biointeractions, which are crucial for applications in the field of biomaterials science. While we demonstrate in this contribution the usefulness of the EPD process to collect collagen-based nanotubes on ITOglass, the technique can be easily adapted to the deposition of NTs on other type of conducting surfaces, such as on metallic implants. Moreover, due to the high versatility of the LbL process used for the preparation of the nanotubes, a huge variety of nanotubes containing different polyelectrolytes, biomacromolecules, or even nanoparticles can be prepared and further collected on conducting substrates to modify their functionality and their topography at the nanometer scale.

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ASSOCIATED CONTENT

S Supporting Information *

Current density profiles measured during the EPD collection of protein nanotubes as well as additional results on the characterization of COL-based multilayer films and nanotubes by QCM-D, SEM, and TOF-SIMS. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION Corresponding Author *Tel: +32-10-472702. Fax: +32-10-451593. E-mail: sophie. [email protected].

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ACKNOWLEDGMENTS The authors thank Etienne Ferain and it4ip company for supplying polycarbonate membranes, Dr. Bernard Knoops and members of his team for providing access to cell culture lab, and Emilienne Zuyderhoff and Simon Degand for collaboration related to cell culture. D.M.K. thanks Prof Paul Rouxhet for valuable scientific discussions and Lucas Leprince for collaboration related to electrochemistry. The work was supported by the Belgian Science Policy through the Interuniversity Attraction Pole Programs (P6/27). The financial support of the Belgian National Foundation for Scientific Research is acknowledged. S.D.C. thanks the F.R.S.-FNRS for her Senior Research Associate position.

Figure 7. Fluorescence microscopy image of MC3T3 preosteoblast cells on ITO glass on which (COL/PSS)3(Flu-PAH/PSS)3 nanotubes were deposited using EPD. Actin stress fibres of cells are labeled with phalloidin dye in red, nuclei are stained with DAPI in blue, and nanotubes appear in green owing to the use of Flu-PAH.

spread and extend their filopodia to interact with single nanotubes. These preliminary results already show that the new elaborated biointerfaces are not cytotoxic and that cells interact directly with the nanotubes, which can be attributed to recognition of the collagen molecules present in their outer layer. The impact of protein-based nanotubes deposited on surfaces on cells morphology, viability, and proliferation is currently investigated in more details and will be reported later.

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REFERENCES

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CONCLUSIONS Protein nanotubes with controlled dimensions, synthesized by a template-assisted LbL assembly, were successfully collected on ITO glass surfaces using a fast, cheap, and scalable process: electrophoretic deposition (EPD). By varying some physical parameters, such as applied voltage, nanotubes concentration in the suspension, and time of deposition, a large variety of surfaces coated with tunable degree of protein nanotubes are easily 4110

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