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Nanopores in Single- and Double-Layer Plasma Polymers Used for Cell Guidance in Water and Protein Containing Buffer Solutions Mischa Zelzer and Morgan R. Alexander* Laboratory of Biophysics and Surface Analysis, School of Pharmacy, The UniVersity of Nottingham, Nottingham, NG7 2RD, U.K. ReceiVed: September 3, 2009; ReVised Manuscript ReceiVed: October 23, 2009
To investigate the behavior of plasma polymers in biomaterial applications, we studied plasma polymerized hexane (ppHex) and plasma polymerized allylamine (ppAAm) in pure, buffered, and protein containing aqueous solutions. We report for the first time on nanoscale pores formed on ppHex deposits upon exposure to water and surface blistering of ppHex deposited on ppAAm in water. We demonstrate that the nature of the solution influenced the feature formation. These studies are necessary to monitor the changes of the plasma polymer surfaces applied in cell guidance. The surface chemical and topographical data from single- and double-layer plasma polymer films were compared. We further demonstrate that the differences in surface chemistry and topography that develop during aqueous exposure between the single- and double-layer plasma polymer deposits do not affect the adhesion of cells to the surface. These novel nanostructured surfaces may prove useful as membranes or structured films in biotechnological applications. 1. Introduction Thin polymer coatings are widely used in many industrial and scientific areas such as corrosion, adhesion, catalysis, and biomaterials. In many of these applications, the goal is to modify the surface without changing the properties of the bulk material. The material surface is of particular interest in the field of biomaterials, where a good control over the interaction of the material with the biological environment is desirable and polymers are often the material of choice. For polymer-based biomaterials, surface properties such as wettability and the identity and quantity of functional groups are routinely used to explain and control the response of cells and proteins to the host material.1 It has also been shown that the surface topography, especially on the nanoscale, can influence the biological response,2 overriding the surface chemistry.3 Hence, there is a large interest in techniques that are able to modify the surface of materials to produce well-defined surface chemistries and topographies. Plasma polymerization has emerged as a versatile tool to deposit thin organic films with a variety of different surface chemistries as biomaterial coatings. It has found applications as an intermediate step for further modification of the surface by introducing functional groups4 and has been shown to be useful to control a number of biological responses such as proteinadsorption,5,6 theadhesionandproliferationofcells7–9seither over the whole sample or on spatially defined areas and patternssand as screening tools in the form of surface chemical gradients.3,10–12 The application of plasma polymer films in biological environments exposes the material to conditions that are very different from the ambient conditions and the ultrahigh vacuum usually used in instruments to characterize them. Therefore, if these coatings are to be used to control a biological response to the material surface, it is essential to know how the aqueous environment affects the surface over several days. Gengenbach * Corresponding author. E-mail:
[email protected]. Tel.: +44 (0) 1159515119. Fax: +44 (0) 1159515102.
et al. reported on the long-term stability under ambient conditions of various plasma polymers such as hexane,13 heptylamine,14 methyl-methacrylate,15 ethylenediamine,16 and diaminopropane17 deposited from a plasma generated between two parallel plates using radio frequencies between 100 and 225 kHz. XPS was used to monitor the oxygen content on the surface and revealed that oxidation in plasma polymers generally occurs in three stages. An initial rapid uptake in oxygen occurs within the first few hours after deposition, followed by a more moderate increase in the oxygen content over several days. After this, a slow increase in oxygen could still be seen for several months. In these studies, atomic force microscopy (AFM) was used to show that the plasma polymer films retained their smooth surface during the monitored period. In contrast, when Whittle et al. studied the long-term stability in air of plasma polymers deposited from a continuous wave radio frequency (13.56 MHz) plasma with copper coil electrodes using XPS, they found that few changes occurred over time on allylalcohol and acrylic acid plasma polymers.18 For allylamine and 1,7-octadiene plasma polymers, however, a large initial increase in oxygen was observed after deposition. The oxygen uptake became slower but did not stabilize even after one year. In addition, allylamine demonstrated a loss of nitrogen over time. Comparing their results with those of Gegenbach et al., Whittle et al. concluded that the stability of plasma polymers also depends on the deposition conditions. To date, despite the growing interest in plasma polymer films for biological applications, there are only a small number of published studies investigating the effect of an aqueous environment on plasma polymers. Using neutron reflectometry,19,20 surface plasmon resonance,21 and quartz crystal microbalance22 measurements, plasma polymers produced from the monomers allylamine,19,21 heptylamine,19 N-isopropylacrylamide,22 methylmethacrylate,20 di(ethylene glycol) monovinylether,21 and maleic anhydride21 have been shown to swell up to 15% in volume when immersed in water. The extent of swelling was shown to depend on the deposition conditions.21 Recently, Tarasova et al. monitored the oxidation of plasma polymerized
10.1021/jp908516r 2010 American Chemical Society Published on Web 11/25/2009
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allylamine and heptylamine within the first 24 h after immersion in water and found that the oxidation rate is similar to that observed previously on other plasma polymers in air.23 They reported a conversion of amine and imine groups to amides, while at the same time the number of C-O and CdO groups increased. In most studies, no topographical changes were observed with AFM, although the roughness of plasma polymerized allylamine (ppAAm) was seen to increase in phosphate buffered saline (PBS).24 Vasilev et al. investigated the surface topography of an n-heptylamine plasma polymer after immersion in water and found that pores were formed on the surface with the pore size depending on the deposition parameters.25 The use of layered plasma polymer films formed by sequential deposition of two different plasma polymers has been shown to be a readily accessible and reproducible method to prepare gradients that allow control over the attachment of 3T3 fibroblasts.3,10 Notably, there have been no studies of layered plasma polymers in biological media, despite their great promise.3,8,10 In this work, we investigate the chemical and topographical changes on plasma polymers in aqueous environments with in situ AFM and post exposure WCA and XPS measurements. Single-layer plasma polymerized allylamine (ppAAm) and hexane (ppHex) as well as a double-layer plasma polymer consisting of ppHex deposited on top of ppAAm (ppAAm/ ppHex) are considered. The formation of nanosized pores on ppHex and blisters on ppHex/ppAAm, that were observed here for the first time, will be discussed. The influence of the different surface topographies and chemistries on the 3T3 cell attachment to these surfaces (exposure of up to 4 days) will also be explored. 2. Experimental Methods 2.1. Plasma Polymerization. Deposition of plasma polymers was carried out in a custom built T-shaped borosilicate glass reactor with stainless steel end plates sealed with Viton O-rings. Allylamine (99%, Sigma-Aldrich) and hexane (HPLC grade, FischerScientific)weredegassedbyatleastonefreeze-pump-thaw cycle prior to use. The monomer pressure was measured with a Pirani gauge and controlled with needle valves. The radio frequency (RF) plasma (13.56 MHz) was struck with an external power supply at two capacitively coupled copper band electrodes. The plasma polymers were deposited on glass substrates that were previously cleaned with ultrasound in water, rinsed with acetone, and exposed to an oxygen plasma (20 W, 300 mTorr) for 5 min. Both hexane and allylamine plasma polymers were deposited at a power of 20 W and an operating pressure of 300 mTorr. The thickness of the deposit was monitored via a quartz crystal sensor in the plasma reactor. Both plasma polymers were deposited to a film thickness of 50 nm on the sensor. After deposition, the sample remained in the chamber, maintaining the monomer flow for another 3 min. For the layered plasma polymer, ppAAm was deposited first under the above conditions. The second layer of ppHex was deposited the next day in the cleaned reactor, using the same deposition conditions as above without any previous sample treatment. 2.2. Exposure to PBS. The samples were left to settle for one day after preparation before further treatment. They were placed in untreated polystyrene six-well plates, covered with 3 mL of phosphate-buffered saline (PBS) and stored in an incubator at 37 °C and 5% CO2. Samples were exposed for 1 h, 3 h, 6 h, 12 h, 1 d, 2 d, 3 d, and 4 d, after which they were removed and rinsed in ultrapure water (18 mΩ) and blow dried under nitrogen gas.
Zelzer and Alexander 2.3. Atomic Force Microscopy. Images of the surface topography were obtained on a Dimension 3000 (Digital Instruments, Veeco) atomic force microscope (AFM) controlled with NanoScope (V5.30, 2005). The samples were imaged in tapping mode, and the acquired micrographs had a dimension of 10 × 10 µm. Unless otherwise stated, all images in liquid were obtained within the first 2 h of exposure. Micrographs were taken in ultrapure water, phosphate buffered saline (PBS), a solution of 4 mg/mL bovine serum albumin in PBS (BSA), and cell culture media (DMEM, including supplements as specified for the cell culture experiments). The images were processed with The Scanning Probe Image Processor (SPIP, Version 3.3.6.5, 2005, Image Metrology). The route mean square (rms) surface roughness and the area coverage were calculated after linewise plane corrections. The data presented are an average of at least three measurements. All errors shown represent standard deviations. 2.4. XPS Analysis. XPS data were obtained on a Kratos Axis Ultra instrument with a monochromated Al KR X-ray source (1486.6 eV). It was operated at 15 mA emission current and 10 kV anode potential. The pass energy was 80 and 20 eV for the acquisition of wide and high resolution spectra, respectively. The takeoff angle of the photoelectron analyzer was 90°. Empirically derived sensitivity factors provided by the manufacturer were used to calculate the elemental composition (in atomic percent, at %). The data presented are an average of three measurements. All errors shown represent standard deviations. 2.5. Water Contact Angle Analysis. Static water contact angles (WCA) were measured on a Cam200 Optical Contact Angle Meter (KSV Instruments Ltd.) using ultrapure water. For each drop, 20 images were taken in 1 s intervals, and the average WCA from the left and right side of the drop was calculated using a Young-Laplace fit. To eliminate distortion of the data due to the nonequilibrium state of the drop immediately after deposition, the first value was discarded and the initial WCA at the time point t ) 0 was obtained by extrapolation using a linear regression. The data presented are an average of 22 measurements obtained from 6 samples. All errors shown represent standard deviations. 2.6. Cell Culture. The cell culture conditions and passaging procedures of the NIH 3T3 fibroblasts (obtained from ATCC) were described in detail elsewhere.10 On the plasma polymer samples, 50 000 cells (103 cells per mm2) were seeded in 3 mL of Dulbeccos modified Eagle’s media (DMEM) supplemented with 10% (v/v) fetal calf serum (FCS), 2 mM Lglutamine, 100 units/mL of penicillin, 0.1 mg/mL of streptomycin, and 0.25 mg/mL of amphotericin B. The cells were incubated at 37 °C and 5% CO2 for 3 days. After the first day, the media was removed; the samples were washed with 3 mL of PBS for 5 min; and 3 mL of fresh media was added after removal of the PBS. Microscope images were taken with a Leica DM IRB. For analysis, the images were divided into five square areas of 0.6 × 0.6 mm in which the cell number was determined. Cells were classified as either spread and well adhered or round and not well adhered to the surface. Only spread cell were counted. The data were converted to represent the number of cell per mm2 and are an average over 35 measurements. 3. Results 3.1. Atomic Force Microscopy. To study how various aqueous solutions affect the surface topography of single- and double-layered plasma polymers, in situ AFM imaging was used (Figure 1). The ppHex surface showed the formation of small nanosized pores when immersed in pure water or PBS but
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Figure 1. AFM micrographs (10 × 10 µm) of ppHex, ppAAm, and ppAAm/ppHex obtained in air, deionized water, phosphate-buffered saline (PBS), a solution of bovine serum albumin in PBS (BSA), and cell culture media (Dulbeccos Modified Eagles Media, DMEM, supplemented with 10% fetal calf serum). The insets are 4× magnifications of the features observed in the images.
remained smooth when exposed to protein-containing solutions (BSA and DMEM). The pores had a larger diameter in water (220 ( 34 nm) than in PBS (54 ( 17 nm), and the coverage changed slightly from 9% in water to 7% in PBS. A control measurement on oxygen etched glass imaged in PBS (Supporting Information, Figure 1) did not display any comparable features. Significant differences in the surface topography were observed between ppHex and ppHex deposited on top of ppAAm (ppHex/ppAAm). On ppHex/ppAAm, the smooth coating in air changed to a blistered surface in all aqueous environments, displaying islands of several tens of nanometers in height with diameters between 0.4 and 0.8 µm. A small fracture, which is most distinct in the phase contrast images, was observed in the center of the blisters. In comparison, the images taken on ppAAm showed little differences for all five conditions. The sample was flat and featureless when imaged in air, BSA, or DMEM. In water and PBS, some small circular features with irregular edges were observed. The rms roughness for ppAAm and ppHex determined from the AFM micrographs is used to quantify the formation of the features over the field of view and is plotted in Figure 2. The greatest change in roughness was observed in water where it increased from about 0.5 nm in air to 3.0 and 4.6 nm for ppAAm and ppHex, respectively. While the rms roughness in PBS was
Figure 2. Route mean square (rms) roughness of ppAAm (9) and ppHex (0) in air and different aqueous environments obtained from AFM micrographs (10 × 10 µm).
lower than in water, it was still significantly higher than in air. In BSA and DMEM, no significant change in the rms roughness was observed. For ppHex/ppAAm, the area coverage by the blisters, the z-mean, and the z-range of the features are shown in Figure 3. The area coverage is similar in all four aqueous environments (15-17%). The mean difference between the height of the features (z-mean) and the range between the highest and the lowest point (z-range) is larger in water than in the other three liquids.
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Figure 3. Area coverage of features (0), z-mean (9), and z-range (gray filled square) on ppAAm/ppHex in various environments obtained from AFM micrographs (10 × 10 µm).
Figure 4. Evolution of the area coverage (a) and the z-mean (b) of ppAAm/ppHex in PBS over time. The value at t ) 0 represents the measurement of the sample before exposure to PBS. Error bars represent standard deviations for each image.
To determine if long-term changes in the surface topography occur over time, the three sample formats were imaged in PBS over a period of 12 h. While the changes in topography on ppAAm and ppHex occurred within the time needed to acquire the first measurement (about 15 min), no additional changes were observed within the following 12 h. In contrast, the area coverage of the features on ppHex/ppAAm increased over the first 6 h from 20% to 30% (Figure 4). Over the following 6 h, the coverage did not change significantly. A similar trend was observed for the z-mean. To investigate if the blisters on ppHex/ppAAm are still present after exposure to the aqueous environment, the sample was exposed to PBS for different times and imaged in air after rinsing with ultrapure water (Figure 5). The presence of circular features including the crack in the center was confirmed for dry ppHex/ppAAm up to an exposure time of 6 h, although their height was significantly smaller than in PBS, reducing the z-range from 34 nm to less than 10 nm. The 12 h sample was very different from the other exposure times and did not have any circular features but displayed large flat patches instead.
Zelzer and Alexander 3.2. X-ray Photoelectron Spectroscopy. XPS analysis allowed the quantification of chemical changes on the surface over time. Since biological applications usually require the surface to be exposed to the aqueous environment for several days, we monitored the surface chemistry for a period of 4 days. On all samples, carbon and oxygen but no significant amount of silicon (
