Colloids and Surfaces B: Biointerfaces 71 (2009) 293–299
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Biofouling of dextran-derivative layers investigated by quartz crystal microbalance Justin Dubois a,b , Charles Gaudreault a,b , Patrick Vermette a,b,∗ a Laboratoire de Bioingénierie et de Biophysique de l’Université de Sherbrooke, Department of Chemical and Biotechnological Engineering, Université de Sherbrooke, 2500, blvd de l’Université, Sherbrooke, Québec, Canada J1K 2R1 b Research Centre on Aging, Institut universitaire de gériatrie de Sherbrooke, 1036, rue Belvédère Sud, Sherbrooke, Québec, Canada J1H 4C4
a r t i c l e
i n f o
Article history: Received 21 November 2008 Received in revised form 5 March 2009 Accepted 9 March 2009 Available online 21 March 2009 Keywords: Biofouling Carboxymethyl dextran (CMD) Plasma polymer Amine derivatization XPS Quartz crystal microbalance (QCM)
a b s t r a c t This study reports the fouling of carboxymethyl dextran (CMD) layers in cell culture medium, ﬁbronectin, and serum solutions. CMD layers were covalently immobilized onto amine groups available either on an n-heptylamine plasma polymer (HApp) layer or onto a polyethylenimine (PEI) coating grafted to an acetaldehyde plasma polymer (AApp) layer. The successful immobilization of the graft layers was demonstrated by X-ray photoelectron spectroscopy (XPS). Primary amines on HApp and AApp + PEI surfaces were quantiﬁed using a colorimetric assay. Quartz crystal microbalance (QCM) was used to investigate in realtime the fouling of the graft layers upon incubation in cell culture medium (RPMI), ﬁbronectin, and foetal bovine serum (FBS) solutions. HApp, AApp and AApp + PEI layers exhibited large fouling in ﬁbronectin and FBS solutions, while fouling in RPMI cell culture medium was not signiﬁcant. Protein repellent properties of CMD layers in FBS and ﬁbronectin have been demonstrated compared to the other tested surfaces. QCM has shown that both CMD layers were fouled to a small extent in RPMI medium. © 2009 Elsevier B.V. All rights reserved.
1. Introduction Speciﬁc cell adhesion on biomaterials is a crucial step in many tissue engineering applications. Surface coatings that yield the required high discrimination between areas where cells should, or should not grow can be an efﬁcient strategy to monitor and modulate cell responses. It appears that an initial protein adsorption event is required to allow cell adhesion to biomaterials surfaces [1–3]. Consequently, testing the fouling resistance of biomaterials surfaces used in tissue engineering applications towards cell culture components and adhesive proteins is essential to design surfaces that would modulate selected speciﬁc cell responses. Low-fouling surfaces can be used to limit non-speciﬁc cell responses towards biomaterials surfaces by limiting non-speciﬁc protein adsorption. Graft layers made of poly(ethylene oxide) (PEO) or poly(ethylene glycol) (PEG) [4–6] and dextran derivatives [7–9] have been one of the most successful low-fouling surfaces. However, the mechanisms behind surfaces that limit non-speciﬁc cell attachment are not fully understood. Nevertheless, the anti-fouling properties of these surfaces seem to be related to a combination of steric repulsion, water structuring effect, and/or electrostatic
∗ Corresponding author at: Department of Chemical and Biotechnological Engineering, Université de Sherbrooke, 2500, boul. de l’Université, Sherbrooke, Québec, Canada J1K 2R1. Tel.: +1 819 821 8000x62826; fax: +1 819 821 7955. E-mail address: [email protected]
(P. Vermette). 0927-7765/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2009.03.002
repulsion towards an initial non-speciﬁc protein adsorption [9–11]. Monchaux and Vermette [7,12] have investigated the physicochemical surface properties of carboxymethyl dextran (CMD) layers with the aim to relate these properties to cell responses. However, the fouling of these CMD layers has not been fully studied. Plasma polymerization can be used to deposit thin reactive ﬁlms on inert surfaces, providing good properties, adherence and longterm stability due to their cross-linked structures [13–15]. In this study, CMD layers were covalently immobilized onto amine groups available either on an n-heptylamine plasma polymer (HApp) layer or onto a polyethylenimine (PEI) coating grafted to an acetaldehyde plasma polymer (AApp) layer. PEI is a cationic polymer that can be used to attach other graft layers via a carbodiimide chemistry [16,17]. PEI coatings have also been investigated to test their fouling . However, some discrepancies can be found in the literature concerning PEI cell afﬁnity [19,20]. Biocompatibility assays revealed that high molecular weight PEI could be cytotoxic . Therefore, it is important to use PEI only as a spacer, as in the present study in which PEI is used to immobilize CMD graft layers. The fouling resistance of CMD graft layers immobilized on PEI surfaces was compared to that of CMD layers attached to HApp surfaces. Consequently, primary amines on HApp and AApp + PEI surfaces were quantiﬁed using a colorimetric assay for comparison purposes. The aim of this study was to compare the fouling resistance of CMD graft layers immobilized on two different layers using quartz crystal microbalance (QCM). QCM was used to investigate in realtime the fouling of the graft layers upon incubation in cell culture
J. Dubois et al. / Colloids and Surfaces B: Biointerfaces 71 (2009) 293–299
medium (RPMI), ﬁbronectin, and foetal bovine serum (FBS) solutions. 2. Materials and methods 2.1. Materials N-heptylamine (#126802-100G, 99.5% purity), acetaldehyde (#402788), N-(3-dimethylaminopropyl)-N -ethylcarbodiimide hydrochloride (EDC, #E1769-10G), N-hydroxysuccinimide (NHS, #130672), sodium cyanoborohydride (#S8628), 2-iminothiolane hydrochloride (ITL, #I6256-500MG), 4-(dimethylamino)pyridine (DMAP, #107700-5G), DL-dithiothreitol (DTT, #D9779-1G), cysteine (#168149-2.5G), Triton X-100 (#T-9284), sodium carbonate (#S-2127), bovine ﬁbronectin (#F4759), foetal bovine serum (FBS, #F1051-100 mL), and RPMI-1640 medium (#R8758) were obtained from Sigma–Aldrich (Oakville, CAN). Hydrogen peroxide, sulphuric acid, BCA reagent A (#PI23228), and BCA reagent B (#PI-23224) were purchased from Fisher Scientiﬁc (Mississauga, CAN). Dextran (#17-0280-01, MW of 70 kDa) was obtained from Amersham Biosciences (Piscataway, USA). Polyethylenimine (PEI, 70 kDa, 9002-98-02) was purchased from Polysciences Inc. (Warrington, USA). Sodium bicarbonate was obtained from EMD Chemicals Inc. (Gibbstown, USA). 2.2. Fabrication of CMD graft layers Clean borosilicate and gold-coated quartz crystal resonators were ﬁrst surface-modiﬁed by plasma polymerization in a custombuilt plasma reactor . Surfaces were cleaned using a UV/ozone treatment (PSD-UV, Novascan, IA, USA) for 40 min and then immersed in ethanol, rinsed in water, and blown dry using a highvelocity stream of 0.2-m ﬁltered air. Borosilicate substrates were used once but quartz crystals were reused for few experiments following an additional cleaning step in etching piranha solution (3:1 ratio H2 SO4 /H2 O2 ) for 2 min after UV/ozone cleaning. Plasma polymerization was carried out in a reactor built inhouse. Operational conditions used to deposit HApp layer were an initial monomer pressure of 0.040 Torr, a power of 80 W, an excitation frequency of 50 kHz, a deposition time of 70 s, and a distance between the electrodes of 10 cm. The resulting thickness of this HApp layer was estimated to approximately 40 nm . For AApp layers, the conditions were 50 kHz, 20 W, 0.055 Torr, a deposition time of 70 s, and a distance between the electrodes of 7 cm. The plasma reactor chamber was always under vacuum before each treatment. The reactor was pre-coated with the respective plasma polymer prior to each experiment. PEI (3 mg/mL in Milli-Q water, pH adjusted to 7.4) was attached directly to AApp layers by reductive amination using sodium cyanoborohydride at a concentration of 3 mg/mL. AApp + PEI surfaces were then washed three times with Milli-Q water, immersed
24 h in a 1 M NaCl solution to remove any non-covalently attached molecules, and ﬁnally washed three times with Milli-Q water. CMD with a MW of 70 kDa and a carboxylation degree of 50% was covalently immobilized either on HApp or AApp + PEI layers using carbodiimide chemistry. The detailed protocol to synthesize CMD is described elsewhere [7,9] and was inspired by the work of Johnsson et al. . CMD graft layers were prepared by ﬁrst dissolving CMD in Milli-Q water to a concentration of 2 mg/mL. EDC (19.2 mg/mL) and NHS (11.5 mg/mL) were added in excess to this CMD solution and allowed to react for a period of 10 min . Samples were then immersed in this solution for a period of 24 h. Subsequent washing steps with 1 M NaCl and Milli-Q water were also done twice. For ﬁbronectin and FBS fouling assays, the quartz sensors were immersed in RPMI-1640 medium for 24 h prior to QCM measurements, while for RPMI assays, quartz sensors were pre-incubated in Milli-Q water. 2.3. Elemental composition of CMD layers by X-ray photoelectron spectroscopy (XPS) XPS analysis of the different layers was performed using an AXIS ULTRADLD spectrometer (Kratos Analytical Ltd., Manchester, GB) equipped with a monochromatic Al K␣ source at a power of 225 W. For XPS analyses, thin borosilicate plates (12 mm × 12 mm) from Assistent (Cover Glasses No. 990) were used. The elemental composition of the analysed surface areas was obtained from survey spectra collected at pass energy of 120 eV. High-resolution C 1s spectra were collected at 20 eV. Atomic concentrations of each element were calculated using CasaXPS (Casa Software Ltd.) by determining the relevant integral peak areas, and applying the sensitivity factors supplied by the instrument manufacturer; a Shirley background was used. To compare the high-resolution C 1s peak positions, the spectra were shifted to ensure that the leading edges of the ﬁtted aliphatic CHx component were coincident. All spectral intensities were normalized to a maximal intensity corresponding to the full height of the ﬁtted aliphatic CHx (285.0 eV) component peak. Survey scans and C 1s high-resolution spectra were collected on each spot (at least three samples per condition). 2.4. Derivatization of primary amines Primary amine densities available on HApp or AApp + PEI layers immobilized on borosilicate plates (24 mm × 24 mm, Assistent, Cover Glasses No. 990) were quantiﬁed. The chemistry used to derivatize primary amines has been adapted from elsewhere [23,24] and is presented in Fig. 1. HApp or AApp + PEI layers were incubated for 1 h under agitation in an activation solution (20 mM DMAP, 20 mM ITL, 3% (v/v) Triton X-100 and 0.1 M sodium bicarbonate) adjusted to pH 8.5. Then, the substrates were dried and washed in water several times. Samples were incubated 1 h in a 1 mM DTT solution with slight agitation. Samples were dried, washed twice
Fig. 1. Chemistry involved in the derivatization of primary amines. ITL reacts with primary amines to form sulphyldryl-containing species. Sulphyldryl reduces Cu2+ to Cu+ . BCA and Cu+ form complex adsorbing at 562 nm.
J. Dubois et al. / Colloids and Surfaces B: Biointerfaces 71 (2009) 293–299
in ethanol, and rinsed three times in a 0.25 M sodium carbonate solution. Samples were incubated in a solution containing 1.96 mL of BCA reagent A, 0.04 mL of BCA reagent B and 0.1 mL of a 0.25 M sodium carbonate solution. Samples were agitated 5 min and then incubated 1 h at 60 ◦ C. Thereafter, surfaces were cooled down to room temperature prior to the absorbance reading at 562 nm. The BCA reagents A and B with sodium carbonate were used as blanks while controls have followed the same protocol as those modiﬁed. Absorbance of the controls was subtracted from that of samples to eliminate interference produced by DTT, ITL and Triton X-100. Standard solutions were made with BCA reagents (with sodium carbonate) and by diluting cysteine (0.00125–0.01 mM). 2.5. Fouling of graft layers measured by QCM QCM measurements were performed using an apparatus from Resonant Probes GmbH (Goslar, Germany). The gold-coated quartz resonators (Maxtek Inc., Cypress, CA, No. 149273-1) were placed into a commercial Kynar holder (Maxtek Inc., Cypress, CA, No. CHT100). Data from the impedance analyser (Agilent, Palo Alto, CA, HP4396A) were recorded using a software from Resonant Probes. Prior to each QCM analysis, the quartz was rinsed, dried, and placed into the holder before RPMI or Milli-Q water was injected to serve as baselines for, respectively, ﬁbronectin/FBS or RPMI fouling assays. Harmonics vibrations were then located. 2 mL of either RPMI, ﬁbronectin (0.05 mg/mL in RPMI), or FBS (10%, v/v in RPMI) solutions were injected into the chamber at a ﬂow rate of 0.5 mL/min. The frequency shifts were recorded by the software to monitor changes in layer mass. FBS and RPMI assays were done at 37 ◦ C while RPMI and ﬁbronectin assays were done at room temperature. Data were obtained in triplicate and only data at 15 MHz have been interpreted. Frequency shifts represent average differences between the ﬁrst baseline (prior to FBS, ﬁbronectin, or RPMI injections) and the last baseline (after rinsing with the same solution as that used to obtain the ﬁrst baseline). 3. Results and discussion 3.1. Surface chemical composition by XPS analyses XPS analysis of freshly deposited HApp layers on borosilicate substrates indicated a polymer rich in hydrocarbon- and nitrogencontaining species (Table 1 and Fig. 2A). N/C and O/C atomic ratios are similar to those reported in the literature . The composition of AApp coatings is in agreement with earlier results . The spectrum shows a polymer rich in hydrocarbon and oxygen species (Table 1 and Fig. 2B). We note the presence of a signal from the element Si (ca. 1%), which indicates that the coating thickness is less than 10 nm. The broad C 1s spectrum (Fig. 2B) provides evidence for the presence of C–O and C O functional groups in the coating. The non-aldehyde oxygen containing species within the ﬁlm may be due to reactions within the plasma during ﬁlm deposition, or additionally the result of post-plasma oxidation as described previously [26–29]. Table 1 Elemental compositions of surfaces tested in this study. The data were derived from XPS survey spectra. Surfaces
HApp HApp + CMD AApp AApp + PEI AApp + PEI + CMD
0.02 0.28 0.16 0.16 0.45
0.08 0.07 0.00 0.09 0.08
90.3 73.6 84.3 77.9 65.2
2.2 20.8 13.8 12.3 29.3
7.5 5.2 0.0 6.7 5.0
NS NS 1.2 1.0 0.2
NS: not signiﬁcant. a Atomic concentration.
Fig. 2. High-resolution XPS C 1s spectra recorded at various fabrication stages of the multi-layer construct: (A) n-heptylamine plasma polymer (HApp), HApp bearing carboxymethyl dextran (CMD); (B) acetaldehyde plasma polymer (AApp), AApp bearing covalently immobilized polyethylenimine (PEI), AApp + PEI with grafted CMD layers.
The XPS C 1s spectrum of PEI grafted onto AApp layers is shown in Fig. 2B. The increase in nitrogen content (but not oxygen) shown in Table 1 suggested that the increase in height of the C 1s shoulder at 286.5 eV is largely attributable to the presence of C–N species within the PEI. These observations are in agreement with earlier results . The ratio of atomic concentrations of nitrogen to carbon (N/C) for this coating (ca. 0.09) was considerably smaller than the 0.5 expected for a pure PEI coating. This demonstrates that the PEI coating is either extremely thin (less than 10 nm) or not complete. The latter appears less likely as XPS measurements were reproducible across the surface, as well as from AFM force analyses and streaming potential data . Moreover, XPS analysis is done in an ultrahigh vacuum (UHV) environment and therefore the layers are fully dehydrated and ﬂattened. The elemental composition obtained on CMD layers revealed a signiﬁcant increase in oxygen content relative to those observed either on freshly deposited HApp or AApp + PEI layers, conﬁrming the successful attachment of the polysaccharide. Fig. 2 shows representative examples of high-resolution C 1s spectra indicating the introduction of oxygen containing carbon species C–O (286.5 eV) and C O (287.9 eV), also conﬁrming the grafting of CMD on both HApp and AApp + PEI surfaces. The presence of an N signal on CMD layers can be explained by either an inefﬁcient washing procedure
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Table 2 Surface density of primary amines derivatized on HApp and AApp + PEI surfaces. Surfaces
Surface density of primary amines (groups/nm2 )
HApp (1 day post-modiﬁcation in water) HApp (8 days post-modiﬁcation in water) AApp + PEI (2 days post-modiﬁcation in water)
10.0 ± 1.0 6.8 ± 0.3 7.0 ± 1.0
allowing some N-containing species (e.g., EDC, NHS) to be trapped within the CMD layers or by thinner CMD coatings (in the dry state) than the XPS analysis depth. From AFM force measurements, CMD layers immobilized on HApp layers using similar conditions have, in a hydrated state, an apparent thickness of approximately 20–30 nm . In addition, when comparing O/C ratios and C 1s spectra of CMD layers immobilized on AApp + PEI to those of CMD layers attached on HApp, it appears that the CMD layers attached to AApp + PEI is thicker and/or denser. Fouling of the CMD graft layers by RPMI components was detected by XPS (data not shown). From the XPS survey and C 1s spectra, fouling of HApp + CMD and AApp + PEI + CMD from RPMI resulted in different surface chemical composition changes, allowing us to hypothesize that these two CMD layers either have different initial molecular structure and/or suffer from different molecular reorganisation following the fouling assays. RPMI is rich in various kinds of species (sugars, vitamins, amino acids, inorganic salts, etc.). 3.2. Derivatization of primary amines Data from derivatization experiments presented in Table 2 reveal that both HApp and AApp + PEI surfaces exposed functional primary amines. In this method, the surfaces are immersed in 2imiothiolane (ITL) to form one sulphydryl group for each primary amine. After DTT immersion, sulphydryl groups reduce Cu2+ to Cu+ and form a complex in the presence of bicinchoninic acid (BCA), which absorbs at a wavelength of 562 nm. The use of DTT after surface activation blocks disulphide bonds and maintains sulphydryl groups for further reduction of Cu2+ (present in BCA solution in the CuSO4 form). DTT can interfere at 562 nm. The absorbance measured is then correlated with a cysteine standard curve to obtain the primary amine concentration. Thereafter, surface amine density is calculated by taking the number of primary amines divided by the borosilicate surface area. See Fig. 1 and Section 2.4 for more details on the protocol.
HApp and AApp + PEI-modiﬁed substrates have 10.0 ± 1.0 amine molecules/nm2 and 7.0 ± 1.0 amine molecules/nm2 , respectively. Previous studies using derivatization have indicated much lower amine density with HApp and with other nitrogen-containing monomers (ca. 0.5–2.0 amines/nm2 ) [30,31], while other studies have reached such density with melamine and even higher density with urea plasmas [32,33]. Surface density of primary amines available on HApp decreased to 6.8 ± 0.3 amine molecules/nm2 when HApp layers were incubated in Milli-Q water for a period of 8 days prior to derivatization. This ﬁnding might be the result of postplasma oxidation, which has been shown by Gengenbach et al. to substantially increase the polarity of the surface by adding more oxygen-containing groups in the ﬁrst 3 weeks of aging [26–28]. Such oxidation could affect the derivatization results. Operational conditions have also been reported to affect the resulting amine surface density of, for example, pulsed plasmas [34,35]. Treatment time , power , and the monomer nature  can affect the amine surface density available on the resulting coating. Also, probes or molecules used to derivatize the amines can penetrate to different depths into the coatings, resulting in a different probed volume. Consequently, it is difﬁcult to draw ﬁrm conclusions concerning the observed differences noted between the different studies. It was initially hypothesized that higher surface density of amine groups could have resulted in higher surface concentration of CMD molecules and subsequently affect the fouling of the resulting CMD layers. In Fig. 3, we illustrate how surface coverage of primary amines was approximated. This correlation has been made by considering the molecular radius of a –NH2 molecule. In the best case scenario, the value of 10 amine/nm2 found on HApp represent a surface coverage of approximately 50%. For AApp + PEI surfaces, this hypothesis could result important differences from the reality while for thin HApp surfaces, it could be closer. The amine density found on AApp + PEI (7.0 ± 1.0 amine molecules/nm2 ) is relatively low, if compared with results from another study (up to 66 amine molecules/nm2 for PEI-covered surfaces) . However, primary amine content of PEI coatings appears to depend on the immobilization protocol, as lower values have been reported by colorimetric assays (ca. 3.5 amines/nm2 ) . Since results from derivatization assays between HApp and AApp + PEI surfaces are in the same range, similar reactivity could be expected with CMD. However, our XPS analyses reveal some differences between CMD graft layers immobilized on HApp and AApp + PEI surfaces. Thus, we can assume that the number of amine groups available for CMD attachment could differ due to steric conformation or primary amine density is not the main parameter
Fig. 3. Schematic picture illustrating how surface densities of primary amines were calculated on HApp and AApp + PEI layers. Primary amine molecules were assumed to be spherical with a molecular radius of 0.123 nm. The surface projection was used to approximate the surface density.
J. Dubois et al. / Colloids and Surfaces B: Biointerfaces 71 (2009) 293–299
Table 3 ANOVA table (p-values) comparing frequency shifts (f/n) measured by QCM for layers exposed either to RPMI culture medium, foetal bovine serum (FBS), or ﬁbronectin (Fn) solutions. The results are based on the third overtone harmonic (15 MHz). p-Values of compared f/n
AApp vs. AApp + PEI AApp vs. AApp + PEI + CMD AApp vs. HApp AApp vs. HApp + CMD AApp + PEI vs. AApp + PEI + CMD AApp + PEI vs. HApp AApp + PEI vs. HApp + CMD AApp + PEI + CMD vs. HApp AApp + PEI + CMD vs. HApp + CMD HApp vs. HApp + CMD
0.0071 0.0012 0.0384 0.0001 0.2249 0.2998 0.0027 0.0416 0.0178 0.0006
0.0019 0.0001 0.0001 0.0002 0.0196 0.0001 0.0899 0.0001 0.3548 0.0001
0.0020 0.0001 0.3895 0.0001 0.0001 0.0006 0.0001 0.0001 0.5743 0.0003
dictating either CMD thickness or atomic concentration differences observed by XPS analyses. It is also possible that the PEI molecules could stretch, to some extent, into the incubating solution, while HApp layers are believed to be more rigid and dense . Also, as mentioned above, the penetration depth of the derivatization reagents that were used in these experiments cannot be determined. Derivatization results should be used to have an idea of the order of magnitude of primary amine content of the layers. 3.3. Fouling of graft layers measured by QCM The main reason to produce substrates covered by thin CMD layers is to delay, lower or perhaps eliminate non-speciﬁc cell–material interactions. A good estimation of this property is to monitor protein adsorption on the polymer layers, as this adsorption will dictate further interactions between cells and the solid support covered by these coatings. In the present study, QCM was used to measure the adsorption of the large spectrum of proteins found in FBS on CMD samples, as it is a good way to screen for the surface fouling resistance encountered in cell culture. Also, the fouling of these CMD layers by ﬁbronectin and RPMI cell culture components was tested. Results from the different biofouling assays are listed in Table 3. Fig. 4 presents the adsorbed mass calculated using the Sauerbrey equation: m = −C
C is the mass sensitivity constant and is equal to 0.056 ng/(cm2 Hz) at 5 MHz (this value is based on the information provided by the supplier, RQCM research quartz crystal microbalance, IPN 603800-J, Inﬁcon Inc., 2007) and n is the overtone order. Rechendoff  estimated that the Sauerbrey equation should not be used when dissipation shift is greater than 10−6 . Thus, mass shifts presented here should be used only to get an order of magnitude and for semi-quantitative comparison. After injecting RPMI, FBS, or ﬁbronectin solutions and following rinsing and subtracting initial baseline values, small positive frequency shifts were observed. These shifts upon injection can be explained by a weak interaction and/or even penetration within the graft layers (at least for CMD layers) of some components dissolved in these solutions (e.g., proteins, amino acids). Also, it is possible that these shifts were inﬂuenced by a change in viscosity or in density (water → RPMI and RPMI → FBS or ﬁbronectin solutions). 3.3.1. Fouling by RPMI From Fig. 4A, it can be seen that the adsorption of RPMI components on the layers was very small, as indicated by the small frequency shifts. The statistically signiﬁcant differences
Fig. 4. Adsorbed mass calculated using the Sauerbrey Eq. (1) from frequency shifts measured with QCM at steady state on the different graft layers exposed to (A) RPMI culture medium and to (B) FBS and ﬁbronectin solutions. Overtone 3 (i.e., 15 MHz) is the harmonic for which the results are reported here. Masses calculated from Sauerbrey should be used to get an idea of the order of magnitude of the adsorbed masses.
between the frequency shifts are listed in Table 3. HApp and AApp layers show negligible adsorption from RPMI considering the detection limit of the instrument (ca. ± 2 Hz). CMD graft layers exposed to RPMI showed the largest frequency shifts among all the tested coatings. Penetration/entrapment of RPMI components (e.g., amino acids, vitamins, salts, sugars) may explain the larger shifts measured with AApp + PEI + CMD and HApp + CMD layers. Also, diffusion of the electrolyte solution inside the CMD polymer layer can result in the formation of a swollen ﬁlm in equilibrium with the new contacting medium . Moreover, electrostatic interactions (physisorption) can be observed between some charged species contained in RPMI and the grafted CMD molecules. Nevertheless, XPS analyses and QCM measurements show that CMD layers suffer from a low level of fouling from RPMI and that this level of fouling was not sufﬁcient to affect these CMD layers resistance towards non-speciﬁc protein adsorption and cell attachment [9,40]. Proteins are much bigger molecules that probably cannot penetrate inside the CMD layers due to steric hindrance. 3.3.2. Fouling by serum The statistically signiﬁcant differences between the frequency shifts obtained from the different layers exposed to FBS are listed in Table 3. Fig. 4B reveals that the adsorption of FBS components on HApp layers was the largest among the tested surfaces. Considering the detection limit of the QCM instrument, it can be assumed that CMD layers immobilized both on HApp and AApp + PEI suffer from very small permanent adsorption following FBS exposure. If there was some fouling, QCM was not sensitive enough to detect it or adsorption would result after longer period of time. These results are in good agreement with the ﬁndings of Monchaux and Vermette  revealing that CMD layers produced in the same way on HApp layers were resistant to cell attachment. Nevertheless, comparing the FBS fouling on CMD graft layers with those measured on other surfaces emphasizes the excellent low-fouling properties of these surfaces. FBS adsorption on biomaterial surfaces is a
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good indicator of the subsequent cell responses to that biomaterial [41,42]. 3.3.3. Fouling by ﬁbronectin Fibronectin is known to bind both integrin protein receptors and extracellular matrix (ECM) components such as collagen, ﬁbrin, and heparan sulphate . Several studies display the effectiveness of ﬁbronectin in cell adhesion [44–48]. Fibronectin must be attached to a substrate or be insoluble to promote cell adhesion [49,50]. Also, to efﬁciently promote cell attachment on biomaterials surfaces through ﬁbronectin, cells must be able to remodel this protein via endogeneous matrix protein production . Fibronectin is not the only protein responsible for cell attachment. Other proteins such as vitronectin, ﬁbrinogen, and laminin, only to name a few, can affect cell attachment. Nevertheless, ﬁbronectin is one of the most studied proteins in cell culture and tissue engineering. The present tests reveal that ﬁbronectin adsorption was the highest on AApp + PEI layers, when compared to the other tested surfaces (Fig. 4B). Nevertheless, frequency shifts are much less than those observed with FBS, probably due to the lower protein concentration (0.05 mg/mL for ﬁbronectin and 30–45 mg/mL for FBS). Table 3 reveals signiﬁcant differences between all surfaces except between AApp and HApp surfaces and between the two tested CMD layers. CMD layers show no signiﬁcant ﬁbronectin adsorption, while the other tested surfaces were all fouled. Based on previous studies, CMD layers inhibited cell adhesion and spreading [8,9]. It can be hypothesized that the resistance of CMD graft layers to cell attachment can be explained by their ability to resist both serum and ﬁbronectin (and probably other cell adhesive molecules) fouling. Half-band-half-width (HBHW) graphs have not been shown in this study because of the absence of any statistically signiﬁcant differences observed between the tested surfaces. It was also difﬁcult to obtain repeatable HBHW results. At this stage, this issue will require further investigation. QCM uses an oscillating potential difference (AC) to induce mechanical oscillations by the converse piezoelectric effect . The adsorption of foreign mass onto QCM crystals results in frequency shifts detectable by an impedance analyser. However, since many experimental conditions (e.g., rheological properties, surface stress and surface roughness, free interfacial energy, slip conditions, etc.) can also induce frequency shifts, interpretation of QCM results can become extremely difﬁcult . Nevertheless, several authors interpret in an audacious way data obtained in complex biological ﬂuids [40,54]. Protein adsorption from complex mixtures on polymeric graft layers should not be quantiﬁed, in the absolute sense, using the Sauerbrey equation, which linearly relates mass changes to frequency shifts for rigid ﬁlms. Other models (e.g., Voigt, Maxwell, and others) have been tested to try to overcome this limitation [55–57]. These models can become complex and need several assumptions. In some studies, results from QCM should be interpreted in a more conservative way, because: • the lateral sensitivity of thickness shear mode resonator is not uniform and vary over the radius [42,53]; • upon adsorption onto a solid surface, protein geometry and conformation could change HBHW shifts; • the structure of hydrated graft layers can contribute to frequency shifts . Despite these notes for caution in the interpretation of QCM data, QCM remains a powerful tool to follow and compare kinetic adsorption in real-time in the ng/cm2 range .
4. Conclusions The aim of this study was to compare the fouling resistance of CMD coatings grafted on two different sublayers. In both cases, CMD layers were covalently bounded on primary amine groups by carbodiimide chemistry. Primary amine groups were available on both HApp layer and on PEI coating, which the latter was grafted to an AApp layer. From XPS results, we showed the successful deposition of all the tested layers. Also, XPS analyses revealed differences between CMD layers immobilized on AApp + PEI and HApp layers in terms of atomic composition. XPS conﬁrmed the presence of RPMI components on CMD graft layers, as also demonstrated by QCM analyses. However, QCM assays conﬁrmed that the frequency shifts on CMD layers exposed to RPMI were very small, while no significant protein adsorption was detected on CMD surfaces from FBS and ﬁbronectin. HApp coatings have exhibited the largest fouling in FBS while the largest fouling from ﬁbronectin was observed on AApp + PEI. Surface densities of primary amines were found to be similar on HApp and AApp + PEI layers, revealing that this factor was not responsible for the differences observed in chemical compositions of the CMD surfaces. Also, surface density of primary amines decreased over incubation time of HApp layers in water. Acknowledgements This work was supported by the Canadian Foundation for Innovation/Ministère de l’Éducation du Québec through an On-going New Opportunities Fund (project # 7918) and by the Université de Sherbrooke. We thank the National Science and Engineering Research Council of Canada (NSERC) for ﬁnancial support through an Undergraduate Student Research Award awarded to Charles Gaudreault. References  N. Nath, J. Hyun, H. Ma, A. Chilkoti, Surf. Sci. 570 (2004) 98.  B. Katz, E. Zamir, A. Bershadsky, Z. Kam, K.M. Yamada, B. Geiger, Mol. Biol. Cell 11 (2000) 1047.  A. Underwood, B.A. Dalton, J.G. Steele, F.A. Bennett, P. Strike, J. Cell Sci. 102 (1992) 843.  F. Brétagnol, M. Lejeune, A. Papadopoulou-Bouraoui, M. Hasiwa, H. Rauscher, G. Ceccone, P. Colpo, F. Rossi, Acta Biomater. 2 (2006) 165.  M. Ulbricht, H. Matuschewski, A. Oechel, H.-G. Hicke, J. Membr. Sci. 115 (1996) 31.  Y. Martin, P. Vermette, Macromolecules 39 (2006) 8083.  E. Monchaux, P. Vermette, J. Biomed. Mater. Res. A 85 (2008) 1052.  S.P. Massia, J. Stark, D.S. Letbetter, Biomaterials 21 (2000) 2253.  K.M. McLean, G. Johnson, R.C. Chatelier, G.J. Beumer, J. Steele, H.J. Griesser, Colloid Surf. B: Biointerf. 18 (2000) 221.  M. Morra, J. Biomater. Sci. 11 (2000) 547.  P. Vermette, L. Meagher, Colloid Surf. B: Biointerf. 28 (2003) 153.  E. Monchaux, P. Vermette, Langmuir 23 (2007) 3290.  Hayakawa, M. Yoshinari, K. Nemoto, Biomaterials 25 (2004) 119.  Y. Martin, D. Boutin, P. Vermette, Thin Solid Films 515 (2007) 6844.  K.S. Siow, L. Britcher, S. Kumar, H.J. Griesser, Plasma Process. Polym. 3 (2006) 392.  J. Wei, D.B. Ravn, L. Gram, P. Kingshott, Colloid Surf. B: Biointerf. 32 (2003) 275.  J. Ji, Q. Tan, J. Shen, Polym. Adv. Technol. 15 (2004) 490.  R. Bahulekar, N.R. Ayyangar, S. Ponrathnam, Enzyme Microb. Technol. 13 (1991) 858.  C.L. Vleggeert-Lankamp, A.P. Pêgo, E.A. Lakke, M. Deenen, E. Marani, R.T. Thomeer, Biomaterials 25 (2004) 2741.  B. Liu, J. Ma, E. Gao, Y. He, F. Cui, Q. Xu, Biosens. Bioelectron. 23 (2008) 1221.  C. Brunot, L. Ponsonnet, C. Lagneau, P. Farg, C. Picart, B. Grosgogeat, Biomaterials 28 (2007) 632.  B. Johnsson, S. Löfås, G. Lindquist, Anal. Biochem. 198 (1991) 268.  M. Ghasemi, M. Minier, M. Tatoulian, F. Areﬁ-Khonsari, Langmuir 23 (2007) 11554.  S.E. Kakabakos, P.E. Tyllianakis, G. Evangelatos, D.S. Ithakissios, Biomaterials 15 (1994) 289.  P. Hartley, S. McArthur, K. McLean, H.J. Griesser, Langmuir 18 (2002) 2483.  T. Gengenbach, Z. Vasic, R. Chatelier, H.J. Griesser, J. Polym. Sci. Part A: Polym. Chem. 32 (1994) 1399.  T. Gengenbach, Z. Vasic, R. Chatelier, H. Griesser, Plasmas Polym. 2 (1996) 91.  T. Gengenbach, R. Chatelier, H.J. Griesser, Surf. Interf. Anal. 24 (1996) 271.
J. Dubois et al. / Colloids and Surfaces B: Biointerfaces 71 (2009) 293–299  P. Vermette, T. Gengenbach, U. Divisekera, P.A. Kambouris, H.J. Griesser, L. Meagher, J. Colloids Interf. Sci. 259 (2003) 13.  H.J. Griesser, R. Chatelier, J. Appl. Polym. Sci.: Appl. Polym. Symp. 46 (1990) 361.  H. Mugurama, EICE Trans. Electon. E91-C 6 (2008) 963.  Ganapathy, X. Wang, F. Denes, M. Sarmadi, J. Photopolym. Sci. Technol. 9 (1996) 181.  F. Basarir, N. Cuong, W.-K. Song, T.-H. Yoon, Macromol. Symp. 249–250 (2007) 61.  G. Calderon, A. Harsch, G.W. Gross, R.B. Timmons, J. Biomed. Mater. Res. 42 (1998) 597.  A. Harsch, J. Calderon, R.B. Timmons, G.W. Gross, J. Neurosci. Methods 98 (2000) 135.  J. Kim, H.K. Shon, D. Jung, D.W. Moon, S.Y. Han, T.G. Lee, Anal. Chem. 77 (2005) 4137.  H.J. Kim, J.H. Moon, J.W. Park, J. Colloids Interf. Sci. 227 (2000) 247.  K. Rechendoff, The inﬂuence of surface roughness on protein adsorption, Ph.D. Thesis, University of Aarhus, Aarhus, 2006.  T. Zhou, K. Marx, M. Warren, H. Schulze, S. Braunhut, Biotechnol. Prog. 16 (2000) 268.  S.P. Massia, J. Stark, J. Biomed. Mater. Res. 56 (2001) 390.  C. Fredriksson, S. Kihlman, M. Rodahl, B. Kasemo, Langmuir 14 (1998) 248.  A.A. Sawyer, K.M. Hennessy, S.L. Bellis, Biomaterials 26 (2005) 1467.  M. Lyon, G. Rushton, J. Biol. Chem. 275 (2000) 4599.
 H. Kuriharaan, T. Nagamune, J. Biosci. Bioeng. 100 (2005) 82.  B. Zhao, Z. Yi, Z. Lu, W. Tian, F. Cui, H. Fen, H. Hu, T. Kawakami, T. Takagi, N. Nagai, J. Hard Tissue Biol. 15 (2006) 65.  S.M. Sagnella, F. Kligman, E.H. Anderson, J.E. King, G. Murugesan, R.E. Marchant, K. Kottke-Marchant, Biomaterials 25 (2004) 1249.  C.C. Larsen, F. Kligman, K. Kottke-Marchant, R.E. Marchant, Biomaterials 27 (2006) 4846.  K. Kottke-Marchant, A.A. Veenstra, R.E. Marchant, J. Biomed. Mater. Res. 30 (1996) 209.  C.M. Allen, W.M. Sharman, C. La Madeleine, J.E. van Lier, J.M. Weber, Photochem. Photobiol. Sci. 1 (2002) 246.  H.L. Hadden, C.A. Henke, Am. J. Respir. Crit. Care Med. 162 (2000) 1553.  M.S. Lord, C. Modin, M. Foss, M. Duch, A. Simmons, F.S. Pedersen, F. Besenbacher, B.K. Milthorpe, Biomaterials 29 (2008) 2581.  J. Wegener, A. Janshoff, C. Steinem, Cell Biochem. Biophys. 34 (2001) 121.  M. Thompson, G.L. Hayward, Proceedings of the 1997 IEEE International, 1997, p. 114.  C. Modin, A.-L. Stranne, M. Foss, M. Duch, J. Justesen, J. Chevallier, L.K. Andersen, A.G. Hemmersam, F.S. Pedersen, F. Besenbacher, Biomaterials 27 (2006) 1346.  B. Du, D. Johannsmann, Langmuir 20 (2004) 2809.  B. Du, I. Goubaidoulline, D. Johannsmann, Langmuir 20 (2004) 10617.  M.V. Voinova, K.B. Johnson, Biosens. Bioelectron. 17 (2002) 835.