Influence of Nanoroughness and Detailed Surface Morphology on

4406

J. Phys. Chem. C 2009, 113, 4406–4412

Influence of Nanoroughness and Detailed Surface Morphology on Structural Properties and Water-Coupling Capabilities of Surface-Bound Fibrinogen Films A. Dolatshahi-Pirouz,† S. Skeldal,‡ M. B. Hovgaard,† T. Jensen,† M. Foss,*,† J. Chevallier,† and F. Besenbacher† Interdisciplinary Nanoscience Center (iNANO), Aarhus UniVersity, DK-8000 Aarhus C, Denmark, and Department of Molecular Biology, Aarhus UniVersity, DK-8000 Aarhus C, Denmark ReceiVed: September 24, 2008; ReVised Manuscript ReceiVed: December 22, 2008

Adsorption of fibrinogen onto nanorough platinum surfaces was investigated using radiolabeling and quartz crystal microbalance with dissipation (QCM-D). In total, four surface topographies were studied with a rootmean-square (rms) roughness value ranging from 1.49 to 9.01 nm, including one surface with sharp whiskerlike surface protrusions and three surfaces with more smoothly shaped surface features. From the radiolabeling results, it is concluded that the fibrinogen adsorption process is influenced by surface roughness, and by combining radiolabeling with QCM-D, we found that the water content of the fibrinogen layers formed on respective surfaces depends significantly on the detailed surface morphology in question. Furthermore, from plots of the dissipation versus frequency shift obtained with QCM-D, the presence of several distinct adsorption phases on nanorough surfaces with more smoothly shaped surface features was observed. This type of multiphased adsorption behavior was not observed on flat reference surfaces or the surface with whiskerlike surface nanofeatures. The results demonstrate that adsorption of fibrinogen onto nanorough topographies is affected by mechanisms which go beyond simple scaling with rms roughness, and that the specific type of surface morphology associated with surface roughness can have a unique impact on the protein adsorption process as well as on the hydration level in the protein films. Introduction In the past decades, a great effort has been made to understand the detailed molecular and cellular mechanisms controlling the biocompatibility of artificial implants.1 Although the biocompatibility of artificial implants has been considerably improved, problems after surgery still arise that influence the long-term function of the implant. Some of the major persistent issues after implantation are the formation of fibrous tissue around the implant and acute inflammation. Both issues are heavily influenced by blood clot formation on the surface.1 In particular, the blood protein, fibrinogen, which has a mass of 340 kDa and an elongated shape with dimensions of 45 nm × 9 nm × 6 nm, is very important in directing the blood coagulation processes on the interface between an implant and surrounding tissue. Fibrinogen has a specific cleavage site that is recognized and cleaved by thrombin, and upon thrombin-catalyzed cleavage of fibrinogen, the blood clot formation is initiated by formation of fibrin monomers, which polymerize into a large scaffold matrix on the surface that eventually leads to clotting.2 In addition to mediating blood clot formation on the surface, fibrinogen is also known to trigger an inflammatory response that can lead to rejection of an implant.3 Thus, by controlling the surface mass uptake and changing the biological activity of fibrinogen on implant surfaces, we may be able to influence the foreign body response as well. Often protein adsorption is controlled by changing the chemical properties of the biomaterial surface. In the past, numerous studies have been carried out to study the influence of surface chemistry on the amount and conformational state * To whom correspondence should be addressed. E-mail: [email protected]. † Interdisciplinary Nanoscience Center (iNANO). ‡ Department of Molecular Biology.

of fibrinogen on a surface.4-14 For instance, in ref 4 the interaction of fibrinogen with tantalum oxide, titanium oxide, and gold surfaces was studied using the QCM-D technique, and it was observed that the fibrinogen surface mass uptake and the structural arrangement of individual fibrinogen molecules on the surface were significantly influenced by substrate chemistry. The same conclusion was drawn in ref 5, where adsorption of fibrinogen onto CH3- and OH-terminated surfaces was examined using the QCM-D technique combined with grazing angle infrared spectroscopy. The effect of nanoroughness on fibrinogen adsorption has also been investigated as an alternative means for controlling the amount and functionality of fibrinogen adsorbed onto surfaces.15-17 Several studies, theoretical and experimental, have been conducted with more simple polymers adsorbing onto silica beads of different sizes. These results have given input to a more fundamental understanding of the interaction between proteins and nanostructured substrates18,19 because proteins can be considered complex polymers. The studies have proven that nanoparticles anchored to a surface can lead to a curvatureinduced enhancement of the amount of polymers adsorbing on top of the structures when appropriate particle sizes are chosen.19 Such curvature-dependent adsorption characteristics have also been observed for proteins such as fibrinogen adsorbed onto hydrophilic and hydrophobic silica beads in free solution.15 Moreover, after using QCM-D and Monte Carlo simulations, it is concluded in ref 16 that the geometrical orientation of fibrinogen is altered because of its anisotropical shape upon adsorption onto tantalum surfaces with a stochastic surface roughness as compared to a flat reference. On the other hand, in ref 17 it was found that the surface roughness did not influence fibrinogen adsorption after a bicinchoninic acid assay was employed to examine the binding of fibrinogen onto

10.1021/jp808488f CCC: $40.75  2009 American Chemical Society Published on Web 02/24/2009

Surface-Bound Fibrinogen Films titanium films with different surface roughness values. Despite recent studies that examine how surface roughness influences adsorption of fibrinogen, no general consensus has been reached on the influence of different sizes and/or types of nanorough surface morphologies on fibrinogen adsorption. Here we investigate using quartz crystal microbalance with dissipation (QCM-D) and radiolabeling how nanorough platinum surfaces, consisting of either sharp whiskerlike surface protrusions or more smoothly shaped nanorough surface features, influence the adsorption characteristics of fibrinogen. Platinum substrates have been chosen because platinum is a biocompatible material frequently used as the conducting material in implant electrodes.20 Materials and Methods Proteins and Antibodies. Human fibrinogen with a purity of 99% was purchased from Kordia Life Sciences (Netherlands) and mouse IgG monoclonal antifibrinogen from Sigma-Aldrich. To test for unspecific binding, rabbit IgG polyclonal anti-bovine serum albumin (BSA) was used from Sigma-Aldrich (Denmark). The fibrinogen was dissolved in a 10 mM Tris buffer with 1 mM CaCl2 and 100 mM NaCl adjusted with HCl and NaOH to pH 7.79 at 22 °C to a concentration of 140 µg/ml. Monoclonal antibodies were dissolved in the same buffer to a concentration of 40 µg/ml. The final protein and antibody solutions were stored at 4 °C, and no solutions were used after more than 10 days. During that period of time no signal degradation was observed as judged by QCM-D (data not shown). The dimensions of a polyclonal IgG antibody are 5.9 nm × 13.1 nm × 14.3 nm with a molecular mass of 150 kDa,21 while fibrinogen has an elongated shape with the dimensions of 45 nm × 9 nm × 6 nm and a molecular mass of 340 kDa.22 Preparation and Characterization of Surfaces. All platinum substrates (Dansk Ædelmetal A/S, DK with a purity of 99.9%) used in the experiments were grown with e-gun evaporation performed at room temperature in a vacuum of about 10-8 mbar, with a distance between the evaporation source and the substrate of 25 cm. Depositions were performed either on gold-coated, AT-cut quartz crystals (Q-Sense AB, Gothenburg, Sweden, model QSX 301) or on silicon (100) wafers (Menzel-Glaser, Germany) precoated with gold by e-gun evaporation. Roughness of the grown platinum film increases when the deposition angle decreases or the surface mass density of the deposited material on the respective samples (representing the total deposited mass per area) increases, which is consistent with previous findings for tantalum and platinum films as described in refs 23 and 24. The morphology and in particular the height variations across the platinum surfaces were determined by atomic force microscopy (AFM) using a commercial Nanoscope IIIA Multimode SPM (Veeco instruments, Santa Barbara, CA). AFM images were acquired in the tapping mode at scan frequencies of 1-2 Hz under ambient conditions using a silicon cantilever (NSG01, NT-MDT, Russia) with a typical resonance frequency around 150 kHz, a spring constant of 5.5 N/m, and a tip radius below 10 nm. The images were quadratic with linear dimensions of 1 µm, 5.5 µm, and 10 µm, and a linear resolution of 512 pixels. To obtain higher lateral resolution images without image artifacts related to the finite radius of the cantilever tip used for the AFM imaging, scanning electron microscopy (SEM) images were obtained on the thin platinum film as well using a Nova NanoSEM 600 (FEI Company).

J. Phys. Chem. C, Vol. 113, No. 11, 2009 4407 Characterization Methods Employed in Surface Morphology Analysis. A standard measure of the surface roughness is the root-mean-square (rms) roughness value, which expresses the variation of the height, h(r,t), over a two-dimensional (2D) substrate with linear size, L, given by

rms(L,t) )




1 L2

∑ [h(r,t) - h¯(t)]2

(1)

where r is the position vector, and the mean height hj(t) is given by

1 h¯(t) ) 2 L

∑ h(r,t)

(2)

In most cases, the rms roughness value depends on the length scale of the surface, and the surface appears rougher as the length scale increases until the length scale reaches the crossover length, Lcrossover, after which the rms surface roughness value saturates.23,24 In this study, saturated rms roughness is used as a measure of the overall roughness of the surfaces employed in the present fibrinogen adsorption experiments and is denoted as the rms value. The crossover length, Lcrossover, is approximately related to the lateral correlation length, ξ, along the individual nanorough surface features on a surface as Lcrossover ∼ 4ξ,25 where ξ defines the length scale at which the surface heights are correlated with one another. The definition of the lateral correlation length, ξ, also implies that it can be used as an approximate measure for the lateral size of the individual surface features on the platinum surfaces.25 Becasue the crossover length does not directly relate to the morphological characteristics of a rough surface, we choose to focus on the saturated rms value and the correlation length, ξ, in the roughness characterization of the respective surfaces. To determine the rms value of the respective surfaces, the AFM images were thoroughly analyzed using a home written plug in to image analyzing software (scanning probe image processor, SPIP),26 and the correlation length, ξ, was found by using the roughness analysis tools in SPIP. Moreover, the height of the individual nanorough surface features was also determined by analyzing AFM images with the grain analysis tools in SPIP, where the height is defined as the horizontal distance between the highest point of a surface feature and the lowest point of the valley separating the surface feature from surrounding ones. Radiolabeling Technique. To investigate the amount of fibrinogen adsorbed onto the respective platinum substrates, gold-coated silicon wafers were precut to measure either 4 mm2 × 4 mm2 or 5 mm2 × 5 mm2 and then coated with platinum using the glancing angle deposition (GLAD) technique. The fibrinogen surface mass density on the respective platinum surfaces was quantified by adsorbing 125I-labeled fibrinogen onto the substrates and subsequently measuring the γ-radiation arising from the 125I decay.12 Labeling of fibrinogen with 125I was performed in a similar manner as described in ref 27, where 22 µL of 0.2 M sodium phosphate buffer, pH 8.1, containing 2 µg of fibrinogen was added to 0.4 mCi of Na125I (PerkinElmer, Denmark), followed by the addition of 4 µL of 0.5 mg/mL chloramin-T in 0.2 M sodium phosphate buffer, pH 8.1, and incubated for 3 min at 21 °C. The reaction was stopped with 100 µL of 0.1 M sodium phosphate buffer, pH 6.5. Protein and free 125I were subsequently separated with a Sephadex G-25 PD-10 column (Amersham Pharmacia Biotech), equilibrated, and run in a washing buffer (10 mM Tris, 1 mM CaCl2, 100 mM NaCl, and 0.22 mg/mL fibrinogen), and 15 300 µL fractions

4408 J. Phys. Chem. C, Vol. 113, No. 11, 2009

Dolatshahi-Pirouz et al.

were collected. Only fractions with a yield of g99.5% were employed for the labeled fibronectin adsorption experiments. Under the assumption that all of the free iodine in the solution actually was bound to the surface, these high-yield fractions ensured that the contribution from free iodine would be less than 5% of the total radioactivity (data not shown). Adsorption tests were carried out at 22 °C by placing a 20 µL drop on substrates situated in a 24-well tissue plate with a thin sponge saturated with a Tris buffer placed at the periphery of the tissue plates to avoid evaporation. After fibrinogen adsorption, the samples were rinsed with a Tris buffer and γ-counted. In total, 3-10 measurements were performed for each substrate type to ensure good statistics. The counts were averaged, and the surface mass density, Γradiolabeling, was determined by12

Γradiolabeling )

counts (cpm) × fibrinogen concentration (3) Asolution(cpm ⁄ mL) × SA

where the counts measure the radioactivity of the surfaceimmobilized samples in counts per minute (cpm), Asolution is the specific activity of the labeled fibrinogen solution, and SA is the sample surface area. Quartz Crystal Microbalance with Dissipation (QCM-D) Technique. The quartz crystal microbalance with dissipation (QCM-D) technique relies on a piezoelectric quartz crystal sensor, which deforms mechanically when exposed to an electrical field. By application of a radio frequency (rf) alternating current (ac) voltage across the crystal, a shear oscillation is induced at the fundamental resonance frequency (first), and at the third, fifth, seventh, etc., overtones; for the crystals used here the fundamental resonance frequency is ∼5 MHz.28 In most cases, an increase in the adsorbed mass on the sensor surface will induce a decrease in resonance frequency and vice versa.29 In the case of a thin nondissipative layer with a no-slip condition, the frequency shift, ∆f, and the surface mass density (ng/cm2), ΓQCM-D, are proportional, according to the simple Sauerbrey equation30

C × ∆fn ΓQCM-D ) n

Elost 2π × Estored

amount of water bound to the protein film (ΓQCM-D/Γdry mass) and the hydrated mass density, Flayer, which is estimated by

1 Flayer

)

(

Fsolvent

(5)

(6)

where Elost is the energy dissipated during one oscillation cycle, and Estored is the total energy of the system. Equation 5 is generally considered to be valid for low-dissipative systems, which is the case if the overtones, ∆fn, overlap after scaling (∆fn/n) with their respective overtone numbers. Throughout this study, eq 5 for the third overtone (∼15 MHz) is used to convert frequency shifts measured using the QCM-D technique to surface mass density values (ng/cm2). A special challenge with respect to the QCM-D technique is that the observed frequency shift is due to the total effective mass coupled to the surface, including both hydrated proteins and any water trapped in the pores of the protein film,31 termed the “wet mass”. In contrast, the radiolabeling technique only detects the protein mass, or the “dry mass”, adsorbed at the interface. The difference in the protein surface mass uptake detected with the QCM-D and radiolabeling techniques can thus be utilized to determine the

)

Vprotein + Vsolvent 1 Γdry mass ) + Mprotein + Msolvent Fprotein ΓQCM-D 1

Here C denotes the mass sensitivity constant (17.7 ng/cm2 Hz-1 for a 5 MHz crystal), and n (1, 3, 5, etc.) is the overtone number. In addition to the frequency shift, the shift in the dissipation factor, ∆D, was also monitored. The dissipation is defined as

D)

Figure 1. Representative AFM images of nanorough platinum surfaces (A, B, C, and D) used in experiments (79 mm × 81 mm, 300 DPI × 300 DPI). (A) Flat reference deposited at a 35° angle with a surface mass density of 8.2 × 10-5 g/cm2. (B) Substrate dominated by nanofeatures with smooth curvatures obtained by depositing 8.2 × 10-5 g/cm2 at 10°. (C) Substrate dominated by nanofeatures with smooth curvatures obtained by depositing 21.5 × 10-5 g/cm2 at 10°. (D) Columnar-shaped surface features obtained by depositing 4.3 × 10-5 g/cm2 at 5°.

(

)

ΓQCM-D - Γdry mass (7) ΓQCM-D

where Mprotein and Msolvent are the dry and solvent mass present in the protein film occupying the respective volumes Vprotein and Vsolvent in the hydrated layer, respectively, Fprotein ) 1.35 g/cm3, and Fsolvent ) 1.00 g/cm3.32 All QCM-D experiments were carried out under static (no flow) conditions at 22 °C using a Q-Sense D300 (Q-Sense AB, Sweden). Crystals applied in the QCM-D measurements were ozone-cleaned with UV for 25-30 min before each experiment. Results and Discussion Characterization of Platinum Surface Morphologies. Using glancing angle deposition (GLAD), the rms value can be controlled by adjusting the deposition angle and the surface mass density of the deposited material of the samples. Moreover, the detailed characteristics of the surface morphology typically change as a function of the deposition angle.23 Figure 1 depicts representative AFM images of the nanorough platinum surfaces synthesized at different deposition times and angles (A, B, C, and D). For surfaces A, B, and C, more smooth nanorough surface features of varying sizes in the nanoscale regime are observed compared to those of the surface morphology on surface D, which is dominated by more sharp columnar protrusions. From the AFM images, the respective rms values of the different substrates are determined and presented in Table 1,

Surface-Bound Fibrinogen Films

J. Phys. Chem. C, Vol. 113, No. 11, 2009 4409

TABLE 1: Different Morphological Characteristics of Respective Surfacesa surface

rms value (nm)

roughness factor

height (nm)

ξ (nm)

A B C D

1.49 ( 0.02 4.62 ( 0.03 7.53 ( 0.04 9.01 ( 0.04

1.030 ( 0.004 1.16 ( 0.03 1.28 ( 0.05 1.323 ( 0.011

6.3 ( 1.2 16 ( 4 29 ( 12 39 ( 9

16.7 ( 0.7 28 ( 5 34.3 ( 0.7 40 ( 4

a

Roughness factor is defined as R ) (Asurface)/(Aflat

surface).

Figure 2. SEM images of surfaces C and D (80 mm × 35 mm, 300 DPI × 300 DPI).

where it is seen that the surface roughness and surface area increase as the depositing angle and the surface mass density of the deposited material on the respective surfaces increase, in accordance with refs 23 and 24. Surface roughness for a deposition angle of 10° was determined to be 4.62 ( 0.03 nm (surface B), increasing to 7.53 ( 0.04 nm (surface C) as the surface mass density of the deposited material on the samples increased from 8.2 to 21.5 × 10-5 g/cm2. Moreover, by adjusting the deposition angle from 35° to 5° (A f D), we observed an increase in the surface roughness from 1.49 ( 0.02 nm (surface A) to 9.01 ( 0.04 nm (surface D). In addition to surface roughness, we also determined the height of the surface features on the different rough surfaces and found that the height was similar for surfaces C and D, corresponding to 29 ( 12 and 39 ( 9 nm, respectively, while the surface feature height was appreciably lower on surfaces B and A, attaining values of 16 ( 4 and 6.3 ( 1.2 nm, respectively. The correlation length, ξ, which gives a rough estimate of the surface feature size is also listed in Table 1. It is observed that the correlation length, ξ, attains almost the same values of 40 ( 4 and 34.3 ( 0.7 nm on surfaces C and D, respectively, while a smaller correlation length, ξ, is found on surfaces A and B, where ξ attains the values of 16.7 ( 0.7 and 28 ( 5 nm, respectively. Interestingly, the surface feature height, correlation length, and rms value for surfaces D and C are similar, despite their different morphological characteristics. Concerning AFM characterization of surface morphologies, it is important to recognize the presence of image artifacts such as the finite resolution of the AFM technique, which depends on the radius of the cantilever tip apex. AFM image artifacts might also, in a worst case scenario, result in distorted images of the true surface topography. Scanning electron microscopy (SEM) was used as a complementary technique to further investigate the morphology of surfaces C and D (Figure 2A,B). SEM images show the same morphological trends as those found by AFM; however, the sharp columnar surface features seen by AFM now appear more evident as whiskerlike surface features on surface D and with more sharp edges present on surface C. Accordingly, the images shown in panels A and B of Figure 2 confirm that the surface morphology on surfaces C and D is indeed quite different.

Results from the Quartz Crystal Microbalance with Dissipation (QCM-D) Technique. The QCM-D technique28 has proven to be very well-suited for in situ dynamic monitoring of mass and mechanical properties such as the viscoelasticity of absorbed biomolecules.34,35 Previously, the technique has successfully been used for determining protein adsorption onto rough surfaces, eliminating some of the challenges that traditional optical methods such as ellipsometry (ELM) and surface plasmon resonance (SPR) are facing16 because of, for example, diffuse light scattering. With respect to the conversion of measured QCM-D frequency shifts to surface mass density (ng/ cm2) values (ΓQCM-D), the simple Sauerbrey equation (eq 5) was employed (Table 2). The Sauerbrey equation has been used for this purpose in several previous studies16,35,36 and is found to be valid for low-dissipative systems, where individual overtones overlap after scaling with their corresponding overtone numbers. In this study, the maximum variation between the scaled overtones within a single QCM-D experiment was always less than 11%, indicating that the Sauerbrey equation provides a reasonable first-order estimate of adsorbed protein surface mass density. From Table 2, it is found that surface mass uptake increases from surface A to surface C. However, for surface D, which has a slightly larger rms value as compared to that of surface C, a decrease in surface mass density is observed. Different mechanisms may explain the correlation between surface mass uptakes determined by the QCM-D technique and substrate morphology: (i) Individual fibrinogen molecules adsorb in different conformations on surfaces D and C, leading to a change in the protein footprints on the respective surfaces. (ii) Less water is bound in the protein film assembled on surface D in comparison to that on surface C. (iii) A combination of (i) and (ii) could also explain the QCM-D results presented in Table 2 because conformational changes in the protein structure arising from the surface morphology upon surface binding may result in a lower hydration level in the protein film formed on surface D, leading to a decrease in the QCM-D frequency response when going from surface C to D. To clarify which of these mechanisms are able to explain the QCM-D results presented in Table 2, further investigations are needed. It is especially important to distinguish between the amount of proteins adsorbed onto the respective surfaces and the hydration level in the respective protein films. Determination of Protein Dry Mass Using Radiolabeling. In order to investigate the impact of bound water on results obtained from the QCM-D technique, we applied the radiolabeling technique. After measuring the γ-activity arising from the 125I-labeled proteins bound to the different surface types (surfaces A, B, C, and D), we converted the number of γ-counts to surface mass densities using eq 3 (Table 2). From these radiolabeling results, we find that the apparent decrease in the fibrinogen surface mass density when going from surface C to D, as observed by the QCM-D technique, is no longer present. With the radiolabeling technique, an increase ranging from 499 ( 27 to 595 ( 8 ng/cm2 in the protein mass uptake is observed, when moving from surface C to D, which suggests the amount of bound water is different in the protein films formed on surfaces C and D. Moreover, the radiolabeling results for surfaces A, B, C, and D all lie between the theoretical full monolayer coverage of fibrinogen in the side-down orientation (Γsurface mass density ) 140 ng/cm2) and the up-right position (Γsurface mass density ) 1050 ng/ cm2).

4410 J. Phys. Chem. C, Vol. 113, No. 11, 2009

Dolatshahi-Pirouz et al.

TABLE 2: Fibrinogen Adsorption Results on Different Nanorough Platinum Surfaces surface

ΓQCM-D (ng/cm2)

Γradiolabeling (ng/cm2)

water factor

Flayer (g/cm3)

antibodies per protein (QCM-D)

∆D3/∆f3 (10-8 Hz-1)

A B C D

1860 ( 31 1912 ( 43 2301 ( 64 2000 ( 69

377 ( 12 451 ( 4 499 ( 27 595 ( 8

4.93 ( 0.18 4.24 ( 0.10 4.6 ( 0.3 3.36 ( 0.12

1.05 ( 0.03 1.06 ( 0.03 1.06 ( 0.04 1.08 ( 0.05

0.99 ( 0.06 0.89 ( 0.09 0.86 ( 0.12 0.77 ( 0.04

4.90 ( 0.10 5.25 ( 0.09 5.4 ( 0.4 3.8 ( 0.3

The water binding capability of a protein film depends on the conformational arrangements of the individual protein molecules on the surface,37-39 which is why additional information can be obtained on protein adsorption by quantifying the hydration level in the protein films formed on the different substrates. The relative amount of coupled water molecules in the different protein layers was determined from the ratios of the observed surface mass densities (ΓQCM-D/Γradiolabeling) (Table 2). These results show that the water factor is nearly the same for the flat substrate (A) and the substrates dominated by nanofeatures with smooth curvatures (B and C), while it is significantly lower for the substrate with the columnar-shaped surface features (D). The water factor found with QCM-D in the present work is close to the water factor found in other QCM-D studies with fibrinogen. For instance Rechendorff et al.16 found a water factor ranging from 4.1 to 4.8, and Ho¨o¨k et al.38 reported a water factor ranging from 2.6 to 3.0, which are both close to the water factor reported in Table 2 (3.4-4.9). Moreover, the density of the hydrated layer was calculated from eq 7, and a tendency toward a small increase from 1.05 to 1.08 g/cm3 (Table 2) is observed when moving from surface A to D in accordance with a lower water content in the protein layer formed on surface D. The results presented in Table 2 thus demonstrate the importance of combining QCM-D with another technique to determine the true protein surface mass uptake. We find that fibrinogen binds in a different configuration on surface D as compared to that of other substrates with lower water factors than surface D, which is in agreement with scenario (iii) discussed above. The results presented in Table 2 are rather surprising because the rms roughness, correlation length, ξ, and height of individual surface features are the same on surfaces C and D, indicating that other surface characteristics, for example, the shapes of individual surface features, may be responsible for the different fibrinogen adsorption seen on these surfaces, or conversely that the roughness data obtained using AFM are not representative for surface D with its many whiskerlike protrusions. Probing the Protein Layer with Monoclonal AntiFibrinogen. Properties of the resulting fibrinogen layer on the respective surfaces were probed with monoclonal anti-fibrinogen, using the QCM-D technique.4,36,37,40 We can estimate the number of antibodies that bind to each fibrinogen molecule, if we assume the water content remains the same in the protein layer after antibodies bind to the protein film. The number of antibodies per fibrinogen molecule is then given as

∆fantibody mfibrinogen Nantibody ) × Nfibrinogen ∆ffibrinogen mantibody

(8)

where mfibrinogen ) 340 kDa, mantibody ) 150 kDa, and Nantibody and Nfibrinogen are the number of antibodies and fibrinogen molecules, respectively. The number of monoclonal antibodies per fibrinogen molecule on the different substrates is depicted in Table 2, and it is shown that the number of antibodies per fibrinogen molecule drops from 0.99 ( 0.06 to 0.77 ( 0.04 (29%) as we go from surface A to D. Furthermore, there is no

significant difference between surfaces A and C in the number of antibodies per protein molecule, and only a small difference (11%) is observed between surfaces A and B. Tests for unspecific monoclonal anti-fibrinogen binding by adsorption of rabbit IgG polyclonal anti-BSA showed unspecific adsorption up to approximately 2% of the saturated anti-fibrinogen uptake. No clear trend was observed between the unspecific monoclonal anti-fibrinogen binding and platinum surface topography. Because it has been reported that the binding capability of antibodies to surface-bound proteins depends on the structural arrangement of the proteins at the solid-liquid interface,4,35,40 one might argue that the smaller amount of antibodies binding to the protein film on surface D, compared with other substrates, is due to differences in the structural properties of the adsorbed fibrinogen films. QCM-D Dissipation Results. To obtain additional information on the structural properties of the adsorbed fibrinogen layer on the respective platinum substrates, we used QCM-D to measure the ratio between the dissipation and frequency shift, ∆D/∆f (Table 2). This type of analysis has previously been performed to gain a deeper understanding of the structural properties of a given surface-bound protein layer.35,36,38,40-44 From these studies, it has been established that a protein film that binds rigidly and forms a compact protein layer on a surface typically leads to lower ∆D/∆f values.40-42 However, it is important to recognize that the ∆D/∆f ratio depends on the dry protein mass and on the water coupled within the adsorbed protein film. It has been shown that the dissipation shift and the ∆D/∆f ratio increase with increasing water content in the protein film.37-39,44 From the results in Table 2, we notice that the ∆D/∆f ratios are almost the same for platinum surfaces A, B, and C within the displayed uncertainties. However, when proceeding from surface A to D, the ∆D/∆f ratio decreases from 4.90 ( 0.10 to 3.8 ( 0.3. Again, it is quite remarkable that the results obtained for surface D differ significantly from the results obtained for surfaces B and C, while surfaces B and C with similar surface morphologies again follow the same trends. It has previously been shown that if the packing density of ferritin molecules is controlled by changing the salt concentration, a 2D dense protein layer leads to very low ∆D/∆f values.42 Also, in refs 38 and 41, it was shown that less water is trapped in a rigid protein film as compared to that of a more viscoelastic and open protein film.38,41 We can, therefore, tentatively conclude from the results depicted in Table 2 that the lower ∆D/∆f ratios on surface D are in accordance with structural changes in the fibrinogen molecule upon surface binding, which in turn lead to a more rigid and compact protein layer on surface D and a decrease in the water content in the protein film. This scenario is also in accordance with the anti-antigen results obtained from QCM-D (see previous section), because such protein films typically are more inaccessible for antibody-antigen recognition.36,40 Moreover, a lower water content coupled to the protein film on surface D also agrees well with the radiolabeling results.

Surface-Bound Fibrinogen Films

J. Phys. Chem. C, Vol. 113, No. 11, 2009 4411 different adsorption phases as well as higher hydration levels in the protein film. Conclusion By using two complementary techniques, QCM-D and radiolabeling, the adsorption characteristics of fibrinogen onto platinum surfaces with well-characterized surface morphologies have been investigated. From the QCM-D and radiolabeling results, we were able to conclude that fibrinogen adsorption is influenced by surface roughness, and in particular, it is observed that adsorption of fibrinogen was influenced by the characteristic shapes and sizes of the nanorough surface features residing on the different nanorough surface morphologies In a broader perspective, such results are important from a fundamental as well as a more applied point of view because we have shown for the first time that not only the surface roughness but also the specific morphology of nanorough surfaces has to be considered when designing the next generation of superior biomaterial implants and biomedical devices.

Figure 3. ∆D/∆f plot for fibrinogen adsorption onto (A) surfaces A and B and (B) surfaces C and D (71 mm × 120 mm, 300 DPI × 300 DPI).

To gain additional insight into the kinetics of the interplay between fibrinogen and the different substrates during the adsorption process, ∆D versus ∆f is plotted in Figure 3. From Figure 3A, we observe that flat reference surface A displays a simple linear relationship between ∆D and ∆f, whereas surface B, dominated by nanofeatures with smooth curvatures, exhibits in contrast a more complex ∆D versus ∆f behavior. This complex behavior closely resembles previous observations for the adsorption of bovine serum albumin (BSA) onto similar platinum surfaces, 40 with an initial rise in the ∆D versus ∆f slope followed by a steeper rise, indicating the structural conformation of the protein layer is altered during the adsorption process onto softer protein film. In Figure 3B, a similarly complex ∆D versus ∆f curve is also observed for surface C, but variations in the ∆D versus ∆f slope are even more pronounced. In particular, a more apparent third intermediate stiffening phase, in which the ∆D versus ∆f slope gradually decreases, is observed as compared to that of surface B. The ∆D versus ∆f curve observed for surfaces B and C reveals that changes in the rigidity of the protein layer occur as adsorption proceeds, with ∆D versus ∆f changes being more noticeable on surface C, which indicates the surface interaction with fibrinogen leads to a more pronounced effect on surface C as compared to that of surface B. The fact that surface D depicts a relatively simple linear ∆D versus ∆f relation, similar to that observed for surface A, indicates that rearrangements in the later phases of the protein adsorption process are not prominent on surface D or not detectable with QCM-D. The results presented in Table 2 and panels A and B of Figure 3 thus imply that fibrinogen adsorption depends on the specific surface morphology. For example, surface D, with whiskerlike surface protrusions, apparently only results in one prominent adsorption phase, while nanorough surfaces B and C, with less sharp smooth surface features, lead to

Acknowledgment. We gratefully acknowledge financial support from the Danish Research Councils to the Centre for NeuroEngineering (CNE) and Interdisciplinary Nanoscience Center (iNANO) and from the European Commission to the FP6 STREP project, NANOCUES. The authors also thank Folmer Lyckegaard for the production of the thin films used in this work. References and Notes (1) Ratner, B. D.; Hoffman, A. S.; Schoen, F. J.; Lemons, J. E. Biomaterials Science, 2nd ed.; Academic Press: San Diego, 2004. (2) Fuss, C.; Palmaz, J. C.; Sprague, E. A. J. Vasc. InterV. Radiol. 2001, 12, 677. (3) Tang, L.; Eaton, J. W. J. Exp. Med. 1993, 178, 2147. (4) Hemmersam, A. G.; Foss, M.; Chevallier, J.; Besenbacher, F. Colloids Surf., B 2005, 43, 208. (5) Roach, P.; Farrar, D.; Perry, C. C. J. Am. Chem. Soc. 2005, 127, 8168. (6) Cacciafesta, P.; Humphris, A. D. L.; Jandt, K. D.; Miles, M. J. Langmuir 2000, 16, 8167. (7) Yongli, C.; Xiufang, Z.; Yandao, G.; Nanming, Z.; Tingying, Z.; Xinqi, S. J. Colloid Interface Sci. 1999, 214, 38. (8) Horbett, T. A.; Brash, J. L. Proteins Interfaces II 1995, 602, 112. (9) Wertz, C. F.; Santore, M. M. Langmuir 2001, 17, 3006. (10) Wertz, C. F.; Santore, M. M. Langmuir 2002, 18, 706. (11) Lewis, K. B.; Ratner, B. D. Colloids Surf., B 1996, 7, 259. (12) Liu, F.; Zhou, M.; Zhang, F. Appl. Radiat. Isot. 1997, 49, 67. (13) Garguilo, J. M.; Davis, B. A.; Buddie, M.; Ko¨ck, F. A. M.; Nemanich, R. J. Diamond Relat. Mater. 2004, 13, 595. (14) Weber, N.; Pesnell, A.; Bolikal, D.; Zeltinger, J.; Kohn, J. Langmuir 2007, 23, 3298. (15) Roach, P.; Farrar, D.; Perry, C. C. J. Am. Chem. Soc. 2006, 128, 3939. (16) Rechendorff, K.; Hovgaard, M. B.; Foss, M.; Zhdanov, V. P.; Besenbacher, F. Langmuir 2006, 22, 10885. (17) Cai, K.; Bossert, J.; Jandt, K. D. Colloids Surf., B 2006, 49, 136. (18) Singh, N.; Karim, A.; Bates, F. S.; Furusawa, K.; Tirrel, M. Macromolecules 1994, 27, 2586. (19) Ligoure, C.; Leibler, L. Macromolecules 1990, 23, 5044. (20) Rozman, J.; Sovinec, B.; Trlep, M.; Zorko, B. J. Biomed. Eng. 1993, 15, 113. (21) Berg, J. M.; Tymoczko, J. L.; Stryer, L. Biochemistry, 5th ed.; W. H. Freeman and Company: New York, 2002. (22) Brown, J. H.; Volkmann, N.; Jun, G.; Henschen-Edman, A. H.; Cohen, C. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 85. (23) Dolatshahi-Pirouz, A.; Hovgaard, M. B.; Rechendorff, K.; Chevallier, J.; Foss, M.; Besenbacher, F. Phys. ReV. B 2008, 77, 115427. (24) Rechendorff, K.; Hovgaard, M. B.; Chevallier, J.; Foss, M.; Besenbacher, F. Appl. Phys. Lett. 2005, 87, 073105. (25) Pelliccione, M.; Karabacak, T.; Gaire, C.; Wang, G.-C.; Lu, T.-M. Phys. ReV. B 2006, 74, 125420.

4412 J. Phys. Chem. C, Vol. 113, No. 11, 2009 (26) Image Metrology A/S. www.imagemet.com (accessed Month Day, Year). (27) Nykjær, A.; Bengtsson-Olivecrona, G.; Lookene, A.; Moestrup, S. K.; Petersen, C. M.; Weber, W.; Beisigel, U.; Gliemann, J. J. Biol. Chem. 1993, 269, 15048. (28) Rodahl, M.; Ho¨o¨k, F.; Krozer, A.; Brzezinski, P.; Kasemo, B. ReV. Sci. Instrum. 1995, 66, 3924. (29) Voinova, M. V.; Jonson, M.; Kasemo, B. Biosens. Bioelectron. 2002, 17, 835. (30) Sauerbrey, G. Z. Phys. 1959, 155, 206. (31) Larsson, C.; Rodahl, M.; Ho¨o¨k, F. Anal. Chem. 2003, 75, 5080. (32) Fisher, H.; Polikarpov, I.; Craivich, A. F. Protein Sci. 2004, 13, 2825. (33) Dolatshahi-Pirouz, A.; Pennisi, C. P.; Skeldal, S.; Foss, M.; Chevallier, J.; Zachar, V.; Yoshida, K.; Besenbacher, F. Nanotechnology, in press. (34) Hovgaard, M. B.; Dong, M.; Otzen, D. E.; Besenbacher, F. Biophys. J. 2007, 93, 2162. (35) Ho¨o¨k, F.; Kasemo, B.; Nylander, T.; Fant, C.; Sott, K.; Elwing, H. Anal. Chem. 2001, 73, 5796.

Dolatshahi-Pirouz et al. (36) Hovgaard, M. B.; Rechendorff, K.; Chevallier, J.; Foss, M.; Besenbacher, F. J. Phys. Chem. B 2008, 112, 8241. (37) Vo¨ro¨s, J. Biophys. J. 2004, 87, 553. (38) Ho¨o¨k, F.; Vo¨ro¨s, J.; Rodahl, M.; Kurrat, R.; Bo¨ni, P.; Ramsden, J. J.; Textor, M.; Spencer, N. D.; Tengvall, P.; Gold, J.; Kasemo, B. Colloids Surf., B 2002, 24, 155. (39) Reimhult, E.; Larsson, C.; Kasemo, B.; Ho¨o¨k, F. Anal. Chem. 2004, 76, 7211. (40) Dolatshahi-Pirouz, A.; Rechendorff, K.; Hovgaard, M. B.; Foss, M.; Chevallier, J.; Besenbacher, F. Colloids Surf., B 2008, 66, 53. (41) Rodahl, M.; Ho¨o¨k, F.; Fredrikson, C.; Keller, C. A.; Krozer, A.; Brzezinski, P.; Voinova, M.; Kasemo, B. Faraday Discuss. 1997, 107, 229. (42) Ho¨o¨k, F.; Rodahl, M.; Brzesinski, P.; Kasemo, B. J. Colloid Interface Sci. 1998, 208, 63. (43) Ho¨o¨k, F.; Rodahl, M.; Kasemo, B.; Brzezinski, P. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 12271. (44) Lubarsky, G. V.; Davidson, M. R.; Bradley, R. H. Biosens. Bioelectron. 2007, 22, 1275.

JP808488F