Aggregation Behavior and Adsorption of Enamel Matrix

Human recombinant cementum attachment protein (hrPTPLa/CAP) promotes hydroxyapatite crystal formation in vitro and bone healing in vivo. Gonzalo ...
6 downloads 0 Views 191KB Size
Download PDF
Langmuir 2006, 22, 2227-2234

2227

Self-Assembly/Aggregation Behavior and Adsorption of Enamel Matrix Derivate Protein to Silica Surfaces Tobias J. Halthur,*,†,‡ Anna Bjo¨rklund,†,§ and Ulla M. Elofsson† YKI, Institute for Surface Chemistry, Box 5607, SE-114 86 Stockholm, Sweden, and Department of Chemistry, Surface Chemistry, Royal Institute of Technology, SE-100 44 Stockholm, Sweden ReceiVed September 14, 2005. In Final Form: January 3, 2006 Adsorption of the amelogein protein mixture enamel matrix derivate (EMD) to silica surfaces has been studied by in situ ellipsometry and quartz crystal microbalance with dissipation (QCM-D). The protein was found to adsorb as nanospheres in mono- or multilayers, depending on the concentration of “free” nanospheres available in solution. The concentration of free nanospheres is determined by the competitive processes of adsorption and rapid aggregation into microscopic particles, measured by dynamic light scattering (DLS). Multilayers could also be formed by sequential injections of fresh EMD solution. At higher temperature, an up to 6 times thicker gel-like film was formed on the substrate surface, and decreasing the pH lead to disruption of the multilayer/aggregate formation and a decreased amount adsorbed.

1. Introduction Amelogenin proteins are secreted by the amelioblasts during the formation of dental enamel. They are believed to self-assemble into nanospheres, which then combine to form the major part (∼90%) of the organic extracellular matrix, facilitating the nucleation of hydroxyapatite crystals and guiding the crystal growth, morphology, orientation, and finally the hardening of the dental enamel (further described in a resent review by Fincham et al.1). The amelogenin is spliced during secretion and degraded by enamelysin during the enamel formation, and, as a result, amelogenin extracted from developing teeth consists of a mixture of several different macromolecules with various chain-lengths. Several studies of a recombinant amelogenin protein, rM179, have proved that the full-length proteins self-assemble into rather monodisperse nanospheres 15-20 nm in diameter.2-5 The size of these self-assembled nanospheres has also been shown to increase with increasing temperature, yielding up to 4 times larger particles at physiological temperatures.3 The full-length protein can be described as a bipolar macromolecule with a hydrophilic, highly charged C-terminal with a pI of approximately 4.2, whereas the rest of the molecule is rather hydrophobic, with a pI of about 8-8.3.1,3,4 The N-terminal of the protein has been identified as the part responsible for the self-assembly process through specific interactions between protein segments,6 whereas the C-terminal is believed to stabilize the nanospheres by being exposed on the surface, and thereby hindering further aggregation.1,4,6 It has further been shown that deletion of part of the N-terminal inhibits * To whom correspondence should be addressed. Phone: (46) 8 5010 60 48. Fax: (46) 8 20 89 98. E-mail: [email protected]. † YKI, Institute for Surface Chemistry. ‡ Royal Institute of Technology. § Currently at PRV, Stockholm, Sweden. (1) Fincham, A. G.; Moradian-Oldak, J.; Simmer, J. P. J. Struct. Biol. 1999, 126, 270-299. (2) Wen, H. B.; Fincham, A. G.; Moradian-Oldak, J. Matrix Biol. 2001, 20, 387-395. (3) Moradian-Oldak, J.; Leung, W.; Fincham, A. G. J. Struct. Biol. 1998, 122, 320-327. (4) Fincham, A. G.; Moradian-Oldak, J.; Simmer, J. P.; Sarte, P.; Lau, E. C.; Diekwisch, T.; Slavkin, H. C. J. Struct. Biol. 1994, 112, 103-109. (5) Fincham, A. G.; Moradian-Oldak, J.; Diekwisch, T. G. H.; Lyaruu, D. M.; Wright, J. T.; Bringas, P.; Slavkin, H. C. J. Struct. Biol. 1995, 115, 50-9. (6) Moradian-Oldak, J.; Paine, M. L.; Lei, Y. P.; Fincham, A. G.; Snead, M. L. J. Struct. Biol. 2000, 131, 27-37.

the protein-to-protein interactions, leading to a disruption in the nanosphere formation, whereas deletion of part of the C-terminal does not hinder the nanosphere formation.6 Deletion of the C-terminal does, however, promote fusion of the formed nanospheres to generate larger aggregates.6 The fact that the amino acid sequence of both the N- and C-terminal has been well conserved, whereas the central segment differs somewhat between species, also gives a strong indication that the function of the protein (i.e., its ability to self-assemble and facilitate enamel formation) is associated with these domains. Evidence of amelogenin nanospheres has also been found in vivo using transmission electron microscopy (TEM) imaging on secretory-stage mouse enamel,5 as well as in vitro by atomic force microscopy (AFM), scanning electron microscopy (SEM), and TEM imaging on precipitated native amelogenin gels consisting of a mixture of full-length 25 kDa (7.4%), 23 kDa (10.7%), 20 kDa (49.5%), and smaller peptides (32.4%).7 The three imaging techniques showed a hierarchical structure, in which nanospheres 8-20 nm in diameter assemble to form small spherical assemblies 40-70 nm in diameter. These assemblies then aggregate further, producing larger spherical assemblies 70-300 nm in diameter. The assembly of small nanospheres into larger spheres was attributed to the fact that the majority of the protein lacked the hydrophilic, highly charged C-terminal stabilizing the nanospheres from further aggregation. Both native mixtures7 and recombinant amelogenin2 have been found to precipitate to gels at neutral pH. These gels can then undergo a thermoreversible transition, from clear at 4 °C to opaque at 24 °C. The transition has been identified as a reversible aggregation mechanism in which nanospheres fuse into much larger entities as the hydrophobic interaction increases with increasing temperature; these larger particles are separated by large liquid cavities, resulting in a heterogeneous opaque microstructure.2,7 The enamel matrix derivate (EMD) used in this study is a native amelogenin mixture extracted with acetic acid from scrapings of mandibular, nonerupted, developing premolars and molars of six-month-old pigs. The freeze-dried EMD product consists of a mixture of 20 kDa (80%), 13 kDa (8%), and 5 kDa (7) Wen, H. B.; Moradian-Oldak, J.; Leung, W.; Bringas, P., J.; Fincham, A. G. J. Struct. Biol. 1999, 126, 42-51.

10.1021/la0525123 CCC: $33.50 © 2006 American Chemical Society Published on Web 02/01/2006

2228 Langmuir, Vol. 22, No. 5, 2006

Halthur et al.

(12%) amelogenin proteins, all lacking the hydrophilic, highly charged C-terminal (full-length amelogenin expressed in pigs has a molecular weight of 25 kDa). This protein mixture is highly aggregating and has been shown to aggregate into rather homogeneous globular spheres and short rods of approximately 0.5 µm in size, and form multilayers on both mineral and protein surfaces.8 An in vitro study demonstrated that EMD enhances proliferation of periodontal ligament (PDL) cells, increases total protein production by PDL cells, and promotes mineral nodule formation of PDL cells.9 In vivo studies in monkeys have also shown that it is possible to induce the regeneration of all the periodontal tissues, acellular cementum, periodontal ligaments, and alveolar bone by the application of EMD in a propylene glycol alginate (PGA) matrix.10 Furthermore, the EMD/PGA formulation Emdogain has been shown to be clinically safe,11 able to regenerate periodontal tissues in human subjects,12 and to increase the bone level by 36% of initial bone loss within 36 months in treatments of intrabony periodontal defects.13 In this study, we wanted to focus on the adsorption of EMD protein onto solid surfaces and how this is affected by the protein self-assembly and aggregation behavior. This was done by measuring the adsorption to silica model surfaces in situ at various concentrations, temperatures, and pH, using ellipsometry and quartz crystal microbalance with dissipation (QCM-D). The aggregation behavior was also investigated using dynamic light scattering (DLS). The deduced information will be crucial for future production of EMD-based surface coatings for biomaterial applications, thereby creating a biodegradable and bioactive surface coating able to trigger and guide biomineralization, and facilitate periodontal bone attachment.

PDC-3XG, Ossining, NY) in low-pressure at 30 W for 5 min immediately before use. The EMD was dissolved in acetic acid (10 mM) to create a stock solution of 0.5-5.0 mg/mL. The stock solution was then diluted to its final concentration directly in the cuvette (ellipsometry) or in an external container just prior to measurements (QCM-D and DLS) in citrate-phosphate buffer (0.15 M) of various pH. 2.2. Ellipsometry. Ellipsometry is an optical method that measures the changes in polarization of light upon reflection at a planar surface.14 The instrument used in this study was a Rudolph thin-film ellipsometer, type 436 (Rudolph Research, Fairfield, NJ), equipped with a xenon arc lamp and high-precision step motors controlled by a personal computer. Measurements were performed at a wavelength of 401.5 nm and an angle of incidence of 67.7°. A more detailed description of the setup of the instrument is given by Landgren and Jo¨nsson.15 Prior to EMD adsorption, four-zone measurements were performed in air and in buffer solution to determine the complex refractive index (N ) n - ik) of the substrate bulk material as well as the refractive index (no) and thickness (do) of the outermost oxide layer.16 EMD was then injected into the cuvette, and the ellipsometric angles ψ and ∆ were recorded in situ every 5 s during the initial adsorption, and then at 60 s intervals during the remaining adsorption. When the optical properties of the substrate and the ambient media are known, the effective mean thickness (df) and refractive index (nf) of the growing film can be solved numerically from the change in the optical angles ψ and ∆ using a four-layer model (bulk surface/ oxide/film/ambient).17 The thickness and the refractive index were then used to calculate the adsorbed amount, Γ (mg/m2), according to the Cuypers formula:18

2. Materials and Methods

where V is the specific volume, and M/A is the ratio of the molar refractivity A to the molar weight M. For our calculations, V ) 0.75 and M/A ) 4.1 (normal values for globular proteins)18 were used. The measurement cell system, in which the substrate surface is emerged vertically in a 5 mL thermostated quartz cuvette, is a noncontinuous flow system with continuous stirring, allowing the cuvette solution to be rinsed between additions. 2.3. QCM-D. The instrument used was a QCM-D device from Q-Sense AB (Go¨teborg, Sweden), with the capacity of simultaneously measuring the resonant frequency shift (∆f) and the change in energy dissipation (∆D) at four different frequencies (fundamental, 3rd, 5th, and 7th overtone). The instrument is described in detail by Rodahl et al.19 In the QCM-D technique, a thin piezoelectric AT-cut quartz crystal with metal electrodes deposited on each side is used as the substrate surface. The quartz crystal can be exited to oscillate in shear mode at its resonant frequency fo (or at an overtone) by applying an AC voltage across the electrodes. Adsorption of a small mass (∆m) onto the crystal induces a decrease in the resonant frequency (∆f). Given that the mass adsorbed is much smaller than the mass of the crystal, is evenly distributed, does not slip on the electrode surface, and is sufficiently rigid and/or thin to have negligible internal friction, the frequency change ∆f is directly proportional to the adsorbed mass ∆m, according to the Sauerbrey equation.20

2.1. Materials. The EMD, a native pig amelogenin [20 kDa (80%), 13 kDa (8%), and 5 kDa (12%)] was kindly provided by Biora AB. Buffer salts, citric acid (C6H8O7‚H2O), and sodium phosphate (Na2HPO4‚12H2O) (pro analysis grade) were purchased from Merck. Solutions were prepared using ultrapure water (Milli-Q; Milli-Q plus system, Millipore). Solutions and buffers were used fresh or stored overnight at 4 °C. Ellipsometry measurements were preformed on silicon surfaces with a silica layer of approximately 300 Å. The surfaces were kindly provided by Dr. Stefan Welin-Klintstro¨m (Linko¨ping University, Sweden). The surfaces were cleaned by first boiling them for 5 min in Milli-Q/NH3 (25%)/H2O2 (30%) (5:1:1), followed by rinsing in Milli-Q, and then boiling for 5 min in Milli-Q/HCl (25%)/H2O2 (30%) (5:1:1). The surfaces were then immensely rinsed in Milli-Q and ethanol (99.7%) and finally stored in ethanol (99.7%) at room temperature. QCM-D measurements were preformed on AT-cut 5 MHz quartz crystal sensors, onto which three different layers had been evaporated: a 10 nm chromium layer to enhance adhesion, a 100 nm gold layer to get good conductivity, and 50 nm of silica as the outermost substrate surface. QCM-crystals were purchased from Q-Sense AB (Go¨teborg, Sweden). All substrate surfaces were rinsed with Milli-Q and ethanol and then treated in a plasma cleaner (Harrick Scientific Corp., model (8) Gestrelius, S.; Andersson, C.; Johansson, A.; Persson, E.; Brodin, A.; Rydhag, L.; Hammarstro¨m, L. J. Clin. Periodontol. 1997, 24, 678-684. (9) Gestrelius, S.; Andersson, C.; Lidstro¨m, D.; Hammarstro¨m, L.; Somerman, M. J. Clin. Periodontol. 1997, 24, 685-692. (10) Hammarstro¨m, L.; Heijl, L.; Gestrelius, S. J. Clin. Periodontol. 1997, 24, 669-677. (11) Zetterstro¨m, O.; Andersson, C.; Eriksson, L.; Fredriksson, A; Friskopp, J.; Heden, G.; Jansson, B.; Lundgren, T.; Nilveus, R.; Olsson, A.; Renvert, S.; Salonen, L.; Sjostrom, L.; Winell, A; O ¨ stgren, A.; Gestrelius, S. J. Clin. Periodontol. 1997, 24, 697-704. (12) Heijl, L. J. Clin. Periodontol. 1997, 24, 693-696. (13) Heijl, L.; Heden, G.; Sva¨rdstro¨m, G.; O ¨ stgren, A. J. Clin. Periodontol. 1997, 24, 705-714.

Γ)

0.3df(nf2 - nbuffer2) (nf2 + 2)[A/M(nbuffer2 + 2) - V(nbuffer2 - 1)]

∆m ) -

∆f nC

(1)

(2)

(14) Azzam, R. M. A.; Bashara, N. M. Ellipsometry and Polarized Light; North-Holland Publishing Company: Amsterdam, 1977. (15) Landgren, M.; Jo¨nsson, B. J. Phys. Chem. 1993, 97, 1656-1660. (16) Tiberg, F.; Landgren, M. Langmuir 1993, 9, 927-932. (17) McCrackin, F. L.; Passaglia, E.; Stromberg, R. R.; Steinberg, H. L. J. Res. Natl. Bur. Stand. 1963, 67A, 363-377. (18) Cuypers, P. A.; Corsel, J. W.; Janssen, M. P.; Kop, J. M. M.; Hermens, W. T.; Hemker, H. C. J. Biol. Chem. 1983, 258, 2426-2431. (19) Rodahl, M.; Ho¨o¨k, F.; Krozer, A.; Brzezinski, P.; Kasemo, B. ReV. Sci. Instrum. 1995, 66, 3924-3930.

Adsorption of EMD Protein to Silica Surfaces

Langmuir, Vol. 22, No. 5, 2006 2229

Figure 1. Kinetics of adsorption (adsorbed mass, Γ) at different concentrations. where C is the mass-sensitivity constant (5.72 m2 Hz mg-1 at fo ) 5 MHz), and n is the overtone number. The dissipation factor (D) provides a measure of energy losses in the system, and contains information about film interactions with the bulk solution. Adsorption/desorption as well as structural changes might lead to changes in dissipation. Generally, flat and/or rigid structures have a minimal effect on the dissipation, whereas thick and/or flexible structures increase the dissipation. Hence, dissipation can be seen as a measure of the rigidity or viscoelasticity of the adsorbed film.19 However, it is difficult to deduce any information about the adsorbed film’s viscoelasticity by looking directly at the changes in dissipation. To gain useful information, it is better to analyze how the dissipation changes in relation to the resonance frequency. This is easily done by plotting the dissipation against the resonance frequency in a so-called D-f plot. A straight line in the D-f plot would suggest that a homogeneous layer is adsorbed, while any deviations from a straight line suggests that the viscoelasticity is changing.21 A lower value for the slope indicates a relatively more rigid adsorbed layer. The quartz crystal is suspended at the bottom of a temperaturecontrolled measurement cell with a volume of 80 µL. Liquid is exchanged by a noncontinuous plug flow, first passing through a temperature-controlled loop. EMD adsorption in the QCM-D cell were made by first diluting the EMD stock solution in buffer to the concentration wanted, outside the chamber. The EMD solution was then let into the temperature-controlled loop and equilibrated before it was finally flushed into the measurement chamber approximately 7 min after dilution. 2.4. DLS. DLS measurements were performed on light-scattering equipment from Brookhaven, with a BI-200SM goniometer, a BI9000AT digital autocorrelator, and an argon ion laser (Lexel laser, model 95-2) emitting vertically polarized light at a wavelength of 514 nm. The scattering was measured at an angle of 90° relative to the in-coming beam. Scattering intensities were collected during 2 min, and then the software calculated a mean value for the hydrodynamic diameter using the Z-mean value of the scattering intensity. Measurements were started 1-1.5 min after the EMD stock solution had been injected into buffer. New measurements were then started 5, 10, 15, and 60 min after injection.

3. Results 3.1. Adsorption. Adsorption was followed in situ with ellipsometry for a variety of concentrations ranging from 1 to 75 µg/mL at 25 °C, pH 7 (Figures 1 and 2). The adsorbed amount increased steeply (exponentially; see Figure 2 inset) with the (20) Sauerbrey, G. Z. Phys. 1959, 155, 206-222. (21) Ho¨o¨k, F.; Rodahl, M.; Brzezinski, P.; Kasemo, B. Langmuir 1998, 14, 729-734.

Figure 2. Adsorption isotherm of the (a) thickness d, (b) refractive index n, and (c) adsorbed mass Γ on a linear and (inset) logarithmic scale, after 100 minutes of adsorption. The lines are not representing any data, but were added as guides for the eye.

concentration, up to a concentration of 10 µg/mL, where it leveled out. Furthermore, as can be seen in Figure 1, the rate of the adsorption also increased with increasing concentration up to ∼10 µg/mL. For concentrations of 10 µg/mL and higher, the adsorbed amount was more or less constant around 7-8 mg/m2, and similar adsorption kinetics was exhibited. In contrast, the thickness of the adsorbed protein layer seemed to increase more or less linearly with the concentration, and, above all, it continued to increase up to a concentration of 50 µg/mL, where it leveled around 40 nm (Figure 2a). As for the refractive index, it started out with values close to pure buffer at low protein concentrations, increased steeply with adsorption to a maximum of around 1.41 at 10 µg/mL, and then decreased to 1.38 for the highest concentrations (Figure 1b). This behavior is further illustrated in Figure 3, where the refractive indexes at 5, 10, and 50 µg/mL are shown.

2230 Langmuir, Vol. 22, No. 5, 2006

Halthur et al.

Figure 3. Evolution of the film refractive index during adsorption for three different concentrations.

We also performed ellipsometry experiments with sequential additions, where the protein concentration was increased stepwise in the cuvette to final total concentrations of 10, 25, and 50 µg/mL (Figure 4). The adsorbed mass and the layer thickness increased significantly with each new addition, clearly reaching higher values for adsorption at the higher concentrations (25 and 50 µg/mL) compared to what was reached in nonsequential adsorption at the same concentration. The refractive index, on the other hand, only increased during the initial adsorption (10 µg/mL), and then decreased for each of the following protein additions. The adsorption at three concentrations (10, 25, and 50 µg/mL) was further studied with QCM-D at 25 °C, pH 7 (Figure 5). The frequency response, and thereby also the Sauerbrey mass, increased with increasing concentration. The dissipation factor, on the other hand, was relatively low for the two lower concentrations, while it increased substantially for the highest concentration. This is also reflected in the D-f plot (Figure 6), which clearly shows three distinct adsorption behaviors. Adsorption at 25 µg/mL exhibits a relatively low slope in the D-f plot compared to the lowest concentration (10 µg/mL), which has a steeper slope. The highest concentration (50 µg/mL) starts out with the same slope as that for 25 µg/mL, which then increases substantially, indicating a much more viscose outer-layer structure. 3.2. Effect of pH. The adsorption of EMD was followed in situ by ellipsometry at pH 7, 5, and 3, at a concentration of 50 µg/mL (Figure 7). By decreasing the pH from 7 to 5, the adsorbed amount only decreased from 8 to 6.5 mg/m2, whereas the film thickness almost decreased by half, from 40 to 22.5 nm. When further decreasing the pH to 3, the adsorbed amount decreased substantially down to 2.5 mg/m2, while the thickness was less affected and only decreased down to 15 nm. Furthermore, the effect of rinsing was clearly reduced as the pH was decreased. 3.3. Effect of Temperature. Increasing the temperature by 10 degrees from room temperature (25 °C) to 35 °C substantially affected the adsorption (Figure 8). The adsorbed amount decreased from 7 and 8 mg/m2 (at 25 °C) to 2.5 and 3.5 mg/m2 (at 35 °C) for the two investigated concentrations 10 and 50 µg/mL, respectively. The thickness, on the other hand, increased from 14 and 38 nm, for the two concentrations at 25 °C, to a thickness of approximately 95 nm at 35 °C. The increase in temperature also lead to a significant decrease in the refractive index. 3.4. Effect of Rinsing. In all experiments, the cuvette was rinsed after approximately 100 minutes of adsorption (3 × 100

Figure 4. Time evolution of sequential adsorption at total concentrations of 10, 25, and 50 µg/mL. (a) Thickness d, (b) refractive index n, and (c) adsorbed mass Γ.

minutes for sequential adsorption) by continuously exchanging the liquid in the cuvette with pure buffer. Both the adsorbed mass and the film thickness decreased substantially during rinsing for films adsorbed at high concentrations (Figures 1, 4, and 8), whereas, for films adsorbed at 10 µg/mL and lower, the adsorbed mass and the refractive index decreased (Figures 1 and 3), but the film thickness was maintained (Figure 8). 3.5. Aggregation. The hydrodynamic diameter of the protein aggregates was measured using DLS (Figure 9). The aggregate size was found to increase with increasing concentration over a large size range (150-800 nm). The aggregate size also increased with time, especially for the higher concentrations (25 and 50 µg/mL) where the size increased steeply during the first 15 min. Furthermore, the aggregate size was found to increase slightly when increasing the temperature to 35 °C.

Adsorption of EMD Protein to Silica Surfaces

Langmuir, Vol. 22, No. 5, 2006 2231

Figure 6. D-f plot. Change in dissipation, ∆D, vs change in resonance frequency, ∆f, for the third overtone (15 MHz).

Figure 5. Time evolution of the change in (a) resonance frequency ∆f for the third (15 MHz), fifth (25 MHz), and seventh (35 MHz) overtones; (b) dissipation of the third overtone ∆D; and (c) adsorbed mass Γ calculated from the third overtone.

4. Discussion 4.1. Adsorption Behavior. As mentioned in the Introduction, both the recombinant rM1792-5 and the native7 amelogenins are known to self-assemble into nanospheres in solution. We suggest that the adsorption behavior identified in this study can be explained by the accumulation of such small self-assembled nanospheres at the solid/liquid interface. Our results clearly show a strong concentration dependence in the adsorption behavior of EMD. At low concentrations, EMD seemed to adsorb as small self-assembled particles ∼15-20 nm in diameter, as indicated by the film thickness, and sparsely spread across the surface, giving a low adsorbed mass and film refractive index (Figure 2). As the concentration was increased, the nanospheres packed more densely, which can be deduced from the increase in both the adsorbed mass and the refractive

index in combination with a constant film thickness. The packing continued until the surface was saturated with a rather densely packed monolayer of nanospheres, corresponding to the peak in the refractive index around 1.41-1.42 at a concentration of 10 µg/mL. The adsorbed amount at this concentration compares well with the ellipsometry measurement previously published for adsorption at 10 µg/mL onto a phosphate-coated aluminum oxide surface.8 At the highest concentrations, the nanospheres started to adsorb in a second layer on top of already-adsorbed particles, yielding film thicknesses around 40 nm, equivalent to one bilayer of 20 nm particles. Since the adsorption kinetics was very fast at these high concentrations (see Figure 1), the packing at the surface would not be expected to be optimal, and, hence, the second layer likely started to accumulate before a full monolayer had formed, which would explain the low refractive index of the film. The onset of the second layer was clearly reflected in the refractive index (as can be seen in Figure 3) for the highest concentration (50 µg/mL). Initially, the refractive index increased steeply during the formation of the first layer, closely following the evolution of the refractive index at 10 µg/mL. However, before a full monolayer was formed, which would be indicated by a high refractive index of 1.41-1.42, a second layer started to form, giving rise to the sudden drop in the refractive index after 5 min of adsorption. As a result, films formed at high concentrations are suggested to consist of two less densely packed layers, which, in adsorbed mass, equals one densely packed monolayer formed at 10-20 µg/mL. As seen in Figure 1, the kinetics of the adsorption at high concentrations was very fast, and there is no indication of the concentration at which a second layer is formed. Although the outer layer was not present at 10 µg/mL (as indicated by the refractive index), the sudden dip in refractive index could be seen in some of the 20 and 25 µg/mL experiments (which is what caused the large variations in the data for these concentrations) and in all of the experiments run at 40 µg/mL and above. Whether a second layer will be formed or not at intermediate concentrations could very well depend on local concentration variations during the injection and mixing of the stock protein solution in the cuvette. The adsorbed mass always decreased somewhat during rinsing, while the thickness only decreased for multilayered films formed at high concentrations, indicating that the outer layer was rather loosely attached. The described adsorption behavior is, to some extent, reflected in the AFM results for recombinant full-length amelogenin rM179 published by Wen et al.2, although at a different concentration

2232 Langmuir, Vol. 22, No. 5, 2006

Figure 7. Changes in (a) thickness, (b) refractive index, and (c) adsorbed mass with pH at a concentration of 50 µg/mL. The lines are not representing any data, but were added as guides for the eye.

range. In their experiments, nanospheres were found sparsely spread over the surface at 25 µg/mL (just as we have described the situation at the very lowest concentrations of 1-5 µg/mL), and they did not obtain a full monolayer until the concentration was raised to 100 µg/mL (10 times higher than in our results). However, the discrepancy might be explained by the fact that they only let the protein adsorb for 5 min before rinsing and fixating, and, according to our measurements, less than oneforth of the protein has adsorbed after 5 min and a substantial amount is rinsed off even after 100 minutes of adsorption (see Figure 1). It should also be kept in mind that their study was conducted using a different protein preparation. The recombinant full-length amelogenin rM179 has a C-terminal that is negatively charged at physiological pH (pI ≈ 4.2),1,3 which is believed to be situated at the surface of the self-assembled nanospheres, thereby stabilizing them from further aggregation.1,4,6 The EMD

Halthur et al.

Figure 8. Difference in adsorption behavior at room temperature (25 °C) and 35 °C of (a) thickness d, (b) refractive index n, and (c) adsorbed mass Γ.

protein used in this study, on the other hand, consists of a mixture of different molecular weights where the charged C-terminal of the native 25 kD full-length protein has been enzymatically cleaved off, yielding shorter entities 20, 13, and 5 kDa in length with relative proportions 80/8/12.10 The 20 kDa proteins (80% of the EMD mixture) still have a strong tendency to self-assemble into nanospheres, since this process is mostly driven by specific interactions between domains in the tyrosine-rich amelogenin polypeptide (TRAP) segment at the hydrophobic N-terminal of the protein, and by hydrophobic interactions in the central domains.4,6 Hence, it is likely that the 20 kDa proteins will selfassemble into nanospheres that are able to pack more densely on the surface, since the electrostatic repulsions between particles has been removed. In the absence of electrostatic repulsion, interparticle hydrophobic interactions may lead to particles that can adsorb on top of each other, thereby creating multilayers and particle aggregates.6,7

Adsorption of EMD Protein to Silica Surfaces

Figure 9. Aggregation of EMD. Evolution of the aggregate size with time at various concentrations, measured at room temperature (25 °C) and 35 °C. The time denotes the time at which the measurement was started; the accumulation of data was then conducted for 2 min. The lines are not representing any data, but were added as guides for the eye.

Evidence supporting the fact that the nanospheres aggregate into even larger particles could be seen in the light scattering data (Figure 9), in which particles with hydrodynamic diameters of well over 100 nm were found directly (1-1.5 min) after EMD injection into the DLS vial. The aggregates then grew rapidly, especially at the higher concentrations, to hundreds of nanometers in size within minutes. Such large EMD aggregates have previously been imaged with SEM, and were found to be rather monodisperse spheres or short rods.8,9 The rapid aggregation into large particles should probably be regarded more as precipitation than a self-assembly process, and these large aggregates cannot be seen adsorbing to the surface. However, the aggregation will lead to the concentration or number of nanospheres in solution rapidly decreasing with time, which could explain why the adsorption of nanospheres to the surface slowed after 10-20 minuets. The rapid aggregation also explains why the adsorbed amount calculated from the QCM-D measurements were much lower at 10 and 25 µg/mL compared to the ellipsometry results (see Figures 1, 2, and 5). Since it is not possible to inject and dilute the EMD stock solution directly in the QCM-D measurement chamber, it had to be diluted in buffer (pH 7) outside the chamber. The solution was then allowed to become temperate in the temperaturecontrolled loop for 5 min before it was finally injected into the adsorption chamber. Thus, the EMD solution was effectively allowed to precipitate into larger nonadsorbing aggregates for approximately 7 min before adsorption could start; hence, the concentration of nanospheres still available in solution would be much lower than that in the ellipsometry measurements. Considering the adsorbed amounts for 10 and 25 µg/mL in QCM-D (3 and 6 mg/m2), they roughly compare to the adsorbed amounts for 5 and 10 µg/mL in the ellipsometry measurements. Although, the QCM-D mass also includes hydration water, so the actual corresponding concentration might be even lower. The adsorbed film at these two concentrations also exhibited a rather low dissipation factor, which is to be expected for such thin films. However, looking at the D-f plot (Figure 6), the two lower concentrations exhibit rather distinct slopes, in which the low slope for 25 µg/mL indicates a rigid layer,21 consistent with a densely packed monolayer (10 µg/mL for ellipsometry), whereas the lowest concentration (10 µg/mL) has a higher slope, indicating a less rigid film where energy is dissipated to the water flowing

Langmuir, Vol. 22, No. 5, 2006 2233

between the sparsely spread particles (5 µg/mL for ellipsometry). For the highest concentration measured with QCM-D (50 µg/ mL), it seems like the residual number or nanospheres in solution was high enough for nanospheres to adsorb in a second layer. This was clearly reflected by a higher adsorbed amount for QCM-D compared to that for ellipsometry, indicating that a rather large amount of water was trapped between the loosely packed nanospheres. A further indication of this is the higher dissipation, which, in the D-f plot, is seen to first follow the 25 µg/mL line with a low slope during the formation of the first layer, and then turn into a significantly higher slope when the second layer is added and more water gets trapped. The depletion of nanospheres with time, due to aggregation, was also evident in the sequential experiment. Here, as well as in the other experiments, the adsorption appeared to have been saturated within 1 h of adsorption, at 10 µg/mL, yet more protein was adsorbed upon the injection of 15 µg/mL, and later on 25 µg/mL, of fresh protein, resulting in adsorbed amounts that were higher than what was adsorbed upon direct additions of 25 and 50 µg/mL, respectively (see Figure 4). Interestingly, it seems like a new layer was started for each of the sequential injections, leading to stepwise thickness increments of close to 15 nm each. It was also apparent that consecutive layers never packed as densely as the first layer, indicated by the relatively low mass added in new layers and by the fact that the refractive index was stepwise reduced. 4.2. Effect of pH. A decrease in pH was previously reported to disrupt the self-assembly of the recombinant full-length protein rM179, yielding smaller nanospheres with a high level of polydispersity, because the N-terminal and the center of the protein are slightly positively charged at acidic pH (pI ≈ 8.3).1,3 There was no evidence in the adsorption behavior suggesting that the self-assembly of EMD had been disrupted at acidic pH. On the other hand, no precipitation could be seen, and the thickness of the adsorbed film decreased to approximately 20 nm at pH 5.0, suggesting that the gain in positive charge stopped nanospheres from interacting with each other to form multilayers and aggregating into larger particles (see Figure 7). This is consistent with the much higher solubility found for the 20 kDa amelogenin protein at pH 6 and below.22 The refractive index increased to approximately 1.39 at pH 5, also indicating that a rather densely packed monolayer of nanospheres was formed. The film thickness was only marginally decreased when the pH was further decreased to 3.0, but the refractive index and adsorbed mass decreased significantly. The decrease in adsorption could be caused by the increase in interparticle electrostatic repulsion, hindering the nanospheres from packing densely on the surface. The fact that the desorption of protein upon rinsing was significantly reduced at acidic pH may be due to positive electrostatic interactions between the positively charged nanospheres and the negatively charged silica surface, which has a pI of approximately 2-3.5.23-25 4.3. Effect of Temperature. The size of the recombinant rM179 self-assembled nanospheres has been reported to increase rather steeply with the temperature from 34 °C and above, yielding a hydrodynamic radius 3-4 times larger (around 60 nm) at physiological temperatures compared to room temperature.3 This is probably related to the thermoreversible transition from clear to opaque in gels formed at neutral pH, reported for both (22) Tan, J.; Leung, W.; Moradian-Oldak, J.; Zeichner-David, M.; Fincham, A. G. J. Dent. Res. 1998, 77, 1388-1396. (23) Rubio, J.; Kitchener, J. A. J. Colloid Interface Sci. 1976, 57, 132-142. (24) Iler, R. K. The Chemistry of Silica: Solubility, Polymerization, Colloid and Surface Properties and Biochemistry; Wiley: New York, 1979. (25) Bolt, G. J. Phys. Chem. 1957, 61, 1166-1169.

2234 Langmuir, Vol. 22, No. 5, 2006

recombinant and native amelogenin.2,7 In agreement with these reports, lower adsorbed mass in combination with a significantly thicker film thickness was measured at 35 °C (Figure 8). Furthermore, DLS indicates the formation of a somewhat larger particle size and faster aggregation at 35 °C (Figure 9). The results would suggest that the EMD nanosphere aggregation was even further increased at elevated temperatures, which may be due to increased hydrophobic interactions.7 As a result, EMD seems to adsorb as very large, poorly packed particles in highly hydrated films, in resemblance with the opaque gels imaged with AFM.2,7

5. Summary The EMD was found to adsorb to silica surfaces as selfassembled nanospheres. At high concentrations (25-75 µg/mL) and neutral pH, these nanospheres are able to form multilayers. The same hydrophobic interaction that facilitates the adsorption of nanospheres in multilayers also leads to their rapid aggregation into larger microscopic particles, thereby effectively decreasing the concentration of “free” nanospheres in solution. Hence, the

Halthur et al.

amount adsorbed and whether they will form multilayers on the surface are mainly governed by the concentration of nanospheres available for adsorption. Multilayers could, however, also be formed by the sequential injection of fresh EMD solution. At a higher temperature (35 °C), EMD aggregated even faster, which also affected the adsorption, yielding up to 6 times thicker, highly hydrated films. Furthermore, the adsorption was seen to decrease with decreasing pH as the protein accumulates positive charges, leading to increased intraparticle electrostatic repulsions. Acknowledgment. Biora AB is acknowledged for providing us with EMD and the information thereof. The authors would also like to thank Per Claesson at the Department of Chemistry, Surface Chemistry, Royal Institute of Technology, for fruitful discussions and support. Financial support was received from the Swedish Foundation of Strategic Research (Collintech program) and the European Commission (Surface Improvement of Metal Implants (SIMI) project). LA0525123