Immobilization of Enamel Matrix Derivate Protein onto Polypeptide

Quantitative Assessment of the Enzymatic Degradation of Amorphous Cellulose by Using a Quartz Crystal Microbalance with Dissipation Monitoring. Miro S...
0 downloads 0 Views 132KB Size
Download PDF
Langmuir 2006, 22, 11065-11071

11065

Immobilization of Enamel Matrix Derivate Protein onto Polypeptide Multilayers. Comparative in Situ Measurements Using Ellipsometry, Quartz Crystal Microbalance with Dissipation, and Dual-Polarization Interferometry Tobias J. Halthur,†,‡ Per M. Claesson,‡ and Ulla M. Elofsson*,† YKI, Institute for Surface Chemistry, Box 5607, SE-114 86 Stockholm, and Surface Chemistry, Department of Chemistry, Royal Institute of Technology, SE-100 44 Stockholm, Sweden ReceiVed March 22, 2006. In Final Form: July 27, 2006 The buildup of biodegradable poly(L-glutamic acid) (PGA) and poly(L-lysine) (PLL) multilayers on silica and titanium surfaces and the immobilization of enamel matrix derivate (EMD) protein was followed by utilizing in situ ellipsometry, quartz crystal microbalance with dissipation, and dual-polarization interferometry (DPI). The use of the relatively new DPI technique validated earlier published ellipsometry measurements of the PLL-PGA polypeptide films. The hydrophobic aggregating EMD protein was successfully immobilized both on top of and within the multilayer structures at pH 5.0. DPI measurements further indicated that the immobilization of EMD is influenced by the flow pattern during adsorption. The formed polypeptide-EMD multilayer films are of interest since it is known that EMD is able to trigger cell response and induce biomineralization. The multilayer films thus have potential to be useful as bioactive and biodegradable coatings for future dental implants.

Introduction The buildup of polyelectrolyte multilayer (PEM) coatings using the layer-by-layer (LbL) deposition technique was first described by Decher et al. in the early 1990s.1 This is a technique in which a PEM film is created by exposing a charged surface to alternately positively and negatively charged polymers, each leading to surface charge reversal, thereby facilitating adsorption of the next oppositely charged polymer from solution. The overcompensation and charge reversal have been illustrated by ζ-potential measurements2 and by direct force measurements.3 The LbL deposition technique is interesting for production of biomaterials, since it can be applied to surfaces of virtually any size and shape; even small glass beads4,5 and colloidal particles2 have been successfully coated. It also holds a great advantage compared to other coating techniques due to its ability to control and fine-tune the thickness and structure of the coating on a nanometer scale, simply by adjusting processing parameters, such as the polymers used, the number of layers deposited, the ionic strength at deposition,6-8 and in the case of weak polyelectrolytes the deposition pH.9-13 * To whom correspondence should be addressed. Phone: (46) 8 5010 60 40. Fax: (46) 8 20 89 98. E-mail: [email protected] † YKI, Institute for Surface Chemistry. ‡ Royal Institute of Technology. (1) Decher, G.; Hong, J. D.; Schmitt, J. Thin Solid Films 1992, 210/211, 831835. (2) Caruso, F. Chem.sEur. J. 2000, 6, 413-419. (3) Blomberg, E.; Poptoshev, E.; Claesson, P.; Caruso, F. Langmuir 2004, 20, 5432-5438. (4) Santos, J. P.; Welsh, E. R.; Gaber, B. P.; Singh, A. Langmuir 2001, 17, 5361-5367. (5) Lee, Y.; Stanish, I.; Rastogi, V.; Cheng, T.-C.; Singh, A. Langmuir 2003, 19, 1330-1336. (6) McAloney, R. A.; Sinyor, M.; Dudnik, V.; Goh, C. Langmuir 2001, 17, 6655-6663. (7) Schlenoff, J.; Dubas, S. Macromolecules 2001, 34, 592-598. (8) Ruths, J.; Essler, F.; Decher, G.; Riegler, H. Langmuir 2000, 16, 88718878. (9) Yoo, D.; Shiratori, S. S.; Rubner, M. F. Macromolecules 1998, 31, 43094318. (10) Boulmedais, F.; Bozonnet, M.; Schwinte´, P.; Voegel, J. C.; Schaaf, P. Langmuir 2003, 19, 440-445.

The interest in building polyelectrolyte multilayer films using charged polypeptides has increased in recent years. The polypeptides poly(L-glutamic acid) (PGA) and poly(L-lysine) (PLL) have been found to be useful for this purpose as demonstrated by us14,15 and others.10,16,17 These polymers have been found to build multilayers that grow more rapidly than linearly with the number of deposited layers in a highly reproducible manner.14 These polypeptides are, since they are biodegradable, of great interest for use as coatings for biomaterial applications.18 Furthermore, it has been shown that it is possible to incorporate and immobilize proteins,18-20 as well as other biomacromolecules,11,21,22 within the multilayer film. Proteins have been shown to retain their internal structure,23,24 and enzymes remain active when embedded inside multilayer films.4,5 Hence, a variety of possible applications within the biotechnology and biomaterials area, such as tissue engineering,21 immunosensing,25 and construction of hollow capsules for drug delivery,2 have been suggested. (11) Burke, S. E.; Barrett, C. J. Biomacromolecules 2003, 4, 1773-1783. (12) Shiratori, S. S.; Rubner, M. F. Macromolecules 1999, 33, 4213-4219. (13) Schoeler, B.; Poptoshev, E.; Caruso, F. Macromolecules 2003, 36, 52585264. (14) Halthur, T. J.; Elofsson, U. M. Langmuir 2004, 20, 1739-1745. (15) Halthur, T. J.; Claesson, P. M.; Elofsson, U. M. J. Am. Chem. Soc. 2004, 126, 17009-17015. (16) Lavalle, P.; Gergely, C.; Cuisinier, F. J. G.; Decher, G.; Schaaf, P.; Voegel, J. C.; Picart, C. Macromolecules 2002, 35, 4458-4465. (17) Chluba, J.; Voegel, J. C.; Decher, G.; Erbacher, P.; Schaaf, P.; Ogier, J. Biomacromolecules 2001, 2, 800-805. (18) Jessel, N.; Atalar, F.; Lavalle, P.; Mutterer, J.; Decher, G.; Schaaf, P.; Voegel, J. C.; Ogier, J. AdV. Mater. 2003, 15, 692-695. (19) Ladam, G.; Schaaf, P.; Cuisinier, F. J. G.; Decher, G.; Voegel, J. C. Langmuir 2001, 17, 878-882. (20) Ladam, G.; Gergely, C.; Senger, B.; Decher, G.; Voegel, J. C.; Schaaf, P.; Cuisinier, F. Biomacromolecules 2000, 1, 674-687. (21) Johansson, J. A.; Halthur, T.; Herranen, M.; So¨derberg, L.; Elofsson, U.; Hilborn, J. Biomacromolecules 2005, 6, 1353-1359. (22) Zhang, J.; Chua, L. S.; Lynn, D. M. Langmuir 2004, 20, 8015-8021. (23) Schwinte´, P.; Voegel, J. C.; Picart, C.; Haikel, P.; Schaaf, P.; Szalontai, B. J. Phys. Chem. B 2001, 105, 11906-11916. (24) Schwinte´, P.; Ball, V.; Szalontai, B.; Haikel, P.; Voegel, J. C.; Schaaf, P. Biomacromolecules 2002, 3, 1135-1143.

10.1021/la0607712 CCC: $33.50 © 2006 American Chemical Society Published on Web 11/08/2006

11066 Langmuir, Vol. 22, No. 26, 2006

Amelogenin proteins are secreted by the amelioblasts during formation of dental enamel. They have been found to selfassemble into monodisperse nanospheres 150-200 Å in diameter.26-29 These nanospheres are believed to play an important role in the formation of the enamel by facilitating the nucleation of hydroxyapatite crystals and guiding the crystal growth, morphology, orientation, and finally hardening of the dental enamel.30 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. The full-length protein can be described as an amphiphilic 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 8-8.3.27,28,30 The N-terminal of the protein has been identified as the part responsible for the self-assembly process through specific interactions such as hydrogen bonding and hydrophobic interactions between protein segments,31 whereas the charged C-terminal is suggested to be localized at the surface of the nanosphere, thereby hindering further aggregation due to repulsive electrostatic forces.28,30,31 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 6 month old pigs. The freeze-dried EMD product consists of a mixture of 20 kDa 80%, 13 kDa 8%, and 5 kDa 12% amelogenin proteins, all lacking the hydrophilic highly charged C-terminal (full-length amelogenin expressed in pigs has a molecular mass of 25 kDa). This protein mixture is highly aggregating and has been shown to aggregate into rather homogeneous globular spheres and short rods approximately 0.5 µm in size. It adsorbs as nanosphere multilayers on silica32 as well as on mineral- and protein-coated surfaces.33 An in vitro study has demonstrated that EMD enhanced proliferation of periodontal ligament cells (PDLs) and increased their total protein production.34 In vivo studies in monkeys have also shown that it is possible to induce regeneration of all the periodontal tissues, acellular cementum, periodontal ligament, and aveolar bone by application of EMD in a propylene glycol alginate matrix.35 Furthermore, the “EMD/propylene glycol alginate” formulation Emdogain has been shown to be clinically safe,36 able to regenerate periodontal tissues in human subjects,37 and to increase the bone level by 36% of initial bone loss within 36 months in treatments of intrabony periodontal defects.38 (25) Caruso, F.; Niikura, K.; Furlong, D. N.; Okahata, Y. Langmuir 1997, 13, 3427-3433. (26) Wen, H. B.; Fincham, A. G.; Moradian-Oldak, J. Matrix Biol. 2001, 20, 387-395. (27) Moradian-Oldak, J.; Leung, W.; Fincham, A. G. J. Struct. Biol. 1998, 122, 320-327. (28) 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. (29) 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, 50-59. (30) Fincham, A. G.; Moradian-Oldak, J.; Simmer, J. P. J. Struct. Biol. 1999, 126, 270-299. (31) Moradian-Oldak, J.; Paine, M. L.; Lei, Y. P.; Fincham, A. G.; Snead, M. L. J. Struct. Biol. 2000, 131, 27-37. (32) Halthur, T. J.; Bjo¨rklund, A.; Elofsson, U. M. Langmuir 2006, 22, 22272234. (33) Gestrelius, S.; Andersson, C.; Johansson, A.; Persson, E.; Brodin, A.; Rydhag, L.; Hammarstro¨m, L. J. Clin. Periodontol. 1997, 24, 678-684. (34) Gestrelius, S.; Andersson, C.; Lidstro¨m, D.; Hammarstro¨m, L.; Somerman, M. J. Clin. Periodontol. 1997, 24, 685-692. (35) Hammarstro¨m, L.; Heijl, L.; Gestrelius, S. J. Clin. Periodontol. 1997, 24, 669-677. (36) Zetterstro¨m, O.; et al. J. Clin. Periodontol. 1997, 24, 697-704. (37) Heijl, L. J. Clin. Periodontol. 1997, 24, 693-696. (38) Heijl, L.; Heden, G.; Sva¨rdstro¨m, G.; O ¨ stgren, A. J. Clin. Periodontol. 1997, 24, 705-714.

Halthur et al.

In this study, we address the buildup behavior of LbLconstructed PLL-PGA films by comparing earlier findings with results obtained using the relatively new technique DPI. Furthermore, we explore the possibility to immobilize the hydrophobic protein EMD on top of, as well as inside, these polypeptide films. Such polypeptide-EMD coatings, where the bioactive EMD protein is immobilized in a biodegradable polypeptide matrix, are of potential interest for dental implants. Materials and Methods Materials. PGA, MW ) 50000-100000 (cat. no. P-4886), PLL, MW ) 30000-70000 (cat. no. P-2636), and buffer salts tris(hydroxymethyl)aminomethane (Tris; cat. no. T-1503) and 2-(Nmorpholino)ethanesulfonic acid (MES; cat. no. M-8250) were all purchased from Sigma-Aldrich. EMD was received from Biora, and sodium chloride (pro analysis grade) was purchased from Merck. All chemicals were used as received, and solutions were prepared using ultrapure water (Milli-Q) (Milli-Q Plus system, Millipore). The polyelectrolytes were dissolved in an MES/Tris buffer (25 mM MES, 25 mM Tris, and 100 mM NaCl), pH 7.4. EMD was dissolved in an MES/Tris buffer with the same ionic strength, but with pH 5.0 or in acetic acid (10 mM). Solutions and buffers were stored at 4 °C and used within 2 days. Ellipsometry measurements were performed on silicon surfaces coated with approximately 1500 Å of titanium, kindly provided by Bo Thuner (Linko¨ping University, Sweden). These surfaces were passivated for at least 20 h in HNO3 (50%)39 and stored in ethanol (99.7%). The passivated titanium-coated surfaces had an oxide layer of approximately 40 Å. Quartz crystal microbalance with dissipation (QCM-D) measurements were performed on AT-cut 5 MHz quartz crystals, onto which three different layers had been evaporated, a 100 Å chromium layer to enhance adhesion, a 1000 Å gold layer to get good conductivity, and 3000 Å of titanium as the outermost substrate surface. The crystals were purchased from Q-Sense AB (Gothenburg, Sweden). Both ellipsometry and QCM-D substrate surfaces were rinsed with ethanol and Milli-Q water and then treated in a plasma cleaner (Harrick Scientific Corp., model PDC-3XG, Ossining, NY) at low pressure at 30 W for 5 min immediately before use. The waveguide chips, composed of nitrogen-doped silica, used for the DPI measurements were provided by Farfield Ltd. (Salford, U.K.). The unmodified silicon oxynitride has a hydrophilic nature with hydroxyl surface chemistry carrying negative charge. The chips were used as received, but they were cleaned inside the instrument by injection of deconex (10%) prior to calibration and adsorption. Ellipsometry. Ellipsometry is an optical method that measures the changes in polarization of light upon reflection at a planar surface.40 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 and controlled by a personal computer. Measurements were performed at a wavelength of 4015 Å 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.41 Prior to multilayer 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. Polyelectrolytes were then injected into the cuvette, and the ellipsometric angles ψ and ∆ were recorded every 10 s. If the optical properties of the substrate and the ambient medium are known, the mean thickness (df) and refractive index (nf) of the growing film can be solved numerically from the change in the optical angles ψ and (39) Brunett, D.; Tengvall, P.; Textor, M.; Thomsen, P. Titanium in Medicine: Material science, Surface science, Engineering, Biological responses and Medical applications; Engineering Materials; Springer: Berlin, 2001. (40) Azzam, R. M. A.; Bashara, N. M. Ellipsometry and polarized light; NorthHolland Publishing Co.: Amsterdam, 1977. (41) Landgren, M.; Jo¨nsson, B. J. Phys. Chem. 1993, 97, 1656-1660.

Immobilization of EMD onto Polypeptide Multilayers

Langmuir, Vol. 22, No. 26, 2006 11067

∆.42 The thickness and the refractive index were then used to calculate the adsorbed amount, Γ (mg/m2). As in our earlier studies,14,15 the adsorbed amount of polyelectrolytes was calculated using the de Feijter formula43 Γ)

df(nf - nbuffer) dn/dc

(1)

A dn/dc value of 0.15 for PGA and PLL was obtained from refractometer measurements (Wyatt Technology Optilab DSP interferometric refractometer). The adsorbed mass is usually determined with a higher accuracy than df and nf due to covariance of the errors in these latter quantities. Just as in our previous study on EMD adsorption,32 the adsorbed amount of immobilized EMD was calculated using the Cuypers formula44 Γ)

3df(nf - nbuffer)2 (nf2 + 2)[(A/M)(nbuffer2 + 2) - n(nbuffer2 - 1)]

(2)

where ν is the specific volume and M/A is the ratio of the molar refractivity (A) to the molar weight (M). For our calculations, ν ) 0.75 and M/A ) 4.1 (normal values for globular proteins)44 were used. The measurement cell system, where the substrate surface is emerged vertically in a 5 mL thermostated quartz cuvette, is a noncontinuous flow system with continuous stirring, allowing the cuvette to be rinsed between additions. DPI. DPI is a relatively new technique, able to measure changes in both the thickness and the refractive index of adsorbed layers in situ. The instrument used was an AnaLight Bio200 from Farfield Sensors Ltd. (Manchester, U.K.). A detailed description of the instrument is provided by Swann et al.45 The heart of this instrument is the substrate surface, which is a sandwich chip structure of two horizontally stacked waveguides made of nitrogen-doped silica. When plane-polarized laser light is shone on the short end of the surface, it splits and travels separately through the two waveguides (sensing and reference). As it emerges on the other side of the chip, the two signals interfere with each other; this interference can be detected by a camera as a fringe pattern in the far field. The evanescent field emitted by the sensing waveguide into the solution is affected by changes in the index of refraction and by adsorption onto the surface. Hence, the light propagating through the sensing waveguide is somewhat changed relative to the light traveling through the reference waveguide. This difference is detected as a shift in the fringe pattern in the far field, and these shifts are alternately and continuously recorded for both horizontally and vertically polarized light. From Maxwell’s equations a number of possible solutions for the thickness and refractive index of the adsorbed film can be solved for each polarization. By combining the results from both polarizations, a unique solution for the thickness and the refractive index can be obtained.46 The adsorbed masses of the polypeptide multilayer film and of EMD are then (just as in ellipsometry experiments) calculated using eqs 1 and 2. A 100 µm thick silicon mask with two slits are placed on top of the waveguide chip. As the chip is mounted in the instrument, these two slits create two separate flow channels, 1 mm wide and 17 mm long, which constitute the actual 1.7 µL measurement chambers. Solution is continuously flowed through the channels by use of a syringe pump, and samples are injected through an HPLC valve and can be directed to flow through both or just one of the two channels. (42) McCrackin, F. L.; Passaglia, E.; Stromberg, R. R.; Steinberg, H. L. J. Res. Natl. Bur. Stand. (U.S.) 1963, 67A, 363-377. (43) De Feijter, J. A.; Benjamins, J.; Veer, F. A. Biopolymers 1978, 17, 17591772. (44) 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. (45) Swann, M. J.; Peel, L. L.; Carrington, S.; Freeman, N. J. Anal. Biochem. 2004, 329, 190-198. (46) Cross, G. H.; Reeves, A.; Brand, S.; Swann, M. J.; Peel, L. L.; Freeman, N. J.; Lu, J. R. J. Phys. D: Appl. Phys. 2004, 37, 74-80.

Before the adsorption is started, the instrument and the waveguide are calibrated by injecting two solutions of known index of refraction, in this case deionized water and ethanol (80%). QCM-D. The instrument used was a QCM-D device from Q-Sense AB (Gothenburg, Sweden), with the capacity of simultaneously measuring the resonant frequency shift (∆f) and the change in energy dissipation (∆D). The instrument is described in detail by Rodahl et al.47 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 mass oscillating with the crystal ∆m according to the Sauerbrey equation48 ∆m ) -∆f/(nC)

(3)

where C is the mass-sensitivity constant (5.72 Hz at fo ) 5 MHz) and n is the overtone number. The dissipation factor (D) provides a measure of energy losses in the system. 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, the dissipation can be viewed as a measure of the rigidity or viscoelasticity of the adsorbed film.47 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. PEM Preparation and Measurements. Polypeptides were deposited directly in the measurement cell at a concentration of 1 mg/mL in MES/Tris buffer, pH 7.4. For the QCM-D and DPI measurements, polypeptide solutions were led directly into the measurement cell, whereas, for the ellipsometry measurements, stock solutions were prepared which then were diluted when added to the continuously stirred cuvette. In all three techniques, the surfaces were rinsed between polypeptide additions by exchanging the solution with pure buffer (MES/Tris, pH 7.4). A more detailed description of the buildup procedure can be found elsewhere.14 In one experiment the buildup procedure was altered to simulate a dipping protocol, which would correspond more closely to how these films would be manufactured by robots in industry production. This was done in the ellipsometer by temporarily turning off the measurement after each adsorption, completely draining the cuvette, and then filling it with rinsing buffer, which was continuously exchanged while the measurements were resumed. The measurements were then paused again, the cuvette was drained, and the next polypeptide was added to the cuvette. No additional drying was performed between the baths. By this procedure, the multilayer film passes through the air/liquid interface in the same manner as if it had been lifted and dipped in alternating baths. For measurements at pH 7.4, EMD was first dissolved (1 mg/mL) in acetic acid (10 mM) and then diluted to 75 µg/mL in MES/Tris, pH 7.4, directly in the measurement cell. When adsorption was done at lower pH, EMD was dissolved (1 mg/mL) directly in MES/Tris, pH 5.0, and allowed to adsorb at this higher concentration. The reason for choosing a lower EMD concentration at pH 7.4 is the low solubility of the protein at this pH. m2

mg-1

Results In Situ Measurements of Multilayer Buildup and EMD Immobilization. The adsorption of EMD to (PLL-PGA)6 (47) Rodahl, M.; Ho¨o¨k, F.; Krozer, A.; Brzezinski, P.; Kasemo, B. ReV. Sci. Instrum. 1995, 66, 3924-3930. (48) Sauerbrey, G. Z. Phys. 1959, 155, 206-222.

11068 Langmuir, Vol. 22, No. 26, 2006

Halthur et al.

Figure 2. Evolution of the calculated Sauerbrey mass Γ (b) and the dissipation change ∆D (O) during the pH decrease and adsorption of EMD to a (PLL-PGA)6 film measured by the QCM-D. For illustrative purposes, the mass has been set to zero at the beginning of the EMD adsorption.

Figure 1. Evolution of the (a) calculated thickness d, (b) refractive index n, and (c) adsorbed mass Γ during adsorption of EMD to multilayers ending with PGA (open symbols) and PLL (filled symbols). The EMD concentration was 75 µg/mL at pH 7.4 and 1 mg/mL at pH 5.0.

polypeptide multilayers was first studied at pH 7.4, which is the same pH used during the multilayer buildup. Virtually no adsorption could be detected onto multilayers terminated with the negatively charged PGA. A small increase in thickness and decrease in refractive index could however be observed on surfaces having the positively charged PLL as the outermost layer (see Figure 1). By decreasing the pH to 5.0, where the protein carries a small positive net charge, the adsorption was increased considerably to approximately 1.4 mg/m2 and a thickness of around 50 Å before rinsing, as seen in Figure 1. The protein desorbed during rinsing to leave 0.8 mg/m2 in a 40 Å thick film. EMD adsorption to (PLL-PGA)6 films at pH 5.0 was also studied with a QCM-D. This resulted in similar adsorption with a calculated Sauerbrey mass of approximately 1.6 mg/m2 before and 1.4 mg/m2 after rinsing (Figure 2). The dissipation factor increased considerably, by two units, during EMD adsorption, and then further to above three units during rinsing. Furthermore, the figure shows a dip in both mass and dissipation during the

buffer exchange from pH 7.4 to pH 5.0. This dip was also seen in the ellipsometry data as a decrease in both thickness and mass. The polypeptide multilayer buildup and EMD adsorption were further investigated using DPI. A good agreement was obtained for the adsorbed mass evaluated by ellipsometry and DPI during the polypeptide multilayer buildup, as demonstrated in Figure 3c. The thickness and refractive index (Figure 3a,b) show less scatter with DPI than with ellipsometry for the initial 6-7 layers, after which both techniques agree and return virtually the same values. Thus, we suggest that DPI provides more reliable data for these quantities for thin layers. The figure also demonstrates that the film properties do not differ between films prepared by the regular liquid exchange method and films prepared by the simulated dipping procedure. This was evaluated in the ellipsometer and clearly showed that the buildup was not at all affected by the passage of the adsorbed film through the air/water interface. This may, however, not be the case for other polyelectrolyte combinations. Both techniques also displayed the distinct dip in thickness and a small decrease in mass as the pH was decreased from 7.4 to 5.0. On the other hand, EMD adsorption was found to be very different in the two techniques where a huge adsorption of more than 15 mg/m2 with a thickness increase of close to 300 Å before rinsing was observed with DPI. A large fraction of the protein desorbed during rinsing, leaving approximately 8 mg/m2 with a thickness of around 130 Å. The corresponding values returned by the ellispometry measurements were 1.4 mg/m2 and 50 Å before rinsing and 0.8 mg/m2 and 40 Å after rinsing. The reason behind these differences will be discussed below. Experiments were also conducted where three layers of EMD were immobilized in the multilayer structure by alternate deposition of EMD and PGA, at pH 5.0, on a (PLL-PGA)3 substrate. This EMD multilayer buildup was followed in situ by both ellipsometry and DPI (Figure 4). The first EMD layer was significantly thicker with a larger mass adsorbed when determined with DPI (14.7 mg/m2, 245 Å) compared to ellipsometry (2.8 mg/m2, 150 Å). On the other hand, this smaller amount adsorbed in the ellipsometry experiment was still twice as much as what was measured with ellipsometry during adsorption on a (PLLPGA)6 substrate (Figure 3). In both techniques a larger amount of EMD adsorbed for each new layer, and after three layers of EMD both thickness and mass data were similar in the ellipsometer and in the DPI instrument. Both techniques showed a relatively small but significant adsorption of PGA of around 1.6 mg/m2

Immobilization of EMD onto Polypeptide Multilayers

Figure 3. Evolution of layer properties during buildup of a (PLLPGA)6-EMD multilayer film: (a) calculated thickness d, (b) refractive index n, and (c) adsorbed mass Γ measured by ellipsometry (circles), DPI (squares), and ellipsometry with simulated dipping (tilted squares).

between the EMD adsorptions, whereas the thickness did not change much and even decreased somewhat in some cases.

Discussion Some Comments on the Techniques Used. In this work we have investigated multilayer buildup using three different techniques, ellipsometry, DPI, and QCM-D measurements. Of these, ellipsometry and DPI register adsorbed amounts, whereas the QCM-D registers the mass associated with the oscillating crystal, i.e., the adsorbed polymer and the associated solvent. Comparing the adsorbed amounts obtained with ellipsometry and DPI for the (PLL-PGA)6 layer with the mass sensed by the QCM-D shows that the latter is larger, and the water content in the layer has been estimated to be approximately 70%.14 For equilibrium situations one would expect that ellipsometry and DPI would report the same adsorbed amount. On the other hand, strongly associating systems are prone to be trapped in

Langmuir, Vol. 22, No. 26, 2006 11069

Figure 4. Evolution of layer properties during the buildup of a (PLL-PGA)3-(EMD-PGA)-EMD multilayer film: (a) calculated thickness d, (b) refractive index n, and (c) adsorbed mass Γ measured by ellipsometry (circles) and DPI (tilted squares).

long-lived nonequilibrium states.49,50 In such cases the plateauadsorbed amount may depend strongly on how the adsorbate is transported to the surface. In this respect ellipsometry and DPI differ strongly. In our ellipsometry experiments a concentrated solution is added to the solution in the cuvette under constant stirring. Previous results have shown that a stagnant layer develops outside the substrate surface in the cuvette, and the stagnant layer is about 20-50 µm thick.51,52 The adsorbing species reach the surface through diffusion through this layer, a process that is relatively slow for high molecular weight substances. In contrast, the solution is constantly pumped through the DPI cell, which in total is 100 µm thick. During the low flow rate used (49) Naderi, A.; Claesson, P. M.; Bergstro¨m, M.; De´dinaite´, A. Colloids Surf., A 2005, 253, 83-93. (50) Pagac, E. S.; Prieve, D. C.; Tilton, R. D. Langmuir 1998, 14, 2333-2342. (51) Elofsson, U. M.; Paulsson, M. A.; Arnebrant, T. Langmuir 1997, 13 (6), 1695-1700. (52) Kop, J. M. M.; Corsel, J. W.; Janssen, M. P.; Coypers, P. A.; Hermens, W. T. J. Phys. 1983, C10, 491.

11070 Langmuir, Vol. 22, No. 26, 2006

in our experiments (50 µL/min) a laminar flow profile is created. The stagnant layer thickness has not been evaluated in the DPI instrument, but it is by necessity significantly smaller than in the ellipsometer. Further, since new solution is entered into the DPI cell continuously, a depletion of adsorbing species outside the surface is, unlike in the ellipsometer, never achieved. Thus, a more rapid mass transport to the surface is expected in the DPI instrument than in the ellipsometer. In the QCM-D no stirring is applied and after the solution is injected the adsorbing species have to diffuse to the surface in order to adsorb. Polypeptide Multilayer Buildup. As mentioned in the Results, the adsorbed mass for polypeptide multilayers agrees nicely when evaluated from ellipsometry and DPI measurements (Figure 3c). Thus, in this case the difference in mass transport to the surface in these two techniques does not play any significant role. The agreement between the two techniques strongly suggests that the adsorbed amount during PLL-PGA multilayer buildup previously published based on in situ ellipsometry measurements is accurate.14,15 The thickness and refractive index for the initial 6-7 layers of these highly hydrated films were more consistently determined using DPI. However, for thicker films a very good agreement was obtained between the two techniques also for the thickness and refractive index. As was suggested in our earlier paper, the widely scattered refractive indices measured by ellipsometry for the initial layers were unreasonably high, thereby yielding inaccurate values also for the film thickness.14 However, the overall trends with a two-regime buildup, where polypeptides in the initial layers adsorb as thinner and denser layers, which then turns into constantly higher increments in film thickness in combination with a close to exponential mass growth, were validated by the DPI measurements. The considerable reduction in thickness and the small decrease in adsorbed mass observed for the (PLL-PGA)6 layer with both ellipsometry and DPI (Figure 3) when the pH is decreased from 7.4 to 5.0 have been studied and discussed previously for this system.15 This pH effect was attributed to a deswelling of the film as PGA, when the pH is nearing its pKa, loses charge and consequently releases counterions. The effect could also be seen in the QCM-D experiments as a decrease in both total mass and dissipation (see Figure 2), which is consistent with loss of water and a deswelling of the film. However, the increase in dissipation found upon rinsing of the system was not reflected in an increase in thickness or decrease in refractive index, as could have been expected. The reason may be that a few loosely attached aggregates have a considerable effect on the dissipation, but would not affect a corresponding homogeneous layer, assumed in the evaluation of both ellipsometry or DPI data, significantly. Immobilization of EMD. Immobilizing EMD on top of (PLLPGA)6 multilayers proved to be somewhat more difficult than expected. At neutral pH this hydrophobic and uncharged protein does not seem to have any affinity for the multilayer film, and virtually no adsorption could be detected (see Figure 1). However, the small increase in thickness in combination with the decrease in the refractive index of PEM films ending with PLL indicates that a small number of EMD nanospheres might have adsorbed to the film. Decreasing the pH for EMD adsorption to 5.0 did however lead to definite adsorption of around 1 mg/m2 EMD, which could readily be measured by both ellipsometry and the QCM-D (see Figures 1 and 2). The increase in adsorption at pH 5.0 is suggested to be related to the fact that the EMD nanospheres carry a slight positive net charge at this pH, and at the same time the terminating PGA layer in the multilayer film loses charge, which would

Halthur et al.

make it more prone to interact with the still mostly hydrophobic protein. The slightly higher adsorbed amount measured by the QCM-D compared to ellipsometry is expected, since the QCM-D measures the total mass on the crystal, including hydration water.53,54 This is further discussed in our earlier study.14 The increase in dissipation during EMD adsorption and during rinsing indicates that the relatively large EMD nanospheres adsorbed on the PLL-PGA film dissipate some energy of the oscillator as water flows between the nanospheres on the surface, as was described earlier for EMD adsorption on silica.32 Comparing the adsorption of EMD to (PLL-PGA)6 films evaluated with ellipsometry and the QCM-D with the DPI results reveals that a considerably thicker layer and larger amount of EMD adsorbed in the DPI instrument as seen in Figure 3. Thus, in this case the more efficient transport to the surface in the DPI instrument results in a significant increase in the adsorption. The DPI measurements indicate that more than a monolayer of EMD nanospheres has adsorbed and that after rinsing 8 mg/m2 EMD stays adsorbed in a 150 Å thick layer. The latter value does according to earlier findings correspond to a full monolayer of EMD nanospheres.32 EMD adsorption measured by ellipsometry was higher for the (PLL-PGA)3 substrates compared to the (PLL-PGA)6 film (Figures 1 and 4). This suggests that EMD is more prone to adsorb to these thinner films with flatter polypeptide layers. It could also be that the surface is not fully covered by the polypeptide multilayers so that the EMD nanosperes have a chance to interact with the substrate surface as well. Upon rinsing, the outer layer of EMD starts desorbing. However, adsorbing a negatively charged PGA layer on top of the EMD layer stopped EMD from further desorbing and thus immobilized the protein inside the film. Rinsing after PGA adsorption did not lead to any desorption, as indicted by the constant mass. A small decrease in thickness and increase in refractive index could however be detected, indicating consolidation of the film. The adsorption of PGA also leads to the possibility that yet another layer of EMD could be adsorbed, and thus, LbL deposition of PGA-EMD could be achieved (see Figure 4). The adsorbed amount as well as the thickness of the EMD layer increased for each layer added (especially for ellipsometry), which in principle could indicate a roughening of the film, leading to a blooming fractal-like buildup behavior as has been reported for some PEM films at high ionic strength.6,8 Such a behavior would however lead to a decrease in the refractive index of the film, which could not be detected in any of the techniques, and we thus disregard this possibility. The increase in mass for each EMD layer is therefore suggested to be facilitated by an increased EMD affinity to the surface. We note that EMD by itself forms multilayer structures, and the data suggest that this process could be further facilitated by the introduction of alternately adsorbed PGA, which effectively can encapsulate part of the surface of the adsorbed EMD particles and reverse the charge of the surface. Finally, we note that immobilization of EMD deep inside the multilayer film does not necessarily mean that it is unavailable for cells if applied on the surface of a medical device. Jessel et al.18 have shown that PLL-PGA films can be locally biologically degraded by cells, thereby allowing them to extend their membrane into the multilayer film. Thus, EMD-PGA multilayers may be useful for improving the surface properties of implant materials. (53) Ho¨o¨k, F.; Vo¨ro¨s, J.; Rodahl, M.; Kurrat, R.; Bo¨ni, J. J. Colloids Surf., B 2002, 24, 155-170. (54) Ho¨o¨k, F.; Rodahl, M.; Brzezinski, P.; Kasemo, B. Langmuir 1998, 14, 729-734.

Immobilization of EMD onto Polypeptide Multilayers

Conclusions PLL-PGA multilayer formation has been evaluated by three different techniques: ellipsometry, QCM-D measurements, and DPI. It is found that the masses detected by ellipsometry and DPI agree excellently. On the other hand, the QCM-D registers a significantly larger mass, which is due to the fact that this technique also registers the solvent oscillating with the layer that in the present case constitutes 70% of the sensed mass. When comparing ellipsometry and DPI data for the thickness and refractive index of the multilayer film, we find more consistent data with DPI for thin films. The adsorption of EMD on and within PLL-PGA multilayer films was also investigated. It is found that EMD can be adsorbed on top of PGA layers but not on top of layers of PLL. The adsorption was found to be higher at pH 5.0 than at pH 7.4. Multilayers of PGA and EMD can also be formed, which facilitates immobilization of a larger amount of EMD on the surface. This is expected to be an advantage for production of biodegradable and bioactive coatings on implant

Langmuir, Vol. 22, No. 26, 2006 11071

surfaces. For instance, immobilized EMD is of interest for dental implants due to the ability of this protein to trigger bone growth and formation of hydroxyapatite. It was found that the adsorption of EMD occurs considerably more extensively in the DPI instrument than in the ellipsometer. This is suggested to be due to the more efficient mass transport to the surface in the DPI that may allow aggregates of EMD nanospheres to attach to the surface. Acknowledgment. This work has been done as a part of the SIMI project (Surface Improvement of Metal Implants) funded by the European Commission. Financial support has also been received from the Swedish Foundation of Strategic Research (Collintech program). We also acknowledge Biora AB for providing us with the EMD protein and Fairfield Sensors Ltd. for lending us a DPI instrument. LA0607712