Mica Surfaces: Relationship

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Immobilization of Metallothionein on Gold/Mica Surfaces: Relationship between Surface Morphology and Protein-Substrate Interaction Elena Casero,† Luis Va´zquez,*,‡ Jaime Martıń-Benito,§ Miguel A. Morcillo,| Encarnacioń Lorenzo,† and Fe´lix Pariente† Departamento de Quı´mica Analı´tica y Ana´ lisis Instrumental, Universidad Auto´ noma de Madrid (UAM), Cantoblanco 28049, Madrid, Spain, Instituto de Ciencia de Materiales de Madrid (CSIC), Campus de la UAM, Cantoblanco 28049, Madrid, Spain, Centro Nacional de Biotecnologıá (CSIC), Campus de la UAM, Cantoblanco 28049, Madrid, Spain, and Unidad de Iso´ topos (AÄ rea de Quı´mica) CIEMAT, Avda. Complutense 22, 28040, Madrid, Spain Received March 8, 2002. In Final Form: April 29, 2002 We have studied the surface coverage as well as the morphological and mechanical properties of metallothionein (MT) layers immobilized on gold and mica surfaces by means of three different strategies based on (1) unspecific adsorption/chemisorption (MT-Au system), (2) covalent binding (MT-modified Au system), and (3) physisorption (MT-mica). These systems have been studied by quartz crystal microbalance and atomic force microscopy techniques. The monolayers of metallothionein obtained by direct adsorption onto gold surfaces are disordered and relatively rough in contrast to those obtained by covalent binding of the protein to a previously functionalized substrate, which are smoother and show a close-packed ordered molecular arrangement. For the MT-mica system the MT layer is as smooth as the MT-modified Au layer although the imaged protein dimensions are larger, indicating a strong protein-substrate interaction. These different properties are explained on the basis of the different MT-substrate interactions governing the MT adsorption process and the MT adsorption sites available in each case. Besides, for the MT-Au system the proteins are imaged in the attractive and repulsive force regimes, the proteins being elastically deformed in the latter regime. Our results also show that covalent binding strategies not only allow the protein-substrate interaction to be controlled but can also lead to the protein layer ordering.

Introduction Metallothioneins (MTs) are ubiquitous and very unusual proteins present in a broad range of species, which have been the focus of interest of chemists and biochemists for over the last three decades. This interest is due to the special characteristics of this family of polypeptides: (1) low molecular weight (6000-7000); (2) high metal content; (3) great number of cysteine residues in the primary structure; (4) arrangement of metal ions in oligonuclear metal thiolate complexes (clusters).1 The mammalian MTs consist of a single polypeptide chain of 61 or 62 amino acid residues, 20 of them being cysteines located in highly conserved positions. All the cysteine residues are coordinated to metal ions via metal-thiolate bonds, giving rise to spectroscopic properties characteristic of metalthiolate clusters. Each protein molecule is able to bind 7 equivalents of divalent metal ions. Due to this structural design, metallothioneins show great stability at high temperatures as well as in moderately high acidic or alkaline solutions.2 Many physiological roles have been proposed for these proteins, including detoxification of heavy metals, homeostasis of essential trace metals, and scavenging of free radicals.3 Since one of the most important features of MT is its capacity to bind essential metals such as copper and * To whom correspondence should be addressed. E-mail: lvb@ icmm.csic.es. † Universidad Auto ´ noma de Madrid. ‡ Instituto de Ciencia de Materiales de Madrid (CSIC). § Centro Nacional de Biotecnologıá (CSIC). | Unidad de Iso ´ topos (A Ä rea de Quı´mica) CIEMAT. (1) Suzuki, K. T.; Imura, N.; Kimira, M. Metallothionein IIIs Biological Roles and Medical Implications; Birkhaüser Verlag: Basel, Switzerland, 1993. (2) Hamer, D. H. Annu. Rev. Biochem. 1986, 55, 913.

zinc and cytotoxic heavy metals such as cadmium, mercury, and silver, they are considered as a potential biomarker of environmental metal contamination.4 Besides, the great versatility of MTs to bind metals or free radicals such as nitric oxide or superoxide5,6 can be very useful to develop biosensors for the determination of these analytes in real samples. The immobilization of MTs on a transducer, which is the first step in biosensor design, can induce structural changes in the protein and, therefore, can alter its chemical properties and capability to bind metals or free radicals. It is known that adsorption of a protein on a flat surface is a process which involves many factors such as the existence and magnitude of van der Waals interactions as well as hydrogen, ionic, and/or covalent bonding.7-11 Besides, it depends also on the metastability of some of the protein properties, particularly those related to the protein conformational flexibility, and on the role played by the water molecules in the thin water layer between the protein and the surface.9,11,12 Thus, it is clear that, to achieve the control of the protein adsorption process required for technological applications, such as biosensor device construction, the knowledge and control (3) Sato, M.; Bremner, I. Free Radical Biol. Med. 1993, 14, 325. (4) Roesijadi, G. Comp. Biochem. Physiol., C 1996, 113, 117. (5) Sexton, D. J.; Muruganandam, A.; McKenney, D. J.; Mutus, B. Photochem. Photobiol. 1994, 59, 463. (6) Wood, P. D.; Mutus, B.; Redmond, R. W. Photochem. Photobiol. 1996, 64, 518. (7) Kinnear, K. T.; Monbouquette, H. G. Anal. Chem. 1997, 69, 1771. (8) Patel, N.; Davies, M. C.; Hartshorne, M.; Heaton, R. J.; Roberts, C. J.; Tendler, S. J. B.; Williams, P. M.. Langmuir 1997, 13, 6485. (9) Moulin, A. M.; O’Shea, S. J.; Badley, R. A.; Doyle, P.; Welland, M. E. Langmuir 1999, 15, 8776. (10) Ohnisi, S.; Murata, M.; Hato, M. Biophys. J. 1998, 74, 455. (11) Zhdanov, V. P.; Kasemo, B. Langmuir 2001, 17, 5407. (12) Ostuni, E.; Chapman, R. G.; Holmlin, R. E.; Takayama, S.; Whitesides, G. M.. Langmuir 2001, 17, 5605.

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of the protein-protein and protein-substrate interactions with nanometer precision will be essential. In this sense, during the last few years special interest has been paid to the study of the nature of the proteinsurface interaction (hydrophobic, electrostatic, or covalent) to control the protein adsorption process or even to inhibit it.12-14 In these investigations atomic force microscopy (AFM) has particular relevance. On one hand, since the development of tapping operation modes under an aqueous environment,15,16 AFM is able to provide morphological data at the molecular level of protein deposits without damaging the sample surface.13,17 Recently, lateral resolution as small as 1-1.5 nm has been reported on twodimensional crystalline protein surfaces.18 On the other hand, AFM provides important information on the protein-protein and protein-surface interactions through force spectroscopy measurements10,14,17,19,20 and on surfaceinduced conformational changes on proteins.9 Furthermore, AFM has been reported to be sensitive to enzyme activity.21 Finally, very recently AFM has also been used to produce nanopatterns of protein deposits.22 One of the main issues addressed in these studies is the relationship between the protein deposit morphology, especially the protein aggregation, and the main protein-surface interaction mechanisms.23,24 However, the molecular arrangement and the degree of ordering of the protein layer have not been specifically addressed yet. These issues are important since due to the continuous miniaturization of the technological devices, particularly biosensors,22 it would be necessary to increase the density of the protein molecules on the electrode and to control their orientation. A first step in such an investigation consists of the characterization at the nanometer level by AFM of the molecular arrangement of the protein deposits. The main goal of this work is to study the influence of the protein-substrate interactions on the mechanical and morphological properties of the protein layer. Three systems are considered: MT adsorbed directly onto a gold substrate (MT-Au), MT immobilized on a functionalized gold surface (MT-DTSP-Au), and MT adsorbed onto mica. The corresponding interactions responsible for MT immobilization are direct (unspecific) adsorption, covalent binding, and electrostatic interactions, respectively. For the MT covalent binding we have used a well-established and standard strategy.23 This approach is based on the covalent attachment of proteins containing lysine residues through the reaction of their primary amine groups with gold surfaces modified with bifunctional molecules containing succinimide functionalities. Thus, this procedure (13) Thomson, N. H.; Smith, B. L.; Almqvist, N.; Schmitt, L.; Kashlev, M.; Kool, E. T.; Hansma, P. K. Biophys. J. 1999, 76, 1024. (14) Feldman, K.; Ha¨hner, G.; Spencer, N. D.; Harder, P.; Grunze, M. J. Am. Chem. Soc. 1999, 121, 10134. (15) Hansma, P. K.; Cleveland, J. P.; Radmacher, M.; Walters, D. A.; Hillner, P. E.; Bezanilla, M.; Fritz, M.; Vie, D.; Hansma, H. G.; Prater, C. B.; Manie, J.; Fukunaya, L.; Gurley, J.; Elings, V. Appl. Phys. Lett. 1994, 64, 1738. (16) Puttman, C. A.; van der Werf, K. O.; de Grooth, B. G.; van Hulst, N. I.; Grene, J. Appl. Phys. Lett. 1994, 64, 2454. (17) Forbes, J. G.; Jin, A. J.; Wang, K. Langmuir 2001, 17, 3067. (18) Mo¨ller, C.; Allen, M.; Elings, V.; Engel, A.; Mu¨ller, D. J. Biophys. J. 1999, 77, 1150. (19) Mueller, H.; Butt, H.-J.; Mamberg, E. Biophys. J. 1999, 76, 1072. (20) Snellings, G. M. B. F.; Vansteenkiste, S. O.; Corneillie, S. I.; Davies, M. C.; Schacht, E. Adv. Mater. 2000, 12, 1959. (21) Radmacher, M.; Fritz, M.; Hansma, H. G.; Hansma, P. K. Science 1994, 265, 1577. (22) Wadu-Mesthrige, K.; Amor, N. A.; Grano, J. C.; Xu, S.; Liu, G.. Biophys. J. 2001, 80, 1891. (23) Wadu-Mesthrige, K.; Amor, N. A.; Liu, G. Scanning 2000, 22, 380. (24) Denis, F. A.; Hanarp, P.; Sutherland, D. S.; Gold, J.; Mustin, C.; Rouxhet, P. G.; Dufreˆne, Y. F. Langmuir 2002, 18, 819.

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is very suitable for the MT covalent binding since it has six lysine residues. Besides, understanding the protein interaction with and response to its natural environment is crucial for biotechnological applications. In particular, the mechanical stability of immobilized proteins on solid surfaces under physiological conditions plays an important role in biosensor development.23 Thus, we have performed a detailed study of the protein nanomechanical properties by AFM operating in a fluid environment. The study of these three different protein-substrate systems can contribute a deeper insight into the relationship between protein-substrate interactions and protein layer properties at the molecular level. These studies are a preliminary and essential step in the design of biosensors based on these proteins. Materials and Methods Reagents. 3,3′-Dithiodipropionic acid di(N-succinimidyl ester) (DTSP) and dimethyl sulfoxide (DMSO) were obtained from Aldrich Chemical Co. (Milwaukee, WI) and used as received. Sodium phosphate (Sigma Chemical Co., Saint Louis, MO) was employed for the preparation of buffer solutions. Other chemicals used in the present work were reagent grade quality. Water was purified with a Millipore Milli-Q system. All solutions were prepared just prior to use. Purification and Characterization of Rat Liver Metallothioneins. Purification. Metallothionein 2 (MT2) used in the present work was isolated from rat liver according to the following procedure: Sprague-Dawley rats were administered subcutaneously with CdCl2 (3.0 mg of Cd/kg of body weight) over 14 days; afterward the animals were sacrificed, and subsequently the livers were removed, minced, and homogenized with 3 volumes of 0.02 M Tris-HCl buffer (pH 8.6) containing 0.25 M sucrose and 1.0 mM 2-mercaptoethanol in a Potter homogenizer at 4 °C. The homogenate was centrifuged, and the supernatant fraction was collected and subsequently heated at 65 °C for 10 min. Heat-denatured samples were centrifuged at 3500g for 15 min at 4 °C to remove the heat-unstable proteins and obtain the heat-stable metallothionein in the soluble fraction. The subsequent purification of metallothionein isoforms was achieved through two-step column chromatographic fractionation of the heat-treated cytosol. An aliquot of this cytosolic crude extract was applied to a Sephadex G-75 column (65 × 2.6 cm). Protein fractions, eluting with a retention coefficient of 1.5-1.8, were collected as the total MT of the sample and were concentrated by ultrafiltration in an Amicon Ultrafiltration cell using a YM5 membrane. The last concentrated solution was applied and adsorbed onto an anion exchange column of DEAE cellulose (Sigma DEAE Sephacel) with 20 mM Tris-HCl buffer (pH 8.6) to separate the isoforms. MTs were eluted with a linear gradient of buffer solution at a flow rate of 30 mL/h. The resulting chromatographic profile yielded two major peaks corresponding to both MT isoforms. The corresponding fractions were pooled, concentrated, and desalted by ultrafiltration under the same conditions as described above. After the concentration step the protein solutions were aliquoted and stored at -80 °C. Under these conditions MTs remain stable for many months. Determination of MT by Sulfydryl Content Assay. The sensitive Ellman’s reagent DTNB [5,5′-dithiobis(2-nitrobenzoic acid)] was used for sulfydryl determination25 according to the following procedure: Aliquots of purified MT solution or 2.0-150 nmol of cysteine (for the calibration (25) Ellman, G. L. Arch. Biochem. Biophys. 1959, 82, 70.

Metallothionein Immobilization on Gold Substrates

curve) were transferred to test tubes, and the final volume was adjusted to 4.0 mL with buffer (100 mM Na2HPO4‚ 2H2O, 20 mM EDTA, and 1% SDS, pH 8.0). Then, 100 µL of a DTNB solution (39.6 mg of DTNB in 10 mL of phosphate buffer) was added. The color developed after 10-30 min was read at 412 nm against a reagent blank. The absorbance was used to calculate the sulfydryl content of the MT samples using cysteine solutions as standards. We have assumed a content of 20 cysteine molecules per MT molecule and a molecular weight for the protein of 6200 Da. Analysis of MT Samples by PAGE. MT determination by means of electrophoretic methods consists of a separation step based on the mobility of either the native or denatured protein in polyacrylamide gels. Native PAGE and SDS-PAGE have been used with the aim to check the purity of MT after its isolation and purification from liver rat. A Bio-Rad Mini-PROTEAN II dual slab cell gel apparatus was used for SDS-PAGE using the Laemmli buffer system as well as for the native gel system. The run was stopped when bromophenol blue reached the bottom of the gel. The gels were stained by placing them in a staining solution containing 0.1% Coomassie blue R-250, 40% methanol, and 10% acetic acid for 1 h. The stained gel was placed in a circulating destaining apparatus (BioRad) containing 40% methanol and 10% acetic acid for at least 24 h. High-Performance Liquid Chromatography. Highperformance liquid chromatography has been applied to the separation of MT isoforms from different tissues for a variety of eukaryotic species.26 We have used two HPLC instruments: a Waters (Waters Associates, Milford, MA) liquid chromatograph and a Waters Alliance 2690 chromatographic system. Chromatography was performed on a Delta Pack C18 column (3.9 i.d. × 150 mm, 5 µm particle size, 30 nm pore diameter; Waters). The mobile phase used to elute MT isoforms from the RP-column consisted of an equilibration buffer (10 mM NaH2PO4, pH 7.0 (buffer A)) and an elution buffer (60% acetonitrile dissolved in buffer A (buffer B)). MT isoforms were eluted from the Delta Pack C18 column with a linear gradient from 0% to 30% B for 30 min at a flow rate of 1 mL/min. Alternatively, an acidic buffer system was applied to the separation of MT isoforms. The acidic buffer system consisted of 0.12% (w/v) trifluoroacetic acid (TFA) as buffer A and acetonitrile dissolved in 0.1% TFA (60:40) as buffer B. MT isoforms were eluted from a Delta Pack C18 column with a twostep, linear gradient consisting of 0-30% B from 0 to 5 min followed by 30-40% B from 5 to 30 min at a flow rate of 1.0 mL/min. Determination of Metal Content by Atomic Absorption Spectroscopy (AAS). Zinc and cadmium are the metals associated with the MT. The determination of these metals was performed with a Varian SpectrAA-640Z atomic absorption spectrometer. Electrothermal heating of samples was carried out with a Varian GTA-100 graphite tube atomizer. The integrated area absorbances, measured during the atomization steps, were used to obtain the calibration curves and to quantify the metal contents. To compensate differences between samples and standards prepared in the laboratory, a previous 1:10 dilution with 0.1 M HNO3 was performed. In the case of heat-treated cytosol samples, an aliquot of 200 µL was made up to 10 mL with HNO3 (0.1 M), and the metals were determined by AAS. Preparation of Immobilized Metallothioneins on Flat Substrates. Direct Immobilization. To follow the (26) Richards, M. P. Methods Enzymol. 1991, 205, 217.

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immobilization of MT2 by direct adsorption onto the quartz crystal microbalance (QCM) electrodes, a conditioned QCM-Au resonator was immersed in a thermostated solution (10 mL of 0.1 M phosphate buffer at different pH values) and the frequency monitored as a function of time. After the temperature and frequency had stabilized, an aliquot of MT2 solution containing 3.8 mg/mL was added to a final concentration of 0.9 µM. The immobilization process was followed by a frequency decrease until equilibrium was reached. For AFM measurements the enzyme adsorption was carried out by immersing gold or mica supports into a solution of MT (0.9 µM in 0.1 M phosphate buffer at pH 7.0). Before being imaged by AFM, the samples were rinsed thoroughly with deionized water to remove any loosely bound residue of MT2. Immobilization of MT2 on DTSP-Functionalized Gold Substrates. A conditioned gold substrate was immersed for 3 h at room temperature in a solution of DTSP (1.0 mM in DMSO). The corresponding monolayer containing the active succinimidyl esters was subsequently rinsed in DMSO and water. Afterward, the functionalized substrate was placed in contact with a solution of MT (0.9 µM in 0.1 M phosphate buffer at pH 7.0). The immobilized protein was rinsed thoroughly in water and immediately used. Experimental Techniques. QCM Measurements. ATcut quartz crystals (5.0 MHz) of 25 mm diameter with Au electrodes deposited over a Ti adhesion layer (Maxtek Inc., Santa Fe Springs, CA) were used for QCM measurements. An asymmetric keyhole electrode arrangement was used, in which the circular electrode geometrical areas were 1.370 (front side) and 0.317 (backside) cm2. The electrode surfaces were overtone polished. Prior to use, the quartz crystals were cleaned by immersion in piranha solution, H2SO4/H2O2 (3:1 v/v). Caution: Piranha solution is extremely reactive! They were subsequently rinsed with water and acetone and dried in air. The quartz crystal resonator was set in a probe (TPS550, Maxtek) made of Teflon in which the oscillator circuit was included, and the quartz crystal was held vertically. The probe was connected to a cell by a homemade Teflon joint which was immersed into a water-jacketed beaker thermostated at the assay temperature with a thermostatic bath (Digital Temperature Controller Haake F6). The frequency was measured with a plating monitor (PM740, Maxtek) and simultaneously recorded by a personal computer. 109 Cd Binding Assay in MT Immobilized Samples. This measurement is based on an assay described previously.27 Immobilized MT2 samples onto gold disks (6.0 cm diameter) were rinsed with Tris buffer (10 mM Tris-HCl, pH 7.4) and subsequently placed in contact with 100 µL of a 109Cd solution (0.5-1.0 µCi/mL, 1.80 µg of Cd/mL as Cl2Cd in Tris buffer) for 2 h at room temperature. The samples were rinsed three times in Tris buffer, and the gold disks were transferred to γ counting test tubes. The Cd binding capacity was estimated from the radioactivity content of the samples. Atomic Force Microscopy Measurements. Two different kinds of supports (mica and gold) were used as substrates for protein attachment in the AFM measurements. The gold substrates consist of glass substrates (1.1 × 1.1 cm) covered with a chromium layer (1-4 nm thick) on which a gold layer (200-300 nm thick) was deposited (Metallhandel Schroër GmbH, Lienen, Germany). Prior to use, gold surfaces were annealed for 2 min in a gas flame to obtain Au(111) terraces. The mica substrates are easily (27) Eaton, D. L.; Cherian, M. G. Methods Enzymol. 1991, 205, 83.

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cleaved to generate atomically flat surfaces, an ideal substrate for AFM studies. The atomic force microscope was a Nanoscope IIIa from Digital Instruments (DI; Santa Barbara, CA) used with a D scanner (maximal scan range of ∼14 µm). Measurements were carried out by using Si3N4 cantilevers (nominal spring constant of 0.38 N/m) from Digital Instruments. The samples were imaged with different cantilevers to ensure that the imaged structures were not due to tip artifacts. Imaging under these conditions led to consistent and reproducible results. Image acquisition times were between 2 and 6 min for images with pixel resolutions of 512 × 512. All measurements were carried out in aqueous buffered solutions since the protein structure may be distorted by the drying process and the biological relevance of dry samples may be questioned. The substrate was first imaged in buffer solution to ensure that the surface was flat and clean before the protein immobilization was carried out. Both the sample and cantilever were located within a Plexiglas fluid cell where extremely small volumes (∼50 µL) of buffer were added. Contact-mode AFM imaging of the samples was unsuccessful since the protein structures were altered and damaged by the cantilever tip to which proteins attached during the imaging process, leading to unreliable and irreproducible images. This adhesion hampers the use of force-distance curves for the simultaneous study of the topographic and mechanical properties of the protein monolayer. However, the contact mode was used for atomic and molecular resolution imaging of the gold and DTSPmodified gold samples, respectively. Tapping-mode AFM imaging has been used to image the different protein deposits and substrates. In this mode the cantilever is oscillated in the aqueous buffers at a frequency close to its resonant frequencies, f0, in the ∼911 kHz range with amplitude A0 above the surface. The probe interacts with the sample surface, leading to a decrease of the cantilever amplitude, A (A < A0). The sample surface is imaged by keeping constant the imaging amplitude A. The lower the ratio A/A0, the higher the average applied force on the sample.28-30 We observed that tapping-mode imaging at A/A0 ) 0.92-0.96 led to reproducible images of the MT layers without any appreciable alteration of the protein structures. In some cases, imaging at different A/A0 ratios was performed on a given sample area to study the mechanical response of the protein structures to the force exerted on them by the probe. Phase contrast imaging was performed simultaneously with the standard topographic tapping-mode imaging on some samples. In these images we measure the phase lag of the cantilever oscillation with respect to the excitation force. This measurement contains information about the interaction between the tip and sample. The origin and nature of the phase contrast has been discussed over the last few years.31-34 However, it seems clear that phase contrast arises mainly from differences in the energy dissipation between the tip and the sample32,35,36 although (28) Garcia, R.; Paulo, A. S. Phys. Rev. B 1999, 60, 4961. (29) Fain, S. C.; Barry, K. A.; Bush, M. G.; Pittenger, B.; Louie, R. N. Appl. Phys. Lett. 2000, 76, 930. (30) Yu, M.; Kowalewski, T.; Ruoff, R. S. Phys. Rev. Lett. 2000, 85, 1456. (31) Anczykowsky, B.; Kruger, D.; Fuchs, H. Phys. Rev. B 1996, 53, 15485. (32) Tamayo, J.; Garcia, R. Appl. Phys. Lett. 1998, 73, 2926. (33) Nony, L.; Boisgard, R.; Aime, J. P. J. Chem. Phys. 1999, 111, 1615. (34) Bar, G.; Brandsch, R.; Bruch, M.; Delineau, L.; Whangbo, M. H. Surf. Sci. 2000, 444, L11.

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some other contributions can appear for tapping-mode experiments performed in liquid.37 Basically, when the tip is far from the surface, it oscillates freely (A ) A0) and the phase shift, ∆Φ, is 0° in DI instruments. When it approaches the sample surface, it undergoes a long-range attractive interaction (noncontact regime), leading to the damping of the oscillation (A < A0) and to ∆Φ < 0° (DI convention). When repulsive forces dominate the tipsample interactions (intermittent contact regime), that is, when the tip is closer and intermittently contacts the sample surface, A < A0 and ∆Φ > 0° (DI instruments). Thus, it can be shown that a more-energy-dissipative feature would appear light in the noncontact regime and dark in the intermittent contact regime.38 Results and Discussion Characterization of Purified Methallothioneins. A preliminary step in our study consists of the characterization of the MT samples obtained after the purification process. Two major isoforms of MTs, MT1 and MT2, have been identified in mammals, which are similar with respect to metal-binding capacities and sulfydryl content, differ slightly in amino acid content, and exhibit a single charge difference at neutral pH.39 They can be separated by high-performance liquid chromatography.26 Figure 1A shows the PAGE analysis, in native conditions, of the heatstable cytosolic fraction (line 1), used as starting material, as well as the fractions corresponding to the purified MT2 (line 2) and MT1 (line 3) after their chromatographic separation. These last two electrophoretic profiles are in agreement with those obtained for commercial MT2 and MT1 from Sigma (lines 4 and 5, respectively). In addition, Figure 1B presents the HPLC analysis corresponding to the MT2 fraction which shows a high degree of purity. Fractions containing MT1 gave similar levels of purity; however, to clarify the results, further studies were carried out using the isoform MT2 exclusively. The amount of MT2 present in the stock solutions was estimated from the sulfydryl content as described in the Materials and Methods. According to these measurements, stock solutions of MT2 in water at a concentration of 3.8 mg/mL were used in further studies. The metal content of these protein samples was estimated from atomic absorption measurements. According to the results, 260 µg/mL Cd and 150 µg/mL Zn were found. These figures are consistent with a stoichiometry of Zn3Cd4-MT2. Study of MT2 Immobilization on Gold Substrates by QCM. We have studied the immobilization process of MT2 on gold substrates by means of QCM measurements. The QCM technique allows the measurement of mass changes at surfaces through changes in the resonant frequency of the quartz crystal. These measurements allow kinetic information about the adsorption process and the surface coverage of the MT2 monolayers to be obtained simultaneously. Figure 2 shows the resulting frequency changes associated with the immobilization process of MT2 on the gold surface in a phosphate buffer solution at pH 3.0. Upon addition of the protein, the frequency decreases gradually during the first 5 min, and then a steady state (35) Cleveland, J. P.; Anczykowski, B.; Schmid, A. E.; Elings, V. B. Appl. Phys. Lett. 1998, 72, 2613. (36) Anczykowski, B.; Gotsmann, B.; Fuchs, H.; Cleveland, J. P.; Elings, V. B. Appl. Surf. Sci. 1999, 140, 376. (37) Tamayo, J. Appl. Phys. Lett. 1999, 75, 3569. (38) James, P. J.; Antognozzi, M.; Tamayo, J.; McMaster, T. J.; Newton, J. M.; Miles, M. J. Langmuir 2001, 17, 349. (39) Klaassen, C. D.; Lehman-McKeeman, L. D. Methods Enzymol. 1991, 205, 190.


Figure 1. (A) Analysis by PAGE, in native conditions, of some purification steps of rat liver MT1 and MT2. Line 1, fraction corresponds to total MT content after heat cytosolic treatment and G-75 filtration; line 2, MT2 fraction after ionic exchange purification in DEAE-cellulose; line 3, MT1 purification after DEAE-cellulose purification; lines 4 and 5, commercial MT2 and MT1 from Sigma, respectively. (B) Reversed-phase HPLC at neutral pH of MT2 purified from rat liver whose electrophoretic profile is presented in line 2.

Figure 2. Time dependence of the frequency changes of a quartz-crystal resonator in a solution containing 0.9 µM MT2 in 0.1 M phosphate buffer, pH 3.0 at 25.0 ( 0.1 °C. The solid line represents data fitted to a first-order kinetic equation.

is reached. Considering that the frequency decrease obtained (∆F ) 12.0 Hz) is only due to the change in mass arising from the adsorption of MT2, one can calculate the amount and surface coverage, Γ, of the adsorbed MT2 monolayer by means of the Sauerbrey equation:

∆m ) -Cf ∆F where ∆m is the mass change (ng cm-2) and Cf (17.7 ng Hz-1 cm-2) is a proportionality constant for the 5.0 MHz crystals used in this study. A value of Γ ) 3.4 × 10-11 mol cm-2 was obtained for the surface coverage. It should be noted that the theoretical surface coverage estimated from

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a monolayer of MT2 according of its molecular size40 is in the range of 5 × 10-12 to 4 × 10-11 mol cm-2, depending on the protein orientation and packing in the monolayer. As mentioned above, the shape of the frequency-time profile can be employed to study the kinetics of the adsorption. The process can be controlled by either transport (diffusion-controlled) or kinetics (activationcontrolled), which predict time dependencies of t1/2 and exp(t), respectively. Assuming that the immobilization process is kinetically controlled, we attempted to fit the data (Figure 2, solid line) to a first-order kinetics equation: ∆F ) ∆Fmax(1 - e-kt), where ∆F is the frequency change (Hz), ∆Fmax is the frequency change between the initial and the final steady-state frequencies, and k is the first-order rate constant (min-1).41 From the fit of the experimental data, values of ∆Fmax ) 12.1 Hz and k ) 1.20 min-1 were obtained. In principle, two main mechanisms can contribute to the direct adsorption of MT2 onto gold surfaces: a chemisorption process involving free thiolate groups and unspecific adsorption through other outer functional groups. The relative balance between them can change with pH. The first one is enhanced at low pH since under these conditions the metal atoms (Cd and Zn in particular) are dissociated from the protein, giving rise to an increase in the free thiolate groups and enabling their chemisorption onto the gold surface. To assess the metal binding capacity of the MT2 monolayer obtained by direct adsorption at low pH, we estimated it by 109Cd exchange experiments. Aliquots of MT2 were placed in contact with gold disks in buffer solution at pH 3.0. Under these conditions, Zn and Cd are readily removed from the protein and the apometallothionein (apo-MT2) is chemiadsorbed onto the gold surface. Afterward, the gold disks, containing the apo-MT2 monolayer, were placed in contact with a solution containing 109 CdCl2 (see the Materials and Methods). In these experiments we have observed that the capacity to bind Cd of the apo-MT2 monolayer, immobilized at pH 3.0, is significantly lower than that observed for no immobilized MT2. The metal/protein ratio was determined using as surface coverage the value estimated from QCM experiments, Γ ) 3.4 × 10-11 mol cm-2, and the Cd surface coverage calculated by the Cd binding assay was found to be 1.72 × 10-11 mol cm-2. From these results the calculated ratio was 0.5 mol of Cd/mol of immobilized apo-MT2. As in mamalian MTs 20 cysteine thiolates serve as ligands of 7 divalent metal ions, only a small fraction of cysteine residues would be free after the apo-MT2 is immobilized. This fact explains the lower metal binding capacity of directly immobilized MTs, especially in solutions at low pH. Thus, under low pH immobilization conditions, the protein is strongly attached to the surface, but its capability for metal binding is significantly reduced, and accordingly, the potential use of these monolayers as metal biomarkers could be strongly restricted. Therefore, higher values of pH are required to retain the metal biomarker capability of the MT monolayer. For this reason, the variation of the surface coverage as a function of pH was studied in the range from pH 2.5 up to pH 7.0, and the results are presented in Table 1. According to these results, a small increase of the surface coverage was obtained at pH around 4.5, which is very close to the isoelectric point (40) Furey, W. F.; Robbins, A. H.; Clancy, L. L.; Winge, D. R.; Wang, B. C.; Stout, C. D. Science 1986, 231, 704. (41) Abad, J. M.; Pariente, F.; Hernańdez, L.; Abrunã; H. D.; Lorenzo, E. Anal. Chem. 1998, 70, 2848.

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Table 1. MT2 Surface Coverage Dependence on pH Conditionsa pH

∆F (Hz)

Γ × 1011 (mol cm-2)

pH

∆F (Hz)

Γ × 1011 (mol cm-2)

2.5 3.8

12.0 8.5

3.4 2.4

4.5 7.0

15.6 2.5

4.4 0.7

a Surface coverage (Γ) and variation in the frequency (∆F) measured once the steady state is reached of a quartz-crystal resonator in a solution containing 0.9 µM MT2 in 0.1 M phosphate buffer at different pH values.

Figure 3. Time dependence of the frequency changes of an unmodified quartz-crystal resonator (top) and a DTSP-modified quartz-crystal resonator (bottom) in the presence of 0.9 µM MT2 in 0.1 M phosphate buffer solution, pH 7.0 at 25.0 ( 0.1 °C.

(IEP) of the protein. This result is consistent with previously reported experiments of protein adsorption.8,23 We have performed QCM and AFM experiments at pH 7.0 since it is a physiologically relevant value and also meets the biomarker requirements. Two model systems have been chosen to study the influence of the proteinsubstrate interactions on the monolayer properties. The first one consists of MT2 adsorption on bare gold substrates. Under these conditions an unspecific adsorption mechanism is predominant. The second one is based on the use of functionalized gold substrates. It is known that exposure of gold surfaces to solutions of dithiobis-Nsuccinimidylpropionate (DTSP) gives rise to monolayers of N-succinimidyl-3-thiopropionate. These exposed succinimidyl monolayers can react with amino groups, allowing for the covalent immobilization of a number of proteins containing external lysine residues.23,42 Similar QCM experiments were carried out to study the immobilization of native MT2 on DTSP-modified gold surfaces. Figure 3 shows the frequency decrease as a function of time when native MT2 solutions are placed in contact with bare gold (top curve) or DTSP-functionalized gold substrates (bottom curve). In the DTSP-Au substrate the immobilization of protein first involves a drastic decay in the frequency over 3 min, followed by a short transient in which the frequency increases slightly, to finally reach a constant frequency value. This profile can be interpreted considering three steps in the immobilization process. The first one involves the fast reaction between the succinimidyl rests present on the surface with amino groups of the protein. The second one involves structural rearrangements in the growing monolayer. The third step is the final steady state. The frequency decrease obtained at the steady state was estimated around ∆F ) 3.5 Hz corresponding to a value of surface coverage of Γ ) 1.1 × 10-11 mol cm-2. This value is higher than that observed for MT2 immobilized on bare

Figure 4. (A) 5.34 µm × 5.34 µm tapping-mode AFM image taken under buffer conditions of the as-prepared gold substrate. (B) 310 nm × 310 nm tapping-mode AFM image taken under buffer conditions obtained on one of the large grains of the as-prepared gold substrate.

gold electrodes under the same experimental conditions (Γ ) 0.7 × 10-11 mol cm-2), suggesting differences in the protein packing as a function of the preliminary treatment of the gold surface. AFM Study of Mechanical and Morphological Properties of the MT2 Monolayers. Bare Gold Substrates. AFM measurements were carried out to study the MT2 monolayers obtained on gold substrates to obtain more information about the different immobilization approaches. Figure 4A shows a 5.34 µm wide image of a bare as-prepared gold substrate. The images reveal smooth grains of size in the 0.5-1.0 µm range with deep grain boundaries leading to a root-mean-square roughness (σ) of 13 nm. However, the surface of the grains is considerably smoother (σ e 0.1 nm), showing the typical step arrangement of a Au(111) surface as can be seen in Figure 4B. It should be noted that the height range of Figure 4B has been adjusted to be the same as those corresponding to the AFM images of the gold surfaces with bound proteins (Figures 5B, 10A, and 12). Furthermore, when this surface was imaged in contact mode on the atomic scale, we observed the atomic arrangement of the Au(111) surface (data not shown). MT2 Directly Adsorbed onto Gold. Figure 5A shows a 452 nm wide image of a sample prepared by direct adsorption of MT2 onto a gold surface as described in the Material and Methods. The image has been taken on one


Figure 5. (A) 452 nm × 452 nm tapping-mode AFM image taken under buffer conditions of a MT2 layer adsorbed onto a gold substrate at pH 7.0. Together with the protein structures the (111) facets of the substrate are still visible. (B) 300 nm × 300 nm tapping-mode AFM image taken under buffer conditions of a MT2 layer adsorbed onto a gold substrate. Note the disordered surface morphology.

of the micro-sized gold grains similar to those observed in Figure 4A. In fact, despite the MT layer deposit, the characteristic triangular facet arrangements of the Au(111) surface are still observed. The overall surface roughness is 0.6 nm. A higher magnification image (300 nm wide) is displayed in Figure 5B. Both images show that the protein layer does not present any degree of ordering, although the layer appears to be quite compact. The typical dimensions of the proteins in the layer are h ) 0.6-1.2 nm (height) and dfwhm ) 4-5 nm (defined as the diameter or full width at half-maximum). The contrast of the MT molecular morphology obtained by AFM differs in some extent from that obtained by scanning tunneling microscopy (STM) operating in a fluid environment.43 However, this fact is not surprising since the probe-sample interactions, and therefore the image contrast, in both techniques are quite different. In fact, for MT2 molecular STM imaging the tunneling current seems to be enhanced by the metal ions present inside the R and β domains. Clearly, this source of contrast does not exist in AFM imaging. (42) Darder, M.; Takada, K.; Pariente, F.; Lorenzo E.; Abrunã, H. D. Anal. Chem. 1999, 71, 5530. (43) Maret, W.; Heffron, G.; Hill, H. A. O.; Djuricic, D.; Jiang, L.; Vallee, B. L. Biochemistry 2002, 41, 1689.

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Figure 6. (A) 84.5 nm × 84.5 nm tapping-mode AFM image of a MT2 layer adsorbed onto a gold substrate at pH 7.0. The surface was imaged at A/A0 ) 0.96. The rectangle indicates the limits of the surface profile shown in Figure 8A. (B) Same area of (A) imaged at A/A0 ) 0.89. (C) Same area of (A) imaged at A/A0 ) 0.77. The rectangle, which corresponds to the same sample area of the rectangle depicted in (A), indicates the limits of the surface profile shown in Figure 8B. (D) Contrast-phase image measured simultaneously with (A). The rectangle indicates the limits of the surface profile shown in Figure 8A. (E) Contrast-phase image measured simultaneously with (B). (F) Contrast-phase image measured simultaneously with (C). The rectangle indicates the limits of the surface profile shown in Figure 8B. The bars indicate 15 nm.

On this sample we have studied the mechanical response of the nanometer protein structures to the force exerted on them by the probe using the tapping-mode AFM imaging. The experiments consisted in imaging the same sample area under different loads (i.e., imaging force), that is, at four A/A0 ratios (0.96, 0.89, 0.82, and 0.77) with the same cantilever. Figure 6A corresponds to the lowest force one, A/A0 ) 0.96, Figure 6B to A/A0 ) 0.89, and Figure 6C to the highest force one, A/A0 ) 0.77. The drift during these measurements, estimated to be 0.15 nm/s along the horizontal direction mainly, accounts for the slight differences between the images. It is worth mentioning that the protein structures are more sharply imaged for A/A0 e 0.89. It is observed that the measured surface roughness decreases (Table 2) and the height distribution function of the images becomes narrower (Figure 7) as the A/A0 ratio decreases (i.e., imaging force increases), indicating the soft nature of the imaged structures. This fact can also be observed in Figure 8, in which the top curves show the topographical surface profile meas-

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Figure 7. Height distributions corresponding to the same area of the MT2 layer adsorbed onto a gold substrate at pH 7.0 imaged at different forces: A/A0 ) 0.96 (solid line), A/A0 ) 0.89 (dotted line), and A/A0 ) 0.77 (dashed line). Table 2. Dependence of the MT2 Layer Roughness on the Applied Force for the MT2-Au System A/A0 ratio

surface roughness (nm)

A/A0 ratio

surface roughness (nm)

0.96 0.89

0.31 0.30

0.82 0.77

0.27 0.24

ured along the proteins contained inside the rectangle displayed in Figures 6A,C, respectively. That is, the top curves of Figures 8A,B are obtained on the same surface region under the lowest and highest force imaging conditions, respectively. From the comparison of these topographical surface profiles it is evident that the imaged structures are deformed by the tip at higher loads. Note that not all the protein structures are deformed to the same extent. We checked experimentally that the protein deformation was reversible since the protein structures, after they had been imaged at high force, recovered their original dimensions when they were finally imaged again at lower forces (data not show). Therefore, the proteins are elastically (not plastically) deformed. Simultaneously with these topographic tapping-mode images, we have recorded the contrast phase images of the same area (Figure 6D-F). At low forces (A/A0 ) 0.96, Figure 6D) the protein structures present positive ∆Φ values since slight protuberances (lighter areas) are detected in the phase images at those locations corresponding to protein structures in the topographic image. While for slightly higher forces (A/A0 ) 0.89) the phase contrast is appreciably reduced (Figure 6E), for higher forces (A/A0 ) 0.77, Figure 6F) the contrast is increased and reversed. Thus, the protein structures appear as small depressions (darker areas) rather than as small bumps. This effect can be better observed in the bottom curves of Figure 8, which are the phase contrast profiles measured simultaneously with the surface profiles shown in the corresponding top curves at the lowest force (Figure 8A) and the highest one (Figure 8B). The comparison of parts A and B of Figure 8 confirms the phase contrast reversal at higher loads. This change of contrast is the behavior expected when an energy-dissipative feature, in our case the protein structures, is imaged first in the noncontact regime, where attractive interactions dominate, and finally in the intermittent-contact regime, in which repulsive interactions are dominant.38 It is important to stress that in the repulsive interaction regime (A/A0 e

Figure 8. (A) Topographic surface profile (top curve) and the corresponding phase contrast profile (bottom curve) measured at A/A0 ) 0.96 inside the rectangles displayed in parts A and D, respectively, of Figure 6. (B) Topographic surface profile (top curve) and the corresponding phase contrast profile (bottom curve) measured at A/A0 ) 0.77 inside the rectangles displayed in parts C and F, respectively, of Figure 6. The vertical dashed lines mark each individual protein structure as a guide to the eye to better identify the changes in the visualization of the protein structures with the applied force.

0.89) the protein structures are better (i.e., sharper) imaged and they are elastically, i.e., reversibly, deformed, at least in the force range sampled in this work. These results differ from those recently reported on tapping-mode imaging, in air conditions, of single antibody molecules.44 In this work it was reported that the antibody molecules were better resolved in the attractive interaction regime. Besides, imaging of these biomolecules in the attractive interaction regime led to their (i.e., plastic) irreversible deformation. However, as the authors of this report discussed, tapping-mode imaging in air implies the existence of relatively strong capillary forces which could contribute to the irreversible sample deformation of the antibody molecules in the repulsive interaction regime. Thus, our results indicate that nondestructive imaging, within nanometer resolution, of protein structures in tapping mode in a buffered environment in the repulsive interaction regime is possible. The surface of the MT2 layer adsorbed onto gold shows a compact and disordered morphology. In fact, the layer is rougher when it is compared to that measured for the MT2-DTSP-Au system (see below and Table 3). Both the rough layer morphology and the different elastic deformation observed for the directly adsorbed MT2 structures suggest that this protein has different adsorp(44) San Paulo, A.; Garcia, R. Biophys. J. 2000, 78, 1599.

Metallothionein Immobilization on Gold Substrates Table 3. Main Morphological Properties of the Three Protein-Substrate Systems Studied in This Work h (nm) dfwhm σ (nm)

MT2-Au

MT2-DTSP-Au

MT2-mica

0.6-1.0 4.0-5.0 ∼0.6

0.5-0.9 2.0-3.0 ∼0.3

0.4-0.9 3.0-6.0 ∼0.3

a Range of height (h) and diameter measured at half of the maximum height (dfwhm) of the protein structures as well as the root-mean-square surface roughness (σ).

Figure 9. 11.8 nm × 11.8 nm contact-mode AFM image of the DTSP-modified gold substrate. Individual DTSP molecules, disorderly arranged, are observed.

tion geometries. This conclusion is consistent with the fact that the main MT2 adsorption mechanism, under our experimental conditions (buffered solutions at neutral pH), is the unspecific one, and therefore, the adsorption takes place through different outer positions of the MT2 structure. MT2 Immobilized onto DTSP-Functionalized Gold Substrates. To fully characterize morphologically the MT2 adsorption process on the DTSP-functionalized gold substrates, we have first imaged the DTSP-gold surface before protein immobilization. At low magnification the sample surface is similar to that reported for the bare gold substrate (Figure 4A). The differences arise when the surface is imaged in contact mode at molecular resolution as can be observed in Figure 9. This figure shows individual DTSP molecules disorderly arranged. The distance between two adjacent molecules is ∼0.5-0.6 nm. This result indicates that the DTSP layer is less tightly packed than the methyl-terminated linear thiol.45 This looser arrangement of the DTSP monolayer can affect the contact-mode AFM imaging since the DTSP molecules could be more easily displaced by the tip. Finally, it is worth noting that we atomically resolve the gold surface when the imaging force is largely increased (data not shown). AFM imaging of an MT2 layer adsorbed onto DTSPgold surfaces for an incubation time of 20 h at pH 7.0 showed ordered regions such as that displayed in the 134 nm wide image of Figure 10A. The surface is quite smooth (σ ) 0.3 nm) and shows an evident ordering of the globular proteins. The protein dimensions are h ) 0.5-0.9 nm and dfwhm ) 2-3 nm. To assess the MT2 layer ordering, we have performed a fast Fourier transform (FFT) analysis of the AFM data. A typical FFT is shown in Figure 10B, in which six spots hexagonally ordered are clearly (45) Terań Arce, F.; Vela, M. E.; Salvarezza, R. C.; Arvia, A. J. J. Chem. Phys. 1998, 109, 5703.

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observed. Since the protein layer is not a two-dimensional protein crystal,46 only six relatively wide spots are obtained in the FFT image. To know the periodicity along the three fundamental directions of the structure, we have performed a cross section analysis of the FFT image (Figure 10C). Despite the different broadnesses and intensities of the spots, it can be concluded that the main periodicity of the structure in the reciprocal space is k0 ) 0.16 nm-1 (i.e., 6 ( 1 nm in the real space). The intensity of the spots is related to the different degrees of order along each direction. To further analyze the protein layer ordering, we have also performed the two-dimensional autocorrelation analysis of the AFM data (Figure 10D). In this figure the short-range quasi hexagonal ordering with a nearest neighbor protein distance of ∼6 nm is clearly observed. The dotted structure of the image all around these six spots indicates that some sort of long-range order, although weaker than the short-range one, is still present. These results indicate that protein structures in the MT2 layer adopt a short-range hexagonal ordering typical of a closepacked structure. In this system we have attempted to perform a study on the dependence of the AFM imaging with the applied force similar to that realized on the MT2-Au layer. However, in this case we only could vary the A/A0 ratio from 0.95 down to 0.90. The application of higher forces (i.e., smaller A/A0 ratios) led to the image distortion. In this narrower force range the images obtained at different forces were very similar. Figure 11A shows the height distribution function of the same area imaged at A/A0 values of 0.95 (solid line) and 0.9 (dashed line). Both distributions are quite similar in contrast to the MT2Au case (Figure 7), where some differences are already observed for the same A/A0 range. In addition, the surface roughness is slightly higher at A/A0 ) 0.9, indicating better imaging conditions for this setting. Figure 11B shows the surface profiles of the same sample region measured at A/A0 values of 0.95 (top curve) and 0.9 (bottom curve), which are quite similar with no appreciable deformation or damaging of the protein structures. In the corresponding phase contrast images (not shown) the protein structures appear as slight bumps (lighter regions), implying imaging in the attractive interaction regime at both A/A0 ratios. There is not an appreciable difference of contrast between both images. However, for the MT2-Au system the contrast was already clearly reduced when A/A0 was changed from 0.96 to 0.89. These results suggest that the tip-sample interaction is different for the MT2-Au and MT2-DTSP systems, which is consistent with a different protein-substrate interaction. These observations indicate that covalent binding is stronger than physical interactions in the case of protein immobilization as already reported for other systems.23 Thus, the MT2-DTSP-Au system presents morphological and mechanical properties different from those observed for the MT2-Au system. The MT2-DTSP-Au layer is more homogeneous and 2 times smoother than the MT2-Au layer. In addition, the imaged protein dimensions are, on average, smaller than in the MT2-Au case. Besides, it shows a close-packed (next neighbor distance of 6 ( 1 nm) ordered structure. Finally, the mechanical response of the protein structures in the MT2DTSP system also seems to be different from that measured in the MT2-Au system. These data suggest both that the protein adsorption geometry is different and that the nature of the protein-substrate interaction is different from those for the MT2-Au system. (46) Mu¨ller, D. J.; Engel, A.; Carrascosa, J. L.; Velez, M. 1997. EMBO J. 1997, 16, 2547.

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Figure 10. (A) 134 nm × 134 nm tapping-mode AFM image of the MT2 layer adsorbed onto the DTSP-modified gold substrate at pH 7.0. Note the ordered arrangement of the protein structures. (B) FFT image corresponding to the AFM images of the MT2 layer adsorbed onto the DTSP-modified gold substrate. The six spots are marked 1-6 to better understand the further analysis. (C) Cross sections measured on the FFT image of (B) along the main three symmetry directions: from spot 1 to spot 4 (dotted line); from spot 2 to spot 5 (solid line); from spot 3 to spot 6 (dashed line). The main frequency in the reciprocal space, k0, is indicated by the vertical lines. The FFT profiles have been low-pass-filtered to better identify the different peaks. (D) Two-dimensional autocorrelation of the AFM image shown in (A). Six spots hexagonally arranged around the central spot indicate the close-packed structure of the MT2 layer. Surrounding these spots, a dotted structure is visible, indicating a weaker long-range ordering.

Taking into account the amino acid sequence of MT2, we observe that it contains only six lysine groups which can form covalent bonds with the succinimidyl groups present on the gold functionalized surface. This fact contrasts with the considerably higher amount of sites available in the MT2 structure for both chemisorption and unspecific protein adsorption on the gold surface. This difference could explain the smoother and more homogeneous MT2-DTSP-Au monolayer morphology observed. Both the different geometric location of the sites responsible for the protein adsorption and the different nature of the lysine-substrate interaction, stronger than that of the unspecific adsorption, agree with both the different morphological and mechanical properties measured by AFM. However, the most striking morphological property of the MT2 monolayer on DTSP-Au substrates is the evident short-range, as well as the weaker longrange, ordering. This ordering suggests that the protein orientation on the DTSP layer tends to be the same as otherwise steric hindrance and the different interactions between neighbor MT2 molecules would probably hamper the self-ordering of the MT2 layer. From the six lysine groups present in the MT2 primary structure, Lys30 and Lys31 are located40,47 between the two, R and β, hydrophobic domains and probably are the most exposed ones. In contrast, the other four sites are relatively more isolated (47) Zangger, K.; O ¨ z, G.; Otvos, J. D.; Armitage, I. M. Protein Sci. 1999, 8, 2630.

and buried by the rest of the MT2 structure. Thus, it seems likely that the sequence Lys30-Lys31 could be the preferential site for the protein covalent bonding to the functionalized Au surface, leading to a preferential MT2 adsorption geometry. MT2 Directly Adsorbed onto a Mica Substrate. Despite gold surfaces having some advantages for protein attachment and AFM imaging, mica substrates have been extensively used in a great number of AFM studies due to the possibility of being easily cleaved to generate atomically flat surfaces. In our case, the study of the MT2mica system contributes to a better knowledge about the nature of the MT2-mica interactions and its relationship with the properties of the MT2 layer. It is well-known that, originally, the mica surface is negatively charged, but when it is exposed to cations, the surface becomes positively charged. Thus, first we have studied the MT2 adsorption on mica in a buffer solution at pH 7.0. AFM imaging of this sample is shown in Figure 12. The image revealed the formation of a smooth (σ ) 0.3 nm) protein layer. The protein dimensions are h ) 0.40.9 nm and dfwhm ) 3-6 nm. The depth of the hole (2 nm), indicated by the arrow, allows us to conclude that the thickness of the protein deposit corresponds to that of a monolayer. These results indicate that a stronger attractive electrostatic interaction exists between the mica


Langmuir, Vol. 18, No. 15, 2002 5919

indicates that the attractive interaction between the mica surface, which is negatively charged, and the protein is very weak. This different behavior can be explained by taking into account the IEP value of the MT2, which is 4.5. When the deposition takes place under neutral pH conditions, the protein is negatively charged (pH > IEP) and the mica surface is positively charged, leading to an appreciable MT2 deposit. The comparison between the MT2-DTSP-Au and MT2-mica systems reveals some interesting features. Despite the fact that the surface roughness is the same for both systems from the AFM images, we can observe some differences (Table 3). By comparing typical images (Figures 10A and 12), we observe that the MT2 layer on mica seems to be less tightly packed than those in the other systems. In fact, compared to those of the MT2DTSP system, h is similar while dfwhm is appreciably larger. These differences indicate that MT2 lies spread on the mica surface, apparently with a larger contact area between the protein and the substrate. This fact is consistent with previous reports on the strong proteinmica interactions which, sometimes, lead to the protein distortion.17 Conclusions

Figure 11. (A) Height distributions corresponding to the same area of the MT2 layer adsorbed onto a DTSP-modified gold substrate at pH 7.0 imaged at different forces: A/A0 ) 0.95 (solid line) and A/A0 ) 0.90 (dotted line). (B) Surface profiles measured at A/A0 ) 0.95 (top curve) and A/A0 ) 0.90 (bottom curve). An offset has been included between both profiles.

Figure 12. 310 nm × 310 nm tapping-mode AFM image of an MT2 layer adsorbed onto a freshly cleaved mica surface after 30 min of incubation at pH 7.0.

surface (this time positively charged due to the cations present in the buffer) and the protein. We have also performed AFM imaging of MT2 deposited onto mica in deionized water (under these conditions the mica surface remains negatively charged). These images (data not shown) reveal a very low adsorption of protein on mica. Furthermore, the imaged protein appears to be weakly bonded to the substrate since the protein was easily attached to the tip, leading to multiple tip imaging artifacts (data not shown). The fact that the adsorption of the MT2 in a deionized water environment is scarce and weak

In this work we have studied the surface coverage and the morphological properties of MT2 layers immobilized on gold and mica surfaces. To analyze the relationship between the MT2-substrate interaction and these properties, we have deposited the MT2 layers following three different approaches based on unspecific adsorption/ chemisorption (MT2-Au system), covalent binding (MT2DTSP-Au system), and physisorption (MT2-mica system). The MT2 layers formed from chemisorption/ unspecific MT2-substrate interactions are significantly disordered and relatively rough in contrast to those formed from MT2-DTSP-Au, which are smoother and show a close-packed molecular arrangement. Besides, our experiments also suggest different mechanical properties of the protein layers for both systems. In particular, for the MT2Au system the proteins are imaged in the attractive and repulsive force regimes, the proteins being elastically deformed in the latter regime. These different properties are explained on the basis of the different MT2-substrate interactions. Thus, for the MT2-Au, there are a large number of sites available in the MT2 structure for the unspecific adsorption. This scenario involves different MT2 adsorption geometries on the Au surface and, therefore, is consistent with the rougher and disordered layer morphology as well as with the different elastic response of the protein structures of the layer. For the MT2DTSP-Au system, MT2 is covalently bonded to the functionalized substrate. Two of the six lysine rests present in MT2 (Lys30-Lys31) could be directly involved in the immobilization process, because of their particular geometrical location in the protein structure. This fact would explain the close-packed ordering of the MT2-DTSP layer. On the other hand, the MT2-mica system, in which MT2 is physisorbed onto the substrate surface, depicts monolayers which are quite smooth, but the protein lateral dimensions are larger, suggesting a larger proteinsubstrate contact area. Finally, we have found that the MT2 layer morphological and mechanical properties are related to the main MT2substrate interaction governing the MT2 adsorption process. Furthermore, our results obtained on the MT2-

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DTSP-Au system show that covalent binding strategies not only allow the protein-substrate interaction to be controlled but can also lead to the protein layer ordering, which could be very interesting for future technological applications.

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Acknowledgment. This work has been partially supported by MCYT (Spain) Grant Nos. BQU2001-0163 and MAT2000-0375-C02. LA025712C

Mica Surfaces: Relationship

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