Langmuir 1997, 13, 6485-6490
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Immobilization of Protein Molecules onto Homogeneous and Mixed Carboxylate-Terminated Self-Assembled Monolayers Nikin Patel,* Martyn C. Davies, Mark Hartshorne, Richard J. Heaton, Clive J. Roberts, Saul J. B. Tendler, and Philip M. Williams Laboratory of Biophysics and Surface Analysis, School of Pharmaceutical Sciences, The University of Nottingham, Nottingham NG7 2RD, U.K. Received August 18, 1997X The attachment of biomolecules, in particular proteins, onto solid supports is fundamental in the development of advanced biosensors, bioreactors, affinity chromatographic separation materials, and many diagnostic techniques. In addition, the effective investigation of biomolecular structure and function with scanning probe microscopy often requires a strong attachment of the biomolecule to a substrate. Here, we investigate the binding of the protein catalase to gold surfaces modified by self-assembled monolayers (SAMs). The chemical and physical adsorption of the protein molecules onto SAMs of 3-mercaptopropanoic acid (3-MPA), 11-mercaptoundecanoic acid (11-MUA), and a mixture of the two acid thiols (mixed) was investigated by utilizing tapping mode atomic force microscopy, scanning tunneling microscopy, surface plasmon resonance (SPR), static secondary ion mass spectrometry, and X-ray photoelectron spectroscopy. The surface concentration of catalase adsorbed on the SAMs decreased in the following order: mixed > 11-MUA > 3-MPA. Utilizing the terminal carboxylic acid functionalities, catalase was immobilized with a water-soluble carbodiimide and N-hydroxysuccinimide (NHS). Immobilization resulted in increased coverage of the protein. SPR studies on silver surfaces modified by these SAMs indicate that immobilization of carbodiimide and NHS decreased in the same order, namely mixed > 11-MUA > 3-MPA.
1. Introduction In recent years, scanning probe microscopy (SPM) has proven to be an invaluable tool in investigating the structure and function of biomolecules.1-3 The scanning tunneling microscope (STM) and atomic force microscope (AFM) have both demonstrated with high resolution, often reaching the molecular level, their ability to study a wide range of biological specimens such as whole cells, proteins, and nucleic acids.4-8 However, for such fragile samples, the effective use of SPM is often limited by interactions between tip and sample.9 In addition, if investigations are performed in a liquid environment, biomolecules need to be attached to the substrate surface to prevent dissolution or resuspension. AFM has also been utilized to measure forces of interactions between biomolecules, a procedure which requires firm attachment of biomolecules to both substrate and tip.10,11 Different methods have been investigated to increase the adhesion of individual molecules to a surface, including * To whom correspondence should be addressed. Telephone: +44 (0) 115 9515063. Fax: +44 (0) 115 9515110. E-mail: paxnp@ unix.ccc.nottingham.ac.uk. X Abstract published in Advance ACS Abstracts, November 1, 1997. (1) Yang, J.; Tamm, L. K.; Somlyo, A. P.; Shao, Z. J. Microsc. 1992, 171 (3), 183. (2) Rabe, J. Ultamicroscopy 1992, 41, 42. (3) Lindsay, S. M. Scanning Tunneling Microscopy and Spectroscopy. Theory, Techniques and Applications, 1st ed.; VCH Publishers: New York, 1993; pp 335-408. (4) Kasas, S.; Gotzos, V.; Celio, M. R. Biophys. J. 1993, 64, 539. (5) Parker, M.-C.; Patel, N.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B.; Williams, P. M. Protein Sci. 1996, 5, 2329. (6) Patel, N.; Davies, M. C.; Lomas, M.; Roberts, C. J.; Tendler, S. J. B.; Williams, P. M. J. Phys. Chem. B 1997, 101, 5138. (7) Guckenberger, R.; Heim, M.; Cevc, G.; Knapp, H. F.; Wiegra¨be, W.; Hillebrand, A. Science 1994, 266, 1538. (8) Hansma, H. G.; Hoh, J. H. Annu. Rev. Biophys. Biomol. Struct. 1994, 23, 115. (9) Weisenhorn, A. L.; Hansma, P. K.; Albrecht, T. R.; Quate, C. F. Appl. Phys. Lett. 1989, 54, 2651. (10) Lee, G. U.; Kidwell, D. A.; Colton, R. J. Langmuir 1994, 10, 354. (11) Allen, S.; Chen, X.; Davies, J.; Davies, M. C.; Dawkes, A. C.; Edwards, J. C.; Roberts, C. J.; Sefton, J.; Tendler, S. J. B.; Williams, P. M. Biochemistry 1997, 36, 7457.
S0743-7463(97)00933-5 CCC: $14.00
modification of the substrate surface and/or modification of the biomolecule. For proteins, attachment can be either physically or chemically mediated, with both methods having advantages. Physical adsorption processes involve hydrophobic or electrostatic interactions between the protein and the solid phase. Such processes are generally experimentally simple and often retain the activity of the protein.12 However, the adsorption can be reversible, with protein being removed by certain buffers or detergents or replaced by other proteins in solution. In contrast, chemical immobilization involves the covalent bonding of the protein to the solid phase. This method is experimentally more difficult and often exposes the protein to a harsher environment. However, the resultant irreversible binding which can be produced with high levels of surface coverage makes this approach more popular, although in some cases covalent binding can alter the conformational structure and active center of the protein, causing a reduction in activity.12 Our previous studies have shown that a significant proportion of immobilized catalase can still remain catalytically active (at least 2031%).5,13 The functionalization of metal and glass surfaces with self-assembled monolayers (SAMs)14,15 is one method used for the physical and chemical attachment of proteins.16,17 The SAM can be tailored to present a hydrophobic or charged surface, hence enabling hydrophobic or electrostatic interactions with proteins.18,19 For covalent im(12) Chibata, I. Immobilized EnzymessResearch and Development, 1st ed.; John Wiley and Sons: New York, 1978; Chapter 2. (13) Parker, M.-C.; Davies, M. C.; Tendler, S. J. B. J. Phys. Chem. 1995, 99, 16155. (14) Ulman, A. An Introduction to Ultrathin Organic Films: From Langmuir-Blodgett to Self-Assembly, 1st ed.; Academic Press: Boston, 1991; pp 236-304. (15) Nuzzo, R. G.; Allara, D. A. J. Am. Chem. Soc. 1983, 105, 4481. (16) Prime, K.; Whitesides, G. M. Science 1991, 252, 1164. (17) Leggett, G. J.; Roberts, C. J.; Williams, P. M.; Davies, M. C.; Jackson, D. E.; Tendler, S. J. B. Langmuir 1993, 9, 2356. (18) Silin, V.; Weetall, H.; Vanderah, D. J. J. Colloid Interface Sci. 1997, 185, 94-103. (19) Lo´pez, G. P.; Biebuyck, H. A.; Ha¨rter, R.; Kumar, A.; Whitesides, G. M. J. Am. Chem. Soc. 1993, 115, 10774.
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Figure 1. Strategy for immobilizing protein onto carboxylateterminated SAMs on gold. Addition of NHS and the watersoluble carbodiimide EDC to the SAMs results in the formation of an NHS ester. Reaction of protein side-chain lysine residues with the ester results in the formation of an amide bond.
mobilization of proteins onto SAMs, the general procedure involves the formation of either a disulfide or an amide. One method for amide bond formation utilizes the production or termination of SAMs with an N-hydroxysuccinimide (NHS) ester.5,13,20-22 Immobilization occurs by displacement of the NHS group by lysine residues of the protein. Frey and Corn have shown that polylysine can be immobilized onto an 11-mercaptoundecanoic acid (11-MUA) SAM by activation of the terminal carboxylate group with an N-hydroxysulfosuccinimide ester intermediate.20 Successful formation of the ester intermediate is reliant on the accessibility of the terminal carboxylate groups; steric packing of these acid groups can limit the rate of intermediate formation, with full conversion of accessible acid groups occurring only after several repeated reaction cycles. In these studies, we have generated SAMs from short- and long-chain carboxylic acid-terminating alkanethiols to assess the effect of chain length on the reactivity of SAMs toward protein immobilization (Figure 1). In addition, the reactivity of a mixed SAM formed by the coadsorption of the two alkanethiols has been studied. The extent of protein attached physically and chemically has been determined at the molecular level by using tapping mode AFM and STM. The study is supported by complementary surface analytical techniques, including static secondary ion mass spectrometry (SSIMS), X-ray photoelectron spectroscopy (XPS), and surface plasmon resonance (SPR). 2. Experimental Section Materials. 11-Mercaptoundecanoic acid was synthesized using the method detailed in ref 23. Absolute ethanol (Haymans, (20) Frey, B. L.; Corn, R. M. Anal. Chem. 1996, 68, 3137-3193. (21) Wagner, P.; Kernen, P.; Hegner, M.; Ungewickell, E.; Semenza, G. FEBS Lett. 1994, 356, 267. (22) Wagner, P.; Hegner, M.; Kernen, P.; Zaugg, F.; Semenza, G. Biophys. J. 1996, 70, 2052.
Patel et al. Essex, U.K.), 3-mercaptopropanoic acid (Aldrich, Dorset, U.K.), 1-decanethiol (Aldrich), N-hydroxysuccinimide (Avocado, Lancashire, U.K.), 1-ethyl-3-[3-(dimethylamino)propyl]carbodiimide hydrochloride (Sigma, Dorset, U.K.), and catalase (EC 1.11.1.6) from bovine liver (Sigma) were all used as received. Preparation of Substrates and Monolayer Formation. Gold films (∼50-100 nm thick) were prepared by slow vapor deposition24 of gold (99.99%, Birmingham Metals, U.K.) onto heated, freshly cleaved mica (Agar Scientific, Essex, U.K.), followed by annealing at 425 °C for 24 h. Gold films were cut into sections, washed with ethanol, and dried under a stream of high-purity argon before use. These samples were immersed into 10 mM ethanolic solutions of 3-MPA, 11-MUA, or a mixed solution of 3-MPA and 11-MUA (10:1) for 24 h before being rinsed in ethanol followed by distilled deionized water. Monolayers were prepared in glass scintillation vials that had been cleaned with chromic acid for 1 h and rinsed exhaustively with distilled deionized water and ethanol. For SPR experiments, electron beam-evaporated silver films (40 nm thick) on glass slides (Johnson & Johnson Clinical Diagnostics, Chalfont St. Giles, Buckinghamshire, U.K.) were used to support 3-MPA, 11-MUA, mixed, and decanethiol SAMs. The SAMs were prepared as described above. Protein Adsorption and Immobilization. For protein immobilization, the carboxylic acid-terminated SAMs were immersed into an aqueous solution of 75 mM EDC and 15 mM NHS for between 10 s and 30 min. The resultant NHS ester monolayers were reacted for 30 min in a solution of catalase (10 µg/mL) in sodium phosphate buffer (10 mM, pH 8.0). A relatively high pH was used to deprotonate the lysine residues on the surface of catalase to enable reaction with the NHS ester. After removal of the SAM from protein solution, the surface was rinsed exhaustively with distilled deionized water and dried under a stream of argon. For protein adsorption studies, SAMs on gold were immersed into the catalase solution for 30 min, rinsed with distilled deionized water, and dried under a stream of argon. For SPR experiments, the EDC/NHS solution was flowed over the SAMs on silver for 4 min, followed by a buffer wash. Subsequently, the interaction of catalase solution (10 µg/mL) with the immobilized EDC/NHS was studied. Instrumentation. (a) SSIMS. Spectra were obtained using a VG Ionex SIMSLAB 3B instrument equipped with a differentially pumped EX05 ion gun and a 12-12 M quadrupole mass spectrometer. Argon atoms at 2 keV were used as the primary source, with an equivalent current of 1 nA. The total dose per sample was approximately 5 × 1012 atoms/cm2, which is within the regime of the static SIMS experiment. (b) XPS. XPS analysis was carried out on a VG Scientific ESCALAB Mk II electron spectrometer employing Mg KR X-rays (hν ) 1253.6 eV) with an electron take-off angle of 35°. The X-ray gun was operated at 12 kV and 20 mA. Survey spectra (0-1000 eV binding energy) were obtained, followed by highresolution spectra of the C 1s, O 1s, N 1s, and S 1s regions. Acquisition and analysis of data were handled using a VGS 5000S data system. (c) Contact Angle Measurements. Contact angles were measured in air on static drops using a goniometer (Ealing Electro-Optics plc, Watford, U.K.). A 2 µL drop of sodium phosphate buffer (10 mM, pH 8.0) was applied to the surface, and contact angle measurements were performed on opposite edges of the drop. Measurements were repeated on at least four drops. (d) AFM Imaging. AFM images of protein samples were obtained in air with a NanoScope IIIa (Digital Instruments, Santa Barbara, CA) in the tapping imaging mode.25 Images were acquired at a scan rate of 2 Hz with a silicon cantilever unit (Nanoprobe, cantilever length 125 µm and resonant frequency 307-375 kHz). A drive amplitude of between 20 and 50 mV was used. (23) Bain, C. D.; Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides, G. M.; Nuzzo, R. J. Am. Chem. Soc. 1989, 111, 321. (24) DeRose, J. A.; Thundat, T.; Nagahara, L. A.; Lindsay, S. M. Surf. Sci. 1991, 256, 102. (25) Zhong, Q.; Innes, D.; Kjoller, K.; Elings, V. B. Surf. Sci. Lett. 1993, 290, 688.
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bound to Ag (II). In comparison, a positive ion spectrum
[AgS(CH2)10COOH + H]+
Figure 2. Positive ion SIMS spectra of SAMs on silver. (A, B) Mixed SAM. (C) 3-MPA SAM. (D) 11-MUA SAM. (e) STM Imaging. STM images were obtained in a variablehumidity chamber with platinum-iridium tips using a VG STM 2000 system (VG Microtech, Uckfield, U.K.). Typical experimental conditions comprised a current of 40 pA and a tip bias voltage of +1.13 V. The relative humidity (RH) of each sample atmosphere was controlled using saturated sodium chloride solutions for hydrating the sample. The humidity of the chamber was routinely checked using a humidity meter (Vaisala HMI 31, R.S. Components, Corby, Northhants, U.K.) housed inside the chamber. (f) SPR Analysis. The SPR instrument (Johnson & Johnson Clinical Diagnostics), which has been described in detail elsewhere,26 monitors interactions at or above a silver-coated glass sensor surface. A laser light source at 780 nm wavelength is used. Sample solutions of 1 mL were introduced through a loop valve system to the SPR instrument at a rate of 4 µL/s by means of a syringe pump. The sample cell allowed two channels to be analyzed simultaneously. This cell was composed of a thermoregulated copper block which was equilibrated to a temperature of 34 °C prior to commencement of the experiment.
3. Results and Discussion Characterization of SAMs. SAMs formed on both gold and silver from solutions of 3-MPA, 11-MUA, and a mixture of 3-MPA/11-MUA (10:1 ratio) were characterized using both XPS and SSIMS. XPS studies on the two homogeneous SAMs detected compositions of carbon, oxygen, and sulfur on the surface close to the theoretical value of these constituents in the SAM (data not shown). SSIMS studies also revealed the successful formation of the 3-MPA27 and 11-MUA SAMs. Molecular ions from each of these SAMs on gold and silver were detected. In addition, the mixed monolayer system was analyzed to ascertain the adsorption of both thiols from solution. Figure 2A shows a positive ion SSIMS spectrum from the mixed monolayer system on silver in the region m/z 200400. The spectrum is dominated by three clusters of peaks, which can be identified as being generated by the silver surface or the mixed SAM. The peaks at m/z 214, 216, and 218 are due to Ag2+. These three peaks are generated from the combination of the two Ag isotopes (m/z 107 and 109). The peaks at m/z 247, 249, and 251 are due to Ag2SH+. The cluster of peaks from m/z 319 to 327 are generated from the coadsorption of 3-MPA and 11-MUA to silver; Figure 2B shows an enlargement of this region. The peaks at m/z 319, 321, and 323 are due to 3-MPA bound to Ag2 (I). Again, three peaks are produced due to
[Ag2SCH2CH2COOH]+
I
the presence of two Ag atoms. The smaller peaks at m/z 325 and 327 correspond to the molecular ion of 11-MUA (26) Green, R. J.; Davies, J.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B. Biomaterials 1997, 18, 405. (27) Leggett, G. J.; Davies, M. C.; Jackson, D. E.; Tendler, S. J. B. J. Phys. Chem. 1993, 97, 5349.
II
in this region (m/z 315-330) generated by a 3-MPA SAM on silver (Figure 2C), shows the presence of species I only. Similarly, a positive ion spectrum (m/z 315-330) generated by an 11-MUA SAM on silver (Figure 2D) shows the presence of species II with an absence of I. In addition, a lower intensity peak occurs at m/z 323 which is probably due to [AgSdCH(CH2)9COOH]+. Contact angle measurements were also performed on the SAMs formed on gold in order to determine the relative hydrophilicity of the surfaces. A low contact angle measurement indicates a relatively hydrophilic surface. All the SAMs were shown to have relatively hydrophilic surfaces, which is not surprising since the terminal carboxylate group is highly polar. The contact angles decreased in the order mixed (41° ( 1°) > 11-MUA (29° ( 4°) > 3-MPA (22° ( 2°). The higher hydrophobicity of the 11-MUA compared with that of the 3-MPA SAM may result from disruptions in the ordered SAM by underlying defaults of the substrate. Areas of increased surface roughness may expose the hydrophobic methylene groups of the alkanethiol. We expect this effect to be more pronounced with the longer chain of 11-MUA. The contact angle from the mixed 3-MPA/11-MUA SAM is significantly greater compared with those of either of the pure components. The mixture of the two thiols is likely to lead to increased exposure of the methylene groups of the 11-MUA. Physical Adsorption of Protein on SAMs. Utilizing both STM and AFM, the adsorption of the protein catalase to the carboxylate-terminated SAMs on gold was studied at the molecular level. Measuring the extent of physical adsorption is important if we are to determine how effective these SAMs are toward covalent immobilization of protein. Figure 3 shows that differing amounts of protein adsorption to the 3-MPA, 11-MUA, and mixed SAMs occur after 30 min exposure to a 10 µg/mL catalase solution. In Figure 3A, the topography image (left) reveals that individual catalase molecules on the 3-MPA SAM surface are observed with an approximate diameter of 12 nm and a height of 6 nm, which correlate reasonably with the dimensions obtained from X-ray crystallography (10.5 nm × 10.5 nm × 6 nm).28 These images were obtained using tapping mode AFM as opposed to contact AFM to avoid the strong probe-sample forces which may displace the physisorbed protein.8 In conventional contact mode AFM, imaging involves dragging the probe tip across the surface. The dragging motion combined with adhesive forces between the tip and the surface can lead to substantial damage and displacement of the sample. In tapping mode, the tip is vibrated at or close to its resonant frequency and, hence, is alternatively placed in and out of contact with the surface so as to avoid dragging the tip across the surface. In addition to the topography images, the corresponding phase detection images are also shown (Figure 3A, right). Contrast in the phase detection imaging is generated from differences in viscoelastic or adhesive properties between protein and non-protein material.29 The catalase molecules appear to be sparsely distributed, with the underlying gold surface visible. The surface coverage of catalase was 38%, calculated by thresholding of the topography image using the Genesis (28) Murthy, M. R. N.; Reid, T. J., III.; Sicignano, A.; Tanaka, N.; Rossmann, M. G. J. Mol. Biol. 1981, 152, 465. (29) Tamayo, A.; Garcia, R. Langmuir 1996, 12, 4430.
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Figure 4. STM image of physisorbed catalase on the mixed SAM on gold. Compared with AFM (Figure 3C), which shows a monolayer of catalase adsorbed to the mixed SAM, STM shows very little protein material on this surface. The higher probesample forces involved with STM imaging may result in sweeping of these physisorbed molecules, thus resulting in the apparent absence of the protein monolayer.
Figure 3. AFM images showing the physisorption of catalase to (A) 3-MPA, (B) 11-MUA, and (C) mixed SAM on gold. Individual catalase molecules are observed with an approximate diameter of 12 nm and a height of 6 nm. Both topography (lefthand images) and the corresponding phase detection (righthand images) images are shown. The white lines observed in the phase image of (C) are not related to the surface topography; instead, they result from instrumentation noise. Scan sizes: 500 nm × 500 nm.
II System, which has been developed in our laboratory.30,31 A similar effect is seen with the 11-MUA SAM (Figure 3B), although the level of protein adsorption is greater (72% surface coverage). Again, the gold substrate surface can be identified between protein molecules, in both the topography and the phase images. A monolayer of catalase is adsorbed onto the mixed SAM (Figure 3C), with the underlying gold surface no longer visible (98% surface coverage). This high level of lateral coverage results in the lack of contrast in the phase imaging. Adsorption on the pure 3-MPA and 11-MUA SAMs probably results from electrostatic interactions between the terminal carboxylate group and protein.18,19 Electrostatic adsorption of proteins onto charged SAMs is dependent on the protein’s isoelectric point (pI) and solution pH;32 here, catalase has a pI ) 5.5, and the solution pH is 8.0. Under these conditions, both the surface of the SAMs and the protein are negatively charged, hence resulting in a sparse physisorption of catalase. However, (30) Williams, P. M.; Davies, M. C.; Jackson, D. E.; Roberts, C. J.; Tendler, S. J. B. J. Vac. Sci. Technol. B 1994, 12 (3), 1456. (31) Williams, P. M.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B. Analyst 1997, 293, 101. (32) Jordan, C. E.; Corn, R. M. Anal. Chem. 1997, 69, 1449.
some protein adsorption nevertheless occurs on these surfaces. Although catalase at pH 8.0 has a net negative charge, adsorption may occur at positive charges residing on the surface.18 In addition, hydrophobic interactions may occur in areas where the SAM is disrupted, such as step edges and edges of gold islands. Greater amounts of adsorption occur on the mixed SAM with full coverage of the surface. This probably results from the exposure of the methylene groups of 11-MUA in the mixed SAM allowing hydrophobic interactions with the protein in addition to the electrostatic interactions via the terminal group. A similar monolayer of adsorbed protein can be observed with a decanethiol SAM which has a highly hydrophobic surface (water contact angle 97° ( 4°) (data not shown). As a control, gold surfaces modified with 3-MPA, 11MUA, or mixed SAMs were immersed in buffer solution (pH 8.0) for 30 min. AFM imaging (data not shown) failed to show any protein-like stuctures on the surface. The monolayer of physisorbed catalase (from pH 8.0 solution) onto the mixed SAM was also analyzed by using STM. Imaging was carried out in a high-humidity environment to increase the samples conductivity.6,7 This is important for the successful imaging of biological samples, which often have poor conductivity. Despite catalase forming a monolayer on the mixed SAM, as confirmed by AFM, STM imaging (Figure 4) fails to shows this. Instead, globular features with diameters of approximately 6 nm are observed, primarily at the gold step edges. The higher probe-sample forces involved with STM imaging may result in sweeping of these physisorbed molecules, thus resulting in the apparent absence of the protein monolayer. Covalent Immobilization of Protein onto SAMs. The immobilization of catalase onto gold involves the formation of an NHS ester with the carboxylate-terminated SAMs using the water-soluble carbodiimide EDC (Figure 1). Side-chain amino groups of lysine residues on the protein surface displace the terminal NHS groups, resulting in covalent immobilization of the protein. SPR was used to monitor the amounts of EDC/NHS and subsequent catalase immobilization onto the 3-MPA, 11MUA, and mixed SAMs. The change in millidegree angle (mDA) is related to the amount of material that is lost or gained on the surface.33 Figure 5 shows a SPR trace of the consecutive adsorption/immobilization profiles of EDC/ NHS followed by catalase onto the mixed SAM on silver. Injection of the EDC/NHS solution over the SAM surface (33) De Bruijin, H. E.; Kooyman, R. P. H.; Grewe, J. Appl. Opt. 1993, 32, 2426.
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Figure 5. SPR signal on injection of EDC/NHS followed by catalase over the mixed SAM. (A) and (B) represent the extent of EDC/NHS and catalase binding, respectively.
Figure 6. SPR angle shifts for immobilization of EDC/NHS and catalase onto SAMs on silver.
results in a sharp, increased mDA due to the change in refractive index at the solid/liquid interface. After approximately 4 min, buffer is washed over the surface, resulting in a decline in mDA shift due to the removal of unbound material. The resultant increased mDA compared to the initial level (A) relates to the amount of EDC/ NHS covalently bound to the SAM. A gradual increase in mDa often occurs after 200 s. This appears not to have an effect on the value of A, indicating the gradual increase results from the adsorption of EDC/NHS with the surface, which is removed following the buffer wash after 4 min. Injection of catalase results in another increased mDA, which fails to decrease following a buffer wash at approximately 1100 s. This second increase (B) correlates to the amount of catalase attached to the surface. Figure 6 shows the extent to which EDC/NHS followed by catalase react with 3-MPA, 11-MUA, and mixed SAMs. A small amount of EDC/NHS (mDA shift < 25) reacts with the 3-MPA and 11-MUA surfaces. The ordered arrangement of the terminal carboxylate groups of the homogeneous SAMs may sterically hinder the reaction and formation of the NHS ester. In comparison, a significantly greater amount of EDC/NHS (mDA shift 152 ( 7) reacts with the mixed SAM surface. Two possible reasons can account for this substantial increase. First, terminal carboxylate groups of the mixed SAM are not as ordered as the homogeneous SAMs. Disorder may result either from phase separation or from full integration of the two thiol components. With increased disorder, the terminal carboxylate groups are more accessible for the immobilization of EDC/NHS. Second, the increased hydrophobicity of the mixed SAM could promote hydrophobic interactions between the exposed methylene groups and EDC/NHS. However, Figure 6 shows that only a small amount of EDC/NHS (mDA shift 33 ( 10) interacts with a methylterminated thiol, decanethiol, the surface of which is highly hydrophobic. This indicates that the substantial increase
Figure 7. AFM images showing catalase immobilization onto (A) 3-MPA, (B) 11-MUA, and (C) mixed SAM on gold. Both topography (left) and the corresponding phase detection (right) images are shown. Scan sizes: 1 µm × 1 µm.
in mDA shift for the mixed SAM surface is due to an increase in immobilization of EDC/NHS and is not a result of a hydrophobic phenomenon. Following EDC/NHS treatment, the amount of catalase which attaches to the SAMs decreased in the order mixed > 11-MUA > 3-MPA, which correlates to the immobilization of EDC/NHS onto the SAMs. Catalase immobilization onto the mixed SAM surface produces a mDA shift 3 times greater than that produced by 3-MPA. This substantial increase is due primarily to increased protein immobilization as opposed to hydrophobic interactions. Significantly lower protein adsorption occurs on the highly hydrophobic decanethiol. These results suggest that the reactivity of the SAMs toward protein immobilization decreases in the order mixed > 11-MUA > 3-MPA. In order to relate these SPR data to the extent of surface coverage of catalase, AFM was utilized. Due to the relatively high surface roughness of the silver SPR slides, AFM imaging at molecular resolution was not possible following SPR experiments. Instead, SAMs were formed on gold films and immersed in the EDC/NHS solution for varying time lengths, with final immersion into a catalase solution (10 µg/mL) for 30 min. The immersion time of the SAMs in the EDC/NHS is an important factor which determines the extent of protein immobilization. Figure 7 shows varying surface coverage of catalase on all three carboxylate-terminated SAMs following 4 min immersion in EDC/NHS. Both topography (left) and phase detection (right) images are shown. A submonolayer of catalase is
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relatively stronger covalent linkage between protein and substrate prevents this. The same tip was used in the imaging of both Figures 4 and 8. Repeated experiments confirmed the differences shown in the figures, indicating the need for a strong covalent linkage between sample and substrate to prevent displacement from probe-sample forces. Densely packed areas of catalase immobilized onto the mixed SAM, as can be seen in Figure 7C, have also been observed with STM, although individual molecules within these areas could not be clearly differentiated. 4. Conclusion Figure 8. STM image of catalase covalently immobilized onto the mixed SAM on gold. The covalent binding of protein to the substrate results in the successful imaging of individual molecules. The molecules have an observed diameter of approximately 10 nm and a height of 6 nm.
attached to the 3-MPA surface (Figure 7A), with small gaps observed between protein molecules. The surface coverage was measured at 38%, which matches that of physisorbed catalase onto the 3-MPA SAM (Figure 3A). SPR data (Figure 6) display a very small mDa shift (10 ( 5 mDA) for EDC/NHS immobilization onto the 3-MPA SAM after 4 min. This indicates that the small amount of EDC/NHS immobilization after 4 min has no significant effect on surface coverage. Using AFM, we have observed that incubation of this SAM for longer periods of time in EDC/NHS results in increased protein immobilization until a full monolayer is attained after approximately 10 min incubation. Figure 7B shows that a monolayer of catalase is immobilized to the 11-MUA surface after 4 min immersion in EDC/NHS. This increased surface coverage (97%) compared to that of the 3-MPA SAM correlates with the SPR data (Figure 6), which show an increase in mDa shift of approximately 82 mDA. We believe this increase in coverage results from the small increase in immobilization of EDC/NHS to the 11-MUA SAM. Figure 7C shows that full protein coverage (100%) of catalase occurs on the mixed SAM after a 4 min immersion into EDC/NHS. In addition, clusters of larger features which have a greater height compared with the surrounding catalase monolayer are observed. The clusters vary in size and shape and are uniformly distributed, covering a large percentage of the surface. These clusters indicate dense packing of catalase molecules, possibly relating to phase separation of the two-component thiols. The presence of densely packed catalase correlates with the high mDA shift (303 ( 40 mDA) resulting from catalase immobilization onto the mixed SAM (Figure 6). STM imaging at 72% RH of the covalently immobilized catalase on the mixed SAMs yielded similar results to those produced by AFM. Figure 8 displays individual molecules (diameter approximately 10 nm) within a monolayer. In comparison to the physisorbed catalase (Figure 4), sweeping of protein by the probe tip was not evident with the covalently immobilized catalase. The
We have demonstrated the successful immobilization of the protein catalase onto carboxylate-terminated SAMs utilizing EDC and NHS. The reactivity of the homogeneous and mixed SAMs toward protein immobilization varied depending on the accessibility of the terminal carboxylate group. SPR has shown that the immobilization of EDC/NHS to the SAMs decreases in the order mixed > 11-MUA > 3-MPA. The same trend is observed for the subsequent catalase immobilization. Disorder created by the coadsorption of both 3-MPA and 11-MUA resulted in the mixed SAM displaying the greatest reactivity toward immobilization. The level of protein adsorption and immobilization onto these surfaces was effectively monitored to the molecular level by utilizing both AFM and STM. The strong covalent linkage between protein and substrate was demonstrated to be essential for the successful imaging of protein by STM, a technique in which high probe-sample forces can lead to displacement of fragile biomolecules. The multitechnique approach used here is imperative for the understanding of protein adsorption and immobilization. Information gained from such processes is important in the development of advanced biosensors, bioreactors, affinity chromotographic separation materials, and many diagnostic techniques, all of which require biomolecular attachment to a solid substrate. The design of novel protein assemblies, which can be incorporated into these systems, requires the patterning and attachment of protein into defined areas. In addition, advanced biosensors and diagnostic techniques which probe the functionality of proteins may require immobilization to both tip and substrate. Acknowledgment. The authors gratefully acknowledge Mike Allen of Digital Instruments and Nicola Forsyth of LOT Oriel for their excellent support and the Royal Society for the funding of a gold evaporator. The authors thank Dr. S. R. Chhabra, Z. Affas, and D. J. Evans for helpful discussions on chemical synthesis and immobilization processes. M.C.D., S.J.B.T., and C.J.R. thank Eli Lilly for providing a studentship for N.P., and Polymer Laboratories and the BBSRC for funding postdoctoral fellowships to M.H. and R.J.H. S.J.B.T. is a Nuffield Foundation Science Research Fellow. LA970933H
