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Enzymatic Proteolysis of a Surface-Bound r-Helical Polypeptide Jasper O. Hardesty,‡ Luis Casca˜o-Pereira,† James T. Kellis,† Channing R. Robertson,‡ and Curtis W. Frank*,‡ Biochemistry Department, Genencor International, Palo Alto, California 94304, and Department of Chemical Engineering, Stanford UniVersity, Stanford, California 94305 ReceiVed June 29, 2008. ReVised Manuscript ReceiVed August 25, 2008 In this work, we studied the interactions of enzymes with model substrate surfaces using label-free techniques. Our model system was based on serine proteases (a class of enzymes that digests proteins) and surface-bound polypeptide substrates. While previous studies have focused on bulk media factors such as pH, ionic strength, and surfactants, this study focuses on the role of the surface-bound substrate itself. In particular, we assess how the substrate density of a polypeptide with an R-helical secondary structure influences surface reactivity. An R-helical secondary structure was chosen based on literature indicating that stable R-helices can resist enzymatic digestion. To investigate the protease resistance of a surface-bound R-helix, we designed an R-helical polypeptide (SS-polypeptide, where SS ) disulfide), used it to form films of varying surface coverage and then measured responses of the films to enzymatic exposure. Using quartz-crystal microbalance with dissipation (QCM-D), angle-resolved X-ray photoelectron spectroscopy (AR-XPS), grazing-angle infrared spectroscopy (GAIRS), and other techniques, we characterized the degradation of films to determine how the lateral packing density of the surface-bound SS-polypeptide substrate affected surface proteolysis. Characterization of pure SS-polypeptide films indicated dense packing of helices that maintained their helical structure and were generally oriented normal to the surface. We found that films of pure SS-polypeptide significantly resisted enzymatic digestion, while incorporation of very minor amounts of a diluent in such films resulted in rapid digestion. In part, this may be due to the need for the enzyme to bind several peptides along the peptide substrate within the cleft for digestion to occur. Only SS-polypeptide films that were densely packed and did not permit catalytic access to multiple peptides (e.g., terminal peptides only) were resistant to enzymatic proteolysis.
Introduction In nature, proteases are ubiquitous enzymes found in all organisms and are essential for cell growth and differentiation. Microbial proteases are mostly extracellular and are used to digest proteins for nutritional purposes. Reaction occurs through hydrolyzing peptide bonds in the substrate, referred to as either hydrolysis or proteolysis. Proteases are classified into four groups based on the essential catalytic amino acid residue of their active site: serine proteases (e.g., trypsin, chymotrypsin, subtilisin), cysteine proteases (e.g., papain and bromelain), aspartic proteases (e.g., pepsin and chymosin), and metalloproteases (e.g., thermolysin and neprilysin).1,2 Subtilisin is a member of the serine proteases and serves as the model enzyme in our experiments. It exhibits greater stability and less specificity than other proteases, allowing subtilisin to digest a broad spectrum of proteins.3 Subtilisin from Bacillus serves as an important model system for protein engineering studies,4,5 and there is extensive literature on proteases, with emphasis on different aspects, including microbial selection, fermentation, production and processing,6-8 * To whom correspondence should be addressed. E-mail: curt.frank@ stanford.edu. † Genencor International. ‡ Stanford University. (1) Rao, M.; Tanksale, A.; Ghatge, M.; Deshpande, V. Microbiol. Mol. Biol. ReV. 1998, 62(3), 597–635. (2) Barrett, A.; Rawlings, N.; Woesneer, J. F. The Handbook of Proteolytic Enzymes; Academic Press: London, 1998. (3) Ottesen, M.; Svendsen, I. The subtilisins; Academic Press: New York, 1970; Vol. 19, pp. 199-215. (4) Strausberg, S. L.; Ruan, B.; Fisher, K. E.; Alexander, P. A.; Bryan, P. N. Biochemistry 2005, 44, 3272–3279. (5) Bryan, P. N. Biochim. Biophys. Acta 2000, 1543, 203–222. (6) Aunstrup, K. Proteinases; Academic Press: New York, 1980; pp 50-114. (7) Anwar, A.; Saleemuddin, M. Bioresour. Technol. 1998, 64, 175–183. (8) Kumar, C.; Takagi, H. Biotechnol. AdV. 1999, 17, 561–594.
natural protease functions and sources,9 commercial applications,10 genetic and molecular biology,11,12 and protein engineering.13,14 Studies of subtilisin catalysis and properties have been aided by a number of 3-D structures from X-ray crystallographic data,15,16 and detailed information on its structural characteristics and enzymatic mechanism has been obtained.17-22 As a promising target for protein engineering, subtilisin and its variants have served as models to study altered catalytic behavior and stability, and they continue to attract attention for expanding applications.5,8 In industry, proteases are valued for many uses, and subtilisins are among the most investigated enzymes.3,5,8,11 Proteases are used for applications such as detergent, food, leather, diagnostics, waste management, silver recovery, and pharmaceuticals. Enzyme production in 1982 was $375 million, up to $720 million in 1990, $1 billion in 1994, $1.9 billion in 1996, $2.5 (9) Ward, O. Proteolytic enzymes; Pergamon Press: Oxford, 1985; Vol. 3, pp 789-818. (10) Kalisz, H. AdV. Biochem. Eng./Biotechnol. 1988, 36, 1–65. (11) Outtrup, H.; Boyce, C. Microbial proteinases and biotechnology; Elsevier: London, 1990; pp 227-254. (12) Siezen, R.; de Vos, W.; Leunissen, J.; Dijkstra, B. Protein Eng. 1991, 4, 719–737. (13) Wells, J. A.; Ferrari, E.; Henner, D. J.; Estell, D. A.; Chen, E. Y. Nucleic Acids Res. 1983, 11, 7911–7925. (14) Takagi, H. Int. J. Biochem. 1993, 25, 307–312. (15) Gallagher, T.; Gilliland, G.; Wang, L.; Bryan, P. Structure 1995, 3, 907– 14. (16) Jain, S.; Shinde, U.; Li, Y.; Inouye, M.; Berman, H. J. Mol. BIol. 1998, 284, 137–144. (17) Ballinger, M. D.; Tom, J.; Wells, J. A. Biochemistry 1995, 34, 13312– 13319. (18) Gron, H.; Breddam, K. Biochemistry 1992, 31, 8967–8971. (19) Gron, H.; Meldal, M.; Breddam, K. Biochemistry 1992, 31, 6011–6018. (20) Rheinnecker, M.; Baker, G.; Eder, J.; Fersht, A. R. Biochemistry 1993, 32, 1199–1203. (21) Bott, R. R.; Chan, G.; Domingo, B.; Ganshaw, G.; Hsia, C. Y.; Knapp, M.; Murray, C. J. Biochemistry 2003, 42(36), 10545–10553. (22) Fersht, A. Enzyme Structure and Mechanism, 2nd ed.; W. H. Freeman and Co.: New York, 1985.
10.1021/la8020386 CCC: $40.75 2008 American Chemical Society Published on Web 11/14/2008
Proteolysis of a Surface-Bound SS-Polypeptide
billion in 2001, and approximately $2.8 billion in 2002.23,24 Bacterial alkaline proteases accounted for approximately 35% of the total industrial enzyme market.25 Market projections show that, in the U.S. enzyme market alone, demand will grow by 6.9% annually through 2010, with both proteases and carbohydrases expected to continue to dominate market growth and demand.26 Subtilisin BPN′ is a wild-type protease from the Bacillus species amyloliquefaciens and serves as a very well-studied model for comparison to other subtilisins.5 The enzyme used in this study was BLS subtilisin from Bacillus lentus.27 BLS differs from BPN′ at 103 of a possible 269 residues, including deletion of six residues in three segments. BLS and BPN′ share a conserved tertiary fold, and with residue numbering by homology with BPN′ the catalytic triad is formed by Asp 32, His 64, and Ser 221. The BLS subtilisin was chosen for this work, as the enzyme itself had been well-studied, and it has been used commercially as a component of detergent formulations for degradation of proteinaceous material at surfaces. The activity of proteases on substrates, whether in solution or at an interface, can be related to a variety of factors. Commonly recognized variables include the specificity of the enzyme toward particular amino acid sequences, environmental conditions (i.e., pH and ionic strength), the stability of polypeptide segments, and the accessibility of those segments. Regarding accessibility, binding of a segment of six to eight amino acid residues at the active site cleft of a protease is required for enzymatic proteolysis of a polypeptide chain.28,29 Thus, for globular proteins in their native conformation, initial digestion by proteases is limited by the amount of flexible substrate chains that are accessible to the enzyme. In general, the interior of the protein is inaccessible and thus is not susceptible to proteolysis until the protein completely denatures, with loss of tertiary structure. Regarding stability, digestion is a consequence of the affinity of the enzyme for the substrate (to remain bound) and the stability of the protein under the given conditions. For all polypeptides, enhanced structural stability is imparted for any given segment that is part of a secondary or tertiary structure. Thus, segments within stable secondary structures (i.e., helices, sheets, bundled helices, sheet barrels, etc.) may retard or prevent degradation by a protease. Two examples of these protease-resistant structures are found in the persistence of prions30 and development of peptide antibiotics.31 Unlike the well-understood catalytic mechanisms of subtilisin in bulk solution, subtilisin’s proteolysis of a protein bound to a surface is not well-understood.17-22 Biocatalysis at surfaces is a growing field, with applications that include surface coatings to prevent fouling, contact lens treatment, conversion of organic material for biofuels, treatment of textiles, detergents, processing of foods, bioremediation of pollutants, synthesis of pharmaceuticals, biodefense systems, sensors, green chemistry, and other (23) Chenel, J.; Tyagi, R.; Surampalli, R. Water Sci. Technol. 2008, 57(5), 639–645. (24) Godfrey, T.; West, S. Introduction to industrial enzymology, 2nd ed.; Macmillan Press: London, 1996; pp 1-8. (25) Cherry, J. R.; Fidantsef, A. L. Curr. Opin. Biotechnol. 2003, 14, 438– 443. (26) Enzymes Industry Forecasts to 2010 & 2015; Freedonia Group, Inc.: Cleveland, OH, 2006. (27) Kuhn, P.; Knapp, M.; Soltis, S. M.; Ganshaw, G.; Thoene, M.; Bott, R. Biochemistry 1998, 37(39), 13446–13452. (28) Fontana, A.; Fassina, G.; Vita, C.; Dalzoppo, D.; Zamai, M.; Zambonin, M. Biochemistry 1986, 25(8), 1847–1851. (29) de Laureto, P.; Scaramella, E.; De Fillipis, V.; Bruix, M.; Rico, M.; Fontana, A. Protein Sci. 1997, 6, 860–872. (30) Vorberg, I.; Chan, K.; Priola, S. J. Virol. 2001, 75(21), 10024–10032. (31) Yamaguchi, H.; Kodama, H.; Osada, S.; Kato, F.; Jelokhani-Niaraki, M.; Kondo, M. Biosci., Biotechnol., Biochem. 2003, 67(10), 2269–2272.
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Figure 1. Graphic of BLS subtilisin enzyme approaching surface-bound protein substrate. Red ) enzyme active site (Ser, His, Asp). (Subtilisin from 1GCI.pdb, and protein film from 1BM0.pdb.)
uses.32-35 Interest in surface catalysis of proteases has increased with the recognition that applications often involve surface-bound substrates, and that enzymatic activity at surfaces does not necessarily mirror the same trends as in bulk solution.36-38 Surface processes between enzymes and substrates at an interface are dominated by noncovalent interactions. For an enzyme approaching a surface-bound substrate, as shown in Figure 1, film properties and features can affect the ability of the enzyme to adsorb to the surface and to engage the substrate with an orientation that allows reaction and surface diffusion.39 Biocatalysis can thus be affected by a variety of physical and chemical factors. For a uniform polypeptide film, the surface accessibility, packing density, and secondary structure represent major physical issues, while surface charge and amino acid sequence represent major chemical considerations. Biocatalysis at surfaces differs from solution conditions in that additional processes are introduced by the presence of an interface.37,40-43 In solution, there is bulk diffusion, binding, and reaction. At an interface, bulk diffusion of the enzyme to the substrate remains as the initial step, but it differs from solution conditions in that the surface-bound substrate is not diffusing. Binding at an interface differs in that a surface-bound substrate requires adsorption with binding. Surfaces may create constraints on enzyme adsorption, such as hydrophobic effects, surface charges, and substrate orientation, any of which can retard or prevent enzyme adsorption and binding to a surface-bound (32) Ahuja, S. K.; Ferreira, G. M.; Moreira, A. R. Crit. ReV. Biotechnol. 2004, 24(23), 125–154. (33) Bommarius, A. S.; Riebel, B. R. Biocatalysis; Wiley-VCH Verlag GmbH & Co.: Weinheim, Germany, 2004. (34) LeJeune, K. E.; Russell, A. J. Biotechnol. Bioeng. 1999, 62(6), 659–665. (35) Xu, P.; Uyama, H.; Whitten, J. E.; Kobayashi, S.; Kaplan, D. L. J. Am. Chem. Soc. 2005, 127(33), 11745–11753. (36) Brode, P.; Erwin, C.; Rauch, D.; Barnett, B.; Armpriester, J.; Wang, E.; Rubingh, D. Biochemistry 1996, 35, 3162–3169. (37) Kim, J. H.; Roy, S.; Kellis, J. T.; Poulose, A. J.; Gast, A. P.; Robertson, C. R. Langmuir 2002, 18(16), 6312–6318. (38) Rubingh, D. Engineering proteases with improVed properties for detergents; IBC Biomedical Library Series: 1996; pp 98-123. (39) Malmsten, M. Biopolymers at Interfaces; Marcel Dekker: New York, 1998; Vol. 75. (40) Gaspers, P. B.; Robertson, C. R.; Gast, A. P. Langmuir 1994, 10(8), 2699–2704. (41) Roy, S.; Thomas, J. M.; Holmes, E. A.; Kellis, J. T.; Poulose, A. J.; Robertson, C. R.; Gast, A. P. Anal. Chem. 2005, 77(24), 8146–8150. (42) Esker, A. R.; Brode, P. F.; Rubingh, D. N.; Rauch, D. S.; Yu, H.; Gast, A. P.; Robertson, C. R.; Trigiante, G. Langmuir 2000, 16(5), 2198–2206. (43) Lee, H. J.; Wark, A. W.; Goodrich, T. T.; Fang, S.; Corn, R. M. Langmuir 2005, 21(19), 4050–4057.
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substrate. Similarly, the reaction step can be altered by the same surface effects as for adsorption, resulting in either an enhancement or retardation of reaction. Finally, surface diffusion is uniquely a surface process not present for free substrate in solution. Surface diffusion involves the transport of enzyme molecules across the interface. When the rate of enzyme adsorption is much greater than the rate of enzyme desorption, “hopping” of the enzyme from one location to another on the surface-bound substrate is minimal.40,41 Under such conditions, the transport of enzyme molecules is restricted to two dimensions, instead of the three dimensions available without a solid surface. Reduction in dimensionality means that an enzyme molecule in bulk solution can access a surface-bound substrate only by diffusing in the bulk to a random location on the surface, adsorbing to the surface, and then diffusing along the surface until it reaches its target.44 After adsorption, the target encounter rate is governed by twodimensional enzyme surface diffusion rather than threedimensional enzyme bulk diffusion, or “hopping”. For surface diffusion to occur, the enzyme must adsorb to the surface with an affinity strong enough to keep it on the bound substrate but weak enough to allow mobility.40 The overall reactivity of an enzyme on a surface-bound substrate is thus determined by a balance of the adsorption, surface reactivity, and surface diffusion processes. For subtilisin at surfaces, a high sensitivity of overall reactivity has been observed for a variety of environmental determinants such as pH, surfactant, and ionic strength of bulk media. In addition, a single point mutation in subtilisin, distant from the active site, can lead to substantial changes in its catalytic properties.37 Previous studies of enzymatic processes at surfaces have been performed using fluorescent tags on surface-bound substrates.37,40,41,45,46 However, use of labels can alter reactivity and adsorption due to steric, hydrophobic, and charge perturbations.47-51 Given the sensitivity of subtilisin, as exhibited by changes in its activity at a surface, it is preferable to avoid fluorescent (or any other) labels; thus, techniques employed in this study are all label-free. Our experimental objective was to understand the reactivity of a subtilisin on a surface-bound polypeptide. Instead of investigating environmental effects, this study considers the surface-bound substrate and its role in regulating surface reactivity. How might surface packing density or secondary structure of a protein substrate affect enzymatic proteolysis of that substrate when bound at a surface? Several studies have pointed to protease-resistant proteins or domains that include R-helical regions.52-54 In this work, we study the reactivity of a single subtilisin type on an engineered surface-bound polypeptide having an R-helical secondary structure. We demonstrate (44) Adam, G.; Delbruck, M. Structural Chemistry and Molecular Biology; Freeman: San Francisco, CA, 1968; p 198. (45) Konash, A.; Cooney, M. J.; Liaw, B. Y.; Jameson, D. M. J. Mater. Chem. 2006, 16, 4107–4109. (46) Moore, C. M.; Akers, N. L.; Hill, A. D.; Johnson, Z. C.; Minteer, S. D. Biomacromolecules 2004, 5, 1241–1247. (47) Brynda, E.; Drobnik, J.; Vacik, J.; Kalal, J. J. Biomed. Mater. Res. 1978, 12(1), 55–65. (48) Drees, B. L.; Rye, H. S.; Glazer, A. N.; Nelson, H. C. M. J. Biol. Chem. 1996, 271(50), 32168–32173. (49) Gajraj, A.; Ofoli, R. Y. Langmuir 2000, 16(21), 8085–8094. (50) Muller, R. H.; Ruhl, D.; Luck, M.; Paulke, B. R. Pharm. Res. 1997, 14(1), 18–24. (51) Sacco, D.; Dellacherie, E.; Prouchayret, F. J. Protein Chem. 1994, 13(1), 1–8. (52) Puntheeranurak, T.; Leetacheewa, S.; Katzenmeier, G.; Krittanai, C.; Panyim, S.; Angsuthanasombat, C. J. Biochem. Mol. Biol. 2001, 34(4), 293–298. (53) Rall, S. C.; Ye, P.; Bu, G. J.; Wardell, M. R. J. Biol. Chem. 1998, 273(37), 24152–24157. (54) VanMelderen, L.; Thi, M. H. D.; Lecchi, P.; Gottesman, S.; Couturier, M.; Maurizi, M. R. J. Biol. Chem. 1996, 271(44), 27730–27738.
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that the R-helix substrate forms well-characterized films and observe that the BLS subtilisin can digest such films having various surface packing densities. Determining conditions by which enzymatic degradation occurs, or does not occur, can provide information about interactions and processes that govern surface reactions by enzymes on surface-bound substrates.
Experimental Section Materials. Reagents. Sodium carbonate and sodium sulfate were purchased from Sigma-Aldrich Chemical. Thiolated methoxy poly(ethylene glycol) (PEG-thiol) was purchased from Polypure AS (Oslo, Norway) and consisted of six poly(ethylene oxide) units with a methoxy at one terminus and a thiol at the other terminus. The designed R-helical peptide (SS-polypeptide, with SS ) disulfide) was constructed by solid phase peptide synthesis (SPPS) based on Fmoc-protection/deprotection chemistry (at the Protein and Nucleic Acid Facility, Beckman Center, Stanford University, CA). The SSpolypeptide sequence is lipoic acid-NA(EAAAR)6A, verified via HPLC and mass spectrometry. Buffers. All buffers were prepared in Milli-Q water (18.2 MΩ). Buffer used for all experiments contained 2 mM sodium carbonate and 15 mM sodium sulfate (pH 9.5). Enzyme. The enzyme used in this study was BLS subtilisin from Bacillus lentus (PDB file 1GCI).27 Enzyme stock solutions were diluted to a concentration of 10 µg/mL in buffer for all experiments, unless noted otherwise. Substrate Surface Preparations. Substrates used for electrical impedance spectroscopy (EIS) experiments consisted of glass slides with a 2-10 nm chromium or titanium adhesion layer under vapordeposited 100 nm gold films. Gold surfaces were cleaned with Milli-Q water and ethanol, and then dried under a stream of nitrogen prior to use. Substrates were prepared on these surfaces by exposure of the gold surfaces to solutions of each respective compound. Due to the sensitivity of the EIS equipment, measurements could not be made of film formation starting with bare gold; thus, these substrates were prepared prior to measurements of pure films. Surfaces used for QCM-D experiments consisted of gold-coated quartz crystals (Q-Sense AB, Gothenburg, Sweden). Immediately prior to use, each crystal was treated with oxygen plasma (GaLa Instrumente GmbH, plasma instrument, Germany) at ∼80 W for 5 min, rinsed with Milli-Q water and ethanol, and then dried under a stream of nitrogen. Substrate films were generated in the QCM-D chamber by exchanging solutions, which were monitored via changes in frequency and dissipation. Methods. Circular Dichroism (CD) Spectroscopy. CD spectroscopy of polypeptides is based on the differential absorption of rightand left-handed circularly polarized light by amide groups in the peptide backbone. In addition to the intrinsic chirality of the amino acids, secondary structure elements have a defined twist; hence, the CD spectra of these structures are distinguishable from each other. Existence of characteristic CD spectra for helix, sheet, or random structures make this the spectroscopic method of choice to monitor conformations of proteins and polypeptides.55 We used CD spectroscopy to monitor subtilisin digestion of the synthetic R-helical peptide (SS-polypeptide, where SS ) disulfide) via measurement of the helical fraction in solution. Measurements were performed using an Aviv model 215 spectrophotometer with a 450 W xenon arc lamp, a double-fused silica prism monochromator (range 165-1200 nm), a MgF2 polarizer, a high-sensitivity end-on photomultiplier tube with preamplifier, and a ThermoNeslab M25 temperature control system. Operation and setup of the spectrophotometer incorporated a PC containing Aviv CDS control software. Scans were conducted at 25 ((0.1) °C in wavelength mode (range 270-190 nm) at 1.0 nm steps, with an averaging time of 5.0 s/datum point, bandwidth/slits at 1.0 nm, averaged over 2 scans. Cuvettes were rectangular, of FUV-grade quartz, with a light path length of 0.1 cm. All samples were measured in carbonate buffer. For analysis, CD scans were corrected by background substitution based on buffer (55) Woody, R. W. Methods Enzymol. 2004, 380, 242–285.
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scans. Spectral analysis for secondary structure composition was performed by CDpro, a suite of CD spectral analysis tools that includes three different algorithms: SELCON 3, CDSSTR, and CONTIN-II.56-58 Quartz Crystal Microbalance with Dissipation (QCM-D). The quartz crystal microbalance is an ultrasensitive mass sensor consisting of a thin piezoelectric quartz disk having electrodes deposited on each side. When connected to an oscillating current, the quartz crystal will oscillate at resonant frequencies that are sensitive to the crystal mass; thus, as material adsorbs to the surface of the crystal surface, there is a decrease in these frequencies. A change of adsorbed mass (∆m) on the crystal results in a frequency shift (∆f) of the shear oscillations. So long as the adsorbed material is elastically (rigidly) attached to the crystal, the mass and frequency changes are related by the Sauerbrey equation:59
∆f ) (-2)
f02 AõqFq
(∆m) ) (-Cf)(∆m)
(1)
where ∆f is the measured change of resonant frequency (Hz), ∆m is the amount of adsorbed mass (g), with ∆m g 0, µq and Fq are the shear modulus and density of quartz, respectively, A is the electrode area, f0 is the resonant frequency, and Cf is the integrated crystal sensitivity factor. For measurements of adsorbed materials that do not behave as elastic masses on the QCM surface, the Sauerbrey equation does not apply, and analysis of the film mass is more complicated. However, the films studied herein exhibit elastic behavior consistent with use of the Sauerbrey relationship, and therefore, it is used throughout this work. To study in situ film formation and changes under conditions of exposure to enzyme, we used a QCM-D apparatus (Q-Sense D300, Q-Sense AB, Gothenburg, Sweden). AT-cut crystals (Q-Sense) 14 mm in diameter with 50 nm gold thermally evaporated over a 10 nm titanium adhesion layer were used for all studies. Each crystal was scanned at various frequencies to determine the primary harmonic frequency as well as the third, fifth, and seventh harmonic frequencies. The temperature of the Q-Sense cell was 25.1 ( 0.5 °C, controlled by a Peltier element. X-ray Photoelectron Spectroscopy (XPS). XPS quantitatively measures the elemental composition of a material at a surface and is sensitive to the upper ∼10 nm of the material surface. The distance an electron travels in a material affects the probability that an electron will be emitted, so collecting electrons at an acute angle to the surface reduces the depth from which a signal can be collected. Angle-resolved (AR)-XPS is based on varying the take-off angle for depth profiling of a film:60
IA ) K
(
)
dz ∫0∞ CA(z) exp λA-z sin θ
(2)
where IA is the intensity of the peak for element A, CA(z) is the concentration of A at depth z, λA is the attenuation length for the electron of the appropriate kinetic energy, θ is the take-off angle (to sample surface), and K is the normalization parameter. A general determination of elemental distribution in a sample can be made by determining its “atomic %” at different angles, such that composition and thickness can be analyzed by graphically plotting the appropriate ratios of those values for elements of interest. This technique uses reduced area fractions instead of areas, which eliminates all angle-dependent instrument factors.60 We used AR-XPS of pure SS-polypeptide films on gold to determine if there were exposed disulfides present at the upper layer of the film. Analysis was conducted on a SSI S-Probe Monochro(56) Provencher, S. W.; Gloeckner, J. Biochemistry 1981, 20(1), 33–37. (57) Sreerama, N. Anal. Biochem. 2000, 287(2), 252–260. (58) Johnson, W. C. Proteins: Struct., Funct., Genet. 1999, 35(3), 307–312. (59) Marx, K. A. Biomacromolecules 2003, 4(5), 1099–1120. (60) Spruytte, S.; Coldren, C.; Harris, J.; Pantelidis, D.; Lee, H.-J.; Bravman, J.; Kelly, M. J. Vac. Sci. Technol., A 2000, 19(2), 603–608.
matized XPS spectrometer, using Al KR radiation (1486 eV), a quartz monochromator, a concentric hemispherical analyzer, and a multichannel detector. Pressure in the measurement chamber during analysis was approximately 1 × 10-9 Torr. Take-off angles of 90° and 30° from the surface were employed to obtain survey spectra from 0 to 1000 at 1.0 eV resolution and high-resolution spectra from 160 to 170 at 0.1 eV resolution. Each survey spectrum was the average of 5 scans, and high-resolution surveys were averaged across 10 scans. Grazing Angle Infrared Spectroscopy (GAIRS). GAIRS identifies chemical bonds in thin films as well as secondary structures and molecular orientation of polypeptides. By reflecting a coherent light beam off of a reflective surface covered by a thin film, a grazing angle provides an improved signal-to-noise ratio by allowing the beam to (1) pass across a greater surface area of the film and (2) interact with the film at an aspect that effectively increases the light path through the film. At incident angles greater than ∼80°, reflection from a surface results in constructive interference of the light vector in the incident plane (Ep) while causing destructive interference of the electrical vector parallel to the incident plane (Es).61 Thus, GAIRS measures only the p-polarized component of reflected light from a surface. With respect to a thin film at a surface, the only active vibrations observed are those in the direction normal to the surface plane. Secondary structure and orientation of adsorbed films on gold substrates were confirmed by GAIRS. FTIR spectra were collected on a Perkin-Elmer Spectrum 2000 FTIR spectrometer (Waltham, MA) at room temperature, using a Specac monolayer grazing-angle reflection accessory (Woodstock, GA). The incident angle was set at 83° from the surface normal. Spectra were collected at 3400-1400 cm-1 in 0.5 cm-1 increments, with 4 cm-1 resolution, accumulating 300 interferograms per spectrum. Dry nitrogen was used to purge the sample chamber for ∼20 min to reduce water and carbon dioxide. Reference gold surfaces were used for background scans, and sample scans were normalized with the background. The orientation of peptide films on gold surfaces was determined via the ratios of the amide-I/amide-II absorbance peak areas (centered at ∼1669/1545 cm-1, respectively) in each spectrum using62
I1 [(3cos2 γ - 1)(3cos2 θ1 - 1) + 2] ) 1.5 I2 [(3cos2 γ - 1)(3cos2 θ2 - 1) + 2]
(3)
where Ii is the observed absorbance (peak area), i is 1 or 2 corresponding to amide-I or amide-II, respectively, γ is the tilt angle of the helix axis (from surface normal), θi is the angle between the transition moment and helix axis (39° for θ1 and 75° for θ2),63 and the prefactor of 1.5 is a proportionality constant used for polypeptides.62 Electrochemical Impedance Spectroscopy (EIS). In an EIS interfacial measurement, an AC potential is imposed on the specimen (film) over a range of frequencies, and the phase and amplitude of the resulting current are measured at each frequency.64 Impedance measurement does not require special reagents and is amenable to label-free operation. For a DC circuit, Ohm’s Law states V ) IR, with electrical potential (V), current (I), and resistance (R). In a DC circuit, the only element that impedes electron flow is a resistor. In contrast, with an AC circuit, the frequency is nonzero, and impedance (Z) is the AC equivalent of resistance:
V ) IZ
(5)
The complex AC impedance is defined as the sum of its real (Z′) and imaginary (Z′′) components, from which the absolute magnitude and phase angle (θ) of the impedance can be determined:
|Z| ) √Z ′ 2 + Z ′′ 2 Z′′ tan θ ) Z′
(6) (7)
Monolayers of SS-polypeptide and PEG-thiol were generated in the EIS setup by placing a clean gold-coated slide (electrode) on a flat
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Figure 2. Circular dichroism spectra of SS-polypeptide in buffer before exposure to enzyme (dashed line) and after exposure to enzyme (dotted line). All spectra are shown with buffer background subtracted.
metal support (∼100 × 50 × 7 mm3), clamping it in place with a second metal piece having two rows of eight sample holes containing Teflon tubes (ID ) 3.6 mm) inserted to provide individual sample cells, and incubating specified wells with solutions of each material. Solutions that consisted of either 0.01 mM SS-polypeptide or 0.1 mM PEG-thiol in buffer were pipetted into specified sample wells and allowed to incubate for 20 h. After the incubation period, each sample cell was rinsed five times with buffer, refilled with buffer, and equilibrated at room temperature prior to collecting spectra. Electrical impedance spectra were collected using a custom built 16-channel impedance analyzer (provided by Australian Membrane Biotechnology Research Institute, Sydney, Australia) configured for running one sensor block of 16 wells and controlled by an IBM compatible PC using custom software.65,66 Scans were performed over frequencies between 0.1 and 1000 Hz over a period of ∼30 s, with a DC bias of 300 mV. A three-terminal AC excitation of 50 mV was applied between a gold (Au) measurement, silver (Ag) reference, and platinum (Pt) counter electrodes. All 16 independent impedance measurements were conducted simultaneously and repeatedly over several hours to verify film stability. Final spectra were averaged from five scans taken in rapid succession for each sample well. Analysis was performed using LEVMW software for CNLS data fitting (by J. Ross Macdonald and Solartron Group Limited, 1999-2005, joint copyright). The system was modeled with an equivalent circuit consisting of a resistor (buffer/film interface) in series with both a resistor and capacitor (actually, a constant phase element, CPE) in parallel to represent each film.
Results This work was designed to investigate the effect of surface packing density of an R-helical secondary structure on its enzymatic digestion. Specifically, we studied the proteolysis of the polypeptide by a single BLS subtilisin. Based on research by others52-54 regarding the protease resistance of R-helices, we elected to construct an R-helical polypeptide substrate. Design of the polypeptide was based on literature regarding R-helical (61) Ulman, A. Fourier Transform Infrared Spectroscopy in Colloid and Interface Science; Scheuing, D. R., Ed.; American Chemical Society: Washington, D.C., 1991; Vol. 447, pp 144-159. (62) Samulski, E. T.; Enriquez, E. P. Mater. Res. Soc. Symp. Proc. 1992, 255, 423–434. (63) Tsuboi, M. J. Polym. Sci. 1962, 59(167), 139. (64) Macdonald, J. R. Impedance Spectroscopy: Emphasizing Solid Materials and Systems; Wiley-Interscience: New York, 1987. (65) Yin, P.; Burns, C. J.; Osman, D. J.; Cornell, B. A. Biosens. Bioelectron. 2003, 18, 389–397. (66) Krishna, G.; Schulte, J.; Cornell, B. A.; Pace, R. J.; Osman, P. D. Langmuir 2003, 19, 2294–2305.
secondary structure stability.67,68 To study surface proteolysis, we designed a helix that was soluble and stable in buffer and that was capable of forming self-assembled monolayers (SAMs) on gold surfaces via covalent bonds. The designed R-helical polypeptide was rich in alanine and included an acid residue and an amine residue at the i+4 positions along the sequence to provide solubility and enhance stability. A length of 33 amino acids was chosen to increase helical content, as ends are generally less stable and lengths over 50 residues tend to aggregate and can result in β-sheet structures.68 This length also provided good yields from the synthesis process. The N-terminus of the helical polypeptide was connected to lipoic acid (5-(1,2-dithiolane-3yl)pentanoic acid), which contains a disulfide in a ring structure (1,2 dithiolane) to provide a binding moiety for SAM formation of the helix on gold surfaces. Hence, we identify this polypeptide as the “SS-polypeptide”. The SS-polypeptide was modeled by numerous structure prediction programs, and all results showed R-helical secondary structure. A structural model of the SSpolypeptide, with the attached disulfide, is available as Figure S-1 in the Supporting Information. Prior to studying the SS-polypeptide at surfaces, it was important to characterize its structure in buffer as well as its response when exposed to enzyme. Circular dichroism scans (Figure 2) of the SS-polypeptide in buffer yielded two minima at λ ) 208 and 222 nm, characteristic of an R-helical secondary structure.69 After exposure of the same solution to enzyme for ∼10 min, there was a marked change in the spectrum, with loss of both minima noted above and a shift toward a curve characteristic of a random coil structure. Analysis indicated that the polypeptide in buffer was roughly 95% helical, 1% β-sheet, and 4% turns and random coil; on exposure to enzyme, the secondary structure changed to roughly 23% helical, 4% β-sheet, and 73% turns and random coil. To characterize the SS-polypeptide films formed, a variety of techniques were employed. For GAIRS measurements, selectivity allows for the determination of molecular orientation, if present. Films composed of the SS-polypeptide have two primary peaks of interest: the amide-I and amide-II peaks. The amide-I peak is primarily generated by the carbonyl stretch along the backbone of the peptide chain, while the amide-II peak is primarily a (67) Huyghuesdespointes, B. M. P.; Scholtz, J. M.; Baldwin, R. L. Protein Sci. 1993, 2(1), 80–85. (68) Scholtz, J. M.; Qian, H.; Robbins, V. H.; Baldwin, R. L. Biochemistry 1993, 32(37), 9668–9676. (69) Fasman, G. D. Circular Dichroism and the Conformational Analysis of Macromolecules; Plenum: New York, 1996; p 738.
Proteolysis of a Surface-Bound SS-Polypeptide
Langmuir, Vol. 24, No. 24, 2008 13949
Figure 3. GAIRS and SS-polypeptide orientation at the surface. Schematic of amide-I and amide-II bond vibrations on SS-polypeptide (left) and processed GAIRS data of SS-polypeptide monolayer on a gold surface (right).
combination of the carbon-amide bond stretch along the peptide backbone plus the bending of the amide N-H bond. Secondary structure is typically identified by evaluating the wavenumber at which the amide-I peak is located. For peptides that form an R-helix, the amide-I peak is frequently cited as occurring at ∼1650-1658 cm-1.70 However, with GAIRS of films on metallic surfaces, there are optical surface effects that result in a spectral shift for some peaks.70,71 Specifically, the amide-I peak has been found to exhibit a positive shift of ∼13-17 cm-1, which is consistent with our measurements at 1668-1670 cm-1, while the amide-II peak has been found to experience a small negative shift of ∼3-5 cm-1, consistent with the peak at ∼1545 cm-1 (compared to common assignment at 1550 cm-1), as shown in Figure 3. The peak at ∼1516 cm-1 is attributed to the NH3+ of arginine.72 The amide-I dipole moment is aligned nearly parallel to the helix axis, while the amide-II dipole moment is more perpendicular to the helix axis, so orientation of the R-helices at surfaces was determined by evaluating the ratio of both amide peaks according to eq 3 (see Methods subsection). Evaluating the spectra of SS-polypeptide SAMs on gold showed that the helices were consistently tilted at 57.2 ( 1.5° from the surface (Figure 3). To ascertain the condition of the uppermost surface for our SS-polypeptide SAMs, we performed AR-XPS to generate an elemental depth profile. By obtaining and comparing XPS spectra at 90° and 30°, we sought to determine if a partial layer of disulfides was present at the exposed (outer) surface of the film. If disulfides were present at the outer surface, we would expect a sulfur peak to be visible in the 30° spectra. However, the 30° broad elemental survey spectra did not show any discernible sulfur peak. Moreover, when we collected a high-sensitivity spectrum at 164.4 eV (sulfur 2p peak) to verify the elemental scan, again no sulfur peak was detected. The 90° spectra showed elemental peaks identical to the 30° spectra, except for a sulfur peak at ∼164 eV and that the gold peak was larger. These peak differences were due to the shorter escape path for photoelectrons from the gold surface in the 90° spectra. We used QCM-D to monitor SS-polypeptide film formation on gold-coated quartz crystals. We found that films were densely packed (∼1.1 × 1014 helices/cm2) and rigid, with a mass of (70) Enander, K.; Aili, D.; Baltzer, L.; Lundstrom, I.; Liedberg, B. Langmuir 2005, 21, 2480–2487. (71) Allara, D. L.; Baca, A.; Pryde, C. A. Macromolecules 1978, 11(6), 1215– 1220. (72) Bieri, M.; Burgi, T. Langmuir 2006, 22, 8379–8386.
∼625 ng/cm2. Following exposure of the film to the BLS subtilisin, a very slow degradation of the SS-polypeptide film occurred. As seen in Figure 4, the initial film mass is stable to rinses, but when exposed to enzyme there is gradual mass loss until the film is reduced to ∼310 ng/cm2 after 210 min of exposure. To generate less densely packed films of SS-polypeptide, we selected a shortchain thiol (PEG-thiol, MW ) 356.4, see Figure 5) to act as a diluent in mixed binary SAMs. Films of pure PEG-thiol were also measured by QCM-D to monitor film formation and response when exposed to enzyme, as a control. Films formed rapidly and were stable to repeated rinses, as shown in Figure 6. Film mass stabilized at ∼270 ng/cm2, indicating that complete films were densely packed (∼4.6 × 1014 PEG/cm2) and stable. After exposure of the PEG-thiol film to the BLS subtilisin, we observed a minor shift of
