pubs.acs.org/Langmuir © 2009 American Chemical Society
In Situ IR Spectroscopic Studies of the Avidin-Biotin Bioconjugation Reaction on CdS Particle Films Aidan G. Young,*,†,§ A. James McQuillan,† and David P. Green‡ †
Department of Chemistry and ‡Department of Anatomy and Structural Biology, University of Otago, P.O. Box 56, Dunedin, New Zealand. § Present address: Industrial Research Ltd, P.O. Box 31-310, Lower Hutt 5040, New Zealand Received January 27, 2009. Revised Manuscript Received March 4, 2009
Avidin-biotin bioconjugation reactions have been carried out on CdS nanoparticle films in H2O and D2O and investigated using in situ ATR-IR spectroscopic techniques. The experimental procedure involved the sequential adsorption of mercaptoacetic acid, the protein avidin, and the subsequent binding of the ligand biotin. The IR spectra of the solution-phase species mercaptoacetic acid, avidin, and biotin, at pH = 7.2 were generally found to be similar in both H2O and D2O, with some minor peak shifts due to solvation changes. The IR spectra of the adsorbed species suggested that avidin may have undergone a conformational change upon adsorption to the CdS surface. In general, adsorptioninduced conformational changes for avidin are likely, but to our knowledge have not been previously reported. The conformation of adsorbed avidin appeared to change again upon the binding of biotin, with the spectral data suggesting partial reversion to its native solution conformation.
Introduction Colloidal semiconductor nanocrystals which have sizedependent optical properties, often called quantum dots (QDs), have received much interest in the last 10 years due to their potential in applications including biological imaging,1,2 lasers,3,4 light emitting devices,5,6 and solar cells.7-9 For biological imaging, in particular, QDs have significant advantages over conventional fluorophores, including greater resistance to chemical and photochemical degradation and a narrow tunable emission wavelength.10 In contrast, conventional fluorophores (typically genetically encoded fluorescent proteins or chemically synthesized fluorescent dyes) often suffer from susceptibility to photobleaching and have broad emission spectra which frequently tail off toward longer wavelengths. In 1998, the first major applications of QDs as biological labels were demonstrated by two different groups. Chan and Nie reported the uptake of transferrin-QD conjugates by live HeLa cells.11 Bruchez et al. reported the labeling of mouse fibroblasts, where red QDs (4 nm core) were modified to *Corresponding author. Fax: +64 4 931 3142; Tel: +64 4 931 3144; E-mail:
[email protected]. (1) Kim, S.; Lim, Y. T.; Soltesz, E. G.; De Grand, A. M.; Lee, J.; Nakayama, A.; Parker, J. A.; Mihaljevic, T.; Laurence, R. G.; Dor, D. M.; Cohn, L. H.; Bawendi, M. G.; Frangioni, J. V. Nat. Biotechnol. 2004, 22, 93–97. (2) Lidke, D. S.; Nagy, P.; Heintzmann, R.; Arndt-Jovin, D. J.; Post, J. N.; Grecco, H. E.; Jares-Erijman, E. A.; Jovin, T. M. Nat. Biotechnol. 2004, 22, 198– 203. (3) Artemyev, M. V.; Woggon, U.; Wannemacher, R.; Jaschinski, H.; Langbein, W. Nano Lett. 2001, 1, 309–314. (4) Klimov, V. I.; Mikhailovsky, A. A.; Xu, S.; Malko, A.; Hollingsworth, J. A.; Leatherdale, C. A.; Eisler, H. J.; Bawendiz, M. G. Science 2000, 290, 314–317. (5) Colvin, V. L.; Schlamp, M. C.; Alivisatos, A. P. Nature (London) 1994, 370, 354–357. (6) Coe, S.; Woo, W.-K.; Bawendi, M.; Bulovic, V. Nature (London) 2002, 420, 800–803. (7) Brown, P.; Kamat, P. V. J. Am. Chem. Soc. 2008, 130, 8890–8891. (8) Kongkanand, A.; Tvrdy, K.; Takechi, K.; Kuno, M.; Kamat, P. V. J. Am. Chem. Soc. 2008, 130, 4007–4015. (9) Weiss, E. A.; Porter, V. J.; Chiechi, R. C.; Geyer, S. M.; Bell, D. C.; Bawendi, M. G.; Whitesides, G. M. J. Am. Chem. Soc. 2008, 130, 83–92. (10) Mattoussi, H.; Medintz, I. L.; Clapp, A. R.; Goldman, E. R.; Jaiswal, J. K.; Simon, S. M.; Mauro, J. M. JALA 2004, 9, 28–32. (11) Chan, W. C. W.; Nie, S. Science 1998, 281, 2016–2018.
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selectively stain cytoskeletal filaments and green QDs (2 nm core) were linked to recognition molecules selective for the cell nucleus.12 Bruchez et al. utilized a bioconjugation reaction where biotin-labeled QDs were coupled to streptavidin-labeled biomolecules, and such coupling reactions have since become widely used in biological imaging studies. Bioconjugation reactions hold much interest, as they provide a method for coupling two biomolecules with a robust chemical linkage. Furthermore, the binding of ligands by proteins is an important biochemical cell process and therefore also of general interest to biochemists. The most widely used and well-known bioconjugation reaction is between avidin and biotin, partly due to widely available reagents that do not require significant chemical modification prior to use. Bioconjugation reactions are frequently used in the development of detection platforms,13,14 with diagnostic platforms for biomolecule detection often utilizing avidin/streptavidin-biotin chemistry, usually at the solid-solution interface.15 Typically, this involves immobilization of one component of the bioconjugate pair on a surface so that the other may be captured and detected subsequently. The structures of avidin and biotin are shown in Figure 1, with the amino acid side chains of avidin omitted for clarity. Avidin is a 68 kDa tetrameric glycoprotein with a reported size of ∼5 nm18 and is composed of four identical subunits, each of which can bind biotin with one of the highest binding constants (12) Bruchez, M.Jr.; Moronne, M.; Gin, P.; Weiss, S.; Alivisatos, A. P. Science 1998, 281, 2013–2016. (13) Medintz, I. L.; Berti, L.; Pons, T.; Grimes, A. F.; English, D. S.; Alessandrini, A.; Facci, P.; Mattoussi, H. Nano Lett. 2007, 7, 1741–1748. (14) Helfrich, M. R.; El-Kouedi, M.; Etherton, M. R.; Keating, C. D. Langmuir 2005, 21, 8478–8486. (15) Perez-Luna, V. H.; O’Brien, M. J.; Opperman, K. A.; Hampton, P. D.; Lopez, G. P.; Klumb, L. A.; Stayton, P. S. J. Am. Chem. Soc. 1999, 121, 6469–6478. (16) Pugliese, L.; Malcovati, M.; Coda, A.; Bolognesi, M. J. Mol. Biol. 1994, 235, 42–46. (17) Weber, P. C.; Ohlendorf, D. H.; Wendoloski, J. J.; Salemme, F. R. Science 1989, 243, 85–88. (18) Misawa, N.; Yamamura, S.; Kim, Y.-H.; Tero, R.; Nonogaki, Y.; Urisu, T. Chem. Phys. Lett. 2006, 419, 86–90.
Published on Web 04/08/2009
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Figure 1. (a) The asymmetric unit of incompletely deglycosylated egg-white avidin (protein data bank (PDB) ID: 1AVE).16 A single N-acetylglucosamine residue is evident as the ball and stick features. Amino acid side chains have been omitted for clarity. (b) The structure of biotin. The binding of biotin by avidin is reported to involve predominantly the ureido portion (RNHCONHR0 ) of biotin.17
(Kb ∼ 1015 L mol-1) known.19 Crystallographic studies of the avidin-biotin complex20,21 have indicated that biotin binding proceeds via hydrogen bonding of the ureido group (RNHCONHR0 ) protons to avidin amino acid side chains (one asparagine and one threonine residue). In addition, the carbonyl oxygen of biotin is hydrogen-bonded to three different avidin protons (two from one tyrosine and one from a serine residue).20,21 The biotin binding site of avidin also contains numerous tryptophan and phenylalanine residues that do not participate in hydrogen bonding with biotin.20,21 Avidin-biotin bioconjugation reactions have most commonly been studied at solid-solution interfaces by allowing adsorption of biotin to a surface with the subsequent introduction of avidin.12,22-24 The carboxylate tail of biotin is typically tailored by functional group conversion to allow adsorption to the interface of interest. Avidin, which is much larger than biotin, then proceeds to bind to a small fraction of the available surfaceimmobilized biotin molecules. This strategy provides a large number of surface-bound biotin and avidin molecules, and high signal-to-noise ratio spectroscopic data of each immobilized component can generally be obtained. While this allows the bioconjugate pairing to be monitored in situ, it provides little information on the specific chemical interactions and changes that occur when biotin binds to avidin, because only a small fraction of the surface-immobilized biotin is bound by avidin. Vibrational spectroscopy is a valuable tool for studying protein chemistry, because it is one of few techniques that allow protein secondary structure to be probed in a nondestructive manner.25,26 Fourier transform infrared (FT-IR) spectroscopic studies of avidin and the avidin-biotin complex in anhydrous conditions have been reported by Swamy et al.27 Spectral deconvolution of the amide I peak was used to investigate changes in secondary (19) Green, N. M. Methods Enzymol. 1990, 184, 51–67. (20) Pugliese, L.; Coda, A.; Malcovati, M.; Bolognesi, M. J. Mol. Biol. 1993, 231, 698–710. (21) Livnah, O.; Bayer, E. A.; Wilchek, M.; Sussman, J. L. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 5076–5080. (22) Ye, L.; Pelton, R.; Brook, M. A. Langmuir 2007, 23, 5630–5637. (23) Yam, C.-M.; Pradier, C.-M.; Salmain, M.; Fischer-Durand, N.; Jaouen, G. J. Colloid Interface Sci. 2002, 245, 204–207. (24) Pradier, C. M.; Salmain, M.; Liu, Z.; Methivier, C. Surf. Interface Anal. 2002, 34, 67–71. (25) Haris, P. I.; Chapman, D. Trends Biochem. Sci. 1992, 17, 328–333. (26) Chittur, K. K. Biomaterials 1998, 19, 357–369. (27) Swamy, M. J.; Heimburg, T.; Marsh, D. Biophys. J. 1996, 71, 840–847.
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structure with respect to both thermal stability and biotin binding. Swamy et al.27 reported that analysis of the amide I peak indicated a secondary structure composed of approximately 66% β-sheet and extended structures with the remainder attributed to β-turns and disordered structures, and that the binding of biotin did not result in any appreciable secondary structure changes. A similar Raman spectroscopic study was reported by Fagnano et al., in which detailed assignments of the vibrational spectra of avidin and the avidin-biotin complex were presented.28 The authors suggested that some relatively minor changes in secondary structure occurred upon binding of biotin. FT-IR studies under physiological conditions (pH = 7.4) including spectral assignments have also been reported by Barbucci et al.,29 where vibrational assignments were used to infer interactions between the avidin and the ureido functional group of biotin. Such studies investigating secondary structure contributions and conformational changes upon biotin binding have been carried out on avidin in its native solution or lyophilized form. To our knowledge, no adsorption-induced conformational changes have been reported for avidin, and investigating such changes is important due to the widespread use of the avidin-biotin bioconjugation reaction in potential applications. Furthermore, a better understanding of the avidin-biotin bioconjugation reaction at solidsolution interfaces is of interest, particularly those utilizing quantum dots where many bioconjugation studies have been reported but fewer mechanistic details of the process are available. Attenuated total reflection infrared (ATR-IR) spectroscopy with thin deposited particle films has in the past decade become widely used to provide detailed information about wet surface chemistry and adsorption processes.30-33 In this study, the binding of unmodified biotin by surface-immobilized avidin has been studied using in situ ATR-IR spectroscopic techniques.31 Avidin was first immobilized onto a CdS nanoparticle film following the initial adsorption of the bifunctional ligand mercaptoacetic acid (MAA). Subsequently, the binding of solution-phase (28) Fagnano, C.; Fini, G.; Torreggiani, A. J. Raman Spectrosc. 1995, 26, 991– 995. (29) Barbucci, R.; Magnani, A.; Roncolini, C.; Silvestri, S. Biopolymers 1991, 31, 827–834. (30) Hug, S. J.; Sulzberger, B. Langmuir 1994, 10, 3587–3597. (31) McQuillan, A. J. Adv. Mater. 2001, 13, 1034–1038. (32) Lefevre, G. Adv. Colloid Interface Sci. 2004, 107, 109–123. (33) Burgi, T.; Baiker, A. Adv. Catal. 2006, 50, 227–283.
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biotin by immobilized avidin was then investigated. All stages in the process were monitored by infrared difference spectroscopy.
Materials and Methods
General Methods. Solution pH measurements were made with a Mettler-Toledo MP220 meter to a precision of (0.1. BET surface area and pore size distribution measurements were performed using a TriStar 3000 N2 adsorption Analyzer at 77 K. The BET surface area of the prepared CdS was found to be 139 m2 g-1, and pore size distribution analysis indicated a BJH adsorption average pore radius of 2.6 nm. SEM imaging was performed with a JEOL 6700F field emission scanning electron microscope (JEOL, Japan). Samples were coated in an Emitech 575 high-resolution sputter coater (EM Technologies Ltd., England) with 5 nm of chromium and viewed at a working distance of 2-8 mm with a 3-7 kV range of accelerating voltages. SEM experiments were conducted on films prepared on glass microscope slides from aqueous suspensions, similarly to those used in the ATR-IR experiments. SEM images of a typical deposited CdS film used in this study are shown in Figure 2. Estimation of the film thickness using SEM images of film cross sections was reported previously, with an average thickness of ∼500 nm observed.34
Materials. Cadmium nitrate tetrahydrate (Scharlau, 99+%), :: sodium sulfide nonahydrate (Riedel-De Haen, 60-62% Na2S), :: mercaptoacetic acid (Riedel-De Haen, 90%), egg white avidin (Sigma-Aldrich, E1%/280), biotin (Sigma-Aldrich, 99%), and D2O (Cambridge Isotope Laboratories, Inc., 99.9%) were all used as received. All H2O used in experiments was deionized (Millipore, Milli-Q RG, resistivity 18 MΩ cm). Preparation of CdS Nanoparticles. Aqueous suspensions of uncapped CdS nanoparticles were prepared by the mixing of equal volumes of 2 10-3 mol L-1 cadmium nitrate and 2 10-3 mol L-1 sodium sulfide solutions, according to a previously reported method.34 The UV-vis absorption spectrum of an aqueous nanoparticle suspension showed a CdS band edge at ∼480 nm (2.59 eV), indicating an average particle size of 4.5 nm.35 An X-ray diffraction pattern obtained from a powder sample of the prepared CdS indicated a unit cell of cubic symmetry, in agreement with literature reports from similar preparations.36 Methods. ATR-IR Spectroscopy. A DuraSampleIR triple-reflection 3-mm-diameter diamond-faced ZnSe prism (ASI SensIR Technologies) and a Horizon accessory containing a thirteen-reflection 50 mm 10 mm 2 mm 45° ZnSe prism (Harrick Scientific) were used to collect spectral data. The observed absorbances are approximately proportional to the number of internal reflections within the accessory. The diamond surface of the triple-reflection prism isolates the CdS and any potentially corrosive solutions (e.g., pH extremes) from the underlying ZnSe surface. Prior to preparation of the CdS films for ATR-IR analysis, the prism surfaces were cleaned by polishing with 0.015 μm γ-alumina (BDH, polishing grade) on a wet polishing microcloth (Buehler) and then rinsed with water. For the triple-reflection prism, the CdS film was formed by depositing 10 μL of 1 10-3 mol L-1 CdS suspension onto the prism. The removal of most of the water using a water pump vacuum (∼50 mbar) for ∼30 min produced a ∼0.25 cm2 film. For the thirteen-reflection prism, the film was formed in a similar manner by depositing 700 μL of 1 10-3 mol L-1 CdS suspension to produce a ∼4.1 cm2 film. All films were deposited on the prisms immediately following the CdS suspension preparation and were initially washed for 20 min with 1 10-2 34 mol L-1 NaOH to remove any surface S2O23 ligands. Each prism was interfaced via a rubber O-ring to a flow cell. The triple- and thirteen-reflection prisms incorporated glass and poly(methyl methacrylate) (PMMA) flow cells, respectively, each of which was manufactured in this department. The solutions were delivered to the flow cell using a Masterflex C/L peristaltic pump and Masterflex Tygon tubing at a constant flow rate of 1 mL min-1. The IR analysis of the CdS films under solution flow was conducted in the absence of laboratory lighting. A Digilab FTS 4000 infrared spectrometer equipped with a KBr beamsplitter, Peltier cooled DTGS detector, and WinIR Pro v 3.4 software was used to collect and analyze spectra. The optical bench was purged with dried air. ATR-IR spectra were obtained from 64 scans at 4 cm-1 resolution and were not corrected for dependence of absorbance on wavenumber. Spectra are shown as recorded at room temperature (22 °C) and were not further modified by baseline correction or subtraction.
Solution IR Spectra of MAA, Avidin, and Biotin in H2O. Figure 3a-c shows the ATR-IR spectra of the solution species MAA, avidin, and biotin, respectively, recorded in H2O at pH = 7.2. In the figures, only the 1800-800 cm-1 range is shown due to the lack of useful information present outside this spectral region. The peak wavenumbers of the C-H stretching modes are relatively insensitive to adsorption. Spectrum (a) of Figure 3 is that of a 1 10-1 mol L-1 MAA solution at pH = 7.2, a detailed analysis of which has been reported previously by this group.34 This spectrum is dominated by the antisymmetric and symmetric carboxylate stretching absorptions at 1569 and 1388 cm-1, respectively. A weaker peak at 1240 cm-1 has been assigned to a CH2 wagging mode in a Raman spectroscopic study.37 Peak wavenumbers and assignments for MAA, avidin, and biotin, in both the solution and adsorbed phases, in H2O and D2O are given in Table 1. Spectrum (b) of Figure 3 is that of a 1 10-3 mol L-1 solution of avidin at pH = 7.2. The spectral features are dominated by the amide I and II peaks centered at 1628 and 1537 cm-1, respectively. The amide I peak arises from the CdO stretching vibration with minor contributions from the C;N stretching vibration, the C;C;N deformation, and the N;H in-plane bend.38 The amide I peak has a clear shoulder at higher wavenumber which reflects contributions from β-turn, β-sheet, and unordered secondary structures of the protein backbone.39 The amide II peak results from a combination of the N;H inplane bend and the C;N stretching vibration with smaller contributions from the C;O in-plane bend and the C;C and N;C stretching vibrations.38 Detailed analysis of the amide IR absorption features of avidin have been reported in numerous studies.27,29,39 After the amide I and II peaks, the next strongest avidin IR absorption is at 1075 cm-1. This peak and a smaller shoulder at 1157 cm-1 are due to glycosylation of avidin, principally from mannose and N-acetylglucosamine residues.40 Each avidin subunit contains 129 amino acid residues, 4 mannose residues, and 3 N-acetylglucosamine residues.41 The sugar residues account for ∼10% w/w of avidin and are all located
(16) Pugliese, L.; Malcovati, M.; Coda, A.; Bolognesi, M. J. Mol. Biol. 1994, 235, 42–46. (17) Weber, P. C.; Ohlendorf, D. H.; Wendoloski, J. J.; Salemme, F. R. Science 1989, 243, 85–88. (34) Young, A. G.; Green, D. P.; McQuillan, A. J. Langmuir 2006, 22, 11106– 11112. (35) Brus, L. E. J. Chem. Phys. 1983, 79, 5566–5571. (36) Olshavsky, M. A.; Allcock, H. R. Chem. Mater. 1997, 9, 1367–1376.
(37) Castro, J. L.; Lopez-Ramirez, M. R.; Centeno, S. P.; Otero, J. C. Biopolymers 2004, 74, 141–145. (38) Barth, A. Biochim. Biophys. Acta 2007, 1767, 1073–1101. (39) Celej, M. S.; Montich, G. G.; Fidelio, G. D. Biochem. Biophys. Res. Commun. 2004, 325, 922–927. (40) Khajehpour, M.; Dashnau, J. L.; Vanderkooi, J. M. Anal. Biochem. 2006, 348, 40–48. (41) DeLange, R. J. J. Biol. Chem. 1970, 245, 907–916.
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Results and Discussion
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Figure 2. SEM images of a typical deposited CdS film used in this study.
in a carbohydrate appendage attached to an asparaginyl residue of the avidin protein backbone.41,42 The carbohydrate appendages are evident as the ball and stick features shown in the structure of avidin in Figure 1.16 This structure was sourced from a report where there had been incomplete removal of glycoslyated forms using lectin columns prior to crystallization, and only a single N-acetylglucosamine residue of the carbohydrate appendage remained.16 Other smaller unassigned peaks at 1449, 1395, and 1235 cm-1 in spectrum (b) of Figure 3 are likely to arise from the overlapping absorptions of various amino acid side chains. For proteins in general, below 1700 cm-1 all amino acid side chain absorptions overlap with those of other side chains or of the protein backbone,43 which complicates assignment of such absorptions to specific residues. Spectrum (c) of Figure 3 is that of a 1 10-1 mol L-1 solution of biotin at pH = 7.2. The dominant spectral features include the ketone stretching, and antisymmetric and symmetric carboxylate stretching absorptions at 1674, 1546, and 1407 cm-1, respectively. A smaller peak at 1480 cm-1 has been assigned to the CH2 deformation.29 A recent computational (gas phase) study of biotin assigned peaks at 1339 and 1272 cm-1 to C-C and C-O stretching vibrations, respectively.44 IR Spectra of MAA Adsorbed on CdS in H2O. In Figure 3d-f, the in situ spectra from the bioconjugation experiment carried out in H2O on a film of deposited CdS nanoparticles are presented. The approach to the bioconjugation reaction involved the sequential adsorption of MAA, avidin, and the subsequent binding of biotin. This stepwise series of flowed solutions gave a progressively more complex surface structure, and difference spectra were used for observing changes at each step. The selected bioconjugation procedure was based on a reported method by Jaiswal et al.45 Their method employed an extra protein, MBP-zb, which competed with avidin for QD surface adsorption sites and provided a means to regulate the number of avidin molecules present on each QD. MBP-zb was not used in the current study, to simplify spectral assignment and maximize the signal-to-noise ratio of adsorbed avidin in H2O. Spectrum (d) of Figure 3 shows the time-evolution over 35 min of IR spectra during the adsorption of MAA to a CdS film from a 1 10-3 mol L-1 MAA solution at pH = 7.2, for which spectral assignments have been reported previously by this group.34 At a solution concentration of 1 10-3 mol L-1, there is no significant MAA solution contribution to the spectrum. This MAA solution concentration is high enough to result in sufficiently fast adsorption kinetics such that adsorption equilibrium was reached well before the allocated 35 min of (42) Huang, T.-S.; DeLange, R. J. J. Biol. Chem. 1971, 246, 686–697. (43) Barth, A.; Zscherp, C. Q. Rev. Biophys. 2002, 35, 369–430. (44) Emami, M.; Teimouri, A.; Chermahini, A. N. Spectrochim. Acta A 2008, 71, 1516–1524. (45) Jaiswall, J. K.; Goldman, E. R.; Mattoussi, H.; Simon, S. M. Nat. Methods 2004, 1, 73–78.
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Figure 3. ATR-IR spectra of (a) 1 10-1 M MAA solution at
pH = 7.2, (b) 1 10-3 M avidin solution at pH = 7.2, and (c) 1 10-1 M biotin solution at pH = 7.2; (d) MAA adsorbed on CdS from 1 10-3 M MAA solution at pH = 7.2 at 1, 2, and 35 min; (e) avidin adsorbed on MAA/CdS from 1 10-3 M avidin/ 1 10-3 M MAA solution at pH = 7.2 at 1, 9, and 60 min; and (f) binding of biotin by avidin/MAA/CdS from a 1 10-4 M biotin/ 1 10-3 M MAA solution at pH = 7.2 at 1, 5, and 55 min. The background spectra were from (a)-(c) H2O on diamond, (d) H2O (pH = 7.2) on CdS, (e) 1 10-3 M MAA (pH = 7.2) on MAA/ CdS, and (f) 1 10-3 M avidin/1 10-3 M MAA (pH = 7.2) on avidin/MAA/CdS. Final spectrum of time-evolution spectra is bolded. All spectra were obtained in H2O using a triple-reflection accessory, from 64 scans at 4 cm-1 resolution.
flow time, and no further MAA increases were expected in the following avidin adsorption stage. MAA adsorbs to the CdS surface via the thiol functional group, and at pH = 7.2 leaves the carboxylate group exposed to the solution and available for further reaction. An adsorption isotherm of MAA on CdS has been reported previously by this group, and a Langmuir adsorption (affinity) constant KL ∼ 105 L mol-1 was obtained.34 Thus, a MAA solution concentration of 1 10-3 mol L-1 corresponds to >99% MAA equilibrium surface coverage on CdS and thus generating a high negative surface charge at pH = 7.2. DOI: 10.1021/la900350s
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Table 1. IR Peak Wavenumbers and Assignments for MAA, Avidin, and Biotin, in Solution and Adsorbed Phase, in H2O and D2Oa H2O (pH = 7.2) solution
adsorbed
D2O (pD = 7.2) solution
adsorbed
MAA νas(CO2) νs(CO2) CH2 wag37
1569 1388 1240
1555 1384 1227
1549 1386 1223
1565 1386 1224
1631 ∼1540 ∼1450
1630 1561 1447 1401
Avidin amide I
1628
1632
amide II
1537 1395 1235 1157 1075
1548 1389 1230
glycosylation40 glycosylation40
1081 Biotin
ν(CdO) νas(CO2) δ(CH2)29 νs(CO2 ) ν(C-C)44 ν(C-O)44
1674 1546 1480 1407 1334 1270
1651 1552 1462 1409
1269 1227 a The symbols ν and δ refer to stretching and deformation, respectively. The subscripts “as” and “s” refer to antisymmetric and symmetric, respectively.
IR Spectra of Avidin Adsorbed on MAA/CdS in H2O. The next step in setting up the bioconjugation reaction was the adsorption of avidin to the carboxylated CdS surface. Reported studies where avidin was immobilized on carboxylated Si and SiO2 surfaces have involved covalent bond formation via N-hydroxysuccinimide (NHS) and 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) activating agents.18,46 No activating agents were used in the present work, and the interaction between the positively charged avidin (pHiep = 10 for avidin47) and the negatively charged CdS surface is electrostatic, rather than covalent.45 The nonspecific electrostatic (or passive) adsorption of avidin has been reported on a wide variety of substrates, including carboxylated Au surfaces (using mercaptoundecanoic acid),48 carboxylated indium tin oxide surfaces (using mercaptopropionic acid),49 tin oxide surfaces,50 perfluorinated nanoparticle surfaces,51 and others.52,53 Protein binding experiments are often carried out in phosphate-buffered saline (PBS) or sodium borate buffer solutions to provide protein stability and constant pH. Such buffers were not used in the current study, due to signal interference by these generally strongly IR absorbing species. The short experimental time-scale of avidin adsorption at room temperature and physiological pH constitutes very mild conditions, which are unlikely to result in significant denaturing of solution-phase avidin. It was necessary to include MAA in the subsequent adsorbing species solutions to maintain the solution-adsorbed MAA interfacial (46) Gu, J.; Yam, C. M.; Li, S.; Cai, C. J. Am. Chem. Soc. 2004, 126, 8098–8099. (47) Blanco, E. M.; Horton, M. A.; Mesquida, P. Langmuir 2008, 24, 2284–2287. (48) Jordan, C. E.; Corn, R. M. Anal. Chem. 1997, 69, 1449–1456. (49) Wei, M.-Y.; Guo, L.-H.; Chen, H. Microchimica Acta 2006, 155, 409–414. (50) Dong, D.; Zheng, D.; Wang, F.-Q.; Yang, X.-Q.; Wang, N.; Li, Y.-G.; Guo, L.-H.; Cheng, J. Anal. Chem. 2004, 76, 499–501. (51) Prosperi, D.; Morasso, C.; Tortora, P.; Monti, D.; Bellini, T. ChemBioChem 2007, 8, 1021–1028. (52) Guo, L.-H.; Yang, X.-Q. Analyst 2005, 130, 1027–1031. (53) Jiang, X.; Xu, Q.; Dertinger, S. K. W.; Stroock, A. D.; Fu, T.; Whitesides, G. M. Anal. Chem. 2005, 77, 2338–2347.
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equilibrium and prevent MAA desorption obscuring spectral regions of interest. Spectrum (e) of Figure 3 shows the time evolution over 60 min of IR difference spectra during the adsorption of avidin to a MAA/CdS film from a solution containing 1 10-3 mol L-1 avidin and 1 10-3 mol L-1 MAA at pH = 7.2. The spectral baseline is distorted by displacement of bulk water from the interface relative to the background spectrum, which is observed in the broad negative absorptions at ∼1650 and
