DOI: 10.1021/la902226b 1029Langmuir 2010, 26(2), 1029–1034 Published on Web 09/09/2009

pubs.acs.org/Langmuir

© 2009 American Chemical Society

On the Adsorption Behavior of Biotin-Binding Proteins on Gold and Silica

Patricia M. Wolny,†,‡,§ Joachim P. Spatz,‡,§ and Ralf P. Richter*,†,‡

†Biosurfaces Unit, CIC biomaGUNE, Paseo Miramon 182, 20009 Donostia-San Sebastian, Spain,‡Department of New Materials and Biosystems, Max-Planck-Institute for Metals Research,Heisenbergstrasse 3, 70569 Stuttgart, Germany, and §Department of Biophysical Chemistry,

University of Heidelberg, INF 253, 69120 Heidelberg, Germany

Received June 20, 2009. Revised Manuscript Received August 22, 2009

Streptavidin (SAv), avidin (Av), and neutravidin (NAv) have become widely used molecular tools in biotechnologythanks to their remarkable affinity for biotin. Their tetravalency renders these molecules particularly interesting for thefunctionalization of solid-liquid interfaces. Using the quartz crystal microbalance with dissipation monitoring, wesystematically investigate the deposition of biotin-binding proteins to two surfaces that are popular in biotechnology:gold and silica. We find that simple physisorption of biotin-binding proteins is a viable method to confer biotin-bindingfunctionality to gold surfaces. Both SAv and Av form dense, stable protein monolayers that retain biotin-bindingactivity and are largely inert to the unspecific binding of bovine serum albumin. Furthermore, we report that SAv resistsadsorption to silica over a wide range of pH and ionic strength. The contrast in the binding behavior of SAv on silica andon gold suggests a simple strategy for the selective biofunctionalization of nano- or microstructured surfaces.

Introduction

The interaction between biotin and streptavidin (SAv) isknown to be one of the strongest noncovalent biomolecularbinding events with an affinity of 1013 M-1.1 In addition, SAvis stable in aqueous solution over a wide range of temperature,pH, and salt concentration.2 These features have rendered SAv,next to other biotin-binding proteins like avidin (Av) and neu-travidin (NAv), very popular for biotechnological and biochem-ical applications.

With a total of four biotin binding sites, located on twoopposing faces of the molecule, such biotin-binding proteins areparticularly attractive for the functionalization of macroscopicsurfaces or nanoparticles. (See ref 3 and references therein.) Knollet al.,4,5 for example, have successfully employed SAv as a“bridge” between a biotinylated surface-a mixed self-assembledmonolayer (SAM) made from biotinylated and nonbiotinylatedalkyl thiols on gold-and a biotinylated target. Other biotin-binding molecules have also been immobilized on layers ofbiotinylated thiols,6,7 and photochemical approaches that useeither photoactivated or photocleaved biotin groups have beenreported.8,9 In all of these approaches, the multivalency of thebiotin-binding proteins is directly exploited to link a biotinylatedtarget to a surface exposing biotin groups.

Such sandwich strategies are not unique to gold surfaces.10,11

Huang et al.12 used assemblies of grafted copolymers of poly-(L-lysine) and partially biotin-derivatized poly(ethylene glycol) toimmobilize biotin-binding proteins on niobium oxide surfaces.Supported lipid bilayers that contain a fraction of biotin-functio-nalized lipids (biotin-SLBs) are attractive for the functionaliza-tion of mica and silica: the biotin concentration can be easilytuned and biotin-binding proteins are immobilized in a highlyoriented manner.13-17

In other strategies, biotin-binding proteins were covalentlyimmobilized. Reznik et al.,18 for example, mutated SAv with aC-terminal cystein tag for immobilization onto maleimid-coatedsurfaces. Primary amines or carboxyl groups on the proteins’surface have also been employed for covalent bonding to suitablyprefunctionalized surfaces of silica19,20 and gold.5 The nature ofthe coupling chemistry seldom allows for the orientation of themolecule on the surface to be controlled. Here, the multivalencyincreases the likelihood for biotin-binding sites to remain acces-sible for target molecules in solution.

All above-described functionalization approaches have incommon that they require the presence of an additional linkerlayer between the solid support and the biotin receptor. Thepreparation of a suitable linker layer is not a trivial task: carefultuning of the layer assembly is often required to optimize

*Corresponding author. E-mail: rrichter@cicbiomagune.es.(1) Wilchek, M.; Bayer, E. A. Biomol. Eng. 1999, 16, 1–4.(2) Green Michael, N. Method. Emzymol. 1990, 184, 51–67.(3) Rusmini, F.; Zhiyuan, Z.; Jan, F. Biomacromolecules 2007, 8, 1775–1789.(4) Knoll, W.; Zizlsperger,M.; Liebermann, T.; Arnold, S.; Badia, A.; Liley,M.;

Piscevic, D.; Schmitt, F.-J.; Spinke, J. Colloids Surf., A 2000, 161, 115–137.(5) Su, X.; Wu, Y.-J.; Robelek, R.; Knoll, W. Langmuir 2005, 21, 348–353.(6) Yue, M.; Stachowiak, J. C.; Lin, H.; Datar, R.; Cote, R.; Majumdar, A.

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C. M. J. Phys. Chem. B 2008, 112, 6708–15.(8) Pritchard, D. J.; Morgan, H.; Cooper, J. M.Angew. Chem., Int. Ed. 1995, 34,

91–93.(9) Kim, K. J. Am. Chem. Soc. 2004, 126, 15368–69.(10) Weisser, M.; Tovar, G.; Mittler-Neher, S.; Knoll, W.; Brosinger, F.;

Freimuth, H.; Lacher, M.; Ehrfeld, W. Biosens. Bioelectron. 1999, 14, 405–411.(11) Birkert, O.; Haake, H. M.; Schutz, A.; Mack, J.; Brecht, A.; Jung, G.;

Gauglitz, G. Anal. Biochem. 2000, 282, 200–8.

(12) Huang, N.-P.; V€or€os, J.; De Paul, S. M.; Textor, M.; Spencer, N. D.Langmuir 2002, 18, 220–230.

(13) Larsson, C.; Rodahl, M.; H€o€ok, F. Anal. Chem. 2003, 75, 5080–5087.(14) Richter, R. P.; Hock, K. K.; Burkhartsmeyer, J.; Boehm, H.; Bingen, P.;

Wang, G.; Steinmetz, N. F.; Evans, D. J.; Spatz, J. P. J. Am. Chem. Soc. 2007, 129,5306–5307.

(15) Bingen, P.; Wang, G.; Steinmetz, N. F.; Rodahl, M.; Richter, R. P. Anal.Chem. 2008, 80, 8880–8890.

(16) Reimhult, E.; Larsson, C.; Kasemo, B.; H€o€ok, F. Anal. Chem. 2004, 76,7211–7220.

(17) Reviakine, I.; Brisson, A. Langmuir 2001, 17, 8293–8299.(18) Reznik, G. O.; Vajda, S.; Cantor, C. R.; Sano, T. Bioconjugate Chem. 2001,

12, 1000–1004.(19) Baselt, D. R.; Lee, G. U.; Hansen, K. M.; Chrisey, L. A.; Colton, R. L.

Proc. IEEE 1997, 85, 672–680.(20) Bhushan, B.; Tokachichu, D. R.; Keener, M. T.; Lee, S. C. Acta Biomater.

2005, 1, 327–341.

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Article Wolny et al.

performance in terms of biotin-binding activity and resistance tounspecific binding. For example and contrary to what intuitionmay suggest, the display of too many biotin groups by the linkerlayer typically has adverse effects on the biotin-binding capacityof the final functionalized surface: excess biotin groups on denselypacked molecular layers impose steric constraints that limit thebinding of biotin receptors, as nicely demonstrated for SAMs;4 toohigh of a biotin density in a layer of flexible linkers can lead to theoccupation of all four of the biotin receptor’s binding sites or theburial of biotin receptors in the interior of the linker layer, therebyrendering the receptors inactive or inaccessible.12 Also, the uncon-trolled orientation of biotin receptors upon covalent linkage has beenreported to limit the functionality of immobilized target molecules.5

Despite the popularity of SAv, Av, and NAv as moleculartools, rather little is currently known about the direct interactionbetween these proteins and surfaces of silica, glass, and gold (i.e.,materials that are commonly employed in biotechnologicalapplications). To the best of our knowledge, for example, thereis only a single study on the adsorption behavior of SAv on baresilica surfaces.20 Spontaneous adsorption of SAv and Av to goldand silver has been reported and suggested for potential immo-bilization applications.21a A quantification of the amount ofbound material and of the retention of biotin-binding activity inthe immobilized state is, however, lacking.

As part of recent efforts to develop selective functionalizationstrategies for nanostructured surfaces, we have investigated theadsorption behavior of three different biotin-binding proteins-SAv, Av, and NAv-on gold and silica in a systematic manner.Here, we report on the characterization, by quartz crystal micro-balance with dissipationmonitoring (QCM-D), of the adsorptionprocesses as well as the stability, morphology, and biotin-bindingactivity of the resulting protein films. QCM-D has previouslybeen demonstrated to be a very sensitive, powerful method for thecharacterization of binding processes at surfaces, providing time-resolved information not only about adsorbed amounts but alsoabout themorphology of surface-confined films.22,23We find thatthe simple physisorption of biotin-binding proteins to gold can,despite its crudeness, represent a viable alternative for thefunctionalization of surfaces with biotin receptors. In particular,we find an interesting contrast when comparing the bindingbehavior of SAv on gold and silica, respectively.

Materials and Methods

Materials. Lyophilized avidin (Av), streptavidin (SAv),FITC-labeled streptavidin (FITC-SAv), bovine serum albumin(BSA), and biotinylated bovine serum albumin (b-BSA) werepurchased from Sigma-Aldrich (Schnelldorf, Germany). Neutra-vidin (NAv) was purchased from Invitrogen (Karlsruhe,Germany). All proteinswere dissolved inPBS (10mMphosphate,137 mM NaCl, 2.7 mM KCl) at a concentration of 1 mg/mL,divided into aliquots, and stored at -20 �C. Before use, theproteins were diluted in working buffer to a final concentra-tion of about 20 μg/mL. If not otherwise stated, PBS wasemployed as a working buffer. In some cases, Hepes buffer(10 mM Hepes, 150 mM NaCl, 3 mM NaN3, and 2 mM CaCl2)was used. The pH of the buffers was 7.4, unless otherwisestated, and adjusted with either HCl or NaOH. All bufferingredients were purchased from Sigma.

Supported lipid bilayers (SLBs) were prepared by the exposureof small unilamellar vesicles (SUVs) to silica-coated QCM-Dcrystals, as described earlier.24 The SLB-formation processwas monitored by QCM-D. SUVs were prepared by sonica-tion from mixtures of dioleoylphosphatidylcholine (DOPC) anddioleoylphosphatidylethanolamine-CAP-biotin (DOPE-CAP-biotin) (Avanti Polar Lipids, Alabaster, AL) in a molar ratioof 9:1.24 They were applied at a final concentration of about50 μg/mL in Hepes buffer.

Surfaces. QCM-D crystals with a resonance frequency of4.95MHz, coated with either gold or silica, were purchased fromQ-Sense AB (V€astra Fr€olunda, Sweden). In some cases, an addi-tional silica layer of 100 nm thickness was evaporated on the gold-coated crystals (G. Albert PVD Beschichtungen, Silz, Germany).Differences between this additional silica layer and the silicacoating provided by O-Sense with respect to protein adsorptionwere not observed.

Gold-coated crystals were cleaned by immersion in a mixtureof NH3/H2O2/H2O in a volume ratio of 1:1:4 at a temperature of70 �C for 5 min, rinsed thoroughly with ultrapure water(Barnstead, model D1193, Dubuque, IA), and blown dry in N2.Silica-coated crystals were immersed in an aqueous solution of3% sodium dodecyl sulfate for 30 min, rinsed abundantly withultrapure water, and blown dry in N2. Prior to use, the crystalswere exposed to oxygen plasma (200 W, 0.35 mbar) for 25 min(TePla 100-E, Feldkirchen, Germany).

Static contact angle measurements (DSA100, Kr€uss GmbH,Hamburg,Germany) revealed that the cleaned silica surfaceswerevery hydrophilic, inducing complete wetting.

Quartz Crystal Microbalance with Dissipation Monitor-

ing (QCM-D). QCM-D is an electromechanical method thatallows adsorption processes on a solid surface to be followed inreal time.25 Briefly, upon interaction of (soft) matter with thesurface of a QCM-D sensor crystal, changes in its resonancefrequency,Δf, related to attachedmass, and in its dissipation,ΔD,related to frictional (viscous) losses in the adlayer, are measuredwith a time resolution of better than 1 s.

As shown by Sauerbrey,26 the mass uptake, Δm, is directlyproportional to the frequency change, Δfn, of the oscillatingsensor crystal, provided that the adsorbed film is sufficiently thin,rigid, and homogeneous

Δm ¼ -CΔfnn

ð1Þ

whereC is themass sensitivity constant (here 18.06 ng/cm2/Hz) andn is the overtone number. In solution, the QCM-D is also sensitiveto water that is coupled to the adsorbed molecular film. The massdetermined by eq 1 therefore typically exceeds the biomolecularmass of the film.15 For a sufficiently rigid biomolecular film at highcoverage, the frequency shift typically corresponds rather well tothe biomolecular mass together with the mass of the solvent that iscontained inside the film.Hence, it appears reasonable to representthe adsorbed film in terms of acoustic thickness:16

d ¼ Δm

FH2O

¼ -C

FH2O

Δfnn

ð2Þ

The acoustic thickness neglects the density differencesbetween thebiomolecules (F ≈ 1.4 g/cm3 for proteins13,27) and water (FH2O

=1.0 g/cm3).With typically 50%ormore water being present in thefilm,15 dmay thus overestimate the geometrical thickness by up to

(21) (a) Ebersole, R. C.; Miller, J. A.; Moran, J. R.; Ward, M. D. J. Am. Chem.Soc. 1990, 112, 3239–3241. (b) Tsortos, A.; Papadakis, G.; Mitsakakis, K.; Melzak,K. A.; Gizeli, E. Biophys. J. 2008, 94, 2706–2715.(22) H€o€ok, F.; Larsson, C.; Fant, C. In Encyclopedia of Surface and Colloid

Science; Hubbard, A. T., Ed.; Marcel Dekker: New York, 2002; pp 774-791.(23) Janshoff, A.; Steinem, C. Piezoelectric Sensors; Springer Series on Chemical

Sensors and Biosensors; Wolfbeis, O. S., Ed.; Springer: Berlin, 2007; Vol. 5.

(24) Richter, R.; Mukhopadhyay, A.; Brisson, A. Biophys. J. 2003, 85, 3035–3047.

(25) Rodahl, M.; H€o€ok, F.; Krozer, A.; Brzezinski, P.; Kasemo, B. Rev. Sci.Instrum. 1995, 66, 3924–3930.

(26) Sauerbrey, G. Z. Phys. A 1959, 155, 206–222.(27) Cuypers, P. A.; Corsel, J. W.; Janssen, M. P.; Kop, J. M. M.; Hermens, W.

T.; Hemker, H. C. J. Biol. Chem. 1983, 258, 2426–2431.

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Wolny et al. Article

20%. In the following text, we will speak in terms of measuredshifts in frequency and dissipation as well as acoustic thickness.

QCM-D measurements were conducted with a Q-Sense E4system. The systemwas operated in flowmode, with a continuousliquid flux of typically 25 μL/min being delivered by a peristalticpump (model Reglo digital, Ismatec, Glattbrugg, Switzerland).To switch between sample liquids, the flow was interrupted for afew seconds without disturbing the QCM-D signal. The workingtemperature was 24.0 �C. The resonance frequency and dissipa-tion were measured at six harmonics (∼15, 25...65 MHz), simul-taneously. For simplicity, only changes in the dissipationand normalized frequency, Δf = Δfn/n, of the seventh overtone(n = 7, i.e., ∼35 MHz) are presented.

Results

The ability of SAv and other biotin receptors to bind biotin-tagged molecules is not necessarily given when the protein isimmobilized on a solid support. It is well known that proteins canundergo structural changes upon adsorption and lose theirbiological activity.28,29 Also, the surface may impose steric con-straints that limit binding. Successful immobilization should thusyield a surface on which (i) the optimum amount of protein isimmobilized, (ii) the protein’s biotin-binding activity is main-tained, and (iii) unspecific binding is limited.

This study focuses on a systematic examination of the immo-bilization of biotin-binding proteins by direct deposition on baregold and bare silica surfaces. We start, however, by recalling andextending data for another immobilization platform: biotin-functionalized supported lipid bilayers. The adsorption of SAvto biotin-SLBs has already been characterized in detail by QCM-Dandothermethods13,15-17 andwas found toperformwell in variousapplications.13,14,30 Particular features of this system are that it canbe preparedwith high reproducibility, that the orientation of biotinreceptor molecules is well defined and optimized in the sense thattwo biotin binding sites point directly into the bulk solution, that itis resistant to unspecific binding, and that the surface density ofbiotin receptors is tunable. These characteristics make biotin-SLBsa well-suited reference system for the present study.Biotinylated SLBs: The Reference System.Upon exposure

to biotinylated SLBs, SAv adsorbed readily, as indicated by themonotonously decreasing frequency (Figure 1A). The adsorptionreached a plateau within less than 15 min, yielding overall changesin frequency of-28Hz,which corresponds to a thickness of 5.1 nmand dissipation of 0.5� 10-6. The adsorption kinetics and the finalQCM-D response are consistent with earlier reports and indicatethe formation of a dense monolayer of SAv on the SLB.13,15,16

Biotinylated BSA (b-BSA), successively added to the SAv-covered SLB, bound with a maximum frequency change of-13.5 Hz (d = 2.4 nm, Figure 1C). Considering the shape ofBSA, a flattened ellipsoid of 3 nm � 4 nm � 8 nm in size,31 thisfrequency shift suggests that the protein adsorbs with its long axisparallel to the surface.As for SAv, equilibriumwas reached ratherquickly, and the change in dissipationwas low, consistentwith theformation of a monolayer of globular proteins. Binding wasirreversible as expected for the strong interaction between biotinand SAv. Native BSA did not bind to the SAv-covered SLB(|Δf| < 0.5 Hz, Figure 1B), confirming that the binding of b-BSAwas specific and SAv was thus active.

Similar responses were observed for other biotin-binding proteins(Figure 1, Table 1). Av and NAv adsorption to biotinylated SLBsequilibrated within about 15 min, with final frequency shifts of-26 Hz (4.7 nm) and -32 Hz (5.8 nm), respectively, indicatingmonolayer formation. b-BSA bound stably to Av (-15 Hz) andNAv (-9.5 Hz) whereas native BSA did not bind, indicating thatthese two proteins retain their activity upon immobilization. How-ever, somepeculiaritieswerenotable forNAv: thedissipation shift forthe NAv film (1� 10-6) was considerably larger than for Av (0.6�10-6) andSAv, andb-BSAbindingwas reduced to-9.5Hz.Wealsoobserved that the adsorption of NAv was not well reproducible. Insome cases, absolute frequency shifts of 40 Hz and more wereobserved. The binding of b-BSA was not affected by this variability.Gold Surfaces. Next, we investigated the adsorption of SAv to

gold.Av andNAvwere tested for comparison.All proteins adsorbedwell to gold-coated QCM-D crystals (Figure 2A, Table 1). Adsorp-tion kinetics and final frequency shifts for SAv (-30 Hz) and Av(-32 Hz) were similar to, albeit slightly higher than, those observedon biotin-SLBs, indicating the formation of a dense monolayer. Thebinding of both molecules was stable upon rinsing in buffer. Theseexperiments support earlier findings of spontaneous and directbinding of Av and SAv to gold and silver surfaces.21a

The dissipation shifts for both proteins on goldwere significantlysmaller (<0.2 � 10-6) than on biotin-SLBs, indicating subtledifferences in the mode of immobilization. Indeed, the unspecificbinding to gold is direct and likely to proceed via tight association ofone of the extended faces of the cuboid-shaped proteins with thesurface, whereas specific binding to biotin-SLBs is mediated by oneor two short but flexible linkers. We suggest that it is this differencein linkage that gives rise to the different dissipation signals.32

Figure 1. (A) Representative QCM-D responses, Δf and ΔD, forthe formation of biotinylated supported lipid bilayers and subse-quent adsorption of biotin-binding proteins SAv (blue triangles),Av (green rectangles), andNAv (red circles). The typical two-phaseresponses inΔf andΔDwith the expected final shifts of-25Hz and<0.5 � 10-6, respectively, were measured for the formation ofSLBs.24,48 (B) Subsequent exposure of native BSA on the samesurfaces reveal no binding. (C) Subsequent binding of b-BSA onthe same surfaces. Each incubation step starts at 0 min, except forthe incubation of SAv, Av, and NAv that starts at 20, 40, and20 min, respectively; rinses in buffer are indicated (arrow heads).

(28) Welzel, P. B. Thermochim. Acta 2002, 382, 175–188.(29) H€o€ok, F.; Rodahl, M.; Kasemo, B.; Brzezinski, P. Proc. Natl. Acad. Sci. U.

S.A. 1998, 95, 12271–12276.(30) Berquand, A.;Mazeran, P.-E.; Pantigny, J.; Proux-Delrouyre, V.; Laval, J.-

M.; Bourdillon, C. Langmuir 2003, 19, 1700–1707.(31) H€o€ok, F.; Rodahl, M.; Brzezinski, P.; Kasemo, B. Langmuir 1998, 14, 729–

734. (32) Johannsmann, D.; Reviakine, I.; Richter, R. P. Anal. Chem., in press.

1032 DOI: 10.1021/la902226b Langmuir 2010, 26(2), 1029–1034

Article Wolny et al.

The adsorption behavior of NAv differed markedly from thatof the other two biotin receptors. Whereas the adsorption of Avand SAv reached a plateau after around 15 min of incubation,NAv continued to bind, albeit more slowly. Frequency shiftsbeyond-60Hzwere reached, which is twice the response for SAvorAv. The large difference is surprising, given that the dimensionsof all proteins are similar. Δf = -60 Hz would correspond to athickness of around 11 nm, or about twice NAv’s moleculardimensions, suggesting that the adsorption of NAv on gold is notrestricted to a monolayer. Notably, very similar responses havealready been reported7 for the adsorption of NAv on goldsurfaces that were coated with a layer of biotinylated thiols. Wespeculated that aggregates in the stock solution may be the causeof increased adsorption. Indeed, centrifugation of freshly thawedaliquots at 10000g for 10 min resulted in decreased QCM-Dresponses (data not shown). The absolute frequency shift remainedthough at least 10Hz higher thanwhat was observed for SAv andAv. It is likely that gentle centrifugation is insufficient to removeall aggregates. At present, we cannot exclude that surface-induceddenaturation of NAv may also contribute to the unexpectedadsorption behavior. We note that the apparent instability of

NAv, in solution and/or upon confinement to a surface, may alsoexplain the variability in the binding behavior, the increaseddissipation, and the reduced activity of NAv that we observedon biotin-SLBs.

Table 1. QCM-D Responses upon Incubation of Surfaces with Biotin-Binding Proteins, BSA, and b-BSAa

on b-SLB on gold on silica

|Δf | (Hz) ΔD (10-6) |Δf | (Hz) ΔD (10-6) |Δf | (Hz) ΔD (10-6)

SAv SAv 28 ( 2 0.5 ( 0.1 30 ( 2 0.1 ( 0.1 <0.5 <0.1BSA <0.5 <0.1 1 ( 1 <0.1 <0.5 <0.1b-BSA 13.5 ( 1 0.3 ( 0.1 12.5 ( 1 0.5 ( 0.1 <0.5 <0.1

Av Av 26 ( 1 0.6 ( 0.1 32 ( 2 0.1 ( 0.1 20 ( 4 0.3 ( 0.2BSA <0.5 <0.1 3 ( 1 <0.2 <0.5 <0.1b-BSA 15 ( 1 0.6 ( 0.1 15.5 ( 2 0.4 ( 0.1 14 ( 2 0.5 ( 0.1

NAv NAv g32 g0.9 >40 >1.2 >60 >3BSA <0.5 <0.1 0.5 ( 0.5 0.1 ( 0.1 <0.5 <0.1b-BSA 9.5 ( 1 0.2 ( 0.1 >10 (0.5 16 ( 2 -0.3 ( 0.2

aResponses after rinsing in buffer are given. Errors correspond to the spread of experimental data, obtained from two to five measurements persample. (See Figures 1-3 for representative measurements.)

Figure 2. (A) Representative QCM-D responses, Δf and ΔD, forthe adsorption of biotin-binding proteins to gold: SAv (bluetriangles), Av (green rectangles), and NAv (red circles). (B) Sub-sequent exposure of native BSA on the same surfaces. (C) Sub-sequent binding of b-BSA on the same surfaces. Each incubationstep starts at 0 min; rinses in buffer are indicated (arrow heads).

Figure 3. (A) Representative QCM-D responses, Δf and ΔD, forthe adsorption of biotin-binding proteins to silica: SAv (bluetriangles), Av (green rectangles), and NAv (red circles). (B) Ex-posure of native BSA on other surfaces, prepared identically tothose shown in A, reveal no binding. (C) Binding of b-BSA on thesurfaces shown in A. Each incubation step starts at 0 min; rinses inbuffer are indicated (arrow heads).

Figure 4. QCM-D responses, Δf and ΔD, for the adsorption ofnative BSA to gold (black squares) and to silica (blue triangles).Incubation starts at 0minwith concentrations of 20μg/mLongoldand 10 mg/mL on silica. Rinses in buffer are indicated (arrowheads).

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We used b-BSA to determine if the immobilized moleculesretain their biotin-binding activity. To reduce and exclude anyunspecific binding, the surface was incubated with native BSAprior to exposing the biotinylated target. Interestingly, the bind-ing of native BSAwas low (|Δf| = 1.0 Hz for SAv, 3.0 Hz for Av,and<1.0 Hz for NAv; Figure 2B). Because BSA adsorbs readilyto bare gold (Figure 4),33 we conclude that the biotin-bindingproteins themselves confer rather good resistance to unspecificprotein binding. In contrast, biotinylated BSA bound stably toAv, SAv, and NAv, with binding rates similar to those found onSLBs (Figure 2C, Table 1). The final frequency shifts did varyfrom-12.5Hz for SAv to-15.5Hz and-18Hz forAv andNAv,respectively. Differences in the arrangement of b-BSAmost likelyexplain these variations: given BSA’s elongated shape, adsorptionin a slightly tilted orientation would easily result in a few Hz ofadditional frequency shift.15

Silica Surfaces. The binding behavior of SAv on silicadiffered strikingly from that observed on gold: no adsorptioncould be observed (Figure 3A). Adsorption was pronounced forAv and NAv (Figure 3A), with adsorption rates and maximumshifts in frequency and dissipation similar to those observed forrespective protein on gold (Table 1). In contrast to gold, however,a fraction of the proteins desorbed upon rinsing with buffer,indicating that binding was not entirely stable.

The lack of adsorption of SAv was unexpected and meritedfurther examination. To this end, we tested the effect of the buffersolution on binding. A large range of pH was sampled (3.5, 4.5,5.5, 6.5, 8.0, 9.0, and10.0) at a salt concentration of 150mMNaCl,and no significant adsorption could be found (|Δf|e 0.5 Hz, datanot shown). The use of Hepes instead of PBS at a similar ionicstrength (150 mM NaCl) and pH did not result in any significantbinding. Variation of the NaCl concentration, in the range of 0 to200 mMNaCl, at neutral pH resulted in no adsorption except forvery low ionic strength. The highest frequency shift of-6 Hz wasattained inHepes buffer withoutNaCl. Even in this case, however,most of the adsorption was reversible, with only-2 Hz remainingafter rinsing (data not shown), corresponding to less than 7%of aSAv monolayer on gold.

The exposure of Av- and NAv-covered surfaces to b-BSAresulted in bindingwith final frequency shifts of-14 and-16Hz,respectively (Figure 3C, Table 1). Native BSA did not bind(Figure 3B), confirming that the immobilized biotin receptorsretained their binding activity.

We stress that b-BSA did not bind to a bare silica surface(Figure 3C). Even native BSA showed very low adsorption(Figure 3B). This may at first be surprising, given that BSA iswidely used as a surface-blocking agent. Our observation, how-ever, is consistent with previous findings.34 It should also benoted that the BSA concentrations that are typically applied forblocking are 1000-fold higher than the concentrations employedhere.35,36 Indeed, when incubating BSA at 10 mg/mL (1% BSAsolution), we found frequency shifts of about-30Hz, confirmingthe formation of a stable protein layer of about 5 nm thickness(Figure 4).

Discussion

We have investigated the immobilization of streptavidin andother biotin receptors to different surfaces. Two main observa-

tions are particularly remarkable. First, SAv as well as Av andNAv bound well to gold and retained their biotin-binding activityafter physisorption. Second, SAv was uniquely resistant toadsorption to silica.Physisorption of Biotin Receptors ontoGold Surfaces.We

found all investigated biotin receptors to adsorb spontaneouslyand with high coverage on gold surfaces. SAv and Av wereparticularly well suited for surface functionalization in the sensethat they formed stable, well-defined monolayers. Tests with themodel protein BSA revealed that the biotin-receptor monolayerswere efficient at preventing unspecific protein binding. Physisorp-tion has so far only rarely been employed to functionalize surfaceswith biotin-binding receptors.20,21a,21b Gold is an interesting andwidely used material for biotechnological applications, includingbiosensors.37 Our results demonstrate that the simple strategy ofphysisorption is more attractive than its crudeness may at firstsuggest. Although unspecific adsorption is likely to lead toadsorption in various orientations, a large fraction of the im-mobilized molecules retained their biotin-binding activity. Weremind the reader, however, that detection schemes that rely onfluorescent detection may find a drawback in the direct immobi-lization of biotin receptors to gold:38 the limited distance betweenthe gold surface and fluorescent biotinylated targets can poten-tially quench the fluorescence.Absence of SAv Binding to Silica Surfaces. No significant

responsewas observed byQCM-Dupon exposure of SAv to silicaover a wide range of pHand ionic strength. Given that the limit ofdetection of our setup is better than 0.5 Hz and considering thatthe solvent contributes about 80% to the measured mass at lowsurface coverage of SAv,15 we can derive an upper limit for thebound protein mass of 2 ng/cm2, or about 1% of a completemonolayer.

Whatmay explain the absence of binding for SAv as comparedto the strong binding of Av and NAv? In the simplest considera-tion, one may solely consider mean-field electrostatic and van derWaals interactions (DLVO theory39). Av has an isoelectric pointof 10.540 and thus carries a net positive charge at neutral pH.Electrostatic interactions would thus promote the adsorption ofAv onto the negatively charged silica surface, which was indeedobserved. The fact, however, that NAv readily bound to silicawhereas SAvdid not, althoughbothhave similar isoelectric points(6.341 and 6.4,42 respectively), argues against electrostatics as thekey interaction that prevents the binding of SAv. Also, SAv didnot adsorb even at pH values between 3.5 and 6, a range in whichsilica and SAv would be expected to carry opposite net charges.A rigorous comparison of themagnitude of the surface charges ofSAv and silica as a function of pH and ionic strength43 and of thebinding behavior of SAv on other metal oxide surfaces, such astitanium,may provide further insight into the role of electrostaticsin SAv’s resistance to binding but is outside the scope of thiswork.Apart from these mean-field considerations, discrete molecularinteractions and steric effects may of course also be at play.39

To the best of our knowledge, the absence of unspecificinteraction between SAv and silica has not yet been reported inthe literature. One earlier work, however, contrasts our finding:

(33) Reimhult, K.; Petersson, K.; Krozer, A. Langmuir 2008, 24, 8695–8700.(34) Svedhem, S.; Pfeiffer, I.; Larsson, C.;Wingren, C.; Borrebaek, C.; H€o€ok, F.

ChemBioChem 2003, 4, 339–343.(35) Pierce Biotechnology; Instructions for Blocker BSA (prod. number 37525).(36) Hedge, P.; Qi, R.; Abernathy, K.; Gay, C.; Dharap, S.; Hughes, J. E.;

Snesrud, E.; Lee, N.; Quackenbush, J. Biotechniques 2000, 29, 548–550.

(37) Knoll, W. Annu. Rev. Phys. Chem. 1998, 49, 569–638.(38) Liebermann, T.; Knoll, W. Colloids Surf., A 2001, 171, 115–130.(39) Israelachvili, J. N. Intermolecular and Surface Forces, 2nd ed.; Academic

Press: London, 1992.(40) Savage, M. D. Avidin-Biotin Chemistry: A Handbook.; Pierce Chemical Co.:

Rockford, IL, 1992.(41) Pierce Biotechnology, Instructions for NeutrAvidin (prod. number 31000).(42) Techniques in Immunocytochemistry; Bullock, G. R., Petrusz, P., Eds.; Aca-

demic Press: London, 1983; Vol. 2, p 236.(43) Parsegian, V. A.; Gingell, D. Biophys. J. 1972, 12, 1192–1204.

1034 DOI: 10.1021/la902226b Langmuir 2010, 26(2), 1029–1034

Article Wolny et al.

Bushan et al.20 observed the binding of FITC-labeled SAv tosilica. UsingQCM-D, we could not detect any significant adsorp-tion to cleaned silica (data not shown). The presence of thefluorescent dye alone thus cannot explain the binding. Instead,we propose that variations in the surface preparation explain thedifferent adsorption behavior: the surfaces employed by Bushanet al. exhibited a contact angle of around 40�whereas our cleanedsurfaces showed complete wetting. Indeed, the preparation andcleaning of surfaces have previously been found to affect proteinadsorption behavior strongly.44

It should be pointed out that the resistance of silica to SAvbinding over a wide range of experimental conditions suggests avery simple yet promising strategy for the biofunctionalization ofnano- ormicropatterned structures that are based on silicon tech-nology. Defined surfaces areas covered, e.g., with gold or anothermaterial/linker with an affinity for SAv could be selectivelyfunctionalized with SAv while leaving remaining surface areasempty and ready for alternative functionalization.

Conclusions

Wehavedemonstrated that thephysisorption ofbiotin-bindingproteins, such as streptavidin and avidin, to gold presents a simpleway to equip a surface with a high density of biotin-binding sites.The functionalized surfaces were stable and exhibited little, if any,unspecific binding of BSA. Given the plethora of availablebiotinylated biomolecules, such a surface represents an interesting

immobilization platform. Physisorption is attractive for its sim-plicity and shouldbe a valuable alternative to established yetmorecomplex strategies that require a linker layer between the solidsupport and the biotin receptor. However, it remains rather crudein that the orientation of the biotin-binding sites on the surface isnot well controlled.

In stark contrast to gold, streptavidin did not adsorb to silicaover a wide range of pHand ionic strength. This contrast suggestsa particularly attractive application of physisorption as a simpleroute for the selective functionalization of nano- or microstruc-tures of gold on silicon-based devices. Such devices include small-scale probes (microcantilevers and atomic force microscopy tips)and nanostructured surfaces,45-47 which are increasingly used forbiophysical studies and biotechnological applications.

Acknowledgment. R.P.R. gratefully acknowledges fundingfrom the Department of Industry of the Basque Governmentand support from theGermanFederalMinistry of Education andResearch (BMBF, project 0315157). Support from the Max-Planck-Society is also acknowledged. We thank Ilya Reviakine(CIC biomaGUNE) for fruitful discussions.

(44) Ying, P.; Yu, Y.; Jin, G.; Tao, Z. Colloids Surf., B 2003, 32.

(45) Hanarp, P.; Sutherland, D. S.; Gold, J.; Kasemo, B. Colloids Surf., A 2003,214, 23–36.

(46) Glass, R.; Arnold,M.; Bl€ummel, J.; K€uller, A.;M€oller,M.; Spatz, J. P.Adv.Funct. Mater. 2003, 13, 569–575.

(47) Dahlin, A.; Z€ach, M.; Rindzevicius, T.; K€all, M.; Sutherland, D. S.; H€o€ok,F. J. Am. Chem. Soc. 2005, 127, 5043–5048.

(48) Keller, C. A.; Glasm€astar, K.; Zhdanov, V. P.; Kasemo, B. Phys. Rev. Lett.2000, 84, 5443–5446.