Partial Denaturation of Silica-Adsorbed Bovine Serum Albumin

Using TREWIFS, the adsorption behavior of 1-anilinonaphthalene-8-sulfonic acid (ANS) complexed with. BSA has been investigated as a function of penetr...
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Langmuir 2002, 18, 9924-9931

Partial Denaturation of Silica-Adsorbed Bovine Serum Albumin Determined by Time-Resolved Evanescent Wave-Induced Fluorescence Spectroscopy Levie Lensun, Trevor A. Smith, and Michelle L. Gee* School of Chemistry, University of Melbourne, Parkville, 3010, Victoria, Australia Received May 20, 2002. In Final Form: September 18, 2002 Time-resolved evanescent wave-induced fluorescence spectroscopy (TREWIFS) has been used in the study the molecular conformation of bovine serum albumin (BSA) adsorbed at the silica/aqueous interface. Using TREWIFS, the adsorption behavior of 1-anilinonaphthalene-8-sulfonic acid (ANS) complexed with BSA has been investigated as a function of penetration depth of the evanescent wave. Restriction of the fluorescent probe’s motion as a function of distance from the interface was also studied by time-resolved evanescent wave-induced fluorescence depolarization measurements. The results are consistent with a model of a partial protein denaturation: at the surface, an adsorbed BSA molecule unfolds, thus optimizing protein-silica interactions and the number of points of attachment to the surface. Further away normal to the surface, the protein molecule maintains its coiled structure.

Introduction Recently, much interest has been focused on the study of molecular interactions and dynamics of adsorbed species at an interface.1 These areas are of great importance to the understanding of a variety of phenomena, including catalysis, lubrication, corrosion, and biocompatibility. For example, in the area of biotechnology and biocompatibility, understanding protein structural dynamics on surfaces, with a view to the development of bioassays and materials suitable for bodily implants, is of primary interest.2 As a consequence, there has been increasing interest in the development of powerful methodologies to enhance and extend surface characterization. Protein adsorption is a widespread event occurring in a vast range of systems from the purely biological to industrial. For example, protein adhesion to the surface of biomedical implants, i.e., bioadhesion or biofouling, can have an adverse effect on a wide range of biomedical devices and prosthetics, in some cases leading to surfaceinduced thrombosis.3 Filtration systems, transport systems, storage systems, and ship hulls are also prone to the destructive effects of biofouling. The strong tendency to adsorption exhibited by proteins is exploited in areas such as food science and the cosmetics industry, where proteins are used as stabilizers and emulsifiers. Whenever a surface is exposed to an aqueous solution containing protein, there is, in most cases, a strong driving force to protein adsorption. At a charged interface, the electrostatic attraction between the surface and the oppositely charged functional groups along the polypeptide chain can be the dominant driving force to adsorption. If electrostatic interactions are weak or absent, there exists the ubiquitous entropic driving force to the adsorption of a protein. This arises due to the hydrophobic effect4 that * Author to whom correspondence should be addressed. E-mail: [email protected]. (1) de Mello, A. J.; Elliott, J. A.; Rumbles, G. J. Chem. Soc., Faraday Trans. 1997, 93, 4723-4731. (2) Crystall, B.; Rumbles. G.; Smith, T. A. J. Colloid Interface Sci. 1993, 155, 247-250. (3) Kidane, A.; Lanyz, G. C.; Jo, S.; Park, K. J. Biomater. Sci., Polym. Ed. 1999, 10, 1089-1105. (4) Tanford, C. In The Hydrophobic Effect; Wiley: New York, 1980.

drives the nonpolar portions of the protein molecule to the surface, to the exclusion of the water. Once at a surface, the protein molecule can rearrange and often denature so as to maximize favorable interactions with both the surface and solvent. Denaturation results in a maximization of favorable protein surface interactions and, in some cases, an increase in the protein’s flexibility compared to the molecule in its native state, thus increasing the protein’s conformational entropy.5 Despite the importance of the surface activity to many systems, little is known about the details of protein adsorption and the resulting adsorption-induced conformational changes. Studies involving the use of indirect experimental methods such as the measurement of dynamic surface tension and surface pressure have provided some information on protein adsorption.6-9 However, with these techniques, information on protein conformation must be inferred. Photon correlation spectroscopy (PCS)10-12 and specular neutron reflection (SNR)13-15 have also been used to study the conformation of adsorbed proteins. However, in both PCS and SNR, data must be interpreted via model fits. Additionally, in SNR, it is necessary to deuterate the water. This limits hydrogen bonding between the protein and the solvent, which might impact on the native structure of the protein and its conformation when adsorbed. (5) Magdassi, S.; Kamyshyny. Surface Activity and Functional Properties of Proteins. In Surface activity of Proteins: Chemical and Physicochemical Modifications; Magdassi, S., Ed.; Marcel Dekker: New York, 1996. (6) Tornberg, E. J. Colloid Interface Sci. 1978, 64, 391-402. (7) Tripp, B. C.; Magda, J. J.; Andrade, J. D. J. Colloid Interface Sci. 1995, 173, 16-27. (8) Wang, J.; McGuire, J. J. Colloid Interface Sci. 1997, 185, 317323. (9) Ybert, C.; di Meglio, J. M. Langmuir 1998, 14, 471-475. (10) Dalgleish, D. G. Colloids Surf. 1990, 46, 141-155. (11) Dickenson, E.; Robson, E. W.; Stainsby, G. J. Chem. Soc., Faraday Trans. 1 1983, 79, 2937-2952. (12) Leaver, J.; Horne, D. S. J. Colloid Interface Sci. 1996, 181, 220224. (13) Lu, T. J.; Su, T. J.; Thirtle, P. N.; Thomas, J. K.; Rennie, A. R.; Cubitt, R. J. Colloid Interface Sci. 1998, 206, 212-223. (14) Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F.; Penfold, J. J. Phys. Chem. B 1998, 102, 8100-8108. (15) Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F. J. Phys. Chem. B 1999, 103, 3727-3736.

10.1021/la020473e CCC: $22.00 © 2002 American Chemical Society Published on Web 10/30/2002

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where E0 is the intensity of the electric light field at the interface and the penetration depth, Λ, is defined as the distance into medium 2 normal to the interface at which the electric field amplitude of the evanescent wave has decreased to 1/e of its interface value. Λ is related to the wavelength of the excitation radiation, λ, the incident angle of the light, θi, the refractive index of medium 1, n1, and the critical angle for the pair of media, θc, through eq 2, viz.:

Λ)

λ 1 4πn1 (sin2 θ - sin2 θ )1/2 i c

(2)

In addition, E0 is also related to the angle of incidence and relative refractive indices of the two media constituting the interface, viz.: Figure 1. Schematic representation of the TIR principle and the resulting standing evanescent wave. Λ is the penetration depth of the evanescent field (defined in text) which is related to the angle of incidence of the excitation light, θi, relative to the normal. x is the distance away from the interface.

The use of nonlinear optical effects (in particular second harmonic and sum-frequency generation) at surfaces and interfaces is another means of achieving a high degree of surface specificity.16-20 These methods can provide useful information regarding orientation and order of species at an interface. Another attractive way to probe interfacial phenomena is through the utilization of total internalreflection-based spectroscopic methods in which the penetrating evanescent field is used to photoexcite an absorbing species placed within the penetrating field.21 These techniques can be used at the interface between many phases (e.g., liquid/solid, liquid/liquid, solid/solid, etc.). Evanescent wave and second harmonic generation methods have even been combined.22 The basic principles behind evanescent wave spectroscopy are illustrated schematically in Figure 1 and are described below. When light transmitted in a medium of refractive index, n1, encounters a medium of lower refractive index, n2, two process can occur: when the angle of incidence is less than the critical angle, θc, defined by Snell’s law as θc ) sin-1(n2/n1), refraction occurs; for angles of incidence greater than θc, total internal reflection occurs. The name total internal reflection (TIR), however, is a misnomer, as it implies that all of the light remains within the more refractive medium. This is not the case, as predicted by Maxwell’s equations of electromagnetic radiation,23 since a small amount of the light field penetrates the lower refractive index medium in the form of a standing wave known as an evanescent wave. The intensity of the evanescent wave, E, decays exponentially with distance from the interface, x, according to eq 1

E ) E0 exp(-x/Λ)

(1)

(16) Bain, C. D. J. Chem. Soc., Faraday Trans. 1995, 91, 1281-1296. (17) Paul, H. J.; Corn, R. M. J. Phys. Chem. B 1997, 101, 4494-4497. (18) Zhang, D.; Gutow, J.; Eisenthal, K. B. J. Phys. Chem. 1994, 98, 13729-13734. (19) Zhao, X.; Subrahmanyan, S.; Eisenthal, K. B. Phys. Rev. Lett. 1991, 67, 2025-2028. (20) Kikteva, T.; Star, D.; Leach, G. W. J. Phys. Chem. B 2000, 104, 2860-2867. (21) Harrick, N. J. In Internal Reflection Spectroscopy; WileyInterscience: New York, 1967. (22) Conboy, J. C.; Daschbach, L. J.; Richmond, G. L. J. Phys. Chem. 1994, 98. (23) Rumbles, G.; Brown, A. J.; Phillips, D. J. Chem. Soc., Faraday Trans. 1991, 87, 825-830.

E0 )

4 cos2 θi 1 - (n2/n1)2

(3)

Variable angle absorption-based TIR techniques24-28 either rely on spectral changes or are otherwise restricted to intensity information that is related to the concentration and extinction coefficient of the absorbing species within the evanescent field. Experiments based on attenuated TIR absorption spectroscopy can suffer from limited absorption-attenuation of the transmitted beam and hence limited sensitivity, but this inherent insensitivity is usually improved by the use of multiple reflections. TIR techniques can be extended to the fluorescence domain: if the absorbing species is also fluorescent, the evanescent field can induce fluorescence in the interfacial region, the intensity, If, of which can be described by

If ∝

∫0∞ φf(x)c(x)I0 exp(- Λx ) dx

(4)

where φf is the fluorescence quantum efficiency of the fluorophore, c(x) is the fluorophore concentration profile in the x direction, and I0 is the fluorescent intensity at x ) 0. The total internal reflection fluorescence technique, or evanescent wave-induced fluorescence spectroscopy (EWIFS),23 is a reasonably surface-specific method with increased sensitivity over transmission-based techniques. EWIFS permits a single reflection geometry with which to probe the interfacial region, although a multiple internal reflection fluorescence geometry has been used in the early work.29 This combination of the evanescent wave principle with conventional fluorescence spectroscopy30,31 takes advantage of the inherent multidimensionality and sensitivity of fluorescence-based techniques. The process of emission yields information that is intrinsically related to the nature of the fluorophore and its immediate microenvironment.32 EWIFS has been used to characterize the interfacial properties of many species, ranging from (24) Neivandt, D. J.; Gee, M. L.; Tripp, C. P.; Hair, M. L. Langmuir 1997, 13, 2519-2526. (25) Neivandt, D. J.; Gee, M. L. Langmuir 1995, 11, 1291-1296. (26) Trau, M.; Grieser, F.; Healy, T. W.; White, L. R. J. Chem. Soc., Faraday Trans. 1994, 90, 1251-1259. (27) Pirinia, A.; Sung, C. S. P. Macromolecules 1991, 24, 6104-6109. (28) Tronin, A.; Blasie, J. K. Langmuir 2001, 17, 3696-3703. (29) Harrick, N. J.; Loeb, G. I. Anal. Chem. 1973, 45, 687-691. (30) Masuhara, H.; Mataga, N.; Tazuke, S.; Murao, T.; Yamazaki, I. Chem. Phys. Lett. 1983, 100, 415-419. (31) Ausserre, D.; Hervet, H.; Rondelez, F. Macromolecules 1986, 19, 85-88. (32) Byrne, C. D.; de Mello, A. J.; Barnes, W. L. J. Phys. Chem. B 1998, 102, 10326-10333.

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small dye molecules to macromolecules such as proteins, polymers, enzymes, and DNA.1,33-36 The surface specificity of the evanescent wave has also been successfully used in many biosensing applications.1 Steady-state EWIFS shares some similar limitations to absorption-based TIR methods. Principally, if there are no spectral changes on adsorption, then the steady-state fluorescence signal can only contain intensity information that will be proportional to both the fluorophore concentration and the fluorescence quantum yield. In much steady-state EWIFS work to date, it has been assumed that the fluorescence quantum efficiencies are known and invariant with distance normal to the interface. Under these interpretation constraints, the EWIF intensity measurements are related directly to the surface excess of an adsorbed species. However, some controversy exists in the area of EWIFS concerning the effects of the surface itself on the photophysical properties of the fluorophore and, more specifically, the fluorescence quantum yield.23 It must be considered that even a “simple” interface will almost certainly contain more than one type of fluorescent center,1 thus providing a heterogeneous environment for molecular species.37 Emission characteristics, such as fluorescence quantum yield, will therefore be dependent on the precise location of individual fluorophores within this environment. Thus, the interpretation of steady-state measurements is restricted, limiting their value.37 The limitations of steady-state EWIFS can be overcome by resolving the emission both spectrally and temporally, a fact that we have exploited in the present work. The incorporation of time resolution into EWIFS, i.e., TREWIFS (time-resolved evanescent wave induced fluorescence), can be achieved with either frequency-domain38 or, more widely used, time-domain methods. Time-resolved fluorescence spectroscopy, and so TREWIFS, has the potential to resolve some of the limitations of steady-state measurements since any changes in the photophysical properties of the fluorophore will be reflected in the fluorescence decay time profile.23 Furthermore, unlike steady-state measurements in which the fluorescence quantum yield of surface adsorbed species is assumed to be equal to that in the bulk solution,31 TREWIFS has the capability to detect any variation of fluorescence quantum yield as a function of distance away from the interface.23,37 This can be done through varying the angle of incidence of the excitation light in order to vary the penetration depth, Λ, of the evanescent wave, as detailed above (see eq 2). Thus, wheng a fluorophore is monitored within an adsorbed layer, the time-resolved fluorescence decay profiles monitored as a function of distance normal to the surface enable depth profiling of the adsorbed species. TREWIFS can be extended to the use of time-resolved fluorescence anisotropy measurements, TRAMS, which enables the motion of macromolecular species on the nanosecond and sub-nanosecond time scales to be monitored with high sensitivity. In principle, the fluorescence decay profile from a fluorophore is monitored, following excitation with vertically polarized light, through polarizers set parallel and perpendicular to the polarization of (33) Liebmann, L. W.; Robinson, J. A.; Mann, K. G. Rev. Sci. Instrum. 1991, 62, 2083-2092. (34) Yao, H.; Ikeda, H.; Kitamura, N. J. Phys. Chem. 1998, 102, 76917694. (35) Parsons, D.; Harrop, R.; Mahers, E. G. Colloids Surf. 1992, 64, 151-160. (36) Watarai, H.; Funaki, F. Langmuir 1996, 12, 6717-6720. (37) de Mello, A. J.; Crystall, B.; Rumbles, G. J. Colloid Interface Sci. 1995, 169, 161-167. (38) Lundgren, J. S.; Bekos, E. J.; Wang, R.; Bright, F. V. Anal. Chem. 1994, 66, 2433-2440.

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the excitation light, i.e., IVV(t) and IVH(t), respectively. The time-resolved fluorescence anisotropy function, r(t), given by

r(t) )

IVV(t) - IVH(t) IVV(t) + 2IVH(t)

(5)

contains information on the motion of the fluorophores on the time scale of the lifetime of the excited state. Essentially, fluorescence depolarizing processes such as chromophore/segmental rotation, energy migration, etc., are monitored. If the photoexcitation is achieved via a penetrating evanescent field, then chromophore motion within the interfacial region is directly monitored, which can provide information on the binding (through rotational dynamics) of an adsorbate at an interface. Furthermore, such binding can be monitored as a function of distance from the interface simply by varying the penetration depth, Λ, of the field. There is vast potential for TRAMS coupled with variable angle, evanescent wave excitation to provide spatial information on the fluorescence depolarizing processes near an interface. However, despite this potential, remarkably few such studies have been undertaken.39-43 An excellent review discussing time-resolved techniques has been published by Bright.44 In this paper we use variable angle TREWIFS and evanescent wave-induced TRAMS to investigate the rotational dynamics and molecular conformation of bovine serum albumin (BSA) adsorbed from aqueous solution onto a silica surface, in situ, and as a function of distance normal to the solid/solution interface. Serum albumins are blood serum proteins of considerable interest since they rapidly form a covering layer on synthetic polymers. This is of a special interest with regards to biocompatibility. Control over this type of nonspecific adsorption of serum proteins, as well as a number of other related deposition processes, must start with an understanding of the nature of protein adsorption. The time-resolved fluorescence decay and protein rotational dynamics behavior were probed using 1-anilinonaphthalene-8-sulfonic acid (ANS) which is a fluorescent probe whose fluorescence decay kinetics depend on the probe’s microenvironment such that, in a polar microenvironment, the fluorescence is rapidly and efficiently quenched. ANS associates with BSA noncovalently by partitioning into hydrophobic pockets within the protein’s tertiary structure where the fluorescence decay of the probe is relatively slow. Experimental Section Instrumentation. The EWIFS apparatus was constructed in-house according to the arrangement shown in Figure 2 and based on the hemicylindrical prism designs reported previously.1,39,45,46 This arrangement is especially useful for depth profiling experiments since this configuration leads to the incident and totally internally reflected light entering and exiting the prism normal to the surface of the cylindrical part of the prism, regardless of the angle of incidence relative to the interfacial (39) Rumbles, G.; Bloor, D.; Brown, A. J.; de Mello, A. J.; Crystall, B.; Phillips, D.; Smith, T. A. Time-Resolved Evanescent Wave Induced Fluorescence Studies of Polymer-Surface Interactions. In Microchemistry: Spectroscopy and Chemistry in Small Domains; Masuhara, H., Schryver, F. C. D., Kitamura, N., Tamai, N., Eds.; North-Holland Delta Series; Elsevier Science B.V.: London, 1994; pp 269-286. (40) Fukumura, H.; Hayashi, K. J. Colloid Interface Sci. 1990, 135, 435-442. (41) Wirth, M. J. Appl. Spectrosc. 1993, 47, 651-653. (42) Piasecki, D. A.; Wirth, M. J. Langmuir 1994, 10, 1913-1918. (43) Wirth, M. J.; Burbage, J. D. Anal. Chem. 1991, 63, 1311-1317. (44) Bright, F. V. Appl. Spectrosc. Rev. 1997, 32, 1-43. (45) Hamai, S.; Tamai, N.; Masuhara, H. J. Phys. Chem. 1995, 99, 4980-4985.

Time-Resolved Evanescent Wave Study of BSA

Figure 2. Schematic representation of the in-house built EWIFS optical arrangement, used for time-resolved EWIFS and EW-TRAMS. region. This helps minimize spurious reflections, which could interfere with the measurements and also causes minimal deviations (refraction) of the beam and hence aids positional stability. The hemicylindrical substrate used was specially constructed (Herbert Groiss Co., Melbourne, Australia) of fused silica (refractive index of 1.46). The height of the prism is 3.0 with a radius of curvature 2.00 cm. The rest of the sample compartment consisted of a standard PMMA fluorescence cuvette that was modified by drilling a hole through one face (6 mm diameter) in order for the bulk solution to be in a direct contact with the flat side of the hemicylindrical prism.23 The hemicylindrical prism thereby acts as the surface to which species can adsorb. The disposable cuvette was clamped against the prism to prevent leakage of the sample solution. The prism and the cell are mounted onto a compact xyz translation stage (OptoSigma) in order to allow accurate positioning of the point of evanescent excitation at the central pivot point of the cell, even when the angle of incidence is varied. All the previous components are mounted on a precision 360° rotational stage (OptoSigma) to allow the measurements as a function of penetration depth through variation of incident angle. The majority of the limited number of TREWIFS instruments reported have principally used the time-correlated single photon counting (TCSPC)47,48 technique, although frequency domain detection38 and the use of an image intensifier/optical multichannel analyzer system have also been used.49,50 TCSPC is a widely used tool, with exceptional sensitivity, dynamic range, and flexibility, and is ideally suited to TREWIFS; therefore, (46) Toriumi, M.; Yanagimachi, M. Time-Resolved Total-Internal Reflection Fluorescence Spectroscopy and its Applications to Solid/ Polymer Interface Layers. In Microchemistry: Spectroscopy and Chemistry in Small Domains; Masuhara, H., Schryver, F. C. D., Kitamura, N., Tamai, N., Eds.; North-Holland Delta Series; Elsevier Science B. V.: London, 1994; pp 257-268. (47) Lakowicz, J. R. In Topics in Fluorescence Spectroscopy; Plenum Press: New York, 1991; Vol. 1-3. (48) O’Connor, D. V.; Phillips, D. In Time Correlated Single Photon Counting; Academic Press: London, 1984.

Langmuir, Vol. 18, No. 25, 2002 9927 TCSPC was used here. The TREWIFS system used is summarized schematically in Figure 2 and described below. The excitation source is a mode-locked titanium/sapphire laser (Coherent Mira 900F-Innova 400) operating at ∼800 nm and a pulse repetition rate of 76 MHz. The output of this laser was pulse picked to ∼2 MHz using a TeO2 Bragg cell (Gooch & Housego) driven by synchronized electronics (CAMAC model CD5000) and frequency doubled to ∼400 nm. The vertically polarized beam was incident on the sample prism at an angle of 69.0° relative to the detection axis, as shown in Figure 2, and focused loosely onto the interfacial region. The emission was imaged with a wide angle telephoto camera lens (Nikon) with a polarizer (King) mounted in front. The emission was passed through suitable filters and onto the slit of a 1/4 m monochromator (CVI Digikrom CM110). The fluorescence was detected by a microchannel plate photomultiplier (Eldy, St. Petersburg, Russia, model EM1-132) and recorded using conventional TCSPC electronics described elsewhere.51 Fluorescence decay data were collected to 20 000 counts at the channel of maximum intensity. For emission anisotropy measurements, the rotatable emission polarization analyzer was manually aligned at 0° and 90° relative to the vertically polarized excitation to measure fluorescence decays with polarization parallel, IVV(t), and perpendicular, IVH(t), to the excitation. Data were transferred from the multichannel analyzer (MCA) to a computer where subsequent analysis of the decays was performed using nonlinear least-squares iterative fitting.52 The experimentally obtained IVV(t) and IVH(t) decays were used to generate the time-resolved anisotropy, r(t) function (eq 5). The instrumental correction factor, G, which is usually included to take into account the bias of the detector system (in particular due to the monochromator) to the polarization was not determined here and assumed to equal unity. Materials and Cleaning Methods. Prior to each experiment, the hemicylindrical prism was cleaned by washing with copious amounts of acetone. The optical material was then further cleaned by soaking in a warm ammonical peroxide (50:50 of hydrogen peroxide and Milli-Q water with the addition of a few milliliters of concentrated ammonia) solution to oxidize any residual contaminants noncorrosively.53 The prism was then washed again in copious amounts of Milli-Q water. All of the chemical reagents were used without any further purification. The BSA and ANS were purchased from Sigma Chemical Co. The solvent used was acetate buffer solution, which has been made by mixing 0.0037 M of sodium acetate solution with 0.0070 M of acetic acid glacial solution to give overall 0.01 M buffer at pH 5, without any added electrolyte. All experiments were carried out at room temperature (approximately 20° C) following an equilibration time of approximately 3 h. Experimental Protocols. The spectroscopic absorption and emission of ANS and BSA-bound ANS in bulk solution were characterized using a Varian Cary 50 Bio steady-state absorption spectrometer and a Cary Eclipse fluorimeter, respectively. Emission spectra were background subtracted and corrected for detector and monochromator response. The excitation wavelength was 385 nm, and emission was scanned between 400 and 600 nm. The excitation and emission bandwidths were set to 2.5 nm, and the PMT voltage was set to 800. Steady-state measurements were performed using ANS at a constant concentration of 1 × 10-5 M as a function of different BSA concentrations, varying from 10 to 100 ppm. For the TREWIFS and TRAMS, solutions of ANS and BSA (BSA concentration ∼100 ppm) were prepared in buffered solution (49) Schneckenburger, H.; Stock, K.; Eickholz, J.; Strauss, W. S. L.; Lyttek, M.; Sailer, R. Time-Resolved Total Internal Reflection Fluorescence Spectroscopy: Application to the Membrane Marker Laurdan. In Proc. SPIE, Laser microscopy; Ko¨nig, K., Tanke, H. J., Schneckenburger, H., Eds.; SPIE: Bellingham, WA, 2000; Vol. 4164, pp 36-42. (50) Schneckenburger, H.; Sailer, R.; Stock, K.; Lyttek, M.; Strauss, W. S. L. Total Internal Reflection Fluorescence Lifetime Imaging (TIRFLIM) of Living Cells. Multidimensional Microscopy 2001, 3rd AsiaPacific International Symposium on Confocal Microscopy and Related Technologies, 2001, Melbourne, Australia. (51) Ghiggino, K. P.; Smith, T. A. Prog. React. Kinet. 1993, 18, 375436. (52) Smith, T. A.; Irwanto, M.; Haines, D. J.; Ghiggino, K. P.; Millar, D. P. Colloid Polym. Sci. 1998, 276, 1032-1037. (53) Gee, M. L. Ph.D. Thesis, University of Melbourne, 1987.

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and contained in the modified plastic cuvette clamped to the hemicylindical prism. The solution was allowed to be in direct contact with the silica substrate for 3 h, after which it is assumed that adsorption was complete and that the fluorescence decay can be observed with little variation. TREWIFS experiments were carried out as a function of angle of incidence to allow depth profiling of the probe. Two sets of experimental conditions were used, referred to as type A and type B. Type A measurements were performed on BSA-ANS adsorbed from solution, maintaining equilibrium with BSAANS in the bulk. A set of fluorescence decay profiles was obtained for a variety of angles of incidence ranging from 69° to 89° with 4° increments. Hence, the penetration depth ranged from 100 to 40 nm (eq 2). The type B measurements were performed after the bulk solution of BSA-ANS had been replaced with acetate buffer solution to remove an nonadsorbed or loosely adsorbed BSA. This removal was done by slowly flushing the cell with buffer solution (with the volume of 10 times of cell’s volume), while simultaneously removing the residual BSA solution by pipet. This procedure ensured that the silica surface was always in contact with aqueous solution. Once the protein solution was replaced with the buffer, emission decays were taken at the angles of incidence specified above. The TRAMS experiment was carried out also as a function of penetration depth of the evanescent wave, but only in the presence of bulk BSA-ANS solution. To characterize the anisotropy of nonadsorbed BSA, TRAMS were also performed in bulk solution with the sample solution contained in a standard silica quartz cuvette, and the emission was collected in the usual right angle geometry. A full description of the instrumentation used for the bulk measurements is given elsewhere.52

Results and Discussion ANS is an extraordinarily sensitive fluorescent probe for determining the microscopic viscosity and polarity of chemical and biological systems. It is well-known that under aqueous conditions, ANS is virtually nonfluorescent (the quantum yield, φf ) 0.004) due to an efficient fluorescence quenching mechanism.54 A decrease in polarity of the ANS environment leads to both an increase in the fluorescence lifetime (and a concomitant increase in fluorescence quantum yield, φf ) 0.98) and a slightly blue shifted fluorescence maximum.54 Despite the widespread interest in this probe and the volume of literature relating to it spanning several decades, there is still an ongoing debate regarding the origin of the remarkable sensitivity of this and related probes to their environment. Monophotonic photoionization was attributed as the most likely dominant nonradiative path in polar solvents by Robinson and co-workers,55 but other explanations have also been postulated, as discussed by Ebbesen and Ghiron.56 These include intersystem crossing to the triplet state, variation in molecular conformation, intramolecular charge transfer, and specific solvent-solute interactions. Most recently, the fluorescence quenching was attributed to polarity-dependent twisted intramolecular charge transfer (TICT).57 While the actual fluorescence quenching mechanism in ANS is not the subject of the present paper, the mechanism may play a role in determining how ANS probes the adsorption behavior of BSA to hydrophobic surfaces from aqueous solution. It is well documented that neither unbound ANS nor BSA alone fluoresce significantly in buffered aqueous solution under the excitation and emission wavelengths (54) Hlady, V.; Andrade, J. D. Colloids Surf. 1988, 32, 359-369. (55) Robinson, G. W.; Robbins, R. J.; Fleming, G. R.; Morris, J. M.; Knight, A. E. W.; Morrison, R. J. S. J. Am. Chem. Soc. 1978, 100, 714550. (56) Ebbesen, T. W.; Ghiron, C. A. J. Phys. Chem. 1989, 93, 71397143. (57) Das, K.; Sarkar, N.; Nath, D.; Bhattacharya, K. Spectrochim. Acta, Part A 1992, 48A, 1701-5.

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Figure 3. Time-resolved fluorescence decay curves of the BSAANS complex as a function of penetration depth, Λ, of the evanescent wave normal to the interface. Two data sets are presented, corresponding to type A and to type B experimental protocols, as described in the Experimental Section. The penetration depth was varied from around 40 nm to around 100 nm within each data set. The data all show that the fluorescence decays more rapidly the smaller the penetration depth of the evanescent field.

used here (λexc ∼ 390 nm, λem ∼ 490 nm). When no BSA is present, no hydrophobic regions exist into which the ANS can partition, and the emission of ANS is quenched very efficiently. BSA is known to intrinsically fluoresce around 344 nm in bulk solution and ∼333 nm in close proximity to surfaces54 when excited in the ultraviolet region (i.e., following direct excitation of the amino acids, especially tryptophan). This spectral shift of the fluorescence is believed to occur on adsorption of BSA onto a hydrophilic surface, due to the BSA molecules reconfiguring to “hide” their tryptophans in nonpolar “pockets”. This mechanism is different from heat denaturation of BSA in solution, where the tryptophan fluorescence maximum shifts to 350 nm upon disruption of the secondary and tertiary structure.54 When the fluorescence of BSA-bound ANS is measured, it is observed that as the concentration of BSA in bulk solution is increased, the fluorescence quantum yield of the ANS increases dramatically. This is a direct result of the increase in the number of hydrophobic regions per unit volume as more BSA is added to solution, thus protecting the ANS from the fluorescence quenching mechanism in aqueous environments. There is negligible spectral shift of the ANS emission profile upon complexation with the protein or upon the adsorption of the BSAANS complex to the silica surface. This lack of spectral change restricts the information provided by steady-state fluorescence measurements since the observed intensity changes are dependent on both the ANS and BSA concentrations, and the fluorescence quantum yield. TREWIFS measurements in variable angle mode were recorded from silica-adsorbed BSA-ANS complex as a function of excitation angle relative to the interface (and hence penetration depth). These TREWIFS measurements were performed under two experimental conditions, referred to as type A and type B, the details of which are outlined in the Experimental Section. The TREWIFS results for each are shown in Figure 3 (λexc ) 400 nm, λem ∼ 470 nm). In type A measurements, the fluorescence decay profiles were measured from surface adsorbed BSAANS complex adsorbed from buffered bulk solution. At the highest angle of incidence employed, the depth of the penetration (i.e., Λ ) 40 nm) will still be longer than the thickness of the interfacial region. Therefore, in these

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measurements, the fluorescence intensity not only will have a contribution from the surface region but also will contain a significant proportion of emission from any free BSA-ANSBSA-ANS complex remaining in bulk solution (referred to as “bulk” in Figure 3). In type B measurements, however, nonadsorbed and any reversibly adsorbed protein was slowly flushed out of the cell. Even though the evanescent field penetrates beyond the interfacial region, the bulk buffer solution will not contribute to the fluorescence intensity, assuming that, after the cell is flushed, any remaining BSA-ANS complex is irreversibly adsorbed at the interface. Hence, in type B measurements, the fluorescence decays are comprised solely of the signal from the surface-adsorbed form of the BSA-ANS complex (referred to as “surface” in Figure 3). The observation of measurable fluorescence intensity under these conditions confirms that the surface excess of BSA remains relatively large, even after flushing the cell. The decay profiles were significantly different to those of the bulk solution, and these decays were reproducible over several hours, indicating that the remaining BSA is, indeed, irreversibly adsorbed. The fluorescence decay profiles of the BSA-bound ANS recorded under both the surface and “bulk” conditions are dependent upon the penetration depth (Figure 3). In the case of a single fluorophore, the fluorescence quantum efficiency of the fluorophore is defined by

φf )

kr ) k r τf kr + knr

(6)

where kr is the radiative deactivation rate coefficient, knr is the nonradiative deactivation rate coefficient, and τf is the fluorescence lifetime. None of the decay profiles follows a single-exponential profile, but rather the decays can be parameterized by the sum of two or three exponential terms. In the case of multiple individual emitting species, the fluorescence quantum yield of each contributing component with fluorescence decay time, τi, and fractional contribution, Ai, is given by:

φi )

Aiτi n

(7)

Ajτj ∑ j)1 In this work we do not attempt to assign any physical significance to the individual components resulting from multiexponential fitting but, rather, compare the relative total emission quantum yield of each of the decay profiles. The total fluorescence quantum yield is proportional to the integrated area under the fluorescence decay curve. Differences between the fluorescence quantum yield of BSA-ANS in the bulk solution and that of BSA-ANS near the surface are clearly observed by a significant shortening of the fluorescence decay curves recorded at shallower penetration depths as compared with those recorded from the bulk solution. The decay curves collected under type B conditions show that the fluorescence decay is most rapid when the penetration depth, Λ, is smallest. This implies that ANS within the adsorbed layer experiences a more polar microenvironment the closer the ANS is to the silica surface. Indeed, the longer the penetration depth, the more hydrophobic the microenvironment for ANS; that is, we see a gradient of hydrophobicity, normal to the interface, experienced by the ANS. Interestingly, fluorescence quantum yields are more often observed to increase when the fluorophore is

adsorbed on a surface compared to bulk solution, due to the restriction of the fluorophore’s vibrational motion.23,58 Our results clearly illustrate that the fluorescence quantum yield cannot be assumed to be constant as a function of distance from the interface, confirming the findings of other TREWIFS measurements23 and again illustrating the limitations of steady-state measurements. Assuming that kr, the radiative deactivation rate coefficient, is independent of environment, then eq 4 above can be rewritten as

If ) kr

∫0∞ τf(x)c(x)I0 exp(- Λx ) dx

(8)

suggesting that a distribution of decay times might be a more appropriate model for interpreting the fluorescence decay profiles than a sum of a discrete number of exponential terms; however, this was not attempted in this work. The change in fluorescence decay observed as a function of incident angle under the “bulk” (type A) conditions is attributed to the contributions in quantum yield variation from the surface-adsorbed species inherent in the decays of the bulk. The adsorption of BSA onto silica has been investigated in detail previously by EWIFS methods using the intrinsic fluorescence of BSA59 and a variety of fluorescence probes including ANS.2,40,54,60-62 The behavior of ANS itself near interfaces has also been investigated.63,64 The work to date has indicated that, upon adsorption to silica, BSA undergoes some degree of conformational change.59,62 Hlady et al.54,60,61 present data that indicate the presence of some sort of surface aggregates at high BSA surface concentrations, in agreement with the existence of two-dimensional aggregates of adsorbed BSA concluded by others. These authors also interpret a change in the distance between energy donors and acceptors as being due to conformational changes (partial unfolding of adsorbed BSA) upon adsorption.60 Fukumura and Hayashi40 also reported that BSA undergoes conformational change upon adsorption to a silica surface. In addition, Hlady and Andrade54 reported that the conformation change involves the tryptophan group of BSA becoming embedded further within more hydrophobic regions of the coil, while the ANS probe groups experience a more polar environment (hence fluorescence quenching) through increased exposure to more hydrophilic regions. Our time-resolved fluorescence decay results, which illustrate a hydrophobicity gradient in the adsorbed layer normal to the surface, are also consistent with the existence of a conformational change of BSA upon adsorption to the silica interface. We provide here the additional information regarding the dependence of the fluorescence decay on penetration depth of the evanescent field, Λ. This allows the conformation of the adsorbed protein to be resolved spatially, normal to the surface. The observed decrease in fluorescence quantum efficiency at small Λ may be explained by the BSA unfolding thus exposing bound-ANS groups to a more polar environment than that experienced by the BSA-ANS complex in bulk (58) Bell, M. A.; Crystall, B.; Rumbles, G.; Porter, G.; Klug, D. R. Chem. Phys. Lett. 1994, 221, 15-22. (59) Rainbow, M. R.; Atherton, S.; Eberhart, R. C. J. Biomed. Mater. Res. 1987, 21, 539-555. (60) Hlady, V.; Andrade, J. D. Colloids Surf. 1989, 42, 85-96. (61) Suci, P.; Hlady, V. Colloids Surf. 1990, 51, 89-104. (62) Burghardt, T. P.; Axelrod, D. Biochemistry 1983, 22, 979-985. (63) Bessho, K.; Uchida, T.; Yamauchi, A.; Shioya, T.; Teramae, N. Chem. Phys. Lett. 1997, 264, 381-386. (64) Hlady, V.; Go¨lander, C.; Andrade, J. D. Colloids Surf. 1988, 33, 185-190.

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solution. It appears that BSA unfolds to the greatest extent at the silica surface so as to optimize its interaction with silica, since further away from the silica surface (as Λ increases) there is a corresponding increase in fluorescence decay time. This indicates that further away from the silica surface, an adsorbed BSA molecule is increasingly more coiled and so retains increasingly more hydrophobic sites where ANS resides. That is to say, BSA only partially denatures upon adsorption and the unfolding that does occur is a direct result of the optimization of BSA-silica interactions. Given the strong binding of BSA at the silica surface, it is possible that two competing processes affecting the fluorescence quantum yield of the ANS are working in opposition against each other: there is a reduction in fluorescence quantum yield due to exposure of ANS to a more hydrophilic microenvironment, as discussed above, but in addition, there might also be some restriction of vibrational motion, which tends to increase quantum yield. Our measurements can only show the net result of these two opposing effects, but nonetheless, we believe our interpretation still holds. An explanation raised by Hlady and Andrade60 is that the binding of ANS to surface-adsorbed BSA may be weaker compared with the binding of the same probe to BSA in solution. If this were the case, then the bound BSA may release some of the ANS probe to the hydrophilic environment and a very marked shortening of the decay profile would then be expected. This is in contrast to our observations. Time-Resolved Fluorescence Anisotropy Measurements (TRAMS). The difficulties in analyzing fluorescence decay profiles of fluorophores near surfaces are well documented, and until further work in this area definitively solves this problem, the value of fluorescence decays per se may be limited. Nonetheless, it is clear that time-resolved measurements are more informative than steady-state measurements alone. Fluorescence anisotropy measurements, while also suffering from drawbacks such as poorer signal quality and complexity of analysis, have the potential to provide additional qualitative and quantitative information on dynamic processes that can contribute to a loss of emission polarization including energy transfer, migration, and fluorophore/segmental rotation motion. As mentioned above, despite the potential of time-resolved fluorescence polarization measurements coupled with variable angle, evanescent wave excitation to provide information on these fluorescence depolarizing processes near an interface, it is surprising how few such studies have been undertaken. Evanescently induced TRAMS (EW-TRAMS) should aid in identifying whether or not the changes observed in the fluorescence decay profiles (quantum yields) of BSA-ANS as a function of penetration depth are in fact related to partial denaturation of the protein upon adsorption, thus exposing the ANS to more aqueous environments. TRAMS were conducted on complexed BSA-ANS in bulk solution (BSA concentration 100 ppm) as described in the Experimental Section. These data are shown in Figure 4 as the curve labeled “bulk”. Analyzing data for the anisotropy measurements can be very difficult, as is well documented elsewhere. For the case of a fluorophore with a single fluorescence decay time, free to rotate through a limited angular range (within a cone of semiangle θ) attached to a large particle that undergoes isotropic rotational diffusion, the anisotropy decay kinetics

Lensun et al.

Figure 4. Time-resolved fluorescence anisotropy profiles of BSA-ANS complex recorded as a function of depth penetration, Λ, of the evanescent wave.

are often well described by a double exponential decay model, i.e.,

r(t) ) r0[β1 exp(-t/τc1) + β2 exp(-t/τc2)]

(9)

This model adequately describes the decay of the fluorescence anisotropy from the BSA-ANS in the current context, implying that the ANS exhibits at least two modes of rotational motion. The two resulting rotational correlation times, τC1 and τC2, were found to be ∼2 and ∼40 ns, respectively. Using the Stokes-Einstein Equation, viz:

τc ) Vmη/RT

(10)

where Vm is the molar volume of the macromolecule-probe conjugate assuming its shape to be spherical, η is the viscosity of the solution, R is the ideal gas constant, and T is the absolute temperature. Assuming that the protein adopts a spherical shape, the 40 ns correlation time corresponds to a hydrodynamic radius of the protein of ∼3.7 nm (using the viscosity 0.00894 P at 20 °C). This is in good agreement with the value for the Stokes radius quoted from the supplier of the BSA (3.48 nm) suggesting that the longer decay component is related to the overall molecular motion of the BSA-ANS coil. This result indicates that at concentrations as high as 100 ppm, the BSA does not appear to be aggregating to any large degree in bulk solution. The shorter component (τC1 ∼ 2 ns) is tentatively attributed to a segmental motion of the BSA peptide chain with complexed ANS chromophores. The overall rotational motion of the BSA-ANS is composed of rapid rotational motion of the probe superimposed upon that of the entire protein molecule. EW-TRAMS measurements on the surface-adsorbed BSA-ANS complex were conducted as a function of penetration depth, under the type B conditions discussed above. The resulting time-resolved fluorescence anisotropy profiles from these experiments are also shown in Figure 4. These decay profiles are not well described by the simple model accounted for by eq 9. It is well-known that fluorescence anisotropy decays can become very complex to analyze even for relatively simple systems,52 and often the quality of the data does not justify the use of complex analysis functions. Anisotropy measurements are further complicated under EWIFS conditions by requiring the simultaneous analysis of all four permutations of the polarization orientations (IVV, IVH, IHV, and IHH), unlike conventional (bulk) polarized excitation, as discussed elsewhere.39 Briefly, the E-field penetration depth is

Time-Resolved Evanescent Wave Study of BSA

independent of the polarization of the incident light; however, the electric field amplitudes at the interface are dependent upon the polarization. Within the evanescent standing wave, a component of the E-field exists in each of the three spatial directions, in contrast with a conventional traveling light wave where only two components exist. In light of these complications, in this work, the time-dependent fluorescence anisotropy decays have not been analyzed analytically, but rather qualitatively at this stage. Another possible complication arises if the adsorbed species is not isotropic. Certain adsorbed species might exhibit spatial alignment relative to the surface leading to anisotropy of the absorption and emission dipole moments.41,43 However, in the BSA-ANS system, this effect is minimized since the partitioning of the ANS into the BSA coil is not likely to show any preferred orientation relative to the plane of the surface. Additionally, anisotropy only affects the values of r(t) and has little or no effect in the time domain, i.e., the rate of depolarization. It is the relative rate of depolarization that we are scrutinizing in the present work. It is clear from the data shown in Figure 4 that as the depth of penetration of the evanescent wave is reduced, the fluorescence anisotropy decays far more slowly than is the case in bulk solution. This indicates that as the protein adsorbs onto the surface, the motion of the parts of the protein to which the ANS is bound is more restricted, as might be expected. Neither the specific short nor long rotational components reported above for BSA-ANS in bulk solution are evident in the decays from the surfaceadsorbed measurements. This indicates that the BSA coil as a whole is undergoing restricted rotation and the more rapid segmental motion of the ANS-bound segments of the protein referred to above is severely hindered. This corroborates the interpretatation of the EWIFS data, discussed above: The shortening of the fluorescence decay profiles observed (Figure 3) does not appear to be the result of a simple opening up of the BSA coil conformation on adsorption, since this might be expected to increase the time scale of the motion of the individual ANS-bound BSA segments rather than decrease it, as observed. One mechanism by which these observations may be explained is an unraveling of the BSA coil at the silica surface leading to an increase in the number of points of attachment. Any ANS bound to portions of the BSA that experience this uncoiling and adsorption to the surface will consequently

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exhibit restricted rotation due to the inhibited coil and chain segment motion. This restriction of the chromophore motion supports the findings discussed above relating to the TREWIFS measurements. Any detachment of the ANS from the BSA upon adsorption, as previously suggested,49 would result in a very marked shortening of the fluorescence decay, and this emission, which may be too weak to be monitored by the EW-TRAMS experiment anyway, would in any case be depolarized very rapidly, contrary to our observations. Conclusions We have shown that variable angle, time-resolved, evanescent wave-induced fluorescence decay measurements and EW-TRAMS can be applied to the study of adsorption-induced protein denaturation and that TREWIFS and EW-TRAMS are able to spatially resolve protein molecular conformational changes normal to the surface. Specifically, we have successfully studied the extent to which BSA denatures upon adsorption at a silica surface, using ANS as a fluorescent probe. When adsorbed, BSA adopts a more “open” structure through which it exposes the BSA-ANS binding sites to a slightly more hydrophilic environment (including the silica surface). The shortening of the fluorescence decay as the emission closer to the interface is probed (Figure 3) is most likely due to a slight decrease in hydrophobicity experienced by the ANS as the protein denatures. The variation of fluorescence quantum yield with distance from the interface suggests a hydrophobicity gradient experienced by the ANS in the region of the interface. The EW-TRAMS shows an increase in the overall time scale of the motion of the protein segments to which the ANS is bound as the emission is probed from closer to the interface. The results are consistent with a model of a partial protein denaturation: at the surface, an adsorbed BSA molecule unfolds, thus optimizing protein-silica interactions and the number of points of attachment to the surface. Further away normal to the surface, the protein molecule maintains its coiled structure. Acknowledgment. The authors would like to thank the Australian Research Council for their generous financial support of the work in the form of an ARC Large Grant. LA020473E