Coverage-Dependent Orientation of Lysozyme Adsorbed on Silica

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Coverage-Dependent Orientation of Lysozyme Adsorbed on Silica Susan M. Daly,† Todd M. Przybycien,†,‡ and Robert D. Tilton*,† Departments of Chemical Engineering and Biomedical Engineering, Center for Complex Fluids Engineering, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213 Received October 13, 2002. In Final Form: February 1, 2003

We use total internal reflection fluorescence (TIRF), streaming current, and optical reflectometry measurements, under a variety of solution conditions, to examine the mechanism of the chicken egg lysozyme reorientation on silica surfaces, originally proposed by Robeson and Tilton (Robeson, J. L.; Tilton, R. D. Langmuir 1996, 12, 6104). The TIRF data suggest that a two-stage reorientation occurs in the lysozyme layer during adsorption. The first stage involves a reorientation to optimize lysozyme interaction with the charged surface and to reduce lateral repulsions between adsorbed protein molecules. This stage occurs when the adsorbed layer reaches a critical surface coverage and indicates the importance of lateral interactions between adsorbed proteins. Moreover, the reorientation rate depends on both the bulk protein concentration and the fluid wall shear rate, indicating that incoming protein molecules from the bulk solution also participate in the reorientation. The preferred orientation puts the active site face toward the solution and the most positively charged amino acid patch into contact with the negative silica surface. A second stage of slow restructuring occurs as the adsorbed layer coverage asymptotically approaches saturation and corresponds to reorientation of the protein molecules that fill surface vacancies made accessible by the first reorientation. Consistent with this proposed two-stage reorientation mechanism, streaming current measurements indicate that adsorbed lysozyme initially causes a significant interfacial potential reversal, followed by a slow relaxation to less positive ζ-potentials as the lysozyme optimizes its orientation. Accordingly, the desorbable fraction of the adsorbed layer after reorientation is significantly smaller than that before reorientation.

Introduction The performance of biomedical devices that come into contact with tissues or biological fluids, ranging from drug delivery devices to contact lenses, tissue culture scaffolds, and biosensors, depends critically upon protein-surface interactions. In these diverse applications, it is the structure of the adsorbed layer that governs its biochemical function and its impact on the technology. It is important to recognize that the “layer structure” extends beyond protein conformation to include orientation relative to the surface, two-dimensional organization, and local electric field gradients. Several techniques have recently been used to determine the preferred orientation of adsorbed proteins, including neutron reflection,2 streaming potential,3 surface force apparatus,4 and vibrational sum frequency spectroscopy.5 Interesting structural changes may occur during the initial stages of protein adsorption, and these changes are often quite rapid.1,6 Techniques that directly report details of the time-dependent structure of the adsorbed layer yield difficult to obtain mechanistic information that ultimately should allow for prediction of structural and biofunctional changes upon adsorption. * Corresponding author. E-mail: [email protected]. † Department of Chemical Engineering. ‡ Department of Biomedical Engineering. (1) Robeson, J. L.; Tilton, R. D. Langmuir 1996, 12, 6104. (2) Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F.; Penfold, J. Langmuir 1998, 14, 438. (3) Ethe`ve, J.; De´jardin, P. Langmuir 2002, 18, 1777. (4) Blomberg, E.; Claesson, P. M.; Forberg, J. C.; Tilton, R. D. Langmuir 1994, 10, 2325. (5) Kim, G.; Gurau, M.; Kim, J.; Cremer, P. S. Langmuir 2001, 18, 2807. (6) Norde, W.; Anusiem, A. C. I. Colloids Surf. 1992, 66, 73.

We previously proposed a dynamic reorientation mechanism for lysozyme adsorption to silica, based on total internal reflection fluorescence (TIRF) experiments using protein labeled with a pH-sensitive fluorophore.1 Here we further test the proposed mechanism for consistency with electrokinetic measurements that are also responsive to protein orientation effects, and also by subjecting the fluorescence model that we use to interpret TIRF results to a wide variety of solution conditions. This investigation has revealed the rich dynamics of the interplay between lateral interactions among adsorbed proteins and interactions among adsorbed proteins and proteins in solution near the surface. We use TIRF,7-11 optical reflectometry,12-16 and streaming current measurements17,18 to decipher the initial orientational dynamics of fluorescein isothiocyanate (FITC) labeled chicken egg white lysozyme at the silica-water (7) Lok, B. K.; Cheng, Y.; Robertson, C. R. J. Colloid Interface Sci. 1983, 91, 104. (8) Lok, B. K.; Cheng, Y.; Robertson, C. R. J. Colloid Interface Sci. 1983, 91, 87. (9) Axelrod, D. Rev. Biophys. Bioeng. 1984, 13, 247. (10) Hlady, V.; Reinecke, D. R.; Andrade, J. D. J. Colloid Interface Sci. 1986, 111, 555. (11) Rondelez, F.; Ausserre, D.; Hervet, H. Annu. Rev. Phys. Chem. 1987, 38, 317. (12) Tilton, R. D. Scanning Angle Reflectometry and its Application to Polymer Adsorption and Coadsorption with Surfactants; Dubin, P., Farinato, R., Eds.; Wiley: New York, 1999; pp 331-363. (13) Velegol, S. B.; Tilton, R. D. Langmuir 2001, 17, 219. (14) Schaaf, P.; De´jardin, P.; Schmitt, A. Langmuir 1987, 3, 1131. (15) Dijt, J. C.; Cohen Stuart, M. A.; Hofman, J. E.; Fleer, G. J. Colloids Surf. 1990, 51, 141. (16) Dijt, J. C.; Cohen Stuart, M. A.; Fleer, G. J. Adv. Colloid Interface Sci. 1994, 50, 79. (17) Braem, A. D. Colloidal and Interfacial Phenomena in Polymer/ Surfactant Mixtures; Carnegie Mellon University: Pittsburgh, 2001. (18) Braem, A. D.; Prieve, D. C.; Tilton, R. D. Langmuir, in press.

10.1021/la026690x CCC: $25.00 © 2003 American Chemical Society Published on Web 03/11/2003

Orientation of Lysozyme Adsorbed on Silica

interface. Lysozyme is a well-characterized model protein that resists major conformational change upon adsorption.1,19 The FITC fluorescence exhibits pH sensitivity20-22 and therefore electrostatic potential sensitivity1 that allows us to track its distance from the charged surface using the TIRF technique. Further, lysozyme’s intrinsically fluorescent, pH-insensitive tryptophan residues can be used to determine surface concentration10,23-25 in the TIRF system in the absence of significant structural changes. By simultaneously monitoring the emission from the tryptophan residues and from the FITC label as a function of time, we observe the interplay between the kinetics of adsorption and reorientation. Independent optical reflectometry adsorption experiments were used to validate the tryptophan fluorescence calibration. Streaming current measurements provided the surface potentials for silica substrates as a function of buffer ionic strength, a quantity needed in order to test the fluorescence model. By correlating variations in the ζ-potential upon adsorption with the TIRF observations, streaming current results supported the proposed reorientation mechanism. Experimental Section Buffers. All experiments were conducted in triethanolamine hydrochloride (TEA; Aldrich) buffers prepared with varying pH and ionic strengths according to the protocol outlined by Bates.26 This is a monovalent system. All buffers were refrigerated and used within a week of preparation. Proteins. We labeled chicken egg white lysozyme (Sigma) with FITC (Molecular Probes) according to the protocol outlined by Robeson and Tilton.1 The FITC labels lysine residues and the amino terminus. The average labeling ratio of lysozyme in stock solution was L ) 0.23 ( 0.05 fluorescein molecules per lysozyme molecule as determined by the ratio of the optical absorbance at 500 and 278 nm.1 For all experiments, the average labeling ratio was decreased to 0.04 by mixing the labeled stock lysozyme solutions with unlabeled lysozyme solutions. Labeled stock solutions were refrigerated and used within 4 days of labeling and purification. Schnaible et al. observed that only lysine residues 33 and 97 and the amino terminus are modified for monosubstituted lysozyme.27 Because our labeling conditions lead to only one label in every four proteins, we assume that only the most reactive primary amino group, the amino terminus, is labeled on average. The amino terminus is located on the face opposite the active site,28 and it resides within the most positively charged patch on the protein. Adsorption Substrates. TIRF, reflectometry, and streaming current experiments were conducted on quartz substrates (Bioelectrospec), surface-oxidized optical grade silicon wafers (Virginia Semiconductor), and acid-treated soda lime glass slides (Fisher Scientific), respectively. We cleaned all surfaces by first rinsing with RBS detergent (Pierce). After rinsing with MilliQpurified water (Millipore), the slides were soaked in Chromerge for 30 min. We purchased Chromerge cleaning solution, ACS grade hydrochloric acid, and ACS grade sulfuric acid from Fisher Scientific. Chromerge was prepared by mixing 25 mL of the Chromerge cleaning solution with 2.5 L of 2.5 N sulfuric acid. After an extensive water rinse, the slides were soaked in 6 M hydrochloric acid for 20 min and thoroughly rinsed with water (19) Arai, T.; Norde, W. Colloids Surf. 1990, 51, 1. (20) Chen, R. F. Arch. Biochem. Biophys. 1969, 133, 263. (21) Klugerman, M. R. J. Immunol. 1966, 95, 1165. (22) Yguerabide, J.; Talavera, E.; Alvarez, J. M.; Quintero, B. Photochem. Photobiol. 1994, 60, 435. (23) Roth, C. M.; Lenhoff, A. M. Langmuir 1995, 11, 3500. (24) Shibata, C. T.; Lenhoff, A. M. J. Colloid Interface Sci. 1992, 148, 469. (25) Buijs, J.; Hlady, V. J. Colloid Interface Sci. 1997, 190, 171. (26) Bates, R. G. Determination of pH Theory and Practice; Wiley: New York, 1964. (27) Schnaible, V.; Przybylski, M. Bioconjugate Chem. 1999, 10, 861. (28) Wilson, K. P.; Malcolm, B. A.; Matthews, B. W. J. Biol. Chem. 1992, 267, 10842.

Langmuir, Vol. 19, No. 9, 2003 3849 again. The slides were then soaked in 10 mM sodium hydroxide for 20 min and rinsed thoroughly with water. After treatment, slides were stored in MilliQ water until use. This treatment left the slides completely wettable by water. The reflectometry and TIRF substrates were cleaned and reused repeatedly. The surface-oxidized silicon wafers used in reflectometry underwent an additional treatment step. After every five treatments with the above procedure, the substrates were soaked in hydrofluoric acid for 10 s to remove the oxide layer from the surface. As a safety precaution, a fresh hydrofluoric acid antidote (Pharma Science) was kept within reach. The substrates were then rinsed, blown dry with a high-purity nitrogen jet, and baked in a 1000 °C oven for 20 min to produce a fresh 20-30 nm thick oxide layer on the surface. Before subjecting the soda lime glass slides that were to be used in streaming current experiments to the above treatment, we first soaked them in concentrated sulfuric acid for 16 h. This gives a surface chemistry that is very similar to that of pure silica.29,30 TIRF. The basic principles of TIRF8,23,31 have been described elsewhere. For this study, we used a modular Spex-Fluorolog-3 spectrofluorometer. A TIRF flow cell manufactured by Bioelectrospec was mounted into the fluorometer. The flow cell consists of a 73° dove-type quartz prism optically coupled to a quartz slide via glycerol. The flow cell is a 16 mm × 24 mm × (15-400 µm) rectangular slit, where the thickness can be varied by the choice of gasket thickness. All flows in this work were laminar; emitted light is collected from a region of the cell that is roughly 12 inlet diameters removed from the inlet, ensuring fully developed laminar flow. The excitation and emission wavelengths were λex ) 488 nm and λem ) 520 nm for FITC and λex ) 295 nm and λem ) 320 nm for tryptophan, with excitation and emission slit widths of 7 nm. The evanescent wave penetration depths were 83 and 138 nm for excitation at 295 and 488 nm, respectively. We used the internal calibration method of Roth and Lenhoff23 to convert the tryptophan fluorescence intensity to a lysozyme surface concentration. As discussed below, the FITC emission provides information about the local electrostatic potential in its immediate vicinity and therefore its proximity to the charged surface. By simultaneously monitoring both FITC and tryptophan emission, we obtain an instantaneous relationship between orientation and surface concentration, provided that structural changes do not perturb the wavelengths of maximum emission for either fluorophore. Tryptophan emission spectra were monitored to assess the extent of any possible tertiary structure change. To examine orientation dynamics, we use the pH-sensitive fluorophore approach that tailors the characteristic length scale for the TIRF surface sensitivity to match the thickness of an adsorbed protein layer. Here we only summarize the salient features of this technique, referring the reader to the original detailed discussion.1 Typically, the TIRF surface sensitivity arises from an evanescent wave generated at the solid-water interface. When the fluorophore emission depends on its degree of protonation (i.e., pH or electrostatic potential sensitivity) and the surface is charged, the characteristic length scale for changes in emission becomes the Debye screening length. The Debye screening length can be adjusted via the solution ionic strength to match the protein monolayer thickness. Thus, the electrostatic potential originating from the charged surface varies strongly over the thickness of the protein layer, and therefore the position of the fluorophore within the layer strongly influences its fluorescence emission. The penetration depth of the evanescent wave, ∼1000 Å, is much greater than the monolayer thickness or Debye length, ∼30 Å, so that orientational variations in the FITC emission intensity are not obscured by the spatial variation of the excitation intensity. FITC emission is sensitive to electrostatic potential. In particular, the fluorescence emission intensity decreases as the electrostatic potential surrounding the FITC molecule becomes increasingly negative due to a shift in the protonation state of (29) Rebar, V. A.; Santore, M. M. J. Colloid Interface Sci. 1996, 178, 29. (30) Fu, Z.; Santore, M. M. Colloids Surf., A 1998, 135, 63. (31) Roth, C. M.; Lenhoff, A. M. Langmuir 1993, 9, 962.

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the fluorophore. According to eq 1,1 the FITC emission intensity

F∝

∫

π/2

-π/2

×

[

{

( ) [

10-pH exp -4 tanh

]

(1 + sin R) 2dp

10-pKa dR exp -D

]}

ψ0J -κD(1 + sin R) exp 4RT 2

+ 10-pKa (1)

for a layer of randomly oriented protein molecules depends on the degree of dissociation of the FITC molecule and on the spatially varying electrostatic potential and evanescent wave intensity. In eq 1, dp is the evanescent wave penetration depth, κ is the reciprocal of the Debye length, J is Faraday’s constant, ψ0 is the surface potential, and pKa is the dissociation constant for FITC, previously determined to be 6.2 via a fluorescence titration for lysozyme-bound FITC.1 The derivation of eq 1 has been presented previously.1 The model treats the approximately ellipsoidal lysozyme molecule as a cylinder of diameter D, with a fluorophore located on its surface. To compare to the ellipsoidal protein, D is taken to match the short axis of the ellipsoid. Lysozyme is approximated as a 3 × 3 × 4.5 nm molecule, so we set D ) 3 nm. The cylinder is free to rotate around an axis parallel to the surface. Thus, the fluorophore may assume any position, represented by the angle R, on a circle of diameter D that rests on the adsorbing surface. Since none of the possible FITC labeling sites are on the “ends” of the lysozyme ellipsoid, it is unlikely that FITC labels will ever be much further than 3 nm from the surface. This equation allows the FITC fluorescence intensity to be computed at various adsorbed lysozyme orientations. The decaying electrostatic potential adjacent to the surface is approximated for simplicity by Gouy-Chapman theory.32 The shortcomings of this simplification were described previously,1 but it suffices to capture the key effects. Equation 2 applies to a layer of proteins in a fixed

( ) ( ) }

y 10 exp dp F∝ (2) ψ0 J 10-pH exp -4 tanh exp(-κy) + 10-pKa 4RT -pKa

{

orientation relative to the surface, such that the FITC sits a distance y from the surface. Reflectometry. Here we use optical reflectometry as an external check on the internal TIRF calibration procedure. Details of the optical reflectometry technique and the particular instrument used here have been described elsewhere.12,13,17,18,33,34 Conversion from raw reflectivity data to surface excess concentrations was based on a two-layer striated interface optical model, consisting of the oxide layer and the adsorbed layer, using dn/dc ) 0.18 cm3/g for the refractive index increment of lysozyme. For both TIRF and reflectometry adsorption experiments, the temperature was held constant at 25 ( 1 °C. Buffer was pumped through the flow cell for at least 30 min prior to baseline collection for all experiments. The wall shear rate for reflectometry experiments was identical to that used in TIRF experiments. Streaming Current. We use streaming current as an independent test of the reorientation mechanism. The streaming current refers to the steady-state electrical current that develops when a pressure-driven flow sweeps counterions along a charged surface. One of the family of electrokinetic phenomena,35 the streaming current is determined by the electrostatic potential at the plane of shear adjacent to the charged surface, that is, the ζ-potential. Instrumental and theoretical issues, particularly concerning the advantage of streaming current measurements (32) Evans, D. F.; Wennerstrom, H. The Colloidal Domain, 2nd ed.; Wiley: New York, 1999. (33) Furst, E. M.; Pagac, E. S.; Tilton, R. D. Ind. Eng. Chem. Res. 1996, 35, 1566. (34) Pagac, E. S.; Prieve, D. C.; Solomentsev, Y.; Tilton, R. Langmuir 1997, 13, 2993. (35) Hunter, R. J. Zeta Potential in Colloid Science; Academic Press: New York, 1981.

Figure 1. FITC fluorescence (solid line) and the intrinsic tryptophan fluorescence were gathered simultaneously in TIRF, for adsorption from 5 mM TEA buffer, pH 7.4, 10 µg/mL lysozyme solutions onto quartz. The surface concentration values (diamonds) were calculated from a calibration of the tryptophan fluorescence. The arrows indicate times at which desorption experiments were conducted. over streaming potential measurements to determine the ζ-potential, are described elsewhere.17,18 The ζ-potential may be calculated from the slope of the streaming current, Is, versus pressure drop, ∆P, according to18

Is )

-0ζwh ∆P µL

(3)

where w, h, and L are the width, height, and length of the rectangular slit flow cell, respectively, µ is the viscosity, is the relative permittivity, and 0 is the permittivity of a vacuum. The instantaneous ζ-potential may also be determined during adsorption at a constant pressure drop according to eq 3, albeit with less precision than the slope-based calculation.

Results and Discussion TIRF and Reflectometry. We monitored the FITC label emission as well as the intrinsic tryptophan emission for 20 min periods while labeled lysozyme adsorbed from a flowing solution to the quartz surface in the TIRF cell. Figure 1 shows an example of the simultaneously recorded FITC emission and tryptophan emission intensities, where the latter is converted to lysozyme surface concentration via the internal calibration. The solution in this experiment is pH 7.4, 5 mM TEA buffer containing 10 µg/mL lysozyme. The wall shear rate is 32 s-1, and the temperature is 25 °C. The overshoots in the FITC emission are the signatures of the reorientation. Unlike FITC, tryptophan fluorescence is not sensitive to the local electrostatic potential. Hence, the tryptophan signal, but not the FITC signal, can be used as a reporter for the lysozyme surface concentration. Initially, the surface fills rapidly with adsorbing lysozyme followed by a slower adsorption phase. It is clear from Figure 1 that there is no overshoot in the surface concentration corresponding to the maxima in the FITC fluorescence response. To characterize the mass transfer during adsorption in this flow cell, we varied the wall shear rate and measured the initial adsorption rate using the tryptophan calibration. Comparing the initial steady-state adsorption rate to that predicted by the well-known Leveˆque analysis,7 we calculated a diffusion coefficient of (1.24 ( 0.04) × 10-6 cm2/s for lysozyme, consistent with the previously measured value.1 Thus, under the conditions of this study, adsorption occurs in fully developed laminar flow and the initial rate of adsorption is transport-limited. Figure 2 shows a comparison of the surface concentrations determined by TIRF tryptophan emission with internal calibration and by reflectometry for adsorption under identical conditions. The reflectometry data have


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Figure 2. Adsorption kinetics of lysozyme adsorbed from 5 mM TEA buffer, pH 7.4, 10 µg/mL solutions, at a shear rate of 32 s-1 for reflectometry (solid line) and calibrated TIRF (circles) experiments. TIRF has a poorer signal-to-noise ratio due to the fairly weak tryptophan fluorescence.

Figure 3. The ζ-potential of acid-treated and cleaned bare glass microscope slides decreases in magnitude with increasing buffer ionic strength. The solid line is generated from GouyChapman theory at a constant 800 Å2/charge. Error bars represent the standard deviation for multiple experiments.

a higher signal-to-noise ratio than that of the TIRF data. Otherwise, the data from the two techniques are consistent. The appeal of the simultaneous monitoring of FITC and tryptophan emission is that it eliminates uncertainties due to subtle differences in surface chemistry, solution composition, and flow conditions between independent TIRF and reflectometry experiments. The most notable features in the FITC emission profile in Figure 1 are the sharp maximum at ∼100 s followed by a minimum and a second broad maximum. Previous investigation of this system showed that the sharp overshoot was consistent with a reorientation of lysozyme on the silica surface.1 The reorientation puts lysozyme’s most positively charged amino acid patch into contact with the negatively charged silica surface. The amino terminus is located in this patch, so we assume the patch is home to the FITC label. (In the event that FITC randomly labels residues 33 and 97 and the amino terminus, the FITC distribution on lysozyme is still biased toward this positive patch.) When the FITC is closer to the negative surface, the local electrostatic potential surrounding it is more negative and protonation of the carboxyl residues on fluorescein is favored. This decreases the FITC fluorescence emission intensity.1 Other phenomena that could possibly have triggered the FITC emission peak, including polarization effects associated with rotation of the fluorescence excitation dipole, concentration quenching, photobleaching, and competitive adsorption of labeled and unlabeled lysozyme, were previously ruled out.1 Extensive conformational changes could also potentially alter FITC fluorescence by altering the local dielectric and electrostatic environment. To probe possible changes in tertiary structure, we monitored the full tryptophan emission spectrum as a function of time during adsorption from 10 µg/mL lysozyme solutions in pH 7.4, 5 mM buffer. The wavelength of maximum emission did not shift, but rather remained constant at 345 nm over the course of the experiment. The wavelength of maximum emission of native lysozyme is 345 nm and that of unfolded lysozyme in 6 M guanidine hydrochloride is 351 nm.36 This indicates that lysozyme does not experience significant tertiary structure changes upon adsorption to silica. The first FITC emission maximum is followed by an emission minimum, followed in turn by a second broad maximum. This behavior is observed for all buffer ionic strengths less than 40 mM studied herein. The second maximum is attributed to reorientation of protein that

adsorbed after completion of the first reorientation. The interactions that drive the reorientation are small at the trough and increase as more protein adsorbs into voids created by the initial reorientation. Eventually, the interactions become strong enough to trigger additional reorientation. These interactions will be discussed in greater detail in subsequent sections. Experimental Tests of the Electrostatic Potential Model. The model described by eq 1 predicts the FITC fluorescence for a layer of randomly oriented lysozyme molecules, where the FITC label can be at any distance from the surface that is consistent with the 3 × 3 × 4.5 nm dimensions of the native molecule, lying with its long axis parallel to the surface with no bias toward any particular rotation angle. To model the random orientation, we set D ) 3 nm in eq 1. To predict the FITC emission intensity for the lysozyme orientation that juxtaposes the FITC against the surface, we set y ) 0 nm in eq 2. The surface potential ψ0 that figures prominently in the model depends on the ionic strength of the solution as described by Gouy-Chapman theory.32 Following common practice, we equate ψ0 to the bare surface ζ-potential measured before protein adsorption. The ζ-potentials of the glass slides at pH 7.4 are shown as a function of buffer ionic strength in Figure 3. The data are consistent with a constant surface charge density of 800 Å2/charge. Since the surface charge density does not change with varying TEA concentrations, we can rule out buffer salt adsorption. The FITC fluorescence during adsorption from 10 µg/ mL lysozyme solutions, in pH 7.4 buffers containing 5-40 mM TEA, is presented in Figure 4. The “peak-to-trough” intensity ratio (defined as the FITC intensity at the first maximum divided by the FITC intensity at the subsequent minimum) decreases with increasing ionic strength and disappears, becoming just a small shoulder in the time profile, by 40 mM ionic strength. The effect of ionic strength on FITC fluorescence during adsorption is not simple. The FITC fluorescence intensity change depends on how much the electrostatic potential surrounding the FITC molecule changes during the reorientation. This of course varies with ionic strength according to its effects on κ-1 and ψ0. As the ionic strength is increased from 5 to 40 mM, κ-1 decreases from 4.3 to 1.5 nm, and ψ0 decreases in magnitude from -80 to -40 mV. The surface potential and Debye length have opposing effects on the dynamic range of the FITC emission intensity: decreases in the magnitude of ψ0 depress the magnitude of the FITC fluorescence change as a function of distance from the surface, while shorter Debye lengths increase the magnitude of the FITC fluorescence change

(36) Morgan, C. J.; Miranker, A.; Dobson, C. M. Biochemistry 1998, 37 (23), 8473.

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Figure 4. Normalized FITC fluorescence during adsorption from pH 7.4, 10 µg/mL lysozyme solutions shows a decrease in the first peak-to-trough ratio with increasing ionic strength. The shear rate is 32 s-1 for all experiments. The intensity is normalized to 1 and intentionally offset for illustration. Table 1. Electrostatic Model Predictions for the Peak to Trough Ratio Compared with the Experimental Values for the First FITC Peaka ionic strength (mM)

surface potential (mV)

experimental peak/trough ratios

electrostatic model predictions

5 20 30 40

-80 ( 4 -55 ( 3 -45 ( 2 -40 ( 2

1.44 ( 0.07 1.29 ( 0.1 1.12 ( 0.09 1

1.29 ( 0.03 1.2 ( 0.01 1.14 ( 0.009 1.12 ( 0.005

Figure 5. Surface concentrations determined by the tryptophan calibration show that more protein adsorbs at lower ionic strength. Circles represent adsorption from 5 mM buffer and squares represent adsorption from 40 mM buffer, both with the bulk protein concentration fixed at 10 µg/mL and pH 7.4. These data were gathered simultaneously with those presented in the FITC fluorescence plots shown in Figure 4.

a The experimental peak/trough ratios are reported as the mean ( standard deviation for multiple experiments. Errors in the model predictions are propagated from the errors in the surface potential.

versus distance by steepening the spatial gradient of the electrostatic potential. These competing effects motivate the use of the model to interpret the results. The experimental peak-to-trough ratios at various ionic strengths are listed in Table 1, along with the electrostatic model predictions. The predictions are calculated simply as the ratio of the fluorescence intensities expected for random orientation (calculated via eq 1 with D ) 3 nm) for the peak to the intensity for a fixed orientation that places FITC immediately adjacent to the surface for the trough (calculated via eq 2 with y ) 0 nm). Surface potentials for each ionic strength are taken from Figure 3. There is reasonably good quantitative and qualitative agreement between the model and the experimental results. Although the model presented considers a single labeling site on the protein, we determined that random labeling of residues 33 and 97 and the amino terminus only acts to mute, not eliminate, the predicted peak-totrough ratio trend. This model does not account for the complex distribution of dielectric constant and charged amino acids in the protein layer, and the notion that FITC moves from a random location to a fixed orientation with y ) 0 is certainly an oversimplification. Nevertheless, the model provides a satisfactory prediction of the FITC emission overshoot in the TIRF experiments for widely varying solution conditions. At 40 mM ionic strength, the overshoot in the FITC fluorescence response is reduced to a small shoulder, rather than a peak and trough, and is sometimes not observed at all in repeat experiments. Although the model predicts a smaller peak-to-trough intensity ratio, it does not predict the complete disappearance of the peak for 40 mM ionic strength. This requires further explanation. As the ionic strength increases to 40 mM, the amount of adsorption decreases in the pseudoplateau region (Figure 5). As will be shown below, the FITC overshoot only occurs when the surface concentration reaches a critical value.

Figure 6. The surface concentration Γ from TIRF tryptophan fluorescence at the time of the peak for various shear rates (circles) and solution concentrations (diamonds) stays constant. Adsorption experiments that did not reach a surface concentration of ∼1.6 mg/m2 did not show an overshoot in the FITC fluorescence. The bulk concentration varied from 1 to 20 µg/ mL, and the shear rate varied from 3 to 117 s-1.

In the 40 mM and higher ionic strength buffers, the surface concentration does not reach that critical value, and the reorientation does not occur; hence the experimental and model predictions differ as expected at higher ionic strengths. Component Interactions that Drive the Initial Reorientation. Interactions between the surface and the protein, between neighboring adsorbed proteins, and between adsorbed proteins and proteins in solution close to the surface can potentially participate in the reorientation mechanism. To examine the contribution of these various interactions to the driving force for lysozyme reorientation, we systematically varied the flux of protein to the surface by changing either the concentration of protein in the bulk solution or the wall shear rate during adsorption. The bulk concentration was varied from 1 to 20 µg/mL with the labeling ratio held constant. We also varied the shear rate from 3 to 117 s-1. Simultaneous monitoring of the FITC and tryptophan fluorescence emissions during adsorption at the various bulk concentrations and shear rates shows that the first peak in the FITC trace always occurs at the same surface concentration, regardless of solution or flow conditions, as shown in Figure 6. There is also no noticeable change in the surface coverage at the time of the first FITC peak,


Langmuir, Vol. 19, No. 9, 2003 3853

Figure 7. The FITC fluorescence adsorption time profiles for different values of the bulk lysozyme concentration and shear rate. In (a), the profiles during adsorption from pH 7.4 solutions of 1, 5, and 10 µg/mL lysozyme show that the sharpness of the peaks depends on the supply rate of protein to the surface. (c) shows the profile for shear rates 3, 32, and 117 s-1. The arrows indicate the direction of decreasing solution concentration and shear rates. (b) and (d) show the data from (a) and (c), respectively, where time is scaled by the time constant defined in eq 4.

over the range of ionic strengths used in our experiments. The value of the surface concentration at the time of the first FITC emission maximum, ∼1.6 mg/m2, is significant because it lies between the random parking limits37 for side-on (1.3 mg/m2) and end-on (1.9 mg/m2) lysozyme adsorption. At this critical surface concentration, lateral repulsions among neighboring protein molecules become severe and cause the proteins to reorient to put the most positively charged patch on the protein’s surface into contact with the negatively charged surface. Lateral interactions are critical for this reorientation to occur. Roth and Lenhoff23,31 have shown that the orientation of an isolated lysozyme molecule relative to a negatively charged surface significantly affects its adsorption energy. Yet, as the surface concentration becomes large, lateral interactions between neighboring proteins also contribute significantly to the adsorption energy landscape. Haggerty and Lenhoff38 have calculated the electrostatic potential contours within an array of lysozyme molecules that are confined to a monolayer. Their calculations indicate that the electrostatic energy is minimized when all the lysozyme molecules assume an orientation that places each molecule’s active site in a plane parallel to the monolayer. This orientation may even allow for favorable lateral interactions between adsorbed lysozyme molecules along one axis.38 An examination of the structure of lysozyme indicates that the most positively charged patch is on a face of the molecule that is directly opposite the face that contains the active site.1 Thus, the preferred orientation that we infer from our experimental observations, with the most positively charged patch juxtaposed against the quartz surface, is entirely consistent with the Haggerty and Lenhoff calculations, since it places the active site in a plane parallel to the surface. Furthermore, it allows for attractive electrostatic interactions between the most positively charged patch and the negatively charged surface. (37) Vigil, R. D.; Ziff, R. M. J. Chem. Phys. 1989, 4, 2599. (38) Haggerty, L.; Lenhoff, A. M. Biophys. J. 1993, 64, 886.

Time trajectories for the FITC emission intensity are shown for varying bulk protein concentration and shear rate in parts a and c of Figure 7, respectively. At higher bulk concentrations and shear rates, the initial peak in the FITC emission intensity occurs sooner. During the initial transport-limited regime, the FITC emission time profiles for different bulk concentrations or shear rates can be collapsed by scaling time in dimensionless form as given in eq 4 (see Figure 7b,d) where γ is the wall shear

τA )

tγ1/3D2/3c0 Γpeak × 1.86

(4)

rate, D is the diffusion coefficient, c0 is the bulk protein concentration, and Γpeak represents the critical surface concentration. The scaling is suggested by the Leveˆque solution to the convective-diffusive equation for adsorption from laminar flow in a rectangular slit.7 In the case of experiments conducted at constant bulk concentration but varying shear rate, the time scaling neatly collapses not only the initial FITC intensity data but also the FITC intensities after the peak. The data after the FITC peak in experiments conducted with varying bulk concentrations also collapse, but not as neatly as in the shear rate experiments. The sharpness of the peak, especially the rate of intensity decrease after the peak, reflects the rate of reorientation. If the reorientation were driven entirely by lateral protein-protein interactions at the critical surface concentration, the supply rate of protein to the surface should not influence the rate of the reorientation. Nevertheless, the FITC emission traces in Figure 7 show a dependence of the reorientation rate on the bulk concentration and shear rate: as the protein flux to the surface decreases, the rate of reorientation also decreases. This indicates that the reorientation involves both neighborneighbor interactions between adsorbed proteins and interactions between adsorbed proteins and proteins newly arriving from bulk solution. Clearly, the incoming bulk protein plays a role in propagating the reorientation.

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Figure 8. (a) The normalized FITC fluorescence (thin line) is overlaid with the ζ-potential (thick line) measured from streaming current. The two plots were gathered under identical adsorption conditions of pH 7.4, 10 µg/mL lysozyme solutions in 5 mM buffer. The shear rate was 2719 s-1. (b) The ζ-potential for adsorption from 10 µg/mL lysozyme in 40 mM buffer is shown. All ζ-potential data are smoothed with the bisquare technique in SigmaPlot, where the sampling proportion is 0.7 and the polynomial degree is 1.

ζ-Potential during Adsorption. The proposed reorientation mechanism is based on electrostatic arguments, where the most positively charged patch on the lysozyme molecule ultimately ends up juxtaposed to the surface. Given that the ζ-potential is the potential at the plane of shear, adjacent to the outer “edge” of the adsorbed layer, it will be sensitive to nonuniform charge distribution in the layer. Streaming current measurements, and their interpretation in terms of ζ-potentials, provide an independent tool by which to judge the proposed mechanism. ζ-Potential time trajectories were obtained during adsorption from 10 µg/mL lysozyme solutions in pH 7.4 buffers at ionic strengths ranging from 5 to 40 mM. We overlaid the ζ-potential results with FITC fluorescence time traces from TIRF experiments conducted with the same protein solution, at an identical wall shear rate. Because of the large pressure drops required in the streaming current measurements, the wall shear rate was rather high. Both streaming current data and additional TIRF data meant to complement streaming current results were collected at a wall shear rate of 2719 s-1. Overlaid data are shown in Figure 8. For all ionic strengths examined, lysozyme causes a net interfacial charge reversal (see Figure 8a,b). A similar effect has been detected by surface force apparatus measurements of lysozyme on mica.39 For protein in 5 mM buffer, the ζ-potential becomes less negative as the protein adsorbs, and it passes through 0 mV near the time that corresponds to the trough after the first FITC peak, where the surface concentration of protein is approximately 2 mg/m2. A complete charge reversal occurs, and the ζ-potential reaches a maximum of approximately +20 mV. The ζ-potential maximum nearly coincides with the second maximum observed in the FITC fluorescence. At this point, the ζ-potential then relaxes to less positive potentials. This is strong evidence that the second peak observed in the FITC fluorescence emission profile does in fact correspond to a reorientation in the lysozyme layer that buries lysozyme’s most positive patch against the silica surface. The initial neutralization can be simply attributed to the adsorption of net positively charged proteins. As described above, the second FITC emission maximum is attributed to continued random adsorption of lysozyme molecules (see Figure 1 for evidence of continuing adsorption) after the first reorientation, followed by a slower reorientation to the preferred side-on arrangement with the positive patch against the silica surface. This is entirely consistent with the ζ-potential time trajectory. Random (39) Tilton, R. D.; Blomberg, E.; Claesson, P. M. Langmuir 1993, 9, 2102.

addition of net positively charged proteins increases the ζ-potential such that it becomes less negative and ultimately becomes positive. Since the reorientation puts more of the positive charge close to the surface, further away from the plane of shear, the ζ-potential then decreases as a result of the reorientation. Thus, both FITC fluorescence and ζ-potential are mutually consistent with a preferred side-on orientation with the most positive patch down. Inspection of protein structure shows that this scenario leaves the active site up.1 We do not observe any notable changes in the rate of ζ-potential evolution at the time that corresponds to the initial reorientation. Effects of that reorientation are likely masked by rapid, continued adsorption of positively charged proteins. For high ionic strengths, the surface concentration never reaches the critical 1.6 mg/m2 value required for the production of a TIRF overshoot. As seen in Figure 5, the surface concentration reaches a plateau near 1.5 mg/m2, just below the critical surface concentration, upon adsorption from 40 mM buffer. Accordingly, the ζ-potential kinetics are far less interesting than at the lower ionic strengths. Under these conditions, the ζ-potential first increases and becomes slightly positive, but does not relax to a less positive value at long times. Likewise, there are no overshoots in the FITC emission, either at short or long times. This is strong evidence that there is no reorientation in the 40 mM ionic strength buffer because the surface concentration never reaches the critical value during the experiments. Other phenomena that could cause the observed ζ-potential maximum include the following: protein desorption; exchange of labeled, less positive protein for unlabeled protein; and counterion condensation. Labeled protein is less positively charged than unlabeled protein because the FITC molecule is negatively charged and the coupling of each FITC molecule consumes one primary amino group. Clearly, we have not protein desorption but rather continued protein adsorption. If exchange were responsible for the long-time relaxation of the ζ-potential, the FITC fluorescence would increase since labeled protein (less positively charged) would have to displace unlabeled protein (more positively charged) on the surface. Instead, the FITC fluorescence is decreasing during this time. Counterion condensation in the adsorbed layer can be ruled out because the average distance between charges is 35.6 nm at the most positive ζ-potential, and this is much greater than the Bjerrum length (0.71 nm). On the basis of the ζ-potential results, the second TIRF maximum most likely corresponds to a reorientation. Reversibility and Surface Optimization. The reorientation process is expected to be reversible if the transition to the preferred orientation is a thermodynamic


Figure 9. The FITC fluorescence intensity divided by the tryptophan fluorescence intensity is shown for increasing and decreasing surface coverage. Arrows pointing right correspond to adsorption of FITC-labeled lysozyme in 30 mM buffer, and the arrow pointing left corresponds to desorption into 30 mM buffer.

phase change. The adsorption/desorption reversibility for lysozyme on silica depends on the buffer ionic strength. As the buffer salt concentration is increased, the amount of protein that desorbs into pure buffer increases. The adsorption is completely reversible at ionic strengths greater than 150 mM. To probe the reversibility of the orientation change mechanism, we used a buffer rinse to induce desorption while monitoring both the FITC and tryptophan emission intensities. If the orientation change were reversible, the FITC-to-tryptophan emission intensity ratio would increase during desorption as the surface concentration fell below the critical surface concentration, that is, the behavior would be the reverse of that observed during adsorption. The key to the experiment was to identify an ionic strength at which the reorientation was observed during adsorption (i.e., less than 40 mM) but that was also sufficient to induce appreciable desorption. TEA buffer with 30 mM ionic strength satisfies these requirements. Figure 9 shows that the ratio of FITC-to-tryptophan fluorescence is initially constant during adsorption from 30 mM buffer and then drops as the protein reorients. The ratio does not increase again when the surface concentration is decreased below the critical surface coverage by desorption. Thus, the lysozyme remains in its preferred orientation with the most positive patch against the surface, and it does not revert to a random orientation. Since the reorientation does not reverse, it must optimize the overall energetics of adsorption, leading to an increase in the tenacity of adsorption. In fact, the FITC-to-tryptophan ratio decreases continuously as the lysozyme desorbs. This would suggest that the bias toward the preferred orientation becomes stronger during desorption. A plausible explanation is that those molecules that have assumed the optimal orientation, corresponding to those with the lowest FITC-to-tryptophan ratios, would have the lowest propensity to desorb. We studied lysozyme desorption from silica using optical reflectometry after adsorbing protein for 1, 4, and 20 min at the shear rate and bulk concentration used in Figure 1. These times correspond to adsorption prior to initial reorientation, after initial reorientation but before the second reorientation, and after the second reorientation, respectively, as indicated by the reference arrows in Figure 1. We induced desorption by rinsing with protein-free buffer of the same pH (7.4) and ionic strength as that used during adsorption. Results show that the longer the adsorption time, the smaller the fraction of protein that may be removed by rinsing with buffer. The fraction of the protein remaining adsorbed after 17 min of attempted desorption is 0.48 ( 0.04, 0.49 ( 0.04, and 0.69 ( 0.03 for

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adsorption times of 1, 4, and 20 min, respectively, where the standard deviation is calculated from multiple experiments. Hence the protein is optimizing its interactions with the surface over the course of 20 min and two FITC emission maxima. We can speculate that the lysozyme in the “positive patch down” orientation is more strongly adsorbed to silica than randomly oriented lysozyme. Additionally, we note that the fraction of protein remaining adsorbed after the buffer rinse was the same for unlabeled and labeled lysozyme. This eliminates the possibility that the FITC emission overshoots were simply artifacts of preferential adsorption of unlabeled protein. Kinetic Model. A mass action kinetic model captures some of the key features of the reorientation mechanism. The model includes bulk protein adsorption into either a “nonoptimized configuration” (drawn as the end-on oriented molecule B in Figure 10a) or an “optimized configuration” (drawn as the side-on oriented molecule) and transitions between the two configurations, as shown by the model schematic in Figure 10a. Equations 5-7 are

dCA )0 dt

(5)

dθB ) dt kACA(1 - θB - θC) - k2fθBCA - kd1θB + k2rθCCA (6) dθC ) dt kACA(1 - θB - θC) + k2fθBCA - kd2θC - k2rθCCA (7) used to model the adsorption and reorientation kinetics, where θ is the surface coverage; CA is the bulk concentration; and kA, k2f, k2r, kd1, and kd2 are rate constants for adsorption, the transition from the B to the C orientation, the transition from the C to the B orientation, desorption of species B, and desorption of species C, respectively. The rate constants were selected to represent adsorption rates similar to those observed in the experimental data. The rate constant kd2 was set to zero, because we have seen that desorption decreases as protein reorients to its optimized configuration. The rate constant kd1 was also set to zero because we observe very little desorption into the 5 mM TEA buffer which gives the strongest overshoot in the FITC kinetics. Additionally, we have observed that the FITC overshoot is irreversible, so k2r was also set to zero. Equation 5 is based upon experimental conditions whereby the bulk solution is continuously replenished, and protein is not depleted from solution. Equations 6 and 7 represent the presence of nonoptimized and optimized protein at the interface, where the second term in both equations represents the rate of reorientation. We used the surface coverage evolution of configurations B and C to predict the time evolution of the fluorescence intensity during adsorption. For this, we assigned different proportionality constants for the emission from each configuration, such that the emission intensity per molecule is higher by a factor of 3.55 for the nonoptimized configuration B. This proportionality constant comes from the results of a fluorescence pH titration.1 As shown in Figure 10b, we are able to capture the initial fluorescence overshoot with the rate constants given in the caption, while the surface coverage profile is monotonic. This model is further able to capture the increase in time of the overshoot with decreasing bulk protein concentration from 10 to 3 µg/mL. Figure 10c shows

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Figure 10. (a) The schematic shows that bulk protein (A) can adsorb into either a “nonoptimized” (B) or “optimized” (C) orientation. The rate constants were ka ) 0.005 (µg/mL)-1 s-1, kd1 ) 0 s-1, kd2 ) 0 s-1, k2r ) 0, k2f ) 0.005, bulk concentration CA ) 3-10 µg/mL, and initial coverage of B, C ) 0. The units of k2f and k2r are s-1 and s-1 µg/mL for CA included and excluded, respectively. (b) The kinetic profiles when CA is included in the reorientation term. (c) The curves in (b) collapsed by scaling. (d) The kinetic profiles when the CA is not included in the reorientation term. (e) The curves in (d) collapsed by scaling.

that curves for various concentrations can be collapsed, both before and after the overshoot, by scaling time as by τB ) tCA, where CA is the solution protein concentration. Above, we showed that the protein in solution near the interface helps to propagate the reorientation. The equations used to generate Figure 10b,c, discussed above, reflect this observation, in that the reorientation rate is first order with respect to the bulk protein concentration CA. For comparison, the fluorescence intensity profile when the rate of reorientation is considered to be independent of bulk concentration (i.e., CA is eliminated from the reorientation term k2fθBCA in eqs 6 and 7) is shown in Figure 10d. The bulk concentration is varied again from 3 to 10 µg/mL. Figure 10e shows the curves in Figure 10d replotted by again scaling time as τB ) tCA. The curves do not collapse after the overshoot as they did when the bulk concentration term was included in the reorientation rate expression. The peak predicted by the model is sharper when the bulk concentration is included in the reorientation rate expression and more closely represents experimental results. The reorientation rate is a function of the flux of additional protein to the surface from the bulk solution. The model results show that a reconfiguration of the adsorbed protein that leads to a more tightly bound and less fluorescent state, on average, than the initially

bound state or ensemble of states will give rise to a peak in the time profile of the emission. This simple model is consistent with our proposed mechanism of reorientation propagated by incoming proteins from solution.

Conclusions We present evidence from two independent experimental techniques, conducted under a variety of conditions, that lysozyme undergoes an irreversible reorientation to a configuration that places its most positively charged patch against the silica surface. Nonmonotonic FITC fluorescence and corresponding ζ-potential time traces indicate that the reorientation occurs in two stages. The final orientation we propose is consistent with the side-on adsorption model that has been proposed to explain neutron reflectivity data.2 This orientation may minimize lateral repulsions among adsorbed protein molecules that may lead to the development of an ordered two-dimensional structure.36 The final protein orientation or any clustering that results from reorientation can feasibly cause the observed increase in the tenacity of adsorption.


In this paper, we have systematically varied solution conditions including ionic strength, bulk lysozyme concentration, and shear rate in order to investigate reorientation dynamics. It is clear that these dynamics are controlled by interactions between the protein and the surface, between adsorbed proteins, and between adsorbed proteins and incoming bulk proteins. An understanding of these interactions in model systems is useful if we hope to eventually make generalized predictions of layer

Langmuir, Vol. 19, No. 9, 2003 3857

structure and protein activity changes upon protein adsorption. Acknowledgment. This material is based on work supported by the National Science Foundation under Grant Number BES-9907504. S.D. acknowledges support from the John E. Swearingen graduate fellowship. LA026690X

Coverage-Dependent Orientation of Lysozyme Adsorbed on Silica

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