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Surface Plasmon Resonance/Surface Plasmon Enhanced Fluorescence: An Optical Technique for the Detection of Multicomponent Macromolecular Adsorption at the Solid/ Liquid Interface Shaunak Roy,† Joon-H. Kim,†,‡ James T. Kellis, Jr.,§ A. J. Poulose,§ Channing R. Robertson,† and Alice P. Gast*,†,| Department of Chemical Engineering, Stanford University, Stanford, California 94305-5025, and Genencor International, Inc., Palo Alto, California 94304 Received January 25, 2002. In Final Form: April 22, 2002 We describe an optical technique for the measurement of macromolecular adsorption at the solid/liquid interface when multiple species are present. The technique combines surface plasmon resonance (SPR) with simultaneous surface plasmon enhanced fluorescence (SPEF). The relative ease of construction and linear correlation between SPR and SPEF signals make the technique amenable for coadsorption studies or multiple ligand binding experiments. Here, we demonstrate the utility of the technique with a biotin/ avidin/BSA “sandwich” experiment. We then apply SPR/SPEF for the simultaneous monitoring of enzyme adsorption and substrate cleavage of a protease interacting with a substrate surface.
Introduction The adsorption of macromolecules to solid surfaces is of importance in a wide variety of technologies, both biological and otherwise. Polymer adsorption is used in such applications as precoating surfaces in biosensors and surface modification for adhesion and lubrication.1 The adsorption of biological polymers, or proteins, plays a role in separation processes and is a limiting factor in the design of virtually any biocompatible material. For example, when blood comes into contact with a biomedical device, various plasma proteins, including albumin, immunoglobulins, and fibrinogen adsorb to the surface and thereby establish the extent of surface biocompatibility.2,3 A precise understanding of the physics of adsorption is necessary to design a surface that is most biocompatible. A particularly interesting aspect of proteins is the potential for retention of their intrinsic function when adsorbed to a surface. Thus, surface-bound antibodies may retain their specificity for antigens, ligands may still bind their proper proteins, and enzymes may catalyze biological reactions. This surface activity is exploited in such applications as diagnostic devices, contact lens cleaners, and detergent enzyme additives. For these reasons, a large volume of work is available on the adsorption of proteins to solid surfaces. Techniques as varied as ellipsometry,4-10 scanning angle reflectometry,11,12 electron spectroscopy for chemical analysis * Corresponding author: tel, 617-253-1403; fax, 617-253-8388; e-mail,
[email protected]. † Stanford University. ‡ Present address: LG Chem, 104-1 Moonji-dong, Yusong-gu, Taejon, Korea. § Genencor International, Inc. | Present address: MIT, 77 Massachusetts Ave., Cambridge, MA 02139-4307. (1) Fu, Z.; Santore, M. M. Colloids Surf., A 1998, 135, 63-75. (2) Green, R. J.; Davies, J.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B. Biomaterials 1997, 18, 405-413. (3) Ostuni, E.; Yan, L.; Whitesides, G. M. Colloids Surf., B 1999, 15, 3-30. (4) Arwin, H. Appl. Spectrosc. 1986, 40 (3), 313-318. (5) Jonsson, U.; Lundstrom, I.; Ronnberg, I. J. Colloid Interface Sci. 1987, 117 (1), 127-138.
(ESCA),13 total internal reflection fluorescence (TIRF),14,15 electron microscopy,16,17 infrared spectroscopy (IR),18 and atomic force microscopy (AFM)19,20 have been used to probe such interactions, and the subject has been reviewed by Andrade et al.21 and more recently by Malmsten.22 Much is now known about the surface properties that govern both the equilibrium and kinetics of adsorption. In many of the systems described above, however, several components are present at the surface at once and thus there may be several events occurring simultaneously. Deconvoluting such events with a single experimental technique can often be a complicated process and is nearly impossible to do in real time. One solution to this problem would be to use two or more of the above-mentioned techniques simultaneously to distinguish multiple components at the surface. Fu et al.23 have used TIRF along with Brewster (6) Golander, C. G.; Lin Y. S.; Hlady, V.; Andrade, J. D. Colloids Surf. 1990, 49 (3-4), 289-302. (7) Krisdhasima, V.; Vinaraphong, P.; McGuire, J. J. Colloid Interface Sci. 1993, 161 (2), 325-334. (8) Lin, Y.-S.; Hlady, V. S. Colloids Surf., B 1994, 2 (5), 481-491. (9) Malmsten, M. J. Colloid Interface Sci. 1994, 166 (2), 333-342. (10) Taylor, G. T.; Troy, P. J.; Sharma, S. K. Mar. Chem. 1994, 45 (1-2), 15-30. (11) Schaaf, P.; Dejardin, P. Colloids Surf. 1988, 31, 89-103. (12) Shirahama, H.; Lyklema, J.; Norde, W. J. Colloid Interface Sci. 1990, 139 (1), 177-187. (13) Ratner, B. D.; Horbetta T. A.; Shuttleworth, D.; Thomas H. R. J. Colloid Interface Sci. 1981, 83 (2), 630-642. (14) Cheng, Y. L.; Darst, S. A.; Robertson, C. R. J. Colloid Interface Sci. 1987, 118 (1), 212-223. (15) Tilton, R. D.; Robertson, C. R.; Gast, A. P. J. Colloid Interface Sci. 1990, 137, 192-203. (16) Gorman, R. R.; Stoner, G. E.; Catlin, A. J. Phys. Chem. 1971, 75 (14), 2103-2107. (17) Feder, J.; Giaever, I. J. Colloid Interface Sci. 1980, 78 (1), 144154. (18) Story, G. M.; Rauch, D. S.; Brode, P. F.; Marcott, C. ACS Symp. Ser. 1991, No. 447, 225-236. (19) Haggerty, L.; Lenhoff, A. M. Biotechnol. Prog. 1993, 9 (1), 1-11. (20) Haggerty, L.; Lenhoff, A. M. Biophys. J. 1993, 64 (3), 886-95. (21) Andrade, J. D. Surface and Interfacial Aspects of Biomedical Polymers; Plenum Press: New York, 1985; Vol. 2. (22) Malmsten, M. Biopolymers at interfaces; Marcel Dekker: New York, 1998; Surfactant Science Series 75. (23) Fu, Z.; Santore, M. M. Macromolecules 1998, 31 (20), 70147022.
10.1021/la025578w CCC: $22.00 © 2002 American Chemical Society Published on Web 07/30/2002
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angle spectroscopy to measure the competitive adsorption of polymers with different molecular weights onto silica glass. We use a similar tandem technique for the measurement of simultaneous surface events, in which both techniques rely on the resonant excitation of surface plasmons.24,25 Briefly, surface plasmons are propagating electromagnetic modes at the interface between a metal (typically either gold or silver) and a dielectric that can be resonantly excited by incoming p-polarized light. In the well-known Kretchmann configuration, a thin metal film is put in contact with the base of a prism such that the film is adjacent to a dielectric. When p-polarized light impinges on the base of the prism slightly above the angle of total internal reflection, the wave vector of the evanescent wave associated with total internal reflection
kin )
ω 1/2 sin θ c p
can be adjusted to match the real component of the wave vector of the surface plasmon
ksp )
(
)
md ω c (m + d)
1/2
(where ω is the frequency of the incoming light, c the speed of light in a vacuum, θ the angle of incidence, and p, m, and d the complex dielectric functions of the prism, metal, and dielectric, respectively). Under these conditions, surface plasmon resonance is achieved. At the resonance angle a sharp drop in reflectivity is caused by the transfer of energy from the incident photons to the surface plasmon. In a conventional surface plasmon resonance (SPR) biosensor, a specific analyte such as a protein or DNA becomes bound to the surface, thereby slightly altering the angle of resonance. This shift in the resonance angle is correlated to an average thickness of the bound layer or an effective surface mass. Kinetic measurements of the changes in amount of bound analyte can be made by monitoring an angle off resonance and measuring temporal changes in reflected intensity. The evanescent wave associated with the surface plasmon is also useful as a surface sensitive probe of fluorescence in the same manner as an evanescent wave formed in total internal reflection. Since energy is coupled into the surface plasmon, a substantial enhancement of field intensity is observed along the metal/dielectric interface compared to that of a TIRF sensor. This phenomenon was first used by Attridge et al.26 to produce an immunoassay. More recently, Liebermann et al.27 reported signal enhancements of up to 16× for gold surfaces and 80× for silver surfaces. In this work, they showed that for analytes large enough to be detected by surface plasmon resonance, both the surface plasmon and fluorescence signals vary linearly with adsorbed amount. We combine the phenomena described above in a technique called surface plasmon resonance/surface plasmon enhanced fluorescence (SPR/SPEF). The idea is that in an experiment with two components on the surface, SPR provides a measure of the total average layer thickness of surface coverage. One of the components is (24) Knoll, W. Annu. Rev. Phys. Chem. 1998, 49, 569-638. (25) Green, R. J.; Frazier, R. A.; Shakeshef, K. M.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B. Biomaterials 2000, 21, 1823-1835. (26) Attridge, J. W.; Daniels, P. B.; Deacon, J. K.; Robinson, G. A.; Davidson, G. P. Biosens. Bioelectron. 1991, 6, 201-214. (27) Liebermann, T.; Knoll, W. Colloids Surf., A 2000, 171 (1-3), 115-130.
Figure 1. (a) SPR/SPEF setup: L, He-Ne laser; NDF, neutral density filter; M, mirror; P, Glan-Tompson polarizer; SF, spatial filter; E, laser collimator; L1, planar cylindrical lens f ) 100 mm; L2, planar cylindrical lens f ) 25 mm; CCD, CCD detector (2048 elements line image detector); MO, microscope objective; PMT, photomultiplier tube detector. (b) Flow cell.
labeled with fluorescent dye and interrogated by SPEF. Liebermann’s demonstration of SPR/SPEF linearity permits calibration of the two signals to one another, thus allowing measurement of the unlabeled component via their difference. Our primary interest in designing the technique is to study the in situ adsorption and reactivity of enzymes interacting with surface-bound substrates. We envision it to be applicable to other instances where multiple species are present at the surface, such as coadsorption studies or immunoassays. Materials and Methods Apparatus. A schematic of the experimental setup is shown in Figure 1a. A 35 mW He-Ne laser is directed by a series of mirrors through a neutral density filter (used to control incident light intensity) and into the optical train. The beam passes through a Glan-Thompson polarizer to ensure p-polarized light for the SPR experiment. A spatial filter removes stray light from the beam, yielding a more homogeneous profile, followed by a beam collimator/expander which expands it to the appropriate diameter (∼2.5 cm). The widened beam is focused onto the sample cell by a vertically mounted planar cylindrical lens, as in the scanning angle reflectometry apparatus employed by Leermakers et al.28 and the SPR apparatus of Matsubara et al.29 An entire SPR spectrum can thus be captured at once on the CCD. In our experiment, we capture a horizontal spectrum over an angular width of ∼8°. This differs from the approach used by Lieberman et al.27 and will be discussed further in the Results and Discussion section. (28) Leermakers, F. A. M.; Gast, A. P. Macromolecules 1990, 24 (3), 718-730. (29) Matsubara, K.; Kawata, S.; Minami, S. Appl. Opt. 1988, 27, 1160.
SPR/SPEF for Macromolecular Adsorption The reflected beam is “vertically averaged” by a horizontally mounted cylindrical lens and directed onto the CCD element of a 1D CCD, which records the reflectivity data. Fluorescence is measured from behind the sample cell. A 5× microscope objective lens collects light emitted from the center of the flow cell and a PMT records a photon count. Both the PMT and camera are connected directly to a PC computer and data recorded using Labview software. Sample Cell. A diagram of the sample cell is shown in Figure 1b. A thin 50 nm gold film with a 2 nm chromium undercoat is evaporated onto a polished SF10 glass (n ) 1.723) slide (Schott Glass Technologies). The surface is functionalized by means of a thiol anchor carrying the selected reactive group. Another glass slide is sandwiched together with the first, with the gap between them defined by a silicone gasket (thickness 0.5 mm). A solution containing the analyte is then flowed through the cell by means of a peristaltic pump. The entire assembly is mounted onto an SF10 glass (n ) 1.723) hemicylindrical prism. Index matching is accomplished via a series M index matching liquid (n ) 1.730) from Cargille Laboratories. This provides a close but not perfect match to the SF10 glass, and as a result, we observe an interference pattern on the CCD image. We eliminate the effects of this pattern by vertically averaging the image using a second focusing lens (L2 in Figure 1a) just before the camera. Substrate Surfaces. Self-assembled monolayers of propionic acid were prepared by dipping gold-coated slides into a 3-mercaptopropionic acid solution (200 µL/100 mL) for 30 min. The carboxyl end groups of the immobilized hydrocarbon chains were activated for peptide bond formation using EDC (1-ethyl-3-(3dimethylaminopropyl)carbodiimide hydrochloride)/NHS (N-hydroxysuccinimide) chemistry. Slides were immersed in a solution of 40 mg/100 mL EDC and 60 mg/100 mL NHS in reaction buffer (2-(N-morpholino)ethanesulfonic acid 20.62 g/L, NaCl 29.2 g/L) for 1 h. The EDC/NHS step produces a water-stable ester able to react with primary amines. Biotinylated surfaces were produced by reaction with biocytin hydrazide (Molecular Probes B-1603, 10 mg/100 mL for 2 h) in reaction buffer. Monolayers of covalently bound fluorescent tagged bovine serum albumin (BSA) were formed by reaction with Texas Red/BSA conjugates (Molecular Probes A23017, 2.5 mg/mL for 2 h) in reaction buffer. In either case, the slides remained overnight in protein or biocytin hydrazide solution to ensure complete reaction. Protein Solutions. Avidin-Texas Red conjugates (A-820) were purchased from Molecular Probes. Biotin in solution was conjugated to BSA in a manner identical to that used to biotinylate the surface. Carboxy side chains of BSA (10 mg/100 mL) were activated by reaction with EDC (400 mg/100 mL) and NHS (600 mg/100 mL) for 1 h and then covalently linked to the amine group in biocytin hydrazide (5 mg/100 mL). Conjugates were separated from unreacted biotin using gel filtration chromatography. B. lentus subtilisin enzyme was obtained from Genencor International, Inc. All proteins were assumed to have a refractive index of n ) 1.57.30
Results and Discussion A typical “raw” reflectivity plot is shown in Figure 2a. The gold/buffer reflectivity curve is normalized on the gold/ air signal at the same angular range. This imageprocessing step allows us to remove the effects of inhomogeneities in the beam profile caused by either the laser or the surface. Normalized curves are shown in Figure 2b. Avidin-Biotin “Sandwich” Experiment. To demonstrate the utility of the SPR/SPEF technique in distinguishing among multiple components at a surface, we conduct a set of biotin/avidin/BSA “sandwich” experiments, shown diagrammatically in Figure 3. In each case, we start by binding a monolayer of biotin to the functionalized layer of hydrocarbons. In the first experiment, we allow (30) Jung, L. S.; Campbell, C. T.; Chinowsky, T. M.; Mar, M. N.; Yee, S. S. Langmuir 1998, 14 (19), 5636-5648.
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Figure 2. Sample gold (n ) 0.1726, k ) 3.4218)/air (n ) 1) (gray line) and gold/buffer (n ) 1.335) (black line) reflectivity signal (where n and k are the real and imaginary parts of the complex refractive index) before (a) and after (b) normalization by the reference gold/air signal. A substantial improvement in the surface plasmon image quality is observed.
a monolayer of fluorescently labeled avidin to bind the biotin (Figure 3a). On the basis of the Liebermann et al.27 results, the SPR and SPEF signals should be proportional to one another. Our use of a planar cylindrical lens to focus a fan of laser light onto the surface further simplifies this analysis. In a conventional single angle kinetics experiment, the amount of energy transfer to the surface plasmon wave changes through the course of the experiment. This will detract from the linearity of the correlation between the SPR and SPEF signals. Liebermann et al.27 discussed this effect and stated that it must be accounted for during the analysis. However, in our experiment, light is impinging on the surface at a range of angles, and thus the intensity transferred to the surface plasmon wave will remain constant throughout the experiment. Thus, no further signal processing is necessary to linearly correlate the SPR and SPEF measurements. In the next experiment, we successively add labeled and unlabeled avidin to the monolayer of biotin (Figure 3b). The SPR signal should rise throughout both additions due to a continued increase in protein layer thickness; however the SPEF signal will only rise following addition of the labeled protein. Thus, the two signals can be used to separate labeled and unlabeled species. The same idea is used to distinguish the presence of two separate proteins on the surface in the second part of a “sandwich” experiment (Figure 3c). The first layer consists of unlabeled avidin, and thus no rise in SPEF signal should be observed. We then add a fluorescently labeled biotinBSA conjugate to this layer. The rise in SPEF signal indicates the formation of the second layer. Once again, the amounts of the two proteins on the surface can be differentiated by use of the SPR and SPEF signals in tandem. The results from these three experiments are shown in Figures 4-6. In Figure 4, fluorescently labeled avidin was flowed over a biotin monolayer and bound forming a 31
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Figure 5. SPEF (black symbols) and SPR (gray symbols) results from the experiment described in Figure 3b. The SPEF units are arbitrary (AU ) 0 corresponds to dark current). Arrows indicate the time of addition. (a) Incomplete monolayer experiment in which fluorescently labeled avidin is passed over the biotin surface and followed by unlabeled avidin. No rise in SPEF follows addition of the unlabeled avidin. (b) The corresponding control experiment in which fluorescently labeled avidin is added at both steps. Figure 3. Biotin/avidin binding experiment. In (a), a monolayer of labeled avidin is bound to a biotin surface. In (b), an incomplete monolayer of labeled avidin is formed followed by completion of the layer with unlabeled avidin. In (c), the first complete avidin monolayer is left unlabeled and another layer of fluorescently labeled biotin-BSA is bound (“sandwiched”) to the unlabeled avidin surface.
Figure 4. SPEF (black symbols) and SPR (gray symbols) results of the experiment depicted in Figure 3a. The SPEF units are arbitrary (AU ) 0 corresponds to dark current). Arrows indicate the time of addition. Avidin bound to an equilibrium thickness of 31 ( 3 Å. The inset shows the linear correlation between the two signals (9, data; s, least-squares fit).
( 3 Å thick layer. The thickness was determined by matching the shift in resonance angle to model Fresnel calculations. The uncertainty reflects the spread in data from repeated measurements. SPR and SPEF signals are essentially identical. The inset clearly shows the linearity of the signals. Another advantage of the tandem experi-
Figure 6. SPEF (black symbols) and SPR (gray symbols) results from experiment depicted in Figure 3c. The SPEF units are arbitrary (AU ) 0 corresponds to dark current). Arrows indicate the time of addition. (a) Biotin/avidin/BSA “sandwich” experiment in which only BSA is fluorescently labeled. The SPEF signal only rises in the second step. The inset shows the amounts of BSA (gray symbols) and avidin (black symbols) present on the surface separately. (b) Corresponding control experiment in which neither component is labeled.
ment is that it discounts the need for an independent calibration of the fluorescence signal. The two signals can
SPR/SPEF for Macromolecular Adsorption
be calibrated with one another and an average layer thickness determined. An example of an incomplete monolayer is shown in Figure 5. Labeled avidin is allowed to bind to a thickness of ∼18 Å. Unlabeled avidin is then added and allowed to complete the monolayer (Figure 5a). This second step illustrates the separation of the two components, as the SPR signal rises but the SPEF signal remains unchanged. The corresponding control experiment is shown in Figure 5b. In this case, labeled avidin is added in both steps demonstrating no distinction between SPR and SPEF signals. Finally in Figure 6, the same idea is applied to the distinction of two unrelated proteins. Fluorescently labeled biotin-BSA is passed over an unlabeled avidin layer and bound to a thickness of ∼10 Å (Figure 6a). A fluorescence signal is only detected in the BSA step, and thus the amounts of the two proteins on the surface at any given time can be determined as shown in the inset. In the corresponding control (Figure 6b), unlabeled BSA is added and no fluorescence signal is detected in either step. Application to Measurement of Adsorption and Reactivity. Our primary goal in developing this technique is to simultaneously measure the adsorption and reaction kinetics of an enzyme interacting with a substrate surface. Our model substrate for this experiment is fluorescently labeled BSA. On average, there are 3.4 fluorescent labels per protein, with the labels dispersed randomly at lysine residues throughout the molecule. The enzyme is the serine protease subtilisin. Subtilisin adsorbs to and cleaves BSA from the surface. A suitable option to this configuration would be to fluorescently label the enzyme and work with unlabeled BSA. Our intention however, is to eventually compare the relative surface reactivities of several variants of subtilisin. It was thus deemed more appropriate to study their interactions with a standardized substrate and avoid any possible alteration of enzyme activity by labeling. Figure 7 shows results from this experiment. At the beginning of the experiment, buffer (carbonate buffer, pH ) 10) is passed over the surface to establish an initial gold/thiol/BSA/buffer surface plasmon. Once enzyme solution is added, it begins to hydrolyze the BSA layer. The drop in the SPR signal indicates a depletion of the total protein layer. This results from a combination of enzyme adsorption (the layer thickens) and BSA hydrolysis (the layer is thinned). The fluorescence signal isolates the loss of BSA or the kinetics of the reaction. Once again, taking advantage of the linearity between the SPR and SPEF signals, the displacement between the two signals relates to the kinetics of adsorption. As shown in Figure 7b, we observe the dynamic adsorption and reactivity behavior of an enzyme interacting with a substrate surface. A complete analysis of our adsorption/reaction results including kinetic modeling is the subject of a subsequent report.
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Figure 7. (a) SPEF (black symbols) and SPR (gray symbols) raw data from reaction of an enzyme (subtilisin) with a monolayer of fluorescently labeled BSA substrate. The SPEF units are arbitrary (AU ) 0 corresponds to dark current). Arrows indicate the time of addition. Calibration and subtraction of the two signals (SPR-SPEF) allows measurement of both surface-bound substrate (black symbols) as well as adsorbed enzyme (gray symbols), as shown in (b).
Conclusion The surface properties of macromolecules are often difficult to discern due to the simultaneous presence of multiple species at the surface. By use of the tandem technique of SPR/SPEF, the amounts of each individual species on the surface can be determined. We demonstrated the utility of the technique by distinguishing two separate proteins using a biotin/avidin/BSA “sandwich” assay. SPR/SPEF can also be used to probe more complicated dynamic behavior, as illustrated by the measurement of concurrent adsorption and reaction of an enzyme interacting with a substrate surface. In such cases, simultaneous surface events are often codependent, and using SPR/SPEF, one can observe the interplay of competing phenomena. Acknowledgment. We thank Dr. Jim Mikkelsen for help with the flow cell design. Subtilisin for the enzyme experiments was provided by Genencor International, Inc. We also thank Genencor International, Inc. for funding. LA025578W
