Langmuir 2008, 24, 12303-12311
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Fluorescence Quantification for Surface Plasmon Excitation Bob E. Feller,† James T. Kellis, Jr.,§ Luis G. Casca˜o-Pereira,‡ Wolfgang Knoll,‡ Channing R. Robertson,† and Curtis W. Frank*,† Department of Chemical Engineering, Stanford UniVersity, Stanford, California 94305-5025, Max-Planck-Institute for Polymer Research, Ackmannweg 10, D-55128 Mainz, Germany, and Genencor, a Danisco DiVision, Palo Alto, California 94304 ReceiVed May 5, 2008. ReVised Manuscript ReceiVed August 1, 2008 Surface plasmon resonance and surface plasmon fluorescence spectroscopy in combination have the potential to distinguish multicomponent surface processes. However, surface intensity variations from resonance angle shifts lead to a nonlinear response in the fluorescence intensity. We report a method to account for surface intensity variations using the experimentally measured relationship between fluorescence and reflectivity. We apply this method to monitor protease adsorption and proteolytic substrate degradation simultaneously. Multilayer protein substrates are prepared for these degradation studies using a layer-by-layer technique.
Introduction Surface plasmon resonance (SPR) is one of the most widely used techniques for characterizing thin films and measuring changes in interfacial properties.1 SPR is commonly employed to determine the kinetics of protein adsorption;2 however, binding events such as DNA hybridization at low target concentration do not produce substantial enough interfacial changes to be monitored using SPR.3 Such sensitivity limitations associated with SPR can be overcome by detecting fluorescence excited by the evanescent field. As a result, surface plasmon fluorescence spectroscopy (SPFS) is currently one of the most sensitive techniques for measuring surface2-4 or solution5 concentrations associated with biomolecular binding events. The sensitivity for SPFS detection has been shown to be in the attomolar concentration range for binding onto a 3D dextran matrix.5 The sensitivity of SPFS can be partially attributed to the surface enhancement of the incoming light as well as to the surface selectivity. The enhancement factor can reach up to 16 for Au and 50 for Ag with respect to the incoming light.2 This is particularly useful for binding events that result in negligible plasmon shifts and are, therefore, undetectable using conventional SPR. SPR has two modes of analysissangular scan and kinetics both of which have been previously described in detail;1 however, they will be briefly reviewed since understanding these concepts is crucial to the quantification. Both modes monitor the reflected and fluorescence light intensity as a function of either the angle or time for the angular scan or kinetic mode, respectively. From the reflectivity scan, both the critical angle and angle of minimum reflectivity are measured, which provide the thickness of the film (for a material of known refractive index). The fluorescence scan, which has a strong angular dependence, shows a maximum * To whom correspondence should be
[email protected]. † Stanford University. ‡ Max-Planck-Institute for Polymer Research. § Genencor.
addressed.
E-mail:
(1) Knoll, W. Annu. ReV. Phys. Chem. 1998, 49, 569–638. (2) Liebermann, T.; Knoll, W. Colloids Surf. A. 2000, 171, 115–130. (3) Liebermann, T.; Knoll, W.; Sluka, P.; Herrmann, R. Colloids Surf. A 2000, 169, 337–350. (4) Ekgasit, S.; Stengel, G.; Knoll, W. Anal. Chem. 2004, 76, 4747–4755. (5) Yu, F.; Persson, B.; Lo¨fås, S.; Knoll, W. J. Am. Chem. Soc. 2004, 126, 8902–8903.
intensity at a slightly smaller angle than the minimum in reflectivity. As the film thickness increases, so does the angle of minimum reflectivity (assuming constant refractive index). Consequently, this causes the fluorescence intensity peak to shift to higher angles as well. The kinetic mode is used to monitor interfacial film thickness as a function of time. The reflectivity and fluorescence intensity are monitored at a constant angle within the linear regime of the plasmon, and the reflectivity can be linearly correlated with the initial and final interfacial film thicknesses determined from the angular scans. The thickness measured from the reflectivity during kinetic mode represents the total thickness, which consists of both fluorescently labeled and unlabeled material. In our case, this is the protein substrate and enzyme, respectively. Alternatively, the fluorescence signal is only related to the fluorescently labeled portion of the film. This study focuses on the quantification of the fluorescence signal excited using SPR. Typically, fluorescence techniques maintain constant illumination intensity throughout the experiment to allow direct comparisons of measured intensities. For example, percent recovery in fluorescence recovery after photobleaching (FRAP) is defined as the ratio of the intensity after complete recovery to the initial intensity. The immobile fraction is calculated as the difference between the fraction recovered and unity in the photobleached region. The constant illumination intensity allows the fluorescence to be linearly correlated to the surface concentration.6 This is also true for kinetic mode SPFS when resonance angle shifts are negligible, resulting in a constant surface intensity. Resonance angle shifts due to more substantial interfacial refractivity differences (those detectable by SPR) lead to a change in the surface intensity. Such resonance angle shifts in kinetic mode are monitored by measuring the reflectivity at a given angle. It is important to note that light reflected from the interface does not excite surface plasmons. Since the laser light intensity directed at the interface remains constant, changes in reflectivity correlate to changes in surface plasmon intensities. For appreciable reflectivity changes, the linear correlation between fluorescence intensity and surface concentration is no longer valid. For example, Liebermann demonstrated that thickness changes as small as 0.8 nm can result in deviations from linearity.2 Together, SPR and (6) Axelrod, D.; Koppel, D.; Schlessinger, J.; Elson, E.; Webb, W. Biophys. J. 1976, 16, 1055–69.
10.1021/la8013943 CCC: $40.75 2008 American Chemical Society Published on Web 10/10/2008
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SPFS can provide sufficient information to decouple simultaneous surface processes, for instance, enzyme adsorption and reaction. These measurements have been demonstrated using multiangle SPR geometries; however, there is no angular dependence to the fluorescence intensity, since light striking the surface does so over a range of angles.7 Single angle SPR and SPFS could also decouple multicomponent association/dissociation events; however, most biomolecular binding events (those leading to substantial reflectivity changes) would result in a nonlinear fluorescence response and therefore incorrect interpretation. Accounting for surface intensity changes would enable SPR and SPFS to be used simultaneously to monitor multiple components for thickness changes on a scale similar to typical protein dimensions. Theoretical understanding of surface plasmons and the angular dependence on surface intensity is well-documented.8-10 However, surface intensity simulations can be tedious, and software is not readily available. Furthermore, simulations may not accurately fit the experimental data precisely throughout the entire range of interest, leading to disparities in the interpretation. In this study, we demonstrate a method to account for surface intensity variations so that the thickness of a fluorescing film can be quantified under surface plasmon excitation. In combination with conventional SPR, we have the ability to decouple simultaneous surface processes. We have applied this method to separate enzyme adsorption and reaction, and we have chosen to use a protease/protein substrate system to address this issue. The protein substrate is prepared using a layer-by-layer technique to create a multilayer architecture to ensure large resonance angle shifts. Since the entire protein film is a substrate for proteolytic hydrolysis, we can be confident that significant resonance angle shifts will occur during substrate degradation. Thus, surface intensity normalization will be necessary to correlate the fluorescence intensity with a film thickness. The adsorption process can also produce resonance angle shifts that result in surface intensity changes. Enzyme adsorption is the first step in the enzyme-catalyzed surface-bound substrate degradation. The adsorption process is complex; therefore, understanding the interactions between the enzyme and surface will allow us to design enzymes with improved catalytic properties at the solid-liquid interface. We chose the enzyme subtilisin for these studies, because of its availability and because detailed studies have previously characterized its solution catalysis and structure.11,12 Subtilisin is a serine protease that catalyzes the hydrolysis of a protein’s peptide backbone. Since fluorescently labeling the enzyme could affect performance, we chose to use a fluorescently labeled protein substrate. Protease binding is interpreted as the difference between the total thickness of the film (from the SPR signal) and substrate thickness (from the fluorescently labeled protein). Nonlinear behavior is accounted for by normalizing the fluorescence signal by the relative surface intensity using the experimentally measured relationship between fluorescence intensity and reflectivity.
Experimental Section Materials. Subtilisin, bovine serum albumin (BSA), dextran sulfate sodium salt Mr ∼ 5000 (DSS), glutaraldehyde (50% by weight), (7) Roy, S; Kim, J.; Kellis, J.; Poulose, A.; Robertson, C.; Gast, A. Langmuir 2002, 18, 6319–6323. (8) Ekgasit, S.; Thammacharoen, C.; Knoll, W. Anal. Chem. 2004, 76, 561– 568. (9) Ekgasit, S.; Thammacharoen, C.; Knoll, W. Anal. Chem. 2004, 76, 2210– 2219. (10) Ekgasit, S.; Yu, F.; Knoll, W. Sens. Actuators B 2005, 104, 294–301. (11) Kuhn, P.; Knapp, M.; Soltis, S. M.; Ganshaw, G.; Theone, M.; Bott, R. Biochemistry 1998, 37, 13446–13452. (12) Ballinger, M.; Jeffrey, T.; Wells, J. Biochemistry 1995, 34, 13312–13319.
Feller et al. N-ethyl-N′-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC), N-hydroxysuccinimide (NHS), phenylmethanesulfonyl fluoride, 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid sodium salt (HEPES sodium salt), and 16-mercaptohexadecanoic acid were purchased from Sigma (St. Louis, MO). Alexa Fluor 647 (AF 647) labeled bovine serum albumin was prepared using an NHS activated Alexa Fluor 647 labeling kit purchased from Molecular Probes (Eugene, OR). The protocol was modified such that a 3-fold molar excess of protein to label was used in order to minimize intramolecular quenching from proteins with multiple fluorescent labels. Water was obtained from a Milli-Q water system (Millipore, Billerica, MA) at 18.2 MΩ · cm. Protein Substrates. We used the layer-by-layer (LbL) procedure described by Brynda and Houska13to prepare multilayer protein films. The cationic layer was BSA at a pH below its pI, while the anionic layer was a low molecular weight polymer, dextran sulfate (Mr ∼ 5000). The protein layers were cross-linked with glutaraldehyde, which does not react with the anionic polymer. Brynda showed that when the pH of the solution was then increased above the pI of the protein, electrostatic repulsion drove the low molecular weight polymer out of the film. Previous studies using this method contained only one step to cross-link the entire film.13 Interestingly, during degradation studies with one final cross-link step, we noticed that only the outermost layer of the multilayer seemed to have any significant degree of cross-linking. Once this layer was degraded, the entire multilayer disintegrated due to electrostatic repulsion among the un-cross-linked proteins, leaving behind only the first covalently bound monolayer. For this reason, we adopted a modified protocol using multiple cross-linking steps to give a more uniformly crosslinked film, as described in the following paragraph. The crosslinking reagent glutaraldehyde reacts with amines in the protein to form a Schiff base. The pKa of the Schiff base is approximately 3 pH units lower than the primary amine of the lysine.14,15 Depending on the pH of the experiment, the cross-linking step may change the surface charge of the thin protein film with respect to the native protein. The pH of interest for these experiments was pH 8, which is nearly equivalent or slightly higher than the pKa of the Schiff base.14,15 Therefore, the surface charge of the film is expected to be more negative than the native protein, which may influence the reactivity and adsorption of the enzyme. Protein films ranging from one to 10 layers of BSA were prepared for fluorescence quenching studies, while enzyme studies were performed on films prepared using a five-layer protocol. SPR substrates were prepared by thermal evaporation of Cr (2 nm) and Au (50 nm) onto LaSFN9 (Schott glass) followed immediately by placement into a solution of 2 mM 16-mercaptohexadecanoic acid for at least 24 h. Slides were removed from solution, rinsed with ethanol, and dried with nitrogen. The first layer of protein was covalently bound to the surface using EDC and NHS, and the slides were activated prior to being placed into a HMS programmable slide stainer (Carl Zeiss). A 500 µL portion of 128 mM EDC and 32 mM NHS in Milli-Q H2O were placed on the slide for approximately 15-20 min. Slides were rinsed with Milli-Q H2O, dried with nitrogen, and then placed into the slide holder. The activated slides were transferred to a protein solution (0.5 mg/mL BSA in 30 mM citrate buffer pH 4 with 0.5% BSA AF 647 protein content) for 60 min. Higher fluorophore loading showed evidence of intermolecular self-quenching. Slides were rinsed twice in 30 mM citrate pH 4 for 5 min to remove excess protein and cross-linked using glutaraldehyde (5 mL of 50 wt % glutaraldehyde in 500 mL of citrate, pH 4). Slides were then rinsed in 30 mM citrate (pH 4) for 10 min and transferred to a solution of dextran sulfate (1 mg/mL DSS in 30 mM citrate pH 4) for 20 min followed by two more 5-min rinses with 30 mM citrate. This cycle was repeated to obtain the appropriate number of layers until the final cycle, where the DSS bath was omitted and slides were transferred directly from the (13) Brynda, E.; Houska, M. Macromol. Rapid Commun. 1998, 19, 173–176. (14) Johnson, T. J. Electron Microsc. Tech. 1985, 2, 129–138. (15) French, D.; Edsall, J. AdV. Protein Chem. 1945, 2, 277–335.
Fluorescence Quantification for SP Excitation glutaraldehyde to PBS (pH 7.4) for 1 h. During each step of the assembly process, the slides were agitated every 5 s. Thin Film Dielectric Constant. The dielectric constant of the protein film was determined using a two-wavelength SPR approach.16 The dielectric constant is equal to the square of the refractive index. All measurements were made in 5 mM Hepes (pH 8) plus 2.5 mM NaCl, the same buffer used for enzyme studies. SPR angular scans were collected at wavelengths of 632.8 and 980 nm. Solutions of BSA in 5 mM Hepes (pH 8) plus 2.5 mM NaCl were also prepared, and the dielectric constant of the solution was measured by the angle of attenuated total reflection against a LaSFN9 prism. The volume fractions of BSA in the solution were calculated using a density of 1.41 g/cm3.17 SPR/SPFS. A brief description of the apparatus is presented here. A more detailed description of the SPR/SPFS technique as well as a schematic of the experimental setup can be found elsewhere.2 Light from a He-Ne laser (632.8 nm) was passed through a chopper and reflected off the sample placed on a θ-2θ goniometer. The infrared laser light (980 nm) was reflected off a flip mirror placed between the He-Ne laser and the chopper. The sample was mounted using the Kretchmann configuration with index-matching fluid (Cargille, refractive index 1.7) between the slide and prism. The reflected laser light was measured with a photodiode and analyzed using a lock-in amplifier. The sample slide was pressed against the flow cell, which was constructed from a PDMS gasket (∼500 µm thick) and a BK7 microscope slide. An inlet and outlet hole were drilled in the microscope slide and connected to the Teflon tubing using PDMS. The fluorescence intensity was measured from the backside of the flow cell through BK7 glass using a photomultiplier tube (PMT) and a bandpass filter mounted on the sample stage. The goniometer and detectors were enclosed within a black structure to block external light. A shutter was placed between the laser and the dark enclosure to minimize fluorophore photobleaching between data points. Film thickness was analyzed by fitting the reflectivity data using Winspall software (Max Planck Institute for Polymer Research). Kinetic experiments were performed in four steps. First, a baseline was established by flowing buffer to ensure the film stability. Next, enzyme was added to the buffer at the concentration of interest and allowed to react with the substrate. Once the enzyme had adequate time to react, the enzyme was removed. Because bound enzyme was not removed by buffer rinse alone, it was necessary to add 150 mM sodium chloride to the original buffer to attenuate the attractive electrostatic interactions, thus permitting bound enzyme to be removed. Finally, the original buffer was added to return the solution to the original refractive index. Enzyme solutions were prepared at 2 µg/mL immediately before use. Enzyme inhibition was induced using phenylmethanesulfonyl fluoride (PMSF). PMSF forms a sulfonic ester with the serine residue of the catalytic triad, rendering the enzyme inactive. A stock solution of PMSF was prepared at 100 mM in ethanol and was added to the enzyme solution at a final concentration of 2 mM. Fluorescence Quenching. BSA films were prepared using the layer-by-layer technique described above. The thickness of each sample was determined using SPR. The experimental setup described in the SPR/SPFS section was used to measure the fluorescence for the distance-dependent (surface plasmon) excitation. After the SPR/ SPFS measurement, the flow cell was carefully disassembled so that buffer remained on the slide and the samples were never exposed to air between the SPR/SPFS and fluorescence microscope measurements. For the distance-independent excitation, a fluorescence microscope equipped with a 40× water immersion objective and CCD camera (Nikon Eclipse E800, 40× Nikon Fluor, Photometrics CoolSnap HQ) were used to measure the fluorescence intensity. An average pixel count for each image was determined by averaging over the entire image using Metamorph image analysis software (Universal Imaging). Equations were fit to experimental data by minimizing the sum of the squared error between the measured fluorescence intensity (surface plasmon and fluorescence microscope (16) Peterlinz, K.; Georgiadis, R. Opt. Commun. 1996, 130, 260–266. (17) Fischer, H.; Polikarpov, I.; Craievich, A. Protein Sci. 2004, 13, 2825– 2828.
Langmuir, Vol. 24, No. 21, 2008 12305 excitation) and the numerical integration. The measured intensities, in counts per second, were significantly higher for the PMT (SPFS) compared to the CCD camera (fluorescence microscope). Unfortunately, this caused the fit for the fluorescence microscope data to be poor, since the sum error squared was essentially determined by the data set with the higher count rate. Therefore, it was necessary to normalize each data set prior to fitting the data. The coefficient of determination, R2, for each data set was calculated using the most general definition (1 minus the ratio of the residual sum of squares to the total sum of squares).
Results and Discussion Thin Film Dielectric Constant. SPR is a technique to monitor interfacial refractive index, which is related to both the thickness and refractive index of the film. Typically, if the film thickness is of interest, it is necessary that the refractive index of the film be estimated or measured using another technique. However, it is possible to determine both the thickness and the refractive index using either a multisolvent or a multiwavelength approach.16 Each approach exploits the change in evanescent decay length caused by either the difference in refractive index of the bulk medium (multisolvent) or the wavelength of excitation (multiwavelength). The multisolvent approach can be used for films where it is unlikely that the solvent affects the structure or thickness of the film. The protein films used in this study most likely contain a substantial amount of water, especially under conditions where BSA is highly charged and proteins within the film repel one another. As a result, the properties of the protein film are likely to be significantly influenced by the solvent; thus, the multisolvent approach would not provide an accurate determination of film parameters. The multiwavelength approach allows us to determine the thickness and refractive index without changing the solvent; however, since the refractive index of both components of the film (buffer and protein) are weakly wavelength-dependent, the refractive index of the hydrated BSA film will also be wavelength-dependent to some degree. It is possible to determine the thickness and dielectric constant of the film using the multiwavelength approach not knowing the dielectric constant at either wavelength if we can estimate the difference in the dielectric constant at the two wavelengths.16 Assuming this difference can be measured or calculated theoretically, film properties can be determined from the angular scans at both wavelengths. First, we must generate a curve that represents all possible solutions (dielectric constant and thickness) for the angular reflectivity scan at each wavelength. Typically, the thickness of a film is determined by matching the reflectivity minimum of a surface plasmon simulation to the measured reflectivity data assuming a value for the dielectric constant. However, this interpretation is ambiguous because it does not describe a unique solution for the thickness and dielectric constant. Had we initially chosen a different dielectric constant, the thickness where the simulation matched the experimental data would have been different. Thus, we can generate a curve that represents all possible solutions of dielectric constant and thickness. Alternatively, this curve can be generated by selecting the thickness and adjusting the dielectric constant of the simulation to match the data. We chose the latter because the dielectric constant increment in the simulation software led to a smaller angle shift in the simulated reflectivity minimum and, therefore, greater accuracy matching the data. These data are shown in Figure 1 for a 10-layer BSA film at each of the wavelengths, 632.8 and 980 nm. The intersection of these two curves corresponds to the thickness and dielectric constant of the film, assuming that the difference in dielectric constants at the two wavelengths is negligible.
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Figure 1. Determination of dielectric constant using the two-wavelength approach. Thickness as a function of dielectric constant for 632.8 nm (9), 980 nm (0), and the adjusted values for 980 nm (O) given the difference of dielectric constants at 632.8 and 980 nm. Table 1. Dielectric Constant of Solutions at Different Volume Fractions of BSA Measured at 632.8 and 980 nm dielectric constant volume fraction BSA
at 632.8 nm (ε632.8)
at 980 nm (ε980)
difference (ε632.8 - ε980)
0 0.012 0.057 0.33
1.773 1.781 1.808 1.991
1.757 1.765 1.791 1.972
0.016 0.016 0.017 0.019
The difference in dielectric constant for our film is estimated by measuring the dielectric constant of solutions at different volume fractions of BSA. As seen in Table 1, the difference in dielectric constant (ε632.8 - ε980) is only weakly dependent on the BSA volume fraction. For example, the difference in dielectric constant only changes 0.003 from pure buffer to the highest volume fraction of BSA. Using the difference in dielectric constant at the highest BSA volume fraction, 0.019, we calculate the film’s dielectric constant. Since the difference in dielectric constant of the film is independent of thickness, the curve generated at 980 nm is shifted by this measured difference, 0.019, resulting in a new intersection point. From this intersection, shown in Figure 1, the dielectric constant of the film at 632.8 nm is determined to be 2.042. Comparing the calculated dielectric constant for the film at 632.8 nm, 2.042, to the value for the highest volume fraction of BSA at 632.8 nm, 1.991, we see that this difference is less than one-fourth of that between the highest volume fraction, 1.991, and the pure buffer, 1.773, at 632.8 nm. Since the difference in dielectric constant (ε632.8 - ε980) over the larger dielectric change at 632.8 nm, 1.773-1.991, was only 0.003, we assume that the smaller dielectric change at 632.8 nm, 1.991-2.042 (between the highest volume fraction and the BSA film) leads to a considerably smaller change than 0.003. Therefore, we can be confident that the difference in dielectric constant (ε632.8 - ε980) at the highest volume fraction of BSA is an accurate approximation for the difference in dielectric constant of the BSA thin film at these wavelengths. The measured dielectric constant of the BSA film, 2.042, in combination with angular scans made at 632.8 nm allows us to determine the thickness of the protein film before and after the reaction. The enzyme layer is assumed to have the same dielectric constant as the substrate. Protein Multilayer. Layer-by-layer deposition is a desirable approach for fabricating the protein substrates because it gives
precise control over a number of the film properties. Most importantly, LbL deposition permits control of both the film thickness and the cross-linking density. The thickness is directly related to the number of protein deposition steps, while the crosslinking density is determined by the concentration and time in the glutaraldehyde solution. Control over these properties is essential, because we are interested in interpreting the interaction between an enzyme and immobilized protein substrate. As the digested multilayer film approaches monolayer to submonolayer coverage, it is likely that portions of the underlying solid support become exposed. By generating sufficiently thick films with an appropriate cross-linking density and terminating the reaction with a substrate thickness of greater than 20 nm, we can ensure that enzyme-thiol interactions are not present. Thicker substrates have an added benefit of increased signal-to-noise because metal quenching is reduced. Finally, this particular LbL approach allows us to create a uniform protein film by removing any unwanted polyelectrolyte present in the deposition. SPR/SPFS. SPR is used to determine the thickness of the entire protein film (substrate plus enzyme). From the reflectivity angular scans, the substrate thickness is measured before and after the reaction. For the kinetic experiments, the reflectivity is monitored at a constant angle below resonance near the upper bound of the linear regime because film degradation will shift resonance to lower angles. The thicknesses measured from the reflectivity scans are used to correlate reflectivity data from the kinetic experiment to film thickness. The fluorescence signal is also monitored as a function of angle and time for the angular scan and kinetic experiment, respectively. The fluorescence intensity is a measure only of the labeled protein substrate, allowing us to decouple adsorption from the reaction phenomena. Quantification of the fluorescence signal is described in the following section. Angular scans before exposure, during exposure, and after removal of inhibited enzyme are shown in Figure 2a. Clearly, inhibited enzymes do not react with the surface since the resonance angles before exposure and after removal are identical. During exposure, the resonance angle shifts to a higher value as the inhibited enzyme binds to the protein multilayer. Fluorescence data are normalized to the maximal value of the initial scan to account for the small amount of photobleaching occurring during the scan while the shutter is open. Kinetic information obtained by monitoring at a constant angle will resemble the schematics in Figure 2b,c. Thus, it is reasonable that binding will decrease the fluorescence intensity even though the fluorescent substrate remains unaffected due to a decrease in surface intensity. Figure 3 shows the raw and quantified data for active and inhibited subtilisin interacting with the BSA multilayer substrate. The change in reflectivity after the introduction of high ionic strength buffer is due to a combination of removal of bound enzyme and an increase in the solution refractive index. The reflectivity drop observed after returning to the original buffer is due purely to a refractive index change. Because the reflectivity signal returns to its original value after the high ionic strength buffer in the inhibited enzyme experiment, we are confident that the high ionic strength buffer is sufficient to remove all adsorbed enzyme. Initial and final values for the quantified reflectivity and fluorescence intensity are fixed to the value measured from the appropriate angular scan. A description of the procedure to quantify the fluorescent signal is given in the following section. From Figure 3d, one can determine the amount of enzyme adsorbed and the rate of substrate removal at any given time during the reaction, as shown in Figure 4. It is interesting to note that the rate of reaction begins to level off after 10 min of reaction while adsorption continues to increase until around the time
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Figure 2. (a) Reflectivity (9) and fluorescence intensity (0) angular scans of BSA multilayer. Reflectivity (b) and fluorescence intensity (O) angular scans during exposure to inhibited subtilisin. Reflectivity (2) and fluorescence intensity (4) angular scans of BSA multilayer after exposure and removal of inhibited subtilisin. Note: the values after enzyme removal are difficult to discern because they lie beneath the scan before enzyme exposure. Schematic of constant angle kinetic response of (b) reflectivity and (c) fluorescence intensity to enzyme binding. Θobs represents the observation angle, while ∆R and ∆F represent the change in reflectivity and fluorescence intensity for plasmon shifts at a constant angle. The arrows show the movement over time.
Figure 3. Raw reflectivity (9) and fluorescence intensity (O) data for (a) inhibited subtilisin and (b) active subtilisin exposure to BSA multilayer, and total thickness (9) and substrate thickness (O) as measured from reflectivity and fluorescence intensity for (c) inhibited subtilisin and (d) active subtilisin exposure to BSA multilayer: (1) 5 mM Hepes (pH 8) plus 2.5 mM NaCl, (2) 2 µg/mL subtilisin in 5 mM Hepes (pH 8) plus 2.5 mM NaCl, (3) 5 mM Hepes (pH 8) plus 150 mM NaCl, and (4) 5 mM Hepes (pH 8) plus 2.5 mM NaCl.
when the reaction is quenched. We propose two possible explanations for this observation. First, the cross-linking density of the film changes as a function of thickness. As mentioned in
the Experimental Section, we observed that BSA films prepared using only one cross-linking step following the multilayer deposition of BSA gave poor cross-linking uniformity. Once the
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is randomly distributed and oriented during the deposition process, leading to an even distribution of fluorophore throughout the film. In addition to the number of fluorophores within the film, changes in plasmon excitation intensity and distance-dependent excitation and de-excitation pathways affect the total fluorescence. Excitation occurs via energy transfer from the exponentially decaying evanescent field, which has its maximum at the metal-dielectric interface. The amplitude of the evanescent field also has an angular dependence corresponding to the magnitude of surface plasmon excitation and consequently its relation to the resonance angle. The excitation intensity, Iex(z), is proportional to the square of the evanescent field4,8-10,20,21
Iex(z)/Io ) (Ez/Eo)2 ) (e-z/ld)2 ) e-2z/ld
(1)
where z is the distance from the metal, Io is the intensity at the metal/dielectric interface, and the decay length1,8,22 Figure 4. Enzyme adsorption (nm) (9) and substrate removal rate (nm/ min) (O) obtained from data in Figure 2d: (1) 5 mM Hepes (pH 8) plus 2.5 mM NaCl, (2) 2 µg/mL subtilisin in 5 mM Hepes (pH 8) plus 2.5 mM NaCl, (3) 5 mM Hepes (pH 8) plus 150 mM NaCl, and (4) 5 mM Hepes (pH 8) plus 2.5 mM NaCl.
outmost layer of BSA is degraded, the film disintegrates due to repulsion between relatively un-cross-linked underlying proteins. To increase cross-linking uniformity, we chose to cross-link the film after each protein deposition step. If underlying protein layers are completely isolated from the cross-linker, we would expect this to generate a uniformly cross-linked film. It is possible that the cross-linker interacts primarily with the outmost layer and to a much lower extent with the underlying protein layers of the film. In this case, we would expect the underlying layers to have a slightly higher degree of cross-linking. The reactivity would then seem to plateau due to the competing effects of increased enzyme and higher cross-linking density. While crosslinking uniformity has been improved with respect to a single cross-linking step, we cannot rule out the possibility that slight differences might still exist. Second, this could be attributed to crowding that occurs at high surface coverage. Crowding has been shown to affect certain enzyme properties, such as diffusion on different surfaces.18,19 It is possible that crowding or decreased mobility due to crowding will affect the specific activity of each adsorbed enzyme molecule. Fluorescence Quantification. Surface intensity variations associated with SPFS cause a nonlinear response in the fluorescence intensity, uncharacteristic of many fluorescence techniques. Conventional SPR measures resonance angle shifts to determine film thickness; however, these resonance angle shifts also cause surface intensity changes. Thus, to extract useful information from both signals independently, it is necessary to account for the changes in surface intensity. We have developed a method to account for such changes and have demonstrated this using a protease/protein substrate system. The layer-by-layer technique described herein is ideal for preparing the protein substrate for this study. The degradation of this multilayer film will lead to significant resonance angle shifts, and therefore, the applied method is required for accurate interpretation of the fluorescence signal. Fluorescence for this type of system becomes difficult to quantify due to the multiple factors that affect the fluorescence signal. One such factor is the number of fluorophores within the protein film. We assume that the fluorescently labeled protein (18) Sonesson, A.; Elofsson, U.; Brismar, H.; Callisen, T. Langmuir 2006, 22, 5810–5817. (19) Sonesson, A.; Brismar, H.; Callisen, T.; Elofsson, U. Langmuir 2007, 23, 2706–2713.
ld )
λ 2π√εp sin2(θp) - εd
(2)
where λ is the wavelength of the light, εp and εd are the dielectric constants of the prism and dielectric medium, and θp is the angle of the light within the prism. While fluorophore excitation is greatest at the metal/dielectric interface, the measured fluorescence intensity is negligible due to alternative de-excitation pathways that dominate close to the metal. Analogous to energy transfer efficiency, the efficiency of fluorescence in the presence of an acceptor is given by22-24
Φ(z)/Φo )
1 1 + (β/z)n
(3)
where Φo is the yield at infinite separation from the metal film, β is the characteristic distance, and n is the exponential dependence. Equation 3 is used to describe all forms of energy transfer that do not result in fluorescence, including Fo¨rster transfer,24 surface energy transfer,24 and surface-plasmon-coupled emissions.23,25-27 If this were a theoretical study on the mechanism of energy transfer to the surface, each of these phenomena would play a different role depending on factors such as separation distance, orientation, etc.23,25-28 Here we are only interested in representing the alternative de-excitation pathways such that the fluorescence signal can be quantified. The total measured fluorescence under surface plasmon excitation of an infinitely thin film of fluorophores at a finite distance from a metal film is the product of the excitation intensity (eq 1) and the efficiency of fluorescence (eq 3)
I(z) ) R
1 e-2z⁄ld 1 + (β ⁄ z)n
(4)
where R is a proportionality depending on fluorophore loading, Io, Φo, and the surface plasmon intensity. Therefore, integration of Equation 4 over the film thickness should characterize a uniformly labeled fluorescent film such as that constructed using the layer-by-layer approach. (20) Liedberg, B.; Lundstro¨m, I.; Stenberg, E. Sens. Actuators B 1993, 11, 63–72. (21) Jung, L.; Campbell, C.; Chinowsky, T.; Mar, M.; Yee, S. Langmuir 1998, 14, 5636–5648. (22) Tawa, K.; Morigaki, K. Biophys. J. 2005, 89, 2750–2758. (23) Lakowicz, J. Anal. Biochem. 2005, 337, 171–194. (24) Yun, C.; Javier, A.; Jennings, T.; Fisher, M.; Hira, S.; Peterson, S.; Hopkins, B.; Reich, N.; Strouse, G. J. Am. Chem. Soc. 2005, 127, 3115–3119. (25) Lakowicz, J. Anal. Biochem. 2004, 324, 153–169. (26) Vasilev, K.; Knoll, W.; Kreiter, M. J. Chem. Phys. 2004, 120, 3439– 3445. (27) Hellen, E.; Axelrod, D. J. Opt. Soc. Am. B. 1987, 4, 337–350. (28) Chance, R.; Prock, A.; Silbey, R. AdV. Chem. Phys. 1978, 37, 1–65.
Fluorescence Quantification for SP Excitation
Figure 5. Average maximum fluorescence intensity (cps) of the SPR angular scan (9) versus film thickness. Error bars represent one standard deviation. Fluorescence intensity trend line (cps), ∫I(z) (- -), versus film thickness. Normalized excitation intensity, Iex(z)/Io (0), versus separation distance. Normalized fluorescence intensity, I(z)/R (O), versus separation distance.
Figure 6. Average fluorescence intensity (cps) of the fluorescence microscope image (9) versus film thickness. Error bars represent one standard deviation. Fluorescence intensity trend line (cps), ∫Φ(z) (- -), versus film thickness. Normalized fluorescence intensity, Φ(z)/Φo (O), versus separation distance.
The integral of eq 4 as well as the maximum fluorescence intensity from the SPR/SPFS angular scans are plotted as a function of film thickness in Figure 5. Similarly, Figure 6 shows the integral of eq 3 as well as the intensity measured using a fluorescence microscope versus film thickness for this distanceindependent excitation. Each point represents the average total thickness and average maximum fluorescence intensity for three samples of a multilayer protein film ranging from zero to 10 layers. Error bars represent one standard deviation. Since the measurements are taken using the same protein substrates, the thickness and standard deviation in Figures 5 and 6 are identical. Qualitatively, the integrals of eqs 3 and 4 closely represent the measured fluorescence intensity using both fluorescence microscope and surface plasmon excitation. R2 values calculated from the experimental data and the integral of eqs 3 and 4 were 0.995 and 0.987, respectively. Equations 1 and 4 are also shown in Figure 5, normalized such that the excitation intensity at the
Langmuir, Vol. 24, No. 21, 2008 12309
interface, Io, and the yield at infinite separation, Φo, are unity. Similarly, eq 3 is shown in Figure 6; however, since the excitation is distance-independent, the normalized excitation at all separations is unity. Equation 1 shows the exponential dependence of the excitation intensity using surface plasmons, which approaches zero at large separations. In contrast, the efficiency of fluorescence (eq 3) is negligible at close proximities to the metal, while it approaches unity at larger separations. Since eq 4 is the product of eqs 1 and 3, the intensity will have a maximum at intermediate separations but will approach zero at both extremes. In addition to the good qualitative fit and high R2 values, the normalized fluorescence intensity of eq 4 from Figure 5 closely resembles previous experimental data22,26 of fluorescence intensity versus separation; thus, we are confident we have accurately represented the distance dependence relationship of the fluorescence. To obtain kinetic information on the substrate thickness from the fluorescence signal, we must first normalize the data by the changing surface intensity and, second, account for variation in the fluorescence yield due to its distance dependence from the metal surface. We first account for the surface plasmon intensity, as this affects the fluorescence intensity proportionately, independent of the distance from the metal. The reflectivity and fluorescence angular scans shown in Figure 7a were measured before and after enzyme exposure (see Figure 3b). From the reflectivity measurements, we determine the thickness of the protein substrate. The substrate has clearly been degraded, as seen by the lower resonance angle. From the fluorescence measurements, we see that the peak intensity is much lower, consistent with the loss of fluorescent material. As seen in Figure 7a, the fluorescence intensity varies with the sample angle while during each scan the film thickness remains constant. The goal is to decouple the changes in fluorescence intensity caused by surface intensity and those caused by film removal. At this point, we would like to establish a relationship between a measurable quantity and the surface plasmon intensity such that the kinetic data can be normalized using this relationship. The surface plasmon intensity is visualized through the fluorescence intensity. From Figure 7a, the maximum intensities of the two fluorescence scans occur at considerably different angles; thus, no connection can be inferred between the surface plasmon intensity and the incident angle. However, to a first-order approximation, the fluorescence increases with decreasing reflectivity, a consequence of using plasmon excitation. Since directly reflected light does not excite surface plasmons, it does not contribute to fluorophore excitation. The slight disparity between the angle of minimum reflectivity and the angle of maximum fluorescence intensity arises from interference between directly reflected light and re-emitted surface plasmons. Of course, fluorescence intensity is an indirect method of visualizing the surface plasmon intensity and, therefore, only provides proportionality, not an absolute assessment of the intensity. The plot of fluorescence intensity versus reflectivity is shown in Figure 7b. The upper curve represents 100% substrate and corresponds to the thickness of the film before enzyme exposure (43.4 nm), as determined from the reflectivity scan. The same is true for the lower curve, which corresponds to the thickness of the protein substrate after enzymatic degradation and enzyme removal (28.4 nm). The fluorescence intensity of the thicker film is obviously greater; however, this should not affect the proportionality R at constant surface plasmon intensity. At this point, the data will show a small angular dependence caused by the angular dependence of the evanescent decay length, since the data points are collected at different angles. This dependence can be removed by dividing the measured fluores-
12310 Langmuir, Vol. 24, No. 21, 2008
Feller et al.
Figure 7. (a) Reflectivity (9) and fluorescence intensity (0) angular scan of BSA multilayer. Reflectivity (b) and fluorescence intensity (O) angular scan of BSA multilayer exposed to active subtilisin. (b) Fluorescence intensity versus reflectivity from data (a) before reaction (9) and after reaction (O). (c) R versus reflectivity before reaction (9) and after reaction (O) and polynomial trend line of R (s). (d) R versus reflectivity before reaction (9), fluorescence intensity (normalized to values similar to R) versus reflectivity before reaction (O), and polynomial trend line of R (s).
cence intensity by the integral over the fluorescently labeled film using the appropriate decay length from eq 2 for each data point.
R(R) )
Ifluor 1 e-2z/ld dz n zfluor 1 + (β/z)
∫
(5)
Figure 7c shows R as a function of reflectivity, R, which we will call the normalization curve. As stated previously, R accounts for quantities such as Io and Φo, which are affected by the concentration and yield of the fluorophore, as well as surface plasmon intensity. Thus, we can now predict the relative surface plasmon intensity from the measured reflectivity. The data, R versus reflectivity, are fit to a sixth-order polynomial, also shown in Figure 7c. The fluorescence signal of the kinetic mode is divided by the appropriate R value for the given reflectivity, yielding a fluorescence value free of surface intensity contributions. The next step is to account for the distance dependence of the fluorescence intensity due to evanescent wave excitation and alternative de-excitation pathways. The de-excitation process of a fluorophore in the presence of a thin metal film is complex. At short distances, much of the energy is dissipated as heat in the metal film. Slightly larger distances will generate red-shifted plasmons in the metal that can re-emit through the prism. These red-shifted plasmons are called surface plasmon couple emissions (SPCE) and are well-
documented throughout the literature.23,25-27,29,30 SPCEs have been shown to reach a maximum slightly above 30 nm and have been visualized at thicknesses up to 60-70 nm.26,29 Theoretically, these reemitted surface plasmons are predicted to be present to a moderate extent at distances of 80 nm.26 Separations larger than 80 nm primarily result in fluorescence emission. Figures 5 and 6 corroborate the existence of fluorescence quenching using both surface plasmon and fluorescence microscope excitation. Since the films used in this study are thinner than the distance over which quenching acts, fluorescence intensity will vary with distance from the metal surface. The normalized fluorescence from the kinetic mode is equal to the product of the quenching relationship and the exponentially decaying evanescent field integrated over the thickness of the film.
Ifluor/R(R) )
∫z
fluor
1 e-2z/ld dz n 1 + (β/z)
(6)
The fluorescence signal is not required to be normalized in the previous step, since R could have been placed in the integral. We did this to emphasize that R is not distance-dependent and (29) Gryczynski, I.; Malicka, J.; Nowaczk, K.; Gryczynski, Z.; Lakowicz, J. J. Phys. Chem. B 2004, 108, 12073–12083. (30) Gryczynski, I.; Malicka, J.; Gryczynski, Z.; Lakowicz, J. J. Phys. Chem. B 2004, 108, 12568–12574.
Fluorescence Quantification for SP Excitation
therefore can be accounted for at any point of the quantification. During the kinetic mode, all reflectivity and fluorescence intensity values are collected at a constant angle; therefore, the decay length will remain constant throughout this portion of the experiment. The lower limit of the integral depends on the attachment of the fluorescent film to the gold. For our studies, we are using a thiol linker to covalently attach the first layer of BSA; therefore, the lower limit of the integration will be the thickness of the thiol monolayer. The upper limit of the integration that yields a value equal to the measured fluorescence intensity will be the thickness of the substrate film. This calculated upper limit, substrate thickness, can be seen in the adsorption and reaction data of inhibited and active enzyme in Figure 3c,d. Deviations between the raw and quantified data become obvious when comparing Figure 3b,d. The deviation becomes most apparent at the end of the reaction, where the change in surface plasmon intensity is the most pronounced. Near the end of the reaction, the raw data corresponds to an enzyme adsorption value of approximately 8.9 nm, whereas the corrected value is approximately 4.8 nm, giving an 85% error in the measurement. Simplifying Assumptions. It is possible to simplify the quantification procedure by ignoring the angular dependence. This is equivalent to assuming a constant decay length of the evanescent field, which is commonly employed. Therefore, instead of using R(R) from Figure 7c to obtain a normalization curve for the surface intensity, we will use the fluorescence intensity versus reflectivity from Figure 7b. The data from Figure 7b prior to enzyme exposure have been adjusted proportionately to values similar to R(R) so that the effect of assuming a constant decay length can be seen in Figure 7d. The kinetic data can be adjusted, as was done previously to yield values free from surface intensity changes. To simplify the analysis further, we chose to fit the measured fluorescence intensity as a function of film thickness to a fifth-order polynomial trend line instead of the integral of eq 4. It is no longer necessary to separate the yield due to quenching and the exponential decay, since we are assuming a constant decay length. This trend line then accounts for both the energy transfer to the metal film as well as the exponentially decreasing excitation intensity. The entire film has been fluorescently labeled; therefore, this relationship represents the integral of the fluorescence intensity of an infinitely thin film of fluorophores as a function of separation analogous to the integral
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of eq 4. We chose to set the coefficient of the linear term in the fifth-order polynomial equal to zero because the derivative of this trend line should become zero as the separation distance between the fluorophore and the metal approaches zero. Physically, this is interpreted as 100% energy transfer between the excited fluorophore and the metal film. This has been shown both experimentally and theoretically.22,26-28 Once the kinetic data have been adjusted for the surface intensity using the normalization curve, the thickness can be found using the polynomial trend line. The substrate thickness is calculated from the fluorescence intensity as described above. The maximum difference in substrate thickness throughout the experiment is 0.18 nm, leading to an adsorption difference of 4.6% at this point in the reaction compared with the previous quantification. If it is experimentally difficult to obtain fluorescence data as a function of thickness, there are certain regimes where this can be approximated using previous experimental data. For instance, if the experiment occurs over a range of 15-30 nm, the fluorescence yield increases approximately linearly with separation distance, as seen in Figure 5 as well as previous experiments.22,26 Alternatively, the distance dependence can be ignored if the fluorescence yield remains constant over the range of thicknesses measured during the experiment. This approximation is reasonable where the increase in fluorescence due to greater separation from the metal is balanced by the decaying evanescent field, which occurs from about 45 to 60 nm.20,26
Conclusion We provide a method to account for surface intensity variations in SPFS measurements. From this, the thickness of a fluorescently labeled substrate can be measured. This method allows multicomponent surface processes to be decoupled for kinetic mode SPR/SPFS over significant thickness changes. We have also used this method to measure the total film thickness and substrate thickness while a protease reacts with and degrades a protein substrate. From these measurements, it is possible to determine both the thickness of adsorbed enzyme and the rate of substrate degradation. To our knowledge, this is the first account of a means to compensate for surface intensity variations, thereby making these techniques applicable to numerous other multispecies processes. LA8013943
