Anal. Chem. 1996, 68, 1451-1455
Dynamic Surface Events Measured by Simultaneous Probe Microscopy and Surface Plasmon Detection Xinyong Chen,†,‡ Martyn C. Davies,*,§ Clive J. Roberts,*,§ Kevin M. Shakesheff,§ Saul J. B. Tendler,*,§ and Philip M. Williams§
Department of Applied Physics, Chongqing University, Sichuan, 630044, China, and Laboratory of Biophysics and Surface Analysis, Department of Pharmaceutical Sciences, The University of Nottingham, Nottingham NG7 2RD, U.K.
The in situ study of dynamic surface events has significant implications for developing a deeper understanding of macromolecular interactions at or near the solid-liquid interface. Here we describe a new approach to studying this interface using a novel combined surface plasmon resonance and surface force microscope (SPR-SFM) which allows the simultaneous in situ acquisition of the surface kinetic and topographic data. We illustrate the potential of this instrumentation in two key fields of biomedical research, polymer surface degradation and surface adsorption phenomena. The instrument allows the determination of the kinetics of nanometric changes in the thickness of the polymer films and the visualization of the corresponding surface topographical changes on polymer erosion. The kinetics of the adsorption of the protein fibrinogen to a polymer interface are realized to molecular resolution. In conclusion, we believe that the synergistic combination of SPR and SFM offers an elegant approach to the quantitative study of dynamic surface events at the molecular level. Surface biomolecular interactions are the principle feature of many important molecular recognition events within fundamental and applied fields of biomedical science including the study of receptor-ligand association and the design of thin-film technologies for novel immunosensors.1 In order to gain some understanding of the structure-activity relationships for such recognition events, a range of biophysical and surface analytical techniques2 has been employed to define the interfacial chemistry and interactions of both biomolecular systems and synthetic materials. In this regard, scanning force microscopy (SFM) has emerged over the last five years as a powerful tool for highresolution three-dimensional imaging of surfaces exposed to a variety of conditions including air and water.3 SFM studies have made important contributions to our understanding of biomolecular structure4 and association,5 the forces of interactions between †
Chongqing University. Professor Chen is a visiting scientist at the LBSA, The University of Nottingham. § The University of Nottingham. (1) Malmquist, M. Nature 1993, 361, 186-187. (2) Andrade, J. D. In Surface and Interfacial Aspects of Biomedical Polymers; Andrade, J. D., Ed.; Plenum Press: New York, 1985. (3) Drake, B.; Prater, C. B.; Weisenhorn, A. L.; Gould, S. A. C.; Albrecht, T. R.; Quate, C. F.; Cannell, D. S.; Hansma, H. G.; Hansma, P. K. Science 1989, 243, 1586-1589. (4) Yang, J.; Mou, J.; Shao, Z. FEBS Lett. 1994, 338, 89-92. ‡
0003-2700/96/0368-1451$12.00/0
© 1996 American Chemical Society
ligands and receptors,6-8 and the real-time conformational changes of an individual enzyme.9 One concern in employing SFM for the imaging of molecular interactions is the lack of simultaneous quantitative and complementary biophysical data. Here, we report on the design and performance of a new SFM instrument which incorporates a surface plasmon resonance (SPR) device. SPR is currently routinely employed within the biomedical sciences for the study of biomolecular interactions10-12 by monitoring changes in surface refractive index. The new combined instrument integrates the sensitivity of the SPR for the detection of mass transport of picomole quantities to and from a surface with the nanometer resolution imaging of surface events with the SFM. We show that the synergistic combination of SPR and SFM allows the simultaneous in situ real-time quantification and imaging of surface events including surface-molecule and molecule-molecule interactions. We illustrate the potential of the simultaneous in situ acquisition of SPR and SFM data in two key areas of surface science at the solid-liquid interface: polymer surface degradation and surface adsorption phenomena. In the former case, we describe the study of the interfacial erosion of a biodegradable polymer which undergoes surface-mediated acid-catalyzed hydrolysis.13 Such biodegradable polymers are finding important medical, environmental, and biotechnological applications.14 The nature of their degradation has been the subject of considerable interest.15 In the latter case, we illustrate the dynamic adsorption of a plasma protein to a polymeric interface. The protein conditioning of surfaces is of major importance in determining the biocompatability of medical implants16 and in the development of novel biosensors and immunoassays.17 In addition, this approach provides fundamental information on proteins. (5) Rees, W. A.; Keller, R. W. Vesenka, J. P.; Yang, G.; Bustamante, C. Science 1993, 260, 1646-1649. (6) Lee, G. U.; Chrisey, L. A.; Colton, R. J. Science 1994, 266, 771-773. (7) Moy, V. T.; Florin, E. L.; Gaub, H. E. Science 1994, 266, 257-258. (8) Dammer, V.; Popescu, O.; Wagner, P.; Anselmetti, D.; Gu ¨ ntherodt, H.-J.; Misevic, G. N. Science 1995, 267, 1173. (9) Radmacher, M.; Fritz, M.; Hansma, H. G.; Hansma, P. K. Science 1994, 265, 1577-1579. (10) Rahn, J. R.; Hallock, R. B. Langmuir 1995, 11, 650-654. (11) Bondeson, K.; Frostell-Karlsson, A° .; Fa¨gerstam, L.; Magnusson, G. Anal. Biochem. 1993, 214, 145-151. (12) Fischer, B.; Heyn, S. P.; Egger, M.; Gaub, H. E. Langmuir 1993, 9, 136140. (13) Heller, J. J. Controlled Release 1985, 2, 167-177. (14) Peppas, N. A.; Langer, R. Science 1994, 263, 1715. (15) Go ¨pferich, A.; Langer, R. Macromolecules 1993, 26, 4105.
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EXPERIMENTAL SECTION Instrumentation. A schematic diagram of the combined SPR and SFM instrument is shown in Figure 1, and the major design features are described in the corresponding figure caption. From a design perspective, the combination of the SPR with an SFM is attainable since the SPR signal is accessed from below the sample surface and the SFM tip probes the surface topography from above. With this in mind, an SPR with an open sample architecture using the Kretschmann configuration18 was designed and constructed (Johnson & Johnson Clinical Diagnostics, Amersham, U.K.) (Figure 1A). The SFM (Topometrix, Saffron Walden, U.K.) was modified to allow the scanning of the probe and cantilever within a flowing liquid environment through the construction of an enclosed cell. This cell has a volume of ∼300 µL. During experiments, the SPR data were acquired continuously from seven adjacent 3.2 × 104 µm2 sampling areas (Figure 1B) sited 200 µm apart. SFM images, with dimensions of 3 µm × 3 µm or 1 µm × 1 µm, were obtained at selected time intervals. All imaging was performed with silicon nitride probes on triangular cantilevers and a pixel resolution of 200 × 200. To minimize the influence of the flowing liquid on SFM imaging, the syringe pump that initiates fluid flow was switched off during the short time of image acquisition. SFM images are displayed in left shading mode in which the topography is shown as if illuminated at an angle of 45° from the left side of the image. The imaging of polymer degradation was performed in contact mode with a scan rate of 5 Hz. The contact forces between the SFM probe and polymer surface were minimized using data from force-distance curves. The repulsive force between probe and surface was estimated to be less than 1 nN. During the protein adsorption experiments, noncontact mode imaging was utilized to achieve molecular resolution of fibrinogen. For noncontact imaging, a modulation frequency of 124 kHz, an amplitude of 15-18 nm, and a scan frequency of 2 Hz were employed. Sample Preparation. A glass slide coated with a thin silver film (∼50 nm) is required as a sample substrate to ensure the excitation of surface plasmons. Surface phenomenon may be studied on either the native metal film or on overlayer which is adsorbed, covalently coupled, or, as in this case, coated onto the silver surface. The polymer sample used for both the degradation and protein adsorption experiments was a poly(ortho ester) (POE) synthesized by the copolymerization of 3,9-bis(ethylidene)-2,4,8,10-tetraoxaspiro[5.5]undecane with 1,5-pentanediol as described previously.13 Coating was achieved by dissolving the POE in chloroform (10 mg/mL) and spin casting the solution onto the SPR sensor. Continuous films with thickness of ∼100 nm were generated, as confirmed by SFM, X-ray photoelectron spectroscopy, and ellipsometry, and used within 1 h of preparation. Surface Degradation. Preliminary stability studies were undertaken with the polymer film exposed to a 10 mM phosphate buffer, pH 7.4. The degradation experiment was then initiated by adjusting the solution under constant flow to pH 4.0 and subsequently to pH 4.5 after 27 min (potassium hydrogen phthalate buffer adjusted with 0.1 M HCl).
Figure 1. Schematic diagram of the SPR/SFM configuration (A, top) (with inset showing SFM probe) and an illustration of the relative sampling areas of SFM and SPR (B, bottom). The key to the diagram A is as follows: A, SFM laser; B, split diode detector; C, SFM approach mechanism; D, SFM scanner tube; E, liquid inlet; F, liquid outlet; G, silicon O-ring; H, semicylindrical prism; I, silver-coated SPR slide; J, SPR top plate; K, SPR laser; L, polarizer; M, ccd detector. An SPR with open sample architecture was designed and constructed from an instrument supplied by Johnson & Johnson Clinical Diagnostics (Amersham, U.K.) to allow access to the sample for the SFM scanner and cantilever. The SPR was positioned on the underside of a heavy gauge adaptor plate at a point where an opening for the silver-coated sample slide was located. The mass of the plate helped to reduce mechanical vibration induced noise in the SFM data. The SPR power supply, cooling system, and other electrical components were also positioned external to the combined instrument in a remote housing to further diminish noise effects. Slots were milled into the SPR top plate to allow the precise location of the AFM and hence probe over the SPR slide. The SFM component consists of a modified TopoMetrix Explorer (Topometrix, Saffron Walden, U.K.), a stand-alone system which scans the cantilever as opposed to the sample. The macor housing on the piezo scanner was beveled at its edges to secure a soft silicone O-ring on the scanner tube. This makes contacts with the SPR slide thus forming an enclosed liquidtight cell (SPR/SFM sample cell). The softness of the O-ring avoids excessive transmission of vibrations from the SPR to the SFM. Two small channels were drilled through the piezo scanner macor housing, one allowing fluid to flow in and the other out of the SPR/SFM sample cell with a maximum flow rate during imaging of 10 µL/s. The SFM is capable of imaging in either contact or noncontact modes using one of two scanner heads with ranges of 150 µm × 150 µm or 3 µm × 3 µm for low- and high-resolution studies, respectively.
(16) Williams, D. F. In Materials Science and Technology, 14. Medical and Dental Materials; Cahn, R. W., Haasen, P., Kramer, E. J., Eds.; Vol. Ed. Williams, D. F.; VCH: Weinheim, Germany, 1992. (17) Nylander, C.; Liedberg, B.; Lind, T. Sens. Actuators 1982, 3, 79. (18) Kretschmann, E. Z. Phys. 1971, 241, 313-317.
Protein Adsorption. A bovine fibrinogen solution (10 or 100 µg/mL) in 10 mM phosphate buffer, pH 7.4, was pumped at 1 µL/s over the POE polymer film.
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A
Figure 2. SPR/SFM analysis of polymer degradation. SPR data were acquired continuously from seven spots on the sample surface and plotted as (A) the averaged SPR angle as a function of time. SFM images B-J (all 3 µm × 3 µm) were acquired at selected time intervals [indicated by arrows on (A)]. (B-J) reduced 50% of original for publication.
RESULTS AND DISCUSSION Polymer Degradation. The simultaneous SPR and SFM analysis of a POE13 polymer film undergoing hydrolytic degradation is shown in Figure 2. The polymer was initially studied at pH 7.4 for over 4 h to confirm the stability of the film by both techniques under nondegrading conditions. A pH 4.0 aqueous buffer solution was then introduced into the sample cell under constant-flow conditions. The SPR data (Figure 2A) show that the surface degradation of the polymer is immediately initiated within this acidic environment. A continuous linear reduction in the SPR shift is observed which suggests the thinning of the polymer film. On erosion, the hydrolytic cleavage of the ortho ester bonds generates water-soluble oligomeric molecules leading to film disruption. At 27 min, the pH of the solution was increased from 4.0 to 4.5. The anticipated reduction in the rate of erosion is clearly evident. The hydrolysis continues at this slower rate until the film is exhausted. The SFM data in Figure 2B-J show clear evidence of the nature of the topographical degradation of the polymer within the acid environment. The polymer erosion
is characterized from early stages in the experiment by the emergence of erosion pits distributed over the otherwise smooth featureless polymer surface. These features act as foci for further local erosion, leading to a widening and an increase in depth of pits with time,19 resulting in the gradual exposure of the underlying silver substrate. The pit continued to widen until (Figure 2J) all polymeric material has been removed and the topography is consistent with that expected of the silver substrate. In order to quantify the interfacial erosion process, the kinetics of surface degradation were derived from the SPR data. For a 1 µL flow rate, the SPR data yielded a resonance peak shift rate of 0.084 and 0.021°/min (Figure 2A) for pH 4.0 and 4.5, respectively, which corresponds to a 4-fold reduction in polymer degradation rate on changing to the more alkaline pH. The reduction in polymer degradation rate at higher pH is expected from the mechanism of hydrolysis of the poly(ortho esters), which is promoted by the protonation of the oxygen atom linking the diol and the diketene acetal monomer units. The rapid identification of the pH dependence of the hydrolysis of the poly(ortho ester) confirms the potential of the SPR technique to quantify erosion kinetics. It should be noted that the SPR data provide no information on the nature of the surface topography changes occurring during degradation. In contrast, the SFM images reveal clearly the heterogeneity of this process with preferential erosion occurring at points on the polymer surface creating pitted structures. The mechanism by which certain points on the polymer surface degrade to form pits is not currently known. Comparison of the position of pit formation and the underlying silver topography provides no evidence of silver morphological features promoting pit formation. Kinetic analyses of polymer degradation may be obtained from the SFM data20 but are limited due to the small sampling area within the image. Replicate SFM experiments from different regions revealed a range of kinetic erosion values around the mean of the SPR data, reflecting the heterogenous nature of the polymer degradation across the surface. Hence, the simultaneous acquisition of the SPR and SFM data allows us to pinpoint not only the kinetics of the polymer erosion but also the mechanism of surface degradation at each time point. Such an approach may be extended to forms of polymer treatments and synthesis including plasma technology. The three-dimensional nature of the SFM data provides a novel method of comparing the kinetics of polymer degradation revealed by both the SFM and SPR techniques. We previously described computational methods of calculating the change in polymer matrix volume during degradation from SFM data.20 Applying these methods to the SFM data of poly(ortho ester) degradation displayed in Figure 2 reveals a good agreement between SFM and SPR recorded changes in the kinetics of polymer film erosion. Figure 3 displays the SFM recorded change in volume plotted over the SPR angle change. This graph highlights that both SFM and SPR data reveal the slowing of polymer hydrolysis when the environment becomes more basic in pH. Protein Adsorption. Data from passing fibrinogen solutions (10 and 100 µg/mL) through the solution cell at a flow rate of 1 µL/s are presented in Figure 4 to illustrate how the SPR-SFM (19) Shakesheff, K. M.; Davies, M. C.; Domb, A.; Roberts, C. J.; Shard, A. G.; Tendler, S. J. B. Langmuir 1994, 10, 2654-2655. (20) Shakesheff, K. M.; Davies, M. C.; Heller, J.; Roberts, C. J.; Tendler, S. J. B.; Williams, P. M. Langmuir 1995, 11, 2547-2553.
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A
Figure 3. Comparison of rate of polymer erosion as measured by SPR (solid line) and volume analysis of SFM data (black squares). The methodology of volume analysis is detailed in ref 20.
instrument may elucidate the different kinetics of adsorption and lateral surface distribution. The concentrations of the protein solutions were selected to achieve monolayer and submonolayer adsorption over the time scale of the experiment. Any difference in adsorption behavior for the two solutions will reflect the influence of the increasing concentration gradient promoting the rate of adsorption. It is evident from Figure 4A that the SPR data display the expected increase in the rate of adsorption with increasing protein concentrations. For the 100 µg/mL solution, a rapid Langmuirian adsorption profile is observed. The maximum SPR angle shift of ∼0.35° is in good agreement with previous studies of protein adsorption.21 As before, SFM images were taken at selected time intervals during the absorption profile. In agreement with the SPR data for the high protein concentration solution, a densely packed layer of protein molecules was observed in the SFM data from early time points over all regions examined and this layer was continuous (Figure 4B). In contrast, the data for the lower protein concentration show a slower linear increase in the SPR shift (Figure 4A), attaining a maximum shift after 3 h of around 0.170°, some 60% below the value from the higher protein solution. These findings were confirmed by a series of SFM images where a submonolayer coating of the protein was observed at all stages of the experiment (Figure 4C). The individual fibrinogen molecules display an average diameter of 40 nm 22 and are clearly observed on the smooth polymer surface in Figure 4C. The lateral distribution across the surface appears to be random with some trimer and dimer clusters. The surface density of protein adsorption varied from region to region, particularly at early time points. Again, in agreement with the SPR data, the SFM images did show a gradual increase in surface adsorption with time during the course of the experiment.
Figure 4. Study of fibrinogen adsorption to the POE surface. Averaged SPR angle as a function of time is displayed (A). The arrows labeled B and C indicated the times at which the SFM data in Figure (B) and (C) were acquired.
CONCLUSIONS In this paper, we have demonstrated that the simultaneous acquisition of the surface refractive index and topographic data provides a unique insight into the dynamic changes in the interfacial chemistry associated with two important biomedical systems. The wider implications of the exploitation of this instrument in both fundamental and applied surface science are profound. The SPR-SFM could be employed to examine many
important systems which exploit dynamic surface interactions including ligand-receptor recognition in immunosensors,1 epitope mapping,23 formation and stability of self-assembled monolayers,24 development of nanoengineered devices, and assessment of stimuli sensitive (or so-called “smart”) materials.25 The instrument may also be simply adapted to include other forms of scanning probe technology including SNOM26 or its potential extended by new applications of SFM including measuring forces of molecular interaction or real-time imaging. In conclusion, we believe that the synergistic combination of SPR and SFM offers an elegant approach to the quantitative study of dynamic surface events at the molecular level.
(21) Davies, J.; Allen, A.; Burrus, Y.; Bruce, I.; Heaney, P. J.; Hemmings, F. A.; Nunnerley, C. S.; Skelton, L. In Surface Properties of Biomaterials; West, R., Batts, G., Eds.; Butterworth-Heinemann Ltd.: Oxford, U.K., 1993. (22) Wigren, R.; Eluring, H.; Erlandsson, R., Welin, S.; Lundtrom, I. FEBS Lett. 1991, 280, 225-228.
(23) Briggs, S.; Price, M. R.; Tendler, S. J. B. Immunology 1991, 73, 505-507. (24) Abbott, N. C., Gorman, C. B.; Whitesides, G. M. Langmuir 1995, 11, 1622. (25) Chen, G.; Hoffman, A. S. Nature 1995, 373, 49-52. (26) Vanhulst, N. F.; Deboar, N. P.; Bolger, B. J. Microsc. 1991, 163, 117-130.
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ACKNOWLEDGMENT The authors acknowledge the support of the BRITE/Euram programme, the BBSRC Biotechnology Directorate, the EPSRC/ DTI Nanotechnology LINK programme, VG Microtech, Kodak Limited, and Oxford Molecular Group plc. X.C. thanks the State Education Commision of China for fundings his sabbattical within the Laboratory of Biophysics and Surface Analysis. Thanks to (27) Guthold, M., Bezanilla, N.; Erie, D. A.; Jenkins, B.; Hansma, H. G.; Bustamante, C. Proc. Natl. Acad. Sci. U.S.A. 1994, 91, 12927-12931.
Dr. J. Davies (Johnson & Johnson Clinical Diagnostics Ltd., Amersham, U.K.), Dr. S. Sharma (Amersham International Plc., Amersham, U.K.), and Dr. R. Gamble (Topometrix Corp., Santa Clara, CA) for their valuable help in instrument design. Received for review August 21, 1995. Accepted January 26, 1996.X AC950844E X
Abstract published in Advance ACS Abstracts, March 1, 1996.
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