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Irreversible Adsorption of Lysozyme to Polishing Marks on Silica Cozette M. Cuppett, Leon J. Doneski, and Mary J. Wirth* Department of Chemistry & Biochemistry, University of Delaware, Newark, Delaware 19716 Received November 26, 1999. In Final Form: April 17, 2000 Fluorescence microscopy, in combination with atomic force microscopy (AFM), reveals that rhodaminelabeled lysozyme adsorbs strongly from aqueous solution to polishing marks of nanometer depths on fused silica surfaces. The fluorescence and topographical images correspond closely. Fluorescence intensity varies by as much as 2-fold over the same region where the topography varies over 2.5 nm. Desorption of lysozyme from the surface occurs on rinsing with solutions of increased ionic strength or decreased pH, showing that reducing the Coulombic attraction enhances desorption. Different polishing marks behave differently with respect to desorption: the fluorescence pattern on the surface changes when the ionic strength of the rinsing solution reaches 0.1 M, indicating that some polishing marks are more strongly adsorptive than are others. The desorption depends on pH, which is consistent with the known variation in the charge of silica surface. A superpolished silica photomask, which has significantly fewer polishing marks on the nanometer scale, has significantly less adsorption of lysozyme, suggesting that the topography on the nanometer scale influences adsorption.
Introduction Understanding how proteins adsorb to surfaces is critical to their economical purification and their analysis in pharmaceutical, clinical, and research laboratories. The adsorption of proteins to surfaces is presently the reason capillary electrophoresis has not fully realized its potential as a method for rapid analysis of proteins.1 A considerable effort is underway to devise protective, hydrophilic coatings for capillaries2-4 and microchips5 to be used for fast electrophoresis of proteins. Another opportunity impeded by the lack of understanding of protein-surface interactions is the design of protein-based biosensors that require protein orientation.6-9 Protein adsorption is crucial to directing cell growth on surfaces because the attachment of cells occurs through a protein-surface interactions.10 Protein adsorption has caused long-standing problems in a diverse set of technologies. One example is the corrosion of heat-exchangers, where biofilms form on metal surfaces and secrete corrosive acids.11 Another is the development of biocompatible materials for medical implants, which either fail or must frequently be serviced because of adsorption of fibrinogen or other proteins.12 If all factors influencing the adsorption of proteins were understood, many technologies would be advanced. Lysozyme is an antimicrobial protein that is prevalent in ocular fluid, thus adsorption of lysozyme to contact * Corresponding author. (1) Watzig, H.; Degenhardt, M.; Kunkel, A. Electrophoresis 1998, 19, 2695-2752. (2) Hjerte´n, S. J. Chromatogr. 1985, 347, 191-198. (3) Bruin, G. J. M.; Chang, J. P.; Kuhlman, R. H.; Zegers, K.; Kraak, J. C.; Poppe, H. J. Chromatogr. 1989, 471, 429-436. (4) Towns, J. K.; Regnier, F. E. J. Chromatogr. 1990, 516, 69-78. (5) He, B.; Tan, L.; Regnier, F. E. Anal. Chem. 1999, 71, 1464-1468. (6) Jones, V. W.; Kenseth, J. R.; Porter, M. D.; Mosher, C. L.; Henderson, E. Anal. Chem. 1998, 70, 1233-1241. (7) Edmiston, P. L.; Lee, J. E.; Cheng, S. S.; Saavedra, S. S. J. Am. Chem. Soc. 1997, 119, 560-570. (8) Keating, C. D.; Kovaleski, K. M.; Natan, M. J. J. Phys. Chem. B 1998, 102, 9404-9413. (9) Ratner, B. D. J. Biomat. Sci.-Polym. Ed. 1992, 4, 3-11. (10) Boncheva, M.; Duschl, C.; Beck, W,; Jung, G.; Vogel, H. Langmuir 1996, 12, 5636-5642. (11) Flemming, H. C. Water Sci. Technol. 1993, 27, 1-10. (12) Tang, L. P.; Eaton, J. W. Am. J. Clin. Path. 1995, 103, 466-471.
lenses is a considerable problem in ophthalmology.13 Lysozyme is a strongly cationic protein at neutral pH, it has dimensions of 3.0 × 3.0 × 4.5 nm3, and it is a hard protein up to ionic strengths of at least 0.4, based on studies of crystal structures.14 The adsorption of lysozyme to silica at neutral pH has been studied previously, using such varied methods as in situ ellipsometry,15 neutron reflection,16,17 fluorescence with total internal reflection,18 and ultraviolet (UV) absorbance.19,20 In each study, lysozyme adsorbed to silica surfaces to concentrations on the order of a monolayer or more, and increased ionic strength reduced the amount adsorbed. Also, in each study, the adsorption of most of the lysozyme was thermodynamically irreversible (i.e., on rinsing with a solution of the same composition minus the protein, a significant amount of lysozyme remained adsorbed). The exact amount of irreversibly adsorbed lysozyme varies from one report to the next. Three studies of flat silica surfaces reported some fraction is reversibly adsorbed,15-18 whereas one group studying colloidal silica reported none of the adsorption was reversible.19,20 Two studies reported that some of the irreversibly adsorbed lysozyme remained after rinsing under more aggressive conditions, such as higher ionic strength or higher concentration of protonated base,15,18 whereas two others studies showed that none remained.16,17,19,20 The disagreement among these studies indicates that there are unknown variables that affect the adsorption of lysozyme on silica. Recently, there have been numerous reports that surface topography influences the attachment and growth of biological cells. These studies have recently been reviewed, (13) Garrett, Q.; Garrett, R. W.; Milthorpe, B. K. Invest. Ophth. Vis. Sci. 1999, 40, 897-903. (14) Bell, J. A.; Wilson, K. P.; Zhang, X. J.; Faber, H. R.; Nicholson, H.; Matthews, B. W. Proteins 1991, 10, 10-21. (15) Wahlgren, M.; Arnebrant, T.; Lundstro¨m, I. J. Colloid Interface Sci. 1995, 175, 506-514. (16) Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F.; Penfold, J. J. Colloid Interface Sci. 1998, 203, 419-429. (17) Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F.; Penfold, J. Langmuir 1998, 14, 438-445. (18) Robeson, J. L.; Tilton, R. D. Langmuir 1996, 12, 6104-6113. (19) Norde, W.; Anusiem, A. Colloids Surf. 1992, 66, 73-80. (20) Norde, W.; Favier, J. P. Colloids Surf. 1992, 64, 87-93.
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and when taken together, micron-scale topography appears to be the important factor in microbial adhesion.21 Because microbial adhesion occurs through proteinsurface interactions, it is conceivable that topography also influences the adsorption of proteins. Large proteins, such as fibronectin, have been observed to adsorb to grooves and ridges on the micron scale.22 A relation between protein adsorption and surface topography is indicated by studies of plasma proteins, such as fibronectin and albumin, adsorbed in different amounts to colloidal latex spheres with different morphology.23 These various studies suggest that the adsorption of proteins can be influenced by nanoscale surface topography; therefore, topography could be a factor in the adsorption of lysozyme. In this work, to investigate the role of topography in the irreversible adsorption of lysozyme on silica, lysozyme is labeled with a fluorescent dye, enabling the use of fluorescence microscopy. Fluorescence imaging is used to observe the spatial heterogeneity of the adsorbed lysozyme, and atomic force microscopy (AFM) is used to determine the surface topography of the same region, allowing adsorptivity and surface topography to be compared. Both ionic strength and pH are varied in the rinsing steps to investigate desorption and its spatial heterogeneity.
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Figure 1. Schematics for optical experiments. (a) Upright optical microscope using evanescent-wave excitation of fluorescence. The fluorescence was collected with a 10X objective and imaged onto a CCD camera. (b) Inverted optical microscope combined with an atomic force microscope. A 40X microscope objective was used to collect fluorescence and to observe the position of the AFM tip. The AFM tip was positioned to scan over the same region as the fluorescence.
Experimental Section Protein Tagging. Three-times crystallized, dialyzed, and lyophilized lysozyme from chicken egg white (Sigma) was labeled with a fluorescent probe, tetramethylrhodamine isothiocyanate (TRITC). The procedure used to conjugate the terminal amino group of lysozyme with tetramethylrhodamine-5-isothiocyanate (Molecular Probes, 5-TRITC; G isomer) was a variation of the methods described by Brinkley.24 A 10 mg/mL solution of lysozyme was made in a pH 7.5 phosphate buffer of ∼50 mM in ionic strength. With stirring, TRITC dissolved in dimethylformamide (DMF; Fisher) was added slowly to the protein solution such that the final concentration of dye in solution was 0.17 mg/mL. A buffer solution at pH 7.5 was used to selectively react the probe with the R-amino groups of the protein.24 The reaction was allowed to proceed with stirring at room temperature for ∼1.5 h. The reaction was stopped by adding 0.1 mL of 1.5 M hydroxylamine (Aldrich) dissolved in phosphate buffer. The solution was stirred for another hour and then separated on a Sephadex G25 Medium column (Amersham Pharmacia Biotech). The protein solution was made to be 10 mM in sodium azide (Aldrich) to avoid bacteria growth. Protein solutions were protected from light and refrigerated until further use. UV absorbance maxima of separate solutions of lysozyme and TRITC were used to create a calibration curve from which the concentration of the protein, as well as the dye-to-protein ratio, in the protein solution was determined. The dye-to-protein ratio was 1.2, which is consistent with tagging just the terminal amino group of the protein. This procedure provided a stock solution of 80 µM in labeled lysozyme, and this stock solution was used to prepare fresh solutions of typically 100 nM. All water used in this work was purified to a resistance of 18-MΩ-cm using a commercial purification system (Barnstead E-pure). The ionic strengths of the final 100-nM solutions of lysozyme were calculated to be 2.5 µM because of the dilution of the sodium azide. Silica. The fused silica (ESCO Products) consisted of prisms and coverslips with scratch-dig specifications of 20-10 and 6040, respectively. A fused-silica photomask was donated by Dupont Photomasks, Inc. All silica samples were cleaned by first wiping with lens tissue to remove debris, then by immersing in boiling methanol, and finally by boiling the specimen in solution made by mixing equal volumes of pure water and concentrated nitric (21) Curtis, A.; Wilkinson, C. Biomaterials 1997, 18, 1573. (22) den Braber, E. T.; de Ruijter, J. E.; Ginsel, L. A.; von Recum, A. F.; Jansen, J. A. J. Biomed. Mater. Res. 1998, 40, 291-300. (23) Luck, M.; Paulke, B. R.; Schroder, W.; Blunk, T.; Muller, R. H. J. Biomed. Mater. Res. 1998, 39, 478-485. (24) Brinkley, M. Bioconjugate Chem. 1992, 3, 2-13.
acid. The cleaned silica surfaces were rinsed with pure water, after which it was confirmed that a film of the water adhered tightly to the surface in each case. As a check to determine whether the experimental results are unique to the procedure of cleaning with nitric acid, new silica samples were cleaned in an ozone plasma, and comparable results were observed. Microscopy. Two types of fluorescence experiments were performed. Evanescent-wave excitation, using an upright microscope (Leica, DMRE), illustrated in Figure 1a, was used to monitor adsorption of lysozyme onto silica as a function of time. A newly purchased equilateral silica prism was used both to control the angle of excitation and to serve as the silica surface under study. The excitation beam had polarization parallel to the plane of the interface (s-polarization) and an 80° angle of incidence, which limits the depth of penetration of the intensity to 110 nm into the aqueous solution. For the filters, a 50:50 beam splitter was used to observe laser scatter, which enabled focusing of the microscope to the silica/water interface. A TRITC filter set was used to monitor fluorescence. Transmission microscopy with an inverted optical microscope (Leica, DMIRBE), illustrated in Figure 1b, was used to measure the topography of the same region observed by fluorescence. For both optical experiments, an aircooled argon-ion laser (Spectra-Physics) was used for excitation light, with a reflection grating used to isolate the 514.5-nm line. The power density was kept below 5 W/cm2 to avoid photobleaching. For both optical microscopes, a liquid-nitrogen cooled CCD camera (Princeton Instruments, TEK 512 × 512B) was used. The topography was characterized with a tip-scanning atomic force microscope (Topometrix, Explorer), operating in contact mode. The AFM was mounted over the sample on the inverted optical microscope, as illustrated in Figure 1b. To minimize vibration of the coverslip, the inverted microscope was mounted on air-cushions (BPG, Stabl-Levl) and on a vibrationisolation table (Newport I-2000 series), and the xy translation stage of the microscope was tightly clamped in place during the topographical measurement.
Results and Discussion Fluorescence images were acquired successively in time after 1 mL of a 100-nM aqueous solution of lysozyme was deposited onto the equilateral silica prism depicted in Figure 1a. The intensity in each image was integrated after subtraction of the small background, and Figure 2 shows a typical plot of the integrated fluorescence intensity as a function of time. The fluorescence intensity rises steadily in time, and reaches a plateau on the time scale
Lysozyme Adsorption to Polishing Marks on Silica
Figure 2. Plots of fluorescence intensity (squares) and relative standard deviation (circles) as a function of time for accumulation of labeled lysozyme onto silica surface. The excitation was polarized in the plane of the surface and no polarization discrimination was used in the emission.
of an hour, in agreement with previous work.15-20 The fluorescence image for the data point in Figure 2 corresponding to 24 s of equilibration, is shown in Figure 3a. This 100 × 100-µm image of Figure 3a reveals that the accumulation of lysozyme is spatially heterogeneous, with numerous lines evident. It is established in a subsequent section of this paper that these lines correspond to nanometer indentations imparted by the polishing process. Figure 3b shows an image of scattered laser light from exactly the same region as for Figure 3a but acquired prior to exposure of the surface to lysozyme. There are vastly fewer lines evident in the image of scattered light and there are more spots. The intense line in Figure 3a, indicated by the black arrow, is close to but not coincident with the line in Figure 3b, indicated by the white arrow. There are no lines common to both images. The spatial pattern of adsorbed lysozyme thus appears to highlight polishing marks that are invisible with scattered light even at near-grazing incidence. The heterogeneity in the fluorescence images was evaluated by calculation of the relative standard deviation of the intensity in each image, after background subtraction, and this quantity is included in the plot in Figure 2. The time-dependence of the relative standard deviation shows that the visibility of the polishing marks develops within the 24-s equilibration time before the first image was acquired, and the visibility gradually diminishes but never goes to zero. To investigate the relation between topography and adsorption, 20 × 20-µm fluorescence and AFM images were acquired for the same region of the surface, using the apparatus depicted in Figure 1b. The AFM cantilever was observed through the microscope eyepieces during its scan to confirm it swept the same region as was viewed by fluorescence. The sample was prepared by equilibrating the silica coverslip for 10 min with a 100-nM aqueous solution of lysozyme, and then rinsed with pure water. A typical fluorescence image is shown in Figure 4a, which again shows that the adsorption is spatially heterogeneous. Figure 4c provides a line analysis of the fluorescence micrograph for the line in Figure 4a, revealing that the fluorescence intensity from pixel to pixel varies by as much as a factor of 2. The fluorescence intensity is higher in the middle of the image because the laser beam has a Gaussian profile. Figure 4b shows the atomic force micrograph of the same region of the surface, after removing the lysozyme with strong acid and then drying the surface with clean
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methanol. Removing the lysozyme was required for minimizing adhesion of the AFM tip to the surface. Comparison of Figures 4a and 4b shows that the fluorescence and AFM images correlate strikingly. Figure 4d is a line scan of the AFM image, showing that the height variations on the surface are on the scale of a few nanometers, explaining why these features were not detectable using light scattering even at near-grazing incidence. The polishing marks on the nanometer scale are more adsorptive to lysozyme than are the polishing marks visible on the optical scale. Some features near the top of the two images of Figure 4 appear in noticeably different places in each image. The reason for this result is that the AFM measurement suffers from hysteresis arising from the piezoelectric crystals that control the x and y positions. Distortion in both the optical and atomic force images was tested by the imaging of a calibration grid. The optical microscope gave the expected image, whereas the atomic force microscope gave an image with distortions as much as 15% of the image size. It is concluded that the main features of the fluorescence and AFM images of Figure 4a and 4b are the same, and that the level of disagreement in the placement of the features is expected based on the hysteresis in the AFM instrument. From a chemical viewpoint, the underlying reason for the topographical influence on adsorption is of the utmost interest. The essential question is whether the adsorptivities of the polishing marks differ from that of flat silica, or rather, whether the polishing marks simply have higher surface area. Comparison of Figures 4a and 4b shows there is not a one-to-one correspondence between surface topography and fluorescence. Some features that are more prominent in one image are less prominent than in the other. One example is a ridge that is clearly visible in the AFM image, marked by the X on the left of Figure 4b. The ridge is not visible in the fluorescence image of Figure 4a. The cause of the correlation, and occasional lack thereof, between topography and adsorptivity is curious. The low spatial resolution of AFM for surfaces that are not atomically smooth causes a considerable loss of topographical detail on the nanometer scale. The estimated radius of the AFM tip is 50 nm, washing out any detail that would provide insight into the topography of the silica on the 3-nm size scale of the protein. A priori, the increased adsorption at polishing marks could be due either to a locally increased surface area or to a locally unique adsorptivity. The fact that the contrast in fluorescence intensity is only 2-fold, as shown in Figure 4c, would make an argument for increased surface area quite plausible. However, the faster rate of adsorption by the polishing marks, shown in Figure 2, indicates that the phenomenon is not simply due to higher surface area. The kinetics of adsorption would be uniform if small variations in the local surface area were the origin of the phenomenon. The desorption behavior of the protein lends further support for the conclusion that the polishing marks have different adsorptive properties than the flat regions of silica. The desorption of lysozyme from a silica coverslip was studied with the inverted fluorescence microscope, and images were obtained for the same region already detailed in Figure 4. The adsorption and rinsing steps were done in the same way as for the experiment that generated Figure 4. Figure 5 shows a series of images, with the top row of images corresponding to the surface prepared each time by the same procedure used for Figure 4a. The similarity among the three images in the top row (Figures 5a, c, and e), which are each on the same intensity scale, illustrates the level of reproducibility of the adsorption of lysozyme. The bottom row shows the resulting
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Figure 3. Images of (a) fluorescence after 24 s of exposure of silica to solution of labeled lysozyme and (b) scattered excitation light. The images are 100 × 100 µm. The arrow in each image denotes a line that appears similar to a line in the other image, but the lines are not coincident (see text).
Figure 4. Comparison of fluorescence and AFM images of the same region, and line scans of each. (a) Fluorescence image of a 20 × 20 µm region. (b) Fluorescence intensity along the horizontal line drawn in part a. The positions of the Xs in part a correspond to the positions of the vertical lines in part b. (c) AFM image of the same region as in part a, nominally 20 × 20 µm, although there is some distortion due to hysteresis of the piezoelectrics. (d) Plot of topography along the horizontal line drawn in part c. The positions of the Xs in part c correspond to the positions of the vertical lines in part d.
images after rinsing the respective surface a second time with a solution with an ionic strength of 0.10 M, adjusted by addition of KCl, and a variable pH. The bottom row of images is on a 15X more sensitive intensity scale than the top row. To examine the case where only ionic strength is varied, a comparison Figures 5a and 5b shows that
rinsing with a solution the same pH of 7 but an ionic strength of 0.10 M gives a dramatic change in the image. Different polishing marks are visible after rinsing with the solution of higher ionic strength. Two lines, denoted by arrows in Figure 5b, are not visible in Figure 5a. The lower of the two lines in Figure 5b is oriented at 42° from
Lysozyme Adsorption to Polishing Marks on Silica
Figure 5. Images before and after desorption of labeled lysozyme using varying pH and ionic strength. (a), (c), (e) images from replicate experiments where adsorbed lysozyme was rinsed with solution of pH 7 and ionic strength of 0.01, (b) image after further rinsing the surface shown in panel a with solution of pH 7 and ionic strength of 0.10, (d) image after further rinsing the surface shown in panel c with solution of pH 4 and ionic strength of 0.10, (f) image after further rinsing the surface shown in panel e with solution of pH 2 and ionic strength of 0.10. Images b, d, and f are on an intensity scale that is 15 times more sensitive than that for images a, c, e. The size scale for each image is 20 µm × 20 µm.
the vertical. This line is not coincident with the line of Figure 5a, which is oriented at 37° and is also denoted by an arrow. The fluorescence signal contributing to these two new lines of Figure 5b must be present in Figure 5a, but the lines could not be seen on the less sensitive intensity scale of Figure 5a. The AFM image matches Figure 5a much better than it does Figure 5b, although the upper line of Figure 5b is readily visible in the AFM image as the ridge that was previously noted as missing. The lower line of Figure 5b is not visible in the AFM, and efforts to find subtle evidence of this line through shadowing and three-dimensional viewing did not reveal its presence. There are numerous other small lines in the AFM image that are not visible in the fluorescence image. Desorption using an even higher ionic strength, 1.00 M KCl, does not reveal any new marks, instead, it removes virtually all lysozyme from the surface. These studies of desorption establish that different regions of the surface have different adsorptivities toward lysozyme, again supporting the conclusion that the adsorption to polishing marks is not simply an effect of increased surface area. The adsorptive behavior of polishing marks thus differs from that of the rest of the surface, and the question is whether this is due to the chemical composition or to the topography of the surface on the scale of the protein. Desorption with rinse solutions with the same ionic strength of 0.10 M but varying pH shows that lower pH enhances desorption of lysozyme. Comparison of Figures 5b, d, and f shows that reducing pH from 7 to 4 to 2 gives progressively more desorption. No new marks are uncovered by varying pH. Fused silica has a pKa on the order of 5,25-27 thus the solutions at pH 2 and 4 would significantly reduce the charge on the silica surface. The significant drop in fluorescence between pH 7 and 4 indicates that the adsorption to these polishing marks is affected by the expected charge on silica. This result argues against the possibility that unusually acidic silanols at edges are the cause for the strong adsorption. The pH (25) Iler, R. The Chemistry of Silica: Solubility, Polymerization, Colloid and Surface Properties, and Biochemistry; Wiley: New York, 1979. (26) Kohr, J.; Englehardt, H. J. Chromatogr. 1993, 652, 309-316. (27) Huang, X.; Kovaleski, J. M.; Wirth, M. J. Anal. Chem. 1996, 68, 4119-4123.
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dependence of desorption is consistent with the polishing marks having the same chemical composition as the flat silica surface. However, the possibility remains that some surface species, with the same pKa as silica but a greater ability to adsorb protein, exists at the polishing marks. Ceria, which is the material used for polishing, has a pKa on the order of 7,28 so the possibility that the adsorptive sites are due to ceria is not eliminated. However, it has been shown that there is a negligible amount of ceria remaining after chemical-mechanical polishing of silica.28 Also, the entire surface has been in contact with the ceria, not just the polishing marks. The possibility that ceria impurities collect at the edges of polishing marks and irreversibly adsorbs lysozyme is absolutely not eliminated by these measurements. The interactions contributing to irreversible adsorption of lysozyme would be short-range attractive interactions between the protein and the surface, such as van der Waals interactions, hydrogen bonding, acid-base, or conformational changes that reduce the solubility of the protein in water. It is possible that topography itself contributes to irreversible adsorption through a higher area of contact between the surface silanols and the hydrogen-bonding groups prevalent on the exterior of the protein or by the local topography promoting denaturation. Nanometerscale topographical measurements with a sharper tip will ultimately be able to test directly the amount by which nanometer morphology affects the adsorption of proteins. One means of testing the importance of the sizes of the indentations is to study a surface that has fewer adsorption sites on the order of the size of the protein. The superpolishing of silica, which is an emerging capability in the semiconductor industry, greatly reduces indentations that are g1 nm in depth by using a ceria slurry with a more narrow particle size distribution. Collaborative attempts with a polishing company to achieve a superpolish on the coverslips were unsuccessful because of the fragility of the coverslips; however, Dupont Photomasks, Inc. provided us with a sample of a fused silica photomask that had been superpolished. Lysozyme was adsorbed to this superpolished surface and rinsed by the same procedure as before. It was more difficult to focus the microscope onto the superpolished surface because of the weaker adsorption of lysozyme. A 75 × 75 µm fluorescence image is shown in Figure 6a, and a 10 × 10 µm AFM image is shown in Figure 6b. The fluorescence and AFM images could not be obtained for the same region because of the 1-cm thickness of the sample. Several AFM images were obtained for different regions of the sample, and the image shown is typical. The fluorescence is not from a typical region; instead, it is from a region that has an unusually large number of features in the fluorescence image. Despite this, the fluorescence image for the photomask still shows much more uniformity than images observed for any coverslip, and the average fluorescence intensity over a typical region of observation is >2-fold lower than for coverslips. The AFM image confirms the much smoother polish. The brightness of the fluorescence along the polishing lines is comparable to that for the coverslips, however, the difference is that the lines are much more rare on the photomask. These polishing lines visible in the fluorescence image appear to be defects caused by ceria aggregates that did not break apart in the polishing machine. The intensity would thus depend on how many points on a ceria aggregate contacted the surface, rather than simply the local adsorptivity. Further, the intensity is an average over the 140 nm dimension of the pixel, (28) Tesar, A. A.; Fuchs, B. A.; Hed, P. P. Appl. Opt. 1992, 31, 71647172.
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Figure 6. (a) Fluorescence image of a 75 × 75 µm area of a silica photomask after adsorption of labeled lysozyme and rinsing with pure water. (b) AFM image of a 10 × 10 µm region of the same surface, although not the same region.
which is much larger than the size of the protein. This averaging allows for the possibility that the local adsorptivity is much greater than two times the average adsorptivity of the flat surface. The weaker and more uniform fluorescence from the superpolished surface suggests that nanometer topography of the polishing marks contributes to the irreversible adsorption of lysozyme to polishing marks. Again, experiments with sharper AFM tips are needed to test this directly. Conclusions The evidence presented here indicates there are two factors contributing to the adsorption of lysozyme to polishing marks. First, charge is a factor, as shown by increased desorption of the lysozyme at both higher ionic strength and lower pH. Second, the fact that different levels of polish greatly change the adsorptive behavior suggests that topography itself is a factor that affects adsorption. The topographical influence on the adsorption of lysozyme to silica offers a possible explanation for the disagreement in the literature on the fractional amount of irreversibly adsorbed lysozyme on silica because surface roughness had not previously been taken into account.
Specifically, the colloidal silica could be rougher on the nanometer scale than the polished silica surfaces. For protein chromatography and for biocompatible materials, where irreversible adsorption of proteins in both cases is an expensive problem that has persisted for decades, these results suggest that manufacturers might improve performance by taking steps to control the nanometer-scale topography of the surface. In devising new surfaces to control protein adsorption on biosensors, topography of the surface is a variable that might be used to control adsorption and orientation. Acknowledgment. The assistance and advice given by Derrick Swinton is greatly appreciated. This research was supported by the Center for Nanomachined Surfaces and the National Science Foundation. We are grateful to Dupont Photomasks, Inc. for donation of the samples from a superpolished silica photomask. Funds for the atomic force microscope were donated by the W. M. Keck Foundation. LA9915419
