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Fibrinogen Adsorption on Three Silica-Based Surfaces: Conformation and Kinetics Alicia Toscano and Maria M. Santore* Department of Polymer Science and Engineering, UniVersity of Massachusetts, Amherst, Massachusetts 01003 ReceiVed June 17, 2005. In Final Form: NoVember 23, 2005
Using AFM (atomic force microscopy) to probe protein conformation and arrangement, and TIRF (total internal reflectance fluorescence) to monitor kinetics, fibrinogen adsorption on three different silica-based surfaces was studied: the native oxide on silicon, acid-etched microscope slides, and acid-etched polished glass. The three are chemically similar, but the microscope slide is rougher and induces AFM tip instabilities that appear as high spots on the bare surface. Fibrinogen’s conformation and transport-limited adsorption kinetics are found to be quantitatively similar on all three surfaces. Further, the number of adsorbed proteins in progressive AFM micrographs quantitatively match the coverages measured by TIRF during early adsorption. Surfaces appear full, via AFM, when adsorbed amounts are about an order of magnitude below their true saturation levels (via TIRF) because, above about 0.26 mg/m2, individual proteins cannot be discerned. The results demonstrate how the appearance of AFM micrographs can be misleading regarding surface saturation. On all three surfaces, fibrinogen is, at most, slightly aggregated, showing limited, if any, surface mobility. The complexities of the microscope slide’s surface landscape minimally impact adsorption.
Introduction Atomic force microscopy (AFM) has provided means to visualize proteins on surfaces, a research topic considered exciting because of the importance of adsorbed proteins in biomaterial interactions. Fibrinogen, in particular, has frequently been a model system in these studies, not only because of its relevance to thrombotic interactions with blood-contacting materials but also because its distinctive shape facilitates unambiguous protein identification in the complex surface landscapes explored by AFM. Fibrinogen is a flexible 340 kDa molecule that is predominantly R-helical and roughly 47 × 4.5 × 4.5 nm3. Its trinodular structure was first visualized by SEM1,2 and decades later using AFM.3 Figure 1 reproduces a schematic of fibrinogen,4 which consists of two symmetrical halves, each containing three polypeptide chains (AR, Bβ, γ). These six polypeptides form fibrinogen’s structure, which is comprised of a central e-domain and two terminal d-domains. The RC arms are extensions of the AR peptides from the d-domains and are negatively charged. The N-terminus of all six polypeptides resides in the central e-domain, giving this domain an overall positive charge which allows electrostatic binding of the RC arms. The cleavage of the end sequences of the AR and Bβ peptides by thrombin in the e-domain results in the formation of fibrin, an activated state of fibrinogen able to associate and polymerize. The past 15 years have seen AFM studies of fibrinogen on a variety of surfaces. Large scanning areas5-7 and high fibrinogen (1) Fowler, W. E.; Erickson, H. P. J. Mol. Biol. 1979, 134, 241-249. (2) Hall, C. E.; Slayter, H. S. J. Biophys. Biochem. Cytol. 1959, 5, 11. (3) Marchant, R. E.; Barb, M. D.; Shainoff, J. R.; Eppell, S. J.; Wilson, D. L.; Siedlecki, C. A. Thromb. Haemostasis 1997, 77, 1048-1051. (4) Feng, L.; Andrade, J. D. In Proteins at Interfaces II: Fundamentals and Applications; Horbett, T. A., Brash, J. L., Eds.; American Chemical Society: Washington, DC, 1994; Vol. 602, pp 66-79. (5) Ta, T. C.; McDermott, M. T. Anal. Chem. 2000, 72, 2627-2634. (6) Ta, T. C.; Sykes, M. T.; McDermott, M. T. Langmuir 1998, 14, 24352443. (7) Cacciafesta, P.; Hallam, K. R.; Watkinson, A. C.; Allen, G. C.; Miles, M. J.; Jandt, K. D. Surf. Sci. 2001, 491, 405-420.
Figure 1. Schematic reproduced from ref 4.
surface loading8-10 often obscured the identification of single molecules, unless investigators deliberately adsorbed proteins from extremely dilute solutions and terminated the adsorption process well before the surface saturated. Only then were coverages sufficiently low to distinguish isolated fibrinogen molecules on surfaces such as mica,11-14 poly-L-lysine-coated mica,15 graphite,11,12 and titanium.16 Of these model substrates, the trinodular structure of fibrinogen was consistently seen on hydrophobic surfaces;11,12 however, its structure on mica has varied between groups: The Berrie lab reported it to have a predominately globular conformation11 on mica whereas Agnihotri and Siedlecki12 and Sit and Marchant14 both found its trinodular form to dominate. Fibrinogen images were also reported on less ideal, more application-relevant surfaces such as OTS-,14 APTES-,14 and hydrophobically modified-silicon wafers,17 and (8) Ortega-Vinuesa, J. L.; Tengvall, P.; Lundstrom, I. J. Colloid Interface Sci. 1998, 207, 228-239. (9) Ortega-Vinuesa, J. L.; Tengvall, P.; Lundstrom, I. Thin Solid Films 1998, 324, 257-273. (10) Choi, K. H.; Friedt, J. M.; Frederix, F.; Campitelli, A.; Borghs, G. Appl. Phys. Lett. 2002, 81, 1335-1337. (11) Marchin, K. L.; Berrie, C. L. Langmuir 2003, 19, 9883-9888. (12) Agnihotri, A.; Siedlecki, C. A. Langmuir 2004, 20, 8846-8852. (13) Agnihotri, A.; Siedlecki, C. A. Ultramicroscopy 2005, 102, 257-268. (14) Sit, R.; Marchant, R. E. Thromb. Haemostasis 1999, 82, 1053-1060. (15) Taatjes, D. J.; Quinn, A. S.; Jenny, R. J.; Hale, P.; Bovill, E. G.; McDonagh, J. Cell Biol. Int. 1997, 21, 715-726. (16) Cacciafesta, P.; Humphris, A. D. L.; Jandt, K. D.; Miles, M. J. Langmuir 2000, 16, 8167-8175.
10.1021/la051641g CCC: $33.50 © 2006 American Chemical Society Published on Web 02/09/2006
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even on PTFE fibers,18 though resolution and identification of the different domains was more difficult, if not impossible. Though AFM typically employs smooth surfaces so that molecules are easily identified, the use of phase imaging has made it possible to identify fibrinogen on rough materials such as PDMS and PE.19 Both ambient and fluid conditions have been used and consistent molecular proportions are reported.11-16 Of the AFM studies of proteins reported thus far, most have allowed protein adsorption to proceed in a quiescent solution,11,12,14-16,19 with poor transport characteristics due to disturbances during the initial substrate contact and removal of adsorbate solution. While problematic in studies targeting dynamics, this is not an issue in studies whose goal is simply to report the surface-induced conformations of isolated proteins. The recent availability of flow through in situ (wet) measurement chambers has provided a more systematic approach to protein adsorption, in response to interest in the conformations of proteins during the adsorption process itself. The commercially available flow cells are not, however, designed with controlled mass transport in mind and an effective mass-transfer coefficient for these cells has not yet been reported. The commercially available chambers also suffer from a lag time between initial protein introduction and the time at which viable images can be obtained. Even so, their use has provided important insight into protein behavior during adsorption, a relatively recent development. Among the first groups to report progressive images during deposition, the McDermott lab published micrographs where graphite surfaces become increasingly filled with time though features on individual molecules are not resolved, as a result of the large scan area.5,6 The ability to probe progressive protein adsorption has brought several key scientific questions to the forefront. For instance, in a very recent work, Gettens et al. adsorbed fibrinogen from a standing solution onto graphite and mica and employed AFM software to determine the percent area covered with increasing adsorption time.20 While at short times, individual fibrinogen molecules were distinguished; at longer times this level of detail was lost, with only occupied area reported. While this work is important from the kinetic perspective, without efforts to deconvolute the AFM tip size, the significance of the area reported by the AFM software is unclear, particularly its relation to molecular size. Indeed, other methods have been more effective at interpreting the protein footprint in terms of excluded surface area experienced by approaching proteins, with clear results for the area per molecule and its dependence on surface residence and surface chemistry.21-23 The appearance of the fullness or the emptiness of a surface, as viewed by AFM, is still an open issue, as is the ability of AFM to measure surface-induced protein denaturing. This latter process, likely to increase the area per molecule, was addressed effectively in the Siedlecki lab,12 focusing on protein height rather than area. While the AFM method still faces challenges to quantify occupied protein area, it can provide perspective into the issues of protein surface mobility and clustering, not directly accessible by other methods. Recently, based on a comparison of OWLS (17) Wigren, R.; Elwing, H.; Erlandsson, R.; Welin, S.; Lundstrom, I. FEBS Lett. 1991, 280, 225-228. (18) Rasmusson, J. R.; Erlandsson, R.; Salaneck, W. R.; Salaneck, M.; Schott, M.; Clark, D. T.; Lundstrom, I. Scanning Microsc. 1994, 8, 481-490. (19) Holland, N. B.; Marchant, R. E. J. Biomed. Mater. Res. 2000, 51, 307315. (20) Gettens, R. T. T.; Bai, Z.; Gilbert, J. L. J. Biomed. Mater. Res. 2005, 72A, 246-257. (21) Wertz, C. F.; Santore, M. M. Langmuir 1999, 15, 8884-8894. (22) Wertz, C. F.; Santore, M. M. Langmuir 2001, 17, 3006-3016. (23) Wertz, C. F.; Santore, M. M. Langmuir 2002, 18, 706-715.
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data for protein adsorption to a statistical mechanics model, Tie and Van Tassel argued in favor of aggregation of fibronectin and lysozyme on Si(Ti)O2 surfaces, during adsorption.24 The TiO2 substrate used by Cacciafesta et al. is similar to that in the Van Tassel study and Cacciafestia et al. do report clusters of fibrinogen molecules in their AFM images.16 While this finding agrees with Van Tassel’s interpretation of kinetics, clustering is not accepted as a ubiquitous feature of protein adsorption, with several groups observing single molecules over a relatively broad range of surface loadings.11-15 Related to clustering is the question of protein surface mobility which, while dependent on chemistry-specific protein-surface interactions, may be diffusion- or tip-induced. For instance, adsorbed fibrinogen is commonly found clustered on the edges of graphite planes:11,20 Most likely, initial adsorption occurs uniformly and the proteins subsequently migrate to the edges diffusively or under the AFM tip. Conversely, on hydrophilic surfaces such as mica, some groups report protein movement as a result of tip-surface interactions,12 though protein heights, relative to those of neighboring molecules on sequential scans, appear not to be influenced by the AFM tip. On hydrophilic silicon, evidence argues against restructuring of fibrinogen during drying.8 (This may be the case if “drying” leaves surface-bound water that facilitates protein relaxation processes “in air” that would have otherwise occurred under water.) The current work combines TIRF (total internal reflectance fluorescence)-measured kinetics of fibrinogen adsorption on silica with, to the extent possible, AFM measurements on the same surfaces. (The best AFM images are obtained on the native oxide of silicon wafers, a substrate not amenable to TIRF because it is opaque.) This work is the first to quantitatively confirm agreement between protein levels measured by the two techniques in progressive kinetic runs. More importantly, the work demonstrates how the appearance of the surface (per AFM images) relates to the kinetic adsorption regimes, showing how the kinetic surface capacity relates to the visual fullness of the surface on micrographs. The results also demonstrate the extent to which fibrinogen clusters on the surface and when fibrinogen-fibrinogen interactions on the surface become important. Results also argue against substantial protein mobility on these surfaces. Experimental Methods Materials. Bovine plasma fibrinogen, type IV, was purchased from Sigma and was 95% clottable. In experiments requiring fluorescent traces, fluorescein isothiocyanate (Sigma) was covalently attached to fibrinogen by reaction at room temperature in carbonate buffer (0.004 M Na2CO3 and 0.046 M NaHCO3) for several hours, according to established procedures.25 Free fluorescein and other potential contaminants (including protein aggregates) were removed from the protein solution using size exclusion chromatography with a BioGel P-6 polyacrylamide gel column (Biorad). The column eluent was phosphate buffer (0.008 M Na2HPO4 and 0.002 M KH2PO4), such that the purified, labeled product was at pH 7.4 rather than the pH 9 corresponding to the reaction solution. Buffer salts were purchased from Fisher Scientific. The extent of fluorescein labeling was measured with absorbance at 494 nm and, for different labeling batches, was between 0.7 and 1.3 per fibrinogen molecule. Microscope slides were purchased from Fisher Scientific, and silicon wafers and polished glass were obtained from International Wafer Service. All surfaces were cleaned for 30 min in a 70 vol % H2SO4/30 vol % H2O2 solution and thoroughly rinsed with our reverse osmosis (RO) purified water before using. (24) Tie, Y.; Calonder, C.; Tassel, P. R. V. J. Colloid Interface Sci. 2003, 268, 1-11. (25) Robeson, J. L.; Tilton, R. D. Biophys. J. 1995, 68, 2145-2155.
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Figure 2. Tapping mode AFM images (2 × 2 µm2 each) of bare surfaces: (A) silicon wafer, (B) polished glass, and (C) acid-etched microscope slide. Roughness traces are shown below each figure. Methods. Adsorption experiments were conducted using a TIRF (total internal reflectance fluorescence) setup built inside a Spex Fluorolog II Fluorescence Spectrometer. This differs from our laserbased instrument26 in that the SPEX employs a Xenon lamp with a single monochromator to provide the excitation light at a chosen wavelength (488 nm), uses a double monochromator to sort the emissions, and mounts the flow cell vertically instead of horizontally. Fluorescein emissions were measured at 519 nm. The slit shear flow cell, used in TIRF27 and reflectometry studies,28 but also employed here for controlled protein deposition, in some cases without optical monitoring, continually replenishes the bulk solution and maintains a constant bulk protein concentration. The cell was made of a Teflon block into which a 1.3 mm deep channel was machined, per a modification of Shibata’s design.29 A microscope slide (or precut silicon wafer or polished glass), comprising the substrate, was then clamped against the Teflon surface, and a peripheral O-ring was used to prevent leaks. The calibration to convert fluorescence to the adsorbed amount of protein identifies regions of transport-limited protein adsorption kinetics21 and employs known values for the free solution protein diffusion coefficients.30 Also, when using a fluorescence tracer, two issues must be addressed: the potential invasiveness of the fluorescent label and the proper interpretation of the fluorescent signal. A thorough discussion based on previous work has shown the fluorescein to be noninvasive in the particular cases of fibrinogen adsorbing to hydrophobic and hydrophilic surfaces at the concentrations currently employed,21-23 with the fluorescence signal proportional to the interfacial mass of tagged protein. Once the flow cell was assembled, phosphate buffer solution was passed through it for 5 min at a wall shear rate of 5 s-1. The phosphate buffer solution was switched over to a solution of phosphate buffered fibrinogen for a set amount of time (depending on the experiment) before being switched back to buffer for 5 min. When protein adsorption was monitored via TIRF, this procedure established the fluorescence baseline prior to protein adsorption, and the buffer flush clarified any contributions of free protein to the signal, as the
evanescent penetration depth of 100 nm is typically greater than the adsorbed layer thicknesses. AFM imaging was done in tapping mode on a Multimode Nanoscope IV system (Digital Instruments). The system was suspended via a bungee cord on a cement slab to minimize unwanted vibrations during imaging. Adsorption experiments using quiescent conditions were carried out in 20 mL vials, where the substrate was placed before adding a 1 ppm protein solution. After 10 min, the sample was removed under a stream of water and dried with nitrogen before imaging. Most of the surfaces studied by AFM in this work were prepared by protein deposition in the TIRF flow cells. In this case, specimens were removed from the flow cell under a continuous stream of RO water after protein deposition. Since samples can only be produced one at a time in the flow cell system, finished samples were placed in a custom-built glass slide holder submersed in MilliQ water until that batch of 6 samples was complete (up to 1 h). Following this, the samples were dried in an oven at 120 °C for 1 h prior to imaging. 2 × 2 cm2 samples were cut from the middle of each glass slide or silica surface so they would fit onto the sample stage of the AFM. Imaging was performed on at least 2 samples at each deposition condition and images were obtained at two scan areas: 2 × 2 µm2 scans as well as 500 × 500 nm2 for each sample. Imaging was carried out in tapping mode, with scan rates of 1-2 Hz. Imaging on silicon wafers was done with standard silicon cantilevers with a resonance frequency of 300 kHz and spring constant of 40 N/m (Digital Instruments), whereas imaging on microscope and polished glass surfaces was done with silicon cantilevers with a resonance frequency of 150 kHz and spring constant of 5 N/m (Digital Instruments). These different cantilevers were chosen to give the best images on their respective substrates. Image resolution was 1026 × 1026 samples/line and the target RMS amplitude was set at 0.5 V for all samples. Captured images were processed offline with a first-order flatten function and analyzed with section analysis. AFM images were produced using surfaces on which tagged or untagged fibrinogen was adsorbed, and no difference was observed.
Results (26) Kelly, M. S.; Santore, M. M. J. Appl. Polym. Sci. 1995, 58, 247-263. (27) Rebar, V. A.; Santore, M. M. Macromolecules 1996, 29, 6262-6272. (28) Fu, Z. G.; Santore, M. M. Colloids Surfaces, A 1998, 135, 63-75. (29) Shibata, C. T.; Lenhoff, A. M. J. Colloid Interface Sci. 1992, 148, 469484. (30) Wojciechowski, P. W.; Brash, J. L. Colloids Surf., B 1993, 1, 107.
Substrate Characterization. While the goal of this work was to compare AFM data to kinetic traces measured by TIRF, the two techniques have different requirements for the substrates. AFM requires relative surface smoothness while TIRF requires
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Table 1. XPS Data (at 15° Take-off Angle) and Oxide Thickness glass slide
polished glass
Si wafer
O 1s C 1s Si 2p Na 1s
68.5% 4.9% 25.9% 0.6%
72.2% 3.0% 24.3% 0.6%
61.0% 2.0% 37.9% 0.0%
oxide thickness
9 nm
2.3 nm
optical clarity, motivating our consideration of three types of silica surfaces. Figure 2 presents 2 × 2 µm2 AFM images of acid-etched silicon, polished glass, and glass microscope slides. There is little apparent difference between the silicon and polished glass, whose RMS roughnesses approach 0.19 nm, significantly less than the small 4.5 nm dimension of fibrinogen, suggesting that imaging fibrinogen via AFM on theses surfaces should not be difficult. (As a referee points out, RMS values of roughness do overlook isolated surface features which may approach the size of fibrinogen; however, as long as these features are relatively few in number relative to the adsorbed fibrinogen molecules, as is the case in our study, the fact that RMS roughness is an order of magnitude smaller than the smallest anticipated protein dimension is a favorable indicator of the ease of AFM imaging.) By contrast the microscope slide contains distinct snowy features, as tall as 8 nm and averaging about 35 nm in width, on top of a “base roughness” that is only slightly greater than that of the silicon wafer or polished glass. These tall features potentially mask the topography of adsorbed fibrinogen. Rather than being a true surface feature, the snowy tall spots on the microscope slides more likely result from an AFM tipsurface interaction, not present for the polished glass and silicon substrates, an argument continued later in this paper. The tall spots differ greatly in characteristic heights and widths from one slide to another, but are always discretely spaced, with relatively smoother areas in between. They never translate laterally on the surface. These tall features depend greatly on the effective tapping force and can be made to reversibly disappear.31 Though we employ AFM tips with cantilever resonance frequencies of 150 and 300 kHz for glass and silicon surfaces, respectively, similar AFM parameters are used on all surfaces, chosen to facilitate, ultimately, protein imaging. The Target Amplitude was set at 0.5 V for all experiments, resulting in an amplitude setpoint of around 0.27 V. Images of silicon or glass with and without proteins were taken at setpoint values of 0.17-0.23 V (or setpoint to free amplitude ratio, Rsp ) 0.63-0.85), producing a relatively weak tip force. On microscope slides, images containing instability signatures such as those in Figure 2C were produced using this weak tip force (setpoint value of around 0.25 V, or Rsp ) 0.93). In one case the weak tip force allowed a tip instability that resulted in the appearance of tall surface features (up to 15 nm height and 150 nm in width), whereas tuning the AFM to a stronger tapping interaction (0.05 V, or Rsp ) 0.185) with the same surface nearly eliminated the instability, with the height and size of the asperities reVersibly reduced down to 5 nm in height and 35 nm in width. Repeated scans over the same area with a strong tapping force did not move or remove these features from the surface, and subsequent scans of the same area with reduced force recovered the larger and taller surfaces features in their original locations. These instability features are discussed again, below, in the context of protein adsorption. Surfaces were also characterized using XPS and ellipsometry, in Table 1. Results were generally similar, with greater carbon (31) Toscano, A., Ph.D. Thesis, University of Massachusetts, expected 2006.
Figure 3. Fibrinogen adsorbed from quiescent 1 ppm solution for 10 min on polished glass. (A) 2 × 2 µm2 image; (B) close-up of linear conformation; (C) close-up of extended conformation; (D) close-up of bent conformation. On the profile plots below parts (B-D), the distance axis follows the contour indicated by the straight or bent line in the scan image.
contamination on the glass slides, and trace sodium on the glasses, not present on silicon oxide. The oxide layer on the glass side, characterized previously by reflectometry,28 is thicker than that on the silicon. In summary, the polished glass, silicon wafer, and acid-etched microscope slide surfaces present substantial chemical and topographical similarity, suggesting that meaningful results may be obtained by using these different types of silica surfaces in different experimental instruments in fibrinogen adsorption studies. The key chemical differences are the levels of carbon contamination and the unstable tip interactions with the microscope slide. Adsorbed Fibrinogen Morphology. Figure 3 presents AFM images of fibrinogen molecules adsorbed from a quiescent 1 ppm solution onto polished glass for 10 min followed by a gentle rinse in pH 7.4 buffer and air-drying. Below the 2 × 2 µm2 image are close-ups taken from the main figure that provide examples of different surface conformations. The trinodular structure of fibrinogen is generally apparent, and the figures confirm its expected 47 nm molecular length. The fibrinogen width exceeds the native protein dimension by about a factor of 2 (10 nm rather than 5 nm) and may be a result of protein denaturing during the initial adsorption or drying steps, in addition to any tip contributions. One interesting feature of the fibrinogen adsorbed in Figure 3 is the variety in overall protein conformation: Some molecules adopt a linear trinodular conformation in Figure 3B, while others are unfolded in Figure 3C with the AR and RC domains extended
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Figure 4. AFM data for fibrinogen adsorption from gentle (5 s-1) shearing flow and a bulk solution of 1 ppm on a silicon wafer at 30 s. (A) shows a 2 × 2 µm2 scan while (B-D) show close-ups of straight and bent conformations. Table 2. Relative Proportions of Adsorbed Fibrinogen Conformations percent of total molecules 2-domains 3-domains-line 4-domains 3-domains-bent
polished glass
silicon wafer
23% 44% 8% 25%
15% 54% 15% 15%
away from the center E domain, and still others, for instance in Figure 3D, are bent. (A referee points out that an alternate explanation for the extended conformation in Figure 3C is splitting of the D-domain into BR and R parts.) Table 2 summarizes the relative proportions in which we found these conformations for the polished glass and the experimental conditions in Figure 3. These straight and bent conformations have been previously observed16 and their proportions on Ti are not inconsistent with Table 2. On polished glass (as well as microscope slides and silicon wafers) we observe that the middle node containing the e-domain was shorter and smaller than the outer nodes that contain the d-domain, though the overall node heights appeared to be substrate-dependent, perhaps as a result of intrinsic surface roughness. These findings agree with the relative node heights measured in aqueous conditions on mica by Agnihotri and Siedlecki12 but differ substantially from ref 9 which reports more globular structure for fibrinogen on mica, where the three domains tended to be hidden. This raises issues as to the extent to which one can translate results between silica and mica, two frequently studied model hydrophilic surfaces. Figure 4 presents AFM micrographs of fibrinogen adsorbed on a substrate cut from a silicon wafer. Here, adsorption was conducted in a flow cell from 1 ppm buffered fibrinogen solution flowing with a wall shear rate of 5 s-1, sufficiently gentle to avoid any hydrodynamic alterations of the fibrinogen structure. Following a period of flowing buffer, the dilute fibrinogen solution
Figure 5. TIRF measurements of fibrinogen adsorption kinetics from 5 s-1 wall shear rate laminar flow onto acid-etched glass slides. (A) Uninterrupted runs of (pink) 25 ppm fibrinogen solution on microscope slide, (gray) 100 ppm fibrinogen solution on microscope slide, and (red) 100 ppm fibrinogen solution on polished glass are compared with a (blue) run employing 25 ppm fibrinogen solution which is interrupted by switching to buffer after 5 s of flow. (B) shows a high-resolution close-up of the same data.
was allowed to flow for only 30 s before switching the flow back to buffer, to limit the density of absorbing molecules. Individual trinodular fibrinogen molecules are evident in the main 2 × 2 µm2 image, and a sampling of the different fibrinogen conformations observed is also provided. On oxidized silicon, the length of straight trinodular conformations is 47 nm, as it was on polished glass, though the molecular width is 8 nm, slightly smaller than that on the polished glass, which may result from use of a different tip for the silicon surfaces. The heights of fibrinogen molecules on silicon wafers, such as those in Figure 4, are consistently lower than that seen on polished glass, such as that in Figure 3 (and on glass microscope slides, as will be shown later), about 2 nm on silica as opposed to about 3 nm on glass. This result is not expected since the fibrinogen dimensions are otherwise similar on glass and silicon, suggesting minimal differences in spreading of the molecule on the two substrates. The observation certainly suggests that on glass, fibrinogen does not settle into crevices and low regions of the substrate. Table 2 compares the fibrinogen conformations on glass to those on silica, demonstrating the similarities between adsorption on the two substrates. Given the similarities between our results on glass and silicon, and the difference between our results and the published literature on mica and graphite, we conclude that the similarities between glass and silicon are not coincidental. Rather, despite certain topographical and chemical differences in the bare silicon and glass surfaces, they are actually very nearly similar in their interactions with fibrinogen. Adsorption Kinetics in Flow. While AFM data, in Figures 3 and 4, argue for the similarity of fibrinogen adsorption onto
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Figure 6. Time progression of 2 × 2 µm2 AFM tapping mode scans for fibrinogen adsorption (from 25 ppm 5 s-1) on a silicon wafer: (A) 15 s (included is an example of an annotated arbitrary box for counting purposes); (B) 30 s; (C) 45 s; (D) 1 min; (E) 3 min; (F) 5 min; (G) 7 min; (H) 14 min; (I) 35 min.
polished glass and silicon at low surface loadings, this section examines adsorption dynamics on optically clear surfaces. Microscope slides are the substrate of choice for our TIRF experiments because they are inexpensive and our cell accommodates their shape. We have, however, conducted a limited number of TIRF runs with polished glass, which was hand-cut to fit our TIRF cells. Figure 5 presents TIRF data for fibrinogen adsorption onto microscope slides and polished glass, with a wall shear rate of 5 s-1. These data are consistent with previous fibrinogen adsorption studies in our lab,22 albeit on self-assembled monolayers of varied hydrophobicity, some of which were substantially hydrophilic, though not so much so as glass. As expected, the initial slopes of the kinetic traces for two different bulk solution concentrations are proportional to those concentrations, a signature of transport-limited adsorption kinetics. The data also demonstrate higher ultimate coverages (in the time frame of this study) for adsorption from higher solutions concentrations. Prior studies21-23 demonstrated that this behavior results from protein-denaturing kinetics that compete with the transport-limited arrival rate of proteins to the surface, producing different jammed final states depending on the ultimate extent of protein denaturing. Most important to the question of substrate chemistry is that fibrinogen adsorption kinetics are quantitatively identical on microscope slides and glass, despite the complexity of the AFM-imaged glass surface.
The key feature of the fibrinogen adsorption kinetics in Figure 5 is the sustained transport-limited kinetics up to rather large surface coverages of approximately 2.5 and 3 mg/m2 for bulk protein concentrations of 25 and 100 ppm, respectively. Transportlimited kinetics generally result from rapid protein adhesion (compared with diffusive time scales of interfacial protein arrival), and relatively irreversible adsorption. Indeed, the relative irreversibility, which is a requirement for transport-limited kinetics, is consistent with the robustness of the adsorbed fibrinogen samples in Figures 3 and 4. In those micrographs, randomly deposited molecules appeared not to be rearranged on the surface by the drying processes (which could involve an air-water contact line traveling over the surface, generating substantial local forces that drag molecules into aggregates that appear macroscopically like water spots or coffee rings) or by the AFM tip (as the same micrograph would be reproducibly scanned at least 2-3 times.) While the TIRF flow cell apparatus is useful for measuring adsorption kinetics, it also can be employed, as shown in Figure 5B, to controllably deposit very small amounts of protein on a surface, as was done in Figure 4. In Figure 5B, a 25 ppm fibrinogen solution was allowed to flow for only 5 s before the flow was switched back to buffer. The adsorbed molecules are retained on the surface after buffer reinjection, and in this case give a coverage of 0.025 mg/m2, an extremely low level which is easily discernible by our instrument. The short time kinetics in Figure 5B do not
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Figure 7. Time progression of 500 × 500 nm2 AFM tapping mode scans for fibrinogen adsorption on a silicon wafer: (A) 15 s; (B) 30 s; (C) 45 s; (D) 1 min. The left side of each image shows the raw tapping image which is annotated on the right side to indicate protein identification for later counting.
display the expected Leveque transport-limited slope because the protein solution exposure time (5 s) was shorter than that needed to establish a pseudo-steady-state concentration profile in the interfacial fluid.32 Interruption of adsorption traces (by flowing buffer) at times longer than that needed to establish the linear protein profile in the boundary layer will result in deposition of proteins in proportion to the protein solution flow time and the bulk solution concentration. We exploit this approach below to produce adsorbed fibrinogen samples for AFM study. AFM Studies Intermediate to Kinetic Runs. While substrate opacity confounds the TIRF measurement of fibrinogen adsorption kinetics on silicon wafer surfaces, insight into these adsorption kinetics and other dynamic interfacial processes during adsorption can be gained through AFM studies of surfaces generated by controlled protein deposition per Figure 5B. Indeed, the similarity of AFM images of fibrinogen on silicon (Figure 4) and polished glass (Figure 3) and the quantitative agreement between fibrinogen adsorption on microscope slide glass and polished glass (Figure 5) motivate an ultimate comparison of fibrinogen on silicon and microscope slides. Figure 6 presents a series of 2 × 2 µm2 AFM images that illustrate the progression of surface states during the adsorption (32) Lok, B. K.; Cheng, Y.-L.; Robertson, C. R. J. Colloid Interface Sci. 1983, 91, 104-116.
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of fibrinogen from 25 ppm buffered solution and a wall shear rate of 5 s-1. Here, 8 different silicon billets were employed in identical kinetic experiments, but the runs were arrested by the introduction of flowing buffer at different times: 15 s, 30 s, 45 s, 1 min, 3 min, 5 min, 7 min, 14 min, and 35 min. (Hence, the TIRF flow cell was used for deposition, but TIRF measurements were obviously not made.) Figure 7 shows additional 500 × 500 nm2 scans, imaged separately from the 2 × 2 µm2 scans, for the same protein deposition history. The second half of Figure 7 annotates the images to demonstrate how individual molecules are identified for subsequent analysis, in runs where individual molecules could be identified. In Figures 6 and 7, the density of surface features in both figures increases with protein deposition time, consistent with transport-limited adsorption. Individual molecules can be distinguished for protein adsorption times of a minute or less, though the surfaces appear slightly randomly aggregated at 45 s and 1 min. At shorter times the molecules generally appear not to touch each other. At 2 min and beyond, the amount of fibrinogen has increased to the point that individual molecules can no longer be distinguished. We will argue later that from 2 to 20 min the surface coverage increases substantially, and simply note here that during this time frame the appearance of the surface does continue to evolve, albeit in ways that are difficult to quantitate. We note here that the “surface aggregation” increases with surface loading and that at the lowest coverages are dominated by individual protein molecules. Thus, the mild surface aggregation is predominantly (but not exclusively) a percolation effect, since random deposition of molecules with an aspect ratio of about 10 will lead to “intersections” at low coverage levels. Figures 8 and 9 repeat the same experiment as that in Figures 6 and 7, but with a series of glass microscope slide substrates. (With the glass microscope slides, AFM micrographs correspond exactly to snapshots taken intermediate to the 25 ppm run in Figure 5, and the TIRF-measured coverage levels apply to the micrographs.) Figure 9 also shows, in addition to height and annotated height versions of surfaces with deposition times of a minute or less, phase contrast images. (Phase contrast has become an established means of identifying proteins on surfaces whose roughness exceeds the small dimensions of the protein.19) Generally, the images for the microscope slides are very similar to those obtained on silicon in Figures 6 and 7: The density of surface features generally increases with protein exposure times, with individual protein molecules being discernible for deposition times of a minute or less (corresponding to coverages of 0.26 mg/m2 or less, in Figure 5A.) Also, in Figures 8 and 9, molecules appear to touch each other in the 45 s and 1 min runs while molecules are isolated at 20 and 30 s (0.065 and 0.12 mg/m2). At times greater than 1 min, the surface appearance continues to evolve in ways that are difficult to quantitate, and the surfaces at 14 and 35 min are representative of saturated fibrinogen layers and indicate no substantial surface evolution. The most dramatic difference between fibrinogen adsorption on glass and silicon is the appearance of the surfaces at short times, corresponding to small amounts of adsorbed fibrinogen and reflecting differences in the underlying substrates in Figure 2. With AFM, tapping parameters were chosen to avoid disturbing the adsorbed proteins. Unstable tip interactions with the bare microscope slides are apparent at low protein coverages and complicate identification of individual protein molecules. It is fascinating, however, that the images corresponding to 45 s or more of protein adsorption (0.18 mg/m2 of fibrinogen) are nearly identical on silica and on microscope slides. That is, the asperities seen on the bare microscope slides and microscope slides with
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Figure 8. Time progression of 2 × 2 µm2 AFM tapping mode scans for fibrinogen adsorption (from 25 ppm 5 s-1) on acid-etched slides: (A) 20 s; (B) 30 s; (C) 45 s; (D) 1 min; (E) 3 min; (F) 5 min; (G) 7 min; (H) 14 min; (I) 35 min. On A and D, height profiles for representative fibrinogen molecules are shown.
very low protein coverages are absent at higher fibrinogen coverages. This suggests, as one possible explanation, that proteins adsorb onto these hot spots or asperities, or rapidly diffuse and adhere to them within seconds of first contacting the surface. Indeed, with the exception of what clearly are adsorbed fibrinogen molecules, the surfaces of Figure 6.D.1 and 9.D.1 appear relatively smooth. Further, the heights of the fibrinogen molecules adsorbed in Figures 8A-8D and 9A-9D are in the range of 3 nm (graph insert Figures 8A and 8D), consistent with their heights on the snow-free polished glass in Figure 3. The apparent disappearance of “high spots” on the glass microscope slides, caused by protein adsorption, is consistent with the previous speculation that the snowy features were not actually topographical. While it seems that, on glass, fibrinogen may be adsorbing onto small regions with higher than average surface energy, these “hot spots” are apparently not topographically elevated in the ballpark of 10 nm, as they appeared in Figure 2C. This conclusion follows from the “normal” fibrinogen height of 3 nm on the microscope slides, and even suggests an electrostatic contribution to adsorption: If the original asperities were electrostatic in nature, they would be eliminated by adsorption of oppositely charged regions of protein. Another possibility is that the asperities are somehow representative of the greater carbon contamination found by XPS on the microscope slides. Adsorption of fibrinogen onto hot spots on the glass also has kinetic implications, discussed below.
Discussion of Adsorption Kinetics: TIRF Vs AFM. While TIRF can measure kinetics over the full course of an adsorption run on microscope slides, these kinetics should also be apparent with AFM (with the protocols employed to produce Figures 6-9), as long as individual protein molecules are distinguishable in the micrographs. The right-hand sides of Figures 7 and 9 illustrate how we identified proteins on the surfaces at relatively low coverages. (Surface features that counted as proteins were based on the findings of Figures 3 and 4: Features that were trinodular, bent, or straight with the appropriate length and height. Some 2-node objects also counted as proteins, when they resembled the objects in Figures 3 and 4.) Counting was done on both 2 × 2 µm2 and 500 × 500 nm2 micrographs by choosing an arbitrary box and counting the proteins within it. An example of this is shown in Figure 6A, where a 650 × 650 nm2 box has been drawn. Numerous boxes of random sizes were drawn and counted for each picture. The number of proteins intersecting the box boundary were also counted but divided by two, being half in and half out. This procedure was adopted simply because of the difficulty in identifying proteins at the edge of the actual micrograph. In the case of low protein coverage (corresponding to less than 30 s of adsorption) on the microscope slides it was difficult to identify proteins, and here phase images were also employed to aid in our judgment. Figure 10 displays a comparison of the numbers of adsorbed proteins on microscope slide and silicon wafer substrates over
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Figure 9. Time progression of 500 × 500 nm2 AFM scans of fibrinogen adsorption on to acid-etched glass slides from 25 ppm solution and 5 s-1 wall shear rate flow. (A) 20 s; (B) 30 s; (C) 45 s; (D) 1 min. Each part of the figure contains (1) tapping-mode image on the left, (2) phase-contrast image in the middle, and (3) tapping mode image on the right annotated to indicate how proteins were identified for later counting.
Figure 10. Summary of early fibrinogen adsorption kinetics measured by TIRF on acid-etched microscope slide (solid line) and counting fibrinogen molecules on AFM images from samples where fibrinogen was adsorbed from a flow cell before it was removed and dried (blue [) on silicon wafer pieces and (pink 9) on acid-etched glass slides.
the time range where individual molecules could be distinguished. Imposed on the data is a straight line corresponding to the transport-limited behavior observed with TIRF for the 25 ppm run in Figure 5. There is excellent agreement among all the data. The agreement between TIRF and AFM measures of fibrinogen adsorption kinetics on the microscope slides is expected since the substrates for the various procedures were identical. The agreement further confirms that sample preparation for AFM does not remove protein from the surface or redistribute it from one region to another. The similar behavior on the silicon wafer
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demonstrates that fibrinogen adsorption onto this surface is also transport-limited, an observation which should not come as a surprise if one believes in the similarity of the underlying substrates in Table 1 and Figure 2. Comparison of TIRF and micrograph data also reveals some surprises, or at least force adjustments to one’s intuition. For instance, the micrographs of Figures 8E-8G, corresponding to adsorption times beyond 2 min of protein flow, show the appearance of a crowded surface in which it is difficult to see that substantial amounts of fibrinogen continue to adsorb, though it is clear that changes occur beyond 2 min. The TIRF data, however, indicate that even when the micrographs give the appearance of surface crowding (for instance, at 2 min), the surfaces are actually substantially open for further protein adsorption at the transport-limited rate! What appears to the eye as crowded is actually the opposite: approaching fibrinogen molecules in the interfacial fluid can diffuse to empty surface sites with a high probability of finding an open spot, so that the transport-limited adsorption kinetics are preserved. We emphasize here that this behavior should not be confused with multilayer adsorption. The ultimate coverages in the TIRF runs of Figure 5 are consistent with molecular footprints of 160 nm2/molecule,21 corresponding to the side-on conformation of fibrinogen and inconsistent with multilayer coverage. This conclusion, concerning the actual (large) capacity of surfaces that appear (to the AFM) as nearly full, provides a different perspective from the messages conveyed in studies such as that by Gettens et al.20 The micrographs for silicon and microscope slide substrates also suggest a small amount of surface mobility for fibrinogen in its early stages of adsorption. The extremely low coverage levels presented in Figure 2 show randomly deposited molecules, regardless of the adsorption conditions being flow or quiescent. Figure 8D and to a lesser extent Figure 6D, however, show what looks like slight fibrinogen aggregation, perhaps beyond expectations for the random deposition of elongated objects. Likewise, the low coverage micrographs of Figures 8 and 9 suggest selective adsorption onto hot spots. The mild degree of aggregation is still substantially less than that reported by Cacciafesta on TiO216 or by McDermott on graphite.6 Both of those prior works, on different substrates than ours, described larger aggregates whose heights exceeded the side-on dimensions of fibrinogen. Both the surface aggregation and adsorption onto the instabilityproducing regions of the microscope slide are processes requiring a small amount of surface diffusion: Within seconds or less of fibrinogen adsorbing to the microscope slide, it appears to be able to diffuse a small distance, less than 100 nm to a preferred adhesion site before becoming permanently attached. (That the fibrinogen molecules are indeed attached with relative permanence is upheld by protein displacement studies,33 and by our unpublished FRAPP studies with these systems.) Note that it is unlikely that the surface structures in Figures 6-9, C and D, were produced by adsorption directly onto hot spots or spots contiguous with previously adsorbed protein: The direct adsorption no-diffusion model would require that some adsorption attempts be rejected, leading to adsorption kinetics substantially lower than the observed transport limit. Figure 10 also confirms transport-limited adsorption kinetics of fibrinogen on silicon-wafer (silicon oxide) surfaces, for short exposure times when individual proteins can be identified. This result is not a surprise, given that both acid-etched microscope slides and silicon wafers present similar silica surfaces. We might further extrapolate our interpretation of the adsorption kinetics (33) Wertz, C. F.; Santore, M. M. Langmuir 2005, 21, 10172-10178.
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on silicon based on the similarities of the AFM micrographs at times longer than 2 min. Even though individual proteins are no longer discernible, the microscope slide and wafer images evolve in a similar fashion with continued protein exposure, and it would be fair to guess that, on silicon wafers, the transport-limited kinetics proceed to longer exposure times as they did for the microscope slides. We know, however, of no reports in the literature for the quantification of the adsorption kinetics of fibrinogen on silicon. Armstrong et al., however, describe rapid adsorption on fibrinogen on silicon wafers and pronounced linearity in the initial kinetics, consistent with the expected time scales of transport-limited behavior.34
Conclusions This work demonstrated quantitative similarities between fibrinogen adsorption on three silica surfaces: acid-etched glass microscope slides, acid-etched polished glass, and silicon wafers. At short times, when individual proteins could be distinguished by AFM, the relative populations of straight, extended, and bent conformations were substrate-independent. Transport-limited adsorption kinetics were observed on all surfaces, and it was noted that surfaces may appear full to the AFM at relatively short times when, in fact, they actually contain only about 10% of their full protein capacity (as deposited in the TIRF cells) and further adsorption can occur at the transport-limited rate to produce (34) Armstrong, J.; Salacinski, H. J.; Mu, Q.; Seifalian, A. M.; Peel, L.; Freeman, N.; Holt, C. M.; Lu, J. R. J. Phys. Condens. Matter 2004, 16, S2483-S2491.
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protein monolayers. We found no evidence of substantial longrange fibrinogen mobility on any of the surfaces. There was no evidence that proteins were perturbed by light AFM tapping or by the drying process, in which a three-phase contact line could potentially drag proteins over many micrometers but does not. (We note that we do not make claims concerning the areal footprint per molecule, which surely is affected by drying.) At coverages of 0.1-0.3 mg/m2 we find mild surface aggregation of the fibrinogen into a loose surface floccs requiring protein translation of less than 100 nm after adsorption. This mild aggregation is distinctly different from the dense aggregates reported by others on different surfaces. Fibrinogen adsorption on acid-etched microscope slides was remarkably similar to that on the smoother surfaces despite the “hot spots” or “snow” on the bare glass AFM images. While proteins appeared to have migrated to these surface features to, ultimately, obscure them, the appearance of microscope slides carrying small amounts of fibrinogen, and the relatively flat (
