Adsorption and Relaxation Kinetics of Albumin and Fibrinogen on

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Adsorption and Relaxation Kinetics of Albumin and Fibrinogen on Hydrophobic Surfaces: Single-Species and Competitive Behavior Christian F. Wertz and Maria M. Santore* Department of Chemical Engineering, Lehigh University, Bethlehem, Pennsylvania 18015 Received January 29, 1999. In Final Form: July 20, 1999 We report the kinetic behavior of albumin and fibrinogen adsorption and relaxation from gentle shearing flow and phosphate buffer onto C16 self-assembled monolayers. The adsorption kinetics were generally transport-limited; however, the ultimate coverages depended on the rates at which protein molecules arrived at the surface, suggesting that interfacial relaxations determined the ultimate coverage. Of particular note was a dependence of the ultimate coverage of both proteins on the wall shear rate, in addition to the influence of the bulk solution concentration. Analysis of single protein experiments revealed interfacial protein relaxation rates of 0.12 and 0.15 nm2 molecule-1 s-1 for albumin and fibrinogen, respectively. These rates were constant over the range of experimental conditions and represent the initial relaxation rates after protein adhesion to the surface. The initial protein footprints were consistent with the free solution protein dimensions and, in the case of albumin, grew over a factor of 5 as the protein relaxed. For fibrinogen, relaxations were less extensive, increasing the footprint by a factor of 3. The extents of relaxation and the sizes of the protein footprints during the linear regime of spreading suggest that interfacial denaturing contributes significantly to the relaxation process, in addition to simple reorientations. The albumin relaxation behavior was shown, in addition to its influence on albumin coverage, to affect the coverage of fibrinogen in competitive situations. When the C16 layer was passivated with albumin prior to fibrinogen adsorption, short albumin exposures (still sufficient to cover the C16 surface) were ineffective at preventing fibrinogen adsorption. Prolonged incubation of albumin layers in albumin solution or buffer dramatically reduced subsequent fibrinogen adhesion.

Introduction Protein adsorption at liquid-solid interfaces has been a focus of much past research and continues to be a focus of research because of its importance to the development of biocompatible materials, diagnostic kits, and protein separations. Where blood compatibility is concerned, one is typically interested in which proteins adsorb and the time scales that dominate their adsorption and surface activity. A veritable library of work is available on blood protein-surface interactions, with most of the studies employing a variety of polymer and silica substrates.1-15 Results from these studies were sometimes contradictory because of complications stemming from the particular assays for adsorbed proteins. For instance, methods that require removal of the surface from the protein solution and exposure to air can give nonclassical results.16 The (1) Arai, T.; Norde, W. Colloids Surfaces 1990, 51, 17. (2) Shiraham, H.; Lyklema, J.; Norde, W. J. Colloid Interface Sci. 1990, 139, 177. (3) Norde, W.; Lyklema, J. J. Colloid Interface Sci. 1978, 66, 257. (4) Norde, W.; Lyklema, J. J. Colloid Interface Sci. 1978, 66, 277. (5) Norde, W.; Lyklema, J. J. Colloid Interface Sci. 1978, 66, 295. (6) Norde, W.; Lyklema, J. J. Colloid Interface Sci. 1981, 82, 77. (7) Ortega-Vinuesa, J. L.; Hidalgo-Alvarez, R. Biotech. Bioeng. 1995, 47, 633. (8) Ishida, K. P.; Griffiths, P. R. J. Colloid Interface Sci. 1993, 160, 190. (9) Park, K.; Mosher, D. F.; Cooper, S. J. Biomed. Mater. Res. 1986, 20, 589. (10) Grasel, T. G.; Cooper, S. L. Biomaterials 1986, 7, 315. (11) Grasel, T. G.; Cooper, S. L. J. Biomed. Biomater. Res. 1989, 23, 311. (12) Takahara, A.; Okkema, A. Z.; Cooper, S. L.; Coury, A. J. Biomaterials 1991, 12, 324. (13) O’Connor, S. M.; Patuto, S. J.; Gehrke, S. H.; Retzinger, G. S. Polym. Repr. Div. Polym. Chem. ACS 1997, 38, 559. (14) Zembala, M.; Voegel, J. C.; Schaff, P. Langmuir, 1998, 14, 2167. (15) Lu, J. R.; Su, T. J.; Thomas, R. K. J. Phys. Chem. B 1998, 102, 10307.

most consistent findings are those from in-situ methods including fluorescence, ellipsometry or reflectivity (or surface plasmon), in-situ radiotracers, or acoustic waves. Most recently, the development of self-assembled monolayers has facilitated precise and reproducible control of surface chemistry, further increasing the consistency between results from different research groups.17-22 Some features of protein adsorption from blood parallel polymer adsorption and even the adsorption of small molecules: When multiple species adsorb from a multicomponent solution, the evolving surface composition is influenced by the diffusion of the various species from the solution to the interface. We have shown that, for adsorption of multiple polymer populations, the initial surface composition follows the transport-limited adsorption of the individual species up to the point when the surface appears full to at least one of the species (onset of lateral interactions).23-25 Beyond this time, some of the initially adsorbed molecules may be displaced as those ultimately preferred on the surface continue to adsorb. A similar phenomenon, the Vroman effect, was reported for (16) Brash, J. L.; Horbett, T. A. in Proteins at Interfaces II: Fundamentals and Applications; ACS Symposium 602; Brash, J. L., Horbett, T. A., Eds.; American Chemcial Society: Washington, DC, 1995; p 1. (17) Nygren, H.; Stenberg, M.; Karlsson, C. J. Biomed. Mater. Res. 1992, 26, 77. (18) Nygren, H.; Stenberg, M. J. Biomed. Mater. Res. 1988, 22, 1. (19) Prime, K.; Whitesides, G. M. Science 1991, 252, 1164. (20) Sheller, N. B.; Petrash, S.; Foster, M. D.; Tsukruk, V. V. Langmuir 1998, 14, 4535. (21) Petrash, S.; Liebmann-vinson, A.; Foster, M. D.; Lander, L. M.; Brittain, W. J.; Majkrzak, C. F. Biotechnol. Prog. 1997, 13, 635. (22) Liebmann-Vinson, A.; Lander, L. M.; Foster, M. D.; Brittain, W. J.; Vogler, E. A.; Majkrzak, C. F.; Satija, S. Langmuir 1996, 12, 2256. (23) Santore, M. M.; Fu, Z. Macromolecules 1997, 30, 8516. (24) Fu, Z.; Santore, M. M. Macromolecules 1998, 31, 7014. (25) Fu, Z.; Santore, M. M. Langmuir 1998, 14, 4300.

10.1021/la990089q CCC: $18.00 © 1999 American Chemical Society Published on Web 10/07/1999

Albumin and Fibrinogen on Hydrophobic Surfaces

the adsorption of fibrinogen from blood on hydrophilic surfaces: Fibrinogen that initially adsorbs is later displaced by more surface-active proteins such as kininogen that are present in solution at lower concentrations and reach the surface more slowly.26-32 This overshoot in the amount of adsorbed fibrinogen is most pronounced on hydrophilic surfaces and is reduced or eliminated on hydrophobic surfaces due to tighter protein adhesion on the latter.33,34 Besides kininogen, which is strongly surface active, there have been reports that abundant proteins such as albumin yield unusual interfacial competitive kinetics on hydrophobic surfaces.35 Other features of protein adsorption are very different from the adsorption of small molecules or even highmolecular-weight polymers. In addition to the apparent irreversibility of protein adsorption, with minimal exchange of molecules between the surface and the bulk solution (relative to the time scales for polymers), proteins dramatically change their configuration upon adsorption.1,36 Because with proteins this reconfiguration goes beyond the random coil to heterobrush transition observed for homopolymers, protein adsorption is complex: Denaturing on surfaces can reveal groups internal to the native protein which may provide surface sites for additional protein adsorption.30 There are several studies which report changes in the ordering (amount of R-helix or β-sheet) in adsorbed versus native proteins, and currently there is disagreement about whether the native form is recovered when the proteins are desorbed.36,37 Also, there are a few reports on the kinetics of protein unfolding. Of particular interest to us are two simple models: one that involves an intrinsic protein relaxation rate from its native configuration to some final form on the surface38 and a second which also includes the amount of available surface area in the spreading rate law.39 The competitive adsorption of albumin with other proteins such as fibrinogen is of general importance because it is prevalent in blood. Indeed, standard practice includes passivating surfaces with a layer of adsorbed albumin prior to contact with blood or blood proteins, to minimize surface-blood interactions. In this paper, we report the kinetics of albumin and fibrinogen on a model hydrophobic self-assembled monolayer (SAM) surface and explain the competitive adsorption kinetics in the context of the individual noncompetitive protein adsorption behavior. In our investigation of the individual protein (26) Cornelius, R. M.; Brash, J. L. J. Biomed. Mater. Res. 1997, 37, 314. (27) Wojciechowski, P. W.; Brash, J. L. Colloids Surf., B: Biointerfaces 1993, 1, 107. (28) Dejardin, P.; ten Hove, P.; Yu, X. J.; Brash, J. L. Langmuir 1995, 11 (1), 4001. (29) Brash, J. L.; ten Hove, P. J. Biomed. Mater. Res. 1989, 23, 157. (30) Le, M. T.; Dejardin, P. Langmuir 1998, 14, 3356. (31) Breemhaar, W.; Brinkman, E.; Ellens, D. J.; Beugeling, T.; Bantjes, A. Biomaterials 1984, 5, 269. (32) Dejardin, P.; Le, M. T. Langmuir 1995, 11, 4008. (33) Wojciechowski, P.; tenHove, P.; Brash, J. L. J. Colloid Interface Sci. 1986, 111, 455. (34) Slack, S. M.; Horbett, T. A. J. Colloid Interface Sci. 1989, 133, 148. (35) Baszkin, A.; Boissonnade, M. M. In Proteins at Interfaces II: Fundamentals and Applications; ACS Symposium 602; Brash, J. L., Horbett, T. A., Eds.; American Chemcial Society: Washington, DC, 1995; p 209. (36) Norde, W.; Haynes, C. A. In Proteins at Interfaces II: Fundamentals and Applications; ACS Symposium 602; Brash, J. L., Horbett, T. A., Eds.; American Chemcial Society: Washington, DC, 1995; p 26. (37) Kondo, A.; Oku, S.; Kigashitani, J. J. Colloid Interface Sci. 1991, 143, 214. (38) van Eijk, M. C. P.; Cohen Stuart, M. A. Langmuir 1997, 13, 5447. (39) Seigel, R. R.; Harder, P.; Dahint, R.; Grunze, M.; Josse, F.; Mrksich, M.; Whitesides, G. M. Anal, Chem. 1997, 69, 3321.

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adsorption, we propose that relaxations, rather than a fundamental isotherm shape, are critical in establishing the ultimate coverage and play a key role in the competition between the two proteins. In the current paper, we restrict our competitive adsorption studies to sequential behavior, to lay a groundwork for later reports of competitive coadsorption. Also, the current work focuses on short and intermediate time kinetics, rather than extremely long time behavior. Therefore, we use the term “irreversible” in the context of the time scale of our studies. Our study employs an in-situ technique, total internal reflectance fluorescence (TIRF), chosen for its ability for precise measurement of kinetics without interruptions that could change the interfacial chemistry. We also conduct our studies in gentle shearing flow to mediate the mass transport of proteins to the surface and maintain the bulk solution concentration at a fixed level. Other studies run in batch mode or employing quiescent conditions have the added complication that the bulk solution may become depleted of the species that one wishes to study. Also, with quiescent adsorption studies, the rate of protein arrival to the interface varies throughout an experiment. Use of shearing flow maintains a constant rate of interfacial protein arrival, facilitating quantitative interpretation of adsorption kinetics. Experimental Section Bovine serum albumin (BSA) was purchased from Sigma (catalog #A-0281). It was essentially fatty acid free (no more than 0.005%) and globulin free (no more than 0.1 µg of IgG per mg of BSA as determined by HPLC).40 Gel electrophoresis showed the BSA content exceeded 99%.40 In experiments requiring fluorescent tracers, fluorescein isothiocyanate was covalently attached by reaction at room temperature in a carbonate buffer overnight, according to established procedures.41 Free fluorescein and other potential contaminants 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, such that the labeled protein product was in phosphate buffer. We tested all stock solutions to be sure the dominant ions, corresponding to a pH just above 7, were those of the phosphate buffer, rather than the pH 9 carbonate buffer. The extent of fluorescein labeling was assessed with absorbance at 494 nm and, for the runs presented here, was between 0.7 and 1.1 labels per albumin molecule, for different labeling batches. Bovine plasma fibrinogen, type IV, was purchased from Sigma (catalog # F-4753) and was 95% clottable.40 Gel electrophoresis on a different lot of this fibrinogen product revealed no detectable impurities.42 The current study used the same fibrinogen product from several different lots and found consistent results. Fibrinogen labeling with fluorescein isothiocyanate followed the procedure for albumin,41 including purification in the Biogel P-6 column. For the runs presented here, the labeling densities were between 0.6 and 1.5 labels per fibrinogen molecule, for different labeling batches. The buffer solutions employed salts from Fisher Scientific. The phosphate buffer was comprised of 0.008 M Na2HPO4 and 0.002 M KH2PO4. The carbonate buffer was made of 0.004 M Na2CO3 and 0.046 M NaHCO3. A number of other investigators, also interested in protein adsorption and techniques similar to ours, purified their materials (after fluorescent labeling) on columns such as our P-6 gel column41,43-45 or used the proteins as provided by the supplier.1,2,9-12,15,17-22,27-29,33,38,39,42,46-49 Other groups (40) Protein characterization data provided by Sigma Chemical Co. (41) Robeson, J. L.; Tilton, R. D. Biophys. J. 1995, 68, 2145. (42) Malmsten, M.; Johansson, J.-A.; Burns, N. L.; Yasuda, H. K. Colloids Surf., B: Bioninterfaces 1996, 6, 191. (43) Lok, B. K.; Cheng, Y.; Robertson, C. R. J. Colloid Interface Sci. 1983, 91, 87. (44) Liu, J.; Hlady, B. Colloids Surf., B: Biointerfaces 1996, 8, 25. (45) Tremsina, Y. S.; Sevastianov, V. I.; Petrash, S.; Dando, W.; Foter, M. D. J. Biomater. Sci. Polym. Ed. 1998, 9, 151.

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filtered13,14,30,39 or dialyzed their proteins for at least 1 day.3-6,26,35 As a reviewer suggested to us, such proteins may not be sufficiently pure to justify the conclusions in a study such as ours. Therefore, we took great care to demonstrate quantitatively how contaminants would affect our data and its interpretation. For these control studies, we chose bovine plasma fibronectin (Sigma catalog #F4759, lyophilized from pH 7.5 0.05 M tris-buffered saline) as a deliberate contaminant which was added to the proteins of interest in control studies. Fibronectin homegeneity was evaluated by immunoelectrophoresis and was cell culture tested.40 The appendix summarizes the control study and demonstrates why our protein-handling procedures yield materials of adequate purity. C16 (hexadecyltrichlorosilane, from Huls) self-assembled monolayers (SAMs) served as a model hydrophobic surface. These were formed on microscope slides following a well-established procedure:50 The slides were first cleaned by soaking for 30 min in a 70 vol % H2SO4/30 vol % H2O2 solution. They were then thoroughly rinsed with deionized water and blown dry with nitrogen. After exposure in a Harrick plasma chamber for 20 s, they were placed in a desiccator under vacuum for 30 min, resting over a hexadecyltrichlorosilane-containing Petri dish, balanced on the dish edges. Water contact angles on these surfaces ranged from 109° to 111°, ensuring complete coverage of the SAM.51 TIRF (total internal reflectance fluorescence) was used to measure protein adsorption kinetics from gentle shearing flow of protein solutions. In TIRF, a light beam is totally internally reflected inside an optically clear adsorption substrate, setting up an evanescent wave in the nearby solution. This evanescent wave, with an exponential decay length near 100 nm, but depending on the light wavelength and its angle of incidence, excites fluorescent labels on molecules in solution near or on the surface. In adsorption studies, the fluorescence signal is often dominated by the adsorbed molecules which are at a much greater concentration and in the most intense part of the evanescent field relative to those free in solution near the surface. The fluorescence signal, measured in real time, therefore facilitates kinetic measurements of polymer or protein adsorption. Our TIRF setup, which has been described in detail previously,52,53 employs an Ar+ ion laser at 488 nm for excitation and a photon-counting detection system. The microscope slide substrates which comprise one wall of the adsorption flow cell are optically coupled to a waveguide prism using index-matching oil. TIRF can be used in single-component studies to track adsorption kinetics directly, or labeled populations can be monitored in more complex mixtures to probe competitive behavior.24 Adsorption experiments were conducted in a slit shear flow cell which continually replenishes the bulk solution and maintains a constant bulk protein concentration. The cell was made of a black Teflon block into which a 0.5 mm-deep channel was machined, per a modification of Shibata’s design.54 A microscope slide, comprising the substrate and optical cell window, was then clamped against the Teflon surface, and a peripheral O-ring was used to prevent leaks. Using TIRF to quantitatively determine the adsorbed amount of a protein or polymer requires careful calibration. Internal methods, which assume that the fluorescent labels on adsorbing molecules have the same quantum yield as those in free solution have been developed,55-57 and we have implemented these (46) Golander, C. G.; Kiss, E. J. Colloid Interface Sci. 1987, 121, 240. (47) Shibata, C. T.; Lenhoff, A. M. J. Colloid Interface Sci. 1992, 148, 469. (48) Voegel, J. C.; DeJardin, P.; Strasser, C.; deBaillou, N.; Schmitt, A. Colloids Surf. 1987, 25, 139. (49) Retzinger, G. S.; Cook, B. C.; Danglis, A. P. J. Colloid Interface Sci. 1994, 168, 514. (50) Chaudhury, M. K.; Whitesides, G. M. Langmuir 1991, 7, 1013. (51) Heid, S.; Effenberger, F. Langmuir 1996, 12, 2118. (52) Kelly, M. S.; Santore, M. M. Colloids Surf., A: Physiochem. Eng. Aspects 1995, 96, 199. (53) Rebar, V. A.; Santore, M. M. Macromolecules 1996, 29, 6263. (54) Shibata, C. T.; Lenhoff, A. M. J. Colloid Interface Sci. 1992, 148, 485. (55) Rondelez, F.; Ausserre, D.; Hervet, H. Annu. Rev. Phys. Chem. 1987, 38, 317. (56) Hlady, V.; Reinecke, D. R.; Andrade, J. D. J. Colloid Interface Sci. 1986, 111, 555.

Wertz and Santore approaches with moderate success.53 We have also used a nearBrewster internal reflection optical reflectometer, which operates on the same surfaces in the same flow cell as TIRF, to externally calibrate the adsorbed amount of poly(ethylene oxide) on silica,58 and had planned a similar strategy for the current protein study. It turns out, however, that the chemical treatment of the microscope slide prior to deposition of the SAM generates a layer on the glass surface beneath the SAM which, through interference, minimizes the sensitivity and linearity of the optical reflectometer. This has motivated us to develop an alternate method for the TIRF calibration. In our prior work with polymer adsorption, we had found conditions which give rise to transport-limited behavior and adsorption which proceeds at a constant rate for an extended period of time, as a substantial fraction of the surface becomes filled.58 In this case, knowledge of the free solution diffusion coefficient D facilitates calculation of the adsorption rate dΓ/dt, according to the Leveque solution to the convection-diffusion equation for transport-limited adsorption from gentle shearing flow:59

dΓ ) 0.538(γ/L)1/3D2/3C dt

(1)

Here L is the distance from the cell entrance to the point of observation, C is the bulk solution concentration, and γ is the wall shear rate, all of which are known. In our prior polymer adsorption studies, where the adsorbed amount was determined from fundamental reflectivity optics, the free solution diffusivity was back-calculated from eq 1 with great accuracy.58 In the protein adsorption studies presented here, we employed reported diffusivity values of 6 × 10-7 and 2 × 10-7 cm2/s for albumin and fibrinogen,27 respectively, to determine the adsorbed amount in protein runs that were transport-limited. Establishing those runs (or the portions of runs) that were indeed transportlimited was conducted with great care, as discussed below, and tested by examining the concentration and wall shear rate dependence of the initial kinetics. The conversion of fluorescence counts to mass of protein was then applied uniformly to all runs. Of particular note for the data presented here, careful control of instrument settings (e.g. filter positions and gains) gave us highly reproducible sets of data where the relative fluorescence for various runs represented the relative adsorbed masses. Then application of the calibration, using the diffusion coefficient and transport-limited kinetics, amounted to a simple multiplicative constant. The adsorbed amounts we measured, using this type of calibration, were consistent with the coverage levels reported in the literature for albumin and fibrinogen adsorption onto hydrophobic surfaces.18,35,59 When conducting fluorescence tracer experiments, one must address two issues: the potential invasiveness of the fluorescent label and the proper interpretation of the fluorescent signal. We addressed the potential invasiveness of fluorescein tags in our previous work on polymer adsorption. For adsorption onto negatively charged surfaces such as silica, the fluorescein labels are actually repelled from the surface because, near neutral bulk pH’s, they carry two negative charges.60 As a result, unlabeled polymers are selectively adsorbed over labeled ones,25 a feature which is very pronounced for mobile adsorbed layers (those that exhibit rapid and continued molecular exchange between the adsorbed layer and free solution, such as poly(ethylene oxide)). The influence of the fluorescein repulsion from the surface is dramatically reduced at the moderate ionic strengths in our current study.25 Further, for molecules such as hydroxyethylcellulose (HEC) that adsorb tightly and do not rapidly exchange with the bulk solution, the polymer backbone, rather than the fluorescein label, dominates the adsorption behavior.61 Because (57) Shibata, C. T.; Lenhoff, A. M. J. Colloid Interface Sci. 1992, 148, 469. (58) Fu, Z.; Santore, M. M. Colloids Surf., A: Physiochem. Eng. Aspects 1998, 135, 63. (59) Lok, B. K.; Cheng, Y.-L.; Robertson, C. R. J. Colloid Interface Sci. 1983, 91, 104. (60) Liu, X.; Kim, D. J.; and Santore, M. M. Phys. Rev. Lett., in press. (61) Mubarekyan, E.; Santore, M. M. Submitted to J. Colloid Interface Sci.

Albumin and Fibrinogen on Hydrophobic Surfaces the apparent irreversibility and nonexchangeability of albumin and fibrinogen, below, parallel those of HEC, fluorescein labels would also be expected to minimally alter protein adsorption. Two additional observations in the current work confirm the noninvasiveness of the fluorescein labeling: First, no fluorescein adsorption onto the C16 SAM was observed in control runs. Second, repeat runs with several albumin and fibrinogen samples with different labeling densities, up to and including at least 1.5 tags/molecule, gave identical kinetics. We are therefore confident that the results presented below are the true protein adsorption kinetics. Even though fluorescent labeling may not alter the physics that one attempts to study, care must be taken when interpreting the fluorescent signal. In studies such as ours, one intends that the fluorescence be directly proportional to the interfacial mass of the tagged species. This does not require that the fluorophores on adsorbed proteins possess the same quantum yield as those on proteins in free solution. Instead, the average quantum yield from labels in adsorbed layers must not change over the course of an experiment or from one experiment to another. We found that, for fluorescein tags on polymers adsorbing onto silica, this was often the case. Most important, the quantum yield within adsorbed layers was relatively constant during the initial stages of rapid transport-limited polymer adsorption; and for the case of PEO adsorbing onto silica, this requirement was also met for several hours following the initial adsorption.25 Therefore, fluorescein could be used as an effective tracer for PEO. Fluorescein’s quantum yield for HEC adsorbing onto silica also appeared constant during the adsorption of that polymer.61 In the Results and Discussion section below, the transport-limited adsorption regime for albumin and fibrinogen adsorption is equally well-defined, arguing against any drift in the fluorescein quantum yield in the initial stages of adsorption of these proteins. We are also aware, however, that fluorescein is a pH sensitive probe and its quantum yield is reduced at slightly acidic pH’s.60,62 This is especially important when studying adsorption onto negatively charged surfaces such as silica, because the nearby fluid is locally acidic. We have demonstrated the use of fluorescein as an interfacial pH sensor and have also demonstrated how its fluorescence can be used to measure the interfacial conformational changes of charged polymers in response to changes in ionic strength. When fluorescein-tagged molecules relax on silica, slow reductions in fluorescence are observed. We have found this to be the case for HEC adsorbed onto silica, where, after the initial adsorption, slow relaxations (at fixed interfacial mass) reduce the fluorescence at a rate of about 3%/h.61 Similar longtime fluorescence reductions were observed upon the adsorption of fluorescein-tagged albumin on silica63 and fluorescein-tagged lysozyme on silica.64 In the case of albumin, these fluorescent decays were eliminated by switching from a silica to a hydrophobic surface. The C-16 SAM surfaces in the current study apparently avoid the pH-quenching of the fluorescein during protein relaxation. In this study, which focuses on short- and intermediate-time protein adsorption kinetics, we interpret the fluorescence change primarily in the context of the evolution of the adsorbed mass of the labeled species. This interpretation appears especially robust at short times, up to half an hour, in cases where transportlimited behavior was observed. Neglecting the influence of interfacial relaxations on the quantum yield also seems reasonable, because, for several test cases in which adsorbed labeled protein was allowed to age on the surface for extended times, minimal fluorescence evolution was observed. A closer examination of the influence of protein relaxation on quantum yield will be part of a study of the long-time interfacial kinetic behavior.

Results and Discussion Interpreting TIRF Data. Figure 1 illustrates a typical trace for albumin adsorption on a C16 monolayer. Initially, phosphate buffer is flowing through the cell, and following a valve turnover, labeled protein solution flows through (62) Rebar, V. A.; Santore, M. M. J. Colloid Interface Sci. 1996, 178, 29. (63) Lorenz, D. Lehigh University research report. (64) Robeson, J. L.; Tilton, R. Langmuir 1996, 12, 6104.

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Figure 1. Typical albumin adsorption run for a bulk solution concentration of 10 mg/L and a wall shear rate of 5 s-1. After half an hour of flowing labeled albumin solution, unlabeled protein is injected. Curve “a” is for unlabeled albumin at a concentration of 100 mg/L, and curve “b” is for unlabeled fibrinogen at a concentration of 100 mg/L.

the cell, with adsorption commencing at time zero. After half an hour, the surface is nearly saturated and the adsorption is substantially slower than the initial rate. At this point, a more concentrated albumin solution (curve a) is injected into the cell and the signal no longer rises, since no additional fluorescent protein is being provided. A second run, with the same initial steps has been superposed on this graph. In the second run, after half an hour of labeled albumin exposure, unlabeled fibrinogen has been injected. In this case the signal decreases slightly. Figure 1 emphasizes several important points. First, there is minimal contribution of the labeled bulk solution to the fluorescent signal, as there is no abrupt signal drop when the labeled protein is removed from the flow channel. This was found to be the case for runs with up to 20 times more concentrated solutions than in Figure 1. Second, the signal is steady when unlabeled albumin is placed in the flow cell after adsorption of a fluorescein-tagged layer. The steadiness of the signal suggests that, for the time scales of interest to us, the albumin is almost irreversibly bound. The labeled protein is not displaced by the unlabeled protein. Also, the lack of any signal drift suggests that any albumin relaxations do not alter the fluorescence from the adsorbed layer. Finally, since there is minimal exchange between labeled and unlabeled albumin, one would also expect no desorption of albumin into flowing phosphate buffer, a process which is less energetically favored than exchange. Also, in Figure 1, there is a slight decay of the fluorescence when unlabeled fibrinogen challenges an adsorbed layer of labeled albumin. This likely results from a small displacement of the albumin by the fibrinogen. The main conclusions from Figure 1 are that there is no bulk solution contribution to the signal and no influence of relaxations on the fluorescence signal. This is also the case for fibrinogen adsorption experiments. Single-Protein Adsorption Kinetics. The adsorption of single proteins from buffer solutions onto the hydrophobic monolayers was investigated to determine the fundamental adsorption kinetics, the levels of coverage, and any apparent history dependence. Figure 2 shows the results for albumin, testing the observed adsorption behavior against transport-limited adsorption kinetics. In Figure 2A, the bulk solution concentration is varied (at a fixed wall shear rate of 5 s-1), while, in Figure 2B, the wall shear (flow) rate is varied (with a fixed bulk solution concentration of 10 mg/L). The inset in Figure 2A illustrates that, at low bulk solution concentrations, the initial adsorption rate dΓ/dt is proportional to the bulk solution concentration C, in agree-

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Figure 3. Apparent adsorption isotherm for albumin.

eq 1. This latter case will give rise to slower adsorption than described by eq 1; however, the transient convectiondiffusion-adsorption phenomena may still be kinetically limited by the diffusion step. The characteristic time τ for establishing the pseudo-steady-state profile is59

τ)

( ) L2 γ2D

1/3

(2)

If the time to saturate the surface at the transportlimited rate

τsat-TL )

Figure 2. (A) Albumin adsorption kinetics at a wall shear rate of 5 s-1 and varying concentrations of 200, 100, and 25 mg/L. In the inset, the bulk solution concentrations are 3, 6, and 12 mg/L and the x-axis is multiplied by concentration to superpose the runs. (B) Albumin adsorption kinetics at wall shear rates of 41, 8.5, and 1.1 s-1 and a bulk solution concentration of 10 mg/L. The same runs are superposed in the inset by multiplying the x-axis by γ1/3. (C) Initial adsorption rates for albumin scaled per the Leveque form.

ment with eq 1. Equation 1 also predicts that, for diffusionlimited adsorption, the initial adsorption rate will also be proportional to the wall shear rate to the 1/3 power. This is confirmed in the inset of Figure 2B, for the initial stages of adsorption, beyond which the different curves peel away at lower adsorption rates, from the transport-limited behavior. A few interesting features, concerning the comparison of the observed initial adsorption kinetics with eq 1, are worth mentioning. First, eq 1 applies when the surface is of sufficient capacity that it does not saturate before a pseudo-steady-state concentration gradient is achieved in the nearby fluid. While the concentration gradient near the surface is fixed, the constant adsorption rate, equal to the rate of interfacial protein arrival, is that given in eq 1. If the surface is of low capacity, or if conditions are such that the surface is filled too quickly, then the transient concentration gradient near the surface will never be as steep as the pseudo-steady-state profile corresponding to

Γsat 0.538(γ/L)1/3D2/3C

(3)

is less than τ from eq 2, then the diffusion-limited adsorption will not follow eq 1. Instead, the apparent dependence of dΓ/dt on concentration will be less than power unity, and the dependence on the wall shear rate will be less than power 1/3. Indeed, we found this to be the case in our initial adsorption kinetic studies with albumin near physiological concentrations. At concentrations of 100 mg/L or more, we were unable to collapse the initial adsorption kinetics by multiplying the time axis by C. Figure 2C summarizes the initial adsorption kinetics for different bulk solution concentrations and wall shear rates. Transport-limited adsorption follows eq 1 for values of γ1/3C less than about 75 mg/(L s1/3). At higher concentrations and/or wall shear rates, the initial adsorption kinetics are probably still diffusion limited; however, they do not follow eq 1 because the surface saturates too quickly. Our conclusion of initially transport-limited albumin adsorption onto C16 monolayers parallels Lok’s observations for albumin on poly(dimethylsiloxane).59 Lok also found a breakdown of the Leveque form as a result of rapid surface saturation in runs with high albumin concentrations. Thus far we have addressed only the initial adsorption kinetics. Turnover from the fast adsorption rate in eq 1 to slower kinetics signals the influence of adsorbed proteins on the adsorption kinetics of later-arriving molecules (since the bare hydrophobic surface has an intrinsically rapid adsorption rate.) For instance, for the runs in Figure 2A, a round shoulder in the kinetics sets in near 75% of the coverage. This is in contrast to our observations of polymer (poly(ethylene oxide)) adsorption on silica, where the adsorption rate from eq 1 applied up to full surface coverage.58 Also of interest in Figure 2A is that different ultimate coverages are achieved with different bulk solution concentrations. Normally one would interpret these ultimate coverages as a measure of the adsorption isotherm, in Figure 3. Indeed, similar adsorption isotherms (with comparable levels of coverage) have been reported


for albumin adsorption on heterogeneous hydrophobic surfaces such as poly(dimethylsiloxane).59 An alternate interpretation of the different ultimate coverage levels in Figure 2A is, however, that surface relaxations determine the ultimate albumin footprint on the surface. It is possible that when albumin initially adheres to a surface, it occupies a region whose size is similar to that of albumin free in solution. With time, however, the albumin molecule may increase its number of segment-surface contacts, and unfold or reorient on the surface, to occupy a larger area and exclude late-arriving molecules. If this were to occur, one would expect that experiments with slow rates of albumin arrival to an interface would exhibit lower coverages than experiments with fast rates of interfacial albumin arrival. The early arriving molecules would have time to relax and exclude late-arriving proteins. As the interfacial rate of albumin arrival is given by eq 1, one expects that runs with high bulk solution concentrations and wall shear rates would give higher ultimate coverages. The extent to which the albumin spreading is reversible can greatly complicate the situation. Figure 2A and B confirms that the ultimate albumin coverage (on moderate experimental time scales) is dependent on wall shear rate as well as concentration, suggesting that interfacial relaxations rather than the underlying isotherm shape are the reason for the variation in the ultimate coverage. Notably, the kinetic turnovers and dependence of the ultimate coverage on C and γ are a striking contrast to our prior work with PEO adsorption on silica,58 which provides a benchmark for fundamental interpretation of adsorption kinetics. This difference is a strong indicator that the fundamental interfacial behavior of proteins such as albumin is dominated by relaxations. Figure 4 illustrates the adsorption kinetics of fibrinogen, similar to the albumin study above. In Figure 4A, the concentration dependence of the adsorption kinetics (at a fixed wall shear rate of 5 s-1) is shown. The initial adsorption rate, in the inset, scales as the bulk solution concentration, per eq 1, signaling mass-transport-limited adsorption. There is, however, a significant dependence of the ultimate coverage on the bulk concentration, either as a result of the fundamental isotherm shape (shown in Figure 5) or due to interfacial (nearly irreversible) fibrinogen relaxations which affect the protein footprint. In Figure 4B, the wall shear rate is shown to affect both the initial adsorption rate and the long-time coverage. In the inset the initial adsorption rate is shown to scale as γ1/3, confirming that the initial adsorption kinetics are transport-limited. The dependence of the ultimate coverage on the wall shear rate again suggests that interfacial relaxations rather than the underlying shape of the adsorption isotherm determine the ultimate protein footprint and, hence, the ultimate coverage. Figure 4C summarizes the dependence of the initial adsorption kinetics on bulk concentration and wall shear rate. In the case of fibrinogen, the Leveque form in eq 1 applies to all the data, as a result of fibrinogen’s slower diffusivity and higher ultimate surface coverage. The transport-limited adsorption of fibrinogen on C16 monolayers parallels findings for fibrinogen on poly(dimethylsiloxane),59 as does the apparent adsorption isotherm in Figure 5. A few interesting features of albumin and fibrinogen adsorption onto C16 monolayers are worth comparing. First, the ultimate fibrinogen coverage exceeds that of albumin by a factor of ∼2 for similar concentrations and flow conditions. Our reported coverages are quite similar to those reported by Lok for albumin and fibrinogen on poly(dimethylsiloxane);59 however, Lok’s apparent isotherms had plateaus that were flatter than what we report

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Figure 4. (A) Fibrinogen adsorption kinetics at a wall shear rate of 5 s-1 and varying concentrations of 100, 50, and 25 mg/L. In the inset, the x-axis is multiplied by concentration to superpose the same runs. (B) Fibrinogen adsorption kinetics at wall shear rates of 22, 5.0, and 1.5 s-1 and a bulk solution concentration of 25 mg/L. The same runs are superposed in the inset by multiplying the x-axis by γ1/3. (C) Initial adsorption rates for fibrinogen scaled per the Leveque form.

Figure 5. Apparent adsorption isotherm for fibrinogen.

on the C16 surface. Also of interest is that the initial fibrinogen adsorption kinetics follow eq 1 within a particular run to a greater percent (∼90%) of the ultimate coverage than is the case for albumin. The rounder appearance of the albumin runs (e.g. Figure 2A as opposed to Figure 4A) may suggest that the relaxation and albumin

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arrival rates to the interface are more nearly competitive than is the case for fibrinogen or that the extent of albumin relaxation is greater than that of fibrinogen. Alternate explanations for the relatively more rounded albumin adsorption kinetics may include dimers in solution65 and immobile and laterally mobile interfacial albumin populations.66 This section established transport-limited adsorption for albumin and fibrinogen from single-protein solutions, concepts used as points of departure for interpreting competitive protein adsorption and interfacial relaxations in subsequent sections of this paper. Particularly important is the amount of tagged protein that adsorbs to the surface at the transport-limited rate and the apparent plateau levels of the kinetic runs. These two features of adsorption may change in competitive situations, or as a result of relaxations, or as a result of contaminant proteins present in the mixture. We therefore attempted to quantify the influence of potential contaminants on the rate and extent of transport-limited adsorption and the ultimate coverages obtained. The details of these studies are described in the Appendix. In brief, 10 wt % of the model contaminant, fibronectin, was added to test protein solutions of interest, and the adsorption behaviors were re-evaluated. The conclusions are that the initial transport-limited adsorption rates of the test proteins are unaltered by the presence of at least 10% contaminant. While the ultimate coverages of the test proteins are reduced by the presence of contaminants, the reduction in test protein coverage is roughly proportional to the amount of contaminant. Also, contaminant levels of 10% or less have no visible impact on relaxations and protein desorption. Since the materials we employed were pure to within 1% or better, we conclude that any slight contamination of our samples had a minimal influence on the observed adsorption rates and coverages. We therefore proceeded to interpret competitive and protein adsorption and relaxation behavior in the context of the adsorption behavior of the pure proteins, described in the next sections. Sequential Competitive Adsorption. Since the early albumin adsorption history influences its subsequent adsorption kinetics and the ultimate albumin coverage, one would expect that the albumin adsorption history would also influence its interfacial competition with other proteins. This issue was investigated in a series of runs in which a hydrophobic surface was first coated with adsorbing albumin and later challenged with fibrinogen. To determine the evolving surface composition, repeated runs were conducted using fluorescently tagged fibrinogen or tagged albumin. Figure 6 illustrates the adsorption of labeled fibrinogen for a 2 h period following the adsorption of unlabeled albumin for various amounts of time. All portions of all runs have a wall shear rate of 5 s-1. The unlabeled albumin concentration is 25 mg/L in all runs, and its adsorption commences at time zero. In Figure 6, however, the albumin adsorption is invisible because it is not fluorescently labeled. Therefore, for reference, the inset of Figure 6 illustrates the adsorption of labeled albumin for 4 h. The initial adsorption kinetics are consistent with Figure 1, and at long times, the coverage creeps upward very gradually. In the main part of Figure 6, several runs are superposed. In each, the albumin adsorption begins at time (65) Peters, T., Jr. All About Albumin; Academic Press: New York, 1996; p 26. (66) Tilton, R. D.; Robertson, C. R.; Gast, A. P. J. Colloid Interface Sci. 1990, 137, 192.

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Figure 6. Adsorption of labeled fibrinogen from a bulk solution concentration of 50 mg/L, following the adsorption of unlabeled albumin (invisible, starting at time zero) from a bulk solution concentration of 25 mg/L. The arrows indicate the addition of phosphate buffer solution. The wall shear rate is 5 s-1. In the inset, the albumin adsorption run is repeated with labeled albumin to illustrate the kinetics.

zero. Then, after a prescribed adsorption time, fibrinogen is passed through the flow cell at a concentration of 50 mg/L. The fibrinogen adsorption is shown on the y-axis and is greatly dependent on the prior albumin adsorption history. The beginning of each signal increase corresponds to a time on the x-axis, indicating prior albumin exposure/ adsorption. Experiments with only brief albumin adsorption periods, on the order of 30 min, exhibited transportlimited fibrinogen adsorption with an ultimate fibrinogen coverage of 2.6 mg/m2, about 75% of the coverage expected for fibrinogen adsorption onto a bare C16 SAM for a fibrinogen concentration of 50 mg /L and a flow rate of 5 s-1. Note that the brief 30 min adsorption period is sufficient to saturate the surface with albumin, yet this offered little protection from subsequent fibrinogen adsorption. For runs with longer albumin adsorption times prior to fibrinogen exposure, the fibrinogen adsorption was slower and less extensive. For albumin exposure times of 4 h, there was barely any subsequent fibrinogen adsorption at all over the following 2 h. In Figure 6, the fibrinogen exposure was kept to 2 h, as a result of our current focus on kinetics at short and intermediate times. During this period, in most cases, the fibrinogen coverage was still increasing slowly at the end of the run. Our long-time pilot studies indicated that some of these increases can be quite protracted, exceeding 12 h, and therefore extreme care must be taken to achieve quantitatively reproducible long-time results, a focus of our ongoing work. Also, with slow signal changes, one must take care to interpret results in the context of evolving interfacial mass or evolving quantum yield of the label as a result of slow protein unfolding, another focus of our ongoing work. For the results presented here, the signal changes are all generally consistent with evolving interfacial adsorbed amounts of the labeled protein. Ending the runs near 2 h of fibrinogen exposure did, however, reveal some additional information: In each run, flowing phosphate buffer was injected into the cell to replace the adsorbing fibrinogen, as indicated by the arrows. For the cases where there was substantial fibrinogen adsorption despite the albumin, the fibrinogen appears to be rather loosely bound and desorbs on a time scale similar to that of the slow fibrinogen adsorption. (In these runs, the bulk solution contribution to the fluorescence signal is negligible.) Since we have established that albumin adsorption itself is influenced by interfacial relaxations, it is worth asking whether the amount of adsorbed albumin or its extent of relaxation is more important in controlling subsequent


Figure 7. Adsorption of labeled fibrinogen following albumin adsorption of different histories. All albumin exposures are at 25 mg/L and a wall shear rate of 5 s-1. Curve “a” directly follows a 1/2 h albumin adsorption run. Curve “b” follows a 1/2 h albumin exposure and a subsequent 3 1/2 h wash in phosphate buffer. Curve “c” follows a 4 h albumin adsorption run.

Figure 8. Experiments complimentary to those in Figure 6, here with labeled albumin and unlabeled fibrinogen.

fibrinogen adsorption. This question is particularly relevant, since the inset in Figure 6 indicates only small increases in albumin coverage beyond the first 30 min of adsorption (from a 25 mg/L albumin solution at a wall shear rate of 5 s-1). To probe the importance of albumin relaxation, we compared two runs in which the albumin adsorption time was 30 min, long enough to cover most of the surface. In run “a” of Figure 7, already presented as the first of the 4 runs in Figure 6, fibrinogen was adsorbed immediately after the 30 min albumin exposure. In run “b”, after the 30 min albumin adsorption period, the albumin layer was exposed to flowing buffer for 3 1/2 h, so the total albumin layer age was about 4 h prior to fibrinogen exposure. There is a significant difference in the fibrinogen adsorption kinetics onto a fresh albumin layer as opposed to an aged albumin layer with the same initial albumin coverage. Aging the albumin significantly reduces the ability of fibrinogen to adhere to the surface. After 2 h of fibrinogen exposure, the fibrinogen coverage on the aged albumincontaining surface is less than half that on the surface containing the fresh albumin. Also, on the surface containing the aged albumin, the initial fibrinogen adsorption kinetics are a factor of 5 slower than the transport-limited rate. As a point of reference, in Figure 7, we present case “c”, in which fibrinogen adsorbs onto a surface that has been incubated in flowing albumin for 4 h. Clearly, this last case is the most effective at preventing fibrinogen adsorption: It may be that, at 30 min into the albumin adsorption run, there are still substantial bare surface patches available for a small amount of albumin or fibrinogen. Figure 8 presents the fate of the preadsorbed albumin for the runs shown in Figure 6. Figure 8 was generated

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from experiments complimentary to those in Figure 6, where labeled albumin was preadsorbed for various amounts of time from a 25 mg/L solution at a wall shear rate of 5 s-1. After different albumin adsorption periods, corresponding to those in Figure 6, unlabeled fibrinogen was injected at a concentration of 50 mg/L. Figure 8 reveals only modest albumin desorption, which is the least dramatic for the oldest albumin layers. Most of the albumin is retained on the surface, and the fibrinogen adsorption, already shown in Figure 6, occurs on sites not occupied by the albumin or on the surface of the adsorbed albumin that is facing the solution. Notably, the initial albumin adsorption kinetics are highly reproducible, so that the front ends of all the runs in Figure 8 superposed quite well. Presumably this is due to the highly reproducible hydrophobic nature of the C16 monolayers. Interfacial Protein Relaxation Kinetics. So far we have discussed the kinetics and extent of protein adsorption directly and expressed the importance of interfacial relaxations. The same relaxations that give rise to varied albumin and fibrinogen coverages in noncompetitive situations also play a role in competitive adsorption. The data in the previous sections can be interpreted in a semiquantitative fashion to reveal the extent and rate of interfacial relaxation in terms of spreading and/or reconfiguration. (By spreading, we mean the unfolding of adsorbed proteins and their continued flattening down onto the surface to yield an overall greater footprint per molecule. By reconfiguration, we mean processes in which the protein reorients or “rolls over” on the surface, without denaturing, to occupy a different amount of surface area per molecule.) In making this interpretation, we invoke the simple picture of an adsorbed protein as a blob occupying some area of surface which excludes other proteins from that portion of the surface.38 The initial protein footprint would be close to the size of the protein in solution, and there should be a maximum area that a single protein could occupy, after complete relaxation. The relaxation process might, however, be stunted by filling of the surface and lateral interactions between proteins. A complete treatment of protein relaxations must account for the fact that, during adsorption experiments such as ours, proteins continually bombard and adhere to the surface, such that the surface contains a mixture of proteins with different surface aging times and potentially a distribution of footprints. The surface coverage should, then, be the convolution of the individual protein relaxation function and the time-dependent protein flux toward the surface.38 Reversibility of the adsorption or relaxation processes will further complicate the model. In a simplified treatment, one could interpret the various single-protein adsorption kinetic runs in the context of an average surface residence time for each experiment. Protein relaxations occur during that period, to an ultimate footprint that determines the plateau coverage of a particular run. Runs that slowly saturate the surface (at low bulk solution concentrations or wall shear rates) would have a higher average protein residence time (a greater time where relaxations are allowed) and greater extents of relaxation (lower coverages) than runs where the surface is saturated rapidly (from high bulk solution concentrations and high wall shear rates). One could define this characteristic time per eq 3 or simply measure the time needed to reach some fraction (e.g. 75%) of the total coverage (τ-75). Using the second criteria (arbitrary coverage level) has the advantage of not being forced to assume transport-limited behavior. (Also note that a threshold coverage level somewhat lower than the maximum amount is used, so that the characteristic times do

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Figure 9. (A) Evolution of the adsorbed albumin protein footprint based on the runs (varied concentration and wall shear rate) in Figure 2. (B) Evolution of the adsorbed fibrinogen footprint based on the runs (varied concentration and wall shear rate) in Figure 4.

not depend on the tailing at the end of the run.) This analysis was applied to all the data in Figures 2 and 4 for albumin and fibrinogen, respectively. The results are summarized in Figure 9. The plots are nearly linear, indicating a constant rate of protein footprint growth, 0.12 and 0.15 nm2 molecule-1 s-1 for albumin and fibrinogen, respectively, for the range of experimental conditions in Figures 2 and 4. Notably, the overall extent of albumin relaxation is greater, with the footprint increasing just over a factor of 5 for the range of conditions studied, while, for fibrinogen, the overall increase is about a factor of 3. Figure 9 indicates that, for the experimental conditions we employ, the surface saturates and protein footprint growth is stunted by lateral interactions long before the proteins achieve their maximum footprints on the surface. Therefore, the protein relaxation rates we report are the initial footprint growth rates shortly after adsorption. Determination of the maximum footprints would require much lower bulk solution concentrations (rates and levels of interfacial chain arrival) than those in this study. Extrapolating the data in Figure 9 to time zero reveals the initial protein footprint prior to relaxation. In the case of albumin, this is about 25 nm2/molecule while for fibrinogen the initial value is about 75 nm2/molecule. These initial protein footprints are consistent with the reported protein sizes. Albumin is a small, soft globular protein with reported dimensions of 4.0 × 4.0 × 14 nm3.65,67 The end-on and side-on unperturbed dimensions are 16 nm2 (yielding a close-packed surface coverage of 7.2 mg/m2) and 56 nm2 (yielding a close-packed surface coverage of 2.1 mg/m2), respectively. From our experimentally determined zero-time albumin footprint of 25 nm2, one would conclude that the state of the interface, prior to protein relaxation or spreading, is one in which albumin molecules (67) Sigal, G. B.; Mrksich, M.; Whitesides, G. M. J. Am. Chem. Soc. 1998, 120, 3464.

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are adsorbed in a mixture of side-on and end-on orientations. For fibrinogen, the reported dimensions are 5.0 × 5.0 × 47.0 nm3,67 yielding an end-on molecular area of 16 nm2 (and a corresponding packed monolayer coverage of 22.6 mg/m2) and a side-on molecular area of 235 nm2 (corresponding to a close-packed monolayer coverage of 2.4 mg/m2). The extrapolated zero-time footprint of 75 nm2 for fibrinogen also suggests that, were the surface to fill without any spreading or protein reconformation, a mixture of side-on and end-on conformations would be found on the C16 surface, with significant amounts of fibrinogen adsorbed end-on. During the relaxation process, the albumin and fibrinogen footprints in Figure 9 grow at rates of 0.12 and 0.15 nm2 molecule-1 s-1, respectively, ultimately reaching molecular areas of 140 and 275 nm2, respectively, in 20 min. For albumin, the relaxations cause the molecular area to increase by a factor of 5, and the footprint of 140 nm2 does not correspond to a final protein size (because the relaxations in Figure 9A are linear, showing no signs of leveling off in the 20 min of relaxation times studied). The fact that the albumin surface area is 140 nm2 and growing suggests that the relaxation process involves more than a reconformation from end-on to side-on orientations. The occupied albumin area at a relaxation time of 20 min is about 3 times greater than that calculated for the sideon orientation, the maximum footprint calculated for native albumin. Clearly, substantial interfacial unfolding of albumin is responsible for the 0.12 nm2 molecule-1 s-1 footprint growth. Though some reorientation may be involved, this relaxation processes is dominated by a spreading or denaturing of the albumin on the surface. Similar arguments apply to the fibrinogen relaxations, where the molecular footprint at 20 min is 275 nm2, slightly larger than the footprint for side-on adsorbed fibrinogen. The fact that when the footprint is 275 nm2, the fibrinogen is still spreading at its initial rate (rather than leveling off) suggests that denaturing and spreading also contribute substantially to fibrinogen relaxation on C16 surfaces, though reconformations may also be important. The influence of albumin incubation time on the subsequent fibrinogen adsorption kinetics, in Figure 6, provides additional information about the albumin relaxation process. As albumin adsorbs and relaxes on the surface, the area available for fibrinogen decreases. Granted, the large rod-shaped fibrinogen molecule is a poor probe of open surface area; however, it is worth considering what the data in Figure 6 imply for albumin relaxation. First, we will only consider the area occupied by fibrinogen that has adsorbed rapidly at the transportlimited rate, since fibrinogen adsorption onto the bare C16 SAM is transport-limited. For fibrinogen adsorption onto a bare C16 SAM at the flow conditions of 5 s-1 and a fibrinogen concentration of 50 mg/L, the transportlimited regime of adsorption is complete at a coverage of 2.7 mg/m2, corresponding to a fibrinogen footprint of 210 nm2/molecule. This fibrinogen footprint could be considered the “probe size” for the runs in Figure 6. In Figure 6, deviation from transport-limited fibrinogen kinetics sets in at fibrinogen coverages of 1.6, 0.8, and 0.14 mg fibrinogen/m2 for prior albumin exposure times of 0.5, 1, and 2 h, respectively. One might imagine, then, that there are bare (or accessible) patches on the order of 1.6, 0.8, and 0.14 mg/m2 in the aging albumin layer (at the various times). With a nearly constant albumin coverage of 2.1 mg/m2 after the first 1/2 h of exposure (in the inset of Figure 6), one can calculate the amount of bare surface per albumin molecule, on the ordinate in Figure 10. As the albumin layer ages, the area available for fibrinogen


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Figure 10. Tracking the albumin relaxation according to the area remaining for fast (transport-limited) fibrinogen adsorption in Figure 6.

adsorption decreases, until, at 4 h, there is no surface to which the fibrinogen can quickly adsorb. The kinetics in Figure 10 show another perspective on albumin relaxation, though a direct quantitative comparison to the albumin spreading kinetics in Figure 9A is difficult. First, two different representations of albumin relaxation are indicated in Figures 9A and 10. (Figure 9A shows the growth of the area occupied by the albumin while Figure 10 shows the consumption of the free surface.) Figure 9A compares a series of albumin layers where the mass coverage varies significantly from datum to datum. Figure 10 essentially indicates the relaxation of a single albumin layer toward a final state of surface crowding. (Of course, a separate experiment and a separate layer were used for each datum in Figure 10, but all those layers had identical adsorption histories.) Then the comparison between Figure 9A and 10 is further complicated by the use of albumin as a self-probe in Figure 9A and fibrinogen as a probe in Figure 10. Even with these complications, however, the time evolutions in Figures 9A and 10 are consistent: In Figure 9A the longest time tested (corresponding to the slowest albumin run) was 1200 s. Up to this first 20 min, the albumin relaxation still persists at a constant rate, because the protein relaxation in runs such as those in Figure 2 is stunted by lateral interactions at 20 min. Figure 10 goes on to show that the relaxations and footprint growth continue for a period of several hours, with a characteristic time near 2 h. Up to the first hour in Figure 10, the albumin relaxation appears linear. The relaxations in Figure 10 are not well-described by a singleexponential.

Figure 11. Fibrinogen adsorption from a 50 mg/L solution in phosphate buffer, with a wall shear rate of 5.0 s-1. Curve “a” uses fibrinogen from Sigma, labeled with fluorescein and with the free fluorescein isothiocyanate removed on a column as described in the Experimental Section. Curve “b” uses the same adsorption conditions as curve “a”, but with 5 mg/L fibronectin added to a 50 mg/L fibrinogen solution. Protein adsorption starts at time zero, and the adsorbed layers are exposed to flowing buffered saline after 1 h.

cantly to the growth of the molecular footprint, in addition to simple reconformations or orientations. We have yet to conduct experiments with complete protein relaxation at low coverage levels where there is always bare surface and minimal lateral protein interactions. Because interfacial relaxations determine the ultimate coverage in single-protein adsorption experiments, they also determine the competitive adsorption between albumin and fibrinogen. Immediately after albumin adsorption on a surface, substantial amounts of fibrinogen are able to adsorb readily, even when the surface is nearly saturated with albumin. As the albumin layer is incubated in buffer or albumin solution for up to an additional 4 h, it becomes increasingly difficult for fibrinogen to adsorb. Saturated albumin layers incubated about 4 h almost completely exclude fibrinogen from the interface. The ability of fibrinogen to adsorb onto an albumin-coated surface provides a second measure of albumin relaxation kinetics, consistent with the albumin relaxation kinetics calculated from single-component albumin runs. Acknowledgment. This work was supported through a grant from The Whitaker Foundation.

Conclusions

Appendix

Careful measurements of albumin and fibrinogen adsorption from gently flowing phosphate buffer onto a model C16 SAM reveal transport-limited kinetics in the initial stages of adsorption, with the final coverage determined by the extent of relaxation of the proteins. Despite the fact that a variety of wall shear rates and concentrations were used, all the data for each protein collapsed onto a single kinetic curve for the interfacial protein relaxation: 0.12 and 0.15 nm2/s for albumin and fibrinogen, respectively. These values indicate the initial protein relaxation rates, and the spreading process is ultimately stunted by lateral interactions between proteins. The self-consistent form of the relaxation kinetics for the variety of experimental conditions also upholds the hypothesis that it is the relaxation kinetics rather than an intrinsic isotherm form that leads to the influence of bulk solution concentration on the ultimate coverage. (Faster adsorption is associated with high bulk solution concentrations.) For both proteins, especially albumin, interfacial spreading (denaturing) contributed signifi-

In response to questions concerning the influence of contaminants on the rigor of our data interpretation and the significance of our conclusions, we executed a series of test runs with a model contaminant protein, bovine fibronectin. This material was mixed with labeled test proteins to yield solutions containing 10% fibronectin contaminant (relative to the test proteins). Comparing the labeled test protein behavior to the case with deliberate contamination revealed the effects of contamination on the underlying interfacial behavior. Figure 11 shows an example pair of runs. Here, in run “a”, a 50 mg/L solution of labeled fibrinogen is adsorbed onto a C16 monolayer from phosphate buffer at a wall shear rate of 5.0 s-1. After 1 h, protein solution is flushed from the cell by flowing phosphate buffer. In curve “b” the run is repeated with a solution into which fibronectin has been added at a level of 10% of the test fibrinogen. Both the raw data and the fibrinogen coverages are shown on the y-axes to emphasize the fact that we did not have to rescale or adjust the data when comparing the two runs.

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The initial fibrinogen adsorption kinetics are identical for the two runs in the transport-limited regime (which accounts for 80% of the total coverage). Only when the surface approaches saturation are the affects of contamination apparent. The run with the fibronectin shows a lower tagged fibrinogen coverage because some portion of the surface is occupied by fibronectin and not available to the fibrinogen. There is no evidence for displacement of one protein by the other in the late stages of the runs because the slow portions of the adsorption curves run nearly parallel. The slow desorptions of loosely bound fibrinogen during buffer flow also run parallel for the two experiments, signaling no unusual influence of the fibronectin on desorption or fibrinogen relaxation. Test runs such as these, with huge (10%) amounts of contaminants, illustrated that the effect of competing

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proteins in the adsorption mixture has no influence on the transport-limited adsorption rate of the individual proteins. Further, the ultimate coverage levels of the test proteins in the presence of contaminants are reduced in predictable ways, taking into account the concentration of contaminant in the bulk solution. Considering that our original test mixtures have

Adsorption and Relaxation Kinetics of Albumin and Fibrinogen on

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