Planar Supported Lipid Bilayer Polymers Formed by Vesicle Fusion. 2

Planar supported lipid bilayers (PSLBs) composed of phosphorylcholine (PC) lipids are known ..... using the vesicle fusion method, whereas DAPC PSLBs ...
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Planar Supported Lipid Bilayer Polymers Formed by Vesicle Fusion. 2. Adsorption of Bovine Serum Albumin† Eric E. Ross, Tony Spratt, Sanchao Liu, Lynn J. Rozanski, David F. O’Brien, and S. Scott Saavedra* Department of Chemistry, University of Arizona, Tucson, Arizona 85721-0041 Received July 8, 2002. In Final Form: November 22, 2002 Planar supported lipid bilayers (PSLBs) composed of phosphorylcholine (PC) lipids are known to be highly resistant to nonspecific adsorption of soluble proteins. However, these structures lack the stability desired for implementation in molecular devices (e.g., biosensors). In part 1 of this report [Ross, E. E.; Spratt, T.; Liu, S.; Rozanski, L. J.; O’Brien, D. F.; Saavedra, S. S. Langmuir 2003, 19, 1752-1765], we described preparation and characterization of (poly)PSLBs composed of diene-functionalized lipids. Here we have examined nonspecific adsorption of bovine serum albumin (BSA) to these (poly)PSLBs and several reference surfaces, using total internal reflection fluorescence and atomic force microscopy. The results show that the protein resistance of a cross-linked (poly)PSLB composed of bis-sorbyl phosphatidylcholine (bis-SorbPC) is equivalent to that of a fluid PSLB composed of 1-palmitoyl-2-oleoylphosphatidylcholine, even after the former has been dried and rehydrated. Since the lipids in a (poly)bis-SorbPC PSLB are laterally immobile, these results also demonstrate that fluidity is not required for a PC lipid bilayer to resist nonspecific protein adsorption. Relative to (poly)bis-SorbPC, BSA adsorption to other types of (poly)PSLBs was significantly greater, including PSLBs formed from a diacetylene-functionalized lipid. The increased protein adsorption is attributed to a higher density of structural defects.

Introduction Toward the goal of creating “biocompatible” surface coatings that prolong the useful life of devices intended to operate in contact with biological fluids, several chemical functionalities have been identified that minimize the nonspecific adsorption of soluble proteins. Poly(ethylene glycol) (PEG) is one of the most extensively characterized of these functionalities.1 Recently, other functional groups that are comparably effective in reducing protein adsorption have been identified.2,3 For example, Holmlin et al.3a reported that mixed self-assembled monolayers (SAMs) composed of equimolar positively and negatively charged tailgroups, that is, a “zwitterionic” SAM, adsorbed less than 1% of a monolayer of fibrinogen and lysozyme from a 1 mg/mL solution, which is comparable to the protein resistance of oligo(ethylene glycol)-terminated SAMs. This finding is consistent with the well-known protein resistance of the zwitterionic phosphorylcholine (PC) moiety. The native protein resistance of the plasma membranes of erythrocytes and platelets has long been associated with the PC headgroup, which comprises approximately 90% of the lipid headgroups in the outer leaflet of these membranes.4 Due in part to their inherent protein resistance, lipid coatings composed of natural and/ * To whom correspondence should be addressed. Voice: (520) 621-9761. Fax: (520) 621-8407. E-mail: [email protected]. † Part of the Langmuir special issue entitled The Biomolecular Interface. (1) (a) Poly(ethylene glycol) Chemistry; Harris, J. M., Ed.; Plenum Press: New York, 1992. (b) Bailey, F. E., Jr.; Koleske, J. Y. Poly(Ethylene oxide); Academic Press: New York, 1976. (2) Luk, Y.-Y.; Kato, M.; Mrksich, M. Langmuir 2000, 16, 96049608. (3) (a) Holmlin, R. E.; Chen, X.; Chapman, R. G.; Takayama, S.; Whitesides, G. M. Langmuir 2001, 17, 2841-2850. (b) Ostuni, E.; Chapman, R. G.; Holmlin, R. E.; Takayama, S.; Whitesides, G. M. Langmuir 2001, 17, 5605-5620. (c) Ostuni, E.; Chapman, R. G.; Liang, M. N.; Meluleni, G.; Pier, G.; Ingber, D. E.; Whitesides, G. M. Langmuir 2001, 17, 6336-6343. (4) (a) Zwall, R. F. A.; Hemker, H. C. Haemostasis 1982, 11, 12. (b) Chapman, D. Langmuir 1993, 9, 39.

or synthetic PC lipids have recently been used in several biosensing applications.5 An additional advantage of this strategy is that a planar supported lipid bilayer (PSLB) can be readily derivatized with molecules that impart a desired functionality, for example, chromophores, ligands, and proteins, including transmembrane receptor proteins.5-7 The protein resistant properties of fluid PC lipid films have been described by several groups. For example, (i) Malmstem observed that multilayer PC lipid films adsorbed significantly less fibrinogen, IgG, and human serum albumin than methylated silica;8 (ii) Vogel and co-workers observed no adsorption of bovine serum albumin (BSA) (5) (a) Terrettaz, S.; Stora, T.; Duschl, C.; Vogel, H. Langmuir 1993, 9, 1361-1369. (b) Puu, G. Anal. Chem. 2001, 73, 72-79. (c) Stelzle, M.; Weissmu¨ller, G.; Sackmann, E. J. Phys. Chem. 1993, 97, 2974-2981. (d) Song, X. D.; Swanson, B. I. Anal. Chem. 1999, 71, 2097-2107. (e) Schmidt, C.; Mayer, M.; Vogel, H. Angew. Chem., Int. Ed. 2000, 39, 3137-3140. (f) Cornell, B. A.; Braach-Maksvytis, V.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wieczorek, L.; Pace, R. J. Nature 1997, 387, 580583. (g) Tien, H. T.; Ottova, A. L. Colloids Surf., A 1999, 149, 217-233. (6) (a) Sackmann, E. Science 1996, 271, 43. (b) Edmiston, P. L.; Saavedra, S. S. Biophys. J. 1998, 74, 999-1006. (c) Edmiston, P. L.; Saavedra, S. S. J. Am. Chem. Soc. 1998, 120, 1665-1671. (d) Fischer, B.; Heyn, S. P.; Egger, M.; Gaub, H. E. Langmuir 1993, 9, 136. (e) Duschl, C.; Liley, M.; Corradin, G.; Vogel, H. Biophys. J. 1994, 67, 1229. (7) (a) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 14773-14781. (b) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. Langmuir 1998, 14, 2467-2475. (c) Bieri, C.; Ernst, O. P.; Heyse, S.; Hofmann, K. P.; Vogel, H. Nat. Biotechnol. 1999, 17, 1105-1106. Naumann, R.; Jonczyk, A.; Hampel, C.; Ringsdorf, H.; Knoll, W.; Bunjes, N.; Gra¨ber, P. Bioelectrochem. Bioenerg. 1997, 42, 241-247. (d) Fisher, M. I.; Tja¨rnhage, T. Biosens. Bioelectron. 2000, 15, 463-471. (e) Puu, G.; Gustafson, I.; Artursson, E.; Ohlsson, P. Å. Biosens. Bioelectron. 1995, 10, 463-476. (f) Puu, G.; Aartursson, E.; Gustafson, I.; Lundsro¨m, M.; Jass, J. Biosens. Bioelectron. 2000, 15, 31-41. (g) Graff, A.; Winterhalter, M.; Meier, W. Langmuir 2001, 17, 919-923. (h) Naumann, R.; Schmidt, E. K.; Jonczyk, A.; Fendler, K.; Kadenback, B.; Liebermann, T.; Offenha¨usser, A.; Knoll, W. Biosens. Bioelectron. 1999, 14, 651662. (i) Schmidt, E. K.; Liebermann, T.; Kreiter, M.; Jonczyk, A.; Naumann, R.; Offenha¨usser, A.; Neumann, E.; Kukol, A.; Maelicke, A.; Knoll, W. Biosens. Bioelectron. 1998, 13, 585-591. (j) Se´vin-Landais, A.; Rigler, P.; Tzartos, S.; Hucho, F.; Hovius, R.; Vogel, H. Biophys. Chem. 2000, 85, 141-152. (8) (a) Malmsten, M. J. Colloid Interface Sci. 1995, 172, 106-115. (b) Malmsten, M. Colloids Surf., A 1999, 159, 77-87.

10.1021/la026198d CCC: $25.00 © 2003 American Chemical Society Published on Web 01/11/2003

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from a 1.75 µM solution to a SAM-supported 1-palmitoyl2-oleoylphosphatidylcholine (POPC) monolayer;5a (iii) Puu5b observed negligible adsorption of albumin to supported bilayers of mixed lipid compositions containing PC lipids, as measured by the resonant mirror technique. More recently, Glasma¨star et al.9 quantitatively evaluated the adsorption of several different proteins to fluid PC lipid bilayers, showing that in the majority of cases, irreversible adsorption was undetectable. However, in a fluid PSLB formed from naturally occurring lipids, the molecules are only weakly associated with each other and the underlying substrate surface.10 Consequently, these structures lack the robustness required for implementation in many technological applications (e.g., as a nonfouling coating in a washable, reusable biosensor). Improving the structural stability of supported lipid films while maintaining their inherent biocompatibility has therefore been a focus of research efforts for at least two decades.4b,11 One strategy, pursued since the early 1980s, is covalent polymerization of the lipid monomers. Despite the considerable scientific attention this topic has received, only a few publications have addressed the protein adsorption properties of polymerized, supported mono- and bilayer lipid films. Chapman and co-workers4b,11a were among the first to use polymerizable PC lipids to modify the surface properties of substrates. Langmuir-Blodgett mono- and multilayers of diacetylene-functionalized PC lipids were deposited on plastic surfaces and polymerized by UV irradiation. This procedure increased surface hydrophilicity and substantially decreased platelet adhesion and activation.4b More recently, Marra et al.11d fused and radically polymerized a mono-acroyloyl-functionalized PC lipid monolayer on an alkyl SAM film, producing a surface that significantly decreased platelet adhesion relative to bare glass. Both of these works indirectly demonstrate that polymerized PC lipids suppress protein adsorption since this process always precedes platelet adhesion and activation. However, in neither case was a comparison to fluid lipid PC layers performed, so the effect of stabilizing the film on its protein resistant properties is unknown. Several other strategies have been used to create PCterminated thin films on planar solids. For example, Nakabayashi and others incorporated PC moieties in hydrogel polymers, generally using monomers of PCmethacrylate polymerized with other vinyl components, to produce robust coatings.12 Further development of this strategy13 has demonstrated that thin films of a PCcontaining copolymer are effective in reducing protein adsorption relative to uncoated substrates. However, these films lack the degree of structural control available from Langmuir-Blodgett (LB) and self-assembly methods, meaning that the copolymer strategy does not allow presentation of a uniform array of PC groups to the (9) Glasma¨star, K.; Larsson, C.; Ho¨o¨k, F.; Kasemo, B. J. Colloid Interface Sci. 2002, 246, 40-47. (10) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 25542559. (11) (a) Hayward, J.; Chapman, D. Biomaterials 1984, 5, 135. (b) Albrecht, O.; Johnston, D. S.; Villaverde, C.; Chapman, D. Biochim. Biophys. Acta 1982, 687, 165-169. (c) Foltynowicz, Z.; Yamaguchi, K.; Czajka, B.; Regen, L. R. Macromolecules 1985, 18, 1394. (d) Marra, K. G.; Winger, T. M.; Hanson, S. R.; Chaikof, E. L. Macromolecules 1997, 30, 6483. (e) Marra, K. G.; Kidani, D. D. A.; Chaikof, E. L. Langmuir 1997, 13, 5697. (f) Orban, J. M.; Faucher, L. M.; Dluhy, R. A.; Chaikof, E. L. Macromolecules 2000, 33, 4205. (12) (a) Ishihara, K.; Fukumoto, K.; Iwasaki, Y.; Nakabayashi, N. Biomaterials 1999, 20, 1545-1551. (b) Ishihara, K.; Fukumoto, K.; Iwasaki, Y.; Nakabayashi, N. Biomaterials 1999, 20, 1553-1559. (13) (a) Murphy, E. F.; Lu, J. R.; Lewis, A. L.; Brewer, J.; Russell, J.; Stratford, P. Macromolecules 2000, 33, 4545. (b) For a review, see: Lewis, A. L. Colloids Surf., B 2000, 18, 261-275.

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overlying medium. Thus another strategy is to create a PC-terminated SAM3c,14 or a grafted monolayer.15 Whitesides and co-workers3a,c measured less protein adsorption on PC SAMs relative to saturated alkyl SAMs, but the degree of protein resistance was less than that of oligo(ethylene glycol)-terminated SAMs, the “standard” for protein resistance. This may be due in part to the packing arrangement in a PC SAM, which is not likely to assemble with the same surface area per PC moiety as that found in a fluid PC lipid bilayer. Consequently the dipole orientation of the PC group and associated water molecules,16 which may be an important aspect of its protein resistance,9,17 may be altered in a PC SAM relative to a PC lipid bilayer. In summary, if one seeks to create a thin film in which the packing geometry of the PC termini mimics that found in a natural PC lipid bilayer, both the copolymer and SAM strategies appear to be problematic. Furthermore, neither film type is a suitable architecture for reconstitution and stabilization of transmembrane protein receptors, which is of interest for development of biosensors and devices to screen pharmaceutical candidates [e.g., ref 7c]. For these reasons, we are working to develop methods to stabilize lamellar PC lipid structures that retain the characteristic protein resistance of fluid PC lipid bilayers. In part 1 of this report,18 we described the polymerization of PSLBs composed of diene-functionalized lipids. Different polymerization conditions produced films with different structural features, but all were found to increase PSLB stability relative to unpolymerized PSLBs. Here we investigate the protein adsorption behavior of these PSLBs. BSA adsorption behavior is compared among film types, including fluid PSLBs as well as reference surfaces that are not protein resistant, and is correlated to the density of structural defects in the films described in part 1. Experimental Section Materials. The polymerizable lipids bis-sorbyl phosphatidylcholine (bis-SorbPC), mono-sorbyl phosphatidylcholine (monoSorbPC), bis-dienoyl phosphatidylcholine (bis-DenPC), and dienoyl sorbyl phosphatidylcholine (DenSorbPC) were synthesized as described previously.19,20 1,2-Bis(10,12-tricosadiynoyl)sn-glycero-3-phosphocholine (DAPC), phosphatidyl-ethanolamine labeled with lissamine rhodamine B (Rh-DOPE), and POPC were purchased from Avanti Polar Lipids and used without further purification (see http://www.avantilipids.com for structures). Arachidic acid was purchased from Aldrich. BSA (A 3803), rhodamine isothiocyanate labeled BSA (RITC-BSA), and fibrinogen from human serum (F 3879) were purchased from Sigma and used as received. The RITC/BSA labeling ratio was between 1.1 and 1.4 (mol/mol). Proteins were dissolved in 50 mM phosphate (14) Tegoulia, V. A.; Rao, W.; Kalambar, A. T.; Rabolt, J. F.; Cooper, S. Langmuir 2001, 17, 4396-4404. (15) (a) Wang, Y.; Su, T. J.; Green, R.; Tang, Y.; Styrkas, D.; Danks, T. N.; Bolton, R.; Lu, J. R. Chem. Commun. 2000, 587. (b) Lu, J. R.; Murphy, E. F.; Su, T. J.; Lewis, A. L.; Stratford, P. W.; Satija, S. K. Langmuir 2001, 17, 3382-3389. (16) Hauser, H.; Pascher, I.; Pearson, R. H.; Sundell, S. Biochim. Biophys. Acta 1981, 650, 21-51. (17) (a) Harder, P.; Grunze, M.; Dahint, R.; Whitesides, G. M.; Laibinis, P. E. J. Phys. Chem. B 1998, 102, 426-436. (b) Feldman, K.; Hahner, G.; Spencer, N. D.; Harder, P.; Grunze, M. J. Am. Chem. Soc. 1999, 121, 10134-10141. (c) Wang, R. L. C.; Kreuzer, H. J.; Grunze, M. J. Phys. Chem. B 1997, 101, 9767-9773. (d) Zolk, M.; Eisert, F.; Pipper, J.; Herrwerth, S.; Eck, W.; Buck, M.; Grunze, M. Langmuir 2000, 16, 5849-5852. (18) Ross, E. E.; Spratt, T.; Liu, S.; Rozanski, L. J.; O’Brien, D. F.; Saavedra, S. S. Langmuir 2003, 19, 1752-1765. (19) (a) Lamparski, H.; Liman, U.; Frankel, D. A.; Barry, J. A.; Ramaswami, V.; Brown, M. F.; O’Brien, D. F. Biochemistry 1992, 31, 685-694. (b) Dorn, K.; Klingbiel, R. T.; Specht, D. P.; Tyminski, P. N.; Ringsdorf, H.; O’Brien, D. F. J. Am. Chem. Soc. 1984, 106, 1627-1633. (20) Liu, S.; Sisson, T. M.; O’Brien, D. F. Macromolecules 2001, 34, 465-473.

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buffer, pH 7.4, prepared using water obtained from a Barnstead Nanopure system. Planar Supported Lipid Bilayers. The methods that were used to prepare unpolymerized and polymerized PSLBs on fused silica and silicon wafer substrates were described in part 1.18 Briefly, all PSLBs except those composed of DAPC were deposited using the vesicle fusion method, whereas DAPC PSLBs were deposited using the Langmuir-Blodgett-Schaefer (LBS) method. PSLBs composed of reactive lipids were subjected to either UV photopolymerization or redox-initiated radical polymerization; they were kept under water until the polymerization reaction was complete. PSLBs composed of nonreactive lipids, or reactive lipids that were deliberately not polymerized, were always maintained under water or buffer. Arachidic acid (AA) monolayers were deposited on fused silica substrates using a standard LB method.21 Briefly, AA was vertically transferred on the first upstroke at a surface pressure of 55 mN/m from a subphase of 10 mM CaCl2, producing a “tailgroup-out” monolayer. The abbreviations used to refer to the various types of (poly)PSLBs examined in this work were stated in part 1 and are briefly reviewed here, using bis-SorbPC as an example: (i) a bis-SorbPC PSLB treated with potassium persulfate and sodium bisulfate to initiate radical polymerization is denoted as redox(poly)bis-SorbPC and (ii) a bis-SorbPC PSLB polymerized by direct UV irradiation is denoted as UV(poly)bis-SorbPC. Total Internal Reflectance Fluorescence (TIRF) Spectroscopy. RITC-BSA adsorption to unpolymerized and polymerized PSLBs composed of reactive lipids was measured using TIRF spectroscopy. Measurements were also made at three reference surfaces: a fluid POPC PSLB, an AA monolayer, and clean (bare) fused silica. Excitation was provided by the 514 nm line from an argon ion laser. The laser beam was coupled into a fused silica slide (Dynasil) using a right angle prism, propagated through the length of the slide via total internal reflection, and outcoupled via another right angle prism. The outcoupled beam was directed onto a reference diode detector to monitor fluctuations in source intensity. The slide was mounted in a small volume flow cell (approximately 50 µL),22 which was in turn mounted on a Nikon Diaphot inverted microscope. Fluorescence emission was back-collected through the fused silica slide using a 4× objective focused on a total reflection of the laser beam that occurred within the cell volume. The emission was passed through a 568 ( 20 nm band-pass filter and detected with a photomultiplier tube. The incident laser beam was mechanically chopped at 2.5 kHz, and phase sensitive detection was used to retrieve the fluorescence intensity. The experiment was interfaced to a PC using Lab Windows for data collection. All experiments were performed at ambient room temperature, ca. 25 °C. To perform TIRF measurements on PSLBs without exposing them to air, small unilamellar vesicles (SUVs) of the appropriate lipid composition were injected into the flow cell, resulting in vesicle fusion.18 After 15 min, the cell was rinsed with approximately 250 µL of buffer to remove unfused vesicles before proceeding to the protein adsorption experiment. In some cases, TIRF measurements were conducted on PSLBs that were polymerized in situ, followed by rinsing with deionized water and buffer; protein adsorption experiments were then performed without exposing the PSLB to air. TIRF measurements were also conducted on polymerized PSLBs that had been dried. The (poly)PSLB was formed on a silica slide, rinsed with deionized water, dried with a flowing nitrogen stream, mounted in the TIRF flow cell, and rehydrated. In all cases, before the initial RITC-BSA solution was injected into the flow cell, the background intensity was measured for subsequent subtraction. After the protein solution was injected and allowed to equilibrate for 30 min, the fluorescence intensity was measured. The cell was flushed with 500 µL of buffer, and the intensity was measured again. The procedure was then repeated using solutions of sequentially greater protein concentration. Making measurements before and after flushing the cell with buffer allowed the fluorescence emission from strongly (21) Edmiston, P. L.; Lee, J. E.; Cheng, S.-S.; Saavedra, S. S. J. Am. Chem. Soc. 1997, 119, 560-570. (22) Conboy, J. C.; McReynolds, K. D.; Gervay-Hague, J.; Saavedra, S. S. J. Am. Chem. Soc. 2002, 124, 968-977.

Ross et al. adsorbed proteins to be distinguished from that of dissolved proteins. The latter quantity, measured with the flow cell filled with a known concentration of dissolved RITC-BSA, provided the means to normalize the surface fluorescence data measured on different substrate surfaces, accounting for fluctuations in experimental variables such as prism coupling efficiency and optical collection efficiency.22,23 Sequential measurements were performed on several surfaces without observing a decline in signal intensity, indicating that under the conditions in which the measurements were performed, photobleaching was minimal. To further compare RITC-BSA adsorption on different surfaces, the fluorescence data (intensity versus protein concentration) were fit to one-site and two-site Langmuir adsorption models. The respective forms of the Langmuir isotherm were

F ) [FmaxcKa][(1 + Ka)c]-1

(1)

F ) [Fmax1cKa1][(1 + Ka1)c]-1 + [Fmax2cKa2][(1 + Ka2)c]-1 (2) where F is the measured fluorescence, c is the bulk concentration of BSA injected into the flow cell, Ka is the apparent binding constant, and Fmax is the maximum fluorescence at apparent saturation. The numeric subscripts on Ka and Fmax in eq 2 refer to the respective binding sites. Both Ka and Fmax were determined by a nonlinear least-squares analysis of the experimental data, using software Origin version 6.0 implemented on a PC. Isotherms measured on different substrate surfaces were normalized as described above. Atomic Force Microscopy (AFM). Atomic force microscopy was performed in tapping mode on a Digital Instruments Nanoscope III with silicon oxide tips (Digital) tuned between 300 and 400 kHz. Images were acquired on dried samples before and after incubation for 30 min in a 1 mg/mL protein solution, either BSA or fibrinogen, prepared in 50 mM phosphate buffer, pH 7.4. After incubation, the samples were briefly rinsed with deionized water and dried under a stream of N2. A minimum of three separate samples for each type of PSLB and reference surface were prepared and imaged; for each sample, images were acquired at several different locations on each substrate. The images and linescans presented below are typical and representative of the measurements on the respective samples. Roughness parameters were determined on a 1 µm2 scan area using Digital Instruments software. Bearing analysis, which provides a surface height histogram, was performed using Digital Instruments software to estimate adsorbed protein surface coverage on PSLB films.

Results and Discussion In part 1 of this report,18 we investigated the effects of the number and location of the diene moieties in polymerizable lipids and the polymerization method on the physical characteristics of polymerized PSLBs. Here we quantitatively examine the nonspecific adsorption of the serum protein BSA to these films relative to several reference surfaces, including fluid PSLBs, which is the architecture after which the polymerized films are modeled. TIRF Measurements of Adsorbed BSA Surface Coverage. TIRF was used to measure the degree of RITC-BSA adsorption to a variety of PSLB and reference surfaces. Surface coverages obtained after incubating a 1 mg/mL (15 µM) RITC-BSA solution with each type of adsorbent surface are listed in Table 1. These coverages were determined by calibrating and normalizing the respective TIRF responses to that measured for protein adsorption on a tailgroup-out AA monolayer, which was used as a hydrophobic reference surface.3 Protein adsorp(23) Hlady, V.; Reinecke, D. R.; Andrade, J. D. J. Colloid Interface Sci. 1986, 111, 555-569.

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Table 1. RITC-BSA Adsorption to Various PSLBs and Reference Surfaces from a 1 mg/mL Solution type of film or reference surfacea

rel adsorbed amountb,c

arachidic acid monolayer fused silica POPC bilayer, not dried redox(poly)bis-SorbPC, dried and rehydrated UV(poly)bis-SorbPC, dried and rehydrated UV(poly)bis-SorbPC, not dried redox(poly)DenSorbPC, dried and rehydrated DenSorbPC, unpolymerized, not dried UV(poly)DenSorbPC, not dried UV(poly)DAPC, dried and rehydrated UV(poly)DAPC, not dried

100 ( 14, n ) 4 60 ( 12, n ) 5 6 ( 7, n ) 4 6 ( 4, n ) 4 24 ( 4, n ) 2 7, n ) 1 27 ( 8, n ) 3 8 ( 3, n ) 2 23 ( 4, n ) 2 40 ( 11, n ) 3 20 ( 6, n ) 2

a UV and redox refer to the method of PSLB polymerization. Values expressed as percentages normalized to the amount of BSA that was adsorbed on an arachidic acid monolayer. c n refers to the number of independent measurements.

b

tion is highly favored at hydrophobic surfaces,24 and as expected, the AA monolayer was observed to adsorb the greatest amount of RITC-BSA. Under the assumption that a monolayer of RITC-BSA was adsorbed on the AA monolayer under the stated conditions (i.e., apparent saturation), the percentages listed in Table 1 represent percent monolayer coverages.25 Since protein adsorption on silica and glasses has been extensively studied,24 clean fused silica was also used as a reference surface. About 60% of a RITC-BSA monolayer was formed, which is in reasonably good agreement with published data. At a dissolved concentration of 1 mg/mL, the surface coverage of BSA adsorbed on bare glass and silica surfaces is reported to be ca. one monolayer [e.g., refs 13a and 24a]. Nonspecific protein adsorption on most surfaces is frequently greatest at a pH near its pI,24b,c,28 which is approximately 4.8 for BSA. At neutral pH (at which our measurements were performed), BSA has a net charge of about -18.29 Assuming monolayer coverage occurs at ca. pH 4.8, then due to electrostatic repulsion, submonolayer coverage is more likely at neutral pH. Thus our results are reasonably consistent with published data. The third reference surface was a fluid PSLB composed of POPC, at which the surface coverage of nonspecifically adsorbed RITC-BSA was 6% of that on the AA monolayer. Other groups have reported that nonspecific adsorption of albumins and other proteins (e.g., fibrinogen) to fluid PC lipid layers is negligible and sometimes undetectable, as measured using quartz crystal microbalance, surface (24) Some reviews on protein adsorption are: (a) Brash, J. L.; Horbett, T. A. In Proteins at Interfaces II; Horbett, T. A., Brash, J. L., Eds.; ACS Symposium Series 602; American Chemical Society: Washington, DC, 1995; pp 1-23. (b) Norde, W. Adv. Colloid Interface Sci. 1986, 25, 267340. (c) Haynes, C. A.; Norde, W. Colloids Surf., B 1994, 2, 517-566. (d) Hlady, V.; Buijs, J. Curr. Opin. Biotechnol. 1996, 7, 72-77. (25) As discussed previously by a number of authors (refs 22, 23, 26, and 27), we note that using TIRF to determine the surface coverage of an adsorbed protein film can be problematic. There are several contributing factors. Of most concern here is self-quenching of RITC fluorescence due to energy transfer, which may occur when RITC-labeled protein molecules are closely packed in an adsorbed film. This will cause the quantum yield to decrease as the surface coverage increases. Considering the fractional surface coverages presented in Table 1, the net effect of self-quenching is that the values listed for the most proteinresistant surfaces (e.g., POPC) are probably overestimated. Cognizant of these limitations, the data can still be quantitatively evaluated, allowing the protein resistance of (poly)PSLBs to be compared to the reference surfaces. (26) Rebar, V. A.; Santore, M. M. Macromolecules 1996, 29, 62626272. (27) Shibata, C. T.; Lenhoff, A. M. J. Colloid Interface Sci. 1992, 148, 469. (28) Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F. J. Phys. Chem. B 1999, 103, 3727-3736. (29) Peters, T., Jr. Adv. Protein Chem. 1985, 37, 161-245.

plasmon resonance, and resonance mirror techniques.5a,b,9 The discrepancy with our results is likely due to the greater sensitivity of TIRF, which can easily detect less than 0.1% monolayer coverage of a protein labeled with a strongly fluorescent dye.30,31 The results obtained with the reference surfaces were used to assess the protein resistance of polymerized PSLBs, specifically, the degree to which a (poly)PSLB mimics a fluid PSLB after the former is dried and rehydated. BSA adsorption from a 1 mg/mL solution to a redox(poly)bis-SorbPC PSLB that was dried and rehydrated produced a relative surface coverage of 6%, equivalent to that measured on a fluid POPC bilayer (Table 1). In part 1 of this report,18 redox-initiated polymerization of bis-SorbPC PSLBs was found to best preserve the bilayer structure after removal of the film from water. Structural characterization by ellipsometry and AFM revealed a highly uniform, nearly defect-free film of approximate bilayer thickness. The observation of protein resistance equivalent to that of a POPC bilayer provides additional evidence that the structure of a bis-SorbPC PSLB is maintained throughout redox polymerization, rinsing and drying, and subsequent rehydration. Cross-linking polymerization of the monomers in a bisSorbPC PSLB eliminates lateral lipid diffusion. This was qualitatively observed by epifluorescence microscopy; scratches formed with a syringe needle in a hydrated redox(poly)bis-SorbPC PSLB doped with 1 mol % RhDOPE did not “heal” (data not shown). In contrast, scratches formed in a hydrated, fluid POPC PSLB doped with 1% Rh-DOPE were observed to disappear in several minutes, which is qualitatively consistent with measurements of lateral lipid diffusion in fluid PSLBs.10,32 In a recent paper,9 it was suggested that lipid lateral mobility may be a significant aspect of the protein resistance characteristic of a fluid PC lipid bilayer. However our results indicate otherwise; that is, immobilization of the lipid monomers does not affect the protein resistant properties of a PC PSLB. Neither extensive buffer rinses nor rinses with high ionic strength buffer (500 mM NaCl) caused measurable desorption of the 6% monolayer coverage of RITC-BSA that adsorbed on redox(poly)bis-SorbPC BSA. However, approximately two-thirds (4% coverage) could be removed by rinsing the film with the surfactant Triton X-100 (2% v/v). These observations suggest that the small degree of nonspecific protein adsorption observed on redox(poly)bis-SorbPC is hydrophobically driven. Re-exposing the surfactant-washed PSLB to 1 mg/mL RITC-BSA reproduced the 6% monolayer coverage initially measured, demonstrating the stability of the PSLB to surfactant exposure. Equivalent experiments could not be conducted with fluid POPC PSLBs, since exposure to Triton X-100 causes desorption of nonpolymerized lipids from a solid support. As described in part 1,18 polymerization of a bis-SorbPC PSLB by direct UV irradiation produced a dried film that was only 28 Å thick and more hydrophobic than redox(poly)bis-SorbPC (sessile water contact angles of ca. 59° and 32°, respectively). AFM imaging of dried UV(poly)bis-SorbPC films revealed a significant density of defects which was attributed to the production of smaller cross(30) Nie, S.; Zare, R. N. Annu. Rev. Biophys. Biomol. Struct. 1997, 26, 567-596. (31) (a) Kang, S. H.; Shortreed, M. R.; Yeung, E. S. Anal. Chem. 2001, 73, 1091-1099. (b) Funatsu, T.; Harada, Y.; Tokunaga, M.; Saito, K.; Yanagida, T. Nature 1995, 374, 555-559. (32) Cremer, P. S.; Groves, J. T.; Kung, L. A.; Boxer, S. G. Langmuir 1999, 15, 3893-3896.

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linked domains that are more easily desorbed upon removal of the film from water, relative to the polymer units generated by redox-initiated polymerization. The higher defect density in UV(poly)bis-SorbPC PSLBs was accompanied by an increase in protein adsorption. The relative RITC-BSA surface coverage was 24%, 4-fold greater than on POPC and redox(poly)bis-SorbPC (Table 1). The increase is attributed to hydrophobically driven adsorption at the defects where the interior of the bilayer is exposed to the overlying solution. However, we note that a 24% monolayer coverage still represents a significant improvement in protein resistance in comparison to the underlying bare silica substrate and the highly adsorbing AA monolayer. RITC-BSA adsorption to UV(poly)bis-SorbPC was also measured on a film that was never dried. In this case, the PSLB was formed in the TIRF flow cell by fusion, rinsed, UV polymerized, and rinsed again before proceeding to protein adsorption measurements. The relative BSA surface coverage was only 7%, equivalent to that measured on POPC and redox(poly)bis-SorbPC bilayers (Table 1). This result shows that the greater protein adsorption measured on dried UV(poly)bis-SorbPC occurs as a result of removing the PSLB from water; in other words, the increase cannot be a consequence of the UV photopolymerization reaction. These results therefore provide additional support for our contention18 that the structural defects present in a dried UV(poly)bis-SorbPC PSLB are created by desorption of lipid material when the film is dried. In part 1,18 it was found that redox-initiated polymerization of DenSorbPC produced dried PSLBs that were structurally equivalent to redox(poly)bis-SorbPC PSLBs, as evaluated by ellipsometry, AFM, and stability to rinsing with Triton X-100. However, a significant difference in protein adsorption behavior was observed. The relative RITC-BSA surface coverage on redox(poly)DenSorbPC PSLBs was 27%, about 4.5 times greater than on POPC and redox(poly)bis-SorbPC and comparable to that measured for dried UV(poly)bis-SorbPC films. The greater degree of protein adsorption was determined to be a consequence of the polymerization reaction. The relative RITC-BSA coverage measured on an unpolymerized DenSorb PSLB that was never exposed to air was only 8%, comparable to that of POPC (Table 1). After in situ UV polymerization, without exposing the PSLB to air, the relative RITC-BSA surface coverage increased about 3-fold to 23%. Although polymerization of a DenSorb PSLB clearly lessens its protein resistance, the structural origin of this change is not clear at this time. A possible explanation is that creation of a cross-linked network adjacent to the glycerol backbone alters the orientation of the PC headgroup and the structure of its solvation shell. Although we are not aware of any published studies that relate protein adsorption behavior to PC dipole orientation and solvation, these factors are likely to be an important aspect of the protein resistance of PC films. This explanation is based on the theoretical and experimental studies of Grunze and co-workers,17 who found that differences in orientation and water solvation of PEG SAMs on Au and Ag substrates resulted in significant differences in protein adsorption resistance. In another section of part 1, we showed that the UV photopolymerization of DAPC, a diacetylene PC lipid, produced (poly)PSLBs with a relatively high defect density after drying, similar to UV(poly)bis-SorbPC. The relative surface coverage of RITC-BSA adsorbed on UV(poly)DAPC was 40% (Table 1), intermediate between the values

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measured for fused silica and UV(poly)bis-SorbPC. The greater degree of protein adsorption relative to UV(poly)bis-SorbPC may reflect the more hydrophobic nature of the DAPC tailgroups relative to those of bis-SorbPC, which contain a ester group near their termini. This explanation assumes that the degree of protein adsorption is correlated with the hydrophobicity of the defect sites in these PSLBs. Although it is well established that protein adsorption is favored at hydrophobic surfaces,24 the underlying mechanism(s) are still poorly understood. RITC-BSA adsorption to UV(poly)DAPC was also measured on PSLBs that were not dried. In this case, the relative protein surface coverage was 20% (Table 1), half of that measured on bilayers that were dried. These results indicate that removing the PSLB from water creates structural defects, probably due to desorption of lipid material, that strongly adsorb protein. Comparison of Protein Adsorption Isotherms. RITC-BSA adsorption isotherms were measured on several PSLB and reference surfaces at dissolved protein concentrations ranging from nM to µM. In an effort to quantitatively compare the isotherms, the data were fit to one-site and two-site Langmuir models (eqs 1 and 2). At the outset, we note the assumptions of the Langmuir model are not likely met in our experiments. Specifically, the inherent assumptions that only one type of surface site is present, that lateral interactions between adsorbed molecules are absent, and that the adsorption process is reversible and measured at equilibrium are probably not valid.24a,33 Therefore, the Ka and Fmax values extracted from fitting the data are used here only as empirical parameters to enable apparent binding constants and relative binding site densities to be compared. Isotherms for RITC-BSA adsorption to an AA monolayer, clean fused silica, dried and rehydrated redox(poly)bis-SorbPC and redox(poly)DenSorbPC PSLBs, and a POPC PSLB are plotted in Figure 1a. Adsorption to the AA monolayer rises to an asymptote at relatively low dissolved protein concentrations, qualitatively indicative of strong binding. The isotherm on fused silica is less steep. Both data sets were fit well using a single-site Langmuir model (eq 1). Table 2 lists the parameters recovered from the respective fits: (i) The Ka values were 23 × 105 and 2.1 × 105 for the AA monolayer and fused silica, respectively. (ii) The corresponding Fmax values, which represent the relative density of binding sites at apparent saturation, were 0.98 and 0.79. These data are consistent with the relative surface coverage measurements discussed above. We turn now to a comparison of the redox(poly)bisSorbPC and POPC data. Although when exposed to 1 mg/ mL RITC-BSA, the surface coverages on these films are equivalent (Table 1), the isotherms are clearly different. As the expanded view in Figure 1b reveals, at dissolved protein concentrations less than 2.5 µM, the (poly)PSLB adsorbed more protein. The shape of the isotherm more closely resembles that of arachidic acid (i.e., comparatively strong binding) than that of POPC (i.e., weaker binding). Fitting the data sets to a one-site Langmuir model produced the following: (i) For POPC, the fit was adequate and returned a Ka of 1.12 × 105 and a Fmax of 0.10. The latter quantity indicates a site density of approximately 10% relative to that of an AA monolayer, which is consistent with the surface coverage data listed in Table 1. (ii) In contrast, for redox(poly)bis-SorbPC, a one-site model produced a relatively poorer fit (R2 of 0.969) with a Ka of 2.9 × 106 and a Fmax of 0.06. Fitting the data to a two-site model was statistically better (R2 ) 0.997), (33) Norde, W.; Amusiem, A. C. I. Colloids Surf. 1992, 66, 73.

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Langmuir, Vol. 19, No. 5, 2003 1771 Table 2. Parameters Obtained from Fitting BSA Adsorption Data to a Single-Site Langmuir Adsorption Model type of film or reference surfacea arachidic acid monolayer fused silica POPC bilayer, not dried redox(poly)bis-SorbPC, dried and rehydrated UV(poly)bis-SorbPC, not dried redox(poly)DenSorbPC, dried and rehydrated

Ka × 10-5 b

R2

Fmaxb

23 ((2.7) 2.1 ((0.19) 1.12 ((0.073) 29 ((5.6)

0.991 0.996 0.998 0.969

0.98 ((0.028) 0.79 ((0.031) 0.10 ((0.0032) 0.06 ((0.007)

2.0

0.981 0.08

6.3 ((0.46)

0.997 0.29 ((0.0065)

a UV and redox refer to the method of PSLB polymerization. Values in parentheses are estimated standard deviations obtained from fitting the data. Where standard deviations are listed, at least three independent sets of isotherm data were measured for that surface.

b

Figure 1. (a) TIRF isotherms for BSA adsorption on an arachidic acid monolayer (4), a redox(poly)DenSorbPC PSLB (O), clean fused silica (1), a redox(poly)bis-SorbPC PSLB ()), and a fluid POPC PSLB (9). Error bars were computed from at least three independent measurements. At concentrations less than 2.5 µM, error bars were not plotted to enhance visualization of the data. (b) Normalized BSA adsorption isotherms, taken from (a) and plotted on an expanded scale, for redox(poly)bis-SorbPC (b) and fluid POPC PSLBs (9).

returning Ka1 and Ka2 values of 8 ((2.6) × 105 and 2 ((1.5) × 107 and Fmax values of 0.04 ((0.0068) and 0.02 ((0.0053). This analysis suggests that there is a more heterogeneous distribution of protein adsorption sites on a rehydrated redox(poly)bis-SorbPC PSLB, relative to a POPC PSLB. Additional support for site heterogeneity is provided by the finding, discussed above, that only two-thirds of the RITC-BSA adsorbed to a redox(poly)bis-SorbPC PSLB could be removed by rinsing the film with 2% Triton X-100. We conjecture that the PSLB contains at least two classes of sites: (i) a minor population of higher affinity sites that dominate the adsorption behavior at low protein concentrations and (ii) a majority population of lower affinity sites, similar in binding strength to those on a POPC PSLB. An adsorption isotherm was also measured on a bisSorbPC PSLB that was polymerized in the TIRF flow cell by UV irradiation and never dried. In this case, fitting the data to a one-site Langmuir model returned a Ka of 2.0 × 105 and a Fmax of 0.08 (R2 ) 0.981). The fit could not be

improved using a two-site model. In light of the data presented above, this result indicates that the higher affinity adsorption sites present in a rehydrated redox(poly)bis-SorbPC PSLB are produced when the film is dried. However, regardless of whether the film remains hydrated or is dried, the total density of adsorption sites was approximately 6-8% relative to that of an AA monolayer, which is consistent with the surface coverage data listed in Table 1. A possible origin for the higher affinity sites in a dried redox(poly)bis-SorbPC PSLB is that unreacted monomers or lower molecular weight polymer fragments are desorbed when the film is removed from water, exposing the hydrophobic interior of the bilayer. As in all radically initiated chain polymerizations, redox-initiated polymerization of bis-SorbPC will generate a polydisperse range of molecular weight fragments.34 Although evidence from part 1 of this report18 suggests that redox(poly)bis-SorbPC behaves as a cross-linked, semicontinuous molecular film, small defects arising from desorption of a small number of monomers and/or low weight molecular oligomers would likely not be observable by the methods used to interrogate the film structure in part 1. In other words, protein adsorption measurements are more sensitive to the presence of a small number of physically minute defects than ellipsometry and AFM. The isotherm for RITC-BSA adsorption to a redox(poly)DenSorbPC PSLB is also plotted in Figure 1a. With respect to shape, the isotherm is more similar to that of fused silica than to that of redox(poly)bis-SorbPC. Fitting the data to a one-site model returned a Ka of 6.3 × 105 and a Fmax of 0.29. Relative to the adsorption behaviors discussed above, these data are indicative of relatively weak binding to a relatively high density of adsorption sites. This finding provides additional support for our interpretation of the higher RITC-BSA surface coverage on (poly)DenSorbPC relative to (poly)bis-SorbPC (Table 1). As discussed above, the difference in protein adsorption on these two surfaces is not thought to arise from a greater degree of exposure of the hydrophobic bilayer interior of (poly)DenSorbPC. If protein adsorption on (poly)DenSorbPC is predominately hydrophobically driven, then the isotherm would likely mimic the shape of the isotherm on the hydrophobic AA monolayer, albeit with a lower density of sites. Rather, protein adsorption on (poly)DenSorbPC more closely mimics the relatively weaker adsorption on POPC, although the site density is about (34) Odian, G. Principles of Polymerization; John Wiley & Sons: New York, 1991.

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Figure 2. AFM images and linescans obtained on surfaces before and after incubation with a 1 mg/mL BSA solution: (i) A clean Si wafer before (a) and after (b) BSA adsorption. (ii) A dried redox(poly)bis-SorbPC PSLB, formed on a Si wafer, before (c) and after (d) BSA adsorption. (iii) A redox(poly)DenSorbPC PSLB, formed on a Si wafer, before (e) and after (f) BSA adsorption. The image areas are 1 µm2 with a height scale of 3 nm. The inset in (f) is a zoomed image showing BSA molecules adsorbed on the redox(poly)DenSorbPC film.

5-fold greater. In summary, the comparison of isotherms provides additional support for our proposal that crosslinking polymerization of the dienoyl groups in DenSorbPC may alter PC headgroup orientation, resulting in increased protein adsorption. AFM Measurements on Adsorbed Proteins. BSA adsorption to redox(poly)bis-SorbPC and redox(poly)DenSorbPC films was also examined by AFM. AFM has been used extensively to image protein films on a variety of surfaces, including supported lipid layers.35-37 In some (35) (a) Bernard, A.; Renault, J. P.; Michel, B.; Bosshard, H. R.; Delamarche, E. Adv. Mater. 2000, 12, 1067-1070. (b) Yang, J.; Tamm, L. K.; Tillack, T. W.; Shao, Z. J. Mol. Biol. 1993, 229, 286-290. (c) Viitala, T.; Vikholm, I.; Peltonen, J. Langmuir 2000, 16, 4953-4961. (d) Raab, A.; Han, W.; Badt, D.; Smith-Gill, S. J.; Lindsay, S. M.; Schindler, H.; Hinterdorfer, P. Nat. Biotechnol. 1999, 17, 902-905. (36) Mueller, H.; Butt, H.-J.; Bamberg, E. J. Phys. Chem. B 2000, 104, 4552-4559.

cases, individual protein molecules have been imaged, providing information on protein adsorption behavior, surface structure, and activity [e.g., ref 36]. AFM measurements can also provide an estimate of surface coverage when the amount of adsorbed protein is relatively low (i.e., submonolayer).37 In Figure 2 are shown AFM images and example linescans of three surfaces measured before and after a 30 min incubation in a 1 mg/mL BSA solution: a clean Si wafer, a Si wafer coated with a dried redox(poly)bisSorbPC PSLB, and a Si wafer coated with a dried redox(poly)DenSorbPC PSLB. For the SiO2 surface, a change in surface topography occurred as a result of BSA adsorption (Figure 2a,b). The root-mean-square (rms) roughness increased from 0.10 to 0.35 nm, as illustrated (37) Coen, M. C.; Lehmann, R.; Gro¨ning, P.; Bielmann, M.; Galli, C.; Schlapbach, L. J. Colloid Interface Sci. 2001, 233, 180-189.

PSLBs Formed by Vesicle Fusion

by comparing the linescans. This result is qualitatively consistent with the high degree of protein adsorption to fused silica measured by TIRF (Table 1). In contrast, BSA adsorption to the redox(poly)bis-SorbPC PSLB (Figure 2c,d) produced negligible changes in surface topography and rms roughness (the latter increased from 0.125 to 0.133 nm). However new surface features, presumably BSA molecules, are clearly visible in the postadsorption image. These features were widely distributed, making it evident that most of the PSLB surface was protein-free. A greater degree of BSA adsorption to redox(poly)DenSorbPC is apparent by comparing the last set of images and linescans (Figure 2e,f). The surface features were visibly more dense than on the redox(poly)bis-SorbPC PSLB, and the rms roughness increased moderately, from 0.15 to 0.20 nm. On both types of PSLBs, the adsorbed proteins appeared randomly distributed. The apparent size of the adsorbates was fairly uniform, although a minority was slightly larger, both in the substrate plane and normal to the plane. With respect to the smaller, more uniform features, it is not clear if these are aggregates of protein molecules or possibly individual molecules. However, a “zoomed” image (inset in Figure 2f) shows that many of these features have the shape of a prolate ellipsoid, with apparent lateral dimensions of 15-25 nm by 5-15 nm. Thus the aspect ratio is approximately 2 to 1, which is similar to the molecular dimensions of BSA (140 Å by 40 Å29). This similarity suggests that a significant fraction of the adsorbates imaged on PSLBs are individual protein molecules. The discrepancy between the apparent lateral dimensions and the known molecular dimensions is due to “tip broadening” effects. It is well documented that lateral resolution in AFM is limited by convolution of the probe tip radius with features on the surface of the sample that are smaller than the tip radius.37,38 Consequently, molecular-scale features are often artificially enlarged. To compare and contrast with the TIRF measurements, the surface coverage of BSA adsorbed to the PSLB surfaces was estimated using a bearing analysis, which provides a histogram of surface feature height or depth.37 (A bearing analysis was not performed on the BSA/Si wafer. Simulations with model surfaces and adsorbates have shown that protein coverages of greater than 50% cannot be accurately determined by AFM due to tip broadening effects.37) Features corresponding to adsorbed protein molecules in each image were highlighted by selecting an appropriate threshold height. The analysis returned coverages of 1.2% ((0.4%, n ) 4) for redox(poly)bis-SorbPC and 7.8% ((1.1%, n ) 4) for redox(poly)DenSorbPC, expressed as a percentage of the total surface area on the respective PSLB. These surface coverages are lower than the corresponding estimates obtained from TIRF measurements (see above). Several factors probably contributed to this discrepancy: (i) In the bearing analysis, it is assumed that all adsorbed proteins are detectable as topographic features. Since the substrate surface has an inherent, finite roughness, it is impossible to select a height threshold that highlights 100% of the protein features and simultaneously rejects all of the substrate features. However, the PSLB surfaces examined here were very smooth relative to the dimensions of BSA, which provides confidence that the percentage of undetected protein was minimal. (ii) Tip broadening effects exaggerate the size and thus the surface area of a protein feature. (iii) For reasons discussed above, when the absolute surface (38) Howland, R.; Benatar, L. A Practical Guide to Scanning Probe Microscopy; Park Scientific Instruments: Sunnyvale, CA, 1993.

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coverage is much less than a monolayer, the TIRF method is likely to generate an overestimate of relative surface coverage. However, the TIRF and bearing analysis results are in good agreement when the ratio of BSA surface coverages on the two types of PSLBs is considered. From both methods, a ratio of approximately 0.20-0.25 was obtained for the coverage on redox(poly)DenSorbPC relative to the coverage on redox(poly)bis-SorbPC. Thus in summary, the discrepancy between the results obtained using AFM and TIRF is attributed to systematic differences between the two methods. Finally, fibrinogen adsorption to redox(poly)bis-SorbPC PSLBs was also examined by AFM, to assess if the resistance to nonspecific BSA adsorption discussed above applies to other proteins. Imaging was performed on PSLBs before and after a 30 min incubation in a 1 mg/mL fibrinogen solution (images not shown). The surface coverage of adsorbed fibrinogen, estimated using a bearing analysis, was 2% ((1%, n ) 3), comparable to the results obtained with BSA. Thus the protein adsorption resistance of a redox(poly)bis-SorbPC appears to be a general phenomenon, which is consistent with studies of protein adsorption to fluid PC lipid bilayers [e.g., ref 9]. Conclusions In part 1 of this report,18 we demonstrated that redoxinitiated radical polymerization of bis-SorbPC produces a nearly defect-free, cross-linked PSLB. Here we have demonstrated that a dried redox(poly)bis-SorbPC bilayer, upon exposure to a 1 mg/mL BSA solution, exhibits a resistance to nonspecific protein adsorption that is equivalent to that of a fluid POPC PSLB. This finding provides additional support to the evidence presented in part 1 showing that the structure of a redox(poly)bis-SorbPC PSLB is not compromised upon removal from water or rinsing in surfactant solutions or organic solvents, which are conditions that destroy a fluid bilayer. Furthermore, since the lipids in a cross-linked bis-SorbPC PSLB are laterally immobile, our results demonstrate that fluidity is not required for a PC lipid bilayer to be protein resistant. TIRF measurements provide an upper estimate of the amount of BSA adsorbed from a 15 µM solution; for both fluid POPC and redox(poly)bis-SorbPC PSLBs, the surface coverage was approximately 6% of a monolayer. The protein resistance of redox(poly)bis-SorbPC compares favorably to published data on protein adsorption to PCfunctionalized thin films. Murphy et al.13a measured BSA adsorption on polymer films with incorporated PC groups. Adsorption from solutions that were 0.05 and 2 g/L in dissolved protein produced coverages of 20% and 36%, respectively, relative to the coverages measured on clean SiO2. Subsequent work with PC-bearing organic monolayers generated similar results.15b Whitesides and coworkers examined protein adsorption to PC-terminated SAMs on Au.3c At dissolved protein concentrations of 1 mg/mL, they measured surface coverages of 32% and 9% for fibrinogen and lysozyme, respectively, relative to the amounts of these proteins adsorbed on a methylterminated SAM. Relative to redox(poly)bis-SorbPC, the protein resistance of other types of (poly)PSLBs was weaker. Specifically, PSLBs polymerized under different conditions or formed from different monomers contained a significant density of structural defects18 that resulted in a greater degree of protein adsorption. (This includes (poly)DAPC, which historically has been considered to be a biocompatible material4b,11a). However, regardless of monomer

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type or polymerization method, all of the (poly)PSLB films examined here exhibited some degree of protein resistance relative to a hydrophobic AA monolayer and a clean SiO2 surface. This observation is consistent with numerous studies showing that the PC moiety resists irreversible protein adsorption.3,4b,8-15 In conclusion, our results suggest that with further development, redox(poly)bis-SorbPC PSLBs may be useful as transducer coatings in receptor-based biosensors, for the following reasons: (i) On the basis of their observed protein resistance, they may be resistant to “biofouling” in complex biological matrixes. (ii) They are highly stable structures, which may allow rinsing, regeneration, and/ or reuse under conditions that would compromise a fluid bilayer (e.g., surfactant washing). (iii) A (poly)lipid bilayer formed from bis-SorbPC may prove to be a suitable

Ross et al.

architecture for reconstitution and stabilization of transmembrane protein receptors. Acknowledgment. The work described herein was partially supported by the National Science Foundation under Grant Numbers CHE-0108805 to S.S.S. and DMR-9619887 to D.F.O. and by Grant Number DE-FG0302ER15378 from Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Research, U.S. Department of Energy. E.E.R. was supported in part by a Graduate Research Fellowship in Analytical Chemistry from the Analytical Research & Development Department of Pfizer Global Research & Development. LA026198D