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In Situ Formation and Characterization of Poly(ethylene glycol)-Supported Lipid Bilayers on Gold Surfaces Jeffrey C. Munro and Curtis W. Frank* Department of Chemical Engineering, Stanford University, Stanford, California 94305-5025 Received June 30, 2004. In Final Form: August 23, 2004 Inclusion of a polymer cushion between a lipid bilayer membrane and a solid surface has been suggested as a means to provide a soft, deformable layer that will allow for transmembrane protein insertion and mobility. In this study, mobile, tethered lipid bilayers were formed on a poly(ethylene glycol) (PEG) support via a two-step adsorption process. The PEG films were prepared by coadsorbing a heterofunctional, telechelic PEG lipopolymer (1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-poly(ethylene glycol)2000-N-[3-(2-(pyridyldithio)propionate]) (DSPE-PEG-PDP) and a nonlipid functionalized PEG-PDP from an ethanol/water mixture, as described in a previous paper (Munro, J. C.; Frank, C. W. Langmuir 2004, 20, 3339-3349). Then a two-step lipid adsorption strategy was used. First, lipids were adsorbed onto the PEG support from a hexane solution. Second, vesicles were adsorbed and fused on the surface to create a bilayer in an aqueous environment. Fluorescence recovery after photobleaching experiments show that this process results in mobile bilayers with diffusion coefficients on the order of 2 µm2/s. The mobility of the bilayers is decreased slightly by increasing the density of tethered lipids. The formation of bilayers, and not multilayer structures, is also confirmed by surface plasmon resonance, which was used to determine in situ film thickness, and by fluorimetry, which was used to determine quantitatively the fluorescence intensity for each 18 by 18 mm sample. Unfortunately, fluorescence microscopy also shows that there are large defects on the samples, which limits the utility of this system.
Introduction Poly(ethylene glycol) (PEG) has several useful characteristics that have made it suitable for a variety of biological applications. As discussed in several reviews, many studies have shown that surface-grafted PEG substantially reduces the adsorption of proteins and cells.2-4 In addition, PEG is an inert, water-soluble polymer that does not display any antigenic activity.3 These properties have made platforms such as PEG-grafted liposomes popular for drug delivery and other therapeutic treatments.5 PEG provides a barrier around the liposomes, protecting them from rupture and thereby increasing their circulation time.6 Despite the ability of grafted PEG chains to protect liposomes against aggregation and fusion, solubilized PEG has been shown to promote fusion of lipid vesicles by creating a large osmotic gradient from inside to outside the vesicles.7,8 The hydrophilic character of PEG along with its weak interactions with proteins and lipids has made PEG attractive for use as a cushion for supported lipid bilayers. Such supported lipid bilayers have been * To whom correspondence should be addressed. E-mail
[email protected]. (1) Munro, J. C.; Frank, C. W. Langmuir 2004, 20, 3339-3349. (2) Merrill, E. W. In Poly(Ethylene Glycol): Biotechnical and Biomedical Applications; Harris, J. M., Ed.; Plenum Press: New York, 1992; pp 199-220. (3) Golander, C.-G.; Herron, J. N.; Lim, K.; Claesson, P.; Stenius, P.; Andrade, J. D. In Poly(Ethylene Glycol): Biotechnical and Biomedical Applications; Harris, J. M., Ed.; Plenum Press: New York, 1992; pp 221-245. (4) Currie, E. P. K.; Norde, W.; Cohen Stuart, M. A. Adv. Colloid Interface Sci. 2003, 100-102, 205-265. (5) Harris, J. M.; Zalipsky, S. Poly(Ethylene Glycol): Chemistry and Biological Applications; American Chemical Society: Washington, DC, 1997. (6) Woodle, M. C. In Poly(Ethylene Glycol): Chemistry and Biological Applications; Harris, J. M., Zalipsky, S., Eds.; American Chemical Society: Washington, DC, 1997; pp 60-81. (7) Hui, S. W.; Kuhl, T. L.; Guo, Y. Q.; Israelachvili, J. Colloids Surf., B 1999, 14, 213-222. (8) Proux-Delrouyre, V.; Laval, J. M.; Bourdillon, C. J. Am. Chem. Soc. 2001, 123, 9176-9177.
proposed for use in applications such as biosensors9 or as models to study the structure and function of biomembranes.10 Solid-supported lipid bilayers have been a standard since they were first studied.10 They are able to selfassemble when vesicle suspensions are brought in contact with materials such as hydrophilic glass.11 However, this simple system has two disadvantages that have prompted others to develop more complicated systems. First, while the presence of a thin (1-2 nm) lubricating layer of water between the bilayer and the substrate leads to good fluidity on materials such as hydrophilic glass,12-14 lipid mobility is often suppressed on many other solid substrates, such as metals and metal oxides.15-17 Unlike glass, metal substrates offer a means to transduce electrical signals, which can be useful in biosensor applications.18,19 Second, large transmembrane proteins inserted into solid-supported bilayers can interact with the surface, leading to a loss of their lateral mobility.20-22 To prevent such interactions, many authors have suggested incorporating (9) Sackmann, E. Science 1996, 271, 43-48. (10) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105-113. (11) Brian, A. A.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 6159-6163. (12) Bayerl, T. M.; Bloom, M. Biophys. J. 1990, 58, 357-362. (13) Johnson, S. J.; Bayerl, T. M.; McDermott, D. C.; Adam, G. W.; Rennie, A. R.; Thomas, R. K.; Sackmann, E. Biophys. J. 1991, 59, 289294. (14) Koenig, B. W.; Krueger, S.; Orts, W. J.; Majkrzak, C. F.; Berk, N. F.; Silverton, J. V.; Gawrisch, K. Langmuir 1996, 12, 1343-1350. (15) Groves, J. T.; Ulman, N.; Boxer, S. G. Science 1997, 275, 651653. (16) Groves, J. T.; Ulman, N.; Cremer, P. S.; Boxer, S. G. Langmuir 1998, 14, 3347-3350. (17) Keller, C. A.; Kasemo, B. Biophys. J. 1998, 75, 1397-1402. (18) Cornell, B. A.; Braach-Maksvytis, V. L. B.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wieczorek, L.; Pace, R. J. Nature 1997, 387, 580583. (19) Raguse, B.; Braach-Maksvytis, V.; Cornell, B. A.; King, L. G.; Osman, P. D. J.; Pace, R. J.; Wieczorek, L. Langmuir 1998, 14, 648659. (20) Poglitsch, C. L.; Sumner, M. T.; Thompson, N. L. Biochemistry 1991, 30, 6662-6671.
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a soft, deformable hydrophilic polymer cushion between the lipid bilayer and solid substrate, as described in a review by Sackmann.9 In addition, since the self-assembly of thiols and disulfides onto gold surfaces has been studied extensively,23 many strategies have been devised to utilize this facile method to construct polymer-supported bilayer structures on gold. In a previous publication,1 we discussed many of the strategies used to construct bilayers on gold surfaces, including hybrid bilayers constructed with alkanethiol self-assembled monolayers (SAMs),24,25 oligo(ethylene glycol) (OEG)-containing SAMs,26 and multicomponent SAMs.18,19,27-30 All of these systems present well-organized, hydrophobic surfaces for the deposition of the free lipids that form the top leaflet. These systems are easy to construct and have been shown to reproduce the electrical properties (capacitance and impedance) of biological membranes.26 In addition, several polymeric systems have been proposed with the idea that hydrophilic polymeric spacers can provide a soft, deformable, water-swollen cushion.31-37 Several authors have used self-assembly of disulfide-containing lipopolymers to gold surfaces, followed by vesicle fusion to form bilayers on top of the polymer films.38-42 Self-assembly has the advantage of simplicity of monolayer and bilayer fabrication. Unlike LangmuirBlodgett transfer where planar substrates are used, films can be self-assembled on surfaces with different geometries, including within closed channels. However, the studies performed on these self-assembled lipopolymers have not demonstrated that bilayers with good lateral mobility or electrical barrier properties have been formed. Studies that have demonstrated good bilayer formation and mobility have used Langmuir-Blodgett techniques to first organize the amphiphilic polymer molecules at (21) Hinterdorfer, P.; Baber, G.; Tamm, L. K. J. Biol. Chem. 1994, 269, 20360-20368. (22) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 14773-14781. (23) Ulman, A. Chem. Rev. 1996, 96, 1533-1554. (24) Plant, A. L. Langmuir 1993, 9, 2764-2767. (25) Meuse, C. W.; Niaura, G.; Lewis, M. L.; Plant, A. L. Langmuir 1998, 14, 1604-1611. (26) Schiller, S. M.; Naumann, R.; Lovejoy, K.; Kunz, H.; Knoll, W. Angew. Chem., Int. Ed. 2003, 42, 208-211. (27) Stora, T.; Dienes, Z.; Vogel, H.; Duschl, C. Langmuir 2000, 16, 5471-5478. (28) Lahiri, J.; Kalal, P.; Frutos, A. G.; Jonas, S. T.; Schaeffler, R. Langmuir 2000, 16, 7805-7810. (29) Krishna, G.; Schulte, J.; Cornell, B. A.; Pace, R.; Wieczorek, L.; Osman, P. D. Langmuir 2001, 17, 4858-4866. (30) Krishna, G.; Schulte, J.; Cornell, B. A.; Pace, R. J.; Osman, P. D. Langmuir 2003, 19, 2294-2305. (31) Majewski, J.; Wong, J. Y.; Park, C. K.; Seitz, M.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1998, 75, 2363-2367. (32) Wong, J. Y.; Majewski, J.; Seitz, M.; Park, C. K.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1999, 77, 1445-1457. (33) Wong, J. Y.; Park, C. K.; Seitz, M.; Israelachvili, J. Biophys. J. 1999, 77, 1458-1468. (34) Luo, G. B.; Liu, T. T.; Zhao, X. S.; Huang, Y. Y.; Huang, C. H.; Cao, W. X. Langmuir 2001, 17, 4074-4080. (35) Kuhner, M.; Tampe, R.; Sackmann, E. Biophys. J. 1994, 67, 217-226. (36) Dietrich, C.; Tampe, R. Biochim. Biophys. Acta 1995, 1238, 183191. (37) Elender, G.; Kuhner, M.; Sackmann, E. Biosens. Bioelectron. 1996, 11, 565-577. (38) Spinke, J.; Yang, J.; Wolf, H.; Liley, M.; Ringsdorf, H.; Knoll, W. Biophys. J. 1992, 63, 1667-1671. (39) Erdelen, C.; Haussling, L.; Naumann, R.; Ringsdorf, H.; Wolf, H.; Yang, J. L.; Liley, M.; Spinke, J.; Knoll, W. Langmuir 1994, 10, 1246-1250. (40) Marra, K. G.; Kidani, D. D. A.; Chaikof, E. L. Langmuir 1997, 13, 5697-5701. (41) Hausch, M.; Zentel, R.; Knoll, W. Macromol. Chem. Phys. 1999, 200, 174-179. (42) Munro, J. C.; Frank, C. W. Polymer 2003, 44, 6335-6344.
Munro and Frank
the air-water interface along with free lipids, followed by transfer to a solid substrate and deposition of a top leaflet of free lipids via vesicle fusion or horizontal Langmuir-Schaeffer transfer.43-46 Langmuir-Blodgett transfer has advantages over self-assembly for monolayer and bilayer fabrication. Langmuir films containing amphiphilic lipopolymers along with free lipids to control the anchoring density can be organized at the air-water interface and then transferred to a solid substrate. In addition, the Langmuir-Blodgett approach provides a homogeneous hydrophobic surface, independent of the tethered lipid density, which is amenable to vesicle fusion. Our goal is to develop an in situ formation strategy for application in situations where an air-water interface is unavailable. One strategy used to overcome the lack of an air-water interface is to chemisorb PEG-grafted vesicles, followed by triggered vesicle fusion.47,48 One problem that arises in this tethered vesicle approach is the existence of grafted PEG chains that do not chemisorb to the surface, but remain exposed at the top of the bilayer.48 These free PEG chains could prevent adsorption of proteins, which could limit their utility in some applications. In a previous publication, we reported on the adsorption of heterofunctional, telechelic PEGs to gold surfaces.1 This approach took advantage of the benefits of both the SAM-based and polymeric support methods to create an easy-to-construct self-assembled structure that incorporates tethering to both the gold surface and the lipid bilayer and provides a substantial hydrophilic polymer cushion. The density of lipid anchoring groups can be controlled by coadsorbing the lipopolymer with nonlipid functionalized PEGs. In addition, by adsorbing from the correct solvent, the orientation of these multifunctional polymers can be controlled, eliminating the need for organizational methods such as the Langmuir-Blodgett techniques that have been used on a similar PEG system.43 Unfortunately, we found that vesicle fusion does not take place on the PEG surfaces studied here, likely due to their lack of purely hydrophilic or hydrophobic character. Therefore, a new approach taken in this study (depicted in Scheme 1) was to attempt first to create a surface more likely to promote vesicle fusion by adsorbing free lipids from hexane, a solvent that has been used to create planar bilayers.49 Then, vesicle fusion of standard small unilamellar vesicles was performed. The deposition of the lipids from hexane was monitored by ellipsometry and wetting measurements. Then, the characteristics of the bilayers created by vesicle fusion were studied using fluorescence recovery after photobleaching (FRAP) to determine their mobility and surface plasmon resonance (SPR) to determine their thickness. Experimental Section Materials. The heterofunctional PEG, 1,2-distearoyl-snglycero-3-phosphoethanolamine-N-poly(ethylene glycol)-2000-N[3-(2-(pyridyldithio)propionate] (DSPE-PEG-PDP), and lipids, L-R-phosphatidylcholine from egg (egg-PC), were purchased from Avanti Polar Lipids (Alabaster, AL). The egg-PC purchased is (43) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400-1414. (44) Shen, W. W.; Boxer, S. G.; Knoll, W.; Frank, C. W. Biomacromolecules 2001, 2, 70-79. (45) Naumann, C. A.; Prucker, O.; Lehmann, T.; Ruhe, J.; Knoll, W.; Frank, C. W. Biomacromolecules 2002, 3, 27-35. (46) Purrucker, O.; Fortig, A.; Jordan, R.; Tanaka, M. ChemPhysChem 2004, 5, 327-335. (47) Berquand, A.; Mazeran, P. E.; Pantigny, J.; Proux-Delrouyre, V.; Laval, J. M.; Bourdillon, C. Langmuir 2003, 19, 1700-1707. (48) Rossi, C.; Homand, J.; Bauche, C.; Hamdi, H.; Ladant, D.; Chopineau, J. Biochemistry 2003, 42, 15273-15283. (49) Hanke, W.; Schlue, W. R. Planar Lipid Bilayers: Methods and Applications; Academic Press: New York, 1993.
PEG-Supported Bilayers on Gold Surfaces Scheme 1. Two-Step Adsorption of Lipids to Chemisorbed PEG Film To Form Tethered Polymer-Supported Lipid Bilayer
a mixture of PC lipids containing approximately 90% saturated lipids (67% dipalmitoyl and 23% distearoyl), 10% unsaturated lipids (6% oleoyl, 2% palmitoleoyl, 1% linoleoyl), and 1% other PC lipids. The monofunctional PEG, methoxy-poly(ethylene glycol)-2000-N-[3-(2-(pyridyldithio)propionate] (MeO-PEG-PDP), was custom synthesized by Avanti Polar Lipids. The fluorescent lipid, N-(Texas Red sulfonyl)-1,2-dihexadecanoyl-sn-glycero-3phosphoethanolamine triethylammonium salt (Texas Red DHPE), was purchased from Molecular Probes (Eugene, OR). Rossville ethanol (200 proof) was purchased from Gold Shield Chemical Co. (Hayward, CA). Water was obtained using a Milli-Q water system (18.2 MΩ, Millipore Corp., Bedford, MA). Sample Preparation. Several different substrates were used, depending on the experimental technique. Silicon wafers, used in the ellipsometry and wetting measurements, were coated (in an Edwards Auto 306 evaporator, BOC Edwards, West Sussex, UK) with approximately 2 nm of chromium and 50 nm of gold (Plasmaterials, Livermore, CA). BK7 glass coverslips (VWR, 18 by 18 mm), used for FRAP experiments, were coated with approximately 2 nm of chromium and 35 nm of gold. In addition, hydrophilic BK7 glass coverslips were used for bilayer-onglass controls. Slides were heated until clear for 20 min in a 4:1 distilled water:7X cleaning solution (ICN Biomedicals, Inc., Aurora, OH), rinsed thoroughly with distilled water, and baked at 400 °C for 4 h. Finally, LASFN9 glass microscope slides, used in the SPR experiments, were coated with approximately 50 nm of gold. LASFN9 is a high refractive index glass that provides the necessary contrast with water. Gold adheres to LASFN9 glass sufficiently well such that no chromium adhesion layer is required. After removal from the evaporator, substrates were placed directly in polymer solutions. Mixtures of DSPE-PEG-PDP and MeO-PEG-PDP were prepared at a concentration of 0.1 mM in a 90/10 v/v % ethanol/water mixture with DSPE-PEG-PDP contents of 2, 10, 20, 30, and 100 mol %. Following adsorption, the substrates were removed from the polymer solutions, rinsed with ethanol, and dried under a stream of N2. Samples were then placed in a 0.5 mg/mL egg-PC (with or without 1 mol % Texas Red DHPE) in hexane solution for 30 min. Samples were subsequently removed, rinsed with
Langmuir, Vol. 20, No. 24, 2004 10569 hexane, and dried under a stream of N2. Finally, samples were incubated on a drop of approximately 100 µL of buffer (100 mM NaCl, 10 mM Tris, pH 8) mixed with 100 µL of vesicles in water. After a 15-min incubation period, the sample was rinsed vigorously with water and remained under water for all subsequent analysis. Vesicle Preparation. Vesicles were prepared by the standard extrusion method, as described elsewhere.50,51 Briefly, approximately 5 mg of egg-PC in chloroform, with or without 1 mol % Texas Red DHPE, was placed in a round-bottom flask. The chloroform was evaporated under a stream of N2, resulting in a thin lipid film on the flask. After at least 2 h in a vacuum desiccator, approximately 1 mL of water was added to the dry lipids. The flask was vortexed for about 5 min to dissolve all of the lipids in the water. The lipids were allowed to hydrate for at least 1 h. Finally, the lipid solution was extruded a minimum of 15 times through a 100 nm polycarbonate membrane using a Mini-Extruder (Avanti Polar Lipids, Alabaster, AL), resulting in a translucent solution of small unilamellar vesicles in water. Vesicles were stored in the dark at 4 °C until use. Ellipsometry. Dry thicknesses of adsorbed PEG films on gold, before and after the adsorption of lipids from hexane, were determined with a Gaertner L116C ellipsometer (Gaertner Scientific Corp., Chicago, IL) equipped with a He-Ne laser (λ ) 632.8 nm) at an incident angle of 70°, assuming an refractive index of 1.44.3,52 Measurements were taken at four different locations on each sample and averaged. Wetting Measurements. Advancing and receding water contact angles were measured using a FTÅ 200 contact angle goniometer (First Ten Angstroms, Portsmouth, VA). Water was advanced or withdrawn (0.5 µL/s) prior to measurement using a stepper-driven syringe pump. The syringe tip remained in the drop during the measurements. Images of each drop were acquired via a CCD camera and video capture card. Contact angles were determined using both sides of each of three drops placed at different locations on each sample. Fluorescence Recovery after Photobleaching. After vesicle fusion onto the samples, a sandwich was formed under water using a glass coverslip. Samples were imaged on a Nikon Eclipse E800 epifluorescent microscope using a 40× objective and Texas Red filter cube set. The samples were illuminated with 532-587 nm light by a super high pressure mercury arc lamp. Emitted fluorescence at 608-683 nm was collected by a Photometrics CoolSnap HQ CCD camera. Images were collected and analyzed using the Metamorph program (Universal Imaging Corp., Downingtown, PA). All measurements were conducted at room temperature, about 20 °C. Diffusion coefficients were determined using the Axelrod method assuming a circular disc beam spot.53 Samples were first bleached for 30 s and subsequently imaged with an exposure time of 50-300 ms at one-quarter the light intensity to monitor the recovery. Briefly, the fluorescence intensity of the bleached spot was determined as a function of time, correcting for background counts and photobleaching caused by illumination during image capture. Then the fractional fluorescence, f(t), was plotted versus time:
f(t) )
F(t) - F(0) F(∞) - F(0)
(1)
where F(0) is the initial fluorescence intensity (estimated as the intensity at the center of the bleaching area immediately following photobleaching) and F(∞) is the fluorescence intensity at infinite time (estimated as the intensity before bleaching, which assumes complete recovery). From the plot of f(t) versus time, the time at which f(t)τ1/2) ) 1/2 was determined, from which the diffusion coefficient, D, could be calculated: (50) Hope, M. J.; Bally, M. B.; Webb, G.; Cullis, P. R. Biochim. Biophys. Acta 1985, 812, 55-65. (51) Mayer, L. D.; Hope, M. J.; Cullis, P. R. Biochim. Biophys. Acta 1986, 858, 161-168. (52) Picard, F.; Buffeteau, T.; Desbat, B.; Auger, M.; Pezolet, M. Biophys. J. 1999, 76, 539-551. (53) Axelrod, D.; Koppel, D. E.; Schlessinger, J.; Elson, E.; Webb, W. W. Biophys. J. 1976, 16, 1055-1069.
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0.22r2 τ1/2
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where r is the radius of the bleaching spot, approximately 35 µm. The mobile fraction was determined by fitting f(t) to an exponential function and extrapolating to infinity. Note that there are significant errors in our analysis due to the necessity of estimating F(0) and F(∞). However, since the analysis is consistent throughout this paper and comparable to estimates made by others in the literature, the conclusions based on our results are reasonable. Fluorimetry. Quantitative fluorescence intensities were collected on an Edinburgh Instruments (Livingston, UK) fluorimeter. Samples were excited with 543 nm light, and emission was collected from 560 to 750 nm. The FRAP samples were rinsed thoroughly with 3.2 mL of a 1% N,N-dimethyldodecylamine N-oxide (LDAO) in water detergent solution to remove all the free lipids, including the Texas Red DHPE molecules. This solution was then placed in a cuvette and the fluorescent counts were measured. The fluorescence intensity was determined from the area of the emission peak minus the counts for a 1% LDAO control solution. Surface Plasmon Resonance. A more detailed description of SPR as well as a schematic of the experimental setup can be found elsewhere.54 A brief summary of the technique is provided here. Surface plasmons were excited using p-polarized light from a He-Ne laser beam at a wavelength of 632.8 nm. The chopped laser beam was reflected off the back side of the gold-coated glass slide, which was mounted on a θ/2θ goniometer in the Kretchmann configuration. The reflected intensity was then monitored by means of a photodiode and read out by a lock-in amplifier. By varying the angle of incidence, reflectivity-versusangle scans could be recorded. The observed minimum corresponds to the excitation of a surface plasmon; i.e., momentum and energy of the laser beam and the surface plasmon excitation are matched. The angle of excitation depends on the thickness and refractive index of the dielectric materials on top of the gold film. The resulting scans were fit to Fresnel calculations, where the different layers were represented by a simple box model.
Results Ellipsometry. Ellipsometry was used to determine dry film thicknesses. Table 1 shows the thicknesses of the PEG films before and after deposition of free lipids from hexane. The PEG film thicknesses before lipid deposition are in agreement with those we have published previously for the same system.1 As discussed in that publication, the PEG coils in the DSPE-PEG-PDP and MeO-PEG-PDP molecules are the same size. Therefore, the increase in thickness for the 100 mol % DSPE-PEG-PDP case is due to the presence of the DSPE groups that reside at the top of the adsorbed films. Independent of the DSPE-PEGPDP film content, there is significant deposition of eggPC to all of the PEG films. However, unlike the adsorbed PEG films, there is significant variation in the adsorbed egg-PC thickness, which was seen on a majority of the samples tested. Therefore, instead of an average value, ranges in the thickness are listed in Table 1. A typical lipid molecule is about 2.5 nm in length.55 Based on an average thickness for each DSPE-PEG-PDP density, the adsorbed egg-PC thicknesses correspond to a minimum of 0.6 lipid monolayer at 100 mol % DSPE-PEG-PDP to 1.7 lipid monolayers at 2 mol % DSPE-PEG-PDP. There is an apparent decreasing trend in adsorbed egg-PC thickness with increasing DSPE-PEG-PDP content. Wetting Measurements. Contact angle measurements provide evidence of the location of the tethered DSPE groups in the chemisorbed PEG films and of the (54) Knoll, W. Annu. Rev. Phys. Chem. 1998, 49, 569-638. (55) Nagle, J. F.; Tristram-Nagle, S. Biochim. Biophys. Acta 2000, 1469, 159-195.
Table 1. Ellipsometric Thickness of Dry PEG Films Before and After Lipid Deposition from Hexane DSPE-PEG-PDP, mol %
before
after
difference
2 10 20 30 100
3.1 ( 0.1a 2.9 ( 0.1 3.1 ( 0.1 3.2 ( 0.3 4.1 ( 0.4
6.4-10.6b 6.1-6.2 5.4-8.3 4.6-6.1 4.6-7.3
3.3-7.5 3.2-3.3 2.3-5.2 1.4-2.9 0.5-3.2
a All thicknesses in nanometers. Errors represent one standard deviation. b Thickness range based on multiple spots on two samples.
Figure 1. Advancing and receding water contact angles on adsorbed PEG films before and after deposition of egg-PC from hexane. Errors bars represent one standard deviation.
organizational state of the physisorbed egg-PC after its deposition from hexane. Figure 1 shows advancing and receding water contact angles before and after the deposition of egg-PC for a variety of tethered lipid densities. The contact angles for the chemisorbed PEG films are comparable to those previously published.1 The advancing contact angles do not change significantly upon deposition of egg-PC from hexane. For all but the lowest tethering density of 2 mol % DSPE-PEG-PDP, there is a slight decrease of 5-10° in the advancing contact angles. Meanwhile, all of the receding contact angles are below 10°, indicating the ability of the physisorbed egg-PC to rearrange in the presence of water. Fluorescence Recovery after Photobleaching. Following egg-PC deposition from hexane and subsequent vesicle fusion, samples were imaged by fluorescence microscopy. Independent of the tethered lipid density of the chemisorbed PEG films, each individual sample had two types of characteristic regions: bilayer regions and defect regions. Figure 2 shows an example of a bilayer region and its recovery on a 10 mol % DSPE-PEG-PDP film. The bright spots on the sample are physisorbed vesicles or other aggregates that remain on the surface even after vigorous rinsing. These aggregates were visible on the bilayer regions of every sample examined. The images shown in Figure 2 are 220 by 170 µm, but continuous bilayer regions on the order of several square millimeters were visible on all of the samples examined. These bilayer regions were used to determine the diffusion coefficients and mobile fractionssdetermined using the Axelrod approach described above in the Experimental Sectionsdisplayed in Figure 3a,b. Bilayer regions on at least three samples were examined for each tethered lipid density listed and several spots on each sample were measured. Also included in Figure 3 are values for a bilayer on glass (D ) 0.5 ( 0.1 µm2/s and mobile fraction ) 85 (
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Figure 2. Fluorescence recovery of 99/1 egg-PC/Texas Red DHPE bilayer on a PEG film with 10 mol % DSPE-PEG-PDP at (a) 0, (b) 1, (c) 3, and (d) 15 min after photobleaching. Photobleached spot is 70.2 µm in diameter with a diffusion coefficient of 2.1 µm2/s and a mobile fraction of 87%. Images are 220 by 170 µm.
2%), which provides a control for comparison. Note that a range of diffusion coefficients have been reported for a bilayer on glass, from 1 to 8 µm2/s.10,11,48,56 The value reported here falls near the bottom of the range of reported values and provides a quantitative means for comparing the diffusion coefficients determined for the bilayers on the PEG films. The lipids in hexane solution and the vesicles can each be made with or without the Texas Red DHPE probes. By including the probes in only one of the two lipid deposition steps, we can attempt to determine where the lipids deposited in each separate step reside in the final bilayer structure. There are two basic possibilities for how the lipids are deposited. First, the free lipids dissolved in hexane may form the bottom lipid leaflet and the lipids from the vesicles subsequently may form the top lipid leaflet. Second, lipids from the hexane solution and vesicles may mix in both leaflets in patches or in a completely homogeneous fashion. To help determine which of these situations occurs, two fluorescence cases were studied. In the first case, Texas Red DHPE was dissolved in hexane along with the egg-PC lipids and the vesicles also contained Texas Red DHPE. In the second case, Texas Red DHPE was dissolved in hexane along with the egg-PC lipids, but the vesicles were made from egg-PC only. Figure 3 includes diffusion and immobile fractions for both of these cases. Unfortunately, in addition to the bilayer regions, every sample examined had large defect regions visible under the fluorescence microscope. Similar defect regions are seen on all samples, independent of the tethered lipid density used. Figure 4 shows a few examples of these defect
regions for two different tethered lipid densities. As shown, there are grainy regions that when photobleached show no recovery. In Figure 4a,c the defect regions are separated by lines of bilayer region that may suggest the defects originate from some dewetting or crystallization phenomena. However, the defect regions formed come in all shapes and sizes making it difficult to determine exactly what the origin of the defects is, as discussed below in the Discussion section. The percentage of a sample covered with defect regions was generally in the range of 20-40% of the total area. Meanwhile, the bilayer-on-glass controls had no visible defect regions. Fluorimetry. Fluorimetry was used to determine the quantitative fluorescence intensity of each sample examined. The fluorescence intensity for each sample is reported in Figure 3c. Again, results for the two fluorescence cases are shown. The fluorescence intensities are normalized by the intensity for the control, a bilayer on glass. Each sample was examined under a fluorescence microscope before being rinsed with detergent solution. To ensure that the detergent rinsing was sufficient to remove all of the free lipids, select samples were reexamined on the microscope after rinsing to confirm that there was no detectable fluorescence. Also, the fluorescence intensities for the bilayer-on-glass samples were very repeatable, as shown by the relatively small standard deviation reported in Figure 3c. As noted above, unlike the bilayer-on-glass samples, PEG samples all had (56) Stelzle, M.; Miehlich, R.; Sackmann, E. Biophys. J. 1992, 63, 1346-1354.
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Figure 3. Fluorescence data for bilayers on PEG films as a function of tethered lipid density: (a) diffusion coefficients; (b) mobile fractions as determined from FRAP experiments; (c) quantitative fluorescence intensity as determined from fluorimetry. (The fluorescence intensity is normalized by the value for the bilayer-on-glass control.)
significant defect regions. Since the area covered by defects was not consistent from sample to sample and was not determined, there was significant variation in the quantitative fluorescence intensity from sample to sample, as indicated by the large error bars in Figure 3c. However, two important conclusions can be drawn from the fluorimetry data. First, multilayer structures are not formed on our samples. That is, the bilayer regions that show good mobility are not multilayer structures. If they were, a much larger fluorescence intensity would be detected. Second, when Texas Red DHPE is included in both lipid deposition steps, the fluorescence intensity is relatively constant, within the error of the measurement, as a
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Figure 4. Immobile defect regions after photobleaching on (a) a 30 mol % DSPE-PEG-PDP film with Texas Red DHPE included only in the lipids deposited from hexane and (b, c) a 10 mol % DSPE-PEG-PDP film with Texas Red DHPE included in both the lipids deposited from hexane and vesicles. Examples of defect and bilayer regions are labeled in each frame. The defect regions are grainy and show no recovery. Frames a and c also contain bilayer regions that are homogeneous and show recovery as indicated by the lack of a defined boundary between bleached and unbleached regions. Images are 220 by 170 µm.
function of tethered lipid density, indicating that similar bilayer structures are forming independent of the tethering density. Surface Plasmon Resonance. SPR was used to determine the in situ thicknesses of the bilayers on the
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PEG films. Samples were first examined under the fluorescence microscope to determine where good bilayer regions were. Then, SPR curves were collected for the sample, focusing the 1 mm2 laser spot on those bilayer regions. A two-layer box model was used to model the films. The water-swollen PEG film was assumed to have a refractive index of 1.36,3 while the lipid bilayer was assumed to have a refractive index of 1.45.52 The results of the modeling indicated that the two-step lipid adsorption process resulted in the addition of 6.6 ( 0.8 nm and 4.6 ( 0.7 nm, for 10 and 100 mol % DSPE-PEG-PDP films, respectively. These values are close to the expected bilayer thickness of about 5 nm.55 Note there is significant error in our measurement because we use separate samples to determine the before and after in situ thicknesses. There are slight differences in the evaporated gold thickness from sample to sample that can lead to errors in the calculated thicknesses beyond those captured in the standard deviation values shown above. Discussion Deposition Strategy. To form a stable polymersupported lipid bilayer, a few thermodynamic conditions must be met.9,57 First, the spreading pressure for the lipid bilayer on the polymer film must be positive. Second, the interactions between the bilayer and solid substrate must be weak to prevent dewetting. By tethering lipids to the polymer chains, the stability of the system is increased.9,45,46,58 Wagner and Tamm construct a similar PEG-supported system by organizing the lipopolymer and free lipids of the bottom leaflet at the air-water interface and then transferring this ordered film to a solid substrate via Langmuir-Blodgett methods.43 They report that defect-free bilayers are created using this strategy, indicating that there are no negative interactions between the lipids and PEG that cause the system to be unstable. Unfortunately, we found that a single vesicle fusion step was unsuccessful at forming a bilayer on our chemisorbed PEG films (data not shown). Vesicle fusion has been shown to be an easy and effective means of forming bilayers in situ on both hydrophilic9,15,59 and hydrophobic24,60 surfaces. However, vesicles have been shown to adsorb, but not fuse, to surfaces with intermediate water contact angles.61 Several strategies have been used in the literature to promote vesicle fusion to surfaces, including PEG-induced fusion,8,47 temperature,59 increased osmotic stress,62 vesicle composition,63 and vesicle size.59,64,65 However, none of these strategies was successful in producing any bilayer regions on the chemisorbed PEG films studied here. There may not be a strong enough driving force for the vesicles to spread on these PEG films under water. Therefore, as depicted in Scheme 1, a step in which free lipids were deposited from hexane was added (57) Elender, G.; Sackmann, E. J. Phys. II 1994, 4, 455-479. (58) Beyer, D.; Elender, G.; Knoll, W.; Kuhner, M.; Maus, S.; Ringsdorf, H.; Sackmann, E. Angew. Chem., Int. Ed. Engl. 1996, 35, 1682-1685. (59) Reimhult, E.; Hook, F.; Kasemo, B. Langmuir 2003, 19, 16811691. (60) Kalb, E.; Frey, S.; Tamm, L. K. Biochim. Biophys. Acta 1992, 1103, 307-316. (61) Baumgart, T.; Kreiter, M.; Lauer, H.; Naumann, R.; Jung, G.; Jonczyk, A.; Offenhausser, A.; Knoll, W. J. Colloid Interface Sci. 2003, 258, 298-309. (62) Seitz, M.; Ter-Ovanesyan, E.; Hausch, M.; Park, C. K.; Zasadzinski, J. A.; Zentel, R.; Israelachvili, J. N. Langmuir 2000, 16, 60676070. (63) Nollert, P.; Kiefer, H.; Jahnig, F. Biophys. J. 1995, 69, 14471455. (64) Reviakine, I.; Brisson, A. Langmuir 2000, 16, 1806-1815. (65) Reimhult, E.; Hook, F.; Kasemo, B. J. Chem. Phys. 2002, 117, 7401-7404.
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to attempt to create a surface more likely to promote vesicle fusion. As noted in our previous publication,1 slightly higher tethered lipid densities can be achieved by adsorbing 100 mol % DSPE-PEG-PDP films from ethanol alone. Nonetheless, those films still do not form wellpacked lipid monolayers and do not promote vesicle fusion (data not shown). A method that has been used to deposit lipid films onto solid substrates is the so-called painting method.66 Lipids are deposited onto hydrophobic surfaces from ethanol or an alkane solvent, such as hexane or decane, and then exposed to water. To reduce the surface energy that arises from the interactions of the hydrophobic surface with water, the free lipids self-assemble into an upper leaflet where the hydrophilic headgroups are now exposed to the water.19,66-68 In this study, we attempt to use a similar driving force to deposit the bottom lipid leaflet. Since hexane is a poor solvent for PEG,69 there is a driving force to reduce the contact between hexane and PEG, which can be achieved by the deposition of free lipids from hexane. In addition, to reduce the interaction energy, the alkyl tails of the lipids would prefer to be exposed to the hexane solvent over the headgroups. The SPR, fluorimetry, and FRAP results support the conclusion that the two-step lipid deposition strategy used here leads to the formation of mobile bilayer regions on the chemisorbed PEG films. The SPR results indicate that approximately 5 nm of lipids is added onto the PEG films during the two-step lipid adsorption procedure. The fluorimetry results indicate that the fluorescence intensity for the PEG samples is close to that for a bilayer on glass. The fluorimetry results certainly rule out the formation of any multilayer structures, which ensures that the diffusion coefficients determined from the FRAP measurements are indeed for a bilayer and not some multilayer structure. However, the experimental results also make it clear that the formation of these bilayers is complex and may vary depending on the tethered lipid density. Note that while the two-step bilayer formation strategy studied here has proven successful in creating tethered bilayer regions, there are still a few disadvantages to this strategy. First, as illustrated by Figure 4, there are still large defect regions on the PEG films. This may result from the inhomogeneity of the lipid deposition from hexane. Second, because hexane is used as the deposition solvent for egg-PC in the first step, hexane may be incorporated into the tethered bilayer film, which is not ideal for biological applications.68 Mechanism of Bilayer Formation. Although mechanisms have been postulated for vesicle fusion processes on both hydrophilic and hydrophobic surfaces, experimental evidence is not conclusive.64,70,71 In this work, there is the added complexity of an inhomogeneous physisorbed lipid film with neither purely hydrophilic nor hydrophobic character on which vesicle fusion is to take place. The ellipsometry results shown in Table 1 indicate that, depending on the tethered lipid density, between 0.6 and 1.7 egg-PC monolayers are deposited on top of the PEG films from hexane. Unlike the ideal scenario depicted in (66) Florin, E.-L.; Gaub, H. E. Biophys. J. 1993, 64, 375-383. (67) Steinem, C.; Janshoff, A.; Ulrich, W. P.; Sieber, M.; Galla, H. J. Biochim. Biophys. Acta 1996, 1279, 169-180. (68) Ding, L.; Li, J.; Dong, S.; Wang, E. J. Electroanal. Chem. 1996, 416, 105-112. (69) Yuan, Q. W. In Polymer Data Handbook; Mark, J. E., Ed.; Oxford University Press: New York, 1999; pp 542-552. (70) Keller, C. A.; Glasmastar, K.; Zhdanov, V. P.; Kasemo, B. Phys. Rev. Lett. 2000, 84, 5443-5446. (71) Johnson, J. M.; Ha, T.; Chu, S.; Boxer, S. G. Biophys. J. 2002, 83, 3371-3379.
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Scheme 1 where a single organized egg-PC monolayer is shown, the ellipsometric thicknesses and wetting measurements indicate that these physisorbed lipids are not well organized. There are large thickness ranges across individual samples. Also, if the physisorbed lipids were well-organized, with their alkyl tails facing the air, the advancing water contact angles would be closer to 100°, indicating a hydrophobic surface, which is not the case, as shown by Figure 1. Furthermore, the ellipsometric thicknesses and advancing water contact angles after the egg-PC deposition from hexane vary depending on the tethered lipid density, indicating that the physisorbed lipids do not form a well-organized bottom leaflet. Finally, the low receding water contact angles indicate that the physisorbed lipids are easily rearranged in the presence of water, which may have important consequences for the final vesicle fusion step. During vesicle fusion, an aqueous vesicle solution is introduced to the surface. Water is a good solvent for PEG. In addition, the lipids will rearrange such that the headgroups and not the alkyl tails interact with the water. The dynamics of the physisorbed lipid rearrangement in water along with vesicle adsorption and fusion are unknown. It is possible that, when the surface is wet by the aqueous vesicle solution, the physisorbed lipids rearrange before and during any vesicle adsorption and fusion. In addition, the dynamics of this process may vary with changes in the tethered lipid density. As the differences in the advancing water contact angles show, the physisorbed lipids are not organized consistently for all of the tethered lipid densities used. Therefore, the vesicles may adsorb and/or fuse quite differently on the surfaces depending on the DSPE-PEG-PDP content. Differences in the formation dynamics with changes in the tethered lipid density are highlighted by the fluorimetry and FRAP results, where the two lipid deposition steps could be followed separately by including the Texas Red DHPE probes in only one or both steps. For the 10 mol % DSPE-PEG-PDP case, the fluorimetry results (see Figure 3c) show that when only the lipids in hexane included Texas Red DHPE the fluorescence intensity was 51%, compared to 86% when both lipid deposition steps included Texas Red DHPE. Therefore, slightly more than half of the bilayer lipids come from the hexane deposition step and a little less than half come from the vesicle fusion step. On the other hand, for the 30 and 100 mol % DSPE-PEG-PDP cases, the fluorescence intensities decrease only a few percent when the Texas Red DHPE probes are left out of the vesicles. This would seem to indicate that most of the lipids that form the bilayer regions come from the hexane deposition step. However, if the 30 and 100 mol % DSPE-PEG-PDP samples are immersed only in water instead of a vesicle solution following the hexane deposition step, there are no signs of bilayer formation, indicating that the presence of vesicles in the last step is important to bilayer formation. However, there is a significant problem with the fluorimetry results that cannot be overlooked. As shown in Figure 4, there are large defect regions on each sample. The percentage of the substrate covered by defects varies from sample to sample. Without systematically acquiring images of the entire substrate, this percentage cannot be determined. In addition, it is unclear what the nature of the defects is. Since similar defect regions appear whether or not the vesicles used include Texas Red DHPE, they likely consist of lipid aggregates that, at least in part, result from the hexane deposition step and subsequent exposure to water. It is unclear if vesicles also adsorb in these defect regions. Either way, the fluorimeter results
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average over the bilayer and defect regions, which may lead to a significant error in these measurements. Diffusion Coefficients. The diffusion coefficients and immobile fractions are consistent with measurements on similar telechelic polymer supported bilayers.43,45,48 However, to compare previous results with those from this study, we need to translate the DSPE-PEG-PDP mole percent values used here to a tethered lipid density, which can be defined as the percent of total lipids in the bottom leaflet that are tethered to PEG chains. In studies where the lipopolymers are organized at the air-water interface, the tethered lipid density is simply controlled by adjusting the ratio of free lipids to lipopolymer added to the airwater interface. Note that if one transfer pressure is used, then this mixing ratio also determines the conformation of the PEG chains.43 In this study, we control the tethered lipid density by diluting our lipoPEG with nonlipid functionalized PEG. As shown in our previous paper,1 the conformation of the PEG chains on the surface is determined by the adsorption solvent. For the ethanol/water mixture used here, the PEG chains are in mushroom conformations with a projected area of approximately (4.3 nm)2 ) 18.5 nm2.1 Based on Langmuir films of these lipoPEG/lipid mixtures, the projected area of a lipid is approximately 0.65 nm2.72 Therefore, the 100 mol % DSPE-PEG-PDP film corresponds to a tethered lipid density of approximately 0.65 nm2/18.5 nm2 ) 3.5 mol %. The other values of DSPE-PEG-PDP mole percent of 2, 10, 20, and 30 mol % correspond to tethered lipid densities of 0.1, 0.4, 0.7, and 1.1 mol %, respectively. The diffusion coefficients and mobile fractions shown in Figure 3a,b are similar to those measured by Wagner and Tamm on their PEG-supported bilayers.43 Wagner and Tamm observed a decrease in diffusion coefficient and mobile fraction at a tethered lipid density of about 4 mol %. The same trend is seen here, where only at the highest tethered lipid density of 3.5 mol % is there a significant decrease in the diffusion coefficient and mobile fraction. Unfortunately, we were not able to measure the inner and outer leaflet diffusion coefficients independently because of the apparent lipid mixing that takes place in our two-step deposition process, as discussed above. Therefore, we are unable to determine how the increase in tethered lipid density affects the diffusion of the inner and outer leaflets. Results from the literature have shown that high tethering densities can decrease the mobility of both leaflets.45 Conclusions This study has demonstrated the feasibility of creating tethered polymer-supported lipid bilayers in situ. Chemisorbed PEG films are formed from a mixture of a heterofunctional, telechelic PEG lipopolymer (DSPE-PEG-PDP) and a nonlipid functionalized PEG-PDP (MeO-PEG-PDP), as described in a previous publication.1 While vesicle fusion does not occur directly on these chemisorbed PEG films, we have demonstrated that a two-step lipid adsorption process can create bilayer regions on the PEG films with a range of tethered lipid densities from 0.1 to 6 mol %. After chemisorption of the PEG films to the gold surface, lipids are physisorbed on the PEG films from a hexane solution. Finally, vesicles are adsorbed and fused to the sample in an aqueous environment, resulting in tethered lipid bilayers. FRAP measurements have shown that these bilayer regions are mobile with diffusion coefficients comparable to other polymer-supported systems. Unfortunately, fluorescence microscopy has also shown that (72) Ke, P. C.; Naumann, C. A. Langmuir 2001, 17, 5076-5081.
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there are large defects present for all tethered lipid densities. Due to these large defects, these bilayers have poor electrical barrier properties (high capacitance and low resistance) compared to biological membranes. (Electrical properties were determined by electrochemical impedance spectroscopy measurements, but the results are not shown or discussed.) While the two-step lipid deposition process used here was successful in creating mobile lipid bilayer regions, the need for the deposition of free lipids from hexane complicates matters. The physisorbed lipid films deposited from the hexane solution are poorly organized, and the mechanism of bilayer formation during the subsequent vesicle fusion step is not fully understood. Given that PEG
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is such a complex polymer with many odd characteristics (such as solubility in hydrophilic solvents like water and hydrophobic solvents such as toluene), we suggest that a polymer with a more pure hydrophilic character may be beneficial. Acknowledgment. Financial support for the project was provided by a NSF Graduate Research Fellowship (J.C.M.), the NSF XYZ-on-a-chip program under DMR9980799, and the Center on Polymer Interfaces and Macromolecular Assemblies (CPIMA), which is sponsored by the NSF-MRSEC program under DMR-9808677. LA048378O
