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Atomic Force Microscopy Assisted Immobilization of Lipid Vesicles Holger Scho¨nherr,* Dorota I. Rozkiewicz, and G. Julius Vancso Materials Science and Technology of Polymers, MESA+ Institute for Nanotechnology and Faculty of Science and Technology, University of Twente, 7500 AE Enschede, The Netherlands Received January 13, 2004. In Final Form: May 3, 2004 We report on a new approach to direct the immobilization of unilamellar lipid vesicles on substratesupported lipid bilayers in a spatially confined manner. The adsorption of vesicles from solution is limited to areas of disorder in the bilayers, which is induced by scanning a pattern in situ with an atomic force microscopy (AFM) tip using high imaging forces. Lines of vesicles with a length exceeding 25 µm and a width corresponding to that of a single surface-immobilized vesicle have been fabricated. The adsorbed vesicles are effectively immobilized and do not desorb spontaneously. However, AFM with forces of several nanoNewtons allows one to displace vesicles selectively. The novel methodology described, which may serve as a platform for research on proteins incorporated in the lipid bilayers comprising the vesicles, does not require chemical labeling of the vesicles to guide their deposition.
Introduction Substrate-supported lipid bilayers serve as valuable model systems for biological membranes1 and for studies of membrane constituents and their function.2 Among the different methods to fabricate substrate-supported lipid bilayers, the process of vesicle fusion3 has attracted interest owing to its ease, reproducibility, and widespread applicability on substrates, including glass,4 mica,5 and hydrophilized gold.6 Although proteins have been successfully incorporated and shown to be functional in substrate-supported lipid bilayers,7 the requirement of a water layer between bilayer and substrate to protect sensitive proteins from malfunction or denaturation has been realized and spurred significant research in the area of, for example, polymer-tethered lipid membranes.8 For applications, including molecular separation,9 lipid bilayer compartments or patterned bilayers have been * Corresponding author. Tel: ++31 53 489 3170. Fax: ++31 53 489 3823. E-mail:
[email protected]. (1) (a) Plant, A. L. Langmuir 1993, 9, 2764-2767. (b) Sackmann, E. Science 1996, 271, 43-48. (2) (a) McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A. Biochim. Biophys. Acta 1986, 864, 95-106. (b) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 14773-14781. (c) Grakoui, A.; Bromley, S. K.; Sumen, C.; Davis, M. M.; Shaw, A. S.; Allen, P. M.; Dustin, M. L. Science 1999, 285, 221-227. (d) Wagner, M. L.; Tamm, L. K. Biophys. J. 2001, 81, 266-275. (e) Cho, W. W.; Bittova, L.; Stahelin, R. V. Anal. Biochem. 2001, 296, 153-161. (3) (a) Watts, T. H.; Brian, A. A.; Kappler, J. W.; Marrack, P.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 7564-7568. (b) Watts, T. H.; Gaub, H. E.; McConnell, H. M. Nature 1986, 320, 179-181. (4) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 25542559. (5) Mou, J. X.; Yang, J.; Shao, Z. F. J. Mol. Biol. 1995, 248, 507-512. (6) (a) Williams, L. M.; Evans, S. D.; Flynn, T. M.; Marsh, A.; Knowles, P. F.; Bushby, R. J.; Boden, N. Supramol. Sci. 1997, 4, 513-517. (b) Cheng, Y. L.; Boden, N.; Bushby, R. J.; Clarkson, S.; Evans, S. D.; Knowles, P. F.; Marsh, A.; Miles, R. E. Langmuir 1998, 14, 839-844. (7) Jenkins, A. T. A.; Boden, N.; Bushby, R. J.; Evans, S. D.; Knowles, P. F.; Miles, R. E.; Ogier, S. D.; Scho¨nherr, H.; Vancso, G. J. J. Am. Chem. Soc. 1999, 121, 5274-5280. (8) See e.g.: (a) Ringsdorf, H.; Schlarb, B.; Venzmer, J. Angew. Chem., Int. Ed. Engl. 1988, 27, 113-158. (b) Beyer, D.; Elender, G.; Knoll, W.; Kuhner, M.; Maus, S.; Ringsdorf, H.; Sackmann, E. Angew. Chem., Int. Ed. 1996, 35, 1682-1685. (c) Heibel, C.; Maus, S.; Knoll, W.; Ru¨he, J. ACS Symp. Ser. 1998, 695, 104-118. (9) van Oudenaarden, A.; Boxer, S. G. Science 1999, 285, 10461048.
utilized.10 Different approaches to obtain patterned bilayers have been recently described, including photopolymerization,11 mechanical manipulation,12 or the use of prepatterned supports.13 Similarly, the immobilization of intact vesicles has been pursued for applications in biotechnology14-17 and, for instance, for the development of chemosensors.18 The approaches by the groups of Boxer14 and Ho¨o¨k15 rely on the interaction of complementary DNA fragments that are exposed on the surface and the vesicle membrane, respectively. Jung et al. have utilized streptavidin-biotin interactions for vesicle immobilization.16 The fabricated architectures possess the advantage that the underlying substrate interferes only marginally or not at all with the membrane properties of the immobilized vesicles.19 We have been involved in interfacial studies of unilamellar lipid vesicles interacting with various surfaces in the vesicle fusion process,20 patterned bilayer architectures,7,21 and the measurement of interactions of proteins with lipid bilayer membranes by means of atomic force microscopy (AFM). In an extension of these studies, (10) For recent reviews, see: (a) Boxer, S. G. Curr. Opin. Chem. Biol. 2000, 4, 704-709. (b) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149-157. (11) Morigaki, K.; Baumgart, T.; Offenhausser, A.; Knoll, W. Angew. Chem., Int. Ed. 2001, 40, 172-174. (12) (a) Cremer, P. S.; Groves, J. T.; Kung, L. A.; Boxer, S. G. Langmuir 1999, 15, 3893-3896. (b) Hovis, J. S.; Boxer, S. G. Langmuir 2000, 16, 894-897. (c) Hovis, J. S.; Boxer, S. G. Langmuir 2001, 17, 3400-3405. (13) (a) Groves, J. T.; Ulman, N.; Boxer, S. G. Science 1997, 275, 651-653. (b) Jenkins, A. T. A.; Bushby, R. J.; Evans, S. D.; Knoll, W.; Offenhausser, A.; Ogier, S. D. Langmuir 2002, 18, 3176-3180. (c) Cremer, P. S.; Yang, T. J. Am. Chem. Soc. 1999, 121, 8130-8131. (14) Yoshina-Ishii, C.; Boxer, S. G. J. Am. Chem. Soc. 2003, 125, 3696-3697. (15) Svedhem, S.; Pfeiffer, I.; Larsson, C.; Wingren, C.; Borrebaeck, C.; Hook, F. ChemBioChem 2003, 4, 339-343. (16) Jung, L. S.; Shumaker-Parry, J. S.; Campbell, C. T.; Yee, S. S.; Gelb, M. H. J. Am. Chem. Soc. 2000, 122, 4177-4184. (17) Stamou, D.; Duschl, C.; Delamarche, E.; Vogel, H. Angew. Chem., Int. Ed. 2003, 42, 5580-5583. (18) Kim, J.-M.; Ji, E.-K.; Woo, S. M.; Lee, H. W.; Ahn, D. J. Adv. Mater. 2003, 15, 1118-1121. (19) (a) Boukobza, E.; Sonnenfeld, A.; Haran, G. J. Phys. Chem. B 2001, 105, 12165-12170. (b) Stanish, I.; Santos, J. P.; Singh, A. J. Am. Chem. Soc. 2001, 123, 1008-1009. (20) Scho¨nherr, H.; Johnson, J. M.; Lenz, P.; Frank, C. W.; Boxer, S. G. Langmuir, submitted. (21) Morigaki, K.; Scho¨nherr, H.; Frank, C. W.; Knoll, W. Langmuir 2003, 19, 6994-7002.
10.1021/la0498915 CCC: $27.50 © 2004 American Chemical Society Published on Web 07/14/2004
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we discovered a simple yet efficient way to direct vesicle adsorption to preformed substrate-supported lipid bilayers. The approach was refined to serve as a means to fabricate bilayer-based platforms that serve as a generic interface between artificial surfaces, such as sensor surfaces, and biologically relevant molecules embedded in a near-natural environment. As shown in this communication, the adsorption of vesicles on lipid bilayers can be spatially controlled and directed in situ, in principle, with nanometer scale precision using an AFM-based approach. This strategy enables us to fabricate patterned vesicle arrays without the need to implement molecular recognition units in the vesicles and hence is applicable to a broad range of systems. Experimental Section Vesicle Preparation. The lipid (1.2-dimyristoyl-sn-glycero3-phosphatidylcholine, DMPC) was obtained from Avanti Polar Lipids, Alabaster AL. DMPC was suspended by vortexing in 0.1 M NaCl at a concentration of 2 mg/mL. After hydrating for 60 min, the vesicles were extruded through a polycarbonate membrane with 50 nm diameter pores using a Lipsofast basic extruder (Glen Creston Ltd., Stanmore, Middlesex, U.K.). After the liposomes had been passed 11 times through the extruder, they were diluted to 0.4 mg/mL and were ready for use. Substrate Preparation. All glassware used to prepare monolayers was immersed in piranha solution (70% concentrated sulfuric acid and 30% hydrogen peroxide) for 15 min and rinsed with large amounts of high-purity water (Millipore Milli-Q water). Caution: Piranha solution should be handled with extreme caution; it has been reported to detonate unexpectedly. Gold substrates used in the experiments were obtained from Metallhandel Schro¨er GmbH (Lienen, Germany). The substrates were cleaned immediately before use by oxygen plasma (5 min, 30 mA, 60 mTorr) (SPI Supplies, Plasma Prep II), followed by soaking in ethanol. The freshly cleaned gold substrates were immersed with minimal delay into a 1.0 mM solution of 2-mercaptoethanol (Aldrich, C2OH) in ethanol. After >12 h of assembly time, the substrates were removed from the solutions and rinsed extensively with ethanol to remove any physisorbed material. Bilayers were prepared in situ in the AFM liquid cell as described in Results and Discussion. Atomic Force Microscopy. The AFM experiments were carried out using a NanoScope II (Digital Instruments (DI), Santa Barbara, CA) in contact mode (CM). V-Shaped Si3N4 cantilevers were used (Nanoprobes, DI), with kc ) 0.30 N/m calibrated by the reference lever method22 and tip radii of 20-40 nm (estimated from images of a calibration standard; Silicon-MDT, Moscow, Russia). Measurements were performed in 0.1 M NaCl using a DI liquid cell. The cell was carefully cleaned by rinsing with chloroform and ethanol; further, all components of the tubing system and O-rings were cleaned by repeated sonication in ethanol followed by exhaustive flushing with Milli-Q water and subsequent drying in a stream of high-purity argon. For all experiments, poly(dimethylsiloxane)-based rubber O-rings were used to allow for frequent exchange of larger volumes (a few milliliters) of the liquid medium. To avoid problems caused by relaxation of sheared pressure-sensitive adhesive, glass substrates were glued with epoxy to the sample holder disk. After an initial equilibration period that ensured minimized thermal and instrumental drift, the AFM experiments were carried out with controlled forces. Typical scan rates of 3-5 Hz were used to acquire the data. All the images shown were subjected to a first-order plane-fitting procedure to compensate for sample tilt and, if necessary, to a zeroth-order flattening.
Results and Discussion DMPC bilayers were prepared inside the AFM liquid cell on granular gold substrates modified with a selfassembled monolayer (SAM) of C2OH by exposing the substrates to aqueous NaCl solutions of DMPC vesicles. (22) Tortonese, M.; Kirk, M. Proc. SPIE 1997, 3009, 53-60.
Figure 1. Contact mode AFM height image of DMPC vesicles adsorbed on a C2OH SAM (top) and AFM height image of the neat C2OH SAM on granular gold (bottom). The AFM data were acquired in 0.1 M aqueous NaCl solution at minimized forces (∼50 pN).
As shown in Figure 1 (top), the presence of nearly round elevated features was observed on top of the SAM after injection of vesicles. These features are attributed to adsorbed vesicles, since they possess typical heights of 10-30 nm, thus significantly exceeding the corrugations observed on the gold substrate (4-8 nm). The root-meansquare (rms) roughness of the granular gold was 1.5 ( 0.4 nm (scan size 25 µm2). Owing to the low imaging forces (e75 pN) utilized to ensure nonevasive imaging conditions,20 the tip frequently did not follow the vesicles accurately once the top of the vesicle was traversed. The corresponding horizontal profile hence showed a halo in the fast scanning direction.23 These observations are consistent with earlier reports on adsorbed vesicles studied with AFM approaches.20,24 At low concentrations, the vesicles were found to adsorb intact on the SAM, while at higher concentrations and longer interaction times, lipid bilayers were formed. The morphology of the lipid bilayers was indistinguishable from the morphology of the underlying SAM-modified gold substrate; the rms roughness was found to be the same to within the experimental error. The presence of the bilayers could be inferred from the step height measured at intentionally introduced scratches, where the underlying SAM is exposed (see the Supporting Information). We measured 4.5 ( 0.5 nm, which is in good agreement with previous reports for DMPC bilayer thicknesses as measured by intermittent contact mode AFM,25 when possible compression of the bilayer by the imaging force is considered.26 The data also agree with data from X-ray studies.27 (23) The halo appeared always in the horizontal direction, i.e., along the fast scan axis, independent of the scan angle utilized, while the imaged features rotated consistently. (24) (a) Egawa, H.; Furusawa, K. Langmuir 1999, 15, 1660-1666. (b) Reviakine, I.; Brisson, A. Langmuir 2000, 16, 1806-1815. (c) Kumar, S.; Hoh, J. H. Langmuir 2000, 16, 9936-9940. (d) Pignataro, B.; Steinem, C.; Galla, H. J.; Fuchs, H.; Janshoff, A. Biophys. J. 2000, 78, 487-498. (25) A thickness of 4.2 nm was reported: Tokumasu, F.; Jin, A. J.; Feigenson, G. W.; Dvorak, J. A. Ultramicroscopy 2003, 97, 217-227. (26) See e.g.: Dufrene, Y. F.; Barger, W. R.; Green, J. B. D.; Lee, G. U. Langmuir 1997, 13, 4779-4784. (27) A value of ∼5.3 nm has been reported depending on temperature and hydration level: (a) Smith, G. S.; Sirotra, E. B.; Safinya, C. R.; Plano, R. J.; Clark, N. A. J. Chem. Phys. 1990, 92, 4519-4529. (b) Harroun, T. A.; Heller, W. T.; Weiss, T. M.; Yang, L.; Huang, H. W. Biophys. J. 1999, 76, 937-945.
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Scho¨ nherr et al. Scheme 1. Schematic of the AFM-Tip-Assisted Immobilization Drawn Approximately to Scalea
a Bilayer thickness, ∼4 nm; tip radius, 20 nm; vesicle diameter, ∼40 nm. (A) An intact defect-free DMPC bilayer is formed on a C2OH SAM on gold (the SAM is omitted from the schematic); (B) subsequent scanning with an AFM tip at high forces leads to local damage of the bilayer; (C) in these areas (the schematic drawing does not imply any molecular detail concerning the damage created in step B), vesicles will adsorb from the solution and stay immobilized.
Scanning the DMPC bilayers on C2OH SAMs in the presence of DMPC vesicles with forces below a threshold force of ∼80-120 nN could be utilized to guide the immobilization of vesicles from solution to the SAMsupported bilayer.28 In a “writing step”, two lines comprising an angle of 60° were scanned subsequently using normal forces of 30-50 nN. Each line was scanned repeatedly, before the imaging force was minimized to 10 nN (A) and
