Stability of Lipid Films Formed on γ-Aminopropyl Monolayers

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Stability of Lipid Films Formed on γ-Aminopropyl Monolayers Todd W. McBee and S. Scott Saavedra* Department of Chemistry, University of Arizona, Tucson, Arizona 85721 Received September 21, 2004. In Final Form: December 15, 2004 The stability of supported lipid membranes (SLMs) deposited on planar substrates derivatized with (γ-aminopropyl)silane (GAPS) was examined. Ellipsometry, fluorescence microscopy, and atomic force microscopy were used to characterize SLMs exposed to repeated drying and rehydration. Vesicle fusion on GAPS-coated substrates produced SLMs with a thickness significantly greater than that of a single lipid bilayer. Exposure to even one cycle of drying/rehydration significantly decreased the thickness of a SLM on GAPS, and repeated drying/rehydration resulted in near quantitative lipid desorption. Thus SLMs on GAPS do not appear to be significantly more stable than the single bilayer SLM that is formed on bare glass or SiO2 under equivalent conditions.

Introduction Supported lipid membranes (SLMs) have been widely implemented as cell membrane mimics in molecular devices and bioassay systems (e.g., refs 1-4). The welldefined and controllable architecture of a SLM allows for favorable orientation and minimal denaturation of tethered water-soluble proteins,5-9 as well as a biocompatible, two-dimensionally fluid environment for reconstitution of transmembrane receptors with retained bioactivity (e.g., refs 1, 3, and 10-14). Furthermore, the phosphorylcholine (PC) lipid headgroup is highly resistant to protein adsorption;15,16 thus PC-capped SLMs minimize the undesirable adsorption of nontarget protein molecules that are invariably present in biological matrixes. Recent developments in micropatterning and array deposition techniques illustrate the potential of SLM-based biochips * To whom correspondence should be addressed: phone, (520) 621-9761; fax, (520) 621-8407; e-mail, [email protected]. (1) Cornell, B. A.; Braach-Maksvytis, V.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wieczorek, L.; Pace, R. J. Nature 1997, 387, 580-583. (2) Song, X. D.; Nolan, J.; Swanson, B. I. J. Am. Chem. Soc. 1998, 120, 4873-4874. (3) Schmidt, C.; Mayer, M.; Vogel, H. Angew. Chem., Int. Ed. 2000, 39, 3137-3140. (4) Stelzle, M.; Weissmuller, G.; Sackmann, E. J. Phys. Chem. 1993, 97, 2974-2981. (5) Edmiston, P. L.; Saavedra, S. S. Biophys. J. 1998, 74, 999-1006. (6) Edmiston, P. L.; Saavedra, S. S. J. Am. Chem. Soc. 1998, 120, 1665-1671. (7) Fischer, B.; Heyn, S. P.; Egger, M.; Gaub, H. E. Langmuir 1993, 9, 136-140. (8) Duschl, C.; Se`vin-Landais, A.-F.; Vogel, H. Biophys. J. 1996, 70, 1985-1995. (9) Yeung, C.; Purves, T.; Kloss, A. A.; Kuhl, T. L.; Sligar, S.; Leckband, D. Langmuir 1999, 15, 6829-6836. (10) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 14773-14781. (11) Graneli, A.; Rydstrom, J.; Kasemo, B.; Hook, F. Langmuir 2003, 19, 842-850. (12) Naumann, R.; Schmidt, E. K.; Jonczyk, A.; Fendler, K.; Kadenbach, B.; Liebermann, T.; Offenha¨usser, A.; Knoll, W. Biosens. Bioelectron. 1999, 14, 651-662. (13) 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. (14) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. Langmuir 1998, 14, 2467-2475. (15) Ross, E.; Rozanski, L.; Spratt, T.; Liu, S.; O’Brien, D. F.; Saavedra, S. S. Langmuir 2003, 19, 1766-1774. (16) Glasma¨star, K.; Larsson, C.; Ho¨o¨k, F.; Kasemo, B. J. Colloid Interface Sci. 2002, 246, 40-47.

with parallel arrays of sensing elements for high throughput biological or pharmaceutical screening.17-23 A key problem associated with using SLMs composed of natural lipids in molecular devices is their instability.24,25 The noncovalent forces responsible for self-organization of the lipid lamellar phase are insufficient to maintain the bilayer structure when, for example, it is exposed to surfactants, organic solvents, or transfer from water into air. Enhanced stability would be advantageous for further development of SLM-based technologies. Interest in developing stabilized SLMs has consequently received considerable scientific attention. Strategies include covalent tethering of lipids to the underlying support, linear and cross-linking lipid polymerization, the use of bolaamphiphiles to eliminate bilayer delamination, and biospecifically adsorbed protein coatings.15,24,26-32 Recently, Fang et al.18 reported that both fluid- and gel-phase SLMs deposited on glass substrates derivatized with (γ-aminopropyl)silane could be repeatedly dried and rehydrated without apparent loss of lipid. Furthermore, they showed that G protein-coupled receptors could be reconstituted in arrays of these SLMs with retention of (17) Morigaki, K.; Kiyosue, K.; Taguchi, T. Langmuir 2004, 20, 77297735. (18) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124, 2394-2395. (19) Fang, Y.; Frutos, A. G.; Lahiri, J. Chembiochem 2002, 3, 987991. (20) Hovis, J. S.; Boxer, S. B. Langmuir 2001, 17, 3400-3405. (21) Cremer, P. S.; Groves, J. T.; Kung, L. A.; Boxer, S. G. Langmuir 1999, 15, 3893-3896. (22) Kam, L.; Boxer, S. G. J. Am. Chem. Soc. 2000, 122, 1290112902. (23) Groves, J. T.; Mahal, L. K.; Bertozzi, C. R. Langmuir 2001, 17, 5129-5133. (24) Ross, E.; Rozanski, L.; Spratt, T.; Liu, S.; O’Brien, D. F.; Saavedra, S. S. Langmuir 2003, 19, 1752-1765. (25) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 25542559. (26) Kim, J. M.; Patwardhan, A.; Bott, A.; Thompson, D. H. Biochim. Biophys. Acta, Biomembr. 2003, 1617, 10-21. (27) Halter, M.; Nogata, Y.; Dannenberger, O.; Sasaki, T.; Vogel, V. Langmuir 2004, 20, 2416-2423. (28) Holden, M. A.; Jung, S. Y.; Yang, T. L.; Castellana, E. T.; Cremer, P. S. J. Am. Chem. Soc. 2004, 126, 6512-6513. (29) Yang, Z.; Yu, H. Langmuir 1999, 15, 1731. (30) Regen, S. L.; Kirszensztejn, P.; Singh, A. Macromolecules 1983, 16, 335-338. (31) Orban, J. M.; Faucher, K. M.; Dluhy, R. A.; Chaikof, E. L. Macromolecules 2000, 33, 4205-4212. (32) Hayward, J. A.; Chapman, D. Biomaterials 1984, 5, 135-142.

10.1021/la047646f CCC: $30.25 © 2005 American Chemical Society Published on Web 03/11/2005

Lipid Film Stability

nativelike ligand binding activity. Presented here are studies performed on GAPS-supported SLMs to further characterize their structure and stability to drying and rehydration. Experimental Section Materials and Substrates. γ-(Aminopropyl)triethoxysilane (GAPS) was purchased from Aldrich and used without further purification. Silicon wafers (1,1,1; Wacker) and glass microscope slides (Gold Seal) were cleaned using a mixture of 70:30 concentrated H2SO4/H2O2 for 20-30 min, followed by a thorough rinsing with deionized water (18 MΩ‚cm, Barnstead Nanopure) and drying at 105 °C for approximately 30 min. For Si wafers, this treatment generated a native oxide layer about 20 Å thick.33 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), and 1,2-dipalmitoylsn-glycero-3-phosphocholine (DPPC) in chloroform were purchased from Avanti Polar Lipids. N-(6-Tetramethylrhodaminethiocarbamoyl)-1,2-dihexadecanoyl-sn-glycero-3- phosphoethanolamine, triethylammonium salt (TMR-DHPE), Texas Red 1,2dihexadecanoyl-sn-glycero-3- phosphoethanolamine, triethylammonium salt (TR-DHPE), and 5-carboxyfluorescein (CF) were purchased from Molecular Probes. Sephadex G-50 (50-150 µm) was purchased from Aldrich. GAPS-coated microscope slides manufactured by Corning (part no. Z713856, denoted here as GAPS-II) were purchased from Sigma and used as received. For comparison to GAPS-II, GAPS coatings were formed on silicon wafers and glass microscope slides (hereby denoted as GAPS-I). Clean substrates were incubated in 1% GAPS/99% dry toluene solution for 1 h at room temperature and sonicated twice in 100% methanol for 10 min, followed by rinsing in ethanol and drying under a nitrogen stream. Lipid films were deposited on GAPS-I-coated substrates within 60 min after their preparation. Formation of Lipid Films. Supported lipid membranes (SLMs) were prepared by fusion of small unilamellar lipid vesicles (SUVs) composed of either DOPC or a mixture of DPPC and DMPC (4:1, mol/mol). To perform fluorescence experiments on SLMs, the SUVs also contained either TMR-DHPE or TR-DHPE (0.5 mol %). In all cases, lipids were dried from stock chloroform solutions under flowing Ar, followed by drying 4-5 h in a vacuum. The lipids were then resuspended in deionized water to a final concentration of 0.5 mg/mL and then vortexed and sonicated to clarity in a Branson sonicator fitted with a cup horn (Sonifier 450). For those experiments involving entrapped CF, the SUVs were formed in a buffer (20 mM phosphate buffer, pH 7.36) containing 2 mM CF. To form SLMs, a few drops (approximately 100 µL) of SUV solution were deposited on the substrate. Lipids were fused at room temperature (approximately 22-25 °C) for 30 min and maintained under water thereafter, except where noted. In a number of experiments, the SLM was removed from water to assess its stability in air. In these cases, after fusion the SLM was kept hydrated by submersion into a large volume (∼100 mL) of deionized water. To dehydrate it, the SLM was slowly withdrawn across the air/water interface and allowed to dry at room temperature (as indicated by an absence of visually observable liquid). In addition to experiments on extended SLMs, experiments were also performed on arrays of SLM spots. The arrays were formed by placing 10 µL of SUV solution, dispensed from a syringe, onto 10 spatially disparate regions of each microscope slide. The spot diameter was 1.5-2.5 mm. In some cases (noted below), 20 mM phosphate buffer, pH 7.36, was used in place of deionized water for the entire experiment (i.e., from resuspension of dried lipids through multiple drying/rehydration cycles). Characterization. Ellipsometric measurements of organic film thicknesses deposited on oxidized Si wafers were made with a Sentech model SE 400 ellipsometer, using the 632.8 nm line of a HeNe laser at a 70° incident angle. The elliptical spot size was approximately 1.5 mm × 4 mm. In all cases, the thickness of the oxide layer and the GAPS layer formed on the oxide layer (33) Du, Y.-Z.; Wood, L. L.; Saavedra, S. S. Mater. Sci. Eng. C 2000, 7, 161-169.

Langmuir, Vol. 21, No. 8, 2005 3397 was measured before the deposition of the SLM. A uniform refractive index of 1.46 was assumed for all native oxide, GAPS, and lipid layers. Static water contact angles were measured with a Kruss model DSA 10 MK2 drop shape analysis system. Deionized water droplets (2 µL) were delivered with a motordriven syringe at multiple sites on each sample. Drop shapes were analyzed with the software Drop Shape Analysis (DSA) for Windows, version 1.70. Data are reported herein as the mean of measurements on at least three independently prepared samples for each film type, made at three different physical locations on each sample. Epifluorescence measurements were performed using Nikon Diaphot microscope with the fluorescence signal directed to a monochromator and detected using a photomultiplier tube.34 The surface morphology of dried films was examined using atomic force microscopy (AFM) performed in tapping mode on a Digital Instruments Dimension 3100 scanning probe microscopy system. The height and phase images were acquired simultaneously and represent the same 1 µm × 1 µm region. Oxidesharpened silicon nitride tips were used. To minimize distortion by contact forces between the tip and the sample, the minimum force required to obtain optimum resolution was used. The driving amplitude applied was the automated instrument default, between 100 and 400 mV. After the tip was engaged and imaging commenced, the setpoint value was increased until contact with the surface was lost and then incrementally decreased until optimal resolution was achieved. Setpoint values scaled with drive amplitude values and were generally between 0.2 and 1.2 V, which usually resulted in “softer” imaging than the default engagement settings. The images herein are representative of scans from different locations on the sample, different samples, and using different tips to image the surfaces.

Results and Discussion Si wafers coated with GAPS-I were characterized by ellipsometry, AFM, and contact angle measurements. The ellipsometric thickness was 8.0 ( 0.8 Å (n ) 13), consistent with the expected value of 9 Å for a GAPS monolayer based on known bond lengths, assuming an extended, alltrans conformation perpendicular from the surface. The static water contact angle for GAPS-I was 41 ( 1.3°, considerably greater than that of a freshly cleaned silicon wafer (12 ( 3.5°). The contact angle for GAPS-II is 47 ( 1.4°. The origin of this consistently higher value, relative to GAPS-I, is likely due to differences in the preparation methods for the two surfaces (information on preparation of GAPS-II slides is proprietary19), although differences in storage conditions may also play a role.35,36 AFM images of GAPS-I monolayers were featureless on the 1 × 1 µm scale (see Supporting Information), with a root mean square surface roughness of 0.18 nm, slightly greater than that of a clean Si wafer (0.11 nm).33 SLMs were formed from 4:1 DPPC/DMPC SUVs in deionized water on the surface of oxidized silicon wafers and wafers coated with GAPS-I. Ellipsometry was used to measure the thickness of these films as a function of the number of times that they were immersed and then withdrawn from deionized water. Room temperature is very close to the phase transition temperature of DMPC (23 °C) and well below that of DPPC (41 °C); thus this mixture is predominately in the gel phase. Figure 1 shows the results of these measurements. The initial thickness (dried once) of the films on oxidized Si wafers was 41 ( 8.7 Å, which is approximately that expected for one bilayer (45-55 Å),24 but repeated passage across the air/water interface caused near quantitative desorption. (34) Phimphivong, S.; Kolchens, S.; Edmiston, P. L.; Saavedra, S. S. Anal. Chim. Acta 1995, 307, 403-417. (35) Petri, D. F. S.; Wenz, G.; Schunk, P.; Schimmel, T. Langmuir 1999, 15, 4520-4523. (36) Flink, S.; van Veggel, F.; Reinhoudt, D. N. J. Phys. Org. Chem. 2001, 14, 407-415.

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Figure 1. Ellipsometric thickness of dried SLMs measured as a function of the number of times the SLM was immersed and then withdrawn from water. The combinations of SLM composition and substrate were 4:1 DPPC/DMPC and GAPS-Icoated Si wafer (diagonal lines), 4:1 DPPC/DMPC and oxidized Si wafer (vertical lines), DOPC and GAPS-I-coated Si wafer (filled), and DOPC and oxidized Si wafer (unfilled). The error bars represent three measurements on independently prepared samples. In those cases where data are not plotted, a lipid film could not be detected.

Figure 2. Representative height AFM image and line scan of a lipid film formed on an oxidized Si wafer coated with GAPS-I. The 1 µm × 1 µm image was acquired after the film was dried once. The ellipsometric thickness of the film is 160 ( 25 Å, and the root mean square roughness of the surface is 2.76 nm. The step heights in this image are approximately 5 nm (the step marked by the arrows has a height of 4.5 nm), which corresponds to the height of a single lipid bilayer. The film is clearly multilamellar. The corresponding phase image is provided in Supporting Information.

DPPC/DMPC films formed on GAPS-I were much thicker. The initial thickness (dried once) was 190 ( 47 Å (Figure 1). Clearly the lipids form a multilayer structure, the surface of which is topographically very rough as shown in the AFM image displayed in Figure 2, as well as in the corresponding phase image in Supporting Information. However, similar to that observed for the films on Si wafers, after five to six cycles of rehydration/drying, the film thickness dropped to well below that expected for one bilayer.

McBee and Saavedra

Figure 3. Fluorescence intensities measured on hydrated SLMs composed of 4:1 DPPC/DMPC, doped with 0.5% of a fluorescent lipid, as a function of the number of times that the SLM-coated substrate was withdrawn and then reimmersed in deionized water. The data were normalized to the initial intensity measured after the SLM was rinsed to remove unfused vesicles, without exposing it to air. The combinations of fluorescent lipid and substrate were TMR-DHPE and GAPS-I (diagonal lines); TR-DHPE and GAPS-I (vertical lines); TRDHPE and GAPS-II (filled); and TMR-DHPE and GAPS-II (unfilled). The error bars represent three measurements on independently prepared samples. In those cases where data are not plotted, the residual fluorescence was not detectable above the background signal. After four rinses, residual fluorescence could not be detected above the background signal for any of the four combinations.

Analogous experiments were also performed using DOPC, a fluid-phase lipid; the results are also shown in Figure 1. On oxidized Si wafers, a thickness less than that of one bilayer was observed after drying the film once, consistent with prior studies showing that fluidphase lipid bilayers desorb from SiO2 and bare glass when the substrate is withdrawn from water.24,25 On substrates coated with GAPS-I, the thickness after drying once was 94 ( 5.2 Å, approximately twice that expected for a bilayer structure; however near quantitative desorption was observed after three cycles. In summary, fusion of both gel-phase and fluid-phase lipids on the GAPS-I coating results in formation of a multilayer lipid film. Repeated drying/rehydration of these SLMs results in near quantitative lipid desorption, which is inconsistent with the results reported by Fang et al.18 On the basis of measurements of relative fluorescence intensity of TR-DHPE doped into both gel-phase and fluid-phase SLMs, they reported that lipid films coated on GAPS-II slides are stable to repeated drying and reimmersion in buffer. The discrepancy between their results and those in Figure 1 may be attributable to differences in substrate surface chemistry (i.e., GAPS-I vs GAPS-II) and/or the analytical method. To assess these possibilities, epifluorescence measurements were performed on SLMs deposited on GAPS-II and GAPS-I glass slides. SLMs were doped with either TR-DHPE or TMR-DHPE. Retention of fluorescence intensity was measured as a function of the number of times that the SLM-coated substrates were withdrawn and then reimmersed in deionized water. Results from experiments on 4:1 DMPC/DPPC are shown in Figure 3. The decline of intensity shows that repeated drying and rehydration causes near quantitative desorption of the fluorescent lipids from both GAPS-II and GAPS-I, which is consistent with the ellipsometry data (Figure 1). Thus differences in the method of preparation and composition of the GAPS-II and GAPS-I coatings did not measurably affect SLM stability. Analogous experiments were performed on DOPC SLMs and on DMPC:DPPC SLMs that were fused, rinsed, and

Lipid Film Stability

Figure 4. Arrays of SLMs composed of 4:1 DPPC/DMPC and doped with 0.5% TR-DHPE were deposited on GAPS-II slides. Fluorescence intensities were measured on hydrated arrays as a function of the number of times that the array-coated substrate was withdrawn and then reimmersed in deionized water. The data were normalized to the initial intensity measured on spots that were rinsed to remove unfused vesicles, without exposing them to air. The error bars represent three measurements on independently prepared samples.

reimmersed in 20 mM phosphate buffer, pH 7.36 (rather than deionized water). In all cases, the results were similar to those shown in Figure 3 (see Supporting Information). Epifluorescence measurements were also performed on GAPS-II slides coated with arrays of SLM spots having a diameter of 1.5-2.5 mm. Results obtained for spots composed of 4:1 DMPC/DPPC doped with TR-DHPE are plotted in Figure 4. The fluorescence intensity declined to 6 ( 6% after four drying/reimmersion cycles, consistent with the data plotted in Figure 3 for extended SLMs. Finally, experiments were performed to assess if the multilayer SLMs formed on GAPS-II contained intact (unruptured) vesicles. Vesicles composed of 4:1 DPPC/

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DMPC with entrapped CF37 were prepared by the methods outlined above and separated from unentrapped CF on a Sephadex G-50 column (equilibrated with 20 mM phosphate buffer, pH 7.36). SLMs were formed by fusion onto GAPS-II slides and retention of fluorescence intensity was measured as described above. The results (see Supporting Information) were similar to those obtained when using fluorescent lipid probes. A significant decline of the initially strong emission intensity was observed, showing that (a) intact vesicles were present in the SLM, and (b) that repeated drying and rehydration caused the vesicles to rupture, resulting in loss of CF from the SLM. In conclusion, fusion of SUVs on GAPS-coated substrates produces SLMs with a thickness significantly greater than that of a single bilayer. The mechanism of multilayer formation is not clear. Exposure to repeated cycles of drying/rehydration results in near quantitative lipid desorption, regardless of whether the substrate is coated with GAPS-II or GAPS-I. Thus the reported stability18 of SLMs on GAPS appears to be a consequence of their multilayer structure. Acknowledgment. This work was supported in part by the National Science Foundation under Grant Number CHE-0108805 and in part by Grant Number DE-FG0302ER15378 from Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Research, U.S. Department of Energy. Supporting Information Available: Additional AFM images of bare and SLM-coated GAPS-I substrates, additional data on the stability of SLMs on GAPS-I and GAPS-II substrates to drying and rehydration, as measure by retention of TR-DHPE fluorescence. This material is available free of charge via the Internet at http://pubs.acs.org. LA047646F (37) Johnson, J. M.; Ha, T.; Chu, S.; Boxer, S. G. Biophys. J. 2002, 83, 3371-3379.