Preparation of Solid-Supported Lipid Bilayers by Spin-Coating

Page 1. Preparation of Solid-Supported Lipid Bilayers by. Spin-Coating. Ulrike Mennicke and Tim Salditt*. Universita¨t des Saarlandes, Im Stadtwald 3...
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Preparation of Solid-Supported Lipid Bilayers by Spin-Coating Ulrike Mennicke and Tim Salditt* Universita¨ t des Saarlandes, Im Stadtwald 38, Postfach 15 11 50, 66041 Saarbru¨ cken, Germany Received April 25, 2002. In Final Form: July 5, 2002 A new method to deposit lipid bilayers on solid substrates by spin-coating of lipid solutions is presented, leading to highly aligned and homogeneous bilayers probably nucleating at the interface. Importantly, the total number of bilayers can be controlled by the deposition parameters. The samples have been characterized by X-ray reflectivity, both at partial and at full hydration in the presence of excess water.

Introduction Solid-supported lipid bilayers are used as biomimetic systems to study the structure as well as various biochemical and biophysical aspects of the interaction between membranes and membrane-active biomolecules under well-controlled conditions.1 Apart from the predominant role that lipid bilayers play in the organization of the biological cells, they are also expected to become a key component of novel biomolecular materials. In particular, solid-supported lipid bilayers may provide a way to biofunctionalize solid-state and semiconductor surfaces, providing a compatible interface between the inorganic and the biomolecular world. The preparation and characterization of solid-supported bilayers is therefore of great importance. To date, several alternative routes to the preparation are known: (a) vesicle fusion,2,3 (b) Langmuir-Blodgett and Langmuir-Schaeffer,4-6 and (c) spreading from organic solution.7 The latter technique is attractive for structural investigations by scattering or NMR methods because it allows for the deposition of multilamellar oriented bilayers by spreading from a lipid solution in organic solvent. Compared to single-supported bilayers, multilamellar arrangements may also present important advantages in some applications. If functional proteins are incorporated, the yield in sensing, catalytic, light harvesting, or drug delivery applications can in most cases be expected to scale linearly with the volume of bilayers. Just as mother nature does (e.g., in the thylakoid membrane of the plant chloroplast), multilamellar architectures may present an efficient packaging solution. At the same time, thermal stability and precise control both of the total number of layers (N) and of the orientational alignment with respect to the substrate is desirable. In highly oriented samples, the degree of membrane mosaicity achieved (i.e., orientational distribution with respect to the solid substrates) is typically better than 0.02°. The bilayer structure and fluctuations have been probed by interface-sensitive scattering methods such as X-ray reflectivity, diffuse * Corresponding author e-mail: [email protected]. (1) Sackmann, E. Science 1996, 271, 43-48. (2) Brian, A. A.; Mc Connell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81 6159-6163. (3) Kalb, E.; Frey, S.; Tamm, L. K. Biochim. Biophys. Acta 1992, 1103, 307-316. (4) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 104-113. (5) Fragneto, G.; Charitat, T.; Graner, F.; Mecke, K.; Perino-Gallice, L.; Bellet-Amalric, E. Europhys. Lett. 2001, 53, 100-106. (6) Merkel, R.; Sackmann, E.; Evans, E. J. Phys. Fr. 1989, 50, 1535. (7) Seul, M.; Sammon, M. J. Thin Solid Films 1990, 185, 287.

Figure 1. (A) Sketch illustrating the spin-coating procedure showing a pipet with lipid solution above the rotating substrate. (B) Sketch of a spin-coated stack of membranes with incoming and reflected beams in a reflectivity setup. Ri and Rf denote the angles between the sample surface and the incoming and reflected beam, respectively. qz denotes the momentum transfer in z-direction, and q|| marks the lateral momentum transfer.

(nonspecular) scattering, and grazing incidence diffraction8-11 (Figure 1). Importantly, experiments can be carried out at physiological conditions of hydration, temperature, and ionic strength. Typically, a few hundred to thousands of bilayers are prepared by this method, depending on the concentration of the lipid solution. However, it is impossible to prepare just a few bilayers with this method, let alone to achieve a well-controlled number of bilayers. Contrarily, in the Langmuir-Schaeffer technique where floating monolayers are repeatedly deposited on a substrate, N is known and controlled exactly. However, only up to two bilayers can be prepared. Similar restrictions in N apply to vesicle fusion,2,3 where the lipids are deposited by adsorption and fusion of unilamellar vesicles to substrates. In comparison to Langmuir-Schaeffer, vesicle fusion has the advantage that the lipids and other components are in a hydrated phase during the whole preparation process. Both techniques and also combinations of them have proven to be very useful for the preparation of thin samples such as single-supported membranes. However, they are not suitable for the preparation of N g 3 bilayers. (8) Salditt, T.; Mu¨nster, C.; Lu, J.; Vogel, M.; Fenzl, W.; Souvorov, A. Phys. Rev. E 1999, 60, 7285. (9) Vogel, M.; Mu¨nster, C.; Fenzl, W.; Salditt, T. Phys. Rev. Lett. 2000, 84, 390-393. (10) Mu¨nster, C.; et al. Europhys. Lett. 1999, 46, 486. (11) Lyatskaya, Y.; Liu, Y.; Tristram-Nagle, S.; Katsaras, J.; Nagle, J. Phys. Rev. E 2001, 63, 11907.

10.1021/la025863f CCC: $22.00 © 2002 American Chemical Society Published on Web 08/30/2002

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Table 1. List of Investigated Lipid and Peptide Systems lipids/peptide

solvent

etching

size, mm2

amount, µL

step 1,a rpm

step 2,b rpm

DMPC, DLPC, DOPC, OPPC DMPC DMPG, DPPG DMPE, DPPEc PE:PC, 1:1-1:10d PG:PC, 1:1-1:10d POPS DPPC, DOPC DMPC/alamethicin, 10:1-200:1d DLPC/alamethicin, 10:1-200:1d

2-propanol chloroform chloroform chloroform chloroform chloroform chloroform TFE 2-propanol 2-propanol

yes no no no no no no yes yes yes

15 × 25 15 × 25 15 × 25 15 × 25 15 × 25 15 × 25 15 × 25 15 × 25 2 in. L 15 × 25

100-200 100-200 100-200 100-200 100-200 100-200 100-200 100-200 500 100-200

500

3000 3000 3000 3000 3000 3000 3000 3000 3000 3000

500 1000 500

a Step 1: rotational frequency of first second. b Step 2: rotational frequency for further 30 s. c DMPE and DPPE solutions yielded multiple phase layers. d Mixtures are indicated by molar ratios.

Here we present a technique by which stacks of a small and well-controlled number of bilayers can be prepared on a substrate. N is essentially uniform over the area of the sample and can be varied from N ) 1, 2 up to j30. For this purpose, the lipids and other components of the membranes are being dissolved in an organic solvent and are pipetted onto a cleaned, hydrophilic or hydrophobic substrate that is accelerated to rotation using a spincoater. This procedure yields highly oriented homogeneous stacks of bilayers with a well-defined total film thickness. Spin-coating is very well-established in organic thin film deposition but has so far not been used for lipids. It is important to realize that the bilayers form only while or after the deposition (when in contact with water vapor), probably nucleating at the interfaces. X-ray and neutron reflectivity have been employed for a long time to investigate phospholipid membranes supported on solid surfaces and to deduce detailed structural parameters of the bilayers.12 Different N ) 1 bilayer systems with and without polymer cushions have been studied by neutron reflectometry.13,14 Beyond pure lipid bilayers, solid-supported membranes have further been used to study fundamental structural aspects of lipid-peptide interaction (see refs 15-18). The lipid bilayer samples prepared by the spin-coating method presented here exhibit a mosaic spread (variation of tilt angles) on the order of a typical instrumental resolution of about 0.01°, as measured in a rocking scan. In other words, the membranes are essentially “single crystalline” with respect to the layer orientation. This allows for the application of specular and nonspecular (diffuse) X-ray or neutron reflectivity as quantitative methods to investigate normal and lateral structural parameters of single or multiple interfaces.19,20 Note that the most important structural features such as bilayer periodicity (d) and N can be directly extracted from the data without modeling and data fitting (e.g., from the so-called Kiessig fringes). (12) Katsaras, J.; Raghunathan, V. A. Aligned Lipid-Water Systems. In Lipid bilayers: structure and interactions; Katsaras, J., Gutberlet, T., Eds.; Springer: Berlin, 2000. (13) Wong J. Y.; et al. Biophys. J. 1999, 77, 1445-1457. (14) Meuse, C. W.; et al. Biophys. J. 1998, 74, 1388-1398. (15) Ludtke, S. J.; He, K.; Wu, Y.; Huang, H. W. Biochim. Biophys. Acta 1994, 1190, 181-184. (16) Ludtke, S. J.; He, K.; Huang, H. W. Biochemistry 1995, 34, 16764-16769. (17) Ludtke, S. J.; He, K.; Heller, W. T.; Harroun, T. A.; et al. Biochemistry 1996, 35, 13723-13728. (18) Wu, Y.; He, K.; Ludtke, S. J.; Huang, H. W. Biophys. J. 1995, 68, 2361-2369. (19) Daillant, J., Gibaud, A., Eds. X-ray and neutron reflectivity: principles and applications; Lecture Notes in Physics 58; Springer: New York, 1999. (20) Tolan, M. X-ray scattering from soft-matter thin films; Tracts in Modern Physics 148; Springer: New York, 1999.

The structural details of the systems will be analyzed and presented elsewhere. Materials and Methods Lipids and Peptides. The lipids 1,2-dimyristoyl-sn-glycero3-phosphocholine (DMPC), 1,2-dilauroyl-sn-glycero-3-phosphocholine (DLPC), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2dimyristoyl-sn-glycero-3-phosphoethanolamine (DMPE), 1,2dimyristoyl-sn-glycero-3-[phospho-rac-(1-glycerol)] (DMPG), 1-oleoyl-2-palmitoyl-sn-glycero-3-phosphocholine (OPPC), and 1-palmitoyl-2-oleoyl-sn-glycero-3-[phospho-L-serine] (POPS) were bought from Avanti Lipids (Alabaster, AL), and the peptide alamethicin was bought from Sigma Aldrich. Both lipids and peptides were used without further purification. Poly(ethylene glycol) (PEG) 300 was bought from Fluka. Highly aligned oligo membranes were prepared by spin-coating whereby lipids and peptides are pipetted from a solution onto the rotating substrate. For this purpose all components (i.e., lipids, lipid-peptide, or lipid-lipid mixtures) must be co-dissolved in the desired ratio in an organic solvent. The solvent should simultaneously meet the requirements of solvation and wettability of the substrate. 2-Propanol is a suitable solvent for phosphatidylcholine such as DMPC and also for the antibiotic peptide alamethicin, while phosphatidylethanolamine and phosphatidylglycerol are best soluble in chloroform. The choice is mainly dependent on the solubility of the bilayer components. Optical inspection of the samples shows that the choice of the solvent and the associated wetting properties influence the homogeneity on the scale of 10100 µm. Chloroform solutions on hydrophobic silicon substrates, for example, yielded particularly uniform layers. Table 1 gives an overview about all lipid and lipid-peptide solutions examined in this work together with spin-coating parameters. For hydrophilic solvents such as 2-propanol or trifluoroethanol (TFE), the substrate surface has to be rendered hydrophilic in an etching process, while for hydrophobic solvents such as chloroform the surface must be hydrophobic and needs no further treatment after the cleaning. Substrate Cleaning. Samples were prepared on rectangular silicon or glass substrates cut from standard commercial silicon wafers (diameter 5 in. and thickness ≈0.5 mm) and standard glass object slides (as used for optical microscopy) to a size of about 15 × 25 mm2. In addition, larger round silicon wafers (diameter 2 in. and thickness 0.635 mm) were used for sample deposition. The substrates were carefully cleaned in an ultrasonic methanol bath for about 10 min and subsequently were rinsed in methanol and two bechers of ultrapure water (specific resistivity g18.2 MΩ cm, Millipore, Bedford, MA). Finally, the substrates were dried with a flow of nitrogen. Hydrophilic etching was achieved by the following three alternative procedures: (a) washing the substrates in a 5 M solution of KOH in water (for glass and quartz substrates) or in a saturated KOH solution in ethanol (for silicon substrates), respectively, for about 1 min. (KOH etching has to be followed by extensive rinsing in ultrapure water (Millipore).); (b) plasma etching in a plasma cleaner (Harrick Scientific, NY) for 30 s; or (c) in case of silicon substrates, ozone etching by irradiation from a ozone-producing UV lamp

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Figure 2. Open symbols: X-ray reflectivity measurements of four spin-coated DMPC samples prepared from lipid solutions of the concentrations 20, 10, 5, and 1 mg/mL (top to bottom) along with simulations based on semi-kinematic theory. The lowest curve (without simulation) was measured at room temperature ≈20 °C and ≈60% RH. All other curves were measured in LR phase at a temperature of 40.2 °C and 97-99% RH. Insert: Electron density profile models corresponding to the reflectivity simulations: dark gray, 5 mg/mL curve; gray, 10 mg/mL curve; and light gray, 20 mg/mL curve. (UVP) for 30 min. Each of these procedures results in a hydrophilic substrate surface. Typical values of 8-14 Å rms roughness were obtained. Spin-Coating. For the spin-coating process a certain amount of solution (for small samples of 15 × 25 mm2, 0.1 mL for concentrations of 1-15 mg/mL up to 0.2 mL for a concentration of 20 mg/mL) was carefully pipetted centrally onto the cleaned substrate (Figure 1A). The solution was spread over the surface, and immediately after the solution had wetted the whole substrate, the sample was accelerated to rotation. Typically the rotation is performed in two steps. While a first, slower rotation allows the solution to cover the surface homogeneously, the second step enforces quick drying and tossing of the spare solution (Table 1). For solutions with chloroform, it has proven useful to skip the first step and start the rotation with the final frequency of 3000 rpm immediately after the solution has been pipetted onto the substrate. After fast solvent evaporation, a well-defined number of bilayers nucleates at the substrate surface with water from the surrounding air. To remove all traces of organic solvent, the samples were exposed to high vacuum over 20 h. The samples were kept refrigerated until the measurement. Sample Environment. The films were hydrated in a closed temperature-controlled chamber with water vapor atmosphere. For measurements presented in this work, two different sample chambers were used. Both consist of a cylindrical stainless steel chamber with Kapton windows that can be cooled or heated by a flow of oil connected to a temperature-controlled reservoir (Julabo, Germany). The samples were mounted in an inner stainless steel chamber with a water reservoir to keep the relative humidity close to 100%. Temperature was measured at the inner chamber by a Pt100 sensor, with a stability of better than 0.03 K over several hours. For experiments with the membranes immersed in excess water, an additional chamber that can be filled with water21 was used inside the temperature-controlled chamber. X-ray Experiment. The X-ray reflectivity measurements presented here were carried out at the bending magnet beamline D4 of HASYLAB/DESY in Hamburg, Germany. At D4, a single Si(111) monochromator was used to select a photon energy of 19.92 keV after passing a Rh-mirror to reduce higher harmonics. (21) Vogel, M. Ph.D. Dissertation, Universita¨t Potsdam, 2000.

The chamber was mounted on the z-axis diffractometer, and the reflected beam was measured by a fast scintillation counter (Cyberstar, Oxford). Incident and exit beams were defined by a system of several motorized slits. To record a reflectivity curve, the incident beam with wave vector ki has to be collimated to less than a few hundredths of a degree and directed on the sample at glancing incidence angle Ri. The reflected intensity is then measured as a function of Ri under specular conditions (e.g., at an exit angle Rf ) Ri), with the wave vector of the exit beam denoted by kf. Thus, the momentum transfer of the elastic scattering q ) kf - ki is always along qz, with the z-axis parallel to the sample normal (Figure 1B). Typically, the reflectivity can be recorded over 7-8 orders of magnitude (after correction for diffuse scattering and background), as measured in a so-called offset scan. To get correct results when fitting the reflectivity, the correction for diffuse background at higher angles is essential.22Interface-sensitive scattering techniques applicable on highly oriented samples have been extensively used in studies of lipid monolayers or monomolecular films at the air-water interfaces23-27 and single solidsupported bilayers.5,28

Results and Discussion Figure 2 shows X-ray reflectivity measurements on four DMPC samples in the fluid LR phase (upper three curves), along with simulations based on the semi-kinematic theory22 and the corresponding electron density profiles (22) Salditt, T.; Li, C.; Spaar, A.; Mennicke, U. Eur. Phys. J. E 2002, 7, 105-116. (23) Als-Nielsen, J.; Mo¨hwald, H. In Handbook of Synchrotron Radiation; Ebashi, S., Rubinstein, E., Koch, K., Eds.; North Holland: Amsterdam, 1992; Vol. 4, p 3. (24) Helm, C. A.; Mo¨hwald, H.; Kjaer, K.; Als-Nielsen, J. Europhys. Lett. 1987, 4, 697-703. (25) Baltes, H.; Schwendler, M.; Helm, C. A.; Mo¨hwald, H. J. Colloid Interface Sci. 1996, 178, 135-143. (26) Kuhl, T. L.; Majewski, J.; Wong, J. Y.; Steinberg, S.; Leckband, D. E.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1998, 75, 2352-2362. (27) Schalke, M.; Kru¨ger, P.; Weygand, M.; Lo¨sche, M. Biochim. Biophys. Acta 2000, 1464, 113-126. (28) Gliss, C.; Clausen-Schaumann, H.; Gu¨nther, R.; Odenbach, S.; Randl, O.; Bayerl, T. M. Biophys. J. 1998, 74, 2443-2450.

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Table 2. Simulation Parameters for Reflectivity Curves in Figure 2 in Semi-Kinematic Theory As Described in Ref 24 DMPC5 DMPC10 DMPC20

σsuba

σtopa

d

d0b

N

∆maxc

f1

f2

f3

f4

f5

f6

10 14 11

2.1 2 2

50.2 53.9 50.4

18 20 18

9 10 22

0.09 0.08 0.08

1 1 1

-0.6 -0.7 -0.6

-0.4 -0.27 -0.38

-0.15 -0.01 -0.19

0.06 0.1 0.06

0.01 0 0.016

a σ sub and σtop denote the rms roughness of the substrate and the top of the membrane stack consisting of N membranes with periodicity d. b d0 is the distance between substrate and first membrane. c ∆max multiplied with the difference in electron density from substrate to air ∆12 ) -0.699 e-/Å3 denotes the amplitude of the corresponding electron density profile that is expressed with normalized dimensionless Fourier coefficients fi where f1 ≡ 1.

in the insert. The curves were measured at D4 (HASYLAB/ DESY) at a relative humidity (RH) of approximately 9799% and at a temperature of T ) 40.2 °C. All curves have been corrected for illumination, background, and storage ring current. They were shifted vertically for clarity. Each sample was prepared from a DMPC solution in 2-propanol at decreasing concentrations (20, 10, and 5 mg/mL, top to bottom). In the top curve (squares) six multilamellar Bragg peaks are observed, indicating a periodicity of d ) 50.8 ( 0.2 Å in good agreement with literature values for the thickness of partially hydrated DMPC bilayers with a water layer (≈97% RH in water vapor atmosphere).29 From the Bragg peak width, one can estimate the total lipid film thickness of the sample. Considering a simple multilamellar structure factor for N bilayers, s(qz, N) ) N ∑m)1 eiqzdm, one would expect a Bragg peak width of fwhm ∆qz = 5.57/Nd. Given the constraints of the resolution, we can deduce that N J 22 bilayers for this sample. The simulation and corresponding electron density profile Fe(z) shown in the insert (light gray) leads to a bilayer thickness of 34 Å and a water layer thickness of 16 Å. The second curve (triangles) shows Bragg peaks up to the fifth order. This sample was prepared from a stock solution of 10 mg/mL concentration. Note that the Bragg peaks have shifted to smaller qz values, which indicate a thicker repeat distance (53.9 ( 0.2 Å) and a higher hydration of the sample. This can also be deduced from the electron density profile in the insert (gray), giving a water layer thickness of 21 Å for this sample whereas the bilayer thickness is unchanged. The fourth Bragg peak has almost disappeared, which indicates a form factor minimum. From the intensity oscillations between the Bragg peaks (Kiessig fringes), one can determine the total thickness (L) of the DMPC layer on the substrate L ) 2π/∆qz with ∆qz being the distance between two minima. The observation of the Kiessig fringes indicates a well-defined film thickness and thus a well-defined number (N ) 10) of bilayers. In the top curve only very weak Kiessig fringes are observed between the Bragg peaks since they are at the limit of the instrumental resolution. The next curve below (circles) shows similar features to the second curve with a repeat distance of d ) 50.2 ( 0.2 Å. The fringes exhibit eight minima between two neighboring Bragg peaks, which indicate a total number of nine bilayers on the sample. This sample was prepared from a stock solution of 5 mg of DMPC/1 mL of 2-propanol. The lowest curve (stars) shows the reflectivity of a DMPC sample made of a 1 mg/ mL solution measured at room temperature (≈20 °C) and ≈60% RH. From the oscillations of the curve and from the fact that the curve does not exhibit multilamellar Bragg peaks, one can deduce that this sample consists of less than three lipid bilayers. Simulations for this curve have so far not been successful, which may be due to the fact that N is not uniform over the whole area of this sample and that the sample consists of domains of one and two bilayers, respectively. (29) Nagle, J. F.; Katsaras, J. Phys. Rev. E 1999, 59, 7018.

Figure 3. Number (N) of DMPC bilayers on silicon or glass substrates of 15 × 25 mm2 as a function of the concentration of the lipid solution used for the spin-coating process. Solid symbols: N determined from Kiessig fringes; open symbols: N determined from Bragg peak width.

All simulations in Figure 2 show systematic deviations from the measured curves that may be due to a too simplistic or idealistic model. However, fundamental parameters such as the bilayer thickness and water layer thickness can be extracted. The parameters used for the simulations as described in ref 22 are given in Table 2. Note that the number of bilayers on the sample increases with the concentration of the lipid solution that was used for sample preparation. This relationship has also been observed for other lipids examined in this study (DLPC, DOPC, not shown) and is quantitatively plotted in Figure 3 for DMPC bilayers on glass and silicon substrates of 15 × 25 mm2. The filled symbols represent the N values calculated from Kiessig fringes, while the open symbols show the number estimated from the peak fwhm. The number increases almost linearly (line: linear fit) with the concentration. The samples represented in this plot were prepared in different series proving the reproducibility of the method. Because of solvent evaporation during the whole preparation process, the concentration is marked with large error bars. The number of bilayers deposited on a substrate depends not only on the concentration but also on the rotational frequency. Larger numbers of bilayers can be deposited on substrates by decreasing the rotational frequency (e.g., to values between 100 and 200 rpm). In this case there is no tossing of lipid solution from the substrate. Thus, the full amount of lipids pipetted onto the substrate remains on the sample leading to a higher sample thickness. This “slow-spin” method can be used as an alternative to the spreading method for the preparation of thick stacks with N ≈ 102-103 (not shown). However, the focus of this work is the preparation of thin films with N in the range of 1-30; therefore, we have concentrated on one rotational frequency of 3000 rpm (step 2) and have systematically varied the parameter of concentration. A dependency of the exact number on

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Figure 4. Reflectivity curves of one DMPC sample consisting of 10 bilayers on silicon in different phases, measured at e98% RH and at temperatures of 40.2, 23.0 and 14.3 °C (top to bottom). The insert shows an angular scan at the position of the first Bragg peak in LR phase. The peak fwhm of 0.012° (mosaicity) indicates that the misalignment of the sample is negligible.

ambient conditions such as temperature or relative humidity during the preparation process has not been observed. Figure 3 visualizes clearly the possibility to control the number of deposited bilayers by varying the concentration of the lipid solution in the above-described spin-coating process. Although these principles seem to apply to all lipids examined in this study, one “standard sample” (prepared from DMPC dissolved in 2-propanol) has been chosen for the characterization section in this work for several reasons. First, the preparation parameters such as solvent concentration and rotational frequency are directly comparable. Second, the sensitivity for structural changes because of phases and hydration can be better visualized. Other lipids and lipid-peptide mixtures that have been used for this work are listed in Table 1. Figure 4 shows the reflectivity curves of 10 DMPC bilayers in three different phases: LR phase (circles), P′β phase (triangles), and Lβ phase (diamonds). All curves have been obtained from the same sample measured at e98% RH in water vapor at different temperatures (40.2 °C LR phase, 23 °C P′β phase, and 14.3 °C Lβ phase). These curves exhibit significant structural differences. Compared to the LR curve (same as in Figure 2) with a d spacing of 53.9 ( 0.2 Å, the peak positions in the Lβ curve have shifted to smaller values of qz (d ) 61.4 ( 0.2 Å) as the lipid bilayer thickness increases in the phase where the hydrocarbon chains are stretched. While the LR curve exhibits strong Kiessig fringes, they are weak in the Lβ phase. In addition, the fourth Bragg peak becomes visible in the Lβ phase. But most significantly, the shape of the curve between the peaks has changed strongly. A pronounced modulation of the form factor is observed in the Lβ phase between the relatively weak Bragg peaks, which has not been observed before in thick samples. The Bragg peaks of the P′β phase indicate a higher repeat distance d ) 66.2 Å. The third and fourth Bragg peaks are strongly reduced, and there is not much structure visible between the peaks, possibly indicating a frustration of the ripple structure. The presence of a flat substrate may modify the formation of the ripple structure, depending on the number of bilayers. This is to be investigated in further studies. Quantitative analysis of T-dependent specular and nonspecular (diffuse) reflectivity can also shed some light on the precritical swelling just above the main phase transition, which is still a matter of debate.5,30 All these

Mennicke and Salditt

Figure 5. Bright-field optical micrograph of a defect in a spincoated DMPC sample (20 mg/mL 2-propanol solution) on silicon, imaged at room temperature and without hydrating vapor. The sample was in gel phase. The defect consists of a dewetted patch of bare silicon.

features demonstrate clearly the sensitivity of reflectivity measurements for structural differences in the abovedescribed oligo membrane samples. Note that for quantitative reflectivity analysis it is very important to distinguish between specular and off-specular intensity. This requires a very good alignment of the sample, which means a very low mosaicity. The angular scan (insert in Figure 4) demonstrates a very low misalignment of the spin-coated sample (in the range of 0.01°) close to the resolution limit of the measurement. Sample Homogeneity. While a reflectivity curve with Kiessig fringes indicates a well-defined number of bilayers, the samples may still exhibit a defect structure on various length scales. Experimentally, X-ray diffuse scattering gives access to the lateral film structure on the mesoscale between a few nanometers and several micrometers (limited by the coherence length of the X-ray beam). The level of diffuse scattering at the first Bragg peak was found to be 4 orders of magnitude lower than the specular peak (see the rocking curve in the insert of Figure 5). This indicates both a suppression of thermal fluctuations in the thin film and a high degree of lateral homogeneity on length scales up to the X-ray coherence length, which is on the order of a few micrometers. Contrarily, defects are observed in optical and AFM microscopy. These show defects consisting of dewetted patches (bare substrate) of typical lateral length scales larger than 10 µm (Figure 5). Between the defects, however, the sample thickness is almost uniform, in contrast to samples prepared with the spreading method, which exhibits thickness variations of a few tens of microns.24 The defect density and pattern in spin-coated samples seems to depend strongly on sample history, sample phase, and also whether the sample is fully or partially hydrated.31 Furthermore, substrate inhomogeneities or impurities can trigger dewetting during the spin-coating procedure itself. Full Hydration. While for some applications partial hydration is sufficient or even desirable,22 for other experiments full hydration is needed with the membranes immersed in water.9 Oligo membranes prepared with the spin-coating procedure can also be measured at full (30) Chen, F. Y.; Hung, W. C.; Huang, H. W. Phys. Rev. Lett. 1997 79, 4026. (31) Perino-Gallice, L.; Fragneto, G.; Mennicke, U.; Salditt, T.; Rieutord, F. Eur. Phys. J. E 2002, 8, 275-282.

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has decreased to 56.7 ( 0.2 Å. Importantly, the Kiessig fringes indicate that all bilayers remained on the sample. The influence of salt solutions, pH value, or osmotic pressure on the membranes shall be examined in further experiments.

Figure 6. Reflectivity measurement of DMPC membranes immersed in an aqueous PEG solution at an osmotic pressure of 8.9 × 105 Pa (open symbols). Reflectivity curves of one DMPC sample immersed in ultrapure water (solid symbols) measured 2:45, 5:35, and 8:26 h after the sample has been immersed in water (T ≈ 40 °C).

hydration in excess water. However, they seem to be less stable under water than thick multimembrane systems as prepared by spreading.9 Figure 6 shows the reflectivity measurements of 12 DMPC membranes on silicon measured in ultrapure water at a temperature of 43 °C (solid symbols). All curves have been corrected for illumination, background, and storage ring current. They have been shifted vertically in chronological order (top to bottom). Measurements have been taken in equidistant time intervals, beginning with 2 h 45 min after the sample has been set under water (top) to 8 h 26 min after immersion (bottom). After 5 h 36 min the membranes detached, and only one single membrane remained on the substrate. Obviously the stacks of membranes are less stable in water against unbinding from the substrate than thick spreaded samples (as examined in ref 9). The detailed mechanism of unbinding remains unclear.32 The systematic influence of temperature, time, and sample thickness and the fact that a single bilayer remains on the substrate suggest that the experimentally observed unbinding is an intrinsic instability of multilamellar bilayers on solid support rather than just a loss of loosely bound bilayers due to hydrodynamic perturbations or other secondary effects as suggested in ref 33. At the same time, it can be ruled out that the bilayers do peel off one by one, as predicted by simulations,34 since this would result in a measurable broadening of the Bragg peaks. A swelling of the membranes has also not been observed. The d-spacing remained constantly at 63.4 ( 0.3 Å. The membrane stacks can be stabilized on the substrate by applying osmotic pressure. The top curve in Figure 6 (open symbols) shows the reflectivity of 10 DMPC membranes on silicon in a PEG solution with a corresponding osmotic pressure of ≈8.9 × 105 Pa. Compared to the samples in pure water, the peak distance has shifted to higher values, indicating that d (32) Hartung, J.; Helfrich, W.; Klo¨sgen, B. Biophys. Chem. 1994, 49, 77-81. (33) Pabst, G.; Katsaras, J.; Raghunathan, V. A. Phys. Rev. Lett. 2002, 88, 128101. (34) Netz, R. Ph.D. Dissertation, Universita¨t Ko¨ln, 1994.

Summary and Outlook Diffraction from highly aligned, multilamellar samples can be used as a sensitive probe to study structure and interactions in multicomponent bilayers. Important examples are bilayers containing membrane-active molecules (such as stereols, peptides, and proteins) or membranes containing certain lipid mixtures that lead to a lateral organization on the nanometer to micrometer scale (so-called lipid rafts).35,36 Systems with amphiphilic peptides that act as antibiotic and fungicidal agents in the immune system of vertebrates are relevant both to biological and to pharmaceutical sciences. While the peptide function in the natural organism is often wellestablished, many of the underlying biochemical and structural mechanisms remain unknown, and valuable insight can be derived from simple model systems composed only of one or two lipid components and peptides at various concentrations.37 Complementary to NMR, diffraction techniques can be used to study the structural basis of such systems.15-17,38 To this end, small changes of the bilayer form factor because of the peptide contribution have to be detected. Note that in a reflectivity experiment, structural information is derived not only from the Bragg peaks but from the fitting of a complete curve. In samples of a few bilayers with a well-controlled number (such as presented in Figure 2), the advantages of probing the form factor between the peaks is particularly pronounced, since the structure factor does not lead to a complete destructive interference between the Bragg peaks. For this purpose all components can be co-dissolved in the desired ratio for one stock solution. Series with different peptide-lipid ratios of the antibiotic peptide alamethicin in DMPC membranes have been successfully prepared and measured (not shown). The results of the ongoing analysis will be discussed elsewhere. Beyond structure determination, the method presented here may be useful for a number of analytical techniques. Furthermore, the solid-supported membranes with functional proteins are currently developed and studied for device applications. In a number of future applications, a stack of a well-controlled number may help to overcome stability and efficiency problems. Acknowledgment. We thank Greg Smith for sharing his ideas and experience in solid-supported membranes with us and thank Franz Pfeiffer, Alexander Hildenbrand, Chenghao Li, and Ansgar Jarre for their help in the preparation and the measurements at HASYLAB. Financial aid by Deutsche Forschungsgemeinschaft through Grants SA 772/3 and SA-772/4 is gratefully acknowledged. LA025863F (35) Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 1417-1428. (36) Leidy, C.; Wolkers, W. F.; Jørgensen, K.; Mouritsen, O. G.; Crowe, J. H. Biophys. J. 2001, 80, 1819-1828. (37) Bechinger, B. J. Membr. Biol. 1997, 156, 197-211. (38) Mu¨nster, C.; Lu, J.; Schinzel, S.; Bechinger, B.; Salditt, T. Eur. Biophys. J. 2000, 28, 683-688.