Bilayer Lipid Membrane Formation on a ... - ACS Publications

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Bilayer Lipid Membrane Formation on a Chemically Modified S-Layer Lattice A. Schrems,† A. Kibrom,‡ S. K€upc€u,† E. Kiene,† U. B. Sleytr,† and B. Schuster*,† †

Department for NanoBiotechnology, University of Natural Resources and Life Sciences, Vienna (BOKU), Muthgasse 11, 1190 Vienna, Austria ‡ Austrian Institute of Technology (AIT), Muthgasse 11, 1190 Vienna, Austria ABSTRACT: The present paper describes the generation of a biomimetic model lipid membrane on bacterial surface (S-) layer which covered the entire surface of various sensors. The S-layer lattice allows one to be independent from the underlying solid material and provides a biological surface and anchoring structure for lipid membranes. S-layer proteins were chemically modified via binding of two amine-terminated phospholipids. Subsequently, a bimolecular lipid membrane anchored to the previously generated viscoelastic lipid monolayer was generated by the rapid solvent exchange technique. Characterization of the intermediate (monolayer) and final membrane structures (bilayer) was performed by imaging, surface-sensitive, and electrochemical techniques. This bilayer lipid membrane generated on an S-layer lattice revealed a thickness of ∼6 nm and constitutes a stable supported model membrane system with highly isolating properties showing a membrane resistance of 8.5 MΩ  cm2.

1. INTRODUCTION Although investigations on supported lipid membranes started approximately 25 years ago,1 there is still sound interest in these model lipid membranes. In particular, nature-inspired, so-called biomimetic supported lipid membranes are currently receiving increasing attention.2
6 In general, they have proven to be highly valuable for studying the properties and function of membranebound peptides and proteins and investigations on membranemediated processes such as cell
cell interactions1,7 and biological signal transduction.8 The intrinsically low degree of nonspecific adsorptivity of supported membranes makes them interesting as an interface between the nonbiological materials on the surface of a sensor or implant and biologically active fluids.1,8
10 Potential applications include the acceleration and improvement of medical implant acceptance, programmed drug delivery, production of catalytic interfaces, as a platform to study transmembrane proteins and membrane-active peptides, and as biosensors.11
16 In basic research, as it is in the focus of the present work, this model lipid membrane is well suited for studying the adsorption and incorporation of anionic peptides and genetically modified tyrosine kinases. Furthermore, a broad range of surface-sensitive techniques are now available to study all of the previously mentioned issues.3,8,14
17 The present contribution deals with the formation and characterization of a biomimetic model lipid membrane generated on crystalline bacterial cell surface layers (S-layers). It has been shown that S-layer lattices constitute stabilizing scaffoldings for many different types of lipid membranes.2,5 The highly porous S-layer can be recrystallized on a broad spectrum of organic and inorganic substrates and if necessary be cross-linked to enhance the stability. Interestingly, the recrystallization of an S-layer on the substrate commonly caused a reduction of the surface roughness.18 Furthermore, the S-layer is utilized as (1) a r 2011 American Chemical Society

tethering structure to decouple the lipid membrane from the solid support, (2) an ion reservoir necessary for electrochemical measurements like impedance spectroscopy, and (3) a chemically modifiable binding matrix for hydrophobic lipid anchor molecules. For the generation of artificial lipid membranes on electrodes, the S-layer protein SbpA from Lysinibacillus sphaericus CCM 2177 represents a convenient building block with a chemically modifiable surface. A common method in particular to build up a lipid double layer structure is the vesicle rupture/fusion mechanism.19,20 Beside this, another method can be utilized, the rapid solvent exchange (RSE) technique.21 In the past, the generation of lipid bilayers on SbpA has been attempted by various techniques, for instance the Langmuir
Blodgett and Langmuir
Schaefer methods, liposome fusion, and mixed micelle adsorption. All of these techniques are time-consuming, and hence, a simpler and faster process like RSE is preferred. In the present study, SbpA is modified by a sparsely hydrophobic lipid anchor region under physiological conditions. These linchpins can be constructed by linking phosphatidyl ethanolamines to surface exposed carboxyl groups on S-layer proteins. Subsequently, bilayer formation is initiated by flushing the chemically modified, hydrophobic S-layer lattice with an ethanolic phospholipid solution. In a final step, the excess of lipidic material is removed by flushing with HEPES buffer resulting in an assembly of the lipids into a dense bimolecular lipid membrane. Received: October 28, 2010 Revised: February 3, 2011 Published: March 14, 2011 3731

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2. EXPERIMENTAL SECTION 2.1. Materials. Following gold-coated substrates were utilized for the present study: quartz crystals (QSX 301, Q-Sense, Gothenburg, Sweden), microscope slides (fcc 111, Sigma-Aldrich), and SF-10 glass (Phasis, Switzerland). Protein recrystallization buffer was prepared with 0.5 mM TRIS base (Sigma) and 10 mM CaCl2 (98%, Sigma) and adjusted to pH 9 with 0.1 M NaOH. Ethanol absolute (Sigma-Aldrich), Milli-Q water (Millipore, Molsheim, France; resistivity: 18.2 MΩ cm
1), NaOH (Sigma-Aldrich), hydrogen peroxide (30%, Fluka), ammonia (25%, Merck), and Hellmanex II (2%, Hellma) were used as cleaning solutions. 400 mM 1-ethyl3-(3-dimethylaminopropyl)carbodiimide (EDC) (Sigma-Aldrich) and 200 mM sulfo-N-hydroxysulfosuccinimide sodium salt (S-NHS) (Sigma-Aldrich) were stored as aliquots in 50 mM PBS buffer at pH 6.5 at
20 °C. PBS buffer for the anchoring procedure was prepared by mixing monobasic and dibasic sodiumdihydrogenphosphate (Merck) solutions in distinct ratios and brought to the desired pH by titration with HCl (Riedel-de Haen) and 1 mg/mL sulfo-succinimidyl acetate (Pierce) in 50 mM PBS pH 7.4 for blocking of primary amines.22 All used lipids were purchased from Avanti polar lipids. For the anchoring procedure 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamineN-(hexanoylamine) (L1) and 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (L2) were used, containing 0.003% Triton X-100 (Fluka) in 50 mM PBS buffer at pH 7.4. For the rapid solvent exchange method the lipids 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1,2-dioleoyl-3-trimethylammonium-propane (chloride salt) (DOTAP) were solved in 80% ethanol in a ratio 3:1. As dilution buffer for the RSE method and in all electrochemical experiments 10 mM HEPES buffer (Gerbu) containing 150 mM NaCl and adjusted to pH 7.4 was used. As refractive index matching liquid in the surface plasmon resonance measurements 1-iodonaphtalene from Sigma-Aldrich was used. 2.2. Methods. 2.2.1. Cleaning of Substrates. For QCM-D, EIS and electro kinetic measurements 5 MHz quartz crystals coated with a 50 nm thick gold layer were used. For the contact angle measurements a gold coated microscope slide with a 10 nm thick gold layer was used (Sigma Aldrich). All gold surfaces were cleaned intensively before every use according to Kern et al.23 Briefly, the sensor was first immersed in a MilliQ water/H2O2/NH3 (5:1:1) at 75 °C for 10 min and rinsed two times with Milli-Q water/ethanol/Milli-Q water. After this procedure the sensor was dried under a stream of nitrogen. Finally, all gold surfaces were cleaned immediately previous use by ozone plasma treatment in a plasma cleaner to remove any organic contaminations from the surface (Plasma Prep2, Gala, Gabler Labor Instruments, Germany). 2.2.2. Coating of Substrate. The bacterial cell surface layer protein SbpA (molecular weight 127 kDa) was isolated from Lysinibacillus sphaericus CCM 2177 according to the previously reported procedure.24 The protein solution (1 mg/mL) was diluted 1:10 with recrystallization buffer before incubation onto the substrates. This step was done very quickly so that no self-assemblies occurred. The protein was incubated for 3
12 h at room temperature and afterward the substrates were rinsed with Milli-Q water in order to remove excess protein. 2.2.3. Anchoring Procedure of Lipids on SbpA. The lipids L1 and L2 were covalently linked via its primary amines to free-standing carboxyl groups of the S-layer protein SbpA. This technique is similar to that described by Sinner et al.25 The lipids were solubilized at a concentration of 0.5 μmol/mL in 0.003% (w/v) Triton X-100 and bound to the SbpA surface via a reactive ester coupling mechanism. EDC (400 mM) and S-NHS (100 mM) were each dissolved in 5 mL PBS buffer (10 mM, pH 6.5
6.8) and stored at
20 °C until use. For activation of the proteinaceous surface EDC and S-NHS were mixed in equal amounts and SbpA was incubated for 30 min. After the activation procedure the lipid solution (0.5 μmol/mL, 0.003% Triton X-100, in 50 mM PBS pH 7.4) was added and the reaction took place for 1 h. After this step the surface was rinsed with buffer or Milli-Q water.

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2.2.4. Rapid-Solvent Exchange (RSE) Method. A modified version of the RSE technique reported elsewhere was used for generating the model lipid membrane.21,26 Briefly, the S-layer surface with the bound L1 and L2 was first flushed with 80% ethanol and then a 10 mM solution of lipids (DOPC: DOTAP, 3:1) in 80% ethanol was allowed to incubate on the modified S-layer surface substrate for 10 min at room temperature. Higher content of ethanol was not used due to the loss of SbpA structure as previously described.27 Afterward, the ethanol lipid mixture was rapidly displaced by a large amount of HEPES buffer. Care was taken in all steps to avoid the formation of air bubbles at the surface that could disturb the bilayer formation. 2.2.5. Transmission Electron Microscope (TEM) Preparation. The preparation was similar as described previously.28 Briefly, images of S-layer lattices were taken by a FEI Tecnai G2 20 TEM at 120 kV. Hydrophilized carbon-coated piloform supported 300-mesh copper grids were incubated with the carbon-coated side on a drop of the SbpA solution. The attached S-layer proteins were chemically fixed with 0.5% glutaraldehyde in 0.1 M potassium dihydrogen phosphate buffer (pH 7.4) for 10 min and after three washing steps with Milli-Q water, negatively staining with 1.0% uranyl acetate in Milli-Q water for approximately 1 min was performed. The samples were dried overnight and then used for TEM studies. 2.2.6. Quartz Crystal Microbalance with Dissipation Monitoring (QCM-D) and Electrochemical Impedance Spectroscopy (EIS) Measurements. QCM-D measurements were carried out with a Q-Sense E4 device (electronic unit) equipped with four standard flow modules (Qsense AB, Gothenburg, Sweden). The QSX 301 gold-covered crystal sensors have a fundamental frequency of ∼5 MHz (Q-Sense AB, Gothenburg, Sweden) and were cleaned as previously described (2.2.1). Afterward they were immediately mounted in the flow modules of the instrument. Frequency (Δf) and dissipation (ΔD) shifts were recorded using Q-Soft 401 software, version 2.5.7.505 from Q-Sense AB. The presented results correspond to the seventh overtone. All experiments were done at a temperature of 21 °C ( 0.02 °C. S-layer protein binding and recrystallization was analyzed by the Sauerbrey relation.9 The data Δf and ΔD at the overtones n = 3, 5, 7 measured at the binding of the lipidic anchor molecules and the bilayer formation have been fitted to the Voigt
Kelvin-based viscoelastic model. The fitting routine is included in the Q-Tools 3 software, version 3.0.9.268 from Q-Sense AB. Furthermore, a flow module allowing simultaneous QCM-D and EIS measurements has been used (QCM-D E1, Q-Sense AB, Sweden). In this cell, the quartz crystal sensor is connected to a potentiostat (CH Instruments, CHI660c, Austin, U.S.A.) and used as working electrode. The counter electrode is a platinum plate at the top of the chamber whereas the Ag/AgCl reference electrode is located in the outlet of the flow cell. The frequency range was 1 mHz to 100 kHz. An AC potential of 15 mV was applied at a DC bias voltage of 100 mV versus the used reference electrode. All impedance data were fitted by the CHI software in a parallel equivalent circuit R
R ZCPE, whereas ZCPE was in all cases a constant phase element. 2.2.7. Zeta-Potential Measurements on QCM-D Sensors. The zeta potential of the sample was determined by streaming potential measurements using an adjustable gap cell for QSX sensors in an electrokinetic analyzer (SurPASS, Anton Paar GmbH, Graz, Austria). Two planar samples are separated by a gap of 90 to 100 μm and form a channel. The streaming potential was sensed by Ag/AgCl electrodes. A background electrolyte of 1 mM KCl solution was used and the pH was varied in the range of 2.5
6.5 with 0.1 M HCl. Higher pH regions were not investigated because the zeropoint charge was the important value for the characterization. The zeta potential, ζ, was determined from the measured streaming potential based on the Fairbrother
Mastin approach29 ξðUstr Þ ¼

ηkB dUstr ε0 εr dp

where Ustr is the streaming potential, dp is the pressure drop across the streaming channel, ε0 is the vacuum permittivity (8.854  10
12 J
1 C2 m
1), εr is the dielectric constant of the solution (78.3), η is the solution 3732

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Figure 1. Chemical drawing of the used anchor lipids for S-layer modification. (a) L1, 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamineN-(hexanoylamine) and L2, 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine. (b) Schematic sketch of the binding reaction of lipids (L1 and L2) on an S-layer lattice on a gold substrate. viscosity (0.8902 mPa s), and κB is the electrical conductivity of the bulk solution. 2.2.8. Contact Angle Measurements. The contact angle of water on gold surfaces with different coatings was obtained using a goniometer (Easy Drop, DSA15, Kr€uss, Hamburg). The data were evaluated by the enclosed analysis software. First the functionalized surface was dried under a nitrogen stream (5 min), and then a drop of Milli-Q water (5 μL) was deposited on the surfaces and immediately afterward the static contact angle was determined. On each surface tree drops were analyzed on different positions. 2.2.9. Thickness Determination by Combined Ellipsometry and Surface Plasmon Resonance Spectroscopy (SPR). The ellipsometry platform EP3 SE SPR (Accurrion, G€ottingen) was used to determine the thickness of each building up step in a liquid environment. All measurements and the reference on the bare gold surface were done in HEPES buffer. The setup consists of a 60° prism (SF-10, Hellma Optik), a substrate, and a copper temperature unit, whose temperature can be controlled and adjusted by a thermostat. Gold coated SF-10 glass was used to tune the SPR angle to a value which can be measured by a working range between 49° and 59°. The angular position of the smallest angle, the matching condition between the evanescent electromagnetic waves and surface plasmons, is a function of the refractive index and the thickness of the layer adsorbed or attached onto the gold surface. If the angle shifts after adsorption, the thickness of the organic layer can be calculated.30 The thickness of the titanium oxide and gold layer is known from the provider. Prism and substrate were arranged in a Kretschmann configuration.31 The EP3
SE/SPR device is equipped with a 658 nm solid state laser (coherent cube) and is operated in a nulling ellipsometer configuration. An angle spectrum was recorded and thickness values were fitted with the EP3View V2.01 software using a three layer model with the following parameters: n = 0.188 and k = 3.579 for the titanium/gold layer and n = 1.450 and k = 0 for SbpA. In the literature, the refractive index values for solid supported lipid bilayers vary from 1.45 to 1.5.32
34 In this study n = 1.490 and k = 0 was used for the calculation of the thickness of the lipid layer.35

3. RESULTS AND DISCUSSION 3.1. Modification of the S-Layer (SbpA) Lattice Surface. The aim of this modification is to supply the surface of SbpA with hydrophobic anchor molecules. SbpA provides a variety of functional groups that can be chemically modified to change the surface

properties.36,37 In the present study, the carboxylic acid groups were activated by the well-known reactive ester reaction with EDC and S-NHS. Free amino groups are able to bind to this activated functionality and form an amide bond. Hence, the binding of the two lipids 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(hexanoylamine) (L1) and 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (L2) (see Figure 1a) on an activated SbpA surface has been explored. Weigert et al. determined the amount of carboxyl groups on an S-layer protein very similar to SbpA and found 1,6 carboxyl groups per nm2.36,37 However, the distribution of these groups exposed on the surface or located in the pores of the S-layer lattice is not known. Hence, the two lipids L1 and L2 with different distances of the free amino groups to their hydrophobic tails were applied to obtain a higher number of bound lipids since the S-layer lattice is a highly porous structure, and therefore, the chance to bind the lipids on the surface but also somewhat deeper in the pore region of the S-layer lattice might be higher. The success in modification was assessed by static contact angle measurements, QCM-D combined with streaming potential measurements, and SPR. Because L1 and L2 bound via its polar head groups to the porous S-layer lattice and expose its alkyl chains, an increase of the surface hydrophobicity can be expected. Hence, first screening for highest hydrophobicity was performed by static contact angle determination at different pH values during the activation and binding of different ratios of these two lipids. The highest hydrophobicity could be observed at pH 6.5 during the reactive ester formation although the contact angle dropped slightly down from 61.3° for the unmodified SbpA to 56.5°. The pH was then set to 7.4, so that the anchoring reaction of L1 and L2 occurred at approximately pH 7.22 Subsequent binding of L1, L2, and mixtures of L1 and L2 resulted in the highest hydrophobicity of SbpA at a 1:1 lipid ratio (78.8°). Therefore the lipid ratio of 1:1 was utilized in all further experiments. Table 1 summarizes the data of the static contact angle measurements. At this point it is important to note that this modification did not form a dense lipid monolayer on the S-layer surface, which would have led to a higher contact angle (>90°) and a thickness of approximately 1.5 nm. The above-mentioned values are typical for classical SAM monolayers where the headgroup has a hydrophobic functionality.38
41 3733

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Table 1. Contact angle determination on functionalized gold substrates. The values are the average of 3 single measurements surface

contact angle

Au

71.1 ( 0.58

SbpA

61.3 ( 0.73

SbpA (EDC/S-NHS activated)

56.4 ( 0.49

L1:L2 = 1:0

58.4 ( 2.32

L1:L2 = 2:1 L1:L2 = 1:1

76.1 ( 0.43 78.8 ( 1.17

L1:L2 = 1:2

71.5 ( 0.42

L1:L2 = 0:1

69.9 ( 0.35

Figure 3. (A) Scheme of the experimental performance of the streaming potential measurements (picture provided by Anton Paar GmbH, Graz, Austria). (B) Dependence of the zeta potential as determined by streaming potential measurements on the pH value for each building up step of the anchoring procedure.

Figure 2. TEM images of SbpA coated grids. (A) Unmodified SbpA and (B) SbpA with anchored lipids (L1:L2 = 1:1). The p4 lattice symmetry of SbpA is visible on both specimen.

McGillifray et al. assured lipid bilayer generation by utilizing RSE on sparsely anchor region on SAMs of low hydrophobicity.21 Sinner et al. utilized a similar modification procedure of thiopeptides and they also observed an increase of the contact angle of approximately 20°.25 However, the main advantage of binding hydrophobic anchors to the SbpA lattice in contrast to other randomly and sparsely arranged anchor molecules might be the provided defined spatial distribution of the linchpins finally leading to nano patterned lipid membranes.5 TEM was used to investigate the lattice structure of the SbpA coating before and after the anchoring procedure (Figure 2). TEM is a powerful technique for imaging and resolving structures in the nanometer range. As shown in Figure 2, small crystalline patches can be seen on both specimen; hence, the p4 lattice is not affected by the surface modification. This finding is also in accordance with previously published data.37 Because on the S-layer lattice very different functionalities are exposed, the surface charge may also play an important role in the generation of a bound lipid bilayer. For this reason, an ex situ combination of QCM-D and streaming potential measurements was performed to obtain on the one hand information about the changes in mass and on the other hand the changes in zeta potential during each S-layer modification step. The changes of the zeta potential at different pH values are shown in Figure 3, whereas the data of the QCM-D measurements and the determined pI values are listed in Table 2. After each building up step, the observed titration graphs showed different behaviors according to the change of charged groups on the surface (Figure 3b). Whereas the bare gold surface remained negative in the whole

pH range, the zeta potential of SbpA coated substrates changed to more positive values. The pI of SbpA was calculated to 4.69,42 whereas the apparent pI value determined by isoelectric focusing was determined to be 4.2.43,44 Hence, the presently determined pI value of SbpA (Figure 3b; Table 2) is in good agreement with previously published data.43,44 Protection of free amino groups resulted in a shift of the pI value to a more acidic value because the amount of carboxylic acid groups was higher with respect to the amino groups. The shape of the graph of zeta potential
pH dependence changed to a smaller slope in the alkaline range. After binding of the lipid anchor molecules, the acid groups were converted to amides, and hence, the pI was shifted to a more basic value. This experiment showed that the surface charge is altered after each building up step. The adsorbed mass could be determined by QCM-D as indicated in Table 2. The thickness of SbpA was calculated by the Sauerbrey equation and determined to approximately 11 nm (see Table 2), which is in good agreement to previously published data.26 The coupling procedure was followed by two independent measurements. The coupling procedure resulted in a thickness of the formed layer of dQCM-D = 0.33 ((0.13) calculated by the Voigt-Kelvin viscoelastic model and a thickness of dSPR = 0.6 ( 0.3, determined by QCM-D and SPR, respectively. Both data, together with the contact angle measurements pointed to the formation of a viscoelastic and loosely packed lipid monolayer. 3.2. Formation of Lipid Bilayer by the Rapid Solvent Exchange (RSE) Method. The RSE technique was preferred to the conventional liposome fusion technique because of a higher reproducibility and faster generation of the lipid bilayer in particular for larger surfaces.45 Liposome fusion of large unilamellar vesicles was tried but no generation of a closed lipid bilayer could be observed on SbpA. Though, fusion reagents like divalent ions were added. Nevertheless, the outcomes of the liposome fusion studies was that positively charged liposomes 3734

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Table 2. Outcomes of the Adsorption Behavior Measured by QCM-D and Zero-Point Charges Determined by ex Situ Transfer of QCM-D Gold Sensors in an Elektrokinetic Analyzera surface

Δf (Hz)

ΔD  10-6 (a.u.)

mass (ng cm-2)

dQCM-D (nm)

pI

dSPR (nm)

Au

n. d.

n. d.

n. d.

2.87

200

52.8b

SbpA protected NH2

84.00 ( 5.34 0

2.22 ( 0.41 0

1486.80 ( 94.52 0

4.18 3.92

11.11 ( 0.83 0

8.0 ( 0.2 0

anchored lipids

3.17 ( 1.61

12.70 ( 6.11

4.21

0.33 ( 0.13c

0.6 ( 0.3d

2.33 ( 0.58

b

d

The number of QCM-D experiments was five. Values given by provider. Calculated by viscoelastic model. On gold coated SF-10 glass, dSPR values are related to specific refractive index of each layer. a

b

Figure 4. Rapid solvent exchange method monitored by QCM-D in combination with electrochemical impedance spectroscopy. (a) Start of addition of SbpA, (b) rinsing with Milli-Q water, (c) rinsing with HEPES buffer, (d) EDC/S-NHS activation, (e) addition of anchor lipids, (f) rinsing with 80% EtOH, (g) addition of lipid solution in 80% EtOH, and (h) flushing with HEPES buffer.

were likely to adsorb onto the sparsely hydrophobic surface under physiological conditions (data not shown). Hence, RSE was performed with the same lipid composition than the previously mentioned liposomes, i.e. DOPC and DOTAP at the ratio of 3:1. The combination of QCM-D and EIS revealed the formation of a dense lipid bilayer. In Figure 4, the frequency and the dissipation change during the building up steps are monitored. From a previous study it is known that the time required for S-layer recrystallization on gold to form a closed monolayer is 3 h.46 Therefore this time was assessed in all of these experiments. Moreover after this time, the frequency as well as the dissipation reached a plateau and no further changes could be observed. The chemical binding of lipid anchors was described in section 3.1. Before the addition of the RSE solution, the surface was flushed with 80% EtOH solution, which shifts the frequency to very negative values. Such an ethanol shift has been previously examined, and it turned out that the magnitude of this shift depended on the overall ethanol content.47 Control experiments have been performed by flushing the crystalline SbpA with 80% EtOH. No change in mass could be determined by this treatment as proven by QCM-D (data not shown). After the addition of the lipid solution, the frequency shifted back to more positive values. Only during flushing with HEPES buffer, the frequency and dissipation changed dramatically. Table 3 summarizes all results measured by the electrochemical QCM-D for each modification step. After RSE, the frequency remained constant with a shift in

c

d

the range of
50 Hz. Due to the different solvents used, the shift in frequency and dissipation can only be compared after the final rinsing with HEPES buffer. Moreover, in contrast to studies on lipid bilayer formation on inorganic surfaces by liposome fusion,19,20 the shift in dissipation was not close to zero as a shift in dissipation of ∼11  10
6 a.u. has been observed (Table 3). However, lipid membranes facing a solid support might be more rigid than those on SAM’s (∼3  10
6 a.u.).31 It is conceivable that both above-mentioned membranes are, due to the tighter surface interaction, more rigid than a lipid membrane sparsely anchored to a hydrophilic proteinaceous lattice. The thickness of the lipid bilayer was calculated by the VoigtKelvin viscoelastic model and was calculated to dQCM-D = 6.30 ( 0.43 nm. In comparison, the thickness dSPR determined with the SPR setup is with a value of 5.80 ( 0.20 nm in good agreement. Although the anchor region contributes to the measured thickness, these values are on the upper limit for lipid bilayers. However, similar values were determined by ellipsometry as 5.70 ( 1.00 nm have been reported for DOPC lipid bilayers in the same buffer solution.48 DOPC/DOTAP membranes on bare electrodes determined by AFM revealed a thickness of 4.7 ( 0.11 nm.49 Hence, the thickness determined for the presently formed membrane on the S-layer lattice is also in a good agreement with data from the literature. Moreover, another reason for the elevated thickness might be the interaction of HEPES buffer with the lipid bilayer. Peiro-Salvador et al. analyzed the influence of HEPES and PEPES buffer as well as sodium chloride on DOPC liposomes by smallangle X-ray scattering, and it turned out that these buffers caused membrane swelling.50 After each building up step within the EQCM-D setup, EIS was performed to determine the membrane capacitance as well as the membrane resistance. The results of these measurements are listed in Table 3. The value of the constant phase element refers to an electrical element with an impedance of ZCPE = 1/Q(iω)R, where Q is the constant phase element coefficient with the unit Siemens per unit area times Hz
R respectively Farad per unit area times sR
1. The exponent R may vary in the range from 0 (ideal resistor) to 1 (ideal capacitor). This element was used in the fitting procedure because the S-layer itself as well as the S-layer supported lipid bilayer do not behave like an ideal capacitor. Moreover, it is important to take into account that the S-layer lattice, onto which the anchored lipid membrane is resting, is a highly hydrated structure with water-filled pores.2 For SbpA (thickness is 9 nm), ∼40% of the volume of the S-layer lattice is occupied by protein and 60% consists of water.51 Hence, one can not assume that Cblm is equally distributed over the whole surface. Therefore, the coefficient of the constant phase element of the membrane Qblm can not be set equal to Cblm because this fit parameter is coupled to the electrolyte resistance Rel (in this case 8.5 Ω cm2).52 This dependence is given by the 3735

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Table 3. Summary of the Results and Fitted Data of the EQCM-D, EIS, and Adsorption Kinetics Measurementsa surface

R

Cblm (μF cm
2)

R (MΩ cm2)

Δf (
Hz)

ΔD  10
6 (a.u.)

SbpA

30.95 ( 1.25

0.964

22.56 ( 0.92

0.55 ( 0.27

89.87 ( 14.31

2.53 ( 0.72

anchored lipids

24.85 ( 3.34

0.920

12.01 ( 1.61

1.10 ( 0.48

5.04 ( 2.72

1.92 ( 0.80

7.05 ( 0.32

0.915

2.88 ( 0.13

8.50 ( 2.09

54.00 ( 10.59

11.31 ( 1.33

lipid bilayer a

Q (μF Hz
R cm
2)

The number of E-QCM-D experiments was four.

following equation: Cm ¼ ½QRel ð1
RÞ 1=a The capacitance of the bare gold sensor surface in HEPES buffer was determined ∼30 μF cm
2. After recrystallization of SbpA the capacitance drops down (∼ 22.6 μF cm
2) indicating that the SbpA layer itself was acting as a first barrier for charged molecules. The capacitance after the chemically binding of the anchor lipids was significantly lower than for SbpA which is a sign for a sparsely hydrophobic region. Finally, the capacitance dropped down to a value of 2.9 μF cm
2 after formation of the lipid bilayer (see Table 3). This determined capacitance is higher than that of bilayers formed by a rigid surface-bound lipid- or SAM-monolayer and a second leaflet composed of phospholipids.32,48,53 However, whenever a low capacitance for the composite double layer membrane was observed, the first layer (i.e., a bound monolayer or SAM) revealed already a very low capacitance, and the addition of the second phospholipid monolayer did not have a pronounced effect.32,48,53 Furthermore, the described RSE technique required ethanol for the preparation of the bilayer; hence, low amounts entrapped ethanol with a dielectric constant of 25 (at 298 K)54 might also be responsible for the increased capacitance. However, the presently described bilayer lipid membrane on SbpA having a thickness of approximately 6 nm revealed a significant lower specific capacitance than peptide tethered and protein tethered bimolecular lipid membranes with the advantage of a much higher specific membrane resistance.6 Interestingly, SbpA with or without anchor lipids on gold revealed a rather low resistance, but bilayer formation by RSE caused a pronounced increase of the membrane resistance to 8.5 MΩ cm2 (see Table 3). Hence, the present lipid membrane, generated by the RSE technique, is well suited for electrochemical measurements.

4. SUMMARY AND CONCLUSION The present paper reports on a straightforward approach to generate a biomimetic S-layer supported lipid membrane. This composite structure is composed of a closed S-layer protein on a sensor surface. Functional groups on the proteinaceous layer have been utilized to bind anchor molecules presenting its hydrophobic moieties. The presence of these sparsely bound linchpins led to an increased contact angle compared to unmodified SbpA indicating an increased hydrophobicity of the surface. This anchor layer is soft and viscoelastic resulting in an average thickness of 0.33 and 0.6 nm as determined by QCM-D and SPR, respectively. By the application of the RSE-technique a (bimolecular) lipid membrane, anchored via the bound linchpins on the S-layer lattice, could be formed. The shift in frequency and dissipation is higher compared to bilayer lipid membranes directly deposited on a sensor surface. However, it is evident that a tight surface interaction of the adjacent lipid leaflet results in a higher overall membrane rigidity compared to a lipid membrane sparsely

anchored by hydrophobic interaction to a hydrophilic proteinaceous lattice. Moreover, the thickness of the generated membrane (∼6 nm) is in good agreement with published data and supports a bimolecular structure. Evaluation of the S-layer supported lipid membrane by EIS spectra resulted in a membrane capacitance of approximately 2.9 μF cm
2 and a membrane resistance of 8.5 MΩ cm2. These values are in good agreement with other electrically tight tethered membranes. Although in biological systems membrane-associated biomolecules prefer neutral or negatively charged membranes, the present model membrane has its qualification e.g. for the binding of filamentous viruses,55 DNA,56 or the transport-related Naþ and Kþ-activated ATPase from kidney.57 Furthermore, positively charged membranes have also proven to be useful for studying the host
guest complex of valinomycin.58 The presently characterized lipid membrane will be utilized for studies on the adsorption and incorporation of anionic peptides and genetically modified tyrosine kinases. Furthermore, both the fluidity of the lipid molecules and of incorporated membrane proteins will be investigated by several surface sensitive techniques including fluorescence and infrared spectroscopy to support the performance of this platform.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected].

’ ACKNOWLEDGMENT Financial support from the Austrian Science Fund (FWF), Project 20256-B11 is gratefully acknowledged. The authors thank Jacqueline Friedmann for her skilful assistance in the AFM work and Denise Schach for helpful and stimulating discussions on the EIS data. ’ REFERENCES (1) McConnell, H. M.; et al. Supported Planar Membranes in Studies of Cell-Cell Recognition in the Immune-System. Biochim. Biophys. Acta 1986, 864 (1), 95–106. (2) Schuster, B.; Pum, D.; Sleytr, U. B. S-layer stabilized lipid membranes (Review). Biointerphases 2008, 3 (2), FA3–FA11. (3) Sackmann, E.; Tanaka, M. Supported membranes on soft polymer cushions: Fabrication, characterization and applications. Trends Biotechnol. 2000, 18 (2), 58–64. (4) Vockenroth, I. K.; et al. Incorporation of alpha-Hemolysin in Different Tethered Bilayer Lipid Membrane Architectures. Langmuir 2007, 24 (2), 496–502. (5) Schuster, B.; Sleytr, U. B. Composite S-layer lipid structures. J. Struct. Biol. 2009, 168 (1), 207–216. (6) Sinner, E.-K.; et al. Self-Assembled Tethered Bimolecular Lipid Membranes. In Advances in Clinical Chemistry: Elsevier: Amsterdam, 2009; Chapter 7, pp 159
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