Langmuir 1993,9, 2764-2767
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Self-Assembled Phospholipid/Alkanethiol Biomimetic Bilayers on Gold Anne L.Plant Biotechnology Division, National Institute of Standards and Technology, Gaithersburg, Maryland 20899 Received July 2,1993. I n Final Form: August 23,1993 Alkanethiols can act as hydrophobic monolayer substrates for the formation of phospholipid-containing bilayers. Phospholipid vesicleswere allowed to fuse to alkanethiolmonolayers, resulting in stable, solventfree lipid bilayers on gold electrodes. The capacitance of these bilayers has been determined by impedance measurements, and the effect of the pore-forming peptide melittin on the bilayer has been determined by the measurementof faradaic current. These supportedself-assemblingphospholipid/alkanethiolbilayers demonstrate properties consistent with those of fluid membranes, and provide a useful way to study the electrical characteristics of membranes in the absence of solvent.
Introduction Planar phospholipid bilayer membranes are of great interest as model systems for the study of cell membrane transporter phenomena and other forms of membrane signal transduction. The interest in forming phospholipid/ alkanethiol bilayers on gold is to identify a model system that exhibits membrane mimetic behavior like LangmuirBlodgett (LB) films and black (bilayer) lipid membranes (BLMs), but with the additional advantages of ease and reproducibility of preparation, long-term stability, formation in the absence of solvent, and formation on a support that is conducive to surface analysis and is electrically conductive. Recently there has been a great deal of interest in the characterization of self-assembled monolayers (SAMs) of alkanethiols on gold. Since the report that showed that dialkyl disulfides form oriented monolayers on gold,l monolayers of alkanethiols have been examined in detail. These monolayers have been determined to be tightly packed and structurally well-defined.2 Contact angle measurements confirm that these monolayers make the gold surface extremely hydr~phobic.~ Several approaches to using hydrophobic surfaces for the formation of structurally stable supported phospholipid bilayers have been reported. One approach involved transferring LB films to surfaces made hydrophobic by s i l a n i ~ a t i o n In . ~ another ~~ approach, phospholipid vesicles were fused with an LB monolayer which was transferred to a quartz slide? In an effort to ensure the formation of a fluid membrane, an acyl chain-terminated thiol-containing polymer was developed to derivatize gold surfaces and allow fusion of sonicated vesicles to form a bilayer.' (1) Nuzzo,R. G.; Fusco, F. A.; Allara, D. L. Spontaneously organized molecular assemblies. 3. Preparationandpropertiesof solutionadsorbed monolavers of organic disulfides on gold surfaces. J. Am. Chem. SOC. 1987,109,2358-5368, (2)Porter. M. D.; Bright, T. B.; Allara, D. L.: Chidsey, C. E. D. Spontaneously organized-molecular assemblies. 4. Structkal characterization of n-alkyl thiol monolayers on gold by optical ellipsometry, infrared spectroscopy, and electrochemistry. J. Am. Chem. SOC.1987, 109,3559-3568. (3) Bain, C. D.; Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides,G. M.;Nuzzo, R. G. Formation of monolayer films by spontaneous assembly of organic thiols from solution onto gold. J. Am. Chem. SOC.1989,111, 321-335. (4) Egger, M.; Heyn, S.-P.; Gaub, H. E. Synthetic lipid-anchored receptors based on the binding site of a monoclonal antibody. Biochim. Biophys. Acta 1992,1104,45-54. (5) Miller, C.;Cuendet,P.;Gratzel,M. K+ sensitive bilayer supporting electrodes. J. Electroanal. Chem. 1990,278, 175-192. (6) Kalb, E.; Frey, S.; Ta", L. K. Formation of supported planar bilayers by fusion of vesicles to supported phospholipid monolayers. Biochim. Biophys. Acta 1992,1103,307-316.
Lipids in organic solvent have also been exposed to silanizeds or alkanethioP-coated surfaces. This report shows that self-assembled alkanethiol monolayers formed directly on gold surfaces can also provide a hydrophobic substrate for the formation of stable self-assembledlipid bilayers by phospholipid vesicle fusion. The bilayer formed by this process provides a model of phospholipid membranes with which permeability and structural characteristics of membranes can be studied electrochemically on a planar, supported, structurally defined, and relatively rugged surface. The advantage associated with the use of alkanethiols is their selfassembly, their efficiency of packing and coverage of the metal substrate, and their relatively long-term stability; these characteristics have been established through careful characterization of these l a y e r ~ . ~ aThe formation of phospholipidlalkanethiol bilayers by fusion of phospholipid vesicles has the important advantage of being easy and reproducible in the absence of solvent.
Materials and Methods The fabrication of the gold electrodesused in these experimenta has been previously des~ribed.~ Gold was sputterdeposited onto chromium-coated 2-cm X 2-cm silicon wafers to a nominal thicknessof 2000 A. Monolayers were formed by placing freshly prepared gold electrodes into a 1 mM ethanolic solution of hexanethiol,decanethiol,or octadecanethiolfor several days at room temperature. Before assembly of the electrochemicalcell, the electrodes were rinsed thoroughly with ethanol and dried under a stream of nitrogen. The electrochemical cell used was athree-electrodesystem with a Ag/AgCl reference electrode. The body of the cell was a Delrin or polyethylene cylinder which was positioned on top of the planar gold electrode and a flattened gold wire lead. The exposed electrode surfacearea was 0.32 om2, and the volume of the cell was approximately 0.6 mL. Phospholipid/alkanethiolbilayers were formed by adding a solution of phoepholipid vesiclesto the self-assembledalkanethiol monolayer (Figure1A)to allow self-assembly of a phospholipid/ alkanethiol bilayer (Figure 1B). Figure 1 is not intended to
indicate the absence of tilt of the alkane chains of the bilayer; evidence from IR measurements indicates that alkanethiol monolayers exhibit a chain tilt of 2+30°.2 The thermodynamic driving force for formation of a bilayer from an alkanethiol monolayer and phospholipid vesicles is the increase in entropy that is achieved when water is excluded from (7) Spinke, J.; Yang, J.; Liley, M.; Ringsdorf,H.; Knoll, W. Polymersupported bilayer on a solid substrate. Biophys. J . 1992,63,1667-1671. (8)Florin, E.-L.; Gaub, H. E. Painted supported lipid membranes. Biophys. J. 1993,64,375-383. (9) Tarlov, M. J. Silver metalization of octadecanethiol monolayers self-assembled on gold. Langmuir 1992,8, 80-89.
This article not subject to U.S.Copyright. Puiblished 1993 by the American Chemical Society
The Langmuir Lectures
Langmuir, Vol. 9,No.11,1993 2765
I/II IIII IIIIII
A
R
C
s s s s s s s s s s s s s s Au
~
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103
104
Frequency (Hz) I I
Au
I
Figure 2. Representativeplot of measuredimpedancevs applied
ac frequency (e),and the result of nonlinear regression analysis (-1. Capacitancewas calculatedby assuming a simpleequivalent circuit of a resistor in series with a capacitor.
L
Figure 1. A schematic representation of an alkanethiol mono-
layer on gold (A) and the bilayer formed by addition of phospholipid vesicles (B).This picture does not intend to depict lack of chain tilt or the stoichiometry of the phospholipid/ alkanethiol bilayer. the hydrophic chains of the alkanethiol and the hydrophobic acyl chainsof the phospholipid. In the absenceof a hydrophobic surface, most phospholipid molecules will form lipid vesicIes or liposomes spontaneously when dispersed in water. The amphiphilic nature of phospholipids and their geometry result in the incorporation of the lipid molecules into spherical vesicles composed of lipid bilayers. For thisstudy, vesiclesof l-palmitoyl2-oleoylphosphatidylcholine (POPC) were prepared by two methods, injection'O and extrusion.11 To prepare lipid vesiclesby either method,a solution of POPC in chloroform was placed in a glass test tube or flask, and the solvent was evaporatedunder a stream of nitrogen,leavinga film of dried lipid in the vessel. The sample was then placed in a vacuum desiccator overnight to remove residual chloroform. Injected vesicles were prepared by resolubilizing2 pmol of lipid in 0.05 mL of dry 2-propanol, and injecting this solution with a syringe into 1 mL of 20 mM Tris and 150 mM NaCl, pH 7.3 (TBS), which was being mixed by vortex. On the basis of the measured values for diffusion coefficients of the vesicles as determined by autocorrelation of scattered this method consistentlyresults in vesicles which are approximately 0.01 pm in diameter and have a narrow size distribution. Vesicles were also prepard by extrusion in the absence of any solvent. Addition of TBS directly to a dried lipid film and mixing by vortex or shaking result in multilammelarliposomes. These large liposomes were converted to small unilamellar vesicles by extrudingthe samplethroughpolycarbonatemembranes. Three filtration steps were performed using filters with pore sizes of 0.6,0.4, and 0.2pm sequentiallyto achievea population of vesicles of 0.2-pm diameter. Vesicles prepared by either method were added to the cell in TBS at a fiial lipid concentrationof 0.2 mM. The vesicle solution was left in contact with the alkanethiol-coatedelectrode for at least 1 h and as long as 16 h. Formation of the bilayer was monitored by impedance measurements in the presence of vesicles, and was considered complete when the capacitance reached a stable value. The cell was then rinsed several times with water or electrolyte solution to remove excess vesicles. (10) Batzri, S.;Korn, E. D. Single bilayer liposomes prepared without sonication. Biochim. Biophys. Acta 1973, 901, 157. (11)Olson, F.;Hunt, C. A;Szoka, F. C.;Vail, W. J.; Papahadjopoulos, D. Preparation of liposomes of defined size distribution by extrusion through polycarbonate membranes. Biochim. Biophys. Acta 1979,557, 9-23. (12) Selaer, J. C.;Yey, Y.; Baskin, R. J. A lighbscattering characterization of membrane vesicles. Biophys. J. 1976, 16, 337.
Capacitanceof the alkanethiolmonolayers and phospholipid/ alkanethiolbilayers was determinedby measuringimpedanceas a function of frequency. Impedance measurements were made with a Solartron 1286electrochemicalinterface end a Solartron 1250frequency responseanalyzer (Schlumberger). A sinusoidal ac signal was applied at frequencies between 10 and 63 OOO Hz. Nonlinear least-squares analysis was used to fit the impedance amplitude as a function of frequency to a simple model for an equivalentcircuit consistingof a resistor (theelectrolytesolution) in serieswitha capacitor(theelectrodemembrane). The collected data fit this simplistic model well, aa shown in Figure 2. In this frequencyrange, the contribution of membrane resistance to the total impedance amplitude could be ignored. Comparison of these impedance data with data simulated using an equivalent circuit containing a parallel RC circuit (the membrane) in series with a resistor (the solution) indicated that the membrane resistance was at least 1 X 10s Q. Measurements were made in 10 mM KCl or 10mM NaF with 0.1 or 0.01-V ac amplitude and 0.0 V vs Ag/AgCl. No difference was seen for the different ac amplitudes. Cyclic voltammetry was performed with solutions of 1 mM KJ?e(CN)ein 1M KC1. Cyclic voltammetrywas performedwith either the Solartron 1286 interface or a BAS100 (Bioanalytical Systems).
Results and Discussion Figure 3 compares the capacitance values measured for alkanethiol monolayers and phospholipid/alkanethiolbilayers as a function of the chain length of the alkanethiol. We observed electrical characteristics for the monolayers which are in agreement with those previously published by others. A plot of the inverse capacitance of the monolayers as a function of alkanethiolchain length shows a slope (0.05) and intercept (0.1)which are very similar to those reported for alkanethiol monolayers by Porter et aL2 For bilayers formed from POPC vesicles prepared by injection, the slope is similar, but not identical to that seen for the monolayers (0.0581,suggesting that the phospholipid interacts similarly with monolayers of different chain lengths, but may form a more defect-free bilayer with the longer chain thiols. As has been previously observed? we saw that the capacitance determined for the hexanethiol monolayers was sensitive to the supporting electrolyte, and a larger capacitance was measured in KC1 than in NaF'. While monolayers of long-chain alkanethiols are highly ordered and resistant to penetration by ions, the shorter chain thiols are less ordered and presumably contain packing defects2 The larger hydrated F-ion may be less likely to
2766 Langmuir, Val. 9,No. 11, 1993
.
00
,
The
. 5
0
10
20
15
NUMBER OF ALKANETHIOL (CH 2 ) UNITS
Figure 3. Inverse capacitance vs alkanethiol chain length for alkanethiol monolayers (e),and bilayers formed by the addition of POPC vesicles prepared by injection (B). Regression analysis of the monolayer data: slope, 0.05; intercept, 0.1. For the bilayer: slope, 0.058. Table I. Effect of Solvent on POPC/Octadecanethiol Bilayer Capacitance. membrane system
Cm(rrF/cm2)
Cm-PL
octadecanethiol monolayer + POPC (injected) POPC (extruded) POPC/decane initial (15-30 min) final (16 h) BLM of PC in decane solvent-free bilayers of PC in hexane
0.96 0.13b 0.68 f O.Olb 0.64
1.92* 1.76
0.42 0.60 0.39 0.72d
0.12 1.48 0.78 1.44
+
+
a The contribution to the specific capacitance of the bilayer, C,, which is due to the phospholipid half-bilayer is reported as C--PL. b Average and standard deviation of data collected on at least three different d a y with at least two different electrodes. Data taken from ref 15. Data taken from ref 14.
penetrate defect sites than the smaller C1- ion. In contrast, capacitance measured for bilayers, even those prepared from hexanethiol monolayers, was the same in NaF as in KC1. This is an indication of the effectiveness of the phospholipid in coating the electrode, even over an alkanethiol monolayer containing defects. The electrical capacitance of the phospholipid halfbilayer (Cm-pL) was calculated from the value for the capacitanceof the alkanethiol monolayer (Cm-monohyer) and the capacitance of the phospholipidJalkanethio1bilayer (Cm-bhyer) by the following relationship: 1 -=-Cm-PL
1
1
Cm-bilayer
Cm-monolayer
Values for the specific capacitance of bilayers prepared from octadecanethiol monolayers and different preparations of phospholipid vesicles are shown in Table I, along with the capacitance values calculatedfor the phospholipid half-bilayers. Since water can penetrate the polar head group region and into the acyl chain region to a t least the C2 position,13the phospholipid monolayer can be modeled as a large capacitor (the polar headgroups) in series with a smaller capacitor (the alkane chains).14 The hydrocarbon (13) Griffith, 0.H.; Dehlinger, P. J.; Van,S.P.J . Membr. Biol. 1974, 15, 159. (14) Ben, R.;Frohlich, 0.;Lauger, P.; Montal, M. Electrical capacity of blacklipidfii8and of lipid bilayersmade from monolayers. Biochim. Biophys. Acta 1975,394,323-334.
Langmuir Lectures
portion of the phospholipid monolayer is therefore the predominant contributor to the overall half-bilayer capacitance. The data in Table I show the effect of the solvent used in the preparation of liposomes on the capacitance of the resulting membrane. For this study, phospholipidvesicles were prepared by two different methods. As seen in Table I, the capacitance measured for half-bilayers (Cm-PL) prepared from phospholipid vesicles made by injection with 2-propanol, 1.92 pF/cm2, is slightly larger than the value for half-bilayers prepared from extruded vesicles, 1.76 pF/cm2. both of these capacitance data indicate halfbilayers with a larger capacitance than has been previously reported for BLMs14J5which are formed by painting a decane solution of phospholipid over a septum. These values are also larger than the capacitance measured for so-called %olvent-free’’ bilayers, which are also formed across a septum but are prepared from a monolayer of lipid spread a t an air-water interface from a solution of lipid in organic s01vent.’~In an effort to understand this discrepancy, decane was added to the phospholipid for some vesicle preparations, a t a decane:phospholipidmolar ratio of 51, prior to solubilization and injection with 2-propanol. Fusion of these decane-containing vesicles with a decanethiol monolayer resulted in a significantly lower half-bilayer capacitance which increased from 0.72 to 1.48 pFJcm2over 16 h, as can be seen in the table. An increase in capacitance over time for decane-containing bilayers has been reported pre~ious1y.l~Recently, timedependent solvent loss from a similar bilayer system has been directly observed by plasmon resonance imaging.8 The effect of organic solvent on bilayer capacitance has been studied.l4J6 Benz et a1.14 compared BLMs with solvent-free bilayers. While the solvent-free method did not result in bilayerswhich were completelyfree of solvent, the amount of solvent present compared to BLMs was greatly reduced, resulting in an increase in the membrane capacitance. As shown in Table I, from their data one can calculate a half-bilayer specific capacitance for solventfree bilayers of dioleoylphosphatidylcholine of 1.42 pF/ cm2. In contrast, the half-bilayer capacitance calculated from data reported by Dilger et al.15for BLMs made from a solution of phospholipid in decane was 0.78pF/cm. As seen in Table I, capacitance data for phospholipid/ uctadecanethiol bilayers prepared in the presence of decane appear to correspond well to literature values for bilayers prepared with solvent. On the other hand, for phospholipidlalkanethiolbilayers prepared in the absence of any solvent, i.e., from vesicles prepared by extrusion, the capacitance values are larger than values reported for planar membrane systems where decane or hexane were used in the preparation. Consider the relationship among capacitance, dielectric constant, and thickness of the membrane:
cm= q,u/d where C m is the specific capacitance (normalizedby surface area), EO is the permittivity of free space, K is the dielectric constant of the material comprising the membrane, and d is the thickness of the membrane. It is apparent that there are two parameters that may be responsible for this inconsistency, the thickness, d , of the layer, and the dielectric constant, K , of the membrane. At this point, neither of these parameters can be known unambiguously, and furthermore, they can both change in response to changes in membrane composition. For example, there (15) Dilger, J. A.; McLaughlin, S. G. A.; McIntosh, T. J.; Simon,S. A. The dielectric conetant of phospholipid bilayers and the permeability of membranes to ions. Science 1979,206,1196-1198.
The Langmuir Lectures
Langmuir, Vol. 9, No. 11, 1993 2767
E (mV) va AgAgCl
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voltammetric r&ponse of 1 mM KsFa (CN)e in 1 M KCl at a decanethiol monolayer (a),and a POPCI decanethiol bilayer (v). (B)Aftar 30-min incubations of the bilayer with 1 X 1 P M melittin in the presence ( 0 )of 500 mM phoephate buffer, pH 7.0, and in TBS only (H). Inset: cyclic voltammetric response from a bare electrode. Scan rate 50 mV/ 8.
Figure 4. (A) Cyclic
are X-ray diffraction datal6*" which suggest that phosphatidylcholine bilayers prepared in the presence of a 1:l molar ratio of decane have a smaller chain tilt, and therefore a larger hydrocarbon thickness, compared to bilayers without decane. Compared to an alkanethiol monolayer, phospholipid/ alkanethiol bilayers exhibit not only greatly reduced capacitance but also an enhanced ability to block the electrode to redox species. Figure 4a shows cyclic voltammograms of 1 mM Fe(CN)& in 1 M KC1 with a decanethiol monolayer-modified electrode and a bilayermodified electrode. For comparison, a cyclic voltammogram of this ion a t a bare gold electrode is shown in the inset. The data show that the decanethiol monolayer significantly reduces oxidation and reduction of the ion. These data are consistent with published data of the effective barrier that alkanethiol monolayers create at electrode surfaces.2 The subsequent formation of a bilayer with POPC further reduces the amount of faradaic current measured compared to the monolayer, indicating the formation of an even more effective barrier to electron transfer. Figure 4B indicatss the sensitivityof the bilayer integrity to the pore-forming peptide melittin. Melittin in TBS was added to the electrochemical cell for 30 min at room (16) Tardieu, A.; Luzzati, V.; Reman, F. C. Structureand polymorphism of the hydrocarbon chaine of lipids: a study of lecithin-waterphases. J. Mol. Biol. 1973, 75, 711-733. (17) McIntosh, T. J. Differences in hydrocarbon chain tilt between hydrated phosphatidylethanolamine and phosphatidylcholine bilayers. A molecular packing model. Biophys. J. 1980,29,237-246.
temperature, after which the melittin solutionwas replaced with 1mM Fe(CN)6&in 1M KCl. Cyclic voltammograms are shown for the response of K3Fe(CN)eafter incubation with 1X 1o-B M melittin in the presence and absence of 500mM phosphate. Incubation of melittin with the bilayer in the presence of a high concentration of phosphate ion resulted in little increase in the Fe(CN)& response. This is consistent with the inhibitory effect of phosphate on melittin activity. High ionic strength has been shown to promote self-association of melittin in solution, thereby reducing the association of melittin monomer with the bilayer as is required for expression of ita lytic activity.18 However, in the absence of a high concentration of phosphate ion, incubation with melittin resulted in a significant increase in bilayer permeability, rendering the bilayer nearly as permeable as the monolayer alone. This effect of melittin on the bilayer took place in the presence of 6 mM ethylenediaminetetraacetate (EDTA),which was added to inhibit any residual phospholipase activitywhich may be present in melittin preparati~ns.'~Only a negligible change in membrane capacitance accompanied the large increase in faradaic current, suggestingthat a change in permeability occurred that was not due to a significant change in membrane dielectric constant. PhospholipicValkanethiolbilayer-modified electrodes were very stable in aqueous solution and exhibited consistent and stable responses even after being used for days for scores of measurements and many changes of electrolyte solutions. The phospholipid layer could be disturbed by removing allwater from the electrode surface. It could be entirely removed by rinsing the electrode with ethanol; a new bilayer could then be formed from this recovered monolayer.
Conclusion Interest in supported lipid bilayers which mimic cell membranes has led to the study of phospholipid thiolmor silanez1analogs for the preparation of cell-membrane-like monolayers on metal or glass surfaces. As an alternative to that approach, this report shows that stable bilayers of phospholipids and alkanethiols can be formed on planar gold surfacesusing phospholipidvesicles. Two important characteristics of a good model membrane system are ion impermeability and lipid fluidity. Capacitance values presented here indicate that phospholipicValkanethio1 bilayers exhibit very low permeability, and their response to melittin indicates that these bilayers are sufficiently flexible to accommodate the pore-forming protein. This simple method produces supported planar bilayer membranes in the absence of solvent, and will be useful in the study of reconstituted protein/lipid systems, in studies of membrane interfacialphenomena, and in sensor applications.
Acknowledgment. Thanks are extended to Dr. Michael Tarlov for providing the gold electrodes, Dr. Baldwin Robertson for providing the electrochemicalinterface and software, and Dr. P. C. Pandey for technical assistance. (18) Hider, R.; Khader, F.; Tatham, A. Lytic activity of monomeric and oligomeric melittin. Biochim. Biophys. Acto 1983, 72.9,206-214. (19) Dempeey, C. E. The actions of melittin on membranes. Biochim. Biophys. Acto 1990, 1031,143-161. (20) Diem, T.; Czajka, B.; Weber, B.; Regen, S. Spontaneous aasembly of phospholipid monolayers via adsorption onto gold. J.AM.CheM. SOC. 1986,108,6094-6095. (21) Kallury, K. M. R.; Lee,W. E.;Thompson, M. Enhancement of the thermal and storage stability of urease by covalent attachment to phospholipid-bound silica. Anal. Chem. 1992,64,1062-1068.
