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Formation of Planar Solvent-Free Phospholipid Bilayers by Langmuir-Blodgett Transfer of Monolayers to Micromachined Apertures in Silicon Thor D. Osborn and Paul Yager* Molecular Bioengineering Program, Center for Bioengineering, University of Washington, Seattle, Washington 98195 Received December 7,1993. In Final Form: October 3, 1994@ Solvent-free lipid bilayers were reproducibly formed across 100 pm apertures in a hydrophilic micromachined silicon support. The bilayers were assembled from mixed phosphatidylcholinelcholesterol monolayers at the air-water interface by the Langmuir-Blodgett method. "he presence and longevity ofthe bilayerswere determined by fluorescence microscopy. "he bilayer seal impedance was also measured. Such a mass producible device for supportinglipid bilayers is of potential value for biosensor development, for receptor and drug assays, and for biophysical studies of ion channels. Planar lipid bilayers are regularly employed as model systems for study of transport across biomembranes. In such studies a bilayer is suspended across an aperture in an electrical insulator that separates two aqueous compartments; changes in transmembrane potential and/or current can be measured by electrodes on either side of the bilayer. Both hydrophobic and hydrophilic supports have been employed, but the hydrophobic supports invariably require the presence of an organic solvent on the support surface,1*2 and hydrophilic supports have been limited to glass pipet with openings generally no larger than a few micrometers, although 30-pm openings have been used successfully.3 The formation of the bilayers is not particularly reproducible because of crude methods of forming the bilayers over the orifices and because the orifices in the substrates are neither particularly smooth nor reproducibly made. In the case of hydrophobic supports, the bilayer-bearing apertures are generally rough on a microscopic scale and are smoothed with a nonvolatile solvent. The bilayer is usually formed either by painting a lipid solution over the aperture followed by spontaneous thinning to a bilayer membrane4 or by forming a monolayer a t the air-water interface on both sides of the aperture and raising the water level first on one side, then the other.5 In either case the presence of the solvent torus surrounding the bilayer may perturb the membrane properties. Bilayers may be formed on hydrophilic glass pipet tips by dipping the tip 2 or more times through a collapsed lipid monolayer;6 however, the bilayer formation mechanism and resulting lipid structure are not well understood. Bilayer formation rates are generally rather poor with existing support methods (about 20%)and considerable operator technique is essential for success. If a superior, automated, method were available for supporting planar lipid bilayers, research into ion channels would be more widely accessible, and new technologies such as biosensors based on the permeability of transmembrane ion channels7-" would be possible. e Abstract published in Advance ACS Abstracts, November 1, 1994. m i t e , S.H.; Petersen, D. c.;Simon, s.;~ a f ~ sM. o ,Biophys. J. 1976,16,481-489. (2) White, S. H. Biophys. J . 1972, 12, 432-445. (3)Andersen, 0. S.Bwphys. J . l98S, 41, 119-133. (4) Mueller, P.: Rudin. D. 0.:Tien. H. T.: Wescott. W. C. J. Phvs. Chem. 1983,67,534-535. . ( 5 ) Montal, M.; Mueller, P. Proc. Natl. Acud.Sci. ( U S A . ) 1972,69, 3561-3566. (6) Coronado, R.; Latorre, R. Biophys. J. 1983,43, 231-236. (7) Arya, A.; Krull, U. J.; Thompson, M.; Wong, H. E. Anal. Chim. Acta 1986, 173, 331-336.
(6
Reasonable steps in this direction include the development of a geometrically simple and reproducible hydrophilic (i.e.,solvent-free)support and the application ofthe bilayer through the controllable and relatively well-understood Langmuir-Blodgett transfer method. A number of attempts at producing a bilayer support for biosensor research have been made. Notably, in every case Langmuir-Blodgett transfer was employed. Two separate designs employing a hydrogel to support the bilayer were tried with some ~ u c c e s sand , ~ ~a~multihole plastic device was demonstrated;'* however, those devices were not readily amenable to batch fabrication. A device employing a porous silicon support for the bilayer has been reportedlo that is well suited for batch fabrication but should suffer from the gradual decay in bilayer impedance seen on other porous support^.'^ Platinum rods have also been used as substrates for bilayers and multilayers.20 This approach has promise but does not provide a space between the support and the membrane, a limitation that may interfere with transmembrane protein function. Our approach to a mass-producible bilayer support device (BSDl2l is shown schematically in Figure 1. The BSD consists of an array of cavities in a silicon substrate sealed on the back side by an anodically bonded piece of Pyrex glass bearing AgIAgC1 electrodes and tracings to (8) Hongyo, K.;Joseph, J.; Huber, R. J.; Janata, J. Langmuir 1987, 3,827-830. (9) Krull, U. J.; Thompson, M. Trends Anal. Chem. 1985,4,90-96. (10) Ligler, F. S.;Fare, T. L.; Seib, K. D.; Smuda, J. W.; Singh, A.; Ahl, P.; Ayers, M. E.; Dalziel, A.; Yager, P. Med. Znstrum. 1988, 22, 247-256. (11)Thompson, M.; Krull, U. J.; Worsfold, P. J. Anal. Chim. Acta 1980,117,133-145. (12) Tedesco, J. L.; Krull, U. J.; Thompson, M. Biosensors 1989, 4 , 135-167. (13) Uto, M.; Michaelis, E. K.; Hu, I . F.; Umezawa, Y.;Kuwana, T. Anal. Sci. 1990, 6, 221-225. (14) Eldefrawi,M. E.; Sherby, S.M.;Andreou, A. G.;Mansour,N. A.; Annau, Z.; Blum, N. A.; Valdes, J. J. Anal. Lett. 1988,21,1665-1680. (15) Gotoh,M.;Tamiya,E.;Momoi,M.;Kagawa,Y.;Karube,I.AnaZ. Lett. 1987,20, 857-870. (16) Rogers, K. R.; Eldefrawi, M. E.; Menking, D. E.; Thompson, R. G.;Valdes, J. J. Biosens. Bioelectron. 1991, 6, 507-516. (17) Taylor, R. F.; Marenchic, I . G.; Cook, E.J. Anal. Chim. Acta 1988,213, 131-138. (18) Vodyanoy, V. In ZEEE Engineering in Medicine and Biology Society 10th Annual International Conference; Institute of Electrical and Electronics Engineers: New Orleans, 1988; pp 997-998. (19) Drachev, L. A.; Kaulen, A. D.; Semenov, A. Y.; Severina, I . I.; Skulachev, V. P. Anal. Biochem. 1979,96, 250-262. (20) Fare, T. L. Langmuir 1990, 6, 1172-1179. (21)Osbom, T. D.; Person, J. J.; Yager, P. InZEEE-Engineering in Medicine and Biology Society 11th Znternational Conference; IEEE: Seattle, USA, 1989; pp 1375-1376.
0743-746319512411-0008$09.0010 0 1995 American Chemical Society
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Figure 1. (A) Schematic side cutaway view of one of the 16 cavities of the BSD. The micromachined silicon chip is represented by light angled hatching and the Pyrex by horizontal hatching. The recessed tracing is in black and the Ag/AgCl electrode is represented by vertical hatching. The silicon wafer is 225 pm thick and the membrane is 3 pm thick. (B) Scanning electron micrograph of the interior of one micromachined cavity. The walls of the pyramidal cavity are bounded by the { 111) crystallographic faces of the silicon wafer. The cavity shown here is identical to those in the BSD, except that the aperture is approximately 50 pm in diameter (scale bar = 100pm). (C) Tracing pattern of the BSD. Dotted lines show borders of Si chip and the bases of the pyramidal cavities.
external contact pads. The surface oxide of silicon wafers is extraordinarily smooth,22providing a highly reproducible hydrophilic substrate. Phospholipids are necessary for the function of many membrane proteins, but many investigators have found the Langmuir-Blodgett (L-B) assembly of phospholipidsto be substantially more difficult than that of fatty a ~ i d s . ~Successfid ~ - ~ ~ deposition of more than a single phospholipid monolayer has generally required high monolayer surface pressures (above40 mN/ m) because of poor adhesion of polar lipid headgroups to such substrates. We have recently demonstrated that bilayers of a phospholipid-cholesterol mixture can be quantitatively and reproducibly transferred to solid oxidized Si wafers by L-B assembly as long as the surface pressure of the monolayer is carefully ~ o n t r o l l e d .We ~~ report here our success in using this method to form solvent-freesuspended bilayers spanning holes in the BSD. A 2:l molar ratio mixture of the lipids 1-stearoyl, 2-oleoyl-sn-glycero-3-phosphocholine (SOPC) and cholesterol was chosen for this work because of its high vesicle lysis tension and toughness28and our success in forming headgroup-out bilayers of this mixture on oxidized Si.27 SOPC was purchased from Avanti Polar Lipids (Alabaster, AL)and cholesterol was purchased from ICN Biomedical (Cleveland, OH). The purity of the lipids was verified by thin-layer chromatography on silica gel using a solvent system of 65:25:4 CHCl&H30WH20 by volume and 12 detection. The solvents and 12 were all of reagent grade or better. One mass-percent of the fluorescent probe DiI (l,l’-dioctade~yl-3,3,3’,3’-tetramethylindocarbocyanine perchlorate, Molecular Probes, Eugene, OR) was added to the SOPC/cholesterol mixture. The BSDs were made by the anodic bonding29 of micromachined silicon chips to electrode-bearing 2.5 cm by 3.7 cm by 0.3 cm Pyrex glass (Corning,code 7740)plates. A simplified BSD (S-BSD)for optical microscopy was also made using an unprocessed glass plate without electrodes. Micromachining was performed using established techn i q u e ~ .Substrates ~~ were formed from double-side polished (100) orientation wafers (Virginia Semiconductor, Fredericksburg, VA). A 4 x 4 array of circular 100 pm diameter apertures was defined by heavily boron doping one side of the wafer except in the areas of the intended apertures. Pyramidal cavities were then etched from the oppositeside using an etchant solution of ethylenediamine, pyrocatechol, pyrazine, and water.31 The heavily boron doped layer (about 3pm thick) blocked the etchant, leaving membranes of silicon with circular apertures as shown in Figure 1. The silicon wafer was oxidized to an oxide thickness of 60 nm before anodic bonding. The electrode-bearing glass plates were produced by a series of commonly-employedmicrolithographic steps. The tracing pattern (see Figure 1)was etched into the surface of the glass through use of a photoresist mask and 1O:l buffered hydrofluoric acid. A 150-nm layer of silicon nitride was deposited by ion-milling. Next, three metal (22) Ducker, W. A.; Senden, T. J.; Pashley, R. M. Langmuir 1992,8, 1831-1836. (23) Hasmonay, H.; Caillaud, M.; Dupeyrat, M. Biochem. Biophys. Res. Commun. 1979,89,338-344. (24) Honig, E. P.; Hengst,J. H. T.;Engelsen, D. d. J. Colloid Interface Sci. 1973,45, 92-102. (25) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985,47,105-113. (26) Langmuir-Blodgett Films; Roberts, G.,Ed.;Plenum Press: New York and London, 1990,425 pp. (27) Osborn, T. D. Doctoral Thesis, University of Washington, 1994. (28) Needham, D.; Nunn, R. S. Biophys. J. 1990,58,997-1009. (29) Wallis, G.;Pomerantz,D. I. J. Appl. Phys. 1969,40,3946-3949. (30) Bassous, E. IEEE Trans. Electron Devices 1978, ED-25,11781185. (31) Wu, X.-P.; Wu, Q.-H.; KO,W. H. Sens. Actuators 1986,9,333343.
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Figure 2. Time-sequence of motion of fluorescent particles: (A) initial configuration, (B) after 2 min; (C)after a total of 4 min. All of the particles within the aperture moved. Those within the center of the aperture moved in small, abrupt jumps on the order of 1pm, reminiscent of Brownian motion. The particles in contact with the edge of the aperture were relatively sluggish. The darkening of the images and loss of contrast in sequence resulted from photobleaching of the fluorophore. The scale bar is 50 pm.
layers were deposited in sequence by thermal evaporation in a single vacuum step: 15 nm of Cr, 85 nm of Pd, and 500 nm of Ag. Ultrasonically-assisted liftoff in acetone removed the siliconnitride and metal layers except within the tracing pattern. Another lithographic step was performed to etch away the Ag in Fe(N03)3solution except at the desired electrode locations. The electrodes were chloridizedin FeCl3 solution. Anodic bonding to the silicon wafers was carried out for 10 s at 350 "C. Before use, a 2-cm portion of a glass capillary tube was glued with 5 min epoxy (Devcon) to one end of the Pyrex plate so that the silicon face of the S-BSD would not touch a flat surface when laid face down. The glue did not touch the silicon chip. The S-BSDwas then cleaned in 0 2 plasma (a 10-min exposure to an 0 2 plasma (300 W, 1Torr 0 2 ) ) . The cavities of the S-BSD were filled by placing it in buffer
and alternately applying and releasing a vacuum. The buffer used for filling the cavities (and as the subphase for monolayers) was 100 mM NaCl and 10 mM TRIS in 18-MQ deionized water at 20 f 1"C, adjusted to pH 7.4 with HCl, and filtered to 0.2 pm. A small single-barrier trough was built for L-B transfers to the BSDs. Monolayers were spread on the aforementioned buffer from 10 mg/mL lipid solution in CHCl3. Two to five minutes was allowed for solvent evaporation prior to compression. The dipping speed was 100 pmh. Monolayer surface pressures of 46.6 and 47.6 mN/m, which are approximately 4 mN/m below the monolayer collapse pressure, were chosen for bilayer formation based on our prior theoretical work and experiments with nonporous Si substrate^.^^ Following bilayer transfer and while still in the subphase, the S-BSD
Langmuir, Vol. 11, No. 1, 1995 11
Letters was placed face down in a submerged plastic Petri dish for transfer to an optical microscope. The bilayer transfer experiment was repeated 3 times with the same S-BSD. After each experiment the S-BSD was rinsed in methanol and soaked for an hour in deionized water. This was done to minimize trapping of buffer salts within the micromachined cavitiesbefore cleaning and refilling with buffer. Microscopic observations of substrates with adherent monolayers were performed with a Zeiss ICM 405inverted fluorescence microscope (Oberkochen, Germany) using a mercury arc lamp and a Zeiss filter set (450-490,FT 510, and LP 520). Images were recorded with a DAGE-MTI (Michigan City, IN) Model 66 SIT camera and either saved on S-VHSvideotape or transferred directly to a Macintosh I1 computer equipped with a Data Translation (Marlboro, MA) Quickcapture frame grabber. Images were processed using the public domain Image software package (Version 1.47)from NIH. Electrical impedance measurements were carried out in a Faraday cage using a Dagan 8900 amplifier with a 100-Mi2 feedback resistance headstage. Data were stored using a videotape adaptor for later observation. The complete BSDs were cleaned and filled in the same manner as the S-BSD and bilayers were transferred similarly, but the BSDs were left submerged in the L-B trough. One of the 16 apertures of the S-BSD was malformed due to errors in processing. In three successive experiments fluorescence intensity above the background was seen in 14, 13, and 9 of the 15 functional apertures, respectively, indicating that aperture-spanning bilayers were present. In the first experiment the S-BSD was stored for 13 days in buffer at 20 f 3 “C and observed again. Of the 14 apertures having above-background intensity within minutes of transfer, 9 had maintained that intensity, indicating that this may be a very longlived bilayer preparation. The fluorescence within most of the apertures was relatively featureless. However, in some cases features were seen that confirmed the formation of bilayers. In the first experiment highly fluorescent domains or particles were observed within and near some of the apertures, as illustrated in Figure 2. These particles probably resulted from segregation or crystallization of the fluorescent probe. The particles on the silicon dioxide surface did not move, but those within the aperture were in constant motion. Real-time observation indicated that the speed of motion of the particles was inversely related to their size, consistent with Brownian motion. Trajectory plotting shows a large drift component in the particle motions, however, obfuscating measurement of the diffusivity (Figure 3). The aperture shown in Figure 2 was observed until the fluorophores in the particles became severely photobleached and could no longer be distinguished (about 1 h). Notably, throughout all of their motions the particles never went out of focus, indicating that they were trapped in the plane of the aperture. Investigators have generally found that formation of model membranes from monolayers5 requires spreading an excess of lipid at the air-water interface,32forcing the monolayers to their collapse pressures and causing partial monolayer collapse. The “tip-dip” formation of bilayers on patch-clamp pipet from lipids at an airlwater interface is usually performed with a collapsed monolayer.33 These aperture-spanning bilayers were formed from monolayers at 4 mN/m below their collapse pressure, which should have avoided forming collapsedregions on the monolayers. (32)Tancrhde, P.;Paquin, P.; Houle, A.; Leblanc, R. M. J.Biochem. Biophys. Methods 1983,7,299-310. (33)Coronado, R.Biophys. J. 1986,47,851-857.
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In the second experiment some of the apertures were observed to be filled with large domains of two distinct fluorescent intensities, as shown in Figure 4A. Unlike the irregular and unchanging shapes of the particles seen in the first experiment, these domains formed circles or arcs, suggesting that the light and dark regions were coexistingimmiscible fluid phases. These domains slowly moved within the aperture, and, in one dramatic instance, an arc-shaped domain that spanned the aperture was seen to detach from one edge and rapidly contract into a more rounded shape, as seen in Figure 4B. A two-phase coexistence region with two immiscible liquid phases has been demonstrated for mixtures of 1,2dipalmitoyl-sn-glycero-3-phosphocholine(DPPC) and cholester01.~~ The phase diagram shows this two-phase coexistenceregion for moderate cholesterol concentrations a t temperatures above the main-chain melting transition temperature of pure DPPC. Further, fluorescence micrographs of monolayers formed from mixtures of 1,2dimyristoyl-sn-glycero-3-phosphocholine and cholesterol show features similar to those of parts A and B of Figure 3 under conditions where two immiscible liquid phases are believed to form.35 Apparently, bilayers of SOPC and cholesterol can also form two immiscible liquid phases, although we have observed such features in only a few apertures. This two-phase coexistence may play a role in the enhanced mechanical stability of such membranes.36 The differences in features observed in separate experiments are probably the result of slight variations in experimental conditions, such as the surface pressure of the monolayers at the time of transfer to the S-BSD. This will be explored further in future work. Preliminary electrical data on the BSD include a seal resistance of approximately 30 f 10 MQ, a bilayer breakdown potential of about 1 V, and an apparent formation rate (Le., a seal between bilayer and substrate) of about 10% for each aperture. The seal resistance is rather low compared to the GQ-range seals often observed on pipets, although the much greater circumference of the BSD apertures may account for some of this discrepancy. The sealed bilayer formation rate is disappointing but within the neighborhood of success rates for other techniques. The low formation rate may be due in part (34) Vist, M. R.; Davis, J. H. Biochemistry 1990,29,451-464. (35)Hirshfeld, C. L.; Seul, M. J . Phys. (Paris) 1990,51,1537-1552. (36)Bloom, M.; Evans, E.; Mouritsen, 0. G. Q. Rev. Biophys. 1991, 24,293-397.
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Figure 4. Metamorphosis of immiscible fluid domains: (A) initial configuration showing a central arc spanningthe aperture; (B) less than 1 min later, the central arc-shaped domain has detached from the right side and contracted to the left. The scale bar is 50 pm.
to poorly defined electrode potentials resulting from the anodic bonding procedure, which causes migration of Ag+ ions into the glass. The susceptibility of the bilayer to rupture from these fluctuating potentials could be responsible for the low observed seal rate. Redesign of the BSD assembly process to yield better electrodes and optimization of the L-B transfer speed could improve these results. The electrical capacitance of the bilayer is not measurable with the BSD as the built-in capacitance of the device ranges from 0.5 to 1.1nF, depending on the tracing length, and is therefore much greater than the approximately 100 pF expected for the bilayer membrane in the aperture. This capacitance could be greatly reduced by employing a thicker oxide layer on the Si chip and reducing the tracing width; however, a thicker oxide would interfere with anodic bonding. The use of other bonding techniques will be considered in future embodiments of the BSD. The BSD structure is capable of supporting stable aperture-spanning bilayers when an appropriate formation protocol is observed. The BSD is completely amenable to the batch-fabrication methods of the semiconductor industry. One micromachined silicon wafer and one processed Pyrex wafer could be anodically bonded and then diced to form dozens or hundreds of BSDs. The aperture size of the BSD may be readily varied within the limits of semiconductor process line widths. Currently, apertures as small as 1pm are possible. The BSD design allows 16 independent model membrane experiments to be carried out simultaneously. A BSD could be made with any number of electrodes and apertures through the same processes, however, merely by changing the photolithographic mask patterns. The availability of an off-the-shelfbilayer support could
spur biophysical research by allowing investigators to spend less time preparing pipets or black lipid membrane apparatus. The planar surface of the BSD also allows use of fluorescent or radiographic methods for determination of the number and distribution of proteins and associated ligands after the electrical measurements. The ability to perform many experiments simultaneously may also reduce the time required to obtain results. The relative reproducibility of the BSD and the L-B method may also reduce the impact of variability in operator technique. The availability of a reproducible bilayer support is also a crucial step in the development of biosensors based on the conductive properties of ligandgated ion channels.37
Acknowledgment. We gratefully acknowledge the NIH for support under a Predoctoral Training Grant in Biotechnology GM08437 (T.D.O.), the National Science Foundation for Grant DIR-9024971 (P.Y.) and for a Graduate Research Fellowship (T.D.O.), the Washington Technology Center 09-1044 for funding through the Microsensors Center (P.Y.), and the Center for Bioengineering for continuing support. We also thank Professor John C. Berg for use of his Langmuir-Blodgett trough for portions of this study, and Professors Viola Vogel and Bertil Hille for numerous helpful conversations. All micromachiningwas performed in the Washington Technology Center Microfabrication Laboratory on the UW campus. LA930715Y (37) Yager, P. In United States Statutory Invention Registration;U.S. Patent Office: Washington, DC, 1987.
