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Ion channel reconstitution into lipid bilayer membranes on glass patch pipettes
329
Bioelectrochemistry and Bioenergetics, 12 (1984) 329-339 A section of J. Electroanal. Chem., and constituting Vol. 173 (1984) Elsevier Sequoia S.A., Lausanne - Printed in The Netherlands
651 --ION CHANNEL RECONSTITUTION INTO LIPID BILAYER MEMBRANES ON GLASS PATCH PIPETTES l
W. HANKE,
C. METHFESSEL,
Arbeitsgruppe Biophysikalische D - 4630 Bochum (F. R. G.) (Revised
manuscript
received
U. WILMSEN
and G. BOHEIM
Chemie von Membranen,
December
L.ehrstuhl ftir Zellphysiologie,
Ruhr-Universitiit,
29th 1983)
SUMMARY Planar lipid bilayer membranes were formed at the tip of glass patch clamp pipettes, and ion channels reconstituted by fusion of protein-containing vesicles with the bilayer. The advantages of the method over conventional bilayer techniques are numerous and include better current resolution, improved bilayer stability, compatibility with patch clamp apparatus and simple handling. Completely solvent-free lipid bilayers without any pretreatment can be prepared and only small amounts of materials are required.
INTRODUCTION
Living cell membranes control ion flow by integral membrane proteins that form ion-selective channels. The ion currents carried by single channels can be measured by patch clamp recording or by lipid bilayer reconstitution. In the patch clamp developed by Neher and co-workers [l] a patch of cell membrane, small enough to give good signal resolution, is electrically isolated with a glass micropipette. Although the technique is very successful, channels can be identified only by their electrical and pharmacological properties and it is difficult to assign them to specific biochemical proteins. Since artificial lipid bilayers became models for biological membranes [2], various methods have been tried to reconstitute ion channel proteins into bilayers. Promising is the fusion of lipid vesicles containing a protein with bilayers made from suitable lipids [3,4] or below the lipid phase transition temperature [5]. Acetylcholine receptor channels [5], a protein from sarcoplasmic reticulum [3] and Paramecium cilia channels [6] were reconstituted by this method. Reconstitution allows the study of biochemically defined proteins and much freedom in the choice of lipids, and so holds promise for biochemical assays and the study of
Presented 1983.
l
at the 7th International
0302-4598/84/$03.00
Symposium
0 1984 Elsevier Sequoia
on Bioelectrochemistry,
S.A.
Stuttgart
(F.R.G.),
18-22
July
330
lipid-protein interactions. Yet some drawbacks limit the technique’s usefulness. Planar bilayer membranes span holes in hydrophobic Teflon foils whose diameter cannot be made much smaller than 50 pm. The capacitance of such large membranes leads to electrical noise and limits current resolution. Solvent-free bilayers are fragile and exhibit microphonics. Moreover, really pure lipid membranes are impossible to make because the Teflon foil must be treated with a non-volatile hydrocarbon, e.g. hexadecane. Such solvents diffuse into and modify the lipid membrane. To overcome these limitations, we have developed the methods described in this paper to form artificial lipid bilayer membranes at the tips of glass patch pipettes. MATERIALS
AND
METHODS
Synthetic lecithins 1-oleoyl-2-palmitoyl-3-phosphatidylcholine (OPPC) and lstearoyl-2-myristoyl-3-phosphatidylcholine (1,2-SMPC) were purchased 99 % pure from Avanti Biochemicals. Soybean lipid (asolectin) was purchased from Sigma and washed with acetone before use. Soybean phosphatidylethanolamine (Soy PE) was separated from asolectin by column chromatography on silica gel and purified on DEAE. l-Stearoyl-3-myristoyl-2-phosphatidylcholine (1,3SMPC) was synthesized and kindly provided by H. Eibl, Gottingen. Synthetic alamethicin F-30 was kindly provided by G. Jung, Tubingen. Phallolysin was a gift from H. Faulstich, Heidelberg, and cilia fragment vesicles from Paramecium tetraurelia were prepared and supplied by J.E. Schultz, Tubingen. Salts, reagents and solvents were of analytical grade and water was distilled twice in a quartz still. All salt solutions were filtered through microfilters (Satorius SM 16529, pore size 0.2 pm). Pipettes were made in the same way as for patch clamping [l]. Both flint glass (CEEBEE hematocrit tubes) and borosilicate glass gave good results. Fire polishing of the tip is not essential, but makes bilayer formation easier. Pipettes were not pretreated or coated in any way. Sylgard-coated pipettes were not used in order to work with grease-free lipid bilayers. In the case of the droplet method, the pipette tip was stopped up irreversibly, because some components of Sylgard were probably dissolved by hexane. After filling, the pipettes were inserted into a patch clamp holder [l] with an outlet for applying suction. Electrical contact was made with Ag ]AgCl electrodes. The head stage amplifier was mounted on a hydraulic microdrive (Narishige) so that the pipette could be dipped slowly into the bath solution. Electrical apparatus was also the same as that for patch clamping [l]. We used an operational amplifier (Burr-Brown OPA103AM) in a current-voltage converter circuit with a 1 GQ feedback resistor. Current resolution was better than 0.6 pA (peak) at 1 kHz. A commercial patch clamp system (e.g. List-Medical EPC-5) would have served equally as well. To monitor bilayer formation electrically, a square wave test pulse 1 mV in amplitude was applied to the bath electrode. The resistance was measured by the amplitude of the square current response. Very high resistances require a larger test
331
pulse. Bilayer capacitance was measured by the amplitude of the transient current spikes resulting from the test pulse. This capacitive response was calibrated using known test capacitors. The voltage sign convention is as follows. The cis side is that to which the channel-forming preparation was added, and applied potentials are referred to the truns side as ground. A positive current is a flow of cations from the cis side to ground. RESULTS AND DISCUSSION
Formation of bilayers Lipid membranes on glass micropipettes were first reported by Mueller [7] and then by Andersen and Muller [8], who used pipettes to excise portions of decanecontaining bilayers. For several reasons, we decided not to follow this approach. First, our goal was to form solvent-free lipid bilayers. Secondly, the method should be simple and not involve bent glass pipettes or the problem of contacting a preformed membrane. Finally, we wanted to obtain bilayers directly on the pipette without first making a conventional planar lipid film. Tank et al. [9] have applied the patch clamp technique directly to large liposomes made by the freeze-thaw method with Torpedo electroplax proteins. This ensures solvent-free membranes but is restricted to lipids that form large stable vesicles and to proteins that are not denatured by the freeze-thaw procedure. We wanted to find a more generally applicable method. We have found it surprisingly easy to make lipid bilayers directly at the tip of patch pipettes. Following our preliminary reports [lO,ll], other groups also have used patch pipettes in bilayer studies [12-141. Here, we describe the procedures that we have found particularly useful. We emphasize that all of our methods involve no preconditioning with non-volatile alkanes, which is needed when Teflon septa are used. (a) Making bilayers from a lipid solution A simple and reliable method is the droplet technique, shown in Fig. 1. The lipid is dissolved in a volatile solvent such as hexane (typically l-2 mg lipid per cm3 of solution). The pipette is dipped slightly into the salt solution and lo-20 mm3 of lipid solution are allowed to run down the pipette shaft. The lipid spreads out and also seals the pipette tip. The solvent rapidly diffuses into the water phase and in a few minutes evaporates. The pipette is left closed with almost solvent-free lipid, which thins out into a lipid bilayer. This can occur spontaneously, but is usually induced by gentle suction on the pipette holder. Bilayer formation is observed by an increasing capacitance signal (Fig. lb). The bilayer capacitance is much larger than expected from the area of the pipette opening and can vary over a wide range. Assuming a specific bilayer capacitance of 0.8 pF/cm2, the membrane areas he between 10 and 300 pm2 or more. Thus, the bilayers form some distance inside the pipette (in some cases this is actually visible
332
(b)
suction 2pF I
Fig. 1. (a) Method for obtaining a bilayer from a lipid solution. From left to right: the pipette is dipped into the bath solution; a droplet of hexane llipid solution is spread on the surface, sealing the pipette; the solvent has evaporated, leaving a monolayer on the bath surface and a lipid droplet sealing the pipette; with gentle suction, the lipid thins out into a bilayer. (b) The development of the bilayer is monitored by the capacitance increase in response to suction. The final value of 2.5 pF corresponds to a bilayer area of 300 um2.
under a microscope). This protected position of droplet bilayers gives them excellent mechanical stability. The pipette can be immersed deeply or even removed from the bath without damaging the membrane. However, since the pipette contacts the lipid solution we cannot claim with certainty that the bilayers are solvent-free. Even highly volatile solvents (e.g. pentane) may become trapped in the hydrophobic bilayer core, in very small amounts. With a very high yield (> 80 %), bilayers are stable for hours before breaking. The seal resistance amounts l-10 GLI. Usually the membranes tolerate voltages up to 400 mV without showing bursts of spike-like current events. (b) Making bilayers from a surface monolayer The risk of solvent residues is reduced or entirely eliminated in the contact method shown in Fig. 2. A solvent-free lipid monolayer is made on the surface of the bath containing the pipette tip. The pipette is removed briefly from the bath and lowered until it gently touches the surface. As the pipette tip passes through the water surface it becomes coated with a monolayer of lipid molecules. By gentle contact of the two monolayers a bilayer is formed. Two variants of this method differ only in the manner in which the monolayer is made. One way is to spread lipid solution on the bath surface as before, but with the pipette immersed too deeply to contact the lipid solution. The solvent evaporates, leaving a lipid monolayer on the surface. Such monolayers are virtually solvent-free,
333
contact
1
---_l-
I’“’
10ms
Fig. 2. (a) The contact method of forming solvent-free bilayers. From left to right: first, a monolayer of lipid is made on the bath surface; lifting the pipette from the bath causes a monolayer to cover the pipette tip also; by carefully bringing together the two monolayers, a bilayer is formed: repeating the procedure and applying gentle suction leads to more stable bilayers. (b) Capacitance signal during formation of a bilayer by the contact method. Before contact, small spikes due to parasitic capacitance are seen. After contact, a larger capacitance, composed of the pipette and bilayer capacitances, is measured.
but residual traces of solvent cannot be excluded in principle. It is therefore better to obtain the monolayer from liposomes suspended in the bath solution. Since liposomes can be made without any solvents, bilayers made by this route are completely solvent-free. Because the surface monolayer is continuously regenerated, bilayers are stabilized and form more readily than from solvent-spread monolayers. In contrast to Tank et al. [9], the liposomes do not need to be large or well formed, and sonicated and subsequently freeze-thawed vesicles (diameters 0.1-1.0 pm) suffice to establish a suitable monolayer on the bath surface. The capacitances of contact bilayers are small compared to the pipette capacitance and so cannot be measured accurately. This suggests that these bilayers form at the extreme tip of the pipette. Although just as simple in principle, the contact method is more demanding than the droplet procedure. Contact must be made gently and the tip may not be immersed deeper than a few pm into the bath. Therefore, a good micromanipulator must be used and the apparatus must stand on a vibration-free table. On a fresh pipette, solvent-free contact bilayers form reluctantly and the first two or three attempts frequently do not seal the pipette. Thereafter, bilayers form more readily. Repeating the procedure several times, and applying gentle suction leads to more stable bilayers as a coat of lipid is gradually built up on the tip. The bilayers break if they are pushed into the subphase more deeply than a few pm. Thus, in several minutes, evaporation can lower the bath surface sufficiently to lose contact, and the pipette must occasionally be adjusted downwards in compensation. Moreover, the bilayer temperature can differ from that of the bulk liquid. Nevertheless, the contact
334
method has important advantages. Due to their small area, the bilayers give good electrical resolution comparable with the patch clamp, and the bilayers can be guaranteed to be completely solvent-free. The stability of the bilayers is not as good as that of the bilayers obtained by the droplet method. As a consequence of evaporation, the electrolyte interface drops and contact between the pipette tip and water interface is interrupted. During this process of contact loosening, bursts of spike-like current events are observed. (c) General remarks Lipid bilayers on patch pipettes show excellent mechanical and electrical stability. In general, droplet bilayers are even more stable than those made by the contact method. This is due to their protected position inside the pipette, but may also be attributed to traces of residual solvent. Droplet bilayers show more electrical noise because of their larger capacitance. The specific resistance of a pure lipid bilayer is typically 10’ Q cm2 (in 1 M NaCl). The resistance of a bilayer membrane 10 pm2 in area would therefore be far too large to measure with our equipment. The observed resistance can vary instead from several GO up to the resistance of the glass pipette itself. We attribute this to leakage currents associated with the glass-lipid seal. An electrically stable membrane can withstand large potentials without electrical breakthrough. Although the breakdown voltage of good solvent-free bilayers is generally held to lie between 150 and 250 mV, bilayers on patch pipettes are often stable to 500 mV or more [8]. This suggests that electrical breakthrough generally occurs at the membranes’ rim or at defects, and that, intrinsically, lipid bilayers may be able to withstand much higher potentials. In contrast to patch clamping on biological membranes [l], pipettes can be used more than once. If a bilayer breaks, another is made by simply raising and lowering the pipette, provided that a sufficient monolayer still covers the bath surface. Otherwise, more lipid must be spread first. To ensure clean electrical measurements, the pipette must be checked to be open before the bilayer is made. This is done by observing the square current pulses following the test signal. If bilayers are leaky or the open pipette conductance is much reduced, this is usually due to lipid build-up. If brief application of strong suction or overpressure does not clear the opening, a fresh pipette must be used. Just as with conventional solvent-free bilayers, cleanliness is decisive. Traces of surface-active contaminants cause spurious leakage currents or prevent bilayer formation. Fortunately, the newly made pipette surface is certainly clean and the bath solution can be kept in a small disposable glass cup. Cleanliness is therefore much easier to maintain than with a conventional bilayer set-up involving Teflon foils and reusable cuvettes. Solutions must be prepared carefully because small particles or dust can obstruct the pipette tip or, just as in patch clamp work, prevent a good glass-membrane seal. Also, bacteria and fungal cells must be excluded from the pipette since they can secrete pore-forming substances [15]. If such cells drift down onto the bilayer, their secretions may become incorporated and lead to
335
unwanted channel before use. Reconstitution
fluctuations.
For
these
reasons
all solutions
must
be filtered
of ion channels
(a) Incorporation of water-soluble peptides Small water-soluble peptides are best introduced from the aqueous phase. They diffuse to the bilayer and partition into it. Peptides can be added to the bath or to the pipette solution. Addition to the bath allows prior study of the unmodified bilayer and the concentration of peptide is easily varied during the experiment. This is more difficult if the peptide is added to the pipette solution. However, less peptide is usually required on the pipette side. Here, we illustrate both options with alamethicin added to the pipette solution and phallolysin incorporated from the bath. The peptide antibiotic alamethicin forms distinctive voltage-dependent multi-level pore fluctuations in bilayer membranes. Following the addition of a small amount of alamethicin to the bath, these fluctuations quickly appear in pipette bilayers. Their appearance, amplitude and kinetics are just as in conventional bilayers under comparable conditions. Because alamethicin exhibits these characteristic pore states only in true bilayer lipid membranes, this proves that the pipettes are indeed sealed by a single lipid bilayer and not by other unspecified lipid agglomerations. In analysing alamethicin pore properties, the enhanced current resolution of the pipette method allows the study of sub-levels or inhomogeneities in the conductance values of the individual pore-level states [16]. Also, due to the small membrane area, single pore fluctuations persist under conditions where normally many additional pores would become active, making it possible to observe the tenth and eleventh levels of a single alamethicin pore. For the recording shown in Fig. 3, alamethicin was added to the pipette solution only, as in previous experiments on biological muscle-cell ‘membranes [17]. The pipette contained 10 mm3 of solution with only 1 ng of alamethicin, illustrating that very small amounts of peptide are needed for pipette bilayer experiments. Also, only 50 mm3 of lipid solution, containing 25 pg of l,ZSMPC, was required to form the bilayer.
250 pA
Fig. 3. Alamethicin pore fluctuation in a bilayer made by the droplet method from 1,2SMPC. The salt solution was 1 M KCI buffered with 10 mM Hepes to pH 7.2. Synthetic alamethicin was added to the pipette filling solution at a concentration of 0.1 ag/cm3. The applied potential was 100 mV on the pipette side (pipette outward current positive) and the temperature was 25 QC, which is below the lipid transition temperature t, = 33 o C of 1,2-SMPC.
336
*n.
*I
1-1
5pA 200
ms
Fig. 4. Current fluctuations of the phallolysin channel in a 1,3-SMPC bilayer. The salt solution was 1 M KCI buffered with 10 mM Hepes to pH 7.2; phallolysin concentration, 0.16 ag/cm3 in the bath. A potential of 72 mV was applied to the bath side (pipette inward current positive). The temperature was 22 OC (the transition temperature of 1,3-SMPC is ca. 30 “C).
As a larger water-soluble protein, we’have incorporated phallolysin, a channelforming mushroom toxin from Amanita phalloides [18] with a m.w. of ca. 30000 g/mol. After bilayer formation, the peptide was added to the bath solution at a concentration of 5 nM. The resulting current fluctuations in Fig. 4 are similar to those in conventional bilayers. (b) Biological channel proteins Large hydrophobic channel proteins are not water-soluble and so cannot be reconstituted from the water phase. However, channel proteins incorporated into lipid vesicles will be present in a lipid film formed from such vesicles on the bath surface. A bilayer made from this film can then contain the reconstituted protein [19]. In the contact method, any proteins present on the surface can become incorporated into the bilayer. This method of reconstitution was used for acetylcholine receptor channels [13]. In general, contact with the air interface may modify or denature channel-forming proteins although some integral membrane proteins at the surface may be protected from air contact by excess lipid in a multiple layer. The method of incorporation least likely to damage proteins is direct fusion of proteincontaining vesicles with the bilayer. If the bilayer is made on a patch pipette, the vesicles can be present in the pipette-filling solution or added to the bath after the bilayer is made. It seems particularly convenient to put the protein-containing vesicles into the pipette. In conventional bilayer fusion experiments, a problem is to ensure contact between the vesicles and the bilayer. Next to the membrane, there is an unstirred layer which the vesicles must traverse by Brownian motion. When loaded osmotically to encourage fusion, the vesicles are denser than the surrounding solution and tend to sink to the bottom of the measuring chamber rather than diffuse to the bilayer. With pipettes, this is no problem since the vesicles will drift naturally down onto the preformed bilayer and, if conditions are otherwise appropriate, fuse with it. To exhibit a biological channel incorporated by fusion into bilayers on patch
337
1PA -
1s
Fig. 5. Cilia channels from Paramecium tefruurelia incorporated by fusion into a 1,3-SMPC bilayer. The salt solution was 100 mM KCI, 1 mM CaCI,, 1 mM MgCI,, buffered with 10 mM Hepes to pH 7.4. Cilia fragments incubated with 100 m M additional sucrose were added to the pipette solution before bilayer formation. The bilayer was made by the contact method. After spreading of the lipid monolayer, ca. 30 min was allowed to ensure complete evaporation of the solvent before the bilayer was made. A potential of 50 mV was applied to the pipette; temperature 22 o C. The total amount of cilia protein in the pipette was only 4 ng, illustrating the sensitivity of this method.
pipettes, we present recordings of the cationic channel from Paramecium tetraurelia cilia [20]. We have previously incorporated this channel into bilayers made from 1,3-SMPC below its phase transition temperature, which are known to be very suitable for fusion with lipid vesicle preparations [6]. A 1,3-SMPC bilayer below its phase transition temperature was made at the tip of a patch pipette containing osmotically loaded cilia fragment vesicles. Figure 5 shows the resulting current fluctuations which closely resemble those of the same channel in conventional planar bilayers [6]. Because no cilia fragment vesicles were in the bath, it can be assumed that incorporation was by fusion with the 1,3-SMPC bilayer. The total amount of cilia proteins in the pipette was only 4 ng, again illustrating the great sensitivity of pipette reconstitution. We have also incorporated the cilia channel into bilayers made by the contact method from a Soy PE/OPPC vesicle suspension. Similar lipid mixtures are known to be highly favourable for liposome fusion. As shown in Fig. 6, channel fluctuations were again seen closely resembling those found previously. (c) Discussion We have shown that protein-containing ported bilayers under suitable conditions.
vesicles can be fused into pipette-supUnfortunately, so far we cannot control
I
2~A
Fig. 6. Cilia channels incorporated into a bilayer made by the contact method. The salt solution was 100 mM KCI, 1 m M CaCI,, buffered with 5 mM Hepes to pH 7.2. The bath solution contained 5.5 mg/cm3 of lipid as freeze-thawed vesicles made from Soy PE/OPPC in a ration of 70/30. Cilia fragment vesicles were added to the pipette solution only. A potential of 100 mV was applied to the pipette. The temperature was 23 o C, which is above the main phase transition temperature of the lipid components.
338
the number of fusion events and with time the number of channels in a bilayer can increase until it finally ruptures. This problem is opposite to that encountered with conventional planar bilayers, where fusion is hard to initiate but easily suppressed by perfusion of the cuvette [3]. It is not certain that protein incorporation into pipette bilayers is always by direct fusion with the bilayer. From fusion rates with conventional bilayers, fusion with a pipette bilayer should be a quite rare occurrence in view of the small membrane area. Prolonged and intensified contact of the vesicles with the bilayer may explain the improved fusion rates. But it is also possible that proteins become incorporated into a lipid film on the glass surface, which presumably is also of bilayer character, and then diffuse laterally into the bilayer membrane. Since it is easy to incorporate channels uia the surface monolayer, this possibility cannot be excluded if proteincontaining vesicles are present in the bath. It seems quite possible for lipids or proteins to diffuse from the surface film into the bilayer along a lipid film on the glass surface. CONCLUSION
We have shown that lipid bilayers can be formed easily on glass patch pipettes, with significant improvements over conventional bilayer techniques. Due to the small membrane area, current resolution becomes competitive with patch clamp methods. With pipette methods, measurements can be extended to extreme conditions that are difficult or impossible to attain otherwise. Stable bilayers are formed over a wide range of ionic strength, and we have observed alamethicin fluctuations in 5 mM as well as in 5 M NaCl [ll]. Pipette bilayers are mechanically less sensitive, far less susceptible to microphonics, and can withstand higher voltages. Since there is no pretreatment of the pipettes, bilayer membranes for the first time can be guaranteed to be solvent-free, ending all discussion about possible detrimental effects of solvent residues or pretreatment hydrocarbons on the reconstituted ion channels. There are also technical advantages. Pipettes are cheap and simple to reproduce. Although many membranes can be made using the same pipette, it is quickly replaced if needed. The same apparatus as that for patch clamp recording is used so that bilayer experiments can now be performed in any laboratory equipped with a patch clamp stand. Finally, the methods use very small amounts of material. In principle, it may even suffice to fill only the extreme tip of the pipette with the protein preparation and back-fill with plain salt solution. Also, only small volumes of bath solution are needed, and we have used as little as 50 mm3. This is helpful if suspensions of rare or costly lipids are used. However, in very small bath volumes, evaporation can change the salt concentration in a short time. In the past, reconstitution techniques have shown much promise, but suffered from the limitations outlined in the Introduction. The new method of making bilayers on patch pipettes overcomes most of these problems, and also is much simpler to handle while extending the range of possible applications. Particularly
339
attractive are the extremely small amounts of materials required. This should encourage biochemists working with ion channel proteins and who have hesitated to employ electrochemical reconstitution methods to turn to them now as an assay for channel protein activity. ACKNOWLEDGEMENTS
We thank Drs. H. Eibl, G. Jung, H. Faulstich and J.E. Schultz for kindly providing us with 1,3-SMPC, synthetic alamethicin, phallolysin, and Paramecium cillia fragments, respectively. This work was supported by the Minister fur Wissenschaft und Forschung des Landes Nordrhein-Westfalen (IVB4-FA9273). REFERENCES 1 O.P. Hamill, A. Marty, E. Neher, B. Sakmann and F.J. Sigworth, Pfltugers Arch., 391 (1981) 85. 2 P. Mueller and D.O. Rudin in Current Topics in Bioenergetics, D.R. Sanadi (Editor), Academic Press, New York, 1969, Vol. 3, pp. 157-249. 3 C. Miller, J. Membr. Biol., 40 (1978) 1. 4 F.S. Cohen, J. Zimmerberg and A. Finkelstein, J. Gen. Physiol., 75 (1980) 251. 5 G. Boheim, W. Hanke, F.J. Barrantes, H. Eibl, B. Sakmann, G. Fels and A. Maelicke, Proc. Natl. Acad. Sci U.S.A., 78 (1981) 3586. 6 W. Hanke, H. Eibl and G. Boheim, Biophys. Struct. Mech., 7 (1981) 131. 7 P. Mueller, Ann. N.Y. Acad. Sci., 264 (1975) 247. 8 O.S. Andersen and R.U. Muller, J. Gen. Physiol., 80 (1982) 403. 9 D.W. Tank, C. Miller and W.W. Webb, Proc. Nat]. Acad. Sci. U.S.A., 79 (1982) 7749. 10 G. Boheim, W. Hanke and G. Jung in Proceedings of the 4th U.S.S.R./F.R.G. Symposium on Chemistry of Peptides and Proteins, Tuebingen, June 1982, in press. 11 U. Wilmsen, C. Methfessel, W. Hanke and G. Boheim in Physical Chemistry of Transmembrane Ion Motions (Proceedings of the 36th International Meeting of the Socitte de Chimie Physique, Paris, 1982), G. Spach (Editor), Elsevier, Amsterdam, 1983, pp. 479-485. 12 T. Schuerholz and H. Schindler, FEBS Lett., 152 (1983) 187. 13 B.A. Suarez-Isla, K. Wan, J. Lindstrom and M. Montal, Biochemistry, 22 (1983) 2319. 14 R. Coronado and R. Latorre, Biophys. J., 41 (1983) 56a. 15 S.J. Schein, B.L. Kagan and A. Finkelstein, Nature (London), 276 (1978) 159. 16 G. Jung, G. Becker, H. Schmitt, K.P. Voges, G. Boheim and S. Griesbach in Peptides, Structure and Function, Proceedings of the 8th American Peptide Symposium, V.J. Hruby and D.H. Rich (Editors), Pierce Chem. Comp., Rockford, IL, 1983, pp. 491-494. 17 B. Sakmann and G. Boheim, Nature (London), 282 (1979) 336. 18 H. Faulstich, S. Zobeley and M. Weckauf-Bloching, Hoppe-Seylers Z. Physiol. Chem., 355 (1974) 1495. 19 H. Schindler and U. Quast, Proc. Nat]. Acad. Sci. U.S.A., 77 (1980) 3052. 20 G. Boheim, W. Hanke, C. Methfessel, H. Eibl, U.B. Kaupp, A. Maelicke and J.E. Schultz in Transport in Biomembranes: Model Systems and Reconstitution, R. Antolini, A. Gliozzi and A. Gorio (Editors), Raven Press, New York, 1982, pp. 87-97.
Bioelectrochemistry and Bioenergetics, 12 (1984) 329-339 A section of J. Electroanal. Chem., and constituting Vol. 173 (1984) Elsevier Sequoia S.A., Lausanne - Printed in The Netherlands
651 --ION CHANNEL RECONSTITUTION INTO LIPID BILAYER MEMBRANES ON GLASS PATCH PIPETTES l
W. HANKE,
C. METHFESSEL,
Arbeitsgruppe Biophysikalische D - 4630 Bochum (F. R. G.) (Revised
manuscript
received
U. WILMSEN
and G. BOHEIM
Chemie von Membranen,
December
L.ehrstuhl ftir Zellphysiologie,
Ruhr-Universitiit,
29th 1983)
SUMMARY Planar lipid bilayer membranes were formed at the tip of glass patch clamp pipettes, and ion channels reconstituted by fusion of protein-containing vesicles with the bilayer. The advantages of the method over conventional bilayer techniques are numerous and include better current resolution, improved bilayer stability, compatibility with patch clamp apparatus and simple handling. Completely solvent-free lipid bilayers without any pretreatment can be prepared and only small amounts of materials are required.
INTRODUCTION
Living cell membranes control ion flow by integral membrane proteins that form ion-selective channels. The ion currents carried by single channels can be measured by patch clamp recording or by lipid bilayer reconstitution. In the patch clamp developed by Neher and co-workers [l] a patch of cell membrane, small enough to give good signal resolution, is electrically isolated with a glass micropipette. Although the technique is very successful, channels can be identified only by their electrical and pharmacological properties and it is difficult to assign them to specific biochemical proteins. Since artificial lipid bilayers became models for biological membranes [2], various methods have been tried to reconstitute ion channel proteins into bilayers. Promising is the fusion of lipid vesicles containing a protein with bilayers made from suitable lipids [3,4] or below the lipid phase transition temperature [5]. Acetylcholine receptor channels [5], a protein from sarcoplasmic reticulum [3] and Paramecium cilia channels [6] were reconstituted by this method. Reconstitution allows the study of biochemically defined proteins and much freedom in the choice of lipids, and so holds promise for biochemical assays and the study of
Presented 1983.
l
at the 7th International
0302-4598/84/$03.00
Symposium
0 1984 Elsevier Sequoia
on Bioelectrochemistry,
S.A.
Stuttgart
(F.R.G.),
18-22
July
330
lipid-protein interactions. Yet some drawbacks limit the technique’s usefulness. Planar bilayer membranes span holes in hydrophobic Teflon foils whose diameter cannot be made much smaller than 50 pm. The capacitance of such large membranes leads to electrical noise and limits current resolution. Solvent-free bilayers are fragile and exhibit microphonics. Moreover, really pure lipid membranes are impossible to make because the Teflon foil must be treated with a non-volatile hydrocarbon, e.g. hexadecane. Such solvents diffuse into and modify the lipid membrane. To overcome these limitations, we have developed the methods described in this paper to form artificial lipid bilayer membranes at the tips of glass patch pipettes. MATERIALS
AND
METHODS
Synthetic lecithins 1-oleoyl-2-palmitoyl-3-phosphatidylcholine (OPPC) and lstearoyl-2-myristoyl-3-phosphatidylcholine (1,2-SMPC) were purchased 99 % pure from Avanti Biochemicals. Soybean lipid (asolectin) was purchased from Sigma and washed with acetone before use. Soybean phosphatidylethanolamine (Soy PE) was separated from asolectin by column chromatography on silica gel and purified on DEAE. l-Stearoyl-3-myristoyl-2-phosphatidylcholine (1,3SMPC) was synthesized and kindly provided by H. Eibl, Gottingen. Synthetic alamethicin F-30 was kindly provided by G. Jung, Tubingen. Phallolysin was a gift from H. Faulstich, Heidelberg, and cilia fragment vesicles from Paramecium tetraurelia were prepared and supplied by J.E. Schultz, Tubingen. Salts, reagents and solvents were of analytical grade and water was distilled twice in a quartz still. All salt solutions were filtered through microfilters (Satorius SM 16529, pore size 0.2 pm). Pipettes were made in the same way as for patch clamping [l]. Both flint glass (CEEBEE hematocrit tubes) and borosilicate glass gave good results. Fire polishing of the tip is not essential, but makes bilayer formation easier. Pipettes were not pretreated or coated in any way. Sylgard-coated pipettes were not used in order to work with grease-free lipid bilayers. In the case of the droplet method, the pipette tip was stopped up irreversibly, because some components of Sylgard were probably dissolved by hexane. After filling, the pipettes were inserted into a patch clamp holder [l] with an outlet for applying suction. Electrical contact was made with Ag ]AgCl electrodes. The head stage amplifier was mounted on a hydraulic microdrive (Narishige) so that the pipette could be dipped slowly into the bath solution. Electrical apparatus was also the same as that for patch clamping [l]. We used an operational amplifier (Burr-Brown OPA103AM) in a current-voltage converter circuit with a 1 GQ feedback resistor. Current resolution was better than 0.6 pA (peak) at 1 kHz. A commercial patch clamp system (e.g. List-Medical EPC-5) would have served equally as well. To monitor bilayer formation electrically, a square wave test pulse 1 mV in amplitude was applied to the bath electrode. The resistance was measured by the amplitude of the square current response. Very high resistances require a larger test
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pulse. Bilayer capacitance was measured by the amplitude of the transient current spikes resulting from the test pulse. This capacitive response was calibrated using known test capacitors. The voltage sign convention is as follows. The cis side is that to which the channel-forming preparation was added, and applied potentials are referred to the truns side as ground. A positive current is a flow of cations from the cis side to ground. RESULTS AND DISCUSSION
Formation of bilayers Lipid membranes on glass micropipettes were first reported by Mueller [7] and then by Andersen and Muller [8], who used pipettes to excise portions of decanecontaining bilayers. For several reasons, we decided not to follow this approach. First, our goal was to form solvent-free lipid bilayers. Secondly, the method should be simple and not involve bent glass pipettes or the problem of contacting a preformed membrane. Finally, we wanted to obtain bilayers directly on the pipette without first making a conventional planar lipid film. Tank et al. [9] have applied the patch clamp technique directly to large liposomes made by the freeze-thaw method with Torpedo electroplax proteins. This ensures solvent-free membranes but is restricted to lipids that form large stable vesicles and to proteins that are not denatured by the freeze-thaw procedure. We wanted to find a more generally applicable method. We have found it surprisingly easy to make lipid bilayers directly at the tip of patch pipettes. Following our preliminary reports [lO,ll], other groups also have used patch pipettes in bilayer studies [12-141. Here, we describe the procedures that we have found particularly useful. We emphasize that all of our methods involve no preconditioning with non-volatile alkanes, which is needed when Teflon septa are used. (a) Making bilayers from a lipid solution A simple and reliable method is the droplet technique, shown in Fig. 1. The lipid is dissolved in a volatile solvent such as hexane (typically l-2 mg lipid per cm3 of solution). The pipette is dipped slightly into the salt solution and lo-20 mm3 of lipid solution are allowed to run down the pipette shaft. The lipid spreads out and also seals the pipette tip. The solvent rapidly diffuses into the water phase and in a few minutes evaporates. The pipette is left closed with almost solvent-free lipid, which thins out into a lipid bilayer. This can occur spontaneously, but is usually induced by gentle suction on the pipette holder. Bilayer formation is observed by an increasing capacitance signal (Fig. lb). The bilayer capacitance is much larger than expected from the area of the pipette opening and can vary over a wide range. Assuming a specific bilayer capacitance of 0.8 pF/cm2, the membrane areas he between 10 and 300 pm2 or more. Thus, the bilayers form some distance inside the pipette (in some cases this is actually visible
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(b)
suction 2pF I
Fig. 1. (a) Method for obtaining a bilayer from a lipid solution. From left to right: the pipette is dipped into the bath solution; a droplet of hexane llipid solution is spread on the surface, sealing the pipette; the solvent has evaporated, leaving a monolayer on the bath surface and a lipid droplet sealing the pipette; with gentle suction, the lipid thins out into a bilayer. (b) The development of the bilayer is monitored by the capacitance increase in response to suction. The final value of 2.5 pF corresponds to a bilayer area of 300 um2.
under a microscope). This protected position of droplet bilayers gives them excellent mechanical stability. The pipette can be immersed deeply or even removed from the bath without damaging the membrane. However, since the pipette contacts the lipid solution we cannot claim with certainty that the bilayers are solvent-free. Even highly volatile solvents (e.g. pentane) may become trapped in the hydrophobic bilayer core, in very small amounts. With a very high yield (> 80 %), bilayers are stable for hours before breaking. The seal resistance amounts l-10 GLI. Usually the membranes tolerate voltages up to 400 mV without showing bursts of spike-like current events. (b) Making bilayers from a surface monolayer The risk of solvent residues is reduced or entirely eliminated in the contact method shown in Fig. 2. A solvent-free lipid monolayer is made on the surface of the bath containing the pipette tip. The pipette is removed briefly from the bath and lowered until it gently touches the surface. As the pipette tip passes through the water surface it becomes coated with a monolayer of lipid molecules. By gentle contact of the two monolayers a bilayer is formed. Two variants of this method differ only in the manner in which the monolayer is made. One way is to spread lipid solution on the bath surface as before, but with the pipette immersed too deeply to contact the lipid solution. The solvent evaporates, leaving a lipid monolayer on the surface. Such monolayers are virtually solvent-free,
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contact
1
---_l-
I’“’
10ms
Fig. 2. (a) The contact method of forming solvent-free bilayers. From left to right: first, a monolayer of lipid is made on the bath surface; lifting the pipette from the bath causes a monolayer to cover the pipette tip also; by carefully bringing together the two monolayers, a bilayer is formed: repeating the procedure and applying gentle suction leads to more stable bilayers. (b) Capacitance signal during formation of a bilayer by the contact method. Before contact, small spikes due to parasitic capacitance are seen. After contact, a larger capacitance, composed of the pipette and bilayer capacitances, is measured.
but residual traces of solvent cannot be excluded in principle. It is therefore better to obtain the monolayer from liposomes suspended in the bath solution. Since liposomes can be made without any solvents, bilayers made by this route are completely solvent-free. Because the surface monolayer is continuously regenerated, bilayers are stabilized and form more readily than from solvent-spread monolayers. In contrast to Tank et al. [9], the liposomes do not need to be large or well formed, and sonicated and subsequently freeze-thawed vesicles (diameters 0.1-1.0 pm) suffice to establish a suitable monolayer on the bath surface. The capacitances of contact bilayers are small compared to the pipette capacitance and so cannot be measured accurately. This suggests that these bilayers form at the extreme tip of the pipette. Although just as simple in principle, the contact method is more demanding than the droplet procedure. Contact must be made gently and the tip may not be immersed deeper than a few pm into the bath. Therefore, a good micromanipulator must be used and the apparatus must stand on a vibration-free table. On a fresh pipette, solvent-free contact bilayers form reluctantly and the first two or three attempts frequently do not seal the pipette. Thereafter, bilayers form more readily. Repeating the procedure several times, and applying gentle suction leads to more stable bilayers as a coat of lipid is gradually built up on the tip. The bilayers break if they are pushed into the subphase more deeply than a few pm. Thus, in several minutes, evaporation can lower the bath surface sufficiently to lose contact, and the pipette must occasionally be adjusted downwards in compensation. Moreover, the bilayer temperature can differ from that of the bulk liquid. Nevertheless, the contact
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method has important advantages. Due to their small area, the bilayers give good electrical resolution comparable with the patch clamp, and the bilayers can be guaranteed to be completely solvent-free. The stability of the bilayers is not as good as that of the bilayers obtained by the droplet method. As a consequence of evaporation, the electrolyte interface drops and contact between the pipette tip and water interface is interrupted. During this process of contact loosening, bursts of spike-like current events are observed. (c) General remarks Lipid bilayers on patch pipettes show excellent mechanical and electrical stability. In general, droplet bilayers are even more stable than those made by the contact method. This is due to their protected position inside the pipette, but may also be attributed to traces of residual solvent. Droplet bilayers show more electrical noise because of their larger capacitance. The specific resistance of a pure lipid bilayer is typically 10’ Q cm2 (in 1 M NaCl). The resistance of a bilayer membrane 10 pm2 in area would therefore be far too large to measure with our equipment. The observed resistance can vary instead from several GO up to the resistance of the glass pipette itself. We attribute this to leakage currents associated with the glass-lipid seal. An electrically stable membrane can withstand large potentials without electrical breakthrough. Although the breakdown voltage of good solvent-free bilayers is generally held to lie between 150 and 250 mV, bilayers on patch pipettes are often stable to 500 mV or more [8]. This suggests that electrical breakthrough generally occurs at the membranes’ rim or at defects, and that, intrinsically, lipid bilayers may be able to withstand much higher potentials. In contrast to patch clamping on biological membranes [l], pipettes can be used more than once. If a bilayer breaks, another is made by simply raising and lowering the pipette, provided that a sufficient monolayer still covers the bath surface. Otherwise, more lipid must be spread first. To ensure clean electrical measurements, the pipette must be checked to be open before the bilayer is made. This is done by observing the square current pulses following the test signal. If bilayers are leaky or the open pipette conductance is much reduced, this is usually due to lipid build-up. If brief application of strong suction or overpressure does not clear the opening, a fresh pipette must be used. Just as with conventional solvent-free bilayers, cleanliness is decisive. Traces of surface-active contaminants cause spurious leakage currents or prevent bilayer formation. Fortunately, the newly made pipette surface is certainly clean and the bath solution can be kept in a small disposable glass cup. Cleanliness is therefore much easier to maintain than with a conventional bilayer set-up involving Teflon foils and reusable cuvettes. Solutions must be prepared carefully because small particles or dust can obstruct the pipette tip or, just as in patch clamp work, prevent a good glass-membrane seal. Also, bacteria and fungal cells must be excluded from the pipette since they can secrete pore-forming substances [15]. If such cells drift down onto the bilayer, their secretions may become incorporated and lead to
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unwanted channel before use. Reconstitution
fluctuations.
For
these
reasons
all solutions
must
be filtered
of ion channels
(a) Incorporation of water-soluble peptides Small water-soluble peptides are best introduced from the aqueous phase. They diffuse to the bilayer and partition into it. Peptides can be added to the bath or to the pipette solution. Addition to the bath allows prior study of the unmodified bilayer and the concentration of peptide is easily varied during the experiment. This is more difficult if the peptide is added to the pipette solution. However, less peptide is usually required on the pipette side. Here, we illustrate both options with alamethicin added to the pipette solution and phallolysin incorporated from the bath. The peptide antibiotic alamethicin forms distinctive voltage-dependent multi-level pore fluctuations in bilayer membranes. Following the addition of a small amount of alamethicin to the bath, these fluctuations quickly appear in pipette bilayers. Their appearance, amplitude and kinetics are just as in conventional bilayers under comparable conditions. Because alamethicin exhibits these characteristic pore states only in true bilayer lipid membranes, this proves that the pipettes are indeed sealed by a single lipid bilayer and not by other unspecified lipid agglomerations. In analysing alamethicin pore properties, the enhanced current resolution of the pipette method allows the study of sub-levels or inhomogeneities in the conductance values of the individual pore-level states [16]. Also, due to the small membrane area, single pore fluctuations persist under conditions where normally many additional pores would become active, making it possible to observe the tenth and eleventh levels of a single alamethicin pore. For the recording shown in Fig. 3, alamethicin was added to the pipette solution only, as in previous experiments on biological muscle-cell ‘membranes [17]. The pipette contained 10 mm3 of solution with only 1 ng of alamethicin, illustrating that very small amounts of peptide are needed for pipette bilayer experiments. Also, only 50 mm3 of lipid solution, containing 25 pg of l,ZSMPC, was required to form the bilayer.
250 pA
Fig. 3. Alamethicin pore fluctuation in a bilayer made by the droplet method from 1,2SMPC. The salt solution was 1 M KCI buffered with 10 mM Hepes to pH 7.2. Synthetic alamethicin was added to the pipette filling solution at a concentration of 0.1 ag/cm3. The applied potential was 100 mV on the pipette side (pipette outward current positive) and the temperature was 25 QC, which is below the lipid transition temperature t, = 33 o C of 1,2-SMPC.
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*n.
*I
1-1
5pA 200
ms
Fig. 4. Current fluctuations of the phallolysin channel in a 1,3-SMPC bilayer. The salt solution was 1 M KCI buffered with 10 mM Hepes to pH 7.2; phallolysin concentration, 0.16 ag/cm3 in the bath. A potential of 72 mV was applied to the bath side (pipette inward current positive). The temperature was 22 OC (the transition temperature of 1,3-SMPC is ca. 30 “C).
As a larger water-soluble protein, we’have incorporated phallolysin, a channelforming mushroom toxin from Amanita phalloides [18] with a m.w. of ca. 30000 g/mol. After bilayer formation, the peptide was added to the bath solution at a concentration of 5 nM. The resulting current fluctuations in Fig. 4 are similar to those in conventional bilayers. (b) Biological channel proteins Large hydrophobic channel proteins are not water-soluble and so cannot be reconstituted from the water phase. However, channel proteins incorporated into lipid vesicles will be present in a lipid film formed from such vesicles on the bath surface. A bilayer made from this film can then contain the reconstituted protein [19]. In the contact method, any proteins present on the surface can become incorporated into the bilayer. This method of reconstitution was used for acetylcholine receptor channels [13]. In general, contact with the air interface may modify or denature channel-forming proteins although some integral membrane proteins at the surface may be protected from air contact by excess lipid in a multiple layer. The method of incorporation least likely to damage proteins is direct fusion of proteincontaining vesicles with the bilayer. If the bilayer is made on a patch pipette, the vesicles can be present in the pipette-filling solution or added to the bath after the bilayer is made. It seems particularly convenient to put the protein-containing vesicles into the pipette. In conventional bilayer fusion experiments, a problem is to ensure contact between the vesicles and the bilayer. Next to the membrane, there is an unstirred layer which the vesicles must traverse by Brownian motion. When loaded osmotically to encourage fusion, the vesicles are denser than the surrounding solution and tend to sink to the bottom of the measuring chamber rather than diffuse to the bilayer. With pipettes, this is no problem since the vesicles will drift naturally down onto the preformed bilayer and, if conditions are otherwise appropriate, fuse with it. To exhibit a biological channel incorporated by fusion into bilayers on patch
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1PA -
1s
Fig. 5. Cilia channels from Paramecium tefruurelia incorporated by fusion into a 1,3-SMPC bilayer. The salt solution was 100 mM KCI, 1 mM CaCI,, 1 mM MgCI,, buffered with 10 mM Hepes to pH 7.4. Cilia fragments incubated with 100 m M additional sucrose were added to the pipette solution before bilayer formation. The bilayer was made by the contact method. After spreading of the lipid monolayer, ca. 30 min was allowed to ensure complete evaporation of the solvent before the bilayer was made. A potential of 50 mV was applied to the pipette; temperature 22 o C. The total amount of cilia protein in the pipette was only 4 ng, illustrating the sensitivity of this method.
pipettes, we present recordings of the cationic channel from Paramecium tetraurelia cilia [20]. We have previously incorporated this channel into bilayers made from 1,3-SMPC below its phase transition temperature, which are known to be very suitable for fusion with lipid vesicle preparations [6]. A 1,3-SMPC bilayer below its phase transition temperature was made at the tip of a patch pipette containing osmotically loaded cilia fragment vesicles. Figure 5 shows the resulting current fluctuations which closely resemble those of the same channel in conventional planar bilayers [6]. Because no cilia fragment vesicles were in the bath, it can be assumed that incorporation was by fusion with the 1,3-SMPC bilayer. The total amount of cilia proteins in the pipette was only 4 ng, again illustrating the great sensitivity of pipette reconstitution. We have also incorporated the cilia channel into bilayers made by the contact method from a Soy PE/OPPC vesicle suspension. Similar lipid mixtures are known to be highly favourable for liposome fusion. As shown in Fig. 6, channel fluctuations were again seen closely resembling those found previously. (c) Discussion We have shown that protein-containing ported bilayers under suitable conditions.
vesicles can be fused into pipette-supUnfortunately, so far we cannot control
I
2~A
Fig. 6. Cilia channels incorporated into a bilayer made by the contact method. The salt solution was 100 mM KCI, 1 m M CaCI,, buffered with 5 mM Hepes to pH 7.2. The bath solution contained 5.5 mg/cm3 of lipid as freeze-thawed vesicles made from Soy PE/OPPC in a ration of 70/30. Cilia fragment vesicles were added to the pipette solution only. A potential of 100 mV was applied to the pipette. The temperature was 23 o C, which is above the main phase transition temperature of the lipid components.
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the number of fusion events and with time the number of channels in a bilayer can increase until it finally ruptures. This problem is opposite to that encountered with conventional planar bilayers, where fusion is hard to initiate but easily suppressed by perfusion of the cuvette [3]. It is not certain that protein incorporation into pipette bilayers is always by direct fusion with the bilayer. From fusion rates with conventional bilayers, fusion with a pipette bilayer should be a quite rare occurrence in view of the small membrane area. Prolonged and intensified contact of the vesicles with the bilayer may explain the improved fusion rates. But it is also possible that proteins become incorporated into a lipid film on the glass surface, which presumably is also of bilayer character, and then diffuse laterally into the bilayer membrane. Since it is easy to incorporate channels uia the surface monolayer, this possibility cannot be excluded if proteincontaining vesicles are present in the bath. It seems quite possible for lipids or proteins to diffuse from the surface film into the bilayer along a lipid film on the glass surface. CONCLUSION
We have shown that lipid bilayers can be formed easily on glass patch pipettes, with significant improvements over conventional bilayer techniques. Due to the small membrane area, current resolution becomes competitive with patch clamp methods. With pipette methods, measurements can be extended to extreme conditions that are difficult or impossible to attain otherwise. Stable bilayers are formed over a wide range of ionic strength, and we have observed alamethicin fluctuations in 5 mM as well as in 5 M NaCl [ll]. Pipette bilayers are mechanically less sensitive, far less susceptible to microphonics, and can withstand higher voltages. Since there is no pretreatment of the pipettes, bilayer membranes for the first time can be guaranteed to be solvent-free, ending all discussion about possible detrimental effects of solvent residues or pretreatment hydrocarbons on the reconstituted ion channels. There are also technical advantages. Pipettes are cheap and simple to reproduce. Although many membranes can be made using the same pipette, it is quickly replaced if needed. The same apparatus as that for patch clamp recording is used so that bilayer experiments can now be performed in any laboratory equipped with a patch clamp stand. Finally, the methods use very small amounts of material. In principle, it may even suffice to fill only the extreme tip of the pipette with the protein preparation and back-fill with plain salt solution. Also, only small volumes of bath solution are needed, and we have used as little as 50 mm3. This is helpful if suspensions of rare or costly lipids are used. However, in very small bath volumes, evaporation can change the salt concentration in a short time. In the past, reconstitution techniques have shown much promise, but suffered from the limitations outlined in the Introduction. The new method of making bilayers on patch pipettes overcomes most of these problems, and also is much simpler to handle while extending the range of possible applications. Particularly
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attractive are the extremely small amounts of materials required. This should encourage biochemists working with ion channel proteins and who have hesitated to employ electrochemical reconstitution methods to turn to them now as an assay for channel protein activity. ACKNOWLEDGEMENTS
We thank Drs. H. Eibl, G. Jung, H. Faulstich and J.E. Schultz for kindly providing us with 1,3-SMPC, synthetic alamethicin, phallolysin, and Paramecium cillia fragments, respectively. This work was supported by the Minister fur Wissenschaft und Forschung des Landes Nordrhein-Westfalen (IVB4-FA9273). REFERENCES 1 O.P. Hamill, A. Marty, E. Neher, B. Sakmann and F.J. Sigworth, Pfltugers Arch., 391 (1981) 85. 2 P. Mueller and D.O. Rudin in Current Topics in Bioenergetics, D.R. Sanadi (Editor), Academic Press, New York, 1969, Vol. 3, pp. 157-249. 3 C. Miller, J. Membr. Biol., 40 (1978) 1. 4 F.S. Cohen, J. Zimmerberg and A. Finkelstein, J. Gen. Physiol., 75 (1980) 251. 5 G. Boheim, W. Hanke, F.J. Barrantes, H. Eibl, B. Sakmann, G. Fels and A. Maelicke, Proc. Natl. Acad. Sci U.S.A., 78 (1981) 3586. 6 W. Hanke, H. Eibl and G. Boheim, Biophys. Struct. Mech., 7 (1981) 131. 7 P. Mueller, Ann. N.Y. Acad. Sci., 264 (1975) 247. 8 O.S. Andersen and R.U. Muller, J. Gen. Physiol., 80 (1982) 403. 9 D.W. Tank, C. Miller and W.W. Webb, Proc. Nat]. Acad. Sci. U.S.A., 79 (1982) 7749. 10 G. Boheim, W. Hanke and G. Jung in Proceedings of the 4th U.S.S.R./F.R.G. Symposium on Chemistry of Peptides and Proteins, Tuebingen, June 1982, in press. 11 U. Wilmsen, C. Methfessel, W. Hanke and G. Boheim in Physical Chemistry of Transmembrane Ion Motions (Proceedings of the 36th International Meeting of the Socitte de Chimie Physique, Paris, 1982), G. Spach (Editor), Elsevier, Amsterdam, 1983, pp. 479-485. 12 T. Schuerholz and H. Schindler, FEBS Lett., 152 (1983) 187. 13 B.A. Suarez-Isla, K. Wan, J. Lindstrom and M. Montal, Biochemistry, 22 (1983) 2319. 14 R. Coronado and R. Latorre, Biophys. J., 41 (1983) 56a. 15 S.J. Schein, B.L. Kagan and A. Finkelstein, Nature (London), 276 (1978) 159. 16 G. Jung, G. Becker, H. Schmitt, K.P. Voges, G. Boheim and S. Griesbach in Peptides, Structure and Function, Proceedings of the 8th American Peptide Symposium, V.J. Hruby and D.H. Rich (Editors), Pierce Chem. Comp., Rockford, IL, 1983, pp. 491-494. 17 B. Sakmann and G. Boheim, Nature (London), 282 (1979) 336. 18 H. Faulstich, S. Zobeley and M. Weckauf-Bloching, Hoppe-Seylers Z. Physiol. Chem., 355 (1974) 1495. 19 H. Schindler and U. Quast, Proc. Nat]. Acad. Sci. U.S.A., 77 (1980) 3052. 20 G. Boheim, W. Hanke, C. Methfessel, H. Eibl, U.B. Kaupp, A. Maelicke and J.E. Schultz in Transport in Biomembranes: Model Systems and Reconstitution, R. Antolini, A. Gliozzi and A. Gorio (Editors), Raven Press, New York, 1982, pp. 87-97.