LETTER pubs.acs.org/NanoLett
Simple Reconstitution of Protein Pores in Nano Lipid Bilayers Joanne L. Gornall,†,§ Kozhinjampara R. Mahendran,‡,§ Oliver J. Pambos,† Lorenz J. Steinbock,† Oliver Otto,† Catalin Chimerel,† Mathias Winterhalter,‡ and Ulrich F. Keyser*,† † ‡
Cavendish Laboratory, University of Cambridge, J J Thomson Avenue, Cambridge, CB3 0HE, U.K. School of Engineering and Science, Jacobs University Bremen, Campus Ring 1, 28759 Bremen, Germany
bS Supporting Information ABSTRACT: We developed a new, simple and robust approach for rapid screening of single molecule interactions with protein channels. Our glass nanopipets can be fabricated simply by drawing glass capillaries in a standard pipet puller, in a matter of minutes, and do not require further modification before use. Giant unilamellar vesicles break when in contact with the tip of the glass pipet and form a supported bilayer with typical seal resistances of ∼140 GΩ, which is stable for hours and at applied potentials up to 900 mV. Bilayers can be formed, broken, and reformed more than 50 times using the same pipet enabling rapid screening of bilayers for single protein channels. The stability of the lipid bilayer is significantly superior to that of traditionally built bilayers supported by Teflon membranes, particularly against perturbation by electrical and mechanical forces. We demonstrate the functional reconstitution of the E. coli porin OmpF and R-hemolysin in a glass nanopipet supported bilayer. Interactions of the antibiotic enrofloxacin with the OmpF channel have been studied at the single-molecule level, demonstrating the ability of this method to detect single molecule interactions with protein channels. High-resolution conductance measurements of protein channels can be performed with low sample and buffer consumption. Glass nanopipet supported bilayers are uniquely suited for single-molecule studies as they are more rigid and the lifetime of a stable membrane is on the scale of hours, closer to that of natural cell membranes. KEYWORDS: Nanopore, bilayer, OmpF, antibiotics, hybrid nanopore, single molecule
C
ell membranes are essential for compartmentalization in living cells and distribution of function. Understanding how the cell controls the translocation of molecules across lipid bilayer membranes, and the role of protein channels, has received considerable research attention due to the fundamental importance of this process. Over the past 3 decades, the standard approach to the study of the biophysical properties of these channels has been to insert them into an artificial lipid bilayer that separates two electrolyte solutions. Measuring the ion current through purified membrane proteins can provide information about a number of structural and functional properties of the channel, such as pore size and selectivity. Traditionally, planar lipid bilayers are supported across holes, tens of micrometers in diameter, in Teflon membranes.1 Bilayers supported on such large orifices are sensitive to pressure fluctuations and mechanical vibrations. There has been significant interest in developing new methods for studying protein channel properties, such as ion conductance, ion selectivity, and voltage gating, which have advantages over traditional planar bilayer techniques. Significant advances in stability of the lipid bilayer have been achieved by sandwiching the bilayer between polymer gels; however, temporal resolution is limited in these systems because the analyte must diffuse through the gel to be detected.2,3 Several investigators have developed techniques for producing lipid bilayers on apertures significantly smaller than that formed in traditional r 2011 American Chemical Society
Teflon membranes. Bilayer stability can be enhanced by decreasing the area over which the bilayer spans. This has the additional advantage that the current noise is reduced, since the capacitance of the bilayer is decreased. Kitta et al. developed a method for producing Teflon membranes with micrometer-sized pores that could support lipid bilayers that were stable at high voltages.4 Planar glass chips containing micrometer-sized pores have been used to support planar lipid bilayers with gigaohm resistances.5,6 Glass nanopore membranes,7,8 the surfaces of which are chemically modified with (3-cyanopropyl)dimethylchlorosilane, have also been used to support lipid bilayers for single ion-channel recordings, with extremely high seal resistances (>teraohm). Micrometersized pores in silicon nitride membranes are able to successfully support artificial lipid bilayers,9 and hybrid nanopores have been created by inserting a single, preassembled, R-hemolysin pore alone into a nanopore in a silicon nitride membrane, with no lipid bilayer.10 However, fabrication of silicon nitride nanopores with precise dimensions is challenging, requiring use of a transmission electron microscope to ablate the surface.10 Since its invention in the 1970s, patch-clamping has been used to study ion-channels in their native state.11,12 In the last 10 years a number of techniques have been developed for incorporating Received: May 20, 2011 Revised: July 8, 2011 Published: July 13, 2011 3334
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Nano Letters protein channels in a functional state into giant unilammelar vesicles (GUVs).5,1315 By incorporation of protein channels into GUVs, protein channels in artificial bilayers can be studied using patch-clamping techniques.16,17 Typically patch-clamp pipets are 13 μm in diameter. To form an artificial bilayer, the pipet tip is carefully apposed to the surface of a GUV, which is anchored to a surface, using a micromanipulator and gentle suction is applied to the back of the pipet to form a seal.15 If one wishes to form an excised patch, in order to have access to both sides of the lipid bilayer, the pipet must be carefully withdrawn from the surface of the GUV, quickly passing the tip through an air/water interface if necessary.17 A simple, rapid and reusable method for the formation of stable, planar, lipid bilayers would greatly increase the efficiency of the characterization of protein channels in the bilayers. In this work we report the use of glass nanopipet supported lipid bilayers for ion channel recordings. Glass nanopipets are fabricated using a laser pipet puller, typically used to make microelectrodes, patch pipets, and microinjection needles from glass capillaries. In contrast to other methods, glass nanopipets can be made in a matter of minutes, without requiring use of clean room facilities and transmission electron microscopes. Glass nanopipets do not require further modification before use, such as surface coating. Simply by tuning the parameters of the pipet puller, glass nanopipets can be fabricated with a range of tip diameters from several micrometers to as small as 45 nm.18 We are able to form supported lipid bilayers using glass nanopipet tips with diameters as small as ∼230 nm, corresponding to an open area of 0.04 μm2, which is 100000 times smaller than traditional methods using Teflon membranes1 and 60 times smaller than patch clamping techniques.16,17 The open area of the glass nanopipet tip, over which the bilayer spans, is comparable to that of glass nanopore membranes,7,8 paralene nanopore membranes,19 and glass chips.5 Bilayers formed using glass nanopipets offer several advantages compared to those formed by traditional planar lipid bilayer techniques and patch clamping.1,16,17 To form an artificial bilayer, GUVs are simply added to the bath solution; they are free in solution and do not need to be anchored to a surface. Gentle suction is applied to the back of the pipet and the vesicles are drawn toward the tip. When GUVs come into contact with the tip of the glass pipet, they break and readily form a bilayer. In comparison to conventional patch clamping, the level of motor skill and handeye coordination necessary for formation of a bilayer is significantly reduced, since the nanopipet does not need to be apposed to the surface of an immobile GUV. After addition of GUVs to the system, a bilayer can be formed within seconds, producing high-resolution ion-current recordings, with typical seal resistances of ∼140 GΩ. As we show here, our glass nanopipets are also reusable; bilayers can be formed more than 50 times using the same glass nanopipet without cleaning, allowing for rapid screening of bilayers for single protein channels. Bilayers formed by this approach are stable, even at high transmembrane voltages of a few hundred millivolts, enabling longterm recordings lasting several hours. The formation of artificial lipid bilayers using glass nanopipet tips is very simple, reliable, efficient, and low cost. Here, we have established three methods of protein insertion into lipid bilayer and successfully recorded single channel recordings of membrane protein OmpF and R-hemolysin. OmpF is well-characterized in terms of structure and channel activity20 and is, therefore, used as our model system. OmpF is a
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nonspecific transport channel that allows for the passive diffusion of small, polar molecules (600700 Da in size) through the cell’s outer membrane. To demonstrate that our method is applicable to other membrane proteins, we inserted R-hemolysin into nanopipet supported bilayers, a protein that has been extensively studied in order to elucidate the mechanisms of peptide and DNA translocation.21 Materials and Methods. Borosilicate glass capillaries (Hilgenberg GmbH, Germany) were used for all experiments, with an outer diameter of 0.5 mm and a wall thickness of 0.064 mm. Pipets were prepared by drawing the glass capillaries with a laser pipet puller (P-2000, Sutter Instruments). Prior to use, the capillaries were thoroughly cleaned by sonicating in acetone and ethanol for 5 min. Residual ethanol from the cleaning process was removed with gaseous nitrogen. Following cleaning, the capillary was mounted in the laser pipet puller where the glass was heated and pulled to form two virtually identical pipets. By tuning the parameters of the pull, such as temperature and velocity, we produced nanopipets with tip diameters between ∼230 and ∼785 nm. In order to verify the shape and diameter of the tip, pipets were imaged using scanning electron microscopy (SEM). Prior to visualization in the SEM, the nanopipets were coated with a 10 nm thick layer of palladium/gold (Pd/Au). GUVs were prepared by electroformation22 in an indium tin oxide (ITO)-coated glass chamber connected to the Nanion Vesicle Prep Pro setup (Nanion Technologies, Munich, Germany). The ITO layers on the two glass slides are electrically conductive and therefore serve as electrodes. A 5 mM solution of 1,2diphytanoyl-sn-glycero-3-phosphatidylcholine (DPhPC; Avanti Polar Lipids, Alabaster, AL) with 10% cholesterol in chloroform was deposited onto the ITO-coated slides, and the chamber was filled with a 1 M solution of sorbitol (Sigma-Aldrich) in ddH2O. Electroformation was controlled by the Vesicle Prep Pro setup, and all parameters for the electroformation were programmed in the VesicleControl software (Nanion Technologies, Munich, Germany). Typical values of the amplitude, frequency, and duration of the potential applied across the chamber were 3 Vpp, 5 Hz, and 2 h, respectively. Vesicle preparation was performed at 37 °C. The formation of GUVs was highly reproducible and by adjusting the concentration of lipids used, GUVs may be formed with diameters in the range 1100 μm. Typically, the diameter of the GUVs in solution required to form a bilayer on the glass pipet was between 5 and 50 μm. Purified wild-type OmpF (1.5 mg/mL) in 1% n-octyl-polyoxyethylene (octyl-POE; Bachem, Bubendorf, Switzerland) was reconstituted into GUVs as described previously.6 The reconstitution of a single hydrophobic membrane protein is more complex than the reconstitution of water-soluble peptides.23 Our protocol for micellar insertion of membrane proteins into lipid bilayers involves the removal of detergents. In the case of nanopipet supported lipid bilayers, addition of even small quantities of detergent leads to immediate disruption of the membrane (see Figure 1 in the Supporting Information). Therefore, following incubation of the GUVs with the porin solution, octyl-POE was removed using Bio-Beads (Bio-Rad, Munich, Germany). The mixture was incubated at 4 °C overnight, and the Bio-Beads were removed afterward by centrifugation. Direct insertion of membrane proteins into GUVs is challenging, and the protocol needs to be optimized for each individual protein due to the relative instability of GUVs. The average 3335
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Figure 1. (a) Experimental setup. (b) Typical borosilicate glass capillary after pulling with a laser pipet puller. The nanopipet was coated with a 10 nm thick Pd/Au layer to prevent charging effects. The inner diameter of this tip is 243 nm.
number of proteins in each GUV can be efficiently optimized by the varying protein concentration and the time of incubation. Proteo-GUVs were used directly for lipid bilayer formation, and when kept at 4 °C, the proteo-GUVs could be used for over a week successfully. Since R-hemolysin is soluble in water, its incorporation into the lipid bilayer was achieved by adding the protein to the solution and mixing. The experimental setup is shown in Figure 1a and is based around a custom-built inverted microscope which allows the vesicles and nanopipet to be imaged during the experiment. Single-channel current measurements were performed using an Axopatch 200B amplifier (Axon Instruments, Foster City, CA) in voltage clamp mode. The signal was filtered using a four-pole low-pass Bessel filter at a frequency of 2 kHz and sampled at 10 kHz. The signals were acquired with a NI-PCIe-6251 card (National Instruments, USA), and data were recorded using custom written LabVIEW code. Pipets and the surrounding bath were filled with a buffer solution containing 150 mM KCl and 10 mM MES (pH 6). Chlorinated (Ag/AgCl) silver electrodes (200 μm diameter) were placed in the nanopipet and bath. Ag/AgCl electrodes were prepared by chlorination of bare silver wires in a 1 M KCl solution at 1.5 V for 20 s. For bilayer measurements the bath electrode was defined as the ground. The nanopipet was attached to the headstage (CV203BU, Axon Instruments) of the amplifier via an adaptor and attached to a micromanipulator (PatchStar Micromanipulor, Scientifica). This allows for precise control of the nanopipet with an accuracy of 100 nm. Before each experiment, the electrode offset was set to zero, and the nanopipet was tested for stable currentvoltage characteristics. In order to from a lipid bilayer, 10 μL of the vesicle solution was pipetted into the bath solution and a negative
Figure 2. (a) Currentvoltage curves for open 229 nm (O) and 784 nm (0) diameter pipet tips in 150 mM KCl 10 mM MES buffer at pH 6.0 (right axis) and for a 784 nm tip sealed by a DPhPC bilayer (b, left axis). The open tip resistances are 24 and 6.3 MΩ for the 229 and 784 nm pipet tips, respectively. The sealed bilayer tip resistance is 653 GΩ in this case. Note that the axis on the left is scaled by a factor of 102 as compared to the right axis. Error bars represent 1 standard deviation from the mean value. The inset shows a current trace for a sealed pipet tip held stably at 10 mV for over 10 h. (b) Cumulative frequency distribution of the time taken to complete a form, break, and re-form cycle for 50 successive bilayers formed on a single nanopipet tip. The distribution is fit with a Weibull distribution (solid line), which is differentiated to give the probability density function shown inset. The PDF peaks at a cycle time of 6.9 s.
pressure of approximately 1 Pa was applied to the back of the nanopipet, using the attached syringe to draw the vesicles to the orifice of the nanopipet. When a vesicle came into contact with the tip of the capillary, it broke and a bilayer with a high seal resistance formed immediately. Characterization of Nanopipets and Bilayer Formation. An SEM image of a typical ∼230 nm nanopipet is shown in Figure 1b. An analysis of the diameters of the tips using ImageJ showed the glass nanopipets used here had tip diameters between 229 ( 24 nm and 784 ( 83 nm depending on the pulling parameters used. Before each experiment the open pipet response was checked for linear currentvoltage characteristics. The ionic current between the Ag/AgCl electrodes was measured as the voltage is increased stepwise. At each voltage step, 30000 current measurements were taken with 10000 samples/s. The results for open pipets with tip sizes of 229 and 784 nm are 3336
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shown in Figure 2. The average open pipet resistance was found to be 6.5 ( 0.7 and 23.5 ( 0.7 MΩ for the 784 and 229 nm diameter pipet tips, respectively. It is possible to estimate the inner diameter, di, of the pipet tip by using a simplified equation for the conductance of a conical nanocapillary18,24 di ¼
4Gl πgdb
ð1Þ
where l is the length of the conical part of the pipet (2 mm), db the diameter of the back of the pipet before it starts to converge to form the tip (0.372 mm), g is the specific conductance of the buffer used (1.7 S/m), and G is the measured conductance of the pipet. This equation gives an inner diameter of 600 nm for the largest capillaries used and 170 nm for the smallest. Equation 1 is, therefore, in good agreement with the tip diameters determined by SEM. On application of a negative pressure (∼1 Pa) to the back of the pipet, GUVs are drawn toward the tip where they break and form a bilayer with a high seal resistance, which develops almost immediately. The sealed pipet response is shown in Figure 2a. The average seal resistance was found to be 139 ( 49 GΩ for the largest tips used, based on nine independent measurements. Maintaining the bath and pipet solutions at an acidic pH between 5.5 and 6 was found to promote bilayer formation, as observed for patch-clamp recording measurements.25 H + ions increase the adhesion forces between lipid bilayer and glass by screening the electrostatic repulsion between the negatively charged lipids and glass, resulting in tighter seals between bilayer and glass.26 Decreasing the pH also promotes the spreading of the lipid bilayer up the inner surface of the pipet, increasing the surface area of the seal.26 In addition, there exists a thin layer of water molecules between the lipid bilayer and the glass, several molecules thick which promotes spreading of the lipid bilayer up the walls of the pipet.26 We would also like to emphasize, that the bilayers are stable up to voltages of 900 mV (Figure 2a), which demonstrates the robustness of our simple approach. In addition, it is possible to keep stable bilayers for up to 10 h, as shown by the inset in Figure 2a. Until now, the only limiting factor for bilayer stability was the evaporation of the bath solution in our open setup configuration. Having established the characteristics of our system, we discuss now the reproducibility of the bilayer formation. Each lipid bilayer seal can be broken by applying a positive pressure to the back of the pipet using the attached syringe, some requiring applying a voltage of up to 1.3 V in addition. It is important to note, that the bilayer can then be easily re-formed by applying a negative pressure to draw another vesicle to the tip of the pipet. In order to determine how quickly successive bilayers can be screened for protein channels and how many can be formed on a single nanopipet tip, the process of applying negative pressure to form a bilayer and then positive pressure to break the bilayer was repeated continuously. Figure 2b shows the cumulative frequency distribution of the time taken to form, break, and re-form a bilayer. A small potential, of the order of a few millivolts, was applied and successful bilayer formation is ascertained by observing current level drop to zero and flow ceasing through the capillary. The cycle of forming and breaking of the bilayer can be repeated more than 50 times with the same pipet. The distribution is fit with a Weibull distribution, since the distribution is positively skewed due to a small number of cycles with a long completion time, most likely due to a depletion of
Figure 3. (a) Currentvoltage curve for a 784 nm pipet sealed using GUVs containing the E. coli porin OmpF (150 mM KCl, 10 mM MES, pH 6). The conductance, determined from the slope of the iV curve, is 0.8 ( 0.01 nS, consistent with a single trimeric OmpF channel. Error bars represent 1 standard deviation from the mean value. (b) Measured conductance of 30 successive bilayers formed on a single nanopipet tip using proteo-GUVs containing a low concentration of OmpF. The horizontal shading is a guide to the eye to indicate typical range of conductance values for one or two trimeric OmpF channels in a bilayer. Some of the bilayers formed exhibit conductance values of approximately 0.2 nS due to the presence of a single monomer of OmpF.
vesicles in the area surrounding the nano pipet tip. The distribution is then differentiated to give the probability density function (PDF), shown inset in Figure 2b. The PDF peaks at a cycle time of only 6.9 s, which could be further reduced by automating the application of the negative pressure to the back of the nanopipet, currently performed by hand. Obviously, the ability to reuse the pipets enables rapid screening of bilayers for single protein channels. To further demonstrate that a lipid bilayer was the structure sealing the end of the capillary, the detergent octyl-POE was added to the bath solution. As expected, following addition of the detergent the noise in the bilayer recording increases and subsequently the bilayer breaks, indicated by the resistance of the pipet dropping to open pipet values (see Figure 1 in the Supporting Information). Single Ion-Channel Recordings. In order to characterize the response of single protein channels, various transmembrane 3337
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Figure 4. (a) Ion-current recording of a single trimeric OmpF channel in a nano lipid bilayer at an applied potential of 150 mV. (b) Application of a potential of 200 mV results in gating of the channel in three stages, revealing its trimeric structure. Measurements were performed in 150 mM KCl, 10 mM MES at pH 6.
potentials were applied and the corresponding ion-current traces were recorded. Figure 3a shows typical currentvoltage characteristics for lipid bilayers containing OmpF channels. OmpF is a trimeric protein with three independent pores, as shown in Figure 4b. The X-ray structure of OmpF reveals a homotrimer, where each monomer is folded as a β-barrel formed by 16 antiparallel β-strands. A key feature in the structure of OmpF is the presence of the L3 loop that folds into the channel to form a constriction zone characterized by a strong electrostatic field.20,27 The single trimeric channel conductance of OmpF has been reported to be ∼1 nS in 150 mM KCl;28 therefore, the trace shown in Figure 3a with conductances of 0.8 ( 0.01 nS is consistent with a bilayer containing one trimeric channel. Stable bilayers containing single OmpF channels were also formed successfully using nanopipets with tip diameters of 229 nm (see Figure 2 in the Supporting Information). The OmpF channel, reconstituted in the lipid bilayer, showed asymmetric conductance with respect to the polarity of the applied potential at higher potentials, suggesting an asymmetric conformation of the channel.28 While there is no firm evidence as to whether the OmpF porin inserts with the periplasmic loops inward or outward into the membrane, it is certain that it always inserts in the
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same direction. The OmpF channel also exhibits the same directionality when inserted into liposomes and solvent-free planar lipid bilayers.28 By tuning the concentration of OmpF added to the GUVs and the incubation time, it was possible to regularly form bilayers containing a single OmpF channel. Figure 3b shows the conductance of 30 bilayers formed successively on the tip of a single nanopipet. The bilayers were formed from proteo-GUVs containing a low concentration of the E. coli porin OmpF. Five of the bilayers in this experimental run exhibit increased conductance values consistent with the presence of one, two, or three trimeric OmpF channels in the bilayer. Bilayers can also be repeatedly formed using proteo-GUVs containing high concentrations of OmpF, leading to the presence of up to 50 channels in the nanopipet supported bilayer (See Figure 3 in the Supporting Information). This clearly demonstrates the reproducibility of our approach over a wide range of protein concentrations. It is also very important to emphasize that reconstitution of single channels like OmpF is still a major challenge in the field. Figure 4a shows ion-current recordings, as a function of time, of a single trimeric OmpF channel. The channel is highly stable and maintains its open configuration for the lifetime of the bilayer at low voltages up to 150 mV. Applications of high transmembrane potentials above 200 mV, as shown in Figure 4b, enhance the closure of the channel in three steps revealing trimeric organization of the channel, in agreement with previous studies.5,28,29 Gating of the channel strongly depends on various external parameters like ionic strength of salt solution, applied transmembrane voltage, and pH, and this mechanism is not fully understood. In addition, we observe a fast flickering activity with short conductance drops in the ion-current traces, which is termed “subconductance”. The amplitude of conductance drop is lower than that for a monomer closure of a single trimeric channel (∼330 pS), which indicates this process is distinct from the gating process. The most striking feature of this subconductance is that the frequency of events is strongly dependent on the applied potential: at low potentials events are rare whereas at high potentials the number of events becomes large. The frequency of channel subconductance events and the duration of events deviates between two channels of the same type and also from the same stock of the protein channel. However, the qualitative behavior of subconductance states is comparable in all cases. One hypothesis is that the subconductance states originate from the purification and reconstitution of the porin.30 Reconstitution of channels into lipid membranes is influenced by the presence of lipopolysaccharides (LPS) that contain negative charges. The OmpF channels used in this study were extracted and solubilized with octyl-polyoxyethylene and the amount of LPS varies from protein to protein, which may influence the channel closure. To best of our knowledge, the molecular mechanism underlying the subconductance states is still unclear. Our experiments are not limited to the reconstitution of OmpF into the bilayer. In addition, we have been able to successfully incorporate wild-type R-hemolysin. R-Hemolysin is one of the best known and characterized channels in the single-molecule sensing field. However, in the same buffer used to investigate the OmpF channel (150 mM KCl, 10 mM MES buffer at pH 6.0), its single channel conductance is low, ∼0.2 nS (see Figure 4 in the Supporting Information), as found previously.31 Our measurements also show that wild-type R-hemolysin displays high noise in the bilayer and fast flickering activity (see Figure 4 in the 3338
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Figure 5. Antibiotic interactions with OmpF. (a) Ion-current traces for single trimeric OmpF at an applied potential of 50 mV in the absence (top) and presence (bottom) of the antibiotic enrofloxacin (0.1 mM). Inset shows a single antibiotic binding event. (b) Power spectrum analysis of the ion-current traces in the presence and absence of enrofloxacin, at applied potentials of 25 and 50 mV. Enrofloxacin measurements were performed using 150 mM KCl, 10 mM MES at pH 6.
Supporting Information), making single-molecule detection difficult at this buffer concentration.21,32 However, at an increased
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salt concentration, 300 mM, the noise level is significantly reduced (see Figure 4 in the Supporting Information). The reconstitution of a protein channel like OmpF from a detergent solution is substantially more challenging than reconstitution of water-soluble proteins optimized for membrane insertion like R-hemolysin. The techniques demonstrated here open up the possibility of simple and robust single channel measurements to a much wider range of protein channels. In combination with the successful experiments on OmpF, our results on R-hemolysin demonstrate the wide applicability of our approach. Antibiotic Interactions with OmpF. In order to prove the single molecule detection capabilities of our experiments, we carried out high-resolution ion current recordings in the presence of antibiotics that enabled us to quantify the antibiotic channel interactions. In this work we focused on the interaction of enrofloxacin, a quinolone antibiotic, with OmpF channels. A single OmpF trimer was reconstituted into a lipid bilayer, and the ion current through the channel was recorded. Addition of enrofloxacin to the system caused transient blockages of the ionic current in a voltage-dependent manner. The number of blockage events depends on the applied transmembrane potential and concentration of the antibiotics used. Figure 5a shows that enrofloxacin interacts with OmpF resulting in frequent ion current blockages at 50 mV, as expected. At negative voltages, ion blockages are 10 times more frequent than at positive voltages, irrespective of the concentration of the antibiotic used.33 As shown previously34,35 such interactions of the antibiotic with this channel can be assumed to be a two-state Markovian process that generates an excess ion current noise and whose power spectrum can be fitted by a single Lorentzian, shown in Figure 5b. Noise spectra obtained at 50 mV reveal significant excess noise compared to those obtained at 25 mV, indicating stronger antibioticchannel interactions at higher potentials. The average residence time of enrofloxacin blockage was calculated by dwell time analysis (see Figure 5 in the Supporting Information). The single exponential fit of the blockage time histogram revealed the kinetic constants for antibiotic interaction with the channel surface. Kinetic constants of enrofloxacin blocking were obtained from two experimental parameters, the open time τo (the average time interval between two successive blockages events) and the closed time, τc (the average residence time of channel closure). The open time was calculated to be 25 ( 2 ms and closed time (residence time) was calculated to be 0.8 ( 0.2 ms. Conclusions. We have successfully used glass nanopipets as a support for artificial lipid bilayers. Pipets can be fabricated simply by drawing glass capillaries using a benchtop laser pipet puller in a matter of minutes and do not require further modification before use. On application of a negative pressure of 1 Pa to the back of the pipet, giant unilamellar vesicles were drawn to the nanopipet tip where they break onto form a supported bilayer with typical seal resistances of ∼140 GΩ. The reduction in the area over which the bilayer spans, as compared to traditional methods using Teflon membranes, leads to enhanced mechanical and voltage stability. Over 50 bilayers can be formed on a single nanopipet tip, in less than 10 min, enabling bilayers to be rapidly screened for the presence of single protein channels. Bilayers are successfully formed containing OmpF and R-hemolysin protein channels. We successfully demonstrate blocking of the OmpF channel with the antibiotic enrofloxacin, proving the single molecule detection capabilities of our setup. The simplicity of 3339
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’ ASSOCIATED CONTENT
bS Supporting Information. Additional figures showing breaking of bilayer by detergent, insertion of OmpF, repeated formation of bilayers with high protein concentrations, recordings with Rhemolysin, and analysis of antibiotics blocking events. This material is available free of charge via the Internet at http://pubs.acs.org. ’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected]. Author Contributions §
These authors contributed equally to the work.
’ ACKNOWLEDGMENT We thank A. H€ufner for assistance with measurements. This work was supported by a grant from the Nanoscience E+ initiative (through DFG WI 2279/18-1). O. J. Pambos acknowledges funding by the EPRSC, through the Physics of Medicine Initiative, University of Cambridge. L. J. Steinbock and O. Otto thank the Deutsche Telekom Foundation and the Boehringer Ingelheim Fonds, respectively, for a PhD scholarship. U. F. Keyser and C. Chimerel acknowledge funding by the Emmy Noether-Program of the DFG.
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