Membrane Surface Dynamics of DNA-Threaded Nanopores Revealed

Dec 25, 2003 - Department of Chemistry, Wheaton College, Wheaton, Illinois 60187, ... and Measurements, 203 Kellogg Pl., Wheaton, Illinois 60187,...
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Langmuir 2004, 20, 898-905

Membrane Surface Dynamics of DNA-Threaded Nanopores Revealed by Simultaneous Single-Molecule Optical and Ensemble Electrical Recording Emily L. Chandler,†,‡ Alyssa L. Smith,† Lisa M. Burden,†,‡ John J. Kasianowicz,§ and Daniel L. Burden*,†,‡ Department of Chemistry, Wheaton College, Wheaton, Illinois 60187, Biomolecular Nanotechnologies and Measurements, 203 Kellogg Pl., Wheaton, Illinois 60187, and National Institute of Standards and Technology, Biotechnology Division, Gaithersburg, Maryland 20899-8313 Received September 15, 2003. In Final Form: November 17, 2003 We describe a method for simultaneous single-molecule optical and electrical characterization of membrane-based sensors that contain ion-channel nanopores. The technique is used to study the specific and nonspecific interactions of streptavidin-capped DNA polymers with lipid bilayers composed of diphytanoyl phosphatidylcholine and diphytanoyl phosphatidylglycerol. Biotinylated DNA that is bound to fluorescently labeled streptavidin is electrophoretically driven into, or away from, the lumen of alpha hemolysin (RHL) ion channels by an external electric field. Confocal microscopy simultaneously captures single-molecule fluorescence dynamics from the membrane interface at different applied potentials. Fluorescence correlation analysis is used to determine the surface number density and diffusion constant of membrane-associated complexes. The dual optical and electrical approach can detect membrane-associated species at a surface coverage below 10-5 monolayers of streptavidin, a sensitivity that surpasses most other in vitro surface analysis techniques. By comparing the change in transmembrane current to the number of fluorescent molecules leaving the bilayer when the electrical potential is reversed, we demonstrate the general utility of the approach within the context of nanopore-based sensing and discuss a mechanism by which DNA-streptavidin complexes can be nonspecifically retained at the membrane interface.

Introduction Sensors based on planar lipid bilayer membranes have been widely investigated for their analytical utility.1 A wide variety of applications exist for immunoassays,2 toxicology analysis,3-5 ion sensing,6-8 environmental pollutants,9 and medical diagnostics.8 To work effectively, these devices must be selective, rapid, reproducible, and highly sensitive. Because interfacial chemistry plays a critical role in defining the sensor’s performance, a detailed understanding of the surface molecular dynamics is warranted. Naturally existing or modified ion channels incorporated into lipid bilayers have been used in a variety of sensing applications.10-26 Analyte detection is usually based on changes in the electrical properties of the channel * Corresponding author. Telephone: (630) 752-5065. Fax: (630) 752-5996. E-mail: [email protected]. † Wheaton College. ‡ Biomolecular Nanotechnologies and Measurements. § National Institute of Standards and Technology. (1) Trojanowicz, M. Fresenius’ J. Anal. Chem. 2001, 371, 246-60. (2) Reiken, S. R.; Van Wie, B. J.; Sutisna, H. Biosens. Bioelectron. 1996, 11, 91-102. (3) Andreou, V. G.; Nikolelis, D. P.; Tarus, B. Anal. Chim. Acta 1997, 350, 121-9. (4) Andreou, V. G.; Nikolelis, D. P. Anal. Chem. 1998, 70, 2366-71. (5) Siontorou, C. G.; Andreou, V. G.; Nikolelis, D. P.; Krull, U. J. Electroanalysis 2000, 12, 747-51. (6) Sato, H.; Wakabayashi, M.; Ito, T.; Sugawara, M.; Umezawa, Y. Anal. Sci. 1997, 13, 437-46. (7) Salamon, Z.; Tien, H. T. Electroanalysis 1991, 3, 707-10. (8) Tien, H. T.; Ottova, A. L. In Membrane biophysics: As viewed from experimental bilayer lipid membranes (planar lipid bilayers and spherical liposomes); Elsevier: Amsterdam, 2000. (9) Nikolelis, D. P.; Siontorou, C. G.; Andreou, V. G. Lab. Rob. Autom. 1997, 9, 285-96. (10) Thompson, M.; Krull, U. J.; Venis, M. A. Biochem. Biophys. Res. Commun. 1983, 110, 300-4.

recognition element. The alpha hemolysin ion channel (RHL), which self-assembles from seven identical monomers to form a pore with an inner diameter ranging from 14 to 46 Å,27 creates a nanoscale structure that is of optimal size for detecting the passage of small molecules and polymers. Under the proper solution conditions, this nanopore can also be made unusually stable in the open state, with virtually no random gating events.17 These (11) Minami, H.; Sugawara, M.; Odashima, K.; Umezawa, Y.; Uto, M.; Michaelis, E. K.; Kuwana, T. Anal. Chem. 1991, 63, 2787-95. (12) Adachi, Y.; Sugawara, M.; Taniguchi, K.; Umezawa, Y. Anal. Chim. Acta 1993, 281, 577. (13) Sugawara, M.; Hirano, A.; Rehak, M.; Nakanishi, J. Biosens. Bioelectron. 1997, 12, 425. (14) Nikolelis, D. P.; Siontorou, C. G.; Krull, U. J.; Katrivanos, P. L. Anal. Chem. 1996, 68, 1735-41. (15) Cornell, B. A.; Braach-Maksvytis, V. L.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wieczorek, L.; Pace, R. J. Nature 1997, 387, 580-3. (16) Kasianowicz, J.; Burden, D.; Han, L.; Cheley, S.; Bayley, H. Biophys. J. 1999, 76, 837-45. (17) Kasianowicz, J.; Bezrukov, S. Biophys. J. 1995, 69, 94-105. (18) Gu, L. Q.; Braha, O.; Conlan, S.; Cheley, S.; Bayley, H. Nature 1999, 398, 686-90. (19) Kasianowicz, J.; Henrickson, S.; Weetall, H.; Robertson, B. Anal. Chem. 2001, 73, 2268-72. (20) Movileanu, L.; Howorka, S.; Braha, O.; Bayley, H. Nat. Biotechnol. 2000, 18, 1091-5. (21) Meller, A.; Nivon, L.; Brandin, E.; Golovchenko, J.; Branton, D. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 1079-84. (22) Howorka, S.; Movileanu, L.; Braha, O.; Bayley, H. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 12996-3001. (23) Kasianowicz, J.; Brandin, E.; Branton, D.; Deamer, D. W. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 13770-3. (24) Akeson, M.; Branton, D.; Kasianowicz, J. J.; Brandin, E.; Deamer, D. W. Biophys. J. 1999, 77, 3227-33. (25) Henrickson, S.; Misakian, M.; Robertson, B.; Kasianowicz, J. J. Phys. Rev. Lett. 2000, 85, 3057-60. (26) Vercoutere, W.; Winters-Hilt, S.; Olsen, H.; Deamer, D.; Haussler, D.; Akeson, M. Nat. Biotechnol. 2001, 19, 248-52. (27) Song, L.; Hobaugh, M. R.; Shustak, C.; Cheley, S.; Bayley, H.; Gouaux, J. E. Science 1996, 74, 1859-66.

10.1021/la035728i CCC: $27.50 © 2004 American Chemical Society Published on Web 12/25/2003

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features have made RHL especially useful for detecting compounds at the single-molecule or single-ion level.16-23,26 Such exquisite detection power has been demonstrated for H+/D+ discrimination,17 heavy-metal cation sensing,16 polynucleotides,23 proteins,20 and small molecules in solution.15,18 In one particular strategy,19 analytes react with a binding site that is covalently linked to the end of a singlestranded polynucleotide. In the presence of an electric field, the negative charge residing on the polymer causes the complex to thread through the RHL nanopore. However, if the analyte bound to the polymer end is larger than the pore lumen, the polynucleotide-analyte complex cannot completely pass through the channel. Instead, translocation is halted and a portion of the polymer remains trapped within the pore, occluding the flow of ionic current for an extended period of time. By designing polynucleotides that produce a characteristic electrical signature during pore penetration, it should be possible to expand the detection methodology for simultaneous identification of a number of soluble species.19,21 However, in both ideal and complex media, nonspecific adsorption to the lipid membrane could ultimately limit the sensor’s utility, if adsorption is left uncharacterized and uncontrolled. In this study, we combine single-molecule confocal fluorescence microscopy with ion-channel current recording to monitor RHL nanopores and lipid membrane adsorbates. The surface dynamics in bilayers composed of diphytanoyl phosphatidylcholine and diphytanoyl phosphatidylglycerol are compared. Patterned after the detection strategy discussed above, we utilize single-stranded biotinylated DNA and streptavidin to reduce ion flow across a channel-containing membrane; however, in this work, the streptavidin is fluorescently labeled. By controlling the polarity of the transmembrane potential, polynucleotides are electrophoretically threaded into or ejected from RHL pores on command, effectively producing ion channels with “switchable” fluorescent states. When threaded through the channel interior, the fluorescent DNA-streptavidin complex enables ion-channel optical recording. When the complex is ejected into bulk solution, the channels lose their fluorescent reporter and become invisible to the microscope detector. Only those complexes that are nonspecifically bound to the membrane surface continue to produce detectable signatures. At both positive and negative polarities, we use fluorescence correlation analysis to determine the lateral diffusion constants and the surface coverage of membrane-associated complexes. Simultaneous fluorescence intensity and electrical measurements have been used in the past to determine structural properties of ion channels in planar bilayers.28 More recent advances in optical microscopy have greatly enhanced the sensitivity and information-gathering power of fluorescence-based measurements. For example, array detectors have been employed for evaluating singlemolecule diffusion by probing large membrane areas with 10-50 ms time resolution.29,30 Similar optical approaches have been used for simultaneous optical and electrical measurement of single channels in planar membranes31-34 and living cells.35,36 Confocal microscopy enables single-molecule detection at better than 10-µs time resolution and also provides a (28) Veatch, W. R.; Mathies, R.; Eisenberg, M.; Stryer, L. J. Mol. Biol. 1975, 99, 75-92. (29) Schmidt, Th.; Schu¨tz, G. J.; Baumgartner, W.; Gruber, H. J.; Schindler, H. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 2926-9. (30) Schmidt, Th.; Schu¨tz, G. J.; Baumgartner, W.; Gruber, H. J.; Schindler, H. J. Phys. Chem. 1995, 99, 17662-8.

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level of spatial discrimination in the axial direction that is not available with other approaches. Unlike other optically based surface analytical tools that require a solid-liquid interface (e.g., surface plasmon resonance, ellipsometry, and various evanescent wave techniques), the far-field nature of confocal microscopy permits solutes to traverse the membrane in an uninhibited fashion. In addition, most traditional surface analysis techniques cannot generate single-molecule detection limits. Thus, the combination of single-molecule confocal microscopy with ion-channel current recording in planar bilayers creates a unique tool for membrane characterization that enables new surface dynamics to be described. As we demonstrate in this study, the dual-recording methodology confirms nonspecific interactions between the DNAstreptavidin complexes and the membrane surface that can only be indirectly detected by either technique alone. Experimental Section Membrane Formation. Unsupported lipid bilayers were formed on a small Teflon aperture with a modified version of the Mueller-Rudin technique.37 We employed a horizontal bilayer chamber similar to that described elsewhere,24,38 but with geometrical alterations to enable viewing under a high numerical aperture (NA) microscope objective. We formed a long and narrow constriction between the upper and lower solution-filled chambers (1 M KCl, 5 mM HEPES, pH 7.5) by forcing a thin metal needle through a Teflon plug until the sharpened tip broke through the surface on the opposite side. Extraction of the needle created a tapered hydrophobic channel that varied from 20 to 100 µm in diameter and was typically 8-12 mm in length. The long narrow channel helped minimize force imbalance due to pressure differentials across the membrane and reduced the likelihood of problematic membrane movement. Lipids composed of diphytanoyl acyl chains and either a phosphatidylcholine (PC) or a phosphatidylglycerol (PG) headgroup were purchased from Avanti Polar Lipids (Alabaster, AL). All lipid materials were dissolved in either hexadecane (Aldrich, Milwaukee, WI) or pentane (VWR, Bridgeport, NJ). Fluorescently labeled lipid (TRITC-DHPE, Molecular Probes, Eugene, OR) was mixed with diphytanoyl PC at a molar ratio of 1:106 for lipid diffusion measurements. Bilayers were prepared by allowing lipid in pentane solution to dry on the aperture surface, filling the upper and lower chambers with aqueous buffer, and then stroking lipid in hexadecane across the aperture until a membrane spontaneously formed. Solution exchange was performed via gravity feed using flow rate controllers with optical feedback (Spectrum Chromatography, Houston, TX). This permitted delivery and withdrawal of buffer to and from the top chamber at programmable rates. The solution exchange removed excess protein or DNA from the membrane surface without destabilizing the membrane. RHL Synthesis. RHL was amplified by PCR from a DNA template (pBR322 containing a 3 kb EcoRI/HindIII insert including RHL) and subcloned into pET-2T-parallel39 using the NcoI and SpeI sites. The entire sequence of RHL was confirmed (31) Ide, T.; Yanagida, T. Biochem. Biophys. Res. Commun. 1999, 265, 595-9. (32) Harms, G.; Orr, G.; Montal, M.; Thrall, B.; Colson, S.; Lu, H. Biophys. J. 2003, 85, 1826-38. (33) Ide, T.; Takeuchi, Y.; Yanagida, T. Single Molecules 2002, 3, 33-42. (34) Borisenko, V.; Lougheed, T.; Hesse, J.; Fu¨reder-Kitzmu¨ller, E.; Fertig, N.; Behrends, J. C.; Woolley, G. A.; Schu¨tz, G. J. Biophys. J. 2003, 84, 612-22. (35) Sonnleitner, A.; Mannuzzu, L.; Terakawa, S.; Isacoff, E. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 12759-64. (36) Harms, G.; Cognet, L.; Lommerse, P.; Blab, G.; Kahr, H.; Gamsja¨ger, R.; Spaink, H.; Soldatov, N.; Romanin, C.; Schmidt, T. Biophys. J. 2001, 81, 2639-46. (37) Mueller, P.; Rudin, D. O.; Tien, H. T.; Wescott, W. C. Nature 1962, 194, 979-80. (38) Burden, D.; Kasianowicz, J. J. Phys. Chem. B 2000, 104, 61037. (39) Sheffield, P.; Garrard, S.; Derewenda, Z. Protein Expression Purif. 1999, 15, 34-9.

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using DNA sequencing. Expression of the (His6)-fusion protein was in BL21(DE3) cells (Novagen, Madison, WI). Briefly, cells at an OD600 of 0.6-0.7 were induced with 1 mM IPTG after which they were grown for 3 h at 37 °C. Pelleted cells were lysed in BPER buffer (Pierce, Rockford, IL) followed by centrifugation at 14 000 rpm for 30 min. The soluble His-tagged RHL was purified using HiTrap resin (Amersham Biosciences, Piscataway, NJ). After the resin was washed with buffer (20 mM Tris pH 8.0, 100 mM KCl), fusion protein was eluted with buffer containing imidazole. Removal of the His6 tag was accomplished by rTEV protease (Invitrogen, Carlsbad, CA) during dialysis in 20 mM Tris pH 8.0, 100 mM KCl, 1 mM DTT, 0.5 mM EDTA. SDSPAGE verified the molecular weight of RHL and revealed that the RHL accounted for ∼70% of the protein present in the sample. The yield of RHL was ∼1 mg from 50 mL of cell culture. Another source of RHL, which was produced by in vitro transcription/translation synthesis,40 was also used. Both sources of RHL gave identical average single-channel current rectification ratios (1.3-1.4), were more than 70% pure (as judged by SDSPAGE), and behaved identically in simultaneous single-molecule optical and electrical experiments. Electrical Recordings. Electrical potentials were applied across the aperture using a DC-10 MHz waveform generator (Hewlett-Packard 33120A, Palo Alto, CA) and Ag/AgCl electrodes (In Vivo Metric, Ukiah, CA) placed above and below the orifice. The ionic current was amplified by a factor of 106-109 using a custom-built current-to-voltage converter and low-pass filtered using an 8-pole Bessel active filter (Frequency Devices 9002, Haverhill, MA). Data were digitized at 10 Hz using a 12-bit analog-to-digital converter (PCI-MIO-16E-1, National Instruments, Austin, TX) and were analyzed with software written in LabVIEW 6.1. The bilayer chamber and the microscope were isolated from sound and light, as well as external electrical fields, by an enclosed Faraday cage. Capacitance Recordings. Membrane capacitance recordings served both to indicate bilayer formation and to provide a measure of bilayer area. A triangle wave (400 Hz, 100 mV peak-peak) generated a corresponding output square wave upon membrane formation, the amplitude of which was converted into capacitance at 0.5 Hz. Typical capacitance values for properly formed bilayers ranged from 15 to 30 pF, depending upon membrane curvature and aperture size. Given the area of the aperture as determined via image analysis, these values result in a specific capacitance of ∼0.7 µF/cm2 for both PC and 5% PG membranes. Others have reported specific capacitance figures that range from 0.4 to 1 µF/cm2 for similar membranes.41,42 Current Recordings. Electrical potential is defined with respect to the top solution compartment. Thus, a negative applied voltage drives anions from the top to the bottom chamber and is identified as positive current flow. We studied the properties of membranes containing between 1 and the conductance equivalent of ∼35 000 channels. A linear extrapolation from the average single-channel current measurement (129 ( 7 pA at -120 mV, 112 measurements) was used to determine the total number of channels present in the membrane. If independent electrical behavior and a homogeneous spatial distribution of the membrane-incorporated channels are assumed, calculated channel densities for simultaneous optical and electrical measurements ranged between 0.5 and 20 channels/µm2. Channel Insertion and Occlusion. A 0.5-µL aliquot of RHL (0.4 mg/mL for ensemble electrical measurements and 0.01 mg/ mL for single-channel electrical measurements) mixed with >2 M KCl (1:1 v/v ratio) was delivered via pipet to the top solution chamber in close proximity to the membrane-supporting aperture. Upon diffusion to the bilayer, RHL monomers self-assembled to form transmembrane channels. Fluorescently labeled streptavidin (tetramethylrhodamine conjugate, TMR) was obtained from Molecular Probes, and biotinylated polynucleotide [5′-biotinylated poly(deoxyadenylic acid), bT-poly(dA)100] was purchased from Midland Certified Reagents (Midland, TX). The streptavidin and polynucleotide were mixed in buffer (10 mM Trizma base, 10 mM EDTA, pH 8.0) to create a DNA-protein complex. Experiments were conducted using either a 400-nM streptavidin-TMR/ (40) Walker, B.; Krishnasastry, M.; Zorn, L.; Kasianowicz, J.; Bayley, H. J. Biol. Chem. 1992, 267, 10902-9.

Chandler et al. 400-nM bT-poly(dA)100 or a 400-nM streptavidin-TMR/4-µM bTpoly(dA)100 solution. All solutions were filtered three times with a 10-kD Microcon centrifugal filter to remove residual free dye. Using fluorescence correlation techniques and the single-molecule optical detection capability of the microscope, the free-dye content in the third filtrate was measured and found to be equivalent to that of a filtered blank. After channels appeared in the membrane, 0.5 µL of the DNA-protein complex mixed with >2 M KCl (1:1 v/v ratio) was added over the membrane surface. Optical Recordings. The instrument consists of a confocal microscope that was enhanced from previous work38 to enable simultaneous electrical and optical recordings. The microscope was configured in an upright geometry and used water-immersion optics to interrogate membranes formed on a horizontal bilayer sample chamber. Briefly, the 514-nm line from an argon ion laser (Melles Griot, Carlsbad, CA) was sent through a 5× beam expander and the internal optics in the microscope body (Carl Zeiss, Inc., Thornwood, NY). The laser beam was directed to the microscope objective by a dichroic mirror (532 DRLP, Omega Optical, Inc., Brattleboro, VT). Our instrument used a 150×, NA ) 1.25 water-immersion objective to focus the laser beam to a diffraction-limited spot (10-40 µW) and collect the fluorescence emission. This resulted in a power density of 5-20 kW/cm2, which virtually eliminates biasing of the fluorescence burst signature due to photobleaching.38 Fluorescence photons were transmitted through the first dichroic and one barrier filter (530 EFLP, Omega Optical). A second dichroic (565 DCSP, Omega Optical) was employed to direct light between 580 and 720 nm to a CCD camera (XC-75, Sony) for image analysis of the membrane aperture. The remaining fluorescent band (532-580 nm) was transmitted to a 100-µm pinhole. A small lens collected light emerging from the pinhole and focused the photons onto the sensitive area of an avalanche photodiode single-photon counting module (SPCMAQR-15, PerkinElmer Optoelectronics, Fremont, CA). Pulse trains from the photon counter were monitored with a multichannel scalar card (MCS-plus, EG&G Ortec, Oak Ridge, TN) operated by a dedicated personal computer. The 3D confocal detection volume can be approximated by a cylinder.43 The characteristic dimensions in our apparatus (∼2 µm axial, 230-260 nm radial) were determined by 3D fluorescence correlation analysis and the known diffusion constant for R6G in water43 under 20-40 µW illumination. All correlation analyses were performed by recording real-time photon-burst signatures to disk and autocorrelating between 10 and 60 s of data with software written in LabVIEW 6.1. The resulting correlation curves were fit using the Levenburg-Marquardt nonlinear least squares algorithm and standard correlation models used in fluorescence correlation spectroscopy (FCS). Computer Automation and Image Analysis. Input and output signals from the stage drivers, function generator, current amplifier, CCD camera, and APD detector were coordinated and controlled by a computer. Two 3D translation stages used in tandem, both capable of operation in either a manual mode or via automated feedback from image analysis software, positioned the membrane aperture in the laser focus. A 3-axis stage actuated by stepper motors (Nanomotion II, Melles Griot, Rochester, NY) provided a range of motion of up to 25 mm in the X and Y directions with a precision of ∼50 nm. The Z axis spanned 20 mm, with a precision of ∼500 nm. The large range of motion enabled membrane formation using a low-magnification, long-workingdistance objective and quick sample repositioning for interrogation with the 150×, short-working-distance objective. A second 3-axis piezoelectric-actuated stage (Nanoblock, Melles Griot) under capacitive feedback control performed small position adjustments, with a range of 20 µm in the X and Z directions and 30 µm in the Y direction. All axes were capable of 5-nm step resolution. National Instruments IMAQ Vision software was developed to visualize the aperture, calculate its area and centroid location, and provide spatial feedback for automated positioning of the confocal detection volume on the bilayer. (41) Niles, W. D.; Levis, R. A.; Cohen, F. S. Biophys. J. 1988, 53, 327-35. (42) White, S. H.; Thompson, T. E. Biochim. Biophys. Acta 1973, 323, 7-22. (43) Rigler, R.; Mets, U.; Widengren, J.; Kask, P. Eur. Biophys. J. 1993, 22, 169-75.

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Figure 1. (A) CCD image of an aperture supporting a lipid bilayer. The rim location is enhanced by tracing that is used for locating the membrane centroid. The box represents the region over which 3D confocal imaging is performed, and the inset shows a cross-sectional view of the bilayer chamber. (B) Confocal image of a bilayer containing DNA-threaded channels. Membrane curvature results from the stroking technique used in bilayer formation. A Teflon shard is visible in the rear. (C) Real-time fluorescence recording of single-molecule bilayer dynamics. The photon-burst signature immediately disappears upon rupturing the bilayer and exactly correlates with a large change in current.

Results and Discussion Figure 1A shows an aperture at the interface between the top solution chamber and the Teflon plug. The aperture is coated with a lipid/hexadecane mixture and suspends a membrane bilayer. Transillumination by a low-wattage incandescent lamp generates enough contrast between the coated Teflon and buffer solution to locate the rim. Scanning 3D confocal imaging enables investigation of the spatial distribution of fluorescent material across the bilayer. Figure 1B displays a 3D image of a membrane

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containing ∼12 000 RHL channels that are threaded with fluorescent DNA-streptavidin complexes. As can be seen, fluorescent material is uniformly distributed over the bilayer surface. The cutaway image of the membrane indicates no discernible thickness increase in the region of the membrane torus, an expected result given that only the top surface of the bilayer contains fluorescent material. Apertures were occasionally found to contain thin shards, presumably composed of Teflon, extending inwardly from the rim over a distance of several microns. A typical shard can be seen in Figure 1B, supporting the membrane on either side. These protrusions, which were not readily visible in the CCD images, do not appear to interfere with the formation or stability of the lipid membrane. However, during optical interrogation, inadvertent placement of the detection volume on a lipid-coated protrusion produces a fluorescence time record that does not reflect the true dynamic behavior of a bilayer surrounded by solution on each side. Placement of the detection volume in the center of the aperture greatly reduces the likelihood of data collection from a lipid-coated shard. As an additional precaution, we terminated all membrane measurements by recording an induced breakage event (Figure 1C). Application of a 1 V dc pulse exceeds the membrane breakdown potential, causing the immediate rupture of the bilayer. Figure 1C shows a real-time single-molecule optical recording taken from a membrane that contained threaded ion channels. The discrete nature of the photon bursts (i.e., individual flashes separated by short dark periods) indicates that single fluorescent molecules are being detected as they traverse the confocal detection circle on the membrane surface. Upon membrane breakage, cessation of the photon-burst pattern and an instantaneous change in current are simultaneously observed. Recording a membrane rupture at the end of each measurement confirmed that optical data were generated by a bilayer that was bordered by solution on either side, rather than bulk material on a liquid/solid interface. Although the geometry of the confocal detection volume can be approximated by a cylinder, the position of the actual hourglass spatial profile leads to measurement errors in both the diffusion coefficients and label number densities if the bilayer is not coplanar with the volume waist. Errors introduced by axial displacements can be minimized by using a pinhole diameter that is smaller than the diffraction-limited laser spot when demagnified into the object plane; however, this practice attenuates the photon collection efficiency. To estimate the magnitude of the error associated with axial displacement in our apparatus, optical recordings were conducted at several locations above and below a PC bilayer that contained a trace quantity of TRITC-DHPE (molar ratio of 106:1). Fluorescence correlation analysis of the real-time optical recording was performed using a 2D correlation model with correction for background,44-48

G2D(τ) )

[〈F(t)〉 + 〈B(t)〉]2 1 g (τ) + 1 〈N〉 2d [〈F(t)〉]2

(

g2d(τ) ) 1 +

)

4Dτ ω12

(1)

-1

(2)

where 〈F(t)〉 and 〈B(t)〉 are the time-averaged fluorescence signal and background, 〈N〉 is the time-averaged number density of fluorescent labels occupying the spot, ω1 is the radius of the beam waist, and D is the apparent diffusion constant. Background data were acquired immediately

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Figure 2. Axial profile of the apparent lipid diffusion coefficient, D, and G(0) values produced by correlation analysis of a TRITC-DHPE containing membrane. The beam-waist intersection with the membrane plane is located at ∼16.7 µm (relative to the stage zero position) and yields a lipid diffusion measurement of (1.4 ( 0.2) × 10-7 cm2/s over an ∼0.8-µm distance. Similar contours were obtained for membranes containing DNA-threaded nanopores but yield smaller values for D.

following the intentional rupture of the bilayer (Figure 1C). Holding ω1 fixed at 245 nm produces the G(0) and D values displayed in Figure 2. The profile reveals a (0.4-µm distance on either side of the membrane where diffusion coefficient measurements are relatively insensitive to axial displacements. The positioning reproducibility and membrane stability within our system are estimated to be less than (25 nm, a value that lies well within the total 0.8-µm tolerance range. Thus, accurate and reproducible diffusion coefficient measurements can be accomplished without exactly replicating the position of the membrane at the beam waist. Displacement of the membrane just outside of the (0.4-µm axial range results in an ∼40% decrease in the calculated value for D because of the hourglass geometry of the detection volume. Erroneously large diffusion coefficients result from the correlation of fluorescent background material that is freely moving in solution above and below the bilayer, where no portion of the volume intersects the membrane plane. For the 0.8-µm range depicted in Figure 2 that produces accurate diffusion measurements, correlation analysis gives a lipid diffusion coefficient of (1.4 ( 0.2) × 10-7 cm2/s. This value is similar to measurements reported elsewhere.49 G(0) values are much more sensitive to axial displacement, varying by ∼20% over the same 0.8-µm region. Because G(0) values are related to the amplitude of the fluorescent bursts, monitoring the burst amplitude in real time is an effective means to detect small membrane displacements. In the event of slow membrane drift, a change in burst amplitude is quickly detected and the membrane position can be adjusted to reestablish coplanarity with the beam waist. For all experiments reported here, time recordings were carefully monitored (44) Fahey, P.; Koppel, D.; Barak, L.; Wolf, D.; Elson, E.; Webb, W. Science 1977, 195, 305-6. (45) Thompson, N. L. Topics in Fluorescence Spectroscopy; Plenum Press: New York, 1991; Chapter 6, pp 337-378. (46) Magde, D.; Elson, E. L.; Webb, W. W. Phys. Rev. Lett. 1972, 29, 705-8. (47) Elson, E. L.; Magde, D. Biopolymers 1974, 13, 1-27. (48) Magde, D.; Elson, E. L.; Webb, W. W. Biopolymers 1974, 13, 29-61. (49) Sonnleitner, A.; Schu¨tz, G.; Schmidt, Th. Biophys. J. 1999, 77, 2638-42.

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to ensure that no detectable membrane displacements occurred during data acquisition. Figure 3 displays simultaneous optical and electrical recordings of threaded ion channels incorporated into a PC membrane. The current trace (Figure 3A) reveals a rapid rise after the addition of RHL, indicating thousands of channel insertions. Upon the addition of DNAstreptavidin complexes, a slight rise in current is immediately evident due to the slightly elevated salt level in the added aliquot. This is soon followed by a rapid current decrease that indicates channel occlusion by the polymer complex. The aqueous solution in the top chamber is subsequently rinsed free of DNA-streptavidin by continuously exchanging the buffer solution. Figure 3C illustrates the various axial positions of real-time optical recording, measured with respect to the position that produces a maximum photon-burst amplitude. Figure 3D shows real-time photon-burst recordings at various axial locations before the potential is reversed. The polarity change is evident in both the current (Figure 3A) and the potential recordings (Figure 3B), indicating the ejection of DNA from the membraneincorporated nanopores. However, evidence of the polarity reversal cannot be directly observed in the optical record. Burst recordings from the membrane surface before, during, and after the potential switch (Figure 3D,E) are indistinguishable, even though the electrical record clearly shows that DNA-streptavidin complexes no longer block ion flow through the channels. Fluorescence correlation analysis of the optical records also reveals no significant change in surface coverage after the potential switch. Figure 4 shows a correlation curve from the photon-burst record prior to the potential switch. Correlation analysis conducted after the potential switch produces a curve that is identical (within experimental uncertainty). These results indicate that a nonspecific membrane interaction causes the DNA-streptavidin complexes to remain localized at the membrane-solution interface. Fitting the data before and after the potential switch with a 2D, single-component model (eqs 1 and 2) produces diffusion coefficients of (5.9 ( 0.2) × 10-8 cm2/s. This value is consistent with large membrane-associated complexes when compared to the diffusion constant measured for freely diffusing lipids ((1.4 ( 0.2) × 10-7 cm2/s). In an attempt to reduce nonspecific binding, we also conducted measurements using membranes of binary lipid composition (95% PC/5% PG, molar ratio). The negative charge residing on the PG headgroup repels the negatively charged polynucleotides from the membrane interface. We also increased the molar concentration of DNA, elevating the DNA-streptavidin ratio to 4:1. Because streptavidin has four biotin binding sites, a larger fraction of the occlusion complexes carry an increased negative charge. Thus, when the polarity is reversed, association with the lipid membrane is less energetically favorable, and occlusion complexes can move into bulk solution more readily. Figure 5A shows electrical data from a typical experiment. Here, ∼23 000 channels are incorporated into the bilayer, followed by occlusion with DNA-streptavidin. After rinsing to remove excess fluorescent material (9501070 s), the detection volume is centered on the membrane and the photon-burst record is acquired. A photonburst record is also acquired after the polarity reversal at 1838 s. Fluorescence correlation analysis of the photon record before and after the potential reversal are shown in Figure 5B. Here, the increased amplitude of the curve after the polarity switch clearly indicates that many of the DNA-

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Figure 3. Simultaneous optical and electrical recording of channel dynamics in a PC membrane. (A) Current recording shows channel insertions, amplifier gain changes, and channel occlusions after the addition of DNA-streptavidin (the current axis is calibrated for the final gain setting). The chamber is then rinsed to remove free DNA-streptavidin, and the confocal detection volume is positioned on the membrane. (B) Time course of the applied potential. (C) Confocal micrograph of a typical lipid bilayer. Red indicates the Teflon aperture; blue indicates lipid. (D) Real-time photon-burst recordings at various axial positions before the potential reversal. Each burst corresponds to one or two fluorescent complexes traversing the laser spot. (E) No change in the optical recording before, during, and after the potential switch reveals nonspecific binding that keeps DNA-streptavidin localized at the membrane interface, even though the large negative current (A) indicates that nearly all channels are open.

Figure 4. Autocorrelation of the photon-burst recording taken before the potential switch. The data are fit using eqs 1 and 2 and yield a diffusion constant of (5.9 ( 0.2) × 10-8 cm2/s. Correlation analysis following the potential switch produces a curve that is statistically identical to the one shown here.

streptavidin complexes have been ejected from the interface. As in the 100% PC measurement, the large increase in negative current upon polarity reversal (at 1838 s) indicates that many ion channels are no longer occluded. Quantitative comparison of the value for 〈N〉 from a fit of the data in Figure 5B with eqs 1 and 2 provides an estimate of the relative surface coverage of fluorescent material. The values of 〈N〉 before and after the polarity reversal are 1.28 ( 0.07 and 0.148 ( 0.003, respectively. If a homogeneous distribution of material across the bilayer is assumed, the optical detection area of 0.189 µm2 (245-nm radius) can be utilized to estimate the total

number of fluorescent molecules residing over the entire bilayer interface. Using a value for the total membrane area (∼2500 µm2) derived from the bilayer capacitance measurement (17.8 pF) and the specific membrane capacitance (∼0.7 µF/cm2), a number density of 〈N〉 ) 1.28 indicates that ∼17 000 fluorescent molecules are spread over the membrane surface before the polarity switch. After reversal, 〈N〉 ) 0.148 indicates that only ∼2000 fluorescent molecules remain attached to the membrane and that ∼15 000 DNA-streptavidin complexes have been ejected into bulk solution. The ∼2000 adhering molecules represent 3.5 × 10-5 monolayers of streptavidin (assuming a monolayer packing density of ∼230 ng/cm2).50 This level is well below the quantity detectable by other surface analysis techniques, such as surface plasmon resonance.50,51 Previous ellipsometric52 and surface-force measurements53 have indicated that negligible levels of streptavidin adhere to neutral phospholipids. Although the level of nonspecific binding quantified here is very small, it is highly significant for membrane-based sensors, particularly when signals are generated by individual or small numbers of ion channels within the bilayer. In addition, the trace level of streptavidin adsorption measured in our experiments is not representative of the lowest detectable quantity. The signal-to-noise ratio of the correlation curves generated by trace streptavidin adsorption indicates that confocal microscopy is capable of (50) Jung, L. S.; Nelson, K. E.; Stayton, P. S.; Campbell, C. T. Langmuir 2000, 16, 9421-32. (51) Jung, L. S.; Campbel, C. T.; Chinowsky, T. M.; Mar, M. N.; Yee, S. S. Langmuir 1998, 14, 5636-48. (52) Reiter, R.; Motschmann, H.; Knoll, W. Langmuir 1993, 9, 24305. (53) Sheth, S.; Leckband, D. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 8399-404.

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Figure 5. Simultaneous optical and electrical recording of dynamics in a PC membrane doped with 5% PG. (A) Ensemble current recording shows channel insertion, occlusion after the addition of DNA-streptavidin complexes, and reopening after potential reversal. (B) Autocorrelation of single-molecule photon-burst signatures. The data are fit using eqs 1 and 2. Before the polarity reversal 〈N〉 ) 1.28 ( 0.07. After the polarity switch 〈N〉 ) 0.148 ( 0.003. Diffusion constants, which are reported as an average from three different membrane measurements, give values before and after potential reversal of D ) (3.3 ( 2.7) × 10-8 cm2/s and D ) (7.7 ( 6.4) × 10-8 cm2/s.

quantifying surface coverage by a factor that is ∼10-fold less than 3.5 × 10-5 monolayers. Electrically, the current recording in Figure 5A indicates an initial insertion of ∼23 000 channels. Based on the stepwise decrease in current measured for more than 200 single-channel occlusions, blockage by DNA-streptavidin results in an ∼85% drop in conductance for each pore. Thus, using 129 pA for the average fully open channel, a leakage current of ∼21 pA remains for an occluded pore. Given an ∼108 pA drop per channel, the ensemble current level measured shortly after addition of DNA-streptavidin reveals that ∼20 000 channels are occluded and ∼3000 channels remain in an open state. In this experiment, nearly all of the available DNA-streptavidin complexes have threaded ion channels, leaving behind ∼3000 open nanopores. The gradual rise in current during the rinsing period can be attributed to a combination of two factors. First, rinsing redistributes protein and lipid from the Teflon surface into the aperture, enabling more channels to be incorporated into the bilayer. The increase in negative

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current between potential reversals at 800 and 1910 s is consistent with the insertion of additional channels within the membrane. The difference between these current levels corresponds to an extra ∼1200 fully open channels. Second, rinsing disturbances might also remove threaded DNAstreptavidin complexes from RHL. Although the turbulence is likely to be small, removal can occur more readily than in the 100% PC membrane due to the electrostatic repulsion between the DNA and the PG. After accounting for the ∼1200 additional channel insertions, the remaining difference between current levels at 1000 and 1800 s (∼300 nA) can be attributed to rinsing (∼2800 channel openings). Thus, at the time just prior to the polarity reversal, the original value of ∼20 000 occluded channels is reduced to ∼17 000. This figure matches the value produced by fluorescence correlation analysis prior to the polarity change. After the polarity switch, the current difference measured between 1910 and 1830 s indicates that ∼17 000 channels open to ionic flow in the reverse direction. This figure is also in close agreement with the number of fluorescent molecules leaving the surface (∼15 000), as determined from fluorescence correlation analysis. The fluorescent complexes that remain at the surface (∼2000) apparently adhere nonspecifically, albeit to a much lesser extent than when the membrane is composed of 100% PC. Optical experiments using 100% PC bilayers without incorporated RHL indicate that fluorescently labeled streptavidin (with no complexed DNA) remains on the membrane surface following extensive rinsing at both polarities. Diffusion constant measurements were performed and found to be within experimental error of those obtained from nanopores threaded with DNA-streptavidin. It is possible that the covalently linked fluorescent label enhances the association with the membrane. Control single-channel electrical measurements using bTpoly(dA)100 reveal a drastic decrease in translocation event frequency following a chamber rinse. Similar DNA results have also been observed in vertically oriented bilayers (Kasianowicz and Henrickson, unpublished). Taken together, these data suggest that the labeled streptavidin plays a more significant role than DNA in the retention of complexes at the membrane surface and suggest that the primary mode of nonspecific association is between the lipid membrane and the streptavidin. These data lead to a simple diagram of the possible surface interactions before and after potential reversal (Figure 6A,B). Three types of surface interaction with the DNA-streptavidin complexes are depicted. At negative potential (Figure 6A), type 1 structures involve nonspecifically bound DNA-streptavidin molecules that have not threaded nanopores but undergo lateral movement with diffusion constants typical of peripherally associated complexes. Because the data from Figure 5A indicate that nearly all the DNA-streptavidin complexes have threaded nanopores, very few of these structures are likely to be present before the potential switch. Type 2 structures are occluded nanopores that are formed by DNA-streptavidin access from free solution; no membrane mediation is involved. Conversely, type 3 occlusion complexes arise via membrane mediation. These structures likely involve a diffusive collision between type 1 molecules and open channels, enabling streptavidin to remain nonspecifically associated with the lipid. At constant negative potential, dual optical and electrical measurements cannot discern between these three types. Figure 6B illustrates the interface after the application of a positive potential. Any type 1 complexes present before the potential switch remain associated with the membrane. Type 1 complexes are also formed by the ejection

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Figure 6. Model of surface interactions under positive and negative potentials. Streptavidin caps contain a fluorescent tag. (A) Under negative potential, DNA-streptavidin is driven toward the bilayer interface and might associate with the surface in three characteristic ways (see discussion). (B) Application of a positive potential expels the DNA-streptavidin from the nanopores, enabling ionic current to flow uninhibited. Nonspecifically bound complexes remain localized at the membrane interface, particularly in 100% PC membranes. Membranes containing PG exhibit a much greater predominance of full dissociation (type 2).

of DNA from the lipid-mediated occlusion structures depicted in Figure 6A. This type of nonspecific association dominates in 100% PC membranes, where no electrostatic repulsion from the lipid headgroups is present. DNAstreptavidin can also be expelled into free solution (type 2), causing a decrease in the photon-burst frequency and an increase in ionic current in the negative direction. For membranes containing negatively charged lipids, electrostatic repulsion gives rise to a much more notable change in the bilayer optical signature upon potential reversal, indicating that nonspecific binding has been significantly reduced over the neutral lipid counterpart. Last, DNA-streptavidin might also be ejected from pores but remain bound to the interface through a channelmediated mechanism (type 3 complex). However, at constant positive potential (Figure 6B), dual optical and electrical measurements cannot discern between types 1 and 3. Conclusions Our data indicate that nonspecific adsorption of a DNAprotein complex occurs at the interface of PC membranes.

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These interactions might interfere with the use of membrane-incorporated nanopores for sensing applications, particularly in screening assays that require complete washing of the sample chamber before delivery of new analyte. However, experiments designed to alter the electrostatic charge residing at the membrane surface suggest that it is possible to mitigate adsorption by tailoring the membrane composition to meet the demands of specific applications. Alternatively, non-protein-based polynucleotide caps might also be considered as a means of reducing nonspecific membrane binding. The addition of a rapid optical detection modality should enhance the potential utility of nanopore-based sensors by enabling the simultaneous characterization of both the membrane and the pores. This general approach will also enable the creation of new sensing schemes that are designed to capitalize on the two orthogonal signalproduction mechanisms. Similar techniques will be useful in deciphering the specific mechanisms that cause current blockades in nanopores19-26 and will be applicable to high-throughput screening assays that are designed for selecting drug targets against ion channels or other membrane receptors.54,55 Although the purposes of this study do not demand better than 1-ms time resolution, the high signal-to-noise ratios (>75) generated in the realtime optical record indicate that single-molecule dynamics in lipid bilayers can be acquired with ∼10-µs time resolution. Such rapid data acquisition rates should prove useful for the study of rapid transient ion-channel gating mechanisms and/or the transport of molecules through nanoscale pores. Acknowledgment. We thank Dr. John J. Iandolo (Kansas State University) for the generous gift of the plasmid containing the RHL gene and Dr. Hagan Bayley (Texas A&M University) for supplying RHL produced by in vitro transcription/translation. Funding for this work was provided by Research Corporation, the donors of the Petroleum Research Fund, administered by the American Chemical Society, the Camille and Henry Dreyfus Faculty Start-Up Grant Program for Undergraduate Institutions, the Wheaton College Alumni Association, and the National Institute of Standards and Technology (NIST). Commercial names of materials and apparatus are identified only to specify the experimental procedures. This does not imply a recommendation by NIST, nor does it imply that they are the best available for the purpose. LA035728I (54) Ru¨diger, M.; Haupts, U.; Moore, K.; Pope, A. J. Biomol. Screening 2001, 6, 29-37. (55) Mirzabekov, T.; Silberstein, A.; Kagan, B. Methods Enzymol. 1999, 294, 661-74.