Lipid Nanobilayers to Host Biological Nanopores for DNA Translocations

Dec 7, 2012 - ABSTRACT: We characterize a recently introduced novel nanobilayer technique. [Gornall, J. L.; Mahendran, K. R.; Pambos, O. J.; Steinbock...
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Lipid Nanobilayers to Host Biological Nanopores for DNA Translocations Kerstin Göpfrich,†,‡,§ Chandrashekhar V. Kulkarni,†,‡ Oliver J. Pambos,†,‡ and Ulrich F. Keyser*,‡ ‡

Biological and Soft Systems, Cavendish Laboratory, University of Cambridge, JJ Thomson Avenue, Cambridge CB3 0HE, United Kingdom § Friedrich-Alexander-Universität Erlangen-Nürnberg, Staudtstraße 7, 91058 Erlangen, Germany S Supporting Information *

ABSTRACT: We characterize a recently introduced novel nanobilayer technique [Gornall, J. L.; Mahendran, K. R.; Pambos, O. J.; Steinbock, L. J.; Otto, O.; Chimerel, C.; Winterhalter, M.; Keyser, U. F. Simple reconstitution of protein pores in nano lipid bilayers. Nano Lett. 2011, 11 (8), 3334−3340] and its practical aspects for incorporating the biological nanopore α-hemolysin from Staphylococcus aureus and subsequent studies on the translocation of biomolecules under various conditions. This technique provides advantages over classical bilayer methods, especially the quick formation and extended stability of a bilayer. We have also developed a methodology to prepare a uniform quality of giant unilamellar vesicles (GUVs) in a reproducible way for producing nanobilayers. The process and the characteristics of the reconstitution of α-hemolysin in nanobilayers were examined by exploiting various important parameters, including pH, applied voltage, salt concentration, and number of nanopores. Protonation of α-hemolysin residues in the low pH region affects the translocation durations, which, in turn, changes the statistics of event types as a result of electrostatics and potentially the structural changes in DNA. When the pH and applied voltage were varied, it was possible to investigate and partly control the capture rates and type of translocation events through α-hemolysin nanopores. This study could be helpful to use the nanobilayer technique for further explorations, particularly owing to its advantages and technical ease compared to existing bilayer methods.

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alumina (Al2O3),12 polyethylene terephthalate (PET),13 or graphene.8,14 Despite mechanical and durable strength, entirely synthetic nanopores lack simplistic and low-cost preparation methodology, especially when atomic-level precision is required. Hybrid nanopores evolved from the above two types of nanopores facilitate high-throughput applications for biotechnological and potential genomic applications.1,7,15 Nanopores largely serve as bio- and chemical sensors. Their applications are not limited to mere identification of molecules but have traversed well beyond this to account for their quantification, sequencing, structural depiction (e.g., length and secondary structures), folding states,16 residual/ligand-directed binding, orientation-specific discrimination, and biomolecular separation.1 Biological nanopores embedded in lipid bilayers are ideally suited for the above applications. Our group has recently developed a novel “nanobilayer technique”,9 where biological nanopores are incorporated into a free-standing lipid bilayer situated on the nanotip of a borosilicate or quartz capillary. This method provides certain advantages over conventional Teflon-based black lipid membranes (BLMs).17,18 First, the bilayer diameter can be tuned down

INTRODUCTION An emerging yet rapidly popularizing field of “nanopores” involves purely biological, purely synthetic, and intermediate nanopores.1−3 Even though the biological nanopores originate from natural sources, they are prone to a high degree of tunability, thereby offering many more benefits for understanding cellular processes as well as for advanced research applications, such as biomolecular sensing and sequencing.1−3 Biological nanopores are based on multimeric proteins, usually transmembrane proteins, which assemble into a channel embedded in a lipid bilayer. The activity of these highly specific nanopores is regulated by a single or a combination of parameters, such as voltage, osmotic gradient, ligands, or mechanical force. These nanopores, naturally occurring in cellular and subcellular membranes, play a vital role for cell functioning. There are more than 300 biological channels;4 however, α-hemolysin (α-HL) from Staphylococcus aureus is one of the best-studied5,6 and most widely used biological nanopores. In the lipid bilayer, α-hemolysin monomers assemble into a heptameric complex.7,8 Other biological pores employed for nanopore applications include outer membrane porin F (OmpF)9 and Mycobacterium smegmatis porin A (MspA) channels. Synthetic, solid-state nanopores are normally fabricated using finely focused ion or electron-beam sculpting10 upon a base material made of silicon nitride (SiN),3 silicon,11 © 2012 American Chemical Society

Received: October 19, 2012 Revised: December 6, 2012 Published: December 7, 2012 355

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to ∼200 nm (for borosilicate nanocapillaries) or 30−40 nm (for quartz nanocapillaries)19 in contrast to micrometer length scales for BLM techniques. Second, the bilayer stability is significantly enhanced. Third, the bilayer can be formed and broken within seconds (repeatedly up to ∼50 times). Details of this method have been published earlier.9 In this report, we explore the nanobilayer technique further to optimize the properties of a biological nanopore for translocation of biomolecules, namely, single-stranded DNA homopolymers. We start with a description of technological modifications for the production of giant unilamellar vesicles (GUVs), the basis for bilayer formation. To gain more control over the translocation process and to prove its at least equal suitability and potential advantages with respect to other conventional nanopore methods, we have tested our nanobilayer technique by varying many important parameters over a wide range, including the salt concentration, applied voltage, pH, and type of DNA.

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EXPERIMENTAL SECTION

Materials. Lipids, 1,2-diphytanoyl-sn-glycero-3-phosphatidylcholine (DPhPC), and cholesterol (Ch), used for the preparation of GUVs, were purchased from Avanti Polar Lipids (Alabaster, AL) and stored at −20 °C. Other chemicals, namely, chloroform, potassium chloride (KCl), 2-(N-morpholino)ethanesulfonic acid (MES) (a buffer for pH 6.0), Tris(hydroxymethyl)aminomethane (Tris) (a buffer for pH 8.0), ethylene glycol, and sorbitol, were obtained from SigmaAldrich, Ltd. (Dorset, U.K.). The ion channel protein α-hemolysin in powder form was also purchased from Sigma-Aldrich. Single-stranded DNA homopolymers of adenine [poly(dA)75] and thymine [poly(dT)75] were ordered from Invitrogen Life Technologies, Ltd. (Paisley, U.K.). Water (ddH2O) used for all experimental studies was distilled and deionized using a Millipore unit (Millipore Corporation, Billerica, MA). Producing GUVs for Nanobilayers. GUVs for the preparation of nanobilayers were produced in a custom-built unit (Figure 1) using a method suggested by Angelova et al.20 For this, we designed (using AutoCAD software, AutoDesk, Inc., Novi, MI) a lipid spreading unit and a GUV preparation assembly, which was three-dimensionally (3D) printed in polymer material (material type: confidential from the company) by 3dprintuk (Oxford, U.K.). A 10 μL solution (50 mg/ mL) of lipids (DphPC/Ch ∼ 10:1) dissolved in chloroform was poured on an indium tin oxide (ITO)-coated glass slide (Visiontek Systems, Ltd., Chester, U.K.), which was placed in the bottom part of the lipid spreading unit. The top part with chamfer (tapered end) can slide over to form a practically uniform thin lipid film on the ITO surface (Figure 1a). Two such ITO slides were placed in the GUV preparation assembly (Figure 1b) filled with 1 M sorbitol (with the lipid-coated surfaces facing each other). The ITO slides were then connected to a function generator via a conducting copper tape. The tiny (1 mm thick) slot inside the chamber of the GUV preparation assembly allowed the slides to be positioned such that they did not touch each other. The whole assembly was kept in an oven at 37 °C for 2 h, while an alternating current (AC) amplitude of 2 Vp−p and a frequency of 5 Hz were applied. This method produces ∼1.8 mL solution of GUVs (size range of 1−100 μm), which can be used for up to 1 week when stored at 4 °C (Figure 1c). Formation of Nanobilayers. Nanobilayers were formed on the nanoscale tip of glass capillaries (Figure 2). Such nanocapillaries (typical diameters of ∼200 nm) were obtained by pulling borosilicate glass capillaries (outer diameter of 0.5 mm and wall thickness of 0.064 mm from Hilgenberg GmbH, Germany) with a laser pipet puller (P2000, Sutter Instruments, Novato, CA), as shown by schematics in Figure 2a. GUVs were added to the salt solution (KCl, concentration of 0.3 M unless mentioned) placed on a glass coverslip. The latter was positioned on the custom-built microscope, which allows for direct observation of GUVs and a capillary. After the Ag/AgCl electrode was

Figure 1. Laboratory-scale GUV production unit for the preparation of nanobilayers: (a) ITO slide is placed in the 3D-printed unit, where a 10 μL drop of a lipid solution is spread into a thin uniform lipid film. (b) Lipid-coated dry ITO slides are then placed in a GUV preparation assembly containing 1 M sorbitol. A capacitor is formed when the conducting sides of ITO slides are connected to a function generator. GUVs are thus produced at 37 °C within 2 h. (c) GUVs stored in plastic eppendorf tubes at 4 °C are large enough (∼1−100 μm) to produce locally flat lipid nanobilayers in nanocapillaries. immersed, the capillary was backfilled with the same concentration of salt solution and then connected to an amplifier head stage (CV203BU, Axon Instruments, Inverurie, Scotland, U.K.) attached to a semi-automated micromanipulator (PatchStar Micromanipulor, Scientifica, U.K.). Nanobilayers were formed primarily by inducing a suction force via a plastic syringe, as shown in Figure 2b. Bilayer 356

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conditions. A good nanobilayer shows a characteristic current of 6−10 pA at 100 mV in 0.3 M KCl solution at pH 6.0 and is considered to be ready for reconstitution of biological nanopores, such as α-hemolysin. An insertion of a single αhemolysin can be detected as a jump in a current value, as shown in Figure 3a, because the protein channel facilitates the passage of ions through the nanobilayer. An insertion typically happened in about 20−40 min after the addition of protein solution. A resistance of ∼109 Ω and rectification in the I−V curve (plot is shown in Figure S1 of the Supporting Information) were seen, as expected for single α-hemolysin nanopore insertions. We can keep a single α-hemolysin pore stable in the nanobilayer for at least 1 h (corresponding plot is shown in Figure S2 of the Supporting Information). This enables us to use the same pore to perform various tests and experiments. Although it is a nanobilayer (area ∼ 31 400 nm2), we found that it can hold more than ∼400 α-hemolysin pores (ca. area occupied ∼ 2000 nm2) stably, whose insertion kinetics approximates to an exponential curve (Figure 3b). The concentration of 1 mg/L (for α-hemolysin solubilized in a 50:50 mixture of ethylene glycol and ddH2O) appears to be suitable to reconstitute fewer than ∼3 pores (note that 10 μL of the above solution was added to 50 μL of salt solution on a glass coverslip). After insertion of each α-hemolysin nanopore, a conductance was measured at 100 mV. It increases linearly with the number of α-hemolysin nanopores (Figure 3c) as well as with the salt concentration (Figure 3d). However, with pH, the conductance decreases marginally (Figure 3f). The change in conductance (ΔG) is higher at low pH values, which is ascribed to a protonation of amino residues at the lumen of an ion channel,23 for example, as elucidated in Figure 4. In general, ion channel conductance is altered by the pH values, as illustrated by Kasianowicz et al.24 Some of the aforementioned parameters directly affect the capture rates of charged analytes. We used well-characterized single-stranded DNA homopolymers [poly(dA)75 and poly(dT)75] as charged analytes to study their translocation properties through α-hemolysin nanopores as a function of these parameters. As shown in Figure 4, the capture rate of poly(dA)75 is more than an order of magnitude higher (150 s−1 μM−1) at pH 6.0 than at pH 8.0 (4.9 s−1 μM−1). The amino acid residues His 48, His 144, Lys 205, and Lys 266 are very likely to be protonated at lower pH values25 (Figure 4c). Although there is no significant lateral separation of the anionic and cationic permeation pathways, as observed for OmpF,25,26 the protonation is sufficient to influence the capture rates of charged analytes. The capture rate virtually follows this mechanism (protonation at low pH), thereby attracting more negatively charged analytes [poly(dA)75] at low pH values. The increase in the capture rate at lower pH values is clearly related to the increase in the number of protonated residues, as shown in Figure 4c. The capture rate does not only depend upon the pH but can also be significantly improved by applying increasing potentials. It is essential to acquire efficient capturing of analytes to facilitate molecular analysis, especially where biological nanopores are employed. A combination of low pH and high potential show the highest capture rates in our experiments (Figure 5a). At high pH, however, an increase in potential does not have a strong effect on capture rates, as demonstrated by the red data in Figure 5a. Only at voltages of 150 mV do we observe a noticeable increase in the capture rate at pH 8.0.

Figure 2. Preparation of nanobilayers: (a) Nanocapillaries were prepared by pulling thin glass capillaries with a laser pipet puller. Such a capillary was then mounted on the headstage attached to a micromanipulator setup, as shown in panel b. Subsequently, a solution of GUVs was added to the glass coverslip. Within a few seconds, a vesicle bursts onto the nanotip to form a nanobilayer. Sometimes suction (negative pressure) is applied using a plastic syringe to facilitate the nanobilayer formation. Sealing of the nanotip was detected by a sudden current drop, which confirms the bilayer formation, as shown in panel c. Upon bilayer formation, the open nanocapillary conductance (G) decreased from 80 to 0.1 nS. formation was clearly detected by the drop in a current and the typical seal resistance of 10−100 GΩ21,22 (Figure 2c). For more details, readers are directed to earlier published work.9

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RESULTS AND DISCUSSION Properties of α-Hemolysin in Nanobilayers. Once a nanobilayer is formed, it is stable for hours under experimental 357

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Figure 3. Properties of α-hemolysin in a nanobilayer: (a) Insertion of a single α-hemolysin pore in the nanobilayer. The current step clearly indicates the process of insertion. (b) Subsequent current steps designate further insertions up to ∼400 α-hemolysin pores in the nanobilayer. The inset shows distinct steps in the current that can be attributed to consequent insertion of α-hemolysin nanopores. Up to 13 steps are shown here. (c) Conductance measured for up to 5 pores shows a linear increase, which is also the case for the salt concentration, as shown in panel d. (e) Histograms of single pore conductances in 0.3, 0.5, and 1 M KCl (+10 mM MES at pH 6.0) determined for 181, 33, and 44 individual measurements, respectively, define Gaussian fits, as indicated by black curves. (f) Low pH values seem to facilitate passage of more ions, in turn increasing the relative conductance (ΔG = ΔI/V) of α-hemolysin nanopores. Dotted lines are drawn as a guide for the eyes.

“capture” is not essentially a translocation event. It can also be a collision or an interaction with the luminal sites of the pore (Figure 6). Previous studies suggest that ∼0.1% of DNA that collides with the pore is translocated27 at the voltages close to the threshold. In our study, current drop signals as a result of partial blockade of a pore (as evident from Figures 4 and 6) are called “events”. The amplitude of the spikes IBase − IEvent reflects the level of pore blockade, while the width is directly related to event duration tDwell. Those events with a large amplitude (>25% of the baseline current), crossing the blue block in Figure 6a are deemed to be translocations (for instance, I and III in Figure 6).27,28 The remaining low blockade events are collisions. The DNA is captured in the entrance of the pore, the so-called lumen, and exits from the same side, which causes a low-amplitude event, e.g., as specified by II in Figure 6. The software “Clampfit” (Molecular Devices, Inc., Sunnyvale, CA) was used to extract pore blockade and event duration from

The ionic strength is another important parameter that alters capture rates. We demonstrate this by increasing the concentration of KCl in our experiments from 0.3 to 1 M (Figure 5b). We find that at values above 0.3 M, the capture rates are considerably higher, usually sufficient for various translocation studies (Figure 5b). We can conclude that an increase from 0.5 to 1 M has only a small effect on the capture rate. It is interesting to note here that the nanobilayers are stable over this wide range of concentrations. Also, in this case, high potential and low pH values are favored to achieve better capture rates. Nonetheless, the voltage cannot be increased beyond 125 mV, otherwise “gating” may occur, closing the pore (see Figure S3 of the Supporting Information). Optimization of Translocations over Collisions. The capture rate of charged analytes by the α-hemolysin nanopore can be improved by optimizing certain parameters, as discussed earlier. It is now important to note that the detection or 358

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Figure 4. Effect of pH on the capture rate: Plots a and b represent current traces showing the frequency of DNA translocations. Dashed red lines indicate minimum threshold, above which events were considered as a “capture”. It is clear to note that the capture events counted per unit time are almost 2 orders of magnitude higher at (a) pH 6.0 than at (b) pH 8.0. (c) Schematic cross-section of the α-hemolysin pore elucidates the location of four known residues, which become protonated (indicated by red positive signs) because of the decrease in pH (from 8.0 to 6.0). The graph on the right side presents experimental data for the capture rate (at 1 M KCl and 125 mV) obtained at varying pH values. A corresponding number of charged residues is also plotted on the same graph. It is evident that the capture rate is significantly altered by the pH.

Figure 5. Capture rates regulated by increasing potential and salt concentration: (a) Capture rate is radically enhanced by applying potential, which causes major changes at lower pH values. (b) Changing the KCl concentration from 0.3 to 0.5 M increases the capture rate by about an order of magnitude, which slightly increases with a further increase in the salt concentration. The trend is also similar at pH 8.0. Error bars correspond to the standard deviation of the event count within a 30 s time interval. Dotted lines are drawn to guide the eyes.

the current traces. The software performs a threshold-based search by setting the values for the baseline current, the trigger level, and the re-arming value (see Figure 6a) and then scans the file for data that crosses the threshold. An “event” starts when the trace crosses the trigger and ends when the trace recrosses the trigger or crosses the re-arming level. Short current spikes that were affected by the cutoff frequency were rejected by the software. Resulting data imported into Origin

(Origin Lab Corporation, Northampton, MA) is plotted, as shown in Figure 7. A histogram of the dwell time distribution (panels a−d of Figure 7) typically has a slow exponential decay after the peak value (thus, a Poisson distribution). For collisions, which usually have shorter durations, the peak of the histogram is shifted to shorter times. The relative frequency of the event types (either collision or translocation) depends 359

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Effect of pH on Relative Pore Blockade. Although translocation events are clearly distinct from collisions, the blockade levels of translocation events are not necessarily monotonic concerning the relative blockades. The relative blockade, IB is defined as IB = Ievent/IBase. This normalization eliminates drift and allows for events to be compared independent of the pore conductivity and the applied voltage. A typical scatter plot for the translocation events is shown in Figure 8. Each point represents an event that was characterized by its duration, tDwell, and relative blockade, IB. These translocation events can be separated into two groups of translocations, indicated in red, group 1, and green, group 2. The translocations with lower average currents (IB = 0.110) are referred to as group 1 translocations, while translocations with an average current blockade (IB) of 0.186 are referred as group 2. Thus, the histogram of the relative blockades has two peaks that were well-fitted by the sum of two Gaussians (see Figure 8c). These results are in excellent agreement with previous findings.36 At pH 8.0 and an applied voltage of V = 125 mV, group 1 accounts for about 80% of the total number of events (Figure 8b). For the same experiment, when performed at pH 6.0, we observed that group 1 events decreased dramatically, whereas group 2 events increased explicitly (Figure 8a). However, the peak positions are not affected much by the change in pH. The blockade for group 1 events peaks at IB = 0.11, and the blockade for group 2 events peaks at IB = 0.2 (data not shown). Other intermediate pH values (pH 6.5, 7.0, and 7.5) fill the gap between the two pH values from above, as shown in Figure 8d, where the percentage of group 1 events shows a well-defined trend with respect to increasing pH. An analyte, as used in the studies described above, i.e., poly(dA), is prone to structural changes, for instance, with regards to variation in pH. These structural changes can be detected with nanopore translocation measurements. Here, we mention some of the well-known features: (1) The doublestranded structure (duplex) is favored under acidic conditions,32−34 because the protonation allows two adenine heterocycles to combine via base stacking.31 (2) At alkaline pH, poly(dA) tends to occur as a less structured single-stranded helix.31,37,38 (3) At intermediate pH, i.e., near the neutral pH range, mere base stacking could be expected, because the stacking does not reach toward complete duplex formation. However, there are more parameters that can affect the structural rearrangements, for instance, the length and type of DNA or salt concentration,32,39,40 which are beyond the scope of our current focus on the nanobilayer technique. The most likely cause for the two groups of translocation events in our experiments is due to base stacking of the DNA (feature 3, as described above). We can exclude the formation of a doublestranded DNA structure (feature 1), because it cannot be translocated through the narrow stem of the α-hemolysin pore. Any existing stacked structure must be broken before the translocation, because the diameter for poly(dA) is known to be larger than the diameter of the nanopore. The additional time for this process would shift the peak of the dwell time distribution to longer times. The observed time shift of around 80 μs between the peaks at pH 6.0 and 8.0 (see Figure 7f) is in good agreement with the time scale for unstacking poly(dA).40,41 Moreover, an energy barrier has to be overcome to enable unstacking of the bases. This would broaden the dwell time distribution, which results in larger decay constants (τ), as observed at acidic pH (about an order of magnitude higher for pH 6.0 compared to pH 8.0, as discussed in the last section).

Figure 6. Event distinction for DNA capture by α-hemolysin nanopore: (a) Ionic current trace with downward spikes corresponding to the interaction of single-stranded DNA with α-hemolysin. The red line indicates baseline current IBase corresponding to the open current of a pore. Dotted and dashed lines represent re-arming and trigger levels, respectively, that were used for event detection. As indicated by the sketches on the left, events with large amplitudes are considered as translocations (downward arrow), while others are more likely to be collisions (curved arrow). (b) Zoom onto typical events from panel a. Position of a dashed line along the y axis indicates the event current IEvent, and the width (along the x axis) of a spike corresponds to the dwell time tDwell.

upon various parameters,29,30 for instance, the applied potential and pH, as elucidated in Figure 7. The translocation of 75 bases of adenine [poly(dA)75] was studied at pH 8.0 and 6.0 for applied voltages between 50 and 125 mV (Figure 7). Other plots at 75 and 100 mV for pH 8.0 and at 75, 100, and 150 mV for pH 6.0 are shown in Figure S4 of the Supporting Information. At lower potentials, the proportion of collisions (with blockade currents of less than 25% of the open pore current) was higher than the proportion of translocations. At pH 8.0, 125 mV appears to be a threshold value, below which translocations were effectively suppressed. Only 2.8% of events were full translocations, while 48.9% translocations were found at low pH 6.0 for an applied potential of 50 mV. In other words, the translocations became more likely at much lower potentials under slightly acidic conditions. However, the dwell times are higher at low pH, although the event duration decreases upon applying higher potentials, as expected (Figure 7f). The increase of the translocation times at lower pH could be attributed to base stacking of DNA [poly(dA)75] as a result of protonation at low pH, causing longer times of translocation. This additional time needed during the translocation process corresponds to the DNA unstacking process happening before the actual translocation event. Stacking of adenine at low pH has been suggested earlier by Saenger et al.31 and many others.32−34 At both pH values of 6.0 and 8.0, the peaks of the dwell time distribution shift with increasing potential, as seen from panels a−d of Figure 7. This dependency is in accordance with Kasianowicz et al.35 The decay constant (τ) obtained by fitting an exponential curve to the dwell time distributions decreases strongly (δτ = −10.7 μs mV−1) at pH 6.0, while it has only a weak dependence (δτ = −1.8 μs mV−1) at pH 8.0 at increasing applied potentials (plot is shown in Figure S5 of the Supporting Information). Our results demonstrate that we can control the translocation time in our nanobilayer system by varying the pH and applied voltage. 360

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Figure 7. Enhancement of translocations over collisions: Histograms showing the dwell time distribution of translocation (red) and collision (blue) events. “n” indicates the number of events. (a) At 50 mV, the frequency of translocations is very low. It is clearer from the inset picture. It increases considerably at higher voltages, for instance, at 125 mV, as shown in panel b. (c) At low pH (pH 6.0), a high percentage of translocations was observed, even at low voltage (50 mV). The amount of translocations relative to collisions was increased even further at 125 mV, as shown in panel d. (e) Percentage of translocations was higher at low pH, while at pH 8.0, it shows a threshold value of 100 mV, above which more than 50% of the events were translocations. (f) Examination of the dwell time values shows that the events happen faster at high pH and high potentials. Dotted lines are drawn to guide the eyes.

DNA structure is purely driven by entropy for poly(dT). Camerman et al. proposed a single-stranded helical structure for poly(dT), with the bases pointing away from axis of the helix, which prevents base stacking.44 In contrast to our results for poly(dA) (Figure 9a), for poly(dT), there is only one DNA structure in solution, which results in a single peak in the blockade current histogram, as observed in our experiments (Figure 9b). We note here that the capture rates for poly(dA) and poly(dT) were practically the same for different pH values as checked for their 1 M KCl solutions at 125 mV. The data are shown in Figure S6 of the Supporting Information. The unstacked poly(dT) polymer can pass the α-hemolysin pore without having to undergo a structural change before the translocation. This leads to the observed shorter dwell times and small decay constants of the dwell time distribution. The fact that poly(dT) translocates even quicker and causes a lower

Thus, group 1 events could be interpreted as translocations of the linear poly(dA)75, while group 2 events could be interpreted as translocations of the stacked structure. This would explain the dominance of group 2 events at low (acidic) pH values. The possibility that the group 1 and 2 events are related to either of the two possible orientations of the phosphate backbone,42 3′ → 5′ or 5′ → 3′, can be ruled out in our case because we did not observe distinct events in the case of a different type of an analyte, poly(dT) (see Figure 9b). poly(dT) is known to show fewer structural changes under pH variation, as seen for poly(dA). Effect of the Type of Nucleic Acid on the Translocation Process. We will now compare and discuss our results for translocations of poly(dA) and poly(dT) to clarify the origin of the two groups in the poly(dA) results (Figure 9). Goddard et al.43 showed that poly(dA) displays an enthalpic rigidity consistent with base stacking, whereas a single-stranded 361

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Figure 8. pH effect on translocation event types: Scatter plot of translocation events for poly(dA)75 at (a) pH 6.0 and (b) pH 8.0. The number of group 1 events decreases and the number of group 2 events increases significantly at pH 6.0 compared to pH 8.0 and vice versa. (c) There is a clear increasing trend of the percentage of group 1 events as a function of pH, as indicated by the dotted line joining individual points.

Figure 9. Effect of type of DNA, poly(dA) and poly(dT), on translocation events: At low pH, poly(dA) undergoes structural changes, resulting in two distinct event types: group 1 and group 2, typical for unstacked and stacked structures, respectively, as shown in panel a. However, poly(dT) shows only one type of event, as shown in panel b, characteristic of the absence of a significant secondary structure changing the relative blockade level.

controlling reconstitution kinetics and process automation. The properties of α-hemolysin in the nanobilayer display monotonic behavior, for example, with regards to the conductance change as a result of the number of nanopores and salt concentration. We studied the translocation of singlestranded DNA as a function of the applied voltage, salt concentration, and pH by employing the novel nanobilayer technique. Our results regarding the dependence of translocations upon the applied voltage at pH 8.0 are in accordance with the literature, confirming the suitability of the nanobilayer technique. Some important observations for increasing voltages are as follows: the dwell time for the DNA translocation follows an exponential decay distribution with decreasing decay

relative blockade than the helical structure of poly(dA) may be directly related to the diameter of the helix.45

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CONCLUSIONS AND PERSPECTIVES Here, we showed that we can effectively reconstitute αhemolysin into lipid nanobilayers supported across 200 nm sized glass nanocapillaries in the pH range of 6.0−8.0. A single α-hemolysin pore could be kept stable in the membrane for a long time (1 h), enabling single-channel recordings. A nanobilayer technique used for the reconstitution of αhemolysin provides several benefits in terms of its simplicity of reconstitution, fast rate of repeatability, possibility of controlling the number of nanopores, and potentially 362

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constants; the peak of the dwell time distribution shifts to shorter times linearly; and the capture rate and the proportion of translocations relative to collisions increase as the potential increases. The capture rate is also enhanced by increasing the salt concentration. During our studies using the nanobilayer technique, we gained some interesting insights regarding the pH dependence of translocation. First of all, the overall capture rate increases by more than 1 order of magnitude as the pH is reduced from pH 8.0 to 6.0. At the same time, the rate of translocation is enhanced. This has been attributed to a change in the protonation state of α-hemolysin, which introduces positive charges at the cis entrance and at the constriction of the pore. In the case of pH 8.0, above 125 mV, the translocations are dominant and split into two groups that can be distinguished by their characteristic current blockades. Group 1 translocations with a blockade of around 90% are more probable, and their peak shifts to shorter durations relative to group 2 events. On the contrary, when the pH is reduced to 6.0, the group 2 events become dominant and the dwell time peaks at longer durations and decays slower. The existence of two secondary structures of poly(dA) appears to be the most suitable explanation for the observations at low pH. Group 1 events could correspond to the translocation of an unfolded DNA, which is a favored conformation of poly(dA) at non-acidic pH values. Group 2 events may then be related to a stacked structure, which must be unstacked before translocation. This fact was confirmed by replacing adenine by thymine, i.e., a different type of nucleic acid, poly(dT), which does not form stacked structures at low pH. This work adds important points to the knowledgebase of biological nanopores; moreover, it facilitates use of nanobilayers to study protein nanopores more efficiently and effectively.

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ASSOCIATED CONTENT

* Supporting Information S

Additional features of α-hemolysin reconstituted in the nanobilayer presented with corresponding graphs, enhancement of translocations over collisions at remaining voltages, trends of the decay constant (τ) at various applied voltages, and capture rate comparison of poly(dA) and poly(dT). This material is available free of charge via the Internet at http:// pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Telephone: +44-1223-33-7272. Fax: +44-1223-33-7000. Email: [email protected]. Author Contributions †

These authors contributed equally to this work.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by a grant from the Nanoscience E+ initiative and the Engineering and Physical Sciences Research Council (EPSRC). Oliver J. Pambos thanks the EPRSC for funding through the Physics of Medicine Initiative, University of Cambridge. Kerstin Göpfrich acknowledges Dr. Klaus Mecke from Friedrich-Alexander-Universität Erlangen-Nürnberg, Germany, for this collaboration work with Cambridge, U.K. Ulrich F. Keyser was partly supported by an Emmy Noether grant of the Deutsche Forschungsgemeinschaft. 363

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