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* Department of Physics and Astronomy, and Department of Biochemistry, University of British Columbia, Vancouver, British Columbia, Canada
Correspondence: Address reprint requests to Andre Marziali, Tel.: 604-822-5489; Fax: 604-822-5324; E-mail: andre{at}physics.ubc.ca.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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-hemolysin) housing a DNA probe with an avidin anchor at the 5' end and a nucleotide sequence designed to noncovalently bind a specific single-stranded oligonucleotide at the 3' end. The 3' end of the DNA probe is driven to the opposite side of the pore by an applied electric potential, where it can specifically bind to oligonucleotides. Reversal of the applied potential withdraws the probe from the pore, dissociating it from a bound oligonucleotide. The time required for dissociation of the probe-oligonucleotide duplex under this force yields identifying characteristics of the oligonucleotide. We demonstrate transmembrane detection of individual oligonucleotides, discriminate between molecules differing by a single nucleotide, and investigate the relationship between dissociation time and hybridization energy of the probe and analyte molecules. The detection method presented in this article is a candidate for in vivo single-molecule detection and, through parallelization in a synthetic device, for genotyping and global transcription profiling from small samples. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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), uncovering the temporal dynamics of such networks will necessitate time-resolved, single-cell measurement methods that can be carried out in vivo. As the operation of such networks cannot always be synchronized over a large ensemble of cells, ensemble average measurements will not be sufficient for many systems. The challenges in developing the desired assays are numerous: sufficient signal/noise ratio must be obtained, perturbations to cell function must be minimized, and a large variety of molecules must be detected with great specificity.
Although these arguments point to the need for in vivo detection and measurement techniques, near-term improvements to measurement of gene expression using DNA microarrays would also represent progress toward these goals. At present, nonspecific hybridization in microarrays results in low signal/noise ratios, even under highly stringent conditions (Schuchhardt et al., 2000). In addition, practical constraints on fluorescence detection methods used in microarrays set a lower limit on the number of molecules required for detection.
We present a first demonstration of a transmembrane single molecule sensor (Kasianowicz, 2002) with implications to a broad range of engineered transmembrane molecule detectors. By employing electronic molecule detection within a nanopore, we achieve a high signal/noise ratio and specific detection of oligonucleotides with single base resolution. Although many technical issues must be addressed to develop this sensor for in vivo applications, it has a number of advantages that make it a compelling candidate. The sensor is assembled in a lipid bilayer, and detects molecules on the side opposite to assembly, opening the possibility of assembling the sensor on the membrane of a living cell and assaying molecules in the cytoplasm. Also, unlike other in vivo detection techniques currently employed (e.g., GFP-tagging; Tsien, 1998, or lacZ-reporters; Miller, 1972), the nanosensor does not require genetic modification of the cells being observed, nor does it require introduction of foreign molecules into the cell. Development of synthetic nanopores (Li et al., 2001; Storm et al., 2003) and nanoporous membranes (Siwy and Fulinski, 2002) may allow extension of this work to the development of hybrid organic-synthetic nanosensor devices for transcript profiling or genotyping.
Our prototype nanosensor takes advantage of the geometry of the -hemolysin (a-HL) nanopore (Song et al., 1996): the narrowest part of its aqueous channel allows passage of single-stranded DNA (ssDNA), but is impassable to double-stranded DNA (dsDNA) (Kasianowicz et al., 1996). As described in Materials and Methods, a single a-HL pore is formed in a lipid membrane, and a probe designed to bind specifically to the molecule of interest is inserted into the pore by an applied electric potential, so that the binding region of the probe protrudes through the opposite side of the pore. Throughout the process, the ionic current is measured and monitored. A sudden step-like decrease in the current through the pore signals successful insertion of the probe. Binding of the analyte molecule to the probe traps the probe in the pore, even when the applied potential is reduced or reversed. Reversal of the potential tends to pull the probe from the pore, eventually forcing probe-analyte dissociation and allowing the probe to exit the pore, thus returning the ionic current to the open pore value. By observing kinetics of unbinding events, we are able to quantify parameters of the interaction between probe and analyte molecules.
Previous work on DNA detection in the a-HL pore has focused on analyzing the ionic current signature as DNA translocates through the pore (Kasianowicz et al., 1996, Akeson et al., 1999, Meller et al., 2001), a very difficult task given the translocation rate (
1 nt/µs at 100 mV) and the inherent noise in the ionic current signal. Higher specificity has been achieved in nanopore-based sensors by incorporation of probe molecules permanently tethered to the interior of the pore (Howorka et al., 2001a,b; Movileanu et al., 2000). However, the probe-analyte dimer must be small enough to enter the pore lumen. Similar recent work requires the probe and analyte molecules to be initially present as a duplex on one side of the membrane (Sauer-Budge et al., 2003). Other schemes require the molecule being analyzed to be of a specific shape or configuration to increase analysis times (Vercoutere et al., 2001, 2003).
Our sensor differs from previous approaches in several important details: limitations from translocation rate are avoided in this work since detection is based on the presence or absence of the probe strand inside of the pore, with a signal/noise ratio >50. The probe molecule extends across the membrane to access target molecules on the opposite side, allowing for true trans-membrane detection. The probe is also separate from the pore, allowing for easy modifications in probe design with unmodified biological or synthetic pores; the analyte molecule does not translocate through the pore, allowing the analyte-probe duplex to be larger than the pore. The probe in our sensor is uniformly charged, yielding greater control over the dissociation force and giving greater access to information on the energetics of interaction between probe and analyte.
Unlike the vast majority of biochemical assays that are colorimetric, fluorescent, or chemiluminescent, nanopore detection only requires electrical measurement of ionic current through the pore, and requires no optics or fluorescent labels. The ability to detect single unmodified analytes makes the nanosensor less likely to interfere with cell function by depleting or altering analytes, making it an excellent candidate for in vivo measurement. With further adaptation of probes, it may be possible to assay specific protein or other biomolecule levels by incorporating sequences for RNA aptamers into the probes. Although many challenges need to be overcome for eventual in vivo application of this sensor, it should be noted that single channel recordings have been successfully taken from -hemolysin incorporated into Lettre cells (Korchev et al., 1995). The general technique we describe may also be amenable to in vitro applications such as genotyping or transcription profiling with availability of suitable nanopore membranes.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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; Nakane et al., 2003; see also Fig. 1, this article). A lipid bilayer is formed across a 50-µm hole in a Teflon tube which connects two reservoirs. The reservoirs and Teflon tube are filled with microfiltered buffered 1 M KCl (10 mM HEPES, 1 mM EDTA, pH 8.0) as the conducting medium. Each reservoir contains a silver chloride-coated electrode from a patch-clamp amplifier (Axopatch 200B, Axon Instruments, Foster City, CA). Data acquisition software and hardware (Labview, National Instruments, Austin, TX) is used to record the applied electric potential and ionic current data from the experiment and to apply various electric potential profiles across the pore. All data is low-pass filtered at 10 kHz and digitized at 50 kHz for analysis. The pore is formed by addition of monomeric -hemolysin (CalBioChem, San Diego, CA) to one side of the bilayer (the cis side). Pore formation is detected by applying a 100-mV electric potential across the bilayer with the anode on the trans side of the membrane, and waiting for a stepwise increase in the measured current from 0 pA to 100 pA. The cis chamber is then rinsed with fresh buffer solution to prevent formation of other pores. Under these conditions, the open pore shows a forward resistance of 1 G
. For tests of the sensor, 14-mer single-stranded DNA analyte (MWG Biotech, High Point, NC) was added to the solution on the trans side of the pore before bilayer formation at 2 µM.
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4 G. The probe is held in this position (1 s) to allow time for an analyte molecule to hybridize to the probe on the trans side of the pore. To determine whether analyte is bound to the probe, we lower the potential across the bilayer to +10 mV. At this potential, unbound probes tend to quickly exit the pore, resulting in a return to a low-resistance state (1 G). This step is performed only to avoid collecting data on events that do not result in analyte capture. Although some unbound probes remain in the pore beyond the duration of the 10-mV hold time, this results in very short-lived events that do not contribute to the average event lifetime as calculated based on dominant timescales. Accumulation of statistics on the number of probe escapes during the 10-mV hold can yield information on analyte concentration, although this is not pursued in this article.
Probes bound to analyte molecules, however, remain trapped in the pore by the probe-analyte duplex. The applied potential is then reversed (anode on cis side) to –30 mV to –90 mV, which adds a force tending to separate the probe from the analyte and withdraw the former from the pore. In the reversed state, the blocked channel impedance is 4–10 G whereas the open channel impedance is 1.5 G. With continued application of this potential, the bonds forming the analyte-probe duplex eventually dissociate, allowing the probe to escape and returning the pore to a low impedance state.
The analyte molecules used in the study consisted of fully complementary 14-mer oligonucleotides and various other strands with single nucleotide mismatches within the sequence as shown in Table 1.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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), which in this case is assumed to be the direction of the pore axis:
 | (1) |
xbarrier. xbarrier is the energy barrier width and thus related to the distance by which the molecules must be separated along the reaction coordinate for dissociation to occur. Repeated measurements of toff for the same molecule under different applied forces f (controlled by the applied reverse potential) will yield details of the dominant free energy barrier in the analyte-probe bond energy landscape in one dimension along the pore axis. For large values of f, it is possible that outer barriers will be sufficiently lowered to uncover inner barriers (Evans, 2001).
Probe-pore interaction
Before applying the sensor to oligonucleotide detection, it was necessary to calibrate it by measuring the applied potential that corresponds to zero force applied to the probe-analyte duplex. The free energy cost associated with confinement of the probe adds an entropic recoil force (Turner et al., 2002), so that at zero applied potential we expect a net force tending to remove the probe from the pore. To estimate this entropic recoil force we measured the probe escape time as a function of applied potential without analyte molecules present. We captured the probe at +200 mV, decreased the potential to a small preset forward voltage, and measured the time to probe escape. The mean escape time for each voltage tested is shown in Fig. 3.
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where U is the increment in the barrier height resulting from the applied potential, and U0 is the entropic cost of confining the probe to the pore. Of the 65 nucleotides composing the probe, we estimate from a-HL dimensions (Song et al., 1996) that 40 are on the trans side of the membrane, whereas 12 are located in the narrow region of the pore, across which the majority of the electric potential falls (Meller et al.). A first approximation of the energy barrier height increment is 
where z is the fractional average charge per nucleotide, and 
is (25 mV)–1 at 20°C. Comparing this to the measured exponent 0.815 mV–1 and assuming that U = –U0 at Vt = 10.3 mV, yields the values z = 0.4, tD 3 ns, and U0 8 kbT for the free energy cost of confining the probe to the pore, which is in close agreement with other experimental estimates (Henrickson et al., 2000). Considering that at Vt 10 mV the free energy cost of probe confinement and the electrostatic energy gain of probe escape are balanced, we use this potential as a baseline for the effective force applied to the probe-analyte bond.
Detection and identification of oligonucleotides differing by a single nucleotide
To test the sensor's ability to detect and distinguish oligonucleotides, we exposed it to solutions of the analytes listed in Table 1 and recorded the binding event characteristics associated with each solution to see whether the characteristics of the fully complementary sequence (14pc) were distinguishable from oligonucleotides that differed from this sequence by a single nucleotide. We assembled the sensor as previously described with probe molecules on the cis side of the membrane and analyte molecules on the trans side. Typical experimental measurements of unsuccessful and successful analyte captures are shown in Fig. 2.
Fig. 4 shows the cumulative distribution of event lifetimes for the 7c analyte at –55 mV, representing the probability Pesc(t) that the event will end (resulting in escape of the probe), within a given time. We used a non-negative least-squares error fit with regularization (Whitall and MacKay, 1989) on a multiexponential expansion of Pesc to find the dominant timescales for each molecule and potential, with Pesc taking the form
 | (2) |
The function minimized for the fit is
 | (3) |
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Although the multiplicity of timescales appearing in the data may suggest multiple processes for probe escape after analyte binding, the identification of these processes will require additional data and is beyond the scope of this article. We assume for now that the dominant process required for probe escape is unbinding of the probe-analyte duplex. Analyte molecules incorrectly hybridized to the probe may contribute to short timescale events, as do the occasional delayed escapes of unbound probe molecules. Occasional long-lived events may result from the probe or probe-analyte duplex becoming lodged in the pore leading to blockages lasting seconds to minutes, as previously observed in studies of ssDNA translocation through the a-HL pore (Kasianowicz et al., 1996).
Averaged dominant timescales for the molecules in Table 1 are plotted as a function of the applied potential in Fig. 5.
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where j is the set of coefficients pertaining to dominant timescales. Spurious, poorly resolved long and short-lived timescales with small amplitudes of the coefficient are not counted in this average under the assumption that they belong to events unrelated to probe-analyte dissociation as described above. An example of selection of timescales for this average is shown in the inset of Fig. 5. Each point in this plot represents a timescale for the 10c molecule, with the diameter of the circle indicating the relative amplitude of the coefficient. All points below the dotted line are rejected; those above the dotted line are included in the average.
Analyte molecules differing in sequence by a single nucleotide yield measurably different binding event lifetime versus potential characteristics, and can be discriminated using this sensor if enough events are detected. Lifetimes decrease exponentially with applied potential as expected, and, in general, are shorter for the molecules that are not perfectly matched to the probe. These lifetime trends are supported by earlier unzipping experiments using nanopores with much longer complementary regions (50 nt) and much larger mutations (four mismatches) (Sauer-Budge et al., 2003). The lifetime versus potential curve for the 10c molecule near 80 mV deviates from a simple exponential relationship. This may indicate that multiple energy barriers are associated with unbinding of 10c from the probe, and that near 80 mV the outer barrier has been lowered below an inner barrier, changing the slope of the lifetime-potential relation beyond that point.
The natural logarithm of the expected event lifetime from Eq. 1 is
 | (4) |
l, the effective charge per nucleotide is z, and there are N nucleotides present in the region over which the potential drop occurs (l).
The slope of the lifetime versus potential relationship for each molecule is proportional to the width of the energy barrier for probe escape projected along the reaction coordinate, with steeper slope implying a wider barrier (Evans, 2001). Assuming that each nucleotide of the probe strand is separated by 0.5 nm, N/l = 2 nm–1 and the slope is zx/12.5 mV–1. We propose that fracture of the duplex occurs as cooperative unbinding of N bonds in series (Strunz et al., 1999); the width of the energy barrier x then corresponds to Nxb, where xb is the width of the individual bond barriers and the distance traveled by the probe with each bond dissociation. Using our estimate of z = 0.4 we find x 4–6 nm for 14pc and 1a, and xb 0.35 nm. Given a duplex base separation of 0.32 nm and a ssDNA base separation of 0.5 nm, we expect xb 0.2 nm. Previous studies of short DNA duplex dissociation under force (Strunz et al., 1999) yield smaller values (0.12 nm), although under different conditions.
For the 10c and 7c molecules, the calculated energy barrier widths are x 1.5–4 nm. This is possibly a result of the fact that 7c and 10c contain two shorter regions of complementary sequence, whereas 14pc and 1a have a single contiguous region of complementary sequence. It is plausible, particularly given the high reverse potential behavior seen in the 10c data, that fitting a single exponential to the lifetime-potential curve of the 7c and 10c molecules over the entire measurement range obscures details of the energy barrier.
The relationship between the intercepts of the curves in Fig. 5 and the binding energies expected from Mfold (Zuker, 2003) calculations is shown in Fig. 6. Based on our estimates of the applied potential necessary to compensate for the free energy of probe confinement, intercepts are calculated at a forward potential of +10 mV, where U = 0. Based on Eq. 4, the relationship between these intercept values and binding energies in units of kbT (at 20°C) should yield a slope of 1 with an intercept of ln(tD). The slope obtained from this plot is 0.75 +0.38, –0.3, and the large uncertainty in the intercept values precludes any useful estimate of tD.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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The data collected in this initial demonstration contains many details that have not been addressed in this manuscript. In particular, the multiple timescales for the binding event duration (that manifest themselves in the escape probability of the probe) hint at the possibility of multiple processes for unbinding of the probe-analyte duplex and escape of the probe. Investigation of these processes and their biophysical implications is an exciting priority for future work, and a stepping stone to redesign of the probe to reduce the number of timescales for probe escape—thereby increasing the likelihood that the sensor will be successful at distinguishing oligonucleotides that are present in mixes of other molecules.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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We gratefully acknowledge the Natural Sciences and Engineering Research Council of Canada for funding.
Submitted on January 19, 2004; accepted for publication March 29, 2004.
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The amplitude of the coefficients ai is plotted on the vertical axis, whereas the timescales ti used in the fit are plotted on the horizontal axis. For the 7c molecule data shown in this figure, the dominant timescales are 2.7 ms (with ai = 0.58) and 8.4 ms (ai = 0.24). Inset b shows a similar fit obtained on the Pesc data for the 14pc molecule at –55 mV. Dominant timescales are 100 ms (ai = 0.39) and 1.3 s (ai = 0.34).
Plotting this against the molecule binding energy in units of kbT as predicted by Mfold is expected to yield a linear relationship with a slope of 1. A linear fit through the data yields a slope of 0.75 +38, –0.3.