Article pubs.acs.org/ac
A Method for Determining the Actual Rate of Orientation Switching of DNA Self-Assembled Monolayers Using Optical and Electrochemical Frequency Response Analysis J. Casanova-Moreno†,‡ and D. Bizzotto*,†,‡ †
Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, BC V6T 1Z1, Canada Advanced Materials and Process Engineering Laboratory, University of British Columbia, 2355 East Mall, Vancouver, BC V6T 1Z4, Canada
‡
S Supporting Information *
ABSTRACT: Electrostatic control of the orientation of fluorophore-labeled DNA strands immobilized on an electrode surface has been shown to be an effective bioanalytical tool. Modulation techniques and later time-resolved measurements were used to evaluate the kinetics of the switching between lying and standing DNA conformations. These measurements, however, are the result of a convolution between the DNA “switching” response time and the other frequency limited responses in the measurement. In this work, a method for analyzing the response of a potential driven DNA sensor is presented by calculating the potential effectively dropped across the electrode interface (using electrochemical impedance spectroscopy) as opposed to the potential applied to the electrochemical cell. This effectively deconvolutes the effect of the charging time on the observed frequency response. The corrected response shows that DNA is able to switch conformation faster than previously reported using modulation techniques. This approach will ensure accurate measurements independent of the electrochemical system, removing the uncertainty in the analysis of the switching response, enabling comparison between samples and measurement systems.
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to probe the kinetics of the switching in these systems. By varying the frequency of the applied AC potential, the DNA reorients between the lying down and standing up states at low frequencies, while at high frequency regimes the switching can no longer occur. At these high frequencies, the DNA remains essentially static regardless of the change in potential.11 To characterize this frequency-dependent response, Rant defines a cutof f f requency (fc) where the fluorescence signal amplitude becomes half of its value at low frequencies. Due to the drag of the solution surrounding the DNA, the cutoff frequency is influenced by the hydrodynamic radius of the “switching” molecule. Thus, modification of the labeled DNA strand alters the response, shifting the cutoff frequency. Besides using this technology to determine the concentration of a given analyte, it is also used to determine the size of bound target molecules.8 In this analysis, it is assumed that the hydrodynamic drag is the main factor limiting how fast the DNA can “switch” its configuration. Since the movement of the DNA is a response to a change in the charge (and hence in potential) at the electrode surface, the time taken to charge the electrode (electrochemical time constant, τE) can be important. Typically, low electrolyte concentrations are used in order for the electric field to
lectrochemical based biosensors for nucleic acids, proteins, and small target molecules have been demonstrated based on the formation of self-assembled monolayers of DNA on an electrode surface. Transduction of the binding or recognition event can involve a change in the redox, impedance, or optical characteristics, each requiring control over the electrode potential to improve signal-to-noise. One example of this motif is the creation of a SAM using DNA with a methylene blue (MB) redox moiety.1−5 Hybridization with the complementary strand can turn on or off the redox signal depending on the detection strategy required. Modification of the DNA with a fluorescent moiety has also been used for detecting recognition events based on a change in the fluorescence intensity with electrode potential.6−10 In this case, a potential is applied to an electrode modified with a fluorophore-labeled single DNA strand inducing a change in fluorescence intensity which originates from a combination of two effects: (i) a change in conformation of the DNA molecule as the negative charged phosphate backbone is either repelled or attracted from the electrode depending on its charge and (ii) fluorescence quenching by the metal surface which is dependent on the distance between the fluorophore and the metal. This system was originally proposed by Rant et al. as a sensing transductor in which the specific binding (either by the complementary DNA strand11 or using epitope protein tags8) produced a change in the rate and the extent of DNA reorientation or “switching”. Sinusoidal potential perturbations have been used © 2015 American Chemical Society
Received: October 20, 2014 Accepted: January 11, 2015 Published: January 12, 2015 2255
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propagate far enough away from the interface so as to exert the greatest amount of torque on the DNA. Large optical signal modulation is realized at the expense of large electrochemical time constants. Under these conditions, if the electrode charging time is comparable or longer than the time needed for the DNA to change its configuration, the frequency dependent fluorescence response will be convoluted with charging rather than purely reporting on the DNA hydrodynamics. Since the analytical signal is based on the change in frequency response caused by a change in the size of the molecule after binding, electrochemical effects must be ameliorated. Although initially very promising, this frequency based approach has failed to distinguish differences in target molecule size under certain conditions (e.g., high viscosity). Currently, a time-resolved evolution of the fluorescence signal is measured (corrected for the charging time), providing separate information on the “lifting up” and “lying down” processes.12−14 Removing the influence of the electrochemical time constant can be accomplished through the use of frequency response analysis (FRA) methods applied to the optical and electrochemical systems. In this FRA methodology, a modulated perturbation (potential) at a given frequency is applied to the system under study which, for linear systems, creates a modulated response (current or intensity) at the same frequency.15 A transfer function (i.e., the ratio of the modulated output and input signals) is typically used to characterize the system when working in frequency space. The transfer function is the Laplace transform of the impulse response function (IRF) for the system under study.16 Typically, the experimental system is composed of multiple components each having a transfer function. The net transfer function measured is the product of the transfer functions for each individual component. To determine the transfer function for the reorientation of the DNA SAM independent of the measurement system, deconvolution of the other components in the system (e.g., electrochemical charging time constant) is required. In this contribution, we employ a FRA methodology to study the dynamics of an electrode modified with a mixed selfassembled monolayer (SAM) containing a fluorophore-labeled single DNA strand. The electrochemical system is perturbed with a sinusoidal potential applied to the electrode, and the measured response is the change in fluorescence intensity. In a recent publication,17 we have pointed out that modulation techniques still have some advantages over their time-resolved counterparts. In particular, differences in the phase of the transfer function between different regions of the SAM-covered electrode can be used for characterization. In this Article, a procedure for accurately determining the frequency response curve of the DNA orientation changes with modulated potential on the electrode surface is outlined. We demonstrate that, by calculating the potential expressed at the electrode interface, the effect of the electrochemical time constant can be deconvoluted enabling accurate measurement of DNA reorientation rates for comparison across experimental setups. This approach is demonstrated for a low coverage DNA SAM, showing that the actual rate of DNA reorientation is much faster than previously reported.
Article
METHODOLOGY
Experimental Section. Preparation of the modified electrodes, as well as the in situ fluorescence measurement setup, has been described in detail recently.17 Briefly, multicrystalline Au beads created through the melting of a 0.5 mm diameter Au wire (Goodfellow, 99.95% purity) were used as working electrodes (WEs) in a spectroelectrochemical cell equipped with a 250 μm thick glass window as its bottom. These beads were modified with a 1 mM solution of 6mercapto-1-hexanol (MCH) in a pH 7.5 buffer solution composed of 10 mM 2-amino-2-hydroxymethylpropane-1,3diol (Tris, J.T. Baker) and 100 mM NaCl for 90 min. Subsequently, the beads were immersed in a solution of the DNA sequence (from 5′ to 3′) HS-C6-CTG-TAT-TGA-GTTGTA-TCG-TGT-GGT-GTA-TTT-AlexaFluor 488. This sequence was chosen such that secondary structures were not stable. Furthermore, the oligonucleotide was long enough to ensure separation from the surface which would yield a measurable change in fluorescence. The electrochemical and spectroelectrochemical measurements were performed in a three-electrode arrangement with Au bead working, SCE reference, and Pt coil counter electrodes, respectively. The potential was controlled with an Autolab PGSTAT30 potentiostat equipped with the FRA2 module. On the “high speed” bandwidth setting, the PGStat30 can provide potential waveforms with an attenuation of less than 15% up to 100 kHz. The electrolyte used was 10 mM Tris, pH ∼ 7.5 adjusted with HNO3 with the addition of supporting electrolyte (10 or 100 mM KNO3). The measurements done in low ionic strength introduce significant problems in the control of the WE potential at higher frequencies (>10 kHz) when using a salt bridge. A “fourth electrode”18 was introduced to prevent artifacts resulting from the high resistance in the salt bridge. This was implemented by connecting a Pt wire in the electrolyte solution to the reference electrode (RE) connector through a 7.17 nF capacitor (details in ref 17). The spectroelectrochemical cell was placed on the inverted microscope such that the electrode was at the focal point of the objective (5× and 40× water immersion). The surface was illuminated with wavelengths specific to excite Alexafluor488 (450−490 nm), and the fluorescence emission was collected using the same objective and filtered using a dichroic mirror (495 nm) and an optical filter (500−550 nm). The filtered light was directed through two collimated pinholes into a Newport 77348 photomultiplier tube (PMT). The fluorescence signal from the PMT was conditioned with a Stanford Research Systems SR570 low-noise current preamplifier. Low-pass filters of 100 Hz and 1 MHz were employed for the cyclic voltammetry (CV) and FRA experiments, respectively. The fluorescence FRA measurements were obtained after determining the appropriate DC potential and the amplitude of the AC potential perturbation. To ensure the measurements were within the linear response regime, cyclic voltammetry experiments were performed to find the potential range at which the fluorescence variation with potential is linear. Furthermore, these experiments also served to determine the potential range of stable response to ensure that desorption does not occur. The potential was cycled between +0.35 and −0.3 V at a rate of 0.100 V/s while continuously recording the signal from the PMT and the electrochemical current. Fluorescence FRA spectra were obtained by applying a DC potential of +0.150 V/SCE and superimposing an AC 2256
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Figure 1. (a) Fluorescence image of the DNA modified Au bead electrode using a 5× objective outlining the region of interest. (b) Fluorescence image of the region of interest at 40× with apertured illumination. (c) Fluorescence intensity and electrochemical current during potential cycling for two concentrations of electrolyte within a restricted potential range where the SAM is stable (100 mV/s, 0.18 cm2 estimated from optical measurements).
effects of the solution resistance in electroreflectance experiments.21−23 The measured transfer function is defined in eq 1, and with the ratio of the interface potential to the cell potential (EAC,int( f)/EAC,cell(f)), a modified transfer function Hint(f) can be defined
perturbation (25 frequencies between 30 Hz and 100 kHz with an amplitude of 0.200 V peak to peak). The measured experimental output is the transfer function (Hcell) defined as
Hcell(f ) =
FAC(f ) EAC,cell(f )
(1)
in which the fluorescence response (FAC( f)) is divided by the potential perturbation, EAC,cell(f) (both complex quantities resulting in a complex transfer function). EIS were collected immediately after the fluorescence FRA experiments using the same experimental configuration but employing a 5 mV rms perturbation. Fitting of the EIS data was performed using the Autolab Nova software. In order to simply and unambiguously represent an equivalent circuit, we adopt the Circuit Description Code (CDC). Originally proposed by Boukamp,19 it is composed of letters to represent the electrical components and nested parentheses to indicate connectivity. Elements enclosed in square brackets are connected in series, while those in parentheses are connected in parallel.20 Measurement of the optical transfer function of the measurement system using FRA was performed by replacing the spectroelectrochemical cell by a Panasonic LNG992CFBW blue LED and obtaining FRA spectra by applying, through the potentiostat, a DC potential of 3 V and an AC amplitude of 200 mV (p−p). Approach to Data Analysis and Deconvolution. To determine the actual transfer function for the DNA SAM, the electrochemical response must be deconvoluted from the experimental response. Our approach acknowledges that the rate at which the electrical interface develops a potential depends on its relative impedance and the frequency of the perturbation. Therefore, at some frequencies, only a portion of the potential applied to the electrochemical cell (Ecell) would be expressed at the electrode interface (Eint). The remaining potential is dropped across other components of the cell, such as the solution resistance. Since Eint is responsible for driving the DNA movement, the fluorescence modulation transfer function (H) should be recalculated using the potential that is present at the electrochemical interface, not the applied or cell potential. A similar approach has been used to eliminate the
H int(f ) ≡
FAC(f ) EAC,int(f )
−1 FAC(f ) ⎛ EAC,int(f ) ⎞ ⎜ ⎟ = ·⎜ ⎟ EAC,cell(f ) ⎝ EAC,cell(f ) ⎠
⎛ EAC,int(f ) ⎞−1 ⎟⎟ = Hcell(f ) ·⎜⎜ ⎝ EAC,cell(f ) ⎠
(2)
It is important to note that all of these quantities are complex numbers and thus they have associated with them a specific phase for each frequency which is needed to accurately calculate Hint. As part of the routine data analysis, the transfer function is divided by the average fluorescence intensity measured during the fluorescence impedance (IDC). This is useful for comparing different electrodes or regions on the electrode surface, since DNA coverage may be quite variable. Determining (EAC,int(f)/EAC,cell(f)) can be accomplished by measuring EIS after each fluorescence FRA acquisition and fitting the electrochemical response to a simple equivalent circuit. In order for the correction to be successful, it is imperative to have high quality impedance data and an appropriate equivalent circuit which requires optimizing the cell configuration.17
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RESULTS AND DISCUSSION The DNA modified electrode surface was analyzed using fluorescence imaging to identify regions on the electrode surface to be studied. A typical fluorescence image of this interface, taken at 0 V/SCE, is shown in Figure 1a. It shows a wide variety of intensities, with some regions containing a significant amount of DNA (large fluorescence) and other regions devoid of fluorescence, presumably due to a low surface concentration of DNA. Details regarding this difference are 2257
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Figure 2. (a) Fluorescence FRA measurements of a fluorophore-labeled DNA-containing SAM under the following set of conditions: f = 30 Hz to 100 kHz, EDC = +150 mV, EAC = 200 mV p−p, 10 mM Tris + 10 mM KNO3. (b) Electrochemical impedance measurements under the same set of conditions except EAC = 5 mV rms. Symbols are measured data, lines are fit to either (Cst[Rs(Qint[RintCint])]) or (Cst[Rs(Cint[RintQint])]) equivalent circuit.
given in a previous publication.17 The region of interest (ROI) outlined in Figure 1a was specifically studied by reducing the illumination aperture to 95 μm spot, as shown in Figure 1b. CVs of the electrochemical response show a typical RC type behavior without evidence of a faradaic reaction. The interface is stable in this potential window, as evidenced by the consistent capacitive current. The change in fluorescence intensity with potential (Figure 1c) is also stable, showing a nonlinear response characteristic over the large potential range. A linear potential region exists from 0.05 to 0.250 V/SCE which was used for the FRA measurements. Correction of Measured Fluorescence FRA. The magnitude and phase of the fluorescence transfer function (Hcell(f)) and EIS for a typical set of measurements are shown in parts a and b, respectively, of Figure 2. The Hcell( f) has a roll off with a cutoff frequency at ∼3 kHz in agreement with previous publications (Figure 5 in ref 11). At low frequencies, the phase of the fluorescence response is π, which is expected, since the fluorescence increases as the potential gets more negative. At high frequencies, the phase decreases first to π/2 and tends toward zero at the highest frequencies. The magnitude and phase of the electrochemical impedance is presented in Figure 2b. An equivalent circuit model describing the electrochemical cell is needed to calculate the value of EAC,int( f). A series RC circuit is typical for these nonfaradaic systems. The resistor represents solution resistance (Rs) and the capacitor represents the electrode surface, which we will represent as a generalized impedance (Zint) that is not defined. In addition to these components, stray capacitance Cst is also present in the measurement, indicated by the nonzero phase of the impedance at high frequencies. The source of this capacitance has been described by Fletcher.24 The general equivalent circuit used in the EIS analysis is shown in Figure 3a.
Calculating the value of (EAC,int(f)/EAC,cell(f)) requires that the values of Rs and Cst be extracted from the EIS data. This requires that the impedance of the interface Zint be described in terms of an equivalent circuit. A constant phase element (CPE) (Q) alone does not fit the EIS (Figure S1 and Table S1, Supporting Information). Placing a [RC] component parallel to this CPE (Figure 3b, Q[RC]) enables an excellent fit for all the systems measured (Figure 2b). The relative errors in the fitting were 40 kHz. The data sets for 10 and 100 mM KNO3 before and after correction are compared by normalizing the transfer function at 30 Hz (Figure 7). The EIS data show system responses that have time constants for charging the interface which vary almost 1 order of magnitude. The fluorescence FRA data sets, measured on the same electrode and on the same 95 μm spot, when corrected for the potential at the interface (Hint(f)) and normalized to the response at 30 Hz, result in a comparable transfer function. Hint( f) does not to show a significant decrease in fluorescence intensity modulation below 10 kHz. Furthermore, the measurements at 100 mM KNO3, while only showing 20% of the fluorescence modulation of the 10 mM case, have no significant drop off with frequency until close to 30 kHz. If characterized by the phase of the fluorescence response, the frequency where the phase crosses π/2 is just under 100 kHz. It is important to note that the electrochemical fitting errors are equally distributed across the whole frequency range, never exceeding 2% deviation (Figure S2 in the Supporting Information), too small to create artifacts in the calculated transfer function. Hint( f) is therefore a better representation of the DNA response to potential perturbation, demonstrating that the procedure used is able to correct the experimental data for the electrochemical time constant differences that would be expected when comparing different samples on different days or setups. These results also show that the DNA reorientation rates are rapid and, for the low surface concentration interface analyzed, do not show a substantial modulation decrease except for frequencies above 2260
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Figure 8. Comparison of the corrected fluorescence FRA response of a DNA containing SAM (symbols) with the response obtained using an LED as a potential-dependent light source. The different symbols represent 10 and 100 mM experiments depicted in Figure 7. The phase for the LED was shifted by π to facilitate comparison.
Figure 7. Fluorescence FRA as measured (filled symbols) and corrected (open symbols) for potential across the interface (solid lines). Magnitudes and phases of these transfer functions are shown in parts a and b, respectively. Results for 10 and 100 mM KNO3 concentrations. Fluorescence FRA magnitudes have been normalized using their values at 30 Hz. The corrected data for measured signals that are below 5% (dashed line) of the applied potential are not shown.
electrical time constant >100 kHz (EIS for LED shown in Figure S4, Supporting Information). Within the uncertainty of the experiments, the observed DNA switching response (Hint( f)) is similar to the LED response. Since LEDs can be modulated at frequencies greater than 1 MHz, the decrease in the modulation amplitude and phase must be a consequence of the optical measurement system. Work is currently underway to increase this bandwidth. Nevertheless, the results show that, if Hint( f) is corrected for the optical measurement transfer function, the rate of the DNA switching response is >100 kHz. Relevance to Published Data. Frequency domain analysis of the DNA switching response to potential perturbations has been published8,11 for an electrochemical system that uses gold disc electrodes with a Pt quasi reference electrode. The SAM was composed of a low surface concentration of ssDNA (72mer) with the surface passivated with MCH. The complementary DNA strand modified with a digoxigenin protein binding tag was also used to make a dsDNA SAM that would bind the protein. The cutoff frequencies for dsDNA switching with and without digoxigenin were determined to be 18 and 11 kHz, respectively, values that are comparable with the uncorrected response presented in this work. Also included in that paper was the EIS of the interface fitted to a simple [RC] circuit. Figure 9 shows that data from refs 8 and 11 compared to the result after correcting for the electrochemical time constant following the methods used in this work. Using a simple [RC] circuit results in a fluorescence modulation (circles) that closely follows the potential drop at the interface (line) in the absence of sheep immunoglobulin G (IgG). The corrected response starts to decrease at f > 20 kHz. Notably, the cutoff frequency value obtained after correction is more in line with the kinetics reported using their most recent timeresolved methods (which compensate for the electrochemical
20 kHz in contrast to some published results8,11 but in line with recent time-resolved measurements.26 It has been previously reported that the frequency response of these layers may not be homogeneous over the whole surface and notably depends on the density of DNA in that local region. The question arises then, whether the values calculated using spatially averaging electrochemical measurements are useful to estimate the potential drop across distinct regions of the interface. The Rs and Cst values reflect the solution resistance and geometric arrangements of the leads and connections, and are not expected to vary with the layer DNA content. The capacitance of the interface, on the other hand, is more likely to be affected by the layer composition, but at the low DNA mole fractions employed in this work (40 kHz). This lack of influence of electrolyte concentration suggests that the high frequency measurements may be limited by the optical measurement system. Transfer function characteristics were performed in which the DNA-modified electrode was replaced by a lightemitting diode (LED). Figure 8 presents the collection of Hint(f) values for the DNA SAM (symbols) over top of which is shown the transfer function determined for the LED (line, shifted by π). The modulation of the LED was performed with the potentiostat, and the EIS analysis showed the LED had an 2261
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CONCLUSIONS Measurement of the frequency response of a potential driven change in DNA−SAM orientation over a wide frequency range (30 Hz to 100 kHz) using fluorescence microscopy and FRA was demonstrated. A small region (r ≈ 43 μm) of the electrode was studied which had a low density of DNA coverage, resulting in large changes in potential modulated fluorescence. The response of the fluorescence to the frequency of the potential perturbation was studied with FRA measurements characterized by a transfer function. This transfer function was corrected for the frequency dependent variation of the potential at the interface. This corrected transfer function showed little change over the frequency range investigated, only decreasing at the highest frequencies measured in contrast to other reports. The veracity of the correction method was tested by adding a resistor between the WE and the corresponding potentiostat lead artificially increasing the charging time constant. After applying the correction, the fluorescence response was shown to be the same as without the added resistor. Moreover, an increase in the electrolyte concentrationthough decreasing the fluorescence modulationalso resulted in a corrected fluorescence response that did not decrease significantly until above 10 kHz. In all cases studied, the fluorescence modulation after correction for the potential across the interface was the same in both the magnitude and phase of the response. The limiting factor in the high frequency measurement was found to be the optical system electronics which means that the rate at which DNA can reorient closely follows the potential that drives these reorientations and occurs on a fast time scale (>100 kHz). As these reorientations are used for bioanalytical determinations of DNA hybridization and sizing of proteins, the correction for the driving potential should result in more reliable and accurate estimates of the reorientation kinetics.
Figure 9. Correction applied to data published in ref 8 showing data for digoxigenin-labeled DNA in the absence and presence of sheep immunoglobulin G (IgG) antibody. The potential across the interface (continuous line) is calculated using the fit to an [RC] circuit with R = 1.6 kΩ and C = 9.6 nF reported. Uncorrected (filled symbols) and corrected (open symbols) FAC( f) values are shown, evidencing the increase in the frequency response shift upon IgG binding if the correction is used.
charging time)26 than with the results using modulation techniques.8,11,27 The phase of the transfer function was not used, as phase information was not available in the original publication. Upon addition of IgG antibody, a frequency shift is observed in the response slowing down the “switching” process (triangles). This is also reflected in the corrected curve which starts decreasing at ∼1 kHz. A comparison of the response before and after the IgG binding reveals that, while a discernible shift is observed even without the correction (filled symbols), this difference is significantly amplified after taking into account the actual potential at the electrode interface (open symbols). Moreover, comparison between different measurements, performed on different electrodes and electrochemical setups, will be facilitated without the need to worry about the electrochemical measurement conditions. Another approach was taken by Rant et al. to mitigate the τE problem.8 The experimental conditions are maintained constant to reduce variations in the electrochemical time constant for experiments to be compared. Additionally, to correct for changes to the electrochemical time constant induced by the binding process itself, an electrochemical impedance spectrum was measured before and after binding. After determining the corresponding electrochemical time constants (τE1 and τE2, respectively), the fluorescence cutoff frequency is multiplied by the correction factor τE2/τE1. Although this simple normalization method of the electrochemical time constants may help reduce its variations due to binding events, the analysis shown above makes it evident that those results are still strongly influenced by the electrochemistry of the cell, potentially leading to inaccurate and unreproducible results even in simple arrangements. The use of the method outlined above which decouples the electrochemical time constant results in data that is more representative of the hydrodynamic-limited “switching” regime. It must be noted, however, that large values of τE will be reflected in low amplitudes which will translate into large noise at the higher frequencies. Despite not being able to separate the upward and downward motion processes like the time-resolved measurements,26 the proposed correction method can help reposition frequency-resolved measurements in the analysis of “switching” DNA.
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ASSOCIATED CONTENT
S Supporting Information *
Details on the component values obtained from EIS fitting, comparison of impedance results at the two different potential amplitudes used, as well as the EIS data for the system calibration with a LED are provided. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The authors thank Brian Ditchburn (UBC, Chemistry, Glassblowing) for his efforts in creating the spectroelectrochemical cell used in this work and Dr. Jamie Noel for useful discussions. J.C.-M. was supported by the National Council of Science and Technology of Mexico (CONACYT) through the scholarship 207929. This research was funded by NSERC (Canada).
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