Incorporation of a DNAzyme into Au-coated nanocapillary array membranes with an internal standard for Pb(II) sensing

Abstract

A Pb(II)-specific DNAzyme has been successfully incorporated into Au-coated polycarbonate track-etched (PCTE) nanocapillary array membranes (NCAMs) by thiol–gold immobilization. Incorporation of the DNAzyme into the membrane provides a substrate-bound sensor using a novel internal control methodology for fluorescence-based detection of Pb(II). A non-cleavable substrate strand, identical to the cleavable DNAzyme substrate strand except the RNA-base is replaced by the corresponding DNA-base, is used for ratiometric comparison of intensities. The cleavable substrate strand is labeled with fluorescein, and the non-cleavable strand is labeled with a red fluorophore (Cy5 or Alexa 546) for detection after release from the membrane surface. This internal standard based ratiometric method allows for real-time monitoring of Pb(II)-induced cleavage, as well as standardizing variations in substrate size, solution detection volume, and monolayer density. The result is a Pb(II)-sensing structure that can be stored in a prepared state for 30 days, regenerated after reaction, and detect Pb(II) concentrations as low as 17 nM (3.5 ppb).

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Introduction

Toxic metal ions, such as Pb(II), are found in the environment and they have severe adverse health effects to human beings, especially to children.^1,2 Designing sensors may allow on-site, real-time detection and quantification of these metal ions. Toward this goal, a number of metal ion sensors have been reported.^2–10 Among these sensors, DNAzyme-based sensors are particularly attractive, because DNAzymes that are selective for any specific metal ion can be obtained through a combinatorial biology method called in vitro selection,^11,12 and these metal ion-specific DNAzymes can be converted into fluorescent biosensors by attaching fluorophores and quenchers to the DNAzymes.^8,13–15 For example, an in vitro selected Pb(II)-specific DNAzyme (Fig. 1(a))¹⁶ has been converted into a fluorescent sensor for Pb(II) with a solution detection range of ∼10 nM to 4 µM.⁷ However, several limitations have to be overcome in order to transform these DNAzyme sensors from laboratory experiment to field test. One major limitation is high background fluorescence due to dehybridization of the substrate from the enzyme strands and/or slow cleavage of the substrate during storage.¹⁷ To decrease the high background fluorescence from dehybridization, we have explored a dual-quencher method that improves the signal-to-noise ratio by 10-fold.¹⁷ To decrease the high background fluorescence from slow non-specific cleavage of the substrate, we used a method for immobilization of the Pb(II)-cleavable DNAzyme on planar Au surfaces.¹⁸ Immobilizing the DNAzyme on a solid surface allows rinsing of the surface to remove cleaved products, thus reducing background fluorescence, and resulting in an even wider linear dynamic range (1 nM < Pb(II) < 10 µM) than in solution.

Fig. 1 Schematic representation of immobilization protocol for DNAzyme on a Au surface. (a) Thiolated enzyme strand is tethered to the Au surface, (b) shorter chain mercaptohexanol displaces non-specific association of enzyme strand with the surface, (c) fluorophore-labeled substrate is hybridized, and (d) after Pb(II) reaction, the substrate is cleaved and fluorophore released from the surface into solution. The released fluorophore is then detected in solution.

Initial strategies for immobilization of DNAzyme on a solid support were based on minor modifications of previously reported methods for tethering oligonucleotides to Au.^19–22 Briefly, the enzyme strand (HS-(7)17E-Dy) of the DNAzyme complex is immobilized on a Au surface utilizing well-known thiol–Au chemistry. A short-chain alkanethiol, mercaptohexanol (MCH), is then used to displace non-thiol interaction of the DNA backbone with the Au surface, forcing the enzyme strand into an extended conformation necessary for maximum hybridization efficiency. The cleavable substrate strand (Dy-(7)17DS-Fl) is then hybridized to the immobilized enzyme strand forming the DNAzyme complex. Reaction with the target analyte, Pb(II), causes cleavage and release of a portion of the substrate strand that is labeled with a fluorophore which is then detected in solution. This approach, which is amenable to chemical sensing, avoids the complex and expensive instrumentation commonly used for direct Pb(II) detection.^23–35 The immobilization method and reaction, shown schematically in Fig. 1, also minimize the background arising from incomplete hybridization, seen in solution. In addition, solid-based sensors offer several further advantages including sensor regeneration and long-term storage that are not possible with solution-based approaches.

While immobilization of the DNAzyme for Pb(II) assays resolves many of the challenges associated with bulk solution detection methods,^7,17,36 it creates its own set of demands. First, variable monolayer packing densities can lead to variability in the fluorescence intensity for identical Pb(II) concentrations, making sample standardization based only on fluorophore intensity problematic. To eliminate this potentially large source of variability, a non-destructive in situ method to quantify the monolayer density is needed. Second, mass transport of Pb(II) to the functionalized surface yields a 10-fold increase in reaction time for the immobilized sensor compared to bulk solution assays. Increasing the surface-to-volume ratio of the structure presenting active DNAzyme should enable faster development of Pb(II)-induced fluorescence signal. Substrates in which DNAzymes are immobilized within nanoscale features represent one method to achieve this goal. In particular, the interior pores of Au-coated nanocapillary array membranes (NCAMs) can be utilized for this purpose, and previous work has shown that oligonucleotides tethered in these environments retain their activity.³⁷

This work puts forward two major advances in the use of surface-immobilized DNAzymes for chemical sensing. The first is the development of a novel ratiometric internal standard that allows both real-time monitoring of the Pb(II) cleavage reaction and simultaneous quantification of the total mass of immobilized DNAzyme for signal standardization. A second advance is incorporation of the DNAzyme into the nanometre scale pores of Au-coated NCAMs fashioned from polycarbonate track-etched membranes. The purpose of this immobilization scheme is to provide an increase in surface-to-volume ratio without an increase in geometric sample size. Furthermore, utilizing the Au-coated membranes as supports for DNAzyme immobilization is a natural complement to the use of NCAMs as switchable fluidic elements in multidimensional lab-on-a-chip schemes^38–40 where microfluidic separations are coupled to post-processing for the identification of biothreat agents or the removal of trace contaminants in water purification.^41–43 Finally, the use of these DNAzyme membrane NCAMs lend themselves to facile storage and regeneration.

Experimental

Reagents and oligonucleotides

All reagents were purchased from Sigma Aldrich. Buffer solutions were prepared with as-received reagents, chelated with Chelex 100 beads for 1 h to remove contaminating divalent metal ions, filtered, and titrated with glacial acetic acid to adjust the pH as desired. Concentrated Pb(II) solutions were prepared using Pb(OAc)₂ salt in 10 mM acetic acid to assist in solubility. Oligonucleotides with modification were purchased from Integrated DNA Technologies, Inc. with HPLC purification and were used without additional purification. Enzyme strand, HS-(7)17E-Dy, was labeled with 5′-thiol and 3′-Dabcyl (quencher) label. Cleavable substrate strand, Dy-(7)17DS-Fl, was labeled with 5′-fluorescein and 3′-Dabcyl. Non-cleavable internal control strand, (7)17DSnc-Cy5 or (7)17DSnc-Alexa546, was labeled with 5′-Cy5 or Alexa546 fluorophore, respectively. Oligonucleotide sequences and modification are shown in Table 1, while sequence and modification justification were reported previously.^17,18

Table 1 Names and sequences of modified oligonucleotides used

HS-(7)17E-Dy	5′-(C6Thiol)-TTTTTAAAGAGACATCTCTTCTCCGAGCCGGTCGAAATAGTGAGT-Dabcyl-3′
Dy-(7)17DS-Fl	5′-Fluorescein-ACTCACTATrAGGAAGAGATGTCTCTTT-Dabcyl-3′
(7)17DSnc-Cy5	5′-Cy5-ACTCACTATAGGAAGAGATGTCTCTTT-3′
(7)17DSnc-Alexa546	5′-Alexa546-ACTCACTATAGGAAGAGATGTCTCTTT-3′

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Evaporation of gold on glass slides

Planar Au surfaces were produced by vapor deposition onto glass microscope slides. Glass microscope slides were first cleaned in piranha solution (70% H₂SO₄, 30% H₂O₂) for 30 min followed by thorough rinsing with deionized (18 MΩ cm) water. (CAUTION: Piranha is a vigorous oxidant and should be used with extreme caution!) After cleaning, the glass slides were rinsed with isopropyl alcohol and dried with a dry N₂ stream. Prior to Au film deposition, Cr was deposited as an adhesion layer at a rate of 0.1 Å s⁻¹ to a final thickness of 50 Å. Au was then deposited at a rate of 1 Å s⁻¹ to a final thickness of 500 Å. Freshly prepared Au films were stored in a dry N₂ atmosphere until use. Immediately prior to DNAzyme immobilization, Au films were cleaned with piranha solution for 20 min and rinsed with deionized water for 5 min.

Electroless deposition of Au on NCAMs

Electroless deposition of Au on commercial polycarbonate track-etched NCAMs (Osmonics, Inc.), was performed by modifying previously reported methods.⁴⁴ Membranes with pore sizes of 10 nm and 650 nm (internal diameter, i.d.) were cleaned by soaking for 5 min in CH₃OH. Membranes were sensitized with Sn²⁺ by soaking for 45 min in 0.022 M SnCl₂ and 0.067 M trifluoroacetic acid in a 1 ∶ 1 mixture of water and methanol. After rinsing with water, the membranes were activated with Ag⁺ by immersing in an aqueous solution of 0.035 M AgNO₃ for 5 min and subsequently rinsed with water. The final solution used to deposit Au was 0.023 M NaHCO₃, 0.118 M Na₂SO₃, 0.68 M 37% formaldehyde in water, and 0.007 M Oromerse gold solution, part B. The membranes were soaked in this solution at 4 °C for 3 hours. The coated membranes were rinsed with water and cleaned in 25% HNO₃ overnight, then rinsed with water and dried with dry N_2(g). The Au-coated NCAMs were stored in a dry nitrogen atmosphere until use. Immediately prior to DNAzyme immobilization, the Au-coated NCAMs were cleaned by exposure to O_3(g) for 20 min.

Immobilization procedure for DNAzyme

Assembly of thiolated-DNA on Au and hybridization of complementary DNA followed previously reported methods^19,20,45 and is portrayed in Fig. 1. Immobilization of HS-(7)17E-Dy, or enzyme, on Au was achieved by soaking cleaned Au surfaces (ca. 0.5 × 0.5 cm² for Au–glass, ca. 0.3 × 0.3 cm² for Au–NCAM) in 1 M potassium phosphate buffer (pH = 6.9), 100 µM tris(2-chloroethyl) phosphate (TCEP), and 1 µM HS-(7)17E-Dy for 90 min. TCEP was added to disrupt the formation of disulfide bonds or other forms of oxidized thiols.⁴⁵ Surfaces were then thoroughly rinsed in deionized (DI) water and immediately soaked in 1 mM mercaptohexanol (MCH) for 5 min. Subsequently, surfaces were thoroughly rinsed in 50 mM tris acetate buffer (pH = 7.4) and 1 M NaCl. Hybridization of substrate and internal control was accomplished by soaking the immobilized enzyme and MCH mixed monolayer in 50 mM tris acetate buffer (pH = 7.4) and 1 M NaCl containing 1 µM Dy-(7)17DS-Fl and 125 nM (7)17DSnc-Alexa546 (or (7)17DSnc-Cy5) and heating in a 70 °C water bath for 60 min. The bath was then allowed to cool to room temperature over 60 min, cooled to 4 °C for 30 min, and again allowed to come to room temperature, resulting in a DNAzyme coated NCAM. It should be noted that early characterization was performed using Cy5 for the non-cleavable strand label due to its relative inexpensiveness when compared to the more time- and pH-stable Alexa label.

Prior to using the prepared DNAzyme coated NCAM for Pb(II) sensing, it was soaked in a 50 mM tris acetate buffer (pH = 7.7) and 50 mM NaCl solution for 5 min in an effort to remove any physisorbed substrate strands adsorbed in spite of MCH-passivation, and to rinse away any dissociated or partially dissociated substrate strands at the lower NaCl concentration. Measurements were made by placing the assembled DNAzyme in 600 µL of freshly prepared Pb(II)-containing 50 mM tris acetate buffer (pH = 7.7) and 50 mM NaCl. The DNAzyme surface was allowed to react with the Pb(II) solution, after which it was removed and rinsed with the reaction solution. Fluorescence intensity of the cleaved DNA in solution was determined using a 0.5 × 0.5 cm² quartz cell in a Jobin Yvon Fluoromax-P fluorimeter (λ_ex = 491 nm; λ_em = 518 nm, λ_ex = 555 nm; λ_em = 571 nm and λ_ex = 648 nm; λ_em = 665 nm) to measure fluorescein, Alexa546 and Cy5 fluorescence intensity, respectively.

Regeneration procedure

For determination of regeneration capability, the surface-immobilized DNAzyme was first prepared as described above. After an initial reaction of the sensor with 10 µM Pb(II), the activity was determined by fluorescence measurements. The reacted sensors were subsequently soaked individually in 18 MΩ cm DI water for 18 h in closed sample vials. Upon emersion the samples were rinsed with fresh 18 MΩ cm DI water for 5 min. Rehybridization of Dy-(7)17DS-Fl and internal control, (7)17DSnc-Alexa546, occurred by soaking the reacted sensors in 50 mM tris acetate buffer (pH = 7.4) and 1 M NaCl with 1 µM Dy-(7)17DS-Fl and 125 nM (7)17DSnc-Alexa546 with the same heating and cooling protocol described above. Non-regenerated control samples were soaked in identical buffer solutions and subjected to identical heating conditions, but without oligonucleotide. The controls and regenerated substrates were reacted with 600 µL of freshly prepared 10 µM Pb(II) in 50 mM tris acetate buffer (pH = 7.7) and 50 mM NaCl for 60 min. The reaction solution was rinsed over the surface of the sensor, and fluorescence intensity was determined as described above.

Dry storage method

Upon complete assembly of DNAzyme on Au-coated NCAMs, the NCAMs were dried using a gentle N_2(g) stream, and placed in a capped vial for storage in a dark drawer. Prior to use, the DNAzyme tab was soaked in 50 mM tris acetate buffer (pH = 7.7) and 50 mM NaCl for 5 min to rehydrate the sensor followed by transfer to the reaction solution. The NCAMs were then soaked in 10 µM Pb(II) in 50 mM tris acetate buffer (pH = 7.7) and 50 mM NaCl for 30 min, and the fluorescence was measured.

Results and discussion

Surface area and internal standardization

To determine relative surface area available for DNAzyme tethering, SEM images of evaporated planar gold and electrolessly deposited 10 nm and 650 nm membranes (internal diameters prior to coating with Au) were obtained. The insets in Fig. 2(a) reveal a relatively smooth surface for the planar gold, in contrast to the much greater surface roughness of the electrolessly deposited samples. The effective pore size for the 10 nm membrane after coating with Au is 0 nm, because electroless deposition of gold at the experimental pH completely blocks the pores, rendering them inaccessible to fluid. In contrast, the 650 nm pore diameter membranes allow access of the DNAzyme to the interior of the pores as well as the top and bottom surfaces of the membrane. Greater surface area for DNAzyme assembly translates into greater fluorescence intensity after reaction with Pb(II), as seen in Fig. 2(a). Electrolessly deposited gold (0 nm effective i.d.), has a fluorescence intensity that is 4.38 ± 1.45 times higher than the planar gold surface due to the enhanced surface area available for DNAzyme binding. In addition, the 500 nm pores (measured i.d. after Au deposition) show a 2.8 ± 0.7 times increase in intensity compared to the blocked pore membrane. This increase is due to accessible Au-coated surface in the walls of the pores and confirms DNAzyme immobilization within the pores. However, this difference is only 54% ± 14% of that calculated for full coverage within the pores, indicating incomplete pore coverage.

Fig. 2 (a) Fluorescence intensities obtained from samples of identical geometric area for each type of surface with inset scanning electron micrographs of each surface. Inset scale bars are 1 µm in length. (b) A cross-sectional SEM image of the porous membrane reveals incomplete gold coating within pores of the polycarbonate.

Full coverage within the pores is estimated by scaling fluorescent intensity from blocked pore NCAMs with the increase in surface area contributed by pore walls in the 500 nm effective diameter, (3 × 10⁷ pores cm⁻²; 10 µm thick), NCAMs. The assumption made for this calculation is that the DNAzyme coverage density within the pores should be the same as on the face of the NCAM. This assumption is supported by two observations. (1) The oligonucleotides do not form a true monolayer, but instead are part of a mixed monolayer with MCH, resulting in approximately an adlayer with ca. 1% the maximum density of close-packed oligonucleotides. That means the DNAzymes are not close-packed and, thus, should not be affected by the curvature of the pore. (2) Pore diameters of 500 nm are large relative to the extended length of the DNAzyme (∼7 nm), and with an average spacing of 10 nm between each DNAzyme duplex, it is unlikely that steric crowding decreases the oligonucleotide density observed on the face. The lower coverage within the pores is then attributed to incomplete coating of the pore wall by Au, not by insufficient access of the DNAzyme to the pores, as supported by the SEM cross-section image shown in Fig. 2(b). Nevertheless, these data furnish proof that the amount of surface immobilized DNAzyme increases when moving from evaporated Au to electrolessly-deposited Au on essentially planar surfaces to electrolessly deposited Au in the interior pores of NCAMs.

In the surface-immobilized DNAzyme-mediated assay, the Pb(II) concentration is related to the intensity of released fluorophore-bearing strand, which in turn depends on the amount of accessible DNAzyme. Thus, variations in DNAzyme surface coverage from sample-to-sample suggest that a method for standardizing fluorescence intensity between sample generations is needed. Previous work on immobilized DNAzymes for Pb(II) detection was based solely on single-wavelength fluorescence intensity, in combination with a separate simultaneously monitored control sample for comparison.¹⁸ To simplify the assay and render it more robust to all factors affecting sample-to-sample reproducibility, a ratiometric approach utilizing an internal standard was developed. Use of an internal standard enables DNAzyme surface loadings to be determined as well as allowing real-time analyte detection without the need for a parallel control sample. Standardization was accomplished by incorporating an internal control strand, identical to the cleavable substrate strand, but rendered uncleavable by replacement of the RNA base with a DNA base. The non-cleavable strand is labeled with a different fluorophore from that used on the cleavable strand, providing a distinguishable parallel tag by which the two fluorescent signals may be compared (see Fig. 3).

Fig. 3 Schematic representation of the use of an internal standard to implement ratiometric monitoring of nonspecific dehybridization. (a) Ratio of release of both cleavable (dark circles) and uncleavable (light circles) substrate is constant in the absence of Pb(II) resulting in a constant fluorescein-to-Cy5 fluorescence intensity ratio. (b) Cleaved fragment release in the presence of Pb(II) increases the release of signal from the cleavable strand causing an increase in the ratio.

Even in the absence of analyte, nonspecific dissociation (or dehybridization) of the two strands generates a background signal, which ultimately limits the smallest detectable concentration of analyte. Introduction of analyte then produces an additional increment to the fluorophore signal due to cleavable substrate. However, when the internal standard is used, the presence of Pb(II) does not alter the amount of fluorescence arising from the uncleavable strand, since it can only be released from the surface by nonspecific dehybridization. Because the fraction of strands undergoing nonspecific dehybridization is a constant characteristic of the particular DNAzyme used, the fluorescence intensity of the labeled uncleavable strand is proportional to the DNAzyme surface coverage. Thus, the ratio of fluorescence intensities arising from cleavable and uncleavable strands is independent of sample size or monolayer density, thereby standardizing individual DNAzyme NCAMs.

For each system investigated, the fluorescence intensity arising from the fluorophore on cleavable substrate, Dy-(7)17DS-Fl, was ratioed to that arising from the fluorophore on uncleavable substrate, (7)17DSnc-Cy5, and this ratio was monitored over time. In the absence of Pb(II) this ratio remained constant after an initial equilibration time, as seen in Fig. 4. However, in the presence of Pb(II), the fluorescence intensity arising from the cleavable strand increases due to the specific cleavage of the substrate strand from the DNAzyme complex. In the example shown (100 nM Pb(II)), the ratio between Dy-(7)17DS-Fl and (7)17DSnc-Cy5 fluorescence intensities increases ∼2-fold, which is easily discernible from the original baseline ratio. The raw fluorescence intensity from the uncleavable strand fluorophore quantitates the amount of DNAzyme complex on the NCAMs, while the change in ratio of the two intensities provides real-time analysis of Pb(II)-induced cleavage. Although this ratiometric approach works well over reasonable sampling periods, at extremely long times (>20 h), an artificial increase in background intensity ratio is observed. This increase occurs because the internal standard strand, which is present at lower surface concentration than the cleavable DNAzyme, is completely removed. Thus, long-time operation clearly demands a robust method for sample regeneration.

Fig. 4 Time dependence of the fluorescein-to-Cy5 fluorescence ratio in the internal standard method for background (Pb(II)-absent, open circles) and analyte-containing samples (100 nM Pb(II), filled circles).

To further illustrate the capabilities of the internal standard method, replicate experiments were performed on NCAMs with areas of 6.25 mm² and 22.5 mm², a difference in area of 3.6 times, to determine the ability to standardize measured Pb(II) concentrations. NCAMs were placed in 10 µM Pb(II) solution for 30 min and fluorescence intensities recorded. The large tab showed a fluorescein intensity that was 3.50 ± 0.50 times larger than that of the smaller tab, as expected. However, the ratio of fluorescence intensities, (I_Fl/I_Cy5)_{large tab}/(I_Fl/I_Cy5)_{small tab} was 1.21 ± 0.36, i.e. unity within experimental error, thereby illustrating the ability of the internal standard to correct for differences in substrate area and DNAzyme loading in Pb(II) determinations.

LOD and dynamic range

The tab-based method for introduction of DNAzyme into analyte solution is only useful if a detection limit below the EPA threshold of 75 nM Pb(II) for water is maintained.⁴⁶ A Pb(II) calibration curve was constructed for reacted NCAMs, with ratio sampling at one hour to allow sufficient time to distinguish low Pb(II) concentrations from background, viz.Fig. 5, and a limit of detection of 17 nM was obtained with the tab based system. Operation at concentrations near the detection limit required long sampling times, so the DNAzyme NCAMs were exhausted in this time frame at relatively modest concentrations, [Pb(II)] ∼500 nM, thereby decreasing the dynamic range of the sensor. However, it was possible to differentiate among higher concentrations of Pb(II) by decreasing the reaction time, thereby avoiding total exhaustion of the DNAzyme on the tab, and providing a tunable detection range and detection limit. The reaction rate is faster for DNAzyme NCAMs compared to DNAzyme immobilized on planar Au, as demonstrated by the fact that on comparable substrate areas 500 nM Pb(II) exhausts DNAzyme bound to NCAMs in 1 h, while 10 µM Pb(II) is required to exhaust the DNAzyme bound to planar Au in 1 hr.¹⁸ Since there is little reason to believe the intrinsic cleavage reaction kinetics are different on the two surfaces, the differences in gross rate likely reflect the more efficient mass transport possible within the nanopores.

Fig. 5 Calibration curve for Pb(II) reaction on DNAzyme NCAMs for one hour reaction time showing a detection limit of 17 nM Pb(II). The detection limit can be tuned to differentiate higher Pb(II) concentrations by decreasing the reaction time.

Regeneration and storage

To optimally exploit this method of DNAzyme-based analyte detection in a variety of sensing applications, it will be necessary to store prepared NCAMs until they are ready for use and to regenerate the surface architectures after each use. To test the ability to regenerate the active surface-bound DNAzyme across multiple generations, sequential experiments were performed in which DNAzyme NCAMs were reacted with 10 µM Pb(II) for 1 h to ensure complete reaction of the tab for analysis of total released fluorophore, fluorescence was measured, reacted NCAMs were rinsed, and then rehybridized with a mixture of cleavable and uncleavable substrate. The intensity from Pb(II) reaction with DNAzyme on the surface in successive generations, shown in Fig. 6(a), remains unchanged over four regenerations. Total enzyme loss is negligible, and the overall activity depends more on the hybridization ratio of cleavable and uncleavable strands for a given regeneration than on the regeneration number, cf.Fig. 6(b), as evidenced by constant fluorescein intensity, with the only differences arising from variations in the intensity of the internal standard strand. The observed decrease in ratio is attributed to incomplete rinsing away of internal standard due to the stability of the duplex in the uncleaved state. This gradually lowers the ratio of cleavable to uncleavable substrate and would likely be corrected by more stringent rinsing conditions (e.g. heating, urea, etc.) It should also be noted that the ratiometric determinations reduced the magnitude of the deviation (error bars), while increasing the difference between Pb(II)-absent and present values compared to fluorescein intensity measurements.

Fig. 6 Multi-regeneration study of DNAzyme NCAMs. (A) Total fluorescence intensity, and (B) ratiometric analysis using an internal standard. In both panels gray bars represent the background (no Pb(II)) signal, and black bars represent signal in the presence of 10 µM Pb(II) after 1 h reaction time.

To study the retention of sensing activity over long periods of dry storage a sheet of immobilized DNAzyme was prepared and stored, and NCAMs were removed and sampled at various time intervals. Drying of the NCAMs did not reduce the activity of the DNAzyme, and in fact, increased the ratio of cleavable (Fl) to uncleavable (Alexa) from 28.5 ± 3.00 to 119 ± 15.9, (data not shown), by reducing the surface loading of the internal standard. During storage the fluorescein/Alexa fluorescence intensity ratio decayed exponentially over a span of 30 days of dry storage, viz.Fig. 7. Despite the decay in signal intensity ratio, activity is maintained with 10 µM Pb(II) producing a six-fold increase in intensity over background even after 30 days. This is a notable advantage made by moving from solution-based to immobilized DNAzyme sensors, yielding a sensor that may be prepared ahead of time and ready to use with minimal additional preparation.

Fig. 7 Fluorescence intensity ratio of fluorescein and Alexa as a function of storage time; (filled circles) 10 µM Pb(II), (open circles) no Pb(II) present.

Conclusion

The DNAzyme-immobilized Au-coated nanocapillary array membranes, or DNAzyme NCAMs, are a powerfully general and robust platform for DNAzyme-based sensing of environmental agents, such as Pb(II). Use of these structures as sensor platforms does not require lengthy hybridization procedures, handling of DNA-containing solutions, or volumetric measurements, because the assembled DNAzyme NCAM may be prepared ahead of time and stored. In addition, this approach provides a very simple method for the introduction of analyte-sensing DNAzyme into solutions containing the target analyte. This NCAM-based method maintains a low detection limit of 17 nM, allows for long-term storage of the prepared DNAzyme sensor and the DNAzyme molecular recognition motifs may be regenerated over several cycles. The ratiometric method of signal generation overcomes a key practical problem, sample-to-sample variability due to background variation, by constructing the DNAzyme NCAMs using an internal standard strand, which effectively standardizes fluorescent signals by correcting for the major source of background: nonspecific dehybridization. This enables real-time detection of Pb(II)-induced cleavage.

Finally, the combination of immobilizing the DNAzyme on a Au-coated NCAM support and the internal standard method of background correction are well-poised for extension to more sophisticated sensor and micrototal analytical system applications in which the sensor-bearing NCAMs would be incorporated directly into nanofluidic–microfluidic hybrid architectures for multidimensional chemical analysis. Admittedly, these more complicated analytical systems may necessitate more involved calibration and mathematical signal processing strategies, due to the dependence of the signal on the length of time the DNAzyme–NCAM has been stored and other environmental factors. In a straightforward approach, a multichannel microfluidic device would use one channel for calibration and a second, measurement, channel to quantify Pb(II) concentrations. The major challenge that remains, then, is to incorporate these NCAMs into hybrid microfluidic systems, a very viable goal given advances in portable fluorimeters, microfluidic devices, and integrated circuits.

Acknowledgements

This work was supported by the National Science Foundation Science and Technology Center for Advanced Materials for Water Purification and by the Strategic Environmental Research and Development Program.

References

1. H. L. Needleman, Human Lead Exposure, CRC Press, Boca Raton, 1992.
2. H. A. Godwin, Curr. Opin. Chem. Biol., 2001, 5, 223–227.
3. R. Y. Tsien, in Fluorescent Chemosensors for Ion and Molecule Recognition, ed. A. W. Czarnik, American Chemical Society, Washington, DC, 1993, pp. 130–146.
4. A. W. Czarnik, Chem. Biol., 1995, 2, 423–428.
5. S. C. Burdette, G. K. Walkup, B. Spingler, R. Y. Tsien and S. J. Lippard, J. Am. Chem. Soc., 2001, 123, 7831–7841.
6. S. Deo and H. A. Godwin, J. Am. Chem. Soc., 2000, 122, 174–175.
7. J. Li and Y. Lu, J. Am. Chem. Soc., 2000, 122, 10466–10467.
8. Y. Lu, Chem.—Eur. J., 2002, 8, 4588–4596.
9. C. J. Fahrni and T. V. O'Halloran, J. Am. Chem. Soc., 1999, 121, 11448–11458.
10. M. M. Henary, Y. Wu and C. J. Fahrni, Chem.—Eur. J., 2004, 10, 3015–3025.
11. R. R. Breaker and G. F. Joyce, Chem. Biol., 1994, 1, 223–229.
12. Y. Li and R. R. Breaker, Curr. Opin. Struct. Biol., 1999, 9, 315–323.
13. G. M. Emilsson and R. R. Breaker, Cell. Mol. Life Sci., 2002, 59, 596–607.
14. R. Nutiu, S. Mei, Z. Liu and Y. Li, Pure Appl. Chem., 2004, 76, 1547–1561.
15. J. C. Achenbach, R. Nutiu and Y. Li, Anal. Chim. Acta, 2005, 534, 41–51.
16. J. Li, W. Zheng, A. H. Kwon and Y. Lu, Nucleic Acids Res., 2000, 28, 481–488.
17. J. Liu and Y. Lu, Anal. Chem., 2003, 75, 6666–6672.
18. C. B. Swearingen, D. P. Wernette, D. M. Cropek, Y. Lu, J. V. Sweedler and P. W. Bohn, Anal. Chem., 2005, 77, 442–448.
19. R. Levicky, T. M. Herne, M. J. Tarlov and S. K. Satija, J. Am. Chem. Soc., 1998, 120, 9787–9792.
20. T. M. Herne and M. J. Tarlov, J. Am. Chem. Soc., 1997, 119, 8916–8920.
21. A. B. Steel, T. M. Herne and M. J. Tarlov, Anal. Chem., 1998, 70, 4670–4677.
22. A. B. Steel, R. L. Levicky, T. M. Herne and M. J. Tarlov, Biophys. J., 2000, 79, 975–981.
23. R. Ma, W. V. Mol and F. Adams, Anal. Chim. Acta, 1994, 285, 33–43.
24. J. Liu, H. Chen, X. Mao and X. Jin, Int. J. Environ. Anal. Chem., 2000, 76, 267–282.
25. M. Ochsenkuhn-Petropoulou and K. M. Ochsenkuhn, Fresenius’ J. Anal. Chem., 2001, 369, 629–632.
26. H. Elfering, J. T. Andersson and K. G. Poll, Analyst, 1998, 123, 669–674.
27. Z. Fang, M. Sperling and B. Welz, J. Anal. At. Spectrom., 1991, 6, 301–306.
28. K. Crowley and J. Cassidy, Electroanalysis, 2002, 14, 1077–1082.
29. M. R. Ganjali, A. Rouhollahi, A. R. Mardan, M. Hamzeloo, A. Mogimi and M. Shamsipur, Microchem. J., 1998, 60, 122–133.
30. T. Sokalski, A. Ceresa, T. Zwickl and E. Pretsch, J. Am. Chem. Soc., 1997, 119, 11347–11348.
31. A. Sussell and K. Ashley, J. Environ. Monit., 2002, 4, 156–161.
32. N. A. Yusof and M. Ahmad, Talanta, 2002, 58, 459–466.
33. D. A. Blake, R. C. Blake II, M. Khosraviani and A. R. Pavlov, Anal. Chim. Acta, 1998, 376, 13–19.
34. R. Wilson, D. J. Schiffrin, B. J. Luff and J. S. Wilkinson, Sens. Actuators, B: Chem., 2000, B63, 115–121.
35. C. E. Reese and S. A. Asher, Anal. Chem., 2003, 75, 3915–3918.
36. J. W. Liu and Y. Lu, J. Fluoresc., 2004, 14, 343–354.
37. P. Kohli, C. C. Harrell, Z. Cao, R. Gasparac, W. Tan and C. R. Martin, Science, 2004, 305, 984–986.
38. T. C. Kuo, D. M. Cannon, M. A. Shannon, P. W. Bohn and J. V. Sweedler, Sens. Actuators, A: Phys., 2003, 102, 223–233.
39. T. C. Kuo, D. M. Cannon, Y. N. Chen, J. J. Tulock, M. A. Shannon, J. V. Sweedler and P. W. Bohn, Anal. Chem., 2003, 75, 1861–1867.
40. H. C. Yang, P. F. Kuo, T. Y. Lin, Y. F. Chen, K. H. Chen, L. C. Chen and J. I. Chyi, Appl. Phys. Lett., 2000, 76, 3712–3714.
41. K. Kosutic, L. Furac, L. Sipos and B. Kunst, Sep. Purif. Technol., 2005, 42, 137–144.
42. P. Horsch, A. Gorenflo, C. Fuder, A. Deleage and F. H. Frimmel, Desalination, 2005, 172, 41–52.
43. N. Hilal, G. Busca, N. Hankins and A. W. Mohammad, Desalination, 2004, 167, 227–238.
44. S. F. Yu, N. C. Li, J. Wharton and C. R. Martin, Nano Lett., 2003, 3, 815–818.
45. T. Aqua, R. Naaman and S. S. Daube, Langmuir, 2003, 19, 10573–10580.
46. http://www.epa.gov/safewater, 2005.

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