Alma D.
Castañeda
,
Donald A.
Robinson
,
Keith J.
Stevenson‡
and
Richard M.
Crooks
*
Department of Chemistry, Center for Electrochemistry, and the Center for Nano- and Molecular Science and Technology, The University of Texas at Austin, 105 E. 24th St., Stop A5300, Austin, TX 78712-1224, USA. E-mail: crooks@cm.utexas.edu; Tel: +1-512-475-8674
First published on 1st July 2016
We report a new and general approach that will be useful for adapting the method of electrocatalytic amplification (ECA) to biosensing applications. In ECA, individual collisions of catalytic nanoparticles with a noncatalytic electrode surface lead to bursts of current. In the work described here, the current arises from catalytic electrooxidation of N2H4 at the surface of platinum nanoparticles (PtNPs). The problem with using ECA for biosensing applications heretofore, is that it is necessary to immobilize a receptor, such as DNA (as in the case here) or an antibody on the PtNP surface. This inactivates the colliding NP, however, and leads to very small collision signatures. In the present article, we show that single-stranded DNA (ssDNA) present on the PtNP surface can be detected by selectively removing a fraction of the ssDNA using the enzyme Exonuclease I (Exo I). About half of the current associated with collisions of naked PtNPs can be recovered from fully passivated PtNPs after exposure to Exo I. Experiments carried out using both Au and Hg ultramicroelectrodes reveal some mechanistic aspects of the collision process before and after treatment of the ssDNA-modified PtNPs with Exo I.
We are interested in using ECA to detect small numbers of biological molecules such as DNA. Previous work from our lab has shown, however, that when DNA is present on colliding NPs, very little current results.2 This is, of course, a consequence of DNA-induced blocking of catalytically active sites on the NP surface. This finding presents a clear problem for integration of ECA into biosensing schemes: specifically, if a biological recognition element is immobilized on the NPs, then little or no current will be observed upon impact. Here we present a strategy that at least partially addresses this difficulty. Specifically, we start with catalytically inactive DNA-modified NPs, and then detect the presence of DNA using an enzyme (an exonuclease) that degrades DNA sufficiently to reactivate the catalytic properties of the NPs. Importantly, the goal of the present work is to introduce a general methodology that could be useful for a range of future biosensing applications. At this early stage in our research, we do not intend to suggest that the metrics presented here are competitive with the many other DNA sensing methods that have been reported.
There are a number of different experiments that fall into the general category of detection of collisions between particles and an electrode. The first of these3–5 was reported by Scholz and coworkers who began a study of collisions between liposomes and Hg electrodes in 2002. Two years later Lemay and coworkers observed collisions between individual micron-scale, carboxylated latex spheres and an electrode surface arising from partial masking of the electroactive area of the electrode by the insulating spheres.6 Compton and coworkers used a different approach, in which current pulses resulted from oxidation of AgNPs when they struck an electrode surface.7,8 As mentioned earlier, ECA occurs when a colliding NP initiates a catalytic reaction. Two forms of this collision method have been described. In the first reports of ECA, the NPs were tethered to an electrode surface, but this did not result in observation of individual collisions.9,10 In 2007, however, Xiao and Bard showed that when the catalytic NPs were free in solution, individual ECA collisions could be discerned.11 As discussed next, the experiment reported here is most similar to the latter approach. Finally, we note that the research groups of Unwin,12,13 Zhang,14,15 Koper,12,16 Andreescu,17 Alpuche-Aviles,18 Crooks,2,19–21 Stevenson,22–24 and MacPherson25 have all made important contributions to the general field of single-particle collision electrochemistry that have influenced the findings reported here.
For the present study, we chose N2H4 oxidation (eqn (1)) as the redox indicator reaction, because this inner-sphere electron-transfer reaction is catalyzed by PtNPs but not by Au or Hg ultramicroelectrodes (UMEs).
N2H4 = N2 + 4H+ +4e− | (1) |
Accordingly, when an appropriate interfacial potential is selected for the UME, no current due to eqn (1) is observed until a PtNP strikes the electrode surface. The current transients that result from these collisions are step-shaped for Au UMEs26 and spike-shaped for Hg UMEs.22–24 The rapid current decrease observed for Hg UMEs is due to deactivation of the PtNPs resulting from Hg poisoning.
Enzymes have heretofore not been integrated into ECA sensing schemes, but we were inspired by the surface-enzyme chemistry reported by Corn and coworkers27,28 over the past several years, and we thought coupling the two methods could be quite powerful. This first report focuses on the use of nucleases, which are a family of enzymes capable of hydrolyzing the phosphodiester bonds in DNA chains.29 The difference between endo- and exonucleases lies in the way each class initiates hydrolysis. Endonucleases cleave phosphodiester bonds at a specific site within the middle section of the oligonucleotide, while exonucleases initiate cleavage at a free –OH group on either the 5′ or 3′ end.30 The 5′ to 3′ exonucleases are commonly used in biology to remove RNA primers,31,32 whereas 3′ to 5′ exonucleases are used to help repair DNA mismatches.33 A number of analytical assays incorporate the use of nucleases for operating on nanoparticles conjugated with DNA. These include schemes involving detection via colorimetry,34,35 fluorescence,36 and electrochemistry.37 For this study, we used Exonuclease I (Exo I), a nuclease extracted from E. coli, which is selective for denatured or single-strand DNA (ssDNA). Exo I initiates cleavage at a free 3′-hydroxyl end of ssDNA and cuts nucleotides in a stepwise fashion.38
In the present work, we modified 22 nm-diameter PtNPs with 25-mer ssDNA. Consistent with an earlier report from our group,2 these PtNP@ssDNA conjugates yield few collision signals (Scheme 1b), because DNA restricts access of N2H4 to the electrocatalytic PtNP surface. After incubation of PtNP@ssDNA with 30 U of Exo I, however, about 50% of the collision activity (compared to naked PtNPs) returns (Scheme 1c). This is because Exo I removes much of the ssDNA originally present on the PtNPs. The key result is that these findings point to a general approach for using ECA to detect small molecules, proteins, and DNA.
A 1.0 mL aliquot of the PtNP seed solution was added to 29.0 mL of H2O at room temperature. With stirring, 0.023 mL of a 0.40 M H2PtCl6 solution and 0.50 mL of a solution containing 1% sodium citrate and 1.25% L-ascorbic acid was added. The solution was then heated to boiling at the rate of 10 °C min−1. The total reaction time was 30 min. After cooling to room temperature, the solution was transferred to a 35 mm dialysis sack (12000 Da MWCO, Sigma-Aldrich) and submerged in 4 L of DI H2O for 24 h to remove excess salts. The PtNPs were characterized by transmission electron microscopy (TEM, FEI Tecnai TEM), and found to have an average diameter of 22 ± 4 nm. A representative TEM image and a histogram showing the NP size distribution are provided in the ESI (Fig. S1†).
These materials were used for all experiments reported herein, except for those relating to Fig. 2 where lower ratios of PtNP:ssDNA were used to gauge the effect of ssDNA surface concentration on ECA measurements. In those experiments the PtNP@ssDNA were resuspended in 50 mM PB (pH 7) rather than Taq buffer.
Hg UMEs were prepared by electrodepositing Hg onto a Pt UME according to a previously reported method.42 Briefly, the Pt UMEs were polished via wet sanding for 1 min. Hg was then electrodeposited (−0.10 V vs. Ag/AgCl, 3.4 M KCl) for 300 s from a solution containing 5.7 mM Hg2(NO3)2, 0.5% conc. HNO3, and 1 M KNO3. Finally, the Hg UMEs were rinsed with H2O immediately before use.
To estimate the average number of ssDNAs per PtNP, a previously reported technique, which is based on fluorescence quenching by the PtNPs, was used.40,43 For this purpose, the thiolated ssDNA was modified at the 3′ end with the fluorescent dye Cy3 (ssDNA–Cy3), which has a maximum absorption wavelength at 550 nm and a maximum emission wavelength at 570 nm. Adsorption of ssDNA–Cy3 to the PtNP brings the dye sufficiently close to the NP surface to quench its fluorescence. Therefore, after modification of the PtNPs with Cy3-tagged ssDNA, residual fluorescence in solution will arise primarily from unbound ssDNA.
To carry out this analysis we first recorded a calibration curve for ssDNA–Cy3 in Taq buffer (Fig. S2, ESI†), and then measured the fluorescence of solutions containing PtNP@ssDNA–Cy3. By comparison of the residual fluorescence in the latter solution to the calibration curve, the concentration of bound DNA can be determined. Dividing this value by the concentration of PtNPs (determined by NTA) yields a rough estimate of the number of ssDNA–Cy3 per PtNP, which we find to be ∼35. The conditions used to modify PtNPs with unlabeled DNA (e.g., PtNP@ssDNA) were the same as for PtNP@ssDNA–Cy3 to ensure similar coverages.
The poor colloidal stability of nanoparticles in ECA experiments is a very serious problem,16,44,45 and therefore we evaluated this parameter for the PtNP conjugates used in this study prior to carrying out collision experiments. These NTA results (Fig. 1a) indicate a size distribution of 46 ± 27 nm for nominally naked PtNPs (black trace) in 50 mM PB. This value can be compared to the size measured in water (no buffer): 28 ± 20 nm, indicating that the electrostatic shield effect of the buffer leads to some aggregation.46,47 After modification with ssDNA (PtNP@ssDNA), the size of the resulting conjugates in 50 mM PB is a little larger than the nominally naked PtNPs: 57 ± 31 nm (red trace) and 53 ± 29 nm (blue trace) before and after treatment with Exo I, respectively. Note that the aforementioned size distributions are the averages of three independent measurements.
It has previously been shown that the presence of N2H4 can increase the degree of aggregation of PtNPs,16 and therefore we repeated the measurements shown in Fig. 1a except included 10.0 mM N2H4 in the solutions (Fig. 1b). The size distribution for naked PtNPs (black trace) in the presence of N2H4 confirms the earlier report. In this case the average PtNP diameter is 100 ± 45 nm. After addition of the ssDNA shell, however, the conjugates are stabilized, and the degree of N2H4-induced aggregation is reduced both before (PtNP@ssDNA, 52 ± 31 nm, red trace) and after (PtNP@ssDNA, 55 ± 35 nm, blue trace) exposure to Exo I. In other words, N2H4 has no significant effect on the colloidal stability of PtNP@ssDNA.
Fig. 2a shows representative current–time (i–t) traces for N2H4 + 50 mM PB solutions in the absence of PtNPs (black trace), and in the presence of naked PtNPs (red trace), PtNP@ssDNA prepared using a PtNP:ssDNA ratio of 1:10 (blue trace), and PtNP@ssDNA prepared using a PtNP:ssDNA ratio of 1:100 (green trace). Obviously, no collisions are observed in the absence of PtNPs. Naked PtNPs produced an average ECA current transient frequency of 0.048 ± 0.006 Hz and current magnitude of 51 ± 41 pA (see Fig. S3† for additional i–t curves for collisions of nominally naked PtNPs with Au UMEs). This current is lower than that predicted by eqn (2) (237 pA), which has previously been shown to yield reasonable agreement with experimental measurements involving smaller NPs (∼3–4 nm).1
i = 4π(ln2)nFDCr | (2) |
The average ECA current and transient frequency resulting from collisions of PtNP@ssDNA (1:10), 51 ± 48 pA and 0.043 ± 0.006 Hz, respectively, were almost identical to those of the naked PtNPs. However, PtNPs with the highest ssDNA modification ratio revealed a much larger decrease in signal. Specifically, the average collision current for PtNP@ssDNA (1:100) was 17 ± 11 pA, a ∼66% decrease relative to naked PtNPs. Additionally, the frequency decreased from 0.048 ± 0.006 Hz to 0.0072 ± 0.004 Hz.
Fig. 2b is a histogram showing the distribution of ECA currents as a function of the PtNP:ssDNA solution ratio used to prepare the conjugates. These data reveal the quantitative attenuation in collision current as a function of increasing ssDNA coverage. Although not shown here, the collision frequency also decreases with increasing ssDNA coverage. This trend is consistent with data previously reported for 4.6 nm PtNPs.2
Fig. 1c presents normalized NTA data for both naked PtNPs (black trace) and PtNP@ssDNA (red trace). These data were obtained by resuspending the NPs in Taq buffer and incubating at room temperature for 1 h. An aliquot of the NP solution was then further diluted in Taq buffer to an appropriate concentration for NTA measurements (∼0.5–1.0 pM). The important result is that the average size of naked PtNPs in the Taq buffer is 155 ± 119 nm compared to 44 ± 40 nm for those stabilized with ssDNA. The latter size can be compared to 57 ± 31 found for PtNP@ssDNA in 50 mM PB (Fig. 1a). We conclude that PtNP@ssDNA are sufficiently stable in the Taq buffer to carry out the Exo I cleavage. NTA plots for the individual components of Taq buffer (75 mM Tris–HCl (pH 8.8), 2.0 mM MgCl2, 0.01% Tween 20, and 20 mM (NH4)2SO4) can be found in Fig. S4.†
A histogram showing the frequency of collisions as a function of the ECA current is shown in Fig. 3c. The average collision current for the post-Exo I PtNP–ssDNA is 24 ± 15 pA, which can be compared to the value for naked PtNPs: 51 + 41 pA. Because only about half of the original current is recovered after exposure to the enzyme, we conclude that only about that same fraction of the PtNP surface is available for N2H4 oxidation. The most likely scenario is that a substantial fraction of the ssDNA is reduced in length by Exo I, thereby provide access of N2H4 to the PtNP surface. We wish to emphasize that the ssDNA incorporates a 6-carbon alkylthiol on its 5′ (proximal) end. These alkyl chains will not be removed by Exo I, so it is unlikely that additional catalytic sites are exposed on the Pt surface. Rather, removal of some of bases reduces mass transfer hindrance of N2H4.
Exo I will only initiate cleavage in ssDNA having a free 3′-hydroxyl end. To confirm that the action of Exo I is responsible for the observation of collisions, we immobilized ssDNA on the PtNPs such that only the 5′-hydroxyl end was accessible. For this control experiment, only the orientation of the DNA on the PtNPs was reversed: the ssDNA sequence was unchanged. In this case, no collisions were observed after treatment with Exo I (Fig. S5†). This important finding confirms that the specific enzymatic activity of Exo I for cleavage of 3′-hydroxyl DNA is responsible for the collision currents shown in Fig. 3.
In addition to the collision current, we also studied the collision frequency for PtNP@ssDNA post-Exo I and found that it correlates linearly with concentration in the range of 0.58–11.7 pM (Fig. 3d). This observation is consistent with previous findings for collisions of naked PtNPs at Au UMEs.20 Here, we also found that the collision frequency is higher for PtNP@ssDNA post-Exo I than for naked PtNPs. For example, at a concentration of 11.7 pM, the frequency for PtNP@ssDNA post-Exo I was 0.190 ± 0.033 Hz, compared to 0.048 ± 0.006 Hz for naked PtNPs. We have observed this same trend previously for ssDNA-modified PtNPs colliding with a Au microband electrode.2
The factor-of-three difference in collision frequency may be caused by the difference in the rates of mass transport between the slower diffusing aggregates of naked PtNPs and the smaller, more colloidally stable PtNP@ssDNA. Another observation worth considering is the high noise level observed for collisions arising from PtNP@ssDNA post-Exo I (Fig. 3b). This might suggest that the ssDNA-modified NPs reside on or near the UME surface longer than naked PtNPs, and hence each particle may collide with the electrode multiple times.
Fig. 4a shows i–t results for collisions of PtNP@ssDNA with an Hg UME before (black trace) and after (red trace) Exo I digestion of PtNP@ssDNA. The inset in this frame is an expanded view of the black trace over a limited time window. These data show that in the absence of Exo I exposure, PtNP collisions yield small and infrequent collisions having a magnitude of ∼5 pC. Note that due to the spike-shaped i–t transients that occur at Hg UMEs, it is conventional to report their magnitude in terms of charge rather than current.22 After enzymatic digestion, more frequent, spike-shaped collisions are observed as shown in the expanded view of the i–t trace in Fig. 4b. Comparison of the red traces in Fig. 3a (Au UME) and 4a (Hg UME) also shows that the baseline current is much more stable for the latter after digestion.
Fig. 4c is a histogram showing the frequency of collisions as a function of their magnitude for post-Exo I PtNP@ssDNA. The average collision charge for these experiments is 63 ± 98 pC, which is ∼50% less than for naked PtNPs: 118 ± 128 pC. This trend of recovering roughly half the original collision signal is consistent with that observed for Au UMEs. Moreover, the broader distribution of collision charges (determined from the integrated current transients) on Hg is consistent with results reported by Stevenson and coworkers.22
The collision frequency of PtNP@ssDNA post-Exo I as a function of concentration at an Hg UME is shown in Fig. 4d. As for the Au UME data (Fig. 3d), this plot is linear over most of the concentration range studied. The interesting result, however, is that the collision frequencies are significantly lower at every concentration than for the Au UME results (Table 1). This finding is in contrast to previously reported observations in which the larger surface area of the hemispherical Hg UME led to higher collision frequencies.21 It has been found previously that deposition of Hg on Pt yields a hemispherical Hg drop.24 If we make that assumption here too, then the geometric surface area of the Hg electrode is 157 μm2, compared to 123 μm2 for the Au disk. In other words, the relative surface areas of the two electrodes does not account for the observation of lower collision frequencies on the Hg UME. As suggested earlier, however, this difference might be due to specific interactions between the ssDNA-coated NPs and the Au UME that lead to multiple collisions per PtNP at the Au UME. In contrast, Hg deactivates every PtNP upon impact,22 and thus each PtNP@ssDNA post-Exo I will only produce one current transient per collision event.
PtNP–ssDNA concentration (pM) | Frequency (Hz) Au UME | Frequency (Hz) Hg UME |
---|---|---|
0.58 | 0.047 ± 0.014 | 0.03 ± 0.08 |
1.17 | 0.077 ± 0.012 | 0.057 ± 0.008 |
4.0 | 0.084 ± 0.011 | 0.072 ± 0.010 |
5.8 | 0.12 ± 0.01 | 0.083 ± 0.010 |
11.7 | 0.19 ± 0.03 | 0.11 ± 0.01 |
Finally, there are three specific conclusions that can be drawn from the results presented here. First, PtNPs modified with ssDNA are not irreversibly passivated, and at least ∼50% of the original current can be recovered after treatment with Exo I. Second, PtNP@ssDNA remain colloidally stable under the conditions required for enzymatic digestion (e.g., high salt concentration). Third, the lower collision frequency observed at Hg UMEs, compared to Au UMEs, may be indicative of single NPs producing more than one signal per collision event at Au electrodes. This is an important point, because practical applications of ECA require quantitative correlation of the concentration of a target with the collision frequency and/or signal magnitude. These findings set the stage for future biosensing applications of ECA, and we will report the results of those studies in due course.
Footnotes |
† Electronic supplementary information (ESI) available: Supporting data including size distribution analysis of the PtNPs by TEM, a fluorescence calibration curve for estimating the coverage of ssDNA on PtNPs, ECA of PtNP@ssDNA modified with thiol on the 3′-end, and i–t curves for collisions of naked PtNPs at Au UMEs. See DOI: 10.1039/c6sc02165d |
‡ Current address: Skolkovo Institute of Science and Technology, 3 Nobel Street, Moscow, Russia 143026. |
This journal is © The Royal Society of Chemistry 2016 |