DT-Diaphorase as a Bifunctional Enzyme Label That Allows Rapid

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DT-Diaphorase as a Bifunctional Enzyme Label That Allows Rapid Enzymatic Amplification and Electrochemical Redox Cycling Cheolho Kang,† Juyeon Kang,† Nam-Sihk Lee,‡ Young Ho Yoon,‡ and Haesik Yang*,† †

Department of Chemistry and Chemistry Institute for Functional Materials, Pusan National University, Busan 46241, Korea EOne Laboratories, Incheon 22014, Korea

‡

S Supporting Information *

ABSTRACT: The most common enzyme labels in enzyme-linked immunosorbent assays are alkaline phosphatase and horseradish peroxidase, which, however, have some limitations for use in electrochemical immunosensors. This Article reports that the small and thermostable DT-diaphorase (DT-D) and electrochemically inactive 4-nitroso-1-naphthol (4-NO-1-N) can be used as a bifunctional enzyme label and a rapidly reacting substrate, respectively, for electrochemical immunosensors. This enzyme−substrate combination allows high signal amplification via rapid enzymatic amplification and electrochemical redox cycling. DT-D can convert an electrochemically inactive nitroso or nitro compound into an electrochemically active amine compound, which can then be involved in electrochemical−chemical (EC) and electrochemical−enzymatic (EN) redox cycling. Six nitroso and nitro compounds are tested in terms of signal-to-background ratio. Among them, 4-NO-1-N exhibits the highest signal-to-background ratio. The electrochemical immunosensor using DT-D and 4-NO-1-N detects parathyroid hormone (PTH) in phosphate-buffered saline containing bovine serum albumin over a wide range of concentrations with a low detection limit of 2 pg/mL. When the PTH concentration in clinical serum samples is measured using the developed immunosensor, the calculated concentrations are in good agreement with the concentrations obtained using a commercial instrument. Thus, the use of DT-D as an enzyme label is highly promising for sensitive electrochemical detection and point-ofcare testing.

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back to the original substrate. However, during electrochemical reduction, H2O2 reduction can also occur, making it difficult to obtain low background levels.9 Moreover, the use of H2O2 makes HRP unsuitable for point-of-care testing because H2O2 cannot exist in a stable dried form. DT-diaphorase [NAD(P)H dehydrogenase (quinone) (EC 1.6.99.2, currently transferred to EC 1.6.5.2)] (DT-D) is a redox enzyme that catalyzes a two-electron reduction of quinone in the presence of NADH or NADPH,10,11 and can reduce nitro and nitroso groups to amine groups even under aerobic conditions.12,13 Importantly, DT-D from Bacillus stearothermophilus (EC 1.6.99.-) has a low molecular weight (30 kDa) and high thermal stability.10,14 Because of its unique characteristics along with a high catalytic performance, DT-D has been used to obtain amplified signals via electrochemicalenzymatic (EN) redox cycling2 of the enzyme product produced by enzyme labels, such as ALP15,16 and βgalactosidase.17 Nevertheless, DT-D has never been employed as an enzyme label in ELISAs. Reduction of a nitro group to an amine group requires the transfer of 6 electrons, whereas reduction of a nitroso group to an amine group requires the transfer of 4 electrons.18 To obtain high signal levels via rapid catalytic conversion, it is better to

nzyme-linked immunosorbent assays (ELISAs) have been widely employed for sensitive detection of various targets, such as small molecules, biomolecules, and microorganisms.1−3 In ELISAs, an enzyme label that can rapidly produce many signaling chemical species plays a crucial role in obtaining high sensitivity and reproducibility. The enzyme label should catalyze its enzymatic reaction with a high reaction rate and exhibit long-term stability under both dry and physiological buffer conditions. The most common enzyme labels that meet such requirements are alkaline phosphatase (ALP) and horseradish peroxidase (HRP).4 ALP is a nonredox (hydrolytic) enzyme that removes the phosphate group from a phosphorylated substrate.4 In electrochemical detection, this dephosphorylation converts an electrochemically non- or less-active substrate into an electrochemically active product. However, in some cases, low stability of the phosphorylated substrate (e.g., 4-aminophenyl phosphate) in solution5 and a small difference in the formal potentials between the substrate and the product (e.g., 4aminophenyl phosphate and 4-aminophenol)6 make obtaining low background levels difficult. Moreover, the large size of ALP (molecular weight = 150 kDa)4 hinders low nonspecific adsorption and multiple conjugation to the probe antibody. HRP (molecular weight = 40 kDa)4 is a redox enzyme that catalyzes the oxidation of various organic and organometallic substrates in the presence of H2O2.7,8 In electrochemical detection, the oxidized product is electrochemically reduced © XXXX American Chemical Society

Received: April 2, 2017 Accepted: June 20, 2017

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DOI: 10.1021/acs.analchem.7b01223 Anal. Chem. XXXX, XXX, XXX−XXX

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Analytical Chemistry

Figure 1. (a) Nitroso and nitro compounds selected as potential DT-D substrates. (b) Schematic diagram of an electrochemical immunosensor using DT-D as an enzyme label. A high signal level is obtained using (i) enzymatic amplification, (ii + iii) EC redox cycling, and (ii + iv) EN redox cycling.

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EXPERIMENTAL SECTION Chemicals and Solutions. DT-D from Bacillus stearothermophilus (EC 1.6.99.-) and diaphorase (EC 1.8.1.4) were obtained from Nipro Co. (Osaka, Japan). Nitroreductase from Escherichia coli (N9284), 4-nitroso-1-naphthol (4-NO-1-N), 1nitroso-2-naphthol (1-NO-2-N), 4-nitrophenol (4-NO2-1-P), NADH (β-nicotinamide adenine dinucleotide reduced dipotassium salt), bovine serum albumin (BSA), tris(hydroxymethyl)aminomethane (tris), phosphate buffered saline (PBS), and all the reagents used for the optimization experiment and electrode pretreatment were purchased from Sigma-Aldrich Co. 4-Nitrosophenol (4-NO-1-P), 4-nitro-1-naphthol (4-NO21-N), and 1-nitro-2-naphthol (1-NO2-2-N) were obtained from Tokyo Chemical Industry Co., Ltd. Two goat polyclonal antiPTH IgGs (70-XG67 and 70-XG68) and recombinant PTH (30R-2670) were obtained from Fitzgerald, Inc. (Acton, MA, USA). Sulfosuccinimidyl-4-[N-maleimidomethyl]cyclohexane1-carboxylate (sulfo-SMCC), N-succinimidyl-S-acetylthiopropionate (SATP), and sulfosuccinimidyl-6-[biotinamido]-6-hexanamidohexanoate (EZ-link sulfo-NHS-LC-LC-biotin) were obtained from Thermo Fisher Scientific Inc. (Meridian, Rockford, USA). Tris buffers (pH 7.5 and 9.0) were prepared using 50 mM tris and 1.0 M HCl. The phosphate-buffered saline (PBS buffer, pH 7.4) contained 10 mM phosphate, 0.138 M NaCl, and 2.7 mM KCl. PBSB contained all of the ingredients in PBS with 1% (w/v) BSA. The rinsing buffer (pH 7.6) contained 50 mM tris, 40 mM HCl, 0.05% (w/v) BSA, 0.05% Tween 20, and 0.5 M NaCl. ITO electrodes were purchased from Corning Co. (Daegu, Korea). Conjugation. DT-D-conjugated anti-PTH IgG was prepared by cross-linking the amine group of anti-PTH IgG and the amine group of DT-D. Anti-PTH IgG (100 μg/mL) in 1 mL of PBS and SATP (2 mg/mL) in 10 μL of PBS were mixed for 30 min at room temperature. Afterward, the solution was mixed with 20 μL of deacetylation solution containing 0.012 g/ mL ethylenediaminetetraacetic acid and 0.044 g/mL hydroxylamine·HCl for 2 h at room temperature, and the SATP-

use a nitroso compound as a DT-D substrate rather than a nitro compound. However, a nitroso compound might react with NADH in the absence of DT-D, which increases the background levels. Nitro and nitroso groups are strongly electron-withdrawing, but amine groups are strongly electrondonating. When a nitro or nitroso compound is converted to an amine compound, there is a big decrease in formal potential. It is much easier to oxidize the amine compound than the nitro or nitroso compound, which allows low background levels at an applied potential where amine compound is oxidized. Electrochemical redox cycling combined with enzymatic amplification by an enzyme label allows higher signal amplification than enzymatic amplification alone.2 Redox cycling can be obtained both without or with a redox enzyme. Electrochemical−chemical (EC) redox cycling19−21 is obtained in the presence of a reductant or oxidant without a redox enzyme, whereas EN redox cycling22−24 is obtained with a redox enzyme. When DT-D is used as an enzyme label, it can act both as a redox enzyme for EN redox cycling of an electrochemically active species and as a redox enzyme for the enzymatic amplification that converts an electrochemically inactive species into an active one. However, an immunosensor using such a bifunctional enzyme label has not been reported previously. Here, we report an electrochemical immunosensor that uses DT-D as the redox enzyme label. The enzyme converts an electrochemically inactive nitroso compound to an electrochemically active amine compound, which is then involved in EC and EN redox cycling. Six nitroso and nitro compounds (Figure 1a) were tested and compared in terms of signal-tobackground ratio, and optimum conditions were determined. Finally, the developed immunosensor using the bifunctional DT-D was applied for the detection of parathyroid hormone (PTH)25 in buffer and human serum (Figure 1b). B

DOI: 10.1021/acs.analchem.7b01223 Anal. Chem. XXXX, XXX, XXX−XXX

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Analytical Chemistry conjugated anti-PTH IgG was then filtered by centrifugation for 20 min at 12 000 rpm. The filtrate was dissolved in 1 mL of PBS. DT-D (100 μg/mL) in 1 mL of PBS and sulfo-SMCC (1 mg/mL) in 50 μL of PBS were mixed and incubated for 30 min at room temperature. Sulfo-SMCC-conjugated DT-D was filtered by centrifugation for 20 min at 12 000 rpm. SATPconjugated anti-PTH IgG (100 μg/mL) in 1 mL of PBS was then mixed with sulfo-SMCC-conjugated DT-D (100 μg/mL) in 1 mL of PBS at a molar ratio of 1:1, and the mixture was incubated for 2 h at room temperature. The mixture was then centrifuged for 20 min at 12 000 rpm to filter the DT-D conjugated PTH antibody. The filtrate was dissolved in 1 mL of PBSB. Any noticeable change in the activity of DT-D was not observed one month after its conjugation (Figure S-1). Biotinylated anti-PTH IgG was obtained by cross-linking EZlink sulfo-NHS-LC-LC-biotin and the amine group of antiPTH IgG. Anti-PTH IgG (100 μg/mL) dissolved in 1 mL of PBS was mixed with 1.33 μL of 10 mM EZ-link sulfo-NHS-LCLC-biotin and incubated overnight at 4 °C. The mixture was centrifuged for 20 min at 12 000 rpm to filter biotinylated antiPTH IgG. The filtrate was dissolved in 1 mL of PBSB. Preparation of Sensing Electrodes. ITO electrodes were pretreated with a 5:1:1 solution of H2O, H2O2 (30%), and NH4OH (30%) at 70 °C for 1 h.26 To obtain the avidinmodified ITO electrodes (avidin/ITO electrodes), 70 μL of a carbonate buffer solution (20 mM, pH 9.6) containing 10 μg/ mL avidin was dropped onto the ITO electrodes and incubated for 2 h at 20 °C.20 Next, 70 μL of PBSB solution was dropped onto the avidin/ITO electrodes to obtain the BSA/avidin/ITO electrodes. Biotinylated anti-PTH IgG was immobilized by dropping 70 μL of PBSB solution containing 10 μg/mL biotinylated anti-PTH IgG onto the BSA/avidin/ITO electrodes. The sensing electrodes were maintained in the treated state for 30 min at 4 °C. Procedure for PTH Detection. PBSB containing different concentrations of PTH (or clinical serum samples) (70 μL) was dropped onto a sensing electrode and incubated for 30 min at 4 °C, and the sensing electrode was then washed with rinsing buffer. Subsequently, 70 μL of PBSB containing 10 μg/mL DTD-conjugated anti-PTH IgG was dropped onto the electrodes and incubated for 30 min at 4 °C, followed by washing with rinsing buffer. Tris buffer (pH 7.5) containing 1.0 mM NADH and 0.1 mM 4-NO-1-N was injected into a Teflon electrochemical cell that consisted of the sensing electrode, an Ag/ AgCl (3 M NaCl) reference electrode, and a platinum counter electrode. The exposed geometric area of the sensing electrode was approximately 0.28 cm 2. Cyclic voltammetry and chronocoulometry were performed using a CHI 708C system (CH Instruments, Austin, TX, USA). PTH in the clinical serum samples was measured using the Cobas e602 analyzer (Roche Diagnostics GmbH, Mannheim, Germany). The study protocol using clinical serum samples was approved by the Institutional Review Board of EOne Laboratories (#128477-201611-BR011). The UV−vis spectrum was obtained using a UV-1650 (SHIMADZU, Kyoto, Japan).

should be very slow in the absence of DT-D while it should be very fast in the presence of DT-D. Therefore, it is of primary importance to choose an appropriate substrate containing a nitro or nitroso group that exhibits such reaction requirements. To obtain a high electrochemical signal-to-background ratio, the reduced product containing an amine group should have a formal potential near 0 V and should be readily electro-oxidized at an electrode. Because aminonaphthols and aminophenols meet these requirements,20 six compounds containing a nitro or nitroso group (4-NO-1-N, 1-NO-2-N, 4-NO2-1-N, 1-NO2-2-N, 4-NO-1-P, and 4-NO2-1-P shown in Figure 1a) were selected as potential DT-D substrates. To investigate the possibility of a direct (or ITO-mediated) reaction between the substrate and NADH, cyclic voltammograms were obtained 10 min after mixing one of the six substrates with NADH (Figures 2a and S-2a). The currents in

Figure 2. Cyclic voltammograms obtained (at a scan rate of 20 mV/s) at bare ITO electrodes after an incubation period of 10 min at 25 °C (a) in (i) tris buffer (pH 7.5) and (ii, iii, iv) tris buffer (pH 7.5) containing 1.0 mM NADH and 0.1 mM (ii) 1-NO-2-N, (iii) 4-NO-1N, or (iv) 4-NO-1-P and (b) in tris buffer (pH 7.5) containing 1.0 mM NADH, 10 μg/mL DT-D, and 0.1 mM (i) 4-NO-1-N or (ii) 4-NO-1P.

the cyclic voltammogram for 1-NO-2-N (curve ii of Figure 2a) were much larger than those for the buffer solution (curve i of Figure 2a). The increased currents indicate that an electroactive species such as 1-amino-2-naphthol (1-NH 2 -2-N) was produced as a result of the direct (and/or ITO-mediated) reaction between 1-NO-2-N and NADH. The near-limitingcurrent behavior observed in the voltammogram was due to the EC redox cycling of 1-NH2-2-N by NADH. Consequently, 1NO-2-N was excluded as a DT-D substrate. The direct (and/or ITO-mediated) reaction between 4-NO-1-P and NADH was also substantial (curve iv of Figure 2a). In case of the other four substrates, only capacitive currents were observed up to 0.3 V.

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RESULTS AND DISCUSSION DT-D can catalyze the reduction of nitro and nitroso groups by NAD(P)H. However, nitro and nitroso groups can also react directly (and/or by ITO mediation) with NAD(P)H in the absence of DT-D. To obtain a high signal-to-background ratio in an immnosensor using DT-D as an enzyme label, the reaction between the nitro or nitroso group and NAD(P)H C

DOI: 10.1021/acs.analchem.7b01223 Anal. Chem. XXXX, XXX, XXX−XXX

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Analytical Chemistry Unlike that for 1-NO-2-N, the direct (and/or ITO-mediated) reaction between 4-NO-1-N and NADH was very slow or did not occur (curve iii of Figure 2a). As the next step to find a good substrate offering a high signal-to-background ratio, cyclic voltammograms were obtained 10 min after mixing one of the five remaining substrates with NADH and DT-D (Figures 2b and S-2b). High anodic currents were observed for 4-NO-1-N (curve i of Figure 2b), whereas much lower anodic currents were observed for 4-NO1-P (curve ii of Figure 2b). The high currents indicate that 4NO-1-N was catalytically and rapidly converted to 4-amino-1naphthol (4-NH2-1-N). Importantly, the high currents near 0.2 V were obtained even at a low electrocatalytic ITO electrode without modification with an electrocatalytic material. In case of the three nitro compounds, anodic currents were much lower (Figure S-2b). 4-NO-1-N was better than the other substrates in terms of signal-to-background ratio. Consequently, 4-NO-1-N was chosen as a DT-D substrate. Time-course absorbance data (Figure S-3) were also obtained to confirm the electrochemical results. The absorbance for 4-NO-1-N did not change for 10 min in the absence of DT-D, whereas the absorbance rapidly decreased with time in the presence of DT-D because 4-NH2-1-N exhibits an extinction coefficient much lower than 4-NO-1-N. The absorbance data clearly show that the direct (and/or ITOmediated) reaction between 4-NO-1-N and NADH was negligible or did not occur but that the catalytic reduction of 4-NO-1-N by DT-D was very fast. Diaphorases and nitroreductases, as well as DT-D, might be used as an enzyme label for the reduction of 4-NO-1-N.27,28 When a diaphorase (EC 1.8.1.4) was tested, no noticeable current increase due to the catalytic reduction of 4-NO-1-N was observed after an incubation period of 10 min (curve i of Figure 3a). In case of a nitroreductase from Escherichia coli, a substantial increase in current was observed (curve ii of Figure 3a). However, this increase was much lower than that for DT-D (curve i of Figure 2b). Thus, DT-D was much better as an enzyme label than the tested diaphorase and nitroredutase. The product of the catalytic reduction of 4-NO-1-N by DTD (4-NH2-1-N) can be oxidized by the oxygen dissolved in solution, even though DT-D is insensitive to oxygen. Nevertheless, the oxidized species can be rapidly reduced back to 4-NH2-1-N by the excess NADH. For this reason, the cyclic voltammogram obtained in O2-saturated PBS containing 4-NO-1-N, NADH, and DT-D was only slightly lower than that obtained in air-saturated PBS (Figure S-4). Enzyme activity and electrochemical reactions highly depend on pH and buffer composition. In general, electrochemical detection and enzymatic amplification are carried out in PBS (pH 7.4) or tris buffer (∼pH 9). Although a high electrochemical signal was obtained in tris buffer of pH 9.0 (curve i of Figure 3b), it was lower than that obtained in tris buffer of pH 7.5 (curve i of Figure 2b). In PBS of pH 7.4, high overpotential was required to oxidize 4-NH2-1-N (curve ii of Figure 3b). Therefore, tris buffer at pH 7.5 was used in detection experiments. In many cases, charge data allow better signal-to-background ratios than current data. Chronocoulometric data were also obtained in the absence and presence of DT-D (Figure 4a). The difference between the two charge data was very high, because the direct (and/or ITO-mediated) reaction between 4NO-1-N and NADH was negligible and the catalytic reduction of 4-NO-1-N by DT-D was very fast. To determine an

Figure 3. Cyclic voltammograms obtained (at a scan rate of 20 mV/s) at bare ITO electrodes after an incubation period of 10 min at 25 °C (a) in tris buffer (pH 7.5) containing 0.1 mM 4-NO-1-N, 1.0 mM NADH, and 10 μg/mL (i) diaphorase or (ii) nitroreductase and (b) in (i) tris buffer (pH 9.0) containing 0.1 mM 4-NO-1-N, 1.0 mM NADH, and 10 μg/mL DT-D and (ii) PBS (pH 7.4) containing 0.1 mM 4-NO-1-N, 1.0 mM NADH, and 10 μg/mL DT-D.

optimum applied potential in chronocoulometry, the charge data obtained at four applied potentials (0.10, 0.15, 0.20, and 0.25 V) were compared (Figure 4b). The charge measured in the absence of DT-D corresponds to a background value, whereas the charge measured in the presence of DT-D corresponds to a signal level. A high signal-to-background ratio was obtained at 0.20 and 0.25 V. Because a lower potential is better for obtaining more reproducible results, 0.20 V was selected as the applied potential. A sandwich-type electrochemical immunosensor using a DTD label and the optimum conditions was then applied for PTH detection (Figure 1b). During the detection, PTH was captured by a biotinylated capture IgG, and a DT-D-conjugated detection IgG was then attached to PTH. The DT-D label catalyzed the reduction of 4-NO-1-N to 4-NH2-1-N by NADH. After an incubation period of 10 min for the enzymatic reaction, 4-NH2-1-N was electrochemically oxidized at the ITO electrode. The oxidized species was reduced back to 4-NH2-1N directly by NADH and with the help of DT-D. The regenerated 4-NH2-1-N was again electrochemically oxidized. This redox cycling significantly increases the electrochemical signal. When the electrochemical oxidation (reaction ii of Figure 1b) and direct reduction (reaction iii of Figure 1b) are combined, this coupled reaction corresponds to EC redox cycling. When the electrochemical oxidation (reaction ii of Figure 1b) and catalytic reduction (reaction iv of Figure 1b) are combined, this coupled reaction corresponds to EN redox cycling. EC and EN redox cycling along with enzymatic amplification allows a highly amplified electrochemical signal. D

DOI: 10.1021/acs.analchem.7b01223 Anal. Chem. XXXX, XXX, XXX−XXX

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Analytical Chemistry I1 = nFAC4‐NH2‐1‐N D4‐NH2‐1‐Nk1C NADH

(1)

I2 = nFAC4‐NH2‐1‐N D4‐NH2‐1‐Nk 2C DT‐D

(2)

C4‑NH2‑1‑N, CNADH, and CDT‑D (0.32 μM) are the concentrations of 4-NH2-1-N, NADH, and DT-D, respectively, and D4‑NH2‑1‑N is the diffusion coefficient of 4-NH2-1-N (6.3 × 10−6 cm2/s).32 n is the number of electrons involved: two electrons are assumed to be involved in both reactions. F and A are Faradaic constant and electrode area (0.28 cm2), respectively. The apparent k1 and k2 values were 2.2 and 3.3 × 106 M−1 s−1, respectively. Because the apparent k1 and k2 values were measured using air-saturated solutions and dissolved oxygen could oxidize 4-NH2-1-N, the real k1 and k2 values would be larger. The apparent k2 value for EN redox cycling using DT-D was comparable to the reported values (6 × 105−8 × 108 M−1 s−1).33 Although the apparent k1 value for EC redox cycling and the current I1 were considerable, the contribution of EC redox cycling to the total current (I1 + I2) obtained in the presence of 4-NH2-1-N, DT-D, and NADH was much smaller than that of EN redox cycling because k1 was much smaller than k2. In immunosensing experiments, the amount of the DT-Dconjugated IgG that is affinity-bound on the immunosensing electrode is very low, and the signal amplification due to EC redox cycling is independent of the amount. Under this condition, the contribution of EC redox cycling to signal amplification becomes much larger whereas the contribution of EN redox cycling becomes much lower. Even in immunosensing experiments for very low concentrations of PTH, the signal amplification due to EN, as well as EC, redox cycling is expected to be substantial because k2 is very high. Figure 5a shows chronocoulograms obtained at the sensing electrodes for various concentrations of PTH. The charge values increased with increasing the concentration of PTH. The calibration plot obtained from the charge values measured at 100 s is shown in Figure 5b. The calculated detection limit was ∼2 pg/mL, which is very low. Importantly, PTH was detected over a wide range of concentrations. Table S-1 shows comparison of PTH detection methods reported previously. Only our method could detect PTH with a short detection time and a low detection limit over a very wide range of concentrations. To validate the developed immunosensor, the concentration of PTH in clinical serum samples was measured. The results for 10 clinical serum samples are shown in Figure 6. The concentration calculated using the calibration plot in Figure 5b was in good agreement with the concentration measured using a commercial instrument except for three high concentration data. This good agreement indicates that the developed immunosensor is practically appealing for the detection of PTH in clinical samples.

Figure 4. (a) Chronocoulograms obtained at 0.20 V at avidin-modified ITO electrodes after an incubation period of 10 min at 25 °C in tris buffer (pH 7.5) containing (i) 0.1 mM 4-NO-1-N and 1.0 mM NADH or (ii) 0.1 mM 4-NO-1-N, 1.0 mM NADH, and 10 μg/mL DT-D. (b) Histogram of the signal-to-background (S/B) ratios calculated from charge values measured at 100 s in the chronocoulograms obtained at avidin-modified ITO electrodes at four applied potentials (0.10, 0.15, 0.20, and 0.25 V) after an incubation period of 10 min at 25 °C in tris buffer (pH 7.5) containing (i) 0.1 mM 4-NO-1-N and 1.0 mM NADH or (ii) 0.1 mM 4-NO-1-N, 1.0 mM NADH, and 10 μg/mL DT-D.

To investigate nonspecific binding of DT-D, cyclic voltammograms and chronocoulograms were obtained after an incubation period of 10 min in a solution containing 4-NO1-N and NADH at bare ITO and BSA/avidin/ITO electrodes on which DT-D was nonspecifically adsorbed and then washed (Figure S-5). The nonspecific binding was significantly decreased on BSA/avidin/ITO electrode compared to bare ITO electrode. To investigate the relative contribution of EC and EN redox cycling to signal amplification, the rate constants for EC and EN redox cycling were calculated using limiting currents obtained at 50 s in chronoamperograms (Figure S-6). When the concentration of NADH (5.0 mM) is much higher than that of 4-NH2-1-N (0.05 mM) and the applied potential in chronoamperometry (0.35 V) is much higher than the formal potential of 4-NH2-1-N, the difference between the limiting current obtained in the presence of 4-NH2-1-N and NADH and the limiting current obtained in the presence of NADH only (I1, 7.13 × 10−7 A) is related to the rate constant of EC redox cycling (k1 in Figure 1). The difference between the limiting current obtained in the presence of 4-NH2-1-N, DT-D, and NADH and the limiting current obtained in the presence of 4NH2-1-N and NADH (I2, 7.01 × 10−6 A) is related to the rate constant of EN redox cycling (k2 in Figure 1). Under the experimental condition, I2 was approximately 10 times higher than I1. I1 and I2 have the following relationships with k1 and k2, respectively:19,29−31

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CONCLUSIONS We developed an electrochemical immunosensor using the small, thermostable, and bifunctional DT-D and rapidly reacting 4-NO-1-N as a new enzyme label and substrate, respectively. DT-D allowed high signal amplification via fast enzymatic amplification and fast EC and EN redox cycling. DTD converts the electrochemically inactive 4-NO-1-N into the electrochemically active 4-NH2-1-N, which is then involved in EC and EN redox cycling. PTH was detected in PBSB over a wide range of concentrations with a low detection limit of 2 pg/ mL. When the concentration of PTH in clinical serum samples E

DOI: 10.1021/acs.analchem.7b01223 Anal. Chem. XXXX, XXX, XXX−XXX

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Cyclic voltammograms for DT-D stability, different DTD substrates, O2 effect, and nonspecific binding and chronoamperograms for the measurement of rate constants (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Fax: (+82)-51-516-7421. ORCID

Haesik Yang: 0000-0001-7450-5915 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research was supported by the National Research Foundation of Korea (2015R1A2A2A01002695 and 2016M3A7B4910538). This research is also supported by the Ministry of Trade, Industry & Energy (MOTIE, Korea) under Industrial Technology Innovation Program (No. 10062995).

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Figure 5. (a) Chronocoulograms obtained at 0.20 V in tris buffer (pH 7.5) containing 0.1 mM 4-NO-1-N and 1.0 mM NADH, using sensing electrodes treated with PBSB containing various concentrations of PTH. (b) Calibration plot of the charge measured at 100 s in the chronocoulograms of panel a. Each experiment at different concentrations was carried out with three different electrodes for assaying the same sample. All charge data were subtracted by the mean value obtained from seven measurements at zero concentration. The dashed line corresponds to 3 times the charge standard deviation (SD) at zero concentration.

Figure 6. Comparison between the PTH concentration measured with the immunosensor and the PTH concentration measured with a commercial instrument for 10 clinical serum samples.

was measured, it was found to be in good agreement with that measured using a commercial instrument. Thus, the electrochemical immunosensor using DT-D is highly promising for sensitive point-of-care testing.

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REFERENCES

(1) Wild, D. The Immunoassay Handbook, 4th ed.; Elsevier: Great Britain, 2013. (2) Labib, M.; Sargent, E. H.; Kelley, S. O. Chem. Rev. 2016, 116, 9001−9090. (3) Yang, H. Curr. Opin. Chem. Biol. 2012, 16, 422−428. (4) Savage, D.; Mattson, G.; Desai, S.; Nielander, G.; Morgensen, S.; Conklin, E. Avidin−Biotin Chemistry: A Handbook, 2nd ed.; Pierce; Rockford, IL, 1994; pp 147−160. (5) Wilson, M. S.; Rauh, R. D. Biosens. Bioelectron. 2004, 20, 276− 283. (6) Aziz, M. A.; Selvaraju, T.; Yang, H. Electroanalysis 2007, 19, 1543−1546. (7) Liu, Z.-M.; Yang, H.-F.; Li, Y.-F.; Liu, Y.-L.; Shen, G.-L.; Yu, R.-Q. Sens. Actuators, B 2006, 113, 956−962. (8) Kasai, S.; Yokota, A.; Zhou, H.; Nishizawa, M.; Niwa, K.; Onouchi, T.; Matsue, T. Anal. Chem. 2000, 72, 5761−5765. (9) Kang, H. J.; Aziz, M. A.; Jeon, B.; Jo, K.; Yang, H. Electroanalysis 2009, 21, 2647−2652. (10) Ogino, Y.; Takagi, K.; Kano, K.; Ikeda, T. J. Electroanal. Chem. 1995, 396, 517−524. (11) Takagi, K.; Kano, K.; Ikeda, T. J. Electroanal. Chem. 1998, 445, 211−219. (12) Koga, N.; Hokama-Kuroki, Y.; Yoshimura, H. Chem. Pharm. Bull. 1990, 38, 1096−1097. (13) Sarlauskas, J.; Dickancaite, E.; Nemeikaite, A.; Anusevicius, Z.; Nivinskas, H.; Segura-Aguilar, J.; Cenas, N. Arch. Biochem. Biophys. 1997, 346, 219−229. (14) https://www.nipro.co.jp/en/business/other/enzymes/di-1.pdf (accessed on March 26, 2017). (15) Campàs, M.; de la Iglesia, P.; Le Berre, M.; Kane, M.; Diogène, J.; Marty, J.-L. Biosens. Bioelectron. 2008, 24, 716−722. (16) Limoges, B.; Marchal, D.; Mavré, F.; Savéant, J.-M. J. Am. Chem. Soc. 2006, 128, 6014−6015. (17) Rochelet-Dequaire, M.; Djellouli, N.; Limoges, B.; Brossier, P. Analyst 2009, 134, 349−353. (18) Blaser, H.-U. Science 2006, 313, 312−313. (19) Das, J.; Jo, K.; Lee, J. W.; Yang, H. Anal. Chem. 2007, 79, 2790− 2796. (20) Akanda, M. R. H.; Aziz, M. A.; Jo, K.; Tamilavan, V.; Hyun, M. H.; Kim, S.; Yang, H. Anal. Chem. 2011, 83, 3926−3933. (21) Park, S.; Kim, J.; Ock, H.; Dutta, G.; Seo, J.; Shin, E.-C.; Yang, H. Analyst 2015, 140, 5481−5487. (22) Singh, A.; Park, S.; Yang, H. Anal. Chem. 2013, 85, 4863−4868. (23) Dutta, G.; Kim, S.; Park, S.; Yang, H. Anal. Chem. 2014, 86, 4589−4595.

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Analytical Chemistry (24) Dutta, G.; Park, S.; Singh, A.; Seo, J.; Kim, S.; Yang, H. Anal. Chem. 2015, 87, 3574−3578. (25) Delmas, P. D.; Vergnaud, P.; Arlot, M. E.; Pastoureau, P.; Meunier, J. P.; Nilssen, M. H. L. Bone 1995, 16, 603−610. (26) Choi, M.; Jo, K.; Yang, H. Bull. Korean Chem. Soc. 2013, 34, 421−425. (27) Valiauga, B.; Williams, E. M.; Ackerley, D. F.; Č ėnas, N. Arch. Biochem. Biophys. 2017, 614, 14−22. (28) Chen, J.; Dai, R. J.; Tong, B.; Xiao, S. Y.; Meng, W. Chin. Chem. Lett. 2007, 18, 10−12. (29) Bard, A. J.; Faulkner, L. R. Electrochemical Methods: Principles and Applications, 2nd ed.; John Wiley & Sons: New York, 2001; pp 501−503. (30) Chen, L.; Gorski, W. Anal. Chem. 2001, 73, 2862−2868. (31) Ogino, Y.; Takagi, K.; Kano, K.; Ikeda, T. J. Electroanal. Chem. 1995, 396, 517−524. (32) Sharma, L.; Kalia, R. J. Chem. Eng. Data 1977, 22, 39−41. (33) Limoges, B.; Marchal, D.; Mavré, F.; Savéant, J.-M. J. Am. Chem. Soc. 2006, 128, 2084−2092.

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DOI: 10.1021/acs.analchem.7b01223 Anal. Chem. XXXX, XXX, XXX−XXX