Preparative Biochemistry and Biotechnology
ISSN: 1082-6068 (Print) 1532-2297 (Online) Journal homepage: http://www.tandfonline.com/loi/lpbb20
Comparative Evaluation of an Electrochemical Bioreporter for Detecting Phenolic Compounds Hae Ja Shin & Woon Ki Lim To cite this article: Hae Ja Shin & Woon Ki Lim (2014): Comparative Evaluation of an Electrochemical Bioreporter for Detecting Phenolic Compounds, Preparative Biochemistry and Biotechnology, DOI: 10.1080/10826068.2014.979207 To link to this article: http://dx.doi.org/10.1080/10826068.2014.979207
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Date: 05 November 2015, At: 16:40
Comparative Evaluation of an Electrochemical Bioreporter for Detecting Phenolic Compounds Hae Ja Shin1, Woon Ki Lim2 1
Division of Energy and Bio-engineering, Dongseo University, Busan, Republic of Korea, 2 Department of Molecular Biology, College of Natural Sciences, Pusan National University, Busan, Republic of Korea
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Corresponding author Hae Ja Shin, E-mail: [email protected]
Abstract In the present study, we constructed an Escherichia coli-based electrochemical bioreporter (EB) harboring pLZCapR, which encodes the CapR regulatory protein (for phenol degradation) along with β-galactosidase, and examined its ability to detect phenolic compounds as compared with previously reported optical bioreporters (OBs) controlled by CapR and detected using a luminometer (OB-lum) or spectrophotometer (OB-spec). The recombinant E. coli bioreporter cells were immobilized in polyvinyl alcohol (PVA); p-aminophenyl-β-D-galactopyranoside (PAPG) was used as the enzymatic substrate; and electrochemical measurements were taken. The peak current obtained on cyclic voltammetry (CV) was used to measure the redox response of PAPG degradation. Our results revealed that the EB system showed a detection range of 10 nM to 10 mM phenol with a good lower detection limit (30 nM phenol). Furthermore, the detection time was dramatically lower for the EB system (15-20 min) compared to the OBs (~6 h). These responses were reliably repeatable with an acceptable standard deviation (± 2.7%; n = 6), and the system showed good stability without loss of activity over 7 h of operation or following 2 weeks of cold storage. Together, these results show
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that the EB system is faster and has a lower detection limit than the existing optical techniques.
KEYWORDS: Electrochemical Escherichia coli bioreporter, Phenolic compounds
INTRODUCTION
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Microbial bioreporters have been used as first-step monitoring tools to detect target chemicals and physiological signals in diverse fields.[1] Numerous recombinant microbial bioreporters have been constructed by placing a reporter gene (i.e., luc, lacZ or gfp) under the control of regulatory genes that are responsive to the target chemicals and signals.[2] The biological signals from these cellular components are dose-dependent and can generate measurable responses via various transducers. The transducers that have been exploited in the construction and monitoring of microbial bioreporters include electrochemical transducers, which can convert biological signals to electrochemical signals (e.g., currents, potentials and conductivities). Among the electrochemical transducers, amperometers have been widely used to visualize signal responses to target chemicals and physiological signals, such as biochemical oxygen demand (BOD)[3], heavy metals[4], and organophosphates.[5,6]
Potentiometers have also been exploited,
such as for detecting organophosphates with pH electrodes[7], sucrose and BOD with oxygen electrodes[8], and urea with NH4+ ion-selective electrodes.[9] Electrochemical transducers may be cost-effective, portable and miniaturized[10], although some systems require electroactive mediators[11], such as anthraquinone[12] or ferricyanide[13], to improve their electrochemical responses. Overall, the electrochemical microbial
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bioreporters typically show acceptable sensitivity and stability, but their selectivity tends to be relatively poor.[14] In terms of optical transducers, luminometers, spectrophotometers, fluorometers, and flow cytometer are commonly used in microbial bioreporter systems. The luminometer is a sensitive, rapid, and portable transducer that may be monitored on-line through optic fibers. Luminescence-based systems are common, but their sensitivity can be decreased by light scattering, such as in turbid
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solutions and under anaerobic conditions.[4] Spectrophotometers and fluorometers are widely used to take sensitive measurements in ordinary laboratories, but they are unwieldy and impractical for on-site measurements. Currently, the flow cytometer has been used to monitor the inner cell information and its kinetic behaviors at individual cell level. [15] To date, no study has made a direct comparison of transducers, even though this could help researchers choose the one most suitable for their purposes in monitoring target chemicals.
In our previous work, we have extensively studied microbial bioreporters that used optical transducers (e.g., luminometers and spectrophotometers) to sense benzene, toluene, ethylbenzene and xylene (BTEX)[16], phenolic compounds[17-19], and salicylate.[20-22] We found that these microbial bioreporters could quantitatively detect aromatic compounds at relatively high concentration ranges (i.e., μM~mM) in ~6 h.[18,20,21]
However, such tests should be speed up for field use. Furthermore,
environmental contamination can involve lower concentrations of aromatic hydrocarbons, necessitating more sensitive detection methods for the monitoring of such compounds.
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Accordingly, it would be useful to compare the analytical performance of bioreporters that use different transducers to detect aromatic compounds.
In the present study, we constructed an amperometer-based electrochemical bioreporter (EB) for the monitoring of phenolic compounds, and compared its ability with those of previously described optical bioreporters (OBs) that detected such compounds by
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luminometry (OB-lum) and spectrophotometry (OB-spec). All of the studied bioreporters were under the control of the CapR regulatory protein, which directs phenol degradation. For the construction of EB, recombinant E. coli cells harboring the plasmid pLZCapR (capR::lacZ) were immobilized in polyvinyl alcohol (PVA). For electrochemical detection of phenolic compounds, p-aminophenyl-β-D-galactopyranoside (PAPG; a substrate of β-galactosidase) was added, and the oxidation of p-aminophenol (PAP; a catabolite of PAPG) was monitored by cyclic voltammetry (CV). This EB system was validated, characterized, and compared with OBs.
EXPERIMENTAL Chemicals The media components were purchased from Difco (MO, USA). The PAPG, PVA, and boric acid were purchased from Sigma (MO, USA). The genomic DNA isolation kit, plasmid isolation spin kit, and gel extraction kit were purchased from Q-BIO Gene (CA, USA), Qiagen (Hilden, Germany), and Cosmogenetech (Seoul, Korea), respectively. AgeI, KpnI, SalI, HindIII, and T4-DNA ligase were purchased from Takara (Shiga, Japan). The pSV-beta-gal plasmid and pGL3b basic vector were purchased from Promega (WI, USA).
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All other utilized reagents were of analytical grade and were used as received without further purification.
Bacteria And Culture Conditions Escherichia coli DH5α (hsdR, recA, Thi-1, relA1, gyrA96) was used to maintain plasmids. P. putida KCTC1452 was obtained from the Korean Collection for Type Cultures (KCTC)
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and grown in tryptic soy broth at 25°C. E. coli DH5α cells harboring plasmids were cultured at 37°C in TYS media (1% tryptone, 0.5% yeast extract, and 0.5% NaCl) containing 30 μg ml-1 ampicillin. The number of colony forming units (CFU) per ml of culture sample was determined by serially diluting the sample and plating the cells on Luria-Bertani agar.
Construction Of Plasmid Plzcapr The capR gene, its own promoter (Pr), and the Po promoter were cloned from the genomic DNA of P. putida KCTC1452, as described previously.[17] Briefly, the capR/Pr/Po fragment was PCR amplified with specific primers that each harbored KpnI sites at their ends (forward, 5´-CCC AAG CTT GGT ACC GCT TAC ATG CCG ATC AAG TAC-3´ and reverse, 5´-CGG GGT ACC AAG CTT TAA CGA GTG AGC TGA TCG AAA-3´). The PCR products were gel-isolated and digested with KpnI. A HindIII/SalI restriction fragment from pGL3 basic and a 3749-bp HindIII/SalI restriction fragment from pSV-beta-gal were ligated with T4-DNA ligase. The resulting 6610-bp pGLβ-GAL fragment was digested with HindIII/AgeI to remove the gpt promoter from pSV-beta-gal; the fragment was then blunt-ended and ligated to generate the promoterless
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pGLβ-GAL. Finally, a fragment of KpnI-digested capR/Pr/Po was inserted at the KpnI site of the 6392-bp pGLβ-GAL to construct the 8538-bp pLZCapR (Fig. 1). This plasmid was transformed into E. coli DH5α by the CaCl2 method. Immobilization Of Bacterial Cells E. coli cells harboring pLZCapR were grown for 16 h at 37℃ with shaking, and then harvested by centrifugation at 10,000 rpm for 15 min. The supernatant was discarded and
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the cells were washed with 50 mM PBS buffer (pH 7.4). To optimize the immobilization of these cells in PVA beads, various amounts (0.3, 0.5, 0.7, 0.9 or 1 gram) of PVA (nominal degree of polymerization = 1750; approx. molecular weight 75,000-80,000) were solubilized in 5 ml of distilled hot water, dissolved completely in a boiling water bath, and then cooled to 40℃. An equal volume of cells (approximately 4.2 x 109 CFU/ml) was mixed with the dissolved PVA solution, and beads were formed by dropping the mixture into a saturated boric acid solution. The resulting beads were soaked in 0.5 M sodium orthophosphate for 1 h, washed with saline, and then either directly subjected to the electrochemical assay for β-galactosidase activity (see below), or stored at 4℃ until use. Proper immobilization was confirmed by scanning electron microscopy (SEM) of control PVA and cell-immobilized PVA (magnification 6000 x).
Apparatus And Procedure For Electrochemical Measurement Of ß-Galactosidase Activity CV measurements were performed using a potentiostat/galvanostat (Cosentech, Korea). All operations, including data acquisition, were completely controlled by the Physiolab kst-p1 software (Cosentech, Korea). The three-electrode system comprised a platinum (Pt)
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working electrode (diameter, 3 mm; working area, 0.071 cm2), an Ag/AgCl reference electrode, and a Pt auxiliary electrode. Microbial cells were immobilized and loaded into the workstation, and β-galactosidase activity was measured using PAPG as the substrate. The three electrodes were placed in contact with the PVA-immobilized bacterial cells in the wells of a 24-well plate containing appropriate volumes of test solution. For CV measurements, bacterial cells that had been treated with or without phenolic compounds
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were placed in PBS buffer containing 15 mM PAPG for 15 min, and then cyclic voltammograms were recorded (cyclic scan range, -100 mV to +500 mV; scan rate, 100 mV s-1). All experiments were carried out at room temperature, and were assessed under the optimum pH (7.4), amount of cell loading (~ 4.2 x 109 CFU/ml), induction time (2 h), and concentrations of phenolic compounds. To examine selectivity, we assessed the biosensor activities obtained in response to 0.1 mM of 2-chlorophenol, 2-methylphenol, 2-nitrophenol, 3-methylphenol, 4-chlorophenol, catechol, resorcinol, and 2,5dimethylphenol. These phenolic compounds were dissolved in ethanol and spiked into either 50 mM PBS buffer (pH 7.4) or untreated hospital wastewater from several sources in Busan City. To test response repeatability, we examined the standard deviation from six consecutive measurements. Response stability was monitored over 7 h, and storage stability was monitored after storage for 2 weeks at 4℃.
RESULTS AND DISCUSSION EB System And Measurements To generate the EB system, we transformed cells with pLZCapR (Fig. 1A), immobilized them in PVA to increase their stability and long-term use (Fig. 1B), and examined their
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bioreporting activity (i.e., β-galactosidase activity) by CV (Fig. 1C). Typical cyclic voltammograms obtained from PBS buffer alone (solid line) and PBS buffer containing EB cells treated with (dashed line) or without (dotted line) phenolic compounds are presented in Figure 1C. The PBS buffer alone did not significantly contribute to the magnitude of the signal, indicating it did not interfere with the redox responses of PAP. Consistent with a previous report[23], EB-cell-free PAPG also failed to show any current
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response (data not shown). After hydrolysis of PAPG by β-galactosidase, phenol-induced EB cells (Fig. 1C, dashed line) showed peaks that were increased and shifted to approximately +220 mV and +120 mV due to the oxidation of PAP. This is in good agreement with the findings of other groups[4,23], although it is known that oxidation and reduction potentials can vary depending on the scan rate and experimental conditions.[10] Biran et al. (2000) previously performed electrochemical bioreporting of cadmium by monitoring electrochemical β-galactosidase activity with PAPG as the substrate, using disposable graphite working electrodes (+220 mV) and Ag/AgCl ink in the reference electrodes.[4] Matsui et al. (2006) reported that the oxidation currents for PAP and PAPG are first observed at 0.0 and 0.3 V versus Ag/AgCl, respectively, with the magnitude of the PAP current reaching approximately 2~3 times that of PAPG; they did not find any other electroactive species around this potential.[23] In the present work, we found that non-induced EB cells treated with PAPG (Fig. 1C, dotted line) showed redox peaks at approximately +200 mV and +100 mV; this was taken as the blank test for each experimental day. This could reflect some “leaky” production of ß-galactosidase even in the absence of phenol, as the utilized promoter is not completely silent.[24] Furthermore, bacteria can reportedly transfer electrons to an electrode via various bacterially produced
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redox mediators or direct electron transfer by cytochrome-rich membranes.[25]
The
potential values of PAP redox (+220 mV and +120 mV) are relatively close to, but distinct from, those of PAPG (+200 mV and +100 mV). Therefore, we designated redox peaks with signal:blank ratios of ≥ 1.45 at approximately +220 mV and +120 mV as reflecting data on PAP redox responses.
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Sensitivity In the EB system, PAP is oxidized at the Pt electrode, where it can be directly converted to a current signal without needing time to accumulate. Thus, we hypothesized that this system would be more sensitive and rapid than the conventional OB measurements. We assessed EB measurements at various concentrations (10 nM~10 mM) of phenol (Fig. 2). The current responses appeared at 10 nM phenol, peaked at 0.05 mM (occasionally 0.1 mM) phenol, and decreased thereafter (Fig. 2). We tested 1 nM and 5 nM phenol, but the signals were not sufficiently different from those obtained from the blank (signal:blank ratio ≤ 1.45), and the data were therefore excluded. When we used the equation 3S/m (three times standard deviation/slope of regression curve) to calculate the lower detection limit for EB, we obtained a lower detection limit of 30 nM. Our calibration plot for the electrochemical measurement of EB cells showed relatively good linearity with regard to phenol concentrations from 0.001 to 0.1 mM (logarithmic regression curve; y = 5.06 log x + 58.1; R2 = 0.9).
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The EB system showed a maximum response at approximately 0.05 mM of phenol, whereas the OB systems showed their maximum responses at approximately 1 mM.[17-19] The result suggests that the EB system is more responsive to lower concentrations of phenol than the previously reported OB systems.[17-19] The intensities of peak signals compared with minimum detectable signals for EB were approximately 1-2 fold and were very similar to OB based intact cells. [18,19] Our findings are comparable to those of Biran
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et al. (2000), who found that, under optimum conditions, their amperometric system had lower limits of detection of 50 nM Cd2+ after 30 min and 25 nM Cd2+ after 1 h, and that the current signal decreased at concentrations higher than 10 μM Cd2+ due to metal toxicity.[4] Notably, our PVA-immobilized EB cells showed high levels of phenol tolerance (~10 mM) and operational stability compared with non-immobilized bioreporter cells.[17] This is consistent with the findings of several previous reports.[26-28] Given that luciferase systems are generally more sensitive than β-galactosidase systems[18,29], it is rather remarkable that the EB system achieved a nano-level lower limit of detection. We speculate that the high sensitivity of the EB system may reflect our use of an electrochemical sensing system.
Specificity The specificity of a CapR-controlled bioreporter may be determined by the abilities of the phenolic compounds to activate the binding site of CapR[17], so we hypothesized that all of the CapR-controlled bioreporters would have similar specificities. To test this, we examined the specificity of EB cells against different phenolic compounds and compared with data previously reported for the OBs.[17,18] As expected, similar current responses
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were observed with various phenol-like compounds (unbranched or single-branched). When EB cells harboring pLZCapR were treated with various phenolic compounds (0.1 mM), significant responses were obtained against phenol, 2-chlorophenol, 2methylphenol, 2-nitrophenol, 3-methylphenol, 4-chlorophenol, catechol, and resorcinol, but not 2,5-dimethylphenol (data not shown). We then assessed the detection ranges of the EB cells against various concentrations (from 1 nM to 1 mM) of the phenolic
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compounds that had yielded current responses at 0.1 mM (Fig. 3). The detected concentration ranges varied; catechol, 2-chlorophenol, 3-methylphenol, 2-methylphenol and resorcinol yielded responses in the range of 1~10 nM to 1 mM, whereas 4chlorophenol and 2-nitrophenol yielded responses in the range of 10~100 nM to 1 mM. In terms of maximum responses, catechol, 2-nitrophenol and resorcinol yielded their maximum responses at 0.1 mM, while 2-chlorophenol, 4-chlorophenol and 3methylphenol yielded their maximum responses at 0.01 mM. Interestingly, the response to 2-methylphenol increased dose-dependently up to 1 mM (Fig. 3).
However, the intensities of the responses differed somewhat across the systems: resorcinol and 4-methylphenol were detected by the EB system; OB-spec responded only weakly to 4-chlorophenol and yielded no detectable response to resorcinol[18]; and the EB and OB-spec all failed to detect 2,5-dimethyphenol. Interestingly, the intensities of the responses to various phenolic compounds also varied depending on the reporter type and the state of the biosensor. Resorcinol, 4-methylphenol, and 2,5-dimethyphenol were only weakly detected by OB-lum harboring pCapR[17]; resorcinol was not detected by freezedried cells or free biosensors harboring pLZCapR (OB-spec); and catechol was strongly
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detected by the free cells (OB-lum), but only weakly by the freeze-dried (OB-spec) and immobilized cells (EB and OB-spec).[17,18] We do not yet know whether the responseinducing phenolic compounds act as effectors, or if they are actually substrates for a catabolic enzyme of the CapR-controlled operon. We also do not know which structures are responsible for the activity changes that translate to the bioreporting activity of CapRcontrolled systems. Additional research is therefore warranted to examine the structural
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and functional interactions of CapR with its inducer(s).
We obtained detection ranges against various phenolic compounds in the nM~mM range, whereas those of the OB systems were generally higher, in the μM~mM range.[17-19] Therefore, the detection ranges of the EB system against phenolic compounds were relatively lower than those previously reported for the OB systems.[17-19]
Stability, Repeatability, And Application The long-term use and reliable operation of bioreporters requires that they have good stability and repeatability. Figure 4A shows that the EB system had relatively good operational stability for at least 7 h. In terms of storage stability, which is also important, the PVA-immobilized EB cells retained 80~100% of their initial current responses after 2 weeks of storage at 4℃ (data not shown). These results are consistent with the previous report showing that PVA-immobilized microbial cells can be stored for long durations (up to 100 days) without large losses of activity.[8] To assess the degree of precision achieved by the EB system, we tested the repeatability by monitoring six consecutive uses of the system to assess phenol. We obtained a relatively good standard deviation (± 2.7%, n = 6).
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Together, these findings show that the EB system showed acceptable operational stability and retained its activity during storage.
To further validate the EB system, we simulated the occurrence of phenol contamination in wastewater. Immobilized EB cells were exposed to buffer (control) or untreated hospital wastewater spiked with 0.1 mM phenol (spiking was performed because the
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hospital wastewater samples obtained from several sources in Busan City, South Korea, were free of phenolic contamination). The EB cells showed similar or slightly lower responses to phenol-spiked hospital wastewater compared to phenol-spiked buffer (Fig. 4B), suggesting that impurities in the field samples might have interfered with the current responses of the EB system. Bioreporters may be affected by the toxicity of the environment and the bioavailability of the target.[30] Thus, further instrumental analysis may be needed to examine the detailed compositions and concentrations of phenolic compounds from field samples. However, our present results suggest that PVAimmobilized EB cells may prove useful for the rapid preliminary detection of phenolic compounds in field samples.
Comparative Evaluation Table 1 summarizes our comparison of the EB and OB systems, which included free, freeze-dried, and immobilized (in PVA or agarose) cells. β-Galactosidase activities were observed in response to concentration ranges of approximately 10 nM to 10 mM in the EB system and 0.1~10 μM to 10~100 mM in the OB systems. The lower detection limit of the EB system was 1~3 orders of magnitude lower than those of the OB systems.
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Overall, the EB cells showed good sensitivity against lower concentrations of phenol, whereas agarose-immobilized OB cells showed good sensitivity against higher concentrations of phenol (Table 1). Bioreporter cells immobilized in PVA or agarose showed higher tolerances for toxic concentrations of phenol (~100 mM phenol) compared to free OB-lum cells (~1 mM phenol), supporting the notion that immobilization might protect bioreporter cells against environmental toxicity.[26-28,31] The
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biosensor cells were lysed for use in the OB-lum system, suggesting that the presence of an intact bioreporter cell membrane might help guard against environmental-toxicityinduced enzymatic inactivation or cellular dysfunction.[16] Bioreporter cells in the freezedried state may be conveniently stored for months without loss of activity.[32,33] The PVA beads utilized in our EB system dispersed freely in the reaction solution and allowed the free electrons generated from PIQ (the final product of PAPG) to quickly access the electrodes for current generation. This merit of the EB system was reflected in a much shorter detection time (15-20 min) compared with the OB systems (1~6 h), which require time for substrate reaction/accumulation, addition of the stop solution, and measurement of the bulk concentration change (Table 1).
CONCLUSIONS The responses of microbial bioreporters may be visualized with several types of transducers. To choose a suitable transducer for a given use, researchers may need to compare the analytical ability of bioreporters that pair the same regulatory circuit with different transducers. Here, we developed an amperometer-based EB for the detection of phenolic compounds, and compared its analytical capabilities with those of comparable
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luminometry-[17] and spectrophotometry-[18,19] based OB systems. E. coli DH5α cells were transformed with pLZCapR and immobilized in PVA beads, and the abilities of these cells to sense phenolic compounds were assessed by electrochemical measurement of β-galactosidase activity. The EB system showed a relatively low detection range (10 nM to 10 mM phenol) compared to the OB systems (0.1-10 μM to 10-100 mM phenol).[17-19] Furthermore, the detection time required by the EB system (15-20 min) was
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clearly shorter than that required by the OB systems (1-6 h). We herein show that the novel EB system offers several advantages over the previously described OB systems, including more rapid processing and a higher sensitivity to phenolic compounds at low (nanomolar) concentration ranges.
ACKNOWLEDGEMENTS This work was supported by the Basic Science Research Program of the National Research Foundation of Korea (NRF) funded by the Ministry of Education, Science and Technology (2011-0007702), by a grant (2013) from the Innovation Center for Engineering Education of Dongseo University, and by a grant (2014) from Dongseo University, Republic of Korea. The authors thank Jin Woo Lee, Su Min Yeom, So Hee Kang, Jeong Ha Park, and Chang Bin Shon for their help in performing the research.
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29. Keane, A.; Phoenix, P.; Ghoshal, S.; Lau, P.C.K. Exposing culprit organic pollutants: a review. J. Microbiol. Meth. 2002, 49, 103-119. 30. Kelsey, J.W.; Kottler, B.D.; Alexander, M. Selective chemical extractants to predict bioavailability of soil-aged organic chemicals. Environ. Sci. Technol. 1997, 31, 214–217. 31. Liu, L.; Zhai, J.; Zhu, C.; Gao, Y.; Wang, Y.; Han, Y.; Dong, S. One-pot synthesis of 3-dimensional reduced grapheme oxide-based hydrogel as support for microbe immobilization and BOD biosensor preparation. Biosens. Bioelectron. 2015, 63, 483-489. 32. Leslie, S.B.; Israeli, E.; Lighthart, B.; Crowe, J.H.; Crowe, I.M. Trehalose and sucrose protect both membranes and proteins in intact bacteria during drying. Appl. Environ. Microbiol. 1995, 61, 3592–3597. 33. Perry, S.F. Methods in Molecular Biology: Cryopreservation and freeze-drying protocols. Human Press, Totowa, 1995.
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Table 1. Comparing the abilities of various biosensors to detect phenol Reporter
β-
Transducer
Amperometer
Immobilization Detection
PVA
Galactosidase Firefly
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Reference
range
time
10 nM-10
15-20 min
This study
1h
[17]
5-6 h
[18]
5-6 h
[19]
mM Luminometer
10 μM -1
None
luciferase
mM
β-
Spectrophotomete
Galactosidase
r
β-
Spectrophotomete
Galactosidase
r
a
Detectiona
Freeze-dry
0.1 μM-10 mM
Agarose
10 μM-100 mM
Detection time included the processing time but excluded the induction time.
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FIGURE 1. EB system and measurements. E. coli DH5α cells harboring pLZCapR were constructed, immobilized in PVA and treated with PAPG for 15 min, and their βgalactosidase activities were assessed by CV, as described in the Materials and Methods section. (A) In the presence of phenolic compounds, β-galactosidase is expressed and hydrolyzes PAPG to PAP, which is further oxidized to PIQ at the Pt working electrode, generating a current. (B) SEM images of PVA control and cell-immobilized PVA were
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taken at 6,000 x. (C) Cyclic voltammograms of 50 mM PBS buffer (solid line), and EB cells treated with (dashed line) or without phenol (0.1 mM; dotted line) in 50 mM PBS buffer plus 15 mM PAPG. Scan rate, 100 mV s-1.
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FIGURE 2. EB cells respond to various concentrations of phenol. EB cells were prepared as described in the Materials and Methods section. Immobilized cells (approximately 4.2 x 109 CFU/ml) were treated with 10 nM to 10 mM phenol for 2 h and then exposed to 15 mM PAPG for 15 min, and CV was used to assess β-galactosidase activity at a scan rate of 100 mV s-1. The peak currents from each voltammogram were plotted as a function of the phenol concentration. The regression curve (y = 5.06 log x + 58.1; R2 = 0.9) is shown
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in the inset; linearity is seen between 0.001 and 0.1 mM phenol.
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FIGURE 3. The response of EB cells to various concentrations of phenolic compounds. EB cells were incubated for 2 h in the presence of various concentrations (1 nM to 1 mM) of phenolic compounds, and β-galactosidase activity was assessed by CV at a scan rate of
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100 mV s-1.
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FIGURE 4. Operational stability of the EB system and its use to assess phenol in wastewater. (A) EB cells were treated with 0.1 mM phenol for 2 h, and β-galactosidase activity was assessed hourly for 7 h, using CV at a scan rate of 100 mV s-1. (B) EB cells were exposed to buffer or wastewater from several sources (a~f) in Busan City spiked with 0.1 mM phenol, and β-galactosidase activity was measured by CV at a scan rate of
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100 mV s-1.
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ISSN: 1082-6068 (Print) 1532-2297 (Online) Journal homepage: http://www.tandfonline.com/loi/lpbb20
Comparative Evaluation of an Electrochemical Bioreporter for Detecting Phenolic Compounds Hae Ja Shin & Woon Ki Lim To cite this article: Hae Ja Shin & Woon Ki Lim (2014): Comparative Evaluation of an Electrochemical Bioreporter for Detecting Phenolic Compounds, Preparative Biochemistry and Biotechnology, DOI: 10.1080/10826068.2014.979207 To link to this article: http://dx.doi.org/10.1080/10826068.2014.979207
Accepted author version posted online: 30 Oct 2014.
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Date: 05 November 2015, At: 16:40
Comparative Evaluation of an Electrochemical Bioreporter for Detecting Phenolic Compounds Hae Ja Shin1, Woon Ki Lim2 1
Division of Energy and Bio-engineering, Dongseo University, Busan, Republic of Korea, 2 Department of Molecular Biology, College of Natural Sciences, Pusan National University, Busan, Republic of Korea
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Corresponding author Hae Ja Shin, E-mail: [email protected]
Abstract In the present study, we constructed an Escherichia coli-based electrochemical bioreporter (EB) harboring pLZCapR, which encodes the CapR regulatory protein (for phenol degradation) along with β-galactosidase, and examined its ability to detect phenolic compounds as compared with previously reported optical bioreporters (OBs) controlled by CapR and detected using a luminometer (OB-lum) or spectrophotometer (OB-spec). The recombinant E. coli bioreporter cells were immobilized in polyvinyl alcohol (PVA); p-aminophenyl-β-D-galactopyranoside (PAPG) was used as the enzymatic substrate; and electrochemical measurements were taken. The peak current obtained on cyclic voltammetry (CV) was used to measure the redox response of PAPG degradation. Our results revealed that the EB system showed a detection range of 10 nM to 10 mM phenol with a good lower detection limit (30 nM phenol). Furthermore, the detection time was dramatically lower for the EB system (15-20 min) compared to the OBs (~6 h). These responses were reliably repeatable with an acceptable standard deviation (± 2.7%; n = 6), and the system showed good stability without loss of activity over 7 h of operation or following 2 weeks of cold storage. Together, these results show
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that the EB system is faster and has a lower detection limit than the existing optical techniques.
KEYWORDS: Electrochemical Escherichia coli bioreporter, Phenolic compounds
INTRODUCTION
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Microbial bioreporters have been used as first-step monitoring tools to detect target chemicals and physiological signals in diverse fields.[1] Numerous recombinant microbial bioreporters have been constructed by placing a reporter gene (i.e., luc, lacZ or gfp) under the control of regulatory genes that are responsive to the target chemicals and signals.[2] The biological signals from these cellular components are dose-dependent and can generate measurable responses via various transducers. The transducers that have been exploited in the construction and monitoring of microbial bioreporters include electrochemical transducers, which can convert biological signals to electrochemical signals (e.g., currents, potentials and conductivities). Among the electrochemical transducers, amperometers have been widely used to visualize signal responses to target chemicals and physiological signals, such as biochemical oxygen demand (BOD)[3], heavy metals[4], and organophosphates.[5,6]
Potentiometers have also been exploited,
such as for detecting organophosphates with pH electrodes[7], sucrose and BOD with oxygen electrodes[8], and urea with NH4+ ion-selective electrodes.[9] Electrochemical transducers may be cost-effective, portable and miniaturized[10], although some systems require electroactive mediators[11], such as anthraquinone[12] or ferricyanide[13], to improve their electrochemical responses. Overall, the electrochemical microbial
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bioreporters typically show acceptable sensitivity and stability, but their selectivity tends to be relatively poor.[14] In terms of optical transducers, luminometers, spectrophotometers, fluorometers, and flow cytometer are commonly used in microbial bioreporter systems. The luminometer is a sensitive, rapid, and portable transducer that may be monitored on-line through optic fibers. Luminescence-based systems are common, but their sensitivity can be decreased by light scattering, such as in turbid
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solutions and under anaerobic conditions.[4] Spectrophotometers and fluorometers are widely used to take sensitive measurements in ordinary laboratories, but they are unwieldy and impractical for on-site measurements. Currently, the flow cytometer has been used to monitor the inner cell information and its kinetic behaviors at individual cell level. [15] To date, no study has made a direct comparison of transducers, even though this could help researchers choose the one most suitable for their purposes in monitoring target chemicals.
In our previous work, we have extensively studied microbial bioreporters that used optical transducers (e.g., luminometers and spectrophotometers) to sense benzene, toluene, ethylbenzene and xylene (BTEX)[16], phenolic compounds[17-19], and salicylate.[20-22] We found that these microbial bioreporters could quantitatively detect aromatic compounds at relatively high concentration ranges (i.e., μM~mM) in ~6 h.[18,20,21]
However, such tests should be speed up for field use. Furthermore,
environmental contamination can involve lower concentrations of aromatic hydrocarbons, necessitating more sensitive detection methods for the monitoring of such compounds.
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Accordingly, it would be useful to compare the analytical performance of bioreporters that use different transducers to detect aromatic compounds.
In the present study, we constructed an amperometer-based electrochemical bioreporter (EB) for the monitoring of phenolic compounds, and compared its ability with those of previously described optical bioreporters (OBs) that detected such compounds by
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luminometry (OB-lum) and spectrophotometry (OB-spec). All of the studied bioreporters were under the control of the CapR regulatory protein, which directs phenol degradation. For the construction of EB, recombinant E. coli cells harboring the plasmid pLZCapR (capR::lacZ) were immobilized in polyvinyl alcohol (PVA). For electrochemical detection of phenolic compounds, p-aminophenyl-β-D-galactopyranoside (PAPG; a substrate of β-galactosidase) was added, and the oxidation of p-aminophenol (PAP; a catabolite of PAPG) was monitored by cyclic voltammetry (CV). This EB system was validated, characterized, and compared with OBs.
EXPERIMENTAL Chemicals The media components were purchased from Difco (MO, USA). The PAPG, PVA, and boric acid were purchased from Sigma (MO, USA). The genomic DNA isolation kit, plasmid isolation spin kit, and gel extraction kit were purchased from Q-BIO Gene (CA, USA), Qiagen (Hilden, Germany), and Cosmogenetech (Seoul, Korea), respectively. AgeI, KpnI, SalI, HindIII, and T4-DNA ligase were purchased from Takara (Shiga, Japan). The pSV-beta-gal plasmid and pGL3b basic vector were purchased from Promega (WI, USA).
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All other utilized reagents were of analytical grade and were used as received without further purification.
Bacteria And Culture Conditions Escherichia coli DH5α (hsdR, recA, Thi-1, relA1, gyrA96) was used to maintain plasmids. P. putida KCTC1452 was obtained from the Korean Collection for Type Cultures (KCTC)
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and grown in tryptic soy broth at 25°C. E. coli DH5α cells harboring plasmids were cultured at 37°C in TYS media (1% tryptone, 0.5% yeast extract, and 0.5% NaCl) containing 30 μg ml-1 ampicillin. The number of colony forming units (CFU) per ml of culture sample was determined by serially diluting the sample and plating the cells on Luria-Bertani agar.
Construction Of Plasmid Plzcapr The capR gene, its own promoter (Pr), and the Po promoter were cloned from the genomic DNA of P. putida KCTC1452, as described previously.[17] Briefly, the capR/Pr/Po fragment was PCR amplified with specific primers that each harbored KpnI sites at their ends (forward, 5´-CCC AAG CTT GGT ACC GCT TAC ATG CCG ATC AAG TAC-3´ and reverse, 5´-CGG GGT ACC AAG CTT TAA CGA GTG AGC TGA TCG AAA-3´). The PCR products were gel-isolated and digested with KpnI. A HindIII/SalI restriction fragment from pGL3 basic and a 3749-bp HindIII/SalI restriction fragment from pSV-beta-gal were ligated with T4-DNA ligase. The resulting 6610-bp pGLβ-GAL fragment was digested with HindIII/AgeI to remove the gpt promoter from pSV-beta-gal; the fragment was then blunt-ended and ligated to generate the promoterless
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pGLβ-GAL. Finally, a fragment of KpnI-digested capR/Pr/Po was inserted at the KpnI site of the 6392-bp pGLβ-GAL to construct the 8538-bp pLZCapR (Fig. 1). This plasmid was transformed into E. coli DH5α by the CaCl2 method. Immobilization Of Bacterial Cells E. coli cells harboring pLZCapR were grown for 16 h at 37℃ with shaking, and then harvested by centrifugation at 10,000 rpm for 15 min. The supernatant was discarded and
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the cells were washed with 50 mM PBS buffer (pH 7.4). To optimize the immobilization of these cells in PVA beads, various amounts (0.3, 0.5, 0.7, 0.9 or 1 gram) of PVA (nominal degree of polymerization = 1750; approx. molecular weight 75,000-80,000) were solubilized in 5 ml of distilled hot water, dissolved completely in a boiling water bath, and then cooled to 40℃. An equal volume of cells (approximately 4.2 x 109 CFU/ml) was mixed with the dissolved PVA solution, and beads were formed by dropping the mixture into a saturated boric acid solution. The resulting beads were soaked in 0.5 M sodium orthophosphate for 1 h, washed with saline, and then either directly subjected to the electrochemical assay for β-galactosidase activity (see below), or stored at 4℃ until use. Proper immobilization was confirmed by scanning electron microscopy (SEM) of control PVA and cell-immobilized PVA (magnification 6000 x).
Apparatus And Procedure For Electrochemical Measurement Of ß-Galactosidase Activity CV measurements were performed using a potentiostat/galvanostat (Cosentech, Korea). All operations, including data acquisition, were completely controlled by the Physiolab kst-p1 software (Cosentech, Korea). The three-electrode system comprised a platinum (Pt)
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working electrode (diameter, 3 mm; working area, 0.071 cm2), an Ag/AgCl reference electrode, and a Pt auxiliary electrode. Microbial cells were immobilized and loaded into the workstation, and β-galactosidase activity was measured using PAPG as the substrate. The three electrodes were placed in contact with the PVA-immobilized bacterial cells in the wells of a 24-well plate containing appropriate volumes of test solution. For CV measurements, bacterial cells that had been treated with or without phenolic compounds
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were placed in PBS buffer containing 15 mM PAPG for 15 min, and then cyclic voltammograms were recorded (cyclic scan range, -100 mV to +500 mV; scan rate, 100 mV s-1). All experiments were carried out at room temperature, and were assessed under the optimum pH (7.4), amount of cell loading (~ 4.2 x 109 CFU/ml), induction time (2 h), and concentrations of phenolic compounds. To examine selectivity, we assessed the biosensor activities obtained in response to 0.1 mM of 2-chlorophenol, 2-methylphenol, 2-nitrophenol, 3-methylphenol, 4-chlorophenol, catechol, resorcinol, and 2,5dimethylphenol. These phenolic compounds were dissolved in ethanol and spiked into either 50 mM PBS buffer (pH 7.4) or untreated hospital wastewater from several sources in Busan City. To test response repeatability, we examined the standard deviation from six consecutive measurements. Response stability was monitored over 7 h, and storage stability was monitored after storage for 2 weeks at 4℃.
RESULTS AND DISCUSSION EB System And Measurements To generate the EB system, we transformed cells with pLZCapR (Fig. 1A), immobilized them in PVA to increase their stability and long-term use (Fig. 1B), and examined their
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bioreporting activity (i.e., β-galactosidase activity) by CV (Fig. 1C). Typical cyclic voltammograms obtained from PBS buffer alone (solid line) and PBS buffer containing EB cells treated with (dashed line) or without (dotted line) phenolic compounds are presented in Figure 1C. The PBS buffer alone did not significantly contribute to the magnitude of the signal, indicating it did not interfere with the redox responses of PAP. Consistent with a previous report[23], EB-cell-free PAPG also failed to show any current
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response (data not shown). After hydrolysis of PAPG by β-galactosidase, phenol-induced EB cells (Fig. 1C, dashed line) showed peaks that were increased and shifted to approximately +220 mV and +120 mV due to the oxidation of PAP. This is in good agreement with the findings of other groups[4,23], although it is known that oxidation and reduction potentials can vary depending on the scan rate and experimental conditions.[10] Biran et al. (2000) previously performed electrochemical bioreporting of cadmium by monitoring electrochemical β-galactosidase activity with PAPG as the substrate, using disposable graphite working electrodes (+220 mV) and Ag/AgCl ink in the reference electrodes.[4] Matsui et al. (2006) reported that the oxidation currents for PAP and PAPG are first observed at 0.0 and 0.3 V versus Ag/AgCl, respectively, with the magnitude of the PAP current reaching approximately 2~3 times that of PAPG; they did not find any other electroactive species around this potential.[23] In the present work, we found that non-induced EB cells treated with PAPG (Fig. 1C, dotted line) showed redox peaks at approximately +200 mV and +100 mV; this was taken as the blank test for each experimental day. This could reflect some “leaky” production of ß-galactosidase even in the absence of phenol, as the utilized promoter is not completely silent.[24] Furthermore, bacteria can reportedly transfer electrons to an electrode via various bacterially produced
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redox mediators or direct electron transfer by cytochrome-rich membranes.[25]
The
potential values of PAP redox (+220 mV and +120 mV) are relatively close to, but distinct from, those of PAPG (+200 mV and +100 mV). Therefore, we designated redox peaks with signal:blank ratios of ≥ 1.45 at approximately +220 mV and +120 mV as reflecting data on PAP redox responses.
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Sensitivity In the EB system, PAP is oxidized at the Pt electrode, where it can be directly converted to a current signal without needing time to accumulate. Thus, we hypothesized that this system would be more sensitive and rapid than the conventional OB measurements. We assessed EB measurements at various concentrations (10 nM~10 mM) of phenol (Fig. 2). The current responses appeared at 10 nM phenol, peaked at 0.05 mM (occasionally 0.1 mM) phenol, and decreased thereafter (Fig. 2). We tested 1 nM and 5 nM phenol, but the signals were not sufficiently different from those obtained from the blank (signal:blank ratio ≤ 1.45), and the data were therefore excluded. When we used the equation 3S/m (three times standard deviation/slope of regression curve) to calculate the lower detection limit for EB, we obtained a lower detection limit of 30 nM. Our calibration plot for the electrochemical measurement of EB cells showed relatively good linearity with regard to phenol concentrations from 0.001 to 0.1 mM (logarithmic regression curve; y = 5.06 log x + 58.1; R2 = 0.9).
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The EB system showed a maximum response at approximately 0.05 mM of phenol, whereas the OB systems showed their maximum responses at approximately 1 mM.[17-19] The result suggests that the EB system is more responsive to lower concentrations of phenol than the previously reported OB systems.[17-19] The intensities of peak signals compared with minimum detectable signals for EB were approximately 1-2 fold and were very similar to OB based intact cells. [18,19] Our findings are comparable to those of Biran
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et al. (2000), who found that, under optimum conditions, their amperometric system had lower limits of detection of 50 nM Cd2+ after 30 min and 25 nM Cd2+ after 1 h, and that the current signal decreased at concentrations higher than 10 μM Cd2+ due to metal toxicity.[4] Notably, our PVA-immobilized EB cells showed high levels of phenol tolerance (~10 mM) and operational stability compared with non-immobilized bioreporter cells.[17] This is consistent with the findings of several previous reports.[26-28] Given that luciferase systems are generally more sensitive than β-galactosidase systems[18,29], it is rather remarkable that the EB system achieved a nano-level lower limit of detection. We speculate that the high sensitivity of the EB system may reflect our use of an electrochemical sensing system.
Specificity The specificity of a CapR-controlled bioreporter may be determined by the abilities of the phenolic compounds to activate the binding site of CapR[17], so we hypothesized that all of the CapR-controlled bioreporters would have similar specificities. To test this, we examined the specificity of EB cells against different phenolic compounds and compared with data previously reported for the OBs.[17,18] As expected, similar current responses
10
were observed with various phenol-like compounds (unbranched or single-branched). When EB cells harboring pLZCapR were treated with various phenolic compounds (0.1 mM), significant responses were obtained against phenol, 2-chlorophenol, 2methylphenol, 2-nitrophenol, 3-methylphenol, 4-chlorophenol, catechol, and resorcinol, but not 2,5-dimethylphenol (data not shown). We then assessed the detection ranges of the EB cells against various concentrations (from 1 nM to 1 mM) of the phenolic
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compounds that had yielded current responses at 0.1 mM (Fig. 3). The detected concentration ranges varied; catechol, 2-chlorophenol, 3-methylphenol, 2-methylphenol and resorcinol yielded responses in the range of 1~10 nM to 1 mM, whereas 4chlorophenol and 2-nitrophenol yielded responses in the range of 10~100 nM to 1 mM. In terms of maximum responses, catechol, 2-nitrophenol and resorcinol yielded their maximum responses at 0.1 mM, while 2-chlorophenol, 4-chlorophenol and 3methylphenol yielded their maximum responses at 0.01 mM. Interestingly, the response to 2-methylphenol increased dose-dependently up to 1 mM (Fig. 3).
However, the intensities of the responses differed somewhat across the systems: resorcinol and 4-methylphenol were detected by the EB system; OB-spec responded only weakly to 4-chlorophenol and yielded no detectable response to resorcinol[18]; and the EB and OB-spec all failed to detect 2,5-dimethyphenol. Interestingly, the intensities of the responses to various phenolic compounds also varied depending on the reporter type and the state of the biosensor. Resorcinol, 4-methylphenol, and 2,5-dimethyphenol were only weakly detected by OB-lum harboring pCapR[17]; resorcinol was not detected by freezedried cells or free biosensors harboring pLZCapR (OB-spec); and catechol was strongly
11
detected by the free cells (OB-lum), but only weakly by the freeze-dried (OB-spec) and immobilized cells (EB and OB-spec).[17,18] We do not yet know whether the responseinducing phenolic compounds act as effectors, or if they are actually substrates for a catabolic enzyme of the CapR-controlled operon. We also do not know which structures are responsible for the activity changes that translate to the bioreporting activity of CapRcontrolled systems. Additional research is therefore warranted to examine the structural
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and functional interactions of CapR with its inducer(s).
We obtained detection ranges against various phenolic compounds in the nM~mM range, whereas those of the OB systems were generally higher, in the μM~mM range.[17-19] Therefore, the detection ranges of the EB system against phenolic compounds were relatively lower than those previously reported for the OB systems.[17-19]
Stability, Repeatability, And Application The long-term use and reliable operation of bioreporters requires that they have good stability and repeatability. Figure 4A shows that the EB system had relatively good operational stability for at least 7 h. In terms of storage stability, which is also important, the PVA-immobilized EB cells retained 80~100% of their initial current responses after 2 weeks of storage at 4℃ (data not shown). These results are consistent with the previous report showing that PVA-immobilized microbial cells can be stored for long durations (up to 100 days) without large losses of activity.[8] To assess the degree of precision achieved by the EB system, we tested the repeatability by monitoring six consecutive uses of the system to assess phenol. We obtained a relatively good standard deviation (± 2.7%, n = 6).
12
Together, these findings show that the EB system showed acceptable operational stability and retained its activity during storage.
To further validate the EB system, we simulated the occurrence of phenol contamination in wastewater. Immobilized EB cells were exposed to buffer (control) or untreated hospital wastewater spiked with 0.1 mM phenol (spiking was performed because the
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hospital wastewater samples obtained from several sources in Busan City, South Korea, were free of phenolic contamination). The EB cells showed similar or slightly lower responses to phenol-spiked hospital wastewater compared to phenol-spiked buffer (Fig. 4B), suggesting that impurities in the field samples might have interfered with the current responses of the EB system. Bioreporters may be affected by the toxicity of the environment and the bioavailability of the target.[30] Thus, further instrumental analysis may be needed to examine the detailed compositions and concentrations of phenolic compounds from field samples. However, our present results suggest that PVAimmobilized EB cells may prove useful for the rapid preliminary detection of phenolic compounds in field samples.
Comparative Evaluation Table 1 summarizes our comparison of the EB and OB systems, which included free, freeze-dried, and immobilized (in PVA or agarose) cells. β-Galactosidase activities were observed in response to concentration ranges of approximately 10 nM to 10 mM in the EB system and 0.1~10 μM to 10~100 mM in the OB systems. The lower detection limit of the EB system was 1~3 orders of magnitude lower than those of the OB systems.
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Overall, the EB cells showed good sensitivity against lower concentrations of phenol, whereas agarose-immobilized OB cells showed good sensitivity against higher concentrations of phenol (Table 1). Bioreporter cells immobilized in PVA or agarose showed higher tolerances for toxic concentrations of phenol (~100 mM phenol) compared to free OB-lum cells (~1 mM phenol), supporting the notion that immobilization might protect bioreporter cells against environmental toxicity.[26-28,31] The
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biosensor cells were lysed for use in the OB-lum system, suggesting that the presence of an intact bioreporter cell membrane might help guard against environmental-toxicityinduced enzymatic inactivation or cellular dysfunction.[16] Bioreporter cells in the freezedried state may be conveniently stored for months without loss of activity.[32,33] The PVA beads utilized in our EB system dispersed freely in the reaction solution and allowed the free electrons generated from PIQ (the final product of PAPG) to quickly access the electrodes for current generation. This merit of the EB system was reflected in a much shorter detection time (15-20 min) compared with the OB systems (1~6 h), which require time for substrate reaction/accumulation, addition of the stop solution, and measurement of the bulk concentration change (Table 1).
CONCLUSIONS The responses of microbial bioreporters may be visualized with several types of transducers. To choose a suitable transducer for a given use, researchers may need to compare the analytical ability of bioreporters that pair the same regulatory circuit with different transducers. Here, we developed an amperometer-based EB for the detection of phenolic compounds, and compared its analytical capabilities with those of comparable
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luminometry-[17] and spectrophotometry-[18,19] based OB systems. E. coli DH5α cells were transformed with pLZCapR and immobilized in PVA beads, and the abilities of these cells to sense phenolic compounds were assessed by electrochemical measurement of β-galactosidase activity. The EB system showed a relatively low detection range (10 nM to 10 mM phenol) compared to the OB systems (0.1-10 μM to 10-100 mM phenol).[17-19] Furthermore, the detection time required by the EB system (15-20 min) was
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clearly shorter than that required by the OB systems (1-6 h). We herein show that the novel EB system offers several advantages over the previously described OB systems, including more rapid processing and a higher sensitivity to phenolic compounds at low (nanomolar) concentration ranges.
ACKNOWLEDGEMENTS This work was supported by the Basic Science Research Program of the National Research Foundation of Korea (NRF) funded by the Ministry of Education, Science and Technology (2011-0007702), by a grant (2013) from the Innovation Center for Engineering Education of Dongseo University, and by a grant (2014) from Dongseo University, Republic of Korea. The authors thank Jin Woo Lee, Su Min Yeom, So Hee Kang, Jeong Ha Park, and Chang Bin Shon for their help in performing the research.
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2. Shin, H.J. Genetically engineered microbial biosensors for in situ monitoring of environmental pollution. Appl. Microbiol. Biotechnol. 2011, 89, 867-877. 3. Jia, J.; Tang, M.; Chen, X.; Qi, L.; Dong, S. Co-immobilized microbial biosensor for BOD estimation based on Sol-gel derived composite material. Biosens. Bioelectron. 2003, 18, 1023–1029. 4. Biran, I.; Babai, R.; Levcov, K.; Rishpon, J.; Ron, E. Online and in sutu monitoring of
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Table 1. Comparing the abilities of various biosensors to detect phenol Reporter
β-
Transducer
Amperometer
Immobilization Detection
PVA
Galactosidase Firefly
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Reference
range
time
10 nM-10
15-20 min
This study
1h
[17]
5-6 h
[18]
5-6 h
[19]
mM Luminometer
10 μM -1
None
luciferase
mM
β-
Spectrophotomete
Galactosidase
r
β-
Spectrophotomete
Galactosidase
r
a
Detectiona
Freeze-dry
0.1 μM-10 mM
Agarose
10 μM-100 mM
Detection time included the processing time but excluded the induction time.
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FIGURE 1. EB system and measurements. E. coli DH5α cells harboring pLZCapR were constructed, immobilized in PVA and treated with PAPG for 15 min, and their βgalactosidase activities were assessed by CV, as described in the Materials and Methods section. (A) In the presence of phenolic compounds, β-galactosidase is expressed and hydrolyzes PAPG to PAP, which is further oxidized to PIQ at the Pt working electrode, generating a current. (B) SEM images of PVA control and cell-immobilized PVA were
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taken at 6,000 x. (C) Cyclic voltammograms of 50 mM PBS buffer (solid line), and EB cells treated with (dashed line) or without phenol (0.1 mM; dotted line) in 50 mM PBS buffer plus 15 mM PAPG. Scan rate, 100 mV s-1.
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FIGURE 2. EB cells respond to various concentrations of phenol. EB cells were prepared as described in the Materials and Methods section. Immobilized cells (approximately 4.2 x 109 CFU/ml) were treated with 10 nM to 10 mM phenol for 2 h and then exposed to 15 mM PAPG for 15 min, and CV was used to assess β-galactosidase activity at a scan rate of 100 mV s-1. The peak currents from each voltammogram were plotted as a function of the phenol concentration. The regression curve (y = 5.06 log x + 58.1; R2 = 0.9) is shown
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in the inset; linearity is seen between 0.001 and 0.1 mM phenol.
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FIGURE 3. The response of EB cells to various concentrations of phenolic compounds. EB cells were incubated for 2 h in the presence of various concentrations (1 nM to 1 mM) of phenolic compounds, and β-galactosidase activity was assessed by CV at a scan rate of
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100 mV s-1.
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FIGURE 4. Operational stability of the EB system and its use to assess phenol in wastewater. (A) EB cells were treated with 0.1 mM phenol for 2 h, and β-galactosidase activity was assessed hourly for 7 h, using CV at a scan rate of 100 mV s-1. (B) EB cells were exposed to buffer or wastewater from several sources (a~f) in Busan City spiked with 0.1 mM phenol, and β-galactosidase activity was measured by CV at a scan rate of
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100 mV s-1.
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