Development of an Electrochemical DNA Biosensor to Detect a Foodborne Pathogen


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A protocol for the development of an electrochemical DNA biosensor comprising a polylactic acid-stabilized, gold nanoparticles-modified, screen-printed carbon electrode to detect Vibrio parahaemolyticus is presented.

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Nordin, N., Yusof, N. A., Radu, S., Hushiarian, R. Development of an Electrochemical DNA Biosensor to Detect a Foodborne Pathogen. J. Vis. Exp. (136), e56585, doi:10.3791/56585 (2018).


Vibrio parahaemolyticus (V. parahaemolyticus) is a common foodborne pathogen that contributes to a large proportion of public health problems globally, significantly affecting the rate of human mortality and morbidity. Conventional methods for the detection of V. parahaemolyticus such as culture-based methods, immunological assays, and molecular-based methods require complicated sample handling and are time-consuming, tedious, and costly. Recently, biosensors have proven to be a promising and comprehensive detection method with the advantages of fast detection, cost-effectiveness, and practicality. This research focuses on developing a rapid method of detecting V. parahaemolyticus with high selectivity and sensitivity using the principles of DNA hybridization. In the work, characterization of synthesized polylactic acid-stabilized gold nanoparticles (PLA-AuNPs) was achieved using X-ray Diffraction (XRD), Ultraviolet-visible Spectroscopy (UV-Vis), Transmission Electron Microscopy (TEM), Field-emission Scanning Electron Microscopy (FESEM), and Cyclic Voltammetry (CV). We also carried out further testing of stability, sensitivity, and reproducibility of the PLA-AuNPs. We found that the PLA-AuNPs formed a sound structure of stabilized nanoparticles in aqueous solution. We also observed that the sensitivity improved as a result of the smaller charge transfer resistance (Rct) value and an increase of active surface area (0.41 cm2). The development of our DNA biosensor was based on modification of a screen-printed carbon electrode (SPCE) with PLA-AuNPs and using methylene blue (MB) as the redox indicator. We assessed the immobilization and hybridization events by differential pulse voltammetry (DPV). We found that complementary, non-complementary, and mismatched oligonucleotides were specifically distinguished by the fabricated biosensor. It also showed reliably sensitive detection in cross-reactivity studies against various food-borne pathogens and in the identification of V. parahaemolyticus in fresh cockles.


A major topic of public and scientific debate in recent years, food poisoning is mainly associated with 3 agents: microorganisms1, chemicals2, and parasites3. Contaminated food can cause serious health consequences in humans, especially in the higher risk group of those with weak immune systems, the elderly, pregnant women, babies, and young children4. With more than a million cases of acute diarrhea occurring annually in children under 5 years old in Africa, Asia, and Latin America, food poisoning is a major global disease5,6 and the World Health Organization has established microorganisms as the most important contributor7. Vibrio parahaemolyticus stands out amongst the most widely recognized virulent strains. Usually found in coastal, estuarine, and marine environments8, it is a Gram-negative bacterium, which becomes active in high salt environments, and causes serious human gastroenteritis when eaten in inadequately cooked, mishandled or raw marine products9. Additionally, existing medical conditions in some people make them prone to wound infection, septicemia or ear infection arising from V. parahaemolyticus10. The virulence factors of V. parahaemolyticus hemolysins are divided into two types which contribute to the disease pathogenesis: thermostable direct hemolysin (TDH) coded by tdh genes, and TDH-related hemolysin coded by trh genes11. The virulence markers (tdh and trh genes) of V. parahaemolyticus are mostly found in clinical specimens rather than in environmental specimens.

V. parahaemolyticus possesses the ability to survive under a broad range of conditions, rapidly responding to environmental changes12. Its proliferation mechanism escalates its hazard potential as its toxicity increases in parallel with cell mass13. Even worse, climate change is providing these bacteria with ample conditions to accelerate their cell population growth14. Due to its high frequency, V. parahaemolyticus needs to be monitored along the food supply chain, particularly in the trade and production of seafood since those products are where they are found in enormous quantities15,16 throughout the world. Currently the bacteria are identified and isolated using a range of methods including biochemical tests, enrichment and selective media17, enzyme-linked immunomagnetic sorbent assay (ELISA)18, pulse-field gel electrophoresis (PFGE)19, latex agglutination tests, and polymerase chain reaction (PCR) tests20. These methods usually require qualified personnel, sophisticated instruments, and laborious techniques which do not provide information about contamination immediately. This severely limits the likelihood of promptly detecting harmful contamination and on-site applications. Rapid detection tools remain an outstanding challenge.

Biosensing is emerging as a promising option for the detection of foodborne pathogens because it offers a time-saving, cost-effective, practical, and real-time analysis method21,22,23,24. However, although there have been many positive results of analyte detection in spiked samples and standard solution using biosensors, there is still a lack of research applied to real samples either in aqueous mixtures or organic extracts25. Recently, electrochemical biosensors using direct and/or indirect deoxyribonucleic acid (DNA) detection have received increased attention among scientists, due to their specific detection of the complementary target via a hybridization event26,27,28,29. These unique approaches are more stable in comparison with enzyme-based biosensors, thus offering a promising technology for miniaturization and commercialization. The target of the study reported here is to construct a fast tool that can detect V. parahaemolyticus with high selectivity, sensitivity, and practicality, based on the DNA sequence specificity during hybridization. Identification strategies involve the combination of polylactic acid-stabilized gold nanoparticles (PLA-AuNPs)30 and screen-printed carbon electrodes (SPCEs) in the presence of the hybridization indicator, methylene blue (MB). The potential of the developed detection construct is further explored using bacteria DNA lysate and fresh cockle samples.

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NOTE: All the chemical and biochemical reagents to be used should be of analytical grade and used without further purification. Prepare all solutions using sterile deionized water. Autoclave all glassware prior to sterilization.

Caution: Please use all appropriate safety practices when performing laboratory activities including the use of engineering controls (fume hood, glovebox) and personal protective equipment (safety glasses, gloves, lab coat, full length pants, closed-toe shoes).

1. Fabrication and Characterization of Modified Electrode using PLA-AuNPs

  1. Preparation and characterization of PLA-AuNPs
    1. Quickly add 10 mL of sodium citrate solution (38.8 mM) to 100 mL of boiling aqueous chloroauric acid solution (1 mM) and cool it down to ambient temperature. Observe the changes in color in the prepared AuNPs solution which starts as yellow, changes to blackish and finally ends as dark ruby red.
    2. Place the PLA pellets in a stainless-steel mold for the melting process and preheat them at a temperature of 190 °C for 10 min. Follow with the pressing procedure: put them under a pressure of 2.2 MPa for 1 min as part of the PLA sheet preparation. The PLA sheet will be ready for use after cooling for about 3 h.
    3. Stirring vigorously, dissolve 0.68 mg of the PLA sheets in 5 mL of chloroform at room temperature. Mix the dissolved PLA with 10 mL of the previously prepared AuNPs solution and homogeneously stir it at room temperature. The homogeneous mixtures can then be denoted as PLA-AuNPs and characterized using UV-Vis, XRD, TEM, and FESEM with EDX.
  2. Preparation and characterization of modified screen-printed carbon electrode
    NOTE: The SPCE used in this study comprises a three-electrode system: a counter electrode, a carbon working electrode, and an Ag/AgCl reference electrode.
    1. Quickly pipette 25 µL of the homogenous solution of PLA-AuNPs onto the SPCE and then air dry it for 24 h prior to use.
    2. Electrochemically characterize the modified electrodes, SPCE/PLA-AuNPs, in potassium ferro cyanide (K3[Fe(CN)6]3-/4-) to measure active surface area, electrochemical impedance spectroscopy, repeatability, reproducibility, and stability30.

2. Development of the Electrochemical DNA Biosensor

  1. Probe preparation
    Note: The sequences of the ssDNA probe and complementary DNA were chosen based on information from the National Center for Biotechnology Information (NCBI) database.
    1. Purchase synthetic oligonucleotides (20-mer ssDNA probe, 20-mer complementary DNA, 20-mer mismatched DNA, and 21-mer non-complementary DNA) as lyophilized powder from commercial laboratories, based on the following sequences:
      thiolated ssDNA probe: 5′- / 5ThioMC6-D/CGGATTATGCAGAAGCACTG - 3′
      complementary DNA: 5′ - CAGTGCTTCTGCATAATCCG - 3′
      one-base mismatched DNA: 5′ - CAGTGCTTCTGC ṪTAATCCG - 3′
      three-base mismatched DNA: 5′ - CAGTGCTTCT Ċ ṪṪTAATCCG - 3′
      non-complementary DNA: 5′ - CGCACAAGGCTCGACGGCTGA - 3′
    2. Prepare stock solutions of all oligonucleotides (100 µM) using a sterile TE solution (10 mM Tris-HCl, 1 mM EDTA, pH 7.5), divided into analytical portions. Keep this at -4 °C and then make the appropriate dilutions just prior to use.
  2. Optimization of Immobilization and Hybridization
    Note: An SPCE modified with PLA-AuNPs, denoted as SPCE/PLA-AuNPs, was used for this study and a drop casting method was used for the DNA immobilization and hybridization.
    1. First, immobilize 25 µL of thiolated ssDNA probe on the SPCE/PLA-AuNPs and then air dry it for 24 h at room temperature, after which it will be finally denoted as SPCE/PLA-AuNPs/ssDNA. Determine optimization of the immobilization condition by using three factors: the concentration of ssDNA (ranging from 0.2 to 1.4 µM), time (ranging from 30 to 210 min), and temperature (ranging from 25 to 75 °C).
    2. Pipette 25 µL of the complementary DNA on to the surface of SPCE/PLA-AuNPs/ssDNA to carry out the hybridization event after which it can be finally denoted as SPCE/PLA-AuNPs/dsDNA. Determine optimization of the hybridization condition via the two factors of time (ranging from 5 to 30 min) and temperature (ranging from 25 to 75 °C).
    3. Immerse the immobilized and hybridized electrodes in 20 µM MB for 30 min. Remove the non-specifically adsorbed DNA and excess MB by washing with 0.5 M ABS/20 mM NaCl (pH 4.5) and then rinsing with deionized water. Measure the peak current of MB reduction using DPV technique. Execute the DPV measurement using 0.1 M PBS (pH 7) containing no indicator.
  3. Characterization of the fabricated DNA biosensor
    1. Immerse the SPCE/PLA-AuNPs in 20 µM MB for 30 min, wash with 0.5 M ABS/20 mM NaCl (pH 4.5), and then rinse with deionized water prior to measuring. Follow a similar procedure for all interactions including the probe DNA, complementary DNA, mismatched DNA, and non-complementary DNA samples.
    2. Measure the electrochemical reduction.
      NOTE: We measured DPV using a commercially manufactured Autolab with interface software.
      The DPV measurements of the MB electrochemical reduction performed at a potential ranging from -0.5 V to 0.25 V, with a step potential of 0.005 V, modulation amplitude of 0.05 V, and a scan rate of 7.73 mVs-1 in 0.1 M PBS (pH 7), containing no indicator. The selectivity, sensitivity, reproducibility, and heat stability of the fabricated DNA biosensor were further studied. Reported results were presented as mean value measurements in three replicates.

3. Validation of the Fabricated DNA Biosensor Using Real Samples

  1. Preparation of bacterial strains
    1. Select bacterial samples based on their significant contamination of seafood31,32 .
      NOTE: V. parahaemolyticus as reference strains and 8 other bacterial strains (Campylobacter jejuni, Listeria monocytogenes, Salmonella typhimurium, Salmonella enteritis, Klebsiella pneumonia, Escherichia coli O157:H7, Bacillus cereus, and Vibrio alginolyticus) of the common foodborne pathogen were employed for the electrochemical DNA biosensor validation in this study.
    2. Conduct a routine sub-culture of bacterial strains on their respective agar every two weeks to maintain their viability, refer Table 1 for culture conditions.
    3. Maintain long-term preservation of the bacterial strains in their respective growing broths containing 20% (v/v) glycerol at -80 °C as glycerol protects bacteria by preventing the formation of ice crystals that can damage them during the freezing process. Next, inoculate a loop of preserved bacterial cell culture in glycerol stocks into the respective broths. Incubate the broths according to their respective growth time and temperature when needing to revive the bacteria.
    4. Determine the quantity of V. parahaemolyticus cells using a spread plate technique33, a standard and a well-known method for enumerating microorganisms derived from a series of dilutions.
      1. Culture V. parahaemolyticus in Trypticase Soy Broth (TSB + 3% NaCl) at 37 °C in a 140-rpm shaking condition. Then, transfer 1 mL of the bacteria cell culture into 9 mL of TSB + 3% NaCl to obtain a 10-1 dilution factor.
    5. Repeat this step nine times to get a full series of up to 10-10 dilution factor. Next, spread 0.1 mL of each dilution factor (starting from 10-1 to 10-10) on a Vibrio's selective agar plate for colony counting, respectively. Incubate the plates incubated at 37 °C for 24 h.
    6. Use a colony counter to count individual colonies and then back-calculate (count the CFU/pipetted amount x dilution factor) to get a colony-forming unit per milliliter density (CFU mL-1). Derive the concentration of the colony forming units (CFUs) in the bacteria cell suspensions from the calibration plot of CFU mL-1 against absorbance.
  2. Preparation of the PCR assay
    1. Extract genomic DNA following a modified boiled lysis procedure34. First, streak an individual bacterial strain on agar media and inoculate it in broth media, followed by incubating it at its relative growth condition, a 140 rpm shaking condition.
    2. Pipette a 1 mL culture of broth into a micro centrifuge tube. Centrifuge this at 4830 x g and 4 °C for 1 min to obtain pellets. Use about 500 µL of sterile TE buffer for re-suspension with the pellet. Hold the resulting suspension for 10 min at 98 ± 2 °C in a heating block and immediately chill it at -18 °C for 10 min to lyse the cells and release the DNA.
    3. Centrifuge the genomic DNA at 4830 x g and 4 °C for 3 min to obtain a clear suspension and keep the supernatant at -20 °C for further use. Use a biophotometer measurement with UV absorption at a wavelength of 260 nm (as nucleic acids absorbing wavelength of light) and also at a wavelength of 280 nm (as the proteins absorbing wavelength of light) to determine the concentration and purity of the extracted genomic DNA and then identify all strains using standard biochemical assays35 and verifying them by 16S rRNA gene sequencing36. Finally, denature the genomic DNA at 92 °C for 2 min and cool it rapidly in ice water prior to application to the biosensor.
    4. Further confirm the presence V. parahaemolyticus using PCR to target the toxR gene. Perform this procedure in a thermocycler using a primer pair (5'-GTCTTCTGACGCAATCGTTG-3' and 5'-ATACGAGTGGTTGCTGTCATG-3'). Optimize the PCR amplification in a total reaction volume of 25 µL consisting of sterile water (15.5 µL), buffer (5 µL), primer (1 µL, 10 µM), dNTP mix (1 µL), DNA template (2 µL), and Taq DNA polymerase (0.2 µL, 10x).
    5. Mix the components well and arrange the PCR amplification of the target sequence in a thermocycler programmed for 30 cycles of amplification. In our study, each cycle consisted of three-step reactions i.e., initial denaturation (94 °C, 3 min) followed by 30 cycles of denaturation (94 °C, 1 min), annealing (63 °C, 1.5 min) and extension (72 °C, 1.5 min), followed by final extension (72 °C, 7 min).
    6. Determine the amplified products and their sizes via electrophoresis on 1.5% agarose gel. Capture the gel images with a commercially produced gel-documentation system.
  3. Preparation of cockle samples
    1. Obtain fresh samples.
      NOTE: Our fresh cockles (Anadara granosa) were obtained from the wet market in Serdang, Selangor, and quickly brought to the laboratory in an ice cooler box for analysis. Divide the samples into 2 groups, namely the treated and untreated group, assuming that the cockles were harvested uniformly from the start of the harvest until placing them into cold storage.
    2. Pre-treat cockles in the treated group by storing them at -20 °C for 24 h, followed by exposure to UV light at 20 °C for 4 h prior to DNA extraction.
      NOTE: The freezing temperature and UV exposure used for the controlled group kills or at least limits the naturally accumulating V. parahaemolyticus in the cockles. Do not apply a higher pasteurization regime of 70 °C as the aim of the controlled condition in this study is to mimic the actual situation of the fresh cockles. Meanwhile, analyze samples directly from the untreated group without any pre-treatment as soon as they arrive in the laboratory.
    3. Subculture a strain of V. parahaemolyticus ATCC 17802 in TSB (3% NaCl). Determine the level of viable cells in the inoculum via plating of appropriate dilutions of TSB (3% NaCl) on CA in order to obtain an inoculum of 104 CFU mL-1 of V. parahaemolyticus. Wash each cockle in distilled water and scrub it free of dirt before using sterile forceps in a laminar flow cabinet to remove the tissues from the shell.
    4. Homogenize about 10 g of cockle tissue samples with a homogenizer in 90 mL of sterile TSB (3% NaCl) for 60 s. Add a known amount of V. parahaemolyticus to 9 mL of the homogenized sample broth for the spiked samples. Use the unspiked samples as a negative control. Estimate the cell counts of V. parahaemolyticus spiked on to the cockle samples by plating 0.1 mL of the samples on CA, and subsequently, pipetting 1 mL of samples into a microcentrifuge tube for DNA sample extraction, to be used in the biosensor and PCR assay.
    5. Extract the genomic DNA of the V. parahaemolyticus from the spiked and unspiked samples following a modified boiled lysis procedure36. Finally, determine the DNA concentration and purity using a bio photometer measurement with UV absorption at a wavelength of 260 nm (as nucleic acids absorbing wavelength of light) and also at a wavelength of 280 nm (as the proteins absorbing wavelength of light).

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Representative Results

Formation of AuNPs was revealed through the change in color of the aqueous solution with sodium citrate present. This caused the color to change from light yellow to a deep ruby red. The generation of PLA-AuNPs was confirmed from the UV-vis spectra (Figure 1) where the growth of the surface plasmon resonance (SPR) peak was found at around 540 nm. The formation and existence of PLA-AuNPs was indicated at 500-600 nm wavelength ranges, depending on particle size37. The XRD patterns of AuNPs and PLA-AuNPs are shown in Figure 2. All crystallite peaks were observed at the 2θ of 31.7, 38.2, 44.4, 64.7, and 77.7°. They were attributed to the (100) (111) (200) (220), and (311) crystallographic planes of cubic-FCC (face-centered cubic) gold (metal) nanostructures (JCPDS file no.-00-004-0783). The PLA-AuNPs crystallite peak intensities also increased as a broader diffraction peak centered at 31.7°, implying that the AuNPs were embedded in the PLA and suggesting the formation of polyphasic gold nanostructures38. Furthermore, all AuNP diffraction peaks shown on the PLA-AuNPs indicated that the AuNPs had a stable structure. The mean size of PLA-AuNPs was calculated using Scherrer's equation (L = kλ/βcosθ), by determining the width of the (100) Bragg's reflection. From the FWHM and d-spacing (Table 2), the crystallite size of PLA-AuNPs was calculated as 27 nm.

The dimensions of the PLA-AuNPs and its morphology were further investigated using TEM. From the TEM particle distribution curve and images of PLA-AuNPs, it was seen that gold nanoparticles were almost spherical in shape (Figure 3). The size of the nanoparticles was in the range of about 37 nm, slightly bigger than obtained from Scherrer's formula, suggesting a polycrystalline nature of the as-synthesized nanoparticles39. Smaller size resulting from XRD in contrast to TEM has also been observed in previous research, most likely because the particle structure is polycrystalline in nature because of the significant amount of crystallites that are smaller (sub-particles)40. Figure 4 shows the FESEM image of as-prepared PLA-AuNPs on SPCE. It is evident from the FESEM images and EDX spectra that the SPCE/PLA-AuNPs had the highest weight percentage of gold, greatest density, and largest surface of nanoparticle coverage. It can also be noted, particularly, that Figure 4(a) shows the FESEM image of a bare SPCE. The FESEM image in Figure 4(b) shows well-distributed PLA-AuNPs. To affirm the synthesized PLA-AuNPs' chemical composition, EDX was employed to investigate the chemical composition of the as-produced PLA-AuNPs, as per Figure 4(a)and 4(b). The EDX spectrum confirmed that the as-produced PLA-AuNPs were composed of gold (Au) only, and displayed no other element except the peak corresponding to carbon (C) and oxygen (O). The presence of the C peak was due to the carbon electrode on to which the sample was drop cast.

The EDX spectrum also showed the presence of O indicating that oxygen was adsorbed on the carbon electrode because the film was exposed to air. This confirms that oxygen is adsorbed on carbon nanotubes if they are exposed to air for a long time41. Although there are various gas molecules in air, oxygen was believed to be the main adsorbate on the air-exposed carbon nanotubes42, possibly due to its faster adsorption compared with other molecules43. This was based on finding that oxygen was the main adsorbate on the carbon electrode during overnight air exposure in sample preparation and reinforced the chemical purity of the PLA-AuNPs. The ratio of anodic and cathodic peak (ipa/ipc) was almost 1, which gave constant peak potential as the scan rate increased44. Moreover, the anodic and cathodic peak potentials were consistent at different scan rates45 suggesting that the electrochemical reaction was reversible based on peak separation (Figure 5). The active surface area of the modified electrode was calculated using the Randles-Sevick equation (ipc = (2.69×105) n3/2 AD1/2 Cv1/2. From the slope of the plot ipc versus v1/2 in Figure 5(a) and (b), the surface area of the electrodes was calculated as 0.26 cm2 and 0.41 cm2 for the bare SPCE and PLA-AuNPs, respectively. This result confirmed that the improvement afforded by PLA-AuNPs on the active surface area was relatively higher compared with bare SPCE. The derived result was comparable to that of polymer-embedded gold nanoparticles (β-CD-EDAS(N-[3-(trimethoxysilyl)propyl]ethylene-diamine matrix encapsulated gold nanoparticles), which revealed that a high active surface area enhanced the process of electron-transfer and therefore resulted in better sensitivity46.

The modified SPCE electrochemical reaction was in a process of controlled diffusion, shown by a reduction in the K3[Fe(CN)6]3-/4- peak current, which in turn relied on the current of the scan rate square root (v1/2). To get a better view of the modified electrode properties, an electrochemical impedance spectroscopy (EIS) study was performed to correlate with the performance of the modified electrode presented in terms of CV measurements. The semicircle section seen at high frequencies correlated with the limiting process of the electron transfer in the EIS. The charge transfer resistance (Rct) was measured directly as the diameter of the semicircle. This result tallied with the value of Rct of the bare SPCE, which was 1932 Ω and where the modification of SPCE/PLA-AuNPs decreased to 1444 Ω (Figure 6). The decrease of Rct value in SPCE/PLA-AuNPs may have resulted from a decrease in local dielectric constant and/or an increase in the thickness of the electric double layer47, suggesting that K3[Fe(CN)6]3-/4- function by adsorption at the metal/solution interface. These outputs complied with the ones derived from the measurements of CV (Figure 7). The peak currents increased progressively with the modification of the SPCEs using PLA-AuNPs, which showed that a higher sensitivity was the inverse of the lower Rct48. Thus, the increase in the value of CV of the modified electrodes and the decrease in Rct values may have been due to the gradual replacement of water molecules (the volume of the water molecules is 27.19 Å3) by the adsorption of the K3[Fe(CN)6]3-/4- on the metal surface, decreasing the extent of the dissolution reaction49.

Table 3 indicates the relative standard deviation (RSD) of the repeatability and reproducibility of the modified electrode. Routine CV recording of K3[Fe(CN)6]3-/4- was made for six consecutive sets to investigate the repeatability of the modified electrodes. The RSD of SPCE/PLA-AuNPs was derived as 4.26%. The electrode with PLA-AuNPs modification gave better RSD after several measurements. Moreover, using the same procedure, six modified electrodes were independently fabricated, which gave RSD values of 4.05% for SPCE/PLA-AuNPs, indicating better reproducibility of the electrode fabrication for PLA-AuNPs. The modified substrate depending on PLA-AuNPs was kept at 37 °C for an extended duration. It was also utilized for the record of 20 cycles and a decrease of 18% in signal quality. In order to achieve the maximum output from the biosensor, immobilization period, temperature, together with ssDNA concentration and criteria for hybridization were examined. These conditions needed to be optimized, as they influence the peak current of DPV. To optimize the ssDNA concentration, SPCE/PLA-AuNPs was pipetted with different concentration (0.2, 0.4, 0.6, 0.8, 1.0, 1.2 and 1.4 µM) of ssDNA for 60 min so that the optimum concentration of ssDNA was determined. Based on Figure 8, the DPV peak current increased as higher DNA concentrations were used. The study also found that the optimized concentration of the DNA was 1.2 µM ssDNA.

To optimize the immobilization period involving the single-stranded DNA in SPCE/PLA-AuNPs, 25 µL of ssDNA (1.2 µM) were tested over different times (30, 60, 90, 120, 150, 180 and 210 min). In Figure 9, the findings show an increase in immobilization period resulting in the continuous increase of the DPV peak current. Thus, the optimum immobilization time of the ssDNA was selected as 180 min. To optimize the immobilization temperature, the SPCE/PLA-AuNPs was pipetted with ssDNA (1.2 µM) in various temperatures (75, 65, 55, 45, 35, and 25 °C) for 180 min. We found that when the temperature of immobilization was increased to 55 °C, the DPV peak current initially escalated followed by a subsequent depreciation (Figure 10); thus, 55 °C was selected as the optimized immobilization temperature to be used in subsequent experiments. Separation of ssDNA that is immobile from surfaces of the electrode and hybridization disablement was caused by increased temperatures. There have been other reports with similar results50. Faster splitting and degeneration of DNA from the surfaces of nanoparticles of gold occurred at higher temperatures51. Research conducted recently reported that the bond between gold and thiol oxidizes degrades quickly and simply in a situation that is ambient, for example when exposed to water, air, and light52,53,54. The effect of hybridization time was also investigated in this study.

The fabricated SPCE/PLA-AuNPs/ssDNA was drop-cast with a solution of target DNA (0.2 µM) at different hybridization times (5, 10, 15, 20, 25, and 30 min). The response of the DPV rose obviously, with a 5-min increase in time of incubation (from 5 to 10 min) (Figure 11). Once this value was obtained, a 10 to 30 min downward pattern was observed so the desired time for hybridization was chosen as 10 min. The efficiency of hybridization increased with an increase in temperature from 25 °C to 35 °C (Figure 12). The high current was seen to slowly decrease at a temperature higher than 35 °C. The reason for this was attributed to sandwich structure denaturation reducing the detection signal. Hence, the optimal hybridization temperature selected was 35 °C. The process we used in detecting V. parahaemolyticus comprised AuNPs to enhance the active surface area of the working electrode, PLA to mediate the surface modification and the redox complex, MB, to reduced it to LB. The outcome of this oxidation process was a better affinity between ssDNA and MB55. The DPV current was decreased due to the lower number of attached MB molecules during hybridization of ssDNA with its complementary sequence.

We conducted further investigations into the performance of the developed biosensor. The DPV results in Figure 13 show that it was able to selectively distinguish probe DNA, complementary DNA, one-base mismatched DNA, three-base mismatched DNA, and non-complementary DNA in the presence of MB as the detection label56,57. The lowest peak current (0.73 µA) was exhibited by complementary DNA, which slowly increased by one-base mismatched DNA (0.94 µA), three-bases mismatched DNA (1.05 µA), non-complementary DNA (3.25 µA), and probe DNA (4.79 µA). The target DNA current was the smallest of all the currents of oligonucleotide sequences allowing our biosensor to discriminate between the sequences of oligonucleotide sensitively and specifically. MB bound strongly with ssDNA of immobilized probe DNA producing a large DPV peak current. SPCE/PLA-AuNPs/non-complementary DNA reduced the immobilized probe DNA current by almost half as only a small amount of MB assembled on the surface of SPCE/PLA-AuNPs/dsDNA. This is likely to be due to inaccessible interactions between MB and guanine bases. The interaction modes of DNA and MB are strongly dependent on such experimental conditions as pH, temperature, DNA concentration, ionic strength, and type of buffer58,59. MB commonly links to dsDNA by groove and intercalative modes, resulting in MB accumulation on dsDNA surface60. The exposure of our capture probe DNA to one-base mismatched DNA and three-base mismatched DNA consistently reduced DPV peak currents and the complementary target DNA continued this trend. Table 4 shows the percentage of selectivity rate for MB peak current. Our overall finding was that hybridization of the constructed DNA biosensor was highly selective via immobilized probe DNA on the SPCE/PLA-AuNPs' surface.

The sensitivity of the developed DNA biosensor was further investigated with various concentrations of complementary target oligonucleotide. Figure 14 depicts the gradual depletion of DPV peak current of MB on the SPCE/PLA-AuNPs as the complementary DNA concentration increased. Figure 15 exhibits a linear regression coefficient of 0.96 which was generated between the log of reduced MB peak current and the log of several concentration of target DNA (0.2 pM to 0.02 mM). A graph of the peak current changes (ΔP) was plotted using the 3σ/m formula (σ being the standard deviation of blank solution and m being the slope of the linear curve)61,62,63. The detection limit of the fabricated DNA biosensor was calculated to be 4.43 pM which was greater than our smallest actually measured sample. This outcome was in agreement with the fact that ΔP for 0.2 pM and 2 pM (2.65×10-6 A and 2.81×10-6 A) with a standard deviation of 1.19×10-7 A and 1.06×10-7 A, respectively, overlapped.

To compute the LOQ for the fabricated DNA biosensor, we used the 10σ/m formula in which (σ is the blank solution standard deviation and m is the slope of the linear curve) and found the outcome to be a gain of 14.8 pM. Repeated denaturation of the complementary DNA was performed to assess the DNA biosensor in terms of its regeneration capacity. We did this by incubating 0.2 µM complementary DNA-modified electrode in hot water at 86 °C for 8 min to denature the double-stranded DNA, followed by rapid cooling in an ice bath to maintain the denatured single-stranded DNA64.

The hybridization activity was maintained at 91% of its initial response after 8 efforts of regeneration (n = 5, RSD = 4.05%). The DNA biosensor based on SAM method for DNA probe immobilization on to SPCE/PLA-AuNPs was highly stable, achieving a good response in detecting complementary DNA after repeated denaturation processes. The stability of our DNA biosensor regarding heat was further explored. To do this, we stored the PLA-AuNPs/SPCE/ssDNA electrode at temperatures of 4 °C, 25 °C, and 45 °C in a sealed aluminum pouch containing silica gels. At the end of 6 months, we assessed the complementary DNA and estimated the percentage of recovery of signal production. Figure 16 shows that the probe SPCE/PLA-AuNPs/ssDNA was highly stable with more than 80% of its initial signal still recoverable after 6 months' storage. The strong reaction between PLA-AuNPs and ssDNA was found to have long-term stability at a temperature below 45 °C.

Nine different species of bacteria (B. cereus, L. monocytogenes, C. jejuni, E. coli O157:H7, V. alginolyticus, S. typhimurium, S. enteritis, K. pneumonia and V. parahaemolyticus) were screened by PCR detection. The PCR amplification products for species-specific detection of V. parahaemolyticus from the bacteria culture are shown in Figure 17. The toxR gene of V. parahaemolyticus was utilized as PCR primer for V. parahaemolyticus identification due to its presence in pathogenic isolates. From the PCR analysis, V. parahaemolyticus showed positive results due to the specificity of the PCR primer of the toxR gene towards V. parahaemolyticus. No PCR products from other bacterial strains were detected. The DPV peak current obtained after a cross-reactivity assessment showed very clear detection of V. parahaemolyticus in comparison with other bacteria species, as depicted in Figure 18. The DPV signal increased as the number of base pairs decreased, due to the larger amount of MB molecules bound to the probes. V. parahaemolyticus were clearly able to be discriminated from the other foodborne pathogens which mimicked the environment of the food sample.

Cockles were selected as real samples for the DNA biosensor validation in this study since they are popular ingredients in several types of local food. It is well known that raw cockles often carry pathogenic V. parahaemolyticus and can transmit these bacteria to humans, especially if they are inadequately refrigerated during storage after harvesting65. The treated group of cockle samples was stored at -20 °C for 24 h, and exposed to UV light at 20 °C for 4 h prior to DNA extraction during our pre-treatment step. Figure 19 shows that the PCR analysis found V. parahaemolyticus in both the treated and untreated (spiked) samples. This is shown in Lane 7, 8, 9, 10, 11, and 12 correlating with the colony count in the previous discussion. No V. parahaemolyticus appears in the treated and untreated (unspiked) samples, as shown in Lane 1, 2, 3, 4, 5 and 6 in the PCR results although the colony count did indicate its presence in the samples. A possible reason for this is the presence of contaminating metabolites such as protein in the extracted DNA66.

Figure 20 summarizes the detection results using the developed DNA biosensor, which successfully determined the presence of V. parahaemolyticus in the treated group consisting of spiked and unspiked samples. These results positively correlate with the previously discussed PCR results. Every sample of the PCR that showed positive results was discovered using the PLA-stabilized electrochemical biosensor technique, which is AuNP-based, and the DPV peak area clearly corresponded with the resulting PCR intensity bands (Figure 19 and Figure 20). These results indicate that the suggested nanoparticle-based electrochemical sensor in this study detected V. parahaemolyticus successfully in the real samples, thus proving that it is superior to PCR with no effect on the authenticity of the results.

Figure 1
Figure 1: Absorbance of PLA-AuNPs Please click here to view a larger version of this figure.

Figure 2
Figure 2: X-ray diffraction spectrum of PLA-AuNPs. Adapted with permission from ref. [30]. Copyright (2016) Sociedade Brasileira de Química. Please click here to view a larger version of this figure.

Figure 3
Figure 3: TEM frequency diameter distribution and images of PLA-AuNPs at a measurement scale of 50 nm. Reprinted with permission from ref. [30]. Copyright (2016) Sociedade Brasileira de Química. Please click here to view a larger version of this figure.

Figure 4
Figure 4: EDX spectra and FESEM images of (a) bare SPCE and (b) SPCE/PLA-AuNPs. Reprinted with permission from ref. [30]. Copyright (2016) Sociedade Brasileira de Química. Please click here to view a larger version of this figure.

Figure 5
Figure 5: The plot of peak current oxidation against the scan rate square root and cyclic voltammograms: (a) bare SPCE and (b) SPCE/PLA-AuNPs in 1.0 mmol L-1 K3[Fe(CN)6] (pH 7). The scan rates were 300, 250, 200, 150, 100, and 50 mV s1. Adapted with permission from ref. [30]. Copyright (2016) Sociedade Brasileira de Química. Please click here to view a larger version of this figure.

Figure 6
Figure 6: Impedance spectra of modified electrodes in 1.0 mmol L-1 K3[Fe(CN)6] (pH 7). Amplitude: 5 mV, and frequency range: 0.1-105 Hz. Adapted with permission from ref. [22]. Copyright (2016) Elsevier. Please click here to view a larger version of this figure.

Figure 7
Figure 7: Anodic peak current of modified electrodes in 1.0 mmol L-1 K3[Fe(CN)6] (pH 7) at a scan rate of 0.1 V s-1 using cyclic voltammetry measurement Please click here to view a larger version of this figure.

Figure 8
Figure 8: Effect of concentration for ssDNA immobilization on SPCE/PLA-AuNPs in 0.1 mol L-1 PBS (pH 7) at scan rates of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement Please click here to view a larger version of this figure.

Figure 9
Figure 9: Effect of ssDNA immobilization time on SPCE/PLA-AuNPs in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement Please click here to view a larger version of this figure.

Figure 10
Figure 10: Effect of ssDNA immobilization temperature on SPCE/PLA-AuNPs at a scan rate of 0.1 V s-1 at 0.1 mol L-1 PBS (pH 7) after an incubation period of 30 min in 20 µM of MB using differential pulse voltammetry measurement Please click here to view a larger version of this figure.

Figure 11
Figure 11: Effect of hybridization time on SPCE/PLA-AuNPs/ssDNA in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement Please click here to view a larger version of this figure.

Figure 12
Figure 12: Effect of hybridization temperature on SPCE/PLA-AuNPs/ssDNA in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement. Reprinted with permission from ref. [22]. Copyright (2016) Elsevier. Please click here to view a larger version of this figure.

Figure 13
Figure 13: Histogram of the MB reduction peak current using different types of DNA in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB. The inset illustrates differential pulse voltammograms in the same conditions. Reprinted with permission from ref. [22]. Copyright (2016) Elsevier. Please click here to view a larger version of this figure.

Figure 14
Figure 14: Histogram of the MB reduction peak current using different DNA concentration in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement. Reprinted with permission from ref. [22]. Copyright (2016) Elsevier. Please click here to view a larger version of this figure.

Figure 15
Figure 15: A linear plot of the reduction peak current of MB against the log of different DNA concentration in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement. Reprinted with permission from ref. [22]. Copyright (2016) Elsevier. Please click here to view a larger version of this figure.

Figure 16
Figure 16: Effect of different hybridization temperatures on SPCE/PLA-AuNPs/ssDNA at 6 months of storage interval (n = 3). Measurements were made in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement. Reprinted with permission from ref. [22]. Copyright (2016) Elsevier. Please click here to view a larger version of this figure.

Figure 17
Figure 17: Agarose gel electrophoresis-PCR amplification using 16S rRNA gene sequencing products of selected bacteria from bacteria culture. Lane M: 1kb DNA ladder, Lane C−: negative control (sterile distilled water), Lane C+: positive control (DNA extracted from E. coli is used as template), Lane EC: E. coli O157:H7, Lane CJ: C. jejuni, Lane BC: B. cereus, Lane KP: K. pneumonia, Lane ST: S. typhimurium, Lane LM: L. monocytogenes, Lane SE: S. enteritis, Lane VP: V. parahaemolyticus and Lane VV: V. alginolyticus Please click here to view a larger version of this figure.

Figure 18
Figure 18: Histogram of the MB reduction peak current for cross-reactivity study of the DNA biosensor against various foodborne pathogens in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement. Reprinted with permission from ref. [23]. Copyright (2017) Springer. Please click here to view a larger version of this figure.

Figure 19
Figure 19: Agarose gel electrophoresis-PCR amplification products of V. parahaemolyticus from fresh cockle samples. Lane M: 2ul of 100 bp DNA ladder, Lane 1-3: untreated unspiked samples, Lane 4-6: treated unspiked samples, Lane 7-9: untreated spiked samples, Lane 10-12: treated spiked samples, Lane C+: positive control (V. parahaemolyticus ATCC 17802), Lane C-: negative control (sterile distilled water). Reprinted with permission from ref. [23]. Copyright (2017) Springer. Please click here to view a larger version of this figure.

Figure 20
Figure 20: Histogram of the MB reduction peak current for detection of V. parahaemolyticus in fresh cockle samples using the DNA biosensor in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB using differential pulse voltammetry measurement. Reprinted with permission from ref. [23]. Copyright (2017) Springer. Please click here to view a larger version of this figure.

Electrode composition Incubation time (h) Temperature (°C) Growth media
Campylobacter jejuni 48 42 BA/ BB
Listeria monocytogenes 48 30 TSA/ TSB
Salmonella typhimurium 24 37 TSA/ TSB
Salmonella enteritidis 24 37 TSA/ TSB
Klebsiella pneumonia 24 37 EMBA/ TSB
Escherichia coli O157:H7 24 37 MA/ TSB
Bacillus cereus 24 37 MYPA/ TSB
Vibrio alginolyticus 24 37 TSA+3% NaCl/TSB+3% NaCl
Vibrio parahaemolyticus 24 37 CA/TSA+3% NaCl/TSB+3% NaCl

Table 1: Culture conditions of selected bacteria.

Sample Pos. [°2Th.] FWHM [°2Th.] d-spacing [Å] Crystallite size (nm)
PLA-AuNPs 31.682 0.3149 2.8243 27

Table 2: The size of the PLA-AuNP crystallites. Adapted with permission from ref. [30]. Copyright (2016) Sociedade Brasileira de Química.

Modified electrode % RSD (n = 6)       Signal reduction (%) (n = 20)
Repeatability Reproducibility
SPCE/PLA-AuNPs 4.26 4.05 18

Table 3: Properties of modified electrodes relating to AuNPs and PLA-AuNPs modification in K3[Fe(CN)6] with 1.0 mmol L-1 concentration at a scan rate of 0.1 V s-1. Adapted with permission from ref. [30]. Copyright (2016) Sociedade Brasileira de Química.

Electrode composition ipa (µA) Epa Selectivity rate (%)
(n = 3) (V)
SPCE/PLA-AuNP/probe DNA 4.79 ± 0.10 -0.46 -
SPCE/PLA-AuNP/non-complementary DNA 3.25 ± 0.12 -0.44 67.85
SPCE/PLA-AuNP/3-bases mismatched DNA 1.05 ± 0.11 -0.45 21.92
SPCE/PLA-AuNP/1-base mismatched DNA 0.94 ± 0.12 -0.46 19.62
SPCE/PLA-AuNP/complementary DNA 0.73 ± 0.10 -0.45 15.24

Table 4: DPV selectivity of MB peak current in 0.1 mol L-1 PBS (pH 7) at a scan rate of 0.1 V s-1 after 30 min incubation in 20 µM MB

Composition of the electrodes Detection method Linear range (M) Detection limit (M) References
3D AuNPs DPV 1.5×10-6 - 7.0×10-9 2.0×10-10 69
(+)AuNPs DPV 1.0×10-9 - 1.5×10-12 2.6×10-12 70
AuNPs/GR DPV 1.3×10-9 - 2.5×10-11 8.3×10-12 71
AuNPs-ADH-GSs DPV 5.0×10-8 - 3.0×10-13  3.0×10-13 72
AuNPs-MPTS DPV 5.0×10-9 - 1.0×10-11  5.0×10-11 73
AuNPs–GO DPV 1.0×10-9 - 1.0×10-14 3.5×10-15 74
CoS/AuNPs DPV 1.0×10-9 - 1.0×10-12 7.0×10-13 75
PLA-AuNPs DPV 2.0×10-8 - 2.0×10-13 5.3×10-12 This work

Table 5: Comparison of some of the recently developed electrochemical DNA nanosensors. Reprinted with permission from ref. [22]. Copyright (2016) Elsevier.

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The critical steps in a framework for successful development of this type of electrochemical biosensor are selection of appropriate biological recognition elements for the transducer (nucleic acid or DNA here); chemical approach for constructing the sensing layer of the transducer; transduction material; optimization of DNA immobilization and hybridization; and validation of the developed biosensor using real samples.

Core to the successful development of a sensitive and selective electrochemical DNA biosensor, is the optimization of the immobilization and hybridization condition. In this work, we used MB as the redox complex. There are various principles of MB-DNA interactions: (i) by electrostatic interaction of the MB cationic charge with the phosphate backbone anionic charge of both ssDNA and dsDNA; (ii) by MB intercalation through successful insertion of MB between alternating G-C base pairs in dsDNA; (iii) by covalent bond to the end of the target ssDNA, which produces a low label density (one or two MB molecules per DNA strand); and (iv) by MB interactions with an unbound guanine base in ssDNA. We applied the fourth principle, which accurately identifies V. parahaemolyticus from cross reactivity of bacteria and in fresh cockles. After oxidization of particular interest, there was a stronger affinity between MB and ssDNA. Hybridization of ssDNA with its complementary sequence replaces the bonded MB molecules, and thus reduces the signal.

The series of experimental results presented above focuses on the development of a rapid method of detecting V. parahaemolyticus with high selectivity and sensitivity using the principles of DNA hybridization. The first step is the synthesis and characterization of polylactic acid-stabilized gold nanoparticles (PLA-AuNPs) for modification of the electrode (Figures 1-6 and Table 2). The quality of the gold nanoparticles was evaluated using Ultraviolet-visible Spectroscopy (UV-Vis), X-ray Diffraction (XRD), Field-emission Scanning Electron Microscopy (FESEM), Transmission Electron Microscopy (TEM), Cyclic Voltammetry (CV) and Impedance analysis. The second part of the development involved an electrochemical characterization of the modified electrode (Figures 7-12 and Table 3-4) which determined the successful enhancement of the modified electrode relative to the bare electrode in terms of detecting the hybridization event.

The third step of the DNA based biosensor development was based on the optimization of the experimental conditions of the hybridization event and the application of the developed biosensor to the detection of the actual pathogen (Figures 13-20). The hybridization event which determined the success of the detection of the pathogen was assessed using differential pulse voltammetry (DPV). The positive existence of the targeted pathogen was indicated through reduction of the current. The sensitivity of the developed biosensor was evaluated through the construction of a calibration plot and the calculation of LOD (Figure 15). The fabricated biosensor was able to specifically distinguish V. parahaemolyticus from other pathogens based on the different levels of current reduction (Figures 17 and 18). The evaluation of the feasibility of the developed biosensor for detection of V. parahaemolyticus in actual food sample is presented in Figures 19 and 20 and the positive existence of the targeted pathogen is indicated by reduction of the current. The successful development of DNA-based biosensors is highly dependent on DNA probe selection, enhancement of the modified electrode as well as the optimization of the experimental conditions. The DNA probe here has been adapted from the previous work of Nakano67. Enhancement of the modified electrodes was performed by using gold nanoparticles stabilized with polylactic acid to enable the standardization of the nanosize. The gold nanoparticles function as electro catalytic material68 and act to increase the transfer rate of the electron and also provide a surface on which to anchor the DNA probe. The experimental parameters were optimized in terms of temperature and incubation time which is crucial for DNA to form duplex. The hybridization temperature affects the separation between the non-specific adsorption and this increases the selectivity of the sensor. Incubation time of the hybridization increases the chances of the duplex formation. The immobilization procedure must be optimized in order to make sure the DNA probe will be in suitable orientation for the duplex formation to form.

Table 5 compares a sample of recently developed electrochemical DNA biosensors69,70,71,72,73,74,75,. DNA-based biosensing is a promising method for replacing molecular based techniques although there are a few limitations that must be addressed before the method can be portable enough for onsite usage. The main limitation is sample processing DNA extraction as the purity and the amount of DNA extracted on site is not likely to be as good as that extracted by the standard lab-based method. It is, however, a promising alternative if the PCR and gel electrophoresis can be replaced with sample pretreatment system such as microfluidic.

DNA-based biosensors are potentially useful for many applications including medical diagnostics, environmental monitoring and food monitoring. This technique will enable online monitoring of quality control processes as well as point-of-care monitoring of patients. The portability of the whole sensor system will enable the decentralization of diagnostic processes where the consumer can use the detection system in the comfort of their home.

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The authors have nothing to disclose.


The authors would like to acknowledge the support of Universiti Putra Malaysia.


Name Company Catalog Number Comments
Acetic acid Merck 100056
Chloroform Merck 102445
Diaminoethane tetraacetic acid Promega E5134
Dibasic sodium phosphate  Sigma-Aldrich S9763
Disodium hydrogen phosphate Sigma-Aldrich 255793
Ethanol  Sigma-Aldrich 16368
Gold (III) chloride trihydrate Sigma-Aldrich 520918
Hydrochloric acid Merck 100317
Methylene blue Sigma-Aldrich M44907
Monobasic sodium phosphate, monohydrate Sigma-Aldrich S3522
Phosphate-buffered saline Sigma-Aldrich P5119
Poly(lactic acid) resin, commercial grade 4042D NatureWorks 4042D
Potassium chloride R&M Chemicals 59435
Potassium dihydrogen phosphate Sigma-Aldrich P9791
Potassium hexacyanoferrate III R&M Chemicals 208019
Sodium acetate anhydrous salt Sigma-Aldrich S2889
Sodium chloride Sigma-Aldrich S9888
Trisodium citrate Sigma-Aldrich S1804
Tris(hydroxymethyl) aminomethane Fisher Scientific T395-100
Tris-Base Fisher Scientific BP152-500
2X PCR Master Mix with Dual-Dye Norgen Biotek 28240
Agarose gel Merck 101236
Bolton Agar Merck 100079
Bolton Broth Merck 100079
CHROMagar Vibrio CHROMagar VB910
dNTPs Promega U1511
Nuclease-free water Thermo Scientific R0581
Eosin methylene blue agar  Merck 101347
GelRed Biotium 41001
Glycerol Merck 104092
Go Taq Buffer Promega M7911
Loading dye 100 bp DNA ladder Promega G2101
Loading dye 1kb DNA ladder Promega G5711
Magnesium chloride Promega 91176
Mannitol egg yolk polymyxin agar Merck 105267
McConkey Agar Merck 105465
Nutrient Broth Merck 105443
Taq polymerase Merck 71003
Trypticase Soy Broth Merck 105459
Trypticase Soy Agar Merck 105458



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