A ZnO–CNT nanocomposite based electrochemical DNA biosensor for meningitis detection

Abstract

A zinc oxide (ZnO) and multiwalled carbon nanotube (CNT) based nanocomposite matrix has been fabricated onto indium tin oxide (ITO) coated glass (ITO/glass) platform via. A chemical route. The thiolated oligonucleotides probe sequence consisting of 23 bases (single-stranded DNA) has been immobilized onto the ZnO–CNT/ITO electrode using a physical adsorption technique. The fabricated bioelectrode (ssDNA/ZnO–CNT/ITO) is used to specifically detect N. meningitidis with a high sensitivity of 20.20 μA per decade and 45 s of hybridization time by differential pulse voltammetry (DPV) using methylene blue as an electroactive indicator. The prepared DNA biosensor exhibits linearity in a wide range (5–180 ng μl⁻¹). The electrochemical impedance response behaviour of the ssDNA/ZnO–CNT/ITO bioelectrode with an increase in the target DNA concentration has also been studied using [Fe(CN)₆]^3−/4− as the hybridization indicator. The results indicate the possibility of fabricating a handheld device for meningitis detection at an early stage.

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1. Introduction

Meningitis is an inflammation of the membranes covering the brain and spinal cord, called the meninges. It is caused when germs infect the cerebral spinal fluid (CSF) that circulates around the brain and spinal cord. The infection can be caused by three kinds of germs namely virus, bacteria and fungus. Viral meningitis is the most common and the least severe type. Almost all patients suffering from viral meningitis, recover without any permanent damage, although full recovery might sometimes take many weeks. Bacterial meningitis is more aggressive and can lead to permanent damage or death in extreme cases. It is fatal in around 50% of cases if untreated, and accounts for several deaths globally each year.¹ The other causes is Fungal meningitis, which causes severe infections but occurs less frequently than viral or bacterial meningitis.

Meningitis can develop quickly, over a matter of hours. Until the cause of meningitis has been established, it should be regarded as a medical emergency requiring prompt diagnosis and urgent treatment. Meningitis diagnosis is carried out using a very popular technique called lumbar puncture (spinal tap). It involves inserting a needle into the middle of the lower back and collecting some drops of spinal fluid. The spinal fluid is then subjected to biochemical analysis to find out if there is any change in white blood cells count, sugar level or proteins. The fluid is stained using Gram staining^2,3 and also, bacteria obtained from these cerebral spinal fluid (CSF) samples can be grown on suitable culture medium which will confirm if the meninges are infected.⁴ Blood cultures are also used for carrying out diagnosis of this life-threatening disease. These tests are time consuming as well as very expensive. Certain imaging tests such as X-rays, CT scan, MRI and ultrasound are also employed to reveal any swelling or inflammation in different areas of body such as brain, chest or spinal cord but does not provide any confirmatory results that the infection is associated with meningitis.⁵ Latex agglutination test, immunological test, biochemical test and PCR are some other tests which are also used.^6–17 It can be seen that these tests involve utilization of costly machines, well trained manpower and are time consuming and they sometimes give non-confirmatory results.

Recently, nucleic acid biosensors based on electrochemical technique have emerged as a promising alternative as they offer fast, cost-effective, and simple detection.^18–20 Hence, an electrochemical DNA hybridization biosensor with low detection limit and high sensitivity is desired for specific DNA sequence detection.^21–23 In the recent past there have been few reports on detection of Neisseria meningitidis (N. meningitidis) using electrochemical DNA sensor.¹ Patel et al. have utilized gold surface to immobilize thiolated probe DNA. Though chemical bonding of the thiol group of DNA probe with gold was utilized to the immobilize DNA probe onto a gold surface by self-assembly but for further improving its sensitivity and selectivity, an immobilization platform which offers high adsorption ability, fast electron communication feature along with biocompatibility for higher stability of the biosensor is required.

Metal oxide thin films and nanostructures are known to have unique ability to promote faster electron transfer kinetics and offer excellent biosensing response characteristics towards a number of biomolecules. Among the metal oxides, nanostructured zinc oxide (ZnO) attained technological importance for biosensors because of its biocompatible properties including high catalytic efficiency, electron transfer characteristics, high chemical stability, high isoelectric point (∼9.5) and strong adsorption ability etc.²⁴ The high isoelectric point (IEP) of ZnO provides a platform to immobilize DNA having low isoelectric point (∼) through electrostatic interaction. Several reports are available in literature using ZnO for biosensing purposes like, Satriano et al. has reported utilization of ultrathin and nanostructured ZnO-based films for fluorescence biosensing; Khun et al. has utilized ZnO nanorods and thin films for biosensing applications and ZnO/chitosan hybrid nanostructure based biosensor was developed by Zhao et al.^25–27 All the above mentioned reports showed good biosensing results but in order to further improve the performance of a biosensor we need to incorporate some modifications into the matrix which helps in enhancing the biosensing characteristics. Recently, multiwalled carbon nanotubes (CNT), have attracted much attention for biosensors due to their unique structure, mechanical, electrical and thermal properties.^28–35 A number of reports are available on incorporating MWCNT for biosensing exhibiting excellent properties though stability and shelf-life are some of the issues.^36–38 The amalgamation of MWCNT with metal oxide (ZnO in the present case) can be an attractive route to obtain electronic properties based on morphological modification and electronic interactions between the two components which has not yet been much explored. Hence, ZnO–CNT based matrix has been fabricated in the present work using chemical route. The DNA probes for meningitis detection have been immobilized on the fabricated matrix by physical technique and the bioelectrode has been utilized for efficient detection of complementary N. meningitis DNA.

2. Experimental

2.1. Chemicals and reagents

N. meningitidis oligonucleotides probes, zinc acetate dihydrate (C₆H₆O₄Zn·2H₂O, 99.0%) and Triton X-100 are procured from Sigma-Aldrich (USA). Ammonia solution (min 25% GR) was obtained from Merck India Pvt. Ltd., India. Multiwalled carbon nanotubes (MWCNT, diameter 20–40 nm) are procured from Shenzhen Nanotech port Co., Ltd. Sodium phosphate monobasic anhydrous and sodium phosphate dibasic dihydrate, Tris buffer, ethylene diamine tetra acetic acid (EDTA) are obtained from Sisco chemical, India. All chemicals were used without further purification. 50 mM phosphate buffer saline (PBS), pH 7.0 (0.9% NaCl) solution is prepared by adjusting the proportion of monobasic sodium phosphate and dibasic sodium phosphate solution and then adding 0.9% NaCl to the solution. Deionized water (resistance ∼ 18.2 MΩ cm) is used for the preparation of aqueous solutions. All oligonucleotides solutions of varying concentrations are prepared in Tris–EDTA (TE) buffer (10 mM Tris, pH 8.0, 1 mM EDTA). All the oligonucleotide sequences used in the present work procured from Sigma-Aldrich and have been used as it is, without any further purifications or chemical treatment.

The various sequences of DNA oligomers used for carrying out electrochemical studies for DNA hybridization detection are tabulated below (Table 1):

Table 1 Various oligonucleotide sequences used in the present study

Probe DNA	5′-HS-GATACGAATGTGCAGCTGACACG-3′
Complementary target	3′-CTATGCTTACACGTCGACTGTGC-5′
Non-complementary target	3′-CTTAGCAAACTGGTGCACACACG-5′

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2.2. Preparation of ZnO–CNT (ZnO–CNT/ITO) electrode

The multiwalled carbon nanotubes (CNT) were functionalized by refluxing them in 1 : 3 v/v mixture of sulphuric acid (10 ml, 95–97%) and nitric acid (30 ml, 65%) followed by washing with deionized water until the pH of the solution became approximately 7.0 and then finally dried in oven at 60 °C.

0.455 M zinc acetate solution is prepared in distilled de-ionized water (D.I.) at room temperature. Ammonia solution (25%) was added dropwise to the above solution while constant stirring leading to the formation of milky white precipitate (pH = 9–10). After continuous stirring for 4–5 hours at room temperature, wash the precipitates well with D.I. water until neutral pH = 7.0 was obtained. Subsequently, 1 M dilute HNO₃ was added yielding a transparent solution (pH = 1). Functionalized CNTs (0.1 wt%) were added to the transparent sol and then ultrasonicated for 3 hours. The concentration of CNTs was decided based on our earlier work where 0.1 wt% CNTs in ZnO are found to be exhibiting efficient redox properties without being leached out. Ultrasonication of the ZnO–CNT composite solution for 1 hour and 2 hours was also done, however, after some time, the CNTs settled down at the bottom of the beaker containing composite solution resulting in the non-homogeneous solution which could not be used for film preparation. But, ultrasonication of the solution for 3 hours or higher resulted in formation of homogeneous solution of ZnO–CNT in which CNTs were uniformly dispersed. The solution thus obtained was spin coated on the ITO coated glass (ITO/glass) platform. Prior to spin coating, ITO coated glass was precleaned. In order to achieve uniform coating a surfactant Triton X-100 was also added to the prepared composite solution. The film so formed was pyrolysed for 10 minutes at 300 °C. The process was repeated several times for getting desired thickness. Finally, the prepared films were annealed at 300 °C for 3 hours in atmospheric air.

2.3. Fabrication of DNA biosensor

Prior to immobilization of single stranded DNA probe (ssDNA), the ZnO–CNT/ITO electrode was washed with D.I. water followed by drying at room temperature. 10 μl of DNA probe solution (ssDNA, 5 ng/10 μl) prepared in TE buffer solution (10 mM Tris, pH 8.0, 1 mM EDTA) was immobilized onto ZnO–CNT/ITO electrode and kept in a humid chamber for about 3 h at 25 °C. The unbound probe was removed by several washings with TE buffer and dried. These ssDNA/ZnO–CNT/ITO bioelectrodes were incubated with complementary and non-complementary target solutions in particular concentrations for 45 s at 25 °C for DNA hybridization reaction to occur. After hybridization the double stranded DNA (dsDNA) containing bioelectrode was again washed several times with buffer to remove the unbound ssDNA. The schematic of immobilization of probe and hybridization with target DNA has been shown in Scheme 1.

Scheme 1 Schematic representation of SH labeled probe immobilized on ZnO–CNT/ITO electrode and hybridization of complementary DNA with immobilized probe.

2.4. Experimental characterizations

The structural properties of ZnO–CNT films were studied using X-ray diffraction (XRD) [Bruker D8 Discover]. The Surface morphology of the ZnO–CNT/ITO electrode and ssDNA/ZnO–CNT/ITO bioelectrode was studied using scanning electron microscopy (SEM) [Quanta FEI 200]. Transmission electron microscope (TEM) [TECHNAI G² T30, u-TWIN] was employed to characterize the prepared ZnO–CNT nanocomposite solution. Cyclic voltammetric (CV) measurements were carried out on a potentiostat/galvanostat [Gamry Inc. reference 600] using a three-electrode system in PBS solution (50 mM, pH 7.0, 0.9% NaCl) containing 5 mM [Fe(CN)₆]^3−/4− with Ag/AgCl as the reference electrode. The hybridization studies of ssDNA/ZnO–CNT/ITO bioelectrode were carried out using Differential Pulse Voltammetry (DPV) studies in phosphate buffer saline (PBS, 0.05 M, pH 7.0, 0.9% NaCl) solution containing methylene blue as hybridization indicator (20 μM) for given concentration of target DNA.

3. Results and discussion

3.1. Transmission electron microscopy (TEM) studies

The transmission electron microscopy (TEM) has been adopted to characterize non-functionalized CNTs and a nanocomposite of functionalized CNTs co-existing with ZnO solution. TEM images of non-functionalized CNTs as shown in Fig. 1(a) indicates nanotubes of about 20 to 40 nm average diameter. In the nanocomposite of ZnO–CNT (Fig. 1(b)), it can be clearly seen that the inner walls are present in CNTs of 10 nm diameter. After functionalization decreased weight (loss of carbon content) of the CNTs is visible form TEM image indicating better hydrophilicity.³⁹ From TEM images of the ZnO–CNT nanocomposite (Fig. 1(b)), it appears that ZnO is attached non-uniformly onto the surface of the nanotubes at different places along its length. Furthermore, ZnO particles inside the tubes can also be seen in the TEM image of higher magnification (Fig. 1(c)). It can be clearly seen from this image that some black colour material (ZnO) is deposited onto the walls of the CNTs along its length. The mechanism for the formation of ZnO which is deposited on the CNTs is as follows:⁴⁰

NH₃ + H₂O ↔ NH⁴⁺ + OH⁻ (1)

Zn²⁺ + NH₃ → Zn(NH₃)₄²⁺ (2)

Zn(NH₃)₄²⁺ + OH⁻ → ZnO (3)

or Zn²⁺ can also form complex ion Zn(OH)₄²⁺ to give ZnO as follows:

Zn²⁺ + OH⁻ → Zn(OH)₄²⁺ (4)

Zn(OH)₄²⁺ → ZnO (5)

Fig. 1 TEM image of (a) non-functionalized bare CNTs and (b), (c) ZnO–CNT nanocomposite.

3.2. XRD studies

Fig. 2 shows the XRD spectra of ZnO–CNT nanocomposite thin film. The reflections from (100), (002), (101), (102) and (110) crystal planes of ZnO (Fig. 2) have been indexed under JCPDS card no. 36-1451 and corresponds to the formation of hexagonal wurtzite structure of ZnO and shows the polycrystalline nature of the deposited films. A small diffraction peak at 2θ ∼ 26° is also observed which corresponds to the (002) reflection from the graphitic planes of CNTs. The intensity of the diffraction peak is weak due to the very small concentration of multiwalled carbon nanotubes into ZnO. Similar XRD curves have been reported by Chen et al., also for the ZnO–CNT thin films grown by chemical route.⁴¹ The average crystallite size of ZnO in ZnO–CNT nanocomposite corresponding to (002) crystallite planes has been estimated to be 7 nm using Scherrer formula.⁴²

Fig. 2 XRD spectra of ZnO–CNT nanocomposite thin film.

3.3. Scanning electron microscopy (SEM) studies

Surface morphology of the ZnO–CNT/ITO and ssDNA/ZnO–CNT/ITO electrodes was studied using scanning electron microscope (SEM) and the results are shown in Fig. 3. The SEM images of ZnO–CNT/ITO electrode clearly indicate the clusters of CNTs and ZnO particles having nanostructured and highly porous surface (Fig. 3(a) and (b)). However, after immobilization of ssDNA on ZnO–CNT/ITO electrode, the surface morphology changes and becomes denser (Fig. 3(c) and (d)). This is attributed to the immobilization of thiolated DNA molecules present onto ZnO–CNT/ITO electrode.

Fig. 3 (a), (b) SEM images of ZnO–CNT/ITO electrode and (c), (d) ssDNA/ZnO–CNT/ITO bioelectrode.

3.4. Electrochemical studies

3.4.1 Cyclic voltammetric studies. Fig. 4 shows the cyclic voltammograms obtained for differently modified electrodes in PBS (50 mM, pH 7.0, 0.9% NaCl) containing 5 mM [Fe(CN)₆]^3−/4− at the scan rate of 100 mV s⁻¹. Curve (i) shows the CV curve obtained for bare ITO glass electrode. Well-defined redox peaks are observed with the anodic (E_pa) and cathodic (E_pc) peak potential of 0.47 V and 0.0 V, respectively and peak oxidation current about 417 μA (curve (i)). After the deposition of ZnO–CNT nanocomposite thin film onto ITO, the peak oxidation current is increases about 683 μA while the peak to peak potential separation (ΔE_p) is decreased significantly to 0.27 V (E_pa ∼ 0.37 V and E_pc ∼ 0.11 V) (curve (ii)), indicating a better redox behaviours of [Fe(CN)₆]^3−/4− on the ZnO–CNT/ITO electrode. This may be attributed to the presence of MWCNTs in ZnO–CNT nanocomposite thin film which provides a conducting path and rapid electron transfer between electrode and the electrolyte. When ssDNA is immobilized on the surface of ZnO–CNT nanocomposite electrode, the peak oxidation current reduces to 530 μA (curve (iii)). It is due to the fact that the self-assembled layer of ssDNA is insulating in nature and it blocks conducting sites of the ZnO–CNT/ITO electrode thus, reducing the peak oxidation current. When complementary target DNA is hybridized with the thiolated single stranded DNA immobilized on the ZnO–CNT nanocomposite matrix, a further decrease in the peak oxidation current to 240 μA is observed along with an increased ΔE_p to 0.39 V (E_pa ∼ 0.56 V and E_pc ∼ −0.17 V) (curve (iv)). This is in accordance with the earlier reports on DNA biosensor and is attributed to the fact that the introduction of complementary DNA increases the negative charge responsible for the increased repellence of redox species and thus obstructs the charge communication hence, reducing the current.

Fig. 4 Cyclic voltammetry curves of (i) ITO/glass platform, (ii) ZnO–CNT/ITO electrode, (iii) ssDNA/ZnO–CNT/ITO bioelectrode, and (iv) dsDNA/ZnO–CNT/ITO bioelectrode.

3.4.2 Electrochemical impedance studies. Electrochemical impedance analysis was carried out to study the charge transfer characteristics of the fabricated electrode and bioelectrodes. Fig. 5 shows the results of EIS measurements carried out as a function of frequency on ITO, ZnO–CNT/ITO electrode, ss DNA/ZnO–CNT/ITO and dsDNA/ZnO–CNT/ITO bioelectrodes in the presence of PBS (50 mM, pH 7.0, 0.9% NaCl) solution containing 5 mM [Fe(CN)₆]^3−/4−. The charge transfer resistance (R_ct) of the ITO electrode is found to be 390 Ω (curve (i)) which further decreases to 161 Ω for the ZnO–CNT/ITO electrode (curve (ii)). Lower value of R_ct obtained for the ZnO–CNT/ITO electrode indicates that the matrix provides an increased electroactive surface and thus facilitates the electron transfer. On the other hand, when ssDNA is immobilized onto ZnO–CNT/ITO electrode, R_ct value increases to 293 Ω (curve (iii)). This is attributed to the presence of electro-negative phosphate species in ssDNA that hinders the flow of ions from electrolyte to the electrode during redox reaction and hence the charge transfer characteristics. The results further confirm the immobilization of ssDNA on the ZnO–CNT/ITO electrode. On hybridization with the complementary DNA, R_ct of the dsDNA/ZnO–CNT/ITO bioelectrode increases to 458 Ω (curve (iv)). This is again due to the presence of the negatively charged surface of the dsDNA/ZnO–CNT/ITO bioelectrode that prevents [Fe(CN)₆]^3−/4− present in the electrolyte from reaching the electrode surface for electron exchange. The results are in accordance with the CV studies where a lower charge transfer resistance promotes the electron flow and hence gives a high peak oxidation current. An increased R_ct value is followed by the hindrance in charge transfer from electrolyte to the electrode and hence a low value for peak oxidation current.

Fig. 5 EIS curves of (i) ITO, (ii) ZnO–CNT/ITO electrode, (iii) ssDNA/ZnO–CNT/ITO bioelectrode, and (iv) dsDNA/ZnO–CNT/ITO bioelectrode.

3.5. Biosensor response studies

3.5.1 Amperometric response. Fig. 6(a) shows the results of differential pulse voltammetry (DPV) measurements carried out on the (i) ssDNA/ZnO–CNT/ITO (ii) non-complementary DNA/ZnO–CNT/ITO and (iii) dsDNA/ZnO–MWCNT/ITO bioelectrodes in 0.05 M PBS (pH 7.0, 0.9% NaCl) containing 20 μM methylene blue (MB) which is used as an electroactive redox indicator to investigate DNA hybridization at the electrode surface. MB is associated with the unpaired nitrogenous guanine bases on single stranded DNA (ssDNA) as compared to the double stranded DNA and is responsible for the current in DPV measurements.^43–45 It may be seen from Fig. 6(a) that the bioelectrode (ssDNA/ZnO–CNT/ITO) on hybridization with non-complementary DNA (non-complementary DNA/ZnO–CNT/ITO) does not show any change in the peak current, however, the peak current is seen to be drastically reduced to 120 μA on hybridization with the complementary DNA (dsDNA/ZnO–CNT/ITO). MB has strong affinity towards the free guanine bases at the ssDNA/ZnO–CNT/ITO bioelectrode giving a high peak current of 47 μA (curve i) in Fig. 6(a). When non-complementary DNA comes in contact with the ssDNA of meningitis, no hybridization takes place and hence the free guanine bases remain unchanged for MB giving almost similar value of peak current in DPV measurement (∼450 μA). We also tried to do sensing using different concentrations of non-complementary target DNA (80 ng μl⁻¹ and 100 ng μl⁻¹) which has two mutated nucleotides but no change in the DPV current was found after hybridization with the probe meningitis DNA which shows that the presently fabricated ZnO–CNT based DNA biosensor is highly sensitive as well as selective towards target meningitis DNA.

Fig. 6 (a) Differential pulse voltammograms of (i) ssDNA/ZnO–CNT/ITO bioelectrode, (ii) treated with non-complementary target DNA and (iii) complementary DNA in 0.05 M phosphate buffer of pH 7.0 containing 0.9% NaCl and methylene blue (MB, 20 μM); (b) response of the ssDNA/ZnO–CNT/ITO bioelectrode after hybridization with complementary target probe in varying concentration from 5 to 180 ng μl⁻¹; (c) plot of DPV peak current as a function of logarithm of target DNA concentration.

On the contrary, when complementary DNA comes in contact with the ssDNA, duplex formation takes place, reducing the free guanine bases and hence drastically reduce the peak current to 120 μA (Fig. 6(a)). The results indicate the specificity of the prepared bioelectrode (ssDNA/ZnO–CNT/ITO) towards complementary DNA only.

Fig. 6(b) shows the DPV response studies of ssDNA/ZnO–CNT/ITO bioelectrode after hybridization with different concentrations of target DNA of N. meningitidis varying from 5 ng μl⁻¹ to 180 ng μl⁻¹. It can be seen that the DPV peak current obtained for ssDNA/ZnO–CNT/ITO bioelectrode decreases with an increase in the target DNA concentration. As the concentration of target DNA increases, the number of the duplexes formed at the surface of the bioelectrode also increases leading to continuously decreasing free guanine bases and as a result peak current in DPV decreases. The magnitude of the peak DPV current obtained is plotted against the corresponding logarithmic concentration value of the complementary target sequence hybridized with the DNA bioelectrode (ssDNA/ZnO–CNT/ITO) in the range 5 ng μl⁻¹ to 180 ng μl⁻¹ and is found to exhibit good linearity.

Detection limit of the ssDNA/ZnO–CNT/ITO bioelectrode is found to be 5 ng μl⁻¹ as for concentrations lower than this no decrease in DPV current has been observed indicating that no significant hybridization takes place. It is found that 45 s is the minimum required hybridization time for the given reaction to take place. Sensitivity of the ssDNA/ZnO–CNT/ITO bioelectrode towards target DNA is calculated from the linearity curve and is found to be 20.20 μA per decade along with wide linear range (5–180 ng μl⁻¹), indicating that the ZnO–CNT nanocomposite based matrix has significantly enhanced the performance of nucleic acid biosensor the sensitivity of the biosensor is found to be much superior to already reported value for meningitis detection using metal/metal oxide based matrices.¹ This is attributed to the high surface provided by the nanocomposite matrix to the probe DNA loading and efficient electron communication feature of the prepared matrix. The ss DNA/ZnO–CNT/ITO electrode is found to be giving similar results within ±5% for about 20 weeks when stored at 4 °C indicating high stability of the prepared bioelectrode. Reusability of the ssDNA/ZnO–CNT/ITO bioelectrode has been estimated by dipping dsDNA/ZnO–CNT/ITO electrode in hot water (80 °C) for 5 minutes followed by cooling in an ice bath and then washing with double deionized water (DDI) prior to reuse. The DPV response obtained for the regenerated bioelectrode showed almost similar magnitude of the peak current as that obtained for the ssDNA/ZnO–CNT/ITO prepared bioelectrode prior to hybridization thus ensuring good reusability of the ZnO–CNT nanocomposite based bioelectrode. The ZnO–CNT nanocomposite based provides desirable platform for single stranded probe DNA to immobilize with better orientation thus, giving good stability and improved electron transfer kinetics between the electrolyte and electrode. Table 3 compares the presently fabricated ZnO–CNT based DNA biosensor with earlier reported meningitis detection based DNA biosensors which highlights the importance of the present work.

Table 2 Calculated value of surface concentration of the target DNA onto ssDNA/ZnO–CNT/ITO bioelectrode for the corresponding target DNA concentrations

Target DNA concentration (ng μl^−1)	10	20	40	80	100	140
Surface concentration θ	0.42	0.54	0.68	0.73	0.78	0.80

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Table 3 Comparison table for meningitis DNA based biosensors

Matrix/DNA detected	Detection range (ng μl^−1)	Hybridization time (s)	Stability (months)	Detection technique	Ref.
Gold/meningitis	7–42	120	4	CV	48
Gold/meningitis	10–60	60	4	CV	1
ZnO–CNT nanocomposite/meningitis	5–180	45	5	EIS, DPV	Present work

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3.5.2 Electrochemical impedance spectroscopy (EIS) response studies. The presence of N. meningitidis is also investigated in different concentrations of the target DNA using electrochemical impedance spectroscopy (EIS). Fig. 7(a) shows the impedimetric response of the ssDNA/ZnO–CNT/ITO bioelectrode on hybridization with different concentration of target DNA. It can be seen that diameter of the semicircle indicating the charge transfer resistance, R_ct, keeps on increasing along with the concentration of target DNA. As the concentration of the target DNA increases, more hybridization occurs with the probe DNA immobilized onto the surface of the immobilized nanocomposite matrix. This results in an increase in the concentration of negatively charged phosphate group on the DNA and hence the electrostatic repulsion between these groups and [Fe(CN)₆]^3−/4− ions in electrolyte hinders the flow of charge carriers and increasing the charge transfer resistance. This is in accordance with the earlier reported results.⁴⁶ The charge-transfer resistance, R_ct, continues to increase until a target DNA concentration of 140 ng μl⁻¹ is reached and thereafter it saturates (not shown in Fig. 7(a)). A plot of R_ct as a function of target DNA concentration (Fig. 7(b)), shows that the charge-transfer resistance (R_ct) increases linearly from 495 Ω to 1504 Ω within 5–140 ng μl⁻¹ concentration range of target DNA. The sensitivity of the prepared DNA biosensor calculated from the slope of the linear graph is found to be 7.51 Ω ng⁻¹ μl⁻¹ with a regression coefficient, r = 0.98.

Fig. 7 (a) EIS response of ssDNA/ZnO–CNT/ITO bioelectrode on hybridization with different DNA concentration from 10–140 ng μl⁻¹ and (b) plot of charge transfer resistance (R_ct) with concentration of the target DNA.

The EIS spectra can be used calculated the fraction of the target DNA molecules efficiently hybridized onto the ssDNA/ZnO–CNT/ITO bioelectrode surface known as the surface coverage (θ). The value of ‘θ’ can be calculated using the following eqn:⁴⁷

where ‘R_o’ and ‘R_i’ are is the charge transfer resistance of the ssDNA/ZnO–CNT/ITO bioelectrode and after hybridization with a particular concentration of the target DNA concentration respectively. The values of R_o and R_i are obtained from the EIS measurements (Fig. 7(a)). The values of θ is calculated for different concentrations of the target DNA sequence and are listed in Table 2. It may be seen that the value of θ increases with increasing target DNA concentrations and reaches almost saturated value of 0.80 at the concentration of 140 ng μl⁻¹ indicating that the maximum surface coverage by target DNA strands onto the surface of ssDNA/ZnO–CNT/ITO bioelectrode is about 80% for 140 ng μl⁻¹ of target DNA. The results supported the earlier observations on DPV measurements that the nanocomposite ZnO–CNT matrix support the favourable immobilization of ssDNA probe which give rise to enhanced hybridization with the target DNA.

4. Conclusion

The ZnO–CNT nanocomposite matrix offers an efficient platform for the fabrication of highly sensitive nucleic acid biosensor for detection of N. meningitidis. The fabricated DNA biosensor exhibits low detection limit of target DNA concentration (5 ng μl⁻¹), wide detection range from 5 ng μl⁻¹ to 180 ng μl⁻¹ with a very fast hybridization time of 45 s and a high shelf-life of 20 weeks. The bioelectrode exhibits high sensitivity of 7.51 Ω ng⁻¹ μl⁻¹ and 20.20 μA per decade as measured from the DPV and EIS measurements. EIS measurements highlights the fact that the prepared matrix provides a high surface coverage of 80% towards 140 ng μl⁻¹ concentration of target DNA. The results are much superior to the earlier reports on meningitis nucleic acid biosensors because of the fabricated nanocomposite ZnO–CNT nanocomposite matrix. The results obtained for both amperometric as well as impedimetric DNA sensing points to the fact that, in future, this useful nanocomposite can be utilized for detection of other important bacterial infectious diseases.

Acknowledgements

Authors are thankful to DST and DBT for financial support to carry out this work. One of the authors (MT) is thankful to UGC for granting research fellowship.

References

1. M. K. Patel, P. R. Solanki, A. Kumar, S. Khare, S. Gupta and B. D. Malhotra, Biosens. Bioelectron., 2010, 25, 2586.
2. L. D. Gray and D. P. Fedorko, Clin. Microbiol. Rev., 1992, 5, 130.
3. S. A. Dunbar, R. A. Eason, D. M. Musher and J. E. Clarridge, J. Clin. Microbiol., 1998, 36, 1617–1620.
4. N. Deivanayagam, T. P. Ashok, K. Nedunchelian, S. S. Ahamed and N. Mala, J. Trop. Pediatr., 1993, 39, 284.
5. C. T. Tan and B. B. Kuan, Neuroradiology, 1987, 29, 43.
6. F. O. Finlay, H. Witherow and P. T. Rudd, Arch. Dis. Child., 1995, 73, 160–161.
7. J. E. Sippel, P. A. Hider, G. Controni, K. D. Eisenach, H. R. Hill, M. W. Rytel and B. L. Wasilauskas, J. Clin. Microbiol., 1984, 20, 884–886.
8. K. Surinder, K. Bineet and M. Megha, Indian J. Med. Microbiol., 2007, 25, 395–397.
9. R. Bortolussi, A. J. Wort and S. Casey, Can. Med. Assoc. J., 1982, 127, 489.
10. M. Kilian, I. Sorensen and W. Frederiksen, J. Clin. Microbiol., 1979, 9, 409.
11. P. Radstrom, A. Backman, N. Qian, P. Kragsbjerg, C. Pahlson and P. Olcen, J. Clin. Microbiol., 1994, 32, 2738–2744.
12. J. Neewcombe, K. Cartwright, W. H. Palmer and J. Mcfadden, J. Clin. Microbiol., 1996, 34, 1637–1640.
13. P. Kotilinen, J. Jalava, O. Meurman, O. P. Lehtonen, E. Rintala, O. Pekka, S. La, E. Eerola and S. Nikkari, J. Clin. Microbiol., 1998, 36, 2205.
14. L. F. Baethegan, C. Moraes, L. Weidlich, S. Rios, C. I. Kmetzsh, M. S. N. Silva, M. L. R. Rossetti and A. Zaha, J. Med. Microbiol., 2003, 52, 793.
15. D. C. Richardson, L. Louie, M. Louie and A. E. Simor, J. Clin. Microbiol., 2003, 41, 3851.
16. D. E. Bennett, R. M. Mulhall and M. T. Cafferkey, J. Clin. Microbiol., 2004, 42, 1764.
17. E. D. Carrol, A. P. J. Thomson, P. Shears, S. J. Gray, E. B. Kaczmarski and C. A. Hart, Arch. Dis. Child., 2003, 83, 271.
18. S. R. Mikkelsen, Electroanalysis, 1996, 8, 15–19.
19. E. Palecek and M. Fojta, Anal. Chem., 2001, 73, 74–83.
20. J. Wang, Chemistry, 1999, 5, 1681–1685.
21. T. G. Drummond, M. G. Hill and J. K. Barton, Nature Biotechnology, 2003, 21, 1192–1199.
22. P. Kavanagh and D. Leech, Anal. Chem., 2006, 78, 2710–2716.
23. C. Liu, Y. Hsieh, C. Huang, Z. Lin and H. Chang, Chem. Commun., 2008, 2242–2244.
24. S. Saha, V. Gupta, K. Sreenivas, H. H. Tan and C. Jagadish, Appl. Phys. Lett., 2010, 97, 133704.
25. C. Satriano, M. E. Fragalà and Y. Aleeva, Sens. Actuators, B, 2012, 161, 191–197.
26. K. Khun, Z. H. Ibupoto, C. O. Chey, J. Lu, O. Nur and M. Willander, Appl. Surf. Sci., 2013, 268, 37–43.
27. M. Zhao, J. Huang, Y. Zhou, Q. Chen, X. Pan, H. He and Z. Ye, Biosens. Bioelectron., 2013, 43, 226–230.
28. M. Gao, L. Dai and G. G. Wallace, Electroanalysis, 2003, 15, 1089–1094.
29. J. Wang, Electroanalysis, 2005, 17, 7–14.
30. M. J. O'Connell, S. M. Bachilo, C. B. Huffman, V. C. Moore, M. S. Strano, E. H. Haroz, K. L. Rialon, P. J. Boul, W. H. Noon, C. Kittrell, J. Ma, R. H. Hauge, R. B. Weisman and R. E. Smalley, Science, 2002, 297, 593–596.
31. K. Balasubramanian and M. Burghard, Anal. Bioanal. Chem., 2006, 385, 452–468.
32. M. Zhang, A. Smith and W. Gorski, Anal. Chem., 2004, 76, 5045–5050.
33. S. G. Wang, Q. Zhang, R. Wang and S. F. Yoon, Biochem. Biophys. Res. Commun., 2003, 311, 572–576.
34. Y. Lin, F. Lu, Y. Tu and Z. Ren, Nano Lett., 2004, 4, 191–195.
35. B. Kim and W. M. Sigmund, Langmuir, 2004, 20, 8239–8242.
36. F. Gutierrez, M. D. Rubianes and G. A. Rivas, Sens. Actuators, B, 2012, 161, 191–197.
37. H. Teymourian, A. Salimi and R. Hallaj, Talanta, 2012, 90, 91–98.
38. A. Wong, M. Del Pilar and T. Sotomayor, Sens. Actuators, B, 2013, 181, 332–339.
39. R. Yudianti, H. Onggo, Y. S. Sudirman, Y. Saito, T. Iwata and J. Azuma, Open Mater. Sci. J., 2011, 5, 242–247.
40. S. Baruah and J. Dutta, Sci. Technol. Adv. Mater., 2009, 10, 013001.
41. H. Chen, S. Lin and K. Huang, Appl. Opt., 2014, 53, 242–247.
42. V. Gupta and A. Mansingh, J. Appl. Phys., 1996, 80, 1063.
43. R. Singh, G. Sumana, R. Verma, S. Sood, M. K. Pandey, R. K. Gupta and B. D. Malhotra, J. Biotechnol., 2010, 150, 357–365.
44. W. Yang, M. Ozsoz, D. B. Hibbert and J. J. Gooding, Electroanalysis, 2002, 14, 1299.
45. A. Erdem, K. Kerman, B. Meric, U. S. Akarca and M. Ozsoz, Anal. Chim. Acta, 2000, 422, 139.
46. I. Szymańska, H. Radecka, J. Radecki and R. Kalisza, Electrochem. Commun., 2009, 11, 969–973.
47. I. Szymańska, H. Radecka, J. Radecki and R. Kalisza, Biosens. Bioelectron., 2007, 22, 1955–1960.
48. M. K. Patel, P. R. Solanki, S. Seth, S. Gupta, S. Khare, A. Kumar and B. D. Malhotra, Electrochem. Commun., 2009, 11, 969–973.

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