ELECTROCHEMICAL SENSOR

Abstract

An electrochemical sensor comprising first and second electrodes. The first electrode has a molecular imprinted polymer (MIP) immobilised thereon and the MIP comprises a redox label and is imprinted with an analyte. The disclosure also provides methods of producing such sensors, methods of detecting and determining the concentration of analytes and the use of the electrochemical sensors for the detection of analytes.

Claims

1. An electrochemical sensor comprising first and second electrodes, wherein the first electrode has a molecular imprinted polymer (MIP) immobilized thereon and the MIP comprises a redox label and is imprinted with an analyte.

2. The electrochemical sensor of claim 1, wherein the MIP is a bulk MIP, microMIP, or nanoMIP.

3. The electrochemical sensor according to claim 1, wherein the analyte is selected from the group consisting of: a bioanalyte, a pharmaceutical, a pesticide, and an explosive.

4. The electrochemical sensor according to claim 1, wherein the analyte is selected from the group consisting of; a biological receptor, a nucleic acid, a cell, a spore, a virus, a microorganism, a tissue sample, a carbohydrate, a oligosaccharide, a polysaccharide, a protein, a peptide, a nucleoprotein, a mucoprotein, a lipoprotein, a synthetic protein, a glycoprotein, a glucosaminoglycan, a steroid, a hormone, an immunosuppressant, heparin, an antibiotic, a vitamin, a biomarker of a pathological or disease state, a toxin, a pesticide, an herbicide, an insecticide, a fungicide, an explosive, a nerve agent, a pollutant, an endocrine disrupting compound, a nucleotide, a nucleoside, an oligonucleotide, a metabolite, a secondary metabolite, a drug metabolite, a drug intermediate, and a drug.

5. The electrochemical sensor according to claim 1, wherein the redox label is a dye, a quinone, a free transition metal, a conjugated π-electron molecule, a complex with d- and f-electrons, a carbon nanotube, or graphene.

6. The electrochemical sensor according to claim 1, wherein the redox label is a transition metal compound or complex.

7. The electrochemical sensor according to claim 6, wherein the redox label comprises a metallocene and/or a ferrocene.

8. The electrochemical sensor according to claim 7, wherein the redox label comprises ferrocenylmethyl methacrylate (FMMA).

9. The electrochemical sensor according to claim 1, wherein the MIP is configured to swell and/or shrink upon binding of the analyte.

10. The electrochemical sensor according to claim 1, wherein the MIP is immobilized on a surface of the first electrode due to the presence of one or more covalent bonds, coordinate bonds, or mechanical bonds.

11. The electrochemical sensor according to claim 1, wherein the electrochemical sensor is configured to detect the concentration of redox label moieties exposed to the first electrode by detecting a signal response.

12. An electrode with a molecular imprinted polymer (MIP) immobilized thereon, wherein the MIP comprises a redox label and is imprinted with an analyte.

13. The method of producing the electrode of claim 12, wherein the method further comprises: polymerizing a plurality of monomers in the presence of an analyte to form a polymer; extracting the analyte from the polymer to expose analyte binding sites in the polymer, thereby providing a MIP; and immobilizing the imprinted MIP onto an electrode to produce the electrode of the second aspect, wherein the plurality of monomers are polymerized in the presence of a redox label, or the polymer is contacted with a redox label after polymerized.

14. The method of claim 13, wherein the immobilizing step comprises; mixing the MIP with conductive ink; and immobilizing the MIP-conductive ink mixture onto the surface of the electrode by screen printing, inkjet printing, or 3D printing.

15. The method of claim 13, wherein the polymerization is a living radical polymerization, living anionic or cationic polymerization or controlled polycondensation.

16. The method according to claim 13, wherein the plurality of monomers comprise one or more monomers selected from the group consisting of: vinyl monomers, allyl monomers, acetylenes, acrylates, methacrylates, acrylamides, methacrylamides, chloroacrylates, itaconates, trifluoromethylacrylates, derivatives of amino acids, nucleosides, nucleotides, and carbohydrates.

17. The method according to claim 13, wherein the plurality of monomers are co-polymerized in the presence of the redox label, and wherein the redox label is a molecule comprising a double or triple bond.

18. A method of detecting an analyte, the method comprising: disposing first and second electrodes in a solution comprising an analyte, wherein the first electrode has a molecular imprinted polymer (MIP) immobilized thereon and the MIP comprises a redox label and is imprinted with the analyte; allowing the analyte to bind to the MIP; and measuring a signal response to confirm the presence of the analyte.

19. The method according to claim 18, further comprising the step of comparing the signal response to one or more signal responses caused by a known concentration of the analyte and thereby determining the concentration of the analyte.

20. (canceled)

21. The method according to claim 11, wherein the signal response is a change in potential difference, impedance, capacitance, and/or current between the first and second electrodes.

Description

[0069] For a better understanding of the invention, and to show how embodiments of the same may be carried into effect, reference will now be made, by way of example, to the accompanying Figures, in which:—

[0070] FIG. 1 shows a schematic representation of the change in polymer conformation triggered by interaction with a template;

[0071] FIG. 2 shows a schematic representation of the template-induced change in the quantity of anchored redox markers exposed on an electrode surface;

[0072] FIG. 3a shows the current response of the nanoMIPs with anchored ferrocenylmethyl methacrylate (FMMA) to N-butyryl-L-homoserine lactone (C.sub.4—HSL), wherein the concentration of C.sub.4-HSL is (a) 0 μM, (b) 0.1 μM, (c) 0.2 μM, (d) 0.4 μM, (e) 0.8 μM, (f) 1.6 μM, (g) 3.2 μM, (h) 6.2 μM, (i) 12.5 μM, (j) 25 μM, or (k) 50 μM. The redox potential was measured at 0.23 V (vs AgCl) on screen printed electrodes; and FIG. 3b shows calibration plots showing the sensor response of nanoMIPs with anchored FMMA to NAHSLs in the concentration range 6.25-400 μM, wherein the NAHSLs are (a) C.sub.4-HSL, (b) C6-HSL, (c) GBL and (d) 3-oxo-C6-HSL;

[0073] FIG. 4a shows DPV measurements on a nanoMIPs modified gold electrode for trypsin solution, wherein the concentration of trypsin in 5 mM PBS is (a) 0 μM, (b) 0.006 μM, (c) 0.012 μM, (d) 0.025 μM, (e) 0.05 μM or (f) 100 μM; and FIG. 4b shows the response of nanoMIPs using DPV measurements for (1) Trypsin, (2) Avidin and (3) Pepsin in a concentration ranging from 6.5 to 100 nM in 5 mM PBS;

[0074] FIG. 5a shows DPV measurements on nanoMIPs immobilized on gold electrodes, wherein the concentration of DPV in 5 mM PBS is (a) 0.8 mM, (b) 1.6 mM, (c) 3.2 mM, (d) 6 mM, (e) 12.5 mM, (f) 25 mM, or (g) 50 mM; and FIG. 5b shows calibration curves for glucose, response for the (a) nanoMIPs imprinted with glucose and (b) nanoMIP imprinted with dopamine; and

[0075] FIG. 6 shows a nanoMIPs glucose sensor response for (a) glucose, (b) fructose, (c) maltose and (d) lactose in linear concentration range from 0.8 to 50 mM in 5 mM PBS

EXAMPLE 1—PREPARATION OF NANOMIPS VIA SOLID-PHASE SYNTHESIS

[0076] 200 g of glass beads were incubated in boiling 4.0 M sodium hydroxide for 15 min, washed with deionised water and dried at 150° C. Activated glass beads were incubated for 10 hour in a solution 2% (v/v) of N-[3-(trimethoxysilyl)propyl]ethylenediamine (DAMO), and 0.33% of 1,2-bis(triethoxysilyl)ethane (BTSE) in toluene at 70° C., washed with acetone and dried at 150° C. Glass beads were immersed into 0.1 M MES buffer, pH 6.0 containing 0.01 M N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC) and 3.6 mM N-hydroxysuccinimide (NHS), and mixed with 0.03 M dodecanedioic acid dissolved in DMF for 20 min. After that beads were washed with 50% DMF and acetone. The template was immobilised on the glass beads by incubation in 0.43 mM (S)-(−)-α-amino-γ-butyrolactone hydrobromide and 0.43 mM of sodium carbonate. Glass beads with immobilised template were washed with water and dried under vacuum. Monomer mixture was prepared by mixing 1.44 g of methacrylic acid (MAA), 1.62 g of ethylene glycol dimethacrylate (EGDMA), 1.62 g of trimethylolpropane trimethacrylate (TRIM), 0.37 g of N,N-diethyldithiocarbamic acid benzyl ester (DABE), 0.09 g of pentaerythritol-tetrakis-(3-mercaptopropionate) (PETMP), and 0.14 g of ferrocenylmethyl methacrylate (FMMA) and deoxygenating with nitrogen for 10 min. Glass beads with immobilised template (25 g) were degassed in vacuum for 20 min and coated with monomer mixture. Polymerisation was initiated by exposing the mixture to UV-light for 2 min (Philips model HB/171/A, 4×15 W/amps). After polymerisation, the crude of reaction was transferred into a solid phase extraction (SPE) cartridge fitted with a polyethylene frit (20 mm porosity) and washed with cold acetonitrile at 0° C. in order to remove monomers, residues and low affinity nanoparticles. The high affinity nanoMIPs were extracted by elution at 6° C.

[0077] In examples 2 to 4 the template was tetrahydrocannabinol (THC), morphine or N-butyryl-L-homoserine lactone (C.sub.4—HSL).

EXAMPLE 2—DLS ANALYSIS OF NANOMIPS

[0078] 700 μL of nanoMIPs (0.02 mg/mL) in water, prepared according to example 1, which were imprinted with either THC (THC nanoMIPs) or morphine (morphine nanoMIPs) were mixed with 350 μL of water, and optionally 95 μM THC or morphine and briefly sonicated. The hydrodynamic size of nanoMIPs was measured by dynamic light scattering (DIS) with a ZetaSizer Nano ZS (Malvern Instruments Inc, UK). NanoMIPs conformational changes triggered by the analyte are shown in Table 1.

TABLE-US-00001 TABLE 1 DLS results for nanoMIPs and nanoNIPs in solution and loaded with THC and Morphine molecules. Diameter Nanoparticle sample (nm) PDI* THC nanoMIPs 307.3 ± 2.5 0.323 THC nanoMIPs in the presence of THC 379.2 ± 6.4 0.295 THC nanoMIPs in the presence of morphine 265.8 ± 3.2 0.261 Morphine nanoMIPs 195.4 ± 4.4 0.156 Morphine nanoMIPs in the presence of THC 179.3 ± 8.7 0.278 Morphine nanoMIPs in the presence of morphine 208.3 ± 4.5 0.363 *Polydispersity index (PDI)

[0079] As shown in the table, the diameter of the nanoMIPs increases in the presence of the molecule with which it has been imprinted, i.e. the template molecule. This is shown schematically in FIGS. 1 and 2.

EXAMPLE 3—IMMOBILIZATION OF NANOMIPS

[0080] NanoMIPs prepared according to example 1, were immobilised on gold electrodes using one of two methods.

[0081] (i) Gold electrodes (Drop-Sense gold electrodes DRP-250AT (aux.: Pt ref.: AgCl) were functionalized by incubation in 3 mM solution of crystamine in ethanol for 8 hours. Functionalised electrodes were incubated for 30 min in 100 μL solution containing 0.03 mg/mL of nanoMIPs, 0.4 M N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC) and 0.1M N-hydroxysuccinimide (NHS) in 50 mM PBS.

[0082] (ii) NanoMIPs (3.6 mg/mL in acetonitrile) were mixed with 200 mg of gold polymer electrode paste (C2041206P2, Gwent Group U.K.) and spread over gold electrodes (Drop-Sense gold electrodes DRP-250AT (aux.: Pt ref.: AgCl). Electrodes were cured at 8° C. for 30 minutes.

EXAMPLE 4—ELECTROCHEMICAL MEASUREMENTS OF SMALL MOLECULES

[0083] NanoMIPs imprinted with C.sub.4-HSL were prepared according to example 1 and immobilised on gold electrodes by covalent attachment according to example 3.

[0084] All experiments were carried out using an Autolab11 Instruments, Netherland. Differential pulse voltammetry (DVP) was performed under the following experimental conditions: potential was recorded in the range of −0.4 to +0.8 V and the step potential and modulation amplitude at 50 mV. The current signal was measured at the redox potential of the redox marker in nanoMIPs (0.22 V vs AgCl).

[0085] DPV responses of nanoMIP-coated electrodes to C.sub.4-HSL solutions at concentrations between 0 and 50 μM are presented in FIG. 3a. The nanoMIPs with anchored FMMA showed good response to the template with a sensitivity of 47.5 μA/μM (R.sup.2=0.998) and LOD of 0.13 μM. The sensor response of nanoMIPs-coated electrode to different homoserine lactones is shown in FIG. 3b. The calibration plots demonstrate good selectivity of sensor for C.sub.4-HSL.

EXAMPLE 5—SYNTHESIS OF NANOMIPS IMPRINTED WITH TRYPSIN

[0086] For the polymerization mixture the following monomers were dissolved in 100 ml of water: 39 mg (0.214 mmol) of N-isopropylacrylamide (NIPAM), 6 mg (0.078 mmol) of N,N-methylene-bis-acrylamide (MBA), 2.2 μL (0.0224 mmol) of acrylic acid (AAc), 6 mg (0.0211 mmol) of ferrocenylmethyl methacrylate (FMAA) and 5.8 mg (0.0325 mmol) of N-(3-aminopropyl) methacrylamide hydrochloride (NAPMA). Additionally, 33 mg (0.264 mmol) of tert-butyl acrylamide (TBAM) dissolved in 2 mL ethanol were added to the aqueous mixture. The solution was sonicated for 5 min, and then purged with nitrogen for 30 min. 50 mL of the polymerization mixture were added to 60 g of trypsin-derivatized glass beads. Polymerization was initiated by addition of 0.5 mL of (60 mg/mL) ammonium persulfate and (30 μL/mL) tetramethylethylenediamine and continued at room temperature for 1 hour. Beads were decanted into solid phase extraction cartridge and washed with water (10 bead volumes, 50 mL). The SPE cartridge was then placed in water bath at 60° C. for 7 min and high affinity nanoMIPs eluted with water at 6° C. (5 bead volumes, 20 mL). The concentration of the nanoMIPs fractions was evaluated by freeze-drying of solution aliquots and weighing.

EXAMPLE 6—ELECTROCHEMICAL MEASUREMENTS OF TRYPSIN

[0087] All experiments were carried out using an Autolab11 Instruments, Netherland using a gold electrode prepared using nanoMIPs prepared according to example 5 and the method described in Example 3(i). Differential pulse voltammetry (DVP) was performed under the following experimental conditions: potential was recorded in the range of −0.4 to +0.8 V and the step potential and modulation amplitude at 50 mV. The current signal was measured at the redox potential of the redox marker in nanoMIPs (0.22 V vs AgCl).

[0088] DPV responses of nanoMIP-coated electrodes to trypsin solutions are presented in FIG. 4a. The calibration curve shows the good sensitivity for trypsin 0.25 nM/μA (R.sup.2=0.998) and LOD of 0.15 μM. The responses from avidin (3.95×10.sup.−8 nM, R.sup.2=0.897) and pepsin (8.8×10.sup.−8 nM, R.sup.2=0.577) are negligible, six orders of magnitude lower compared to the trypsin as shown in FIG. 4b. This shows that the system exhibits good sensitivity for trypsin.

EXAMPLE 7—SYNTHESIS OF NANOMIPS IMPRINTED WITH GLUCOSE

[0089] The monomer mixture was composed by 39 mg (0.214 mmol) of N-isopropylacrylamide (NIPAM), 6 mg (0.078 mmol) of N,N-methylene-bis-acrylamide (MBA), 33 mg (0.264 mmol) of tert-butyl acrylamide (TBAM), 2.2 μL (0.022 mmol) acrylic acid (AAc), 39 mg of N-iopropylacrylamide (NIPAM), 7 mg (0.028 mmol) of ferrocenylmethyl methacrylate (FMAA), 5.8 mg (0.033 mmol) of N-(3-aminopropyl) methacrylamide hydrochloride (NAPMA) and 30 mg (0.167 mmol) of glucose. The components were dissolved in 100 mL 2% ethanol in water (v/v), sonicated for 5 min, then degassed using nitrogen for 30 min. The polymerisation was initiated by the addition of a solution comprising 30 mg (0.132 mmol) ammonium persulfate (APS) and 30 μL (0.2 μmol) of N,N,N′,N′-tetramethylethane-1,2-diamine (TEMED) in 0.5 mL of water. The monomer mixture was allowed to polymerize at ambient temperature (25° C.) for 1 hour. Subsequently, MIP was washed with water 7 times in a centrifuge cartridge filter (10 kDa) to remove template and unreacted monomers. Next, the fractions of high affinity nanoparticles were collected.

EXAMPLE 8—ELECTROCHEMICAL MEASUREMENTS OF GLUCOSE

[0090] All experiments were carried out using an Autolab11 Instruments, Netherland using a gold electrode prepared using nanoMIPs prepared according to example 7 and the method described in Example 3(i). Differential pulse voltammetry (DVP) was performed under the following experimental conditions: potential was recorded in the range of −0.4 to +0.8 V and the step potential and modulation amplitude at 50 mV. The current signal was measured at the redox potential of the redox marker in nanoMIPs (0.22 V vs AgCl).

[0091] DPV responses of nanoMIP-coated electrodes to glucose solutions are presented in FIG. 5a. For control measurements a polymer imprinted with dopamine, which was prepared using the method of example 7 but replacing the glucose with dopamine, was used. The control polymer known as non-specific imprinted polymer (NIP) reflects as expected a negligible response (24 times lower), see FIG. 5b. Additionally, the glucose sensor does not show high level of cross reactivity for fructose, maltose and lactose, see FIG. 6.

CONCLUSION

[0092] The inventors have shown that it is possible to synthesise MIPs imprinted with an analyte and comprising ferrocenylmethyl methacrylate (FMAA), a redox label. The inventors have shown that the diameter of the MIPs changes depending upon whether or not the analyte is present, as shown schematically in FIG. 1.

[0093] The MIPs may be immobilised on a first electrode. The first electrode and a second electrode may then be placed in a solution and a potential difference applied across them. As the concentration of the analyte varies the volume of the MIPs also vary, changing the number of redox labels which contact the electrode, as shown in FIG. 2. This in turn causes the current which passes between the first and second electrodes to vary, allowing the concentration of the analyte to be accurately calculated.

REFERENCES

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