**9.2. NAD and ferrocene polymeric mediator redox couple**

As a coenzyme for hundreds of oxidoreductases, NAD has attracted great attention for biosensor development. Because its direct oxidation at electrode surface requires high applied potential and involves side reactions leading to the formation of electrochemically inactive by-products (see the preceding section), NAD is normally coupled with mediators for its application in biosensors. In the literature, ferrocenium and its derivatives have been reported as mediator for oxidation of NADH to NAD+. [60-63] As mentioned in the preceding section, small molecular mediators are not normally suitable for continuous monitoring biosensors. The effective immobilization of the ferrocene polymeric mediator in the screenprinted carbon electrode developed at LifeScan Scotland Limited holds potentials for development of continuous monitoring biosensors using NAD-dependant oxidoreductases. To investigate electrochemical communication between NAD and the immobilized ferrocene polymeric mediator, 0.3 V potential (ECOChemie PGSTat Autolab, Type III) was applied to an electrochemical cell containing 3.0 mL 0.01M pH7.4 PBS and equipped with a screen-printed carbon electrode (approximately dimension length x width = 10 x 5 mm) with immobilized ferrocene polymeric mediator (about 12.5% wt/wt), an Ag/AgCl reference electrode and a platinum counter electrode to oxidize the ferrocene moieties of the polymeric mediator to ferrocenium moieties. After the current of ferrocene oxidation reached a steady level, PBS in the cell was replaced with 0.3 mM NADH (from Sigma-Aldrich) in 3.0 mL 0.01M pH7.4 and a step increase in current was detected, which indicated oxidation of NADH by the ferrocenium moieties of the polymeric mediator. The NADH oxidation was followed by measuring absorbance of the solution at 340 nm wavelength by UV/vis spectroscopy (Cecil Instruments, CE9500) at different time intervals. The result shown in Figure 16 illustrates gradual decrease in NADH concentration (E340 = 6,330) with the oxidation time, which can be attributed to continuous oxidation of NADH by the ferrocene polymeric mediator immobilized in the carbon electrode.

0.65 mL above oxidized NADH solution was collected and mixed with 0.25 mL 99.5% ethanol (EtOH, Sigma-Aldrich) in a UV cuvette (l = 1cm). UV spectra of the solution in the cuvette were recorded before and after addition of 0.4 mg alcohol dehydrogenase (ADH, Sigma-Aldrich). As shown in Figure 17, the addition of EtOH and ADH increased the absorbance peak at around 340 nm to almost the same level as the control solution which contained 0.3 mM NADH in 0.01M pH7.4 PBS and did not subject to the oxidation process. This suggests that NADH oxidation by the immobilized ferrocene polymeric mediator predominantly, if not completely produced electrochemically active NAD+ that was reduced to NADH by EtOH in the presence of ADH.

stability of the biosensors by recording sensor response change with time at a fixed glucose concentration proved to be unreliable. Nevertheless, closely aligned data points to the linear regression line in Figure 15 illustrate stable biosensor response during the 54-hour test,

As a coenzyme for hundreds of oxidoreductases, NAD has attracted great attention for biosensor development. Because its direct oxidation at electrode surface requires high applied potential and involves side reactions leading to the formation of electrochemically inactive by-products (see the preceding section), NAD is normally coupled with mediators for its application in biosensors. In the literature, ferrocenium and its derivatives have been

section, small molecular mediators are not normally suitable for continuous monitoring biosensors. The effective immobilization of the ferrocene polymeric mediator in the screenprinted carbon electrode developed at LifeScan Scotland Limited holds potentials for development of continuous monitoring biosensors using NAD-dependant oxidoreductases. To investigate electrochemical communication between NAD and the immobilized ferrocene polymeric mediator, 0.3 V potential (ECOChemie PGSTat Autolab, Type III) was applied to an electrochemical cell containing 3.0 mL 0.01M pH7.4 PBS and equipped with a screen-printed carbon electrode (approximately dimension length x width = 10 x 5 mm) with immobilized ferrocene polymeric mediator (about 12.5% wt/wt), an Ag/AgCl reference electrode and a platinum counter electrode to oxidize the ferrocene moieties of the polymeric mediator to ferrocenium moieties. After the current of ferrocene oxidation reached a steady level, PBS in the cell was replaced with 0.3 mM NADH (from Sigma-Aldrich) in 3.0 mL 0.01M pH7.4 and a step increase in current was detected, which indicated oxidation of NADH by the ferrocenium moieties of the polymeric mediator. The NADH oxidation was followed by measuring absorbance of the solution at 340 nm wavelength by UV/vis spectroscopy (Cecil Instruments, CE9500) at different time intervals. The result shown in Figure 16 illustrates gradual decrease in NADH concentration (E340 = 6,330) with the oxidation time, which can be attributed to continuous oxidation of NADH by the

0.65 mL above oxidized NADH solution was collected and mixed with 0.25 mL 99.5% ethanol (EtOH, Sigma-Aldrich) in a UV cuvette (l = 1cm). UV spectra of the solution in the cuvette were recorded before and after addition of 0.4 mg alcohol dehydrogenase (ADH, Sigma-Aldrich). As shown in Figure 17, the addition of EtOH and ADH increased the absorbance peak at around 340 nm to almost the same level as the control solution which contained 0.3 mM NADH in 0.01M pH7.4 PBS and did not subject to the oxidation process. This suggests that NADH oxidation by the immobilized ferrocene polymeric mediator predominantly, if not completely produced electrochemically active NAD+ that was reduced

[60-63] As mentioned in the preceding

suggesting good FAD-GDH stability during the continuous measurement.

**9.2. NAD and ferrocene polymeric mediator redox couple** 

ferrocene polymeric mediator immobilized in the carbon electrode.

to NADH by EtOH in the presence of ADH.

reported as mediator for oxidation of NADH to NAD+.

**Figure 16.** Variation of UV absorbance at 340 nm of 0.3 mM NADH in 0.01 M pH7.4 PBS by carbon electrode with immobilized ferrocene polymeric mediator at 0.3 V potential, by using an Ag/AgCl reference electrode and a platinum counter electrode, at room temperature

**Figure 17.** UV spectroscopy monitoring reduction of oxidized NADH in the presence of ADH and EtOH, 0.65ml NADH (0.2 mM) + 0.25ml EtOH (99.5%) + 20ul ADH (control) (solid line), 0.65ml oxidized NADH + 0.15mL EtOH (before reduction) (broken line), 0.65ml oxidized NADH + 0.25ml EtOH + 20 ul ADH (4mg/ml) (dotted line)

For continuous monitoring, use of native NAD as a coenzyme apparently is not ideal due to its low molecular weight and high water-solubility. Direct immobilization of free NAD at electrode has been a long-standing challenge. [64, 65] There are a large number of publications in the literature concerned with NAD retention for various purposes. In the field of biosensors, one promising approach is covalently attaching NAD moieties to polymer chains which can be immobilized at electrode by different means. This can be achieved by directly coupling NAD to an electrode modified with a polymer bearing functional groups,[66] entrapping polymeric NAD in semipermeable membranes [67, 68] or synthesizing a charged polymeric NAD which was then physically adsorbed at electrode surface in conjunction with counter-charged polymer(s) in a manner of layer-by-layer. [69] At LifeScan Scotland Limited, we intend to develop continuous monitoring biosensors using NAD-dependent enzymes by immobilizing all the redox species, i.e. an enzyme(s), the ferrocene polymeric mediator and a polymeric NAD in an electrode with the 3 dimentional nano-porous structure (see Figure 10). A polymeric NAD is normally synthesized by two routes. One is coupling NAD or NAD analogue with a polymer bearing functional groups. [66, 70-73] The other route involves synthesis of a NAD monomer and then its copolymerisation with another co-monomer(s). One of the challenges for NAD immobilization is to keep NAD coenzymic activity while achieving effective retention at electrode. Yamazaki *et al* synthesized three NAD monomers (N6-[N-(6-methacrylamidohexyl)carbamoylmethyl]-, N6[N-[2-[N-(2 methacrylamidoethyl)carbamoyl]ethyl] carbamoylmethyl]-, and N6 -[N-[N-(2-hydroxy-3 methacrylamidopropyl) carbamoylmethyl] carbamoylmethyl]-NAD) and then copolymerized them with various co-monomers (acrylamide, N-(2-hydroxyethyl)-, N-ethyl-, N,N-diethyl-, and N,N-dimethylacrylamide, acrylic acid, and 6-methacrylamidohexylammonium) by free radical polymerisation to form a series of polymeric NADs. [74] Their studies revealed that hydrophilicity and length of the spacers linking NAD moieties and the polymer backbone had the most important effects on coenzymic activity of the polymeric NADs. This suggests that keeping mobility of the NAD moieties covalently attached to a polymer chain is critical to NAD coenzymic activity. Chemically modification to native NAD is required to tailor chemical properties of the spacers. N6-amino group on the adenine ring of NAD (see Figure 18) is normally selected as the site for this purpose. [75, 76] Lindberg *et al* reported alkylation of NAD+ with iodoacetic acid followed by alkaline rearrangement to give N6-carboxymethyl-NAD+. [75] However, the reaction between NAD+ and iodoacetic acid took 10 days in the dark at room temperature. We successfully synthesized N6-carboxymethyl-NAD+ by a modified method with a dramatic reduction in the reaction time.

Amperometric Glucose Sensors for Whole Blood Measurement Based on Dehydrogenase Enzymes 347

**9.5. Synthesis of N6-carboxymethyl-NADH (compound 2 in Figure 19)** 

**9.6. Oxidation of N6-carboxymethyl-NADH to N6-carboxymethyl-NAD+**

3.99g crude N6-carboxymethyl-NAD+ as a tan coloured hygroscopic solid.

column of Dowex 1-X2 ion exchange resin (Cl-

The reaction mixture containing N6-carboxymethyl-NADH was treated with 17.5 mL 3M Tris buffer (pH7.0) and the pH was adjusted to 7.5 using 5M HCl aqueous solution. 3.5 mL acetaldehyde (62.6mmol) was added, immediately followed by 10.5 mg yeast alcohol dehydrogenase (s*accharomyces cerevisiae*) (~300U/mg) before allowing agitating at ambient temperature to deoxidize the nicotinamide moiety. After 18 hours, the reaction mixture (c.a. 485 mL) was concentrated *in vacuo* (30°C/10-15bar) to approximately 1/3 volume and poured into 1800 mL pre-cooled (-5°C) mixture of acetone/IMS (1:1). The fine slurry was left to age for 18 hours at 3°C. The resulting precipitate was collected by centrifugation and washed on a glass sinter with 40 mL IMS then 120 mL dry diethyl ether before air-drying under dry nitrogen for 10 minutes. Further drying overnight in a desiccator over fused CaCl2 afforded

1.0 g of the above-prepared crude N6-carboxymethyl-NAD+ was taken up in 20 mL water and passed through a Sephadex G10 gel filtration column (2x10cm, 20 mL). All eluted fractions containing UV active material were combined (60 mL total volume) and added to a

equilibrated with water. A linear gradient of 0-50 mM LiCl (buffered to pH 3.0), at 10 mL per minute over 65 minutes, was applied using "*Presearch Combiflash Companion"* chromatography equipment. The fractions eluted between 25-35 mM were combined (c.a. 100 mL), neutralized to pH 7.0 with 5M LiOH and evaporated to approximately 1/3 volume and poured into 300 mL pre-cooled (-5°C) mixture of acetone/IMS (1:1). The fine slurry was left to age for 18 hours at 3°C. The resulting precipitate was collected by centrifugation and washed on a glass sinter with 30 mL IMS then 50 mL dry diethyl ether before air-drying under dry nitrogen for 10 minutes. Further drying overnight in a desiccator over fused CaCl2 afforded 0.307g purified N6-carboxymethyl-NAD+ as a cream coloured hygroscopic

, 4x50cm, 200 mL) which had been pre-

carboxymethyl-NADH.

solid.

**(compound 3 in Figure 19)** 

9.1g (c.a. 10.57mmol) above prepared crude N1-carboxymethyl-NAD+ was dissolved in 1.3% w/v NaHCO3 in 450 mL aqueous solution and the solution was deoxygenated by sparging with nitrogen for 10 minutes. 3.5 g (20.1mmol) sodium dithionite was added in one portion and the mixture was stirred at ambient temperature to affect reduction of the nicotinamide moiety. After 1.0 hour, the solution colour changed from pink to yellow. The solution was then sparged with air for 10 minutes to destroy any excess dithionite and the pH was brought to 11.0 by using 5M NaOH aqueous solution. The mixture was heated at 70°C for 90 minutes, to promote Dimroth rearrangement to N6-carboxymethyl-NADH, before cooling to 25°C. Thin-layer chromatography (silica gel, isobutryic acid/water/32% NH4OH (aq), 66/33/1.5 by volume) showed no evidence for the presence of N1-

**Figure 18.** Chemical structure of NAD+

#### **9.4. Synthesis of N1-carboxymethyl-NAD+ (compound 1 in Figure 19)**

1.0g NAD+ (1.51mmol) was dissolved in 3.5mL 0.1M pH 7.0 sodium phosphate buffer in a 5ml Biotage microwave reaction tube. Then, 1.5 g (8.06mmol, 5.34eq) iodoacetic acid was added and pH was adjusted to 7.0 by using 5.0M NaOH aqueous solution. The reaction vessel was sealed and the mixture was heated to 50°C for 10 minutes by using microwave irradiation. After that, the pink solution (c.a. 5.0 mL) was acidified to pH3.0 using 5M HCl aqueous solution before being poured into 25 mL pre-cooled (-5°C) mixture of acetone/IMS (1:1). The resulting precipitate was filtered, washed first with 5.0 mL IMS, then 15 mL dry diethyl ether before air drying under dry nitrogen for 10 minutes. Further drying overnight in a desiccator over fused CaCl2 gave 1.62 g crude N1-carboxymethyl-NAD+ as a pink amorphous solid.
