Structured biotinylated poly(3,4-ethylenedioxypyrrole) electrodes for biochemical applications

Abstract

The immobilization of biotin on transducer surfaces is a very important step for the fabrication of biosensors for many applications (immunoassay, DNA-hybridization assays, targeted imaging). Biotinylated polypyrroles have been studied and tested but gave rise to problems of polymerization and stability due to the intrinsic properties of pyrrole. As an alternative, biotinylated pyrroles were often used in a copolymerization with pyrrole or with an amphiphilic pyrrole derivative in a copolymerization to reduce the problems due to the pyrrole substitution. To find a new strategy, this paper presents the homopolymerization, instead of the use of a copolymerization, by replacing pyrrole by 3,4-ethylenedioxypyrrole bearing biotinylated substituent. We report the synthesis, characterization and electrochemical properties of two biotinylated 3,4-ethylenedioxypyrroles differing by the length of the alkyl spacer (ethyl or dodecyl) as well as the characterization of the corresponding polymer films. We successfully show, by cyclic voltammetry, that these monomers polymerize perfectly and give relatively stable polymer films. The increase of the alkyl spacer improves the polymerization and increases the polymer stability. For the first time, we also studied the surface morphology of an electrodeposited biotinylated polymer. The electrodeposition of these biotinylated derivatives gave rise to the ability to modulate the surface microstructuration, which consists of microspheres or cauliflower-like microstructures according to the length of the alkyl spacer.

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Introduction

The very high non-covalent interactions between biotin and glycoproteins such as avidin and streptavidin makes biotin an ideal candidate for many applications, for example in immunoassays, DNA-hybridization assays, targeted imaging or drug delivery.^1–3 For some of these applications, biotin needs to be immobilized on surfaces for further bio-recognition for instance.

Classical methods of immobilization are based on grafting or polymer deposition. Electrochemical polymerization is a very fast method for the deposition of conducting polymers using soft conditions, which can be employed, in certain conditions, with biological species such as biotin.^4–19 Among all the electronically conducting polymers, polypyrrole is often used because of its biocompatibility and soft electropolymerization conditions.^4–16 Usually, biotin is either incorporated inside electroactive polymers as a doping compound thanks to its carboxylic group,¹⁴ or grafted on the monomer before polymerization.^4–13 The biotin doping allowed a maximum of one biotin per three–four monomer units and presented polymer stability problems or doping loss with time. To avoid this problem, biotin can be chemically grafted on the monomer. In this method, the substituent size (biotin + spacer + connector) has a very significant influence on the polymerization ability and has a consequence on polymer chain lengths, dispersion in size and solubility. In other cases, to facilitate the polymerization, biotin was grafted on the polymer using a post-treatment,⁴ which leads to lower concentration of biotin.

In all the publications reported in the literature on biotin-substituted pyrrole, biotin was chemically linked to pyrrole in the N-position. Many pyrrole derivatives containing biotin at the N-position and with various spacers were synthesized and analyzed by many groups, as represented in Scheme 1A. However, the polymerization of pyrrole derivatives was very sensitive to the substituent presence and in the case of pyrrole with substitution at the N-position, very high steric hindrances were induced^20,21 giving rise to problems of stability as previously reported by Cosnier et al.⁸ To reduce steric hindrances and to make the electrodeposition easier, a biotinylated pyrrole was often mixed with pyrrole or an amphiphilic pyrrole derivative in a copolymerization process.^4–10,16N-Biotinylated polypyrroles showing biological activity have been reported where the carboxylic group of biotin was not much involved in the molecular recognition.

Scheme 1 (A) Biotinylated pyrroles reported in the literature and (B) schematic representation of the biotinylated 3,4-ethylenedioxypyrroles studied in this paper.

One of the methods to improve the electropolymerization would be to put the substituent in the 3-position. However, it is now established that polypyrrole films often present many surface defects such as wrinkles which are not wanted for many applications.^22–24 Very recently, a new family of polypyrrole derivatives, known as poly(3,4-alkylenedioxypyrroles), showing exceptional electronic and optical properties, has been developed.^25–29 In this series, the presence of the 3,4-alkylenedioxy bridge has many advantages. First of all, if the 3- and 4-position are occupied, no structural defects are possible. Moreover, the electron donating properties of the bridge highly decrease the monomer and polymer oxidation potentials rendering the electropolymerization much easier. Fluorinated poly(3,4-ethylenedioxypyrroles) (PEDOP) and poly(3,4-propylenedioxypyrroles) (ProDOP) with exceptional electrochemical properties and stability and with various surface morphologies have been reported.^30–31

The aim of this work is to report the synthesis and electrochemical properties of two biotinylated EDOP monomers represented in Scheme 1B. The two monomers (B₂ and B₁₂) differ by the length of the alkyl spacer between the polymerizable core and the biotin. Their ability to form polymer films electrochemically was analyzed by cyclic voltammetry. The polymer films were analyzed by imaging infrared, optical profilometry and scanning electron microscopy (SEM) in order to determine their morphology. Because the presence of nano/microstructures is very important for the elaboration of biosensors,^32–35 due to the large specific surface area, three different surfaces were chosen for the electrodeposition: smooth gold plates, microstructured gold plates and nanostructured gold plates. The evaluation of the surface structuration effect on the molecular recognition power of the biotinylated surfaces will be evaluated in a future project.

Experimental

1. Monomer synthesis and characterization

All chemical were purchased from Sigma-Aldrich. EDOP, 2-tert-butyldimethylsilyloxyethyltosylate (Tos₂) and 2-(2H-[1,4]dioxino[2,3-c]pyrrol-6(3H)-yl)ethanol (OH₂) were obtained using a procedure already reported in the literature (Scheme 2).³⁰ The NMR spectra were obtained with a Bruker Avance 200 MHz. The mass spectra of the molecules were realized either by electron ionisation at 70 eV with a Thermo TRACEGC of Thermofischer Corp. fitted with an Automass III Multi spectrometer (GC-MS) or with a Thermo LCQ Classic « Ion Trap » spectrometer of Thermofischer Corp. fitted with ESI Type API.1 (ESI-SM).

Scheme 2 Synthetic route to the monomers.

1.1 Synthesis of 2-(tert-butyldimethylsilyloxy)dodecanol. Triethylamine (5 mL, 48.2 mmol) and 4-dimethylaminopyridine (DMAP) (50 mg) were added to a solution of 1,12-dodecanediol (12 g, 59.4 mmol) in 200 mL of anhydrous dichloromethane. After cooling to 0 °C, a solution of tert-butyl(chloro)dimethylsilane (5.4 g, 35.6 mmol) in 50 mL of anhydrous dichloromethane was slowly added. The reaction mixture was then heated at 40 °C. After stirring for one day, the solution was washed with hydrogen carbonate saturated aqueous solution. The organic layer was separated and dried on Na₂SO₄. Removal of the solvent and purification by column chromatography (silica gel; eluent: chloroform/cyclohexane 1 : 1) gave the product (retention time: 16.4 min).

Yield 54%; δ_H(250 MHz, CDCl₃) 0.04 (6 H, s), 0.89 (9 H, s), 1.44 (20H, m) and 3.62 (4H, m).

1.2 Synthesis of 2-tert-butyldimethylsilyloxydodecyltosylate (Tos₁₂). Tosyl chloride (7 g, 22.1 mmol) and DMAP (50 mg) were added to a solution of 2-(tert-butyldimethylsilyloxy)dodecanol (6.33 g, 33.2 mmol) in anhydrous pyridine at 0 °C. After stirring for 12 h at 0 °C, 200 mL of dichloromethane was added and the reaction mixture was washed with 1 N HCl. The organic layer was separated and dried on Na₂SO₄. Removal of solvent and purification by column chromatography (silica gel; eluent: dichloromethane/hexane 1 : 4) gave the product.

Yield 45%; δ_H(250 MHz, CDCl₃) 0.04 (6 H, s), 0.88 (9 H, s), 1.39 (20 H, m), 2.44 (3 H, s), 3.59 (2 H, t, ³J_HH 6.5), 4.01 (2 H, t, ³J_HH 6.5), 7.34 (2 H, d, ³J_HH 8.2) and 7.79 (2 H, d, ³J_HH 8.2); δ_C(50 MHz, CDCl₃) −5.27, 18.36, 21.61, 25.31, 25.78, 25.97, 28.80, 28.91, 29.36, 29.41, 29.45, 29.51, 29.58, 32.87, 63.32, 70.69, 127.87, 129.77, 133.26 and 144.57.

1.3 Synthesis of 12-(2H-[1,4]dioxino[2,3-c]pyrrol-6(3H)-yl)dodecan-1-ol (OH₁₂). Dry sodium hydride (0.50 g, 20 mmol) was carefully added to a solution of EDOP (1 g, 8 mmol) in anhydrous THF. After stirring for 30 mn, Tos₁₂ (4.7 g, 10 mmol) was added and the reaction mixture was refluxed for 24 h. After cooling at room temperature and solvent evaporation, aqueous ammonium chloride was carefully added and the solution was extracted with diethyl ether and dried on Na₂SO₄. Removal of solvent and purification by column chromatography (silica gel; eluent: dichloromethane) gave a mixture of the product (r.t. 28.8 mn) and 2-tert-butyldimethylsilyloxydodecyl tosylate. The mixture was used without further purification.

To this mixture, was added tetrabutylammonium fluoride in THF. After stirring during 24 h at 60 °C and solvent evaporation, the purification by column chromatography (silica gel; eluent: dichloromethane) afforded the product as crystalline solid.

Yield 38%; crystalline solid; m.p. 29.5 °C; δ_H(250 MHz, CDCl₃) 1.25 (16 H, s), 1.56 (2 H, quint, ³J_HH 6.35), 1.66 (2 H, quint, ³J_HH 6.57), 3.63 (4 H, m), 4.17 (4 H, s) and 6.05 (2H, s); δ_C(50 MHz, CDCl₃) 25.70, 26.66, 29.18, 29.38, 29.44, 29.48, 29.53, 31.30, 32.78, 50.51, 63.05, 65.83, 100.96 and 131.62; m/z 309 (M⁺, 1), 278 (C₁₇H₂₈NO₂⁺, 1), 264 (C₁₆H₂₆NO₂⁺, 2), 250 (C₁₅H₂₄NO₂⁺, 3), 236 (C₁₄H₂₂NO₂⁺, 4), 222 (C₁₃H₂₀NO₂⁺, 6), 208 (C₁₂H₁₈NO₂⁺, 6), 194 (C₁₁H₁₆NO₂⁺, 11), 180 (C₁₀H₁₄NO₂⁺, 15), 166 (C₉H₁₂NO₂⁺, 6), 152 (C₈H₁₀NO₂⁺, 14), 139 (C₇H₉NO₂⁺, 100), 138 (C₇H₈NO₂⁺, 61) and 125 (C₆H₇NO₂⁺, 36).

1.4 Synthesis of B₂ and B₁₂. N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC) (0.36 g, 1.8 mmol) and DMAP (30 mg) were added to a solution of biotin (0.44 g, 1.8 mmol) in dichloromethane. After stirring for 30 mn at room temperature, OH₂ (0.25 g, 1.5 mmol) or OH₁₂ (0.46 g, 1.5 mmol) was added. After a day, the solvent was removed and the crude was purified by column chromatography (silica gel; eluent: methanol). After solvent removal, the monomer was dissolved in acetonitrile and filtrated with 0.45 μm syringe filter.

(B₂) Yield 65%; white solid; δ_H(250 MHz, CD₃OD) 1.28–1.78 (6 H, m), 2.34 (2 H, t, ³J_HH 7.16), 2.70 (1 H, d, ²J_HH 12.70), 2.94 (1 H, dd, ²J_HH 12.70, ³J_HH 4.43), 3.20 (1 H, m), 3.92 (2 H, t, ³J_HH 5.21), 4.10 (4 H, s), 4.23 (2 H, t, ³J_HH 5.21), 4.29 (1 H, dd, ³J_HH 7.87, ³J_HH = 4.64), 4.50 (1 H, dd, ³J_HH 7.87, ³J_HH 4.43) and 6.09 (2 H, s); δ_C(50 MHz, CD₃OD) 25.87, 29.43, 29.60, 34.78, 41.08, 56.89, 61.67, 63.40, 65.36, 66.98, 102.48, 133.54, 165.98, 175.11; ν_max(film)/cm⁻¹ 3280 (NH), 2925, 2864, 1731 (COO), 1694 (NHCONH), 1551, 1391, 1362 and 1055; ESI-MS m/z 418.1 (M+Na).

(B₁₂) Yield 59%; white solid; δ_H(250 MHz, CD₃OD) 1.28–1.78 (26 H, m), 2.34 (2 H, t, ³J_HH 7.21), 2.70 (1 H, d, ²J_HH 12.70), 2.93 (1 H, dd, ²J_HH 12.70, ³J_HH 4.80), 3.20 (1 H, m), 3.65 (2 H, t, ³J_HH 6.87), 4.10 (6 H, m), 4.30 (1 H, dd, ³J_HH 7.84, ³J_HH 4.40), 4.49 (1 H, dd, ³J_HH 7.84, ³J_HH 4.80) and 6.04 (2 H, s); δ_C(50 MHz, CD₃OD) 25.87, 25.93, 26.00, 27.03, 27.67, 29.46, 29.68, 29.72, 30.27, 30.29, 30.58, 32.50, 34.84, 34.88, 41.02, 56.96, 61.45, 61.61, 63.38, 65.60, 66.95, 101.96, 133.04, 166.10 and 175.51; ν_max(film)/cm⁻¹ 3228 (NH), 2923, 2852, 1733 (COO), 1699 (NHCONH), 1552, 1392, 1361 and 1056; ESI-MS m/z 558.3 (M+Na).

2. Elaboration of micro-patterned gold plates

The structures were micro-patterned using optical lithography based on a Microposit positive resist S1813-G2. The resist film was spin-coated onto the substrate at 3000 rpm resulting in thickness of approximately 1.5 μm. The photo resist was structured using a UV source illuminating a Digital Mirror Device with about 600 000 individually switchable micro mirrors. The resulting pattern was projected onto the substrate. The illuminated part of the substrate was removed in a development step, while the unexposed part remained as a structure of mesa-shaped objects with interconnections (cf.Fig. 7). In a subsequent step the structure was metal coated.

3. Elaboration of nano-structured gold/SiOx plates using nano-template fabrication

The gold/SiOx nano-templates were fabricated by using a procedure already described elsewhere^33,34 and illustrated in Fig. 1. Briefly, the fabrication consisted of the creation of a 2D hexagonal crystalline monolayer mask of self assembled polystyrene nano-beads (nominal diameter 500 nm) (Sigma-Aldrich) on the gold surface by spin coating. An oxygen plasma etching was then performed to reduce the bead diameter and to define the size of the nano-mask. Afterwards, an insulating SiOx layer was deposited through the nano-mask by Plasma Enhanced Chemical Vapour Deposition to form the nano-templates. Finally the beads were removed in water/ultrasound baths to obtain the SiOx nano-template and form gold nano-electrodes array. The mean height of the SiOx nano-template was about 14.5 nm (Fig. 2).

Fig. 1 Schematic representation of the fabrication of the gold/SiOx nano-templates.

Fig. 2 AFM pictures of the gold/SiOx nano-templates. The scale bars represent 2 μm and 500 nm.

4. Electrochemical polymerization

For the electrochemical experiments, 1 mM of the monomer and 0.1 M of tetrabutylammonium hexafluorophosphate (Bu₄NPF₆) were dissolved in anhydrous acetonitrile and degassed. A three-electrode system was used: a working electrode (platinum disk for cyclic voltammetry experiments, smooth or pattern gold plates for chronoamperometry experiments), a graphite counter-electrode and a saturated calomel reference electrode (SCE).

Results and discussion

1. Synthesis

The two EDOP derivatives containing a hydroxyl group and differing by the alkyl spacer (ethyl or dodecyl) represented in Scheme 2 were obtained in ten steps from iminodiacetic acid following a synthetic route reported in previous work.³⁰ The synthesis route includes obtaining EDOP in eight steps, the introduction of a hydroxylated substituent at the N-position using a molecule containing both a silyl and a tosylate group (2-tert-butyldimethylsilyloxyethyltosylate for B₂ and 2-tert-butyldimethylsilyloxydodecyltosylate for B₁₂). The successful synthesis of these two derivatives was confirmed by ¹H and ¹³C NMR, infrared and mass spectrometry (cf. ESI). Afterwards, the biotinylated monomer was obtained by coupling in dichloromethane with EDC and DMAP.

2. Electrochemical characterization

Each biotinylated monomer was separately introduced inside a cell containing a solution of tetrabutylammonium hexafluorophosphate (0.1 M) in anhydrous acetonitrile for the electrodeposition. A monomer concentration of 1 mM was chosen (a higher monomer concentration can lead to a decrease in the electrochemical stability). First of all, their oxidation potential, E_ox = 0.91 V vs. SCE for B₂ and 0.86 V for B₁₂, was determined by potential scanning between 0 and 1.2 V. Therefore, the increase of the alkyl spacer seemed to have a beneficial effect on their oxidation. Then, their electropolymerization ability was studied by cyclic voltammetry. Their cyclic voltammograms (10 scans) are represented in Fig. 3A for B₂ and 3C for B₁₂. These graphs exhibited a reversible system, which corresponds to the oxidation and the reduction of the electroactive polymer. The increase in the intensity after each scan means that the amount of electrodeposited polymer regularly increases when the monomer oxidation potential is reached. Then, after ten scans, the working electrode was washed and put in a solution without monomer, and the stability of the films and was determined by performing 100 scans by cycling voltammetry. First of all, the half-wave potential of the polyB₂ (Fig. 3B), about 0.29 V, was higher than that of non-substituted PEDOP (E_1/2 = −0.24 V) and semi-fluorinated PEDOP (E_1/2 = 0.24 V), whereas that of polyB₁₂ (Fig. 3D), about 0.16 V, was much lower. As a consequence, the alkyl chain increase favoured electropolymerization, increasing the polymer chains length, while the presence of the biotin induced higher steric hindrance than semi-fluorinated tails (F-octyl) giving rise to a shortening of the polymer chains. Indeed, the biotin should be sufficiently far from the polymer backbone to not impede the intra- and intermolecular electronic exchanges. As a consequence, the cyclic voltammogram of polyB₁₂ was an extremely well-defined doping system with three oxidation and reduction peaks, possible only if the polymer backbone is not highly disrupted,^30a while that of polyB₂ displayed only one oxidation and reduction peak. The polymer oxidation potentials were far from the monomer oxidation potentials, which indicated the polymer chain lengths were relatively important. Indeed, the polymers were completely insoluble in acetonitrile and no soluble part was observed during the deposition, which is a very important point in the polymer electrodeposition. Finally, the 100 scans revealed that just a slight decrease in the intensity of the polymer oxidation was detected for polyB₁₂ (Fig. 3D) and a higher decrease for polyB₂ (Fig. 3B), confirming the higher steric hindrances in this film. We also observed the very high adherence of the film on the working electrode and the difficulty to remove it after the experiments.

Fig. 3 Cyclic voltammograms of (A) B₂ and (C) B₁₂ (1 mM) in 0.1M Bu₄NPF₆/CH₃CN: 10 scans vs. SCE. Cyclic voltammograms of (B) polyB₂ and (D) polyB₁₂ in 0.1M Bu₄NPF₆/CH₃CN without monomer: 100 scans vs. SCE.

In a recent publication, the group of Reeman reported an electrochemical method to characterize the bioaffinity of biotinylated polyterthiophenes in the presence of avidin.^17a They showed that the exposure of the polymers to a buffer solution containing avidin can lead to drastic changes in the cyclic voltammograms because the presence of avidin decreases the electronic exchanges. This method was used to characterize polyB₂ and polyB₁₂. More precisely, the working electrodes containing the polymers were, first, put in a buffer solution (1 M NaCl, 10 mM EDTA) during 15 mn and a scan was performed to evaluate their resistance to the buffer. Unfortunately, polyB₂ was affected by the buffer confirming its lower stability while polyB₁₂ was completely unaffected. Then, polyB₁₂ was added to a buffer solution containing different amounts of avidin (from 10⁻¹² to 10⁻⁸ M) and the film was characterized by cyclic voltammetry after each exposure. The cyclic voltammograms are gathered in Fig. 4. This figure shows that the cyclic voltammogram of polyB₁₂ was not affected after these experiments. Hence, either the bioaffinity of these polymers is not very important or the presence of avidin does not affect the electronic exchange of these polymers. Other experiments will be undertaken in the near future to determine the bioaffinity of these surfaces.

Fig. 4 Cyclic voltammogram of polyB₁₂ (0.01 M) after exposure to different amounts of avidin.

3. Surface characterization

For the study of their surface properties, polymer films were electrodeposited on smooth and microstructured gold substrates by imposing a constant potential of 0.84 V vs. SCE for B₂ and 0.80 V for B₁₂ and with a deposition charge of 100, 200 and 300 mC cm⁻². The films were characterized by infrared imaging in reflection mode. In this method, the infrared beam penetrates the polymer film and is reflected by the gold plate. Therefore, the polymer film should not be too thick or opaque. We also chose to remove the doping anions by electrochemical reduction (−0.5 V vs. SCE during 15 min) because of their absorbance in the infrared. First of all, the infrared spectra of the polymers were obtained (Fig. 5A). These spectra confirmed the presence of the polymers thanks especially to two bands: a relatively large absorption between 1680 and 1750 cm⁻¹, due to the superposition of the ester band (1730 cm⁻¹) and the carbamide band (1696 cm⁻¹) of the biotin, and a large band at about 3300 cm⁻¹ due to the N–H stretching of the carbamides. The alkyl band at 2920 and 2848 cm⁻¹ were more intense for polyB₁₂ due to the presence of the dodecyl spacer. Secondly, infrared imaging was used to obtain the total absorbance of the surface. According to Fig. 5B, domains where the polymer is concentrated are visualized showing that the surface seems to be structured.

Fig. 5 Cyclic (A) infrared spectra of dedoped polyB₂ and polyB₁₂ obtaining by imaging infrared and (B) total absorption of the de-doped polyB₁₂ (Qs = 200 mC cm⁻²).

The roughness of the surfaces was evaluated by optical profilometry (cf.Fig. 6 and Table 1). Table 1 gathers the roughness data of polyB₂ and poly B₁₂ as a function of the deposition charge (Qs). An increase of the surface roughness with Qs was measured for the two polymers. Electrodeposited polyB₁₂ films were rougher, with the same polymerization conditions, than polyB₂ films.

Fig. 6 Cyclic optical profilometry images of polyB₁₂ deposited on smooth gold plates for a deposition charge of (A) 100, (B) 200 and (C) 300 mC cm⁻².

Table 1 Roughness data obtained by optical profilometry

Polymer	Qs [mC cm^−2]	Roughness parameters
		Ra [nm]	Rq [nm]
PolyB2	100	67.4	125.0
	200	117.5	279.4
	300	556.4	1165.0
PolyB12	100	162.1	266.1
	200	312.4	457.6
	300	893.9	1550.1

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To better evaluate the influence of the alkyl spacer on the surface roughness and morphology, the polymer films were analyzed by SEM (Fig. 7). First of all, the polymer films were very structured. The same surface morphology was observed when the polymers were electrodeposited on smooth or micro-patterned gold surfaces. The two polymer films were very structured but differences in surface morphology were observed. PolyB₂ films were composed of an assembly of micrometric spherical particles of different sizes. The spherical structures were relatively smooth as shown in Fig. 7C. On the other hand, polyB₁₂ consisted of cauliflower-like microstructures. The structures had micro and sub-micro dimensions, as shown in Fig. 7F, confirming the higher roughness of these films.

Fig. 7 SEM images of polyB₂ (scale bar: (A) 200 μm, (B) 10 μm and (C) 3 μm) and polyB₁₂ (scale bar : (D) 200 μm, (E) 10 μm and (C) 3 μm) electrodeposited on micro-patterned gold plates; Qs = 300 mC cm⁻².

The possibility of having nano-structured films of the polymer opens the possibility of many applications, as for example biosensors with increased sensitivity.³⁵ Many trials using electrodes consisting of arrays of insulating SiO_x nano-pillars showed the possibility of producing nanostructured biotinylated poly(3,4-ethylenedioxypyrrole) surfaces with a deposition charge below 5 mC per cm² of projected area (the local density is much higher in nanostructured surfaces) (Fig. 8A). For a deposition above 5 mC cm⁻², the polymer growth covered the SiOx nano-pillars as shown in Fig. 8B. Therefore, the thickness of the electrodeposited polymer is about 15 nm for a deposition charge of 5–7.5 mC cm⁻².

Fig. 8 AFM pictures of polyB₂ electrodeposited on nano-structured gold/SiOx plates with a deposition charge of (A) 5 and (B) 7.5 mC cm⁻²; the area represents 5 × 5 μm².

Conclusions

In aiming to improve the fabrication of biosensors based on the biotin-avidin system by electrodeposition, two biotinylated monomers based on the 3,4-ethylenedioxypyrrole core and differing by the alkyl spacer (ethyl or dodecyl) were synthesized, characterized and electrodeposited.

First of all, the two monomers perfectly polymerized and the polymer films were very adherent, and microstructured. We showed that a long alkyl spacer (dodecyl for example) improves the polymerization and the electrochemical stability. The alkyl spacer length also has a noticeable effect on the surface morphology. The polymerization was applied to micro and nanostructured surfaces and was successfully in creating spatially controlled and structured films which may offer new opportunities for the production of novel structured sensor surfaces.

Acknowledgements

The research leading to these results has received funding from the European Community's Seventh Framework Programme (FP7/2007-2013) under grant agreement no CP-FP 214478-2.

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Footnote

† Electronic Supplementary Information (ESI) available: NMR analyses. See DOI: 10.1039/c1ra00392e/

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