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Molecularly imprinted polymer for recognition of 5fluorouracil by the RNA-type nucleobase pairing Tan-Phat Huynh, Piotr Pieta, Francis D'Souza, and Wlodzimierz Kutner Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/ac401598k • Publication Date (Web): 26 Jul 2013 Downloaded from http://pubs.acs.org on July 27, 2013

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Analytical Chemistry

Molecularly imprinted polymer for recognition of 5-fluorouracil by the RNA-type nucleobase pairing Tan-Phat Huynh,a,b Piotr Pieta,a Francis D’Souza,b,* and Wlodzimierz Kutnera,c,* a

Department of Physical Chemistry of Supramolecular Complexes, Institute of Physical Chemistry, Polish Academy of Sciences, Kasprzaka 44/52, 01-224 Warsaw, Poland

b

Department of Chemistry, University of North Texas, Denton, TX 76203-5017, USA

c

Faculty of Mathematics and Natural Sciences, School of Science, Cardinal Stefan Wyszynski University in Warsaw, Wóycickiego 1/3, 01-815 Warsaw, Poland KEYWORDS 5-fluorouracil, molecularly imprinted polymer, electropolymerization, capacitive impedimetry, piezoelectric microgravimetry ABSTRACT: A 6-aminopurine (adenine) derivative of bis(2,2’-bithienyl)methane, the 4-[2-(6-amino-9H-purin-9yl)ethoxy]phenyl-4-[bis(2,2’-bithienyl)methane], Ade-BTM, was designed and synthesized for recognition of 5fluorouracil (FU), an antitumor chemotherapy agent, by the RNA-type nucleobase pairing driven molecular imprinting. The pre-polymerization complex stoichiometry involved one FU molecule and two molecules of the Ade-BTM functional monomer. Molecular structure of this complex was thermodynamically optimized using DFT at the B3LYP/3-21G* level. The stability constant of the FU and Ade-BTM complex of the 1:2 stoichiometry was K = 2.17(±0.07)×107 M-2, as determined by titration with quenching of fluorescence of the bis(2,2’-bithienyl)methane moiety of Ade-BTM by the FU titrant, in benzonitrile, at the 352-nm excitation. Next, FU-templated molecularly imprinted polymer (MIP-FU) films were deposited on the ITO or Au film coated glass slides, the Pt disk electrodes, or the 10-MHz quartz crystal resonators by potentiodynamic electropolymerization from solution of FU, Ade-BTM, and the tris([2,2’-bithiophen]-5-yl)methane, TTM, cross-linking monomer at the FU: Ade-BTM: TTM = 1:2:3 mole ratio. Then, the UV-vis and FTIR spectra of the MIP-FU films were recorded to confirm the FU template presence in the MIP-FU film and its subsequent release by extraction with methanol from this film. For determination of the stability constant of the complex of the MIP cavity and FU, piezoelectric microgravimetry (PM) under both batch- and flow-injection analysis conditions was used. For sensing application, three different transduction platforms, i.e., DPV, CI, and PM were integrated with the MIP-FU recognition unit. The limit of detection (LOD) was 56 nM, 75 nM, and 0.26 mM, for these chemosensors, respectively, indicating suitability of the former two for the FU determination in blood plasma or serum (~500 nM). Moreover, the CI chemosensor was appreciably more sensitive to FU than to their common interferences.

The 5-fluorouracil (FU), a fluorine derivative of uracil, contains pseudo-amide moieties like the parent uracil. Therefore, it is highly soluble in polar solvents, such as dimethylformamide (DMF), methanol, etc. FU dissociates in two steps with pKa,1 = 8.0 and pKa,2 = 13.0.1 For the last 50 years, FU has successfully been used in antitumor chemotherapy as a cytostatic agent for inhibition of the colorectal cancer as well as for treatment of head and neck malignancies.2 The antitumor activity of FU is based on the anabolic way with which FU is converted to its metabolites, vis., 5-fluoro-2’-deoxyuridine-5’monophosphate, 5-fluoro-2’-deoxyuridine-5’-diphosphate, or 5-fluorouridine-5’-diphosphate, thus inhibiting the thymidylate synthase, ultimately preventing deoxyribonucleic acid (DNA) replication. However, only 20% FU is activated for the anabolic way while 80% FU is

mainly catabolically degraded to 5,6-dihydro-5fluorouracil in liver by dihydropyrimidine dehydrogenase.3-4 Concentration of FU in, e.g., a blood serum or plasma of a patient after FU infusion is usually maintained in the range of 0.1 to 1.0 μM.3, 5 Therefore, sufficiently sensitive and selective sensors for determination of FU in body fluids are needed to improve the chemotherapy and predict some possible side-effects from the FU remains in human body. Most commonly, FU is determined using separation techniques under flow analytical conditions. First, capillary electrophoresis (CE) with the UV spectroscopic detection was applied for that.6 However, the 13-μM limit of detection (LOD) reached was insufficient for clinical applications. Therefore, liquid chromatography with mass spectrometry (LC-MS)3, 7 or the ultraviolet irradiation (LC-

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UV)8 detection was then used because of relatively low LOD, i.e., in the range of tens of nM. Recently, more sensitive CE procedures using large-volume sample stacking9 or FU enrichment by solid-phase extraction (SPE) on a hydrophobic column10 were developed. However, these separation techniques suffer from disadvantages, such as poor selectivity with respect to regioisomers and similar polarity interferences, relatively high solvent consumption, and necessity of taking safety precautions against the high-voltage operated CE. In batch FU determinations, a hundred-nM LOD was reached using fluorescence emission generated by reacting FU with the KMnO4 oxidant in the presence of formaldehyde,11 or fluorescence quenching of quantum dots or 9-anthracene carboxylic acid by FU.12-13 For the recent two decades, application of molecularly imprinted polymers (MIPs) for biomimetic recognition in selective chemosensing has steadily been increasing14-17 because of high MIP detectability, selectivity, and reproducibility. However, only few studies were performed towards FU recognition and detection.18-20 For this recognition, some new functional monomers were recently synthesized, including 2,6bis(acrylamido)pyridine and 2-(trifluoromethyl)acrylic acid, 2-hydroxyethylmetacrylate and acrylic acid or methacrylic acid and ethylene glycol dimethacrylate.18, 20 The affinity of these monomers to the FU target was evaluated by measuring the FU concentration in solution by UV-vis spectroscopy or by swelling of the MIP hydrogels after releasing FU from them. For fabrication of the MIP chemosensors for FU determination, the MIP using the 1,3-diacryloyl urea functional monomer was applied with differential pulse anodic stripping voltammetry for signal transduction.19 For selective FU recognition, we adopted herein the Watson-Crick type RNA nucleobase pairing of adenine and uracil.21-22 For that, first, we developed a preparation procedure and synthesized 4-[2-(6-amino-9H-purin-9yl)ethoxy]phenyl-4-[bis(2,2’-bithienyl)methane] AdeBTM, which served as the functional monomer. Structure of the pre-electropolymerization complex of Ade-BTM and FU was computationally optimized. Moreover, we experimentally determined the stability constant of this complex by fluorescence titration of Ade-BTM with the FU titrant in benzonitrile. Then, we deposited an MIP-FU film on a Pt, ITO glass slide electrode, or Au-coated quartz crystal resonator (QCR) by electropolymerization from a solution of FU, Ade-BTM, and the tris([2,2’-bithiophen]-5yl)methane TTM cross-linking monomer. Subsequently, the FU template was extracted from the MIP-FU film under dynamic methanol-extraction conditions. Then, the stability constant of the complexes of cavities of the FU-extracted MIP-FU film and the FU molecules was determined by piezoelectric microgravimetry (PM). Finally, we applied the capacitive impedimetric (CI), differential pulse voltammetric (DPV), and PM transduction for the FU determination under batch and flow-injection analysis conditions.

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EXPERIMENTAL SECTION Chemicals. Acetonitrile (ACN), 1,2-dichlorobenzene (DCB), benzonitrile (BN), FU, 5-fluorouridine (FUrid), thymine, cytosine, and all chemicals for syntheses were purchased from Sigma-Aldrich. Tetra-n-butylammonium perchlorate, (TBA)ClO4, was supplied by SACHEM. Ethanol, methanol, and potassium chloride (KCl) were from PHARMCO-AAPER, EMD, and Fisher, respectively. The 10-Ω/cm2 ITO (indium-tin oxide) glass slides were supplied by Delta Technologies. The cross-linking monomer, TTM, was prepared according to our described procedure.23 The procedure of synthesis of the functional monomer, Ade-BTM, is described in Appendix 1S and Scheme 1S in Supporting Information. Instrumentation and procedures. A broadband two-channel 400-MHz Varian NMR spectrometer, composed of a highly homogeneous superconducting magnet (9.4 T) and 4 nuclear (1H/9F/13C/31P) probes, was used for the 1H-NMR spectra recording. The 5-mm diameter NMR quartz tubes for samples were purchased from Wilmad-LabGlass. The NMR spectra were processed by the VNMR software of Varian installed on a workstation computer. A PARSTAT 2273 computerized electrochemistry system of PAR Princeton Applied Research (Tennessee, USA), equipped with the potentiostat/galvanostat/frequency-response-analyzer and controlled by the PowerSuite software of the same manufacturer, was used for the cyclic voltammetry (CV), DPV, and CI measurements. The PerkinElmer FTIR spectrophotometer was used for the IR spectra recording. It was configured to run in a single-beam, ratio, or interferogram mode. This spectrophotometer was capable of data acquisition over the wavenumber range of 370 to 7800 cm-1 with as low as 0.1 cm-1 resolution using a fast recovery deuterated triglycine sulfate (DTGS) detector with KBr splitting. Its signal-to-noise ratio was 35,000:1. The spectrophotometer was controlled by the Spectrum One HP software of Hewlett Packard, which allowed for recording of both qualitative and quantitative data. The UV-visible spectra were recorded with 0.1-nm resolution by using a UV-2550 spectrophotometer of Shimadzu Corp. (Tokyo, Japan). The Cary Eclipse fluorimeter was used for fluorescence spectra recording. The fluorimeter performance was validated using the instrument built-in test software. The recognition film was imaged by atomic force microscopy (AFM) using a MultiMode® 8 AFM under control of Nanoscope V controller of Bruker. The Tapping Mode mode was utilized for sample imaging with the use of the RTESP probes provided by Bruker. The films for imaging were deposited on Au-coated glass slides. The slides were cleaned with acetone, and then dried in an Ar stream prior to use. A model EQCM 5610 and EQCM 5710 quartz crystal microbalance24-25 controlled by the EQCM 5710-S2 software, all of IPC PAS, served to perform the PM experiments under either the batch or flow-injection


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Analytical Chemistry solvent in order to remove the supporting electrolyte solution. Then, the FU template was extracted from the film with methanol under vigorous magnetic stirring for 8 h. The extraction completeness was confirmed by the UV-vis and FTIR spectroscopic measurements. The same procedure was used to deposit a control nonimprinted polymer (NIP) film, except of no FU template in the solution for the electropolymerization. For the FTIR spectra recording, an ITO glass slide was drop-coated with an FU film from the 10-mM FU methanol solution. Calculations and data fitting. Molecular structure, electron density distribution, and values of thermodynamic functions of formation of the complex of FU with the functional monomer Ade-BTM in vacuum were optimized and calculated, respectively, by using the density functional theory (DFT) at the B3LYP/321G* level with the Gaussian 2009 software.26 For fitting of the FU sorption isotherms, three different (Langmuir, Freundlich, and Langmuir-Freundlich) models were adopted (Appendix 2S).

analysis (FIA) conditions, respectively. The resonant frequency change was measured with 0.1-Hz resolution using the 14-mm diameter AT-cut plano-plano 10-MHz resonant frequency QCRs with 5-mm diameter and ∼100nm thick circular Au-film electrodes evaporated over Ti underlayers on both sides of QCRs. However, only one QCR side was wetted by a working solution and the Aufilm electrode of this side was used as both the working electrode and the substrate for deposition of an MIP recognition film. For the FIA experiments, the FUextracted MIP-FU film-coated Au-QCRs were examined by PM. The carrier liquid was pumped at the 20 μL/min flow rate through the EQCM 5610 holder with a model NE-500 syringe pump of New Era Pump Systems. Sample solutions were injected with a model 7725i rotary six-port valve of Rheodyne. The FU samples were dissolved in water, i.e., in the same medium as that of the carrier liquid. For deriving the FU sorption isotherms, the FUextracted MIP-FU film-coated Au-QCRs were examined by PM with the EQCM 5710 microbalance under batchinjection analysis conditions. The MIP-FU film-coated QCRs were soaked in 30 mL of water under dynamic magnetic stirring condition. For determination of the sorption stability constant under FIA conditions, the EQCM 5610 microbalance was used. Volume of the injected sample was 500 μL. For FU determination under FIA conditions using PM, volume of each injection of the FU samples of different concentrations was 200 μL. For the batch analysis DPV determinations, the 1-mm diameter Pt disk electrodes, coated with the FU-extracted MIP-FU films, were immersed for 20 min in solutions of different concentrations of FU in ACN. Then, DPV curves were recorded for solution of 0.1 M K4[Fe(CN)6] in 0.1 M KCl. The potential scan range, potential step, pulse amplitude, and pulse duration was 0 to 0.5 V, 5 mV, 50 mV, and 50 ms, respectively. Moreover, the above FU-extracted MIP-FU film-coated Pt electrodes were used to determine FU by CI under batch-injection analysis conditions. A Pt wire and an Ag/AgCl, KCl (satd.) electrode served as the counter and reference electrode, respectively. The FU of different concentrations in 0.1 M KCl in water:ethanol (1:1, v:v), was used for injections. The applied frequency and potential was kept constant at f=20 Hz and E=0.50 V vs. Ag/AgCl, respectively. At this potential, no faradaic process occurred. Fabrication of the FU film as well as the imprinted (MIP-FU) and non-imprinted (NIP) films. The MIP-FU films were prepared by potentiodynamic electropolymerization with the potential linearly cycled between 0.50 and 1.60 V (for the Pt and ITO glass slide electrodes) or 0.50 and 1.30 V (for the Au-coated glass slides and QCR) vs. Ag/AgCl at the scan rate of 50 mV/s. The counter electrode was a 1-mm diameter Pt wire. The film growth on the electrodes was controlled with the number of potential cycles. After electropolymerization, the MIP-FU films were rinsed with the abundant ACN

RESULTS AND DISCUSSION Molecular modeling of interactions between binding sites of FU and recognizing sites of 1. Four binding sites of two pseudo-amide groups of FU can be recognized by two 9H-purine-6-amine moiety of two AdeBTM molecules (Scheme 1a), according to the electron density distribution in an isolated FU molecule (Scheme 2S). This distribution involves the electron enriched -C=O and electron depleted -N-H groups. The optimized structure of the FU:Ade-BTM = 1:2 complex (Scheme 1b) revealed four hydrogen bonds of two pseudoamide groups (oxygen atoms 6 and 10 as well as hydrogen atoms 8 and 11) of FU with two 9H-purine-6-amine substituents (nitrogen atoms 67 and 131 and hydrogen atoms 72 and 136) of the molecule Ade-BTM and AdeBTM’. That way, all recognition sites of FU were captured. Moreover, the N, H, and O atoms engaged in these hydrogen bonds lie in a straight line obeying the Corey-Pauling rules.27 Here, Ade-BTM’ stands for AdeBTM interacting with another pseudo-amide group of FU. The binding of FU and Ade-BTM’ is the exact mimic of the Watson-Crick nucleobase pairing in RNA. The Gibbs energy gain due to formation of the structurally optimized complex was calculated to be as high as-124.5 kJ/mol indicating higher stability of the complex molecule than that of the molecules of individual complex components alone. The total Gibbs energy gain consists of contributions of the gains of two 1:1 complexes of FU with Ade-BTM and Ade-BTM’. By separately calculating the gain of the FU-(Ade-BTM) and FU-(AdeBTM’) complexes, we found that contributions of two complexes are different,28 and equal to -65.7 kJ/mol (~53%) and -58.8 kJ/mol (~47%) for the FU-(Ade-BTM) and FU-(Ade-BTM’) complex, respectively. Furthermore, the Gibbs energy gain corresponding to the (Ade-BTM)-(Ade-BTM) dimer (Scheme 3S) was -38.6 kJ/mol, i.e., much lower than that of the 1:1 complex of FU


Analytical Chemistry with Ade-BTM or Ade-BTM’, given above. Conclusively, the exhaustive H-bond nucleobase pairing of Ade-BTM and FU is strong and very promising for molecular imprinting. Therefore, Ade-BTM was chosen as the functional monomer for the FU recognition.

inset). From the plot in Fig. 1b, stoichiometry of the FU: Ade-BTM complex was determined to be 1:2, as followed from the / Ade BTM mole ratio of ~0.36 (Fig. 4Sa). and Ade BTM is the equilibrium concentration of FU and Ade-BTM, respectively. The equilibrium of the complexation of FU and Ade-BTM can be presented, as follows.

S S

S S S

2Ade BTM FU ⇆ Ade BTM-FU-Ade BTM′ 1

TTM S S

S S

The complex stability constant was calculated according to the Scatchard dependence, Equation (2), for the 1:2 complex (Appendix 3S).30

S

S O O

N

H

N

S

2

N H

Ade-BTM

N

N

N

O H N

H

N

O

H

N

S

Ade-BTM'

where I0 and I is the fluorescence intensity of 1 before and after FU addition, respectively. Ade BTM$ is the initial concentration of Ade-BTM, here equal to 1 mM. Ks is the stability constant of the 1:2 complex. ∆&Ade BTM-FU-Ade BTM′ &Ade BTM-FU-Ade BTM′ &Ade BTM & , where &Ade BTM-FU-Ade BTM′ , &Ade BTM, and & is the molar absorptivity of the (Ade-BTM)-FU-(AdeBTM’) complex, Ade-BTM, and FU, respectively.

N

N N

FU

Ade-BTM

!"#$ Δ& !"#-- !"#' 2

H

F

136 10 131 11

67 8

Ade-BTM’ 4

72

0.05 mM FU

6

FU

Scheme 1. (a) Structural formula and (b) molecular structure of the complex of one molecule of FU and two molecules of Ade-BTM optimized by the B3LYP density functional theory method using the 3-21G* basis set. Ade-BTM’ is another AdeBTM interacting with another pseudo-amide group of FU.

3

1.26

3.6

Intensity / a.u.

S

Intensity / a.u.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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3.4 3.2 3.0 2.8 2.6 2.4 0.0 0.2 0.4 0.6 0.8 1.0 1.2

2

FU concentration / mM 1

Complexation of FU with Ade-BTM in benzonitrile. Although the concept of RNA nucleobase pairing is well understood, application of an artificial adenine derivative Ade-BTM as the functional monomer for imprinting of the uracil derivative remains challenging. This is because of high polarity of both those counterparts making them insoluble in most organic solvents. By linking 9H-purin6-amine to bis(2,2’-bithienyl)methane through the ethyl bridge (Scheme 1S), we decreased its polarity to make it more soluble in solvent of low polarity. Therefore, BN was chosen as a solvent for fluorescent titration. Moreover, this ethyl makes the molecule of Ade-BTM more flexible. Hence, the steric hindrance is lowered and molecules of the FU target analyte can more readily interact with the imprinted cavities. After recording the UV-vis spectrum for 0.25 mM AdeBTM in BN (Fig. 3S), we recorded the emission spectrum of Ade-BTM by exciting Ade-BTM at the 352-nm absorption peak maximum of bis(2,2’-bithienyl)methane. Then, titration of Ade-BTM with the FU titrant resulted in quenching of the emission intensity of excited AdeBTM likely due to the electron transfer from the bis(2,2’bithienyl)methane moiety of Ade-BTM to FU (Fig. 1).29 In this titration, Ade-BTM played a role of the analyte in view of no fluorescence of the FU titrant and the complex at the 352-nm excitation. The emission intensity of AdeBTM decreased during the titration and virtually reached saturation after making the solution ~0.8 mM in FU (Fig 1,

0

400

450

500

550

Wavelength / nm

Figure 1. The steady-state fluorescence spectra of 1 quenched by the FU titrant, in BN; excitation at 352 nm. Concentration of Ade-BTM was 1 mM and that of FU ranged from 0.05 to 1.26 mM. Inset presents a plot of the change in the fluorescence intensity with the change of the FU concentration.

By linear fitting to the data points, according to Equation (2), of the plot of /2 versus , we arrived to the linear regression equation of /2 2.167 , 10. 2.390 , 10. with the correlation coefficient of 0.97 and the slope equal to the Ks value of 2.17(±0.07)×107 M-2 (Fig. 4Sb). In view of this appreciably high value,31 the 1:2 complex of FU and AdeBTM could be transferred, in a form of the FU-templated cavities of the MIP-FU film, onto the electrode. Deposition of the MIP-FU films on electrodes by potentiodynamic electropolymerization. First, electrochemical behavior of FU and Ade-BTM in 0.1 M (TBA)ClO4, in ACN:DCB (1:1, v:v) was characterized by CV. Accordingly, FU was not oxidized in the positive potential range (curve 1 in Fig. 5S). However, an anodic peak of irreversible Ade-BTM oxidation was observed at ~1.25 V because of radical electropolymerization of the


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bis(2,2’-bithienyl)methane moiety (curve 2 in Fig. 5S). Therefore, an MIP-FU film was deposited at this more positive potential on the one hand and FU was not oxidized on the other. Hence, the MIP-FU film was templated, desirably, with FU and not with its oxidation product.

FU film, the more positively shifted was the anodic peak potential, and the more decreased was the anodic peak current of the bis(2,2’-bithienyl)methane moiety oxidation. These changes were due to the growth of the MIP-FU film, which played a role of the self-barrier of increasing resistance, which prevented further electrooxidation of the bis(2,2’-bithienyl)methane moiety. The color of the Pt electrode changed from silver-white to green-yellow after the MIP-FU film deposition. Similarly, the FU-templated MIP-FU film was deposited on the Au-coated QCR from the same electropolymerization solution above (Fig. 2a). However, potential was limited in the range of 0.50 to 1.30 V to prevent electro-oxidation of the Au electrode layer. Apparently, the anodic current and potential of bis(2,2’bithienyl)methane electro-oxidation decreased and positively shifted, respectively, after each consecutive potential cycle because of formation of a resistive film. Moreover, the growth of the MIP-FU film was manifested by appreciable resonant frequency decrease (~10 kHz) of the QCR (Fig. 2b). Furthermore, the decrease of the resonant frequency change after each potential cycle was observed indicating the more resistive the MIP-FU film was, the smaller amount of MIP was deposited. According to the Sauerbrey equation,25 one can calculate mass of the FU-templated MIP-FU film from the resonant frequency change, ∆1,

Current / A

0.4

a

1

0.3 0.2

10

0.1

Resonant frequency change / kHz

0.0

Dynamic resistance change / Ω

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


b 0 -3 -6 -9

c 0 20 40 60 0.5

0.6

0.7

0.8

0.9

1.0

1.1

1.2

1.3

∆1

Potential / V vs. (Ag/AgCl)

Figure 2. Simultaneously recorded curves of (a) the potential dependence of current as well as the change of (b) resonant frequency, and (c) dynamic resistance recorded using the Au-film electrode of the 10-MHz QCR for deposition of the MIP-FU film by potentiodynamic electropolymerization from 0.5 mM FU, 1 mM Ade-BTM, and 1.5 mM TTM in 0.1 M (TBA)ClO4 in ACN:DCB (1:1, v:v), during 10 potential cycles. The cycle numbers are indicated at curves. The potential scan rate was 50 mV/s.

21 ∆2 3 345 67 87 9/

where f0 is the fundamental frequency of the resonator (here 10 MHz), Aac is acoustically active area of the 2 resonator (here 0.1963 cm ), µq is the shear modulus of 11

-1

quartz (2.947×10 g s-2 cm ), and ρq is the quartz density -3

(2.648 g cm ). For such a ~161(±4)-nm thin film (Fig. 4a), changes in visco-elasticity with respect to the dynamic resistance change of ~52 Ω (Fig. 2c) can be neglected.34 As a result, the calculated total mass of the FU-templated MIP-FU film deposited was ~9.15 μg. For that, density of the MIP-FU film, ρ = Δm/δA, was calculated to be 1.08±0.03 g cm-3; δ and A is the film thickness and area, respectively. The film thickness was determined from the AFM imaging by measuring the height of the step formed by removing some part of the film from the electrode surface. Moreover, density as well as pre-concentration of FU bound to the FU-extracted MIP-FU film was determined with respect to adsorption isotherm (discussed below). Topography of the 161-nm MIP-FU film was analyzed by using AFM (Fig. 3). It appeared that the FU-templated MIP-FU film was composed of 20-nm diameter well distinguishable grains. The polymer film is highly homogenous with respect to the determined film roughness of 1.1(±0.1) nm.

Then, a solution for the MIP-FU film deposition was prepared. Its composition was adjusted to be similar to that used for the fluorescent titration, i.e., it was 0.5 mM in FU, 1 mM in Ade-BTM, 1.5 mM in TTM, and 0.1 M in (TBA)ClO4 in ACN:DCB (1:1, v:v). Here, the solution was spiked with 50% DCB not only to pull back polarity of ACN to be closer to that of BN for strengthening the FU(Ade-BTM) hydrogen bonds but also to dissolve TTM completely. Furthermore, a large excess of the crosslinking monomer TTM was added in order to interconnect molecules of the functional monomer AdeBTM. Moreover, a rigid MIP-FU matrix with molecular cavities not shrinking after template extraction was formed that way.32 Besides, the (TBA)ClO4 supporting electrolyte was used both for eliminating ionic migration during the electropolymerization and, due to large molecular size of its ions, for incurring porosity of the MIP-FU film.17, 33 Next, the FU-templated MIP-FU films were deposited on the Pt disk electrodes (Fig. 6S) and on the ITO glass slides (Fig. 7S) by potentiodynamic electropolymerization. Furthermore, the thicker the MIP-



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of different concentrations of FU were injected. The stirring enhanced transport of FU in the solution and, indirectly, that in the MIP-FU film. After each injection, resonant frequency decreased because FU molecules entered cavities of the FU-extracted MIP-FU film increasing its mass (Fig. 10Sa). By knowing the initial mass and volume of the FU-extracted MIP-FU film and mass change of the film after injection of each FU sample solution, one can calculate mass and concentration of the FU bound in the MIP-FU film (Fig. 10Sb) (see Eqn. (3) above). For that, the maximum concentration of the FU imprinted cavities in the MIP-FU film accessible for FU, ,; , was determined as ~30.7 mM, that is much higher than that in solution. At this concentration, the isotherm of the FU sorption by the MIP-FU film reached plateau, indicating that the FU molecules filled all imprinted cavities available in the MIP-FU film.

Figure 3. Atomic force microscopy (AFM) images of the MIP-FU film deposited on the Au-coated glass slide of 1×1 2 μm surface area.

Extraction of the FU template from the MIP-FU film. The FU template was extracted from the FUtemplated MIP-FU films with methanol because it is well soluble in this solvent. Completeness of this extraction was confirmed by the UV-vis and FTIR spectroscopy measurements, as described below. The extracting methanol solution was examined by UVvis spectroscopy (curve 1 in Fig. 8S). The absorption peak of the extracted FU appeared at 265 nm, as confirmed by the control spectrum recorded for 1 mM FU in methanol (Fig. 8S, inset). After the extraction, the FU peak was absent for another fresh extracting solution (curve 2 in Fig. 8S), i.e., no more FU was extracted from the MIP-FU film. Moreover, the FTIR spectra of different polymer films deposited onto the ITO glass slides were recorded in the range of 450 to 4000 cm-1. The spectra in the range of 500 to 2000 cm-1 were examined for easy detection of FU in the MIP-FU film (Fig. 9S). The control spectrum of the FU film deposited by drop-casting onto the ITO glass slide was characterized by the peak at 1646 cm-1 corresponding to stretching vibrations of the C=O bond of the pseudo-amide groups (curve 1 in Fig. 9S).19, 35 Because only the FU component of the MIP-FU film contained the C=O groups, the signal at 1646 cm-1 was used as the internal marker to confirm the presence of FU in the film. Besides, the sharp high-intensity peak at 1684 cm-1 in the spectrum for the NIP film was assigned to bending vibration of the N-H bond of the primary amine group of the adenine moiety of Ade-BTM (curve 2 in Fig. 9S).36 Expectedly, the peak at 1646 cm-1 was clearly seen in the spectrum of the MIP-FU film before FU extraction (curve 3 in Fig. 9S) confirming the presence of FU. Evidently, the complex of FU and Ade-BTM was successfully transferred onto the electrode by the electropolymerization. After FU extraction, the 1684 cm-1 peak reappeared in expense of the peak at 1646 cm-1, which vanished (curve 4 in Fig. 9S), proving that FU was completely extracted from the MIP-FU film. Characterization of the FU sorption by the MIP-FU film under batch- and flow-injection analysis conditions Derivation of the FU sorption isotherms for the MIP-FU film under batch analysis conditions. The FU-extracted MIP-FU film deposited on the 10-MHz QCR was soaked in water under vigorous stirring conditions. Then, solutions

Table 1. Isotherm-fitting parameters for the FU sorption by the FU-extracted MIP-FU film with different adsorption models. Fitting models

K/M

-1

-1

N / μmol g

Correlation coefficient

(±st.d.) Freundlich

—

—

0.955

Langmuir

402.1±41.6

39.5

0.990

LangmuirFreundlich

481.6±41.0

69.5

0.994

K – stability constant of the complex of the FU molecule and the imprinted cavity determined by different adsorption isotherm models; N – density of the imprinted molecular cavities.

For curve fitting, three commonly used sorption isotherms, i.e., those of Langmuir, Freundlich, and Langmuir-Freundlich,37 were used (Appendix 2S and Fig. 10Sb) and the fitted parameters were summarized in Table 1. Among them, the Langmuir-Freundlich isotherm with its correlation coefficient exceeding 0.994 better fits experimental data than that of Langmuir and Freundlich with correlation coefficient of 0.990 and 0.955, respectively (Fig. 10Sb). This indicates that our MIP-film is not homogenous, i.e., there may be defects in the MIPFU film incurred by electropolymerization or template extraction; for instance, by impurity imprinted during the electropolymerization or by shrinking of the film caused by the FU extraction. Sorption of FU by the MIP-FU film under FIA conditions using PM. Another FU-extracted MIP-FU film deposited on the 10-MHz QCR was prepared to measure resonant frequency changes under dynamic (FIA) conditions.38-39 Herein, large volume of the injected sample (500 μL) was used to reach equilibrium of FU loading. After each injection of the FU solution of different concentration, decrease of ∆f (Fig. 11S) is similar to that in batch-analysis condition. However, shape of the frequency change was peak-wise due to reversible binding and rebinding of FU to MIP cavities. The ∆f decrease was proportional to the FU concentration in solution in the range of 0.5 to 2.0 μM with the linear regression equation of ∆f / Hz =


0.17(±0.09)−1.27(±0.16) / mM. The sensitivity and correlation coefficient was (1.27±0.16) Hz μM-1 and 0.94, respectively. Moreover, the sorption stability constant determined from the FIA measurements23 was 319 M-1, i.e., it was lower than that derived from the analysis of all isotherms under the batch-injection conditions. Moreover, the initial slope of the dependence of the FU concentration in film on that in solution (dash line in inset to Fig. 10Sb) indicates nearly tenfold preconcentration of FU of FU in the MIP-FU film. Determination of FU using MIP-FU films and different transduction platforms. For application of the MIP-FU film for determination of FU, three different transduction schemes were used, vis., DPV, CI, and PM under the batch, batch-injection, and flow-injection analysis conditions, respectively. Differential pulse voltammetry (DPV) under batch conditions. For the FU determination under the batch analysis conditions, an indirect DPV procedure was adopted. It was similar to that used in our former determination of adrenaline.23 The FU-extracted 161-nm thick MIP-FU film-coated Pt electrode was used for recording the change of the DPV peak current of the 0.1 M K4Fe(CN)6 redox probe (Fig. 12S). This procedure exploits the increase of the MIP-FU film resistance due to the analyte occupation of the imprinted cavities causing the decrease of the electro-oxidation DPV peak of the probe.40 As expected, this peak was progressively lower after each 20-min film soaking in the ACN solutions of the progressively higher concentration of FU. That is, the higher the fraction of imprinted cavities occupied by the FU molecules the more resistive was the film and the lower was the DPV peak. With respect to the linear dynamic range of the FU concentration, which extended from 20 to 400 nM, the linear regression equation of the calibration plot (Fig. 10S, inset) and the correlation coefficient of the calibration plot was (IDPV,e – IDPV,s) / μA = 0.075(±0.03) + 0.791(±0.015)×10-3cFU / μM and 0.88, respectively. Here, the IDPV,e and IDPV,s is the DPV peak current for the MIPFU film with the FU template extracted and for the FUextracted MIP-FU film soaked with the FU analyte, respectively. The sensitivity of the MIP-FU film to FU was 0.791(±0.015) µA nM-1. At the signal-to-noise ratio, S/N = 3, the detectability was appreciably high reaching LOD of 56 nM FU. Therefore, this DPV chemosensor can readily detect FU at a ~500 nM biological level. Capacitive impedimetry (CI) under the batch-injection analysis conditions. The same FU-extracted MIP-FU filmcoated Pt electrode as that described above, was prepared for the CI determination of FU under batch-injection analysis conditions. The double-layer capacity, Cdl, was determined using Equation (4) from the measured imaginary component of impedance, Z'', for the electrode of the surface area, A (~0.8 mm2), at the constant potential and angular frequency, ω=2πf, applied. 1 ?′′ 4 @AB 3

Expectedly, this capacity increased after each consecutive injection of a sample of the FU solution (Fig. 4a). In view of almost no contribution of the capacity of the diffuse layer to the total interfacial capacity measured at the relatively high (0.1 M KCl) concentration of the supporting electrolyte solution used, the measured capacity practically corresponded to the capacity of the compact part of the electrical double layer.41 Therefore, the compact layer was approximated by the Helmholtz model with its capacity solely depending on electric permittivity, &, of the MIP-FU film described by Equation (5). && AB 5 D where ε0 is electric permittivity of free space and d is the thickness of the compact part of the double layer. Time / min 0

20

40

60

80

a -2

Capacity / mF cm

100

2

0.1010

120

3 µM FU

1.6 1.2 0.9 0.6 0.4

0.1000

0.1 0.0995

FU concentration / µM 0.0

0.5

1.0

1.5

2.0

2.5

3.0

-2

0.1005

1.5 1.2 0.9 0.6 0.3

1.5

b

-2

Capacity change / µF cm

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Capacity change / µF cm

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1

1.2

0.9

2 3

0.6

4 0.3

0.0 0.0

0.5

1.0

1.5

2.0

Analyte or interference concentration / µM

Figure 4. (a) The double-layer capacity change with time during injections of 0.1 M KCl solutions of different FU concentrations (final FU concentrations in solution are indicated with numbers at each step) under batch-injection analysis conditions, measured at 0.50 V and 20 Hz for the FU-extracted MIP-FU film-coated 1-mm diameter Pt disk electrode. The MIP film was prepared as described in caption to Fig. 3a. Inset shows the resulting FU calibration plot. The initial volume of 0.1 M KCl was 5 mL. (b) Calibration plots for injections of 0.1 M KCl solutions of (1) FU, (2) cytosine, (3) thymine, and (4) FUrid, measured under the conditions described in (a).

The permittivity change is expected to be proportional to the MIP-FU film resistance, which in turn is proportional to the decrease of the DPV peak current


Analytical Chemistry when the FU molecules enter the imprinted empty cavities of this film. In effect, the capacity increases due to the ingress of the FU molecules to the FU-extracted MIP-FU film. Moreover, this capacity increase was steprather than peak-wise, the latter being typical for determination of an analyte reversibly bound to an MIP under the FIA conditions, because the FU concentration in the MIP film increased and stayed constant with the increase of the FU concentration in solution. The linear dynamic concentration range extended from 0.1 to 1.6 μM (Fig. 4a and 4b, inset) obeying the regression equation of Cdl / (µF cm-2) = 0.15(±0.05) + 0.76(±0.05) cFU / μM. The sensitivity and correlation coefficient was (0.76±0.02) µF cm-2 μM-1 and 0.98, respectively. At S/N=3, LOD of this CI chemosensor was 75 nM FU, i.e., sufficiently low for determination of FU in biological systems (~500 nM). The CI chemosensor was selective with respect to the most typical interferences (Scheme 4S). That is, the sensitivity to FU was more than five times that to FUrd, (0.14±0.02) µF cm-2 μM-1, three times that to thymine (0.28±0.02) µF cm-2 μM-1, and two-and-a-half times that to cytosine, (0.3±0.03) µF cm-2 μM-1 (curves 2-4 in Fig. 4b). The high selectivity of the MIP-FU film to FUrid is most plausibly because the FUrd molecules are too big to enter the cavities of the FU-extracted MIP-FU film. Piezoelectric microgravimetry (PM) under flow-injection analysis conditions. Another MIP-FU film was deposited on the Au film electrode of the 10-MHz QCR for PM determination of FU in solution. Herein, the change of the dynamic resistance (less than two ohms) upon interaction of FU with the MIP surface was negligible. Apparently, the FU binding in the film was fully reversible (Fig. 5). The linear dynamic concentration range was 0.25 to 2.00 mM obeying the regression equation of with the ∆f / Hz = 0.33(±0.04) − 6.47(±0.39)cFU / mM correlation coefficient of 0.98 (curve 1 in Fig. 5, inset). At S/N = 3, detectability of the PM chemosensor, LOD = 0.26 mM, was much higher than that of the DPV and CI chemosensors with the 6.47(±0.39) Hz/mM sensitivity. To confirm the imprinting, an NIP film was used for the FU determination in a control experiment (curve 2 in Fig. 5, inset). In view of no imprinted cavities emptied by the template extraction, the FU binding by NIP was much weaker than that by the FU-extracted MIP-FU, as manifested by only minor decrease of the resonant frequency with the FU concentration increase in solution. That is, the Δf for the former was proportional to cFU between 0.25 to 2.00 mM obeying the linear regression equation of ∆f / Hz = –0.42(±0.19) – 1.89(±0.16)cFU / mM; the correlation coefficient being 0.97. By calculating the ratio of slopes of the calibration curves for the MIP-FU and NIP film, one arrives to the apparent imprinting factor, which here was ~3.6.

Time / min 50

100

150

200

2.00 mM 0.25 0.50 0.75 1.00 1.25 1.50 1.75 FU

3

0

-3 FU concentration / mM 0.5

1.0

1.5

2.0 0

2

-3 -6 -9

1 -12

-6

-9

-12

Resonant frequency change / Hz

0

Resonant frequency change / Hz

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 12

Figure 5. The resonant frequency change with time for the FU-extracted MIP-FU film coated 10-MHz QCR, after injection of the 200-μL samples of aqueous solutions of FU of different concentrations indicated with numbers at the curve. Inset is the calibration plot for FU recorded by (1) MIP-FU and (2) NIP films. The flow rate of water used as the carrier liquid was 20 μL/min.

CONCLUSIONS High selectivity of the Watson-Crick RNA nucleobase pairing was successfully implemented for interaction of the artificial adenine derivative, a moiety of Ade-BTM, with FU. The synthetic route of Ade-BTM opened new possibilities of preparation of other nucleobase derivatives, i.e., thymine, guanine, cytosine, and uracil, of the bis(2,2’-bithienyl)methane for extensive sensing applications, e.g., determination of the nucleic acid cancer biomarkers. The functional monomer Ade-BTM is unique due to high selectivity of the recognizing adenine moiety, flexibility of the ethyl chain bridge, and electropolymerizability of the bis(2,2’-bithienyl)methane moiety. Complexation of Ade-BTM and FU was theoretically modeled by the DFT calculations and experimentally verified by the fluorescence titration. Although relatively polar solvent, benzonitrile, was used for this titration, binding of FU and Ade-BTM was quite appreciable leading to the stability constant of 2.17(±0.07)×107 M-2 for the FU-(Ade-BTM) complex of the 1:2 stoichiometry. Potentiodynamic electropolymerization conveniently lead to deposition of the FU-templated MIP-FU film with no need of the use of any polymerization initiator, heat, or UV-light irradiation. Moreover, this polymerization featured high reproducibility, well-adhering film formation, and easy control of the film thickness. The FTIR and UV-vis spectra proved that structure of the pre-polymerization complex self-assembled in solution was preserved during the electropolymerization and that the FU template was then exhaustively extracted from the resulting MIP-FU film, respectively. The stability constants of complexes of FU with the MIP-FU imprinted cavities, determined under steadystate conditions by adopting the Langmuir, Freundlich, or Langmuir-Freundlich models, were compared to that determined under PM dynamic conditions without


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Analytical Chemistry 4. Casale, F.; Canaparo, R.; Serpe, L.; Muntoni, E.; Pepa, C. D.; Costa, M.; Mairone, L.; Zara, G. P.; Fornari, G.; Eandi, M., Pharmacol. Res. 2004, 50, 173-179. 5. Furuya, Y.; Yamamoto, K.; Kohno, N.; Yamamoto, M.; Saitoh, Y., Cancer Lett. 1995, 94, 207-211. 6. Prochazkova, A.; Liu, S.; Friess, H.; Aebi, S.; Thormann, W., J. Chromatogr. A 2001, 916, 215-224. 7. Remaud, G.; Boisdron-Celle, M.; Morel, A.; Gamelin, A., J. Chromatogr. B 2005, 824, 153-160. 8. Micoli, G.; Turci, R.; Arpellini, M.; Minoia, C., J. Chromatogr. B 2001, 750, 25-32. 9. Yang, Y.; Liu, Q.; Tao, W.; Nie, L.; Yao, S., J. Sep. Sci. 2007, 30, 3296-3301. 10. Mahnik, S. N.; Rizovski, B.; Fuerhacker, M.; Mader, R. M., Anal. Bioanal. Chem. 2004, 380, 31-35. 11. Sun, H.; Li, L.; Chen, X., J. Clin. Lab. Anal. 2007, 21, 213-219. 12. Zhu, D.; Chen, Y.; Jiang, L.; Geng, J.; Zhang, J.; Zhu, J.-J., Anal. Chem. 2011, 83, 9076-9081. 13. Khot, M. S.; Bhattar, S. L.; Kolekar, G. B.; Patil, S. R., Spectrochim. Acta A 2010, 77, 82-86. 14. Alexander, C.; Andersson, H. S.; Andersson, L. I.; Ansell, R. J.; Kirsch, N.; Nicholls, I. A.; O’Mahony, J.; Whitcombe, M. J., J. Mol. Recognit. 2006, 19, 106-180. 15. Suriyanarayanan, S.; Cywinski, P. J.; Moro, A. J.; Mohr, G. J.; Kutner, W., in Top. Cur. Chem. Springer, 2010, vol. 325, pp 165266. 16. Sharma, P. S.; D'Souza, F.; Kutner, W., Trends Anal. Chem. 2012, 34, 59-77. 17. Sharma, P. S.; Pietrzyk-Le, A.; D'Souza, F.; Kutner, W., Anal. Bioanal. Chem. 2012, 402, 3177-3204. 18. Kugimiya, A.; Mukawa, T.; Takeuchi, T., Analyst 2001, 126, 772-774. 19. Prasad, B. B.; Kumar, D.; Madhuri, R.; Tiwari, M. P., Electrochim. Acta 2012, 71, 106-115. 20. Singh, B.; Chauhan, N., Acta Biomater. 2008, 4, 1244-1254. 21. Watson, J. D.; Crick, F. H. C., Nature 1953, 171, 737-738. 22. Huynh, T.-P.; Pietrzyk-Le, A.; C., C. B. K.; Noworyta, K.; Sobczak, J. W.; Sharma, P. S.; D’Souza, F.; Kutner, W., Biosens. Bioelectron. 2013, 41, 634-641. 23. Huynh, T.-P.; K.C., C. B.; Lisowski, W.; D'Souza, F.; Kutner, W., Bioelectrochemistry 2012, DOI: 10.1016/j.bioelechem.2012.07.003. 24. Kochman, A.; Krupka, A.; Grissbach, J.; Kutner, W.; Gniewinska, B.; Nafalski, L., Electroanalysis 2006, 18, 2168–2173. 25. Buck, R. P.; Lindner, E.; Kutner, W.; Inzelt, G., Pure Appl. Chem. 2004, 76, 1139–1160. 26. Frisch, M. J.; al., e. Gaussian, Inc.: Wallingford CT, 2009. 27. Pauling, L.; Corey, R. B., Proc. Natl. Acad. Sci. 1951, 37, 235240. 28. Ran, J.; Hobza, P., J. Phys. Chem. B 2009, 113, 2933-2936. 29. Barbarella, G.; Zambianchi, M.; Sotgiu, G.; Ventola, A.; Galeotti, M.; Gigli, G.; Cazzato, A.; Capobianco, M. L., J. Noncryst. Solids 2006, 352, 2465-2467. 30. Atwood, J. L.; Davies, J. E. D.; Macnicol, D. D.; Vogtle, F., Comprehensive supramolecular chemistry. 1st ed.; Pergamon: Oxford, 1999; Vol. 8. 31. Rai, R.; Pandey, P. S., Bioorg. Med. Chem. Lett. 2005, 15, 2923-2925. 32. Karim, K.; Breton, F.; Rouillon, R.; Piletska, E. V.; Guerreiro, A.; Chianella, I.; Piletsky, S. A., Adv. Drug Deliver. Rev. 2005, 57, 1795– 1808. 33. Roncali, J., Chem. Rev. 1992, 92, 711-738. 34. Pieta, P.; Obraztsov, I.; Sobczak, J. W.; Chernyayeva, O.; Das, S. K.; D’Souza, F.; Kutner, W., J. Phys. Chem. C 2013, 117, 1995-2007.

adopting any sorption model. This comparison prompts us to speculate that the derived under the latter conditions stability constant, smaller than those derived from the isotherms, may indicate some inhomogeneity in the film. Due to low LOD, both the DPV and CI chemosensors are suitable for determination of FU in the human blood plasma. Between them, detectability of the indirect DPV chemosensing of FU is higher revealing the 56-nM LOD. The CI chemosensing with its 75-nM LOD, is also stable with respect to signal transduction and, moreover, it is practical in use. Both these detectability values suffice for clinical FU assays. Furthermore, the CI chemosensor was substantially more sensitive to FU than to its FUrid metabolite as well as thymine and cytosine interferences. Therefore, the CI chemosensor is applicable for determination of FU in biological fluids. Despite that LOD, for the PM chemosensor is relatively high (~0.26 mM), this LOD is sufficient for determination of FU in real samples by for the MIP-FU film of the 3.6-imprintingfactor.

ASSOCIATED CONTENT Supporting Information available: Preparation of 4-[2-(6amino-9H-purin-9-yl)ethoxy]phenyl-4-[bis(2,2’-bithienyl) methane] Ade-BTM; isotherm models and data; NMR, mass, and UV-vis spectra of Ade-BTM, deposition and FTIR spectra of MIP films, DPV data. This material is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author

* Francis D'souza, e-mail adresses: [email protected], tel.: +940-369-8832, fax: +(48 22) 343 3333 * Wlodzimierz Kutner, e-mail adresses: [email protected], tel.: +48 22 343 3217, fax: 940 565 4318

ACKNOWLEDGMENT We thank M.Sc. Chandra B. KC. for helpful discussion on the synthesis of Ade-BTM. The present research was financially supported by the Foundation for Polish Science (Project No. MPD/2009/1/styp19) to TPH, the European Regional Development Fund (Project ERDF, No. POIG.01.01.02-00008/08 2007-2013) to WK, the European Union 7.FP (Grant REGPOT-CT-2011-285949-NOBLESSE) to PP, and the US National Science Foundation (Grant No. 1110942) to FD. Access to AFM was funded by the Foundation for Polish Science under the FOCUS Grants no FG 3/2010.

REFERENCES 1. Dawson, R. M. C., Data for Biochemical Research. 3rd ed.; Clarendon Press: New York, 1989; p 592. 2. Heidelberger, C.; Chaudhuri, N. K.; Danneberg, P.; Mooren, D.; Griesbach, L.; Duschinsky, R.; Schnitzer, R. J.; Pleven, E.; Scheiner, J., Nature 1957, 179, 663-666. 3. Breda, M.; Barattè, S., Anal. Bioanal. Chem. 2010, 397, 11911201.



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35. Silverstein, R. M.; Webster, F. X.; Kiemle, D. J., Spectrometric Identification of Organic Compounds. 7th ed.; John Wiley & Sons: Westford MA, 2005. 36. Mathlouthi, M.; Seuvre, A.-M., Carbohyd. Res. 1984, 131, 115. 37. Garcıa-Calzon, J. A.; Dıaz-Garcıa, M. E., Sens. Actuators, B 2007, 123, 1180–1194. 38. Skladal, P., J. Braz. Chem. Soc. 2003, 14, 491-502.

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39. Pietrzyk, A.; Suriyanarayanan, S.; Kutner, W.; Maligaspe, E.; Zandler, M. E.; D'Souza, F., Bioelectrochemistry 2010, 80, 6272. 40. Gong, J.-L.; Gong, F.-C.; Zeng, G.-M.; Shen, G.-L.; Yu, R.-Q., Talanta 2003, 61, 447-453. 41. Bard, A. J.; Faulkner, L. R., Electrochemical methods: Fundamentals and Applications. 2nd ed.; John Wiley & Sons: New Jersey, 2001.


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