ARTICLEElectron Transfer in Methylene Blue-labelled G3 Dendrimers Tethered to GoldIsabel Álvarez-Martos[a], Andrey Kartashov[a] and Elena E. Ferapontova*[a]

Redox-modified branched 3D dendrimeric nanostructures are considered as a proper tool for wiring of redox enzymes, by providing both the enzyme-friendly environment and exquisite electron transfer (ET) mediation. ET rates in G3 poly(amido)amine (PAMAM) dendrimers, covalently attached to gold electrodes and labelled with methylene blue (MB), approach 267 s-1 and decrease with the increasing packing density of dendrimers on the electrode surface. Mechanistic analysis of the ET kinetics and its fitting to the Marcus relation showed that with the increasing PAMAM surface coverage the ET mechanism switches from surface-confined ET (electron tunneling) in diluted monolayers to diffusional ET (electron hopping) at higher surface populations of dendrimers. Structural changes in positively charged dendrimers electrostatically compressed at negative charges of the electrode surface and their dependence on the dendrimer surface packing contribute to both. Electrical wiring of horseradish peroxidase and hexose oxidase by MB-labelled dendrimers allowed bioelectrocatalytic reduction of H2O2

and oxidation of glucose by those enzymes. Demonstrated here electrical communication between MB groups, localised on the periphery of dendrimers and 2-3 nm distanced from the electrode surface, and the electrodes open new ways of their perspective biosensor and bioelectronic applications.

Introduction

Redox-labelled branched dendrimeric nanostructures, with their sizes compatible with those of biological macromolecules, may be considered as an excellent tool both for studies of electron transfer (ET) reactions in hybrid macromolecular assemblies and for programmable design of bioelectronic and biosensor devices exploiting their co-assemblies at electrodes.[1] Numerously discussed bio-compatibility of dendrimers[2] allows saving the structural integrity of enzymes at dendrimer-modified electrodes and, as a result, their bioelectrocatalytic functioning, though at a higher complexity level compared to direct electronic wiring (enzyme-electrode coupling with no mediators).[3] In particular, the possibility of electronic wiring of the active sites of redox enzymes by ferrocene (Fc)- [4] and methylene blue (MB)-labelled

dendrimers,[5] as alternative to “reagentless” wiring by conductive polymers [6], Os complex-conjugated polymers [7], and electrode-tethered redox relays [8] has been demonstrated. From strategic point of view, efficient wiring of enzymes by the dendrimer relays may be of a special biotechnological value and allow design of artificial ET chains and metabolic enzymatic cascades with required properties,[9] once a proper electronic inter-communication between enzymes, dendrimers and electrodes is established. Redox-modified branched 3D dendrimeric nanostructures are indeed perfectly suited for that, with their controllable topology, precise position and number of a variety of redox labels that can be used in their modification, and straightforward labelling and handling protocols.[1]

In this respect, mechanisms of ET in redox-labelled dendrimers should play an important if not critical role in their ability to electrically wire biomolecules to electrodes. By ultrafast voltammetric analysis of Ru (II) terpyridine ([Ru(tpy)2]2+)-modified synthetic G4 poly(amido)amine (PAMAM) dendrimers adsorbed on ultramicroelectrodes Amatore et al. demonstrated that electron hopping between the terminal [Ru(tpy)2]2+ groups of dendrimers is the dominant way the electron transport occurred over the spherical dendrimer shell.[10] To allow random hopping of electrons between all terminal groups involved in ET, over the dendrimer shell, the dendrimer branches should be essentially mobile to enable direct contact/collisions between the redox sites. That was shown to be facilitated in the distorted dendrite structures, transformed from the spherical to rather hemispheric shape on the electrode surface,[11] the latter being confirmed by scanning tunneling microscopy[12] and AFM[13] studies. A partial modification of the dendrimer peripheral groups by redox pendants or protein molecules coming into direct contact with the dendrimer shell can affect the ET and wiring efficiency. [4]

The ET reaction may be then confined to the electron exchange solely between the redox groups in the electrode proximity, affecting bioelectrocatalysis as well. Here, we have studied ET reactions in third generation (G3) PAMAM dendrimers covalently coupled to gold electrodes by alkanethiol-peptide chemistry and “on-surface” modified with methylene blue (MB) redox groups (Figure 1). Kinetics of ET in the MB-terminated dendrimers was studied by cyclic voltammetry (CV), and the results were assessed within the Butler-Volmer[14] and the Marcus semi-classical theory taking into account the reorganization energy contribution to the activation energy of the ET reaction.[15] Bioelectrocatalysis of two redox enzymes, heme-containing horseradish peroxidase (HRP)[16] and FAD-dependent hexose oxidase (HOX),[17] wired by MB-dendrimers was interrogated.

[a] Dr. Isabel Álvarez-MartosMr. Andrey KartashovDr. Elena FerapontovaInterdisciplinary Nanoscience Center (iNANO), Aarhus UniversityGustav Wieds Vej 14, DK-8000 Aarhus C, Denmark E-mail: elena.ferapontova@inano.au.dk

Supporting information for this article is given via a link at the end of the document.

ARTICLE

Figure 1. Schematic representation of immobilization and redox labeling of the 3G-PAMAM dendrimer on the gold electrode (more details are in the

Experimental section).

Results and Discussion

Dendrimer immobilization and modification. Gold electrodes were modified with di(N-succinimidyl)-3,3´-dithiodipropionate (DTSP) and PAMAM by their stepwise immersion in the DTSP and PAMAM solutions. First, the adsorbed monolayer of N-succinimidyl-3-thiopropionate (NSTP) was formed by dissociative chemisorption of DTSP,[18] which was then PAMAM modified by further reaction of NSTP with peripheral amine groups on the PAMAM dendrimer shell (Figure 1). Formation of a monolayer of NSTP could be followed through a substantial decrease in the gold capacitive currents (Figure 2). The reductive desorption of the NSTP SAM in alkaline solutions produced a well-defined peak at -0.73 V (Figure S1, ESI), consistent with the previous reports.[19]

E vs Ag/AgCl (3M KCl), V

-0.4 -0.2 0.0 0.2

I, µA

-4

-3

-2

-1

0

1

2

Bare Au NSTP/Au PAMAM/NSTP/Au MB/PAMAM/NSTP/Au

Figure 2. Representative cyclic voltammograms (CV) of (dotted line) the bare gold electrode successively modified with (NSTP/Au) NSTP, (PAMAM/NSTP/Au) PAMAM dendrimer and (MB/PAMAM/NSTP/Au) NHS-MB, recorded in N2-saturated 50 mM PBS, pH 7, scan rate 50 mV s -1. Electrochemically active surface area of the gold electrodes was 0.084±0.006 cm2, the PAMAM concentration used for modification was 0.2 mM.

After covalent attachment of the PAMAM dendrimer to the gold surface, the rest of the amine groups on the dendrimer shell were conjugated to succinimide ester-activated MB (NHS-MB) by peptide chemistry (Figure 1). A pronounced couple of peaks characteristic of MB, with a mid-potential at -189±8 mV, was then observed in CVs recorded with MB/PAMAM/NSTP/Au electrodes (Figure 2). The peak potential was 60 mV more positive than the MB redox potential on mercaptohexanol SAMs[20], consistent with the MB coupling to the hydrophobic, positively charged layer. No MB signals were observed after NHS-MB reaction with the NTSP-modified surface, in the absence of dendrimers (Figure S2, ESI). The MB peak shape and position approached the “reversible peaks” case (the peak separation of 17 ± 1 mV at 50 mV s-1), with a high symmetry of the peaks and the full-width-at-half maximum (ΔEfwhm) of 92 ± 16 mV. The latter value is close both to the theoretical 90.6 mV expected for a reversible one electron transfer system (ΔEfwhm=3.53RT/nF) and to the widths of cathodic (62.5/αn mV) and anodic (62.5/(1-α)n mV) peaks for a quasi-reversible system.[21]

S

N

N N S

N

N N

+1 e

+H

Methylene Blue (MB)

Leucomethylene Blue (LB)

S

HN

N N

+1 e

S

HN

N N

Scheme 1. Schematic representation of the redox reactions of MB at pH 7. [20,

22]

It is important to note that the ET reaction of MB is more complex than just one ET characteristic of Fc and Ru complexes studied in previous works.[10-11, 13, 23] In aqueous solutions MB undergoes two consecutive e- transfers, with a chemical protonation step following the first electron transfer (Scheme 1). However, upon MB conjugation and interaction with such biomolecules as DNA, at sufficiently high potential scan rates the redox reaction of MB starts to follow a one electron transfer pathway, though always being accompanied by protonation.[24] In our previous study of MB-modified PAMAM on graphite, the MB redox reaction was mostly consistent with a one ET. [5] In the current work, the number of electrons involved in the reaction, n, was always evaluated as a function of experimental conditions, from the ΔEfwhm of cathodic and anodic peaks in CV,[21] and was found to approach theoretical 2e- at low scan rates and 1e- at scan rates higher than 1 V s-1 (Table 1).Thus, electrochemistry of MB at high scan rates appears to be a single ET reaction, similar to the previously obtained data on DNA-mediated ET between the electrode and DNA-intercalated MB.[24]

Table 1. Number of electrons transferred in ET (n), ET coefficient (α), ET rate constants, MB surface concentration (ΓMB) and the corresponding surface density of G3-PAMAM (ΓG3-PAMAM), evaluated for different immobilization conditions.

ARTICLE

[PAMAM] (mM)

n a evaluated at different v (V s-1) n calculated from trumpet

plotsα

ks , s-1

(ΔE<200 mV)

ks , s-1

(ΔE>200 mV)

ΓMB,c pmol cm-2

ΓG3-PAMAM,d

pmol.cm-2ΓG3-PAMAM,e

pmol.cm-2

0.05-0.1 5-14

0.2 1.9±0.1 1.0±0.1 1.0 ± 0.1 0.46 108±6 267±1 24 ± 1 0.8 1.1

1 1.6±0.1 0.9±0.1 0.8 ± 0.1 0.47 92±13 211±5 106 ± 5 3.4 5.1

5 1.7±0.1 0.9±02b n.a. n.a. 34±7* n.a. 224 ± 8 7.2 10.7

18 1.7±0.1 0.8±0.2b n.a, n.a. 26±9* n.a. 267 ± 7 8.6 12.7

a The number of electrons was calculated from the widths of the cathodic peak (equal to 62.5/αn mV) and of the anodic peak (62.5/(1 -α)n mV),[21] and b according to the equation for the diffusion process Epeak-E1/2=2.218RT/nF;[14] c ΓMB was evaluated at 0.1 V s-1 and referred to the corresponding number of electrons. d ΓG3 was calculated assuming that 31 amines were substituted by MB. e ΓG3 was calculated assuming that 21 surface amines were substituted by MB; *evaluated for conditions fitting the criteria of a surface-controlled ET.

The dendrimer surface coverage as a function of the dendrimer concentration in immobilization solutions. A G3 PAMAM dendrimer is 3.6 nm in diameter (d) and bears 32 peripheral amine groups.[13] The theoretical limiting surface coverage Γmax of a such size dendrimer is either 16.9 pmol cm -2, if calculated in assumption of a spherical size of the dendrimer [25]

Γmax=(1/πR02NA), (1)

with R0 as the PAMAM radius and NA as Avogadro’s number, or 12.8 pmol cm-2 if one assumes that each dendrimer occupies a d×d square. Those surface coverages correspond to ca. 410-541 pmol cm-2 MB molecules if all 32 peripheral amines of the dendrimer are modified with MB. In practice, the exact number of modified dendrimer’s surface groups is unknown, though modification was performed under conditions ensuring their maximal number. Thus, the evaluation of the dendrimer surface coverage from the MB signals can be done only under certain assumptions, such as that maximal number of amine groups that can be modified is 31 (with one functional group used for the PAMAM surface attachment).Immobilization of dendrimers was performed from solutions with different PAMAM concentrations. After the MB modification step the MB surface coverage (ΓMB) was estimated from the charge (Q) consumed in the ET reaction, determined by integration of the MB anodic and cathodic peak currents recorded at low potential scan rates:[21]:

ΓMB=Q/nFA , (2)

where F is the Faraday constant and A is the electroactive surface area of the electrode. For the lowest PAMAM concentration the ΓMB was found to be 24±1 pmol cm-2, which corresponds to a diluted monolayer of 0.8 pmol cm -2 PAMAM dendrimers, if all MB moieties of the dendrimer are assumed to be electroactive (Table 1). With increasing PAMAM concentrations in immobilization solutions, the ΓMB also increased, as could be followed from the intensity of the MB voltammetric signals (Figure 3), and the maximum ΓMB of 267±7 pmol cm-2 was obtained for the highest 18 mM PAMAM concentration used, which correlated with 8.61 pmol cm -2 of dendrimer molecules immobilized on the electrode surface.

E vs Ag/AgCl (3M KCl), V

-0.6 -0.4 -0.2 0.0 0.2

I, µA

-100

0

100

200

E vs Ag/AgCl (3M KCl), V

-0.6 -0.4 -0.2 0.0 0.2

I, µA

-20

0

20

40

E vs Ag/AgCl (3M KCl), V

-0.6 -0.4 -0.2 0.0 0.2

I, µA

-4

-2

0

2

4 A B C

ab

c

d

a

b

cd

a

b

c d

Figure 3. Representative CVs recorded in 50 mM PBS, pH 7, with gold electrodes modified with (a) 0.2, (b) 1, (c) 5, and (d) 18 mM PAMAM at scan rates: (A) 0.1, (B) 1, and (C) 10 V s-1.

The PAMAM concentration dependence of the ΓMB followed the simplest case of the Langmuir adsorption isotherm (Figure 4) with the maximum (saturating) ΓMB of 297 ± 8 pmol cm-2, which roughly correlated with 9.6 pmol cm-2 of PAMAM dendrimers immobilized on the electrode if we consider that all 31 peripheral amines of the dendrimer are modified with MB. These 9.6 pmol cm-2 represent 51%-67% of the theoretical monolayer (12.8 - 16.9 pmol cm-2, see above). In reality, not all amine groups on the dendrimer shell may be modified with MB, thus the true surface coverage can approach the compact monolayer.

[PAMAM], mM

0 5 10 15 20

MB, p

mol

cm

-2

0

50

100

150

200

250

300

k s, s

-1

0

50

100

150

200

250

300

Figure 4. Dependence of the (circles) MB surface coverage (ΓMB) and (squares) ET rate constant (ks) on the concentration of PAMAM in immobilization solutions. The ΓMB was evaluated by integrating the MB peak currents recorded in N2-saturated 50 mM PBS, pH 7, scan rate of 0.1 V s-1. ET rates were calculated from the iR-drop corrected peak potentials using the high potential scan rate segments of E-log v plots for diluted monolayers and the Laviron peak separation procedure for compact monolayers. [21] Solid black line is fitting to the Langmuir isotherm; blue line - to the exponential decay function.

ARTICLEAnother phenomenon that may affect the MB/dendrimer surface concentration is the shape[10-11, 23a] and the size of the adsorbed dendrimers that apparently differ from the sphere with a theoretical diameter.[13, 23b] Both surface studies [13, 23b] and quantitative modelling analysis[10-11] evidence that the adsorbed dendrimer molecules are flattened and their shape is rather hemispherical than spherical known in solution. It was discussed that dendrimer molecules changed their conformation on the surface to compensate the negative energy of adsorption,[23b]

and that resulted in the truncated sphere/hemisphere shapes of the surface-attached dendrimers[11] (Scheme 2A). That may have its implications for the MB-on-surface modification process: the dendrimer immobilization then should involve more than one amine group of the dendrimer shell covalently linked to the NSTP-modified electrode (Figure 1), thus resulting in less number of sites modified with MB. Based on the Figure 4 data, we believe that at the highest PAMAM concentration we have a compact monolayer of tightly packed dendrimers partially modified with MB (16-21 groups of 32, if refer to the theoretical monolayer values)), while the rest of amine groups are either covalently coupled to NSTP or resting at the bottom of the dendrimer hemisphere on the electrode surface (Scheme 2).Along with that, the G3-PAMAM-modified electrodes showed both good chemical and mechanical stability due to the amide bond formation between PAMAM and NSTP (NB: stability of electrodes without covalent coupling of G3-PAMAM to gold was insufficient for kinetic studies). The MB signals recorded with the MB/PAMAM/NSTP/Au electrodes remained quite stable during at least 10 days of storage in PBS, pH 7, at 4°C (Figure S3, ESI), with a 6% decrease in the initial MB peak intensity after 10 days. Such stability of a monolayer is superior to that commonly observed with SAMs of alkanethiols and thiolated DNA tethered to gold electrodes, demonstrating as a rule 30% decrease in the surface coverage in 4 days.[24c, 26] That may be both due to a specific interfacial behavior of bulky and hydrophobic dendrimer molecules not prone to surface detachment and due to multiple peptide bond formation with the modified surface of the electrodes.

Kinetic analysis of ET in MB-modified PAMAM dendrimers. On scrutinizing CVs recorded with the MB-PAMAM-modified electrodes (such as in Figure 3) it became evident that with the increasing PAMAM surface coverage the mechanism of ET was changing, and that could be followed both from the shapes and intensities of the voltammetric signals and peak positions. Basic analysis of the dependencies of the peak current intensities Ip on the potential scan rate v (Figure 5) showed that independently of the dendrimer surface coverage, for all studied systems for the v not exceeding 1 V s-1 both anodic (Ip,a) and cathodic (Ip,c) peak currents increased linearly with the potential scan rate, formally designating a surface-controlled ET reaction[14] (Figure 5, insets).

Figure 5. Representative CVs recorded in N2-saturated 50 mM PBS, pH 7, with electrodes modified with (A) 0.2 mM and (C) 18 mM PAMAM; scan rates: 0.025, 0.05, 0.1, 0.2, 0.30, 0.4, 0.5, 0.75, 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 12, 14, 16 and 18 V s−1. Insets: Dependencies of the CV peak currents Ip on the potential scan rate v for different scan rate ranges. (B, D): Corresponding trumpet plots for the (full circles) anodic and (empty circles) cathodic peak potentials vs. the logarithmic scan rate. All potentials were corrected for the iR-drop.

The ratio between the Ip,c and Ip,a was close to 1 (0.85±0.02) and it was a single couple of peaks in CVs, implying that the ET is almost reversible and all redox moieties on the dendrimer shell are equivalent, namely, the MB groups that are further from the electrode surface have the same potential as those in the vicinity. However, at sufficiently high v (between 1 and 90 V s-1) the surface-controlled ET process (the linear Ip-v dependence) remained the same only for diluted PAMAM monolayers (Figure 5A, inset, similar tendency data were also obtained for 1 mM PAMAM), while for denser packed dendrimers the surface-controlled process switched for the diffusion-controlled ET reaction characterized by the Ip-v0.5 dependence (Figure 5C, inset, similar tendency data were also obtained for 5 mM PAMAM). Earlier, ultrafast cyclic voltammetry analysis performed under conditions of a few nm thickness of the diffusion layer (smaller than the size of studied G4-PAMAM dendrimers), showed that electron hopping between the Ru redox centers is the dominant mechanism of ET in Ru-labelled G4-PAMAM dendrimers.[10-11] Therewith, both the dendrimer deformation on the electrode surface and motions of the redox sites near their average positions were shown to contribute to electron hopping between the Ru redox sites (Scheme 2A), and the apparent diffusion coefficient Dhop in the studied system was determined to be 5×10-6 cm2 s-1.[11]

Those data were obtained at positive charges of the electrode surface, which might imply certain repulsive interactions between the electrode surface and positively charged ([Ru2+/3+

(tpy)2]) terminal groups of the dendrimer. In our current work, ET kinetics is studied at negative charges of the electrode surface that provide conditions for attractive interactions between the electrode surface and positively charged terminal groups of the dendrimer, positively charged MB and unmodified NH2-groups. Those conditions may promote a higher mobility of the redox

ARTICLEsites/ability of dendrimer branches bending toward the electrode surface in the applied electric field, which should promote closer approach of MB groups to each other and to the negatively charged electrode surface (Scheme 2B). A similar effect of the applied electric field and its duration on the surface state and as a result on ET kinetics was earlier reported for redox-labelled surface-tethered DNA duplexes, whose electrochemistry depended both on the electrode surface and redox label charges. [27]

Basic analysis of our results (Figure 5) shows that ET in the MB-dendrimer system may proceed both via electron hopping (at high scan rates) and by electron tunneling, (at low scan rates, providing conditions for MB redox sites approaching each other close enough to collide). As discussed above, the ET reaction in the overall positively charged dendrimer molecules occurs at negative charges of the electrode surface, providing conditions for a higher (than at positive charges of the electrode surface, the Ru-dendrimer system)[11] degree of the dendrimer electrostatic compression in the applied electric field. We suggest that at low scan rates there is enough time to place the MB redox sites in a closer proximity to each other and the electrodes surface indeed, either via dendrimer branches bending to the electrode or due to the overall conformational changes in the dendrimer structure, both making it compact enough for electron tunneling between the colliding centers (Scheme 2B). This situation is different from that observed with the Fc- and Ru-complex labelled dendrimers,[10-11, 13, 23b, 28] which ET reactions proceed at positive charges of the electrode surface that electrostatic unfavorable for a such dendrimer compression. In compact dendrimer monolayers, the motion/bending of the MB-modified dendrimer branches in the electric field is expected to be sterically hindered, and that should impede intimate interactions between MB moieties and with the electrode necessary for the electron tunneling pathway. Then, the system behavior transforms into the Scheme 2A case, earlier studied in detail with Ru-complex labelled dendrimers,[11] in which minor fluctuations of the Ru redox sites around their positions in the dendrimer shell allow electron hopping as the only way of redox centers electronic communication with the electrode. Another possible reason for switching of the ET mechanism in more compact monolayers might be incomplete or a lesser degree chemical modification of the dendrimer with MB groups (a lesser density of MB redox centers in the dendrimer shell). However, very similar results were obtained with dendrimers labelled prior the immobilization step with the thiolated methyl-viologen pendant groups, so this possibility should be excluded.[29]

The rate of the heterogeneous ET (the ET rate constant ks of surface-confined ET) was assessed according to Laviron [21] both from the peak-to-peak separations ΔE (for ΔE<200/n mV) using the tabulated values of the ks–related parameter m, and from the E-log (v) trumpet plots (Figure 5B) for the ΔE exceeding 200/n mV:

ks=αnFνc/RT=(1-α)nFνa/RT (3)

where νc and νc are the scan rates obtained by extrapolating the linear part of the Epeak vs. log (ν) plots to ΔE=0, with the slopes of the linear regions equal to -2.3RT/αnF (the cathodic branch) and 2.3RT/(1-α)nF (the anodic branch) that gave the values of α close to 0.5 (a symmetrical redox process) and n approaching a 1e- limit at high scan rates (Table 1). Lower potential scan rates

provided enough time to complete a 2e-/1H+ transfer reaction (Scheme 1, Table 1). Those results are consistent with previous reports on ET reactions of MB either intercalated into or conjugated to the DNA duplex[5, 24c, 24d, 27a] and in MB-dendrimers adsorbed on graphite. [5]

In most diluted dendrimer monolayers, ET was quite fast, with the ks of 267±1 s-1 (Table 1). In MB-G3-PAMAM dendrimers tethered to graphite, ET was characterized by a quite moderate ks of 7 s-1, which may be ascribed to the properties of graphite electrodes different from gold: hydrophilic graphite can be considered as a “slow” electrode material for the hydrophobic PAMAM dendrimer nanostructures.[5] This ks of 267 s-1 was also essentially higher than 33 s-1 shown for Fc-terminated G3-PAMAM dendrimers immobilized on multi-walled carbon-nanotube-modified electrodes via polyazedine prepolymer[28] (at positive charges of the electrode surface).Therewith, the ks values appear to be highly reliant on the dendrimer surface density and decrease exponentially with the increasing [PAMAM] (Figure 4). This decrease in the ET rates with the increasing ΓG3-PAMAM resembles the reported earlier drop in ET rates in the DNA-electrode systems composed of surface-tethered DNA duplexes and redox indicators bound to them, [24c,

30] ascribed to the interference from the neighboring, closely packed molecules. In the present case it may be primarily connected with the lesser extent of electrostatic compression of dendrimers in the applied electric field in compact dendrimer monolayers discussed earlier.

Scheme 2. Schematic representation of the MB-modified dendrimer attached to the surface, (A) either with no applied field or at positive charges of the electrode surface and (B) in the applied field at negative charges of the electrode surface, the dendrimer shell being positively charged (MB is denoted by blue balls and amine groups are not shown).

Diffusional ET in compact dendrimer monolayers. In all studied dendrimer systems, at low scan rates the duration of the potential scan apparently exceeded the time of “diffusion” θ ≈ RT/Fv [11] that allowed the placement of the redox centers in close proximity to each other and their collisions, not the least due to the deformation of the dendrimer shape in the electric field, making dendrimer more compact (Scheme 2B). When dendrimer molecules were spaced on the electrode surface (the low ΓG3-PAMAM), at any scan rate the dendrimers had enough conformational freedom to respond to the electric field assisting the sequential collisions between redox centers and the electrode. In compact dendrimer monolayers, at higher scan rates the MB motion became restricted by the time the potential was applied for and ET started to formally follow the diffusion-controlled case. In theory, at very high potential scan rates (not used here) ET in diluted monolayers should be restricted by diffusion as well, the same way as reported for Fc-terminated DNA duplexes, negatively charged, at positive charges of the electrode surface:

ARTICLEunder conditions of such an electrostatic compatibly the θ should be low indeed, below 0.06-0.3 ms.[27b, 27c]

Figure 6. Dependence (red squares) of the peak potential separations and (black circles) of the relationship kETdif/D1/2 on the scan rate v evaluated for the 18 mM dendrimer system according to Nicholson [31] using the dimensionless kinetic parameter Ψ tabulated for peak separations fitting the 61-212 mV range. Solid lines are SigmaPlot fittings to (left axis) the exponential decay function (y=y0 + ae-bx) with y0=3.29 s-0.5 and (right axis) to the exponential rise to maximum.

It is understood that the MB redox sites covalently attached to the terminal groups on the molecular surface of the dendrimer cannot diffuse to the electrode surface in its classical sense.[11]

Alternatively, electron hopping between the redox centers that are involved in the fluctuating movements around their average positions on the dendrimer shell may be formally exhibited as a diffusion-limited ET, characterized by the apparent coefficient of diffusion DMB (or Dhop) and the apparent ET rate constant kETdif, which evaluations can provide valuable mechanistic information.To address that, the behavior of the compact-monolayer dendrimer system at high scan rates was approximated by the diffusion-limited ET in a nanosize layer in the vicinity of the electrode surface with a thickness corresponding to the dendrimer size (Scheme 2A). We analyzed the system within the routinely used planar-diffusion analytical scheme applicable for anodic and cathodic peak potential separations fitting the 61 - 212 mV range.[31] Therewith, the data referred to the surface-confined process (scan rates below 1 V s-1 (Figure 5C, inset) have not been taken into account. Such macroscopic approximation, not taking into account specific molecular level distributions of/reactions between the redox sites,[11, 32] is quite rough, but it was already shown to be useful for analysis and description of diffusional movements of few-nm length DNA towards the negatively charged electrode surface.[24d, 27d, 27e]

The scan rate dependence of the kETdif, characteristic of the diffusion-limited ET, related to the square root of the DMB

(kETdif/DMB1/2) was evaluated according to Nicholson[31] from the

tabulated kinetic parameter Ψ referred, via the peak potential separation, to the specific potential scan rate (kETdif/DMB

1/2 ~

const×Ψ×v1/2). As can be seen (Figure 6), the kETdif/DMB1/2 - v

dependence shows that at high potential scan rates, the kETdif/DMB

1/2 levels at its minimal value of 3.29 s-0.5. This value is essentially lower than 12,000 s-0.5 evaluated for Ru(3+/2+)-labelled

G4 PAMAM dendrimers in acetonitrile. [11] That is somehow an expected result since the rate of the outer-sphere reactions such as that of the [Ru(tpy)2]3+/2+ couple should be much faster than the inner-sphere electron transfer as in our case. Therewith, the apparent DMB evaluated with the Randles-Sevčik equation (with a surface concentration of MB moieties CMB approximated by m/(A×l), where m is the amount of electrochemically active MB, in moles, A is the electrode surface area, and l is the thickness of the dendrimer layer in which the MB molecules are localized, l = 2R0, where R0 is the radius of the dendrimer sphere, Figure 7),[14] was in the order of 10-13 cm2 s-1, which is a very low value, unless it is referred to the nanoscale diffusion distances, as in the studied system.

The results obtained by approximating the nanoscale system behavior by the macroscopic diffusion approach were validated by engagement of a model taking into account individual ET reactions between neighboring redox centers.[11, 32] The variation of the ΔE with the logarithmic potential scan rate (the slope of the dependence of 124±2 mV, within the 3 – 30 V s -1 range, Figure 7) corresponded to the semi-infinite diffusion accompanied by slow kinetics of ET, for which, according to Amatore et al.,[11] the relationship between the voltammetric peak currents Ip and the Dhop for electron hopping between the redox moieties distributed over the nanometric dendrimer shell (case I in [11], Scheme 2A) can be described as:

Ip/(nFm)=0.496(α*nFvDhop/R02 RT)1/2tan(φ0/2), (4)

where φ0 is the half angle of the cone the part of the dendrimer sphere contacts the surface (Figure 7) and α* is α for the cathodic and (1-α) for anodic processes, correspondingly. The Dhop values were evaluated for the φ0 changing from 11o to 90o

(the hemi-sphere case). As can be seen in Figure 7, consistent with the model (Figure 7 inset), they decrease with the increasing φ0 (equivalent to the decreasing diffusion distance) and the scan rate (a power decay, Dhop~v-0.733), tending to level at some minimal values. Interestingly, though the apparent Dhop

values (Figure 7, inset) are slightly lower, they appear to be reasonably close to the DMB values evaluated using the macroscopic model approximation that does not take into account specific molecular features of nanoscale dendrimeric structures at the electrode surface and pathways of electron hopping between the redox moieties. That finding does not eliminate the necessity of detailed kinetic analysis of such nanoscale systems using the models accounting for their molecular level complexity, but allows using macroscopic models as a first, rough but still valid approximation for basic description of ET kinetics in such systems.

ARTICLE

Figure 7. Scan rate dependence of the anodic and cathodic peak potential separation ΔE. Inset: the apparent Dhop values calculated using Eq. (4) for the φ0:11, 23, 33, 46, 53, 62, 70, 77, 84, and 90o. In the schematically represented dendrimer attached to the electrode surface, its radius R0 and the half angle φ0

of the cone the truncated part of the dendrimer sphere contacts the surface are denoted.

To summarize, kinetic analysis suggests that ET in differently packed MB-modified dendrimer monolayers can proceed both due to collisions between MB moieties in the outer shell of the positively charged dendrimers electrostatically compressed on the negatively charged electrode surface and as electron hopping, when intimate contacts of MB redox centers are sterically impeded. Those two cases are experimentally exhibited via the surface-confined and diffusion-limited ETs evaluated from the CV data. In the latter case, switching from the surface to diffusion-controlled ET is triggered by the steric hindrance in tightly packed dendrimer monolayers, restricting the motion of the dendrimer brunches and close contacts/collisions between MB groups that are essential for electron tunneling.

ET in dendrimers in the context of the Butler-Volmer and Marcus models. It was of a particular interest to understand if the surface-confined ET in the dendrimer systems may fit the existing models for the electrode ET kinetics and be described as a function of activation and reorganization energies of ET, more specifically, within the Butler-Volmer and the modified Marcus theories. Therewith, it was understood that the overall complex MB reaction involves transfers of two electrons, with an intermediate protonation step (Scheme 1), and further only the data fitting the criteria for a one electron transfer reaction and the surface-confined ET mechanism (diluted dendrimer monolayers) were analyzed.

In general, kinetics of an electrode reaction, specifically, the heterogeneous rate constant ks, exponentially depends on the reaction driving force, i.e. reaction overpotential η (equal to the difference in the applied and formal potential of the reaction (E-Eo´), and that is reflected by the Butler-Volmer model:[14]

k red=k0 exp (−αnF ( E−E0 )RT

)

(5)

k ox=k0 exp ((1−α)nF ( E−E0 )

RT)

(6)

with the k0 as the standard ET rate constant.The Marcus theory takes into account the ks dependence on the nuclear reorganization energy λ and the extent of the electronic coupling between the donor and acceptor in the transition state of the reaction,[15] with an electrode replacing one of the reaction partners when the ET reaction is electrochemically induced:[33]

k ox/k red=kmax √ RT4 π ∫

−∞

∞

exp ¿¿¿, (7)

where kmax is the maximum rate of ET at F(E - E0) significantly exceeding λ. Here, we examined the applicability of Eqs. (5)-(7) for description of kinetics of ET between MB centers attached to the dendrimer periphery and the electrode.The ks were evaluated at different overpotentials from the CV data collected at different scan rates, by the procedure proposed in [34]. According to Murray et al. [34] for a simple reversible ET reaction, the η-dependence of the rate constants (kRed,η and kOx,η) for the oxidation and reduction half-reactions of the surface-attached redox species can be expressed as:

kOx , η≅iOx ,η

nFA ΓOx ,η=

iOx ,η

nFAQOx, η

nFA

=iOx ,η

QOx, η;k Red, η≅

iRed,η

nFA Γ Red, η=

iRed , η

nFAQRed ,η

nFA

=¿iRed ,η

QRed , η

(8)

Where iη , Γ,η, Q,η are the peak intensity, surface coverage and charge of the oxidized and reduced redox species, respectively. The dependence of the obtained ks on the η was fitted to Eqs.(5) -(7) (Figure 8). As can be seen, the studied ET cannot be approximated by the Butler-Volmer dependence (Figure 8B), while the Marcus theory reasonably well reflects the ks changes with η and the ks tendency to level at a limiting value with the increasing η (Figure 8A). A reasonable fitting of the experimental data was obtained for all η, except of │η│<20-30 mV where the error of the ks determination introduced by the background correction could be high; also, quite low values of the reorganization energy λ approaching 0.1 eV were obtained. Overall, those data support our hypothesis that ET in diluted dendrimer monolayers proceeds via electron tunneling between colliding MB redox centers, which are otherwise spatially separate in dendrimer molecules. That means that the conformation of the spherical dendrimer molecule should be essentially distorted indeed, being electrostatically compressed at negative charges of the electrode surface in order to allow closer approach and collisions between the MB redox sites and the electrode (Scheme 2B).

ARTICLE

Figure 8. Dependence of the ks on the overpotential η for different dendrimer surface coverages: gold electrodes modified using (1) 0.2 and (2) 1 mM solutions of G3-PAMAM. Dots are experimental data, and lines correspond to (A) fitting to Eq.(6), (1) kmax = 1199 s-1;(dashed line) λ =0.08 eV and (solid line) λ =0.1 eV; (2) kmax = 610 s-1; (dashed line) λ =0.06 eV and (solid line) λcathodic=0.08 eV and λanodic =0.06 eV; and (B) fitting to Eqs. (4-5).

Enzyme wiring to electrodes by MB-modified dendrimers. An efficient electrical communication between MB redox centers in G3-PAMAM dendrimers was used to design reagentless enzyme electrodes exploiting redox dendrimers as electrical wires mediating the bioelectrocatytic activity of the enzymes. [3, 35] Two enzymes were studied here, heme-containing HRP, which bioelectrocatalysis of H2O2 reduction has been intensively investigated,[16] and FAD-dependent HOX capable of bioelectrocatalytic oxidation of sugars.[17] The FAD active site in HOX is covalently bound, and its bioelectrocatalysis is not compromised by the FAD loss from the protein, in contrast to the widely used glucose oxidase.[5]

Native HRP did not show any redox activity on the MB/PAMAM/NSTP/Au modified electrodes. In HRP, its heme redox center is deeply buried inside the protein, and the existing efficient ET pathways through the protein media [36] are not easily accessible in the glycosylated native protein,[37] which is consistent with our result. In contrast, de-glycosylated recombinant rHRP interacted productively with the MB-PAMAM, which could be followed from the potentials of the MB wire (Figure 9A), which indicates the MB-PAMAM ability of shuttling electrons between the redox center of rHRP and the electrode.The bioelectrocatalytic current of H2O2 reduction increased with the increasing concentration of the substrate, and the dependence of the current on the [H2O2] followed a typical Michaelis-Menten behavior (Figure 9A).

E vs Ag/AgCl (3M KCl), V

-0.4 -0.2 0.0 0.2

I, µA

cm

-2

-20

-15

-10

-5

0

5

10

E vs Ag/AgCl (3M KCl), V

-0.4 -0.2 0.0 0.2 0.4

I, µA

cm

-2

-20

-15

-10

-5

0

5

1040 mM glucose

2 mM glucose

A B

5 µM H2O2

90 µM H2O2

[H2O2], µM

0 20 40 60 80 100

I-I0,

µA.c

m-2

0.0

0.5

1.0

[Glucose], mM

0 10 20 30 40 50

I-I0,

µA.c

m-2

0.0

0.1

0.2

0.3

Figure 9. Representative CVs of (A) rHRP/MB-PAMAM/NSTP/Au recorded in N2-saturated PBS, pH 7, in the presence of increasing concentrations of H 2O2

(0, 5, 10, 15, 20, 30, 50, 70, and 90 µM) and (B) HOx/MB-PAMAM/NSTP/Au in O2-saturated PBS, pH 7, in the presence of increasing concentrations of glucose (0, 2, 4, 6, 8, 10, 20, 40 mM). CVs were recorded at a scan rate of 20 mV s-1; ΓMB = 24 ± 1 pmol cm-2. Inset: plot of I-I0 vs. [analyte], data collected at -0.3 V for H2O2 and at +0.5 V for glucose. (Solid lines) Fitting to the Michaelis-Menten dependence I0=Imax×[substrate]/(KM+[substrate]) was performed with Sigma Plot software.

Oriented immobilization of rHRP via genetically introduced surface tags (His or Cys) resulted in ET rates between the HRP heme and the electrodes exceeding 100 s-1, at pH 7, (those ET rates are comparable with ET rates in diluted MB-PAMAM monolayers, Table 1) and, as a result, in particularly efficient direct ET-based bioelectrocatalytic reduction of H2O2.[36] The main disadvantage of such approach was very low, sub-picomolar-level amounts of electronically wired peroxidase molecules, representing less than 1% of the theoretical monolayer.[36, 38] With our current system, a higher efficiency of the dendirmer-wired HRP bioelectrocatalysis was achieved [36], both due to more efficient electrical wiring and higher amount of rHRP molecules wired. Thus, the protein immobilization on dendrimer wires results in a higher efficiency bioelectrocatalysis indeed. Electrically wired HOX, with its FAD potential at -307 mV, pH 7,[17] demonstrated bioelectrocatalytic oxidation of glucose in the air-saturated PBS, however, with an essential overpotential both relative the potentials of HOX’s FAD and a MB wire (Figure 9B). Therewith, a typical change in the shape of the voltammogram, characteristic of the bioelectroctalytic oxidation process,[39] with the increasing bioelectrocatalytic currents, at 100-400 mV, and decreasing of the mediating species reduction peak, at -240 mV, could be followed (Figure 9B). This suggests a typical enzyme-catalyzed reaction that involves reduction of the FAD by glucose followed by re-oxidation of FAD by the MB mediator regenerating the catalytically active oxidized form of the enzyme. [40]. As a matter of fact, HOX was wired by the MB-modified dendrimer, and its bioelectrocatalytic activity followed the Michaelis-Menten dependence (Figure 9B, inset). However, glucose oxidation started at potentials 400 mV higher than the potential of HOX’s FAD. Similar phenomenon has been observed with HOX wired by MB-PAMAM to graphite, with a 200 mV overpotential (the ks

for MB-PAMAM on graphite was 7 s-1),[5] and with HOX directly wired to graphite, also with a 200 mV overpotential (the ks for FAD in HOX on graphite was 29 s-1).[17] Even higher overvoltage was reported for glucose oxidation by glucose oxidase wired by gold nanoparticle relay[41] providing high ET rates between the FAD redox center and the electrode. Thus, the tendency of increasing the reaction overpotential with increasing the ET rates between the redox center and the electrode can be followed (no

ARTICLEbioelectrocatalysis overpotential was observed for soluble MB operating as a redox mediator for HOX catalysis).[17] We believe that in the case of a fast ET reaction between the HOX’s FAD and the electrode, provided either by the direct ET reaction or ET wired by the MB-modified dendrimer (i.e. when a small molecular substrate was replaced either by an electrode or a nanoscale wire), some additional reaction step in the bioelectrocatalytic transformation of glucose starts to limit the overall reaction rate (for example, the conformational changes in the FAD center that start to gate the ET and bioelectrocatalysis) and that results in the increased overvoltage of bioelectrocatalysis. As in the case of HRP, those aspects are of fundamental importance for understanding the electrode functioning of such biotechnologically important enzymes as glucose oxidase and HOX and would be studied in more detail in future.

Conclusions

Here, we show that electrical wires based on MB-labeled G3 PAMAM dendrimer structures allow shuttling electrons over the MB-modified dendrimer crown with ET rates approaching 267 s-

1. ET in diluted dendrimer monolayers follows the features of surface-confined ET reaction that occurs by electron tunneling in positively charged dendrimeric structures becoming more compact at negative charges of the electrode surface, thus enabling a closer approach and collisions between the MB redox centers located in the outer shell of the dendrimer structure. It is shown that the ET is strongly influenced by the molecular arrangement of the dendrimer at the electrode surface, being more efficient in the case of less crowded monolayers. In compact dendrimer monolayers, the ET mechanism formally switches to the diffusion-controlled ET reaction, as a result of strong interactions between the neighboring dendrimers, restricting the dendrimer branches motional freedom and affecting the conformational state of the dendrimers on the electrode surface. In agreement with earlier discussions [11] this apparent diffusion-controlled ET proceeds via electron hopping between PAMAM terminal redox groups brought in a sufficient for electron hopping proximity by the flexible movements of the dendrimer branches. Demonstrated fast ET rates between the electrodes and remote MB groups the peripheral groups at the top of the dendrimers, separated from the electrode surface by the 2-3 nm distance, outline the applicability of the system for challenging bioelectronic and biosensor applications, exemplified here by wiring two biotechnologically important enzymes, H2O2-reducing HRP and glucose-oxidizing HOX, both demonstrating dendrimer mediated bioelectroacatlytic activities. The efficiency of bioelectroacatlysis was though less than expected, which might be connected with a complicated electronic marriage between dendrimers and proteins used in the current work, and will be further studied in more detail.

Experimental Section

Materials. G3-poly(amidoamine) dendrimer (PAMAM), di(N-succinimidyl)-3,3´-dithiodipropionate (DTSP), dimethyl sulfoxide (DMSO), components of buffer solutions and methanol of analytical grade or of ultra-high purity were purchased from Sigma-Aldrich (Germany) and used as received. N-hydroxysuccinimide ester (NHS)-activated

monocarboxymethylene blue (NHS-MB) was acquired from empBiotech (Berlin, Germany). Horseradish peroxidase (HRP, 330 U mg-1) was from Sigma Aldrich; the recombinant form of HRP of wild type expressed in E. coli[36] (rHRP, 600 U mg-1) was kindly provided by Prof. Irina Gazaryan, Burke Medical Research Institute, New York City, USA. Hexose oxidase (HOX) expressed in Hansenula polymorpha was kindly provided by Prof. Charlotte Poulsen, Dupont Biosciences, Denmark. All over the work, deionised Milli-Q reference A+ water purification system (18 MΩ, Millipore, Bedford, MA, USA) was used for preparation of solutions.

Preparation of the dendrimer-modified electrode.

Gold cleaning. Gold disk electrodes (0.2 cm Ø) were electrochemically cleaned by cycling in 0.5 M NaOH at 0.05 Vs-1. Then they were hand-polished to a mirror luster with 1µm diamond and 0.1 µm alumina slurries (Struers, Denmark) on microcloth pads (Buehler, Germany) and sonicated for 30 min in EtOH/water (1:1) ultrasonic bath. Finally, they were electrochemically polished by cycling in 1 M H2SO4 and 0.5 M H2SO4/10 mM KCl at 0.3 Vs-1. The electrode surface area was determined by integrating the reduction peak of gold surface oxide during the final scan in 0.1 M H2SO4 and assuming a theoretical value of 400 µC cm-2 for a monolayer of chemisorbed oxygen on gold electrode[42]. The electrochemical surface area of the bare gold electrode was typically of 0.087 ± 0.004 cm2. Cleaned electrodes were washed in water and kept in EtOH for 30 min prior modification. N-succinimidyl-3-thiopropionate self-assembled monolayer (NSTP/Au). The conditioned electrode was immersed in 250 µL solution of 4 mM DTSP in dimethyl sulfoxide (DMSO) for 1 h at room temperature[19]. Afterwards, the electrode was thoroughly rinsed with DMSO (10 min). Electrodes were employed immediately after preparation.

G3-poly(amidoamine) dendrimer covalent bonding (PAMAM/NSTP/Au). NSTP/Au electrode was immersed for 2 h at room temperature in 250 µL solution of PAMAM in methanol[43] (to prevent hydrolysis of the activated ester group). Afterwards, the modified electrode was rinsed with 50 mM phosphate buffer pH 7 in order to remove the weakly attached dendrimer and kept in 0.1 M Tris-HCl pH 7.4 to hydrolyze the unreacted succinimide ester on the surface.

Monocarboxymethylene blue NHS ester coupling (MB/PAMAM/NSTP/Au). Amine terminated PAMAM/NSTP/Au electrode was incubated with 10 µL drop containing 0.38 µmol of NHS-MB [44] in DMSO overnight at 4 °C under the lid. The electrodes were finally washed with a buffer solution and kept in it at 4 °C when not in use.

Instrumentation. Cyclic voltammetry (CV) was performed in a three-electrode cell, with a gold disk working electrode (CH Instruments, Austin, TX), platinum wire as a counter electrode, and Ag/AgCl (3 M KCl) as a reference electrode (Metrohm, Denmark). All measurements were carried out with a potentiostat AUTOLAB PGSTAT 30 (Eco Chemie B.V., Utrecht, the Netherlands) equipped with NOVA 1.10 software. Working solutions were de-aerated with N2 for at least 30 min prior data recording and was kept under N2 during the entire experiment. All experiments were carried out in 50 mM phosphate buffer solution (PBS), pH 7, at 22±1 °C. In kinetic analysis of ET reactions, the potentials were corrected for the iR- drop as follows: Ecorrected=Ecathodic/anodic±iR, where i is the peak current in amperes and R is the solution resistance in ohms determined by the electrochemical impedance spectroscopy (EIS) at -0.1 - 0 V with the modified electrodes in the corresponding working solutions.

Calculations and simulations. Simulation of dependences of rate constants on overpotential and fitting to data, calculation of rate constants and reorganization energies were accomplished using script written in R (a programming language for statistical computing) and run on an open source integrated development environment – R studio. [45] A typical calculation/simulation execution was based on the integrated version of the Marcus equation[15] for the oxidation and reduction processes,[33] (Eq. 6) and considered either for different reorganization energies, λ, and ET rate constants for oxidation and reduction of MB, kox

ARTICLEand kred, or their similar values (symmetrical processes). For identification of the reorganization energy value, simulated data were fitted to the experimental ones by the least squares method, assuming that the maximal experimental rate constant equals to kmax. Data fitting to Eqs. (5) - (6)[14] was performed using the standard exponential growth fitting with two parameters in SigmaPlot 12.5 (Systat Software, San Jose, CA) software package.

Acknowledgements

The work was supported by the Danish National Research Foundation through their support to the Center for DNA Nanotechnology (CDNA-2). IAM acknowledges the EU’s support under H2020-MSCA-IF-2014 grant agreement 660339 (eADAM).

Keywords: Methylene blue • G3 PAMAM dendrimer • Electron transfer • Enzyme electrode • Hexose oxidase • Peroxidase

[1] D. Astruc, Nature Chem. 2012, 4, 255-267.[2] M. A. Mintzer, M. W. Grinstaff, Chem. Soc. Rev. 2011, 40, 173-

190.[3] E. E. Ferapontova, in Encyclopedia of Sensors, Vol. 10 (Eds.: C.

A. Grimes, E. C. Dickey, M. V. Pishko), ASP, Stevenson Ranch, California, 2006, pp. 391-422.

[4] M. Frasconi, D. Deriu, A. D'Annibale, F. Mazzei, Nanobiotechnology 2009, 20, 505501 (505508p).

[5] V. Castaing, I. Álvarez-Martos, E. E. Ferapontova, Electrochim. Acta 2016, 10, 263-272.

[6] S. Cosnier, Biosens. Bioelectron. 1999, 14, 443-456.[7] a) S. C. Barton, H.-H. Kim, G. Binyamin, Y. Zhang, A. Heller, J.

Am. Chem. Soc. 2001, 123, 5802-5803; b) N. Mano, V. Soukharev, A. Heller, J. Phys. Chem. B 2006, 110, 11180-11187.

[8] a) E. Katz, D. D. Schlereth, H.-L. Schmidt, A. J. Olsthoorn, J. Electroanal. Chem. 1994, 368, 165-171; b) I. Willner, V. Heleg-Shabtai, R. Blonder, E. Katz, G. Tao, A. F. Bückmann, A. Heller, J. Am. Chem. Soc. 1996, 118, 10321-10322.

[9] a) T. Nöll, G. Nöll, Chemical Society Reviews 2011, 40, 3564-3576; b) B. Willner, E. Katz, I. Willner, Curr.Opin. Biotechnol. 2006, 17, 589-596.

[10] C. Amatore, Y. Bouret, E. Maisonhaute, J. I. Goldsmith, H. D. Abruña, ChemPhysChem 2001, 2, 130-134.

[11] C. Amatore, Y. Bouret, E. Maisonhaute, J. I. Goldsmith, H. D. Abruña, Chem. Eur. J. 2001, 7, 2206-2226.

[12] D. J. Diaz, G. D. Storrier, S. Bernhard, K. Takada, H. D. Abruña, Langmuir 1999, 15, 7351-7354.

[13] K. Takada, D. J. Diaz, H. D. Abruna, I. Cuadrado, C. Casado, B. Alonso, M. Moran, J. Losada, J. Am. Chem. Soc. 1997, 119, 10763-10773.

[14] A. J. Bard, L. R. Faulkner, Electrochemical Methods - Fundamentals and Applications, Wiley, New York, 1980.

[15] R. A. Marcus, N. Sutin, Biochim. Biophys. Acta 1985, 811, 265-322.

[16] E. E. Ferapontova, Electroanalysis 2004, 16, 1101-1112.[17] D. Fapyane, C. H. Poulsen, E. E. Ferapontova, Electrochem.

Commun. 2016, 65, 1-4.[18] H. A. Biebuyck, C. D. Bain, G. M. Whitesides, Langmuir 1994, 10,

1825-1831.[19] M. Darder, K. Takada, F. Pariente, E. Lorenzo, H. D. Abruña, Anal.

Chem. 1999, 71, 5530-5537.[20] E. E. Ferapontova, K. V. Gothelf, Electroanalysis 2009, 21, 1261-

1266.[21] E. Laviron, J. Electroanal. Chem. 1979, 101, 19-28.[22] R. Zhan, S. Song, Y. Liu, S. Dong, J. Chem. Soc., Faraday Trans.

1990, 86, 3125-3127.[23] a) C. Amatore, F. Grün, E. Maisonhaute, Angew. Chem. Int. Ed.

2003, 42, 4944-4947; b) K. Takada, G. D. Storrier, M. Moran, H. D. Abruna, Langmuir 1999, 15, 7333-7339.

[24] a) I. Alvarez-Martos, R. Campos, E. E. Ferapontova, Analyst 2015, 140, 4089-4096; b) I. Álvarez-Martos, E. E. Ferapontova, Anal. Chem. 2016, 88, 3608-3616; c) R. Campos, A. Kotlyar, E. E. Ferapontova, Langmuir 2014, 30, 11853–11857; d) E. Farjami, R. Campos, E. E. Ferapontova, Langmuir 2012, 28, 16218-16226.

[25] J. Lahiri, L. Isaacs, B. Grzybowski, J. D. Carbeck, G. M. Whitesides, Langmuir 1999, 15, 7186-7198.

[26] N. Phares, R. J. White, K. W. Plaxco, Anal. Chem. 2009, 81, 1095-1100.

[27] a) A. Abi, E. E. Ferapontova, J. Am. Chem. Soc. 2012, 134, 14499-14507; b) A. Anne, A. Bouchardon, J. Moiroux, J. Am. Chem. Soc. 2003, 125, 1112-1113; c) A. Anne, C. Demaille, J. Am. Chem. Soc. 2006, 128, 542-557; d) E. Farjami, R. Campos, E. E. Ferapontova, Langmuir 2016, 32, 928; e) E. Farjami, L. Clima, K. Gothelf, E. E. Ferapontova, Anal. Chem. 2011, 83, 1594-1602; f) L. Kékedy-Nagy, S. Shipovskov, E. E. Ferapontova, Anal. Chem. 2016, 88, 7984–7990.

[28] D. Deriu, G. Favero, A. D'Annibale, F. Mazzei, ECS Transactions 2008, 16, 105-113.

[29] I. Alvarez-Martos, N. Plumere, E. E. Ferapontova, unpublished.[30] R. Campos, E. E. Ferapontova, Electrochim. Acta 2014, 126, 151-

157.[31] R. S. Nicholson, Anal. Chem. 1965, 37, 1351-1355.[32] E. Laviron, J. Electroanal. Chem. 1980, 112, 1-9.[33] L. J. C. Jeuken, Biochim. Biophys. Acta 2003, 1604, 67-76.[34] L. Tender, M. T. Carter, R. W. Murray, Anal. Chem. 1994, 66,

3173-3181.[35] E. E. Ferapontova, S. Shleev, T. Ruzgas, L. Stoica, A.

Christenson, J. Tkac, A. I. Yaropolov, L. Gorton, In Electrochemistry of Nucleic Acids and Proteins: Towards Electrochemical Sensors for Genomics and Proteomics, E. Paleček, F. Scheller, J. Wang, Ed. Elsevier B.V. 2006, 1, 517-598.

[36] A. V. Kartashov, G. Serafini, M. Dong, S. Shipovskov, I. Gazaryan, F. Besenbacher, E. E. Ferapontova, Phys. Chem. Chem. Phys. 2010, 12, 10098-10107.

[37] a) E. Ferapontova, K. Schmengler, T. Börchers, T. Ruzgas, L. Gorton, Biosens. Bioelectron. 2002, 17, 953-963; b) E. E. Ferapontova, V. G. Grigorenko, A. M. Egorov, T. Börchers, T. Ruzgas, L. Gorton, Biosens. Bioelectron. 2001, 16, 147-157.

[38] a) R. Andreu, E. E. Ferapontova, L. Gorton, J. J. Calvente, J. Phys. Chem. B. 2007, 111, 469-477; b) E. E. Ferapontova, J. Castillo, D. Hushpulian, V. Tishkov, T. Chubar, I. Gazaryan, L. Gorton, Electrochem. Commun. 2005, 7, 1291-1297.

[39] S. Shipovskov, E. Ferapontova, Biocat. Biotrans. 2008, 26, 455-465.

[40] J. Wang, Chem. Rev. 2008, 108, 814-825.[41] Y. Xiao, F. Patolsky, E. Katz, J. F. Hainfeld, I. Willner, Science

2003, 299, 1877-1881.[42] J. C. Hoogvliet, M. Dijksma, B. Kamp, W. P. van Bennekom, Anal.

Chem. 2000, 72, 2016-2021.[43] H. Tokuhisa, M. Zhao, L. A. Baker, V. T. Phan, D. L. Dermody, M.

E. Garcia, R. F. Peez, R. M. Crooks, T. M. Mayer, J. Am. Chem. Soc. 1998, 120, 4492-4501.

[44] M. N. Hansen, E. Farjami, M. Kristiansen, L. Clima, S. U. Pedersen, K. Daasbjerg, E. E. Ferapontova, K. V. Gothelf, J. Org. Chem. 2010, 75, 2474-2481.

[45] R. C. Team, Vienna, Austria, 2015, pp. http://www.R-project.org/.

ARTICLE

Entry for the Table of Contents

ARTICLEElectron transfer (ET) in methylene blue (MB) labelled G3 PAMAM dendrimers dramatically depends on the dendrimer surface coverage and switches from electron tunneling, in loosely packed dendrimers electrostatically compressed at negative electrode charges, to electron hopping at their higher surface coverage. Demonstrated wiring of redox enzymes by MB-labelled dendrimers allows their bioelectrocatalysis and outlines challenging perspectives for their bioelectronic applications.

Dr. Isabel Álvarez-Martos, Mr. Andrey Kartashov, Dr. Elena Ferapontova*

Page No. – Page No.

Electron Transfer in Methylene Blue-labelled G3 Dendrimers Tethered to Gold