desorption of biomacromolecules involved

Bioelectrochemistry 120 (2018) 87–93

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Bioelectrochemistry journal homepage: www.elsevier.com/locate/bioelechem

Adsorption/desorption of biomacromolecules involved in catalytic hydrogen evolution Slađana Strmečki a,⁎, Emil Paleček b a b

Ruđer Bošković Institute, Division for Marine and Environmental Research, Bijenička 54, 10 000 Zagreb, Croatia Institute of Biophysics, Academy of the Sciences of the Czech Republic, v.v.i., Kralovopolska 135, 612 65 Brno, Czech Republic

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Article history: Received 25 May 2017 Received in revised form 23 November 2017 Accepted 25 November 2017 Available online xxxx In memory of Dr. Michael Heyrovsky. Keywords: Adsorption/desorption Alternating current voltammetry Catalytically active biomacromolecules Mercury electrode Tensammetric minimum

a b s t r a c t Previously, it has been shown that proteins and some polysaccharides (PSs) catalyse hydrogen evolution, producing electrochemical signals on mercury electrodes. The catalytic hydrogen evolution reaction (CHER) of the above-mentioned biomacromolecules was studied by voltammetric and chronopotentiometric stripping (CPS) methods. To obtain more information about electrode processes involving CHER, here we used protein such as BSA, and chitosan as a PS; in addition, we investigated dextran as a control PS not involved in CHER. We studied biomacromolecules by phase-sensitive alternating current (AC) voltammetry. Using phase-in AC voltammetry, for CHER-involved biomacromolecules we observed a CHER peak at highly negative potentials, similar to that observed with other voltammetric and CPS methods. On the other hand, by means of the adsorption/desorption processes studied in phase-out AC voltammetry, we uncovered a sharp and narrow decrease of capacitive current in the potential range of the CHER peak, denominated as the tensammetric minimum. This minimum was closely related to the CHER peak, as demonstrated by similar dependences on specific conditions affecting the CHER peak such as buffer capacity and pH. A tensammetric minimum was not observed for dextran. Our results suggest specific organization of biopolymer layers at negative potentials observed only in biomacromolecules involved in CHER. © 2017 Elsevier B.V. All rights reserved.

1. Introduction Since the beginning of the 1970s, electrochemistry of proteins has dealt predominantly with conjugated proteins containing redox-active non-protein components (e.g., in metalloproteins), producing very interesting results. This research was, however, limited to a relatively small number of proteins as compared to the thousands of proteins in nature [1]. Recently it has been shown that practically any protein can be analysed using the constant current chronopotentiometric stripping (CPS) peak H [2]. This peak is due to the catalytic hydrogen evolution reaction (CHER), and allows structure-sensitive analysis of proteins [3,4], including those important in biomedicine and cancer research [3,5]. Some amino acid residues such as arginine (Arg), lysine (Lys), cysteine (Cys) and histidine (His) have been found to be responsible for CHER observed for proteins [6]. Several years ago it was shown that some polysaccharides (PSs, considered earlier as electrochemically inactive compounds), including chitosan, are involved in CHER, and produce electrochemical signals at highly negative potentials [7,8], similar to protein peak H. Chitosan can be prepared from chitin (composed of Nacetylated glucosamine residues) by chemical deacetylation [9]. ⁎ Corresponding author. E-mail address: [email protected] (S. Strmečki).

https://doi.org/10.1016/j.bioelechem.2017.11.013 1567-5394/© 2017 Elsevier B.V. All rights reserved.

Chitosan is a biodegradable polymer, containing N 50% glucosamine residues (usually 70–90%), attracting great attention in various fields, including industry, pharmacy and medicine [10]. Proteins and PSs involved in CHER have been studied predominantly by CPS and square wave (SW) voltammetry methods. Here, we applied phase-sensitive alternating current (AC) voltammetry, in an attempt to obtain additional information about the adsorption/desorption behaviour of the biomacromolecules studied. AC voltammetry is a frequency-dependent electrochemical method characterized by a small amplitude potential superimposed on a linear time ramp as an excitation signal [11]. It allows independent separation of faradaic (If) and capacitive (Ic) current component as a function of electrode potential. If is due to the electron transfer between a depolarizer and an electrode, while Ic is due to the charged electrode double layer. In a technical sense, If is phase-in (phase angle ϕ = 0°), while Ic is phase-out (ϕ = 90°) with the applied sinusoidal potential [12]. Measurements in the latter mode are usually referred to as tensammetric, and include change in Ic in a double layer of electrode-electrolyte interface due to the adsorption/desorption processes of electrically neutral or charged molecules [11]. Adsorption/desorption characteristics of many biologically relevant molecules such as nucleic acids and their components on mercury and other metal electrodes have been studied by tensammetry [13–15]. Furthermore, tensammetry coupled with a

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S. Strmečki, E. Paleček / Bioelectrochemistry 120 (2018) 87–93

mercury electrode has been utilized by marine scientists in the study of surface-active substances (SAS) present in natural water systems, in order to elucidate their biogeochemical fate and role. Hence, SAS have been characterized in seawater [16,17], the sea surface microlayer [18], lake water [19] and river water [20,21], as well as in rain [22,23] and aerosols [24]. Many organic molecules relevant to natural water systems, such as fulvic and humic acids, PSs and proteins have also been studied tensammetrically in the frame of similar experiments [25,26]. In general, tensammetric voltammograms show characteristic lowering of Ic (i.e. capacitance) below the background electrolyte in the less negative potential range between about 0.0 and −1.0 V, and a specific increase in Ic at potentials more negative than around −1.0 V. The former effect indicates adsorption of organic molecules at the electrode-electrolyte interface, and the latter may suggest desorption or reorientation of the molecules at the electrode surface [27]. The behaviour described derives from the electrical charge of the groups in molecules and from variation in the interfacial tension of the mercury electrode in certain electrolytes with a potential difference imposed across the interface (electrocapillary curve) [28]. AC voltammetry is generally considered as the most common method used in the electrochemical study of surface-related processes. Recently, it was theoretically introduced and experimentally confirmed that adsorption and desorption of model electroinactive surfactants (such as Triton-X-100 and sodium dodecyl sulfate) on a mercury electrode can be explored in SW voltammetry too [29,30], although SW voltammetry is commonly considered as highly sensitive toward faradaic processes because of its insensitivity to Ic [12]. In this paper, we used bovine serum albumin (BSA) together with some other proteins, and chitosan (containing free NH2 groups) as a PS showing intensive CHER signals [8] to study the relationship between CHER signals and the adsorption/desorption behaviour of these biomacromolecules. Dextran was used as a control PS, as it adsorbs strongly on a mercury electrode [27] but does not contain catalytic groups and does not produce any CHER signal [31]. Using phase-in AC voltammetry to observe CHER signals, and phase-out AC voltammetry to study adsorption/desorption behaviour of these biomacromolecules, we found that proteins and chitosan involved in CHER (and producing peak H in the CPS method) displayed a sharp minimum in Ic (denominated as the tensammetric minimum, Tmin), observed in the potential region of the reduction signals due to CHER. In contrast, dextran, which does not produce a CHER signal, did not show any Tmin. These results show a new phenomenon in the behaviour of biomacromolecules involved in CHER on mercury electrodes.

background electrolyte in 25 mL volumetric flasks, and then measured in a glass electrochemical cell under air at room temperature. In order to denature BSA, 2 mg/mL of BSA was incubated with 8 M urea at 4 °C overnight. AC voltammetric measurements were done in a freshly diluted solution in the presence of 80 mM non-denaturing concentration of urea that did not influence the AC voltammograms of BSA. Mixtures with dextran were prepared by direct addition of dextran to the electrochemical cell containing chitosan, BSA or ConA solution. 2.2. Instrumentation and electrochemical methods Electrochemical research was conducted using a μAutolab analyser type III connected to a 663VA Stand electrode system (MetrohmAutolab, Utrecht, The Netherlands). The three electrode system consisted of a hanging mercury drop working electrode (HMDE, surface area 0.40 mm2), a glassy carbon counter electrode and an Ag/AgCl (3 M KCl) reference electrode in relation to which all potentials were expressed. Measurements were guided using the GPES 4.9 software. An AC voltammetry method with phase-in (phase angle ϕ = 0°) and phase-out (ϕ = 90°) modes was used under the following conditions: accumulation potential, EA − 0.10 V, accumulation time, tA 60 s, step potential, ES 4.95 mV and amplitude, a 25 mV. Accumulation was achieved by stirring the solution using a Teflon stirrer adjusted to 1500 rpm, followed by cathodic scan from − 0.10 V until final potential of − 1.98 V was reached. The voltammograms obtained were analysed with ECDSOFT software (ElectroChemical Data SOFTware, Zagreb, Croatia), while numerical data were further processed in Origin. We did not convert capacity current values into capacitance (an internationally recognized SI derived quantity), as our intention was to show both phase-in and phase-out AC voltammetric results on the same current scale. The adsorptive transfer (AdT) procedure included: accumulation of chitosan or BSA for tA 60 s from 0.55 M NaCl and 0.5 M acetate buffer (pH 5.14) on HMDE at different EA between − 0.10 and − 1.80 or − 1.66 V, respectively, followed by a transfer of the analyte-modified HMDE into the blank background electrolyte where the AC voltammetry was applied. A bench pH meter equipped with a glass body-Ag/AgCl electrode (Hanna Instruments, Croatia) calibrated against Hanna buffer standards (pH 4.01 and 7.01) was used for pH measurements. 3. Results 3.1. Phase-in versus phase-out AC voltammetry of biomacromolecules

2. Experimental 2.1. Chemicals and solutions Chitosan (medium Mw), dextran (Mw 2 × 106), BSA, human serum albumin (HSA), Concanavalin A (ConA), lectin from Helix pomatia and urea were purchased from Sigma-Aldrich, Germany. Prostate specific antigen (PSA) was bought from Lee Biosolutions, USA. Chitosan stock solutions were prepared by weighing and dissolving powder in 0.4 M acetic acid as it is not soluble in water [32], while all other chemicals were dissolved in Milli-Q water with a resistance of 18.2 MΩ cm prepared in Milli-Q filter apparatus (Millipore, USA). Background electrolyte consisted of 0.55 M NaCl (BioXtra, ≥ 99.5%, Sigma-Aldrich) buffered with 0.5 M acetate buffer (consisting of CH3COOH and CH3COONa; Merck-Millipore, Darmstadt, Germany), if not stated otherwise. Phosphate buffer (0.05 M, pH 7.00; consisting of NaH2PO4·H2O, ACS reagent, 98.0–100.0%, Sigma-Aldrich, Germany and Na2HPO4·7H2O, Merck-Millipore, Darmstadt, Germany) and 0.1 M McIlvaine buffer (consisting of citric acid, ≥ 99.5%, Sigma-Aldrich, Germany and Na2HPO4·7H2O, Merck-Millipore, Darmstadt, Germany) were also used. PS and protein solutions were prepared in buffered

Biomacromolecules of proteins and PSs were measured at concentrations which, according to the adsorption isotherms (not shown), assure complete surface coverage of the HMDE. Fig. 1A–C shows phase-in and phase-out AC voltammograms for chitosan, BSA and dextran in 0.55 M NaCl and 0.5 M sodium acetate buffer, pH 5.14. After 60 s accumulation at − 0.1 V, both chitosan and BSA produced a phase-in AC voltammetric faradaic peak at −1.83 V and −1.67 V (Fig. 1A and B), respectively. Considering previous chronopotentiometric and voltammetric results for chitosan [7,8] and BSA [5], we may conclude that these peaks show characteristics of CHER peaks. Compared to chitosan and BSA, dextran (PS composed of glucose, not containing any catalytic group), even at a concentration five times higher (100 μg/mL), did not produce any CHER peak (Fig. 1C). That was in accordance with data obtained previously for the same electrolyte, but using a more sensitive CPS method [31]. Tensammetric measurements for dextran reported that it strongly adsorbed on the electrode in the potential range between − 0.10 and − 1.1 V, accompanied by significant lowering of Ic (Fig. 1C). At potentials more negative than − 1.0 V dextran was desorbed. Maximum adsorption of dextran occurred not far from the point of zero charge potential (p.z.c. around −0.47 V [33]) under conditions that favoured strong adsorption of electrically neutral molecules


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or their parts. Similar adsorption features were observed for chitosan, BSA (Fig. 1A and B) and all other proteins investigated (HSA, ConA, lectin from Helix pomatia and PSA) in a very broad potential segment from −0.1 V to about −1.5 V. Opposite to that observed for dextran, instead of desorption, only biomacromolecules involved in catalytic hydrogen evolution on the HMDE showed a sharp and narrow decrease in Ic between − 1.50 and − 1.90 V. We named it the tensammetric minimum (Tmin). Tmin appeared at a particular potential: for chitosan at −1.81 V (Fig. 1A) and for BSA at −1.66 V (Fig. 1B). The amount of Ic decrease represents the difference between the background electrolyte Ic and the minimum Ic at a particular potential (ΔIc in Fig. 1A and B inset). Relative standard deviation (RSD) calculated for 10 consecutive, independent and identical measurements of Tmin for 20 μg/mL chitosan or BSA was b 1%, implying high reproducibility in detection. AC voltammetry was not able to differentiate between native and denatured BSA because native BSA undergoes denaturation during a relatively slow potential scan up to −1.98 V [34]. In agreement with secondary BSA denaturation at the electrode surface, both native and denatured BSA produced peak S (at −0.49 V) of the same height due to reduction of the Hg\\S bond [35]. Thus, it is not surprising that the same CHER peak and corresponding Tmin were observed regardless of the native or denatured form of BSA that was originally put into the electrochemical cell. Very characteristic sharp lowering in capacitance-potential curves, called a capacitance ‘pit’ has previously been observed on mercury electrodes for the 2D condensation layer of nucleic acid bases [15,36], pyrimidine oligonucleotides [37] and amyloid peptides [38], as well as for alkaloids [39]. Such capacitance pits appear in the less negative potential domain and are quite wide, while Tmin described in this paper appeared at highly negative potentials, was relatively narrow and its shape did not correspond to the pit described previously [40]. Quite similar to Tmin, a considerable AC polarographic capacity curves electrode admittance in the region of the catalysed evolution of hydrogen, called a ‘dip’, has been reported for some oxytocin and vasopressin synthetic analogues [41]. Furthermore, a minimum Ic at very negative potentials has been observed for different types of SAS present in seawater, diatom and dinoflagellate exudates measured at two different pHs (acidic and basic) [17], without any interpretation of why or how it appears. Nevertheless, our results suggested that the Tmin observed might be related to the corresponding CHER peak. Such a view is strongly supported by the fact that both effects appeared at almost the same negative potential, and that catalytically active biomacromolecules took part in both effects. The appearance of peak S at −0.49 V in solutions of BSA (Fig. 1B), HSA, lectin from Helix pomatia and PSA (not shown) confirmed immobilization of the biomacromolecules on the HMDE surface. To prove a meaningful link between two surface-related processes, we performed parallel series of phase-in and phase-out AC voltammetric experiments with chitosan and BSA under conditions that significantly alter CHER peak, such as buffer concentration, pH, analyte concentration and tA. Our results are presented below. 3.2. Effect of acetate buffer concentration

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E / V (vs. Ag/AgCl) Fig. 1. Phase-in (red line 1) and phase-out (black line 2) AC voltammograms for (A) 20 μg/ mL chitosan, (B) 20 μg/mL BSA and (C) 100 μg/mL dextran in 0.55 M NaCl and 0.5 M acetate buffer, pH 5.14 (background electrolyte, dashed lines). Inset: enlarged part (A) between −1.50 V and −1.90 V of the phase-out AC voltammogram for chitosan and (B) between − 1.50 V and −1.75 V for BSA. Blue line 3 in (B) is a phase-out AC voltammogram for 20 μg/mL BSA in unbuffered 0.55 M NaCl (voltammogram of background electrolyte was similar to the dashed black line). ΔIc in (A) and (B) represents Tmin. AC voltammetric conditions were: EA − 0.1 V, tA 60 s, ES 4.95 mV, a 25 mV. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

CHER peak and Tmin of BSA were not produced in unbuffered 0.55 M NaCl solution, where only desorption of BSA from the electrode at potentials from −1.3 to −1.9 V was observed (Fig. 1B). After the addition of 0.05 M acetate buffer, Tmin and a corresponding CHER peak appeared. The presence of buffer is mandatory for CHER, as the acid component of the buffer efficiently regenerates the catalyst by proton donation, while in the unbuffered electrolyte H3O+ ions act as only possible, but poorly effective, proton donors [6,31,42,43]. Accordingly, with increasing acetate buffer concentration from 0.05 to 0.5 M (in 0.055 M NaCl at constant pH 5.14), the CHER peak and Tmin for chitosan (Fig. 2A and B) and BSA (Fig. 2C and D) around −1.75 V increased. These results represent strong evidence that Tmin is significantly dependent on the presence of buffer.

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Fig. 2. Phase-in and phase-out AC voltammograms of increased acetate buffer concentration for (A) 20 μg/mL chitosan and (C) 20 μg/mL BSA. Dependence of Ep and height of the CHER peak (If, –○–) and of Tmin (Ic, __●__) for (B) chitosan and (D) BSA on acetate concentration. Regression equation for the CHER peak height of chitosan was If = −31.5[acetate buffer]2 + 33.7[acetate buffer] + 1.3 (r2 = 0.989) and for Tmin was Ic = 8.45[acetate buffer] + 0.03 (r2 = 0.995), while for the CHER peak of BSA, the equation was If = 61.2[acetate buffer]-0.55 (r2 = 0.999) and for Tmin was Ic = 10.2[acetate buffer] + 0.88 (r2 = 0.992). Background electrolyte was 0.055 M NaCl. The height of the chitosan peak was defined with regard to the exponential baseline. For BSA, the linear baseline was used, while Ic decrease was for both analytes calculated as the difference between Ic of the background electrolyte and the maximum decrease in Ic around −1.80 V for chitosan and around −1.70 V for BSA. Other details are given in Fig. 1.

The strong dependence on buffer concentration is in agreement with previous chronopotentiometric and voltammetric analysis of various proteins [43,44] and chitosan [7] producing CHER peaks. Thus, our results suggest that Tmin is closely related to the CHER peak. The dependence of Tmin on acetate buffer concentration for both chitosan and BSA was linear (Fig. 2B and D). Dependence of the CHER peak height of BSA on acetate buffer concentration was also linear (Fig. 2D), while for chitosan it was second order polynomial (Fig. 2B). Similar results were obtained for chitosan and BSA in solutions of increased concentration (0.05–0.5 M) of phosphate buffer, pH 7.00 (not shown). CHER peak potential, Ep and accompanying potential of Tmin shifted toward more negative values for higher acetate buffer concentrations, probably due to a stabilization effect [42]. The shift of Tmin for chitosan was from −1.76 V in 0.05 M acetate to −1.83 V in 0.5 M acetate, and for BSA it was from − 1.62 V to − 1.68 V (Fig. 2B and D). The chitosan CHER peak shifted more toward the ascending part of the baseline, so the

peak was asymmetrically shaped (Fig. 2A), while the peak for BSA was symmetrical (Fig. 2C). 3.3. Effect of pH and ionic strength The CHER peak and Tmin of chitosan and BSA showed a crucial dependence on pH in 0.1 M McIlvaine buffer. With increasing pH of McIlvaine buffer from pH 6.16 to 7.95, the CHER peak and Tmin decreased linearly (Fig. 3 for chitosan, not shown for BSA). At pH 7.95, neither CHER peaks nor Tmin were observed. At higher pH, there are fewer hydrogen ions available to undergo CHER, and therefore the rate of protonation decreases and CHER peak height reduces [42]. Moreover, these experiments were performed in McIlvaine buffer, implying that Tmin is little affected by buffer composition, as we demonstrated the appearance of the Tmin together with the CHER peak in three different buffers (acetate, phosphate and McIlvaine buffer).

S. Strmečki, E. Paleček / Bioelectrochemistry 120 (2018) 87–93 5

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3.4. Influence of chitosan and BSA concentration and tA on CHER peak and Tmin Dependence of CHER peak height and Tmin on chitosan concentration in the range from 0.2 to 57 μg/mL (tA 60 s, Fig. 4B) and on tA from 10 to 180 s (20 μg/mL chitosan, Fig. 4B inset) was studied. A notable difference between CHER peak and Tmin was observed at concentrations of chitosan lower than 10 μg/mL. Tmin was a linear function of chitosan concentration in the range 0.2–10 μg/mL, while a CHER peak appeared only at concentrations higher than 6 μg/mL, increased also linearly up to 10 μg/mL and then levelled off. The results pointed out that the sensitivity of tensammetry in the detection of chitosan reached the sensitivity of the CPS method where a CHER peak of chitosan might be also detected down to 0.2 μg/mL (after 60 s accumulation at − 0.1 V and stripping current −50 μA). At lower CPS current densities, it was possible to detect even lower chitosan concentrations. CHER peak potential, Ep − 1.83 V, shifted negligibly with chitosan concentration (Fig. 4A). In contrast, the potential of Tmin sharply shifted from − 1.76 V at 0.2 μg/mL to −1.82 V at 10 μg/mL, and retained that value at higher concentrations (Fig. 4A). At longer tA, the CHER peak and Tmin of chitosan changed linearly and levelled off at tA ≥ 60 s (Fig. 4B inset). Regarding BSA, the CHER peak and Tmin increased up to 2 μg/mL BSA and then levelled off. It is evident from Fig. 2 that BSA is a more efficient catalyst, producing a higher CHER peak than chitosan under the same measurement conditions. AC voltammetry appears to be equally sensitive for the detection of CHER peak and Tmin of BSA. 3.5. AdT AC voltammetric dependence on EA To check whether chitosan and BSA were attached to the HMDE at the potentials of their Tmin (− 1.80 and − 1.66 V, respectively), we used an AdT (ex situ) method in which the accumulation step was experimentally separated from the CHER [45]. We performed this experiment with 6 and 60 μg/mL of chitosan; only the latter concentration

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The effect of ionic strength on the CHER peak and Tmin of chitosan and BSA was studied in a solution of 0.05 M acetate buffer, pH 5.14, at concentrations of NaCl ranging from 0.1 to 1 M. CHER peak height and Tmin for chitosan (Fig. 3 inset) and BSA (not shown) changed negligibly at higher concentration of NaCl, suggesting that ionic strength variation little influences both surface processes monitored. It appears that electrostatic interactions in the solution are crucial neither for CHER nor for Tmin.

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c(chitosan) / µg/mL Fig. 4. Dependence of (A) Ep and (B) CHER peak height (If, –○–) and Tmin (Ic, —●—) on chitosan concentration. A solution of 0.55 M NaCl and 0.5 M acetate buffer, pH 5.14 was the background electrolyte. Inset: the effect of tA on CHER peak and Tmin for 20 μg/mL chitosan. Other conditions are given in Fig. 1.

assured full HMDE surface coverage. Chitosan produced the same Tmin and CHER peak regardless of different EA (not shown), fully confirming its immobilization on the HMDE at all applied EAs, including at −1.80 V where the Tmin appeared. BSA (20 μg/mL) showed the same results only for EAs less negative than − 1.66 V. Using AdT at EA − 1.66 V, both Tmin and the CHER peak of BSA diminished by 18%. That happened most probably due to the presence of negatively charged groups such as carboxyl groups in BSA that may repel this protein from the negatively charged HMDE and therefore disturb adsorption and/or orientation of BSA on HMDE. Such behaviour is related to the dynamic character of interfacial interactions of large protein biomacromolecules, especially pronounced on the highly negative HMDE surface [46]. Chitosan does not carry any negatively charged group in its structure so its CHER peak and Tmin were not disturbed at very negative EAs. Our AdT experiments confirmed adsorption of BSA on the HMDE at the potential of Tmin.

3.6. AC voltammetric detection of catalytically active biomolecules in a mixture with catalytically inactive PS The tensammetric results described in Section 3.1 (Fig. 1) suggest that chitosan and BSA were strongly adsorbed, while dextran (an otherwise catalytically inactive PS) fully desorbed from the electrode at potentials more negative than − 1.2 V. We were interested in whether different adsorption/desorption features at the negatively charged HMDE might serve to discriminate catalytically active from catalytically inactive types of biomacromolecule in the mixture. Citosan-dextran and BSA-dextran mixtures were measured by AC voltammetry. Our results showed that the CHER peak of chitosan changed by 31% and Tmin by 14% when in a mixture with 100 μg/mL dextran (Fig. 5A). On the other hand, no significant change in CHER peak and Tmin of BSA was observed upon addition of dextran (Fig. 5B). Furthermore, we wondered if specific interactions between dextran and an electrocatalytically active protein would change the AC voltammograms of such a protein. For that purpose, voltammograms of 15 μg/mL of ConA, lectin from Canavalia ensiformis (binding to α-D-glucose- and α-

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E / V (vs. Ag/AgCl) Fig. 5. AC voltammograms of (A) 20 μg/mL chitosan, (B) 20 μg/mL BSA and (C) 15 μg/mL ConA alone (solid lines) and in a mixture with 100 μg/mL dextran (bold solid lines). Red lines (1) present phase-in and black (2) phase-out AC voltammograms. Dashed lines were recorded in the corresponding background electrolyte 0.55 M NaCl and 0.5 M acetate buffer, pH 5.14. Other conditions are given in Fig. 1. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

D-mannose [47]), with and without addition of 100 μg/mL dextran, were

recorded. Specific lectin-carbohydrate interactions in solution are in general stronger than nonspecific protein-carbohydrate interactions [48]. At the HMDE, dextran lowered the CHER peak of ConA by 16% and changed Tmin by 7% (Fig. 5C), which is comparable to the nonspecific influence on chitosan we observed. Our results thus suggest that both the CHER peak and Tmin yielded by chitosan or proteins can be observed even in the presence of dextran, i.e. a PS not producing electrocatalytic signals, and that the adsorption/desorption behaviour of dextran differed greatly from that of the electrocatalytically active biomacromolecules studied. At this stage, our results do not allow us to offer a detailed explanation of the interaction of the studied biomacromolecules at the HMDE, which may involve competitive adsorption as well as mutual interactions of the biomacromolecules in solution followed by biomacromolecule adsorption. In this short section, we only showed that the adsorption/desorption behaviour of electrocatalytically active biomacromolecules differs greatly from that of dextran, thus allowing analysis of mixtures of these compounds.

4. Discussion Our results suggest that Tmin is a consequence of strong adsorption of positively charged species making a specially oriented layer on HMDE. We found participants in CHER as reasonable candidates that might be a part of such a layer. AdT study implied that the catalysts are anchored on the HMDE at the potential of its Tmin. The other possibility for formation of Tmin could be a high excess of hydrogen ions that are attracted to the HMDE at a particular negative potential in order to be reduced into hydrogen molecules. However, we could not exclude the possibility that both catalyst and hydrogen ions are mandatory for formation of Tmin. The effect or participation of hydrogen molecules, as a product of CHER, on the adsorbed layer in slow scan AC voltammetry may appear less probable. On the other hand, catalyst structure appears to be of paramount importance for Tmin, as it is for CHER peak. ConA is a lectin that does not contain any Cys [49], so only nitrogen-containing amino acid residues such as Arg, Lys and His appear to be involved in CHER. It is known that chitosan is positively charged under mildly acidic conditions, and this characteristic allows deposition (i.e. ‘self-assembling’) of a thin chitosan layer on a negative electrode under an applied voltage [50]. Therefore, it might be that amino groups in chitosan, apart from being catalytic groups [8], also have an important

role in the formation of a specifically oriented layer on HMDE, resulting in Tmin. Nevertheless, we observed that the PSs carrageenans, although having negatively charged sulfate groups, produce similar Tmin (unpublished). It is crucial to conduct more experiments to elucidate the real composition of the adsorbed layer and the role of specific catalytic groups in this layer. 5. Conclusions In this paper, we have described for the first time a narrow decrease in Ic at highly negative potentials, called the tensammetric minimum, Tmin. Tmin was observed in phase-out AC voltammograms using HMDE, and it appears to be very specific for biomacromolecules involved in CHER such as proteins and some PSs. Detailed AC voltammetric study of BSA and chitosan in a buffer solution showed that Tmin is significantly related to CHER. A quite similar dependence of CHER peak and Tmin on buffer concentration (Fig. 2) and pH (Fig. 3) was observed, representing the main experiments highly characteristic for CHER. Furthermore, Tmin was increased linearly with analyte concentration (at tA 60 s) or tA (at constant analyte concentration) until full electrode surface coverage was reached (Fig. 4), in accordance with the behaviour of the CHER peak, again showing their close relationship. It seems that Tmin, besides the CHER peak, could serve as new voltammetric evidence for the catalysed reduction of hydrogen ions on a mercury electrode. Tmin may find a purpose in distinction of biomacromolecules that are involved in CHER from those showing catalytic inactivity (Fig. 1), gaining information about the catalyst structure, i.e. the presence of possible catalytic groups in the molecule. Furthermore, it seems that Tmin is related to the adsorption/desorption surface processes of biomacromolecules involved in CHER on a mercury electrode, and it is influenced by tight interplay of the electrode surface charge with biomacromolecular structure and charge. Acknowledgements The authors thank to Drs. V. Dorčak and M. Plavšić for critical reading of the manuscript and Dr. M. Lovrić for fruitful discussion. Financial support for this work was provided from the EU Seventh Framework Programme under grant agreement no. 291823 Marie Curie FP7-PEOPLE2011-COFUND (NEWFELPRO, to S.S.). This work was also supported by Czech Science Foundation 15-15479S project (to E.P.) and Croatian Science Foundation project No. 8607 (AMBIOMERES).


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