Abstract
Electrochemical methods can be used not only for the sensitive analysis of proteins but also for deeper research into their structure, transport functions (transfer of electrons and protons), and sensing their interactions with soft and solid surfaces. Last but not least, electrochemical tools are useful for investigating the effect of an electric field on protein structure, the direct application of electrochemical methods for controlling protein function, or the micromanipulation of supramolecular protein structures. There are many experimental arrangements (modalities), from the classic configuration that works with an electrochemical cell to miniaturized electrochemical sensors and microchip platforms. The support of computational chemistry methods which appropriately complement the interpretation framework of experimental results is also important. This text describes recent directions in electrochemical methods for the determination of proteins and briefly summarizes available methodologies for the selective labeling of proteins using redox-active probes. Attention is also paid to the theoretical aspects of electron transport and the effect of an external electric field on the structure of selected proteins. Instead of providing a comprehensive overview, we aim to highlight areas of interest that have not been summarized recently, but, at the same time, represent current trends in the field.
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Introduction
Proteins are structural, functional, and regulatory elements of cells and tissues. Current knowledge about proteins is closely connected with progress in structural biology research and development and the determination of the structure of a number of proteins [1], as well as in analytical instrumentations and methodologies. Interest in the electrochemical analysis of proteins was initiated only 6 years after J. Heyrovský’s invention of polarography [2] by discovering a new phenomenon manifested by the ability of proteins to catalyze hydrogen evolution at a dropping mercury electrode (DME). Circumstances leading to this significant discovery (in the absence or presence of cobalt ions), along with early polarographic investigations of proteins, are described in the literature [3, 4].
Later, the DME was gradually replaced with gold, silver, platinum, and graphite electrodes modified with various adsorbates to study a relatively small group of conjugated proteins, usually containing non-proteinaceous redox-active metal centers (prosthetic groups or cofactors) that provided fast reversible redox electrode processes. Great interest in this branch of protein electrochemistry (also called protein film voltammetry or protein film electrochemistry) was induced by a very important finding, showing that appreciable electron transport between the electrode and protein redox-active centers, which are not accessible to the electrode surface, can be achieved using an effective electron transfer mediator [5, 6]. Mostly, proteins containing heme, iron-sulfur, or copper redox centers are investigated, and methodologies for their attachment onto a variety of conducting surfaces and assemblies of them that are useful for probing biological redox processes have been recently reviewed [7,8,9].
The isolation and structural elucidation of key membrane proteins, both transporters and receptors, gave a new impulse to electrochemical studies of proteins connected with the development of various biomimetic membranes or simple detergent and lipid layers [10, 11]. Furthermore, due to advances in the construction of various electrochemical (bio)sensors and proteomic approaches [12, 13], labeling proteins with redox-active probes has been established, and electrochemiluminescence has been utilized for protein analysis [13]. Today’s trends are also directed toward the electrochemical sensing of proteins at the atomic level and taking advantage of computational tools to study protein interactions with electrode surfaces and electron transfer phenomena in general [14]. Finally, it is worth mentioning that close attention has to be paid to the adsorption of proteins onto electrode surfaces, as well as to the structural and functional changes that may occur in them when exposed to an electric field, to avoid misinterpretation of the results obtained, particularly in investigations of protein structures and interactions with other bio(macro)molecules or substances [15,16,17,18]. As for future trends in single-molecule analysis and sequencing, the application of proteins as nanopores, and nanopore technologies in general, are also very promising [19,20,21,22]. A schematic overview of general electrochemical approaches useful for protein studies is shown in Fig. 1.
This text does not aim to provide a comprehensive overview, but instead to point out individual trends, especially in research on the electroactivity of non-conjugated and membrane proteins. Furthermore, we aim to describe the effects that occur after the exposure of proteins to an electric field, and advanced computational tools are also highlighted. In addition, we summarize the basic strategies in protein microanalysis and labeling (or electrochemically promoted labeling) with redox-active probes. Particular attention is paid to areas not reviewed in this form in recent years.
For further study, we recommend the following comprehensive reviews on proteomics and glycomics [18], membrane proteins [11], and on the electrochemical research of peptides [15]. Historical aspects in this regard have also been recently reported [31].
Intrinsic electroactivity of proteins
Over the last few decades, significant progress in the electrochemical analysis of proteins has been made using constant-current chronopotentiometric stripping (CPS) in combination with mercury-containing electrodes (reviewed in [17, 18, 25, 27]). Under conditions close to physiological, proteins containing Arg, Lys, Cys, or His amino acid (aa) residues [32,33,34] produce a well-developed peak (so-called peak H) due to the catalytic hydrogen evolution reaction (CHER) [18, 35]. Methods based on CPS peak H can be applied for the label-free reagentless structure-sensitive analysis of practically any protein, since proteins that do not contain any of these residues, if any, are extremely rare. An important condition for obtaining protein peak H is the accessibility of the catalytically active aa residues for the electrode process. In a native folded protein, aa residues buried in the interior of the molecule and/or those located far from the electrode surface can remain catalytically silent. On the other hand, they may become involved in CHER after the protein’s denaturation [25, 36]. Even the first works utilizing CPS peak H demonstrated the possibility of studying local and global changes in protein structures [25, 36] due to the ability of surface-attached proteins to retain their folded structures close to the potential of zero charge, but undergo time-dependent denaturation/unfolding at negatively charged surfaces [17, 37, 38]. The denaturation of surface-attached proteins can be minimized by adjusting the duration of the protein exposure to the electric field to milliseconds [39], as well as other experimental conditions, such as solution temperature [37] and ionic strength [40]. The high sensitivity of the CPS peak H to structural changes in proteins can be utilized not only for monitoring protein denaturation [41,42,43], oligomerization and aggregation [44, 45], posttranslational modifications [46,47,48], oxidative damage [49, 50], and single-aa replacements [51, 52] but also for investigating protein interactions with DNA [53,54,55], peptides [56], and other proteins [57,58,59,60]. CPS peak H appeared to be particularly useful for analyzing water-soluble and membrane proteins [11, 61,62,63,64,65]. All these analyses are based on utilizing the different accessibilities of electroactive residues, which is influenced by the adsorption and/or structural stability of the given protein or its complex. Peak H appears at highly negative potentials close to –1.7 V (vs. Ag|AgCl|3M KCl). At such negative electrode potentials, an extremely high electric field (109 V/m) [66] can affect the electric double-layer with adsorbed biomolecules. Moreover, it can cause DNA melting, protein denaturation, complex disaggregation, etc. [18].
CPS peak H was also utilized to study proteins in complex media [53] as well as to analyze clinical samples, such as human serum albumin (HSA) samples isolated from blood serum [67]. The surface distribution of aa residues active in CHER in the HSA molecule is shown in Fig. 2A–C. HSA isolated from healthy volunteers gave a CPS peak H which decreased after modification of the sample with methylglyoxal (MGO); see Fig. 2D. MGO is a reactive metabolite that is able to modify the same aa residues that are involved in the CHER. Based on the results of this study, the coefficient of variation for the native albumin samples was estimated to be 8.5%, while that for the inter-individual binding capacity variations, evaluated using the artificial-glycation/carbonylation approach, was 23.2 % (Fig. 2E). Recently, interactions of HSA with fatty acids and their nitro-derivatives were also investigated using CPS peak H [68, 69]. In addition to hanging mercury drop electrode (HMDE), CPS analysis in combination with an Ag-amalgam electrode microdevice can be effectively applied for CHER monitoring, which was demonstrated in bovine serum albumin sensing; see Fig. 3 [70].
In addition to proteins, also some peptides, oligo- and poly-nucleotides, and oligo- and polysaccharides have been found to be catalytically active in the hydrogen evolution reaction at mercury-containing electrodes (reviewed in the literature [15, 17, 18, 27]). Nevertheless, despite some attempts to better understand CHER at the fundamental level [32,33,34,35, 71,72,73,74], methodologies utilizing CHER are still mostly based on empirical findings, and further research is necessary to exploit the full potential of this electrocatalytic phenomenon in the research of various bio(macro)molecules and their mutual interactions.
Besides the electrocatalytic reduction occurring at mercury-containing electrodes, the oxidation reactions of proteins at carbon electrodes have also been found to be useful for their label-free analysis [18, 25, 75,76,77]. Even though some aa’s, namely Trp, Tyr, Cys, His, and Met, are oxidizable at carbon electrodes [18, 78,79,80], it is predominantly Tyr and Trp residues that have yielded well-developed oxidation peak/s with proteins [78]. Peptides and small proteins yielded separated peaks of Tyr and Trp residues [75, 81]. However, larger proteins, in most cases, only produced a single peak. The well-developed Tyr and/or Trp peak/s, in contrast to the poor or absent peaks of Met, His, and Cys residues in proteins, could be due to the stronger interactions of Tyr and Trp residues with the electrode surface than those of other electroactive residues, as a theoretical study showed [82]. A well-developed His peak was observed for oncoprotein AGR2 modified with a His-tag [80], since the six linked His residues on its terminus are more accessible for oxidation than those buried inside the protein structure. Aa residues of proteins are more exposed after protein denaturation. Denaturation of the AGR2 protein led to the appearance of a negligible His peak and an increase in the Tyr and Trp peak [80]. Severalfold higher Tyr and Trp peaks for denatured forms than those for native ones were also reported for other proteins [78, 83,84,85]. Oxidation of the Tyr and/or Trp residues was also found to be useful for studying the oligomerization and aggregation of alpha-synuclein and amyloid peptides [86,87,88]. Similarly, the posttranslational modification of peptide and protein Tyr and Trp residues, such as phosphorylation [89] and nitration [90], as well as oxidative damage [50] and ligand binding [69, 91], had an impact on the oxidation responses of Trp or Tyr residues.
Electroactive redox labels in protein sensing
Protein labeling is generally performed via Lys residues with N-hydroxysuccinimide-activated esters, sulfonyl chlorides, or iso(thio)cyanates. Another option is Cys labeling by Michael reaction or reactions that target electron-rich Tyr or Trp residues [92, 93]. The electrochemically promoted Tyr-modification of peptides and proteins with labeled urazoles was studied at the low potential of +0.36 V (vs. Ag|AgCl|sat. KCl). Under these conditions, the urazole anchors could be activated without oxidizing the sensitive aa residues in the protein. Protocols were successfully performed in the electrosynthesis of peptides and proteins, such as oxytocin, angiotensin, BSA, and epratuzumab. An electrochemically promoted labeling approach was also developed for Tyr-containing proteins with phenothiazine derivatives [94]. The electro-oxidation of phenothiazine produces a nitrogen radical cation, which reacts with the ortho position of the Tyr phenol. Two proteins, which contain Tyr on the protein surface, insulin, and myoglobin, were modified with phenothiazine [94]. Similar to the electrochemically promoted Tyr-click reaction, a bioconjugation reaction for selective Trp labeling in peptides and proteins has been developed (Fig. 4A) [100].
Boronic acid functionalized compounds have been utilized for biosensing glycoproteins. A schematic representation of the interaction of glycoprotein with modified boronic acid is shown in Fig. 4B. Boronic acids interact with 1,2- or 1,3-diols of saccharide to create five/six-membered cyclic complexes and also interact with Lewis bases to form boronate anions [101]. An amperometric sensor was constructed for monitoring fructosyl valine, the glycosylated part of hemoglobin, based on soluble ferrocenylboronic acid [102]. The glycosylated part of hemoglobin was investigated at the carbon electrode via a ferrocene moiety. An electrochemical method for glycoprotein detection based on 4-mercaptophenylboronic acid (MBA)/biotin-modified gold nanoparticles (AuNPs) was used for the study of recombinant human erythropoietin (rHuEPO) as a model protein. In more detail, rHuEPO was first captured by an electrode covered with anti-rHuEPO aptamer and then derivatized with MBA-biotin-AuNPs. The MBA-biotin-AuNPs interact with streptavidin-conjugated alkaline phosphatase to produce electroactive p-aminophenol [7]. Electrochemical biosensors based on MBA-capped AuNPs were also used for monitoring a prostate-specific antigen and avidin. A sub-picomolar limit of detection of avidin/prostate-specific antigen was achieved [103].
There have been several studies focused on utilizing osmium complexes for the electrochemical analysis of peptides and proteins. A complex composed of osmium tetraoxide and 2,2´-bipyridine was used for the labeling and electrochemical detection of Trp residues of salmon and human luteinizing hormone [104], avidin, streptavidin, and lysozyme [105]. Osmium(VI) complexes (ligands: 2,2′-bipyridine and N,N,N′,N′-tetramethylethylenediamine) were also used for labeling the sugar part of glycoproteins with an electrochemical detection endpoint. This approach was applied to determine RNase B and avidin, with the limit of detection (LOD) ranging between 25 and 50 nM. Electrochemical signals were monitored at a pyrolytic graphite electrode by adsorptive transfer stripping square-wave voltammetry [106, 107].
The most commonly used detectors coupled to separation techniques in proteomics are mass spectrometry and laser-induced fluorescence detection. An alternative to the above-mentioned detectors is electrochemical detection, especially amperometric and pulse amperometric detection [95]. The direct detection of a redox-active aa on carbon electrodes or the utilization of metal-based solid electrodes is limited by the LOD [76]. The most commonly used derivatization agents in terms of aa’s and peptides are o-phthaldialdehyde (OPA) and naphatalene-2,3-dicarboxyaldehyde (NDA) in the presence of a nucleophile (sulfur derivatives or CN−), reviewed in the literature [95]. Other agents utilized in the derivatization of aa’s are 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate (6-AQC) [96, 97] and p-nitrophenol-2,5-dihydroxyphenylacetate bis-tetrahydropyranyl ether [98, 99]. The protein interaction with the most common derivatization agents is demonstrated in Fig. 4C.
A peptide-1 probe (RNRCKGTDVQAW) was designed as an electroactive label of daunomycin for ovalbumin protein recognition. The peak current of the daunomycin moiety decreased with increasing concentration of ovalbumin due to the interaction between ovalbumin and the electroactive peptide probe. Differential pulse voltammograms of daunomycin and labeled peptides in the presence or absence of ovalbumin were obtained using a glassy carbon electrode. According to this protocol, the concentrations of ovalbumin in the egg whites were measured with a detection limit at the 10−11 M level.
An electrochemical sensor was also developed [108] for monitoring the following protein kinases: sarcoma-related kinase, extracellular signal-regulated kinase 1, and cyclin A-dependent kinase 2. The approach is based on the ability of kinases to transfer a redox-labeled phosphoryl group, the specific substrate for the protein kinase, to surface-bound peptides. Voltammetric and electrochemical impedance spectroscopic detection was enabled due to 5′-γ-ferrocenoyl-ATP, a co-substrate for peptide phosphorylation. The labeling strategies are schematically summarized in Fig. 4.
Nano/micro materials in protein electrochemistry: double-surface technique
In the last decade, there has been a significant increase in published studies in which the authors use a variety of nano- and micromaterial technologies for the study and sensitive electrochemical analysis of proteins. Very often, these are applications of carbon or metal nanoparticles, which are used to decorate the surfaces of the electrodes, often complex multi-layer or multi-component modifications [109]. These systems are effectively applied to increase the working (active) surface of the sensor, improve the sensitivity/selectivity of the determination, or increase the electron transfer rate between the protein and the electrode, e.g., in the development of enzyme electrodes or biofuel cells [110]. On the other hand, a certain disadvantage of these procedures could be the poor reproducibility of the preparation of such complex electrode architectures. In fact, for fundamental research, it is usually best to use an unmodified (bare) electrode with a well-defined and reproducible surface. In addition, the preparation of complicated (e.g., sandwich configuration) structures on the surfaces of electrodes goes against the main added value of electrochemical determinations, which is the simplicity (“elegance”) of the experimental setup, possibly even the minimal financial demands of performing such analysis or research; see a recent historical review [111].
One of the other possibilities where we can use nano/micro technologies in the electrochemistry of biomacromolecules is the concept of the double-surface technique (DST) [112, 113]. This is based on the application of microparticles or a selected nano/micro material (“first surface”) for the manipulation or purification of the investigated proteins before their adsorption onto the electrode detection surface (“second surface”) and subsequent electrochemical analysis. After the release of the protein from the first surface, the adsorptive transfer (AdT) technique [114] can be used. This method allows the protein to be adsorbed onto the surface of the electrode from microliter volumes, and after washing the electrode, the biopolymer-modified electrode is inserted into an electrochemical cell containing an already pure supporting electrolyte. For DST purposes, magnetic nano- or micromaterials can be beneficial. Originally, these approaches were applied to the study of DNA interactions and hybridization. In this particular case, magnetic beads (magnetoseparation) were used [115, 116]. DNA was bound (anchored) to their surface, most often via a terminal oligo(A) sequence or a biotinylated terminus to the oligo(T) chain or (strept)avidin immobilized on the surface of magnetic beads, e.g., ref. [117]. The immobilized DNA (but also RNA) can be easily purified (washing step) and further incubated. The washing is based on the repetitive magnetic attraction of the beads to the wall of the plastic microtube and the consequent resuspension step, which allows multiple purification cycles [115]. Subsequently, the target biopolymer is released from the magnetic carrier and dissolved in a buffer or medium that is fully compatible and optimized for analysis.
This technique is applicable to the research of protein-DNA interactions [118, 119] and can also be used for the electrochemical analysis of proteins, both labeled and unlabeled, according to the approaches mentioned above in the text. In such cases, proteins can be bound to magnetic particles using antibodies or aptamers (Fig. 5) [120].
Electro-manipulation of protein structure and function
An electric field (EF) acts as a direct force on charged groups in the proteins, and an EF can also act on a protein indirectly through its action on the charges of the surrounding ions and solvent. The presence of ions and (polarizable) solvent also decreases the effective EF strength by Coulombic screening [121]. The most trivial effect is the net translation (electrophoretic) force on a protein (see Fig. 6) used in a variety of separation and detection techniques. An EF, even an intrinsic protein EF, also naturally acts on electrostatic interactions (including Coulomb interactions) in the protein [122] and on protein-solvent interactions. The electric double-layer around the protein, a simplified picture of the complex charge distribution at the interface of the protein and solvent [123], can be potentially also perturbed by an external EF affecting the balance of the forces of the protein. The charge distribution on the protein itself forms an effective dipole, on which the EF acts as a torque, leading to the rotation of the protein [124, 125]. Electric forces can also cause overall deformation [126] of the protein, leading to a change in the secondary structure and, provided the electric force is high enough, ultimately to unfolding [127,128,129]. All the above-mentioned effects of EF on protein structure can lead to a plethora of functional effects.
Electric fields in the form of short (nanosecond-microsecond) intense pulses (pulsed electric field, PEF) are of particular interest for the electro-manipulation of proteins for several reasons. First, very high electric fields (>units and tens of MV/m) are strong enough to affect the protein structure [126]. Second, intense electric pulses of nanosecond-microsecond duration can only carry a small amount of energy, so they cause little to no heating. Appropriate guidance by theories and models is needed to rationally guide the formulation of hypotheses for experiments. Computational molecular dynamics (MD) simulation is such a modeling tool, furthermore with the ultimate spatial and temporal resolution, so far unmatched by any experimental technique. Hence, MD simulations enable exploration of the effects of an EF on molecules and proteins at the atomistic level and with time resolution down to femtoseconds [130, 131]. Using MD simulations, it has been demonstrated that an intense EF can rotate the protein, affect (i) the protein’s secondary and tertiary structure [132,133,134], (ii) the radius of the gyration [135,136,137], (iii) dipole moment [138, 139], and ultimately lead to unfolding [140, 141]. At the level of peptide and protein ensembles, the EF causes disaggregation and the detachment of subunits from multimeric protein complexes [142, 143].
Here, we highlight effects which most commonly appear in the literature analyzing the effect of an intense EF on proteins in silico. One of the direct effects of EF on proteins is the rotation of the protein by a torque exerted by the EF. The dipole moment of a protein arises from the charge distribution in the protein, and the effects of EFs on protein dipole moments have been extensively studied using MD simulations [130, 143,144,145,146]. Myoglobin polarization under pulsed/static EFs exhibited a fast transition with high-intensity EFs and an increase in dipole moment with lower-intensity EFs, despite minimal impact on the protein’s structure or geometry [140]. Tubulin proteins, with a high charge and dipole moment, exhibited polarization-induced changes in shape and orientation under EFs, influencing binding sites and potential applications in protein-drug interactions and ion channels [133]. Recent work has highlighted the non-linear effects of high-intensity continuous-wave EFs and emphasized the importance of operating within a “weak field condition intensity range” in MD simulations to avoid significant non-linear and saturation effects [147, 148]. Manipulating protein orientation through their dipole moment using EFs also has promising implications in the X-ray imaging of single molecules, allowing structure determination with smaller sets of diffraction data [149]. Simulations demonstrated an “orientation window” of field strengths in which proteins maintained intact structures, while longer exposure times shifted the window toward lower fields, suggesting “orientation before destruction” [125, 149].
Other substantial effects of a strong EF on proteins are the change in the protein’s secondary structure and unfolding. For example, extensive MD simulations on myoglobin showed that both static and nanosecond pulsed EFs disrupt about 70% of its α-helical secondary structure [140]. However, EF intensities below 100 MV/m have no observable impact on the secondary structure or geometry of myoglobin [140]. Insulin’s response to EFs varied based on the type and intensity, with 500 MV/m causing more significant disruption than static fields of the same intensity [150]. Studies on hen egg white lysozyme revealed denaturation under oscillating EFs, while high field strengths induced similar unfolding pathways [144, 145]. MD simulations of the SOD1 enzyme showed that 100 MV/m had no effect on secondary structures, 500 MV/m caused partial denaturation, and 700 MV/m led to complete unfolding [151]. The unfolding of ubiquitin protein using static EFs exhibited an intensity-dependent speed, with medium and high strengths inducing rapid unfolding, where deliberate unfolding using EFs provided valuable insights into protein stability [141]. There is also growing experimental evidence for a variety of these simulation predictions. For example, it was demonstrated that an intense EF affects the secondary structure of BSA [152], whey proteins [153], and lysozyme [154] using circular dichroism spectroscopy.
At the level of larger protein structures and polymers, it was shown, for example, that an intense EF can significantly affect the microtubule (MT) lattice. It was found that a nanosecond-scale intense electric field can induce a longitudinal opening of the cylindrical shell of the MT lattice, modifying the structure of the MT. This effect is field strength- and temperature-dependent and occurs on the cathode side [143]. MD simulations suggest that a high EF strength (at least tens or even a few hundreds of MV/m) is required to affect protein structures.
It is often difficult to achieve such values of field strength while being able to observe the behavior of proteins at the same time. To address this challenge, there is an ongoing development of new technological platforms (Fig. 7) which integrate planar electromagnetic chips into advanced light microscopy and spectroscopy systems [157,158,159]. These chips enable the controlled delivery of intense short electric pulses to protein samples, while the microscopes make it possible to observe the response of proteins to the EF in situ, in real time, and in a biologically relevant chemical environment. For instance, the delivery of 6 MV/m 11 ns pulses to biosamples on such a chip integrated into a structured illumination microscope has been recently demonstrated [156]. In that study, it was shown that these electric pulses can remodel the cellular microtubule network in rat basophilic leukemia (RBL) cells. In a follow-up work, it was revealed that the pulsed EF (PEF) also exerts similar effects on the microtubule network in human osteosarcoma (U2OS) cells as well as retinal pigment (RPE1) cells [160]. Furthermore, several works showed that μs and ns electric pulses affect the microtubule cytoskeleton in a variety of cells; see more in a recent review [161]. These effects of intense PEF delivered in short pulses on the microtubule network in cells are very promising for potential therapeutic applications, but the inevitable side effects of ns PEF on cellular complexity and the cell membrane obscure the mechanism of action. In short, the effects observed on microtubules in cells could be an effect of downstream signaling due to the primary action of PEF on the membrane (causing electroporation) or on membrane voltage-gated ion channels.
Therefore, there is an ongoing effort to understand the effects of PEF on well-defined reconstituted systems, such as giant unilamellar vesicles [162], vesicles with actin [163], or isolated protein structures [164,165,166]. For example, a study was conducted to examine the direct impact of PEF on tubulin [109]. There, nanosecond PEF was applied to isolated unpolymerized tubulin, and it changed the tubulin’s self-assembly capability. The change was reversible or irreversible, depending on the pulse parameters.
We foresee that the combination of chips enabling pulsed electric/electromagnetic field delivery with a variety of advanced spectroscopy and microscopy systems might enable breakthroughs in several fields:
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physical chemistry and electrochemistry: providing insight into the mechanisms of action of an electric field on non-covalent interactions in proteins,
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spectroscopy: aligning dipolar proteins with an electric field enables enhanced signals in X-ray scattering for single-molecule imaging, microwave, THz, and IR spectroscopy,
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structural biology: an electric field represents a physical handle for protein unfolding,
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physical biochemistry: the controlled activation of proteins in solution or a gaseous phase to probe the protein’s stability under different conditions,
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biotechnology: controlling the enzymatic activity of proteins in bulk,
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bioelectromagnetics: understanding in broad terms how an electric field affects biological systems at the molecular level,
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biosensing and analytics: controlling mass transport in biosensor applications.
Theory and computation of electron transfer in proteins
Electron transfer (ET) at heterogeneous bio/metallic interfaces is traditionally studied by electrochemical methods, for example, protein film voltammetry [167,168,169,170]. Redox-active biomolecules such as metalloproteins are adsorbed onto the electrode surfaces, where their charge-transfer properties are probed by measuring current-voltage responses [171, 172]. Besides fundamental studies focused on the electronic behavior of the biomolecules, many interesting applications exploiting the natural biocompatibility, high selectivity, and enzymatic activity of suitable redox proteins have been developed, including manufacturing accurate biosensors, fuel cells, or enzyme-based biocatalysts [173,174,175,176,177]. Motivated by the efficient ET capabilities of metalloproteins, these biomolecules have started to be also incorporated into vacuum-based nanoelectronics, where solid-state protein junctions are created between metal contacts to form devices such as bio-based transistors or memristors [178,179,180,181]. However, unexpected quantum phenomena emerged at such bio/metallic interfaces, which soon attracted the attention of the broader research community [182, 183].
In aqueous solutions, redox-active proteins are known to transfer charge by the so-called incoherent hopping mechanism, theoretically described by the Marcus theory [184, 185]. The electron is localized to a redox site, where it stays for a long enough time to allow the relaxation of the molecular environment to the perturbed electrostatic potential. The energy needed to overcome the free energy barriers separating the individual redox sites is provided by the fluctuating electrostatic fields arising from the thermal motions of the protein and nearby hydration layers. Therefore, the hopping mechanism is strongly temperature-dependent. However, when single-protein junctions began to be probed by scanning tunneling microscopy or its electrochemical variant [186,187,188,189], unexpectedly high electric currents were detected, exhibiting practically no dependence on temperature [190]. Surprisingly, these data indicated that electrons could coherently tunnel through the protein, which is a fundamentally different charge-transport mechanism not typical for soft biomatter. Even more puzzlingly, the redox activity of the proteins, necessary for ET in their native environments, was shown to not affect the protein conductance when incorporated into metallic solid-state junctions [191, 192].
Knowledge of the adsorption structure of a protein on electrode surfaces at atomistic and electronic resolutions is essential for understanding the ET at the interfaces [9, 193, 194]. However, such details are hardly obtainable by experimental measurements. Therefore, computer simulations are often utilized to elucidate the structural data and the transport mechanism. Classical MD based on empirical potentials is used to predict representative adsorption structures on model surfaces, where image-charge interactions at the highly polarizable metal surfaces or covalent interactions (i.e., chemisorption) must be treated with special care [195,196,197,198,199,200,201,202]. Adsorption of the proteins into the desired conformation on the surface is often controlled by chemical modifications (for example, by introducing a reactive group to the biomolecular structure by protein engineering methods or coating the surface with suitable linkers) and can be enhanced by the application of external fields [195, 203,204,205].
The electron localization typical for the hopping mechanism enables the application of combined quantum-mechanical/molecular mechanical (QM/MM) methods or their semi-empirical variants, such as the perturbed matrix method (PMM), where only the redox sites undergoing the oxidation/reduction processes are treated at the quantum level of theory, while the rest of the system is described by less-demanding empirical potentials [14, 206,207,208]. These methods are used to sample vertical ionization energy on the MD trajectories of oxidized/reduced systems, from which redox potentials and reorganization free energies are obtained by reconstructing Marcus parabolic free energy surfaces (Fig. 8A). The system response to the change in charge is often linear, which simplifies the calculation of these ET parameters [209,210,211]. The electronic coupling elements needed for determining the transfer rate constants can be obtained by different approaches, of which the most popular are the generalized Mulliken-Hush method [212, 213], fragment charge and energy difference techniques [214, 215], or DFT-based approaches such as FODFT [216, 217], CDFT [218,219,220,221], and POD [222,223,224,225]. For the interfacial ET between the electrode and the molecule, Marcus-Hush-Chidsey integrals have to be evaluated to obtain the rate constants [226, 227]. These are then brought together into a kinetic master equation for the site populations (Fig. 8B). By solving the equation, either by iterative techniques or the kinetic Monte Carlo method, the desired electronic flux is obtained [228,229,230].
Investigation of the coherent tunneling processes in protein junctions is, from the computational point of view, a much more challenging task. Transport at molecular junctions between metal contacts is usually studied within the framework of non-equilibrium Green’s function theory (NEGF) combined with tight-binding models or DFT [231,232,233,234]. Although this approach is formally accurate, in practice, it is only applicable to small systems because of its high computational demand. Studies of protein junctions are scarce because they require a full quantum description of relatively large proteins and electrode contacts. Such large-scale DFT calculations were recently demonstrated on the blue-copper protein azurin [235] and the small-tetraheme cytochrome [236, 237]. In the latter case, the tunneling currents were computed within the Landauer-Büttiker formalism (Fig. 8C), where the transmission function was treated in the Breit-Wigner approximation [238,239,240], regarding the transferred electrons as independent. Despite these necessary simplifications, the computed current-voltage curves agree well with experimental data, and the visualized conduction channels (Fig. 8D) helped with understanding the incoherent tunneling transport in such large molecules [236, 237].
Knowledge of the correct ET mechanism for the specific system is thus crucial for interpreting the measured data and performing computer simulations. While electron hopping is typical for electrochemical interfaces at one electrode, the tunneling mechanism can occur in protein junctions with two electrode contacts. The key factor controlling the ET mechanism in bio/metallic junctions is the electronic level alignment between the electrode and the molecule [182]. When there is a significant difference between the molecular redox states and the electrode Fermi level, the electron (hole) injection/ejection at the interface becomes the limiting step for the hopping mechanism, substantially lowering its efficiency, and the electronic charge is transported by coherent tunneling. For weakly adsorbed systems, which is usually the case with proteins, the alignment can be computationally estimated, for example, by the DFT+Σ technique, in which the Kohn-Sham DFT states obtained in generalized gradient approximation (GGA) are corrected for self-interaction error and missing image-charge interactions [241, 242]. These corrections can reach magnitudes of up to 1 eV and are essential for quantitative calculations of the interfacial ET [197]. However, as molecular electronics is a rapidly developing research field, new methods and approaches are being designed and optimized for these kinds of calculations.
Besides these fundamental concerns about the transport mechanism, the geometrical arrangements of the interfaces involving proteins and their electronic structures are affected by local EFs, as discussed in the previous section. These are dominated by their intrinsic components stemming from the structural and chemical composition, for example, by the location of various charged or polarized atomic groups [243,244,245,246,247]. The intrinsic fields are thus highly localized, oriented, and relatively strong. Therefore, they typically control the adsorption orientations and confinement [195, 248, 249]. On the other hand, the external fields, induced by applied bias potentials, are considerably weaker; however, they can vary over time. In computations, these fields are involved via a Lorentz force acting on the partial atomic charge in classical simulations or affecting the electronic density in quantum calculations, thus polarizing the affected structures [130]. While the structural field effects are usually studied by non-equilibrium MD techniques, NEGF can be utilized to capture electronic transport [231,232,233,234, 250]. Nevertheless, these approaches are computationally demanding and hardly applicable for large protein models, requiring more approximative treatments, as explained above.
The structure and functionalization of the electrode surfaces at nanoscales thus play a crucial role in protein electrochemistry. The electrode material, its surface facet orientation, and the surface dipole induced by reconstruction and termination processes determine the work function [251, 252] of the specific electrode and, as a result, the efficiency of the charge transfer mediated by the adsorbed biomolecules [182, 197, 237, 253]. Atomistic computer simulations can be employed to explore these effects and suggest key parameters for the desired adsorption and transport properties of specific systems. For example, suitable protein mutations with simultaneous surface functionalization by molecular linkers can be designed in silico to achieve the desired adsorption of enzymes onto biologically active surfaces [254, 255]. Furthermore, the incorporation of metallic nanoparticles at the surfaces has become popular in the last few decades due to their ability to induce locally enhanced fields. These techniques are often combined with plasma-polymeric surface coatings, ensuring good adsorption of both the nanoparticles as well as the proteins [256,257,258,259]. However, detailed knowledge of the atomistic details of these complex interfaces is required for further tuning and control of measurements and devices. Although such details are hard to obtain experimentally, they can be provided by computer simulations, which have proven to be useful tools for such applications.
Conclusions and further prospects
Investigation of the intrinsic electroactivity of proteins is based on the reduction or oxidation of individual aa residues in their structure. These redox-active aa residues can be located on the surface of proteins, where they are fully accessible to the electrode surface. If another substance interacts with the surface of the protein, these aa residues can be modified (covalent bond) or blocked (non-covalent association), and the exchange of electrons between the protein and the electrode surface cannot take place. This can be clearly observed electrochemically at the surfaces of both mercury and carbon electrodes. Metal electrodes (such as gold or mercury) can also be used to investigate the oxidation or chemical modification of Cys residues, which is crucial to the function and structure of a whole range of proteins or peptides [18, 25]. In addition, a selective electrochemical method for analyzing His residues has recently been developed and demonstrated on various model peptides and proteins [23]. For these purposes, a mercury electrode was used, which is a very effective tool not only for the sensitive analysis of proteins but also for the analysis of their interactions and structural changes, such as aggregation, folding, or oxidative damage. In this sense, the monitoring of protein interactions and structural changes is based on electrochemically active aa residues inside the protein structure. These aa residues can be exposed to the surface of the electrode due to a structural (relaxation) change, e.g., unfolding. Today, the discontinuation of mercury electrodes in electrochemical research is a global trend that we perceive very negatively (for more details, see the literature [17, 260, 261]) since liquid mercury electrodes with an atomically smooth surface are excellent for evaluating and characterizing the electrochemical behavior of various compounds. Today, the electrochemistry of proteins and peptides is also increasingly connected with research on membrane systems and research on membrane proteins and their interactions [11, 15]. One of the important prerequisites in this research field is the fact that membranes can be anchored to electrode surfaces. Also, very often, the components of the lipid bilayer are not electrochemically active and thus do not directly interfere with the analysis of proteins that are reconstituted in these membranes. In our opinion, studies of the effect of the electric field on the structure and intermolecular interactions of proteins with other molecules or conductive (or biomimetic) surfaces are interesting research areas. In this review, we demonstrated this on cytoskeletal proteins using chip technologies and computational methods [133, 143, 155, 156, 160]. Intense short electrical pulses can modulate the network of non-covalent interactions of proteins and their components and thus interfere with their self-assembly processes, which can be utilized in protein molecular manipulation approaches.
In addition, we also point out the importance of computer simulations of processes associated with the structure of peptides and proteins immobilized on an electrically charged surface [190, 209, 210, 236]. Simultaneously, the simulation of electron transfer significantly helps to understand the biological function of redox-active proteins. In addition to ET, electrochemical research is also interested in proton transfer in proteins and research on other proton-dependent processes [262,263,264]. It would be beneficial to combine approaches based on the analysis of intrinsic electroactivity with approaches targeting ET at non-protein redox-active centers of the metalloproteins [91]. Also, the infrequent application of advanced computing techniques prevents an expansion of the interpretive framework of experimental studies. As for protein labeling, electrochemiluminescence approaches are considered to have a lot of potential [12, 13]. In general, the combination of optical (spectral) detection methods (in situ spectroelectrochemistry [265]), microscopic techniques, and electrochemistry (including electrochemical impedance spectroscopy [266]) has considerable potential for the future.
In this review, we have shown selected applications in protein electroanalysis. However, it is important to not only describe the advantages but also take into account the experimental difficulties and obstacles that can limit the application of electrochemistry in the research of protein interactions, both protein-low-molecular-weight-ligand interactions and protein-protein-DNA or -membrane interactions. In this sense, it is very important to understand the importance of adsorption effects and “protein surface denaturation” phenomena [267], which can lead to artifacts in interaction studies. At the same time, it is important to pay close attention to the influence of the electric field on the native structure of proteins for the correct interpretation of electrochemical data [18]. The above could help to orient oneself in the above-mentioned areas and, at the same time, see all the possibilities that electrochemistry offers for protein research and bioanalysis.
Abbreviations
- 6-AQC:
-
6-aminoquinolyl-N-hydroxysuccinimidyl carbamate
- aa :
-
amino acid
- AdT:
-
adsorptive transfer
- AuNPs:
-
gold nanoparticles
- CHER:
-
catalytic hydrogen evolution reaction
- CPS:
-
chronopotentiometric stripping
- CV:
-
coefficient of variation
- DFT:
-
density functional theory
- DME:
-
dropping mercury electrode
- DNA:
-
deoxyribonucleic acid
- DST:
-
double-surface technique
- ECL:
-
electrochemiluminescence
- EF:
-
electric field
- ET:
-
electron transfer
- GGA:
-
generalized gradient approximation
- HMDE:
-
hanging mercury drop electrode
- HSA:
-
human serum albumin
- LOD:
-
limit of detection
- MBA:
-
4-mercaptophenylboronic acid
- MD:
-
molecular dynamics
- MGO:
-
methylglyoxal
- MIP:
-
molecularly imprinted polymer
- MT:
-
microtubule
- NDA:
-
naphatalene-2,3-dicarboxyaldehyde
- NEGF:
-
non-equilibrium Green’s function theory
- OPA:
-
o-phthaldialdehyde
- PEF:
-
pulsed electric field
- PMM:
-
perturbed matrix method
- PNP:
-
protein nanopore
- QM/MM:
-
quantum-mechanical/molecular mechanical
- RBL:
-
rat basophilic leukemia
- rHuEPO:
-
recombinant human erythropoietin
- RPE1:
-
retinal pigment cells
- SNP:
-
solid nanopore
- U2OS:
-
human osteosarcoma cells
References
Curry S (2015) Structural biology: a century-long journey into an unseen world. Interdiscip Sci Rev 40(3):308–328. https://doi.org/10.1179/0308018815z.000000000120
Koryta J (1990) The origin of polarography. J Electroanal Chem 296(2):293–297. https://doi.org/10.1016/0022-0728(90)87254-H
Heyrovsky M (2004) Early polarographic studies on proteins. Electroanalysis 16(13-14):1067–1073. https://doi.org/10.1002/elan.200403008
Zuman P, Palecek E (2005) Polarography of proteins: a history. In: Palecek E, Scheller F, Wang J (eds) Perspectives in Bioanalysis. Elsevier, 1, pp 755–771. https://doi.org/10.1016/S1871-0069(05)01020-7
Eddowes MJ, Hill HAO (1977) Novel method for the investigation of the electrochemistry of metalloproteins: cytochrome c. J Chem Soc, Chem Commun 21:771b–7712b. https://doi.org/10.1039/C3977000771B
Blanford CF (2013) The birth of protein electrochemistry. Chem Commun 49(95):11130–11132. https://doi.org/10.1039/C3CC46060F
Liu J, Chakraborty S, Hosseinzadeh P, Yu Y, Tian S, Petrik I et al (2014) Metalloproteins containing cytochrome, iron–sulfur, or copper redox centers. Chem Rev 114(8):4366–4469. https://doi.org/10.1021/cr400479b
Sarkar A, Chattopadhyay S, Mukherjee M, Ghosh Dey S, Dey A (2022) Assembly of redox active metallo-enzymes and metallo-peptides on electrodes: abiological constructs to probe natural processes. Curr Opin Chem Biol 68:102142. https://doi.org/10.1016/j.cbpa.2022.102142
Yates NDJ, Fascione MA, Parkin A (2018) Methodologies for “wiring” redox proteins/enzymes to electrode surfaces. Chem Eur J 24:12164–12182. https://doi.org/10.1002/chem.201800750
Laftsoglou T, Jeuken LJC (2017) Supramolecular electrode assemblies for bioelectrochemistry. Chem Commun 53(27):3801–3809. https://doi.org/10.1039/c7cc01154g
Vacek J, Zatloukalova M, Novak D (2018) Electrochemistry of membrane proteins and protein–lipid assemblies. Curr Opin Electrochem 12:73–80. https://doi.org/10.1016/j.coelec.2018.04.012
Vacek J, Hrbac J (2020) Sensors and microarrays in protein biomarker monitoring: an electrochemical perspective spots. Bioanalysis 12(18):1337–1345. https://doi.org/10.4155/bio-2020-0166
Van Gool A, Corrales F, Colovic M, Krstic D, Oliver-Martos B, Martínez-Cáceres E et al (2020) Analytical techniques for multiplex analysis of protein biomarkers. Exp Rev Proteom 17(4):257–273. https://doi.org/10.1080/14789450.2020.1763174
Blumberger J (2015) Recent advances in the theory and molecular simulation of biological electron transfer reactions. Chem Rev 115:11191–11238. https://doi.org/10.1021/acs.chemrev.5b00298
Sęk S, Vacek J, Dorcak V (2019) Electrochemistry of peptides. Curr Opin. Electrochem 14:166–172. https://doi.org/10.1016/j.coelec.2019.03.002
Suprun EV (2021) Direct electrochemistry of proteins and nucleic acids: the focus on 3D structure. Electrochem Commun 125:106983. https://doi.org/10.1016/j.elecom.2021.106983
Palecek E, Heyrovsky M, Dorcak V (2018) J. Heyrovský’s oscillographic polarography. Roots of present chronopotentiometric analysis of biomacromolecules. Electroanalysis 30(7):1259–1270. https://doi.org/10.1002/elan.201800109
Palecek E, Tkac J, Bartosik M, Bertok T, Ostatna V, Palecek J (2015) Electrochemistry of nonconjugated proteins and glycoproteins. Toward sensors for biomedicine and glycomics. Chem Rev 115(5):2045–2108. https://doi.org/10.1021/cr500279h
Li J, Hu R, Li X, Tong X, Yu D, Zhao Q (2017) Tiny protein detection using pressure through solid-state nanopores. Electrophoresis 38(8):1130–1138. https://doi.org/10.1002/elps.201600410
Liu W, Yang CN, Yang ZL, Yu RJ, Long YT, Ying YL (2023) Observing confined local oxygen-induced reversible thiol/disulfide cycle with a protein nanopore. Angew Chem Int Ed 62(27). https://doi.org/10.1002/anie.202304023
Luan B, Stolovitzky G, Martyna G (2012) Slowing and controlling the translocation of DNA in a solid-state nanopore. Nanoscale 4(4):1068–1077. https://doi.org/10.1039/c1nr11201e
Maglia G, Heron AJ, Stoddart D, Japrung D, Bayley H Analysis of single nucleic acid molecules with protein nanopores. Method Enzymol 4752010:591–623. https://doi.org/10.1016/S0076-6879(10)75022-9
Havran L, Vacek J, Dorcak V (2022) Free and bound histidine in reactions at mercury electrode. J Electroanal Chem 916:116336. https://doi.org/10.1016/j.jelechem.2022.116336
Sumitha MS, Xavier TS (2023) Recent advances in electrochemical biosensors – a brief review. Hybrid Adv 2:100023. https://doi.org/10.1016/j.hybadv.2023.100023
Palecek E, Ostatna V (2007) Electroactivity of nonconjugated proteins and peptides. Towards electroanalysis of all proteins. Electroanalysis 19:2383–2403. https://doi.org/10.1002/elan.200704033|ISSN
Mostafa AM, Barton SJ, Wren SP, Barker J (2021) Review on molecularly imprinted polymers with a focus on their application to the analysis of protein biomarkers. TrAC Trends Anal Chem 144:116431. https://doi.org/10.1016/j.trac.2021.116431
Palecek E, Dorcak V (2017) Label-free electrochemical analysis of biomacromolecules. Appl Mater Today 9:434–450. https://doi.org/10.1016/j.apmt.2017.08.011
Ramsden JJ (1994) Experimental methods for investigating protein adsorption kinetics at surfaces. Q Rev Biophys 27(1):41–105. https://doi.org/10.1017/S0033583500002900
Randviir EP, Banks CE (2022) A review of electrochemical impedance spectroscopy for bioanalytical sensors. Anal Methods 14(45):4602–4624. https://doi.org/10.1039/D2AY00970F
Arrigan DWM, Hackett MJ, Mancera RL (2018) Electrochemistry of proteins at the interface between two immiscible electrolyte solutions. Curr Opin Electrochem 12:27–32. https://doi.org/10.1016/j.coelec.2018.07.012
Suprun EV, Budnikov HC (2022) Bioelectrochemistry as a field of analysis: historical aspects and current status. J Anal Chem 77(6):643–663. https://doi.org/10.1134/S1061934822060168
Vargova V, Zivanovic M, Dorcak V, Palecek E, Ostatna V (2013) Catalysis of hydrogen evolution by polylysine, polyarginine and polyhistidine at mercury electrodes. Electroanalysis 25(9):2130–2135. https://doi.org/10.1002/elan.201300170
Zivanovic M, Aleksic M, Ostatna V, Doneux T, Palecek E (2010) Polylysine-catalyzed hydrogen evolution at mercury electrodes. Electroanalysis 22(17-18):2064–2071. https://doi.org/10.1002/elan.201000088
Dorcak V, Vargova V, Ostatna V, Palecek E (2015) Lysine, arginine, and histidine residues in peptide-catalyzed hydrogen evolution at mercury electrodes. Electroanalysis 27(4):910–916. https://doi.org/10.1002/elan.201400644
Doneux T, Ostatna V, Palecek E (2012) On the mechanism of hydrogen evolution catalysis by proteins: a case study with bovine serum albumin. Electrochim Acta 56(25):9337–9343. https://doi.org/10.1016/j.electacta.2011.08.017
Ostatna V, Dogan B, Uslu B, Ozkan S, Palecek E (2006) Native and denatured bovine serum albumin. D.c. polarography, stripping voltammetry and constant current chronopotentiometry. J Electroanal Chem 593:172–178. https://doi.org/10.1016/j.jelechem.2006.03.037
Cernocka H, Ostatna V, Palecek E (2015) Protein structural transition at negatively charged electrode surfaces. Effects of temperature and current density. Electrochim Acta 174:356–360. https://doi.org/10.1016/j.electacta.2015.06.009
Ostatna V, West RM (2020) Effects of ex situ chronopotentiometric analysis on stability of bovine serum albumin on mercury electrodes. J Electroanal Chem 860:113884. https://doi.org/10.1016/j.jelechem.2020.113884
Cernocka H, Ostatna V, Palecek E (2015) Fast-scan cyclic voltammetry with thiol-modified mercury electrodes distinguishes native from denatured BSA. Electrochem Commun 61:114–116. https://doi.org/10.1016/j.elecom.2015.10.017
Palecek E, Ostatna V (2009) Ionic strength-dependent structural transition of proteins at electrode surfaces. Chem Commun 13:1685–1687. https://doi.org/10.1039/B822274F
Ostatna V, Cernocka H, Palecek E (2010) Protein structure-sensitive electrocatalysis at DTT-modified electrodes. J Am Chem Soc 132(27):9408–9413. https://doi.org/10.1021/ja102427y
Ostatna V, Kuralay F, Trnkova L, Palecek E (2008) Constant current chronopotentiometry and voltammetry of native and denatured serum albumin at mercury and carbon electrodes. Electroanalysis 20:1406–1413. https://doi.org/10.1002/elan.200804206
Ostatna V, Palecek E (2008) Native, denatured and reduced BSA - enhancement of chronopotentiometric peak H by guanidinium chloride. Electrochim Acta 53(11):4014–4021. https://doi.org/10.1016/j.electacta.2007.10.035
Rimankova L, Cernocka H, Tihlarikova E, Nedela V, Ostatna V (2022) Chronopotentiometric sensing of native, oligomeric, denatured and aggregated serum albumin at charged surfaces. Bioelectrochemistry 145:108100. https://doi.org/10.1016/j.bioelechem.2022.108100
Palecek E, Ostatna V, Masarik M, Bertoncini CW, Jovin TM (2008) Changes in interfacial properties of alpha-synuclein preceding its aggregation. Analyst 133(1):76–84. https://doi.org/10.1039/b712812f
Ostatna V, Kasalova V, Kmetova K, Sedo O (2018) Changes of electrocatalytic response of bovine serum albumin after its methylation and acetylation. J Electroanal Chem 821:97–103. https://doi.org/10.1016/j.jelechem.2017.11.044
Izadi N, Cernocka H, Trefulka M, Ostatna V (2020) Influence of protein modification and glycosylation in the catalytic hydrogen evolution reaction of avidin and neutravidin: an electrochemical analysis. ChemPlusChem 85(6):1347–1353. https://doi.org/10.1002/cplu.202000298
Havlikova M, Zatloukalova M, Ulrichova J, Dobes P, Vacek J (2015) Electrocatalytic assay for monitoring methylglyoxal-mediated protein glycation. Anal Chem 87(3):1757–1763. https://doi.org/10.1021/ac503705d
Borsarelli CD, Falomir-Lockhart LJ, Ostatna V, Fauerbach JA, Hsiao HH, Urlaub H et al (2012) Biophysical properties and cellular toxicity of covalent crosslinked oligomers of alpha-synuclein formed by photoinduced side-chain tyrosyl radicals. Free Radic Biol Med 53(4):1004–1015. https://doi.org/10.1016/j.freeradbiomed.2012.06.035
Vargova V, Gimenez RE, Cernocka H, Trujillo DC, Tulli F, Zanini VIP et al (2016) Label-free electrochemical detection of singlet oxygen protein damage. Electrochim Acta 187:662–669. https://doi.org/10.1016/j.electacta.2015.11.104
Kasalova V, Hrstka R, Hernychova L, Coufalova D, Ostatna V (2017) Chronopotentiometric sensing of anterior gradient 2 protein. Electrochim Acta 240:250–257. https://doi.org/10.1016/j.electacta.2017.04.090
Palecek E, Ostatna V, Cernocka H, Joerger AC, Fersht AR (2011) Electrocatalytic monitoring of metal binding and mutation-induced conformational changes in p53 at picomole level. J Am Chem Soc 133(18):7190–7196. https://doi.org/10.1021/ja201006s
Cernocka H, Fojt L, Adamik M, Brazdova M, Palecek E, Ostatna V (2019) Interfacial properties of p53-DNA complexes containing various recognition elements. J Electroanal Chem 848:113300. https://doi.org/10.1016/j.jelechem.2019.113300
Palecek E, Cernocka H, Ostatna V, Navratilova L, Brazdova M (2014) Electrochemical sensing of tumor suppressor protein p53–deoxyribonucleic acid complex stability at an electrified interface. Anal Chim Acta 828:1–8. https://doi.org/10.1016/j.aca.2014.03.029
Ostatna V, Kasalova-Vargova V, Kekedy-Nagy L, Cernocka H, Ferapontova EE (2017) Chronopotentiometric sensing of specific interactions between lysozyme and the DNA aptamer. Bioelectrochemistry 114:42–47. https://doi.org/10.1016/j.bioelechem.2016.12.003
Ostatna V, Kasalova V, Sommerova L, Hrstka R (2018) Electrochemical sensing of interaction of anterior gradient-2 protein with peptides at a charged interface. Electrochim Acta 269:70–75. https://doi.org/10.1016/j.electacta.2018.02.152
Belicky S, Cernocka H, Bertok T, Holazova A, Reblova K, Palecek E et al (2017) Label-free chronopotentiometric glycoprofiling of prostate specific antigen using sialic acid recognizing lectins. Bioelectrochemistry 117:89–94. https://doi.org/10.1016/j.bioelechem.2017.06.005
Vargova V, Helma R, Palecek E, Ostatna V (2016) Electrochemical sensing of concanavalin A and ovalbumin interaction in solution. Anal Chim Acta 935:97–103. https://doi.org/10.1016/j.aca.2016.06.055
Cernocka H, Vonka P, Kasalova V, Sommerova L, Vandova V, Hrstka R et al (2021) AGR2-AGR3 hetero-oligomeric complexes: identification and characterization. Bioelectrochemistry 140:107808. https://doi.org/10.1016/j.bioelechem.2021.107808
Ostatna V, Hason S, Kasalova V, Durech M, Hrstka R (2019) Anterior gradient-3 protein-antibody interaction at charged interfaces. Label-free chronopotentiometric sensing. Electrochim Acta 297:974–979. https://doi.org/10.1016/j.electacta.2018.12.049
Novak D, Viskupicova J, Zatloukalova M, Heger V, Michalikova S, Majekova M et al (2018) Electrochemical behavior of sarco/endoplasmic reticulum Ca-ATPase in response to carbonylation processes. J Electroanal Chem 812:258–264. https://doi.org/10.1016/j.jelechem.2018.01.036
Svrckova M, Zatloukalova M, Dvorakova P, Coufalova D, Novak D, Hernychova L et al (2017) Na+/K+-ATPase interaction with methylglyoxal as reactive metabolic side product. Free Radic Biol Med 108:146–154. https://doi.org/10.1016/j.freeradbiomed.2017.03.024
Vacek J, Zatloukalova M, Geleticova J, Kubala M, Modriansky M, Fekete L et al (2016) Electrochemical platform for the detection of transmembrane proteins reconstituted into liposomes. Anal Chem 88(8):4548–4556. https://doi.org/10.1021/acs.analchem.6b00618
Vacek J, Zatloukalova M, Havlikova M, Ulrichova J, Kubala M (2013) Changes in the intrinsic electrocatalytic nature of Na+/K+ ATPase reflect structural changes on ATP-binding: electrochemical label-free approach. Electrochem Commun 27:104–107. https://doi.org/10.1016/j.elecom.2012.11.020
Zatloukalova M, Nazaruk E, Novak D, Vacek J, Bilewicz R (2018) Lipidic liquid crystalline cubic phases for preparation of ATP-hydrolysing enzyme electrodes. Biosens Bioelectron 100:437–444. https://doi.org/10.1016/j.bios.2017.09.036
Neumann E (1986) Chemical electric-field effects in biological macromolecules. Prog Biophys Mol Biol 47(3):197–231. https://doi.org/10.1016/0079-6107(86)90014-3
Vacek J, Svrckova M, Zatloukalova M, Novak D, Proskova J, Langova K et al (2018) Electrocatalytic artificial carbonylation assay for observation of human serum albumin inter-individual properties. Anal Biochem 550:137–143. https://doi.org/10.1016/j.ab.2018.04.025
Hernychova L, Alexandri E, Tzakos AG, Zatloukalova M, Primikyri A, Gerothanassis IP et al (2022) Serum albumin as a primary non-covalent binding protein for nitro-oleic acid. Int J Biol Macromol 203:116–129. https://doi.org/10.1016/j.ijbiomac.2022.01.050
Zatloukalova M, Mojovic M, Pavicevic A, Kabelac M, Freeman BA, Pekarova M et al (2019) Redox properties and human serum albumin binding of nitro-oleic acid. Redox Biol 24:101213. https://doi.org/10.1016/j.redox.2019.101213
Juskova P, Ostatna V, Palecek E, Foret F (2010) Fabrication and characterization of solid mercury amalgam electrodes for protein analysis. Anal Chem 82(7):2690–2695. https://doi.org/10.1021/ac902333s
Doneux T, Dorcak V, Palecek E (2010) Influence of the interfacial peptide organization on the catalysis of hydrogen evolution. Langmuir 26(2):1347–1353. https://doi.org/10.1021/la9024603
Dorcak V, Palecek E (2019) Catalytic deuterium evolution and H/D exchange in DNA. ChemElectroChem 6(4):1032–1039. https://doi.org/10.1002/celc.201801214
Ilimbi D, Buess-Herman C, Doneux T (2019) Chronopotentiometry as a sensitive interfacial characterisation tool for peptide aptamer monolayers. Electroanalysis 31(10):2041–2047. https://doi.org/10.1002/elan.201900285
Rimankova L, Hason S, Danhel A, Fojta M, Ostatna V (2020) Catalytic and redox activity of nucleic acids at mercury electrodes: roles of nucleobase residues. J Electroanal Chem 858:113812. https://doi.org/10.1016/j.jelechem.2019.113812
Brabec V, Vetterl V, Vrana O (1996) Electroanalysis of biomacromolecules. In: Brabec V, Walz D, Milazzo G (eds) Experimental Techniques in Bioelectrochemistry. Birghauser Verlag, Basel, p 287
Herzog G, Arrigan DW (2007) Electrochemical strategies for the label-free detection of amino acids, peptides and proteins. Analyst 132(7):615–632. https://doi.org/10.1039/B701472D
Baluchova S, Danhel A, Dejmkova H, Ostatna V, Fojta M, Schwarzova-Peckova K (2019) Recent progress in the applications of boron doped diamond electrodes in electroanalysis of organic compounds and biomolecules - a review. Anal Chim Acta 1077:30–66. https://doi.org/10.1016/j.aca.2019.05.041
Ostatna V, Cernocka H, Kurzatkowska K, Palecek E (2012) Native and denatured forms of proteins can be discriminated at edge plane carbon electrodes. Anal Chim Acta 735:31–36. https://doi.org/10.1016/j.aca.2012.05.012
Enache TA, Oliveira-Brett AM (2013) Peptide methionine sulfoxide reductase A (MsrA): direct electrochemical oxidation on carbon electrodes. Bioelectrochemistry 89:11–18. https://doi.org/10.1016/j.bioelechem.2012.08.004
Ostatna V, Vargova V, Hrstka R, Durech M, Vojtesek B, Palecek E (2014) Effect of His6-tagging of anterior gradient 2 protein on its electro-oxidation. Electrochim Acta 150:218–222. https://doi.org/10.1016/j.electacta.2014.10.125
Cai XH, Rivas G, Farias PAM, Shiraishi H, Wang J, Palecek E (1996) Potentiometric stripping analysis of bioactive peptides at carbon electrodes down to subnanomolar concentrations. Anal Chim Acta 332(1):49–57. https://doi.org/10.1016/0003-2670(96)00189-4
Hughes ZE, Walsh TR (2015) What makes a good graphene-binding peptide? Adsorption of amino acids and peptides at aqueous graphene interfaces. J Mater Chem B 3(16):3211–3221. https://doi.org/10.1039/c5tb00004a
Oliveira SCB, Santarino IB, Oliveira-Brett AM (2013) Direct electrochemistry of native and denatured anticancer antibody rituximab at a glassy carbon electrode. Electroanalysis 25(4):1029–1034. https://doi.org/10.1002/elan.201200552
Fernandes IPG, Oliveira-Brett AM (2017) Calcium-induced calmodulin conformational change. Electrochemical evaluation. Bioelectrochemistry 113:69–78. https://doi.org/10.1016/j.bioelechem.2016.10.002
Topal BD, Özkan SA, Uslu B (2014) Direct electrochemistry of native and denatured alpha-2-Macroglobulin by solid electrodes. J Electroanal Chem 719:14–18. https://doi.org/10.1016/j.jelechem.2014.02.008
Lopes P, Xu M, Zhang M, Zhou T, Yang YL, Wang C et al (2014) Direct electrochemical and AFM detection of amyloid-beta peptide aggregation on basal plane HOPG. Nanoscale 6(14):7853–7857. https://doi.org/10.1039/c4nr02413c
Suprun EV, Khmeleva SA, Radko SP, Archakov AI, Shumyantseva VV (2016) Electrochemical analysis of amyloid-beta domain 1-16 isoforms and their complexes with Zn(II) ions. Electrochim Acta 187:677–683. https://doi.org/10.1016/j.electacta.2015.11.111
Vestergaard M, Kerman K, Saito M, Nagatani N, Takamura Y, Tamiya E (2005) A rapid label-free electrochemical detection and kinetic study of Alzheimer’s amyloid beta aggregation. J Am Chem Soc 127(34):11892–11893. https://doi.org/10.1021/ja052522q
Kerman K, Vestergaard M, Chikae M, Yamamura S, Tamiya E (2007) Label-free electrochemical detection of the phosphorylated and non-phosphorylated forms of peptides based on tyrosine oxidation. Electrochem Commun 9(5):976–980. https://doi.org/10.1016/j.elecom.2006.11.033
Suprun EV, Zharkova MS, Morozevich GE, Veselovsky AV, Shumyantseva VV, Archakov AI (2013) Analysis of redox activity of proteins on the carbon screen printed electrodes. Electroanalysis 25(9):2109–2116. https://doi.org/10.1002/elan.201300248
Novak D, Mojovic M, Pavicevic A, Zatloukalova M, Hernychova L, Bartosik M et al (2018) Electrochemistry and electron paramagnetic resonance spectroscopy of cytochrome c and its heme-disrupted analogs. Bioelectrochemistry 119:136–141. https://doi.org/10.1016/j.bioelechem.2017.09.011
Alvarez-Dorta D, Thobie-Gautier C, Croyal M, Bouzelha M, Mével M, Deniaud D et al (2018) Electrochemically promoted tyrosine-click-chemistry for protein labeling. J Am Chem Soc 140(49):17120–17126. https://doi.org/10.1021/jacs.8b09372
Willwacher J, Raj R, Mohammed S, Davis BG (2016) Selective metal-site-guided arylation of proteins. J Am Chem Soc 138(28):8678–8681. https://doi.org/10.1021/jacs.6b04043
Song C, Liu K, Wang Z, Ding B, Wang S, Weng Y et al (2019) Electrochemical oxidation induced selective tyrosine bioconjugation for the modification of biomolecules. Chem Sci 10(34):7982–7987. https://doi.org/10.1039/C9SC02218J
Sierra T, Crevillen AG, Escarpa A (2017) Derivatization agents for electrochemical detection in amino acid, peptide and protein separations: the hidden electrochemistry? Electrophoresis 38(21):2695–2703. https://doi.org/10.1002/elps.201700167
Li G-D, Krull I, Cohen S (1996) Electrochemical activity of 6-aminoquinolyl urea derivatives of amino acids and peptides. Application to high-performance liquid chromatography with electrochemical detection. J Chromatogr A 724(1-2):147–157. https://doi.org/10.1016/0021-9673(95)00941-8
Pappa-Louisi A, Nikitas P, Agrafiotou P, Papageorgiou A (2007) Optimization of separation and detection of 6-aminoquinolyl derivatives of amino acids by using reversed-phase liquid chromatography with on line UV, fluorescence and electrochemical detection. Anal Chim Acta 593(1):92–97. https://doi.org/10.1016/j.aca.2007.04.044
Rose MJ, Lunte SM, Carlson RG, Stobaugh JF (1999) Hydroquinone-based derivatization reagents for the quantitation of amines using electrochemical detection. Anal Chem 71(11):2221–2230. https://doi.org/10.1021/ac981236c
Rose MJ, Lunte SM, Carlson RG, Stobaugh JF (2003) Amino acid and peptide analysis using derivatization with p-nitrophenol-2, 5-dihydroxyphenylacetate bis-tetrahydropyranyl ether and capillary electrophoresis with electrochemical detection. J Pharm Biomed Anal 30(6):1851–1859. https://doi.org/10.1016/s0731-7085(02)00528-9
Toyama E, Maruyama K, Sugai T, Kondo M, Masaoka S, Saitoh T et al (2019) Electrochemical tryptophan-selective bioconjugation. https://doi.org/10.26434/chemrxiv7795484
Li M, Zhu W, Marken F, James TD (2015) Electrochemical sensing using boronic acids. Chem Commun 51(78):14562–14573. https://doi.org/10.1039/C5CC04976H
Chien HC, Chou TC (2011) A nonenzymatic amperometric method for fructosyl-valine sensing using ferroceneboronic acid. Electroanalysis 23(2):402–408. https://doi.org/10.1002/elan.201000426
Xia N, Deng D, Zhang L, Yuan B, Jing M, Du J et al (2013) Sandwich-type electrochemical biosensor for glycoproteins detection based on dual-amplification of boronic acid-gold nanoparticles and dopamine-gold nanoparticles. Biosens Bioelectron 43:155–159. https://doi.org/10.1016/j.bios.2012.12.020
Billova S, Kizek R, Palecek E (2002) Differential pulse adsorptive stripping voltammetry of osmium-modified peptides. Bioelectrochemistry 56(1-2):63–66. https://doi.org/10.1016/S1567-5394(02)00008-7
Fojta M, Billova S, Havran L, Pivonkova H, Cernocka H, Horakova P et al (2008) Osmium tetroxide, 2, 2′-bipyridine: electroactive marker for probing accessibility of tryptophan residues in proteins. Anal Chem 80(12):4598–4605. https://doi.org/10.1021/ac800527u
Trefulka M, Dorcak V, Krenkova J, Foret F, Palecek E (2017) Electrochemical analysis of Os(VI)-modified glycoproteins and label-free glycoprotein detection eluted from lectin capillary column. Electrochim Acta 239:10–15. https://doi.org/10.1016/j.electacta.2017.04.045
Trefulka M, Palecek E (2014) Direct chemical modification and voltammetric detection of glycans in glycoproteins. Electrochem Commun 48:52–55. https://doi.org/10.1016/j.elecom.2014.08.011
Martic S, Labib M, Kraatz H-B (2011) Enzymatically modified peptide surfaces: towards general electrochemical sensor platform for protein kinase catalyzed phosphorylations. Analyst 136(1):107–112. https://doi.org/10.1039/C0AN00438C
Ramya M, Senthil Kumar P, Rangasamy G, Umashankar V, Rajesh G, Nirmala K et al (2022) A recent advancement on the applications of nanomaterials in electrochemical sensors and biosensors. Chemosphere 308: 136416. https://doi.org/10.1016/j.chemosphere.2022.136416
Mishra A, Bhatt R, Bajpai J, Bajpai AK (2021) Nanomaterials based biofuel cells: a review. Int J Hydrogen Energy 46(36):19085–19105. https://doi.org/10.1016/j.ijhydene.2021.03.024
Smutok O, Katz E (2023) Electroanalytical instrumentation—how it all started: history of electrochemical instrumentation. J Solid State Electrochem. https://doi.org/10.1007/s10008-023-05375-3
Palecek E, Fojta M (2007) Magnetic beads as versatile tools for electrochemical DNA and protein biosensing. Talanta 74(3):276–290. https://doi.org/10.1016/j.talanta.2007.08.020
Palecek E, Fojta M, Jelen F (2002) New approaches in the development of DNA sensors: hybridization and electrochemical detection of DNA and RNA at two different surfaces. Bioelectrochemistry 56:85–90. https://doi.org/10.1016/S1567-5394(02)00025-7
Palecek E, Postbieglova I (1986) Adsorptive stripping voltammetry of biomacromolecules with transfer of the adsorbed layer. J Electroanal Chem 214(1-2):359–371. https://doi.org/10.1016/0022-0728(86)80108-5
Palecek E, Kizek R, Havran L, Billova S, Fojta M (2002) Electrochemical enzyme-linked immunoassay in a DNA hybridization sensor. Anal Chim Acta 469(1):73–83. https://doi.org/10.1016/S0003-2670(01)01605-1
Wang J, Xu D, Erdem A, Polsky R, Salazar MA (2002) Genomagnetic electrochemical assays of DNA hybridization. Talanta 56(5):931–938. https://doi.org/10.1016/S0039-9140(01)00653-1
Vacek J, Mozga T, Cahova K, Pivonkova H, Fojta M (2007) Electrochemical sensing of chromium-induced DNA damage: DNA strand breakage by intermediates of chromium(VI) electrochemical reduction. Electroanalysis 19(19-20):2093–2102. https://doi.org/10.1002/elan.200703917
Masarik M, Cahova K, Kizek R, Palecek E, Fojta M (2007) Label-free voltammetric detection of single-nucleotide mismatches recognized by the protein MutS. Anal Bioanal Chem 388(1):259–270. https://doi.org/10.1007/s00216-007-1181-7
Palecek E, Masarik M, Kizek R, Kuhlmeier D, Hassmann J, Schulein J (2004) Sensitive electrochemical determination of unlabeled mutS protein and detection of point mutations in DNA. Anal Chem 76(19):5930–5936. https://doi.org/10.1021/ac049474x
Kawde AN, Rodriguez MC, Lee TMH, Wang J (2005) Label-free bioelectronic detection of aptamer-protein interactions. Electrochem Commun 7(5):537–540. https://doi.org/10.1016/j.elecom.2005.03.008
Matthew JB (1985) Electrostatic effects in proteins. Ann Rev Biophys Biophys Chem 14(1):387–417. https://doi.org/10.1146/annurev.bb.14.060185.002131
Park JW, Rhee YM (2016) Electric field keeps chromophore planar and produces high yield fluorescence in green fluorescent protein. J Am Chem Soc 138(41):13619–13629. https://doi.org/10.1021/jacs.6b06833
Henderson D, Boda D (2009) Insights from theory and simulation on the electrical double layer. Phys Chem Chem Phys 11(20):3822–3830. https://doi.org/10.1039/B815946G
Schönknecht T, Pörschke D (1996) Electrooptical analysis of α-chymotrypsin at physiological salt concentration. Biophys Chem 58(1):21–28. https://doi.org/10.1016/0301-4622(95)00082-8
Sinelnikova A, Mandl T, Agelii H, Grånäs O, Marklund EG, Caleman C et al (2021) Protein orientation in time-dependent electric fields: orientation before destruction. Biophys J 120(17):3709–3717. https://doi.org/10.1016/j.bpj.2021.07.017
Hekstra DR, White KI, Socolich MA, Henning RW, Srajer V, Ranganathan R (2016) Electric-field-stimulated protein mechanics. Nature 540(7633):400–405. https://doi.org/10.1038/nature20571
Fernandez-Diaz MD, Barsotti L, Dumay E, Cheftel JC (2000) Effects of pulsed electric fields on ovalbumin solutions and dialyzed egg white. J Agric Food Chem 48(6):2332–2339. https://doi.org/10.1021/jf9908796
Liu Y-Y, Zhang Y, Zeng X-A, El-Mashad H, Pan Z-L, Wang Q-J (2014) Effect of pulsed electric field on microstructure of some amino acid group of soy protein isolates. Int J Food Eng 10(1):113–120. https://doi.org/10.1515/ijfe-2013-0033
Wu L, Zhao W, Yang R, Chen X (2014) Effects of pulsed electric fields processing on stability of egg white proteins. J Food Eng 139:13–18. https://doi.org/10.1016/j.jfoodeng.2014.04.008
English NJ, Waldron CJ (2015) Perspectives on external electric fields in molecular simulation: progress, prospects and challenges. Phys Chem Chem Phys 17(19):12407–12440. https://doi.org/10.1039/C5CP00629E
Noble BB, Todorova N, Yarovsky I (2022) Electromagnetic bioeffects: a multiscale molecular simulation perspective. Phys Chem Chem Phys 24(11):6327–6348. https://doi.org/10.1039/D1CP05510K
Alizadeh H, Davoodi J, Rafii-Tabar H (2017) Deconstruction of the human connexin 26 hemichannel due to an applied electric field; a molecular dynamics simulation study. J Mol Graph Model 73:108–114. https://doi.org/10.1016/j.jmgm.2017.02.006
Marracino P, Havelka D, Prusa J, Liberti M, Tuszynski J, Ayoub AT et al (2019) Tubulin response to intense nanosecond-scale electric field in molecular dynamics simulation. Sci Rep 9(1):10477. https://doi.org/10.1038/s41598-019-46636-4
Wang J, Vanga SK, Raghavan V (2020) Structural responses of kiwifruit allergen Act d 2 to thermal and electric field stresses based on molecular dynamics simulations and experiments. Food Funct 11(2):1373–1384. https://doi.org/10.1039/C9FO02427A
Lugli F, Toschi F, Biscarini F, Zerbetto F (2010) Electric field effects on short fibrils of Aβ amyloid peptides. J Chem Theory Comput 6(11):3516–3526. https://doi.org/10.1021/ct1001335
Singh A, Orsat V, Raghavan V (2013) Soybean hydrophobic protein response to external electric field: a molecular modeling approach. Biomolecules 3(1):168–179. https://doi.org/10.3390/biom3010168
Todorova N, Bentvelzen A, Yarovsky I (2020) Electromagnetic field modulates aggregation propensity of amyloid peptides. J Chem Phys 152(3):035104. https://doi.org/10.1063/1.5126367
Astrakas L, Gousias C, Tzaphlidou M (2011) Electric field effects on chignolin conformation. J Appl Phys 109(9):094702. https://doi.org/10.1063/1.3585867
Prusa J, Cifra M (2019) Molecular dynamics simulation of the nanosecond pulsed electric field effect on kinesin nanomotor. Sci Rep 9(1):19721. https://doi.org/10.1038/s41598-019-56052-3
Marracino P, Apollonio F, Liberti M, d’Inzeo G, Amadei A (2013) Effect of high exogenous electric pulses on protein conformation: myoglobin as a case study. J Phys Chem B 117(8):2273–2279. https://doi.org/10.1021/jp309857b
Sinelnikova A, Mandl T, Östlin C, Grånäs O, Brodmerkel MN, Marklund EG et al (2021) Reproducibility in the unfolding process of protein induced by an external electric field. Chem Sci 12(6):2030–2038. https://doi.org/10.1039/D0SC06008A
Baumketner A (2014) Electric field as a disaggregating agent for amyloid fibrils. J Phys Chem B 118(50):14578–14589. https://doi.org/10.1021/jp509213f
Prusa J, Ayoub AT, Chafai DE, Havelka D, Cifra M (2021) Electro-opening of a microtubule lattice in silico. Comput Struct Biotechnol J 19:1488–1496. https://doi.org/10.1016/j.csbj.2021.02.007
English NJ, Mooney DA (2007) Denaturation of hen egg white lysozyme in electromagnetic fields: a molecular dynamics study. J Chem Phys 126(9). https://doi.org/10.1063/1.2515315
English NJ, Solomentsev GY, O'Brien P (2009) Nonequilibrium molecular dynamics study of electric and low-frequency microwave fields on hen egg white lysozyme. J Chem Phys 131(3):035106. https://doi.org/10.1063/1.3184794
Toschi F, Lugli F, Biscarini F, Zerbetto F (2009) Effects of electric field stress on a β-amyloid peptide. J Phys Chem B 113(1):369–376. https://doi.org/10.1021/jp807896g
Amadei A, Marracino P (2015) Theoretical-computational modelling of the electric field effects on protein unfolding thermodynamics. RSC Adv 5(117):96551–96561. https://doi.org/10.1039/c5ra15605j
Marracino P, Paffi A, d’Inzeo G (2022) A rationale for non-linear responses to strong electric fields in molecular dynamics simulations. Phys Chem Chem Phys 24(19):11654–11661. https://doi.org/10.1039/d1cp04466d
Marklund EG, Ekeberg T, Moog M, Benesch JLP, Caleman C (2017) Controlling protein orientation in vacuum using electric fields. J Phys Chem Lett 8(18):4540–4544. https://doi.org/10.1021/acs.jpclett.7b02005
Budi A, Legge FS, Treutlein H, Yarovsky I (2005) Electric field effects on insulin chain-B conformation. J Phys Chem B 109(47):22641–22648. https://doi.org/10.1021/jp052742q
della Valle E, Marracino P, Pakhomova O, Liberti M, Apollonio F (2019) Nanosecond pulsed electric signals can affect electrostatic environment of proteins below the threshold of conformational effects: the case study of SOD1 with a molecular simulation study. PLoS ONE 14(8):e0221685. https://doi.org/10.1371/journal.pone.0221685
Wang R, Wen QH, Zeng XA, Lin JW, Li J, Xu FY (2022) Binding affinity of curcumin to bovine serum albumin enhanced by pulsed electric field pretreatment. Food Chem 377:131945. https://doi.org/10.1016/j.foodchem.2021.131945
Sun WW, Yu SJ, Zeng XA, Yang XQ, Jia X (2011) Properties of whey protein isolate-dextran conjugate prepared using pulsed electric field. Food Res Int 44(4):1052–1058. https://doi.org/10.1016/j.foodres.2011.03.020
Zhao W, Yang R (2010) Experimental study on conformational changes of lysozyme in solution induced by pulsed electric field and thermal stresses. J Phys Chem B 114(1):503–510. https://doi.org/10.1021/jp9081189
Havelka D, Zhernov I, Teplan M, Lansky Z, Chafai DE, Cifra M (2022) Lab-on-chip microscope platform for electro-manipulation of a dense microtubules network. Sci Rep 12(1):2462. https://doi.org/10.1038/s41598-022-06255-y
Havelka D, Chafai DE, Krivosudsky O, Klebanovych A, Vostarek F, Kubinova L et al (2020) Nanosecond pulsed electric field lab-on-chip integrated in super-resolution microscope for cytoskeleton imaging. Adv Mater Technol 5(3):1900669. https://doi.org/10.1002/admt.201900669
Casciola M, Liberti M, Denzi A, Paffi A, Merla C, Apollonio F (2017) A computational design of a versatile microchamber for in vitro nanosecond pulsed electric fields experiments. Integration 58:446–453. https://doi.org/10.1016/j.vlsi.2017.03.005
Dalmay C, Villemejane J, Joubert V, Silve A, Arnaud-Cormos D, Français O et al (2011) A microfluidic biochip for the nanoporation of living cells. Biosens Bioelectron 26(12):4649–4655. https://doi.org/10.1016/j.bios.2011.03.020
Merla C, Liberti M, Marracino P, Muscat A, Azan A, Apollonio F et al (2018) A wide-band bio-chip for real-time optical detection of bioelectromagnetic interactions with cells. Sci Rep 8(1):5044. https://doi.org/10.1038/s41598-018-23301-w
Chafai DE, Vostarek F, Draberova E, Havelka D, Arnaud-Cormos D, Leveque P et al (2020) Microtubule cytoskeleton remodeling by nanosecond pulsed electric fields. Adv Biosyst 4(7):e2000070. https://doi.org/10.1002/adbi.202000070
Graybill PM, Davalos RV (2020) Cytoskeletal disruption after electroporation and its significance to pulsed electric field therapies. Cancers (Basel) 12(5):1132. https://doi.org/10.3390/cancers12051132
Dimova R, Riske KA, Aranda S, Bezlyepkina N, Knorr RL, Lipowsky R (2007) Giant vesicles in electric fields. Soft Matter 3(7):817–827. https://doi.org/10.1039/B703580B
Perrier DL, Vahid A, Kathavi V, Stam L, Rems L, Mulla Y et al (2019) Response of an actin network in vesicles under electric pulses. Sci Rep 9(1):8151. https://doi.org/10.1038/s41598-019-44613-5
Ho SY, Mittal GS, Cross JD (1997) Effects of high field electric pulses on the activity of selected enzymes. J Food Eng 31(1):69–84. https://doi.org/10.1016/S0260-8774(96)00052-0
Jin W, Wang Z, Peng D, Shen W, Zhu Z, Cheng S et al (2020) Effect of pulsed electric field on assembly structure of α-amylase and pectin electrostatic complexes. Food Hydrocolloids 101:105547. https://doi.org/10.1016/j.foodhyd.2019.105547
Rodrigues RM, Avelar Z, Machado L, Pereira RN, Vicente AA (2020) Electric field effects on proteins - novel perspectives on food and potential health implications. Food Res Int 137:109709. https://doi.org/10.1016/j.foodres.2020.109709
Armstrong FA (2002) Insight from protein film voltammetry into mechanism of complex biological electron-transfer reactions. Dalton Trans:661-671. https://doi.org/10.1039/B108359G
del Barrio M, Fourmond V (2019) Redox (in)activations of metalloenzymes: a protein film voltammetry approach. ChemElectroChem 6:4949–4962. https://doi.org/10.1002/celc.201901028
Gulaboski R, Lovric M, Mirceski V, Bogeski I, Hoth M (2008) Protein-film voltammetry: a theoretical study of the temperature effect using square-wave voltammetry. Biophys Chem 137:49–55. https://doi.org/10.1016/j.bpc.2008.06.011
Meyer T, Melin F, Xie H, von der Hocht I, Choi SK, Noor MR et al (2014) Evidence for distinct electron transfer processes in terminal oxidases from different origin by means of protein film voltammetry. J Am Chem Soc 136:10854–10857. https://doi.org/10.1021/ja505126v
Bostick CD, Mukhopadhyay S, Pecht I, Sheves M, Cahen D, Lederman D (2018) Protein bioelectronics: a review of what we do and do not know. Rep Prog Phys 81:26601. https://doi.org/10.1088/1361-6633/aa85f2
Leger C, Bertrand P (2008) Direct electrochemistry of redox enzymes as a tool for mechanistic studies. Chem Rev 108:2379–2438. https://doi.org/10.1021/cr0680742
Gorton L, Lindgren A, Larsson T, Munteanu FD, Ruzgas T, Gazaryan I (1999) Direct electron transfer between heme-containing enzymes and electrodes as basis for third generation biosensors. Anal Chim Acta 400:91–108. https://doi.org/10.1016/S0003-2670(99)00610-8
Cho I-H, Kim DH, Park S (2020) Electrochemical biosensors: perspective on functional nanomaterials for on-site analysis. Biomater Res 24:6. https://doi.org/10.1186/s40824-019-0181-y
Kornienko N, Ly KH, Robinson WE, Heidary N, Zhang JZ, Reisner E (2019) Advancing techniques for investigating the enzyme-electrode interface. Acc Chem Res 52:1439–1448. https://doi.org/10.1021/acs.accounts.9b00087
Szczesny J, Markovic N, Conzuelo F, Zacarias S, Pereira IAC, Lubitz W et al (2018) A gas breathing hydrogen/air biofuell cell comprising a redox polymer/hydrogenase-based bionanode. Nat Commun 9:4715. https://doi.org/10.1038/s41467-018-07137-6
Wong TS, Schwaneberg U (2003) Protein engineering in bioelectrocatalysis. Curr Opin Biotechnol 14:590–596. https://doi.org/10.1016/j.copbio.2003.09.008
Ha TQ, Planje IJ, White JRG, Aragones AC, Diez-Perez I (2021) Charge transport at the protein-electrode interface in the emerging field of biomolecular electronics. Curr Opin Electrochem 28:100734. https://doi.org/10.1016/j.coelec.2021.100734
Kayser B, Fereiro JA, Guo C, Cohen SR, Sheves M, Pecht I et al (2018) Transistor configuration yields energy level control in protein-based junctions. Nanoscale 10:21712–21720. https://doi.org/10.1039/C8NR06627B
Lee T, Kim S, Kim J, Park S-C, Yoon J, Park C et al (2020) Recent advances in biomolecule-nanomaterial heterolayer-based charge storage devices for bioelectronic applications. Materials 13:3520. https://doi.org/10.3390/ma13163520
Zhang L, Lu JR, Waigh TA (2021) Electronics of peptide- and protein-based biomaterials. Adv Colloid Interface Sci 287:102319. https://doi.org/10.1016/j.cis.2020.102319
Cahen D, Pecht I, Sheves M (2021) What can we learn from protein-based electron transport junctions? J Phys Chem Lett 12:11598–11603. https://doi.org/10.1021/acs.jpclett.1c02446
Kumar KS, Pasula RR, Lim S, Nijhuis CA (2016) Long-range tunneling processes across ferritin-based junctions. Adv Mater 28:1824–1830. https://doi.org/10.1002/adma.201504402
Marcus RA (1956) On the theory of oxidation-reduction reactions involving electron transfer. I. J Chem Phys 24(5):966–978. https://doi.org/10.1063/1.1742723
Marcus RA (1956) Electrostatic free energy and other properties of states having nonequilibrium polarization. I. J Chem Phys 24(5):979–989. https://doi.org/10.1063/1.1742724
Artes JM, Diez-Perez I, Sanz F, Gorostiza P (2011) Direct measurement of electron transfer distance decay constants of single redox proteins by electrochemical tunneling spectroscopy. ACS Nano 5:2060–2066. https://doi.org/10.1021/nn103236e
Elliott M, Jones DD (2018) Approaches to single-molecule studies of metalloprotein electron transfer using scanning probe-based techniques. Biochem Soc Trans 46:1–9. https://doi.org/10.1042/BST20170229
Nazmutdinov RR, Zinkicheva TT, Shermukhamedov SA, Zhang J, Ulstrup J (2018) Electrochemistry of single molecules and biomolecules, molecular scale nanostructures, and low-dimensional systems. Curr Opin Electrochem 7:179–187. https://doi.org/10.1016/j.coelec.2017.11.013
Salvatore P, Zeng D, Karlsen KK, Chi Q, Wengel J, Ulstrup J (2013) Electrochemistry of single metalloprotein and DNA-based molecules at Au(111) electrode surfaces. ChemPhysChem 14:2101–2111. https://doi.org/10.1002/cphc.201300299
Garg K, Ghosh M, Eliash T, van Wonderen JH, Butt JN, Shi L et al (2018) Direct evidence for heme-assisted solid-state electronic conduction in multi-heme c-type cytochromes. Chem Sci 9:7304–7310. https://doi.org/10.1039/C8SC01716F
Agam Y, Nandi R, Kaushansky A, Peskin U, Amdursky N (2020) The porphyrin ring rather than the metal ion dictates long-range electron transport across proteins suggesting coherence-assisted mechanism. Proc Nat Acad Sci USA 117:32260–32266. https://doi.org/10.1073/pnas.2008741117
Zhang B, Song W, Brown J, Nemanich R, Lindsay S (2020) Electronic conductance resonance in non-redox-active proteins. J Am Chem Soc 142:6432–6438. https://doi.org/10.1021/jacs.0c01805
Hitaishi VP, Clement R, Bourassin N, Baaden M, de Poulpiquet A, Sacquin-Mora S et al (2018) Controlling redox enzyme orientation at planar electrodes. Catal 8:192. https://doi.org/10.3390/catal8050192
Mazurenko I, Hitaischi VP, Lojou E (2020) Recent advances in surface chemistry of electrodes to promote direct enzymatic bioelectrocatalysis. Curr Opin Electrochem 19:113–121. https://doi.org/10.1016/j.coelec.2019.11.004
Biriukov D, Futera Z (2021) Adsorption of amino acids at the gold/aqueous interface: effect of an external electric field. J Phys Chem C 125:7856–7867. https://doi.org/10.1021/acs.jpcc.0c11248
Feng J, Slocik JM, Sarikaya M, Naik RR, Farmer BL, Heinz H (2012) Influence of the shape of nanostructured metal surfaces on adsorption of single peptide molecules in aqueous solution. Small 8(7):1049–1059. https://doi.org/10.1002/smll.201102066
Futera Z (2021) Amino-acid interactions with the Au(111) surface: adsorption, band alignment, and interfacial electronic coupling. Phys Chem Chem Phys 23:10257–10266. https://doi.org/10.1039/D1CP00218J
Futera Z, Blumberger J (2019) Adsorption of amino acids on gold: assessing the accuracy of the GolP-CHARMM force field and parametrization of Au-S bonds. J Chem Theory Comput 15:613–624. https://doi.org/10.1021/acs.jctc.8b00992
Hoefling M, Iori F, Corni S, Gottschalk K-E (2010) Interaction of amino acids with the Au(111) surface: adsorption free energies from molecular dynamics simulations. Langmuir 26(11):8347–8351. https://doi.org/10.1021/la904765u
Iori F, Di Felice R, Molinari E, Corni S (2009) GoIP: an atomistic force-field to describe the interaction of proteins with Au(111) surfaces in water. J Comput Chem 30:1465–1476. https://doi.org/10.1002/jcc.21165
Wright LB, Rodger PM, Corni S, Walsh TR (2013) GoIP-CHARMM: first-principles based force fields for the interaction of proteins with Au(111) and Au(100). J Chem Theory Comput 9:1616–1630. https://doi.org/10.1021/ct301018m
Wright LB, Rodger PM, Walsh TR, Corni S (2013) First-principle-based force field for the interaction of proteins with Au(100)(5x1): an extension of GolP-CHARMM. J Phys Chem C 117:24292–24306. https://doi.org/10.1021/jp4061329
Brusatori MA, Tie Y, Van Tassel PR (2003) Protein adsorption kinetics under an applied electric field: an optical waveguide lightmode spectroscopy study. Langmuir 19:5089–5097. https://doi.org/10.1021/la0269558
Mulheran PA, Connell DJ, Kubiak-Ossowska K (2016) Steering protein adsorption at charged surfaces: electric fields and ionic screening. RSC Adv 6:73709–73716. https://doi.org/10.1039/C6RA16391B
Xie Y, Liao C, Zhou J (2013) Effects of external electric fields on lysozyme adsorption by molecular dynamics simulations. Biophys Chem 179:26–34. https://doi.org/10.1016/j.bpc.2013.05.002
Amadei A, Daidone I, Bortolotti CA (2013) A general statistical mechanical approach for modeling redox thermodynamics: the reaction and reorganization free energies. RSC Adv 3:19657–19665. https://doi.org/10.1039/C3RA42842G
Blumberger J (2008) Free energies for biological electron transfer from QM/MM calculation: method, application and critical assessment. Phys Chem Chem Phys 10:5651–5667. https://doi.org/10.1039/B807444E
Daidone I, Amadei A, Zaccanti F, Borsari M, Bortolotti CA (2014) How the reorganization free energy affects the reduction potential of structurally homologous cytochromes. J Phys Chem Lett 5:1534–1540. https://doi.org/10.1021/jz5005208
Jiang X, Futera Z, Blumberger J (2019) Ergodicity-breaking in thermal biological electron transfer? Cytochrome C Revisited. J Phys Chem B 123:7588–7598. https://doi.org/10.1021/acs.jpcb.9b05253
Kontkanen OV, Biriukov D, Futera Z (2022) Reorganization free energy of copper proteins in solution, in vacuum, and on metal surfaces. J Chem Phys 156:175101. https://doi.org/10.1063/5.0085141
Tipmanee V, Oberhofer H, Park M, Kim KS, Blumberger J (2010) Prediction of reorganization free energies for biological electron transfer: a comparative study of ru-modified cytochromes and a 4-helix bundle protein. J Am Chem Soc 132:17032–17040. https://doi.org/10.1021/ja107876p
Cave RJ, Newton MD (1996) Generalization of the Mulliken-Hush treatment for the calculation of electron transfer matrix elements. Chem Phys Lett 249:15–19. https://doi.org/10.1016/0009-2614(95)01310-5
Cave RJ, Newton MD (1997) Calculation of electronic coupling matrix elements for ground and excited state electron transfer reactions: comparison of the generalized Mulliken-Hush and block diagonalization methods. J Chem Phys 106:9213–9226. https://doi.org/10.1063/1.474023
Hsu C-P (2009) The electronic couplings in electron transfer and excitation energy transfer. Acc Chem Res 42:509–518. https://doi.org/10.1021/ar800153f
Voityuk AA, Rosch N (2002) Quantum chemical modeling of electron hole transfer through pi stacks of normal and modified pairs of nucleobases. J Phys Chem B 106:3013–3018. https://doi.org/10.1021/jp013417f
Oberhofer H, Blumberger J (2010) Insight into the mechanism of the Ru2+-Ru3+ electron self-exhchange reaction from quantitative rate calculations. Angew Chem Int Ed 49:3631–3634. https://doi.org/10.1002/anie.200906455
Senthilkumar K, Grozema FC, Bickelhaupt FM, Siebbeles LDA (2003) Charge transport in columnar stacked triphenylenes: effects of conformational fluctuations on charge transfer integrals and site energies. J Chem Phys 119(18):9809–9817. https://doi.org/10.1063/1.1615476
Gillet N, Berstis L, Wu X, Gajdos F, Heck A, de la Lande A et al (2016) Electronic coupling calculations for bridge-mediated charge transfer using constrained density functional theory (CDFT) and effective Hamiltonian approaches at the density functional theory (DFT) and fragment-orbital density functional tight binding (FODFTB) level. J Chem Theory Comput 12:4793–4805. https://doi.org/10.1021/acs.jctc.6b00564
Oberhofer H, Blumberger J (2010) Electronic coupling matrix elements from charge constrained density functional theory calculations using a plane wave basis set. J Chem Phys 133:244105. https://doi.org/10.1063/1.3507878
Wu Q, Van Voorhis T (2006) Extracting electron transfer coupling elements from constrained density functional theory. J Chem Phys 125:164105. https://doi.org/10.1063/1.2360263
Wu Q, Van Voorhis T (2006) Direct calculation of electron transfer parameters through constrained density functional theory. J Phys Chem A 110:9212–9218. https://doi.org/10.1021/jp061848y
Futera Z, Blumberger J (2017) Electronic couplings for charge transfer across molecule/metal and molecule/semiconductor interfaces: performance of the projector operator-based diabatization approach. J Phys Chem C 121:19677–19689. https://doi.org/10.1021/acs.jpcc.7b06566
Ghan S, Kunkel C, Reuter K, Oberhofer H (2020) Improved projection-operator diabatization schemes for the calculation of electronic coupling values. J Chem Theory Comput 16:7431–7443. https://doi.org/10.1021/acs.jctc.0c00887
Kondov I, Cizek M, Benesch C, Wang H, Thoss M (2007) Quantum dynamics of photoinduced electron-transfer reactions in dye-semiconductor systems: first-principles description and application to coumarin 343-TiO2. J Phys Chem C 111:11970–11981. https://doi.org/10.1021/jp072217m
Ziogos OG, Blumberger J (2021) Ultrafast estimation of electronic couplings for electron transfer between pi-conjugated organic molecules. II. J Chem Phys 155:244110. https://doi.org/10.1063/5.0076555
Henstridge MC, Laborda E, Rees NV, Compton RG (2012) Marcus-Hush-Chidsey theory of electron transfer applied to voltammetry: a review. Electrochim Acta 84:12–20. https://doi.org/10.1016/j.electacta.2011.10.026
Chidsey CED (1991) Free energy and temperature dependence of electron transfer at the metal-electrolyte interface. Science 251(4996):919–922. https://doi.org/10.1126/science.251.4996.919
Breuer M, Rosso KM, Blumberger J (2014) Electron flow in multiheme bacterial cytochromes is a balancing act between heme electronic interaction and redox potentials. Proc Nat Acad Sci USA 111(2):611–616. https://doi.org/10.1073/pnas.1316156111
Byun HS, Pirbadian S, Nakano A, Shi L, El-Naggar MY (2014) Kinetic Monte Carlo simulations and molecular conductance measurements of the bacterial decaheme cytochrome MtrF. ChemElectroChem 1(11):1932–1939. https://doi.org/10.1002/celc.201402211
Polizzi NF, Skourtis SS, Beratan DN (2012) Physical constraints on charge transport through bacterial nanowires. Faraday Discuss 155:43–61. https://doi.org/10.1039/C1FD00098E
Cuevas JC, Scheer E (2017) Molecular electronics: an introduction to theory and experiment. World Scientific Publishing
Datta S (1995) Electronic Transport in Mesoscopic Systems. Cambridge University Press
Landauer R (1989) Conductance determined by transmission: probes and quantised constriction resistance. J Phys: Condens Matter 1:8099–8110. https://doi.org/10.1088/0953-8984/1/43/011
Papior N, Lorente N, Frederiksen T, Garcia A, Brandbyge M (2017) Improvements on non-equilibrium and transport green function techniques: the next-generation transiesta. Comp Phys Comm 212:8–24. https://doi.org/10.1016/j.cpc.2016.09.022
Romero-Muniz C, Ortega M, Vilhena JG, Diez-Perez I, Perez R, Cuevas JC et al (2021) Can electron transport through a blue-copper azurin be coherent? An ab initio study. J Phys Chem C 125:1693–1702. https://doi.org/10.1021/acs.jpcc.0c09364
Futera Z, Ide I, Kayser B, Garg K, Jiang X, van Wonderen JH et al (2020) Coherent electron transport across a 3 nm bioelectronic junction made of multi-heme proteins. J Phys Chem Lett 11:9766–9774. https://doi.org/10.1021/acs.jpclett.0c02686
Futera Z, Wu X, Blumberger J (2023) Tunneling-to-hopping transition in multiheme cytochrome bioelectronic junctions. J Phys Chem Lett 14:445–452. https://doi.org/10.1021/acs.jpclett.2c03361
Carey R, Chen L, Gu B, Franco I (2017) When can time-dependent currents be reproduced by the Landauer steady-state approximation? J Chem Phys 146:174101. https://doi.org/10.1063/1.4981915
Nitzan A (2014) Chemical dynamics in condensed phases: relaxation, transfer, and reactions in condensed molecular systems. Oxford University Press
Valianti S, Cuevas J-C, Skourtis SS (2019) Charge-transport mechanism in azurin-based monolayer junctions. J Phys Chem C 123:5907–5922. https://doi.org/10.1021/acs.jpcc.9b00135
Egger DA, Liu Z-F, Neaton JB, Kronik L (2015) Reliable energy level alignment at physisorbed molecule-metal interfaces from density functional theory. Nano Lett 15:2448–2455. https://doi.org/10.1021/nl504863r
Neaton JB, Hybertsen MS, Louie SG (2006) Renormalization of molecular electronic levels at metal-molecule interfaces. Phys Rev Lett 97:216405. https://doi.org/10.1103/PhysRevLett.97.216405
Biava H, Schreiber T, Katz S, Voller J-S, Stolarski M, Schulz C et al (2018) Long-range modulations of the electric fields in proteins. J Phys Chem B 122:8330–8342. https://doi.org/10.1021/acs.jpcb.8b03870
Bim D, Alexandrova AN (2021) Electrostatic regulation of blue copper sites. Chem Sci 12:11406–11413. https://doi.org/10.1039/D1SC02233D
Bradshaw RT, Dziedzic J, Skylaris C-K, Essex JW (2020) The role of electrostatics in enzymes: do biomolecular force fields reflect protein electric fields? J Chem Info Model 60:3131–3144. https://doi.org/10.1021/acs.jcim.0c00217
Stuyver T, Ramanan R, Mallick D, Shaik S (2020) Oriented (local) electric fields drive the millionfold enhancement of the H-abstraction catalysis observed for synthetic metalloenzyme analogues. Angew Chem Int Ed 59:7915–7920. https://doi.org/10.1002/anie.201916592
Suydam IT, Snow CD, Pande VS, Boxer SG (2006) Electric fields at the active site of an enzyme: direct comparison of experiment with theory. Science 313:200–204. https://doi.org/10.1126/science.1127159
Htwe EE, Nakama Y, Yamamoto Y, Tanaka H, Imanaka H, Ishida N et al (2018) Adsorption characteristics of various proteins on a metal surface in the presence of an external electric potential. Colloid Surf B 166:262–268. https://doi.org/10.1016/j.colsurfb.2018.03.035
Martin LJ, Akhavan B, Bilek MMM (2018) Electric fields control the orientation of peptides irreversibly immobilized on radical-functionalized surfaces. Nat Comm 9:357. https://doi.org/10.1038/s41467-017-02545-6
Lakshmi S, Dutta S, Pati SK (2008) Molecular electronics: effect of external electric field. J Phys Chem C 112:14718–14730. https://doi.org/10.1021/jp800187e
De Renzi V, Rousseau R, Marchetto D, Biagi a, Scandolo S, del Pennino U. (2005) Metal work-function changes induced by organic adsorbates: a combined experimental and theoretical study. Phys Rev Lett 95:46804. https://doi.org/10.1103/PhysRevLett.95.046804
Derry GN, Kern ME, Worth EH (2015) Recommended values of clean metal surface work functions. J Vac Sci Technol A 33:60801. https://doi.org/10.1116/1.4934685
Amdursky N, Ferber D, Bortolotti CA, Dolgikh DA, Chertkova RV, Pecht I et al (2014) Solid-state electron transport via cytochrome c depends on electronic coupling to electrodes and across the protein. Proc Nat Acad Sci USA 111:5556–5561. https://doi.org/10.1073/pnas.1319351111
Casalini S, Berto M, Kovtun A, Operamolla A, Di Rocco G, Facci P et al (2015) Surface immobilized his-tagged azurin as a model interface for the investigation of vectorial electron transfer in biological systems. Electrochim Acta 178:638–646. https://doi.org/10.1016/j.electacta.2015.07.156
Zanetti-Polzi L, Daidone I, Bortolotti CA, Corni S (2014) Surface packing determines the redox potential shift of cytochrome c adsorbed on gold. J Am Chem Soc 136:12929–12937. https://doi.org/10.1021/ja505251a
Jensen PS, Chi Q, Grumsen FB, Abad JM, Horsewell A, Schiffrin DJ et al (2007) Gold nanoparticle assisted assembly of a heme protein for enhancement of long-range interfacial electron transfer. J Phys Chem C 111:6124–6132. https://doi.org/10.1021/jp068453z
Liu S, Vareiro MMLM, Fraser S, Jenkins ATA (2005) Control of attachment of bovine serum albumin to pulse plasma-polymerized maleic anhydride by variation of pulse conditions. Langmuir 21:8572–8575. https://doi.org/10.1021/la051449e
Onoda A, Taniguchi T, Inoue N, Kamii A, Hayashi T (2016) Anchoring cytochrome b562 on a gold nanoparticle by a heme-heme pocket interaction. Eur J Inorg Chem:3454-9. https://doi.org/10.1002/ejic.201600301
Wieland F, Bruch R, Bergmann M, Partel S, Urban GA, Dincer C (2020) Enhanced protein immobilization on polymers - a plasma surface activation study. Polymers 12:104. https://doi.org/10.3390/polym12010104
Vacek J, Zatloukalova M, Kabelac M (2022) Redox biology and electrochemistry. Towards evaluation of bioactive electron donors and acceptors. Curr Opin. Electrochem 36:101142. https://doi.org/10.1016/j.coelec.2022.101142
West RM, Janata J (2020) Praise of mercury. J Electroanal Chem 858:113773. https://doi.org/10.1016/j.jelechem.2019.113773
Dorcak V, Kabelac M, Kroutil O, Bednarova K, Vacek J (2016) Electrocatalytic monitoring of peptidic proton-wires. Analyst 141(15):4554–4557. https://doi.org/10.1039/c6an00869k
Dorcak V, Novak D, Kabelac M, Kroutil O, Bednarova L, Veverka V et al (2018) Structural stability of peptidic His-containing proton wire in solution and in the adsorbed state. Langmuir 34(24):6997–7005. https://doi.org/10.1021/acs.langmuir.7b04139
Kroutil O, Kabelac M, Dorcak V, Vacek J (2019) Structures of peptidic H-wires at mercury surface: molecular dynamics study. Electroanalysis 31(10):2032–2040. https://doi.org/10.1002/elan.201900314
Murgida DH (2021) In situ spectroelectrochemical investigations of electrode-confined electron-transferring proteins and redox enzymes. ACS Omega 6(5):3435–3446. https://doi.org/10.1021/acsomega.0c05746
Miao P, Wang B, Han K, Tang Y (2014) Electrochemical impedance spectroscopy study of proteolysis using unmodified gold nanoparticles. Electrochem Commun 47:21–24. https://doi.org/10.1016/j.elecom.2014.07.013
Holtz B, Wang Y, Zhu X-Y, Guo A (2007) Denaturing and refolding of protein molecules on surfaces. Proteomics 7(11):1771–1774. https://doi.org/10.1002/pmic.200700053
Acknowledgements
This work was supported by the Czech Science Foundation via projects 19-21237Y and Palacky University Young Researcher Grant (UP JG_2023_006, M.Z.), 20-02067Y (Z.F.), 22-26590S (V.O.), and 20-06873X (M.C.). M.C. rephrased a part of the text of “Electro-manipulation of protein structure and function” section using OpenAI’s ChatGPT 3.5. Daniel Havelka, PhD, is gratefully acknowledged for preparing the material for Fig. 7. The authors are indebted to Ben Watson-Jones MEng for language correction.
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Jan Vacek: conceptualization, methodology – literature search, writing – original draft, writing – review and editing – special focus on the “Intrinsic electroactivity of proteins,” “Electroactive redox labels in protein sensing,” and “Nano/micro materials in protein electrochemistry: double-surface technique” sections, supervision, project administration. Veronika Ostatná: methodology – literature search, writing – special focus on the “Intrinsic electroactivity of proteins” section, writing – review and editing. Michal Cifra: methodology – literature search, writing – special focus on the “Electro-manipulation of protein structure and function” section, writing – review and editing. Zdeněk Futera: conceptualization, methodology – literature search, writing – special focus on the “Theory and computation of electron transfer in proteins” section, writing – review and editing. Martina Zatloukalová: methodology – literature search, writing – special focus on the “Electroactive redox labels in protein sensing” section, writing – review and editing. Vlastimil Dorčák: methodology – literature search, writing – special focus on the “Intrinsic electroactivity of proteins” section, writing – review and editing.
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Vacek, J., Zatloukalová, M., Dorčák, V. et al. Electrochemistry in sensing of molecular interactions of proteins and their behavior in an electric field. Microchim Acta 190, 442 (2023). https://doi.org/10.1007/s00604-023-05999-2
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DOI: https://doi.org/10.1007/s00604-023-05999-2