Construction of Uniform Monolayer-and Orientation-Tunable Enzyme

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Energy, Environmental, and Catalysis Applications

Construction of Uniform Monolayer- and Orientation Tunable Enzyme Electrode by a Synthetic Glucose Dehydrogenase without Electron Transfer Subunit via Optimized Site-Specific Gold Binding Peptide Capable of Direct Electron Transfer Yoo Seok Lee, Seung Woo Baek, Hyeryeong Lee, Stacy Simai Reginald, Yeongeun Kim, Hyunsoo Kang, Ingeol Choi, and In Seop Chang ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b08876 • Publication Date (Web): 01 Aug 2018 Downloaded from http://pubs.acs.org on August 3, 2018

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ACS Applied Materials & Interfaces

Construction of Uniform Monolayer- and Orientation Tunable Enzyme Electrode by a Synthetic Glucose Dehydrogenase without Electron Transfer Subunit via Optimized Site-Specific Gold Binding Peptide Capable of Direct Electron Transfer

Yoo Seok Lee1,‡, Seungwoo Baek2,‡, Hyeryeong Lee1, Stacy Simai Reginald1, Yeongeun Kim1, Hyunsoo Kang1, In-Geol Choi2* and In Seop Chang1* 1

School of Earth Sciences and Environmental Engineering, Gwangju Institute of Science and

Technology (GIST), 123 Cheomdan-gwagiro, Buk-gu, Gwangju 61005, Republic of Korea 2

Department of Biotechnology, College of Life Sciences and Biotechnology, Korea University,

Seoul 02841, Republic of Korea

Keywords: direct electron transfer, synthetic glucose dehydrogenase, gold binding peptide, orientation, electron tunneling distance

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Abstract The direct electron transfer (DET) between enzymes and electrodes is a key issue for practical use of bioelectrocatalytic devices as a bioenergy process, such as enzymatic electrosynthesis, biosensors and enzyme biofuel cells. To date, based on the DET of bioelectrocatalysis, less than 1% of the calculated theoretical current was transferred to final electron acceptor due to energy loss at enzyme-electrode interface. This study describes the design and construction of a synthetic glucose dehydrogenase (GDH; α and γ subunits) combined with a gold binding peptide at its amino- or carboxy- terminus for direct contact between enzyme and electrode. The fused gold binding peptide facilitated stable immobilization of GDH and constructed uniform monolayer of GDH onto an Au electrode. Depending on the fused site of binding peptide to the enzyme complex, 9 combinations of recombinant GDH proteins on the electrode show significantly different direct electron transfer efficiency across the enzyme-electrode interface. The fusion of site-specific binding peptide to the catalytic subunit (α subunit, carboxy-terminus) of the enzyme complex enabled apparent direct electron transfer (DET) across the enzyme– electrode interface even in the absence of electron transfer subunit (i.e., β subunit having cytochrome domain). The catalytic glucose oxidation current at an onset potential of c.a. (−) 0.46 V vs. Ag/AgCl was associated with the appearance of an FAD/FADH2 redox wave and a stabilized bioelectrocatalytic current of more than one hundred microamperes, determined from chronoamperometric analysis. Electron recovery was 7.64%, and the catalytic current generation was 249 µA per GDH enzyme loading unit (U), several orders of magnitude higher than the values reported previously. These observations corroborated that the last electron donor facing to electrode was controlled to be in close proximity without electron transfer intermediates and the native affinity for glucose was preserved. The design and construction of the site-specific “sticky-ended” proteins without loss of catalytic activity could be applicable to other redox enzymes having a buried active site.


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INTRODUCTION Effective redox bioelectrocatalysts are key to the design and development of next generation microscale electrochemical biosensors, biomedical devices, and biological fuel cells.1-6 Efficient device operation requires operational parameter optimization. First, the enzymes should have high catalytic activities and stabilities, and they should be economically feasible to produce.7,8 Second, bioelectrocatalysis processes require enzyme immobilization methods that optimize the electron transfer (ET) kinetics between the enzyme and the electrode.10-13 Ideally, a biocatalyst should enable fuel oxidation at a low redox potential, without interference from background redox mediators, through direct electron transfer (DET), such that the difference between the operating potentials of the anode and cathode is maximized (i.e., the driving force of the fuel cell is maximized). Use of an external redox mediator (or shuttle) for electron transfer introduces a potential difference between the enzyme cofactor/active site and the mediator, resulting in a loss in potential.14,15 Most biofuel cells employed redox enzymes described to date utilize mediated electron transfer (MET) because this approach can produce current densities in the range of 200 µA/cm2 to 3500 µA/cm2, much higher than the current densities achieved using DET;7-11 however, MET has inherent disadvantages, particularly associated with the thermodynamic losses resulting from a negative change in the Gibbs free energy associated with ET between the enzyme and the mediator. Additionally, the diffusion, toxicity and cost issues of the mediator, as well as the system instability can detract from a MET system’s performance.14,16,17 The formation of direct enzyme–electrode electrical connections depends strongly on the orientation of the immobilized protein and the distance between the enzyme redox-active cofactor and the electrode surface.9,10,12,13,15,17 Most immobilization techniques rely on intermolecular forces, such as dipole–dipole interactions, hydrophobic interactions, or Coulombic interactions. Chemical coupling methods have also been tested, for example, crosslinking and encasing an enzyme within a polymeric hydrogel on the electrode.19-21 Recently, we reported the immobilization of flavin adenine dinucleotide (FAD)-dependent glucose dehydrogenase and glucose oxidase (GOx) in free-standing graphitized carbon nanofiber paper modified via non-covalent functionalization with a biofunctional linker reagent.7,12,22 The pyrene moiety engaged in π–π stacking interactions with the nanofiber wall that, coupled with a reactive end-amine reaction, immobilized the proteins and yielded a significant glucose catalytic



current. The extent to which these methods controlled the protein orientations on the electrode surfaces was limited. The enzyme cofactor FAD and the active site of the enzyme were embedded inside the enzyme, precluding close contact between the catalytic subunit and the electrode surface. Therefore, no current could be generated in the absence of a mediator, even in enzymes with a very high activity. Efforts geared toward overcoming these drawbacks have suggested that the enzyme–electrode interface could be improved by attaching mediator molecules to the electrode. These molecules, such as pyrroloquinolines, quinones, and benzoquinones, should have appropriate redox potentials that provide a driving force for pulling electrons from the cofactor to the electrode.13,23-26 Electron transfer molecules, such as gold nanoparticles and cytochrome c, have been successfully attached to enzymes, enabling DET across the enzyme–electrode interface.11,27 These approaches confirmed that the specific attachment of electron transfer intermediates, metal nanoparticles, and redox mediators, which supplies wiring and bridges the gap between enzyme and electrode, behaves in a manner similar to the electron transfer subunit and can transfer electrons from a buried active site to the electrode. Despite the presence of a DET signal, the potential range shifted as a result of the additional charge transfer resistance. These findings indicated that although these enzymes were immobilized on the electrodes, their orientations and/or the distances were unfavorable for electron tunneling. Another approach introduced sitespecific wiring by inserting non-canonical amino acids at desired sites to covalently link a linker bound to the electrode.28 The liner assisted immobilization in the proper orientation and provided close proximity to the electrode. On the other hand, the complicated linking process involved harsh chemical conditions that may compromise enzymatic activity29 and increase the device cost. These results suggested a need to optimize the electron tunneling conditions by minimizing electron transfer intermediates, removing external molecules, and minimizing energy loss during electron transfer at the enzyme–electrode interface. The FAD-GDH complex from the Burkholderia cepacia is a thermostable, oxygenindependent DET enzyme, rendering it useful for the development of bioelectrocatalytic enzyme electrodes.14,18,27,30,31 The native enzyme complex consists of three subunits: a catalytic FAD binding α subunit, a small chaperon-like γ subunit, and a c-type cytochrome β subunit.29 The γ subunit acts as a chaperon to correct the overall folding and maturation, as well as enabling FAD cofactor binding.30 The γ subunit contains a twin arginine translocase (Tat) pathway motif that


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localizes the mature protein to the periplasm of the bacteria. The β subunit is a 43 kDa c-type multi-heme cytochrome capable of transferring electrons from glucose oxidation at the α subunit to an electron acceptor. The entire FAD-GDH complex displays DET with relatively low bioelectrocatalytic currents and a redox potential of –460 mV (vs. Ag/AgCl) at pH 7.0. The size of the β subunit, which enables ET in the native system,33 as well as the random orientations of the enzyme, reduced the efficiency of ET to the electrodes in previous studies.14,18 Therefore, we reasoned that the construction of an FAD-GDH complex with a site-specific gold-binding peptide at the active center of the α subunit and the omission of the β subunit, to reduce the number of electron transfer intermediates at the enzyme–electrode interface, would promote optimal enzyme orientation on electrode35-37 and, at the same time, shorten the enzyme–electrode distance for efficient DET. In this study, a synthetic enzyme complex was designed by combining the binding properties of a gold-binding peptide (GBP) co-expressed with the FAD-GDH complex. The enzymatic and binding activities of the resulting GDH enzyme construct were evaluated using biochemical, spectroscopic, and imaging protocols. The synthetic enzyme constructs immobilized on an Au surface were imaged by atomic force microscopy (AFM). The enzymatic association/dissociation rates and the surface coverage over the electrode were analyzed by SPR spectroscopy. The electron transfer mechanism and rate were evaluated using cyclic voltammetry and the potentiostatic polarization curve, respectively, whereas continuous catalytic oxidation current development at a negative potential was analyzed by chronoamperometry.



RESULTS AND DISCUSSION Design and construction of the synthetic GDHs. We hypothesized that fusion of the metal binding peptide to the GDH α or γ subunit would provide close contact between the catalytic active site and the electrode, leading to oriented immobilization of the protein on the electrode (Fig. 1). The GBP properties were exploited to design recombinant GDH α and γ subunits. The GBP was fused to the N- or C- terminus of each GDH subunit. After fusion of the GBP to the GDH subunit, a six histidine tag (for affinity chromatography), a maltose binding protein (protein solubility enhancer), and a TEV cleavage site (to separate the GDH-GBP fusion protein from the MBP) were added to the N-terminus of the GDH protein (Fig. 2a). Without the maltose binding protein, the GDH protein was not soluble (data not shown). The MBP fusion protein shifted the size of the GDH protein by approximately 40 kDa compared to the original protein (original protein size: GDH α subunit was 60 kDa, GDH γ subunit was 15 kDa) (Fig. 2b). We confirmed that the MBP fusion at the N-terminus did not affect the enzyme activity. Release of the MBP from the GDH protein by TEV protease treatment (Supplementary Fig. S1) did not affect the enzyme activity (Supplementary Tables S2 and 3). All experiments were performed using the MBP-cleaved proteins.

Catalytic activity and Au binding kinetics of the synthetic GDHs. The GDH activities and electrode binding capacities, factors that favor DET, were evaluated using biochemical, spectroscopic, and molecular imaging methods. The 2,6-dichloroindophenol (DCIP) method was used to measure the activities of the GDH constructs. The absorbance decreased with increasing enzyme concentration (Supplementary Tables S2 and 3). The binding kinetics of the native GDH and synthetic GDH proteins bound to the gold electrode surfaces were characterized by SPR spectroscopy, and the enzyme coverage on the electrode surfaces was determined using the Langmuir adsorption model (Supplementary Fig. S1). All measurements were collected under identical conditions, making the results directly comparable. The concentrations of the enzymes were 0, 0.125, 0.25, 0.5, 1, or 2 µM, and their respective SPR sensograms were recorded. After a stable baseline signal was established by allowing 0.1 M PBS buffer, pH 7.4, to flow over the Au surface, solutions of the wild type or genetically engineered proteins were allowed to flow at a rate of 30 µL/min over the surface. The amount of enzyme immobilized on the Au surface increased over time until it reached a steady state, at which point the surface coverage was in


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equilibrium with the enzyme concentration in the solution. The apparent adsorption rate (ka) of the native GDH complex on the gold surfaces was low, a finding likely due to the non-specific affinity of the native GDH for the gold surface (Fig. 3a). This result suggested that the native GDH enzyme did not form a stable immobilization layer. By contrast, the synthetic GDH enzymes containing either an N- or C-terminal GBP at the α and γ subunits displayed significantly higher gold binding activities than the native GDH (Supplementary Fig. S2). Among the synthetic GDH constructs (Fig. 2a. lists the acronyms of the synthetic constructs), GDHαCG-γ, in which GBP was expressed in the α subunit only and a native γ subunit was present, showed an 8.6-fold higher adsorption rate (ka of 6822 M–1s–1) compared to the native GDHα-γ (Figure 3a). Measurements of enzyme desorption from the Au surface as the running buffer flowed over the enzyme-saturated interface revealed that the fusion protein desorbed from the surface at a 1.7-fold higher rate (kd of 4.01×10–5 s–1) compared to the native GDH. The synthetic GDHαCG-γ bound to the Au surface with binding affinity (equilibrium constant) that was 3.2-fold higher than that of the native GDH (Keq of 1.70×108 vs. 5.33×107), suggesting that the synthetic GDHαCG-γ formed a rigid biomolecular interface. A Langmuir adsorption model fit of both sets of SPR data revealed that the surface coverage, calculated using Equation 2, was higher for the synthetic GDH than for the native GDH (Fig. 3b): kobs= ka C+kd

(2)

where ka is the association rate constant, kd is the dissociation rate constant, and C is the running concentration of the enzyme. The equilibrium constant, Keq, could be calculated as the ratio ka/kd. At a protein concentration of 55 µg/mL (0.5 µM), almost 90% of the gold surface (0.9 cm2) was covered by the synthetic construct, GDHαCG-γ. This observation suggested that the surface coverage (90%) via protein immobilization was comparable to that (82%) obtained via conventional chemical functionalization using alkanethiol on the Au surface.39 It should be noted that the other fusion constructs displayed higher binding affinities for the gold electrode than the native GDH (Supplementary Figs. S2 and 3) (Supplementary Table S4).

Binding characterization of the synthetic GDH on the Au substrate. The oriented immobilization and morphologies of the native or synthetic constructs (GDHαCG-γ) on a gold electrode were characterized using high resolution AFM in the noncontact mode (Figs. 4a–4d).



The GDH-GBP units were more tightly packed and more evenly distributed across the electrode surface (Figs. 4a and b) compared to the wild-type GDH units (Figs. 4c and d). The two enzyme films displayed similar thicknesses, but the variations in the enzyme layer height of the native GDHα-γ (7 ± 6 nm) was almost triple the corresponding value obtained from the synthetic GDHαCG-γ (7 ± 2 nm), indicating that the wild type GDH without GBP formed multiple layers, possibly resulting from agglomeration along the direction perpendicular to the surface. By contrast, the synthetic GDHαCG-γ formed a closely packed and planiform monolayer with a thickness very close to the height of a single GDH molecule (~6 nm), suggesting that the GBP unit oriented the enzyme molecules in a uniform manner with the subunit facing the electrode. Furthermore, we observed undulating plateau-like features about 150 nm in width across the surface, indicative of the polycrystalline gold nanoparticle (AuNP) structure of the underlying surface (Figures 4a and c). Taken together, these findings indicated that the synthetic GDHs formed a surface that was more evenly distributed and had a more oriented protein layer than the native GDH (Supplementary Fig. S4).

DET-based bioelectrocatalysis by the synthetic GDH. The DET capacities of the synthetic GDHαCG-γ, in which GBP was expressed only at the catalytic subunit, the α subunit, and of the native GDHα-γ on a gold electrode were examined using cyclic voltammetry in PBS buffer (100 mM, pH 7.0) at 30°C. (Figs. 5a–f). Following the addition of glucose, the fusion construct showed strong redox events at –455 mV (vs. Ag/AgCl electrodes, Fig. 5a). The significant increase in the oxidative current was a response to enzyme-catalyzed glucose oxidation. The starting potential of –455 mV provided evidence for efficient DET between the FAD center and the electrode, which had a formal potential (E0) of –460 mV (vs. Ag/AgCl) at pH 7.0. By contrast, the wild type GDH displayed almost no anodic peak upon the addition of glucose, similar to the background current. These findings, that only the synthetic GDHαCG-γ exhibited an apparent DET signal, indicated that direct contact between the FAD center and the electrode was facilitated by the engineered GBP. Similar approaches to efficient DET have been reported. For example, the attachment of a metal nanoparticle to an enzyme enabled DET of the enzyme by bridging the gap between the enzyme and the electrode.37 Another system consisted of a GOx mutant bearing a single free sulfhydryl group (cysteine) on its surface and a AuNP addition, attachment of a redox mediator to an enzyme was found to enhance the efficient electron transfer


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of the enzyme electrode.10 The major difference between these approaches and the approach described here is that we used a genetically designed synthetic protein. This engineered GDHGBP had a site-specific 12 base pairs long sticky end (metal binding peptide, 1.3 kDa) without electron transfer intermediates. The construct enabled significant DET labelled with a single maleimide attached to the surface of the protein near the FAD center, enabling a redox relay via cysteine.9 In addition, attachment of a redox mediator to an enzyme was found to enhance the efficient electron transfer of the enzyme electrode.13 This construct enabled significant DET across the enzyme–electrode interface and stable current generation that maintained a negative potential under various loads. This protein played a dual role, acting as both a binding and an orienting agent for direct electric communication between the active center and the electrode. In the presence of glucose, an electrocatalytic current was observed to begin at –455 mV, forming a large sigmoidal catalytic wave corresponding to FAD/FADH2. In earlier studies, however, an enzyme-catalyzed glucose oxidation current was observed without apparent signaling of the FAD/FADH2 redox wave.11,40 Although the oxidative current began at –400 mV, its shape was slanted toward the x-axis. The shift in the redox potential and the disappearance of the redox wave may have resulted from an overpotential caused by the high internal resistance to electron transfer at the enzyme electrode. These results indicated that site-specific expression of the GBP at the active center could facilitate contact between the enzyme and the electrode for DET, producing a relatively low additional resistance. Recent significant advances in the development of DET-based bioelectrocatalysts are tabulated in Table 3. Amazingly, compared to other previous reports as well as native and synthetic GDHs in this study, a synthetic GDHαCGγ electrode only produced significantly high catalytic current, 249 µA per GDH enzyme loading unit (U), which is several orders of magnitude high. Additionally, the electron recovery calculated based on the current max monitoring (Imax, monitored using amperometry detection, GDHαCG-γ electrode conditions listed in Table 1 and 2) and the theoretical current calculations reached to 7.64% indicating that this engineered GDH-GBP (GDHαCG-γ: GBP expressed in carboxy-terminus in α-subunit with γ-subunit) is an exact combination to be desired electron transfer across GDH enzyme-gold electrode interface. Most previous studies reported electron recoveries of less than 0.1%. This high electron recovery was supported by the calculation of the current generation per GDH enzyme unit, which was several orders of magnitude higher than the corresponding values reported in previous studies.



The development of an open circuit potential (OCP) in the presence of synthetic GDHαCG-γ or the native GDH is shown in Fig. 5c. Introduction of 100 mM glucose shifted the redox potentials of both the synthetic GDH and the native GDH to negative values, and they stabilized at –370 mV and –180 mV (vs. Ag/AgCl), respectively. In principle, the pure catalytic FAD/FADH2 reaction should generate a negative OCP in the absence of glucose, such that the change in potential is attributed to glucose oxidation, as estimated by the Nernst equation. The 60 mV potential change in response to a 100 mM increase in the glucose concentration stabilized the FAD/FADH2 onset potential at –460 mV (vs. Ag/AgCl) at pH 7.9. Although the OCP observed in the fusion protein electrode was close to its theoretical maximum, it did not perfectly coincide with the theoretical Nernstian behavior. Moreover, no direct evidence was available to suggest that all synthetic GDH molecules were fully immobilized on the electrode surface with the same orientation. The observed behavior may indicate that a fraction of the enzymes were immobilized in an orientation unfavorable for DET. By contrast, the stabilized OCP was very close to the formal potential of FAD/FADH2, indicating that the sample covered most of the electrode surface with the enzymes in an orientation suitable for DET. The thermostability, oxygen-independent activity, and substrate abundance in nature have made GDH a near-ideal bioelectrocatalytic system, the performances of which depend significantly on enzymatic bioelectrocatalysis. The inverse relationship between the durability of an oxidative current and the magnitude of a negative potential directly determines the performance of these systems. To date, DET signals resulting from substrate oxidation have been reported only over short loading times due to insufficient electron relay and significant potential shift. High current generation at a low scan rate, durable current generation at a negative potential, and long loading times should be assessed to determine whether DET is strong. The potentiostatic polarization curve of a synthetic GDHαCG-γ electrode (Fig. 5d) revealed the generation of a catalytic oxidation current, starting at –450 mV, with a maxi-mum current at –300 mV (vs. Ag/AgCl), at a scan rate of 1 mVs–1. Amperometry curves of the synthetic construct (GDHαCG-γ) revealed that at a constant potential, an electron relay from the enzyme to the electrode was continuously maintained at various poised potentials (i.e., –400, –300, –200, and –100 mV), with a stabilized current density of 107 µA at –200 mV (Fig. 5e). In addition, the electric current output increased with increasing glucose concentration (Fig. 5f), confirming that the catalytic current was indeed proportional to the physiological concentration of glucose in the


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blood. This system avoided problems associated with toxic mediators or mediator diffusion. These results suggested that the synthetic GDH constructs might be useful in on-body devices for health and medical applications. Although electrons were transferred directly from the enzyme to the electrode under various conditions, the initial electrode performance decreased by 68% after continuous maintenance of an electron relay at a poised potential (i.e., 334–107 µA at –200 mV), suggesting that the biological template material-based bioelectrode might require modifications to maintain the initial DET efficiency at the enzyme–electrode interface. Electrochemical studies of this redox protein electrode revealed that the orientations of the protein and the shortest distance between the active center and the electrode surface were of the utmost importance for facilitating direct electrochemistry across the enzyme–electrode interface. These results suggested that the GBP expression site in the three-dimensional globular structure of the redox enzyme is an important factor determining DET efficiency. The DET activity of the construct depended on the expression of the GBP unit in the catalytic subunit, for example, at either terminus of the α subunit (αNG-γ) or in a different GDH subunit, for example at either terminus of the γ subunit (α-γCG and α-γNG) (Table 1 and 2). The synthetic GDHαNG-γ displayed strong redox events at –385 mV (vs. Ag/AgCl electrodes), with a large sigmoidal catalytic wave corresponding to FAD/FADH2 (Supplementary Fig. S5a). This significant increase in the oxidative current was a response to enzyme-catalyzed glucose oxidation. The onset potential of –455 mV provided evidence of efficient DET between the FAD center and the electrode, which had a formal potential (E0) of –460 mV (vs. Ag/AgCl) at pH 7.0. The CV profile of GDHαNG-γ was shifted toward more positive values and the charge transfer resistance was 960 Ω higher than that of GDHαCG-γ (Supplementary Fig. S5b), indicating that electron tunneling through GDHαCG-γ was more favorable than it was through GDHαNG-γ. On the other hand, GDHα-γCG and GDHα–γNG showed no catalytic current upon the addition of glucose (Supplementary Figs. S6a and b). The finding that only the synthetic constructs (GDHαCG-γ and GDHαNG-γ) exhibited an apparent DET signal indicated that the enzyme–electrode interface favored DET. In addition, expression of GBP at either subunit (GDHαCG-γ (C or N) and GDHαNG-γ (C or N)) produced DET performances similar to those obtained when GBP was expressed at the α or native γ subunits, indicating that expression of GBP at the γ subunit did not affect the DET efficiency (Supplementary Figs. S6c and d).



CONCLUSIONS This study described the genetic engineering of GDH enzymes using GBP as a molecular binder to enable efficient DET. The enzymes genetically fused with GBP were expressed in E. coli cells. The catalytic activity of the enzyme toward its substrate was conserved in the fusion enzymes, with the latter having strong binding activities, as confirmed spectroscopically and biochemically. Immobilization of the fusion GDH onto an electrode produced significant DET across the enzyme–electrode interface and stable current generation, maintaining a negative potential under various loads. The findings of this study reveal the potential utility of integrated technologies involving a combination of protein engineering, biotechnology, and energy technologies. These results may lead to new and important insights into the development of biological catalyst-based techniques, including enzymatic electrosynthesis, photosynthesis systems, biosensors and biofuel cells, in research fields that rely on DET.


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METHODS Design of the α and γ subunits of the FAD-GDH construct with the GBP. The synthetic constructs were created using the α and γ subunits of FAD-GDH from the proteobacteria Burkholderia cepacia (GenBank ID: AF430844.1). The GBP sequence (LKAHLPPSRLPS) was obtained from Nam et al.31 The subunit constructs were codon-optimized and synthesized (GeneScript Inc., USA). The GBP was inserted at either the N- or C-terminus of the GDH gene by PCR (Supplementary Table S1).

Cloning, overexpression, and purification of the synthetic constructs. Various synthetic constructs (the GDH α subunit, GDH α subunit with GBP at the N- or C-terminus, GDH γ subunit, GDH γ subunit with GBP at the N- or C-terminus) were cloned into a modified pET21a plasmid using the ligation independent cloning method.35 The modified pET21a plasmid included a 6×histidine tag and a maltose binding protein (MBP) fusion tag at the Nterminus for solubilization and purification. The fusion tag could be removed using the Tobacco Etch Virus (TEV) protease cleavage site integrated into the linker. All clones were sequenced and confirmed prior to transforming for expression into the host E. coli BL21 - CodonPlus (DE3) - RIL (Agilent Technologies, USA).

Enzyme activity Assay. The activities of the enzymes in solutions containing Tris–HCl (10 mM, pH 7.5) and MgCl2 (10 mM) were measured in the presence of pNPP (5.5 mM) for 30 min. A 96-well plate reader recorded the absorbance changes of a DCIP solution at 600 nm. The enzyme activity was then calculated from the Beer–Lambert law, as follows (Eq. 1): Enzyme activity (mmol/min) = [V (mL) × OD600nm (cm–1)] / [ε × incubation time (min)]

(1)

where ε is the molar extinction coefficient (M–1 cm–1), which, for 2,6- dichloroindophenol (DCIP), was 16.3 mM–1 cm–1, calculated from the standard DCIP absorbance curve measured at 600 nm using a UV-visible spectrophotometer (DU730, Beckman Coulter Inc., Fullerton, CA, U.S.A); V is the final assay volume; OD600nm (cm–1) is the absorbance change at 600 nm divided by the light path length (1 cm).

TEV protease treatment. Purified GDH protein and the TEV protease were mixed in a 10 to 1 (w/w) ratio and were incubated at 4°C for 16 h. After TEV protease treatment, the purity of the resulting product was checked on a 12% SDS-PAGE gel.

AFM measurements. As recommended by the manufacturer, gold substrates were cleaned with 1:3 (v/v) 30% H2O2/H2SO4 (piranha solution) for 5 min at room temperature, rinsed with deionised water, and dried under nitrogen gas flow to remove any organic contaminants while retaining pristine gold surfaces. To prepare enzyme electrodes, the gold substrates were cut into 1 cm2 squares and fully submerged in 0.5 µM synthetic or native GDH for 120 min. The gold plates with immobilised enzyme were removed from the solution and sprayed with a stream of deionised



water from a soft water bottle. The surfaces were rinsed for 5 min, thoroughly blow dried in argon and scanned by atomic force microscopy (AFM) using contact and non-contact modes with an XEP system controller (AFM, XE200, PSIA Corporation). To minimize feedback artifacts in the non-contact mode, a 125-µm long silicon/aluminium coated cantilever (PPP-NCHR 10M, Park Systems), with a resonance frequency of 200–400 kHz, a nominal force constant of 42 N/m and a scanning velocity of ~0.2 Hz, was used. For high-resolution imaging, cantilevers with a nominal radius of 5 nm were used to assess the topography of enzyme immobilized on the Au surface, ~ 400 nm in size, with one ~125 nm magnified to analyze the detailed characteristics of each enzyme’s position. Images were analyzed in height or amplitude display mode, and average feature widths were quantified by cross-sectional analysis of the peak-to-peak distance over multiple immobilized proteins, using the XEI software provided by the manufacturer.

SPR experiments. The affinity of wild-type and fusion proteins to the gold electrodes was assessed by surface plasmon resonance (SPR) using a Biacore 300 system (GE Healthcare). Immediately prior to each experiment, the Au sensors were cleaned with 1:3 (v/v) 30% H2O2/H2SO4 for 5 min at room temperature, rinsed with deionised water, and dried under a flow of nitrogen gas. To establish the baseline, the reaction buffer was injected first and the analytes were injected at a flow rate of 10 µL/min until full saturation was achieved. The buffer was injected again to monitor dissociation behavior. During the experiments, sensograms were recorded and analysed using BIAevaluation 4.1 software. The Au surface of the sensor chip was regenerated after each binding experiment by injection of NaOH (50 mM), iso-propanol (50%), and CHAPS (0.01%). Association and dissociation rate constants were determined from the experimental SPR data using BIAevaluation 4.1 software, by assuming 1:1 binding with a Langmuir isotherm model. The apparent binding rate (kobs) was calculated using the equation kobs = ka C + kd, where ka and kd are the association and dissociation constants, respectively, and C is the enzyme concentration. The equilibrium constant (Keq) was calculated using the equation Keq = ka/kd, and the equilibrium surface coverage (θ) was determined using the equation θ = C/(C + Keq–1). Electrochemical measurement. Electrochemical measurements, including measurements of cyclic voltammetry, potentiostatic polarization and chronoamperometry, were performed using a potentiostat (Metrohm AutoLab, Utrecht, Netherlands). All tests were carried out using a three-electrode system, with plain gold as the working electrode, Pt wire as the counter electrode and Ag/AgCl (3 M KCl) as the reference electrode. The CV for enzymeelectrode characterization was performed in 0.1 M PBS buffer with 100 mM glucose at scan rates of 1, 10, 20, 30, 40, 50, 60, 70, 80, 90 and 100 mV s–1, with the potential of the working electrode ranging from −0.6 V to 0 V compared with the reference electrode. Potentiostatic polarization curves of the anodes were obtained at a scan rate of 1 mV s–1 was performed in 0.1 M PBS, applying a negative poised potential (–400, –300, –200 and –100 mV vs Ag/AgCl) for 10 min. Chronoamperometry results were used in the form of the experimental current value to obtain the B/A ratio of the experimentally measured current (B) to the calculated theoretical current (A) (Table 1 and 2). The open circuit potential of the enzyme electrode was measured using the Pt counter electrode and the Ag/AgCl reference electrode (MF-2052; Bioanalytical Systems Inc., West Lafayette, IN), which were placed as close as


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possible to the counter electrode (around 0.2 cm). Data were recorded for 60 min at time intervals of 10 s using a data acquisition system (Multimeter 2700, Keithley Co., Cleveland, OH, U.S.A.).



Figure 1. Schematic diagram depicting possible enzyme–cofactor orientations proximal to the electrode, as facilitated by a site-specific gold binding peptide, resulting in direct electron transfer (top) and non-specific immobilization of the enzyme (bottom).


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Figure 2. Recombinant GDH proteins and SDS-PAGE gel image of the purified GDH proteins. (a) Schematic diagram showing the recombinant GDH proteins, (b) SDS-PAGE gel image of the purified GDH proteins, L: protein ladder, α: GDH α subunit, γ: GDH γ subunit, C: GBP fusion at the C-terminus of GDH, N: GBP fusion at the N-terminus of GDH.



Figure 3. (a) SPR spectroscopy results obtained from 0.125, 0.25, 0.5, 1, and 2 µM synthetic GDHαCG-γ or wild-type GDH binding to bare gold surfaces. mRIU corresponds to the change in the dip position of the SPR spectrum. (b) Surface coverage of the synthetic GDH αCG-γ or wildtype GDH, calculated using the Langmuir isotherm model based on the results of the SPR experiments. The schematic diagrams depict possible scenarios for directed and non-specific immobilization of the enzymes.


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Figure 4. AFM images showing the surface topographies of the synthetic GDHαCG-γ (a and b) and native GDHα-γ (c and d) films prepared at 55 µg/mL (0.5 µM). The areas correspond to 500 nm X 500 nm scans. The images in (b and d) represent digitally magnified areas of the red squares in (a and c) and are presented as pseudo-three-dimensional images of 125 nm X 125 nm areas to show the surface topographies following immobilization.



Figure 5. (a) CVs of the synthetic construct (GDHαCG-γ) and wild-type protein (control sample, native GDHα-γ) immobilised on a gold electrode in the presence of 100 mM glucose (pH 7.0, 1 mV/s). The synthetic proteins exhibited enzymatic glucose oxidation starting at –455 mV, corresponding to the direct transfer of electrons from the enzyme. Inset: CVs collected from the wild-type GDH or a bare electrode in the presence of 100 mM glucose. (b) Dependence of direct electron transfer in the synthetic protein on the scan rate. Inset: CV of the fusion protein at a scan rate of 1 mV/s, clearly showing the direct transfer of electrons from/to the enzyme. (c)


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Potentiostatic response of the open-circuit potential over time for the synthetic GDHαCG-γ and wild-type proteins upon addition of glucose (arrow). (d) Potentiostatic polarization curves of the synthetic GDHαCG-γ and wild-type GDH on a gold electrode in the presence of 100 mM glucose in 0.1 M PBS buffer. (e) Amperometric responses of the synthetic GDHαCG-γ to glucose oxidation at the applied potentials of –0.4, –0.3, –0.2 and –0.1 V. (f) Current dependence of the immobilized synthetic GDHαCG-γ on the glucose concentration (0–200 mM).



Table 1 | Protein clones and their enzymatic activities.

C: GBP at the C-terminus; N: GBP at the N-terminus; Control: TEV protease untreated; TEV: TEV protease treated.


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Table 2 | Protein clones and their electrochemical activities.

C: GBP at the C-terminus; N: GBP at the N-terminus; Control: TEV protease untreated; TEV: TEV protease treated; Theoretical current: theoretical generated current obtained from the used enzyme mass and the specific activity results; Imax: maximum current obtained from cyclic voltammetry measurement (ca. –0.21 V vs Ag/AgCl, scan rate: 10 mV/s), and chronoamperometry measurement (set potential: –0.2 V); B/A: the ratio between the experimental current measured using chronoamperometry (B) and the calculated theoretical current (A); Rct: charge transfer resistance measured using electrochemical impedance spectroscopy (EIS); N.M.: Not measured.



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Table 3 | DET-based bioelectrocatalysis and performance differences at various enzyme electrodes. Biocatalyst

Information of used enzyme

Electrolyte

Interface/electrode

1 nM GOx-AuNP solution

1 M glucose in 0.1 M PBS

GOx mutant-AuNP conjugate/ Gold electrode

PQQ dependent glucose degydrogenase

N.M.

0.01 M glucose in 0.02 M MOPS

PQQ dependent glucose degydrogenase

N.M.

Glucose oxidase

Glucose oxidase

Glucose oxidase

Cholesterol oxidase, cholesterol esterase

Ref

-

11

PQQ-sGDH/ 1,4-benzoquinone-SWBP

-

CV test: 100 µA of redox peak was obtained at ca. 0.2 V (vs Ag/AgCl), 10 mV/s. Potentiostatic polarization curve: ca. 80 µA/cm2 was measured at 0.3 V of set potential (vs Ag/AgCl). (Data was extracted from graph)

13

0.01 M glucose in 0.02 M MOPS

PQQ GDH/ CNT paper

-

CV test: 50 µA of redox peak was obtained at ca. 0.0 V (vs Ag/AgCl) after addition of glucose (scan rate: 150 mV/s). Potentiostatic polarization curve: N.M. (Data was extracted from graph)

23

500 U/ml

0.5 mM glucose in PBS (pH 7.4)

GOx/ Nafion-Pt-CNT

13.3

CA test: 41.4 µA/cm2.

41

5 µl of 20-40 mg/mL (50 U)

1 mM glucose in 0.01 M PBS, 0.1 M KCI

GOx/ Prussian Blue-GCE

8.10 · 10-5

CA test: 0.18 µA/cm2.

42

0-1 M glucose in 0.1 M PBS

GOx/ PEI-Nanomesh-Au

-

CV test: strong DET redox peaks at ca. -0.35 V (vs Ag/AgCl) was appeared with ca. 100 µA of peak current, 400 mV/s. (Current was extracted from graph)

43

0.1 mM ATCl in PBS

AChE/ Au–Ppy–rGO/GCE

6.00

CA test: 0.9 µA/cm2 was obtained at 0.65 V of applied potential CV test: oxidation peak potential obtained at ca. 0.2 V (vs Ag/AgCl)

44

COx/ GNS-nPt

0.426

CA test: 0.23 mA was obtained at 0.4 V of applied potential. CV test: 0.16 V of peak-to-peak separation (∆ ∆ Ep) was obtained on the GNS-nPt electrode, 100 mV/s EIS test: 19 kΩ (Rct).

45

CV test: anodic current observed at -0.35 V (vs Ag/AgCl), 5 mV/s.

46

CA test: 3.49 µA of steady current was measured at ca. 0.1 V of the applied poteintal.

47

3 µl of 50 U/mL (500 U/mg)

Cholesterol 5 µM cholesterol oxidase (54 U/mg), and 5 µM cholesterolesterase cholesteryl stearate (13.2 U/mg) in 0.1 M PBS

FAD dependent glucose dehydrogenase

3 mL

0.06 M glucose in 0.1 M PBS

GDH/ AuNPs composite

-

Glucose oxidase

2U


GOx/ PDDA-ZnO-MWNTs

1.75

Native GDH(α-γ)/ gold electrode FAD dependent glucose degydrogenase

Additional electrical test CA test: 0.05 µA/cm2 was shown with 0.6 V (vs Ag/AgCl) of applied potential. CV test: peak current > 50 µA/cm2 was shown at ca. -0.2 V (vs Ag/AgCl), 10 mV/s. Potentiostatic polarization curve: 0.25 µA/cm2 was shown at ca. 0 V of set potential. (Data was extracted from graph)

Glucose oxidase 5 µl of 100 mg/mL

Acetylcholinesterase

Current/ enzyme loading (µA/U)

0.43 U (0.0775 mg)


Synthetic GDH(α-γNG )/ gold electrode Synthetic GDH(αCG-γ )/ gold electrode

No catalytic current observed CA test: 0.249 mA/U was shown at -0.2 V of applied potential. CV test: 0.376 mA/U was obtained at ca. -0.21 V (vs Ag/AgCl), This No catalytic 10 mV/s study current observed Potentiostatic polarization curve: 0.155 mA/U was measured at -0.31 V of set potential (vs Ag/AgCl), 1 mV/s. 249.00

Current/enzyme loading: amperometric results/used enzyme unit; CA test: Chronoamperometry test; N.M.: Not mentioned.


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ASSOCIATED CONTENT Supporting Information Details of the materials, instruments and comparison of experimental results; cloning method; growth and purification of the enzymes; release of the fusion tag by TEV protease treatment; measurement of the enzyme activity; SPR characterization of the synthetic GDH binding kinetics; morphology of a fusion enzyme-immobilized electrode surface; DET capacity of the synthetic GDHs depended on the GBP expression site.

AUTHOR INFORMATION Corresponding Author *

Co-correspondence to In-Geol Choi ([email protected]) or In Seop Chang

([email protected]) Author contributions Y.S.L.: data acquisition, data analysis and manuscript preparation. S.B.: data acquisition, data analysis and manuscript preparation; H.L., S.R., Y.K., and H.K.: data acquisition and data analysis; I.G.C. ([email protected]) and I.S.C. ([email protected]) take responsibility for the integrity of this work. All authors have given approval to the final version of the manuscript.

‡

These authors are contributed equally to this study. Notes The authors declare that there is no competing financial interest.

ACKNOWLEDGEMENTS This work was supported by grants from the National Research Foundation of Korea (NRF), funded by the Korean Government (2016R1A2B3015426), and the GIST Research Institute (GRI) in 2018. Special thanks to Prof. Hohjai Lee for insightful discussions of electrochemistry.



Page 26 of 31

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Table of Contents


Construction of Uniform Monolayer-and Orientation-Tunable Enzyme

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