Investigation of the pH-Dependent Electron Transfer Mechanism of

Article pubs.acs.org/Langmuir

Investigation of the pH-Dependent Electron Transfer Mechanism of Ascomycetous Class II Cellobiose Dehydrogenases on Electrodes Wolfgang Harreither,†,‡ Peter Nicholls,§ Christoph Sygmund,† Lo Gorton,⊥ and Roland Ludwig*,†,⊥ †

Food Biotechnology Laboratory, Department of Food Sciences and Technology, BOKU − University of Natural Resources and Life Sciences, Muthgasse 18, A-1190 Vienna, Austria ‡ Department of Chemistry and Molecular Biology, University of Gothenburg, SE-421 96 Göteborg, Sweden § Department of Biological Sciences, University of Essex, Colchester, Essex CO4 3SQ, U.K. ⊥ Department of Analytical Chemistry/Biochemistry and Structural Biology, Lund University, P.O. Box 124, SE-221 00 Lund, Sweden ABSTRACT: Cellobiose dehydrogenase (CDH) is capable of direct electron transfer (DET) on various carbon and thiol-modified gold electrodes. As a result, these systems have been utilized as biocatalyst in biosensors and biofuel cell anodes. Class I CDHs, from basidiomycetous fungi, are highly specific to cellulose or lactose, and DET is only observed at pH values below 5.5. To extend the applicability of CDH-based electrodes, the catalytic properties and the behavior on electrode surfaces of ascomycetous class II CDHs from Chaetomium attrobrunneum, Corynascus thermophilus, Dichomera saubinetii, Hypoxylon haematostroma, Neurospora crassa, and Stachybotrys bisbyi were investigated. We found that class II CDHs have diverse properties but generally show a lower substrate specificity than class I CDHs by converting also glucose and maltose. Intramolecular electron transfer (IET) and DET at neutral and alkaline pH were observed and elucidated by steady-state kinetics, pre-steady-state kinetics, and electrochemical measurements. The CDHs ability to interact with the electron acceptor cytochrome c and to communicate with electrode surfaces through DET at various pH conditions was used to classify the investigated enzymes. In combination with stopped-flow measurements, a model for the kinetics of the pH-dependent IET is developed. The efficient glucose turnover at neutral/alkaline pH makes some of these new CDHs potential candidates for glucose biosensors and biofuel cell anodes.

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INTRODUCTION In the past decade, the main efforts in bioelectronics focused on the design and development of implantable microbiosensors1−3 and miniature biofuel cells.4−6 The employed redox enzymes must exhibit a stable and efficient electronic communication under human physiological conditions. There are two different ways to establish electric contact between the active site of redox enzymes and electrodes: mediated electron transfer (MET) employing soluble redox mediators or redox polymers and direct electron transfer (DET). Direct coupling of biocatalyst active sites to electrodes allows the development of selective biosensors and environmentally sound biofuel cells.7 In cellobiose dehydrogenase (CDH), DET occurs between the enzyme’s active site, located in a flavodehydrogenase domain (DHCDH), and the electrode via its N-terminal cytochrome domain (CYTCDH), which acts as a relay (“a built-in mediator”8) (Scheme 1). CDH is a flavocytochrome, with the cofactor flavin adenine dinuceotide (FAD) located in the DHCDH domain and heme b in the CYTCDH domain.9,10 In this scheme, the catalytic mechanism consists of a reductive half-reaction, in which two electrons and two protons are abstracted from the anomeric carbon atom of cellobiose or cello-oligosaccharides and transferred to the FAD. The © 2012 American Chemical Society

oxidative half-reaction proceeds by electron transfer to either a two-electron acceptor or two equivalent one-electron acceptors. It is not necessary that the electron acceptors also accept the protons. The reduction of electron acceptors mainly takes place at DHCDH by a ping-pong mechanism, but some one-electron acceptors (e.g., Fe3+ or cytochrome c (cyt c)) can also be reduced by CYTCDH. In this case, the electrons are shuttled sequentially from the reduced FAD on the DHCDH domain to heme b via the intramolecular electron transfer (IET).11 Recent studies have provided strong evidence to the fact that CDH transfers electrons to copper-dependent polysaccharide monooxygenases from the glycoside hydrolase 61 (GH61) family rather than to small molecular-weight electron acceptors like oxygen, iron, or quinones.12,13 The capability of CYTCDH to efficiently transfer electrons to biological macromolecules (like GH 61 enzymes or the redox protein cyt c) is also observed with a polarized electrode. It has been unequivocally proven that CYTCDH is responsible for DET and that DHCDH has never shown observable DET characteristics.14,15 Both IET and DET are pH-dependent Received: February 6, 2012 Revised: March 27, 2012 Published: April 3, 2012 6714

dx.doi.org/10.1021/la3005486 | Langmuir 2012, 28, 6714−6723

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Scheme 1. Electron Transfer Pathways in CDHa

fractions of 1 (1 indicates a homogeneous preparation22): CDH from Chaetomium atrobrunneum IIA (CaCDH) with a DCIP pH optimum at 6.0 and a specific activity of 11.5 U mg−1 (purity = 0.65), CDH from Corynascus thermophilus IIB (CtCDH) with a DCIP pH optimum of 5.0 and a specific activity of 17.8 U mg−1 (purity = 1.0), CDH from Dichomera saubinetii IIA (DsCDH) with a DCIP pH optimum of 5.5 and a specific activity of 13.2 U mg−1 (purity = 0.87), CDH from Hypoxylon haematostroma IIB (HhCDH) with a DCIP pH optimum of 5.0 and a specific activity of 9.0 U mg−1 (purity = 0.62), CDH from Neurospora crassa IIA (NcCDH) with a DCIP pH optimum of 5.5 specific activity of 10.3 U mg−1 (purity = 0.33), and CDH from Stachybotrys bisbyi IIA2 (SbCDH) with a DCIP pH optimum of 4.5 and a specific activity of 2.7 U mg−1 (purity = 0.20). Buffers used in the experiments were 50 mM sodium citrate between pH 3.0 and 6.5, 50 mM sodium hydrogen phosphate between pH 6.0 and 8.0, 50 mM aminoethanol phosphate buffer between pH 8.0 and 10.5, and 50 mM phosphate buffered saline (PBS) adjusted to pH 7.4. The buffers were prepared using deionized water (18 MΩ cm) purified with a Milli-Q system (Millipore, Bedford, MA). All buffers were degassed before use to prevent the formation of air bubbles in the flow system. Amperometric Measurements. A flow-through amperometric wall jet cell with three electrodesa working electrode (graphite electrode modified with CDH), a reference electrode (Ag|AgCl in 0.1 M KCl), and a counter electrode made of a platinum wirewas used. The cell was connected to a potentiostat (Zäta Elektronik, Höör, Sweden). The enzyme-modified electrode was press-fitted into a Teflon holder and inserted into the wall jet cell and kept at a constant distance (ca. 1 mm) from the inlet nozzle.23 The response currents were recorded on a strip chart recorder (Kipp & Zonen, Delft, The Netherlands). The electrochemical cell was connected online to a single line flow injection system, in which the carrier flow was maintained at a constant flow rate of 0.5 mL min−1 by a peristaltic pump (Gilson, Villier-le-Bel, France). The injector was an electrically controlled six-port valve (Rheodyne, Cotati, CA), and the injection loop volume was 50 μL. CDH was immobilized through simple adsorption onto the surface of solid spectroscopic graphite electrodes (o.d. 3.05 mm, Ringsdorff Spektralkohlestäbe, SGL Carbon Sigri Greatlakes Carbon Group Ringsdorff-Werke GmbH, Bonn Germany). The electrode was cut, polished on wet emery paper (Tufbak, Durite, P1200), rinsed with deionized water, and dried. A 5 μL aliquot of enzyme solution (protein concentration = 5 mg mL−1) was spread across the active surface of the electrode. The electrode was dried at room temperature and stored overnight at 4 °C. Before using the electrode, it was thoroughly rinsed with deionized water to remove weakly adsorbed enzyme. The electrode was plugged into the buffer filled wall jet cell. Prior to substrate injection, the electrode was polarized with an applied potential until a stable background current was achieved. The current densities were calculated with respect to the geometric electrode area of 0.0731 cm2. Heterogeneous Kinetics and pH Profiles on Graphite Electrodes. The influence of different substrates, the substrate concentration, and pH on the adsorbed enzyme was electrochemically determined with the flow injection system. The carrier solution was prepared by diluting a carbohydrate stock solution with the appropriate buffer. This solution was allowed to reach the mutarotational equilibrium before it was used. The kinetic parameters for the heterogeneous electron transfer KM,app (apparent Michaelis−Menten constant) and Imax (maximum current response at infinite substrate concentration) were determined for a number of substrates in the DET mode. The kinetic constants were calculated by fitting the observed data to the Michaelis−Menten equation using nonlinear least-squares regression. The substrate concentration values were corrected with the dispersion factor of the flow system. According to the Ruzicka and Hansen relationship, the factor was determined by dividing the steady state current registered for the 50 mM ferrocyanide solution by the peak current for an equal concentration of ferrocyanide. The applied potential was +300 mV. In our case, the dispersion factor (D) based on the 1 mm distance between the inlet nozzle and the electrode and the 0.5 mL min−1 flow was 1.25.24 The applied potential used in the heterogeneous kinetic experiments was

a

Schematic presentation of the catalytic reaction (CAT) followed by the intramolecular electron transfer step (IET) and the direct electron transfer step (DET) of the electrons to the electrode The flavodehydrogenase domain DHCDH is left, and right is the cytochrome domain (CYTCDH).

processes. While in the well-investigated class I CDHs the IET is switched off at pH values above pH ∼5.5−6.0,16 the ascomycetous class II CDHs from Humicola insolens17,18 and Chaetomium sp. INBI (2-26-)19 have shown IET with cyt c as electron acceptor at neutral/alkaline pH and therefore also DET in this pH range seems to be achievable with some class II CDHs.20,21 This was our motivation to search for new CDHs in ascomycetous strains with an IET in the neutral/alkaline pH range. A series of ascomycetous fungi were screened, and six new class II CDHs were found and characterized, including CDHs from Chaetomium atrobrunneum (CaCDH IIA), Corynascus thermophilus (CtCDH IIB), Dichomera saubinetii (DsCDH IIA), Hypoxylon haematostroma (HhCDH IIB), Neurospora crassa (NcCDH IIA), and Stachybotrys bisbyi (SbCDH IIA).22 Besides the presence (denoted by A) or the absence (B) of a C-terminal carbohydrate binding domain (CBM), we found the pH optima for DHCDH and the IET vary from acidic (pH 4.5) to the alkaline (pH 7.5) range. Another unique feature of class II CDHs is their broad substrate spectra. Several class II CDHs convert maltose and especially glucose at considerable rates. Here we report on a detailed kinetic characterization of these new ascomycetous CDHs under two conditions, in solution (homogeneous) and adsorbed on graphite electrodes (heterogeneous). Special interest is given to the mechanism and kinetics of IET and DET as well as the adsorption of CDH to the electrode surface. The results are used to develop an initial model of the electron transfer kinetics and to elucidate whether these CDHs are useful candidates for DET-based glucose biosensors and biofuel cell anodes.

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MATERIALS AND METHODS

Chemicals and Enzymes. All chemicals used were of analytical grade. 2,6-Dichloroindophenol sodium salt hydrate (DCIP) and D(+)lactose monohydrate were from Fluka (Buchs, Switzerland). D(+)Cellobiose, D(+)-glucose, maltose, orthophosphoric acid (85%), citric acid monohydrate, 2-aminoethanol, cytochrome c, and 1,4-benzoquinone were purchased from Sigma-Aldrich Chemicals (Steinheim, Germany). Sodium hydroxide and sodium hydrogen phosphate monohydrate were from VWR (Darmstadt, Germany). Six ascomycetous CDHs were produced according to previous protocols,22 (partially) purified, and studied. The purity was determined by measuring the absorbance ratio 420 nm/280 nm and indicated by 6715


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Figure 1. Dependence of the biocatalytic current of class II CDHs in DET-mode (black line) and MET-mode using 1,4-benzoquinone (gray line) on the pH and the buffer composition (filled squares: 50 mM sodium citrate buffer; filled triangles: 50 mM sodium hydrogen phosphate buffer; filled circles: 50 mM aminoethanol buffer) for 5 mM lactose. The applied potential was +400 mV vs Ag|AgCl in 0.1 M KCl, and the flow rate was 0.5 mL min−1. Below: pH profiles of the relative activities of the investigated CDHs using the two-electron acceptor DCIP (gray line) and the one-electron acceptor cyt c (black line) with lactose as electron donor. equal to +300 mV vs Ag|AgCl in 0.1 M KCl reference electrode. The current versus pH profile was estimated for both DET and MET. The used substrate was in both cases 5 mM lactose, but in the case of the MET measurements 20 μM 1,4-benzoquinone (BQ) was added as an electron acceptor (redox mediator). The BQ stock solution was prepared fresh daily. To guarantee an efficient electrochemical reoxidation of the reduced benzoquinone, the applied potential was set to +400 mV vs Ag|AgCl in 0.1 M KCl.25 Voltammetric Measurements. All measurements were performed at 25 °C. Cyclic voltammetry (scan rate 10 mV s−1) was performed using a Gamry Reference 600 (Gamry Instruments, Warminster, PA). A standard three-electrode configuration was used with an Ag|AgCl reference electrode in 3 M KCl (Gamry Instruments) and a platinum wire as counter electrode. The buffer was carefully degassed under vacuum and purged with argon prior to experiments. To maintain the inert atmosphere during the measurements, argon was blown over the sample solution. CDH-Modified SAM Gold Electrodes. The preparation of thiolmodified disk gold electrodes (BAS, West Lafayette, IN, ϕ = 1.6 mm, geometric surface area = 0.0201 cm2) for cyclic voltammetry started with dipping the electrodes into piranha solution (H2SO4:H2O2 = 3:1) for 10 min, followed by cycling in 0.1 M NaOH between 0 and −1000 mV vs NHE with a scan rate of 100 mV s−1 (10 cycles). Afterward, the electrodes were cleaned mechanically by polishing on Microcloth (Buehler, Lake Bluff, IL) in a Masterprep polishing suspension (0.05 μm, Buehler). The electrodes were rinsed with deionized water, sonicated for 10 min in deionized water, followed by cycling in 0.5 M H2SO4 for 20 cycles with a scan rate of 200 mV s−1 between 0 and +1950 mV vs NHE, and finally rinsed again with deionized water. The

thiol self-assembled monolayer (SAM) formation at the electrode surface was done by immersing the electrode in a 10 mM thioglycerol solution at room temperature overnight. Before exposure to CDH, the electrode was carefully rinsed with deionized water. Then the electrodes were press fitted in a Teflon cap, which formed a cell volume of 30 μL on the thiol-modified gold electrode. Modification with CDH was made by filling the Teflon cell with enzyme solution. A dialysis membrane (Carl Roth, Karlsruhe, Germany, molecular weight cutoff = 14 kDa) was used to trap the enzyme in the cell.26 The dialysis membrane (presoaked in buffer) was pressed onto the Teflon cell and fixed tightly to the electrode with a rubber O-ring. Before the measurements the enzyme preparation was diafiltrated against the buffer used as electrolyte. Enyzme Assays and Protein Determination. The activity of the CDH preparations was specifically determined by following the reduction of 50 μM cytochrome c (ε550 = 19.6 mM−1 cm−1) at 30 °C in the appropriate buffer, containing 30 mM lactose.27 One unit of enzymatic activity was defined as the amount of enzyme that oxidizes 1 μmol of lactose per min under the assay conditions. Alternatively, the DCIP assay measuring both the CDH and DHCDH activity was performed by measuring the time-dependent reduction of 300 μM DCIP at 520 nm and 30 °C in appropriate buffer containing 30 mM lactose.28 The absorption coefficient for DCIP at 520 nm was determined to be 6.8 mM−1 cm−1. Even though the absorbance is pH dependent, the absorbance varied only about 3% between pH 3.0 and 8.0.19 The protein concentration was determined by the Bradford method using a prefabricated assay from Bio-Rad Laboratories (Hercules, CA) and bovine serum albumin as a standard. 6716


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Table 1. Steady-State Kinetic Constants of Class II CDHs for the Electron Acceptors DCIP and Cyt ca enzyme origin (purity factor)

electron acceptor

stoichiometryb

assay pH

KM (μM)

kcat (s−1)

C. atrobrunneum (1.54)

DCIP Cyt c DCIP Cyt c DCIP Cyt c DCIP Cyt c DCIP Cyt c DCIP Cyt c

1 2 1 2 1 2 1 2 1 2 1 2

6.0 5.0 5.0 7.5 5.0 5.0 5.0 5.0 5.5 6.0 5.0 5.0

9.2 8.0 69.8 4.6 5.8 5.7 5.6 25.9 8.7 53.7 2.4 4.9

17.0 (26.2) 21.3 (32.8) 53.8 21.4 22.6 (26.0) 25.2 (27.0) 13.5 (21.9) 18.1 (29.3) 24.8 (74.4) 28.7 (86.1) 4.0 (20.0) 4.9 (26.2)

C. thermophilus (1.0) D. saubinetii (1.15) H. haematostroma (1.62) N. crassa (3.0) S. bisbyi (5.0)

kcat/KM (M−1 s−1) 1.8 2.7 2.2 3.1 3.9 4.4 2.4 7.0 2.9 5.4 1.7 1.0

× × × × × × × × × × × ×

106 106 105 106 106 106 106 105 106 105 106 106

(2.8 × 106) (4.1 × 106)

(4.5 (5.1 (3.9 (1.1 (8.6 (1.6 (8.3 (4.7

× × × × × × × ×

106) 106) 106) 106) 106) 106) 106) 106)

a

Values calculated for a homogeneous enzyme preparation are given in parentheses. bMoles of electron acceptor reduced per mole of lactose oxidized.

Table 2. Steady-State Kinetic Constants of Class II CDHs for Carbohydrates Using Cyt c as Electron Acceptora enzyme origin (purity factor)

assay pH

substrate

KM (μM)

C. atrobrunneum (1.54)

5.0

cellobiose lactose maltose glucose cellobiose lactose maltose glucose cellobiose lactose maltose glucose cellobiose lactose maltose glucose cellobiose lactose maltose glucose cellobiose lactose maltose glucose

2 18 7.8 1.7 3 14 2.4 2.1 12 48 1.3 7.7 5 50 7.1 2.6 7 28 3.1 1.7 7 32 2.4 2.3

C. thermophilus (1.0)

D. saubinetti (1.15)

H. haematostroma (1.62)

N. crassa (3.0)

S. bisbyi (5.0)

a

7.5

5.0

5.0

6.0

5.0

KI (mM)

× 103 × 105

3.3 × 106 39 52 877

× 103 × 104

× 104 × 105

1.6 × 103

× 103 × 105

3.3 × 103

× 104 × 105

× 104 × 105

kcat (s−1) 7.7 (11.9) 8.2 (12.6) 1.0 (1.5) 7.1 (10.9) 11.8 10.9 5.0 10.0 9.4 (10.8) 9.2 (10.6) 1.2 (1.4) 16.2 (18.6) 3.9 (6.3) 3.6 (5.8) 0.5 (0.8) 3.7 (6.0) 3.3 (9.9) 3.4 (10.2) 0.3 (0.9) 1.9 (5.7) 2.1 (10.5) 2.3 (11.5) 0.2 (1.0) 1.6 (8.0)

kcat/KM (M−1 s−1) 3.3 × 106 (6.0 4.5 × 105 (6.0 128 (197) 40 (64) 4.0 × 106 7.8 × 105 208 48 7.6 × 105 (9.0 1.9 × 105 (2.2 90 (108) 21 (24) 8.5 × 105 (1.3 7.1 × 104 (1.2 67 (110) 14 (23) 4.5 × 105 (1.4 1.2 × 105 (3.6 10 (30) 11 (33) 3.2 × 105 (1.5 7.1 × 104 (3.5 9 (42) 7 (35)

× 106) × 106)

× 105) × 105)

× 106) × 105)

× 106) × 105)

× 106) × 105)

Values calculated for a homogeneous enzyme preparation are given in parentheses. following concentrations at pH 7.4: 500, 250, 50, and 12.5 μM. The rates for the fast electron transfer reactions are averages of at least three experiments.

Homogeneous Kinetics. Initial rates for the determination of steady-state catalytic constants were determined spectrophotometrically at 30 °C in the indicated buffer. For kinetic measurements, carbohydrate stock solutions used were prepared in the appropriate buffer and allowed to stand for 12 h until mutarotational equilibrium was reached. Stock solutions of electron acceptors were prepared in deionized water and immediately used. The reaction stoichiometry is 1 for the two-electron acceptor DCIP but 2 for the one-electron acceptor cyt c. All kinetic constants were calculated using nonlinear least-squares regression (Sigma Plot 11, Systat Software, San Jose, CA) by fitting the observed data to the Michaelis−Menten equation. An Applied Photophysics (Leatherhead, UK) SX-20 stopped-flow apparatus in either diode array or single wavelength mode was used to measure the pre-steady-state kinetics of FAD and heme b reduction at 447 and 564 nm, respectively. The experiments were carried out with an enzyme concentration of 6 μM using cellobiose as substrate at 25 °C. Both enzyme and cellobiose solution were buffered in 80 mM potassium phosphate, pH 5.0 or 7.4. Cellobiose was varied in the

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RESULTS

Homogenous Kinetics. The pH-dependent activity of all CDHs was measured with the one-electron acceptor cyt c and the two-electron acceptor DCIP (Figure 1). The pH optima found with cyt c (between pH 5.0 and 7.5) and DCIP (5.0− 6.0) were used to determine the apparent catalytic constants for both electron acceptors (Table 1). Additionally, the kcat values were calculated for a homogeneous CDH preparation by normalizing with the determined purity factor (see Materials and Methods). It has to be considered that turnover rates are given for the electron acceptors, and in the case of cyt c two molecules are necessary for the oxidation of a carbohydrate 6717


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Table 3. Heterogeneous, Apparent Catalytic Constants of Class II CDHs Adsorbed on Graphite Electrodes for Carbohydrates enzyme origin C. thermophilus

D. saubinetii

H. haematostroma

substrate

pH

cellobiose lactose maltose glucose cellobiose lactose maltose glucose cellobiose lactose maltose glucose

8.5

5.0

5.0

Imax (μA cm−2) 4.9 4.1 1.7 2.6 25.9 33.1 2.5 7.7 2.2 2.1 0.75 0.22

KM,app (μM) 170 370 2.4 × 1.5 × 215 410 1.2 × 9.3 × 130 190 1.6 × 3.9 ×

Imax/KM,app (μA M−1 cm−2)

103 105

104 104

103 104

2.9 1.1 7.0 1.7 1.2 8.1 1.9 8.3 1.7 1.1 1.4 1.9

× × × × × × × × × × × ×

104 104 102 101 105 104 102 101 104 104 102 101

pH 7.4

7.4

7.4

Imax (μA cm−2)

KM,app (μM)

7.1 6.6 1.8 3.5 8.3 8.5 3.0 4.8 0.79 1.1

800 232 1.9 × 2.4 × 80 120 5.4 × 3.8 × 78 260

0.38

5.2 × 104

103 104

103 104

Imax/KM,app (μA M−1 cm−2) 8.8 2.9 9.3 1.5 1.0 7.1 5.6 1.2 1.0 4.1

× × × × × × × × × ×

103 104 102 102 105 104 102 102 104 103

7.4 × 10°

optimal working range was between pH 4.5 and 8.0. The highest DET currents were observed for DsCDH (22 μA cm−2) followed by CtCDH (5.0 μA cm−2), SbCDH (2.5 μA cm−2), HhCDH (2.2 μA cm−2), NcCDH (0.7 μA cm−2), and CaCDH (0.7 μA cm−2). However, when considering the purity of the CDH preparations, higher DET current density values for SbCDH (∼12.5 μA cm−2), HhCDH (∼3.5 μA cm−2), and NcCDH (∼2 μA cm−2) can be expected for pure enzyme preparations. In the presence of the soluble mediator 1,4benzoquinone the efficiency of the DET current between CYTCDH and the graphite electrode can be estimated and compared to the MET current from the DHCDH to the electrode. The ratios of MET:DET currents were calculated from the currents measured at the pH optimum of the DET current and are 14.3 for CaCDH (pH 6.0), 4.4 for CtCDH (pH 8.0), 1.8 for DsCDH (pH 5.0), 13.9 for HhCDH (pH 8.0), 5.8 for NcCDH (pH 5.0), and 7.0 for SbCDH (pH 5.0). The most efficient DET communication is thus found for DsCDH, CtCDH, and NcCDH. Heterogeneous Steady-State Kinetics. The three most interesting CDHs with DET properties, the ones with low MET:DET ratios and high observed currents, i.e., CtCDH, DsCDH, and HhCDH, were selected for further investigation. The steady-state kinetics of these selected modified enzyme electrodes for cellobiose, lactose, maltose, and glucose at the optimum pH for DET was obtained (Table 3, left columns). Similar to the homogeneous kinetics, cellobiose was the most efficiently oxidized substrate. However, compared to the KM measured in solution, the apparent KM values (KM,app) at the optimum pH increased by a factor of 70, 20, and 30 for CtCDH, DsCDH, and HhCD, respectively. The same was observed for lactose. The Imax values, comparable with the kcat values in homogeneous solution, were similar for both substrates. Again, for maltose greatly reduced Imax values, 10− 39% relative to cellobiose, were found; this is in good agreement with the results obtained in the homogeneous solution. The Imax values were higher for glucose, but not as high as the cellobiose Imax values observed when the CDHs were adsorbed onto the electrode. The three immobilized CDHs were also tested at physiological pH using PBS buffer (Table 3, right columns). The physiological pH environment had a positive impact on the DET efficiency of CtCDH, which resulted in 1.5-, 1.6-, 1.1-, and 1.3-fold higher current densities for cellobiose, lactose, maltose, and glucose, respectively. The KM,app values were altered as well. The KM,app for cellobiose was raised by a factor of 5, while the

molecule. Apparent catalytic constants for the disaccharides cellobiose (4-O(β-D-glucopyranosyl)-D-glucopyranose), lactose (4-O(β-D-galactopyranosyl)-D-glucopyranose), maltose (4-O(αD-glucopyranosyl)-D-glucopyranose), and the monosaccharide D(+)-glucose were determined with the one-electron acceptor cyt c (Table 2). The highest catalytic efficiency of all six CDHs was found for the natural substrate cellobiose, which is between 3.75 and 12 times higher than that for lactose. This difference is a result of the higher KM values for lactose since the kcat values of both substrates are very similar. In contrast, the unfavorable α-linkage of maltose was not only responsible for a KM value 800−4400 times higher than for cellobiose ranging from 2.4 to 31 mM, but it also reduced the kcat to 9−13% relative to cellobiose, except CtCDH (42%). The lowest catalytic efficiencies were observed for glucose, which can only be bound by either the binding (B-) site or the catalytic (C-) site of CDH. The KM values vary from 21 mM (CtCDH) to 770 mM (DsCDH). For CaCDH, DsCDH, and HhCDH substrate inhibition was observed. Contrary to maltose, the kcat values for glucose were almost as high as those for cellobiose. Pre-steadystate kinetic experiments were performed with CtCDH and cellobiose at pH 5.0 and 7.4. The highest turnover rate (kobs) of FAD reduction was observed with the highest applied cellobiose concentration (500 μM). It is 62 s−1 at pH 5.0 and 50 s−1 at pH 7.4. The turnover rate for heme b reduction is independent of the cellobiose concentration (except at 12.5 μM where the FAD reduction becomes rate limiting). At pH 5.0 kobs is 4 s−1 and at pH 7.4 kobs is 9 s−1. Heterogeneous pH Profiles. Heterogeneous pH profiles (enzyme adsorbed onto graphite electrodes) were measured using the MET (with 1,4-benzoquinone as redox mediator) and the DET approach. Figure 1 compares the pH profiles, i.e., homogeneous case using either cyt c or DCIP as electron acceptor and heterogeneous case using either a MET or a DET approach. The influence of the pH on the interaction between CYTCDH and the polarized graphite electrode was diverse. In case of CaCDH, DsCDH, NcCDH, and SbCDH the highest electron transfer rates were found around pH 5.0, while HhCDH and especially CtCDH exhibited the highest catalytic currents in the alkaline region, around pH 7.5. Also, the pH range in which at least half-maximal DET currents are reached varies between the species. On the one hand, CaCDH, DsCDH, and SbCDH show a narrow window between pH 4.5 and pH 6.5. On the other hand, CtCDH, HhCDH, and NcCDH show a broad DET range with remarkable currents a higher pH values. This effect is most pronounced in case of HhCDH, where the 6718


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Figure 2. Cyclic voltammograms (scan rate was 10 mV s−1) obtained with CtCDH (20 mg mL−1) in the absence (upper row) and presence (lower row) of 5 mM lactose with McIllvaine buffer pH 3.5, 5.5, and 7.5 on thioglycerol-modified gold electrodes.

KM,app for glucose was lowered by the same factor. Also, DsCDH generally showed lower KM,app values for the chosen substrates in the physiological buffer system. Current densities losses for cellobiose and lactose were due to the pH shifting within the expected range. Only the losses with maltose and glucose were lower than expected, with 80% and 60% of the current densities retained, respectively. The pH influence on HhCDH was not consistent; the KM,app values for cellobiose, lactose, and glucose were increased, despite lower current densities for the cellobiose measurement. No catalytic currents were measured for maltose. The pH dependence of the CtCYTCDH’s redox potential and DET current on a thioglycerol-modified gold electrode was investigated in presence and absence of lactose (Figure 2). The best communication between CYTCDH and the SAM-modified gold electrode was found at pH 7.5, where the peak separation of the CV was around 75 mV with a midpoint potential of 100 mV vs SHE. At pH 5.5, the oxidation and reduction peaks were separated by 120 mV and the reduction peak was not as clearly defined as at pH 7.5. At pH 3.5, the electron transfer between CYTCDH and the gold electrodes is poor, and the oxidation and reduction peaks are no longer visible. In the presence of lactose the behavior followed the same pattern. The highest catalytic currents were found at pH 7.5, with a 2-fold higher catalytic current than at pH 5.5. No catalytic current was observed at pH 3.5.

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suddenly decreases at pH 7.0. The previously studied CDH from M. thermophilum is an example of an acidic class II CDH.15,25 Intermediate CDHs have their optima also close to pH 5.0, but they show rather high DET currents up to pH 8.5. This behavior is most pronounced in the case of HhCDH, but also CaCDH, NcCDH, and SbCDH can deliver electrons efficiently over a broad pH range. Since the measured DET current is modulated by three sequential reactions, the pH dependency of the reaction and the electron transfer steps are discussed in the following order: (1) the catalytic step of carbohydrate oxidation at the FAD in the active site, (2) the intramolecular electron transfer (IET) step between FAD of the DHCDH and heme b of the CYTCDH, and (3) the DET step from CYTCDH to the electrode. pH Dependency of the Catalytic Step. The catalytic reaction in DHCDH is catalyzed by the FAD cofactor. We expect this reaction to be pH-dependent. For class I CDHs the pH optimum for the reductive half-reaction is acidic. Pre-steadystate investigations with class I Phanerochaete chrysosporium CDH16 show that FAD reduction is most efficient between pH 4.0 and 5.0 (kobs ∼ 60 s−1). At pH 7.0 the reduction rate decreased to a quarter of this value. However, the oxidation of carbohydrates can still be observed in the alkaline range, e.g., with Sclerotium rolfsii CDH at pH 9.0.19 The pre-steady-state experiments with the alkaline class II CtCDH in this work also indicate an acidic pH optimum for FAD reduction (kobs = 62 s−1 at pH 5.0), but it is still high at neutral/alkaline pH (kobs = 50 s−1 at pH 7.4). The obtained steady-state kinetic data for the other five investigated class II CDHs indicate a similar behavior. The interaction of the employed two-electron acceptors, DCIP and 1,4-benzoquinone with the FAD of the DHCDH, in the reductive half-reaction at neutral and alkaline pH proceeds fast, but it is difficult to discriminate its contribution from the oxidative half-reaction in steady-state turnover rates. The effect of pH on the oxidative half-reaction of the enzyme bound FAD by various electron acceptors is evident. For the partially

DISCUSSION

Harreither et al. recently classified ascomycetous CDHs based on their pH behavior in solution.22 This classification is also valid for the CDHs in this investigation and can be applied to the DET-related pH profiles. Alkaline class II CDHs, such as CtCDH and CDH from Humicola insolens,29 deliver electrons from the CYTCDH to the electrode most efficiently under alkaline pH conditions. For acidic class II CDHs, like DsCDH, DET efficiency reaches a maximum around pH 5.0 and 6719


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CtCDH showed also a quite efficient DET with a current density of 5.0 μA cm−2 and a MET:DET ratio of 1.9. A moderate to poor DET efficiency was found for HhCDH (2.2 μA cm−2, MET:DET = 10.0) and SbCDH (2.5 μA cm−2, MET:DET = 6.7), CaCDH (0.7 μA cm−2, MET:DET = 14.3), and NcCDH (0.7 μA cm−2, MET:DET = 5.8). For NcCDH and SbCDH the low purity of the preparation has to be considered. From the MET:DET ratios it is obvious that acidic DsCDH has the most efficient DET at pH 5.0 and that the MET:DET ratio is greatly reduced at pH 7.0 (MET:DET = 10). The alkaline CtCDH has a very low MET:DET ratio at its optimum DET pH 7.0−8.0, which becomes less favorable at pH 5.0 (MET:DET = 8.0). The intermediate class II CDHs HhCDH, NcCDH, and SbCDH show a high MET:DET ratio6−15 over a relatively broad pH range. The amount of adsorbed, active CDH on spectrographic graphite electrodes seems to differ considerably for the tested CDHs by investigation of the measured MET currents, which range from 3.3 to 45 μA cm−2 at pH 6.0. Of course, pH 6.0 is not the optimal pH for all six CDHs, and some CDH preparations are less pure than others. When taking the purity of the enzymes into account (by normalizing with the purity number), the recalculated currents are CaCDH 11 μA cm−2, CtCDH 18 μA cm−2, DsCDH 50 μA cm−2, HhCDH 38 μA cm−2, NcCDH 10 μA cm−2, and SbCDH 95 μA cm−2, which still differ considerably. To approximate the relative amount of absorbed CDH, it is assumed that the surface area of the electrode, the amount of protein applied to the electrodes (25 μg), and the mass transfer in the flow cell are equal for all CDH modified graphite electrodes. Thus, the specific activities of these CDHs in solution at pH 6.0 (taken from ref 22 and normalized by the respective purity number: CaCDH 17.7 U mg−1, CtCDH 12.5 U mg−1, DsCDH 14.4 U mg−1, HhCDH 10.2 U mg−1, NcCDH 24.7 U mg−1, and SbCDH 10.2 U mg−1) give the approximate substrate turnover per mg bound CDH. Whereas the specific activities differ only by a factor of 2.5, the current densities differ by a factor of 9.5. This indicates that more active CDH molecules are adsorbed on the graphite electrodes in case of SbCDH, HhCDH, and DsCDH. As a result, higher currents are measured despite their low specific activity. Contrary, CaCDH, CtCDH, and NcCDH show a lower binding efficiency although the enzyme samples would have a higher specific activity. Judged by these values, the surface coverage on the graphite electrode is different for CDHs. However, efficient binding of a CDH does not go hand in hand with high a DET efficiency as is demonstrated by the high MET:DET ratios (indicating high DET efficiency) for HhCDH and SbCDH. It is, however, observed that the DET pH profile of CDHs with a higher DET current fitted better to the data observed with cyt c (Figure 1). This indicates that DET is rate limiting for the less efficient CDHs (CaCDH, NcCDH) instead of IET being rate limiting for CDHs with efficient DET (DsCDH, CtCDH). The efficient DET observed at neutral and alkaline pH differentiates alkaline and intermediate class II CDHs from all known basidiomycetous class I CDHs.33,34 What is obvious from the results of this and previous studies is that the obtained current density depends very much on the particular applied CDH. So far, DsCDH exhibits the highest current densities not only among the new investigated CDHs, but among all investigated CDHs. It even supersedes the class I CDHs from P. sordida and T. villosa,33−36 which were regarded to exhibit the

negatively charged two-electron acceptor, DCIP, the pH optima fall between pH 5.0 and 6.0 and the relative activities at pH 7.0 are only 20−55%. For the uncharged two-electron acceptor, 1,4-benzoquinone, the pH optima were between 5.0 and 8.0. When compared to the pH classification for cyt c and DET, it can be seen that the pH optimum of the catalytic reaction does not strictly follow the classification: alkaline CtCDH (pH optimum for 1,4-benzoquinone: 8.0), acidic DsCDH (8.5), and CaCDH (6.5), intermediate HhCDH (8.0), NcCDH (8.0), and SbCDH (5.0). This stresses that the catalytic step in class II CDHs is (1) more efficient in slightly acidic or slightly alkaline pH and (2) is not the governing factor for the pH dependency associated with the DET and MET measurements. pH Dependency of the IET Step. It is known that the one-electron acceptor, cyt c, interacts solely with CYTCDH. The efficiency of IET between FAD of the DHCDH and heme b of the CYTCDH is indicative of the cyt c reduction rate. Therefore, it is used to determine the activity of the intact enzyme in fermentation broths, where proteolytic cleavage produces catalytically active DHCDH fragments.30 The two electrons stored in the FADH2 are transferred sequentially to the heme b via IET. In vivo, the heme b is presumably reoxidized by polysaccharide monoxygenases of the GH61 family.12,13 In vitro, the one-electron acceptor cyt c (12.4 kDa, pI = 10.5) or a polarized electrode can serve as a terminal electron acceptor. Since the bimolecular reduction rate of cyt c at CYTCDH is very fast (1.75 × 107 M−1 s−1,31 6.0 × 106 M−1 s−1 32), it is not rate limiting. Therefore, the turnover rate with cyt c can be used to measure the much slower IET. On the basis of pH-dependent activities measured with cyt c, it is known that the IET is strongly affected by pH. For example, in class I CDHs the IET is switched off at pH values above pH 6.0. This finding was confirmed with pre-steady-state experiments.16 The pHdependent activity of the six investigated class II CDHs varies from acidic to alkaline. This observation is most likely connected to varying pH conditions encountered by fungus during growth. To investigate whether the IET of CtCDH is alkaline as predicted, stopped-flow measurements at pH 7.4 and 5.0 were performed and IET rates of 9 and 4 s−1 were measured. The results from the steady-state and pre-steadystate experiments are comparable. The cyt c activity at pH 7.4 is about 2.6 times higher than that at pH 5.0, and the IET rate at pH 7.4 is about 2.25 times higher than that at pH 5.0. By measuring four different substrate concentrations in the stopped-flow apparatus, it was found that the rate of FAD reduction depends on the substrate concentration (a KM of 30 μM for cellobiose is calculated from the stopped-flow data). However, the IET rate is independent of the substrate concentration, as long as the catalytic rate of the FAD reduction is not the rate-limiting factor for IET. pH Dependency of the DET Step. The final electron transfer step, i.e. from the heme b of the CYTCDH to the electrode, is difficult to investigate on spectrographic graphite electrodes because of the unknown surface coverage by CDH. It is, however, possible to investigate the DET efficiency and compare it to the total bound catalytic activity. The total bound catalytic activity can be determined by using 1,4-benzoquinone as redox mediator and by contacting all the active sites of adsorbed and catalytically active CDH molecules by MET.33 Relative to the mediated activity, the most efficient DET was observed for DsCDH. It has a specific current density of 22 μA cm−2, which is only 1.7 times lower than the corresponding MET current density measured at the DET pH optimum. 6720


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improved for it to be competitive to existing mediated systems. One possible strategy that can be employed to increase the current density and optimize the electrode−enzyme interface is by the construction of 3D-nanostructured surfaces using carbon nanotubes (CNTs).34 Alternatively, the enzyme's orientation was much improved by immobilization on diazonium-activated CNTs.41 The use of carbon nanotube fibers,42 nanostructured graphite films,43 thiol-modified gold nanoparticles,39 deglycosylation of the enzyme,44 or the use of cross-linking agents45 has also been shown to be beneficial. The optimization of the DET characteristics of CYTCDH can be envisaged as a complementary strategy. The CDHs presented in this study vary strongly in their DET efficiencies and are currently studied to determine the key properties to achieve the highest DET rate on electrodes. From the varying substrate specificities of the investigated CDHsfrom class I like strictly cellodextrin/ lactose specific ones to class II with a much lower specificity the most suitable biocatalyst for biofuel cell anodes or biosensors can be selected or the protein scaffold used as template for further protein engineering.

most efficient with the highest DET currents among all published CDHs. Behavior on Thiol-Modified Gold Electrodes. By using gold electrodes, it was possible to observe DET currents in the absence (no catalytic turnover) and presence of substrate (turnover conditions) by cyclic voltammetry (CV). The midpoint potential of the CYTCDH for CtCDH was found to be 100 mV vs NHE at pH 7.5, which is in good agreement with the results obtained from carbon nanotube modified graphite electrodes.37 In comparison the acidic ascomycetous CDH from M. thermophilum exhibited a higher midpoint potential of 190 mV at pH 4.5 and 130 mV at pH 7.0.15 The alkaline H. insolens CDH showed a midpoint potential of 195 mV at pH 3.0 and 140 mV at pH 7.0.17 For basidiomycetous CDHs similar midpoint potentials were reported.16,17,36,38 It is obvious that the low potential of the CYTCDH of CtCDH makes it an interesting candidate as a biocatalyst for biofuel cell anodes.39 For the alkaline CtCDH the best communication between the CYTCDH and the electrode was found at pH 7.5, where the peak separation was only 75 mV, indicating a quasi-reversible process. At pH 5.5 the reversibility of the redox process was limited with a peak separation of almost 120 mV, and the nonturnover electrochemistry was totally untraceable at pH 3.5. An asymmetric peak separation was found at pH 5.5 compared to pH 7.5: while the oxidation peak was shifted by 10 mV, the reduction peak was shifted by 30 mV to a more negative potential. Using the thioglycerol-modified gold electrodes very high catalytic currents, ∼75 μA cm−2, were obtained at pH 7.5. This indicates a very efficient DET. A thiol with a hydroxy functionalized headgroup was chosen because it does not exhibit any change over the chosen pH range. Additionally, previous investigations performed using various thiols with different headgroups have shown that a hydroxy headgroup agrees very well with efficient DET.15,40 Application in Biosensors and Biofuel Cell Anodes. The applicability of CtCDH and DsCDH as biocatalysts for biofuel cell anodes has been previously demonstrated.20,21,39 These were the very first reports of glucose/O2-based membrane-, compartment-, and mediator-less enzymatic biofuel cells working with direct electrochemical communication between the enzymes and the electrodes for both the anode and the cathode. CtCDH is so far the only example of a CDH, which has both efficient IET and DET in the alkaline pH region. Humicola insolens CDH, which exhibits also an alkaline pH optimum with cyt c,18 was shown previously to give very low current densities in the presence of substrate.29 The less pronounced substrate specificity of class II CDHs, especially of CtCDH, allows the conversion of glucose in vivo. This is especially useful for biofuel cell anodes where substrate specificity is not of major concern. For a glucose biosensor a high selectivity for the analyte is however of utmost importance. The enhanced glucose turnover of CtCDH has stimulated initial studies for its use as a glucose recognition element in DET-based biosensors,37 but for the application in complex carbohydrate matrices the enzyme’s substrate specificity has to be further optimized by protein engineering. Especially CtCDH is a promising engineering template and might become an excellent alternative to glucose oxidase, and the various glucose dehydrogenases employed glucose biosensors. The limitations associated with the use of ascomycetous class II CDHs in bioelectronics has to be mentioned as well. Although the production of mediatorless devices is interesting, the performance of DET-based enzyme electrodes must be

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CONCLUSIONS Six novel ascomycete cellobiose dehydrogenases were characterized with respect to their substrate specificity as well as their IEF and DET efficiency. According to their cyt c and DETrelated pH profiles, these enzymes can be classified into three groups: acidic (DsCDH), intermediate (CaCDH, HhCDH, NcCDH, and SbCDH), and alkaline (CtCDH). The described CDHs present a variable toolbox for the building of engineered variants with improved properties for various electrode applications at neutral or alkaline pH.

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AUTHOR INFORMATION

Corresponding Author

*Phone +431 47654 6149; e-mail [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS Elvira Lackner, Magarita Keinrath, and Manfred Augustin are acknowledged for their superb technical support. The authors thank the following agencies for financial support: the Austrian Science Foundation (translational project FWF L395-B11), the University of Natural Resources and Life Sciences University (KUWI-travel grant to E. Lackner and M. Keinrath), the Swedish Research Council (project 621-2010-5031), and the European Commission (project “3D-Nanobiodevice” NMP4SL-2009-229255).

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ABBREVIATIONS CDH, cellobiose dehydrogenase; cyt c, cytochrome c; CYTCDH, cytochrome domain of CDH; DHCDH, dehydrogenase domain of CDH; DET, direct electron transfer; GH61, glycoside hydrolase 61; IET, intramolecular electron transfer; MET, mediated electron transfer.

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Investigation of the pH-Dependent Electron Transfer Mechanism of

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