- Email: [email protected]
Nafion modified glassy carbon electrodes
Electrochimica Acta 82 (2012) 214–217
Contents lists available at SciVerse ScienceDirect
Electrochimica Acta journal homepage: www.elsevier.com/locate/electacta
Ubiquinol-cytochrome c reductase (Complex III) electrochemistry at multi-walled carbon nanotubes/Nafion modified glassy carbon electrodes Lindsey N. Pelster a,b , Shelley D. Minteer a,b,∗ a b
Department of Chemistry, University of Utah, 315 S. 1400 E Rm 2020, Salt Lake City, UT 84112, United States Department of Chemistry, Saint Louis University, 3501 Laclede Ave, Saint Louis, MO 63103, United States
a r t i c l e
i n f o
Article history: Received 4 October 2011 Received in revised form 9 November 2011 Accepted 10 November 2011 Available online 14 February 2012 Keywords: Ubiquinol-cytochrome c reductase Electron transport chain Bioelectrocatalysis Cytochrome c
a b s t r a c t Electron transport chain complexes are critical to metabolism in living cells. Ubiquinol-cytochrome c reductase (Complex III) is responsible for carrying electrons from ubiquinol to cytochrome c, but the complex has not been evaluated electrochemically. This work details the bioelectrochemistry of ubiquinol-cytochrome c reductase of the electron transport chain of tuber mitochondria. The characterization of the electrochemistry of this enzyme is investigated in carboxylated multi-walled carbon nanotube/tetrabutyl ammonium bromide-modified Nafion® modified glassy carbon electrodes by cyclic voltammetry. Increasing concentrations of cytochrome c result in a catalytic response from the active enzyme in the nanotube sandwich. The experiments show that the enzyme followed Michaelis–Menten kinetics with a Km for the immobilized enzyme of 2.97 (±0.11) × 10−6 M and a Vmax of 6.31 (±0.82) × 10−3 mol min−1 at the electrode, but the Km and Vmax values decreased compared to the free enzyme in solution, which is expected for immobilized redox proteins. This is the first evidence of ubiquinol-cytochrome c reductase bioelectrocatalysis. © 2012 Elsevier Ltd. All rights reserved.
1. Introduction Mitochondria are considered the powerhouse of the living cell, because they contain several critical metabolic pathways including the electron transport chain. The electron transport chain is made up of five enzyme complexes that are primarily responsible for converting NADH to ATP, which is the biological energy currency in the living cell. The electron transport chain carries out several functions and is a source of a number of diseases in the mitochondria [1,2]. For this reason, the five enzyme complexes are of particular interest in fundamental bioelectrochemistry. The electron transport chain complexes shuttle electrons through the cytochrome c protein mediator and small molecule cofactors of NADH and ubiquinone, producing a proton gradient along the inner membrane of the mitochondria [3]. Ubiquinol-cytochrome c reductase, or Complex III (E.C. 1.10.2.2), of the electron transport chain, is a multi-subunit, membrane bound oxidoreductase protein that employs ubiquinol and a well-known redox protein, cytochrome c, to transfer electrons to neighboring complexes [4]. Cytochrome c is an important part of the chain that carries electrons from ubiquinol-cytochrome c reductase to cytochrome c oxidase, where it is oxidized and
∗ Corresponding author at: Department of Chemistry, University of Utah, 315 S. 1400 E Rm 2020, Salt Lake City, UT 84112, United States. Tel.: +1 801 587 8325; fax: +1 801 581 8181. E-mail address: [email protected] (S.D. Minteer). 0013-4686/$ – see front matter © 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.electacta.2011.11.121
takes part in the reaction of oxygen reduction to water. Ubiquinolcytochrome c reductase contains heme groups, like cytochrome c, that allow for electron transfer that can be detected electrochemically [4,5]. However, its electrochemistry has not been directly investigated. For this reason, this paper investigates the bioelectrochemistry of ubiquinol-cytochrome c reductase. Few have tried to characterize electron transport proteins, because of their transmembrane nature and importance of appropriate orientation on electrode surfaces. Since Complex III is a membrane protein, protein immobilization and stabilization is critical for the complex to maintain activity at the electrode surface. Enzyme immobilization and stabilization is an advanced field with a variety of different common methods for immobilization [6]. Several different methods have been developed to immobilize an active protein enzyme and maintain its activity at an electrode surface. These include: physical adsorption, incorporation into monolayers and bilayers, and entrapment into polymers. Physical adsorption normally leads to the denaturing of membrane proteins as well as protein leaching, so it is seldom used for kinetic analysis. On the other hand, one group has been able to electrochemically study an electron transport protein in a lipid bilayer on an electrode, cytochrome c oxidase [7]. Lipid bilayers are a replica of the inner membrane and help to stabilize membrane proteins in an environment that is similar to their cellular environment. Previous research in our group has demonstrated active protein immobilization in tetrabutylammonium bromide (TBAB) modified Nafion® at electrode surfaces [8–10] via both encapsulation and
L.N. Pelster, S.D. Minteer / Electrochimica Acta 82 (2012) 214–217
sandwich techniques. This polymer has successfully immobilized both membrane proteins and cytosolic proteins [10–12] due to the biocompatible amphiphilic environment that is formed. Glassy carbon electrodes are low surface area electrodes, so it is difficult to load sufficient electron transport chain complex onto the electrode for reproducible evaluation. For this reason, we added an additional component to the electrode to increase the effective surface area for protein loading. Wang and Davis have shown that the heme groups often have facile electron transport at carbon nanotube modified electrodes [5,13], so we have utilized multi-walled carbon nanotubes for improved loading of Complex III onto glassy carbon electrodes. In order to improve the surface area and electron transport, carbon nanotubes were incorporated onto the electrode followed by a layer of TBAB modified Nafion® membrane to form a stable protein modified electrode and then evaluated with cyclic voltammetry in the presence and absence of the substrate cytochrome c to understand the bioelectrochemistry of ubiquinol-cytochrome c reductase. 2. Experimental 2.1. Reagents All reagents purchased were used as received from Sigma–Aldrich, unless otherwise noted. All solutions were prepared with 18 M deionized water. 2.2. Preparation of enzyme Mitochondria was harvested by juicing organic tubers (potatoes) with a Kempo Green Power Juicer into 100 L of homogenization buffer containing: 2.6 M NaCl, 100 mM Tris–HCl pH 8.0, 50 mM EDTA, 0.4% bovine serum albumin (BSA), 26 mM dithiothreitol, and 0.1% l-cysteine. The juiced material was centrifuged at 4 ◦ C at 500 × g for 10 min, 800 × g for 10 min, and 5250 × g for 60 min. The supernatant was retained for the first two spins, discarding the starch pellets. The last pellet was used for the rest of the purification as it contained mitochondria and the supernatant was discarded. This protocol was modified from Scotti et al. [14]. All preparation, solubilization, and chromatography of the mitochondrial membranes was followed per the procedure of Weiss and Juchs omitting the sucrose density gradient step and gel filtration [15]. The active affinity fractions were concentrated with 100 kDa Amicon centrifugation filtration cells and combined for assays. Affigel 10® was used in place of CNBr-activated Sepharose® 4B for better retention of cytochrome c ligand through subsequent use of the column. The column media was washed with three volumes of 10 mM pH 4.5 sodium acetate buffer. The media was added to a flask with 0.1 M pH 8.0 sodium bicarbonate buffer containing 0.5 M NaCl and 25 mg/ml cytochrome c to a total of 250 mg cytochrome c for 10 ml of media. The mixture was mixed at 4 ◦ C for 4 h. A blocking agent of 100 l of 0.1 M ethanolamine HCl at pH 8 was added into the mixture and allowed to react at 4 ◦ C for 1 h. The media was packed into a column of 1.0 cm × 10.0 cm. The column was washed with a cycle of 1 M NaCl in 0.1 M pH 4 sodium acetate buffer and 1 M NaCl in 0.1 M pH 8 sodium borate buffer. Following elution of protein of interest, the wash buffers with 0.05% Triton X-100 and 1 mM K3 [Fe(CN)6 ] were used to clean the column, and the column was stored in 0.05 M Tris–acetate pH 7, 0.02 M NaCl, and 1 mM EDTA at 4 ◦ C until use.
215
pH 7.4 Tris–acetate buffer, 20 mM EDTA, 50 mM KCl, 5–100 M cytochrome c, and 15 M hydroquinone with 0.010 ml enzyme solution [16]. The change in absorbance for each sample was measured at room temperature over 2 min at 550 nm on a Shimadzu UV-VIS Spectrophotometer. Final specific activity of purified and concentrated Complex III was 1.82 g × ml−1 × mg−1 . For activity characterization, concentration additions of 5, 10, 15, 20, 25, 40, 50, 75, 100 M cytochrome c were added into reaction mixture for each of the assays. 2.4. Preparation of electrodes with enzyme Multi-walled carbon nanotubes (MWCNTs) were purchased from CheapTubes. The MWCNTs have an outer diameter of 20–30 nm, inner diameter of 5–10 nm, length of 10–30 m, a purity of >95 wt%, and carboxylated functional content of 1.23%. The multi-walled carbon nanotubes were sonicated and washed in 100 mM nitric acid and centrifuged at 5000 × g for 15 min. This surface preparation procedures was repeated twice. The nanotubes were washed with deionized water and centrifuged twice to remove all excess acid. The nanotubes were dried by vacuum desiccation overnight. 5 mg of washed COOH-MWCNTs, 50 l of ethanol and 100 l of deionized water were added together and sonicated for 30–45 min. 100 l of 10 mM pH 7.4 sodium phosphate buffer was added to the nanotubes and sonicated for 5 min. 50 l of purified and concentrated enzyme solution was added to the cooled nanotube mixture. 50 l of 10 mM pH 7.4 sodium phosphate buffer was added to a separate mixture instead of enzyme as a control electrode. The mixture was vortexed for 1 min to ensure a homogeneous mixture. CH Instruments glassy carbon electrodes (3 mm diameter) were polished with 1.0 m and 0.05 m Buehler alumina on Buehler polishing clothes and dried. The glassy carbon electrodes have a working area of 0.0707 cm2 (5% by wt). TBAB-modified Nafion® suspension was prepared as previously described in Moore et al. [11]. 10 l of enzyme/nanotube mixture was pipetted onto the top of the polished glassy carbon electrodes and dried under a small fan. After the mixture was completely dry on top of the electrode, 1.5 l of TBAB-modified Nafion® suspension was pipetted onto the top of the mixture and allowed to dry completely. The electrodes were soaked in degassed 10 mM pH 7.4 sodium phosphate buffer for an hour to equilibrate before electrochemical measurements were performed. 2.5. Characterization of electrodes The electrochemical cell setup included a Ag/AgCl (saturated KCl, CH Instruments) reference electrode and platinum mesh counter electrode. The working volume of the experiments remained constant at 5 ml of degassed solution. The electrochemical cell was kept under a nitrogen blanket throughout the experiments. Cyclic voltammetry was utilized to characterize the enzyme on the electrode with a CH Instruments potentiostat interfaced to a PC. A blank was run in the presence of buffer only, as well as control experiments without enzyme present. Oxidized cytochrome c from bovine heart was added in concentrations from 5 M to 50 M for enzyme bioelectrocatalysis experiments. Cyclic voltammetry scans were swept between 0.6 V to −0.6 V at a scan rate of 0.002 V/s, unless otherwise noted. During scan rate studies, the following scan rates were tested: 0.3 mV/s, 1 mV/s, 2 mV/s, and 10 mV/s in 25 M cytochrome c.
2.3. Characterization of the enzyme 3. Results and discussion For ubiquinol-cytochrome c reductase activity, a spectrophotometric assay was performed to study the change in oxidation state of cytochrome c. The reaction mixture consisted of 50 mM
The purification of the Complex III membrane protein followed previously published literature procedures with a few minor
216
L.N. Pelster, S.D. Minteer / Electrochimica Acta 82 (2012) 214–217
Fig. 1. (A) Enzyme activity assay results for free enzyme in solution with increasing substrate concentration. This plot shows a non-linear regression with an equation of y = (4.94 (±0.49) × 10−2 mol min−1 × [x])/(1.17 (±0.18) × 10−4 M + [x]) and R2 of 0.9934. (B) A linear Lineweaver–Burke plot produces an equation of y = (3.12 (±0.12) × 10−3 )x + (2.16 ± 6.16) and a R2 of 0.9906.
modifications described above [15]. Affi-gel® 10 was used in place of CNBr-Sepharose® 4B for better retention of cytochrome c after subsequent uses. The better retention allowed for more binding sites of cytochrome c substrate for the enzyme, therefore, providing more purified protein. Protein purity was confirmed with native PAGE electrophoresis. A spectroscopic activity assay evaluating the redox state of cytochrome c as a function of time was employed to study the kinetics of the purified enzyme. Fig. 1A shows the spectroscopic data as a function of concentration of cytochrome c. The final specific activity is 1.82 mol min−1 mg−1 . Nonlinear regression was employed to calculate the catalytic parameters for the free enzyme in solution. Km was determined to be 1.17 (±0.18) × 10−4 M and Vmax was determined to be 4.94 (±0.49) × 10−2 mol min−1 for the purified protein in solution. These values show that the Km , or affinity, for the substrate in solution is lower for tubers compared to published Km value of 1.3 × 10−5 M for bovine complex III [17]. Cyclic voltammetry was employed to investigate the electrochemical activity of the oxidoreductase enzyme with changing substrate (cytochrome c) concentrations. Literature has shown that cytochrome c is one of the most studied electrochemically active proteins [5,13,18]. The fact that the substrate is electrochemically active in the same potential window as the enzyme complicates the electrochemistry. It can be assumed that the enzyme is active by the comparison of data with and without enzyme and in the presence of cytochrome c. The increasing concentrations
Fig. 2. (A) Electrochemical cytochrome c reduction studies shows Michaelis–Menten kinetics with the enzyme immobilized on the electrode in a MWCNT/TBAB modified Nafion sandwich. This plot shows a non-linear regression with an equation of y = (6.31 (±0.04) × 10−3 mol min−1 × [x])/(2.97 (±0.11) × 10−6 M + [x]) and R2 of 0.9971. Cyclic voltammetry without enzyme produces a linear response with concentration and an equation of y = (0.085 ± 0.002)x + (4.15 ± 0.04) and R2 of 0.9975. (B) A linear Lineweaver–Burke plot produces an equation of y = (0.29 ± 0.002)x + (98642 ± 311) and R2 of 0.9982.
show catalytic response of enzyme activity rather than just the electrochemical reduction peaks of cytochrome c. Fig. 2A shows the difference between electrode preparations with and without enzyme. The electrochemical reduction of cytochrome c is shown to be linear with concentration as expected for a redox species diffusing in a polymer film to an electrode surface, where the catalytic reduction follows enzymatic behavior with communication to the electrode by the reduced substrate. The enzyme follows Michaelis–Menten kinetics while on the electrode. The Km for the immobilized enzyme is 2.97 (±0.11) × 10−6 M. Shelimov and Jarrold found that cytochrome c has a charge at pH 7 of +5 in reduced form and +6 in the oxidized form [19], which means that cytochrome c may preconcentrate in the Nafion membrane, since it is a cation exchange membrane. The restricted containment of the enzyme in the polymer and the extraction/pre-concentration of the substrate into the polymer enables for a tighter affinity of the enzyme to the substrate. The Vmax of the immobilized enzyme is 6.31 (±0.82) × 10−3 mol min−1 at the electrode. Both Vmax and Km are lower for the immobilized enzyme than the free enzyme in solution. Fig. 3 displays the representative cyclic voltammetric scans of the enzyme electrodes with increasing substrate concentrations. The first scan without cytochrome c shows the electrode in
L.N. Pelster, S.D. Minteer / Electrochimica Acta 82 (2012) 214–217
217
Table 1 Peak reduction potentials versus Ag|AgCl reference electrode as a function of concentration of cytochrome c.
Fig. 3. Representative cyclic voltammograms of ubiquinol-cytochrome c reductase in MWCNT/TBAB Nafion on glassy carbon electrodes with increasing cytochrome c concentrations (scan rate 2 mV/s).
[Cytochrome c]/mol L−1
Epc /V (versus Ag|AgCl)
5 10 15 25 40 50
−0.082 −0.081 −0.079 −0.083 −0.088 −0.080
± ± ± ± ± ±
0.019 0.004 0.017 0.027 0.033 0.041
1.33 (±0.37) × 10−8 moles of enzyme per cm2 of glassy carbon electrode area was found to be loaded and electrochemically accessible. Table 1 displays the reduction potentials for cytochrome c on the electrode with the enzyme, showing very little shift between concentrations of the substrate. Scan rate studies were performed on these ubiquinolcytochrome c reductase modified glassy carbon electrodes to understand if they were under mass transport control or surface confined behavior. When peak current was plotted versus square root of scan rate, there was not a linear correlation (R2 = 0.9783). However, when peak current was plotted versus scan rate, there was a linear relationship (R2 = 0.9942). This linear relationship between peak current and scan rate demonstrates thin layer voltammetry, as shown in Fig. 4. 4. Conclusions Ubiquinol-cytochrome c reductase is an oxidoreductase enzyme that resides in the inner membrane of mitochondria as part of the electron transport chain. This is the first time that this enzyme has been electrochemically characterized, showing a catalytic response with cytochrome c substrate at the electrode and an ability for direct electron transfer to carbon electrodes. A multi-walled carbon nanotube/TBAB modified Nafion® layered electrode structure was used to immobilize the enzyme complex at a glassy carbon electrode for kinetic evaluation. This research provides a new way to study the electron transport chain electrochemically. Acknowledgement The authors would like to thank the Air Force Research Laboratory for funding this project. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11]
Fig. 4. (A) Representative cyclic voltammograms of increasing scan rates with enzymatic electrodes. (B) Linear relationship between peak current and scan rate produces an equation of y = (1.45 × 10−6 )x + 4.30 × 10−6 with a R2 of 0.9942.
buffer, and the characteristics and capacitance of the MWCNTs. It is important to note that this cyclic voltammogram shows the ability Complex III to do direct electron transfer with the electrode. Using these results to determine enzyme loading, an average of
[12] [13] [14] [15] [16] [17] [18] [19]
S. DiMauro, Biochim. Biophys. Acta 1658 (2004) 80. S. DiMauro, E.A. Schon, N. Engl. J. Med. 348 (2003) 2656. J.N. Siedow, A.L. Umbach, Plant Cell 7 (1995) 821. E.A. Berry, L.S. Huang, V.J. DeRose, J. Biol. Chem. 266 (1991) 9064. J. Wang, M. Li, Z. Shi, N. Li, Z. Gu, Anal. Chem. 74 (2002) 1993. M.J. Cooney, V. Svoboda, C. Lau, G.P. Martin, S.D. Minteer, Energy Environ. Sci. 1 (2008) 320. J.D. Burgess, M.C. Rhoten, F.M. Hawkridge, Langmuir 14 (1998) 2467. S.D. Minteer, S. Topcagic, B. Treu, in: U.P.T. Office (Ed.), Immobilized Enzymes in Biocathodes, St. Louis University, U.S., 2005. B.L. Treu, S.D. Minteer, Bioelectrochemistry 74 (2008) 73. B.L. Treu, R.L. Arechederra, S.D. Minteer, J. Nanosci. Nanotechnol. 9 (2009) 2374. C.M. Moore, N.L. Akers, A.D. Hill, Z.C. Johnson, S.D. Minteer, Biomacromolecules 5 (2004) 1241. R.L. Arechederra, B.L. Treu, S.D. Minteer, J. Power Sources 173 (2007) 156. J.J. Davis, R.J. Coles, H. Allen, O. Hill, J. Electroanal. Chem. 440 (1997) 279. N. Scotti, T. Cardi, L. Marechal-Drouard, Plant Mol. Biol. Report. 19 (2001) 67a. H. Weiss, B. Juchs, Eur. J. Biochem. 88 (1978) 17. P. Linke, H. Weiss, Methods Enzymol. 126 (1986) 201. W.D. Engel, H. Schagger, G. von Jagow, Biochim. Biophys. Acta 592 (1980) 211. M.J. Eddowes, H.A.O. Hill, J. Chem. Soc. Chem. Commun. (1977) 771. K.B. Shelimov, M.F. Jarrold, J. Am. Chem. Soc. 118 (1996) 10313.
Contents lists available at SciVerse ScienceDirect
Electrochimica Acta journal homepage: www.elsevier.com/locate/electacta
Ubiquinol-cytochrome c reductase (Complex III) electrochemistry at multi-walled carbon nanotubes/Nafion modified glassy carbon electrodes Lindsey N. Pelster a,b , Shelley D. Minteer a,b,∗ a b
Department of Chemistry, University of Utah, 315 S. 1400 E Rm 2020, Salt Lake City, UT 84112, United States Department of Chemistry, Saint Louis University, 3501 Laclede Ave, Saint Louis, MO 63103, United States
a r t i c l e
i n f o
Article history: Received 4 October 2011 Received in revised form 9 November 2011 Accepted 10 November 2011 Available online 14 February 2012 Keywords: Ubiquinol-cytochrome c reductase Electron transport chain Bioelectrocatalysis Cytochrome c
a b s t r a c t Electron transport chain complexes are critical to metabolism in living cells. Ubiquinol-cytochrome c reductase (Complex III) is responsible for carrying electrons from ubiquinol to cytochrome c, but the complex has not been evaluated electrochemically. This work details the bioelectrochemistry of ubiquinol-cytochrome c reductase of the electron transport chain of tuber mitochondria. The characterization of the electrochemistry of this enzyme is investigated in carboxylated multi-walled carbon nanotube/tetrabutyl ammonium bromide-modified Nafion® modified glassy carbon electrodes by cyclic voltammetry. Increasing concentrations of cytochrome c result in a catalytic response from the active enzyme in the nanotube sandwich. The experiments show that the enzyme followed Michaelis–Menten kinetics with a Km for the immobilized enzyme of 2.97 (±0.11) × 10−6 M and a Vmax of 6.31 (±0.82) × 10−3 mol min−1 at the electrode, but the Km and Vmax values decreased compared to the free enzyme in solution, which is expected for immobilized redox proteins. This is the first evidence of ubiquinol-cytochrome c reductase bioelectrocatalysis. © 2012 Elsevier Ltd. All rights reserved.
1. Introduction Mitochondria are considered the powerhouse of the living cell, because they contain several critical metabolic pathways including the electron transport chain. The electron transport chain is made up of five enzyme complexes that are primarily responsible for converting NADH to ATP, which is the biological energy currency in the living cell. The electron transport chain carries out several functions and is a source of a number of diseases in the mitochondria [1,2]. For this reason, the five enzyme complexes are of particular interest in fundamental bioelectrochemistry. The electron transport chain complexes shuttle electrons through the cytochrome c protein mediator and small molecule cofactors of NADH and ubiquinone, producing a proton gradient along the inner membrane of the mitochondria [3]. Ubiquinol-cytochrome c reductase, or Complex III (E.C. 1.10.2.2), of the electron transport chain, is a multi-subunit, membrane bound oxidoreductase protein that employs ubiquinol and a well-known redox protein, cytochrome c, to transfer electrons to neighboring complexes [4]. Cytochrome c is an important part of the chain that carries electrons from ubiquinol-cytochrome c reductase to cytochrome c oxidase, where it is oxidized and
∗ Corresponding author at: Department of Chemistry, University of Utah, 315 S. 1400 E Rm 2020, Salt Lake City, UT 84112, United States. Tel.: +1 801 587 8325; fax: +1 801 581 8181. E-mail address: [email protected] (S.D. Minteer). 0013-4686/$ – see front matter © 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.electacta.2011.11.121
takes part in the reaction of oxygen reduction to water. Ubiquinolcytochrome c reductase contains heme groups, like cytochrome c, that allow for electron transfer that can be detected electrochemically [4,5]. However, its electrochemistry has not been directly investigated. For this reason, this paper investigates the bioelectrochemistry of ubiquinol-cytochrome c reductase. Few have tried to characterize electron transport proteins, because of their transmembrane nature and importance of appropriate orientation on electrode surfaces. Since Complex III is a membrane protein, protein immobilization and stabilization is critical for the complex to maintain activity at the electrode surface. Enzyme immobilization and stabilization is an advanced field with a variety of different common methods for immobilization [6]. Several different methods have been developed to immobilize an active protein enzyme and maintain its activity at an electrode surface. These include: physical adsorption, incorporation into monolayers and bilayers, and entrapment into polymers. Physical adsorption normally leads to the denaturing of membrane proteins as well as protein leaching, so it is seldom used for kinetic analysis. On the other hand, one group has been able to electrochemically study an electron transport protein in a lipid bilayer on an electrode, cytochrome c oxidase [7]. Lipid bilayers are a replica of the inner membrane and help to stabilize membrane proteins in an environment that is similar to their cellular environment. Previous research in our group has demonstrated active protein immobilization in tetrabutylammonium bromide (TBAB) modified Nafion® at electrode surfaces [8–10] via both encapsulation and
L.N. Pelster, S.D. Minteer / Electrochimica Acta 82 (2012) 214–217
sandwich techniques. This polymer has successfully immobilized both membrane proteins and cytosolic proteins [10–12] due to the biocompatible amphiphilic environment that is formed. Glassy carbon electrodes are low surface area electrodes, so it is difficult to load sufficient electron transport chain complex onto the electrode for reproducible evaluation. For this reason, we added an additional component to the electrode to increase the effective surface area for protein loading. Wang and Davis have shown that the heme groups often have facile electron transport at carbon nanotube modified electrodes [5,13], so we have utilized multi-walled carbon nanotubes for improved loading of Complex III onto glassy carbon electrodes. In order to improve the surface area and electron transport, carbon nanotubes were incorporated onto the electrode followed by a layer of TBAB modified Nafion® membrane to form a stable protein modified electrode and then evaluated with cyclic voltammetry in the presence and absence of the substrate cytochrome c to understand the bioelectrochemistry of ubiquinol-cytochrome c reductase. 2. Experimental 2.1. Reagents All reagents purchased were used as received from Sigma–Aldrich, unless otherwise noted. All solutions were prepared with 18 M deionized water. 2.2. Preparation of enzyme Mitochondria was harvested by juicing organic tubers (potatoes) with a Kempo Green Power Juicer into 100 L of homogenization buffer containing: 2.6 M NaCl, 100 mM Tris–HCl pH 8.0, 50 mM EDTA, 0.4% bovine serum albumin (BSA), 26 mM dithiothreitol, and 0.1% l-cysteine. The juiced material was centrifuged at 4 ◦ C at 500 × g for 10 min, 800 × g for 10 min, and 5250 × g for 60 min. The supernatant was retained for the first two spins, discarding the starch pellets. The last pellet was used for the rest of the purification as it contained mitochondria and the supernatant was discarded. This protocol was modified from Scotti et al. [14]. All preparation, solubilization, and chromatography of the mitochondrial membranes was followed per the procedure of Weiss and Juchs omitting the sucrose density gradient step and gel filtration [15]. The active affinity fractions were concentrated with 100 kDa Amicon centrifugation filtration cells and combined for assays. Affigel 10® was used in place of CNBr-activated Sepharose® 4B for better retention of cytochrome c ligand through subsequent use of the column. The column media was washed with three volumes of 10 mM pH 4.5 sodium acetate buffer. The media was added to a flask with 0.1 M pH 8.0 sodium bicarbonate buffer containing 0.5 M NaCl and 25 mg/ml cytochrome c to a total of 250 mg cytochrome c for 10 ml of media. The mixture was mixed at 4 ◦ C for 4 h. A blocking agent of 100 l of 0.1 M ethanolamine HCl at pH 8 was added into the mixture and allowed to react at 4 ◦ C for 1 h. The media was packed into a column of 1.0 cm × 10.0 cm. The column was washed with a cycle of 1 M NaCl in 0.1 M pH 4 sodium acetate buffer and 1 M NaCl in 0.1 M pH 8 sodium borate buffer. Following elution of protein of interest, the wash buffers with 0.05% Triton X-100 and 1 mM K3 [Fe(CN)6 ] were used to clean the column, and the column was stored in 0.05 M Tris–acetate pH 7, 0.02 M NaCl, and 1 mM EDTA at 4 ◦ C until use.
215
pH 7.4 Tris–acetate buffer, 20 mM EDTA, 50 mM KCl, 5–100 M cytochrome c, and 15 M hydroquinone with 0.010 ml enzyme solution [16]. The change in absorbance for each sample was measured at room temperature over 2 min at 550 nm on a Shimadzu UV-VIS Spectrophotometer. Final specific activity of purified and concentrated Complex III was 1.82 g × ml−1 × mg−1 . For activity characterization, concentration additions of 5, 10, 15, 20, 25, 40, 50, 75, 100 M cytochrome c were added into reaction mixture for each of the assays. 2.4. Preparation of electrodes with enzyme Multi-walled carbon nanotubes (MWCNTs) were purchased from CheapTubes. The MWCNTs have an outer diameter of 20–30 nm, inner diameter of 5–10 nm, length of 10–30 m, a purity of >95 wt%, and carboxylated functional content of 1.23%. The multi-walled carbon nanotubes were sonicated and washed in 100 mM nitric acid and centrifuged at 5000 × g for 15 min. This surface preparation procedures was repeated twice. The nanotubes were washed with deionized water and centrifuged twice to remove all excess acid. The nanotubes were dried by vacuum desiccation overnight. 5 mg of washed COOH-MWCNTs, 50 l of ethanol and 100 l of deionized water were added together and sonicated for 30–45 min. 100 l of 10 mM pH 7.4 sodium phosphate buffer was added to the nanotubes and sonicated for 5 min. 50 l of purified and concentrated enzyme solution was added to the cooled nanotube mixture. 50 l of 10 mM pH 7.4 sodium phosphate buffer was added to a separate mixture instead of enzyme as a control electrode. The mixture was vortexed for 1 min to ensure a homogeneous mixture. CH Instruments glassy carbon electrodes (3 mm diameter) were polished with 1.0 m and 0.05 m Buehler alumina on Buehler polishing clothes and dried. The glassy carbon electrodes have a working area of 0.0707 cm2 (5% by wt). TBAB-modified Nafion® suspension was prepared as previously described in Moore et al. [11]. 10 l of enzyme/nanotube mixture was pipetted onto the top of the polished glassy carbon electrodes and dried under a small fan. After the mixture was completely dry on top of the electrode, 1.5 l of TBAB-modified Nafion® suspension was pipetted onto the top of the mixture and allowed to dry completely. The electrodes were soaked in degassed 10 mM pH 7.4 sodium phosphate buffer for an hour to equilibrate before electrochemical measurements were performed. 2.5. Characterization of electrodes The electrochemical cell setup included a Ag/AgCl (saturated KCl, CH Instruments) reference electrode and platinum mesh counter electrode. The working volume of the experiments remained constant at 5 ml of degassed solution. The electrochemical cell was kept under a nitrogen blanket throughout the experiments. Cyclic voltammetry was utilized to characterize the enzyme on the electrode with a CH Instruments potentiostat interfaced to a PC. A blank was run in the presence of buffer only, as well as control experiments without enzyme present. Oxidized cytochrome c from bovine heart was added in concentrations from 5 M to 50 M for enzyme bioelectrocatalysis experiments. Cyclic voltammetry scans were swept between 0.6 V to −0.6 V at a scan rate of 0.002 V/s, unless otherwise noted. During scan rate studies, the following scan rates were tested: 0.3 mV/s, 1 mV/s, 2 mV/s, and 10 mV/s in 25 M cytochrome c.
2.3. Characterization of the enzyme 3. Results and discussion For ubiquinol-cytochrome c reductase activity, a spectrophotometric assay was performed to study the change in oxidation state of cytochrome c. The reaction mixture consisted of 50 mM
The purification of the Complex III membrane protein followed previously published literature procedures with a few minor
216
L.N. Pelster, S.D. Minteer / Electrochimica Acta 82 (2012) 214–217
Fig. 1. (A) Enzyme activity assay results for free enzyme in solution with increasing substrate concentration. This plot shows a non-linear regression with an equation of y = (4.94 (±0.49) × 10−2 mol min−1 × [x])/(1.17 (±0.18) × 10−4 M + [x]) and R2 of 0.9934. (B) A linear Lineweaver–Burke plot produces an equation of y = (3.12 (±0.12) × 10−3 )x + (2.16 ± 6.16) and a R2 of 0.9906.
modifications described above [15]. Affi-gel® 10 was used in place of CNBr-Sepharose® 4B for better retention of cytochrome c after subsequent uses. The better retention allowed for more binding sites of cytochrome c substrate for the enzyme, therefore, providing more purified protein. Protein purity was confirmed with native PAGE electrophoresis. A spectroscopic activity assay evaluating the redox state of cytochrome c as a function of time was employed to study the kinetics of the purified enzyme. Fig. 1A shows the spectroscopic data as a function of concentration of cytochrome c. The final specific activity is 1.82 mol min−1 mg−1 . Nonlinear regression was employed to calculate the catalytic parameters for the free enzyme in solution. Km was determined to be 1.17 (±0.18) × 10−4 M and Vmax was determined to be 4.94 (±0.49) × 10−2 mol min−1 for the purified protein in solution. These values show that the Km , or affinity, for the substrate in solution is lower for tubers compared to published Km value of 1.3 × 10−5 M for bovine complex III [17]. Cyclic voltammetry was employed to investigate the electrochemical activity of the oxidoreductase enzyme with changing substrate (cytochrome c) concentrations. Literature has shown that cytochrome c is one of the most studied electrochemically active proteins [5,13,18]. The fact that the substrate is electrochemically active in the same potential window as the enzyme complicates the electrochemistry. It can be assumed that the enzyme is active by the comparison of data with and without enzyme and in the presence of cytochrome c. The increasing concentrations
Fig. 2. (A) Electrochemical cytochrome c reduction studies shows Michaelis–Menten kinetics with the enzyme immobilized on the electrode in a MWCNT/TBAB modified Nafion sandwich. This plot shows a non-linear regression with an equation of y = (6.31 (±0.04) × 10−3 mol min−1 × [x])/(2.97 (±0.11) × 10−6 M + [x]) and R2 of 0.9971. Cyclic voltammetry without enzyme produces a linear response with concentration and an equation of y = (0.085 ± 0.002)x + (4.15 ± 0.04) and R2 of 0.9975. (B) A linear Lineweaver–Burke plot produces an equation of y = (0.29 ± 0.002)x + (98642 ± 311) and R2 of 0.9982.
show catalytic response of enzyme activity rather than just the electrochemical reduction peaks of cytochrome c. Fig. 2A shows the difference between electrode preparations with and without enzyme. The electrochemical reduction of cytochrome c is shown to be linear with concentration as expected for a redox species diffusing in a polymer film to an electrode surface, where the catalytic reduction follows enzymatic behavior with communication to the electrode by the reduced substrate. The enzyme follows Michaelis–Menten kinetics while on the electrode. The Km for the immobilized enzyme is 2.97 (±0.11) × 10−6 M. Shelimov and Jarrold found that cytochrome c has a charge at pH 7 of +5 in reduced form and +6 in the oxidized form [19], which means that cytochrome c may preconcentrate in the Nafion membrane, since it is a cation exchange membrane. The restricted containment of the enzyme in the polymer and the extraction/pre-concentration of the substrate into the polymer enables for a tighter affinity of the enzyme to the substrate. The Vmax of the immobilized enzyme is 6.31 (±0.82) × 10−3 mol min−1 at the electrode. Both Vmax and Km are lower for the immobilized enzyme than the free enzyme in solution. Fig. 3 displays the representative cyclic voltammetric scans of the enzyme electrodes with increasing substrate concentrations. The first scan without cytochrome c shows the electrode in
L.N. Pelster, S.D. Minteer / Electrochimica Acta 82 (2012) 214–217
217
Table 1 Peak reduction potentials versus Ag|AgCl reference electrode as a function of concentration of cytochrome c.
Fig. 3. Representative cyclic voltammograms of ubiquinol-cytochrome c reductase in MWCNT/TBAB Nafion on glassy carbon electrodes with increasing cytochrome c concentrations (scan rate 2 mV/s).
[Cytochrome c]/mol L−1
Epc /V (versus Ag|AgCl)
5 10 15 25 40 50
−0.082 −0.081 −0.079 −0.083 −0.088 −0.080
± ± ± ± ± ±
0.019 0.004 0.017 0.027 0.033 0.041
1.33 (±0.37) × 10−8 moles of enzyme per cm2 of glassy carbon electrode area was found to be loaded and electrochemically accessible. Table 1 displays the reduction potentials for cytochrome c on the electrode with the enzyme, showing very little shift between concentrations of the substrate. Scan rate studies were performed on these ubiquinolcytochrome c reductase modified glassy carbon electrodes to understand if they were under mass transport control or surface confined behavior. When peak current was plotted versus square root of scan rate, there was not a linear correlation (R2 = 0.9783). However, when peak current was plotted versus scan rate, there was a linear relationship (R2 = 0.9942). This linear relationship between peak current and scan rate demonstrates thin layer voltammetry, as shown in Fig. 4. 4. Conclusions Ubiquinol-cytochrome c reductase is an oxidoreductase enzyme that resides in the inner membrane of mitochondria as part of the electron transport chain. This is the first time that this enzyme has been electrochemically characterized, showing a catalytic response with cytochrome c substrate at the electrode and an ability for direct electron transfer to carbon electrodes. A multi-walled carbon nanotube/TBAB modified Nafion® layered electrode structure was used to immobilize the enzyme complex at a glassy carbon electrode for kinetic evaluation. This research provides a new way to study the electron transport chain electrochemically. Acknowledgement The authors would like to thank the Air Force Research Laboratory for funding this project. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11]
Fig. 4. (A) Representative cyclic voltammograms of increasing scan rates with enzymatic electrodes. (B) Linear relationship between peak current and scan rate produces an equation of y = (1.45 × 10−6 )x + 4.30 × 10−6 with a R2 of 0.9942.
buffer, and the characteristics and capacitance of the MWCNTs. It is important to note that this cyclic voltammogram shows the ability Complex III to do direct electron transfer with the electrode. Using these results to determine enzyme loading, an average of
[12] [13] [14] [15] [16] [17] [18] [19]
S. DiMauro, Biochim. Biophys. Acta 1658 (2004) 80. S. DiMauro, E.A. Schon, N. Engl. J. Med. 348 (2003) 2656. J.N. Siedow, A.L. Umbach, Plant Cell 7 (1995) 821. E.A. Berry, L.S. Huang, V.J. DeRose, J. Biol. Chem. 266 (1991) 9064. J. Wang, M. Li, Z. Shi, N. Li, Z. Gu, Anal. Chem. 74 (2002) 1993. M.J. Cooney, V. Svoboda, C. Lau, G.P. Martin, S.D. Minteer, Energy Environ. Sci. 1 (2008) 320. J.D. Burgess, M.C. Rhoten, F.M. Hawkridge, Langmuir 14 (1998) 2467. S.D. Minteer, S. Topcagic, B. Treu, in: U.P.T. Office (Ed.), Immobilized Enzymes in Biocathodes, St. Louis University, U.S., 2005. B.L. Treu, S.D. Minteer, Bioelectrochemistry 74 (2008) 73. B.L. Treu, R.L. Arechederra, S.D. Minteer, J. Nanosci. Nanotechnol. 9 (2009) 2374. C.M. Moore, N.L. Akers, A.D. Hill, Z.C. Johnson, S.D. Minteer, Biomacromolecules 5 (2004) 1241. R.L. Arechederra, B.L. Treu, S.D. Minteer, J. Power Sources 173 (2007) 156. J.J. Davis, R.J. Coles, H. Allen, O. Hill, J. Electroanal. Chem. 440 (1997) 279. N. Scotti, T. Cardi, L. Marechal-Drouard, Plant Mol. Biol. Report. 19 (2001) 67a. H. Weiss, B. Juchs, Eur. J. Biochem. 88 (1978) 17. P. Linke, H. Weiss, Methods Enzymol. 126 (1986) 201. W.D. Engel, H. Schagger, G. von Jagow, Biochim. Biophys. Acta 592 (1980) 211. M.J. Eddowes, H.A.O. Hill, J. Chem. Soc. Chem. Commun. (1977) 771. K.B. Shelimov, M.F. Jarrold, J. Am. Chem. Soc. 118 (1996) 10313.