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Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades
BBABIO-47518; No. of pages: 4; 4C: 3, 4 Biochimica et Biophysica Acta xxx (2015) xxx–xxx
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Biochimica et Biophysica Acta journal homepage: www.elsevier.com/locate/bbabio
Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades Shelley D. Minteer Departments of Chemistry and Materials Science & Engineering, University of Utah, 315 S 1400 E Rm 2020, Salt Lake City, UT 84112, USA
a r t i c l e
i n f o
Article history: Received 2 May 2015 Accepted 20 August 2015 Available online xxxx Keywords: Metabolic pathway Bioelectrocatalysis Metabolon Enzyme cascade Biofuel cell
a b s t r a c t Anodic bioelectrodes for biofuel cells are more complex than cathodic bioelectrodes for biofuel cells, because laccase and bilirubin oxidase can individually catalyze four electron reduction of oxygen to water, whereas most anodic enzymes only do a single two electron oxidation of a complex fuel (i.e. glucose oxidase oxidizing glucose to gluconolactone while generating 2 electrons of the total 24 electrons), so enzyme cascades are typically needed for complete oxidation of the fuel. This review article will discuss the lessons learned from natural metabolic pathways about multi-step oxidation and how those lessons have been applied to minimal or artificial enzyme cascades. © 2015 Published by Elsevier B.V.
1. Introduction Enzymatic fuel cells are a type of bioelectronic device focused on bioenergetic conversion of chemical energy to electrical energy via the utilization of enzymes as electrocatalysts on the anode and/or the cathode. They were first discovered in the 1960s when a glucose oxidase anode was combined with a platinum cathode to produce a positive open circuit potential [1]. That early work invigorated a field that was primarily focused on harnessing electrical energy from biofuels present in living organisms (i.e. glucose in the blood stream) [2–5]. However, as the field of enzymatic fuel cells (often called biofuel cells) advanced, it became clear that there were alternative applications for enzymatic fuel cells. As stability and power density/current density performance increased, there became an interest in biofuel cells for portable power applications [6]. This interest spurred engineering of anodes from the original single oxidoreductase enzyme systems to cascades of enzymes responsible for catalyzing sequential reactions. This started with early work by Palmore and Whitesides, which incorporated the three enzyme cascade (NAD-dependent alcohol dehydrogenase, NAD-dependent aldehyde dehydrogenase, and NAD-dependent formate dehydrogenase) into the bioanode compartment of a methanol/oxygen biofuel cell [7]. This work showed the increase in performance observed with deeper degrees of oxidation of the fuel and postulated an increase in efficiency that would lead to increased energy density of biofuel cells/biobatteries. This early inspiration in enzyme cascades has led to a variety of research efforts in enzyme cascades that will be discussed below. They include
E-mail address: [email protected].
natural metabolic pathways, natural metabolons, synthetic metabolons, and minimal enzyme cascades. 2. Natural metabolic pathways Metabolic pathways are cascades of enzymes responsible for energy conversion in the living organism (oxidation of food/fuel or production of energetic molecules in the cell). These pathways include the glycolytic pathway, the Krebs cycle, the pentose phosphate pathways, fatty acid metabolism, amino sugar metabolism, purine biosynthesis, amino acid synthesis, and sucrose/starch metabolism. These pathways are responsible for catalyzing sequential reactions and therefore have been of interest to the field of bioelectrochemistry, since Suzuki and coworkers first used invertase and glucose oxidase to electrochemically monitor sucrose concentrations [8]. This novel experiment was not a bioanode for a biofuel cell, but it showed the possibility of utilizing sequential enzymes of natural pathways in a bioelectrochemical application. Over the last decade, there has been a wealth of evaluation of natural metabolic pathways for fuel oxidation. The most common natural metabolic pathway evaluated for bioelectrocatalysis is the methanol metabolic pathway utilizing alcohol dehydrogenase to oxidize methanol to formaldehyde, aldehyde dehydrogenase to oxidize formaldehyde to formate, and formate dehydrogenase to oxidize formate to carbon dioxide. This enzyme cascade has been used for oxidation of methanol in biofuel cells [7,9–11], as well as reduction of carbon dioxide to methanol in bioelectrosynthetic processes [12]. This pathway has been interesting for study, because it contains only three enzymes, all three enzymes are oxidoreductase enzymes capable of generating electrons, and the three enzymes have quite different specific activities. The results of this work have shown
http://dx.doi.org/10.1016/j.bbabio.2015.08.008 0005-2728/© 2015 Published by Elsevier B.V.
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
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that the enzyme cascade functions to completely oxidize methanol or to produce methanol from carbon dioxide and that degree of oxidation alters biofuel cell/biobattery performance (current density and power density). It also provided the system for the first work at modeling an enzyme cascade at an electrode for bioelectrocatalysis [9]. This model showed that there are many engineering parameters to consider when designing an enzyme cascade-based bioanode, including transport of fuel and cofactor, individual enzyme kinetics, and cofactor electrocatalysis. The Krebs cycle (commonly called the citric acid cycle) has been evaluated as a classic example of a natural metabolic pathway. Sokic-Lazic et al. evaluated the fabrication of NAD(P)-dependent dehydrogenasebased bioanodes containing the Krebs cycle enzymes in ethanol, pyruvate, and lactate biofuel cells [13–15]. Since oxidoreductase enzymes in the Krebs cycle are NAD(P)-dependent, then a mediated bioanode is necessary to transfer the electrons from the oxidoreductase enzyme to the electrode surface. The diffusional cofactor NAD(P) is used at the mediator, but it has poor electrochemistry on carbon current collectors (i.e. carbon cloth, carbon papers, and carbon felt). Therefore, an additional electrocatalyst is needed to decrease the overpotential of NAD(P)H oxidation. The electrocatalyst used in this study was methylene green, which was electropolymerized into a conducting polymer layer on the carbon current collector. As shown in Fig. 1, these studies show that the degree of oxidation improves the performance (current density and power density) of a biofuel cell. Theoretically, we would expect as we add additional oxidoreductase enzymes we would see an increase in performance that is a characteristic of the increase degree of oxidation (i.e. one oxidoreductase enzymes gives 2 electrons, but three oxidoreductase enzymes give 6 electrons, so we would expect a 3 fold increase in current and power density). However, due to the metabolic control within the Krebs cycle, performance is quite low until the entire cascade/cycle is immobilized on the electrode surface and then large enhancements (26 fold) in performance are observed due to venting of carbon dioxide and elimination of the build-up of individual byproducts of the cycle that can inhibit different enzymes of the cycle. Therefore, this study showed that natural enzyme cascades can improve efficiency of transformation or in the case of a biofuel cell efficiency of current generation, but due to the metabolic control present in many of these natural pathways, then a complete pathway is needed for optimal performance. Beilke et al. also evaluated the natural glycolytic process in-vitro [16, 17]. However, the glycolytic pathway is not particularly interesting from a bioelectrochemical perspective, because it only contains one oxidoreductase (electron producing) enzyme, whereas the Krebs cycle contains
5 oxidoreductase enzymes. Therefore, it showed the importance of high oxidoreductase enzyme to total enzyme ratios to ensure high performance (i.e. at least 50% of the enzyme on the electrode needs to be an oxidoreductase enzyme). Beilke et al. had an interest in combining the glycolytic pathway cascade and the Krebs cycle cascade together in a bioanode, but due to the incompatibility of two enzyme systems, no single immobilization material was discovered that could immobilize both sets of enzymes at a single bioanode. Beyond these three metabolic pathways, there are several other examples of subsets of metabolic pathways used to study enzyme cascades for bioelectrocatalysis. For instance, Moehlenbrock et al. evaluated the cascade of two enzymes of the pentose phosphate pathway to understand the effects of sequential enzyme proximity on bioelectrocatalytic performance [18], while Nguyen et al. studied the two enzyme sucrose metabolic system (invertase and glucose oxidase) while also evaluating the effect of sequential enzyme proximity [19]. Hickey et al. evaluated a slightly different sucrose sub-metabolic system utilizing invertase, glucose oxidase, and fructose dehydrogenase [20] to get 4 electrons per molecule of sucrose. All of these natural metabolic pathways (either as complete pathways or subsets of metabolic pathways) led researchers to realize the importance of enzyme cascades for deep or complete oxidation of biofuels. 3. Metabolons Paul Srere et al. postulated that most metabolic pathway enzymes exist in supercomplexes that provide proximity between sequential enzyme active sites and may even channel substrate between active sites [21–23]. He termed these supercomplexes as metabolons. After evaluating the performance of Krebs cycle enzyme cascades in biofuel cells and comparing them to mitochondrial bioanodes in biofuel cells, it was clear that for improved performance there was a need for structural proximity between neighboring enzymes. Fig. 2 shows a representative set of power curves for pyruvate/air biofuel cells where the bioanode is either an intact mitochondrial electrode or a lysed mitochondrial electrode, where the loading and therefore the volumetric catalytic activity are the same on both bioanodes [24]. This data makes it quite clear that the structure in the mitochondria is different than the structure in the lysed mitochondria and that the structure of the Krebs cycle enzymes in the mitochondria is critical to high metabolic flux. This work on mitochondrial bioelectrocatalysis led to an attempt to isolate and purify intact metabolons for bioelectrocatalysis. Moehlenbrock et al. developed a technique for crosslinking metabolons in-vivo in
Fig. 1. (Left) Schematic of the complete biofuel cell. Ethanol is oxidized serving as the fuel source at the anode (dark red lettering represents dehydrogenase enzymes, whereas the light red/pink lettering represents other non-energy producing enzymes). Oxygen is reduced to water at the 20% Pt on carbon GDE cathode. Potentiostat is used to measure open circuit potential and linear sweep polarization curves. (Right) Representative power curves of ethanol/air biofuel cells with different enzymatic cascades at the bioanode. All solutions are 100 mM ethanol and 1 mM NAD+ in pH 7.5 phosphate buffer and all measurements were made at room temperature. Anode electrode area is 1 cm2. Reproduced with permission from Elsevier [13].
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
S.D. Minteer / Biochimica et Biophysica Acta xxx (2015) xxx–xxx
Fig. 2. Representative power curves for unmediated whole mitochondria bioanodes and poly(methylene green) mediated mitochondrial lysate bioanodes in a 100 mM pyruvate/air biofuel cell at room temperature. Reproduced with permission from Elsevier [24].
isolated mitochondria with glutaraldehyde and dimethyl suberimidate, lysing the mitochondria, and isolating these metabolons. Those studies showed that the metabolon has considerably higher flux (current density) than the isolated enzyme without the metabolon structure [25], even though there is a decrease in enzyme activity for the individual enzymes of the Krebs cycle. The Krebs cycle metabolon had been proposed by Paul Srere, but never structurally evaluated [26]. Wu et al. utilized invivo crosslinking in isolated mitochondria, trypsin protein digestion, and mass spectrometry to evaluate the structure of the metabolon and verify structural evidence of a substrate channel between enzymes [27]. This evidence along with evidence of the need for proximity and substrate channeling in other metabolic systems led to the consideration of how to form metabolons ex-situ (hereafter referred to as synthetic metabolons or artificial metabolons). 4. Synthetic metabolons After showing that the metabolon structure is important for bioelectrocatalysis, efforts have focused on developing synthetic metabolon structures. Although synthetic metabolons have been popular in synthetic biology [28–30], there are not many examples in bioelectrocatalysis. Most of the work has focused on providing proximity to sequential enzymes of the cascade. Moehlenbrock et al. evaluated proximity utilizing heterobifunctional crosslinkers to crosslink two enzymes of the pentose phosphate cascade [31]. This evaluation showed that proximity is important in terms of current density performance, but does not engineer substrate channeling into the synthetic metabolon. Nguyen et al. used the sucrose oxidation enzymes discussed above to utilize DNA as a structural scaffold for altering the separation between sequential enzymes [19]. Although these studies have focused on proximity, non-bioelectrocatalytic studies have shown that substrate channeling is important [32]. Therefore, the future of enzyme cascade work will need to focus on developing synthetic metabolons that both provide proximity between active sites, and also provide for substrate channeling. 5. Minimal and artificial enzyme cascades Metabolic pathways are complex for biological reasons that are unnecessary for electrochemical applications (fuel cells and electrosynthesis). In an electrochemical application of an enzyme cascade, the only reactions that are important are oxidation reactions catalyzed by oxidoreductase enzymes and breaking of carbon–carbon bonds catalyzed by lyases. Therefore, a number of artificial enzyme cascades have been designed and engineered to work more efficiently for bioelectrocatalysis applications. For instance, the normal metabolic pathway for oxidizing
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glycerol requires N20 enzymes and utilizes the Krebs cycle and glycolytic pathway enzymes. Many of these enzymes catalyze electrochemically unnecessary reactions (phosphorylating a substrate and then dephosphylating it), whereas the electrochemical applications are only interested in breaking carbon–carbon bonds and generating electrons. All other enzymes take up unnecessary and needed space on the electrode without providing electrochemical benefit. Therefore, Arechederra et al. developed a minimal enzyme cascade for oxidizing glycerol to carbon dioxide utilizing three promiscuous enzymes (PQQ-dependent alcohol dehydrogenase, PQQ-dependent aldehyde dehydrogenase, and oxalate oxidase) [33]. These three enzymes do multi-step oxidization and carbon–carbon bond cleavage due to their inherent promiscuity. Xu et al. expanded on this original cascade to develop a 6 enzyme cascade for glucose oxidation to carbon dioxide [34,35]. This cascade utilized a PQQdependent glucose dehydrogenase and a PQQ-dependent gluconate dehydrogenase to oxidize glucose, then an aldolase to break the sugar ring into two glycerol derivatives that can be oxidized by the 3 enzyme glycerol cascade. This cascade was shown to have high coulombic efficiency (63%), faradaic efficiency (95%), and product efficiency (45%). This is considerably higher than cascades with higher numbers of enzymes in the pathway (i.e. Krebs cycle). Another example of cascade engineering is the 3-enzyme cascade for methanol. Banta et al. developed an artificial three enzyme cascade for methanol [11]. This cascade is considered an engineered cascade, because it is engineered to form a protein hydrogel for ensuring high concentration of protein on the electrode surface and short distances between enzyme active sites. This resulted in remarkably high current and power density methanol biobatteries. Chen and Banta also utilized this same cascade with a different immobilization strategy [36]. Their technology expressed the three enzyme cascade on the surface of yeast cells with specific sequential orientation via cohesion and dockerin pairs. This was not used in a biofuel cell, but did result in a 5 fold increase in NADH production with the three enzyme proximity, therefore showing its potential utility in a biofuel cell. These two examples of enzyme engineering were used to produce synthetic metabolons as discussed above. One of the most interesting works in artificial enzyme cascades is the recent work of the Zhang group. Zhang et al. have also utilized cell-free synthetic biology to design and engineer artificial enzyme cascades for sugar oxidation [37,38]. This particular artificial enzymatic cascade contains 13 enzymes and is responsible for complete oxidation of maltodextrin to carbon dioxide in a biobattery. This particular pathway is ATP and CoA-free compared to the natural pathway for sugar oxidation, therefore, eliminating many of the issues with previous work with the natural glycolytic pathway and Krebs cycle pathway. This resulted in greater than 90% faradaic efficiency for the system, which is extremely impressive for an enzyme cascade as large and complex as this cascade. Greater than 90% of biofuel cell anodes contain glucose oxidase to oxidize glucose to gluconolactone, but there are several inherent problems with this enzyme system, including the production of peroxide, difficulty in achieving direct electron transfer, and the production of a byproduct that is difficult to further oxidize. The Gorton research group has been pioneers in alternative glucose oxidizing enzymes [39–44] and has been evaluating and utilizing these alternative enzymes for sugar oxidation cascades. Gorton et al. developed an artificial 2 enzyme cascade for deep oxidation of sugars utilizing promiscuous enzymes [45]. The bi-enzyme cascade of cellobiose dehydrogenase and pyranose dehydrogenase can provide for multi-step oxidation of a variety of simple sugars therefore increasing electrochemical performance, efficiency, and energy density. 6. Conclusions Overall, enzyme cascades are critical for deep oxidation of complex biofuels and deep oxidation is critical for high energy density biofuel cells or biobatteries, where energy density is directly proportional to
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
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extended use time of the battery. Recent research has evaluated the engineering of those enzyme cascades utilizing structural scaffolds to ensure enzyme active site proximity. Recent research has also artificially engineered enzyme cascades for improved coulombic, faradaic, and product efficiencies. Overall, the last decade has seen considerable improvements in performance of enzyme cascade-based bioanodes for biofuel cells and biobatteries, but future research will need to focus on stability of artificial cascades as well as the design of substrate channeling and not just proximity into artificial metabolons. Transparency Document The Transparency document associated with this article can be found, in the version. Acknowledgements The author would like to acknowledge the funding of the Air Force Office of Scientific Research (FA950-12-10112). References [1] A.T. Yahiro, S.M. Lee, D.O. Kimble, Bioelectrochemistry. I. Enzyme utilizing biofuel cell studies, Biochim. Biophys. Acta 88 (1964) 375–383. [2] S.C. Barton, J. Gallaway, P. Atanassov, Enzymatic biofuel cells for implantable and microscale devices, Chem. Rev. 104 (2004) 4867–4886. [3] P. Cinquin, C. Gondran, F. Giroud, S. Mazabrard, A. Pellissier, F. Boucher, J.-P. Alcaraz, K. Gorgy, F. Lenouvel, S. Mathe, P. Porcu, S. Cosnier, A glucose biofuel cell implanted in rats, PLoS One 5 (2010) (No pp. given). [4] L. Halámková, J. Halámek, V. Bocharova, A. Szczupak, L. Alfonta, E. Katz, Implanted biofuel cell operating in a living snail, J. Am. Chem. Soc. 134 (2012) 5040–5043. [5] A. Szczupak, J. Halamek, L. Halamkova, V. Bocharova, L. Alfonta, E. Katz, Living battery — biofuel cells operating in vivo in clams, Energy Environ. Sci. 5 (2012) 8891–8895. [6] W. Gellett, M. Kesmez, J. Schumacher, N. Akers, S.D. Minteer, Biofuel cells for portable power, Electroanalysis 22 (2010) 727–731. [7] G. Palmore, H. Bertschy, S.H. Bergens, G.M. Whitesides, A methanol/dioxygen biofuel cell that uses NAD+-dependent dehydrogenases as catalysts: application of an electro-enzymatic method to regenerate nicotinamide adenine dinucleotide at low overpotentials, J. Electroanal. Chem. 443 (1998) 155–161. [8] I. Satoh, I. Karube, S. Suzuki, Enzyme electrode for sucrose, Biotechnol. Bioeng. 18 (1976) 269–272. [9] P. Kar, H. Wen, H. Li, S.D. Minteer, B.S. Calabrese, Simulation of multistep enzymecatalyzed methanol oxidation in biofuel cells, J. Electrochem. Soc. 158 (2011) B580–B586. [10] P.K. Addo, R.L. Arechederra, S.D. Minteer, Evaluating enzyme cascades for methanol/ air biofuel cells based on NAD+-dependent enzymes, Electroanalysis 22 (2010) 807–812. [11] Y.H. Kim, E. Campbell, J. Yu, S.D. Minteer, S. Banta, Complete oxidation of methanol in an enzymatic biofuel cell by a self-assembling hydrogel created from three modified dehydrogenase, Angew. Chem. 52 (2013) 1437–1440. [12] P.K. Addo, R. Arechederra, A. Waheed, J.D. Shoemaker, W.S. Sly, S.D. Minteer, Methanol production via bioelectrocatalytic reduction of carbon dioxide: role of carbonic anhydrase in improving electrode performance, Electrochem. Solid-State Lett. 14 (2011), E9. [13] D. Sokic-Lazic, S.D. Minteer, Citric acid cycle biomimic on a carbon electrode, Biosens. Bioelectron. 24 (2008) 945–950. [14] D. Sokic-Lazic, S.D. Minteer, Pyruvate/air enzymatic biofuel cell capable of complete oxidation, Electrochem. Solid-State Lett. 12 (2009) F26–F28. [15] D. Sokic-Lazic, A.A.R. de, S.D. Minteer, Utilization of enzyme cascades for complete oxidation of lactate in an enzymatic biofuel cell, Electrochim. Acta 56 (2011) 10772–10775. [16] M.C. Beilke, S.D. Minteer, Immobilization of the glycolysis enzymes in hydrophobically modified Nafion, Polym. Mater. Sci. Eng. 94 (2006) 556–557. [17] C.E. Menius, M.C. Beilke, D.S. Minteer, Immobilization of glycolysis enzymes in modified chitosan, abstracts of papers, 235th ACS National Meeting, New Orleans, LA, United States, April 6–10, 2008, 2008.
[18] M.J. Moehlenbrock, M. Meredith, S.D. Minteer, Bioelectrocatalytic oxidation of glucose in CNT impregnated hydrogels: advantages of synthetic enzymatic metabolon formation, ACS Catal. 2 (2012) 17–25. [19] K. Van Nguyen, F. Giroud, S.D. Minteer, Improved bioelectrocatalytic oxidation of sucrose in a biofuel cell with an enzyme cascade assembled on a DNA scaffold, J. Electrochem. Soc. 161 (2014) H930–H933. [20] D.P. Hickey, F. Giroud, D.W. Schmidtke, D.T. Glatzhofer, S.D. Minteer, Enzyme cascade for catalyzing sucrose oxidation in a biofuel cell, ACS Catal. 3 (2013) 2729–2737. [21] P.A. Srere, The metabolon, Trends Biochem. Sci. 10 (1985) 109–110. [22] P.A. Srere, Complexes of sequential metabolic enzymes, Annu. Rev. Biochem. 56 (1987) 89–124. [23] P.A. Srere, C.K. Mathews, Purification of multienzyme complexes, Methods Enzymol. 182 (1990). [24] R.L. Arechederra, K. Boehm, S.D. Minteer, Mitochondrial bioelectrocatalysis for biofuel cell applications, Electrochim. Acta 54 (2009) 7268–7273. [25] M.J. Moehlenbrock, T.K. Toby, A. Waheed, S.D. Minteer, Metabolon catalyzed pyruvate/air biofuel cell, J. Am. Chem. Soc. 132 (2010) 6288–6289. [26] C. Velot, M.B. Mixon, M. Teige, P.A. Srere, Model of quinary structure between Krebs TCA cycle enzymes: a model for the metabolon, Biochemistry 36 (1997) 14271–14276. [27] F. Wu, S. Minteer, Krebs cycle metabolon: structural evidence of substrate channeling revealed by cross-linking and mass spectrometry, Angew. Chem. Int. Ed. 54 (2015) 1851–1854. [28] C. You, Y.H.P. Zhang, Self-assembly of synthetic metabolons through synthetic protein scaffolds: one-step purification, co-immobilization, and substrate channeling, ACS Synth. Biol. 2 (2013) 102–110. [29] S. Schoffelen, J.C.M. van Hest, Multi-enzyme systems: bringing enzymes together in vitro, Soft Matter 8 (2012) 1736–1746. [30] C. Singleton, T.P. Howard, N. Smirnoff, Synthetic metabolons for metabolic engineering, J. Exp. Bot. 65 (2014) 1947–1954. [31] S. Schoffelen, J.C.M. van Hest, Chemical approaches for the construction of multi-enzyme reaction systems, Curr. Opin. Struct. Biol. 23 (2013) 613–621. [32] J.-L. Lin, L. Palomec, I. Wheeldon, Design and analysis of enhanced catalysis in scaffolded multienzyme cascade reactions, ACS Catal. 4 (2013) 505–511. [33] R.L. Arechederra, S.D. Minteer, Complete oxidation of glycerol in an enzymatic biofuel cell, Fuel Cells 9 (2009) 63–69. [34] S. Xu, S.D. Minteer, Characterizing efficiency of multi-enzyme cascade-based biofuel cells by product analysis, ECS Electrochem. Lett. 3 (2014) H24–H27. [35] S. Xu, S.D. Minteer, Enzymatic biofuel cell for oxidation of glucose to CO2, ACS Catal. 2012 (2012) 91. [36] F. Liu, S. Banta, W. Chen, Functional assembly of a multi-enzyme methanol oxidation cascade on a surface-displayed trifunctional scaffold for enhanced NADH production, Chem. Commun. 49 (2013) 3766–3768. [37] Z. Zhu, T.K. Tam, F. Sun, C. You, Y.-H.P. Zhang, A high-energy-density sugar biobattery via a synthetic enzymatic pathway, Nat. Commun. 5 (2014) 3026. [38] Z. Zhu, Y. Wang, S.D. Minteer, Y.H. Percival Zhang, Maltodextrin-powered enzymatic fuel cell through a non-natural enzymatic pathway, J. Power Sources 196 (2011) 7505–7509. [39] L. Stoica, R. Ludwig, D. Haltrich, L. Gorton, Third-generation biosensor for lactose based on newly discovered cellobiose dehydrogenase, Anal. Chem. 78 (2006) 393–398. [40] L. Stoica, T. Ruzgas, R. Ludwig, D. Haltrich, L. Gorton, Direct electron transfer—a favorite electron route for cellobiose dehydrogenase (CDH) from Trametes Villosa. Comparison with CDH from Phanerochaete Chrysosporium, Langmuir 22 (2006) 10801–10806. [41] F. Tasca, L. Gorton, W. Harreither, D. Haltrich, R. Ludwig, G. Noll, Direct electron transfer at cellobiose dehydrogenase modified anodes for biofuel cells, J. Phys. Chem. C 112 (2008) 9956–9961. [42] R. Ludwig, W. Harreither, F. Tasca, L. Gorton, Cellobiose dehydrogenase: a versatile catalyst for electrochemical applications, ChemPhysChem 11 (2010) 2674–2697. [43] M.N. Zafar, F. Tasca, S. Boland, M. Kujawa, I. Patel, C.K. Peterbauer, D. Leech, L. Gorton, Wiring of pyranose dehydrogenase with osmium polymers of different redox potentials, Bioelectrochemistry 80 (2010) 38–42. [44] O. Spadiut, D. Brugger, V. Coman, D. Haltrich, L. Gorton, Engineered pyranose 2oxidase: efficiently turning sugars into electrical energy, Electroanalysis 22 (2010) 813–820. [45] M. Shao, M. Nadeem Zafar, C. Sygmund, D.A. Guschin, R. Ludwig, C.K. Peterbauer, W. Schuhmann, L. Gorton, Mutual enhancement of the current density and the coulombic efficiency for a bioanode by entrapping bi-enzymes with Os-complex modified electrodeposition paints, Biosens. Bioelectron. 40 (2013) 308–314.
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
Contents lists available at ScienceDirect
Biochimica et Biophysica Acta journal homepage: www.elsevier.com/locate/bbabio
Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades Shelley D. Minteer Departments of Chemistry and Materials Science & Engineering, University of Utah, 315 S 1400 E Rm 2020, Salt Lake City, UT 84112, USA
a r t i c l e
i n f o
Article history: Received 2 May 2015 Accepted 20 August 2015 Available online xxxx Keywords: Metabolic pathway Bioelectrocatalysis Metabolon Enzyme cascade Biofuel cell
a b s t r a c t Anodic bioelectrodes for biofuel cells are more complex than cathodic bioelectrodes for biofuel cells, because laccase and bilirubin oxidase can individually catalyze four electron reduction of oxygen to water, whereas most anodic enzymes only do a single two electron oxidation of a complex fuel (i.e. glucose oxidase oxidizing glucose to gluconolactone while generating 2 electrons of the total 24 electrons), so enzyme cascades are typically needed for complete oxidation of the fuel. This review article will discuss the lessons learned from natural metabolic pathways about multi-step oxidation and how those lessons have been applied to minimal or artificial enzyme cascades. © 2015 Published by Elsevier B.V.
1. Introduction Enzymatic fuel cells are a type of bioelectronic device focused on bioenergetic conversion of chemical energy to electrical energy via the utilization of enzymes as electrocatalysts on the anode and/or the cathode. They were first discovered in the 1960s when a glucose oxidase anode was combined with a platinum cathode to produce a positive open circuit potential [1]. That early work invigorated a field that was primarily focused on harnessing electrical energy from biofuels present in living organisms (i.e. glucose in the blood stream) [2–5]. However, as the field of enzymatic fuel cells (often called biofuel cells) advanced, it became clear that there were alternative applications for enzymatic fuel cells. As stability and power density/current density performance increased, there became an interest in biofuel cells for portable power applications [6]. This interest spurred engineering of anodes from the original single oxidoreductase enzyme systems to cascades of enzymes responsible for catalyzing sequential reactions. This started with early work by Palmore and Whitesides, which incorporated the three enzyme cascade (NAD-dependent alcohol dehydrogenase, NAD-dependent aldehyde dehydrogenase, and NAD-dependent formate dehydrogenase) into the bioanode compartment of a methanol/oxygen biofuel cell [7]. This work showed the increase in performance observed with deeper degrees of oxidation of the fuel and postulated an increase in efficiency that would lead to increased energy density of biofuel cells/biobatteries. This early inspiration in enzyme cascades has led to a variety of research efforts in enzyme cascades that will be discussed below. They include
E-mail address: [email protected].
natural metabolic pathways, natural metabolons, synthetic metabolons, and minimal enzyme cascades. 2. Natural metabolic pathways Metabolic pathways are cascades of enzymes responsible for energy conversion in the living organism (oxidation of food/fuel or production of energetic molecules in the cell). These pathways include the glycolytic pathway, the Krebs cycle, the pentose phosphate pathways, fatty acid metabolism, amino sugar metabolism, purine biosynthesis, amino acid synthesis, and sucrose/starch metabolism. These pathways are responsible for catalyzing sequential reactions and therefore have been of interest to the field of bioelectrochemistry, since Suzuki and coworkers first used invertase and glucose oxidase to electrochemically monitor sucrose concentrations [8]. This novel experiment was not a bioanode for a biofuel cell, but it showed the possibility of utilizing sequential enzymes of natural pathways in a bioelectrochemical application. Over the last decade, there has been a wealth of evaluation of natural metabolic pathways for fuel oxidation. The most common natural metabolic pathway evaluated for bioelectrocatalysis is the methanol metabolic pathway utilizing alcohol dehydrogenase to oxidize methanol to formaldehyde, aldehyde dehydrogenase to oxidize formaldehyde to formate, and formate dehydrogenase to oxidize formate to carbon dioxide. This enzyme cascade has been used for oxidation of methanol in biofuel cells [7,9–11], as well as reduction of carbon dioxide to methanol in bioelectrosynthetic processes [12]. This pathway has been interesting for study, because it contains only three enzymes, all three enzymes are oxidoreductase enzymes capable of generating electrons, and the three enzymes have quite different specific activities. The results of this work have shown
http://dx.doi.org/10.1016/j.bbabio.2015.08.008 0005-2728/© 2015 Published by Elsevier B.V.
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
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that the enzyme cascade functions to completely oxidize methanol or to produce methanol from carbon dioxide and that degree of oxidation alters biofuel cell/biobattery performance (current density and power density). It also provided the system for the first work at modeling an enzyme cascade at an electrode for bioelectrocatalysis [9]. This model showed that there are many engineering parameters to consider when designing an enzyme cascade-based bioanode, including transport of fuel and cofactor, individual enzyme kinetics, and cofactor electrocatalysis. The Krebs cycle (commonly called the citric acid cycle) has been evaluated as a classic example of a natural metabolic pathway. Sokic-Lazic et al. evaluated the fabrication of NAD(P)-dependent dehydrogenasebased bioanodes containing the Krebs cycle enzymes in ethanol, pyruvate, and lactate biofuel cells [13–15]. Since oxidoreductase enzymes in the Krebs cycle are NAD(P)-dependent, then a mediated bioanode is necessary to transfer the electrons from the oxidoreductase enzyme to the electrode surface. The diffusional cofactor NAD(P) is used at the mediator, but it has poor electrochemistry on carbon current collectors (i.e. carbon cloth, carbon papers, and carbon felt). Therefore, an additional electrocatalyst is needed to decrease the overpotential of NAD(P)H oxidation. The electrocatalyst used in this study was methylene green, which was electropolymerized into a conducting polymer layer on the carbon current collector. As shown in Fig. 1, these studies show that the degree of oxidation improves the performance (current density and power density) of a biofuel cell. Theoretically, we would expect as we add additional oxidoreductase enzymes we would see an increase in performance that is a characteristic of the increase degree of oxidation (i.e. one oxidoreductase enzymes gives 2 electrons, but three oxidoreductase enzymes give 6 electrons, so we would expect a 3 fold increase in current and power density). However, due to the metabolic control within the Krebs cycle, performance is quite low until the entire cascade/cycle is immobilized on the electrode surface and then large enhancements (26 fold) in performance are observed due to venting of carbon dioxide and elimination of the build-up of individual byproducts of the cycle that can inhibit different enzymes of the cycle. Therefore, this study showed that natural enzyme cascades can improve efficiency of transformation or in the case of a biofuel cell efficiency of current generation, but due to the metabolic control present in many of these natural pathways, then a complete pathway is needed for optimal performance. Beilke et al. also evaluated the natural glycolytic process in-vitro [16, 17]. However, the glycolytic pathway is not particularly interesting from a bioelectrochemical perspective, because it only contains one oxidoreductase (electron producing) enzyme, whereas the Krebs cycle contains
5 oxidoreductase enzymes. Therefore, it showed the importance of high oxidoreductase enzyme to total enzyme ratios to ensure high performance (i.e. at least 50% of the enzyme on the electrode needs to be an oxidoreductase enzyme). Beilke et al. had an interest in combining the glycolytic pathway cascade and the Krebs cycle cascade together in a bioanode, but due to the incompatibility of two enzyme systems, no single immobilization material was discovered that could immobilize both sets of enzymes at a single bioanode. Beyond these three metabolic pathways, there are several other examples of subsets of metabolic pathways used to study enzyme cascades for bioelectrocatalysis. For instance, Moehlenbrock et al. evaluated the cascade of two enzymes of the pentose phosphate pathway to understand the effects of sequential enzyme proximity on bioelectrocatalytic performance [18], while Nguyen et al. studied the two enzyme sucrose metabolic system (invertase and glucose oxidase) while also evaluating the effect of sequential enzyme proximity [19]. Hickey et al. evaluated a slightly different sucrose sub-metabolic system utilizing invertase, glucose oxidase, and fructose dehydrogenase [20] to get 4 electrons per molecule of sucrose. All of these natural metabolic pathways (either as complete pathways or subsets of metabolic pathways) led researchers to realize the importance of enzyme cascades for deep or complete oxidation of biofuels. 3. Metabolons Paul Srere et al. postulated that most metabolic pathway enzymes exist in supercomplexes that provide proximity between sequential enzyme active sites and may even channel substrate between active sites [21–23]. He termed these supercomplexes as metabolons. After evaluating the performance of Krebs cycle enzyme cascades in biofuel cells and comparing them to mitochondrial bioanodes in biofuel cells, it was clear that for improved performance there was a need for structural proximity between neighboring enzymes. Fig. 2 shows a representative set of power curves for pyruvate/air biofuel cells where the bioanode is either an intact mitochondrial electrode or a lysed mitochondrial electrode, where the loading and therefore the volumetric catalytic activity are the same on both bioanodes [24]. This data makes it quite clear that the structure in the mitochondria is different than the structure in the lysed mitochondria and that the structure of the Krebs cycle enzymes in the mitochondria is critical to high metabolic flux. This work on mitochondrial bioelectrocatalysis led to an attempt to isolate and purify intact metabolons for bioelectrocatalysis. Moehlenbrock et al. developed a technique for crosslinking metabolons in-vivo in
Fig. 1. (Left) Schematic of the complete biofuel cell. Ethanol is oxidized serving as the fuel source at the anode (dark red lettering represents dehydrogenase enzymes, whereas the light red/pink lettering represents other non-energy producing enzymes). Oxygen is reduced to water at the 20% Pt on carbon GDE cathode. Potentiostat is used to measure open circuit potential and linear sweep polarization curves. (Right) Representative power curves of ethanol/air biofuel cells with different enzymatic cascades at the bioanode. All solutions are 100 mM ethanol and 1 mM NAD+ in pH 7.5 phosphate buffer and all measurements were made at room temperature. Anode electrode area is 1 cm2. Reproduced with permission from Elsevier [13].
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
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Fig. 2. Representative power curves for unmediated whole mitochondria bioanodes and poly(methylene green) mediated mitochondrial lysate bioanodes in a 100 mM pyruvate/air biofuel cell at room temperature. Reproduced with permission from Elsevier [24].
isolated mitochondria with glutaraldehyde and dimethyl suberimidate, lysing the mitochondria, and isolating these metabolons. Those studies showed that the metabolon has considerably higher flux (current density) than the isolated enzyme without the metabolon structure [25], even though there is a decrease in enzyme activity for the individual enzymes of the Krebs cycle. The Krebs cycle metabolon had been proposed by Paul Srere, but never structurally evaluated [26]. Wu et al. utilized invivo crosslinking in isolated mitochondria, trypsin protein digestion, and mass spectrometry to evaluate the structure of the metabolon and verify structural evidence of a substrate channel between enzymes [27]. This evidence along with evidence of the need for proximity and substrate channeling in other metabolic systems led to the consideration of how to form metabolons ex-situ (hereafter referred to as synthetic metabolons or artificial metabolons). 4. Synthetic metabolons After showing that the metabolon structure is important for bioelectrocatalysis, efforts have focused on developing synthetic metabolon structures. Although synthetic metabolons have been popular in synthetic biology [28–30], there are not many examples in bioelectrocatalysis. Most of the work has focused on providing proximity to sequential enzymes of the cascade. Moehlenbrock et al. evaluated proximity utilizing heterobifunctional crosslinkers to crosslink two enzymes of the pentose phosphate cascade [31]. This evaluation showed that proximity is important in terms of current density performance, but does not engineer substrate channeling into the synthetic metabolon. Nguyen et al. used the sucrose oxidation enzymes discussed above to utilize DNA as a structural scaffold for altering the separation between sequential enzymes [19]. Although these studies have focused on proximity, non-bioelectrocatalytic studies have shown that substrate channeling is important [32]. Therefore, the future of enzyme cascade work will need to focus on developing synthetic metabolons that both provide proximity between active sites, and also provide for substrate channeling. 5. Minimal and artificial enzyme cascades Metabolic pathways are complex for biological reasons that are unnecessary for electrochemical applications (fuel cells and electrosynthesis). In an electrochemical application of an enzyme cascade, the only reactions that are important are oxidation reactions catalyzed by oxidoreductase enzymes and breaking of carbon–carbon bonds catalyzed by lyases. Therefore, a number of artificial enzyme cascades have been designed and engineered to work more efficiently for bioelectrocatalysis applications. For instance, the normal metabolic pathway for oxidizing
3
glycerol requires N20 enzymes and utilizes the Krebs cycle and glycolytic pathway enzymes. Many of these enzymes catalyze electrochemically unnecessary reactions (phosphorylating a substrate and then dephosphylating it), whereas the electrochemical applications are only interested in breaking carbon–carbon bonds and generating electrons. All other enzymes take up unnecessary and needed space on the electrode without providing electrochemical benefit. Therefore, Arechederra et al. developed a minimal enzyme cascade for oxidizing glycerol to carbon dioxide utilizing three promiscuous enzymes (PQQ-dependent alcohol dehydrogenase, PQQ-dependent aldehyde dehydrogenase, and oxalate oxidase) [33]. These three enzymes do multi-step oxidization and carbon–carbon bond cleavage due to their inherent promiscuity. Xu et al. expanded on this original cascade to develop a 6 enzyme cascade for glucose oxidation to carbon dioxide [34,35]. This cascade utilized a PQQdependent glucose dehydrogenase and a PQQ-dependent gluconate dehydrogenase to oxidize glucose, then an aldolase to break the sugar ring into two glycerol derivatives that can be oxidized by the 3 enzyme glycerol cascade. This cascade was shown to have high coulombic efficiency (63%), faradaic efficiency (95%), and product efficiency (45%). This is considerably higher than cascades with higher numbers of enzymes in the pathway (i.e. Krebs cycle). Another example of cascade engineering is the 3-enzyme cascade for methanol. Banta et al. developed an artificial three enzyme cascade for methanol [11]. This cascade is considered an engineered cascade, because it is engineered to form a protein hydrogel for ensuring high concentration of protein on the electrode surface and short distances between enzyme active sites. This resulted in remarkably high current and power density methanol biobatteries. Chen and Banta also utilized this same cascade with a different immobilization strategy [36]. Their technology expressed the three enzyme cascade on the surface of yeast cells with specific sequential orientation via cohesion and dockerin pairs. This was not used in a biofuel cell, but did result in a 5 fold increase in NADH production with the three enzyme proximity, therefore showing its potential utility in a biofuel cell. These two examples of enzyme engineering were used to produce synthetic metabolons as discussed above. One of the most interesting works in artificial enzyme cascades is the recent work of the Zhang group. Zhang et al. have also utilized cell-free synthetic biology to design and engineer artificial enzyme cascades for sugar oxidation [37,38]. This particular artificial enzymatic cascade contains 13 enzymes and is responsible for complete oxidation of maltodextrin to carbon dioxide in a biobattery. This particular pathway is ATP and CoA-free compared to the natural pathway for sugar oxidation, therefore, eliminating many of the issues with previous work with the natural glycolytic pathway and Krebs cycle pathway. This resulted in greater than 90% faradaic efficiency for the system, which is extremely impressive for an enzyme cascade as large and complex as this cascade. Greater than 90% of biofuel cell anodes contain glucose oxidase to oxidize glucose to gluconolactone, but there are several inherent problems with this enzyme system, including the production of peroxide, difficulty in achieving direct electron transfer, and the production of a byproduct that is difficult to further oxidize. The Gorton research group has been pioneers in alternative glucose oxidizing enzymes [39–44] and has been evaluating and utilizing these alternative enzymes for sugar oxidation cascades. Gorton et al. developed an artificial 2 enzyme cascade for deep oxidation of sugars utilizing promiscuous enzymes [45]. The bi-enzyme cascade of cellobiose dehydrogenase and pyranose dehydrogenase can provide for multi-step oxidation of a variety of simple sugars therefore increasing electrochemical performance, efficiency, and energy density. 6. Conclusions Overall, enzyme cascades are critical for deep oxidation of complex biofuels and deep oxidation is critical for high energy density biofuel cells or biobatteries, where energy density is directly proportional to
Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008
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extended use time of the battery. Recent research has evaluated the engineering of those enzyme cascades utilizing structural scaffolds to ensure enzyme active site proximity. Recent research has also artificially engineered enzyme cascades for improved coulombic, faradaic, and product efficiencies. Overall, the last decade has seen considerable improvements in performance of enzyme cascade-based bioanodes for biofuel cells and biobatteries, but future research will need to focus on stability of artificial cascades as well as the design of substrate channeling and not just proximity into artificial metabolons. Transparency Document The Transparency document associated with this article can be found, in the version. Acknowledgements The author would like to acknowledge the funding of the Air Force Office of Scientific Research (FA950-12-10112). References [1] A.T. Yahiro, S.M. Lee, D.O. Kimble, Bioelectrochemistry. I. Enzyme utilizing biofuel cell studies, Biochim. Biophys. Acta 88 (1964) 375–383. [2] S.C. Barton, J. Gallaway, P. Atanassov, Enzymatic biofuel cells for implantable and microscale devices, Chem. Rev. 104 (2004) 4867–4886. [3] P. Cinquin, C. Gondran, F. Giroud, S. Mazabrard, A. Pellissier, F. Boucher, J.-P. Alcaraz, K. Gorgy, F. Lenouvel, S. Mathe, P. Porcu, S. Cosnier, A glucose biofuel cell implanted in rats, PLoS One 5 (2010) (No pp. given). [4] L. Halámková, J. Halámek, V. Bocharova, A. Szczupak, L. Alfonta, E. Katz, Implanted biofuel cell operating in a living snail, J. Am. Chem. Soc. 134 (2012) 5040–5043. [5] A. Szczupak, J. Halamek, L. Halamkova, V. Bocharova, L. Alfonta, E. Katz, Living battery — biofuel cells operating in vivo in clams, Energy Environ. Sci. 5 (2012) 8891–8895. [6] W. Gellett, M. Kesmez, J. Schumacher, N. Akers, S.D. Minteer, Biofuel cells for portable power, Electroanalysis 22 (2010) 727–731. [7] G. Palmore, H. Bertschy, S.H. Bergens, G.M. Whitesides, A methanol/dioxygen biofuel cell that uses NAD+-dependent dehydrogenases as catalysts: application of an electro-enzymatic method to regenerate nicotinamide adenine dinucleotide at low overpotentials, J. Electroanal. Chem. 443 (1998) 155–161. [8] I. Satoh, I. Karube, S. Suzuki, Enzyme electrode for sucrose, Biotechnol. Bioeng. 18 (1976) 269–272. [9] P. Kar, H. Wen, H. Li, S.D. Minteer, B.S. Calabrese, Simulation of multistep enzymecatalyzed methanol oxidation in biofuel cells, J. Electrochem. Soc. 158 (2011) B580–B586. [10] P.K. Addo, R.L. Arechederra, S.D. Minteer, Evaluating enzyme cascades for methanol/ air biofuel cells based on NAD+-dependent enzymes, Electroanalysis 22 (2010) 807–812. [11] Y.H. Kim, E. Campbell, J. Yu, S.D. Minteer, S. Banta, Complete oxidation of methanol in an enzymatic biofuel cell by a self-assembling hydrogel created from three modified dehydrogenase, Angew. Chem. 52 (2013) 1437–1440. [12] P.K. Addo, R. Arechederra, A. Waheed, J.D. Shoemaker, W.S. Sly, S.D. Minteer, Methanol production via bioelectrocatalytic reduction of carbon dioxide: role of carbonic anhydrase in improving electrode performance, Electrochem. Solid-State Lett. 14 (2011), E9. [13] D. Sokic-Lazic, S.D. Minteer, Citric acid cycle biomimic on a carbon electrode, Biosens. Bioelectron. 24 (2008) 945–950. [14] D. Sokic-Lazic, S.D. Minteer, Pyruvate/air enzymatic biofuel cell capable of complete oxidation, Electrochem. Solid-State Lett. 12 (2009) F26–F28. [15] D. Sokic-Lazic, A.A.R. de, S.D. Minteer, Utilization of enzyme cascades for complete oxidation of lactate in an enzymatic biofuel cell, Electrochim. Acta 56 (2011) 10772–10775. [16] M.C. Beilke, S.D. Minteer, Immobilization of the glycolysis enzymes in hydrophobically modified Nafion, Polym. Mater. Sci. Eng. 94 (2006) 556–557. [17] C.E. Menius, M.C. Beilke, D.S. Minteer, Immobilization of glycolysis enzymes in modified chitosan, abstracts of papers, 235th ACS National Meeting, New Orleans, LA, United States, April 6–10, 2008, 2008.
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Please cite this article as: S.D. Minteer, Oxidative bioelectrocatalysis: From natural metabolic pathways to synthetic metabolons and minimal enzyme cascades, Biochim. Biophys. Acta (2015), http://dx.doi.org/10.1016/j.bbabio.2015.08.008