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Letter
Improving the Performance of Methanol Biofuel Cells Utilizing an Enzyme Cascade Bioanode with DNA Bridged Substrate Channeling Lin Xia, Khiem Van Nguyen, Yaovi Holade, Han Han, Kevin Dooley, Plamen Atanassov, Scott Banta, and Shelley D. Minteer ACS Energy Lett., Just Accepted Manuscript • Publication Date (Web): 17 May 2017 Downloaded from http://pubs.acs.org on May 18, 2017
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ACS Energy Letters
Improving the Performance of Methanol Biofuel Cells Utilizing an Enzyme Cascade Bioanode with DNA Bridged Substrate Channeling Lin Xia1, Khiem Van Nguyen,1,2 Yaovi Holade,1,3 Han Han4, Kevin Dooley5, Plamen Atanassov6, Scott Banta5, and Shelley D. Minteer*,1 1 Departments of Chemistry and Materials Science and Engineering, University of Utah, 315 S 1400 E Room 2020, Salt Lake City, Utah 84112, United States 2 VietnamNTT Hi‐Tech Institute, Nguyen Tat Thanh University, Ho Chi Minh City, Vietnam 3 Institut Européen des Membranes (IEM) UMR 5635, ENSCM, CNRS, UM, France 4Department of Biochemistry, University of Utah, Salt Lake City, Utah, United States 5Department of Chemical Engineering, Columbia University, New York City, United States
6Department of Chemical and Nuclear Engineering, University of New Mexico, Albuquerque, NM, United States
ABSTRACT: The development of enzymatic biofuel cells has been plagued by the high cost of enzyme purification and low efficiency of fuel oxidation. Here, we demonstrate a protein purification‐free approach to assemble an alcohol dehydrogenase and aldehyde dehydrogenase enzyme cascade‐based bioanode for use in a methanol biofuel cell. Each enzyme was fused to a different sequence‐specific, zinc finger DNA‐binding protein. The zinc finger domains both serve as tags to isolate the en‐ zymes from crude cell lysates as well as anchors to immobilize the enzymes on DNA modified multiwalled carbon nanotubes. The biofuel cells based on the enzyme cascade bioanodes show a maximum power output of 24.5 ± 3.2 μW cm−2, which is comparable to fuel cells utilizing purified enzymes. Further analysis of kinetic behavior revealed a significant increase in the reactivity of the complexes due to substrate channeling of the aldehyde intermediate.
Recent advancements in genetic engineering and nucleic acid nanotechnology have brought artificial multi‐enzyme complex assembly into the forefront of biocatalysis research1‐3. In na‐ ture, multi‐enzyme complexes offer significant advantages in catalytic efficiency over isolated enzymes during sequential multistep catalysis. Many of these enzyme systems are spatially organized to control the mass transport of reactants and inter‐ mediates between enzymes from one active site to another which decreases diffusion to the bulk environment. This phe‐ nomenon has been termed substrate channeling4‐6.
Promoting proximity of active sites that catalyze multi steps of sequential reactions is one of the simplest strategies that cells employ to achieve substrate channeling between multi‐enzyme complexes. Inspired by nature, assembling artificial cascade enzyme complexes has attracted great research interest. Vari‐ ous scaffolds including peptides7, nucleic acids8‐9, and poly‐ mers10‐11 have been explored for the spatial organization of en‐ zymes of a cascade. The programmability of DNA nanostruc‐ tures and the versatility of DNA‐protein conjugation strategies makes DNA an ideal scaffold for this purpose2, 12. Moreover, in‐ troducing sequence‐specific DNA‐binding proteins, specifically
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zinc finger proteins, into the target enzyme as an anchor offers a controllable way to attach enzyme onto DNA scaffolds while retaining the enzymes activity9, 13. By using this DNA based multi‐enzyme complex assembly tech‐ nique to advance enzymatic biofuel cell (EBFC) applications, we determined that the system addresses two of the major is‐ sues existing in EBFCs. First, by fusing sequence‐specific DNA‐ binding proteins onto alcohol dehydrogenase (ADH) and alde‐ hyde dehydrogenase (ALDH), we demonstrate the ability to capture recombinant enzymes directly from cell lysates, which can potentially reduce the high costs of enzyme purification necessary for EBFC fabrication. Second, the site‐specific assem‐ bly of cascade enzymes on the anode not only decreased the time for the intermediate to move between enzymes, but also the extent of methanol oxidation to formate.
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Figure 1. A) Enzyme activity assay of DNA bridged ADH‐DNA‐ ALDH complex (black curve) and isolated ADH/ALDH mixture (blue curve); B) plot of lag time versus NADH yield at 1 min, blue square: isolated ADH/ALDH mixture as a control; black triangle: DNA bridged complex.
To achieve our goal of using zinc finger domains to organize en‐ zymes on DNA decorated carbon nanotubes, the zinc finger protein genes Zif268 and PBSII were, respectively, fused to ADH and ALDH gene by using the Gibson assembly method. Then, the constructed plasmids were expressed in E.coli BL21 strain to produce Zif 268 fused ADH enzyme (Zif268‐ADH) and PBSII fused ALDH enzyme (PBSII‐ALDH) (detailed enzyme con‐ struction, expression, and purification procedures can be found in the Supporting Information). In natural metabolic pathways, sequential enzymes are ori‐ ented in close proximity with each other to promote substrate channeling of intermediates from one enzyme active site to the next. This substrate channeling improves flux through the met‐ abolic pathways. To investigate if there is a substrate channel‐ ing effect in the as‐assembled ADH‐DNA‐ALDH complexes, ki‐ netic analyses of supramolecular DNA–enzyme complexes were performed in comparison with ADH and ALDH mixture without the DNA bridge. Prior to the enzyme activity assay, the concentration of the purified enzyme complex and enzyme mixture was normalized by the BCA assay. Details regarding to the preparation of purified enzyme complex can be found in ESI. For the protein‐DNA complex binding study, the bienzyme complex is too large for native gel analysis, so the bienzyme complex was studied by gel‐filtration chromatography. Through the analysis of the size‐exclusion chromatography re‐ sults (Figure S3), it was determined that the dual binding yield of protein‐DNA complex is >60% when assembled with an op‐ timized DNA binding site length. Figure 1A shows the kinetic difference between the DNA–enzyme complexes and isolated enzyme mixture. Clearly the DNA bridged ADH and ALDH com‐ plex shows higher activity towards methanol, indicated by a re‐ duction in the lag time required to reach steady state, where lag time is defined as the time required for intermediate to transfer between sequential active sites. By calculating the lag time of this two cascade catalytic reaction system, a plot of lag time versus NADH yield was given to further demonstrate the DNA bridged complex offers distinct improvement in the time for the intermediate to transfer between active sites over isolated enzymes during the sequential catalysis of methanol to formal‐ dehyde, and then to formic acid (Figure 1B).
and a single stranded tail was attached on multiwalled carbon nanotubes (MWCNTs) via sonication. The DNA scaffolds bind to MWCNTs through the single‐stranded toehold domain and the double stranded domain after hybridization is exposed to enable binding of the zinc finger proteins. Although DNA scaf‐ folds have been widely used to assemble enzyme cascades, the non‐conductivity of this biological scaffold is a huge drawback for application of the system in bioelectrocatalysis. With the ex‐ cellent conductivity of the MWCNTs, our design also intro‐ duced a new method for site‐specific assembly of an enzyme cascade on a DNA scaffold that is particularly suitable for bioe‐ lectrocatalytic applications. The modified MWCNTs were cast on Toray carbon paper electrodes and dried. Finally, the modi‐ fied electrode was incubated in cell lysate solution to capture target Zif268‐ADH and PBSII‐ALDH enzyme. The specific‐bind‐ ing between ADH/ALDH and its corresponding DNA sequence that modified on electrode was studied by capture efficiency studies. After optimization, the capture efficiency of the specific DNA‐MWCNTs modified electrode was about 6 times higher than that of non‐specific DNA‐MWCNTs modified electrode. The fabrication detail of the different assembled anodes and the capture efficiency of the DNA/MWCNTs modified Toray carbon paper electrode was calculated and optimized, as shown in Supporting Information.
Figure 2. A cartoon illustration of the enzyme cascade anode fabrication process
The fabrication of the ADH/DNA/ALDH complex‐based cas‐ cade bioanode is illustrated in Figure 2. Detailed electrode fab‐ rication procedures can be found in the Supporting Infor‐ mation. Briefly, the DNA scaffold containing Zif268 and PBSII binding domains
After assembling the enzyme cascade anode, we then investi‐ gated the possibility of using this method for assembling bio‐ anodes directly from unpurified proteins in cell lysates for
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ACS Energy Letters purified enzymes for assembling the anode. From the Lin‐ eweaver‐Burke plot, an apparent Km value of 20 ± 0.8 mM for methanol for the zinc‐finger capture method assembled bi‐en‐ zyme anode and 38 ± 1.2 mM for the purified complex modified bi‐enzyme anode were determined (Figure 3D). The lower ap‐ parent Km observed from capture method assembled anode may be attributed to the specific binding of enzyme onto the electrode which provides an improved electrode interface and so better enzyme affinity.
methanol oxidation via bioelectrocatalysis. As both ADHII and ALDH are NAD‐dependent enzymes, anodic bioelectrocatalysis resulting from the enzymatic oxidation of methanol is electro‐ chemically related to the oxidation of NADH at the carbon elec‐ trode. To lower the overpotential of NADH oxidation, 1,2‐naph‐ thoquinone‐4‐sulfonic acid was employed to mediate NADH oxidation. Cyclic voltammetric (CV) analysis was performed to evaluate the catalytic performance of different electrodes to‐ wards methanol oxidation. Figure 3A shows the bioelectrocat‐ alytic current of the bioanode formed via the zinc‐finger cap‐ ture method of assembly resulting from the enzymatic oxida‐ tion of methanol. The as‐prepared anode were capable of gen‐ erating 0.052 ± 0.007 mA cm−2 in the presence of 60 mM meth‐ anol at pH 7.5 and room temperature. As a comparison, when using non‐specific DNA modified electrodes for enzyme cap‐ ture, the bioelectrocatalytic current of the resulting bioanode was barely observed (Figure S4A), because of the low capture efficiency without specific DNA‐zinc finger protein binding. In addition, the CVs of the DNA/MWCNTs modified Toray carbon paper incubated with purified enzymes was shown in Figure 3B. The catalytic current density generated was 0.044 ± 0.005 mA cm−2 higher than that of the purification‐ free method as‐ sembled cascade anode, presumably because of a much higher enzyme loading on the anode. However, the isolated purified enzyme (no DNA bridged enzyme complex formed) modified electrode (Figure S4B) yield 0.021 ± 0.003 mA cm−2 less than the zinc‐finger capture method of assembly from purified en‐ zyme.
Figure 4. Representative power curves (to the right Y axis) and polarization curves (to the left Y axis) obtained from the BFCs constructed with enzyme captured by MWNTs/TP anode (without DNA) ,black curve : from cell lysate; blue curve: from purified enzyme mixture; and enzyme captured by DNA/MWNTs/TP anode, red curve : from cell lysate and dark golden curve: from purified enzyme mixture. All tests were per‐ formed with BOD cathode in one compartment cell, 0.1 M Po‐ tassium phosphate buffer, pH 7.5, 2 mM mediator, 5 mM NAD+ and 120 mM methanol. Subsequently, EBFCs were assembled based on differently as‐ sembled cascade anodes. The cathode was assembled from bil‐ irubin oxidase (EC 1.3.3.5 BOD), Nafion and MWCNTs mixture casted on Toray carbon paper electrode, as described in the Supporting Information. The fuel was methanol at the anode and oxygen from the air was reduced at the cathode. The fuel cell was a single compartment configuration and the PBS elec‐ trolyte solution was bubbled with air. The performance of the biofuel cell with different anodes is shown in Figure 4. As ex‐ pected, the maximum power output from the zinc‐finger cap‐ ture method assembled bioanodes based BFCs is 24.5±3.2 μW cm−2 (red curve), while the purified enzyme complex modified anode based BFC shows a maximum power output of 33.4 ± 4.5 μW cm−2 (dark golden curve ). However, the enzyme captured by MWCNTs without DNA modified anode based BFCs has shown a lower power output of 17.0 ± 2.3 μW cm−2 from cell lysate( black curve), and 19.2 ± 3.3 μW cm−2 from purified en‐ zyme mixture solution. This clearly shows that the DNA bridge orientation of the enzymes on the carbon nanotubes improved performance by 73%and that high performance bioanodes can be formed without the need to purify enzyme in advance of cre‐ ating the BFCs.
Figure 3. Cyclic voltammograms of (A) the enzyme zinc finger capture method of anode assembly and (B) purified enzyme complex modified anode; black curve: before adding methanol, red curve: upon the addition of 60 mM methanol, scan rate 5 mV/s, , 0.1 M Potassium phosphate buffer, pH 7.5, 2 mM medi‐ ator, 5 mM NAD+ C) The amperometric i‐t curve of the zin finger capture method assembled anode (black) and purified enzyme complex modified anode (red), applied potential 0.05V; D) the methanol concentration versus responsive current density plot. The amperometric response of the modified electrodes upon the addition of methanol was investigated at an applied poten‐ tial of 0.05 V vs SCE (Figure 3C). The zinc finger capture method assembled an anode from cell lysate that demonstrated compa‐ rable amperometric performance to an anode fabricated using
In conclusion, we have successfully demonstrated a zinc finger mediated approach to capture enzyme directly from cell lysate for fabricating enzyme cascade‐based bioanodes. The as‐pre‐ pared anode displayed comparable bioelectrocatalytic perfor‐
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mance to purified enzyme complex modified electrode. Moreo‐ ver, by using a designed DNA duplex sequence with a predicted 4 nm between enzyme binding site, evidence of apparent sub‐ strate channeling between active sites of cascade enzymes was observed and a 73% increase in the power density of the re‐ sulting biofuel cell.
4. Huang, X.; Holden, H. M.; Raushel, F. M. Channeling of Substrates and Intermediates in Enzyme‐Catalyzed Reactions. Annu. Rev. Biochem 2001, 70, 149‐180. 5. Milani, M.; Pesce, A.; Bolognesi, M.; Bocedi, A.; Ascenzi, P. Substrate channeling: Molecular bases. Biochemistry and Molecular Biology Education 2003, 31, 228‐233.
ASSOCIATED CONTENT
6. Spivey, H. O.; Ovádi, J. Substrate Channeling. Methods 1999, 19, 306‐321.
Supporting Information. Experimental procedures, chroma‐ tographic analysis, and spectroscopic analysis.
7. Dueber, J. E.; Wu, G. C.; Malmirchegini, G. R.; Moon, T. S.; Petzold, C. J.; Ullal, A. V.; Prather, K. L. J.; Keasling, J. D. Syn‐ thetic protein scaffolds provide modular control over meta‐ bolic flux. Nat Biotech 2009, 27, 753‐759.
AUTHOR INFORMATION
8. Erkelenz, M.; Kuo, C.‐H.; Niemeyer, C. M. DNA‐ Mediated Assembly of Cytochrome P450 BM3 Subdomains. J. Am. Chem. Soc. 2011, 133, 16111‐16118.
Corresponding Author *
[email protected] Funding Sources
9. Ngo, T. A.; Nakata, E.; Saimura, M.; Morii, T. Spatially Organized Enzymes Drive Cofactor‐Coupled Cascade Reac‐ tions. J. Am. Chem. Soc. 2016, 138, 3012‐3021. 10. Vriezema, D. M.; Garcia, P. M. L.; Sancho Oltra, N.; Hat‐ zakis, N. S.; Kuiper, S. M.; Nolte, R. J. M.; Rowan, A. E.; van Hest, J. C. M. Positional Assembly of Enzymes in Polymersome Nano‐ reactors for Cascade Reactions. Angew. Chem. Int. Ed. 2007, 46, 7378‐7382.
Army Research Office and Air Force Office of Scientific Re‐ search
ACKNOWLEDGMENT The authors would like to thank the Army Research Office MURI and Air Force Office of Scientific Research for funding this project.
11. Wang, X.; Li, Z.; Shi, J.; Wu, H.; Jiang, Z.; Zhang, W.; Song, X.; Ai, Q. Bioinspired Approach to Multienzyme Cascade System Construction for Efficient Carbon Dioxide Reduction. ACS Catal. 2014, 4, 962‐972.
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