Anal. Chem. 1998, 70, 5062-5071
Fast-Scan Cyclic Voltammetry of Protein Films on Pyrolytic Graphite Edge Electrodes: Characteristics of Electron Exchange Judy Hirst and Fraser A. Armstrong*
Inorganic Chemistry Laboratory, South Parks Road, Oxford, OX1 3QR, England
The rapid electron-exchange characteristics of metalloproteins adsorbed at a pyrolytic graphite “edge” electrode have been studied by analog dc cyclic voltammetry at scan rates up to 3000 V s-1. The voltammetry of four proteins, azurin (a “blue” copper protein) and three 7Fe ferredoxins, reveals oxidation and reduction peaks that display only modest increases in width and peak separation as the scan rate is raised. This is indicative of a substantially homogeneous population of noninteracting centers which undergo rapid electron exchange with the electrode. Both the Butler-Volmer and Marcus models have been tested. The electrochemical kinetics, as reflected by k0 (the rate at zero overpotential), are too fast to allow the determination of reorganization energies by this method. Nonetheless, the rapid and energetically coherent nature of the electron transfer enables the cyclic oxidation and reduction of protein redox centers to be examined on a time scale sufficiently short to recognize coupled processes occurring in the millisecond time domain, which are characteristic of the protein under investigation. Two of the ferredoxins display increasingly asymmetric voltammetry as the scan rate is increased, which is attributed to the coupling of electron transfer to conformational (or orientational) changes. For azurin, the use of higher electrolyte concentrations enables studies to be made at scan rates up to 3000 V s-1, from which a standard electron-transfer rate constant in the region of 5000 s-1 is obtained. At these high scan rates, azurin still shows very symmetrical voltammograms but with peak shapes displaying a more gradual decrease in current, at increasing overpotential, than is predicted using realistic values of the reorganization energy. The ability to measure even faster rate constants and access coupled reactions occurring in shorter time domains is likely to be limited by complex processes occurring on the graphite surface. Protein film voltammetry1 affords new, and as-yet largely unexploited, opportunities for the detailed study of electron transfer and coupled reactions in proteins. The principle is that the protein sample under investigation is confined to the electrode surface, ideally as a homogeneous population of noninteracting molecules at monolayer coverage or less, retaining full biological (1) Armstrong, F. A.; Heering, H. A.; Hirst, J. Chem. Soc. Rev. 1997, 26, 169179.
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activity and exhibiting fast interfacial electron transfer. The inherent usefulness of the technique stems from several features. First, immobilization minimizes complications arising from sluggish and irreproducible protein diffusion,2 and in the simplest case of reversible electron transfer, sharp voltammetric signals at characteristic potentials are produced which are akin to spectral signals and which thus serve to identify particular redox centers.1 Second, the experiment addresses just a minuscule quantity of sample, creating sensitivity to extremely low levels of reacting species in the contacting electrolytesthe supply and removal of which can be controlled by induced hydrodynamics. Third is the ability to induce and monitor even the most complex redox reactions under the strictest conditions of controlled potential. We are particularly interested in elucidating the mechanisms by which electron transfer is coupled, a topic of major importance in biology.3 By coupling, we allude to how electron-transfer rates may actually be controlled (gated) by chemical events4 or how electron transfer drives other reactions such as conformational changes or ion (e.g., proton) transfer.3 In the past, efforts to study the kinetics of these complex coupling processes in proteins have been restricted to bringing together various pieces of data from separate kinetic and thermodynamic experiments conducted in solution. However, exciting opportunities are afforded by electrochemical methods. Cyclic voltammetry, although regarded generally as a qualitative technique with regard to electron-transfer kinetics, is very well suited to visualizing and analyzing coupling processes. In effect, the technique (as applied to a protein film) drives and monitors the rapid injection of an electron (or hole) into a protein redox center and its retrieval within a short, defined time scale. Rate-determining coupled reactions are revealed because they distort the symmetry of the exchange, and quantitative information can be extracted by simulation. So far, we have used protein film voltammetry to probe the complex redox chemistry of a variety of systems ranging from labile Fe-S clustersswhere the voltammetric signal has served to show the presence of a particular cluster and reveal its redox (2) Armstrong, F. A.; Bond, A. M.; Hill, H. A. O.; Psalti, I. S. M.; Zoski, C. G. J. Phys. Chem. 1989, 93, 6485-6493. (3) See, for example: Rich, P. R. In Protein Electron Transfer; Bendall, D. S., Ed.; Bios Scientific Publishers: Oxford, 1996; pp 217-248. Brzezinski, P. Biochemistry 1996, 35, 5611-5615. Malmstro¨m, B. G. Acc. Chem. Res. 1993, 26, 332-338. (4) Bond, A. M.; Oldham, K. B. J. Phys. Chem. 1993, 87, 2492-2502. O’Connell, K. M.; Evans, K. B. J. Am. Chem. Soc. 1983, 105, 1473-1481. Brunschwig, B. S.; Sutin, N. J. Am. Chem. Soc. 1989, 111, 7455-7465. Hoffman, B. M.; Ratner, M. A. J. Am. Chem. Soc. 1987, 109, 6237-6243. 10.1021/ac980557l CCC: $15.00
© 1998 American Chemical Society Published on Web 10/28/1998
and chemical properties5-7sto enzymes where both nonturnover (no-substrate) electron exchange and catalytic electron transport have been examined in detail.8 In these studies, we have made extensive use of the pyrolytic graphite “edge” (PGE) electrode, the surface of which is hydrophilic due to the presence of oxide functionalities generated during the polishing procedure. Electroactive protein films of up to monolayer coverage are obtainable by optimizing conditions of temperature and electrolyte and (where necessary) by the inclusion of cationic coadsorbates such as polymyxin or an aminocyclitol.1,9 As judged from their retention of native catalytic activity and the high rates of many independently defined coupled processes, the adsorbed proteins are usually able to react freely with small-molecule reagents in solution. In many cases it has been possible to achieve well-defined oxidation and reduction peaks which have wave shapes close to those predicted for an ideal (nondispersed, noninteracting) ensemble of adsorbed molecules.10 Indeed, throughout our studies we have been intrigued by the quality and reproducibility of results obtainable without any sophisticated pretreatment or modification of the electrode surface, particularly since graphite is acknowledged to be a far from ideal electrode material.11-13 Simplicity and efficiency have been important since our focus throughout has been on the potential applications to mechanistic bioinorganic chemistry and enzymology rather than interfacial electrochemistry.1 For comparison, there have been several accounts of the electrochemistry of cytochrome c adsorbed on self-assembled headgroup (X)-functionalized alkanethiol (X(CH2)nS) monolayers on gold electrodes.14,15 This more sophisticated approach to (5) (a) Butt, J. N.; Armstrong, F. A.; Breton, J.; George, S. J.; Thomson, A. J.; Hatchikian, E. C. J. Am. Chem. Soc. 1991, 113, 6663-6670. (b) Butt, J. N.; Sucheta, A.; Armstrong, F. A.; Breton, J.; Thomson, A. J.; Hatchikian, E. C. J. Am. Chem. Soc. 1991, 113, 8948-8950. (c) Butt, J. N.; Sucheta, A.; Armstrong, F. A.; Breton, J.; Thomson, A. J.; Hatchikian, E. C.; J. Am. Chem. Soc. 1993, 115, 1413-1421. (d) Butt, J. N.; Fawcett, S. E. J.; Breton, J.; Thomson, A. J.; Armstrong, F. A. J. Am. Chem. Soc. 1997, 119, 97299737. (6) Butt, J. N.; Sucheta, A.; Martin, L. L.; Shen, B.; Burgess, B. K.; Armstrong, F. A. J. Am. Chem. Soc. 1993, 115, 12587-12588. (7) Duff, J. L. C.; Breton, J. L. J.; Butt, J. N.; Armstrong, F. A.; Thomson, A. J. J. Am. Chem. Soc. 1996, 118, 8593-8603. (8) Sucheta, A.; Ackrell, B. A. C.; Cochran, B.; Armstrong, F. A. Nature 1992, 356, 361-362. Sucheta, A.; Cammack, R.; Weiner, J.; Armstrong, F. A. Biochemistry 1993, 32, 5455-5465. Mondal, M. S.; Fuller, H. A.; Armstrong, F. A. J. Am. Chem. Soc. 1996, 118, 263-264. Hirst, J.; Sucheta, A.; Ackrell, B. A. C.; Armstrong, F. A. J. Am. Chem. Soc. 1996, 118, 5031-5038. Hirst, J.; Ackrell, B. A. C.; Armstrong, F. A. J. Am. Chem. Soc. 1997, 119, 74347439. Heering, H. A.; Weiner, J. H.; Armstrong, F. A. J. Am. Chem. Soc. 1997, 119, 11628-11638. Heering, H. A.; Hirst, J.; Armstrong, F. A. J. Phys. Chem. B 1998, 102, 6889-6902. Mondal, M. S.; Goodin, D. B.; Armstrong, F. A. J. Am. Chem. Soc. 1998, 120, 6270-6276. (9) Armstrong, F. A. In Bioelectrochemistry of Biomacromolecules. Bioelectrochemistry: Principles and Practice; Lenaz, G., Milazzo, G., Eds.; Birkhauser Verlag: Basel, 1997; pp 205-255. Armstrong, F. A.; Butt, J. N.; Sucheta, A. Methods Enzymol. 1993, 227, 479-500. Armstrong, F. A. Adv. Inorg. Chem. 1992, 38, 117-163. (10) Laviron, E. In Electroanalytical Chemistry; Bard, A. J., Ed.; Marcel Dekker: New York, 1982; Vol. 12, pp 53-157. (11) (a) Randin, J.-P.; Yeager, E. J. Electroanal. Chem. 1975, 58, 313-322. (b) Engstrom, R. C. Anal. Chem. 1982, 54, 2310-2314. (c) Engstrom, R. C.; Strasser, V. A. Anal. Chem. 1984, 56, 136-141. (d) Cabaniss, G. E.; Diamantis, A. A.; Murphy, W. R.; Linton, R. W.; Meyer, T. J. J. Am. Chem. Soc. 1985, 107, 1845-1853. (12) Muller, M.; Kastening, B. J. Electroanal. Chem. 1994, 374, 149-158. (13) McDermott, M. T.; McDermott, C. A.; McCreery, R. L. Anal. Chem. 1993, 65, 937-944. McCreery, R. L.; Cline, K. K.; McDermott, C. A.; McDermott, M. T. Colloids Surf. 1994, 93, 211-219. Cline, K. K.; McDermott, M. T.; McCreery, R. L. J. Phys. Chem. 1994, 98, 5314-5319. Chen, P.; Fryling, M. A.; McCreery, R. L. Anal. Chem. 1995, 67, 3115-3122.
protein film voltammetry has complemented pioneering studies on self-assembled monolayers (SAMs) of small molecules. Notably the groups of Chidsey,16 Creager,17 Murray,18 and Finklea19 have investigated the electron-transfer characteristics of metal complex-terminated alkanethiol SAMs adsorbed on gold electrodes, while Forster and co-workers20 have studied osmium complexes adsorbed onto platinum. These efforts have established more rigorous theoretical models for interfacial electron transfer by evaluating the predictions of Marcus theory and seeking to explain the deviations that are observed. For example, rate constants measured for cytochrome c adsorbed on (X(CH2)nS) SAMs show the expected dependence on distance.15 Another approach, used particularly by Rusling and co-workers, has aimed at immobilizing proteins within membrane-mimetic assemblies, typically consisting of liquid crystal surfactants.21,22 This has produced very stable electroactive arrays, although the electrontransfer kinetics have not so far been studied in detail. Given this background, we now seek to define what is achievable with our own systems, which in the past have been restricted to experiments performed at slow scan rates where reversibility is approached. The aim is to establish a foundation upon which to study the kinetics and energetics of processes such as proton transfer that are coupled to and may gate electron transfer.23 In this paper, we investigate the extent to which the voltammetry of small redox proteins remains interpretable at high scan rates by well-established theorysselecting examples and conditions for which simple uncoupled electron transfer ought to be observed. We use as our basis the Marcus24 and ButlerVolmer models.10 Four protein active sites have been studieds the blue Cu center (Cu2+/1+) of Pseudomonas aeruginosa azurin,25 and the [3Fe-4S]1+/0 cluster in Azotobacter vinelandii ferredoxin I (Av Fd I), a mutant form of A. vinelandii ferredoxin I (D15N)6,26 and Sulfolobus acidocaldarius 7Fe ferredoxin (Sa Fd).7 (14) Song, S.; Clark, R. A.; Bowden, E. F.; Tarlov, M. J. J. Phys. Chem. 1993, 97, 6564-6572. Nahir, T. M.; Clark, R. A.; Bowden, E. F. Anal. Chem. 1994, 66, 2595-2598. Nahir, T. M.; Bowden, E. F.; J. Electroanal. Chem. 1996, 410, 9-13. Clark, R. A.; Bowden, E. F.; Langmuir 1997, 13, 559-565. Kasmi, A. E.; Wallace, J. M.; Bowden, E. F.; Binet, S. M.; Linderman, R. J. J. Am. Chem. Soc. 1998, 120, 225-226. (15) Feng, Q. F.; Imabayashi, S.; Kakiuchi, T.; Niki, K. J. Electroanal. Chem. 1995, 394, 149-154. (16) Chidsey, C. E. D. Science 1991, 251, 919-922. (17) Weber, K.; Creager, S. E. Anal. Chem. 1994, 66, 3164-3172. (18) Tender, L.; Carter, M. T.; Murray, R. W. Anal. Chem. 1994, 66, 31733181. (19) Finklea, H. O.; Liu, L.; Ravenscroft, M. S.; Punturi, S. J. Phys. Chem. 1996, 100, 18852-18858. (20) (a) Forster, R. J.; Faulkner, L. R. J. Am. Chem. Soc. 1994, 116, 5444-5452. (b) Forster, R. J.; Faulkner, L. R. J. Am. Chem. Soc. 1994, 116, 54535461. (c) Forster, R. J.; O’Kelly, J. P. J. Phys. Chem. 1996, 100, 3695-3702. (21) Nassar, A. E. F.; Willis, W. S.; Rusling, J. F. Anal. Chem. 1995, 67, 23862392. Tominaga, M.; Yanagimoto, J.; Nassar, A. E. F.; Rusling, J. F.; Nakashima, N. Chem. Lett. 1996, 7, 523-524. Zhang, Z.; Rusling, J. F. Biophys. Chem. 1997, 63, 133-146. Kong, J.; Lu, Z.; Lvov, Y. M.; Desamero, R. Z. B.; Frank, H. A.; Rusling, J. F. J. Am. Chem. Soc. 1998, 120, 73717372. (22) Bianco, P.; Haladjian, J. Electrochim. Acta 1997, 42, 587-594. (23) Hirst, J.; Duff, J. L. C.; Jameson, G. N. L.; Kemper, M. A.; Burgess, B. K.; Armstrong, F. A. J. Am. Chem. Soc. 1998, 120, 7085-7094. (24) Marcus, R. A.; Sutin, N. Biochim. Biophys. Acta 1985, 811, 265-322. (25) Adman, E. T.; Jensen, L. H. Isr. J. Chem. 1981, 21, 8-12. Adman E. T.; Canters, G. W.; Hill, H. A. O.; Kitchen, N. A. FEBS Lett. 1982, 143, 287292. Nar, H.; Messerschmidt, A.; Huber, R.; Van de Kamp, M.; Canters, G. W. J. Mol. Biol. 1991, 221, 765-772. (26) Shen, B.; Martin, L. L.; Butt, J. N.; Armstrong, F. A.; Stout, C. D.; Jensen, G. M.; Stephens, P. J.; La Mar, G. N.; Gorst, C. M.; Burgess, B. K. J. Biol. Chem. 1993, 268, 25928-25939.
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SUBJECTS OF STUDY Pseudomonas aeruginosa Azurin. Azurin is a small (128 residues) “blue”-copper protein, in this case isolated from P. aeruginosa. The crystal structure is well defined,25 and the copper site (Cu2+/1+) is buried ∼7 Å beneath the solvent-accessible surfacesan area of hydrophobic residues believed to be important for interaction with enzyme redox partners such as nitrite reductase.27 All experiments were carried out at pH 8.5, where the reduction potential is almost independent of pH, to minimize any complications due to protonation equilibria.28 Azotobacter vinelandii Ferredoxin I: Wild Type and D15N Mutant. Ferredoxin I from A. vinelandii is a structurally defined 7Fe ferredoxin of 106 residues, containing one [3Fe-4S] and one [4Fe-4S] cluster.29 Extensive studies have already been made using conventional scan rates (
