Cytochrome c Oxidase Immobilized in Stable Supported Lipid Bilayer

Apr 3, 1998 - Planar Supported Lipid Bilayer Polymers Formed by Vesicle Fusion. 2. Adsorption of Bovine Serum Albumin. Eric E. Ross, Tony Spratt, Sanc...
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Langmuir 1998, 14, 2467-2475

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Cytochrome c Oxidase Immobilized in Stable Supported Lipid Bilayer Membranes James D. Burgess,† Melissa C. Rhoten, and Fred M. Hawkridge* Department of Chemistry, Virginia Commonwealth University, Box 842006, Richmond, Virginia 23284 Received November 3, 1997. In Final Form: February 17, 1998 The voltammetry of cytochrome c oxidase immobilized in lipid bilayer membranes on gold electrodes and amperometric data of cytochrome c reacting at these electrodes under flow conditions are reported. A submonolayer of octadecyl mercaptan formed on electrodeposited silver anchors and becomes a part of the lipid bilayer membrane on the gold electrode. The supported lipid bilayer membrane containing cytochrome c oxidase is formed during a deoxycholate dialysis procedure. Slow scan rate cyclic voltammograms (20 mV/s) taken at the oxidase-modified electrodes show well-defined anodic waves. Fast scan rate cyclic voltammograms (200 mV/s) taken at the oxidase-modified electrodes show well-defined anodic and cathodic waves. Cyclic voltammograms taken at the oxidase-modified electrodes under 0.1 mM sodium cyanide show an increase (ca. 300%) in electrode capacitance and well-defined anodic and cathodic waves irrespective of scan rate. The voltammetric data are consistent with electron transfer of cytochrome c oxidase coupled with changes in nonfaradaic current and possibly diffusion of cytochrome c oxidase in a lipid multilayer structure. Quartz crystal microbalance data of cytochrome c binding to lipid bilayer membranes containing no cytochrome c oxidase under flow conditions are presented.

Introduction There has been progress in elucidating the conditions necessary for direct electron transfer between small heme proteins and solid electrodes.1 However, the use of voltammetry to study electron-transfer reactions of enzymes, without redox mediators or electron relays within the enzyme, is not common. Immobilization of enzymes in supported lipid bilayer membranes is one strategy for achieving direct electron transfer between solid electrodes and enzymes.2,3 Results reported by Cullison et al.3 showed that cytochrome c oxidase can be immobilized in a lipid bilayer membrane on a gold electrode so that it undergoes direct electron transfer with the gold electrode. Voltammetric and spectroelectrochemical results showed that the oxidase enzyme is able to mediate electron transfer between the gold electrode and solution-resident cytochrome c, its native redox partner. In this initial work the oxidase-modified electrode was formed by depositing a submonolayer of octadecyl mercaptan (OM) onto a gold electrode using thiol self-assembly chemistry and then constructing a lipid bilayer membrane containing the oxidase enzyme using deoxycholate dialysis. The OM submonolayer becomes part of the subsequently formed bilayer, anchors this bilayer to the electrode, and increases the interfacial hydrophobicity at the gold surface without blocking access of the oxidase enzyme to the gold electrode. Dialysis tubing containing the thiol-modified electrodes and a solution of lipids, deoxycholate, and the oxidase enzyme was placed in a beaker filled with dialysis buffer. As deoxycholate was removed from the oxidase-lipid solution, a lipid bilayer membrane containing cytochrome † Current address: Microanalytical Instrumentation Center, Ames Laboratory-USDOE, and Department of Chemistry, Iowa State University, Ames, IA 50011. * To whom correspondence should be addressed.

(1) Hawkridge, F. M.; Taniguchi, I. Comments Inorg. Chem. 1995, 17, 163. (2) Salamon, Z.; Hazzard, J. T.; Tollin, G. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 6420. (3) Cullison, J. K.; Hawkridge, F. M.; Nakashima, N.; Yoshikawa, S. Langmuir 1994, 10, 877.

c oxidase on the electrode formed. This membraneforming procedure was based on previously reported dialysis procedures for reconstituting cytochrome c oxidase into solution-resident vesicles.4 Unfortunately, only about 20% of these electrodes were successfully modified.3 Better control of both the OM electrode surface coverage and the dialysis step of the modification scheme first reported by Cullison et al.3 is reported here. A quartz crystal microbalance (QCM) was used to control the OM self-assembly reaction at electrodeposited silver on gold.5 Reproducing thiol self-assembly kinetics was the sole reason for using an electrodeposited silver surface. Two other advantages of the silver-sulfur bond over the goldsulfur bond can be cited. The silver-sulfur bond accommodates an orientation of the thiol hydrocarbon tail that is closer to normal with respect to the surface plane than does the gold-sulfur bond.6,7 A near normal orientation of OM with respect to the surface plane may maximize hydrophobic interactions between the hydrocarbon tails of the lipids and the thiol hydrocarbon tails which anchor the lipid bilayer to the electrode. Also, for a given OM surface coverage with the hydrocarbon tails oriented near normal with respect to the surface plane, more underlying silver would be exposed for close approach by the oxidase enzyme than with a tilted orientation of the OM molecules. Second, the silver-sulfur bond is believed to be stronger than the gold-sulfur bond,7-9 perhaps yielding a more stable lipid membrane. Stable lipid bilayer membranes have been formed on platinum by liposome fusion onto Langmuir-Blodgett lipid monolayers, and this work is consistent with the method described here for forming stable lipid bilayer membranes on electrodeposited silver (4) Hinkle, P. C.; Kim, J. J.; Racker, E. J. Biol. Chem. 1972, 247, 1338. (5) Burgess, J. D.; Hawkridge, F. M. Langmuir 1997, 13, 3781. (6) Sellers, H.; Ulman, A.; Shnidman, Y.; Eilers, J. E. J. Am. Chem. Soc. 1993, 115, 9389. (7) Bryant, M. A.; Pemberton, J. E. J. Am. Chem. Soc. 1991, 113, 8284. (8) Jennings, G. K.; Laibinis, P. E. Langmuir 1996, 12, 6173. (9) Jennings, G. K.; Laibinis, P. E. J. Am. Chem. Soc. 1997, 119, 5208.

S0743-7463(97)01199-2 CCC: $15.00 © 1998 American Chemical Society Published on Web 04/03/1998

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derivatized with an OM submonolayer.10 Lipid monolayers have also been self-assembled onto alkanethiol monolayers on gold from lipid vesicle solution.11 The original dialysis procedure was improved through use of a dual-chambered electrochemical cell. The volume of the sample, 0.5 mL, and the distance between the electrode and the dialysis membrane are fixed with this cell arrangement. Also, the volume of buffer that the electrodes are dialyzed against can be easily controlled. Another advantage of this cell is that dialysis, voltammetry, and flow experiments12 (QCM and amperometric) can be conducted without further handling of the electrode. Complete reaction of alkanethiols with gold and silver surfaces results in a chemisorbed monomolecular film with the hydrocarbon tails extending out from the plane of the metal surface and with the sulfur bound to the metal substrate, presumably as a thiolate.6,7,13,14 Some insight into the mechanism of thiol film formation on gold15 and silver16 has been gained from scanning tunneling microscopy. Also, atomic force microscopy (AFM)17 has been used to study alkanethiol self-assembly on gold. Thiol domain formation has been reported on gold15,17 and silver16 when self-assembled from ethanol. Lateral hydrophobic interactions between thiol molecules have been proposed to explain this ordering event.17 At low coverages the thiol molecules are believed to undergo lateral diffusion on the metal surfaces.15-17 Alkanethiol submonolayers formed on gold under potential control have been shown to stimulate physisorption of thiol multilayers from acetonitrile.18 OM submonolayers formed on the electrodeposited silver surfaces used in this work may stimulate lipid bilayer formation on the electrode by allowing hydrophobic interactions between the thiol octadecyl tails and the lipid hydrocarbon tails. Similar interactions between surface confined OM and deoxycholate are expected. On the basis of this background, the OM molecules that anchor the lipid bilayer to the electrode are envisioned as being oriented with the hydrocarbon tail near normal with respect to the surface plane. If thiol islands exist on the substrates used in this work prior to lipid deposition, lateral hydrophobic interactions between the OM hydrocarbon tails and the lipids, deoxycholate, and cytochrome c oxidase may disperse the islands, leading to a thiol submonolayer with the thiol molecules more evenly spaced over the electrode surface. Beef heart cytochrome c oxidase has 13 subunits and a molecular weight of ca. 204 000 amu.19 It is the terminal enzyme of oxidative phosphorylation and is located in the inner mitochondrial membrane, where it reduces molecular oxygen to water. Cytochome c oxidase also pumps protons against a concentration gradient across the inner mitochondrial membrane to support the production of ATP.20-28 The enzyme contains four redox-active sites in (10) Puu, G.; Gustafson, I.; Artursson, E.; Ohisson, P.-Å Biosensors Bioelectron. 1995, 10, 463. (11) Plant, A. L. Langmuir 1993, 9, 2764. (12) Ruzicka, J. Analyst 1994, 119, 1925. (13) Widrig, C. A.; Chung, C.; Porter, M. D. J. Electroanal. Chem. 1991, 310, 335. (14) Laibinis, P. E.; Fox, M. A.; Folkers, J. P.; Whitesides, G. M. Langmuir 1991, 7, 3167. (15) Poirier, G. E.; Pylant, E. D. Science 1996, 272, 1145. (16) Dhirani, A.; Hines, M. A.; Fisher, A. J.; Ismail, O.; GuyotSionnest, P. Langmuir 1995, 11, 2609. (17) Tamada, K.; Hara, M.; Sasabe, H.; Knoll, W. Langmuir 1997, 13, 1558. (18) Schneider, T. W.; Buttry, D. A. J. Am. Chem. Soc. 1993, 115, 12391. (19) Tsukihara, T.; Aoyama, H.; Yamashita, E.; Tomizaki, T.; Yamaguchi, H.; Shinzawa-Itoh, K.; Nakashima, R.; Yaono, R.; Yoshikawa, S. Science 1996, 272, 1136. (20) Wikstro¨m, M. Nature 1977, 266, 271.

Burgess et al.

two transmembrane subunits (I and II) and has 28 transmembrane R helices.19 The binuclear Cua redox site of subunit II may be the primary electron acceptor from ferrocytochrome c,23 and it is contained in a hydrophilic portion of the enzyme that extends out of the membrane into the cytoplasm.28 The other three redox sites (heme a, heme a3, and Cub) are contained within the transmembrane portion of subunit I.28 Negative cooperative coupling between the redox sites has been reported,29-32 and this cooperativity may play a central role in electron gating, which is believed to be a requirement for proton translocation.22 Possible proton, water, and oxygen channels have been identified from the crystal structure,19 and a proton pump model involving two conformation states has been proposed.22 The position of the oxidase enzyme relative to the inner mitochondrial membrane has been assessed. Electron microscopy of two-dimensional vesicle crystals,33-36 and of platinum/carbon replicas of freezefractured vesicle surfaces containing cytochrome c oxidase37 show a dimeric form of the oxidase enzyme asymmetrically positioned in the lipid bilayer with a large domain (containing the Cua redox site) extending into what would be the cytoplasm and with only a small portion the enzyme exposed on the other side of the membrane (matrix). The three-dimensional X-ray structure of cytochrome c oxidase at 2.8 Å resolution has been reported for crystals formed in the presence of decylmaltoside using poly(ethylene glycol) as the precipitate.19,28 In these structures, the oxidase enzyme is also present as a dimer but a more symmetric orientation of the enzyme in the lipid bilayer is predicted. In the three-dimensional crystals, the location of the Cua site is 8 Å above the cytoplasmic membrane surface and the other three redox sites are 13 Å below the cytoplasmic membrane surface in the 48 Å thick transmembrane portion of the enzyme.28 These locations of the metal sites are in general agreement with locations of the metal sites in prokaryotic cytochrome c oxidase as inferred from site-directed mutagenesis studies37 and X-ray crystal structures.39,40 Earlier work (21) Malmstro¨m, B. G. Chem. Rev. 1990, 90, 1247. (22) Babcock, G. T.; Wikstro¨m, M. Nature 1992, 356, 301. (23) Ramirez, B. E.; Malmstro¨m, B. G.; Winkler, J. R.; Gray, H. B. Proc. Natl. Acad. Sci. USA 1995, 92, 11949. (24) Winkler, J. R.; Malmstro¨m, B. G.; Gray, H. B. Biophys. Chem. 1995, 54, 199. (25) Malatesta, F.; Antonini, G.; Sarti, P.; Brunori, M. Biophys. Chem. 1995, 54, 1. (26) Einarsdo´ttir, O Ä . Biochim. Biophys. Acta 1995, 1229, 129. (27) Ferguson-Miller, S.; Babcock, G. T. Chem. Rev. 1996, 96, 2889. (28) Tsukihara, T.; Aoyama, H.; Yamashita, E.; Tomizaki, T.; Yamaguchi, H.; Shinzawa-Itoh, K.; Nakashima, R.; Yaono, R.; Yoshikawa, S. Science 1995, 269, 1069. (29) Ellis, W. R.; Wang, D. F.; Blair, D. F.; Gray, H. B.; Chan, S. I. Biochemistry 1986, 25, 161. (30) Blair, D. F.; Ellis, W. R., Jr.; Wang, H.; Gray, H. B.; Chan, S. I. J. Biol. Chem. 1986, 261, 11524. (31) Moody, A. J.; Rich, P. R. Biochim. Biophys. Acta 1990, 1015, 205. (32) Nilsson, T. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 6497. (33) Henderson, R.; Capaldi, R. A.; Leigh, J. S. J. Mol. Biol. 1977, 112, 631. (34) Deatherage, J. F.; Henderson, R.; Capaldi, R. A. J. Mol. Biol. 1982, 158, 487. (35) Deatherage, J. F.; Henderson, R.; Capaldi, R. A. J. Mol. Biol. 1982, 158, 501. (36) Valpuesta, J. M.; Henderson, R.; Frey, T. G. J. Mol. Biol. 1990, 214, 237. (37) Tihova, M.; Tattrie, B.; Nicholls, P. Biochem. J. 1993, 292, 933. (38) Hosler, J. P.; Ferguson-Miller, S.; Calhoun, M. W.; Thomas, J. W.; Hill, J.; Lemieux, L.; Ma, J.; Georgiou, C.; Fetter, J.; Shapleigh, J.; Tecklenburg, M. M. J.; Babcock, G. T.; Gennis, R. B. J. Bioenerg. Biomembr. 1993, 25, 121. (39) Wilmanns, M.; Lappalainen, P.; Kelly, M.; Sauer-Eriksson, E.; Saraste, M. Proc. Natl. Acad. Sci. U.S.A. 1995, 92, 11955. (40) Iwata, S.; Ostermeier, C.; Ludwig, B.; Michel, H. Nature 1995, 376, 660.

Cytochrome c Oxidase in Lipid Bilayer Membranes

has the heme a3 site ca. 12 Å closer to the matrix side of the membrane relative to the heme a site.41-43 Cytochrome c oxidase can exist in several forms in solution,44 and conformational heterogeneity is of concern.45,46 Cyanide binds to the oxygen binding site of the oxidase enzyme and this binding is lethal.47 The kinetics of cyanide binding to cytochrome c oxidase have been used to characterize the various states of the enzyme.44,46 Reported here are voltammetric and potential step data of cytochrome c oxidase-modified electrodes formed under optimized dialysis conditions on electrodeposited silver surfaces derivatized with a controlled amount of OM.5 Controlled potential amperometry under sample injection conditions using a wall-jet configuration was used to detect reduced cytochrome c undergoing an oxidation reaction at the oxidase-modified electrodes. Although not strictly a flow-injection analysis (FIA)12 experiment because a wall-jet configuration is used, for conciseness this will be termed an FIA experiment. QCM data were collected during some of these experiments (QCM-FIA), and no incorporation of cytochrome c into the membrane was detected.

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Figure 1. Cell used for the oxidase experiments. The dimensions of the cell are 2 cm × 5 cm × 5 cm. The quartz crystal microbalance electrode is clamped to the sample chamber. The dialysis membrane is clamped between the flow chamber and the sample chamber.

Experimental Section Instrumentation. The cyclic voltammetric experiments were performed using a potentiostat built in-house that was controlled by a computer equipped with a data acquisition board (National Instruments) and LabView graphical programming software (National Instruments). The square wave voltammetry and potential step experiments were performed using a computercontrolled potentiostat (1090-CS, Cypress Systems). The differential pulse experiments were performed using an PAR 174 polarographic analyzer (BAS). The FIA current responses were measured using the potentiostat built in-house. The QCM was built in-house based on a circuit published by Bruckenstein et al.48 The difference frequency of the QCM is passed through a 7414 NAND gate Schmitt Trigger before being counted using the MIO-16DE-10 DAQ board and LabView graphical programming software. Electrochemical Dialysis Cell. The cell was machined from two separate pieces of Lucite. One piece houses the reference (Ag/AgCl, 1 M KCl) and auxiliary (coiled platinum wire) electrodes. The gold QCM electrode is clamped to this piece to form one wall of the sample chamber (volume ca. 0.5 mL). The other piece contains a buffer flow chamber with a volume of ca. 0.5 mL. A dialysis membrane (molecular weight cutoff 3500, Spectrapor, Spectrum Medical Industries, Inc.) is placed between the two cell pieces separating the two chambers. The cell assembly is then clamped together as shown in Figure 1. The Ag/AgCl, 1 M KCl reference electrode was calibrated using a platinum wire immersed in a saturated solution of quinhydrone (Eastman) of known pH.49 All potentials are reported versus the normal hydrogen electrode (NHE). Substrate and Lipid Preparation. The preparation of the OM submonolayer on electrodeposited silver5 and the preparation of the deoxycholate-lipid stock solution3 have been reported elsewhere. All experiments shown were conducted at gold QCM electrodes derivatized with 1.6 monolayers of electrodeposited (41) Holm, L.; Saraste, M.; Wikstro¨m, M. EMBO J. 1987, 6, 2819. (42) Ohnishi, T.; LoBrutto, R.; Salerno, J. C.; Bruckner, R. C.; Frey, T. G. J. Biol. Chem. 1982, 257, 14821. (43) Blasie, J. K.; Pachence, J. M.; Tavormina, A.; Dutton, P. L.; Stamatoff, J.; Eisenberger, P.; Brown G. Biochim. Biophys. Acta 1982, 679, 188. (44) Palmer, G. J. Bioenerg. Biomembr. 1993, 25, 145. (45) Malmstro¨m, B. G. Arch. Biochem. Biophys. 1990, 280, 233. (46) Lodder, A. L.; Gelder, B. F. Biochim. Biophs. Acta 1994, 1186, 67. (47) Lee, S. C.; Scott, M. J.; Kauffmann, K.; Mu¨nck, E.; Holm, R. H. J. Am. Chem. Soc. 1994, 116, 401. (48) Bruckenstein, S.; Michaliski, M.; Fensore, A.; Li, Z.; Hillman, A. R. Anal. Chem. 1994, 66, 1847. (49) Bates, R. G. Determination of pH: Theory and Practice; JohnWiley & Sons: New York, 1985; p 299.

Figure 2. Cyclic voltammograms taken at three cytochrome c oxidase-modified electrodes (unlabeled scans). Scan (a) is a cyclic voltammogram taken at a lipid bilayer modified electrode containing no cytochrome c oxidase. The scan rate is 20 mV/s, the buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2.

Figure 3. Scan rate dependence and stability of the cyclic voltammetry taken at a cytochrome c oxidase-modified electrode. The scan rates are (a) 20 mV/s, (b) 10 mV/s, (c) 30 mV/s, and (d) 20 mV/s. The buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2. silver, except for the cyclic voltammograms shown in Figure 4 where a monolayer of electrodeposited silver was the substrate. Higher silver coverages were used to further improve the reproducibility of both the OM self-assembly reactions and the oxidase experiments. Only slow scan rates were used to characterize the oxidase-modified electrodes formed on 1.6 monolayers of electrodeposited silver in order to minimize the potential window encompassed by the cyclic voltammetry experiments and to improve the FIA reproducibility from electrode to electrode. The deoxycholate-lipid stock solution contained 40.2 mM sodium deoxycholate (Sigma), 11.2 mM (DOPE) l-phosphatidylethanolamine, dioleoyl (Sigma, 99%), and 2.5 mM (DOPC) L-phosphatidylcholine, dioleoyl (Sigma, 99%) in phosphate buffer (0.1 M, ACS reagent grade, pH 7.4).3 Two changes in the preparation of the deoxycholate-lipid stock solution are noted; the lipid storage solvent and ether washes were

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Figure 4. Scan rate dependence and stability of the cyclic voltammetry taken at an oxidase-modified electrode. The scan rates are; (a) and (b) 250 mV/s, (c) and (d) 200 mV/s, (e) and (f) 150 mV/s. The buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2. removed by evaporation with a stream of prepure nitrogen instead of using a rotary evaporator, and the lipids were treated with Chelex-100 (ion-exchange resin, BioRad) for ca. 12 h at 4 °C without agitation to remove metal cation contaminants. The Chelex-100 was removed by filtration (UNIFLO plus 0.2 µm CA and glass fiber) of the deoxycholate-lipid solution. The water used for all experiments was deionized and further purified with a Milli RO-4/Milli-Q system (Millipore Corp.) to exhibit a resistivity of 17-18 MΩ‚cm. Atomic force microscopy (AFM, Seiko 3700) was used to characterize the surface roughness of the bare gold quartz crystal microbalance electrodes. Dialysis Experiment. For the preparation of a cytochrome c oxidase-modified electrode, an aliquot of the stock deoxycholatelipid solution was mixed with an equal volume of buffer (0.01 M Tris/HCl pH 7.6, 0.1% Triton X-100) containing the oxidase enzyme (2 mg/mL) and the mixture was added to the sample chamber of the cell. For the preparation of the lipid bilayer modified electrodes containing no cytochrome c oxidase (control experiments), an aliquot of the stock deoxycholate-lipid solution was mixed with an equal volume of 0.1 M phosphate buffer (ACS reagent grade, pH 7.4) and added to the sample chamber of the cell. Phosphate buffer was pumped (syringe pump, Harvard Apparatus, Model 944) through the flow chamber of the cell at a rate of 50 µL/min for g18 h. The sample chamber was then flushed by pumping phosphate buffer through the wall jet (see Figure 1) for ca. 20 h at a rate of 5 µL/min. For studies over several days, phosphate buffer was pumped through the sample chamber of the cell at night at a flow rate of 5 µL/min. Bovine cytochrome c oxidase was isolated from fresh beef hearts essentially following the procedure published by Soulimane and Buse.50 Flow Experiments. All FIA experiments were carried out at a flow rate of 0.5 mL/min using the Harvard Apparatus syringe pump. The linear flow rate of the buffer through the tip of the wall jet is ca. 6 cm/s, and the wall-jet tip is positioned ca. 3 mm from the working electrode. Phosphate buffer 0.1 M, pH 7.4 was used for all FIA experiments shown except for the QCM-FIA experiments probing the electrostatic association of cytochrome c with lipid bilayer modified electrodes containing no oxidase and with oxidase-modified electrodes where 0.01 M phosphate buffer, pH 7.4 was used. Horse heart cytochrome c (Sigma Chemical Co., 98%) was purified with a cation-exchange column (mono-S, Pharmacia) using an FPLC chromatography system (Pharmacia). The cytochrome c samples were reduced with dithionite (Sigma Chemical Co.) and desalted using a 5 mL, HighTrap size-exclusion column (Pharmacia). The protein concentrations were determined with a Hewlett-Packard Model 8452A diode array UV/vis spectrophotometer using a molar absorptivity for reduced cytochrome c of 29 500 M-1 cm-1 at 550 nm.51 Two syringes were held in the pump, one containing buffer and the other containing reduced cytochrome c in buffer. Injections were (50) Soulimane, T.; Buse, G. Eur. J. Biochem. 1995, 227, 588. (51) Van Gelder, B. F.; Slater, E. C. Biochim. Biophys. Acta 1962, 58, 593.

Burgess et al. performed manually using a six-way valve to direct either buffer or buffer containing cytochrome c through the wall-jet inlet of the sample chamber. All FIA experiments probing the oxidation of cytochrome c at the modified electrodes were conducted at 472 mV vs NHE. The QCM-FIA experiments were conducted using reduced or oxidized cytochrome c and at an applied potential of 472 mV vs NHE or under open circuit. Experimental Procedure. The process of immobilizing cytochrome c oxidase in a lipid bilayer membrane on a gold electrode is described as follows. A submonolayer of OM formed on silver electrodeposited onto gold QCM electrodes is used as the substrate.5 The thiol-modified electrode is clamped to the dual-chambered electrochemical dialysis cell (see Figure 1). The sample chamber is filled with a solution containing deoxycholate, biological amphiphiles, buffer, and the oxidase enzyme. Buffer is passed through the flow chamber, which is separated from the sample chamber by a dialysis membrane, to dialyze the enzyme solution. This procedure drives the formation of a lipid bilayer membrane containing cytochrome c oxidase on the thiol-modified electrode. After dialysis, the sample chamber is flushed with buffer to remove remaining solution-resident species. This enzyme immobilization scheme produces a stable lipid membrane environment that allows the study of heterogeneous electron transfer between cytochrome c oxidase and the gold electrode and between solution-resident cytochrome c and the immobilized oxidase.

Results and Discussion All of the cyclic voltammograms shown were taken after the sample chamber of the cell had been flushed with phosphate buffer for at least 20 h after dialysis. The voltammetric waves observed at the oxidase-modified electrodes grow during this time. The time dependence may be due to the removal of deoxycholate from the lipid membrane and/or to the desorption of lipid multilayers from the electrode surface.3 All of the cyclic voltammograms shown were taken at 10 min intervals, and the working electrode was held at the initial potential of 122 mV vs NHE between scans. The magnitude of the cyclic voltammogram peak currents observed at the oxidase-modified electrodes varied with the time elapsed between scans, increasing for delays up to ca. 30 min, and the best stability was obtained for delays of 10-15 min. This time dependence may be due to the reductive removal of oxygen from the membrane. FIA data (not shown) show that oxygen present in buffer is reduced at the oxidase-modified electrode at 122 mV (i.e., the voltammetric initial potential). After ca. 20 h of flushing and then repeated cycling (ca. 4 scans) of the potential at 10 min intervals, about half of the oxidasemodified electrodes exhibit reasonable stability. Other oxidase-modified electrodes require additional equilibration for days (2-3) before voltammetric stability is observed. Figure 2 shows the reproducibility obtained for three consecutive oxidase experiments carried out under fixed conditions and a control experiment for only a lipid bilayer membrane containing no oxidase enzyme (see Experimental Section). All four of the experiments shown in Figure 2 were conducted at separate gold electrodes. Scans taken at the lipid bilayer modified electrodes containing no oxidase are essentially flat, as shown in Figure 2a. Cyclic voltammetric scans taken at various oxidase-modified electrodes formed under the same conditions differ somewhat in peak current, peak potential, and wave shape (see Figure 2). These variations are likely due to differences in the substrate, the structure of the lipid membrane, and the position of the oxidase enzyme relative to the electrode surface. The anodic waves observed in the cyclic voltammetry data obtained at the oxidase-modified electrodes are apparent for days under phosphate buffer at room temperature although there is

Cytochrome c Oxidase in Lipid Bilayer Membranes

some change in peak shape and peak position from day to day. Further removal of dexoycholate from the lipid membrane and desorption of lipid multilayers may occur over time.3 One of the cyclic voltammograms shown in Figure 2 has a relatively broad anodic wave compared to the other two scans. As mentioned above, Figure 2 shows three consecutive oxidase experiments and about half of the oxidase-modified electrodes exhibit broad anodic waves such as the one shown in Figure 2. About 10-15% of the oxidase-modified electrodes show anodic peak potentials that are more negative (e.g., as much as ca. 50 mV) than those shown in Figure 2. AFM was used to characterize the surface roughness of the bare gold QCM electrodes. The images of the gold surfaces show rolling hills with a valley to peak height of about 5 nm and a valley to valley distance of about 300 nm. Also, some narrow deep defects were apparent, covering no more than ca. 5% of the surface. This structure may not significantly affect the packing of the lipid bilayer membrane. Figure 3 shows the scan rate dependence and the stability of the cyclic voltammetry data taken at an oxidase-modified electrode. Four scans are shown and the sequence of the scans is 20 mV/s (Figure 3a), 10 mV/s (Figure 3b), 5 mV/s (data not shown), 30 mV/s (Figure 3c), and 20 mV/s (Figure 3d). Thus the time elapsed between the first and last 20 mV/s scans was 40 min. Scans (a) and (d) of Figure 3 are nearly superimposed, a result that is consistent with complete immobilization of the oxidase enzyme in the supported lipid bilayer membrane. As mentioned above, some oxidase-modified electrodes show either an increase or a decrease in peak current (e.g., 10% of the peak current) for the same scan rate conducted first and last in a comparable set of scans and such electrodes will often stabilize upon additional equilibration (days). The voltammetric peak shape is dependent on the hydrodynamic history of the electrode (e.g., flowing buffer through the wall jet causes a decrease in the anodic voltammetric peak current). This observation may be consistent with the reported pressure-induced effects on the oxidase enzyme.52 A striking feature of the voltammograms taken at the oxidase-modified electrodes is the large amount of charge passed during the anodic scans. If cytochrome c oxidase is immobilized on the electrode surface in a single lipid bilayer with its large hydrophilic domain extending out into solution and its hydrophobic domain stabilized in the lipid membrane,53 then the majority of the charge under the oxidative wave results from nonfaradaic current coupled to electron transfer. The estimated charge associated with a 50% electrode coverage of oxidase, assuming a molecular diameter of 80 Å,28 4 electrons per molecule, and no contribution from nonfaradaic current is ca. 130 nC. However, the charge passed during one of the anodic scans shown in Figure 2 is ca. 33 µC. The charge passed during the anodic scan of the control experiment shown in Figure 2a is ca. 4 µC. As mentioned above, the enzyme cycles between two conformational states during turnover.22,54,55 The two conformational states are believed to be associated with the proton pump input and output states,21 and the apparent nonfaradaic contribution to the charge passed during anodic voltam(52) Kornblatt, J. A.; Hoa, G. H. B.; Heremans, K. Biochemistry 1988, 27, 5122. (53) Creighton, T. E. Proteins: Structures and Molecular Properties; W. H. Freeman and Co.: New York, 1984; Chapter 4. (54) Copeland, R. A. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 7281. (55) Einarsdo´ttir, O Ä .; Georgiadis, K. E.; Sucheta, A. Biochemistry 1995, 34, 496.

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metric scans may be the result of the conformational change that occurs in cytochrome c oxidase during turnover. This conformational transition may significantly change the capacitance of the membrane by allowing ion movement into and out of the lipid bilayer. On the other hand, assuming that there is no nonfaradaic current coupled to electron transfer, a surface coverage of oxidase is estimated, from the difference between the charge passed during the anodic scan of a cyclic voltammogram taken at an oxidase-modified electrode (see Figure 2) and the charge passed during the anodic scan taken at a control membrane (see Figure 2a) (e.g., the difference charge is ca. 29 µC), to be ca. 110 monolayers. However, FIA data of cytochrome c reacting at the oxidase-modified electrodes are not consistent with a thick oxidase membrane (vide infra). The capacitive nature of the lipid membranes formed in this work may support the possibility that nonfaradaic current is coupled to electron transfer. The capacitance of the lipid membranes (e.g., 50 µF/cm2) is much higher than the capacitance values reported for different supported lipid bilayer membranes (ca. 1 µF/cm2).56,57 The presence of cyanide in solution causes an increase of ca. 300% in electrode capacitance as measured by cyclic voltammetry experiments conducted at both lipid bilayer modified electrodes containing no cytochrome c oxidase and at oxidase-modified electrodes (vide infra). The conformational change that occurs in cytochrome c oxidase upon reduction, which is strongly coupled to electron transfer,58 is apparently followed by a slower conformational change and/or reorientation of the enzyme in the lipid bilayer membrane. Figure 4 shows six voltammograms taken at an oxidase-modified electrode at three different scan rates. The 200 mV/s scan rate experiments (Figure 4c,d) were conducted first and last, and the six scans shown were taken at 10 min intervals. As shown in Figure 4, a cathodic wave is observed at scan rates faster than ca. 100 mV/s (compare Figure 3 and Figure 4) and the ratio of the cathodic peak current to the anodic peak current increases with increasing scan rate (see Figure 4). Much smaller cathodic waves have been observed at a few oxidase-modified electrodes. On the time scale probed by the cyclic voltammetry experiments, the oxidase enzyme apparently reorients and/or undergoes a slow conformational change upon being reduced such that electron transfer between the electrode and the enzyme is unfavorable and slow. If the oxidase enzyme is free to diffuse in a lipid multilayer membrane on the electrode, the observed time dependence of the cathodic wave likely results due to a slow conformational change of the oxidase enzyme that follows the fast conformational change associated with its reduction.58 If the oxidase is immobilized in a lipid bilayer, reorientation of the oxidase enzyme relative to the electrode surface may be involved in the observed time dependence of the cathodic wave. At a scan rate of 20 mV/s, about 10 s elapse after oxidation of the enzyme and before a reductive potential is reached, giving the oxidase time to reorient or change conformation. At faster scan rates (i.e., 200 mV/s), the time elapsed after oxidation of the enzyme and before a reducing potential is reached is much shorter (ca. 3 s). Thus, complete change of conformation and/or reorientation is not allowed and reduction is favored. The possibility that the conformational transition that is strongly coupled to electron transfer58 is slow in this supported membrane is not consistent with the large current observed in the voltam(56) Tien, H. T.; Salamon, Z. J. Electroanal. Chem. 1989, 276, 211. (57) Lang, H.; Duschl, C.; Vogel, H. Langmuir 1994, 10, 197. (58) Michel, B.; Bosshard, H. R. Biochemistry 1989, 28, 244.

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Figure 5. Potential step experiments conducted at an oxidasemodified electrode (a) and at a lipid bilayer modified electrode containing no cytochrome c oxidase (b). The initial potential is 272 mV vs NHE, and the final potential is 522 mV vs NHE. The buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2.

metric waves (this conformational transition is proposed to cause nonfaradaic current, which contributes to the observed voltammetric waves). Figure 5 shows potential step experiments taken at an oxidase-modified electrode (a) and at a lipid bilayer membrane containing no cytochrome c oxidase (b). These data are consistent with a slow structural change of the oxidase-lipid membrane that results in a change in electrode capacitance or diffusion of the oxidase enzyme in a lipid multilayer membrane. The former possibility is consistent with the cyclic voltammetry experiments, which show that cytochrome c oxidase undergoes a change in structure and/or orientation within a few seconds after being reduced. Again, this slow change is believed to be in addition to the well-characterized change in conformation that occurs on the time scale of turnover.58-60 Slow electron-transfer kinetics may explain the smaller charge passed during the anodic cyclic voltammetric scans with increasing scan rate (i.e., ca. 41 µC at 10 mV/s, ca. 33 µC at 20 mV/s, and ca. 9 µC at 30 mV/s) shown in Figure 3. Faster potential scan conditions may limit conformational changes as discussed above, thereby reducing the total charge passed. The positive shift in peak potential of the anodic wave with increasing scan rate (see Figure 3) is consistent with a kinetic barrier to electron transfer.61,62 The decrease in charge passed with increasing scan rate may also reflect diffusion of the oxidase enzyme within a lipid multilayer structure. More than a monolayer of cytochrome c oxidase present in a lipid multilayer membrane on the electrode could explain the amount of charge passed during the anodic voltammetric scans. Diffusion of the oxidase enzyme to the electrode during the anodic voltammetric scans would have the result of more enzyme reacting at the electrode during the slower scan rate experiments relative to the faster scan rate experiments. Perhaps the voltammetric response of the oxidase-modified electrode is the result of a combination of faradaic current from more than a monolayer of oxidase and nonfaradaic current coupled to electron transfer. Attempts were made to follow the membrane deposition process by QCM measurements. However, only a few experiments resulted in a stable QCM frequency response during dialysis. The magnitudes of (59) Copeland, R. A.; Smith, P. A.; Chan, S. I. Biochemistry 1987, 26, 7311. (60) Copeland, R. A.; Smith, P. A.; Chan, S. I. Biochemistry 1988, 27, 3552. (61) Laviron, E. J. Electroanal. Chem. 1979, 101, 19. (62) Nicholson, R. S. Anal. Chem. 1965, 37, 1351.

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the frequency shifts that were observed varied and corresponded to masses that ranged from that of ca. one lipid bilayer to that of ca. two lipid bilayer membranes. These measurements may be erroneous due to possible viscoelastic properties of the deposited lipid membranes that can influence the resonant frequency of the QCM crystals.63 The data have been interpreted on the basis of two models of the lipid bilayer membrane containing the oxidase enzyme: the oxidase immobilized in a single lipid bilayer on the electrode and the oxidase in a membrane consisting of several lipid bilayers on the electrode. The model involving the oxidase immobilized in a single lipid bilayer on the electrode is supported by the FIA experiments of reduced cytochrome c reacting at these oxidasemodified electrodes. As shown in Figure 5a, current flows several seconds after the anodic potential step experiments. Assuming this current is faradaic due to diffusion of the oxidase to the electrode through a thick lipid membrane, enzyme-mediated electron transfer from solution-resident cytochrome c to the electrode would require several seconds (i.e., the oxidase enzyme would have to be reduced at the membrane-solution interface by the hydrophilic protein (cytochrome c) and then diffuse to the electrode through the membrane). Most FIA experiments of reduced cytochrome c reacting at the oxidase-modified electrodes show no delay between the arrival of cytochrome c at the membrane-solution interface and onset of current flow (vide infra). Therefore, most of the oxidase lipid bilayer membranes must be sufficiently thin to account for this observation. QCM-FIA data for the exposure of cytochrome c to these oxidase-modified electrodes show that cytochrome c does not penetrate or bind to the lipid bilayer under the conditions of the amperometric FIA experiments (vide infra). Assuming a multilayer oxidase membrane, fast homogeneous electron transfer from oxidase enzyme to oxidase enzyme is not consistent with the potential step experiments, which show current several seconds after the potential step. The scan rate dependence of the cathodic wave observed in the voltammetry conducted at the oxidase-modified electrodes shows that a structural change of the oxidase and/or of the lipid membrane occurs following oxidation of the enzyme. The model involving the immobilization of the oxidase in a single lipid bilayer on the electrode and nonfaradaic current coupled to the structural changes of the membrane that occur following oxidase turnover is assumed. A typical cyclic voltammogram taken at an oxidasemodified electrode under 0.1 mM sodium cyanide is shown in Figure 6b. Figure 6a is a cyclic voltammogram taken at the oxidase-modified electrode before the addition of cyanide. The cathodic wave observed in the cyclic voltammogram taken at the oxidase-modified electrode in the presence of solution-resident cyanide may be due to faster reduction relative to the apparent reduction rate of the cyanide free oxidase enzyme on the electrode. The oxidized state of heme a3 is stabilized in cyanide-bound oxidase,31,64,65 and Cub has been titrated in cyanide-bound oxidase.64,65 The binding of cyanide to the heme a3-Cub site may be involved in the changes in wave shape that are observed in the oxidase voltammetry under solutionresident cyanide (see Figure 6). However, cyanide causes a ca. 300% increase in electrode capacitance perhaps due to incorporation of the cyanide into the lipid membrane (compare Figure 6a,b at 130 mV of the positive scan). (63) Buttry, D. A.; Ward, M. D. Chem. Rev. 1992, 92, 1355. (64) Nicholls, P.; Chanady, G. A. Biochem. J. 1982, 203, 541. (65) Goodman, G. J. Biol. Chem. 1984, 259, 15094.

Cytochrome c Oxidase in Lipid Bilayer Membranes

Figure 6. Cyclic voltammograms taken at an oxidase-modified electrode under phosphate buffer (a) and under phosphate buffer containing 0.1 mM sodium cyanide (b). The scan rate is 20 mV/s. The buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2.

This increase in capacitance is observed at control membranes containing no oxidase enzyme (data not shown), and the observed changes in wave shape of the oxidase voltammetry in the presence of cyanide may be coupled to the structure of the lipid membrane in the presence of solution-resident cyanide. Cyanide is in the form HCN at the pH used (7.4). The Ka for HCN66 is 6.2 × 10-10. The anodic peak potentials observed in the cyclic voltammograms, the square wave voltammograms (data not shown), and the differential pulse voltammograms (data not shown) are all near the potentials reported by Cullison et al.3 and by Salamon et al.2 for cytochrome c oxidase immobilized in supported lipid bilayer membranes and these studies are consistent with titration measurements.29-32 The voltammetry reported here is also consistent with a voltammetric study of cytochrome c oxidase immobilized on self-assemble monolayers of 3-mercaptopropionic acid on gold.67 If the oxidase enzyme is unidirectionally immobilized in a single supported lipid bilayer with its Cua redox site (i.e., the cytochrome c binding site) held away from the electrode surface, then the faradaic component of the oxidative wave observed in the cyclic voltammetry experiments results due to intramolecular electron transfer from the Cua redox site to heme a or heme a3-Cub (i.e., the oxygen binding site) and from heterogeneous electron transfer from one of these sites to the electrode. Based on the crystal structure,28 heme a, heme a3, and Cub may be approximately the same distance from the electrode (ca. 35 Å) and, though the membrane topography may be different on the electrode, it is not appropriate to assign the voltammetric peak potentials to a specific redox center. Salamon et al.68 have shown that electron transfer between cytochrome c and cytochrome c peroxidase occurs across a conductive lipid bilayer membrane. This work demonstrates that complex formation is not required for electron transfer. Fast electron transfer occurs between Cua and heme a3 (2 × 105 s-1)24 at a distance of 19 Å,28 and electron-transfer distances up to ca. 30 Å have been reported in zinc/ ruthenium-modified myoglobins.69 Also, an electron(66) Analytical Chemistry An Introduction, 6th ed.; Skoog, D. A., West, D. M., Holler, F. J., Eds.; Harcourt Brace College Publishers: Philadelphia, 1994; p A-4. (67) Li, J.; Cheng, G.; Dong, S. J. Electroanal. Chem. 1996, 416, 97. (68) Salamon, Z.; Vitello, L. B.; Erman, J. E.; Butko, P.; Tien, H. T. J. Electroanal. Chem. 1989, 275, 213. (69) Axup, A. W.; Albin, M.; Mayo, S. L.; Crutchley, R. J.; Gray, H. B. J. Am. Chem. Soc. 1988, 110, 435.

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transfer distance of ca. 30 Å through ferrocene-terminated self-assembled monolayers has been reported.70 In light of these measurements, a 35 Å separation between the electrode and the closest redox site (heme a or heme a3Cub) may be sufficiently close to allow heterogeneous electron transfer. The following sequence of electron transfer has been generally accepted.24 Electrons are first injected into the enzyme at the Cua redox site from cytochrome c. Electrons are then transferred from the Cua redox site to the heme a redox site. This chargetransfer process is followed by electron transfer from the heme a redox site to the binuclear heme a3-Cub redox site where molecular oxygen is reduced to water.24 Recently, a second path of electron transfer (cytochrome c f Cua f cytochrome a3-Cub) (i.e., heme a is bypassed) has been proposed to explain a dependence of the proton pump H+/ e- ratio on the rate of electron transfer through the oxidase enzyme.71 Electron transfer from Cua to heme a3-Cub has also been demonstrated by others.25 The direction of electron transfer upon oxidation of the enzyme in the supported lipid bilayer membrane may be in the same direction as the proposed paths for electron transfer occurring through cytochrome c oxidase in vivo.25 This possibility is supported by FIA experiments (vide infra). The possibility that silver chemistry is involved in the observed voltammetry conducted at the oxidase-modified electrodes under both phosphate buffer and phosphate buffer containing 0.1 M sodium cyanide has been ruled out on the basis of experiments conducted at gold QCM electrodes with no electrodeposited silver. Following the dialysis procedure discussed above, lipid bilayer membranes containing cytochrome c oxidase were formed on cleaned bare gold QCM electrodes. The voltammograms taken at these electrodes showed anodic waves comparable in size to the anodic waves that are observed at oxidasemodified electrodes formed on electrodeposited silver. In the presence of solution-resident cyanide, a cathodic wave was apparent in the cyclic voltammograms taken at the oxidase membranes formed on bare gold QCM electrodes. Thus, silver chemistry is not believed to contribute to the voltammetry described here and this proposal is consistent with the control experiments shown in Figure 2a. The reproducibility and stability of the voltammograms taken at the oxidase membranes formed on bare gold electrodes is not good, as reported earlier.3 The stability of this oxidase membrane allows amperometric FIA of cytochrome c through enzyme-mediated electron transfer from the solution-resident protein to the metal electrode. Figure 7 shows three sequential aerobic FIA experiments for reduced cytochrome c reacting at an oxidase-modified electrode. The peak heights observed upon injection of the three plugs of cytochrome c are nearly the same, and this reproducibility is typical of most of the oxidase-modified electrodes. Under decreasing ionic strength (i.e., 0.02 M phosphate buffer), the peak currents were only slightly less (e.g., 5%) than those observed under 0.1 M phosphate buffer, indicating that the current responses do not reflect large capacitive changes (i.e., nonfaradaic current). The reproducibility of the current peak heights from oxidase-modified electrode to oxidasemodified electrode is ca. 30%. Figure 8 shows aerobic FIA experiments in which three different reduced cytochrome c concentrations are detected at the same oxidase-modified electrode. Figure 8d is a control experiment showing the current measured at a lipid bilayer modified electrode (70) Chidsey, C. E. D.; Bertozzi, C. R.; Putuinski, T. M.; Mujsce, A. M. J. Am. Chem. Soc. 1990, 112, 4301. (71) Capitanio, N.; Capitanio, G.; Demarinis, D. A.; De Nitto, E.; Massari, S.; Papa, S. Biochemistry 1996, 35, 10800.

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Figure 7. FIA data of cytochrome c reacting at a cytochrome c oxidase-modified electrode. Three sequential 83 µL injections of 100 µM reduced cytochrome c. The electrode potential is 472 mV vs NHE; the buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2.

Figure 8. FIA data of different cytochrome c concentrations reacting at a cytochrome c oxidase-modified electrode (a-c). Response (d) is FIA of cytochrome c reacting at a lipid bilayer modified electrode containing no cytochrome c oxidase. The concentrations of cytochrome c injected are 150 µM (a), 75 µM (b), 37.5 µM (c), and 75 µM (d). The electrode potential is 472 mV vs NHE; the buffer is 0.1 M phosphate, pH 7.4, and the electrode area is 0.2 cm2.

containing no oxidase upon injection of reduced cytochrome c using the same electrode at which the cyclic voltammetric scan shown in Figure 2a was taken. The response of 75 µM cytochrome c at the lipid bilayer modified electrode (Figure 8d) is 2.5% of the 75 µM cytochrome c response at the oxidase-modified electrode (Figure 8b). The current observed during control experiments (lipid bilayer modified electrodes containing no oxidase) probably results from electron transfer at defects in the membrane. The possibility that the oxidase-modified electrodes are defect rich and the current observed upon injection of reduced cytochrome c is not enzyme mediated is not consistent with the comparable capacitance exhibited by the lipid bilayer membrane containing no oxidase and the oxidasemodified electrodes (see Figure 2 near 130 mV of the positive scan). The comparable capacitance of the two electrodes used for the experiments shown in Figure 8 suggest that the lipid bilayer membrane containing no oxidase enzyme and the oxidase-modified electrode have nearly the same number of defects. Figure 8 compares the 75 µM cytochrome c FIA current response at the lipid bilayer modified electrode containing no oxidase enzyme (Figure 8d) with one of the smallest 75 µM cytochrome c FIA current responses observed at an oxidase-modified electrode (Fgiure 8b). Under anaerobic conditions, reduced cytochrome c FIA experiments conducted at the oxidase-modified electrodes

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Figure 9. QCM-FIA data of oxidized cytochrome c at a lipid bilayer modified electrode containing no cytochrome c oxidase: (a) 10 µM cytochrome c in 0.1 M phosphate buffer; (b) 2.5 µM cytochrome c in 0.01 M phosphate buffer; (c) 5 µM cytochrome c in 0.01 M phosphate buffer; (d) 10 µM cytochrome c in 0.01 M phosphate buffer. The phosphate buffer is pH 7.4, and the electrode area is 0.2 cm2. The experiments were conducted under open circuit.

show kinetic transitions involving an increase in the rate of oxidase mediated electron transfer during turnover (i.e., the current responses are biphasic). Lowering the cytochrome c concentration that is reacting with the oxidase results in kinetic transitions involving a decrease in the rate of oxidase mediated electron transfer.72 These data are consistent with the literature describing both activation of the resting oxidase state to the pulsed oxidase state during turnover and decay of the pulsed oxidase state to the resting oxidase state.25,73 The time required for the injected cytochrome c sample to reach the electrode (16 s) was determined by injection of potassium ferricyanide at a clean gold QCM electrode. There was no detectable delay of the current response at most of the oxidase-modified electrodes upon injection of reduced cytochrome c. This result is consistent with cytochrome c oxidase being immobilized in a single lipid bilayer on the electrode with its cytochrome c binding site exposed to solution. The immediate rise times of the current responses observed at the oxidase-modified electrodes following injections of reduced cytochrome c were inconsistent (e.g., see Figure 8 at 12 s) and some electrodes (20%) exhibit a delay of ca. 2-3 s upon injection of cytochrome c. This rise time variation may be due to differing amounts of oxygen in the membrane from injection to injection. However, reordering of the oxidase in the lipid membrane upon exposure to cytochrome c cannot be ruled out nor can the possibility that lipid multilayers exist on some of the modified electrodes. QCM-FIA experiments were conducted to probe the electrostatic association between cytochrome c and both lipid bilayer modified electrodes containing no oxidase enzyme and oxidase-modified electrodes. Under 0.1 M phosphate buffer no binding of 100 µM cytochrome c was detected at either the lipid bilayer modified electrodes containing no cytochrome c oxidase or the oxidase-modified electrodes. However, under reduced ionic strength conditions (e.g., 0.01 M phosphate buffer) cytochrome c was found to bind to both the lipid bilayer modified electrodes containing no oxidase and the oxidase-modified electrodes. Figure 9 shows a set of QCM-FIA experiments conducted at a supported lipid bilayer membrane containing no oxidase. Figure 9a shows the QCM frequency upon (72) Burgess, J. D.; Rhoten, M. C.; Hawkridge, F. M. J. Am. Chem. Soc., in press. (73) Antonini, E.; Brunori, M.; Colosimo, A.; Greenwood, C.; Wilson, M. T. Proc. Natl. Acad. Sci. USA 1977, 74, 3128.

Cytochrome c Oxidase in Lipid Bilayer Membranes

injection of 10 µM cytochrome c under 0.1 M phosphate buffer. No binding of cytochrome c is detected. Figure 9d shows the QCM frequency shift observed upon injection of 10 µM cytochrome c under 0.01 M phosphate buffer. The decrease in QCM frequency corresponds to an increase in mass. The reproducibility from injection to injection at the same electrode is ca. 2-3 Hz. However, the reproducibility from electrode to electrode is not good. About half of the lipid bilayer modified electrodes containing no oxidase showed frequency shifts comparable in magnitude to those shown in Figure 9 only when much higher cytochrome c concentrations (e.g., 150 µM) were injected. The oxidase-modified electrodes showed QCM frequency shifts comparable in magnitude to the frequency shifts shown in Figure 9. The phosphate buffer concentration dependence of the binding of cytochrome c to the supported lipid membranes is consistent with electrostatic association of the protein and lipids. However, hydrophobic forces may also be involved.74 Cytochrome c was stripped from the modified electrodes upon flushing the cell with 0.01 M phosphate buffer for ca. 30 min. These QCM-FIA data suggest that no cytochrome c is incorporated into the lipid membranes of either the lipid bilayer modified electrodes containing no oxidase or the oxidasemodified electrodes under 0.1 M phosphate buffer (the FIA experiments probing the oxidation of reduced cytochrome c at the modified electrodes were conducted under 0.1 M phosphate buffer). The data shown in Figure 9 were collected at open circuit and there was no detectable potential dependence at 472 mV vs NHE (the FIA experiments probing the oxidation of reduced cytochrome c at the modified electrodes were conducted at 472 mV vs NHE). There was no detectable difference in the binding of reduced and oxidized cytochrome c to the modified electrodes as measured by QCM-FIA. From comparison of the currents from cytochrome c FIA data obtained at the oxidase-modified electrodes and from cytochrome c FIA data acquired at gold QCM electrodes modified with bis(4-pyridyl) disulfide, a wellknown promoter,75 the active area of the oxidase-modified (74) Salamon, Z.; Tollin, G. Biophys. J. 1996, 71, 848. (75) Taniguchi, I.; Toyosawa, K.; Yamaguchi, H.; Yasukouchi, K. J. Electroanal. Chem. 1982, 140, 187.

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electrodes is estimated to be about one-half of the geometric area (i.e., the cytochrome c current peak height observed at the oxidase-modified electrode was in general ca. onehalf of the cytochrome c current peak height observed at the promoter-modified gold electrode). This estimation (50% oxidase coverage on the electrode) is indeed qualitative due to the reproducibility of the two measurements. The possibility that cytochrome c undergoes slower electron transfer at the oxidase-modified electrodes than it does at the promoter-modified electrodes cannot be ruled out. Conclusions Stable lipid bilayer membranes and lipid bilayer membranes containing cytochrome c oxidase were immobilized on gold electrodes using an OM submonolayer on electrodeposited silver. The OM submonolayer becomes part of the bilayer membrane as it forms under dialysis conditions and it anchors the bilayer membrane to the electrode. Results from cyclic voltammetry, differential pulse voltammetry, and square wave voltammetry experiments indicate that the oxidase enzyme undergoes direct electron transfer with the gold electrode. The voltammetric and FIA (amperometric and QCM) experiments are consistent with cytochrome c oxidase being unidirectionally immobilized on the electrode in a single lipid bilayer membrane with its cytochrome c binding site accessible to solution-resident cytochrome c. Experiments probing the dynamics of oxidase-mediated electron transfer are underway. This enzyme immobilization scheme contributes to the field of biosensor development and should be applicable to the study of other transmembrane enzymatic systems. Acknowledgment. Dr. Bertha King, Dr. Zoia Nikolaeva, and Professor Mikhail Smirnov are gratefully acknowledged for isolation of cytochrome c oxidase. Professor Isao Taniguchi graciously provided the AFM measurements as described herein. The assistance of Professor Stanley Bruckenstein in constructing the QCM circuit is much appreciated. We also acknowledge the National Science Foundation (Grant NSF CHE-9508640) for support of this research. LA9711995