Cytochrome c and Cytochrome c Oxidase: Monolayer Assemblies

J. Phys. Chem. B 2001, 105, 11351-11362

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Cytochrome c and Cytochrome c Oxidase: Monolayer Assemblies and Catalysis Alan S. Haas,† Denis L. Pilloud,† Konda S. Reddy,† Gerald T. Babcock,‡ Christopher C. Moser,† J. Kent Blasie,§ and P. Leslie Dutton*,† The Johnson Research Foundation, Department of Biochemistry and Biophysics, UniVersity of PennsylVania, Philadelphia, PennsylVania 19104, Department of Chemistry, Michigan State UniVersity, East Lansing, Michigan, and Department of Chemistry, UniVersity of PennsylVania, Philadelphia, PennsylVania 19104 ReceiVed: May 14, 2001; In Final Form: August 7, 2001

A novel electrochemical system has been designed and assembled to study the kinetic activity of cytochrome c oxidase. Gold electrodes coated with 3-mercapto-1-propanol formed the surface for the physisorption of monolayers of cytochrome c and cytochrome c oxidase or a preformed cytochrome c-cytochrome c oxidase complex. The films were investigated by cyclic voltammetry at scanning at rates slow enough to permit near redox equilibrium between electrode and redox protein and hence obtain redox midpoint potentials. Cytochrome c monolayers alone displayed a reversible midpoint potential at pH 8 (Em8 vs NHE) at +240 mV, close to the native cytochrome c value observed in solution. In contrast, oxidase monolayers alone failed to support any detectable redox contact between electrode and protein, implying that the distances between the oxidase redox cofactors in the adsorbed oxidase are too far away from the electrode to promote significant electron transfer rates. However, adsorption of a preformed cytochrome c-cytochrome c oxidase complex promoted effective redox contact, demonstrating electron transfer with an apparent onset halfpoint potential at +225 mV. This effect is consistent with the mandatory requirement for cytochrome c to mediate electrons from the electrode to cytochrome c oxidase and presumably in a way reflecting the physiological pathway. Cyclic voltammetric measurements arranged to determine the rates of electron transfer between electrode and the complex showed that at scan rates up to 50 mV/s, extraordinary kinetic turnover is displayed attributable to the catalysis of oxygen reduction. Thus it is established that the protein complex can be assembled and enable the natural mediation of electron transfer from the electrode by cytochrome c to the enzyme at a rate fast enough for catalysis to be observed and controlled.

Introduction Cyclic voltammetry (CV) applied to proteins adsorbed as films on electrodes is offering new analytical opportunities to dissect both equilibrium and kinetic properties of intraprotein electron transfer and coupled chemistry in the large oxidoreductase family of enzymes.3 CV applied to immobilized protein films is a natural way to observe vectorial electronic and charged chemical events common in the mechanisms of redox enzymes. The importance of CV has been further heightened by recent demonstrations of natural substrate oxidation-reduction catalysis by several structurally complex oxidoreductases while adsorbed to electron accepting/donating electrodes.22 These include glucose oxidase,2 cytochrome c peroxidase,31 succinate dehydrogenase,26,45 and fumarate reductase.46 The CV signature of enzymatic activity is the “catalytic wave” seen as a significant and continuous stream of current as the supporting electrode is scanned over a potential range characteristic of the oxidation and reduction of the enzyme cofactors and substrate.22 Thus, the presence of substrates in the solution promotes multiple turnovers and a steady state transfer of electrons through the * Corresponding author address: 1005 Stellar-Chance Labs, Department of Biochemistry and Biophysics, University of Pennsylvania, Philadelphia, PA 19104. E-mail: [email protected]. Fax: 215-573-2235. † Department of Biochemistry and Biophysics, University of Pennsylvania. ‡ Michigan State University. § Department of Chemistry, University of Pennsylvania.

electrode-associated enzyme to and from the electrode. But the method is particularly informative in rapid scan CV studies when it can reveal critical quantitative information on the time-scales of multistep electron transfer within the enzyme, and on the nature of its coupling to protons, substrate/product exchange, and to other chemical events. Redox protein films can be created using Langmuir-Blodgett methods and deposited on transparent substrates that allow characterization of the spectroelectrochemical properties of the redox centers and monitoring of catalytic electron transfer reactions. However, the routine creation of redox protein films on electrodes and the application of CV to their analysis require a sufficiently strong binding interaction between electrode and protein to create a stable film without denaturation that often occurs at electrode surfaces. In principle, this danger can be overcome through coating of electrodes with monolayers of selected functionalized spacers. An appropriately charged or polar molecular film that overlays the electrode surface can conserve the native state and promote order in the protein films.42 For redox proteins, a good indicator that a film has been successfully constructed without denaturation is the retention in the film of the native redox electrochemistry that is commonly found in solution. Thus for hemes that have high in vivo potentials, such as cyt c, denaturation can lead to ligation change6 or increased exposure of the heme to water and cause a dramatic drop in redox potential. Thus, cyt c adsorbed on a carboxylated surface retains the native redox potential value,47

10.1021/jp011834m CCC: $20.00 © 2001 American Chemical Society Published on Web 10/16/2001

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Figure 1. Designs and challenges associated with constructing cyt c and CcO monolayer films on electrodes. (A) Cyt c with heme (black bar) oriented for electrical contact with the electrode surface, in this case coated with an organic acid terminated surface monolayer. Hydroxyl-terminated surfaces can provide a similar orientation. (B) CcO adsorbed directly on electrode but out of electrical contact. (C) Cyt c in electrical contact with organic acid terminated electrode surface, but heme faces electrode and is out of electrical contact with CcO. (D) CcO and cyt c complexes in electrical contact with electrode through matrix side. (E) Cyt c partially denatured on thiol terminated electrode surface. (F) CcO and cyt c complex in electrical contact with hydroxyl terminated electrode surface.

whereas on other surfaces such as bare gold the potential is hundreds of millivolts lower.41 The success of rapid scan CV rests critically on minimizing the distance between the electrode and the closest redox cofactor bound in the protein.37 Small redox proteins such as cytochrome c, azurin or ferredoxin may naturally lend themselves to establish good communication with electrodes25,24,44,20,10,49 because their redox cofactors are typically located near the surface for rapid electron transfer with other redox proteins. By directing the surface of optimal electron entry/egress on the small protein toward the electrode by a specific interaction, very rapid electron transfer can be achieved. Cyt c, arranged as in Figure 1A, provides a well-established example.4,44 When no linker is used, as with pyrolytic graphite edge electrodes,25,24 or when gold electrodes are used in combination with substances such as conducting polyions,27 then communication may be fast but the orientation unknown. However, for the larger oxidoreductase proteins including transmembrane proteins such as cytochrome c oxidase (CcO), redox cofactors are often buried well within the interior. In these cases, establishment of close contact and rapid electron transfer between redox cofactor and electrode becomes far less certain.9,13,23,35 Depending on the binding and orientation, the surface topologies around physiological electron entry/egress areas may produce a gap between protein and the

electrode or place all redox cofactors at such distances to render electron transfer rates too slow for analytical purposes (Figure 1B). We have approached this problem by employing adsorbed natural, smaller native redox protein partners to form a specific and physiological connection for electron transfer between the cofactors in large oxidoreductases and supporting electrode in order to explore how well a multicomponent system using only unmodified natural proteins can communicate with the electrode. The potential challenges of the method are illustrated in Figure 1C where cyt c is shown bound in the native state with its heme cleft dominated by positive lysines that are strongly attracted to the negative carboxylates on the surface. This configuration mimics the natural attraction of this cleft region for the carboxylates at the cytochrome binding sites on photosynthetic reaction centers or cyt bc1 complex or cytochrome c oxidase.29 While this orientation promotes rapid electron transfer between cyt c with the electrode, it is an orientation expected neither to favor binding and self-assembly of CcO onto such a preexisting cyt c monolayer nor to create the desired electron transfer link to the electrode. In this assembly, even the closest approach between the cyt c heme edge and CuA, is about 30 Å, far too long to promote significant electron transfer rates. An alternative arrangement inverts the direction of the cyt c (Figure 1F). Construction involves either the stepwise self-

Electrochemical Study of Cyt c Oxidase Kinetics assembly of a monolayer film of cyt c bound to a coated electrode surface at the side remote from the heme, leaving the heme cleft exposed for ready adsorption of CcO, or direct onestep assembly of a preformed cyt c-CcO complex on the electrode surface. This physiological orientation of the cyt c in the cyt c-CcO assembly resembles monolayers of CcO and covalently tethered yeast cyt c-CcO complex films fabricated on silanized quartz surfaces whose structure was probed by X-ray scattering.1,17,16 This structural orientation offers a marked improvement over that shown in Figure 1C, because the intervening heme of the cyt c is more centrally disposed in the long span from the electrode to CuA and considerably accelerates the rate between cyt c and CuA, which is ∼104 s-1 under physiological conditions. The rate of transfer from the electrode to the heme through the reverse side of the cyt c need not be prohibitive. For example, electron transfer along a similar path from a flavin tethered at the native Cys 102 on the reverse side of yeast cyt c to the heme has been measured at a rate of 103 s-1.51 With a short linker between electrode and protein, electron tunneling over ∼22 Å from electrode, through linker and protein to the heme should be in the 10-100 s-1 range. Although this rate may be slower than optimal catalytic rates (kcat ∼103 s-1), our goal with these studies is to demonstrate that a rational, step-by-step construction of a physiologically related electron transfer link between the cytochrome oxidase and the electrode can be achieved. Materials and Methods Protein Preparations. Bovine cytochrome c oxidase was prepared according to the method of Yoshikawa,56 with slight modifications. After preparation in 100 mM sodium phosphate, pH 7.4 buffer, the enzyme was dialyzed to remove excess detergent, n-dodecyl-β-D-maltoside detergent (Calbiochem, La Jolla, CA), used in the isolation. Aliquots of 30 µL volume of a 200 µM enzyme solution were stored in liquid nitrogen until use. Crystalline cytochrome c from Saccharomyces cereVisiae was purchased from Sigma (St. Louis, MO) and stored at -20 °C. A 10 mM stock solution of ferro cyt c was prepared by reduction with dithiothreitol (Kodak, Rochester, NY) followed by purification on a G-25 column (Pharmacia Biotech, Piscataway, NJ) in order to minimize dimer-forming disulfide bonds and to remove excess salts and peptide fragments. The cyt c-CcO complex was formed in solution by the addition of a several-fold excess of cyt c to a 20 µM solution of CcO in 5 mM sodium phosphate buffer, pH 7.4, containing 0.03% n-dodecyl-β-D-maltoside detergent (Calbiochem, La Jolla, CA). Excess cyt c and buffer were removed by passing through a Sephadex G-50 bead (Sigma, St. Louis, MO) column equilibrated with 1 mM sodium phosphate, pH 8.0, and 0.03% n-dodecyl-β-D-maltoside. A 1:1 cyt c-CcO ratio was verified by optical spectroscopy after these procedures.30 Preparation of Langmuir-Blodgett Films. Glass microscope slides (VWR Scientific, San Francisco, CA) were cut into 12 by 25 mm strips and used as substrates for film deposition. They were cleaned by sonication for 30 min in concentrated RBS 35 detergent (Pierce, Rockford, IL) and by soaking for 2 h in Nochromix sulfuric acid (Aldrich, Milwaukee, WI). They were then rinsed thoroughly with deionized water, dried with an argon stream, and used immediately for deposition. The subphase of the Langmuir-Blodgett trough (Lauda Filmbalance FW2, Sybron/Brinkman, Westbury, NY) was deionized water containing 1 mM CdCl2 (Aldrich) and 1 mM HEPES, pH 7.4 buffer (Aldrich). All trough work was done at 10 °C under an argon atmosphere. The enzyme was spread onto

J. Phys. Chem. B, Vol. 105, No. 45, 2001 11353 the surface of the trough via the glass rod method. The 1 ul drops were added at 5 s intervals to half the area of trough until the surface pressure reached 5-8 mN/m. The trough was then fully expanded and more material was added to the same pressure again. The trough was allowed to equilibrate for 30 min, with minor losses in surface pressure, and then slowly compressed to an operating pressure between 21 and 25 mN/m for deposition. The glass substrates were then inserted rapidly into the subphase of the trough to a depth of 20 mm and withdrawn slowly at a rate of 10 mm/minute by a mechanical arm. The samples were dried with a gentle stream of argon as they were removed. They were stored at 4 °C under an argon atmosphere and used within 48 h. Optical Spectroscopy and Redox Titrations of LangmuirBlodgett Films. All spectra and redox titrations were performed in 50 mM MOPS buffer, pH 7.4 (Aldrich). Reduced-state spectra were obtained by adding 1 mM sodium dithionite (Aldrich) to degassed buffer and by flowing this over the sample; carbon monoxide spectra were obtained by bubbling this same solution with pure CO for several minutes. Optical spectra were taken at a 45° angle of incidence with a Johnson Foundation dual beam spectrophotometer in a thin layer cell. The cell contains positions for the coated slide and a blank glass sample so that absorbance from redox mediators and sodium dithionite could be subtracted. Redox mediators for titrations were 30 µM 2,3,5,6-tetramethyl-p-phenylenediamine (DAD, Aldrich), phenazine ethosulfate (PES, Sigma), phenazine methosulfate (PMS, Sigma), and 2-hydroxy-1,4-naphthoquinone (OHNQ, Sigma). All redox titrations were performed in the oxidative direction, starting from the fully reduced state. Sodium dithionite was the reductant and potassium ferricyanide was used as the oxidant. The potential of the redox solution was measured and controlled in an Erlenmeyer flask with a platinum and calomel electrode. The solution was pumped through the cell and over the slides with a peristaltic pump. A one minute scan of the cytochrome oxidase slide and the reference slide was taken at every potential for the redox titration. The references were subtracted from the monolayer spectra to obtain final spectra. The values at 443 and 460 nm were determined by averaging the data within a 2 nm wide region of these points. Electrode Preparation. Glass microscope slides (Sigma, St. Louis, MO) were cut into 8 × 25 mm pieces. They were first cleaned by sonication for 30 min in FL-70 detergent (Fisher Scientific, Springfield, NJ) and by rinsing with Millipore water (18 MΩ‚cm). The substrates were then soaked for 2 h in a sulfuric acid (Aldrich, Milwaukee, WI) and hydrogen peroxide (Fisher Scientific, Springfield, NJ) solution (4:1). After thoroughly rinsing with water again and drying with an argon stream, the glass substrates were used immediately for the preparation of the electrode by metal vapor deposition. Metal deposition was performed in a Denton DV-502A highvacuum evaporator (Denton, Cherry Hill, NJ). All depositions were performed at < 2 × 10-6 Torr. An adhesion layer of 150 Å of Ti was evaporated from titanium wire (0.127 mm diameter, Aldrich, St. Louis, MO) heated in a tungsten wire basket (Polysciences Inc., Warrington, PA). Immediately thereafter, without opening the chamber, a 1000 Å thick layer of gold was deposited from another tungsten basket. Self-Assembly of Substituted Alkanethiol and Protein Films. After gold vapor deposition, the gold electrodes were quickly submerged in a 10 mM solution of 3-mercapto-1propanol (Aldrich, Milwaukee, WI) in 2-propanol (HPLC grade, J. T. Baker, Phillipsburg, NJ). The alkanethiol self-assembly

11354 J. Phys. Chem. B, Vol. 105, No. 45, 2001 was allowed to proceed for 2-3 h, typical for SAM construction with short chain alkanethiols. Afterward, the modified electrodes were thoroughly rinsed with 2-propanol and dried with an argon stream. The fresh alkanethiol SAMs were then immediately immersed into protein solutions for incubation. Cyclic Voltammetry. Cyclic voltammetry (CV) was performed on a BAS 100B electrochemical workstation (Bioanalytical Systems Inc., W. Lafayette, IN). The reference electrode was Ag/AgCl in 3M KCl and the counter electrode was a platinum foil. CV was performed at room temperature in 12 mL of 1 mM sodium phosphate, pH 8.0, buffer that was degassed for 1 h with oxygen scrubbed argon prior to use and maintained under argon throughout. The area of the working electrode was between 0.7 and 0.8 cm2. The solution resistance, typically in the 0.5-1.5 KΩ range, was measured automatically by the BAS software in a potential region without faradaic currents. This fixed resistance value was entered into the software to prevent small instabilities that can come with dynamic compensation of the resistance. After insertion of protein films into the cell, degassing with argon was continued for an additional 30 min. The degassing period was extended to 1 h before using carbon monoxide. All films were stable for up to at least 2 h. CV scanning of the protein films between +500 mV and -100 mV was not deleterious to the films, but scanning below -100 mV led to some removal of the protein from the electrode. For near-redox equilibrium measurements, CV scan rates were done at 200 mV/s. Three sweeps were typically taken to establish reversibility and reproducibility. The current voltage relationship of the first scan was usually slightly different from subsequent ones on all protein coated electrodes, manifest as a higher current below the Em of cyt c and attributable we believe to some rearrangement of the protein film or irreversible reduction of residual oxygen by the porphyrin. Thus it was usual to use the second scan for analysis. However, for analyses done at 1 mV/s, only a single scan was taken because of the length of time required. Data Analysis. CV data were analyzed using BAS software. All potentials were converted relative to the normal hydrogen electrode (NHE) by adding +196 mV to values measured with the Ag/AgCl electrode at 25 °C. Protein midpoint potentials were determined by drawing a baseline under reductive and oxidative sweep waves and locating peaks; the average of the two peak potentials was taken to be the Em. Coverages were quantified by integration of the area under reductive and oxidative waves. The electron transfer rates between the electrode and cyt c were calculated taking into account the electronic distribution about the electrode Fermi levels and using Marcus theory. This method is described in detail in the work of Tender et al.48 and was performed using a simple computer program as in similar situations.38 Results Depostion of Langmuir-Blodgett Films of CcO. Although CV of monolayer films on electrodes permits great freedom to probe electron transfer reactions, spectroscopy of these monolayers is difficult. On the other hand, Langmuir-Blodgett films on transparent substrates have long been used to create electron transfer active monolayers and multilayers that permit spectroscopic verification of native properties and activity. We describe deposition of Langmuir-Blodgett films of CcO onto unmodified hydrophilic glass surfaces and characterize single monolayers by optical spectroscopy and with redox titrations, testing them for catalytic activity using reduced cyt c. We show that CcO can maintain native properties in the film state.

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Figure 2. Langmuir-Blodgett compression isotherm for CcO.

CcO formed a stable film at the air-water interface. After spreading onto a Langmuir-Blodgett trough, the monolayers were stable for hours with minor losses in surface pressure attributable primarily to the loss of the solubilizing detergent in the form of micelles into the subphase. In the LangmuirBlodgett compression isotherm for CcO shown in Figure 2, a liquid-like phase is exhibited at higher surface areas and then a transition to a solid-like phase takes place when slowly compressed. The creation of the solid-like state is interpreted as the formation of a densely packed monolayer on the airwater surface. Based upon the quantity of biological material added to the trough, we concluded that the transition to this state occurs at an area of about 5000 Å2/molecule. This value is near to the expected area of a CcO monomer (∼60 by 80 Å) as determined from a crystal structure.15,50 After deposition onto glass substrates, the surface area of the trough had decreased by approximately the same area as the surface of the substrates, indicating that a single monolayer of material was transferred to the glass. Optical Characterization of Langmuir-Blodgett Films of CcO. Optical spectra for CcO in solution in the Soret region are shown in Figure 3A. The two hemes contribute an equal absorption (50/50) in this region.55 The hemes of the oxidized enzyme display a peak at 421 nm and in the reduced form with a higher extinction coefficient at 443 nm with a ratio that depends on the quality of the preparation. In principle, using second-derivative spectroscopy each of these bands can be resolved into two slightly differing components to reveal the individual hemes.12 Carbon monoxide binds to the reduced enzyme on heme a3, shifting its absorption maximum to 430 nm, as can be observed in Figure 3A. For the weaker alpha band at 605 nm (not shown), there is an 80% contribution from heme a and 20% from heme a3. The Soret spectra of a single Langmuir-Blodgett monolayer of CcO (both sides of a slide) are shown in Figure 3B for comparison with solution spectra. It displays distinct oxidized, reduced, and reduced plus carbon monoxide bound forms of the enzyme. It was clear that the enzyme retained optical properties qualitatively similar to the protein in solution. However, the peak wavelengths and amplitudes of the film spectra differed slightly from solution values. These reproducible peak measurements are summarized in Table 1. The apparent reduced Soret peak at 440 nm in the monolayer films was nearly equivalent in magnitude to the oxidized peak at 419 nm,

Electrochemical Study of Cyt c Oxidase Kinetics

J. Phys. Chem. B, Vol. 105, No. 45, 2001 11355 TABLE 1: Optical Soret Peak Wavelengths and Redox Midpoint Potentials for CcO in Solution vs LB Monolayer

Figure 3. CcO optical spectra in the Soret region for solution (A) vs monolayer (B). Oxidized as prepared (thin line), reduced with sodium dithionite (thick line), and reduced plus carbon monoxide (medium line).

differing slightly from solution studies where the value at 443 nm of the reduced spectrum is typically expected to be 20% greater than the oxidized value at 421 nm. A smaller amplitude effect is observed as well in the carbon monoxide spectrum. These differences may in part be caused by a slope in the background due to light scattering off the film on the slide. In the case of the presence of carbon monoxide, it was evident that the CO bound to heme a3 because there was little change in the alpha region absorbance at 605 nm (not shown due to the low signal-to-noise ratio), of which 80% of the absorbance is attributed to heme a. As determined from the optical measurements, coverages of CcO on glass substrates approximated that of a densely packed monolayer. An absorbance at 440 nm of 0.9 (0.4 mOD/ monolayer yields an estimate of approximately 1.8-4.6 pmol/ cm2 using an extinction coefficient of 100 mM-1cm-1/heme.55 This is very near to the value of 2-3 pmol/cm2 expected for a densely packed monolayer of CcO as estimated from the dimensions of the crystal structure.21,50 Redox Titrations of Langmuir-Blodgett Films of CcO. Classic redox titrations of CcO solution fits to two equivalent magnitude n ) 1 Nernsts,14,53 with the potential of the a hemes of Em7.4 ) +210 mV and Em7.4 ) +370 mV (open circles, Figure 4A). These are the physiologically expected values for the two a hemes. Cyt c displays an Em7.4 at +260 mV (data not shown).

CcO

oxidized (nm)

reduced (nm)

reduced + CO (nm)

Em7.4 (heme a/a3) (mV)

solution monolayer

421 419

443 440

430 436

210/370 190/340 ((30)

For the titration of a single monolayer of CcO, the signalto-noise level was low because of the small quantity of biological material on the surface and also due to large interfering absorbances from redox mediators. Nevertheless, reliable redox titrations were plotted. A representative film titration is shown in Figure 4A in comparison with a solution titration. Redox titrations of the single monolayer consistently showed two native-like high potential components similar to the solution study. The average best fit potentials observed for a group of six different films were 190 mV for the lower a heme component (( 30 mV) and 340 mV for the higher a heme component (( 30 mV), as listed in Table 1. These are only slightly below the solution Em7.4 values measured at 210 mV and 370 mV. Infrequently, a markedly lower component between approximately 0 and 80 mV was measured, though this usually did not contribute to more than 30% of the entire optical change. The absolute absorption redox spectra for solution are shown in Figure 4B; the contribution from the redox mediators can be noticed as the slope in the background. For comparison, the redox difference spectra for the monolayer are displayed in Figure 4C. Redox titrations of multilayer films containing four monolayers showed similar behavior to the single monolayers, as displayed in Figure 5A. In these films, high components were consistently measured at potentials near 210 and 340 mV. The absolute redox absorption spectra are shown in Figure 5B. Activity of Langmuir-Blodgett Films of CcO. To test for enzymatic catalytic activity, the films were inserted into a solution of reduced cyt c in the presence of O2 at the partial pressure of air. Some oxidation of cyt c by the CcO films was observed. The initial expected rate of autoxidation of cyt c by dissolved dioxygen is followed by a further increase in the steady-state diminishment of the reduced alpha peak of cyt c (550 nm) after replacement of an uncoated glass substrate in the solution with one covered with a single monolayer of CcO. A rate of oxidation of 0.1-1 e- s-1/CcO was measured. Although this is much less than the ideal maximal rate of oxidase turnover (up to 1000 s-1), it did establish that oxidase can be immobilized on a film to retain native redox and spectral properties, with at least partial catalytic activity, as required for the next step of immobilization of oriented CcO on electrodes for CV. Attachment of Cyt c to Electrodes and Linkers. Our initial strategies to create the cyt c-CcO assembly on nontransparent electrode surfaces for CV studies involved attempts to covalently attach the reverse side of the cyt c to the electrode or spacer coating. We made use of yeast cyt c with its natural, conveniently positioned cysteine 102 located on the reverse side of the protein for covalent linkage to thiols or other reactive surfaces. It was considered that the yeast cyt c tethered in this way, with its heme cleft binding domain exposed, would set the stage for the second step in which CcO would specifically bind to the domain with high affinity and complete the construction of the physiological complex as shown. It has long been known that cyt c is rapidly denatured on contact with gold and exhibits dramatically lowered redox potentials, so no attempt was made to react the cys102-cyt c directly with the gold surface. Instead, gold electrodes were

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Figure 4. (A) Redox titration of Langmuir-Blodgett monolayer film of CcO (solid circles) vs in solution (open circles). The value at 445 nm minus the isosbestic point at 460 nm was plotted as a percent of the total change for a 5 µM CcO solution. The concentration of redox mediators (DAD, PES, PMS, OHNQ) for the titrations was 10 µM. (B) Representative absolute spectra in solution from 65 mV to 183, 252, 306, and 370 mV, respectively. (C) Redox difference spectra in monolayer for reduced minus 160, 228, 269, 341, and 358 mV.

treated with n-alkanedithiols to form a thiolate labeled monolayer coating38 to react with the cys-102 of cyt c. However, it was evident that the yeast cyt c underwent rapid adsorption to the thiolated surface, accompanied by massive modification of the electrochemistry. A typical CV analysis of the monolayer film is shown in Figure 6A. As listed in Table 2, the potential of the cytochrome dropped dramatically from Em8 ) +240 mV in solution32 to between -200 and -240 mV, a value expected for a heme with considerable exposure to water. Thus it seems likely that the cyt c undergoes considerable denaturation as depicted in Figure 1E, perhaps resulting from the relatively hydrophobic thiol terminated surface drawing the interior of the protein onto the surface disrupting ligation and exposing the heme. Further attempts at covalent attachment were continued using surfaces that were considered less likely to denature or deleteriously modify the cyt c. Surfaces comprising mixed mercaptopropionic acid with hexanedithiol were systematically tested. Protein incubations were performed in buffered solutions at high ionic strength (200-400 mM KCl) and pH 8.0 to minimize electrostatic binding to the heme cleft side and favor the desired covalent tethering. However, as shown in Table 2, when the

mercaptopropionic acid:hexanedithiol ratio on the surface was below an estimated 50:1, denaturation occurred. On the other hand, when the ratio was higher than 50:1, the cyt c bound and retained its native high redox potential, indicating that there were spaces on the surface of sufficient area to accommodate electrostatic interactions. Although this ratio of carboxylates to free thiols corresponds to approximately 1 thiol per 900 Å2, about the area of one cyt c, the protein proved to be readily removable by soaking in 1 M KCl solutions, showing that binding was only electrostatic and that a covalent linkage had not formed. Testing Other Polar Surfaces for Cyt c Binding Interactions. Other polar but non-carboxyl terminated surfaces tested for cyt c binding tended to avoid binding to the net positively charged (lysine) region around the heme cleft. We found that while binding was evident in low ionic strength buffers, denaturation still occurred with a positively charged electrode coating surface formed with cysteamine and also with the neutral mercaptopropanol coatings (Table 1). However, it was clear from the examination of the CV of cyt c on the hydroxyl surfaced linker that the adsorption of populations of low potential redox states was slow. This agrees with work showing

Electrochemical Study of Cyt c Oxidase Kinetics

Figure 5. Redox titration of 4-layer CcO film (A) and absolute redox spectra (B).

that hydrophilic surfaces tend to resist the adsorption and denaturing of proteins.39 However, and also in agreement with the previous studies with yeast cyt c on optically transparent thiolated and hydroxylated surfaces, binding rates are slow and coverage was found to be low.15,54 Effect of Detergent on Binding to Hydroxylated Surfaces. The addition of the CcO solubilizing detergent (n-dodecyl-βD-maltoside) to cyt c incubation solutions was found to improve considerably the formation of cyt c monolayer coverage while maintaining native-like potentials. As fully displayed in Figure 6B, the inclusion of detergent in incubating solutions clearly aided in the formation of a cyt c protein film with a high nativelike potential at +240 mV ((5 mV variance between different electrode preparations), a value matching the potential observed for native yeast cyt c in solution. Furthermore, it is also evident that scanning to potentials as low as -400 mV revealed little or none of the denatured component. Coverages for these new cyt c films was typically between 2 and 3 pmol/cm2. As listed and compared to the other surfaces in Table 2, this coverage is several times lower than measured on mercaptopropionic acid linkers and values ranging between 10 and 17 pmol/cm2 for densely packed monolayers of cyt c on carboxylated surfaces reported in the literature.44 Therefore, while a relatively low submonolayer coverage was achieved, the natural electrochemistry was retained without relying on negatively charged surfaces that would obligatorily force binding in the undesired orientation. Electron Transfer Rates to Cyt c. The electron transfer rate to the cyt c in films on hydroxylated surfaces was determined using the method of Chidsey11 for electron transfer between

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Figure 6. Cyclic voltammograms display dependency of redox potential for bound cyt c on surface and self-assembly conditions. (A) Denaturing occurs on a propanedithiol modified gold electrode after only a 90 min immersion in a 75 µM cyt c solution (10 mM HEPES, 10 mM KCl, pH 8.0) (solid line), and a reference modified electrode (dashed line). (B) Mercaptopropanol-modified electrodes display native potentials after 20 h assembly in 10 µM cyt c in the presence of detergent as detailed in Table 2 (solid line), and reference modified electrode (dashed line). Some noncatalyzed reduction of O2 is shown at low potentials. Scan rates were 200 mV/s.

and electrode and redox couples secured to the electrode by a covalent linker. In Figure 7, the overpotentials (E-E°) vs scan rate are plotted and fitted using these equations to reveal the ET rate to cyt c. The ket between the electrode and the cyt c heme was determined to be 20 s-1, a rate consistent with the relatively long distance between the electrode and heme when the heme cleft faces away from the electrode. This rate is orders of magnitude slower than that observed for cyt c bound to carboxy-terminated linkers of similar length, where presumably the heme cleft faces the electrode. This 20 s-1 rate is also only slightly faster than the CV derived rate observed for cyt c bound to carboxy-terminated linkers of much longer length (11mercapto-undecanoic acid, X. Chen, personal communication), where presumably the heme cleft also faces the electrode. Binding of CcO to Cyt c Films. Figure 8A compares the reversible Em8 of +240 mV of the cyt c monolayers with films after submergence in a CcO solution resulted in a shift in the cyt c midpoint potential. Some broadening of the reductive and oxidative waves and slight increase in the electroactive current is evident. These factors were an indication of the presence of CcO. The apparent overall 10-20 mV shift down in the cyt c midpoint potential relative to cyt c films without CcO is

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TABLE 2: Surfaces and Buffer Conditions Determine Cyt c Monolayer Redox Potential, Binding Mode, and Coverage alkanethiol(s) HS(CH2)3COOH HS(CH2)3-8SH HS(CH2)3COOH/(CH2)6SH 50:1 HS(CH2)3NH3 HS(CH2)3OH HS(CH2)3OH HS(CH2)3OH HS(CH2)3OH

buffera 10 mM HEPES, pH 8, 10 mM KCl 10 mM HEPES, pH 8, 1-200 mM KCl 10 mM HEPES, pH 8, 200-400 mM KCl 10 mM HEPES, pH 8, 200-400 mM KCl 10 mM HEPES, pH 7, 10 mM KCl 1 mM NaPO4-, pH 8, with up to 200 mM KCl 1 mM NaPO4-, pH 8, 0.03% dodecyl-maltoside 1 mM NaPO4-, pH 8, 0.03% dodecyl-maltoside + inc. in 10 µM CcO 1 mM NaPO4-, pH 8, 0.03% dodecyl-maltoside 10 µM cyt c-CcO complex

adsorption

coverageb

Em (mV)

electrostatic

+230

1

hydrophobic

-200 to -240, +50

1, negligible

hydrophobic

-220

up to 1

electrostatic

+225

0.1-0.4

hydrophobic

-130

1

hydrophobic or electrostatic/polar electrostatic/polar

-220, +20, +240

0.3, 0.1-0.2,