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An anaerobic spectroelectrochemical cell for studying the spectral and redox properties of flavoproteins
ANALYTICAL
BIOCHEMISTRY
An Anaerobic
109. 295-308
(1980)
Spectroelectrochemical and Redox Properties MARIAN
Cell for Studying of Flavoproteinsl
the Spectral
T. STANKOVICH
An improved spectroelectrochemical cell has been developed in which both coulometric and potentiometric titrations of flavoproteins can be performed. Simultaneous acquisition of spectral data in these titrations enables measurements of molar absorptivity, number of electrons transferred and redox potentials. This cell has an oxygen leak rate of0.02 nmolimin, a factor of 20 slower than that reported for the cell of Hawkridge and Kuwana (R. Szentrimay . R.. P. Yeh. and T. Kuwana. 1977. in Electrochemical Studies of Biological Systems. American Chemical Society Symposium No. 38 (Sawyer. D. T., ed.). pp. 142- 169, American Chemical Society, Washington. D. C.) in which similar measurements can be made. Oxygen leakage from the electrodes was minimized by pretreating the electrodes with dithionite and by including glucose and glucose oxidase in the auxiliary electrode. Oxygen in the atmosphere was scrubbed by the glucose-glucose oxidase system in a sidearm. The initial oxygen concentration is comparable to that of Hawkridge and Kuwana. The cell is simple in design. easy to construct, and fits into an unmodified spectrophotometer. The cell was tested on methyl viologen where n = I.00 was obtained, and on flavodoxin where n = 1.04 t- 0.02 and n = 1.03 ? 0.02 were obtained for the two electrons. The two electrons of glucose oxidase which are transferred within 2 mV of each other were transferred with 95-100s efficiency. Using the potentiometric titration and four mediator dyes, we were able to measure redox potentials of both electrons of flavodoxin in pH range from 5.3 to 9.3. Redox potentials for glucose oxidase were measured at pH 5.3. Values for both flavoproteins agreed with published data.
The most commonly used method for quantitatively reducing flavoproteins is anaerobic titration with standardized solutions of sodium dithionite t l-4). Data obtainable from dithionite titrations include molar absorptivity (E). number of electrons transferred (n ). and redox potential value (E”‘) (5). Dithionite is used because it has a relatively negative potential; the titration is rapid, requiring 30 to 60 min: and it is colorless in the visible wavelength region. However. dithionite has several disadvantages (6): (a) the redox potential of dithionite changes with pH, therefore, it may not be negative enough to quantitatively reduce all I This work was supported by a Cottrell Research Corporation Starter Grant and a University of Massachusetts Faculty Research Grant #2-03875.
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enzymes at all pH values (e.g., the flavoprotein dehydrogenases), (b) dithionite is unstable at low pH values, so it cannot be used in redox titrations below pH 6.5, (6) its oxidation product, bisulfite, forms complexes with flavoprotein oxidases (7,8) (d) dithionite is very reactive with oxygen, so it must be prepared and stored anaerobically. Because of these problems, the use of dithionite titrations is subject to severe restrictions. As a result, the spectroelectrochemical properties of these classesof enzymes have not been well characterized. An electrochemical reductive coulometric titration can provide the sameinformation as the dithionite titration, but in order to be a practical alternative. the electrochemical reduction 0003-2697180118029514$02.00/O Copyright All right,
C 1980 by Academic Press. Inc. of reproduction in any form rexwed.
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must be rapid, efficient, and oxygen leakage into the system must be minimized. Electrochemical methods have been applied to a wide range of enzymes participating in biological electron transfers (9), but have not been used extensively for flavoproteins. Of the methods reported, most measure changes in absorbance (AA ) or the number of coulombs (AQ) added as a function of the potential of the system (IO- 15). The potential of the system (E) may be a potentiostatically controlled value or a measured value of a system at equilibrium. These methods yield redox data directly for simple systems whereAA/A or AQiQ is a valid measure of the total change in concentration of one member of a redox couple. For more complex systems involving two or more electron transfers, where redox potentials are separated by less than 100 mV, computer programs are used to resolve A vs E data (E is applied or measured potential) (9,16). They have been applied to membranebound cytochromes in which sequential electron transfers are separated by a minimum of 30 mV having a deviation of 10 mV (16). However, the minimum potential difference between electron transfers which can be resolved by these programs is not stated (9,16). However, even if the computer programs are applicable for flavoproteins, the spectral information provided by the coulometric titration is still required. Since we are studying purified enzymes isolated in the oxidized state (17), both coulometric and potentiometric titrations can yield valuable complementary information (9). This twopronged approach is especially important for flavoproteins which can exhibit three spectrally distinct oxidation states (18). In the coulometric experiment, AA is measured as a function of AQ. One thing that coulometry can do that potentiometry cannot do is that with a purified system it can give quantitative information in the actual micromolar amount of redox groups present. As a result, the coulometric titration can directly
yield two pieces of information which could be obtained only indirectly from potentiometric titrations. (a) The molar absorptivity of all species, oxidized, one electron reduced. and two electron reduced is obtained. By definition, E is AA/AC, in the coulometric titration both AA and AC (concentration change) are measured directly. (b) The number of electrons (n ) transferred to the system is obtained directly. This value can also be obtained in potentiometry from the slope of the Nernst equation. However, in the case where an enzyme contains two independent one-electron transfer sites with similar absorbance and redox properties, the slope of the Nernst plots will yield a value of II = 1 rather than II = 2 (19). The actual micromolar amount of the redox groups present could be obtained from a potentiometric titration if the number of reducing equivalents used to reduce the mediators could be obtained independently and subtracted from the total number of reducing equivalents transferred to the enzyme-mediator system at any point in the titration ( 15). A second advantage of coulometry, at the present time, in the case involving flavoproteins, is the lack of adequate transparent mediators in the spectral absorbance range used for these enzymes. The coulometric experiments on membrane-bound enzyme or unpurified enzyme would yield accurate data only if the redox potentials of other electron acceptors differ significantly from the enzyme of interest. Initially the species of interest should be totally in the oxidized form. If these conditions could be met, the spectral data described above could be provided for membrine-bound enzymes. Hawkridge and Kuwana (20) have developed an anaerobic spectroelectrochemical cell in which coulometric, potentiometric, and absorbance measurements can all be made. The primary difficulty in designing cells of this type is the maintenance of an oxygen-free system over extended periods
SPECTROELECTROCHEMICAL
of time (1 to 3 h). Other considerations are complexity of construction and compatibility with conventional spectrophotometers. We have developed an electrochemical cell which has an oxygen leak rate 20fold slower than Hawkridge’s and Kuwana’s cell. Unlike their cell, ours can be used in a standard spectrophotometric cuvette holder, and can be easily constructed. This cell was tested with flavodoxin, glucose oxidase, and methyl viologen. all of which have well-characterized spectral properties and redox potentials. These test cases were chosen to examine the performance of our system under conditions which have proved difficult with the approaches discussed above, such as dithionite titration for the enzymes and the Hawkridge-Kuwana cell for methyl viologen. Difficulties with the dithionite titration have been set forth previously. When the Hawkridge-Kuwana cell was used in coulometric reductions of benzyl viologen, 8% errors were obtained; when it was used to reduce enzyme systems with more positive potentials 2-3s errors were obtained (21,22). Quantitative reduction of glucose oxidase is difficult for the following reasons. The redox potentials of the two electrons of glucose oxidase are separated by 2 mV (23). thus all three oxidation states are present in the reaction mixtures throughout the titration. The equilibration among these three species is slow. requiring 30 min (23), therefore. the cell must be extremely anaerobic to allow accurate determination of the E values. Last. glucose oxidase forms a bisulfite complex at low pH. Quantitative reduction of flavodoxin is difficult for a different reason. The redox potential for its second electron transfer is very negative (24). Therefore. dithionite cannot be used to determine this redox potential at most pH values (24). Methyl viologen has a redox potential even more negative than flavodoxin (21). Our cell yields accurate data for both of the flavo-
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proteins and for methyl viologen. As such, it represents a marked advance in the development of spectroelectrochemical cells. Other authors have developed cells which are perfectly adequate for the tasks for which they are designed. This particular cell design is optimized for parallel spectral and electrochemical studies of flavoproteins which may have negative redox potentials, potentials that are separated by only 2 mV, that may require long times to equilibrate and for which mixtures of optically transparent mediators do not exist. THEORY Coulonzetric titration. In coulometric experiments involving redox-active proteins. a mediator is necessary to transfer electrons from the electrode to the protein. This mediator is required to have: (a) the ability to rapidly transfer an electron to the reactive site of the protein, (b) the capability of being rapidly reduced at the electrode surface, and (c) a redox potential considerably more negative than that of the protein. This last requirement is in accord with Marcus theory (25,26), which predicts that the log of the rate of homogeneous electron transfer is proportional to the redox potential difference between the electron donor and electron acceptor. Viologen dyes are the best known electron transfer mediators for this purpose (21). Methyl viologen (MV)’ (E”’ = -0.450 V) was used in this work because it has the most negative redox potential of the commercially available dyes and therefore should transfer electrons to protein most rapidly. When the potential of the system is controlled at a value (-0.480 V) negative ’ Abbreviations used: MV. methyl viologen; IDS. indigo disulfonate: IDS,,.,,. reduced indigo disulfonate: BV. benzyl viologen: DHNQ. 1,4-dihydroxynaphthoquinone: IDS,,,. oxidized indigo disulfonate; R2G. Rosinduline 2G: FMN. flavine mononucleotide; SHE, standard hydrogen electrode; EFL,,, oxidized enzyme: EFIH”. one electron reduced enzyme: EFl,,,,,H-. two electron reduced enzymes.
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enough to reduce MV during a coulometric experiment, the transfer of electrons from the electrode to flavoproteins occurs in the following sequence of reactions: MVii
+ eme MV+”
Ill
MV+’ + EFl,,, + H’ + MV++ + EFlHO MV+O + EFlHO = MV++ + EFl,,,H-
[2a] [2b]
(EFI,, is oxidized flavoprotein, EFIH” is the one-electron-reduced flavoprotein, a radical: EFlrcdH- is the two-electronreduced flavoprotein). Reactions [2a] and [2b] represent the process by which MV++ , the electrochemically active species, is regenerated in the solution. Because of this cyclic mechanism of regeneration, a high concentration of MV++ is stabilized at the electrode surface and a constant, fast rate of electron transfer occurs (Eq. [I]) (21.27,28). Since the current is integrated, the number of coulombs (Q) transferred to the protein can be accurately determined. In the case of flavoproteins, spectra can be periodically recorded throughout the reduction; thus, absorbance can be obtained as a function of (!. The endpoint is determined by the appearance of completely reduced enzyme. Calculations made from these measurements yield the number of equivalents transferred to a mole of protein at a controlled potential and in agiven time interval. Potentiometric titration. The potentiometric titration gives optimal results when performed in the presence of two kinds of mediators. an electron transfer mediator, e.g., methyl viologen and redox mediator (9.20,29.30) or a mixture of three to five redox mediators (9.12.13). The redox mediators are chosen to have redox potentials very near that of the protein. When in equilibrium with the protein, the redox mediator is capable of poising the potential of the indicator electrode. As in the coulometric titration, electrons are transferred from the electrode to MV at very negative potential (Eq. [I]). The concepts and pro-
cedures involved are illustrated by the following description of the first electron transfer to a flavoprotein. Initially. electrons from MV+” are transferred to both the protein (Eq. [2a] and [2b]) and the redox mediator (Eq. [3]). As the concentration of the one-electron-reduced form of the protein (EFIH”) increases. it equilibrates with the redox mediator (e.g., indigo disulfonate, IDS) (Eq. [4]). 2MV+(’ + IDS,,, $ 2MV’ + + IDS,,,1 2EFlH” + IDS ox = 2EFLx + IDS,,,,
[3] [41
(IDS,,, is oxidized indigo disulfonate; IDS,,, is reduced indigo disulfonate.) At each point in the titration the system is allowed to equilibrate. Then both the potential (E) and the spectrum of the system are recorded. The concentration of EFl,,, and EFlH” are calculated from the spectra, corrected for the absorbance of the redox mediator (see Results. potentiometric titration) using published values of molar absorptivity. For flavodoxin. EFIH” has E-,,x,,= 4500. and e$15= 2 100 M ' cm-‘, and EFI,,, has l qq5= 10,200 Mm' cm-’ (17). Redox potential (,!?,‘I) is obtained from the Nernst equation (Eq. IS]) by plotting E versus log ((EFI,,,)/(EFlH”). 0.059 (EFlox) E = E;’ + log ~ n (EFlHO)
[51
(E is the measured potential; Ey’ is the redox potential: n is the number of electrons transferred in the reduction step.) MATERIALS
AND METHODS
Flavodoxin was prepared from Megcrsphera efsdenii (formerly Peptostrrptoc,oc.(‘us elsdenii) according to the method of Mayhew and Massey (17). Glucose oxidase (EC I. 1.3.4) was prepared using the method of Swoboda and Massey (31). Methyl viologen (MV), and benzyl viologen (BV), were purchased from British Drug House, Poole. England; 1.4-dihydroxynaphthoquinone (DHNQ) and Rosinduline 2G
SPECTROELECTROCHEMICAL
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(R2G) were purchased from K & K Labs (Division ICN Life Sciences, Plainview. N. Y.). Indigo disulfonate (IDS), purified, was kindly supplied by Dr. Fred Guengerich. Vanderbilt University. The buffers used were pH 5.25, 0.1 M sodium acetate: lpH 7.25, 0.1 M sodium phosphate; for pH 23 8.26, 0.044 M sodium pyrophosphate-HCI; 4 for pH 9.25-O. 1 M glycine. Deionized water, 18 Mohm resistance was used to make 5 solutions. Prepurified nitrogen was passed over Ridox scrubber (Fisher Scientific Co., Springfield. N. J.) for further oxygen removal. A potentiostat. Princeton Applied Research, Inc. (Princeton, N. J.) (PAR) 6 Model No. 173 was used to control the I+--7 potential: a PAR No. 179 coulometer was 6 used to integrate the current, and a PAR FIG. 1. Spectroelectrochemical cell; main body is No. 174A polarographic module was used constructed of quartz. (A) Front view. electrodes are to determine the potential of the dye Rosinin the plane of the paper. (1) Main connection of nitroduline 2G. A Varian Cary 219 UV-visible gen line; (2) sidearm; (3) gold foil working electrode: chloride reference electrode; (5) recording spectrophotometer was used (4) silver-silver silver-silver chloride auxiliary electrode: (6) standard to record spectra. quartz cuvette; (71 glass stirrer. (B) Side view. All potentials are reported versus the 3-ml cell is rotated 90” to give a clearer view of (I) and (2). standard hydrogen electrode (SHE). The working electrode was a gold foil 2.6 x 0.6 cm. The auxiliary electrode was a silver erence electrode was checked before and wire, contained in a fritted (frit is fine after each experiment against standard porosity) compartment which was plugged ferro-ferricyanide (32). The anaerobic with agar and was filled with KCI solution spectroelectrochemical cell was made from (Fig. I). The silver wire becomes coated a standard 3-ml quartz cuvette by adding with silver chloride during the course of sidearms with standard taper ground quartz electrochemical reduction reaction. Re- joints. The stirrer was a magnet sealed in action at the auxiliary electrode is glass or polyethylene. The auxiliary and reference electrodes Ag + Cl- - AgCl + (J-. were stored anaerobically in 5 mM diThis electrode is used to prevent the gen- thionite solution for 12 h prior to the eration of 0, at the auxiliary electrode, experiment. Shortly before the experiment a during electrochemical reduction. This degassed solution of 10 mM KCI, buffered electrode can be used for an oxidative to pH 5.25, was placed in the auxiliary titration; first the AgCl will be reduced electrode compartment (Fig. I). To this then H,O will be reduced to produce H, solution was added SO ~1 of 4 M glucose as at a platinum auxiliary electrode. and 10 ~1 of 1 mM glucose oxidase (oxygen The reference electrode was a silverscrubber). The compartment was sealed by silver chloride electrode. Electrical contact inserting the electrode, and was stored in was made to the solution through an dithionite until the experiment started. Only asbestos fiber: a small agar plug was used to the tips of the electrodes are immersed in minimize diffusion. The potential of the ref- dithionite. The absorbance of the solution
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FIG. 2. Controlled potential coulometric titration of flavodoxin. x lo-” M and 4.86 x IO-” MV in 4.42 ml of 0. I M sodium phosphate (2) n = 0.5 electronsimol FMN, (3) n = 1.0 electronimol FMN (5) n = I .89 electronimol FMN.
E = -0.480 V. Flavodoxin 3.52 pH 7.25. (1) Oxidized flavodoxin, (4) n = 1.49 electronsimol FMN.
was monitored before the degassing pro- pressure. At each point in the reduction, cedure and the addition of electrodes. After the spectrum was recorded within 2 min the addition of electrodes, the absorbance after the desired number of coulombs was transferred. No spectra1 changes occurred and the potential were monitored for 3045 min to insure that dithionite was not after 2 min time, this indicated that equilibleaking out, reducing the enzyme. rium was established within 2 min of each Coulometric experiment. The coulometric transfer of coulombs to the solution. Values experiments on flavodoxin and MV were for 11 and the molar absorptivity of the performed at room temperature at pH 7.25, flavodoxin radical were obtained indeas outlined in the Theory section. A solution pendently at 445 and 580 nm by extra(4.40 ml) containing tlavodoxin or MV, polating the linear parts of the coulometric mediator dye, and buffer was placed in the titration curves (Figs. 2 and 3). This will cell. Flavodoxin is isolated and stored as be discussed more fully in the Results the oxidized enzyme (17). There are no re- section. For methyl viologen, the efficiency ducing agents or oxidizing agents present of electrogeneration in terms of the II value to contaminate the enzyme in the final steps was determined from the slope of a plot of of the isolation procedure. Glucose and A versus Q, using known values of molar glucose oxidase in pH 5.3 buffer (same absorptivity and spectra1 path length (23). concentrations as above) were placed in the The reductive titrations were not corroborated with oxidative titrations. Ferrosidearm to scrub the 0, in the atmosphere. The system was closed and subjected to cyanide is the mediator titrant with positive I I cycles of evacuation and nitrogen potential most commonly used to carry out equilibration. The electrodes were then oxidations. Reportedly it complexes methyl placed in the cell under positive nitrogen viologen the reducing mediator titrant (9).
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P 445nm
'2.
d!--ieYo
115
n per mole FMN FIG. 3. Controlled potential coulometric vs amount ofcharge added to the system to residual 0, in the system. Absorbance
titration of flavodoxin. Plot of absorbance at 445 and 580 nm (n equals equivalents of charge). The nonzero intercept is due values have been corrected for methyl viologen absorbance.
Therefore, it should not be in the system. This should be checked in future experiments. Potentiornetric titrutions. The potentiometric titration was performed as outlined in the Theory section. Measurements were performed at a temperature of 25 +- 0S”C.
The process represented by Eq. [4] was allowed to come to equilibrium at open circuit. The equilibration time ranged from 5 to 15 min. The criterion used for equilibration was the stabilization of the potential reading. The potential was measured at the indicator electrode (the working elec-
302
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trode in the controlled potential mode) and the spectrum of the system was recorded. In control experiments the electron transfer mediators (MV and BV) and the redox mediators (IDS, DHNQ, and R2G) were all titrated; the concentration and pH values were the same as those used in the protein experiments. The distributions of the oxidized and reduced forms of the dyes were obtained as a function of potential. The total absorbance (contributed by both oxidized and reduced species of the dye) at each wavelength can be measured as a function of potential. Wavelengths of interest are usually 580 and 445 nm, where EFIH” and EFI,,, absorb maximally. These measurements were later used to correct for mediator contribution to the combined flavoprotein-mediator spectrum at each measured potential value. This method was used for IDS at pH values other than 7 where no convenient isosbestic point existed. At pH 5.3, the isosbestic point for IDS is at 416 nm. The difference in molar absorptivity between the EFIH” and EFl,,, at this wavelength is not great enough to allow independent spectrophotometric measurement of concentrations. However, the absorbance of IDS at 445 nm is small and directly proportional to the potential of the system. (The total L!A,,, = 0.04A for total reduction of the dye). The equilibrium potential and total absorbance at 445 nm are measured for the system containing enzyme and dye. The contribution of dye to the absorbance at 445 nm is subtracted and the concentrations of EFI,,, and EFIH” are calculated from the change in absorbance t 17). For IDS. at pH 7.0. a convenient isosbestic point exists at 460 nm. The change in absorbance at this wavelength was used to calculate the concentration of oxidized and reduced enzyme as described previously (20). DHNQ has a very small absorbance at 445 nm ttJJs = 2000 M-’ cm-‘): it has no absorbance at wavelengths longer than 550 nm. Therefore the concentration of EFIH” can be
directly measured from the spectrum at 580 nm t 17). Therefore when DHNQ was used as a mediator, no spectral correction was needed. The contribution of R2G to the glucose oxidase spectrum was subtracted as described previously (23). The viologens were used as both electron transfer and redox mediators for the second electron of flavodoxin. The reduced form of the viologens absorbs at long wavelengths where flavodoxin does not. From the absorbance at 725 nm, the entire MV”’ spectrum was calculated and subtracted from the combined MV-protein spectrum. Titrations of flavodoxin were performed at pH values 5.25, 7.25, 8.25, and 9.3. At pH 7.25 the E”’ for flavodoxin was measured using two dyes for each of the electron transfers to show that there was no binding of enzyme to mediator. Undetected binding of mediator to flavoprotein will cause appreciable error in redox measurements (33). The E"' for the first electron transfer was measured using the redox mediators DHNQ and IDS; both MV and BV were used to measure E”’ for the second electron transfer. Anaerobic precautions were the same as those described for the coulometric experiment. Glucose oxidase was titrated at pH 5.3. Anaerobic procedures were less rigorous than those described for flavodoxin and MV. The auxiliary and reference electrodes were not soaked in dithionite prior to the and no atmospheric oxygen experiment. scrubber was used. Glucose oxidase (1.6 x lop5 M) and R2G (4 x IO-” M) were coulometrically reduced at a potential of -0.155 V. The R2G (E"' = 0.080 V) acted as both electron transfer mediator and redox mediator. Because of the low concentration of mediator. electron transfer to the enzyme was slow (20 to 40 min) and an additional 20 to 40 min was required for equilibration (23). The efficiency of the reduction pro cedure was determined by comparing the coulombs of charge t Q) transferred to the system, with the coulombs of charge
SPECTROELECTROCHEMICAL
theoretically needed to produce the ured change in concentration of and EFI,,,,H. Concentrations of and EF&Hwere calculated from bance measurements and published absorptivities (23).
CELL
measEFIH” EFIH
absormolar
RESULTS Coulometric
Titmtiorl
The anaerobicity of the cell was tested by coulometrically titrating 2.5 x 10d5 M MV. A plot of Acio;: versus Q was linear with a nonzero intercept. The nonzero intercept corresponded to the amount of current consumed by oxygen initially in the system. On the average, 1.2 x lo-” M oxygen was initially present. The slope yielded the published value for the molar absorptivity of MV (12,400 M-’ cm-‘) with less than 1% error (assuming II = 1) (21). No background correction was required. When the absorbance of 50% reduced MV was monitored as a function of time. the oxygen leak rate was found to be 0.02 nmot per minute or 0.13 mC per hour. Figure 2 shows the spectral change which occurred upon coulometric reduction of 3.25 x IO-: M flavodoxin. The concentration of flavodoxin was calculated from the measured absorbance and the known molar absorptivity (17). The absorbance spectrum of reduced MV interfered with the flavodoxin spectrum only at values of II > 1.75 (n = equivalents of charge per FMN). The molar absorptivities of the species EFI,,,; EFIH”. and EFl,.,,,H- differ substantially at 580 and 445 nm. Figure 3 is a graph of A 5xoand A,,, as a function of the number of reducing equivalents added per flavin. Absorbance values and Q values were corrected for the contribution from MV. (Absorbance corrections were performed as described under Materials and MethodsPotentiometric titrations). From the absorbance values for MV the concentration of the reduced form was calculated. From the concentration and volume the number of
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coulombs of charge transferred to MV was calculated and subtracted from the total number of coulombs transferred to the MV -flavodoxin system. The nonzero intercept corresponded to the titration of residual oxygen in the system (0.5 x IO-” M). The value of ALx,, increased linearly with the number of equivalents added to the system up to 1~= I equivalent/FMN. Between II = 1 and II = 2 it decreased linearly. ‘The A,,, decreased linearly with the number of equivalents added. up to f~ = 1 equivalentsi FMN: from )I = 1 to II = 2 it still decreased. but the slope was much less steep. The A5H,I maximum occurred at the same value of II as did the inflection in the A,.,, plot. These data were not corrected for oxygen leakage. The calculated E;,~,)for EFIH” was in perfect agreement with the literature value (17). Table I summarizes the coulometric reduction experiments on flavodoxin. Our spectral data are identical to those obtained previously (17). In both cases a stable spectrally distinct one-electronreduced species is formed. Spectrum 3 of Fig. -3 is essentially the spectrum of the electron-reduced species (EFlH”I. The spectra of EFIH” and EFI,,, fully oxidized species differ most at wavelengths 580 nm where only EFIH” absorbs and at 445 nm where EFI,,, absorbs much more strongly (E = 10.200 M ’ cm-‘) than EFIH” (E = 2100 hr-’ cm- ‘). Therefore between II = 0 and II = I. there is an equilibrium between EFI,,, and EFIH”: this is retlected in the spectra (Fig. 2) and in the plots (Fig. 3). That is. A Sx,)increases from II = 0 to II = I. while A,,, decreases. This equilibrium is broken at II = I. From PI = 1 to 11 = 2, the equilibrium is between EFIH” and EFI,.,,,H,. EFI,.,,,,H, does not absorb at 580 nm but has molar absorptivity of E = 1600- 1800 at 445 nm ( 17). Therefore. A,,,, decreases tI = 1 to tz = 2 as does A,,,. From experiment I in Table I. it can be seen that 100% EFIH” was formed at tr = 1.08 equivalents of FMN. an 8% error: likewise the two-electron reduction was
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T. STANKOVICH TABLE
CONTROLLED
POTENTIAL
COULOMETRIC
I TITRATION or; FL4VoDoXIN.
pH 7.25
Experiments I Concentration enzyme (PM) Concentration MV (FM) Ratio dye/enzyme
35.2 48.6 1.4
2
3
53.8 25.0 0.46
55.3 25.0 0.45
4 47.1 100 2. I
First electron Average current (PA) Q calculated (n = 1) (mC) Q observed (mC) n measured n corrected Average corrected value
12.0 15.0 16.3 1.08 I .03 1.04 _f 0.02
6 22.8 25.6 1.12 1.06
6 23.6 26.7 1.13 1.06
25 20.0 21.2 1.06 I .02
Second electron Average current +A) Final current @A) Q observed (n = 2) (mC) n measured n corrected Average corrected value
5.5 1.5 15.5 1.03 0.98 1.03 -t 0.02
3.0 I.0 24.6 I .08 1.02
3.0 I.0 26.5 1.12 1.05
18.0 6.0 21.8 1.09 1.05
predicted observed Q initial 0, reduction (mC) Estimated 0, contribution to flavodoxin (%) Ass,, A,,,
0.158 0.158 0.6
0.242 0.241 0
0.249 0.246 0.2
0.212 0.212 1.6
5
6
6
3.5
complete at n = 2.12 an overall error of 6%. The average error in n was 1.10 & 0.02 for the first electron and 1.08 ? 0.02 for the second electron. The error for MV was less than 1%. This difference in percentage error between the flavodoxin and MV experiments can be ascribed to the timescale of the experiments (Table 3) and the difference in reactivity of the reduced species with oxygen. MV+O reacts with oxygen rapidly, therefore any oxygen initially present will be reduced before MV++ starts to reduce (34). EFIH” reacts more slowly with oxygen, therefore EFIH” and O2 can exist in solution simultaneously (35). It was found that for flavodoxin solutions an average of 0.85 mC was required to reduce the “initial oxygen” (Table 1) as opposed to the 2.5 mC for MV++. This initial difference of 2.5-0.85 mC, plus the leak rate
of 0.12 mC/h provided the value captioned “Estimated 0, contribution to flavodoxin” in Table 1. This value (approximately 5%) was used to correct the n value for the first and second electron transfers, resulting in 1.04 & 0.02 and 1.03 * 0.02, respectively.
Potentiornetric
Titrution
Figure 4 shows the results of a series of potentiometric titrations on flavodoxin. The individual values (Table 2) were obtained as described under Materials and Methods. Our data are in excellent agreement with those of Mayhew et crl. (24) as is shown by the solid lines which are the best fit to their redox data. The slope of the potential versus pH curve for the first electron transfer is 60 mV/pH unit. Ey’ values for the first electron at pH values
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DISCUSSION
0I\ .
-0.4
CELL
6
7
PH
8
9
10
We have developed a spectroelectrochemical cell in which potentiometric, coulometric, and spectroscopic experiments on oxygen-sensitive samples may be conveniently performed. The primary advantage of our system lies in the degree of anaerobiosis which can be maintained. The oxygen leak rate for our cell is 20-fold lower than that of Hawkridge and Kuwana (21. 22) (see Table 3). Data for the 0, leak rate are not published for other systems in Table 3. Thus we are able to successfully engage in studies under conditions where oxygen interference can cause the most problems, i.e., at potentials more negative than -0.400 V and for systems that require several hours to reach equilibrium.
FIG. 4. Effect of pH on the redox potential of flavodoxin. (0) Experimentally determined values for E”’I and E;‘. (0) experimental point for Ey’. previously determined value. The solid lines in the figure are the best fits from the data of Mayhew rf al.
(24).
5.25, 8.26, and 9.25 have not been directly measured before. E;’ for the second electron above pH 7.0 was constant at -0.373 V. Figure 5 shows the individual Nernst plots. The potentiometric titration experiments with glucose oxidase show that the agreement between charge transfer and spectral data depended on the time required for the experiment. In an experiment where electron transfer was relatively rapid and only two experimental points were measured (29 and 47% reduced), agreement was 100%. In a second experiment where seven points were measured, agreement was 95% for the first four points (2.5 h); 77% for the last three points (4 h). The redox potential data for both electrons of glucose oxidase at pH 5.3 are given in Table 2. The redox data obtained from three experiments were again in excellent agreement with values obtained by conventional methods (23).
‘lredl
FIG. 5. Representative Nemst plots for the twoelectron transfers of Ravodoxin at different pH values. The y axis on the right is used for curves (a) and (b), (shown in dashed lines). (a) First electron. pH 5.3: (b) first electron pH 7.25. The y axis on the left is used for curves (c) through(f). (c) First electron, pH 8.26; (d) first electron, pH 9.3: (e) second electron. pH 5.25: (f) composite curve for second electron, pH values 7.25-9.25. pH 7.25 (0): pH 8.25 (A); pH 9.25 (a,.
306
MARIAN
T. STANKOVICH TABLE
REDOX
-0.024 -Il.(lXh
2
POTENTIAI.
t 0 WI + KIM?
5 4
DA-I A
1).33ll
0 iIll?
I) 327 + 0 ,Y)? 7.??
I). 130 -0 II4 -0.144 0.127 -0 IX
+ + f f +
U.uu? 0 uul U.uuI u (XI? O.WI
7 K 7 6 Y
196 1 U.UU3
>
DHNQ
h 4
BV BV
X.26
-0
Y 25
Cl.??3 f O.(XK15 -Il.??7 - O.(KI
NOW2 IDS DHNQ DHNQ IDS
0 I I I z U.WI -0 171 + O.wu~ -
-
To demonstrate the capacity of our cell, we have determined the redox and spectral properties of three previously investigated systems. The molar absorptivity for methyl viologen was determined coulometrically to better than 1% precision. This despite the fact that reduced methyl viologen is extremely oxygen reactive. with an E”’ of -0.450 v. The redox and spectral properties for both electron transfers of flavodoxin were determined with greater than 95% precision, at several pH values. These results include redox potentials for the first electron transfer at pH 5.25. 8.26. and 9.25 which have not been directly measured before. Each experiment required 2-3 h to complete, thus demonstrating the performance of our cell over periods of prolonged anaerobiosis while in the presence of an oxygen-reactive
6 >
,,
-
0.371 -0.373
f u IWII + (1 ,“,I -
? 7 5
nv
-,I 3’iu
HV MV -.
Il.350 ll.44~ -~
UV BV “V
-
HV MV DHNQ MV MV
IMV
0 440
MV
HV MV
U.350 0 440
HV HV MV
compound of notably negative redox potential (-0.373 V). Finally. we measured the redox potential for both electron transfers of glucose oxidase at pH 5.3. Glucose oxidase is an enzyme for which the redox potentials foreach of the two electron transfers are almost identical (23). Yet we were able to accurately determine both values, even though the total experiment required 6.5 h to complete. These three tests clearly demonstrate the effectiveness of our technique. Table 3 affords a comparison of the capabilities of our system with the three other comparable systems currently in the literature. The importance of performing the coulometric titration and thus measuring the spectral properties of flavoproteins was pointed out in the introduction. By performing two experiments our cell enables us to
SPECTROELECTROCHEMICAL
CELL TABLE
C~MPARISION
OF ELECTROCHEMICAL
FOR
ENZYME
3 CELL
CHARACTERISTICS
I,, ml” x-30
x-30 -
-
307
REDUCTION
Ill,” ml,”
IT h XI
30
0 75
0.9x + 0 05 I Of3 T KO!
4 hNo. of rlectron\ for each compound
-
70- l4lJ
I hl
A, 1.
” Exprssxd m term\ of 10~s of abaorbancc due tu reduced viulugen ” II = equivalenfr. Q = coulombs: E = potential: .4 - ;lhcorbance.
measure accurately II, E, E”’ for purified flavoproteins as shown in Table 3. As shown in Table 3. the Hawkridge-Kuwana cell is designed to perform all three measurements (20-22). Hawkridge’s and Kuwana’s cell has an error of 2 to 3% when measuring E or II values for the cytochrome systems for which it was designed (these have positive potential value). However, for 1 mM benzyl viologen alone there is an 8% error in determining II or E. In our present cell, methyl viologen was reduced with less than I% error. However, because of the longer times scales of the experiment, coulometric titration of flavodoxin has a 3 to 4% error. The Hawkridge-Kuwana cell would be expected to yield less accurate data for flavodoxin due to: (a) its higher error with BV, (b) its higher 0, leak rate. Watt’s ( 15) method was designed primarily for the study of purified enzymes with negative redox potentials with no spectral properties. It yields n and I?’ values with accuracy, but no spectral data can be obtained using this system (Table 3). The system of Hendler et ~(1. (12) was designed to yield absorbance data as a function of potential
for membrane-bound cytochrome systems and in mixtures of optically transparent mediator dyes. The presence of other redox active species in this membrane system makes accurate coulometric titrations in this system very difficult. The potential of their system is computer controlled and the spectral data are computer analyzed. Their experimental design yields accurate data for their complex system. However, it is too complex and inaccessible to other workers: simpler approaches yield more data and equivalent data for purified enzymes. In addition, even if this computer-controlled system were available, coulometric titration is important for flavoenzymes because optically transparent dyes are not available. Therefore, as these other experimental designs and cells were optimized for data collection for particular systems, our system is optimized for measuring spectral (E) and electrochemical data (n, E”‘) for flavoproteins. However, the lower leak rate of our cell would be an advantage in any of the other systems. We are further improving our system by increasing the electrode area. the redox mediator concentra-
308
MARIAN
T. STANKOVICH
tion. and the stirring rate in order to accelerate charge transfer and decrease equilibration time. CONCLUSION
In conclusion, all four methods outlined in Table 3 can provide redox potential data on enzymes for which e values are known or for those enzymes where successive electron transfers are sufficiently separated. Only our cell and that of Hawkridge and Kuwana were designed to enable complete characterization of purified enzyme. Of these latter two, our cell clearly excels at maintaining anaerobiosis. In addition, it is simple to construct and maintain, and it will fit into a standard spectrophotometer. The accuracy of our system has clearly been demonstrated by reproducing values of E, E"', and II for the well-characterized proteins, glucose oxidase and flavodoxin, and also for methyl viologen. ACKNOWLEDGMENTS My thanks to V. Massey. A. Bard, L. Schopfer for helpful suggestions.
E. Moore,
and
REFERENCES 1. Foust. G. P., Burleigh, B. D., Jr., Mayhew. S. G.. Williams. C. H., Jr., and Massey. V. (1969) And. Biochem. 27, 530-535. 2. Burleigh. B. D.. Foust, G. P.. and Williams, C. H.. Jr. (1969) And. Biochrm. 27, 536-544. 3. Lambeth, D. 0.. and Palmer, G. (1973) J. Bid. Chum. 248, 6095-6103. 4. Dutton, L. (1975) in Methods in Enzymology (Fleischer, S., and Packer, L.. eds.), Vol. 54, Part E. pp. 411-435. Academic Press, New York. 5. Moore, E. G.. Ghisla. S.. and Massey, V. (1979) J. Bid. Chem. 254, 8173-8178. 6. Mayhew. S. G. (1978) Eur. J. Biochem. 85, 535-547. 7. Massey. V., Muller. F., Feldberg, R.. Schuman, M., Sullivan, P. A., Howell, L. G., Mayhew. S. G.. Matthews, R. G., and Faust. G. P. (1969) J. Bid. Chrm. 244, 3999-4006. 8. Muller. F., and Massey. V. (1969) J. Bid. Chrm. 244, 4007-4016. 9. Wilson, G. S. in Methods in Enzymology (1978) (Fleischer, S., and Packer, L.. eds.), Vol. 54, pp. 396-410, Academic Press, New York.
10. Barman, B. G., and Tollin, G. (1972) Biochemi.ctr~ 25, 4755-4759. 11. Swartz. D. B., and Wilson. G. S. (1971) Anal. Biochem. 40, 392-400. 12. Hendler, R. W.. Songco. D.. and Clem. T. R. (1977) And. Chrm. 49, 1908- 1913. 13. Hendler. R. W. ( 1977)Anal. Chrm. 49, l914- 1918. 14. Hendler. R. W.. and Shrager, R. I. (1979) J. Bid. Chrm. 254, 11288- 11299. 15. Watt, G. D. (1979) And. Biochrm. 99, 399-407. 16. Hendler. R. W. Towne, D. W.. and Shrager, R. I. (197.5) B&him. Biophys. Acta 376, 42-62. 17. Mayhew. S. G.. and Massey. V. (1969) J. Bid. Chrm. 244, 794-802. 18. Singer, T. P. and Edmondson. D. E. (1978) in Methods in Enzymology (Fleischer. S., and Packer. L.. eds.). Vol. 53, Part D. pp. 397418, Academic Press, New York. 19. Iyanagi, T.. Makino. N., and Mason, H. S. (1974) Bkxhemisrry 13, 1701- 1710. 20. Hawkridge. F. M.. and Kuwana. T. (1973) And. Chef?. 45, IO?]- 1027. 21. Szentrimay, R.. Yeh, P., and Kuwana, T. (1977) in Electrochemical Studies of Biological Systems. ACS Symposium No. 38, (Sawyer, D. T.. ed.). pp. 142-169. American Chem. Sot.. Washington. D. C. 22. Heineman, W. R.. Kuwana, T., and Hartzell. C. R. ( 1972) Biochum. Biophys. Rrs. Commun. 49, 1-8. 23. Stankovich. M. T.. Schopfer. L. M.. and Massey, V. (1978) J. Bid. Chrm. 253, 4971-4979. 24. Mayhew. S. G., Foust. G. P., and Massey. V. (1969) J. Bid. Chem. 244, 803-810. 25. Marcus. R. A. (1965) J. Chem. Phys. 43, 679701. 26. Marcus, R. A. (1964) Annu. Rev. Phys. Chem. 15, 155- 196. 27. Kuwana. T., and Winograd, N. (1974) in Electroanalytical Chemistry (Bard, A. J.. ed.), Vol. 7, pp. l-78, Dekker. New York. 28. Bard, A. J.. and Santhanam. K. S. V. (1970) in Electroanalytical Chemistry, (Bard. A. J., ed.). Vol. 4, pp. 215-315. Dekker. New York. 29. Moreno. C. G., Choy, M.. and Edmondson. D. (1979) J. Bid. Chem. 254, 7630-7635. 30. Anderson, J. L.. Kuwana. T.. and Hartzell. C. t 1976) Biochemistry 15, 3847-3855. 31: Swoboda. B. P., and Massey. V. (1965) J. Bid. Chrm. 240, 2209-2215. 32. O’Reilly, J. E. (1973) Biochim. Biophys. Acfu 292, 509-515. 33. Stombaugh. N. A., Sundquist, J. E., Burris. R. H., and Orme-Johnson, W. H. (1976) Biochemisfo 15, 2633-2641. 34. Small. R. D., and Scaiano. J. C. ( 1977) J. Phys. Chrm. 81, 2126-2131. 35. Mayhew. S. G. (1971) Biochim. Biophys. Acln 235, 282- 302.
BIOCHEMISTRY
An Anaerobic
109. 295-308
(1980)
Spectroelectrochemical and Redox Properties MARIAN
Cell for Studying of Flavoproteinsl
the Spectral
T. STANKOVICH
An improved spectroelectrochemical cell has been developed in which both coulometric and potentiometric titrations of flavoproteins can be performed. Simultaneous acquisition of spectral data in these titrations enables measurements of molar absorptivity, number of electrons transferred and redox potentials. This cell has an oxygen leak rate of0.02 nmolimin, a factor of 20 slower than that reported for the cell of Hawkridge and Kuwana (R. Szentrimay . R.. P. Yeh. and T. Kuwana. 1977. in Electrochemical Studies of Biological Systems. American Chemical Society Symposium No. 38 (Sawyer. D. T., ed.). pp. 142- 169, American Chemical Society, Washington. D. C.) in which similar measurements can be made. Oxygen leakage from the electrodes was minimized by pretreating the electrodes with dithionite and by including glucose and glucose oxidase in the auxiliary electrode. Oxygen in the atmosphere was scrubbed by the glucose-glucose oxidase system in a sidearm. The initial oxygen concentration is comparable to that of Hawkridge and Kuwana. The cell is simple in design. easy to construct, and fits into an unmodified spectrophotometer. The cell was tested on methyl viologen where n = I.00 was obtained, and on flavodoxin where n = 1.04 t- 0.02 and n = 1.03 ? 0.02 were obtained for the two electrons. The two electrons of glucose oxidase which are transferred within 2 mV of each other were transferred with 95-100s efficiency. Using the potentiometric titration and four mediator dyes, we were able to measure redox potentials of both electrons of flavodoxin in pH range from 5.3 to 9.3. Redox potentials for glucose oxidase were measured at pH 5.3. Values for both flavoproteins agreed with published data.
The most commonly used method for quantitatively reducing flavoproteins is anaerobic titration with standardized solutions of sodium dithionite t l-4). Data obtainable from dithionite titrations include molar absorptivity (E). number of electrons transferred (n ). and redox potential value (E”‘) (5). Dithionite is used because it has a relatively negative potential; the titration is rapid, requiring 30 to 60 min: and it is colorless in the visible wavelength region. However. dithionite has several disadvantages (6): (a) the redox potential of dithionite changes with pH, therefore, it may not be negative enough to quantitatively reduce all I This work was supported by a Cottrell Research Corporation Starter Grant and a University of Massachusetts Faculty Research Grant #2-03875.
295
enzymes at all pH values (e.g., the flavoprotein dehydrogenases), (b) dithionite is unstable at low pH values, so it cannot be used in redox titrations below pH 6.5, (6) its oxidation product, bisulfite, forms complexes with flavoprotein oxidases (7,8) (d) dithionite is very reactive with oxygen, so it must be prepared and stored anaerobically. Because of these problems, the use of dithionite titrations is subject to severe restrictions. As a result, the spectroelectrochemical properties of these classesof enzymes have not been well characterized. An electrochemical reductive coulometric titration can provide the sameinformation as the dithionite titration, but in order to be a practical alternative. the electrochemical reduction 0003-2697180118029514$02.00/O Copyright All right,
C 1980 by Academic Press. Inc. of reproduction in any form rexwed.
296
MARIAN
T. STANKOVICH
must be rapid, efficient, and oxygen leakage into the system must be minimized. Electrochemical methods have been applied to a wide range of enzymes participating in biological electron transfers (9), but have not been used extensively for flavoproteins. Of the methods reported, most measure changes in absorbance (AA ) or the number of coulombs (AQ) added as a function of the potential of the system (IO- 15). The potential of the system (E) may be a potentiostatically controlled value or a measured value of a system at equilibrium. These methods yield redox data directly for simple systems whereAA/A or AQiQ is a valid measure of the total change in concentration of one member of a redox couple. For more complex systems involving two or more electron transfers, where redox potentials are separated by less than 100 mV, computer programs are used to resolve A vs E data (E is applied or measured potential) (9,16). They have been applied to membranebound cytochromes in which sequential electron transfers are separated by a minimum of 30 mV having a deviation of 10 mV (16). However, the minimum potential difference between electron transfers which can be resolved by these programs is not stated (9,16). However, even if the computer programs are applicable for flavoproteins, the spectral information provided by the coulometric titration is still required. Since we are studying purified enzymes isolated in the oxidized state (17), both coulometric and potentiometric titrations can yield valuable complementary information (9). This twopronged approach is especially important for flavoproteins which can exhibit three spectrally distinct oxidation states (18). In the coulometric experiment, AA is measured as a function of AQ. One thing that coulometry can do that potentiometry cannot do is that with a purified system it can give quantitative information in the actual micromolar amount of redox groups present. As a result, the coulometric titration can directly
yield two pieces of information which could be obtained only indirectly from potentiometric titrations. (a) The molar absorptivity of all species, oxidized, one electron reduced. and two electron reduced is obtained. By definition, E is AA/AC, in the coulometric titration both AA and AC (concentration change) are measured directly. (b) The number of electrons (n ) transferred to the system is obtained directly. This value can also be obtained in potentiometry from the slope of the Nernst equation. However, in the case where an enzyme contains two independent one-electron transfer sites with similar absorbance and redox properties, the slope of the Nernst plots will yield a value of II = 1 rather than II = 2 (19). The actual micromolar amount of the redox groups present could be obtained from a potentiometric titration if the number of reducing equivalents used to reduce the mediators could be obtained independently and subtracted from the total number of reducing equivalents transferred to the enzyme-mediator system at any point in the titration ( 15). A second advantage of coulometry, at the present time, in the case involving flavoproteins, is the lack of adequate transparent mediators in the spectral absorbance range used for these enzymes. The coulometric experiments on membrane-bound enzyme or unpurified enzyme would yield accurate data only if the redox potentials of other electron acceptors differ significantly from the enzyme of interest. Initially the species of interest should be totally in the oxidized form. If these conditions could be met, the spectral data described above could be provided for membrine-bound enzymes. Hawkridge and Kuwana (20) have developed an anaerobic spectroelectrochemical cell in which coulometric, potentiometric, and absorbance measurements can all be made. The primary difficulty in designing cells of this type is the maintenance of an oxygen-free system over extended periods
SPECTROELECTROCHEMICAL
of time (1 to 3 h). Other considerations are complexity of construction and compatibility with conventional spectrophotometers. We have developed an electrochemical cell which has an oxygen leak rate 20fold slower than Hawkridge’s and Kuwana’s cell. Unlike their cell, ours can be used in a standard spectrophotometric cuvette holder, and can be easily constructed. This cell was tested with flavodoxin, glucose oxidase, and methyl viologen. all of which have well-characterized spectral properties and redox potentials. These test cases were chosen to examine the performance of our system under conditions which have proved difficult with the approaches discussed above, such as dithionite titration for the enzymes and the Hawkridge-Kuwana cell for methyl viologen. Difficulties with the dithionite titration have been set forth previously. When the Hawkridge-Kuwana cell was used in coulometric reductions of benzyl viologen, 8% errors were obtained; when it was used to reduce enzyme systems with more positive potentials 2-3s errors were obtained (21,22). Quantitative reduction of glucose oxidase is difficult for the following reasons. The redox potentials of the two electrons of glucose oxidase are separated by 2 mV (23). thus all three oxidation states are present in the reaction mixtures throughout the titration. The equilibration among these three species is slow. requiring 30 min (23), therefore. the cell must be extremely anaerobic to allow accurate determination of the E values. Last. glucose oxidase forms a bisulfite complex at low pH. Quantitative reduction of flavodoxin is difficult for a different reason. The redox potential for its second electron transfer is very negative (24). Therefore. dithionite cannot be used to determine this redox potential at most pH values (24). Methyl viologen has a redox potential even more negative than flavodoxin (21). Our cell yields accurate data for both of the flavo-
CELL
FOR
ENZYME
REDUCTION
397
proteins and for methyl viologen. As such, it represents a marked advance in the development of spectroelectrochemical cells. Other authors have developed cells which are perfectly adequate for the tasks for which they are designed. This particular cell design is optimized for parallel spectral and electrochemical studies of flavoproteins which may have negative redox potentials, potentials that are separated by only 2 mV, that may require long times to equilibrate and for which mixtures of optically transparent mediators do not exist. THEORY Coulonzetric titration. In coulometric experiments involving redox-active proteins. a mediator is necessary to transfer electrons from the electrode to the protein. This mediator is required to have: (a) the ability to rapidly transfer an electron to the reactive site of the protein, (b) the capability of being rapidly reduced at the electrode surface, and (c) a redox potential considerably more negative than that of the protein. This last requirement is in accord with Marcus theory (25,26), which predicts that the log of the rate of homogeneous electron transfer is proportional to the redox potential difference between the electron donor and electron acceptor. Viologen dyes are the best known electron transfer mediators for this purpose (21). Methyl viologen (MV)’ (E”’ = -0.450 V) was used in this work because it has the most negative redox potential of the commercially available dyes and therefore should transfer electrons to protein most rapidly. When the potential of the system is controlled at a value (-0.480 V) negative ’ Abbreviations used: MV. methyl viologen; IDS. indigo disulfonate: IDS,,.,,. reduced indigo disulfonate: BV. benzyl viologen: DHNQ. 1,4-dihydroxynaphthoquinone: IDS,,,. oxidized indigo disulfonate; R2G. Rosinduline 2G: FMN. flavine mononucleotide; SHE, standard hydrogen electrode; EFL,,, oxidized enzyme: EFIH”. one electron reduced enzyme: EFl,,,,,H-. two electron reduced enzymes.
298
MARIAN
T.STANKOVICH
enough to reduce MV during a coulometric experiment, the transfer of electrons from the electrode to flavoproteins occurs in the following sequence of reactions: MVii
+ eme MV+”
Ill
MV+’ + EFl,,, + H’ + MV++ + EFlHO MV+O + EFlHO = MV++ + EFl,,,H-
[2a] [2b]
(EFI,, is oxidized flavoprotein, EFIH” is the one-electron-reduced flavoprotein, a radical: EFlrcdH- is the two-electronreduced flavoprotein). Reactions [2a] and [2b] represent the process by which MV++ , the electrochemically active species, is regenerated in the solution. Because of this cyclic mechanism of regeneration, a high concentration of MV++ is stabilized at the electrode surface and a constant, fast rate of electron transfer occurs (Eq. [I]) (21.27,28). Since the current is integrated, the number of coulombs (Q) transferred to the protein can be accurately determined. In the case of flavoproteins, spectra can be periodically recorded throughout the reduction; thus, absorbance can be obtained as a function of (!. The endpoint is determined by the appearance of completely reduced enzyme. Calculations made from these measurements yield the number of equivalents transferred to a mole of protein at a controlled potential and in agiven time interval. Potentiometric titration. The potentiometric titration gives optimal results when performed in the presence of two kinds of mediators. an electron transfer mediator, e.g., methyl viologen and redox mediator (9.20,29.30) or a mixture of three to five redox mediators (9.12.13). The redox mediators are chosen to have redox potentials very near that of the protein. When in equilibrium with the protein, the redox mediator is capable of poising the potential of the indicator electrode. As in the coulometric titration, electrons are transferred from the electrode to MV at very negative potential (Eq. [I]). The concepts and pro-
cedures involved are illustrated by the following description of the first electron transfer to a flavoprotein. Initially. electrons from MV+” are transferred to both the protein (Eq. [2a] and [2b]) and the redox mediator (Eq. [3]). As the concentration of the one-electron-reduced form of the protein (EFIH”) increases. it equilibrates with the redox mediator (e.g., indigo disulfonate, IDS) (Eq. [4]). 2MV+(’ + IDS,,, $ 2MV’ + + IDS,,,1 2EFlH” + IDS ox = 2EFLx + IDS,,,,
[3] [41
(IDS,,, is oxidized indigo disulfonate; IDS,,, is reduced indigo disulfonate.) At each point in the titration the system is allowed to equilibrate. Then both the potential (E) and the spectrum of the system are recorded. The concentration of EFl,,, and EFlH” are calculated from the spectra, corrected for the absorbance of the redox mediator (see Results. potentiometric titration) using published values of molar absorptivity. For flavodoxin. EFIH” has E-,,x,,= 4500. and e$15= 2 100 M ' cm-‘, and EFI,,, has l qq5= 10,200 Mm' cm-’ (17). Redox potential (,!?,‘I) is obtained from the Nernst equation (Eq. IS]) by plotting E versus log ((EFI,,,)/(EFlH”). 0.059 (EFlox) E = E;’ + log ~ n (EFlHO)
[51
(E is the measured potential; Ey’ is the redox potential: n is the number of electrons transferred in the reduction step.) MATERIALS
AND METHODS
Flavodoxin was prepared from Megcrsphera efsdenii (formerly Peptostrrptoc,oc.(‘us elsdenii) according to the method of Mayhew and Massey (17). Glucose oxidase (EC I. 1.3.4) was prepared using the method of Swoboda and Massey (31). Methyl viologen (MV), and benzyl viologen (BV), were purchased from British Drug House, Poole. England; 1.4-dihydroxynaphthoquinone (DHNQ) and Rosinduline 2G
SPECTROELECTROCHEMICAL
CELL
FOR
ENZYME
REDUCTION
299
(R2G) were purchased from K & K Labs (Division ICN Life Sciences, Plainview. N. Y.). Indigo disulfonate (IDS), purified, was kindly supplied by Dr. Fred Guengerich. Vanderbilt University. The buffers used were pH 5.25, 0.1 M sodium acetate: lpH 7.25, 0.1 M sodium phosphate; for pH 23 8.26, 0.044 M sodium pyrophosphate-HCI; 4 for pH 9.25-O. 1 M glycine. Deionized water, 18 Mohm resistance was used to make 5 solutions. Prepurified nitrogen was passed over Ridox scrubber (Fisher Scientific Co., Springfield. N. J.) for further oxygen removal. A potentiostat. Princeton Applied Research, Inc. (Princeton, N. J.) (PAR) 6 Model No. 173 was used to control the I+--7 potential: a PAR No. 179 coulometer was 6 used to integrate the current, and a PAR FIG. 1. Spectroelectrochemical cell; main body is No. 174A polarographic module was used constructed of quartz. (A) Front view. electrodes are to determine the potential of the dye Rosinin the plane of the paper. (1) Main connection of nitroduline 2G. A Varian Cary 219 UV-visible gen line; (2) sidearm; (3) gold foil working electrode: chloride reference electrode; (5) recording spectrophotometer was used (4) silver-silver silver-silver chloride auxiliary electrode: (6) standard to record spectra. quartz cuvette; (71 glass stirrer. (B) Side view. All potentials are reported versus the 3-ml cell is rotated 90” to give a clearer view of (I) and (2). standard hydrogen electrode (SHE). The working electrode was a gold foil 2.6 x 0.6 cm. The auxiliary electrode was a silver erence electrode was checked before and wire, contained in a fritted (frit is fine after each experiment against standard porosity) compartment which was plugged ferro-ferricyanide (32). The anaerobic with agar and was filled with KCI solution spectroelectrochemical cell was made from (Fig. I). The silver wire becomes coated a standard 3-ml quartz cuvette by adding with silver chloride during the course of sidearms with standard taper ground quartz electrochemical reduction reaction. Re- joints. The stirrer was a magnet sealed in action at the auxiliary electrode is glass or polyethylene. The auxiliary and reference electrodes Ag + Cl- - AgCl + (J-. were stored anaerobically in 5 mM diThis electrode is used to prevent the gen- thionite solution for 12 h prior to the eration of 0, at the auxiliary electrode, experiment. Shortly before the experiment a during electrochemical reduction. This degassed solution of 10 mM KCI, buffered electrode can be used for an oxidative to pH 5.25, was placed in the auxiliary titration; first the AgCl will be reduced electrode compartment (Fig. I). To this then H,O will be reduced to produce H, solution was added SO ~1 of 4 M glucose as at a platinum auxiliary electrode. and 10 ~1 of 1 mM glucose oxidase (oxygen The reference electrode was a silverscrubber). The compartment was sealed by silver chloride electrode. Electrical contact inserting the electrode, and was stored in was made to the solution through an dithionite until the experiment started. Only asbestos fiber: a small agar plug was used to the tips of the electrodes are immersed in minimize diffusion. The potential of the ref- dithionite. The absorbance of the solution
300
MARIAN
T. STANKOVICH
FIG. 2. Controlled potential coulometric titration of flavodoxin. x lo-” M and 4.86 x IO-” MV in 4.42 ml of 0. I M sodium phosphate (2) n = 0.5 electronsimol FMN, (3) n = 1.0 electronimol FMN (5) n = I .89 electronimol FMN.
E = -0.480 V. Flavodoxin 3.52 pH 7.25. (1) Oxidized flavodoxin, (4) n = 1.49 electronsimol FMN.
was monitored before the degassing pro- pressure. At each point in the reduction, cedure and the addition of electrodes. After the spectrum was recorded within 2 min the addition of electrodes, the absorbance after the desired number of coulombs was transferred. No spectra1 changes occurred and the potential were monitored for 3045 min to insure that dithionite was not after 2 min time, this indicated that equilibleaking out, reducing the enzyme. rium was established within 2 min of each Coulometric experiment. The coulometric transfer of coulombs to the solution. Values experiments on flavodoxin and MV were for 11 and the molar absorptivity of the performed at room temperature at pH 7.25, flavodoxin radical were obtained indeas outlined in the Theory section. A solution pendently at 445 and 580 nm by extra(4.40 ml) containing tlavodoxin or MV, polating the linear parts of the coulometric mediator dye, and buffer was placed in the titration curves (Figs. 2 and 3). This will cell. Flavodoxin is isolated and stored as be discussed more fully in the Results the oxidized enzyme (17). There are no re- section. For methyl viologen, the efficiency ducing agents or oxidizing agents present of electrogeneration in terms of the II value to contaminate the enzyme in the final steps was determined from the slope of a plot of of the isolation procedure. Glucose and A versus Q, using known values of molar glucose oxidase in pH 5.3 buffer (same absorptivity and spectra1 path length (23). concentrations as above) were placed in the The reductive titrations were not corroborated with oxidative titrations. Ferrosidearm to scrub the 0, in the atmosphere. The system was closed and subjected to cyanide is the mediator titrant with positive I I cycles of evacuation and nitrogen potential most commonly used to carry out equilibration. The electrodes were then oxidations. Reportedly it complexes methyl placed in the cell under positive nitrogen viologen the reducing mediator titrant (9).
SPECTROELECTROCHEMICAL
CELL
FOR
ENZYME
REDUCTION
301
P 445nm
'2.
d!--ieYo
115
n per mole FMN FIG. 3. Controlled potential coulometric vs amount ofcharge added to the system to residual 0, in the system. Absorbance
titration of flavodoxin. Plot of absorbance at 445 and 580 nm (n equals equivalents of charge). The nonzero intercept is due values have been corrected for methyl viologen absorbance.
Therefore, it should not be in the system. This should be checked in future experiments. Potentiornetric titrutions. The potentiometric titration was performed as outlined in the Theory section. Measurements were performed at a temperature of 25 +- 0S”C.
The process represented by Eq. [4] was allowed to come to equilibrium at open circuit. The equilibration time ranged from 5 to 15 min. The criterion used for equilibration was the stabilization of the potential reading. The potential was measured at the indicator electrode (the working elec-
302
MARIAN
T. STANKOVICH
trode in the controlled potential mode) and the spectrum of the system was recorded. In control experiments the electron transfer mediators (MV and BV) and the redox mediators (IDS, DHNQ, and R2G) were all titrated; the concentration and pH values were the same as those used in the protein experiments. The distributions of the oxidized and reduced forms of the dyes were obtained as a function of potential. The total absorbance (contributed by both oxidized and reduced species of the dye) at each wavelength can be measured as a function of potential. Wavelengths of interest are usually 580 and 445 nm, where EFIH” and EFI,,, absorb maximally. These measurements were later used to correct for mediator contribution to the combined flavoprotein-mediator spectrum at each measured potential value. This method was used for IDS at pH values other than 7 where no convenient isosbestic point existed. At pH 5.3, the isosbestic point for IDS is at 416 nm. The difference in molar absorptivity between the EFIH” and EFl,,, at this wavelength is not great enough to allow independent spectrophotometric measurement of concentrations. However, the absorbance of IDS at 445 nm is small and directly proportional to the potential of the system. (The total L!A,,, = 0.04A for total reduction of the dye). The equilibrium potential and total absorbance at 445 nm are measured for the system containing enzyme and dye. The contribution of dye to the absorbance at 445 nm is subtracted and the concentrations of EFI,,, and EFIH” are calculated from the change in absorbance t 17). For IDS. at pH 7.0. a convenient isosbestic point exists at 460 nm. The change in absorbance at this wavelength was used to calculate the concentration of oxidized and reduced enzyme as described previously (20). DHNQ has a very small absorbance at 445 nm ttJJs = 2000 M-’ cm-‘): it has no absorbance at wavelengths longer than 550 nm. Therefore the concentration of EFIH” can be
directly measured from the spectrum at 580 nm t 17). Therefore when DHNQ was used as a mediator, no spectral correction was needed. The contribution of R2G to the glucose oxidase spectrum was subtracted as described previously (23). The viologens were used as both electron transfer and redox mediators for the second electron of flavodoxin. The reduced form of the viologens absorbs at long wavelengths where flavodoxin does not. From the absorbance at 725 nm, the entire MV”’ spectrum was calculated and subtracted from the combined MV-protein spectrum. Titrations of flavodoxin were performed at pH values 5.25, 7.25, 8.25, and 9.3. At pH 7.25 the E”’ for flavodoxin was measured using two dyes for each of the electron transfers to show that there was no binding of enzyme to mediator. Undetected binding of mediator to flavoprotein will cause appreciable error in redox measurements (33). The E"' for the first electron transfer was measured using the redox mediators DHNQ and IDS; both MV and BV were used to measure E”’ for the second electron transfer. Anaerobic precautions were the same as those described for the coulometric experiment. Glucose oxidase was titrated at pH 5.3. Anaerobic procedures were less rigorous than those described for flavodoxin and MV. The auxiliary and reference electrodes were not soaked in dithionite prior to the and no atmospheric oxygen experiment. scrubber was used. Glucose oxidase (1.6 x lop5 M) and R2G (4 x IO-” M) were coulometrically reduced at a potential of -0.155 V. The R2G (E"' = 0.080 V) acted as both electron transfer mediator and redox mediator. Because of the low concentration of mediator. electron transfer to the enzyme was slow (20 to 40 min) and an additional 20 to 40 min was required for equilibration (23). The efficiency of the reduction pro cedure was determined by comparing the coulombs of charge t Q) transferred to the system, with the coulombs of charge
SPECTROELECTROCHEMICAL
theoretically needed to produce the ured change in concentration of and EFI,,,,H. Concentrations of and EF&Hwere calculated from bance measurements and published absorptivities (23).
CELL
measEFIH” EFIH
absormolar
RESULTS Coulometric
Titmtiorl
The anaerobicity of the cell was tested by coulometrically titrating 2.5 x 10d5 M MV. A plot of Acio;: versus Q was linear with a nonzero intercept. The nonzero intercept corresponded to the amount of current consumed by oxygen initially in the system. On the average, 1.2 x lo-” M oxygen was initially present. The slope yielded the published value for the molar absorptivity of MV (12,400 M-’ cm-‘) with less than 1% error (assuming II = 1) (21). No background correction was required. When the absorbance of 50% reduced MV was monitored as a function of time. the oxygen leak rate was found to be 0.02 nmot per minute or 0.13 mC per hour. Figure 2 shows the spectral change which occurred upon coulometric reduction of 3.25 x IO-: M flavodoxin. The concentration of flavodoxin was calculated from the measured absorbance and the known molar absorptivity (17). The absorbance spectrum of reduced MV interfered with the flavodoxin spectrum only at values of II > 1.75 (n = equivalents of charge per FMN). The molar absorptivities of the species EFI,,,; EFIH”. and EFl,.,,,H- differ substantially at 580 and 445 nm. Figure 3 is a graph of A 5xoand A,,, as a function of the number of reducing equivalents added per flavin. Absorbance values and Q values were corrected for the contribution from MV. (Absorbance corrections were performed as described under Materials and MethodsPotentiometric titrations). From the absorbance values for MV the concentration of the reduced form was calculated. From the concentration and volume the number of
FOR
ENZYME
REDUCTION
303
coulombs of charge transferred to MV was calculated and subtracted from the total number of coulombs transferred to the MV -flavodoxin system. The nonzero intercept corresponded to the titration of residual oxygen in the system (0.5 x IO-” M). The value of ALx,, increased linearly with the number of equivalents added to the system up to 1~= I equivalent/FMN. Between II = 1 and II = 2 it decreased linearly. ‘The A,,, decreased linearly with the number of equivalents added. up to f~ = 1 equivalentsi FMN: from )I = 1 to II = 2 it still decreased. but the slope was much less steep. The A5H,I maximum occurred at the same value of II as did the inflection in the A,.,, plot. These data were not corrected for oxygen leakage. The calculated E;,~,)for EFIH” was in perfect agreement with the literature value (17). Table I summarizes the coulometric reduction experiments on flavodoxin. Our spectral data are identical to those obtained previously (17). In both cases a stable spectrally distinct one-electronreduced species is formed. Spectrum 3 of Fig. -3 is essentially the spectrum of the electron-reduced species (EFlH”I. The spectra of EFIH” and EFI,,, fully oxidized species differ most at wavelengths 580 nm where only EFIH” absorbs and at 445 nm where EFI,,, absorbs much more strongly (E = 10.200 M ’ cm-‘) than EFIH” (E = 2100 hr-’ cm- ‘). Therefore between II = 0 and II = I. there is an equilibrium between EFI,,, and EFIH”: this is retlected in the spectra (Fig. 2) and in the plots (Fig. 3). That is. A Sx,)increases from II = 0 to II = I. while A,,, decreases. This equilibrium is broken at II = I. From PI = 1 to 11 = 2, the equilibrium is between EFIH” and EFI,.,,,H,. EFI,.,,,,H, does not absorb at 580 nm but has molar absorptivity of E = 1600- 1800 at 445 nm ( 17). Therefore. A,,,, decreases tI = 1 to tz = 2 as does A,,,. From experiment I in Table I. it can be seen that 100% EFIH” was formed at tr = 1.08 equivalents of FMN. an 8% error: likewise the two-electron reduction was
304
MARIAN
T. STANKOVICH TABLE
CONTROLLED
POTENTIAL
COULOMETRIC
I TITRATION or; FL4VoDoXIN.
pH 7.25
Experiments I Concentration enzyme (PM) Concentration MV (FM) Ratio dye/enzyme
35.2 48.6 1.4
2
3
53.8 25.0 0.46
55.3 25.0 0.45
4 47.1 100 2. I
First electron Average current (PA) Q calculated (n = 1) (mC) Q observed (mC) n measured n corrected Average corrected value
12.0 15.0 16.3 1.08 I .03 1.04 _f 0.02
6 22.8 25.6 1.12 1.06
6 23.6 26.7 1.13 1.06
25 20.0 21.2 1.06 I .02
Second electron Average current +A) Final current @A) Q observed (n = 2) (mC) n measured n corrected Average corrected value
5.5 1.5 15.5 1.03 0.98 1.03 -t 0.02
3.0 I.0 24.6 I .08 1.02
3.0 I.0 26.5 1.12 1.05
18.0 6.0 21.8 1.09 1.05
predicted observed Q initial 0, reduction (mC) Estimated 0, contribution to flavodoxin (%) Ass,, A,,,
0.158 0.158 0.6
0.242 0.241 0
0.249 0.246 0.2
0.212 0.212 1.6
5
6
6
3.5
complete at n = 2.12 an overall error of 6%. The average error in n was 1.10 & 0.02 for the first electron and 1.08 ? 0.02 for the second electron. The error for MV was less than 1%. This difference in percentage error between the flavodoxin and MV experiments can be ascribed to the timescale of the experiments (Table 3) and the difference in reactivity of the reduced species with oxygen. MV+O reacts with oxygen rapidly, therefore any oxygen initially present will be reduced before MV++ starts to reduce (34). EFIH” reacts more slowly with oxygen, therefore EFIH” and O2 can exist in solution simultaneously (35). It was found that for flavodoxin solutions an average of 0.85 mC was required to reduce the “initial oxygen” (Table 1) as opposed to the 2.5 mC for MV++. This initial difference of 2.5-0.85 mC, plus the leak rate
of 0.12 mC/h provided the value captioned “Estimated 0, contribution to flavodoxin” in Table 1. This value (approximately 5%) was used to correct the n value for the first and second electron transfers, resulting in 1.04 & 0.02 and 1.03 * 0.02, respectively.
Potentiornetric
Titrution
Figure 4 shows the results of a series of potentiometric titrations on flavodoxin. The individual values (Table 2) were obtained as described under Materials and Methods. Our data are in excellent agreement with those of Mayhew et crl. (24) as is shown by the solid lines which are the best fit to their redox data. The slope of the potential versus pH curve for the first electron transfer is 60 mV/pH unit. Ey’ values for the first electron at pH values
SPECTROELECTROCHEMICAL
.-s
5
FOR
ENZYME
REDUCTION
305
DISCUSSION
0I\ .
-0.4
CELL
6
7
PH
8
9
10
We have developed a spectroelectrochemical cell in which potentiometric, coulometric, and spectroscopic experiments on oxygen-sensitive samples may be conveniently performed. The primary advantage of our system lies in the degree of anaerobiosis which can be maintained. The oxygen leak rate for our cell is 20-fold lower than that of Hawkridge and Kuwana (21. 22) (see Table 3). Data for the 0, leak rate are not published for other systems in Table 3. Thus we are able to successfully engage in studies under conditions where oxygen interference can cause the most problems, i.e., at potentials more negative than -0.400 V and for systems that require several hours to reach equilibrium.
FIG. 4. Effect of pH on the redox potential of flavodoxin. (0) Experimentally determined values for E”’I and E;‘. (0) experimental point for Ey’. previously determined value. The solid lines in the figure are the best fits from the data of Mayhew rf al.
(24).
5.25, 8.26, and 9.25 have not been directly measured before. E;’ for the second electron above pH 7.0 was constant at -0.373 V. Figure 5 shows the individual Nernst plots. The potentiometric titration experiments with glucose oxidase show that the agreement between charge transfer and spectral data depended on the time required for the experiment. In an experiment where electron transfer was relatively rapid and only two experimental points were measured (29 and 47% reduced), agreement was 100%. In a second experiment where seven points were measured, agreement was 95% for the first four points (2.5 h); 77% for the last three points (4 h). The redox potential data for both electrons of glucose oxidase at pH 5.3 are given in Table 2. The redox data obtained from three experiments were again in excellent agreement with values obtained by conventional methods (23).
‘lredl
FIG. 5. Representative Nemst plots for the twoelectron transfers of Ravodoxin at different pH values. The y axis on the right is used for curves (a) and (b), (shown in dashed lines). (a) First electron. pH 5.3: (b) first electron pH 7.25. The y axis on the left is used for curves (c) through(f). (c) First electron, pH 8.26; (d) first electron, pH 9.3: (e) second electron. pH 5.25: (f) composite curve for second electron, pH values 7.25-9.25. pH 7.25 (0): pH 8.25 (A); pH 9.25 (a,.
306
MARIAN
T. STANKOVICH TABLE
REDOX
-0.024 -Il.(lXh
2
POTENTIAI.
t 0 WI + KIM?
5 4
DA-I A
1).33ll
0 iIll?
I) 327 + 0 ,Y)? 7.??
I). 130 -0 II4 -0.144 0.127 -0 IX
+ + f f +
U.uu? 0 uul U.uuI u (XI? O.WI
7 K 7 6 Y
196 1 U.UU3
>
DHNQ
h 4
BV BV
X.26
-0
Y 25
Cl.??3 f O.(XK15 -Il.??7 - O.(KI
NOW2 IDS DHNQ DHNQ IDS
0 I I I z U.WI -0 171 + O.wu~ -
-
To demonstrate the capacity of our cell, we have determined the redox and spectral properties of three previously investigated systems. The molar absorptivity for methyl viologen was determined coulometrically to better than 1% precision. This despite the fact that reduced methyl viologen is extremely oxygen reactive. with an E”’ of -0.450 v. The redox and spectral properties for both electron transfers of flavodoxin were determined with greater than 95% precision, at several pH values. These results include redox potentials for the first electron transfer at pH 5.25. 8.26. and 9.25 which have not been directly measured before. Each experiment required 2-3 h to complete, thus demonstrating the performance of our cell over periods of prolonged anaerobiosis while in the presence of an oxygen-reactive
6 >
,,
-
0.371 -0.373
f u IWII + (1 ,“,I -
? 7 5
nv
-,I 3’iu
HV MV -.
Il.350 ll.44~ -~
UV BV “V
-
HV MV DHNQ MV MV
IMV
0 440
MV
HV MV
U.350 0 440
HV HV MV
compound of notably negative redox potential (-0.373 V). Finally. we measured the redox potential for both electron transfers of glucose oxidase at pH 5.3. Glucose oxidase is an enzyme for which the redox potentials foreach of the two electron transfers are almost identical (23). Yet we were able to accurately determine both values, even though the total experiment required 6.5 h to complete. These three tests clearly demonstrate the effectiveness of our technique. Table 3 affords a comparison of the capabilities of our system with the three other comparable systems currently in the literature. The importance of performing the coulometric titration and thus measuring the spectral properties of flavoproteins was pointed out in the introduction. By performing two experiments our cell enables us to
SPECTROELECTROCHEMICAL
CELL TABLE
C~MPARISION
OF ELECTROCHEMICAL
FOR
ENZYME
3 CELL
CHARACTERISTICS
I,, ml” x-30
x-30 -
-
307
REDUCTION
Ill,” ml,”
IT h XI
30
0 75
0.9x + 0 05 I Of3 T KO!
4 hNo. of rlectron\ for each compound
-
70- l4lJ
I hl
A, 1.
” Exprssxd m term\ of 10~s of abaorbancc due tu reduced viulugen ” II = equivalenfr. Q = coulombs: E = potential: .4 - ;lhcorbance.
measure accurately II, E, E”’ for purified flavoproteins as shown in Table 3. As shown in Table 3. the Hawkridge-Kuwana cell is designed to perform all three measurements (20-22). Hawkridge’s and Kuwana’s cell has an error of 2 to 3% when measuring E or II values for the cytochrome systems for which it was designed (these have positive potential value). However, for 1 mM benzyl viologen alone there is an 8% error in determining II or E. In our present cell, methyl viologen was reduced with less than I% error. However, because of the longer times scales of the experiment, coulometric titration of flavodoxin has a 3 to 4% error. The Hawkridge-Kuwana cell would be expected to yield less accurate data for flavodoxin due to: (a) its higher error with BV, (b) its higher 0, leak rate. Watt’s ( 15) method was designed primarily for the study of purified enzymes with negative redox potentials with no spectral properties. It yields n and I?’ values with accuracy, but no spectral data can be obtained using this system (Table 3). The system of Hendler et ~(1. (12) was designed to yield absorbance data as a function of potential
for membrane-bound cytochrome systems and in mixtures of optically transparent mediator dyes. The presence of other redox active species in this membrane system makes accurate coulometric titrations in this system very difficult. The potential of their system is computer controlled and the spectral data are computer analyzed. Their experimental design yields accurate data for their complex system. However, it is too complex and inaccessible to other workers: simpler approaches yield more data and equivalent data for purified enzymes. In addition, even if this computer-controlled system were available, coulometric titration is important for flavoenzymes because optically transparent dyes are not available. Therefore, as these other experimental designs and cells were optimized for data collection for particular systems, our system is optimized for measuring spectral (E) and electrochemical data (n, E”‘) for flavoproteins. However, the lower leak rate of our cell would be an advantage in any of the other systems. We are further improving our system by increasing the electrode area. the redox mediator concentra-
308
MARIAN
T. STANKOVICH
tion. and the stirring rate in order to accelerate charge transfer and decrease equilibration time. CONCLUSION
In conclusion, all four methods outlined in Table 3 can provide redox potential data on enzymes for which e values are known or for those enzymes where successive electron transfers are sufficiently separated. Only our cell and that of Hawkridge and Kuwana were designed to enable complete characterization of purified enzyme. Of these latter two, our cell clearly excels at maintaining anaerobiosis. In addition, it is simple to construct and maintain, and it will fit into a standard spectrophotometer. The accuracy of our system has clearly been demonstrated by reproducing values of E, E"', and II for the well-characterized proteins, glucose oxidase and flavodoxin, and also for methyl viologen. ACKNOWLEDGMENTS My thanks to V. Massey. A. Bard, L. Schopfer for helpful suggestions.
E. Moore,
and
REFERENCES 1. Foust. G. P., Burleigh, B. D., Jr., Mayhew. S. G.. Williams. C. H., Jr., and Massey. V. (1969) And. Biochem. 27, 530-535. 2. Burleigh. B. D.. Foust, G. P.. and Williams, C. H.. Jr. (1969) And. Biochrm. 27, 536-544. 3. Lambeth, D. 0.. and Palmer, G. (1973) J. Bid. Chum. 248, 6095-6103. 4. Dutton, L. (1975) in Methods in Enzymology (Fleischer, S., and Packer, L.. eds.), Vol. 54, Part E. pp. 411-435. Academic Press, New York. 5. Moore, E. G.. Ghisla. S.. and Massey, V. (1979) J. Bid. Chem. 254, 8173-8178. 6. Mayhew. S. G. (1978) Eur. J. Biochem. 85, 535-547. 7. Massey. V., Muller. F., Feldberg, R.. Schuman, M., Sullivan, P. A., Howell, L. G., Mayhew. S. G.. Matthews, R. G., and Faust. G. P. (1969) J. Bid. Chrm. 244, 3999-4006. 8. Muller. F., and Massey. V. (1969) J. Bid. Chrm. 244, 4007-4016. 9. Wilson, G. S. in Methods in Enzymology (1978) (Fleischer, S., and Packer, L.. eds.), Vol. 54, pp. 396-410, Academic Press, New York.
10. Barman, B. G., and Tollin, G. (1972) Biochemi.ctr~ 25, 4755-4759. 11. Swartz. D. B., and Wilson. G. S. (1971) Anal. Biochem. 40, 392-400. 12. Hendler, R. W.. Songco. D.. and Clem. T. R. (1977) And. Chrm. 49, 1908- 1913. 13. Hendler. R. W. ( 1977)Anal. Chrm. 49, l914- 1918. 14. Hendler. R. W.. and Shrager, R. I. (1979) J. Bid. Chrm. 254, 11288- 11299. 15. Watt, G. D. (1979) And. Biochrm. 99, 399-407. 16. Hendler. R. W. Towne, D. W.. and Shrager, R. I. (197.5) B&him. Biophys. Acta 376, 42-62. 17. Mayhew. S. G.. and Massey. V. (1969) J. Bid. Chrm. 244, 794-802. 18. Singer, T. P. and Edmondson. D. E. (1978) in Methods in Enzymology (Fleischer. S., and Packer. L.. eds.). Vol. 53, Part D. pp. 397418, Academic Press, New York. 19. Iyanagi, T.. Makino. N., and Mason, H. S. (1974) Bkxhemisrry 13, 1701- 1710. 20. Hawkridge. F. M.. and Kuwana. T. (1973) And. Chef?. 45, IO?]- 1027. 21. Szentrimay, R.. Yeh, P., and Kuwana, T. (1977) in Electrochemical Studies of Biological Systems. ACS Symposium No. 38, (Sawyer, D. T.. ed.). pp. 142-169. American Chem. Sot.. Washington. D. C. 22. Heineman, W. R.. Kuwana, T., and Hartzell. C. R. ( 1972) Biochum. Biophys. Rrs. Commun. 49, 1-8. 23. Stankovich. M. T.. Schopfer. L. M.. and Massey, V. (1978) J. Bid. Chrm. 253, 4971-4979. 24. Mayhew. S. G., Foust. G. P., and Massey. V. (1969) J. Bid. Chem. 244, 803-810. 25. Marcus. R. A. (1965) J. Chem. Phys. 43, 679701. 26. Marcus, R. A. (1964) Annu. Rev. Phys. Chem. 15, 155- 196. 27. Kuwana. T., and Winograd, N. (1974) in Electroanalytical Chemistry (Bard, A. J.. ed.), Vol. 7, pp. l-78, Dekker. New York. 28. Bard, A. J.. and Santhanam. K. S. V. (1970) in Electroanalytical Chemistry, (Bard. A. J., ed.). Vol. 4, pp. 215-315. Dekker. New York. 29. Moreno. C. G., Choy, M.. and Edmondson. D. (1979) J. Bid. Chem. 254, 7630-7635. 30. Anderson, J. L.. Kuwana. T.. and Hartzell. C. t 1976) Biochemistry 15, 3847-3855. 31: Swoboda. B. P., and Massey. V. (1965) J. Bid. Chrm. 240, 2209-2215. 32. O’Reilly, J. E. (1973) Biochim. Biophys. Acfu 292, 509-515. 33. Stombaugh. N. A., Sundquist, J. E., Burris. R. H., and Orme-Johnson, W. H. (1976) Biochemisfo 15, 2633-2641. 34. Small. R. D., and Scaiano. J. C. ( 1977) J. Phys. Chrm. 81, 2126-2131. 35. Mayhew. S. G. (1971) Biochim. Biophys. Acln 235, 282- 302.