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approximations for intensities of photoionization bands. Intensities of photoionization bands should be obtained by integrating the area of the band and all of its satellite structures. Since peak intensities and widths are a function of the lifetime of the hole-state, the chemical state of the element, contamination, phonon broadening, satellite structure, excitation energy, etc., these variables can also contiibute to uncertainties. 4) The use of ESCA for quantitative analysis and the handling of systematic errors has recently been discussed (4-9).
(2) (3) (4) (5) (6) (7) (8) (9)
"ESCA, Atomic, Molecular, and Solid State Structure Studied by Means of Spectroscopy", Almqvist and Wiksells, Uppsala. Sweden, 1967. J. H. Scofield, Lawrence Livermore Laboratory Report UCRL-51326 (1973). J. T. Huang, J. W. Rabalais, and F. 0. Ellison, J. Electron. Spectrosc., 6, 85 (1975). W. J. Carter, G. K. Schweitzer, and T. A. Carlson, J. Electron. Spectrosc., 5, 827 (1974). V. I. Nefedov, N. P. Sergushin, I. M. Band, and M. B. Trzhaskovskaya, J. Electron. Spectrosc., 2, 383 (1973). C. A. Tolman. W. M. Riggs, W. J. Linn, C. M. King, and R. C. Wendt, Inorg. Chem., 12, 2770 (1973). C. D. Wagner, Anal. Chem., 44, 1050 (1972). R. S. Swingle, Anal. Chem., 47, 21 (1975). P. C. Kemeny, J. G. Jenkin, J. Liesegang, and R. C. G. Leckey, Phys. Rev., 9, 5307 (1974).
LITERATURE CITED (1) K. Siegbahn, C. Nordling, A. Fahlman. R. Nordberg, K. Hamrin, J. Hedman, G. Johansson. T. Bergmark, S. E. Karlsson, I. Lindgren, 8. Lindberg,
RECEIVEDfor review October 3, 1975. Accepted December 8, 1975. Supported by the U S . Army Research Office.
Spectroelectrochemical Cell for Anaerobic Transfer of Biological Samples for Low Temperature Electron Paramagnetic Resonance Studies James L. Anderson' Department of Chemistry, The Ohio State University, Columbus, Ohio 43210
An electrochemical cell is reported, suitable for generation, optical monitoring, and anaerobic transfer of multiple aliquots of concentrated enzyme or other biological samples for analysis by low-temperature electron paramagnetic resonance (EPR) or other external physical techniques. Results are reported for low-temperature EPR measurements on samples of cytochrome c oxidase in varying redox states. Present address, D e p a r t m e n t of Chemistry, North D a k o t a S t a t e University, Fargo, N.D. 58102.
Cytochrome c oxidase is the oxygen-reducing terminal enzyme of the respiratory chain. Cytochrome c oxidase has four active one-electron metal centers: two heme iron and two copper components. In the fully oxidized state, where Fe(II1) and Cu(I1) should exhibit EPR resonances, only 30-40% of the heme and copper components are detectable ( I ). Low spin Fe(I1) and Cu(1) are also EPR-undetectable. Little evidence has been found for any high-spin Fe(I1). One of the coppers is believed to be EPR-undetectable (I, ANALYTICAL CHEMISTRY, VOL. 48, NO. 6, MAY 1976
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B L
STIRRING PASSAGETO O P T l C l L WINDOW
LUCITE
1
LJCITE SIDE COMPARTMENT
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Figure 1. Electrochemical cell and anaerobic transfer apparatus ( A ) Exploded view of electrochemical cell for anaerobic enzyme titration. Valves V 1 , V2, V3 connect sampling tube, counter electrode, and reference electrode Anaerobic sample transfer apparatus to cell. Lucite side compartment, quartz windows, and lucite cover plate are epoxied to cell body. (8)
2). The high-spin (g = 6) a n d low-spin (g = 3) Fe(II1) heme signals, and the d e t e c t a b l e Cu(I1) (g = 2) signals are discussed.
EXPERIMENTAL Purified cytochrome c oxidase samples were prepared from beef heart mitochondria by Hartzell ( 2 ) . Other materials, procedure, and methods for the coulometric titrations were as described previously ( 3 ) . A new cell, shown in Figure 1(A), was designed for sampling of aliquots of enzyme at varying states of reduction. The three-electrode design “sandwich” cell was similar to a previously described cell ( 4 ) in the use of polymethylmethacrylate body, with three ports terminated by Hamilton valves with Teflon stopcocks, and with a tin oxide working electrode clamped to the cell by means of pressure against a Viion O-ring. Cell design criteria were the following: 1) volume of several ml to enable sampling of up to ten 0.3-ml aliquots; 2) electrodes and ports placed near bottom of cell to maintain electrode/solution contact for continuation of electrolysis of a maximum number of aliquots; 3) short optical path to allow monitoring of optical absorption of concentrated enzyme solutions required for EPR samples; 4) efficient stirring and maximum ratio of working electrode area to sample volume, to minimize electrolysis times for relatively large volumes of concentrated enzyme. The tin oxide working electrode was chosen because of the immunity to hydrogen evolution at potentials required for generation of the reductive titrant used (benzyl viologen, electrolyzed a t -0.62 V vs. Ag/AgC1/1 M KCl). The electrode served as the cell bottom to meet criterion 2 above. The optical path (path length 2.52 mm) consisted of a side compartment ca. 5 mm in diameter, with quartz windows, placed as near to the cell bottom as possible. The stirrer-propeller consisted of a disk of polymethylmethacrylate (1.9-cm diameter, 0.6 cm high) with four slots milled on a rotary table a t 30” angles to the vertical and oriented so that stirring convection moved solution downwards toward the working electrode. A magnetic stirring bar core was epoxied into a hole drilled perpendicular to the axis of the disk. The internal cell diameter (2.1 cm) was slightly larger than the stirrer diameter. A narrow lip a t the bottom of the cell body prevented the stirrer from direct contact with the tin oxide working electrode. The very efficient and smooth stirring ensured thorough mixing and transport of solution to the narrow side window, without any bubbling or frothing. Slots milled into the sides of the cell allowed reproducible positioning in a mating holder for alignment in the optical path of the spectrometer. The volume of the cell described here was 3 ml, sufficient for ca. 8 samples of ca. 0.3 ml without refilling. A greater number of aliquots could readily be accomodated by increasing cell volume, consistent with criterion 4 above. Prior to introduction of sample, the 922
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ANALYTICAL CHEMISTRY, VOL. 48, NO. 6, MAY 1976
entire cell assembly was made initially anaerobic by alternate evacuation and pressurization (via the sampling port, seen in Figure 1A) with nitrogen which had been passed over hot copper turnings to eliminate traces of oxygen. The reference electrode compartment was then filled with deaerated 1 M KC1 while all cell valves were closed. All valves were then reopened, and the reference compartment was made anaerobic by repetition of the vacuum degassing procedure. All valves were closed, while the cell was pressurized with nitrogen, and sample was placed in a bulb fitted with a stirring bar and connected to both vacuum-nitrogen line and the cell sampling port. The sample was vacuum-degassed, and forced through the sampling port into the cell by alternate evacuation and nitrogen pressurization. The sample valve was closed while the cell was under pressure. Valves to the reference and counter electrode compartments were then opened and the compartments briefly vented to atmospheric pressure, allowing pressurized solution to be forced into these compartments. The procedure has been described in greater detail elsewhere ( 4 ) . The very narrow valve passages prevented significant intermixing of solution components between compartments for hours longer than required for these experiments. Anaerobic transfers of enzyme samples to calibrated E P R tubes were carried out by means of the transfer apparatus shown in Figure 1B . The procedure for EPR sampling was as follows. 1) The total absorbance change at 603 nm between oxidized and reduced oxidase for a given concentration was determined for an enzyme sample. 2) Using another sample, the initially-oxidized enzyme was titrated reductively to the desired redox level as determined by the spectral change at 603 nm. Aliquots were anaerobically transferred to E P R tubes (see steps 3-7). 3) After the desired redox level had been attained, the solution outlet port was dried with a paper towel wick to minimize contamination by oxygen dissolved in any remaining liquid. 4) The cell and EPR tube were attached to the anaerobic transfer apparatus, which was repeatedly vacuum-degassed and nitrogen-filled for several minutes. Valves separating cell and counter and reference electrode compartments were closed, the transfer assembly was evacuated, and the system was isolated from the vacuum pump. The side-arm stopcock (SAS) in the transfer assembly was closed. The cell solution outlet valve (closed throughout the above operation) was turned quickly through the open to the closed position, allowing a small portion of the solution to be drawn into the E P R tube. The SAS was opened, and the transfer assembly was pressurized with nitrogen. The latter forced the remainder of the solution into the EPR tube. 5 ) If insufficient solution had been transferred, the procedure was repeated. If sufficient solution had been transferred, the E P R tube was carefully frozen and stored in liquid nitrogen until spec-
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Figure 2. Optical absorption spectra of cytochrome c oxidase samples remaining in titration cell after anaerobic aliquot transfer 75 KM oxidase, 0.44 mM benzyl viologen, 0.89% (v/v) Tween 20, pH 7.0 phosphate buffer, ionic strength 0.15, cell path 2.52 mm. ( A ) initial fully oxidized enzyme, ( E ) 26% reduced, ( C ) 52% reduced, ( D ) 68% reduced, (0 fully reduced
t r a c o u l d be taken. T h e apparatus was pressurized w i t h nitrogen, t h e solution o u t l e t valve was q u i c k l y opened a n d closed t o equalize pressure in t h e cell, a n d valves t o t h e counter a n d reference comp a r t m e n t s were reopened. 6) T h e transfer apparatus was detached a n d washed. I t was vacu u m - d r i e d w i t h t h e n e x t EPR t u b e attached. 7 ) T h e solution in t h e cell was stirred f o r a t least 30 s a n d a spectrum was t a k e n t o determine t h e extent o f oxygen leakage i n t o t h e cell. Samples evidencing significant reoxidation were rejected. Careful adherence t o 1h e above procedure allowed “anaerobic” transfers w i t h less t h a n 3% reoxidation (less t h a n 6 n m o l o f 0 2 ) d u r i n g transfer, as determined by t h e spectrum o f solution remaini n g in t h e t i t r a t i o n cell. Samples were generated, anaerobically transferred, a n d transp o r t e d a t liquid nitrogen temperatures t o t h e E n z y m e Institute, U n i v e r s i t y of Wisconsin. Madison, Wis., where t h e EPR experim e n t s were performed. EPR experiments were run a t a frequency o f 9.206 G H z w i t h 10-gauss modulation. H e m e signals (g = 6, g = 3) were m o n i t o r e d a t 13 K a n d 3 mW power. Copper signals (g = 2) were m o n i t o r e d a t 29 K a n d 300 pW power.
RESULTS AND DISCUSSION Figure 2 illustrates the optically monitored spectra a t room temperature (24 “C) of cytochrome c oxidase samples taken for EPR analysis during a reductive titration. Spectra were taken on material remaining in the titration cell after anaerobic transfer and stirring. Normalized doubly integrated EPR absorption signals are plotted in Figure 3 as a function of percentage reduction of enzyme. Integrations were carried out by Helmut Beinert of the Enzyme Institute according to an unpublished procedure of Tore Vannglrd. The heme g = 3 signal decreases approximately linearly with increasing degree o f reduction and disappears a t ca. 75% reduction. The heme g = 6 signal increases approximately linearly from a negligible value in the fully oxidized state to a maximum in the vi-
0 25
0 50 FRACTION REDUCED
0 75
IO0
Figure 3. Normalized integrated EPR signal intensities Each signal is normalized to its maximum observed value. Points with vertical line at top are from oxidative titration. Points joined by lines indicate uncertainty limits for sample with excess oxygen introduction. All other points are from reductive titration. g = 6 signal is sum of rhombic and axial components
cinity of 50% reduction, and declines toward zero a t complete reduction. The g = 2 (copper) signal initially increases significantly from its value in the fully-oxidized enzyme, reaches a maximum a t ca. 25% reduction, and then declines approximately linearly with increasing reduction. Upon reoxidation, the copper signal returns to the high value observed a t 25% reduction. The variations with degree of reduction of the heme g = 3 and g = 6 signals, and of the copper g = 2 signal, are in general agreement with previous results obtained by chemical titration methods ( I , 5-7) and confirm the validity of the indirect coulometric titration approach (8) for techniques in addition to optical spectrometry (8-20). These results demonstrate the utility o f the electrochemical generation and the anaerobic transfer technique.
ACKNOWLEDGMENT The author is deeply grateful to Theodore Kuwana, The Ohio State University, and Helmut Beinert, University of Wisconsin Enzyme Institute, for their hospitality, encouragement and assistance in this work. The author also thanks Charles R. Hartzell (The Pennsylvania State University) for providing samples of cytochrome c oxidase.
LITERATURE CITED C. R. Hartzell, R. E. Hansen, and H. Beinert, Proc. Nat. Acad. Sci. U.S.A., 70, 2477 (1973). C. R . Hartzell and H. Beinert, Biochim. Biophys. Acta, 368, 318 (1974). J. L. Anderson, T. Kuwana, and C. R. Hartzell. submitted to Biochemistry. F. M. Hawkridge, Jr., and T. Kuwana, Anal. Chem., 45, 1021 (1973). B. F. Van Gelder and H. Beinert, Biochim. Biophys. Acta, 189, 1 (1969). D. F. Wilson, J. S. Leigh, Jr., J. G. Lindsay, and P. L. Dutton. in “Oxidases and Related Redox Systems”, Vol. 2, T. E. King, H. S. Mason, and M. Morrison, Ed., Williams and Wilkins, Baltimore, Md., 1973, pp 715-26. J. S. Leigh, Jr., D. F. Wilson, C. S. Owen, and T. E. King, Arch. Biochem. Biophys., 160, 476 (1974). T. Kuwana and W. R. Heineman, Bioelectrochem. Bioenerg., I,389 (1974). M. A. Cusanovich and D. C. Wharton, University of Arizona and Cornell University, private communication, 1973. R . H. Tiesjema, G. P. M. A. Hardy, and B. F. Van Gelder. Biochim. Biophys. Acta, 357, 24 (1974).
RECEIVEDfor review October 10, 1975. Accepted December 8, 1975. This work was supported by PHS-NIH Grant No. 19181.
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