50

Biochimica et Biophysica Acta, 1122(1992) 50-56 © 1992 Elsevier Science Publishers B.V. All rights reserved 0167-4838/92/$05.00

BBAPRO 34239

EPR and redox properties of Desulfovibrio vulgaris Miyazaki hydrogenase: Comparison with the Ni-Fe enzyme from Desulfovibrio gigas Marcel Asso a, Bruno Guigliarelli a, Tatsuhiko Yagi b and Patrick Bertrand a Laboratoire d'Electronique des Milieux Condens(s - URA CNRS 784, Unh'ersit~de Procence, Centre St J~r~me, Marseille (France) and b Department of Chemistry, Shizuoka Unh'ersitY, Shizuoka, (Japan) (Received 28 November 1991)

Key words: Hydrogenase;EPR; Nickel center; Iron-sulfur cluster We have carried out a detailed redox titration monitored by EPR on the hydrogenase from Desulfocibrio culgaris Miyazaki. Typical 3Fe and nickel signals have been observed, which are very similar to those given by Desulfocibrio gigas hydrogenase in all the characteristic redox states of the enzyme. This confirms that D. vulgaris Miyazaki hydrogenase is a Ni-Fe enzyme closely related to that from D. gigas, as was recently proposed on the basis of sequence comparisons (Deckers, H.M., Wilson, F.R. and Voordouw, G. (1990) J. Gen. Microb. 136, 2021-2028).

Introduction

Hydrogenases are bacterial enzymes that catalyze the reversible oxidation of molecular hydrogen. They are found in a number of bacterial genera, and present a great variety of biochemical and physicochemical properties [1]. Two classes of enzymes have been defined in bacteria of the genus Desulfovibrio: Fe and Ni-Fe hydrogenases. In the former class, the catalytic site is an iron-sulfur cluster whose structure is not completely elucidated, whereas it is a nickel center in the latter[l]. The properties of the hydrogenase from Desulfocibrio vulgaris Miyazaki are not well understood: although earlier results were interpreted in terms of the absence of nickel [2], several characteristics like its amino-acid composition and its subunit structure are typical of Ni-Fe hydrogenases [3,4]. Moreover, a recent molecular biology study has revealed a great sequence homology between this enzyme and the Ni-Fe hydrogenase from Desulfovibrio gigas [5]. To clarify this apparent discrepancy, we have carried out a detailed redox titration monitored by EPR on the hydrogenase from D. vulgaris Miyazaki. Our study demonstrates conclusively that this enzyme belongs to the Ni-Fe class, and that the structure and spatial organi-

Correspondence: P. Bertrand, Laboratoire d'Electronique des Milieux Condens~s-URA CNRS 784, Universit~ de Provence, Centre St. J~r6me, 13397 Marseille CEDEX 13, France.

zation of its metal centers are closely related to those of D. gigas hydrogenase. Detailed spin-lattice relaxation time measurements have also been performed in order to gain information on the distance between the 3Fe and nickel centers. Materials and Methods

Hydrogenase was solubilized from the particulate fraction of D. vulgaris Miyazaki by tryptic digestion [3] and purified by the published procedure [6], except that Ampholine Ele~rophocusing (pM 5-7) was employed instead of DEAE-Toyopearl chromatography. The preparation was homogenous on polyacrylamide gel electrophoresis. Determination of the metal content by plasma emission spectroscopy (Jobin Yvon JY 38 apparatus) yielded 1 :i: 0.1 g atom of nickel and 13 _+ 2 g atom of iron per 89000 g of protein. Samples of D. gigas and D. fructosovorans hydrogenases were kindly provided by Dr C. Hatchikian. They were purified and concentrated as previously described [4,7]. The redox titration was performed at 20°C in a 50 mM Tris-HCl (pH 7.7) solution under argon atmosphere. The enzyme concentration was 110 p.M. Potentials were adjusted with small amounts of a 20 mM dithionite solution, in the presence of the following mediators (26 ~M concentration each): dichlorophenolindophenol (220 mV), 1-2 naphthoquinone (145 mV), phenazine methosulfate (80 mV), methylene blue

51 (11 mV), indigo-disulfonate ( - 125 mV), 2-hydroxy-l-4 naphthoquinone ( - 1 4 5 mV), phenosafranine ( - 2 5 5 mV), benzyl viologen ( - 3 5 0 mV), methyl viologen ( - 4 4 0 mV). The redox potential was measured with an Ag/AgCI-KCI (3M) electrode. In the text, all values are given with respect to the standard hydrogen electrode. Stable potentials were achieved in a few minutes, and samples were anaerobically transferred into calibrated EPR tubes which were rapidly frozen. EPR spectra were recorded on a Bruker ESP 300 spectrometer equipped with the ESP 1620 data processing unit. The samples were cooled with an Air Products gas-flow system. The temperature was measured with a calibrated thermocouple (chromel vs. Au/ 0.07% Fe) placed in an EPR tube partially filled with water. For spin quantitations, the second integral value of the spectrum was compared to that given by a CuSP 4 standard recorded at the same temperature. Spin-lattice relaxation time experiments were carried out on the 3Fe and nickel signals given by a fully oxidized sample (160/.tM concentration), and on the nickel signal given by a sample poised at - 2 0 0 mV (110 /zM concentration). Relaxation times T~ were measured by the continuous wave saturation method between 10 K and 30 K for the 3Fe center, and between 10 K and 50 K for the nickel center. We assumed that the saturated spectra could be approximated by the convolution product between an unsaturated spectrum and saturated Lorentzian spin packets [8]. Saturation curves, representing the variations of the signal amplitude as a fonction of the component BI of the microwave fields, were numerically computed for different half-widths ABp of the spin packets. By comparison with the experimental saturation curves, we derived the two parameters ABp and Bt/2, which are related to T 1 by Ref. 8: ABp TI

~(BI/2)

2

For temperatures higher than 30 K, the spin-lattice relaxation time of the 3Fe center was deduced from the broadening of the EPR spectrum. T 1 was then determined by comparison of the experimentally observed broadening with that simulated by numerical convolution of the low-temperature spectrum with broadened Lorentzian packets [9]. Results

At low temperature, the isolated enzyme gives a nearly isotropic signal at g -- 2.02, with a broad bump on the high-field side (Fig. la). This bump is less easily saturated than the rest of the signal, suggesting that the spectrum is in fact composite. This spectrum is identical to that published previously [2], and is re-

/ f

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b

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326

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/ I

330

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Magnetic Field/roT Fig. I. EPR spectra of the 3Fe centers from as isolated hydrogenases from (a) 1). culgaris Miyazaki,(b) D. gigas, and (c) D. fructosororans. Experimental conditions: Temperature, 20 K; microwavefrequency, 9.305 GHz; microwave power, 0.1 mW; modulation frequency, 100 kHz; modulation amplitude, 0.1 mT.

markably similar to the 3Fe signal observed in the isolated enzyme from D. gigas (Fig. lb and Ref. 10). Numerical integration of the signal yields 1 + 0.1 spin per molecule. Another signal can be detected at lower fields, which is more readily observed at high-temperature where it is not saturated and where the 3Fe signal has disappeared due to relaxation broadening (Fig. 2a). Two main components can be distinguished according to their g-values at 2.316, 2.235, 2.015 and 2.33, 2.158, 2.015. These are identical to the signals associated with the Ni-A and Ni-B species in D. gigas hydrogenase, respectively [10,11]. Numerical integration of the whole spectrum at 120 K shows that it accounts for 0.6 + 0.1 spin per molecule. Progressive reduction of the enzyme by sodium dithionite leads to the successive disappearance of the 3Fe, Ni-B and Ni-A signals (Figs. 2, 3). Fitting the variations of the 3Fe signal amplitude to a simple Nernst equation yields midpoint potential of - 7 0 + 10 inV. However, the departure of the experimental data from the theoretical law (Fig. 3) suggests some heterogeneity in the redox properties of the centers. Such redox heterogeneity has already been observed for 3Fe centers in other proteins [12]. Simultaneously to the disappearance of the g = 2.02 signal, a broad signal develops in the g = 12 region (data not shown). In D.

,oo[

52 2.4 I

2.2 I

2.0

I

I

I

L

i

i

"V/'"

-500

/"

-200

100

Fig. 3. Normalized amplitude of the EPR signals given by the Ni-A, Ni-B and oxidized 3Fe centers as a function of the applied potential. Amplitudes of the Ni-A and Ni-B signals were measured at gyA = 2.235 and gya = 2.158 respectively. All measurements were carried out at 15 K in non saturating conditions, Theoretical lines represent Nernst curves with the following midpoint potentials: 3Fe center, - 7 0 mV; Ni-B, -230 mV; Ni-A, -310 inV. The arrows indicate the redox potentials of the two samples that were studied in the T1 measurements. I

!

I

I

290

I

I

310

Magnetic

I

330

Field/aT

Fig. 2. Representatwe EPR spectra displayed by the nickel center during the redox titration of D. vulgaris Miyazaki hydrogenase. Spectrum (d) was recorded after light irradiation at 30 K of the sample giving the spectrum represented In Fig. 4a. After subsequent warming above 200 K, the same sample in the dark gave spectrum (e). Experimental conditions: Temperature, 100 K; microwave power, 10 roW; microwave frequency 9.305 GHz; modulation amplitude, 1 roT.

gigas hydrogenase, a similar signal was also observed, and was attributed to a S = 2 state of the reduced 3Fe center [13]. I

I

I

I

a

I

.~~A._

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I

J

The redox behaviour of the Ni-A and Ni-B signals is well described by Nernstian plots, with midpoint potentials of - 3 1 0 + 10 mV and - 2 3 0 + 10 mV respectively (Fig. 3). This wide difference of redox potential enables the observation of a nearly pure Ni-A signal at - 2 9 5 mV (Fig. 2b). When this signal is compared to that given by the fully oxidized enzyme, it appears that a new feature exists at g = 2.265, suggesting that the Ni-B signal is composite. For potentials more negative than approx. - 3 7 0 mV, new signals become apparent. A first type can be detected at low temperature and low microwave power I

--~. . . . . .

!

I

--_-,-~°mw

__. 50 K

10 K

IOmT, I

2.6

I

I

I

4

I

2.2

I

2.0

I

I

I

1.6

g - sc a le

Fig. 4. Microwavepower dependenceof the signalgiven by a sample of D. vulgaris Miyazakihydrogenasepoised at - 380 mV. Spectrum(a) was recorded at ambientlight.Other conditionsas in Fig.2.

53

100 . . . . . . o-.e.~---e . . . . . . . . . . . . . . . . . . a

l

50 !

2OO

i

3 0 Magnetic

400 Field/roT

Fig. 5. EPR-spectrum given by a sample of 13. culgaris Miyazaki hydrogenasepoised at -420 inV. Experimentalconditions:temperature 10 K; microwavepower 10 roW.Other conditionsas in Fig.2.

(Fig. 4b), but is more readily observed above 40 K (Fig. 4a). It comprises a major component with g-values at 2.196, 2.144 and a minor component at 2.30, 2.117, 2.049. These signals are very similar to those given by the Ni-C and the light-induced Ni-C* species observed in D. gigas hydrogenase, respectively [15]. Light irradiation at 30 K results in an increase of the Ni-C* signal and a concomitant decrease of the Ni-C one, which is illustrated in Fig. 4a and Fig. 2d. We have checked that the full Ni-C species can be recovered by warming the sample above 200 K and by recording the spectrum in the dark (Fig. 2e). Both signals can be observed only between - 3 6 0 mV and - 4 3 0 mV, and reach a maximum around - 4 0 0 mV. At low temperature, they are easily saturated and a complex spectrum with features at 2.21, 2.10, 1.84, 1.70 becomes visible at high microwave power (Fig. 4). This spectrum appears for potentials lower than - 3 0 0 mV, and is well developed at - 4 2 0 mV (Fig. 5). A similar signal was also observed in D. gigas hydrogenase, and was attributed to magnetic interactions between the nickel center and one or two [4Fe4S] + clusters [13,14]. In order to better characterize the EPR signal of the oxidized 3Fe center, we have carried out a detailed study of its temperature dependence. In Fig. 6 the variations of the integrated intensity between 10 K and 55 K are represented. A significant departure from the Curie law is observed for temperatures higher than 30 K. This reflects the existence of low-lying excited levels, as in typical 3Fe centers [15]. For temperatures higher than about 30 K, the spectrum becomes relaxation broadened. Detailed measurements reveal that T t is about two orders of magnitude longer in the 3Fe center of D. vulgaris Miyazaki hydrogenase than in the 3Fe centers of Azotobacter vinelandii FdI and D. gigas FdlI [16] (Fig. 7). Actually, in the range 10-60 K, the relaxation rate of the 3Fe center in D. vulgaris Miyazaki hydrogenase is quite similar to that of the [2Fe-2S] + center from Spirulina maxima ferredoxin [17]. This

0

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I

I

10

I

I

30

I

50

T(K)

Fig. 6. Temperature dependence of the integlatedintensity I of the g = 2.02 signal. Spectra were recorded with a non-saturating power rangingfrom 10/zW at 10 K to 1 mW at 55 K.

illustrates the great versatility of the relaxation properties of the 3Fe centers, and also the difficulties one may encounter when trying to identify iron-sulfur centers solely on the basis of their relaxation properties. A detailed comparison of the relaxation behaviour of [2Fe-2S]+I and [4Fe-4S] +] centers had led to the same conclusion [18]. Owing to the wide difference between the redox potentials of the 3Fe center and of the Ni-A species (Fig. 3), it is possible to examine whether the relaxation behaviour of Ni-A is dependent upon the redox state

I

(8.1)

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e

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oml

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20

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Fig. 7. Temperature-dependence of the spin-lattice relaxation rate I / T l for the followingsignals: (o), 3Fe signal at g = 2.02 (sample poised at + 200 mV); (o) and (E3), Ni-A signal at gyA----2.235 (sample poised at +200 mV and -200 mV, respectively).T1 measuremenls were carried out as described in Materials and Methods. The modulationfrequencywas 1.56 kHz. The dashed line recalls the temperature dependenceof 1/T t for the 3Fe center~of A. cinelandii Fd I and D. gigas Fd It ferredoxins,as reported in Ref. 16.

54 of the 3Fe center. We have then determined the spinlattice relaxation time T t of the Ni-A species, by studying the saturation of the gy = 2.24 peak between 10 K and 50 K (see Materials and Methods). Two samples were studied: in the first one the 3Fe centers were fully oxidized, while the other poised at - 2 0 0 mV contained only reduced 3Fe centers (Fig. 3). Within experimental errors, identical values of T t were measured for both samples in the full temperature range (Fig. 7). The same experiment led to a qualitatively similar result at 31 K in D. gigas hydrogenase [22]. Discussion

Although Ni-A and Ni-B type EPR signals were previously reported in as prepared D. culgaris Miyazaki hydrogenase, they were attributed to adventitious nickel [2]. A striking result of our study is the observation of EPR signals very similar to those given by the nickel center of D. gigas hydrogenase in all characteristic redox states of the enzyme. This provides strong evidence that D. vulgaris Miyazaki hydrogenase is a Ni-Fe enzyme, which is structurally and functionally closely related to the D. gigas one. In the following, a detailed comparison of the EPR and redox properties of these enzymes is developed. The EPR spectra given by the oxidized 3Fe centers of D. vulgaris Miyazaki, D. gigas and D. fructosovorans hydrogenases are compared in Fig. 1. Note that a small modulation amplitude was used in order to avoid distortion of narrow spectral features. The first two spectra have essentially the same shape, characterized by a quasi-symmetrical structure and a broad bump on the high-field side. This shape is markedly different from that given by the Ni-Fe enzyme from D. fructosovorans, which is more reminiscent of the asymmetrical lineshape given by the 3Fe centers of Azotobacter vinelandii and D. gigas ferredoxins [19,20]. We have previously proposed a model to describe the magnetic properties of oxidized 3Fe centers, and have applied it to the simulation of EPR spectra given by a variety of proteins [19,20]. According to this model, the lineshape is essentially determined on the one hand by the magnetic parameters of each ferric site (g-tensor, zero-field splitting tensor), and on the other hand by the exchange interactions between the three sites. These exchange parameters and their differences are expected to be very sensitive to the strains exerted by the proteins, so that their variations could account for the wide diversity of spectral shape exhibited by these centers. Likewise, the great similarity between the spectra given by D. vulgaris Miyazaki, D. gigas and also D. desulfuricans [21] hydrogenases indicates very similar environments for these clusters. Two different values, - 35 _+ 10 mV [22] and - 70 _+ 10 mV [10] have been reported for the midpoint poten-

tial of the 3Fe center in D. gigas hydrogenase at pH 7.0. This probably reflects slight differences in the preparation of the samples, and illustrates the great sensitivity of the redox properties of these centers with respect to their environment. Therefore, the value - 70 +_ 10 mV measured in the D. t,ulgaris Miyazaki enzyme strongly supports the conclusion drawn above about the great similarity of the 3Fe sites in those proteins. The amount of nickel detected by EPR in the isolated enzyme (0.6 spin per molecule) is similar to that usually found in other Ni-Fe hydrogenases [10,22]. From the comparison of the nickel signals at 220 mV (Fig. 2a) and - 2 9 5 mV (Fig. 2b), the intensity ratio of the Ni-B to Ni-A signals can be evaluated to 0.3. This value is intermediate between those observed in D. fructosocorans [7] and D. gigas [10] hydrogenases. In the latter protein, this ratio varies somewhat depending on the protein preparation [13], and can also be modified by reduction and reoxidation [10,11]. Those Ni-A and Ni-B species are currently considered as Ni(III)-centers corresponding to oxygenated (or unready) and deoxygenated (or ready) forms of the enzyme [13,14], respectively. Interestingly, the midpoint potential of the Ni-A species, - 3 1 0 +_ 10 mV at pH 7.7 is notably more negative than the value - 150 + 10 mV at pH 7 reported for D. gigas hydrogenase, even if a - 6 0 m V / p H unit variation is taken into consideration [22]. The exact potential of the Ni-B species is unknown in D. gigas, but it was estimated slightly more negative than - 1 5 0 mV at pH 7 [13]. This is presumably more positive than the value - 2 3 0 _ 10 mV we measured in D. vulgaris Miyazaki hydrogenase at pH 7.7. The Ni-C type signal is also virtually identical to that given by other Ni-Fe hydrogenases [7,13]. This signal is currently attributed to a nickel-hydride species which might be an intermediate in the catalytic cycle. It is only observed in an active form of the enzyme, obtained by reduction with hydrogen or other strong reductants [7,13]. In Ni-Fe enzymes, this Ni-C signal is converted to the so-called Ni-C* one upon illumination at low temperature [14]. A noteworthy particularity of D. vulgaris Miyazaki hydrogenase is to give Ni-C and Ni-C*type signals which can simultaneously be observed in an EPR spectrum recorded at ambient light. Thus, although the interconversion mechanism of those two species seems to be very similar to that reported for D. gigas hydrogenase, the Ni-C species appears to be more photosensitive in D. vulgaris hydrogenase. The range of potentials at which the Ni-C signal can be observed appears to be narrower and more negatively shifted in D. vulgaris Miyazaki hydrogenase at pH 7.7 than in D. gigas at pH 7 [13]. However, this range is strongly pH-dependent in the latter protein [14], so that a more meaningful comparison must await further studies.

55 The origin of the complex spectrum with features at g = 2.21, 2.10 and broad components at higher fields, which appears at potentials more negative than - 3 0 0 mV in D. gigas hydrogenase, has been a subject of discussions [13,14]. The results of recent EPR and Mossbauer experiments provide strong arguments in favour of a spectrum reflecting magnetic interactions between the nickel center and one or two [4Fe4S] + clusters [13]. The shape of such a spectrum is expected to be very sensitive to the distances and the relative orientation between the interacting magnetic centers. Therefore, the close similarity, of the spectra given by D. vulgaris Miyazald and D. gigas hydrogenases implies a quasi identical spatial organization of the interacting centers in these enzymes. The range of potentials at which this spectrum can be observed appears also shifted towards more negative potentials in D. vulgaris Miyazaki hydrogenase by comparison with D. #gas [13,14]. The relaxation time of the oxidized 3Fe center is muct~' shorter than that of the Ni-A species (Fig. 7). In these circumstances, the relaxation rate of the latter might be enhanced by dipolar coupling with the former. Theoretical expressions describing this enhancement are available in the case of point centers characterized by isotropic g-tensors, for a particular value of the angle 0 between the intercenter vector and the applied magnetic field [23]. Among the different terms involved in those expressions, the dominant one is expected to be the following: 1

TI

I

t134(1 - (3 cos 2 0 ) ) "S(S + 1)g'fg i"

T,f

6h2R 6

1+ Aw'T2f

T°s +

(I)

In the second-hand side, Tt°s is the spin-lattice relaxation time of the slow relaxing species in the absence of dipolar interaction, and the second term represents the relaxation enhancement due to dipolar coupling with the fast relaxing species characterized by the spin-spin relaxation time T2f'g s and gf are the g-values, R is the intercenter distance and Ato the difference between the resonance angular frequencies of the two centers. We have used Eqn. 1 to evaluate the relaxation enhancement of Ni-A due to dipolar coupling with the oxidized 3Fe center, by making the following simplifications: (i) both centers were approximated by point centers; (ii) the orientation factor was replaced by its average value 0.8 [23] gf was taken equal to 2.02 and gs to the g value gyg = 2.24 at which T~ measurements were performed. By taking into consideration all numerical factors, we obtain the following expression: 1

r,, =

1

412

+ R"T,----S

(2)

where R is given in ,g,. In the relaxation broadening regime, the spin-spin and spin-lattice relaxation times are approximately equal [18]. Actually, it is the spin-spin relaxation time that is measured in relaxation broadening studies, so that the relevant values of T2f are those plotted in Fig. 7 for temperatures higher than 30 K In the reduced state, the 3Fe center gives a broad EPR signal in the g--- i2 region, which has been attributed to a S---2 manifold [13]. Careful measurements of its linewidth did not reveal any relaxation broadening between 20 K and 50 K. This result, together with Eqn. 1, can be used to estimate an upper bound for the relaxation enhancement given by the reduced 3Fe center. It turns out that, at 50 K this enhancement is expected to be at least an order of magnitude smaller than that given by the oxidized 3Fe center. Therefore, the lack of any appreciable relaxation enhancement of the nickel center in the fully oxidized state implies that the corrective term in Eqn. 2 is less than the uncertainty in the measurement of T~s, which is about + 20%. Using in Eqn. 2 the T~s and T2f values measured at 50 K, we finally conclude that the nickel and 3Fe centers are separated by more than 10A. All the results presented in this study concern samples prepared from enzyme solutions that were rapidly frozen after the purification and concentration steps, and stored in liquid nitrogen during a few months. Generally speaking, the enzyme appeared very sensitive to oxidative degradation, and it was necessary to maintain strict anaerobic conditions during the redox titration. Some samples prepared from lyophilized powder or from solutions subjected to several thawing/freezing cycles have also been studied, and have been found to exhibit some distinct redox and spectral properties. In particular, the purity index Aaoo/h28 o was lower, and the characteristics of the oxidized 3Fe center were modified; content as low as 0.5 spin per molecule, slightly different shape of the EPR spectrum in the high-field region, marked departure from the Nernst equation. On the other hand, the redox and spectral properties of the nickel center appeared well conserved in the different preparations. All these observations lead to the suggestion that the 3Fe center is located in a protein part that can undergo important conformational changes. Our spin-relaxation experiments have shown that it is separated by more than 10 from the nickel center, a result consistent with the absence of any magnetic splitting or broadening of the nickel spectrum in the fully oxidized state. Owing to its high redox potential, the 3Fe center can hardly be involved as a redox intermediate in the reversible electron exchange between the active site of hydrogenase and the low-potential heroes of its redox partner, cytochrome c 3. Furthermore, a recent study of the effect of several transition metals on the activity of D. gigas

56

hydrogenase has led to the conclusion that the 3Fe center is not essential for H2-uptake activity with methyl viologen [24]. More studies are required in order to elucidate the role of this center. Conclusion

The results presented in this study demonstrate conclusively that D. vulgaris Miyazaki hydrogenase is a Ni-Fe enzyme closely related to that from D. gigas. This similarity concerns the structure of the different metal centers, their spatial organization in the protein, and also their general redox behaviour. This conclusion is in agreement with the high level of homology between the two enzymes that was deduced from a recent molecular biology study [5]. However, some significant differences are observed, like the midpoint potentials of the different nickel species and possibly of the 4Fe4S clusters, and also the photosensitivity of the Ni-C form of the nickel center. This probably reflects the subtle influence exerted by these centers' environments on their physicochemical properties. Acknowledgements

We acknowledge Dr. E.C. Hatchikian for providing samples of D. gigas and D. fructosovorans hydrogenases, and Dr. J.C. Germanique for performing the plasma emission spectroscopy experiments. References 1 Adams, M.W.W. 0990) Biochim. Biophys. Acta 1020, 115-145. 2 Yagi, T., Kimura, K. and Inokuchi, N. (1985) J. Biochem. 97, 181-187. 3 Yagi, T., Kimura, K., Daidoji, H., Sakai, F., Tamura, S. and Inokuchi, N. (1976) J. Biochem. 79, 661-671. 4 Hatehikian, C.E., Bruschi, M. and Legall, J. (1978) Biochem. Biophys. Res. Commun. 82, 451-461.

5 Deckers, N.M., Wilson, F.R. and Voordouw, G. (1990) J. Gen. Microb. 136, 2021-2028. 6 Higuchi, Y., Yasuoka, N., Kakudo, M., Katsube, Y., Yagi, T. and Inokuchi, H. (1987) J. Biol. Chem 262, 2823-2825. 7 Hatchikian, C.E., Traore, A.S., Fernandez, V.M. and Cammack, R. (1990) Eur. J. Biochem. 187, 635-643. 8 Castner, T.G. (1959) Phys. Rev. 115, 1506-1515. 9 Bertrand, P., Roger, G. and Gayda, J.P. (1980) J. Magn. Res. 40, 539-549. 10 Teixeira, M., Mourn, I., Xavier, A.V., Huynh, B.H., Dervartanian, D.V., Pepck, H.D., Legall, J. and Mourn, J.J.G. (1985) J. Biol. Chem. 260, 8942-8950. 11 Fernandez, V.M., Hatchikian, E.C., Patil, D.S. and Cammack, R. (1986) Biochim. Biophys. Acta 883, 145-154. 12 Guigliarelli, B., Bertrand, P., More, C., Papavassiliou, P., Hatchikian, E.C. and Gayda, J.P. (1985) Biochim. Biophys. Acta, 810, 319-324. 13 Teixeira, M., Moura, I., Xavier, A.V., Moura, J.J.G., Legail, J., Dervartanian, D.V., Peck, H.D. and Huynh, B.H. (1989) J. Biol. Chem. 264, 16435-16450. 14 Cammack, R., Patil, D.S., Hatchikian, E.C. and Fernandez, V.M. (1987) Biochim. Biophys. Acta 912, 98-109. 15 Gayda, J.P., Bertrand, P., Guigliarelli, B. and Meyer, J. (1983) J. Chem. Phys. 79, 5732-5733. 16 Bertrand, P., Guigliarelli, B., Meyer, J. and Gayda, J.P. (1984) Biochimie, 66, 77-79. 17 Gayda, J.P., Bertrand, P., Deville, A., More, C., Roger, G., Gibson, J.F. and Cammack, R. (1979) Biochim. Biophys. Acta 581, 15-26. 18 Bertrand, P., Gayda, J.P. and Rao, K.K. (1982) J. Chem. Phys. 76, 4715-4719. 19 Guigliarelli, B., Gayda, J.P., Bertrand, P. and More, C., (1986) Biochim, Biophys. Acta 871, 149-155. 20 Guigliarelli, B.~ More, C., Bertrand, P. and Gayda, LP. (1986) J. Chem. Phys. 85, 2774-2778. 21 Kruger, H.J, Huynh, B.H., Ljungdahi, P.O., Xavier, A.V., Dervartanian, D.V., Moura, I., Peck, H.D., Teixeira, M., Mourn, J.J.G. and Legall, J. (1982) J. Biol. Chem. 257, 14620-14623. 22 Cammack, R., Patil, D., Aguirre, R. and Hatehikian, E.C. (1982) FEBS Lett. 142, 289-292. 23 Brudwig, G.W., Blair, D.F. and Chan, S.I. (1984) J. Biol. Chem. 259, 11001-11009. 24 Fernandez, V.M., Rua, M.L., Reyes, P., Cammack, R. and Hatchikian, E.C. (1989) Europ. J. Bioehem. 185, 449-454.