Biochimica et Biophysica Acta. 1119(1992) 157-168

157

© 1992 Elsevier Science Publishers B.V. All rightsreserved 0167-4838/92/$~15.00

BBAPRO 34113

Distinct redox behaviour of prosthetic groups in ready and unready hydrogenase from Chromatium vinosum J.M.C.C. Coremans, J.W. van der Zwaan and S.P.J. Albracht E.C. Slater Institute for Biochemical Research and Biotechnological Centre. Unicersityof Amsterdam, Amsterdam (The Netherlands)

(Received 16 September 1991)

Key words: Hydrogenase;Nickel; Iron-sulfurcluster; Redoxpotential;(C. cmosum)

The redox behaviour of the N i ( I I I ) / N i ( H ) transition in hydrogenase from Chromatium vinosum is described and compared with the redox behaviour of the nickel ion in the F4zo-nonreducing hydrogenase from Methanobacterium thermoautotrophicum. Analogous to the situation in the oxidised hydrogenase of Desulfocibrio gigas (Fernandez, V.M., Hatchikian, E.C., Patil, D.S. and Cammack, R. (1986) Biochim. Biophys. Acta 883, 145-154), the C. vinosum enzyme can also exist in two forms: the 'unready' form (EPR characteristics of Ni(lll): gx.y.z - 2.32, 2.24, 2.01) and the 'ready' form (EPR characteristics Ni(lll): gx~,z= 2.34, 2.16, 2.01). Like in the oxidised e~z~iJie of M. thermoautotrophicum the N i ( I H ) / N i ( H ) transition for the unready form titrated z~mpietely reversible (both at pH 6.0 and pH 8.0). In contrast, the reversibility of the N i ( H I ) / N i ( l l ) transition in the ready enzyme was strongly dependent on pH and temperature. At pH 6.0 and 2°C reduction of Ni(lll) in ready enzyme was completely irreversible, whereas at pH 8.0 and 30°C Ni(lll) in both ready and unready enzyme titrated with E~ = - 1 1 5 mV (n = 1). Hampered redox equilibration between the ready enzyme and the mediating dyes is interpreted in terms of an obstruction of the electron transfer from nickel at the active site to the artificial electron acceptors in solution. The origin of this obstruction might be related to possible changes in the protein structure induced by the activation process. The E~-value of the Ni(HI)/Ni(II) equilibrium was pH sensitive ( - 6 0 m V / , ~ p H ) indicating that reduction of nickel is coupled to a protonation. A similar pH-dependence was observed for the titration of the spin-spin interaction of Ni(lll) and a special form of the [3Fe-4S] + cluster (E o = + 150 mV, pH 8.0, 30°C). Redox equilibragion of this coupling was extremely sensitive to pH and temperature. The uncoupled [3Fe-4S] ÷ cluster titrated pH-independently with E~ = - 1 0 mV (pH 8.0, 30°C).

Introduction The enzyme hydrogenase (hydrogen: (acceptor) oxidoreductase catalyzes the reaction: H 2 ~ 2 H + + 2e-. Hydrogenase from the purple-sulphur bacterium Chromatium vinosum (DSM i85) is a water-soluble protein that consists of two subunits of about 60 and 30 kDa [1]. Per 90 kDa protein 0.8-1.1 gatom of nickel is found in addition to 8.8-11.9 gatom of Fe and 8.6-9.2 gatom of acid-labile sulphur [1]. Nickel is the hydro-

Abbreviations: DCIP, 2,6-dichlorophenolindophenol;TMPD, 2,3,5,6tetramethyl-p-phenylenediaminc dihydrochloride; PMS phenazine methosulphate; DTF, dithiothreitol. Correspondence: S.P.J. Albracht, E.C. Slater Institute for Biochemical Research, Universityof Amsterdam, Plantage Muidergracht 12, 1018 TV Amsterdam,The Netherlands

gen-binding site and, like in other hydrogenases [2], iron and acid-labile sulphur are arranged in Fe-S clusters [3,4]. In the enzyme, as isolated, the valence state of the nickel ion is 3 + and with EPR spectroscopy two rhombic spectra of Ni(lll) can be observed [5,6]. in hydrogenase from the sulphate-reducing bacterium Desulfot'ibrio gigas similar Ni(lll) spectra are observed. In 1986 Fernandez et al. [7] demonstrated with the enzyme from D. gigas that these two forms of Ni(IlI) represented two forms of oxidised hydrogenase molecules that differed in their ability to active H 2. One form, (EPR characteristics of Ni(lll): gx.r.z = 2.31, 2.23, 2.02) was unable to active H2; it was 'unready' to oxidize H2. The other form (EPR characteristics of Ni(lll): gx.r,~ = 2.33, 2.16, 2.0) could 'readily' react with H 2. The conformations of nickel in 'ready' and 'unready' hydrogenase molecules in D. gigas were indicated as Ni-B and Ni-A, respectively [7,8]. In the past

158 the Ni ion in the 'ready' form of the C chmsum enzyme with gx.r.= = 2.34, 2.16, 2.01 and the 'unready form with g.,..,.: = 2.32, 2.24, 2.01 have been referred to as Ni-a and Ni-b, respectively [9]. To avoid confusion in nomenclature, we will now indicate the two nickel forms as: Ni,(lll) for the 'ready' enzyme and Ni~(lll) for the "unready' enzyme. Incubation under H z readily reduces both ready and unready enzyme from C. cinosum. This reduction process is accompanied by various changes in the valence state of the nickel ions. The simplest description is a transformation of Ni(lll) via Ni(ll) to Ni(l) and possibly even to Nil0). When hydrogenase is reduced to the Nil1) state, H , is bound directly to the nickel ion [3,6]. in aerobically isolated preparations of the F4,0-nonreducing hydrogenase from Methanobacterium thermoautotrophicum (strain Marburg) only unready enzyme molecules are found [10]. In agreement with this, the EPR characteristics of Ni(lll) in these molecules (gx.y.==2.30, 2.23, 2.01) closely resemble those of Ni~(llI) in C. t'inosum and D. gigas. In order to gain insight into the working mechanism of hydrogenase from M. thermoautotrophicum we have earlier studied the redox behaviour of nickel in relation to Ha-uptake activity [11]. Anaerobic titration of the Niu(IlI)/Ni,(ll) transition yielded a midpoint potential of - 140 mV at pH 6.0 and 15°C; the nickel ion could not be reduced beyond the Ni(ll) state and no H2-uptake activity was induced. The enzyme could be activated by incubation under H , at pH 6.0 and 45°C and a reversible Ni(ll) Ni(l)o'Ni(0)' equilibrium existed in the active enzyme. (Nickel in active enzyme will be referred to as Niacr) Upon controlled anaerobic reoxidation of active enzyme Ni~(Ill) did not reappear, provided that the temperature was kept low (T ~< 15°C). Even at high redox potentials the enzyme remained in a Ni~t(ll) state and Ha-uptake activity was maintained. Anaerobic oxidation of active enzyme by a large excess of potassium ferricyanide, or the high-potential redox mediator porphyrexide (E~ = + 700 mV), revealed both Ni(lll) forms: Niu(IIl) and Nir(lll). Therefore, like in the D. gigas and C. cinosurn hydrogenase, enzyme from M. thermoautotrophicum can exhibit the Ni,(lll) form in oxidised 'ready' hydrogenase and the Ni,(lll) form in 'unready' hydrogenase. The Ni~(lll) form in M. thermoautotrophicurn, however, was very unstable and rapidly converted into the Ni~(IlI) form, thereby preventing determination of the Ni,(llI)/Ni,(lI) equilibrium. The results of redox titrations of nickel in hydrogenase from M. thermoautotrophicum are summarized in Scheme I. Form these studies we made the assumption that in the ready and active state of the enzyme, the Eli-values of both the Ni(lll)/Ni(lI) transition and the Ni(ll)/Ni(l) transition would be considerably higher than those in the unready enzyme. The presence in C, cinosum hydrogenase of both Niu(lll)

UNREADY

READY

. . . /~ Nrtill~- Nir[rnl ,r-.......................

Eh[mV| -400 .

-200

/

-o

Niulml-~-~ NiIlll |

-200

ACTIVE

-400



N~tnl : ........................ ._, ,

Niatt[lll~ N~(I I "N~t(0 I'.~-- Ni.~(I I NiullJ---~

Scheme !. Redox behaviour of the nickel ion in hydrogenase from

M. thermoautotrophicum in relation to enzyme activity. The drawn boxes enclose the potential regions that have been covered during redox titrations at 15°C (left panel) and 45°C (right panel) [11]. The dashed boxes show the proposed redox equilibria in potential regions which could not be reached under the experimental conditions used.

and Ni,(lll) offered the possibility to verify the assumed difference in midpoint potentials of nickel in the two forms of the enzyme. In this paper we demonstrate that, under some conditions (pH 8.0, 30°C), Nit(Ill) and Ni,(Ill) in hydrogenase from C. t'inosum titrate with identical midpoint potentials. Under other conditions both forms of nickel exhibited quite distinct redox properties, similar to the behaviour observed in the M. thermoautotrophicum enzyme. These findings led us to propose an alternative explanation for the redox behaviour of nickel in these hydrogenases. Materials and Methods

Purification procedure In the first publication on hydrogenase from (7. cinosum by this laboratory in 1981 [12], C. t,inosum strain D .was used. Purification of the H2-methylene blue reductase activity of this strain resulted in a hydrogenas¢ preparation which apparently consisted of a single polypeptide chain with a molecular weight of 61-63 kDa, as judged from SDS gelelectrophoresis, gelfiltration and equilibrium-sedimentation studies. Since 1982, however, we have worked with C. t'inosum strain DSM 185. The enzyme purified from this strain, using the method described below (now routinely assayed by the H2-benzyl viologen assay), consistently shows two subunits of about 30 and 60 kDa. Chemical analyses of these hydrogenase preparations gave values for Ni, Fe and acid-labile sulphur within the ranges determined earlier [1]. The EPR spectra in the Fe-S region (type of spectra, relative intensities, spin-coupling) of the oxidised hydrogenase preparations from C. rinosum strain DSM 185 are indistinguishable from the initial D-strain preparations.

159 C. t'inosum (strain DSM 185) was grown in a 700 1 batch culture in a medium essentially as described by Hendley [5,12,13]. Cells were harvested by means of a Sharpies MP4 continuous-flow centrifuge (Sharpies Centrifuge, Chamberley, U.K.). An acetone-extraction procedure (3 I acetone o f . - 20°C per 500 g cells wet weight) was used to remove lipids and photosynthetic pigments [14]. These hydrophobic substances appeared in the supernatant after centrifugation of the cellacetone suspension (2 rain, 1000xg). Hydrogenase resided in the pellet. This pellet was subjected to acetone treatment as frequently as necessary to extract all pigment. Acetone was finally removed from the pellet by evaporation under a stream of air. The resulting powder was suspended in 20 mM potassium phosphate buffer (pH 7.5) and stirred for 15 min at room temperature. Non-soluble material was removed by low-speed centrifugation (10 min, 1000 x g) and a subsequent high-speed centrifugation (45 min, 35 000 × g). About 75% of the hydrogenase activity appeared in the supernatant. The pellet was extracted three more times (15 min) with 20 mM potassium phosphate buffer (pH 7.5). From this point on all steps in the purification were performed at 4°C (aerobic conditions). The combined supernatants from 1-1.5 kg of cells (wet weight) were applied to a Fraktogel TSK-DEAE-650(M) (Merck) column (2.5 × 40 cm) which was equilibrated with 50 mM Tris-HCl buffer (pH 7.4). Hydrogenase was eluted by a linear NaCI gradient (0-0.4 M) in 50 mM Tris-HCl buffer (pH 7.4) (1000 ml; 0.75 ml/min). Two distinct peaks of H 2-production activity were found at 0.12 M and 0.27 M NaCl. Upon further purification of the Hz-production activity eluting at 0.12 M NaCI, an enzyme preparation was obtained with the following characteristics: (i) main subunits 47 and 24 kDa; (ii) almost no H z-uptake activity; (iii) a trypsin-sensitive NADH-K3Fe(CN) 6 reductase activity, (iv) no, or only very small, E P R signals of Ni(llI) in the oxidised state; and (v) a radical signal and an EPR signal with gx,r.z = 1.895, 1.935, 2.035 in the NADH-reduced enzyme (J.W. van der Zwaan, unpublished data). The enzyme eluting at 0.27 M NaCi predominantly exhibited H2-uptake

activity and this enzyme was used for further purification and study. The solution was concentrated by ultrafiltration (Amicon PM 30 filter) after which KCI was added to a final concentration of 2 M. The solution was then applied to a phenyi-Sepharose CL-4B (Pharmacia) column (2.5 x 40 cm) equilibrated with 50 mM Tris-HCI, 2 M KC! (pH 8.0). in this case hydrogenase was eluted by a linear gradient of 1 M KCI, 0% ethylene glycol to 0 M KCI, 80% ethylene glycol (1000 ml; 0.6 ml/min). The enzyme eluted at 0.12 M KCI, 70% ethylene glycol. Hydrogenase fractions were pooled and concentrated by ultrafiltration (Amicon PM 30 filter) to a volume of less than 10 ml. Subsequently the concentrated phenyl-Sepharose fraction was diluted 4 times in 50 mM Tris-HC! buffer (pH 7.4) in order to lower the ionic strength of the solution to enable binding of the enzyme to a second Fraktogel TSK-DEAE-650(M) column. This column was treated exactly like the first TSK column. After concentration of the pooled fractions with H2-uptake activity to + 10 ml (uitrafiitration Amicon PM 30) 0.1 mg chymotrypsin per mg protein was added and the solution was incubated for 2 h at room temperature. Hereafter lowmolecular-weight material was removed by washing with 50 mM Tris-HCl buffer (pH 8.0) in an ultrafiltration cell (Amicon PM 30 filter) and the protease treatment was repeated with a new addition of chymotrypsin, this time for 2 h at 37°C. Thereafter the hydrogenase was further purified by gel filtration on an Ultrogel ACA-44 (LKB) column (5.0 × 160 cm), equilibrated with 50 mM Tris-HCI buffer (pH 8.0: elution at 1.6 ml/min). With this large scale purification procedure we were able to obtain preparations of 85% purity with average Hz-uptake specific activities of 72 U / m g in a 5-20% overall yield (table I). Hydrogenase activity was measured essentially as described in Ref. 11. H z-consumption of the enzyme preparation subjected to a redox titration was measured in 50 mM Tris-HCl buffer, 2.2 M KCI (pH 8.0) at 30°C. The T-dependence and the pH-dependence of H z-uptake activity were both investigated in 50 mM potassium phosphate buffers. In all

TABLE 1

Purification of the Hz-uptake hydrogenase from C cinosum used for the ¢rperiments described in this paper ! U = 1 /zmol/min. The purification factor is calculated from the Hz-uptake activity of activated enzyme. Step

Protein (mg)

Potassium phosphate extract Fraktogel TSK-DEAE-650(M) Phenyl-Sepharose CL4B Fraktogel TSK-DEAE-650(M ) Ultrogel ACA-44

44392 3 333 361 ! 72 !7

Specific activity (U/mg) H z-uptake

H 2-production

0.6 0.9 5.3 11.2 72.3

0.1 0.1 0.3 0.3 2.0

Purification factor I

i.5 8.8 18.7 120.5

Yield (%) 1011 il 7 7 5

160 cases 100/~M H 2, 4.2 mM benzyl viologen and 80 mM glucose plus glucose oxidase (16 U / m l ) were present in the activity assay.

Redox titrations Redox titrations were carried out in an apparatus similar to that of Dutton [15], under a flow of watersaturated gas. The redox potential was measured with a platinum electrode versus a standard calomel electrode. All potentials are quoted relative to the standard hydrogen electrode, taking into account the effect of temperature in the Nernst equation. Correction for the temperature dependence of the reference electrode was made according to Ref. 16. Two sets of redox titration experiments were performed which differed in the presence of H 2 in the gas phase and the number and concentration of redox mediators in the mediator cocktail: (1) Argon was used as the sparging gas and the potential was set by addition of aliquots of anaerobic solutions of sodium dithionite (100 mM), potassium ferricyanide (250 mM) or DCIP (100 mM). In these titrations the following mediators (mediator cocktail 1; final concentration 50/~M) were added to the enzyme solution: 2,3,5,6-tetramethyl-p-phenylenediamine dihydrochloride (TMPD; E 0' -- +275 mV), 2,6-dichlorophenol-indophenol (DCIP; E~ = + 230 mV), 1,2-naphthoquinone-4-sulphonic acid (E~ = +215 mV), pheriazinc methosulphate (PMS; E~ = + 80 mV), 1,4-naphthoquinone ( E ~ = +36 mV), methylene blue (E~--+ 11 mV), duroquinone (E~ = - 5 / 3 5 mV), pyocyanin perchlorate (E~ = - 34 mV), indigodisulphonate (E~ --- 1 2 5 mV), 2-hydroxy-l,4-naphthoquinone ( E ~ = -139/-152 mV), lapachol ( E ~ = - 1 7 9 mV), anthraquinone-2-sulphonic acid ( E ~ - - - 2 2 5 mV), safranin (E~ --- - 289 mV), neutral red (E~ = - 329 mV), benzyl viologen ( E ~ = - 3 5 8 mV) and methyl viologen (E~ = - 449 mV). (2) A mixture of He and H2, produced by a homebuilt gas mixer, was used as the sparging gas. In these titrations an anaerobic ferricyanide solution (250 mM) was used as oxidant and sodium dithionite was used to adjust the potential to a 10% reduction degree of Ni(lll). The low-potential redox mediators were absent and the concentration of several high-potential mediators was increased in order to create a redox buffer at the pre-set potential. For redox titrations at pH 6.0 and 2°C, TMPD, DCIP, 1,2-naphthoquinone-4-sulphonic acid and PMS were present at a concentration of 250 /~M, whereas 1,4-naphthoquinone, methylene blue, duroquinone, pyocyanine and indigodisulphonate were present at a final concentration of 50 /~M (mediator cocktail 2). For redox titrations at pH 8.0 and 2°C (mediator cocktail 3) the concentrations of pyocyanin, duroquinone and indigodisulphonate were raised to 250 ~M. The final concentrations of TMPD, DCIP,

1,2-naphthoquinone-4-sulphonic acid, PMS, 1A-naphthoquinone, methylene blue, 2-hydroxy-l,4-naphthoquinone, lapachol and anthraquinone-2-sulphonic acid were maintained at 50/~M. For redox titrations at pH 8.0 the enzyme was dissolved in 100 mM Tris-HCl buffer. For titrations at pH 7.0, 100 mM Mops buffer was chosen and the redox titrations at pH 6.0 were conducted in 100 mM Mes buffer. After equilibration of hydrogenase at the appropriate potential, samples were withdrawn with a gas-tight syringe through a subaseal stopper and rapidly frozen in E P R tubes in a cold isopentane (133 K) bath. The E P R tubes were sealed with latex tubing and thoroughly flushed with the equilibrating gas mixture of the redox cuvet before filling. E P R measurements were performed on a Varian E-9 spectrometer equipped with a home-built He-flow cryostat [17]. Spectra were stored on a personal computer using a 12 bits A / D converter. In order to quantify Ni(III) signal intensities recorded with a modulation amplitude of 1.25 mT, simulated lineshapes of Nir(III) (gx.y,z = 2.337, 2.1637, 2.01, and widths ( x , y , z ) = 1.45, 1.02, 1.3 mT) and Niu(IIl) (gx,y,z=2.321, 2.2411, 2.015, and widths ( x , y , z ) = 1.55, 1.29, 1.3 mT), optimized with regard to the positions and the widths at half-height of the gx and gy lines, were fitted to an experimental spectrum of oxidised hydrogenase recorded at T > 40 K. The spin. concentrations were then determined by double integration of the simulated spectra [18,19]. A solution of 10 mM C u S O 4 - 5 H 2 0 in 2 M NaCIO 4 and 10 mM HCI was used as concentration standard. Once the double-integral value of a standard Ni(III) spectrum was known, the amplitude of its gy-line served as a direct measure for the total spin concentration in any experimental spectrum. The concentration of the [3Fe4S] + cluster was determined by direct double integration of its E P R spectrum recorded at 13.5 K in a sample with a [3Fe-4S] + signal of maximal intensity. The amplitude of the g = 2.02 line in the spectra then served as measure for the concentration of the [3Fe4S] ÷ cluster obtained at various redox potentials. Under certain conditions spin-spin interaction of Ni(IIl) and a modified [3Fe-4S] + cluster with different g-values, possibly a [4Fe-4S] 3+ cluster [9], was observed. The signal of the [3Fe-4S] ÷ cluster was, however, never lost completely. Due to the complexity of the resulting signals [9] we have not tried to quantify their intensity. In the redox titrations the coupling-signal intensity was expressed as the amplitude of the trough at g = 1.98 in any experimental spectrum recorded at 13.5 K relative to the amplitude in a spectrum with maximal coupling. Results

Redox titrations at 30°C Enzyme as isolated from C. vinosum was subjected to a redox cycle (2 h incubation under H 2 at room

161 2

J

1.0

A

y

z

~0s 4--

.... I ......... 2.4

I ......... 2.3 G

-

I .... , .... 2.= VALUE

2.

I... 'l

..,

....

I . . . . . . . . . =.O G --

I

O.C~

*

*0 2 B

-

-200

=

-100

medj

0

t..9

100

i

200

3oo r l,,,Vl

VALUE

Fig. 1. Effect of redox mediators at pH 8.0 and :30°C on the EPR spectra of oxidised hydrogenase in the nickel region (panel 1) and Fe-S regions (panel 2). (A) In the absence of redox mediators. (B) 90 rain. after addition of mediator cocktail 1 (see Materials and Methods). EPR conditions: microwave fi-equency, 9268.5 MHz; temperalure, 13.5 K; microwave power incident to the cavity, 2.0 mW; modulation amplitude, 1.25 mT. Relative gains of spectra IA, IB, 2A and 2B are consecutively x40, x216, ×1 and x4.3.

B 1.0

l=a ¢.

*=ed

0 °0 z

temperature followed by 21 h H 2 at 4°C, 30 min Ar at 30°C and 30 rain 0 2 at room temperature). In this way a preparation was obtained with almost twice as much Nir(III) as Ni,(III)(Fig. 1A). An additional advantage of this consecutive reductive and oxidative treatment was that the spin-spin coupling of Ni(III) and an Fe-S cluster, assumed to be a [4Fe-4S] 3+ cluster formed out of the [3Fe-4S] + cluster, was abolished. Such a spin coupling is frequently encountered in oxidised enzyme preparations and can only be observed below 20 K [9]. Quantification of the relative contributions of Nir(III) and Niu(III) in these spin-coupled low-temperature spectra is impossible and therefore uncoupled enzyme preparations were preferred for redox titrations. In preparations where the spin-spin interaction of Ni and the Fe-S cluster was abolished, three EPR signals due to independent non-interacting S = ~1 systems of Nir(III), Niu(III) and a [3Fe-4S] + cluster were observed [9]. The uncoupled preparation was used as starting material for redox titrations at oH 8.0 and 30°C. Unfortunately addition of the redox-mediator cocktail to the enzyme under these conditions (pH 8.0, 30°C) resulted in rapid reconstitution of the spin coupling (Fig. 1B). Coupling signals could be observed already within 5 rain after addition of the redox mediators. Thereafter the intensity of the coupling gradually developed to a maximum. As a consequence, the titration curves of the Niu(III) and Nir(III) signals did not resemble simple Nernst-curves (Fig. 2A,B). The coupling, which is responsible for loss of Ni(III) signal intensity at high redox potentials, titrated according to a reversible n = 1 redox process with a midpoint potential of + 150 mV (pH 8.0, 30°C) (Fig. 3B). From the

O0

10 ¸

-200

-100

0

160

2(}0

300 Eh[mV]

C

._=z

+_..

*med

T

~05

V

~Oll'

-200

-100

I00

200

*02

300Eh[mV]

Fig. 2. Effect of the redox potential at pH 8.0 and 30°C, in the presence of redox mediator cocktail 1 (Materials and Methods), on the Ni(IIl) signal intensity at 42 K. The enzyme was poised at various redox potentials by addition of small aliquots of anaerobic solutions of sodium dithionite (100 mM), DCIP (100 raM) or K3Fe(CN)6 (250 mm). Ni(lII) signal intensity is plotted as a fraction of maximally EPR-detectable Ni(llI) intensity as present in the oxidised aerobic preparation in the absence of redox mediators. EPR conditions for recording of Ni(Ill) signals: microwave frequency, 9.27 GHz; temperature, 42 K; microwave power, 2.0 roW; modulation amplitude, 1.25 mT. Ar was used as sparging gas. Before the start of the experiment the redox curet was flushed with Ar for 1.5 h. For each titration curve closed symbols reflect data points obtained during the reductive titration and open symbols reflect data points obtained during the subsequent oxidative titration. (A) Ni,(Ill) signal intensity (11, D). (B) Ni,(lIl) signal intensity (e,o). (C) Total Ni(III) intensity ( = Niu(III)+ Nir(IlI))( v , v ) as a function of the applied redox potential. For E h < 20 mV the solid line is a simulation of an n = 1 redox process with a midpoint potential of -115 inV. After admittance of air to the titration vessel, El, 0 , v represent Niu(lll), Nir(III) and total Ni(IlI) intensity, respectively.

162

B

10

0 -02@

J~

g05

O0

° 02

.~.'6

16oz6o300EhtmVl

O~

IO.Q -200 -100- -0 - 100 200 300FanlmI V

Fig. 3. Effect of the redox potential on the [3Fe-4S] + signal and the coupling signal at oH 8.0 and 30°C. (A) [3Fe-4S] * signal intensity ( • , C , ) is plotted as a fraction of maximally detected [3Fe-4S] ÷ intensity. For E h < 75 mV the solid line is a simulation assuming an n = 1 redox process with a midpoint potential of - 10 inV. (B) The coupling signal (e,o) was measured by the amplitude of the trough at g = 1.98 and is plotted as fraction of the maximally observed amplitude. The solid line is a simulation of an n = I redox process with a midpoint potential of + 150 mV. The redox titration was performed as described in the legend to Fig. 2. Closed symbols represent samples taken during progressive reduction of the enzyme; open symbols were measured during reoxidation. After admittance of air to the titration vessel, @ represents the [3Fe-4S] ~ intensity and e the amplitude of the coupling signal. EPR conditions: microwave frequency, 9.27 GHz; temperature, 13.5 K; microwave power incident to the cavity. 2.0 roW; modulation amplitude 1.25 roT.

resulting independent S = ½ systems of Ni(lll) and the [3Fe-4S] ÷ cluster, the latter titrated reversibly with E~ -- - 10 mV, n = 1 (Fig. 3A). The redox behaviour of the two Ni(IlI) forms was somewhat more complicated. The titration curves of both Nir(lll) and Ni,(IlI) were not completely reversible. During reoxidation of the reduced preparation, Ni,(lll)yielded less signal intensity (Fig. 2A), whereas the Nir(IlI)form had gained in intensity (Fig. 2B). It looked as if some conversion of the Ni,(Ill) form to the Nir(lll) form had taken place at low redox potentials ( E h < 0 mV). H2-uptake activity of this preparation, measured simultaneously with the redox titrations, showed indeed some increase. Under the applied conditions (pH 8.0, 30°C, E h < 0 mV) transformation of unready hydrogenase into ready hydrogenase is not unexpected. The activation process of hydrogenases is known to proceed under similar conditions [20], If a conversion of Ni.(llI) to Nir(lll) has occurred at low redox potentials in this experiment, and if both nickel forms titrate with the same midpoint potential, then the sum of the signal intensities of Ni.(lll) and Nir(IIl) should titrate according to a reversible n-- 1 redox process. Indeed, an excellent fit of the reductive and oxidative titration curves of total Ni(IIl) intensity was obtained with an n = 1 Nernst curve for Eh < - 5 0 mV. Both Ni,,(lll) and Nir(IlI) thus titrated with a midpoint potential of - 115 mV at pH 8.0 and 30°C (Fig. 2C).

Redox titrations at 2°C To enable a more direct comparison with the experiments of M. thermoautotrophicum hydrogenase (these were all conducted at pH 6.0 [11]) also redox titrations at pH 6.0 were performed. The temperature of these titrations was lowered from 30 to 2°C in order to decrease the rate of conversion from Ni,(llI) into

Nir(lll). During the titrations at pH 6.0 and 2°C, either before or after addition of redox mediators, no spinspin coupling of Ni(IlI) and a [4Fe-4S] 3+ was observed at high redox potentials. When at pH 6.0 reduced enzyme was oxidised anaerobically during the course of a titration at T >/20°C, such a coupling was observed. Under these conditions, however, development of the interaction signal was extremely slow. Even after 1.5 h the coupling signal had not reached its maximal intensity. Subsequent exposure to air recovered maximal magnetic interaction of Ni(lll) and a [4Fe-4S] 3+ cluster within a couple of minutes. The redox titrations at pH 6.0 and 2°C were performed in two different ways. One procedure (Fig. 4) was designed to test the possibility of strong preferential binding of H~ to the ready Ni(II) state; this would result in an elevation of the apparent midpoint potential of the Nir(lll)/Nir(II) transition. Sodium dithionite was used to bring the preparation to a potential where approximately 10% of the nickel was reduced ( E h -- + 120 mV). In this experiment the concentration of the high potential redox mediators (TMPD, DCIP, 1,2-naphthoquinone-4-sulfonic acid and PMS) was increased to 250/zM in order to establish a strong redox buffer. Then 0.3% H 2 was added to the gasphase. Although 0.3% H a in the gasphase sufficed to reduce Nir(lIl) , simultaneous reduction of Ni,(lll) was observed. Along with these very slow reductions (complete reduction of Ni(lll) to Ni(ll) took 3.5 h) the redox potential of the preparation drifted to - 17 mV. Reoxidation was performed by addition of small amounts of an anaerobic potassium ferricyanide solution. During reoxidation 0.3% H 2 remained present in the gasphase. Under these conditions the [3Fe-4S] + signal titrated reversibly with E L = +95 mV (n = 1). The Niu(lll) form titrated reversibly with a midpoint

163 mlZ Iz

1.o, ,

w .-j I

~_ o.~.

Y

m 10 .B

o

/

Q

QQ Q



° 6

60 mm O

l

60 rain z~ 32mm~

S

#

oo

m

m

m

a

#

,

,

l

.

0.0

400 Eh[mV! Fig. 4. Effect of the redox potential on the prosthetic groups of hydrogenase at pH 6.0 and 2°C in the presence of high potential mediators (mediator cocktail 2) as descn'bed in Materials and Methods. Helium was used as sparging gas. The enzyme was reduced to ~ + 120 mV by addition of a sodium dithionite solution (100 raM). Thereafter 0.3% H 2 was added to the gasphase, which resulted in an autonomous, slow reduction of the preparation with a simultaneous decrease of the potential to - 17 mV. (The drift in potential from = + 120 mV to - 17 mV ranged over 3.5 h.) Anaerobic K3Fe(CN)u solution (250 raM) was used for reoxidation. During reoxidation 0.3% H z was kept in the gasphase. The data points at En < - 2 0 0 mV were obtained during reductive activation under 100% H _,. (A) Ni u(lll)signal intensity (Ii, ra ). The solid and dashed lines are simulations of an n = 1 and an n = 2 redox transition with E~ + 20 inV, respectively. (B) Nit(lit) signal intensity (®, o). The solid line is a simulation of an n = 1 redox transition with Et~ = +32 inV, Closed symbols represent samples taken during the reductive titration; open symbols represent samples taken during the consecutive oxidative titration. After reoxidation of the preparation at 2°C, the temperature of the titration vessel was increased to 45°(2. At this elevated temperature the enzyme was reduced by incubation under 100% H 2 for I h. Reoxidation to = + I0 mV was achieved by incubation under He for 50 rain and subsequent addition of an anaerobic K3Fe(CN) ~ solution. Hereafter air was admitted to the titration vessel. The Ni~(lll) and Nit(Ill) signal intensities of the preparation after 32 and 60 min exposure to air are depicted [] and @, respectively. For EPR conditions see legend to Fig. 2. -200

0

200 z:tVa"h'm"" 400

potential of + 20 mV (Fig. 4A). Whether the titration curve of the Niu(lll)/Niu(ll)transition corresponded to an n = 1 or an n = 2 redox process, could not be established accurately. In contrast, titration of the Ni~(lll) form was completely irreversible. Reduction of the Nit(Ill) signal could be fitted with an n = 1 Nernst curve E~ = +32 mV (Fig. 4B). Reoxidation of the reduced preparation up to + 250 mV did not retrieve any Nir(llI) signal. When the enzyme preparation was subsequently subjected to a redox cycle at 45°C (i.e. 1 h H 2 for complete reduction, followed by 50 min incubation under He and oxidation to + 10 mV by addition of an anaerobic K3Fe(CN) 6 solution) both the Ni~(IlI) and Niu(llI) signal reappeared. When redox titrations at pH 6.0 and 20°C were performed with sodium dithionite as only reductant and potassium ferricyanide as oxidant, somewhat different results were obtained. Whereas the [3Fe-4S] ÷ duster titrated reversibly with a midpoint potential of + 100 + 10 mV, the titration curve more closely corresponded to an n = 2 redox process. The titration curve of the Ni,(III) form could now best be fitted with an n = 2 redox process with E~ = +78 + 20 mV. This midpoint potential of the Niu(III) form deviated considerably from the midpoint potential determined in the titration with 0.3% H 2 present (E~ = + 2 0 mV). The Nir(llI) form again titrated irreversibly. Upon progressive reduction of the sample the Nir(llI) signal

-200

0

200

was lost completely within a potential span of only 40 mV, indicating poor redox equilibration. Upon reoxidation of reduced enzyme no Ni,(lll) signal intensity was retrieved. During the course of all redox titrations at pH 6.0 and 2°C the H z-uptake activity of the enzyme preparation varied drastically. For the titration in the presence of 0.3% H2, reduction of the Nit(Ill) form was accompanied by a drop in the H2-uptake activity to 35% of the starting activity. This decrease in H 2-uptake activity must somehow be related to reduction of the Ni r(lll) form as the contribution of the Niu(III) form to the H2-uptake activity is negligible. Moreover, progressive reoxidation of reduced enzyme by KaFe(CN) 6 led to a further decrease of H 2-uptake activity (to 10%), while Ni,,(lll) reappeared with the same initial intensity. Apparently we have come across an inactivated reduced state of the ready enzyme. Inactivation did not result from deterioration of the enzyme preparation during the course of this time-consuming redox titration. At the start of the experiment of Fig. 4 the ratio of the Nir(llI) form to the sum of Ni,(lll) and Niu(lll) is a measure for the expected increase in hydrogenase activity upon activation of the sample [6,7]. In this particular enzyme preparation an increase in H2-uPtake activity of 1.67 times could be expected. At the end of the titration incubation of the inactivated enzyme under H 2 at 45°C yielded an increase in H2-uP-

164

ulO 3

l

61 E _=

) 2

i

o-',2

o~, ' o'.6

|

2

-2 ! Vl[~(¢'1

Fig. 5. The pH-dependenceof H z-uptake activity(nmol per min per ml) of active hydrogenase at 30 and 2°C. H 2-uptake activitywas determined as described in Materials and Methods and plotted as a function of c-[H + ]. The buffer systemwas 50 mM potassium phosphate with 100/zM H 2. 4.2 mM benz'ylviologenand 80 mM glucoseplus glucose oxidase (16 U/ml). The H 2- and benzylviologen-concentrationswere saturating under all conditions. The stock solution of enzymewas in 100 mM Mes buffer (pH 6.0). (A) Activityassayperformed at 30°Cand pH 4.99, 5.45, 5.90, 6.21, 6.66, 6.94, 7.55 and 8.00, (B) at 2°C and pH 4.50, 5.02, 5.41, 6.00, 6,51, 6.98, 7.50, 7.99 and 8.58. take activity of 1,59 times the initial activity, ruling out any doubts on the intactness of the enzyme preparation. A redox titration of the Ni(III)/Ni(ll) transition conducted at pH 8.0 and 2°C was not reversible either. Under these conditions reduction of Ni(III) to Ni(II) was accompanied by a loss in H2-uptake activity to 76% of the activity of the starting preparation. This implied that at pH 8.0 and 2°C, like at pH 6,0, reduction of Nir(III) has led to a partially inactivated Ni(II) state. pH- and T-dependence o f H2-uptake activity

In general, the activation of H 2 by hydrogenase is supposed to involve the heterolytic splitting of dihydrogen into a hydride and a proton [21]. In the absence of an electron acceptor the hydride remains bound to the nickel ion and a neighbouring basic group (X) on the enzyme might accept the proton, thereby facilitating the dissociation reaction [22]. Under the assumption that this group X is redox active, the reduction degree of X might dictate the ability of the enzyme to active H 2. From this point of view, the ready hydrogenase form might be considered to contain Nir(III)" X - and the unready hydrogenase form might then contain Niu(lII)" X. Reduction of X might also influence the coordination of nickel, resulting in the different EPR spectra for Ni,(III) and Ni,(III). The idea is in line with the fact that reducing conditions are required for activation of unready to ready en~me. The observed decrease in activity of reduced ready enzyme might be related to the physico-chemical properties of the group X. If X - functions as a base which accepts the proton resulting from the heterolytic cleavage of H2, an effect of the ionization of - XH on enzyme kinetics might be expected. Provided that enzyme activity is directly dependent on the ionization of - X H , a pKa of 6.1 for hydrogenase activity at 2°C can be predicted from the

drop in activity of the Ni,(ll). X - form at pH 6.0 and pH 8.0 (to 35 and 76% of the initial activity, respectively) [23,24]. In order to verify this prediction, we have investigated the effect of pH on H2-uptake activity of H 2activated enzyme and of oxidised enzyme (in air), both at 30°C and 2°C. At 30°C hydrogenase activity was clearly dependent on the enzyme being in the basic form as a plot of v against v. [H ÷] revealed a straight line with slope - 1 / K a (Fig. 5A) [24]. The PKa for H2-uptake activity at 30°C was = 5.7 and did not appreciably change with pre-treatment of the enzyme (Table ll). At 2°C the pH-dependence of hydrogenase activity could not be explained simply in terms of only one protonic form of an acid or base being catalytically active. Instead, a graph of v as a function of v - [ H +] (Fig. 5B) suggested the existence of two ionization processes with apparent p K a values of 6.0 and 4.7. The distinct pH-dependence of hydrogenase activity at 2 and 30°C demonstrated the T-dependence of ionizing groups which are either directly involved in catalysis at the active site, or are responsible for maintaining the active conformation of the enzyme. Anyway, the marked differences in H2-uptake activities during the course of redox titrations at 2°C versus titrations at 30°C did not result from changes in the activation TABLE II pK a of"H2-uptake activity at 30°C

The pKa was determined from the slope ( - 1/K a) of a plot of v against c . [ H + ]. Enzyme preparation H 2-activated pH 6 pH 8

pK a

Air oxidised

5.64 5.45

pH 6 pH 8

5.79 5.77

165

vinosum, we would like to correct the results of an

energy of catalysis (Fig. 6). There were no break points in the Arrhenius plots of temperature profiles from 4°C to 35.5°C. For H 2-activated enzyme at pH 7.95 and 6,05 the activation energy was 44 kJ mol-= and 47 kJ tool -~, respectively. The activation energy for airoxidised enzyme was 66 El tool- ~, both at pH 7.95 and 6.05.

earlier redox titration reported by this laboratory [3]. In that particular experiment (pH 7.3) a sudden disappearance of the spin-spin interaction of Ni(llI) and a [4Fe-4S] 3÷ cluster was observed upon lowering the redox potential from - 1 9 mV to - 3 8 mV. The midpoint potential for the reduction of Ni(lll) was estimated to be - 175 mV and for the [3Fe-4S] ÷ cluster a midpoint potential of - 165 mV (pH 7.3) was reported. The pronounced changes in signal intensities of the Ni and Fe-S signals within narrow potential regions in this titration indicate that the composition and concentration of the dyes in the mediator cocktail did not ensure redox equilibration with hydrogenase. For the redox titrations described in this paper we have used a wider

Discussion

Oxidation-reduction potentials of prosthetic groups in unready hydrogenase The midpoint potentials of redox equilibria of Ni and Fe-S in several nickel hydrogenases are summarized in Table III. Regarding the enzyme from C.

T A B L E III

Midpoint potentials o f Ni(lll), tile [3Fe-4S] + cluster and the interaction o f Ni(lll) and a [4Fe-4S] 3 + chtster in oxidised hydrogenases The midpoint-potential values are expressed relative to the standard hydrogen electrode. To enable comparison of the midpoint potential values of redox titrations performed at different pH values, the Era. 7 was calculated. W h e n the temperature of the titration experiment was not reported, room temperature (RT, 20°C) was assumed, In case the pH-dependence of a redox transition was not determined (n.d.), a AE m / p H value of - 6 0 mV was assumed for reduction of Ni(IIl) and for removal of the s p i n - s p i n interaction of Ni(Ill) and a [4Fe-4S] 3+ cluster. Reduction of the [3Fe-4Sl + cluster was considered pH-independent. Assignment of pH-dependence was based on the presumed analogy of a transition in hydrogenases isolated from various microorganisms, No attempt was made to convert the midpoint potentials of titrations performed at 2°C to Em.7-values based on r o o m temperature (20°C). Redox event Niu(Ill)+e - ~ Ni.(ll)

pH

T

A Era/pH

145 - 110 115 75 + 20 +78

8.0 6.0 8.5 7.2 8.1 8.0 7.0 6.0 6.0

25 15 RT RT RT 30 30

n.d. -60 n.d. -60 n.d. -60 - 60

- 115 + 32

8.0 6.0

30

- 75 35 70 30 - 15 + 25 - 20 l0 + 10 +95 + 100 + 40 - 50 20 + 25

8.0 7.0 8.5 7.0 7.0 8. ! 8.5 8.0 7.0 6.0 6.0 7.0 7.0 7.0 7.0

25 RT RT RT RT RT 25 30 30

RT RT RT RT

n.d. n.d. n.d. n.d.

8.1 7.0 8.5 8.0 7.0 7.0

RT 25 25 30 30 RT

n.d. -60 - 60 - 60 - 60 n.d.

E~ -410 -140 -

2

2

0

-

-

-

N i t ( I l l ) + e - ~ Nir(II)

[3Fe-4SI ÷ + e - ~ [3Fe-4S] °

-

-

-

-

-

Ni(llI)- - -[4Fe-4S] 3+

+ + + + = + +

105 160 100 150 220 160

2 2 2

2 2

~ 9

Era,7

25

-65

C. cinosum (185) C. cinosum (185)

this work this work

- 75 - 35 - 70 - 30 - 15 + 9,25 - 20 - 10 + 10

M. formicicum D, gigas 1), gigas D, gigas 19. desulfuricans T. roseopersicina C vinosum (D) C cinosum (185) C vinosum (185) C cinosum (185) C cinosum (185) A. eutrophus mb. A. eutrophus sol. A. eutrophus rob. E. coli

25 28

T. roseopersicina

27 26 26 this work this work 29

9

n.d. 0 n.d. n.d. n.d. n.d. 0 0 0

~ 9

Ref.

M. formicicum M. therm. D. gigas D. gigas T. roseopersichla C cinosum (185) C cinosum (185) C. cinosum (185) C, cinosum (185)

~ 9

-60

q

Bacterium

-350 -210 - 130 - 132 - 44 -65 - 75

9 ~ + 40 - 50 - 20 + 25 + + = + +

171 160 190 210 220 160

C. cinosum (D) C vinosum (D)

C vinosum (185) C vinosum (185) A. eutrophas rob.

II

30 28 27 this work this work this work this work

30 34 34 27 26 this work this work this work this work 29 34 34 34

166 6

O!

3.0

'

'

3.2

'

'

3.,5

'

'

3.6

.

.

.

38

. lIT

.

A,.O 111°1(=10 "31

; 4.2

Fig. 6. Arrhenius plots of temperature profiles of Hz-oxidation by enzyme at pH 6.05 {m,c]} and at pH 7.95 (o,o). Open symbols correspond to datapoints obtained with activated hydrogenase (24 h incubation under H, at room temperature) and closed symbols correspond to data points obtained with oxidised hydrogenase in air. Hz-uptake activity was measured as described in Materials and Methods. The buffer system was identical to the one given in the legend to Fig. 5.

selection of redox mediators which enabled us to vary the redox potential in a controlled way. With improved experimental conditions the Niu(Ill)/Niu(ll) transition, the [3Fe-4S]+/[3Fe-4S] ° transition and the spinspin interaction of Ni(III) and a [4Fe-4S]3+ cluster all titrated completely reversibly with midpoint potentials (pH 7.0) of - 7 0 mV, 0 mV and +210 mV, respectively (Table III). Herewith the discrepancy in results obtained with hydrogenase from C. cinosum strain DSM 185 and those from strain D [26] is removed. When all available information on the midpoint potentials of the redox processes that involve Ni(III) and the [3Fe-4S] + cluster in hydrogenase is compared (Table Ill), several marked characteristics of these prosthetic groups in enzymes from different sources can be recognized. The midpoint potential of the Niu(III)/Ni.(II) transition appears to be strongly pHdependent [11,27,28] which implies that reduction of Ni(llI) is accompanied by addition of a proton, either to Ni or to any other nearby group ( A E 6 = - 6 0 m V / p H unit). A similar pH-dependence is observed for the reduction process that cancels the spin-spin interaction of Ni(lll) and a [4Fe-4S] 3+ cluster [26,27,29]. For pH >/7.0, the redox behaviour of the [3Fe-4S] ÷ cluster is not influenced by changes in pH [26,28]. In spite of some variation in the values of the midpoint potentials of the Ni,(lll)/Ni.(II) transition, this 'unready' Ni(llI) form exhibits similar redox behaviour in hydrogenases isolated from different sources. It may be worth noticing that the midpoint potentials of the Niu(IlI)/Niu(II ) redox equilibria in hydrogenases from various microorganisms can be classified consistent with the division of these microorganisms in their taxonomic groups.

Oxidation-reduction potentials of prosthetic groups in ready hydrogenase When oxidised hydrogenase is in the 'ready' form, the redox behaviour of the nickel ion is less well characterized. Considering the F4z0-nonreducing hydrogenases from M. thermoautotrophicum and M. formicicum, where the nickel ion remains predominantly in the Ni(lI) valence state upon anaerobic oxidation of 'ready' enzyme [11,25], either the Nir(lll)/Nir(II) equilibrium might not be enclosed within the experimental potential range of the redox titrations, or the oxidation of nickel by redox mediators is disturbed in ready enzyme. Reoxidation of H 2-activated D. gigas hydrogenase by DCIP at pH > 7.0 for instance, showed a considerable delay in oxidation of the nickel centre ( E r a . 7 = - 130 mV [28,30]) compared with oxidation of the [3Fe-4S] + cluster (Em, 7 = - 3 5 m W [28]). The [3Fe4S] + cluster could be oxidised completely before Ni(llI) signals appeared [7]. In addition, the total spin concentration of Nit(Ill) and Niu(Ill) in fully oxidised D. gigas hydrogenase also does not equal the chemically determined nickel concentration [7]. The soluble hydrogenases from Desulforibrio salexigenes, Desulfovibrio desulfuricans and from the aerobic H2-oxidizing bacterium Alcaligenes eutrophus do not show any Ni(lll) signals in the oxidised state [31,32]. Even in the presence of 0 2, the Ni(ll) valence state is maintained and H 2 is readily oxidised. In general it appears to be more difficult to oxidize nickel to the Ni(lll) valence state in 'ready' hydrogenase than in 'unready' hydrogenase. The results of a redox titration of C. vinosum hydrogenase at pH 8.0 and 30°C show that both the Nir(lll) form and the Ni,(llI) form titrate according to an n = 1 Nernst curve with a midpoint potential of - 1 1 5 mV. The assumed positive shift in midpoint potential of the Ni(lll)/Ni(II) transition upon activation of hydrogenase from M. thermoautotrophicum [11], is therefore absent in the C. vinosum enzyme. The experiment in Fig. 4 argues against a strong preferential binding of H 2 to the Nir(II) state. Such a binding would induce an elevation of the apparent midpoint potential of the Nir(ll)/Nir(IlI) transition, which is not observed. The marked differences in redox behaviour of the nickel ion in ready and unready enzyme at 2°C is not understood. For a meaningful explanation of the redox behaviour of nickel in hydrogenase the catalytic center and its direct ligands cannot be disengaged from the rest of the protein structure and also the interaction of the redox couple with the mediating dyes in solution should be considered. H2-uptake activity is dependent on an ionizable basic group in the protein The pH-dependence of the H z-uptake activity at 30°C (Fig. 5A) shows that catalytic activity of hydroge-

167 nase is dependent on the presence of a basic group in the protein with a p K a of = 5.7. Whether this ionizable group is directly involved in catalysis or that it maintains the active protein conformation can not be concluded from these data. In any case, the heterolytic cleavage of H_, into a hydride and a proton is expected to be facilitated by the presence of a basic group ( X - ) near the Ha-activating site [21,22]. At 2°C hydrogenase activity could not be explained in terms of only the deprotonated form of one acidic group being catalytically active. It appeared as if at 2°C (Fig. 5B) two activity related ionization processes could be resolved with p K a values of 6.0 and 4.7. From the decrease of H 2-uptake activity in reduced ready enzyme during the titrations at 2°C, a p K , of 6.1 was predicted for hydrogenase activity. Considering experimental uncertainties, this p K a value reasonably well approximates the estimated pK~ of 6.0 obtained from the pH-dependence of H2-uptake activity at 2°C in the kinetic experiment. Comparison of the kinetic studies with the activity measurements during the redox titrations, however, is not directly feasible as in the kinetic studies either completely reduced or completely oxidised enzyme was used in the activity assays, whereas intermediate reduction levels of the enzyme were used during the activity measurements of the redox titrations. In spite of this reservation, indistinguishability of steady-state kinetic parameters derived from experiments with either completely reduced or oxidised enzyme, demonstrated that the processes associated with activation of the enzyme did not change the PKa of hydrogenase activity. The observed decrease in activity of reduced ready enzyme during the redox titrations at 2°C was not due to the temperature jump for the enzyme from the titration vessel to the activity assay at 30°C, since the activation energy of catalysis was not altered between 4 and 35.5°C. Although these experiments did not provide a straightforward clue to the process which led to inactivation of the reduced ready enzyme, the postulated protonation of a basic group X - near the H 2-activating site seems a reasonable possibility.

remarkable. No redox equilibrium could be established between Nir(llI) and the mediator cocktail when the preparation was reduced by NazS204. With H 2 in the gasphase, reduction of Ni~(IlI)proceeded according to an n = 1 Nernst-curve, indicating that the reducing equivalents liberated by dissociation of H z at the active site did equilibrate with the redox mediators in solution. This implies direetedness of the electron transfer pathway between the active site in the protein and the surrounding medium. In this light irreversibility of the Nir(lll)/Nir(II) transition can be interpreted as hampered equilibration of nickel on one side and the oxidants (K3Fe(CN) 6, DCIP) and mediator cocktail on the other side; the Ni~(lll)/Ni~(lI) couple might just feel the H2-potentiai. Irrespective of the reductant used, redox titrations of Ni,(lll) were completely reversible. Yet, the midpoint potential in a titration with H 2 as reductant ( + 2 0 mV) differed considerably from the El) = value calculated from a titration with Na2S204 ( + 7 8 mV). This distinct redox behaviour of Ni,(IlI) when H 2 was used as reductant, compared with that when NaaSaO 4 was the reductant, supports the view that Ni.(lll) can be reduced to Ni(lI) by H 2.

Concluding remarks The redox behaviour of nickel in hydrogenases from a wide variety of microorganisms all seems to converge to the same pattern (Scheme 11). In unready enzyme the Ni(lll)/Ni(ll) transition is reversible, but in ready enzyme it is usually not. Redox equilibration of nickel in ready enzyme and of the spin-spin interaction of Ni(lll) with a [4Fe-4S] 3÷ cluster is strongly dependent on pH and temperature, in active enzyme both the Ni(ll)/Ni(I) and the Ni(1)/'Ni(0)' redox equilibria are completely reversible. The origin of hampered redox equilibration between the prosthetic groups in ready hydrogenase and the mediating dyes might be due to

Effect of 1t2 on the redox equilibration of hydrogenase with redox mediators The results of redox titrations conducted at pH 1> 7.0 (Table III) indicate that reduction of Ni(llI) is accompanied by a protonation (Era, 7 -':" - 7 0 mY; AEg -- - 6 0 m V / p H unit) and that the midpoint potential of the [3Fe-4S] ÷ cluster is not affected by pH-changes (E~ = - 1 0 mV). The rather positive midpoint potentials of both the Niu(lll) + e-+-, Ni,(II) equilibrium (E~ either +20 mV or +78 mV) and the [3Fe-4S]÷+ e-+-, [3Fe4S] ° equilibrium (Et~ = + 100 mV) at pH 6.0 and 2°C clearly deviate from the general pattern. In addition, sensitivity of the redox behaviour of the Ni-ion to the reduetant used in the titrations at pH 6.0 and 2°C is

unready

~-~-~

ready

',,\ Eh

I

17

', \

a,e-

,,\

,/ /

active

,~NiaciITII ~eScheme II. Apparent redox behaviour of nickel in hydrogenase.

168 an obstruction in the electron transfer from nickel at the active site to the artificial electron acceptors in solution. Acknowledgements

We would like to thank Mrs. M. Van Veenhuizen and Mrs. E.C.M. Bouwens for the continuous supply of purified hydrogenase. We are indebted to The Netherlands Organization for the Advancement of Pure Research (N.W.O.) for financial support, supplied via the Netherlands Technology Foundation (S.T.W.) and the Netherlands Foundation for Chemical Research (S.O.N.). References 1 Van der Zwaan, J.W. (1987) On the Active Site of Nickel Hydrogenases, Ph.D. Thesis, pp. 50-52, Kaal Boek, Amsterdam. 2 Cammack, R., Fernandez, V.M. and Schneider, K. (1988) in The Bioinorganic Chemistry of Nickel (Lancaster, J.R., Jr., ed.), pp. 167-190, VCH Verlagsgesellschaft, Weinheim. 3 Van der Zwaan, J.W., Albracht, S.P.J., Fontijn, R.D. and Slater, E.C. (1985) FEBS Lett. 179, 271-277. 4 Van der Zwaan, J.W., Albracht, S.P.J., Fontijn, R.D. and Roelofs, Y.B.M. (1986) Biochim. Biophys. Acta 872, 208-215. 5 Albracht, S.P.J., Kalkman, M.L. and Slater, E.C. (1983) Biochim. Biophys. Acta 724, 309-316. 6 Van der Zwaan, J.W., Coremans, J.M.C.C., Bouwens, E.C.M. and Albracht S.P.J. (1990) Biochim. Biophys. Acta 1041, 101-110. 7 Fernandez, V.M., Hatchikian, E.C., Patil, D.S. and Cammack, R. (1986) Biochim. Biophys. Acta 883, 145-154. 8 Teixeira, M. Moura, I., Xavier, A.V., Huynh, B.H., DerVartanian, D.V., Peck, H.D. Jr., LeGall, J. and Moura, JJ.G. (1985) J. Biol. Chem. 260, 8942-8950. 9 Albracht, S.P.J., Van der Zwaan, J.W. and Fontijn, R.D. (1984) Bioehim. Biophys. Acta 766, 245-258. 10 Albracht, S.P.J., Graf, E.-G. and Thauer, R.K. (1982) FEBS Lett. 140, 311-313. 11 Coremans, J.M.C.C., Van der Zwaan, J.W. and AIbracht, S.P.J. (1989) Biochim. Biophys. Acta 997, 256-267.

12 Van Heerikhuizen, H., Albracht, S.PJ., Slater, E.C. and Van Rheenen, P.S. (1981) Biochim. Biophys. Acta 657, 26-39. 13 Hendley, D.D. (1955) J. Bacteriol. 70, 625-634. 14 Kovacs, K.L. (1985) Preparative Biochemistry 15, 321-334. 15 Dutton, P.L. (1971) Biochim. Biophys. Acta 226, 63-80. 16 Ives, D.J.G. and Janz, GJ. (eds.) (1961) Reference Electrodes, Theory and Practice, Academic Press, New York. 17 Luudin, A. and Aasa, R. (1972) J. Magn. Reson. 8, 70-73. 18 Beinert, H. and AIbracht, S.P.J. (1982) Biochim. Biophys. Acta 683, 245-277. 19 AIbracht, S.P.J. (1984) in Current Topics in Bioenergetics (Lee, C.P., ed.), Vol. 13, p. 79-106, Academic Press, New York. 20 Fernandez, V.M., Hatchikian, E.C. and Cammack, R. (1985) Biochim. Biophys. Acta, 69-79. 21 Krasna, A.I. and Rittenberg, D. (1954) J. Am. Chem. Soc. 76, 3015-3020. 22 Krasna, A.I. (1979) Enzyme Microb. Technol. 1, 165-172. 23 Mahler, H.R. and Cordes, E.H. (1967) Biological Chemistry, Harper and Row, New York. 24 Fersht, A. (1985) Enzyme Structure and Mechanism, W.H. Freeman and Company, New York. 25 Adams, M.W.W., Jin, S.-L.C., Chen, J.-S. and Mortenson, L.E. (1986) Biochim. Biophys. Acta 869, 37-47. 26 Cammack, R., Rao, K.K., Serra, J. and Llama, M.J. (1986) Biochimie 68, 93-96. 27 Cammack, R., Bagyinka, C. and Kovacs, K.L. (1989) Eur. J. Biochem. 182, 357-362. 28 Cammack, R., Patil, D., Aguirre, R. and Hatchikian, E.C. (1982) FEBS Lett. 142, 289-292. 29 Schneider, K., Patti, D.S. and Cammack, R. (1963) Biochim. Biophys. Acta 748, 353-361. 30 Teixeira, M., Moura, I., Xavier, A.V., DerVartanian, D.V., LeGall, J., Peck, H.D., Jr., Huynh, B.H. and Moura, J.J.G. (1983) Eur. J. Biochem. 130, 481-484. 31 Teixeira, M., Moura, I., Faugue, G., Czechowski, M., Berlier, Y., Lespinat, P.A., LeGall, J., Xavier, A.V. and Moura, JJ.G. (1986) Biochimie 68, 75-84. 32 Cammack, R., Fernandez, V.M. and Schneider, K. (1986) Biochimie 68, 85-91. 33 Clark, W.M. (1960) Oxidation-Reduction Potentials of Organic Systems, Williams and Wilkins Company, Baltimore, MD. 34 Cammack, R., Lalla-Maharajh, W.V. and Schneider, K. (1982) in Electron Transport and Oxygen Utilization, (Ho, C., ed.), pp. 411-415, Elsevier North Holland, New York.

Distinct redox behaviour of prosthetic groups in ready and unready hydrogenase from Chromatium vinosum.

The redox behaviour of the Ni(III)/Ni(II) transition in hydrogenase from Chromatium vinosum is described and compared with the redox behaviour of the ...
1MB Sizes 0 Downloads 0 Views