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A Glutamate Is the Essential Proton Transfer Gate during the Catalytic Cycle of the [NiFe] Hydrogenase

2004; Elsevier BV; Volume: 279; Issue: 11 Linguagem: Inglês

10.1074/jbc.m312716200

1083-351X

Sébastien Dementin, Bénédicte Burlat, António L. De Lacey, Alejandro G. Pardo, Géraldine Adryanczyk‐Perrier, Bruno Guigliarelli, Vı́ctor M. Fernández, Marc Rousset,

Electrocatalysts for Energy Conversion

Kinetic, EPR, and Fourier transform infrared spectroscopic analysis of Desulfovibrio fructosovorans [NiFe] hydrogenase mutants targeted to Glu-25 indicated that this amino acid participates in proton transfer between the active site and the protein surface during the catalytic cycle. Replacement of that glutamic residue by a glutamine did not modify the spectroscopic properties of the enzyme but cancelled the catalytic activity except the para-H2/ortho-H2 conversion. This mutation impaired the fast proton transfer from the active site that allows high turnover numbers for the oxidation of hydrogen. Replacement of the glutamic residue by the shorter aspartic acid slowed down this proton transfer, causing a significant decrease of H2 oxidation and hydrogen isotope exchange activities, but did not change the para-H2/ortho-H2 conversion activity. The spectroscopic properties of this mutant were totally different, especially in the reduced state in which a non-photosensitive nickel EPR spectrum was obtained. Kinetic, EPR, and Fourier transform infrared spectroscopic analysis of Desulfovibrio fructosovorans [NiFe] hydrogenase mutants targeted to Glu-25 indicated that this amino acid participates in proton transfer between the active site and the protein surface during the catalytic cycle. Replacement of that glutamic residue by a glutamine did not modify the spectroscopic properties of the enzyme but cancelled the catalytic activity except the para-H2/ortho-H2 conversion. This mutation impaired the fast proton transfer from the active site that allows high turnover numbers for the oxidation of hydrogen. Replacement of the glutamic residue by the shorter aspartic acid slowed down this proton transfer, causing a significant decrease of H2 oxidation and hydrogen isotope exchange activities, but did not change the para-H2/ortho-H2 conversion activity. The spectroscopic properties of this mutant were totally different, especially in the reduced state in which a non-photosensitive nickel EPR spectrum was obtained. Many microorganisms use molecular hydrogen in their metabolic routes as an energy source or for evacuating an excess of electrons. The enzymes that catalyze reversibly the conversion of molecular hydrogen to two electrons and two protons are known as hydrogenases. Although this is the simplest chemical reaction, the catalytic mechanism of these enzymes is quite complicated, and its details are still a matter of debate (1Cammack R. Cammack R. Frey M. Robson R. Hydrogen as a Fuel. Taylor & Francis, London, New York2001: 159-180Google Scholar). Hydrogen isotope exchange experiments indicate that the H2 cleavage reaction is heterolytic; thus, a hydride and a proton are formed in the first step (2Yagi T. Tsuda M. Inokuchi H. J. Biochem. (Tokyo). 1973; 73: 1069-1081Crossref PubMed Scopus (65) Google Scholar, 3Krasna A.I. Rittenberg D. J. Am. Chem. Soc. 1954; 76: 3015-3020Crossref Scopus (95) Google Scholar). In the second step, the two electrons of the hydride are extracted, and a second proton is formed. Subsequently, the two electrons have to be transported, via the intramolecular electron transfer chain, from the active site to the redox partner of the hydrogenase (a redox protein or NAD+(P)) in vivo, or a redox dye in vitro; the two protons have to be transferred to the protein environment as well. These steps are reversed in the case of H2 production activity (1Cammack R. Cammack R. Frey M. Robson R. Hydrogen as a Fuel. Taylor & Francis, London, New York2001: 159-180Google Scholar). How do all these steps take place in hydrogenases? These proteins are metalloenzymes that all contain iron, and in many cases, also nickel. The crystallographic structures of several [Fe] hydrogenases (4Peters J.W. Lanzilotta W.N. Lemon B.J. Seefeldt L.C. Science. 1998; 282: 1853-1858Crossref PubMed Google Scholar, 5Nicolet Y. Piras C. Legrand P. Hatchikian C.E. Fontecilla-Camps J.C. Structure Fold. Des. 1999; 7: 13-23Abstract Full Text Full Text PDF Scopus (1240) Google Scholar) and [NiFe] hydrogenases (6Higuchi Y. Yagi T. Biochem. Biophys. Res. Commun. 1999; 255: 295-299Crossref PubMed Scopus (32) Google Scholar, 7Volbeda A. Charon M.H. Piras C. Hatchikian E.C. Frey M. Fontecilla-Camps J.C. Nature. 1995; 373: 580-587Crossref PubMed Scopus (1362) Google Scholar) have been obtained by x-ray diffraction studies. In both types of enzymes, the active site is a deeply buried bimetallic center, in which the metals are bridged by thiol groups and have CO and CN- as ligands. This type of coordination favors the binding of molecular hydrogen or hydride to the active site (1Cammack R. Cammack R. Frey M. Robson R. Hydrogen as a Fuel. Taylor & Francis, London, New York2001: 159-180Google Scholar, 8Darensbourg M.Y. Lyon E.J. Smee J.J. Coord. Chem. Rev. 2000; 206-207: 533-561Crossref Scopus (341) Google Scholar). The crystal structures also indicate that Fe-S clusters are located between the active site and the protein surface, which are thought to form the intramolecular electron pathway in the H2 production/oxidation mechanism (9Page C.C. Moser C.C. Chen X. Dutton P.L. Nature. 1999; 402: 47-52Crossref PubMed Scopus (1473) Google Scholar). In [NiFe] hydrogenases, one nickel and one iron atom form the bimetallic center. The nickel is coordinated to four cysteine ligands via their thiol groups. Two of them are terminal ligands, and the other two are bridging ligands that also coordinate the iron atom (7Volbeda A. Charon M.H. Piras C. Hatchikian E.C. Frey M. Fontecilla-Camps J.C. Nature. 1995; 373: 580-587Crossref PubMed Scopus (1362) Google Scholar). One CO and two CN-, which are detected by FTIR, 1The abbreviations used are: FTIR, Fourier transform infrared; MES, 4-morpholineethanesulfonic acid; CAPS, 3-(cyclohexylamino)propanesulfonic acid.1The abbreviations used are: FTIR, Fourier transform infrared; MES, 4-morpholineethanesulfonic acid; CAPS, 3-(cyclohexylamino)propanesulfonic acid. are also ligands of the iron atom (10Volbeda A. Garcin E. Piras C. de Lacey A.L. Fernandez V.M. Hatchikian C.E. Frey M. Fontecilla-Camps J.C. J. Am. Chem. Soc. 1996; 118: 12989-12996Crossref Scopus (579) Google Scholar, 11Happe R.P. Roseboom W. Pierik A.J. Albracht S.P. Bagley K.A. Nature. 1997; 385: 126Crossref PubMed Scopus (391) Google Scholar). In addition, an oxygen species also bridges both metals in the inactive oxidized states (10Volbeda A. Garcin E. Piras C. de Lacey A.L. Fernandez V.M. Hatchikian C.E. Frey M. Fontecilla-Camps J.C. J. Am. Chem. Soc. 1996; 118: 12989-12996Crossref Scopus (579) Google Scholar), whereas it disappears in the active reduced states and is likely replaced by a hydride bridge (8Darensbourg M.Y. Lyon E.J. Smee J.J. Coord. Chem. Rev. 2000; 206-207: 533-561Crossref Scopus (341) Google Scholar, 12Carepo M. Tierney D.L. Brondino C.D. Yang T.C. Pamplona A. Telser J. Moura I. Moura J.J. Hoffman B.M. J. Am. Chem. Soc. 2002; 124: 281-286Crossref PubMed Scopus (114) Google Scholar, 13Higuchi Y. Ogata H. Miki K. Yasuoka N. Yagi T. Structure Fold. Des. 1999; 7: 549-556Abstract Full Text Full Text PDF Scopus (308) Google Scholar, 14Garcin E. Vernede X. Hatchikian E.C. Volbeda A. Frey M. Fontecilla-Camps J.C. Structure Fold. Des. 1999; 7: 557-566Abstract Full Text Full Text PDF Scopus (407) Google Scholar). The redox chemistry of [NiFe] hydrogenases is very rich, and at least seven redox states of the active site are detected by FTIR (15de Lacey A.L. Hatchikian E.C. Volbeda A. Frey M. Fontecilla-Camps J.C. Fernandez V.M. J. Am. Chem. Soc. 1997; 119: 7181-7189Crossref Scopus (250) Google Scholar). Three of these states are EPR-active due to paramagnetic properties of the nickel. These states are named Ni-A or unready (an inactive oxidized state that needs a long activation period to catalyze H2 oxidation), Ni-B or ready (an inactive oxidized state that can be quickly activated upon reduction), and Ni-C or active (an active reduced state) (16Moura J.J. Moura I. Huynh B.H. Kruger H.J. Teixeira M. DuVarney R.C. DerVartanian D.V. Xavier A.V. Peck Jr., H.D. LeGall J. Biochem. Biophys. Res. Commun. 1982; 108: 1388-1393Crossref PubMed Scopus (97) Google Scholar, 17Cammack R. Patil D.S. Hatchikian E.C. Fernandez V.M. Biochim. Biophys. Acta. 1987; 912: 98-109Crossref Scopus (135) Google Scholar). The EPR-silent states are named SU (an unready reduced state), SI (a silent active state, which has two species in acid-base equilibrium), and R (the super-reduced state) (15de Lacey A.L. Hatchikian E.C. Volbeda A. Frey M. Fontecilla-Camps J.C. Fernandez V.M. J. Am. Chem. Soc. 1997; 119: 7181-7189Crossref Scopus (250) Google Scholar). As indicated before, the identification of the electron transfer pathway in hydrogenases is quite straightforward from the crystallographic structure. In standard [NiFe] hydrogenases, it is formed by two [4Fe4S] clusters and one [3Fe-4S] cluster (7Volbeda A. Charon M.H. Piras C. Hatchikian E.C. Frey M. Fontecilla-Camps J.C. Nature. 1995; 373: 580-587Crossref PubMed Scopus (1362) Google Scholar), but the relative value of their midpoint potential raised questions about their influence on the electron transfer kinetics (18Bertrand P. Dole F. Asso M. Guigliarelli B. J. Biol. Inorg. Chem. 2000; 5: 682-691Crossref PubMed Scopus (27) Google Scholar, 19Rousset M. Montet Y. Guigliarelli B. Forget N. Asso M. Bertrand P. Fontecilla-Camps J.C. Hatchikian E.C. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11625-11630Crossref PubMed Scopus (174) Google Scholar). However, the distances between these redox clusters are adequate for fast electron transfer through the protein (9Page C.C. Moser C.C. Chen X. Dutton P.L. Nature. 1999; 402: 47-52Crossref PubMed Scopus (1473) Google Scholar). A hydrophobic channel that connects the protein surface with the active site for H2 transport has also been detected by x-ray diffraction studies in Desulfovibrio fructosovorans [NiFe] hydrogenase (20Montet Y. Amara P. Volbeda A. Vernede X. Hatchikian E.C. Field M.J. Frey M. Fontecilla-Camps J.C. Nat. Struct. Biol. 1997; 4: 523-526Crossref PubMed Scopus (303) Google Scholar). There is much less evidence regarding the proton transport pathway. Protons are transported inside proteins via motions of water molecules and amino acid residues with acid-base properties, such as histidine, glutamic acid, and aspartic acid (21Williams R.J. Nature. 1995; 376: 643Crossref PubMed Scopus (35) Google Scholar). Several routes have been proposed for [NiFe] hydrogenases, and most likely, proton transport during catalysis does not stick to a single one (7Volbeda A. Charon M.H. Piras C. Hatchikian E.C. Frey M. Fontecilla-Camps J.C. Nature. 1995; 373: 580-587Crossref PubMed Scopus (1362) Google Scholar, 14Garcin E. Vernede X. Hatchikian E.C. Volbeda A. Frey M. Fontecilla-Camps J.C. Structure Fold. Des. 1999; 7: 557-566Abstract Full Text Full Text PDF Scopus (407) Google Scholar, 22Fontecilla-Camps J.C. Frey M. Garcin E. Higuchi Y. Montet Y. Nicolet Y. Volbeda A. Cammack R. Frey M. Robson R. Hydrogen as a Fuel. Taylor & Francis, London, New York2001: 93-109Google Scholar, 23Matias P.M. Soares C.M. Saraiva L.M. Coelho R. Morais J. Le Gall J. Carrondo M.A. J. Biol. Inorg. Chem. 2001; 6: 63-81Crossref PubMed Scopus (182) Google Scholar). 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A glutamic acid residue conserved in [NiFe] hydrogenases (Glu-25 in D. fructosovorans and Glu-18 in Desulfovibrio gigas hydrogenases) is H-bonded to the mentioned terminal cysteine bound to nickel. Thus, it is a potential candidate for the next step of the proton transport pathway (14Garcin E. Vernede X. Hatchikian E.C. Volbeda A. Frey M. Fontecilla-Camps J.C. Structure Fold. Des. 1999; 7: 557-566Abstract Full Text Full Text PDF Scopus (407) Google Scholar, 22Fontecilla-Camps J.C. Frey M. Garcin E. Higuchi Y. Montet Y. Nicolet Y. Volbeda A. Cammack R. Frey M. Robson R. Hydrogen as a Fuel. Taylor & Francis, London, New York2001: 93-109Google Scholar, 23Matias P.M. Soares C.M. Saraiva L.M. Coelho R. Morais J. Le Gall J. Carrondo M.A. J. Biol. Inorg. Chem. 2001; 6: 63-81Crossref PubMed Scopus (182) Google Scholar). In this work, we report the spectroscopic and kinetic characterization of mutants produced at the Glu-25 position of D. fructosovorans [NiFe] hydrogenase, demonstrating that this residue has an important role in hydrogenase catalysis. Bacterial Strains, Plasmids, and Growth Conditions—Escherichia coli strain DH5α, F-, endA1, hsdR17 (rK- mK+), supE44, thi-1, λ-, recA1, gyrA96, relA1, Δ (argF- lacZYA) U169, φ80dlacZΔM15 was used as a host in the cloning of recombinant plasmids. The bacterium was routinely grown at 37 °C in LB medium. Ampicillin at 100 μg/ml or gentamycin at 20 μg/ml was added when cells harbored pUC18 or pBGF4 derivatives, respectively. The pBGF4 plasmid, which is a new shuttle vector of the pBM family, reporting the gentamycin resistance gene (33Rousset M. Casalot L. Rapp-Giles B.J. Dermoun Z. de Philip P. Belaich J.P. Wall J.D. Plasmid. 1998; 39: 114-122Crossref PubMed Scopus (43) Google Scholar), was used in this study to carry the [NiFe] hydrogenase operon from D. fructosovorans as described previously (19Rousset M. Montet Y. Guigliarelli B. Forget N. Asso M. Bertrand P. Fontecilla-Camps J.C. Hatchikian E.C. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11625-11630Crossref PubMed Scopus (174) Google Scholar). D. fructosovorans strain MR400 [hyn::npt ΔhynABC] carrying a deletion in the [NiFe] hydrogenase operon (34Rousset M. Dermoun Z. Chippaux M. Belaich J.P. Mol. Microbiol. 1991; 5: 1735-1740Crossref PubMed Scopus (38) Google Scholar) was grown anaerobically at 37 °C in SOS medium (33Rousset M. Casalot L. Rapp-Giles B.J. Dermoun Z. de Philip P. Belaich J.P. Wall J.D. Plasmid. 1998; 39: 114-122Crossref PubMed Scopus (43) Google Scholar). Large culture volumes were performed as described previously (19Rousset M. Montet Y. Guigliarelli B. Forget N. Asso M. Bertrand P. Fontecilla-Camps J.C. Hatchikian E.C. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11625-11630Crossref PubMed Scopus (174) Google Scholar). Kanamycine at 50 μg/ml was present routinely, and 50 μg gentamycin/ml were added only when cells harbored the plasmid pBGF4. Site-directed Mutagenesis—The QuikChange™ XL site-directed mutagenesis kit (Stratagene, Amsterdam, The Netherlands) was used to generate point mutations in the large subunit gene hynB. The PstI-AatII fragment from pBGF4 was subcloned into pUC18 to generate pUCbn that was used as a template in mutagenesis experiments. After mutagenesis, the PstI-AatII fragment was fully sequenced and inserted in the PstI-AatII-digested pBGF4. The recombinant plasmid was introduced into D. fructosovorans MR400 by electrotransformation (33Rousset M. Casalot L. Rapp-Giles B.J. Dermoun Z. de Philip P. Belaich J.P. Wall J.D. Plasmid. 1998; 39: 114-122Crossref PubMed Scopus (43) Google Scholar). Protein Purification—The Strep tag II sequence (IBA Gmbh, Göttingen, Germany) was introduced in the hydrogenase genes, and the tagged protein was purified in one step on a Strep-Tactin® column (IBA Gmbh). The yield of purification was in a range between 0.3 and 0.5 mg of pure hydrogenase/liter of culture. Activity Measurements—H2 uptake activity was measured with 1 mm methyl viologen as described (35Fernandez V.M. Hatchikian E.C. Cammack R. Biochim. Biophys. Acta. 1985; 209: 69-79Crossref Scopus (179) Google Scholar) modified as follows for pH dependence. pH was titrated between 6 and 11 with HCl or NaOH in a buffer containing 100 mm MES/Tris/CAPS and 2 mm methyl viologen as mediator. Oxygen was removed under vacuum, and the cuvette was flushed with argon. Enzyme (10 μg for wild type and E25D and 1 mg for E25Q) was allowed to activate for 1 h under H2 in the buffer at pH 8. The activity was measured in an H2-flushed UV-cuvette, containing 1 ml of buffer, in which residual oxygen was eliminated by adding 1 μl of a 1 m dithionite solution. The reaction was started by the addition of 5-30 μl of activated enzyme, and the kinetics of the reduction of methyl viologen was measured at 604 nm in a UV 1601 spectrophotometer (Shimadzu) at 30 °C. H+/D+ exchange activity was measured as described (36De Lacey A.L. Fernandez V.M. Rousset M. Cavazza C. Hatchikian E.C. J. Biol. Inorg. Chem. 2003; 8: 129-134Crossref PubMed Scopus (52) Google Scholar). Para-ortho hydrogen conversion was measured as described (37De Lacey A.L. Santamaria E. Hatchikian E.C. Fernandez V.M. Biochim. Biophys. Acta. 2000; 1481: 371-380Crossref PubMed Scopus (21) Google Scholar). Molecular hydrogen consisting of 50% para hydrogen and 50% ortho hydrogen was prepared as described by Hartman et al. (38Hartmann G.C. Santamaria E. Fernandez V.M. Thauer R.K. J. Biol. Inorg. Chem. 1996; 1: 446-450Crossref Scopus (28) Google Scholar). Redox Titration and EPR Spectroscopy—The redox titrations of the wild type and mutant enzymes were carried out in a specially designed anaerobic cell at 25 °C in 50 mm Hepes buffer at pH = 8.0 under an argon atmosphere. The redox potentials were measured with a combined Pt-Ag/AgCl/KCl (3 m) microelectrode in the presence of the following mediators at 5 μm each: 1,2 naphtoquinone, methylene blue, phenazine methosulfate, phenosafranin, neutral red, methyl viologen. The reductive titration was conducted by stepwise additions of small quantities of 20 mm sodium dithionite in the same buffer. EPR spectra were recorded on a Bruker ESP 300E spectrometer fitted with an Oxford Instruments ESR 900 helium flow cryostat. When necessary, sample illumination was performed directly in the EPR cavity by using a fiber optic and a 250-watt quartz-tungsten-iodine lamp. FTIR Spectroscopy—The infrared experiments were done in a spectroelectrochemical cell as described (39De Lacey A.L. Stadler C. Fernandez V.M. Hatchikian E.C. Fan H.J. Li S. Hall M.B. J. Biol. Inorg. Chem. 2002; 7: 318-326Crossref PubMed Scopus (85) Google Scholar). Biochemical Properties—The activity of the mutants was measured by three different assays, which give complementary information. H2 uptake in the presence of a redox dye involves the steps of H2 splitting at the active site as well as the steps of hydron and electron transfer. Isotopic exchange reactions involve both the splitting of H2 (or its isotopes) and the hydron transfer steps but not the electron transfer step. Finally, conversion of para-H2 to ortho-H2 involves only the step of splitting and recombination of H2 at the active site. Fig. 1 shows the pH dependence of the H2 uptake activity values for wild type hydrogenase and the mutants E25D and E25Q. E25D has a similar pH profile to wild type enzyme, and its activity is roughly half of the native hydrogenase activity. Instead, the other mutant, E25Q has only residual activity (less than 0.1% of the wild type one). Rather similar results were obtained when comparing the D2/H+ exchange activity of the hydrogenase mutants with the wild type one (Fig. 2). Virtually no activity could be detected with E25Q, whereas E25D had activity values that were 30-60% of those of the wild type at acidic and neutral pH. In all measurements, the rate of evolution of the double exchange product (H2) was significantly lower than the rate of evolution of the single exchange product (HD), which is the typical behavior of standard [NiFe] hydrogenases (40Fauque G. Peck Jr., H.D. Moura J.J. Huynh B.H. Berlier Y. DerVartanian D.V. Teixeira M. Przybyla A.E. Lespinat P.A. Moura I. FEMS Microbiol. Rev. 1988; 4: 299-344Crossref PubMed Google Scholar). An interesting result is that at basic pH, the D2/H+ exchange activity of E25D drops down, whereas the H2 uptake activity maintains high values at pH 8 and 9. To determine whether this discrepancy was due to an isotopic effect or to a change in a rate-limiting step at high pH, the D2 uptake activities of the wild type and E25D mutant were compared, and the H2 production activity of the wild type and mutant enzyme was tested at pH 9.0. The D2 uptake activity values of 63 and 36 units/mg, respectively, were very similar to those obtained with H2 uptake activity, but the H2 production activity of the mutant exhibited a 80% decrease at pH 9.0 relative to the wild type enzyme (not shown). The results indicate that the different pH profiles for the D2/H+ exchange probably originate from the H2 production step becoming rate-limiting at high pH in the E25D mutant. The E25Q mutant had negligible activity in these assays. The ortho-para hydrogen conversion experiments show that the mutations had little effect on this activity assay (Fig. 3). After a 200-min reaction, the two mutants and wild type hydrogenase reached similar levels of hydrogen conversion, whereas the blank measurement hardly gave hydrogen conversion (O2 traces cause a small increase of the ortho-H2/para-H2 ratio). EPR Spectroscopy—To investigate the influence of Glu-25 mutations on the active site of the enzyme, the various paramagnetic states of the [NiFe] center were studied by EPR spectroscopy. The two mutants were titrated anaerobically, and the [NiFe] EPR signals were compared with those given by the wild type enzyme poised at the same potentials. In the oxidized state, the EPR spectra of E25Q are composed of a mixture of the Ni-A (g = 2.31, 2.24, 2.01) and Ni-B (g = 2.33, 2.16, 2.01) signals identical to those of the wild type enzyme (Fig. 4, a and b). The total amount of Ni-A + Ni-B signals represents 0.8 spin/mol of enzyme in E25Q preparations, with a proportion of the Ni-B signal corresponding to 40% of the total nickel signal, a value slightly higher than those usually found for the wild type enzyme. In the oxidized E25D mutant, the nickel EPR signal is mainly composed of the usual Ni-A signal (50-55%), but a new species with g = 2.286, 2.238, 2.01 is also present (Fig. 4c). This species, which we termed Ni-A′, corresponds to about 40% of the total nickel signal, which represents typically 0.3-0.4 spin/mol of enzyme. The rest of the spectrum is composed of a minor fraction (5-10%) of Ni-B signal. In both mutants, all these nickel signals disappear upon reduction and are replaced by other nickel signals in the -350 mV to -400 mV potential range (Fig. 5). The reduced E25Q mutant gives the usual Ni-C signal (Fig. 5b), which is split at low temperature ( 1 h). Interestingly, at very low temperature (< 4 K), this complex EPR spectrum exhibits additional splitting (Fig. 5e), which reveals the magnetic coupling between the [NiFe] center and the proximal [4Fe-4S]+1 center. This coupling is much smaller than that observed for the split Ni-C signal (Fig. 5c) and leads to the larger splitting on the gY type lines. These features are similar to those found for the Ni-L species where the intercenter exchange interaction with the [4Fe-4S]1+ cluster is cancelled, and the magnetic coupling results from dipolar coupling only (42Dole F. Medina M. More C. Cammack R. Bertrand P. Guigliarelli B. Biochemistry. 1996; 35: 16399-16406Crossref PubMed Scopus (42) Google Scholar). Thus, the new nickel species found in the reduced E25D mutant are reminiscent of the Ni-L species; they show a similar g-tensor rhombicity and weakness of the spin-coupling with the proximal [4Fe-4S]1+ cluster. For the two Glu-25 mutants, the EPR signals of the FeS clusters are unaffected by mutation (not shown).Fig. 5EPR spectra of hydrogenase Glu-25 mutant in the reduced state. a, wild type enzyme poised at -359 mV. b and c, E25Q mutant poised at -370 mV. d and e, E25D mutant poised at -400 mV. Conditions were as follows: temperature of 30 K (a, b, and d), temperature of 6 K (c), temperature of 3.2 K (e) K; microwave power of 10 milliwatts (a-d); microwave power of 0.4 milliwatts (e); modulation amplitude of 1 milliteslas at 100 kHz (a-d), modulation amplitude of 1 milliteslas at 1 kHz (e). Spectrum e is a 10-scan accumulation.View Large Image Figure ViewerDownload Hi-res image Download (PPT) FTIR Spectroscopy—Infrared measurements were also performed to complete the characterization of the active site of Glu-25 mutants (Fig. 6). The spectra of the oxidized samples of both mutants show the intense band at 1947 cm-1, corresponding to the stretching vibrational mode of the CO ligand in the Ni-A state of D. fructosovorans [NiFe] hydrogenase (39De Lacey A.L. Stadler C. Fernandez V.M. Hatchikian E.C. Fan H.J. Li S. Hall M.B. J. Biol. Inorg. Chem. 2002; 7: 318-326Crossref PubMed Scopus (85) Google Scholar). At 2096 and 2084 cm-1, the less intense bands appear due to the CN- ligands in the same state. The shoulders at 2091 and 2081 cm-1 correspond without doubt to the CN- bands of the Ni-B state. These later ones are more evident for the E25Q mutant, which is in agreement with the EPR results. The frequencies of the bands are equal to those of wild type enzyme (39De Lacey A.L. Stadler C. Fernandez V.M. Hatchikian E.C. Fan H.J. Li S. Hall M.B. J. Biol. Inorg. Chem. 2002; 7: 318-326Crossref PubMed Scopus (85) Google Scholar). The shoulder that is clearly observed at 1953 cm-1 for E25D could correspond to the CO band of the Ni-A′ state or to an EPR-silent state, which is in high proportion in the oxidized state, as the EPR spectrum indicates. This shoulder often appears, although in lower proportion, in some native samples of [NiFe] hydrogenases (15de Lacey A.L. Hatchikian E.C. Volbeda A. Frey M. Fontecilla-Camps J.C. Fernandez V.M. J. Am. Chem. Soc. 1997; 119: 7181-7189Crossref Scopus (250) Google Scholar, 43Albracht S.P. Biochim. Biophys. Acta. 1994; 1188: 167-204Crossref PubMed Scopus (444) Google Scholar). Thus, it probably corresponds to an EPR-silent state that is sometimes present in their oxidized samples. Fig. 6 also shows the FTIR spectra of the mutants after the reducti