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J. Biol. Chem., Vol. 280, Issue 12, 11007-11017, March 25, 2005

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Mechanistic Studies of Rhodobacter sphaeroides Me₂SO Reductase^*

Nathan Cobb, Thomas Conrads, and Russ Hille¶

From the Department of Molecular and Cellular Biochemistry, The Ohio State University, Columbus, Ohio 43210-1218

Received for publication, October 25, 2004 , and in revised form, December 29, 2004.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
Studies of the molybdenum-containing dimethyl sulfoxide reductase from Rhodobacter sphaeroides have yielded new insight into its catalytic mechanism. A series of reductive titrations, performed over the pH range 6–10, reveal that the absorption spectrum of reduced enzyme is highly sensitive to pH. The reaction of reduced enzyme with dimethyl sulfoxide is found to be clearly biphasic throughout the pH range 6–8 with a fast, initial substrate-binding phase and substrate-concentration independent catalytic phase. The intermediate formed at the completion of the fast phase has the characteristic absorption spectrum of the established dimethyl sulfoxide-bound species. Quantitative reductive and oxidative titrations of the enzyme demonstrate that the molybdenum center takes up only two reducing equivalents, implying that the two pyranopterin equivalents of the molybdenum center are not formally redox active. Finally, the visible spectrum associated with the catalytically relevant "high-g split" Mo(V) species has been determined. Spectral deconvolution and EPR quantitation of enzyme-monitored turnover experiments with trimethylamine N-oxide as substrate reveal that no substrate-bound intermediate accumulates and that Mo(V) content remains near unity for the duration of the reaction. Similar experiments with dimethyl sulfoxide show that significant quantities of both the Mo(V) species and the dimethyl sulfoxide-bound complex accumulate during the course of reaction. Accumulation of the substrate-bound complex in the steady-state with dimethyl sulfoxide arises from partial reversal of the physiological reaction in which the accumulating product, dimethyl sulfide, reacts with oxidized enzyme to yield the substrate-bound intermediate, a process that significantly slows turnover.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
Dimethyl sulfoxide reductase is a bacterial enzyme responsible for the reduction of dimethyl sulfoxide (Me₂SO)¹ to dimethyl sulfide (DMS). Substrate arises via breakdown of dimethylsulfoniopropionate, the principal osmolyte of marine algae, and is relatively abundant in the environment. The significance of the reaction rests in the important role of DMS in modulating global solar albedo (1), DMS is one of several molecules that has been traced specifically in recent iron-seeding experiments in the equatorial Pacific Ocean (2). In organisms such as Escherichia coli, Me₂SO reductase is a membrane-bound terminal respiratory oxidase consisting of separate molybdenum- and iron-sulfur-containing subunits, as well as a membrane anchor. In organisms such as Rhodobacter sphaeroides and Rhodobacter capsulatus, on the other hand, the enzyme is a soluble periplasmic protein of 85 kDa possessing only a molybdenum center (3); it appears to be involved in dissipating excess reducing equivalents in the absence of oxygen and is maximally expressed when the organism is grown photoheterotrophically on a relatively reduced carbon source such as malate. Me₂SO reductases from both sources are very similar, and members of the same family of mononuclear molybdenum enzymes, and share a common active site structure in which the metal is coordinated by 2 eq of a pyranopterin cofactor (common to all molybdenum enzymes other than nitrogenase, and also found in all tungsten-containing enzymes) via an enedithiolate linkage.

The Rhodobacter enzyme catalyzes the reduction of Me₂SO according to the following stoichiometry, using the pentaheme c-type cytochrome DorC (4) as the physiological source of reducing equivalents: Me₂S = O + DorC_2e– + 2H⁺ Me₂S + DorC_ox + H₂O. Me₂SO reductase has been the subject of a number of spectroscopic studies because of the distinctive absorption features of its molybdenum center and the absence of other redox-active chromophores (5–13). Its structure has also been extensively examined by x-ray crystallography (3, 14–17). Although initially there was some controversy concerning the active site structure, the crystal structure of R. sphaeroides Me₂SO reductase has recently been determined at 1.3-Å resolution (17), where it is found that the oxidized enzyme is heterogeneous at the molybdenum center with two alternate conformations: a pentacoordinate dioxo monodithiolene formed when a molecule of HEPES buffer binds near the active site, as well as a hexacoordinate mono-oxo bisdithiolene ligand set (with the sixth ligand position occupied by Ser¹⁴⁷ from the protein). This latter structure is thought to represent the catalytically functional form of the oxidized active site (18), whereas the former is known to form upon aerobic incubation of oxidized enzyme in HEPES-containing buffer (18). Possessing the serinate, two oxygen, and one (bidentate) dithiolene ligand to the molybdenum center, this "HEPES-modified" enzyme unsurprisingly displays a visible spectrum analogous to that of oxidized Arabidopsis thaliana sulfite oxidase (SO); a mononuclear molybdenum enzyme with a similar pentacoordinate active site (20).

Although Me₂SO reductase from both R. sphaeroides and R. capsulatus has been extensively studied spectroscopically and crystallographically, much remains to be learned concerning its reaction mechanism. Utilizing UV-visible and EPR spectrometry, the present work seeks to elucidate more clearly the mechanism of Me₂SO reductase activity for pre-steady-state turnover in the reductive and oxidative directions as well as enzyme behavior in the steady-state. A study of the oxidative half-reaction of reduced Me₂SO reductase with Me₂SO shows biphasic kinetics over the pH range 6–8, with a kinetically competent intermediate formed with an extremely fast, substrate concentration-dependent rate (extrapolated at infinite [Me₂SO] to a value in excess of 1000 s^–1); this intermediate subsequently breaks down in a substrate-concentration independent manner. The intermediate exhibits a spectrum dominated by two strong absorption bands at 470 and 550 nm and represents the Me₂SO-bound, reduced form of the enzyme that has been characterized by resonance Raman spectroscopy (11) and x-ray crystallography (14). Above pH 8 the kinetics of E_red·Me₂SO decay become increasingly non-ideal, yet show that the rate of enzyme oxidation slows with an apparent pK_a of 8.5, the pH-dependent behavior observed is similar to that determined for forward steady-state activity in both the R. capsulatus (5) and R. sphaeroides enzymes (7). Additionally, we have determined that the visible spectrum of fully reduced enzyme is particularly sensitive to changes in pH as compared with the spectra for either oxidized or Me₂SO-bound Me₂SO reductase. As in the oxidative half-reaction, the pK_a associated with the spectral changes observed for reduced enzyme were found to be in good agreement with that reported for steady-state activity (5, 7). In ensuring that neither pterin moiety is formally redox active, quantitative reductive titrations, performed with the water-soluble phosphine PTA, as well as DMS and sodium dithionite show that the enzyme is indeed a frank 2-electron system. Enzyme-monitored turnover experiments of the reductive half-reaction using sodium dithionite over the pH range 6–10 have revealed the visible spectrum of the 1-electron reduced enzyme. EPR spectrometry confirms that this catalytic intermediate represents the Mo(V) state giving rise to the so-called "high-g split" EPR signal (9). The visible spectrum of this Mo(V) species displays negligible pH sensitivity and is characterized by two absorption bands at 550 and 675 nm, similar to the glycerol-inhibited Mo(V) form of the enzyme, generated by exposure of reduced Me₂SO reductase to 50% (v/v) glycerol as described by Finnegan et al. (6). Finally, spectral deconvolution via multiple component analysis (MCA) of enzyme-monitored turnover experiments (in which the absorption spectrum of enzyme is followed in the course of steady-state turnover with substrate) reveal no accumulation of the substrate-bound intermediate in the course of steady-state turnover with trimethylamine N-oxide (TMAO). However, significant accumulation of this intermediate is observed using Me₂SO as substrate, increasing to 80% late in the course of the reaction. This behavior is the result of product-inhibition by DMS, which, unlike trimethylamine, is able to rebind the oxidized enzyme to yield the E_red·Me₂SO complex. This behavior presumably reflects the thermodynamic stability of the Me₂SO-bound, reduced form of the enzyme.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
R. sphaeroides was grown photoheterotrophically at 30 °C using Omerod's medium with malate as the principle carbon source and in the presence of 60 mM Me₂SO. Bacteria were harvested in late log phase and stored until needed at –20 °C. Me₂SO reductase was purified as follows, with all purification steps performed at 4 °C unless otherwise specified. Approximately 100 g (wet weight) of R. sphaeroides cells were thawed and washed three times with 20 mM Tris-HCl, 0.1 mM phenylmethylsulfonyl fluoride, 0.6 mM EDTA, pH 8.0, containing 0.9% NaCl. The washed cells were then resuspended in 10 mM Tris-HCl, 0.1 mM phenylmethylsulfonyl fluoride, 0.6 mM EDTA, pH 8.4, to which 100 mg of lysozyme was added and incubated for 2 h at room temperature. The cell suspension was next sonicated for three 2-min bursts on a Branson Sonifier 450. Spheroplasts and cell debris were pelleted by ultracentrifugation at 45,000 x g for 45 min. Conductivity of the supernatant was reduced to 3 mmho and pH was adjusted to 8.4 by overnight dialysis in the same buffer. This periplasmic extract was loaded on a 2.5 x 45-cm DEAE column and eluted with a linear gradient of 0 to 600 mM NaCl. Fractions containing Me₂SO reductase were next brought to 20% (NH₄)₂SO₄ and run on a phenyl-Sepharose column; enzyme was eluted with 5% (NH₄)₂SO₄ and concentrated. Finally, Me₂SO reductase was further purified by S-200 gel filtration. All reagents and buffers were of the highest quality commercially available and were used without purification. Prior to use, freshly isolated Me₂SO reductase was subjected to "redox cycling" as previously described (18) to eliminate any structural heterogeneity at the active site and stored in liquid nitrogen until needed.

Ultraviolet visible absorption spectra were obtained using a Hewlett-Packard 8452 diode-array spectrophotometer. Rapid reaction, pH jump, and steady-state experiments were performed using an Applied Photophysics Inc. SX.18MV stopped-flow apparatus. Enzyme-monitored turnover, reduction, and pH jump experiments were performed using a photo-diode array detector that allows detection of the full visible spectrum, with absorbance changes monitored using Applied Photophysics software. Transients observed in rapid reaction experiments were collected as transmittance voltages by a high-speed A/D converter and absorbance changes were obtained using Applied Photophysics software. Time courses thus obtained were fitted to sums of exponentials using an iterative nonlinear least-squares Levenberg-Marquardt algorithm. For rapid reaction studies, hyperbolic plots of observed rate constant versus substrate concentration dependence were fitted using the equation, k_obs = k_ox[S]/(K_d + [S]) to obtain the limiting rate constant, k_ox, and the dissociation constant, K_d. The Arrhenius activation energy, E_A, was determined from plots of ln(k_obs) versus 1/T for data obtained over the temperature range 15–35 °C. The slope of the resulting line is equal to –E_A/R (19); H was calculated by the relation E_A = H + RT. S was calculated using the relation: A = k_BT/h exp (S/R).

Anaerobic solutions of oxidized Me₂SO reductase were prepared by placing enzyme in a tonometer equipped with a side-arm cuvette and three-way stopcock fitted with a male Luer connector, followed by repeated evacuation and flushing with O₂-free argon (passed over an oxygen-scrubbing BASF catalyst). The enzyme was then reduced by titrating with sodium dithionite, and monitoring the reaction spectrophotometrically. Steady-state reactions required quantitation of sodium dithionite, which was accomplished by calibration of concentrated dithionite stock solutions against dichloroindophenol. Loss of absorbance at 600 nm, because of dichloroindophenol reduction, was followed spectrophotometrically and the concentration of reducing equivalents thus calculated. The glycerol-inhibited enzyme was prepared as described by Finnegan et al. (6) where dithionite-reduced enzyme in 50 mM Tris-HCl, 0.6 mM EDTA, pH 8.0, is incubated aerobically with 50% (v/v) glycerol. For pH jump experiments, enzyme was first equilibrated in either 10 mM MES, pH 6, or 10 mM CHES, pH 10. The solution was made anaerobic and reduced as described above and then mixed with 100 mM buffer of the opposing pH in a stopped-flow device at room temperature. The kinetics of enzyme reduction was carried out by mixing 100 µM Me₂SO reductase with 0.5 or 25 mM concentrations of sodium dithionite in 50 mM phosphate, 0.6 mM EDTA, pH 6.0, at 5 °C. Again, reactions were monitored using a photo-diode array and rate constants from these studies were obtained by Applied Photophysics software using an iterative nonlinear least-squares Levenberg-Marquardt algorithm. For enzyme-monitored turnover experiments, 100 µM Me₂SO reductase was mixed with a solution of substrate and dithionite in a stopped-flow apparatus. The basic solution for all reaction mixtures used (including that for the enzyme) was 50 mM KH₂PO₄, 0.6 mM EDTA, pH 6.0. The absorption changes seen in the course of the reaction were obtained and analyzed by multicomponent analysis using the Matlab software package (The Math Works Inc.) installed on an SGI computer. The Moore-Penrose pseudoinverse of a matrix of parent spectra (M) was calculated by the equation: C = (M'M)^–1xM'. Relative contributions of parent spectra (P) are calculated by the equation: P = CA, where A represents a matrix of the experimental data (children spectra). Further manipulations were performed using SigmaPlot version 8.0. Because sodium dithionite (which absorbs strongly in the near UV) was used as the reductant, the analysis was restricted to wavelengths above 400 nm. Parallel steady-state reactions for quantitation of EPR intensity were performed in a cooled water bath (5 °C), with aliquots were removed at specific times and quickly frozen in a dry ice/acetone bath (about 2 s). EPR measurements were carried out on a Brüker ESP 300 spectrometer with the following instrument settings: modulation frequency of 100 kHz, modulation amplitude of 5 gauss, sweep width of 300 G (from 3250 to 3550 G), and 2 milliwatt of power. Quantitation of Mo(V) intensity was performed using a bovine xanthine oxidase-2-hydroxy-6-methylpurine standard. Finally, simulations of these reactions were performed using Applied Photophysics software.

	RESULTS AND DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 
pH Dependence of the Absorption Spectrum of Me₂SO Reductase—Prior to examining the pre-steady-state pH dependence of the kinetics of Me₂SO reductase oxidation by Me₂SO, reductive titrations at differing pH values were performed to determine any pH dependence on the visible spectra associated with the three primary intermediates of the enzyme. Fig. 1 shows absorbance spectra of oxidized, substrate-bound, and reduced Me₂SO reductase at pH 6, 8, and 10. The absorption spectrum of oxidized Me₂SO reductase is essentially insensitive to pH (Fig. 1A), displaying only a slight increase in absorbance near the feature observed at 350 nm in going from pH 6 to 10. By contrast, the spectrum of reduced enzyme exhibits a pronounced loss of absorption features as the pH is increased, with clear maxima at 375 and 650 nm seen at pH 6.0 progressively lost until a final, quite different spectrum is observed at pH 10 (Fig. 1C). This pH-induced spectral change observed for the reduced Me₂SO reductase is fully reversed upon equilibration of enzyme in pH 6.0 buffer. The kinetics of the pH-dependent spectral change seen with reduced Me₂SO was followed in experiments in which enzyme in dilute buffer at one extreme in pH was mixed with more concentrated buffer at the other extreme. In the Supplemental Materials, Fig. S1 shows representative traces seen upon changing the pH, following the reaction at 375 nm; rate constants for the low-to-high and high-to-low pH experiments were determined to be 2.10 and 14.9 s^–1, respectively. Additionally, the absorption spectrum for the E_red·Me₂SO complex (generated by treatment of oxidized enzyme with DMS) is found to be pH-dependent, although the absorption changes observed upon equilibration in higher pH buffer are more subtle than those observed in the reduced enzyme. The overall spectral line shape of the substrate-bound species is maintained, with modest bleaching observed centered at 550 nm and a slight increase in long wavelength absorbance above 600 nm as pH is raised (Fig. 1B). The pK_a values associated with the spectral changes observed for the reduced and E_red·Me₂SO intermediates are 9.5 and 8.5, respectively.

View larger version (13K): [in this window] [in a new window]   FIG. 1. pH dependence of UV-visible spectra for Me2SO reductase intermediates. UV-visible absorption spectra obtained for oxidized (panel A), Ered·Me2SO complex (panel B), and reduced (panel C) Me2SO reductase. Spectra were recorded in 50 mM KH2PO4, 0.6 mM EDTA, pH 6.0 (solid line), 50 mM Tris-HCl, 0.6 mM EDTA, pH 8.0 (dashed line), and 100 mM glycine, 0.6 mM EDTA, pH 10.0 (dotted line). In the case of the substrate-bound and reduced species, additional spectra are shown recorded in 100 mM CHES, 0.6 mM EDTA, pH 9.0 (dotted-dashed line). Spectra for oxidized and reduced enzyme were obtained by anaerobic titration using sodium dithionite as reductant, whereas spectra for the Ered·Me2SO complex were obtained by the addition of a stoichiometric excess of dimethylsulfide to the aerobic, oxidized enzyme.

Several previous crystallographic and spectroscopic studies of the reduced Me₂SO reductase provide precedence for dissociation of the Q pterin. The crystal structure determined for reduced enzyme (3) shows the S2' thiolate of the Q pterin at the non-bonding distance of 3.1 Å from the molybdenum, suggesting at least partial dissociation of the Q pterin although it is unclear whether the structure represents the physiologically relevant reduced enzyme. Additionally, EXAFS analysis of R. sphaeroides Me₂SO reductase performed by George et al. (10, 12) suggests only 3 thiolate ligands (in addition to two molybdenum-O/N interactions) to the molybdenum in the reduced enzyme, although the results are admittedly ambiguous. Finally, recent mutagenesis of the active site residue Trp¹¹⁶ to phenylalanine in the R. capsulatus enzyme (31) results in catalytically impeded Me₂SO reductase with an oxidized visible spectrum akin to that of the HEPES-modified enzyme leading the authors to propose a similar monodithiolene, pentacoordinate molybdenum geometry.

The Reaction of Reduced Me₂SO Reductase with Me₂SO— With the pH dependence of the absorption spectra established, the rapid reaction kinetics of reduced Me₂SO reductase with Me₂SO was investigated over the pH range 6–10. The reaction was found to be biphasic under all reaction conditions investigated at pH 8.0 and below, although non-ideal² behavior following substrate binding was observed at high pH that suggests an additional, unresolved phase in the oxidative half-reaction. The fast phase of the reaction could only be monitored easily at relatively low concentrations of Me₂SO, low temperature, and was best observed at pH 6.0, where large extinction changes were associated with the reaction. Representative kinetic transients from the initial, fast phase of the reaction at pH 6.0, as followed at 374 and 550 nm can be seen in Supplemental Materials Fig. S2. Hyperbolic fits to the data at pH 6.0 and 5 °C yielded an extrapolated limiting rate constant at high [Me₂SO] of 1000 s^–1 and an apparent K_d^Me₂SO of 155 µM. This indicates that the E_red·Me₂SO intermediate formed upon addition of Me₂SO results from breakdown of a prior enzyme-substrate complex and does not itself represent the initial Michaelis complex for the reaction. Because of the fast reaction rates and relatively small extinction changes observed at higher pH values, it was not possible to accurately determine the pH dependence of the fast phase of the reaction at higher pH values.

The intermediate formed at the end of the initial phase of the reaction was essentially identical to that observed upon reaction of oxidized enzyme with excess DMS (5, 11, 14, 32), which has been shown to represent the Me₂SO-bound complex. Decay of the E_red·Me₂SO intermediate is much slower than its formation and could be conveniently followed spectrophotometrically at all pH values. Loss of the substrate-bound intermediate (and subsequent gain of oxidized enzyme) was found to be substrate-concentration independent as well as being highly dependent on pH, with a single apparent pK_a of 8.5, the observed pH dependence of the pre-steady-state rate of Me₂SO reduction in good agreement with existing data describing forward steady-state reactivity (5, 7). Rate constants of 38 and 35 s^–1 were obtained for this process at pH 6 and 8, respectively, but a marked decrease in the reaction rate was observed upon further increases in pH, the rate of Me₂SOR_ox formation slowing to 0.5 s^–1 at pH 9.0. Decay of the E_red·Me₂SO complex is largely rate-limiting for the oxidative half-reaction over pH range 6–10 and is undoubtedly the kinetic event measured in the forward steady-state assay. As mentioned previously, the kinetic behavior of this second phase of the reaction becomes increasingly non-ideal as pH increases above 8, concurrent with the observed bleaching of the spectrum for E_red. It appears most likely that this kinetic complexity, which is suggestive of an additional, poorly resolved kinetic process, results from dissociation of the Q pterin from the metal at high pH. This is best evidenced by loss of the clear isosbestic points observed for the decay of the E_red·Me₂SO complex to oxidized enzyme (Fig. 2) above pH 8.0, and may indicate the existence of an additional intermediate complex; possibly E_ox·DMS.

View larger version (22K): [in this window] [in a new window]   FIG. 2. Analysis of the second phase of the reaction of reduced Me2SO reductase with Me2SO. Panel A, seen in the spectra course of the second phase of the reaction of reduced Me2SO reductase with a stoichiometric excess of Me2SO in M 50 m Tris-HCl, 0.6 mM EDTA, pH 8.0, at 25 °C. The Ered·Me2SO is represented by the species complex showing maximal absorbance at 550 nm, which is directly converted to oxidized enzyme as substrate is reduced with clear isosbestic points at 478 and 592 nm. Panel B, semi-log plot of [Ered·Me2SO] versus time for the reaction shown in panel A, where the slope of the line corresponds to kobs (35 s–1). The concentration of the substrate-bound complex was determined by monitoring the fractional absorbance change at 551 nm. Panel C, Arrhenius plot of the second phase of the reaction. All reactions were carried out in 50 mM Tris-HCl, 0.6 mM EDTA, pH 8.0, as described under "Experimental Procedures."

Quantitative Reductive Titrations of Me₂SO Reductase—It has been suggested that the pyranopterin cofactor of molybdenum-containing enzymes may be redox-active, either in the pterin ring itself (i.e. interconverting between di- and tetrahydropterin forms) or in the enedithiolate side chain (22). To ascertain the redox stoichiometry of the molybdenum center of Me₂SO reductase, quantitative titrations were performed with the water-soluble phosphine, PTA, which is known to react with Me₂SO reductase (26) as well as with oxomolybdenum model complexes by a mechanism involving direct oxygen transfer to PTA (24, 25). The reductants, DMS, and sodium dithionite were also used to determine the number of reducing equivalents reversibly taken up by the enzyme and Me₂SO was used as well in the oxidizing direction. Fig. 3 shows the results of titrations using PTA where reduction of enzyme at pH 8.0 was followed at 374, 640, and 720 nm (wavelengths corresponding to absorption bands that possess relatively large extinction changes). All results indicate that both reductive and oxidative processes are strict 2-electron events whether they occur via chemical reduction or through reaction with substrate. That changes in the absorption spectrum are not observed upon addition of additional reducing equivalents is consistent with the recent results of Burgmayer et al. (23) on model systems, and indicates that neither pterin cofactor of Me₂SOR is formally redox-active under physiological conditions, and specifically not during turnover. It was also found that when exposed to oxygen for up to 2 h, the PTA-reduced enzyme formed a distinct, inactive intermediate previously characterized as being formed by prolonged aerobic incubation of DMS-reduced enzyme with excess dimethylsulfide.³ Titrations performed using PTA, however, were performed anaerobically eliminating the possibility of this secondary process altering results.

View larger version (13K): [in this window] [in a new window]   FIG. 3. Quantitative titration of Me2SO reductase with PTA. Reductive titration of Me2SO reductase with PTA was performed in 100 mM Tris-HCl, 0.6 mM EDTA, pH 8.0. Titration was performed by the slow addition of two reducing equivalents of PTA/molybdenum active site using a tonometer made anaerobic as described under "Experimental Procedures." Percent absorbance change is followed at 374 (•), 640 (), and 716 () nm.

Enzyme-monitored Turnover Kinetics of Me₂SO Reductase with TMAO—A spectrophotometric analysis of Me₂SO reductase during turnover with the alternate substrate TMAO was next carried out. These enzyme-monitored turnover experiments were performed using a stopped-flow spectrophotometer equipped with a photodiode array detector (see "Experimental Procedures") and involved mixing anaerobic solutions of oxidized Me₂SO reductase (100 µM) with buffer containing 6 mM TMAO and 25 mM dithionite (5-fold excess TMAO) at 5 °C. Under these conditions, oxidized enzyme is first reduced by dithionite allowing for substrate binding and catalysis, which results in reoxidation of enzyme for a subsequent catalytic cycle. The reaction continues until the limiting substrate (TMAO in the present experiments) has been depleted and the spectrum at completion is expected to be that of fully reduced enzyme. During the extended steady-state that is achieved in the course of the reaction, the absorption spectrum of the enzyme is monitored so that spectra obtained at any given time in the course of the reaction can be deconvoluted into the component (or parent) spectra of oxidized and reduced enzyme, as well as the spectra of any intermediates that may accumulate in the course of the reaction. In this way time courses for each enzyme form can be obtained.

Quantitation of the EPR signal in parallel experiments in which aliquots were withdrawn from the reaction mixture, frozen quickly, and analyzed by EPR, reveal that upon mixture of Me₂SO reductase with 6 mM TMAO and 25 mM dithionite, 100% Mo(V) accumulation persists throughout the majority of the steady-state. The inset of Fig. 4 shows a representative EPR sample generated as above, which gives the catalytically relevant "high-g split" signal (9). The EPR line shape was essentially unchanged throughout the reaction, indicating that no other paramagnetic species accumulated to any significant degree. Significantly, the absorption spectra seen in this portion of turnover thus represents that of the high-g split Mo(V) species.⁴ As seen in Figs. 4 and 5, this spectrum displays broad bands at 379, 550, and 675 nm with extinction coefficients of 5250, 2340, and 1800 M^–1 cm^–1, respectively. The visible absorption features of the high-g split species are similar to those observed for the glycerol-inhibited Mo(V) form of the enzyme, suggesting similar structures (Fig. 4). The glycerol-inhibited enzyme shows bands at 379, 567, and 681 nm with calculated extinction coefficients of 5300, 2120, and 1820 M^–1 cm^–1. Furthermore, the EPR signal of the glycerol-inhibited species is nearly indistinguishable from that of the high-g split signal when the latter is generated in D₂O (6, 12). The loss of hyperfine splitting is because of lack of protonation at the labile oxygen site, present as a Mo-OH at the Mo(V) oxidation state in the catalytic intermediate. Currently, there is some debate as to the structure of the glycerol-inhibited enzyme. Finnegan et al. (6) have concluded that the glycerol-inhibited enzyme has molybdenum coordinated in a bidentate manner to a single pterin moiety (presumably the P pterin), with 1 eq of glycerol (also coordinated in a bidentate fashion through its hydroxyl groups), a Mo = O group and possibly also the serinate ligand. However, a later report (11) describes the molybdenum ligand set as consisting of bidentate coordination to both pterin moieties as well as one glycerol molecule, having displaced the serine ligand contributed by the peptide as well as the molybdenum = O of oxidized enzyme. Based upon the similarity of the UV-visible and EPR spectra of the glycerol-inhibited and high-g split species, we consider the latter interpretation to be more likely. This spectrum is very similar to the spectrum of glycerol-inhibited enzyme (which also has the metal in the Mo(V) oxidation state).

View larger version (17K): [in this window] [in a new window]   FIG. 4. UV-visible spectrum of the high-g split Mo(V) species. Appropriately scaled absorption spectra of oxidized (solid line), glycerol-inhibited Me2SOR-[Mo(V)] (dotted line), and the physiological high-g split Mo(V) species (dashed line) obtained by mixing 100 µM oxidized enzyme with 1 mM sodium dithionite at pH 6.0; absorbance at wavelengths 400 nm and shorter is greater than expected because of small contributions by the reductant. The glycerol-inhibited enzyme was prepared as described by Finnegan et al. (6) in 50 mM Tris-HCl, 0.6 mM EDTA, pH 8.0. Inset, representative EPR spectrum obtained in EMT experiments with TMAO and Me2SO as well as by reduction with sodium dithionite. Instrument settings were modulation frequency of 100 kHz, modulation amplitude of 5 gauss, sweep width of 300 G (from 3250 to 3550 G), and 2 milliwatt of power.

View larger version (16K): [in this window] [in a new window]   FIG. 5. pH dependence of the absorbance spectrum for the Me2SO reductase high-g split Mo(V) intermediate. UV-visible spectra obtained for the high-g split intermediate of Me2SO reductase. Spectra were recorded in 50 mM KH2PO4, 0.6 mM EDTA, pH 6.0 (solid line), 50 mM Tris-HCl, 0.6 mM EDTA, pH 8.0 (dashed line), and 100 mM glycine, 0.6 mM EDTA, pH 10.0 (dotted line), by monitoring the reductive half-reaction using 1 mM sodium dithionite on a stopped-flow apparatus as described under "Experimental Procedures." The spectra shown represent the point of maximal 550 nm absorbance in the course of reaction; spectral deviations are presumed to be a result of the transient nature of the paramagnetic species. The wavelength range used for the Mo(V) intermediate spectra is constrained by dithionite absorbance in the near-UV as well as the long wavelength cut-off inherent to the diode array detector (732.9 nm). Inset, kinetic trace of Me2SOR reduction by 25 mM sodium dithionite at pH 6.0 and 5 °C. The transient was obtained by monitoring the reaction at 551 nm subsequent to rapid mixing of oxidized Me2SO reductase with sodium dithionite at pH 6.0 and 5 °C as described under "Experimental Procedures." Maximal accumulation of 551 nm absorbance corresponds to near complete 1-electron reduction to the high-g split Mo(V) species. Because of non-ideal (Footnote 3) behavior, only the final time points were used to estimate the rate constant of Mo(V) decay (bold line).

View larger version (20K): [in this window] [in a new window]   FIG. 6. Enzyme monitored turnover of Me2SO reductase with TMAO and Me2SO. Panel A, MCA of enzyme-monitored turnover using 50 µM Me2SOR with 3 mM TMAO and 25 mM dithionite showing relative abundance of oxidized (•), reduced (), and the high-g split Mo(V) species () intermediates over the course of reaction. Panel B, MCA of enzyme-monitored turnover using 50 µM Me2SOR with 2 mM Me2SO and 25 mM dithionite showing relative abundance of oxidized (•), reduced (), high-g split Mo(V) species (), and Ered·Me2SO () intermediates over the course of the reaction. Both reactions were carried out in 50 mM KH2PO4, 0.6 mM EDTA at pH 6.0.

Enzyme-monitored Turnover Kinetics of Me₂SO Reductase with Me₂SO—For the deconvolution of the spectra obtained in the course of the reaction of Me₂SO reductase with the physiological substrate Me₂SO (again using dithionite as reductant), a parent spectrum for the E_red·Me₂SO complex, in addition to those for E_ox, E_red, and the high-g split Mo(V) species, was necessary, and was obtained by reaction of oxidized enzyme with excess dimethylsulfide. The fractional abundance of each of the now four parent spectra over the course of the reaction was calculated as described under "Experimental Procedures." It was found, however, that these four spectra were insufficient to accurately describe the spectra observed in the course of the reaction and it was determined that the spectrum used for the Me₂SO-bound enzyme was most likely incorrect. The original parent spectrum for Me₂SO-bound enzyme was obtained by mixing oxidized Me₂SO reductase and up to 100 mM DMS, taking the transient spectrum with maximal 550 nm absorbance as being that for the E_red·Me₂SO species. The spectrum obtained, however, lacked the intense 550-nm absorbance observed in the latter part of the steady-state reaction, which obviously contained the highest proportion of DMS-reduced enzyme on the basis of the absorbance in the 425–610-nm range. Baseline shifts could not account for this discrepancy in the course of turnover. This appeared to be the result of incomplete product-induced reduction as observed by Bray et al. (27). In the eventuality, it was determined that the spectrum obtained at maximal E_red·Me₂SO accumulation in the steady-state reaction more accurately represents the actual spectrum for this species.

As with experiments using TMAO as substrate, parallel EPR reactions were performed, with aliquots of reaction mixture removed and frozen at various time intervals. In contrast to the results seen with TMAO, turnover using Me₂SO as substrate resulted in substantially less accumulation of the high-g split species, an accumulation of 20% occurring near the very end of the steady-state (20 min). Based on an analysis of the UV-visible spectra it was evident that neither oxidized nor reduced enzyme accumulate to any significant degree in the steady-state. The contribution of the high-g split Mo(V) species, using the spectrum determined from the TMAO experiments, was subtracted from the spectrum observed at 20 min and the resulting spectrum was then normalized to obtain the spectrum for the "pure" E_red·Me₂SO intermediate. In the end, the resulting spectrum was taken to best represent the spectrum for authentic E_red·Me₂SO. The difference between this calculated Me₂SO-bound spectrum and that obtained by mixing excess DMS with oxidized enzyme is shown in the inset to Fig. 7. Slightly more absorbance is observed in the 460–575 nm range in the calculated spectrum, with lower extinction seen at wavelengths below 460 nm and above 575 nm. Also shown is the subtraction of fractional amounts (18%) of the oxidized parent from the DMS-reduced intermediate and normalization, demonstrating that a spectrum more closely resembling that estimated from the above multicomponent analysis can be readily obtained, this point further supports the conclusion that there is incomplete reduction of oxidized enzyme by the product dimethylsulfide. The new Me₂SO-bound spectrum allowed for much better fits; the parent spectra used in the eventual multicomponent analysis are shown in Fig. 7.

View larger version (19K): [in this window] [in a new window]   FIG. 7. UV-visible spectra of parents used in MCA of Me2SO reductase. Absorption spectra for oxidized (solid line), reduced (dotted line), Ered·Me2SO (dashed line), and high-g split Mo(V) species (dotted-dashed line), and parents used in MCA of enzyme-monitored turnover using either TMAO or Me2SO. Inset, Ered·Me2SO absorption spectra are as follows: DMS-reduced oxidized enzyme reacted with 100 mM DMS (dotted line), normalized spectrum calculated by subtracting 18% oxidized parent from DMS-reduced (dashed line), and the normalized spectrum used for MCA (solid line), calculated by subtracting 20% of the Mo(V) parent from the spectrum representing the point of substrate exhaustion in EMT with Me2SO.

The calculated abundance of each species from the above analysis reveal that, as in the TMAO reaction, oxidized enzyme is depleted very early in the course of the reaction and that reduced enzyme does not accumulate until the Me₂SO has been exhausted at the completion of the reaction. The 1-electron reduced and substrate-bound enzyme forms accumulate, reflecting the rate-limiting nature of their decay in the catalytic cycle. The pre-steady-state oxidation of enzyme by Me₂SO is biphasic at pH 6.0 and 5 °C, with substrate binding occurring at a rate on the order of 1000 s^–1 and catalysis at 11 s^–1. Both steps in the oxidative half of the catalytic cycle occur much faster than either step in the reductive half of the reaction. For the E_red·Me₂SO complex to accumulate to a greater extent than the high-g split Mo(V) species, it is necessary that the rate of catalysis be no greater than 0.2 s^–1, i.e. the rate of Mo(V) reduction to Mo(IV). Simulation of the reaction mechanism using the rates obtained here suggests Me₂SO turnover at a rate of 0.05 s^–1, which would account for the 80% accumulation of the E_red·Me₂SO complex observed near the completion of the experiment, and reflects a very significant degree of product inhibition. Inhibition of turnover by binding of the product DMS to oxidized enzyme has been reported previously (5, 27) but the present work is the first to quantify the effect of this product inhibition on forward turnover. Accumulation of E_red·Me₂SO by reaction of oxidized enzyme with DMS as the latter accumulates in the course of the reaction is consistent with the known K_d for DMS with oxidized enzyme: concentrations comparable with the literature values for K ^DMS_d of about 120 µM (5, 27), respectively, would be achieved in the present experiments after only a few turnovers, resulting in the accumulation of inhibitory quantities of product. Furthermore, DMS binding to oxidized enzyme has also been reported to occur at rates >500 s^–1 (27), significantly faster than the observed rate of Mo(VI) reduction. We conclude that the inhibition observed here is quantitatively consistent with the known manner in which DMS interacts with oxidized enzyme and emphasizes the thermodynamic stability of the substrate-bound species.

Our mechanistic results can be incorporated into a comprehensive kinetic mechanism for the enzyme, as shown in Scheme 1. The sequential reduction of Mo(VI) to Mo(V) and Mo(V) to Mo(IV) by sodium dithionite occurs with rate constants of 1 (k₁) and 0.2 s^–1 (k₂), respectively, at pH 6.0 and 5 °C. The rate-limiting nature of Mo(V) reduction is reflected in near 100% accumulation of the EPR-active (high-g split) intermediate during turnover with TMAO. Scheme 1A shows the reaction with TMAO where no substrate-bound intermediate was detected, reduction of substrate proceeding directly from the Michaelis complex. Scheme 1B summarizes the results for the reaction with Me₂SO at low pH where the mechanistic behavior is more easily rationalized. Me₂SO binding to form the substrate-bound species proceeds via the Michaelis complex and is rapid, with an estimated rate constant on the order of 1000 s^–1 for k₄. At lower pH, the kinetics of Me₂SO reduction following substrate binding (k₅) are conveniently followed, with a rate constant of 11 s^–1 determined at pH 6.0 and 5 °C. As the concentration of DMS increases in the enzyme-monitored turnover experiment with Me₂SO, the back reaction (k_–5) inhibits forward catalysis as evidenced by the accumulation of the Me₂SO-bound intermediate. Asterisks in Scheme 1B indicate likely deviations in enzyme behavior at high pH. At higher pH the reduction to Mo(V) occurs at approximately the same rate as at pH 6.0 with reduction of the Mo(V) species, k₂, slowing modestly to 0.08 s^–1. At pH 10, the enzymatic reduction of Me₂SO (k₅) slows to 0.08 s^–1 at 5 °C, although the catalytic mechanism is less clear because of the non-ideal kinetic behavior of the system. Our results suggest that structural changes in the reduced enzyme impedes catalytic activity at high pH. Dissociation of the Q pterin would thus necessitate accommodation of an additional oxygen ligand, presumably by maintaining coordination to the labile oxo group. The structural change in reduced Me₂SO reductase may perturb the equilibrium between reduced, Me₂SO-bound and oxidized, DMS-bound enzyme upon substrate binding resulting in slower rates of turnover and product release (k₅).

View larger version (21K): [in this window] [in a new window]   SCHEME 1. Proposed mechanistic schemes for reaction of Me2SO reductase with TMAO and Me2SO. Literature values of Km and kcat (k3) determined for the steady-state reaction of enzyme with TMAO are 194 µM and 134.5 s–1, respectively (29). Asterisks in B indicate sites of likely deviation between turnover at low and high pH.

	CONCLUSIONS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

In rapid kinetic work, we find the reaction of reduced Me₂SO reductase with Me₂SO to be biphasic at low pH, with the absorption spectrum of the intermediate seen at the conclusion of the fast phase being that of the well characterized E_red·Me₂SO complex. This fast phase of the reaction has not been previously observed; the rate constant for this phase of the reaction has a hyperbolic dependence on [Me₂SO] with a limiting rate constant for complex formation approaching 1000 s^–1 and a dissociation constant for the preceding Michaelis complex of 155 µM. This behavior is consistent with the formation of a prior E·S complex, implying that the spectroscopically distinct E_red·Me₂SO complex does not itself represent the Michaelis complex for the oxidative half-reaction of the catalytic cycle but instead lies downstream from it in the catalytic sequence. Once formed, the E_red·Me₂SO complex decays with a rate constant that is independent of [Me₂SO] but highly pH-dependent, slowing from 38 s^–1 at pH 6 to 0.5 s^–1 at pH 9.0, with pK_a of 8.5 comparable with that previously determined for overall steady-state turnover (5, 7). The temperature dependence of the decay of the E_red·Me₂SO complex yields an entropy and enthalpy of activation comparable with the values seen in the reactions of inorganic model systems for molybdenum-containing enzymes such as Me₂SO reductase, with an "early" transition state involving little formal oxidation of the molybdenum.

In enzyme-monitored turnover experiments, following the absorption and EPR spectra of enzyme in the course of turnover with dithionite as reductant, we have succeeded in fully defining the distribution of enzyme species with both Me₂SO and TMAO as oxidizing substrates. In the case of reaction with TMAO, the physiological relevant EPR-active species giving rise to the high-g split signal predominates in the steady-state, providing a convenient method for generating samples with consistently high Mo(V) content. This species is also seen in the course of turnover with Me₂SO, although it accumulates to a lesser degree. In the latter experiments, DMS accumulates in the course of turnover and eventually reaches concentrations sufficient to appreciably retard turnover by rebinding to oxidized enzyme and shifting the E_ox·DMS = E_red·Me₂SO equilibrium toward the latter species.

	FOOTNOTES

 
^* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains Figs. S1–S4.

Current address: Laboratory of Proteomics and Analytical Technologies, SAIC-Frederick, Inc., National Cancer Institute at Frederick, P. O. Box B, Frederick, MD 21702-1201.

^¶ To whom correspondence should be addressed: Dept. of Molecular and Cellular Biochemistry, The Ohio State University, 333B Hamilton, 1645 Neil Ave., Columbus, OH 43210. Tel.: 614-292-3545; Fax: 614-292-4118; E-mail: hille.1{at}osu.edu.

^{1 1} The abbreviations used are: Me₂SO, dimethyl sulfoxide; Me₂SOR, dimethyl sulfoxide reductase; Rc, Rhodobacter capsulatus; CAPS, 3-(cyclohexylamino)-1-propanesulfonic acid; DMS, dimethylsulfide; PTA, phosphatriazaadamantane (1,3,5-triaza-7-phosphatricyclo[3.3.1.1]decane); MCA, multiple component analysis; MES, 4-morpholineethanesulfonic acid; TMAO, trimethylamine N-oxide.

² Above pH 8 the behavior of the oxidative transient for E_red·Me₂SO decay become increasingly non-ideal. This is also evidenced by loss of the tight isosbestic points as seen at lower pH values (Fig. 2). The rate of substrate binding appears not to be effected as pH increases in that the characteristic substrate-bound intermediate is observed in the dead-time of the diode-array instrument. This suggests the possibility of an additional kinetic process in the reduction of Me₂SO, which becomes rate-limiting as pH increases above 8. Presumably, this would be best represented by product release from an E_ox·DMS complex and further examination of this phenomenon will be carried out.

³ When Me₂SO reductase was reacted with a pseudo first-order excess of PTA under aerobic conditions, the enzyme rapidly became fully reduced, and in the presence of molecular oxygen, converted to a form exhibiting the absorption spectrum shown in Supplemental Materials (Fig. S4) within 2 h. Clear isosbestic points are observed in the formation of the PTA-modified enzyme indicating that no intermediates accumulate in the course of reaction. This modified form of the enzyme is nearly identical to the previously characterized, DMS-modified (Me₂SOR_modD) form, generated when oxidized enzyme is aerobically incubated with excess DMS for 22 h (27). The structure of Me₂SOR_modD determined crystallographically shows a pentacoordinate molybdenum center in a distorted square pyramidal geometry with the four dithiolene sulfurs forming the base of the pyramid and the serinate ligand at the apex, the molybdenum being in the +VI oxidation state (27). Me₂SOR_modD is catalytically inert but can be readily redox-cycled via reduction in the presence of methyl or benzyl viologen to regenerate functional enzyme. A more detailed treatment of the data may be found in the Supplemental Materials.

⁴ Bastian et al. (33) describe a similar visible spectrum corresponding to the high-g split Mo(V) intermediate formed by incubation of enzyme with NO and ascorbate. However, rather than a "physiologically relevant" catalytic intermediate, this species is proposed to be the result of NO binding to either the molybdenum or some protein sulfhydryl group. A second spectrum, observed after incubation of reduced enzyme with substrate, Me₂SO, is also similar to that defined here as the high-g split species but was said to be only 30% EPR active. Comparison of the documented extinction coefficients with our own show that the local maxima described herein are appreciably higher, indicative of incomplete formation of 1-electron reduced enzyme by Bastian et al. (33), which may have been caused by contamination of the reaction mixture by molecular oxygen.

⁵ In following the reduction of oxidized Me₂SO reductase with sodium dithionite at 551 nm, an initial increase in absorbance is observed (reflecting formation of the Mo(V) species) followed by a transient loss in absorbance (as enzyme accepts a second reducing equivalent to become fully reduced). At pH 6.0, decay of the high-g split Mo(V) species to reduced enzyme containing Mo(IV) displays a distinct lag phase. Still, once the reaction has progressed beyond the lag phase, the subsequent decay of the Mo(V) species is exponential, and fits to the data can be used to obtain an effective rate of reduction to the Mo(IV) state. The possibility exists that this behavior reflects some artifact of reduction (HSO^–₃ dissociation after initial reductive event), although some structural rearrangement of the molybdenum center or within the pterin cofactors, allowing for a more thermodynamically favorable reduction to Mo(IV), cannot be ruled out and will be the focus of future investigations.

	ACKNOWLEDGMENTS

 
We thank Prof. R. H. Holm, Department of Chemistry, Harvard, for the generous gift of the substrate PTA.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION CONCLUSIONS REFERENCES

 

1. Turner, S. M., Nightingale, P. D., Spokes, L. J., Liddicoat, M. I., and Liss, P. S. (1996) Nature 383, 513–517
2. Coale, K. H., Johnson, K. S., Fitzwater, S. E., Gordon, R. M., Tanner, S., Chavez, F. P., Feroli, L., Sakamoto, C., Rogers, P., Millero, F., Steinberg, P., Nightingale, P., Cooper, D., Cochlan, W. P., Landry, M. R., Constantinou, J., Rollwagen, G., Trasvina, A., and Kudela, R. (1996) Nature 383, 495–501[CrossRef]
3. Schindelin, H., Kisker, C., Hilton, J., Rajagopalan, K. V., and Rees, D. C. (1996) Science 272, 1615–1621[Abstract]
4. Shaw, A. L., Hochkoeppler, A., Bonora, P., Zannoni, D., Hanson, G. R., and McEwan, A. G. (1999) J. Biol. Chem. 274, 9911–9914[Abstract/Free Full Text]
5. Adams, B., Smith, A. T., Bailey, S., McEwan, A. G., and Bray, R. C. (1999) Biochemistry 38, 8501–8511[CrossRef][Medline] [Order article via Infotrieve]
6. Finnegan, M. G., Hilton, J., Rajagopalan, K. V., and Johnson, M. K. (1993) Inorg. Chem. 32, 2616–2617[CrossRef]
7. Bastian, N. R., Kay, C. J., Barber, M. J., and Rajagopalan, K. V. (1991) J. Biol. Chem. 266, 45–51[Abstract/Free Full Text]
8. Benson, N., Farrar, J. A., McEwan, A. G., and Thomson, A. J. (1992) FEBS Lett. 307, 169–172[Medline] [Order article via Infotrieve]
9. Bennett, B., Benson, N., McEwan, A. G., and Bray, R. C. (1994) Eur. J. Biochem. 225, 321–331[Medline] [Order article via Infotrieve]
10. George, G. N., Hilton, J., Temple, C., Prince, R. C., and Rajagopalan, K. V. (1999) J. Am. Chem. Soc. 121, 1256–1266[CrossRef]
11. Garton, S. D., Hilton, J., Hiroyuki, O., Crouse, B. R., Rajagopalan, K. V., and Johnson, M. K. (1997) J. Am. Chem. Soc. 119, 12906–12916[CrossRef]
12. George, G. N., Hilton, J., and Rajagopalan, K. V. (1996) J. Am. Chem. Soc. 118, 1113–1117[CrossRef]
13. Baugh, P. E., Garner, C. D., Charnock, J. M., Collison, D., Davies, E. S., McAlpine, A. S., Bailey, S., Lane, I., Hanson, G. R., and McEwan, A. G. (1997) J. Biol. Inorg. Chem. 2, 634–643[CrossRef]
14. McAlpine, A. S., McEwan, A. G., and Bailey, S., (1998) J. Mol. Biol. 275, 613–623[CrossRef][Medline] [Order article via Infotrieve]
15. Schneider, F., Lowe, J., Huber, R., Schindelin, H., Kisker, C., and Knablein, J. (1996) J. Mol. Biol. 263, 53–69[CrossRef][Medline] [Order article via Infotrieve]
16. McAlpine, A. S., McEwan, A. G., Shaw, A. L., and Bailey, S. (1997) J. Biol. Inorg. Chem. 2, 690–701[CrossRef]
17. Li, H. K., Temple, C., Rajagopalan, K. V., and Schindelin, H. (2000) J. Am. Chem. Soc. 122, 7673–7680[CrossRef]
18. Bray, R. C., Adams, B., Smith, A. T., Bennett, B., and Bailey, S. (2000) Biochemistry 39, 11258–11269[Medline] [Order article via Infotrieve]
19. Tinoco, I., Sauer, K., and Wang, J. C. (1985) in Physical Chemistry, Principles and Applic