PCR amplification of Rv2540c DNA sequence (putative aroF gene) from genomic M. tuberculosis DNA yielded a fragment with the expected length (1206 bp). The fragment was cloned into pET23a(+) expression vector, and the aroF gene was sequenced, which confirmed its identity and the absence of PCR (Polymerase Chain Reaction)-introduced mutations. A number of E. coli host strains were employed to express MtCS, including BL21(DE3), Origami(DE3), and BL21trxB(DE3), but no expression could be obtained. Different cultivating temperatures (25°, 30° and 37°C) and presence or absence of isopropyl β-D-thiogalactoside (IPTG) were also employed to no avail (data not shown). MtCS overexpression could only be achieved in E. coli Rosetta(DE3) host cells, which provide tRNAs for codons that are rarely used by E. coli. In agreement, ten rare codons were identified on M. tuberculosis aroF sequence (7 × CCC, AUA, CUA, GGA). Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) analysis revealed a cell extract containing a significant amount of a soluble protein with an apparent molecular weight in agreement with the predicted value based on amino acid sequence for MtCS (41.8 kDa) (Figure 2A).

Interestingly, recombinant protein expression was achieved in E. coli Rosetta(DE3) host cells grown for 24 h at 37°C in the absence of IPTG induction. It has been previously shown that the lac-controlled systems, including the pET system, could have a high-level protein expression, in the absence of inducer. This phenomenon occurs in the cell stationary phase, when there is the presence of a complex medium, cyclic AMP, acetate and low pH 19. Recently, another shikimate pathway enzyme from M. tuberculosis, 3-Deoxy-D-arabino-heptulosonate 7-phosphate (DAHP) synthase, was reported to be expressed only in the absence of IPTG 20. The three-step purification protocol yielded approximately 55 mg of MtCS from 6 liters of cell culture. No other protein can be observed on SDS-PAGE, which indicates the high level of MtCS purity (Figure 2B). The MtCS activity before the purification can hardly be measured, probably due to interfering compounds, and thus only the amount of total recombinant protein could be assessed. The ammonium sulfate precipitation of purified MtCS resulted in total inactivation of the enzyme (data not shown). Homogeneous recombinant protein was thus instantaneously frozen in liquid nitrogen in the absence of ammonium sulfate, which proved to be a convenient protocol for active homogeneous recombinant MtCS protein storage.

The ESI-MS of homogeneous MtCS revealed a subunit molecular mass of 41,804 Da, consistent with the theoretical molecular mass (41,792 Da). The first 16 N-terminal amino acid residues were identified as MLRWITAGESHGRALV, which confirmed the identity of homogeneous MtCS and presence of N-terminal methionine residue. Analytical gel filtration chromatography revealed a single peak of approximately 104 kDa, suggesting that MtCS is a dimer in solution, which is in agreement with the hydrodynamic properties of the recombinant protein assessed by sedimentation velocity and sedimentation equilibrium 18.

The synthesis of EPSP was carried out in a vial containing the enzymes MtSK and MtEPSPS, which convert shikimate, ATP and phosphoenolpyruvate (PEP) to 5-enolpyruvylshikimate-3-phosphate (EPSP), ADP and Pi. The equilibrium of the forward reaction was displaced using Purine Nucleoside Phosphorylase (PNP), which consumes Pi, increasing the final concentration of EPSP in the reaction mixture. The enzymes were removed by ultrafiltration to avoid any residual MtEPSPS enzyme activity that could release Pi in solution, which would thus interfere with specific measurements of MtCS enzyme activity. Conversion of EPSP to chorismate and Pi catalyzed by MtCS enzyme activity (Figure 1) was determined by measuring the release of Pi using PNP, which converts 2-amino-6-mercapto-7-methylpurine ribonucleoside (MESG) and Pi to ribose-1-phosphate and 7-methyl-6-thio-guanine base, monitoring absorbance of the latter at 360 nm. The enzymatic activity of homogenous MtCS was dependent on enzyme volume added to the reaction mixture (data not shown), showing that the initial velocity is proportional to total enzyme concentration and that true initial velocities are being measured. The conversion of EPSP to chorismate in aerobic conditions by the addition of FMN_ox and NADH is the first solid evidence that MtCS is bifunctional, since the monofunctional CSs can only be assayed under strictly anaerobic conditions in the presence of chemically or enzymatically reduced flavin 16. The specific activity of bifunctional MtCS (0.004 U.mg^-1) is approximately 175-fold lower than the specific activity of bifunctional Neurospora crassa CS (NcCS; 0.7 U.mg^-1) 21. In the presence of oxygen the CS activity is limited by the reoxidation of FMN_red cofactor; but in bifunctional CSs, NAD(P)H is used in consecutive cycles to maintain the FMN in its reduced form (Figure 1). Moreover, the structures of CS complexed with FMN from Helicobacter pylori 22 and Streptococcus pneumoniae 23 have shown that there are a number of positively charged amino acids in the FMN binding pocket, which could increase the reduction potential of FMN_ox/FMN_red couple, likely permitting the decrease of the rate of reoxidation. Indeed, in bifunctional NcCS the redox potential of the couple FMN_red/FMN_ox was determined to be -167 mV, i.e. 40 mV more positive than that of the free couple in solution (-207 mV) 24.

The NADH-dependent reduction of FMN_ox was measured in the forward reaction monitoring the decrease in NADH concentration. The NADH:FMN-oxidoreductase activity of homogenous MtCS is dependent on the enzyme volume added to the reaction mixture; therefore the initial velocity is proportional to total enzyme concentration, and true initial velocities are being measured. To verify the stability of the MtCS-FMN_ox complex, MtCS was incubated with excess FMN_ox and loaded on an anion exchange column. Two peaks were observed and collected into separated fractions measuring absorbance at 280 nm and 445 nm, and pooled (Figure 3). To roughly estimate the presence of protein in the two pools, coomassie-blue was added to them and only the second pool was shown to contain protein (indicating presence of recombinant MtCS), while the first was not stained (indicating absence of recombinant MtCS). The UV-Vis spectrum for the second pool containing recombinant MtCS protein showed that all the characteristic absorbance peaks of FMN_ox (267, 373, and 445 nm) could be observed (Figure 3 inset). These results suggest that MtCS and FMN_ox form a stable binary complex.

Although the FMN_ox usually appears as a prosthetic group in NAD(P)H:FMN oxidoreductases from other organisms 25, it was not clear whether in MtCS the FMN is covalently bound. CSs from other organisms, such as that from E. coli, showed low affinity for FMN_ox; but the value of Kd for FMN_ox depends strongly on EPSP binding, decreasing ca. 1000-fold in the presence of this substrate (from 30 μM to 20 nM) 26. On the other hand, EPSP has a much smaller effect on the affinity of bifunctional NcCS to FMN_ox 21. Interestingly, the purified MtCS is FMN-free, which could be an effect of the insufficient intracellular concentration of FMN, probably due to MtCS overexpression into the host cell. As described below, FMN appears to have a value lower than 20 μM for the overall dissociation constant for MtCS-FMN binary complex formation at equilibrium. Accordingly, it appears to be more likely that FMN is lost during the protein purification protocol because of a rather weak affinity of MtCS for FMN. Accordingly, the kinetic data were collected considering the holoenzyme as the MtCS-FMN_ox complex brought about by incubating MtCS and near-saturating FMN_ox concentration.

The oxidoreductase activity is functionally independent of CS activity, since the consumption of NADH occurs in the absence of EPSP. Considering the low specific activity of NADH:FMN-oxidoreductase in MtCS, all measurements were made in triplicate, including the controls. Accordingly, the apparent kinetic parameters for the oxidoreductase activity were determined in the absence of EPSP in steady-state kinetic experiments. No activity was observed when NADPH was used at concentrations ranging from 100 to 500 μM (data not shown). Kinetics of reduction of the MtCS-bound FMN_ox by NADH was determined using fixed amount of holoenzyme and varying levels of NADH (Figure 4). The data were fitted to equation 1, and yielded the following values: K_NADH = 36 ± 4 μM, k_cat = 8.3 (± 0.3) × 10^-3 s^-1 and k_cat/K_NADH = 231 M^-1 s^-1. The K_NADH observed for MtCS is similar to the K_NADPH determined for NcCS 24 and ScCS 27 (43 μM and 70 μM, respectively). In other NAD(P)H:FMN oxidoreductases (EC 1.5.1.30), which are present in various organisms, the apparent kinetic constants show significant variation. For instance, the K_NAD(P)H can vary from 208 μM in Rhodococcus erythropolis 28 to 0.85 μM in Bacillus subtilis 29.

The binding of FMN_ox to MtCS was assessed by UV-visible difference spectra as described by Kitzing et al. 21. The observed spectral changes are characterized by hypochromic effects on the flavin absorbance at both 378 nm and 450 nm up to 20 μM FMN_ox (Figure 5). At FMN_ox concentrations larger than 30 μM there is a hyperchromic effect that is probably due to free oxidized FMN in solution (Figure 5 – inset). The spectral changes at low FMN_ox concentrations yielded low values for the difference spectra and thus no reliable data could be obtained at concentrations lower than 10 μM. Accordingly, the data presented here allow only to estimate an upper limit value of 20 μM for the FMN_ox equilibrium dissociation constant. We have also tried to determine the dissociation constant of FMN_ox from protein fluorescence titration as recently described by Broco et al 30, which could allow reliable measurement of oxidized FMN binding to MtCS at low concentrations of FMN_ox. However, there was no quench/enhancement in protein fluorescence upon FMN_ox binding to MtCS (data not shown). At any rate, the UV-visible difference spectra show MtCS-FMN_ox binary complex formation.

The equilibrium binding of NADH to MtCS was assessed by fluorescence spectroscopy. There is a quench in protein fluorescence upon NADH binding to MtCS (Figure 6 – inset). A titration curve showing the quench in protein fluorescence upon the binding of NADH to MtCS is given in Figure 6. The data were fitted to a hyperbolic equation yielding a dissociation constant value of 156 ± 12 μM. Even though the NADH-binding site remains unknown, the results presented here clearly indicate that there is indeed an NADH binding site on MtCS enzyme.

To probe for rate-limiting steps and determine the stereospecificity of hydride transfer, primary deuterium kinetic isotope effects were determined. Initial velocity data were collected using either [4S-¹H]NADH or [4S-²H]NADH (Figure 7), and values of 3.5 ± 0.2 and 3.0 ± 0.4 were determined for, respectively, ^DV and ^DV/K. The magnitude of the primary isotope effects when the [4S-²H]NADH substrate is used indicates that C₄-proS hydrogen is being transferred during the reduction of FMN_ox catalyzed by MtCS. Isotope effects on V are related to events following the formation of the complex capable of undergoing catalysis (MtCS-FMN_ox-NADH in the case studied here), including chemical steps, possible enzyme conformational changes, and product release. Isotope effects on V/K report on steps in the reaction mechanism from the binding of the labeled substrate to the first irreversible step, usually product release 31. The values for primary deuterium isotope effects of biochemical interest usually range from 3 to 7 32, and steps such as conformational changes accompanying hydride transfer and product release may account for the lower values. In particular, primary deuterium kinetic isotope effects typically range from 1 to 3 for enzyme reactions involving NAD(P)H oxidation 33. The ^DV value obtained for MtCS-FMN_ox (3.5) suggests that the hydride transfer contributes significantly to the rate-limiting step of the FMN reduction reaction. The value of 3.0 for the ^DV/K indicates that NADH is not sticky, since ^DV/K values for sticky substrates are around unity 34. It should be remembered that a substrate is sticky if it reacts to give products as fast as, or faster than, it dissociates from the enzyme. The stickiness of a substrate depends on the external part of its commitment: the larger the external commitment, the sticker the substrate 34.

Solvent isotope effects were employed to assess the contribution of solvent proton transfer to the rate of flavin reduction. Initial velocity data were collected in either H₂O or 91 atom % D₂O (Figure 8), and values of 1.7 ± 0.3 and 1.3 ± 0.1 were obtained for ^D2OV and ^D2OV/K, respectively. The magnitude of the solvent isotope effects indicate that proton transfer partially limit the rate of FMN reduction reaction. In order to determine the number of kinetically important protons transferred during the reaction, a proton inventory on V was carried out. The mole fraction of D₂O was varied from 0 to 80%, and a linear relationship between V and the mole fraction of D₂O was observed (Figure 8, inset). This result indicates that a single proton transfer gives rise to the observed solvent isotope effect 35.

Significantly different magnitudes between primary and solvent kinetic isotope effects may indicate that they are reporting on distinct steps of a reaction, as reported for the NADPH-dependent ketoacyl-ACP reductase from S. pneumoniae 36. The primary isotope effect values (^DV = 3.5 and ^DV/K = 3.0) are larger than those for solvent isotope effects (^D2OV = 1.7 and ^D2OV/K = 1.3), suggesting that proton and hydride transfer may take place in two distinct transition-states, with the latter being more rate-limiting for the MtCS-catalyzed reaction. Multiple isotope effects studies are able to distinguish whether two different isotopic substitutions affect the same or different chemical steps. Theory predicts that if protonation and hydride transfer occur in the same transition state, the primary isotope effects will be larger or unchanged with D₂O as compared to H₂O. On the other hand, if hydride transfer and protonation occur in distinct steps, the primary isotope effects will be smaller with D₂O as solvent, as proton transfer will become more rate limiting 3738. Multiple isotope effects were thus evaluated to distinguish whether two different isotopic substitutions affect the same or different chemical steps, i.e., if the reduction of FMN_ox catalyzed by MtCS occurs, respectively, in a concerted or in a stepwise mechanism. Initial velocity data were collected and values of 1.3 ± 0.1 and 2.2 ± 0.1 were obtained for ^D2OV_[4S-2H]NADH and ^D2OV/K_[4S-2H]NADH, respectively (Figure 7, inset). The values for the primary kinetic isotope effects on V and V/K in D₂O were smaller than the effects in H₂O, thereby suggesting a stepwise mechanism for the reduction of FMN_ox. A mechanism for bifunctional NcCS has been proposed and involves an electron transfer from FMN_red to C1 EPSP and C-O bond cleavage, protonation of the leaving phosphate group by His17, tautomerization of the resulting C4(a)-neutral flavin semiquinone to a radical species with concomitant abstraction of the C-(6proR) hydrogen of the dephosphorylated substrate intermediate, and reduced flavin deprotonation restores the initial state of the cofactor 24. More recently, it has been shown for NcCS that the carboxylate group of Asp367 participates in the water-mediated deprotonation of the N(5) atom of the isoalloxazine ring system of FMN_red that is involved in abstraction of C-(6proR) hydrogen of EPSP 39. A comparison of the primary sequences of CSs from different sources has shown that these residues (His11 and Asp343, M. tuberculosis numbering) are conserved 18. Interestingly, His11 and Asp343 are present in monofunctinal and bifunctional CSs and across all known species 40, and these residues are part of the characteristic CS signature sequence 23. The role of His11 and Asp343 in the mode of action of bifunctional MtCS should await site-directed mutagenesis studies that are currently underway in our laboratory.

Pfu DNA polymerase was from Stratagene. The pET23a(+) expression vector and E. coli Rosetta(DE3) host cell were from Novagen. All chromatographic supports, including the molecular weight calibration kits, were purchased from GE Healthcare. The protease inhibitor cocktail was from Roche. Purine nucleotide phosphorylase (PNP, EC 2.4.2.1) and 2-amino-6-mercapto-7-methylpurine ribonucleoside (MESG) were purchased from Molecular Probes. NADH, NAD⁺, oxidized FMN (FMN_ox), ATP, phosphoenolpyruvate (PEP), Leuconostoc mesenteroides glucose-6-phosphate dehydrogenase (type XXIII) and yeast hexokinase (type C-300) were from Sigma Chemical Co. D-glucose-1-d (97 atom % D) was from Aldrich and deuterium oxide (99.9 atom % D) was from Cambridge Isotope Laboratories.

Synthetic oligonucleotide primers complementary to amino-terminal coding and carboxy-terminal noncoding strands of aroF (Rv2540c) gene (5' ggtcatatgttgcgctggatcaccgcgg 3' and 5' cggatcctcaaccggagacccgcgcggc 3', respectively) were designed based on the complete genome sequence of M. tuberculosis 13. These primers, containing 5'NdeI and 3'BamHI restriction sites, in bold, were used to amplify aroF-CS encoding gene (1,206 bp) from M. tuberculosis genomic DNA, using Pfu DNA polymerase and standard PCR conditions. The amplified fragment was purified by low melting agarose electrophoresis, digested with NdeI and BamHI, and cloned into pET23a(+) expression vector previously digested with the same restriction enzymes. The aroF gene identity and the absence of PCR-introduced mutations were confirmed by DNA sequencing. The pET23a (+)::aroF was transformed by electroporation into E. coli Rosetta(DE3) host cells and selected on LB agar plates containing 50 μg.mL^-1 carbenicillin and 34 μg.mL^-1 chloramphenicol. Single colonies were used to inoculate 1 L of Luria-Bertani medium containing the same antibiotics and grown for 24 h at 37°C at 180 rpm with no isopropyl β-D-thiogalactoside (IPTG) induction. The cells (4.5 g) were harvested by centrifugation at 3,000 g for 30 min at 4°C.

All purification procedures were performed at 4°C. Cells (25 g) were suspended in 75 mL of 50 mM Tris-HCl, pH 7.8 (buffer A) containing proteinase inhibitor cocktail and 0.2 mg.mL^-1 lisozyme; and the mixture was stirred for 30 min. The cells were disrupted by sonication and centrifuged at 48,000 g for 30 min to remove cell debris. Streptomycin sulfate was added to the supernantant to a final concentration of 1% (w/v) and the mixture was stirred for 30 min. The soluble fraction was collected by centrifugation at 48,000 g for 30 min and dialyzed 3 times against 2 L of buffer A. This sample was clarified by centrifugation (48,000 g for 30 min), loaded on a Q-Sepharose Fast Flow (2.6 cm × 8.2 cm) anion exchange column previously equilibrated with buffer A and the proteins eluted using a linear gradient from 0.0 to 0.5 M NaCl. The fractions containing MtCS were pooled and ammonium sulfate was added to a final concentration of 0.8 M. This sample was loaded on a High Load Phenyl-Sepharose (1.6 cm × 10 cm) hydrophobic interaction column pre-equilibrated with 50 mM Tris-HCl, pH 7.8, 0.8 M (NH₄)₂SO₄(buffer B). The proteins were fractionated using a linear gradient from 0.8 to 0.0 M (NH₄)₂SO₄. The active fractions were pooled and dialyzed 3 times against 2 L of buffer A. The sample was loaded on a MonoQ (1.6 cm × 10 cm) anion exchange column previously equilibrated with buffer A, and the proteins eluted using a linear gradient of NaCl (0.0 – 0.5 M). The fractions containing homogeneous MtCS were pooled, quickly frozen in liquid nitrogen and stored at -80°C. Samples of each purification step were analyzed by SDS-PAGE 44. Protein concentration was determined by Bradford method 45, using Bio-Rad protein assay kit (Bio-Rad) and bovine serum albumin as standard.

Recombinant MtCS was analyzed using mass spectrometry in an adaptation of Chassaigne and Lobinski systems 46. Samples were analyzed on a triple quadrupole mass spectrometer, model QUATRO II, equipped with a standard electrospray (ESI) probe (Micromass, Altrinchan) adjusted to ca. 250 μL.min^-1. The source temperature (80°C) and needle voltage (3.6 kV) were maintained constant throughout the data collection, applying a drying gas flow (nitrogen) of 200 L.h^-1 and a nebulizer gas flow of 20 L.h^-1. The equipment was calibrated with intact horse heart myoglobin. Approximately 50 pmol of sample was injected into electrospray transport solvent to determine the subunit molecular mass of MtCS. The N-terminal amino acid residues of purified recombinant MtCS were identified by automated Edman degradation sequencing using a PPSQ 21A gas-phase sequencer (Shimadzu).

The molecular mass of native MtCS was determined by gel filtration using Superdex S-200 (10 mm × 30 cm) column eluted with buffer A containing 200 μM NaCl at 0.4 mL.min^-1. The protein elution was monitored at 280 nm. The protein molecular weight standards were from Low Molecular Weight and High Molecular Weight Calibration kits (GE Healthcare).

Since there is no ESPS commercially available, EPSP was synthesized using the enzymes shikimate kinase (MtSK) and EPSP synthase (MtEPSPS) from M. tuberculosis 4748, and PNP 4950. The synthesis was carried out using 9.6 mM shikimate, 2.4 mM ATP, 3 mM PEP, 0.4 mM MESG. This reaction mixture was pre-incubated in 50 mM Tris-HCl, 2.5 mM MgCl₂, 2.5 mM KCl pH 7.6 for 3 min at 25°C. Then the three enzymes were added (2.2 U MtSK, 0.7 U MtEPSPS, 2 U PNP) and the reaction mixture was incubated further at 25°C for 30 min. The enzymes were then removed by ultrafiltration using a Centricon 3000 Da cut-off (Amicon). The filtrated solution, containing the substrate EPSP, was collected and used to measure chorismate synthase activity. The reaction catalyzed by EPSP synthase releases an inorganic phosphate molecule, which is consumed by PNP. This coupled reaction changes the equilibrium constant of ESPS synthase reaction, allowing synthesis of higher concentration of EPSP. One unit (U) of enzyme activity for all enzymes cited in this work is defined as the amount of enzyme that catalyzes the conversion of 1 μmol of substrate per minute at 25°C in an optical path of 1 cm.

The MtCS assay was performed in the forward direction using 15 μL of ESPS-containing solution (see above), 0.04 mM FMN_ox, 0.3 mM NADH, 0.2 mM MESG, 1 U PNP in buffer A. The reaction mixture was incubated for 4 min at 25°C for total consumption of Pi by PNP, and MtCS was then added. The CS reaction was monitored using a coupled reaction with the PNP enzyme 50. This enzyme catalyses the cleavage of MESG by the inorganic phosphate molecule which is released by MtCS, and formation of 2-amine-6-mercapto-7-methylpurine is monitored at 360 nm (ε = 11.0 × 10³M^-1 cm^-1). The assay was performed at 25°C and monitored using a Shimadzu UV-2550 spectrophotometer. All measurements, including the blanks, were carried out in triplicate to ensure that reliable data were being collected.

The stability of the interaction between FMN_ox and MtCS was tested by incubating MtCS with excess FMN_ox in buffer A for 12 hours at 4°C in the dark. The mixture was injected into a HiTrapQ HP anion exchange column previously equilibrated with buffer A. The sample was eluted with a linear gradient from 0.0 to 1 M NaCl. Absorption spectrum of the pool of fractions containing FMN_ox and MtCS was measured at wavelengths values ranging from 210 to 600 nm by a Shimadzu UV-2550 spectrophotometer.

The binding of FMN_ox to MtCS (35 μM) was assessed as described by Kitzing et al. 21. UV-visible difference absorbance spectra were monitored at 25°C in a double beam spectrophotometer UV-2550 (Shimadzu) with photometric accuracy of ± 0.002 absorbance units in the range 0 – 0.5 absorbance. The difference spectra were recorded using quartz cells with two chambers (Hellma GmbH & Co) placing identical arrangements in both the sample (sample cuvette) and reference beam (reference cuvette). Spectra were recorded from 300 to 600 nm. Differences between solutions were minimized by filling both sides (sample and reference cuvettes) from single stock solutions of FMN_ox. Microliter additions of FMN_ox to both one chamber of the reference cuvette and to the sample cuvette, followed by mixing of enzyme and FMN_ox solutions for the latter, allowed direct measurements of the difference spectra. This procedure is almost mandatory when change in absorbance signal is relatively small on interaction. All solutions were in 50 mM Tris-HCl, pH 7.8 (buffer A).

Fluorescence titration was performed at equilibrium in a RF-5301PC Spectrofluorophotometer (Shimadzu) at 25°C by making microliter additions of NADH stock solution (10 mM) to 2 mL of 1 μM MtCS (active site concentration) in 50 mM Tris-HCl, pH 7.8 (buffer A). Controls were determined, following exactly the same procedure, by microliter additions of buffer A to account for changes in protein fluorescence due to dilution. In order to assess the NADH inner-filter effect in the fluorimeter, two cuvettes were placed in series so that the contents of the first cuvette acted as a filter of the excitation light and the light emitted from the second cuvette detected. To the first cuvette, NADH was added, while the second cuvette contained MtCS. In this manner, NADH inner-filter effects upon the protein fluorescence could be assessed. The binding of NADH to MtCS causes a quench in protein fluorescence (λ_exc = 290 nm; 310 ≤ λ_em ≤ 510 nm), and fluorescence values at the maximum emission wavelength (345.5 nm) were plotted against increasing NADH concentrations. The data were fitted to a hyperbolic function, yielding a value of 156 (± 12) μM for the dissociation constant of MtCS-NADH binary complex formation.

FMN-reductase activity of MtCS was monitored for the forward reaction 25°C in buffer A. The activity measurements were based on the decrease of NADH concentration upon FMN reduction. Owing to the high absorption coefficient of FMN at 360 nm, the consumption of NADH was monitored at 380 nm (ε = 0.893 × 10³ M^-1 cm^-1). The apparent steady state kinetic parameters Km and Vmax of FMN reductase activity were determined for holoenzyme MtCS-FMN_ox in the presence of varying concentrations of NADH (10, 25, 50, 75, 100, 200, 300 μM). The reaction was started with the addition of 60 nmol of homogeneous MtCS previously incubated with 40 μM FMN_ox.

[4S-²H]NADH was synthesized as described by Ottolina et al. 51. The substrates, [4S-4-¹H]- or [4S-4-²H]-NADH, were purified on a MonoQ column as previously reported 52, and the fractions with absorbance ratios A_{260 nm}/A_{340 nm} ≤ 2.3 were pooled. Both kinetic isotope effects and proton inventory were determined in buffer A with homogeneous MtCS previously incubated with 40 μM FMN_ox. Primary deuterium kinetic isotope effects were determined from measurements of initial rates in the presence of varying concentrations of either [4S-4-¹H]- or [4S-4-²H]-NADH, and solvent kinetic isotope effects from measurements of initial rates in the presence of varying concentrations of NADH in either H₂O or 91 atom % D₂O. Multiple isotope effects were determined by measuring initial velocities in the presence of varying concentrations of either [4S-4-¹H]- or [4S-4-²H]-NADH in 90 atom % D₂O. The pD of buffers used on solvent and multiple isotope effects were measured on a pH meter considering pD = pH + 0.4. Proton inventory was carried out at saturating NADH concentrations at various mole fractions of D₂O. All measurements were performed in triplicate. The nomenclature proposed by Northrop 31 and Cook and Cleland 34 was used to express isotope effects.

Values for the apparent kinetic parameters and their standard errors were obtained by fitting the data to the appropriate equations using the non-linear regression function of SigmaPlot 2000 (SPSS, Inc.). The initial rate measured at seven different NADH concentrations were fitted to eq. 1.

v = VA/(K + A)

Isotope effect data were fitted to eq 2, which assumes isotope effects on both V/K and V. In equations 1 and 2, V is the maximal velocity, K is the Michaelis constant, A is substrate concentrations, E_V/K and E_V are the isotope effects minus 1 on V/K and V, respectively, and F_i is the fraction of deuterium label.

v = VA/[K(1 + F_iE_V/K) + A(1 + F_iE_V)]