Magnesium-dependent ATPase Activity and Cooperativity of Magnesium Chelatase from Synechocystis sp. PCC6803 - J. David Reid; C. Neil Hunter (2004)
The first committed step in chlorophyll biosynthesis is catalyzed by magnesium chelatase, a complex enzyme with at least three substrates, cooperative Mg2+ activation, and free energy coupling between ATP hydrolysis and metal-ion chelation. A detailed functional study of the behavior of the intact magnesium chelatase has been performed, including characterization of magnesium cooperativity and the stoichiometry of ATP consumption in relation to the magnesium porphyrin produced. It is demonstrated that, in vitro, this catalyzed reaction requires hydrolysis of ∼15 MgATP2– and that the chelation partial reaction is energetically unfavorable, under our assay conditions, with a ΔG°′ of 25–33 kJ mol–1. Given the likely metabolite concentrations in vivo, this results in the chelatase reaction operating far from equilibrium. We have also determined the steady-state kinetic behavior of the intact enzyme and have compared the kinetic parameters obtained with those observed for the partial reactions of individual subunits. KDIX (where DIX represents deuteroporphyrin IX) is estimated to be 3.20 μm, and KMgATP2– is 0.45 mm. kcat for chelation is estimated to be 0.8 min–1, suggesting that the ATP hydrolysis catalyzed by the isolated ChlI subunit is substantially slower in the intact chelatase. The magnesium-rich form of the chelatase is a more effective catalyst of the chelation reaction; magnesium activation of the chelatase increases V, as well as the specificity constant for the reaction of MgATP2– and DIX, possibly as a result of a magnesium-triggered conformational change. The first committed step in chlorophyll biosynthesis is catalyzed by magnesium chelatase, a complex enzyme with at least three substrates, cooperative Mg2+ activation, and free energy coupling between ATP hydrolysis and metal-ion chelation. A detailed functional study of the behavior of the intact magnesium chelatase has been performed, including characterization of magnesium cooperativity and the stoichiometry of ATP consumption in relation to the magnesium porphyrin produced. It is demonstrated that, in vitro, this catalyzed reaction requires hydrolysis of ∼15 MgATP2– and that the chelation partial reaction is energetically unfavorable, under our assay conditions, with a ΔG°′ of 25–33 kJ mol–1. Given the likely metabolite concentrations in vivo, this results in the chelatase reaction operating far from equilibrium. We have also determined the steady-state kinetic behavior of the intact enzyme and have compared the kinetic parameters obtained with those observed for the partial reactions of individual subunits. KDIX (where DIX represents deuteroporphyrin IX) is estimated to be 3.20 μm, and KMgATP2– is 0.45 mm. kcat for chelation is estimated to be 0.8 min–1, suggesting that the ATP hydrolysis catalyzed by the isolated ChlI subunit is substantially slower in the intact chelatase. The magnesium-rich form of the chelatase is a more effective catalyst of the chelation reaction; magnesium activation of the chelatase increases V, as well as the specificity constant for the reaction of MgATP2– and DIX, possibly as a result of a magnesium-triggered conformational change. Chlorophyll is synthesized via a series of tetrapyrrole intermediates; the path toward this pigment diverges from heme biosynthesis at the chelation steps. The chelation step in chlorophyll biosynthesis, catalyzed by magnesium chelatase (E.C. 6.6.1.1), requires ATP hydrolysis and has long been presumed to be energetically unfavorable. The minimum catalytic unit of magnesium chelation requires three proteins, I (38–42 kDa), D (60–74 kDa), and H (140–150 kDa), both in (bacterio) chlorophyll a-producing prokaryotes (1Gibson L.C.D. Willows R.D. Kannangara C.G. von Wettstein D. Hunter C.N. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 1941-1944Crossref PubMed Scopus (154) Google Scholar) and in chlorophyll a-synthesizing bacteria and higher plants (2Jensen P.E. Willows R.D. Petersen B.L. Vothknecht U.C. Stummann B.M. Kannangara C.G. von Wettstein D. Henningsen K.W. Mol. Gen. Genet. 1996; 250: 383-394PubMed Google Scholar, 3Papenbrock J. Gräfe S. Kruse E. Hänel F. Grimm B. Plant J. 1997; 12: 981-990Crossref PubMed Scopus (76) Google Scholar). Studies on purified proteins show that I is the ATPase subunit (4Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1999; 339: 127-134Crossref PubMed Scopus (86) Google Scholar), that it contains a Mg2+ binding site (5Reid J.D. Siebert C.A. Bullough P.A. Hunter C.N. Biochemistry. 2003; 42: 6912-6920Crossref PubMed Scopus (49) Google Scholar), and that it has a fold characteristic of a member of the AAA+ superfamily (6Fodje M.N. Hansson A. Hansson M. Olsen J.G. Gough S. Willows R.D. Al Karadaghi S. J. Mol. Biol. 2001; 311: 111-122Crossref PubMed Scopus (135) Google Scholar). The D subunit forms a stable complex with the I subunit (4Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1999; 339: 127-134Crossref PubMed Scopus (86) Google Scholar) and contains a metal binding motif (6Fodje M.N. Hansson A. Hansson M. Olsen J.G. Gough S. Willows R.D. Al Karadaghi S. J. Mol. Biol. 2001; 311: 111-122Crossref PubMed Scopus (135) Google Scholar), although no metal binding has yet been demonstrated. The H subunit binds porphyrins (7Karger G.A. Reid J.D. Hunter C.N. Biochemistry. 2001; 40: 9291-9299Crossref PubMed Scopus (73) Google Scholar, 8Willows R.D. Beale S.I. J. Biol. Chem. 1998; 273: 34206-34213Abstract Full Text Full Text PDF PubMed Scopus (53) Google Scholar) and presumably also contains the active site for chelation. As the intact chelatase is a complex enzyme with at least three substrates, cooperative Mg2+ activation, and free energy coupling between ATP hydrolysis and metal-ion chelation, existing mechanistic studies have only provided an overview of the behavior of this enzyme (9Gibson L.C.D. Jensen P.E. Hunter C.N. Biochem. J. 1999; 337: 243-251Crossref PubMed Scopus (61) Google Scholar, 10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). In light of recent studies clarifying the role of individual subunits (4Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1999; 339: 127-134Crossref PubMed Scopus (86) Google Scholar, 5Reid J.D. Siebert C.A. Bullough P.A. Hunter C.N. Biochemistry. 2003; 42: 6912-6920Crossref PubMed Scopus (49) Google Scholar, 6Fodje M.N. Hansson A. Hansson M. Olsen J.G. Gough S. Willows R.D. Al Karadaghi S. J. Mol. Biol. 2001; 311: 111-122Crossref PubMed Scopus (135) Google Scholar, 7Karger G.A. Reid J.D. Hunter C.N. Biochemistry. 2001; 40: 9291-9299Crossref PubMed Scopus (73) Google Scholar, 8Willows R.D. Beale S.I. J. Biol. Chem. 1998; 273: 34206-34213Abstract Full Text Full Text PDF PubMed Scopus (53) Google Scholar), a detailed functional study of the behavior of the intact magnesium chelatase is now required. For example, there has been no detailed analysis of the cooperativity shown by this enzyme to magnesium, which is separate from the requirement for magnesium to form the MgATP2– substrate (10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). One compelling reason for requiring such information arises from the demonstration of mutations in the H subunit of magnesium chelatase that impair plastid-to-nucleus signaling (11Mochizuki N. Brusslan J.A. Larkin R. Nagatani A. Chory J. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 2053-2058Crossref PubMed Scopus (508) Google Scholar) and the revelation that GUN4, an additional factor involved in this interaction, activates magnesium chelatase (12Larkin R.M. Alonso J.M. Ecker J.R. Chory J. Science. 2003; 299: 902-906Crossref PubMed Scopus (384) Google Scholar). A regulatory role for magnesium chelatase has long been suspected from the highly cooperative response of this enzyme to magnesium (10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). In view of the likely importance of this cooperativity, we will describe this phenomenon in much greater detail, clarifying the effect of the other substrates. This kinetic information will prove essential in assessing how the GUN4 protein acts to accelerate the rate of magnesium chelation. As intraplastid free magnesium concentration is an effective signal of [ATP]:[ADP]·[Pi] status, this sensitivity links photosynthetic performance to activity of the gateway enzyme for photosynthetic pigment production. Another aspect of this enzyme that now requires quantitative examination is the stoichiometry of ATP consumption in relation to the magnesium protoporphyrin produced. In this report, we demonstrate that, in vitro, this catalyzed reaction requires hydrolysis of ∼15 MgATP2– and that the chelation reaction is energetically unfavorable, under our assay conditions, with a ΔG°′ of 25–33 kJ mol–1. Given the likely metabolite concentrations in vivo, this results in the chelatase reaction operating far from equilibrium. We have also determined the steady-state kinetic behavior of the intact enzyme and can compare the kinetic parameters obtained with those observed for the partial reactions of individual subunits. Protein Purification—The expression vectors pET9a-ChlI, pET9a-His6ChlD, and pET9a-His6ChlH (4Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1999; 339: 127-134Crossref PubMed Scopus (86) Google Scholar) were used to produce recombinant protein essentially as described previously (13Jensen P.E. Gibson L.C.D. Henningsen K.W. Hunter C.N. J. Biol. Chem. 1996; 271: 16662-16667Abstract Full Text Full Text PDF PubMed Scopus (116) Google Scholar). ChlI was purified by anion-exchange chromatography on a Q-Sepharose (Amersham Biosciences) column with a gradient from 250 to 600 mm NaCl over five column volumes in 50 mm Tricine 1The abbreviations used are: Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; MOPS, 4-morpholinepropanesulfonic acid; DIX, deuteroporphyrin IX; MgDIX, magnesium deuteroporphyrin IX; HPLC, high pressure liquid chromatography./NaOH, 0.3 m glycerol, 1 mm dithiothreitol, pH 7.9 (buffer A). The resulting material, which is more than 95% ChlI by SDS-PAGE, was concentrated to ∼5 ml, and monomeric ChlI was purified in buffer A, with 200 mm NaCl, by gel filtration on a Sephacryl S-200 HiPrep 26/60 column (Amersham Biosciences) at 0.5 ml/min. Pure protein was flash-frozen in and stored under liquid nitrogen. Nucleotide-free protein had a λmax of 280 nm, and protein concentration was determined using the calculated ϵ280 of 11,290 m–1 cm–1 (14Gill S.C. von Hippel P.H. Anal. Biochem. 1989; 182: 319-326Crossref PubMed Scopus (5034) Google Scholar). His6ChlD and His6ChlH were purified as described previously except that protein concentration was determined using ϵ280 values of 44,560 m–1 cm–1 and 163,330 m–1 cm–1, respectively, which were calculated and corrected as described (14Gill S.C. von Hippel P.H. Anal. Biochem. 1989; 182: 319-326Crossref PubMed Scopus (5034) Google Scholar). Metal Ion Control—When studying the properties of ATPases, it is important to control the cation/nucleotide equilibrium as metal-bound nucleotide is generally the substrate (15Morrison J.F. Methods Enzymol. 1979; 63: 257-294Crossref PubMed Scopus (80) Google Scholar). As the free species (e.g. ATP4–, Mg2+) often act as activators or inhibitors, it is useful to investigate the effects of varying their concentration (15Morrison J.F. Methods Enzymol. 1979; 63: 257-294Crossref PubMed Scopus (80) Google Scholar); this is particularly important in an enzyme that processes magnesium ions. In our reaction mixtures, essentially all ATP is maintained in the MgATP2– form by keeping at least a 1 mm excess of MgCl2 over ATP. The effectiveness of this procedure can be calculated from the stability constants for MgATP2– and the pKa of the terminal phosphate of ATP (16O'Sullivan W.J. Smithers G.W. Methods Enzymol. 1979; 63: 294-336Crossref PubMed Scopus (187) Google Scholar), and the concentration of free nucleotide in our experiments is always less than 5% of total nucleotide. Magnesium Chelation and ATPase Assays—Assay conditions were essentially as described previously (10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). Reactions were run at 34 °C, I 0.1 and pH 7.7, in 50 mm MOPS/KOH, 0.3 m glycerol, 1 mm dithiothreitol. Assays were initiated by the addition of enzyme to give a final concentration of 0.2 μm ChlI, 0.1 μm His6ChlD, and 0.4 μm His6ChlH. This subunit ratio was previously shown to give an optimal enzyme activity (10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). Substrate concentrations are given in the appropriate figures. Detection systems are described below. Porphyrin solutions were made weekly in assay buffer and discarded if precipitated. Magnesium deuteroporphyrin and deuteroporphyrin were determined spectrophotometrically (ϵ398 433,000 m–1 cm–1 in 0.1 m HCl), ATP and ADP were determined spectrophotometrically (ϵ260 15,400 m–1 cm–1). Malachite Green Phosphate Assays—The method of Lanzetta (17Lanzetta P.A. Alvarez L.J. Reinach P.S. Candia O.A. Anal. Biochem. 1979; 100: 95-97Crossref PubMed Scopus (1810) Google Scholar) was used except that Triton X-100 was used as a detergent. Samples (25 μl) were taken from a reaction every 5 or 10 min and quenched in an equal volume of malachite green solution. The quenched reaction was incubated for 2 min at 34 °C to allow color to develop and then read in a Bio-Tek F2 plate-reader in absorbance mode through a 630-nm bandpass filter. Standard curves were generated using a KPi solution in the same buffer and at the same ATP concentrations as used in the assays. A 100-ml stock solution contained 2g of ammonium molybdate, 0.3 g of malachite green·HCl, 0.5 g of Triton X-100, and was stored shielded from light at 4 °C. HPLC Ion-exchange ADP Assays—Nucleotide analysis was essentially as described (18Guranowski A. Brown P. Ashton P.A. Blackburn G.M. Biochemistry. 1994; 33: 235-240Crossref PubMed Scopus (40) Google Scholar) except that a 1-ml Resource Q column (15-μm beads, Amersham Biosciences) was used on a Waters HPLC system. Samples (50 μl) were taken from a reaction every 5 or 10 min and quenched in 100 μl of ice-cold 95% EtOH; 75 μl of these quenched reactions was applied to the ion-exchange column. The column was equilibrated (1 ml/min) with 50 mm ammonium bicarbonate, pH 8.5, the samples were applied, the column was washed with 2 ml of starting buffer, and then the nucleotides were resolved with a 20-ml linear gradient (50–1000 mm) of the same buffer. Nucleotides were detected by their absorbance at 260 nm. The peak area was estimated, by integration, using the software supplied by the manufacturer (Millenium32). Continuous Fluorimetric Assays for Magnesium Deuteroporphyrin— 100-μl reactions were observed in a Bio-Tek F2 microplate reader with excitation through a 420 ± 25 nm filter, and emission was observed through a 590 ± 17.5 nm filter (7Karger G.A. Reid J.D. Hunter C.N. Biochemistry. 2001; 40: 9291-9299Crossref PubMed Scopus (73) Google Scholar, 9Gibson L.C.D. Jensen P.E. Hunter C.N. Biochem. J. 1999; 337: 243-251Crossref PubMed Scopus (61) Google Scholar). Steady-state rates were estimated from chelatase progress curves using the instrument software. Synchronous Scanning Non-enzymic Fluorimetric Assays for Magnesium Deuteroporphyrin at a Low Magnesium Deuteroporphyrin, Deuteroporphyrin Ratio—Solutions of magnesium deuteroporphyrin (1–5 μm) were incubated overnight at 34 °C in 50 mm MOPS/KOH, 0.3 m glycerol, 1 mm dithiothreitol, I 0.1, pH 7.7. No further reaction was seen when longer incubations were used. Sample (1 ml) fluorescence was determined by synchronous scanning with a 181-nm gap between excitation and emission, with excitation scanned between 350 and 600 nm on a SPEX Fluorolog spectrofluorimeter. Magnesium deuteroporphyrin can be determined from the 400-nm peak and, with ∼90-fold less sensitivity, deuteroporphyrin can be determined from the 495-nm peak Kinetic and Thermodynamic Data Analysis—Non-linear regression analysis was performed using Sigmaplot 8.0 (SPSS). At a fixed free Mg2+ concentration, apparent kinetic parameters for the other two substrates (Vapp, V/KMgATP-app, V/KDIX-app, V/KMgATP: DIX-app) were estimated from hyperbolic fits to primary and secondary plots as usual. The dependence of apparent kinetic parameters on free magnesium was evaluated graphically and by the Hill equation. Apparent kinetic parameters derived from primary plots have an app′ suffix, whereas those derived from secondary plots have an app suffix. A standard state free energy for the uncatalyzed chelation reaction was derived from the porphyrin concentrations observed at equilibrium (Equation 1), assuming that free magnesium and deuteroporphyrin concentrations are identical. Literature estimates of the porphyrin, metallo-porphyrin, and free magnesium concentrations were used to estimate the free energy required for the chelation reaction in vivo (Equation 2). The free energy change associated with the coupled reaction was calculated from literature values of the free energy available from ATP hydrolysis, ΔG′p, and, n, our observed ATPase stoichiometry (Equation 3). ΔGchelation∘′=−RTln([MgDIX][Mg2+][DIX])equilibrium (Eq. 1) ΔGchelation′=ΔGchelation∘′+RTln([MgDIX][Mg2+][DIX])observed (Eq. 2) ΔG′=ΔGchelation′+nΔGp′ (Eq. 3) The Stoichiometric Relationship between MgATP2–Hydrolysis and Mg2+-Deuteroporphyrin Chelation—We have investigated the relationship between magnesium chelation and ATP hydrolysis catalyzed by magnesium chelatase. We have measured the time-dependent production of all three reaction products, magnesium deuteroporphyrin IX (MgDIX), ADP, and Pi. Magnesium chelation is readily followed by a fluorimetric assay using either protoporphyrin IX (10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar), the native substrate, or deuteroporphyrin IX (7Karger G.A. Reid J.D. Hunter C.N. Biochemistry. 2001; 40: 9291-9299Crossref PubMed Scopus (73) Google Scholar), a more water-soluble analogue. Although ATP hydrolysis can be detected by an extensive range of methods, a continuous absorbance-based assay (19Webb M.R. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 4884-4887Crossref PubMed Scopus (486) Google Scholar) has proved a convenient choice for magnesium chelatase (4Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1999; 339: 127-134Crossref PubMed Scopus (86) Google Scholar). This system is, however, not readily applicable to assays containing deuteroporphyrin (DIX) as the Soret band of the pigment significantly overlaps the product absorbance band from the linked Pi assay system. We therefore followed ATP hydrolysis by magnesium chelatase with two alternative assay systems: the well established malachite green assay for inorganic phosphate and an HPLC ion-exchange assay for ADP. The release of Pi, ADP, and MgDIX is coincident when a scaling factor is applied to the ATPase curves. This shows that the chelatase reaction and the ATPase reactions have a similar time course and, in particular, both show the same lag phase. The steady-state rates (vss) of both reactions show similar, apparently cooperative, responses to free magnesium (Fig. 1). Some divergence of the two curves is seen at low Mg2+ concentrations. The ratio of ATPase reaction products to metalloporphyrin, 15, reflects the stoichiometry of the reaction. At first sight, this stoichiometry seems quite large; a preliminary hypothesis is that this expenditure reflects the in vivo thermodynamic requirements of the chelation reaction. Without a detailed understanding of in vivo chelation conditions, such as pH and steady-state reactant concentrations, the thermodynamic constraints may prove rather difficult to assess. A reasonable starting point would be the thermodynamics of the chelation partial reaction, assessed under our assay conditions in the absence of enzyme. The product MgDIX is unstable and will slowly decay to a mixture of DIX and Mg2+. As the spectra of both the porphyrin and the metallo-porphyrin overlap extensively, we used synchronous scanning to determine the exceptionally low concentrations of metallo-porphyrin observed at equilibrium. The equilibrium constant for the uncatalyzed dechelation was observed to be of the order of 10–6, and thus a ΔG°′ of 25–33 kJ mol–1 is observed (Equation 1). The relationship between these thermodynamic parameters and the steady-state ratios of substrates and products that have physiological significance are explored under "Discussion." A Steady-state Approach to Understanding Magnesium Cooperativity—We have explored the steady-state behavior of magnesium chelatase across a range of substrate concentrations. Initial plots of steady-state rate (vss) against DIX or MgATP2– (e.g.Fig. 2) were fitted by the Michaelis-Menten equation. All initial rate versus [S] plots not shown in Fig. 2 are available in the supplementary material (Fig. S1). The resulting parameters (VDIX-app′ and V/KDIX-app′) were then plotted against a second substrate concentration and fitted to the expected and observed hyperbolic relationship. For example, fitted values of VDIX-app′ and V/KDIX-app′ at fixed concentrations of the other two substrates were plotted against [MgATP2–]. The resulting plots (Fig. 3) were then analyzed to provide kinetic parameters. The analogous plots when VMgATP-app′ and V/KMgATP-app′ are plotted against [DIX] are shown in the supplementary material (Fig. S3). The parameters observed at each individual Mg2+ concentration are: Vapp, the apparent maximum rate at saturating MgATP2–; DIX, V/KDIX-app, V/KMgATP-app, the apparent pseudo-first order rate constants for the productive reaction of enzyme with one substrate at saturating concentrations of the other; and V/KMgATP·Dix-app, the apparent pseudo-second order rate constant for the productive reaction of enzyme with both substrates.Fig. 3MgATP2– dependence of the deuteroporphyrin apparent kinetic parameters at a range of magnesium concentrations.A, dependence of VDIX-app′ on MgATP2– at, from the top,9,7,5,4,3,and 1 mm Mg2+. The fitted lines are hyperbolic with characterizing parameters shown in Fig. 4. B, dependence of V/KDIX-app′ on MgATP2– at, from the top, 9, 7, 5, 4, and 3 mm Mg2+. The fitted lines are hyperbolic with characterizing parameters, Vapp, V/KMgATP-app, V/KDIX-app, and V/KMgATP·DIX-app, shown in Fig. 4.View Large Image Figure ViewerDownload Hi-res image Download (PPT) It is immediately obvious from inspection of Fig. 3 that varying the concentration of magnesium ions affects the MgATP2– dependence of the observed VDIX-app′ and V/KDIX-app′. This effect can be characterized from the magnesium dependence of the four apparent kinetic parameters. i.e. Vapp, V/KDIX-app, V/KMgATP-app, and V/KMgATP·Dix-app (Fig. 4). These curves are not hyperbolic and appear to be positively cooperative. These data have been fitted to the Hill equation in which nH, the Hill coefficient, provides a convenient estimate of apparent cooperativity. Kinetic parameters for MgATP2– and DIX can readily be estimated from Fig. 4. The apparent maximum rate (Vapp) at saturating MgATP2– and DIX is plotted against Mg2+ (Fig. 4A). At saturating magnesium, the true maximum rate, V, is 0.31 ± 0.03 μm min–1, a value similar to those estimated previously (10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). The apparent pseudo-first order rate constant (V/KMgATP-app) for the reaction of MgATP2– with an enzymeporphyrin complex is predicted to reach a limiting value of (6.9 ± 1) × 10–4 min–1 at high magnesium concentrations. KMgATP can be estimated to be 0.45 mm, significantly higher than that obtained from a study of the isolated ChlI subunit (5Reid J.D. Siebert C.A. Bullough P.A. Hunter C.N. Biochemistry. 2003; 42: 6912-6920Crossref PubMed Scopus (49) Google Scholar). In this work, Mg2+ saturation of the ATPase reaction catalyzed by ChlI gave a value of K « 0.14 mm. In the Mg2+-unbound form, ChlI gave a value of K ∼1.12 mm (5Reid J.D. Siebert C.A. Bullough P.A. Hunter C.N. Biochemistry. 2003; 42: 6912-6920Crossref PubMed Scopus (49) Google Scholar). The apparent pseudo-first order rate constant (V/KDIX-app) for the reaction of DIX with an enzyme-nucleotide complex also reaches a limit as Mg2+ reaches saturation, giving V/KDIX 0.097 ± 0.004 min–1. KDIX can therefore be estimated to be 3.20 μm, similar to the disassociation constant for the H-DIX complex (4.0 μm) in the presence of MgCl2 and ATP (7Karger G.A. Reid J.D. Hunter C.N. Biochemistry. 2001; 40: 9291-9299Crossref PubMed Scopus (73) Google Scholar). The cooperative response to magnesium cannot be eliminated by saturating with DIX, as shown by the response of V/KMgATP-app to Mg2+ (Fig. 4B), by saturating with MgATP2–, as shown by the response of V/KDIX-app (Fig. 4C), or by saturating with both MgATP2– and DIX, as shown by the response of Vapp to Mg2+ (Fig. 4A). As the response of V/KMgATP·DIX-app to magnesium is linear (Fig. 4D) and does not show magnesium saturation, magnesium binding to the free chelatase is considerably weaker than to chelatase bound to either (Fig. 4, B and C) or both (Fig. 4A) of the other substrates. From our data, it is not possible to determine whether this response is cooperative. Magnesium chelatase is an enzyme of quite formidable complexity. Although it is most often described as a three-substrate enzyme (20Reid J.D. Hunter C.N. Biochem. Soc. Trans. 2002; 30: 643-645Crossref PubMed Scopus (28) Google Scholar, 21Walker C.J. Willows R.D. Biochem. J. 1997; 327: 321-333Crossref PubMed Scopus (177) Google Scholar), this is a convenient simplification. As more than one MgATP2– moiety is hydrolyzed in each catalytic cycle, each of these can be considered as a separate substrate. The kinetic consequences of this view may help us understand how up to 15 MgATP2– are hydrolyzed in a single catalytic cycle. The well established cooperativity with respect to Mg2+ demonstrates that magnesium activates catalysis as well as acting both as a substrate and as a ligand to ATP, forming MgATP2– (9Gibson L.C.D. Jensen P.E. Hunter C.N. Biochem. J. 1999; 337: 243-251Crossref PubMed Scopus (61) Google Scholar, 10Jensen P.E. Gibson L.C.D. Hunter C.N. Biochem. J. 1998; 334: 335-344Crossref PubMed Scopus (76) Google Scholar). The non-hyperbolic magnesium kinetic profiles could arise either from multiple, physically distinct, interacting magnesium binding sites or from magnesium binding at many different stages in a catalytic cycle effectively creating multiple, temporally distinct, sites (22Sweeny J.R. Fisher J.R. Biochemistry. 1968; 7: 561-565Crossref PubMed Scopus (57) Google Scholar). Our data clearly demonstrate that, at high magnesium, chelation of 1 magnesium requires hydrolysis of 15 MgATP2–. Magnesium chelation therefore appears to be one of the more expensive reactions coupled to ATP hydrolysis. This stoichiometry can be compared with nitrogenase, an enzyme of similar complexity, that requires at least 16 ATPase cycles to reduce dinitrogen to ammonia. This coupled reaction is associated with a large free energy change of –117 kJ mol–1 for each N2 reduced (23Thorneley R.N.F. Philos. Trans. R. Soc. Lond. B Biol. Sci. 1992; 336: 73-82Crossref PubMed Scopus (14) Google Scholar). It is interesting to note that under some conditions, the Kp2OX(MgADP+Pi)2Kp1 complex of nitrogenase offers a leak pathway for ATP hydrolysis that can increase the observed stoichiometry. Apart from these mechanistic similarities, nitrogenase and magnesium chelatase are unrelated. Other members of the AAA+ superfamily offer interesting examples of ATPase coupling related to the magnesium chelatase system (24Vale R.D. J. Cell Biol. 2000; 150: F13-F19Crossref PubMed Google Scholar). The ATP-dependent protease ClpXP, a member of the Clp/HSP100 family of AAA+ proteins, denatures and degrades SsrA-tagged proteins (25Wojtkowiak D. Georgopoulos C. J. Biol. Chem. 1993; 268: 22609-22612Abstract Full Text PDF PubMed Google Scholar, 26Gottesman S. Roche E. Zhou Y. Sauer R.T. Genes Dev. 1998; 12: 1338-1347Crossref PubMed Scopus (646) Google Scholar). This protease system hydrolyzes at least 100 ATP molecules/protein substrate, with observed stoichiometries depending more on the C-terminal stability of the substrate (27Kenniston J.A. Baker T.A. Fernandez J.M. Sauer R.T. Cell. 2003; 114: 511-520Abstract Full Text Full Text PDF PubMed Scopus (232) Google Scholar) than the relatively low, in vitro, energy barrier (ΔGd ∼4kJmol–1) to unfolding (28Burton R.E. Siddiqui S.M. Kim Y.I. Baker T.A. Sauer R.T. EMBO J. 2001; 20: 3092-3100Crossref PubMed Scopus (122) Google Scholar). Katanin, a heterodimeric member of the AAA+ superfamily, severs and disassembles microtubules in an ATP-dependent manner. This requires around 50 ATP hydrolysis events/tubulin dimer (29Hartman J.J. Vale R.D. Science. 1999; 286: 782-785Crossref PubMed Scopus (182) Google Scholar). Such examples reflect the more extreme end of ATPase stoichiometries observed in this superfamily. In contrast, the AAA+ component of Escherichia coli DNA polymerase III, subunits γ and δ′, assembles and clamps the holoenzyme onto DNA at a cost of 2–3 ATP (30Bertram J.G. Bloom L.B. Hingorani M.M. Beechem J.M. O'Donnell M. Goodman M.F. J. Biol. Chem. 2000; 275: 28413-28420Abstract Full Text Full Text PDF PubMed Scopus (56) Google Scholar). Thermodynamic details of these AAA+ superfamily catalyzed reactions are not available, so ΔG′ cannot be estimated. A recent structur