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1 School of Cell and Molecular Biosciences, Catherine Cookson Building, University of Newcastle upon Tyne, NE2 4HH, UK
2 Division of Structural Biology, The Henry Wellcome Building for Genomic Medicine, University of Oxford, Oxford OX3 7BN, UK
3 Department of Biological Sciences Centre for Molecular Microbiology and Infection, Imperial College London, London, SW7 2AZ, UK
4 Department of Chemistry, University of Glasgow, Glasgow, G12 8QQ, UK
5 Arrow Therapeutics, London, SE1 1DA, UK
Reprint requests to: Alastair R. Hawkins, School of Cell and Molecular Biosciences, Catherine Cookson Building, University of Newcastle upon Tyne, Framlington Place, NE2 4HH, UK; e-mail: a.r.hawkins{at}ncl.ac.uk; fax: 44-191-2227424.
(RECEIVED February 23, 2004; FINAL REVISION April 28, 2004; ACCEPTED April 28, 2004)
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Abstract |
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-elimination, carbonyl reduction, ring opening, and intramolecular aldol condensation. Kinetic analysis of the isolated DHQS domains with the AROM protein showed that for the substrate DAHP the difference in Km is less than a factor of 3, that the turnover numbers differed by 24%, and that the Km for NAD+ differs by a factor of 3. Isothermal titration calorimetry revealed that a second (inhibitory) site for divalent metal binding has an approximately 4000-fold increase in KD compared to the catalytic binding site. Inhibitor studies have suggested the enzyme could act as a simple oxidoreductase with several of the reactions occurring spontaneously, whereas structural studies have implied that DHQS participates in all steps of the reaction. Analysis of site-directed mutants experimentally test and support this latter hypothesis. Differential scanning calorimetry, circular dichroism spectroscopy, and molecular exclusion chromatography demonstrate that the mutant DHQS retain their secondary and quaternary structures and their ligand binding capacity. R130K has a 135-fold reduction in specific activity with DAHP and a greater than 1100-fold decrease in the kcat/Km ratio, whereas R130A is inactive. Keywords: dehydroquinate synthase; AROM; biocalorimetry; site-directed mutagenesis
6 Present address: Vernalis (Cambridge) Ltd., Granta Park, Abington, Cambridge, Cambridgeshire, CB1 6GB, UK. 
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.04705404.
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Introduction |
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In many microbial eukaryotes the AROM protein shares a common pool of the substrates dehydroquinate and dehydroshikimate with the type II dehydroquinase and the dehydroshikimate dehydratase of the quinic acid utilization pathway. Metabolic control analysis has shown that the AROM protein is very leaky, and that the quinic acid pathway dehydroshikimate dehydratase has a flux control coefficient of -1 in the shikimate pathway. These observations prompted the proposal that the AROM protein may have a low level channelling function to protect against substrate depletion by dehydroshikimate dehydratase under conditions where the exogenous supply of quinic acid is rapidly diminished (Lamb et al. 1992; Wheeler et al. 1996). This putative channelling function could simply be due to the close juxtaposition of the five active sites within AROM. However, it is also possible that each of the enzyme domains has evolved to function with optimal catalytic activity only as part of the AROM protein.
The N-terminal domain of the AROM protein is the enzyme dehydroquinate synthase (DHQS), and this catalyzes the conversion of DAHP to dehydroquinate (DHQ; Giles et al. 1967; Lumsden and Coggins 1977; Charles et al. 1986; Bentley 1990; Hawkins et al. 1993a; Moore et al. 1994). The isolated DHQS domain of the AROM protein from Aspergillus nidulans has been crystallized and its structure determined (Carpenter et al. 1998; Nichols et al. 2003). The enzyme DHQS has generated interest because it apparently catalyzes five individual reactions (alcohol oxidation, phosphate -elimination, carbonyl reduction, ring opening, and intramolecular aldol condensation) in a single active site (Srinivasan et al. 1963; Widlanski et al. 1989). The proposed reaction mechanism is summarized in Figure 1
. The molecular mechanism of the overall reaction has been studied extensively by the use of substrate analogs (Bartlett and Satake 1988; Bender et al. 1989b; Knowles 1989; Widlanski et al. 1989; Bartlett et al. 1994), leading to the suggestion that the enzyme may be acting as a simple oxido-reductase with several of the reactions occurring spontaneously.
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) is further divided into an N-terminal NAD+ binding 
/ domain containing a Rossmann fold ("N" domain) and a C-terminal -helical domain containing most of the residues involved in catalysis, substrate and Zn2+ binding ("C" domain). Formation of each of the two active sites within the dimer requires the interaction of amino acids from the "N" and "C" domains of one monomer together with the side chain R130 from the other monomer’s "N" domain. DHQS, in the absence of the substrate analog CBP (i.e., with NAD+, ADP, or unliganded), is in an open form where a relative rotation of 11°–14° between N- and C-terminal domains occurs. Overlapping 21 different copies of the individual N- and C-terminal DHQS domains revealed a series of pivot points about which the domain closure occurs. This analysis suggested that the structural mechanism for domain closure involved an ordered sequence of substrate binding, local rearrangement within the active site, and a propagation of torque inducing closure of the active-site cleft (Nichols et al. 2001, 2003). A preliminary report of a further open form structure of DHQS in a binary complex with NAD+ has been published (Brown et al. 2003). The crystal structures of DHQS suggested that the enzyme is actively involved in all of the five steps of the reaction and suggested that several highly conserved residues including R130, K152, R264, and H275 played an essential role in catalysis and domain closure (Carpenter et al. 1998; Nichols et al. 2003).
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summarizes the purification protocol, and Figure 3 shows the extent of purification at each stage. AROM was isolated to greater than 98% purity, with a yield of 2.7 mg per liter of cell culture. Purified AROM and the separately expressed DHQS domain were kinetically characterized with respect to their substrate DAHP and the cofactor NAD+, and analyzed using GRAFIT; the results are summarized in Table 2. The comparison (see Table 2) shows that for the substrate DAHP the difference in Km is less than a factor of 3, that the turnover numbers differ by 24%, and that the Km for NAD+ differs by a factor of 3. These values demonstrate that isolation of DHQS domain from the rest of the AROM protein has overall modest effects, and does not drastically compromise enzyme efficiency.
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, and the values in parentheses in the turnover number column are those reported for the equivalent AROM protein from N. crassa (Lambert et al. 1985; Coggins et al. 1987a). With the exception of shikimate dehydrogenase, which has an apparent 35-fold difference, the turnover numbers differ by less than a factor of 4. The shikimate dehydrogenase activity in the A. nidulans AROM is sensitive to substrate inhibition, making accurate measurement of Vmax problematic, and possibly accounts for the difference from turn over number reported for the N. crassa enzyme. Interpretation of the Km values and turnover number for shikimate dehydrogenase shown in Table 2 are therefore subject to this caveat.
Measuring the affinity of an inhibitory metal binding site
We have previously shown that in addition to Zn2+, other metals such as Co2+, Fe2+, Ni2+, Eu3+, and Sm3+ can all reactivate metal-depleted A. nidulans DHQS to differing extents (Moore et al. 1998). In common with DHQS from prokaryotic sources, the A. nidulans DHQS appears to have two metal binding sites, one of which is of low affinity, and acts in an inhibitory manner when occupied. To determine the relative affinities of the two metal binding sites we generated metal-depleted wild-type DHQS and attempted to measure the binding affinities of the two putative Zn2+ sites by ITC. However, the affinity was higher than can be measured by this technique. Therefore, we repeated the experiments using the much weaker binding Ni2+ metal, and found that we were able to measure the affinity of both sites by ITC. Figure 4A shows the isotherm generated, and Figure 4B shows the data after peak integration, subtraction of blank titration data (not shown), concentration normalization, and analysis by the Origin (version 5) suite of programs. A sequential two-site binding model adequately described the data. Analysis of these data showed that the primary metal binding site has a KD of around 245 nM for Ni2+ binding, and the secondary site has KD of 1 mM, an approximately 4000-fold difference. It must be borne in mind that derivation of the KD values for the secondary site from the fit to the model shown is only approximate, and that the possibility of multiple weaker binding sites cannot be excluded. However, the single secondary site model is consistent with the known kinetic characteristics of the wild-type enzyme from A. nidulans and Escherichia coli in the presence of increasing concentrations of metal ions (Bender et al. 1989a; Moore et al. 1998).
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). Table 3 shows the oligonucleotides used to generate these mutants.
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) equate to a greater than an 1100-fold decrease in the kcat/Km ratio.
The mutation R264A is predicted to disrupt the PE2 "pincer-pair," preventing strain propagation through the PE2 loop and 8 transfer helix movement. The effect of this disruption should be to prevent correct domain closure and abolish enzyme activity. Confirming this prediction, the R264A mutant enzyme lacked detectable activity (see Table 2).
Steps 1 and 3 of the complex reaction mechanism are a reversible oxidation of the substrate C5 by NAD+, and H275 is proposed to be part of a proton-shuttling system. Substitution of H275 with leucine is therefore predicted to disrupt the putative proton shuttling system necessary for steps 1 and 3 of the DHQS reaction mechanism. Step 2 is -elimination of the substrate phosphate group, and H275 and K152 are proposed to be part of a phosphate-binding pocket completed by N162, N268, and K356. Steps 4 and 5 consist of ring opening and intramolecular aldol condensation. K152A forms half of the PE1 pincer-pair involved in the initiation of domain closure; thus, this mutation should stop the fold-in of PE1 and strain propagation, so preventing the hinge closure and hence reducing binding of the substrate and potentially abolishing enzyme activity. The side chains of K152 (and possibly R264) are also implicated in preventing epimerization at C2 by interacting with the carboxylic acid group on C2 of the proposed intermediate 4 shown in Figure 1. The side chain of K152 may be involved in stabilizing the negative charge generated during the intramolecular aldol condensation shown in intermediate 5 in Figure 1. Consistent with these predictions, the mutant enzymes H275L and K152A lacked detectable activity.
Biophysical characterization of wild-type and mutant DHQS domains
The disruption of enzymatic activity seen in the mutant DHQS domains shown in Table 2 is consistent with the view that the wild-type residues are all involved in the reaction mechanism described above. However, the data are also consistent with the view that the lack of activity is because the mutations cause some change in the secondary, tertiary, or quaternary structure, or alternatively, that the changes compromise substrate, cofactor, or Zn2+ binding. To address these different possibilities we determined the native molecular weights, near- and far-UV circular dichroism (CD) spectra of the proteins, and characterized their substrate, metal, and cofactor binding ability by differential scanning calorimetry (DSC) and isothermal titration calorimetry (ITC).
CD spectroscopy and native molecular weight determinations
The near- and far-UV CD spectra of the wild-type and mutant DHQS enzymes were compared to look for any evidence that the mutations had caused disruption to the secondary or tertiary structure of the proteins. The near-UV spectra of the wild-type and mutant enzymes were featureless in the region 270 nm to 310 nm, and examples of far-UV spectra are shown in Figure 5. Comparison of the spectra in Figure 5 reveals minimal differences between them, implying that the mutations were not disrupting the DHQS secondary structure.
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and Fig. 6). Taken together, these data indicate than the single-site mutations do not disrupt the folding of the enzyme, and that the reduction in enzyme activity observed in these mutants more likely arises from disruption of specific protein–substrate and/or protein–protein interactions.
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, and typical DSC thermograms are shown in Figure 6. As a control, we also compared the KD for Ni2+ binding to wild-type enzyme as determined by ITC to the value determined by DSC. It should be borne in mind that KD values determined by DSC are likely to be higher than those measured by ITC, as the former measurements are carried out at a higher temperature. The KD value of 435 nM determined by DSC compares favorably with the value of 245 nM determined by ITC for the primary Ni2+ binding site. This reasonable match between the two techniques gives confidence that the use of DSC for a comparison of the relative ligand binding properties of wild-type and mutant DHQS enzymes is valid. The data in Table 4 also show that all the mutant DHQS enzymes tested were able to bind Zn2+ under these conditions, which suggests that the loss of catalytic activity does not arise from perturbation of other residues involved in zinc binding.
Substrate and cofactor binding
Given the success of DSC for the comparative analysis of metal binding to DHQS we chose to use this technique for an analysis of substrate and cofactor binding. The use of DSC has the advantage that much smaller amounts of protein are required for the analysis compared to ITC. Table 4 summarizes the results of the comparative DSC analysis of wild-type and mutant DHQS enzymes with respect to substrate DAHP and the cofactor NAD+. Figure 6 shows typical data for the wild-type and R130K mutant DHQS enzymes. The data in Table 4 show that the KD values for DAHP and NAD+ binding are comparable, with the greatest difference between the wild-type and mutant enzymes being a factor of 5 for DAHP with respect to wild-type and R130A and 6 for NAD+ with respect to wild-type and R264A. This compares sharply with the undetectable enzyme activity of the H275L, K152A, R130A, and R264A mutants and the approximately 135-fold reduction in the activity of the R130K mutant. These data show that the lack of enzymatic activity cannot be ascribed to an inability to bind DAHP or NAD+.
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Discussion |
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Dehydroshikimate dehydratase from the quinic acid utilization pathway has flux control coefficient of -1 in the shikimate pathway, and experiments with A. nidulans have shown that dehydroshikimate dehydratase is present in mycelium at approximately half the quinic acid induced level 2 h after removal of quinic acid (Lamb et al. 1992; Wheeler et al. 1996). Consequently, under these conditions the enzyme can divert flux from the shikimate pathway into the quinic acid utilization pathway and it may be advantageous to the organism to have a low level channelling function associated with the AROM protein (Lamb et al. 1992). However, it could be that the organism derives a selective advantage from having the five enzymatic activities associated with AROM produced in equimolar amounts (Coggins et al. 1987b), and that any low level channelling is simply a consequence of having the five active sites closely juxtaposed.
The site-directed mutagenesis studies we present here show that when the side chains of any of the four amino acids R130, K152, H275, and R264 are changed, DHQS activity is sharply reduced or abolished. On the basis of structural studies these four residues have been identified as important for correct domain closure and catalysis in DHQS. Spectroscopic studies found no changes in the secondary, tertiary, or quaternary structures of the mutant DHQS proteins, and microcalorimetry showed that all the mutant proteins retained their full ligand binding properties. As the mutant enzymes are all able to bind Zn2+, their substrate DAHP, and the cofactor NAD+, the lack of enzyme activity implies that the mutations have disrupted the molecular mechanism of the reaction. Our experiments represent the first rational mutational studies of this enzyme, and show the value of a combined structural and thermodynamic approach in probing complex reaction mechanisms (Carpenter et al. 1998; Nichols et al. 2003).
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Materials and methods |
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-D-thiogalactoside (IPTG) were from Melford Labs UK. Growth media that included Bactotryptone yeast extract and agar were from Beckton Dickinson UK Ltd. Dehydroquinic acid was made as previously described (Grewe and Haendler 1968). SDS PAGE was as previously described (Laemmli 1970).
Kinetic measurements
Assay of the individual activities of the AROM protein were as follows: Dehydroquinate synthase was measured by linking the product of the reaction dehydroquinate to the type 1 dehydroquinase from Salmonella typhi (Moore et al. 1993; Gourley et al. 1999) and measuring the appearance of dehydroshikimate at 240 nm. The standard assay was 1 mL containing 12.5 mM Bis-Tris propane/acetate (pH 7.0), 40 µM ZnSO4, 50 µM NAD+, 340 µM DAHP, and 2 units of Salmonella typhi type 1 dehydroquinase. The type 1 dehydroquinase activity was measured at 240 nm in 1 mL of 12.5 mM Bis-Tris propane/acetate (pH 7.0), containing 300 µM dehydroquinate. Shikimate dehydrogenase was assayed by measuring the NADPH-dependant oxidation of dehydroshikimate at 340 nm in 1 mL containing 50 mM potassium phosphate (pH 7.0), 50 µM NADPH, and 600 µM dehydroshikimate. Shikimate kinase was assayed by measuring the shikimate-dependant oxidation of NADH at 340 nm in 1 mL containing 50 mM triethanolamine/KOH (pH 7.0), 50 mM KCl, 5 mM MgSO4, 0.1 mM shikimic acid, 2.5 mM ATP, 0.5 mM PEP, 50 µM NADH, 6.6 units of pyruvate kinase mL–1, and 13.5 units of lactate dehydrogenase mL–1. All assays were carried out at 25°C, and enzyme kinetic data were analyzed using nonlinear regression with the program GRAFIT (Erithacus software).
Circular dichroism
Circular dichroism spectra were recorded from 10 accumulative scans at 20°C between 180 nm to 250 nm (far-UV) and 250 nm to 320 nm (near-UV) using a Jasco J-810 spectropolarimeter. Scans were recorded at standard sensitivity with a band width of 2 nm, scanning speed of 50 nm min–1and a protein concentration of 0.5 mg mL–1. Specific ellipticity values were calculated using concentrations derived from the absorption measurements at 280 nm on a Shimadzu UV-1601 (
280 = 36,160). Calculations and data analysis were carried out using Jasco Spectra analysis software. Each spectrum was corrected by subtraction of a comparative blank.
Differential scanning calorimetry
Protein thermal transition characteristics in solution were determined by differential scanning calorimetry (DSC) using a Micro-Cal VP-DSC (cell volume 0.52 mL). Samples were scanned over a range from 25°C–65°C, with a scan rate of 90°C h–1, and a filtering period of 16 sec with protein at 20 µM–23 µM and an appropriate buffer in the reference cell. Solutions were gently degassed prior to loading. Thermograms were corrected for instrumental baseline, using control buffer:buffer scans under identical conditions, and analyzed using standard MicroCal ORIGIN software. Protein concentrations were measured spectrophotometrically using the molar absorption coefficient calculated from the amino acid sequence by the vector NTI suite of programs as previously described (Gill and von Hippel 1989). Ligand-induced shifts in thermal transition midpoint temperatures (Tm) were used to estimate approximate ligand affinities using standard thermodynamic methods as previously described (Cooper 1999; Cooper et al. 2000).
Isothermal titration calorimetry
Protein/ligand binding constants were also determined by ITC using a MicroCal VP-ITC microcalorimeter. ITC measurements were carried out at 25°C, and the titrations were carried out using a 250-µL syringe, with stirring at 300 rpm. Each titration consisted of a preliminary injection of 2 µL, followed by 24 injections of 10 µL into a cell containing approximately 1.4 mL of protein sample at 50 µM–100 µM. To correct for dilution and mixture effects a series of baselines were carried out, where injections of substrate/ ligand were carried out into buffer alone. Data were analyzed using MicroCal ORIGIN software and baselines subtracted from data to obtain accurate heat exchanges. Protein solutions for calorimetry were extensively dialyzed against buffer, and the equilibrated dialysis buffer used for ligand solutions and controls to minimize heats of dilution effects.
Site-directed mutagenesis
Site-directed mutagenesis was carried out as previously described (Hemsley et al. 1989).
Native molecular weight determinations
A Superdex 200 FPLC column in conjunction with an AKTA explorer protein purifier was used to determine the molecular weight/oligomeric state of wild-type and mutant DHQS. The void volume of the column was determined with blue dextran and cytochrome c (12 kDa), carbonic anhydrase (29 kDa), bovine serum albumin (66 kDa), and alcohol dehydrogenase (150 kDa) were used to compile a standard curve from which the sample molecular weights were extrapolated.
Purification of the AROM protein
Overproduction of the A. nidulans AROM enzyme was carried out in E. coli expression strain Codon Plus containing the plasmid pEKA15 (Moore 1992). A 200 mL broth starter culture containing 100 µg mL–1ampicillin and 35 µg mL–1chloramphenicol was grown through the day at 27°C and then used to seed 18 x 2-liter flasks containing 500 mL of drug supplemented broth. These cultures were grown at 27° C to an attenuance (D550) of 0.75, and then induced by the addition of IPTG to a final concentration of 0.2 mg mL–1 followed by a further 6-h growth at 27°C. Cells were harvested by centrifugation and stored frozen at –18°C until needed. Cell paste (50 g) was thawed at room temperature, and cells disrupted by sonication at 15 µ in 450 mL of buffer containing 0.1 M Tris HCl, 1 mM DTT, 1 mM benzamidine (pH 7.5), 40 µM ZnSO4 (buffer 1). A cell free extract was obtained by removing cellular debris by centrifugation at 10,000g for 42 min at 4°C. The supernatant was made 30% saturated with ammonium sulphate, and the precipitated proteins removed by centrifugation at 10,000g for 45 min at 4°C. The supernatant was then made 55% saturated with ammonium sulphate and the precipitated protein recovered by centrifugation centrifugation at 10,000g for 45 min at 4°C. The recovered protein pellet was dissolved in a 50 mM potassium phosphate (pH 7.2), 1.0 M ammonium sulphate, 150 mM NaCl, 1 mM DTT (buffer 2) to a final volume of 400 mL. The supernatant was applied to a phenyl Sepharose column pre equilibrated with buffer 2. Following a 500-mL wash with buffer 2, the column was eluted with a 1-L linear reverse ammonium sulphate gradient made by using 500 mL of buffer 2 and 500 mL of 50 mM potassium phosphate (pH 7.2), 150 mM NaCl, 1 mM DTT (buffer 3), and collecting 10 mL fractions. A further 300 mL of buffer 3 was applied and fractions monitored for AROM protein by assaying the DHQS and type 1 dehydroquinase activities. Sample loading and elution of the phenyl Sepharose column were carried out at the maximum flow rate without the use of a pump. Fractions were also monitored by SDS PAGE (7.5% separating gel) and appropriate fractions were pooled and made 60% saturated with ammonium sulphate. The precipitated AROM protein was harvested by centrifugation at 10,000g for 45 min, then resuspended in the minimum required volume of 50 buffer 4. The sample was then applied to a Sephacryl S300 column (125 x 2.5 cm) at a flow rate of 0.5 mL min–1while collecting 10-mL fractions. Fractions containing AROM were identified by enzyme assay and analyzed by SDS PAGE. Appropriate fractions were pooled, dialyzed against 2 x 5l of 25 mM potassium phosphate (pH 7.2), 1 mM DTT (buffer 5) and, following filtration through a 0.45-µm filter, applied in halves to two ceramic hydroxyapatite columns (9 x 5 cm). The columns were washed with 500 mL each of buffer 5 and then eluted with a 1-L gradient of 25–400 mM potassium phosphate (pH 7.2), 1 mM DTT collecting 10-mL fractions. AROM was located by enzyme assay and SDS PAGE. For kinetic analysis the AROM protein was dialyzed into 50 mM potassium phosphate (pH 7.2), 1 mM DTT, 40 µM ZnSO4, and further purified by FPLC using a 5-mL DEAE FF Hi-trap column. AROM protein was eluted with a 160-mL 0 to 0.5-M NaCl gradient collecting 2-mL fractions. AROM was located by enzyme assay and SDS PAGE with a typical yield of 2.7 mg L–1at greater than 98% purity. The wild-type and mutant DHQS domains of the AROM protein were purified as previously described (Moore et al. 1994).
Metal depletion of buffers and enzymes
A 100-mL chelating resin column was prepared by sequentially washing with two volumes of 1 N HCl, 5 volumes of deionized water, 2 volumes of 1 N NaOH, followed by deionized water until the column effluent was at neutral pH. Once neutrality was reached, buffers previously prepared in deionized water were passed through the column to remove divalent cations. All glass and plasticware used was rinsed in buffer containing 10 mM EDTA and then rinsed five times in metal-free water. Metal-free enzyme was prepared by incubation with 1 mM EDTA for 1 h at 4°C while gently mixing the solution. EDTA was removed from the samples by dialysis once into buffer containing 50 µM EDTA and then into three changes of metal-free buffer (typically, a 20-mL sample in 5 L of dialysis buffer) at 4°C, with at least 12 h between each change of buffer.
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The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.
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