The Roles of His-167 and His-275 in the Reaction Catalyzed by Glutamate Decarboxylase from Escherichia coli
1998; Elsevier BV; Volume: 273; Issue: 4 Linguagem: Inglês
10.1074/jbc.273.4.1939
ISSN1083-351X
AutoresAngela Tramonti, Daniela De Biase, Anna Giartosio, Francesco Bossa, Robert A. John,
Tópico(s)Enzyme Structure and Function
ResumoTwo histidine residues in glutamate decarboxylase from Escherichia coli, potential participants in catalysis because they are conserved among amino acid decarboxylases and because they are at the active site in the homologous enzyme ornithine decarboxylase, were mutated. His-275 is shown to bind the cofactor pyridoxal 5′-phosphate but not to contribute directly to catalysis. The H275N enzyme was unable to bind the cofactor whereas the H275Q mutant contained 50% of the normal complement of cofactor and its specific activity (expressed per mole of cofactor) was 70% of that of the wild-type enzyme. The H167N mutant bound the cofactor tightly, its specific activity was approximately half that of the wild-type enzyme and experiments in D2O showed that it catalyzed replacement of the carboxyl group with retention of configuration as does the wild-type enzyme. Comparison of reaction profiles by observing changes in the absorbance of the cofactor after stopped-flow mixing, revealed that a slow reaction, in which approximately one-third of the wild-type enzyme is converted to an unreactive complex during catalysis, does not occur with the H167N mutant enzyme. This reaction is attributed to a substrate-induced conformational change, a proposal that is supported by differential scanning calorimetry. Two histidine residues in glutamate decarboxylase from Escherichia coli, potential participants in catalysis because they are conserved among amino acid decarboxylases and because they are at the active site in the homologous enzyme ornithine decarboxylase, were mutated. His-275 is shown to bind the cofactor pyridoxal 5′-phosphate but not to contribute directly to catalysis. The H275N enzyme was unable to bind the cofactor whereas the H275Q mutant contained 50% of the normal complement of cofactor and its specific activity (expressed per mole of cofactor) was 70% of that of the wild-type enzyme. The H167N mutant bound the cofactor tightly, its specific activity was approximately half that of the wild-type enzyme and experiments in D2O showed that it catalyzed replacement of the carboxyl group with retention of configuration as does the wild-type enzyme. Comparison of reaction profiles by observing changes in the absorbance of the cofactor after stopped-flow mixing, revealed that a slow reaction, in which approximately one-third of the wild-type enzyme is converted to an unreactive complex during catalysis, does not occur with the H167N mutant enzyme. This reaction is attributed to a substrate-induced conformational change, a proposal that is supported by differential scanning calorimetry. Glutamate decarboxylase is a member of a large family of pyridoxal-phosphate (PLP) 1The abbreviation used is: PLP, pyridoxal 5′-phosphate.-dependent enzymes which catalyze a wide variety of different reactions on their amino acid substrates (1Sandmeier E. Hale T.I. Christen P. Eur. J. Biochem. 1944; 221: 997-1002Crossref Scopus (217) Google Scholar). Although the enzyme from Escherichia coli is the most studied of the amino acid decarboxylases, its three-dimensional structure has not been solved. However, sequence comparisons show that it is homologous to ornithine decarboxylase fromLactobacillus 30a, the three-dimensional structure of which (2Momany C. Ghosh R. Hackert M.L. Protein Sci. 1995; 4: 849-854Crossref PubMed Scopus (83) Google Scholar, 3Momany C. Ernst S. Ghosh R. Chang N.-L. Hackert M.L. J. Mol. Biol. 1955; 252: 643-655Crossref Scopus (111) Google Scholar) shows two histidine residues, His-223 and His-354, close to and on either side of the coenzyme (2Momany C. Ghosh R. Hackert M.L. Protein Sci. 1995; 4: 849-854Crossref PubMed Scopus (83) Google Scholar, 3Momany C. Ernst S. Ghosh R. Chang N.-L. Hackert M.L. J. Mol. Biol. 1955; 252: 643-655Crossref Scopus (111) Google Scholar). These histidines are conserved in the sequences of most, if not all, of the PLP-dependent amino acid decarboxylases (2Momany C. Ghosh R. Hackert M.L. Protein Sci. 1995; 4: 849-854Crossref PubMed Scopus (83) Google Scholar). There is no doubt that His-275 ofE. coli glutamate decarboxylase aligns with His-354 of ornithine decarboxylase since, in both enzymes as in other amino acid decarboxylases, it immediately precedes the lysine residue that forms an imine with the coenzyme (Fig. 1). In ornithine decarboxylase, the imidazole ring of His-354 contributes to binding the 5′-phosphate of the cofactor and is considered unlikely to participate in catalysis unless it is displaced by a substrate-induced conformational change (2Momany C. Ghosh R. Hackert M.L. Protein Sci. 1995; 4: 849-854Crossref PubMed Scopus (83) Google Scholar, 3Momany C. Ernst S. Ghosh R. Chang N.-L. Hackert M.L. J. Mol. Biol. 1955; 252: 643-655Crossref Scopus (111) Google Scholar). Alignment of His-167 of glutamate decarboxylase is less clear because sequence similarity between the two enzymes is weak in this region (Fig. 1). However, the presence, in ornithine decarboxylase, of the imidazole ring of His-223 just below the imine of the cofactor on the re face, suggests that this arrangement may be a common feature of the amino acid decarboxylases. His-167 in glutamate decarboxylase is the only histidine residue that can reasonably occupy this position unless there are major differences between the folds of the two proteins. Several stages in the catalytic mechanism of amino acid decarboxylation involve proton transfers for which histidine residues might be responsible (Scheme ins;1821s1}1). The formation of an external aldimine (III) requires multiple proton transfers and, after decarboxylation to give the quinonoid intermediate (IV), the carboxyl group is replaced by a proton. Further proton transfers, analogous to those occurring in external aldimine formation, are required for liberation of the product 4-aminobutyrate. An additional protonation occurs in a side reaction where a proton is added to C4′ of the cofactor rather than Cα of the substrate to give an external aldimine of pyridoxamine phosphate with succinic semialdehyde. This reaction, which occurs only once in 3 × 105 turnovers in E. coli glutamate decarboxylase (4Almazov V.P. Morozov Iu.V. Savin F.A. Sukhareva B.S. J. Mol. Biol. U.S.S.R. 1985; 2: 359-370Google Scholar), is a feature of most amino acid decarboxylases (5Sukhareva B.S. Braunstein A.E. Mol. Biol. (Engl. Transl.). 1971; 5: 241-252PubMed Google Scholar, 6O'Leary M.H. Baughn R.L. J. Biol. Chem. 1977; 252: 7168-7173Abstract Full Text PDF PubMed Google Scholar, 7Minelli A. Charteris A.T. Borri-Voltattorni C. John R.A. Biochem. J. 1979; 183: 361-368Crossref PubMed Scopus (23) Google Scholar, 8Grant P.L. Basford J.M. John R.A. Biochem. J. 1987; 241: 699-704Crossref PubMed Scopus (14) Google Scholar, 9Martin D.L. Prog. Biophys. Mol. Biol. 1993; 60: 17-28Crossref PubMed Scopus (22) Google Scholar). In some of these enzymes this abortive transamination is kinetically more prominent and is almost certainly metabolically important because it inactivates the enzyme by leaving the cofactor as pyridoxamine phosphate. Considerable attention has been given to the stereochemistry of amino acid decarboxylation and subsequent protonation. Decarboxylation requires the carboxyl group to be orthogonal to the plane comprising the cofactor pyridinium ring and the imine double bond (10Dunathan H. Proc. Natl. Acad. Sci. U. S. A. 1966; 55: 712-716Crossref PubMed Scopus (343) Google Scholar). The proton which replaces the carboxyl group at Cα arrives from the same direction in which the carboxyl group has left (11Yamada H. O'Leary M.H. Biochemistry. 1978; 17: 669-672Crossref PubMed Scopus (39) Google Scholar). Retention of configuration has also been observed in all other amino acid decarboxylases in which the question has been investigated (12Stevenson D.E. Akhtar M. Gani D. Biochemistry. 1990; 29: 7660-7666Crossref PubMed Scopus (10) Google Scholar, 13Stevenson D.E. Akhtar M. Gani D. Biochemistry. 1990; 29: 7631-7647Crossref PubMed Scopus (23) Google Scholar, 14No Z. Sanders C.R. Dowhan W. Tsai M.-D. Bioorg. Chem. 1988; 16: 184-188Crossref Scopus (7) Google Scholar). Kinetic evidence (15Akhtar M. Stevenson D.E. Gani D. Biochemistry. 1990; 29: 7648-7660Crossref PubMed Scopus (25) Google Scholar) indicates that, in methionine decarboxylase, a histidine residue protonates at Cα, whereas a lysine residue protonates at C4′ and it has been argued that lysine and histidine perform the corresponding protonations in glutamate decarboxylase (16Tilley K. Akhtar M. Gani D. J. Chem. Soc. Perkin Trans. 1994; 1: 3079-3087Crossref Scopus (15) Google Scholar). In the present work, we have examined the effect of mutations at His-167 and at His-275 on the kinetic and structural properties of the enzyme. 4-Aminobutyrate aminotransferase was prepared as described (17De Biase D. Barra D. Bossa F. Pucci P. John R.A. J. Biol. Chem. 1991; 266: 20056-20061Abstract Full Text PDF PubMed Google Scholar). D2O was from Sigma, Vent polymerase was from New England Biolabs. Restriction enzymes, T4 DNA ligase, the agarose gel DNA extraction kit, and Gabase were from Boehringer. The T7 sequencing kit, DEAE-Sepharose, and Sephadex G-25 were from Pharmacia. [α-35S]dATP (1000 Ci/mmol) was from NEN Life Science Products. Ingredients for bacterial growth were from Difco. Oligonucleotides were from Genenco. Other chemicals were from BDH. Site-directed mutagenesis was performed by overlap extension polymerase chain reactions (18Higuchi R. Innis M.A. Gelfand D.H. Snisky J.J. White T.J. Higuchi R. Innis M.A. Gelfand D.H. Snisky J.J. White T.J. PCR Protocols: A Guide to Methods and Applications. Academic Press Inc., San Diego1990: 177-183Google Scholar). External primers annealing over the N- and C-terminal sequences were those used in the construction of the expression plasmid containing thegadB gene (19De Biase D. Tramonti A. John R.A. Bossa F. Protein Exp. Purif. 1996; 8: 430-438Crossref PubMed Scopus (90) Google Scholar). Mutagenic primers for H167N were 5′-GCTGGAATAAATTCGCCC-3′ and its complementary sequence. Those for H275N and H275Q were 5′-GCTTCAGGCCAGAAATTCG-3′ and 5′-GCTTCAGGCAATAAATTCG-3′ and their complementary sequences, respectively. Plasmid pQgadB was used as template. The products of polymerase chain reactions (25 cycles), carried out with 2.5 units of Vent polymerase with denaturation at 95 °C for 1 min, annealing at 45 °C (H167N) or 48 °C (H275N and H275Q) for 1 min, and extension at 74 °C for 2 min, were used as templates with the external primers to generate the complete coding sequence of glutamate decarboxylase. Fragments (NcoI/EcoRV for H167N and EcoRV/HindIII for H275N and H275Q) were subcloned into pQgadB. The newly inserted parts of the expression constructs, pQgadH167N, pQgadH275N, and pQgadH275Q, were sequenced and the plasmids were used to transform E. coli JM109 carrying the plasmid pREP4. Expression, purification, and assay of mutant forms of glutamate decarboxylase were as described for the wild-type enzyme (19De Biase D. Tramonti A. John R.A. Bossa F. Protein Exp. Purif. 1996; 8: 430-438Crossref PubMed Scopus (90) Google Scholar) except where stated. N-(5′-Phosphopyridoxyl)glutamate was prepared by treating 0.5 m sodium glutamate and 0.5 mmPLP (pH 4.5) with sodium cyanoborohydride (10 mm). Apo-forms of wild-type and mutant forms of glutamate decarboxylase (10 mg/ml in 0.1 m piperazine-HCl, pH 4.5, containing 0.1m dithiothreitol) were reconstituted with a 5-fold molar excess of N-(5′-phosphopyridoxyl)glutamate for 1 h (25 °C). Samples were concentrated, separated on Sephadex G-25, and the proteins dialyzed against 20 mm sodium acetate (pH 3.6) containing 0.1 mm dithiothreitol. Thermal unfolding of degassed samples (1.5–2.0 mg/ml) was analyzed under nitrogen pressure on a MicroCal MC-2D differential scanning calorimeter (MicroCal, Inc., Northampton, MA). Results were corrected for instrumental baseline and normalized for protein concentration. No reversibility was observed in a second heating cycle. Enzyme samples were brought into 99.5% D2O by repeated concentration and dilution. The enzyme (0.8 μm) was mixed with glutamate (14.25 mm). DCl was added to maintain constant pD of 4.6 (reading on pH meter = 4.2). At the end of the reaction (2 h) the solution was neutralized by adding solid Tris, lyophilized, and redissolved in 0.5 ml of D2O. NMR spectra were determined using a Bruker AMX 360 spectrometer. The stereochemistry of the deuterated 4-aminobutyrate was determined by repeating the NMR analysis after an overnight incubation in the presence of 4-aminobutyrate aminotransferase (0.36 mg/ml). Stopped-flow experiments were performed on a SF-1 stopped-flow spectophotometer (Hi-Tech, Salisbury, United Kingdom). Product formation during the period from 0.2 to 7 s was measured using a quenched flow apparatus (8Grant P.L. Basford J.M. John R.A. Biochem. J. 1987; 241: 699-704Crossref PubMed Scopus (14) Google Scholar, 20Eccleston J.F. Messerschmidt R.G. Yates D.W. Anal. Biochem. 1980; 106: 73-77Crossref PubMed Scopus (8) Google Scholar). Reactions lasting longer than 7 s were stopped manually. Curve fitting and statistical analyses were performed using the data manipulation software Scientist (Micromath, Salt Lake City, UT). Absorption spectra were measured with a Hewlett-Packard model 8452 diode-array spectophotometer. CD spectra were recorded as the average of 3 scans on a Jasco 710 spectropolarimeter equipped with a DP 520 processor at 25 °C using a 2-mm quartz cell. 4-Aminobutyrate was measured by high performance liquid chromatography (8Grant P.L. Basford J.M. John R.A. Biochem. J. 1987; 241: 699-704Crossref PubMed Scopus (14) Google Scholar) or using a commercial preparation containing 4-aminobutyrate aminotransferase and succinic semialdehyde dehydrogenase (19De Biase D. Tramonti A. John R.A. Bossa F. Protein Exp. Purif. 1996; 8: 430-438Crossref PubMed Scopus (90) Google Scholar). The far UV CD spectra of the H167N and H275Q mutants were identical with that of the wild-type enzyme (Fig. 2) indicating that these mutations had not introduced major changes in the global structure of the enzyme. The H275N mutant precipitated too rapidly at low concentrations to permit Far UV CD analysis. Yields of the H167N mutant enzyme after the standard purification, which does not include added PLP, were as high as those of the wild-type enzyme. This form, like the wild-type enzyme, was stable for many months at 4 °C and for several hours at room temperature. Its absorption and CD spectra (Fig. 2) were almost identical with those of the wild-type enzyme, indicating that one molecule of cofactor was bound per monomer. The specific activity (126 μmol min−1 mg−1) was approximately half that of the wild-type enzyme. Differential scanning calorimetry of the wild-type enzyme (Fig. 3) showed that the transition temperature of the apoenzyme reconstituted with the covalent substrate-cofactor adduct,N-(5′-phosphopyridoxyl)glutamate, was 8 °C higher than that of the native holo-enzyme (51 °C) suggesting that binding of glutamate at the active site stabilizes the structure considerably. When the same experiment was carried out on the H167N enzyme, the holoenzyme was found to be 4 °C more stable than the wild-type enzyme but the mutant was not further stabilized when the cofactor was replaced by N-(5′-phosphopyridoxyl)glutamate (Fig. 3).Figure 3Differential scanning calorimetry of wild-type and H167N mutant glutamate decarboxylase. Thermal denaturation profiles of (a) unliganded holoenzyme and (b) apoenzyme reconstituted with the covalent adduct of cofactor and substrate, N-(5′-phosphopyridoxyl)glutamate. The samples were scanned at a heating rate of 60 °C/h.View Large Image Figure ViewerDownload Hi-res image Download (PPT) Purification of the H275N enzyme according to the standard protocol resulted in a yield of enzyme protein 63% of that normally achieved with the wild-type enzyme. The absorption spectrum showed that this form contained no cofactor. Inclusion of PLP (0.1 mm) in solutions used for dialysis gave an enzyme preparation that clearly bound the cofactor but with an absorption spectrum (Fig. 2 a) which showed more 340 nm chromophore than is present in the wild-type enzyme. The instability of this form of the enzyme prevented measurement of its kinetic properties. The H275Q mutant was more stable, although, after the standard purification, its absorption spectrum (Fig. 2 a) showed that its coenzyme content was lower than that of the wild-type enzyme. Treatment with 0.1 m NaOH to release the PLP showed it to contain 0.5 mol of cofactor per mol of subunit. Its specific activity was 80 units/mg (35% that of the wild-type enzyme). Thus, per mole of cofactor, the specific activity of this mutant is 70% that of the wild-type enzyme. Glutamate decarboxylase undergoes a sharp, pH-dependent, transition in which a 420-nm absorbing form, presumed to be the protonated internal aldimine, is converted to a 340-nm form. The transition involves uptake of multiple protons and lasts several seconds, demonstrating that a slow, protonation-dependent reaction is involved as well as the protonation itself (21Shukuya R. Schwert G.W. J. Biol. Chem. 1960; 235: 1653-1657Abstract Full Text PDF PubMed Google Scholar, 22O'Leary M.H. Brummund W. J. Biol. Chem. 1974; 249: 3737-3745Abstract Full Text PDF PubMed Google Scholar). The midpoint of the transition (pH 5.5) is within the range expected for protonation of a histidine imidazole. Comparison of absorption spectra of the wild-type enzyme at different pH values with those of the H167N and H275Q mutants showed that both mutants behaved as the wild-type, demonstrating that neither His-167 nor His-275 is responsible for the pH-dependent transition. In ornithine decarboxylase, His-223 which we hypothesize to be equivalent to His-167 in glutamate decarboxylase, occupies a position on the re face of the cofactor. Lys-355 in ornithine decarboxylase is on the siface and the equivalent residue in glutamate decarboxylase is Lys-276. In the aminotransferases from the same family, the proton transfers occur on the si face and are mediated by the equivalent of Lys-276 (23Julin D.A. Kirsch J.F. Biochemistry. 1989; 28: 3825-3833Crossref PubMed Scopus (53) Google Scholar). It is known that decarboxylation of glutamate occurs with retention of configuration (11Yamada H. O'Leary M.H. Biochemistry. 1978; 17: 669-672Crossref PubMed Scopus (39) Google Scholar) but it is not known from which side the carboxyl group leaves. Quantum mechanical calculations (4Almazov V.P. Morozov Iu.V. Savin F.A. Sukhareva B.S. J. Mol. Biol. U.S.S.R. 1985; 2: 359-370Google Scholar) confirm that the Cα-COO− bond has two positions of maximal lability, each perpendicular to the coenzyme ring as predicted by Dunathan (10Dunathan H. Proc. Natl. Acad. Sci. U. S. A. 1966; 55: 712-716Crossref PubMed Scopus (343) Google Scholar), but pointing in opposite directions. It is possible therefore that the carboxyl group leaves from the re face and that protonation of Cα is mediated from this face by His-167. To test this possibility, the 4-aminobutyrate produced by wild-type and H167N glutamate decarboxylase in D2O was analyzed by NMR. In both cases the signal from protons at C4 was halved, indicating that the 4-aminobutyrate produced by each form of the enzyme was monodeuterated at C4 (equivalent to Cα of glutamate). When the products were treated in D2O with 4-aminobutyrate aminotransferase, which labilizes the pro-S proton at C4 of 4-aminobutyrate exclusively (24Bouclier M. Jung M.J. Lippert B. Eur. J. Biochem. 1979; 98: 363-368Crossref PubMed Scopus (40) Google Scholar), the signal from C4 protons was lost in each case. This indicated incorporation of a second deuterium at C4. Significant differences between the H167N mutant and the wild-type enzyme were observed when changes in the absorption of the cofactor were measured after stopped-flow mixing with glutamate. The reaction with the most readily interpreted kinetic behavior was that of the H167N mutant. Changes in absorbance were largest at 322 nm. A large increase, complete within the mixing time (2 ms), was followed by a smaller increase which followed a single exponential (Fig. 4a, k = 30 ± 1 s−1). Thereafter, absorbance fell as expected for a system in which the concentration of ES complex returns to zero as substrate is consumed in the reaction (Fig. 4 b). A single experiment of this kind, in which the pseudo-equilibrium mixture of enzyme-substrate complexes is observed throughout the full course of the reaction from [S] ≫ K m to [S] = 0, provides an accurate estimate of the steady state constantsk cat and K m . (The system is not subject to product inhibition since the kinetic constants were independent of initial substrate concentration and inclusion of 4-aminobutyrate had no effect.) After exclusion of the first 0.2 s because of the fast pre-steady state reaction, the profile gave an excellent fit to the Michaelis equation and provided values for the steady state constants k cat = 27.8 ± 0.1 s−1, K m = 10.6 ± 0.1 mm. The similarity of the values obtained fork cat and the rate constant characterizing the pre-steady state transient, suggests that the reaction reported by the latter is largely rate-determining in the overall process. At 420 nm the absorbance changes observed were much smaller (total absorbance change 0.02) and in the opposite direction but they were governed by the same rate constants. Because of the unusual kinetic behavior of the wild-type enzyme (see later), product formed during the first 10 s of reaction was measured using the quenched-flow apparatus. Conditions for the enzyme-catalyzed reaction were exactly as in the stopped-flow experiment. Fig. 4 c shows the result. The line of best fit through the experimental points from the quenched flow experiment is 0.98 mm s−1. The values ofk cat and K m determined from the stopped flow experiment predict 0.93 mms−1. The absorbance changes observed with the wild-type enzyme were more complex (Fig. 5a). Large changes were observed at 322, 360, and 420 nm. At 322 and 360 nm an initial increase was complete within the mixing time. This was followed by an increase lasting approximately 5 s. After exclusion of the first 5 s, the reaction showed A 322 andA 360 declining as expected for a system which is monitoring ES complex as substrate is consumed. A fit to the Michaelis equation gave values of k cat = 55 s−1 and K m = 6.7 mm. At 420 nm, changes were in the opposite direction but were characterized by the same rate constants. Inclusion of the product 4-aminobutyrate (50 mm) did not affect the kinetic profile, confirming earlier observations that product inhibition is insignificant (5Sukhareva B.S. Braunstein A.E. Mol. Biol. (Engl. Transl.). 1971; 5: 241-252PubMed Google Scholar). The complete kinetic profile was fitted, by nonlinear regression using differential equations, to a scheme in which, after initial rapid reaction to form a pseudo-equilibrium mixture of complexes (ES), a species (EX) is formed which does not lie on the path leading to product formation (Scheme 2). The data (Fig. 5 a) fit well to Scheme 2 and give kinetic constants of k 1 = 0.32 ± 0.02 s−1, k 2 = 0.4 ± 0.1 s−1, k c = 98 ± 32 s−1, K s = 11 ± 2 mm. The fit at 322 nm assigned extinction coefficients to the speciesES and EX of 2748 ± 58m−1 cm−1 and 4290 ± 810m−1 cm−1, respectively. The unusual feature of the reaction profile, namely the transient phase occurring in the first few seconds, is absent from the profile of the His-167 mutant enzyme. When this experiment was conducted with H275Q glutamate decarboxylase, the kinetic profile was very similar to that observed with wild-type enzyme. The initial transient phase was present and the results fitted well to Scheme 2 with best fit valuesk 1 = 0.12 ± 0.01 s−1,k 2 = 0.53 ± 0.02 s−1,K s = 12.4 ± 0.3 mm, andk c = 49 ± 1 s−1.Figure 5Changes in cofactor absorbance and product concentration during the reaction catalyzed by wild-type glutamate decarboxylase. Reactions were conducted (a) in H2O and (b) in D2O. Conditions as in Fig. 4. The solid lines are those of best fit to Scheme 2 using the constants given in the text. Only one in 10 of the data points collected and used in fitting is shown so that the fit of experimental points to the theoretical lines can be seen.View Large Image Figure ViewerDownload Hi-res image Download (PPT) The absence of the transient phase from the H167N mutant enzyme is the most striking difference between this mutant and the wild-type enzyme. Thus the nature of the chromophore formed in the process is important in understanding the role played by His-167. The approximately equal values of k 1 and k 2 and the approximately 2-fold difference between k catand k c suggest that, during the first few seconds of reaction, approximately half of the enzyme converts into a form which cannot react further. Such a process should therefore affect the rate of product formation and result in a burst at the beginning of the reaction. A quenched-flow apparatus was therefore used to measure the formation of 4-aminobutyrate during the first 15 s of reaction. The results (Fig. 5 a, inset) show a clear burst which fits well to the same rate constants and confirms that formation of the 345-nm chromophore is effectively inhibitory. During the investigations aimed at measuring the incorporation of deuterium into the product 4-aminobutyrate (see "Deuterium Exchange Reactions"), we noticed large differences in the rates at which acid needed to be added to maintain constant pH in the reactions catalyzed by the wild-type enzyme. An earlier study of the effects of D2O on the reaction catalyzed by glutamate decarboxylase has already shown large solvent isotope effects (25O'Leary M.H. Yamada H. Yapp C.J. Biochemistry. 1981; 20: 1476-1481Crossref PubMed Scopus (32) Google Scholar) and these have been cited in support of a proposal that, after the decarboxylation step, protonation at Cα is mediated by a histidine residue (16Tilley K. Akhtar M. Gani D. J. Chem. Soc. Perkin Trans. 1994; 1: 3079-3087Crossref Scopus (15) Google Scholar). To investigate these effects further, we conducted rapid mixing experiments in D2O with wild-type enzyme and with the H167N mutant. We measured the steady state constants by analyzing the complete reaction profile after omitting the first 5 s containing the transient. Our results for the wild-type enzyme (Fig. 5 b) are in broad agreement with those published earlier (25O'Leary M.H. Yamada H. Yapp C.J. Biochemistry. 1981; 20: 1476-1481Crossref PubMed Scopus (32) Google Scholar). We found k cat(26.3 ± 0.8 s−1) to be 2.7 times smaller andk cat/K m (2.3 ± 0.1 mm−1 s−1) to be 3.5 times smaller than in H2O. We also observed a pronounced solvent isotope effect on the slow transient, the amplitude of which was greatly increased both at 420 and 322 nm. The whole process, including the slow transient, fitted well to Scheme 2 and gave constants of best fitk 1 = 0.84 ± 0.37 s−1,k 2 = 0.4 ± 0.1 s−1,k c = 74 ± 38 s−1, andK s = 32 ± 12 mm. The fit assigned extinction coefficients to the species ES and EX of 1982 ± 130 m−1 cm−1 and 4320 ± 610 m−1 cm−1, respectively. The increased size of the transient suggested that, if it is due to slow formation of a species off the reaction pathway, the course of product formation should be characterized by a more pronounced burst when the reaction is carried out in D2O. Fig. 5 b (inset) shows the course of 4-aminobutyrate production measured by quenched-flow over the first 10 s of reaction and confirms that a pronounced burst is present. The constants of best fit for these data were k 1= 2.3 ± 1.7 s−1, k 2 = 0.65 ± 0.15 s−1, k c = 42 ± 19 s−1, K s = 32 ± 12 mm. Although the constants derived from the different types of experiment are not as closely similar as those from the corresponding experiments in H2O, we consider that, in view of the large standard deviations, Scheme 2 also provides a satisfactory explanation. When the H167N mutant enzyme was used to catalyze the deuterium exchange reaction, the amount of acid required to maintain constant pH in unbuffered solution in D2O or H2O was the same. The reaction profile monitored at 322 nm after stopped-flow mixing was also similar to that obtained in H2O but analysis showed that k cat (38.2 ± 0.1 s−1) was increased 1.4-fold andk cat/K m (1.66 mm−1 s−1) was decreased 1.6-fold. The slow conversion to an unreactive intermediate does not occur with this form of the enzyme so that this complication is eliminated as a contribution to the solvent isotope effects. We note that, in the absence of the side reaction, a positive isotope effect onk cat was observed rather than the larger negative effect seen in the reaction of the wild-type enzyme. The increased amplitude of the slow transient phase in D2O and the accompanying more marked nature of the burst in product formation strengthen the evidence that an unreactive complex is formed in this phase. The 345-nm chromophore formed in the slow side reaction has been proposed (4Almazov V.P. Morozov Iu.V. Savin F.A. Sukhareva B.S. J. Mol. Biol. U.S.S.R. 1985; 2: 359-370Google Scholar) to arise by tautomerization of the quinonoid structure formed after decarboxylation (Scheme FS1; IV). In this proposal, the proton on N of the cofactor-substrate imine has transferred to O3′ of the cofactor to give an intermediate which was considered to be more likely to protonate at C4′ than at Cα and thereby to lead to the abortive transamination reaction. The H167N mutant provid
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