Reaction Mechanism of Fructose-2,6-bisphosphatase
A MUTATION OF NUCLEOPHILIC CATALYST, HISTIDINE 256, INDUCES AN ALTERATION IN THE REACTION PATHWAY*

Hiroyuki Mizuguchi, Paul F. CookDagger , Chia-Hui TaiDagger , Charles A. Hasemann§, and Kosaku Uyeda

From Research Service, Dallas Veterans Affairs Medical Center, Dallas, Texas 75216, the § Departments of Biochemistry and Internal Medicine, University of Texas Southwestern Medical Center at Dallas, Dallas, Texas 75216, and the Dagger  Department of Chemistry and Biochemistry, University of Oklahoma, Norman, Oklahoma 73019

    ABSTRACT
Top
Abstract
Introduction
References

A bifunctional enzyme, fructose-6-phosphate,2-kinase/fructose 2,6-bisphosphatase (Fru-6-P,2-kinase/Fru-2,6-Pase), catalyzes synthesis and degradation of fructose 2,6-bisphosphate (Fru-2,6-P2). Previously, the rat liver Fru-2,6-Pase reaction (Fru-2,6-P2 right-arrow Fru-6-P + Pi) has been shown to proceed via a phosphoenzyme intermediate with His258 phosphorylated, and mutation of the histidine to alanine resulted in complete loss of activity (Tauler, A., Lin, K., and Pilkis, S. J. (1990) J. Biol. Chem. 265, 15617-15622). In the present study, it is shown that mutation of the corresponding histidine (His256) of the rat testis enzyme decreases activity by less than a factor of 10 with a kcat of 17% compared with the wild type enzyme. Mutation of His390 (in close proximity to His256) to Ala results in a kcat of 12.5% compared with the wild type enzyme. Attempts to detect a phosphohistidine intermediate with the H256A mutant enzyme were unsuccessful, but the phosphoenzyme is detected in the wild type, H390A, R255A, R305S, and E325A mutant enzymes. Data demonstrate that the mutation of His256 induces a change in the phosphatase hydrolytic reaction mechanism. Elimination of the nucleophilic catalyst, H256A, results in a change in mechanism. In the H256A mutant enzyme, His390 likely acts as a general base to activate water for direct hydrolysis of the 2-phosphate of Fru-2,6-P2. Mutation of Arg255 and Arg305 suggests that the arginines probably have a role in neutralizing excess charge on the 2-phosphate and polarizing the phosphoryl for subsequent transfer to either His256 or water. The role of Glu325 is less certain, but it may serve as a general acid, protonating the leaving 2-hydroxyl of Fru-2,6-P2.

    INTRODUCTION
Top
Abstract
Introduction
References

A bifunctional enzyme, Fru-6-P,2-kinase1/Fru-2,6-Pase catalyzes the synthesis and degradation of Fru-2,6-P2, the most potent activator of phosphofructokinase known (1). The rat testis enzyme is a homodimer with a subunit Mr of 55,000. The enzyme consists of two distinct catalytic domains, a Fru-6-P,2-kinase domain in the N-terminal half and a Fru-2,6-Pase domain in the C-terminal half. The crystal structure of the enzyme has recently been determined (2), and it indicates the kinase domain is structurally similar to nucleoside monophosphate kinases and to the catalytic core of the GTP-binding proteins. The crystal structure (2), as well as sequence similarities of a peptide surrounding an active site histidine, shows that the phosphatase domain belongs to the phosphoglycerate mutase family. Residue His8 in the yeast phosphoglycerate mutase has been shown to be phosphorylated during catalysis (3). Similarly, the homologous His258 of the rat liver Fru-2,6-Pase has been shown to be phosphorylated during the hydrolysis of Fru-2,6-P2 (4). Consistent with the idea that a histidine residue is central to the catalytic mechanism of Fru-2,6-Pase is the observation that mutation of His258 of the rat liver enzyme results in complete loss of the hydrolytic activity (5). Furthermore, the rate of formation of the phosphorylenzyme has been shown to be faster than the overall phosphatase activity, supporting a phosphoenzyme intermediate (6). These results suggested the following scheme (Scheme I) for the reaction pathway for the Fru-2,6-P2 hydrolysis catalyzed by Fru-2,6-Pase.


View larger version (4K):
[in this window]
[in a new window]
 
Scheme I.  

The crystallographic data of the rat testis enzyme (2), as well as a truncated form of the liver enzyme containing only the Fru-2,6-Pase domain (7), have demonstrated that in addition to His256 (corresponding to His258 of the liver isozyme), Arg255, Glu325, Arg305, and His390 (corresponding to Arg257, Glu327, Arg307, and His392, respectively, of the liver isozyme) are also located in the active site of the Fru-2,6-Pase (Fig. 1). A role for the conserved residues in catalysis has been proposed. Residue His392 (corresponding to His390 of the testis enzyme) has been suggested to act as a proton donor (5), and Arg257 and Arg307 (corresponding to Arg255 and Arg305, respectively, of the testis enzyme) are proposed to interact with the reactive C-2 phospho group of Fru-2,6-P2 and are proposed to stabilize the phospho group in the ground (Arg257) and transition states (Arg307) (8). Residue Glu327 (corresponding to Glu325 of the testis enzyme) is thought to maintain His392 in the protonated state (9).


View larger version (41K):
[in this window]
[in a new window]
 
Fig. 1.   Active site of the Fru-2,6-Pase of the bifunctional Fru-6-P,2-kinase/Fru-2,6-Pase. Pi is shown in green. Residues of import to the present study are His256, His390, Arg255, Arg305, and Glu325.

During the course of preparation of a mutant testis enzyme in which His256 (corresponding to His258 of the liver enzyme) was substituted with Ala (which, by analogy to the liver enzyme, should have eliminated Fru-2,6-Pase activity (5)), we found that the H256A mutant enzyme retained the hydrolytic activity. The objectives of this investigation are to obtain an explanation for the Fru-2,6-Pase activity of the mutant enzyme and to re-evaluate the significance of the phosphoryl-His256 as a reaction intermediate.

    EXPERIMENTAL PROCEDURES

Materials

[gamma -32P]ATP (3000 Ci/mmol) was purchased from Amersham Pharmacia Biotech. The cDNA encoding rat testis Fru-6-P,2-kinase/Fru-2,6-Pase (RT2K) was prepared as reported previously (10). The cDNA encoding the Trp-less mutant (RT2K-Wo), in which all four Trp residues of RT2K have been altered to Phe by site-directed mutagenesis was prepared as described previously (11). The pT7-7 RNA polymerase/promoter plasmid (12) was a gift from S. Tabor (Harvard Medical School). Restriction enzymes and T4 polynucleotide kinase were from New England Biolabs (Beverly, MA). The DNA ligation kit was from Takara Shuzo (Kyoto, Japan). The Muta-Gene M13 in vitro mutagenesis kit was from Bio-Rad. The Sequenase version 2.0 sequencing kit was from U.S. Biochemical Corp. [2-32P]Fru-2,6-P2 was prepared as described previously (13) using pure rat testis Fru-6-P,2-kinase/Fru-2,6-Pase. All other chemicals were of reagent grade and obtained from commercial sources.

Assay Methods for Fru-2,6-Pase

Fru-2,6-Pase activity was assayed using two methods.

Assay A (Fru-6-P Formation)-- This assay measures the formation of Fru-6-P fluorometrically coupled to NADPH formation using phosphoglucose isomerase and Glu-6-P dehydrogenase as described previously (14). The reaction mixture in a final volume of 1 ml contained 100 mM Tris HCl, pH 7.5, 0.2 mM EDTA, 100 µM NADP, 10 mM dithiothreitol, 0.4 unit of Glu 6-P dehydrogenase, 1 unit of phosphoglucose isomerase, and the indicated amounts of Fru-2,6-P2. The reaction was initiated with addition of enzyme and followed at 25 °C by measuring NADPH formation continuously at 452-nm emission and excitation at 350 nm using a Ratio-2 fluorometer (Optical Technology Devices Co.).

Assay B (Pi Formation)-- This assay measures the release of 32Pi from [2-32P]Fru-2,6-P2. The reaction mixture in a final volume of 0.1 ml contained 100 mM Tris HCl, pH 7.5, 0.2 mM EDTA, 100 µM NADP, 10 mM dithiothreitol, 0.4 unit of Glu-6-P dehydrogenase, 1 unit of phosphoglucose isomerase, and the indicated amount of [2-32P]Fru-2,6-P2. As found earlier (15), the Fru-6-P-depleting system using the coupled enzymes was necessary, since Fru-6-P is a potent inhibitor of Fru-2,6-Pase. The reaction was initiated by the addition of enzyme, and the reaction mixture was incubated at 30 °C for 10 min. The reaction was terminated by adding 10 µl of 1 N NaOH, and the solution was heated at 85 °C for 90 s. Water (1 ml) was added to the heated reaction mixture, and the sample was applied to a Dowex 1-Cl- column (0.5 × 2 cm), which had been equilibrated with 20 mM NH4OH. The column was washed with 5 ml of 20 mM NH4OH, and 32Pi was then eluted with 5 ml of 20 mM NH4OH containing 0.15 M NaCl. The eluate was diluted with 15 ml of scintillation mixture and counted in a scintillation counter.

Assay Method for Fru-6-P,2-kinase

The assay was based on the determination of the amount of Fru-2,6-P2 produced and is the same as that described previously (16) with slight modification. The reaction mixture in a final volume of 50 µl contained 100 mM Tris-HCl, pH 7.5, 0.1 mM EDTA and the indicated concentrations of Fru-6-P, MgCl2, and ATP. The reaction was initiated by the addition of enzyme, and the reaction mixture was incubated at 30 °C for 10 min. After the incubation, the reaction was terminated by adding 50 µl of 0.1 N NaOH, and the mixture was incubated at 85 °C for 90 s. Suitable aliquots were assayed for Fru-2,6-P2 as described by Uyeda et al. (17). One unit of enzyme activity is defined as the amount of enzyme that catalyzes the formation of 1 µmol of Fru-2,6-P2 per min under these conditions.

Site-directed Mutagenesis

Oligonucleotide-directed in vitro mutagenesis was performed as described by Kunkel (18), using the Muta-Gene M13 in vitro mutagenesis kit. Plasmid RT2K-Wo/pT7-7 containing the RT2K-Wo gene, cloned in a pT7-7 vector (11), was digested with XbaI and HindIII, and the isolated 1.7-kilobase pair fragment containing the RT2K-Wo gene was ligated into the XbaI-HindIII site of M13mp18 (M13/RT2K-Wo). The phage harboring M13/RT2K-Wo was transfected into Escherichia coli CJ236 (dut-,ung-). The recombinant M13/RT2K-Wo phage was purified and used to prepare uracil-containing single-stranded template. Synthetic oligonucleotide primers used for constructing various mutants are listed in Table I. The synthesized double-stranded DNA was used to transform E. coli MV1190. Mutant derivatives were checked by DNA sequencing (19). The mutant DNAs were digested with NdeI and HindIII, and the DNA fragments containing a mutated RT2K-Wo gene were subcloned into the NdeI-HindIII site of pT7-7 (RT2K-Wo/pT7-7). H256A/R305S, H256A/E325A, and H256A/H390D double mutant plasmids were constructed using the same method as that used for making single mutants with the exception of using a uracil-containing single-stranded template prepared from the H256A mutant plasmid instead of the RT2K-Wo plasmid DNA.

                              
View this table:
[in this window]
[in a new window]
 
Table I
Oligonucleotide sequences
The mutated nucleotides are underlined. These oligonucleotides are the complementary DNA of the wild type RT2K at positions 790-811 (R255A), 794-815 (H256A), 940-961 (R305S), 1000-1020 (E325A), and 1196-1216 (H390A).

The obtained expression plasmids were transformed into E. coli BL21(DE3), and recombinant mutant enzymes were purified using the same procedure as that for the wild type enzyme (14). All the enzymes were purified to apparent homogeneity as judged by SDS-polyacrylamide gel electrophoresis (SDS-polyacrylamide gel electrophoresis). Before assay, all enzymes were desalted by column centrifugation (20) in 50 mM Tris-phosphate (pH 7.5) containing 0.5 mM EDTA, 0.5 mM EGTA, 2 mM dithiothreitol, 1% polyethylene glycol (Mr = 300), and 5% glycerol.

Assay Method for Phosphoenzyme Intermediate (E-P) Formation

Previously, El-Maghrabi et al. (21) used phosphocellulose under acidic conditions to isolate the phosphoenzyme intermediate. Using the phosphocellulose method, recovery of the phosphoryl-His intermediate is less than 50%. Consequently, we have employed the following alkaline method developed by Wei and Matthew (22) using Nytran filters resulting in quantitative recovery. Nytran filters (Schleicher & Schuell) were pretreated with 10 mM sodium pyrophosphate, pH 10, for 1 h at 4 °C to reduce the background and then air-dried. At given time intervals, aliquots were removed from the reaction mixture, and the reaction was terminated by adding <FR><NU>1</NU><DE>10</DE></FR> volume of 1 N NaOH. A portion of the aliquot was transferred onto the air-dried Nytran filters, and the filters were washed with 10 mM cold sodium pyrophosphate, pH 10, for 10 min (200 ml, three times). After washing, the filters were air-dried and subjected to scintillation counting.

Mass Spectrophotometric Analysis

The enzyme was desalted by reverse phase high pressure liquid chromatography, and a 2-4-µg sample was introduced into a VG30250 quadruple mass spectrometer (Micromass Inc., Altrinckam, United Kingdom) and ionized by electrospray. The mass analyzer was scanned over the m/z range of 600-1600, and the data were transformed automatically using the manufacturer's LAB-BASE protocol.

31P NMR Experiments

All 31P NMR spectra were recorded using a Varian 500-MHz NMR spectrometer operating at 202.46 MHz for 31P. All spectra were collected using 5-mm quartz NMR tubes with a 2.5-s acquisition time, a 45° pulse, and broad band proton decoupling (2.5-3.5 watts). All 31P chemical shifts are referenced to 85% H3PO4 at 0.00 ppm. A small amount of D2O (10%, v/v) was added to the reaction mixture and served as a field/frequency lock. In all cases, about 1000-3000 scans were accumulated. An exponential line broadening of 10 Hz was applied prior to Fourier transformation. Positive chemical shifts are downfield from the reference 85% H3PO4.

Other Methods

The protein concentration was determined by the method of Bradford (23) using bovine serum albumin as a standard. SDS-polyacrylamide gel electrophoresis was carried out according to the procedure of Laemmli (24), using the Phast system (Pharmacia Biotech Inc., Uppsala, Sweden).

    RESULTS

In this study, a mutant enzyme termed Wo has been used in which all four Trp have been substituted with Phe because of its high expression compared with the wild enzyme in E. coli. The W right-arrow F mutations did not alter kinetic properties of either Fru-6-P,2-kinase or Fru-2,6-Pase activities significantly (11).

Mass Spectrometric Analysis of the Wo and H256A Mutant Enzymes-- Unexpectedly, a pure H256A enzyme was found to have Fru-2,6-Pase activity. In order to rule out the possibility that the preparation of the mutant enzyme was contaminated with a trace amount of the wild type enzyme (in this study, Wo), the masses of both the Wo and H256A mutant enzymes were determined using a mass spectrometer. Results showed that both enzymes have two components; the major protein (over 90%) in the samples is the expected enzyme, but a minor protein (less than 10%) exhibited a slightly higher mass. The minor protein in the Wo and H256A gave Mr values of 53,924 and 53,850, respectively. The minor protein in the H256A sample is not likely the Wo enzyme, since its Mr is much lower than that of the Wo enzyme (Mr = 53,872).

Steady-state Kinetic Parameters of Wo and Mutant Enzymes-- The kinetic parameters of Fru-6-P,2-kinase activity of various single and double mutants used in this study were not significantly changed (Table II). Table III provides a summary of the kinetic parameters for the Fru-2,6-Pase activity of the Wo and the mutant enzymes. The H256A mutant enzyme exhibits a kcat of 5.5 × 10-3 s-1 for Fru-2,6-Pase (17% of Wo enzyme activity), while kcat/KFru2,6-P2 decreases by almost an order of magnitude. Previously, Tauler et al. (5) reported that the H258A (corresponding to H256A of the testis enzyme) mutant of the rat liver bifunctional enzyme has no Fru-2,6-Pase activity. It is unlikely that this discrepancy is due to the difference in the isozymic forms, since the three-dimensional structures of the Fru-2,6-Pase active sites for these isozymes are identical (2, 7). The other histidine (His390) thought to act as a catalytic group, gives a similar residual kcat (12% compared with Wo) and kcat/Km (decreased by 2 orders of magnitude compared with Wo) when changed to alanine, while the H390D mutant enzyme exhibits no detectable activity at an enzyme concentration more than 2 orders of magnitude higher than that used for Wo (data not shown). Double mutants in which both histidines are changed to alanine have very little residual activity. The H256A/H390A mutant exhibits an activity 106-fold lower than Wo, while the H256A/H390D mutant exhibits no detectable activity at a 30-fold higher concentration than that used for Wo (data not shown).

                              
View this table:
[in this window]
[in a new window]
 
Table II
Kinetic parameters for the Fru-6-P,2-kinase activity of Wo and mutant enzymes
Fru-6-P,2-kinase activities were assayed as described under "Experimental Procedures" using 0.25 µg of enzyme. The range of substrate concentrations used was 20-500 µM for Fru-6-P and 40-500 µM for MgATP saturation curves, respectively. The concentrations of the fixed substrate and MgCl2 were 2 and 5 mM, respectively. The kcat values reported were calculated from the Fru-6-P saturation curves. S.E. values were ±10%.

                              
View this table:
[in this window]
[in a new window]
 
Table III
Kinetic parameters for Fru-2,6-Pase activity of Wo and the mutant enzymes
Fru-2,6-Pase activity was assayed as discussed under "Experimental Procedures" using 5 µg of enzyme. The range of substrate concentrations used was 0.02-10 µM. S.E. values were ±10%.

Three other positions (Arg255, Arg305, and Glu325) have been changed either singly or in combination with the putative active site nucleophile, His256. Of the other mutants tested, the R255A gives a kcat 10% and a kcat/KFru2,6-P2 70-fold lower than the corresponding values of Wo. The kcat values of the R305S and E325A mutant enzymes are 10- and 50-fold lower than those measured for Wo, while values for the H256A/R305S and H256A/E325A double mutants are decreased 60- and 500-fold, respectively. Interestingly, mutation of either Arg305 or His256 gives a multiplicative effect and 10-fold decrease for R305S. A 60-fold decrease for the H256A/R305S double mutant is more than additive of 6-fold decrease for H256A and a 10-fold decrease for R305S. A much more substantial effect is observed for the H256A/E325A double mutant, suggesting a more important role for Glu325 than for Arg305.

Effect of MgADP on the Fru-2,6-Pase Activity of Wo and the H256A Mutant Enzymes-- In order to rule out a possibility that the observed Fru-2,6-P2 hydrolytic activity is due to the reverse reaction of Fru-6-P,2-kinase in the absence of MgADP, the effect of MgADP on the Fru-2,6-Pase activity of Wo and H256A mutant enzyme was investigated using assay B (Pi formation). If the hydrolysis of Fru-2,6-P2 were catalyzed by the kinase, one would expect that a saturating concentration of MgADP would inhibit Pi formation by the reverse of the kinase reaction and that the inhibition should not be affected by mutation of His256. As shown in Table IV, no inhibition of the phosphatase activity was observed for either Wo or the H256A mutant enzymes using up to 1 mM MgADP. The range of MgADP concentrations used was saturating, since the KADP for the kinase in the reverse reaction has been reported as 62 µM (15), and the concentrations of Fru-2,6-P2 used were subsaturating for the reverse reaction (KFru2,6-P2 = 8 µM; Ref. 15) but near the Km for the Fru-2,6-Pase reaction. Under these conditions, the rate of the reverse reaction should be much slower than that of the phosphatase reaction. Nevertheless, Fru-2,6-P2 hydrolytic activity observed with the H256A mutant enzyme was unchanged. These results are consistent with the Fru-2,6-Pase activity of H256A arising from the phosphatase and not the kinase active site.

                              
View this table:
[in this window]
[in a new window]
 
Table IV
Effect of MgADP on Fru 2,6-Pase activity
Assay B (Pi formation) was used. Enzyme (10 ng of Wo or 100 ng of H256A) was incubated with 0.08 µM (8600 cpm/pmol for Wo) or 1 µM (950 cpm/pmol for H256A) of [2-32P]Fru-2,6-P2 in the presence of the above concentrations of ADP. The concentration of MgCl2 was 2 mM. The S.E. values were ±15%.

Effect of Fru-6-P or Pi on Fru-2,6-Pase Activity of Wo and the H256A Mutant Enzyme-- It is known that Fru-6-P and Pi are noncompetitive and competitive inhibitors, respectively, of the Fru-2,6-Pase activity (15), consistent with Scheme I. The possibility that the Fru-6-P or Pi inhibition pattern might be altered by the mutation was investigated. Inhibition of the H256A mutant enzyme by Fru-6-P gives a noncompetitive inhibition pattern (Fig. 2A (top)), qualitatively identical to that observed for Wo. However, an estimated Ki value of 86 µM is measured for H256A compared with the value of 51 nM obtained for Wo (Fig. 2B), indicating a 1000-fold decrease in affinity for Fru-6-P resulting from the mutation. As shown in Fig. 2 (bottom), Pi is a competitive inhibitor for both Wo and the H256A mutant enzyme, with the Ki values of 0.74 and 0.22 mM, respectively. Thus, very little difference in affinity for phosphate is observed as a result of the H256A mutation.


View larger version (42K):
[in this window]
[in a new window]
 
Fig. 2.   Effect of Fru-6-P (top) and Pi (bottom) on Fru-2,6-Pase activity of Wo (A) and the H256A mutant enzyme (B). Fru-2,6-Pase activity was assayed using assay B, with reaction mixtures containing varying concentrations of [2-32P]Fru-2,6-P2. (The Fru-6-P-depleting system was eliminated.) The concentrations of Fru-6-P used are 0 () for Wo and H256A; 0.04 µM (open circle ) and 0.08 µM (triangle ) for Wo and 0.08 mM (open circle ) and 0.16 mM (triangle ) for H256A. The reaction was initiated by the addition of 20 ng of Wo or 400 ng of H256A. For the effect of Pi, the assay conditions were the same with the exception that 0.4 mM (open circle ) and 0.8 mM (triangle ) Pi was added to the reaction mixture for both enzymes and that 10 ng of Wo and 150 ng of His256 were used.

Time Courses for Fru-6-P and Pi Formation for the Wo, H256A, and H390A Mutant Enzymes-- Fig. 3 shows time courses for Fru-6-P production as a function of the concentration of Wo and the H256A and H390A mutant enzymes. The formation of Fru-6-P was monitored continuously using the coupled fluorimetric assay (assay A). Production of Fru-6-P in the reactions catalyzed by both Wo and the H256A mutant enzyme is linear at all enzyme concentrations, Fig. 3A. On the other hand, the time course for Fru-6-P formation catalyzed by H390A is biphasic with a rapid initial burst of 5-min duration, followed by a slow linear rate. The initial rate of the burst phase of the H390A mutant is approximately 14 milliunits/mg, and the extent of the initial reaction is proportional to enzyme concentration.


View larger version (30K):
[in this window]
[in a new window]
 
Fig. 3.   Kinetics of Fru-6-P and Pi formation by Wo and the H256A and H390A mutant enzymes. A, Fru-6-P formation. Different amounts of enzyme were incubated with 10 mM Fru-2,6-P2, and Fru-6-P formation was monitored fluorometrically using assay A. Enzyme concentrations used were 1.5, 3.0, and 4.5 µg of Wo; 7.9, 15.7, and 23.6 µg of H256A; and 9.34, 14.0, 18.7, and 28.0 µg of H390A. The arrow indicates the time at which the enzyme was added to initiate the reaction. B, Pi and Fru-6-P formation. Enzyme (4.5 µg of Wo, 23.6 µg of H256A, or 37.4 µg of H390A) was incubated with 10 µM [32P]Fru-2,6-P2 (190 cpm/pmol). Fru-6-P formation (open circle ) was fluorometrically monitored using assay A. The formation of Pi (bullet ), was measured at given intervals using 100-µl aliquots and determining 32Pi using assay B.

The time courses for Pi formation catalyzed by the above enzymes were determined by measuring the 32Pi produced from [2-32P]Fru-2,6-P2 (assay B), while following the formation of Fru-6-P simultaneously using the fluorometric assay, so that the formation of both products could be determined in the same reaction mixture. The results (Fig. 3B) exhibit a linear production of Pi for Wo and the H256A mutant enzyme with rates of 42 and 7 milliunits/mg, respectively; identical rates were measured for Fru-6-P. The rate of Pi formation by H390A is linear with a calculated rate of 3.6 milliunits/mg, in contrast to the biphasic curve for Fru-6-P formation (Fig. 3A), and the rate of Pi formation appears to be slightly slower than that of the slow linear rate of the Fru-6-P production (3.8 milliunits/mg).

Quantitation of Enzyme-bound Inorganic Phosphate for Wo, H256A, and H390A-- Formation of phosphoenzyme (E-P) as a reaction intermediate by the H256A mutant enzyme was investigated using [2-32P]Fru-2,6-P2 at high enzyme concentration to allow detection of 32P-labeled enzyme. The amount of radiolabeled enzyme was determined using the alkaline filter procedure of Wei and Matthew (22) developed to detect the acid-labile phosphohistidine. The phosphocellulose filter method employed previously (21) using acidic conditions resulted in less than 50% recovery of E-P in our hands, while the alkaline method (22) yields an amount that is nearly stoichiometric to the Fru-6-P produced. For the H256A mutant enzyme, no detectable 32P-labeled enzyme was observed (Fig. 4B), but approximately 1 nmol of phosphorylated intermediate is observed for the 2 nmol of Wo subunits (Fig. 4A). Under the same conditions, H390A incorporated 1.5 nmol of 32P/2 nmol of subunit (Fig. 4C). Attempts to detect an E-P intermediate of H256A using other methods such as SDS-polyacrylamide gel electrophoresis and rapid gel filtration by centrifugation also failed (data not shown). The negative results indicate that either a phosphoryl-His390, if formed, is extremely labile and escaped detection or that one has direct hydrolysis of the 2-phosphate of Fru-2,6-P2 without formation of a phosphoenzyme intermediate.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 4.   Kinetics of formation of Fru-6-P, Pi, and phosphoenzyme intermediate for Wo (a), H256A (b), and H390A (c) mutant enzymes. The enzymes (110 µg) were incubated with 10 µM [2-32P]Fru2,6-P2 (150 cpm/pmol). The formation of Fru-6-P () was monitored by using assay A. At given intervals, 150-µl aliquots were removed, and the reaction was stopped by the addition of 15 µl of 1 N NaOH. A portion of the aliquot was spotted on a NYTRAN membrane (27.5 µl × 2), and phosphoenzyme formation (bullet ) was assayed as described under "Experimental Procedures." The remainder of the reaction mixture was heated for 2 min at 85 °C and used to determine Pi formation (triangle ).

The Wo enzyme shows a rapid initial formation of E-P, reaching a steady state between 50 and 150 s (Fig. 4A), while the formation of Fru-6-P and Pi proceeds linearly (Fig. 3). Stoichiometry of phosphate to the three enzymes at steady state is given in Table V. The maximum steady-state level of phosphoenzyme intermediate detected is 0.47 mol/mol subunit for Wo, while a somewhat higher steady-state concentration of 0.79 mol/mol of subunit is observed for the H390A mutant enzyme (Fig. 4C and Table V). As discussed above, Pi production lags behind Fru-6-P formation in the first 4 min of the time course for the H390A mutant enzyme, and this can be attributed to the formation of phosphoryl-His256 as an intermediate.

                              
View this table:
[in this window]
[in a new window]
 
Table V
Stoichiometry of steady-state phosphoenzyme intermediate
Phosphoryl-enzyme intermediate was calculated from data in Figs. 5 and 6, where the ratio reached a plateau region.

Quantitation of Enzyme-bound Inorganic Phosphate for R305S, R255A, and E325A-- As shown in Fig. 5, the time courses for Fru-6-P and E-P formation by the R305S mutant enzyme are identical, and no Pi formation is observed until after 20 min. Results suggest that the R305S-catalyzed reaction is practically single turnover and that the R305S mutation results in stabilization of the phosphoryl-His256 against hydrolysis. In support of this interpretation, the stoichiometry of phosphoenzyme intermediate as a function of R305S concentration is 0.98 (Fig. 5, inset). In the case of the R305S/H256A double mutant, no stable phosphoryl-His is detected, indicating that if formed, hydrolysis of phosphoryl-His390, does not require Arg305. A very slow hydrolysis of the E-P intermediate has been reported for the homologous Arg309 mutant of liver isozyme (8), but a comparison is difficult since Fru-6-P formation was not concomitantly measured. Finally, the steady-state concentrations of E-P for the R255A and E325A mutant enzymes is about 0.4 mol/mol of subunit, comparable with that obtained for Wo (Table V).


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 5.   Kinetics of formation of Fru-6-P, Pi, and phosphoenzyme intermediate for the R305S mutant enzyme. The concentration of R305S was 110 µg, incubated with 10 µM [2-32P]Fru2,6-P2 (150 cpm/pmol). The formation of Fru-6-P (), phosphoenzyme (bullet ), and Pi (triangle ) was assayed as described in the legend to Fig. 3, and the inset shows the stoichiometry of the Fru-6-P formation as a function of the amount of R305S.

The 10-fold difference in the kcat of the H256A mutant and the H256A/R305S double mutant enzymes (Table IV) lies in the respective roles of the two catalytic residues. Based on the studies presented, Arg305 is thought to be responsible for neutralizing charge on the 2-phosphate of Fru-2,6-P2 as it is transferred to His256, the nucleophilic catalyst, i.e. transition state stabilization. A minimal kinetic scheme for the kcat of the phosphatase is given below, where k3 represents phosphoryl transfer from Fru-2,6-P2 to His256, k4 represents the reverse of the reaction, and k5 is a net rate constant that represents the hydrolysis of the phosphohistidyl intermediate and release of products. Residue Arg305 is thought to have a role in stabilizing the transition states for both phosphoryl transfer reactions depicted above. Mutation of His256 gives only a 6-fold decrease in kcat because His390 compensates for its absence as the mechanism changes from nucleophilic/general acid catalysis to general base/general acid catalysis. Mutation of Arg305 results in a loss of transition state stabilization in both phosphoryl transfer steps, resulting in a 10-fold decrease in kcat. Mutation of both residues results in a loss of both functions, nucleophilic catalysis of His256 and the transition state stabilization of Arg305, giving an overall decrease in kcat that is the product of the two effects, 60-fold.
E:<UP>Fru-2,6-P</UP><SUB>2</SUB> <LIM><OP><ARROW>⇄</ARROW></OP><LL>k<SUB>4</SUB></LL><UL>k<SUB>3</SUB></UL></LIM> E-<UP>P:Fru-6-P</UP> <LIM><OP><ARROW>→</ARROW></OP><UL>k<SUB>5</SUB></UL></LIM>
<UP><SC>Reaction</SC> 1</UP>

NMR Studies-- Fig. 6 shows 31P NMR spectra taken after a 1-2-h incubation of Fru-2,6-Pase with Fru-2,6-P2 at pH 7.4 and 25 °C. Experiments are the same as those carried out previously (25) for detection of the phosphohistidine intermediate at -7.1 ppm at pH 8 for the wild type rat liver enzyme. Spectrum C shows the reaction mixture for Wo. The extent of the reaction was in the order of 3%. The pH of the solution for NMR experiments is best determined using the pH dependence of the 31Pi chemical shift (26). Inorganic phosphate has a pK of about 6.8, and the chemical shifts for the monoanionic and dianioic forms of Pi are 0.76 and 3.23, respectively. At pH 8, a value very close to 3.2 is expected and is observed in Fig. 6. Spectra reported by Okar et al. (25), however, give a chemical shift of 1.88 for 31Pi, consistent with a pH of about 6.5, not 8 as reported. The lower pH must be taken into account when comparing data for the rat liver and rat testis enzymes.


View larger version (14K):
[in this window]
[in a new window]
 
Fig. 6.   31P NMR experiments carried out under turnover conditions for the phosphatase activity of the bifunctional fructose-6-phosphate 2-kinase/fructose-2,6-bisphosphatase. Spectra are obtained beginning with 7.2 mM Fru-2,6-P2 at pH 8 for the H256A (A; 500 µM), H390A (B; 390 µM), and Wo (C; 190 µM) enzymes. Assignments of resonances are based partially on control spectra and partly on previous work (25). Assignments are as follows. Peak 1, Fru-6-P bound to the phosphatase; peak 2, contaminant present in the Fru-2,6-P2 (probably Fru-1,6-P2); peak 3, 6-phosphate of Fru-2,6-P2 and Fru-6-P; peak 4, Pi; peak 5, 2-phosphate of Fru-2,6-P2; inset, phosphohistidine. Spectra were collected for 1-2 h (1000-3000 scans) after the addition of enzyme to start the reaction, using a Varian 500-MHz NMR operating at 202.46 for 31P. An exponential line broadening of 10 Hz was applied prior to Fourier transformation. Chemical shifts are relative to 85% H3PO4.


    DISCUSSION

A reaction mechanism has been proposed (4-6) for the rat liver fructose 2,6-bisphosphatase reaction involving a phosphoenzyme intermediate (Scheme I), and His258 (corresponds to His256 of the testis enzyme) acts as a phosphoryl acceptor. The evidence in support of their reaction mechanism is as follows: (a) when the enzyme was incubated with [2-32P]Fru-2,6-P2, the enzyme became labeled with 32P on His258; (b) the amino acid sequence around His258 of the bifunctional enzyme is homologous to that of phosphoryl-glycerate mutase, and formation of a similar phosphoryl-His as a reaction intermediate by the mutase has been demonstrated (3); (c) mutation of His258 to Ala causes complete loss of the Fru-2,6-Pase activity, and no phosphoryl-enzyme intermediate by this mutant enzyme was detected (5); and (d) substitution of His392 (corresponds to His390 of the testis enzyme) to Ala retains 5% of the wild type activity, and the level of the E-P formation was almost the same as that of the wild type enzyme (5).

Characterization of Rat Testis H256A and H390A Mutant Enzymes-- It is interesting to note that whether His256, the nucleophilic catalyst, or His390, thought to act as a general base, are changed to Ala, the activity of the phosphatase is decreased by only 6-8-fold. That the groups are catalytically important is substantiated by changing both residues to Ala (H256A/H390A), which eliminates Fru-2,6-Pase activity. All of the data obtained in these studies are consistent with the assignment of His256 as a nucleophilic catalyst. The H256A mutant enzyme exhibits no phosphate covalently bound to the enzyme, while Wo and H390A have 0.5 and 0.8 phosphates per subunit, respectively. Data obtained from 31P NMR experiments corroborate the presence of the phosphoryl histidine, based on high field resonances at -6.5 ppm for Wo and H390A but not for H256A. It is unusual to have activity remaining upon elimination of an enzymic nucleophilic catalyst, let alone the approximately 20% activity observed for H256A. Data suggest that an alternative route is available to the phosphatase, utilizing His390 either as an alternative nucleophile or as a base to activate water for direct attack on the 2-phosphate of Fru-2,6-P2.

It is evident from the kinetic studies of the mutants, compared with Wo, that the activity remaining in the case of the H256A and H390A mutant enzymes reflects the phosphatase and not the kinase active site. Affinity of the phosphatase for Fru-6-P is decreased by 1000-fold compared with Wo, while the affinity for Pi is virtually unaffected. In addition, the presence of MgADP has no effect on the phosphatase activity, as would be expected if it were due to a reversal of the kinase reaction.

The time courses for Wo and H256A are linear with respect to the production of phosphate and Fru-6-P, with equal initial rates obtained monitoring the appearance of either. Interestingly, the H390A mutant enzyme gives a burst of Fru-6-P formation, followed by a slow steady state rate, which is equal to the rate of Pi production. Data are consistent with a rapid formation of phosphoryl-His256 in the case of the H390A mutant enzyme, and the lack of (or instability of) a phosphoenzyme intermediate in the case of H256A. The lack of a burst rate in the case of Wo suggests a more rapid turnover of the phosphoenzyme intermediate, consistent with the 6-8-fold faster overall rate of Wo compared with the mutant enzymes.

The results presented herein differ significantly from those presented by Tauler et al. (5), who reported that mutation of His258 of the rat liver enzyme to Ala resulted in complete loss of the Fru-2,6-Pase activity and that no phosphoryl-enzyme intermediate was detected in this mutant enzyme. It is highly unlikely that the discrepancy between the previously reported work (5) and that reported in this manuscript is due to isozymic differences between the liver and testis enzymes, because the catalytic domains, especially the essential amino acid residues at the active sites of all the bifunctional enzymes, are identical (9). More importantly, the crystal structures of the native testis enzyme (2) and a truncated form of the liver enzyme (missing the kinase domain and 30 amino acids at the C terminus) (7) reveal that the arrangement of the essential amino acids at the active site of the phosphatase, including His256 (His258 in the liver enzyme) and His390 (His392 in the liver enzyme), is identical.

An important question is whether a phosphoryl-His intermediate could form with either His256 or His390 as the active site nucleophile. It is clear that H390A uses a reaction pathway involving phosphoryl-His256, as does the wild type enzyme, and evidence in support of this pathway is well established (Ref. 4 and the present study). On the other hand, evidence in support of the involvement of a phosphoryl-His390 intermediate has not been obtained. Several unsuccessful attempts were made to detect phosphoryl-His390 in the H256A mutant protein, including rapid filtration under alkaline conditions (22), and NMR studies under the conditions in which the phosphoryl-His256 resonance was observed in Wo and the H390A mutant enzyme. Results do not rule out formation of phosphoryl-His390, since it is possible that the intermediate is extremely labile and escapes detection with the methods thus far employed. However, it is also possible, and more likely, that the mechanism of the reaction catalyzed by H256A differs from that catalyzed by Wo and H390A and that the H256A mutant enzyme hydrolyzes Fru-2,6-P2 directly without the formation of a phosphoenzyme intermediate. In the alternative mechanism, His390 would act in its normal proposed capacity as a general base, required to activate H2O for nucleophilic attack on the 2-phosphate of Fru-2,6-P2. The initial rates of formation of Fru-6-P, phosphoenzyme, and Pi by H256A and H390A are different, and the differences are consistent with the idea of two different pathways employed by these mutant enzymes. The only results that are potentially inconsistent with direct hydrolysis of Fru-2,6-P2 are the identical noncompetitive (Fru-6-P) and competitive (Pi) product inhibition patterns exhibited by H256A and H390A, suggesting that both enzymes follow the same kinetic mechanism. However, there is no need to invoke a phosphoenzyme intermediate based on the product inhibition patterns, which simply suggest an ordered kinetic mechanism with Fru-6-P released prior to Pi. Thus, it is proposed that Wo and the H390A mutant enzyme exhibit a nucleophilic catalytic mechanism with His256 as the active site nucleophile, while the H256A mutant enzyme uses His390 as a general base catalyst to activate water in an SN2 reaction (see below).

In support of the above, the crystal structure of the testis enzyme (2) shows two phosphates bound at the active site of the phosphatase, corresponding to the 6- and 2-phosphates of Fru-2,6-P2. One of the P-O bonds of the phosphate corresponding to the 2-phosphate is directly in line with Nepsilon of His256. On the other hand, Nepsilon of His390 is situated 4 Å away from the phosphate. That the enzyme utilizes the direct hydrolytic route during the normal operation of Wo is unlikely but has not been ruled out. The stoichiometry of phosphate/enzyme at steady state is always about 0.5 mol/mol of subunit, suggesting that at least 50% of time the nucleophilic reaction pathway is used, but this cannot be used as evidence that the direct hydrolysis pathway exists, because of the kinetic lability of the phosphoenzyme intermediate. Clearly, additional experiments are required to sort out these interesting possibilities.

Characterization of Other Mutant Enzymes-- Previously, the kcat and KFru2,6-P2 values of the R257A mutant of the liver enzyme (corresponding to Arg255 of the testis enzyme) were reported to be 10- and 12,500-fold, respectively, higher than those of wild type enzyme (8). They also showed that R307A (corresponding to Arg305 of the testis enzyme) decreased in Vmax by a factor of 700 (8). In addition, a human placental bifunctional enzyme was found in which the amino acid residue corresponding to Arg305 of the testis enzyme was naturally substituted with Ser, and this isozyme showed very low Fru-2,6-Pase activity (Vmax = 0.1 milliunit/mg) (13). The active site is shown in Fig. 1, where Arg255 and Arg305 are within hydrogen bonding distance to the presumed 2-phosphate and may participate by neutralizing negative charge on the phosphate and polarizing the phosphoryl to facilitate nucleophilic attack by His256, and Glu325 is near the 2-phosphate. The two arginine residues and Glu325 were thus changed in the rat testis enzyme to determine whether they might, as suspected, have catalytic function.

In all three residues of the mutant enzymes, R255A, R305S, and E325A, a phosphoenzyme intermediate was observed using the alkaline isolation method (22). Indeed, the phosphoenzyme intermediate appears to be quite stable (for about 20 min) in the R305S mutant and is present at a stoichiometry equal to that observed for the H390A mutant discussed above. Results suggest that the R305S-catalyzed reaction is practically single turnover and that the R305S mutation results in stabilization of the phosphoryl-His256 against hydrolysis. In the case of the R305S/H256A double mutant, only a slight further decrease in activity is observed. The kcat values of the R305S and H256A/R305S mutant enzymes are 10- and 60-fold lower than those measured for Wo, so that a multiplicative effect is observed. Data are consistent with the role proposed above for the two arginine residues in neutralizing negative charge on phosphate, allowing nucleophilic attack by His256. The lack of polarization of the phosphohemiketal of Fru-2,6-P2 and the phosphorylhistidine would result in a decreased rate of hydrolysis. The R255A mutation also gives a 1-order of magnitude decrease in kcat and a 70-fold value of kcat/KFru2,6-P2 compared with Wo, consistent with the above proposed role of the arginines in catalysis.

Results suggest a more important role for a base Glu325. A decrease in kcat of 50-fold is obtained for the E325A mutant enzymes, and a further 10-fold decrease is obtained as His256 is changed to alanine. Thus, data are consistent with the role of Glu325 as a general base in deprotonating His390 and then as a general acid protonating the leaving 2-hydroxyl. None of the putative catalytic residues give substantial decreases in activity upon elimination of their side chain functional group, with the largest change observed for the R305S mutant. Even double mutants give less than a 3-order of magnitude decrease in activity. The number of ionizable functional groups that impinge on Fru-2,6-P2 are thus important, with redundancy built in to pairs of histidines and pairs of arginines.

Proposed Chemical Mechanism-- A proposed chemical mechanism is given in Fig. 7. Fru-2,6-P2 is bound at the active site of the phosphatase (interactions other than those at the 2-phosphate are not shown) with hydrogen bonds from Arg305 and Arg255 to phosphate oxygens. The nucleophilic histidine (His256) attacks the 2-phosphate as a base protonates the leaving 2-hydroxyl to generate a phosphohistidine intermediate. The other histidine (His390) then acts as a general base and activates water for the hydrolysis reaction.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 7.   Proposed chemical mechanism for wild type Fru-2,6-Pase.

The mechanism is proposed to differ, dependent on the mutant enzyme studied. In the case of H390A (Fig. 8), the nucleophilic catalytic portion of the reaction probably proceeds as for the wild type enzyme, but another enzyme residue (perhaps Glu325) plays the role of general base catalyst, activating water for the hydrolysis reaction. For H256A, however, direct attack by water on the 2-phosphate probably occurs (Fig. 9). In the case of direct attack, His390 acts as a general base, its normal capacity, activating water for nucleophilic attack on phosphate.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 8.   Proposed chemical mechanism for the H390A mutant enzyme.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 9.   Proposed chemical mechanism for the H256A mutant enzyme.


    ACKNOWLEDGEMENTS

We thank the following for excellent technical assistance: Cu Nguyen and Yang Li for preparation of various enzymes and Eric Enwall for 31P experiments. We also thank Dr. Clive Slaughter (Howard Hughes Medical Institute/University of Texas Southwestern, Dallas) for performing mass spectrometric analysis.

    FOOTNOTES

* This work was supported by a grant from the Department of Veterans Affairs (to K. U.) and National Institutes of Health Grants DK16194 (to K. U.) and GM36799 (to P. F. C.), and a grant from the Robert A. Welch Foundation (to C. A. H.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Dallas VA Medical Center, 4500 S. Lancaster Rd., Dallas, TX 75216. Tel.: 214-857-0318; Fax: 214-372-9534; E-mail: kuyeda6400{at}aol.com.

The abbreviations used are: Fru-6-P, 2-kinase, fructose 6-phosphate,2-kinase; Fru-2, 6-Pase, fructose-2,6-bisphosphatase; Glu-6-P, beta -D-glucose 6-phosphate; Fru-6-P, beta -D-fructose 6-phosphate; Fru-2, 6-P2, fructose 2,6-bisphosphate; RT2K, rat testis isozyme of fructose-6-phosphate,2-kinase/fructose-2,6-bisphosphatase; RT2K-Wo, RT2K with all four tryptophans mutated to phenylalanine; H256A, a mutant form of RT2K-Wo with an additional histidine 256 to alanine mutation.
    REFERENCES
Top
Abstract
Introduction
References

  1. Uyeda, K. (1991) in Enzyme Catalysis (Kurby, S. A., ed), Vol. II, pp. 445-456, CRC Press, Inc., Boston
  2. Hasemann, C. A., Istavan, E. S., Uyeda, K., and Deisenhofer, J. (1996) Structure 4, 1017-1029[Medline] [Order article via Infotrieve]
  3. Han, C.-H., and Rose, Z. B. (1979) J. Biol. Chem. 254, 8836-8840[Abstract]
  4. Pilkis, S. J., Lively, M. O., and El-Maghrabi, M. R. (1987) J. Biol. Chem. 262, 12672-12675[Abstract/Free Full Text]
  5. Tauler, A., Lin, K., and Pilkis, S. J. (1990) J. Biol. Chem. 265, 15617-15622[Abstract/Free Full Text]
  6. Stewart, H. B., El-Magfrabi, M. R., and Pilkis, S. J. (1985) J. Biol. Chem. 260, 12935-12941[Abstract/Free Full Text]
  7. Lee, Y.-H., Ogata, C., Pflugrath, J. W., Levitt, D. G., Sarma, R., Banaszak, L. J., and Pilkis, S. J. (1996) Biochemistry 35, 6010-6019[CrossRef][Medline] [Order article via Infotrieve]
  8. Lin, K., Li, L., Correia, J. J., and Pilkis, S. J. (1992) J. Biol. Chem. 267, 19163-19171[Abstract/Free Full Text]
  9. Lin, K., Li, L., Correia, J. J., and Pilkis, S. J. (1992) J. Biol. Chem. 267, 6556-6562[Abstract/Free Full Text]
  10. Sakata, J., Abe, Y., and Uyeda, K. (1991) J. Biol. Chem. 266, 15764-15770[Abstract/Free Full Text]
  11. Watanabe, F., Jameson, D. M., and Uyeda, K. (1996) Protein Sci. 5, 904-913[Abstract/Free Full Text]
  12. Tabor, S., and Richardson, C. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 1074-1078[Abstract]
  13. Sakakibara, R., Kato, M., Okamura, N., Nakagawa, T., Koyama, Y., Tominaga, N., Shimojo, M., and Fukasawa, M. (1997) J. Biochem. 122, 122-128[Abstract]
  14. Tominaga, N., Minami, Y., Sakakibara, R., and Uyeda, K. (1993) J. Biol. Chem. 268, 15951-15957[Abstract/Free Full Text]
  15. Kitajima, S., Sakakibara, R., and Uyeda, K. (1984) J. Biol. Chem. 259, 6896-6903[Abstract/Free Full Text]
  16. Furuya, E., and Uyeda, K. (1981) J. Biol. Chem. 256, 7109-7112[Abstract/Free Full Text]
  17. Uyeda, K., Furuya, E., and Luby, L. J. (1981) J. Biol. Chem. 256, 8394-8399[Abstract/Free Full Text]
  18. Kunkel, T. A. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 488-492[Abstract]
  19. Sanger, F., Nicklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5463-5467[Abstract]
  20. Penefsky, H. S. (1977) J. Biol. Chem. 252, 2891-2899[Abstract]
  21. El-Maghrabi, M. R., Correia, J. J., Heil, P. J., Pate, T. M., Cobb, C. E., and Pilkis, S. J. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 5005-5009[Abstract]
  22. Wei, Y.-F., and Matthews, R. H. (1990) Anal. Biochem. 190, 188-192[Medline] [Order article via Infotrieve]
  23. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
  24. Laemmli, U. K. (1970) Nature 227, 680-685[Medline] [Order article via Infotrieve]
  25. Okar, D. A., Kakalis, L. T., Narula, S. S., Armitage, I. M., and Pilkis, S. J. (1995) Biochem. J. 308, 189-195[Medline] [Order article via Infotrieve]
  26. Sclinackerg, K. D., and Waldwann, G. (1991) in Enzymes Dependent Pyridoxal-P and Other Carboxyl Compounds as Covactors: Proceedings of the Eighth International Symposium (Fukui, T., Kagamiyama, H., Soda, K., and Wada, H., eds), pp. 149-152, Pergamon Press, Tokyo
  27. Yuen, M.-H., Mizuguchi, H., Lee, Y.-H., Cook, P. F., Uyeda, K., and Hasemann, C. A. (1998) J. Biol. Chem. 274, 2176-2184[Abstract/Free Full Text]


Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.