Alteration of the specificity of the cofactor-binding pocket of Corynebacterium 2,5-diketo-D-gluconic acid reductase A

Scott Banta1,2, Barbara A. Swanson3,4, Shan Wu3, Alisha Jarnagin3 and Stephen Anderson2,,5,6

1 Departments of Chemical and Biochemical Engineering 5 Molecular Biology and Biochemistry, Rutgers, The State University of New Jersey, 2 Center for Advanced Biotechnology and Medicine, 679 Hoes Lane, Piscataway, NJ 08854 3 Genencor International, 925 Page Mill Road, Palo Alto, CA 94304, USA


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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The NADPH-dependent 2,5-diketo-D-gluconic acid (2,5-DKG) reductase enzyme is a required component in some novel biosynthetic vitamin C production processes. This enzyme catalyzes the conversion of 2,5-DKG to 2-keto-L-gulonic acid, which is an immediate precursor to L-ascorbic acid. Forty unique site-directed mutations were made at five residues in the cofactor-binding pocket of 2,5-DKG reductase A in an attempt to improve its ability to use NADH as a cofactor. NADH is more stable, less expensive and more prevalent in the cell than is NADPH. To the best of our knowledge, this is the first focused attempt to alter the cofactor specificity of a member of the aldo–keto reductase superfamily by engineering improved activity with NADH into the enzyme. Activity of the mutants with NADH or NADPH was assayed using activity-stained native polyacrylamide gels. Eight of the mutants at three different sites were identified as having improved activity with NADH. These mutants were purified and subjected to a kinetic characterization with NADH as a cofactor. The best mutant obtained, R238H, produced an almost 7-fold improvement in catalysis with NADH compared with the wild-type enzyme. Surprisingly, most of this catalytic improvement appeared to be due to an improvement in the apparent kcat for the reaction rather than a large improvement in the affinity of the enzyme for NADH.

Keywords: aldo/keto reductase/cofactor specificity/ 2,5-diketo-D-gluconic acid reductase/2-keto-L-gulonic acid/site-directed mutagenesis


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
L-Ascorbic acid, or vitamin C, is an essential part of the human diet and a valuable food preservative. The health benefits of supplementing the diet with antioxidants such as vitamin C were recently underscored by an increase in the daily recommended allowance (RDA) of vitamin C from 60 to 75 mg per day for women and 90 mg per day for men. Currently, vitamin C is predominantly manufactured by the modified Reichstein–Grussner synthesis (Reichstein and Grussner, 1934Go), which involves five chemical steps and a single fermentation step.

This process could be dramatically simplified if more of the chemical steps occurred within the cell during the fermentation step. This goal was achieved when Anderson et al. cloned an NADPH-dependent 2,5-diketo-D-gluconic acid reductase (2,5-DKG reductase A, EC 1.1.1.–, AKR5C) from Corynebacterium sp. and expressed it in Erwinia herbicola (Anderson et al., 1985Go). E.herbicola cells are able to catalyze the multi-step conversion of D-glucose to 2,5-diketo-D-gluconic acid (2,5-DKG) in the periplasm via the action of three membrane-bound dehydrogenases. 2,5-DKG can then be transported into the cytoplasm where it encounters the recombinantly expressed Corynebacterium 2,5-DKG reductase, which is capable of converting 2,5-DKG into 2-keto-L-gluonic acid (2-KLG) (Lazarus et al., 1989Go, 1990Go). 2-KLG can be collected from the fermentation broth and chemically lactonized at low pH directly into L-ascorbic acid. Thus, a metabolically engineered Erwinia can produce the immediate precursor to vitamin C from glucose in a single fermentation step.

Several improvements were subsequently added to this process. A second 2,5-diketo-D-gluconic acid reductase with different kinetic properties (2,5-DKG reductase B, AKR5D), also from Corynebacterium, was identified (Sonoyama and Kobayashi, 1987Go). Mutant strains of the host cell were also isolated that had altered metabolism of carbon precursors or by-products, which resulted in improved yields of 2-KLG (Grindley et al., 1988Go). Additionally, a recent protein engineering effort resulted in catalytic improvements in the original 2,5-DKG reductase A enzyme (Powers, 1996Go). In this work, substitutions based in part on residues found in the predicted substrate-binding pocket of the B form of the enzyme were engineered into the A form and residues in the C-terminal tail of the enzyme were also mutated. An F22Y/A272G double mutant of 2,5-DKG reductase A, which exhibited significantly improved activity with 2,5-DKG compared with the wild-type enzyme, resulted (Powers, 1996Go).

An in vitro 2-KLG production system employing Corynebacterium 2,5-DKG reductase has also been described (Boston and Swanson, 2000Go). Erwinia cells, genetically lesioned to knock out native glucose dehydrogenase activity, were grown to stationary phase and permeabilized and then combined with an NADP+-dependent glucose dehydrogenase and the Corynebacterium NADPH-dependent 2,5-DKG reductase in a bioreactor. NADP+, glucose and 2,5-DKG were then added and the two enzymes were shown to be capable of recycling the cofactor as 2-KLG was produced from glucose.

NADH differs structurally from NADPH solely in the absence of a phosphate group attached to the 2' site of its adenine moiety. The concentration of NADH in a cell is generally an order of magnitude higher than that of NADPH. Lundquist and Olivera reported values of 1.3 µM for NAD(H) and 0.39 µM for NADP(H) for exponentially growing Escherichia coli (Lundquist and Olivera, 1971Go). In addition, NADH is roughly an order of magnitude less expensive than is NADPH and it is more stable. Therefore, a 2,5-DKG reductase that was active with NADH would be a valuable catalyst for use in industrial vitamin C production, either in vivo or in vitro.

The re-engineering of the cofactor specificity of an enzyme has already been accomplished in some protein families. For example, the residues governing cofactor specificity in the cofactor-binding pocket known as the `Rossman fold' have been thoroughly studied (Scrutton et al., 1990Go; Hedstrom, 1994Go). The 2,5-DKG reductase from Corynebacterium belongs to the aldo–keto reductase (AKR) superfamily, which is characterized by a unique cofactor-binding pocket that places the bound cofactor in an extended conformation (Jez et al., 1997Go). This cofactor-binding pocket has been studied in several members of the AKR superfamily. Site-directed mutations have been made in the pocket in order to determine the effects on catalysis in the presence of the native cofactor (Bohren et al, 1991Go.; Yamaoka et al., 1992Go; Kubiseski and Flynn, 1995Go; Ratnam et al., 1999Go).

The 2,5-DKG reductase enzyme is a 34 kDa monomer and, like other members of the AKR superfamily, it appears to follow the ordered bi bi kinetic mechanism, with cofactor binding first and leaving last. The native enzyme has a 170-fold preference for NADPH over NADH as a cofactor (Miller et al., 1987Go). We undertook a concerted protein engineering effort aimed at improving the 2,5-DKG reductase A's ability to utilize NADH, the first such member of the AKR superfamily to be studied with this goal in mind.

The crystal structure of the 2,5-DKG reductase A complexed with NADPH has recently been solved at 2.1 Å (Khurana et al., 1998Go). From this structure, it is possible to delineate every residue that has an interaction with the bound NADPH molecule. To explore the feasibility of increasing the ability of the enzyme to use NADH as a cofactor, we decided to mutate all of the residues that interact with the 2'-phosphate group of the bound NADPH. There appear to be five such residues: Lys232, Ser233, Val234, Arg235 and Arg238 (Figure 1Go). These residues were identified using the Ligand–Protein Contacts program and the characteristics of the interactions were determined using the Contacts of Structural Units program (Sobolev et al., 1999Go). Both programs use the atomic coordinates deposited at the Protein Data Bank (1A80) (Khurana et al., 1998Go).



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Fig. 1. Residues that make a direct interaction with the 2'-phosphate of the bound NADPH molecule. The coordinates for the 2,5-diketo-D-gluconic acid reductase A have been deposited at the Protein Data Bank as 1A80 (Khurana et al., 1998Go).

 
The effects of mutating some of these cofactor-binding pocket residues have previously been studied in other members of the AKR superfamily. Table IGo shows sequence alignments for several members of the superfamily (Jez et al., 1997Go). The lysine corresponding to position 232 occurs in almost every member. It had previously been identified as important in the catalytic mechanism of both aldose reductase and aldehyde reductase. Mutagenesis at this site was shown to affect all of the measured steady-state kinetic parameters for these enzymes (Bohren et al., 1991Go; Yamaoka et al., 1992Go).


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Table I. Sequence alignments of the 2'-phosphate binding residues for several AKR superfamily members using 2,5-DKG reductase A numbering
 
The arginine corresponding to position 238 has also been studied in this family. Kubiseski and Flynn demonstrated that mutating the arginine to methionine in human aldose reductase increased the KM for NADPH, but left the other measured kinetic parameters unchanged (Kubiseski and Flynn, 1995Go).

This idea was further extended by Ratnam et al. with recent work done on rat 3{alpha}-hydroxysteroid dehydrogenase (3{alpha}-HSD), which can naturally use both NADH and NADPH as cofactors, with a preference for NADPH (Ratnam et al., 1999Go). Pre-steady-state kinetic transients were observed when the wild-type enzyme bound NADPH, but not when the wild-type enzyme bound NADH or when an R276M mutant (analogous to R238M in 2,5-DKG reductase A) bound NADPH. The effect on the steady-state kinetic parameters for this mutant was minor except for a dramatic increase in the KM for both NADPH and NADP+. The authors described the interaction between Arg276 and NADPH as an `anchoring' that was required to orient the cofactor in the proper position for optimal catalysis. They suggested that the lack of an anchor in the wild-type enzyme for the NADH molecule could explain the higher KM and Kd for NADH and thus the diminished activity of the native enzyme with this cofactor.

Cofactor dissociation constants were measured for the 3{alpha}-HSD R276M mutant and it was shown that this mutation resulted in an increased affinity for NADH. However, based on apparent kcat/KM values, this mutant enzyme nonetheless exhibited markedly reduced activity with NADH (Ratnam et al., 1999Go).

In the present work, 40 single mutations were made at five cofactor-binding pocket residues that interact with the 2'-phosphate of NADPH in the 2,5-DKG reductase A enzyme in an attempt to enhance activity with NADH, regardless of the impact on NADPH activity. We pursued a systematic `scanning mutagenesis' strategy (Cunningham and Wells, 1989Go) in order to determine exactly which residues in the putative cofactor specificity loop (Figure 1Go) were important. Also, in order not to miss anything due to compensating energetic effects (Bigler et al., 1993Go), we scanned with a range of different substitutions at each position, representing differences in charge, hydrophobicity, size, etc., of the respective mutant vs wild-type residues. The effects of these mutations with both cofactors were assessed using activity-stained native polyacrylamide gels. Mutants that demonstrated an apparent improvement in activity with NADH were then purified to homogeneity and subjected to further kinetic analyses. This allowed for a detailed comparison of the effects of the different mutations on the activity of the enzyme with NADH. This information will be invaluable as a first step in our studies aimed at engineering a mutant 2,5-DKG reductase enzyme with the best possible NADH-mediated catalytic behavior.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Materials

Oligonucleotide primers were from Life Technologies or from Integrated DNA Technologies. XL 1 Blue and JM109 E.coli cells and Pantoea citrea (Acetobacter cerinus, #39140) cells were from the American Type Culture Collection. NADPH, NADH, Bis-Tris, sodium acetate, sodium azide, sodium chloride, ampicillin, IPTG and DEAE-cellulose were from Sigma. DEAE Ceramic Hyper D F ion-exchange resin was from BioSepra. Bug Buster cell lysis detergent was from Novagen. Complete Protease Inhibitor tablets were from Roche. Restriction enzymes and T4 DNA ligase were from New England Biolabs. TaqPlus Precision polyerase was from Stratagene. Precast polyacrylamide gels were from Novex. PM-10 ultrafiltration membranes were from Millipore.

The ptrp1-35.xb vector

The starting point for this work was the ptrp1-35 vector, which contains the 2,5-DKG reductase gene (Anderson et al., 1985Go) and a cassette mutagenesis strategy was employed. Cassette mutagenesis requires unique endonuclease restriction sites that immediately flank the region that is to be mutated. Three rounds of PCR mutagenesis were required to produce the vector ptrp1-35.xb by the method of Ho et al. (Ho et al., 1989Go). The first set of PCR reactions introduced a unique Bsu36I site upstream of the region of interest with three silent mutations. The oligonucleotides used for this were as follows, with the silent mutations in italics: A-5'GAC GTG AAG ATC GAA TCG TGG GG3', B-5'G GTG CCA CCT GAG ACG GCC T3', C-5'AG GCC GTC CTC AGG TGG CAC C3' and D-5'CG ACC CGA ACC GTC GCC CGG3'. The second set of PCR reactions introduced a XhoI site downstream of the region of interest with a single silent mutation. The oligonucleotides used for this were A-5'GTG AAG ATC GAA TCG TGG3', B-5'G GTT CTC CTC GAG GCG3', C-5'CGC CTC GAG GAG AAC C3' and D-5'CGA ACC GTC GCC CGG3'. This new XhoI site was not unique; therefore, a natural XhoI site was removed during a third set of PCR reactions by making a single silent mutation with the following oligonucleotides: A-5'A AGT TCT CGT AAA AAG GGT A3', B-5'AC GAT GCG TTC GAG GTG3', C-5'CAC CTC GAA CGC ATC GT3' and D-5'TC GTA CTT GCC CTG ACC3'. The accuracy of all mutagenesis procedures was verified by DNA sequencing of the resultant mutant plasmids.

The pATP003.xb vector

The new 2,5-DKG reductase A gene was digested out of ptrp1-35.xb and then ligated into the pATP003 vector, using unique EcoRI and HindIII sites, creating the final expression vector pATP003.xb. The newly engineered cassette mutagenesis restriction sites were still unique and this new vector contains a GroES fragment fused to the N-terminus of the protein. The new vector allows for high levels of controlled expression in E.coli (onelly et al., 2001Go)

Cassette mutagenesis

The pATP003.xb vector was doubly digested with Bsu36I and XhoI to liberate a 57 bp fragment that was separated away from the linearized vector by agarose gel electrophoresis. DNA oligonucleotides were designed and used to make synthetic mutagenic cassettes to replace this fragment. Each cassette contained a silent mutation that eliminated a naturally existing PstI site, to permit screening against the wild-type. The two complimentary master oligonucleotides are as follows, with the codons to be mutated in bold and the silent mutation removing the PstI in italics: A-5'TC AGG TGG CAC CTT CAG AAG GGT TTC GTG GTC TTC CCG AAG TCG GTC CGC CGC GAG CGC C3' and B-5'TC GAG GCG CTC GCG GCG GAC CGA CTT CGG GAA GAC CAC GAA ACC CTT CTG AAG GTG CCA CC3'. The codons used for each mutant are as follows: K232G (GGC), K232H (CAC), K232M (ATG), K232Q (CAG), K232R (CGG), K232S (AGC), K232T (ACC), S233E (GAG), S233K (AAG), S233M (ATG), S233N (AAC), S233T (ACC), S233V (GTG), V234D (GAC), V234E (GAG), V234I (ATC), V234M (ATG), V234N (AAC), V234Q (CAG), V234S (TCG), R235C (TGC), R235D (GAT), R235E (GAA), R235G (GGT), R235H (CAT), R235M (ATG), R235N (AAT), R235Q (CAA), R235S (AGT), R235T (ACG), R235Y (TAC), R238D (GAT), R238E (GAA), R238F (TTC), R238G (GGC), R238H (CAT), R238N (AAC), R238Q (CAG) and R238Y (TAT). The complementary primers were annealed together and ligated into the pATP003.xb vector. The ligation reactions were transformed into E.coli cells (XLI-blue) and colonies were picked and used to innoculate 5 ml `mini prep' cultures (LB broth containing 0.1 g/l of ampicillin). Plasmid DNA was extracted from the cultures (High Pure Plasmid Isolation Kit, Roche) and a sample of each culture was screened by restriction digestion with PstI in order to verify that the plasmid was the proper size and the mutagenic cassette was present. Screened mutants were then subjected to DNA sequencing across the entire cassette region.

Activity-stained native gels

Mutant plasmids were transformed into JM109 E.coli cells for protein expression. Cultures of 50 ml were grown and induced with 1 mM IPTG when the A600 of the culture reached 1.0. Pellets were harvested 4 h post-induction and frozen for later use. Thawed pellets were lysed using 2 ml of Bug Buster detergent in the presence of Complete Protease Inhibitor Cocktail tablets. The soluble lysate was loaded directly on to native polyacrylamide gradient gels with a loading dye. Identical gels were run at 3 W per gel at 4°C and were then soaked in 50 mM Bis-Tris, pH 7.0, and either 1 mM NADH or 1 mM NADPH. After rinsing the gels under running water, they were overlaid with filter-paper soaked with 50 mM Bis-Tris, pH 7.0, and 20 mM 2,5-DKG, as described by Seymour and Lazarus (Seymour and Lazarus, 1989Go). Images were taken under UV light at various times after staining, for up to 2 h. The gels were then stained with Coomassie Brilliant Blue to observe protein content. All mutants were assayed at least twice by this method.

Protein expression and purification

Mutants with apparent NADH activity were scaled up to 1l cultures, which were grown and induced as described above. The crude lysate obtained after removal of the cell debris using Bug Buster was buffered with 20 mM sodium acetate, pH 5.5. This was subjected to a batch weak ion-exchange step with DEAE-cellulose resin. The material that eluted with 0.5 M NaCl was diluted to contain 0.15 M NaCl with 20 mM sodium acetate, pH 5.5, and loaded on to a hyper D DEAE F resin column using an FPLC apparatus. The column was eluted with a linear sodium chloride gradient and the active fractions were pooled. These were concentrated using a PM-10 ultrafiltration membrane in a stirred cell unit (Amicon). The concentrate was then loaded on to a Superdex-75 FPLC gel filtration column (Pharmacia Biotech). Active fractions from a single peak were pooled and the homogeneity of the purified protein was verified using SDS–PAGE under reducing conditions. The concentrations of the purified proteins were obtained from their absorbance at 280 nm using 43 430 M–1 cm–1 as an extinction coefficient (Pace et al., 1995Go). These values were then verified using the method outlined by Bradford (Bradford, 1976Go).

Substrate preparation

2,5-DKG was prepared from P.citrea cells that had been induced during growth with sodium gluconate. The induced cells were harvested and washed and stored at 4°C for up to several months. They were then used to convert sodium gluconate enzymatically to sodium 2,5-DKG. The cells were removed by centrifugation, the solution was filtered and the final 2,5-DKG concentration was determined enzymatically. This was accomplished by combining dilutions of the 2,5-DKG stock solution with an excess of NADPH and 2,5-DKG reductase at pH 7.0. The enzyme has been demonstrated to be essentially irreversible at this pH (Miller et al., 1987Go). The concentration of NADPH was monitored for 16 h by measuring the absorbance at 375 nm. A kinetic model was created that accounted for the loss of NADPH due to reaction and for the natural degradation of NADPH over time and this model was used to solve for the initial 2,5-DKG added to each reaction. These values were obtained in triplicate and averaged together to obtain a final 2,5-DKG concentration of the solution.

In order to verify this concentration, ~1 mM samples were converted to 2-KLG using an excess of NADPH and 2,5-DKG reductase. These samples were analyzed with a 4 mm IonPac AS10 Dionex HPLC column with 50 mM sodium acetate, pH 4.5, as the mobile phase and the 2-KLG concentration was compared with known standards. The final quantified 2,5-DKG product was used directly in the kinetic reactions, without further purification.

Enzyme kinetics

Kinetic reactions were performed in 96-well microtiter plates and read with a temperature-controlled SpectraMax 190 plate reader (Molecular Devices). All reactions were performed in a volume of 200 µl, at 25°C and buffered with 50 mM Bis-Tris, pH 7.0. The oxidation of NADH was followed at 375 nm and only initial rates were collected. This wavelength was used instead of the standard 340 nM because the absorbance remains linear at the high concentrations of NADH that were used. Both substrate and cofactor concentrations were varied and the data were fitted to the rate equation for the ordered bi bi kinetic mechanism. The equation was eventually simplified as described in the Results section. Each measurement was done in at least triplicate and the non-linear regression of the averaged unweighted data was performed with the NLREG program using two independent variables (Dennis et al., 1981Go).


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Structural details of residues chosen for mutagenesis

All of the residues that interact with the 2'-phosphate of NADPH in the cofactor-binding pocket of 2,5-DKG reductase A were chosen as preliminary targets for site-directed mutagenesis (Figure 1Go). The crystal structure of the protein allows for an analysis of the interactions between the residues of the native enzyme and the NADPH molecule.

The lysine at position 232, which is buried beneath the bound cofactor, has several interactions with this molecule. The backbone amide nitrogen of the residue forms a hydrogen bond with the adenosine side of the pyrophosphate group. The aliphatic carbon atoms of the side chain lie directly beneath the adenosine ribose. Finally, the positively charged nitrogen of the {varepsilon}-NH3+ group forms an ionic bond with the 2'-phosphate of NADPH and a hydrogen bond with 3'-hydroxyl of the adenosine ribose. A lysine is found at position 232 in almost every member of the AKR superfamily (Table IGo).

The highly conserved serine at position 233 also interacts with the 2'-phosphate group. This residue is buried beneath the bound cofactor. There is a hydrogen bond between the side chain hydroxyl and the 2'-phosphate group. In addition, the C{alpha}, Cß and backbone carbonyl carbons exhibit close van der Waals interactions with the adenosine ribose.

Although Val234 lies adjacent to Ser233, it is a much more solvent accessible residue. Its backbone amide makes a hydrogen bond with the 2'-phosphate group. One of the {gamma}-carbons lies very close to the 2'-phosphate. In other members of the AKR superfamily, this residue is found as an alanine, isoleucine, threonine, serine, asparagine, phenylalanine or tyrosine (Table IGo).

Arg235 was identified using the Ligand–Protein Contacts program, but a role for this residue in cofactor binding in 2,5-DKG reductase has not been noted previously. Its {varepsilon}-nitrogen and one of its terminal guanido nitrogens form a hydrogen bond and an ionic bond, respectively, with the 2'-phosphate. Like Val234, Arg235 is also fairly solvent accessible and its interactions with the rest of the cofactor are minor. In other members of the family it is replaced by threonine, asparagine, aspartate, lysine, serine and histidine. Because an arginine at position 235 is very rare in the AKR superfamily (Table IGo), it is an obvious target for mutagenesis.

The final residue identified was a highly conserved arginine found at position 238. It is positioned adjacent to Arg235 in the three-dimensional structure and the two side chains together resemble fingers cradling the 2'-phosphate of NADPH. One of the terminal guanido nitrogens is found in position to make an ionic bond with the 2'-phosphate. In addition, both terminal nitrogens can make hydrogen bonds with nitrogens in the adenine ring. The aliphatic carbons of the residue are found to lie adjacent to the adenine ring. There is also another hydrogen bond between the {varepsilon}-nitrogen and a nitrogen in the adenine ring. It appears that Arg238 is important for aligning the position of the adenine ring and the positive charge of the guanido group favors NADPH as a cofactor.

Mutant construction and activity-stained native gel assay results

The oligonucleotide cassettes were annealed and ligated into the pATP003.xb vector. The accuracy of each mutation was verified by sequencing across the entire cassette region. While trying to produce the V234M mutant, the double mutant V234M/R235C (ATG/TGC) was obtained incidentally and was retained for analysis. A total of 40 mutants were constructed and transformed into E.coli strain JM109 for expression.

Lysates from cells expressing the mutant proteins were prepared (see Materials and methods) and loaded directly on to native polyacrylamide gels which were electrophoresed at 4°C in a cold room. Identical gels were soaked in a solution of either NADPH or NADH and then overlaid with filter-paper soaked in 2,5-DKG. The gels were then examined under UV light at different times for up to 2 h to see if the activity of the different mutants could be detected. Reduced cofactor fluoresces in the UV, but oxidized cofactor will not. Also, by using the native gel assay, the active reductase migrates away from contaminating proteins in the gel, thereby allowing for a rapid and qualitative way to compare the activities of the mutant and the wild-type enzymes with either NADH or NADPH (Seymour and Lazarus, 1989Go). Once the activities were recorded, the gels were stained with Coomassie Brilliant Blue to assess the relative levels of expressed protein. The native gel results with the 40 site-directed mutants can be found in Table IIGo and Figure 2Go shows an example of the native gels.


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Table II. Native gel results for cofactor-binding pocket mutants
 


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Fig. 2. Example of activity-stained native polyacrylamide gels. By running crude lysates of bacteria expressing the mutant enzymes on native polyacrylamide gels, the 2,5-DKG reductase enzyme could be migrated away from contaminating activity. Gels were prepared as described in Materials and methods. The gels were then viewed under UV light to detect areas where the cofactor had been consumed. (a) Image of a gel using NADH as a cofactor after 2 h. (b) Image of the same crude lysates using NADPH as a cofactor after 20 min. (c) The same crude lysates with a Coomassie Brilliant Blue stain to visualize the total protein content on the gels. The wild-type was run in lane 1, lane 2 contains the R238D mutant, lane 3 is the R238E mutant, lane 4 is the K232G mutant and lane 5 contains the R238H mutant reductase. The arrows indicate the location of the reductase enzyme.

 
Some generalizations can now be made about each of the sites chosen for mutagenesis. Mutations at Val234 did not appear to have significant impact on the catalytic properties or expression of the enzyme. The active protein levels and the activity with NADPH were similar to wild-type for all of the mutants and no activity with NADH was detected for any of the mutants. This is in contrast to the drastic effect seen with mutations made at Ser233. All of the mutations tested at Ser233, other than the conservative S233T mutant, appeared to decrease active protein levels and eliminate all catalytic activity with NADPH as a cofactor. This suggests that a ß-hydroxyl group at this position is crucial for both catalytic activity and for proper folding or thermodynamic stability. None of the mutations at Ser233 led to detectable activity with NADH. Therefore, Ser233 and Val234 do not appear to be useful sites for site-directed mutagenesis in order to improve activity with NADH.

Mutagenesis of the remaining three contact residues studied did result in the discovery of mutants exhibiting improved catalysis with NADH as a cofactor. Increased activity with NADH was observed in the native gel assay when Lys232 was mutated to glycine, methionine, glutamine and serine. Increased activity with NADH was also detected when Arg235 was mutated to glycine and threonine and when Arg238 was mutated to glutamate and histidine, with R238H producing the most intense activity band of all of the mutants on the NADH-stained native gel (Table IIGo).

To characterize further the effects of this mutagenesis effort, the NADPH activity of each mutant was also examined on the native gel assay. NADPH activity was retained in all of the mutants except for the previously mentioned non-conservative mutations at Ser233 and the charge reversal mutants to aspartic and glutamic acids at Arg238 (Table IIGo, Figure 2Go). Interestingly, these same charge reversal mutations at Arg235 did retain NADPH activity.

Expression and purification

The eight single mutants with apparent activity with NADH and the wild-type enzyme were expressed in 1 l cultures and purified by two ion-exchange steps and one gel filtration step. Previous purification protocols have been reported for this enzyme, but they relied on an affinity chromatography step with colored dye resins (Miller et al., 1987Go; Powers, 1996Go). Since mutations are being made in the cofactor-binding site, a new purification protocol was developed that would not be affected by the mutations. The homogeneity of each purified mutant was verified using SDS–PAGE. Protein concentrations were determined spectrophotometrically and verified by the Bradford method (Bradford, 1976Go). The new purification protocol yielded 2–4 mg of reductase per liter of culture.

Steady-state kinetics

The activities of the purified enzymes were assayed in triplicate in 96-well microtiter plates using a plate reader. Both the substrate concentration and the NADH concentration were varied and the data were used to fit the rate equation for an irreversible, ordered bi bi mechanism in the absence of products:


(1)
where dP/dt is the initial rate of product formation, Et is the total enzyme concentration, kcat is the turnover number, A is the concentration of NADH, B is the concentration of 2,5-DKG, Kia is the dissociation constant for the enzyme–NADH complex, Ka is the Michaelis constant for NADH and kb is the Michaelis constant for 2,5-DKG (Cleland, 1963Go).

It soon became clear that the parameters could not be adjusted in this model to give a satisfactory fit, owing to the large apparent value of KM for 2,5-DKG and the low concentrations of 2,5-DKG that were used in the study. The 2,5-DKG concentrations were not large enough to saturate the enzyme (Figure 3BGo), so a further mathematical simplification was required. Preliminary fitting of the model suggested that both kcat and kb were very large in comparison with Ka and Kia. Therefore, the rate equation could be simplified to the following form by dividing through by Kb:



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Fig. 3. Sample kinetic data of wild-type and mutant enzymes with apparently improved activity with NADH. Initial rate measurements were performed for the wild-type and each mutant enzyme in triplicate at 25°C and pH 7.0 with varying concentrations of NADH and 2,5-DKG. (a) Initial rates with 18 mM 2,5-DKG and varying NADH concentrations. Note that mutants K232M, K232Q and K232S appear to be inferior to the wild-type enzyme. (b) Initial rates with 2 mM NADH and varying concentrations of 2,5-DKG. Note that all of the plots are linear (best linear fits are shown for illustration), thus permitting a simplification in the kinetic rate equation. Wild-type (x), K232G (•), K232M ({circ}), K232Q ({blacksquare}), K232S ({square}), R235G ({blacktriangleup}), R235T ({triangleup}), R238E ({lozenge}) and R238H ({blacklozenge}).

 

(2)

This equation is only valid when B < < Kb (Dixon and Webb, 1979Go). It can be used to capture the Michaelis–Menten behavior with NADH as a cofactor (Figure 3AGo), when the reaction rate is directly proportional to the concentration of the substrate, 2,5-DKG (Figure 3BGo). Using this equation, it was possible to achieve a much better fit of the kinetic data for the purified enzymes, as can be seen in Figure 4Go.



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Fig. 4. Complete kinetic data for wild-type and the R238H mutant using a simplified kinetic rate equation (Equation 2Go). Initial rate measurements were performed for the wild-type (a) and the R238H mutant enzyme (b). Double reciprocal plots of the same data and linearized fits are shown as insets. Data were collected in triplicate at 25°C and pH 7.0 with varying concentrations of NADH and 2,5-DKG concentrations of 4.5 (•), 9.0 ({circ}), 13.5 ({blacksquare}), 18 ({square}), 27 ({blacktriangleup}) and 45 mM ({triangleup}). Note the difference in the ordinate scales and the excellent fit obtained with only two adjustable parameters in the kinetic rate equation. The error bars are the SDs of the averaged rate measurements. When not visible, the error bars are smaller than the data symbol used.

 
Other comparisons can be made between the different mutants using these parameters. When NADH is the varied substrate, the term kcatB/Kb becomes the apparent kcat and Kia or the dissociation constant for the EA complex (Kd), becomes the apparent Ka. Therefore, the term kcatB/KbKia becomes the apparent kcat/Ka or apparent specificity constant, which is generally a good way to compare the kinetic performance of the different mutants with the NADH cofactor (Fersht, 1985Go). Dividing these parameters by the wild-type values allows them to be readily compared (Table IIIGo).


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[in this window]
[in a new window]
 
Table III. Kinetic parameters with NADH as a cofactor using the modified rate Equation (2)Goa
 
Upon completion of the kinetic characterization using purified enzymes, three of the four Lys232 mutants identified on the native gel assay appeared to be no better than wild-type with NADH. K232Q, K232M and K232S all had an apparent kcat below that found with wild-type and an apparent Ka that was higher than wild-type. The only mutation at Lys232 that appeared to confer increased activity (albeit slight) compared to the wild-type enzyme was the surprising K232G mutation. This mutant has both a slightly higher apparent kcat and a lower apparent Ka, as compared with the wild-type. Given the extremely non-conservative nature of the Lys to Gly substitution, it is perhaps remarkable that the impact on enzyme activity was so modest.

The two purified mutants with Arg235 substitutions exhibited increases in both the apparent kcat and the apparent Ka. Both mutations increased the apparent Ka more than 3-fold. The R235T mutation increased the apparent kcat almost 2-fold, while the R235G mutant increased it almost 5-fold.

The two mutations at Arg238 had very different effects. The R238E mutation increased the apparent Ka more than 3-fold and it also doubled the apparent kcat. Hence the R238E mutant appeared kinetically very similar to the Arg235 mutants. The R238H mutant was clearly the best obtained in this study. This mutant exhibited a slightly lower apparent Ka than the wild-type and showed a 5-fold improvement in the apparent kcat. Combined, these parameters resulted in an apparent specificity constant (kcatB/KiaKb) that was improved almost 7-fold over the wild-type.

Changes in cofactor binding energies

The Kia values obtained in the modified kinetic rate equation are the dissociation constants for the bound NADH molecule for each mutant. These values can be used to compare the effects of the mutations on the ground and transition-state binding energies (Fersht, 1985Go). The change in the ground-state binding energy for NADH can be calculated by:


(3)

The change in the binding energy for the transition-state of the reaction is given by:


(4)

As mentioned above, the apparent kcat/Ka for the mutants is given by kcatB/KiaKb in the modified rate equation. Therefore, the change in the transition-state binding energy can be estimated by the following:


(5)
where R is the gas constant and T is the absolute temperature (Table IVGo).


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[in this window]
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Table IV. Changes in the cofactor binding energies of the NADH-dependent mutants
 
The ground-state binding energy changes were very minor, ranging from a loss of 0.72 kcal/mol for R235T to a gain of 0.16 kcal/mol for K232G. The changes in the transition-state binding energies were also small except for the R238H mutant, which gained 1.1 kcal/mol of binding energy (Table IVGo).


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Worldwide consumption of vitamin C now exceeds 50000 tons per year. The bulk of the vitamin C is produced through the complex Reichstein and Grussner synthesis (Reichstein and Grussner, 1934Go). For economic reasons and to lessen the impact of the production process on the environment, considerable effort has been expended in a quest for a more efficient process based on enzymatic biosynthesis (Anderson et al., 1985Go; Lazarus et al., 1989Go, 1990Go; Boudrant, 1990Go; Boston and Swanson, 2000Go). Two particularly promising biosynthetic production schemes that employ the 2,5-diketo-D-gluconic acid reductase enzyme as a catalyst have been described (Anderson et al., 1985Go; Boston and Swanson, 2000Go). However, the Corynebacterium 2,5-DKG reductases that have been employed exhibit virtually absolute specificity for NADPH; little or no activity is detectable with NADH. Altering the specificity of this enzyme to enable it to use NADH more effectively as a cofactor has the potential to improve both processes. Therefore, we used mutagenesis along with kinetic analyses to elucidate the molecular determinants of cofactor specificity in 2,5-DKG reductase A.

A variety of single mutations, encompassing side chain substitutions spanning a range of different chemical properties, were constructed and used to probe five different sites in the cofactor-binding pocket of the 2,5-DKG reductase enzyme in order to find mutations that promoted catalytic activity with NADH. The mutant proteins with apparent increased activity with NADH from the native gel assay were purified to homogeneity and kinetically characterized by examining initial rates and the data were fit to a simplified rate equation for an irreversible ordered bi bi mechanism in the absence of products (Equation 2Go). This allowed a comparison of the effects of the different mutations on the kinetic and energetic parameters for the enzyme. Upon completion of this initial analysis, some of the mutants did not appear to be improved over the wild-type enzyme. The reason for their tentative identification as having activity with NADH on the native gels may have been due to expression levels or other effects unrelated to intrinsic cofactor specificity. Because the gel-based assay was designed to be extremely sensitive, such `false positives' were not unexpected.

Only at three positions did mutagenesis result in enzymes with bona fide increases in NADH activity. Several of the mutants exhibited improved apparent kcat values, but only three mutants yielded apparent kcat/Ka values (kcatB/KiaKb) that were better than the wild-type values. K232G and R235G exhibited a slight 1.5-fold increase and the most successful mutant, R238H, exhibited an almost 7-fold improvement over the wild-type 2,5DKG reductase with NADH as a cofactor (Table IIIGo).

The results with some of these mutations were surprising. In the crystal structure, the native lysine at position 232 lies beneath the bound cofactor and appears to orient the molecule (Figure 1Go). The non-conservative K232G mutation would presumably abolish this putative orientation function and might even permit more flexibility in the NADH-binding pocket. Regardless of this, only small changes in both the ground-state and apparent transition-state cofactor-binding energies were observed for this mutant as compared with the wild-type enzyme. The K232G mutation results in a small decrease in the apparent Ka and a small increase in the apparent kcat. Highly non-conservative contact mutations that impart only minor changes in energy parameters have been observed previously and have been termed `isofunctional' (Bigler et al., 1993Go). There are no known members of the AKR superfamily with a glycine at this location (Table IGo).

The arginine at position 235 appears to interact with the cofactor solely via the 2'-phosphate of NADPH. The non-conservative R235G mutation, which should completely remove this interaction, produced a gain in the apparent kcat, but it also resulted in a concomitant gain in the apparent Ka for NADH. This implies that Arg235 forms energetically comparable interactions with the cofactor in both the ground and transition states. There are a variety of residues found at this position in the AKR superfamily, but a glycine at this location has not yet been reported (Table IGo). Surprisingly, none of the mutations that we made abolished activity with NADPH (Table IIGo), indicating that the apparent Arg235 side chain guanido group interaction with the NADPH 2'-phosphate is not critical for enzyme activity.

The best mutant obtained in this study, from the standpoint of activity with NADH, contained a histidine at position 238. From the crystal structure of the native enzyme, arginine at this position appears to lie flush against the adenine ring of the cofactor and form a salt bridge with the 2'-phosphate group of NADPH (Figure 1Go). The Arg -> His substitution is relatively conservative when protonated and may slightly enhance the interaction with the adenine ring via a stacking interaction. This mutation led to a decrease in the apparent Ka for NADH that was similar to that seen with the K232G mutation, yet the mutant enzyme also exhibited an increase in the apparent kcat that exceeded the gain achieved by the R235G mutation. Energetically, this mutation resulted in only a minor gain in ground-state binding energy, yet it demonstrated the largest gain of any of the mutations in apparent transition-state binding energy: 1.1 kcal/mol (Table IVGo). A histidine has not been reported in this position in any member of the AKR superfamily (Table IGo).

The natural substrate for 2,5-DKG reductase has yet to be definitively identified. Recent molecular modeling efforts with this enzyme, using NADPH as a cofactor, have suggested that the natural substrate is a molecule larger than 2,5-DKG and that the active site pocket could be optimized to catalyze the 2,5-DKG -> 2-KLG reaction more efficiently (Khurana et al., 2000Go). The catalytic improvements seen with the R238H mutant may be the result of improving the geometry of the cofactor in relation to the substrate in the transition state. This is reflected in the apparent kcat value, which is the actual specificity constant (kcat/Kb) for 2,5-DKG. In the R238H mutant, this ratio was improved more than 5-fold.

All of the mutants that were identified to have NADH activity retained activity with NADPH except for R238E. This mutant and the R238D mutant both abolished NADPH activity, as assayed on the native gels. It has been demonstrated in rat 3-{alpha}HSD that the arginine residue analogous to R238 is required for pre-steady-state kinetic transients to be observed upon NADPH binding. These kinetic transients were attributed to an anchoring of NADPH by the arginine (Ratnam et al., 1999Go). It is now apparent that a charge reversal at this site can prevent catalytic activity with NADPH, which is consistent with the anchoring hypothesis. However, the cofactor-binding pocket must still be intact, as seen by the retention of activity with NADH in the R238E mutant. By eliminating activity with NADPH and increasing activity with NADH, this mutation effectively reverses the cofactor specificity, albeit with a low resultant level of overall activity.

There has recently been a great deal of interest in the potential of combinatorial methods for engineering new proteins. However, these methods may work best when certain sites are targeted for mutagenesis while others are left unchanged (Miyazaki and Arnold, 2000Go). In this work we initially used structural information to identify five promising sites for mutagenesis in the residue 232–238 loop of the enzyme. By using both conservative and non-conservative site-directed mutations we were able to eliminate two of these potential residues, leaving three good choices for combinatorial mutagenesis in this loop.

Other work in the cofactor-binding pocket of the 3{alpha}-HSD enzyme has shown that the loop anchoring the 2'-phosphate of NADPH is not the sole determinant of cofactor specificity in the AKR superfamily and that NADPH and NADH may exhibit different modes of binding (Ma et al., 2000Go). This suggests that mutations made at many locations in the cofactor-binding pocket may be required to produce improvements in NADH-mediated catalysis. Although the kinetic improvements made in the present study are modest, we are now beginning to understand the structural determinants of the cofactor specificity in the 2,5-DKG reductase and this information will be used to guide future mutagenesis efforts aimed at improving our understanding of this enzyme


    Notes
 
4 Present address: Applied Molecular Evolution, 3520 Dunhill Street,San Diego, CA 92121, USA Back

6 To whom correspondence should be addressed. E-mail: anderson{at}cabm.rutgers.edu Back


    Acknowledgments
 
We thank Jim Kellis at Genencor International for reviewing the manuscript and Matt Boston at Genencor International for assistance with the HPLC. This work was funded in part by the Advanced Technology Program of NIST through a shared-funding project grant to Genencor International. S.B. was a trainee of the NIH Predoctoral Training Grant in Biotechnology (5T32 GM08339).


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 Top
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 Introduction
 Materials and methods
 Results
 Discussion
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Received June 6, 2001; revised November 8, 2001; accepted November 9, 2001.





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