Analysis of stability and catalytic properties of two tryptophanases from a thermophile

Hiromi Kudo1, Ryo Natsume, Makoto Nishiyama2,3 and Sueharu Horinouchi

Department of Biotechnology and 2 Biotechnology Research Center, The University of Tokyo, Bunkyo-ku, Tokyo 113-8657, Japan


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Two tryptophanases, Tna1 and Tna2, both of which were cloned from the thermophile Symbiobacterium thermophilum, differ in their enzymatic properties, such as thermal stability, catalytic efficiency and activation energy of catalysis, despite the great similarity (92%) in their amino acid sequences. Chimeric tryptophanases were constructed by recombination of the two genes to try to elucidate the molecular basis for the difference. The stability of each chimeric enzyme was roughly proportional to the content of amino acid residues from Tna1. Three regions, tentatively named regions 2, 4 and 5, which contained the amino acid residues 70–129, 192–298 and 299–453, respectively, were especially important for the increase in thermal stability. Site-directed mutagenesis revealed that V104 in region 2 and Y198 in region 4 of Tna1 were involved in the increase in thermal stability of Tna1. Amino acid residues contributing to the higher catalytic efficiency of Tna1 were similarly analyzed, using the chimeric tryptophanases, and found to be located in region 5. Site-directed mutagenesis revealed that I383 and G395 in Tna1, which were presumably located close to the putative active center, played an active role in the increase of catalytic efficiency of Tna1. The activation energy of catalysis was proportional to the content of amino acid residues from Tna2, suggesting the amino acid residues responsible for the difference were dispersed over the whole molecule.

Keywords: activation energy/catalytic efficiency/thermal stability/thermostable enzyme/tryptophanase


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Tryptophanase (EC 4.1.99.1) is a tetrameric pyridoxal enzyme that catalyzes the conversion of L-tryptophan to indole, pyruvate and ammonia through the {alpha},ß-elimination reaction (Wood et al., 1947Go; Snell, 1975Go; Terasawa et al., 1991Go). In addition to the {alpha},ß-elimination, tryptophanase also catalyzes the reversal of {alpha},ß-elimination (Watanabe and Snell, 1972Go) and ß-replacement (Newton and Snell, 1964Go). The enzyme, therefore, can be used as a biocatalyst for the synthesis of L-tryptophan through the reversal of {alpha},ß-elimination.

We isolated from soil an obligately symbiotic bacterium, Symbiobacterium thermophilum, which grew only in co-culture with a Bacillus strain, and acts as a potent thermostable tryptophanase producer (Suzuki et al., 1988Go). Cloning and nucleotide sequence analysis revealed that S.thermophilum possesses two highly homologous tryptophanase genes, tna1 and tna2, each of which encodes an enzyme of 453 amino acid residues, giving 92% identity in amino acid sequence (Figure 1Go) (Hirahara et al., 1992Go). Since the two genes are located close to each other in tandem, it is likely that a gene duplication event has occurred. Despite the great similarity, however, the two enzymes markedly differed in thermal stability and activation energy of catalysis (Hirahara et al., 1992Go). When compared to Tna2, Tna1 is more stable at high temperature, shows higher activity and requires a lower activation energy for catalysis. This indicated that a limited number of the amino acids exchanged were responsible for the differences. In order to elucidate the molecular basis for the differences in the enzymatic properties of the two tryptophanases, we constructed chimeric enzymes between Tna1 and Tna2 and carried out site-directed mutagenesis of Tna2, and analyzed the effect of the amino acid substitutions on the thermal stability and catalytic profiles. In this paper, we describe the enzymatic properties of the chimeric tryptophanases and the mutants resulting from site-directed mutagenesis and discuss the amino acid residues responsible for the differences.



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Fig. 1. Amino acid sequence alignments of tryptophanases from P.vulgaris and S.thermophilum. Tryptophanase from P.vulgaris is shown as PvTnase. The amino acid residues identical to those of Tna1 are indicated by asterisks. Secondary structures determined by X-ray analysis of P.vulgaris tryptophanase are also shown. Open arrows, boxes, Ts and Gs, represent ß-strands, {alpha}-helices, ß-turns and 310-helices, respectively. Five regions which tentatively we named are also shown.

 

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

All the restriction enzymes, AmpliTaq DNA polymerase and T4 DNA ligase were purchased from Takara Shuzo Co. (Otsu, Japan). Oligonucleotides were synthesized using a DNA synthesizer, 8900 Nucleic Acid Synthesis System (Millipore Japan Co., Tokyo).

Gene manipulation

General techniques for DNA manipulation were based on Sambrook et al. (1989). Nucleotide sequence was determined by the dideoxy chain-terminating method (Sanger et al., 1977Go) using M13 vectors (Yanisch-Perron et al., 1985Go). Site-directed mutagenesis was carried out by the method of Kunkel (1985).

Bacterial strains and plasmids

Escherichia coli JM105 was used as the host for plasmid construction and M13 phage propagation. Escherichia coli CY15077 (W3110 tnaA2 {Delta}trpEA2) (Kamath and Yanofsky, 1992Go), which was kindly supplied by Dr C.Yanofsky (Stanford University), was used as the host for the expression of the tna genes.

For the efficient expression of tna1 and tna2, two plasmids, pUC18-tna1 and pUC18-tna2, were constructed by PCR. For construction of pUC18-tna1, PCR was performed using the following two oligonucleotides as primers: 5'-CCGAATTCCTGAAAAGGAGGAGTTCACATGCCCAAG-3', which was designed to introduce an EcoRI site just upstream of the translational initiation codon ATG for tna1, and 5'-CACCGCCATGGCCTCTAGATCTCGCCC-3', which contains a BglII site; pTNA15 (Hirahara et al., 1992Go) was used as the template. PCR conditions were as follows: 94°C for 1 min, 50°C for 1 min and 72°C for 1.5 min, in a total of 25 cycles. The amplified DNA fragment was digested with EcoRI and BglII. Additional PCR was carried out with the same template and two oligonucleotide primers: 5'-GGGCGAGATCTAGAGGCCATGGCGGTG-3', which has the sequence complementary to that of the oligonucleotide shown above and therefore possesses a BglII site, and 5'-GGTGCCGAAAGCTTTGGTTCAAGCCGGGTCGAA-3', which was designed to introduce a HindIII site just downstream of the stop codon. The amplified DNA fragment was digested with BglII and HindIII. Both restriction fragments were purified by agarose gel electrophoresis, and ligated with pUC18 digested with both EcoRI and HindIII. The resulting plasmid was named pUC18-tna1. pUC18-tna2 was constructed as follows: PCR was carried out under the conditions shown above using two oligonucleotides, 5'-CCGAATTCCTGAAAGGAGGAACCAGTGATGCCAA-AG-3', which were designed to introduce an EcoRI site just upstream of the translational initiation codon ATG for tna2, and 5'-GTCCAGCAGCAGGTCGATCGGCCGGCCGGC-3'. The amplified fragment was then digested with EcoRI and PmaCI. PCR was also carried out using the oligonucleotides, 5'-GGCCGGGATCTGGAGGCCATGGCGGTG-3' and 5'-CGCGTGAAGCTTGAGCTAGACCAGATCGAA-3', which were designed to introduce a HindIII site just downstream of the stop codon, and the amplified DNA fragment was digested with BamHI and HindIII. The two restriction fragments were purified by agarose gel electrophoresis and ligated with the PmaCI–BamHI fragment encoding a central portion of Tna2 and pUC18 digested with both EcoRI and HindIII to yield pUC18-tna2.

Chimeric tryptophanases between Tna1 and Tna2 were constructed by exchanging regions of the genes using six restriction sites, EcoRI, SplI, PmaCI, BalI, NcoI and HindIII, all of which were conserved in both of the expression plasmids, pUC18-tna1 and pUC18-tna2. Each of the regions, tentatively named regions 1, 2, 3, 4 and 5, contain the amino acid sequence of 1–69, 70–129, 130–191, 192–298 and 299–453 (1–71, 72–138, 139–203, 204–311 and 312–466 in Proteus vulgaris tryptophanase numbering), respectively.

Site-directed mutagenesis of Tna2

Site-directed mutagenesis of TNA2 were generated as follows. The EcoRI–BamHI fragment from pUC18-tna2 carrying the regions 1, 2, 3 and 4, and a small portion of region 5, was subcloned into M13mp18. The BamHI and HindIII fragment from pUC18-tna2, carrying mostly region 5, was subcloned into M13mp18. These two RFI DNAs were used as the template for site-directed mutagenesis (Kunkel, 1985Go). The following oligonucleotide primers were used: 5'-GGTGATCAATTGCGCGAAGGCGACCTTCTC-3' for S104V, 5'-GTACTTCCTCGCGATGCGGTACGTCTCGCG-3' for S198Y, 5'-GGTGCGCACGCCGCCCTCGAGGTAGAGCGCCACCGT-3' for Q369L, 5'-GGTCGCGACCCATCATGAGCGAGCC-3' for V382M, 5'-GTCGCGGCCGATCACTAGTGAGCCGACCTC-3' for M383I, and 5'-GAACTCGAACGGGCCCCGGACGTT-3' for S395G. For construc- tion of the V382M/M383I/S395G mutant, the M13mp18 derivative containing S395G mutation was used as the template for further mutations, V382M/M383I, using the primer, 5'-GCGGCCGATCATGAGGGATCCGACCTC-3'. After confirmation of the mutation by DNA sequencing, the mutations were introduced into the corresponding positions of pUC18-tna2 using several restriction endonucleases. The gene encoding the S104V/S198Y/Q369L mutant was constructed by combining each mutation using restriction sites.

Production of tryptophanases

Escherichia coli CY15077 cells harboring the expression plasmids were precultured overnight in 10 ml 2xYT medium (Sambrook et al., 1989Go) containing 50 µg/ml ampicillin at 37°C. The preculture was transferred into fresh 2xYT medium (1 l) containing 50 µg/ml ampicillin and incubated for an additional 20 h at 37°C. The cells were harvested by centrifugation, suspended in 10 ml 50 mM potassium phosphate buffer (pH 7.0) and disrupted by sonication. After centrifugation, the supernatant was dialyzed overnight at 4°C against 2 l of 50 mM potassium phosphate buffer (pH 7.0) supplemented with 0.1 mM 5'-pyridoxal phosphate (PLP) and 0.1 mM serine protease inhibitor, Pephabloc SC (Merck, Rahway, NJ). The dialysate was applied to a DEAE-Toyopearl (Toso, Tokyo, Japan) column (40 mm x 30 cm) and eluted with a linear gradient of KCl (0–400 mM). Tryptophanase activities in the fractions were assayed using Kovack's reagent composed of 3:1 n-amylalcohol/conc. HCl containing 5% (w/v) p-dimethylbenzaldehyde (Smibert and Krieg, 1970Go). Active fractions were dialyzed against 2 l of 50 mM potassium phosphate buffer (pH 7.0) supplemented with 0.1 mM PLP and 0.1 mM Pephabloc SC, applied to FPLC equipped with a Mono Q anion-exchanger column (HR 10/10, Pharmacia, Japan), and eluted with a linear gradient of KCl (0–400 mM). Tryptophanase showed homogeneity to give a single band on SDS–PAGE (Laemmli, 1970Go). Proteins were determined by the method of Bradford (1976).

Enzyme assay

The reaction mixture (4 ml) containing 1 mM L-tryptophan, 0.1 mM PLP in 50 mM potassium phosphate buffer were preincubated for 10 min at 50°C. An appropriate amount (about 10 µg) of tryptophanase was added to the mixture and incubated for an additional 30 min at 50°C. The reaction was terminated by boiling for 5 min. After rapid cooling in iced water, the amount of pyruvate produced was determined with lactate dehydrogenase as follows. A portion (1.2 ml) of the reaction mixture was mixed with 1.5 ml 100 mM potassium phosphate buffer (pH 8.0), 0.2 ml 4.5 mM NADH and 0.1 ml lactate dehydrogenase (5 U; Sigma Co, type II from rabbit muscle). A decrease in the absorbance of NADH at 340 nm was monitored. The amount of pyruvate in the reaction mixture was calculated from the calibration curve, [pyruvate] versus [{Delta}absorbance340/min].

Kinetic parameters were determined by the method which was principally the same as described above but the reaction mixture contained various concentrations (0.1–0.6 mM) of L-tryptophan. Vmax and Km values were calculated using the application package installed in the spectrophotometer (Milton Roy SPECTRONIC 300 ARRAY, Milton Roy, Tokyo). The kcat value was obtained by dividing the maximum concentrations of pyruvate formed per second, which were calculated from the calibration curve and the Vmax value, by the concentrations of the enzyme used.

Activation energies for each of the chimeric tryptophanases were calculated from the Arrhenius plots of the rate constants, kcat values, determined at 40, 45, 50 and 55°C, as described above.

Thermal stability of chimeric tryptophanases

Purified tryptophanase (0.1 mg/ml) was incubated at various temperatures for 5 min. After immediate cooling in iced water for 15 min, residual tryptophanase activity was measured by the above method.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Construction of chimeric tryptophanases

To elucidate the molecular basis of the difference in the properties of two tryptophanases from S.thermophilum, we constructed 15 chimeric enzymes of the two tryptophanases using six restriction sites. The structures of the chimeric enzymes constructed in this study are schematically shown in Figure 2Go. All the enzymes were purified to homogeneity as demonstrated by SDS–PAGE. In most cases, approximately 10 mg tryptophanases was purified from a 1 l culture. The only exception was chimera 4, containing regions 1–4 from Tna1 and region 5 from Tna2, which was poorly expressed in E.coli cells. Because of the low protein yield (below 0.1 mg from a 1 l culture), several of the properties were not analyzed in the case of chimera 4.



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Fig. 2. Schematic representation of chimeric tryptophanase structures. Regions from Tna1 and Tna2 are shown by black and white bars, respectively. N- and C-termini are indicated as N and C.

 
Thermal stability of chimeric tryptophanases

Because the thermal denaturation process of tryptophanases was irreversible under the conditions used in this study and the thermodynamic properties therefore could not be determined easily, we analyzed thermal stability by measuring the remaining activities of chimeric tryptophanases after heat treatment at elevated temperatures. Tna1 retained significant activity even after heat treatment at 90°C, whereas Tna2 was completely inactivated after heat treatment at 80°C (Figure 3Go). Figure 3Go also shows several examples of chimeric enzymes carrying N- and C-terminal portions derived from Tna1 of various lengths. Thermal stability of the chimeras increased, as the content of the N-terminal portion of Tna1 in the chimera increased (see Tna1, Tna2, chimeras 1, 3 and 4). Consistent with the thermal stability of chimeras carrying N-terminal portions from Tna1, thermal stability increased as the content of the C-terminal portion from Tna1 increased (see Tna1, Tna2, chimeras 5 and 7). For further elucidation, we constructed several chimeras comprising regions from Tna1 and Tna2 in various combinations. When the half-inactivation temperature was plotted against the content of the amino acid residues found in Tna1 in the chimera at 38 positions where amino acid replacements were observed between Tna1 and Tna2, a loose but obvious correlation was observed (Figure 4AGo). These results indicated that various portions of the primary structure of Tna1 were involved in the higher thermal stability of Tna1. Regions 2, 4 and 5 from Tna1 were especially important for the higher stability, because chimera 14, possessing regions 2, 4 and 5 from Tna1, showed a thermal stability comparable with that of Tna1. Since the addition of region 2 from Tna1 to chimera 6, carrying regions 3, 4 and 5 from Tna1, caused a distinct increase in thermal stability (see chimeras 5 and 6), region 2 among the three regions may contribute most to the higher stability. However, chimera 2, carrying regions 1 and 2 from Tna1, and chimera 11, possessing region 2 from Tna1, had only somewhat higher thermal stability, when compared with Tna2. These results suggested that the effect of region 2 from Tna1 on the thermal stability was most effective in the presence of both regions 4 and 5.



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Fig. 3. Thermal stability of chimeric tryptophanases. Remaining activities were measured after heat treatment for 5 min at the indicated temperatures. {circ}, Tna1; •, Tna2; {square}, chimera 1; {blacklozenge}, chimera 3; {lozenge}, chimera 4; {blacksquare}, chimera 5; and {triangleup}, chimera 7.

 


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Fig. 4. (A) Relationship between half-inactivation temperature and the content of the Tna1-type amino acid residues in chimera. The contents of the amino acid residues found in Tna1 (Tna1-type amino acid residues) in chimeras were calculated at 38 positions where amino acid exchanges were observed between Tna1 and Tna2. (B) Relationship between activation energy and the content of the Tna2-type amino acid residues in the chimeras. The contents of the amino acid residues found in Tna2 (Tna2-type amino acid residues) in chimeras were calculated at 38 positions where amino acid exchanges were observed between Tna1 and Tna2.

 
Activities of chimeric tryptophanases

Tna1 possessed the catalytic efficiency (kcat/Km) of 70 (mM–1 s–1), whereas Tna2 had a lower value of 14 (Table IGo). This indicated that the amino acid substitutions also affected the catalytic activity as well as the thermal stability. When the kcat/Km values were compared among the chimeras, only the chimeric tryptophanases with region 5 from Tna1 had a significantly higher kcat/Km value. This indicated that region 5 from Tna1 contained the amino acid residue(s) which enhanced the catalytic activity.


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Table I. Enzymatic properties of chimeric tryptophanases
 
We next examined the activation energies of several chimeras, because a large difference in the activation energy was also observed between Tna1 (49 kJ/mol) and Tna2 (91 kJ/mol) (Table IGo). The analysis using chimeric tryptophanases, however, did not identify the specific regions, which are responsible for the higher or lower activation energy. We then plotted the activation energy against the content of amino acid residues found in Tna2 in the chimera at 38 positions. Similarly in Figure 4A, Goa loose but apparent correlation was again observed (Figure 4BGo). This indicated that various portions of the primary structure of Tna2 were involved in the elevated activation energy.

Site-directed mutagenesis of Tna2

Thermal stability When the hydrophobicity of the two tryptophanases was compared using parameters by Kyte and Doolittle (1982), an increase in the hydrophobicity was observed in regions 2 (0.6 kcal/mol), 4 (4.4 kcal/mol) and 5 (4.1 kcal/mol) in Tna1, although overall hydrophobicity of Tna1 was 17.6 kcal/mol lower than that of Tna2 due to a great decrease in the hydrophobicity in regions 1 (13.3 kcal/mol) and 3 (13.4 kcal/mol). The amino acid sequence alignment between Tna1 and Tna2 in regions 2, 4 and 5, in which the amino acid replacements responsible for the increased thermal stability of Tna1 were assigned, revealed that in most cases similar amino acid residues occupied corresponding positions in both enzymes. However, the amino acids at three positions, 104, 198 and 369, were obviously different from each other: positions 104, 198 and 369 are occupied by Val, Tyr and Leu in Tna1, and Ser, Ser and Gln in Tna2. We therefore conducted site-directed mutagenesis of Tna2 at these three positions and examined the effect on stability. As shown in Figure 5Go, mutations S104V and S198Y were effective in increasing the thermal stability of Tna2, while the mutation Q369L caused little effect on stability. A mutant Tna2 carrying all three mutations showed further increased thermal stability, suggesting a contribution by the introduced amino acid residues, at least V104 and Y198, to the higher thermal stability of Tna1.



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Fig. 5. Thermal stability of Tna2 mutants. Remaining activities were measured after heat-treatment for 5 min at the indicated temperatures. {circ}, Tna1; •, Tna2; {lozenge}, S104V; {blacktriangleup}, S198Y; {square}, Q369L and {blacklozenge}, S104V/ S198Y/Q369L.

 
Catalytic efficiency Activity enhancement resulted from introduction of region 5 of Tna1 into Tna2 (Table IGo). This indicated that region 5 from Tna1 contains amino acid residue(s) which enhance catalytic activity. As for tryptophanase, however, its crystal structure complexed with a substrate analog is not yet available. We therefore could not assess the possible role of amino acid substitutions found in tryptophanases from S.thermophilum in the catalytic function using the P.vulgaris tryptophanase structure. On the other hand, tyrosine phenol-lyase (TPL), which is also a tetrameric pyridoxal enzyme that catalyzes the conversion of L-tyrosine to phenol, pyruvate and ammonia through {alpha},ß-elimination, shares significant identity in its amino acid sequence with tryptophanase. The X-ray structure of TPL from Citrobacter freundii demonstrated that secondary, ternary and quaternary structures were also highly conserved between these two enzymes (Antson et al., 1993Go). In the case of TPL, substrate recognition was analyzed by its crystal structure complexed with a substrate analog (Sundararaju et al., 1997Go). According to the X-ray structures of tryptophanase and the TPL–substrate analog complexes, among 13 substitutions between the regions 5 of Tna1 and Tna2, only three amino acid residues at positions 382, 383 and 395 were suggested to be located close to the substrate recognition site. We then examined the effect of the amino acid replacements at these positions on the catalytic efficiency. Although the replacement of V382M did not increase the kcat/Km value, other replacements, M383I and S395G, caused a small increase in efficiency (Figure 6Go). Furthermore, a triple mutant containing all three mutations showed a much increased kcat/Km value compared with each single site mutant, although full enhancement of the catalytic efficiency was not achieved by introduction of only the three amino acid residues into Tna2.



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Fig. 6. Catalytic efficiency of Tna2 mutants. The kcat/Km values of Tna1, Tna2, V382M, M383I, S395G and a triple mutant with replacements V382M/M383I/S395G are indicated by bars.

 

    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Two tryptophanases from S.thermophilum showed considerably different enzymatic profiles, despite the remarkable high identity in amino acid sequence. Although three-dimensional structures of these tryptophanases were not determined, the structure of their mesophilic counterpart from P.vulgaris has recently been determined by X-ray crystallography at 2.1 Å resolution (Isupov et al., 1998Go). Significant amino acid sequence identity of tryptophanases from S.thermophilum to the enzyme from P.vulgaris (53.9% for Tna1 and 53.4% for Tna2) suggested that the tryptophanases from S.thermophilum could take conformations similar to that of the Proteus enzyme, and therefore allowed assignment of the positions of the amino acid residues, which were different between Tna1 and Tna2 on the three-dimensional structure. In this study, we found that the amino acid residues, V104 and Y198, may contribute to the increase in the thermal stability of Tna1. In P.vulgaris tryptophanase, I106, corresponding to the residue at position 104 in both tryptophanases from S. thermophilum, is surrounded by the hydrophobic residues, I95, V276, I278, F285 and V299, all of which are conserved or replaced by hydrophobic residues in both Tna1 and Tna2, and may therefore stabilize the hydrophobic core of the tryptophanase. Similarly, the amino acid residues at positions 210 and 382 in Proteus tryptophanase, corresponding to 198 and 369 in S.thermophilum tryptophanases, were also found in other hydrophobic cores. This suggested that the lower thermal stability of Tna2 was in part due to the lower hydrophobic packing by the replacements at the three positions, although the effect of Q369L was negligible. As described above, the effect of region 2 from Tna1 on the increase in thermal stability was most effective in the presence of regions 4 and 5. However, the three regions as well as the three residues were located apart from each other in the three-dimensional structure of P.vulgaris tryptophanase. At present, we have assumed that long range effects worked additively towards the increase in stability.

When compared with Tna2, chimeras 4, 9 and 10 also had somewhat increased thermal stability, although these chimeras did not contain all the three regions, 2, 4 and 5, from Tna1. According to the X-ray structure of the Proteus enzyme, tryptophanase is composed of two structurally independent domains, the small and large domains, each of which is roughly formed by the residues in regions 1 and 5, and by the residues in regions 2, 3 and 4, respectively. Chimera 4 possessed the large domain from Tna1 and chimeras 9 and 10 did the small domain from Tna1. This may suggest that a native-like and therefore a stable structure in either of the domains from Tna1 may partially increase the thermal stability of the whole molecule in these chimeras.

The structural analysis of TPL from C.freundii revealed that R404, corresponding to R401 of Tna1 and Tna2, served to bind the {alpha}-carboxylate group of the substrate analog and its phenol ring of the analog was in a hydrophobic pocket stabilized by several hydrophobic residues, M379, F448, F123 and F36 in one subunit, and V283, V284, M288 and Y291 in the neighboring subunit (Figure 7Go). In TPL, involvement of R381, which is located close to the phenolic hydroxyl of the substrate analog, in substrate recognition was elucidated by site-directed mutagenesis (Antson et al., 1993Go). The Arg residue and most of the residues forming a hydrophobic substrate-binding pocket are even conserved in both Tna1 and Tna2. The amino acid residues at positions 385 and 386 in TPL, which correspond to 382 and 383 in S.thermophilum tryptophanases respectively, form the bottom edge of the substrate binding cleft, which could affect the catalytic efficiency through a subtle change in the hydrophobic environment of the substrate binding site. Consistent with this, site-directed mutagenesis in this study revealed that I383 in Tna1 served to partially enhance the catalytic efficiency. Although no positive effect in the catalytic efficiency was introduced by the single mutation, V382M, it may be possible that the mutation was effective in increasing the efficiency in combination with M383I, because the simultaneous mutations at the three sites caused a remarkable increase in the efficiency. The amino acid residue at position 408 (corresponding to 395 in S.thermophilum tryptophanases), on the other hand, is not located as close to the catalytic site. Therefore this residue would not be directly involved in substrate recognition nor catalysis. According to the structure of P.vulgaris tryptophanase possessing Ala at this position, a Gly (Tna1) to Ser (Tna2) substitution could form a new hydrogen bond from the hydroxyl side chain of Ser to E398-OE2 or S380-N. The fact that the S395G mutation increased the efficiency may suggest that the presence or absence of a possible hydrogen bond might affect the catalytic efficiency.



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Fig. 7. Stereo view of the substrate binding site in TPL. The bound substrate analog, 3-(4'-hydroxyphenyl)propionic acid, is also shown as HPP. Residues from an adjacent subunit are marked by asterisks. The figure was drawn using a software WebLabTM ViewerLite 3.1 (Molecular Simulation Inc., San Diego, CA).

 
These two tryptophanases may become a good model for the study of the structure–activity–stability relationships of proteins, because the two proteins are highly homologous. X-Ray analysis of Tna1 and Tna2 and further site-directed mutagenesis based on their structures will help further the understanding of the molecular basis of the differences in properties of the two proteins.


    Acknowledgments
 
We are grateful to Dr C.Yanofsky (Stanford University) for providing us with a tryptophanase-deficient mutant strain, E.coli CY15077.


    Notes
 
1 Present address: Lead Generation Laboratory, Medical Research Laboratories, Taisho Pharmaceutical Co., Ltd, 403, Yoshino-cho 1-chome, Omiya-shi, Saitama, 330-8530, Japan Back

3 To whom correspondence should be addressed Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Antson,A.A., Demidkina,T.V., Gollnick,P., Dauter,Z., von Tersch,R.L., Long,J., Berezhnoy,S.N., Phillips,R.S., Harutyunyan,E.H. and Wilson,K.S. (1993) Biochemistry, 32, 4195–4206.[ISI][Medline]

Bradford,M.M. (1976) Anal. Biochem., 72, 248–254.[ISI][Medline]

Hirahara,T., Suzuki,S., Horinouchi,S. and Beppu,T. (1992) Appl. Environ. Microbiol., 58, 2633–2642.[Abstract]

Isupov,M.N., Antson,A.A., Dodson,E.J., Dodson,G.G., Dementieva,I.S., Zakomirdina,L.N., Wilson,K.S., Dauter,Z., Lebedev,A.A. and Harutyunyan,E.H. (1998) J. Mol. Biol., 276, 603–623.[ISI][Medline]

Kamath,A.V. and Yanofsky,C. (1992) J. Biol. Chem., 267, 19978–19985.[Abstract/Free Full Text]

Kunkel,T.A. (1985) Proc. Natl Acad. Sci. USA, 82, 488–492.[Abstract]

Kyte,J. and Doolittle,R.F. (1982) J. Mol. Biol., 157, 105–132.[ISI][Medline]

Laemmli,U.K. (1970) Nature, 227, 680–685.[ISI][Medline]

Newton,W.A. and Snell,E.E. (1964) Proc. Natl Acad. Sci. USA, 51, 382–389.[ISI][Medline]

Sambrook,J., Fritsch,E.F. and Maniatis,T. (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor.

Sanger,F., Nicklen,S. and Coulson,A.R. (1977) Proc. Natl Acad. Sci. USA, 74, 5463–5467.[Abstract]

Smibert,R.M. and Krieg,N.R. (1970) Manual of Methods for General Bacteriology. American Society for Microbiology, Washington, DC.

Snell,E.E. (1975) Adv. Enzymol. Relat. Areas Mol. Biol., 42, 287–333.[ISI][Medline]

Sundararaju,B., Antson,A.A., Phillips,R.S., Demidkina,T.V., Barbolina,M.V., Gollnick,P., Dodson,G.G. and Wilson,K.S. (1997) Biochemistry, 36, 6502–6510.[ISI][Medline]

Suzuki,S., Horinouchi,S. and Beppu,T. (1988) J. Gen. Microbiol., 134, 2353–2362.[ISI]

Terasawa,M., Inui,M., Uchida,Y., Kobayashi,M., Kurusu,Y. and Yukawa,H. (1991) Appl. Microbiol. Biotechnol., 34, 623–627.[ISI][Medline]

Watanabe,T. and Snell,E.E. (1972) Proc. Natl Acad. Sci. USA, 69, 1086–1090.[Abstract]

Wood,W.A., Gunsalus,I.C. and Umbreit,W.W. (1947) J. Biol. Chem., 170, 313–321.[Free Full Text]

Yanisch-Perron,C., Vieira,J. and Messing,J. (1985) Gene, 33, 103–119.[ISI][Medline]

Received April 5, 1999; revised May 18, 1999; accepted May 21, 1999.