A thermostable variant of fructose bisphosphate aldolase constructed by directed evolution also shows increased stability in organic solvents

Jijun Hao and Alan Berry1

School of Biochemistry and Microbiology and the Astbury Centre for Structural Molecular Biology, University of Leeds, Leeds LS2 9JT, UK

1 To whom correspondence should be addressed. E-mail: a.berry{at}leeds.ac.uk


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Thermostable variants of the Class II fructose bisphosphate aldolase have been isolated following four rounds of directed evolution using DNA shuffling of the fda genes from Escherichia coli and Edwardsiella ictaluri. Variants from all four generations of evolution have been purified and characterized. The variants show increased thermostability with no loss of catalytic function at room temperature. The temperature at which 50% of the initial enzyme activity is lost after incubation for 10 min (T50) of the most stable variant, 4-43D6, is increased by 11–12°C over the wild-type enzymes and the half-life of activity at 53°C is increased ~190-fold. In addition, variant 4-43D6 shows increased stability to treatment with organic solvents. DNA sequencing of the evolved variants has identified the mutations which have been introduced and which lead to increased thermostability, and the role of the mutations introduced is discussed.

Keywords: aldolase/directed evolution/solvent stability/thermostability


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Two features of enzymes that may limit their industrial uses are denaturation at elevated temperatures and their need for an aqueous environment for function. Engineering of proteins is therefore crucial to ensure the more widespread use of enzymes and recombinant proteins in biotechnology. Thermal denaturation generally involves two phases. First, at lower temperatures, the equilibrium between the folded state and its unfolded counterpart is tipped towards unfolding, and secondly, at higher temperatures, irreversible aggregation and covalent modifications may occur. Thermophilic organisms have overcome these problems to maintain active conformations of proteins at higher temperatures. However, a complete understanding of the adaptive mechanisms of enzyme thermostability has proved elusive, because these mechanisms are both many and complex since a folded protein is only slightly more stable than its unfolded counterpart and net stability is equivalent to only a few weak interactions out of the hundreds that are present in any given molecule (Arnold, 2001Go). Comparison of the sequences and structures of thermophilic and mesophilic proteins has revealed that Nature has found many routes to avoid thermal denaturation (for reviews see Querol et al., 1996Go; Vogt et al., 1997Go; Jaenicke and Bohm, 1998Go; Vieille and Zeikus, 2001Go). Thermophilic enzymes show marked preferences for some amino acids (Cambillau and Claverie, 2000Go; Fukuchi and Nishikawa, 2001Go); they often possess shorter surface loops than their mesophilic counterparts (Wintrode et al., 2001Go); and disulphide bonds, salt bridges and metal coordination are commonly used to produce thermostability (Querol et al., 1996Go; Vieille and Zeikus, 2001Go). Rational attempts at engineering thermostability have met with some success, either by incorporating stable secondary structure elements into the engineered protein (Villegas et al., 1996Go) or by mimicking the methods used by Nature to produce thermophilic enzymes. For example, the thermostability of cyclodextrin glycosyltransferase was increased by the introduction of a salt bridge (Leemhuis et al., 2004Go), whereas that of triose phosphate isomerase was increased by changing the amino acid composition to avoid the potential for irreversible changes to structure caused by deamidation of asparagines at elevated temperatures (Ahern et al., 1987Go). Despite such successes, rational protein engineering suffers from the drawback of requiring accurate structural data for the enzyme under study and other approaches have been adopted to produce thermostable enzymes. It has been recognized that the consensus sequence obtained from the multiple sequence alignment of homologous proteins shows higher thermostability than the natural proteins and this so-called ‘consensus concept’ has been used to produce new thermostable proteins (Lehmann and Wyss, 2001Go). Finally, the power of directed evolution and high-throughput screening methods have also been successfully used both to increase protein thermostability (Giver et al., 1998Go; Song and Rhee, 2000Go; Wintrode and Arnold, 2000Go) and resistance to organic solvents (Arnold, 1990Go; Song and Rhee, 2001Go).

Our interest lies with the redesign of aldolases for new functions, in particular fructose 1,6-bisphosphate aldolase (FBP-aldolase; EC 4.1.2.13). Aldolases are key enzymes in cellular carbon–carbon bond formation or breakage and they have important potential for use in synthetic organic chemistry because of their exquisite stereochemical control of the reaction (Wong and Whitesides, 1994Go; Fessner, 1998Go). However, the use of aldolases is often restricted by low thermostability and lack of organic solvent tolerance. Typically FBP-aldolases are classified into two classes, I and II, based on their structural and catalytic properties (Rutter et al., 1966Go). Both classes are members of the ({alpha}/ß)8 barrel family (Sygusch et al., 1987Go; Blom et al., 1996Go; Cooper et al., 1996Go) and naturally catalyse the reversible aldol condensation of dihydroxyacetone phosphate (DHAP) and glyceraldehyde-3-phosphate (G3P) to produce fructose 1,6-bisphosphate (FBP). FBP-aldolases have been used in the synthesis of a range of sugar analogues (Bednarski et al., 1986Go; Durrwachter and Wong, 1988; Henderson et al., 1994Go; Chenevert et al., 1997Go; Schoevaart et al., 1999Go; Azema et al., 2000Go), although the higher stability of the bacterial Class II enzyme is an advantage in its use in synthesis (von der Osten et al., 1989Go).

We have already used directed evolution to alter the stereochemistry and substrate specificity of the Escherichia coli Class II FBP-aldolase (Williams et al., 2003Go). Here we report the directed evolution of a Class II FBP-aldolase with high thermostability by family DNA shuffling (Crameri et al., 1998Go) and random mutagenesis. In addition, the evolved variant was also shown to be more tolerant towards organic solvents. The results are interpreted by analysis of mutations, in an attempt to shed light on the relationship between the sequence of the protein and its activity and thermostability.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Materials

Escherichia coli KM3 [{Delta}(his gnd), {Delta}lac, araD, fda, ptsF, ptsM, rpsL, pro, NAR/F' pro A+B+, lacIq, lacZ {Delta}M15] was as reported previously (Berry and Marshall, 1993Go). E.coli XL1-Blue [recA1 endA1 gyrA96 thi-1 hsdR17 supE44 relA1 lac[F' proAB lacIq Z {Delta}M15 Tn10(Tetr)]] was purchased from Stratagene and the expression vector pKK223-3 was from Pharmacia. Edwardsiella ictaluri (NCIMB 13272) was obtained from NCIMB (Aberdeen, UK) and was grown on Oxoid Brain Heart Infusion agar (Oxoid CM375) at 25°C for 48–72 h.

D-Fructose 1,6-bisphosphate, dihydroxyacetone phosphate (DHAP) and nicotinamide adenine dinucleotide (NADH) were purchased from Sigma. Glycerol-3-phosphate dehydrogenase/triose phosphate isomerase and glycerol-3-phosphate dehydrogenase were obtained from Boehringer Mannheim (Mannheim, Germany). The WizardTM Miniprep DNA Purification System, WizardTM Miniprep PCR Purification System, the endonucleases EcoRI and HindIII, T4 DNA ligase and DNase I were supplied by Promega. QiaExII gel extraction kits and QIAquick Gel extraction kits were obtained from QIAgen. Diethylaminoethylcellulose (DE-52) was purchased from Whatman Biosystems (Maidstone, UK). The SuperdexTM 200 prep grade Hi-LoadTM 16/60 column was supplied by Pharmacia (Milton Keynes, UK).

PCR and site-directed mutagenesis

PCR reactions were performed in a Techne Cyclogene DriBlock Cycler using a 50 µl reaction volume consisting of 20 mM Tris–HCl (pH 8.8), 0.1% Triton® X-100, 10 mM KCl, 2 mM MgSO4, 100 mg/ml nuclease-free BSA, 10 mM (NH4)2SO4, 200 µM each dNTP, 100 pmol of each primer, 0.5 µg template DNA and 2.5 units Pfu DNA polymerase. Amplification involved an initial denaturation step at 95°C for 5 min followed by cycling at 95°C for 1 min, 55°C for 1 min, 72°C for 2.5 min for 35 cycles before a final extension at 72°C for 5 min. PCR products were purified from agarose gels using a QIAquick gel extraction kit.

The mutants K71/V210 and I71/A210 were constructed using the variant 4-43D6 plasmid as a template by megaprimer PCR (Reikofski and Tao, 1992Go). External flanking forward and reverse primers were 5'-AGG ACA GAA TTC ATG TCT AAG ATT TTT GAT-3' (EcoRI site underlined) and 5'-GAA AGG AAG CTT TTA CAG AAC GTC GAT CGC-3' (HindIII site underlined). The mutagenic oligonucleotides (mutagenic sites underlined) used in the individual mutagenesis reactions were K71/V210: 5'-TAT ACC GGC CTT GAA GGC AGC-3' and I71/A210: 5'-TTC ACC AAG CTG AGC GCC ATC AGC CCG-3'. The mutant genes were digested with EcoRI and HindIII before the mutant fda gene was ligated into the expression vector pKK223-3 previously treated with the same restriction enzymes. In all cases the resultant plasmids were transformed into the fda-deficient E.coli KM3 strain (Berry and Marshall, 1993Go).

Cloning the fda gene of Ed.ictaluri

Flanking primers used to amplify the full-length fda gene from Ed.ictaluri (Moore et al., 2002Go) were (Ed.ictaluri forward) 5'-AGG ACA GAA TTC ATG TCT AAA ATC TTT GAC-3' (EcoRI restriction site underlined) and (Ed.ictaluri reverse) 5'-GAA AGG AAG CTT TTA CAG CAC GTC GAT ACA-3' (HindIII restriction site underlined). Total genomic DNA from Ed.ictaluri was prepared for use as template by a rapid boiling method (Van Eys et al., 1989Go) and the PCR reaction was conducted as described above, except that the annealing temperature was lowered to 42°C for the Ed.ictaluri reaction. The amplified, full-length products were restricted with EcoRI and HindIII and ligated into the vector pKK223-3 treated with the same enzymes. The resulting plasmid, designated pKeictfda, was transformed into E.coli KM3 cells.

DNA shuffling and thermostability screening

In vitro DNA shuffling was conducted using a slight modification of the protocol described by Stemmer (1994)Go. The fda genes from E.coli and Ed.ictaluri were amplified by PCR as described above using the previously cloned plasmids pKfda2 (Berry and Marshall, 1993Go) and pKeictfda as templates, respectively, in the presence of two external universal primers, pKKfor (5'-CAT CGG CTC GTA TAA TGT G-3') and pKKrev (5'-CTG CGT TCT GAT TTA ATC TG-3'). About 1.5 µg of each fda gene were then pooled and submitted to DNase I digestion in the presence of 1 mM MgCl2. DNA fragments of between 50 and 150 bp were then excised from 0.8% agarose gel and purified. The purified fragments were reassembled in a PCR-like reaction using Taq DNA polymerase in the absence of primers. The reassembly PCR reaction consists of an initial denaturation step at 95°C for 5 min followed by cycling at 95°C for 1 min, 50°C for 1 min, 72°C for 2.5 min for 30 cycles, before a final extension at 72°C for 5 min. The full-length, reassembled library of fda genes was amplified from this mixture in the presence of the pKKfor and pKKrev primers using identical PCR conditions. The products were then gel purified, restricted with EcoRI and HindIII and ligated into pKK223-3, previously restricted with the same enzymes. The ligation mixture (~5 ng DNA) was transformed by electroporation into 50 µl of E.coli XL1-Blue electrocompetent cells using a BioRad electroporator set at 1.8 kV with 0.1 cm gap cuvettes. After growth on 2xTY (10 g/l bactotryptone, 10 g/l yeast extract, 5 g/l NaCl) 1.5% (w/v) agar plates, individual colonies were picked and separately grown in 96-well microtitre plates containing 150 µl of 2xTY medium supplemented with 50 µg/ml ampicillin and 0.3 mM ZnCl2. After growth at 37°C overnight, 100 µl of each culture were transferred to a fresh microtitre plate for processing. The cells were pelleted by centrifugation (20 min at 1100 g at 4°C) and cleared supernatants were prepared by re-suspension of cells in 100 µl of 50 mM Tris–HCl pH 8.0 containing 0.1 M potassium acetate, 1 mg/ml lysozyme and 0.1% (v/v) Triton X-100. The plate was frozen at –80°C and then thawed at room temperature. Cell debris was removed by centrifugation at 1100 g for 20 min at 4°C and the supernatant (50 µl) was transferred to flat-bottomed microtitre plates for assay.

Screening was carried out by incubating the plates for 10 min in a water-bath at the desired screening temperature followed by 20 min of chilling at 4°C. The residual enzyme activity was then determined by the absorbance change at 340 nm in a coupled enzyme assay (Blostein and Rutter, 1963Go). A volume of 50 mM Tris–HCl (pH 8.0) buffer containing 3 mM FBP, 0.6 mM NADH, 0.1 M potassium acetate and 100 mg/ml glycerol-3-phosphate dehydrogenase equal to the volume of lysate was added to each well and A340 was measured over 10 min in a FLUOstar Galaxy plate-reader.

Purification of FBP-aldolases

The wild-type and mutant enzymes were expressed and purified to homogeneity and protein concentration was determined as described previously (Berry and Marshall, 1993Go).

Aldolase coupled assay

A coupled enzymatic assay was used for FBP-aldolase as described (Blostein and Rutter, 1963Go). The assays were performed at 30°C and the decrease in absorbance at 340 nm was recorded. In order to determine steady-state kinetic parameters, data were fitted to the appropriate rate equation using Sigmaplot.

Hydrazine assay of FBP-aldolase

The assay was performed at 30°C in 1 ml of 50 mM Tris–HCl buffer pH 8.0 containing 0.1 M potassium acetate, a suitable aliquot of aldolase, 4 mM fructose 1,6-bisphosphate and 2 mM hydrazine dichloride hydrate. The increase in absorbance at 240 nm (Jagannathan et al., 1956Go) was recorded as the measure of enzyme activity.

Half-lives of the thermal inactivation

Purified FBP-aldolases were incubated at 53°C in a PCR thermocycler for precise temperature control. Aliquots were taken at various time intervals to measure the residual activities using the coupled enzyme assay system described above. The half-lives, t1/2, of the enzyme activity were calculated from plots of log(residual activity) versus time. These plots were linear and the inactivation rate constant, kinact, was obtained from the slope; t1/2 was calculated as ln2/kinact.


    Results and discussion
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Cloning the fda gene from Ed.ictaluri

A number of methods exist for construction of random libraries of variants of proteins for directed evolution experiments, generally based either on error-prone PCR methods (Cadwell and Joyce, 1992Go) or on DNA shuffling methodologies (Stemmer, 1994Go). We chose to use DNA shuffling but, to allow larger areas of sequence space to be explored, family DNA shuffling, in which a number of homologous genes are subjected to DNA shuffling together, was employed (Crameri et al., 1998Go). A search of the protein sequence database using the E.coli Class II FBP-aldolase as target sequence revealed eight homologues with >50% sequence identity. Of these, the Ed.ictaluri protein (Moore and Maranas, 2000Go), with 87% sequence identity with the E.coli aldolase, was chosen for family shuffling. Primers, containing EcoRI and HindIII restriction sites, were designed against the N- and C-termini of the Ed.ictaluri fda gene, the gene was amplified from a genomic preparation from Ed.ictaluri and was cloned into pKK223-3 to yield the expression plasmid pKeictfda. The Ed.ictaluri Class II FBP-aldolase was then expressed and purified in the same manner as the E.coli enzyme (Berry and Marshall, 1993Go). The subunit molecular mass of the Ed.ictaluri enzyme determined by electrospray ionization mass spectrometry was 39 026.0 Da, consistent with the mass calculated from the gene sequence (39 022.3 Da), assuming that the N-terminal methionine had been removed. Analytical ultracentrifugation studies revealed the native molecular mass to be 77 760 ± 1310 Da, consistent with the enzyme being dimeric. Steady-state kinetics of the Ed.ictaluri enzyme showed that, in common with other Class II FBP-aldolases, the enzyme is activated 2- to 3-fold in the presence of potassium ions (data not shown). The kinetic parameters of the purified Edwardsiella and E.coli enzymes are given in Table I.


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Table I. Steady-state kinetics of the wild-type E.coli and Ed.ictaluri Class II FBP-aldolases and the evolved variants in each generation

 
Evolution of thermostable aldolases

Equal amounts of the cloned fda genes from E.coli and Ed. ictaluri were shuffled to generate the first generation library (Crameri et al., 1998Go) and the resulting library of genes was cloned into pKK223-3 and transformed into E.coli strain XL1-Blue. The transformants from the first generation library were picked into 96-well microtitre plates and crude cell extracts were prepared as described in Materials and methods.

The thermostabilities of both the E.coli and Ed.ictaluri wild-type enzymes were measured by assaying the enzyme activity at 30°C after incubation at various temperatures for 10 min. Both enzymes rapidly lost activity at temperatures above 48°C and had no activity after incubation at 55°C (Figure 1). Thermostable FBP-aldolase variants were therefore detected by the retention of activity after incubation of the crude cell extracts at 55°C for 10 min. After screening 5000 colonies, two positive variants, 1-37D6 and 1-44F2, were identified and the improved thermostability was confirmed by measuring their residual activity after incubation at various temperatures. In the same way, equal amounts of both 1-37D6 and 1-44F2 fda genes were used to parent the second round of directed evolution by DNA shuffling to recombine the beneficial mutations from both parents. Approximately 2000 colonies from the second generation library were subjected to screening at the higher temperature of 58°C for 10 min, at which temperature the first generation ‘parents’ are inactive (Figure 1). This resulted in a more thermostable second generation variant, 2-15B2. Verification of the increased thermostability was again carried out and 2-15B2 was then used to parent the third generation of directed evolution by DNA shuffling. Two third-generation FBP-aldolases with increased thermostability, 3-2C6 and 3-4C10, were identified after screening at 60°C. These were used in a final fourth round of directed evolution in which 6400 transformants were screened at 62°C, to yield two fourth-generation variants, 4-4C10 and 4-43D6.



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Fig. 1. Thermal inactivation curves of purified wild-type E.coli, Ed.ictaluri and evolved variant Class II FBP-aldolases. A total of 15 µg of each enzyme was used to measure the residual activities by using the coupled enzyme system after heat treatment for 10 min. Values of activities are the average of triple measurements with standard deviation <7%. The curves are coloured: 4-43D6 in black, 4-4C10 in green, 3-4C10 in blue, 2-15B2 in pink, 1-44F2 in cyan, 1-37-D6 in dark green, the wild-type E.coli Class II FBP-aldolase in yellow and the Ed.ictaluri FBP-aldolase in red.

 
Thermostabilities and kinetic properties of the evolved variants

Expression plasmids were purified from each of the selected variants and were transformed into the fda deficient strain of E.coli, KM3 (Berry and Marshall, 1993Go) and the variants were all expressed and purified to at least 95% homogeneity. The thermostabilities of the purified, evolved variant FBP-aldolases were assessed by measuring the residual activities at 30°C using the coupled enzyme assay (see Materials and methods) after heat treatment of the pure enzyme at various temperatures for 10 min. Figure 1 shows that thermostability of the evolved variants was improved systematically during directed evolution compared with that of the wild-type parental E.coli and Ed.ictaluri Class II FBP-aldolases. The T50 values (the temperature at which 50% of the initial enzyme activity is lost on incubation for 10 min) of each purified variant and the wild-type parents are shown in Table I. The T50 of the variant 4-43D6 was increased more than 11°C compared with that of the wild-type E.coli FBP-aldolase and over 12°C compared with that of the wild-type Ed.ictaluri FBP-aldolase. In addition, the half-lives, t1/2, at 53°C of the evolved 4-43D6 variant and the parent E.coli and Ed.ictaluri FBP-aldolases were determined by measuring the activity remaining at various times of incubation (Figure 2). The t1/2 of the fourth-generation variant 4-43D6 FBP-aldolase was increased ~190-fold (1490 min) compared with that of E.coli Class II FBP-aldolase (7.7 min) and over 360-fold compared with that of the Ed.ictaluri FBP-aldolase (4.1 min).



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Fig. 2. Kinetics of thermal inactivation of 4-43D6 and the parent enzymes at 53°C. The residual activity of variant 4-43D6 (triangles) and the ‘parent’ wild-type E.coli (closed circles) and Ed.ictaluri (open circles) Class II FBP-aldolases were measured at various times after incubation at 53°C using the coupled enzyme system (Blostein and Rutter, 1963Go). All activities are relative to those measured at 30°C. Note: the y-axis is a logarithmic scale.

 
The kinetic parameters of the purified variants were next determined with fructose 1,6-bisphosphate (FBP) as substrate and the results are shown in Table I. In general, the kinetic properties of the evolved enzymes showed little significant change from the wild-type enzymes. Thus, a significant increase in thermostability of the Class II FBP-aldolases has been evolved in four rounds of directed evolution, with no detrimental effect on the kinetic parameters of the enzyme. This result indicates that enzyme thermostability and catalytic activity at low temperatures are not incompatible and can be evolved independently. This notion has been supported by other studies using directed evolution (Giver et al., 1998Go; Song and Rhee, 2000Go; Uchiyama et al., 2000Go) and the low activity of naturally occurring thermostable enzymes at low temperature probably does not reflect physical, but rather evolutionary, constraints (Arnold, 2001Go).

Tolerance to organic solvents

It has been observed previously that an increase in the thermostability of an enzyme can be accompanied by increased resistance to inactivation by organic solvents (Cowan, 1997Go). To test if the evolved thermostable FBP-aldolase exhibited higher tolerance to organic solvents, the activity of the variant 4-43D6 was measured using a hydrazine-based assay (Jagannathan et al., 1956Go) with FBP as substrate, after incubation of the enzyme in the presence of 20% (v/v) of a range of different organic solvents. Three polar organic solvents [methanol, acetonitrile and N,N-dimethyformamide (DMF)] and three non-polar organic solvents (toluene, chloroform and hexane) were chosen for analysis. For the non-miscible mixtures, the mixture was shaken during the incubation and the enzyme in the aqueous phase was assayed after the incubation. Two different assays were carried out: (i) measurement of the enzyme activity directly in the presence of the organic solvent (Figure 3A) and (ii) residual enzyme activity was measured in an essentially aqueous system (50 mM Tris–HCl pH 8.0 containing 0.1 M potassium acetate) after the 24 h of incubation of enzyme in the presence of 20% (v/v) organic solvent to assess the irreversible inactivation caused by the organic solvent (Figure 3B). Upon assay in the organic solvent mix, the evolved variant 4-43D6 showed significantly more activity than the wild-type enzymes in every solvent system tested, although the level of activity in DMF was very low for all variants. In addition, when the treated enzyme was recovered into purely aqueous media, the variant regained almost 100% of the initial activity whereas the wild-type enzymes permanently lost between 40 and 90% of their initial activity.



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Fig. 3. Activity of FBP-aldolases after 24 h of incubation in 20% (v/v) organic solvents. (A) Enzyme was incubated in the presence of 20% (v/v) organic solvent and activity was then measured in a 20% (v/v) mixture of that solvent with 50 mM Tris–HCl buffer pH 8.0 containing 0.1 M potassium acetate (for water-immiscible solvents saturated buffer was used). (B) Activity measured in 50 mM Tris–HCl buffer pH 8.0 containing 0.1 M potassium acetate in the absence of the organic solvent. All the activities were measured in triplicate using a hydrazine-based assay (Jagannathan et al., 1956Go; Budde and Khmelnitsky, 1999Go) and are relative to that of each enzyme in water at 0 h. White bars, Ed.ictaluri; grey bars, E.coli; black bars, 4-43D6.

 
Sequence and structural analysis

In order to understand how the amino acid changes produced during the directed evolution have impacted on the thermostability, the DNA of the four generations of variants was sequenced. This showed that the genes encoding the first generation variant FBP-aldolases, 1-37D6 and 1-44F2, were both chimeras of the ‘parental’ E.coli and Ed.ictaluri fda genes and that both variants contained three crossovers in the gene sequences (Figures 4 and 5). Comparison of the wild-type parental protein sequences of the E.coli and Ed.ictaluri Class II FBP-aldolases reveals that the sequences are 87% identical but that the Ed.ictaluri protein contains a single residue deletion at position 82 of the E.coli protein. This deletion is carried into all of the evolved variant enzymes (Figure 4). Thus, for comparison, identical residues in the variant and wild-type E.coli enzymes at sequence positions above 82 will differ by one residue number; for example, residue His-110 in the E.coli sequence is aligned with His-109 in the variant sequences.



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Fig. 4. Protein sequence alignment of the selected thermostable variants. The amino acid sequences of the wild-type E.coli (yellow) and Ed.ictaluri (red) aldolases and the selected thermostable variants are aligned. The DNA sequences of the first generation variants were used to determine which of the parental genes encoded which section of the protein and the amino acid sequences of the first generation variants are colour coded accordingly. Later generations were derived from the first generation. Point mutations introduced during the shuffling procedure are highlighted in cyan. All numbering is relative to the wild-type E.coli sequence.

 


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Fig. 5. Location of mutations mapped onto the structure of the E.coli Class II FBP-aldolase. (A) The colour-coding scheme of Figure 4 is used to map the location of sections of the polypeptide of the first generation variant 1-44F2 derived from the E.coli (yellow) and Ed.ictaluri (red) parents on to the structure of the E.coli enzyme [PDB file 1B57.PDB (Hall et al., 1999Go)]. Point mutations are shown in cyan and the catalytic zinc ion in white. (B) The structure of the Class II FBP-aldolase to illustrate the variations between the wild-type parental enzymes and variant 1-44F2. Residues common to both parents and variant 1-44F2 are coloured white, residues common only between the E.coli and the variant are coloured yellow, those common only between the Ed.ictaluri and variant are coloured red and point mutations not found in either parent are coloured cyan.

 
Variant 1-37D6 contained one point mutation which altered Pro9 to a leucine residue (P9L) and another which altered a residue encoded by a section of the Edwardsiella gene into the residue normally found in the E.coli protein (lysine at position 19). Overall, 1-37D6 is 90.8% identical with the E.coli protein sequence and 96.1% identical with the Ed.ictaluri sequence. Similarly, 1-44F2 contained new point mutations found in neither ‘parent’—Q79L and Q321 M (K321 in the Ed.ictaluri enzyme). 1-44F2 is 90.2% identical with the E.coli enzyme and 96.4% identical with the Ed.ictaluri enzyme. It is interesting that the evolved, thermostable first generation variants are more similar to the Ed.ictaluri enzyme, the ‘parent’ enzyme of lower thermostability. The regions of the polypeptide encoded by the two parent genes are shown in Figure 5A. In both 1-44F2 and 1-37D6, helix {alpha}11, part of helix {alpha}5, strand ß4, helix {alpha}6 and strand ß5 are all contributed by sections of the E.coli gene, whereas in variant 1-37D6 the rest of {alpha}5, strand ß3 and half of {alpha}4 are also from E.coli. However, given the high sequence identity between the two parent proteins, the distribution of differences between the parents and the first generation variants is fairly evenly spread throughout the structure (Figure 5B) but with a slight ‘hot spot’ of changes located in helix {alpha}11. This may be significant since helix {alpha}11 interacts with helix {alpha}10 of the opposite subunit and together helices {alpha}10 and {alpha}11 form 921Å2 (33%) of the intersubunit surface area. Mutations in {alpha}11 might therefore subtly alter the packing of these helices and affect the temperature at which an active enzyme conformation is lost on heating.

The relatively large number of amino acid differences between the wild-type enzymes and the first generation variants (e.g. 1–44F2 has 35 mutations compared with the E.coli enzyme) makes it difficult to assess the individual contributions of each mutation to the thermostability of each protein. However, two factors can be ruled out. There is no change in the overall length of the encoded protein, so major changes to loop lengths can be ruled out, although minor changes to the lengths of individual secondary structural elements could occur. Similarly, no cysteine residue has been introduced and the formation of new disulphide bonds in the variant proteins is therefore impossible.

After the first generation, analysis becomes more straightforward as only one or two new point mutations are found in each generation and of particular interest are those which are conserved through subsequent rounds of directed evolution. A comparison of the mutations found in all of the later rounds of directed evolution is given in Table II, along with the location of these residues within the secondary structure elements of the enzyme, the surface accessibility of the residue in the E.coli enzyme dimer [1B57.PDB (Hall et al., 1999Go) calculated using the program DSSP (Kabsch and Sander, 1983Go)] and the surface area of the residue buried during dimer formation.


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Table II. Summary of the point mutations introduced during rounds two to four of directed evolution of the thermostable Class II FBP-aldolases

 
The fourth generation variant 4-43D6 has seven mutations compared with the first generation 1-44F2 (V45A, K71I, A111T, A124V, A210V, M321K and K346R) (Table II) and these must be responsible for the 9.5°C increase in the T50 value between 1-44F2 and 4-43D6. Structures are not yet available for the variant enzymes, but the variation in amino acid sequence observed between these variants is mapped on to the structure of the wild-type E.coli enzyme (Figure 6). This shows that, of the seven positions, four lie in {alpha}-helices and the remaining three lie in the loops. No mutation was found in any of the ß-strands in any generation of the current study, perhaps reflecting the importance of the ß-strands in catalysis in (ß/{alpha})8 barrels. All of the mutations are at least 11 Å from the nearest active site zinc atom and five of the seven residues are significantly exposed on the surface of the enzyme dimer. Only one position, 71, is directly involved in the dimer interface. A similar distribution of mutations across the whole protein has also been observed in the directed evolution of a thermostable Bacillus subtilis subtilisin E mutant and is thought partly to reflect the fact that surface mutations are more easily accommodated in the protein structure than buried mutations, which are unlikely to increase stability without compensating changes elsewhere (Zhao and Arnold, 1999Go).



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Fig. 6. Location of point mutations in variant 4-43D6. The seven point mutations introduced between the first and fourth generation variants 1-44F2 and 4-43D6 are mapped on to the structure of the E.coli Class II FBP-aldolase [PDB file 1B57.PDB (Hall et al., 1999Go)]. The two subunits of the enzyme dimer are coloured orange and green.

 
Site-directed mutagenesis

In order to investigate further the role of some of these substitutions in the evolution of thermostability, we targeted residues 71 and 210 since the two fourth generation variants, 4-43D6 and 4-4C10, differ only at these two positions and yet show a difference of ~2°C in their T50 values (Table I). Variant 4-43D6 has isoleucine and valine at positions 71 and 210, respectively; whereas 4-4C10 has the same residues as the wild-type Ed.ictaluri enzyme, namely lysine and alanine (Table II). To assess the role of each of these substitutions in thermostability, two mutants were constructed by site-directed mutagenesis whereby Ile71 and Val210 in variant 4-43D6 were separately mutated to Lys71 and Ala210. These mutants were designated Lys71/Val210 and Ile71/Ala210 and the two mutant aldolases were characterized in terms of their steady-state kinetics and thermostabilities (Table III). The mutations described have no significant effect on the Km of the enzyme for fructose bisphosphate, but a lysine a position 71 in the chimeric enzyme variants has a slightly detrimental effect on the kcat of the reaction (about 0.5-fold) compared with either of the two wild-type parental enzymes. It is clear that this mutation has an extremely subtle effect on the structure and function of the enzyme, since the wild-type Ed.ictaluri enzyme also possesses lysine at position 71 and yet maintains the higher level of kcat for FBP.


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Table III. Kinetic parameters of site-directed mutants and evolved variants of the Class II FBP-aldolase

 
The thermostabilities of variants 4-4C10, 4-43D6 and the Lys71/Val210 and Ile71/Ala210 site-directed mutants show an interesting progression. Variant 4-43D6 and the I71/A210 site-directed mutant differ only at position 210 (where 4-43D6 contains valine) and these enzymes show the same thermostability (Table III), seeming to suggest that the change of alanine to valine at position 210 has no role on the thermostability of the enzyme. However, a comparison of variant 4-4C10 with site-directed mutant K71/V210, which again only differ by the residues present at position 210 (valine in K71/V210 and alanine in 4-4C10) shows a different picture. Here, the replacement of the alanine in 4-4C10 results in a drop in T50 of almost 9°C. Valine at residue 210 thus appears detrimental to the thermostability of the enzyme, but only if residue 71 is lysine rather than isoleucine. Mutations at positions 71 and 210 of the Class II FBP-aldolases (which are ~40 Å apart) therefore do not appear to fit in with the generally well-established principle that mutations causing an increase in thermostability act independently and, to a large extent, additively (Jaenicke et al., 1996Go; Giver et al., 1998Go; Zhao and Arnold, 1999Go).

Mechanism of stabilization and comparison with thermophilic aldolases

The recently solved crystal structure of the naturally thermophilic Class II FBP-aldolase from Thermus aquaticus (Izard and Sygusch, 2004Go) shows that, in line with general considerations of mechanisms of thermostabilization, the presence of additional salt bridges and hydrogen bonds is a major contributing factor (Querol et al., 1996Go; Xirodimas and Lane, 1999Go). In particular, an intricate network of intramolecular electrostatic interactions exists between {alpha}-helices {alpha}1 and {alpha}8 in the T.aquaticus enzyme (equivalent to {alpha}2 and {alpha}11 in the E.coli enzyme nomenclature). In the E.coli enzyme, this network is replaced by two electrostatic interactions between Lys51 and Glu346 and Glu47 and Arg335. The T.aquaticus enzyme structure also has shorter loops than the corresponding E.coli enzyme. For example, the loops formed by residues 70–79 and 127–137 in the E.coli protein are reduced to only three residues or are missing entirely in the thermophilic enzyme. Finally, the T.aquaticus enzyme also contains an extra 17 residues forming an {alpha}-helix ({alpha}7a) and loop inserted between strand ß7 and helix {alpha}9 of the E.coli enzyme. The large number of hydrophobic amino acids from this extra helix and loop pack against the helices equivalent to {alpha}10 and {alpha}11 in the E.coli enzyme and the enhanced interactions with this pair of helices is thought to stabilize both the enzyme subunit and the dimer interface, resulting in thermostability (Izard and Sygusch, 2004Go). While direct comparison of the mechanism of thermostabilization between the naturally thermostable T.aquaticus enzyme and the evolved variants is complicated by the low sequence identity (27%) between the T.aquaticus and E.coli enzymes, it is, nevertheless, interesting that similar regions of the protein have been found to be mutated during our directed evolution. So, for example, two of the sequence variations introduced (V45A and K346R) lie in helices 2 and 11, three of the mutated positions (K71I, Q79L and A124V) lie in the region of the shorter loops and two (residues 321 and 346) are found in the {alpha}10–{alpha}11 region. Residue 321 lies very near the ‘tip’ of the loop between helices 10 and 11 (Figure 6) and it may be that the stabilizing effect noted here between variants 1-44F2 and subsequent generations is the removal of the previously introduced methionine residue (Table II) since methionine residues can undergo thermally induced modifications which can lead to aggregation, denaturation and/or loss of enzyme activity (Russell et al., 1997Go). Residue 346, lysine in the E.coli enzyme, lies in helix {alpha}-11 and is mutated to arginine in variants 3-4C10 and both fourth-generation variants. The mutation of lysine to arginine is often found in thermophilic proteins compared with their related homologous mesophilic proteins (Menendez-Arias and Argos, 1989Go) and this mutation is often used to generate thermostable proteins (Mrabet et al., 1992Go) since the longer side chain of Arg can interact with more distant amino acids without the requirement for a bridging solvent molecule and without affecting overall charge and the resonance capacity of the guanidinium moiety favours formation of salt bridges and hydrogen bonding with more than one residue. Such changes would therefore stabilize this important region of the dimer interface and could account, in part, for the increased thermostability of variant 4-43D6.

Taken as a whole, our directed evolution experiments present a novel solution to the problem of providing a thermostable aldolase, with combinations of mutations providing small improvements at many positions including regions of the protein known to be important for thermostability in the naturally occurring thermostable enzymes as well as in residue types and positions not previously found in natural evolution.


    Acknowledgments
 
We thank Ms Diana Hall, School of Biochemistry and Microbiology, for help with the growth of Ed.ictaluri and Gavin Williams and Sheena Radford for useful discussions. This work was supported by the BBSRC, EPSRC and The Wellcome Trust and is a contribution from The Astbury Centre for Structural Molecular Biology at the University of Leeds.


    References
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
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Received August 23, 2004; revised October 12, 2004; accepted October 13, 2004.

Edited by Dek Woolfson





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