Directed evolution of Thermotoga neapolitana xylose isomerase: high activity on glucose at low temperature and low pH

Dinlaka Sriprapundh1,2,3, Claire Vieille2 and J.Gregory Zeikus2,4

1Department of Food Science and Human Nutrition and 2Department of Biochemistry and Molecular Biology, Michigan State University, 410 Biochemistry Building, East Lansing, MI 48824, USA 3Present address: Department of Pharmaceutical Chemistry and Biochemistry/Biophysics, University of California, San Francisco, CA 94143, USA

4 To whom correspondence should be addressed. e-mail: zeikus{at}msu.edu


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The Thermotoga neapolitana xylose isomerase (TNXI) is extremely thermostable and optimally active at 95°C. Its derivative, TNXI Val185Thr (V185T), is the most active type II xylose isomerase reported, with a catalytic efficiency of 25.1 s–1 mM–1 toward glucose at 80°C (pH 7.0). To further optimize TNXI’s potential industrial utility, two rounds of random mutagenesis and low temperature/low pH activity screening were performed using the TNXI V185T-encoding gene as the template. Two highly active mutants were obtained, 3A2 (V185T/L282P) and 1F1 (V185T/L282P/F186S). 1F1 was more active than 3A2, which in turn was more active than TNXI V185T at all temperatures and pH values tested. 3A2 and 1F1’s high activities at low temperatures were due to significantly lower activation energies (57 and 44 kJ/mol, respectively) than that of TNXI and V185T (87 kJ/mol). Mutation L282P introduced a kink in helix {alpha}7 of 3A2’s ({alpha}/ß)8 barrel. Surprisingly, this mutation kinetically destabilized 3A2 only at pH 5.5. 1F1 displayed kinetic stability slightly above that of TNXI V185T. In 1F1, mutation F186S creates a cavity that disrupts a four-residue network of aromatic interactions. How the conformation of the neighboring residues is affected by this cavity and how these conformational changes increase 1F1’s stability still remain unclear.

Keywords: directed evolution/random mutagenesis/xylose isomerase


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Xylose isomerase (XI) (EC 5.3.1.5) is an intracellular enzyme found in bacteria that can utilize xylose as a carbon substrate for growth (Chen, 1980Go). As a result of its ability to use glucose as substrate and convert it to fructose, XI is often referred to as glucose isomerase, and it is widely used in the industrial production of high fructose corn syrup (Meng et al., 1993Go; Bhosale et al., 1996Go). XI is one of the three highest tonnage value enzymes, amylases and proteases being the other two (Bhosale et al., 1996Go). Most industrially used XIs are isolated from mesophilic organisms (e.g. Streptomyces spp. and Actinoplanes spp.). The reaction temperature used in the current industrial glucose isomerization process is limited to 60°C because of by-product and color formation that occur at high temperature and alkaline pH, and because the isomerases themselves are not highly thermostable (Lee and Zeikus, 1991Go; Vieille and Zeikus, 2001Go). Thermostable XIs with neutral or slightly acidic pH optima have a potential for industrial applications. Performing isomerization at higher temperature and neutral or slightly acidic pH with thermo-acid stable XIs would allow for faster reaction rates, higher fructose concentrations at equilibrium, higher process stability, decreased viscosity of substrate and product streams, and reduced by-product formation (Lee and Zeikus, 1991Go).

The XI from the hyperthermophile Thermotoga neapolitana (TNXI) has been studied extensively in our laboratory. The gene encoding TNXI (xylA) was cloned, sequenced and over-expressed in Escherichia coli (Vieille et al., 1995Go). TNXI’s active site was engineered by site-directed mutagenesis to increase its activity on glucose (Sriprapundh et al., 2000Go). The TNXI Val185Thr (V185T) mutant derivative is more active, more glucose-efficient, and as stable as the wild-type TNXI. It is also the most active type II XI ever reported. Although TNXI V185T is highly thermostable and highly active at 97°C, it is very poorly active (10% of maximal activity) at the current industrial isomerization temperature (60°C) and it requires a neutral pH for optimal activity.

Rules for engineering protein activity and stability by rational design are likely to be protein-specific, and any such design effort would require prior detailed structural information. Numerous and intensive site-directed mutagenesis studies have probed this issue. Despite these efforts, considerable disagreement remains over which forces dominate stabilization mechanisms, and no generally applicable rules have been established (Giver et al., 1998Go; Vieille and Zeikus, 2001Go). Although protein chemists continue to study the relationships between the sequence, structure and function of proteins, the extensive knowledge that is necessary for the application of rational engineering approaches is available for only a tiny fraction of known enzymes. Directed evolution, on the other hand, has proved to be useful for modifying enzymes in the absence of such knowledge (Kuchner and Arnold, 1997Go). In directed evolution, the process of natural evolution is accelerated in a test tube for selecting proteins with the desired properties (Moore and Maranas, 2000Go). A typical experimental cycle of directed evolution begins with the creation of a library of mutated genes. Among the methods that introduce mutations randomly along the entire length of a gene (Leung et al., 1989Go; Stemmer, 1994Go; Shao et al., 1998Go; Zhao et al., 1998Go; Ostermeier et al., 1999Go; Zhao and Arnold, 1999Go), error-prone PCR has been used the most extensively. The mutated genes are then ligated into an expression vector and transformed into suitable bacterial cells. A screening procedure is next employed to identify the few transformants expressing proteins/enzymes with improved properties. Random mutagenesis and screening are repeated several times depending on the extent to which the properties of the protein should be altered and on the effects of mutations observed in each generation. Interest in engineering enzymes using directed evolution has grown significantly in the past few years. It has been used to increase enzyme thermostability, activity on novel substrates, substrate specificity and enantioselectivity. For example, six generations of random mutagenesis, recombination and screening stabilized Bacillus subtilis p-nitrobenzyl esterase significantly (>14°C increase in Tm) without compromising its catalytic activity at lower temperatures (Giver et al., 1998Go).

Here we use the TNXI V185T-encoding gene as the template for directed evolution to develop an enzyme active at 60°C and acidic pH. We show that activity can be increased significantly at low temperature and acidic pH without cost to the enzyme thermal stability.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Random mutagenesis

Random mutations were introduced into the TNXI V185T-encoding gene cloned between the NdeI and HindIII restriction sites of pET23a(+) (Novagen, Madison, WI). PCR was performed with primers 5'-CGACTCACTATAGGGAGAC-3' and 5'-GGTGGTGCTCGAGTGCG-3' encoding sequences upstream of the NdeI site and downstream of the HindIII site, respectively, in pET23a(+). The reaction mixture contained 100 ng of plasmid DNA, 50 pmol of each primer, 50 mM KCl, 10 mM Tris-HCl (pH 8.3), 1.5 mM MgCl2, 0.4 mM dCTP, 0.4 mM dTTP, 0.08 mM dATP, 0.08 mM dGTP and 2.5 U Taq DNA polymerase (Hoffmann-LaRoche, Nutley, NJ) in a 50 µl reaction volume. Cycling parameters were 36 cycles of 95°C for 45 s, 50°C for 45 s and 72°C for 3 min. Amplification of the 1.4 kb product was checked by running a small aliquot of the reaction on a 1% agarose gel. The PCR product was purified using a Geneclean III kit (Bio101, Carlsbad, CA) and cloned back into the NdeI and HindIII sites of pET23a(+) using standard molecular biological techniques (Ausubel et al., 1993Go). For the second round of random mutagenesis, the gene encoding TNXI 3A2 was used as the template.

Construction of a mutant library

The plasmids resulting from random mutagenesis were transformed into electrocompetent E.coli HB101(DE3) cells (XI-deficient) created using the {lambda}DE3 lysogenation kit (Novagen, Madison, WI). Transformants were plated on Luria–Bertani (LB) agar containing 100 µg/ml ampicillin. After 16 h of growth, single colonies were picked with sterile toothpicks and transferred into 24-well plates, each well containing 2 ml of LB plus 100 µg/ml ampicillin. Plates were then incubated overnight at 37°C on a shaker at 175 r.p.m. to allow for cell growth. One hundred and fifty microliters of each culture were transferred to sterile 96-well plates. These plates were used to quantify bacterial growth by reading the bacterial suspensions’ OD595 in a microplate reader (Dynatech, McLean, VA), before being stored at 4°C to save the original cultures. The rest of the cultures were pelleted by centrifugation at 1000 g for 10 min and resuspended in 200 µl of 50 mM MOPS (pH 7.0) containing 5 mM MgSO4 and 0.5 mM CoCl2 (i.e. buffer A). Bacterial suspensions were incubated with 50 µl of a 1% lysozyme solution at 37°C for 1 h before being subjected to three freeze–thaw cycles (5 min in a dry ice–ethanol bath and 5 min in a 50°C water bath) to break the cells and release the enzymes into the supernatant. Cell-free crude extracts were then obtained by centrifugation at 1000 g for 10 min and stored at 4°C for further use.

Screening the mutant library for increased activity on glucose at 60°C and low pH

The crude extracts were assayed for glucose isomerization in two conditions: 60°C (pH 7.0) and 80°C (pH 5.2). Assays were performed in microtiter plates with 150 µl of 100 mM MOPS (pH 7.0) or 100 mM sodium acetate (pH 5.2) containing 1 mM CoCl2, 0.4 M glucose and 10 µl of crude extract. The plates were incubated at 60°C (pH 7.0) or 80°C (pH 5.2) for 10 min and placed on ice to stop the reactions. The fructose produced was assayed using the resorcinol–ferric ammonium sulfate–hydrochloric acid method (Schenk and Bisswanger, 1998Go). Ten microliters of each reaction were transferred to a new set of microtiter plates and mixed with 40 µl of distilled water and 150 µl of a freshly prepared 1:1 (v/v) mixture of solution A (0.05% resorcinol in ethanol) and solution B [0.216 g of FeNH4(SO4)2.12H2O in 1 l of HCl]. The plates were incubated in an 80°C water bath for 30 min to develop the color. The OD490 was measured with a microplate reader (Dynatech) with 0–2.5 mM fructose as standards. A crude extract of HB101(DE3)pET23a(+) was used as the negative control on each plate. Crude extracts of HB101(DE3) expressing TNXI V185T and TNXI 3A2 were the positive controls in screening rounds one and two, respectively. Mutants with potentially higher activity on glucose than the positive control were selected on the basis of increases in both OD490 and OD490/OD595 relative to the positive controls in the two rounds of mutagenesis. Mutants showing increased activity were screened a second time using crude extracts prepared from 5 ml cultures. These crude extracts were prepared as described above, before being heat-treated at 80°C for 15 min and centrifuged.

Oligonucleotide synthesis and DNA sequencing

PCR primers were synthesized by the Macromolecular Structure Facility, Department of Biochemistry and Molecular Biology at Michigan State University (MSU). DNA sequences were determined either manually using the Thermosequenase kit (USB, Cleveland, OH) or automatically at the MSU Genomics Technology Support Facility.

Protein purification

Recombinant enzymes were purified as described (Vieille et al., 1995Go), followed by an additional ion-exchange chromatography step. Partially purified enzymes were applied to a DEAE–Sepharose column (2.5x15 cm) equilibrated with buffer A, and enzymes were eluted using a 500 ml linear 0–300 mM NaCl gradient in buffer A. The pooled fractions from the DEAE–Sepharose column were concentrated in a stirred ultrafiltration cell (30 kDa MW cut-off) (Amicon, Beverly, MA) and dialyzed twice against buffer A. Concentrated, homogeneous enzymes were dispensed and stored frozen at –70°C.

Glucose isomerase assays

TNXI and its mutants were assayed routinely with glucose as the substrate. The enzyme (1–1.5 mg/ml) was incubated in 100 mM MOPS (pH 7.0) [or 100 mM sodium acetate (pH 5.5)] containing 1 mM CoCl2 and 0.4 M glucose at 80°C for 10 min. The reaction was stopped by transferring the tube to an ice bath. The amount of fructose produced was determined by the resorcinol–ferric ammonium sulfate–hydrochloric acid method (Schenk and Bisswanger, 1998Go). To determine the effect of temperature on activity, the enzymes were incubated in the reaction mixture at the temperatures of interest in a heated water (45–95°C) or oil bath (95–110°C) for 10 min. The effect of pH on activity was determined using the routine assay described above except that the MOPS buffer was substituted with 100 mM sodium acetate (pH 4.3–5.8), 100 mM PIPES (pH 6.1–7.0) or 100 mM EPPS (N-[2-hydroxyethyl]piperazine-N'-[3'-propanesulfonic acid]) (pH 7.2–8.1). All pH values were adjusted at room temperature, and the {Delta}pKa/{Delta}T values for acetate, PIPES and EPPS (0, –0.0085 and –0.011 K–1, respectively) (USB) were taken into account for the results. To determine the kinetic parameters, assays were performed in 50 mM MOPS (pH 7.0) containing 10–1500 mM glucose and 1 mM CoCl2. One unit of glucose isomerase activity is defined as the amount of enzyme that produces 1 µmol of fructose per minute under the assay conditions.

Thermal inactivation assays

To obtain the apo-enzymes (metal-free enzymes), the purified enzymes were incubated overnight at 4°C in 50 mM MOPS (pH 7.0) containing 10 mM EDTA. They were then dialyzed twice against 50 mM MOPS (pH 7.0) containing 2 mM EDTA, and they were finally dialyzed twice against 50 mM MOPS (pH 7.0) without EDTA. CoCl2 (0.5 mM) was added to the apo-enzymes and equilibrated at 4°C overnight before thermoinactivation assays. The time course of irreversible thermoinactivation was measured by incubating the enzymes (0.1–0.2 mg/ml) in either 10 mM MOPS (pH 7.0) or 10 mM sodium acetate (pH 5.5) at various temperatures for different periods of time in a heated water bath. Residual glucose isomerase activity was measured at 80°C as described above. Non-linear curve fitting of the inactivation data was performed using the {chi}2 minimization procedure of the software Origin (Microcal Software, Northampton, MA).

Analysis of three-dimensional structures of TNXI and its variants

Enzymes were visualized on an IRIS-4D25 computer (Silicon Graphics Computer System, Mountain View, CA) using the InsightII graphics program (Biosym Technologies, San Diego, CA). The TNXI pdb file (1A0E) was obtained from the Protein Data Bank (www.rcsb.org/pdb).


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Construction of mutant TNXI libraries and screening for activity on glucose at low temperature and low pH

At pH 7.0, TNXI V185T is optimally active at 95–97°C, but its activity at 60°C does not exceed 10% of its optimal activity (Sriprapundh et al., 2000Go). It retains only 20% of its optimal activity at pH 5.2. To increase this enzyme’s activity at 60°C and at acidic pH, and to gain insight into the factors determining the effects of temperature and pH on activity, we subjected the TNXI V185T-encoding gene to sequential random mutagenesis and to low-temperature/low-pH activity screening. Random mutations were introduced into the gene by error-prone PCR. The PCR conditions used were suggested to yield an average of one to two mutations per gene, conditions deemed optimal for the improvement of specific properties by mutagenesis and screening (Arnold and Moore, 1997Go). After the first round of random mutagenesis, 1000 transformants were screened for their activity on glucose at low temperature (60°C, pH 7.0) and at low pH (pH 5.2, 80°C). Thirty mutants were identified that showed significantly higher activity (>30% increase) than TNXI V185T in both screening conditions. The phenotype of these mutants was tested again with heat-treated crude extracts prepared from 5 ml cultures. Higher activity on glucose was confirmed in only 11 out of the 30 crude extracts. The XI expression level in these 11 crude extracts was checked by SDS–PAGE. Ten crude extracts showed higher levels of XI expression than the TNXI V185T control (data not shown). These 10 mutants were discarded. The remaining mutant, 3A2, was purified to homogeneity. Once it was verified that 3A2 was significantly more active than TNXI V185T at 60°C and at pH 5.2, the gene encoding 3A2 was used as the template in a second round of error-prone PCR and activity screening at low temperature and low pH. A library of ~1500 transformants was screened using 3A2 as the positive control. A single mutant, 1F1, showed 80 and 40% increases in activity on glucose at 80°C (pH 5.2) and 60°C (pH 7.0), respectively, based on assays with heat-treated crude extracts. 3A2 and 1F1 were then purified to homogeneity. Their catalytic properties were studied as a function of temperature and pH, and their thermostability was determined.

Effects of temperature and pH on TNXI 3A2 and 1F1 activities

The effect of temperature on 3A2 and 1F1 glucose isomerase activities is shown in Figure 1A in comparison with the activities of TNXI and TNXI V185T. Both 3A2 and 1F1 show significantly higher specific activity on glucose than TNXI and TNXI V185T do at all temperatures. At their optimal temperatures of activity (i.e. 90°C for 1F1 and 95°C for 3A2), both mutants are ~3-fold more active than TNXI V185T. Activation energies (Ea) for activity on glucose were calculated from the linear regressions shown in Figure 1B, using the equation A = A0eEa/RT. Whereas TNXI V185T shows the same activation energy as TNXI (i.e. 87 kJ/mol), 3A2 and 1F1 show significantly decreased Ea values (57 and 44 kJ/mol, respectively). These lower Ea values explain why 3A2 and 1F1 are as much as 7.3 and 12.3 times more active, respectively, than TNXI at 60°C, but only 4.2 and 4.8 times more active, respectively, than TNXI at 90°C.



View larger version (25K):
[in this window]
[in a new window]
 
Fig. 1. Effect of temperature on the specific activities of TNXI and its mutant derivatives on glucose at pH 7.0. TNXI (squares); TNXI V185T (diamonds); 3A2 (circles); 1F1 (triangles). (A) Specific activity versus temperature. (B) Ln(specific activity) versus 1/temperature. All linear regressions had r2 values above 0.97.

 
The effect of pH on the activities of TNXI and its mutant derivatives is shown in Figure 2. 3A2 and 1F1 show significantly increased specific activity on glucose compared with TNXI and TNXI V185T over the entire active pH range. The activity increase is so significant that 3A2 and 1F1 are more active at pH 5.5 than TNXI and TNXI V185T are at pH 7.0.



View larger version (19K):
[in this window]
[in a new window]
 
Fig. 2. Effect of pH on the specific activities of TNXI and its mutant derivatives on glucose at 80°C. Symbols are the same as in Figure 1.

 
Kinetic parameters of TNXI 3A2 and 1F1

The kinetic parameters on glucose of TNXI V185T, 3A2 and 1F1 were compared in different conditions of temperature and pH (Table I). In all conditions tested, 3A2 and 1F1 showed higher Km and Vmax values than TNXI V185T did. At pH 7.0 (both at 60 and 80°C), 3A2 and 1F1’s Vmax values increased more significantly than their Km values for glucose, yielding important increases in catalytic efficiencies [up to 2.3-fold for 1F1 at 60°C (pH 7.0)]. At 80°C (pH 5.5), the increases in 3A2 and 1F1’s Vmax values did not compensate for the major increases in their Km values for glucose (i.e. 3.0-fold for 3A2 and 4.6-fold for 1F1). In these conditions, 3A2 and 1F1 showed catalytic efficiencies that were approximately half that of TNXI V185T. At 60°C (pH 5.5), TNXI 3A2’s increase in Vmax did not compensate for a poor glucose affinity (high Km), resulting in a lower catalytic efficiency than that of TNXI V185T. Unlike 3A2, 1F1 had a higher catalytic efficiency on glucose than TNXI V185T did due to a dramatic increase (5-fold) in its Vmax that surpassed the increase in its Km (3.7-fold) in these conditions. Its 5-fold increase in Vmax makes 1F1 a 1.7-fold more active enzyme at 60°C (pH 5.5) than TNXI V185T is at 80°C (pH 7.0).


View this table:
[in this window]
[in a new window]
 
Table I. Kinetic parameters of TNXI and its mutant derivatives on glucosea
 
Thermal stability of 3A2 and 1F1

To determine whether the mutations present in 3A2 and 1F1 affected the kinetic stability of the mutated enzymes, the relative residual activities of 3A2 and 1F1 were compared with those of TNXI and TNXI V185T after heat treatment at 80°C (pH 7.0) and 80°C (pH 5.5) for various lengths of time (Figure 3). Stability experiments were performed with the EDTA-treated enzymes in 10 mM MOPS (pH 7.0) containing 0.5 mM CoCl2. All inactivation data were best fit by a two-exponential-terms equation of the form



View larger version (23K):
[in this window]
[in a new window]
 
Fig. 3. Inactivation curves of TNXI and its mutant derivatives at (A) 80°C (pH 7.0) and (B) 80°C (pH 5.5). Symbols are the same as in Figure 1. Non-linear least-squares fit regressions were of the form y = C1ek1t + C2ek2t + C3. All these regressions had r2 values above 0.98.

 
y = C1ek1t + C2ek2t + C3(1)

where y is the relative residual activity, t is the time in minutes and C1, C2, C3, k1 and k2 are constant parameters (Figure 3). This result suggests that TNXI and its derivatives inactivate following the series deactivation scheme:

where a, a1 and a2 are the specific activities of enzyme states E, E1 and E2, respectively; k1 and k2 are first-order inactivation rate constants; and ß1 and ß2 are the ratios a1/a and a2/a, respectively. The rate constants k1 and k2 are directly obtained from Equation 1; the relative activity of state E2, ß2, corresponds to C3 in Equation 1; and the relative activity of state E1, ß1, is obtained from the equation

ß1 = 1 – C1C2[1 + (k2k1)/k1](2)

as described (Sadana, 1991Go). First-order inactivation rate constants k1 and k2 and relative activities ß1 and ß2 are listed in Table II, as well as the {chi}2 and r2 values for the non-linear least-squares fit regressions.


View this table:
[in this window]
[in a new window]
 
Table II. Inactivation parameters of TNXI and its mutant derivatives at 80°C
 
Although the curve fit of our inactivation data does not identify an inactivation mechanism and does not indicate the number of intermediate enzyme forms, we could still see that, with the exception of 1F1 at pH 7.0 (whose E2 form was inactive), the four enzymes inactivated at both pH values to a partially active form E2 (with relative remaining activity between 0.11 and 0.39) that remained stable in these conditions of temperature and pH. At pH 7.0, both 3A2 and 1F1 were kinetically slightly more stable than TNXI and TNXI V185T (Figure 3A). Although the inactivation rate constants k1 and k2 for 3A2 obtained from the curve fit were higher than those of TNXI and TNXI V185T, 3A2 appeared slightly more stable because its E1 and E2 forms had the highest relative residual activities ß1 and ß2 (Table II). 1F1 was the most stable enzyme at pH 7.0 due to its lower inactivation rate constants k1 and k2 and to a ß1 value higher than those of TNXI and TNXI V185T. At pH 5.5, only 1F1 remained more stable than TNXI and TNXI V185T; 3A2 was less stable. Despite having inactivation rate constants slightly lower than those of TNXI and TNXI V185T, 3A2 was the least stable enzyme in long-term incubations due to the low relative residual activity ß2 of its E2 form. In our 5 h inactivation experiment, 1F1’s higher stability was due to its low inactivation rate constants k1 and k2. Because the E2 form of TNXI V185T was ~40% higher than that of 1F1, though, TNXI V185T would be the more stable enzyme in longer-term inactivation experiments.

Amino acid substitutions in 3A2 and 1F1

The mutations present in 3A2 and 1F1 were identified by DNA sequencing. In addition to V185T already present in TNXI V185T, 3A2 contained a single additional mutation, L282P. The L282P mutation is located in helix {alpha}7 of the ({alpha}/ß)8 barrel structure, at ~12–14 Å from the catalytic center (Figure 4). Helix {alpha}7 is located at the surface of a monomer and at the interface of the tight dimer. Neither Leu nor Pro’s side chain can form hydrogen bonds with neighboring residues. Whenever a proline occurs in a peptide chain, it interrupts {alpha}-helices and creates a kink or bend (Lehninger, 1970Go). Detailed analysis of the L282P mutation modeled into the TNXI structure (Figure 5) shows that Pro282:C{delta} is in close contact with the main chain Phe278:O and Gln279:O atoms (1.73 and 2.21 Å, respectively). With van der Waals’s radii of 1.87 and 1.35 Å for C and O atoms, respectively, optimal van der Waals interactions between Pro282:C{delta} and Phe278:O or Gln279:O would take place at ~3.2 Å. The unfavorable van der Waals clashes created by Pro282 probably lead to local conformational rearrangements. These changes might, in turn, affect the active site structure and dynamics, and the enzyme’s interaction with the substrate. They could possibly also affect inter-subunit interactions within the tight dimer.



View larger version (66K):
[in this window]
[in a new window]
 
Fig. 4. Three-dimensional model of the 1F1 monomer showing the positions of mutations V185T, F186S and L282P. {alpha}-Helices are represented by cylinders and ß-sheets by flat arrows; residues 185, 186 and 282 are in Corey–Pauling–Koltun space-filling (CPK) representation.

 


View larger version (22K):
[in this window]
[in a new window]
 
Fig. 5. Three-dimensional model of 3A2 around mutation L282P. The backbone structure of residues 278–283 is shown in light gray. (A) Leu282 and (B) Pro282 side chains are shown in black.

 
1F1 contains the same two mutations as 3A2, plus mutation F186S. This last mutation is located at the bottom of the active site, adjacent to mutation V185T. Residue 186’s side chain points away from the active site cavity, and it forms aromatic interactions with Tyr184, Phe228 and Phe262 (Figure 6), as well as hydrophobic interactions with Leu229 (data not shown). Serine is a much smaller residue than phenylalanine. The F186S mutation creates a large void in the back of the active site (Figure 6). This void probably leads to a rearrangement of the neighboring residues and to changes in the catalytic site’s dynamics. These changes are in turn probably responsible for the large increase in low temperature activity of mutant 1F1.



View larger version (39K):
[in this window]
[in a new window]
 
Fig. 6. Cavity created by mutation F186S in mutant 1F1. Residues surrounding the cavity occupied by residue 186 are shown in TNXI (left) and 1F1 (right) in CPK representation. Carbon atoms (green); nitrogen atoms (blue); oxygen atoms (red); residue 186 (yellow).

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Thermostable enzymes are generally barely active at low temperature, but they are as active at their optimal growth temperature as their mesophilic counterparts (Varley and Pain, 1991Go). We showed here that the activity of a hyperthermostable enzyme at low temperature and low pH can be increased without a loss in thermostability. Our results have shown that the quality of our library of random TNXI mutants is sufficient to isolate mutants with increased activity at low temperature and low pH. Using two sequential rounds of random mutagenesis, we were able to obtain a TNXI mutant derivative, 1F1, showing high activity at low temperature and low pH. 1F1 is not only more active overall than its parental enzymes, but this activity difference is most pronounced at low temperatures. Since 1F1 is more active at low temperature and has a lower temperature optimum for activity, but is more stable than the wild-type TNXI, we suggest that the molecular determinants of this enzyme’s activity and thermostability are, in fact, not the same. This assumption has also been previously observed in the study of B.subtilis p-nitrobenzyl esterase, in which the laboratory-evolved mutant enzyme had a 14°C increase in Tm but still maintained its catalytic activity at low temperature (Giver et al., 1998Go).

Recent studies (Aguilar et al., 1997Go; Zavodszky et al., 1998Go; Kohen et al., 1999Go) suggested that the reduced flexibility of thermostable enzymes was the factor impairing their catalytic activity at low temperatures. Particularly striking is the potential of single point mutations to significantly increase low-temperature activity. Recent studies of psychrophilic enzymes have suggested that, despite the many differences observed between mesophilic and psychrophilic enzymes, single point mutations may be capable of conferring most psychrophilic characteristics (Somero, 1995Go; Feller and Gerday, 1997Go). Other studies using random mutagenesis and screening/selection succeeded in increasing the activity (3-fold at 20°C for Pyrococcus furiosus ß-glucosidase and 17-fold at 37°C for Sulfolobus solfataricus indoleglycerol phosphate synthase) of hyperthermophilic enzymes at mesophilic temperatures with changes in temperature optima (Lebbink et al., 2000Go; Merz et al., 2000Go). In our study, 1F1 showed 4.5- and 2.2-fold increases in Vmax at 60°C (pH 7.0) and 80°C (pH 5.5), respectively, with an optimal temperature for activity only 5°C lower than that of TNXI V185T. The Arrhenius plots of activity of TNXI and its mutants (Figure 1B) revealed that TNXI and TNXI V185T require higher activation energy for activity than either TNXI 3A2 or 1F1. The difference is more pronounced with 1F1, with an ~2-fold decrease in the Ea value compared with those of TNXI and TNXI V185T. The reduction in Ea for activity observed in TNXI 3A2 and 1F1 suggests increased flexibility in the active site of these enzymes even at low temperature; thus, their activity at low temperature is vastly enhanced. Although TNXI V185T has a higher catalytic efficiency on glucose than TNXI, due to an increased glucose binding affinity and a higher catalytic rate (Sriprapundh et al., 2000Go), its Ea of activity remained similar to that of TNXI, suggesting that TNXI V185T’s active site dynamics remained unchanged. This observation is in good agreement with the assumption of Lönn et al. (Lönn et al., 2002Go) that mutations underlying the adaptation of enzymes to temperatures lower than their optimum allow a higher degree of flexibility in areas that move during catalysis. This higher flexibility, in turn, reduces the free energy of activation compared with the wild-type enzymes. The higher flexibility in areas that move during catalysis increases the kcat of the reactions catalyzed by the cold-adapted enzymes. A study of lactate dehydrogenase’s cold adaptation (Fields and Somero, 1998Go) also found that mutations that increase flexibility in regions of the enzyme involved in catalytic conformational changes may reduce energy barriers to these rate-governing conformational changes and thereby increase kcat. TNXI 1F1 has higher kcat and Km values than TNXI V185T does. This observation was rationalized in terms of localized increases in conformational flexibility; mutations that reduce the energetic barriers between different active site conformations (thus allowing for more rapid interconversion among them) will lead to higher kcat values. These same mutations, however, will allow the enzyme to populate conformations that bind substrate poorly more easily, leading to increases in Km. Our study provides obvious support for this hypothesis.

Our sequential random mutagenesis and screening approach with TNXI resulted in the identification of amino acids or local structural conformations that are critical for low-temperature/low-pH catalytic activity. Two mutations were identified in TNXI 1F1 in addition to V185T, namely L282P and F186S. Leu282 is close to the inter-subunit interface of the enzyme tight dimer. While the L282P mutation improved the enzyme’s low temperature and low pH activities, the detailed analysis of the modeled three-dimensional structure of 3A2 revealed unfavorable van der Waals contacts between Pro282’s pyrrolidine ring and the local backbone structure. The changes in local dynamics created by mutation L282P are most probably responsible for the 35% decrease in activation energy for the activity of 3A2 on glucose. It is interesting to note that the kink introduced by mutation L282P in helix {alpha}7 of 3A2’s ({alpha}/ß)8 barrel does not destabilize the enzyme at pH 7.0, and it only decreases the relative activity of 3A2’s E2 form at pH 5.5. These results suggest that mutation L282P occurred in an area of the enzyme structure whose stability is not limiting for the stability of the whole enzyme. In other words, the first steps of TNXI inactivation involve regions of the protein that are less stable than the region surrounding helix {alpha}7. The second mutation, F186S, is located in the back of the active site, adjacent to Thr185. Since serine’s side chain is considerably smaller than phenylalanine’s, this mutation creates a cavity that probably increases mobility in the active site resulting in a dramatic improvement of 1F1’s low-temperature activity. Because mutation F186S disturbs a potentially stabilizing four-residue network of aromatic interactions, the mechanism underlying the increase in kinetic stability of 1F1 remains unknown. Determination of the crystal structures of 3A2 and 1F1 is under way. These structures should provide insightful information on these enzymes’ increased activity and on their stability properties.

There is some evidence that demonstrates the effect of positions where mutations occur on activity and stability of laboratory and naturally evolved enzymes. A study of psychrophilic enzymes revealed that point mutations distant from the catalytic center or in the major substrate-binding site of enzymes could lead to cold adaptation (Feller and Gerday, 1997Go). In the studies of Lebbink et al. (Lebbink et al., 2000Go), all mutants containing subunit interface substitutions were less stable and had lower temperature optima than the wild-type P.furiosus ß-glucosidase, suggesting that subunit interfaces also play an important role in thermoadaptation. Our results showed that even without selective pressure to maintain thermostability, it is possible to obtain a mutant thermozyme with a mutation in the active site that has comparable thermostability to the wild-type enzyme while its low-temperature and low-pH activity are vastly enhanced.

The only random mutagenesis performed on XI so far was by Lönn et al. (Lönn et al., 2002Go). The thermophilic type I Thermus thermophilus XI was subjected to one round of random PCR mutagenesis; the mutant library was screened for increased XI activity at low temperature. Three mutants were isolated, containing point mutations E372G/V379A in the first, F163L/E372G in the second and E372G in the third mutant. These three mutant enzymes showed increased kcat values (up to 9-fold increases on both xylose and glucose) with up to 26 times higher Km values on xylose but relatively unchanged Km values for glucose. All enzyme variants’ relative activities on xylose were higher than that of the wild-type at low temperatures, but the mutants were kinetically less thermostable. These results, together with ours, suggest that point mutations either distant from or inside the catalytic center can lead to cold adaptation. The only difference between their work and ours was that we were able to maintain the thermostability of our mutant enzyme while increasing its activity at low temperatures.

An alignment of different XIs (data not shown) revealed that neither Pro282 nor Ser186 is present in any XI. Because it creates a cavity at the back of the catalytic site and because it disrupts a four-residue network of aromatic interactions, mutation F186S could not have been rationally designed based on the structure of TNXI and on modeling. Similarly, the effect of mutation L282P, in the middle of an {alpha}-helix and 12–14 Å from the active site, was completely unpredictable. These mutations could only be obtained through directed evolution. With a vast improvement in specific activity at 60°C, at pH 5.5, a higher catalytic efficiency on glucose than TNXI V185T in all conditions tested, and a thermostability comparable with that of TNXI V185T, 1F1 could be an interesting candidate for industrial applications. Further study of 1F1’s potential usefulness in conditions used in the industrial production of high fructose syrup in comparison with a commercially available glucose isomerase is under way.


    Acknowledgement
 
This work was supported by the National Science Foundation, grant no. BES-0115754.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Aguilar,C.F., Sanderson,I., Moracci,M., Ciaramella,M., Nucci,R., Rossi,M. and Pearl,L.H. (1997) J. Mol. Biol., 271, 789–802.[CrossRef][ISI][Medline]

Arnold,F.H. and Moore,J.C. (1997) Adv. Biochem. Eng. Biotechnol., 58, 1–14.[Medline]

Ausubel,F.M., Brent,R., Kingston,R.E., Moore,D.D., Seidman,J.G., Smith,J.A. and Struhl,K. (eds) (1993) Current Protocols in Molecular Biology. Greene Publishing and Wiley-Interscience, New York.

Bhosale,S.H., Rao,M.B. and Deshpande,V.V. (1996) Microbiol. Rev., 60, 280–300.[ISI][Medline]

Chen,W. (1980) Process Biochem., 15, 30–35.

Feller,G. and Gerday,C. (1997) Cell Mol. Life Sci., 53, 830–841.[CrossRef][ISI][Medline]

Fields,P.A. and Somero,G.N. (1998) Proc. Natl Acad. Sci. USA, 95, 11476–11481.[Abstract/Free Full Text]

Giver,L., Gershenson,A., Freskgard,P. and Arnold,F.H. (1998) Proc. Natl Acad. Sci. USA, 95, 12809–12813.[Abstract/Free Full Text]

Kohen,A., Cannio,R., Bartolucci,S. and Klinman,J.P. (1999) Nature, 399, 496–499.[CrossRef][ISI][Medline]

Kuchner,O. and Arnold,F.H. (1997) Trends Biotechnol., 15, 523–530.[CrossRef][ISI][Medline]

Lebbink,J.H., Kaper,T., Bron,P., van der Oost,J. and de Vos,W.M. (2000) Biochemistry, 39, 3656–3665.[CrossRef][ISI][Medline]

Lee,C.Y. and Zeikus,J.G. (1991) Biochem. J., 273, 565–571.[ISI][Medline]

Lehninger,A.L. (1970) Biochemistry, 1st edn. Worth, New York, NY.

Leung,D.W., Chen,E. and Goeddel,D.V. (1989) Technique, 1, 1–15.

Lönn,A., Gárdonyi,M., van Zyl,W., Hahn-Hägerdal,B. and Otero,R.C. (2002) Eur. J. Biochem., 269, 157–163.[Abstract/Free Full Text]

Meng,M., Bagdasarian,M. and Zeikus,J.G. (1993) Proc. Natl Acad. Sci. USA, 90, 8459–8463.[Abstract/Free Full Text]

Merz,A., Yee,M.C., Szadkowski,H., Pappenberger,G., Crameri,A., Stemmer,W.P., Yanofsky,C. and Kirschner,K. (2000) Biochemistry, 39, 880–889.[CrossRef][ISI][Medline]

Moore,G.L. and Maranas,C.D. (2000) J. Theor. Biol., 205, 483–503.[CrossRef][ISI][Medline]

Ostermeier,M., Nixon,A.E. and Benkovic,S.J. (1999) Bioorg. Med. Chem., 7, 2139–2144.[CrossRef][ISI][Medline]

Sadana,A. (1991) Biocatalysis: Fundamentals of Enzyme Deactivation Kinetics. Prentice-Hall, Englewood Cliffs, NJ.

Schenk,M. and Bisswanger,H. (1998) Enzyme Microb. Technol., 22, 721–723.[CrossRef][ISI]

Shao,Z., Zhao,H., Giver,L. and Arnold,F.H. (1998) Nucleic Acids Res., 26, 681–683.[Abstract/Free Full Text]

Somero,G.N. (1995) Annu. Rev. Physiol., 57, 43–68.[CrossRef][ISI][Medline]

Sriprapundh,D., Vieille,C. and Zeikus,J.G. (2000) Protein Eng., 13, 259–265.[Abstract/Free Full Text]

Stemmer,W.P. (1994) Proc. Natl Acad. Sci. USA, 91, 10747–10751.[Abstract/Free Full Text]

Varley,P.G. and Pain,R.H. (1991) J. Mol. Biol., 220, 531–538.[ISI][Medline]

Vieille,C. and Zeikus,J.G. (2001) Microbiol. Mol. Biol. Rev., 65, 1–43.[Abstract/Free Full Text]

Vieille,C., Hess,J.M., Kelly,R.M. and Zeikus,J.G. (1995) Appl. Environ. Microbiol., 61, 1867–1875.[Abstract]

Zavodszky,P., Kardos,J., Svingor,A. and Petsko,G. (1998) Proc. Natl Acad. Sci. USA, 95, 7406–7411.[Abstract/Free Full Text]

Zhao,H. and Arnold,F.H. (1999) Protein Eng., 12, 47–53.[Abstract/Free Full Text]

Zhao,H., Giver,L., Shao,Z., Affholter,J.A. and Arnold,F.H. (1998) Nat. Biotechnol., 16, 258–261.[ISI][Medline]

Received February 21, 2003; revised June 24, 2003; accepted July 17, 2003.