Báo cáo Y học: Cold adaptation of xylose isomerase from Thermus thermophilus through random PCR mutagenesis Gene cloning and protein characterization

Eur. J. Biochem. 269, 157–163 (2002) (cid:211) FEBS 2002

Cold adaptation of xylose isomerase from Thermusthermophilus through random PCR mutagenesis Gene cloning and protein characterization

Anna Lo¨ nn1, Ma´ rk Ga´ rdonyi1, Willem van Zyl2, Ba¨ rbel Hahn-Ha¨ gerdal1 and Ricardo Cordero Otero2,*

1Department of Applied Microbiology, Lund University, Sweden; 2Department of Microbiology, University of Stellenbosch, Matieland, South Africa

enzyme at 60 (cid:176)C, but they did not show any increase in catalytic e(cid:129)ciency (kcat/Km). For D-glucose, both the kcat and the kcat/Km values for the mutants were increased compared with the wild-type enzyme. Furthermore, the mutant enzymes exhibited up to 255 times higher inhibi- tion constants (Ki) for xylitol than the wild-type, indicat- ing that they are less inhibited by xylitol. The thermal stability of the mutated enzymes was poorer than that of the wild-type enzyme. The results are discussed in terms of increased molecular flexibility of the mutant enzymes at low temperatures.

Keywords: xylose isomerase; cold adaptation; random mutagenesis; Saccharomyces cerevisiae; xylose fermentation.

Random PCR mutagenesis was applied to the Thermus thermophilus xylA gene encoding xylose isomerase. Three cold-adapted mutants were isolated with the following amino-acid substitutions: E372G, V379A (M-1021), E372G, F163L (M-1024) and E372G (M-1026). The wild- type and mutated xylA genes were cloned and expressed in Escherichia coli HB101 using the vector pGEM(cid:210)-T Easy, and their physicochemical and catalytic properties were determined. The optimum pH for xylose isomeriza- tion activity for the mutants was (cid:25) 7.0, which is similar to the wild-type enzyme. Compared with the wild-type, the mutants were active over a broader pH range. The mutants exhibited up to nine times higher catalytic rate constants (kcat) for D-xylose compared with the wild-type

the prime

cerevisiae is

D-fructose [6]. The physiological function of the enzyme in vivo is, however, the isomerization of the pentose D-xylose to D-xylulose. XI genes (xylA) from several bacteria have been introduced into S. cerevisiae, including xylA from Escherichia coli [7,8], Actinoplanes missouriensis [9], Bacillus subtilis [9], Lactobacillus pentosus [10] and Clostridium thermosulfurogenes [11]. However, none of these attempts generated an active XI.

3-isopropylmalate-dehydrogenase

The use of ethanol from renewable raw materials is an attractive alternative for meeting increasing global demand for liquid fuels because its combustion does not contribute to the greenhouse effect. For the industrial production of ethanol from pretreated and hydrolysed lignocellulose, the yeast Saccharomyces choice (reviewed in [1]). Between 10 and 40% of lignocellulosic raw materials consists of pentoses [2], where xylose is the predominant portion. However, S. cerevisiae can not metabolize xylose, only D-xylulose, an isomerization product of D-xylose. Xylose reductase (EC 1.1.1.21) and xylitol dehydrogenase (EC 1.1.1.9) from the xylose-fer- menting yeast Pichia stipitis, have been introduced into S. cerevisae to allow xylose fermentation to ethanol [3–5]. Fermentations resulted in low ethanol yields and consid- erable xylitol by-product formation. Xylose isomerase (XI) (EC 5.3.1.5) is used in the production of high-fructose corn syrup, where it catalyses the conversion of D-glucose to

The only xylA gene successfully expressed in S. cerevi- siae was cloned from T. thermophilus [12]. This thermo- philic XI, with a temperature optimum at 85 (cid:176)C, has a low activity at 30 (cid:176)C [12] which is the optimal growth temperature for S. cerevisiae. It would therefore be desirable to generate mutants of XI with improved kinetic properties at low temperatures. Random chemical muta- genesis has been used recently to obtain variants of the T. thermophilus [13], Sulfolobus solfataricus indolglycerol phosphate synthase [14] and the mesophilic protease subtilisin BPN¢ [15–17], with increased activity at low temperatures. Error-prone PCR followed by DNA shuffling resulted in the artificial evolution of cold-adapted mutants of a b-glycosidase from Pyrococcus furiosus [18] and a subtilisin-like protease from Bacillus sphaericus [19].

Here, we report on random PCR mutagenesis to create cold-adapted T. thermophilus XI. The character- ization of the physicochemical and catalytic properties of three cold-adapted XIs that exhibited up to 9 times higher kcat for xylose than the wild-type enzyme at 60 (cid:176)C are described.

Correspondence to B. Hahn-Ha¨ gerdal, Department of Applied Microbiology, Lund University, PO Box 124, SE-221 00 Lund, Sweden. Fax: + 46 46 2224203, Tel.: + 46 46 2228428, E-mail: Barbel.Hahn-Hagerdal@tmb.lth.se Abbreviations: XI, xylose isomerase. *Present address: Institute for Wine Biotechnology, University of Stellenbosch, Private Bag XI, Matieland 7602, South Africa. (Received 28 May 2001, revised 23 October 2001, accepted 25 October 2001)

(Eur. J. Biochem. 269)

(cid:211) FEBS 2002

158 A. Lo¨ nn et al.

Protein determination

M A T E R I A L S A N D M E T H O D S

Chemicals

Protein concentration was determined using the Pierce protein reagent with bovine serum albumin as standard [23].

Page

SDS/PAGE was performed as previously described [24].

All chemicals were obtained from commercial suppliers and used as described by the manufacturer. D(+)-xylose was obtained from Sigma (Steinheim, Germany) and sorbitol dehydrogenase from Boehringer Mannheim (Mannheim, Germany).

Immunochemical determination of XI

Strains and plasmids

Escherichia coli HB101(F-hsdS20ara-1 recA13 proA12 lacY1 galK2 rspL20 mtl-1xyl-5) [20] was used for cloning of the mutated XIs using pGEM(cid:210)-T Easy vector (Promega, Madison, WI, USA).

PCR mutagenesis

Rabbit antiserum against XI from Streptomyces rubiginosus was prepared by Antibody AB (So¨ dra Sandby, Sweden) and immunoblotting was performed as described previously [25]. Briefly, 2 lg of cell-free extract together with 2–50 ng of purified XI from S. rubiginosus were resolved by SDS/ PAGE and were then electrophoretically transferred onto a poly(vinylidene difluoride) membrane (Bio-Rad, Hercules, CA, USA). The blotted proteins were identified immuno- chemically by sequential addition of anti-XI serum followed by goat anti-(rabbit IgG) Ig conjugated with alkaline phosphatase (Bio-Rad, Hercules, CA, USA). The secondary antibody was detected with a Storm 860(cid:210) (Pharmacia Amersham, Uppsala, Sweden) using a chemifluorescent substrate ECF (Pharmacia Amersham). Data analysis was performed using IMAGE QUANT(cid:210) software (Pharmacia Amershamm), giving a quantitative measurement of the amount of XI in the cell-free extracts. These data were used with the maximum velocity (Vmax) to calculate kcat.

Enzyme assays

Random mutagenesis of the XI gene (xylA) was performed under conditions described previously [21] using the PCR primers 5¢-TGATCAATGTACGAGCCCAAACC-3¢ and 5¢-TGATCACCCCCGCACC-3¢, which directly flank the xylA gene. Both primers contained the restriction endonuc- lease site for BclI (underlined). The PCR contained: 1 · PCR buffer (BIOTAQ(cid:228)), 0.2 mM dATP, 0.2 mM dGTP, 1 mM dCTP, 1 mM dTTP, 1.5 mM MgCl2, 0.5 mM MnCl2, 0.15 lM of both primers, 0.02 nM template DNA and 5 U Taq DNA polymerase (BIOTAQ(cid:228)) in a total volume of 100 lL. PCR was performed in a Thermal Cycler (PerkinElmer 2400) for nine cycles: 30 s at 94 (cid:176)C, 30 s at 50 (cid:176)C and 45 s at 68 (cid:176)C. The PCR products were then purified using High Pure(cid:228) PCR Product (Boehringer Mannheim).

DNA sequencing

Analysis of the mutated sequences was carried out using ABI PRISM(cid:210) Big Dye(cid:228) Terminator cycle sequencing ready reaction kits with an ABI PRISM(cid:228) 377 DNA sequencer (PE/Applied Biosystems). Both the coding and the noncoding strands were sequenced to ensure the reliable identification of all mutations.

Growth conditions and preparation of cell extract from E.coli

A two-step XI standard assay (0.5 mL) was modified from [26]. A substrate concentration of 700 mM D-xylose was used at 60 (cid:176)C in 200 mM triethanolamine at pH 7.0 in the presence of 10 mM MnCl2 and crude enzyme preparations. Glucose isomerase activity was assayed under the same reaction conditions as those used in the XI assay, except that glucose instead of xylose was used in the reaction mixture. The reactions were stopped by adding 150 lL 50% trichloroacetic acid, and then 2 M Na2CO3 was added to neutralize the solutions. The isomerization products, xylu- lose or fructose, were reduced at pH 7.0 (37 (cid:176)C) with 0.04 U sorbitol dehydrogenase (SDH) or 0.5 U SDH, respectively, and 0.15 mM NADH using a COBAS MIRA plus (Roche, Mannheim, Germany). The rate of disappearance of NADH was followed at 340 nm and the amount of D-xylulose and D-fructose determined from calibration curves. One unit of isomerase activity was defined as the amount of crude enzyme required to produce 1 lmol of product per minute under the assay conditions employed. The specific activity (UÆmin)1Æmg)1) was determined from the activity and the protein concentration of the crude enzyme preparations.

Kinetic parameters

E. coli HB101 harbouring the plasmids pGEM(cid:210)-T Easy containing the wild-type and the mutated XI genes were grown at 37 (cid:176)C in 50 mL Luria–Bertani medium [22] containing 100 lgÆmL)1 ampicillin. The cells were har- vested by centrifugation in the stationary phase of growth and washed once with ice-cold distilled water. Washed resuspended in 100 mM triethanolamine, cells were pH 7.0, 65 kUÆmL)1 lysozyme, 0.25 mgÆmL)1 DNAse and 1 mM phenylmethanesulfonylfluoride in dimethylsulf- oxide. The solutions were kept at room temperature for 1 h and then on ice for 2 h before storing in a freezer at )20 (cid:176)C. Cell extracts were thawed on ice, cell debris was removed by centrifugation (15 000 g for 15 min at 4 (cid:176)C) and the supernatant was used as the crude enzyme preparation.

(lmolÆmin)1Æmg)1) and The kinetic parameters, Vmax (Km, mM), were determined from Michaelis constant Michaelis–Menten plots of specific activities at various substrate concentrations. Typically, duplicate measure- ments at 6–10 concentrations of substrate spanning the value of Km were used to determine the value of Km. The

(cid:211) FEBS 2002

Mutant xylose isomerases (Eur. J. Biochem. 269) 159

concentration of XI in the cell-free extracts was determined immunochemically using a molecular mass of 44 000 kDa [27], to allow calculation of the catalytic rate constant (kcat) from the relationship kcat (cid:136) Vmax/[E0], where [E0] (cid:136) total enzyme concentration [28].

The Ki (mM) for xylitol was determined by incubating crude enzyme preparations in different xylose concentra- tions (20–600 mM) at different fixed xylitol concentrations. By plotting the specific activities for each xylitol concentra- tion against the xylose concentrations, Ki was determined using the equation Km¢ (cid:136) KmÆ(1 + i/Ki) [29], where i is the xylitol concentration (mM) and Km¢ the apparent Km value at a certain concentration of xylitol.

PH profile

The effect of pH on the activity of the wild-type and mutated enzymes was investigated in the pH range 5–10 in 700 mM xylose, 10 mM MnCl2 and a buffer prepared by mixing acetate, Pipes, Hepes and glycine, to a final concentration of 50 mM each [30]. The pH was adjusted at 60 (cid:176)C with NaOH. Above pH 7.0 corrections were made for the chemical isomerization of D-xylose.

Temperature profile

medium allowed detection and quantification of red acid- producing colonies. Three candidate mutants, termed M-1021, M-1024 and M-1026 were identified. Colonies of these three were a deeper red on the McConkey/xylose medium than were wild-type xylA colonies (suggesting higher XI activity). DNA sequencing revealed that the mutants exhibited approximately 80% transitions (T to C) and 20% transversions (A to C or T).

The temperature profiles for the wild-type XI and mutated XIs were measured at temperatures between 30 and 95 (cid:176)C. Above 60 (cid:176)C corrections were made for the chemical isomerization of D-xylose.

Preparation of metal-free XI and metal ion effects on enzyme activity

Metal-free enzymes were prepared as previously described [26]. No isomerase activity was observed in the absence of Mn2+, Mg2+ or Co2+. The effect of metal ions on XI activity was determined by adding 10 mM final concentra- tion of either CoCl2, MnCl2 or MgCl2 to the metal-free enzyme preparations in the assay mixture.

Enzyme stability

XI from T. thermophilus is a homotetrameric enzyme with a 387-residue subunit. Each monomer comprises two domains: the larger N-terminal domain (domain I, residues 1–321), which folds into a (b/a)8 barrel, and the smaller C-terminal domain (domain II, residues 322–387), which consists of loops and helices (Fig. 1) [31]. Domain II extends from domain I and makes extensive contacts with a neighbouring subunit. M-1021 contained two mutations in domain II; E372G and V379A. M-1024 possessed two mutations, one in domain I (F163L) and one in domain II (E372G). M-1026 carries one mutation in domain II that is shared by M-1024 and M-1021; E372G. The locations of the amino-acid substitutions in the original tertiary structure of XI are shown in Fig. 1. Neither the substrate-binding sites (H53, D56 and K182) nor the metal-binding sites (E180, E216, H219, D244, D254, D256 and D286) were affected by the mutations in the mutant enzymes.

Properties of the mutant enzymes

The temperature stability of the wild-type XI and mutated XIs was investigated by incubating metal-free crude enzyme preparations in 200 mM triethanolamine, pH 7.0 with 10 mM MnCl2 in airtight tubes at 70 (cid:176)C. At different times, 100-lL samples were withdrawn and stored on ice until the residual activity was determined.

Fig. 1. Structure of one subunit of T. thermophilus XI. The amino acids 372, 379 and 163 are identified to show the position of the mutations.

R E S U L T S

Isolation of XI mutants with increased activity at low temperatures

Temperature profiles. XI from T. thermophilus has a temperature optimum around 95 (cid:176)C [30]. To investigate whether the mutations caused any change in the tempera- ture optimum the temperature profiles were investigated from 30 to 95 (cid:176)C (Fig. 2). The temperature optimum for M-1024 and M-1026 was around 5 (cid:176)C higher than the optimum for the wild-type (90 (cid:176)C). For M-1021 the temperature optimum was somewhat lower, (cid:25) 75 (cid:176)C. At 30 (cid:176)C the specific activity was higher for the mutants than for the wild-type XI. Due to the overall low activity of the enzymes at this temperature, the physicochemical and the wild-type and mutant kinetic characterization of enzymes was carried out at 60 (cid:176)C.

PH profiles. XI from T. thermophilus shows a pH optimum around 7.0 [30]. To examine whether the mutations altered the pH dependence for xylose isomerization, the activity of

One-step mutagenesis was used to screen for mutant XIs with improved activity at low temperatures. The mutated XI fragments were cloned into the vector E. coli pGEM(cid:210)-T Easy and transformed into the E. coli HB101 (xyl-5) strain to generate a mutant library. Transformants were replica plated on McConkey agar plates, complemented with 1% xylose and cultivated at 37 (cid:176)C overnight. After a further 2 days of incubation at 30 (cid:176)C, the pH indicator in the

(Eur. J. Biochem. 269)

(cid:211) FEBS 2002

160 A. Lo¨ nn et al.

120

100

)

80

Table 1. E(cid:128)ect of various bivalent cations (10 mM) on the activity of EDTA-treated enzymes. The % relative activity is shown compared to the specific activity with 10 mM MnCl2 at 60 (cid:176)C which was set to 100% for each enzyme.

i

60

y t i v i t c a

m u m x a m

40

e v i t

f

l

o %

a e R

20

(

Enzyme Co2+ Mg2+

0

20

30

40

80

90

100

50

70

60 Temperature ( oC)

100% activity [30]. Metal ions are not only essential for the catalytic mechanism, but they also contribute to the stabilization of the native structure, which is especially important for thermophilic enzymes.

Wild type M-1021 M-1024 M-1026 87.9 7.4 33.6 34.6 73.8 23.2 22.7 38.9

The effect of different bivalent metal ions (Mn2+, Mg2+ and Co2+) on the EDTA-treated enzymes was investigated (Table 1). The wild-type and mutated XIs were most effectively activated by Mn2+ and, to a smaller degree, by Mg2+ and Co2+. The wild-type showed 88 and 74% of the maximum activity with Co2+ and Mg2+, respectively. The mutants, on the other hand, were less activated by Co2+ and Mg2+.

each mutant enzyme was measured as a function of pH (Fig. 3). The activity of each enzyme relative to the maximum activity was plotted as a percentage against pH. The pH dependence of the enzyme activity was examined at a substrate concentration well above Km, where the velocity of the reaction is proportional to kcat. The pH activity profiles of the mutants were broader, and extended into the alkaline region, compared with the wild-type XI. The wild- type showed no XI activity at pH 9 and 10. For M-1024 and M-1026 the specific activity at pH 9 and 10, was 66 and 45%, and 62 and 31%, of the maximum, respectively. The pH optima for the mutant XIs were not significantly different from that of the wild-type, i.e. around 7.0.

Kinetic properties of D-xylose and D-glucose isomeriza- tion. The kinetics of D-xylose and D-glucose isomerization were determined from crude enzyme preparations at 60 (cid:176)C, pH 7.0, and at metal-ion saturation (Mn2+) (Table 2). The Km values for D-xylose were up to 26 times higher for the mutants, and the catalytic rate constants (kcat) were up to nine times higher than for the wild-type enzyme. The catalytic efficiency (kcat/Km) for D-xylose for M-1026 was 6% higher than that of the wild-type, while for the other mutants it was lower.

E(cid:128)ect of metal ions. XIs require two metal ions to be bound to the active site of each monomer in order to exhibit enzyme activity [32]. However, XIs from different organisms require different metals for optimal activity [33], and XI from T. thermophilus requires either Mg2+ or Mn2+ for

As for the wild-type XI, the mutants had a lower Km and higher kcat for D-xylose than for D-glucose. The Km for glucose for M-1021 and M-1024 was lower, by as much as three times, than for the wild-type enzyme. For M-1026, on the other hand, the Km was higher than that of the wild-type enzyme. The kcat and the kcat/Km values for D-glucose were up to five and seven times higher, respectively, for all the mutants, than for the wild-type XI.

Inhibition by xylitol. The extended acyclic forms of the substrates xylose and glucose have binding closely resem- bling that observed for the acyclic polyol inhibitor xylitol [34,35]. Competitive inhibition is thus expected and has previously been reported [36,37]. Ki for xylitol for the three mutant enzymes was between seven (M-1021) and 255 (M-1024) times higher, than for the wild-type enzyme (Table 2), indicating that the mutant enzymes are not inhibited by xylitol to the same extent as the wild-type enzyme.

Fig. 2. The relative activity at di(cid:128)erent temperatures for the mutated XIs and the wild-type XI: (e) wild-type; (d) M-1021; (,) M-1024; and (j) M-1026. The scale of relative activity (%) indicates the percentage of experimental values at various temperatures relative to the maxi- mum value of each enzyme.

Thermal stability. To determine whether the mutations producing a change in the temperature dependence of XI activity also affected the thermal stability of the mutated enzymes, the residual activities were measured after heat treatment at 70 (cid:176)C for various lengths of time (Fig. 4). Investigations of the metal-free enzyme preparations in buffer at saturated metal concentration (Mn2+) showed that the wild-type XI and the mutated XIs retained almost

Fig. 3. The relative activity at di(cid:128)erent values of pH for the mutated XIs and the wild-type XI: (e) wild-type; (d) M-1021; (,) M-1024; and (j) M-1026. The scale of relative activity (%) indicates the percentage of experimental value at various pH relative to the maximum value of each enzyme.

(cid:211) FEBS 2002

Mutant xylose isomerases (Eur. J. Biochem. 269) 161

Table 2. Kinetic properties of wild-type XI and mutated XIs.

Xylose Glucose

)1)

)1)

Km (mM) kcat (s)1) kcat/Km (s)1ÆmM Km (mM) kcat (s)1) kcat/Km (s)1ÆmM Xylitol K i (mM)

4.6 33.2 1174 3.44 (cid:139) 0.4 25.1 (cid:139) 4.0 89.4 (cid:139) 8.4 28.7 (cid:139) 3.3 46.6 257.5 381.6 412.4 13.6 10.3 4.3 14.4 146.8 (cid:139) 12.3 52.0 (cid:139) 4.9 130.8 (cid:139) 17.1 171.8 (cid:139) 10.0 16.3 39.9 66.9 88.7 0.11 0.77 0.51 0.47 68.7 Wild-type M-1021 M-1024 M-1026

changes, these effects should play an important role [39]. The effect of mutation in a single amino acid on the kinetic properties reported here has been seen before. There are reports that almost all the psychrophilic character of some cold-adapted enzymes is due to a single amino-acid substitution. A single difference in the sequence at a subunit contact site was the cause of differences in the temperature– Km relationship or stability between closely related fish LDH [40]. In addition, nearly all the improvement in the catalytic efficiency of a mutated Vibrio marinus triosephos- phate isomerase was due to replacement of a completely conserved Ser in the phosphate binding helix by Ala in the psychrophilic enzyme [41]. There are, however, no structural features that can be correlated exclusively to cold adapta- tion. Structural explanations for cold adaptation can not be generalized. There is no single structural characteristic that accounts for the simultaneously appearing low stability and increased catalytic efficiency, proposed to be a consequence of high molecular flexibility. The origin of the increased enzyme activity and reduced stability lies in a particular region of the molecule rather than, for example, a general reduction in intramolecular interactions. A clear correlation seems to exist between cold adaptation and a reduction in the number of interactions between structural domains or subunits [42].

full activity after 8 h of incubation. The mutants showed a drop in residual activity after 24 h, and after 56 h of incubation between 54 and 74% of their maximum activity remained. The wild-type still had 95% residual activity after 56 h of incubation. Clearly, the mutated XIs were more sensitive to heat treatment at 70 (cid:176)C than the wild-type XI.

Fig. 4. Thermal stability of wild-type XI and mutated XIs. Metal-free enzyme preparations were incubated at 70 (cid:176)C in 200 mM triethanol- amine, pH 7.0, 10 mM MnCl2, and residual activities of aliquots were recorded as a function of time using xylose as a substrate: (e) wild- type; (d) M-1021; (,) M-1024; (j) M-1026.

D I S C U S S I O N

E372G/V379A (M-1021),

The goal of the present study was to generate efficient cold- adapted XIs from T. thermophilus, with improved kinetic properties at low temperatures. Random PCR mutagenesis was performed in the gene encoding the enzyme (xylA) and a mutant library was constructed. When the resulting proteins were screened, we obtained three cold-adapted mutants: E372G/F163L (M-1024) and E372G (M-1026), with higher kcat values than the wild-type XI for D-xylose at 60 (cid:176)C.

There is a close relationship between molecular flexibility and function. Thermophilic enzymes are rigid and require elevated temperatures in order to gain sufficient molecular flexibility for activity. Their molecular structure must thus be balanced between the requirements for stability and dynamics. We propose that the sequence changes underly- ing the adaptation of T. thermophilus XI mutants to temperatures lower than their optimal temperature, allow a higher degree of flexibility in areas that move during catalysis. Higher flexibility in these areas should increase kcat by reducing the energetic cost of a conformational change from the apoenzyme to the holoenzyme. By increasing kcat and Km, the catalytic efficiency of most cold-adapted enzymes increases, compared with the warm-adapted ones. kcat increases because of the ability of cold-adapted enzymes to reduce the free energy of activation compared with warm- adapted homologues. The increased Km is the result of a more flexible conformation [39]. Kinetic analysis demon- strated that the increase in the relative activity in the mutated XIs for xylose at low temperatures was indeed caused by an increase in kcat and not by a decrease in the Km value. This suggests that the mutant enzymes did not acquire higher affinity for the substrate than the wild-type enzyme at lower temperatures. The kcat/Km values for xylose for the mutated XIs only improved for M-1026. This was due to the large increase in the Km values for xylose. The

All mutations obtained were located on the enzyme surface, and not close to the active site. Amino-acid substitution distant from the catalytic centre or in the major substrate binding site of enzymes can lead to cold adaptation [38]. It has been proposed that variations in the enthalpy and entropy of conformational changes of impor- tance in binding and catalysis can be due to sequence changes outside the active sites. In the evolutionary adaptation of kcat and Km in response to acute temperature

(Eur. J. Biochem. 269)

(cid:211) FEBS 2002

162 A. Lo¨ nn et al.

R E F E R E N C E S

specific activity, or turnover number, kcat, reflects the catalytic potential at saturated substrate concentrations. The quantity, kcat/Km, is the catalytic efficiency that reflects the overall conversion of substrate to product. It has been suggested that the catalytic efficiency, kcat/Km, provides a better approximation of catalytic activity at physiological substrate concentrations, which are usually below satura- tion [43].

1. Hahn-Ha¨ gerdal, B., Wahlbom, F., Ga´ rdonyi, M., van Zyl, W.H., Cordero Otero, R. & Jo¨ nsson, L.J. (2001) Metabolic engineering of Saccharomyces cerevisae for xylose utilisation. Adv. Biochem. Eng Biotechnol. 73, 53–84.

2. Ladisch, M.R., Lin, K.W., Voloch, M. & Tsao, G.T. (1983) Process consideration in enzymatic hydrolysis of biomass. Enzyme Microb. Technol. 5, 82–102. 3. Ko¨ tter, P. & Ciriacy, M.

(1993) Xylose fermentation by Saccharomyces cerevisiae. Appl. Microbiol. Biotechnol. 38, 776– 783.

4. Tantirungkij, M., Nakashima, N., Seki, T. & Yoshida, T. (1993) Construction of xylose-assimilating Saccharomyces cerevisiae. J. Ferment. Bioeng. 75, 83–88.

In lignocellulosic hydrolysate the concentration of xylose can vary considerably. The concentration of xylose inside the cell, on the other hand, remains unknown, and is probably dependent on the xylose transporters. In natural xylose fermenting yeasts, the first xylose converting enzyme (XR) has a Km for xylose between 10 and 100 mM [44–46]. Recombinant S. cerevisiae expressing XR from P. stipitis has been shown to ferment xylose [3–5]. Therefore it is reasonable to assume that the mutated XIs with Km for xylose between 25 and 89 mM will be able to support a functional xylose metabolic pathway.

5. Walfridsson, M., Hallborn, J., Penttila¨ , M., Kera¨ nen, S. & Hahn- Ha¨ gerdal, B. (1995) Xylose-metabolizing Saccharomyces cerevisiae strains overexpressing the TKL1 and TAL1 genes encoding the pentose phosphate pathway enzymes transketolase and trans- aldolase. Appl. Environ. Microb. 61, 4184–4190. 6. Mermelstein, N.H. (1975) Immobilized enzymes produce high- fructose corn syrup. Food Technol. 29, 20–26.

For glucose, all mutated XIs had both higher kcat and kcat/Km values. These results indicate that we obtained improved kinetic constants at 60 (cid:176)C for D-glucose isomer- ization, but not to the same extent for D-xylose isomeriza- tion.

7. Ho, N., Stevis, P., Rosenfeld, S., Huang, J.J. & Tsao, G.T. (1983) Expression of the Escherichia coli isomerase gene by a yeast pro- moter. Biotechnol. Bioeng. Symp 13, 245–250.

8. Sarthy, A.V., McConaughy, B.L., Lobo, Z., Sundstrom, J.A., Furlong, C.E. & Hall, B.D. (1987) Expression of the Escherichia coli xylose isomerase gene in Saccharomyces cerevisiae. Appl. En- viron. Microbiol. 53, 1996–2000.

9. Amore, R., Wilhelm, M. & Hollenberg, C.P. (1989) The fermen- tation of xylose – an analysis of the expression of Bacillus and Actinoplanes xylose isomerase genes in yeast. Appl. Microbiol. Biotechnol. 30, 351–357.

10. Hallborn, J. (1995) Metabolic Engineering of Saccharomyces cerevisiae: Expression of Genes Involved in Pentose Metabolism. Lund University, Lund, Sweden.

11. Moes, C.J., Pretorius, I.S. & van Zyl, W.H. (1996) Cloning and expression of the Clostridium thermosulfurogenes D-xylose isom- erase gene (xylA) in Saccharomyces cerevisiae. Biotechnol. Lett. 18, 269–274.

Clearly, the mutated XIs were also thermally sensitive at 70 (cid:176)C, indicating that these mutations might confer ther- molabile characteristics on the enzyme. It has been reported previously that the thermostability of proteins can be altered by single amino-acid substitution [47,48], but it is not yet clear which these amino acids are [49]. It has also been suggested that higher catalytic efficiency in naturally occurring cold-adapted enzymes is associated with lower thermal stability, due to the higher molecular flexibility at lower temperatures [43,50,51]. The low stability at high temperatures is therefore regarded as a necessary conse- quence of cold adaptation. The reduced thermal stability of the mutated XIs is not a problem for xylose fermentation because fermentation occurs at moderate (30–40 (cid:176)C) tem- peratures and the yeast is continuously producing the enzyme during the fermentation process. However, the higher kcat at moderate temperatures is essential for obtaining xylose fermentation rates compatible with indus- trial processes [12].

All mutants showed a dramatic increase in Ki for xylitol, which is an inhibitor of XI. This may be a very important trait in the fermentation of xylose to ethanol, as S. cerevisiae produces xylitol from xylose via unspecific aldose reductases [52,53].

12. Walfridsson, M., Bao, X., Anderlund, M., Lilius, G., Bu¨ low, L. & Hahn-Ha¨ gerdal, B. (1996) Ethanolic fermentation of xylose with Saccharomyces cerevisiae harboring the Thermus thermophilus xylA gene, which expresses an active xylose (glucose) isomerase. Appl. Environ. Microbiol. 62, 4648–4651.

13. Suzuki, T., Yasugi, M., Arisaka, F., Yamagishi, A. & Oshima, T. (2001) Adaptation of a thermophilic enzyme, 3-isopropylmalate dehydrogenase, to low temperatures. Protein Eng. 14, 85–91. 14. Merz, A., Yee, M.C., Szadkowski, H., Pappenberger, G., Crameri, A., Stemmer, W.P., Yanofsky, C. & Kirschner, K. (2000) Improving the catalytic activity of a thermophilic enzyme at low temperatures. Biochemistry. 39, 880–889.

Together the improved kinetic properties at 60 (cid:176)C for the mutated XIs make them promising for xylose fermentation. To evaluate the physiological consequence of the changed kinetic properties of the wild-type and mutated xylA genes must, however, be expressed in S. cerevisiae.

15. Kano, H., Taguchi, S. & Momose, H. (1997) Cold adaptation of a mesophilic serine protease, subtilisin, by in vitro random muta- genesis. Appl. Microbiol. Biotechnol. 47, 46–51.

16. Taguchi, S., Ozaki, A. & Momose, H. (1998) Engineering of a cold-adapted protease by sequential random mutagenesis and a screening system. Appl. Environ. Microbiol. 64, 492–495.

A C K N O W L E D G E M E N T S

17. Taguchi, S., Ozaki, A., Nonaka, T., Mitsui, Y. & Momose, H. (1999) A cold-adapted protease engineered by experimental evo- lution system. J. Biochem. (Tokyo). 126, 689–693.

18. Lebbink, J.H., Kaper, T., Bron, P., van der Oost, J. & de Vos, W.M. (2000) Improving low-temperature catalysis in the hyper- thermostable Pyrococcus furiosus beta-glucosidase CelB by directed evolution. Biochemistry. 39, 3656–3665.

We would like to thank Jonas Fast for his technical assistance, and the Department of Biochemistry, Lund University, Sweden, for the use of the Storm 860(cid:210). This work was financially supported by The Swedish National Energy Administration (Energimyndigheten), the Swedish Foundation for International Cooperation in Research and Higher Education (STINT) and the National Research Foundation, South Africa (NRF). 19. Wintrode, P.L., Miyazaki, K. & Arnold, F.H. (2000) Cold adap- tation of a mesophilic subtilisin-like protease by laboratory evo- lution. J. Biol. Chem. 275, 31635–31640.

(cid:211) FEBS 2002

Mutant xylose isomerases (Eur. J. Biochem. 269) 163

20. Boyer, H.W. & Roulland-Dussoix, D. (1969) A complementation analysis of the restriction and modification of DNA in Escherichia coli. J. Mol. Biol. 41, 459–472. 37. Yamanaka, K. & Takahara, N. (1977) Purification and properties of D-xylose isomerase from Lactobacillus xylosus. Agric. Biol. Chem. 41, 1909–1915. 38. Feller, G. & Gerday, C. (1997) Psychrophilic enzymes: molecular basis of cold adaptation. Cell. Mol. Life Sci. 53, 830–841.

21. Shafikhani, S., Siegel, R.A., Ferrari, E. & Schellenbergger, V. (1997) Generation of large libraries of random mutants in Bacillus subtilis by PCR-based plasmid multimerization. Biotechniques. 23, 304–310.

39. Fields, P.A. & Somero, G.N. (1998) Hot spots in cold adaptation: localized increases in conformational flexibility in lactate dehy- drogenase A4 orthologs of antarctic notothenioid fishes. Proc. Natl Acad. Sci. USA 95, 11476–11481. 22. Sambrook, J., Fritsch, E.F. & Maniatis, T. (1989) Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 40. Somero, G.N. (1995) Proteins and temperature. Annu. Rev. Physiol. 57, 43–68.

41. Alvarez, M., Rentier-Delrue, F., Goraj, K. & Mainfroid, W. (1997) Vibrio marinus TIM: a cold adapted enzyme. Arch. Physiol. Biochem. 105, B17–B47. 42. Smala˚ s, A.O., Schroder Leiros, H.-K., Os, V. & Peder, N. (2000) 23. Bradford, M.M. (1976) A rapid and sensitive method for the quantitation of micro-organism quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. 24. Laemmli, U.K. (1970) Cleavage of structure proteins during assembly of the head of bacterophage T4. Nature (London). 227, 680–685. Cold adapted enzymes. Biotechn Annu. Rev. 6, 1–57. 43. Hochachka, P.W. & Somero, G.N. (1984) Biochemical Adaptation. Princeton University Press, Princeton, NJ.

25. Towbin, H., Staehelin, T. & Gordon, J. (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc. Natl Acad. Sci. USA 76, 4350–4354.

44. Mayr, P., Bruggler, K., Kulbe, K.D. & Nidetzky, B. (2000) D-Xylose metabolism by Candida intermedia: isolation and characterisation of two forms of aldose reductase with di(cid:128)erent coenzyme specificities. J. Chromatogr. B Biomed. Sci. Appl. 737, 195–202. 26. Callens, M., Kersters-Hilderson, H., van Opstal, O. & De Bruyne, C.K. (1986) Catalytic properties fo D-xylose isomerase from Streptomyces violaceoruber. Enzyme Microb. Technol. 8, 696–700.

45. Rizzi, M., Erlemann, P., Bui-Thanh, N.-A. & Dellweg, H. (1988) Xylose fermentation by yeasts 4. Purification and kinetic studies of xylose reductase from Pichia stipitis. Appl. Microbiol. Biotechnol. 29, 148–154.

27. Dekker, K., Yamagata, H., Sakaguchi, K. & Udaka, S. (1991) Xylose (glucose) isomerase gene from the thermophile Thermus thermophilus: cloning, sequencing, and comparison with other thermostable xylose isomerases. J. Bacteriol. 173, 3078–3083. 28. Fersht, A.R. (1984) Enzyme Structure and Mechanism, 2nd edn. 46. Verduyn, C., Frank, J., Van Dijken, J.P. & Sche(cid:128)ers, W.A. (1985) Multiple forms of xylose reductase in Pachysolen tannophilus CBS 4044. FEMS Microbiol. Lett. 30, 313–318. Freeman, San Francisco, CA. 29. Dixon, M. & Webb, E.C. (1964) Enzymes, 2nd edn. Longmans Green and Co Ltd, London. 47. Yutani, K., Ogasawara, K., Sugino, Y. & Matsushiro, A. (1977) E(cid:128)ect of a single amino acid substitution on stability of confor- mation of a protein. Nature 267, 274–275.

30. Dekker, K., Sugiura, A., Yamagata, H., Sakaguchi, K. & Udaka, S. (1992) E(cid:129)cient production of thermostable Thermus thermo- philus xylose isomerase in Escherichia coli and Bacillus brevis. Appl. Microbiol. Biotechnol. 36, 727–732.

48. Matsumura, M., Katakura, Y., Imanaka, T. & Aiba, S. (1984) Enzymatic and nucleotide sequence studies of a kanamycin- inactivating enzyme encoded by plasmid from thermophilic bacilli in comparison with that encoded by plasmid pUB110. J. Bacteriol. 160, 413–420.

31. Chang, C., Park, B.C., Lee, D.S. & Suh, S.W. (1999) Crystal thermostable xylose isomerases from Thermus structures of caldophilus and Thermus thermophilus: possible structural deter- minants of thermostability. J. Mol. Biol. 288, 623–634.

49. Imanaka, T., Shibazaki, M. & Takagi, M. (1986) A new way of enhancing the thermostability of proteases. Nature 324, 695–697. 50. Smala˚ s, A.O., Hiemstad, E.S., Hordvik, A., Willassen, N.P. & Male, R. (1994) Cold adaptation of enzymes: structural com- parison between salmon and bovine trypsins. Proteins 20, 149–166. 32. Collyer, C.A., Henrick, K. & Blow, D.M. (1990) Mechanism for aldose–ketose interconversion by D-xylose isomerase involving ring opening followed by a 1,2-hydride shift. J. Mol. Biol. 212, 211–235. 33. Chen, W.P. (1980) Glucose isomerase (a review) – part 2. Process Biochem. 15, 36–41.

51. Davail, S., Feller, G., Narinx, E. & Gerday, C. (1994) Cold adaptation of proteins – purification, characterization, and sequence of the heat-labile subtilisin from the antarctic psychro- phile Bacillus TA41. J. Biol. Chem. 269, 17448–17453. 34. Faber, C.K., Glasfeld, A., Tiraby, G., Ringe, D. & Petsko, G.A. (1989) Crystallographic studies of the mechanism of xylose isomerase. Biochemistry. 28, 7289–7297.

52. Kuhn, A., van Zyl, C., van Tonder, A. & Prior, B.A. (1995) Purification and partial characterization of an aldo-keto reductase from Saccharomyces cerevisiae. Appl. Environ. Microbiol. 61, 1580–1585.

35. Whitlow, M., Howard, A.J., Finzel, B.C., Poulos, T.L., Winborne, E. & Gilliland, G.L. (1991) A metal-mediated hydride shift mechanism for xylose isomerase based on the 1.6 A˚ Streptomyces rubiginosus structures with xylitol and D-xylose. Proteins 9, 153–173. 36. Yamanaka, K. (1969) Inhibition of D-xylose isomerase by penti- 53. Tra¨ (cid:128), K., Codero Otero, R.R., van Zyl, W.H. & Hahn-Ha¨ gerdal, B. (2000) Genetically engineered yeast and mutants thereof for the e(cid:129)cient fermentation of the lignocellulose hydrolysates. Interna- tional patent application, WO 99/54477. tols and D-lyxose. Arch. Biochem. Biophys. 131, 502–506.