Trehalose phosphorylase from Pleurotus ostreatus: Characterization and stabilization by covalent modification, and application for the synthesis of α,α-trehalose

Journal of Biotechnology 129 (2007) 140–150

Trehalose phosphorylase from Pleurotus ostreatus: Characterization and stabilization by covalent modification, and application for the synthesis of ␣,␣-trehalose Alexandra Schwarz a , Christiane Goedl a , Alphonse Minani a , Bernd Nidetzky a,b,∗ a

Institute of Biotechnology and Biochemical Engineering, Graz University of Technology, Petersgasse 12, A-8010 Graz, Austria b Research Centre Applied Biocatalysis, Petersgasse 14, A-8010 Graz, Austria Received 22 December 2004; received in revised form 7 July 2006; accepted 18 July 2006

Abstract Trehalose phosphorylase from the basidiomycete Pleurotus ostreatus (PoTPase) was isolated from fungal fruit bodies through ∼500-fold purification with a yield of 44%. Combined analyses by SDS-PAGE and gelfiltration show that PoTPase is a functional monomer of ∼55 kDa molecular mass. PoTPase catalyzes the phosphorolysis of ␣,␣-trehalose, yielding ␣-d-glucose 1-phosphate (␣Glc 1-P) and ␣-d-glucose as the products. The optimum pH of PoTPase for ␣,␣-trehalose phosphorolysis and synthesis is 6.8 and 6.2, respectively. Apparent substrate binding affinities (Km ) were determined at pH 6.8 and 30 ◦ C: ␣,␣-trehalose (79 mM); phosphate (3.5 mM); d-glucose (40 mM); ␣Glc 1-P (4.1 mM). A series of structural analogues of d-glucose were tested as glucosyl acceptors for the enzymatic reaction with ␣Glc 1-P, and robust activity with d-mannose (3%), 2-deoxy d-glucose (8%), 2-fluoro d-glucose (15%) and 2-keto-d-glucose (50%) was detected. Arsenate replaces, with 30% relative activity, phosphate in the conversion of ␣,␣-trehalose, and vanadate strongly inhibits the enzyme activity (Ki ∼4 ␮M). PoTPase has a half-life (t0.5 ) of approximately 1 h at 30 ◦ C in the absence of stabilizing compounds such as ␣,␣-trehalose (300 mM; t0.5 = 11.5 h), glycerol (20%, w/v; t0.5 = 6.5 h) or polyethylenglycol (PEG) 4000 (26%, w/v; t0.5 = 70 h). Covalent modification of PoTPase with activated derivatives of PEG 5000 increases the stability by up to 600-fold. Sucrose was converted to ␣,␣-trehalose in ∼60% yield using a coupled enzyme system composed of sucrose phosphorylase from Leuconostoc mesenteroides, glucose isomerase from Streptomyces murinus and the appropriately stabilized PoTPase. © 2006 Elsevier B.V. All rights reserved. Keywords: ␣,␣-trehalose synthesis; Substrate specificity; Enzyme stability; Polyethylene glycols; Chemical modification

Abbreviations: ␣Glc 1-P, ␣-d-Glucose 1-phosphate; LmSPase, sucrose phosphorylase from Leuconostoc mesenteroides; m-PEG, methoxypolyethylene glycol p-nitrophenyl carbonate-5000; PEG, polyethylenglycol-4000; PoTPase, trehalose phosphorylase from Pleurotus ostreatus; SC-PEG, succinimidyl carbonate polyethylene glycol monomethyl ether-5000; SmGIase, glucose isomerase from Streptomyces murinus; TNBS, 2,4,6-trinitrobenzenesulfonic acid; TPase, trehalose phosphorylase ∗ Corresponding author at: Institute of Biotechnology and Biochemical Engineering, Graz University of Technology, Petersgasse 12, A-8010 Graz, Austria. Tel.: +43 316 873 8400; fax: +43 316 873 8434. E-mail address: [email protected] (B. Nidetzky).

0168-1656/$ – see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.jbiotec.2006.11.022

A. Schwarz et al. / Journal of Biotechnology 129 (2007) 140–150

1. Introduction ␣,␣-Trehalose is a non-reducing disaccharide (␣d-glucopyranosyl ␣-d-glucopyranoside) that is widely distributed in nature and found in bacteria, yeast, fungi, insects and plants. In addition to serving a function as a storage carbohydrate much alike glycogen, the disaccharide is proposed to have number of other important physiological roles. Most significantly, trehalose is known for its outstanding ability to protect biomolecules, whole cells and tissues against various environmental stresses, such as desiccation, freezing, and osmotic pressure (Elbein, 1974; Elbein et al., 2003; Eleutherio et al., 1993; Page-Sharp et al., 1999; Strom and Kaasen, 1993; Van Laere, 1989). Recent studies indicate that in insects, trehalose has a role in anoxic tolerance and development (Chen et al., 2002). In bacteria, trehalose is involved in the formation of glycolipids such as the potently immunogenic trehalose 6,6 -dimycolate (cord factor) in mycobacteria (Chatterjee, 1997). Trehalose has high thermostability and a wide pHstability range. Due to the absence of reducing ends it is resistant to Maillard reactions in the presence of amino groups of proteins. Furthermore, ␣,␣-trehalose has high water-binding capacity (Crowe et al., 1984) which upon drying results in a local concentration of residual water next to compounds to which trehalose binds (Belton and Gil, 1994). These remarkable properties of ␣,␣-trehalose therefore suggest and let foresee a wide range of possible applications of the disaccharide, for example, as a stable sweetener in foodstuffs or as a stabilizer of liposomes and proteins in cosmetics and pharmaceuticals (Benaroudj et al., 2001; Colaco et al., 1992; Guo et al., 2000; Higashiyama, 2002; Ikegami et al., 1995; Roser, 1991; Singer and Lindquist, 1998). There are presently at least four biocatalytic routes reported for the synthesis of ␣,␣-trehalose which have been reviewed recently (Schiraldi et al., 2002). The biosynthetic system of trehalose 6-phosphate synthase and trehalose 6-phosphate phosphatase is widely distributed in prokaryotes and eukaryotes but not very well explored for the biotechnological production of trehalose (De Smet et al., 2000; Giaever et al., 1988; Vandercammen et al., 1989). A novel trehalose-synthesizing glycosyltransferase was recently discovered in Pyrococcus horikoshii (Ryu et

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al., 2005). An intriguing dual enzyme system displaying glycosyltransferase and hydrolase activities occurs in some mesophilic bacteria such as Arthrobacter sp. (Nakada et al., 1995) and in extremophilic archaea such as Sulfolobus sp. (Nakada et al., 1996). It converts starch or maltodextrins directly into trehalose by catalyzing a two-step reaction that involves a trehalosyl maltodextrin intermediate product. A trehalose synthase has been reported from various organisms such as Pimelobacter sp. (Nishimoto et al., 1996) and Thermus sp. (Koh et al., 2003; Tsusaki et al., 1997) which promotes intramolecular transglucosylation from maltose. Finally, trehalose phosphorylase (TPase) catalyzes glucosyl transfer to and from phosphate and uses d-glucose 1-phosphate as a glucosyl donor for the synthetic reaction with d-glucose to yield ␣,␣-trehalose (Eq. (1)). d-glucose 1-phosphate + d-glucose ↔ ␣, ␣-trehalose + phosphate

(1)

The thermodynamic equilibrium of Eq. (1) at neutral pH favours the synthesis of trehalose although under the physiological conditions phosphorolysis is clearly preferred (Eis and Nidetzky, 1999). The concentration of free glucose in the cell is normally low, and phosphate is always present in excess over glucose 1-phosphate. TPases have been purified and characterized from a range of bacteria, fungi and algae including Micrococcus varians (Kizawa et al., 1995), Catellatospora ferruginea (Aisaka et al., 1998), Grifola frondosa (Saito et al., 1998), Agaricus bisporus (Wannet et al., 1998), Pleurotus ostreatus (Kitamoto et al., 2000), Pleurotus sajor-caju (Han et al., 2003), Schizophyllum commune (Eis and Nidetzky, 1999), and Euglena gracilis (Belocopitow and Marechal, 1970; Marechal and Belocopitow, 1972). The known TPases are classified according to whether they convert ␣,␣trehalose into ␣-d-glucose 1-phosphate (␣Glc 1-P) or the corresponding ␤-anomer. It appears that the fungal enzymes are specific for ␣Glc 1-P. TPases can be used for trehalose production according to Eq. (1) or by using a coupled enzyme approach in which the substrates for TPase are generated in situ by the action of another phosphorylase. Maltose has been converted into trehalose using maltose phosphorylase and bacterial TPase both of which are specific for ␤d-glucose 1-phosphate (Aisaka et al., 2000). Sucrose

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would be another interesting disaccharide to be transformed into trehalose. The energy of the glycosidic bond of sucrose (27 kcal/mol) is much higher than that of ␣,␣-trehalose (1 kcal/mol), implying a substantial thermodynamic driving force for trehalose formation (Schiraldi et al., 2002). Because sucrose phosphorylase produces ␣Glc 1-P, fungal TPases are needed in a coupled enzymatic process that is based on sucrose. However, a major shortcoming of ␣Glc 1-P-specific TPases is the rapid loss of enzyme activity in buffer and under conditions of biocatalytic synthesis which has been reported by several authors (Eis and Nidetzky, 1999; Kitamoto et al., 2000; Klimacek et al., 1999; Marechal and Belocopitow, 1972; Wannet et al., 1998). Immobilization was shown to substantially improve the stability of TPase from S. commune (Klimacek et al., 1999) but for a process in which two or more enzymatic reactions are coupled (see later) the use of a stable soluble TPase is clearly preferred. The present work was undertaken with the aim of identifying a novel TPase biocatalyst that is useful for the conversion of sucrose into trehalose. We describe isolation and characterization of TPase from P. ostreatus (PoTPase) and report on a novel approach by covalent modification that very efficiently stabilizes the activity of the soluble enzyme. TPase with improved stability is applied for the production of ␣,␣-trehalose.

2. Materials and methods

tomyces murinus (SmGIase) was from Novo Nordisk. All other chemicals were of highest purity available and were purchased from Sigma-Aldrich. 2.2. Cultivation of P. ostreatus and enzyme production Fresh fruit bodies of P. ostreatus were obtained from the local market. The strain P. ostreatus was obtained from a culture collection maintained at this institute. The fungus was cultivated at 30 ◦ C in 1 l baffled shaken flasks using 400 ml media that contained 20 g/l d-glucose, 5 g/l peptone from soy bean, 5 g/l yeast extract, 1 g/l malt extract and 1 g/l K2 HPO4 . The pH of the medium was adjusted to 6.0. The flasks were inoculated with 40 ml of fungal mycelium grown for 3 days on the same medium, and incubated at a constant agitation rate of 120 rpm for 7 days. The fungal biomass was washed twice with 20 mM MES buffer, pH 6.8, and harvested by centrifugation at 8000 × g for 30 min at 4 ◦ C. The pellet was resuspended in (1:3, w/v) 20 mM MES buffer, pH 6.8, containing 4 mM EDTA, 2 mM DTT and 300 mM trehalose, and disrupted by repeated passage through a French press. Fruit bodies of P. ostreatus (2.5 kg) were suspended in 5 l of the same MES buffer, and disrupted for 15 min at 4 ◦ C, by using an Ultra Turrax T25 homogenizer (IKA Labortechnik, Germany), with an instrument setting of 24,000 rpm. Crude cell extracts were obtained by centrifugation at 7000 × g for 20 min followed by ultracentrifugation of the supernatant at 30,000 × g for 30 min at 4 ◦ C.

2.1. Materials 2.3. Purification of trehalose phosphorylase 2,4,6-Trinitrobenzenesulfonic acid (TNBS) and succinimidyl carbonate polyethylene glycol monomethyl ether-5000 (SC-PEG) were obtained from Calbiochem. Trehalose dihydrate and polyethylenglycol4000 (PEG) were from Roth, and methoxypolyethylene glycol p-nitrophenyl carbonate-5000 (m-PEG) was from Sigma. DEAE-Sepharose Fast Flow, Superose 12 (prep grade and HR 10/30), Sephacryl S-300, Superdex 75 (prep grade) and Mono Q (HR 5/5) were obtained from GE Healthcare, and Fractogel EMD-Phenyl I (S) was from Merck. Recombinant sucrose phosphorylase from Leuconostoc mesenteroides (LmSPase) was produced and purified by reported procedures (Schwarz and Nidetzky, 2006). Glucose isomerase from Strep-

Unless noted otherwise, 20 mM MES buffer, pH 6.8, containing 4 mM EDTA, 2 mM DTT and 300 mM trehalose was used. The clear cell extract (15 g of protein) was applied to a DEAE-Sepharose Fast Flow column (5 cm × 25 cm) and bound protein was eluted by using a step gradient, with steps of 0, 75, 150, 225 and 300 mM sodium chloride. PoTPase was eluted at 75 mM sodium chloride under these conditions. The enzyme solution was brought to 30% saturated ammonium sulfate and precipitated proteins were separated by ultracentrifugation. The clear supernatant was applied to a Fractogel EMD-Phenyl S column (2.6 cm × 16 cm), equilibrated with buffer containing 30% saturated

A. Schwarz et al. / Journal of Biotechnology 129 (2007) 140–150

ammonium sulfate. Bound protein was eluted by using a step gradient, with steps at 22.5, 15, 7.5 and 0% saturated ammonium sulfate. PoTPase was eluted at 0% saturated ammonium sulfate. Fractions containing phosphorylase activity were pooled and concentrated by ultrafiltration using Macrosep centrifugal concentrator tubes with a molecular mass cut-off of 30 kDa (Pall Filtron). A gelfiltration step was carried out using three columns (16 mm diameter) in series. The first column contained 140 ml of Sepharose 12, the second contained 180 ml of Sephacryl S-300, and the third contained 140 ml of Superdex 75. Each column was equilibrated with 20 mM MES buffer, pH 6.8, containing 300 mM trehalose, 200 mM NaCl and 2 mM DTT. The concentrated protein solution (5 ml) was loaded on to the first column and elution was carried out at a constant flow rate of 0.5 ml/min. After concentrating and desalting of the pooled fractions, the protein was loaded on to a Mono Q column (0.5 cm × 5 cm) equilibrated with 20 mM MES buffer, pH 5.6, containing 2 mM DTT and 300 mM trehalose. Elution of bound protein was carried out at room temperature and with a linear gradient from 0 to 300 mM sodium chloride in the same buffer. The active fractions were pooled, concentrated and brought to 20 mM MES buffer, pH 6.8, containing 4 mM EDTA, 2 mM DTT and 300 mM trehalose, by ultrafiltration. 2.4. Enzymatic assays and determination of molecular mass If not indicated otherwise, reported assays for enzyme activity and reaction products were used (Eis and Nidetzky, 1999). Trehalose phosphorylase activity was assayed in direction of phosphorolysis in a coupled assay with phosphoglucomutase and glucose 6-phosphate dehydrogenase. Initial rates of the phosphorolysis and synthesis reaction were determined by using discontinuous assays, and by measuring the release of ␣Glc 1-P and phosphate, respectively. dGlucose was determined by using NAD+ -dependent glucose dehydrogenase (Eis and Nidetzky, 1999). Protein concentration was determined by a dye-binding assay (Bio-Rad) referenced against BSA. The molecular mass of PoTPase was determined by SDS-PAGE and gelfiltration analysis, carried out on Superose 12 HR 10/30 (GE Healthcare) as described by Eis and Nidetzky (1999).

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2.5. Stabilization and chemical modification of PoTPase Stability of purified PoTPase was examined in the presence of trehalose (300 mM), glycerol (20%, w/v), sucrose (200 mM) and PEG (9 and 26%, w/v), respectively. The enzyme (1.9 U/ml; 1.09 U/mg) was incubated at 25 ◦ C in 20 mM MES buffer, pH 6.8, containing 2 mM DTT, and the change in phosphorylase activity with time was monitored. This was done by taking samples from the enzyme solution in regular intervals and diluting the sample immediately into the assay solution. Chemical modification of purified PoTPase (2.5 U/ml; 0.4 mg/ml) was performed at 4 ◦ C for 2 h, with low stirring (300 rpm) in 20 mM MES buffer, pH 6.8, containing 2 mM DTT. Different molar ratios of active PEG (SC-PEG or mPEG) and 55 kDa protein were tested: 3:1, 10:1 and 30:1. PEG-modified PoTPase was incubated at 25 ◦ C, and changes in activity were recorded over time as described above. The extent of modification of PoTPase was measured through colorimetric titration of free protein lysine groups with 2,4,6-trinitrobenzenesulfonic acid (TNBS) as described by Fields (1971). Typically, a ≥1000 molar excess of TNBS over protein (≈2.8 ␮M) was used, and the incubation time was 60 min at 30 ◦ C and pH 9.5. The adsorption of TNBS-modified protein was measured at 420 nm (molar extinction coefficient, ε = 19.2 mM−1 cm−1 ). 2.6. Trehalose synthesis and determination of reaction products Trehalose synthesis was carried out using PoTPase stabilized by the addition of 20% PEG or through modification with a 30-fold molar excess of m-PEG, respectively. Using d-glucose (25 mM) and ␣Glc 1P (25 mM) as substrates, PoTPase (1-3.5 U/ml) was incubated at 25 ◦ C in 20 mM MES buffer, pH 6.8, containing 2 mM DTT. Incubation at 25 ◦ C rather than 30 ◦ C was chosen to minimize enzyme inactivation. In another approach, the coupled enzyme system consisting of LmSPase, SmGIase and PoTPase was used to convert sucrose into ␣,␣-trehalose. The reaction mixtures contained 20 mM sucrose, 10 mM phosphate, 2.2 U/ml LmSPase, 7.15–29.5 U/ml SmGIase and 0.75–1.3 U/ml PoTPase, and were incubated at 25 ◦ C in 20 mM MES buffer, pH 6.8, containing

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10 mM MgCl2 , 2 mM CoCl2 and 2 mM MnCl2 . Reaction products of trehalose synthesis were analysed by HPLC (d-fructose, sucrose, ␣,␣-trehalose and dglucose) as well as by enzymatic (␣Glc 1-P) and colorimetric assays (phosphate) (Eis and Nidetzky, 1999; Klimacek et al., 1999). The HPLC system consisted of a Merck Model L 6200 A pump and a Model AS 2000 A autosampler. An Supelcosil LCNH2 (250 mm × 4.6 mm; 5 ␮m particle size) column (Supelco), together with a high-sensitivity refractive index detector (Erma Optical Works), were used. The column was kept at 35 ◦ C, and acetonitrile:potassium phosphate buffer (75:25, v/v) was used as eluent at a flow rate of 1.5 ml/min.

PoTPase. It is similar to the reported specific activity of TPase from P. sajor-caju (Han et al., 2003), but about four-fold those published for purified TPase from G. frondosa (Saito et al., 1998) and A. bisporus (Wannet et al., 1998). Substantial losses of original enzyme activity, about 36% during gelfiltration and anionic exchange with Mono Q, are documented in Table 1. They were mainly caused by the required concentration and desalting steps. SDS-PAGE of the purified enzyme gave a single protein band with an apparent molecular mass of 55 kDa (Fig. 1A). The molecular mass of native PoTPase was determined by analytical gel filtration to be 58.3 kDa (Fig. 1B). Interestingly, while a dimeric structure was suggested for PoTPase in earlier work (Kitamoto et al., 2000), the enzyme purified in this study is proposed to exist as functional monomer.

3. Results and discussion 3.1. Purification of PoTPase

3.2. PoTPase is an anomeric configuration-retaining enzyme

After cultivation of P. ostreatus for 7 days in shaken flasks at 30 ◦ C, approximately 22 g/l wet mycelial biomass were obtained. The specific trehalose phosphorylase activity in cell extracts prepared from the mycelia was 0.13 U/mg. Comparable specific activities (0.03 U/mg) were found in cell extracts of commercial fruit bodies of P. ostreatus. For reasons of convenient availability the fruit bodies were selected as the source of PoTPase throughout this work. PoTPase was purified 470-fold by using a fourstep protocol, which is summarized in Table 1. The overall yield was 44% which is high in comparison to results obtained by other authors (Kitamoto et al., 2000), and the specific activity of the purified enzyme was 15.7 U/mg of protein. This value agrees with findings of Kitamoto et al. (2000) who first reported on

The coupled enzymatic assay of trehalose phosphorylase activity which is described under Section 2 is specific for the detection of ␣Glc 1-P, suggesting that the ␣-anomer of glucose 1-phosphate is the product of PoTPase-catalyzed phosphorolysis of ␣,␣trehalose. Production of ␣Glc 1-P upon the enzymatic conversion of ␣,␣-trehalose was confirmed by HPLC analysis referenced to an authentic standard of the phospho-sugar, according to Eis and Nidetzky (1999). Enzymatic assays were performed in the direction of disaccharide synthesis at pH 6.8 (20 mM MES buffer) and 30 ◦ C using ␤-d-glucose 1-phosphate (200 mM) as a substrate replacing ␣Glc 1-P. Within limits of detection of the analytical procedures for measuring phosphate release (which was 0.1% of the basal activity recorded with ␣Glc 1-P and d-glucose at concentra-

Table 1 Purification of PoTPase Purification step

Total volume (ml)

Total activity (U)

Crude cell extract DEAE-Sepharose Fractogel EMD-Phenyl Superdex 75, Sephacryl S-300, Superose 12 Mono Qa

8000 502 218 86

496 427 399 325

52

220

All steps were carried out at 4 ◦ C, unless otherwise specified. a Carried out at room temperature.

Specific activity (U/mg) 0.033 0.649 2.04 9.8 15.7

Yield (%)

Purification (-fold)

100 86 80 66

1 19 61 297

44

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Fig. 1. Determination of the molecular mass of purified PoTPase using analysis by SDS-PAGE (A) and gelfiltration (B). (A) Lane 1, molecular mass standard proteins; lane 2, PoTPase after purification on Mono Q. Staining of protein bands was done with Coomassie Blue. (B) Partition coefficients (Kav ) of the standard proteins (aldolase, 158 kDa; hexokinase, 100 kDa; BSA, 68 kDa; egg albumin, 43 kDa; ␤-lactoglobulin, 35 kDa; myoglobulin, 17.6 kDa and cytocrome c, 12.4 kDa) were calculated according to Kav = (Ve − V0 )/(Vt − V0 ) (Ve , elution volume; V0 , void volume; Vt , total volume of the gel bed) and plotted against the corresponding molecular masses. The molecular mass of purified PoTPase was calculated using the calibration curve equation: log(Mr ) = 3.032 − 3.230Kav .

tions of 200 mM), ␤-d-glucose 1-phosphate was not turned over by the enzyme. In the phosphorolysis direction under the same conditions, ␣,␤-trehalose and ␤,␤-trehalose were inactive in the presence of 20 mM phosphate as were sucrose, cellobiose, lactose, maltose, lactulose and melibiose. In the presence of ␣Glc 1-P (200 mM), ␤-d-glucose was converted by PoTPase with a clear time lag of several minutes that was not observed when an equilibrated mixture of the ␣ and ␤anomers was used as the glucosyl acceptor (for the used methods, see Eis et al. (1998)). Therefore, ␤-d-glucose is not a substrate of PoTPase. The results emphasize the high specificity of PoTPase for reaction with distinct (␣) anomeric forms of the substrates.

at pH 6.8 (20 mM MES buffer) under conditions in which one substrate was varied and the other substrate was present at a constant concentration that was saturating at the steady state. Data were fitted to the Michaelis–Menten equation using non-linear regression, and apparent kinetic parameters are summarized in Table 2. Comparison of values for Km reveal 10–20fold stronger apparent binding of phosphate and ␣Glc 1-P than trehalose and glucose. This was also noted in prior studies of TPase from S. commune (Eis and Nidetzky, 1999). A general observation of PoTPase and related TPase enzymes is that the catalytic efficiencies are low in comparison with other enzymes (Wolfenden and Snider, 2001) and far below a value of 108 M−1 s−1 expected for diffusion-limited enzymatic

3.3. Enzyme kinetics and substrate specificity The pH-dependence of PoTPase activity was measured in the pH range 4.8–7.8, using 20 mM MES buffer between pH 4.8 and 6.5, and 20 mM TES buffer between pH 6.6 and 7.8. Each buffer contained 20% (w/v) glycerol. Results are shown in Fig. 2, indicating that the optimum pH for phosphorolysis is 6.8–7.2 whereas that for synthesis is 5.8–6.4. As the measurement time was <5 min in the assays, the observed pH-dependence probably reflects the variation of the enzymatic rate with the pH and is not strongly clouded by pH effects on enzyme stability. Initial rates of trehalose phosphorolysis and synthesis were recorded

Fig. 2. pH profiles for ␣,␣-trehalose phosphorolysis (䊉) and synthesis ( ) catalyzed by PoTPase.

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Table 2 Apparent kinetic parameters for PoTPase Reaction type

Substrate

Km (mM)

kcat (s−1 )

kcat /Km (M−1 s−1 )

Synthesis

d-Glucose ␣Glc 1-P

40 ± 10 4.1 ± 0.4

16.8 ± 1.2 16.0 ± 0.4

420 3902

Phosphorolysis

␣,␣-Trehalose Phosphate

79 ± 11 3.5 ± 0.4

18.3 ± 0.6 17.6 ± 0.6

232 5029

Initial rates were recorded at 30 ◦ C in 20 mM MES buffer, pH 6.8. Calculation of kcat was based on a molecular mass of PoTPase of 55 kDa. Results are means ± standard error.

reactions. Interestingly, the Km values 40 and 4.1 mM for d-glucose and ␣Glc 1-P, respectively, were significantly lower than the values of 505 and 38 mM for the same substrates previously reported by Kitamoto et al. (2000). Differences in work-up procedures of the fungal fruit bodies may be responsible for the observed variation in structural and functional properties of PoTPase found here and by other authors (Kitamoto et al., 2000). The specificity of PoTPase for glucosyl acceptors that replace d-glucose in the direction of disaccharide synthesis was determined by recording initial rates of phosphate release in the presence of ␣Glc 1-P (200 mM). Discontinuous assays were used, and enzyme-catalyzed product formation by the enzyme (0.2 U/ml) was monitored over approximately 15 min. PoTPase was active with d-mannose (kcat = 0.42 s−1 ; kcat /Km = 2.0 M−1 s−1 ), 2-deoxy d-glucose (kcat = 0.27 s−1 ; kcat /Km = 6.8 M−1 s−1 ), 2-fluoro d-glucose (kcat = 0.89 s−1 ; kcat /Km = 52 M−1 s−1 ) and 2-ketoglucose (kcat = 4.5 s−1 ; kcat /Km = 97 M−1 s−1 ). The decrease in catalytic efficiency, compared to kcat /Km for reaction with d-glucose, when the original C2(R) hydroxyl is replaced or switches its stereochemical position in the substrate analogues suggests that one or more functional groups on the enzyme and the 2-OH interact, probably by hydrogen bonding, to provide transition state stabilization. The observed loss of binding energy (G#) can be calculated by using the relationship G# = − RT ln(kcat /Km )2-deoxy-Glc /(kcat /Km )Glc where R is the gas constant and T is the temperature in Kelvin. G# has a value of approximately 10.4 kJ/mol which is within the expected range for the change in G# upon removal of a hydrogen bond. No measurable activity was found with darabinose, d-fructose, d-fucose, d-galactose, d-lyxose, d-mannoheptaose, d-ribose and d-xylose.

In the direction of phosphorolysis of ␣,␣-trehalose (200 mM), phosphate can be replaced by arsenate (kcat = 1.15 s−1 ; kcat /Km = 550 M−1 s−1 ). As described previously (Eis and Nidetzky, 1999; Nidetzky et al., 2000) the resultant ␣-d-glucose 1-arsenate is unstable and hydrolyses spontaneously so that the overall reaction is the conversion of 1 mole ␣,␣-trehalose into 2 moles d-glucose. The reported kinetic parameters are based on measurements of the rate of d-glucose formation and corrected for the stoichiometry of the reaction. Vanadate has been described as a partial transition state mimic of the reaction catalyzed by S. commune TPase (Nidetzky and Eis, 2001). A steady-state kinetic analysis for the inhibition of PoTPase by vanadate was carried out by recording initial rates of phosphorolysis in the absence and presence of several concentrations of the oxyanion in the range 0.5–20 ␮M. The results reveal that vanadate is a strong competitive inhibitor against phosphate, and at saturating concentrations of ␣,␣-trehalose it binds with inhibition constants of approximately 4.0 ␮M. In conclusion, the pattern of substrate specificity and inhibitor binding affinity of PoTPase is similar to that observed for TPase from S. commune (Nidetzky and Eis, 2001). Moreover, both enzymes exhibit significant similarity in subunit size (∼60 kDa) and quaternary structure (monomer), thus emphasizing their close relationship. 3.4. Stability of PoTPase Purified PoTPase is a very unstable protein, which has a half-life (t0.5 ) in 20 mM MES buffer, pH 7.0, at 25 ◦ C of no more than approximately 1.3 h. Like TPase from S. commune (Eis and Nidetzky, 1999), PoTPase can be stabilized on addition of ␣,␣-trehalose (300 mM) which yields a t0.5 value of 11.5 h. In the presence of 20% (w/v) glycerol, and 9 and 26% (w/v)

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PEG-4000, t0.5 is 6.5, 16 and 70 h, respectively. It is noteworthy that phosphate (20–50 mM) does not exercise a stabilizing effect on the activity of PoTPase which is contrast to the significant extra stability conferred by added phosphate to TPases from A. bisporus (Wannet et al., 1998), E. gracilis (Marechal and Belocopitow, 1972) and C. ferruginea (Aisaka and Masuda, 1995). We achieved large improvements in PoTPase stability by covalent protein modification with activated derivatives of PEG-5000. The enzyme was incubated for up to 2 h at 4 ◦ C with m-PEG or SC-PEG at three different molar ratios of PEG and protein: 3, 10, and 30. The progress and the final extent of the modification was monitored by titrating the available lysyl groups in PoTPase. Complete derivatization of all 12 lysines that are reactive towards TNBS in native PoTPase requires a molar excess of m-PEG of 10 or greater. Under all the different conditions used, the modification of PoTPase was accompanied by an only moderate decrease in enzyme activity (≤13%). The stability of the modified enzyme was examined in comparison to that of native PoTPase in the presence of 300 mM trehalose, and results are shown in Fig. 3. Modification with m-PEG yields better stabilization than modification with SC-PEG. The best improved stability of PoTPase is found after reaction with 30-fold molar excess of m-PEG. The value of t0.5 for the m-PEGmodified enzyme is increased by factors of 55 and 600 in comparison to corresponding t0.5 values of

Fig. 3. Half-lives of native and modified PoTPase at 25 ◦ C in 20 mM MES buffer, pH 6.8, containing 300 mM trehalose. Chemical modification of PoTPase was performed as described in Section 2.

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Fig. 4. Time course of ␣,␣-trehalose synthesis from d-glucose and ␣Glc 1-P catalyzed by PoTPase stabilized with 20% PEG ((䊉) 3.5 U/ml) and m-PEG (( ) 1 U/ml). The reaction mixtures, which contained ␣Glc 1-P (25 mM), d-glucose (25 mM) and PoTPase were incubated in 20 mM MES buffer, pH 6.8, containing 2 mM DTT, at 25 ◦ C for the indicated periods, and the amounts of ␣,␣-trehalose were determined by HPLC. Keq represents the thermodynamic equilibrium constant for ␣,␣-trehalose synthesis under these conditions.

native PoTPase in the presence and absence of 300 mM trehalose. 3.5. Application of stabilized PoTPase for the synthesis of α,α-trehalose A number of studies have shown that the equilibrium for phosphorolysis of ␣,␣-trehalose is on the side of the reactants (Eis and Nidetzky, 1999; Kitamoto et al., 2000; Marechal and Belocopitow, 1972). Equilibrium constants (Keq ) of 4–10 at pH 7.0 and 8–17 at pH 6.4 have been reported, suggesting that: (1) the synthesis of ␣,␣-trehalose should generally proceed with acceptable yields, and (2) decreased pH values will increase the attainable level of substrate conversion into disaccharide. Fig. 4 displays typical time courses of ␣,␣-trehalose production using native enzyme and mPEG-modified PoTPase, and illustrates the limitations in product yield that are due to Keq . Under the conditions used, approximately 75% of the substrate reacts to give ␣,␣-trehalose. Note that the enzymatic reaction of PoTPase derivatized with m-PEG is not intrinsically slower than the corresponding reaction of the native enzyme stabilized with 20% PEG but reflects different levels of initial enzyme activity used in the two experiments reported in Fig. 4. The modified TPase shows good retention of activity after 20 h of incuba-

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Fig. 5. Coupled enzyme system for ␣,␣-trehalose synthesis from sucrose. At pH 6.8, equilibrium constants for sucrose conversion (Keq ≈ 44; Goedl et al., this issue), trehalose synthesis (Keq ≈ 6.2; Eis and Nidetzky, 1999) and interconversion of d-fructose and d-glucose (≈1) imply an overall reaction equilibrium constant of ≈275.

tion, suggesting that it is much more a useful biocatalyst than the native enzyme. The fact that addition of large amounts of glycerol or PEG which is required for stability of the native enzyme is eliminated in PoTPase derivatized with m-PEG is considered to be advantageous for both synthesis and downstream processing of ␣,␣-trehalose. The coupled enzymatic process pursued in this study is shown in Fig. 5. In the presence of SmGIase, sucrose is converted into trehalose through the combined action of LmSPase and PoTPase. The enzymatic reaction involves recycling of phosphate which can therefore be present at levels substoichiometric to the initial concentration of sucrose. Fig. 6A shows a time course of ␣,␣-trehalose production under conditions

in which the activity of PoTPase was stabilized by the addition of 20% (w/v) PEG-4000. After approximately 70 h, sucrose is consumed exhaustively but only about 8 mM ␣,␣-trehalose has been produced, corresponding to a yield of 40%. The accumulation of d-glucose up to concentrations of approximately 6 mM while ␣Glc 1-P is also present at the same or even higher concentrations indicates that PoTPase is essentially inactive after approximately 30–40 h. If the enzyme were active, further formation of ␣,␣trehalose would be expected from the value of Keq . The presence of high concentrations of d-fructose (13 mM) from 30 h until the end of the reaction indicates that isomerization of d-fructose does not proceed efficiently. This may indicate that under the conditions used the activity of the isomerase is limiting, or the thermodynamic equilibrium for the conversion of d-fructose into d-glucose provides insufficient driving force. The sharp increase in the concentration of phosphate after 45 h when the synthesis rate of ␣,␣-trehalose is essentially zero requires explanation because it cannot be due to the canonical phosphorylase activity. However, it is known from literature and experience (Schwarz and Nidetzky, 2006) that LmSPase displays significant hydrolase activity towards ␣Glc 1P which corresponds to approximately 1–2% of the glucosyltransferase activity of this enzyme with the same substrate. Note that enzymatic ‘error hydrolysis’ is not due to contamination but reflects the true catalytic promiscuity of LmSPase. It is thus conceivable (although not proven by the present results) that

Fig. 6. Time course of ␣,␣-trehalose synthesis from sucrose catalyzed by LmSPase, SmGIase and PoTPase, stabilized with: (A) 20% PEG and (B) m-PEG. The reaction mixtures, which contained sucrose (20 mM), phosphate (10 mM), LmSPase (2.2 U/ml), SmGIase (A: 7.15 U/ml, B: 29.5 U/ml) and PoTPase (A: 1.3 U/ml, B: 0.75 U/ml) were incubated in 20 mM MES buffer, pH 6.8, containing 10 mM MgCl2 , 2 mM CoCl2 and 2 mM MnCl2 , at 25 ◦ C for the indicated periods, and the amounts of d-glucose (dashed line), phosphate (dotted line), sucrose (䊉), ␣,␣-trehalose ( ), ␣Glc 1-P () and d-fructose () were measured as described in Section 2.

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formation of phosphate uncoupled from disaccharide synthesis reflects the hydrolytic conversion of ␣Glc 1-P by LmSPase. A decrease in the concentration of ␣Glc 1-P from 10 mM at 46 h to 4 mM at 70 h is in accordance with this notion, because trehalose is not produced at appreciable rates in this late phase of the reaction. The results in Fig. 6A were used as a point of departure for some optimization of the coupled enzymatic reaction, as shown in Fig. 6B. The m-PEG-modified PoTPase was employed to improve the stability of the trehalose-producing enzyme. In addition, the activity of SmGIase was increased four-fold in comparison to the reaction discussed above to achieve accelerated isomerization of d-fructose. The results depicted in Fig. 6B reveal that in comparison to data shown in Fig. 6A, there is substantial increase in the rate of sucrose conversion concomitant with an improvement of the trehalose yield from 40% to approximately 60%. It was also possible to maintain a comparably low level of d-glucose and ␣Glc 1-P, reflecting the persistence of the activity of PoTPase after modification with m-PEG. After sucrose has been nearly depleted (>8 h), the increase in the concentration of phosphate parallels the formation of ␣,␣-trehalose which is line with the expected action of PoTPase. The rate of trehalose formation decreases sharply after about 3 h when the level of disaccharide product reaches a value of about 8 mM, suggesting inhibition of PoTPase activity. We speculate that as in TPase from S. commune (Eis and Nidetzky, 2002) phosphate and glucose can add to PoTPase to form a relatively stable and abortive ternary complex. Thermodynamic consideration for the overall conversion of sucrose into trehalose (Fig. 5, legend) suggest that a yield for trehalose near 100% should be possible at equilibrium (e.g., Saito et al., 1998). Reaction times in panel B of Fig. 6 have obviously been too short to achieve equilibration. In conclusion, therefore, modified PoTPase seems to be a promising biocatalyst for the synthesis of trehalose in single or multi-enzyme processes. The production of trehalose from sucrose could be further enhanced by studying effects of the substrate concentrations and in particular the ratio of enzyme activities on conversion efficiency. The present evidence suggests that the activity of TPase should be increased to improve productivity.

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Acknowledgements Financial support from the Austrian Science Funds (P-15118 and P18038-B09 to B.N., and the DK Molecular Enzymology W901-B05) is gratefully acknowledged. Elisabeth Bauer is thanked for assistance in the determination of kinetic parameters.

References Aisaka, K., Masuda-Kato, T., Chikamune, T., Kamitori, K., Uosaki, Y., Saito, Y., 2000. Enzymatic synthesis of novel disaccharides using disaccharide phosphorylases. J. Biosci. Bioeng. 90, 208–213. Aisaka, K., Masuda, T., 1995. Production of trehalose phosphorylase by Catellatospora ferruginea. FEMS Microbiol. Lett. 131, 47–51. Aisaka, K., Masuda, T., Chikamune, T., Kamitori, K., 1998. Purification and characterization of trehalose phosphorylase from Catellatospora ferruginea. Biosci. Biotechnol. Biochem. 62, 782–787. Belocopitow, E., Marechal, L.R., 1970. Trehalose phosphorylase from Euglena gracilis. Biochim. Biophys. Acta 198, 151–154. Belton, P.S., Gil, A.M., 1994. IR and Raman spectroscopic studies of the interaction of trehalose with hen egg white lysozyme. Biopolymers 34, 957–961. Benaroudj, N., Lee, D.H., Goldberg, A.L., 2001. Trehalose accumulation during cellular stress protects cells and cellular proteins from damage by oxygen radicals. J. Biol. Chem. 276, 24261–24267. Chatterjee, D., 1997. The mycobacterial cell wall: structure, biosynthesis and sites of drug action. Curr. Opin. Chem. Biol. 1, 579–588. Chen, Q., Ma, E., Behar, K.L., Xu, T., Haddad, G.G., 2002. Role of trehalose phosphate synthase in anoxia tolerance and development in Drosophila melanogaster. J. Biol. Chem. 277, 3274–3279. Colaco, C., Sen, S., Thangavelu, M., Pinder, S., Roser, B., 1992. Extraordinary stability of enzymes dried in trehalose: simplified molecular biology. Biotechnology 10, 1007–1011. Crowe, J.H., Whittam, M.A., Chapman, D., Crowe, L.M., 1984. Interactions of phospholipid monolayers with carbohydrates. Biochim. Biophys. Acta 769, 151–159. De Smet, K.A., Weston, A., Brown, I.N., Young, D.B., Robertson, B.D., 2000. Three pathways for trehalose biosynthesis in mycobacteria. Microbiology 146, 199–208. Eis, C., Albert, M., Dax, K., Nidetzky, B., 1998. The stereochemical course of the reaction mechanism of trehalose phosphorylase from Schizophyllum commune. FEBS Lett. 440, 440–443. Eis, C., Nidetzky, B., 1999. Characterization of trehalose phosphorylase from Schizophyllum commune. Biochem. J. 341, 385–393. Eis, C., Nidetzky, B., 2002. Substrate-binding recognition and specificity of trehalose phosphorylase from Schizophyllum commune examined in steady-state kinetic studies with deoxy and

150

A. Schwarz et al. / Journal of Biotechnology 129 (2007) 140–150

deoxyfluoro substrate analogues and inhibitors. Biochem. J. 363, 335–340. Elbein, A.D., 1974. The metabolism of ␣,␣-trehalose. Adv. Carbohydr. Chem. Biochem. 30, 227–256. Elbein, A.D., Pan, Y.T., Pastuszak, I., Carroll, D., 2003. New insights on trehalose: a multifunctional molecule. Glycobiology 13, 17–27. Eleutherio, E.C., Araujo, P.S., Panek, A.D., 1993. Protective role of trehalose during heat stress in Saccharomyces cerevisiae. Cryobiology 30, 591–596. Fields, R., 1971. The measurement of amino groups in proteins and peptides. Biochem. J. 124, 581–590. Giaever, H.M., Styrvold, O.B., Kaasen, I., Strom, A.R., 1988. Biochemical and genetic characterization of osmoregulatory trehalose synthesis in Escherichia coli. J. Bacteriol. 170, 2841–2849. Guo, N., Puhlev, I., Brown, D.R., Mansbridge, J., Levine, F., 2000. Trehalose expression confers desiccation tolerance on human cells. Nat. Biotechnol. 18, 168–171. Han, S.E., Kwon, H.B., Lee, S.B., Yi, B.Y., Murayama, I., Kitamoto, Y., Byun, M.O., 2003. Cloning and characterization of a gene encoding trehalose phosphorylase (TP) from Pleurotus sajorcaju. Protein Expr. Purif. 30, 194–202. Higashiyama, T., 2002. Novel functions and applications of trehalose. Pure Appl. Chem. 74, 1263–1269. Ikegami, S., Kubota, M., Sugimoto, T. and Miyake, T. 1995. Thermostable trehalose-releasing enzyme, and its preparation and uses. European Patent 0688866. Kitamoto, Y., Osaki, N., Tanaka, H., Sasaki, H., Mori, N., 2000. Purification and properties of ␣-glucose 1-phosphate-forming trehalose phosphorylase from basidiomycete, Pleurotus ostreatus. Mycoscience 41, 607–613. Kizawa, H., Miyagawa, K., Sugiyama, Y., 1995. Biosci. Biotechnol. Biochem. 59, 1908–1912. Klimacek, M., Eis, C., Nidetzky, B., 1999. Continuous production of ␣,␣-trehalose by immobilised fungal trehalose phosphorylase. Biotechnol. Tech. 13, 243–248. Koh, S., Kim, J., Shin, H.J., Lee, D., Bae, J., Kim, D., Lee, D.S., 2003. Mechanistic study of the intramolecular conversion of maltose to trehalose by Thermus caldophilus GK24 trehalose synthase. Carbohydr. Res. 338, 1339–1343. Marechal, L.R., Belocopitow, E., 1972. Metabolism of trehalose in Euglena gracilis. J. Biol. Chem. 247, 3223–3228. Nakada, T., Ikegami, S., Chaen, H., Kubota, M., Fukuda, S., Sugimoto, T., Kurimoto, M., Tsujisaka, Y., 1996. Purification and characterization of thermostable maltooligosyl trehalose synthase from the thermoacidophilic archaebacterium Sulfolobus acidocaldarius. Biosci. Biotechnol. Biochem. 60, 263–266. Nakada, T., Maruta, K., Tsusaki, K., Kubota, M., Chaen, H., Sugimoto, T., Kurimoto, M., Tsujisaka, Y., 1995. Purification and properties of a novel enzyme, maltooligosyl trehalose synthase, from Arthrobacter sp. Q36. Biosci. Biotechnol. Biochem. 59, 2210–2214. Nidetzky, B., Eis, C., 2001. ␣-Retaining glucosyl transfer catalysed by trehalose phosphorylase from Schizophyllum commune:

mechanistic evidence obtained from steady-state kinetic studies with substrate analogues and inhibitors. Biochem. J. 360, 727–736. Nidetzky, B., Eis, C., Albert, M., 2000. Role of non-covalent enzyme-substrate interactions in the reaction catalysed by cellobiose phosphorylase from Cellulomonas uda. Biochem. J. 351, 649–659. Nishimoto, T., Nakano, M., Nakada, T., Chaen, H., Fukuda, S., Sugimoto, T., Kurimoto, M., Tsujisaka, Y., 1996. Purification and properties of a novel enzyme, trehalose synthase, from Pimelobacter sp. R48. Biosci. Biotechnol. Biochem. 60, 640–644. Page-Sharp, M., Behm, C.A., Smith, G.D., 1999. Involvement of the compatible solutes trehalose and sucrose in the response to salt stress of a cyanobacterial Scytonema species isolated from desert soils. Biochim. Biophys. Acta 1472, 519–528. Roser, B., 1991. Trehalose, a new approach to premium dried foods. Trends Food Sci. 2, 166–169. Ryu, S.I., Park, C.S., Cha, J., Woo, E.J., Lee, S.B., 2005. A novel trehalose-synthesizing glycosyltransferase from Pyrococcus horikoshii: molecular cloning and characterization. Biochem. Biophys. Res. Commun. 329, 429–439. Saito, K., Kase, T., Takahashi, E., Horinouchi, S., 1998. Purification and characterization of a trehalose synthase from the basidiomycete Grifola frondosa. Appl. Environ. Microbiol. 64, 4340–4345. Schiraldi, C., Di Lernia, I., De Rosa, M., 2002. Trehalose production: exploiting novel approaches. Trends Biotechnol. 20, 420–425. Schwarz, A., Nidetzky, B., 2006. Asp-196–>Ala mutant of Leuconostoc mesenteroides sucrose phosphorylase exhibits altered stereochemical course and kinetic mechanism of glucosyl transfer to and from phosphate. FEBS Lett. 580, 3905–3910. Singer, M.A., Lindquist, S., 1998. Multiple effects of trehalose on protein folding in vitro and in vivo. Mol. Cell 1, 639–648. Strom, A.R., Kaasen, I., 1993. Trehalose metabolism in Escherichia coli: stress protection and stress regulation of gene expression. Mol. Microbiol. 8, 205–210. Tsusaki, K., Nishimoto, T., Nakada, T., Kubota, M., Chaen, H., Fukuda, S., Sugimoto, T., Kurimoto, M., 1997. Cloning and sequencing of trehalose synthase gene from Thermus aquaticus ATCC33923. Biochim. Biophys. Acta 1334, 28–32. Van Laere, A., 1989. Trehalose, reserve and/or stress metabolite? FEMS Microbiol. Lett. 63, 201–209. Vandercammen, A., Francois, J., Hers, H., 1989. Characterization of trehalose-6-phosphate synthase and trehalose-6-phosphate phosphatase of Saccharomyces cerevisiae. Eur. J. Biochem. 182, 613–620. Wannet, W.J., Op den Camp, H.J., Wisselink, H.W., van der Drift, C., Van Griensven, L.J., Vogels, G.D., 1998. Purification and characterization of trehalose phosphorylase from the commercial mushroom Agaricus bisporus. Biochim. Biophys. Acta 1425, 177–188. Wolfenden, R., Snider, M.J., 2001. The depth of chemical time and the power of enzymes as catalysts. Acc. Chem. Res. 34, 938–945.