- Email: [email protected]
Purification and characterization of trehalose phosphorylase from the commercial mushroom Agaricus bisporus
Biochimica et Biophysica Acta 1425 (1998) 177^188
Puri¢cation and characterization of trehalose phosphorylase from the commercial mushroom Agaricus bisporus Wim J.B. Wannet a , Huub J.M. Op den Camp a; *, Hendrik W. Wisselink a , Chris van der Drift a , Leo J.L.D. Van Griensven b , Godfried D. Vogels a a
Department of Microbiology, Faculty of Science, University of Nijmegen, Toernooiveld 1, NL-6525 ED Nijmegen, Netherlands b Mushroom Experimental Station, Postbus 6042, NL-5960 AA Horst, Netherlands Received 30 March 1998; revised 11 June 1998; accepted 16 June 1998
Abstract Trehalose phosphorylase (EC 2.4.1.64) from Agaricus bisporus was purified for the first time from a fungus. This enzyme appears to play a key role in trehalose metabolism in A. bisporus since no trehalase or trehalose synthase activities could be detected in this fungus. Trehalose phosphorylase catalyzes the reversible reaction of degradation (phosphorolysis) and synthesis of trehalose. The native enzyme has a molecular weight of 240 kDa and consists of four identical 61-kDa subunits. The isoelectric point of the enzyme was pH 4.8. The optimum temperature for both enzyme reactions was 30³C. The optimum pH ranges for trehalose degradation and synthesis were 6.0^7.5 and 6.0^7.0, respectively. Trehalose degradation was inhibited by ATP and trehalose analogs, whereas the synthetic activity was inhibited by Pi (Ki = 2.0 mM). The enzyme was highly specific towards trehalose, Pi , glucose and K-glucose-1-phosphate. The stoichiometry of the reaction between trehalose, Pi , glucose and K-glucose-1-phosphate was 1:1:1:1 (molar ratio). The Km values were 61, 4.7, 24 and 6.3 mM for trehalose, Pi , glucose and K-glucose-1-phosphate, respectively. Under physiological conditions, A. bisporus trehalose phosphorylase probably performs both synthesis and degradation of trehalose. ß 1998 Elsevier Science B.V. All rights reserved. Keywords: Trehalose; Trehalose phosphorylase; Carbohydrate metabolism; (Agaricus); (Fungus)
1. Introduction Abbreviations: SDS, sodium dodecyl sulfate; IEF, isoelectric focussing; EDTA, ethylenediamine-tetraacetic acid; DTT, dithiothreitol; MES, 4-morpholineethane-sulfonate; TES, tris(hydroxymethyl)methyl-2-aminoethanesulfonate ; HEPES, 4-(2-hydroxyethyl)1-piperazineethanesulfonate ; PMSF, phenylmethylsulfonyl £uoride; NADP, nicotinamide adenine dinucleotide phosphate; CHAPS, 3-(3-cholamidopropyl)-dimethyl-ammonio propanesulfonate; HPLC, high-performance liquid chromatography; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide; KP, potassium phosphate * Corresponding author. Fax: +31 (243) 553450; E-mail: [email protected]
The carbon metabolism of Agaricus bisporus, especially its role in the process of fructi¢cation, is still unclear despite the great economic importance of this fungus. Thus far some striking di¡erences were observed between the carbon metabolism of the fruit body and the vegetative mycelium of A. bisporus. During the development of the fruit body, large amounts of mannitol accumulate. This compound probably acts as an osmoticum during active growth [1,2]. In contrast, glycogen and trehalose accumulate
0304-4165 / 98 / $ ^ see front matter ß 1998 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 4 1 6 5 ( 9 8 ) 0 0 0 6 6 - X
BBAGEN 24660 2-9-98
178
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
in the mycelium [3]. The latter two compounds may serve as storage carbohydrates [4,5], which are degraded under speci¢c circumstances, e.g. stress [6] or fructi¢cation [7]. Some evidence exists that glycogen and trehalose supply a signi¢cant proportion of the carbon for growth of fruit bodies and that their peak levels in mycelium are related to the productivity of the emerging £ush [3]. This paper focusses on the synthesis and degradation of trehalose in A. bisporus. The non-reducing disaccharide trehalose (K-D-glucopyranosyl-[1,1]-K-D-gluco-pyranoside) consists of two K-1,1-linked glucose molecules. It is widespread in nature and has been isolated from bacteria, algae, fungi, insects, invertebrates and plants [8^13]. The function of trehalose varies from providing energy required for £ying (insects) [14] as to withstand extreme desiccation (desert plants) [15]. In fungi, trehalose is thought to function either as a reserve carbohydrate [9,10] or as an e¡ective protectant against various kinds of stress, such as extreme temperatures, osmotic stress and radiation [11,16, 17]. Synthesis of trehalose in bacteria and yeasts occurs predominantly via the trehalose synthase complex (trehalose-6-phosphate synthase and trehalose-6phosphate phosphatase). This pathway has been studied extensively [10,18^21]. The same counts for the hydrolytic cleavage of trehalose by acid and neutral trehalases [8,10,13,22,23]. An alternative for trehalose synthesis and degradation is provided by trehalose phosphorylase (K,K-trehalose:orthophosphate D-glucosyl-transferase; EC 2.1.4.64). This rare enzyme catalyzes both synthesis and degradation (phosphorolysis) of trehalose: Trehalose Pi 3glucose K-glucose-1-phosphate The enzyme was ¢rst described for the alga Euglena gracilis [24,25] and has thus far been found in a rather limited group of bacteria and fungi [26^29]. Trehalose phosphorylase appears to be a key enzyme in A. bisporus carbon metabolism. This paper describes the puri¢cation and characterization of trehalose phosphorylase from A. bisporus. To our knowledge it is the ¢rst time this enzyme has been puri¢ed to homogeneity from a fungus.
2. Materials and methods 2.1. Materials Sephacryl S-300 HR, Sepharose-6B, Q-Sepharose, Superose-12 and MonoQ (HR 5/5) were obtained from Pharmacia (Uppsala, Sweden). Butyl Toyo-pearl was purchased from Tosohaas (Montgomeryville, PA, USA). MES, TES, HEPES, PMSF, 3-(4,5^dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide (MTT) and phenazine methosulfate (PMS) were obtained from Sigma (St. Louis, MO, USA). Hexokinase, glucose-6-phosphate dehydrogenase, NADP, ATP, ADP and AMP were purchased from Boehringer (Mannheim, FRG). DTT was from Serva (Heidelberg, FRG). KH2 PO4 , K2 HPO4 , NaOH (analyzed reagent) and sodium acetate (analyzed reagent) were obtained from J.T. Baker (Deventer, NL). CHAPS and the trehalose analog 1deoxynojirimycin were purchased from Merck (Darmstadt, FRG). The trehalose analog validamycin A was obtained from Duchefa (Haarlem, NL). HPLC equipment was from Hewlett Packard (Cupertino, CA, USA). The Carbopack PA1 ion-exchange and guard columns were obtained from Dionex (Sunnyvale, CA, USA). All other chemicals used were of analytical grade and of the highest purity. 2.2. Microorganism and culture conditions Fruit bodies of A. bisporus strain Horst U1 were obtained from mycelium cultivated on commercially prepared compost and harvested at growth stage 3 [1]. Mycelium of A. bisporus was grown in static 200 ml liquid cultures in Fernbach £asks on Dijkstra medium [30] with 50 mM glucose as the carbon source (unless stated otherwise), and harvested after 21 days of growth at 24³C. 2.3. Enzyme assays Trehalase activity was determined colorimetrically by measuring the glucose liberated, using a glucose assay kit (Sigma) [31], from 250 mM trehalose as a substrate in 50 mM K2 HPO4 /KH2 PO4 (KP bu¡er, pH 7.0) at 30³C.
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Trehalose-6-phosphate synthase activity was assayed by using 50 mM UDP-glucose and 50 mM glucose-6-phosphate as substrates in 50 mM KP buffer (pH 7.0) or 100 mM MES-KOH bu¡er (pH 6.5). After incubation at 30³C samples were analyzed for trehalose-6-phosphate by HPLC. Trehalose-6-phosphate phosphatase activity was measured by HPLC using 20 mM trehalose-6-phosphate as a substrate in 50 mM KP bu¡er (pH 7.0) or 100 mM MES-KOH bu¡er (pH 6.5), containing 5 mM MgCl2 . Incubations took place at 30³C. Throughout the puri¢cation process, trehalose phosphorylase activity was assayed in the direction of trehalose degradation. The reaction mixture contained 100 mM trehalose in 50 mM KP bu¡er (pH 7.0). After incubation at 30³C for 15^30 min, the reaction was stopped by heating (5 min at 100³C) and the reaction mixture was centrifuged (10 min; 10 000Ug; 4³C). The glucose liberated was measured with the glucose oxidase-peroxidase method (glucose assay kit, Sigma). In other experiments the glucose and glucose-1-phosphate liberated were assayed by HPLC. For trehalose synthesis, the reaction mixture contained 50 mM glucose and 10 mM K-glucose1-phosphate in 100 mM MES-KOH bu¡er (pH 6.5). After incubation at 30³C for 15^60 min, the Pi liberated was measured by the method of Bartlett [32]. Alternatively, the trehalose formed was assayed by HPLC. One unit of enzyme activity was, in both directions, de¢ned as 1 Wmol product formed per min under standard assay conditions. 2.4. Protein assay Protein concentrations were determined with the Bio-Rad (Richmond, CA, USA) protein microassay kit, using Q-globulin as a standard. Absorbance at 280 nm was used for monitoring protein in column eluates. 2.5. E¡ect of temperature and pH To measure the e¡ect of temperature on the activity of trehalose phosphorylase, puri¢ed enzyme was incubated at temperatures between 4 and 60³C using the standard incubation conditions. The temperature stability was determined by preincubating the en-
179
zyme at the various temperatures during 60 min, whereafter the samples were cooled immediately on ice. Residual activity was measured at room temperature. The e¡ect of pH on the enzyme activity was determined by incubating the puri¢ed enzyme between pH 4 and 9 using the standard incubation conditions. The pH stability was determined by preincubating the enzyme at the various pH values during 60 min. After immediately cooling on ice, residual activity was measured at room temperature. The bu¡ers used were 0.1 M MES-KOH (pH 4.0^6.5), 0.1 M TES-KOH (pH 6.0^8.0) and 0.1 M HEPES (pH 7.5^9.0). For all phosphorolysis reactions, 10 mM potassium phosphate was included in the assay mix. 2.6. Electrophoresis Polyacrylamide gel electrophoresis (PAGE) methods were performed on precast slab gels using Phastsystem equipment (Pharmacia). Native PAGE was performed on homogeneous gels (7.5 and 10%). A high molecular weight marker kit (Bio-Rad) was used as standard proteins. Estimation of the molecular weight of the enzyme subunits was done with a 10^15% gradient gel (Pharmacia). A low molecular weight marker kit (Bio-Rad) was used as calibration standard. Activity staining, after native PAGE, was done by an agar overlay method. The gel was incubated overnight (at room temperature, in the dark) with 2 ml of 0.5% agar containing 100 mM trehalose, 15 U hexokinase, 15 U glucose-6-phosphate dehydrogenase, 10 mg ATP, 2 mM MgCl2 , 5 mg NADP, 2 mg PMS and 5 mg MTT. In a control experiment, the trehalose was omitted from the staining procedure. 2.7. Stabilization of trehalose phosphorylase Because of the relatively great loss in enzyme activity during initial puri¢cation procedures, several compounds were tested for stabilization. A complexing agent (EDTA, 1^5 mM), sulfhydryl-protective agents (1 mM L-mercaptoethanol, 1^5 mM DTT), a proteolytic inhibitor (PMSF, 0.1^1 mM), glycerol (10^20% (v/v)), as well as the substrates/products (trehalose, 10^50 mM; Pi , 2^200 mM, K-D-glucose-
BBAGEN 24660 2-9-98
180
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
1-phosphate, 1^10 mM; glucose, 10^50 mM) were added to the standard KP bu¡er. 2.8. Puri¢cation of trehalose phosphorylase All procedures were carried out at 4³C, unless stated otherwise. Enzyme activity was assayed as trehalose degradation throughout the puri¢cation process. The KP bu¡ers used were always supplemented with DTT (2 mM), PMSF (0.5 mM) and glycerol (20% (v/v)), and adjusted to pH 7.0. 2.8.1. Step 1: preparation of cell-free extract Harvested fruit bodies were frozen in liquid nitrogen and subsequently ground with glass beads (o.d. 0.10^0.11 mm) in a mortar. The powdered frozen material was suspended in an equal amount of cold extraction bu¡er (50 mM KP bu¡er). The suspension was centrifuged at 40 000Ug (30 min, 4³C) and the clear supernatant (800 ml) was concentrated 5^10fold by a Minitan apparatus (Millipore, membrane cut-o¡ 30 kDa) and used as a cell-free extract. Alternatively, cell-free extracts from fruit bodies were prepared with a Braun food-processing machine, resulting in extracts with similar enzyme activities as obtained when using mortar and pestle. 2.8.2. Step 2: ammonium sulfate fractionation Grounded solid (NH4 )2 SO4 was added to the cellfree extract to reach 30% saturation. After centrifugation at 30 000Ug (30 min, 4³C), the supernatant was brought to 50% saturation by adding more solid (NH4 )2 SO4 . The resulting precipitate was collected and dissolved in 50 mM KP bu¡er. Subsequently, the solution was dialyzed with the same bu¡er using the Minitan apparatus. 2.8.3. Step 3: ultracentrifugation The dialyzed enzyme solution was centrifuged at 200 000Ug (1 h, 4³C), using an ultracentrifuge (Damon/IEC-B60) equipped with a swing-out rotor (SB 283, Beckman) to spin down membrane fragments. The clear supernatant was used in the subsequent experiments. 2.8.4. Step 4: Q-Sepharose fast £ow column chromatography The supernatant was divided into several portions
and each was loaded on a Q-Sepharose fast £ow column (1.5U30 cm) equilibrated with 0.3 M KP bu¡er. After the column was washed, the enzyme was eluted with a linear gradient of 0.3^0.7 M KP bu¡er. The active fractions were pooled and concentrated by ultra¢ltration (Dia£o YM30, Amicon). 2.8.5. Step 5: Butyl-Toyopearl 650M column chromatography To the enzyme solution KP bu¡er (4 M) was added to reach a ¢nal concentration of 1 M. This mixture was put on a Butyl-Toyopearl 650M column (1.5U30 cm) equilibrated with 1 M KP bu¡er. The column was washed, whereafter the enzyme was eluted with a linear gradient of 1^0.3 M of KP buffer. After being pooled the active fractions were concentrated and dialyzed by ultra¢ltration (30 kDa cuto¡) against 2 mM KP bu¡er. 2.8.6. Step 6: trehalose a¤nity column chromatography The enzyme solution was loaded on a TrehaloseSepharose-6B a¤nity column (1.0U12 cm), prepared according to Teunissen et al. [33] with cellobiose replaced by trehalose. The column was kept at a temperature of 1³C and was pre-equilibrated with 2 mM KP bu¡er. After protein was allowed to bind to the column for 30 min, unbound protein was washed from the column with 30 ml of 2 mM KP bu¡er. Subsequently the enzyme was eluted by increasing the NaCl concentration, in the KP bu¡er used, stepwise to 20 mM. The active fractions were pooled, concentrated and brought to 50 mM KP bu¡er by ultra¢ltration (30 kDa cut-o¡). 2.8.7. Step 7: MonoQ column chromatography The enzyme preparation was applied on a MonoQ HR5/5 column (5U50 mm) equilibrated with 50 mM KP bu¡er. Elution of the enzyme took place by a linear gradient of 0.05^0.5 M KP bu¡er. The active fractions were pooled and concentrated to 1.5 ml by ultra¢ltration (30 kDa cut-o¡), and stored at 320³C. 2.9. Molecular weight measurement After partial puri¢cation by anion-exchange chromatography or hydrophobic interaction chromatography, concentrated active fractions (1.0^1.5 ml)
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
181
were applied on a Sephacryl S-300 HR column (1.5U96 cm, Pharmacia) and a Superose-12 column (1.6U50 cm, Pharmacia), both pre-equilibrated in 0.1 M KP bu¡er (pH 7.0) and eluted at a £ow rate of 0.5 ml/min. The columns were calibrated in separate runs with a molecular weight marker kit (Sigma). 2.10. Isoelectric point (pI) measurements The pI was measured by isoelectric focussing at 4³C on precast IEF-gels (pH range 3^9) using a pI marker kit (Pharmacia) of standard proteins. The pI was also determined with a Rotofor apparatus (BioRad), using an ampholyte solution covering a pH range of 3^10. The temperature was maintained at 2³C and 5 mM CHAPS was added to avoid protein precipitation during isoelectric focussing. 2.11. High-pressure liquid chromatography Trehalose, glucose and glucose-1-phosphate were determined quantitatively by HPLC. Brie£y, aliquots of incubation mixtures were boiled for 5 min, followed by dilution of the samples with milliQ water to a ¢nal volume of 0.5 ml. After cooling on ice, samples were centrifuged at 12 000Ug (10 min, 4³C). Samples (20 Wl) of the supernatant were analyzed for carbohydrates with a Hewlett Packard Ti 1050 series HPLC equipped with a 4U250 mm Carbopack PA1 column (Dionex) and a 4U50 mm Carbopack PA-guard column (Dionex). Elution took place with a non-linear gradient of 0.05 M NaOH and 0.4 M sodium acetate at ambient temperature and a £ow of 1 ml/min. Carbohydrates were detected by a pulsed electrochemical detector (Dionex). Commercially available sugars were used as a standard. Galactose was used as the internal standard. 3. Results 3.1. Detection of trehalose phosphorylase activity in A. bisporus The trehalose synthesizing enzymes trehalose6-phosphate synthase and trehalose-6-phosphate phosphatase could not be detected in crude extracts prepared from fruit bodies or mycelium of A. bispo-
Fig. 1. HPLC chromatograms demonstrating the degradation (A) and synthesis (B) of trehalose by trehalose phosphorylase in dialyzed crude extract from Agaricus bisporus fruit bodies after 0 and 60 min of incubation. 1, trehalose; 2, glucose; 3, K-glucose-1-phosphate. The peak at 16.5 min represents the internal standard (galactose).
rus. Furthermore, no evidence for the presence of acid or neutral trehalases was found. Using dialyzed crude extract from A. bisporus fruit bodies and mycelium, degradation of trehalose resulted in formation of equimolar amounts of glucose and K-glucose1-phosphate (Fig. 1A) and appeared to be phosphate dependent. Additionally, incubation of dialyzed crude extract with glucose and K-glucose-1-phosphate resulted in trehalose synthesis (Fig. 1B). This indicates the presence of an alternative route for both synthesis and degradation of trehalose, namely through the action of trehalose phosphorylase.
BBAGEN 24660 2-9-98
182
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
In mycelium extracts, the speci¢c trehalose phosphorylase activities were between 0.005 and 0.010 U/ mg, whereas in fruit body extracts, the activities were twice as high. In the fruit body, highest speci¢c activities of the enzyme were found in extracts from stipe and pileus (up to 0.025 U/mg), whereas gill extracts showed the lowest speci¢c activities (up to 0.010 U/mg). In extracts from mycelia, no signi¢cant di¡erences in trehalose phosphorylase activity were observed when A. bisporus was cultivated on glucose, fructose, trehalose or mannitol, as the carbon source. 3.2. Stabilization of trehalose phosphorylase In cell-free extracts trehalose phosphorylase from A. bisporus remained fairly stable. However, during its puri¢cation there was a considerable loss (up to 95% in 8 h) in enzyme activity. The best stabilizing results were obtained by addition of a combination of Pi (2^100 mM), DTT (2^5 mM), PMSF (0.5 mM) and glycerol (200 mg/ml), which resulted in a maximum loss of 70% in enzyme activity during puri¢cation and storage. 3.3. Puri¢cation of trehalose phosphorylase Initially the puri¢cation of trehalose phosphorylase was hampered by its instability. Incubation of the puri¢ed enzyme at 4³C for 24 h resulted in a loss of enzyme activity of 60 and 70% for trehalose degradation and synthesis, respectively, despite the presence of stabilizing compounds (see above). Enzyme activities remained stable during storage at 380³C for at least 6 months. Trehalose phosphorylase was puri¢ed from cell-
free extract prepared from the fruit bodies of the basidiomycete A. bisporus strain Horst U1. The use of fruit bodies for the puri¢cation of the enzyme was merely practical, since higher activities of the enzyme were observed in fruit bodies compared to mycelium. The results of the puri¢cation are summarized in Table 1. The enzyme was puri¢ed 174-fold with a yield of 0.8%. The speci¢c activity was 4.35 U/mg of protein. Though the total activity of the enzyme decreased signi¢cantly during trehalose a¤nity- and MonoQ column chromatography, these steps were necessary to remove the last impurities. Su¤cient activity remained to perform further experiments. When a gel permeation chromatography step (Sephacryl S-300 HR) was included after the hydrophobic interaction column, activity of the enzyme was completely lost. The puri¢ed enzyme showed a single band on native PAGE of about 240 kDa. This band was also visible upon activity staining of the gel using either puri¢ed enzyme or cell-free extract which was dialyzed and concentrated. The molecular weight of the native trehalose phosphorylase estimated from the elution of a preparation obtained after Q-Sepharose on both Superose-12 and Sephacryl S-300 gel ¢ltration was 240 kDa (Fig. 2). After SDS-PAGE, only a single band with a molecular weight of 61 kDa was found (Fig. 3). The isoelectric point of the enzyme determined by isoelectric focussing and the Rotofor apparatus was 4.7 and 4.8, respectively. 3.4. E¡ects of temperature and pH The e¡ect of temperature on trehalose phosphory-
Table 1 Summary of the puri¢cation procedure of trehalose phosphorylase from Agaricus bisporus strain Horst U1, measured in the direction of trehalose degradation Step
Total protein (mg)
Total activity (U)
Speci¢c activity (U/mg)
Puri¢cation factor (n-fold)
Yield (%)
Cell-free extract Ammonium sulfate Ultra centrifugation Q-Sepharose fast £ow Butyl-Toyopearl Trehalose a¤nity MonoQ HR5/5
11 775 2 190 1 927 468 35.4 4.11 0.53
291 238 230 139 61 15 2.3
0.025 0.109 0.119 0.297 1.72 3.65 4.35
1 4.4 4.8 12.1 68.9 146 174
100 81.8 79.1 47.8 20.9 5.1 0.8
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Fig. 2. Gel ¢ltration (Sephacryl S-300 HR) of the Agaricus bisporus trehalose phosphorylase. a (molecular weight markers): thyroglobulin (669 000), apoferritin (443 000), alcohol dehydrogenase (150 000) and carbonic anhydrase (29 000); arrowhead indicates puri¢ed trehalose phosphorylase.
lase activity is shown in Fig. 4a,c. For both trehalose degradation and synthesis an optimum temperature of 30³C was found. At higher temperatures the enzyme was unstable. Preincubation for 60 min at 40³C resulted in an almost complete loss of activity. Fig. 4b,d shows the e¡ect of pH on the enzyme activity. For trehalose degradation activity was found in the range of pH 6.0^7.5. For trehalose synthesis the optimum pH range was slightly smaller, at pH 7.5 activity decreased.
183
1 mM CuSO4 and 39% by 1 mM CoCl2 . The addition of the trehalose analogs 1-deoxynojirimycin (1 mM) and validamycin A (10 mM) resulted in an inhibition of 88 and 99%, respectively. Addition of 1 and 10 mM ATP led to 11 and 89% inhibition, respectively. When 10 mM ADP was included in the assay mix, the phosphorolytic activity decreased by 34%. The activity was increased to 132% after addition of 10 mM AMP. The synthesis of trehalose was only inhibited by the addition of 10 mM ADP and AMP, resulting in activity losses of 47 and 54%, respectively. Addition of 10 mM ATP stimulated the synthetic activity by 20%. 3.7. Reaction mechanism and kinetics To study the stoichiometry of trehalose phosphorolysis and synthesis, HPLC analyses of trehalose, glucose and K-glucose-1-phosphate were compared with Pi levels measured by the method of Bartlett
3.5. Substrate speci¢city The substrate speci¢city of trehalose phosphorylase was determined in both directions. In the degradation reaction trehalose could not be substituted by neotrehalose, sucrose, maltose, lactose, cellobiose, starch or glycogen, and Pi could not be replaced by arsenate. The following substances did not act as substitutes for glucose: fructose, mannose, galactose, xylose, glucosamine, 2-deoxyglucose, 6-deoxyglucose and glucono-N-lactone. L-Glucose-1-phosphate, glucose-6-phosphate, fructose-1-phosphate, fructose6-phosphate and mannitol-1-phosphate could not replace K-glucose-1-phosphate. 3.6. E¡ects of inhibitors The e¡ects of various reagents and metal ions on the enzyme activity in both directions are shown in Table 2. Trehalose degradation was inhibited 37% by
Fig. 3. SDS-PAGE analysis of trehalose phosphorylase from Agaricus bisporus. Separation was performed by a 10^15% gradient gel. Lane 1, molecular weight markers: phosphorylase b (97 000), serum albumin (66 000), ovalbumin (45 000) and carbonic anhydrase (29 000). Lane 2, silver-stained puri¢ed trehalose phosphorylase (5 Wg).
BBAGEN 24660 2-9-98
184
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Fig. 4. E¡ects of temperature and pH on trehalose phosphorylase activity (F) and stability (E) during trehalose degradation (a,b) and trehalose synthesis (c,d); 100% of enzyme activity is equal to 2.5 (a), 1.6 (b), 1.5 (c) and 1.5 (d) U/mg, respectively.
[32]. It was found that in 1 h, the puri¢ed enzyme (4 U) converted 0.95 þ 0.19 Wmol of trehalose to 0.97 þ 0.20 Wmol glucose and 0.91 þ 0.16 Wmol of K-glucose-1-phosphate. Synthesis of trehalose took place by conversion of 0.70 þ 0.20 Wmol of glucose and 0.65 þ 0.17 Wmol of K-glucose-1-phosphate to 0.67 þ 0.15 Wmol of trehalose and 0.73 þ 0.16 Wmol of Pi . From these data a stoichiometry between trehalose, Pi , glucose and glucose-1-phosphate can be deduced of 1:1:1:1 (molar ratio). The kinetic parameters for both reactions are shown in Table 3. The apparent Km and Vmax values were calculated from double reciprocal plots of the initial velocities and substrate concentrations (1/v versus 1/s), and by using direct linear plots (v against v/s). The Km values for trehalose, Pi , glucose and K-glucose-1-phosphate were 61, 4.7, 24 and 6.3 mM, respectively. To test the e¡ects on the equilibrium, reaction products were added to the assay mixture. The trehalose degradation reaction was not inhibited by the addition of 5^50 mM glucose. Addition of 5, 20 and 50 mM K-glucose-1-phosphate led to an inhibition of
8, 21 and 46%, respectively. Furthermore, the degradation of trehalose appeared to be strictly dependent on Pi , with a Km of 4.7 mM (Fig. 5). High concentrations of Pi (up to 200 mM) were not inhibitory. However, the synthesis of trehalose was strongly inhibited by Pi (Ki = 2.0 mM); at 50 mM Pi an inhibition of 90% was found. Addition of 5, 20 and 50 mM trehalose did not signi¢cantly inhibit the synthesis reaction. 4. Discussion For more than two decades, the metabolism of trehalose in the white button mushroom, A. bisporus, has been studied [1,24,25,34,35]. It was concluded that trehalose accumulates in the mycelium until a threshold concentration is reached. At the onset of fructi¢cation, the trehalose is translocated to the emerging fruit bodies, where it would be degraded by trehalase yielding two glucose molecules serving as an energy source for further growth of the fruit bodies. Based on these studies, it was postulated that
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188 Table 2 E¡ect of metal ions and reagents on the activity of trehalose phosphorylase from Agaricus bisporus during degradation and synthesis of trehalose Reagent None CaCl2 CoCl2 CuSO4 FeCl3 HgCl2 KCl KCN MgCl2 MnSO4 NaCl NH4 Cl (NH4 )2 SO4 NiCl2 ZnCl2 ZnSO4 ATPa ADP AMP DTT EDTA 2-Deoxyglucose 1-Deoxynojirimycin Fructose-1-phosphate Fructose-6-phosphate Fructose-1,6-diphosphate Glucono-N-lactone K-Glucose-1-phosphate Glucose-6-phosphate UDP-glucose Validamycin A
Concentration Relative activity (%) (mM) Phosphorolysis Synthesis 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 10.0 1.0 10.0 1.0 10.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0
100 72 61 63 79 92 94 80 96 107 97 101 99 89 71 74 89 11 99 66 97 132 101 75 98 12 95 96 94
100 104 103 95 100 96 101 110 98 97 98 103 102 97 92 94 106 120 106 53 101 46 100 89 101 95 98 100 98
1.0 1.0 1.0 1.0 1.0 10.0
68 84 105 91 36 1
96 103 98 99 97 94
Enzyme activity was measured in the presence of metal ions or reagents under the standard assay conditions. The relative activity was expressed as a percentage of the enzyme activity in the absence of reagents. a Hexokinase activity, which could lead to a false impression of ATP inhibition, was absent in the enzyme preparation.
trehalose might serve as a triggering substrate in the process of periodic fruiting in A. bisporus. So far, only suggestive data were provided about the synthesis of trehalose in A. bisporus, assuming a situa-
185
tion identical to Saccharomyces cerevisiae [7]. In contrast, data presented in this article exclude the presence of trehalases and the trehalose synthase complex (trehalose-6-phosphate synthase and trehalose-6-phosphate phosphatase) in the trehalose metabolism of A. bisporus. The results described here clearly show that trehalose degradation and synthesis is catalyzed by a single enzyme: trehalose phosphorylase. Literature about this enzyme is scarce and to our knowledge this is the ¢rst trehalose phosphorylase puri¢ed to homogeneity from a fungus. In contrast to trehalase, which is a rather abundant enzyme, Aisaka and Masuda only found trehalose phosphorylase activity in seven (all actinomycetes) out of 482 (i.e. 1.5%) microorganisms tested [29]. This could be an underestimation since most authors who study trehalase do not perform HPLC experiments to distinguish the enzyme from trehalose phosphorylase. Trehalase is only capable of hydrolyzing trehalose into two glucose molecules, whereas trehalose phosphorylase splits trehalose in one glucose and one glucose-1-phosphate molecule. The latter reaction is energetically more favorable, since no expenditure of ATP is needed to form the phosphorylated sugar. Also, the phosphorylated sugar cannot freely di¡use out of the cell. Besides this, trehalose phosphorylase can also perform the reverse reaction to synthesize trehalose. The attempts to purify trehalose phosphorylase from A. bisporus were initially hampered by the rapid inactivation of the enzyme, especially when the enzyme solution was dialyzed. However, in the presence of Pi , DTT, PMSF and glycerol, loss of enzyme activity, as measured in the direction of trehalose degradation, was markedly decreased. There was a tendency to increased inactivation of the enzyme during the ¢nal steps of the puri¢cation procedure and during gel ¢ltration chromatography. As in the ascomycete Pichia fermentans, this might be caused by Table 3 Kinetic parameters for the substrates of trehalose phosphorylase from Agaricus bisporus Substrate
Km (mM)
Vmax (Wmol/min/mg)
Trehalose Pi Glucose K-Glucose-1-phosphate
61 4.7 24 6.3
1.24 0.90 0.64 0.60
BBAGEN 24660 2-9-98
186
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Fig. 5. E¡ects of phosphate on trehalose phosphorylase activity during trehalose degradation (E) and synthesis (F); 100% of enzyme activity is equal to 2.5 and 1.5 U/mg for trehalose degradation and synthesis, respectively.
dilution, presumably resulting in dissociation of the enzyme into its less active monomeric subunits [36]. Instability of phosphorylase enzymes during puri¢cation procedures has been described before [24,25,36,37]. From SDS-PAGE, activity staining after PAGE and from gel permeation chromatography it could be concluded that the enzyme has a homotetrameric structure with 61-kDa subunits, and a native molecular weight of 240 kDa. The existence of tetrameric forms is a common feature for sucrose phosphorylases [37], whereas cellobiose phosphorylase exists in a monomeric form [38]. The trehalose phosphorylase from Micrococcus varians consists of six identical subunits [28]. The activity staining with the puri¢ed enzyme resulted in a single intense black band. The same unique protein band was seen after activity staining with cell-free extract, although it was faint. The coloring of only one band when using cell-free extract suggests that there was no other trehalose degrading or glucose synthesizing activity present in A. bisporus. The temperature and pH optima for the enzyme, in both directions, are comparable to those found for other organisms [25^28]. The latter also applies for its high substrate speci¢city [27,28]. The result of the ultracentrifugation step indicates that trehalose phosphorylase from A. bisporus is not membrane-bound. This is in contrast to the work of Mare¨chal and Belocopitow [25], who recovered a substantial part (60%) of the enzyme activity from Euglena gracilis in the precipitate after ultracentrifugation. However, these authors suggested binding of the enzyme to heavy cellular components as a probable cause for the precipitation.
The mycelia grown on di¡erent carbon sources all showed trehalose phosphorylase activity, within the same range (0.005^0.010 U/mg). This suggests that the trehalose phosphorylase protein is synthesized constitutively and that the activity of the enzyme might be regulated by post-translational modi¢cation. For instance, regulation of the enzyme by allosteric binding of adenosine phosphates might be possible, implying that the metabolic status of the cell (re£ected by levels of ATP, ADP and AMP) can be involved in the regulation of A. bisporus trehalose phosphorylase. The trehalose analogs 1-deoxynojirimycin and validamycin A inhibit the enzyme through competition with trehalose [28]. Thus far, similar results were obtained with the trehalose phosphorylase from Micrococcus varians [28] and with trehalases [5,22]. The kinetic data of the enzyme fall into the same range as those of other trehalose phosphorylases known sofar [25,27,28]. The relatively low Km values for Pi and K-glucose-1-phosphate compared to trehalose and glucose, indicate binding of the substrates in an ordered sequence. An ordered Bi Bi mechanism (ping pong), proceeding via intermediary complexes, has been proposed for the trehalose phosphorylase from M. varians [28]. In vivo inhibition of trehalose degradation by K-glucose-1-phosphate probably plays no role, since its Ki value is approximately 50 mM. Regulation of the in vivo activity of A. bisporus trehalose phosphorylase by Pi will be studied in more detail. Intracellular concentrations of free inorganic phosphate in S. cerevisiae cells were estimated to be 15 mM [39]. In A. bisporus, the free Pi concentration varied from 2^5 mM in the mycelium to 5^20 mM in the fruit bodies. These data imply that, when the enzyme is cytosolic, synthesis of trehalose can particularly occur in the mycelium, whereas trehalose degradation preferably occurs in the fruit bodies. The properties of catalytic and regulatory sites in phosphorylases have been reviewed by Palm et al. [40] and remain to be studied in A. bisporus. Therefore, it is uncertain to predict the physiological role of the trehalose phosphorylase from A. bisporus. In E. gracilis [25], the enzyme can perform both synthesis and degradation of trehalose, in Flammulina velutipes [27], trehalose phosphorylase seems active only in degradation, and in Catellatospora ferruginea [29] under physio-
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
logical conditions there is probably only trehalose synthesis. For A. bisporus, the Km values and the preliminary data on the regulation of the enzyme indicate that both synthesis and degradation of trehalose can be performed, depending on the physiological conditions in vivo. In summary, the trehalose phosphorylase from A. bisporus resembles other trehalose phosphorylases with respect to its structural and kinetic properties. Besides the fundamental knowledge on trehalose metabolism in general, there is now an increasing commercial interest to the potential applications of trehalose metabolizing enzymes. Next to the synthesis of glucose-1-phosphate as an antibiotic or immunosuppressive drug [41,42], the large-scale industrial synthesis of trehalose has received much attention. The commercial applications of trehalose are currently being investigated by the pharmaceutical and agricultural industries [43^47]. An enzyme converting starch into trehalose has already been commercially exploited [48]. To increase knowledge on A. bisporus trehalose phosphorylase, further research will be directed to the role of the enzyme during di¡erent developmental stages of this basidiomycete fungus, to the elucidation of its molecular background and to potential applications of this unique enzyme.
References [1] J.B.W. Hammond, R. Nichols, J. Gen. Microbiol. 93 (1976) 309^320. [2] J.B.W. Hammond, New Phytol. 89 (1981) 419^428. [3] J.B.W. Hammond, in: D.M. Moore, L.A. Casselton, D.A. Wood, J.C. Frankland (Eds.), Developmental Biology of Higher Fungi, Cambridge University Press, Cambridge, UK, 1985, pp. 389^401. [4] A.D. Panek, Arch. Biochem. Biophys. 100 (1963) 422^425. [5] J.M. Thevelein, Microbiol. Rev. 48 (1984) 42^59. [6] A. Wiemken, Antonie van Leeuwenhoek 58 (1990) 209^217. [7] T.K. Wells, J.B.W. Hammond, A.G. Dickerson, New Phytol. 105 (1987) 273^280. [8] A.D. Elbein, Adv. Carbohydr. Chem. Biochem. 30 (1974) 227^256. [9] H.L. Blumenthal, in: J.E. Smith, D.R. Berry (Eds.), The Filamentous Fungi, Edward Arnold, London, UK, 1976, pp. 292^307. [10] J.M. Thevelein, K.A. Jones, Eur. J. Biochem. 136 (1983) 583^587.
187
[11] T. Hottinger, T. Boller, A. Wiemken, FEBS Lett. 220 (1987) 113^115. [12] A. Van Laere, FEMS Microbiol. Rev. 63 (1989) 201^210. [13] A. Wiemken, M. Schellenberg, FEBS Lett. 150 (1982) 329^ 331. [14] A. Becker, P. Schlo«der, J.E. Steele, G. Wegener, Experientia 52 (1996) 433^437. [15] A.D. Elbein, Adv. Carbohydrate Chem. Biochem. 30 (1974) 227^239. [16] P.W. Piper, FEMS Microbiol. Rev. 11 (1993) 339^356. [17] K. Yoshinaga, H. Yoshioka, H. Kurosaki, M. Hirasawa, M. Uritani, K. Hasegawa, Biosci. Biotech. Biochem. 61 (1997) 160^164. [18] J. Londesborough, O.E. Vuorio, J. Gen. Microbiol. 137 (1993) 323^330. [19] A. Vandercammen, J. Francois, H.G. Hers, Eur. J. Biochem. 182 (1989) 613^620. [20] J. Winderickx, J.H. de Winde, M. Crauwels, A. Hino, S. Hohmann, P. Van Dijk, J.M. Thevelein, Mol. Gen. Genet. 252 (1996) 470^482. [21] J.L. Parrou, M.A. Teste, J. Franc,ois, Microbiology 143 (1997) 1891^1900. [22] O.J.M. Goddijn, T.C. Verwoerd, E. Voogd, R.H.W.W. Krutwagen, P.T.H.M. de Graaf, J. Poels, K. van Dun, A.S. Ponstein, B. Damm, J. Pen, Plant Physiol. 113 (1997) 181^190. [23] R. Horlacher, K. Uhland, W. Klein, M. Ehrmann, W. Boos, J. Bacteriol. 178 (1996) 6250^6257. [24] E. Belocopitow, L.R. Mare¨chal, Biochim. Biophys. Acta 198 (1970) 151^154. [25] L.R. Mare¨chal, E. Belocopitow, J. Biol. Chem. 247 (1972) 3223^3228. [26] S.O. Salminen, J.G. Streeter, Plant Physiol. 81 (1986) 538^ 541. [27] Y. Kitamoto, H. Akashi, H. Tanaka, N. Mori, FEMS Microbiol. Lett. 55 (1988) 147^150. [28] H. Kizawa, K. Miyagawa, Y. Sugiyama, Biosci. Biotech. Biochem. 59 (1995) 1908^1912. [29] K. Aisada, T. Masuda, FEMS Microbiol. Lett. 131 (1995) 47^51. [30] J.J.P. Baars, H.J.M. Op den Camp, J.M.H. Hermans, V. Mikes, C. van der Drift, L.J.L.D. Van Griensven, G.D. Vogels, Microbiology 140 (1994) 1161^1168. [31] H.V. Bergmeyer, E. Berndt, in: H.V. Bergmeyer, K. Gawehn (Eds.), Methoden der Enzymatischen Analyse, Verlag Chemie, Weinheim, Germany, 1974, pp. 1250^1257. [32] G.R. Bartlett, J. Biol. Chem. 234 (1959) 466^468. [33] M.J. Teunisssen, H.T.P. Lahaye, H.J. Huis in 't Veld, G.D. Vogels, Arch. Microbiol. 158 (1992) 63^69. [34] D.O. Chanter, J.H.M. Thornley, J. Gen. Microbiol. 106 (1977) 55^65. [35] D.O. Chanter, J. Gen. Microbiol. 115 (1979) 79^87. [36] I. Schick, D. Haltrich, K.D. Kulbe, Appl. Microbiol. Biotechnol. 43 (1995) 1088^1095. [37] T. Sasaki, T. Tanaka, S. Nakagawa, K. Kainuma, Biochem. J. 209 (1983) 803^807.
BBAGEN 24660 2-9-98
188
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
[38] E.J. Vandamme, J. Van Loo, L. Machtelinckx, A. Delaporte, Adv. Appl. Microbiol. 32 (1987) 163^201. [39] A. Rothstein, in: A. Shanes (Ed.), Electrolytes in Biological Systems, American Physiological Society, Washington, DC, 1955, pp. 65^100. [40] D. Palm, R. Goerl, K.L. Burger, Nature 313 (1985) 500^502. [41] C. Parish, W.B. Cowden, D.O. Willenborg, Int. Cl. (1990) A61K 31/725, 31/71, C07H11/04. [42] A. Weinha«usel, B. Nidetzky, M. Rohrbach, B. Blauensteiner, K.D. Kulbe, Appl. Microbiol. Biotechnol. 41 (1994) 510^ 516.
[43] J. Kim, P. Alizadeh, T. Harding, A. Hefner-Gravik, D.J. Klionsky, Appl. Environ. Microbiol. 62 (1996) 1563^1567. [44] J. Londesborough, O. Vuorio, International patent application (1993) PCT/FI93/00049. [45] M. Driessen, K.A. Osinga, M.A. Herweijer, European patent application (1991) 91200686.3. [46] S. Ogawa, C. Uchida, J. Chem. Soc. 1 (1992) 1939^1943. [47] G. Semenza, Annu. Rev. Cell Biol. 2 (1986) 255^260. [48] K. Maruta, T. Nakada, M. Kubota, H. Chaen, T. Sugimoto, M. Kurimoto, Y. Tsujisaka, Biosci. Biotech. Biochem. 59 (1995) 1829^1834.
BBAGEN 24660 2-9-98
Puri¢cation and characterization of trehalose phosphorylase from the commercial mushroom Agaricus bisporus Wim J.B. Wannet a , Huub J.M. Op den Camp a; *, Hendrik W. Wisselink a , Chris van der Drift a , Leo J.L.D. Van Griensven b , Godfried D. Vogels a a
Department of Microbiology, Faculty of Science, University of Nijmegen, Toernooiveld 1, NL-6525 ED Nijmegen, Netherlands b Mushroom Experimental Station, Postbus 6042, NL-5960 AA Horst, Netherlands Received 30 March 1998; revised 11 June 1998; accepted 16 June 1998
Abstract Trehalose phosphorylase (EC 2.4.1.64) from Agaricus bisporus was purified for the first time from a fungus. This enzyme appears to play a key role in trehalose metabolism in A. bisporus since no trehalase or trehalose synthase activities could be detected in this fungus. Trehalose phosphorylase catalyzes the reversible reaction of degradation (phosphorolysis) and synthesis of trehalose. The native enzyme has a molecular weight of 240 kDa and consists of four identical 61-kDa subunits. The isoelectric point of the enzyme was pH 4.8. The optimum temperature for both enzyme reactions was 30³C. The optimum pH ranges for trehalose degradation and synthesis were 6.0^7.5 and 6.0^7.0, respectively. Trehalose degradation was inhibited by ATP and trehalose analogs, whereas the synthetic activity was inhibited by Pi (Ki = 2.0 mM). The enzyme was highly specific towards trehalose, Pi , glucose and K-glucose-1-phosphate. The stoichiometry of the reaction between trehalose, Pi , glucose and K-glucose-1-phosphate was 1:1:1:1 (molar ratio). The Km values were 61, 4.7, 24 and 6.3 mM for trehalose, Pi , glucose and K-glucose-1-phosphate, respectively. Under physiological conditions, A. bisporus trehalose phosphorylase probably performs both synthesis and degradation of trehalose. ß 1998 Elsevier Science B.V. All rights reserved. Keywords: Trehalose; Trehalose phosphorylase; Carbohydrate metabolism; (Agaricus); (Fungus)
1. Introduction Abbreviations: SDS, sodium dodecyl sulfate; IEF, isoelectric focussing; EDTA, ethylenediamine-tetraacetic acid; DTT, dithiothreitol; MES, 4-morpholineethane-sulfonate; TES, tris(hydroxymethyl)methyl-2-aminoethanesulfonate ; HEPES, 4-(2-hydroxyethyl)1-piperazineethanesulfonate ; PMSF, phenylmethylsulfonyl £uoride; NADP, nicotinamide adenine dinucleotide phosphate; CHAPS, 3-(3-cholamidopropyl)-dimethyl-ammonio propanesulfonate; HPLC, high-performance liquid chromatography; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide; KP, potassium phosphate * Corresponding author. Fax: +31 (243) 553450; E-mail: [email protected]
The carbon metabolism of Agaricus bisporus, especially its role in the process of fructi¢cation, is still unclear despite the great economic importance of this fungus. Thus far some striking di¡erences were observed between the carbon metabolism of the fruit body and the vegetative mycelium of A. bisporus. During the development of the fruit body, large amounts of mannitol accumulate. This compound probably acts as an osmoticum during active growth [1,2]. In contrast, glycogen and trehalose accumulate
0304-4165 / 98 / $ ^ see front matter ß 1998 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 4 1 6 5 ( 9 8 ) 0 0 0 6 6 - X
BBAGEN 24660 2-9-98
178
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
in the mycelium [3]. The latter two compounds may serve as storage carbohydrates [4,5], which are degraded under speci¢c circumstances, e.g. stress [6] or fructi¢cation [7]. Some evidence exists that glycogen and trehalose supply a signi¢cant proportion of the carbon for growth of fruit bodies and that their peak levels in mycelium are related to the productivity of the emerging £ush [3]. This paper focusses on the synthesis and degradation of trehalose in A. bisporus. The non-reducing disaccharide trehalose (K-D-glucopyranosyl-[1,1]-K-D-gluco-pyranoside) consists of two K-1,1-linked glucose molecules. It is widespread in nature and has been isolated from bacteria, algae, fungi, insects, invertebrates and plants [8^13]. The function of trehalose varies from providing energy required for £ying (insects) [14] as to withstand extreme desiccation (desert plants) [15]. In fungi, trehalose is thought to function either as a reserve carbohydrate [9,10] or as an e¡ective protectant against various kinds of stress, such as extreme temperatures, osmotic stress and radiation [11,16, 17]. Synthesis of trehalose in bacteria and yeasts occurs predominantly via the trehalose synthase complex (trehalose-6-phosphate synthase and trehalose-6phosphate phosphatase). This pathway has been studied extensively [10,18^21]. The same counts for the hydrolytic cleavage of trehalose by acid and neutral trehalases [8,10,13,22,23]. An alternative for trehalose synthesis and degradation is provided by trehalose phosphorylase (K,K-trehalose:orthophosphate D-glucosyl-transferase; EC 2.1.4.64). This rare enzyme catalyzes both synthesis and degradation (phosphorolysis) of trehalose: Trehalose Pi 3glucose K-glucose-1-phosphate The enzyme was ¢rst described for the alga Euglena gracilis [24,25] and has thus far been found in a rather limited group of bacteria and fungi [26^29]. Trehalose phosphorylase appears to be a key enzyme in A. bisporus carbon metabolism. This paper describes the puri¢cation and characterization of trehalose phosphorylase from A. bisporus. To our knowledge it is the ¢rst time this enzyme has been puri¢ed to homogeneity from a fungus.
2. Materials and methods 2.1. Materials Sephacryl S-300 HR, Sepharose-6B, Q-Sepharose, Superose-12 and MonoQ (HR 5/5) were obtained from Pharmacia (Uppsala, Sweden). Butyl Toyo-pearl was purchased from Tosohaas (Montgomeryville, PA, USA). MES, TES, HEPES, PMSF, 3-(4,5^dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide (MTT) and phenazine methosulfate (PMS) were obtained from Sigma (St. Louis, MO, USA). Hexokinase, glucose-6-phosphate dehydrogenase, NADP, ATP, ADP and AMP were purchased from Boehringer (Mannheim, FRG). DTT was from Serva (Heidelberg, FRG). KH2 PO4 , K2 HPO4 , NaOH (analyzed reagent) and sodium acetate (analyzed reagent) were obtained from J.T. Baker (Deventer, NL). CHAPS and the trehalose analog 1deoxynojirimycin were purchased from Merck (Darmstadt, FRG). The trehalose analog validamycin A was obtained from Duchefa (Haarlem, NL). HPLC equipment was from Hewlett Packard (Cupertino, CA, USA). The Carbopack PA1 ion-exchange and guard columns were obtained from Dionex (Sunnyvale, CA, USA). All other chemicals used were of analytical grade and of the highest purity. 2.2. Microorganism and culture conditions Fruit bodies of A. bisporus strain Horst U1 were obtained from mycelium cultivated on commercially prepared compost and harvested at growth stage 3 [1]. Mycelium of A. bisporus was grown in static 200 ml liquid cultures in Fernbach £asks on Dijkstra medium [30] with 50 mM glucose as the carbon source (unless stated otherwise), and harvested after 21 days of growth at 24³C. 2.3. Enzyme assays Trehalase activity was determined colorimetrically by measuring the glucose liberated, using a glucose assay kit (Sigma) [31], from 250 mM trehalose as a substrate in 50 mM K2 HPO4 /KH2 PO4 (KP bu¡er, pH 7.0) at 30³C.
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Trehalose-6-phosphate synthase activity was assayed by using 50 mM UDP-glucose and 50 mM glucose-6-phosphate as substrates in 50 mM KP buffer (pH 7.0) or 100 mM MES-KOH bu¡er (pH 6.5). After incubation at 30³C samples were analyzed for trehalose-6-phosphate by HPLC. Trehalose-6-phosphate phosphatase activity was measured by HPLC using 20 mM trehalose-6-phosphate as a substrate in 50 mM KP bu¡er (pH 7.0) or 100 mM MES-KOH bu¡er (pH 6.5), containing 5 mM MgCl2 . Incubations took place at 30³C. Throughout the puri¢cation process, trehalose phosphorylase activity was assayed in the direction of trehalose degradation. The reaction mixture contained 100 mM trehalose in 50 mM KP bu¡er (pH 7.0). After incubation at 30³C for 15^30 min, the reaction was stopped by heating (5 min at 100³C) and the reaction mixture was centrifuged (10 min; 10 000Ug; 4³C). The glucose liberated was measured with the glucose oxidase-peroxidase method (glucose assay kit, Sigma). In other experiments the glucose and glucose-1-phosphate liberated were assayed by HPLC. For trehalose synthesis, the reaction mixture contained 50 mM glucose and 10 mM K-glucose1-phosphate in 100 mM MES-KOH bu¡er (pH 6.5). After incubation at 30³C for 15^60 min, the Pi liberated was measured by the method of Bartlett [32]. Alternatively, the trehalose formed was assayed by HPLC. One unit of enzyme activity was, in both directions, de¢ned as 1 Wmol product formed per min under standard assay conditions. 2.4. Protein assay Protein concentrations were determined with the Bio-Rad (Richmond, CA, USA) protein microassay kit, using Q-globulin as a standard. Absorbance at 280 nm was used for monitoring protein in column eluates. 2.5. E¡ect of temperature and pH To measure the e¡ect of temperature on the activity of trehalose phosphorylase, puri¢ed enzyme was incubated at temperatures between 4 and 60³C using the standard incubation conditions. The temperature stability was determined by preincubating the en-
179
zyme at the various temperatures during 60 min, whereafter the samples were cooled immediately on ice. Residual activity was measured at room temperature. The e¡ect of pH on the enzyme activity was determined by incubating the puri¢ed enzyme between pH 4 and 9 using the standard incubation conditions. The pH stability was determined by preincubating the enzyme at the various pH values during 60 min. After immediately cooling on ice, residual activity was measured at room temperature. The bu¡ers used were 0.1 M MES-KOH (pH 4.0^6.5), 0.1 M TES-KOH (pH 6.0^8.0) and 0.1 M HEPES (pH 7.5^9.0). For all phosphorolysis reactions, 10 mM potassium phosphate was included in the assay mix. 2.6. Electrophoresis Polyacrylamide gel electrophoresis (PAGE) methods were performed on precast slab gels using Phastsystem equipment (Pharmacia). Native PAGE was performed on homogeneous gels (7.5 and 10%). A high molecular weight marker kit (Bio-Rad) was used as standard proteins. Estimation of the molecular weight of the enzyme subunits was done with a 10^15% gradient gel (Pharmacia). A low molecular weight marker kit (Bio-Rad) was used as calibration standard. Activity staining, after native PAGE, was done by an agar overlay method. The gel was incubated overnight (at room temperature, in the dark) with 2 ml of 0.5% agar containing 100 mM trehalose, 15 U hexokinase, 15 U glucose-6-phosphate dehydrogenase, 10 mg ATP, 2 mM MgCl2 , 5 mg NADP, 2 mg PMS and 5 mg MTT. In a control experiment, the trehalose was omitted from the staining procedure. 2.7. Stabilization of trehalose phosphorylase Because of the relatively great loss in enzyme activity during initial puri¢cation procedures, several compounds were tested for stabilization. A complexing agent (EDTA, 1^5 mM), sulfhydryl-protective agents (1 mM L-mercaptoethanol, 1^5 mM DTT), a proteolytic inhibitor (PMSF, 0.1^1 mM), glycerol (10^20% (v/v)), as well as the substrates/products (trehalose, 10^50 mM; Pi , 2^200 mM, K-D-glucose-
BBAGEN 24660 2-9-98
180
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
1-phosphate, 1^10 mM; glucose, 10^50 mM) were added to the standard KP bu¡er. 2.8. Puri¢cation of trehalose phosphorylase All procedures were carried out at 4³C, unless stated otherwise. Enzyme activity was assayed as trehalose degradation throughout the puri¢cation process. The KP bu¡ers used were always supplemented with DTT (2 mM), PMSF (0.5 mM) and glycerol (20% (v/v)), and adjusted to pH 7.0. 2.8.1. Step 1: preparation of cell-free extract Harvested fruit bodies were frozen in liquid nitrogen and subsequently ground with glass beads (o.d. 0.10^0.11 mm) in a mortar. The powdered frozen material was suspended in an equal amount of cold extraction bu¡er (50 mM KP bu¡er). The suspension was centrifuged at 40 000Ug (30 min, 4³C) and the clear supernatant (800 ml) was concentrated 5^10fold by a Minitan apparatus (Millipore, membrane cut-o¡ 30 kDa) and used as a cell-free extract. Alternatively, cell-free extracts from fruit bodies were prepared with a Braun food-processing machine, resulting in extracts with similar enzyme activities as obtained when using mortar and pestle. 2.8.2. Step 2: ammonium sulfate fractionation Grounded solid (NH4 )2 SO4 was added to the cellfree extract to reach 30% saturation. After centrifugation at 30 000Ug (30 min, 4³C), the supernatant was brought to 50% saturation by adding more solid (NH4 )2 SO4 . The resulting precipitate was collected and dissolved in 50 mM KP bu¡er. Subsequently, the solution was dialyzed with the same bu¡er using the Minitan apparatus. 2.8.3. Step 3: ultracentrifugation The dialyzed enzyme solution was centrifuged at 200 000Ug (1 h, 4³C), using an ultracentrifuge (Damon/IEC-B60) equipped with a swing-out rotor (SB 283, Beckman) to spin down membrane fragments. The clear supernatant was used in the subsequent experiments. 2.8.4. Step 4: Q-Sepharose fast £ow column chromatography The supernatant was divided into several portions
and each was loaded on a Q-Sepharose fast £ow column (1.5U30 cm) equilibrated with 0.3 M KP bu¡er. After the column was washed, the enzyme was eluted with a linear gradient of 0.3^0.7 M KP bu¡er. The active fractions were pooled and concentrated by ultra¢ltration (Dia£o YM30, Amicon). 2.8.5. Step 5: Butyl-Toyopearl 650M column chromatography To the enzyme solution KP bu¡er (4 M) was added to reach a ¢nal concentration of 1 M. This mixture was put on a Butyl-Toyopearl 650M column (1.5U30 cm) equilibrated with 1 M KP bu¡er. The column was washed, whereafter the enzyme was eluted with a linear gradient of 1^0.3 M of KP buffer. After being pooled the active fractions were concentrated and dialyzed by ultra¢ltration (30 kDa cuto¡) against 2 mM KP bu¡er. 2.8.6. Step 6: trehalose a¤nity column chromatography The enzyme solution was loaded on a TrehaloseSepharose-6B a¤nity column (1.0U12 cm), prepared according to Teunissen et al. [33] with cellobiose replaced by trehalose. The column was kept at a temperature of 1³C and was pre-equilibrated with 2 mM KP bu¡er. After protein was allowed to bind to the column for 30 min, unbound protein was washed from the column with 30 ml of 2 mM KP bu¡er. Subsequently the enzyme was eluted by increasing the NaCl concentration, in the KP bu¡er used, stepwise to 20 mM. The active fractions were pooled, concentrated and brought to 50 mM KP bu¡er by ultra¢ltration (30 kDa cut-o¡). 2.8.7. Step 7: MonoQ column chromatography The enzyme preparation was applied on a MonoQ HR5/5 column (5U50 mm) equilibrated with 50 mM KP bu¡er. Elution of the enzyme took place by a linear gradient of 0.05^0.5 M KP bu¡er. The active fractions were pooled and concentrated to 1.5 ml by ultra¢ltration (30 kDa cut-o¡), and stored at 320³C. 2.9. Molecular weight measurement After partial puri¢cation by anion-exchange chromatography or hydrophobic interaction chromatography, concentrated active fractions (1.0^1.5 ml)
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
181
were applied on a Sephacryl S-300 HR column (1.5U96 cm, Pharmacia) and a Superose-12 column (1.6U50 cm, Pharmacia), both pre-equilibrated in 0.1 M KP bu¡er (pH 7.0) and eluted at a £ow rate of 0.5 ml/min. The columns were calibrated in separate runs with a molecular weight marker kit (Sigma). 2.10. Isoelectric point (pI) measurements The pI was measured by isoelectric focussing at 4³C on precast IEF-gels (pH range 3^9) using a pI marker kit (Pharmacia) of standard proteins. The pI was also determined with a Rotofor apparatus (BioRad), using an ampholyte solution covering a pH range of 3^10. The temperature was maintained at 2³C and 5 mM CHAPS was added to avoid protein precipitation during isoelectric focussing. 2.11. High-pressure liquid chromatography Trehalose, glucose and glucose-1-phosphate were determined quantitatively by HPLC. Brie£y, aliquots of incubation mixtures were boiled for 5 min, followed by dilution of the samples with milliQ water to a ¢nal volume of 0.5 ml. After cooling on ice, samples were centrifuged at 12 000Ug (10 min, 4³C). Samples (20 Wl) of the supernatant were analyzed for carbohydrates with a Hewlett Packard Ti 1050 series HPLC equipped with a 4U250 mm Carbopack PA1 column (Dionex) and a 4U50 mm Carbopack PA-guard column (Dionex). Elution took place with a non-linear gradient of 0.05 M NaOH and 0.4 M sodium acetate at ambient temperature and a £ow of 1 ml/min. Carbohydrates were detected by a pulsed electrochemical detector (Dionex). Commercially available sugars were used as a standard. Galactose was used as the internal standard. 3. Results 3.1. Detection of trehalose phosphorylase activity in A. bisporus The trehalose synthesizing enzymes trehalose6-phosphate synthase and trehalose-6-phosphate phosphatase could not be detected in crude extracts prepared from fruit bodies or mycelium of A. bispo-
Fig. 1. HPLC chromatograms demonstrating the degradation (A) and synthesis (B) of trehalose by trehalose phosphorylase in dialyzed crude extract from Agaricus bisporus fruit bodies after 0 and 60 min of incubation. 1, trehalose; 2, glucose; 3, K-glucose-1-phosphate. The peak at 16.5 min represents the internal standard (galactose).
rus. Furthermore, no evidence for the presence of acid or neutral trehalases was found. Using dialyzed crude extract from A. bisporus fruit bodies and mycelium, degradation of trehalose resulted in formation of equimolar amounts of glucose and K-glucose1-phosphate (Fig. 1A) and appeared to be phosphate dependent. Additionally, incubation of dialyzed crude extract with glucose and K-glucose-1-phosphate resulted in trehalose synthesis (Fig. 1B). This indicates the presence of an alternative route for both synthesis and degradation of trehalose, namely through the action of trehalose phosphorylase.
BBAGEN 24660 2-9-98
182
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
In mycelium extracts, the speci¢c trehalose phosphorylase activities were between 0.005 and 0.010 U/ mg, whereas in fruit body extracts, the activities were twice as high. In the fruit body, highest speci¢c activities of the enzyme were found in extracts from stipe and pileus (up to 0.025 U/mg), whereas gill extracts showed the lowest speci¢c activities (up to 0.010 U/mg). In extracts from mycelia, no signi¢cant di¡erences in trehalose phosphorylase activity were observed when A. bisporus was cultivated on glucose, fructose, trehalose or mannitol, as the carbon source. 3.2. Stabilization of trehalose phosphorylase In cell-free extracts trehalose phosphorylase from A. bisporus remained fairly stable. However, during its puri¢cation there was a considerable loss (up to 95% in 8 h) in enzyme activity. The best stabilizing results were obtained by addition of a combination of Pi (2^100 mM), DTT (2^5 mM), PMSF (0.5 mM) and glycerol (200 mg/ml), which resulted in a maximum loss of 70% in enzyme activity during puri¢cation and storage. 3.3. Puri¢cation of trehalose phosphorylase Initially the puri¢cation of trehalose phosphorylase was hampered by its instability. Incubation of the puri¢ed enzyme at 4³C for 24 h resulted in a loss of enzyme activity of 60 and 70% for trehalose degradation and synthesis, respectively, despite the presence of stabilizing compounds (see above). Enzyme activities remained stable during storage at 380³C for at least 6 months. Trehalose phosphorylase was puri¢ed from cell-
free extract prepared from the fruit bodies of the basidiomycete A. bisporus strain Horst U1. The use of fruit bodies for the puri¢cation of the enzyme was merely practical, since higher activities of the enzyme were observed in fruit bodies compared to mycelium. The results of the puri¢cation are summarized in Table 1. The enzyme was puri¢ed 174-fold with a yield of 0.8%. The speci¢c activity was 4.35 U/mg of protein. Though the total activity of the enzyme decreased signi¢cantly during trehalose a¤nity- and MonoQ column chromatography, these steps were necessary to remove the last impurities. Su¤cient activity remained to perform further experiments. When a gel permeation chromatography step (Sephacryl S-300 HR) was included after the hydrophobic interaction column, activity of the enzyme was completely lost. The puri¢ed enzyme showed a single band on native PAGE of about 240 kDa. This band was also visible upon activity staining of the gel using either puri¢ed enzyme or cell-free extract which was dialyzed and concentrated. The molecular weight of the native trehalose phosphorylase estimated from the elution of a preparation obtained after Q-Sepharose on both Superose-12 and Sephacryl S-300 gel ¢ltration was 240 kDa (Fig. 2). After SDS-PAGE, only a single band with a molecular weight of 61 kDa was found (Fig. 3). The isoelectric point of the enzyme determined by isoelectric focussing and the Rotofor apparatus was 4.7 and 4.8, respectively. 3.4. E¡ects of temperature and pH The e¡ect of temperature on trehalose phosphory-
Table 1 Summary of the puri¢cation procedure of trehalose phosphorylase from Agaricus bisporus strain Horst U1, measured in the direction of trehalose degradation Step
Total protein (mg)
Total activity (U)
Speci¢c activity (U/mg)
Puri¢cation factor (n-fold)
Yield (%)
Cell-free extract Ammonium sulfate Ultra centrifugation Q-Sepharose fast £ow Butyl-Toyopearl Trehalose a¤nity MonoQ HR5/5
11 775 2 190 1 927 468 35.4 4.11 0.53
291 238 230 139 61 15 2.3
0.025 0.109 0.119 0.297 1.72 3.65 4.35
1 4.4 4.8 12.1 68.9 146 174
100 81.8 79.1 47.8 20.9 5.1 0.8
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Fig. 2. Gel ¢ltration (Sephacryl S-300 HR) of the Agaricus bisporus trehalose phosphorylase. a (molecular weight markers): thyroglobulin (669 000), apoferritin (443 000), alcohol dehydrogenase (150 000) and carbonic anhydrase (29 000); arrowhead indicates puri¢ed trehalose phosphorylase.
lase activity is shown in Fig. 4a,c. For both trehalose degradation and synthesis an optimum temperature of 30³C was found. At higher temperatures the enzyme was unstable. Preincubation for 60 min at 40³C resulted in an almost complete loss of activity. Fig. 4b,d shows the e¡ect of pH on the enzyme activity. For trehalose degradation activity was found in the range of pH 6.0^7.5. For trehalose synthesis the optimum pH range was slightly smaller, at pH 7.5 activity decreased.
183
1 mM CuSO4 and 39% by 1 mM CoCl2 . The addition of the trehalose analogs 1-deoxynojirimycin (1 mM) and validamycin A (10 mM) resulted in an inhibition of 88 and 99%, respectively. Addition of 1 and 10 mM ATP led to 11 and 89% inhibition, respectively. When 10 mM ADP was included in the assay mix, the phosphorolytic activity decreased by 34%. The activity was increased to 132% after addition of 10 mM AMP. The synthesis of trehalose was only inhibited by the addition of 10 mM ADP and AMP, resulting in activity losses of 47 and 54%, respectively. Addition of 10 mM ATP stimulated the synthetic activity by 20%. 3.7. Reaction mechanism and kinetics To study the stoichiometry of trehalose phosphorolysis and synthesis, HPLC analyses of trehalose, glucose and K-glucose-1-phosphate were compared with Pi levels measured by the method of Bartlett
3.5. Substrate speci¢city The substrate speci¢city of trehalose phosphorylase was determined in both directions. In the degradation reaction trehalose could not be substituted by neotrehalose, sucrose, maltose, lactose, cellobiose, starch or glycogen, and Pi could not be replaced by arsenate. The following substances did not act as substitutes for glucose: fructose, mannose, galactose, xylose, glucosamine, 2-deoxyglucose, 6-deoxyglucose and glucono-N-lactone. L-Glucose-1-phosphate, glucose-6-phosphate, fructose-1-phosphate, fructose6-phosphate and mannitol-1-phosphate could not replace K-glucose-1-phosphate. 3.6. E¡ects of inhibitors The e¡ects of various reagents and metal ions on the enzyme activity in both directions are shown in Table 2. Trehalose degradation was inhibited 37% by
Fig. 3. SDS-PAGE analysis of trehalose phosphorylase from Agaricus bisporus. Separation was performed by a 10^15% gradient gel. Lane 1, molecular weight markers: phosphorylase b (97 000), serum albumin (66 000), ovalbumin (45 000) and carbonic anhydrase (29 000). Lane 2, silver-stained puri¢ed trehalose phosphorylase (5 Wg).
BBAGEN 24660 2-9-98
184
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Fig. 4. E¡ects of temperature and pH on trehalose phosphorylase activity (F) and stability (E) during trehalose degradation (a,b) and trehalose synthesis (c,d); 100% of enzyme activity is equal to 2.5 (a), 1.6 (b), 1.5 (c) and 1.5 (d) U/mg, respectively.
[32]. It was found that in 1 h, the puri¢ed enzyme (4 U) converted 0.95 þ 0.19 Wmol of trehalose to 0.97 þ 0.20 Wmol glucose and 0.91 þ 0.16 Wmol of K-glucose-1-phosphate. Synthesis of trehalose took place by conversion of 0.70 þ 0.20 Wmol of glucose and 0.65 þ 0.17 Wmol of K-glucose-1-phosphate to 0.67 þ 0.15 Wmol of trehalose and 0.73 þ 0.16 Wmol of Pi . From these data a stoichiometry between trehalose, Pi , glucose and glucose-1-phosphate can be deduced of 1:1:1:1 (molar ratio). The kinetic parameters for both reactions are shown in Table 3. The apparent Km and Vmax values were calculated from double reciprocal plots of the initial velocities and substrate concentrations (1/v versus 1/s), and by using direct linear plots (v against v/s). The Km values for trehalose, Pi , glucose and K-glucose-1-phosphate were 61, 4.7, 24 and 6.3 mM, respectively. To test the e¡ects on the equilibrium, reaction products were added to the assay mixture. The trehalose degradation reaction was not inhibited by the addition of 5^50 mM glucose. Addition of 5, 20 and 50 mM K-glucose-1-phosphate led to an inhibition of
8, 21 and 46%, respectively. Furthermore, the degradation of trehalose appeared to be strictly dependent on Pi , with a Km of 4.7 mM (Fig. 5). High concentrations of Pi (up to 200 mM) were not inhibitory. However, the synthesis of trehalose was strongly inhibited by Pi (Ki = 2.0 mM); at 50 mM Pi an inhibition of 90% was found. Addition of 5, 20 and 50 mM trehalose did not signi¢cantly inhibit the synthesis reaction. 4. Discussion For more than two decades, the metabolism of trehalose in the white button mushroom, A. bisporus, has been studied [1,24,25,34,35]. It was concluded that trehalose accumulates in the mycelium until a threshold concentration is reached. At the onset of fructi¢cation, the trehalose is translocated to the emerging fruit bodies, where it would be degraded by trehalase yielding two glucose molecules serving as an energy source for further growth of the fruit bodies. Based on these studies, it was postulated that
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188 Table 2 E¡ect of metal ions and reagents on the activity of trehalose phosphorylase from Agaricus bisporus during degradation and synthesis of trehalose Reagent None CaCl2 CoCl2 CuSO4 FeCl3 HgCl2 KCl KCN MgCl2 MnSO4 NaCl NH4 Cl (NH4 )2 SO4 NiCl2 ZnCl2 ZnSO4 ATPa ADP AMP DTT EDTA 2-Deoxyglucose 1-Deoxynojirimycin Fructose-1-phosphate Fructose-6-phosphate Fructose-1,6-diphosphate Glucono-N-lactone K-Glucose-1-phosphate Glucose-6-phosphate UDP-glucose Validamycin A
Concentration Relative activity (%) (mM) Phosphorolysis Synthesis 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 10.0 1.0 10.0 1.0 10.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0
100 72 61 63 79 92 94 80 96 107 97 101 99 89 71 74 89 11 99 66 97 132 101 75 98 12 95 96 94
100 104 103 95 100 96 101 110 98 97 98 103 102 97 92 94 106 120 106 53 101 46 100 89 101 95 98 100 98
1.0 1.0 1.0 1.0 1.0 10.0
68 84 105 91 36 1
96 103 98 99 97 94
Enzyme activity was measured in the presence of metal ions or reagents under the standard assay conditions. The relative activity was expressed as a percentage of the enzyme activity in the absence of reagents. a Hexokinase activity, which could lead to a false impression of ATP inhibition, was absent in the enzyme preparation.
trehalose might serve as a triggering substrate in the process of periodic fruiting in A. bisporus. So far, only suggestive data were provided about the synthesis of trehalose in A. bisporus, assuming a situa-
185
tion identical to Saccharomyces cerevisiae [7]. In contrast, data presented in this article exclude the presence of trehalases and the trehalose synthase complex (trehalose-6-phosphate synthase and trehalose-6-phosphate phosphatase) in the trehalose metabolism of A. bisporus. The results described here clearly show that trehalose degradation and synthesis is catalyzed by a single enzyme: trehalose phosphorylase. Literature about this enzyme is scarce and to our knowledge this is the ¢rst trehalose phosphorylase puri¢ed to homogeneity from a fungus. In contrast to trehalase, which is a rather abundant enzyme, Aisaka and Masuda only found trehalose phosphorylase activity in seven (all actinomycetes) out of 482 (i.e. 1.5%) microorganisms tested [29]. This could be an underestimation since most authors who study trehalase do not perform HPLC experiments to distinguish the enzyme from trehalose phosphorylase. Trehalase is only capable of hydrolyzing trehalose into two glucose molecules, whereas trehalose phosphorylase splits trehalose in one glucose and one glucose-1-phosphate molecule. The latter reaction is energetically more favorable, since no expenditure of ATP is needed to form the phosphorylated sugar. Also, the phosphorylated sugar cannot freely di¡use out of the cell. Besides this, trehalose phosphorylase can also perform the reverse reaction to synthesize trehalose. The attempts to purify trehalose phosphorylase from A. bisporus were initially hampered by the rapid inactivation of the enzyme, especially when the enzyme solution was dialyzed. However, in the presence of Pi , DTT, PMSF and glycerol, loss of enzyme activity, as measured in the direction of trehalose degradation, was markedly decreased. There was a tendency to increased inactivation of the enzyme during the ¢nal steps of the puri¢cation procedure and during gel ¢ltration chromatography. As in the ascomycete Pichia fermentans, this might be caused by Table 3 Kinetic parameters for the substrates of trehalose phosphorylase from Agaricus bisporus Substrate
Km (mM)
Vmax (Wmol/min/mg)
Trehalose Pi Glucose K-Glucose-1-phosphate
61 4.7 24 6.3
1.24 0.90 0.64 0.60
BBAGEN 24660 2-9-98
186
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
Fig. 5. E¡ects of phosphate on trehalose phosphorylase activity during trehalose degradation (E) and synthesis (F); 100% of enzyme activity is equal to 2.5 and 1.5 U/mg for trehalose degradation and synthesis, respectively.
dilution, presumably resulting in dissociation of the enzyme into its less active monomeric subunits [36]. Instability of phosphorylase enzymes during puri¢cation procedures has been described before [24,25,36,37]. From SDS-PAGE, activity staining after PAGE and from gel permeation chromatography it could be concluded that the enzyme has a homotetrameric structure with 61-kDa subunits, and a native molecular weight of 240 kDa. The existence of tetrameric forms is a common feature for sucrose phosphorylases [37], whereas cellobiose phosphorylase exists in a monomeric form [38]. The trehalose phosphorylase from Micrococcus varians consists of six identical subunits [28]. The activity staining with the puri¢ed enzyme resulted in a single intense black band. The same unique protein band was seen after activity staining with cell-free extract, although it was faint. The coloring of only one band when using cell-free extract suggests that there was no other trehalose degrading or glucose synthesizing activity present in A. bisporus. The temperature and pH optima for the enzyme, in both directions, are comparable to those found for other organisms [25^28]. The latter also applies for its high substrate speci¢city [27,28]. The result of the ultracentrifugation step indicates that trehalose phosphorylase from A. bisporus is not membrane-bound. This is in contrast to the work of Mare¨chal and Belocopitow [25], who recovered a substantial part (60%) of the enzyme activity from Euglena gracilis in the precipitate after ultracentrifugation. However, these authors suggested binding of the enzyme to heavy cellular components as a probable cause for the precipitation.
The mycelia grown on di¡erent carbon sources all showed trehalose phosphorylase activity, within the same range (0.005^0.010 U/mg). This suggests that the trehalose phosphorylase protein is synthesized constitutively and that the activity of the enzyme might be regulated by post-translational modi¢cation. For instance, regulation of the enzyme by allosteric binding of adenosine phosphates might be possible, implying that the metabolic status of the cell (re£ected by levels of ATP, ADP and AMP) can be involved in the regulation of A. bisporus trehalose phosphorylase. The trehalose analogs 1-deoxynojirimycin and validamycin A inhibit the enzyme through competition with trehalose [28]. Thus far, similar results were obtained with the trehalose phosphorylase from Micrococcus varians [28] and with trehalases [5,22]. The kinetic data of the enzyme fall into the same range as those of other trehalose phosphorylases known sofar [25,27,28]. The relatively low Km values for Pi and K-glucose-1-phosphate compared to trehalose and glucose, indicate binding of the substrates in an ordered sequence. An ordered Bi Bi mechanism (ping pong), proceeding via intermediary complexes, has been proposed for the trehalose phosphorylase from M. varians [28]. In vivo inhibition of trehalose degradation by K-glucose-1-phosphate probably plays no role, since its Ki value is approximately 50 mM. Regulation of the in vivo activity of A. bisporus trehalose phosphorylase by Pi will be studied in more detail. Intracellular concentrations of free inorganic phosphate in S. cerevisiae cells were estimated to be 15 mM [39]. In A. bisporus, the free Pi concentration varied from 2^5 mM in the mycelium to 5^20 mM in the fruit bodies. These data imply that, when the enzyme is cytosolic, synthesis of trehalose can particularly occur in the mycelium, whereas trehalose degradation preferably occurs in the fruit bodies. The properties of catalytic and regulatory sites in phosphorylases have been reviewed by Palm et al. [40] and remain to be studied in A. bisporus. Therefore, it is uncertain to predict the physiological role of the trehalose phosphorylase from A. bisporus. In E. gracilis [25], the enzyme can perform both synthesis and degradation of trehalose, in Flammulina velutipes [27], trehalose phosphorylase seems active only in degradation, and in Catellatospora ferruginea [29] under physio-
BBAGEN 24660 2-9-98
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
logical conditions there is probably only trehalose synthesis. For A. bisporus, the Km values and the preliminary data on the regulation of the enzyme indicate that both synthesis and degradation of trehalose can be performed, depending on the physiological conditions in vivo. In summary, the trehalose phosphorylase from A. bisporus resembles other trehalose phosphorylases with respect to its structural and kinetic properties. Besides the fundamental knowledge on trehalose metabolism in general, there is now an increasing commercial interest to the potential applications of trehalose metabolizing enzymes. Next to the synthesis of glucose-1-phosphate as an antibiotic or immunosuppressive drug [41,42], the large-scale industrial synthesis of trehalose has received much attention. The commercial applications of trehalose are currently being investigated by the pharmaceutical and agricultural industries [43^47]. An enzyme converting starch into trehalose has already been commercially exploited [48]. To increase knowledge on A. bisporus trehalose phosphorylase, further research will be directed to the role of the enzyme during di¡erent developmental stages of this basidiomycete fungus, to the elucidation of its molecular background and to potential applications of this unique enzyme.
References [1] J.B.W. Hammond, R. Nichols, J. Gen. Microbiol. 93 (1976) 309^320. [2] J.B.W. Hammond, New Phytol. 89 (1981) 419^428. [3] J.B.W. Hammond, in: D.M. Moore, L.A. Casselton, D.A. Wood, J.C. Frankland (Eds.), Developmental Biology of Higher Fungi, Cambridge University Press, Cambridge, UK, 1985, pp. 389^401. [4] A.D. Panek, Arch. Biochem. Biophys. 100 (1963) 422^425. [5] J.M. Thevelein, Microbiol. Rev. 48 (1984) 42^59. [6] A. Wiemken, Antonie van Leeuwenhoek 58 (1990) 209^217. [7] T.K. Wells, J.B.W. Hammond, A.G. Dickerson, New Phytol. 105 (1987) 273^280. [8] A.D. Elbein, Adv. Carbohydr. Chem. Biochem. 30 (1974) 227^256. [9] H.L. Blumenthal, in: J.E. Smith, D.R. Berry (Eds.), The Filamentous Fungi, Edward Arnold, London, UK, 1976, pp. 292^307. [10] J.M. Thevelein, K.A. Jones, Eur. J. Biochem. 136 (1983) 583^587.
187
[11] T. Hottinger, T. Boller, A. Wiemken, FEBS Lett. 220 (1987) 113^115. [12] A. Van Laere, FEMS Microbiol. Rev. 63 (1989) 201^210. [13] A. Wiemken, M. Schellenberg, FEBS Lett. 150 (1982) 329^ 331. [14] A. Becker, P. Schlo«der, J.E. Steele, G. Wegener, Experientia 52 (1996) 433^437. [15] A.D. Elbein, Adv. Carbohydrate Chem. Biochem. 30 (1974) 227^239. [16] P.W. Piper, FEMS Microbiol. Rev. 11 (1993) 339^356. [17] K. Yoshinaga, H. Yoshioka, H. Kurosaki, M. Hirasawa, M. Uritani, K. Hasegawa, Biosci. Biotech. Biochem. 61 (1997) 160^164. [18] J. Londesborough, O.E. Vuorio, J. Gen. Microbiol. 137 (1993) 323^330. [19] A. Vandercammen, J. Francois, H.G. Hers, Eur. J. Biochem. 182 (1989) 613^620. [20] J. Winderickx, J.H. de Winde, M. Crauwels, A. Hino, S. Hohmann, P. Van Dijk, J.M. Thevelein, Mol. Gen. Genet. 252 (1996) 470^482. [21] J.L. Parrou, M.A. Teste, J. Franc,ois, Microbiology 143 (1997) 1891^1900. [22] O.J.M. Goddijn, T.C. Verwoerd, E. Voogd, R.H.W.W. Krutwagen, P.T.H.M. de Graaf, J. Poels, K. van Dun, A.S. Ponstein, B. Damm, J. Pen, Plant Physiol. 113 (1997) 181^190. [23] R. Horlacher, K. Uhland, W. Klein, M. Ehrmann, W. Boos, J. Bacteriol. 178 (1996) 6250^6257. [24] E. Belocopitow, L.R. Mare¨chal, Biochim. Biophys. Acta 198 (1970) 151^154. [25] L.R. Mare¨chal, E. Belocopitow, J. Biol. Chem. 247 (1972) 3223^3228. [26] S.O. Salminen, J.G. Streeter, Plant Physiol. 81 (1986) 538^ 541. [27] Y. Kitamoto, H. Akashi, H. Tanaka, N. Mori, FEMS Microbiol. Lett. 55 (1988) 147^150. [28] H. Kizawa, K. Miyagawa, Y. Sugiyama, Biosci. Biotech. Biochem. 59 (1995) 1908^1912. [29] K. Aisada, T. Masuda, FEMS Microbiol. Lett. 131 (1995) 47^51. [30] J.J.P. Baars, H.J.M. Op den Camp, J.M.H. Hermans, V. Mikes, C. van der Drift, L.J.L.D. Van Griensven, G.D. Vogels, Microbiology 140 (1994) 1161^1168. [31] H.V. Bergmeyer, E. Berndt, in: H.V. Bergmeyer, K. Gawehn (Eds.), Methoden der Enzymatischen Analyse, Verlag Chemie, Weinheim, Germany, 1974, pp. 1250^1257. [32] G.R. Bartlett, J. Biol. Chem. 234 (1959) 466^468. [33] M.J. Teunisssen, H.T.P. Lahaye, H.J. Huis in 't Veld, G.D. Vogels, Arch. Microbiol. 158 (1992) 63^69. [34] D.O. Chanter, J.H.M. Thornley, J. Gen. Microbiol. 106 (1977) 55^65. [35] D.O. Chanter, J. Gen. Microbiol. 115 (1979) 79^87. [36] I. Schick, D. Haltrich, K.D. Kulbe, Appl. Microbiol. Biotechnol. 43 (1995) 1088^1095. [37] T. Sasaki, T. Tanaka, S. Nakagawa, K. Kainuma, Biochem. J. 209 (1983) 803^807.
BBAGEN 24660 2-9-98
188
W.J.B. Wannet et al. / Biochimica et Biophysica Acta 1425 (1998) 177^188
[38] E.J. Vandamme, J. Van Loo, L. Machtelinckx, A. Delaporte, Adv. Appl. Microbiol. 32 (1987) 163^201. [39] A. Rothstein, in: A. Shanes (Ed.), Electrolytes in Biological Systems, American Physiological Society, Washington, DC, 1955, pp. 65^100. [40] D. Palm, R. Goerl, K.L. Burger, Nature 313 (1985) 500^502. [41] C. Parish, W.B. Cowden, D.O. Willenborg, Int. Cl. (1990) A61K 31/725, 31/71, C07H11/04. [42] A. Weinha«usel, B. Nidetzky, M. Rohrbach, B. Blauensteiner, K.D. Kulbe, Appl. Microbiol. Biotechnol. 41 (1994) 510^ 516.
[43] J. Kim, P. Alizadeh, T. Harding, A. Hefner-Gravik, D.J. Klionsky, Appl. Environ. Microbiol. 62 (1996) 1563^1567. [44] J. Londesborough, O. Vuorio, International patent application (1993) PCT/FI93/00049. [45] M. Driessen, K.A. Osinga, M.A. Herweijer, European patent application (1991) 91200686.3. [46] S. Ogawa, C. Uchida, J. Chem. Soc. 1 (1992) 1939^1943. [47] G. Semenza, Annu. Rev. Cell Biol. 2 (1986) 255^260. [48] K. Maruta, T. Nakada, M. Kubota, H. Chaen, T. Sugimoto, M. Kurimoto, Y. Tsujisaka, Biosci. Biotech. Biochem. 59 (1995) 1829^1834.
BBAGEN 24660 2-9-98