Characterization of a novel β-glucosidase from a Stachybotrys strain

Biochemical Engineering Journal 32 (2006) 191–197

Characterization of a novel ␤-glucosidase from a Stachybotrys strain Bahia Amouri, Ali Gargouri ∗ Laboratoire de G´en´etique Mol´eculaire des Eucaryotes, Centre de Biotechnologie de Sfax, BP “K” 3038-Sfax, Tunisia Received 23 May 2006; received in revised form 15 September 2006; accepted 27 September 2006

Abstract An improved mutant was isolated from the cellulolytic fungus Stachybotrys sp. after nitrous acid mutagenesis. It was fed-batch cultivated on cellulose and its extracellular cellulases (mainly the endoglucanases and ␤-glucosidases) were analyzed. One ␤-glucosidase was purified to homogeneity after two steps, MonoQ and gel filtration and shown to be a dimeric protein. The molecular weight of each monomer is 85 kDa. Besides its aryl ␤-glucosidase activity towards salicin, methyl-umbellypheryl-␤-d-glucoside (MUG) and p-nitrophenyl-␤-d-glucoside (pNPG), it showed a true ␤-glucosidase activity since it splits cellobiose into two glucose monomers. The Vmax and the Km kinetics parameters with pNPG as substrate were 78 U/mg and 0.27 mM, respectively. The enzyme shows more affinity to pNPG than cellobiose and salicin whose apparent values of Km were, respectively, 2.22 and 37.14 mM. This enzyme exhibits its optimal activity at pH 5 and at 50 ◦ C. Interestingly, this activity is not affected by denaturing gel conditions (SDS and ␤-mercaptoethanol) as long as it is not pre-heated. The N-terminal sequence of the purified enzyme showed a significant homology with the family 1 ␤-glucosidases of Trichoderma reesei and Humicola isolens even though these two enzymes are much smaller in size. © 2006 Elsevier B.V. All rights reserved. Keywords: Stachybotrys sp.; ␤-Glucosidase; Cellulase; Purification; Chromatography; Enzyme activity

1. Introduction Cellulose is the most abundant polymer in nature and should give sufficient substrate for human nutrition and bio-industries. A prerequisite to do such is to convert this polymer into glucose, the monomeric end-product of enzymatic degradation carried out by cellulases [1]. Many fungal strains secrete higher amounts of cellulases than bacterial ones, with Trichoderma as the leading one. Other fungal species were shown to be interesting cellulase producers, such as Humicola [2] or Aspergillus [3] or but some unstudied strains could reveal some peculiarities of cellullases since much still to be known on this class of hydrolases. The cellulolytic enzymes are composed of three main activities: endoglucanase, exoglucanase and ␤-glucosidase, are wide spread among bacterial and fungal strains even though the exoglucanases (also called cellobiohydrolases) are rare in the bacterial kingdom. More than hundred of endo and exoglucanase sequences are known [4]. For most of endoglucanases and cellobiohydrolases, a common structure is shared, the cellulose binding domain (CBD) is linked by a flexible hinge to a catalytic

∗

Corresponding author. Fax: +216 74 440 818. E-mail address: [email protected] (A. Gargouri).

1369-703X/$ – see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.bej.2006.09.022

core domain [5,6]. Fungal CBDs are very short and are one of the most conserved sequences in nature. Cellulolytic enzymes are also known to play an important role in some industrial applications, such as in bio-stone washing of jeans, replacing the stones in their abrasive effect on the garments [7]. ␤-Glucosidase (␤-d-glucoside glucohydrolase, EC 3.2.1.21) constitutes a major group among glucoside hydrolase. They occur ubiquitously in all three (archea, eubacteria and eukarya) domains of living organisms. Glycoside hydrolases catalyse the selective hydrolysis of glycosidic bond in oligosaccharides, polysaccharides and their conjugates. The ␤-glucosidase has been the focus of much research recently because of their important roles in a variety of fundamental biological processes [8]. These enzymes split the cellobiose into two monomers of glucose, withdrawing by the way the inhibition exerted by cellobiose on certain cellulase such as cellobiohydrolase I of Trichoderma reesei [9]. It is worth to note that some ␤-glucosidase are only able to split aryl ␤glucoside such as salicin, p-nitrophenyl-␤-d-glucoside (pNPG) and methyl-umbellypheryl-␤-d-glucoside (MUG) but not cellobiose; they are called “aryl-␤-glucosidase” and should not be considered as true ␤-glucosidases [10]. ␤-Glucosidases are also important in the regulation of cellulase genes since they are the key enzyme in the synthesis of sophorose, an efficient

192

B. Amouri, A. Gargouri / Biochemical Engineering Journal 32 (2006) 191–197

inducer of the cellulolytic system of T. reesei [11]. They constitute also the focus of many applied researches since they are not only needed in the cellulose breakdown but also in the synthesis of oligomers and other complex molecules (such as alkyl-glucosides) by trans-glycosylation [8]. In a previous work, we designed a simple method to clone fungal CBD and have shown that one such CBD, despite some sequence particularities, retained its function (adsorption on cellulose) when expressed in Escherichia coli [12]. This CBD was cloned from a novel cellulolytic Stachybotrys sp. strain that we have locally isolated. In this paper, we report the purification and characterization of a novel ␤-glucosidase in the cellulolytic system of Stachybotrys strain. 2. Materials and methods 2.1. Fungal strain The mutant A19 was isolated from the cellulolytic fungus N1 (isolated in our laboratory) and identified as Stachybotrys sp. by the Centraalbureau voor Schimmelcultures (The Netherlands) after nitrous acid mutagenesis. 2.2. Cellulases production For batch cultures, cells were cultivated for 4 days at 30 ◦ C in shaking flasks containing Mandels medium supplemented with 100 ␮g/ml ampicillin and 12.5 ␮g/ml tetracycline, to avoid bacterial contamination. The fed-batch culture was performed in fermentor containing 10 l of the initial medium (Mandels medium with 2% cellulose Avicel) and equipped with automatic control of agitation, temperature, pH and foaming. In all experiments, the pH was controlled by automatic addition of ammonium hydroxide (4N) or hydrochloride acid (4N). Ammonium hydroxide addition also ensured a sufficient supply of nitrogen. Dissolved oxygen was kept at above 20% of the saturation for the medium by varying the agitation rate and flow rate of air. The temperature was maintained at 30 ◦ C. The first feeding was initiated after 4 days of batch culture; 500 ml of cellulose Avicel slurry were added to the fermenter to bring the cellulose concentration to 2%. A second feeding was repeated after 3 days and the culture stopped after 2 days more. Twenty milliliters broth samples were taken regularly during the course of fermentation and examined for growth, contamination and cellulases activities. At the end of fermentation (9 days), the mycelium was harvested by centrifugation at 2500 × g for 10 min and the supernatant was used to purify cellulolytic enzymes. 2.3. Cellulases purification Cellulases were precipitated by ammonium sulfate to 80% saturation and then centrifuged at 5000 × g for 25 min. The pellet was re-suspended in a 50 mM phosphate buffer pH 7, 1 mM phenylmethyl sulfonyl fluoride (PMSF), 10 mM dithiothreitol (DTT, and 10 mM EDTA pH 8; desalted on G25-Sephadex, then fractionated on a MonoQ (Pharmacia) FPLC column

(15 mm × 68 mm) using a gradient 0–1 M NaCl in 20 mM Tris–HCl pH 8 buffer. Some fractions were further purified on a gel filtration column (Bio-Sill® Sec 250 Gel Filtration HPLC Column: 300 mm × 7.8 mm serial 402613 Catalog 125-0062) in 20 mM Tris–HCl pH 8 buffer. 2.4. Enzyme assays Total cellulase (usually named “filter paper” or FP), endoglucanase (CMCases) and ␤-glucosidase activities were determined in the supernatant samples collected after 3, 5 and 7 days culture. The filter paper activity was determined as recommended by the IUPAC biotechnology commission, as follows: 0.5 ml of citrate buffer 50 mM pH 4.8 was added to a test tube, 0.5 ml of enzyme suitably diluted and an assay disc (Ø12.7 mm) of Whatman filter paper (Schleicher and Schuell). The reaction mixture was incubated at 50 ◦ C for 60 min and 3 ml of dinitrosalicylic acid (DNS) reagent added to stop the reaction. The tubes were transferred to boiling water for 10 min. Twenty milliliters of distilled water were added, the contents of the tubes were mixed and the color formed was red in a spectrophotometer at 550 nm. Glucose standard solutions were treated exactly in the same conditions. The endoglucanase and ␤-glucosidase (salicinase) activities were determined using carboxymethylcellulose (CMC, Hercules) or salicin (Sigma) substrate, respectively, as follows: 0.5 ml of 2% substrate was incubated with 0.5 ml enzyme, diluted with citrate buffer at 50 ◦ C for 30 min. DNS reagent (3 ml) was added and the solution was boiled for 10 min. Finally, 20 ml distilled water was added and the absorbance was measured at 550 nm. The ␤-glucosidase activity was also determined using pnitrophenyl-␤-d-glucoside (pNPG) as substrate, and therefore called pNPGase, as follows: 1 ml of 5 mM pNPG (in 100 mM acetate buffer pH 4.8) was incubated with the enzyme solution at 50 ◦ C for 30 min. The reaction was stopped by adding 4 ml glycine buffer 400 mM pH 10.8; the liberated p-nitrophenol was measured at 430 nm. For all these dosages, the 1 unit of activity is the amount of enzyme required to release 1 ␮mol glucose (DNS procedure) or p-nitrophenol per min under the conditions of the assay. 2.5. Determination of pH and temperature optima The optimal pH of the pNPGase activity was determined by incubating the purified enzyme at 50 ◦ C for 30 min in different buffers: citrate (100 mM, pH 3), acetate (100 mM, pH 4.8), phosphate (100 mM, pH 7) and glycine buffer (100 mM, pH 9). To determine the optimal temperature, the enzyme was incubated in acetate buffer (100 mM, pH 4.8) for 30 min at different temperatures: from 4 to 100 ◦ C. 2.6. Thermal stability In order to determine the thermo-stability of cellulase activities, the purified enzyme was incubated at different temperatures (0, 30, 50, 60, 70, 80 and 100 ◦ C) in the absence of substrate. After keeping them for certain periods of time (15, 30 or 60 min),

B. Amouri, A. Gargouri / Biochemical Engineering Journal 32 (2006) 191–197

193

the residual ␤-glucosidase activity was determined as described above. 2.7. Protein determination Protein concentration was determined by the Bradford method [13], using a Bio-Rad protein assay, with bovin serum albumin as a standard. 2.8. Protein electophoresis and zymogram Polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate (SDS-PAGE) was carried out in 10% polyacrylamide gel following the standard procedure [14]: samples (in a buffer containing 100 mM DTT, 2% SDS, 0.1% bromophenol blue, 10% glycerol) were loaded on the gel, either after or without heat treatment (1 min at 100 ◦ C). Two identical gels were usually run, one of which was stained with Coomassie brillant blue G-250 to visualize the protein bands. The second served for the zymogram: the gel was treated with 20 mM Tris–HCl pH 8 for 2 h in order to re-naturate proteins by extruding the SDS. The gel was then covered by another gel of the same dimensions containing 2% agar in 50 mM citrate buffer pH 4.8, 1% CMC and 50 ␮g MUG. The ensemble was incubated for 10 min (for MUGase activity) to 1 h or more (for CMCase) at 37 ◦ C. MUG hydrolysis by the ␤-glucosidase released fluorescent methylumbelliferone, which is visualized under ultraviolet light. The hydrolysis of CMC by endoglucanase was visualized after Congo Red staining of the agar gel followed by de-staining in 1 M NaCl. 2.9. Kinetic properties The Michaelis–Menten constant (Km ) and the maximal reaction velocities (Vmax ) were determined for the ␤-glucosidase by incubating in acetate buffer (pH 4.8) at 50 ◦ C with pNPG, cellobiose, salicin or CMC in concentrations ranging from 1.5 to 5 mM pNPG, 0.25–10 mM cellobiose, 5–20 g/l salicin and from 0.5 to 2% CMC, respectively. The reactions were stopped after different times and p-nitrophenol (pNP), glucose and reducing sugars were measured at standard assays conditions. Values for Km and Vmax were determined from Lineweaver–Burk plots. 2.10. Protein sequencing The purified enzyme was applied to 10% SDS-PAGE followed by electro-blotting onto polyvinylidenedifluoride (PVDF) membrane (Amersham). The protein band was excised and subjected to N-terminal sequence analysis using an automated protein sequencer (Procise 492 cLc, Applied Biosystem). 3. Results and discussion 3.1. Enzyme purification Fed-batch cultivation has been widely used as a successful mode to obtain high levels of product yields [15]. The A19 mutant was examined for cellulases production in such culture system, with two feeding by cellulose Avicel at 4 and 7

Fig. 1. Cellulases activities during the fed-batch culture of the mutant A19. The arrows indicate the first feeding (after 4 days of batch culture) and the second feeding (after 7 days). Operating volume: 10 l (Mandels medium with 2% cellulose Avicel). Temperature 30 ◦ C; pH 5.

days (Fig. 1). The cellulases activities obtained were two to five times more important than those achieved in Erlenmeyer culture. The comparison of activities at day 4 (just before the first feeding) and at the end of the culture shows the effect of feeding on the enzyme production (Fig. 1). Without feeding, we obtain currently about the fourth of the enzymes yield obtained with feeding. The extracellular proteins of the A19 were precipitated by ammonium sulfate dissolved and desalted on a G25-Sephadex column. Protein fractions were pooled and applied on a MonoQ FPLC column. As shown on Fig. 2A, few peaks of proteins were observed and isolated for further characterization and purification steps. The determination of enzyme activities and zymogram analysis showed that three fractions (M4, M9 and M11) were active on pNPG and MUG (␤-glucosidase specific substrates). Fig. 2B shows the zymogram analysis: after migration in denaturing conditions, the gel was soaked in Tris–HCl in order to drain the denaturant (␤-mercaptoethanol and SDS) of the gel and to allow the renaturation of proteins, then it is applied to substrate-agar overlay. It is interesting to note that fractions M4, M9 and M11 recovered MUGase activity only when they were not subjected to heat denaturation, prior to loading on SDSPAGE gel (Fig. 2B). The fractions M4 and M11 contained two pNPGases with small sizes. We do not yet know if they derive from proteolytic cleavage of the large ones or if they are true small ␤glucosidases. Such small ␤-glucosidases are very rare in fungal kingdom, their purification is underway.

194

B. Amouri, A. Gargouri / Biochemical Engineering Journal 32 (2006) 191–197

ing to gel analysis. The molecular weight of the M9 protein was determined by SDS-PAGE to be around 85 kDa (Fig. 3). On native gel, a single band was clearly visible at the same level either on Coomassie stained gel (Fig. 3C) or on the zymogram, in presence of MUG (Fig. 3D). The zymogram of the SDS-PAGE gel revealed another feature of this ␤-glucosidase: once denatured, before loading on gel, the protein is no more able to recover its activity even after an extensive wash of the gel before its transfer on the overlay; whereas the non-heated protein recovered its MUGase activity (Fig. 3B). This observation was already reported for some plant and fungal ␤-glucosidase [16]. It is interesting to note that the corresponding active protein, revealed on the Coomassie stained SDS-PAGE gel, migrated more slowly than the fully denatured M9 protein (Fig. 3B). This migration behavior could be interpreted as a consequence of a dimeric form of the protein, as already observed in some plant and fungal ␤-glucosidase [16]. The M9 fraction was further purified on a gel filtration HPLC column to get rid of minor contaminant proteins and to verify the molecular weight. A major peak named p21 was observed is in full agreement with this conclusion since it has 160 kDa of molecular weight. p21 was collected, characterized and served for the following experiments. Table 1 summarizes the purification assessment of the ␤-glucosidase. Fig. 2. (A) Chromatogram of A19 cellulases using a MonoQ FPLC, previously equilibrated with 20 mM Tris–HCl pH 8. A gradient of 0–1 M NaCl 20 mM Tris–HCl was applied. The proteins were followed by absorbance at 280 nm. (B) Zymogram analysis of different enzyme fraction. After electrophoresis, the acrylamide gel was overlaid with an agar film containing MUG and CMC substrates; incubated 10 min at 37 ◦ C and photographed under UV (to reveal MUGase or ␤-glucosidase activity). SH, ␤-mercaptoethanol.

3.2. Characterization of the fraction M9 In this paper, we focused on the fraction M9 which was roughly pure after a single MonoQ purification step, accord-

3.3. Optimum of pH and temperature of the purified enzyme The purified enzyme was incubated at 50 ◦ C for 30 min in presence of the substrate (pNPG) at different pH, ranging from 3 to 9. The optimum pH of this fraction was determined to be around five (Fig. 4A). The optimum of temperature was also determined and shown to be at 50 ◦ C (Fig. 4B). Such optima are currently encountered in ␤-glucosidases from a wide range of fungi but it is quite interesting to note the partial resistance of this enzyme to alkaline pH. It is worth to note also that at the extreme temperatures (0 and 100 ◦ C); the enzyme lost only the half of its

Fig. 3. Zymogram characterization of the M9 fraction under denaturing and non-denaturing conditions: (A) Coomassie staining of the SDS-PAGE; (B) photo of the agar overlay under in the denaturing conditions; (C) Coomassie staining of the native gel; (D) photo of the agar overlay under ultraviolet in the non-denaturing conditions. UV: T, sample treated at 100 ◦ C for 1 min; U: untreated sample; M: molecular weight marker (kDa).

B. Amouri, A. Gargouri / Biochemical Engineering Journal 32 (2006) 191–197

195

Table 1 Purification of the ␤-glucosidase from A19 strain Purification step

Total protein (mg)

Total activity (U)a

Sp. act (U/mg)

Purif. (fold)

Recovery (%)

G25-Sephadex MonoQ (M9) HPLC (P21)

55 3 0.9

3.3 0.33 0.29

0.06 0.11 0.33

1 1.83 5.50

100 10 8.78

a

The substrate used for enzyme assays was pNPG.

Fig. 4. (A) Determination of the optimum of pH and temperature of the p21 ␤-glucosidase activity. The enzyme activity was tested in presence of pNPG at different pH, ranging from 3 to 9 (A) and at different temperature, from 4 to 100 ◦ C (B). (B) Thermal stability of the p21 fraction after 15, 30 and 60 min incubation at 50 ◦ C.

activity during 30 min of incubation time, a common property for M9 and p21 fraction. We verified that this property was not found in other enzymes that we are currently purifying from the same strain. Such characteristics are not common amongst the majority of the known ␤-glucosidases. The enzyme was found to be stable up to 50 ◦ C after 60 min incubation (Fig. 4C). 3.4. The purified enzyme M9 is a true β-glucosidase To verify if the purified protein is a true ␤-glucosidase, it was incubated with cellobiose as a substrate at pH 5.2 at 50 ◦ C and the reaction products were followed by HPLC analysis. Fig. 5 shows that the cellobiose was indeed converted to glucose monomers in presence of M9 fraction. This activity is very low since only 38% of cellobiose was hydrolyzed after 24 h of incubation.

Fig. 5. Conversion of cellobiose to glucose monomers by the M9 ␤-glucosidase.

3.5. Kinetic properties

Substrate

Vmax (U/mg)

Km (mM)

pNPG Cellobiose Salicin CMC

78 59.4 2 14.8

0.3 2.2 37.1 6.2

Table 2 summarizes the apparent Michaelis–Menten constant and the maximal reaction velocity of the p21 fraction. This ␤-glucosidase shows the highest velocity and affinity towards

Table 2 Kinetic parameters of the ␤-glucosidase p21

196

B. Amouri, A. Gargouri / Biochemical Engineering Journal 32 (2006) 191–197

Fig. 6. Comparison of the N-terminal sequence of p21 with known glycosidase: (A) with a putative cellulase from A. bisporus (GenBank accession no. CAC02964), (B) with the ␤-glucosidase BGLII of H. jecorinae (GenBank accession no. BAA74959) and BGL4 H. grisea (GenBank accession no. BAA74958). The identical amino acids were indicated by an asterisk and the conserved by two or one point.

pNPG, compared to salicin, cellobiose and CMC. The affinity constant for pNPG is comparable to other ␤-glucosidases [10]. 3.6. Amino acid sequencing The purified p21 fraction was applied to a SDS-PAGE, followed by a transfer onto a PVDF membrane, in order to be sequenced. The N-terminal sequence, NH2-YDGENVRIGGRGSFVPGISFHVPTGVNLAY, was Blast compared to the available banks and showed a significant homology with a putative cellulase from Agaricus bisporus (46.6% of identity, Fig. 6A) and family 1 ␤-glucosidases of Hypocrea jecorina and Humicola grisea (23.3 and 20%, respectively, Fig. 6B). It is worth to note that these homologies, as expected for mature and secreted proteins, start just after the signal peptide of the other known glycosidases. The N-terminus sequence determination confirmed, not only that our enzyme is a membership of the ␤-glucosidases, but that it is particular since it resembles to two types of enzymes. The first one, presenting the highest homology, is a putative cellulase, 806 amino acids long, which is not yet identified (Morales and Thurston, unpublished results) but could be an endoglucanase as suggested by its Blast comparison to protein banks. The second class is represented by two ␤-glucosidases belonging to family 1 of glycosyl hydrolases and having a molecular weight of 50 kDa, the first one is an intracellular one encoded by bgl2 gene of Hypocrea jecorina the second one is an extracellular ␤-glucosidase from H. grisea [17]. According to these authors, it would seem likely that H. grisea BGL4 was also originally an intracellular enzyme and that a mutation in its N-terminal region have turned this region into a signal sequence [17]. It is therefore very intriguing that our extracellular enzyme, shares with these family 1 enzymes the N-terminal sequence homology and shares the molecular weight (around 85–90 kDa for the monomer) with the family 3 ␤-glucosidase encoded by bgl1 gene of H. jecorina [18]. 4. Conclusion We describe the purification of one ␤-glucosidase of a novel fungal strain. This fungus possesses a variety of ␤-glucosidases since it has at least three of high molecular weight and two smaller ones; whereas the majority of fungal strains contain usually one or two ␤-glucosidases. After a single FPLC MonoQ step and HPLC gel filtration step, we purified one ␤-glucosidase enzyme, to an electrophoretically homogeneous state. On SDS-PAGE, it presented a single

band of 85 kDa, a molecular weight comparable to the majority of ␤-glucosidases. This enzyme was shown to be dimeric and active on three different ␤-glucosidase specific substrates: pNPG, MUG and cellobiose. Zymogram analysis showed that the purified enzyme resisted to the denaturation by SDS and reducing agents as long as it is not heat denatured. According to its N-terminal sequence, the purified enzyme belongs to the ␤-glucosidases family. It is interesting to note that it shares the molecular weight with family 1 and the N-terminal sequence homology with family 3 of ␤-glucosidases [18]. Acknowledgements This work is dedicated to the memory of our colleagues Basma Hentati, Nejib Trigui and Imed Elloumi. We are indebted to Hafedh Majdoub, Facult´e des Sciences de Sfax, for his help in amino acid sequencing. We deeply thank Najla Masmoudi and H´edi Aouissaoui for technical help and Hafedh Belghith, Raja Mokdad-Gargouri and Noomen Hadj-Ta¨ıeb for their help in manuscript correction and English improvement. This work was supported by the “Contrat-Programme” governmental funds provided by the “Minist`ere de la Recherche Sientifique, de la Technologie et du D´eveloppement des Comp´etences” of Tunisia. References [1] T. Wang, X. Liu, Q. Yu, X. Zhang, Y. Qu, P. Gao, T. Wang, Directed evolution for engineering pH profile of endoglucanase III from Trichoderma reesei, Biomol. Eng. 22 (2005) 89–94. [2] S. Takashima, A. Nakamura, H. Masaki, T. Uozumi, Cloning, sequencing, and expression of a thermostable cellulase gene of Humicola grisea, Biosci. Biotechnol. Biochem. 61 (1997) 245–250. [3] T.M. Enari, in: N.M. Fogarty (Ed.), Microbial Cellulases. I. Microbial Enzymes and Biotechnology, Applied Science Publisher, London, 1983, pp. 183–223. [4] Y. Bourne, B. Henrissat, Glycoside hydrolases and glycosyltransferases: families and functional modules, Curr. Opin. Struct. Biol. 11 (2002) 593–600. [5] R. Bhikhabhai, G. Johansson, G. Pettersson, Cellobiohydrolase from Trichoderma reesei: internal homology and prediction of secondary structure, Int. J. Peptide Protein Res. 25 (1984) 368–374. [6] H. Van Tilbeurg, P. Tomme, M. Claeyssens, R. Bhikhabhai, G. Pettersson, Limited proteolysis of cellobiohydrolase I from Trichoderma reesei, FEBS Lett. 204 (1986) 223–227. [7] H. Belghith, S. Ellouz-Chaabouni, A. Gargouri, Biostoning of denims by Penicillium occitanis (Pol6) cellulases, J. Biotechnol. 89 (2–3) (2001) 257–262. [8] T. Hansson, P. Ablercreutz, Enzymatic synthesis of hexyl glycosides from lactose at low water activity and high temperature using hyperthermostables ␤-glucosidases, Biocatal. Biotrans. 220 (2002) 167–178.

B. Amouri, A. Gargouri / Biochemical Engineering Journal 32 (2006) 191–197 [9] C.S. Gong, M.R. Ladisch, G.T. Tsao, Cellobiase from Trichoderma viride: purification, properties, kinetics, and mechanism, Biotechnol. Bioeng. 19 (7) (1977) 959–981. [10] M. Hidalgo, J. Steiner, J. Eyzaguirre, ␤-Glucosidase from Penicillium purpurogenum: purification and properties, Biotechnol. Appl. Biochem. 15 (2) (1992) 185–191. [11] M. Mandels, F.W. Parrish, E.T. Reese, Sophorose as an inducer of cellulase in Trichoderma viride, J. Bacteriol. 83 (1962) 400–408. [12] B. Amouri, N. Fradi, A. Gargouri, Design of a fungal cellulose-binding domain for PCR amplification and expression in E. coli, Biotech. Lett. 23 (2001) 1883–1888. [13] M.M. Bradford, A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal. Biochem. 72 (1976) 248–254.

197

[14] U.K. Laemmli, M. Favre, Maturation of the head of bacteriophage T4. I. DNA packaging events, J. Mol. Biol. 80 (1973) 575–592. [15] A. Whitaker, Fed-batch culture, Process Biochem. 15 (1980) 10–15. [16] E. Asim, Gunay, Biochemistry and Molecular Biology, in: E. Asim (Ed.), ␤-Glucosidases, Elsevier, Americain Chemical Society, Washington, 1992, pp. 215–239. [17] M. Saloheimo, J. Kuja-Panula, E. Ylosmaki, M. Ward, M. Penttila, Enzymatic properties and intracellular localization of the novel Trichoderma reesei ␤-glucosidase BGLII (cel1A), Appl. Environ. Microbiol. 68 (2002) 4546–4553. [18] C.C. Barnett, R.M. Berka, T. Fowler, Cloning and amplification of the gene encoding an extracellular ␤-glucosidase from Trichoderma reesei: evidence for improved rates of saccharification of cellulosic substrates, Biotechnology 9 (1991) 562–567.