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Glycobiology Advance Access originally published online on July 28, 2006
Glycobiology 2006 16(11):1064-1072; doi:10.1093/glycob/cwl026
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© The Author 2006. Published by Oxford University Press. All rights reserved. For permissions, please e-mail: journals.permissions@oxfordjournals.org

Exo-ß-D-glucosaminidase from Amycolatopsis orientalis: catalytic residues, sugar recognition specificity, kinetics, and synergism

Tamo Fukamizo1,3, Alain Fleury4, Nathalie Côté2,4, Masaru Mitsutomi5 and Ryszard Brzezinski4

3 Department of Advanced Bioscience, Kinki University, 3327-204 Nakamachi, Nara 631-8505, Japan; 4 Centre d’Étude et de Valorisation de la Diversité Microbienne, Département de Biologie, Faculté des Sciences, Université de Sherbrooke, Sherbrooke (Québec) J1K 2R1, Canada; and 5 Department of Applied Biological Sciences, Saga University, Saga 840-8502, Japan

1 To whom correspondence should be addressed; e-mail: fukamizo{at}nara.kindai.ac.jp

2 Present address: ISM Biopolymer Inc., Granby, QC, Canada

Received on May 24, 2006; revised on July 20, 2006; accepted on July 21, 2006


   Abstract
Top
 Abstract
Introduction
 Results and Discussion
 Materials and methods
 Conflict of interest statement
 Acknowledgments
 References
 
Catalytic residues and the mode of action of the exo-ß-D-glucosaminidase (GlcNase) from Amycolatopsis orientalis were investigated using the wild-type and mutated enzymes. Mutations were introduced into the putative catalytic residues resulting in five mutated enzymes (D469A, D469E, E541D, E541Q, and S468N/D469E) that were successfully produced. The four single mutants were devoid of enzymatic activity, indicating that Asp469 and Glu541 are essential for catalysis as predicted by sequence alignments of enzymes belonging to GH-2 family. When mono-N-acetylated chitotetraose [(GlcN)3-GlcNAc] was hydrolyzed by the enzyme, the nonreducing-end glucosamine unit was produced together with the transglycosylation products. The rate of hydrolysis of the disaccharide, 2-amino-2-deoxy-D-glucopyranosyl 2-acetamido-2-deoxy-D-glucopyranose (GlcN-GlcNAc), was slightly lower than that of (GlcN)2, suggesting that N-acetyl group of the sugar residue located at (+1) site partly interferes with the catalytic reaction. The time-course of the enzymatic hydrolysis of the completely deacetylated chitotetraose [(GlcN)4] was quantitatively determined by high-performance liquid chromatography (HPLC) and used for in silico modeling of the enzymatic hydrolysis. The modeling study provided the values of binding free energy changes of +7.0, –2.9, –1.8, –0.9, –1.0, and –0.5 kcal/mol corresponding, respectively, to subsites (–2), (–1), (+1), (+2), (+3), and (+4). When chitosan polysaccharide was hydrolyzed by a binary enzyme system consisting of A. orientalis GlcNase and Streptomyces sp. N174 endochitosanase, the highest synergy in the rate of product formation was observed at the molar ratio 2:1. Thus, GlcNase would be an efficient tool for industrial production of glucosamine monosaccharide.

Key words: catalytic residue / exo-ß-d-glucosaminidase / specificity / subsites / synergism


   Introduction
Top
 Abstract
 Introduction
Results and Discussion
 Materials and methods
 Conflict of interest statement
 Acknowledgments
 References
 
Chitin and chitosan are widely distributed in living organisms, including insects, crustaceans, and fungi. In such organisms, chitinases and chitosanases are directly involved in biological phenomena important for their life, such as moulting, morphogenesis, and aggressive and defensive actions toward the targets (Flach et al., 1992Go). It is now widely accepted that two independent chemical processes catalyzed, respectively, by endochitinases and exo-ß-N-acetyl-D-glucosaminidases are involved in chitin degradation (Fukamizo and Kramer, 1985Go; Horsch et al., 1997Go). A similar mechanism has been considered for chitosan degradation: endochitosanases randomly hydrolyze chitosan polysaccharide, producing oligosaccharides, which are then hydrolyzed into the monosaccharide units by exo-ß-D-glucosaminidase (GlcNase).

For endochitosanases, amino acid sequences have recently been accumulated (http://pages.usherbrooke.ca/rbrzezinski/index.html), and the X-ray crystal structures of several endochitosanases have been reported (Marcotte et al., 1996Go; Saito et al., 1999Go; Adachi et al., 2004Go). On the basis of the structural information, the molecular mechanism of chitosan hydrolysis has been discussed in detail. Site-directed mutagenesis studies of these chitosanases provided information on the catalytic and substrate recognition mechanisms (Boucher et al., 1995Go; Tremblay et al., 2001; Fukamizo et al., 2000Go). On the contrary, very few studies have so far been dedicated to GlcNases. GlcNase was first purified from an actinomycete, Nocardia orientalis (present name: Amycolatopsis orientalis), and characterized by Nanjo et al. (1990)Go. The enzyme was found to specifically hydrolyze the ß-1,4-glucosaminide linkage of the non-reducing-end 2-amino-2-deoxy-D-glucopyranose (GlcN) residue, producing the monosaccharide unit. After this characterization, several GlcNases were purified from culture filtrates of filamentous fungi: Trichoderma reesei PC-3–7 (Nogawa et al., 1998Go), Penicillium funiculosum KY616 (Matsumura et al., 1999Go), and Aspergillus oryzae IAM2660 (Zhang et al., 2000Go), and their enzyme functions were characterized. However, the structural data of the enzymes are very limited; hence, the structure–function relationships of GlcNases have not been understood yet.

On the contrary, Tanaka et al. (2003)Go reported the primary structure and properties of GlcNase from Thermococcus kodakaraensis KOD1. This was the first report on the structure of GlcNase. In our previous article (Côté et al., 2006Go), we reported the deduced amino acid sequence of GlcNase from A. orientalis (GenBank/EBI/DDBJ accession number AY962188 [GenBank] ). The enzyme belongs to family GH-2 according to the CAZy database (http://afmb.cnrs-mrs.fr/CAZY/fam/acc_GH.html) and has a unique modular structure consisting of a sugar-binding domain, an immunoglobulin-like ß-sandwich domain, and a TIM barrel catalytic domain. In addition, these domains are followed by a putative carbohydrate-binding module belonging to family CBM-6. Very recently, the deduced amino acid sequence of GlcNase from T. reesei PC-3-7 was reported (Ike et al., forthcomingGo), and the enzyme was found to belong to family GH-2 as well. The structures of A. orientalis and T. reesei GlcNases are considerably different from that of the T. kodakaraensis GlcNase, which consists of the family GH-35 domain and the family GH-42 domain (Tanaka et al., 2003Go). On the basis of sequence alignments of A. orientalis GlcNase with other GH-2 members, we suggested that Asp469 and Glu541 are the putative catalytic residues (Côté et al., 2006Go). In the present study, to further extend the functional data of GlcNase from A. orientalis, we investigated the catalytic residues, sugar recognition specificity, and kinetic properties of the enzyme, using the wild-type and mutated enzymes. Moreover, using the binary enzyme system consisting of A. orientalis GlcNase and endochitosanase from Streptomyces sp. N174, the rate of product formation from chitosan hydrolysis was measured to examine the synergistic effect resulting from mixing the two enzymes.


   Results and Discussion
Top
 Abstract
 Introduction
 Results and Discussion
Materials and methods
 Conflict of interest statement
 Acknowledgments
 References
 
Catalytic residues
In the previous article (Côté et al., 2006Go), we reported that two GlcNases define a new subfamily within family GH-2 glycosyl hydrolases. Sequence alignment of the GH-2 enzymes revealed that most GH-2 members possess a catalytically important NE diad, which is substituted with an SD diad in the two GlcNases from A. orientalis and T. reesei (Ike et al., forthcomingGo), and also in the SAV1223 protein from Streptomyces avermitilis, shown to possess GlcNase activity. The SD diad corresponds to Ser468 and Asp469 in the Amycolatopsis GlcNase. A strictly conserved glutamic acid was found at the 541st position of A. orientalis GlcNase and is supposed to be the catalytic nucleophile. Thus, we mutated Ser468, Asp469, and Glu541 residues of GlcNase to elucidate their role. We successfully produced and purified the wild-type and five mutated enzymes (D469E, D469A, E541D, E541Q, and S468N/D469E). CD spectra of these enzymes are shown in Figure 1. The profiles of the spectra of D469E and S468N/D469E are different from that of the wild type in the region of 205–215 nm. The mutation of Asp469 to glutamic acid would affect the secondary structure of the enzyme. In the spectra for the other mutated enzymes, however, the profiles are basically identical to that of the wild type, indicating that the global conformation of the enzyme is not affected by these mutations. Enzymatic activities of these enzyme preparations were determined by quantifying the liberated GlcN from the substrate (GlcN)2. The mutation of Asp469 to alanine completely abolished the enzymatic activity as listed in Table I, whereas D469E exhibited very low enzymatic activity. Both the E541-mutated enzymes were found to possess very low activity, but the activity of E541D is significantly larger than that of E541Q. These results clearly demonstrated that Asp469 and Glu541 are the catalytic acid and base, respectively. The double mutation of Ser468 and Asp469 (SD diad) to Asn and Glu (NE diad), respectively, also abolished the GlcNase activity. In most of the members of family GH-2, which possess endo-ß-mannosidase, ß-glucuronidase, and ß-galactosidase activities, the SD diad is substituted with NE (Côté et al., 2006Go). Thus, the double mutant was assayed with the substrates, ß-galactoside, ß-mannoside, and ß-glucuronide, but no activity was detected toward any of these substrates (data not shown). Some other structural factors might participate in the catalytic activity of the family GH-2 enzymes with the NE diad.


Figure 1
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  		Fig. 1. Far-UV CD spectra of the wild-type and mutated exo-ß-glucosaminidase (GlcNase) from Amycolatopsis orientalis. The spectra were recorded in 50 mM sodium acetate buffer pH 5.3 by a Jasco J-720 spectropolarimeter at 20°C. Protein concentration was 3–10 µM.

 

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  		Table I. Enzymatic activities of the wild-type and mutated exo-ß-glucosaminidase from Amycolatopsis orientalis

 

Enzymatic hydrolysis of (GlcN)3-GlcNAc, as determined by TLC and HPLC
When the enzymatic hydrolysis of mono-N-acetylated chitotetraose [(GlcN)3-GlcNAc] was monitored by thin-layer chromatography (TLC), the enzyme was found to produce predominantly GlcN, as shown in Figure 2. If the enzyme acted toward the ß-glucosaminide linkage of the reducing end residue, the enzymatic hydrolysis would not take place, as the N-acetyl group attached to the reducing end of the substrate would interfere with the enzyme attack. Thus, the enzyme appears to split off the monosaccharide unit from the nonreducing end. Tanaka et al. (2003)Go reported a similar cleavage mode for T. kodakaraensis GlcNase using sugar-alcohol derivatives of GlcN oligosaccharides. GlcNase from T. reesei PC-3–7 has been reported to have the same cleavage mode (Nogawa et al., 1998Go).


Figure 2
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  		Fig. 2. TLC profile showing the enzymatic hydrolysis of (GlcN)3-GlcNAc. The enzymatic reaction was performed in 50 mM sodium acetate buffer, pH 5.0, at 37°C. Other details of the reaction conditions are described in the text. GlcN and (GlcN)2–6 were used as the standard. Open and shaded circles represent GlcN and GlcNAc residues, respectively.

 

The enzymatic reaction was monitored by high-performance liquid chromatography (HPLC) using detection by UV absorption (220 nm) originating from the N-acetyl groups of the substrate and the products. The resulting time-dependent HPLC profiles are shown in Figure 3. The initial substrate (GlcN)3-GlcNAc was at first degraded into (GlcN)2-GlcNAc, and then into GlcN-GlcNAc, but no N-acetylglucosamine monomer was produced at this stage. This indicates again that the enzyme hydrolyzes the substrate from the nonreducing end in an exo-splitting manner. After a longer incubation period, the substrate was finally hydrolyzed into monosaccharides, GlcN and GlcNAc (data not shown). To evaluate the sugar recognition specificity at (+1) site, we determined the rate of hydrolysis of GlcN-GlcNAc and compared it with that of (GlcN)2. As shown in Figure 4, similar HPLC profiles were obtained for both substrates, suggesting that the (+1) site is specific to neither acetylated nor deacetylated residue. However, closer examination of the profiles indicated that the rate of the degradation of GlcN-GlcNAc is lower than that of (GlcN)2. The N-acetyl group of the sugar bound to (+1) site appears to partly interfere with the catalytic reaction. Because this enzyme does not act toward (GlcNAc)2, the (–1) site must have absolute specificity for GlcN.


Figure 3
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  		Fig. 3. HPLC profiles of the products obtained from enzymatic hydrolysis of (GlcN)3-GlcNAc. The enzymatic reaction was performed in 50 mM sodium acetate buffer, pH 5.0, at 37°C. Other details of the reaction conditions are described in the text. Open and shaded circles represent GlcN and GlcNAc residues, respectively.

 

Figure 4
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  		Fig. 4. HPLC profiles of the products obtained from enzymatic hydrolysis of GlcN-GlcNAc and (GlcN)2. The enzymatic reaction was performed in 50 mM sodium acetate buffer, pH 5.0, at 37°C. Other details of the reaction conditions are described in the text. Open and shaded circles represent GlcN and GlcNAc residues, respectively.

 

It should be noted that the transglycosylation products, whose molecular weights are larger than that of the initial substrate, were detected in significant amounts (Figure 3). The transglycosylation activity was also reported in GlcNase from P. funiculosum KY616 (Matsumura et al., 1999Go). The transglycosylation reaction catalyzed by the enzyme would be useful for producing GlcN-containing oligosaccharide derivatives with unique biological functions.

Experimental time-course of the enzymatic hydrolysis of (GlcN)4
The experimental time-course of the reaction is shown in Figure 5A. At first, the enzyme predominantly produced GlcN and (GlcN)3, which was further degraded into GlcN and (GlcN)2. The course of the degradation exhibited a typical case of an exo-splitting enzyme. The transglycosylation product, (GlcN)5, was also produced, together with a lesser amount of (GlcN)6. It appears that (GlcN)5 is produced by the glycosyl transfer of the transition state GlcN to the initial substrate (GlcN)4, as shown in Figure 6. Similarly, (GlcN)6 appears to be produced by the transfer action to the product (GlcN)5. The reaction time-course quantitatively determined by HPLC was used for the modeling study described below.


Figure 5
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  		Fig. 5. Time-course of the GlcNase-catalyzed reaction toward (GlcN)4. The enzymatic reaction was performed in 50 mM sodium acetate buffer, pH 5.0, at 37°C. Other details of the reaction conditions are described in the text. Panel A, experimental time-course; panel B, calculated time-course.

 

Figure 6
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  		Fig. 6. Reaction model for GlcNase-catalyzed hydrolysis and transglycosylation reaction. E and Mn represent the enzyme and the substrate with a polymerization degree of n, respectively. Notations of individual complexed states, Ai, Bi,j, and Cn,i, are schematically described in the figure. The individual reaction steps are described in the text. Reaction parameters consist of three rate constants, k+1, k-1, and k+2, and six binding free energy changes of individual subsites, (-2)~(+4). In practical calculation, all the possible binding modes were taken into consideration.

 

In silico modeling of the enzymatic hydrolysis of (GlcN)4
Masaki et al. (1981aGo, 1981bGo) attempted to model in silico the hydrolytic and transglycosylation reactions catalyzed by lysozyme. The modeling studies yielded the optimized values of the binding free energies of the individual subsites, the rate constant for glycosidic bond cleavage, and the relative efficiencies of transglycosylation (Fukamizo et al., 1986, 2001; Fukamizo, Kuhara, and Hayashi, 1982; Fukamizo, Torikata, et al., 1982). In the present study, we tried to model the GlcNase-catalyzed reaction toward completely de-N-acetylated chitotetraose by the method of Fukamizo et al. (1986)Go using the reaction model shown in Figure 6, where the reaction parameters are composed of six binding free energy changes at the individual subsites, (–2), (–1), (+1), (+2), (+3), and (+4), and three rate constants, k+1 (cleavage of glycosidic linkage), k–1 (transglycosylation), and k+2 (hydration).

At first, we experimentally estimated the turnover numbers (kcat) from the maximal velocity data of the individual oligosaccharide substrates at the saturated condition, and then, the kcat values were directly allocated to k+1. The values are listed in Table II. In the previous article, the turnover number of the GlcNase-catalyzed hydrolysis of chitosan polysaccharide was reported to be 2832/min (47.2/s). The k+1 values for the substrates with higher polymerization degree (Table III) are comparable to the reported value of the turnover number. The individual k+1 values were fixed in the modeling calculation. As reported by Masaki et al. (1981b), the values of k–1 and k+2 could not be determined independently, but the ratio k–1/k+2 could be determined based on the time-course data. Thus, the k+2 value was tentatively fixed at 200/sec, and the k–1 value was optimized based on the time-course. Because A. orientalis GlcNase splits off the monomer unit from the nonreducing end of the substrate, (–2) site should be the most unfavorable for binding of the GlcN residue. Thus, a very high positive value (+7.0 kcal/mol) was tentatively allocated to the binding free energy value of (–2) site. In the modeling calculation, the values of k+1 and the binding free energy of (–2) site were kept constant, and the k–1 value (rate constant for transglycosylation) and the binding free energy values of individual subsites from (–1) to (+4) were optimized based on the experimental time-course (Figure 5A).


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  		Table II. Rate constants and binding free energy values used for the calculation of reaction time-course shown in Figure 4B

 

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  		Table III. Primers used for the mutagenesis of Amycolatopsis orientalis csxA

 

The optimization based on the reaction model shown in Figure 6 successfully produced the k–1 value and the free energy values of the individual subsites as listed in Table II. The time-course calculated with these values coincided satisfactorily with the experimental one as shown in Figure 5B. As to the free energy value of (–2) site, the product distribution of the calculated time-course was greatly affected upon descending the free energy value from +7.0 to +6.0 or +5.0 kcal/mol, resulting in the larger value of the cost function (Equation 1). However, the profile of the calculated time-course was hardly affected upon elevating the value from +7.0 kcal/mol. Thus, the free energy value of (–2) site should be larger than or equal to +7.0 kcal/mol but could not be determined accurately. The affinities for sugar residues were found to be highest at subsites (–1) and (+1), whereas remote subsites were found to have lesser affinities. The free energy distribution in the substrate-binding cleft is very similar to those of glucoamylases possessing similar cleavage specificity (Tanaka et al., 1983Go; Sierks et al., 1989Go; Ichikawa et al., 2004Go).

The model predicts that shorter substrate chain lengths correspond to higher k+1 values. In our previous article (Côté et al., 2006Go), we reported that the specific activities toward (GlcN)2~(GlcN)6 are similar to each other. The lower k+1 values for the longer substrates might be compensated by the higher affinity to the binding cleft. In fact, by simply assuming additivity, the affinity for (GlcN)5 binding to (–1)~(+4) is higher than that for (GlcN)4 binding to (–1)~(+3) by 0.5 kcal/mol, and (GlcN)4 binding to (–1)~(+3) is stronger than (GlcN)3 binding to (–1)–(+2) by 1.0 kcal/mol.

Synergism between endochitosanase and GlcNase
It has been recognized that polysaccharides are enzymatically degraded into monomers by a couple of enzymes with different cleavage specificities (Zhang and Lynd, 2004Go). Fukamizo and Kramer (1985)Go reported that endochitinase and ß-N-acetylglucosaminidase from the moulting fluid of tobacco hornworm, Manduca sexta, concertedly acted on chitin chain resulting in a synergistic effect on chitin degradation. In the enzymatic degradation of cellulose, synergism has been intensively studied using various types of cellulases from T. reesei (Henrissat et al., 1985Go; Riedel et al., 1997Go). Synergism might be found also in chitosan degradation upon mixing the endochitosanase with GlcNase. Thus, we measured the rate of product formation from chitosan by the binary enzyme system consisting of A. orientalis GlcNase and Streptomyces sp. N174 endochitosanase. The result is shown in Figure 7. In each experiment, the amount of product formed from the enzymatic reaction was almost equal to or less than 1 mg. Because the initial amount of the substrate was 2.4 mg, the extents of the substrate converted were <40%. Thus, all of these data were regarded as the reaction rate of the initial reaction stage. In the presence of an excess amount of endochitosanase, the rate of product formation was enhanced by about 2-fold of the sum of those obtained by individual enzymes, that is, the synergistic factor was about 2. By increasing the ratio of GlcNase to the endo-splitting enzyme, the synergistic factor was gradually enhanced, and it reached maximum (4.1) when the enzyme molar ratio was 2:1 (exo:endo). In the binary chitinase system (exo-ß-N-acetylglucosaminidase and endochitinase) of tobacco hornworm, M. sexta, which is responsible for the destabilization of old cuticle, the greatest synergism of 6 takes place at a 1:6 (exo:endo) ratio of enzymes, typically found in the moulting fluid secreted from epidermal cells (Fukamizo and Kramer, 1985Go). In the cellulase systems studied thus far, the synergistic factor was reported to be 2~10 (Zhang and Lynd, 2004Go). Thus, our binary chitosanase system exhibits a moderate synergism and is efficient for monomer production. In contrast to the binary chitinase system from M. sexta, a higher amount of the exo-enzyme is required to attain the maximum synergism in our binary chitosanase system. The exo-splitting process might be rate-limiting in the tandem action of the chitosan degradation, whereas the endo-splitting process is rate-limiting in the chitin degradation in insects (Fukamizo and Kramer, 1985Go).


Figure 7
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  		Fig. 7. Synergistic effect obtained from mixing GlcNase with endo-splitting chitosanase from Streptomyces sp. N174. The synergistic factor is equal to the ratio of activity exhibited by mixtures of components divided by the sum of the activities of separate components. The two enzymes dialyzed against 50 mM sodium acetate buffer pH 5.0 were mixed together with various exo/endo ratios, and the enzyme mixture solution (340 µL) was incubated with chitosan solution (15 mg/mL, 160 µL) for 2 h at 37oC.

 

An endo-splitting chitosanase was isolated from A. orientalis and enzymatically characterized (Sakai et al., 1991Go). The enzyme can hydrolyze the ß-1,4-linkage of GlcN-GlcNAc in addition to that of GlcN-GlcN, whereas Streptomyces sp. N174 chitosanase hydrolyzes GlcNAc-GlcN in addition to GlcN-GlcN (Fukamizo et al., 1995Go). Thus, some fraction of the products from A. orientalis endochitosanase would consist in oligosaccharides possessing a GlcNAc residue at the nonreducing end. Because such oligosaccharides could not be hydrolyzed by GlcNase, a lower synergism would be obtained when the endo-splitting chitosanase from A. orientalis is used instead of the Streptomyces sp. N174 chitosanase. Thus, the binary chitosanase system used in this study is an efficient tool for the industrial production of glucosamine monomer from chitosan.


   Materials and methods
Top
 Abstract
 Introduction
 Results and Discussion
 Materials and methods
Conflict of interest statement
 Acknowledgments
 References
 
Materials
All glucosamine oligomers (GlcN)2–6 were purchased from Seikagaku Kogyo Co. (Tokyo, Japan). Chitosan 10B (99% de-N-acetylated) was from Funakoshi Chemical Co. (Tokyo, Japan). The chitosan was partially degraded by nitrous acid by the method of Tømmeraas et al. (2001)Go, and the low-molecular-weight chitosan was used as the substrate for GlcNase. (GlcN)3-GlcNAc and GlcN-GlcNAc were obtained by the method of Mitsutomi et al. (2002)Go. Chitosanase from Streptomyces sp. N174 was produced by the expression system of Streptomyces lividans (Masson et al., 1993Go). Culture media components were from Difco (Detroit, MI). All reagents for the enzyme purification were from Sigma (St. Louis, MO). The other reagents were of analytical grade commercially available.

Enzyme production and purification
Wild-type and mutated GlcNases from A. orientalis were produced using the recombinant system established in the heterologous host S. lividans TK24 (Côté et al., 2006Go). Strains were pregrown in YME medium for 24 h at 30oC. The mycelium was recovered by centrifugation (3000g, 10 min 4oC and inoculated (1 mL of mycelial pellet per 100 mL of medium) into M14 medium (Pagé et al., 1996Go) containing 0.2% glucosamine and 0.8% chitosan as carbon sources. After 6 days of culture, supernatant was adjusted to pH 4.2 with glacial acetic acid and filtered on 0.8- and 0.2-µm filters. Supernatant was loaded on an SP Sepharose (Amersham Biosciences, Piscataway, NJ) column equilibrated with 50 mM sodium acetate buffer, pH 4.2. Elution was performed with an NaCl gradient from 0 to 1 M. Desired fractions were dialyzed against 1 mM sodium phosphate, pH 6.8. Hydroxyapatite column was loaded with the dialyzed protein fraction, and the proteins were eluted with an unbuffered MgCl2 gradient from 0 to 1 M. The enzyme solution was finally dialyzed against 50 mM sodium acetate buffer, pH 5.2, and stored at 4oC.

Site-directed mutagenesis and production of mutated enzymes
Site-directed mutagenesis was performed by the method of Ho et al. (1989)Go involving polymerase chain reaction (PCR) using high-fidelity Pfu DNA polymerase (Fermentas, Burlington, ON, Canada). The procedure exploited the fact that the triplets encoding the residues to be mutated (S468, D469, and E541) were localized between unique restriction sites BsmI and EcoRI. In a first series of amplifications, a common primer adjacent to the BsmI site was used with the reverse primer specific for each mutation (Table III). In parallel series of amplifications, a common primer adjacent to the EcoRI site was used with the forward primer specific for each mutation. After purification of both amplicons on agarose gel, a second series of PCR was performed with the same external primers. The resulting mutated amplicons were cloned into the original vector using unique sites BsmI and EcoRI. All mutants were sequenced in both orientations. The mutant proteins were produced and purified by the same procedure as described above (Côté et al., 2006Go).

Enzyme assay
GlcNase assay was performed combining 100 µL of 2 mM (GlcN)2 in 50 mM sodium acetate buffer, pH 5.3, with 100 µL of diluted enzyme preparation. The reaction mixture was incubated for 10 min at 37oC and terminated by the addition of 25 µL of 0.2 M sodium tetraborate. The GlcN liberated from the dimer was N-acetylated by 25 µL of 1.5% acetic anhydride in acetone, and the resultant N-acetylglucosamine was determined by the method of Reissig et al. (1955)Go.

Protein determination
The protein concentration was determined by means of UV absorption at 280 nm using an extinction coefficient calculated from the equation reported by Pace et al. (1995)Go.

CD spectroscopy
The protein solution was dialyzed against 50 mM sodium acetate buffer pH 5.3, and the CD spectra were recorded using a Jasco (Easton, MD) J-720 spectropolarimeter at 20oC.

Time-course of the enzymatic reaction
Enzymatic reaction toward oligosaccharide substrate was performed in 50 mM sodium acetate buffer pH 5.0 at 37oC. Three microliters of the enzyme solution (0.48 µM) was added to 100 µL of the substrate solution (12.3 mM). After an appropriate reaction time, 10 µL of the reaction mixture was combined with the same volume of 0.1 M NaOH to terminate the enzymatic reaction. The resultant solution was analyzed by TLC or HPLC.

TLC
To qualitatively determine the enzymatic products, the enzymatic reaction mixture was spotted on a TLC aluminum plate (Merck, Darmstadt, Germany; Silica gel 60), and the sugars were developed with the solvent system, 28% ammonia solution:1-propanol (1:2). The sugar spots were visualized by spraying a ninhydrin reagent, followed by heating at 90oC.

HPLC
The oligosaccharide products obtained from enzyme digestion were determined by a gel-filtration HPLC on TSK-GEL G2000PW column (7.5 x 600 mm, Tosoh, Tokyo, Japan) using a Hitachi L-7100 intelligent pump. Elution was achieved with 0.1 M NaCl at room temperature at a flow rate of 0.3 mL/min. Mono-N-acetylated oligosaccharides were monitored by their absorbance at 220 nm using a Hitachi (Tokyo, Japan) L-7405 UV detector, and completely de-N-acetylated oligosaccharides were monitored with a Hitachi L-3350 RI monitor.

In silico modeling of the enzymatic hydrolysis of chitotetraose
In silico modeling of the enzymatic hydrolysis of chitotetraose was carried out by the method of Fukamizo et al. (1986)Go using the reaction model shown in Figure 6. E and Mn represent the enzyme and the substrate with a polymerization degree of n, respectively. Notations of individual complexed states, Ai, Bi,j, and Cn,i, are schematically described in the figure. The initial substrate binds to the enzyme with various binding modes to form Cn,i, which are then converted into Bi,n–i through the cleavage of glycosidic linkage (k+1). The complex Ai is formed by releasing the oligosaccharide fragment bound to subsites (+1), (+2), (+3), and (+4). An activated water molecule attacks Ai (k+2) to complete the hydrolytic reaction. On the contrary, the transglycosylation acceptor can bind again to subsites (+1), (+2), (+3), and (+4) to form the complex Bi,j, and then a new glycosidic linkage is formed (k–1) to complete the transglycosylation reaction. Reaction parameters consist of three rate constants, k+1, k–1, and k+2, and six binding free energy changes of individual subsites, (–2)~(+4). In practical calculations, all the possible binding modes were defined by the suffix, n, i, and j, of the notations, Cn,i, Bi,j, and Ai, and taken into consideration. Because the enzyme hydrolyzes the oligosaccharide substrate from the nonreducing end with an exo-splitting mode, the binding cleft of the enzyme was assumed to consist of the subsites (–2), (–1), (+1), (+2), (+3), and (+4), where the (–2) site should have an unfavorable positive binding free energy. By assuming rapid equilibrium, the concentrations of the ES-complexes (Cn,i, Bi,j, and Ai) were calculated from the binding constants, which were obtained from the binding free energy values of individual subsites occupied with the sugar residues assuming additivity. Details of the calculation method were described by Fukamizo et al. (1986)Go.

To estimate the values of rate constants and binding free energies at individual subsites, an optimization technique based on the modified Powell method (Kuhara et al., 1982Go) was employed using the cost function:



Formula

where e and c represent the experimental and calculated values, respectively, n is the size of the oligosaccharides, and i the reaction time.

Chitosan hydrolysis by the binary enzyme system of GlcNase and endo-splitting chitosanase
To examine synergistic effect resulting from mixing A. orientalis GlcNase with endo-splitting chitosanase from Streptomyces sp. N174, we determined the rate of product formation from the low-molecular-weight chitosan by the binary enzyme system. At first, the two enzymes, dialyzed against 50 mM sodium acetate buffer pH 5.0, were mixed together at various exo/endo ratios, and the enzyme mixture solution (340 µL) was added to the chitosan solution (15 mg/mL, 160 µL) to start the enzymatic reaction. The reaction mixture was incubated for 2 h at 37oC. The enzymatic product was determined by the modified Elson–Morgan method with glucosamine as standards (Rondle and Morgan, 1955Go).


   Conflict of interest statement
Top
 Abstract
 Introduction
 Results and Discussion
 Materials and methods
 Conflict of interest statement
Acknowledgments
 References
 
None declared.


   Acknowledgments
Top
 Abstract
 Introduction
 Results and Discussion
 Materials and methods
 Conflict of interest statement
 Acknowledgments
References
 
This work was supported by a grant-in-aid for scientific research (C, 17580085) from the Japan Society for the Promotion of Science (JSPS) and supported in part by an "Academic Frontier" Project for Private Universities: matching fund subsidy from MEXT (Ministry of Education, Culture, Sports, Science and Technology), 2004–2008. Work at Sherbrooke was supported by a Discovery Grant from the Natural Sciences and Engineering Research Council of Canada.


   Abbreviations
 
(GlcN)3-GlcNAc, mono-N-acetylated chitotetraose (the reducing end residue is N-acetylated); GlcN, 2-amino-2-deoxy-D-glucopyranose; GlcNAc, 2-acetamido-2-deoxy-D-glucopyranose; GlcNase, exo-ß-D-glucosaminidase; HPLC, high-performance liquid chromatography; TLC, thin-layer chromatography


   References
Top
 Abstract
 Introduction
 Results and Discussion
 Materials and methods
 Conflict of interest statement
 Acknowledgments
 References
 
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