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• W2025968524 abstract "The gene encoding family 8 glycoside hydrolases from Bacillus halodurans C-125 (BH2105), an alkalophilic bacterium with a known genomic sequence, was expressed in Escherichia coli. The protein was expressed with the intact N-terminal sequence, suggesting that it did not possess a signal peptide and that it was an intracellular enzyme. The recombinant enzyme showed no hydrolytic activity on xylan, whereas it had been annotated as xylanase Y. It hydrolyzed xylooligosaccharide whose degree of polymerization is greater than or equal to 3 in an exo-splitting manner with anomeric inversion, releasing the xylose unit at the reducing end. Judging from its substrate specificity and reaction mechanism, we named the enzyme reducing end xylose-releasing exo-oligoxylanase (Rex). Rex was found to utilize only the β-anomer of the substrate to form β-xylose and α-xylooligosaccharide. The optimum pH of the enzymatic reaction (6.2–7.3) was found in the neutral range, a range beneficial for intracellular enzymes. The genomic sequence suggests that B. halodurans secretes two endoxylanases and possesses two α-arabinofuranosidases, one α-glucuronidase, and three β-xylosidases intracellularly in addition to Rex. The extracellular enzymes supposedly hydrolyze xylan into arabino/glucurono-xylooligosaccharides that are then transported into the cells. Rex may play a role as a key enzyme in intracellular xylan metabolism in B. halodurans by cleaving xylooligosaccharides that were produced by the action of other intracellular enzymes from the arabino/glucurono-xylooligosaccharides. The gene encoding family 8 glycoside hydrolases from Bacillus halodurans C-125 (BH2105), an alkalophilic bacterium with a known genomic sequence, was expressed in Escherichia coli. The protein was expressed with the intact N-terminal sequence, suggesting that it did not possess a signal peptide and that it was an intracellular enzyme. The recombinant enzyme showed no hydrolytic activity on xylan, whereas it had been annotated as xylanase Y. It hydrolyzed xylooligosaccharide whose degree of polymerization is greater than or equal to 3 in an exo-splitting manner with anomeric inversion, releasing the xylose unit at the reducing end. Judging from its substrate specificity and reaction mechanism, we named the enzyme reducing end xylose-releasing exo-oligoxylanase (Rex). Rex was found to utilize only the β-anomer of the substrate to form β-xylose and α-xylooligosaccharide. The optimum pH of the enzymatic reaction (6.2–7.3) was found in the neutral range, a range beneficial for intracellular enzymes. The genomic sequence suggests that B. halodurans secretes two endoxylanases and possesses two α-arabinofuranosidases, one α-glucuronidase, and three β-xylosidases intracellularly in addition to Rex. The extracellular enzymes supposedly hydrolyze xylan into arabino/glucurono-xylooligosaccharides that are then transported into the cells. Rex may play a role as a key enzyme in intracellular xylan metabolism in B. halodurans by cleaving xylooligosaccharides that were produced by the action of other intracellular enzymes from the arabino/glucurono-xylooligosaccharides. Endo-β-1,4-xylanases (EC 3.2.1.8) are glycoside hydrolases (GHs) 1The abbreviations used are: GH, glycoside hydrolase; Rex, reducing end xylose-releasing exo-oligoxylanase; G, glucose; X, xylose; de, deoxy; Xn, xylooligosaccharide whose degree of polymerization is n; TAPS, 3-{[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]amino}-1-propanesulfonic acid; CAPS, 3-(cyclohexylamino)propanesulfonic acid; HPLC, high pressure liquid chromatography; pNP, p-nitrophenyl. that catalyze the degradation of xylan, a main component of hemicelluloses. The enzyme has been classified mainly into GH families 10 and 11 based on the amino acid sequences (1Henrissat B. Biochem. J. 1991; 280: 309-316Crossref PubMed Scopus (2624) Google Scholar) (Carbohydrate-active Enzymes (CAZy) data base, afmb.cnrs-mrs.fr/CAZY/). GH10 is known to have a (α/β)8 barrel structure to be classified as clan GH-A (2Henrissat B. Bairoch A. Biochem. J. 1996; 316: 695-696Crossref PubMed Scopus (1187) Google Scholar, 3Henrissat B. Callebaut I. Fabrega S. Lehn P. Mornon J.P. Davies G. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7090-7094Crossref PubMed Scopus (516) Google Scholar, 4Dominguez R. Souchon H. Spinelli S. Dauter Z. Wilson K.S. Chauvaux S. Beguin P. Alzari P.M. Nat. Struct. Biol. 1995; 2: 569-576Crossref PubMed Scopus (169) Google Scholar, 5White A. Tull D. Johns K. Withers S.G. Rose D.R. Nat. Struct. Biol. 1996; 3: 149-154Crossref PubMed Scopus (187) Google Scholar, 6Guerin D.M. Lascombe M.B. Costabel M. Souchon H. Lamzin V. Beguin P. Alzari P.M. J. Mol. Biol. 2002; 316: 1061-1069Crossref PubMed Scopus (122) Google Scholar, 7Natesh R. Bhanumoorthy P. Vithayathil P.J. Sekar K. Ramakumar S. Viswamitra M.A. J. Mol. Biol. 1999; 288: 999-1012Crossref PubMed Scopus (61) Google Scholar, 8Schmidt A. Gubitz G.M. Kratky C. Biochemistry. 1999; 38: 2403-2412Crossref PubMed Scopus (65) Google Scholar, 9Andrews S.R. Charnock S.J. Lakey J.H. Davies G.J. Claeyssens M. Nerinckx W. Underwood M. Sinnott M.L. Warren R.A. Gilbert H.J. J. Biol. Chem. 2000; 275: 23027-23033Abstract Full Text Full Text PDF PubMed Scopus (41) Google Scholar, 10Fujimoto Z. Kuno A. Kaneko S. Kobayashi H. Kusakabe I. Mizuno H. J. Mol. Biol. 2002; 316: 65-78Crossref PubMed Scopus (74) Google Scholar, 11Canals A. Vega M.C. Gomis-Ruth F.X. Diaz M. Santamaria R.R. Coll M. Acta Crystallogr. Sect. D Biol. Crystallogr. 2003; 59: 1447-1453Crossref PubMed Scopus (23) Google Scholar), while GH11 has a jelly roll structure to be classified as clan GH-C (12Krengel U. Dijkstra B.W. J. Mol. Biol. 1996; 263: 70-78Crossref PubMed Scopus (160) Google Scholar, 13Fushinobu S. Ito K. Konno M. Wakagi T. Matsuzawa H. Protein Eng. 1998; 11: 1121-1128Crossref PubMed Scopus (122) Google Scholar, 14Gruber K. Klintschar G. Hayn M. Schlacher A. Steiner W. Kratky C. Biochemistry. 1998; 37: 13475-13485Crossref PubMed Scopus (133) Google Scholar, 15McCarthy A.A. Morris D.D. Bergquist P.L. Baker E.N. Acta Crystallogr. Sect. D Biol. Crystallogr. 2000; 56: 1367-1375Crossref PubMed Scopus (52) Google Scholar, 16Wouters J. Georis J. Engher D. Vandenhaute J. Dusart J. Frere J.M. Depiereux E. Charlier P. Acta Crystallogr. Sect. D Biol. Crystallogr. 2001; 57: 1813-1819Crossref PubMed Scopus (37) Google Scholar, 17Hakulinen N. Turunen O. Janis J. Leisola M. Rouvinen J. Eur. J. Biochem. 2003; 270: 1399-1412Crossref PubMed Scopus (180) Google Scholar). Although GH10 and GH11 xylanases have different three-dimensional structures, both of these enzymes can produce xylooligosaccharides from xylan in an endo-splitting manner with anomeric retention. Recently a GH8 endo-β-1,4-xylanase was found in a culture supernatant of Pseudoalteromonas haloplanktis and characterized (18Collins T. Meuwis M.A. Stals I. Claeyssens M. Feller G. Gerday C. J. Biol. Chem. 2002; 277: 35133-35139Abstract Full Text Full Text PDF PubMed Scopus (164) Google Scholar). Although the GH8 xylanase also hydrolyzes the β-1,4 glycosidic bond of xylan in an endo-splitting manner, its reaction proceeds with anomeric inversion. The GH8 family contains various endoglycoside hydrolases, such as chitosanase (EC 3.2.1.132), endoglucanase (EC 3.2.1.4), and licheninase (EC 3.2.1.73) as well as endo-β-1,4-xylanase. These enzymes also hydrolyze corresponding polysaccharides with anomeric inversion. The three-dimensional structures of two GH8 enzymes, the endo-β-1,4-xylanase from P. haloplanktis and an endoglucanase from Clostridium thermocellum, were found to have an (α/α)6 barrel structure (clan GH-M) (6Guerin D.M. Lascombe M.B. Costabel M. Souchon H. Lamzin V. Beguin P. Alzari P.M. J. Mol. Biol. 2002; 316: 1061-1069Crossref PubMed Scopus (122) Google Scholar, 19Van Petegem F. Collins T. Meuwis M.A. Gerday C. Feller G. Van Beeumen J. J. Biol. Chem. 2003; 278: 7531-7539Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar). The GH8 endo-β-1,4-xylanase from P. haloplanktis has the highest amino acid identity (32.6%) with the protein encoded by the BH2105 gene (GenBank™ accession number BAB05824) of Bacillus halodurans C-125 (18Collins T. Meuwis M.A. Stals I. Claeyssens M. Feller G. Gerday C. J. Biol. Chem. 2002; 277: 35133-35139Abstract Full Text Full Text PDF PubMed Scopus (164) Google Scholar), which is annotated as “xylanase Y.” B. halodurans C-125 is a xylanase-producing alkalophilic bacterium (20Honda H. Kudo T. Horikoshi K. J. Bacteriol. 1985; 161: 784-785Crossref PubMed Google Scholar) whose genomic sequence is available (21Takami H. Nakasone K. Takaki Y. Maeno G. Sasaki R. Masui N. Fuji F. Hirama C. Nakamura Y. Ogasawara N. Kuhara S. Horikoshi K. Nucleic Acids Res. 2000; 28: 4317-4331Crossref PubMed Scopus (447) Google Scholar). According to the annotation of the genomic sequence, the alkalophilic microbe supposedly possesses three endo-β-1,4-xylanases (GH8, GH10, and GH11). Enzymatic characterizations of two secreted xylanases, XynA (BH2120, GH10) and XynN (probably BH0899, GH11), indicate that XynA exhibits a broad optimal pH range for activity (pH 4–10), whereas XynN (pH 4–6) is active only at neutral acidity (20Honda H. Kudo T. Horikoshi K. J. Bacteriol. 1985; 161: 784-785Crossref PubMed Google Scholar, 22Nishimoto M. Honda Y. Kitaoka M. Hayashi K. J. Biosci. Bioeng. 2002; 93: 428-430Crossref PubMed Scopus (13) Google Scholar). No other endoxylanases have been reported in B. halodurans C125. Thus, we expressed the BH2105 protein in Escherichia coli and characterized the properties of the enzyme. In this study, we found that the BH2105 protein was not an endo-β-1,4-xylanase but has a novel activity of hydrolyzing xylooligosaccharides, releasing xylose from their reducing ends. This result prompted us to characterize the unique enzymatic properties (substrate specificity and reaction mechanism) in detail. Here we describe the characterization of the newly found enzyme named reducing end xylose-releasing exooligoxylanase (Rex). Materials—The B. halodurans C-125 (9153) strain was obtained from the Japan Collection of Microorganisms (Wako, Japan). Restriction endonucleases were obtained from New England Biolabs (Beverly, MA), and the DNA polymerase from Thermococcus kodakaraensis KOD1 was obtained from Toyobo (Osaka, Japan). Birch wood xylan was prepared by lyophilizing the water-soluble fraction of birch wood xylan (Fluka, Buchs, Switzerland) as described previously (23Dupont C. Roberge M. Shareck F. Morosoli R. Kluepfel D. Biochem. J. 1998; 330: 41-45Crossref PubMed Scopus (49) Google Scholar). Chitosan (Sigma), curdlan (Wako Pure Chemicals, Osaka, Japan), lichenan (Sigma), and carboxymethylcellulose (Nacalai Tesque, Kyoto, Japan) were used as purchased. Cellotriose (G3), cellopentaose, laminaripentaose, chitopentaose, and chitosanpentaose were purchased from Seikagaku Kogyo (Tokyo, Japan). Xylooligosaccharides (Xn where n = degree of polymerization) were purchased from Megazyme (Wicklow, Ireland). p-Nitrophenyl-β-d-xylopyranoside (X-pNP) was purchased from Sigma. p-Nitrophenyl-β-d-xylobioside was prepared as described previously (24Kitaoka M. Haga K. Kashiwagi Y. Sasaki T. Taniguchi H. Kusakabe I. Biosci. Biotechnol. Biochem. 1993; 57: 1987-1989Crossref Scopus (25) Google Scholar). β-(1→4) hetero-d-glucose and d-xylose-based disaccharides and trisaccharides (G-X, X-G, G-X-X, X-X-G, G-X-G, X-G-G, G-G-X, and X-G-X (abbreviations indicate the monosaccharide unit from the non-reducing end: G, glucose; X, xylose)) were prepared as described previously (25Shintate K. Kitaoka M. Kim Y.K. Hayashi K. Carbohydr. Res. 2003; 338: 1981-1990Crossref PubMed Scopus (37) Google Scholar). 1,5-Anhydroxylotriitol (1-deoxy-xylotriose, X3-de) was synthesized by reducing acetobromoxylotriose, prepared by acetylation and bromination of X3 with pyridine-acetic anhydride and TiBr4, respectively, followed by reduction with lithium aluminum hydride as described for 1,5-anhydro-d-glucitol (26Reither F.J. J. Am. Chem. Soc. 1945; 67: 1056-1057Crossref Scopus (11) Google Scholar, 27Fletcher Jr., H.G. Methods Carbohydr. Chem. 1963; : 197-198Google Scholar). Other reagents were of analytical grade and were obtained commercially. DNA Manipulation—Recombinant DNA techniques and agarose gel electrophoresis were performed as described by Sambrook et al. (28Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: A Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989Google Scholar). Plasmid DNA was prepared using a QIAprep Spin Plasmid kit (Qiagen, Hilden, Germany). Digestion by restriction enzymes was carried out in the appropriate buffer at concentrations of 1–10 units/μg of DNA for 0.5–16 h at 37 °C. Completion of the reaction was confirmed by agarose gel electrophoresis. A QIAEX Agarose Gel Extraction kit (Qiagen) was used for the extraction and purification of DNA from agarose gels. Nucleotide Sequence Analysis—The nucleotide sequence was determined by the dideoxynucleotide chain termination method using an automated DNA sequencer (Model 310A, Applied Biosystems, Foster City, CA) with a dRhodamine Terminator kit (PerkinElmer Life Sciences). At least three independent clones of each PCR product were sequenced. Sequence data were analyzed using GENETYX MAC software Version 11.0 (GENETYX Software Development Co., Ltd., Tokyo, Japan). Expression of BH2105 in E. coli—The gene encoding BH2105 was amplified from the genomic DNA of B. halodurans C-125 by the polymerase chain reaction using the forward primer 5′-CCT TCC ATG GAG AAA ACG ACA GAA GGT GCA TTT-3′ (containing an NcoI site, denoted by bold type) and the reverse primer 5′-GAA CTC GAG GTG TTC CTC TCT TGG CCC TCA G-3′ (containing an XhoI site, denoted by bold type). Because of the addition of the NcoI site, the second amino acid residue coded was changed into Glu from Lys. The amplified fragment was digested by the corresponding restriction enzymes. The digested fragment was ligated into pET28b (Novagen, Madison, WI) at the corresponding sites, generating the plasmid pET28b-BH2105 encoding the BH2105 protein with the His6 sequence added to its C-terminal end. Next pET28b-BH2105 was electroporated into E. coli BL21(GOLD)(DE3) cells, and positive colonies were selected. Resulting transformants were incubated in Luria broth (100 ml) containing 0.05 mg/ml kanamycin at 37 °C until the optical density, at 600 nm, reached a level of 0.6. Isopropyl-β-d-thiogalactopyranoside was then added to give a final concentration of 1 mm, and the cultures were incubated for 24 h at 25 °C. The BH2105 protein expressed was extracted from the wet cells (1 g) in 5 ml of 50 mm sodium phosphate buffer (pH 8.0) using a sonicator (Model 250D sonifier, Branson, Danbury, CT). Purification of Recombinant BH2105—The cell-free extract was loaded onto a nickel-nitrilotriacetic acid-agarose (Qiagen) column (1 × 3 cm), and the enzyme was eluted with a stepwise gradient of imidazole (1, 10 mm; 2, 20 mm; 3, 250 mm) in 50 mm sodium phosphate buffer (pH 8.0) containing 0.3 m NaCl. The fraction containing the BH2105 protein was desalted using a PD-10 column (Amersham Biosciences). Next the protein solution was loaded onto a Q-Sepharose column (2.5 × 4 cm), and the enzyme was eluted with a stepwise gradient of NaCl (1, 0.05 m; 2, 0.2 m; 3, 0.3 m) in 50 mm sodium phosphate buffer (pH 7.2). The appropriate fractions were collected, and purity was checked by SDS-PAGE (29Laemmli U.K. Nature. 1970; 227: 680-685Crossref PubMed Scopus (207233) Google Scholar). A 10-kDa protein ladder (Invitrogen) was used as a standard molecular marker for SDS-PAGE. Protein concentrations were determined from the absorbance at 280 nm based on the theoretical molar absorption coefficients (106,210 m–1cm–1) determined from the amino acid composition of BH2105 (30Pace C.N. Vajdos F. Fee L. Grimsley G. Gray T. Protein Sci. 1995; 4: 2411-2423Crossref PubMed Scopus (3452) Google Scholar). The N-terminal amino acid sequence of the purified recombinant BH2105 protein was determined using a G1000A protein sequencer (Hewlett Packard, Palo Alto, CA). Enzyme Assay—The enzyme activity was routinely determined by measuring the increase in xylose during the hydrolysis of X3. The enzymatic reaction was carried out in 50 mm sodium phosphate buffer (pH 7.1) containing various concentrations of X3 at 40 °C. Periodically a portion of the reaction mixture was boiled for 5 min to inactivate the enzyme, and the concentration of xylooligosaccharides (X1-X6) was quantified by high performance ion exchange chromatography on a CARBOPAC PA1 column (4 × 250 mm, Dionex, Sunnyvale, CA) equipped with a pulsed amperometric detector (DX-3, Dionex). Chromatography was performed with a linear gradient of 0–0.2 m sodium acetate in 0.1 m NaOH for 20 min at a flow rate of 1 ml/min. Effect of pH and Temperature on Enzymatic Activity—Enzymatic activity was measured under standard conditions of X3 (0.5 mm) hydrolysis, while pH of the reaction mixture was changed with each 50 mm buffer. The pH stability was determined by incubating the enzyme at 30 °C for 30 min at each pH followed by measuring activity under standard conditions. The buffer systems used were sodium acetate (pH 3.5–5.5) and sodium phosphate (pH 6.0–8.0), TAPS (pH 8.0–9.0), and CAPS (pH 9.7–11.0). The final pH values of the reaction solution were determined after addition of the enzyme and the substrates. The optimum temperature of activity was measured for 10 min under standard conditions except for temperature. The thermostability was determined by incubating the enzyme at each temperature for 30 min in 50 mm sodium phosphate buffer (pH 7.1) followed by measuring the activity under the standard conditions at 40 °C. Analysis of the Products—The reaction products from various substrates were separated by TLC on a silica gel 60 F254 plate (5.0 × 7.5 cm, Merck) with a solvent system of acetonitrile:water (4:1, v/v). Sugars were detected by baking after dipping the plate in 5% sulfuric acid in methanol. When necessary, the amounts of the products were quantified by using high performance ion exchange chromatography as described above. The amounts of reducing sugar liberated in the hydrolysis of polysaccharides by the enzyme were determined using the γ 3,5-dinitrosalicylic acid method (31Miller G.L. Anal. Chem. 1959; 31: 426-428Crossref Scopus (22353) Google Scholar) or the copper-bicinchoninic acid method (32Waffenschmidt S. Jaenicke L. Anal. Biochem. 1987; 165: 337-340Crossref PubMed Scopus (248) Google Scholar). Analysis of the Anomeric Form of the Products—The anomeric form of the hydrolytic product from X3 and X4 (50 mm) was determined by using an isocratic HPLC method (33Koga D. Yoshioka T. Arakane Y. Biosci. Biotechnol. Biochem. 1998; 62: 1643-1646Crossref PubMed Scopus (68) Google Scholar) described below. The enzymatic reaction was carried out in 25 mm sodium phosphate buffer (pH 7.1) at 25 °C with an enzyme concentration of 5.5 μm. After incubation for 1 and 25 min, an aliquot (10 μl) of the reaction solution was immediately loaded onto a TSK-GEL Amide-80 column (4.6 × 250 mm, Tosoh, Japan), and eluted with acetonitrile:water (7:3, v/v) at a flow rate of 1.5 ml/min at 25 °C, separating the xylooligosaccharides anomers. The initial substrate and products were detected using a refractive index monitor (RI Model 504, GL Science, Tokyo, Japan). The retention times of α- and β-xylose were confirmed by loading a solution of α-xylopyranose immediately after preparation, while the retention times of anomers of xylobiose were confirmed by loading the products of 1 and 25 min hydrolyzes of phenyl-β-xylobioside by Cex (34Honda Y. Kitaoka M. Sakka K. Ohmiya K. Hayashi K. J. Biosci. Bioeng. 2002; 93: 313-317Crossref PubMed Scopus (12) Google Scholar), a family 10 xylanase that forms β-xylobiose as its initial product. In addition, the retention times of xylooligosaccharides were evaluated by the proportion of the equilibrated anomers (α:β = ∼4:6) using an equilibrated solution of xylooligosaccharides. For all xylooligosaccharides, the α-anomers had shorter retention times than the corresponding β-anomers. Kinetic Analysis—To determine the apparent kinetic parameters, X3–X6 were subjected to hydrolysis in 50 mm sodium phosphate buffer (pH 7.1) at 40 °C. The initial rates were measured as the increase in xylose by using high performance ion exchange chromatography as described above. The kinetic parameters were calculated by regressing the experimental data (substrate concentration range; 0.2 × Km–3 × Km) with the Michaelis-Menten equation by the curve fit method using Kaleidagraph™ Version 3.51 (Synergy Software). Site-directed Mutagenesis—Site-directed mutagenesis for E70A, D128A, D263A, and D128A/D263A was performed using the PCR overlap extension method (35Higuchi R. Krummel B. Saiki R.K. Nucleic Acids Res. 1988; 16: 7351-7367Crossref PubMed Scopus (2102) Google Scholar). The following mutagenetic oligonucleotide primers were used (the mismatched bases are underlined): 5′-CTC GAT GTG CGG ACT GCA GGA ATG TCC TAC-3′ (E70A), 5′-GCC CCA GCT CCG GCC GGA GAG GAA TAT TTT-3′ (D128A), and 5′-CAC TTT TTT AGC GCT TCT TAT CGT GTG GCT-3′ (D263A). The mutated enzymes were prepared and purified as described above. Characterization of Recombinant BH2105—The recombinant BH2105 protein was expressed in E. coli BL21(GOLD) and purified to yield a 45-kDa protein on SDS-PAGE. The enzyme was purified 2.7-fold from the cell-free extract. The N-terminal sequence of the purified protein was Met-Glu-Lys-Thr-Thr-Glu-Gly-Ala-Phe-Tyr, corresponding to the deduced amino acid sequence from the starting codon. This sequence information suggested that the enzyme has no signal peptide. This enzyme did not hydrolyze birch wood xylan even at high concentrations (10 mg/ml) as shown by measuring the increase in reducing value, clearly indicating that it is not an endo-β-1,4-xylanase even though it was annotated as xylanase Y. Furthermore the enzyme did not hydrolyze any other polymeric substrates for GH8 enzymes (chitosan, lichenan, curdlan, and carboxymethylcellulose) determined by increases in reducing value. Various pentasaccharides (xylopentaose, cellopentaose, laminaripentaose, chitopentaose, and chitosanpentaose) were examined for the substrate, and the enzyme showed hydrolytic activity only on xylopentaose, producing initially X1 and X4 and finally X1 and X2. To obtain further information on the hydrolysis of xylooligosaccharides (Xn where n = 2–6), the products were analyzed by TLC. In the initial stage, the enzyme released X1 and X(n–1) from Xn, and the final products were X1 and X2 when n ≥ 3. It hydrolyzed X2 into X1 at a much slower rate than that of X3 (less than 0.01% rate). On the other hand, the enzyme exhibited no activity on X-pNP and X2-pNP even at extended incubation time, suggesting the possibility of hydrolysis at the reducing end. Oligosaccharides larger than the initial substrates in the enzymatic reaction were not detected, indicating that the enzyme had no transglycosidation activity. Enzymatic properties as a function of pH and temperature were determined with the X3 hydrolysis (Fig. 1). The optimum pH for activity was between 6.2 and 7.3, and the enzyme was completely stable between pH 5.0 and 9.8 at 30 °C for 30 min. The enzyme was stable at temperatures up to 40 °C, and the optimal temperature was 50 °C. Substrate Specificity: Recognition of Sugar Residue—To determine the position of hydrolysis and further substrate specificity of the enzyme, various derivatives of X3 were examined. When β-(1→4) d-glucose and d-xylose-based trisaccharides (G-X-X, X-X-G, G-X-G, X-G-G, G-G-X, X-G-X, and G3) were examined as the substrate, the enzyme hydrolyzed only G-X-X, X-X-G, and G-X-G at the linkage of the reducing end side, judging from the products, with a much slower rate than X3 (Table I). The reducing end specificity is completely different from that of β-xylosidase, which liberates xylose from the non-reducing end. The substitution of the middle xylose unit into glucose caused complete loss of activity, indicating that the sugar moiety located at the –1 subsite (Fig. 2) must strictly be xylose. Substitution at the reducing end and the non-reducing end caused drastic decreases in activity, indicating that both the +1 and –2 subsites (Fig. 2) strongly prefer xylose but were not as strict as the subsite –1. Judging from the specificity, the enzyme is very specific to homo-xylooligosaccharides. It hydrolyzed X3-de into X2 and 1,5-anhydroxylitol (X-de) at a rate that was 3.2% of the hydrolysis of X3 (Table I). This result again confirms that the enzyme hydrolyzes the reducing end glycosyl linkage. The result also suggests that the enzyme recognizes one of the anomeric hydroxyl groups at the reducing end. The small hydrolysis rates for the derivatives of xylotriose were due to their higher Km values evidenced by their linear S-ν relations in the range lower than 2.6 mm, whereas Km for X3 was 2.4 mm.Table IActivity of BH2105 toward various trisaccharidesSubstrateProductSpecific activitys-1X-X-X (X3)X-X + X84.0 (100%)X-X-X-de (X3-de)X-X + X-de2.7 (3.2%)G-X-XG-X + X0.94 (1.1%)X-X-GX-X + G0.42 (0.5%)G-X-GG-X + G0.003 (0.004%)X-G-G—a—, not detectable.G-G-X—X-G-X—G-G-G (G3)—a —, not detectable. Open table in a new tab Anomeric Hydroxyl Group Recognition by the Enzyme—The anomeric composition of the degradation products of X3 and X4 by the BH2105 protein were analyzed by HPLC. Fig. 3 shows the HPLC profiles of the hydrolytic products from X3 and X4 obtained with the enzymes. The standard equilibrium ratio of α:β anomers for X3 and X4 were ∼4:6 (Fig. 3). As shown in Fig. 3A, the enzyme produced β-anomer of X1 and α-anomer of X2 from X3 in the reaction for 1 min. Furthermore the α-anomer of X3 was the predominant anomer remaining in the reaction. Small amounts of α-X1 and β-X2 were also observed, which may have been due to the mutarotation of the initial products over the short term. This result strongly suggests that the enzyme hydrolyzed only the β-anomer of X3 at the linkage of the reducing end side with anomeric inversion to form α-X2 and β-X1. The preference of the β-anomer explains the reason why the enzyme hydrolyzed X3-de at a much slower rate. After 25 min of the reaction, the substrate (X3) disappeared, and the anomeric composition of the product (X and X2) reached equilibrium. This result suggests that the enzyme hydrolyzed α-X3 after the mutarotation that converted α-X3 into β-X3. In the case of X4 hydrolytic reaction after 1 min, β-X1 and α-X3 were produced with a decrease in β-X4 (Fig. 3B). Again small amounts of the opposite anomers were also observed that were probably due to mutarotation. It should be noted that X2, the hydrolytic product of X3, was hardly detected even though half of X4 had already been hydrolyzed in the reaction, suggesting that the enzyme did not act on the α-anomer. The α-X4 remaining and the α-X3 produced must be converted into their β-anomers before the hydrolysis. After 25 min of reaction, the products were equilibrated X1 and X2. The hydrolytic mechanism is schematically summarized in Fig. 4. Kinetic Property—The S-ν curve of X3–X6 hydrolysis by the enzyme indicates a typical Michaelis-Menten type relationship. The kinetic parameters are summarized in Table II. The Km value increased with the increase in degree of polymerization, while kcat/Km decreased with the increase in Km. These results suggest that X3 is the most suitable substrate for the enzyme and that the role of the enzyme is to hydrolyze smaller xylooligosaccharides. Subsites downstream of subsite –2 (Fig. 2) may be postulated, but such subsites must have negative binding energies.Table IIKinetic parameters of xylooligosaccharides hydrolysis catalyzed by BH2105Substratek catKmkcat/Kms-1mms-1 mm-1X3163 ± 52.4 ± 0.266.9 ± 3.5X4162 ± 65.0 ± 0.432.3 ± 1.4X573 ± 24.4 ± 0.316.7 ± 0.5X6175 ± 1918.5 ± 1.318.5 ± 1.3 Open table in a new tab Mutational Analysis—The BH2105 amino acid sequence analysis indicated a 32.6% identity with P. haloplanktis GH8 xylanase (GenBank™ accession number AJ427291). As found in the sequence alignment between these enzymes, the catalytic residues (Glu-70, Asp-128, and Asp-263; BH2105 numbering) proposed for the xylanase were strongly conserved in the BH2105 amino acid sequence (19Van Petegem F. Collins T. Meuwis M.A. Gerday C. Feller G. Van Beeumen J. J. Biol. Chem. 2003; 278: 7531-7539Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar). Therefore, we examined the enzymatic activity of alanine mutants (E70A, D123A, and D263A) and found that, as expected, the hydrolytic activity of E70A and D263A was 10–4 orders less than that of wild type (Table III). On the other hand, D128A retained slightly higher activity than that of E70A and D263A, although it was still approximately three hundredths of wild type (Table III). The residues were predicted to act as an acid catalyst (Glu-70) and a base catalyst (Asp-263) and to stabilize the 2,5B conformation of the sugar moiety bound at subsite –1 (Asp-123) (6Guerin D.M. Lascombe M.B. Costabel M. Souchon H. Lamzin V. Beguin P. Alzari P.M. J. Mol. Biol. 2002; 316: 1061-1069Crossref PubMed Scopus (122) Google Scholar, 19Van Petegem F. Collins T. Meuwis M.A. Gerday C. Feller G. Van Beeumen J. J. Biol. Chem. 2003; 278: 7531-7539Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar). These results suggest that these amino acid residues are conserved and are essential for the catalytic reaction of the enzyme.Table IIISpecific activity of BH2105 mutantsEnzymeSpecific activityRelative activity (mutant/wild type)s-1Wild type84.01.0E70A0.00881.1 × 10-4D128A0.293.5 × 10-3D263A0.0192.3 × 10-4" @default.
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• W2025968524 title "A Family 8 Glycoside Hydrolase from Bacillus halodurans C-125 (BH2105) Is a Reducing End Xylose-releasing Exo-oligoxylanase" @default.
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