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. 2009 Sep;151(1):47–58. doi: 10.1104/pp.109.139436

Structural and Enzymatic Characterization of Os3BGlu6, a Rice β-Glucosidase Hydrolyzing Hydrophobic Glycosides and (1→3)- and (1→2)-Linked Disaccharides1,[C],[W],[OA]

Supriya Seshadri 1, Takashi Akiyama 1, Rodjana Opassiri 1, Buabarn Kuaprasert 1, James Ketudat Cairns 1,*
PMCID: PMC2735989  PMID: 19587102

Abstract

Glycoside hydrolase family 1 (GH1) β-glucosidases play roles in many processes in plants, such as chemical defense, alkaloid metabolism, hydrolysis of cell wall-derived oligosaccharides, phytohormone regulation, and lignification. However, the functions of most of the 34 GH1 gene products in rice (Oryza sativa) are unknown. Os3BGlu6, a rice β-glucosidase representing a previously uncharacterized phylogenetic cluster of GH1, was produced in recombinant Escherichia coli. Os3BGlu6 hydrolyzed p-nitrophenyl (pNP)-β-d-fucoside (kcat/Km = 67 mm−1 s−1), pNP-β-d-glucoside (kcat/Km = 6.2 mm−1 s−1), and pNP-β-d-galactoside (kcat/Km = 1.6 mm−1s−1) efficiently but had little activity toward other pNP glycosides. It also had high activity toward n-octyl-β-d-glucoside and β-(1→3)- and β-(1→2)-linked disaccharides and was able to hydrolyze apigenin β-glucoside and several other natural glycosides. Crystal structures of Os3BGlu6 and its complexes with a covalent intermediate, 2-deoxy-2-fluoroglucoside, and a nonhydrolyzable substrate analog, n-octyl-β-d-thioglucopyranoside, were solved at 1.83, 1.81, and 1.80 Å resolution, respectively. The position of the covalently trapped 2-F-glucosyl residue in the enzyme was similar to that in a 2-F-glucosyl intermediate complex of Os3BGlu7 (rice BGlu1). The side chain of methionine-251 in the mouth of the active site appeared to block the binding of extended β-(1→4)-linked oligosaccharides and interact with the hydrophobic aglycone of n-octyl-β-d-thioglucopyranoside. This correlates with the preference of Os3BGlu6 for short oligosaccharides and hydrophobic glycosides.

β-Glucosidases (EC 3.2.1.21) have a wide range of functions in plants, including acting in cell wall remodeling, lignification, chemical defense, plant-microbe interactions, phytohormone activation, activation of metabolic intermediates, and release of volatiles from their glycosides (Esen, 1993). They fulfill these roles by hydrolyzing the glycosidic bond at the nonreducing terminal glucosyl residue of a glycoside or an oligosaccharide, thereby releasing Glc and an aglycone or a shortened carbohydrate. The aglycone released from the glycoside may be a monolignol, a toxic compound, or a compound that further reacts to release a toxic component, an active phytohormone, a reactive metabolic intermediate, or a volatile scent compound (Brzobohatý et al., 1993; Dharmawardhama et al., 1995; Reuveni et al., 1999; Lee et al., 2006; Barleben et al., 2007; Morant et al., 2008). Indeed, the wide range of glucosides of undocumented functions found in plants suggests that many β-glucosidase functions may remain to be discovered.

Plant β-glucosidases fall into related families that have been classified as glycosyl hydrolase (GH) families GH1, GH3, and GH5 (Henrissat, 1991; Coutinho and Henrissat, 1998, 1999). Of these, GH1 has been most thoroughly documented and shown to comprise a gene family encoding 40 putative functional GHs in Arabidopsis (Arabidopsis thaliana) and 34 in rice (Oryza sativa) in addition to a few pseudogenes (Xu et al., 2004; Opassiri et al., 2006). In addition to β-glucosidases, plant GH1 members include β-mannosidases (Mo and Bewley, 2002), β-thioglucosidases (Burmeister et al., 1997), and disaccharidases such as primeverosidase (Mizutani et al., 2002) as well as hydroxyisourate hydrolase, which hydrolyzes the internal bond in a purine ring rather than a glycosidic bond (Raychaudhuri and Tipton, 2002). The specificity for the glycone in GH1 enzymes varies. Some enzymes are quite specific for β-d-glucosides or β-d-mannosides, while many accept either β-d-glucosides or β-d-fucosides, and some also hydrolyze β-d-galactosides, β-d-xylosides, and α-l-arabinoside (Esen, 1993). However, most GH1 enzymes are thought to hydrolyze glucosides in the plant, and it is the aglycone specificity that determines the functions of most GH1 enzymes.

Aglycone specificity of GH1 β-glucosidases ranges from rather broad to absolutely specific for one substrate and is not obvious from sequence similarity. For instance, maize (Zea mays) ZmGlu1 β-glucosidase hydrolyzes a range of glycosides, including its natural substrate, 2-O-β-d-glucopyranosyl-4-dihydroxy-1,4-benzoxazin-3-one (DIMBOAGlc), but not dhurrin, whereas sorghum (Sorghum bicolor) Dhr1, which is 72% identical to ZmGlu1, only hydrolyzes its natural cyanogenic substrate dhurrin (Verdoucq et al., 2003). Similarly, despite sharing over 80% amino acid sequence identity, the legume isoflavonoid β-glucosidases dalcochinase from Dalbergia cochinchinensis and Dnbglu2 from Dalbergia nigrescens hydrolyze each other's natural substrate very poorly (Chuankhayan et al., 2007). Thus, small differences in the amino acid sequence surrounding the active site may be expected to account for significant differences in substrate specificity.

GH1 is classified in GH clan A, which consists of GH families whose members have a (β/α)8-barrel structure with the catalytic acid/base on strand 4 of the β-barrel and the catalytic nucleophile on strand 7 (Henrissat et al., 1995; Jenkins et al., 1995). As such, all GH1 enzymes have similar overall structures, but it has been noted that four variable loops at the C-terminal end of the β-barrel strands, designated A, B, C, and D, account for much of the differences in the active site architecture (Sanz-Aparicio et al., 1998). The similar structures with great diversity in substrate specificity make plant GH1 enzymes an ideal model system to investigate the structural basis of substrate specificity. To date, seven plant β-glucosidase structures have been reported, including three closely related chloroplastic enzymes from maize (Czjzek et al., 2000, 2001), sorghum (Verdoucq et al., 2004), and wheat (Triticum aestivum; Sue et al., 2006), the cytoplasmic strictosidine β-glucosidase from Rauvolfia serpentine (Barleben et al., 2007), and the secreted enzymes white clover (Trifolium repens) cyanogenic β-glucosidase (Barrett et al., 1995), white mustard (Sinapsis alba) myrosinase (thioglucosidase; Burmeister et al., 1997), and rice Os3BGlu7 (BGlu1; Chuenchor et al., 2008). These enzymes hydrolyze substrates with a range of structures, but they cannot account for the full range of β-glucosidase substrates available in plants, and determining the structural differences that bring about substrate specificity differences in even closely related GH1 enzymes has proven tricky (Verdoucq et al., 2003, 2004; Sue et al., 2006; Chuenchor et al., 2008).

Amino acid sequence-based phylogenetic analysis of GH1 enzymes encoded by the rice genome showed that there are eight clusters containing both rice and Arabidopsis proteins that are more closely related to each other than they are to enzymes from the same plants outside the clusters (Fig. 1; Opassiri et al., 2006). In addition, there are a cluster of sixteen putative β-glucosidases and a cluster of myrosinases in Arabidopsis without any closely related rice counterparts. Comparison with characterized GH1 enzymes from other plants reveals other clusters of related enzymes not found in rice or Arabidopsis, including the chloroplastic enzymes, from which the maize, sorghum, and wheat structures are derived, and the cytoplasmic metabolic enzymes, from with the strictosidine hydrolase structure is derived (Fig. 1). Therefore, although the known structures provide good tools for molecular modeling of plant enzymes, most rice and Arabidopsis GH1 enzymes lack a close correspondence in sequence and functional evolution to these structures, suggesting that the variable loops that determine the active site may be different. It would be useful, therefore, to know the structures and substrate specificities of representative members of each of the eight clusters seen in rice and Arabidopsis. To begin to acquire this information, we have expressed Os3BGlu6, a member of cluster At/Os 1 in Figure 1, characterized its substrate specificity, and determined its structure alone and in complex with a glycosyl intermediate and a nonhydrolyzable substrate analog.

Figure 1.

Figure 1.

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Simplified phylogenetic tree of the amino acid sequences of eukaryotic GH1 proteins with known structures and those of rice and Arabidopsis GH1 gene products. The protein sequences of the eukaryotic proteins with known structures are marked with four-character PDB codes for one of their structures, including Trifolium repens cyanogenic β-glucosidase (1CBG; Barrett et al., 1995), Sinapsis alba myrosinase (1MYR; Burmeister et al., 1997), Zea mays ZmGlu1 β-glucosidase (1E1F; Czjzek et al., 2000), Sorghum bicolor Dhr1 dhurrinase (1V02; Verdoucq et al., 2004), Triticum aestivum β-glucosidase (2DGA; Sue et al., 2006), Rauvolfia serpentina strictosidine β-glucosidase (2JF6; Barleben et al., 2007), and Oryza sativa Os3BGlu7 (BGlu1) β-glucosidase (2RGL; Chuenchor et al., 2008) from plants, along with Brevicoryne brassicae myrosinase (1WCG; Husebye et al., 2005), Homo sapiens cytoplasmic (Klotho) β-glucosidase (2E9M; Hayashi et al., 2007), and Phanerochaete chrysosporium (2E3Z; Nijikken et al., 2007), while those encoded in the Arabidopsis and rice genomes are labeled with the systematic names given by Xu et al. (2004) and Opassiri et al. (2006), respectively. One or two example proteins from each plant are given for each of the eight clusters of genes shared by Arabidopsis (At) and rice (Os) and the Arabidopsis-specific clusters At I (β-glucosidases) and At II (myrosinases), with the number of Arabidopsis or rice enzymes in each cluster given in parentheses. These sequences were aligned with all of the Arabidopsis and rice sequences in ClustalX (Thompson et al., 1997), the alignment was manually edited, all but representative sequences were removed, and the tree was calculated by the neighbor-joining method, bootstrapped with 1,000 trials, and then drawn with TreeView (Page, 1996). The grass plastid β-glucosidases, which are not represented in Arabidopsis and rice, are shown in the group marked “Plastid.” Percentage bootstrap reproducibility values are shown on internal branches where they are greater than 60%. Except those marked by asterisks, all external branches represent groups with 100% bootstrap reproducibility. To avoid excess complexity, those groups of sequences marked with asterisks are not monophyletic and represent more branches within the designated cluster than are shown. For a complete phylogenetic analysis of Arabidopsis and rice GH1 proteins, see Opassiri et al. (2006).

RESULTS

Cloning, Expression, and Purification of Os3BGlu6

The cDNA encoding the Os3BGlu6 protein was isolated from a cold-treated rice library, and the sequence was determined (GenBank accession no. AY129294). This full-length cDNA contained an open reading frame of 1,563 nucleotides encoding 521 amino acids, which were predicted to include an N-terminal secretory signal peptide of 38 residues by SignalP (Bendtsen et al., 2004). The Os3BGlu6 protein was expressed in Escherichia coli as a fusion protein with N-terminal thioredoxin and His6 tags, from which it was released by cleavage of a tobacco etch virus (TEV) protease site that was placed four amino acid residues before precursor residue 38. This allowed a soluble and active Os3BGlu6 β-glucosidase of 55 kD to be conveniently purified to more than 95% purity, based on SDS-PAGE (Supplemental Fig. S1).

The activity of the expressed Os3BGlu6 was highest at pH 4 to 5, dropped by 50% at pH 2.5 and 6.0, and was negligible from pH 7.0 upward. The enzyme had highest activity over the temperature range of 40°C to 55°C, was most stable at 20°C to 30°C, and began to lose activity after 40 min at 40°C or higher. Therefore, standard assays were conducted at 30°C, which is physiologically relevant for rice.

Substrate Specificity of Os3BGlu6

The ability of OsBGlu6 to hydrolyze p-nitrophenyl (pNP) glycosides was tested to assess its glycone specificity (Table I). Among the pNP glycosides, Os3BGlu6 preferred pNP-β-d-fucopyranoside (pNPFuc), with a kcat/Km of 67 mm−1 s−1, due to its relatively low Km of 0.50 mm, followed by pNP-β-d-glucoside (pNPGlc) and pNP-β-d-galactoside (pNPGal), with kcat/Km values of 6.2 and 1.6 mm−1 s−1, respectively. Hydrolysis of α-d-glucoside was negligible, but α-l-arabinoside was hydrolyzed at a low rate.

Table I.

Kinetic parameters of Os3BGlu6 for the hydrolysis of pNP glycosides, oligosaccharides, and alkyl and natural glucosides

nd, Not determined.

Substrate nmol pNP min−1 mg−1 Protein Percent Relative Activity Km kcat kcat/Km
mm s−1 mm−1s−1
pNP-β-d-glucoside 5,844 ± 1.5 100 6.3 ± 0.4 38.9 ± 0.9 6.2
pNP-β-d-fucoside 24,470 ± 1.5 415 ± 0.03 0.50 ± 0.02 33.5 ± 0.7 67
pNP-β-d-galactoside 1,719 ± 1.4 29.4 ± 0.02 6.06 ± 0.5 9.8 ± 0.2 1.6
pNP-α-l-arabinoside 51.9 ± 0.3 0.88 ± 0.005 nd nd nd
pNP-β-d-cellobioside 27.67 ± 0.15 0.46 ± 0.003 nd nd nd
pNP-β-d-xyloside 6.3 ± 0.5 0.10 ± 0.008 nd nd nd
pNP-β-d-mannoside 3.8 ± 0.5 0.06 ± 0.008 nd nd nd
pNP-β-d-glucuronide 2.7 ± 0.3 0.04 ± 0.005 nd nd nd
pNP-α-d-glucoside 0.43 ± 0.008 0.007 ± 0.001 nd nd nd
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Hydrolysis of synthetic and natural glycosides and oligosaccharides was tested to assess its aglycone specificity (Table II). Os3BGlu6 hydrolyzed n-octyl-β-d-glucoside with the highest catalytic efficiency (kcat/Km = 2.7 mm−1 s−1), which was approximately 3-fold higher than that for n-heptyl-β-d-glucoside (0.85 mm−1 s−1). Among the disaccharides, Os3BGlu6 hydrolyzed the β-(1→3)-linked laminaribiose best, followed by β-(1→2)-linked sophorose, β-(1→6)-gentiobiose, and β-(1→4)-linked cellobiose, in order of decreasing catalytic efficiency (kcat/Km). Longer β-(1→3)- and β-(1→4)-linked oligosaccharides were hydrolyzed at decreasing rates. Os3BGlu6 also hydrolyzed natural glycosides, including apigenin-7-glucoside, glycitin, diadzin, genistin, esculin, arbutin, coumaryl alcohol β-d-glucoside, coniferin, and salicin, although at low rates. The cyanogenic glycosides, amygdalin, linamarin, and prunacin, and several other glycosides (listed at the bottom of Table II) were not detectably hydrolyzed after reaction overnight.

Table II.

Hydrolysis of glycosides and oligosaccharides by Os3BGlu6

nd, Not determined.

Substrate nmol Glc min−1 mg−1 Proteina Percent Relative Activitya Km kcat kcat/Km
mm s−1 mm−1 s−1
n-Octyl-β-d-glucoside 2,282 ± 0.9 100 ± 0.04 4.5 ± 0.3 12.4 ± 0.4 2.7
n-Heptyl-β-d-glucoside 673.8 ± 0.9 29.5 ± 0.04 5.0 ± 0.3 4.25 ± 0.10 0.85
Glucooligosaccharides
    Laminaribiose [β-(1→3)-linked] 1,272 ± 1.3 55.7 ± 0.05 3.6 ± 0.3 6.17 ± 0.19 1.7
    Laminaritriose 169.6 ± 0.9 7.43 ± 0.04 8.7 ± 0.7 1.64 ± 0.06 0.18
    Laminaritetraose 2.0 ± 0.01 0.08 ± 0.0004 nd nd nd
    Laminaripentaose 1.4 ± 0.01 0.06 ± 0.001 nd nd nd
    Cellobiose [β-(1→4)-linked] 15.0 ± 0.5 0.65 ± 0.001 15.3 ± 1.2 0.13 ± 0.001 0.0085
    Cellotriose 13.1 ± 0.6 0.57 ± 0.03 nd nd nd
    Cellotetraose 12.9 ± 0.5 0.56 ± 0.02 nd nd nd
    Cellopentaose 8.0 ± 0.3 0.35 ± 0.01 nd nd nd
    Cellohexaose 6.38 ± 0.3 0.28 ± 0.01 nd nd nd
    Sophorose [β-(1→2)-linked] 728.7 ± 1.3 31.9 ± 0.05 9.8 ± 0.8 9.8 ± 0.4 0.96
    Gentiobiose [β-(1→6)-linked] 5.25 ± 0.17 0.23 ± 0.007 14.9 ± 1.1 0.17 ± 0.003 0.011
Natural β-d-glucosidesb
    Apigenin-7-glucoside 15.3 ± 0.2 0.66 ± 0.009 nd nd nd
    Glycitin 4.4 ± 0.07 0.19 ± 0.003 nd nd nd
    Diadzin 4.0 ± 0.04 0.17 ± 0.001 nd nd nd
    Genistin 2.9 ± 0.02 0.13 ± 0.001 nd nd nd
    Esculin 2.4 ± 0.01 0.10 ± 0.005 nd nd nd
    Arbutin 0.59 ± 0.05 0.02 ± 0.002 nd nd nd
    Coumaryl alcohol β-d-glucoside 0.54 ± 0.01 0.02 ± 0.0005 nd nd nd
    Coniferin 0.46 ± 0.007 0.02 ± 0.0003 nd nd nd
    Salicin 0.18 ± 0.001 (79 ± 0.3) × 10−4 nd nd nd
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a

Activities were determined by the Glc oxidase assay, and percent relative rates are set relative to hydrolysis of n-octyl-β-d-glucoside as 100%. Amounts released from disaccharides were divided by two, since two Glc molecules are released from one reaction.

b

The natural glycosides for which relative activities were determined are those for which hydrolysis was detected by TLC. Amygdalin, dalcochinin glucoside, gossypin, linamarin, naringin, phlorizin, prunasin, pyridoxine-5′-O-β-d-glucoside, and quercitin 3-β-d-glucoside were not hydrolyzed, as observed by TLC.

Overall Structure of Os3BGlu6

To understand the structural basis for the substrate specificity of Os3BGlu6, its three-dimensional structures alone and in complexes with 2-deoxy-2-fluoroglucoside (a slowly hydrolyzed covalent intermediate) and n-octyl-β-d-thioglucopyranoside (a nonhydrolyzable substrate analog) were determined by protein crystallography. The data processing and refinement parameters are provided in Table III. The overall structure is a (β/α)8-barrel, similar to those of other known structures of GH1 enzymes, with the catalytic acid/base, Glu-178, and nucleophile, Glu-394, located on β-strands 4 and 7 of the β-barrel (Fig. 2A). The active site forms a slot-like cleft in the side of the protein, the bottom of which is negatively charged (Fig. 2B). A single protein molecule was present in the asymmetric unit, and a lack of close crystal contacts indicated that Os3BGlu6 is monomeric.

Table III.

Data collection, processing, and refinement parameters

Data Set Native Os3BGlu6 Os3BGlu6/n-Octyl-β-d-Thioglucopyranoside Complex Os3BGlu6/G2F Complex
PDB code 3GNO 3GNP 3GNR
Beamline BL13B1 BL13B1 BL13B1
Wavelength (Å) 1.00 1.00 1.00
Space group P212121 P212121 P212121
Unit-cell parameters (Å) a = 56.8 a = 57.2 a = 56.7
b = 90.5 b = 91.1 b = 90.6
c = 101.7 c = 111.4 c = 101.9
Resolution range (Å) 30.0–1.83 30.0–1.80 30.0–1.81
Resolution outer shell (Å) 1.90–1.83 1.86–1.80 1.87–1.81
Completeness (%) 98.9 (96.8)a 91.1 (98.0)a 92.9 (95.1)a
Average redundancy per shell 7.0 (6.9) 6.9 (6.9) 3.7 (4.0)
I/σ(I) 18.9 (4.9)a 30.7 (6.4)a 23.9 (8.7)a
Rmerge (%)b 9.5 (36.6)a 5.2 (29.3)a 4.5 (18.4)a
Rfactor (%)c 16.3 17.8 16.2
Rfree (%)c 20.4 20.7 18.5
No. of residues in protein 1–488 1–488 1–488
No. of waters 466 340 452
Mean B factor
    Protein 12.7 18.5 16.0
    Ligand 28.8 41.2 14.8
    Water 26.2 27.1 27.0
    r.m.s. bond deviations (Å) 0.013 0.015 0.012
    r.m.s angle deviations (°) 1.32 1.44 1.28
Ramachandran plot
    Residues in most favorable regions (%) 89.6 89.3 89.8
    Residues in additional allowed regions (%) 10 10 9.5
    Residues in generously allowed regions (%) 0.5 0.7 0.7
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a

Numbers in parentheses designate to the highest resolution shell.

b

Rmerge is the linear residual for all observations of unique reflections.

c

Rfactor = Σ||Fo| − |Fc||/Σ|Fo|, where |Fo| and |Fc| are the observed and calculated structure factor amplitudes. Rfree is the R calculated from the 5% of the data that were not used in refinement.

Figure 2.

Figure 2.

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Overall and active site structures of Os3BGlu6. A, Overall ribbon diagram of native Os3BGlu6. The catalytic residues Glu-394 and Glu-178 and one molecule of Tris in two conformations are shown in ball-and-stick representation. The α-helices are colored purple, β-strands are green, and loops are cyan. B, Surface view of the Os3BGlu6 structure showing the active site cleft with the ligand n-octyl-β-d-thioglucopyranoside. The surface is colored by electrostatic potential, with positively charged, negatively charged, and neutral regions colored blue, red, and white respectively. C, Close-up of the electron density of Tris in the active site in stereo view. The Fo-Fc omit electron density map for the Tris is shown as a mesh contoured at I/σ = 3. The side chains of the surrounding amino acids are represented by sticks, and the Tris ligand is represented by balls and sticks. In all frames, oxygen atoms are shown in red, nitrogen in blue, sulfur in dark yellow, protein carbons in yellow, Tris carbons in green or pink, and n-octyl-β-d-thioglucopyranoside carbons in green.

N-terminal sequencing of the first eight amino acid residues of the protein recovered from crystals gave a sequence consistent with that predicted from the TEV cleavage site, with four residues from the cloning vector and primers, Ser-1, Phe-2, Thr-3, and Met-4, followed by the first four residues included from the protein sequence, Ala-5, Gln-6, Gln-7, and Ser-8. However, these residues, together with Gly-9 and Gly-10, had no detectable electron density, likely due to high flexibility. The C terminus was well defined, except that the side chains of the last two residues, Lys-487 and Thr-488, had poor density. In addition, poor density was observed for the side chains of residues 334 to 337 in loop C in all three structures, although the Cα backbone was clearly visible. This loop is apparently flexible, since it is exposed at the outer surface of the protein, and the position of this loop was slightly different in the case of the Os3BGlu6/n-octyl-β-d-thioglucopyranoside. No density was observed for six of the atoms of the surface residues in Lys-114, Glu-296, Glu-383, and Lys-432 in the native and Os3BGlu6/G2F structures, so these atoms were deleted from the structures. Due to the high resolution of the model, multiple conformations were observed for the flexible side chains of residues Arg-233, Thr-234, Asn-332, and Glu-451 in native Os3BGlu6 and Asn-241, Ser-231, Arg-293, Asp-361, and Glu-433 in the Os3BGlu6/G2F complex. One disulfide bond was seen linking Cys-197 and Cys-205 in loop B, a feature that is conserved in plant GH1 enzymes. Two cis-peptide bonds were found between Ala-193 and Pro-194 and between Trp-444 and Ser-445, which is also common in GH1.

The active site pocket of the native protein structure had extra electron density, which could not be fit to glycerol or a component of the precipitant but was best fit by a molecule of Tris in two possible conformations (Fig. 2C) with an average B factor of 28.81 Å2. Tris was a component of the protein storage buffer and was confirmed to bind by its competitive inhibition of Os3BGlu6 with a Ki of 5.1 ± 0.2 mm. Six glycerol molecules from the cryoprotectant were also seen on the surface of the protein in the native structure. Four of these were also seen in the Os3BGlu6/n-octyl-β-d-thioglucopyranoside complex and two in the Os3BGlu6/G2F complex.

Among known GH1 structures, Os3BGlu6 was most similar to the plant secreted β-glucosidases white clover cyanogenic β-glucosidase (1CBG; Barrett et al., 1995) and rice Os3BGlu7 β-glucosidase (2RGL; Chuenchor et al., 2008), with the root mean squared deviation (RMSD) of their Cα to those of Os3BGlu6 being 0.81 Å for both structures over 467 and 444 Cα atoms, respectively. For other plant enzymes, the RMSD ranged from 0.83 Å over 445 Cα atoms for white mustard myrosinase (1E4M; Burmeister et al., 2000) to 0.96 Å over 451 atoms for maize ZmGlu1 β-glucosidase (1E1E; Czjzek et al., 2001), which was the same as its RMSD over 423 atoms with human cytosolic β-glucosidase (2E9M; Hayashi et al., 2007). Not surprisingly, the bacterial structures have higher RMSD with Os3BGlu6, such as 1.13 Å over 397 Cα atoms for Bacillus polymyxa β-glucosidase (1BGA; Sanz-Aparicio et al., 1998) and 1.31 Å over 354 atoms for Lactococcus lactis 6-phospho-β-galactosidase (1PBG; Wiesmann et al., 1995). Although all of the above mentioned structures are similar, many differences reside in the loop regions around their active sites, where the glycone- and aglycone-binding sites are located.

Three-Dimensional Structures of Os3BGlu6 Complexes with G2F and n-Octyl-β-d-Thioglucopyranoside

To investigate the structural basis for substrate binding and hydrolysis, the structures of Os3BGlu6 in a covalent intermediate with G2F and noncovalently bound to n-octyl-β-d-thioglucopyranoside were solved at 1.81 and 1.80 Å resolution, respectively. The crystals of the covalent G2F intermediate were isomorphous with the native structure, but the crystal unit cell expanded with n-octyl-β-d-thioglucopyranoside, as seen previously in crystals with n-octyl-β-d-glucopyranoside (McPherson et al., 1986). An increase of 9.5 Å was seen in the c side of the unit cell, due to a displacement of the protein molecules related by symmetry along the z axis by 4.5 to 5 Å in their water-mediated interactions, but the only significant change in the monomer structure was the conformation of loop C, which is not involved in these interactions.

In the intermediate complex, the density for the 2-deoxy-2-fluoroglucosyl residue in a relaxed 4C1 chair conformation covalently bound to the catalytic nucleophile was clearly evident in the glycone-binding subsite, surrounded by the universally conserved plant GH1 residues Gln-31, His-132, Asn-177, Glu-178, Glu-394, Glu-451, and Trp-452 (Fig. 3, A and B). The residues Glu-451, Gln-31, Tyr-321, and Trp-452 formed direct hydrogen bonds with G2F. The Oε1 and Oε2 of Glu-451 were linked to O4 and O6 of the Glc ring at 2.60 and 2.65 Å, which held Glu-451 in a single conformation, compared with the two conformations of Glu-451 seen in the native structure. The Gln-31 Oε1 and Nε2 were hydrogen bonded to the glucosyl O3 and O4 at 2.51 and 2.92 Å, respectively. In addition, the hydroxyl group of Tyr-321 was hydrogen bonded to O5 (2.93 Å) and that of Trp-452 Nε1 was hydrogen bonded to O3 (2.84 Å). A hydrophobic stacking interaction between the sugar and the indole ring of Trp-444 was observed, as seen in other GH1 glycosidases. Superimposition of the G2F complex against the native enzyme (Fig. 3A) showed that the hydroxyl group of Tyr-321, which was 2.75 Å from Oε1 of the catalytic nucleophile Glu-394 in the native structure, twisted away in the G2F complex to form a hydrogen bond to O5 of the sugar (2.93 Å). The nucleophile side chain also moved in producing the covalent intermediate, so that Glu-394 Oε1 and Oε2, which were positioned at 4.59 and 3.88 Å from Oε2 of Glu-178 in the native structure, were distanced at 3.99 and 3.25 Å from it in the G2F complex. In addition, the indole ring of Trp-452 in the G2F complex was shifted slightly inward compared with the native structure to allow its nitrogen to hydrogen bond with O3 of the sugar ring at 2.84 Å.

Figure 3.

Figure 3.

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Binding of 2-fluoroglucoside and n-octyl-β-d-thioglucopyranoside in the Os3BGlu6 active site. A, Stereo view of the superimposition of active site residues at the −1 subsite of the native Os3BGlu6 and Os3BGlu6/G2F complex structures. The residues surrounding the −1 subsite are represented by sticks colored with carbons in blue for native Os3BGlu6 and in yellow for Os3BGlu6/G2F. The G2F moiety bound to the catalytic nucleophile, Glu-394, is represented by balls and sticks with carbon in pink. B, Active site of the Os3BGlu6/G2F complex showing the electron density omit map of G2F and protein-ligand binding interactions. Hydrogen-bonding interactions between G2F and amino acid residues are shown as black dashed lines, while the Fo-Fc omit map of G2F contoured at I/σ = 3.0 is shown as blue mesh. C, Glycone and aglycone interactions of n-octyl-β-d-thioglucopyranoside with Os3BGlu6 residues. Representation of the protein and ligand are as in B, with the Fo-Fc omit map for the ligand contoured at I/σ = 2.5. D, Two views of the superimposition of the G2F and n-octyl-β-d-thioglucopyranoside ligands, showing that the Glc ring is in nearly the same position but is shaped as a relaxed 4C1 chair in G2F (green carbons) and as a 1S3 skew boat in n-octyl-β-d-thioglucopyranoside (pink carbons). In all frames, oxygen is in red, nitrogen in blue, fluorine in cyan, and sulfur in yellow.

In the complex of Os3BGlu6 with the nonhydrolyzable substrate analog n-octyl-β-d-thioglucopyranoside, the inhibitor was located deep in the active site slot (Figs. 2B, 3C, and 4A). The density for the sugar was well defined (Fig. 3C). For the aglycone moiety, the density was strongest for C1′, C4′, and C8′ and was lower for C2′, C3′, C5′, C6′, and C7′, likely due to the flexibility of the carbon chain, which appeared to be able to bind in multiple positions. Although both the relaxed 4C1 chair and less energetically favorable 1S3 skew boat conformations fit the density well, the distance of Oε1 of the acid/base catalyst Glu-178 to the sulfur in the 4C1 chair was 2.13 Å, which was unacceptably close. In contrast, this distance was 3.07 Å in the 1S3 conformation, putting the sulfur within distance to accept a proton from the catalytic acid. The glycone residue was in a similar position to that of G2F in its complex and formed similar hydrogen-bonding interactions, except that interactions of the O2 hydroxyl with His-132, Asn-177, and Glu-394 were also evident (Fig. 3, B and C). The aglycone was flanked by hydrophobic residues and a few polar residues. The n-octyl chain was bound by hydrophobic contacts from Tyr-321 and Trp-366 on one side and from Val-250 and Met-251, which also form the entrance to the active site, on the other side of the chain. The polar residues Thr-322, Thr-181, Asp-249 His-320, Asn-319, and Gln-273 also lay on either side of the aglycone.

Figure 4.

Figure 4.

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Comparison of the Os3BGlu6/n-octyl-β-d-thioglucopyranoside complex active site surface with those of maize ZmGlu1 and rice Os3BGlu7 substrate complexes. A, Active site the Os3BGlu6/n-octyl-β-d-thioglucopyranoside complex structure. The extension of Met-251 to restrict the active site can be seen at lower left. B, The aglycone-binding pocket of the inactive ZmGlu1 E191D mutant/DIMBOAGlc (PDB code 1E56) complex structure. The glucosyl moiety is hidden behind the aglycone. C, Active site of Os3BGlu7 with a docked cellotriose ligand. The smaller Asn-245 (in place of Met-251 in Os3BGlu6) provides a more open cleft for extended oligosaccharide binding. In all frames, active site amino acid residues are shown as sticks behind the gray transparent surface. The ligands are shown in ball-and-stick representations. In the online version, protein carbons are shown in green, n-octyl-β-d-thioglucopyranoside carbons in pink, DIMBOAGlc carbons in blue, cellotriose carbons in yellow, oxygen in red, nitrogen in dark blue, and sulfur in yellow. [See online article for color version of this figure.]

DISCUSSION

Os3BGlu6 Substrate Glycone Specificity

Os3BGlu6 showed a strong preference for β-d-fucoside, followed by β-d-glucoside, and then by β-d-galactoside, as was seen for several other plant β-d-glucosidases (Babcock and Esen, 1994; Srisomsap et al., 1996; Opassiri et al., 2004, 2006). Like maize β-glucosidase (Babcock and Esen, 1994), Os3BGlu6 shows a 10-fold higher kcat/Km for pNPFuc than for pNPGlc. In maize and rice Os3BGlu7 β-glucosidases, the preference for pNPFuc over pNPGlc is due to a higher kcat for pNPFuc (Opassiri et al., 2004), whereas for Os3BGlu6, the difference is due to a 10-fold lower Km for pNPFuc than pNPGlc, similar to D. cochinchinensis β-glucosidase, which had Km values of 0.54 mm for pNPFuc and 5.4 mm for pNPGlc (Srisomsap et al., 1996). In Os3BGlu6, this appears to be partly due to a higher tolerance for the equatorial position of the 4′ hydroxyl group in Os3BGlu6, since it had a Km for pNPGal similar to that for pNPGlc, whereas Os3BGlu7 had a Km over 10-fold higher for pNPGal than pNPGlc. In contrast, Os3BGlu6 had a much lower efficiency (kcat/Km of 0.06 mm−1 s−1) for hydrolysis of pNP-β-d-mannopyranoside than did Os3BGlu7 (kcat/Km of 1.01 mm−1 s−1), indicating that the equatorial position of the 2′ hydroxyl group is more critical to Os3BGlu6. Among natural substrates, Os3BGlu6 would be expected to hydrolyze β-d-glucosides, since β-d-fucosides are generally not found in plants.

In reported GH1 complex structures, the Glu corresponding to Glu-451 hydrogen bonds to both the C4 and C6 hydroxyls of the sugar residue in the −1 site (Burmeister et al., 1997; Sanz-Aparicio et al., 1998; Czjzek et al., 2000), as seen in the Os3BGlu6 complex with G2F. The flexibility of Glu-451 in the native structure may help explain the relatively low Km for pNPFuc, since it may more readily bind to the axial 4-hydroxyl and the loss of entropy that occurs upon its hydrogen bonding to the 4- and 6- hydroxyls may decrease its energetic contribution to binding. In this case, the absence of hydrogen bonding to the 6-hydroxyl in pNPFuc would have less effect than in Os3BGlu7, in which the corresponding Glu-440 residue maintains a similar position in the presence and absence of glucoside (Chuenchor et al., 2008).

Os3BGlu6 Aglycone Specificity

The hydrolysis of glucooligosaccharides and glucosides followed a rather unique pattern in Os3BGlu6, although the Km values of over 3 mm suggest that the substrates tested would only be hydrolyzed efficiently at high concentrations. The hydrolysis of β-(1→3)- and β-(1→2)-linked disaccharides in preference to longer oligosaccharides reflects the relatively short binding cleft, as will be discussed below in comparison with Os3BGlu7. The hydrolysis of the n-octyl-β-d-glucoside with high efficiency suggests that Os3BGlu6 may act on hydrophobic glycosides in the plant, rather than or in addition to disaccharides. Among the natural glycosides tested, the flavonoid glycoside apigenin-7-O-β-d-glucoside was hydrolyzed most rapidly, followed by 7-O-β-linked isoflavonoids, which suggests that a flavonoid-7-O-glucoside could serve as a hydrophobic substrate in the plant. The apoplastic or vacuolar location of action suggested by its low pH optimum and secretory signal sequence is consistent with its action on either disaccharides or flavonoids or other hydrophobic glucosides, which may be found in these compartments.

Among plant GH1 enzymes, the aglycone specificities of the chloroplastic β-glucosidases from maize and sorghum have been most extensively studied (Czjzek et al., 2000, 2001; Verdoucq et al., 2003, 2004). The crystal structures of the inactive maize ZmGlu1 E191D mutant with DIMBOAGlc (Protein Data Bank [PDB] code 1E56, Czjzek et al., 2000) and Os3BGlu6 with n-octyl-β-d-thioglucopyranoside were compared (Fig. 4, A and B). The residues that directly interact with the aglycone in ZmGlu1 include Trp-378 on one side and three Phes, Phe-198, Phe-205, and Phe-466, on the opposite side of the active site slot (Czjzek et al., 2000). Although the Trp was conserved in Os3BGlu6 (Trp-366), the opposing amino acid residues were replaced by Gln-185, Gln-192, and Ala-453 in Os3BGlu6, and the aglycone of n-octyl-β-d-thioglucopyranoside is located on the opposite side of the active site, where it makes hydrophobic contacts with other residues. Among the residues noted to indirectly affect aglycone binding in ZmGlu1, Ala-467 is conserved (Ala-454 in Os3BGlu6) while Tyr-473, which orients the plane of Trp-378 for stacking interaction with the DIMBOA substrate, is replaced by Phe-460, which has no hydroxyl to hydrogen bond to the indole ring of Trp-366 in Os3BGlu6. Thus, Trp-366 is twisted away by 1.13 Å for its interaction with the n-octyl chain of Os3BGlu6. These differences in aglycone binding may, in turn, affect the position and orientation of the glycone, as noted by Verdoucq et al. (2004).

Comparison of Substrate Binding between Os3BGlu6 and Os3BGlu7

Rice Os3BGlu6 and Os3BGlu7 have different catalytic specificities that are reflected in their active site structures. While Os3BGlu6 hydrolyzed n-heptyl and n-octyl β-d-glucosides and sophorose more efficiently (kcat/Km of 0.85, 2.7, and 0.95 mm−1 s−1 for Os3BGlu6 and 0.62, 1.25, and 0.42 mm−1 s−1 for Os3BGlu7, respectively), Os3BGlu7 more efficiently hydrolyzed laminaribiose (kcat/Km of 15.7 mm−1 s−1 versus 1.7 mm−1 s−1 for Os3BGlu6) and cellobiose (kcat/Km of 0.05 mm−1 s−1 versus 0.0085 mm−1 s−1 for Os3BGlu6). More significantly, while Os3BGlu7 hydrolyzed longer β-(1→4)-linked oligosaccharides with increasing efficiency, up to six glucosyl residues (kcat/Km of 153 mm−1 s−1 for cellohexaose; Opassiri et al., 2004), Os3BGlu6 hydrolyzed longer cellooligosaccharides more poorly than cellobiose (Table II).

Comparison of the Os3BGlu6 structure with that of Os3BGlu7 docked with the β-(1→4)-linked substrate cellotriose (Chuenchor et al., 2008) revealed differences in the shape of the active site and the residues lining the aglycone-binding region (Fig. 4, A and C). While Os3BGlu7 has a deep and relatively straight binding cleft, that of Os3BGlu6 is shorter and narrower. The distance from the deepest part of the active site slot to the surface entrance is 14 Å, which is shorter by 4 Å from that of Os3BGlu7 (18 Å), consistent with the preference of Os3BGlu6 for shorter oligosaccharides. The Os3BGlu6 glycone-binding pocket is 7.5 Å deep and 9 Å wide and is surrounded by residues conserved with Os3BGlu7 and other GH1 enzymes. The broadest region of the aglycone-binding site is 7.6 Å, while the narrowest part of the aglycone-binding site (i.e. at the entrance to the active site) is 5.3 Å, at the point where Met-251 extends into what would be the +2 subsite in Os3BGlu7. Met-251, which appears to form hydrophobic interactions with the n-octyl chain of n-octyl-β-d-thioglucopyranoside, replaces Asn-245 of Os3BGlu7, which appears to interact with cellotriose, based on the 40-fold increase in Km in the Os3BGlu7 N245V mutation (Chuenchor et al., 2008). Aside from the Met-251, many differences were observed in the residues involved in the aglycone site, including Trp-133, Thr-181, Ile-184, Gln-185, Gln-192, and Ala-453 in Os3BGlu6 in place of Tyr-131, Ile-179, Leu-182, Leu-183, Asn-190, and Leu-442 in Os3BGlu7, respectively. These differences may help explain the fact that Os3BGlu6 prefers hydrophobic β-glucosides and short oligosaccharides while Os3BGlu7 prefers longer β-(1→4)-linked oligosaccharides.

CONCLUSION

As plant GH1 enzymes exhibit a wide variety of specificities and release products with a wide range of physiological functions, the structural basis for this functional diversity is a significant problem. The enzymatic characterization of Os3BGlu6, along with the structures of Os3BGlu6, its covalent intermediate with G2F, and its noncovalent complex with the substrate analog n-octyl-β-d-thioglucopyranoside, extend the understanding of this problem to a new cluster of GH1 in plants. Small differences in the active site amino acids appear to explain the difference between rice Os3BGlu7, which prefers long, straight glucooligosaccharides, and Os3BGlu6, which prefers shorter hydrophobic glycosides. By extension, the small differences in the active sites of plant GH1 enzymes and their impact on the substrate specificity can help explain the expansion of this family to fulfill a range of functional roles in the plant.

MATERIALS AND METHODS

Cloning and PCR Amplification of cDNA Encoding Mature Os3BGlu6

To produce a cDNA library, 7-d-old rice (Oryza sativa ‘Yukihikari’) seedlings were chilled at 5°C for a further 4 d, and then the RNA was extracted by the method of Bachem et al. (1996). Poly(A) RNA was isolated from the total RNA with magnetic poly-T beads (Dynal). The poly(A) RNA was reverse transcribed and used to produce a library with the λZAP-cDNA/Gigapack cloning kit (Stratagene). This library was differentially screened with cDNA reverse transcribed from cold-stressed seedling and control seedling RNA, which had been labeled by random priming with [α-32P]dCTP with the Rediprime II random prime labeling system (GE Healthcare). The GH1 cDNA encoding Os3BGlu6 was selected as a differentially expressed clone, sequenced thoroughly, and submitted to GenBank as accession number AY129294.

The cDNA encoding the predicted mature Os3BGlu6 rice β-glucosidase was amplified by PCR with the specific primers SapporoMstrtF (5′-CACCATGGCGCAGCAGAGCGGAG-3′) and SapporoStopR (5′-GGAGTTCAGGTCTTCAGGAG-3′). Amplification was performed with 30 cycles of 94°C for 1 min, 60°C for 45 s, and 72°C for 1.5 min, with Pfu DNA polymerase (Promega). The PCR product was cloned into pENTR/TEV/D-TOPO according to the supplier's directions (Invitrogen). The cDNA was subcloned into the pET32a(+)/DEST expression vector (Opassiri et al., 2006) by LR Clonase recombination (Invitrogen). The recombinant pET32a(+)/DEST/Os3BGlu6 expression clone was verified by automated DNA sequencing with the T7 promoter and terminator and specific internal primers (F427, 5′-CTTGACAGGCAGATAGTGGA-3′; R600, 5′-GCAGTAGAGGTGGAGCAG-3′; F969, 5′-GCACAACAACACCAACATC-3′; and R946, 5′-CGTGTAGTAGGTGGTGTAGTG-3′).

Expression of Rice β-Glucosidase Os3BGlu6 in Escherichia coli and Purification

Os3BGlu6 was expressed in E. coli Origami (DE3) as a fusion protein with N-terminal thioredoxin and His6 tags. Expression was induced at 20°C and purified by immobilized metal (Co2+) affinity chromatography (IMAC; on Talon resin [Clontech]) as described previously for rice OsBGlu7 (Chuenchor et al., 2006). The active elution fractions containing pNPGlc hydrolysis activity were pooled and concentrated, and the buffer was changed to 20 mm Tris-HCl, pH 8.0, in a 30-kD cutoff ultrafiltration membrane (Amicon YM-30; Millipore). The N-terminal thioredoxin, His6, and S tags were then excised with TEV protease, produced as described by Kapust et al. (2001), at a ratio of 1:50 (w/w) TEV protease to fusion protein for 12 to 14 h at 4°C. The cleaved N-terminal fusion tag and TEV protease were removed by adsorption to Co2+ IMAC resin. The flow-through and 5 mm and 10 mm imidazole wash fractions containing activity were pooled, the buffer was changed to 20 mm Tris-HCl, pH 8.0, and the pool was concentrated by ultrafiltration (Amicon YM-10; Millipore). Protein concentrations were determined by the Bradford protein assay (Bio-Rad) with bovine serum albumin as the standard.

Os3BGlu6 pH and Temperature Optimum and Stability Studies

The optimum pH was determined by incubating 1 μg of enzyme in a reaction volume of 140 μL containing 100 mm universal buffer (citric acid-disodium hydrogen phosphate), pH 2 to 11, at 0.5-pH-unit increments with 1 mm pNPGlc for 10 min at 30°C. The reaction was stopped with 100 μL of 2 m sodium carbonate. The amount of pNP released was determined from its A405. The enzyme's pH stability was studied by incubating 25 μg of enzyme in 20 μL of universal buffer at the same pH range for time periods of 10 min, 1, 2, 6, 12, and 24 h at 30°C. The enzyme was diluted 400-fold in 100 mm buffer at the optimum pH, and the activity was determined as described above. The temperature optimum was determined by incubating 1 μg of enzyme in 100 mm sodium acetate, pH 5.0, over a temperature range of 5°C to 100°C in increments of 5°C for 10 min, and then pNPGlc was added to the reaction to 1 mm final concentration and incubated for 10 min. Temperature stability was determined by preincubating 1 μg of enzyme over a temperature range of 5°C to 70°C. At 10-min increments from 0 to 60 min, aliquots of the samples were placed on ice and the enzyme activity at 30°C was determined as described above.

Substrate Specificity and Enzyme Kinetics

Os3BGlu6 was tested for hydrolysis of glycosides of pNP and commercially available natural and synthetic substrates (listed in Tables I and II) by measuring release of pNP or Glc from the substrates and by thin-layer chromatography (TLC). Activity was assayed in triplicate in 100 mm sodium acetate, pH 5.0, at 30°C. For TLC, the reactions were stopped after overnight digest by boiling 5 min, and products from the reactions were spotted onto silica gel F254 plates (Merck) and chromatographed with solvents of ethyl acetate:acetic acid:water (2:1:1, v/v) for the oligosaccharides and ethyl acetate:acetic acid:methanol:water (15:2:1:2, v/v) for natural glycosides. The products were visualized under UV light at 254 nm and then by coating the plates with 10% (v/v) sulfuric acid in methanol and heating at 120°C until the spots were visible. For oligosaccharides and glucosides other than pNPGlc that were detectably hydrolyzed on TLC, Glc release was measured by the Glc oxidase/peroxidase-coupled assay, as described previously (Opassiri et al., 2003).

Kinetic parameters were determined for substrates of interest, based on the initial relative activity assays. The initial velocities were determined at time points and enzyme concentrations where the reaction rate was linear and the absorbance value was in the range of 0.1 to1.0 and used to calculate the kinetic constants. Substrate concentrations over a range of approximately one-fifth to five times the apparent Km were included. The kcat, Km, and kcat/Km were calculated by nonlinear regression of Michaelis-Menten plots with Grafit 5.0 (Erithacus Software). Competitive Ki values were determined by incubating 1 μg of enzyme with eight different concentrations of the inhibitors (0–30 mm) in 100 mm sodium acetate buffer, pH 5.0, in the presence of 0 to 20 mm pNPGlc substrate under the reaction conditions described above. Lineweaver-Burk and Dixon plots were used to calculate the inhibition constants.

Screening and Optimization for Os3BGlu6 Crystallization

Purified Os3BGlu6 in 20 mm Tris-HCl, pH 8.0, was filtered through an Ultrafree MC 0.22-μm filter (Millipore) and used to screen for crystallization conditions by the microbatch under the oil method at 15°C with precipitants from the Crystals Screen High Throughput HR2-130 and HR2-134 kits (Hampton Research). Clusters of crystals were seen in the condition with 0.1 m Bis-Tris, pH 6.5, and 20% polyethylene glycol monomethyl ether (PEG MME) 5000 (in the HR2-134 kit) after 200 d. Optimization with microseeding yielded diffraction-quality native crystals of Os3BGlu6 in 21% PEG5000 MME, 0.1 m Bis-Tris, pH 6.5. The crystals were soaked in the cryoprotectant (24.15% PEG5000 MME, 0.115 m Bis-Tris, pH 6.5, and 18% [v/v] glycerol) for 5 to 10 s before flash freezing in liquid nitrogen. To obtain Os3BGlu6 crystals with inhibitors, native crystals were soaked in 5 mm 2-deoxy-2-fluoro-β-d-glucoside or 10 mm n-octyl-β-d-thioglucopyranoside overnight in 19% PEG5000 MME, 0.1 m Bis-Tris, pH 6.5. The cryoprotectants contained the same concentrations of inhibitors present in 21.85% PEG5000 MME, 0.115 m Bis-Tris, pH 6.5, and 18% (v/v) glycerol.

Data Collection, Processing, and Structure Refinement

The data sets for native Os3BGlu6, the Os3BGlu6/G2F complex, and the Os3BGlu6/n-octyl-β-d-thioglucopyranoside complex were collected on the PX 13B beamline at the National Synchrotron Radiation Research Center, Hsinchu, Taiwan, with a 1.0-Å wavelength x-ray beam and an ADSC Quantum 315 CCD detector. The crystals were maintained at 110 K with a cold stream of nitrogen (CryoJet; Oxford Instrument) throughout diffraction. All data sets were indexed, integrated, and scaled with the HKL-2000 package (Otwinowski and Minor, 1997). The crystals belonged to space group P212121 with the unit cell parameters shown in Table III. The crystal structure for native Os3BGlu6 was solved by molecular replacement with the white clover (Trifolium repens) cyanogenic β-glucosidase structural model (PDB code 1CBG; Barrett et al., 1995) with the MolRep program in the CCP4 suite (Vagin and Teplyakov, 1997). A subset of 5% of the structure factor amplitudes was reserved for Rfree determination. Iterative refinement was done with Refmac 5.0 (Murshudov et al., 1997; CCP4 suite), and the structure was built into the electron density map with the coot graphics program (Emsley and Cowtan, 2004). The overall quality of each model was evaluated with the program PROCHECK (Laskowski et al., 1993). The structures of the Os3BGlu6/G2F and Os3BGlu6/n-octyl-β-d-thioglucopyranoside complexes were solved by molecular replacement with the native structure as the template model. Structural figures were generated in Pymol (Delano Scientific). Structural comparisons between Os3BGlu6 and other known structures from GH1 β-glucosidase were executed with the Superpose program of the CCP4 suite (Krissinel and Henrick, 2004).

N-Terminal Amino Acid Sequencing of Os3BGlu6

Os3BGlu6 crystal clusters were dissolved in sample buffer and separated on a 12% polyacrylamide gel by the method of Laemmli (1970). The gel was electroblotted onto a Biotrace polyvinylidene fluoride membrane (Pall Corporation) and stained with Coomassie Brilliant Blue R250. The protein bands were excised and Edman degradation sequencing was performed at the Prince of Songkla University Protein Analysis Center.

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers AY129294, 3GNO, 3GNP, and 3GNR.

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure S1. SDS-PAGE analysis of Os3BGlu6 purification.

Supplementary Material

[Supplemental Data]
pp.109.139436_index.html (1,003B, html)

Acknowledgments

We are grateful to Phimonphan Chuankhayan and Chun-Jung Chen and his laboratory for assistance in crystal screening and to the staff of the National Synchrotron Radiation Research Center PX 13B beamline for assistance in data collection. Robert C. Robinson is thanked for data processing advice. Prof. Jisnuson Svasti is thanked for advice and support on the project and for providing dalcochinin glucoside.

1

This work was supported by the Synchrotron Light Research Institute of Thailand.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: James Ketudat Cairns (cairns@sut.ac.th).

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Some figures in this article are displayed in color online but in black and white in the print edition.

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The online version of this article contains Web-only data.

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Open Access articles can be viewed online without a subscription.

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