The Crystal Structure of Human Cytosolic β-Glucosidase Unravels the Substrate Aglycone Specificity of a Family 1 Glycoside Hydrolase

doi:10.1016/j.jmb.2007.05.034

J. Mol. Biol. (2007) 370, 964–975

The Crystal Structure of Human Cytosolic β-Glucosidase Unravels the Substrate Aglycone Specificity of a Family 1 Glycoside Hydrolase Sandra Tribolo 1 , Jean-Guy Berrin 2 , Paul A. Kroon 1 , Mirjam Czjzek 3 ⁎ and Nathalie Juge 1,2 1

Institute of Food Research, Norwich Research Park, Colney, Norwich NR4 7UA, UK 2

Biosciences FRE-3005-CNRS, Université Paul Cézanne Aix Marseille III, Av. Escadrille Normandie-Niemen, 13397 Marseille Cedex 20, France 3

Centre National de la Recherche Scientifique; Université Pierre et Marie Curie-Paris 6, Unité Mixte de Recherche 7139 “Marine Plants and Biomolecules”, Station Biologique, Place George Teissier, BP74, F-29682 Roscoff Cedex, France

Human cytosolic β-glucosidase (hCBG) is a xenobiotic-metabolizing enzyme that hydrolyses certain flavonoid glucosides, with specificity depending on the aglycone moiety, the type of sugar and the linkage between them. In this study, the substrate preference of this enzyme was investigated by mutational analysis, X-ray crystallography and homology modelling. The crystal structure of hCBG was solved by the molecular replacement method and refined at 2.7 Å resolution. The main-chain fold of the enzyme belongs to the (β/α)8 barrel structure, which is common to family 1 glycoside hydrolases. The active site is located at the bottom of a pocket (about 16 Å deep) formed by large surface loops, surrounding the C termini of the barrel of β-strands. As for all the clan of GH-A enzymes, the two catalytic glutamate residues are located on strand 4 (the acid/base Glu165) and on strand 7 (the nucleophile Glu373). Although many features of hCBG were shown to be very similar to previously described enzymes from this family, crucial differences were observed in the surface loops surrounding the aglycone binding site, and these are likely to strongly influence the substrate specificity. The positioning of a substrate molecule (quercetin-4′-glucoside) by homology modelling revealed that hydrophobic interactions dominate the binding of the aglycone moiety. In particular, Val168, Trp345, Phe225, Phe179, Phe334 and Phe433 were identified as likely to be important in determining substrate specificity in hCBG, and sitedirected mutagenesis supported a key role for some of these residues. © 2007 Elsevier Ltd. All rights reserved.

*Corresponding author

Keywords: human β-glucosidase; aglycone specificity; crystal structure; sitedirected mutagenesis; flavonoid glycosides

Introduction Glycoside hydrolases (GH; EC 3.2.1.–) are a widespread group of enzymes that hydrolyse the glycosidic bond between two or more carbohydrates or between a carbohydrate and a non-carbohydrate moiety. To date, GHs have been classified into 108 families on the basis of amino acid sequence Abbreviations used: GH, glycoside hydrolase; hCBG, human cytosolic β-glucosidase; KL, klotho; KLPH, Klotho-LPH-related; LPH, lactase-phlorizin hydrolase; pNP, para-nitrophenol; wt, wild-type; ZMGlu, maize (Zea mays) β-glucosidase. E-mail address of the corresponding author: [email protected]

similarities (see the CAZy database†).1 In some cases, these families are further grouped into clans, which display the same structural folds and catalytic apparatus. In humans, inheritable deficiencies in certain glycosidases are known to induce a variety of impairments, such as lysosomal storage diseases, Gaucher's and Krabbe's diseases, 2 and lactose intolerance.3 GH family 1 consists of enzymes with varying substrate specificities, such as β-glucosidase (EC 3.2.1.21), β-galactosidase (EC 3.2.1.23), β-mannosidase (EC 3.2.1.25), β-glucuronidase (EC 3.2.1.31), β-D-fucosidase (EC 3.2.1.38), phlorizin hydrolase (EC 3.2.1.62), 6-phospho-β-galactosidase (EC 3.2.1.85),

0022-2836/$ - see front matter © 2007 Elsevier Ltd. All rights reserved.

† http://www.cazy.org

Crystal Structure of Human Cytosolic β-Glucosidase

965

6-phospho- β-glucosidase (EC 3.2.1.86), strictosidine β-glucosidase (EC 3.2.1.105), lactase (EC 3.2.1.108), prunasin β-glucosidase (EC 3.2.1.118), raucaffricine β-glucosidase (EC 3.2.1.125), thioglucosidase (EC 3.2.1.147), β-primeverosidase (EC 3.2.1.149), isoflavonoid 7-O- β-apiosyl β-glucosidase (EC 3.2.1.161) and hydroxyisobutyrate hydrolase (EC 3.-.-.-). These enzymes are widely distributed among the Bacteria, Archaea and Eukaryota. GH1 members operate with a molecular mechanism leading to overall retention of the anomeric configuration, and involving the formation and breakdown of a covalent glycosyl enzyme intermediate. To date, the 3D structures of 18 GH1 β-glucosidases (in either resting, complexed, or transition state) have been determined from Eukaryota,4–12 Bacteria13–18 and Archaea.19–22 Although the extent of sequence identity varies between 17% and 45% in the β-glucosidases, all the enzymes display a common (β/α)8-barrel motif. Apart from plant myrosinases and Klotho (KL) family members, all family 1 β-glucosidases conserve two catalytic glutamate residues located at the C-terminal end of β-strands 4 and 7.23 Interestingly, Eukaryota β-glucosidases 3D structures have been reported only for plant enzymes (from Zea mays,7,12 Triticum aestivum,10 Trifolium repens, 4 Sorghum bicolour, 11 Sinapsis alba,5 Rauvolfia sepentina9 and Oryza sativa6) and for an insect myrosinase (from Brevicoryne brassicae8) and there is a complete lack of structural information on mammalian GH1 enzymes. Human cytosolic β-glucosidase (hCBG) is a GH1 enzyme that hydrolyses certain flavonoid glucosides, with the specificity dependent on the aglycone moiety, the type of sugar and the linkage between them.24,25 This enzyme is present in the liver, kidney, intestine and spleen of humans.26 Other human GH1 enzymes include KL,27–29 βKlotho (βKL),30 Klotho-LPH-related (KLPH)31 and lactase-phlorizin hydrolase (LPH).32 LPH is a major membrane-bound intestinal glycosidase contributing to the metabolism of dietary lactose in mammals.3 KL is a multi-functional protein that regulates phosphate/calcium metabolism as well as aging.33 The human KL members, including KLPH and β-KL, both type I membrane proteins, with 37.5% to 41.2% similarity to KL, have yet to be characterised enzymatically. Recently, a KL member from mouse was classified as a β-glucuronidase on the basis of its substrate specificity and sensitivity against inhibitors,34 but it is uncertain whe-

ther KL family proteins have enzymatic activities in vivo. Although structural predictions have been made, the lack of an experimental 3D structure of a human/mammalian GH1 has hampered attempts to establish the detailed catalytic mechanism (substrate binding/activating, inhibitor binding) for this subgroup of enzymes. Here, we report the X-ray structure of hCBG at 2.7 Å resolution and, using homology modelling of a known substrate for the enzyme, identify residues that are likely to contribute to enzyme specificity. Further, the role of several of the residues identified as likely to be important in substrate binding, including V168, Y308, F225, M172, Q307 and W345 in the substrate binding pocket, were examined via a kinetic analysis of mutant enzymes.

Results and Discussion Preparation of recombinant wild-type human CBG (wt-hCBG) The recombinant wild-type human CBG (wthCBG), produced in Pichia pastoris was purified to homogeneity using two chromatographic steps. The single hydrophobic-interaction purification step used in our previous studies was insufficient to obtain a protein for crystallisation purposes.24,25 An additional ion-exchange step was therefore included, resulting in highly pure recombinant hCBG protein of 45 U mg−1 specific activity toward para-nitrophenol (pNP)Glc, corresponding to a 12-fold purification (data not shown). The additional MonoQ chromatography step eliminated the green pigment that remained associated with the recombinant hCBG in P. pastoris culture supernatant and that, we believed, hampered the crystallisation of the recombinant enzyme. The specific activity of wthCBG on pNPαAra, pNPβGlc, pNPβFuc and pNPβGal was 2.1 U mg−1, 3.5 U mg−1, 3.6 U mg−1 and 5.7 U mg−1, respectively (Table 2), and pNPβFuc was the best substrate with regards to catalytic specificity (kcat/Km) (Table 2). wt-hCBG hydrolysed many flavonoids efficiently when glucose substitution occurred at the 4′ and 7 positions. The highest catalytic efficiencies were obtained for quercetin-4′glucoside and apigenin-7-glucoside (49.7 mM−1 s−1 and 41.8 mM−1 s−1, respectively) (Table 3).

Table 1. Primers used to generate mutant hCBG Mutant

Direction

Primer sequence 5′→3′

Met172→Leu Met172→Leu Gln307→Asn Gln307→Asn Trp345→Met Trp345→Met Trp345→Ala Trp345→Ala

Forward Reverse Forward Reverse Forward Reverse Forward Reverse

AATGTTCTTTCTGTGTTGTCATATGACTTAGGTATGTTTCCTCCG CGGAGGAAACATACCTAAGTCATATGACAACACAGAAAGAACATT GGCACTGCTGATTTTTTTGCTGTGAACTATTATACAACTCGC GCGAGTTGTATAATAGTTCACAGCAAAAAAATCAGCAGTGCC CCATCTTGGAAAAATGTGGATATGATCTACGTGGTACCATGGGG CCCCATGGTACCACGTAGATCATATCCACATTTTTCCAAGATGG CCATCTTGGAAAAATGTGGATGCGATCTACGTGGTACCATGGGG CCCCATGGTACCACGTAGATCGCATCCACATTTTTCCAAGATGG

Residues in bold indicate the mutation site.

Crystal Structure of Human Cytosolic β-Glucosidase

966

Table 2. Specific activity (SA) and kinetic parameters of wt and mutant hCBG on pNP-glycosides Substrates

Enzymes

SA (U mg−1)

Km (mM)

kcat (s−1)

pNP β Fuc

wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A

3.6 3.1 0.47 0.03 0.004 2.0 2.2 0.23 0.02 – 3.5 2.9 0.49 0.03 0.008 5.7 5.1 0.50 0.04 0.003 0.23 0.02 0.02 – –

0.12 ± 0.02 0.13 ± 0.01 0.09 ± 0.01 0.15 ± 0.02 – 0.19 ± 0.01 0.12 ± 0.01 0.27 ± 0.01 0.61 ± 0.02 – 0.38 ± 0.02 0.23 ± 0.02 0.27 ± 0.02 0.33 ± 0.02 0.20 ± 0.02 0.90 ± 0.05 1.20 ± 0.05 0.35 ± 0.08 – – 5.10 ± 0.7 4.80 ± 0.3 7.20 ± 0.01 – –

3.13 ± 0.06 2.69 ± 0.02 0.39 ± 0.01 0.02 ± 0.00 – 1.88 ± 0.02 1.99 ± 0.02 0.25 ± 0.00 0.02 ± 0.00 – 3.12 ± 0.03 2.57 ± 0.03 0.46 ± 0.01 0.03 ± 0.00 0.01 ± 0.00 5.53 ± 0.08 5.20 ± 0.05 0.46 ± 0.02 – – 0.29 ± 0.02 0.33 ± 0.00 0.03 ± 0.00 – –

pNP α Ara

pNP β Glc

pNP β Gal

pNP β Ara

kcat/Km (mM−1 s−1) 26.1 20.7 4.3 0.1 – 9.9 16.6 0.9 0.03 – 8.2 11.2 1.7 0.09 0.05 6.1 4.3 1.3 – – 0.06 0.07 0.005 – –

Specific activities are mean data (n ≥ 2) with typical standard deviation below 10% when applicable. They were determined with a substrate concentration of 10 mM for pNP-glycosides.

Overall structure So far, all Eukaryota GH1 representatives for which a 3D structure has been determined are from plants, including maize,7,12 wheat,10 rice,6 white clover,4 sorghum,11 yellow mustard,5 devil pepper,9 or insect (cabbage aphid myrosinase)8 and hCBG

represents the first tertiary structure from a mammalian species. The structure of the monomeric hCBG molecule is presented as a ribbon model in Figure 1(a). The human CBG exhibits the (β/α)8barrel fold characteristic of GH1 β-glucosidases, with structural differences being confined mainly to loop regions (Figure 1(a)), that have been defined as

Table 3. Specific activity and kinetics parameters of wt and mutant hCBG on flavonoid glucosides Substrates

Enzymes

SA (U mg− 1)

Km (μM)

kcat (s− 1)

kcat/Km (mM− 1 s− 1)

Quercetin-4′-Glc

wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A wt-hCBG M172L Q307N W345M W345A

1.6 1.4 0.18 0.008 0.002 0.56 0.50 0.07 0.08 0.003 0.81 0.81 0.10 0.02 0.01 1.2 1.2 0.26 0.01 0.005 0.64 0.89 0.06 0.01 0.005

30.2 ± 1.7 41.5 ± 2.0 30.2 ± 3.6 20.9 ± 2.3 23.9 ± 3.9 36.1 ± 7.1 41.1 ± 7.3 17.9 ± 4.8 36.9 ± 6.5 39.2 ± 7.7 18.4 ± 2.3 23.7 ± 3.1 22.1 ± 2.8 91.5 ± 10.5 122 ± 9 210 ± 18 148 ± 6 272 ± 37 72.7 ± 8.9 122 ± 32 122 ± 11 120 ± 2 76.7 ± 17.3 98.1 ± 14.6 93.6 ± 14.1

1.50 ± 0.02 1.38 ± 0.02 0.17 ± 0.00 0.01 ± 0.00 0.002 ± 0.00 0.60 ± 0.04 0.55 ± 0.03 0.05 ± 0.00 0.01 ± 0.00 – 0.77 ± 0.03 0.78 ± 0.03 0.12 ± 0.00 0.02 ± 0.00 0.01 ± 0.00 1.40 ± 0.06 1.37 ± 0.03 0.25 ± 0.02 0.01 ± 0.00 0.01 ± 0.00 0.73 ± 0.03 0.98 ± 0.00 0.06 ± 0.00 0.01 ± 0.00 0.01 ± 0.00

49.7 33.3 5.6 0.02 0.08 16.6 13.4 2.8 0.3 – 41.8 33.0 5.4 0.2 0.08 6.7 9.2 0.9 0.1 0.08 6.0 8.1 0.8 0.1 0.1

Quercetin-7-Glc

Apigenin-7-Glc

Naringenin-7-Glc

Eriodictyol-7-Glc

Specific activities are mean data (n ≥ 2) with typical standard deviation below 10% when applicable. They were determined with a substrate concentration of 500 μM for flavonoid glucosides.

Crystal Structure of Human Cytosolic β-Glucosidase

967

Figure 1. (a) Ribbon representation of the monomeric structure of hCBG. The loops surrounding the active site pocket are highlighted in red (loop A), orange (loop B), yellow (loop C) and green (loop D). (b) The final (2Fo–Fc) electron density map calculated at a 1σ level around the glycosylation site showing the presence of two GlcNac units attached to Asn46. (c) Superimposition of the 3D structure of different eukaryotic GH1 β-glucosidases: yellow, Brevicoryne brassica myrosinase (1WCG); blue, Triticum aestivum β-glucosidase (2DGA); red, Zea maize β-glucosidase (1E4N); and green, human cytosolic β-glucosidase. The highest variations are observed in the loop regions A through D. (d) Surface representation of hCBG showing the oblong-shaped depression that forms the active site pocket. The catalytic glutamic acid residues positioned at the centre of the barrel are coloured green. (a) and (c) were drawn with MOLSCRIPT64 and Raster3D.65 (b) was prepared with Turbo.61 (d) was drawn with PyMol [http://pymol.sourceforge.net/].

loop A (residues 34–50 in hCBG), loop B (residues 173–187 in hCBG), loop C (residues 318–348 in hCBG) and loop D (residues 378–385).7 In contrast to several other eukaryotic β-glucosidases, hCBG does not contain any disulfide bridge between any of the eight cysteine residues present in the primary sequence. Electron density was observed for two carbohydrate units at one glycosylation site, Asn46, which is at the tip of loop A (Figure 1(b)). As modelled from the electron density, two β-1,4 linked N-acetylglucosamine (GlcNAc) units are attached to this glycosylation site. Notably, these sugar units are involved

in crystal contacts to a neighbouring hCBG molecule. The superimposition of the hCBG structure with those from maize (PDB 1E4N), sorghum (PDB 1V02) or insect (PDB 1WCG) led to the root-mean-square deviation (r.m.s.d.) values reported in Table 4. As expected, they mostly fitted well, with a high level of similarity at the core of the β-barrel structure and differences occurring at the rim of the active site pocket that is formed by loops A through D (Figure 1(c)). The structural differences in conformations within these loops are concomitant with a low level of sequence similarity (Supplementary Data Figure

Crystal Structure of Human Cytosolic β-Glucosidase

968 Table 4. Root-mean-square deviations between hCBG and eukaryotic β-glucosidases and myrosinases with known 3D structure Protein name (PDB) and organism Myrosinase (1WCG) Brevicoryne brassica Cyanogenic β-glucosidase (1CBG) Trifolium repens Dhurrinase (1V02) Sorghum bicolor Myrosinase (1MYR) Sinapis alba β-Glucosidase (1E4N) Zea maize β-Glucosidase (2DGA) Triticum aestivum

Root-mean-square deviation and the number of Cα atoms compared 0.97 for 441 Cα atoms 1.06 for 443 Cα atoms 1.06 for 436 Cα atoms 1.09 for 434 Cα atoms 1.10 for 436 Cα atoms 1.12 for 436 Cα atoms

1(a)). These results are in agreement with the cladogram (Supplementary Data Figure 1(b)), which also shows that, within the eukaryotic GH1 β-glucosidases of known 3D structure, hCBG is more closely related to the insect myrosinase from Brevicoryne brassicae (PDB 1WCG) than it is to β-glucosidases from plants, such as Triticum aestivum (PDB 2DGA) (Table 4). When compared with the other known structures of eukaryotic GH1 β-glucosidases, hCBG is noticeably different in the structure of the top half of the barrel (i.e. in the connections between the C termini of the barrel strands and the N termini of the barrel helices). Structural variations between the human enzyme and the related glucosidases are concentrated mainly on the four loops (Figure 1(a) and (c)) surrounding the active site, whereas the structure on

the other side of the barrel is well conserved. Interestingly, these four loops are responsible for the overall shape of the aglycone binding pocket and support the crucial residues defining the binding affinities of it. Other differences in the overall structure of hCBG are found in the regions involved in the dimerisation of ZMGlu,7 with a considerably higher content of charged residues in the extended loop region connecting β6 and α6 (loop C). The corresponding region in ZMGlu contains several hydrophobic residues that are covered by the dimer interface. The connection between α5 and β6, forming the extra-barrel helices αE and αF,7 is also notably longer in hCBG. Oligomerisation is a common characteristic of GH1 β-glucosidases, ranging from dimeric to octameric quaternary organisations. However monomeric molecules are found also in all three domains i.e. Eukaryota,8 Bacteria13,16–18 and Archaea20 of living organisms whose 3D structures have been determined, so no simple relationship can be derived. Dimerisation and other levels of quaternary associations of β-glucosidases have been shown to be important for stability and/ or activity of these enzymes.35 The hCBG is active as a monomer,24,36 suggesting that oligomerisation is not a strict requirement for stability/activity of family 1 β-glucosidases. The active site pocket of hCBG and comparison with other GH1 β-glucosidases The active site of hCBG is located on the bottom of an approximately 16 Å deep pocket (Figure 1(d)). It

Figure 2. (a) Detailed view of the residues surrounding the glycone binding site. The position of the modelled glucose-moiety is shown. Residues potentially forming hydrogen bonds with the substrate are drawn in stick representation. The catalytic glutamate residues are coloured red. The single amino acid difference (Gln307) with respect to other GH1 β-glucosidases protrudes into the glycone binding pocket and is coloured blue. For clarity, the aglycone moiety is not represented. (b) Detailed view of the residues lining the walls of the aglycone binding pocket. The most probable position of the aglycone moiety is shown. The glycone moiety is omitted for clarity. The residues forming the hydrophobic cluster on each side of the flavonoid moiety are drawn in stick representation. This Figure was prepared using MOLSCRIPT64 and Raster3D.65

Crystal Structure of Human Cytosolic β-Glucosidase

can be divided into two parts; the recognition site for the glucose moiety located at the bottom of the active site and the binding site for the aglycone (Figure 2(a) and (b)) at the entrance of the pocket. The active site is shaped as an oval or slit-like pocket with a cluster of hydrophobic amino acid sidechains lining the walls of the aglycone binding slot, whereas hydrophobic, polar and charged residues are present in the glycone binding moiety. Enzyme–substrate interactions have been investigated from many family GH1 enzymes in studies involving co-crystallization with inhibitors7,15,17,18,37,38 or natural substrates.4,10,11,39 The glucose binding site has been established using, for example, 2-Fglucose derivatives of substrates and appears to be well-conserved for all GH1 β-glucosidases.5,40,41 The involvement of the two glutamate residues in catalysis, one acting as a nucleophile and the other as an acid/base, has been established using various approaches, including mutagenesis and inhibitor studies,42–44 and confirmed in several GH1 enzymes for which the 3D structure is known.10,12,22,39,45,46 The nature of the amino acids forming the glucose binding site of hCBG is highly similar to other family 1 enzymes. Besides the two catalytic residues Glu165 and Glu373, four mainly hydrophobic amino acids (His120, Phe121, Trp417 and Trp425) and Gln17, Arg75, Asn164, Tyr309 and Glu424, all highly conserved in family 1 enzymes, are grouped around the binding site (Figure 2(a)). All these residues are involved in the interaction with the glycoside. The pattern of specific binding to the glucose moiety has been described extensively in several structural studies on GH1 enzymes in complex with inhibitor or substrate molecules.5,11,40,41,47 The only structural difference in the glucose binding site between the human and the other eukaryotic GH1 β-glucosidases is the presence of Gln307 protruding between the two catalytic residues (Figure 2(a)). In all known structures of family 1 β-glucosidases this amino acid is an Asn residue, and the shorter side-chain prevents it from hydrogen bonding to the glucose OH2 group. In the present study, we produced and tested the activity of a Gln307→Asn mutant against a range of substrates. For pNP-glycosides, the catalytic efficiency of Gln307Asn was reduced by five- to tenfold, compared to the wild-type (Table 2). The mutation also induced a decrease in specific activity of 86.9–91.4%. In the hCBG 3D structure, the elongated side-chain of Gln307 is within the minimum distance required for hydrogen bonding to OH2, which likely explains why this enzyme is more sensitive to the stereochemistry of the OH in position 2, in contrast to other GH1 β-glucosidases. This appears to be a stereochemical determinant of the inhibition strength between hCBG and mannose, which has an axial OH2 group. Taken together, these findings suggest that this residue, uniquely for family 1 β-glucosidases, is essential for catalysis in human CBG. In a previous study, we showed that the Phe225→Ser and Tyr308→Ala mutations and, to a lesser degree, the Tyr308→Phe mutation, resulted in drastic reductions in the specific activities for all the

969 substrates tested. These decreases in activity were interpreted as an indication of an important role of those residues in catalysis. However, as seen in Figure 2(b), the side-chain of Tyr308 is turned away from the pocket and is therefore more likely to influence the overall structural arrangements in this region, including the precise form of the active site aglycone pocket. Phe225, on the other hand, is located within the aglycone pocket and is almost certainly crucial for aglycone recognition and specificity (Figure 2(b) and see below). As mentioned earlier, no other 3D structures of mammalian GH1 β-glucosidases have been reported. In order to obtain greater insights into the relationship between hCBG and its mammalian counterparts, phylogenetic analysis based on the neighbour-joining method was performed by CLUSTAL W software.48 Human CBG was classified into the mammalian subgroup of the GH family 1 with KL, β-KL, LPH and KLPH (Supplementary Data Figure 1(c)). The mature form of LPH is composed of two internal repeats (LPH3 and LPH4, each belonging to GH1) and a transmembrane domain with a short cytoplasmic domain at the carboxyl terminus.32 Each of the LPH repeats possesses the two catalytic glutamic acid residues of GH family 1. KL and β-KL consist of two internal repeats with homology to GH1 glucosidases, whereas KLPH has a single extracellular domain with similarity to the KL1 domain of the Klotho proteins.27–31 In this subgroup, hCBG showed highest similarity with LPH3 (46%) and LPH4 (47%), KL1 and β-KL1 (43%), and KLPH (43%). Interestingly, members from the mammalian KL family do not have the two essential glutamic acid residues, common to other GH1 family β-glucosidases.49 In both KL and β-KL, Glu for the acid-base catalyst is replaced by an Asn in the first internal repeat, while Glu for the nucleophile is replaced by an Ala or a Ser in the second internal repeat.27–30 In mouse KLPH, the nucleophile is conserved but the acid-base catalyst replaced by an Asp.31 However, no structural information is available for these proteins and, although mouse Klotho was shown recently to have weak glycosidase activity towards glucuronylated steroids,34 it is uncertain whether KL family proteins have enzymatic activities in vivo. Comparison of the hCBG 3D structure described here with those of KL family proteins that may be described in the future, should provide insights into the basis of the unusual specificity of KL family proteins, since all previously known β-glucuronidases (EC 3.2.1.31) belong to glycosidase family 2. The occurrence of glycosidases of differing substrate specificity in the same family illustrates a common evolutionary origin. Several glycosidase families are grouped together in a large superfamily called clan GH-A, which includes families 1, 2, 5, 10, 17, 26, 30, 35, 39, 42, 51, 53, 59, 72, 79 and 86 (CAZy database) and covers a wide range of substrate specificities. In addition, novel members structurally related to glycosidases, but not isofunctional to their glycosidase ancestors, have emerged recently.50

970 The aglycone binding pocket of hCBG In contrast to the glycone moiety, the structure of the aglycone binding pocket is extremely variable within the family of GH1 enzymes; this is consistent with the large variety of substrates involved in reactions catalyzed by β-glucosidases. Most of the family GH1 β-glucosidases display the highest level of activity for β-D-glucosides, and the substrate variation is the consequence of the varying aglycone specificity. Similar to all GH1 β-glucosidases, the cleft-like gate to the hCBG active site is formed by four extended, solvent-exposed loops (A–D), highly variable throughout the family (Figure 1(c); Supplementary Data Figure 1(a)). These loops carry the residues responsible for aglycone recognition and binding. In the hCBG 3D structure, loops B, C and D are shorter than in many other GH1 β-glucosidases, resulting in a flatter rim to the pocket entrance. As seen in Figure 1(d), the oblong shaped entrance is not extremely deep, and this might explain the inability of hCBG to hydrolyse large substrates such as glycosphingolipids and mucopolysaccharides.51 The overall shape and hydrophobic character of the aglycone binding subsite of the hCBG structure is in agreement with the substrate specificity of hCBG, which prefers rigid, planar, hydrophobic moieties compared to long flexible alkyl chains.24 To determine other elements involved in substrate binding that are specific for the hCBG β-glucosidase, the enzyme-substrate complex with quercetin-4′glucoside, a substrate for which wt-hCBG displays highest efficiency (Table 3), was modelled as described in Materials and Methods. This approach revealed that, due to the shape of the aglycone binding pocket, only a single principal orientation of the quercetin aglycone was likely to occur, since all other orientations led to unfavourable interactions with the hydrophobic regions of the binding pocket (Figure 3). Hence, it was relatively straightforward

Figure 3. Detailed view of the surface representation of hCBG, showing the position of the modelled quercetin4′-glucoside within the active site pocket. Residues responsible for the specific form of the aglycone binding pocket are highlighted as yellow sticks. This Figure was prepared using PyMol [http://pymol.sourceforge.net/].

Crystal Structure of Human Cytosolic β-Glucosidase

to identify residues that were likely to be involved in aglycone binding. The active site aglycone binding pocket of the hCBG 3D structure comprises a cluster of hydrophobic residues (Phe225, Val227, Tyr308, Tyr309, Phe334, Trp345 and Phe433) representing one wall of the binding pocket and Phe121, Val168, Val171, Met172, Leu176, Met178, Phe179, Phe186, Ile326 and Leu327 forming the opposite wall (Figure 2(b)). Surprisingly, in the model of the complex with quercetin-4′-glucoside, no polar or charged residue falls within a distance compatible with the formation of specific hydrogen bonds between the protein with secondary -OH groups of flavonol and flavone aglycones. This situation is encountered also with plant flavonoid glycosyltransferase,52 where the acceptor molecules have been observed to bind to a highly hydrophobic pocket. In both cases, the protein substrate interactions are dominated by a hydrophobic environment. While Phe121, Val168, Phe225, Trp345, Phe433 and Tyr309 (and to some extent Tyr308) form an inner hydrophobic cluster that surrounds the glycosidic linkage, Val171, Met172, Leu176, Phe179 and Phe334 represent a hydrophobic environment that is further away from the point of cleavage. Previous mutational studies suggested that Val168 was essential for binding of the aglycone moiety, whereas Phe225 and Tyr308 were involved in global catalysis of artificial aryl-glycosides and flavonoid glucosides.25 The present hCBG crystal structure is consistent with the earlier assertion concerning the role of Val168, and indicates that Tyr308 is involved in the overall structure of the binding pocket, whereas Phe225, by its central position within the aglycone pocket, is a key residue for aglycone recognition and specificity (Figure 2(b)). Met172 is unique to hCBG, whereas Phe179 and Phe433 residues are present also in the binding pocket of other GH1 β-glucosidases and are in the same physical position in hCBG. Residues equivalent to Trp345 are present in many GH1 enzymes, especially those that cleave aryl-conjugated glucosides; for example, the plant enzymes dhurrinase11 and ZmGlu1.47 In the human GH1 β-glucosidase LPH, hydrolysis of flavonoid glucosides was shown to occur for 75% at the lactase site (LPH4) and 25% at the phlorizin site (LPH3) of the enzyme.53 Trp345 is conserved in the LPH4 domain, but not in the LPH3, whereas Met172 is replaced by a Leu residue in the LPH3 domain. On the basis of these observations, another set of single mutants (Met172→Leu, Trp345→ Met, Trp345→Ala) was designed in order to further investigate the hCBG aglycone preference. The wildtype hCBG and single mutants were produced in P. pastoris and secreted into the culture medium. All proteins were purified from the culture supernatant using a single chromatographic step (octyl sepharose). The ability of the wild-type hCBG and hCBGmutants to hydrolyse several substrates (pNP-glycosides and flavonoid glucosides) and the associated kinetic parameters Km, kcat, and kcat/Km were determined. The Met172Leu mutant exhibited

Crystal Structure of Human Cytosolic β-Glucosidase

specific activities similar to the wild-type for all the substrates tested. All the other hCBG mutant enzymes exhibited drastic reductions in specific activity toward pNPαAra compared to the wildtype. The Trp345Met and Trp345Ala mutants exhibited a drastic decrease in specific activity towards pNP-glycosides and flavonoid-glucosides. For example, for pNPβGal, the specific activity of Met172Leu was 5.1 U mg−1 compared to 5.7 U mg−1 for the wildtype (Table 2), whereas Trp345Met and Trp345Ala, the specific activities were reduced by 100-fold and 2000-fold, respectively. The apparent affinity (Km) and kcat/Km values of Met172Leu for flavonoid glucosides were similar or slightly better than for wild-type. For example, the kcat/Km values of Met172Leu for naringenin-7-glucoside and eriodictyol-7-glucoside were 9.2 mM−1 s−1 and 8.1 mM−1 s−1, respectively, whereas with wt hCBG the values were 6.7 mM−1 s−1 and 6.0 mM−1 s−1, respectively (Table 3). The hCBG 3D structure revealed that Met172, most likely plays a role in the aglycone binding pocket by contributing to the hydrophobic character of the wall. It is thus not surprising that the Met172Leu mutation, which retains the hydrophobic character at this position, does not affect the specific activity of the enzyme. Trp345 is highly conserved among the GH1 β-glucosidases which hydrolyse bulky aromatic conjugated glycosides. The molecular docking together with the observed increases of Km for Trp345 mutants support the assignment of Trp345 as a determinant of enzyme affinity towards glucosides with an aromatic aglycone. Unexpectedly, our kinetic results demonstrate clearly that the dramatic drop in specificity constant is mirrored by a drop in kcat. On the basis of these novel observations, we conducted a further investigation of the 3D structure. This led us to propose that, since the amine group of the Trp345 indole ring is located near (∼3.9 Å) the O6 of the sugar and can act as a stacking platform in the aglycone pocket (Figure 2(a)), this residue may be a key in ensuring that the glucosidic bond is positioned in an orientation favourable for attack by the two catalytic residues Glu165 and Glu373, as observed also in plant GH1 β-glucosidases.39,47 Structural superimposition shows that it has a similar conformation in ZMGlu1, lining the active site in a way that makes stacking interactions with an aromatic aglycone possible (not shown). Taken together, the results indicate that Trp345 is an important component of the aglycone binding pocket. We have shown that the Val168→Tyr mutation did not affect Km on (pNP)-glycosides, but increased Km fivefold on flavonoid glucosides, providing the first biochemical evidence supporting a role for this residue in binding the aglycone portion of substrates. The present 3D structure, together with the molecular docking data (Figure 2(a) and (b)) support this recent assignment. Indeed, the position of Val168 in the active site pocket with respect to the modelled flavonoid aglycone is such that the replacement with tyrosine will form a pinch-like stacking arrangement with Trp345. This particular configuration most possibly is the explanation for the increase of Km of the Val168→Tyr mutation.

971 As described previously, the wt-hCBG is not capable of hydrolysing 3-conjugated flavonoid glucosides (quercetin-3-glucoside, quercetin-3-galactoside, quercetin-3-rhamnoside, and kaempferol-3glucoside).24 Further, previous alterations to various amino acids in the putative binding regions of hCBG through mutagenesis failed to generate an enzyme with activity towards 3-linked flavonoid glucosides.25 The arrangement of the hydrophobic residues forming the aglycone pocket is a likely explanation of this inability to hydrolyse 3-linked glycosides. Efforts to model the docking of flavonoids other than flavonoid-4′ or 7-glucosides into the binding pocket failed due to sterical hindrance that occurred when attempting to orient the aglycone into a compatible position with respect to the fixed glycone moiety. In particular, the relative positions of Phe179, Met172, Phe225 and Trp345 were incompatible with the binding of flavonoid-3-glucosides.

Conclusion In conclusion, by combining site-directed mutagenesis and crystallographic structure analyses we identified several crucial amino acid residues, Val168, Trp345 and Phe225, for aglycone binding and recognition in hCBG. Several additional residues of the hydrophobic cluster, such as Met172, Phe179, Phe334 or Phe433, may also affect substrate binding and further mutational studies will help elucidate the precise role of each of these residues. In addition, we showed that one amino acid difference in the hCBG active site can affect catalysis; the presence of Gln307, unique within GH1 β-glucosidases, conferred a higher level of specificity versus the glycosyl O2 position to hCBG, making mannose a good inhibitor for hCBG. Generally, specificity in GH1 enzymes is dominated by the aglycone of the substrate. In this respect, the hCBG crystal structure revealed that the particular arrangement of hydrophobic residues supported by the loops A through D that surround the rim of the active site are responsible for the shape of the aglycone binding pocket and that they are essential for flavonoid glucoside binding and catalysis. This assumption is confirmed by the site-directed mutagenesis and substrate–enzyme complex modelling. Furthermore, the reported human 3D structure will also help to understand the basis of the distinct and unusual specificity of other mammalian GH1 β-glucosidases.

Materials and Methods Preparation of wild-type hCBG for X-ray structure A large-scale expression of wt-hCBG was produced in P. pastoris as described.24 Purification of wt-hCBG was optimised by including an extra chromatographic step in the purification procedure used previously.24 The concentrated fraction was loaded onto an anion-exchange MonoQHR5/5column(Amersham,Buckinghamshire,UK)

972 previously equilibrated with 25 mM bis-Tris (pH 6.3). The bound material was eluted with a linear gradient of 0–25% (v/v) 25 mM bis-Tris (pH 6.3) containing 1 M NaCl; the flow-rate was 1 ml/min. The volume of each fraction was 0.5 ml with 0.5 ml of ethylene glycol added to stabilise the activity. The fractions containing β-glucosidase activity were pooled, checked for purity by SDSPAGE and concentrated using a Macrosep centrifugal tube (PALL, Portsmouth, UK) with Omega 3K ultrafiltration membrane. The Mono Q-concentrated-pool (A-C) (12.5 ml) was dialysed in 20 mM Mes buffer (pH 6.0) at 4 °C for 1 h, then concentrated using a Macrosep centrigugal tube. A total of 600 μg of ultrapure wt-hCBG was obtained at 300 μg/ml. Cloning and site-directed mutagenesis Mutations were introduced into the pHIL-S1/cbg-1 using the QuickChange® XL site-directed mutagenesis kit (Stratagene Europe) as described,25 using a pair of overlapping complementary oligonucleotides for each mutation designed to contain the corresponding nucleotide changes. The primers used in this study are given in Table 1. Transformation of the P. pastoris strain (his4)/ GS11554 and screening were carried out as described.24 Briefly, pHIL-S1/cbg-1 mutants (∼1 μg) as well as the pHIL-S1 vector (negative control) were digested with BgIII before transformation by the spheroplast method.55 After screening for methanol-sensitive colonies, Muts colonies were used to inoculate 10 ml of buffered minimal glycerol-complex medium (pH 6). After two days at 250 rpm and 30 °C, the cells were pelleted and resuspended in 2 ml of minimal methanol-complex medium. Following another four days at 30 °C, the culture was centrifuged and the amount of hCBG mutant proteins in the supernatant was estimated by activity measurement assays using pNPGlc and quercetin-4′-glucoside as substrates to select the best secreting clones. Human CBG mutant production and purification Selected transformants carrying the mutation were grown in 40 ml of minimal methanol-complex medium and incubated with shaking to induce the expression for five days in 500 ml flask at 30 °C. hCBG mutants were purified in a single step using an octyl Sepharose column.24 The protein concentrations of the mutants (M172L: 432 μg/ ml, Q307N: 320 μg/ml, W345M: 451 μg/ml, W345A: 380 μg/ml) were similar to that of the wild-type hCBG (311 μg/ml). The purified hCBG variants migrated in SDSPAGE as a single band with Mr = 53,000, identical with that of the recombinant wild-type enzyme. Enzyme activity assay The activity of hCBG mutants was assayed on glycosides (pNP-β-D-glucopyranoside, pNP-β-D-galactopyranoside, pNP-β-D-fucopyranoside, pNP-α– and pNP-β-Larabinopyranoside) using a spectrophotometric assay as described for the wild-type enzyme.24 Briefly, the release of 4-nitrophenol (4NP) from pNP-glycosides was monitored at 400 nm using the molar extinction coefficient for 4-nitrophenol of 18,300 M−1 cm−1, and enzyme reactions were carried out as described.24 Enzyme activities towards flavonoid glycosides were determined by measuring the amount of aglycone released from the substrate (10–500 μM in 50 mM sodium phosphate buffer), with

Crystal Structure of Human Cytosolic β-Glucosidase particular care taken to ensure complete solubility of substrates.56 Sequence analysis in silico Alignments and construction of the cladogram were carried out with the CLUSTALW program using sequences collected from CAZy and Swissprot databases.48 The tree was derived using the neighbor-joining method after aligning these sequences by a multiple sequence alignment program, CLUSTAL W version 1.60.57 Protein crystallization The protein solution (20 mM Mes, pH 6.0) was concentrated to about 4.0 mg/ml. Initial crystallization trials were performed with MDL (Molecular Dimensions Ltd) and Decode Genetics (“Wizard” 1&2, Emerald Biostructures) commercial kits. A total of 192 trials were tested in multi-well crystallization plates from Greiner, set up using a Cartesian crystallization robot, mixing 200 nl of protein with 100 nl of reservoir solution. All crystallization trials were carried out at 20 °C. Crystals grew within several days to an approximate size of 50 μm × 50 μm × 20 μm, in the crystallization trial containing 0.2 M ammonium sulphate, Mes buffer (pH 6.5) and 30% (w/v) polyethyleneglycol-monomethylether (PEG-MME) 5500. All attempts to optimize or reproduce the crystallization conditions were unsuccessful. The crystals obtained in the first place were therefore used directly for the data collection. Data collection and processing X-ray diffraction data of a hCBG crystal were collected at 100 K at the European Synchrotron Research Facilities (ESRF, Grenoble, France) beamline ID14 EH2 using an ADSC Quantum 4R CCD detector. The crystal was flashcooled in a stream of liquid-nitrogen with 10% (v/v) Table 5. Data collection and refinement statistics for the crystallographic analysis of the hCBG native data A. Data collection X-ray source Wavelength (Å) Resolution range (Å) Space group Rsym Completeness (%) Multiplicity I/σ(I) No. unique reflections B. Refinement statistics Resolution range (Å) Rcryst (%) Rfree (%) Overall B-factor (Å2) r.m.s.d. from ideality Bond lengths (Å) Bond angles (deg.) Ramachandran plot Most favoured regions (%) Additionally allowed regions (%)

hCBG ESRF ID14-EH2 0.933 45.0–2.70 P3121 0.18 (0.67)a 99.9 (100.0) 5.6 (5.7) 10.6 (3.1) 17,144 (2470) 42.6–2.7 19.7 28.0 38.04 0.022 1.94 82.5 16.5

Values in parentheses are for the highest resolution bin, 2.85– 2.70 Å.

Crystal Structure of Human Cytosolic β-Glucosidase

973

glycerol as cryoprotectant, using a nylon cryoloop of 0.1 mm diameter. The wavelength of the synchrotron X-rays was 0.933 Å. The crystal was rotated through 120° with a 0.5° oscillation range per frame. The raw data were processed using the program MOSFLM,58 and then merged and scaled using the program SCALA.59 Further data collection statistics are summarized in Table 5.

(QLK1-CT-51627), and M.C. was supported by a grant from the “Conseil Général” in France, Région Bretagne. We are indebted to the ESRF for beamtime allocation (BAG-MX485) for this project and to the staff on beamline ID14-EH2 for technical assistance during data collection.

Structure determination and refinement

Supplementary Data

The molecular replacement with the program AMoRe60 in the resolution range 9–3.1 Å with the model of dhurrinase (PDB 1V02) gave a solution with an overall correlation coefficient of 0.59 and an R-factor of 37.4% The residue changes corresponding to the hCBG primary sequence were performed using the program TURBO.61 Water molecules were added with wARP.62 Graphical inspection of the polypeptide chain and the water molecules was subsequently performed with TURBO. The refinement was performed using REFMAC5, part of the CCP4 package.59 The stereochemistry of the final structures was evaluated using PROCHECK.63 All refinement statistics are summarized in Table 5.

Supplementary data associated with this article can be found, in the online version, at doi:10.1016/ j.jmb.2007.05.034

Modelling of flavonoid-glucoside within the active site of hCBG The model of the position of the quercetin-glucoside substrate molecule within the active site of hCBG was obtained as follows: the coordinates of hCBG after the refinement procedure were superposed onto the 3D structure coordinates of dhurrinase in complex with dhurrin (PDB 1V03). The glucose moiety of the substrate molecule within the hCBG pocket is assumed to be positioned identically with dhurrin, since the residues binding this moiety are strictly conserved throughout the GH1 family. The coordinates of the structure of quercetin were extracted from the 3D structure of flavonoid glycosyl transferase in complex with quercetin.52 The molecule was then graphically positioned within the aglycone pocket of hCBG using the program TURBO, and using the position of the aglycone moiety of dhurrin as a guide. Since hCBG is most efficiently active on quercetin-4′-glucoside,24 the quercetin molecule was oriented with respect to the glucoside to form the glucoside 1-quercetin-4′ glycosidic bond. The final quercetin position was chosen such that all atoms were approximately centred with respect to the amino acid side-chains forming the walls of the aglycone pocket in hCBG at the same time as the 4′ carbon atom was maintained at the correct distance from O1 of the glucoside moiety to form the corresponding glycosidic bond. Protein Data Bank accession codes The coordinates of the crystallographic structure of hCBG have been deposited with the Protein Data Bank with ID codes 2JFE and R2JFESF from the atomic coordinates and the structure factors, respectively.

Acknowledgements S.T. was supported by a Marie Curie Individual Fellowship from the Framework V programme

References 1. Couthino, P. M. & Henrissat, B. (1999). Carbohydrateactive enzymes: an integrated database approach. In Recent Advances in Carbohydrate Bioengineering (Gilbert, H., Davies, G., Henrissat, B. & Svensson, B., eds), pp. 3–12, The Royal Chemistry Society, Cambridge, UK. 2. Schiffmann, R. & Brady, R. O. (2002). New prospects for the treatment of lysosomal storage diseases. Drugs, 62, 733–742. 3. Naim, H. Y. (2001). Molecular and cellular aspects and regulation of intestinal lactase-phlorizin hydrolase. Histol. Histopathol. 16, 553–561. 4. Barrett, T., Suresh, C. G., Tolley, S. P., Dodson, E. J. & Hughes, M. A. (1995). The crystal structure of a cyanogenic beta-glucosidase from white clover, a family 1 glycosyl hydrolase. Structure, 3, 951–960. 5. Burmeister, W. P., Cottaz, S., Driguez, H., Iori, R., Palmieri, S. & Henrissat, B. (1997). The crystal structures of Sinapis alba myrosinase and a covalent glycosyl-enzyme intermediate provide insights into the substrate recognition and active-site machinery of an S-glycosidase. Structure, 5, 663–675. 6. Chuenchor, W., Pengthaisong, S., Yuvaniyama, J., Opassiri, R., Svasti, J. & Ketudat, C., Jr (2006). Purification, crystallization and preliminary X-ray analysis of rice BGlu1 beta-glucosidase with and without 2-deoxy-2-fluoro-beta-D-glucoside. Acta Crystallog. sect. F, 62, 798–801. 7. Czjzek, M., Cicek, M., Zamboni, V., Burmeister, W. P., Bevan, D. R., Henrissat, B. & Esen, A. (2001). Crystal structure of a monocotyledon (maize ZMGlu1) betaglucosidase and a model of its complex with p-nitrophenyl beta-D-thioglucoside. Biochem. J. 354, 37–46. 8. Husebye, H., Arzt, S., Burmeister, W. P., Hartel, F. V., Brandt, A., Rossiter, J. T. & Bones, A. M. (2005). Crystal structure at 1.1 angstroms resolution of an insect myrosinase from Brevicoryne brassicae shows its close relationship to beta-glucosidases. Insect Biochem. Mol. Biol. 35, 1311–1320. 9. Ruppert, M., Panjikar, S., Barleben, L. & Stockigt, J. (2006). Heterologous expression, purification, crystallization and preliminary X-ray analysis of raucaffricine glucosidase, a plant enzyme specifically involved in Rauvolfia alkaloid biosynthesis. Acta Crystallog. sect. F, 62, 257–260. 10. Sue, M., Yamazaki, K., Yajima, S., Nomura, T., Matsukawa, T., Iwamura, H. & Miyamoto, T. (2006). Molecular and structural characterization of hexameric beta-D-glucosidases in wheat and rye. Plant Physiol. 141, 1237–1247.

Crystal Structure of Human Cytosolic β-Glucosidase

974 11. Verdoucq, L., Moriniere, J., Bevan, D. R., Esen, A., Vasella, A., Henrissat, B. & Czjze, M. (2004). Structural determinants of substrate specificity in family 1 betaglucosidases: novel insights from the crystal structure of sorghum dhurrinase-1, a plant beta-glucosidase with strict specificity, in complex with its natural substrate. J. Biol. Chem. 279, 31796–31803. 12. Zouhar, J., Vevodova, J., Marek, J., Damborsky, J., Su, X. D. & Brzobohaty, B. (2001). Insights into the functional architecture of the catalytic center of a maize beta-glucosidase Zm-p60.1. Plant Physiol. 127, 973–985. 13. Guasch, A., Vallmitjana, M., Perez, R., Querol, E., Perez-Pons, J. A. & Coll, M. (1999). Cloning, overexpression, crystallization and preliminary X-ray analysis of a family 1 beta–glucosidase from Streptomyces. Acta Crystallog. sect. D, 55, 679–682. 14. Hakulinen, N., Paavilainen, S., Korpela, T. & Rouvinen, J. (2000). The crystal structure of beta-glucosidase from Bacillus circulans sp. alkalophilus: ability to form long polymeric assemblies. J. Struct. Biol. 129, 69–79. 15. Sanz-Aparicio, J., Hermoso, J. A., Martinez-Ripoll, M., Lequerica, J. L. & Polaina, J. (1998). Crystal structure of beta-glucosidase A from Bacillus polymyxa: insights into the catalytic activity in family 1 glycosyl hydrolases. J. Mol. Biol. 275, 491–502. 16. Wang, X., He, X., Yang, S., An, X., Chang, W. & Liang, D. (2003). Structural basis for thermostability of betaglycosidase from the thermophilic eubacterium Thermus nonproteolyticus HG102. J. Bacteriol. 185, 4248–4255. 17. Wiesmann, C., Beste, G., Hengstenberg, W. & Schulz, G. E. (1995). The three-dimensional structure of 6-phospho-beta-galactosidase from Lactococcus lactis. Structure, 3, 961–968. 18. Zechel, D. L., Boraston, A. B., Gloster, T., Boraston, C. M., Macdonald, J. M., Tilbrook, D. M. et al. (2003). Iminosugar glycosidase inhibitors: structural and thermodynamic dissection of the binding of isofagomine and 1-deoxynojirimycin to beta-glucosidases. J. Am. Chem. Soc. 125, 14313–14323. 19. Aguilar, C. F., Sanderson, I., Moracci, M., Ciaramella, M., Nucci, R., Rossi, M. & Pearl, L. H. (1997). Crystal structure of the beta-glycosidase from the hyperthermophilic archeon Sulfolobus solfataricus: resilience as a key factor in thermostability. J. Mol. Biol. 271, 789–802. 20. Akiba, T., Nishio, M., Matsui, I. & Harata, K. (2004). X-ray structure of a membrane-bound beta-glycosidase from the hyperthermophilic archaeon Pyrococcus horikoshii. Proteins, 57, 422–431. 21. Chi, Y. I., Martinez-Cruz, L. A., Jancarik, J., Swanson, R. V., Robertson, D. E. & Kim, S. H. (1999). Crystal structure of the beta-glycosidase from the hyperthermophile Thermosphaera aggregans: insights into its activity and thermostability. FEBS Letters, 445, 375–383. 22. Kaper, T., Lebbink, J. H., Pouwels, J., Kopp, J., Schulz, G. E., van der, O. J. & de Vos, W. M. (2000). Comparative structural analysis and substrate specificity engineering of the hyperthermostable beta-glucosidase CelB from Pyrococcus furiosus. Biochemistry, 39, 4963–4970. 23. Jenkins, J., Lo, L. L., Harris, G. & Pickersgill, R. (1995). Beta-glucosidase, beta-galactosidase, family A cellulases, family F xylanases and two barley glycanases form a superfamily of enzymes with 8-fold beta/ alpha architecture and with two conserved glutamates near the carboxy-terminal ends of beta-strands four and seven. FEBS Letters, 362, 281–285. 24. Berrin, J. G., McLauchlan, W. R., Needs, P., William-

25.

26.

27.

28.

29.

30.

31.

32.

33. 34.

35.

36.

37.

38. 39.

son, G., Puigserver, A., Kroon, P. A. & Juge, N. (2002). Functional expression of human liver cytosolic betaglucosidase in Pichia pastoris. Insights into its role in the metabolism of dietary glucosides. Eur. J. Biochem. 269, 249–258. Berrin, J. G., Czjzek, M., Kroon, P. A., McLauchlan, W. R., Puigserver, A., Williamson, G. & Juge, N. (2003). Substrate (aglycone) specificity of human cytosolic beta-glucosidase. Biochem. J. 373, 41–48. Nemeth, K., Plumb, G. W., Berrin, J. G., Juge, N., Jacob, R., Naim, H. Y. et al. (2003). Deglycosylation by small intestinal epithelial cell beta-glucosidases is a critical step in the absorption and metabolism of dietary flavonoid glycosides in humans. Eur. J. Nutr. 42, 29–42. Matsumura, Y., Aizawa, H., Shiraki-Iida, T., Nagai, R., Kuro-o, M. & Nabeshima, Y. (1998). Identification of the human klotho gene and its two transcripts encoding membrane and secreted klotho protein. Biochem. Biophys. Res. Commun. 242, 626–630. Shiraki-Iida, T., Aizawa, H., Matsumura, Y., Sekine, S., Iida, A., Anazawa, H. et al. (1998). Structure of the mouse klotho gene and its two transcripts encoding membrane and secreted protein. FEBS Letters, 424, 6–10. Yahata, K., Mori, K., Arai, H., Koide, S., Ogawa, Y., Mukoyama, M. et al. (2000). Molecular cloning and expression of a novel klotho-related protein. J. Mol. Med. 78, 389–394. Ito, S., Kinoshita, S., Shiraishi, N., Nakagawa, S., Sekine, S., Fujimori, T. & Nabeshima, Y. I. (2000). Molecular cloning and expression analyses of mouse betaklotho, which encodes a novel Klotho family protein. Mech. Dev. 98, 115–119. Ito, S., Fujimori, T., Hayashizaki, Y. & Nabeshima, Y. (2002). Identification of a novel mouse membranebound family 1 glycosidase-like protein, which carries an atypical active site structure. Biochim. Biophys. Acta, 1576, 341–345. Mantei, N., Villa, M., Enzler, T., Wacker, H., Boll, W., James, P. et al. (1988). Complete primary structure of human and rabbit lactase-phlorizin hydrolase: implications for biosynthesis, membrane anchoring and evolution of the enzyme. EMBO J. 7, 2705–2713. Nabeshima, Y. (2006). Toward a better understanding of Klotho. Sci. Aging Knowledge. Environ. 2006, e11. Tohyama, O., Imura, A., Iwano, A., Freund, J. N., Henrissat, B., Fujimori, T. & Nabeshima, Y. (2004). Klotho is a novel beta-glucuronidase capable of hydrolyzing steroid beta-glucuronides. J. Biol. Chem. 279, 9777–9784. Esen, A. & Gungor, G. (1993). Stability and activity of plant and fungal beta-glucosidases under denaturing conditions. In β-glucosidases: Biochemistry and Molecular Biology (Esen, A., ed.), pp. 214–239, American Chemical Society (ACS), Washington, DC. Daniels, L. B., Coyle, P. J., Chiao, Y. B., Glew, R. H. & Labow, R. S. (1981). Purification and characterization of a cytosolic broad specificity beta-glucosidase from human liver. J. Biol. Chem. 256, 13004–13013. Burmeister, W. P., Cottaz, S., Rollin, P., Vasella, A. & Henrissat, B. (2000). High resolution X-ray crystallography shows that ascorbate is a cofactor for myrosinase and substitutes for the function of the catalytic base. J. Biol. Chem. 275, 39385–39393. Gloster, T. M., Madsen, R. & Davies, G. J. (2006). Dissection of conformationally restricted inhibitors binding to a beta-glucosidase. ChemBiochem, 7, 738–742. Verdoucq, L., Czjzek, M., Moriniere, J., Bevan, D. R. & Esen, A. (2003). Mutational and structural analysis of

Crystal Structure of Human Cytosolic β-Glucosidase

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

aglycone specificity in maize and sorghum betaglucosidases. J. Biol. Chem. 278, 25055–25062. Gloster, T. M., Macdonald, J. M., Tarling, C. A., Stick, R. V., Withers, S. G. & Davies, G. J. (2004). Structural, thermodynamic, and kinetic analyses of tetrahydrooxazine-derived inhibitors bound to beta-glucosidases. J. Biol. Chem. 279, 49236–49242. White, A., Tull, D., Johns, K., Withers, S. G. & Rose, D. R. (1996). Crystallographic observation of a covalent catalytic intermediate in a beta-glycosidase. Nature Struct. Biol. 3, 149–154. Withers, S. G., Warren, R. A. J., Street, I. P., Rupitz, K., Kempton, J. B. & Aebersold, R. (1990). Unequivocal demonstration of the involvement of a glutamate residue as a nucleophile in the mechanism of a retaining glycosidase. J. Am. Chem. Soc. 112, 5887–5889. Withers, S. G., Rupitz, K., Trimbur, D. & Warren, R. A. (1992). Mechanistic consequences of mutation of the active site nucleophile Glu 358 in Agrobacterium betaglucosidase. Biochemistry, 31, 9979–9985. Witt, E., Frank, R. & Hengstenberg, W. (1993). 6-Phospho-beta-galactosidases of gram-positive and 6-phospho-beta-glucosidase B of Gram-negative bacteria: comparison of structure and function by kinetic and immunological methods and mutagenesis of the lacG gene of Staphylococcus aureus. Protein Eng. 6, 913–920. Moracci, M., Capalbo, L., Ciaramella, M. & Rossi, M. (1996). Identification of two glutamic acid residues essential for catalysis in the beta-glycosidase from the thermoacidophilic archaeon Sulfolobus solfataricus. Protein Eng. 9, 1191–1195. Vallmitjana, M., Ferrer-Navarro, M., Planell, R., Abel, M., Ausin, C., Querol, E. et al. (2001). Mechanism of the family 1 beta-glucosidase from Streptomyces sp: catalytic residues and kinetic studies. Biochemistry, 40, 5975–5982. Czjzek, M., Cicek, M., Zamboni, V., Bevan, D. R., Henrissat, B. & Esen, A. (2000). The mechanism of substrate (aglycone) specificity in beta -glucosidases is revealed by crystal structures of mutant maize beta -glucosidase-DIMBOA, -DIMBOAGlc, and -dhurrin complexes. Proc. Natl Acad. Sci. USA, 97, 13555–13560. Thompson, J. D., Higgins, D. G. & Gibson, T. J. (1994). CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucl. Acids Res. 22, 4673–4680. Henrissat, B., Callebaut, I., Fabrega, S., Lehn, P., Mornon, J. P. & Davies, G. (1995). Conserved catalytic machinery and the prediction of a common fold for several families of glycosyl hydrolases. Proc. Natl Acad. Sci. USA, 92, 7090–7094. Coutinho, P. M., Stam, M., Blanc, E. & Henrissat, B. (2003). Why are there so many carbohydrate-active

975

51.

52.

53.

54. 55. 56.

57. 58. 59. 60. 61. 62.

63. 64. 65.

enzyme-related genes in plants? Trends Plant Sci. 8, 563–565. Glew, R. H., Gopalan, V., Forsyth, G. W. & Vanderjagt, D. J. (1993). The mammalian cytosolic broad-specificity beta-glucosidase. In β-glucosidases: Biochemistry and Molecular Biology (Esen, A., ed.), pp. 83–112, American Chemical Society (ACS), Washington, DC. Offen, W., Martinez-Fleites, C., Yang, M., Kiat-Lim, E., Davis, B. G., Tarling, C. A. et al. (2006). Structure of a flavonoid glucosyltransferase reveals the basis for plant natural product modification. EMBO J. 25, 1396–1405. Day, A. J., Canada, F. J., Diaz, J. C., Kroon, P. A., Mclauchlan, R., Faulds, C. B. et al. (2000). Dietary flavonoid and isoflavone glycosides are hydrolysed by the lactase site of lactase phlorizin hydrolase. FEBS Letters, 468, 166–170. Cregg, J. M., Barringer, K. J., Hessler, A. Y. & Madden, K. R. (1985). Pichia pastoris as a host system for transformations. Mol. Cell Biol. 5, 3376–3385. Hinnen, A., Hicks, J. B. & Fink, G. R. (1978). Transformation of yeast. Proc. Natl Acad. Sci. USA, 75, 1929–1933. Lambert, N., Kroon, P. A., Faulds, C. B., Plumb, G. W., McLauchlan, W. R., Day, A. J. & Williamson, G. (1999). Purification of cytosolic beta-glucosidase from pig liver and its reactivity towards flavonoid glycosides. Biochim. Biophys. Acta, 1435, 110–116. Saitou, N. & Nei, M. (1987). The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol. Biol. Evol. 4, 406–425. Leslie, A. (1990). Mosflm. In Crystallographic Computing, pp. 50–61, Oxford University Press, New York. Collaborative Computing Project Number 4. (1994). The CCP4 suite: programs for protein crystallography. Acta Crystallog. sect. D, 50, 760–763. Navaza, J. (1994). AMoRe—an automated package for molecular replacement. Acta Crystallog. sect. A, 50, 157–163. Roussel, A. & Cambillau, C. (1991). TURBO-FRODO. In Silicon Graphics Geometry Partner Directory, pp. 86–87, Silicon Graphics, Mountain View, CA. Perrakis, A., Sixma, T. K., Wilson, K. S. & Lamzin, V. S. (1997). wARP: improvement and extension of crystallographic phases by weighted averaging of multiplerefined dummy atomic models. Acta Crystallog. sect. D, 53, 448–455. Laskowski, R. A., Moss, D. S. & Thornton, J. M. (1993). Main-chain bond lengths and bond angles in protein structures. J. Mol. Biol. 231, 1049–1067. Kraulis, P. J. (1991). MOLSCRIPT—a program to produce both detailed and schematic plots of protein structures. J. Appl. Crystallog. 24, 946–950. Merritt, E. A. & Murphy, M. E. (1994). Raster3D version 2.0. A program for photorealistic molecular graphics. Acta Crystallog. sect. D, 50, 869–873.

Edited by M. Guss (Received 6 February 2007; received in revised form 27 April 2007; accepted 12 May 2007) Available online 18 May 2007