Title: Phosphorylase recognition and phosphorolysis of its oligosaccharide substrate: answers to a long outstanding question
Abstract: Article1 September 1999free access Phosphorylase recognition and phosphorolysis of its oligosaccharide substrate: answers to a long outstanding question K.A. Watson Corresponding Author K.A. Watson Laboratory of Molecular Biophysics, Department of Biochemistry, University of Oxford, South Parks Road, Oxford, OX1 3QU GB Search for more papers by this author C. McCleverty C. McCleverty Present address: Department of Biochemistry, University of Leicester, Leicester, GB Search for more papers by this author S. Geremia S. Geremia Department of Chemistry, University of Trieste, Trieste, Italy Search for more papers by this author S. Cottaz S. Cottaz Centre de Reserche sur les Macromolécules Végétales (CERMAV-CNRS), BP 53, F-38041 Grenoble, cedex 9, France Search for more papers by this author H. Driguez H. Driguez Centre de Reserche sur les Macromolécules Végétales (CERMAV-CNRS), BP 53, F-38041 Grenoble, cedex 9, France Search for more papers by this author L.N. Johnson L.N. Johnson Oxford Centre for Molecular Sciences, New Chemistry Laboratory, University of Oxford, South Parks Road, Oxford, OX1 3QT GB Search for more papers by this author K.A. Watson Corresponding Author K.A. Watson Laboratory of Molecular Biophysics, Department of Biochemistry, University of Oxford, South Parks Road, Oxford, OX1 3QU GB Search for more papers by this author C. McCleverty C. McCleverty Present address: Department of Biochemistry, University of Leicester, Leicester, GB Search for more papers by this author S. Geremia S. Geremia Department of Chemistry, University of Trieste, Trieste, Italy Search for more papers by this author S. Cottaz S. Cottaz Centre de Reserche sur les Macromolécules Végétales (CERMAV-CNRS), BP 53, F-38041 Grenoble, cedex 9, France Search for more papers by this author H. Driguez H. Driguez Centre de Reserche sur les Macromolécules Végétales (CERMAV-CNRS), BP 53, F-38041 Grenoble, cedex 9, France Search for more papers by this author L.N. Johnson L.N. Johnson Oxford Centre for Molecular Sciences, New Chemistry Laboratory, University of Oxford, South Parks Road, Oxford, OX1 3QT GB Search for more papers by this author Author Information K.A. Watson 1, C. McCleverty2, S. Geremia3, S. Cottaz4, H. Driguez4 and L.N. Johnson5 1Laboratory of Molecular Biophysics, Department of Biochemistry, University of Oxford, South Parks Road, Oxford, OX1 3QU GB 2Present address: Department of Biochemistry, University of Leicester, Leicester, GB 3Department of Chemistry, University of Trieste, Trieste, Italy 4Centre de Reserche sur les Macromolécules Végétales (CERMAV-CNRS), BP 53, F-38041 Grenoble, cedex 9, France 5Oxford Centre for Molecular Sciences, New Chemistry Laboratory, University of Oxford, South Parks Road, Oxford, OX1 3QT GB *Corresponding author. E-mail: [email protected] The EMBO Journal (1999)18:4619-4632https://doi.org/10.1093/emboj/18.17.4619 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Phosphorylases are key enzymes of carbohydrate metabolism. Structural studies have provided explanations for almost all features of control and substrate recognition of phosphorylase but one question remains unanswered. How does phosphorylase recognize and cleave an oligosaccharide substrate? To answer this question we turned to the Escherichia coli maltodextrin phosphorylase (MalP), a non-regulatory phosphorylase that shares similar kinetic and catalytic properties with the mammalian glycogen phosphorylase. The crystal structures of three MalP–oligosaccharide complexes are reported: the binary complex of MalP with the natural substrate, maltopentaose (G5); the binary complex with the thio-oligosaccharide, 4-S-α-D-glucopyranosyl-4-thiomaltotetraose (GSG4), both at 2.9 Å resolution; and the 2.1 Å resolution ternary complex of MalP with thio-oligosaccharide and phosphate (GSG4-P). The results show a pentasaccharide bound across the catalytic site of MalP with sugars occupying sub-sites −1 to +4. Binding of GSG4 is identical to the natural pentasaccharide, indicating that the inactive thio compound is a close mimic of the natural substrate. The ternary MalP–GSG4-P complex shows the phosphate group poised to attack the glycosidic bond and promote phosphorolysis. In all three complexes the pentasaccharide exhibits an altered conformation across sub-sites −1 and +1, the site of catalysis, from the preferred conformation for α(1–4)-linked glucosyl polymers. Introduction Structural, biochemical and time-resolved studies on rabbit muscle glycogen phosphorylase have contributed to an understanding of the catalytic mechanism whereby the enzyme promotes the phosphorolysis of an α(1–4) glycosidic bond in glycogen or oligosaccharide substrates. The mechanisms of control by phosphorylation and by allosteric effectors are also understood from crystallographic studies on phosphorylated and unphosphorylated forms of the mammalian phosphorylase (Barford and Johnson, 1989; Barford et al., 1991; Sprang et al., 1991; Johnson, 1992). One question has remained outstanding. This relates to the recognition and conformation of the oligosaccharide component of the substrate when it is bound across the catalytic site. Numerous attempts to observe binding of oligosaccharide substrates with rabbit muscle glycogen phosphorylase (GP) in the active R state were not successful, probably because of the low affinity of the enzyme for linear oligosaccharides. In order to solve this problem we have turned to the Escherichia coli maltodextrin phosphorylase (MalP). As suggested by the nomenclature, GP and MalP exhibit distinct differences in their preference for substrates. GP has a high affinity for glycogen and branched polysaccharides and low affinity for small oligosaccharides (Hu and Gold, 1975; Kasvinsky et al., 1978). MalP has low affinity for glycogen but exhibits ∼100-fold higher affinity for linear oligosaccharides than GP (Schwartz and Hofnung, 1967; Drueckes et al., 1996). This observation has been exploited to solve the major outstanding problem in phosphorylase catalysis, namely the recognition of the oligosaccharide component of the substrate at the catalytic site. MalP (EC.2.4.1.1) is a 796 amino acid, non-allosteric, dimeric enzyme that is 46% identical in sequence to GP (Palm et al., 1985, 1987). Unlike GP, MalP is not regulated by phosphorylation or by allosteric effectors but is regulated by control of gene expression (Raibaud and Schwartz, 1984). Phosphorylases are highly conserved from bacterial to mammalian enzymes, but share no homology with other carbohydrate degrading enzymes, with the exception of the noted similarity of the core structure around the cofactor pyridoxal phosphate binding site in GP with a DNA glucosyltransferase modifying enzyme (Artymuick et al., 1995; Holm and Sander, 1995). Phosphorylase isozymes exhibit nearly 100% identity in residues at the catalytic site (Watson et al., 1997) and all utilize the 5′-phosphate of the essential cofactor pyridoxal phosphate (PLP) in catalysis (Palm et al., 1990). The bacterial and mammalian enzymes catalyse the phosphorolytic cleavage from the non-reducing end of α-1–4 linked maltodextrins (Glc)n to yield α-D-glucose-1-P (Glc-1-P) (Figure 1). The reaction proceeds with retention of configuration. For MalP, the optimal substrate maltodextrin has a minimum length of five glucosyl units for phosphorolysis and four glucosyl units for the reverse reaction of oligosaccharide synthesis (Becker et al., 1994). Figure 1.The reaction catalysed by MalP and the chemical structure of 4-S-α-D-glucopyranosyl-4-thiomaltotetraose (GSG4). Download figure Download PowerPoint Structure determinations of MalP (Watson et al., 1997) and GP have shown that the catalytic site is buried at the centre of the large subunit. It is accessible to the solvent through a 20 Å long channel that has been proposed as the substrate oligosaccharide binding site. Sub-sites for oligosaccharide recognition are numbered −1 to +4 from the non-reducing end to the reducing end, and the site of phosphorolysis is between sub-sites −1 and +1 (Davies et al., 1995). In both the less active T state and the active R state conformations of GP, the catalytic site (sub-site −1) has been defined through binding studies and observations of catalysis in the crystal with monomeric substrates and products such as Glc-1-P and heptulose-2-P, and by binding studies with inhibitors such as glucose, glucose analogues and transition state analogues (Hajdu et al., 1987; Johnson et al., 1989, 1990, 1992; Martin et al., 1990; Duke et al., 1994; Mitchell et al., 1996). The site is situated at the end of the channel, far removed from the bulk solvent and adjacent to the essential cofactor, pyridoxal phosphate. Co-crystallization studies with MalP and maltohexaose (O'Reilly et al., 1997) showed a disaccharide, maltose, bound at adjacent sites to the catalytic site (numbered sub-sites +1 and +2). Analysis of the oligosaccharide mixture from the co-crystallization experiments indicated that both catalysis and hydrolysis of the oligosaccharide had occurred over the period of crystallization. α(1–4)-O-linked glucosyl oligosaccharides exhibit a preferred conformation in which there is an intramolecular hydrogen bond between the O2 hydroxyl and the O3 hydroxyl of contiguous residues, as observed in maltose (Takusagawa and Jacobson, 1978) and related compounds. Modelling studies from the MalP–maltose complex and from the GP–monosaccharide complexes indicated that an oligosaccharide bound across sub-sites −1 and +1 could not be accommodated in the preferred conformation if the oligosaccharide were to follow the path of the catalytic channel (O'Reilly et al., 1997). Either there must be a significant conformational change in the enzyme structure, or a significant alteration of the conformation about the α(1,4) glycosidic bond between sub-sites −1 and +1. It was therefore of interest to observe an oligosaccharide substrate bound across the phosphorylase catalytic site and to assess the significance of any conformational change for catalysis. Acarbose is a pseudo-tetrasaccharide inhibitor of α-glucosidases that can adopt either the preferred conformation or an alternative conformation about the corresponding N-linked glycosidic bond of its acarviosine moiety. Co-crystallization of MalP with acarbose led to oligosaccharide binding in sub-sites +1 to +4 and the identification of the further sub-sites +3 and +4 (O'Reilly et al., 1999). However, in both the MalP–maltose and the MalP–acarbose binding experiments a glycerol molecule, used as a cryoprotectant for the low temperature data collection, blocked the crucial catalytic sub-site −1. Thus the problem of observing an oligosaccharide bound across the phosphorolytic cleavage site remained. In the present work, we developed a method of freezing crystals that avoided the use of glycerol. By carefully excluding phosphate from the purification and crystallization protocols we have been able to co-crystallize the binary MalP complex with the natural substrate maltopentaose (G5). The X-ray crystal structure of the MalP–G5 complex shows for the first time an oligosaccharide bound across the catalytic site in sub-sites −1 to +4 spanning the point of enzymatic cleavage. In order to observe a ternary complex in the presence of phosphate and elucidate the catalytic mechanism, we have exploited the thio-oligosaccharide, 4-S-α-D-glucopyranosyl-4-thiomaltotetraose (GSG4) which is resistant to enzymatic cleavage (Figure 1). Thio-oligosaccharides (Driguez, 1997) have become useful tools for the study of carbohydrate–enzyme interactions (Sulzenbacher et al., 1996). Solution NMR studies (Bock et al., 1994) indicated that the thio analogues showed greater conformational flexibility than the corresponding O-linked and N-linked glucosyl polymers. Comparative studies with the thio and natural pentasaccharide binary complexes showed no significant differences in binding to MalP. Co-crystallization of MalP with the thio-oligosaccharide in the presence of phosphate has allowed us to determine the structure of a ternary enzyme substrate complex. The complex is consistent with previous site-directed mutagenesis data for MalP and provides further understanding of the catalytic mechanism of phosphorylases. Results The overall architecture of the native maltodextrin phosphorylase has been described, and details of the structure compared with the active and inactive forms of mammalian phosphorylase (Watson et al., 1997). In brief, MalP is an α/β protein consisting of two domains (N-terminal domain 19–482; C-terminal domain 483–829, using the rabbit muscle GP numbering) that form the monomeric subunit. The active site lies between these domains, at the centre of the molecule, and is accessible to the solvent via a channel ∼20 Å in length. There is a non-crystallographic 2-fold axis relating two subunits, giving rise to a dimer in the asymmetric unit. The overall quaternary and tertiary protein structures of the three oligosaccharide complexes reported here are similar to the native MalP structure. However there are significant local conformational changes from the native MalP structure in a loop of residues at the catalytic site upon binding the oligosaccharide. The structures of the MalP–G5, MalP–GSG4 and MalP–GSG4-P complexes A summary of the data processing statistics and refinement parameters for each of the three MalP–oligosaccharide complex structures is given in Table I. Views of the final 2Fo−Fc weighted electron density maps for the three complexes are shown in Figure 2. In each of the complexes the five sugars are clearly defined in sub-sites −1 to +4. Sub-site −1 is the furthest buried in the catalytic site channel and is adjacent to the cofactor PLP. The oligosaccharide binds the length of the channel with sub-site +4 most exposed to the bulk solvent. Figure 2.Electron density maps with weighted 2Fo−Fc coefficients (see text for details of weighting) for (A) MalP–G5 complex, (B) MalP–GSG4 complex and (C) MalP–GSG4-P complex. Contour levels are 1σ corresponding to 0.19 e/Å3, 0.24 Å3 and 0.24 e/Å3 for the G5, GSG4 and GSG4-P maps, respectively. The fifth sugar at sub-site +4 is just out of view. Download figure Download PowerPoint Table 1. Summary of the data collection and refinement statistics for the three complexes Data collection MalP–G5 MalP–GSG4 MalP–GSG4-P Resolution range 50.0–2.9 50.0–2.9 50.0–2.0 No. of measurements used 74 406 55 829 152 192 No. of unique reflections 38 437 38 856 72 681 Completeness (%) All data 79.1 84.5 80.0 Last resolution shell (3.1–2.9 Å) 75.1 (3.1–2.9 Å) 71.5 (2.1–2.0 Å) 64.5 (3.7–3.4 Å) 81.0 (2.4–2.2 Å) 81.3 Multiplicity 2.2 1.7 2.1 Rmergea 0.18 0.075 0.102 Space group P212121 P212121 P21 Cell dimensions (Å, °) a = 74.45 a = 74.74 a = 107.41 b = 105.69 b = 105.02 b = 61.50 c = 214.91 c = 217.96 c = 132.38 β = 105.03 Final refinement parameters No. of reflections usedb 29 646 32 407 69 025 No. of protein atoms 6753 6753 6753 No. of solvent atoms 190 299 377 No. of cofactor atoms 15 15 15 No. of ligand atoms 56 56 61 Rconv/Rfreec 0.245/0.306 0.214/0.277 0.228/0.261 R.m.s. deviation from ideal geometry for the protein Bond lengths (Å) 0.008 0.004 0.004 Bond angles (°) 1.33 0.822 0.911 Planar groups (°) 0.876 0.739 0.492 Average B factors (Å2) Main chain 46.3 19.7 23.7 Side chain 47.9 21.5 25.6 Solvent 65.9 41.5 38.3 Oligosaccharide 58.1 20.4 22.9 Phosphate – – 60.1 R.m.s. deviation on B factors Bond B restraints 5.1 3.9 2.5 Angle B restraints 7.5 5.7 3.8 a , where Ih,i and Ih are the i measurement and mean measurement of reflection h, respectively, and the sum is over all reflections for which more than one measurement is recorded. b This is the number of reflections used in the calculation of Rconv. Rfree, calculated for 5% of the data selected at random at the beginning of the structure refinement. c , where Fobs and Fcalc are the observed and calculated structure factor amplitudes, respectively. Comparison of the isomorphous crystal structures of the MalP–G5 and MalP–GSG4 complexes at 2.9 Å resolution showed that the thio analogue bound in a similar manner to the natural pentasaccharide. The single crystal structure of thio-maltoside (Perez and Vergelati, 1984) demonstrated that the thio analogue has similar structural parameters to those observed for the O-linked maltose with an increase in the distance C1 to C4′ of 0.35 Å. In the MalP–G5 complex, the C1…C4′ distance is 2.4 Å, compared with 2.9 Å in the MalP–GSG4 complex showing a similar small increase. The binary structures are shown superimposed in Figure 4. The root mean square deviation (r.m.s.d.) in Cα coordinates of the protein atoms is 0.3 Å and the r.m.s.d. between the G5 and GSG4 oligosaccharide coordinates is 0.55 Å. The thio-oligosaccharide (GSG4) is a good mimic of the natural substrate both in its conformation and its interactions with the enzyme. Figure 3.Superposition of the MalP–G5 and MalP–GSG4 complexes. The two structures are closely similar. Download figure Download PowerPoint Crystals of the ternary complex MalP–GSG4-P were obtained under crystallization conditions similar to those for the binary complex of MalP–GSG4, with the addition of 5 mM phosphate to the protein drop. The crystals belong to the monoclinic space group P21, a space group that is different from the orthorhombic P212121 space group of the MalP–G5 and MalP–GSG4 complexes. The crystals diffracted to 2 Å resolution but there was some disorder in the direction parallel to the thin edge of the plate-like crystal. Using MOSFLM we were able to extract data to 2.0 Å resolution by using a dynamic resolution cut-off applied to individual frames. All the data were used in the refinement but electron density maps were calculated to 2.2 Å resolution, where the data are more complete (Table I). The five sugars were immediately apparent in difference maps obtained after refinement of the protein atoms and each sugar could be fitted unambiguously, since all sugar hydroxyl groups could be clearly located. After refinement of the protein and GSG4 atoms, the position of the phosphate group was apparent as the highest peak in the difference Fourier synthesis. The final refinement gave an overall B factor of 60 Å2 for the phosphate group. This is higher than expected for a group that makes several contacts to the protein and to the sugars at sub-sites −1 and +1. The position of the phosphate ion is nearly identical (overall shift of 0.8 Å) to the position observed for phosphate in the native MalP structure (Watson et al., 1997) where the B factors were 34 Å2. A water molecule modelled at the phosphate site in the MalP–GSG4-P complex did not refine well. It is concluded that the phosphate is present but less well ordered than the other atoms in the structure, possibly as a result of its proximity to the thio linkage of the oligosaccharide (discussed below). The overall structure of the MalP–GSG4-P complex was very similar to the MalP native structure (r.m.s.d. in Cα coordinates 0.8 Å), where the major difference in structure results from a movement in the 380s loop (discussed below) and, to the MalP–GSG4 complex (r.m.s.d. in Cα coordinates 0.3 Å). The only difference between the MalP–GSG4 and MalP–GSG4-P complexes is the presence of a water molecule in the former complex that is replaced by the phosphate in the latter MalP–GSG4-P complex, and an accompanying 4.5 Å shift in the position of Arg569. The binding of GSG4 is identical in the binary and ternary complexes. A summary of the intermolecular contacts for the MalP–GSG4-P complex is given in Table II and shown in Figure 5. The sugar in sub-site −1, buried close to the cofactor, has the largest number of hydrogen bonds and van der Waals contacts with the enzyme. These contacts are similar to those observed for monomeric glucosyl residues binding to the mammalian GP at sub-site −1. Each of the peripheral hydroxyl groups is involved in at least two hydrogen bonds and there are numerous van der Waals contacts, especially to His377. The other sites involve fewer hydrogen bonds but almost all of these involve hydrogen bonds between the sugars and charged residues (Glu88 and Asp339 at subsite +1; Arg292 and His341 at sub-site +2; Arg292 at sub-site +3; and Glu382 and His571 at sub-site +4). The charged–polar hydrogen bonds might be expected to contribute more binding energy than polar–polar hydrogen bonds (Fersht et al., 1985; Street et al., 1986). The van der Waals contacts at sub-sites +2 and +4 are dominated by glucosyl–tyrosyl interactions (to Tyr280 and Tyr613, respectively) as previously noted (O'Reilly et al., 1997, 1999). The contacts from Tyr280 to the sugar involve the C2, C4, O5 and C6 atoms of sub-site +2. The contacts from Tyr613 to the sugar in sub-site +4 also involve the C2, C4, O5, C6 and O3 atoms of the sugar. Both aromatic groups stack with the A side of the glucopyranosyl ring as expected for α(1,4)-linked glucosyl polymers (Johnson et al., 1988). In addition, there are contacts from Tyr613 to C1, O5 and O6 of the sugar in sub-site +3. Hence the contacts from Tyr613 span the two sub-sites +3 and +4. Finally, we note that sub-site +3 appears extraordinarily wet. There are eight water molecules in van der Waals contact with the sugar, although only four of these are hydrogen bonded to the sugar. Figure 4.Summary of the intermolecular contacts for the MalP–GSG4-P complex. Contacts to Arg569 are slightly obscured by Tyr573 in this view. Intramolecular hydrogen bonds between adjacent sugars are not shown. Sugars labelled sub-sites +4 to −1 correspond to Glc994–Glc998, respectively. Full details are given in Table II. Download figure Download PowerPoint Table 2. Intermolecular hydrogen bonds in the MalP–GSG4-P complex Sub-site Oligosaccharide atom Protein atom Distance (Å) Sub-site −1 998 O2 Tyr573 OH 2.9 Pho900 O1 2.8 Wat1122 2.9 Wat1277 2.6 O3 Glu672 OE2 2.6 Ser674 N 3.3 Gly675 N 3.2 O4 Asn484 ND2 3.3 Gly675 N 3.1 Wat1085 2.7 O6 His377 ND1 2.8 Asn484 ND2 2.7 Sub-site +1 997 O2 Asp339 OD2 2.4 O3 Asp339 OD2 2.8 [S4 Pho900 O1 2.9] O5 Glu88 OE2 2.8 O6 Glu88 OE2 2.9 Gly134 N 3.3 Leu136 N 2.9 Sub-site S+2 996 O2 Arg292 NH2 2.9 Wat1162 2.9 O3 His341 NE2 2.9 Wat1162 2.8 O5 Wat1297 2.9 O6 Asn133 ND2 2.7 Arg569 NH1 2.7 Sub-site S+3 995 O2 Wat1160 2.7 O3 Arg292 NH2 3.2 Wat1205 2.7 Wat1261 2.8 O6 Wat1298 2.8 Arg569 NH1 3.3 Sub-site S+4 994 O1 Wat1166 2.7 O5 Wat1354 3.1 O6 Glu382 OE1 2.5 Glu382 OE2 3.3 His571 NE2 2.9 Phosphate 900 O1 Arg569 NH2 2.7 Tyr573 OH 2.8 Lys574 NZ 3.2 Glc998 O2 2.8 [Glc997 S4 2.9] O2 Lys574 NZ 2.9 PLP OP1 2.7 PLP OP2 3.0 O3 Arg569 NE 2.9 Lys574 NZ 2.8 PLP999 OP1 3.0 Wat1265 2.1 O4 Gly135 N 2.7 [Glc997 S4 2.8] Due to the high B factors associated with the phosphate group, we are less certain of the phosphate oxygen positions than for the carbohydrate atoms. Nevertheless, the dynamic refinement has resulted in stereochemically plausible positions that are consistent with the X-ray evidence. The MalP–phosphate contacts are dominated by contacts to an arginine residue, Arg569, where two phosphate oxygens contact the NE and NH2 atoms of the guanidine side chain in a classic binding mode (Johnson and O'Reilly, 1996) (Table II; Figure 5). One of these oxygen atoms also contacts Lys574. A third oxygen is hydrogen bonded to the main chain nitrogen of Gly135, a residue from a glycine-rich loop at the start of an α-helix (Acharya et al., 1991). There is a direct hydrogen bond between one of the oxygens (O2) of the inorganic phosphate and an oxygen (OP1) of the cofactor phosphate and a further phosphate–phosphate contact through water. The cofactor 5′-phosphate is localized by hydrogen bonds from two of its oxygens to two lysine residues (Lys568 and Lys574), while the third oxygen is hydrogen bonded to the main chain nitrogen of a residue at the start of an α-helix. Each of the 5′-phosphate oxygens is also hydrogen bonded to water molecules. This arrangement is very similar to that observed with phosphate bound to the native MalP structure and also to the mammalian enzyme. Finally, in the ternary MalP–GSG4-P complex there are contacts from the inorganic phosphate oxygens (O1 and O4) to the thio-glycosidic linkage [distances 2.9 and 2.8 Å to the S atom, respectively (Table II)]. In the two binary complexes of MalP with G5 and GSG4, where there is no phosphate present, Arg569 adopts an alternative conformation in which it is turned away from the substrate phosphate binding site. The shift in the arginine residue seen on binding phosphate provides additional support for the presence of the phosphate in the ternary complex. Table 3. van der Waals contacts to sugars and phosphate in MalP–GSG4-P complexa Sub-site van der Waals contacts [residue (number contacts)] Total contacts Sub-site −1 Gly135(3), Leu136(4), His377(15), Thr378(4), Asn484(4), 61 998 Val455(3), Tyr573(1), Glu672(4), Ala673(2), Ser674(1), Gly675(4), Pho900(9), Wat1085(3), Wat1122(2), Wat1277(2) Sub-site +1 Glu88(6), Gly134(6), Gly135(3), Leu136(6), Gly137(1), 49 997 Tyr280(3), Asp339(4), His341(1), His377(2), Thr378(3), Arg569(6), Pho900(6), Wat1122(2) Sub-site S+2 Asn133(6), Tyr280(6), Arg292(3), His341(1), Glu382(4), 33 996 Arg569(7), Wat1205(1), Wat1130(1), Wat1162(2), Wat1297(2) Sub-site S+3 Asp283(1), Glu382(4), His571(2), Arg569(2), Ala610(2), 30 995 Tyr613(8), Wat1009(2), Wat1037(1), Wat1160(2), Wat1190(1), Wat1205(2), Wat1233(1), Wat1261(2), Wat1298(1) Sub-site S+4 Glu382(8), His571(3), Gly612(1), Tyr613(10), Wat1166(4), 31 994 Wat1209(2), Wat1354(3) Phosphate Gly134(1), Gly135(2), Arg569(12), Tyr573(1), Lys574(4), 43 900 Glu672(2), Glc997(4), Glc998(9), PLP999(2), Wat1085(2), Wat1122(1), Wat1265(3) avan der Waals contacts <4.0 Å excluding hydrogen bonds listed in Table IIA. Mutagenesis experiments supported Extensive site-directed mutagenesis experiments have been carried out with E.coli MalP in which the target residues were identified by predicting the likely binding site for an oligosaccharide based on the rabbit muscle GP structure (Drueckes et al., 1996; O'Reilly et al., 1997). The mutagenesis studies have yielded results that are fully consistent with the present direct experimental observations of an oligosaccharide binding to MalP. The mutations Glu88Ala (sub-site +1), Tyr280Ala (sub-site +1, +2), Asp339Ala (sub-site +1), His341Ala (sub-site +1, +2), Thr378Gly (sub-site −1, +1), Glu382Ala (sub-site +2, +3, +4) and His571Leu (sub-site +3, +4) result in mutant enzymes that exhibit significantly reduced activity. Each of these residues is involved in direct contact with the oligosaccharide (Table II). The results indicate the importance of each of the interactions in locating and directing the oligosaccharide for correct binding for catalysis. The mutant Tyr613Phe showed only slightly changed kinetic parameters, a result that can be rationalized from the structure since phenylalanine at position 613 could perform similar stacking interactions with the sugars in sub-sites +3 and +4. The triple mutant Asn282Ala, Asp283Ala, Asn284Ala also had only slightly changed kinetic properties. The present results show that these residues from the 280s loop play only a minor role in binding oligosaccharide. Conformational changes The most significant conformational change in the MalP structure on binding oligosaccharide involves residues 377–384, a region designated as the 380s loop. In the T state (less active) form of GP this loop helps close the catalytic site through hydrophobic and ion pair interactions. In the R state (active) form of GP this loop is displaced from the catalytic site. The movement of the 380s loop in the T–R transition is correlated with the movement of Arg569, the important substrate phosphate binding residue. It has been suggested that the changes in the 380s loop in the mammalian GP provide a means of communication between the catalytic site and the glycogen storage site (Barford and Johnson, 1992; O'Reilly et al., 1997). MalP is constitutively active, and in the native MalP structure, Arg569 is already in its correct position to bind the phosphate substrate, since the crystals were grown from a phosphate-buffered solution (Watson et al., 1997). There is no conserved glycogen storage site in MalP, which provides an explanation for the preference of linear maltodextrin substrates over large branched glycogen substrates. The simplicity of the bacterial enzyme in binding a linear oligosaccharide, allows direct observation of the role of the 380s loop. In MalP the conformational change of this loop results in closure of the catalytic site, which is important both for recognition and correct positioning of the oligosaccharide in the catalytic site (Figure 6). The changes in the pentasaccharide complexes are similar to those observed previously for the maltose complex (O'Reilly et al., 1997), where maltose bound to sub-site +1 and +2. In contrast, in the MalP–acarbose complex where acarbose bound to sub-sites +1 to +4, no closure of the 380s loop is observed. The lack of movement can be rationalized in terms of the presence of the cyclitol ring, and not a glucopyranosyl ring, in sub-site +1 (O'Reilly et al., 1999). Acarbose, although a potent in