Insights into the Mechanism of Drosophila melanogaster Golgi α-Mannosidase II through the Structural Analysis of Covalent Reaction Intermediates
2003; Elsevier BV; Volume: 278; Issue: 48 Linguagem: Inglês
10.1074/jbc.m309249200
ISSN1083-351X
AutoresShin Numao, D.A. Kuntz, Stephen G. Withers, David R. Rose,
Tópico(s)Enzyme Production and Characterization
ResumoThe family 38 golgi α-mannosidase II, thought to cleave mannosidic bonds through a double displacement mechanism involving a reaction intermediate, is a clinically important enzyme involved in glycoprotein processing. The structure of three different covalent glycosyl-enzyme intermediates have been determined to 1.2-Å resolution for the Golgi α-mannosidase II from Drosophila melanogaster by use of fluorinated sugar analogues, both with the wild-type enzyme and a mutant enzyme in which the acid/base catalyst has been removed. All these structures reveal sugar intermediates bound in a distorted 1S5 skew boat conformation. The similarity of this conformation with that of the substrate in the recently determined structure of the Michaelis complex of a β-mannanase (Ducros, V. M. A., Zechel, D. L., Murshudov, G. N., Gilbert, H. J., Szabo, L., Stoll, D., Withers, S. G., and Davies, G. J. (2002) Angew. Chem. Int. Ed. Engl. 41, 2824–2827) suggests that these disparate enzymes have recruited common stereoelectronic features in evolving their catalytic mechanisms. The family 38 golgi α-mannosidase II, thought to cleave mannosidic bonds through a double displacement mechanism involving a reaction intermediate, is a clinically important enzyme involved in glycoprotein processing. The structure of three different covalent glycosyl-enzyme intermediates have been determined to 1.2-Å resolution for the Golgi α-mannosidase II from Drosophila melanogaster by use of fluorinated sugar analogues, both with the wild-type enzyme and a mutant enzyme in which the acid/base catalyst has been removed. All these structures reveal sugar intermediates bound in a distorted 1S5 skew boat conformation. The similarity of this conformation with that of the substrate in the recently determined structure of the Michaelis complex of a β-mannanase (Ducros, V. M. A., Zechel, D. L., Murshudov, G. N., Gilbert, H. J., Szabo, L., Stoll, D., Withers, S. G., and Davies, G. J. (2002) Angew. Chem. Int. Ed. Engl. 41, 2824–2827) suggests that these disparate enzymes have recruited common stereoelectronic features in evolving their catalytic mechanisms. N-Linked glycosylation is a post-translational modification found in eukaryotes whereby a complex oligosaccharide is attached to a surface asparagine residue within the sequence Asn-X-(Ser/Thr) through an N-glycosidic linkage. The initial part of the pathway, occurring in the endoplasmic reticulum, is highly conserved and has been well characterized in various organisms (1Burda P. Aebi M. Biochim. Biophys. Acta. 1999; 1426: 239-257Crossref PubMed Scopus (528) Google Scholar, 2Herscovics A. Biochim. Biophys. Acta. 1999; 1473: 96-107Crossref PubMed Scopus (239) Google Scholar, 3Kornfeld R. Kornfeld S. Annu. Rev. Biochem. 1985; 54: 631-664Crossref PubMed Scopus (3770) Google Scholar). The latter Golgi part is species-, tissue-, and cell-dependent and is the source of the diverse nature of glycosylation. In various tumor cell lines such as those from breast, colon, and skin cancer, the distribution of the cell surface N-linked sugars is altered (4Couldrey C. Green J.E. Breast Cancer Res. 2000; 2: 321-323Crossref PubMed Scopus (86) Google Scholar). Because N-linked glycosylation is associated with cell-cell interactions, this alteration correlates with the progression of tumor metastasis (5Dennis J.W. Granovsky M. Warren C.E. Biochim. Biophys. Acta. 1999; 1473: 21-34Crossref PubMed Scopus (605) Google Scholar, 6Granovsky M. Fata J. Pawling J. Muller W.J. Khokha R. Dennis J.W. Nat. Med. 2000; 6: 306-312Crossref PubMed Scopus (468) Google Scholar). Enzymes of this glycosylation pathway are therefore potential targets for the development of inhibitors as cancer treatments. Golgi α-mannosidase II (GMII 1The abbreviations used are: GMIIGolgi α-mannosidase IIDNP-Man2,4-dinitrophenyl α-d-mannosideDNP2,4-dinitrophenol5FGulF5-fluoro-β-l-gulosyl fluoridedGMIIDrosophila melanogaster GMII2FManF2-deoxy-2-fluoro-α-d-mannosyl fluoridewtwild-typeMES4-morpholineethanesulfonic acidr.m.s.d.root mean square deviationCGTasecyclodextrin glucanotransferase.1The abbreviations used are: GMIIGolgi α-mannosidase IIDNP-Man2,4-dinitrophenyl α-d-mannosideDNP2,4-dinitrophenol5FGulF5-fluoro-β-l-gulosyl fluoridedGMIIDrosophila melanogaster GMII2FManF2-deoxy-2-fluoro-α-d-mannosyl fluoridewtwild-typeMES4-morpholineethanesulfonic acidr.m.s.d.root mean square deviationCGTasecyclodextrin glucanotransferase.; mannosyl-oligosaccharide 1,3–1,6-α-mannosidase II; EC 3.2.1.114), a glycosidase involved in trimming of two mannose residues in the Golgi, has been the target of one such drug candidate. In early clinical trials, the compound swainsonine, a well known GMII inhibitor, was found to reduce tumor growth and metastasis when taken orally (7Goss P.E. Baker M.A. Carver J.P. Dennis J.W. Clin. Cancer Res. 1995; 1: 935-944PubMed Google Scholar, 8Goss P.E. Reid C.L. Bailey D. Dennis J.W. Clin. Cancer Res. 1997; 3: 1077-1086PubMed Google Scholar). Cross-reactivity of swainsonine with lysosomal mannosidase limits its clinical usefulness, and, therefore, a detailed understanding of the mechanism of GMII is necessary to develop drug candidates that specifically target this glycosidase. Golgi α-mannosidase II 2,4-dinitrophenyl α-d-mannoside 2,4-dinitrophenol 5-fluoro-β-l-gulosyl fluoride Drosophila melanogaster GMII 2-deoxy-2-fluoro-α-d-mannosyl fluoride wild-type 4-morpholineethanesulfonic acid root mean square deviation cyclodextrin glucanotransferase. Golgi α-mannosidase II 2,4-dinitrophenyl α-d-mannoside 2,4-dinitrophenol 5-fluoro-β-l-gulosyl fluoride Drosophila melanogaster GMII 2-deoxy-2-fluoro-α-d-mannosyl fluoride wild-type 4-morpholineethanesulfonic acid root mean square deviation cyclodextrin glucanotransferase. From the primary structure, GMII has been classified into the glycosyl hydrolase family 38 (9Henrissat B. Biochem. J. 1991; 280: 309-316Crossref PubMed Scopus (2615) Google Scholar, 10Henrissat B. Bairoch A. Biochem. J. 1993; 293: 781-788Crossref PubMed Scopus (1768) Google Scholar, 11Henrissat B. Biochem. Soc. Trans. 1998; 26: 153-156Crossref PubMed Scopus (151) Google Scholar). Members of this family of glycosidases hydrolyze α-mannosides with net retention of configuration at the anomeric center (12Howard S. Braun C. McCarter J. Moremen K.W. Liao Y.F. Withers S.G. Biochem. Biophys. Res. Commun. 1997; 238: 896-898Crossref PubMed Scopus (39) Google Scholar). By analogy with other retaining glycosidases, these enzymes are postulated to catalyze reactions by a double displacement mechanism (13Koshland D.E. Biol. Rev. 1953; 28: 416-436Crossref Scopus (805) Google Scholar, 14Sinnott M.L. Chem. Rev. 1990; 90: 1171-1202Crossref Scopus (1490) Google Scholar, 15Zechel D.L. Withers S.G. Acc. Chem. Res. 2000; 33: 11-18Crossref PubMed Google Scholar). In this mechanism, a covalent glycosyl-enzyme intermediate is first formed via an oxocarbenium ion-like transition state (glycosylation). A carboxylic acid in the active site acts as the catalytic nucleophile in the formation of this intermediate, assisted by a second carboxylic acid that acts as a general acid catalyst. Hydrolysis of the glycosyl-enzyme intermediate (deglycosylation) occurs in a second step, facilitated by general base catalysis provided by the conjugate base of the acid catalyst from the first step. By using a mechanism-based inactivator, 5-fluoro-β-l-gulosyl fluoride (5FGulF), the covalent glycosyl-enzyme intermediate formed on two representative members of this family has been trapped, and the residue acting as the catalytic nucleophile was identified in each case (16Howard S. He S.M. Withers S.G. J. Biol. Chem. 1998; 273: 2067-2072Abstract Full Text Full Text PDF PubMed Scopus (84) Google Scholar, 17Numao S. He S.M. Evjen G. Howard S. Tollersrud O.K. Withers S.G. FEBS Lett. 2000; 484: 175-178Crossref PubMed Scopus (29) Google Scholar). The structure of the catalytic portion of the GMII from Drosophila melanogaster (dGMII) was determined by x-ray crystallography along with structures of complexes with several ligands, namely swainsonine and deoxymannojirimycin (18van den Elsen J.M.H. Kuntz D.A. Rose D.R. EMBO J. 2001; 20: 3008-3017Crossref PubMed Scopus (183) Google Scholar). The ligand binding site features coordination to a zinc ion and includes a conserved aspartate residue, Asp-204 (dGMII numbering), analogous to the residue identified in the previous trapping studies as the catalytic nucleophile. These results further suggested that the acid/base catalyst is most likely Asp-341. These structures, however, only give a first insight into the structural aspects of the catalytic mechanism. A key element of the catalytic mechanism is the glycosyl-enzyme intermediate formed during catalysis. Determination of the structure of such intermediates has provided more detailed insights, not only into the residues involved in catalysis, but also into the possible role of saccharide distortion along the reaction pathway, as discussed recently (19Davies G. Ducros V.M. Varrot A. Zechel DL. Biochem. Soc. Trans. 2003; 31: 523-527Crossref PubMed Google Scholar), primarily for β-glycosidases. Recent studies of β-glycosidases have revealed substrate distortion in Michaelis (enzyme-substrate) complexes but not in α-glycosyl-enzyme intermediates (with the exception of the family 11 xylanase (20Sidhu G. Withers S.G. Nguyen N.T. McIntosh L.P. Ziser L. Brayer G.D. Biochemistry. 1999; 38: 5346-5354Crossref PubMed Scopus (171) Google Scholar) and of the family 26 β-mannanase (21Ducros V.M.A. Zechel D.L. Murshudov G.N. Gilbert H.J. Szabo L. Stoll D. Withers S.G. Davies G.J. Angew. Chem. Int. Ed. Engl. 2002; 41: 2824-2827Crossref PubMed Scopus (121) Google Scholar)). Until now, no structure of trapped intermediates on a true α-glycosidase has yet been published, though the structure of the β-glycosyl-enzyme intermediate on a family 13 transglycosidase, the cyclodextrin glucanotransferase from Bacillus circulans was reported (22Uitdehaag J.C. Mosi R. Kalk K.H. van der Veen B.A. Dijkhuizen L. Withers S.G. Dijkstra B.W. Nat. Struct. Biol. 1999; 6: 432-436Crossref PubMed Scopus (366) Google Scholar). In this paper we have used both 5FGulF and 2-deoxy-2-fluoro-α-d-mannosyl fluoride (2FManF) to trap the glycosylenzyme intermediates formed on both the wild-type (wt) and D341N mutant of dGMII and examined the crystallographic structures of these intermediates. These are the first structures of covalent intermediates determined for any α-mannosidase. Furthermore, this is the first true α-glycoside hydrolase for which the structure of the covalent intermediate has been determined. As such, these studies add to our understanding of how α-glycosidases catalyze hydrolysis of glycosidic linkages. The 2,4-dinitrophenyl α-d-mannoside (DNP-Man) (23Sharma S.K. Corrales G. Penades S. Tetrahedron Lett. 1995; 36: 5627-5630Crossref Google Scholar), 5FGulF (16Howard S. He S.M. Withers S.G. J. Biol. Chem. 1998; 273: 2067-2072Abstract Full Text Full Text PDF PubMed Scopus (84) Google Scholar), and 2FManF (24Hall L.D. Johnson R.N. Adamson J. Foster A.B. Can. J. Chem. 1971; 49: 118-123Crossref Google Scholar) were synthesized according to published procedures. All chemicals and buffer salts were obtained from Sigma unless noted otherwise. To facilitate mutagenesis, the expression plasmid for dGMII (18van den Elsen J.M.H. Kuntz D.A. Rose D.R. EMBO J. 2001; 20: 3008-3017Crossref PubMed Scopus (183) Google Scholar), which contains a metallothioneine promoter and Bip-secretion sequence, was first reduced in size from 6642 to 4610 bp by SalI digestion followed by re-ligation. The single codon mutation was introduced to this plasmid using the QuikChange site-directed mutagenesis kit (Stratagene). The forward primer had the sequence CTG CTG ATT CCG TTG GGT GAC AAC TTC CGC TTC AAG C. The presence of the mutation was confirmed by sequencing. A 238-bp AdeI/partial SalI fragment, which contained the mutated sequence, was then excised from the plasmid and inserted into an AdeI/partial SalI-digested native expression plasmid. The presence of the mutation and the absence of any frameshifts or secondary mutations were confirmed by sequencing the final construct. This D341N construct and a blasticidin resistance plasmid were then used to co-transfect Drosophila cells. The presence of the secreted, full-length protein in the medium from transfected cells induced with 10 μm cadmium was confirmed by Western blot analysis. Anti-pentaHis antibody (Qiagen) was used as the primary antibody whereas an alkaline phosphatase-conjugated anti-mouse antibody was used as the secondary antibody. Alkaline phosphatase activity was monitored with NBT/BCIP reagent (Sigma). Stable populations of transfectants were selected using blasticidin (16 μg/ml). Single cell clones expressing the highest levels of protein were chosen by Western blot analysis for use in large scale expression. These cells were grown up and adapted to serum-free medium (ExCell 420; JRH Biosciences). Growth of stable S2 cells expressing the D341N mutant dGMII and purification of the mutant protein was similar to the procedure described for the wt enzyme (18van den Elsen J.M.H. Kuntz D.A. Rose D.R. EMBO J. 2001; 20: 3008-3017Crossref PubMed Scopus (183) Google Scholar). Steady State Kinetics—All studies of the dGMII were carried out at 37 °C in 50 mm MES buffer, pH 5.6, containing 0.1 mm ZnSO4 and 0.1% bovine serum albumin unless specified otherwise. Determination of kinetic parameters for the hydrolysis of DNP-Man by wt and D341N mutant dGMII were performed by following the increase in the absorbance at 400 nm upon the addition of enzyme (typically 3 and 30 nm final concentration of enzyme, respectively) at varying substrate concentrations (typically 10–12 points). Measurements were made in 1-cm-path length quartz cuvettes with a UNICAM UV4 UV-visible spectrophotometer attached to a circulating water bath or a Varian CARY 4000 spectrophotometer attached to a temperature control unit. The kinetic parameters were estimated by direct fit of the data to the Michaelis-Menten equation using the program GraFit 4.0.14 (25Leatherbarrow R.J. GraFit, Version 4.0.14. Erithacus Software Limited, Surrey, UK2001Google Scholar). Approximate Ki values for both 5FGulF and 2FManF with the mutant enzyme were determined by measuring the rate of hydrolysis of DNP-Man (0.1 mm) at a number of varying inhibitor concentrations. The data were plotted in the form of a Dixon plot (1/rate versus [inhibitor]) and then fit to a line using the program GraFit 4.0.14. The Ki value was determined from the intersection of this line and the line for y = 1/Vmax, assuming competitive inhibition. The kinetic parameters for the hydrolysis of 5FGulF and 2FManF by wt dGMII were determined by measuring the increase in fluoride ion concentration upon the addition of enzyme in the appropriate buffer (typically 10 and 20 nm enzyme final enzyme concentrations, respectively) at varying "substrate" concentrations (typically 6–8 points). Measurements were made using an ORION 96–04 combination fluoride electrode interfaced to a personal computer running the program LoggerPro (Vernier Software). In the case of 2FManF, saturation was not seen, and as such, the data collected at low substrate concentrations were fit to a line using linear regression to determine the kcat/Km value. Inactivation Studies—The inactivation of the D341N mutant dGMII by 5FGulF was monitored by incubating the enzyme (1.2 mg ml–1) under the above conditions in the presence of various concentrations of 5FGulF at 16 °C. Residual activities were determined at various time intervals by adding 10-μl aliquots of the inactivation mixture to a solution of DNP-Man (1.5 mm, 110 μl) in the above buffer and measuring DNP release for 30 s at 16 °C. Pseudo-first order inactivation rate constants were determined by fitting the data for the residual activity versus time to a first order rate equation using the program GraFit 4.0.14 (25Leatherbarrow R.J. GraFit, Version 4.0.14. Erithacus Software Limited, Surrey, UK2001Google Scholar). The ki/Ki value was extracted from the slope of the plot of kobs against 5FGulF concentration. The inactivation of the wt dGMII by 5FGulF was observed in the same manner except that the enzyme concentration was lower (0.22 mg ml–1), and the temperature of the experiment was reduced to 8 °C. The analyses of protein samples were carried out using a Sciex API-300 mass spectrometer interfaced with an LC-Packings high pressure liquid chromatography system. Briefly, the intact protein (10–20 μg) was pre-incubated for 5 min with either 4 mm 5FGulF (wt or D341N mutant dGMII) or 40 mm 2FManF (D341N mutant dGMII). This sample was introduced into the mass spectrometer through a microbore PRLP column (1 × 50 mm) and eluted with a gradient of 20–100% Solvent B (0.05% trifluoroacetic acid/90% CH3CN in water) in Solvent A (0.06% trifluoroacetic acid/2% CH3CN in water) at a flow rate of 50 μl/min over 10 min. The mass spectrum was scanned over a range of 600 to 2400 Da (4 s/scan) with a step size of 0.5 Da. The ion source voltage was 4.8 kV, and the orifice energy was 50 V. Crystallization of dGMII was carried out using hanging drop vapor diffusion as described previously (18van den Elsen J.M.H. Kuntz D.A. Rose D.R. EMBO J. 2001; 20: 3008-3017Crossref PubMed Scopus (183) Google Scholar). Because of a relatively rapid loss of diffraction quality with time, crystals were less than 24 h old at the time of crystal evaluation and freezing. For the 2FManF and native protein complexes, the crystals were grown in the absence of inhibitor and then soaked with the compound for ∼30 min. Prior to freezing, the crystals were passed through drops containing 10, 15, 20, and 25% methyl-pentanediol in crystallization buffer and 5 mm of appropriate compound. The crystals were mounted in nylon CryoLoops (Hampton Research) and frozen directly in a liquid nitrogen cryostream. All data were collected at 100 K. Data were collected either at the Ontario Cancer Institute on a MAR Research 2300 image plate detector mounted on a rotating anode generator with copper target, operated at 50 kV and 100 mA with beam focusing using Osmic optics, or at the Cornell High Energy Synchrotron Source, beamline F1, using an ADSC Quantum 4 CCD detector in the rapid readout mode. Typically 300–400 frames of 0.5° oscillation were collected for each data set. Data reduction and scaling were carried out using Denzo and Scalepack, respectively (26Otwinowski Z. Minor W. Methods Enzymol. 1997; 276: 307-326Crossref PubMed Scopus (38520) Google Scholar). Data collection statistics are provided in Table I.Table IStatistics for data collection and refinementdGMII compoundwt-5FGulFD341N-2FManFD341N-5FGulFProtein Data Bank code1QWN1QX11QWUCrystal(P212121)(P212121)(P212121) Cell dimensions (Å)69.02/110.02/138.9669.05/109.83/138.9168.85/109.80/138.77 Mosaicity0.1250.430.398Data collection X-ray sourceCHESS-F1CHESS-F1OCI/rotating anode Wavelength (Å)0.95040.95041.54Data processing (hi res shell) Resolution Å30-1.20 (1.22-1.20)30-1.3 (1.32-1.3)20-2.03 (2.08-2.03) Reflections (total/unique)2,449,245/324,6051,589,410/246,051476,418/68,989 I/σ23.7 (3.4)14.7 (2.9)16.8 (2.9) % completeness98.8 (91.4)95.3 (89.7)99.9 (98.7) Rmerge0.075 (0.429)0.106 (0.586)0.11 (0.45)Refinement Rtest/Rfree (reflections for Rfree)0.175/0.192 (2225)0.169/0.188 (2180)0.151/0.189 (2336) Amino acids101410141014 Alternate conformations4030 Water molecules103510421025 r.m.s.d. bonds (Å)0.020.020.015 r.m.s.d. angles (°)1.972.021.72 Average B-factor (Å2)Overall12.8416.0216.92Protein main chain10.2313.0214.78Protein side chain12.2016.3716.69Water22.1925.9025.97Covalent ligand7.7514.3310.35 Open table in a new tab The software program CNS (27Brunger A.T. Adams P.D. Acc. Chem. Res. 2002; 35: 404-412Crossref PubMed Scopus (53) Google Scholar, 28Brunger A.T. Adams P.D. Clore G.M. DeLano W.L. Gros P. Grosse-Kunstleve R.W. Jiang J.S. Kuszewski J. Nilges M. Pannu N.S. Read R.J. Rice L.M. Simonson T. Warren G.L. Acta Crystallogr. 1998; 54: 905-921Crossref PubMed Scopus (8) Google Scholar) was used for the refinement of the structures of the complexes. For the calculation of Rfree, a test set comprising ∼2000 reflections was removed from the data set. The structures of the complexes were solved by molecular replacement. Briefly, rigid body refinement was carried out against the published structure of native dGMII (Protein Data Bank code 1HTY) with Tris and water molecules in the region of the active site removed (18van den Elsen J.M.H. Kuntz D.A. Rose D.R. EMBO J. 2001; 20: 3008-3017Crossref PubMed Scopus (183) Google Scholar). Asp-204 was changed to alanine in the Protein Data Bank file to remove bias around the region of formation of the covalent bond. Rigid body refinement was followed by simulated annealing to 3500 K, group B-factor refinement, and individual B-factor refinement, prior to generation of electron density maps. At this initial stage R-factors were typically in the range of 22%, and the Fo – Fc density maps clearly showed the presence of bound compound and unassigned waters. Refinement of the model involving manually fitting the compounds into the density, fitting waters, and checking side-chains for proper fit to the density was carried out with O (29Jones T.A. Zhou J.-Y. Cowan S.W. Kjeldgaard M. Acta Crystallogr. 1991; 47: 110-119Crossref PubMed Scopus (13009) Google Scholar), with intermittent rounds of energy minimization using CNS. Automated water-picking and deletion of weak waters were carried out using the automated routines from CNS, and the picked waters were checked manually. Once all waters and side chains were fit another round of simulated annealing and B-factor refinement was performed. Clear alternate conformations of side chains were then inserted, followed by individual B-factor refinement. Model correctness was checked with The MolProbity server (kinemage.biochem.duke.edu/molprobity) and the WhatIf server (www.biotech.ebi.ac.uk:8400/). Protein overlays and r.m.s.d. calculations were carried out using ProFit (www.bioinf.org.uk/software/profit). Graphics were generated using Pymol (30DeLano W.L. Pymol. DeLano Scientific, San Carlos, CA2002Google Scholar). Protein Data Bank (31Berman H.M. Westbrook J. Feng Z. Gilliland G. Bhat T.N. Weissig H. Shindyalov I.N. Bourne P.E. Nucleic Acids Res. 2000; 28: 235-242Crossref PubMed Scopus (27305) Google Scholar) codes for the deposited coordinates and diffraction data are given in Table I. Kinetic Analysis—The kinetic parameters for the activated substrate, DNP-Man, were determined for both the wt and D341N mutant enzyme. As might be expected from removal of the acid/base catalyst, the kcat value for the D341N mutant dGMII was lower by roughly 200-fold compared with that of the wt enzyme (kcat values are 0.048 and 8.6 s–1, respectively). The Km value for the mutant enzyme was also 2 orders of magnitude lower compared with the Km value of the wt enzyme (Km value are 0.05 and 5.5 mm, respectively). This lowering of the Km value is unlikely to be because of a tightening of the true binding, but is more likely because of an accumulation of the intermediate species as a result of the deglycosylation step becoming rate-limiting. This behavior is also reflected in the fact that the kcat/Km value, which reflects the first irreversible step, most likely glycosylation, is largely unaltered. This is reasonable, because a good leaving group such as DNP requires very little proton assistance for departure, thus removal of the acid catalyst will not significantly affect the glycosylation step. However, removal of general base catalysis clearly slows down the second step, deglycosylation. Thus the intermediate accumulates. Nonetheless, turnover of the intermediate species was unfortunately sufficiently rapid to prevent us from observing the intermediate in crystals of the D341N mutant dGMII soaked with DNP-Man. When 5FGulF was incubated with the D341N mutant dGMII at 37 °C, the compound was found to act as an apparent reversible inhibitor with an approximate Ki value of 0.6 mm. Considering that the wt enzyme has a Km value of 5 mm for DNP-Man, 5FGulF binds considerably more tightly than might be expected for a very simple substrate analogue, especially considering its minimal aglycone (fluoride) and the inverted configuration at C-5. This result is suggestive, however, of 5FGulF acting as a slow substrate with a rate-limiting deglycosylation step, as is seen with some other 5-fluorosugars and their corresponding glycosidases (16Howard S. He S.M. Withers S.G. J. Biol. Chem. 1998; 273: 2067-2072Abstract Full Text Full Text PDF PubMed Scopus (84) Google Scholar, 17Numao S. He S.M. Evjen G. Howard S. Tollersrud O.K. Withers S.G. FEBS Lett. 2000; 484: 175-178Crossref PubMed Scopus (29) Google Scholar, 32McCarter J.D. Withers S.G. J. Biol. Chem. 1996; 271: 6889-6894Abstract Full Text PDF PubMed Scopus (121) Google Scholar). When 5FGulF was assayed as an inactivator, no inactivation was observed, presumably because the deglycosylation step is still fast relative to the assay time. However, when the mutant enzyme was incubated with 5FGulF, and the residual activity was assayed at 16 °C, time-dependent inactivation was indeed observed with a second order rate constant, ki/Ki value of 33 s–1m–1 (Fig. 1, a and b). When excess 5FGulF was removed the inactivated enzyme was found to reactivate over time, demonstrating the catalytic competence of the intermediate formed upon hydrolysis of 5FGulF by the D341N mutant dGMII. The above results suggest that 5FGulF is acting as a slow substrate for the D341N mutant dGMII, with deglycosylation as the rate-limiting step. Inactivation is only observed when assayed at low temperatures such that deglycosylation is slow relative to the assay time. To confirm whether this also applied to the wt enzyme, 5FGulF was assayed as a substrate using the fluoride ion electrode. At 37 °C, 5FGulF was found to be a substrate of wt dGMII with a kcat value of 5.1 × 10–3 s–1 and a Km value of 0.2 mm. As in the case of the mutant enzyme, the relatively low Km value is likely because of the accumulation of the intermediate species as a result of the deglycosylation step becoming rate-limiting. Consistent with this hypothesis, when the wt dGMII was incubated with 5FGulF and the residual activity assayed at 8 °C, time-dependent inactivation was observed. The reactivation, however, was too rapid for the determination of any reliable kinetic parameters. Although the 2-deoxy-2-fluorosugars have been successfully used to label β-retaining glycosidases, they have proven quite ineffective with α-retaining glycosidases. Therefore, it was not surprising that all attempts at trying to inactivate either wt or D341N mutant dGMII with 2FManF were unsuccessful. Instead, the 2FManF was found to be a substrate of wt dGMII with a pseudo-second order rate constant, kcat/Km, of 8.5 × 10–5 s–1m–1 (where the Km > 25 mm). This high Km value is most likely a reflection of the 2-hydroxyl group being important to substrate binding, which would not be surprising given the previously observed role of the 2- and 3-hydroxyl groups in binding to the active site zinc (18van den Elsen J.M.H. Kuntz D.A. Rose D.R. EMBO J. 2001; 20: 3008-3017Crossref PubMed Scopus (183) Google Scholar). Unfortunately, because of the extremely low kcat value, it was not possible to determine any kinetic parameters for the hydrolysis of 2FManF by the D341N mutant dGMII. However, when tested as a reversible inhibitor, a Ki value of 7.5 mm was determined for this compound with the D341N mutant dGMII. Once again, this value is considerably lower than the Km value for 2FManF hydrolysis with wt dGMII, suggesting that 2FManF is a very slow substrate of the mutant enzyme with a rate-limiting deglycosylation step. Mass Spectrometric Analysis—The kinetic analysis clearly indicates that there is accumulation of an intermediate species in the case of the hydrolysis of 5FGulF by either the wt or D341N mutant dGMII. The results also suggest that the deglycosylation step is rate-limiting for hydrolysis of 2FManF by D341N mutant dGMII, likely resulting in an accumulation of an intermediate species. To determine the nature of these intermediate species, each was analyzed using electrospray ionization-mass spectrometry. Compared with the free D341N mutant enzyme (120,502 ± 20 Da), the mass of the inactivated D341N mutant enzyme (120672 ± 20 Da) was greater by 170 Da (see Fig. 1c for representative data). This value corresponds, within experimental error, to the expected increase of 181 Da for the addition of a covalently bonded 5-fluoro-gulosyl moiety. A similar increase in mass was observed when comparing the mass of the free wt dGMII with that of the inactivated wt enzyme. This demonstrates that a covalent intermediate is also formed during the hydrolysis of 5FGulF by wt dGMII. Incubation of the D341N mutant dGMII with 2FManF yielded a species with a mass (120,634 ± 20 Da) higher by 166 than that of the free enzyme (120,468 ± 20 Da). This corresponds to the covalent addition of a 2-fluoro-mannosyl moiety to the enzyme (theoretical change, 165 Da). Only ∼50% of the enzyme was labeled under these conditions, most likely because of the rapid turnover of the intermediate species at steady state. This is consistent with the failure of 2FManF to inactivate dGMII. However, the observation of a covalent species clearly shows that the hydrolysis of 2FManF by the mutant enzyme goes via a covalent glycosyl-enzyme intermediate and that this intermediate accumulates during steady state hydrolysis. Structure of the Intermediate Formed from wt dGMII Reaction of 5FGulF—We have been able to obtain a very high resolution structure (1.20 Å) stru
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