- Research article
- Open Access
Structure of the dimeric N-glycosylated form of fungal β-N-acetylhexosaminidase revealed by computer modeling, vibrational spectroscopy, and biochemical studies
BMC Structural Biology volume 7, Article number: 32 (2007)
Fungal β-N-acetylhexosaminidases catalyze the hydrolysis of chitobiose into its constituent monosaccharides. These enzymes are physiologically important during the life cycle of the fungus for the formation of septa, germ tubes and fruit-bodies. Crystal structures are known for two monomeric bacterial enzymes and the dimeric human lysosomal β-N-acetylhexosaminidase. The fungal β-N-acetylhexosaminidases are robust enzymes commonly used in chemoenzymatic syntheses of oligosaccharides. The enzyme from Aspergillus oryzae was purified and its sequence was determined.
The complete primary structure of the fungal β-N-acetylhexosaminidase from Aspergillus oryzae CCF1066 was used to construct molecular models of the catalytic subunit of the enzyme, the enzyme dimer, and the N-glycosylated dimer. Experimental data were obtained from infrared and Raman spectroscopy, and biochemical studies of the native and deglycosylated enzyme, and are in good agreement with the models. Enzyme deglycosylated under native conditions displays identical kinetic parameters but is significantly less stable in acidic conditions, consistent with model predictions. The molecular model of the deglycosylated enzyme was solvated and a molecular dynamics simulation was run over 20 ns. The molecular model is able to bind the natural substrate – chitobiose with a stable value of binding energy during the molecular dynamics simulation.
Whereas the intracellular bacterial β-N-acetylhexosaminidases are monomeric, the extracellular secreted enzymes of fungi and humans occur as dimers. Dimerization of the fungal β-N-acetylhexosaminidase appears to be a reversible process that is strictly pH dependent. Oligosaccharide moieties may also participate in the dimerization process that might represent a unique feature of the exclusively extracellular enzymes. Deglycosylation had only limited effect on enzyme activity, but it significantly affected enzyme stability in acidic conditions. Dimerization and N-glycosylation are the enzyme's strategy for catalytic subunit stabilization. The disulfide bridge that connects Cys448 with Cys483 stabilizes a hinge region in a flexible loop close to the active site, which is an exclusive feature of the fungal enzymes, neither present in bacterial nor mammalian structures. This loop may play the role of a substrate binding site lid, anchored by a disulphide bridge that prevents the substrate binding site from being influenced by the flexible motion of the loop.
Fungal β-N-acetylhexosaminidases catalyze the hydrolysis of chitobiose into its constituent monosaccharides. These enzymes are physiologically important during the life cycle of the fungus for the formation of septa, germ tubes and fruit-bodies [1–3]. These processes are important in control of fungal and insect pests  and are relevant to human diseases , lending considerable interest in the catalytic mechanism of these enzymes. The enzymes are also used in chemoenzymatic synthesis of biologically interesting oligosaccharides based on their effective transglycosylation of β-GlcNAc and β-GalNAc [6–9].
Crystal structures are known for several β-N-acetylhexosaminidases from the glycohydrolase 20 family including the monomeric bacterial enzymes from Serratia marcescens [10, 11] and Streptomyces plicatus [12, 13]. The catalytic domain of β-N-acetylhexosaminidase is an α/β TIM-barrel. Crystallization with substrate analogs showed the conserved residues Asp539-Glu540 to be close to the binding site and thus predict them to play a key role in chitobiose hydrolysis with Glu540 acting as a proton donor to the substrate, while Asp539 restrains its acetamido group in a specific orientation by hydrogen bonding with N2 of the nonreducing sugar . β-N-acetylhexosaminidase from Streptomyces plicatus has been co-crystallized with the cyclic intermediate analogue N-acetylglucosamine-thiazoline. The pyranose ring of the analogue is bound in the active site in a conformation close to that of a 4C1 chair. Within the substrate-binding pocket, Tyr393 and Asp313 appear important for positioning the 2-acetamido group of the substrate for nucleophilic attack at the anomeric center and for dispersing the positive charge distributed into the oxazolinium ring upon cyclization . Experiments with two mutated forms of the enzyme (Asp313Ala and Asp313Asn) provided evidence that Asp313 stabilizes the transition states, and assists to correctly orient the 2-acetamido group for catalysis . Recently, the structure of dimeric human lysosomal β-N-acetylhexosaminidase has been solved providing new insight into the mechanism of Sandhoff disease . Most mutations associated with late-onset Sandhoff disease reside near the subunit interface, and are thus proposed to interfere with the correct formation of the enzyme dimer .
The fungal β-N-acetylhexosaminidases are robust enzymes commonly used in our laboratories in chemoenzymatic syntheses of oligosaccharides [6, 7, 9]. We have previously reported the remarkable inducibility of a fungal β-N-acetylhexosaminidase from Aspergillus oryzae by GlcNAc . This enzyme was purified to homogeneity from the culture medium, and its sequence was determined using both direct protein sequencing and DNA sequencing of a genomic clone containing the hexA gene . In the present work, to initiate structural studies of this enzyme we performed sequence alignment and homology modeling, and constructed a molecular model of the enzyme and of its complex with the natural substrate chitobiose and with its non-cleavable analog GlcNAcβ1→4ManNAc . Several experimental approaches provide experimental verification of the overall features suggested by the model. Disulfide bridging, secondary structure, and the mode of subunit assembly were determined experimentally and correlated with the model. We also constructed a model of the N-glycosylated enzyme, and compared it with kinetic properties of the native and deglycosylated enzymes.
Molecular models of β-N-acetylhexosaminidase
We have used the primary structure of the fungal β-N-acetylhexosaminidase determined in our laboratory  to perform homology modeling using the solved structures of these enzymes from Serratia marcescens , Streptomyces plicatus  and Homo sapiens [14, 19]. Alignment of the amino acid sequences of the fungal enzyme and these three structurally solved enzymes reveals both areas of extensive amino acid similarity and segments that appear unique to the fungal enzyme (Fig. 1). The shown alignment is the result of a combination of a structural alignment of the three crystal structures and the sequence alignment generated with ClustalX, to exclude misalignment in sequence variable regions, as especially the chitobiase from Serratia marcescens has additional domains. Starting from this alignment we created a structural model of the catalytic subunit including the small N-terminal zincin-like domain using a restraint-based comparative modeling approach. In this model we grouped the six cysteins into three pairs according to the closest distance. The three cysteine pairs Cys290-Cys351, Cys448-Cys483 and Cys583-Cys590 were adjusted into the model by repeating the modeling procedure with the additional restraints as an input for Modeller. The final model had 82.8% of residues in the most favored regions of the Ramachandran plot and an acceptable overall geometry, both determined with the ProCheck program. The overall g-factor of the structure obtained showed a value of -0.22. The g-factor tries to quantify the overall geometry and its value should be above -0.5; values below -1.0 may indicate a wrong structure. With respect to the general shortcomings of homology modelling especially in the loop and sequence variable regions, we decided to solvate the homology structure in SPC water to refine it by 20 ns of molecular dynamics in a NPT ensemble. According to the root mean square deviation of the Cα atoms the structure gets after 7 ns into an equilibrium state oscillating around a fixed value with a deviation of ± 0.3 Å. The resulting catalytic subunit has a kidney-shaped structure with approximate dimensions of 6.8 × 5.8 × 5.6 nm (Fig. 2A). All amino acids involved in catalysis are concentrated in the central TIM barrel, with the catalytic glutamic acid on the upper border (Fig. 2B, center). An overlay with the corresponding amino acids of the three template structures with our final model shows that their position is conserved among the available structures and that the active site in our model structure was stable during the molecular dynamics refinement of the overall structure (Fig. 3).
Docked into the binding site, the natural substrate N, N'-diacetylchitobiose, participates in the hydrogen bonding between Asp345 and Tyr445 with the acetamido group of the non-reducing GlcNAc moiety. The non-reducing end of the disaccharide is locked into the active site owing to the hydrogen bonds of Arg193, Asp447, Trp482 and Glu519 with the C4 and C5 OH groups. The reducing end of the disaccharide is stabilized by a π-π interaction with the aromatic ring of Trp482. After the substrate being docked and the complex being solvated it shows an initial equilibration phase with rather high fluctuations of the observed binding energies and quite large changes in root mean square deviation of the binding site residues (Fig. 4). However, after 1.6 ns of MD simulation the system seems to have established the binding energy gets stable with only minor fluctuation and the root mean square deviation of the binding site residues arrives back at a stable value for each of the residues that corresponds to the concrete amino acids rigidity (Fig. 5). The mean value of the binding energy over the time interval from 1.6 ns to 3 ns is 447.1 kJ/mol [see Additional file 1]. The root mean square fluctuation of the protein is measured during the last 10 ns of the 20 ns MD simulation, as the structure is at that time already several nanoseconds in a equilibrium state. Figure 6 shows all active site residues belonging to the less flexible part of the protein with minimum fluctuation in time. The root mean square fluctuation of Arg193, Tyr445, Asp447 and Trp482 describes these residues as the most rigid part of the protein and thus these amino acids should play the key role for substrate specificity. Asp345 and the catalytic residue Glu346 show a little less rigidity which can be interpreted that they must be able to orient themselves to the O-glycosidic bond after substrate binding.
Since initial biochemical characterization of β-N-acetylhexosaminidase from Aspergillus oryzae indicated it to be a dimer, we used the physiologically relevant dimeric crystal structure of human β-N-acetylhexosaminidase  (pdb ID: 1o7a) to model the dimeric fungal enzyme according to the physiolocially valid dimerization interface I found in the crystal structure (Fig. 2C). Similar like in crystal structure of human β-N-acetylhexosaminidase, the proposed dimer interface is formed mainly by loop regions at the C-terminal face of the (β, α)8-barrel, as is the active site. Furthermore, the large flexible loop residues anchored by the disulphide bridge of Cys448 with Cys483, an exclusive feature of the fungal enzymes, contributes to the dimer interface.
Detailed examination of the dimer contact surface reveals a buried surface area per monomer of 2373 Å2 and 64 residues containing atoms closer than 3.6 Å to atoms of the interface partner with 19 residues being hydrophobic (Ile, Leu, Val, Met, Tyr, Phe and Trp). We were able to identify 39 hydrogen bonds between both monomers, with the first monomer accepting 22 and donating 17. The presence of two Arg, three His, three Glu and 4 Asp explains the importance of ionic interactions between of Arg and His residues of one subunit and Asp and Glu residues of the opposite subunit. The model thus indicates that the β-N-acetylhexosaminidase dimer may be a reversible function of pH, with acidic environments favoring dissociation into subunits. Indeed, down to pH3.5 the enzyme exists exclusively in the dimeric form, but at pH2.5 it exists only as monomeric, but enzymatically active, subunits as revealed by gel filtration and enzyme assay . When the dissociated enzyme is titrated back to pH5.0, after a short incubation the dimer forms to the original extent . This result also indicates contact areas that are easily re-established depending on the ionization state of residues with pKa values near pH3. The role of the ionic pairs of the above type in the formation the dimeric structure of hexosaminidases is well established. In human hexosaminidase B there is Asp494-Arg533 at the interface between the catalytic subunits in the dimer  (Asp522 and His586 in our structure). Notably, since in human hexosaminidases the dimerization is essential for the full catalytic activitity, mutations in the above "interface" amino acids are among the well documented human mutations that have been found in the less severe (late onset) forms of Sandhoff  and Tay-Sachs [18, 21] diseases. Interestingly, in the dimeric structure the large flexible loop of the opposite monomer is about 1 nm above the C-terminal face of the (β, α)8-barrel, and the active site giving the impression of a substrate binding site lid (Fig. 2D, E).
Six putative sites of N-glycosylation are present in the primary structure of the fungal enzyme . Digestion of β-N-acetylhexosaminidase with N-glycanase results in significant mobility shift on SDS-PAGE (Figure 7A), indicating that one or more glycosylation sites are used on the secreted enzyme. In order to reveal the details of these important structural modifications, we have performed a detailed analysis of the actual occupaccy of all these sites using the standard N-glycanase/H218O technique [22, 23] and mass spectrometry which revealed that all six sites of N-glycosylation were indeed used in the actual hexosaminidase preparation. This conclusion was further supported by complete proteolytic digestion of hexosaminidase followed by isolation of the glycopeptides on immobilized plant lectin Concanavalin A, and identification by these glycopeptides by mass spectrometry. Glycopeptides that covered all six sites of N-glycosylation could be identified on the mass spectra (results not shown).
The fact that all the glycopeptides described above could be efficiently recovered on the immobilized Concanavalin A provided a strong indication for the high-mannose type oligosaccharides being present at the individual N-glycosylation sequences. In order to support this indication, the oligosaccharides were released by N-glycanase and analyzed using a combination of Dionex oligosaccharide profiling, exoglycosidase digestion, and mass spectrometry as described in the Experimental section. When we performed this analysis on oligosaccharides released from the entire hexosaminidase, we obtained the overall glycosylation of the entire enzyme indicating the hexamannosyl oligosaccharide M6 as the predominant structure (Table 1, lane Asnall sites). This overall glycosylation profile was very similar to the one we found on the two particular sites of glycosylation Asn427 and Asn499 (Table 1). As the experimental results show a M6 structure as the predominant oligosaccharide, we choose this structure as the oligosaccharide to be attached to the molecular model of β-N-acetylhexosaminidase. In order to get a preliminary insight into the structural arrangement, and possible biological role(s) of N-glycans in the fungal β-N-acetylhexosaminidase, we have sterically fitted and covalently linked the most prevalent (M6) oligosaccharide to each site of glycosylation on our model structure. This gave rise to the model of a fully glycosylated enzyme shown as the most prevalent glycoform in Fig. 2C. Although the analysis of the actual glycoforms would be a much more demanding task, the model shown in Fig. 2C illustrates nicely the surface exposure of the individual oligosaccharides protecting the enzyme dimer from the effects of the extracellular environment.
Evidence for the status of cysteines using protein chemistry and mass spectrometry
The fungal β-N-acetylhexosaminidase is secreted into the extracellular environment, indicating the possibility for forming disulfide bonds. Since there are six cysteines in the primary structure of β-N-acetylhexosaminidase, the status of their sulfhydryl groups was determined by mass spectrometry . It has been found that the experimentally verified arrangement of the disulfide bridges corresponds exactly to that anticipated by the molecular model (see above). Moreover, these results were further confirmed by differential mapping of pepsin peptides with and without DTT reduction. Using this technique reverse phase HPLC identified only one peak with significant shift in the retention time. One of the components of this chromatographic peak was the peptide with m/z 2538.8 (Fig. 7A). Significant decrease in its intensity with concomitant intensity increase of peaks at m/z 1574.0 and 968.2 (Fig. 7B), observed upon addition of DTT, indicates that this peptide consists of two disulfide linked peptides. These were identified by PSD measurement as peptides Tyr445-Phe453 and Asn474-Thr488 containing the cysteines 448 and 483, respectively. This confirms the identification of the disulfide bond between Cys448 and Cys483.
In the model structure without disulfide bridges, Cys483 is 6.2 Å away from Cys448. The formation of the disulfide bond in the model is leading to a somewhat different position of one of the loops, easily accommodating the experimentally determined disulfide bridge with only a relatively minor change in the overall structure. As Asp447 and Trp482 are both amino acids forming the active site, the flexible loop between Cys448 and Cys483 is right beside the substrate-binding site. The behavior of this loop measured by root mean square fluctuation during the last 10 ns MD simulation shows that it is highly flexible (Fig. 6), which may be functionally important. The nearby disulfide bond thus stabilizes not only this loop but also the active site.
The effect of deglycosylation on enzyme activity and stability
Preliminary deglycosylation experiments indicated the possibility to deglycosylate the enzyme under native conditions (Fig. 8A, cf. lane 1 and 11). We evaluated several protocols for deglycosylation using N-glycanase or endoglycosidase H under conditions compatible with optimal stability of the deglycosylated hexosaminidase (Fig. 8A). The deglycosylated enzyme remains fully active (Fig. 8B), and displays kinetic parameters indistinguishable from those of the native enzyme (Km = 0.71 mM and Vmax = 2.16 nmol/min vs. Km = 0.45 mM and Vmax = 1.55 nmol/min for the native form).
To probe the effects of deglycosylation on enzyme stability, native and deglycosylated enzymes were incubated at a range of pH values. No significant activity differences were found at alkaline or neutral pH. However, the deglycosylated enzyme is significant less stable than the native enzyme at pH values below 4. The activity of the deglycosylated enzyme after incubation at pH3.5 and 3.0 is about 75% of the native enzyme, and at pH2.5 the deglycosylated enzyme is only half as active as the native enzyme (Fig. 8C). This result suggests that N-linked oligosaccharides may be important for the stabilization of β-N-acetylhexosaminidase at low pH.
Structure and stability determined by vibrational spectroscopy
Infrared spectra of β-N-acetylhexosaminidase are characterized by two major bands at 1655 cm-1, and 1543 cm-1 (Fig. 9A), associated with the amide I and II vibrations, respectively. The position and shape of these bands are sensitive to protein conformation and secondary structure content. The second derivative, which can identify overlapping components, reveals three major bands in the amide I region. The largest component at 1655 cm-1 belongs to α-helical and disordered conformations [25, 26]. Two other components at 1642 cm-1 and 1626 cm-1 are assigned to β-sheets [25, 26]. The unresolved band at 1684 cm-1 indicates the presence of antiparallel β-strands , and the second part of the unresolved band at 1675 cm-1 corresponds to β-turns [25, 26]. Characteristic side chain absorption of Tyr and Phe is observable at 1517 cm-1 and 1493 cm-1, respectively . The FTIR spectrum of β-N-acetylhexosaminidase after deglycosylation (Fig. 9B) shows no significant shifts with respect to the native protein.
Raman spectroscopy (Fig. 10) confirms the presence of disulfide bonds by the band at 520 cm-1 due to the S-S stretching vibrations. The frequency of this band is sensitive to the local conformation of a disulfide bridge. The band at 520 cm-1 corresponds to the sulfide bridges in a conformation close to gauche-gauche-trans (GGT) . The bands in the region of stretching CS vibrations 700–745 cm-1 can be hardly resolved due to presence of three disulfide bridges and their probable flexibility with respect to ν CS vibrations. The intensity ratio of the tyrosine Fermi resonance doublet (828 cm-1 and 852 cm-1; I852/I828 = 1.6) indicates that the tyrosine OH group acts as an acceptor of H-bonds suggesting that some of the tyrosines are solvent exposed . The Raman spectrum after deglycosylation (Fig. 10B) shows significant changes with respect to native protein (Fig. 10A) that are clearly revealed by the difference spectrum (A-B). Almost all marker bands of aromatic side chains are affected by deglycosylation, although the secondary structure was not affected, as discussed below. Thus, changes in the Raman spectrum can be attributed to changes in environment of the residues, presumably due to unmasking of large surface areas of the protein by deglycosylation.
The FTIR amide I and II bands and the Raman amide I band were analyzed by LSA (Table 2) to quantify the secondary structure content of β-N-acetylhexosaminidase and its deglycosylated form. Deglycosylation of the protein leads to small changes in secondary structure, as estimated by LSA, connected with slight decrease of β-sheets (about 3%) and slight increase of α-helices – probably on the periphery of the protein. The LSA methods employs two types of α-helical structure for Raman spectra: short helical segment up to eight residues called monohydrogen-bonded or disordered, and helical segments bracketed by at least four residues on each side named bihydrogen-bounded or ordered. Recently we proposed that the vibrations could be affected by oligomerization or complex formation , therefore the amount of disordered α-helix (about 15%) was included to other structure in Table 2. If we agree with interpretation that the signal from disordered α-helix is caused mostly by dimerization of β-N-acetylhexosaminidase then carbohydrate moiety plays important role in the dimer formation. The lower stability of the deglycosylated enzyme upon prolonged incubations at acidic pH (see Fig. 8C) may then be interpreted as the gradual decomposition of the enzyme dimer into the monomeric units that is fully reversible for the native but not for the deglycosylated enzyme.
We describe here results of structural analysis of dimeric, fully N-glycosylated, fungal β-N-acetylhexosaminidase isolated from the culture medium of the public collection strain of Aspergillus oryzae. The unique biology  and important biotechnological applications of this enzyme [8, 9, 17] stimulated this work as a prerequisite for structure-function studies and enzyme engineering. To create a conceptual basis for the design of structural experiments, the amino acid sequence of the Aspergillus β-N-acetylhexosaminidase determined in our laboratory  was used to generate a molecular model of the catalytic subunit based on sequence homology to three solved β-N-acetylhexosaminidase structures [10, 12, 14]. Our major experimental efforts then have been directed towards understanding the principal differences between the intracellular bacterial enzymes and the eukaryotic enzymes that have to undergo posttranslational modifications to survive better under the more aggressive conditions in lysosomes and the extracellular environment in which they function. In particular, we investigated how enzyme dimerization, disulfide bonding, and glycosylation participate in enzyme stabilization.
Multimerization of enzymes represents one strategy for catalytic subunit stabilization that is often used, for example, in thermophilic organisms. Consistent with this trend, the intracellular bacterial β-N-acetylhexosaminidases are monomeric whereas the extracellular secreted enzymes of fungi and humans occur as dimers. Dimerization of the fungal β-N-acetylhexosaminidase appears to be a reversible process that is pH dependent. This result provides strong support for the participation of titratable functional groups at the subunit interface as predicted by the constructed dimeric structure. The experimental finding that oligosaccharide moieties may also participate in the dimerization process may represent a unique feature of the exclusively extracellular enzymes.
Formation of disulfide bridges is another common way of stabilizing proteins that are secreted into the oxidative extracellular environment. The disulfide bridges detected experimentally are in complete accordance with the molecular model. One disulfide may stabilize a hinge region in a small flexible loop close to the substrate binding site. This loop is an exclusive feature of the fungal enzymes, neither present in bacterial nor mammalian structures. We hypothesize that this loop may play the role of a substrate binding site lid, as has been found previously in several enzymes including other TIM barrels [31–34], anchored by a disulfide bridge that prevents the substrate binding site from being influenced by the flexible motion of the loop. This substrate binding site lid is formed in the dimeric structure by the large flexible loop of the opposite monomer and is close above the C-terminal face of the (β, α)8-barrel, and the active site thus indicating the necessity of a functional dimer for full enzyme function.
Yet another posttranslational modification known to stabilize proteins is N-glycosylation. However, experimental investigation of its role in enzyme function and stabilization is technically difficult, and only a few studies report detailed experimental analysis [22, 35]. In the case studied here we had the rare possibility to fully de-N-glycosylate the enzyme under native conditions. To probe the role of extensive glycosylation in the fungal β-N-acetylhexosaminidase experimentally we used two deglycosylation methods, which are based on the use of either N-glycanase or endoglycosidase H. The former enzyme is an asparagines amidase that cleaves the N-glycosidic (amide) bond between the asparagines and the oligosaccharide chains. This cleavage gives rise to the free β-glycosylamine, and aspartic acid that remains in the protein in the original position of the asparagines. On the other hand, the latter enzyme is a typical endoglycosidase that cleaves a glycosidic linkage between the two GlcNAc residues forming the chitobiose core of the N-linked oligosaccharide. Such a cleavage leaves the N-glycosidic bond on asparagines, and the proximal GlcNAc intact.
We employed endoglycosidase H digestion which allows to efficiently remove the mannose branches while retaining the core GlcNAc residues to avoid the potential dangers of complete protein deglycosylation that we have previously reported . Deglycosylation had only limited effect on enzyme activity, but it significantly affected enzyme stability in acidic conditions. This role of N-glycosylation is consistent with the observation that the enzyme completely deglycosylated by N-glycanase has low stability and tends to precipitate.
In summary, we have shown that complementary biochemical and biophysical methods provided structural insight into critical features of fungal β-N-acetylhexosaminidase, even in the absence of data from protein crystallography or nuclear magnetic resonance. The results reported here also contain useful information for designing experiments that will use high resolution methods of protein structure analysis. Dimerization of the fungal β-N-acetylhexosaminidase appears to be a reversible process that is strictly pH dependent. Oligosaccharide moieties may also participate in the dimerization process which might represent a unique feature of the exclusively extracellular enzymes. Deglycosylation had only limited effect on enzyme activity, but it significantly affected enzyme stability in acidic conditions. Dimerization and N-glycosylation are the enzyme's strategy for catalytic subunit stabilization. One disulfide bridge, neither present in bacterial nor mammalian structures, anchors a flexible loop close to the active site that may play the role of a substrate binding site lid in the physiologically relevant dimer.
The molecular model of β-N-acetylhexosaminidase was generated by a combination of energetic and homology modeling. In a first step three homologs from the glycohydrolase family 20 were extracted from the Protein Databank  – PDB entry: 1c7s: Serratia marcenscens , 1jak: Streptomyces plicates , 1now: Homo sapiens . These proteins show a primary sequence identity about 30% and a homology around 45% (Serratia marcenscens 32/48%, Streptomyces plicates 27/42%, Homo sapiens 31/49%) making homology modeling possible. A structural alignment was generated in SwissPDBViewer  showing a root mean square deviation ~1.4 Å between the three structures. Then the complete primary sequence of β-N-acetylhexosaminidase from Aspergillus oryzae  was aligned with the three homologs in ClustalX  keeping the positions of the structural alignment conserved (see Fig. 1). The three-dimensional model constituted by all non-hydrogen atoms was built and examined by the Modeller6 package . All three disulfide bridges were created and refinement was achieved through algorithmic analysis and minimization with the Tripos force field in the Sybyl/Maximin2 (Tripos). Hydrogen atoms were added and the model of the generated structure was minimized to convergence of the energy gradient less then 0.01 kcal/mol using the Powell minimiser. The minimization included electrostatic interactions based on Gasteiger-Hückel partial charge distributions using a dielectric constant with a distance dependent function ε = 4r and a non-bonded interaction cut-off of 8 Å. The tertiary structure model was checked with Procheck .
The final proposed model was solvated in simple point charge water and four chloride counter-ions were added. The production runs were preceded by short equilibration runs of altogether 250 ps with positional restraints applied on the protein atoms to allow the solvent to relax and 250 ps of protein relaxation without positional contraints and a timestep of 2 fs. The simulation box was sized 1 nm in each direction from the protein surface and filled with 18950 water molecules to give a system of 62049 atoms. In a first step the system is minimized by 500 steps of steepest descent minimization, followed by 20 ps of solvent relaxation with a 1 fs timestep. Chloride ions were added by replacing four water molecules to neutralize the systems, followed by 30 ps of ion and solvent relaxation with a 1 fs timestep and 200 ps with a 2 fs timestep. The following production MD simulations, without any restraints, were 20 ns long and were run with GROMACS 3.2 [42, 43] using the gmx force field, with a 5 fs time step (which is possible because dummy hydrogens are used). SETTLE (for water) and LINCS were used to constrain covalent bond lengths, and long-range electrostatic interactions were computed with the Particle-Mesh Ewald method. The temperature was kept at 300 K by separately coupling the protein and solvent to an external temperature bath (τ = 0.1 ps) . The pressure was kept constant at 1 bar by weak coupling (τ = 1.0 ps) to a pressure bath. The protein proved to be stable during simulation.
The three dimensional structure of the N-linked complex glycan (M6 oligosaccharide structure) was calculated with Sweet  and the carbohydrates were sterically fitted by visual analysis and covalently attached to the molecular model of β-N-acetylhexosaminidase in Sybyl. The dimeric structure was created by fitting two monomeric structures onto the dimeric structure from Homo sapiens within YASARA . Then the second monomer was placed in a distance of about 1 nm away from its final position to avoid sterical conflicts of the large flexible loop with the first monomer, followed by 2000 steps of simulated annealing minimization. The second monomer was such moved in 0.1 nm steps to its final position, each step followed by another 2000 steps of simulated annealing minimization.
Chitobiose was build in YASARA, forcefield parameters were assigned using the AutoSMILES approach , in a first step YASARA calculated semi-empirical AM1 Mulliken point charges that were corrected by assignment of AM1BCC atom types and improved AM1BCC charges by fragments of molecules with known RESP charges, to closer resemble RESP charges. Corresponding bond, angle and torsion potential parameters are taken from the General AMBER force field. For the docking experiments our model structure was fitted onto the crystal structure of 1qbb , a bacterial chitobiase complexed with N, N'-diacetylchitobiose. Chitobiose was placed in an arbitrary position according to the ligand co-ordinates in the bacterial chitobiase complex. Exact positioning of the ligand was done by a two-step procedure, energy minimization followed by a molecular dynamics. The ligand-protein system was minimized by 2000 steps followed by a 3 ns MD simulation in aqueous solution using the YAMBER2 force field . The protein structure was placed into a box, which was 1 nm larger than the protein along all three axes. The box was filled with TIP3P water, sodium ions were iteratively placed at the coordinates with the lowest electrostatic potential until the cell was neutral. Molecular dynamics simulations were run with YASARA, using a multiple time step of 1 fs for intra-molecular and 2 fs for intermolecular forces. A 1.2 nm cut-off was taken for Lennard Jones forces and the direct space portion of the electrostatic forces, which were calculated using the Particle Mesh Ewald method  with a grid spacing 0.1 nm, 4th order B-splines, and a tolerance of 10-4 for the direct space sum. The simulation of interaction was then run at 298 K and constant pressure (NPT ensemble) to account for volume changes due to fluctuations of homology models in solution.
Interaction energies were calculated considering the internal energy obtained with the specified force field, as well as the electrostatic and Van der Waals solvation energy obtained. The electrostatic solvation energy estimates the interaction energy between the solvent and the solute by treating the solvent as a continuum without explicit solvent molecules. A first-order boundary element approximation to the solvation energy was used. Van der Waals solvation energy was calculated as a function of the solute's solvent accessible surface area . The entropic cost of fixing the ligand in the binding site is almost impossible to calculate accurately, but fortunately not needed since it mainly depends on characteristics that are constant during the simulation (ligand and protein size, side-chains on the surface etc.). The entropic component is thus a constant factor that can be omitted. The more positive the interaction energy, the more favorable is the interaction in the context of the chosen force field.
Enzyme isolation and characterization
Aspergillus oryzae strain CCF1066 (C zech C ollection of F ungi, Faculty of Science, Charles University in Prague) was grown as described in . Ammonium sulfate enzyme precipitate was purified by hydrophobic chromatography on Phenyl-Sepharose 6 Fast Flow (Amersham) using elution with the reversed ammonium sulfate gradient (0.6 M to 0 M in 20 mM sodium phosphate buffer pH6.8). Partially purified enzyme was concentrated and dialyzed against 20 mM sodium citrate buffer pH3.5. The enzyme was then purified on SP-Sepharose Fast Flow (Amersham) using elution with sodium chloride gradient (0 to 1 M). The enzyme was concentrated, dialyzed against 20 mM piperazine-HCl pH5.4, and purified on MonoQ HR 10/10 column (Amersham) eluted with sodium chloride gradient (0 to 0.3 M). The enzyme was concentrated to approx. 10 mg/mL, and stored in the stabilization buffer composed of 0.5 M ammonium sulphate in 50 mM citrate buffer pH5.0. Immediately before the spectroscopic measurements, the enzyme was transferred to 50 mM bis-Tris buffer pH5.0. The purity of the enzyme was checked by SDS-PAGE, and the concentration was determined by .
Determination of enzymatic activity
Enzymatic activity was measured using 4-nitrophenyl-2-acetamido-2-deoxyglucopyranoside at either saturating concentration (5 mM, used for most determinations), or at concentrations around Km (for the evaluation of enzymatic parameters) according to the procedure of .
Analysis of individual sites of N-glycosylation
Occupancy of sites of N-glycosylation was determined as described previously  by comparison of measured masses of peptides corresponding to the sites of glycosylation. The peptides were generated after SDS-PAGE separation by means of in gel digestion with sequencing grade trypsin (Promega), or sequencing grade Asp-A protease (Roche) either in normal or isotopic water H218O (Fluka). The peptides were extracted from the gel, desalted and concentrated with C-18 microcolumn (ZipTip C18, Millipore) and analyzed by MS. To analyze individual sites of N-glycosylation, the glycoprotein was digested with trypsin, and glycopeptides were captured on concanavalin A – Sepharose resin (Amersham), washed, and eluted with 0.01 M Tris-HCl, pH8.0, 0.15 M NaCl and 0.3 M D-mannose (Sigma). Individual glycopeptides were separated by reverse-phase chromatography on the Vydac C-18 column (Dionex), equilibrated in 0.1% trifluoroacetic acid, and eluted by acetonitrile gradient to 70% over 120 min. The glycopeptides were identified by MALDI MS measurements in their native and or deglycosylated state. Glycans released from glycopeptides or from the intact protein with PNGase F (New England BioLabs) in 50 mM sodium bicarbonate pH8.0 were desalted on mini-columns (10 μL of bead volume) filled with non-porous graphitized carbon , and separated by HPAEC-PAD (DX500, Dionex) on Carbopac PA100 column. Elution was performed by 5 min isocratic step with 0.1 M NaOH (solvent A) followed by linear gradient from 0 to 35% of 0.1 M NaOH, 0.6 M NaOAc (solvent B) over 48 min and another linear gradient to 100% solvent B within 10 min at the flow rate 1 mL/min. The collected peaks were desalted (as above) and analysed by MALDI MS. For further characterization of the isolated oligosaccharides, the digestion with α-mannosidase was performed using 5 U/mL of the enzyme in 50 mM sodium citrate buffer pH5.0 .
Enzymatic deglycosylation under native conditions
μg of β-N-acetylhexosaminidase was fully deglycosylated either in 100 μl of 50 mM ammonium bicarbonate pH8.0 using 500 units of N-glycanase (New England BioLabs), or in 100 μL of 50 mM citrate buffer pH5.0 using 500 units of EndoH (New England BioLabs). The reaction was carried out for 16 h at 37°C.
pH stability test
pH stability test of the native and deglycosylated enzyme was performed as follows: 1 μL of the stock solution of enzyme was mixed with 9 μL of 50 mM sodium citrate buffer pH2.5, 3.0, 3.5, 4.0, or 4.5, or with 9 μL of 50 mM sodium phosphate buffer pH5.5, 6.0, 6.5, or 7.0, or 50 mM Tris-HCl buffer pH7.5, 8.0, 8.5, or 9.0. These mixtures were incubated for 16 h at 4°C, diluted with 40 μL of the substrate solution (50 mM sodium citrate pH5.0 with 5 mM substrate), and the enzymatic activity was determined as described in .
Infrared spectra of protein samples (9.5 mg/mL native and 8.5 mg/mL deglycosylated in 50 mM bis-Tris, pH5.0) were recorded in CaF2 10 μm BioCell™ (BioTools) at room temperature with a Bruker IFS-66/S FTIR spectrometer using a standard source, a KBr beamsplitter and an MCT detector. 4000 scans were collected with 4 cm-1 spectral resolution and Happ-Genzel apodization function. Spectral contribution of a buffer in carbonyl stretching region was corrected following the standard algorithm .
Raman spectra were recorded in a standard 90° geometry on a multichannel instrument based on a 600-mm single spectrograph (Monospec 600, Hilger & Watts) with a 1200-grooves/mm grating and a liquid N2-cooled CCD detection system (Princeton Instruments) having 1024 pixels along dispersion axis. A holographic notch-plus filter (Kaiser Optical Systems) was used to remove elastically scattered light. The effective spectral slit width was set to ~5 cm-1. Samples were excited with 514.5 nm/50 mW line of an Ar+ laser Innova 300 (Coherent). Calibrated wavenumber scale by Ar+ plasma lines was accurate to ± 1 cm-1.
Measurements were made on protein samples (8–13 mg/mL in 50 mM bis-Tris, pH5.0) in a 10-μL capillary microcell. Spectra, measured at 4°C, were averaged from 150 exposures of 120 s. Spectra were treated according to , then they were smoothed using 9-point Savitsky-Golay algorithm and normalized to the 1447 cm-1 δCH2 band as an internal standard.
Fourier transform infrared
high-performance anion exchange chromatography with pulsed amperometric detection
high performace liquid chromatography
matrix-assisted laser desorption/ionization
- PNGase F:
peptide-N4-(N-acetyl-β-D-glucosaminyl) asparagine amidase F
restrained electrostatic potential
sodium dodecyl sulfate
SDS polyacrylamide gel electrophoresis.
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Supports from the Ministry of Education of the Czech Republic (Nos. LC06010, MSM0021620835, MSM6007665808, 1M 4635608802), the Institutional Research Concept of the Academy of Sciences of the Czech Republic (Nos. AVOZ60870520, AVOZ50200510), and the Grant Agency of the Czech Republic (Nos. 203/04/1045 and 204/06/0771) are gratefully acknowledged. The authors also would like to thank Elmar Krieger, Yasara.org, for his helping hand and Janette Carey, Princeton University, for a critical reading of this manuscript.
RE was responsible for all the work on sequence alignments, molecular modeling, and molecular dynamics studies and performed most of it. MK built up the ligand structure and supervised the docking, VK jr., KH and VB performed all the IR and Raman spectroscopy measurements and the data interpretation. KH was also responsible for the hexosaminidase production. PN, PP and PM were responsible for mass spectrometry measurements, and solved the disulfide bonds arrangement, NK performed the computational glycosylation and the binding energy calculations and OP and JS were responsible for the determination of hexosaminidase glycosylation and performed the deglycosylation studies. HR took part in the enzyme purification and primary structure determination, VK conceived the study and provided the strains and advice on hexosaminidase production, KB was responsible for the overall coordination of the project and produced the first draft of the manuscript, and was responsible for hexosaminidase purification. All authors read and approved the final manuscript.
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