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Chitin, a β-1,4-linked polysaccharide of N-acetylglucosamine (GlcNAc), is hydrolyzed by chitinases (EC 126.96.36.199) that are widely distributed in living organisms and are responsible for self-defense, growth, morphogenesis, cuticle destabilization and stress tolerance [1–5]. A number of chitinase and chitinase-like genes have been isolated from living organisms, and the gene products have been characterized. Although chitinases from plant origins were classified into at least five classes (classes I–V) based on their deduced amino acid sequences [6,7], nowadays a simple classification according to the CAZy database is more widely used for all chitinolytic enzymes . The chitinases are classified into families GH18 and GH19, both of which can be subdivided based on their domain arrangements and sequence deletions. The first three-dimensional structure of a plant chitinase was reported for 26 kDa family GH19 chitinase from barley seeds . The barley enzyme is composed of two lobes, both of which are rich in α-helical structures, with the substrate binding cleft between the lobes. Several loops are found at both ends of the binding cleft (‘loopful’). The structure of the ‘loopful’ family GH19 chitinase from papaya in a complex with GlcNAc monomers has also been reported, and two GlcNAc molecules were found to bind separately to subsites −2 and +1, respectively, in the complex structure . This structure has been used to build a plausible model of a complex with (GlcNAc)4. On the other hand, family 19 chitinases lacking the loops at the two ends of the binding cleft (‘loopless’ chitinase) have been isolated from two bacteria, Streptomyces griseus HUT6037 and Streptomyces coelicolor A3(2), and the evergreen conifer Norway spruce [11–13]. Based on the three-dimensional structures, it was suggested that the substrate binding clefts of the ‘loopless’ enzymes are shorter than those of ‘loopful’ enzymes [14,15]. The crystal structure of a ‘loopless’ family GH19 chitinase from S. coelicolor A3(2) and analysis of the products from chitooligosaccharide degradation by this enzyme revealed that the substrate binding cleft consists of four subsites . However, no quantitative data for binding of (GlcNAc)n to the ‘loopless’ family GH19 chitinases have been reported to date. The subsite arrangement of the ‘loopless’ chitinase can be delineated not only from structural data but also from quantitative binding experiments.
Recently, Taira et al.  have isolated and characterized a family GH19 enzyme from a moss species, Bryum coronatum, designated as BcChi-A. They showed that the molecular weight of BcChi-A is relatively small compared with those of ‘loopful’ GH19 chitinases due to the deletion of several loops. Thus, BcChi-A is a ‘loopless’ family GH19 chitinase like that from S. coelicolor A3(2) rather than a ‘loopful’ GH19 enzyme such as the GH19 chitinase from barley seeds. Interestingly, when the shorter oligosaccharide substrate (GlcNAc)4 was used as the substrate, BcChi-A exhibited a much higher hydrolytic activity (1000-fold) than the ‘loopful’ GH19 chitinase from rye seeds . In this study, to gain insight into the mechanism of (GlcNAc)n binding to the ‘loopless’ family GH19 enzymes, (GlcNAc)n binding to BcChi-A was examined by oligosaccharide digestion, thermal unfolding experiments and thermodynamic analysis using isothermal titration calorimetry (ITC).
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Crystal structures of both ‘loopless’ and ‘loopful’ family GH19 chitinases from various plant species, including barley seeds, jack bean, mustard greens, papaya, Norway spruce and rice, have been solved by different research groups [10,15,22,23]. Nevertheless, no crystal structure of a family GH19 enzyme complexed with a GlcNAc oligosaccharide is available at present. The structure of a GH19 chitinase in a complexed state was obtained only for ‘loopful’ papaya chitinase with two separately bound GlcNAc monomers . Furthermore, quantitative analysis of the substrate binding to GH19 chitinases has not been conducted yet, and the mechanism of substrate recognition has never been understood. In this study, we report the first quantitative data on the substrate binding to a ‘loopless’ family GH19 chitinase, which were obtained by kinetic and thermodynamic strategies.
We compared the observed thermodynamic values with those of other carbohydrate binding systems such as glycoside hydrolases and lectins. First, the Kd value for (GlcNAc)6 binding to BcChi-A is approximately 3-fold and 6-fold higher respectively than that observed for the GH18 chitinases Chit-42 from Trichoderma harzianum and ChiB from Serratia marcescens. Moreover, all three chitinases demonstrate a clear proportionality between chain length and binding strength as Kd values increase with decreasing length of the oligomers. In comparison, (GlcNAc)6 binds 100-fold more strongly to BcChi-A than to human lysozyme (0.77 versus 73 μm).
Furthermore, for (GlcNAc)6 binding at pH 7.0 and 30 °C, the driving force is the enthalpy change (ΔHr° = −9.5 ± 0.1 kcal·mol−1) and the solvation entropy change (−TΔSsolv° = −7.6 ± 0.6 kcal·mol−1). The latter is almost negated by an unfavorable conformational entropy change (−TΔSconf° = 6.1 ± 0.6 kcal·mol−1). Such thermodynamic signatures are also observed for methyl 3-O-(α-d-mannopyranosy1)-α-d-mannopyranoside binding to the lectin concanavalin A , xylo-oligosaccharides (XOS) to xylanases  and chitotriose to lysozyme . The latter system deviates to a certain extent with a much more negative conformational entropy change compared with the positive solvation entropy change (−TΔSsolv° = −8 kcal·mol−1 versus −TΔSconf° = 14 kcal·mol−1). When the binding of 18 inhibitors to the family GH1 β-glucosidase TmGH1 was studied, it was shown that, even though 11 had positive entropy changes, all 18 had negative enthalpy changes . An interesting case is the binding of N-acetyl gluconolactam, N-acetylglucosaminono-1,5-lactone O-(phenylcarbamoyl)-(Z)-oxime (PUGNAc) and N-acetylglucosaminono-1,5-lactone (Z)-oxime (LOGNAc) to a bacterial O-GlcNAcase homologue. The lactam and PUGNAc binding is clearly driven by favorable enthalpy changes with little entropic contribution, but the LOGNAc binding is facilitated by an increase in entropy with no contribution by enthalpy . For (GlcNAc)6 binding to a family 18 chitinase (chitinase B of S. marcescens), the thermodynamic signatures differ remarkably from the ones discussed. Here, binding is accompanied by an enthalpic penalty (ΔHr° = 1.2 kcal·mol−1) and it is desolvation (−TΔSsolv° = −13 kcal·mol−1) that drives the binding with a non-contributing conformational entropy change (−TΔSconf° = −0.1 kcal·mol−1) . For allosamidin binding to S. marcescens chitinase B, the enthalpic penalty is even larger (ΔHr° = 3.8 kcal·mol−1) and the strong binding is facilitated by favorable conformation changes (−TΔSconf° = −11 kcal·mol−1) and, to a smaller extent, desolvation (−TΔSsolv° = −5 kcal·mol−1). These two examples appear to be anomalies in carbohydrate interactions with respect to having unfavorable enthalpic terms.
The binding of (GlcNAc)6 to BcChi-A was independent of pH. For interaction, this cannot only be due to the fact that there are no titratable groups on the ligands, but must also imply that the titratable groups remaining in the catalytic center of BcChi-A after mutating the catalytic Glu61 to a non-titratable Ala are not significantly titrated in the pH 6.0–8.0 range.
BcChi-A hydrolyzed chitin oligosaccharides at rates in the order (GlcNAc)6 > (GlcNAc)5 > (GlcNAc)4 >> (GlcNAc)3, suggesting that the longer the chain length of the substrate, the higher the affinity of the oligosaccharides to the enzyme (Fig. 1). A similar suggestion was obtained from the thermal unfolding experiments shown in Fig. 2. The transition temperature of thermal unfolding (Tm) was elevated by the addition of (GlcNAc)n (n = 2, 3, 4, 5 and 6). The Tm elevations were found to be proportional to the polymerization degree of the (GlcNAc)n added. It is clear that the Tm elevation (ΔTm) is derived from an increase in the number of interaction sites between BcChi-A and (GlcNAc)n (n = 2–6). The Tm data listed in Table 1 are also consistent with the substrate-size dependence of binding free energy change listed in Table 2. All of these data support the idea that BcChi-A has at least six subsites.
In a previous paper  it has been shown that (GlcNAc)6 productively binds to the enzyme through three types of productive binding modes, from −4 to +2, from −3 to +3, and from −2 to +4 subsites of BcChi-A. (GlcNAc)5 binds to subsites from −3 to +2 and from −2 to +3. From the oligosaccharide digestion experiments (Fig. 1C), (GlcNAc)4 was found to productively bind to −2 to +2 subsites. (GlcNAc)3 binds to −2 to +1 subsites to be split into (GlcNAc)2 and GlcNAc (Fig. 1E). The binding modes are summarized in Fig. 5 together with the binding free energy changes. Thus, the binding free energy change of subsite +2 is estimated to be −1.6 kcal·mol−1 by subtraction of the free energy change of (GlcNAc)3 (−5.0 kcal·mol−1) from that of (GlcNAc)4 (−6.6 kcal·mol−1). The difference between the binding free energy changes for (GlcNAc)4 and (GlcNAc)6 is calculated to be −1.9 kcal·mol−1, suggesting that there are at least two more subsites that contribute with a ΔGr° ∼ −0.9 kcal·mol−1 each. These values correspond well with XOS binding to xylanase where the free energy change contribution from individual subsites is ∼ −0.7 kcal·mol−1 . Both (GlcNAc)n–chitinase and XOS–xylanase interactions experience a more negative ΔGr° with respect to an increase in substrate length due to a decrease in ΔHr°, in spite of smaller magnitudes for the chitinase, ΔHr° ∼ −0.9 kcal·mol−1 per unit outside the −2 to +2 subsites compared with ΔHr° ∼ −3 kcal·mol−1 per unit for the xylanase outside the −2 and −1 subsite . The binding of (GlcNAc)3 deviates somewhat from the binding of the longer chitooligosaccharides in that the enthalpic contribution is 0.7 kcal·mol−1 (see Table 2) more favorable than observed for (GlcNA)4. This is not surprising as there are likely to be many favorable interactions between the sugars and BcChi-A in the −2 to +1 subsites to overcome the loss of free energy associated with the necessary distortion of the sugar bound in the −1 subsite to allow for hydrolysis. Such thermodynamic signatures have been observed previously for various chitinases by both direct calorimetric measurements  and calculations of binding free energies from kinetic measurements [31,32] as discussed below. A subsite energy mapping has also been done for the (GlcNAc)n–S. marcescens chitinase B interactions. In that work, ΔGr° for each subsite–sugar unit interaction varies, with −2.7 kcal·mol−1 for subsite +2, ∼ −1 kcal·mol−1 for subsites +3 and +4, and −0.3 kcal·mol−1 for subsite −3 . In addition, binding to +2 and +3 is associated with favorable entropy gains and unfavorable enthalpy gains despite some enthalpically favorable stacking interactions with Trp and Phe residues. It is suggested that favorable conformational changes and unfavorable intermolecular bond breaks within the protein are triggered by the binding of substrate to these subsites, which is responsible for the observed thermodynamic values . Moreover, binding free energies have been estimated for individual sugar-binding subsites using a purely kinetic approach for (GlcNAc)n interactions with both a family GH18 chitinase  and a family GH19 chitinase . The former yielded −3.8, +3.1, −2.5, −3.0, +0.8 and −1.8 kcal·mol−1 for the −2 to +4 subsites, respectively, while the latter gave −0.4, −4.7, +3.4, −0.5, −2.3 and −1.0 kcal·mol−1 for the −3 to +3 subsites, respectively. It is interesting to observe that the two subsites outside −2 to +2 of the latter (family GH19 enzyme) contribute with a free energy change upon binding of −1.4 kcal·mol−1, which is close to the −1.9 kcal·mol−1 observed in our work. Moreover, it is apparent that binding to the −1 subsite for both family GH19 and family GH18 chitinases is associated with a positive free energy change that is most probably due to the necessary distortion of the sugar bound in subsite −1.
Figure 5. Binding modes and binding free energy changes of chitin oligosaccharides to a family GH19 chitinase from the moss Bryum Coronatum.
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Zolotnitsky et al.  demonstrated that the ΔCp values can be used to evaluate the contribution of stacking-hydrophobic interactions to xylosaccharide binding to a family GH10 xylanase. They estimated that one aromatic residue located in the catalytic cleft of the enzyme contributes about −100 to −150 cal·K−1·mol−1 to ΔCp. The change in heat capacity (ΔCp = ΔΔH/ΔT) for (GlcNAc)6 binding to BcChi-A-E61A was −105 ± 8 cal·K−1·mol−1, possibly suggesting that an aromatic residue of BcChi-A contributes to binding through a hydrophobic interaction with (GlcNAc)6. In fact, as shown in Fig. 6, one tryptophan residue (Trp103) is found near the catalytic residue Glu61 and its side chain is surface exposed within the putative catalytic cleft of the modeled three-dimensional structure of BcChi-A. Our result is consistent with the participation of this residue in substrate binding. This tryptophan residue appears to be similar to Trp62 of hen egg white lysozyme in its localization (Fig. 6). Trp62 is involved in substrate binding of the lysozyme, and significantly contributes to the enzymatic activity . It is likely that Trp103 is involved in the substrate binding of BcChi-A in a similar manner to that of Trp62 of the lysozyme. To confirm this hypothesis and to obtain more insight into the substrate binding mechanism of BcChi-A, a crystal structure determination of the BcChi-A–(GlcNAc)n complex is currently in progress.
Figure 6. The three-dimensional model of BcChi-A (light green) was superimposed on the crystal structure of hen egg white lysozyme (cyan) in complex with (GlcNAc)6 (grey) (PDB: 1sfb)  (using just the proton donor residues, Glu61 in BcChi-A and Glu35 in hen egg white lysozyme, as reference) with the Pair Fitting wizard of the molecular visualization program pymol.
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