Assessment of the properties of chitin deacetylases showing different enzymatic action patterns

Journal of Molecular Graphics and Modelling 88 (2019) 41e48

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Assessment of the properties of chitin deacetylases showing different enzymatic action patterns Diana Larisa Roman a, Marin Roman a, Havard Sletta b, Vasile Ostafe a, Adriana Isvoran a, * a

Department of Biology - Chemistry and Advanced Environmental Research Laboratories, Faculty of Chemistry, Biology, Geography, West University of Timisoara, Timisoara, Romania b SINTEF, Department of Biotechnology, Trondheim, Norway

a r t i c l e i n f o

a b s t r a c t

Article history: Received 25 October 2018 Received in revised form 6 January 2019 Accepted 7 January 2019 Available online 9 January 2019

Chitin deacetylases are a group of enzymes catalysing the conversion of chitin to chitosan. Obtaining chitosan with established deacetylation degree and pattern is important for biomedical and biotechnological applications. Understandings of the structural properties of chitin deacetylases and the specificity of their interactions with chitin may conduct to the control of the pattern of deacetylation of chitosan. Our study is focused on the characterization and comparison of the structural and physicochemical properties of chitin deacetylases from fungi and marine bacteria. Despite the low sequences identity for the investigated chitin deacetylases, there are amino acids belonging to their active sites that are strongly conserved. Moreover, they reveal an increased structural similarity of their catalytic domains, reflecting the common biological function of these enzymes. The studied enzymes present dissimilar local physicochemical properties of their catalytic cavities that could be responsible of their distinct deacetylation patterns. Molecular docking studies reflect that deacetylation efficiency is also distinct for the chitin and partially deacetylated chitin oligomers and that N-acetylglucosamine units and some partially deacetylated chitin oligomers could have inhibitory effect against chitin deacetylases belonging to fungi and marine bacteria. © 2019 Elsevier Inc. All rights reserved.

Keywords: Chitin deacetylases Deacetylation pattern Structure comparison Physiochemical properties Molecular docking Catalytic cavities

1. Introduction Chitin deacetylases (CDAs; EC 3.5.1.41) represent the group of enzymes that can catalyse the hydrolysis of the acetamido group in N-acetylglucosamine (GlcNAc) units in chitin resulting in the formation of chitosan by chitin deacetylation. CDAs belong to carbohydrate esterase family 4 (CE-4s) and specifically bind a sequence of four GlcNAc units in the substrate, one of which suffers deacetylation and consequently, the resulting chitosan has a more regular deacetylation pattern than a chitosan obtained by treating chitin with hot NaOH [1]. Thus, controlled enzymatic deacetylation of chitin by CDAs could allow the tailoring of chitosans for applications within the biotechnological, biomedical and pharmaceutical market [2]. Chitin deacetylases from fungi and marine bacteria have been characterized. Fungal CDAs are classified into two groups: (i) the intracellular CDAs secreted in periplasm of the cell of the Mucor

* Corresponding author. E-mail address: [email protected] (A. Isvoran). https://doi.org/10.1016/j.jmgm.2019.01.002 1093-3263/© 2019 Elsevier Inc. All rights reserved.

rouxii and Absidia coerulea [3,4] and (ii) extracellular CDAs secreted in the culture media by Aspergillus nidulans (AnCDA) [5] and two strains of Colletotrichum lindemuthianum (ClCDA) [6,7]. The marine Vibrio sp. have as core activity the metabolism of chitin, being able to inhabit the chitin particles and decompose them using the concerted action of both chitinases and chitin deacetylases [8]. Chitin deacetylase secreted by A. nidulans and Vibrio cholerae (VcCDA) are not effective on crystalline chitin, but re-precipitated, glycolated, or depolymerized chitins with small particle size are good substrates [1]. Protein Data Bank (PDB) [9] presently includes only 13 structures of CDAs belonging to marine bacteria Vibrio parahaemolyticus (VpCDA, 1 file), V. cholerae (8 files) and Arthrobacter SP. AW19M34-1 (ArCDA, 2 files), respectively to fungal organisms C. lindemuthianum (1 file) and A. nidulans (1 file). Characteristics of these structural files are presented in Table 1. Kinetic data concerning the activity of CDAs against the oligomers of chitin (GlcNAc)n ¼ 2e6 belonging to these organisms have been extracted from BRENDA database [10] and specific literature. These data illustrate that investigated CDAs reveal dissimilar kinetic properties against (GlcNAc)n¼2-6. ClCDA, AnCDA and ArCDA

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D.L. Roman et al. / Journal of Molecular Graphics and Modelling 88 (2019) 41e48

Table 1 Characterization of the structural files of considered CDAs: Ca, Cs, Cl, Co, Mg, Na, Ni, Zn, GlcNAc are non-proteic parts present in the crystallographic structures. PDB code entry

Organism

Acronym

Ligands

Description

Resolution (Å)

2IW0 [11]

Colletotrichum lindemuthianum

ClCDA

Zn, Ca, acetate ion and phosphate ion

1.81 Å

2Y8U [12] 3WX7 [13] 4NY2 [14] 4NYU [14]

Aspergillus nidulans Vibrio parahaemolyticus Vibrio cholerae

AnCDA VpCDA VcCDA

Co, Na, Cl and phosphate ion Ca, Zn, acetate ion and glycerol Zn, Ca, acetate ion and glycerol Zn, Ca, acetate ion and glycerol

residues 1-254 mutations: H125Q, V126L monomer residues 19e237 homodimer residues 26e427 homodimer residues 23e427 homodimer residues 23-427 monomer residues 23-427 homotetramer residues 23e427 mutation: D39S homodimer residues 23-427 mutation: D39S homodimer residues 23e427 homodimer residues 23e427 homodimer residues 23-427 mutation: D39S homodimer residues 42e241 monomer residues 41e239 monomer

4NYY [14]

Zn, Ca, acetate ion and ethanediol

4NZ1 [14] 4NZ3 [14]

Zn, Ca, Cl, GlcNAc and ethanediol Zn, Mg, GlcNAc and ethanediol

4NZ4 [14] 4NZ5 [14] 4OUI [14]

Zn, Ca, GlcNAc, glycerol and acetate ion Na, Cs, GlcNAc, glycerol and acetate ion Zn, Mg, (GlcNAc)3 and ethanediol

5LFZ [15] 5LGC [15]

Arthrobacter SP. AW19M34-1

ArCDA

Ni ion (GlcNAc)2

are active towards several substrates including (GlcNAc)2e6 but expose higher activity against (GlcNAc)5 revealing that they possess catalytic cavities adapted for longer substrates. There is not available kinetic data concerning the activity of VcCDA and VpCDA against longer oligosaccharides (n ¼ 5e6), but these CDAs reveal improved kinetic properties for shorter oligosaccharides (n ¼ 2e4) than ClCDA, AnCDA and ArCDA. The structures of ClCDA and AnCDA correspond to single compact catalytic domains with a deformed (b/a)8 fold that is similar to the TIM barrel structure and includes a His-His-Asp metal-binding triad (His97, His101, Asp48 in AnCDA and His104, His108 and Asp50 in ClCDA), a catalytic acid (His197 in AnCDA, HIS206 in ClCDA) and a catalytic base (Asp47 in AnCDA, Asp49 in ClCDA) [11,12]. Both the VcCDA and the VpCDA structures include one catalytic domain and two carbohydrate binding domains. Each catalytic domain consists in a deformed (b/a)7 barrel and includes the metal binding triad (His93, His97, Asp36 in VpCDA, His97, His101 and Asp40 in VcCDA), the catalytic acid (His201 in VpCDA, His205 in VcCDA) and the catalytic base (Asp35 in VpCDA, Asp39 in VcCDA) [13,14]. ArCDA reveals a deformed (b/a)8 structure that includes the catalytic triad His105-His109-Asp56 and Asp 55 as the catalytic base [15]. There are some distinct features of the structures of these enzymes that may be responsible for their distinct kinetic efficiency against different chitin oligosaccharides. Andres E. et al., 2014 [14] proposed that the substrate specificity of distinct CDAs is governed by the presence/absence of the individual loops surrounding the catalytic sites and that the flexibility of these loops is also important. Recently, the influence of the length of these loops for CDAs specificity has been shown [16, 17]. The CDAs catalytic abilities have proven to be strongly influenced (with the effect depending on the substrate nature) by divalent cations [1] such as Zn2þ, Ca2þ Co2þ and Mg2þ and they are present in the crystallographic structures of enzymes investigated in this study. Also, many of structures contain the acetate ions, the acetate ion being one of the products of the deN-acetylation reaction. The aim of this study is to compare the global and local structural properties of chitin deacetylases belonging to fungal organisms (C. lindemuthianum, A. nidulans) and to bacterial organisms (V. parahaemolyticus, V. cholerae, Arthrobacter SP. AW19M34-1). We use a molecular docking approach to investigate the interactions of

1.99 Å 1.35 Å 1.88 Å 2.03 Å 2.65 Å 2.05 Å 2.11 Å

1.94 Å 1.87 Å 2.17 Å

1.56 Å 2.09 Å

these enzymes with chitin oligosaccharides containing distinct numbers of GlcNAc units in order to improve the knowledge about the use of chitin deacetylases from fungus and bacteria to produce chitosan. Understandings of the structural properties of these functionally different enzymes and the specificity of their interactions with chitin may lead to the control of the pattern of acetylation of the enzymatic deacetylation products during the enzymatic conversion from chitin to chitosan. The outcomes of this study are important as the use of fungal and bacterial chitin deacetylases for the controlled conversion of chitin to chitosan is prevalent in numerous industrial areas, from biotechnology and biomedicine to cosmetics and pharmaceutics. 2. Methods In order to achieve the structural bioinformatics characterization of chitin deacetylases belonging to fungal organisms and marine bacteria respectively, we used their structural files deposited in the Protein Data Bank (PDB) [9] (Table 1) and extracted their sequences and structures. When only one crystallographic structure has been available in PDB, we have considered it for analysis: 2IW0 for ClCDA, 2Y8U for AnCDA and 3WX7 for VpCDA, respectively. In the case of VcCDA, there are numerous structural files and we have considered in this study the structural file 4NZ4 corresponding to the enzyme in complex with GlcNAc. For ArCDA we have considered the structural file of its complex with (GlcNAc)3 having the PDB ID 5LGC. For VpCDA and VcCDA we have only taken into consideration the structures and sequences corresponding to their catalytic domains: residues 23e331 for VpCDA and residues 27e334 for VcCDA. The extracted sequences have been used for the multiple sequence alignment and the generation of the phylogenetic tree using ClustalOmega software [18]. Analysis of the structures and surfaces characteristics have been performed using: (i) Chimera software [19] for the structures superposition and geometric properties; (ii) PDBFlex [20] for revealing the intrinsic flexibility of the CDAs structures (iii) Fpocket tool [21] for identifying and characterizing the catalytic cavities of investigated enzymes; (iv) CASTp software [22] for computing the volumes and solvent accessible surface area for different probe sphere radii for the catalytic sites. Fpocket tool can be used to compute the polarity score of a

D.L. Roman et al. / Journal of Molecular Graphics and Modelling 88 (2019) 41e48

cavity as a sum of polarity over all residues involved in that cavity using a binary scheme, 1 for polar, 0 for nonpolar. It also calculates the hydrophobicity score based on Monera's residue hydrophobicity scale [23] while the mean hydrophobicity score is calculated for all residues involved in the cavity. Fpocket tool is utilized for computing the charge score as the mean charge for all amino acids in the binding site. In order to illustrate local hydrophobic area in the detected cavity, the local hydrophobic density is computed by Fpocket. For each nonpolar probe sphere, the number of nonpolar sphere neighbours is detected by seeking for overlapping nonpolar spheres. The sum of all nonpolar alpha sphere neighbours is further divided by the total number of nonpolar spheres in the pocket and this score is normalized compared to other identified cavities [21]. The concepts of the fractal geometry [24,25] have been applied to quantitatively characterize the surface roughness for the catalytic cavities respectively for every investigated enzyme. Moreover,

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for the multimeric structures, we only took into consideration one monomer, the A chain. Molecular docking studies were performed to assess the interactions of investigated CDAs with (GlcNAc)n¼2-8 chitin oligomers and with partly deacetylated chitin oligomers, (GlcN-GlcNAc)n¼1-4. They were implemented using both SwissDock webserver [26] that uses the EADock algorithm [27] to compute the pairwise interaction energy between the ligand and the protein and PatchDock server [28] that is based on the surface patch matching and ranks complexes considering the geometric shape complementarity score. When using SwissDock server we have considered blind, accurate and rigid docking and 5000 orientations have been generated. In the case of PatchDock, blind docking and 500 orientations are generated and FireDock [29] refinement and re-scoring of complexes have been used. Preparation of the enzymes structures for the molecular docking by adding hydrogens and charges as well as visualization and analysis of the molecular docking results have been performed using Chimera. The predicted ligandprotein complexes are superimposed to the crystallographic structures. The binding modes (BMs) are selected such as to correspond to the natural ligand binding region and taking into account the catalytic specificity of every enzyme. Among these BMs, the particular BM having the most negative Gibbs free energy is considered.

3. Results and discussions

Fig. 1. The amino acid sequence based phylogenetic tree of investigated CDAs.

The phylogenetic tree generated from the sequences of investigated enzymes is presented in Fig. 1. As mentioned in the Methodology we have considered only the catalytic domains for VpCDA

Fig. 2. Sequence alignment of the catalytic domains of investigated CDAs. The catalytic residues are shown in italics, the metal ion binding residues in bold, the residues belonging to the loops surrounding the catalytic sites are highlighted in light grey and the conserved motifs are indicated in rectangles. The consensus of the secondary structure elements is presented: H e helix, E  beta sheet. Conserved residues are marked by the symbol ”*”, residues with strongly similar physicochemical properties are marked by ”:” and residues reflecting weakly similar properties are marked by ”.”

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D.L. Roman et al. / Journal of Molecular Graphics and Modelling 88 (2019) 41e48

Table 2 Structural similarity between the catalytic domains of considered CDAs structures. CDA 1

Total number of residues

CDA 2

Sequence identity (%)

Total number of residues

Number of aligned residues

RMSD (Å)

ClCDA

223

AnCDA

208

VpCDA

305

VcCDA

305

AnCDA VpCDA VcCDA ArCDA VpCDA VCCDA ArCDA VCCDA ArCDA ArCDA

42 25 24 32 24 25 34 84 25 24

208 305 305 201 305 305 201 305 201 201

190 94 102 158 70 52 153 305 57 57

0.80 1.00 1.01 0.90 0.92 0.90 0.91 0.35 0.85 0.86

(residues 23e331) and VcCDA (residues 27e334). CDAs belonging to Vibrio sp are the most closely related and the highest divergence is presented between these CDAs compared to those belonging to fungal organisms. Fig. 2 shows the multiple sequence alignment as well as the secondary structure elements for the catalytic domains of the investigated CDAs. The catalytic residues are shown in italics and the metal ion binding residues in bold. The residues belonging to the loops surrounding the catalytic sites have been identified by structures superposition using Chimera and are highlighted in light grey and the five conserved motifs are highlighted in rectangles. The multiple sequence alignment of the investigated CDAs shows a low sequence identity, as it is illustrated in Table 2. Despite the low sequences identity, residues involved in the metal ion binding and in the catalytic activity belong to the sequence motifs that are strongly conserved and are characteristics for the carbohydrate esterase family [11]. First sequence motif, T(F/ Y)DD, is found in all the investigated sequences and includes two aspartic acid residues, one belonging to the metal binding triad and the other having catalytic activity and interacting with the acetate ion resulting from the catalytic process. Second motif, HSYDHM, totally conserved among sequences of investigated CDAs, contains two histidine residues that are involved in the metal ion binding and in stabilizing the loop forming the groove of the active site. Third motif, R(L/P/A)PY reveals one residue that differs between the investigated enzymes: leucine residues in the cases of VpCDA and

Fig. 3. Superposition of the structures of the catalytic domains of: (A) VcCDA (black) and VpCDA (dark grey); (B) AnCDA (black), ClCDA (dark grey), and ArCDA (light grey); (C) VcCDA (black) and ClCDA (dark grey).

VcCDA, phenylalanine residues in the cases of AnCDA and ArCDA and alanine residue in the case of ClCDA. Motif 4 is the most different in amino acids composition among investigated enzymes, there is only one aspartic acid that is conserved within this motif. Motifs 3 and 4 are usually forming the two opposite sides of the catalytic grove [11e15] and their distinct compositions in amino acids among considered enzymes may influence the catalytic specificity. Inside motif 5, two residues are totally conserved, a leucine and a histidine, the last one acting as the catalytic base. The structural similarity of the catalytic domains of investigated structures has been determinated by structural alignment using the Chimera computational tool and is shown in Table 2. Structural alignment is built through the superimposition of the atomic coordinate sets of the considered proteins and a root mean square deviation (RMSD) value between any two structures is computed. There are distinct possible subsets of the protein atoms that can be considered when generating a structural alignment and computing the RMSD values, but usually the alpha carbon (CA) positions are taken into account [30]. The RMSD value is a measure of the degree of divergence of two protein structures, a high RMSD value meaning dissimilar structures. Table 2 shows that the catalytic domains of VpCDA and VcCDA reveal high structural similarity in concordance with their high sequences identity. Fig. 3a illustrates

Fig. 4. The structural flexibility of the region 230e249 of VcCDA containing the loop 4 (amino acids 234e249): (A) Visualization of this loop (dark grey) in the structure of VcCDA displayed as solid surface (light grey). The ligands are also shown: acetyl ion in dim grey, Zn ion in dark grey sphere and GlcNAc in black sticks; (B) Average local RMSD variations between different structures of VcCDA revealing that the region 230e249 containing this loop reflects an important structural flexibility; (C) Rearrangement of the side chains of residues PRO235 and TRP238 in the loop 4 of VcCDA when it forms the complex with the acetate ion (code entry 4NYY, black) in comparison to its complex with GlcNAc (PDB code entry 4NZ4, dark grey).

D.L. Roman et al. / Journal of Molecular Graphics and Modelling 88 (2019) 41e48

the superposition between the structures of the catalytic domains of VcCDA and VpCDA. Despite quite low sequence identity between the sequences of the catalytic domains of AnCDA, ArCDA and ClCDA (maximum 42%), their structures show a good structural similarity (Fig. 3b). There is a low structural similarity between the structures of the catalytic domains of CDAs belonging to Vibrio sp. by comparison with the structures of the catalytic domains of the other considered CDAs. Fig. 3c shows the superposition of the structures of the catalytic domains of ClCDA and VcCDA, these structures revealing the highest structural dissimilarity. These results show a good concordance with the results given by the phylogenetic tree. Previous published data shows that the distinct deacetylation patterns exhibited by different CDAs could be cauzed by a loop that has the ability to block the accessible subsites in the binding cavity [11e14]. In the case of VcCDA, the loop 4, containing the residues 234e249, regulates the accessibility of the active site cavity, as shown in Fig. 4a [14]. PDBFlex tool [20] has been used to identify regions of structural flexibility present in the structure of VcCDA. This analysis reveals that the region 230e249 (DVDWAPENWGIPMPANSLTE) containing this loop reflects an important structural flexibility when VcCDA interacts with distinct ligands (Fig. 4b), the two hydrophobic residues in this loop, PRO235 and TRP238 (that is also involved in the interaction with GlcNAc) undergoing important structural rearrangements and modifying the accessibility to the active site (Fig. 4c). The structural files of CDAs show that the enzymes have open active-site structures with few subsites. ClCDA possesses four subsites denoted 2, 1, 0 and þ 1 with the subsite 0 being the catalytic site and subsite þ1 having the larger contribution to the energy of binding [11]. AnCDA illustrates three subsites denoted 1, 0 and þ 1 with the subsite þ1 being important for the effective catalysis [12]. ArCDA structure suggests that there are only two clear subsites, 0 and þ 1 that are involved in the catalytic cavity of this enzyme with a strong binding interaction in the þ1 subsite,

Fig. 5. The catalytic cavity of the VcCDA identified using Fpocket tool displayed as grey continuous surface superposed to the structural file of the enzyme (dim grey carton). The ligands that are present in the structural file of the enzyme are also displayed: GlcNAc (black sticks), acetyl ion (dim grey sticks) and Zn ion (dark grey sphere).

45

but 1 and 2 subsites could be also present [15]. VpCDA and VpCDA also reveal three subsites, 1, 0 and þ 1, the accessibility of subsites 0 and þ 1 depending on the conformational change of the loop 4 that may close, respectively open the accessibility to the subsite þ1 when the protein interacts with distinct oligosaccharides [14]. Subsites denoted 2, 1, 0 and þ 1 refer to the glucose residues upstream and downstream to the one in the centre of the catalytic site (subsite 0). All this data implies that the catalytic cavities of investigated CDAs have different properties: VpCDA and VcCDA reveal narrow active site cavities while ClCDA, AnCDA and ArCDA exhibit shallow and more open active site cavities. Furthermore, kinetic data prove that CDAs show a broader substrate specificity, deacetylating several positions in chitin and/or chitosan. VpCDA and VcCDA act on shorter chain chitin oligosaccharides [31], while ClCDA, AnCDA and ArCDA present a higher activity against longer chain chitin oligosaccharides and on and GlcNAc polymer chitin [4,7]. We have identified the catalytic cavities of the considered enzymes using Fpocket tool. Fig. 5 illustrates the catalytic cavity of the VcCDA identified using Fpocket tool (grey surface) superposed to the structural file of the enzyme (dim grey cartoon). We notice that Fpocket correctly identifies the catalytic cavity as it corresponds to the location of the ligands GlcNAc (black sticks), acetyl ion (dim grey sticks) and Zn ion (dark grey sphere).

Fig. 6. Distribution of the electrostatic potential (A), red regions having a negatively electrostatic potential and blue ones having positively electrostatic potential (from 10 kcalmol-1e1 for red and to þ10 kcalmol1e1 for blue) and of the hydrophobicity (B), blue regions are hydrophilic and orange regions are hydrophobic (dodger blue for the most hydrophilic residue to white at 0.0 and orange red for the most hydrophobic residue) on the surfaces of the catalytic cavities of investigated enzymes. The estimated locations of the subsites 1, 0 and þ 1 are illustrated.

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Our study also investigated the electrostatic and hydrophobic properties of the surfaces of the catalytic cavities of considered enzymes (subsites 1, 0, þ1). The molecular surfaces have been coloured based on the charges of the residues they enclose calculated using Columbic Surface Colouring facility under Chimera software where the solvent accessible surface potentials were set to 10 (red) and þ10 (blue) kcal$mol1$e1 and are shown in Fig. 6a. The hydrophobic properties of the surfaces of the catalytic sites are displayed using the Presets/Interactive 3 (hydrophobicity surface) facility of Chimera and are show in Fig. 6b. The hydrophobic surface is coloured based on the amino acid hydrophobicity in the Kyte-Doolittle scale [32] with blue for the most hydrophilic to orange for the most hydrophobic (dodger blue for the most hydrophilic residue to white at 0.0 and orange red for the most hydrophobic residue). Fig. 6 illustrates that the surfaces of the catalytic cavities reveal quite dissimilar electrostatic and hydrophobic properties. Excepting the ArCDA that presents a combined neutral-negative distribution of charges on the active site surface, the other investigated CDAs show negatively charged cavities. All investigated enzymes have cavities presenting both hydrophilic and hydrophobic regions, but ClCDA and AnCDA show local area with high hydrophobicity. In order to quantitatively characterize the electrostatic and hydrophobic properties of the active sites of these enzymes we have computed their charge, polarity and hydrophobicity scores and the local hydrophobic density using Fpocket tool. The surface areas and the volumes of the active site cavities (subsites 1, 0, þ1) are computed using CASTp computational tool. In order to

quantitatively characterize the roughness of the active site cavities, we have also used CASTp software to compute the accessible surface area of catalytic cavities using probe spheres with different radii and the approaches of the fractal geometry to subtract the surface fractal dimensions. All these data are presented in Table 3 and illustrate that the geometric properties of the catalytic cavities do not significantly differ, these cavities revealing quite similar areas, volumes and surface roughness. The high values of the surface fractal dimensions emphasize the significant complexity of the catalytic cavities. On the contrary, data presented in Table 3 confirm the distinct electrostatic and hydrophobic properties of the active site cavities of investigated enzymes. The catalytic cavity of AnCDA exposes the highest negatively charged surface, indicating that it could interact with deacetylated chitin oligomers. The possible influence of the charge distribution along the binding cavities of CDAs in determining their mode of actions and consequently the deacetylation patterns have been recently mentioned [16]. The hydrophobicity profiles of the catalytic cavities of investigated enzymes are also distinct, fungal CDAs revealing higher both global and local hydrophobicity for the subsites þ1 and the importance of hydrophobic interactions at subsite þ1 have been already noticed [11,12]. Binding of chitin and partially deacetylated chitin containing distinct numbers of units (n ¼ 2e8) to the CDAs has been analysed by molecular docking studies using both SwissDock and PatchDock followed by FireDock algorithms. The binding energies and the surface complementarity scores (SCS) are presented in Table 4 for chitin oligomers and in Table 5 for partially deacetylated chitin oligomers, respectively. We have considered partially deacetylated

Table 3 Geometric and physicochemical characteristic of the catalytic cavities (subsites 1, 0, þ1) of investigated enzymes. Enzyme

Surface area (Å2)

Volume (Å3)

Surface fractal dimension

Hydrophobicity score

Local hydrophobic density

Polarity score

Charge score

ClCDA AnCDA ArCDA VpCDA VcCDA

79.49 86.59 90.98 90.66 81.45

50.81 47.40 56.57 54.17 47.01

2.45 ± 0.08 2.82 ± 0.09 2.75 ± 0.02 2.50 ± 0.02 2.61 ± 0.05

13.29 14.64 10.14 9.79 5.03

31.17 27.39 35.16 22.94 27.03

10 9 9 17 24

2 3 1 3 2

Table 4 Interacting energies between CDAs and (GlcNAc)n¼2-8 units: SCS e surface complementarity score. ClCDA Substrate

(GlcNAc)2 (GlcNAc)3 (GlcNAc)4 (GlcNAc)5 (GlcNAc)6 (GlcNAc)7

SwissDock DG (kcal/mol) 6.79 PatchDock SCS 3596 FireDock DG (kcal/mol) 17.85 AnCDA Substrate

8.28 e e

6.02 4908 25.37

7.15 4776 25.82

8.58 5694 26.30

10.14 7844 35.57

8.21 6074 29.67

8.45 6302 30.92

9.17 7462 33.76

9.95 6940 19.52

6.97 6082 20.63

8.04 6146 48.15

7.37 e e

(GlcNAc)8 11.53 7610 28.86 (GlcNAc)8

(GlcNAc)7 (GlcNAc)8

The binding pocket is not large enough to accommodate the ligand.

(GlcNAc)2 (GlcNAc)3 (GlcNAc)4 (GlcNAc)5 (GlcNAc)6

SwissDock DG (kcal/mol) - 8.91 PatchDock SCS e FireDock DG (kcal/mol) e

(GlcNAc)8 The binding pocket is not large enough to accommodate the ligand. 7998 18.13

The binding pocket is not large enough to accommodate the ligand. 8298 7518 e 24.37 30.43 e

(GlcNAc)2 (GlcNAc)3 (GlcNAc)4 (GlcNAc)5 (GlcNAc)6

SwissDock DG (kcal/mol) 7.32 PatchDock SCS 4128 FireDock DG (kcal/mol) 23.79 VpCDA Substrate

10.94 6002 25.57

Substrate (GlcNAc)3 (GlcNAc)4 (GlcNAc)5 (GlcNAc)6 (GlcNAc)7

SwissDock DG (kcal/mol) 7.34 PatchDock SCS 4728 FireDock DG (kcal/mol) 23.19 VcCDA Substrate

8.13 5042 14.61

(GlcNAc)2 (GlcNAc)3 (GlcNAc)4 (GlcNAc)5 (GlcNAc)6 (GlcNAc)7

SwissDock DG (kcal/mol) 6.27 PatchDock SCS 3774 FireDock DG (kcal/mol) 16.21 ArCDA Substrate

10.06 4528 27.19

(GlcNAc)7 (GlcNAc)8

The binding pocket is not large enough to accommodate the ligand.

D.L. Roman et al. / Journal of Molecular Graphics and Modelling 88 (2019) 41e48

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Table 5 Interacting energies between CDAs and (GlcNAc-GlcN)n¼1-4 units. ClCDA

Substrate SwissDock PatchDock FireDock

AnCDA

SCS DG (kcal/mol)

DG (kcal/mol) SCS DG (kcal/mol)

Substrate SwissDock PatchDock FireDock

VpCDA

DG (kcal/mol)

Substrate SwissDock PatchDock FireDock

VcCDA

SCS DG (kcal/mol)

Substrate SwissDock PatchDock FireDock

ArCDA

DG (kcal/mol)

DG (kcal/mol) SCS DG (kcal/mol)

Substrate SwissDock PatchDock FireDock

DG (kcal/mol) SCS DG (kcal/mol)

GlcN-GlcNAc

(GlcN-GlcNAc)2

(GlcN-GlcNAc)3

5.32 3136 21.28

10.33 4674 37.69

11.14 The binding pocket is not large enough to accommodate the ligand. It does not bind in a correct orientation.

Substrate

(GlcN-GlcNAc)2

(GlcN-GlcNAc)3

6.38 3186 19.73

6.95 2986 13.03

6.12 7.58 It does not bind in a correct orientation

GlcN-GlcNAc

(GlcN-GlcNAc)2

(GlcN-GlcNAc)3

(GlcN-GlcNAc)4

6.80 4410 23.66

7.64 5620 32.11

9.61 7010 27.65

The binding pocket is not large enough to accommodate the ligand. It does not bind in a correct orientation

GlcN-GlcNAc

(GlcN-GlcNAc)2

(GlcN-GlcNAc)3

(GlcN-GlcNAc)4

6.24 4200 20.17

e 6236 26.00

It does not bind in a corect orientation

GlcN-GlcNAc

(GlcN-GlcNAc)2

(GlcN-GlcNAc)3

6.79 e e

The binding pocket is not large enough to accommodate the ligand.

Fig. 7. Results of the molecular docking study; (A) interaction of ClCDA shown as hydrophobicity surface with (GlcNAc)5 shown as cyan sticks, the acetyl ion being shown in yellow sticks and Zn ion in black sphere; (B) interaction of AnCDA shown as hydrophobicity surface with (GlcNAc)5 shown as cyan sticks and the Co ion in black sphere; (C) interaction of ArCDA shown as hydrophobicity surface with (GlcNAc)5 shown as cyan sticks, chitobiose being shown in dark blue sticks; (D) interaction of VcCDA shown as hydrophobicity surface with (GlcNAc)3 shown as cyan sticks, the acetyl ion being shown in yellow sticks, GlcNAc in dark blue sticks and Zn ion in black sphere; (E) interaction of VpCDA shown as hydrophobicity surface with (GlcNAc)3 shown as cyan sticks, the acetyl ion being shown in yellow sticks and Zn ion in black sphere. The hydrophobicity of the surfaces is coloured from dodger blue for the most hydrophilic residue to white at 0.0 and to orange red for the most hydrophobic residue. The estimated locations of the subsites 1, 0 and þ 1 are illustrated. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

(GlcN-GlcNAc)4

(GlcN-GlcNAc)4

(GlcN-GlcNAc)4

chitin with a regular pattern for deacetylation, every odd saccharide unit being deacetylated. When analysing the binding modes resulted from the molecular docking study we have taken into consideration the following: (i) ClCDA deacetylates to the nonreducing end from short chito-oligosaccharides only, occupation of more than two subsites (0 and þ 1) being important for activity and consequently, every second GlcNAc sugar in the oligosaccharides is de-N-acetylated [11]; (ii) AnCDA is able to deacetylate at random positions, but not at the reducing end [12]; (iii) VcCDA and VpCDA firmly deacetylate the sugar next to the non-reducing end [13,14]; (iv) ArCDA also reveals a less preferred deacetylation at the reducing end and occupation of more than two subsites (0 and þ 1) is favourable for activity [15]. Usually there is a good agreement between the results obtained using SwissDock server and those obtained using PatchDock and refined with FireDock. The outcomes of the molecular docking analysis confirm the literature data. ClCDA, AnCDA and ArCDA are able to bind longer chito-oligosaccharides (n ¼ 2e8) and are efficient to deacetylate (GlcNAc)5. VpCDA and VcCDA bind shorter oligosaccharides (n ¼ 2e3) and are efficient against (GlcNAc)3 such as subsite þ1 to be occupied. Molecular docking analysis also suggests that these enzymes are able to bind partially deacetylated chitin oligomers, but the binding energy is usually lower than that of the interactions with chitin oligomers. Taking into account that we have considered partially deacetylated chitin oligomers with every odd unit being deacetylated, these data also suggest the importance of the acetyl group for the interaction with the subsite þ1 for chitin deacetylation process. Fig. 7 illustrates the outcomes of SwissDock computational tool for the interactions of CDAs with chitin oligomers. Molecular docking studies prove that some partially deacetylated chitin oligomers are able to bind to the active sites of these enzymes but not in the orientation needed by the catalysis process. Furthermore, we have also considered molecular docking studies for assessing the interaction of GlcNAc unit with the investigated enzymes (data not shown). GlcNAc is able to bind to the active site of every enzyme. These outcomes illustrate the inhibitory potential of GlcNAc unit and of some partially deacetylated chitin oligomers against CDAs.

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4. Conclusions Here we have compared the structural properties of chitin deacetylases belonging to fungi and marine bacteria respectively, as all of them are used for biotechnological applications. Despite the small sequence identity between chitin deacetylases belonging to fungal organisms and those belonging to marine bacteria, their catalytic domains have similar structural properties and the residues involved in the catalytic processes are strongly conserved. There are some local differences between the physicochemical properties of the catalytic domains of investigated CDAs: (i) charge distribution inside the catalytic cavities and in their surrounding is distinct and (ii) the fungal CADs reveal higher hydrophobicity of the catalytic subsite þ1. These local distinct features, together with the length and flexibility of the loops composing the catalytic cavities, may be responsible of the specificity of these enzymes. Our results also imply that these enzymes are capable to bind and process partially deacetylated chitin oligomers, but with a lower efficiency, and this underlines the importance of the acetyl group for the interaction with the subsite þ1 for chitin deacetylation process. N-acetylglucosamine unit and some partially deacetylated oligomers are able to bind to the active sites of CDAs without leading to a deacetylation process and this illustrates their inhibitory potential against these enzymes. Considering the use of CDAs as biocatalysts to produce chitosan from chitin, knowledge concerning their structure-activity relationship opens opportunities for designing new biotechnological methods for production of chitosan with defined patterns of acetylation. Declarations of interest None. Funding This work was supported by the grant Biotechnological tools implementation for new wound healing applications of byproducts from crustacean seafood processing industry (ChitoWound) [PN3P3-284] and by the grant ‘Cercetare de excelenta in UVT prin sustinerea si dezvoltarea centrelor de cercetare ale universitatii’. References [1] Y. Zhao, R.D. Park, R.A.A. Muzzarelli, Chitin deacetylases: properties and applications, Mar. Drugs 8 (2010) 24e46. [2] V. Ghormade, S. Kulkarni, N. Doiphode, P.R. Rajamohanan, M.V. Deshpande, Chitin deacetylase: a comprehensive account on its role in nature and its biotechnological applications. Current Research, Technology and Education Topics, in: A. Mendez-Vilas (Ed.), Applied Microbiology and Microbial Biotechnology, 2010, pp. 1054e1066. [3] Y. Araki, E. Ito, A pathway of chitosan formation in Mucor rouxii: enzymatic deacetylation of chitin, Biochem. Biophys. Res. Commun. 56 (1974) 669e675. [4] D. Kafetzopoulos, A. Martinou, V. Bouriotis, Bioconversion of chitin to chitosan: purification and characterization of chitin deacetylase from Mucor rouxii, Proc. Natl. Acad. Sci. U.S.A. 90 (1993) 2564e2568. [5] C. Alfonso, O.M. Nuero, F. Santamaria, F. Reyes, Purification of a heat-stable chitin deacetylase from Aspergillus nidulans and its role in cell wall degradation, Curr. Microbiol. 30 (1995) 49e54. [6] I. Tsigos, V. Bouriotis, Purification and characterization of chitin deacetylase from Colletotrichum lindemuthianum, J. Biol. Chem. 270 (1995) 26286e26291. [7] K. Tokuyasu, M. Ohnishi-Kameyama, K. Hayashi, Purification and characterization of extracellular chin deacetylase from Colletotrichum lindemuthianum,

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