Protein engineering of GH11 xylanase from Aspergillus fumigatus RT-1 for catalytic efficiency improvement on kenaf biomass hydrolysis

Enzyme and Microbial Technology 131 (2019) 109383

Contents lists available at ScienceDirect

Enzyme and Microbial Technology journal homepage: www.elsevier.com/locate/enzmictec

Protein engineering of GH11 xylanase from Aspergillus fumigatus RT-1 for catalytic efficiency improvement on kenaf biomass hydrolysis

T

Siti Intan Rosdianah Damisa, Abdul Munir Abdul Muradb, Farah Diba Abu Bakarb, ⁎ Siti Aishah Rashida, Nardiah Rizwana Jaafara, Rosli Md. Illiasa,c, a

School of Chemical and Energy Engineering, Faculty of Engineering, Universiti Teknologi Malaysia, 81310 Skudai, Johor, Malaysia Centre for Biotechnology and Functional Food, Faculty of Science and Technology, Universiti Kebangsaan Malaysia, 43600 Bangi, Selangor, Malaysia c Institute of Bioproduct Development, Universiti Teknologi Makaysia, 81310 Skudai, Johor, Malaysia b

A R T I C LE I N FO

A B S T R A C T

Keywords: Xylanase Kenaf Directed evolution Catalytic efficiency Active site N-terminal region

Enzyme hydrolysis faces a bottleneck due to the recalcitrance of the lignocellulose biomass. The protein engineering of GH11 xylanase from Aspergillus fumigatus RT-1 was performed near the active site and at the Nterminal region to improve its catalytic efficiency towards pretreated kenaf (Hibiscus cannabinus) hydrolysis. Five mutants were constructed by combined approaches of error-prone PCR, site-saturation and site-directed mutagenesis. The double mutant c168 t/Q192H showed the most effective hydrolysis reaction with a 13.9-fold increase in catalytic efficiency, followed by mutants Y7L and c168 t/Q192 H/Y7L with a 1.6-fold increase, respectively. The enhanced catalytic efficiency evoked an increase in sugar yield of up to 28% from pretreated kenaf. In addition, mutant c168 t/Q192 H/Y7L improved the thermostability at higher temperature and acid stability. This finding shows that mutations at distances less than 15 Å from the active site and at putative secondary binding sites affect xylanase catalytic efficiency towards insoluble substrates hydrolysis.

1. Introduction Malaysia contributes at least 168 million tonnes of biomass annually, which includes timber and oil palm waste, rice husks, coconut trunk fibres, municipal waste and sugar cane waste [1]. This abundant biomass can be used as feedstocks for bio-conversion into value-added bio-products such as pulp and paper, bio-fuel, composites, fine chemicals, enzymes and animal feed [2]. Biofuel offer numerous advantages related to energy security, economics and environment [3]. Great attention has been given to biobutanol as biofuel due to its superior properties as an alternative gasoline over bioethanol. As second generation biofuels, biobutanol has better characteristics such as less volatile and explosive, high flash point and energy density, possesses lower vapour pressure and easy in distribution and transportation [4,5]. Acetone-butanol-ethanol (ABE) fermentation involving solvent–producing bacteria is a promising sustainable process for the production of biobutanol. However, there are bottlenecks that need to be improved in the production of biobutanol by ABE fermentation [5]. These include sustainable and cost-effective substrate [3,5], suitable microbial strain and tolerance to butanol toxicity [6], fermentation and downstream processes [4]. Lignocellulosic biomass is a promising source of

fermentable sugar for butanol production. Nonetheless, the main challenge lies in the hydrolysis of lignocellulosic biomass to be converted into fermentable sugars [4]. Thus, the developments of efficient hydrolysis enzymes with improve properties such as stability, specific activity and interaction towards low cost substrate are crucial for industrial production of biobutanol. One of the biomasses, kenaf (H. cannabinus), has potential for bioenergy production, such as bioethanol, biodiesel, biohydrogen, solid fuel and biogas [1]. The degradation of lignocellulose into monosaccharide sugars involves two main constituents: cellulose and hemicellulose [7–9]. The highly complex structure of hemicellulose requires the synergistic action of various enzymes, and endo-1,4-β-xylanase is the ‘main player’ to degrade the most prevalent polymer of hemicellulose, xylan, into linear xylooligosaccharides [10–12]. Hemicellulase enzymes are naturally inefficient to hydrolyse insoluble lignocellulosic biomass owing to their ‘biomass recalcitrance’ [13]. Thus lignocellulosic biomass hydrolysis necessitates expensive enzymes, which could only result in relatively low sugar yields [14]. As a solution, one can increase the efficiency of the enzyme [15] and exploit accessory enzymes, such as xylanase and β-glucosidase, to be used synergistically with cellulases [16]. Recently, improvements of

⁎

Corresponding author. E-mail addresses: [email protected] (S.I.R. Damis), [email protected] (A.M.A. Murad), fabyff@ukm.edu.my (F. Diba Abu Bakar), [email protected] (S.A. Rashid), [email protected] (N.R. Jaafar), [email protected] (R.Md. Illias). https://doi.org/10.1016/j.enzmictec.2019.109383 Received 3 January 2019; Received in revised form 30 June 2019; Accepted 16 July 2019 Available online 17 July 2019 0141-0229/ © 2019 Elsevier Inc. All rights reserved.

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Then, 5% (v/v) overnight culture was added to 400 μl fresh LB medium with 100 μg/ml ampicillin in 96-well Masterblock®, U-bottom (Greiner Bio-One, Kremsmünster, Austria), followed by 1 h incubation at 37 °C and 200 rpm before induction with 0.5 mM IPTG. Protein expression was continued for 24 h at 30 °C with a 200 rpm agitation. Subsequently, a 250 μl cell suspension was transferred into a new 96-well Masterblock®, U-bottom (Greiner Bio-One, Kremsmünster, Austria) containing an equal amount of pretreated kenaf, which was transferred previously using MultiScreen Column Loader (Merck Millipore, Massachusetts, United States) and suspended with 250 μl of 50 mM sodium phosphate buffer, pH 5.0. After 4 h of incubation at 50 °C in a water bath, the reaction was centrifuged at 4 °C, 4000 rpm for 30 min, and 100 μl of the reaction was transferred to a 96-well microplate, F-bottom (Greiner Bio-One, Kremsmünster, Austria) and added with 100 μl of DNS in each well. Finally, after boiling for 5 min and cooling down, 100 μl of the mixture was transferred to a new 96-well microplate, F-bottom (Greiner Bio-One, Kremsmünster, Austria) to obtain reading measurements at 540 nm absorbance in Epoch Microplate Spectrophotometer (BioTek, Vermont, United States). Each screening plate contained blank (buffer and DNS), substrate control (pretreated kenaf, buffer and DNS) and a wild-type (AfxynG1) reaction. Enzyme activity was measured by the absorbance value at 540 nm, and the potential positive clones that retained more than 30% of enzyme activity relative to the control were selected from the library and kept as glycerol stocks.

hydrolytic enzymes towards lignocellulose biomass have focused only on cellulases and the synergetic action of cellulases and hemicellulases [17–19], but there are very few studies concerning the performance of xylanases alone. Directed evolution involving library creation strategies of either mutagenesis (random or semi-rational) or gene recombination has become the preferred engineering approach to generate genetic diversity in enzyme variants [20]. Random mutagenesis as a ‘blind watchmaker’ is ‘fine-tuned’ by site-saturation mutagenesis [21–23] in alterations of enzyme properties, such as substrate specificity, product specificity, selectivity, activity, stability or folding/solubility and specifically enzymatic turnover number or catalytic efficiency [21]. For instance, the combination of N-terminal region replacement and sitedirected mutagenesis at the cord of xylanase from Streptomyces rochei L10904 significantly improved the specific activity (5.3-fold increase), substrate affinity and catalytic efficiency [24]. Another study showed that the fusion of a carbohydrate binding module from GH 6 to GH 11 xylanase of Bacillus subtilis enhanced the catalytic efficiency by 65% and led to a 17% increase in sugar release from pretreated sugarcane bagasse hydrolysis [25]. Additionally, error-prone PCR (epPCR) mutagenesis in combination with site-saturation mutagenesis at the H179 residue improved the kcat/Km of xylanase by 3.46-fold [26]. In a recent study, the GH11 xylanase from a locally isolated Aspergillus fumigatus RT-1, AfxynG1 (GenBank accession no: GQ458016) showed immerse thermostability by retaining 70% of its activity after 30 min incubation at 70 °C [27]. Hence, this study aims to add value to this industry-potential xylanase by improving its catalytic efficiency towards lignocellulose biomass hydrolysis. AfxynG1 residues with the distance of less than 15 Å from the active site and at the N-terminal region were mutated by site-saturation and site-directed mutagenesis. As a result, three mutants showed improvements in catalytic efficiency by exhibiting higher sugar yields from pretreated kenaf. The increased catalytic efficiency of these mutants has strong potential in hydrolysing biomass hydrolysis for broad area of industrial application.

2.3. Low-throughput screening All variants with more than 30% improvement (in terms of absorbance OD at 540 nm) from the high-throughput screening were confirmed through low-throughput screening. The glycerol stock was inoculated into 10 ml LB supplemented with 100 μg/ml ampicillin and incubated overnight at 37 °C. Then, 100 ml fresh LB with 100 μg/ml ampicillin was inoculated with 5% overnight culture and grown until the OD600 reached ˜0.6 before inducing with 0.5 mM IPTG and further incubated at 30 °C for 24 h. The culture was harvested at 8000 rpm for 10 min, and the supernatant was transferred to 50 ml flask containing with 2% (w/v) pretreated kenaf and 50 mM sodium acetate buffer, pH 5.0. The reaction was incubated at 50 °C for 4 h in a water bath. Then 1 ml of the reaction was harvested by centrifugation and 750 μl was pipetted out and mixed with 750 μl DNS in a new microcentrifuge tube, followed by boiling for 5 min and cooling down prior to 540 nm OD measurement. The amount of sugar produced per gram substrate was determined according to the xylose standard graph (mg/ml):

2. Materials and methods 2.1. Reagents, bacterial strains, plasmids and substrates Recombinant xylanase construct AfxynG1 and the epPCR libraries were procured from a previous study [27], which are deposited in Genetic Lab, Universiti Teknologi Malaysia, Johor, Malaysia. DNA polymerase and PCR reagents were purchased from Novagen, Merck (Darmstadt, Germany). DMSO, prestained protein ladder for Western blot analysis and a gel extraction kit were purchased from Thermo Fisher Scientific (Massachusetts, United States). The dried-ground kenaf stem was kindly supplied by a kenaf processing company in Bachok, Kelantan (North-East Malaysia) and was prepared for a two-stage process of alkaline-acid pretreatment [28]. Beechwood xylan and xylose were purchased from Sigma-Aldric, Merck (Missouri, United States). DNA ladder and unstained protein ladder were acquired from New England Biolabs (Massachusetts, United States). Escherichia coli BL21 (DE3) and E. coli JM109 were used as expression hosts and cloning hosts, respectively. All analytical grade chemicals for growth and expression media, DNS assay and protein purification buffer were obtained from Merck (Darmstadt, Germany); Thermo Fisher Scientific (Massachusetts, United States); and Fulka, Honeywell (New Jersey, United States). The plasmid extraction kit was originally obtained from Promega (Wisconsin, United States).

mg mg ⎞ vol reaction (ml) Amount of sugar produced ⎛⎜ × ⎟ = x ml amount of substrate (g ) g ⎝ ⎠

Relativity of sugar production Amount of sugar produced by variants =

( )

Amount of sugar produced by wild type

mg g mg ( g)

2.4. D structure model, molecular docking and structural analysis The xylanase of Penicililum funiculosum (PfXynC; 1TE1) [30] was selected as template to construct the AfxynG1 model by Modeler 9v3. The model with the lowest value of DOPE score and the highest number of GA341 was selected to be energy minimized using GROMACS. Model verification was performed by ERRAT, Verify 3D, Anolea, QMEAN6 and DFire. The 3D structure of AfxynG1 and xylohexaose were subjected to protein-ligand docking by using an automated docking simulation of AutoDock 4.2 programme [31]. The structure visualisation and the distance measurement between atoms of residues were carried out by PyMol. Protein interaction between residues was analysed via Protein Interaction Calculator (PIC) [32].

2.2. High-throughput screening of epPCR and site-saturation libraries EpPCR and site-saturation libraries were screened by means of a high-throughput screening system according to a modified protocol [29]. Each variant from the mutant libraries was picked and incubated in separate wells of TPP® 96-well plates (Sigma-Aldrich Merck, Missouri, United States) containing 200 μl of LB and 100 μg/ml ampicillin. 2

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Table 1 Primers used for site-saturation and site-directed mutagenesis. Mutagenesis

Primers Namea

DNA Sequence (5'–3')b

Site-saturation

Y7X_F Y7X_R S75X_F S75X_R D182X_F D182X_R V186X_F V186X_R G56X_F G56X_R Q192X_F Q192X_R N136X_F N136X_R F146X_F F146X_R N54X_F N54X_R Q192H_F Q192H_R Y7L_F Y7L_R

CGGCTCGGAGCAANDTGTTGAGCTAGCC GGCTAGCTCAACAHNTTGCTCCGAGCCG GGTCACCTACNDTGGCTCCTGGCAGACC GGTCTGCCAGGAGCCAHNGTAGGTGACC GCAATTGGGGAACTTTNDTTATATGATTGTTGC GCAACAATCATATAAHNAAAGTTCCCCAATTGC GGAACTTTGACTATATGATTNDTGCGACGGAGG CCTCCGTCGCAHNAATCATATAGTCAAAGTTCC GGAACAACTGCNDTAACTTTGTTGCTGG CCAGCAACAAAGTTAHNGCAGTTGTTCC GGAGGGGTACNDTAGCAGCGGCTCTGC GCAGAGCCGCTGCTAHNGTACCCCTCC CGACGCGGACGNDTGCGCCGTCCATCC GGATGGACGGCGCAHNCGTCCGCGTCGTCTTGTAGAGG GCACGGCTACTNDTGACCAGTACTGG CCAGTACTGGTCAHNAGTAGCCGTGC GGTCGACTGGAACNDTTGCGGCAACTTTGTTGC GCAACAAAGTTGCCGCAAHNGTTCCAGTCGACC GGAGGGGTACCATAGCAGCGGCTCTGC GCAGAGCCGCTGCTATGGTACCCCTCC GCTCGGAGCAATTAGTTGAGCTAGC GCTAGCTCAACTAATTGCTCCGAGC

Site-directed

a b

Forward and reverse primers represented as F and R, respectively. For all primers, mutagenised positions are underlined.

acid (DNS) method [36]. All assays were performed in triplicate. The enzymatic assay consists of 375 μl of the enzyme and 375 μl 2% beechwood xylan (Merck, New Jersey, United States) in sodium acetate buffer (pH 5), which was incubated at 50 °C for 10 min. The reaction was terminated by adding 750 μl DNS and boiled for 5 min. The assay was measured at 540 nm. One unit of xylanase activity was defined as the amount of enzyme releasing 1 μmole of reducing sugar from xylan per min under the conditions described above (with xylose as the standard).

2.5. Construction of site-saturated and site-directed mutants A modified one-step PCR [33] was performed for mutant amplification. The recombinant plasmid AfxynG1 and silent mutant c168 t were used as templates for site-saturation and site-directed mutagenesis, respectively. Primers from pairs of complementary oligonucleotides containing the desired mutants were designed (Table 1). The 50 μl PCR reaction was carried out with ˜120 ng template, 1 μM primer pair, 200 mM dNTPs, 1 U of KOD Hot Start DNA polymerase, 5.0 μl PCR buffer, 1.0–4.0 mM MgSO4 and 5% DMSO. The PCR programme was set as follows: (i) 95 °C for 2 min, (ii) 12 cycles at 95 °C for 30 s, annealing temperature for 1 min, and synthesis at 70 °C for 6 min (1 min/ kb), (iii) 15 cycles at 95 °C for 30 s and 7 min of synthesis at 70 °C, omitting an individual annealing step to increase the specificity of primer template binding. After PCR, the amplified products were mixed with DpnI to digest the wild-type templates for 1 h at 37 °C, and the reaction was column-purified before transforming into E. coli BL21 (DE3). Randomly selected colonies of the transformants were isolated, and the plasmid extracts were analysed via sequencing. Available clones were analysed through high-throughput and low-throughput screening.

2.8. Characterization of AfxynG1 and mutants xylanase The optimum temperature for the enzyme was measured by performing the xylanase activity assay for 10 min at temperatures ranging from 30 to 70 °C at pH 5.0. The thermostability assay was conducted by 30 min incubation of the enzyme at the indicated temperatures and relative activities were measured under standard assay conditions. The effect of pH on the purified xylanase was evaluated under xylanase activity assay conditions at 50 °C within pH 3–8, using four different buffers at 50 mM concentration: glycine/HCl buffer (pH 3), sodium acetate buffer (pH 4–5) and Sørensen’s phosphate buffer (pH 6–8). The pH stability of xylanase was determined by a 30 min incubation at room temperature in different buffer systems (pH 3–8), and the relative activity was measured under standard assay conditions. The Km, kcat and kcat/Km values were determined by measuring the enzymatic activity using the substrate beechwood xylan at a concentration ranging from 0.5 to 8.0% in sodium acetate buffer pH 5 at 50 °C. The data were estimated according to the Lineweaver-Burk plot [37].

2.6. Expression and purification of wild-type AfxynG1 and mutants The wild-type AfxynG1 and mutants were expressed in E. coli BL21 (DE3) as fusion proteins with 6x-His tags. A single colony was grown overnight at 37 °C in 50 ml LB medium containing 100 μg/ml ampicillin. The overnight culture was mixed with 1 l of autoinduction medium [34] containing 100 μg/ml ampicillin, and the culture was further incubated for 24 h at 25 °C. The expressed enzyme was harvested by centrifugation (4000 rpm, 45 min, 4 °C) and concentrated using a Vivaspin® 15R, 10,000 MWCO ultrafiltration concentrator (Sartorius, Göttingen, Germany) to a 10-fold concentration. Then, the enzyme was purified using a Ni-NTA affinity through column (BioRad, California, United States). Protein purity was verified using SDS-PAGE and the Bradford method.

2.9. Biomass hydrolysis Hydrolysis reactions were carried out by 6 U of partially purified enzymes in 50 mM sodium acetate buffer (pH 5.0) in a 10 ml total volume containing 1% (w/v) pretreated kenaf in 30, 40 and 50 °C water bath at 200 rpm agitation. Aliquots were taken from the reaction mixture at regular intervals for 48 h, and the supernatants were collected by centrifugation at full speed for 5 min. The total reducing sugar from enzymatic reaction was determined as Sec 2.7 by the DNS method. All hydrolysis assays were performed in triplicate and the results are

2.7. Measurement of xylanase activity The xylanase activity was assayed as described previously [35]. The amount of sugars released was determined using the 3,5-dinitrosalicylic 3

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Fig. 1. Flowchart of the directed evolution process. The best-performing mutants at each round of evolution are boxed and the mutations acquired by each mutant are italicized. The bold residues were selected for site saturation mutagenesis libraries construction.

presented as mean values.

3. Results and discussion 3.1. Mutants isolation from epPCR library screening, site-saturation and site-directed mutagenesis Overall directed evolution process for this study was summarized in Fig. 1. Over 5000 clones of epPCR library from the previous study [27] were screened using a 96-well system for the hydrolysis of pretreated kenaf in a 50 °C water bath for 4 h. Mutants with an increase of at least 30% enzyme activity compared to the wild-type were double confirmed by low-throughput screening to eliminate any false-positive results. As a result, three mutants with improved activities relative to the wild-type, L_B12 (112%), E_G3 (105%) and C_F2 (116%), and 1 defective mutant, E_H3 (79%), were selected. Sequencing analysis of these four mutants revealed multiple mutations, as shown in Fig.1. Mutant L_B12 was mutated at one amino acid position (Y7N), mutant E_G3 acquired six mutations (Y7S, N54D, S75 N, N136 T, F146 L, and Q192-), mutant C_F2 showed alterations in five amino acids (L10 P, G56S, S75C, E121 V, and D182Y) and the last mutant, E_H3, showed changes in eight amino acids (G46D, L66 M, H116R, N128I, D169E, F168S, T164S, and V186D). In total, 18 residues were mutated, and further assessment was carried out to investigate which residues are important in enhancing the hydrolysis of pretreated kenaf. By focusing on the residues that might affect the enzyme activity, the first strategy was to select those located within the cleft of the active site. Residues that have a distance of 10–20 Å to the active site appeared to increase the efficiency of catalysis [38]. The model structure of AfxynG1 was generated using P. funiculosum xylanase (PfXynC; 1TE1) [30] as a template. As shown in Fig. 2, AfxynG1 exhibited an overall β-jelly roll shape, typical of GH11 xylanases and similar to a folded right hand, with several main parts: 1) ‘palm’ (the twisted part of the β-sheets and the α-helix with the concave face of the substrate binding site), 2) ‘fingers’ (formed by two β-sheets) and 3) ‘thumb’ (a long loop between two β-sheets that partly closes the cleft) [39,40]. The built model has an extra loop at the N-terminal region which was not covered by the template during the structure modelling due to the longer N-terminal region possessed by AfxynG1. The active site is situated in the extended open cleft, which contains two conserved glutamate residues. The carbonyl oxygens of the predicted catalytic residues, Glu 98 (catalytic nucleophile) and Glu 189 (catalytic acid–base), are located 5.0 Å apart between each other, which is consistent with the catalytic apparatus of a “retaining’’ glycoside hydrolase [41]. The distance of epPCR-mutated residues to the active sites was measured using PyMol (Table 2). By considering both of the mutated residues with a distance less than 15 Å to the catalytic sites and the mutation positioned in the important parts of the enzyme, a total of seven residues were selected: Asp 182, Val 186, Gly 56, Gln 192, Asn 136, Phe 146 and Asn

Fig. 2. Structure of AfxynG1 resembles ‘right hand folded’ with the important parts including ‘thumb’, ‘palm’ and ‘fingers’. Two catalytic residues (Glu 98 and Glu 189) are 5 Å distance between each other. Blue sticks represent extra loop at the N-terminal region. Table 2 Distance between of mutated residues and catalytic sites and their location. Catalytic residues

epPCR-mutated residuesa

Distance (Å)b

Location of residues

Glu 189

Asn 54 Gly 56 Gln 192 Val 186 Gly 46 Asp 182 Phe 146 Asn 136 Leu 66

14.3 8.3 11.6 11.9 19.7 11.7 2.4 13.5 19.9

Phe 168 Thr 164 Asp 169 Glu 121 Asn 128 His 116

11.8 15.1 21.3 27.5 18.6 18.9

Fingers Fingers Fingers Palm/cleft Knuckles Palm/cleft Closer to the thumb Thumb Coil at the entrance of the cleft Alpha-helix Alpha-helix Alpha-helix At the cord At the back of palm At the cord

Glu 98

a

Residues in bold is the one that selected for the site saturation mutagenesis. Distance (Å) was measured according to the closest distance between mutated residues and the active site. b

54. The second strategy applied was to choose the mutation that appeared concurrently in more than one mutant. In a previous study, the point mutation H179Y that shared by two mutants 1-B8 and 2-H6 had 4

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Table 4 Kinetic parameters of the wild-type AfxynG1 and its mutants. Variants

Km (mg ml−1)

kcat (s−1)

kcat/Km (ml mg−1s−1)

WT c168t Q192H Y7L c168 t/Q192H c168 t/Q192 H/Y7L

22.01 26.11 31.04 22.80 31.46 23.26

6.57 × 104 3.23 × 104 7.75 × 104 1.11 × 105 7.12 × 105 1.68 × 105

2987 1239 2496 4863 22633 4865

and purified by single-step purification for subsequent analyses. 3.2. Effects of mutation on catalytic activity The steady state kinetic parameters for the wild-type and mutants were determined at 50 °C, pH 5 by using different concentrations of soluble xylan (beechwood xylan) (0.5–8 % w/v) and plotted according to Lineweaver-Burk (Table 4). The Michaelis constant, Km values for the two mutants (Y7L and c168 t/Q192 H/Y7L) showed subtle changes of approximately 3.6–5.7% increase compared to the wild type, while the other mutants acquired higher Km values increased; c168 t (18.6%), Q192H (41.0%) and c168 t/Q192H (42.9%). None of the mutants showed any improvement in substrate affinity by having lower Km values. Based on the result, it can be suggested that the changes in both mutants Q192H and c168 t, which are located closer to the active site, reduced the substrate affinity except for c168 t/Q192 H/Y7L. The substitution of tyrosine by leucine at residue 7 did not affect the substrate affinity of the single mutant Y7L and might influenced the triple mutant (c168 t/Q192 H/Y7L). The turnover number kcat increased for mutants c168 t/Q192H (10.8-fold), c168 t/Q192 H/Y7L (2.6-fold), Y7L (1.7-fold) and Q192H (1.2-fold) except for the silent mutant c168 t. This indicated that the mutants performed better to convert most of the bound substrate into product [44] as compared to the wild-type and mutant c168 t. Furthermore, the synergistic effect of mutants c168 t and Q192H drastically enhanced the kcat value of the double mutant (c168 t/Q192 H). The increased value of kcat led to the improvement of the kcat/Km value for mutants c168 t/Q192H (13.9-fold increase), Y7L and c168 t/ Q192 H/Y7L (1.6-fold increase, respectively). Based on the kcat/Km value, the double mutant c168 t/Q192H is the most catalytically efficient enzyme compared to the others. However, both single mutants Q192H and c168 t showed a reduction in the kcat/Km value compared to the wild type. Gln 192 is located at the ‘fingers’ with the distance of 11.6 Å from the catalytic residue, Glu 189, and connects to Cys 55 and Gly 56 via hydrogen bonds (Fig. 4A). Histidine substitution at residue 192 changes the direction of the side chain by causing it to face inwards towards the cleft (Fig. 4B). According to PIC analysis (Table 5), Q192H mutation has created new hydrogen bonds between His 192 (ND1) and Gly 56 (O) and also between Asp 33 (OD2) and Asn 57 (N). Besides that, a new ionic interaction was formed to Asp 33 which was also bonded to the substrate binding residue, Asn 57. This substrate binding residue was directly in contact with the xylooligosaccharides via hydrogen bonds during substrate binding and catalysis. These new hydrogen bond and ionic interaction between His 192 and Asp 33 might interrupt the contact between Asp 33 and Asn 57 that eventually jeopardise the Asn 57-substrate interaction. Thus, when a higher Km value was acquired, the substrate affinity was reduced, thereby affecting the catalytic efficiency (kcat/Km). Tyr 7 located at the N-terminal region interacts with Val 8, Ala 11 and Leu 15 via hydrophobic interaction (Table 5). This residue is believed to be a putative secondary binding site (SBS) that is located distant from the active site and has been expected to target longer substrates in hydrolysis [45]. SBS is a relatively flat region with aromatic and polar amino acids which is capable of attracting substrate via

Fig. 3. Location of site-saturation mutagenesis residues of AfxynG1. Red stickscatalytic residues and blue sticks-selected residues for site-saturation mutagenesis.

been selected for the construction of site-saturation mutagenesis of endo-β-1, 4-xylanase; xynA from Geobacillus stearothermophilus [26]. In regard, the same approach was used in this study to select Tyr 7 and Ser 75 that shared by more than one mutant of epPCR library. In total, nine residues: Asp 182, Val 186, Gly 56, Gln 192, Asn 136, Phe 146, Asn 54, Tyr 7 and Ser 75 were selected from both strategies to proceed for further assessments via site-saturation mutagenesis (Fig. 3). The NDT (N = A, C, G or T, D = A, G or T) library was chosen for the site-saturation mutagenesis library construction due to the small clones required (only 35 clones) to achieve 95% coverage of a single amino acid substitution. In addition, the NDT library represents a balanced sample of aromatic, aliphatic, non-polar, polar, negative and positively charged residues. This library has up to 3-fold higher efficiency than the NNK (N = A, C, G or T, K = T or G) library [42,43]. Nine NDT libraries (Y7X, S75X, D182X, V186X, G56X, Q192X, N136X, F146X and N54X) were successfully constructed with a total of forty-six mutants in each set of libraries. After high-throughput and lowthroughput screening and DNA sequencing analyses, only three mutants showed higher enzymatic activity compared to the wild-type, namely, Y7L, Q192H and a silent mutant from the G56X library, c168 t (Table 3). To verify the combination effect among these three SSM mutants, site-directed mutagenesis (SDM) was carried out to generate double and triple mutants. Three double mutants (Q192 H/Y7L, c168 t/ Q192H and c168 t/Y7L) and one triple mutant (c168 t/Q192 H/Y7L) were successfully constructed. However, screening showed only mutants c168 t/Q192H and c168 t/Q192 H/Y7L had higher enzymatic activity towards pretreated kenaf than the wild type (Table 3). In conclusion, five potential mutants: c168 t, Q192H, Y7L, c168 t/Q192H and c168 t/Q192 H/Y7L were selected from site-saturation and sitedirected mutagenesis. These final mutants were large-scale expressed

Table 3 Low-throughput screening of improved mutants from site-saturation and sitedirected mutagenesis. Mutagenesis

Mutants

Relative activity (%)a

Site-saturation

c168t Q192H Y7L c168 t/Q192H c168 t/Q192 H/Y7L

207 150 150 134 138

Site-directed

a Hydrolysis activity of mutants was compared to the control reaction of wild-type which was set as 100%.

5

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Fig. 4. A) Gln 192 interaction with close residues at the ‘fingers’ region. Grey cartoon is xylanase structure. Green sticks represent residues involved. Black dotted line is distance measurement between residues. B) His 192 interaction with close residues. Cyan stick shows xylohexaose being inserted into the active site cleft. Red dotted line is protein-ligand interaction.

c168 t, Q192H and c168 t/Q192H.

aromatic stacking interaction and hydrogen bonding, respectively [46]. The leucine substitution at residue 7 eliminates the aromatic group and removes any aromatic interactions that might be formed between the site and xylose sugar. Therefore, this substituted residue generates stronger hydrophobic interaction within the site. Despite the loss of covalent bond (aromatic interaction), secondary binding site of mutant Y7L may still attract the substrate through non-covalent hydrophobic interaction. Therefore, this SBS can facilitate the substrate to the active site [47] and easily release the substrate for catalysis due to the minimal interaction, hence, speeding up the substrate conversion as shown by the higher kcat value obtained. Even though the Km did not improve, the higher kcat contributes to the increased catalytic efficiency. Even though the mechanism of silent mutation (c168 t) in degrading the Km and kcat values is unknown but the combination of double mutant; c168 t/Q192H has successfully improved the kcat value, thereby yielded a remarkable enhancement of kcat/Km. Essentially, Gln 192 and c168 is connected by the hydrogen bond in which Gly 56 is the amino acid translation of c168. Hence, it can be proposed that there is a synergistic effect between these two single mutants that triggered the better catalytic efficiency. The triple mutant c168 t/Q192 H/Y7L generated lower catalytic efficiency than the double mutant. The incorporation of mutant Y7L might give epistasis effect to the triple mutant as kcat/Km value turned to be similar to the single mutant Y7L. In brief, the kcat and kcat/Km values were not influenced by the location of the mutation either closer or distant from the active site, as supported by the previous study [38]. In contrast, the Km value is more affected by mutations closer to the active site, as shown by mutants

3.3. Effects of temperature and pH on mutants Beside kinetic parameters, thermal and pH profiles of the mutants also important in natural substrate hydrolysis for industrial use. The optimum temperature and thermostability of AfxynG1 and the mutants were determined at pH 5 with a temperature range from 30 to 70 °C for 10 min and 30 min, as shown in Fig. 5A and B. Mutants Q192H, c168 t/ Q192H and c168 t/Q192 H/Y7L have an optimum temperature of 50 °C, which was identical to the wild-type, while Y7L and c168 t had a lower optimum temperature of 40 °C. AfxynG1 and the mutants maintained more than 20% of their activity up to 70 °C. After 30 min of incubation at 30–70 °C, the mutants Q192H and c168 t/Q192 H/Y7L retained more than 40% activity through all the temperatures studied, whilst AfxynG1 and the other mutants (c168 t, Y7L and c168 t/Q192 H) displayed a lower thermostability by retaining more than 40% activity at temperatures up to 50 °C only. Mutant Q192H showed higher thermostability due to the substitution of polar glutamine to the positively, charged histidine at residue 192, which is located at the ‘fingers’ (Fig. 3). This mutation might provide a surface area for charge-charge interactions that is important for protein conformation and lead to protein stability [48,49]. Interestingly, mutant c168 t/Q192 H/Y7L displayed the highest stability at 60 °C and 70 °C compared to the others. In the case of triple mutant c168 t/Q192 H/Y7L, the enhanced thermostability is believed to be contributed by the cumulative effect of all mutations. Study conducted by Torpenholt et al. [50], suggested that the thermostability is not controlled by mutation of one specific residue

Table 5 Protein interaction analysis between residues involved in the wild-type and mutants. Interactions

Hydrogen bonds

Ionic interaction Hydrophobic interaction

Wild-type

Mutants

Residues (Atom)

Distance (Å)

Residues (Atom)

Distance (Å)

Gln 192 (N) —— Cys 55 (SG) Gln 192 (N) —— Gly 56 (O) – Asp 33 (OD1) —— Asn 57 (O) Asp 33 (OD1) —— Asn 57 (N) – Tyr 7 (N) —— Arg 13 (O) – Tyr 7 —— Val 8 Tyr 7 —— Ala 11 Tyr 7 —— Leu 15

3.44 3.05

His 192 (N) —— Cys 55 (SG) His 192 (N) —— Gly 56 (O) His 192 (ND1) —— Gly 56 (O)* Asp 33 (OD1) —— Asn 57 (O) Asp 33 (OD1) —— Asn 57 (N) Asp 33 (OD2) —— Asn 57 (N)* Leu 7 (N) —— Arg 13 (O) His 192 —— Asp 33* Leu 7 —— Val 8 Leu 7 —— Ala 11 Leu 7 —— Leu 15

3.54 3.07 3.07 3.30 2.97 3.27 3.26

3.30 2.97 3.26

Newly introduced bond/interaction is marked with an asterisk (*). 6

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Fig. 5. Effects of temperature on the activity and stability of AfxynG1 and its mutants. A) Effect of temperature on the activity of AfxynG1 and its mutants. The assay was performed at different temperature ranging from 30 to 70 °C for 10 min. B) Effect of temperature on the stability of AfxynG1 and its mutants. The purified enzymes were incubated in 50 mM Naacetate buffer (pH 5.0) without substrate for 30 min at varied temperature (30–70 °C) and the relative activities were measured at the optimal condition for 10 min. The highest activity of each WT and mutants were set as 100%.

showed a high tolerance at the lower pH of 4–5 by retaining approximately 80–100 % of the activity with the highest stability at pH 4 and 5 for c168 t and Q192H, respectively. In contrast, the double mutant c168 t/Q192H displayed acid stability at pH 4–6 by preserving approximately 40–100 % activity. It appears that this mutant encounters the instability of both single mutants at pH 6. Lastly, the triple mutant c168 t/Q192 H/Y7L showed the most tolerable enzyme at all pH values studied (3–6) according to the recorded activity, which was approximately 60–100 % with the highest stability at pH 5. Nevertheless, the remarkable acid stability at the lower pH by the silent mutant c168 t is not understood. Mutant Q192H achieve better stability at pH 4–5 than

yet influenced by combined effect of various attributes. In addition, even small structural changes are capable of affecting the overall stability [50]. Among the structural factors that could influence protein thermostability are compactness, hydrophobicity, buried and exposed surface areas, hydrogen bonds and salt-bridges [51]. The optimal pH was verified at 50 °C for 10 min at different pH values (3–8), whilst the acid stability was performed for 30 min incubation at pH 3–6 (Fig. 6A and B). Both the wild-type and mutants showed optimum activity at pH 5. AfxynG1 and mutant Y7L exhibited a similar profile of acid stability by achieving the highest stability at pH 6 but less stable at lower pH (3–5). Both mutants c168 t and Q192H

Fig. 6. Effects of pH on the activity and stability of AfxynG1 and its mutants. A) Effect of pH on the activity of AfxynG1 and its mutants. The assay was performed in different pH buffer ranging from pH 3 to 8 at 50 °C for 10 min. B) Effect of pH on the stability of AfxynG1 and mutants. The enzyme was incubated in different buffer (pH 3 to 6) at 25 °C for 30 min and the relative activities were measured at the optimal condition for 10 min. The highest activity of each WT and mutants were set as 100%.

7

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

Table 6 Enzyme hydrolysis of pretreated kenaf at three different temperature; 30 °C, 40 °C and 50 °C with 36 h incubation. Variants

Rsmax at 30 °C (mg/g)

Rsmax at 40 °C (mg/g)

Rsmax at 50 °C (mg/g)

WT c168t Q192H Y7L c168 t/Q192H c168 t/Q192 H/Y7L

3.14 2.24 2.94 4.03 3.74 3.75

2.97 3.09 3.12 3.13 3.20 3.18

2.62 2.54 2.58 2.78 2.85 2.71

± ± ± ± ± ±

0.01 0.10 0.04 0.16 0.10 0.19

(1.0) (0.71) (0.94) (1.28) (1.19) (1.19)

± ± ± ± ± ±

0.06 0.09 0.09 0.04 0.07 0.07

(1.0) (1.04) (1.05) (1.05) (1.08) (1.07)

± ± ± ± ± ±

0.03 0.02 0.02 0.12 0.16 0.09

(1.0) (0.97) (0.98) (1.06) (1.09) (1.03)

The reaction was carried out in 50 mM sodium acetate buffer (pH 5.0) containing 1% (w/v) pretreated kenaf at 200 rpm water bath. The average of triplicate determination with SD value is shown. Numbers in parentheses indicate values relative to wild-type xylanase at each temperature.

substrate into the active site cleft and thereby might be the reason which allows the mutant to release the highest number of reducing sugars. However, a detailed study on Tyr 7 should be carried out to confirm its role as a secondary binding site. Mutants c168 t/Q192H and c168 t/Q192 H/Y7L also became better producers of reducing sugars than the wild-type. Both mutants showed improvements in catalytic efficiency, and mutant c168 t/Q192H emerged as the most catalytic efficient enzyme among the others. The remaining two mutants, Q192H and c168 t, showed poor hydrolysis performance compared to the wild-type. These mutants attained mutations closer to the active site and this alteration caused a decrease in catalytic efficiency that eventually limiting the sugar yield. The mutations located at the N-terminal region and the synergistic effect of multiple mutations closer to the active site triggered the better catalytic efficiency of the enzyme and subsequently initiated the good performance of hydrolysis on lignocellulose biomass.

the wild type might be because of the glutamine substitution by a positively charged histidine. For a catalytic reaction to occur, the proton donor (in this case Glu 189) is protonated, and the catalytic nucleophile (Glu 98) attacks and stabilizes the intermediate molecules. However, in the low pH environment, the pKa values of these catalytic residues (Glu 189 and Glu 98) are reduced, limiting the enzyme to function optimally as in pH values of approximately 7 [52]. In this study, the positively charged histidine that substituted glutamine at residue 192 might form a desolvation effect and alter the electrostatic field, thereby shifting the pKa values of the catalytic group upward to a near-neutral pH value so that the mutant Q192H performed better at pH 4–5 than the wild type. The better acid stability performance displayed by the H400R mutant of Bacillus licheniformis α-amylase was also contributed by more hydrophilic, charged residue substitution [53]. 3.4. Pretreated kenaf hydrolysis

4. Conclusion

The hydrolysis of pretreated kenaf by partially purified xylanases was measured according to the amount of reducing sugar produced per gram substrate (mg/g). Maximum sugar produced (Rsmax) of pretreated kenaf hydrolysis was achieved after 36 h incubation of the reaction mixture at different temperatures: 30, 40 and 50 °C (Table 6). These temperatures were selected because AfxynG1 and its mutants showed thermostability at 30–50 °C by maintaining more than 40% enzyme activity (Sec. 3.3). Based on the Rsmax value obtained, mutants c168 t and Q192H worked best at 40 °C, while the wild-type and the other mutants favoured 30 °C. Clearly, AfxynG1 and all mutants displayed the lowest Rsmax at 50 °C. This effect might be due to the enzyme instability at higher temperatures for longer reaction times (> 24 h), as supported by the previous study of pretreated kenaf hydrolysis by Xyn2 from Trichoderma reesei ATCC 58350 [54]. At 30 °C, mutant Y7L produced the highest Rsmax (4.03 mg/g), followed by c168 t/Q192 H/Y7L (3.75 mg/g) and c168 t/Q192H (3.74 mg/g) in comparison to the wild-type that produced 3.14 mg/g of the total sugar. Meanwhile, mutants c168 t produced a lower amount of total reducing sugar than the wild-type and mutant Q192H that exhibited no difference to the wild-type. At 40–50 °C, all of the mutants produced quite a similar amount of total reducing sugar compared to the wild-type. As mentioned in Table 6, the yield of total reducing sugar from pretreated kenaf hydrolysis showed only a little improvement (28%) by mutant Y7L if compared to the previous study [47]. The low sugar production is expected as this study only applied two rounds of directed evolution (SSM and SDM) excluding epPCR which acted as a fine tuner for the next step of mutation. Furthermore, xylanase was acting alone without any accessory enzyme like cellulase in the hydrolysis which can improve the hydrolysis synergistically. However, this result provides a good basis to further improving the enzyme performance for natural substrate hydrolysis. Mutant Y7L improved the lignocellulose biomass hydrolysis due to the substitution of an aromatic, polar tyrosine at residue 7 by leucine (a non-polar and smaller in size). This mutation caused better catalytic efficiency due to the higher kcat value as discussed previously in Sec 3.2. As a putative secondary binding site, residue 7 might guide a longer

The present investigation revealed that the mutation closer to the active site and N-terminal residue substitution produced three industrial-potential mutants that might encounter the resistance of lignocellulose biomass structure for hydrolysis. These mutants displayed improved catalytic efficiency and showed an increased release of reducing sugar from the lignocellulosic substrate, as well as an enhancement in thermostability and acid stability. These improved mutants are beneficial to be used synergistically with cellulases for the bioconversion of lignocellulosic biomass into fermentable sugars in broad industrial applications. Acknowledgements We acknowledge Everise Crimson (M) Sdn. Bhd. for providing us with the kenaf sample. This research did not receive any specific grant from funding agencies in the public, commercial or not-for-profit sectors. References [1] N. Saba, M. Jawaid, K.R. Hakeem, M.T. Paridah, A. Khalina, O.Y. Alothman, Potential of bioenergy production from industrial kenaf (Hibiscus cannabinus L.) based on Malaysian perspective, Renewable Sustainable Energy Rev. 42 (2015) 446–459, https://doi.org/10.1016/j.rser.2014.10.029. [2] H.M.N. Iqbal, G. Kyazze, T. Keshavarz, Advances in the valorization of lignocellulosic materials by biotechnology: an overview, BioResources. 8 (2013) 3157–3176, https://doi.org/10.15376/biores.8.2.3157-3176. [3] P.S. Nigam, A. Singh, Production of liquid biofuels from renewable resources, Prog. Energy Combust. Sci. 37 (2011) 52–68, https://doi.org/10.1016/j.pecs.2010.01. 003. [4] L.D. Gottumukkala, K. Haigh, J. Görgens, Trends and advances in conversion of lignocellulosic biomass to biobutanol: microbes, bioprocesses and industrial viability, Renewable Sustainable Energy Rev. 76 (2017) 963–973, https://doi.org/10. 1016/j.rser.2017.03.030. [5] V. García, J. Päkkilä, H. Ojamo, E. Muurinen, R.L. Keiski, Challenges in biobutanol production: How to improve the efficiency? Renewable Sustainable Energy Rev. 15 (2011) 964–980, https://doi.org/10.1016/j.rser.2010.11.008. [6] X.B. Liu, Q.Y. Gu, X. Bin Yu, Repetitive domestication to enhance butanol tolerance

8

Enzyme and Microbial Technology 131 (2019) 109383

S.I.R. Damis, et al.

[7]

[8]

[9]

[10] [11]

[12]

[13]

[14]

[15]

[16]

[17]

[18]

[19]

[20]

[21]

[22]

[23]

[24]

[25]

[26]

[27]

[28]

[29]

and production in Clostridium acetobutylicum through artificial simulation of bioevolution, Bioresour. Technol. 130 (2013) 638–643, https://doi.org/10.1016/j. biortech.2012.12.121. M. Balat, Production of bioethanol from lignocellulosic materials via the biochemical pathway: A review, Energy Convers. Manage. 52 (2011) 858–875, https:// doi.org/10.1016/j.enconman.2010.08.013. N. Pareek, T. Gillgren, L.J. Jonsson, Adsorption of proteins involved in hydrolysis of lignocellulose on lignins and hemicelluloses, Bioresour. Technol. 148 (2013) 70–77, https://doi.org/10.1016/j.biortech.2013.08.121. K. Ulaganathan, S. Goud, M. Reddy, U. Kayalvili, Genome engineering for breaking barriers in lignocellulosic bioethanol production, Renewable Sustainable Energy Rev. 74 (2017) 1080–1107, https://doi.org/10.1016/j.rser.2017.01.028. D. Shallom, Y. Shoham, Microbial hemicellulases, Curr. Opin. Microbiol. 6 (2003) 219–228, https://doi.org/10.1016/S1369-5274(03)00056-0. Q.K. Beg, M. Kapoor, L. Mahajan, G.S. Hoondal, Microbial xylanases and their industrial applications: a review, Appl. Microbiol. Biotechnol. 56 (2001) 326–338, https://doi.org/10.1007/s002530100704. M. Madadi, Y. Tu, A. Abbas, Recent status on enzymatic saccharification of lignocellulosic biomass for bioethanol production, Electron. J. Biol. 13 (2017) 135–143. M.E. Himmel, S.-Y. Ding, D.K. Johnson, W.S. Adney, M.R. Nimlos, J.W. Brady, T.D. Foust, Biomass recalcitrance: engineering plants and enzymes for biofuels production, Science. 315 (2007) 804–807, https://doi.org/10.1126/science. 1137016. E. Visser, T. Leal, M. de Almeida, V. Guimarães, Increased enzymatic hydrolysis of sugarcane bagasse from enzyme recycling, Biotechnol. Biofuels 8 (2015) 5, https:// doi.org/10.1186/s13068-014-0185-8. A. Morone, R.A. Pandey, Lignocellulosic biobutanol production : gridlocks and potential remedies, Renewable Sustainable Energy Rev. 37 (2014) 21–35, https:// doi.org/10.1016/j.rser.2014.05.009. A. Berlin, N. Gilkes, D. Kilburn, R. Bura, A. Markov, A. Skomarovsky, O. Okunev, A. Gusakov, V. Maximenko, D. Gregg, A. Sinitsyn, J. Saddler, Evaluation of novel fungal cellulase preparations for ability to hydrolyze softwood substrates - Evidence for the role of accessory enzymes, Enzyme Microb. Technol. 37 (2005) 175–184, https://doi.org/10.1016/j.enzmictec.2005.01.039. T.S. Quiñones, A. Retter, P.J. Hobbs, J. Budde, M. Heiermann, M. Plöchl, S.R. Ravella, Production of xylooligosaccharides from renewable agricultural lignocellulose biomass, Biofuels. 6 (2015) 147–155, https://doi.org/10.1080/ 17597269.2015.1065589. Y. Yang, J. Yang, J. Liu, R. Wang, L. Liu, F. Wang, H. Yuan, The composition of accessory enzymes of Penicillium chrysogenum P33 revealed by secretome and synergistic e ff ects with commercial cellulase on lignocellulose hydrolysis, Bioresour. Technol. 257 (2018) 54–61, https://doi.org/10.1016/j.biortech.2018.02.028. J.A. Diogo, Z.B. Hoffmam, L.M. Zanphorlin, J. Cota, C.B. Machado, L.D. Wolf, F. Squina, A.R.L. Damásio, M.T. Murakami, R. Ruller, Development of a chimeric hemicellulase to enhance the xylose production and thermotolerance, Enzyme Microb. Technol. 69 (2015) 31–37, https://doi.org/10.1016/j.enzmictec.2014.11. 006. M. McLachlan, R.P. Sullivan, H. Zhao, Directed enzyme evolution and high throughput screening, Biocatal. Pharm. Ind. Discov., Dev. Manuf (2009) 1–20 http://scs.illinois.edu/-zhaogrp/publications/HZ72.pdf. F. Wen, M. Mclachlan, H. Zhao, Directed Evolution : novel and improved enzymes, Web Encycl. Chem. Biol. (2008) 1–7, https://doi.org/10.1002/9780470048672. wecb125. R.A. Chica, N. Doucet, J.N. Pelletier, Semi-rational approaches to engineering enzyme activity: combining the benefits of directed evolution and rational design, Curr. Opin. Biotechnol. 16 (2005) 378–384, https://doi.org/10.1016/j.copbio. 2005.06.004. F. Valetti, G. Gilardi, Improvement of biocatalysts for industrial and environmental purposes by saturation mutagenesis, Biomolecules. 3 (2013) 778–811, https://doi. org/10.3390/biom3040778. Q. Li, B. Sun, H. Jia, J. Hou, R. Yang, K. Xiong, Y. Xu, X. Li, Engineering a xylanase from Streptomyce rochei L10904 by mutation to improve its catalytic characteristics, Int. J. Biol. Macromol. 101 (2017) 366–372, https://doi.org/10.1016/j.ijbiomac. 2017.03.135. Z.B. Hoffmam, L.M. Zanphorlin, J. Cota, J.A. Diogo, G.B. Almeida, A.R.L. Damásio, F. Squina, M.T. Murakami, R. Ruller, Xylan-specific carbohydrate-binding module belonging to family 6 enhances the catalytic performance of a GH11 endo-xylanase, N. Biotechnol. 33 (2016) 467–472, https://doi.org/10.1016/j.nbt.2016.02.006. Y. Wang, S. Feng, T. Zhan, Z. Huang, G. Wu, Z. Liu, Improving catalytic efficiency of endo-β-1, 4-xylanase from Geobacillus stearothermophilus by directed evolution and H179 saturation mutagenesis, J. Biotechnol. 168 (2013) 341–347, https://doi.org/ 10.1016/j.jbiotec.2013.09.014. M.K.H. Bin Abdul Wahab, M.A. Bin Jonet, R.M. Illias, thermostability enhancement of xylanase Aspergillus fumigatus RT-1, J. Mol. Catal., B Enzym. 134 (2016) 154–163, https://doi.org/10.1016/j.molcatb.2016.09.020. N.I. Wan Azelee, J. Md Jahim, A. Rabu, A.M. Abdul Murad, F.D. Abu Bakar, R. Md Illias, Efficient removal of lignin with the maintenance of hemicellulose from kenaf by two-stage pretreatment process, Carbohydr. Polym. 99 (2014) 447–453, https:// doi.org/10.1016/j.carbpol.2013.08.043. L. Song, S. Laguerre, C. Dumon, S. Bozonnet, M.J. ’Donohue, A high-throughput screening system for the evaluation of biomass-hydrolyzing glycoside hydrolases, Bioresour. Technol. 101 (2010) 8237–8243, https://doi.org/10.1016/j.biortech.

2010.05.097. [30] F. Payan, P. Leone, S. Porciero, C. Furniss, T. Tahir, G. Williamson, A. Durand, P. Manzanares, H.J. Gilbert, N. Juge, A. Roussel, The dual nature of the wheat xylanase protein inhibitor XIP-I: structural basis for the inhibition of family 10 and family 11 xylanases, J. Biol. Chem. 279 (2004) 36029–36037, https://doi.org/10. 1074/jbc.M404225200. [31] G.M. Morris, D.S. Goodsell, R. Huey, A.J. Olson, Distributed automated docking of flexible ligands to proteins: parallel applications of AutoDock 2.4, J. Comput. Aided Mol. Des. 10 (1996) 293–304, https://doi.org/10.1007/BF00124499. [32] K.G. Tina, R. Bhadra, N. Srinivasan, PIC: protein interactions calculator, Nucleic Acids Res. 35 (2007) 473–476, https://doi.org/10.1093/nar/gkm423. [33] L. Zheng, An efficient one-step site-directed and site-saturation mutagenesis protocol, Nucleic Acids Res. 32 (2004) e115, https://doi.org/10.1093/nar/gnh110. [34] F.W. Studier, Protein production by auto-induction in high density shaking cultures, Protein Expr. Purif. 41 (2005) 207–234, https://doi.org/10.1016/j.pep.2005.01. 016. [35] M.J. Bailey, P. Biely, K. Poutanen, Interlaboratory testing of methods for assay of xylanase activity, J. Biotechnol. 23 (1992) 257–270, https://doi.org/10.1016/ 0168-1656(92)90074-J. [36] G.L. Miller, Use of dinitrosalicylic acid reagent for determination of reducing sugar, Anal. Chem. 31 (1959) 426–428, https://doi.org/10.1021/ac60147a030. [37] T.J. Bach, H.K. Lichtenthaler, Application of modified Lineweaver-Burk plots to studies of kinetics and regulation of radish 3-hydroxy-3-methylglutaryl-CoA reductase, Biochim. Biophys. Acta (BBA)/Lipids Lipid Metab. 794 (1984) 152–161, https://doi.org/10.1016/0005-2760(84)90308-4. [38] K.L. Morley, R.J. Kazlauskas, Improving enzyme properties: When are closer mutations better? Trends Biotechnol. 23 (2005) 231–237, https://doi.org/10.1016/j. tibtech.2005.03.005. [39] R. Havukainen, A. Törröen, T. Laitinen, J. Rouvinen, Covalent binding of three epoxyalkyl xylosides to the active site of endo-1,4-xylanase II from Trichoderma reesei, Biochemistry. 35 (1996) 9617–9624, https://doi.org/10.1021/bi953052n. [40] A. Törrönen, J. Rouvinen, Structural and functional properties of low molecular weight endo-1,4-beta-xylanases, J. Biotechnol. 57 (1997) 137–149, https://doi.org/ 10.1016/S0168-1656(97)00095-3. [41] J.D. McCarter, G. Stephen Withers, Mechanisms of enzymatic glycoside hydrolysis, Curr. Opin. Struct. Biol. 4 (1994) 885–892, https://doi.org/10.1016/0959-440X (94)90271-2. [42] R. Martínez, U. Schwaneberg, A roadmap to directed enzyme evolution and screening systems for biotechnological applications, Biol. Res. 46 (2013) 395–405, https://doi.org/10.4067/S0716-97602013000400011. [43] M.T. Reetz, H. Höbenreich, P. Soni, L. Fernández, A genetic selection system for evolving enantioselectivity of enzymes, Chem. Commun. (Camb.) (2008) 5502, https://doi.org/10.1039/b814538e. [44] J.M. Berg, J.L. Tymoczko, L. Stryer, Biochemistry, W.H. Free. 5th, (2002), pp. 319–329. [45] S. Cuyvers, E. Dornez, M.N. Rezaei, A. Pollet, J.A. Delcour, C.M. Courtin, Secondary substrate binding strongly affects activity and binding affinity of Bacillus subtilis and Aspergillus niger GH11 xylanases, FEBS J. 278 (2011) 1098–1111, https://doi.org/ 10.1111/j.1742-4658.2011.08023.x. [46] M.L. Ludwiczek, M. Heller, T. Kantner, L.P. McIntosh, A secondary xylan-binding site enhances the catalytic activity of a single-domain family 11 glycoside hydrolase, J. Mol. Biol. 373 (2007) 337–354, https://doi.org/10.1016/j.jmb.2007.07. 057. [47] S. Cuyvers, E. Dornez, J.A. Delcour, C.M. Courtin, Occurrence and functional significance of secondary carbohydrate binding sites in glycoside hydrolases, Crit. Rev. Biotechnol. 32 (2012) 93–107, https://doi.org/10.3109/07388551.2011.561537. [48] T. Tu, H. Luo, K. Meng, Y. Cheng, R. Ma, P. Shi, H. Huang, Y. Bai, Y. Wang, L. Zhang, B. Yao, Improvement in thermostability of an Achaetomium sp. Strain Xz8 endopolygalacturonase via the optimization of charge-charge interactions, Appl. Environ. Microbiol. 81 (2015) 6938–6944, https://doi.org/10.1128/AEM. 01363-15. [49] M. Irfan, C.F. Gonzalez, S. Raza, M. Rafiq, F. Hasan, S. Khan, A.A. Shah, Improvement in thermostability of xylanase from Geobacillus thermodenitrificans C5 by site directed mutagenesis, Enzyme Microb. Technol. 111 (2018) 38–47, https:// doi.org/10.1016/j.enzmictec.2018.01.004. [50] S. Torpenholt, L. De Maria, M.H.M. Olsson, L.H. Christensen, M. Skjøt, P. Westh, J.H. Jensen, L. Lo Leggio, Effect of mutations on the thermostability of Aspergillus aculeatus β-1,4-galactanase, Comput. Struct. Biotechnol. J. 13 (2015) 256–264, https://doi.org/10.1016/j.csbj.2015.03.010. [51] S. Kumar, C.-J. Tsai, R. Nussinov, Factors enhancing protein thermostability, Protein Eng, Protein Eng. Des. Sel. 13 (2002) 179–191, https://doi.org/10.1093/ protein/13.3.179. [52] J.E. Nielsen, J.A. McCammon, Calculating pKa values in enzyme active sites, Protein Sci. 12 (2003) 1894–1901, https://doi.org/10.1110/ps.03114903. [53] Y. Liu, L. Huang, L. Jia, S. Gui, Y. Fu, D. Zheng, W. Guo, F. Lu, Improvement of the acid stability of Bacillus licheniformis alpha amylase by site-directed mutagenesis, Process Biochem. 58 (2012) 174–180, https://doi.org/10.1016/j.procbio.2017.04. 040. [54] N.I. Wan Azelee, J.M. Jahim, A.F. Ismail, S.F.Z.M. Fuzi, R.A. Rahman, R. Md Illias, High xylooligosaccharides (XOS) production from pretreated kenaf stem by enzyme mixture hydrolysis, Ind. Crops Prod. 81 (2016) 11–19, https://doi.org/10.1016/j. indcrop.2015.11.038.

9