Synergistic action of recombinant accessory hemicellulolytic and pectinolytic enzymes to Trichoderma reesei cellulase on rice straw degradation

Bioresource Technology 198 (2015) 682–690

Contents lists available at ScienceDirect

Bioresource Technology journal homepage: www.elsevier.com/locate/biortech

Synergistic action of recombinant accessory hemicellulolytic and pectinolytic enzymes to Trichoderma reesei cellulase on rice straw degradation Thanaporn Laothanachareon, Benjarat Bunterngsook, Surisa Suwannarangsee, Lily Eurwilaichitr, Verawat Champreda ⇑ Enzyme Technology Laboratory and Integrative Biorefinery Laboratory, National Center for Genetic Engineering and Biotechnology (BIOTEC), National Science and Technology Development Agency, Thailand Science Park, 113 Pahonyothin Road, Pathumthani 12120, Thailand

h i g h l i g h t s

g r a p h i c a l a b s t r a c t


Synergy of accessory hemicellulase

and pectinase to core cellulase was shown.
A varying degree of synergism of ARA, PEC, XYL to ACR cellulase was found.
Removal of ara side chain by arabinofuranosidase enhanced cellulase activity.
Synergy of pectin esterase to cellulase mixture was firstly reported.
The quaternary mixture showed 214% glc obtained/FPU compared to ACR alone.

a r t i c l e

i n f o

Article history: Received 16 July 2015 Received in revised form 12 September 2015 Accepted 14 September 2015 Available online 30 September 2015 Keywords: Accessory enzymes Aspergillus aculeatus Cellulase Lignocellulose Mixture design

a b s t r a c t Synergism between core cellulases and accessory hydrolytic/non-hydrolytic enzymes is the basis of efficient hydrolysis of lignocelluloses. In this study, the synergistic action of three recombinant accessory enzymes, namely GH62 a-L-arabinofuranosidase (ARA), CE8 pectin esterase (PET), and GH10 endo-1,4beta-xylanase (XYL) from Aspergillus aculeatus expressed in Pichia pastoris to a commercial Trichoderma reesei cellulase (AccelleraseÒ 1500; ACR) on hydrolysis of alkaline pretreated rice straw was studied using a mixture design approach. Applying the full cubic model, the optimal ratio of quaternary enzyme mixture was predicted to be ACR:ARA:PET:XYL of 0.171:0.079:0.100:0.150, which showed a glucose releasing efficiency of 0.173 gglc/FPU, higher than the binary ACR:XYL mixture (0.122 gglc/FPU) and ACR alone (0.081 gglc/FPU) leading to a 47.3% increase in glucose yield compared with that from ACR at the same cellulase dosage. The result demonstrates the varying degree of synergism of accessory enzymes to cellulases useful for developing tailor-made enzyme systems for bio-industry. Ó 2015 Elsevier Ltd. All rights reserved.

1. Introduction

⇑ Corresponding author. Tel.: +66 2564 6700x3473; fax: +66 2564 6707. E-mail address: [email protected] (V. Champreda). http://dx.doi.org/10.1016/j.biortech.2015.09.053 0960-8524/Ó 2015 Elsevier Ltd. All rights reserved.

Over the past decade, lignocellulosic plant biomass has attracted attention as a renewable resource for production of biofuels and commodity chemicals. As fossil resources become scarcer and the use of their derivatives generate environmental concern on

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

global warming, biorefinery is considered as a more sustainable platform industry with competitive advantages in the long term (Cherubini, 2010). Extensive research has been conducted to understand the complex natural bioconversion of lignocelluloses, particularly on degradation of recalcitrant plant biomass. Insights into biomass decomposition by means of intricate enzymatic processes may lead to the development of efficient enzyme systems for saccharification and modification of lignocellulosic materials in bioindustries. Plant cell wall is a lignocellulosic material composed of mainly three different types of polysaccharide polymers, i.e. cellulose, hemicellulose and pectin. Cellulose is a linear homopolymer of Dglucose formed into highly organized microfibers, which are intimately associated with an intricate network of hemicellulose, a branched heteropolymer of various pentoses, hexoses, and sugar acids. Pectin, a galacturonic acid containing polysaccharide forms a gel layer between cellulose microfibers. The polysaccharide microstructure is covered by lignin, a complex heteropolymer of phenolic substance, which acts as a protective shield. Lignocelluloses are highly recalcitrant to biological degradation, and thus lignocellulolytic microorganisms employ a variety of cellulolytic, hemicellulolytic, and lignolytic enzymes that act specifically and synergistically to degrade plant biomass (Himmel et al., 2007). The core of cellulase contains three types of enzymes including endoglucanase (EG, endo-1,4-D-glucanohydrolase: EC3.2.1.4), which randomly cleaves internal bonds in the cellulose fiber creating free chains ends, exoglucanase or cellobiohydrolase (CBH, 1,4b-exoglucanase-glucan cellobiohydrolase: EC 3.2.1.91), which sequentially cleaves cellobiose units from free chain ends, and bglucosidase (EC 3.2.1.21), which releases glucose units from cellobiose (Howard et al., 2003). A wide variety of hemicellulolytic enzymes attack the heterogenous hemicellulose structures, including several endo-/exo-acting hydrolytic enzymes that degrade the main polysaccharide chain and a range of accessory enzymes attacking the ‘‘decorated” branches of hemicelluloses, e.g., a-Larabinofuranosidase, a-glucuronidase, a-galactosidase, and bmannanase (Gottschalk et al., 2010). The embedded pectin matrix is degraded by various hydrolytic and non-hydrolytic pectinases (Willats et al., 2001). Many cellulolytic and hemicellulolytic enzymes from aerobic or anaerobic bacteria and fungi display synergistic action (Bayer et al., 2004; Zhang and Lynd, 2006). Different mechanisms have been described to explain the synergy, including: (i) enhancement of upstream enzymes by downstream enzymes acting on smaller substrates e.g. alleviation of cellobiohydrolase inhibition by a bD-glucosidase

which cleaves its inhibitor, cellobiose to glucose, (ii) the endo/exo effect, where endocellulases create new, free cellodextrin chain ends for cellobiohydrolases; (iii) synergism between exocellulases attacking reducing and non-reducing ends; (iv) cooperative action of cellulolytic and hemicellulolytic enzymes which increase accessibility to each other’s respective target substrates, and (v) physical alteration of the substrates e.g. loosening of the crystallised region of cellulose by auxiliary proteins and non-hydrolytic enzymes e.g. expansins and lytic polysaccharide monoxygenases (Bunterngsook et al., 2015; Leggio et al., 2015). This synergistic enzyme action forms the basis of efficient lignocellulose degradation in nature and could be applicable for the development of efficient lignocellulolytic enzyme systems for biotechnological application. It is well accepted that different lignocellulosic biomasses pretreated using different methods require empirically determined combinations of different enzymes for optimal digestibility in biotechnological applications. Identification of the key enzymes and optimization of their relative ratios can thus reduce enzyme usage without sacrificing the rate or yield from substrate

683

hydrolysis. In the present study, the synergistic action of different types of accessory enzymes including an a-L-arabonifuranosidase (GH62; ARA) which acts on removing the arabinosyl side chain from the main-chain of hemicellulose, a pectinesterase (CE8; PET) which functions cooperatively with other pectinolytic enzymes on pectin degradation by cleaving the methyl group from the pectin structure, and an endo-1,4-beta-xylanase (GH10; XYL) which attacks the internal glycosidic bond in main-chain xylan and other xylan-containing hemicellulosic polymers from Aspergillus aculeatus BCC17849 to a Trichoderma reesei cellulase (AccelleraseÒ 1500; ACR) on hydrolysis of alkaline pretreated rice straw was studied using a systematic experimental design approach. The work showed strong synergistic actions of carbohydrate processing enzymes with different specificities on hydrolysis of lignocelluloses. The findings provide a basis for formulation of active enzyme mixtures for efficient biomass saccharification and modification. 2. Methods 2.1. Materials Rice straw was collected from a local field in Pathum Thani province, Thailand. The biomass was physically processed using a cutting mill (Retsch SM 2000, Hann, Germany) and sieved to particles of diameter 250–420 lm 0.21–0.35 mesh). The biomass was pretreated with 10% (w/v) NaOH at 80 °C for 90 min at the solid/liquid ratio of 1:3, washed with water until neutral pH was obtained, and air-dried before use as the substrate for enzymatic hydrolysis. A. aculeatus BCC17849 was obtained from the BIOTEC Culture Collection, Thailand (www.biotec.or.th/bcc) and maintained on potato dextrose agar (PDA). Polysaccharides used as substrates in enzymatic activity analysis including carboxymethylcellulose sodium salt (CMC) (Cat. No. 21101) was purchased from Fluka while beechwood xylan (Cat No. X4252), oat-spelt xylan (Cat. No. X0627), and pectin from citrus peel (Cat No. P9135) were obtained from Sigma–Aldrich. The alkali-pretreated rice straw contained 78.36% cellulose, 10.88% hemicellulose, 9.86% lignin and 1.25% ash according to the standard NREL analysis method (Sluiter et al., 2008). 2.2. Culture preparation and total RNA extraction A. aculeatus BCC17849 was grown aerobically in submerged liquid culture in 250 ml conical flasks. The culture was prepared from the culture on PDA by plunging 4 agar pieces each covered with a profuse mycelia mat using a cock borer number 2 into 50 ml of WS medium (3% (w/v) wheat bran and 1% (w/v) soy bean). The inoculated culture was incubated at 25 °C for 5–7 d with no shaking. The mycelia were harvested by filtration on sterile gauze, and ground up to fine powder in liquid nitrogen until a powdery consistency was achieved. Total RNA was extracted using TRI reagent (Molecular Research Center, Inc., Ohio, USA) following the manufacturer’s instruction. The quality and integrity of RNA was determined by gel electrophoresis in 1% agarose containing 3.5% formaldehyde. 2.3. cDNA synthesis The first-strand cDNA of BCC18949 was synthesized using the total RNA as a template. The 20 ll reaction contained 0.5 lg of oligo(dT)18, 1 ll of 10 mM dNTPs and 10 pg-5 lg of total RNA. The reaction was incubated at 65 °C for 5 min and kept on ice for 1 min. The reaction mixture was then supplemented with 4 ll of 5 First-Strand buffer, 1 ll of 0.1 M DTT, 1 ll of RNaseOUT and 1 ll of SuperScriptIII RT (Invitrogen, Carlsbad, CA, USA) and incu-

684

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

bated at 50 °C for 1 h before the reaction was terminated by incubating at 70 °C for 15 min. The reaction mixture was treated by with 1 ll of RNaseH and incubated at 37 °C for 20 min to remove RNA. The synthesized cDNA was kept at 20 °C and used as the template for gene amplification. 2.4. Gene amplification and plasmid construction The full-length genes including a-L-arabinofuranosidase (ara), pectin esterase (pet) and endo-1,4-beta xylanase (xyl) were amplified from the cDNA of A. aculeatus BCC17849 using primers designed from available genome sequence data of the related strain A. aculeatus ATCC16872 (JGI genome database, accessed via: http://genome. jgi.doe.gov/Aspac1/ Aspac1.home.html); (Table 1). The PCR mixtures of 25 ll total volume contained 12.5 ll of 2 PrimeSTAR GC buffer (+Mg2+), 0.4 mM dNTP, 0.4 lM of each primer, and 0.5 ll of PrimeSTAR HS DNA polymerase (Takara Bio Inc., Shiga, Japan), and 20 ng of cDNA. Reactions were performed in a MyCycler thermal cycler (Bio-Rad Laboratories, Hercules, CA). The temperature profile consisted of 95 °C for 1 min, followed by 30 cycles of denaturation of 98 °C for 10 s, annealing at 58 °C for 15 s and extension at 72 °C for 2 min, and a final extension step at 72 °C for 10 min. The amplicons were purified using a GeneJet Gel Extraction Kit (Thermo Fisher Scientific, Waltham, MA, USA) and ligated to the pJET1.2/blunt vector (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s instruction and transformed into the Escherichia coli DH5a. The nucleotide sequences of recombinant plasmids were confirmed by Sanger sequencing (Macrogen, Korea). Signal peptides were predicted from translated sequences using SignalP 4.1 (http:// www.cbs.dtu.dk/ services/SignalP/). Truncated gene sequences corresponding to the assumed mature polypeptide were then amplified using the plasmid as the template using the same reverse primer and a forward primer designed to remove the signal peptide encoding sequence with an additional EcoRI site for cloning. The PCR mixtures of 50 ll total volume contained 25 ll of 2 PrimeSTAR GC buffer (+Mg2+), 0.2 mM dNTPs, 0.2 lM of each primer, 0.5 ll of PrimeSTAR HS DNA polymerase (Takara Bio Inc., Shiga, Japan), and 20 ng of the recombinant plasmid. The temperature profile was the same as described for amplification of the full-length gene. The amplicons were digested with EcoRI and XbaI and ligated to pPICZaA (Invitrogen, Carlsbad, CA, USA) pre-digested with the respective enzymes. The resultant recombinant plasmids containing the ara, pet, and xyl gene were designated as pPICZaA-ara, pPICZaA-pet, and pPICZaA-xyl, respectively. The molecular weights of the translated proteins were calculated using Compute pI/Mw tool (http://web.expasy.org/compute_pi/). The gene sequences were deposited in the NCBI database under the GenBank numbers KT003533, KT003534, and KT003535 for ara, pet, and xyl, respectively.

Table 1 Primer sequences. Primer

Sequence (50 ? 30 )

TR38_ara_F TR38_ara_R TR38_pet_F TR38_pet_R TR38_xyl_F TR38_xyl_R ARA_pPICZaA_EcoRI_F ARA_pPICZaA_XbaI_R PET_pPICZaA_EcoRI_F PET_pPICZaA_XbaI_R Xyl_pPICZaA_EcoRI_F Xyl_pPICZaA_XbaI_R

ATGAAATTCCTCAAGGCCAAAGCTGGTCTGC TTCATTGCTTCAGAGTGAGCACACCCGG ATGCATCTCTACTCTGCTCTTGTCGCCC TTAGTAAGTCCGATCGATCCAATCCCCCC ATGGTTCAAATCAAAGCAGCTGCTCTGGC CTAGAGAGCGTTCGCAATAGCGGTATAAGC CCGCGAATTCAGCTGCCCTCTTCCTTCTACC CGCGTCTAGATCATTGCTTCAGAGTGAG CAGCGAATTCGCTCCGGCGCCAACCCTCG CGCCTCTAGATTAGTAAGTCCGATCGATCC CAGCGAATTCAACCCCATCGAGCCCCGC CGCGTCTAGACTAGAGAGCGTTCGCAATAGC

2.5. Transformation and expression The plasmid pPICZaA-ara, pPICZaA-pet, and pPICZaA-xyl linearized with PmeI were transformed into Pichia pastoris KM71 (Invitrogen, Carlsbad, CA, USA) using electroporation according to the manufacturer’s instruction. Transformants were grown on YPD agar (1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) glucose, and 2% (w/v) agar) plates containing 100 lg/ml Zeocin (Invitrogen, Carlsbad, CA, USA). The presence of the target gene in the transformants was confirmed by colony PCR. For enzyme production, 3 ml of YPD medium (1% (w/v) yeast extract, 2% (w/v) peptone, and 2% (w/v) glucose) were inoculated with recombinant yeast and incubated at 30 °C, 250 rpm for 24 h. The starter inoculum was transferred at 0.1% (v/v) to 25 ml of BMGY medium (1% (w/v) yeast extract, 2% (w/v) peptone, 1% (v/v) glycerol, 100 mM potassium phosphate buffer, pH 6.0, 4  105% (w/v) biotin and 1.34% (w/v) YNB) and grown at 30 °C, 250 rpm until an OD600nm of 5–6 was reached. The cells were collected by centrifugation at 3000g for 5 min at room temperature and resuspended in 5 ml of BMMY (1% (w/v) yeast extract, 2% (w/v) peptone, 3% (v/v) methanol, 100 mM potassium phosphate buffer, pH 6.0, 4  105% (w/v) biotin and 1.34% (w/v) YNB). Methanol was added to a final concentration of 3% (v/v) every 24 h to maintain induction of the AOX promoter. The culture was incubated at 30 °C for 72 h with rotary shaking at 250 rpm. The supernatant fraction was separated by centrifugation at 6000g for 10 min and used as the crude enzyme preparation for experimental study. Proteins were analyzed by 12% SDS–PAGE and visualized by staining with Coomassie Blue G250. Total protein concentration was determined using BioRad’s Bradford assay (Bio-Rad, Hercules, CA, USA) using bovine serum albumin as a standard. 2.6. Enzyme activity assays 2.6.1. a-L-Arabinofuranosidase, cellulase, and xylanase The hydrolytic enzyme activities were determined based on the amount of reducing sugar released by the 3,5-dinitrosalicylic acid (DNS) method (Miller, 1959). Reactions of 1 ml total volume contained an appropriate dilution of enzyme in 50 mM sodium acetate buffer, pH 5.0 with 1% (w/v) of oat spelt xylan, carboxymethyl cellulose sodium salt, and beechwood xylan for a-Larabinofuranosidase, cellulase, and xylanase assay, respectively and incubated at 50 °C for 10 min. The amount of reducing sugars was determined by the absorbance at 540 nm using a VICTOR3 V microplate reader (Perkin Elmer, Waltham, MA, USA). Sugar concentrations were interpolated from standard curves of arabinose, glucose, and xylose. One unit of enzyme activity is defined as the amount of enzyme required to release 1 lmol of reducing sugars from a substrate in 1 min under the assay condition. 2.6.2. Pectin esterase Pectin esterase (PET) activity was measured by determining the carboxyl groups released by titration with 0.02 M NaOH according to a method modified from Kertez (Kertez, 1955). The 20 ml reaction contained 1% (w/v) pectin from citrus peel in 0.1 M NaCl, which was equilibrated to 50 °C and the pH adjusted to 7.5 with 0.02 M NaOH. The reaction was started by the addition of 200 ll PET. The amount of NaOH used to maintain the pH at 7.5 in the reaction period of 5 min was recorded. One unit of PET activity was defined as the amount of enzyme releasing one microequivalent of ester hydrolysed (carboxyl group) per min at pH 7.5 at 50 °C. 2.7. Synergistic enzyme interaction analysis Interactions among the recombinant enzymes and T. reesei cellulase (AccelleraseÒ 1500, Dupont, Rochester, NY, USA) were stud-

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

ied using experimental mixture design approach (Cornell, 2002). A {4,3}- augmented simplex lattice design implemented in the Minitab 17.0 software (Minitab Inc., State College, PA, USA) was used to define an optimal enzyme mixture for the maximal sugar yield. The design showed 25 experimental points, which were examined in quadruplicate with four components and a lattice degree of 3. In the mixture design, the sum of all components was 100% with the total enzyme loading of 0.5 mg/g biomass. The four components consisted of ACR, ARA, PET, and XYL. The reducing sugar yield was applied as a dependent variable for the analysis and simulation of the respondent model. The hydrolysis reactions contained 5% (w/v) alkaline-pretreated rice straw in 50 mM sodium acetate buffer, pH 5.0 containing 1 mM sodium azide and incubated at 50 °C for 48 h. After regression analysis, the full cubic model was used to simulate the responses to the mixture components and determine the optimal enzyme ratio. The amount of released reducing sugars was analysed using the DNS method. The sugar profile in the liquid fractions of the hydrolysis reaction was determined on a Waters e2695 high performance liquid chromatograph equipped with a differential refractometer using an Aminex HPX87H column (Bio-Rad, Hercules, CA, USA) operating at 65 °C with 5 mM H2SO4 as the mobile phase at a flow rate of 0.5 ml/min. The reactions were performed in quadruplicate. 3. Results and discussion 3.1. Cloning of ara, pet, and xyl genes from A. aculeatus The ara, pet and xyl genes were cloned from A. aculeatus BCC17849 by PCR using primers designed from A. aculeatus ATCC16872, in which it was assumed that the open reading frames are conserved among these strains. The cloned ara, pet and xyl gene sequences of 1008, 984, and 984 bp encode proteins of 336, 328, and 328 amino acid residues, respectively. The predicted proteins were 96.8–99.0% identical to their equivalents in A. aculeatus ATCC16872. The translated protein sequence of ara showed 81% identity to a glycosyl hydrolase family 62 (GH62) a-Larabinofuranosidase (axhA) of Aspergillus niger CBS 513.88 (GenBank No. XP_001389998). The encoded protein from pet had 83% identity to pectin esterase of Aspergillus kawachii IFO 4308 (GenBank No. GAA91050) and was classified in carbohydrate esterase family 8 (CE8). The translated protein of xyl was almost identical (99%) to an glycosyl hydrolase family 10 (GH10) endo-1,4-betaxylanase of A. aculeatus (GenBank No. O59859). N-terminal signal peptide encoding sequences of 19–26 amino acid residues were identified in all genes, suggesting that all of the enzymes are secreted. The predicted mature proteins ARA, PET, and XYL consist of 310, 309, and 309 amino acid residues with theoretical molecular weights of 33.36, 32.51, and 33.43 kDa, respectively. 3.2. Expression of recombinant ARA, PET, and XYL in P. pastoris Induction of the recombinant yeast strains using methanol led to expression of the target polypeptides as extracellular enzymes in the supernatant fraction (Fig. 1). The sizes of these proteins estimated from SDS–PAGE were consistent with the expected theoretical molecular weights of 34, 32, and 31 kDa for mature ARA, PET, and XYL, respectively. As the heterologous proteins expressed in the supernatant were essentially homogeneous, the supernatant was used directly for biochemical characterisation of enzyme activity with no further purification (Fig. 2). XYL worked optimally at 55 °C at pH 6.0 on hydrolysis on beechwood xylan with the activity of 328.91 U/ml in the culture supernatant, equivalent to the specific activity of 510.55 U/mg. The enzyme showed slight activity on hydrolysis of carboxymethylcellulose (0.004 U/mg)

685

Fig. 1. Expression of recombinant accessory enzymes in the crude supernatant fraction. The recombinant P. pastoris KM71 clones were induced with 3% methanol and incubated at 30 °C for 72 h with shaking at 250 rpm. Lanes: M, protein markers; 1, ARA; 2, PET; 3, XYL.

but with no activity on microcrystalline cellulose (Avicel). The optimal temperature of ARA was 50 °C with the highest activity of 41.41 U/ml, equivalent to 103.53 U/mg at pH 5.0. PET activity of 0.81 U/ml was found in the supernatant, equivalent to 2.47 U/mg under the assay conditions at 50 °C. However, the optimal condition of PET was not determined due to limitation in the enzyme assay method which was based on a titration technique under a fixed pH condition. All recombinant enzymes showed no detectable FPase activity under the experimental conditions.

3.3. Cooperative enzyme interaction on pretreated rice straw The mixture design approach was applied to study cooperative interactions among the core T. ressei cellulase and the accessory enzyme components and to optimise the composition of the enzyme components on hydrolysis of pretreated rice straw. The measured response of this method was presumed to depend only on the protein proportion of the complements in the mixture. The different proportion of the four components including ACR, ARA, PET, and XYL in relation to the reducing sugar yield from saccharification of alkaline pretreated rice straw was investigated. A {4,3}-simplex lattice model was created using the Minitab 17.0 software. All 25 experimental points were located inside the triangular graph, in which the sum of the four component loading for every experimental point was always 100%, which is equal to a total protein loading of 0.50 mg/g biomass. According to this design approach, the FPU loading/g substrate at each experimental point was varied according to different proportions of the core T. reesei cellulose, but with a fixed total protein loading in the reaction. Each point was operated in quadruplicate to minimise experimental variation. The relative reducing sugar yield of each experimental point after the hydrolysis of pretreated rice straw for 48 h is shown in Table 2. The hydrolysis reactions were incubated at 50 °C which were the optimal conditions for ACR and ARA while XYL showed >75% of its maximal activity. The T. reesei cellulase (ACR) was considered as the core activity on saccharification of the substrate in the experiment. From the results, the hydrolysis of pretreated rice straw using only ACR at 0.50 mg/g biomass, equivalent to 1.13 FPU/ g biomass under the standard conditions, led to a reducing sugar yield of 177.48 mg/g biomass. The binary combination between ACR and XYL at the ratio 2:1 by weight gave the highest reducing sugar yield of 235.11 mg/g biomass. When XYL was supplemented

686

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

Fig. 2. Effects of temperature and pH on activity of the recombinant enzymes. Reactions contained an appropriate dilution of enzyme with the respective substrate and incubated at the respective temperatures and pHs for 10 min. The enzyme activity was measured in 50 mM sodium acetate buffer, pH 5.0 for optimal temperature study and in 50 mM sodium citrate (pH 3–4); sodium acetate (pH 4–6); sodium phosphate (pH 6–8); and Tris–HCl (pH 8–9) at the optimal temperature of the respective enzyme for optimal pH study. (A) ARA; (B) XYL.

in the reaction containing ACR, the protein loading of ACR can be reduced by 33.4% concomitant with a 32.5% increase in the relative reducing sugar yield at an enzyme dosage of 0.75 FPU/g biomass. A high reducing sugar yield was also achieved by the combination of all enzymes in the ratio ACR:ARA:PET:XYL of 5:1:1:1. This combination enhanced the reducing sugar production to 208.97 mg/g, equivalent to a 17.7% increase from that of the ACR only reaction, whereas the ACR loading declined by 37.4% equivalent to an enzyme dosage of 0.70 FPU/g biomass. By focusing on the treatment with the lowest ACR loading, the ternary combination of ACR, ARA, and XYL at the ratio 1:1:1 resulted in a decrease of ACR loading by 66.7%, whereas the reducing sugar level was slightly decreased by 1.2% (175.32 mg/g) at the markedly reduced enzyme dosage of 0.376 FPU/g biomass. The response data for the reducing sugar yield was then analyzed by multiple regression analysis from the linear to the full cubic model. The full cubic model was found as the best fitted model for the relative sugar yield (R2 = 95.37%, PModel < 0.01). The enzyme loading in the mg/g biomass was fitted to the linear regression model (R2 = 100). The statistical coefficient values from the full cubic model regression analysis are shown in Table 3. These coefficients provide insight into the influence of each component and their interactions on the hydrolysis yield. When considering single factors, the ACR, ARA, PET, and XYL coefficients indicated a positive relation to the reducing sugar yield. For pairwise interactions, a significant synergism between ACR and XYL can be observed (308.6), while minor interactions between ARA and XYL (128.6), ACR and PET (104.1), and ACR and ARA (74.4) were also indicated at statistically signif-

icant values. Moreover, the highest coefficient was detected for the ternary combination of ACR * PET * ARA (968.3) followed by ACR * ARA * XYL (830.1), ARA * PET * XYL (791.5) and ACR * PET * XYL (458.7). Based on this model, most of factors were statistically significant (p < 0.05) except for seven terms including the PET * XYL, PET * ARA, ACR * ARA * (), ACR * PET * (), PET * XYL * (), ARA * PET * (), and ARA * XYL * (). The equation for the relative reducing sugar yield based on the component amount is deduced as:

Reducing sugar ðmg=g biomassÞ ¼361:52  ACR þ 124:55  XYL þ 45:76  PET þ 36:08  ARA þ 1246:82  ACR  XYL þ 395:05  ACR  PET þ 335:11  ACR  ARA þ 480:05  ARA  XYL þ 4462:20  ACR  PET  XYL þ 6366:82  ACR  ARA  XYL þ 7473:21  ACR  ARA  PET þ 7243:50  ARA  PET  XYL þ 1718:37  ACR  XYL  ðÞ

ð1Þ

From the reduced equation for the full cubic model, the response of the reducing sugar yield with respect to the various component combinations was graphically shown as ternary contour plots (Fig. 3). According to the plots, the area encompassing

687

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690 Table 2 Experimental design conditions and results. Mixture

Component (mg/g biomass)

ACR ALA PET XYL ACR1 ALA2 ACR2 ALA1 ACR1 PET2 ACR2 PET1 ACR1 XYL2 ACR2 XYL1 ALA1 PET2 ALA2 PET1 ALA1 XYL2 ALA2 XYL1 PET1 XYL2 PET2 XYL1 ACR1 ALA1 PET1 ACR1 ALA1 XYL1 ACR1 PET1 XYL1 ALA1 PET1 XYL1 ACR1 ALA1 PET1 XYL1 ACR5 ALA1 PET1 XYL1 ACR1 ALA5 PET1 XYL1 ACR1 ALA1 PET5 XYL1 ACR1 ALA1 PET1 XYL5

Reducing sugar

ACR

ARA

PET

XYL

(mg/g biomass)

SD

% Relative

SD

0.5 0 0 0 0.167 0.333 0.167 0.333 0.167 0.333 0 0 0 0 0 0 0.167 0.167 0.167 0 0.125 0.313 0.063 0.063 0.063

0 0.5 0 0 0.333 0.167 0 0 0 0 0.167 0.333 0.167 0.333 0 0 0.167 0.167 0 0.167 0.125 0.063 0.313 0.063 0.063

0 0 0.5 0 0 0 0.333 0.167 0 0 0.333 0.167 0 0 0.167 0.333 0.167 0 0.167 0.167 0.125 0.063 0.063 0.313 0.063

0 0 0 0.5 0 0 0 0 0.333 0.167 0 0 0.333 0.167 0.333 0.167 0 0.167 0.167 0.167 0.125 0.063 0.063 0.063 0.313

177.48 13.54 13.23 57.42 102.43 140.98 98.93 155.33 163.49 235.11 13.33 13.61 74.55 66.93 56.05 43.87 115.41 175.32 146.07 67.86 151.05 208.97 115.48 123.93 129.01

2.12 0.35 0.11 1.7 3.34 9.28 3.29 2.68 5.11 14.37 0.09 0.15 6.31 0.75 4.92 2.46 8.32 2.18 4.88 2.6 9.54 14.88 5.45 2.65 0.91

100 7.6 7.5 32.4 57.7 79.4 55.7 87.5 92.1 132.5 7.5 7.7 42 37.7 31.6 24.7 65 98.8 82.3 38.2 85.1 117.7 65.1 69.8 72.7

1.19 0.2 0.06 0.96 1.88 5.23 1.85 1.51 2.88 8.1 0.05 0.08 3.56 0.42 2.77 1.38 4.69 1.23 2.75 1.46 5.37 8.39 3.07 1.49 0.51

NA: not analyzed; NS: not supplemented; ACR: AccelleraseÒ 1500; ARA: a-L-arabinofuranosidase; PET: pectinesterase; XYL: xylanase. Protein concentration: ACR = 20.01 mg/ml; ARA = 0.40 mg/ml; PET = 0.43 mg/ml; XYL = 0.75 mg/ml/ ACR filter paper activity = 45.0 FPU/ml while other enzymes had no FPase activity. The total enzyme loading = 0.50 mg/g biomass. The arabic number means the ratio of enzymes. For example, ACR5 ARA1 PET1 XYL1 means that the ratio of Accellarase: a-Larabinofuranosidase: pectinesterase: xylanase is 5:1:1:1.

Table 3 The regression model analysis of the {4, 3} full cubic model developed for the relative reducing sugar yield (component proportion). Factor

Coefficient

SE

T

pValue

VIF

ACR XYL PET ARA ACR * ARA ACR * PET ACR * XYL ARA * PET ARA * XYL PET * XYL ACR * ARA * PET ACR * ARA * XYL ACR * PET * XYL ARA * PET * XYL ACR * ARA * () ACR * PET * () ACR * XYL * () ARA * PET * () ARA * XYL * () PET * XYL * () S = 14.7637 R2 = 95.37%

186.6 59.8 14.8 16.5 74.4 104.1 308.6 3.2 128.6 57.7 968.3 830.1 458.7 791.5 132.7 87.8 203.8 52.9 105.6 57.1 PRESS = 25011.6 R2 (pred) = 93.35%

7.353 7.353 7.353 7.353 32.903 32.903 32.903 32.903 32.903 32.903 222.434 222.434 222.434 222.434 67.593 67.593 67.593 67.593 67.593 67.593

* * * * 2.260 3.160 9.380 0.100 3.910 1.750 4.350 3.730 2.060 3.560 1.960 1.300 3.020 0.780 1.560 0.840

* * * * 0.026 0.002 0.000 0.922 0.000 0.084 0.000 0.000 0.042 0.001 0.053 0.198 0.003 0.437 0.122 0.401

3.472 3.472 3.472 3.472 2.783 2.783 2.783 2.783 2.783 2.783 1.730 1.730 1.730 1.730 1.176 1.176 1.176 1.176 1.176 1.176

R2 (adj) = 94.27%

the greatest relative sugar yield was located almost at the top of the ACR axis, in the middle of the XYL vertex, and near the bottom of both the ARA and the PET axes. From these results, a high reducing sugar yield could be accomplished when the concentration of ACR was two times greater than that of XYL while lower amounts of ARA and PET were required. The optimum combinations of the enzyme mixture were in the range of 0.20–0.50 mg/g biomass for ACR, 0.04–0.24 mg/g biomass for XYL, and 0–0.10 mg/g biomass for both ARA and PET.

The optimal mixture combination was determined by the optimal mixture Minitab 17.0 software. The first combination was defined as the point where the maximal sugar yield was achieved, which was predicted to be 0.374 mg/g biomass ACR and 0.126 mg/ g biomass XYL, with the expected relative reducing sugar yield of 231.84 mg/g biomass at 0.84 FPU loading/g (Fig. 4A). The experimentally determined yield under these conditions was 206.43 mg/g biomass that was slightly lower than that of the predicted value (10.96%). Supplementation of XYL resulted in an increase in glucose (102.50 ± 6.76 mg/g) and xylose (31.54 ± 7.13 mg/g) in the hydrolysate, which were equivalent to 45.1% and 255.2% increases, respectively compared with those obtained with ACR alone at the same FPU dosage. This was equivalent to an increase in glucose releasing efficiency from 0.081 gglc/ FPU using ACR alone to 0.122 gglc/FPU by the binary enzyme mixture. Stepwise increase in the total enzyme dosage up to 6-fold showed an increasing trend for sugar yield. The highest reducing sugar yield of 520 ± 20 mg/g was achieved at 6 enzyme loading, equivalent to 5.05 FPU/g biomass. This result thus indicates the effectiveness of the optimized enzyme formulation at a higher cellulase range that can produce a practical yield of sugar for bio-industry. Another optimal point was defined as the enzyme combination where the lowest dosage of FPU was used to give the sugar yield at least the same as that obtained using 100% ACR. This was predicted as 0.171 mg/g biomass of ACR supplemented with 0.150 mg/g biomass XYL, 0.100 mg/g biomass PET, and 0.079 mg/g biomass ARA, which gave a predicted theoretical sugar yield of 180.76 mg/g biomass (Fig. 4B). The experimental sugar yield was 172.69 mg/g biomass. The results thus indicate a 65.8% saving of ACR can be obtained by supplementing the reaction with synergistic accessory enzymes. In the reaction containing ACR and accessory enzymes in the optimal ratio, higher yields were obtained for glucose (66 ± 2 mg/g) and xylose (21 ± 1 mg/g), which were equivalent to

688

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

Fig. 3. The ternary plots of the experimental design optimization of the four enzyme complex. The contour plot of the relative reducing sugar yield of (A) ACR:PET:XYL; (B) ACR:ARA:XYL; (C) ACR:ARA:PET; (D) ARA:PET:XYL.

47.3% and 114.5% increase compared with those obtained using ACR alone at the same FPU dosage. Addition of ARA into the enzyme system also led to the release of a substantial amount of arabinose (19 ± 2 mg/g) into the hydrolysate. This resulted in the increase in glucose releasing efficiency of 0.173 gglc/FPU by the quaternary enzyme mixture. Stepwise increase of the total enzyme loading led to a concomitant increase of sugar yield. The highest reducing sugar yield of 340 ± 10 mg/g was achieved at 6 enzyme loading equivalent to 2.30 FPU loading/g. The result thus indicates that enzyme mixture is effective within a broad range of cellulase loading. Many works have shown the synergistic actions among endo-/exo-acting glycosyl hydrolases in different families or to other carbohydrate processing enzymes, for example, the synergism of cellobiohydrolases, endoglucanases, and b-glucosidase on hydrolysis of cellulosic substrates (Kostylev and Wilson, 2012), synergistic action of xylanase, esterase and mannanase on cellulase hydrolysis of pretreated corn stover (Hu et al., 2011) and softwood (Várnai et al., 2011), the synergism of cellulase and pectinase in the hydrolysis of hemp (Zhang et al., 2013), and the synergistic enzyme cocktail containing endoglucanase I, cellobiohydrolase I and II, bglucosidase, endo-xylanase, and b-xylosidase on digesting of ammonia fiber expansion (AFEX) treated corn stover (Gao et al., 2010). Varying degree of synergy between these enzymes have been demonstrated using statistical methods. Synergisms between the core cellulase (ACR), and the accessory enzymes attacking the non-cellulosic part of the cell wall have been demonstrated in our study. GH10 endo-b-1,4-xylanases are major group of glycosyl hydrolases attacking xylan, the most abundant hemicellulose in plant cell wall. Some GH10 xylanases have been reported to possess a side-activity against cellulosic substrates e.g. CMC and pNP-cellobioside (Biely et al., 1997). The enzyme XYL from A. aculeatus in our work showed relatively

similar optimal working condition to some previously reported xylanases e.g. those from Aspergillus and Trichoderma in industry (Fengxia et al., 2008) as well as the recombinant GH10 xylanase from A. terreus expressed in P. pastoris (Chantasingh et al., 2006). a-L-Arabinofuranosidases (EC 3.2.1.55) are classified into different GH families 43, 51, 54, and 62 according to CAZY. They are exo-acting enzymes that remove L-arabinofuranosyl side chain moieties from the main chain xylan and result in enhancement of the action of the other enzymes such as xylanases and b-xylosidases (Vincent et al., 1997). Several GH62 a-arabinofuranosidases have been isolated from bacterial and fungal sources, particularly from the genus Aspergillus. The ARA enzyme from A. aculeatus showed no carbohydrate binding module 1 (CBM1), which has been found in several homologous enzymes from A. fumigatus (Nierman et al., 2005), and P. fumicolosum (De La Mare et al., 2013). In contrast to hydrolytic glycosyl hydrolases, pectin esterases (EC 3.1.1.11) is classified as a carbohydrate esterase family (CE) member which catalyses the de-esterification of pectin and work cooperatively with polygalacturonases and pectate lyases on degradation of pectin. Pectin esterases from various fungal strains have been studied, Fusarium asiaticum (Glinka and Liao, 2011), and yeast Wickerhanomyces anomalus (Martos et al., 2013). A recombinant pectin methyl esterase from A. aculeatus has been expressed in Saccharomyces cerevisiae under an inducible GAL1 promoter (Christgau et al., 1996) while the enzyme from A. niger has also been produced in P. pastoris under a constitutive GAP promoter with a potential use in fruit juice clarification (Jiang et al., 2013) . The physical and chemical properties of the carbohydrate components in lignocellulosic biomass are dependent on the botanical origin of the agricultural residues and the pretreatment methods used for increasing their digestibility. Rice straw typically contains cellulose as the dominant composition (32–47%) followed by the heterogenous hemicellulose (19–27%) composed of xylose

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

Fig. 4. Validation of mixture design experiment with varying total amount of enzyme loading at the optimised ratios of the composite enzyme components. Reactions contained 5% (w/v) alkaline-pretreated rice straw in 50 mM sodium acetate buffer, pH 5.0 containing 1 mM sodium azide and incubated at 50 °C for 48 h with the total enzyme loading of 0.50 mg/g biomass (1) to 3.0 mg/g biomass (6). (A) the binary enzyme mixture (ACR:XYL = 0.374:0.126) for the maximal sugar yield; (B) the quaternary enzyme mixture (ACR:XYL:ARA: PET = 0.171:0.150:0.100:0.079) for the lowest loading of ACR. The line graph represented the reducing sugars and the bar graph showed the monomeric sugar profiles.

(14.8–20.2%), arabinose (2.7–4.5%), and mannose (1.8%) based on the polysaccharide content (Karimi et al., 2006). Alkaline pretreatment is an efficient method used for increasing the digestibility of lignocelluloses by delignification with slight effects on the hemicellulosic fraction, which overall, results in increased accessibility to the cellulose fibers. Strong synergisms between the core cellulase (ACR) to the accessory hemicellulolytic and pectinolytic enzymes and among the accessory enzymes themselves were

689

shown in this study. The most prominent synergism of XYL to ACR on saccharification of alkaline-pretreated rice straw would reflect the degradation of xylan, the major hemicellulose in most plant biomass by the action of XYL which led to increased accessibility of the core cellulase in ACR to attack the cellulosic fibers in the substrate. This resulted in enhanced reducing sugar yield as well as increased glucose and pentose releasing efficiency from the hydrolysis reaction. Complementing ACR with all accessory enzymes as a quaternary enzyme mixture showed strong synergism among the enzyme components. This could be related to the action of ARA on hydrolyzing the arabinosyl branch on the main chain of xylan-arabinoxylan polymers in rice straw. Although the content of arabinose in rice straw is relatively low, this enzyme could act cooperatively with xylanase and other enzymes on increasing accessibility of the cellulolytic enzymes to the substrate. However, only a limited number of studies on synergism of a-arabinofuranosidases on hydrolysis of lignocellulosic substrates have been reported. These included the enhancement of xylose recovery by arabinofuranosidase mutant from Pleutus ostreatus on hydrolysis of pretreated Arundo donax, corn cobs, and brewer’s spent grains (Marcolongo et al., 2014), synergistic effects of Penicillium arabinofuranosidase on hydrolysis of rice straw by cellulase and xylanase (Lee et al., 2011), and the combination of a-arabinofuranosidase to mannanase and xylanase on hydrolysis of bagasse (Beukes and Pletschke, 2010). In addition to the action of hydrolytic enzymes, PET can also further increase the efficiency of the enzyme systems by de-esterification of the pectin component, which acts as the embedded matrix in the cell wall structure. This finding was different to a previous work on hydrolysis of pretreated sugarcane bagasse by the mixture of T. reesei cellulase and pectinase in which no synergism was found between the two enzymes, resulting in no increasing glucose production (Li et al., 2014). Addition of a commercial pectin methyl esterase was reported to increase ethanol yield from fermentation of sugar beet pulp by engineered E. coli KO11 expressing cellobiase and pectate lyase (Edwards et al., 2011) . The synergy between the enzymes in the developed system led to a marked increase in glucose releasing efficiency of the quaternary enzyme mixture (0.173 gglc/FPU) and the binary ACR + XYL system (0.122 gglc/FPU) compared with the ACR alone reaction (0.081 gglc/FPU). The glucose releasing efficiency of the optimized enzyme system in this study is also considerably higher than the T. reesei enzyme cocktail (0.021 gglc/FPU) comprising T. reesei cellobiohydrolase enriched fraction, endoglucanase, and b-glucosidase previously reported by Bussamra et al. (2015). However, it should be noted that these observations cannot be directly compared owing to variation in the substrates, enzyme combination, and reaction conditions. This study suggests an optimal ratio of accessory enzyme to cellulase enzyme that can promote efficient biomass saccharification. The data thus demonstrated the strong synergism of the enzyme components attacking cellulose, hemicellulose, and pectin in the optimized enzyme mixture.

4. Conclusion A synergistic enzyme system comprising the core cellulolytic enzyme supplemented with accessory hemicellulases and pectinase has been developed in this study. Strong synergism between the composite enzyme components on hydrolysis of rice straw has been demonstrated. The work provides insights into synergistic and cooperative action of core and accessory enzymes on hydrolysis of plant biomass, which can be further applied for developing efficient enzyme systems for saccharification or modification of lignocelluloses in bio-industry.

690

T. Laothanachareon et al. / Bioresource Technology 198 (2015) 682–690

Acknowledgements This project was financially supported by the National Science and Technology Development Agency (Grant number P-1550502). Manuscript proofreading by Dr. Philip J. Shaw is appreciated.

References Bayer, E.A., Belaich, J.P., Shoham, Y., Lamed, R., 2004. The cellulosomes: multienzyme machines for degradation of plant cell wall polysaccharides. Annu. Rev. Microbiol. 58, 521–554. Beukes, N., Pletschke, B.I., 2010. Effect of lime pre-treatment on the synergistic hydrolysis of sugarcane bagasse by hemicellulases. Bioresour. Technol. 101, 4472–4478. Biely, P., Vrsanska, M., Tenkanen, M., Kluepfel, D., 1997. Endo-b-1-4-xylanase families: differences in catalytic properties. J. Biotechnol. 57, 151–166. Bunterngsook, B., Eurwilaichitr, L., Thamchaipenet, A., Champreda, V., 2015. Binding characteristics and synergistic effects of bacterial expansins on cellulosic and hemicellulosic substrates. Bioresour. Technol. 176, 129–135. Bussamra, B.C., Freitas, S., da Costa, A.C., 2015. Improvement on sugar cane bagasse hydrolysis using enzymatic mixture designed cocktail. Bioresour. Technol. 187, 173–181. Chantasingh, D., Champreda, V., Kanokratana, P., Eurwilaichitr, L., 2006. Cloning, expression and characterization of a xylanase 10 from Aspergillus terreus (BCC129) in Pichia pastoris. Protein Expr. Purif. 46, 143–149. Cherubini, F., 2010. The biorefinery concept: Using biomass instead of oil for producing energy and chemicals. Energ. Convers. Manage. 51, 1412–1421. Christgau, S., Kofod, L.V., Halkier, T., Andersen, L.N., Hockauf, M., Dörreich, K., Dalbǿge, H., Kauppinen, S., 1996. Pectin methyl esterase from Aspergillus aculeatus: expression cloning in yeast and characterization of the recombinant enzyme. Biochem. J. 319, 705–712. Cornell, J., 2002. Experiments with Mixtures: Designs, Models, and the Analysis of Mixture Data, third ed. Wiley, USA. De La Mare, M., Guais, O., Bonnin, E., Weber, J., Francois, J.M., 2013. Molecular and biochemical characterization of three GH62 a-l-arabinofuranosidases from the soil deuteromycete Penicillium funiculosum. Enzyme Microb. Technol. 53, 351– 358. Edwards, M.C., Henriksen, E.D., Yomano, L.P., Gardner, B.C., Sharma, L.N., Ingram, L. O., Peterson, J.D., 2011. Addition of genes for cellobiase and pectinolytic activity in Escherichia coli for fuel ethanol production from pectin-rich lignocellulosic biomass. Appl. Environ. Microbiol. 77, 5184–5191. Fengxia, L., Mei, L., Zhaoxin, L., Xiaomei, B., Haizhen, Z., Yi, W., 2008. Purification and characterization of xylanase from Aspergillus ficuum AF-98. Bioresour. Technol. 99, 5938–5941. Gao, D., Chundawat, S.P.S., Krishnan, C., Balan, V., Dale, B.E., 2010. Mixture optimization of six core glycosyl hydrolases for maximizing saccharification of ammonia fiber expansion (AFEX) pretreated corn stover. Bioresour. Technol. 101, 2770–2781. Glinka, E.M., Liao, Y.C., 2011. Purification and partial characterisation of pectin methylesterase produced by Fusarium asiaticum. Fungal Biol. 115, 1112–1121. Gottschalk, L.M.F., Oliveira, R.A., Bon, E.P.D.S., 2010. Cellulases, xylanases, bglucosidase and ferulic acid esterase produced by Trichoderma and Aspergillus act synergistically in the hydrolysis of sugarcane bagasse. Biochem. Eng. J. 51, 72–78.

Himmel, M.E., Ding, S.-Y., Johnson, D.K., Adney, W.S., Nimlos, M.R., Brady, J.W., Foust, T.D., 2007. Biomass recalcitrance: engineering plants and enzymes for biofuels production. Science 315, 804–807. Howard, R.L., Abotsi, E., Jansen van Rensburg, E.L., Howard, S., 2003. Lignocellulose biotechnology: issues of bioconversion and enzyme production. Afr. J. Biotechnol. 2, 602–619. Hu, J., Arantes, V., Saddler, J.N., 2011. The enhancement of enzymatic hydrolysis of lignocellulosic substrates by the addition of accessory enzymes such as xylanase: is it an additive or synergistic effect? Biotechnol. Biofuels 4, 36. Jiang, X., Chen, P., Yin, M., Yang, Q., 2013. Constitutive expression, purification and characterisation of pectin methylesterase from Aspergillus niger in Pichia pastoris for potential application in the fruit juice industry. J. Sci. Food Agric. 93, 375– 381. Karimi, K., Kheradmandinia, S., Taherzadeh, M.J., 2006. Conversion of rice straw to sugars by dilute-acid hydrolysis. Biomass Bioenerg. 30, 247–253. Kertez, Z.I., 1955. Pectic enzymes. Methods Enzymol. 1, 158–166. Kostylev, M., Wilson, D., 2012. Synergistic interactions in cellulose hydrolysis. Biofuels 3, 61–70. Lee, D.-S., Wi, S.-G., Lee, Y.-G., Cho, E.-J., Chung, B.-Y., Bae, H.-J., 2011. Characterization of a new a-L-arabinofuranosidase from Penicillium sp. LYG 0704, and their application in lignocelluloses degradation. Mol. Biotechnol. 49, 229–239. Leggio, L.L., Simmons, T.J., Poulsen, J.C.N., Frandsen, K.E.H., Hemsworth, G.R., Stringer, M.A., Freiesleben, P.V., Tovborg, M., Johansen, K.S., De Maria, L., Harris, P.V., Soong, C.L., Dupree, P., Tryfona, T., Lenfant, N., Henrissat, B., Davies, G.J., Walton, P.H., 2015. Structure and boosting activity of a starch-degrading lytic polysaccharide monooxygenase. Nat. Commun. 6, 5961. Li, J., Zhou, P., Liu, H., Xiong, C., Lin, J., Xiao, W., Gong, Y., Liu, Z., 2014. Synergism of cellulase, xylanase, and pectinase on hydrolyzing sugarcane bagasse resulting from different pretreatment technologies. Bioresour. Technol. 155, 258–265. Marcolongo, L., Ionata, E., La Cara, F., Amore, A., Giacobbe, S., Pepe, O., Faraco, V., 2014. The effect of Pleurotus ostreatus arabinofuranosidase and its evolved variant in lignocellulosic biomasses conversion. Fungal Genet. Biol. 72, 162– 167. Martos, M., Zubreski, E.R., Garro, O.A., Hours, R.A., 2013. Production of pectinolytic enzymes by the yeast Wickerhanomyces anomalus isolated from citrus fruits peels. Biotechnol. Res. Int. 2013, 1–7. Miller, G.L., 1959. Use of dinitrosalicylic acid reagent for determination of reducin sugar. Anal. Chem. 31, 426–428. Nierman, W.C., Pain, A., Anderson, M.J., Wortman, J.R., Kim, H.S., Arroyo, J., et al., 2005. Genomic sequence of the pathogenic and allergenic filamentous fungus Aspergillus fumigatus. Nature 438, 1151–1156. Sluiter, A., Crocker, D., Hames, B., Ruiz, R., Scarlata, C., Sluiter, J., Templeton, D., 2008. Determination of Structural Carbohydrates and Lignin in Biomass. NREL/TP510-42618. Várnai, A., Huikko, L., Pere, J., Siika-Aho, M., Viikari, L., 2011. Synergistic action of xylanase and mannanase improves the total hydrolysis of softwood. Bioresour. Technol. 102, 9096–9104. Vincent, P., Shareck, F., Dupont, C., Morosoli, R., Kluepfel, D., 1997. New a-Larabinofuranosidase produced by Streptomyces lividans: cloning and DNA sequence of the abfB gene and characterization of the enzyme. Biochem. J. 322, 845–852. Willats, W.G.T., McCartney, L., Mackie, W., Knox, J.P., 2001. Pectin: cell biology for functional analysis. Plant Mol. Biol. 47, 9–27. Zhang, J., Pakarinen, A., Viikari, L., 2013. Synergy between cellulases and pectinases in the hydrolysis of hemp. Bioresour. Technol. 129, 302–307. Zhang, Y.H., Lynd, L.R., 2006. A functionally based model for hydrolysis of cellulose by fungal cellulase. Biotechnol. Bioeng. 94, 888–898.