High-loading oil palm empty fruit bunch saccharification using cellulases from Trichoderma koningii MF6

Process Biochemistry 49 (2014) 673–680

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High-loading oil palm empty fruit bunch saccharification using cellulases from Trichoderma koningii MF6 Zunsheng Wang, Huiling Bay, Kityeng Chew, Anli Geng ∗ School of Life Sciences and Chemical Technology, Ngee Ann Polytechnic, Singapore

a r t i c l e

i n f o

Article history: Received 19 June 2013 Received in revised form 31 December 2013 Accepted 21 January 2014 Available online 30 January 2014 Keywords: Cellulose T. koningii MF6 Biomass hydrolysis Oil palm empty fruit bunch High biomass loading

a b s t r a c t Strain Trichoderma koningii D-64 was improved for enhanced cellulase production. A potential mutant MF6 was obtained and its enzymes contained filter paper cellulase (FPase), carboxymethylcellulase (CMCase), ␤-glucosidase and xylanase with respective activities of 2.0, 1.3, 2.0 and 3.0 folds of those for the parental strain. MF6 cellulases showed enhanced hydrolysis performance for the treated lignocellulosic biomass. Hydrolysis of treated oil palm empty fruit bunch (OPEFB), horticulture wastes (HW) and wood chips (WC) resulted in cellulose to glucose conversion of 96.3 ± 2.2%, 98.2 ± 3.0% and 81.9 ± 1.4%, respectively. The corresponding conversions of xylan to xylose were 96.9 ± 1.5%, 95.0 ± 2.2% and 76.1 ± 3.1%. Consistently, high sugar yield of 770–844 mg/g biomass was obtained for high-loading (10–16%, w/v) of OPEFB hydrolysis and sugar titer of 135.1 g/L was obtained for 16% (w/v) OPEFB loading at 96 h. In addition, MF6 enzymes alone performed equally well for high-loading OPEFB hydrolysis compared to the enzyme mixture of ␤-glucosidase from Aspergillus niger and cellulase from T. reesei Rut C30. © 2014 Elsevier Ltd. All rights reserved.

1. Introduction With the growing concern about worldwide shortage of fossil fuels, the emission of greenhouse gases and air pollution by incomplete combustion of fossil fuels, there are continuous interests in the exploration of sustainable bioenergy production. Lignocellulosic biomass from agricultural crop residues, grasses, wood and municipal solid waste is abundant and renewable. It accounts for about 50% of all the biomass available in the world asserting its potential as a feasible raw material for the production of future fuels [1]. Biomass conversion to biofuels involves biomass pretreatment, biomass hydrolysis and hydrolysate fermentation. Availability of highly active cellulolytic enzymes is one of the most important objectives to achieve cost-effective production of cellulosic biofuels. Cellulase is an enzyme complex found in fungi [2] and bacteria [3]. The cellulolytic enzyme complex consists of three types of enzymes that act synergistically in cellulose hydrolysis. Endoglucanases randomly attack cellulose chains and release cello-oligosaccharides, exoglucanases or cellobiohydrolases cleave cellobiose units from the end of cellulose chain and ␤-glucosidase converts the resulted cellobiose to glucose. Filamentous fungi,

∗ Corresponding author at: School of Life Sciences and Chemical Technology, Ngee Ann Polytechnic, 535 Clementi Road, Singapore 599489, Singapore. Tel.: +65 64608617; fax: +65 64679109. E-mail addresses: [email protected], [email protected] (A. Geng). 1359-5113/$ – see front matter © 2014 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.procbio.2014.01.024

typically Trichoderma reesei, are the preferred sources of industrial cellulase preparations because they have excellent capacity for extracellular protein secretion and are therefore the best known cellulolytic enzymes producers [4]. It is well known that T. reesei produces the cellobiohydrolase and endoglucanase components of cellulase enzyme complex in large quantities. However, the amount of ␤-glucosidase secreted is insufficient [5]. As a result, the commercially available cellulase produced from T. reesei strains is often supplemented with ␤-glucosidase to attain good cellulase hydrolysis performance [6]. Filamentous fungus T. koningii produces the cellulolytic enzymes and their characteristics were studied [7]. However, to-date, very few reports can be found on its application for lignocellulosic biomass hydrolysis. Recently, our research group managed to isolate a cellulolytic T. koningii D-64 strain [8]. It secretes significant amount of cellulolytic enzymes with proper ratio of ␤-glucosidase and filter paper cellulase (FPase) making it efficient in lignocellulose hydrolysis. However, the enzyme activity is too low to be practically used. The use of progressive and step-wise mutagenesis selection protocols with varied mutagens has been proven effective in increasing enzyme yield [9–11]. Based on this technique, many fungal mutants for industrial scale cellulase production were developed such as T. reesei Rut C30, T. reesei CL 847 [9,10] and Acremonium cellulolyticus CF2612 [11]. Recently, protoplast-fusion-based genome shuffling was successfully introduced for strain improvement [12] and cellulase production was remarkably increased using this technique [13–15]. For the economical production of biofuel from lignocellulosic

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materials, the improvement of cellulase secreting microorganisms is of vital importance and an efficient approach is the development and selection of cellulase hyper-producing mutants. Trichoderma koningii D-64 is a promising cellulolytic fungus discovered in our research group [8]. In order to further enhance its enzyme production and biomass hydrolysis performance, T. koningii D-64 was treated by random mutagenesis, intra-strain protoplast fusion and genome shuffling. Five mutants were developed and their capacity of producing cellulase was investigated. The hydrolysis performance of cellulases from these mutants was assessed using the pretreated oil palm empty fruit bunch. The hydrolysis characteristics of cellulase from mutant MF6 was further evaluated using oil palm empty fruit bunch, wood chips and horticultural waste. Moreover, OPEFB hydrolysis was conducted with high biomass loading using MF6 cellulases. 2. Materials and methods 2.1. Microorganisms and cultivation conditions Wild strain T. koningii D-64 was isolated from soil samples in Singapore. T. reesei Rut C30 (ATCC 56765) was obtained from the American Type Culture Collection (ATCC). The above fungal strains were maintained in potato dextrose agar (Merck, Germany) plates with 0.5% cellulose. Selection agar medium 1 (SM1) contained basal medium (BM) described by Mandels and Weber [16] and was supplemented with 0.5% (w/v) 2-deoxyglucose, 1.0% (w/v) phosphoric acid-swollen cellulose, 0.1% (w/v) Triton X-100, and 2.0% (w/v) agar. In selection agar medium 2, 0.5% (w/v) 2deoxyglucose in SM1 was replaced with 1% (w/v) glucose (SM2) and in selection agar medium 3 (SM3) it was replaced with 5% (w/v) glucose. Seed medium was based on BM with 1% (w/v) lactose as the carbon source. CWB fermentation medium contained the same amount of minerals in BM and was supplemented with 1% cellulose and 1.4% (w/v) wheat bran. On the other hand, CG fermentation medium contained twice the amount of minerals in BM and was supplemented with 2% cellulose and 0.5% (w/v) glucose. Whenever necessary, 0.08% (w/v) potassium sodium tartrate was added for pH adjustment. One milliliter suspended spores (1 × 108 spores) from agar plates were inoculated in 50 mL of seed medium in 250-mL conical flasks and incubated at 30 ◦ C, 150 rpm for 48 h. Five milliliter inoculants were transferred to 50 mL of fermentation medium in 250-mL conical flasks and incubated at 120 rpm and 30 ◦ C for 5 days in CWB medium and 7 days in CG medium. Samples were centrifuged at 5000 × g at 4 ◦ C for 5 min. Supernatants were used for the enzyme activity and soluble protein content assay. 2.2. Developing diversities of mutants for genome shuffling Fungal spores were treated by UV light or ethyl methane sulfonate (EMS) according to Durand and Clanet [9]. One milliliter spore saline suspension (107 spores/mL, 0.9% NaCl) was exposed to UV light for 3 min (UV light from money detector) and stayed in dark for 1 h. In the meantime, 0.06 mL EMS was mixed with 1 mL 107 spores/mL suspension saline (0.9% NaCl) and incubated at 37 ◦ C for 50 min. The fatal rate for both mutagenesis processes was above 99%. The treated spores were dispersed on SM1 or SM2 agar plates. The colonies which rapidly presented clear zones were evaluated by shaking flask cultivation and enzyme activity assay. They were then selected for further improvement. 2.3. Protoplast preparation Potential mutants were grown in seed medium for 48 h and harvested by filtration with four layer lens wiping paper. They were then washed twice with sterilized distilled H2 O. Five hundred mg cell pellets were suspended in 30 mL protoplast formation buffer 1 (0.1 M phosphate buffer at pH 5.8 with 0.6 M KCl) and mixed with 15 mg/mL snailase (Shanghai Qcbio Science & Technologies Co. Ltd., China). The enzymatically hydrolysed cells were incubated on orbital shaker at 35 ◦ C and 80 rpm for 1–1.5 h. The concentration of the released protoplast was above 1 × 106 /mL. The mixture of enzyme and protoplasts was filtered through four layer lens wiping paper and centrifuged at 2000 × g and 4 ◦ C for 8 min. The supernatant was discarded and the protoplast sediment was washed once with buffer 1 and then was re-suspended in buffer 1 to 1 × 107 /mL. They were then regenerated on the regeneration medium, SM2 or SM3. 2.4. Intra-strain protoplast fusion Protoplasts were isolated from buffer 1 containing snailase as described. Equal numbers of protoplasts from each mutant were mixed and centrifuged at 700 × g for 10 min. The pellet was re-suspended in buffer 2 (10 mM pH7.5 Tris–HCl with 50 mM CaCl2 and 0.6 M sorbitol) with 40% polyethylene glycol 6000 (PEG) and incubated at 35 ◦ C for 40 min. After protoplast fusion, the fused protoplasts were centrifuged

Table 1 The chemical composition of the pretreated biomass. Biomass

Cellulose (%)

Xylans (%)

Lignin (%)

Treated OPEFB Treated HW Treated WC

53.2 ± 3.5 53.4 ± 2.9 39.5 ± 3.9

21.9 ± 1.7 11.4 ± 1.5 16.4 ± 1.6

13.1 ± 2.0 15.1 ± 2.5 23.6 ± 3.5

and washed once with buffer 2, and then re-suspended in 10 mL buffer 2. After this, 0.1 mL proper diluted fused protoplasts was spread on SM2 and regenerated at 30 ◦ C. The regenerated colonies with fast growth and clear halos of cellulose hydrolysis were selected for shaking flask cultivation and enzyme activity assay. 2.5. Genome shuffling Genome shuffling was carried out by the described method with modifications [15]. Equal number of protoplasts from mutants were mixed, divided equally into two parts and inactivated. One part was treated with UV irradiation (light from money detector) for 5 min and the other part was inactivated in 50 ◦ C for 50 min. Both treatments made the protoplasts lose their viability. These protoplasts from different treatment were then mixed and centrifuged at 700 × g for 10 min. The pellet was re-suspended in buffer 2 with PEG and incubated at 35 ◦ C for 40 min. The fused protoplasts were centrifuged and washed once with buffer 2, and then re-suspended in 5 mL buffer 2. Afterwards, 0.1 mL proper diluted protoplasts was spread on SM3 and regenerated at 30 ◦ C. The regenerated colonies with fast growth and clear halos of cellulose hydrolysis were selected for shaking flask cultivation and enzyme activity assay. 2.6. Enzyme activity and protein content assay Filter paper cellulase (FPase) activity was determined as described by incubating 0.5 mL of the suitably diluted culture samples with 1.0 mL 50 mM citrate buffer (pH 4.8) and Whatman No.1 filter paper strips (50 mg, 1 × 6 cm) at 50 ◦ C for 60 min [17]. Endoglucanase or carboxymethylcellulase activity (CMCase) was carried out by using carboxymethylcellulose (CMC) in the total reaction mixture of 1.5 mL containing 0.5 mL of suitably diluted enzyme samples and 1.0 mL 1% (w/v) CMC (lower viscosity, Hercules Inc., USA) solution in 50 mM citrate buffer (pH 4.8) [17]. The mixture was incubated at 50 ◦ C for 30 min. Endoxylanase (xylanase) activity was determined by catalyzing of 1.0 mL of 1% (w/v) xylan (xylan from Birchwood, Sigma, USA) in 50 mM phosphate buffer (pH 6.5) by 0.5 mL of suitably diluted enzymes at 40 ◦ C for 10 min [18]. ␤-Glucosidase activity was estimated using p-nitro phenyl ␤-d-glucopyranoside (pNPG) (Sigma) as the substrate according to the method described by Berghem and Pettersson [19]. The 1 mL assay mixture contained 0.9 mL of 1 mM pNPG and 0.1 mL of suitably diluted enzyme samples and it was incubated at 50 ◦ C for 10 min. The reaction was terminated by the addition of 0.5 mL 1 M sodium carbonate. Reducing sugars liberated during reactions on filter paper, CMC and xylan were quantified by dinitrosalicylic acid (DNS) method using glucose or xylose as the standard [20]. The p-nitrophenol liberated on pNPG was quantified by sodium carbonate using p-nitrophenol as the standard. One enzyme unit (U) was defined as the amount of enzyme that releases 1 ␮mole of the product from their corresponding substrate per min under the assay conditions. Total extracellular protein content was measured by the Lowry Protein Assay using bovine serum albumin as the standard [21]. All analyses were done in triplicate. Results were expressed as mean ± standard deviation. 2.7. Random amplified polymorphic DNA (RAPD) Genomic DNA was isolated from parent strain and its mutants using a commercial DNA extraction kit according to the manufacturer’s instructions (Promega). Random Amplified Polymorphic DNA were obtained by a mixture of custom synthesized primers reported previously [22]. The sequences of the random primers, MAP-06 to MAP-10, were 5 -GCACGCCGGA-3 , 5 -CACCCTGCGC-3 , 5 -CTATCGCCGC3 , 5 -CGGGATCCGC-3 , 5 -GCGAATTCCG-3 , respectively. 2.8. Biomass pretreatment by alkaline hydrogen peroxide Oil palm empty fruit bunch (OPEFB) was obtained from an oil palm plantation in Malaysia. Horticulture waste (HW) was from ecoWise Solution Pvt Ltd. and wood chips (WC) were obtained from the landscape industry in Singapore. Before chemical pretreatment, the collected biomass was washed, stored and milled according to our previous report [23]. Ten percent (w/v) of the milled biomass particles (<500 ␮m) in 1% (w/v) NaOH solution was autoclaved at 121 ◦ C for 30 min. The slurry was washed with distilled water and re-suspended to the original volume. The pH value was adjusted to 11.5 using 4 M NaOH. Hydrogen peroxide was added at final concentration of 4% (v/v). The alkaline hydrogen peroxide treatment was performed at 50 ◦ C overnight. The treated lignocelluloses was washed to neutrality and stored at 4 ◦ C until use. The composition of the pretreated lignocelluloses was determined according to the National Renewable Energy Laboratory (NREL, Golden, CO) biomass analytical methods [24] and the results are listed in Table 1. The amount

Z. Wang et al. / Process Biochemistry 49 (2014) 673–680

3.2. Effects of carbon sources on enzyme production Cellulose was found to be the best carbon source for T. koningii D-64 for cellulase production [8]. In order to evaluate enzyme production capabilities, T. reesei Rut C30, T. koningii D-64 and its mutants were grown in CG and CWB media, and FPase activities

2.0 1.0 RUT C30

MF5

MF13

MF6

MF3

PHVM1

SF18

SF32

E49

EH29

UH

U20-1

D-64

0.0

Mutants Genome shuffling and SM3

6.0 5.0 Mutagenes is and

4.0

Mutagenesis , intra-fusion and SM2

3.0 2.0 1.0 RUT C30

MF5

MF13

MF6

MF3

PHVM1

SF18

SF32

E49

0.0 EH29

T. koningii D-64 strain exhibited proper ratio of FPase and ␤glucosidase activities and this feature makes its enzyme system efficient in converting cellulose to glucose [8]. To improve its cellulase production further, here T. koningii D-64 was treated by random mutagenesis followed by genome shuffling. Two mutants, U20-1 and UH, were obtained after ultra violet irradiation and screening on SM1. The FPase and ␤-glucosidase activities for U20-1 were slightly higher than those for parental strain T. koningii D-64 (Fig. 1). On the other hand, the FPase activity for UH enzymes was less than that for D-64 enzymes (Fig. 1). However, the ␤-glucosidase activity for UH cellulase was enhanced compared to the parent strain. In order to develop a strain with combined improvement in both FPase and ␤-glucosidase, mutants UH and U20-1 were further treated by random mutagenesis and intra-strain protoplast fusion. Mutants EH 29 and E49 were respectively obtained from strain UH and U20-1 through EMS treatment and screening on SM2 plates. Meanwhile, mutants SF32 and SF18 were respectively obtained by intra-strain protoplast fusion of UH and U20-1 and screening on SM2 plates. Both FPase and ␤-glucosidase activities of the four mutants (EH29, E49, SF32 and SF18) were slightly enhanced (1.2–1.8 folds) compared to the parent strain D-64 (Fig. 1). To further develop the strain, genome shuffling method was applied. The protoplasts of the above four mutants were prepared, inactivated, mixed and fused by PEG. The hybrid mutants were screened on SM3 plates and potential mutants with bigger clear zones were further evaluated in shaking flasks containing CG fermentation medium followed by enzyme activity assay. Five potential mutants, MF3, MF5, MF6, MF13 and PHVM1, displayed higher FPase activity (2.0–2.3 folds) (Fig. 1). Furthermore, the ␤-glucosidase activities of these mutants were 2.3–3.7 folds compared to strain D-64. These results prove that the step-wise strain improvement process is effective in cellulase production enhancement.

Mutagenesis, intra-fusion and SM2

3.0

UH

3.1. Strain improvement

Mutagenesi s and SM1

4.0

U20-1

3. Results and discussion

5.0

D-64

Commercial enzymes of Novozym 188 from Aspergillus niger (Novozymes, Denmark) and lab-prepared cellulolytic enzymes complex in CWB fermentation medium from T. reesei Rut C30, T. koningii D-64 and its mutants were used for hydrolysis. The hydrolysis experiments were performed using 1.5% (w/v) biomass with specified enzyme loading in 5 mL 50 mM citrate buffer (pH 4.8) in 50-mL test tubes and incubated in water bath at 50 ◦ C and 100 rpm. The activity ratio of FPase and ␤-glucosidase in the enzyme mixture from T. reesei Rut C30 and Novozym 188 (C30N188) was 1:3. The hydrolysis of high-loading biomass (10–16%, w/v) was carried out in total volume of 10 mL in 25-mL conical flasks with 30 U FPase/g biomass. In all hydrolysis experiments, test tubes or conical flasks containing the same amount of buffer and enzyme without the biomass were used as the control. The hydrolysate was centrifuged at 5000 × g for 5 min. The supernatant was suitably diluted and used to determine the amount of reducing sugar released using the DNS method [20] and HPLC analysis.

6.0

FPase (U/ml)

2.9. Biomass hydrolysis

Genome shuffling and SM3

Beta-glucosidase (U/ml)

of reducing sugars released from enzymatic hydrolysis was measured using the 3, 5-dinitrosalicylic acid (DNS) method [8]. Sugar content in hydrolysate was analyzed by high performance liquid chromatography (HPLC) in an Agilent HPLC system with refractive index detector. HPLC organic acid analysis column (Aminex HPX87H column, 300 mm × 7.8 mm, Bio-Rad) was equilibrated at 55 ◦ C using the mobile phase of 5 mM H2 SO4 at a flow rate of 0.6 mL/min. Under the above analysis conditions, the retention time for glucose, xylose, cellobiose and arabinose were respectively 8.8, 9.4, 7.3 and 10.3 min.

675

Mutants

Fig. 1. Activities of FPase and ␤-glucosidase for T. koningii D-64 and its mutants. Fermentation was performed in 250-mL conical flasks containing 40 mL CG medium for 7 days.

were analyzed (Table 2). Generally speaking, mutants presented higher enzyme titer than their parent strain D-64 in both media and almost all the strains showed higher enzyme titer and specific enzyme activities in CBW medium. However, mutants only displayed enhanced specific activity in CG medium. Compared to CG medium, CWB medium contains wheat bran and soluble cellooligosaccharides which stimulate cellulase production [25]. It is therefore often used as a co-substrate for cellulolytic enzyme production [8]. In addition, consistently, all mutants displayed higher Table 2 FPase activities obtained for T. reesei Rut C30, T. koningii D-64 and its mutants cultivated in CG and CWB media. Medium

Mutants

FPase activity (U/mL)

Specific activity (U/mg)

Protein (mg/mL)

CG

Rut C30 D-64 MF3 MF5 MF6 MF13 PHVM1

4.8 2.1 4.4 4.2 4.5 4.3 4.9

± ± ± ± ± ± ±

0.3 0.1 0.1 0.1 0.2 0.3 0.1

1.12 0.72 1.33 1.20 1.25 1.16 1.23

± ± ± ± ± ± ±

0.07 0.03 0.03 0.03 0.06 0.08 0.02

4.3 2.9 3.3 3.5 3.6 3.7 4.3

± ± ± ± ± ± ±

0.1 0.1 0.5 0.2 0.1 0.1 1.3

CWB

Rut C30 D-64 MF3 MF5 MF6 MF13 PHVM1

5.0 3.6 4.4 4.7 4.8 4.2 5.0

± ± ± ± ± ± ±

0.2 0.1 0.3 0.1 0.3 0.2 0.1

1.19 1.33 1.62 1.31 1.37 1.35 1.32

± ± ± ± ± ± ±

0.05 0.04 0.11 0.03 0.09 0.06 0.03

4.2 2.7 2.7 3.6 3.5 3.1 3.8

± ± ± ± ± ± ±

0.1 0.1 0.2 0.2 0.1 0.1 0.1

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Fig. 4. Random amplified polymorphic DNA (RAPD) profiles of T. koningii D-64 and its mutants using primer mixtures of MAP-06 to MAP-10.

Detailed enzyme profile information can be obtained by secretome analysis [26]. 3.3. Random amplified polymorphic DNA (RAPD)

Fig. 2. SDS PAGE of the secreted proteins by parent strain T. koningii D-64, its mutants, MF13 and MF6, and T. reesei Rut C30 cultivated in CWB medium.

protein content than the parent strain T. koningii D-64. This suggests that the enhanced enzyme production might be due to the improvement in protein secretion in mutants. In addition, it may also be attributed to the enhanced protein expression levels resulted from mutagenesis, revealed by the enhanced specific activity of mutants in CG medium. Such enhancement was insignificant in CWB medium arisen from the complex induction effect by wheat bran. Protein gel of the secreted enzymes from T. koningii D-64, its mutants, MF13 and MF6, and T. reesei Rut C30 cultivated in CWB is shown in Fig. 2, where enhanced protein secretion in mutants is clearly demonstrated. In addition, the variations in protein profile between the D-64 mutants and T. reesei Rut C30 are obvious. Strain T. reesei Rut C30’s enzymes presented protein molecular weight of 60–100 kDa, which was consistent to the earlier report [26]. These include cellobiohydrolase I (Cel 7A), cellobiohydrolyase II (Cel 7B), xyloglucanase (Cel74A), ␤-glucosidase (BGLI), xylosidase (BXLI), and endoglucanase I (Cel7B), to name a few. On the other hand, D-64 enzymes were with molecular size of 50–75 kDa, suggesting that besides those above-mentioned Rut C30 enzymes, D-64 mutants presented significant amount of endoglucanase II (Cel 5A).

Grown on potato dextrose agar (PDA) plates, mutants displayed distinctively different morphology compared to their parent strain D-64 (Fig. 3). While D-64 was able to grow uniformly on PDA plates, mutants MF6 and MF13 grew in circles. In fact, MF6 and MF 13 grew faster and produced more spores than D-64 on PDA plates. These imply the mutagenesis of the genes in MF6 and MF13. Besides morphology variation, genetic variation can be detected by analysing the random amplified polymorphic DNA profiles [27]. In this study, a mixture of random primers MAP-06 to MAP-10 was used to get the RAPD profiles (Fig. 4). It was found that MF3 and MF13 displayed less bright DNA bands and notably different RAPD profiles compared to the parent strain and the rest mutants. One bright DNA band (at position of 0.4 kb) was detected for parent strain D-64 and MF5, however, not for other mutants. The RAPD profile of MF5 was very similar to that of parent strain D-64. On the other hand, PHVM1 exhibited similar RAPD with MF6. The above observation reveals the significant mutagenesis of the genes in these mutants. However, more detailed information on the genetic variation can be found through systematic comparative genome analysis. 3.4. Hydrolysis of pretreated oil palm empty fruit bunch (OPEFB) by mutant enzymes Lignocellulosic materials are abundant worldwide and they can be used for biofuel production [28–30]. Fermentable sugar production is very critical in lignocellulose bioconversion. Despite tremendous efforts and R&D in cellulose hydrolysing enzymes, enzymatic biomass saccharification is still challenging and costly [31]. It is therefore important to develop cellulase-producing

Fig. 3. Morphology of the parent strain T. kongingii D-64and its mutants MF13 and MF6 on potato dextrose agar (PDA) plates. (A) D-64; (B) MF13; (C) MF6.

Z. Wang et al. / Process Biochemistry 49 (2014) 673–680

5

MF6 C30N188

4

FPase (U/mL )

microorganisms so that cellulases can be produced on-site. While Trichoderma strains have been extensively studied for cellulase production and biomass hydrolysis [27,32,33], little was reported on the feature of T. koningii enzymes for lignocellulosic biomass hydrolysis. Southeast Asia accounts for more than 80% global palm oil production and every year more than 30 million tonnes of oil palm empty fruit bunch (OPEFB) are generated [34]. OPEFB has been transported to palm oil mills and it is therefore a potential lignocellulosic biomass that can be used for fuel and chemical production in this region. In this part of the study, the hydrolytic efficiency of cellulases from the potential T. koningii mutants was evaluated using the treated OPEFB (Table 3). Enzymes from CWB cultivated parental strain T. koningii D-64, T. reesei Rut C30 and T. reesei Rut C30 enzymes supplemented with commercial ␤-glucosidase (C30N188) were used as the references. It is noted that enzymes from mutant MF6 generated the highest amount of reducing sugar. Although Rut C30 enzymes resulted in almost the same amount of reducing sugar with MF6 enzymes, the cellobiose content was 3.3 mg/mL, much higher than that from MF6 enzymes. This is because that Rut C30 enzymes lack ␤-glucosidase [8]. However, after supplementing the commercial ␤-glucosidase (C30N188), the cellobiose content was greatly reduced to 0.1 mg/mL and the reducing sugar concentration was further increased to 9.3 mg/mL. The above results conclude that MF6 cellulase is most feasible for OPEFB hydrolysis without the need to supplement commercial ␤glucosidase. Thermo-stability analysis of MF6 enzymes and C30N188 were conducted in a similar way of OPEFB hydrolysis, without the addition of biomass. In the first 24 h, C30N188 enzyme activity was pretty stable, whereas that for MF6 enzymes slightly decreased (Fig. 5). After 24 h, C30N188 enzyme activity decreased faster and leveled off at about 60% of the initial value at 48 h. However, MF6 enzyme activity decreased at a relatively slower speed retaining about 70% of the initial enzyme activity at 48 h and afterwards. The stability of C30N188 in the first 24 h might correlate to its higher reducing sugar titer (Table 3). However, the OPEFB hydrolysis between MF6 and C30N188 enzymes are quite comparable (about 10 percent difference). T. reesei Rut C30 is a commercial cellulase-producing fungal strain. The analogous biomass hydrolysis performance and thermo-stability of MF6 and Rut C30 enzymes suggest that MF6 can be potentially used for commercial cellulase production.

677

3

2

1

0 0

20

40

60

80

100

Time (h) Fig. 5. Theremo-stability profiles of MF6 enzymes and C30N188.

3.5. Effects of enzyme loading on the hydrolysis of three types of lignocelluloses Among the few selected mutants, MF6 showed the best capability in OPEFB hydrolysis (Table 3). However, the hydrolysis efficiency was lower than that of C30N188 under the same conditions. Besides enzyme composition, cellulase loading is another important factor for efficient biomass hydrolysis. Further synergism of the enzyme components could be improved by the increase of enzyme dosage. In this part of the study, three types of lignocellulosic materials (OPEFB, HW and WC) were used for enzymatic hydrolysis with varied enzyme dosage, 20 U to 40 U FPase/g of biomass. Cellulases from the parent strain T. koningii D-64 and T. reesei Rut C30 supplemented with ␤-glucosidase (C30N188) at 20 U FPase/g biomass were used as the controls. As can be seen in Table 4, at enzyme loading of 20 U/g biomass, the enzyme mixture, C30N188, presented the highest sugar yield. MF6 enzymes performed better than D-64 enzymes. However, its hydrolysis efficiency was not as good as that from C30N188, consistent with data in Table 3. In order to further enhance its hydrolysis

Table 3 Hydrolysis of treated OPEFB by enzymes from potential mutants at 72 h. Sugars

D-64

MF5

MF3

MF6

MF13

PHVM1

Rut C30

C30N188a

Reducing sugar (mg/mL) Cellobiose (mg/mL)

6.6 ± 0.3 0.1

6.8 ± 0.3 0.2

7.3 ± 0.3 0.3

8.1 ± 0.3 0.1

6.4 ± 0.3 0.3

7.0 ± 0.4 0.2

8.0 ± 0.5 3.0 ± 0.4

9.3 ± 0.4 0.1

Note: Biomass loading was 1.5% (w/v) and FPase loading was 20 U/g biomass. a Mixture of T. reesei Rut C30 enzymes and ␤-glucosidase (Novozym 188) – enzyme activity ratio of FPase and ␤-glucosidase was kept at 1:3. Table 4 Hydrolysis of lignocellulosic materials by MF6 enzymes. Sugars

FPase loading (U/g biomass)

Fungal strains

Sugar concentration (mg/mL) 96 h OPEFB

Reducing sugar

20 20 20 30 40

Note: Biomass loading was 1.5% (w/v). a Same as denoted in Table 3.

C30N188a D64 MF6 MF6 MF6

12.2 8.6 10.5 11.5 12.0

± ± ± ± ±

HW 0.4 0.3 0.3 0.2 0.3

10.0 6.0 7.7 9.8 10.5

WC ± ± ± ± ±

0.4 0.3 0.3 0.3 0.3

6.9 4.9 6.4 7.1 7.5

± ± ± ± ±

0.3 0.1 0.2 0.2 0.2

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Z. Wang et al. / Process Biochemistry 49 (2014) 673–680

(a)

Glucose 100

10% (C30N188) 10% (MF6) 12% (MF6) 16% (MF6)

140 120

10% (C30N188) 10% (MF6) 12% (MF6) 16% (MF6)

80

100 60

Glucose (g/L)

Reducing sugar (g/L)

(b)

Sugar titer

80 60 40

40

20

20 0

0 -20 0

20

40

60

80

0

100

20

40

60

80

100

Time (h)

Time (h)

(d)

(c) Xylose

Cellobiose

40

10% (C30N188) 10% (MF6) 12% (MF6) 16% (MF6)

35 30

15.0 12.5

Cellobiose (g/L)

25 Xylose (g/L)

10% (C30N188) 10% (MF6) 12% (MF6) 16% (MF6)

20 15 10

10.0 7.5 5.0 2.5

5 0.0

0 0

20

40

60

80

100

Time (h)

0

20

40

60

80

100

Time (h)

Fig. 6. Hydrolysis profiles of high-loading OPEFB with 30 U/g FPase loading. Enzyme activity ratio of FPase and ␤-glucosidase was kept at 1:3 in C30N188.

efficiency, the enzyme dosage for MF6 enzymes was increased from 20 U/g biomass to 40 U/g biomass. Apparently, reducing sugar titer increased with the rise of the enzyme dosage (Table 4). At 40 U FPase/g biomass, almost identical sugar titer was obtained for MF6 enzymes and C30N118 (20 U/g biomass). Among the three types of biomass tested, OPEFB produced the highest sugar titer followed by HW. WC produced the least amount of sugar as it is the most recalcitrant biomass. The rigid structure and the high lignin content (Table 1) impede the performance of enzymes during hydrolysis by creating non-productive enzyme-lignin bonds [35,36]. Hydrolysis of treated oil palm empty fruit bunch (OPEFB), horticulture wastes (HW) and wood chips (WC) at 30 U/g biomass and 96 h resulted in cellulose to glucose conversion of 96.3 ± 2.2%, 98.2 ± 3.0% and 81.9 ± 1.4%, respectively. The corresponding conversions of xylan to xylose were 96.9 ± 1.5%, 95.0 ± 2.2% and 76.1 ± 3.1%. The above results demonstrate that increasing enzyme loading was effective in enhancing the biomass hydrolysis efficiency.

3.6. OPEFB hydrolysis at high biomass loading At 1.5% (w/v) OPEFB loading, sugar concentration was too low to be used for commercial purposes (Table 4). It is very essential to obtain high-concentration sugar for an economic feasible biomass conversion process. In this part of the study, hydrolysis of OPEFB was conducted at high biomass loading, 10% to 16% (w/v), with the enzyme dosage of 30 U FPase/g biomass for both MF6 enzymes and the enzyme mixture, C30N188. The hydrolysis profiles are shown in Fig. 6. As expected, titer of reducing sugar (Fig. 6a), glucose (Fig. 6b) and xylose (Fig. 6c) increased with the rise of biomass loading. At 16% (w/v) biomass loading, the overall sugar titer reached 125.5 ± 5.8 g/L at 72 h and 135.1 ± 2.5 g/L at 96 h. It is notable that increase of the incubation time is effective in improving the sugar yield. This probably associates with the relatively good thermostability of MF6 enzymes (Fig. 5). High solid loading hydrolysis of biomass has been a research focus for biomass conversion. Lu

Z. Wang et al. / Process Biochemistry 49 (2014) 673–680

679

Table 5 High-loading OPEFB hydrolysis at 72 h. Biomass loading (w/v)

Enzymes

Sugar titer (g/L)

1.5% 10% 12% 16% 10%

MF6 MF6 MF6 MF6 C30N188a

12.2 81.1 93.0 125.5 79.8

± ± ± ± ±

0.6 3.2 4.4 5.9 1.0

Sugar yield (mg/g biomass) 773.33 811.0 775.0 784.3 798.0

± ± ± ± ±

40.0 32.0 36.7 36.8 10.0

Cellulose conversion (%) 94.7 97.4 92.3 84.3 94.5

± ± ± ± ±

4.5 5.3 6.2 4.5 1.9

Hemicellulose conversion (%) 87.6 73.2 69.8 72.4 72.7

± ± ± ± ±

5.5 4.9 2.4 2.6 1.6

Note: FPase loading was 30 IU/g biomass. a Same as denoted in Table 3.

and his colleagues studied high solid loading hydrolysis of steam explosion pretreated corn stover using the commercial enzymes and a sugar titer of 103 g/L was obtained with the solid loading of 30% (w/v) [37]. More recently, Kim and his coworkers examined high solid OPEFB hydrolysis using the commercial enzymes and a sugar titer of 84.1 g/L was obtained [38]. For both above cases, biomass was chemically pretreated and xylan was almost completely removed. For our case, we retained glucose (89.2 g/L), xylose (29.4 g/L), cellobiose (14.4 g/L) and arabinose (2.1 g/L), with an overall sugar titer of 135.1 g/L at 96 h, which is so far the highest sugar titer reported for OPEFB. Interestingly, at 10% (w/v) biomass loading, MF6 enzymes performed equally well with C30N188 (Fig. 6a and b). Rut C30 enzymes require the supplement of ␤-glucosidase for efficient biomass hydrolysis; whereas MF6 enzymes can function equally well on its own. Again this concurs with our findings for T. koningii D-64 enzymes [8]. Sugar yield and biomass conversion at high biomass loading obtained at 72 h were listed in Table 5. It is notable that the sugar yield was consistently high and was maintained at around 770–811 mg/g biomass, disregarding the increase of biomass loading. Cellulose conversion was above 84% and that for xylan was above 72%. At 96 h, such sugar yield increased up to 844 ± 14.4 mg/g biomass. Such results are very significant as sugar yield normally suffers from the rise of biomass loading [8]. It is notable that at 10% (w/v) biomass loading, much comparable sugar yield were obtained for MF6 enzymes and C30N188. The high sugar yield at high biomass loading using MF6 enzymes implies that cellulases from MF6 are exceptionally feasible for OPEFB hydrolysis. It is also worthwhile mentioning that at high biomass loading cellobiose concentration reached 5–15 g/L (Fig. 6d). This is quite different from data in Table 3. This can be interpreted by the inhibitory effects from the high glucose content at high biomass loading. It was reported that fungal ␤-glucosidases from Aspergillus strain was more sensitive to glucose inhibition than that from T. reesei strains [39]. The discovery in this study suggests that ␤-glucosidases from T. koningii MF6 might be as sensitive as that from Aspergillus strains (Novozym 188) in terms of glucose inhibition (Fig. 6d). For industrial applications, glucose inhibition can be overcome by simultaneous saccharification and fermentation (SSF) processes. Alternatively, development of cellobiose-fermenting microorganisms can enhance cellobiose utilization and product yield.

4. Conclusion A potential mutant MF6 was developed from T. koningii D-64 and activities of FPase, CMCase, ␤-glucosidase and xylanase were respectively improved to 2.0, 1.3, 2.0 and 3.0 folds. MF6 enzymes performed equally well with the enzyme mixture of T. reesei Rut C30 cellulase and Novozym 188 (C30N188) at high biomass loading. Consistent sugar yield of 770–811 mg/g OPEFB was obtained at 10–16% (w/v) biomass loading. Conversion of cellulose to glucose was above 84% and that of xylan to xylose was above 72%. Reducing sugar titer reached 125.5 at 72 h and 135.1 at 96 h for 16%

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