Isolation, Screening, and Identification of Potential Cellulolytic and Xylanolytic Producers for...

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Isolation, Screening, and Identificationof Potential Cellulolytic and XylanolyticProducers for Biodegradation ofUntreated Oil Palm Trunk and ItsApplication in Saccharification ofLemongrass LeavesS. K. Ang a , Adibah Yahya b , Suraini Abd Aziz c & Madihah Md Sallehb

a Faculty of Biosciences and Medical Engineering (FBME) , UniversitiTeknologi Malaysia , Skudai , Johor , Malaysiab Environmental and Bioengineering (EnvBio) Research Group,Sustainability Research Alliance , Universiti Teknologi Malaysia ,Skudai , Johor , Malaysiac Faculty of Biotechnology and Biomolecular Sciences , UniversitiPutra Malaysia , UPM Serdang , Selangor , MalaysiaAccepted author version posted online: 24 Jun 2014.Publishedonline: 05 Sep 2014.

To cite this article: S. K. Ang , Adibah Yahya , Suraini Abd Aziz & Madihah Md Salleh (2015) Isolation,Screening, and Identification of Potential Cellulolytic and Xylanolytic Producers for Biodegradation ofUntreated Oil Palm Trunk and Its Application in Saccharification of Lemongrass Leaves, PreparativeBiochemistry and Biotechnology, 45:3, 279-305, DOI: 10.1080/10826068.2014.923443

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Isolation, Screening, and Identification of PotentialCellulolytic and Xylanolytic Producers for

Biodegradation of Untreated Oil Palm Trunk and ItsApplication in Saccharification of Lemongrass Leaves

S. K. Ang

Faculty of Biosciences and Medical Engineering (FBME), Universiti Teknologi Malaysia,Skudai, Johor, Malaysia

Adibah Yahya

Environmental and Bioengineering (EnvBio) Research Group, Sustainability ResearchAlliance, Universiti Teknologi Malaysia, Skudai, Johor, Malaysia

Suraini Abd Aziz

Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia,UPM Serdang, Selangor, Malaysia

Madihah Md Salleh

Environmental and Bioengineering (EnvBio) Research Group, Sustainability ResearchAlliance, Universiti Teknologi Malaysia, Skudai, Johor, Malaysia

This study presents the isolation and screening of fungi with excellent ability to degrade untreated

oil palm trunk (OPT) in a solid-state fermentation system (SSF). Qualitative assay of cellulases and

xylanase indicates notable secretion of both enzymes by 12 fungal strains from a laboratory

collection and 5 strains isolated from a contaminated wooden board. High production of these enzymes

was subsequently quantified in OPT in SSF. Aspergillus fumigates SK1 isolated from cow dung gives

the highest xylanolytic activity (648.448 U g�1), generally high cellulolytic activities (CMCase: 48.006,

FPase: 6.860, beta-glucosidase: 16.328 U g�1) and moderate lignin peroxidase activity (4.820 U=g), and

highest xylanolytic activity. The xylanase encoding gene of Aspergillus fumigates SK1 was screened

using polymerase chain reaction by a pair of degenerate primers. Through multiple alignment of the

SK1 strain’s xylanase nucleotide sequences with other published xylanases, it was confirmed that the

gene belonged to the xylanase glycoside hydrolase family 11 (GH11) with a protein size of 24.49

kD. Saccharification of lemongrass leaves using crude cellulases and xylanase gives the maximum

reducing sugars production of 6.84 g=L with glucose as the major end product and traces of phenyl-

propanic compounds (vanillic acid, p-coumaric acid, and ferulic acid).

Address correspondence to Madihah Md Salleh, Environmental and Bioengineering (EnvBio) Research Group,

Sustainability Research Alliance, Universiti Teknologi Malaysia, 81310 Skudai, Johor, Malaysia. E-mail: madihah@

fbb.utm.my

Color versions of one or more of the figures in the article can be found online at www.tandfonline.com/lpbb.

Preparative Biochemistry & Biotechnology, 45:279–305, 2015

Copyright # Taylor & Francis Group, LLC

ISSN: 1082-6068 print/1532-2297 online

DOI: 10.1080/10826068.2014.923443

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Keywords cellulases, oil palm trunk (OPT), phenylpropanic, polyoses, solid-state fermentation,xylanase

INTRODUCTION

Millions of tonnes of solid lignocellulose biomass wastes are being discarded every year

from the agricultural, agro-industrial, and forestry industries without any proper treatment. As

a consequence, not only are the biomass’s potential benefits being underestimated, but its

disposal has also triggered unwanted problems such as pests, air pollution, and other environ-

mental concerns.[1] Rather than being disposed of, such biomass, in fact, is a reservoir of fermen-

ted sugars that can act as the raw material for biofuel production because of the heterogeneous

compositions that its macromolecules (e.g., cellulose and hemicellulose) possess. In this regard,

oil palm trunks are cheap and readily available, and thus have great potential in serving as the

feedstock for the production of high value-added products such as food additives, organic acid,

and enzymes.[2]

However, degradation of lignocellulosic biomass such as oil palm trunk is difficult due to its

strong recalcitrant branched phenolic lignin, which acts as the physical barrier in the outer

layer.[3] Moreover, the cellulose core found in the lignocellulose fibers is also resistant to enzy-

matic and biological attack because of its crystalline structure that comprises up to two-thirds of

the total cellulose content.[4] Though various chemical and physical pretreatment methods[5–7]

have been reported to enhance the susceptibility of oil palm trunk by reducing cellulose crystal-

linity, removing the hemicellulose layers, or partial disruption of lignin layers, these methods

have their own limitations. Generally, the chemicals and equipment used are expensive and

the harsh process may form microbial growth-respiration inhibitors that can potentially damage

the holocellulose polymers.[2] As such, pretreatment with biological synergistic actions between

xylanases and ligninase enzymes has been seen as the alternative to loosen the lignocellulosic

structures with less inhibitors being created and lower cost induced.[8]

The biological hydrolysis of lignocellulosic biomass requires synergistic reactions between

complex ligninolytic and cellulolytic and hemicellulolytic degrading enzymes. In general, degra-

dation of lignin requires the presence of ligninase secreted in an extracellular fashion. This has to

be done in an aerobic condition because the carbon–carbon and ether bonds joining the subunits

can only be cleaved through oxidation hydrolysis mechanisms.[9] Reports on the

lignin-degrading ability of various microorganisms including white rot fungi, brown rot fungi,

soft rot fungi, and filamentous bacteria have been widely published.[10,11] However, most of

these microorganisms are incapable of using lignin as the source of energy or carbon[12]; their

survival is more dependent on the hydrolysis of polysaccharides such as hemicellulose or cellu-

lose in the lignocellulose substrate. Therefore, the primary function of secreted ligninase is to

expose the structure so that these microorganisms can synthesize the nutrients through cellulases

and hemicellulases hydrolysis processes.

Cellulolytic enzymes are comprised of three basic types of enzyme responsible for specific

degrading action. The first type is the endoglucanase, which catalyzes the initial disruption of

internal bonds within cellulose crystalline structure to produce oligosaccharides. The second

is the exoglucanase, which attacks the nonreducing end of oligosaccharide chains to produce

tetrasaccharides or cellobiose (disaccharides). The final type is the b-glucosidase, which is

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responsible of completing the hydrolysis process by converting cellobiose fragments into

glucose.[13] Additionally, hemicellulolytic degradation depends on the xylanolytic endohydroly-

sis of 1,4-b-D-xylosidic linkages in xylan, which is undertaken by the xylanase enzyme, to

produce pentose sugars (xylose and arabinose) as well as hexane sugars (galactose and mannose)

that subsequently become the primary carbon source for cell metabolisms.[14,15]

In the year 2010, the demands for strong, thermostable, and highly specific industrial

enzymes in the global market were valued at $3.6 billion; this is estimated to grow at a com-

pounded annual growth rate (CAGR) of 9.1% to reach $6 billion by 2016.[16] Approximately

75% of these industrial enzymes in the global enzyme market are hydrolases, which include

cellulases, hemicellulases, and pectinase. However, one of the bottlenecks of this technique is

the high preparation cost of pure commercial enzymes, as well as relatively low conversion

efficiency, both of which make it less practicable for a vast industrial scale.[17] Thus, many inno-

vative efforts such as microbiological selection, genetic modification, protein engineering, and

enzymes immobilization have been made to innovate an alternative tuning tool that can enhance

the enzymes features and their production process.[18]

Solid-state fermentation (SSF) is a technique that involves the growth of microorganisms on

moist solid with minimum visibility of water. A point that has so far been overlooked is that

filamentous fungi are generally stronger than yeast and bacteria in terms of producing extracel-

lular cellulase and xylanase strains on complex solid substrates. Moreover, these fungi are cap-

able of enhancing the enzymes’ productivity and adapt easily to SSF since the microscopic and

macroscopic conditions are similar to their natural habitat.[19] SSF is also superior to another

popular fermenting technique, submerged liquid fermentation (SmF), in other ways. For

example, it gives greater volumetric productivity, lower risk of contamination, lower instrumen-

tal cost, higher product stability, and the possibility of producing valuable by-products such as

secondary metabolites.[20] Furthermore, the widely accepted usage of cheap lignocellulosic bio-

mass for value-added product synthesis such as fermentable sugars and ethanol has been proven

as technically and economically viable.[21] In light of this, the aim of this research is to meet

these demands, and with this, an alternative enzymes production method using cheap ingredient

with high enzymes production is introduced.

To date, extensive research studies have been done on saccharification of lignocellulosic bio-

mass through cellulases and xylanase degradation due to its various potential applications. In this

study, lemongrass leaves (Cymbopogon citratus) have been used as the substrate for cellulases

and xylanase mediated saccharification. This plant grows and is widely cultivated throughout the

tropical and subtropical countries, including Malaysia, where it could be easily found in most of

the local home gardens.[22] It has been estimated in Malaysia that 1180 ha of land was planted

with lemongrass and 7612 tonnes of lemongrass leaves were harvested during the year 2012

alone.[23] The medicinal potentials of lemongrass leaves that have so far been discovered include

their antidepressant, antioxidant, antiseptic, astringent, bactericidal, fungicidal, nervine, and

sedative properties.[24] However, the saccharification potential of this easily available lignocel-

lulosic biomass for sugar production has yet to be explored. Saccharification using an enzyme

cocktail that contain cellulases along with xylanases is, in fact, more advantageous than

with purified enzyme since it encompasses several enzymes that are essential for complete

hydrolysis of lignocellulose biomass into sugars.[25] Furthermore, the phenylpropanic com-

pounds (p-coumaric acid, ferulic acid, and vanillic acid) that are produced concomitantly in

the saccharification process could be used in vanillin production. For examples, p-coumaric acid

SCREENING AND IDENTIFICATION OF OPT-DEGRADING FUNGI 281

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could be used as the sole carbon source in bioconversion of ferulic acid into vanillin, where van-

illic acid can be used as starting material in chemical synthesis of vanillin.[26,27]

In this study, the major targets were to identify, screen, and select the isolates that were able

to produce hypercellulytic and xylanolytic enzymes as well as secrete the crucial ligninase for

improving the utilization of robust oil palm trunk. To the best of the authors’ knowledge, there

have been no published papers that have described the screening of cellulase, xylanase, and lig-

ninase producers that are capable of utilizing untreated oil palm trunk through the SSF system.

As such, this article is a pioneer in this regard. In addition, this study also examined the presence

of xylanase gene (GH11) found in the strongest xylanase producer among the isolated fungus,

which was Aspergillus fumigatus SK1. Furthermore, the crude cellulases and xylanase enzymes

produced by Aspergillus fumigatus SK1 through the SSF of oil palm trunk have also been

employed to conduct the saccharification of lemongrass leaves to produce sugar and

phenylpropanic compounds.

EXPERIMENTAL

Microorganisms Isolation and Growth

Five out of 17 fungal strains used in this study were isolated from a contaminated wooden board.

The samples that exhibited visible fungi mycelium were collected and mixed with sterile

distilled water prior to streaking onto potato dextrose agar (PDA). Further separation was done

until pure cultures were obtained. The spores of all pure fungal culture (coded MMS1 to MMS5)

were collected and maintained on Protect Bacterial Preserves (Technical Service Consultants,

Heywood, UK) at �80�C for long-term storage.

Inoculum Preparation

Twelve strains of fungi obtained from laboratory culture collection and five newly isolated

strains from wooden board were cultured on PDA plates for 7 days, incubated at 27�C. Then

1% (v=v) sterile Tween-80 solution was used to collect fungal spores, followed by centrifugation

at 4000 rpm for 20 min. The resulting pellet was mixed with sterile distilled water and further

dilutions were conducted to obtain a spore inoculum of 108 spores=g OPT.

Identification of Fungi Species

The extraction of DNA was initiated by physically disrupting the cell wall of frozen fungi

spores; this was done by grinding the fungi spores with cetyltrimethylammonium bromide

(CTAB) buffer[28] in sterilized mortar and pestle. Crude DNA extract was then collected and

further purified according to the modified Promega Wizard Genomic DNA purification kits

protocols. The samples were incubated at 37�C for 1 hr after being mixed with nucleus lysis

solution.[29] The purified DNA was used as the template for the polymerase chain reaction

(PCR) amplification of fungi 18S rDNA and internal transcribed spacer (ITS) sequences. The

18S rDNA primers used were: NS1 (50-GTA GTC ATA TGC TTG TCT C-30), NS8 (50-TCC

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GCA GGT TCA CCT ACG GA-30),[30] NU 0817 (50-TTA GCA TGG AAT AAT RRA ATA

GGA-30), and NU 1536 (50-ATTGCAATGCYCTATCCCCA-30).[30] The primers for ITS regions

amplification were: ITS1-F (500CTT GGT CAT TTA GAG GAA GTA A-30) and ITS4 (50-TCC

TCC GCT TAT TGA TAT GC-30).[31] The PCR amplification was performed in a total reaction

volume of 50 mL, which consisted of 25 mL Promega PCR Master Mix, 3 mL DNA template,

1 mL of each forward and reverse primer, 5 mL 10� bovine serum albumin (BSA), and 15 mL

of Promega nuclease-free water. Amplification was conducted using Bio-Rad MJ Mini 48-Well

Personal Thermal Cycler by subjecting the sample to a cycle of initial denaturation at 95�C for

5 min, 30 cycles of amplification, and finally a cycle of final extension at 72�C for another 5 min.

The condition of amplification cycles adopted were: NS1–NS8 (95�C for 30 s, 57�C for 30 s, and

72�C for 90 s), NU0817–NU1536 (95�C for 30 s, 58�C for 30 s, and 72�C for 46 s), and ITS1F–

ITS4 (95�C for 30 s, 58�C for 30 s, and 72�C for 30 s). The intensity and approximate sizes of

PCR products were examined using 1% (w=v) agarose gel electrophoresis with ethidium bro-

mide (EtBr) staining. Amplified DNA was purified and sequenced by 1st BASE (Malaysia)

Sdn Bhd. Sequenced data were analyzed using Heracle BioSoft Chromatogram Explorer, and

lastly compared with Genbank data. Closely related sequences were aligned using CLUSTALW

(MEGA 5.0) and phylogenetic studies were conducted using a neighbor-joining method with

1000 bootstrap replicates.

Qualitative Screening Medium

The specific solid medium used for screening the fungi’s abilities to secrete extracellular cellu-

lase and xylanase was a modified composition of Mendel basal medium.[32] The basic medium

contained 3.0 g=L yeast extract, 2.0 g=L KH2PO4, 3.5 g=L peptone, 1.0 g=L MgSO4 � 7H2O,

1.0 g=L (NH4)2SO4, and 20.0 g=L agar. The medium was supplemented with 2% (w=v) carbox-

ymethyl cellulose (CMC) and 1% (w=v) birchwood-xylan, respectively, as the sole carbon

source for cellulytic and xylanolytic qualitative screening. Ten microliters of spores suspended

from the 17 selected fungi strains were inoculated into a precast hole at the central of CMC agar

plates and birchwood–xylan agar plates. The hydrolyzed zones were examined every 24 hr by

flooding the CMC plates with 1% (w=v) Congo red solution followed by 1 M NaCl destaining

and the birchwood–xylan plates with 0.25% (w=v) aqueous I2 and KI.[33]

Substrate

The oil palm trunk (OPT) utilized as sole carbon substrate for SSF was obtained from a local

FELDA oil palm estate in Johor, Malaysia. The OPT was air-dried without chemical pretreatment.

Quantitative Screening

The solid-state fermentation was conducted using 15 g of OPT moistened with 45 mL of

production medium and 5 mL of spores suspension (108 spores=g OPT) to obtain a final moisture

level of 80%. The production medium was a modified Mendel basal medium with 1.4 g=L

(NH4)2SO4, 2.0 g=L KH2PO4, 0.3 g=L urea, 0.3 g=L CaCl2, 0.3 g=L MgSO4, 0.005 g=L FeSO4,


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0.0016 g=L MnSO4 �H2O, 0.0014 g=L ZnSO4 � 7H2O, 0.002 g=L CoCl2, 0.75 g=L peptone, and

2% (v=v) Tween 80.[32] The final moisture level was determined using a moisture analyzer

(MX50, A&D Weighing Co., Ltd., Japan). The cultures were incubated at room temperature

(27�C) and 1 g of sample was taken every 24 hr to analyze the cellulases, xylanase, and lignin

peroxidase activities. The cocktail of crude enzymes was extracted by mixing 1 g of fermented

OPT with 25 mL of 4�C 0.05 M sodium acetate buffer (pH 5) followed by 2 min of vortex.

The solids were separated by centrifugation at 4000 rpm for 30 min, and the clear supernatants

were used to test cellulases, xylanases, and lignin peroxidase activities.

Analysis Procedures

The activities of endoglucanase (carboxymethylcellulase activity, CMCase), exoglucanase

(filter paper activity, FPase), and xylanase were measured using the modified dinitrosalicylic

acid (DNS) method[34] based on the standard procedure recommended by the Commission on

Biotechnology, IUPAC.[35,36] The enzymes activities were assayed in a mixture containing crude

enzyme, reaction substrates (1% [v=v] CMC, birchwood–xylan, or filter paper strip), and 0.05 Msodium acetate buffer. The amount of reducing sugars produced was quantified by spectrophoto-

metrically at 540 nm after boiling the reaction mixture with 1 mL of DNS solution. One unit of

enzyme activity was defined as the release of 1mmol reducing sugar per milliliter enzyme per

minute. b-Glucosidase was measured based on the conversion rate of p-nitrophenyl-b-D-glucoside

(pNPG) into 1mmol of p-nitrophenol per milliliter enzyme per minute in 0.05 M sodium acetate

buffer.[37] Lignin peroxidase activity was estimated based on the ability of the enzyme to oxidize

methylene blue in the presence of hydrogen peroxide and 125 mM sodium tartrate buffer.[38] The

protein content in the supernatant was measured based on Lowry method.[39]

SDS-PAGE

The sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) analysis was con-

ducted using 12% (w=v) polyacrylamide gel as the separating gel; 20 mL of samples with 5 mL of

5� SDS-based sample loading buffer was heated at 100�C for 5 min. The samples and ladder

were loaded into wells. Electrophoresis was then conducted at 200 V for 50 min. The gel was

stained with 0.1% (w=v) Coomassie blue solution for 30 min, followed by destaining using

30% (v=v) methanol until clear visible bands were observed.

Analysis of Oil Palm Trunk Compositions

Extractives include gums, resins, pitch, waxes, sterols, flavinoids, tannins, terpenes, quinones,

nonstructural sugars, chlorophyll, and many other minor building-block reserves.[40] The

amounts of extractives in OPT were determined as the total weight loss after 1 g dried OPT

treated with 60 mL acetone had been incubated at 90�C for 2 hr.[41] The hemicellulose content

was determined as the difference between the sample weight before and after 1 g dried

extractive-free had been treated with 10 mL of 0.5 M sodium hydroxide at 80�C for 3.5 hr.[41]

To determine the amount of lignin, 1 g of dried extractive-free OPT was mixed with 98%

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(v=v) of sulfuric acid at room temperature and held for 24 hr before being boiled at 100�Cfor 1 hr. The weight of residual after being dried at 105�C was recorded as the lignin content.[41]

Assuming that the lignocellulose materials only had cellulose, hemicellulose, lignin, and other

extractives, the amount of cellulose content was determined by excluding all other components

of lignocellulos.[41]

Applications of Crude Cellulases and Xylanase in Saccharification ofLemongrass Leaves

The enzymatic degradation of lemongrass leaves (Cymbopogon citratus) for polyoses production

was done in batch culture. Initially, lemongrass leaves cut to 1 cm were boiled for 60 minutes

followed by sterilization at 121�C and 15 psi for 20 min. Crude cellulases and xylanases were

prepared by mixing 60 g of fermented OPT solids with 100 mL of cold sodium acetate (pH 5)

buffer. The sterilized lemongrasses 8% (w=v) were subsequently mixed with crude enzymes to

perform saccharification at 60�C and 200 rpm. Samples were drawn out every 24 hr and centri-

fuged for 30 min at 4000 rpm. The supernatants were collected and used for further analysis.

The production of phenylpropanic aromatic acids compounds (ferulic acid, p-coumaric acid,

and vanillic acid) due to lignin degradation was analyzed using the Agilent high-performance

liquid chromatograph (HPLC) 1200 equipped with a Zorbax SB-C18 column (5mm, 150�0.5 mm). The mobile phase was an isocratic mixture of 2% (v=v) acetic acid, 10% (v=v) acetoni-

trile, and 88% (v=v) water with a flow rate of 20mL=min. Detection was carried out using a diode

array detector (DAD) at wavelengths of 264 nm and 320 nm. The total reducing sugar concentra-

tions were detected using a standard DNS method.[34] The polyoses (arabinose, maltose, fructose,

xylose, galactose, cellobiose, and glucose) were analyzed using the Agilent HPLC 1100 equipped

with Rezex RPM-Monosaccharide Pbþ2 column and refractive index detector (RID). The mobile

phase was 100% pure water with the flow rate of 0.3 mL=min at 70�C.

RESULTS AND DISCUSSION

PCR Amplification and Sequencing of 18S rDNA and ITS Gene

The genomic DNA of 12 lignocellulose fungal degraders was successfully extracted. Amplifi-

cation of the ribosomal DNA (rDNA) and internal transcribed spacer (ITS) region was essential

to reveal the identified of fungal species used in this study. Two sets of primers (NS1–NS8 and

NU0817–NU1536) were used to specifically amplify the 18S rDNA region. All amplicons were

examined for their approximate nucleotide size using agarose gels (data not shown). The

addition of bovine serum albumin (BSA) into PCR reaction mixture significantly improves

the quality of amplicon obtained.

The 18S rDNA sequences of 12 fungal strains were successfully amplified to give near

full-length amplicons (�1700 bp) for 4 fungal strains using primers NS1–NS8; however, shorter

amplicons (675–753 bp) were obtained for 9 fungal strains amplified using NU0817–NU1536

primers (Table 1). The evolutionary relationship of all strains based on their 18S rDNA and

ITS sequences was elucidated via phylogenetic tress construction, respectively (Figure 1 and

Figure 2). Phylogenetic analyses based on both ITS and 18S rDNA sequences indicated that


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TABLE

1

SpeciesofFungiDeterm

inedbyAmplificationof18SrD

NAandInternalTranscribedSpacer(ITS)PrimerPairs

Cu

ltu

re

Most

sim

ilar

fun

gal

spec

ies

18

SrD

NA

pri

mer

sIT

Sp

rim

ers

NS

1-N

S8

NU

08

17

-NU

153

6IT

S1-I

TS

4

Am

pli

con

size

(bp

)

Mo

stsi

mil

ar

com

par

edsp

ecie

s

(acc

essi

on

num

ber

)

Sim

ilar

ity

(%)

Am

pli

con

size

(bp

)

Most

sim

ilar

com

par

edsp

ecie

s

(acc

essi

on

nu

mb

er)

Sim

ilar

ity

(%)

Am

pli

con

size

(bp

)

Mo

stsi

mil

ar

com

par

edsp

ecie

s

(acc

essi

on

num

ber

)

Sim

ilar

ity

(%)

SK

1Aspergillus

fumigatus

17

21

AB

00

84

01

.19

8—

——

58

0F

J21

43

71

.19

9

EF

B1

Aspergillus

niger

——

—7

44

JX1

12

70

3.1

99

57

6JQ

31

65

22

.19

9

EF

B2

.2Aspergillus

niger

——

—6

90

HQ

39

770

5.1

10

05

39

HQ

28

556

3.1

99

EF

B2

.3Aspergillus

flavus

17

31

GU

95

321

0.1

98

——

—5

49

JX2

322

69

.11

00

EF

B3

Fusarium

proliferatum

——

—4

80

HQ

87

188

0.1

99

51

5H

Q3

32

53

3.1

10

0

EF

B4

Hypocreavirens

——

—5

86

GQ

30

620

0.1

99

57

2H

Q6

08

07

9.1

10

0

AF

Aspergillus

nomius

——

—7

20

JF4

16

646

.19

95

21

FR

73

38

21

.11

00

MM

S1

Penicillium

griseofulvum

17

08

FJ4

58

44

6.1

98

75

3E

F6

08

15

1.1

98

49

7G

U5

65

09

9.1

10

0

MM

S2

Aspergillus

versicolor

——

—7

00

FJ9

41

881

.19

95

17

GU

23

276

7.1

10

0

MM

S3

Trichodermaviride

——

—7

13

FN

666

09

3.1

99

55

0H

M0

37

97

6.1

99

MM

S4

Aspergillus

awam

ori

——

—6

75

HQ

39

387

0.1

99

55

4A

Y3

73

84

0.1

98

MM

S5

Fusarium

oxysporum

17

07

AB

11

09

10

.19

8—

——

51

5E

U83

93

70

.11

00

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most strains were nested within the genus Aspergillus. Other strains were also closely clustered

together with cellulases and xylanase-producing fungi from the genus of Trichoderma (MMS3

and EFB4), Fusarium (EFB3 and MMS 5), and Penicillium (MMS1).

Strain SK1 was most closely related to Aspergillus fumigatus UPSC 177 and Aspergillusfumigatus ASNRR5S with sequence similarity of 98% and 99% based on the 18S rDNA

and ITS sequences, respectively. The 18S rDNA nucleotide sequence of strain SK1 has been

deposited in GenBank and assigned accession number JQ665711.1. Strain SK1 was highlighted

throughout this study due to its prominent ability to degrade untreated OPT using solid-state

fermentation through enzymatic mechanism.

FIGURE 1 Phylogenetic analysis of partial 18S rDNA sequences between 12 fungi clones by neighbor-joining (NJ)

method with bootstrap of 1000.


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FIGURE 2 Phylogenetic analysis of partial internal transcribed spacer (ITS) region between 12 fungi clones by

neighbor-joining (NJ) method with bootstrap of 1000.

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Qualitative Screening for Lignocellulose Degraders

Fifteen strains of fungi obtained from laboratory collections and five strains isolated from

a wooden board were initially compared and observed for their level of cellulolytic and xylano-

lytic enzymes productions under solid-agar screening. Selective solid agar was chosen based on

reliability and rapidness.[42]

The qualitative screenings for all fungi were conducted for 3 days in CMC and

birchwood-xylan plates. All species showed positive clearing zones (Figure 3 and Figure 4).

The diameter of clearing zones for all species on both types of selective agar plate varied from

22 mm to 68 mm (Table 2). All selected Aspergillus species produced small clearing zones ran-

ging from 22 mm to 42 mm for the CMC–agar plates and 21 mm to 33 mm for birchwood–xylan

agar plate after 48 hr. On the contrary, the Trichoderma species showed larger and faster

expanded clearing zones compared to Aspergillus, where five Trichoderma strains had produced

38-mm to 65-mm hydrolysis zones after 48 hr of incubation and this continued to increase until

more than 70% of the agar surface was covered after being incubated for another 24 hr. Interest-

ingly, the Fusarium species showed inconsistent cellulase and xylanase activities on agar plates.

Fusarium oxysporum MMS 5 had small cellulytic (17 mm to 44 mm) and xylanolytic (20 mm to

39 mm) clearing zones after 3 days, but those of Fusarium proliferatum EFB 3 were 65 mm and

58 mm, respectively, for CMC–agar and birchwood–xylan agar during the first 48 hr, and all sur-

faces were fully covered after 72 hr. Penicillium griseofulvum MMS 1, on the other hand,

showed the smallest cellulase and xylanase activities, where the clearing zones only covered

about 30% of the agar surface after 72 hr of incubation as compared to the control plates

(Figure 3 and Figure 4).

FIGURE 3 Qualitative screening for cellulytic activity on CMC–agar plates with Congo red staining. I, Trichoderma

reesei, II, Trichoderma virens, III, Aspergillus niger, IV, Aspergillus nomius AF, V, Aspergillus niger EFB 1, VI,

Fusarium proliferatum EFB 3, VII, Trichoderma koningiopsis EFB 4, VIII, Trichoderma virens EFB 2.1, IX, Aspergillus

niger EFB 2.2, X, Aspergillus flavus EFB 2.3, XI, Aspergillus fumigatus SK1, XII, Aspergillus niger MA, XIII,

Penicillium griseofulvum MMS 1, XIV, Aspergillus versicolor MMS 2, XV, Trichoderma viride MMS 3, XVI,

Aspergillus awamori MMS 4, XVII, Fusarium oxysporum MMS 5, XVIII, Control.


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TABLE 2

The Comparison of Clearing Zones Produced by Cellulases and Xylanase

Organisms

Clearing zones (mm)

Cellulases (CMC agar) Xylanase (birchwood xylan agar)

Day 1 Day 2 Day 3 Day 1 Day 2 Day 3

Trichoderma reesei 20 55 57 18 40 78

Trichoderma virens 30 45 67 17 56 79

Aspergillus niger 28 39 52 20 33 41

Aspergillus nomius AF 17 22 32 18 25 36

Aspergillus niger EFB 1 27 42 53 15 32 51

Fusarium proliferatum EFB 3 30 60 OG� 27 68 OG�

Trichoderma koningiopsis EFB 4 32 65 OG� 32 58 OG�

Trichoderma virens EFB 2.1 31 38 62 8 39 72

Aspergillus niger EFB 2.2 30 40 57 20 25 47

Aspergillus flavus EFB 2.3 25 35 46 20 26 49

Aspergillus fumigatus SK1 21 30 40 10 24 26

Aspergillus niger MA 26 35 55 22 29 39

Penicillium griseofulvum MMS 1 15 22 25 15 23 25

Aspergillus versicolor MMS 2 19 22 25 16 21 26

Trichoderma viride MMS 3 14 41 68 11 54 OG�

Aspergillus awamori MMS 4 15 25 45 19 29 40

Fusarium oxysporum MMS 5 17 30 44 20 31 39

Control (without inoculation) 0 0 0 0 0 0

�OG, Overgrow.

FIGURE 4 Qualitative screening for xylalolytic activity on birchwood–xylan agar plates with iodine staining. I,

Trichoderma reesei, II, Trichoderma virens, III, Aspergillus niger, IV, Aspergillus nomius AF, V, Aspergillus niger

EFB 1, VI, Fusarium proliferatum EFB 3, VII, Trichoderma koningiopsis EFB 4, VIII, Trichoderma virens EFB 2.1,

IX, Aspergillus niger EFB 2.2, X, Aspergillus flavus EFB 2.3, XI, Aspergillus fumigatus SK1, XII, Aspergillus niger

MA, XIII, Penicillium griseofulvum MMS 1, XIV, Aspergillus versicolor MMS 2, XV, Trichoderma viride MMS 3,

XVI, Aspergillus awamori MMS 4, XVII, Fusarium oxysporum MMS 5, XVIII, Control.

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Generally, the xylanolytic clearing zones on birchwood–xylan agar plates were clearer as well

as more distinct and prominent than the cellulolytic clearing zones found on CMC-agar plates.

This showed that xylanase activity was much higher compared to CMCase activities, and

this was supported by the quantitative results shown in Table 3 and Table 4. Furthermore, the

formation of clearing zones proved that all tested fungi species were able to secrete extracellular

cellulases and xylanase enzymes. Most lignocellulose wastes are too large to be transported

across the cell membrane. Due to the insolubility of cellulose, hemicellulose, and lignin, the

fermenting microorganisms, especially fungi, have to directly and extracellularly secrete

hydrolytic enzyme to break down these complex food sources.[3]

The clearing zones of CMC–agar and birchwood–xylan plates were expected to be produced

by endoglucanase and xylanase enzymes.[43] This screening method is commonly taken as the

first basis tool for identifying enzymes-producing and nonproducing strains, though it has

its own potential technical limitations as well. Most profoundly, the size of the clearing zones

is dependent on various factors such as kinetics of isolates, concentration of agar (which affects

the migration speed of enzyme), molecular size of the enzyme, and optimum growth temperature

of the strains.[42] Hence, as a rule of thumb, the results of this routine screening method need

further confirmation with quantitative enzyme measurement under SSF.

Quantitative Screening for Lignocellulose Enzymatic Hydrolysis Strains

Using agriculture wastes to produce value-added products has become the focus of concerned

scientists, the public, and governments nowadays. Scientists have been desperately searching

TABLE 3

Cellulases Production by 17 Strains of Fungi Under Solid-State Fermentation Using Oil Palm Trunk

as the Sole Carbon Source

CMCasea activity

(U=g)

FPasea activity

(U=g)

Beta-glucosidasea

activity (U=g)

Aspergillus niger MA 31.447 2.588 21.902

Trichoderma reesei 41.249 3.004 4.774

Aspergillus flavus EFB 2.3 41.014 2.429 11.571

Aspergillus fumigatus SK1 48.006 6.860 16.328

Aspergillus niger EFB 1 45.230 3.600 24.352

Trichoderma virens 45.200 3.294 4.055

Aspergillus niger 55.100 1.976 14.425

Aspergillus nomius AF 48.608 1.981 10.233

Trichoderma virens EFB 2.1 44.259 2.397 5.675

Aspergillus niger EFB 2.2 40.245 2.237 28.305

Fusarium proliferatum EFB 3 25.557 1.414 7.216

Hypocrea virens EFB 4 49.345 2.231 9.046

Penicillium griseofulvum MMS 1 23.823 0.596 12.012

Aspergillus versicolor MMS 2 4.765 0.277 7.665

Trichoderma viride MMS 3 45.498 2.299 28.473

Aspergillus awamori MMS 4 37.190 2.597 29.255

Fusarium oxysporum MMS 5 23.940 0.468 15.103

aMaximum activity.


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for alternative ways to produce enzymes, especially hydrolases, to meet the demand of the world

market for more stable, highly active, and distinctive enzymes that are able to grow rapidly.

Furthermore, the use of enzymes to catalyze several chemical processes has been identified as

being much cheaper, more efficient, and more environmentally friendlier than traditional chemical

methods. These have made enzyme production into a multimillion-dollar business in this decade.

Profile of Cellulase Production

In this study, all isolates were tested for their ability to synthesize extracellular cellulytic and

xylanolytic hydrolases enzymes by utilizing OPT as the sole carbon and energy source in

SSF (Table 3 and Table 4). Aspergillus niger, Aspergillus nomius AF, and Aspergillus fumigatusSK1 were found to be the highest CMCase (endoglucanase) producers, while Aspergillusfumigatus SK1, Aspergillus niger EFB1, and Trichoderma virens were the highest FPase

(exoglucanase) producers. These findings are consistent with some previous studies that reported

the genus Aspergillus as the most abundant paper-degrading mycoflora that can produce a high

amount of endoglucanase.[44]

At the same time, an interesting trend in cellulases production was found in this study.

The hyper FPase production strains could not synthesize a high amount of CMCase (Table 3).

This was caused by the catabolite repression effect of cellobioses, most significantly the exoglu-

canases (FPase) degradation reactions product, which inhibited the activity of endoglucanase.[45]

Yet the Aspergillus fumigatus SK1 with the strongest FPase production produced a high level

TABLE 4

Xylanase, Lignin Peroxidase, and Total Reducing Sugars Production by 17 Strains of Fungi Under Solid-State

Fermentation Using Oil Palm Trunk as the Sole Carbon Source

Xylanasea

activity (U=g)

Lignin peroxidasea

activity (U=g)

Total reducing

sugarsa (g=L)

Aspergillus niger MA 518.895 3.653 0.162

Trichoderma reesei 156.870 3.431 0.076

Aspergillus flavus EFB 2.3 567.180 4.708 0.154

Aspergillus fumigatus SK1 648.448 4.820 0.084

Aspergillus niger EFB 1 615.805 4.014 0.166

Trichoderma virens 60.850 3.416 0.023

Aspergillus niger 204.575 4.193 0.034

Aspergillus nomius AF 439.666 3.583 0.088

Trichoderma virens EFB 2.1 132.274 3.542 0.074

Aspergillus niger EFB 2.2 603.904 4.472 0.090

Fusarium proliferatum EFB 3 111.195 4.931 0.100

Hypocrea virens EFB 4 103.069 7.083 0.146

Penicillium griseofulvum MMS 1 93.713 3.139 0.077

Aspergillus versicolor MMS 2 74.678 4.375 0.084

Trichoderma viride MMS 3 45.508 5.097 0.089

Aspergillus awamori MMS 4 488.393 4.014 0.091

Fusarium oxysporum MMS 5 60.562 4.819 0.077

aMaximum activity.

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of CMCase. This was due to the beta-glucosidase found concomitantly in the enzyme cocktail

that had produced glucose from cellobiose and reduced the cellobiose inhibition effect.

Most Trichoderma species strains in this study produced less beta-glucosidase compared to

Aspergillus species (Table 3) because the Trichoderma species’ beta-glucosidase enzyme was

more sensitive to product (glucose) inhibition. Not only that, the level of Trichoderma species

enzyme productivity was only sufficient for growth on cellulose but inadequate for in vitro

saccharification of cellulose.[45] Moreover, the Trichoderma species were more susceptible to

thermal inactivation and periplasmic localization, and even a high concentration of cellobiohy-

drolyase enzyme would cause low beta-glucosidase production.[46] On the other hand, the over-

producers for beta-glucosidase, Aspergillus awamori MMS4 and Trichoderma viride MMS3,

which were originally isolated from a wooden board, could produce high levels of beta-

glucosidase. This was because their original isolated habitat was similar to the surface condition

of the hardwood,[45] and thus more efficient conversion of cellobiose to glucose could take place.

All studied strains produced a very low level of total reducing sugar. This showed that the

fungi had limited the release of cellulose hydrolysis products (sugars) to the environment to

support their high cell growth in SSF system. Furthermore, the low residual sugar level also

reduced the level of inhibitors (glucose and cellobioses), thus inducing cellulase production.[45]

Profile of Xylanase Production

Xylanases have received huge attention and gained its place in industries since they are able to

improve the digestibility of animal feed,[47] pulp bleaching,[48] and bioconversion of lignocellu-

lose waste into feedstock and fuels.[49] In this study, almost half of the 17 tested fungi were good

xylanase producers since more than 150 units of xylanase were produced for every gram of solid

substrate. Aspergillus fumigatus SK1, Aspergillus niger EFB1, and Aspergillus niger EFB 2.2

were classified as the hyperxylanase producers since more than 600 U=g of xylanase activity

was recorded. Previous research also showed that several species of Aspergillus have high

xylanolytic potential due to their high xylanases secretion.[50,51] High xylanase activity was

essential for complete hydrolysis of hemicellulose layers, which made up of 29.5% of the total

OPT lignocellulosic materials (see Table 6, shown later). Degradation of hemicellulose could

produce high-economic-value pentose (mainly xylose and arabinose) and hexose (glucose)

sugars essential for microbial biofuel production.[52] Besides that, Penicillium griseofulvumMMS1 and Aspergillus versicolor MMS 2, which are capable of producing high levels of

xylanases with less or no cellulases, are useful for biobleaching in the paper and pulp industries.

These cellulase-less xylanases attack links between lignin and hemicellulose (xylan) to break

down the cellulose pulp and facilitate the removal of lignin-associated hemicellulose fractions

without hydrolyzing the pulp cellulose and losing the pulp’s viscosity and strength.[53]

Profile of Lignin Peroxidase Production

Lignin is an aromatic polymer that is structurally complex, noncarbohydrate, and is built up of

phenylpropene units as the monomer.[9] These complex structures protect the inner parts of the

cell wall from biological and chemical attacks that decompose lignocellulosic materials.[54] Deligni-

fication is necessary to degrade the amorphous sugar polymers (cellulose and hemicelluloses)


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TABLE5

ComparisonBetweenDifferentFungiandSubstratesforCellulaseandXylanaseProductio

nUnderSolid-State

Ferm

entation

Mic

robes

Subst

rate

Chem

ical

Tre

atm

ent

CM

Cas

e

(U=g

)

FP

ase

(U=g

)

b-g

luco

sidas

e

(U=g

)

Xy

lanas

e

(U=g)

Ref

eren

ces

Aspergillus

niger

Whea

tb

ran

No

ne

48

.22

01

3.5

70

21

.69

02

60

4.0

60

[46

]

Aspergillus

fumigates

Whea

tb

ran

No

ne

20

.00

00

.670

99

.40

01

72

2.0

00

[64

]

Aspergillus

fumigates

Bag

asse

No

ne

29

.30

01

.430

48

.80

08

90

.00

0[6

4]

Pha

nerochaete

chrysosporiumþ

Aspergillus

sp.þ

Aspergillus

terreus

PO

MEþ

rice

stra

w0

.5%

(w=v

)

NaO

H

23

.05

41

1.8

15

11

.07

3N=D

[61

]

Aspergillus

sp.

Su

gar

can

eb

agas

seN

on

eN

D0

.400

ND

60

2.0

00

[62

]

Aspergillus

niger

Ric

eb

ranþ

hu

skN

on

e2

5.8

70

ND

ND

52

.82

0[6

3]

Aspergillus

niger

Oil

pal

mtr

unk

No

ne

55

.10

01

.976

14

.42

52

04

.57

5T

his

wo

rk

Aspergillus

niger

EF

B1

Oil

pal

mtr

unk

No

ne

45

.23

03

.600

24

.35

26

15

.80

5T

his

wo

rk

Aspergillus

fumigatus

SK

1O

ilp

alm

tru

nk

No

ne

48

.00

66

.860

16

.32

86

48

.44

8T

his

wo

rk

294

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beneath recalcitrant lignin layers. In this regard, bio-delignification is a more preferable method

than a chemical pretreatment process because the latter synthesizes various inhibitors that may

interfere with cellulase, xylanase, and glucosidase.[55] In this study, all selected fungi could

produce a significant level of lignin peroxidase with their activities marked between 3.13 U=g

and 7.08 U=g (Table 4). Low levels of nitrogen and sulfur available in the medium also allowed

the degradation of lignin as a secondary metabolite.[56] Although the lignin peroxidase activity in

this study was not high as for white rot basidiomycetes, it could be associated with high xylanase

activities that had induced good disruption of lignin-hemicellulose structure without releasing

inhibitors from lignin mineralization in chemical pretreatment.[57,58] In a nutshell, a complete

degradation of lignocellulose wastes not only depends on the high level of hydrolases secretion,

but also on a unique system that has good initial hemicellulose degradation as well as lignin

oxidation to induce other hydrolysis processes such as cellulase production.[3]

Comparative Studies

At present, only a handful of studies have described the direct utilization of untreated oil palm

trunk for cellulase and xylanase production. Studies using rice bran, wheat bran, sugarcane

bagasse, and palm oil mill effluent (POME) were comparatively more frequent. The maximum

endoglucanase enzyme activities (CMCase) by Aspergillus niger was recorded at 55.01 U=g

from the current study (Table 4), and this was higher than that of wheat bran (48.22 U=g) as well

as rice bran and husk mixture (25.87 U=g) (Table 5). Besides Aspergillus niger, Aspergillusfumigatus strain SK1 also showed 2.4-fold and 1.6-fold higher activities compared to

other CMCase production using similar species in wheat bran and bagasse. Furthermore, the

production of endoglucanase (FPase) by Aspergillus fumigatus SK1 was nearly 923% and

380% higher compared to wheat bran and bagasse SSF system (Table 5). In addition, previous

studies had also showed that lignin would affect cellulases production by limiting the access of

cellulases to substrat,[59] directly inhibiting the hydrolysis process,[60] or even causing unproduc-

tive absorption of cellulases into lignin.[4] However, in this case, since OPT had significantly

lower lignin content (9.32%) as compared to wheat bran and bagasse (Table 6), Aspergillusfumigatus SK1 was able to produce more cellulases. Previous reports also stated that mixed

culture of microorganisms could accelerate hydrolysis process and produce higher level of

hydrolysis enzymes.[61] However, Table 5 showed that production of the respective fungi, that

is, Aspergillus fumigatus SK1 and Aspergillus niger EFB1, yields higher CMCase and

beta-glucosidase activities in OPT SSF system as compared to the mixed pretreated substrate

TABLE 6

Chemical Compositions of Various Lignocellulosic Wastes

Cellulose Hemicellulose Lignin Extractive References

OPT 48.3% 29.6% 9.3% 12.8% This work

Wheat bran 29.0–35.0% 26.0–32.0% 16.0–21.0% N=A [52]

Rice straw 32.1% 24.0% 18.0% N=A [52]

POME 38.4% 23.2% 26.7% N=A [3]

Bagasse 43.0% 25.0% 24.0% N=A [38]


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system (0.5% [w=v] NaOH-treated POME and rice straw) with a complex fungi system of

Phanerochaete chrysosporium, Aspergillus sp., and Aspergillus terreus.[61] The current study

revealed that untreated OPT is a good sample substrate that produces a high level of enzymes

in easier, cheaper, and simpler ways.

The reason that all fungi produced a high level of xylanase as compared to cellulases could be

that the hemicellulose had shorter chains, noncrystalline arrangement, lower molecular weight,

and more amorphous and branched structure, which had eased its degradation.[2,3] However,

compared to previous reports, the production of xylanase by the strongest xylanase secretion

strain, Aspergillus fumigatus SK1, was only moderate, whereas it was much higher compared

to Moretti et al. (2012)[62] and Nguyen Anh Tuan and Van Hop (2009),[63] but slightly lower

than for the other studies[46,64] (Table 5). Utilization of substrate and synthesis of corresponding

enzymes, especially cellulases and xylanase, is highly dependent on crystallinity, degree of

polymerization, particle size, pore volume, and accessible surface area of the natural lignocel-

lulose material in an SSF system.[65] Therefore, the high levels of cellulases and xylanase

production in this initial screening revealed that this system has huge potential and is worth

for further optimization.

Sequence Analysis of Glycoside Hydrolyse GH11 Xylanases inAspergillus fumigatus SK1

Xylanase is responsible for degrading heteroxylans, which is the main constituent of hemicellulose.

Xylanases are highly diversified enzymes; many microorganisms can produce different types of

xylanases simultaneously.[66] Due to such high variety, the glycoside hydrolases (GH) classi-

fication system was developed to assort these xylanases into GH5, GH8, GH10, GH11, GH16,

GH26, GH30, GH43, and GH62 families.[66] The GH11 family is considered the true xylanases

due to their high selectivity toward the substrates, high catalytic efficiency, small protein size

(20 kD), intolerance of high substitutions on the xylan backbone, and adaptability to a variety of

pH and temperature.[66,67] Therefore, the strongest xylanase producer, Aspergillus fumigatusSK1, was selected to investigate the GH11 xylanase gene and its presence in crude extract.

The GH11 xylanases gene for Aspergillus fumigatus SK1 was amplified using the

degenerated primers set (AFORF11F and AFORF11R) reported by Jeya et al. (2009).[68] The

parameters of PCR were determined using PRIMER 3 (version 0.4.0) online application (http://

frodo.wi.mit.edu) (Figure 5). The PCR products met the predicted nucleotide length at approxi-

mately 750 bp. According to Jeya et al. (2009), 52 bp of single intron is the fragment of GH11

xylanase gene. In this case, the interrupted intron was removed from 295 bp to 346 bp following

the GT–AG rule.[68] The complete sequence of Aspergillus fumigatus SK1 GH11 was compared

with other xylanase genes in the NCBI GenBank database. The BLAST result indicated that the

Aspergillus fumigatus SK1 GH11 gene was 100% identical to the Aspergillus fumigatus strain

MKU1 endoxylanase (xynf11a) gene with a maximum 98% of coverage. The deduced nucleotide

DNA sequences were then translated into protein sequences using MEGA 5 ClustalW (version

1.6). The deduced amino acid sequences were then analyzed using the ExPASy Protparam

database (http://web.expasy.org/cgi-bin/protparam/protparam). The results indicated that the

Aspergillus fumigatus SK1 GH11 xylanase contained 228 amino acids (Table 7) with a predicted

molecular mass of 24.49 kD and theoretical pi of 6.27. The presence of GH11 xylanase

protein (24.49 kD) was further confirmed by SDS-PAGE analysis (Figure 6).

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Enzymatic Saccharification of Lemongrass Leaf

The crude enzyme cocktail system synthesized by Aspergillus fumigatus SK1 using untreated oil

palm trunk was tested for its efficiency on saccharification of lemongrass leaves, and Figure 7

shows its corresponding reducing sugars produced. The formation of reducing sugars during the

first 24 hr was very rapid, but it decreased after that. This could be due to a depletion in the easily

hydrolyzable fractions of cellulose and hemicellulose or enzyme inactivation.[69] The reducing

sugar production gradually increased until maximum reducing sugars production of 6.84 g=L

was reached at day 6. Further extension in incubation period did not improve the sugar

production because the soluble sugar formation per amount of absorbed enzyme had been

reduced.[70] In addition, the accumulation of excess end products (sugars) could have inactivated

the cellulases by blocking the active site of the enzymes.[71–73] Furthermore, the presence of lig-

nin contents in both tested substrates might have strengthened the resistance of enzymatic

hydrolysis process, especially in terms of cellulose degradation,[74] by providing a shelter to

hinder the attachment of enzymes with substrates.[75] The released lignin fragments could have

FIGURE 5 Predicted PCR conditions by Primer3. ATC, start condon, TAG, stop codon.


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FIGURE 6 SDS-PAGE analysis of Aspergillus fumigatus SK1 crude cellulases and xylanase enzymes. Line 1, Ladder,

line 2, sterile medium (control), line 3, crude cellulases and xylanase enzyme, line 4, concentrated crude cellulases and

xylanase enzyme by vivaspin filtration, line 5, concentrated crude cellulases and xylanase enzyme with vivaspin filtration

þTCA precipitation.

TABLE 7

Amino Acid Composition of Aspergillus fumigatus SK1 GH11 Xylanase

Amino acid Number count Percentage (%)

Ala (A) 17 7.5

Arg (R) 8 3.5

Asn (N) 20 8.8

Asp (D) 3 1.3

Cys (C) 1 0.4

Gln (Q) 3 1.3

Glu (E) 9 3.9

Gly (G) 34 14.9

His (H) 3 1.3

Ile (I) 7 3.1

Leu (L) 9 3.9

Lys (K) 3 1.3

Met (M) 3 1.3

Phe (F) 9 3.9

Pro (P) 8 3.5

Ser (S) 22 9.6

Thr (T) 27 11.8

Trp (W) 7 3.1

Tyr (Y) 20 8.8

Val (V) 15 6.6

Pyl (O) 0 0.0

Sec (U) 0 0.0

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competitively bound to the cellulases, too, and thus caused the enzyme to become inaccessible

to cellulose.[76]

The profile of polyoses (multiple fermented sugars) production was detected using

HPLC analysis (Figure 7). The high concentration of glucose (4.04 g=L) with low amount of

cellobioses (1.43 g=L) revealed the efficiency of cellulases in converting polysaccharides

into monomer with minimal intermediate sugars (cellobioses) that could appear as inhibitors

or unfermentable residual in ethanol fermentation.[77,78] Detection of hexose and pentose sugars

(xylose, arabinose, and maltose) released by lemongrass leaf can be justified with reference to

the high xylanase activity in the crude cocktail enzyme system. Generally, hexoses and pentoses

are the major fermented sugars required to produce ethanol.[77] Previous studies have shown

that Saccharomyces cerevisiae and Zymomonas mobilis could produce 2 moles of ethanol for

every mole of hexose through fermentation.[77] Hence, the crude cellulases and xylanase from

Aspergillus fumigatus SK1 are beneficial to the bioconversion of lemongrass leaf into interesting

biofuel and other industrial applications.

The crude cellulases and xylanase obtained from fermented solid were used to catalyse the

production of phenylpropanic compounds including vanillic acid, p-coumaric acid, and ferulic

acid from lemongrass leaf. These compounds are commonly found in the plant cells and the

lignin layer. They function as the signalling molecules, the phytoalexins compounds, and the

antimicrobial compounds.[79] Figure 8 reveals the detection of these compounds in lemongrass

hydrolysate. The maximum p-coumaric acid production (0.0886 g=L) was observed at day 7,

whereas the maximum production of phenolic acid (0.0417 g=L) was detected on day 8. The pres-

ence of these compounds in the hydrolysate liquid evidenced the mineralization of substrate,

which was triggered by the existence of these compounds released during the degradation of

FIGURE 7 Profile of total reducing sugars and polyoses production from saccharification of lemongrass leaves using

crude cellulases and xylanase produced by Aspergillus fumigatus SK1.


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lignin and other cell wall compounds.[26] Furthermore, the long retained period of these compound

in the medium had eased the recovery process.[80] However, the ferulic acid was different. Ferulic

acid was only observed on day 3 with a production of 0.0230 g=L. This might be due to the natural

characteristic of the compound such that it is sensitive to heat and susceptible to oxidation.[81]

The enzymatic synthesis of these compounds is more attractive than chemical syntheses.[81]

The possibility of producing vanillic acid, p-coumaric acid, and ferulic acid using crude

cellulases and xylanses as shown in the current research shows that enzymatic conversion can

be done by controlling a few variables only, such as the medium’s chemical composition, the

reduction in substrate, and the changes in product content during cells metabolisms or degra-

dation mechanisms.[81] Furthermore, the use of the inexpensive resource of lemongrass as the

raw material with single-step production is attractive for field applications. Compared to that,

the direct bioconversion of pure substrates such as ferulic acid to vanillic acid is affected by

many factors. According to some research, such production in batch fermentation is limited

because the cells may metabolize the vanillic acid to protocatechuic acid through the demethhly-

tion process.[80] Furthermore, the production of vanillic acid is reliant on the type of carbon

sources used. If nonaromatic compound (glucose) is used as the carbon sources, the cells cannot

convert ferrulic acid to vanillic acid.[26] In most cases, an expensive pure p-coumaric acid has to

be used as the inducer and carbon source.[26] The bioconversion, on another note, is dependent

on the ferulic acid concentration. High concentration of ferulic acid is toxic to the cells and tends

to limit the development of biomass.[26]

The production of polyoses as well as phenylpropanic compounds in this study was low

because the natural crude enzymes were facile and easily denatured entities in in-vitro milieu

when mixed with other contamination compounds; and it could only operate in a narrow

optimum range.[82,83] Many other factors also influenced the enzyme-substrate interaction, which

included the physiochemical hydrolysis conditions (e.g., temperature, pH, protein loadings, and

substrate concentration), substrate factors (e.g., lignocellulose structure and accessibility, as well

FIGURE 8 Profile of ferulic acid, p-coumaric acid, and vanillic acid production using crude cellulases and xylanase

produced by Aspergillus fumigatus SK1.

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as changes in substrate characteristic during hydrolysis), and enzymes-related factors (e.g.,

proportions of individual enzymes in the enzymes cocktail mixtures).[84] Moreover, the short

catalytic lifespan, high downstream processing cost, related environmental issues, difficulties

in the recovery of by-product, and limited reusability of these enzymes have further affect their

usefulness in enzyme-based industrial applications.[82,85] Thus, this calls for the immobilization

of enzymes to be practiced.

Immobilization is a preferred choice to improve the performance of cellulase and xylanase

in various applications, and allows the reuse of enzymes for multiple rounds of hydrolysis.

In addition, such use facilitates in situ product recovery, allows continuous mode of operation,

enhances the reaction rate, prevents enzymes aggregation, preserves enzyme properties under

drastic conditions, prevents subunit dissociation in multimeric enzymes, and improves the

enzyme behavior in term of its activity, specificity, and selectivity.[83,84,86] Damasio et al.

reported that the thermal stability of a recombined xylanase GH11 cloned from Aspergillusniveus had successfully improved 91.4% of the residual activity after 180 minutes of incubation

at 60�C.[87] Similarly, Sanchez et al. also successfully employed the immobilization of cellulase

and xylanase on different supports such as chitosan, alginate, carboxymethylcellulase (CMC),

and pectin.[88] Additionally, Gawande and Kamat demonstrated the noncovalent immobilization

of two types of xylanase produced by Aspergillus sp. for saccharification.[89] In that study, the

system had not only retained 70% and 80% of xylanase activities, respectively, but had recycled

the immobilized xylanase from both strains three times without losing the enzyme activity.[89]

Typically, multiple types of enzymes are required for complete hydrolysis, but the chemistry

and functionality of each enzyme is different and unique.[82] Therefore, different optimizing

immobilization methods have to be used for the respective enzyme variants,[82,90] and this

can be achieved by obtaining more detailed information on factors related to the individual

enzymes (enzyme size, isoelectrical point, comformation flexibility, surface functional group,

glycosylation, and presence of hydrophobic and=or hydrophilic region), carrier (surface functio-

nalization, mechanical stability, chemical stability, porosity, and particle size), and reaction

system (medium, inhibition, viscosity, thermodynamics, and diffusion limitations).[85] However,

these come with their own limitations such as high cost; mass transfer limitation; loss of enzyme

activity; loss of enzyme due to leakage and weak specific carrier-enzymes interaction;

operational resistances; additional material and equipment requirement; immobility of enzyme

inside the carrier; and particle erosion.[86] Thus, a more understanding and innovative immo-

bilization method has to be introduced to act as the preparative strategies in commercializing

the immobilized enzymes as an industrial catalyst in the future.

CONCLUSION

Twelve strains of fungi from laboratory collections together with five fungi strains isolated from

a wooden board were selected to investigate their ability to utilize untreated oil palm trunk for

cellulases and xylanase production. These fungi were in the genera of Aspergillus, Trichoderma,Fusarium, and Penicillium. A qualitative solid-agar screening method was used to select the

cellulolytic and xylanolytic secretion strains. All selected strains showed positive clearing zones

on CMC and birchwood–xylan plates. On quantitative screening, all strains produced different

levels of cellulases and xylanase under solid-state fermentation of untreated oil palm trunk.


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The Aspergillus fumigatus SK1 was found to be the best candidate for cellulases and xylanase

production among all other species. It produced the highest xylanase (648.448 U=g) activities

along with high CMCase (48.006 U=g), FPase (6.860 U=g), and beta-glucosidase (16.328 U=g),

which were found concomitantly in this enzyme cocktail system. After molecular and

SDS-PAGE characterization, the xylanase that belonged to xylanase glycoside hydrolase family

11 (GH11) with protein size of 24.49 kD was found among a variety of secreted enzymes.

To the best of our knowledge, this is the first report on the saccharification of lemongrass leaves

for the production of sugar and phenylpropanic compounds.

FUNDING

This work was financially supported by the Research University Grant (Q.J130000.2535.00H88)

of Universiti Teknologi Malaysia and MyBrain15 under the Ministry of Education, Malaysia

(MOE).

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Isolation, Screening, and Identification of Potential Cellulolytic and Xylanolytic Producers for...

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