Journal Search Engine

Download PDF Export Citation Korean Bibliography
ISSN : 1226-9999(Print)
ISSN : 2287-7851(Online)
Korean J. Environ. Biol. Vol.35 No.4 pp.648-653

Evaluation of Cellulolytic Enzyme Production by Indigenous Fungi in Korea

Hanbyul Lee, Young Min Lee, Young Mok Heo, Jaejung Lee1, Jae-Jin Kim*
Division of Environmental Science & Ecological Engineering, College of Life Science & Biotechnology, Korea University, Seoul 02841, Republic of Korea
1Division of Wood Chemistry and Microbiology, National Institute of Forest Science, Seoul 02455, Republic of Korea
Corresponding author : Jae-Jin Kim, 02-3290-3049, 02-2-3290-9753,
20171102 20171212 20171213


The aim of this study was to select various fungal strains indigenous to Korea that have the potential to produce cellulases, including filter paper activity (FPase), endo-β-1,4-glucanase (EG), and β-glucosidase (BGL). Among the 25 species of Ascomycetes and the 32 species of Basidiomycetes tested in this study, the Bjerkandera adusta KUC10565, Heterobasidion orientale KUC10556, Hyphoderma praetermissum KUC10609, and Trichoderma harzianum KUC1716 all exhibited remarkably high FPase activity. In addition, the T. harzianum KUC1716 showed high levels of EG and BGL activity. This strain has been selected for further study because of their enzymatic potential.


    Cellulose is the major constituent of lignocellulosic biomass and thus the most abundant and renewable resource for bioenergy production (Ja’afaru 2013). Chemically, it is simple repeating molecules composed of D-glucopyranose that are covalently linked together to form linear chain via β-1,4-glucan (Khianngamn et al. 2014). Cellulosic can be broken down either enzymatically or chemically into glucose which can be fermented to liquid fuel such as ethanol. Therefore, the utilization of the cellulosic biomass is an important and promising alternative energy production (Devendran et al. 2016).

    The basic enzymatic hydrolysis process of cellulose requires three types of enzymes. Endoglucanases (EGs) hydrolyzes the β-1,4 glucan chain of cellulose internally, thus generating new free ends in the polymer. Cellobiohydrolases (CBHs) release consecutive cellobiose from either the reducing or non-reducing end of oligosaccharide chains (Devendran et al. 2016). β-glucosidases (BGLs) hydrolyze soluble cellobiose molecules to glucose (Teugjas and Väljamäe 2013). These three components are contained in most of the fungal cellulases at different ratios and act synergistically for complete hydrolysis of cellulose (Singhania et al. 2013).

    Although the cellulolytic enzymes are synthesized by many microorganisms, higher enzyme quantities are produced by fungi (Wang et al. 2012). Coughlan (1985) reported a list of the cellulase-producing microorganisms, which established or potential commercial use. According to the report, fungi are good producers of cellulolytic enzymes compared to actinomycetes and bacteria. Although fungi have trouble in mass transfer compared to yeast or bacterial growth, they made technological success for enzyme production (Singhania et al. 2010). Trichoderma reesei is among the best protein secretors known which makes it has been widely used in bioprocessing for cellulase production (Berlin et al. 2007). Other fungal species such as Aspergillus, Penicillium, Mucor, Phanerochaete, Fomitopsis, and Humicola are also widely known to have prominent high cellulolytic activities (Saha 2004; Baldrian and Valaskova 2008; Imran et al. 2016).

    In the industrial-scale cellulosic ethanol process, the cost of cellulases represents significant operational cost (Ellilä et al. 2017). Recent molecular biology technique aims developing at microorganisms which can produce large amounts and more efficient cellulases. Therefore, obtaining high titer cellulase-producing microorganisms is positively necessary process. The present study aims at screening of indigenously isolated potent cellulase-producing filamentous fungi for further biotechnological applications.

    Cultures of filamentous fungi (n=64) were isolated from various sites and substrates in Korea from 2000 to 2013 (Table 1). The cultures were deposited in the Korea University Culture Collection (KUC, Seoul, Korea) and selected for screening. The phylogenetic analysis of the fungi was performed by using ribosomal internal transcribed spacer (ITS) region. The fungal nucleotide sequences were compared to those in the GenBank database using BLAST search. In addition, Bayesian analysis was performed with Markov chain Monte Carlo (MCMC) analysis by MrBayes 3.1.2. All 64 strains of fungi were grown and maintained on plates containing 2% malt extract agar (MEA; BD Difco, USA) medium at room temperature.

    Filamentous fungal isolates were screened for cellulases production in liquid cultivation using the small-scale method described by Lee et al. (2011). Two agar plugs with mycelium collected from MEA plates were inoculated in 10 mL of Mandels’ medium (Juhász et al. 2005) containing 1% (w/v) cellulose, 0.3 g L-1 urea, 1.4 g L-1 (NH4)2SO4, 2.0 g L-1 KH2PO4, 0.3 g L-1 CaCl2, 0.3 g L-1 MgSO4, 0.25 g L-1 yeast extract, 0.75 g L-1 peptone, 5 mg L-1 FeSO4·7H2O, 20 mg L-1 CoCl2, 1.6 mg L-1 MnSO4, and 1.4 mg L-1 ZnSO4.

    Cultures were incubated aerobically at 25°C with shaking at 150 rpm for 7 days. The cultures were prepared in triplicate. After incubation, the fermentation broth was harvested by centrifugation at 4000 rpm for 25 min at 4°C. The supernatants were collected by filtering through 0.45 μm membranes (Sartorius Stedium Biotech, Germany) and used as crude enzyme to measure the cellulase activities.

    Enzyme activity was determined by estimating the released reducing sugar using 3,5-dinitrosalicylic acid (DNS) (Miller 1959) according to the method of Xiao et al. (2004) with modifications. The reaction mixture composed of 40 μL of 50 mM sodium citrate buffer (pH 4.8), 20 μL of enzyme solution, and a 7-mm-diameter filter paper disk (Whatman No. 1) in a 200 μL PCR tube (BR781301, Sigma). The reaction mixture was incubated for 60 min at 50°C, then 120 μL of the DNS was added into each reaction. The reaction was boiled and then cooled for 5 min each. The color was developed and 36 μL of each sample in triplicate was transferred in a 96-well microtiter plate containing 160 μL of distilled water. The absorbance was measured at 540 nm. The EG activity was assayed using carboxymethyl cellulose (CMC) according to the miniaturized enzyme assay of Lee et al. (2011). The released reducing sugar was determined using the DNS method (Bailey 1988), and expressed as glucose equivalent. Each micro PCR tube contained 25 μL of 2% CMC (C5678, low viscosity, Sigma) in 50 mM sodium citrate buffer (pH 4.8) and 25 μL of enzyme solution. After incubation for 30 min at 50°C, 150 μL of DNS reagent was added. The reaction was boiled and then cooled for 5 min each. The color was developed and 33 μL of each sample in triplicate was transferred to individual wells in a 96-well microtiter plate containing 165 μL of distilled water. The absorbance was measured at 540 nm. BGL enzyme activity was carried out using p-nitrophenyl-β-D-glucopyranoside (pNPG, N7006, Sigma) as the substrate according to Lee et al. (2011). Briefly, 20 μL of the appropriate enzyme suspension was mixed with 20 μL of a 1 mM substrate solution in 100 mM sodium acetate buffer (pH 5.0). After incubating the assay mixture for 5 min at 50°C, the reaction was stopped by the addition of 20 μL of 2 M Na2CO3. The absorbance obtained at 405 nm with a microplate spectrophotometer (Bio-Tek, PowerWaveXS, VT, USA). One unit per mL of enzyme activity was defined as the amount of enzyme required to release 1 μmol from the substrates per milliliter of culture medium per minute.

    Cellulolytic activities of the 64 filamentous fungi belonging to 26 genera and 57 species were determined. The results of the cellulase activities are shown in Fig. 1. The strains investigated in this study had FPase activity ranging from 0 to 0.259 U mL-1. The highest FPase activity was determined from Trichoderma harzianum KUC1716. Bjerkandera adusta KUC10565, Heterobasidion orientale KUC10556, and Hyphoderma praetermissum KUC10609 showed similar FPase activities with T. harzianum KUC1716. Above the three basidiomycetes are known as white-rot fungi (Wu 1997; Maijala et al. 2003; Jung et al. 2014). Although white-rot fungi effectively degrade cellulose and hemicellulose (Schmidt 2006), they are more known as the best degraders of lignin in nature (Ten Have and Teunissen 2001). For this reason, many studies have focused on the degradation of lignin and environmental pollutants (Bumpus and Aust 1987; Aust 1995; Youn et al. 1995; Ohkuma et al. 2001). Meanwhile, T. harzianum KUC1716 also produced the highest EG activity (0.753 U mL-1) among the 64 strains, followed by Phanerochaete sp. KUC10530 (0.561 U mL-1) and T. gamsii KUC1747 (0.568 U mL-1). The ascomycetous Trichoderma is important fungi used to produce enzymes by fermentation process and has long been considered as the most productive cellulolytic fungi (Valencia and Chambergo 2013). One of the best known cellulolytic organisms is T. reesei. The present commercial cellulase preparations based on mutant strain of T. reesei have been produced on an industrial scale (Ma et al. 2011). However, BGL from T. reesei is produced in very small quantities, which leads to uncomplete biomass hydrolysis and limits its industrial application (Qian et al. 2016). This pattern is also presented in this study (Fig. 1). Despite Marasmiellus candidus KUC10547 was the highest BGL producer, followed by Penicillium oxalicum KUC5242 and T. harzianum KUC1716 in this study, the activities were generally low (0.020 to 0.298 U mL-1). However, T. harzianum has become a potent microorganism for the secretion of xylanases and cellulases (Theodore and Panda 1995; Seyis and Aksoz 2005; Maeda et al. 2011; Delabona et al. 2012), and the induction of xylanases and cellulases is likely to be under separate regulatory control (Senior et al. 1989). Moreover, the capacity of this fungus to secrete well-balanced cellulase complex, which breakdowns cellulosic compounds into glucose, has been reported in several studies (Kalra et al. 1984; Benoliel et al. 2013; Delabona et al. 2016).

    From the screening results for cellulase activities, T. harzianum KUC1716 revealed the highest FPase and EG activities in this study. Although overall activities of BGL were low, T. harzianum KUC1716 possessed high BGL activity among 64 strains. Based on this study, T. harzianum KUC1716 can be recommended as a promising producer of cellulases, and will be further studied for its enzymatic potentials.



    Phylogenetic tree and cellulolytic enzyme profiles of 64 fungal strains. The numbers above the branches indicate posterior probabilities. The scale bar indicates nucleotide substitutions per position.


    Fungal species isolated from various sites and substrates in Korea.


    1. AustS.D. (1995) Mechanisms of degradation by white rot fungi. , Environ. Health Perspect., Vol.103 ; pp.59-61
    2. BaileyM.J. (1988) A note on the use of dinitrosalicylic acid for determining the products of enzymatic-reactions. , Appl. Microbiol. Biotechnol., Vol.29 ; pp.494-496
    3. BaldrianP. ValaskovaV. (2008) Degradation of cellulose by basidiomycetous fungi. , FEMS Microbiol. Rev., Vol.32 ; pp.501-521
    4. BenolielB. TorresF.A. de MoraesL.M. (2013) A novel promising Trichoderma harzianum strain for the production of a cellulolytic complex using sugarcane bagasse in natura. , Springerplus, Vol.2 ; pp.656
    5. BerlinA. MaximenkoV. GilkesN. SaddlerJ. (2007) Optimization of enzyme complexes for lignocellulose hydrolysis. , Biotechnol. Bioeng., Vol.97 ; pp.287-296
    6. BumpusJ.A. AustS.D. (1987) Biodegradation of environmental-pollutants by the white rot fungus Phanerochaete chrysosporium: Involvement of the lignin degrading system. , BioEssays, Vol.6 ; pp.166-170
    7. CoughlanM.P. (1985) The properties of fungal and bacterial cellulases with comment on their production and application. , Biotechnol. Genet. Eng. Rev., Vol.3 ; pp.39-109
    8. DelabonaP.D. FarinasC.S. da SilvaM.R. AzzoniS.F. PradellaJ.G. (2012) Use of a new Trichoderma harzianum strain isolated from the Amazon rainforest with pretreated sugar cane bagasse for on-site cellulase production. , Bioresour. Technol., Vol.107 ; pp.517-521
    9. DelabonaP.D. LimaD.J. RoblD. RabeloS.C. FarinasC.S. PradellaJ.G. (2016) Enhanced cellulase production by Trichoderma harzianum by cultivation on glycerol followed by induction on cellulosic substrates. , J. Ind. Microbiol. Biotechnol., Vol.43 ; pp.617-626
    10. DevendranS. Abdel-HamidA.M. EvansA.F. IakiviakM. KwonI.H. MackieR.I. CannI. (2016) Multiple cellobiohydrolases and cellobiose phosphorylases cooperate in the ruminal bacterium Ruminococcus albus 8 to degrade cellooligosaccharides. , Sci. Rep., Vol.6 ; pp.35342
    11. ElliläS. FonsecaL. UchimaC. CotaJ. GoldmanG.H. SaloheimoM. SaconV. Siika-ahoM. (2017) Development of a low-cost cellulase production process using Trichoderma reesei for Brazilian biorefineries. , Biotechnol. Biofuels, Vol.10 ; pp.30
    12. ImranM. AnwarZ. IrshadM. AsadM.J. AshfaqH. (2016) Cellulase production from species of fungi and bacteria from agricultural wastes and its utilization in industry: a review. , Adv. Enzyme Res., Vol.4 ; pp.44
    13. Ja ?(tm)afaruM.I. (2013) Screening of fungi isolated from environmental samples for xylanase and cellulase production. , ISRN Microbiol., Vol.2013 ; pp.283423
    14. JuhaszT. SzengyelZ. ReczeyK. Siika-AhoM. ViikariL. (2005) Characterization of cellulases and hemicellulases produced by Trichoderma reesei on various carbon sources. , Process Biochem., Vol.40 ; pp.3519-3525
    15. JungP.E. FongJ.J. ParkM.S. OhS.Y. KimC. LimY.W. (2014) Sequence validation for the identification of the white-rot fungi Bjerkandera in public sequence databases. , J. Microbiol. Biotechnol., Vol.24 ; pp.1301-1307
    16. KalraM.K. SidhuM.S. SandhuD.K. SandhuR.S. (1984) Production and regulation of cellulases in Trichoderma harzianum. , Appl. Microbiol. Biotechnol., Vol.20 ; pp.427-429
    17. LeeY.M. LeeH. KimG.H. KimJ.J. (2011) Miniaturized enzyme production and development of micro-assays for cellulolytic and xylanolytic enzymes. , J. Microbiol. Methods, Vol.86 ; pp.124-127
    18. MaL. ZhangJ. ZouG. WangC.S. ZhouZ.H. (2011) Improvement of cellulase activity in Trichoderma reesei by heterologous expression of a beta-glucosidase gene from Penicillium decumbens. , Enzyme Microb. Technol., Vol.49 ; pp.366-371
    19. MaedaR.N. SerpaV.I. RochaV.A. MesquitaR.A. Santa AnnaL.M. de CastroA.M. DriemeierC.E. PereiraN. JrPolikarpovI. (2011) Enzymatic hydrolysis of pretreated sugar cane bagasse using Penicillium funiculosum and Trichoderma harzianum cellulases. , Process Biochem., Vol.46 ; pp.1196-1201
    20. MaijalaP. HarringtonT.C. RaudaskoskiM. (2003) A peroxidase gene family and gene trees in Heterobasidion and related genera. , Mycologia, Vol.95 ; pp.209-221
    21. MillerG.L. (1959) Use of dinitrosalicylic acid reagent for determination of reducing sugar. , Anal. Chem., Vol.31 ; pp.426-428
    22. OhkumaM. MaedaY. JohjimaT. KudoT. (2001) Lignin degradation and roles of white rot fungi: Study on an efficient symbiotic system in fungus-growing termites and its application to bioremediation., Riken Review, ; pp.39-42
    23. QianY.C. ZhongL.X. HouY.H. QuY.B. ZhongY.H. (2016) Characterization and strain improvement of a hypercellulytic variant, Trichoderma reesei SN1, by genetic engineering for optimized cellulase production in biomass conversion improvement. , Front. Microbiol., Vol.7 ; pp.1349
    24. RubeenaM. NeethuK. SajithS. SreedeviS. PrijiP. UnniK.N. JoshM.K. JishaV.V. PradeepS. BenjaminS. (2013) Lignocellulolytic activities of a novel strain of Trichoderma harzianum. , Adv. Biosci. Biotechnol., Vol.4 ; pp.214
    25. SahaB.C. (2004) Production, purification and properties of endoglucanase from a newly isolated strain of Mucor circinelloides. , Process Biochem., Vol.39 ; pp.1871-1876
    26. SchmidtO. (2006) Wood and tree fungi: biology, damage, protection, and use., Springer Science & Business Media, ; pp.96-102
    27. SeniorD.J. MayersP.R. SaddlerJ.N. (1989) Xylanase production by Trichodermaharzianum E58. , Appl. Microbiol. Biotechnol., Vol.32 ; pp.137-142
    28. SeyisI. AksozN. (2005) Effect of carbon and nitrogen sources on xylanase production by Trichoderma harzianum 1073 D3. , Int. Biodeterior. Biodegradation, Vol.55 ; pp.115-119
    29. SinghaniaR.R. PatelA.K. SukumaranR.K. LarrocheC. PandeyA. (2013) Role and significance of beta-glucosidases in the hydrolysis of cellulose for bioethanol production. , Bioresour. Technol., Vol.127 ; pp.500-507
    30. SinghaniaR.R. SukumaranR.K. PatelA.K. LarrocheC. PandeyA. (2010) Advancement and comparative profiles in the production technologies using solid-state and submerged fermentation for microbial cellulases. , Enzyme Microb. Technol., Vol.46 ; pp.541-549
    31. Ten HaveR. TeunissenP.J. (2001) Oxidative mechanisms involved in lignin degradation by white-rot fungi. , Chem. Rev., Vol.101 ; pp.3397-3413
    32. TeugjasH. ValjamaeP. (2013) Selecting beta-glucosidases to support cellulases in cellulose saccharification. , Biotechnol. Biofuels, Vol.6 ; pp.105
    33. TheodoreK. PandaT. (1995) Application of response surface methodology to evaluate the influence of temperature and initial pH on the production of beta-1,3-glucanase and carboxymethylcellulase from Trichoderma harzianum. , Enzyme Microb. Technol., Vol.17 ; pp.1043-1049
    34. ValenciaE.Y. ChambergoF.S. (2013) Mini-review: Brazilian fungi diversity for biomass degradation. , Fungal Genet. Biol., Vol.60 ; pp.9-18
    35. ViterboA. HaranS. FriesemD. RamotO. ChetI. (2001) Antifungal activity of a novel endochitinase gene (chit36) from Trichoderma harzianum Rifai TM. , FEMS Microbiol. Lett., Vol.200 ; pp.169-174
    36. WangM.Y. LiZ.H. FangX. WangL.S. QuY.B. (2012) Cellulolytic enzyme production and enzymatic hydrolysis for second-generation bioethanol production. Biotechnology in China Iii: Biofuels and Bioenergy., Springer Berlin Heidelberg, ; pp.1-24
    37. WuS.H. (1997) New species and new records of Hyphoderma (Basidiomycotina) from Taiwan. , Bot. Bull. Acad. Sin., Vol.38 ; pp.63-72
    38. XiaoZ.Z. StormsR. TsangA. (2004) Microplate-based filter paper assay to measure total cellulase activity. , Biotechnol. Bioeng., Vol.88 ; pp.832-837
    39. YounH.D. HahY.C. KangS.O. (1995) Role of laccase in lignin degradation by white-rot fungi. , FEMS Microbiol. Lett., Vol.132 ; pp.183-188

    Vol. 40 No. 4 (2022.12)

    Journal Abbreviation 'Korean J. Environ. Biol.'
    Frequency quarterly
    Doi Prefix 10.11626/KJEB.
    Year of Launching 1983
    Publisher Korean Society of Environmental Biology
    Indexed/Tracked/Covered By

    Contact info

    Any inquiries concerning Journal (all manuscripts, reviews, and notes) should be addressed to the managing editor of the Korean Society of Environmental Biology. Yongeun Kim,
    Korea University, Seoul 02841, Korea.
    E-mail: /
    Tel: +82-2-3290-3496 / +82-10-9516-1611