Screening and Identification of Trichoderma Strains Isolated from Natural Habitats with Potential to Cellulose and Xylan Degrading Enzymes Production

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VOLUME 67 , ISSUE 2 (June 2018) > List of articles

Screening and Identification of Trichoderma Strains Isolated from Natural Habitats with Potential to Cellulose and Xylan Degrading Enzymes Production

ROMAN MARECIK * / LIDIA BŁASZCZYK / RÓŻA BIEGAŃSKA-MARECIK / AGNIESZKA PIOTROWSKA-CYPLIK

Keywords : microorganisms screening, Trichoderma species, lignocellulose biomass, cellulolytic activity, xylanolytic activity

Citation Information : Polish Journal of Microbiology. Volume 67, Issue 2, Pages 0-0, DOI: https://doi.org/10.21307/pjm-2018-021

License : (CC-BY-NC-ND-4.0)

Received Date : 22-May-2017 / Accepted: 02-February-2018 / Published Online: 30-June-2018

ARTICLE

ABSTRACT

A total of 123 Trichoderma strains were isolated from different habitats and tested for their ability to degrade cellulose and xylan by simple plate screening method. Among strains, more than 34 and 45% respectively, exhibited higher cellulolytic and xylanolytic activity, compared to the reference strain T. reesei QM 9414. For strains efficiently degrading cellulose, a highest enzyme activity was confirmed using filter paper test, and it resulted in a range from 1.01 to 7.15 FPU/ml. Based on morphological and molecular analysis, the isolates were identified as Trichoderma. The most frequently identified strains belonged to Trichoderma harzianum species. Among all strains, the most effective in degradation of cellulose and xylose was T. harzianum and T. virens, especially those isolated from forest wood, forest soil or garden and mushroom compost. The results of this work confirmed that numerous strains from the Trichoderma species have high cellulose and xylan degradation potential and could be useful for lignocellulose biomass conversion e.g. for biofuel production.

Graphical ABSTRACT

Introduction

Lignocellulose is among the most important components of plant biomass. It represents more than half of the globally produced organic matter during photosynthesis. In spite of its high abundance and energetic potential, this resource has not been fully utilized (Piotrowska-Cyplik, Czarnecki, 2003; Sanchez, 2009; Marecik et al., 2012). One of the reasons is a complex structure of plants biomass components, which mainly comprises polymeric compounds, such as cellulose, hemicelluloses, lignin and pectin (Sun and Cheng, 2002; Taherzadeh and Karimi, 2008; Hendriks and Zeeman, 2009; Sarkar et al., 2012). Regrettably, the presence of compounds with such a high degree of polymerization restricts their use as a carbon or energy source for animals and typical fermentation microorganisms. Finding a cheap, and safe for environment method of lignocellulosic biomass degradation would allow increasing feed digestibility and improve effectiveness of livestock production or simple conversion of plant biomass to biofuels (Harris and Ramalingam, 2010; Marecik et al., 2015; Chakdar et al., 2016).

Efficient use of the lignocellulosic resource as a source of renewable energy requires the employment of processes, which lead to the release of monosaccharides. This allows for obtaining substrates, which are easily assimilated by microorganisms and bioconverted to liquid or gaseous fuels, such as ethanol, methanol, hydrogen, methane and others (Saxena et al., 2009). A wide variety of methods can be employed for the degradation of the lignocellulosic complex, including physical, chemical or biochemical treatment. Especially, combined physical and chemical methods allow for rapid and efficient depolymerization of lignocelluloses; however, considerable energy expenditure is required possessing a notable threat to the environment (Kumar et al., 2009; Park and Kim, 2012).

The development of biotechnological hydrolyzation methods for the lignocellulosic complex is considered to be promising. These methods utilize unique properties of microorganisms to degrade different organic and inorganic or even xenobiotic substances to the simpler or nontoxic ones (Cyplik et al., 2012; Pęziak et al., 2013; Lisiecki et al., 2014). The use of such methods is based on the introduction of specific microorganisms or commercially available enzymatic preparations to the lignocellulosic biomass, what causes release of smaller pentose or hexose components. Enzymatic preparations employed for the decomposition of cellulose or hemicellulose are acquired from the cultivation of selected microbial strains (Aehle, 2007). The complete degradation of cellulose requires cellobiose dehydrogenases (CDHs) enzymes complex containing: endo- and exoglucanases and β-glucosidases. Depending on the producers, CDHs are classified into two classes: class I for CDHs produced by basidiomycetes and class II for CDHs from ascomycetes. Cellobiose dehydrogenases are flavocytochromes and belong to oxidoreductase class of enzymes. The efficient degradation of crystalline cellulose or hemicellulose is strongly related to copper-dependent lytic polysaccharide monooxygenases (LPMOs) (Harreither et al., 2011; Tanx et al., 2015). The preparations used for hemicelluloses hydrolysis are verycomplex, since they usually consists of a mixture of eight enzymes, such as endo-1,4-β-D-xylanase, exo-1,4-β-D-xylanase, α-L-arabinofuranosidase, endo-1,4-β-D-mannase, β-mannosidase, acetyl xylan esterase, α-glucuronidase and α-galactosidase (Clarke, 1997; Jorgensen et al., 2003; Banerjee et al., 2010). However, many different species of microorganisms capable of cellulolytic and hemicellulolytic enzymes synthesis have been discovered, including bacteria and fungi. It is important to note that the efficiency of lignocellulose decomposition is still unsatisfactory (Sun and Cheng, 2002).

Among the microorganisms, which exhibit the ability to produce hemicellulolytic enzymes, the filamentous fungi belonging to the Trichoderma genus attract particular attention (Xu et al., 1998). Due to substrate induction, these fungi produce and secrete considerable amounts of enzymes, which belong to cellulases as well as hemicellulases, which is why they are capable of growth under unfavourable environmental conditions (Sandgren et al., 2005). This is a valuable adaptive trait, which allows them to utilize different carbon and energy sources and grow under different temperature regimes, regardless of the presence of light (Polizeli et al., 2005). Due to their various metabolic activity, fungi belonging to the Trichoderma genus have found numerous practical applications e.g. enzyme producers, used as a biofungicides (Vinale et al., 2006; Wojtkowiak-Gębarowska., 2006; Vinale et al., 2008; Harris and Ramalingam, 2010; Chakdar et al., 2016).

The purpose of this study was to examine the ability of Trichoderma fungi isolated from different habitats to production of cellulose and xylan degrading enzyme and determine the activity of those enzymes.

Experimental

Materials and Methods

Fungal collection. The one hundred and twentythree Trichoderma strains, belonging to eleven species or species complex: Trichoderma atroviride, Trichoderma citrinoviride, Trichoderma hamatum, Trichoderma harzianum, Trichoderma koningii, Trichoderma koningiopsis, Trichoderma longibrachiatum, Trichoderma pseudokoningii, Trichoderma viride, Trichoderma viridescens and Trichoderma virens, were investigated in this study. The one hundred and seven strains were previously identified by Błaszczyk et al. (2011, 2016) and Jeleń et al. (2014) and deposited in the collection of the Institute of Plant Genetics, Polish Academy of Science, Poznań, Poland. Ten Trichoderma isolates were collected from: wheat kernels (Lublin – AN158 isolate), pieces of decaying wood with white or brown rot (Czerwonak – AN109, AN110 isolates; Golęcin Park, Poznań – AN131 isolate; Strzeszyn Park, Poznań – AN177 isolate; Joniec, Warszawa – AN501 isolate) and mushroom compost used for Agaricus bisporus cultivation (Skierniewice – AN186, AN187, AN188 isolates; Poznań – AN204 isolate) in Poland and isolated as described by Błaszczyk et al. (2011). Other strains including T. pseudokoningii (AN219, ITEM 1416), T. koningiopsis (AN222, ITEM2688), T. harzianum (AN220, ITEM 1328) and T. virens (AN267 – ITEM 1357, AN268 – ITEM 1591, AN269 – ITEM 1594) were kindly supplied by dr. Antonio Logrieco, CNR, ISPA, Bari, Italy. Trichoderma reesei QM 9414, sourced from the Czech Collection of Microorganisms (CCM), Brno, Czech Republic was used as the reference strain.

Morphological and molecular analysis. Ten isolates of Trichoderma sourced from wheat grains, compost used for mushroom cultivation and pieces of decaying wood collected from the floor of forests and parks in eastern and central Poland were identified morphologically following the procedure described by Gams and Bisset (1998). Colony characteristics were examined from cultures grown on PDA and SNA after 3–7 days at a temperature of 25°C. Microscopic observations were performed from cultures grown on SNA. Molecular species identification was based on the sequencing of two different phylogenetic markers: a fragment of the ITS1-5.8S – ITS2 rRNA region and a fragment of the translation-elongation factor 1-alpha (tef1) gene. Mycelium for DNA extraction was obtained as described previously (Błaszczyk et al., 2011). Isolation of total DNA was performed using the CTAB method (Doohan et al., 1998). The ITS1 and ITS2 region of the rDNA gene cluster was amplified using primers ITS4 and ITS5 (White et al. 1990). A fragment of the 1.2-kb tef1 gene was amplified using primers Ef728M (Carbone and Kohn, 1999) and TEF1LLErev (Jaklitsch et al., 2005). PCR amplification, DNA sequencing and sequence analysis was carried out under the conditions described by Błaszczyk et al. (2011). The sequences were identified by BLASTn (http://blast.ncbi.nlm.nih.gov/) as well as TrichOKEY and TrichoBLAST (http://www.isth.info; Druzhinina et al., 2005; Kopchinskiy et al., 2005). The sequences were deposited in the NCBI GenBank (https://www.ncbi.nlm.nih.gov/genbank/) and listed in Table I.

Table I

List of isolates originating from the different habitats identified as the Trichoderma species and analyzed for their cellulolytic and xylanolytic activity.

10.21307_pjm-2018-021_tbl1.jpg

Cultivation of Trichoderma and induction of enzyme synthesis. For assessing the capability to celululolytic or hemicellululolytic enzyme production, the fungi were cultivated on medium consisting only of carboxymethylcellulose sodium salt (Akzo Nobel Chemicals) or xylan (10 g/l) as a sole source of carbon. Furthermore, the medium contained: NaNO3 – 3 g/l, K2 HPO4 – 1 g/l, MgSO4 · 7H2O – 0.5 g/l, KCl – 0.5 g/l, FeSO4·7H2O – 0.01 g/l and pH was adjusted to 5.6±0.1. The inducing enzyme synthesis culture was carried out in 300 ml Erlenmeyer flasks, on a rotary shaker (150 rpm) for five days at a temperature of 25 ± 1°C. After the cultivation process the fungal cells were centrifuged (4500 rpm for 10 min) and obtained supernatants containing crude cellulolytic and xylanolytic enzymes were used for determination of the enzymes activity.

Analysis of cellulolytic and xylanolytic activity of Trichoderma fungi – plate method. The analysis of cellulolytic and hemicellulolytic enzymes activity was carried out using the plate screening method described by Hadkin and Anagnostakis (1977). The method is based on the observation of changes (determination of the size of clearance zones), which occur in the solid medium as a result of enzymatic activity. For determination of the cellulolytic activity, the medium including a 1% solution of carboxymethylcellulose sodium salt and 0.1 g/l of chloramphenicol in 2% solution of agar was used. The media were poured into Petri dishes (diameter of 90 mm) and then, after solidification, the central part was removed using a cork borer to create a well. To evaluate the xylanolytic activity the plates were prepared analogously, however a 1% solution of xylan was used instead of carboxymethylcellulose sodium salt.

The cultures of the Trichoderma fungi were centrifuged at 4500 rpm for 15 min, and then 200 μl of supernatants were placed in the wells. The plates were incubated at 37°C for 48 h and rinsed with 5 ml of a 1% Lugol’s iodine solution. After 15 minutes, the excess of the Lugol’s solution was rinsed with 0.1% solution of NaCl. The areas including non-hydrolyzed carboxymethylcellulose sodium salt or xylan were stained with a deep brown colour, whereas the areas in the direct vicinity of the well were characterized by a visible clearance, due to the enzymatic activity. The size of the clearance in each specific sample reflected the activity of cellulolytic or xylanolytic enzymes. The size of the clearance area, which occurred due to the activity of enzymes secreted by a given strain, was compared with the size of the clearance area obtained for the reference strain with known cellulolytic properties – T. reesei QM9414 (Sazci et al., 1986).

Analysis of cellulolytic activity of Trichoderma fungi – a blotting filter paper method. The overall cellulolytic activity (FPU) of selected fungal strains was also determined using the method recommended by Ghose (1987). Blotting filter paper stripes (Whatman No. 1) were placed in test tubes and incubated for 60 minutes at 50°C in the presence of 0.1 mol acetate buffer (pH 4.8) and the post-cultivation medium acquired after cultivation of fungi for 5 days. The amount of reducing sugars released into the supernatant was measured by employing the colorimetric method, using 3,4-dinitrosalicylic acid (DNS) (Miller, 1959). The cellulolytic activity of the post-cultivation medium was expressed as FPU (Filter Paper Unit) according to the International Union of Pure and Applied Chemistry (IUPAC) (Ghose, 1987). The amount of the enzyme, which allowed for the release of 1 μmol of glucose during 1 minute, was adapted as one unit of FPU cellulolytic activity.

Statistical analysis of the results. Each experiment of the enzyme activity analysis was carried out in three replicates. The Levene’s test (the homogeneity of variance test) and Turkey’s test were carried out in order to conduct a statistical verification of the obtained results. The calculations were carried out using Statistica 6.0 software.

Results and Discussion

Trichoderma species identification. Ten isolates of Trichoderma from samples of wheat grains, compost used for mushroom cultivation and decaying wood in Poland were identified at the species level based on morphological as well as ITS1, ITS2 and tef1 sequencing data. Finally, five species or species complex: were found to be: T. harzianum species complexes – 3 strains, T. virens – 4 strains, T. viride – 1 strain, T. viridescens – 1 strain and T. hamatum – 1 strain. The identification, origin and NCBI GenBank accession numbers of all Trichoderma isolates (both of ten isolates identified in this study and isolates previously recognized by Błaszczyk et al. (2011, 2016) and Jeleń et al. (2014) originating from the different habitats in Poland are given in Table I.

Cellulolytic activity of the studied fungal strains. The studies regarding the cellulolytic activity based on the plate method described by Hadkin and Anagnostakis (1977) revealed that among the 123 strains belonging to the Trichoderma genus more than 34% exhibited higher cellulolytic activity compared to the reference strain T. reesei and that these differences were statistically significant (p ≤ 0.05) (Fig. 1A). T. harzianum can be included as a species with high cellulolytic activity. Among the representatives of this species up to 21 out of 39 strains displayed a higher activity compared to the reference T. reesei QM9414 strain. The highest activity was observed for strains AN108, AN133, AN136, and AN360. The activity of these strains exceeded the activity of the reference strain by 2.4 times on the average. An activity exceeding 50% was noted for strains AN101, AN137, and AN367. All of these efficient T. harzianum strains were isolated from different locations of forest wood. Another species, which included very active strains with regard to degradation of cellulose, was T. virens, especially isolated from garden or mushroom compost. Among these species, 12 out of 15 strains were more active compared to the reference strain. The activity exceeding that of the reference strain by 2.6 times was observed for strains AN73, AN187, and AN268. Additionally, the degradation of cellulose was approximately twice as efficient for strains AN68, AN70, and AN188. Higher cellulolytic activity compared to the reference T. reesei strain was also observed in the case of three strains belonging to the T. viride and T. citrinoviride species as well as strain from the T. pseudokoningii (AN219). Among these species, a particularly high activity was exhibited by AN262 belonging to T. citrinoviride species and AN142 belonging to the T. viride species, both collected from forest wood. The cellulolytic activity of strains belonging to the remaining species, identified as T. viridescens, T. hamatum, T. koningii, T. koningopsis and T. atroviride were usually at a much lower level compared to the reference strain. High cellulolytic activity of the selected fungal strains belonging to the Trichoderma genus was also confirmed using the blotting filter paper method described by Ghose (1987). The selected strains characterized by the highest cellulolytic activity were presented in Table II.

Fig. 1.

Cellulolytic (part A) and xylanolytic activity (part B) of the studied fungal strains relative to the reference strain T. reesei QM9414 – the plate method analysis. The studied fungal strains belonged to the following species: A – T. viride, B – T. viridescens, C – T. virens, D – T. harzianum, E – T. hamatum, F – T. atroviride, G – T. longibrachiatum, H – T. citrinoviride, I – T. pseudokoningii, J –T. koningii.

* The value corresponding to difference in clearing zone diameter between analyzed strains

10.21307_pjm-2018-021_f001.jpg
Table II

Total cellulase activities of selected Trichoderma strains measured by the filter paper assay method (FPA).

10.21307_pjm-2018-021_tbl2.jpg

Xylanolytic activity of the studied fungal strains. The studies regarding the xylanolytic activity of the selected fungal strains belonging to the Trichoderma genus revealed that 56 out of 123 studied isolates were characterized by higher activity compared to the reference T. reesei strain (Fig. 1B). T. harzianum exhibited the highest activity. Up to 31 strains of these species displayed higher activity compared to the reference strain. Among these strains the highest activity was observed for strain AN94 obtained from forest soil, which was capable of degrading xylan over 3.5 times more efficiently compared to the reference strain. A notable xylanolytic activity was also observed in the case of strains AN101 and AN205. These strains exhibited activity, which was over 2.5 times higher compared to the reference strain. T. citrinoviride was another species, which included strains with high xylanolytic activity. The strain AN262 that belonged to this species, was capable of degrading xylan over 2 times more efficiently compared to the reference strain. High xylanolytic activity was also noted for AN213, belonging to T. longibrachiatum species, AN69 of T. virens species and AN277 of T. hamatum species.

For both activities analysed, no direct dependence between particular source of fungi strains and their degradative potential was observed; however, the strains isolated from forest wood, forest soil and compost were the most effective.

Filamentous fungi exhibit a broad spectrum of secondary metabolic activity representing important for the people – enzymes or antibiotics production, but also secretion of some dangerous, toxic or cancerogenic substances like mycotoxins (Jae-Hyuk and Keller, 2005; Błaszczyk et al., 2013; Błaszczyk et al., 2016). This is the effect of excellent adaptation ability to different extreme environment condition and the reason why these fungi are very interesting as a source of novel bioactive substances (Altinok, 2009; Chavez, 2015). The strains of Trichoderma used in this study were isolated from different habitats: decaying wood, forest soil, garden and mushroom compost, wheat and maize kernels. These habitats are a reach in carbon source but available only for microorganisms able to degrade lignocellulose compounds. This feature is widespread among the different fungi, including Trichoderma species (Druzhinina et al., 2010; Amore et al., 2013; Kubicek, 2013). Trichoderma genus is very common, diverse and occurs in a wide geographic distribution likewise north regions of Europe; however, the most of T. harzianum species identified in this study are uncommon, known mainly from Europe and North America (Jaklitsch et al., 2011; Chavez, 2015; Qin and Zhuang, 2016). The differences in the Trichoderma occurrence, related to habitats and geographic regions in Poland, were described also in previous studies (Błaszczyk et al., 2011, 2013, 2016)

Participation of cellulases and hemicellulases in global enzymes market has been increasing year to year. It is the effect of the expanding possibilities of their application in industrial practice (Beg et al., 2000). They may be used as a supplement in animal feeding as well as food or wood industry (Harris and Ramalingam, 2010). Developing biofuel industry (biogas, bioethanol) is also the area of cellulose and hemicellulose enzymes application to increase of the fermentation efficiency (Taherzadeh and Karimi, 2008; Ziemniński et al., 2012; Chakdar et al., 2016). These are the reasons that new and more effective sources of these enzymes are still studied. Many of the microorganisms are saprotrophs and contribute to the decay of organic matter exhibiting the possibility to cellulose and hemicellulose enzymes production (Crowther et al., 2012). However, despite that different microorganisms like bacteria, actinomycetes, yeast or even algae or insects are able to secrete these enzymes, filamentous fungi are especially worth of attention (Polizeli et al., 2005). The genus Aspergillus and Trichoderma secrete these enzymes directly into the environment at the remarkably higher than other microorganisms efficiency. The ability of different fungi strains belonging to the Trichoderma to produce cellulolytic and hemicellulolytic enzymes was extensively studied (Clarke, 1997; Xu et al., 1998; Sandgren et al., 2005; Banerjee et al., 2010). Such enzymes are obtained on industrial scale by aerobic cultivation of fungi, such as T. reesei and Humicola insolens or from recombinant strains (Liming and Xueliang, 2004; Wilson, 2009). The strains of filamentous fungi isolated from soil, decaying wood and sawdust were analyzed by Inuwa Ja’afaru (2013). Up to 42.6% of the 110 identified isolates belonged to the Trichoderma genus. The highest xylanolytic activity was exhibited by T. viride Fd18 strain, whereas the highest cellylolytic activity was observed for Trichoderma sp. F4 strain. The high potential of fungi belonging to the Trichoderma to produce cellulolytic and hemicellulolytic enzymes was confirmed in further studies (Wen et al., 2005; Chandel et al., 2013). Additionally, 23 out of 36 fungal isolates originating from compost also displayed cellulolytic activity. The isolates were identified as Trichoderma, Aspergillus, Rhizopus and Penicillium species (Chandel et al., 2013). The ability to synthesize cellulolytic enzymes by the modified T. reesei RUT-C30 strain QM 9414 with the use of cow manure as a substrate was confirmed by Wen et al. (2005). This strain was characterized by a higher production of cellulose compared to the reference T. reesei QM 9414 strain.

In summary, the results obtained in our study confirmed that numerous strains from the Trichoderma species are characterized by high lignocellulose degradation potential. The studies performed on forest soil, decaying wood or different kind of compost indicate a source of effective degraders of cellulose and hemicellulose. Due to potentially benefits related to the production of cellulolytic and hemicellulolytic enzymes and a relatively good growth rate, which is a characteristic trait of such microorganisms; these fungi may be helpful in the industrial practice. For this reason the screening of new producers and study of molecular mechanisms of metabolite secretion regulation should be continued.

Acknowledgements

This work was supported by strategic program of the National (Polish) Center for Research and Development (NCBiR), Advanced Technologies for Energy Generation. Task 4: Elaboration of Integrated Technologies for the Production of Fuels and Energy from Biomass, Agricultural Waste and other Waste Materials.

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  32. Kubicek C.P. 2013. Systems biological approaches towards understanding cellulase production by Trichoderma reesei. J. Biotechnol. 163: 133–142.
    [CROSSREF]
  33. Kumar P., D. M. Barrett, M.J. Delwiche and P. Stroeve. 2009. Methods for pretreatment of lignocellulosic biomass for efficient hydrolysis and biofuel production. Ind. Eng. Chem. Res. 48 (8): 3713–3729.
    [CROSSREF]
  34. Liming X. and S. Xueliang. 2004. High-yield cellulase production by Trichoderma reesei ZU-02 on corn cob residue. Bioresour. Technol. 91(3): 259–262.
    [CROSSREF]
  35. Lisiecki P., Ł. Chrzanowski, A. Szulc, Ł. Ławniczak, W. Białas, M. Dziadas, M. Owsianiak, J. Staniewski, P. Cyplik, R. Marecik and others. 2014. Biodegradation of diesel/biodiesel blends in saturated sand microcosms. Fuel. 116: 321–327.
    [CROSSREF]
  36. Marecik R., R. Dembczyński, W. Juzwa, Ł. Chrzanowski and P. Cyplik. 2015. Removal of nitrates from processing wastewater by cryoconcentration combined with biological denitrification. Desalin. Water Treat. 54(7): 1903–1911.
    [CROSSREF]
  37. Marecik R., J. Wojtera-Kwiczor, Ł. Ławniczak, P. Cyplik, A. Szulc, A. Piotrowska-Cyplik and Ł. Chrzanowski. 2012. Rhamnolipids increase the phytotoxicity of diesel oil towards four common plant species in a terrestrial environment. Water Air Soil Poll. 223(7): 4275–4282.
    [CROSSREF]
  38. Miller G.L. 1959. Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal. Chem. 31: 426–428.
    [CROSSREF]
  39. Park Y.C. and J.S. Kim. 2012. Comparison of various alkaline pretreatment methods of lignocellulosic biomass. Energy. 47(1): 31–35.
    [CROSSREF]
  40. Pęziak D., A. Piotrowska, R. Marecik, P. Lisiecki, M. Woźniak, A. Szulc, Ł. Ławniczak and Ł. Chrzanowski. 2013. Bioavailability of hydrocarbons to bacterial consortia during Triton X-100 mediated biodegradation in aqueous media. Acta Biochim. Pol. 60(4): 789–793.
    [PUBMED]
  41. Piotrowska-Cyplik A. and Z. Czarnecki. 2003. Phytoextraction of heavy metals by hemp during anaerobic sewage sludge management in the non-industrial sites. Pol. J. Environ. Stud. 12(6): 779–784.
  42. Polizeli M., A. Rizzatti, R. Monti, H. Terenzi, J.A. Jorge and D. Amorim. 2005. Xylanases from fungi: properties and industrial applications. Appl. Microbiol. Biotechnol. 67: 577–591.
    [CROSSREF]
  43. Sanchez C. 2009. Lignocellulosic residues: Biodegradation and bioconversion by fungi. Biotechnol. Adv. 27: 185–194.
    [CROSSREF]
  44. Sandgren M., J. Stáhlberg and C. Mitchinson. 2005. Structural and biochemical studies of GH family 12 cellulases: improved thermal stability, and ligand complexes, Prog. Biophys. Mol. Biol. 89: 246–291.
    [CROSSREF]
  45. Sarkar N., S. Kumar, S. Bannerjee and K. Aikat. 2012. Bioethanol production from agricultural wastes: An overview. Renew. Energ. 37: 19–27.
    [CROSSREF]
  46. Saxena R., D. Adhikari and H. Goyal. 2009. Biomass-based energy fuel through biochemical routes: A review. Renew. Sust. Energ. Rev. 13: 168–178.
    [CROSSREF]
  47. Sazci A., A. Radford and K. Erenler. 1986. Detection of cellulolytic fungi by using Congo-red as an indicator: a comparative study with the dinitrosalicilic acid reagent method. J. Appl. Bacteriol. 61: 559–562.
    [CROSSREF]
  48. Sun Y. and J. Cheng. 2002. Hydrolysis of lignocellulosic material for ethanol production: a review. Bioresour. Technol. 96: 673–686.
  49. Taherzadeh M. and K. Karimi. 2008. Pretreatment of lignocellulosic wastes to improve ethanol and biogas production: a review. Int. J. Mol. Sci. 9: 1621–1651.
    [CROSSREF]
  50. Than T.-C., D. Kracher, R. Gandini, Ch. Sygmund, R. Kittl, D. Haltrich, B. M. Hällberg, R. Ludwig and Ch. Divne. 2015. Structural basis for cellobiose dehydrogenase action during oxidative cellulose degradation. Nat. Commun. 6: 7542. doi: 10.1038/ncomms8542.
    [CROSSREF] [URL]
  51. Qin W.T. and W.Y. Zhuang. 2016. Seven wood-inhabiting new species of the genus Trichoderma (Fungi, Ascomycota) in Viride clade. Sci. Rep. 6, 27074. doi: 10.1038/srep27074.
    [CROSSREF] [URL]
  52. Vinale F., R. Marra, F. Scala, E. Ghisalberti, M. Lorito and K. Sivasithamparam. 2006. Major secondary metabolites produced by two commercial Trichoderma strains active against different phytopathogens. Lett. Appl. Microbiol. 43: 143–148.
    [CROSSREF]
  53. Vinale F., K. Sivasithamparam, E.L. Ghisalberti, R. Marra, S.L. Woo and M. Lorito. 2008. Trichoderma-plant-pathogen interactions. Soil Biol. Biochem. 40: 1–10.
    [CROSSREF]
  54. Wen Z., W. Liao and S. Chen. 2005. Production of cellulase by Trichoderma reesei from dairy manure. Bioresour. Technol. 96(4): 491–499.
    [CROSSREF]
  55. White T.J., T. Bruns, S. Lee and J.W. Taylor. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics, pp. 315–322. In: Innis M.A., D.H. Gelfand, J.J. Shinsky, T.J. White (eds). PCR protocols: a guide to methods and applications. Academic, San Diego.
  56. Wilson D.B. 2009. Cellulases and biofuels. Curr. Opin. Biotechnol. 20: 295–299.
    [CROSSREF]
  57. Wojtkowiak-Gębarowska E. 2006. Mechanizmy zwalczania fitopatogenów glebowych przez grzyby z rodzaju Trichodrema. Post. Mikrobiol. 45(4): 261–273.
  58. Xu J., N. Takakuwa, M. Nogawa, H. Okada and Y. Morikawa. 1998. A third xylanase from Trichoderma reesei PC-3-7. Appl. Microbiol. Biotechnol. 49: 718–724.
    [CROSSREF]
  59. Ziemniński K., I. Romanowska and M. Kowalska. 2012. Enyzmatic pretreatment of lignocellulosic wastes to improve biogas production. Waste Manag. 32: 1131–1137.
    [CROSSREF]
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FIGURES & TABLES

Fig. 1.

Cellulolytic (part A) and xylanolytic activity (part B) of the studied fungal strains relative to the reference strain T. reesei QM9414 – the plate method analysis. The studied fungal strains belonged to the following species: A – T. viride, B – T. viridescens, C – T. virens, D – T. harzianum, E – T. hamatum, F – T. atroviride, G – T. longibrachiatum, H – T. citrinoviride, I – T. pseudokoningii, J –T. koningii.

* The value corresponding to difference in clearing zone diameter between analyzed strains

Full Size   |   Slide (.pptx)

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  40. Pęziak D., A. Piotrowska, R. Marecik, P. Lisiecki, M. Woźniak, A. Szulc, Ł. Ławniczak and Ł. Chrzanowski. 2013. Bioavailability of hydrocarbons to bacterial consortia during Triton X-100 mediated biodegradation in aqueous media. Acta Biochim. Pol. 60(4): 789–793.
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  42. Polizeli M., A. Rizzatti, R. Monti, H. Terenzi, J.A. Jorge and D. Amorim. 2005. Xylanases from fungi: properties and industrial applications. Appl. Microbiol. Biotechnol. 67: 577–591.
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  45. Sarkar N., S. Kumar, S. Bannerjee and K. Aikat. 2012. Bioethanol production from agricultural wastes: An overview. Renew. Energ. 37: 19–27.
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  46. Saxena R., D. Adhikari and H. Goyal. 2009. Biomass-based energy fuel through biochemical routes: A review. Renew. Sust. Energ. Rev. 13: 168–178.
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  47. Sazci A., A. Radford and K. Erenler. 1986. Detection of cellulolytic fungi by using Congo-red as an indicator: a comparative study with the dinitrosalicilic acid reagent method. J. Appl. Bacteriol. 61: 559–562.
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  52. Vinale F., R. Marra, F. Scala, E. Ghisalberti, M. Lorito and K. Sivasithamparam. 2006. Major secondary metabolites produced by two commercial Trichoderma strains active against different phytopathogens. Lett. Appl. Microbiol. 43: 143–148.
    [CROSSREF]
  53. Vinale F., K. Sivasithamparam, E.L. Ghisalberti, R. Marra, S.L. Woo and M. Lorito. 2008. Trichoderma-plant-pathogen interactions. Soil Biol. Biochem. 40: 1–10.
    [CROSSREF]
  54. Wen Z., W. Liao and S. Chen. 2005. Production of cellulase by Trichoderma reesei from dairy manure. Bioresour. Technol. 96(4): 491–499.
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  57. Wojtkowiak-Gębarowska E. 2006. Mechanizmy zwalczania fitopatogenów glebowych przez grzyby z rodzaju Trichodrema. Post. Mikrobiol. 45(4): 261–273.
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    [CROSSREF]

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