Native macrofungi that produce lignin-modifying enzymes, cellulases, and xylanases with potential biotechnological applications

Gutiérrez-Soto, G., Medina-González, G. E., Treviño-Ramirez, J. E., and Hernández-Luna, C. E. (2015). "Native macrofungi that produce lignin-modifying enzymes, cellulases, and xylanases with potential biotechnological applications," BioRes. 10(4), 6676-6689.

Abstract

With the aim of identifying and exploiting the mycological resources available in the Mexican Sierra Madre Oriental, the lignocellulolytic and pectinolytic potential of autochthonous fungi were evaluated in the present work. A solid media selection system was established in which 74 isolated strains were tested and compared to six international reference strains. The macrofungi Xylaria sp CS121, Inonotus sp CU7, Basidiomycete CH32, Basidiomycete CH23, Xylaria poitei, and Trametes maxima CU1 showed the highest cellulolytic and pectinolytic potential. The greatest lignolytic capability was exhibited by T. maxima CU1 and Pycnoporus sanguineus CS43. Under stirred submerged culture, T. maxima CU1 (cellulases, cellobiose dehydrogenase, manganese peroxidase (MnP), and laccase, with 200, 359, 51, and 267 U/L, respectively) and Xylaria sp CS121 (198 U/L of xylanases) were the highest enzymatic producers. Under stationary conditions, the best producers were Inonotus sp CU7 for cellulases, P. sanguineus CS43 for cellobiose dehydrogenase and laccase, and T. maxima CU1 for xylanases and MnP (242, 467, 35, 165, and 31 U/L, respectively). These results demonstrate the efficiency of enzymatic profiling as a tool for enzyme discovery with Mexican native fungi.

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Native Macrofungi that Produce Lignin-Modifying Enzymes, Cellulases, and Xylanases with Potential Biotechnological Applications

Guadalupe Gutiérrez-Soto,^a Guadalupe E. Medina-González,^a José E. Treviño-Ramirez,^a and Carlos E. Hernández-Luna ^b*

Keywords: Cellulolytic potential; Enzyme screening; Lignin modifying enzymes; Mexican fungi; Pectinolytic potential

Contact information: a: Universidad Autónoma de Nuevo León, Facultad de Agronomía, Laboratorio de Fitopatología y Microbiología. Francisco Villa S/N Ex Hacienda El Canadá́, Gral. Escobedo, N. L. México. CP. 066054; b: Universidad Autónoma de Nuevo León, Facultad de Ciencias Biológicas, Departamento de Química, Laboratorio de Enzimología, Av. Pedro de Alba esq. Av. Manuel Barragán, Ciudad Universitaria, San Nicolás de los Garza, N. L. México, CP 66451;

* Corresponding author: carlosehlmx@yahoo.com

INTRODUCTION

Lignocellulose is the most plentiful component of photosynthetic plant biomass in the world. Representing almost 50% of total biomass, it is the most abundant organic renewable resource on earth. Plant residues increase slowly in natural ecosystems as trunks, timber, branches, and fallen leaves, while in agricultural ecosystems these accumulations occur over short periods of time and in considerable amounts. For this reason, lignocellulosic-rich residues are seen as a good source of valuable raw matter for paper and fuel production, composts, feed, fodder, and others (Sánchez 2009).

Because of the chemical bonds between plant cell wall components, the disposal of cellulose, hemicellulose, and lignin through artificial degradation is difficult and expensive in terms of the energy costs and pollutant production. Natural decomposition is slow, and in the case of lignin, only occurs with certain microbiota.

Therefore, research on efficient microorganisms able to degrade plant cell wall components is an investigative interest shared by many throughout the world. At first glance, most explorations have been focused on microorganisms such as bacteria and filamentous fungi, due to their ease of cultivation and enzyme concentration (Khokhar et al. 2013). However, a promising source of diverse and efficient enzymatic activities has been discovered in some other fungi, mainly basidiomycetes (Sánchez 2009; Elisashvili et al. 2011). Research on the hydrolytic and oxidative activities of plant cell wall components by this type of fungus involves the exploration of native flora and collection of strains. The basidiomycetes have been used as models to understand plant cell wall degradation processes, but just few of them have been applied in industrial practices.

Therefore, in the last decade, a great number of papers have been published in which native mycoflora are explored. Scientists have been searching for fungi of different physiologies in order to obtain their enzymatic profiles. The objectives of these activities are the research of basic useful information on biosystematics and biodiversity (Xavier-Santos et al. 2004; Atri and Sharma 2011). This adds to the understanding of saprophytic action by this group over fallen leaves and helps explain how chemical composition changes during degradation (Valášková and Baldrian, 2006), creating humic compounds (Steffen et al. 2007).

In the same way, microflora’s potential as enzyme producers has been explored in different areas, which include: degradation of lignocellulose for fertilizers production or bioremediation of dyes (Nazareth and Sampy 2003); bio-bleaching and fibers improvement in the paper industry (Sigoillot et al. 2002; Elisashvili et al. 2011); and on the development of integral biorefineries using agro-industrial wastes (Elisashvili et al. 2009).

In northeastern México, the exploration of the biotechnological potential of native fungi from mountain zones surrounding the city of Monterrey and its metropolitan area has led to a collection of 85 macrofungal strains that have been characterized for their capacity at decolorizing synthetic dyes that are commonly used in industry (Hernández-Luna et al. 2008). From this work, the laccases from Trametes maxima CU1 (Gutiérrez-Soto et al. 2011) and Pycnoporus sanguineus CS2 were purified, and their ability to degrade dyes with high demand in the region were characterized (Salcedo-Martínez et al. 2013).

In the present work, 74 native macrofungi newly isolated from different ecotypes (timber, plant litter, and dung) in northeastern México were explored. The aim was to find new enzyme sources capable of degrading cell wall components of plants, laying a foundation for their subsequent operative and functional characterization relative to the development of digestive aids to improve animal nutrition.

EXPERIMENTAL

Materials

A collection of 74 macrofungal strains from the Enzymology Laboratory at Facultad de Ciencias Biológicas, UANL and six reference strains were used in this work. Isolates were obtained from carpophores collected in oak forests and scrubland around Monterrey, N.L., México, from different ecotypes, mainly lignocellulosic substrates. The reference strains were kindly donated by M. Pickard from the University of Alberta, Canada. Their corresponding codes are: COL1 Berkjandera adusta UAMH 8258, COL2 Coprinus cinereus UAMH 4303, COL3 Coprinus sp UAMH 130509, COL4 Phanerochaete chrysosporium ATCC 24785, COL6 Trametes hirsuta UAMH 8165, and COL7 T. versicolor UAMH 8272. Native and collection strains consisting of vegetative mycelia were preserved at 4 ºC in YMGA medium (10 g·L^-1 malt extract, 4 g·L^-1 glucose, 4 g·L^-1 yeast extract, and 15 g·L^-1agar) as reported by Hernández-Luna et al. in 2008 and were transferred to new plates every three months for conservation and studies.

Methods

Plate screening for degrading enzymes of plant cell wall components

Identification of the main enzymatic activities involved in the modification of the components of plant cell walls was carried out using the medium previously reported by Sin et al. (2002). The base composition of the medium was: 0.1% peptone, 0.01% yeast extract, and 1.6% agar, which was supplemented with different carbon sources, dyes, or chromogenic substrates for revealing the presence of various enzyme activities through degradation of carbon source, discoloration of dye, or color generation. Corresponding supplements to these activities were: 2% carboxymethyl cellulose (CM-cellulose, Sigma) to detect cellulases, xylan (birch wood xylan, Sigma) for xylanases, 0.5% pectin (pectin from citrus peel, Sigma) for pectinases, 0.02% Poly R-478 dye for lignin modifying enzymes (LME), 0.015% Azure B dye for lignin peroxidase (LiP), and 0.02% syringaldazine for laccases. For dye decolorizing and laccase tests, 0.2% glucose was added as a carbon source as well. All media were sterilized by autoclaving at 121 ºC and 1.05 kg.cm^-2 for 15 min. Autoclaved media were poured in three division plates, allowed to solidify, and each division was inoculated with a 0.5 cm diameter cylinder of mycelium taken from the periphery of a 5 day growth colony of each strain growing in YMGA . All media were incubated at 28 ºC; colony growth as well as the existing areas of reaction or discoloration was registered daily as the diameter increased (mm) measured with a vernier caliper. Detection of cellulase, xylanase, and pectinase was conducted on the third day of growth (in most isolates), revealing their degradation by the addition of iodine solution. All assays were performed in triplicate, and a scale was established as support for the recording of results and the selection of the best enzyme producers.

Scale for the interpretation of results

In order to establish a scale for interpretation of the results for carbohydrolases, the ratio considering hydrolysis reaction and the growth diameters was used (Table 1). In this way, the hydrolysis area on the third day of the assay was revealed in the medium supplemented with CM-cellulose, xylan, or pectin.

Table 1. Scale for Carbohydrolases Results Interpretation

A value of one plus symbol (+) was given when the ratio was smaller than 1. Two plus symbols (++) corresponded to a ratio between 1 and 2. Finally, three plus symbols (+++) were assigned when a ratio greater than 2 was achieved (Fig. 1).

Fig. 1. Hydrolysis scale used for cellulases, xylanases, and pectinases detection. Enzyme activity was revealed at the third day of growth with iodine solution. Values given considered the difference between growth and hydrolysis areas.

In the case of Poly R-478, if decolorization was only partial from the seventh day or later, the assigned value was (+). If complete decolorization appeared in the area after the seventh day of growth, the corresponding value was (+ +). Only when decolorization area appeared on the fifth day of or before, a value of (+ + +) was assigned. For Azure B if complete decolorization become evident showing a light blue color on day 7 or before, the value of (+ + +) was given. When decolorization occurred after day 9, the value (+ +) was assigned. In most isolates only a zonal transformation appeared after day 5, showing a pink color; in those cases, a value of (+) was assigned. For laccase activity, just one plus symbol was assigned (+) to strains that showed a mild pink reaction appearing on the syringaldazine after the fifth day, a two plus symbols (+ +) for strains that showed a reaction at the fifth day, and a three plus symbols (+ + +) corresponded only to strains whose reaction was a deep pink before the fifth day of growth; all these are shown graphically in Fig. 2.

Fig. 2. Detection scale used for selection of lignin modifying enzymes producer strains. Poly R-478 decolorization was associated with lignolytic activity of isolates; Azure B decolorization with LiP production; and pink reaction of syringaldazine with laccase activity.

Identification of enzymatic profiles in liquid media

The determination of production curves for cellulases, xylanases, and ligninases was performed in a rich medium for basidiomycetes as reported by Pozdnyakova et al. (2006). Its composition per liter was: 10 g glucose; 0.5 g yeast extract; 10 g peptone; 0.72 g NH₄NO₃; 1.0 g MgSO₄; 0.5 g KCl; and 1 mL of trace solution (1 g/L FeSO₄; 2.8 g/L ZnSO₄; 3.3 g/L CaCl₂). Production was accomplished in 500 mL Erlenmeyer flasks containing 200 mL of medium and 2% wheat straw (m/v). After autoclaving (121 ºC, 1.05 kg·cm^-2) for 15 min and allowing to cool at room temperature, each flask was inoculated with three 0.5 mm diameter cylinders of mycelium taken from the periphery of a 5 day growth fungal colony in YMGA. Flasks were incubated at 28 ºC under stirring (150 rev·min^-1) and stationary conditions; every two days a 2 mL aliquot was taken for the assays of cellulases, xylanases, laccase, LiP, and manganese-dependent peroxidase (MnP). The determination of cellulase (CMCases, Avicelases, and -glucosidase) and xylanase activities was made following the method reported by Miller (1959). The reaction mixture consisted of 0.5 mL of sodium citrate (50 mM at pH 5.0), 0.3 mL of the respective substrate (2% CM-cellulose, Avicel, D-cellobiose, or D-Xylan), and 0.2 mL of sample aliquot. The mixture was incubated for 30 min at 50 °C. For the quantification of glucose or xylose, 0.1 mL of the reaction mixture and 0.1 mL of dinitrosalicylic acid (DNS) were added. After boiling for 5 min, the combination was placed in an ice bath and 1 mL of double distilled water was added. Photometric measurement was performed at 540 nm with a Shimadzu UV-Vis 1240 mini-spectrophotometer. A standard curve of glucose and xylose was used for cellulase and xylanase determinations, respectively. All assays were performed in duplicate. Laccase determinations were conducted using the method described by Abadulla et al. (2000), using 2,6-dimethoxyphenol (DMPO) as a substrate in sodium acetate buffer (200 mM at pH 4.5), reading at 468 nm in the spectrophotometer. For LiP, the method proposed by Tien and Kirk (1988) was used. The reaction mixture consisted of veratryl alcohol (3 mM) and hydrogen peroxide (2 mM) in sodium succinate buffer (50 mM at pH 4.5) and read at 310 nm. The assay for MnP was carried out following the protocol reported by Elisashvili et al. (2011), following the formation of malonate-Mn II complex at 270 nm. The reaction was started with 10 μL of sample, and all assays were made at room temperature per triplicate.

RESULTS AND DISCUSSION

Northeast México is a region with a great biodiversity that has not been well studied yet. Similarly, mycological resources present in this region have been underutilized as study models or novel sources of biomolecules and functional metabolites of biotechnological interest. The white-rot basidiomycetes (WRF) produce two extracellular enzyme systems responsible for degrading polysaccharide and lignin in the plant wall cell by the action of hydrolases and oxidoreductases (Baldrian and Valášková 2008; Sánchez 2009). These fungi obtain their carbon source from lignin (Pointing 2001; Mtui 2010), an evolutionary advantage that allows them to easily access metabolizable carbon sources, such as cellulose and hemicellulose, due to the presence of cellulases and xylanases. Enzyme production, as well as its operational and functional properties, varies depending on the strain and environment (Elisashvili et al. 2011). This has led to the continuous search for new WRF strains able to produce cell wall component degrading enzymes, isolated from different lignocellulose-rich biomass (Xavier-Santos et al. 2004; Valášková and Baldrian, 2006; Elisashvili et al. 2011; Ben Younes et al. 2011; Isikhuemhen et al. 2012). Previous reports of Sin and coworkers (2002), compare the diversity of lignocellulose degrading enzymes from basidiomycetes and filamentous fungi obtained from different substrates, finding a higher lignocellulosic potential in basidiomycetes.

Plate Screening for Degrading Enzymes of Plant Cell Wall Components

The lignocellulolytic, amylolytic, and pectinolytic potential of 74 native isolates obtained from different ecotypes of subtropical regions was determined. Table 2 shows the general results of the lignocellulolytic, pectinolytic, and amylolytic potentials of all strains. Ninety percent of the isolates were cellulase producers, while 96% were xylanase producers. Almost all of the isolates were able to act upon the cellulose and xylan, while 38.5% and the 18.7% showed the greatest hydrolytic activity, respectively. These results are very similar to those reported by Sin et al. (2002), who isolated fungi from different substrates rich in lignocellulose, which resulted in better degradation of cellulose rather than lignin. However, the activity shown by the isolates reported by Sin et al. (2002) was less compared to the positive control, while the best isolates in the present work showed greater activities than the international reference strains (Fig. 4).

In the analysis of the pectinolytic potential of the isolates, 68.7% of these acted on pectin, where 13.7% showed the greatest pectinase activity. Within the species with high pectinolytic potential, strains Pycnoporus sanguineus CS2 and P. sanguineus CS43 stood out. There are few reports about pectinases produced by basidiomycetes (Pericin et al. 1992; Pericin et al. 1997; Levin and Forchiassin 1998; Xavier-Santos et al. 2004), indicating that these results contribute to the validation of basidiomycetes as a potential source of pectinases with more robust operative and functional properties than those commercially available.

In the analysis of the lignolytic potential of the native strains, it was shown that 37.9% of the strains were able to produce LME, from which 9.5% completely discolored Poly R-478 dye on the fifth day of growth. In the test for the detection of LiP producing fungi, only the LA1 fungus was able to discolor Azure B on the fifth day of growth, showing a better result than P. chrysosporium ATCC 24785. Despite this, 40% of the isolates were able to transform Azure B from blue to pink. These same isolates showed a strong reaction over the syringaldazine, suggesting that laccase might be involved in the transformation of Azure B (Arantes and Milagres 2009). In a previous study (Sin et al. 2002), a pair of strains (Periconia sp. 1 y Piricaudia sp.) also showed this partial discoloration of Azure B. The possibility that this transformation of Azure B is associated with the production of high redox potential laccases, the production of MnP-laccase, or the presence of mediators of low molecular weight will be addressed in future investigations.

Table 2. Lignocellulolytic, Pectinolytic, and Amylolytic Potential of Native Macrofungi

The determination of cellulases, xylanases, pectinases, and amylases was conducted on the third day of culture, while the laccase was incubated until the seventh day and the LME for 30 days.

A total of 96.9% of the isolates had the capacity to hydrolyze starch, while 40.9% presented the significant activity over the substrate. Amylases are widely distributed enzymes in plants, animals, and microorganisms, but bacterial enzymes are in the greatest demand at the industrial level (Pandey et al. 2000; Gupta et al. 2003), even though there are reports that indicate that certain fungi are good amylase producers (Das and Sen-Mandi, 1992; Pal et al. 1980). There are studies where amylases of basidiomycetes have been exploited in order to increase the nutritional value of agroindustrial residues in the livestock industry (Han 2003; Han et al. 2005). Thus, knowing the amylolytic potential of the isolates makes it possible to obtain new producers of amylases that can be used for the development of digestive adjuvants.

From this system in solid media, strains Xylaria sp. CS121, Inonotus sp. CU7, Basidiomycete CH32, Basidiomycete CH23, and X. poitei were selected as the candidates with the highest cellulolytic and pectinolytic potential. Additionally, strains T. maxima CU1, P. sanguineus CS2, and P. sanguineusCS43 had lignolytic and amylolytic potential, while the Basidiomycetes CS 52, RS 9 and CH37 only displayed cellulolytic ability (Table 3). T. maxima CU1 showed higher activity than the international reference strains P. chrysosporium ATCC 24785, T. hirsuta UAMH 8156, or T. versicolor UAMH 8272 (Fig. 3).

Table 3. Native Fungal Strains with High Cellulolytic Potential

Fig. 3. Enzyme profiles in solid media. Row I corresponds to lignocellulolytic profile of T. maximaCU1, row II to that of P. chrysosporium ATCC 24785, row III to T. hirsuta UAMH 8156, and IV to T. versicolor UAMH 8272. The first three panels show the activities revealed with Gram iodine at the third day of growth: panel A corresponds to cellulases, B to xylanases, C to pectinases. Panels D, E and F corresponds to LME, LiP and laccase activities respectively. Discoloration of Poly R-478 (D) and Azure B (E) was observed in cultures along 30 days and laccase results were followed from the third to fifth day of growth.

Screening for Cellulase, Xylanase, and Lignolytic Production in Liquid Medium

From the scrutiny of 74 strains in solid media, strains P. sanguineus CS2, P. sanguineus CS43, Armilariella sp. CS134, Xylaria sp. CS121, T. maxima CU1, Inonotus sp. CU7, and Ps. candollenaCU20 were selected for the production of cellulases, xylanases, and laccases in submerged cultures with and without stirring, using a wheat straw based medium. Native strain Armilariella sp CS134 was used as positive control. In stirring conditions, all of the strains had the CMCases titers, with T. maxima CU1 (200 U/flak), Xylaria sp. CS121 (199 U/flask), and Armilariella sp CS134 (190 U/flask) being the greatest producers. The rest of the strains had a production of less than 100 U/flask. The avicelases showed greater amounts of production at the fourth day, where the majority of the isolates produced over 200 U/flask, except for the strains P. sanguineus CS2 (12 U/flask) and Ps. candollenaCU20 (45 U/flask). The greatest production of xylanases was presented at the fourth day, with Xylariasp. CS121 (198 U/flask), and Inonotus sp. CU7 (123 U/flask) being the greatest producers; the rest of the strains produced less than 100 U/flask. -D-glucosidase showed the greatest production on the fourth day of culture, with T. maxima CU1 and P. sanguineus CS43 being the greatest producers with 359 and 326 U/flask, respectively. The best laccase producer strain was P. sanguineus CS2 (435 U/flask); the rest of the isolates produced less than 10 U/flask. All of these results are shown in Table 4.

Table 4. Lignocellulolytic Enzyme Activity in Wheat Straw Agitated Liquid Medium

CMCases and Xylanases at third day; Avicelases and ß-D-Glucosidase at fourth day, and Laccase at seventh day of growth.

* Liquid media on flask = 200 mL total volume

The results of the enzymatic production of all the strains grown in stationary conditions are shown in Table 5. The isolates Inonotus sp. CU7 (242 U/flask), T. maxima CU1 (170 U/flask) and Armilariellasp. CS134 (152 U/flask), were the best producers of CMCases. Strain T. maxima CU1 showed the highest levels of Avicelases (288 U/flask) and xylanases (165 U/flask). For the detection of β-D-glucosidase, strain P. sanguineus CS43 was the greatest producer with 476 (U/flask), whereas the rest of the fungi showed levels of less of 200U/flask. Under these culture conditions, the best laccase producer was P. sanguineus CS2 (59 U/flask); however, the levels were less than those detected in stirring conditions. The rest of the strains produced less than 10 U/flask.

Table 5. Lignocellulolytic Enzyme Activity in Wheat Straw Static Liquid Medium

CMCases and Xylanases on the third day; Avicelases and -D-Glucosidase on the fourth day, and

* Liquid media on flask = 200 mL total volume

^&Laccase on the seventh day of growth.

Regarding the production of LiP, in none of the isolates was the presence of LiP detected in the studied culture conditions. For the quantification of MnP in stirring conditions, Trametes maxima CU1 was the greatest producer with 10.2 U/flask at the seventh day of culture. Strains Armilariella sp CS134, Xylaria sp. CS121 and Inonotus sp. CU7 produced less than 1 U/flask between day 1 and 7 of growth. In stationary conditions, Trametes maxima CU1 presented the lowest concentration (6.2 U/flask) at a shorter time (third day), followed by Inonotus sp. CU7 with 4.3 U/flask. Xylaria sp. CS121 and Armilariella sp CS134 produced 1.6 and 1.0 U/flask, respectively, at day 5.

These results allow one to know the metabolic plasticity of the isolates as a function of the culture conditions, since the same fungus presented differences in the levels of enzymatic production. Isolate T. maxima CU1 was the greatest producer in stirring and stationary conditions, presenting high levels in the majority of the detected activities.

The levels of enzymatic production of the native isolates were similar to those reported for different strains of Lentinus polychrous, L. squarrosulus, and L. sajor-caju (Pukahuta et al. 2004), where differences were observed in the titers of the same strain. In the case of the laccase production, for these strains, greater titles from 0.43 to 3.15 U/mL were reported at day seven, while the native strains studied here had a range from 0.03 to 0.3 U/mL, with the exception of P. sanguineus CS2, which produced 2.17 U/mL. In the present work, low production levels of laccase were also observed in P. sanguineus CS43 (0.11 and 0.025 U/mL). Although these titers are low, it was found in previous reports that the laccases from P. sanguineus CS2 and P. sanguineus CS43 are thermostable enzymes (Salcedo-Martínez et al. 2013; Ramirez-Cárdenas et al. 2014). In addition, remarkably higher production levels (143,000 U/L) have been obtained for P. sanguineus CS43 through Central Composite Design (Ramirez-Cárdenas et al. 2014). Therefore, the exploration of new native strains can drive us to obtain new sources of lignocellulolytic enzymes with operational and functional properties more robust than those reported, despite the low levels of production.

Regarding the production of cellulases and hemicellulases in the work of Pukahuta et al. (2004), the majority of the strains were greater producers of xylanases than cellulases, while in the present work the fungi studied were greater producers of cellulases than xylanases. In the case of the work by Elisahvili and coworkers (2008), the production of CMCases and xylanases was higher than the total of cellulase activity, laccase, and MnP; but also there were differences observed in the titers of enzymatic production in function of the strain, culture conditions, as well as the composition of the medium. This has been reported by different authors, who have used substrates rich in lignocellulose of different origins for the enzymatic production of lignocellulases (Elisahvili et al. 2008, 2009, 2011; Isikhuemhen et al. 2012). These differences can be seen in all these works as a function of the medium and the culture conditions; therefore the selection of native strains with high lignocellulolytic potential can lead to the finding of new sources of novel enzymes, with robust operational and functional properties, despite their low levels of production, as it is the case of the thermostable laccase of P. sanguineus CS2, mentioned above. This possibility will be addressed in further studies of optimization of the enzymatic production, as well as its application in the development of new digestive adjuvants for the livestock industry.

CONCLUSIONS

Nuevo Leon has a great diversity of macromycetes that possess high lignocellulolytic, pectinolytic, and amylolytic potentials that can be exploited in diverse industrial processes.
Seven native strains with cellulolytic, pectinolytic, and amylolytic capacities greater than those showed by six international reference strains were obtained. Two of these native strains (T. maxima CU1 and P. sanguineus CS43), presented the greatest lignolytic potential as well.
In function of the media, as well as the culture conditions, the enzymes detected here can be obtained, or these conditions can be adjusted to obtain one in particular.

ACKNOWLEDGMENTS

The authors are grateful for the financial support provided by the Programa del Mejoramiento del Profesorado (PROMEP 103.5/12/7884) and the Programa de Apoyo a la Investigación Cientifica y Tecnológica de la UANL (CA953-11). We also thank Ing. Olivia Gaona Quintanilla for her critical revision and comments on the manuscript.

REFERENCES CITED

Abadulla, E., Tzanov, T., Costa, S., Robra, K. H., Cavaco-Paulo, A., and Gübitz, G. M. (2000). “Decolorization and detoxification of textile dyes with a laccase from Trametes hirsuta,” Applied and Environmental Microbiology 66(8), 3357-3362. DOI: 10.1128/AEM.66.8.3357-3362.2000

Arantes, V., and Milagres, A. M. (2009). “Relevance of low molecular weight compounds produced by fungi and involved in wood biodegradation,” Química Nova 32(6), 1586-1595. DOI: 10.1590/S0100-40422009000600043

Atri, N. S and Sharma, S. K. (2011). “Qualitative estimation of cellulases and lignin modifying enzymes in five wild Lentinus species selected from North India,” Academic Journal of Plant Sciences 4(4), 105-109. DOI: 10.5829/idosi.wjfpb.2012.3.1.304

Das, G., and Sen-Mandi, S. W. A. T. I. (1992). “Scutellar amylase activity in naturally aged and accelerated aged wheat seeds,” Annals of Botany 69(6), 497-501.

Elisashvili, V., Kachlishvili, E., Tsiklauri, N., Metreveli, E., Kahrdziani, T., and Agathos, S. N. (2009). “Lignocellulose-degrading enzyme production by white-rot Basidiomycetes isolated from the forest of Georgia,” World Journal of Microbiology and Biotechnology 25(2), 331-339. DOI: 10.1007/s11274-008-9897-x

Elisashvili, V., Torok, T., Kachlishvili, E., Khardziani, T., Metreveli, E., Kobakhidze, A., and Berikashvili, I. (2011). “Evaluation and regulation of the lignocellulolytic activity of novel white-rot basidiomycetes,” Global Journal of Biochemistry 2(2), 134-141.

Gutiérrez-Soto, J. G., Salcedo-Martínez, S. M., Contreras-Cordero, J. F., and Hernández-Luna, C. E. (2011). “Characterization of the major laccase from Trametes maxima CU1 and decolorization of nine commercially significant dyes by the enzyme,” Water R&D 1(1), 10-19. DOI: 10.5772/56374.

Han, J. (2003). “Solid-state fermentation of cornmeal with the basidiomycetes Hericium erinaceum for degrading starch and upgrading nutritional value,” International of Food Microbiology 80(1), 61-66. DOI: 10.1016/S0168-1605(02)00122-8

Han, J. R., An, C. H., and Yuan, J. M. (2005) “Solid-state fermentation of cornmeal with the basidiomycetes Ganoderma lucidum for degrading starch and upgrading nutritional value,” Journal of Applied Microbiology 99(4), 910-915.

Hernández-Luna, C. E., Gutiérrez-Soto, G., and Salcedo-Martínez, S. M. (2008). “Screening for decolorizing Basidiomycetes in Mexico,” World Journal of Microbiology and Biotechnology 24(4), 465-473. DOI: 10.1007/s11274-007.9495-3

Isikhuemhen, O. S., Mikiashvili, N. A., Adenipekun, C. O., Ohimain, E. I., and Shahbazi, G. (2012). “The tropical white rot fungus, Lentinus squarrosulus Mont.: Lignocellulolytic enzymes activities and sugar release from cornstalks under solid state fermentation,” World Journal of Microbiology and Biotechnology 28(5), 1961-1966. DOI: 10.1007/s11274-011-0998-6

Khokhar, I., Haider, M. S., Mushtaq, S., and Mukhtar, I. (2013). “Isolation and screening of highly cellulolytic filamentous fungi,” Journal Applied Sciences and Environmental Management 16(3), 223-226.

Levin, L., and Forchiassin, F. (1998). “Culture conditions for the production of pectinolytic enzymes by the white‐rot fungus Trametes trogii on a laboratory scale,” Acta biotechnologica 18(2), 157-166. DOI: 10.1002/abio.370180213

Miller, G. L. (1959). “Use of dinitrosalicylic acid reagent for determination of reducing sugar,” Analytical Chemistry 31(3), 426-428. DOI: 10.1021/ac60147a030

Mtui, G., and Masalu, R., (2010) “Extracellular enzymes from brown-rot fungus Laetiporus sulphureus isolated from mangrove forests of coastal Tanzania,” Scientific Research and Essays 3(4), 154-161.

Nazareth, S. W., and Sampy, J. D. (2003). “Production and characterization of lignocellulases of Panus tigrinus and their application,” International Biodeterioration and Biodegradation 52(4), 207-214. DOI: 10.1016/S0964-8305(03)00051-9

Pal, A., Roy, A., and Das, A. (1980). “Production of amylase by Polyporus ostreiformis,” Mycologia72(6), 1134-1141. DOI: 10.2307/3759567

Pandey, A., Nigam, P., Soccol, C. R. V. T., Soccol, V., Singh, D., and Mohan, R. (2000). “Advances in microbial amylases,” Biotechnology and Applied Biochemistry 31(2), 135-152. DOI: 10.1042/BA19990073

Pericin, D., Kevresan, S., Banka, L., Antov, M., and Škrinjar, M. (1992). “Separation of the components of pectinolytic complex produced by Polyporus squamosus in submerged culture,” Biotechnology Letters 14(2), 127-130.

Pericin, D., Antov, M., Dimic, N., and Vujicic, B. (1997). “Rapid method for detecting low basal activity of exo-pectinase of Polyporus squamosus,” Biotechnology Techniques 11(11), 833-836. DOI: 10.1023/A:1018485510779

Pointing, S. (2001). “Feasibility of bioremediation by white-rot fungi,” Applied Microbiology and Biotechnology 57(1), 20-33. DOI: 10.1007/s002530100745

Pozdnyakova, N. N., Turkovskaya, O. V., Yudina, E. N., and Rodakiewicz-Nowak, Y. (2006). “Yellow laccase from the fungus Pleurotus ostreatus D1: Purification and characterization,” Journal of Applied Biochemistry and Microbiology 42(1), 56-61. DOI: 10.1134/S000368380601008X

Salcedo-Martínez, S. M, Gutiérrez-Soto, G., Rodríguez, C. F., Villarreal, T. J., Contreras, J. F., and Hernández-Luna, C. E. (2013). “Purification and partial characterization of a thermostable laccase from Pycnoporus sanguineus CS-2 with ability to oxidize high redox potential substrates and recalcitrant dyes,” in: Applied Bioremediation – Active and Passive Approaches Y. B. Patil and P. Rao (eds.), InTech, Rijeka, Croatia. DOI: 10.5772/56374

Sánchez, C. (2009). “Lignocellulosic residues: Biodegradation and bioconversion by fungi,” Biotechnology Advances 27(2), 185-194. DOI: 10.1016/j.biotechadv.2008.11.001

Sigoillot, C., Lomascolo, A., Record, E., Robert, J. L., Asther, M., and Sigoillot, J. C. (2002). “Lignocellulolytic and hemicellulolytic system of Pycnoporus cinnabarinus: Isolation and characterization of a cellobiose dehydrogenase and a new xylanase,” Enzyme and Microbial Technology 31(6), 876-883. DOI: 10.1016/S0141-0229(02)00208-9

Sin, M. K. W., Hyde, K. D., and Pointing, S. B. (2002). “Comparative enzyme production by fungi from diverse lignocellulosic substrates,” Journal of Microbiology 40(3), 241-244. ISSN: 1225-8873

Steffen, K. T., Cajthaml, T., Šnajdr, J., and Bladrian, P. (2007). “Differential degradation of oak (Quercus petraea) leaf litter by litter-decomposing basidiomycetes,” Research in Microbiology 158(5), 447-455. DOI: 10.1016/j.resmic.2007.04.002

Valášková, V., and Baldrian, P. (2006). “Degradation of cellulose and hemicelluloses by the brown rot fungus Piptoporus betulinus – Production of extracelular enzymes and characterization of the major cellulases,” Microbiology 152(2), 3613-3622. DOI: 10.1099/mic.0.29149-0

Xavier-Santos, S., Carvalho, C. C., Bonfá, M., Silva, R., Capelari, M., and Gomes, E. (2004). “Screening for pectinolytic activity of wood-rotting basidiomycetes and characterization of the enzymes,” Folia Microbiologica 49(1), 46-52. DOI: 10.1007/BF02931645

Younes, B. S., Karray, F., and Sayadi, S. (2011). “Isolation of thermophilic fungal strains producing oxido-reductase and hydrolase enzymes from various Tunisian biotopes,” International Biodeterioration and Biodegradation 65(7), 1104-1109. DOI: 10.1016/j.ibiod.2011.08.003

Article submitted: January 23, 2015; Peer review completed: July 6, 2015; Revised version received: August 1, 2015; Accepted: August 10, 2015; Published: August 19, 2015.

DOI: 10.15376/biores.10.4.6676-6689