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Montibeller, V. W., Vandenberghe, L. P. S., Amore, A., Soccol, C. R., Birolo, L., Vinciguerra, R., Salmon, D. N. X., Spier, M. R., and Faraco, V. (2014). "Characterization of hemicellulolytic enzymes produced by Aspergillus niger NRRL 328 under solid state fermentation on soybean husks," BioRes. 9(4), 7128-7140.


This manuscript describes the analysis of xylanase production by Aspergillus niger NRRL 328 in solid state fermentation (SSF) of soybean husks. A maximum value of extracellular xylanase activity of approximately 950 U g-1 was achieved after 96 h. Proteomic analyses performed on the enzymatic mixture responsible for the maximum value of xylanase activity in SSF revealed the presence of two xylanases. This xylanolytic mixture was partially purified and characterized. It followed Michaelis-Menten kinetics towards xylan, with a KM of 7.92 ±0.97 mg xylan/mL and a Vmax of 262.2 ± 27.8 g L-1 s-1. The optimum pH for the enzyme is 5.3, and the optimal temperature is 50 °C. The enzyme retains 100% of its activity at 40 °C for at least 1 month. It shows very high stability in a broad pH range, with a half-life of 40 days at pH 5.3, pH 6.0, pH 6.5, pH 7.0, and pH 8.0.

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Characterization of Hemicellulolytic Enzymes Produced by Aspergillus niger NRRL 328 under Solid State Fermentation on Soybean Husks

Valesca Weingartner Montibeller,a Luciana Porto de Souza Vandenberghe,Antonella Amore,b Carlos R. Soccol,a Leila Birolo,b Roberto Vinciguerra,Denise N. Xavier Salmon,a Michele Rigon Spier,a and Vincenza Faraco b,*

This manuscript describes the analysis of xylanase production by Aspergillus niger NRRL 328 in solid state fermentation (SSF) of soybean husks. A maximum value of extracellular xylanase activity of approximately 950 U g-1 was achieved after 96 h. Proteomic analyses performed on the enzymatic mixture responsible for the maximum value of xylanase activity in SSF revealed the presence of two xylanases. This xylanolytic mixture was partially purified and characterized. It followed Michaelis-Menten kinetics towards xylan, with a Kof 7.92 ±0.97 mg xylan/mL and a Vmax of 262.2 ± 27.8 g L-1s-1. The optimum pH for the enzyme is 5.3, and the optimal temperature is 50 °C. The enzyme retains 100% of its activity at 40 °C for at least 1 month. It shows very high stability in a broad pH range, with a half-life of 40 days at pH 5.3, pH 6.0, pH 6.5, pH 7.0, and pH 8.0.

Keywords: Aspergillus niger; Solid state fermentation; Xylanase

Contact information: a: Department of Bioprocess Engineering and Biotechnology, Federal University of Paraná, Coronel Francisco H. dos Santos Avenue, 210, Zip Code 81531-990 Curitiba, Brazil; b: Department of Chemical Sciences, University of Naples “Federico II,” Complesso Universitario Monte S. Angelo, via Cintia, 4 80126 Naples, Italy; *Corresponding author:


Hemicellulose represents 20 to 35% of lignocellulosic biomass and is a very complex polysaccharide composed of a wide variety of sugars, including xylose, arabinose, mannose, and galactose (Saha 2003). The hemicellulose enzymes degrading hemicellulose are divided into two major groups: depolymerizing enzymes, which cleave the backbone, and auxiliary enzymes, which remove substituents. These enzymes act synergistically to hydrolyze lignocellulosic polysaccharides (Van Dyk and Pletschke 2012). Depolymerizing enzymes include endo-β-1,4-xylanases (EC and β-xylosidases (EC, while auxiliary enzymes comprise α-glucuronidases (EC 3.2.1), α-arabinofuranosidases (EC, acetyl esterases or acetyl xylan esterases (EC, and β-mannanases (EC

Most studies on hemicellulases have focused on enzymes that hydrolyze the xylan backbone. These enzymes include, according to the CAZY (Carbohydrate-Active enZYmes) database, GH 3, 30, 39, 43, 52, 54, 116, and 120. Endo-1,4--xylanase (1,4-b-d-xylan xylanohydrolase, EC cleaves the glycosidic bonds in the xylan backbone, releasing shorter xylo-oligosaccharides; -xylosidase (1,4-b-d-xylan xylohydrolase, EC cleaves the small xylo-oligosaccharides and cellobiose into xylose. These enzymes are produced by microorganisms such as fungi, bacteria, yeast, and marine algae. Filamentous fungi are interesting producers because the enzymes are excreted at higher levels than those of yeasts and bacteria.

The use of xylanases is very well established in the food industry, for the digestion of animal feed, in the textile industry, as well as in the paper and pulp industry for bleaching processes to replace the chlorine utilization and to reduce the environmental impacts (Juturu and Wu 2012). Lately, the application of hemicellulases in lignocellulose conversion is increasing because of the progress in lignocellulose biorefinery installation (Kumar and Satyanarayana 2012). In this case, a finely planned cocktail of enzymes, consisting not only of cellulases but also of xylanases, is required to achieve the highest yields of fermentable sugars from the polysaccharidic components of lignocellulosic biomass. However, the high cost of enzymes is one of the major economic bottlenecks for the implementation of economically feasible lignocellulosic biorefineries (Hasunuma et al. 2013).

A major challenge therefore entails the development of a suitable efficient and economically viable hydrolysis process step, through discovery of new enzymes with higher catalytic efficiency, recycling enzymes during the hydrolysis and fermentation process, and on-site enzyme production in the biorefinery facility. In this way, the development of processes, with the employment of agro-industrial residues, is a good alternative for enzyme production. Among processes used for enzyme production, solid-state fermentation (SSF) is an attractive one. SSF systems resemble the natural habitats of microbes and, therefore, may prove to be efficient in producing certain enzymes and metabolites (Haltrich et al. 1996). This biotechnological process allows the use of inexpensive substrates that decrease the cost of enzyme production (Pandey et al. 2000; Soccol and Vandenberghe 2003). The cost of carbon source plays a major role in the economics of xylanase production. An approach to reduce the cost of xylanase production is the use of lignocellulosic materials as substrates rather than opting for the expensive pure xylans (Haltrich et al. 1996).

Brazil is the second largest producer of soybean, with 30.2% of the world production or 86.5 million tons in 2014. According to the Ministry of Agriculture, Livestock and Supply of Brazil (IBGE 2014), the exportation of soybean grains generates more than $5.5 billion dollar per year and its local processing to produce products with high value is of paramount importance for the economic development and job creation in the country (MAPA 2014). Soybean husks (SH) are one of the byproducts/wastes obtained in the process of extraction of seed oil. For each ton of processed soybeans, up to 3% shells, are generated (Karp et al. 2011).

Several innovative processes and bioproducts linked to the productive chains of soybean, which were transferred to the productive sector, have already been developed. Among these, are bioproducts obtainable from soybean hulls and molasses that include bioethanol and acids (Siqueira et al. 2008, Sanada et al. 2009; Karp et al. 2011; Letti et al. 2012), and extracts of soybean meal as bacterial growth inhibitors (Noseda et al. 2012). This substrate could also be used for xylanases and other enzymes production.

This manuscript describes the production of xylanases from the strain Aspergillus niger NRRL 328 through solid state fermentation (SSF) on soybean husks, as well as partial purification, proteomic analyses for enzymatic identification, and functional characterization of extracellular xylanolytic enzymes.




The fungus Aspergillus niger NRRL 328, from the Agricultural Research Service – ARS Culture Collection, was cultivated on potato dextrose agar (Sigma-Aldrich) slants at 28 °C for 168 hand stored at 4 °C. The culture was maintained at the Culture Collection of the Biotechnological Process Laboratory, Bioprocess Engineering and Biotechnology Division, Federal University of Paraná, Curitiba, Brazil. The strain was previously selected as good xylanase producer.


The pre-inoculum of A. niger NRRL 328 was cultivated in Erlenmeyer flasks (125 mL) containing 20 mL of potato dextrose agar (Sigma-Aldrich, Milan, Italy) and incubated at 28 °C for 144 h. Spore suspensions were obtained by mixing pre-cultured fungus in 20 mL of sterile solution Tween 80® 0.01% containing glass pearls in constant agitation at 120 rpm for 10 min. The concentration of spores/mL was determined by counting using a Bürker camera.


SSF for xylanase production analysis

Approximately 10spores/g of A. niger NRRL 328 were inoculated on 2.5 g of soybean husks, between 0.8 and 2.0 mm, in 100-mL Erlenmeyer’s flasks, in triplicate. Previous studies with other lignocellulosic wastes had shown that this is the best dimension for SSF (Soccol and Vandenberghe 2003; Maciel et al. 2008; Iandolo et al. 2011). The initial moisture was adjusted to 70% utilizing a nutritive solution containing (g/L) manganese sulfate (1.0), and urea (0.7). The flasks were incubated at 28 °C for 168 h. Enzymes produced by SSF were extracted by solid-liquid extraction using a 50 mM sodium citrate buffer, pH 5.0, at the proportion of 1:10 (w/v) with vigorous agitation for 10 min. The mix was filtered in TNT paper and centrifuged at 8000 rpm, 4 °C for 15 min. The supernatant was filtered using Whatman nº 1 filter paper and subjected to analyses.

Xylanase activity assay

The xylanase activity assay was performed using a modified version of the methodology reported by Bailey et al. (1992). An aliquot of 20 µL of enzymatic filtrate (appropriately diluted in 50 mM sodium citrate buffer, pH 5.0) was mixed with 180 µL of 1% beech wood xylan and incubated for 5 min at 50 °C. Released reducing sugars were quantified using the dinitrosalicylic acid reagent (DNS) method (Miller 1959), adding 300 µL of DNS solution and then incubating the mixture at 95 °C for 5 min. Absorbance was measured at 540 nm. One unit of enzyme is defined as the amount of enzyme catalyzing the release of 1 μmol of xylose equivalent per min.

Protein concentration determination

Protein concentration was determined by the method of Lowry et al. (1951), using the BioRad Protein Assay (BioRad Laboratories S.r.l.; Segrate, MI, Italy), with bovine serum albumin as a standard.

Enzyme enrichment

Supernatant obtained by extraction of the enzymes produced by SSF was precipitated by the addition of (NH4)2SO4 up to 80% saturation at 4 °C and centrifugation at 10,000×g for 30 min. The precipitate was resuspended in 0.02 M Tris-HCl at pH 7.5 and loaded on HiTrap Phenyl FF high sub (GE Healthcare; Uppsala, Sweden), equilibrated in buffer A (0.02 M Tris-HCl, 1.2 M (NH4)2SO4, pH 7.5), and the proteins were eluted isocratically with buffer B (0.02 M Tris-HCl pH 7.5). Fractions containing activity were combined and concentrated on an Amicon PM-10 membrane (Millipore, Vimodrone, Italy) and analyzed by SDS-PAGE (Sodium Dodecyl Sulphate–Poly-Acrylamide Gel Electrophoresis) according to Laemmli (1970).

Optimal temperature and thermo-resistance

To determine the optimum temperature of the partially purified enzyme, the substrate of the activity assay (1% beech wood xylan) was dissolved in 50 mM sodium citrate buffer at pH 5.3 and the incubation (5 min) was performed at 40, 50, 60, 70, and 80 °C. The thermo-resistance was studied by incubating the partially purified enzyme preparation in sodium citrate buffer at pH 5.3 at 40, 50, and 60 °C. The samples withdrawn were assayed for residual xylanase activity, performing incubation (5 min) at 50 °C. The reported values are representative of three experiments, and each experiment was performed in duplicate.

Optimal pH and pH stability

To determine the optimum pH of the partially purified enzyme, the substrate of the activity assay (1% beech wood xylan) was dissolved in citrate phosphate buffers (McIlvaine 1921) with pH values between 3.0 and 7.0, and in 50 mM Tris-HCl with pH values of 7.0, 8.0, and 9.0, and then incubated (5 min) at 50 °C. The pH stability of the purified enzyme preparation was studied by diluting it in citrate phosphate buffers, pH 5.3 – 6.5 – 7.0 – 8.0, and incubating at 25 °C. From time to time, samples were withdrawn and immediately assayed for residual xylanase activity, performing incubation (5 min) at 50 °C. The reported values are representative of three experiments, and each experiment was performed in duplicate.

Determination of kcat and KM

For determination of the Michaelis-Menten constants KM and kcat, the activity assay was performed at beech wood xylan concentrations ranging from 0.2 mg/mL to 25 mg/mL at pH 5.3, performing incubation for 5 min at 50 °C. The reported values are representative of three experiments, and each experiment was performed in duplicate.

Zymogram analyses

Semi-denaturing gel electrophoresis was carried out by loading non-denatured and not-reduced samples on a SDS polyacrylamide gel, performed as described by Laemmli (1970).

Proteins showing xylanolytic activity purified by HIC were visualized as followsAfter electrophoresis, the gel was soaked in the same buffer used for dissolving proteins and gently shaken to remove SDS and to renature the proteins in the gel.

Protein identification by mass spectrometry

Active fractions were diluted in denaturant buffer (final concentrations: Tris 300 mM pH 8.8, urea 6 M, EDTA 10 mM), and disulfide bridges were reduced with 10 mM dithiothreitol at 37 °C for 2 h and then alkylated by adding 50 mM iodoacetamide at room temperature for 30 min in the dark. Protein sample was desalted by size exclusion chromatography on a Shephadex G-25M column (GE Healtcare, Milan, Italy) equilibrated in 50 mM NH4HCObuffer, pH 8.0. Protein-containing fractions were frozen at -80 °C, and lyophilized by Vacuum Freeze Dryer Lyophilizer system (Lio 5Pascal) at -50 °C and with a forced vacuum of 0.30 mbar. Lyophilized fractions were dissolved in 50 µL of 10 mM NH4HCObuffer, pH 8.0, and enzymatic digestion was performed with 5 µg of trypsin at 37 °C for 16 h.

Peptide mixtures were filtered on a 0.22-µm PVDF membrane (Millipore) and analysed by LC-MS/MS (liquid chromatography coupled with tandem mass spectrometry) on a 6520 Accurate-Mass Q-TOF LC/MS (liquid chromatography mass spectrometry) System (Agilent Technologies, Palo Alto, CA), equipped with a 1200 HPLC (High-Performance Liquid Chromatography) system and a chip cube (Agilent Technologies, Palo Alto, CA). After loading, the peptide mixture was concentrated and washed in a 40-nL enrichment column (Agilent Technologies chip, Palo Alto, CA), with 0.1% formic acid in 2% acetonitrile as the eluent.

The sample was then fractionated on a C18 reverse-phase capillary column (Agilent Technologies chip, Palo Alto, CA) at a flow rate of 400 nL/min, with a linear gradient of eluent B (0.1% formic acid in 95% acetonitrile) in A (0.1% formic acid in 2% acetonitrile) from 7 to 80% in 50 min. Peptide analysis was performed using data-dependent acquisition of one MS scan (mass range from 300 to 1800 m/z) followed by MS/MS scans of the five most abundant ions in each MS scan. The MS/MS spectra were measured automatically when the MS signal was over the threshold of 50,000 counts. Double- and triple-charged ions were preferably isolated and fragmented over single-charged ions. The acquired MS/MS spectra were transformed in mzData (.XML) format and used for protein identification with a licensed version of MASCOT software (, version 2.4.0.

Raw data from nano LC-MS/MS analysis were used to query the NCBInr database (the nr database compiled by the National Center for Biotechnology Information), NCBInr 20121120 (21,582,400 sequences; 7,401,135,489 residues), with taxonomy restriction to Fungi (1,569,912 sequences). The MASCOT search parameters were as follows: trypsin as enzyme; 3, as allowed number of missed cleavages; carboamidomethyl (C) as fixed modification; oxidation of methionine and pyro-Gluformation at N-term Q as variable modifications; 10 ppm as MS tolerance and 0.6 Da as MS/MS tolerance; and peptide charge from +2 to +3.

The peptide score threshold provided from MASCOT software to evaluate the quality of matches for MS/MS data was 40. Spectra with a MASCOT score of < 20 were rejected as having low quality.

Trypsin, dithiothreitol, iodoacetamide, and NH4HCO3 were purchased from Sigma-Aldrich (Milan, Italy). Trifluoroacetic acid (TFA)-HPLC grade was from Carlo Erba (Milan, Italy). All other reagents and solvents were of the highest purity available from Baker (Milan, Italy).


Xylanase Production by Solid State Fermentation

Different substrates were already tested in previous studies by the group for xylanase production (Maciel et al. 2008). Soybean husks were chosen for xylanase production due to their main lignocellulolytic composition (34.5% fibers, 13.2% proteins), which could promote not only the production of xylanases, but the production of other enzymes such cellulases, etc. The choice of the use of only soybean husks was done due to the interest of concomitantly producing mannanases. Seeds present an interesting composition of galactomannans. So, soybean husks certainly would present the favorable composition for that.

Analysis of xylanase activity production by the strain A. niger NRRL 328 showed that a maximum production of xylanase activity of around 950 U g-1was achieved by SSF on soybean husks at 96 h (Fig. 1), with a productivity of 9.89 U g-1h-1.

Fig. 1. Time course of xylanase activity (U g-1) produced by A. niger NRRL 328 in SSFon soybean husks

When compared with xylanase activity produced by other Aspergillus spp., the maximum xylanase activity value obtained for A. niger NRRL 328 was similar or, in some cases, lower than those obtained by SSF with most Aspergillus strains (Table 1). Even so, this process presents good perspective for xylanase production. It must be pointed out that the enzyme production was not optimized and/or concentrated, which could lead to a 5- to 10-fold augmentation of enzyme activity. However, this fact may not restrain the direct application of the fermented material with the enzymes in feed. Rations are generally composed of the most commonly available feed ingredients in alfalfa, corn (grain and silage), grass hay, soybean meal, and pasture (Peters et al. 2014). The mixture of the fermented substrate, soybean husks, containing xylanases and other produced hemicellulolytic enzymes, with ration components can be a good alternative for their employment for animal feed. The complete analysis of other enzymes concomitantly produced will certainly define the real potential for the use of the fermented material. Besides, the direct use of the fermented material with enzymes would eliminate the need and costs of separation and purification steps.

Table 1. Comparison of Xylanolytic Activity Values Reported for Other Aspergillus spp.

Enzyme Identification and Characterization

The xylanolytic enzymes produced by A. nigerNRRL 328SSF were partially purified, as shown by SDS-PAGE (Fig. 2), and subjected to proteomic analyses and functional characterization as described below.

Fig. 2. SDS-PAGE profiling of the partially purified xylanase from A. niger NRRL 328: Lane 1: molecular marker; lane 2: partially purified xylanase

The sample positive for xylanolytic activity, as assessed by zymogram analyses, was enzymatically digested, and the peptide mixture was analyzed by LC-MS/MS. Database searches confidently assessed the presence of an endo-1,4-β-xylanase C and a xylanase that can account for the observed activity. Interestingly, anα-L-arabinofuranosidase axhA, an endoglucanase A, and an endo-β-1,4-glucanase B were also identified (Table 2) in the same sample.

Table 2. Proteins Identified in the Sample with Xylanolytic Activity

The NCBInr database was searched with MS/MS ion search MASCOT software (, with carboamidomethyl (C) as fixed modification and oxidation on Metand cyclization Pyro-Glu of Gln at N-terminus of the peptides as variable modifications. Only proteins identified with at least two peptides were considered significant. Peptides with individual ion score < 20 were rejected

Fig. 3. Effect of pH (A) and temperature (B) on xylanase activity produced by A. niger NRRL 328

Functional characterization of the enriched xylanolytic mixture showed that xylanase activity follows Michaelis-Menten kinetics towards xylan: the KMfor this substrate was 7.92 ±0.97 mg xylan/mL and the Vmax was 262.2 ± 27.8 g L-1 s-1. The optimum pH for the xylanase activity (assayed in a range from 3.0 to 8.0) was 5.3 (Fig. 3a), and the optimal temperature was 50 °C (Fig. 3b).

Both the optimum temperature and pH of A. niger NRRL 328 xylanase activity were similar to those of most xylanases from Aspergillus spp. (Table 3).

Table 3. Properties Reported for Xylanases from Other Filamentous Fungi

The A. niger NRRL 328 xylanase activity showed a half-life of 2 h at 50 °C and 15 min at 60 °C, with a similar behavior to that of most xylanases from Aspergillus spp. (Table 3). It is worth noting that the enzymatic mixture retained 100% of its activity for at least 1 month at 40 °C. It showed very high stability in a broad pH range, with a half-life of 40 days at pH 5.3, pH 6.0, pH 6.5, pH 7.0, and pH 8.0.


  1. The highest level of Aspergillus niger NRRL 328 xylanase activity achieved by SSF on soybean husks was around 950 U g-1 at 96 h.
  2. Proteomic analyses on partially purified enzymatic mixtures produced by A. niger NRRL 328SSF on soybean husks at 96 h revealed the presence of endo-1,4-β-xylanase C and xylanase that can account for the observed xylanase activity.
  3. Functional characterization of the enriched xylanolytic mixture showed that xylanase activity follows Michaelis-Menten kinetics towards xylan: the KM for this substrate is 7.92 ±0.97 mg xylan/mL, and the Vmax is 262.2 ± 27.8 g L-1 s-1.
  4. The optimum pH for the xylanase activity is 5.3, and the optimal temperature is 50 °C.
  5. The xylanase activity shows a half-life of 2 h at 50 °C and 15 min at 60 °C.
  6. The enzymatic mixture retains 100% of its xylanase activity for at least 1 month at 40 °C. It shows very high stability in a broad pH range, with a half-life of 40 days at pH 5.3, pH 6.0, pH 6.5, pH 7.0, and pH 8.0.


This work was supported by a grant from European Commission Marie Curie International Research Staff Exchange Scheme within the 7th European Community Framework Programme: “Improvement of technologies and tools, e.g. biosystems and biocatalysts, for waste conversion to develop an assortment of high added value eco-friendly and cost-effective bio-products” BIOASSORT (contract number 318931). The authors also thank Coordination for the Improvement of Higher Education Personnel (CAPES) and the National Council of Technological and Scientific Development (CNPq) of Brazil.


Ang, S. K., Shaza, E. M., Adibah, Y., Suraini, A. A., and Madihah, M. S. (2013). “Production of cellulases and xylanase by Aspergillus fumigatus SK1 using untreated oil palm trunk through solid state fermentation,” Process Biochemistry 48(9), 1293-1302.

Bailey, M. J., Biely, P., and Poutanen, K. (1992). “Interlaboratory testing of methods for assay of xylanase activity,” Journal of Biotechnology 23(3), 257-270.

Delabona, P. S., Pirota, R. D. P. B., Codima, C. A., Tremacoldi, C. R., Rodrigues, A., and Farinas, C. S. (2013). “Effect of initial moisture content on two Amazon rainforest Aspergillus strains cultivated on agro-industrial residues: Biomass-degrading enzymes production and characterization,” Industrial Crops and Products 42, 236-242.

Farinas, C. S., Loyo, M. M., Junior, A. B., Tardioli, P. W., Neto, V. B., and Couri, S. (2010). “Finding stable cellulase and xylanase: Evaluation of the synergistic effect of pH and temperature,” New Biotechnology 27(6), 810-815.

Haltrich, D., Nidetzky, B., Kulbe, K. D., Steiner, W., and Zupancic, S. (1996). “Production of fungal xylanases,” Bioresource Technol. 58(2), 137-161.

Hasunuma, T., Okazaki, F., Okai, N., Hara, Y., Ishii, J., and Kondo, A. (2013). “A review of enzymes and microbes for lignocellulosic biorefinery and the possibility of their application to consolidated bioprocessing technology,” Bioresour. Technol. 135, 513-522.

Iandolo, D., Piscitelli A., Sannia G., and Faraco V. (2011). “Enzyme production by solid substrate fermentation of Pleurotus ostreatus and Trametes versicolor on tomato pomace,” Applied Biochemistry and Biotechnology. 163(1), 40-51

IBGE, Instituto Brasileiro de Geografia e Estatística (2014) Accessed 28 September 2014

Instituto Brasileiro de Geografia e Estatística – IBGE (2010) Accessed 28 May 2010

Jiang, Z., Cong, Q., Yan, Q., Kumar, N., and Du, X. (2010). “Characterization of a thermostable xylanase from Chaetomium sp. and its application in Chinese steamed bread,” Food Chemistry 120(2), 457-462.

Juturu, V., and Wu, J. C. (2012). “Microbial xylanases: Engineering, production and industrial applications,” Biotechnology Advances 30, 1219-1227.

Kamat, S., Khot, M., Zinjarde, S., RaviKumar, A., and Gade, W. N. (2013). “Coupled production of single cell oil as biodiesel feedstock, xylitol and xylanase from sugarcane bagasse in a biorefinery concept using fungi from the tropical mangrove wetlands,” Bioresource Technology 135, 246-253.

Karp, S. G., Igashiyama, A. H., Siqueira, P. F., Carvalho, J. C., Vandenberghe, L. P. S., Thomaz-Soccol, V., Coral, J., Tholozan, J. L., Pandey, A., and Soccol, C. R. (2011). “Application of the biorefinery concept to produce L-lactic acid from the soybean vinasse at laboratory and pilot scale,” Bioresour. Technol. 102(2), 1765-1772.

Laemmli, U. K. (1970). “Cleavage of structural proteins during the assembly of the head of bacteriophage T4,” Nature 227(5259), 680-685.

Lakshmi, G. S., Rao, C. S., Rao, R. S., Hobbs, P. J., and Prakashman, R. S. (2009). “Enhanced production of xylanase by a newly isolated Aspergillus terreus under solid state fermentation using palm industrial waste: A statistical optimization,” Biochemical Engineering Journal 48, 51-57.

Letti, L. A. J., Karp, S. G., Woiciechowski, A. L., and Soccol, C. R. (2012). “Ethanol production from soybean molasses by Zymomonas mobilis,” Biomass and Bioenergy 44, 80-86.

Liao, H., Xu, C., Tan, S., Wei, Z., Ling, N., Yu, G., Raza, W., Zhang, R., Shen, Q., and Xu, Y. (2012). “Production and characterization of acidophilic xylanolytic enzymes from Penicillium oxalicum GZ-2,” Bioresource Technology 123, 117-124.

López, J. A., Lázaro, C. C., Castilho, L. R., Freire, D. M. G., and Castro, A. M. (2013). “Characterization of multienzyme solutions produced by solid-state fermentation of babassu cake, for use in cold hydrolysis of raw biomass,” Biochemical Engineering Journal 77, 231-239.

Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951). “Protein measurement with the Folinphenol reagent,” J. Biol. Chem. 193, 265-275.

Maciel, G. M., Vandenberghe, L. P., Winson, I. H., Fendrich, R. C., Bianca, B. E. D., Brandalize, T. Q. S., Pandey, A., and Soccol, R. C. (2008). “Xylanase production by Aspergillus niger LPB 326 in solid state fermentation using experimental design,” Food Technol. Biotechnol. 46, 181-187.

MAPA, Ministério da Agricultura, Pecuária e Abastecimento (2014) Accessed 25 September2014

MASCOT software,

McIlvaine, T. C. (1921). “A buffer solution for colorimetric comparison,” J. Biol. Chem. 49, 183-186.

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

Ministério da Agricultura, Pecuária e Abastecimento – MAPA (2010) Accessed 25 September 2014

Moreira, L. R., Campos, M. C., Siqueira, P. H. V. M., Silva, L. P., Ricart, C. A., Martins, P. A., Queiroz, R. M. L., and Filho, E. X. F. (2013). “Two β-xylanases from Aspergillus terreus: Characterization and influence of phenolic compounds on xylanase activity,” Fungal Genetics and Biology 60, 46-52.

Ncube, T., Howard, R. L., Abotsi, E. K., van Rensburg, E. L. J., and Ncube, I. (2012). “Jatropha curcas seed cake as substrate for production of xylanase and cellulase by Aspergillus niger FGSCA733 in solid-state fermentation,” Industrial Crops and Products 37(1), 118-123.

Pal, A., and Khanum, F. (2010). “Production and extraction optimization of xylanase from Aspergillus niger DFR-5 through solid-state-fermentation,” Bioresource Technology 101(19), 7563-7569.

Pal, A., and Khanum, F. (2011). “Purification of xylanase from Aspergillus niger DFR-5: Individual and interactive effect of temperature and pH on its stability,” Process Biochemistry 46(4), 879-887.

Pandey, A., Soccol, C. R. Nigam, P., Soccol, V. T., Vandenberghe, L. P. S., and Mohan, R. (2000). “Biotechnological potential of agroindustrial residues: II-cassava bagasse,” Bioresource Technology 74, 81-87.

Peters, C. J., Picardy, J. A. Darrouzet-Nardi, A., and Griffin, T. S. (2014). “Feed conversions, ration compositions, and land use efficiencies of major livestock products in U.S. agricultural systems,” Agricultural Systems 130, 35-43.

Rodríguez-Fernández, D. E., Rodriguez-Leon, J. A., Carvalho, J. C., Sturm, W., and Soccol, C. R. (2011). “The behavior of kinetic parameters in production of pectinase and xylanase by solid-state fermentation,” Bioresource Technology 102, 10657-10662.

Saha, B.C. (2003). “Hemicellulose bioconversion,” J. Ind. Microbiol. Biotechnol. 30(5), 279-291.

Sanada, S. T. N., Karp, S. G., Spier, M. R., Portella, A. C., Gouvêa, P. M., Yamaguishi, C. T., Vandenberghe, L. P. S., Pandey, A., and Soccol, C. R. (2009). “Utilization of soybean vinasse for a-galactosidase production,” Food Research International 42, 476-483.

Siqueira, P. F., Karp, S. G., Carvalho, J. C., Sturm, W., Rodríguez-León, J. A., Tholozan, J.-L., Singhania, R. R., Pandey, A., and Soccol, C. R. (2008). “Production of bio-ethanol from soybean molasses by Saccharomyces cerevisiae at laboratory, pilot and industrial scales,” Bioresource Technology 99, 8156-8163.

Soccol, C. R., and Vandenberghe, L. P. S. (2003). “Overview of applied solid-state fermentation in Brazil,” Biochemical Engineering Journal 13, 205-218.

Vafiadi, C., Christakopoulos, P., and Topakas, E. (2010). “Purification, characterization and mass spectrometric identification of two thermophilic xylanases from Sporotrichum thermophile,” Process Biochemistry 45(3), 419-424.

Van Dyk, J. S, and Pletschke, B. I. (2012). “A review of lignocellulose bioconversion using enzymatic hydrolysis and synergistic cooperation between enzymes – Factors affecting enzymes, conversion and synergy,” Biotechnology Advances 30(6), 1458-1480.

Article submitted: May 12, 2014; Peer review completed: August 9, 2014; Revised version received: September 29, 2014; Accepted: October 1, 2014; Published: October 10, 2014.