Optimization of Cellulase Production from Agri-Industrial Residues by Aspergillus terreus NIH2624

Abstract

The objective of this study was to assess the cellulase production of four fungi: Aspergillus terreus NIH2624, Aspergillus clavatus NRRL1, Aspergillus versicolor CBS583.65 and Aspergillus phoenicis ATCC3157, under submerged cultivation conditions. When these fungi were cultured in shake flasks using Mandels and Weber’s minimal medium with 1% sugarcane bagasse as a carbon source and 1.8 g/L of rice bran extract as a nitrogen source, A. terreus showed maximum cellulase production (filter paper activity (FPase) 3.35 U/mL; carboxymethyl cellulase activity (CMCase) 1.69 U/mL). Consequently, A. terreus was selected for the optimization study for cellulase production. Among the different tested carbon sources, A. terreus showed higher CMCase activity when it was cultivated on delignified sugarcane bagasse (1.64 U/mL) and higher FPase activity on sugarcane straw (7.95 U/mL). Regarding the nitrogen sources, the maximum FPase activity was observed when using rice bran (FPase, 8.90 U/mL) and soybean meal (FPase, 9.63 U/mL). The optimized fermentation medium (minimal medium with delignified sugarcane bagasse and rice bran as carbon and nitrogen sources, respectively) resulted in an enzymatic cocktail mainly composed of xylanases, with a maximum activity of 1701.85 U/mL for beechwood xylan, 77.12 U/mL for endoglucanase and 21.02 U/mL for cellobiohydrolase. Additionally, the enzymatic cocktail showed efficient activities for β-glucosidase, β-xylanase, arabinofuranosidase and lytic polysaccharide monoxygenases (LPMOs). This cellulase enzyme solution has the potential to efficiently hydrolyze lignocellulosic biomass, producing second-generation sugars in biorefineries.

1. Introduction

The limited availability of gasoline and its consequent impacts on the environment are important concerns that highlight the urgent demand for sustainable gasoline alternatives. One of the main challenges in the renewable industry is the conversion of lignocellulosic biomass into fermentable sugars at a low cost [1]. Among the strategies proposed to reduce the cost of cellulase production, the use of agro-industrial residues from feedstock shows great promise due to their wide availability and low cost [2]. Agricultural residues such as sugarcane bagasse, sugarcane straw, corn cob, rice straw, etc. have been recognized as viable substrates to produce fungal enzymes [3]. These substrates are abundantly available throughout the year in many agricultural countries, including Brazil, the USA, China and India. Therefore, such lignocellulosic biomasses can be utilized to develop sustainable cellulase production processes in fermentation industries, holding significant economic importance [4].

Lignocellulosic biomass usually primarily consists of cellulose, hemicellulose and lignin. However, the interaction between these three components in the plant cell wall renders the biomass rigid, complex and recalcitrant. Consequently, an effective biomass pre-treatment becomes extremely necessary to enable the enzyme access to biomass during saccharification. The sugars derived from enzymatic hydrolysis are referred to as second-generation (2G) and serve as building blocks in lignocellulose biorefineries to produce renewable fuel, biomaterials and other high value-added chemicals [5].

An ideal cellulolytic enzyme cocktail should contain a good amount of exoglucanases, endoglucanases, β-glucosidase and other auxiliary enzymes for efficient enzyme hydrolysis of the lignocellulosic biomass [4]. Cellulase enzymes are not only of great interest in biorefineries but also in the textile, beverage, cleaning product and pulp and paper manufacturing industries. Moreover, cellulases are being used in the degradation and smoothing of fabric fibers, aging of jeans, production of fruit juice, production of detergent and in winemaking, the animal feed industry and in the pulp and paper industries [6].

It is evident that cellulases are enzymes that exhibit different specificities for cellulose degradation, converting into soluble sugars that are then assimilated by microorganisms for fuels and specialty chemical products. The complete enzymatic degradation of cellulose requires the coordinated action of hydrolytic and oxidative enzymes. Hydrolytic enzymes include endoglucanases (responsible for the hydrolysis of the internal regions of the cellulose), exoglucanases (acting at the ends of the microcrystalline cellulose molecule, releasing cellobiose units) and β-glucosidases (which hydrolyze cellobiose and some glucose-soluble oligosaccharides). Oxidative enzymes, such as lytic polysaccharide monoxygenases (LPMOs), act in synergy with cellulases to potentiate cellulose degradation, being activated by enzymes that act as electron donors, such as cellobiose dehydrogenase (CDH) [7,8].

According to Florencio et al. [9], among the microorganisms capable of producing cellulolytic enzymes that efficiently degrade plant biomass, filamentous fungi stand out, especially Trichoderma reesei and Aspergillus niger strains. In this study, we chose to an Aspergillus species, as these fungi are well-known to produce a complete cellulase cocktail, including cellobiohydrolases, endoglucanases, β-glucosidases, β-xylosidases, endoxylanases, xyloglucanases and α-arabinofuranosidases. Aspergillus spp. are robust fungi that thrive on a variety of low-cost carbon and nitrogen sources, yielding high titers of plant biomass-degrading enzymes. It is evident from the literature that A. niger produces more effective hemicellulases and β-glucosidases than Trichoderma spp., with strong resistance to phenolic compounds from lignin and deactivators. Florecio et al. [9] studied the proteomic analysis of the A. niger strain and T. reesei and found that the former showed a secretome with a greater number of identified proteins and greater titers of cellulase enzymes.

The production of cellulase enzymes by Aspergillus sp. and Trichoderma sp. has been well-documented; however, there are only a few reports available on the production of cellulase by A. terreus. To the best of our knowledge, this study is first to report a comparison of enzyme production potentials in four different species viz. Aspergillus clavatus, A. versicolor, A. terreus and A. phoenicis growing on agro-industrial residues as carbon and nitrogen sources in a submerged fermentation process.

2. Materials and Methods

2.1. Microorganisms

Four different filamentous fungi viz. A. clavatus NRRL1, A. terreus NIH2624, A. versicolor CBS583.65 and A. phoenicis ATCC3157 were kindly provided by the Synthetic and Molecular Biology Laboratory at the Department of Biotechnology-EEL/USP, Lorena, São Paulo, Brazil for the production of cellulase.

2.2. Inoculum Preparation

Spores from all four Aspergillus strains were transferred to a pre-inoculum with 50 mL of growth medium (2% D-glucose, 2% malt extract, 1% peptone, pH: 5.5) in an orbital shaker (30 °C and 200 rpm) for 48 h. Subsequently, 1 mL of pre-inoculum was transferred to an Erlenmeyer flask containing 100 mL of the minimal medium (2 g/L of KH₂PO₄, 1.4 g/L of (NH₄)₂SO₄, 0.3 g/L of MgSO₄.7H₂O, 0.3 g/L of CaCl₂, 0.3 g/L of urea, 2 ml/L of trace elements (5 mg/L of FeSO₄.7H₂O, 1.56 mg/L of MnSO₄.H₂O, 1.4 mg/L of ZnSO₄.7H₂O, 2 mg/L of CoCl₂)) with 1% sodium carboxymethylcellulose (CMC) and a pH of 5.5, as described by Mandels and Weber [10], and kept at 30 °C for 96 h under agitation at 200 rpm. Samples were withdrawn after every 24 h with 5 min of rest for decanting, and 2 mL of the supernatant was removed for the cellulase assay.

2.3. Carbon and Nitrogen Sources in the Production Medium

For cellulase production, two different parameters were used: the carbon source and the nitrogen source (defined and non-defined). The carbon sources consisted of sugarcane bagasse, sugarcane straw, corn cob and pequi mesocarp, as these are agro-industrial residues and have different concentrations of cellulose, lignin and hemicellulose. The defined nitrogen sources were based on the medium described by Mandels and Weber [10], where each compound (urea, peptone and ammonium sulfate) was tested separately. The non-defined nitrogen sources consisted of rice straw, soybean bran and rice bran.

2.4. Alkaline Pre-Treatment of Carbon Sources

The native biomass samples were alkaline pre-treated using a 4% NaOH solution (w/v) in an autoclave at 121 °C for 40 min. Briefly, 10% lignocellulosic biomass (sugarcane bagasse, sugarcane straw, corn cob and pequi mesocarp) was added to the 4% NaOH solution. After the end of the pre-treatment, the biomass was filtered under a vacuum pump until most of the liquid was removed. Subsequently, the pre-treated material was washed with distilled water until the yellow color disappeared to ensure the removal of the remaining lignin present and to reach a neutral pH. After this procedure, the biomass was subjected to drying at room temperature and grinding in a knife mill to obtain a size smaller than a 20 mesh (0.841 mm). All samples were stored in a cold room at 4 °C.

2.5. Fermentation Assay under Optimized Conditions

Fermentation conditions, such as the pH (5.5), temperature (30 °C), agitation frequency (200 rpm) and culture medium (except for the carbon and nitrogen sources), were the same in all analyzed fermentations. To prepare the medium to select the best carbon source, the carboxymethyl cellulose (CMC) was replaced with 1% of a single carbon source (sugarcane bagasse or sugarcane straw or corn cob or pequi mesocarp) and the rest of the components followed the minimal medium protocol described above.

To select the best nitrogen source, synthetic nitrogen source substrates were added to the medium with a defined equimolar fraction as well as to the minimal medium already described, obtaining a total weight of 2.25 g/L for urea and ammonium sulfate and 2.7 g/L for peptone. The non-synthetic nitrogen sources were used at a concentration of 1.8 g/L of the total weight. Sugarcane bagasse was used as a major carbon source in this test due to its better performance, as shown in previous assays. For both carbon and nitrogen selection, fermentation was carried out using the same procedure used in the study of potential cellulolytic enzyme-producing fungi but, this time, varying only the carbon and nitrogen sources.

The A. terreus fungus was transferred to a pre-inoculum in 50 mL of medium in an orbital shaker (30 °C, 200 rpm) for 48 h. Subsequently, 2 mL were transferred to an Erlenmeyer flask containing 200 mL of minimal medium with 1% sugarcane bagasse and 1.8 g/L of rice bran, then kept at 30 °C for 96 h while agitated at 200 rpm. After fermentation, the contents of the Erlenmeyer flask were centrifuged, and the liquid fraction was used later for protein quantification and to determine enzyme concentration and activity.

2.6. Carboxymethyl Cellulase (CMCase) Activity

The CMC test indicates the presence of endoglucanases in the enzyme solution. In this step, 2 mL of the sample, obtained after the cultivation of fungi from the best carbon and nitrogen sources, was centrifuged at 10,000 rpm and 4 °C. Then, 1 mL of the supernatant was taken for analysis. The CMCase activity was determined by adding 0.5 mL of the crude enzyme extract into 0.5 mL of the 1% sodium carboxymethylcellulose solution in an acetate buffer (0.05 M, pH 5.0) followed by incubation in a water bath at 50 °C for 60 min. Periodically, the substrate-enzyme system was shaken to keep the cellulose in suspension. Then, 1 mL of a dinitrosalicylic acid (DNS) reagent was added, and the mixture was subjected to a water bath for 10 min at 100 °C. It was then cooled in an ice bath for 5 min, and 3.5 mL of distilled water was added. Finally, the absorbance was read in a spectrophotometer at 540 nm after calibrating the device with a blank composed of DNS reacted with the substrate without crude enzyme, which was replaced by distilled water. The enzyme activity tests of the enzyme solutions obtained from all four fungal strains were performed in triplicate.

Subsequently, CMCase activity tests were also performed on Greiner microplates to obtain enzymatic cocktail production profiles. In this case, 50 µL of the 0.5% CMC solution, 40 µL of a citrate buffer solution (pH 6) and 10 µL of the enzymatic extract were added in microtubes. The same was done for the blank, however, without the addition of the enzyme extract. Then the samples were placed in a dry bath at 50 °C for 30 min. After that, 100 µL of the DNS reagent was added to the samples and 10 µL of the enzymatic extract was added to the blank. The samples were submitted to a dry bath for 5 min at 100 °C and later added to Greiner microplates and analyzed in a plate reader at 540 nm.

The CMCase activity was determined in terms of glucose released (μmoles of reducing sugar produced per minute, using per ml of enzyme extract (U/mL) from the carboxymethylcellulose (substrate; CMC) at a pH of 6.0 and temperature of 50 °C) following the statistical analysis proposed by Triola et al. [11].

2.7. FPase Activity

FPase activity, i.e., filter paper activity, is determined based on the degradation of a strip of Whatman No. 1 filter paper (1.0 × 6.0 cm) by a mixture of exoglucanases and endoglucanases. In the tube containing the reaction assay, 1 mL of sodium citrate buffer solution (pH 4.8 to 50 mM), 0.5 mL of enzymatic extract and a strip of filter paper were added. In another tube, the reaction control was carried out by adding 1 mL of the same buffer solution and 0.5 mL of enzymatic extract; 1.5 mL of buffer solution and a strip of filter paper were added to the third tube, which was compared to the control. The samples were incubated in a water bath at 50 °C for 60 min, and the reaction was stopped with the addition of 3 mL of DNS reagent. The tubes were placed in boiling water for 5 min, and then 20 mL of distilled water was added for subsequent measurement of the absorbance in a spectrophotometer at 540 nm. The FPase activity was determined in terms of glucose released (μmoles of reducing sugar produced per minute, using per ml of enzyme extract (U/mL) from the Whatman filter paper (substrate) at a pH of 4.8 and temperature of 50 °C).

2.8. Biomass Characterization

Native and pre-treated sugarcane bagasse were characterized in terms of structural polymeric fractions (cellulose, hemicellulose, lignin), ash and extractives based on the analytical procedures established by the National Renewable Energy Laboratory (NREL), CO, USA [12].

2.9. Protein Quantification

Protein quantification was performed before and after concentrating the enzymatic extract using the Bio-Rad DC II kit, a colorimetric assay for protein concentration after detergent solubilization [13]. To prepare the blank, in duplicate, 20 µL of the minimal medium were added to microplates with rice bran. Then, 10 µL of reagent A (2% solution of Na₂CO₃ in 0.1 M of NaOH) and 80 µL of reagent B (0.5% solution of CuSO₄.5H₂O and 1% sodium citrate) were added. The plates were left to rest for 15 min, and then the absorbance was measured in the plate reader at 750 nm. As for the samples, in triplicate, the same procedure mentioned above was performed, replacing the minimal medium with 20 µL of the enzymatic extract.

2.10. Concentration of the Crude Enzyme Extract

The enzymatic extract, obtained from the optimized fermentation, was filtered through filter paper and concentrated up to 3 or 10 times by ultrafiltration in an Amicon stirred cell apparatus (Merck-Millipore, São Paulo, Brazil) with a 10 KDa cut-off membrane.

2.11. Xylanase Assay Using Beechwood and Birchwood Xylan

The xylanase activity of the enzymatic solution was performed using beechwood and birchwood as substrates. The substrates beechwood and birchwood were each added to microtubes followed by 50 µL of xylan 1%, 40 µL of citrate buffer (pH 4.8) and 10 µL of the enzyme extract. The same was done for the blank, replacing the enzymatic extract with the buffer. After that, the samples were incubated in a dry bath at 50 °C for 30 min. Then 100 µL of DNS reagent was added to the samples while 100 µL of DNS and 10 µL of enzyme extract were added to the blank. The samples were placed in a dry bath at 100 °C for 5 min and then diluted to a ratio of 1:4 (v/v) using distilled water before being transferred to Greiner microplates and analyzed in a plate reader at 540 nm. The activity value was calculated by considering 1 international unit (IU) equivalent to 1 μmol of xylose released per minute, per ml of enzyme (U/mL) using beechwood and birchwood as substrates at a pH of 4.8 and temperature of 50 °C.

2.12. Assay with Phosphoric Acid Swollen Cellulose (PASC)

For the PASC’s preparation, 1 g of Avicel was added first to 25 mL of phosphoric acid (85% w/v) in an ice bath and stirred for 1 h. Then 400 mL of distilled water was added and stirred for another 1 h. After that, the solution was filtered with filter paper and washed extensively with 2 L of distilled water. The solution was then neutralized with 700 mL of a 1% NaHCO₃ solution and washed again with 1.5 L of distilled water. The PASC was removed with the aid of a spatula and resuspended in a buffer until use.

Samples were made in triplicate by adding 250 µL of PASC (0.5%) to 200 µL of citrate buffer (0.5 M, pH 4.8) with 50 µL of enzyme extract. For the blanks, a sample without substrate and another without the enzymes were made as controls. The citrate buffer was used to replace the substrate and enzymes, respectively, in these two cases. The assay was performed in a shaker bath at 50 °C for 60 min at 1000 rpm. Afterwards, the samples (reaction mixtures) were heated to 100 °C for 5 min to stop the reaction. Thereafter, the samples were cooled and centrifuged at 12,500 rpm for 5 min. After centrifugation, 100 µL of the supernatant was mixed with 100 µL of DNS reagent and incubated in a water bath at 100 °C for 5 min. The absorbance was then read by a spectrophotometer using a plate reader at 540 nm.

For the analysis of lytic polysaccharide monoxygenases (LPMOs), the same procedure as described above was used, with the addition of 1 mM of ascorbic acid in the final concentration. Therefore, 250 µL of PASC, 150 µL of citrate buffer, 50 µL of enzyme extract and 50 µL of ascorbic acid were used for the samples. For these reactions, a reaction containing all the reagents except the enzyme extract was used as a blank.

2.13. Assays with Para-Nitrophenyl (pNP)-Labeled Substrates

The enzymatic extract was used for a multi-enzyme assay. For this, pNP-glucose, pNP-cellobiose, pNP-xylose and pNP-arabinofuranose were taken as substrates to test for the presence of β-glucosidase, cellobiohydrolase, β-xylosidase and arabinofuranosidase, respectively.

For this, 40 µL of the substrate (1 mM), 10 µL of the enzyme extract and 50 µL of the citrate buffer (0.5 M, pH 4.8) were added to the samples. As for the blank, which was made in duplicate, 40 µL of the substrate and 60 µL of the citrate buffer were used. The reaction was incubated at 50 °C for 30 min, then 100 µL of 1 M sodium carbonate was added and the solution was transferred to microplates for absorbance readings at 405 nm by a UV-visible spectrophotometer. After the 30 min incubation, there was a clear linearity in enzyme activity. The same method was used for each substrate and activity calculations were performed considering 1 international unit (IU) as equivalent to 1 nmol of pNP released per minute per ml of enzyme (U/mL) using the specific substrates at a pH of 4.8 and temperature of 50 °C.

3. Results and Discussion

Pre-treated sugarcane bagasse was used in this study as a primary carbon source for the four Aspergillus strains (i.e., A. clavatus NRRL1, A. phoenicis 275 ATCC3157, A. versicolor CBS583.65 and A. terreus NIH2624). The characterization of native sugarcane bagasse and alkaline pre-treated sugarcane bagasse is demonstrated in

Table 1. Chemical composition (% dry wt.) of native and alkaline pre-treated (4% NaOH, 10% bagasse concentration, 120 °C, 30 min) sugarcane bagasse (SCB).

SCB (native)
Sample	Total lignin	Glucan	Xylan	Arabinosyl	Ashes	Total
A	30.88	39.92	22.33	9.73	3.46	106.31
B	33.42	40.62	20.64	10.79	3.66	109.15
C	31.62	41.02	24.52	9.12	3.52	109.81
Media	31.62	40.62	22.33	9.73	3.52	109.15
Pre-Treated SCB
Sample	Total lignin	Glucan	Xylan	Arabinosyl	Ashes	Total
A	6.78	66.65	27.8	6.24	1.53	109
B	7.35	62.25	26.72	6.88	0.79	103.99
C	8.38	54.88	23.77	6.08	0.69	93.8
Media	7.58	60.77	25.78	6.16	0.74	101.03

, showing the amount of total lignin (soluble and insoluble), glucose, xylan, arabinose and ash.

The characterization data of biomass, i.e., native sugarcane bagasse and bagasse pre-treated with sodium hydroxide (NaOH), showed that the concentration of total lignin and arabinose was found to be less in pre-treated sugarcane bagasse compared to native sugarcane bagasse. The cellulosic fraction rationally increased in the substrate as lignin was removed from the pre-treated biomass. The higher concentration of glucose in the pre-treated bagasse is due to lignin removal. Similarly, the concentration of xylan was also found to be higher in pre-treated biomass but cannot be considered significant (variance less than 10%). Table 1 also confirms the presence of a higher composition of glucose in the pre-treated sugarcane bagasse because of the removal of lignin and a fraction of hemicellulose.

The results recorded in the present study showed a resemblance with observations recorded in previous studies [14,15,16] where an alkaline treatment with NaOH was found to be the most efficient method to delignify agricultural residues, increasing the internal surface area and decreasing the degree of polymerization and crystallinity, in addition to separating the structural bonds between lignin and carbohydrates by breaking the lignin structure. The disadvantage of this technique is that it also degrades a fraction of the hemicellulose [15]. Other types of pre-treatments, on the other hand, have little effect on reducing lignin content and do not show a profound impact on cell wall compactness, eventually reducing the accessibility of microorganisms to cellulose entities during cultivation.

The enzymatic activity (CMCase and FPase) of all four test Aspergillus strains (i.e., A. clavatus, A. versicolor, A. terreus and A. phoenicis) was evaluated and is shown in Figure 1.

The analysis demonstrated that A. terreus NIH2624 showed the maximum enzymatic activity (CMCase activity of 1.70 U/mL and FPase activity of 3.36 U/mL) compared to other strains (Figure 1). Therefore, A. terreus NIH2624 was selected for the further optimization of various carbon and nitrogen sources for cellulase production. The FPase activity (3.35 U/mL) reported in the present study was found to be significantly higher compared to previous studies, whereas CMCase activity (1.69 U/mL) was found to be lower. For example, Hui et al. [17] obtained an FPase value of 1.40 U/mL. Shahriarinour et al. [18] obtained an FPase of 0.69 U/mL and a CMCase of 7.41 U/mL for A. terreus, using peptone as a source of nitrogen in the minimum medium and using cellulose as carbon source. Sohail et al. [19] found lower values of FPase (0.15 U/mL) and CMCase (0.295 U/mL) for A. terreus activity with a mineral salt medium while employing solid-state fermentation. The CMCase value (1.32 U/mL) of this study is like the value found by Silva et al. [20] that used the mutant strain of T. reesei RP698.

After the selection of A. terreus NIH2624 as the fungus of study, four different carbon sources, i.e., corn cob, pequi mesocarp, sugarcane straw and sugarcane bagasse, were evaluated for cellulase production. The cellulase production profile of A. terreus NIH2624 growing on these carbon sources is presented in Figure 2.

Figure 2 shows the maximum CMCase (1.64 U/mL) and FPase (7.95 U/mL) production from sugarcane bagasse and sugarcane straw, respectively. Because sugarcane straw showed low CMCase production (0.29 U/mL), this biomass was not used further as a main carbon feedstock for cellulase production and, as the sugarcane bagasse showed a considerably higher production of cellulase, it was further selected as a primary carbon source in fermentation reactions.

The cellulase production by A. terreus NIH2624 can be compared with existing reports documented in the scientific literature. For instance, Corrêa [21] evaluated other A. terreus strains and found that A. terreus BLU24 produced 0.30 U/mL of CMCase and 0.50 U/mL of FPase, whereas Sohail et al. [19] found that A. terreus MS105 produced 0.726 IU/L/h of CMCase activity when grown on sugarcane bagasse. After the selection of A. terreus NIH2624 and the carbon source (sugarcane bagasse), the nitrogen sources were tested separately.

As shown in Figure 3, higher CMCase activity was observed when using rice bran (2.35 U/mL) and higher FPase activity was observed when using soybean bran (9.63 U/mL) while using pre-treated sugarcane bagasse as a carbon source. Considering that an enzyme cocktail must contain, mainly, endoglucanase and exoglucanase, rice bran was chosen as the best source of nitrogen.

The FPase values of this study are promising when compared to the literature. Latifian et al. [22] found FPase values of 1.163 U/mL for T. reesei QM9414 cellulases. In another study using rice bran and husk, a strain of Rhizopus oryzae showed an activity of 0.92 U/mL and 0.43 U/mL for CMCase and FPase, respectively, while the T. reesei strain showed an activity of 0.97 U/mL and 0.32 U/mL for CMCase and FPase, respectively [23]. Aureobasidium pullulans LB83 showed maximum cellulase activity (FPase of 2.27 U/mL; CMCase of 7.42 U/mL) on sugarcane bagasse and, while using soybean meal as a non-defined nitrogen source also showed maximum cellulase activity (FPase of 2.45 U/mL; CMCase of 6.86 U/mL) after 60 h [24].

To have a better understanding of the enzymatic cocktail profile, fermentation was carried out employing optimized conditions (i.e., growing A. terreus on a production medium containing alkaline pre-treated sugarcane bagasse as a carbon source and dilute acid pre-treated rice bran as an organic nitrogen source) (

Table 2. Plant cell wall-degrading enzymes produced by A. terrus NIH2624 growing on a production medium consisting of alkaline pre-treated sugarcane bagasse (carbon source) and rice bran (nitrogen source) and employing standard conditions.

Substrate	U/mL (µmol/(min·mL))	Enzymes
CMC	77.12 ± 2.8	Endoglucanase
Xylan from beechwood	1701.85 ± 98.8	Xylanases
Xylan from birchwood	1559.27 ± 31	Xylanases
Phosphoric acid swollen cellulose	21.03 ± 1.4	Endoglucanase, Cellobiohydrolase
Substrate	U/mL (nmol/(min·mL))	Enzymes
para-nitrophenyl-glucose	23.58 ± 0.1	β-glucosidase
para-nitrophenyl-cellobiose	8.46 ± 0.2	Cellobiohydrolase
para-nitrophenyl-xylose	4.45 ± 0.4	β-xylosidase
para-nitrophenyl-arabinofuranose	10.73 ± 0.4	Arabinofuranosidase

). The enzymatic extract obtained was concentrated from 0.353 µg/µL of protein to 0.575 µg/µL and used to evaluate the presence of different enzyme families through a substrate panel to analyze the auxiliary and plant cell wall-degrading activities.

It can be noted that the concentrated enzyme cocktail has large amounts of xylanases (1701.85 U/mL and 1559.26 U/mL for beechwood xylan and birchwood xylan, respectively). These xylanase values are much higher when compared to the literature. A secretomic analysis study of T. ressei RUT-C30 and A. niger A12, which aimed to produce cellulolytic enzymes and hydrolyze sugarcane bagasse, that was carried out by Florencio et al. [9] showed xylanase activities of 7.9 IU/mL and 7.8 IU/mL for T. reesei RUT-C30 and A. niger A12, respectively. In addition, compared to the study of Florencio et al. [9], the endoglucanase activities shown in the present work were higher than the ones Florencio et al. [9] found of 1.6 IU/mL and 0.6 IU/mL for T. reesei RUT-C30 and A. niger A12, respectively. Zhao et al. [25] found activities for endoglucanase (EG), cellobiohydrolase (CBH) and β-glucosidase of 430.45 mg/h/mL, 10.09 mg/h/mL and 0.25 µmol/min/mL, respectively. Compared to the values obtained in the present work (EG: 832.93 mg/h/mL; CBH: 227.09 mg/h/mL; β-glucosidase: 0.02µmol/min/mL), the activities of EG and CBH were 1.9 and 22.5 times higher, respectively. Thus, A. terreus is notable for its ability to produce cellulolytic enzymes.

The PASC test was also performed by adding ascorbic acid to analyze the presence of LPMOs, important enzymes that have revolutionized our understanding of the biological degradation of plant cell walls. The PASC test can indicate the presence of endoglucanase and cellobiohydrolase [26]. The addition of ascorbic acid to the test and the observation of increased activity strongly indicates the presence of LPMOs since LPMOs are activated in the presence of an electron donor, which is, in this case, ascorbic acid. Comparing the activity with and without ascorbic acid, we noted an increase of 17.84% in the activity (Figure 4), which suggests the presence of LPMOs in the obtained enzymatic cocktail.

The boosting effect of LPMOs acting on cellulose in synergy with cellulases is well known but little explored for A. terreus enzymes. The A. terreus genome harbors 12 genes that encode LPMOs, making it an important source for this type of enzyme [27]. The diversity of proteins produced by A. terreus was also evidenced in SDS-PAGE gels, where it was possible to observe band patterns of different sizes and intensities. The bands with the greatest intensities correspond to approximate sizes of 100 and 25 KDa, which are frequently sizes associated with cellobiohydrolases, beta-glucosidases (~100 KDa) [28], xylanases, endoglucanases and LPMOs (~25 kDa) [29].

4. Conclusions

A. terreus NIH2624 showed a higher production of cellulase and hemicellulose enzymes compared to the other tested fungi. When this microorganism was grown in the production medium containing sugarcane bagasse (as a carbon source) and rice bran (as a nitrogen source), it showed improved production of endoglucanases, exoglucanases and xylanases. Furthermore, the enzyme cocktail from A. terreus NIH2624 showed mainly xylanase activity, with a potential to complement cellulolytic enzyme cocktails of other known fungi, such as T. reesei and other Aspergillus species. A. terreus NIH2624 showed higher endoglucanase and CBH activities compared to the enzyme activity reported with other filamentous fungi. In addition to these enzymes, activities for cellobiohydrolases, β-xylosidases, arabinofuranosidases and LPMOs were also obtained. The use of this fungus shows promising results for holistic cellulolytic enzyme cocktail production in lignocellulose biorefineries, supporting the effective hydrolysis of biomasses to produce second-generation sugars.