Oxidases and hydrolase activities produced by monocultures developed by
Aspergillus sp. and Trametes versicolor solid-state monocultures produced high titers of xylanases and laccases activities (4617 ± 38 and 2759 ± 30 U/gsubstrate, respectively). Fungal biomass was quantified by estimating the ergosterol content of the mycelium, and by a simple material balance the corresponding residual substrate was obtained. Fungal growth and substrate consumption rates showed different behavior for these monocultures (μ = 0.03 and 0.11 h−1; rs = − 0.04 and − 0.0006 gsubstrate/h, respectively). In this case, xylanases production was directly linked to the growth, while laccases were produced during both growth and maintenance phases. Besides xylanases (42% of total Aspergillus enzyme), high titers of cellulases (15%), amylases (34%), and invertases (9%), as well as lignin and manganese peroxidases (10 and 24% of the total Trametes enzyme), were produced on the corresponding monocultures. When both fungi were used in a coculture mode, xylanases and laccases production decreased (around 85 and 70%), and the proportion of the hydrolases and oxidases changed. This suggested the need for most careful coculture design, in order to produce both enzymatic activities simultaneously even though the enzymatic extracts obtained by mono- or cocultures can be applied in several bioprocesses.
- Trametes versicolor
Laccases and xylanases are two of the most important lignocellulases that are employed in several industrial processes. Xylanases (EC 22.214.171.124) are responsible for degrading the xylan, a major polysaccharide in several cereals cell wall, to its constituent monomers. These enzymes are mostly used in textile, pulp and paper, bread making and feed industries, and the production of juice and fruit extracts [1, 2]. Moreover, laccases (EC 126.96.36.199) are multicopper enzymes that catalyze the oxidation of a wide variety of substrates such as mono-, di- and polyphenols, aminophenols, methoxyphenols, aromatic amines, and ascorbic acid. They have several industrial uses: they degrade toxic fungal metabolites and phenolic compounds and also are used in the design of biosensors, the detection of phenols in wastewaters, in the development of biofuel cells, during bleaching and delignification processes in the pulp and paper industry, and for the production of novel paper products . Together, xylanases and laccases can act for boosting bleaching process of several kinds of pulps, generating a cleaner process in which the use of the hazardous chemicals may be considerably reduced [4, 5]. These enzymes can be produced mainly by fungus, either in submerged or in solid-state cultures. The former is the most widely used as it provides a good control of operational parameters, high productivity and easy downstream processing, homogeneity of the culture and pH, and better oxygen supply and agitation speed management . However, solid-state culture could be better for producing this kind of enzymes as it represents the conditions that fungus finds in nature during the invasion of lignocellulosic material. Regarding this, our research group has advanced in optimizing the components of culture media in order to obtain the highest xylanases and laccases activities and yields by
On the other hand, the joint use of fungi which produce xylanase or laccase for developing a coculture system may be considered in order to obtain a mixed enzyme preparation, which has both xylanase and laccase activities, for being applied in biopulping and biobleaching processes. This kind of procedure will provide economic advantages because of the reduction in the overall cost of production .
Therefore, the objective of this work is to characterize solid-state monocultures with respect to growth, substrate consumption, and xylanases or laccases production and to test a coculture for producing both enzymes at the same solid-state fermentation.
2.2. Culture conditions for solid-state monocultures
For developing the corresponding monoculture, each microorganism was cultivated in solid-state fermentation (SSF) using 4 g of wheat bran and sugar cane bagasse (1:1 w/w) as support and substrate. For doing this, the support was moistened with water and autoclaved in 250 mL beakers at 121°C for 20 min.
For laccases production monoculture, an appropriate quantity (around 4 mL) of Kirk medium was added to maintain the desired moisture level of the support (50%) for several experimental units. Each beaker was then inoculated with 4 mycelial plugs (50 mm diameter) taken from the periphery if a
Kirk basal medium composition (in g/L) was as follows: sodium tartrate, 0.275; MgSO4·7H2O, 0.55; K2HPO4, 2.2; CaCl2·2H2O, 0.145; (NH4)2SO4, 0.44; Glucose, 8.2; CuSO4·5H2O, 0.28; trace elements, 11 mL (in g/L: MnSO4·H2O, 0.5; NaCl, 1; FeSO4·7H2O, 0.1; CoCl2·6H2O, 0.185; ZnSO4·7H2O, 0.11; Na2MO4·2H2O, 0.011; H3BO3, 0.011). The pH of the medium was adjusted at 5.0 using 1 M HCl before sterilization.
Basal medium composition employed for this monoculture (in g/L) was as follows: K2HPO4, 2; KH2PO4, 2; (NH4)2SO4, 5.
2.3. Cocultures developing
For developing the cocultures in solid state, the same support employed for monocultures was used. This was moisturized and sterilized as indicated before. In this case, the support was moisturized with Kirk basal medium, and both inoculums (
2.4. Extraction and storage of crude enzymes
After incubation, 80 mL of 100 mM sodium acetate buffer pH 5 was added to each experimental unit, homogenizing with a Multi Braun® mixer and a posterior constant agitation in an ice bath. Afterwards, the enzymatic extracts were recovered by centrifugation at 4000 rpm and stored at 4°C until analyzed. The solids obtained after centrifugation were used for estimate biomass and residual substrate content, as explained below.
2.5. Enzymatic activities quantification
Laccase (Lac) activity was determined by measuring the increase in A470 (ε470 nm = 26,600 M−1 cm−1) due to the oxidation of 10 mM guaiacol in 0.1 M Na-acetate buffer (pH 5.0) after incubation with the crude extract at 25°C for 10 min. One unit of laccase activity (U) was defined as the amount of enzyme required to oxidize 1 μmol of guaiacol per minute of reaction .
Xylanolytic (Xyl) activity was assayed by incubating at 45°C for 20 min using the crude enzyme with 1% (w/v) xylan dissolved in 100 mM acetate buffer pH 5.0, and the release of reducing sugars as xylose was monitored at 575 nm after stopping the reaction by the addition of DNS. The optical density obtained was compared with a 1 g/L xylose standard curve. One unit of xylanolytic activity was defined as the enzyme necessary for the release of 1 μmol of xylose under the described conditions .
Lignin peroxidase (LiP) activity was estimated by incubating at 25 °C for 20 min the crude enzyme with 4 mM veratryl alcohol diluted in 100 mM tartrate buffer pH 3.5 and 0.4 mM H2O2. The formation of veratraldehyde was monitored at 310 nm (ε310 nm = 9.3331 mM−1 cm−1). One unit of lignin peroxidase activity was defined as the enzyme required for oxidize 1 μmol of veratryl alcohol per minute of reaction .
Manganese peroxidase (MnP) activity was assayed by Incubating at 25 °C for 5 min the crude enzyme with 0.1 mM MnSO4, 1 mM H2O2, and 1 mM 2,6-dimethoxyphenol (DMP) as substrate diluted in 0.1 M tartrate buffer pH 4.5, measuring the increase in A469 nm (ε469 nm = 27,500 mM−1 cm−1). One unit of manganese peroxidase activity (U) was defined as the amount of enzyme required to oxidize one μmol of DMP per minute of reaction .
Glucoamylase (Glcamyl) activity was estimated by incubating at 60 °C for 15 min the crude enzyme with 1% (w/v) starch dissolved in 0.15 M sodium chloride buffer pH 5.0. The released reducing sugars as glucose were monitored at 575 nm after stopping the reaction by the addition of DNS. The optical density obtained was compared with a 1 g/L glucose standard curve. One unit of glucoamylase activity was defined as the enzyme necessary for the release of 1 μmol of glucose under the reaction conditions .
α-Amylase (α-amyl) activity was determined by incubating at 37 °C for 20 min the crude enzyme with 1% (w/v) starch dissolved in 0.15 M sodium chloride buffer pH 5. The photometric disappearance of starch was quantified after stopping the reaction by the addition of an iodine (I2/IK) mother solution, and the resultant optical density at 580 nm was registered. One unit of α-amylase activity was defined as the enzyme necessary for hydrolyze 0.1 mg of starch .
Invertase (Inv) activity was determined by incubating at 30 °C for 30 min the crude enzyme with 0.1 M sucrose dissolved in 0.15 M acetate buffer pH 5.5. The reducing sugars as fructose were quantified after stopping the reaction with DNS. The optical density obtained was compared with a 1 g/L glucose standard curve. One unit of invertase activity was described as the enzyme necessary for the release of 1 μmol of reducing sugars per minute of reaction .
Carboxymethyl cellulase (CMCase) activity was quantified By incubating at 50 °C for 5 min the crude enzyme with (1% w/v) carboxymethyl cellulase low viscosity in 50 mM citrate buffer (pH 5.0). The reducing sugars as glucose were quantified after stopping the reaction with DNS. The optical density obtained was compared with a 1 g/L glucose standard curve. One unit of CMCase activity was described as the enzyme necessary for the release of 1 μmol of reducing sugars per minute of reaction .
The total cellulase activity (filter paper activity, FPAse) was assayed by using a 0.5 × 6 cm string of filter paper as the substrate. This was incubated using 100 mM acetate buffer (pH 5.0) and the crude enzyme for 5 min at 45°C, stopping the reaction with DNS. The optical density obtained was compared with a 1 g/L glucose standard curve. One unit of FPAse activity was defined as the amount of enzyme used for the release of 1 μmol of glucose under the assayed conditions .
2.6. Biomass quantification
The quantification of fungal biomass was made after quantifying ergosterol content of the biomass in each sample. For doing this, the solid content of each experimental unit was resuspended in 10 mL of water by vigorous agitation. From this homogeneous solid suspension, 1 mL was withdrawn and filtered, to recover the solids. To this, 3 mL of an alcoholic basic solution (25% w/v of KOH dissolved in methanol) was added, boiling the resultant mixture for 5 h. Afterwards, 1 mL of distilled water and 5 mL of
where 290 and 518 correspond to the molar extinction coefficient (M−1 cm−1) of crystalline ergosterol and 24(28) DHE, respectively. A 10 g/L mycelium (
2.7. Substrate quantification
The solids obtained from each culture were dried at 60°C for 12 h. The biomass content (estimated as explained before) was subtracted to the corresponding dry weight in order to obtain the real substrate content of each sample.
All the experiments were performed in duplicate, and the results are expressed as the mean of these repetitions and the corresponding standard deviation.
3. Results and discussion
3.1. Behavior of solid-state fermentation monocultures during enzymes production
On previous work, we optimized the culture media components in order to obtain high xylanase and laccase productions on solid-state fermentation, using
In this regard, it is worth mentioning that biomass quantification on solid-state cultures is a complicated work. Fungal growth is not easy to quantify because fungus grows as hyphal filaments that cannot be quantified by the usual enumeration techniques, and specifically on cultures in which an insoluble material is used as the only carbon source, complete recovery of fungal biomass from the substrate is very difficult, because the fungal hyphae tend to penetrate into and binds tightly to the solid substrate particles . It is important therefore to have reliable and convenient methods for measuring fungal growth. For this reason, we used the ergosterol content methodology  for quantifying biomass evolution along the culture, and employed a simple mass balance for knowing the corresponding residual substrate. Therefore, this is one of the main contributions of the present work.
With respect to monocultures developed with
With respect to monocultures developed with
Several studies developed on liquid or submerged fermentation have reported that laccases production has a secondary metabolite behavior; it means the activity is produced mainly during the secondary metabolism . On solid-state culture, this relationship is not well known due to the difficulty of quantifying accurately the total biomass grown in the substrate along the culture. However, other studies in which fungal biomass has been quantified by indirect methods, like that used in the present work, have shown that laccases production is growth related, as happened for laccases produced by
The titers obtained on each monoculture for xylanases and laccase activities were high. These results show that solid-state culture is a good alternative for producing high oxidative or hydrolytic activities, in order to employ them for several industrial bioprocesses. At this respect, the results obtained in the present work would represent one basis for developing this process on a full- scale. Thereby, for further characterization of the monocultures, in the next section the production profiles of other oxidative or hydrolytic activities obtained were analyzed.
3.2. Enzymatic profiles of crude enzymes
Total enzyme production obtained along
With respect to oxidases, during the first 72 h of monocultures developed with
|Enzymatic activities (U/gs)|
|Laccases||2759 ± 30||ND|
|LiP||410 ± 10||ND|
|MnP||996 ± 2||ND|
|Xylanases||ND||4617 ± 38|
|FPAses||ND||746 ± 7|
|Carboximetil‐celulases||ND||829 ± 34|
|Glucoamylases||ND||1762 ± 4|
|α-amylases||ND||1898 ± 6|
|Invertases||ND||1023 ± 73|
Taking into account that enzymatic extracts obtained on solid-state cultures by
3.3. Cocultures for joint production of lignocellulases
Solid-state cultures showed high productions of either hydrolase or oxidase activities in the corresponding monocultures. Previous work of this group has shown that bleach boosting of kraft  and jonote  pulp can be improved with the employment of the combined action of xylanases and laccases, produced by solid-state cultures as those described in the present work. So, a greater enzyme production would be desirable in order to have a more efficient bioprocess. At this respect, it has been suggested that the cocultivation of microbes in fermentation can increase the quantity of the desirable components on a cellulose complex . On the other side, some reports have shown that laccases or xylanases production can increase in a coculture mode, as happens with
With respect to hydrolytic activities proportion, it can be seen that while on monocultures xylanase activity was the predominant hydrolytic activity, glucoamylases were in greater proportion for cocultures developed at 30°C, and α-amylases highlighted on cocultures developed at 37°C (Figure 3 A).
Comparing both cocultures, one can see that the highest hydrolytic activities were obtained on that developed at 37°C, which means that temperature could be affecting seriously the
Laccase activities decreased on both cocultures with respect to those values obtained on monocultures. This behavior could be considered unusual, because in general the coculture strategy is used in order to increase these activities. In fact, the addition of soil microorganisms to white rot fungi cultures has increased laccase and other oxidases production . In this case, the diminution on laccase activity can be explained from different points of view. First of all, in the present study both fungi were inoculated at the same time to the support. The decrease in oxidases and hydrolases production could be provoked by problems in fungal growth, considering the differences in specific growth rate of each fungal species. This is because mineral medium used in these cocultures contained glucose, and it has been reported as laccase inductor for
On the other hand, we must remember that enzymes are produced at different rates along the cultures, as described in Section 3.2. In this case, activities analyzed for this experimental phase are those obtained after 10 days, but the enzyme evolution along both cocultures was not registered. This is a great limitation for obtaining valid conclusions. It could be possible that one of these enzymes has been increased at any time during the fermentations, but as we did not quantify the enzyme evolution, we could not know if the reported activity was the highest obtained for the corresponding enzyme in this experiment. So, we can only analyze the enzyme activities at the end of the coculture, and these could not be the highest activities obtained in this case.
Finally, even when some hydrolytic or oxidative enzymes decreased on coculture mode, these extracts can be employed on several processes. Agricultural by-products typically vary in their chemical composition and nutritional value, and sometimes are also higher in low-quality fiber, so a specific enzyme complex is required to break it down in order to be used in ruminant feed. Besides, their nutritional value could be increased by biodegradation methods of fiber in the rumen, through efficient delignification . Therefore, the filtrates obtained on both cocultures could be a good alternative for being employed for using agricultural by-products for ruminant feed.
The indirect technique used for the quantification of fungal biomass content was useful, meaning a great contribution for analyzing solid-state cultures. From this, it could be seen that both fungi have different behaviors along the culture, in which
LSV acknowledges CONACyT for the scholarship 551418. The authors acknowledge Instituto de Ingeniería, from Universidad Nacional Autónoma de México, Tecnológico de Estudios Superiores de Ecatepec and PRODEP, for the economic funding.
Jin N, Ma S, Liu Y, Yi X, He R, Xu H, Qiao DR, Cao Y. Thermophilic xylanase production by Aspergillus nigerin solid state fermentation using wheat straw and corn cob. African Journal of Microbiology Research 2012;6(10):2387–2390. doi: 10.5897/AJMR11.1433
Mohamed SA, Al-Malki AL, Khan JA, Kabli SA, Al-Garni SM. Solid state production of polygalacturonase and xylanase by Trichodermaspecies using cantaloupe and watermelon rinds. Journal of Microbiology. 2013;51(5):605–611. doi: 10.1007/s12275-013-3016-x
Fernández-Fernández M, Sanromán Má, Moldes D. Recent developments and applications of immobilized laccase. Biotechnology Advances. 2013;31(8):1808–1825. doi: 10.1016/j.biotechadv.2012.02.013
Record E, Asther M, Sigoillot C, Pages S, Punt PJ, Delattre M, Haon M, Van den Hondel CA, Sigoillot JC, Lesage-Meessen L, Asther M. Overproduction of the Aspergillus nigerferuloyl esterase for pulp bleaching application. Applied Microbiology and Biotechnology. 2003;62(4):349–355. doi: 10.1007/s00253-003-1325-4
Kapoor M, Kapoor RK, Kuhad RC. Differential and synergistic effects of xylanase and laccase mediator system (LMS) in bleaching of soda and waste pulps. Journal of Applied Microbiology. 2007;103(2):305–317. doi: 10.1111/j.1365-2672.2006.03251.x
Xu P, Ding ZY, Qian Z, Zhao CX, Zhang KC. Improved production of mycelial biomass and ganoderic acid by submerged culture of Ganoderma lucidumSB97 using complex media. Enzyme and Microbial Technology. 2008;42(4):325–331. doi: 10.1016/j.enzmictec.2007.10.016
Domínguez-Morales D. Optimización de la producción de xilanasas y lacasas fúngicas mediante el empleo de la Metodología de Superficie de Respuesta [Thesis]. Tecnológico de Estudios Superiores de Ecatepec; 2013.
Licona-Soto S.R. Bioproceso integrado para el tratamiento biológico de la pulpa de jonote [thesis]. Tecnológico de Estudios Superiores de Ecatepec; 2014.
García-Rivero M, Membrillo-Venegas I, Vigueras-Carmona SE, Zafra-Jiménez G, Zárate-Segura PB, Martínez-Trujillo MA. Enzymatic pretreatment to enhance chemical bleaching of a kraft pulp. Revista Mexicana de Ingeniería Química. 2015;14(2):335–345.
Dwivedi P, Vivekanand V, Pareek N, Sharma A, Singh RP. Co-cultivation of mutant Penicillium oxalicumSAU E-3.510 and Pleurotus ostreatusfor simultaneous biosynthesis of xylanase and laccase under solid-state fermentation. New biotechnology. 2011;28(6):616–626. doi: 10.1016/j.nbt.2011.05.006
Bertrand B, Martínez-Morales F, Tinoco-Valencia R, Rojas S, Acosta-Urdapilleta L, Trejo-Hernández MR. Biochemical and molecular characterization of laccase isoforms produced by the white-rot fungus Trametes versicolorunder submerged culture conditions. Journal of Molecular Catalysis B: Enzymatic. 2015;122:339–347.
Membrillo Venegas I, Fuentes-Hernández J, García-Rivero M, Martínez-Trujillo A. Characteristics of Aspergillus nigerxylanases produced on rice husk and wheat bran in submerged culture and solid?state fermentation for an applicability proposal. International Journal of Food Science and Technology. 2013;48(9):1798–1807. doi: 10.1111/ijfs.12153
Jung H, Xu F, Li K. Purification and characterization of laccase from wood-degrading fungus Trichophyton rubrumLKY-7. Enzyme and Microbial Technology. 2002;30(2):161–168. doi: 10.1016/S0141–0229(01)00485-9
Palma C, Martınez AT, Lema JM, Martınez MJ. Different fungal manganese-oxidizing peroxidases: a comparison between Bjerkanderasp. and Phanerochaete chrysosporium. Journal of Biotechnology. 2000;77(2):235–245. doi: 10.1016/S0168-1656(99)00218-7
Matute L, Bertsch A, Díaz I. Evaluación de la actividad amilolítica de Aspergillus nigerANM-1 en fermentaciones en estado sólido y sumergido para la obtención y caracterización de aditivos enzimáticos. Revista de la Facultad de Agronomía (UCV). 2011;38(1): 9–17.
Romero-Gómez SJ, Augur C, Viniegra-González G. Invertase production by Aspergillus nigerin submerged and solid-state fermentation. Biotechnology Letters. 2000;22(15):1255–1258. doi: 10.1023/A:1005659217932
Miller GL, Blum R, Glennon WE, Burton AL. Measurement of carboxymethylcellulase activity. Analytical Biochemistry. 1960;1(2):127–132. doi: 10.1016/0003-2697(60)90004-X
Ponce-Noyola T, De la Torre M. Isolation of a high-specific-growth-rate mutant of Cellulomonas flavigenaon sugar cane bagasse. Applied Microbiology and Biotechnology. 1995;42(5):709–712. doi: 10.1007/BF00171949
Bhosle SR, Sandhya G, Sonawane HB, Vaidya JG. Ergosterol content of several wood decaying fungi using a modified method. International Journal of Pharmacy and Life Sciences. International Journal of Pharmacy and Life Science (IJPLS). 2011;2(7):916–918.
Abd-Aziz S, Hung GS, Hassan MA, Karim MI, Samat N. Indirect method for quantification of cell biomass during solid-state fermentation of palm kernel cake based on protein content. Asian Journal of Scientific Research 2008;1(4):385–393.
Brijwani K, Rigdon A, Vadlani PV. Fungal laccases: production, function, and applications in food processing. Enzyme Research. 2010 Sep 21;2010. http://dx.doi.org/10.4061/2010/149748
Niladevi KN, Sukumaran RK, Prema P. Utilization of rice straw for laccase production by Streptomyces psammoticusin solid-state fermentation. Journal of Industrial Microbiology and Biotechnology. 2007;34(10):665–674. doi: 10.1007/s10295-007-0239-z
Hasunuma T, Okazaki F, Okai N, Hara KY, Ishii J, Kondo A. A review of enzymes and microbes for lignocellulosic biorefinery and the possibility of their application to consolidated bioprocessing technology. Bioresource Technology. 2013;135:513–522. doi: 10.1016/j.biortech.2012.10.047
Veana F, Martínez-Hernández JL, Aguilar CN, Rodríguez-Herrera R, Michelena G. Utilization of molasses and sugar cane bagasse for production of fungal invertase in solid state fermentation using Aspergillus nigerGH1. Brazilian Journal of Microbiology. 2014;45(2):373–377. http://dx.doi.org/10.1590/S1517-83822014000200002
Mostafa FA, Ahmed SA, Helmy WA. Enzymatic saccharification of pretreated lemon peels for fermentable sugar production. Journal of Applied Sciences Research. 2013;9(3):2301–2310.
Zeng X, Cai Y, Liao X, Zeng X, Li W, Zhang D. Decolorization of synthetic dyes by crude laccase from a newly isolated Trametes trogiistrain cultivated on solid agro-industrial residue. Journal of Hazardous Materials. 2011;187(1):517–525. doi: 10.1016/j.jhazmat.2011.01.068
Kaushal R, Sharma N, Tandon D. Cellulase and xylanase production by co-culture of Aspergillus nigerand Fusarium oxysporumutilizing forest waste. Turkish Journal of Biochemistry/Turk Biyokimya Dergisi. 2012;37(1).
Flores C, Casasanero R, Trejo-Hernández MR, Galindo E, Serrano-Carreón L. Production of laccases by Pleurotus ostreatusin submerged fermentation in co?culture with Trichoderma viride. Journal of Applied Microbiology. 2010;108(3):810–817. doi: 10.1111/j.1365-2672.2009.04493.x
Alegre AC, Polizeli MD, Terenzi HF, Jorge JA, Guimarães LH. Production of thermostable invertases by Aspergillus caespitosusunder submerged or solid state fermentation using agroindustrial residues as carbon source. Brazilian Journal of Microbiology. 2009;40(3):612–622. http://dx.doi.org/10.1590/S1517-83822009000300025
Chan Cupul W, Heredia Abarca G, Martínez Carrera D, Rodríguez Vázquez R. Enhancement of ligninolytic enzyme activities in a Trametes maxima-Paecilomyces carneusco-culture: key factors revealed after screening using a Plackett-Burman experimental design. Electronic Journal of Biotechnology. 2014;17(3):114–121. http://dx.doi.org/10.1016/j.ejbt.2014.04.007
Piscitelli A, Giardina P, Lettera V, Pezzella C, Sannia G, Faraco V. Induction and transcriptional regulation of laccases in fungi. Current Genomics. 2011;12(2):104–112. doi: http://dx.doi.org/10.2174/138920211795564331
Graminha EB, Goncalves AZ, Pirota RD, Balsalobre MA, Da Silva R, Gomes E. Enzyme production by solid-state fermentation: application to animal nutrition. Animal Feed Science and Technology. 2008;144(1):1–22. doi: 10.1016/j.anifeedsci.2007.09.029