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Production and purification of amylolytic enzymes for saccharification of microalgal biomass
Accepted Manuscript Production and purification of amylolytic enzymes for saccharification of microalgal biomass Éllen Francine Rodrigues, Aline Matuella Moreira Ficanha, Rogério Marcos Dallago, Helen Treichel, Christian Oliveira Reinehr, Tainara Paula Machado, Greice Borges Nunes, Luciane Maria Colla PII: DOI: Reference:
S0960-8524(16)31562-0 http://dx.doi.org/10.1016/j.biortech.2016.11.047 BITE 17293
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Bioresource Technology
Received Date: Revised Date: Accepted Date:
2 September 2016 10 November 2016 12 November 2016
Please cite this article as: Francine Rodrigues, E., Matuella Moreira Ficanha, A., Marcos Dallago, R., Treichel, H., Oliveira Reinehr, C., Paula Machado, T., Borges Nunes, G., Maria Colla, L., Production and purification of amylolytic enzymes for saccharification of microalgal biomass, Bioresource Technology (2016), doi: http:// dx.doi.org/10.1016/j.biortech.2016.11.047
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Production and purification of amylolytic enzymes for saccharification of microalgal biomass
Éllen Francine Rodriguesa, Aline Matuella Moreira Ficanhab, Rogério Marcos Dallago b, Helen Treichelc, Christian Oliveira Reinehra, Tainara Paula Machado a, Greice Borges Nunesa, Luciane Maria Collaa*
a
University of Passo Fundo, Campus I, km 171, BR 285, P.O. Box 611, CEP 99001-
970, Passo Fundo, RS, Brazil b
Regional and Integrate University of Upper Uruguay and Missions, URI - Erechim,
Av. Sete de Setembro 1621, Erechim, RS 99700-000, Brazil c
Federal University of Fronteira Sul – Campus de Erechim, ERS 135, km 72, n° 200,
99700-970, Erechim, RS, Brazil * Corresponding author. E-mail: [email protected], [email protected]
Keywords: Fungal amylases, biofuels, immobilization, microfiltration
ABSTRACT The aim of this study was the production of amylolytic enzymes by solid state or submerged fermentations (SSF or SF, respectively), followed by purification using chemical process or microfiltration and immobilization of purified enzymes in a polyurethane support. The free and immobilized enzymes obtained were used to evaluate enzymatic hydrolysis of the polysaccharides of Spirulina. Microfiltration of the crude extracts resulted in an increase in their specific activity and thermal stability at 40 °C and 50 °C for 24 h, as compared to extracts obtained by SSF and SF. Immobilization of polyurethane purified enzyme produced yields of 332% and 205% for the enzymes obtained by SSF and SF, respectively. Free or immobilized enzymes favor the generation of fermentable sugar, being the application of the purified and immobilized enzymes in the hydrolysis of microalgal polysaccharides considered a promising alternative towards development of the bioethanol production process from microalgal biomass.
KEYWORDS: Fungal amylases, biofuels, immobilization, microfiltration
1. Introduction The demand for enzymes, especially amylases, is constantly growing owing to the variety of industrial applications of these hydrolases: hydrolyzed starch processing in the form of glucose and fructose syrups, alcoholic beverage production, textiles, bioethanol production, and others (Joshi and Satyanarayana, 2015; Sherif et al., 2013). This market has been stimulated recently by the production of bioethanol from starch and microalgae. It is estimated that the production and application of enzymes in different markets would increase to 17.5 billion dollars in 2024. The increasing demand for bioethanol and biodiesel in emerging economies including India, Brazil, and Thailand will augment industry growth over the forecast period (Grand View Research, 2016). The need to modify the current energy matrix, based on non-renewable resources like oil, has resulted in increased demand for global production of biofuels, and led to the diversification of biomass used for this purpose (Alemán-Nava, 2014). The use of non-conventional biomass, such as the microalgal biomass, has emerged as a promising alternative (Bahadar and Khan, 2013). The use of microalgae have been reported by many authors, who proposed that algal biomass, especially the carbohydrate- and lipid-rich species (Ruiz et al., 2013), can be used for chemical or biotechnological production of bioethanol and biodiesel (Zhu et al., 2014; Ghasemi et al., 2012). In this context, the use of amylolytic enzymes in bioethanol production appears as a promising technology and the study of enzymes that allow simultaneous hydrolysis and fermentation processes with viable yields are very relevant (Aikawa et al., 2013; Fernandes et al., 2012; Choi et al., 2010).
Among the microalgae used to obtain bioethanol, Spirulina platensis is well known in the scientific community because of its ability to accumulate proteins and other biocompounds. It has been reported to accumulate carbohydrates (up to 70%), when cultured in a nutrient-depleted medium containing added organic sources of carbon (Salla et al., 2016). The production of amylolytic enzymes can be performed using filamentous fungi by bioprocesses, or using by-products derived from the agricultural industry, such as wheat bran, corn bran, soybean meal, sugar cane bagasse, among others (RamosSanchez et al., 2015; Oliveira et al., 2013). Among the filamentous fungi, Aspergillus is the most widely used genus for the production of amylolytic enzymes (Venkat, 2007). Several different techniques have been developed, searching for higher yields, more specific and stable enzymes, making more industrial applications possible (Soccol and Vanderbergue, 2003). In the last decade solid-state fermentation (SSF) has received increased attention from researchers as a bio-based process. This technology allows for the valorization of wastes for the production of valuable commodities such as enzymes (El-Bakry et al., 2015). Solid state fermentation (SSF) is characterized for the growth of the microorganism in the absence of free water and this kind of technique can be used for production of many bio-products. Although important drawbacks are described concerning the SSF scale up largely due to heat transfer, sterilization costs and culture heterogeneity (Casciatori et al., 2016), SSF has several advantages over submerged fermentation (SmF) because of the low capital investment, the simple technique, low energy requirement, end-product inhibition, low waste water output and better product recovery (Pandey at al., 2000).
The development of techniques for concentration, purification, and immobilization of enzymes is essential for obtaining enzymes with high catalytic efficiency. Pure and stable enzymes with high specificity for substrates can be used in industrial processes in a similar way to chemical catalysts, and their recovery and reuse would facilitate a better cost–benefit ratio (Cunha et al., 2014). The purification or concentration of enzymes by membrane separation methods has been proposed for improving production processes owing to their advantages: no addition of chemical products, reduction in process energy consumption, and ease of scale up (Saxena et al., 2009). Downstream of purification/concentration, immobilization of the enzymes is a lucrative option to bypass the limitations of free enzymes by imparting the following properties: greater enzyme stability, operational ease of separation, and possibility of reuse. Immobilization may be achieved by encapsulation, by physical or chemical interactions with a particular media. In the process, following factors should be considered: economic, operational, experimental conditions employed in synthesis, ease of production, duration of the process, and the amount of enzyme incorporated into the support (Ficanha et al., 2015). In this context, we aimed at optimizing the production of amylolytic enzymes from Aspergillus niger using different fermentation processes, followed by their purification, immobilization, and application in the hydrolysis of Spirulina platensis biomass.
2. Material and Methods 2.1. Microorganisms, inoculum preparation, and enzyme production bioprocesses
The enzymes were produced using Aspergillus niger. The isolation and identification of this microorganism were well described in a previous work of our research group. Through the comparison of the partial area of the gene β-Tubulin of the isolated fragments or fungus with other sequences deposited in Genbank was obtained 98.2% of similarity of isolated fungus with Aspergillus niger, enough similarity to consider as the same species for fungus Treichel et al. (2016). The inoculum preparation was carried out by inoculating the microorganism in 1-L flasks, containing 100 mL of PDA medium (Potato Dextrose Agar), followed by incubation at 30 °C for 7 d. Wheat bran was used as a substrate for the bioprocess. In solid-state fermentation (SSF), moisture of the bran was adjusted to 60% with a sterile nutrient solution. For submerged fermentation (SF), the medium was prepared from the baking of 10% (w/v), at 100 °C for 30 min. After the cooking, filtration was carried out through a cotton filter to remove solids. In both culture media, 10% (v/v) of solution was added, which was composed of the following ingredients: 2 g.L-1 KH2PO4, 1 g.L-1 MgSO4, 0.225 g CaCl2, and 10 mL.L1
of trace solution (0.63 mg.L-1 FeSO4.7H2O, 0.01 mg.L-1 MnSO4 and 0.62 mg.L-1
ZnSO4 and distilled water to a volume of 1 L). The medium were sterilized at 121 °C, 101 kPa for 20 min, and the inoculation was performed after cooling the medium to room temperature. Solid-state fermentation experiments were conducted in 1-L beakers, with 100 g of bran inoculated with 107 spores/gmedium, and subsequent incubation at 30 °C for 144 h. The submerged cultivations were carried out in a 5-L bioreactor (Autoclavable
Fermentor Bioreactor SL - 135, SOLAB), containing 3 L of growth medium, inoculated with 4.106 spores/mLmedium, and incubated at 30 °C with shaking at 120 rpm for 72 h. The pH of the medium was around 5.8 and 6.0 at the beginning of fermentation processes in solid-state and submerged cultivation, respectively. 2.2. Purification of enzymes After the solid-state fermentation, the enzymes were extracted using a mediabuffer proportion of 1:17 (g fermented medium:mL phosphate buffer pH 7.0) in an orbital shaker at 35 °C for 60 min and 100 rpm (PARIS et al., 2012). The suspension was filtrated through cotton and centrifuged (Cientec Model CT - 5000) at 910 g for 20 min aiming at removing particulate solids. In this obtained crude extract, ammonium sulfate was added slowly and under stirring on ice bath, until 80% saturation (56.5 g ammonium sulfate in 100 mL of crude extract). After addition of salt, the mixture was kept standing overnight at 4 °C, and centrifuged at 1310 g for 30 min. The pellet was resuspended in distilled water, and the obtained enzyme solution was used in the analytical determinations following the purification process. The extract was called purified extract with ammonium sulfate. The medium of the submerged fermentation culture was filtered through cotton to separate the hyphae, followed by subsequent centrifugation at 910 g for 20 min to remove the particulate solids to yield the crude extract of submerged fermentation. The crude enzyme extract of the submerged fermentation and the purified extract with ammonium sulfate from solid-state fermentation were subjected to microfiltration to separate enzymes of different molecular weights from other low molecular weight compounds from the fermentation. The microfiltration system consisted of an acrylic module, a peristaltic pump, a ball valve to control the flow, pressure gauge, and hose
connector. System operation was at 200 kPa, using a flat microfiltration membrane consisting of polyvinylidene difluoride with a pore diameter of 0.4 µm (GE Water and Process Technologies). The enzymatic activity and the specific enzymatic activity of the purified extracts were determined according to the method described below (2.2.1.). 2.2.1. Determination of enzymatic activity of the extracts The amylolytic activity of the enzyme extracts was determined using starch as a substrate for the saccharification method, which is based on production of reducing sugars (Miller, 1959). One enzyme activity unit (U) was defined as the amount of enzyme capable of releasing 1 µmol of glucose per minute under the conditions of the proposed method. The quantification of proteins in the extracts was performed by the Kjeldahl method (A.O.A.C., 2005), and used for the calculation of specific activities (U.mgprotein-1).
2.2.2. Temperature stability of enzymes purified by microfiltration The thermal stability of amylases obtained from submerged and solid-state fermentation processes was measured by incubating the purified enzyme extracts from 40 ºC to 80 ºC. Aliquots were periodically taken to measure amylolytic activity (according to item 2.2.1.). All experiments were carried out in duplicate. The Arrhenius thermal deactivation and activation energy for thermal destruction constants were calculated according to previously reported methods by Colla et al. (2015), using the Equations 1 to 3 for the calculus of the thermal deactivation constant (Kd), half-lives (t1/2) and activation energy (Ea). The half-lives (t1/2) correspond to the time required, at the temperature tested, so that 50% of the initial enzyme concentration is inactivated. ln = − . ∆
(1)
/ =
,
(2)
= −
(3)
Where: RA = Residual activity of the enzyme; t = Time (min); Kd = Thermal deactivation constant; A = Arrhenius factor (depending, among other things, on the contact area); Ea = Activation energy; −1
−1
R = Ideal gases constant (8.314 J.mol .K ); T = Absolute temperature (K). 2.3. Immobilization of purified enzymes The retained fractions from microfiltered extracts of solid-state and submerged fermentations were used for immobilization in polyurethane. The monomers used for the synthesis of polyurethane are polyol polyether and toluene isocyanate, assigned by Mannes company (Brazil). The additives stannous octoate II, responsible for catalysis of the polymerization reaction, and the triethylenediamine, responsible for catalyzing the reaction between isocyanate and water, in amounts of 1.0% and 0.9% (m/m), respectively, were previously aggregated at polyether polyol. The experimental procedure for immobilization of amylolytic enzymes in polyurethane foam was accomplished in situ rigid polyurethane foam by the polymerization reaction of the monomers, using the volume ratio polyol polyether and toluene isocyanate of 3:3 (v/v), respectively. Previously, an optimization step where different proportions and volumes of monomers were tested (data not show).
The in situ experimental procedure for immobilization of amylolytic enzymes in polyurethane foam involved adding the enzyme prior to the polymerization step. Different volumes of enzyme extracts were used for the immobilization (1 mL, 1.5 mL, or 2 mL). The experiments were codified as E1-SSF, E1.5-SSF, E2-SSF, E1SF, E1.5-SF, E2-SF, corresponding to the volume of microfiltered enzyme extract added in the support. Microfiltrated enzyme extracts were added to polyol polyether and mixed until they were completely solubilized. Following the homogenization step, toluene isocyanate was added to the mixture, held for 30 seconds using a glass rod, and the polymerization reaction was initiated with expansion of the foam. This step was performed in an ice bath to prevent excessive increase in temperature due to the exothermic reaction generated by the mixture of monomers. The maximum temperatures reached by using the monomeric proportions employed in this study (3mL: 3mL) were 57 °C (Treichel et al., 2013.) and 45 °C for tests conducted in the absence and the ice bath presence, respectively. The polyurethane containing the enzyme (immobilized) was maintained for 24 h in a desiccator to minimize the moisture content and ensure complete solidification (Bustamante-Vargas et al., 2016). Thereafter, the support containing the immobilized enzyme was ground and characterized for enzyme activity. The yield of immobilized enzyme was calculated considering the overall activity of the free enzyme in solution (which considers the amount of enzyme extract purified by microfiltration employed in immobilization test) and the overall activity of the immobilized enzyme (which includes the total mass of immobilized produced), according to Equation 7 (Ficanha et al., 2015). % =
!"#$%&%'( )
. 100
(7)
Where: R = Immobilization yield (%); UImobilized = Enzymatic activity in immobilized fraction; U0 = Enzymatic activity of purified extract offered for the immobilization. 2.3.1. Evaluation of reaction recycling capacity of the immobilized amylase The reaction recycling capacity of amylase was tested in starch hydrolysis, considering that the greatest interest in the immobilization of enzymes is the need to reuse with cost reduction purposes. In 125 mL Erlenmeyer flasks, 0.1 g of immobilized enzyme was added to 1 mL of 1% starch solution (prepared in 0.5 M phosphate buffer, pH 7.0). The mixture was incubated for 30 min at 37 °C. After the reaction, the tests were filtered through filter paper to separate the immobilized enzyme, which was washed with distilled water and again subjected to the starch hydrolysis process. Reducing sugars were determined in the filtrate by the colorimetric method, using 3.5 dinitrosalicylic acid (DNS) (Miller, 1959). 2.4. Enzymatic hydrolysis of dry biomass of Spirulina platensis To perform the hydrolysis test, the biomass of Spirulina platensis LEB 52 containing 40% carbohydrate was used, which was cultured in a medium proposed by Margarites (2014). The hydrolysis was performed using concentrated (microfiltration) and immobilized enzymes. The Spirulina biomass used was characterized with respect to the following physicochemical parameters: carbohydrate concentration (Dubois et al., 1956), protein concentration determined by the Kjeldahl method, using a conversion factor of 5.22 that is specific for cyanobacteria (Lourenço et al., 2004), concentration of lipids by a method reported by Colla et al. (2004), humidity and ashes (A.O.A.C., 2005).
The biomass was subjected to thermal pretreatment at 121 °C, 101 kPa, for 30 min to facilitate ease of hydrolysis by disruption of the cell walls of microalgae. The enzymatic hydrolysis process (Table 2) was performed by the addition of 10% (v/v) of enzymes purified by microfiltration (EPM) or 5% (w/v) of enzymes purified by microfiltration and immobilized (EPMI) to a suspension of biomass of Spirulina platensis LEB 52 (100 g.L-1), pH fixed in 5.5, in Erlenmeyer flasks of 125 mL. The temperature was 40 °C for enzymes from solid-state fermentation and 50 °C for enzymes from submerged fermentation, chosen according to the results of thermal stability of the free enzymes, described in section 2.2.1. The enzymatic hydrolysis was carried out for 72 h, samples being collected every 24 h. The samples were then centrifuged at 4400 g for 30 min, followed by determination of reducing sugars using the 3.5 dinitrosalicylic based previously reported method (Miller, 1959).
3. Results and discussion 3.1. Production of amylolytic enzymes In the step of amylase production by solid-state and submerged fermentations, amylolytic activity was 24.63 U.gfermented bran-1 (corresponding to 1.63 U.mL-1 after extraction) after 72 h of fermentation and 0.79 U.mL-1 after 24 h of fermentation. These extracts were used in the purification process and enzyme immobilization. 3.2. Purification of amylolytic enzymes and evaluation of thermal stability of the purified enzyme extract Precipitation by salting out is one of the most widely applied methods for purification of enzymes, separating the proteins from other interfering compounds of the environment and harmless to the enzyme (Amid and Manap, 2014). The enzymatic
extracts from solid-state fermentation, precipitated with ammonium sulfate (EPSA) (Table 1), showed enzymatic activities similar to those from the unpurified crude extract (E), but the specific activity (SA) was higher, indicating that non-protein interfering agents from the crude extract were eliminated (Ugur et al., 2014). Table 1 After microfiltration, the solid-state fermentation extract purified with ammonium sulfate showed an increase in the enzyme activity of the retentate (RM), which can be explained by retention of a fraction of amylolytic enzymes with lower specificity in microfiltration membrane, with an increase by about 38% in the enzymatic activity. According to Fernández et al. (2013), low molecular-weight compounds that inhibited enzyme activity may have been eliminated in the fraction of the permeate. With respect to the specific enzymatic activities of the submerged-fermentation extracts purified by microfiltration, the results obtained for both the permeate extract (79.33±0.99 U.mgprotein-1) and the retentate (103.06±0.71 U.mgprotein-1), were superior to those observed for the crude extract (55.25±0.51 U.mgprotein-1). This shows that the membrane filtration process eliminated enzyme activity inhibitors such as ions or other low molecular-weight compounds that could be affecting the enzymatic activity (Mulder, 1997). Considering the higher specific activity, as compared with to those of the other extracts, the retentate fractions of enzymes from solid-state and submerged fermentations, obtained by microfiltration, were used for the immobilization process. 3.2.1. Evaluation of thermo stability of the purified extracts after purification by membrane filtration technology
Figures 1 and 2 show graphs of thermal stability of retentate extracts of solidstate and submerged fermentations determined at temperatures of 40 °C to 80 °C after microfiltration. Figure 1 Figure 2 Table 2 shows the values of thermal deactivation constants (Kd) and half-life time (t1/2) obtained from each condition. Table 2 Enzyme extracts from solid-state fermentation showed greater thermal stability at 40 °C and 50 °C (Table 2), which can be observed from the high half-life time (t1/2) at a temperature 50 °C (346.5 min) and the stable residual activity when the enzymes were subjected to a temperature of 40 °C (residual activity above 95%). In temperatures above 50 °C, the half-life time of the enzyme decreased considerably. At 60 °C, the enzymatic extract showed 42% residual activity after 10 min, with complete inactivation after 65 min incubation. After 1 min of exposure to temperatures of 80 °C and 70 °C, residual activity of these enzymes was less than 25%. Similarly, amylases obtained by submerged fermentation exhibited greater stability at lower temperatures. However, they showed more stability than enzymes obtained by solid-state fermentation at a temperature of 60 °C, with a half-life time (t1/2) of 330 min, compared to t1/2 of 12.55 min for enzymes in solid-state fermentation at this temperature. The enzymes of the submerged fermentation showed a residual activity of 50% at 80 °C after 20 min of incubation, and a residual activity of 50% at 70 °C after 60 min
of incubation. The enzymes were completely inactivated after 70 min and 240 min at 80 °C and 70 °C, respectively. The purified enzyme extracts from submerged fermentation showed greater thermal deactivation energy (195.02 kJ.mol-1), than those from solid-state fermentation (135.70 kJ.mol-1). The enzymes obtained from solid-state fermentation (SSF) were more unstable than the enzymes from submerged fermentation (SF), owing to the smaller values of thermal deactivation energy. The amylase produced in this study had higher energy thermal deactivation than that obtained by Aguilar et al. (2002), which was 32.60 kJ.mol-1 using Lactobacillus manihotivorans LMG 18010. Pires, Veiga and Finardi Filho (2002) showed that amylases produced by Arracacia xanthorrhiza presented thermal deactivation energies of 171.98 kJ.mol-1. The activation energy reflects the dependence of thermal deactivation constant with respect to temperature (Atkins and Jones, 2001). The enzymes produced in this work present interesting perspectives of application in processes that require temperatures around 40 °C to 50 °C, such as simultaneous saccharification and fermentation processes for the production of bioethanol. An optimum temperature for carrying out the two steps is necessary, since the action of the enzyme may occur at temperatures around 50 °C, and fermentative microorganisms grow between 30 °C and 37 °C. However, the energy consumption of the process is lower than that consumed by the steps conducted separately (Ishola et al., 2013; Binod et al., 2010). 3.3. Immobilization of amylolytic enzymes Table 3 shows the amylolytic activity (AA) of the purified enzyme extracts from both types of fermentation, used for immobilization, as well as the activity of enzymes
immobilized on the support per gram (AS), total amylolytic activity of the support (AT), and the immobilization yields (R) obtained. Table 3 In the E1-SSF test, an increase of 205.99% in the immobilized enzyme activity relative to the free enzyme (purified enzyme extract by microfiltration) was observed. The E1.5-SSF and E2-SSF trials had lower yields than the E1-SSF trial owing to higher amount of enzyme molecules immobilized per gram of support causing limiting effect of diffusion on the support. These values were higher than those reported by Bayramoglu et al. (2004) and Lim et al. (2003), who reported immobilization of less than 80% of amylases in sodium alginate. Day et al. (2003) achieved a α-amylase immobilization yield of about 75% in sodium alginate, using the enzymes produced by a GRS313 strain of Bacillus circulans. Of the immobilized amylases obtained from submerged fermentation, the E1-SF test also showed the highest yield (332.60±26.20%), compared to E1.5-SF (208.70±6.00%) and E2-SF (187.50±12.01%) tests. The increase in the enzymatic activity of amylase on binding to solid supports can be explained by favoring substrate binding to the enzyme due to conformational changes caused by binding of the enzyme to the support. The increase in enzyme stability in the solution reaction medium can also be an additional factor contributing to the same. The results for the tests of E1-SSF and E1-SF, which used 1 mL of the enzyme extract, showed a positive effect of immobilization on enzyme activity. This positive effect, according to Zhang et al. (2008), may be related to factors, such as availability and access to new active sites, and the possibility of reuse of immobilized biocatalyst. These results suggest that the polyurethane support provides protection to the enzyme
from the exterior environment, resulting in an increased amylolytic activity. In immobilized enzymes that can be linked to improvements in the microenvironment created within the support, which minimizes the influence of temperature and mimics the effects of crowding and confinement in a living cell. The use of a hydrophobic support, such as polyurethane for the conservation of the catalytic ability of immobilized enzymes is advantageous, since it simulates the natural environment of the enzyme, and can often promote hyper activation of catalytic activity. It also results in increased enzyme stability, because of the decrease in water retention due to the hydrophobicity of the support, possibly minimizing the denaturing processes that are related to the percentage of hydrating enzymes (Romaškevič et al., 2006). The possibility of reuse of immobilized amylases from Aspergillus niger was determined using the starch hydrolysis reaction as a model. The behavior of residual activity was obtained with reference to the activity of the first reaction (Figure 3). The reuse of enzymes in more than one reaction cycle is one of the main goals of immobilization. This fact is important for enzymes, as the high cost is a major problem in their industrial application. Generally, an enzyme could be reused until its activity is greater than or equal to 50% of the initial activity (Ficanha et al., 2015). Figure 3 From the results obtained for solid-state fermentation (Figure 3a), it is observed that E1-SSF assay permitted 11 recycles, and E1.5-SSF tests and E2-SSF permitted 10 recycles, until they reached a residual activity of about 50%. These immobilized enzymes could be used about 11 times in the same reaction process of enzymatic hydrolysis, thus reducing the process cost, because according to Wang, Liu and Wang
(2011) the cost involved in the immobilization of enzymes together with the use of cycles should be considered in the overall economy of an industrial process. The tests using 1 mL, 1.5 mL and 2 mL of immobilized enzymes from submerged fermentation (Figure 3b) also allowed 11 recycles before reaching a residual activity of about 50%. Thus, both immobilized purified extracts from solid-state fermentation and from submerged fermentation showed similar operational stability with 11 possible recycles. In this work, we demonstrated techniques for obtaining enzymes with high efficiency and using a reduced number of low cost purification steps in comparison with conventional chromatographic techniques. 3.4. Enzymatic hydrolysis of dry biomass of Spirulina platensis LEB 52, using free and immobilized enzymes To evaluate the influence of immobilization process on the catalytic efficiency of amylases produced, we carried out the enzymatic hydrolysis process for Spirulina platensis LEB 52 biomass. Table 4 shows the results of reducing sugars generated during the enzymatic hydrolysis of Spirulina platensis LEB 52 biomass. An increase in the amounts of reducing sugars was observed on comparing the results of initial and final time of hydrolysis. The higher amount of reducing sugar generated was 19.3 g.L-1, from enzymes produced from submerged fermentation and purified by microfiltration. Fermentable sugars around 2 g.L-1 were obtained from acid hydrolysis of Chlorella vulgaris by Klein (2013). However, acid hydrolysis can display effects such as the formation of certain chemical species from the degradation of sugars and other undesirable organic compounds, since they are detrimental to the fermenting agents in bioethanol synthesis. Table 4
Different amounts of enzymes (10% v/v for free enzymes and 5% w/v for immobilized enzymes) were added, as in the hydrolysis process. We calculated the amount of reducing sugars generated by the quantity of enzymes (assessed from the amount of protein in the extract purified from the microfiltration or amount of protein immobilized on polyurethane support), and these results are shown in Figure 4. The values shown above the vertical bars represent the increased amount of reducing sugars released per mg of protein, on comparing free and immobilized enzymes. We found that the enzymes produced via submerged fermentation showed higher reducing sugar released by the amount of protein present. When the enzymes were immobilized, the yields were 11.9 to 18.5 times higher than those obtained using free enzymes. Increased yields of 17.5 to 21.3 fold were observed on comparing the immobilized and free enzymes from solid-state fermentation. The best results of biomass saccharification obtained by submerged fermentation enzymes can be related to higher thermal stability and to higher immobilization yields. Figure 4 The conversion of carbohydrates obtained in the saccharification processes were 3.08% and 3.44 to the enzymatic extracts purified by microfiltration and immobilization, obtained from solid state and submerged fermentations respectively; and 3.49% and 5.33% to the enzymatic extracts purified by microfiltration, obtained from solid state and submerged fermentations, respectively. These conversions are considered low when compared to those reported previously by Choi et al. (2010), who obtained conversion factors of 23.5% using Chlamydomonas reinhardtii UTEX 90. Hernández et al. (2015) showed conversion factors of 18%, 15%, and 11%, using Chlorella sorokiniana gaditana, Nannochloropsis,
and Scenedesmus almeriense, respectively. The results obtained in the present study can be optimized with the study of fermentation medium for increasing the concentration of enzymes produced during the bioprocess, but that was not the main objective of this study. This work contributes with the research area of enzymes and biofuels in many ways. The use of wheat bran as a substrate in the production of amylases contributed to adding economic value to by-products of agribusiness. These enzymes have potential application in simultaneous saccharification and fermentation processes because of their stability at temperatures in the range of 50 °C. Immobilization using polyurethane as support showed a yield of 332% from enzymes in submerged fermentation and 205% from solid-state fermentation. Regarding operational stability, results showed the possible reuse of the immobilized amylases up to 11 times. Enzyme recycling will significantly lower the process cost; one of the main advantages of the purified enzyme extract. The concentrations of reducing sugars generated during the enzymatic hydrolysis of polysaccharides from Spirulina platensis LEB 52, using free (purified enzyme extract by microfiltration) and immobilized enzymes, were increased in all experiments after 72 h.
4. Conclusion The enzymes produced in submerged and solid state fermentations have potential application in simultaneous saccharification and fermentation processes. The use of microalgae technology on an industrial scale for the production of biofuels is still at an early stage. This work presented an alternative to the viability of this process. The purification with microfiltration and immobilization in polyurethane
can be and useful strategy to improve the saccharification of microalgae biomass. The inclusion of immobilization step improved the yields in sugars of 11.9 to 18.5 and of 17.5 to 21.3 times to the purified enzymes of submerged and solid state fermentation, respectively.
5 Acknowledgments The authors are pleased to acknowledge the National Council for Scientific and Technological Development (CNPq) and Coordination of Improvement of Higher Education Personnel (CAPES) for the financial support.
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Figure captions
Figure 1 - Thermal stability of the extracts purified by microfiltration, at temperatures of 60 °C to 80 °C for the extracts obtained from fermentation in solid state, and at 70 °C to 80 °C for the extracts obtained from submerged fermentation.
Figure 2 - Thermal stability of the extracts purified by microfiltration, at temperatures of 40 °C to 50 °C for the extracts obtained from fermentation in solid state, and at 40 °C to 60 °C for the extracts obtained from submerged fermentation.
Figure 3 - Residual amylolytic activity of the operational stability for supports of SSF and SF. (a) SSF, ♦ Free enzyme (purified enzyme extract by microfiltration), ▲ E1, ■ E1.5, × E2; (b) SF, ♦ Free enzyme (purified enzyme extract by microfiltration), ▲ E1, ■ E1.5, × E2.
Figure 4 - Reducing sugars (RS) generated per mg of protein present in the enzymatic extract purified by microfiltration (EPM) or by microfiltration and immobilization (EPMI), produced in submerged (SF) or solid state fermentation (SSF).
Figure 1
Figure 2
Figure 3a
Figure 3b
Figure 4
Table 1 - Amylolytic activities for concentrated/purified enzyme extracts obtained by SSF and SF.
SSF
SF
Determinations
E
EPSA
RM
PM
AA (U.mL-1)
1.63±0.01
1.64±0.03
2.26±0.01
1.03±0.01
PTN (mg.mL-1)
0.039±0.01
0.037±0.01
0.044±0.01
0.030±0.01
SA (U.mg-1)
41.82±0.13
44.09±0.75
50.77±0.19
34.41±0.36
AA (U.mL-1)
0.79±0.01
-
1.13±0.01
1.19±0.01
PTN (mg.mL-1)
0.014±0.01
-
0.011±0.01
0.015±0.01
AE (U.mgprotein-1)
55.25±0.51
-
103.06±0.71
79.33±0.99
AA: Amylolitic activity; PTN: Concentration of proteins in the enzymatic extract; SA: Specific activity; E: Enzymatic extract non purified; EPSA: Enzymatic extract after purification with ammonium sulfate (only to SSF); RM: Retentate of microfiltration process; PM: Permeate of microfiltration process. *Mean ± standard deviation.
Table 2 - Constant thermal deactivation (Kd), the coefficients of determination (R²) and half-life time (t1/2) of the enzymatic extracts obtained by SSF (solid state fermentation) and SF (submerged fermentation) after the concentration/purification by microfiltration. SSF
SF
Temperature (ºC)
Kd (min-1)
R²
t1/2 (min)
Kd (min-1)
R²
t1/2 (min)
50
0.0020
0.9502
346.5
-
-
-
60
0.0552
0.9846
12.55
0.0021
0.9893
330.00
70
0.7133
0.9682
0.97
0.0150
0.9978
46.20
80
0.7632
0.9679
0.90
0.0334
0.9899
20.74
Table 3 - Amylase enzyme immobilization yield purified obtained by SSF and SF polyurethane. E1-SSF
E1,5-SSF
E2-SSF
E1-SF
E1,5-SF
E2-SF
AA (U)*
2.26±0.01
3.39±0.01
4.52±0.01
1.13±0.01
1.69±0.01
2.26±0.01
AS (U)*
0.39±0.01
0.39±0.01
0.39±0.01
0.36±0.02
0.33±0.01
0.39±0.02
AT (U)*
4.67±0.15
3.42±0.15
4.62±0.12
3.78±0.29
3.56±0.10
4.26±0.28
R (%)*
205.99±6.87 103.30±4.60 92.60±2.50 332.60±26.20 208.70±6.00 187.50±12.01
AA: Amylolytic activity present in the mass in the purified enzyme extract by microfiltration added immobilization; AS: Amylolytic activity per gram of support; AT: Total activity in support; SSF: Solid state fermentation; SF: Submerged fermentation. E1; E1,5; E2: volumes of extracts used in the immobilization. *Mean ± standard deviation.
Table 4 - Reducing sugars generated during the enzymatic hydrolysis process of intracellular polysaccharides of microalgae biomass. Reducing sugars (g.L-1)* Experiment 0h
24 h
48 h
72 h
(1) EPM SSF + Sp. LEB 52 1.51±0.13ª 1.53±0.01b 1.71±0.02b 2.09±0.04b (2) EPMI SSF + Sp. LEB 52 1.51±0.13ª 1.26±0.03ª 1.44±0.03ª 1.84±0.06ª (3) EPM SF + Sp. LEB 52
1.51±0.13a 1.87±0.07d 2.71±0.02d 3.19±0.03c
(4) EPMI SF + Sp. LEB 52
1.51±0.13a 1.49±0.07c 1.89±0.03c 2.06±0.05b
EPM: Enzymatic extract purified by microfiltration; EPMI: Enzymatic extract purified by microfiltration and immobilization; SSF: Solid state fermentation; SF: Submerged fermentation; Sp.: Spirulina platensis. *Mean ± standard deviation.
Production and purification of amylolytic enzymes for saccharification of microalgal biomass
Highlights: - Production of enzymes using agro industrial residuals - Proposition of a combined strategy of purification, applying membrane technology and immobilization - The enzymes presented thermal stability at 50 ºC, allowing their use in procedures of simultaneous saccharification and fermentation of microalgal biomass. - We demonstrate the use of these enzymes in the saccharification of lab-produced, carbohydrate-rich biomass.
S0960-8524(16)31562-0 http://dx.doi.org/10.1016/j.biortech.2016.11.047 BITE 17293
To appear in:
Bioresource Technology
Received Date: Revised Date: Accepted Date:
2 September 2016 10 November 2016 12 November 2016
Please cite this article as: Francine Rodrigues, E., Matuella Moreira Ficanha, A., Marcos Dallago, R., Treichel, H., Oliveira Reinehr, C., Paula Machado, T., Borges Nunes, G., Maria Colla, L., Production and purification of amylolytic enzymes for saccharification of microalgal biomass, Bioresource Technology (2016), doi: http:// dx.doi.org/10.1016/j.biortech.2016.11.047
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Production and purification of amylolytic enzymes for saccharification of microalgal biomass
Éllen Francine Rodriguesa, Aline Matuella Moreira Ficanhab, Rogério Marcos Dallago b, Helen Treichelc, Christian Oliveira Reinehra, Tainara Paula Machado a, Greice Borges Nunesa, Luciane Maria Collaa*
a
University of Passo Fundo, Campus I, km 171, BR 285, P.O. Box 611, CEP 99001-
970, Passo Fundo, RS, Brazil b
Regional and Integrate University of Upper Uruguay and Missions, URI - Erechim,
Av. Sete de Setembro 1621, Erechim, RS 99700-000, Brazil c
Federal University of Fronteira Sul – Campus de Erechim, ERS 135, km 72, n° 200,
99700-970, Erechim, RS, Brazil * Corresponding author. E-mail: [email protected], [email protected]
Keywords: Fungal amylases, biofuels, immobilization, microfiltration
ABSTRACT The aim of this study was the production of amylolytic enzymes by solid state or submerged fermentations (SSF or SF, respectively), followed by purification using chemical process or microfiltration and immobilization of purified enzymes in a polyurethane support. The free and immobilized enzymes obtained were used to evaluate enzymatic hydrolysis of the polysaccharides of Spirulina. Microfiltration of the crude extracts resulted in an increase in their specific activity and thermal stability at 40 °C and 50 °C for 24 h, as compared to extracts obtained by SSF and SF. Immobilization of polyurethane purified enzyme produced yields of 332% and 205% for the enzymes obtained by SSF and SF, respectively. Free or immobilized enzymes favor the generation of fermentable sugar, being the application of the purified and immobilized enzymes in the hydrolysis of microalgal polysaccharides considered a promising alternative towards development of the bioethanol production process from microalgal biomass.
KEYWORDS: Fungal amylases, biofuels, immobilization, microfiltration
1. Introduction The demand for enzymes, especially amylases, is constantly growing owing to the variety of industrial applications of these hydrolases: hydrolyzed starch processing in the form of glucose and fructose syrups, alcoholic beverage production, textiles, bioethanol production, and others (Joshi and Satyanarayana, 2015; Sherif et al., 2013). This market has been stimulated recently by the production of bioethanol from starch and microalgae. It is estimated that the production and application of enzymes in different markets would increase to 17.5 billion dollars in 2024. The increasing demand for bioethanol and biodiesel in emerging economies including India, Brazil, and Thailand will augment industry growth over the forecast period (Grand View Research, 2016). The need to modify the current energy matrix, based on non-renewable resources like oil, has resulted in increased demand for global production of biofuels, and led to the diversification of biomass used for this purpose (Alemán-Nava, 2014). The use of non-conventional biomass, such as the microalgal biomass, has emerged as a promising alternative (Bahadar and Khan, 2013). The use of microalgae have been reported by many authors, who proposed that algal biomass, especially the carbohydrate- and lipid-rich species (Ruiz et al., 2013), can be used for chemical or biotechnological production of bioethanol and biodiesel (Zhu et al., 2014; Ghasemi et al., 2012). In this context, the use of amylolytic enzymes in bioethanol production appears as a promising technology and the study of enzymes that allow simultaneous hydrolysis and fermentation processes with viable yields are very relevant (Aikawa et al., 2013; Fernandes et al., 2012; Choi et al., 2010).
Among the microalgae used to obtain bioethanol, Spirulina platensis is well known in the scientific community because of its ability to accumulate proteins and other biocompounds. It has been reported to accumulate carbohydrates (up to 70%), when cultured in a nutrient-depleted medium containing added organic sources of carbon (Salla et al., 2016). The production of amylolytic enzymes can be performed using filamentous fungi by bioprocesses, or using by-products derived from the agricultural industry, such as wheat bran, corn bran, soybean meal, sugar cane bagasse, among others (RamosSanchez et al., 2015; Oliveira et al., 2013). Among the filamentous fungi, Aspergillus is the most widely used genus for the production of amylolytic enzymes (Venkat, 2007). Several different techniques have been developed, searching for higher yields, more specific and stable enzymes, making more industrial applications possible (Soccol and Vanderbergue, 2003). In the last decade solid-state fermentation (SSF) has received increased attention from researchers as a bio-based process. This technology allows for the valorization of wastes for the production of valuable commodities such as enzymes (El-Bakry et al., 2015). Solid state fermentation (SSF) is characterized for the growth of the microorganism in the absence of free water and this kind of technique can be used for production of many bio-products. Although important drawbacks are described concerning the SSF scale up largely due to heat transfer, sterilization costs and culture heterogeneity (Casciatori et al., 2016), SSF has several advantages over submerged fermentation (SmF) because of the low capital investment, the simple technique, low energy requirement, end-product inhibition, low waste water output and better product recovery (Pandey at al., 2000).
The development of techniques for concentration, purification, and immobilization of enzymes is essential for obtaining enzymes with high catalytic efficiency. Pure and stable enzymes with high specificity for substrates can be used in industrial processes in a similar way to chemical catalysts, and their recovery and reuse would facilitate a better cost–benefit ratio (Cunha et al., 2014). The purification or concentration of enzymes by membrane separation methods has been proposed for improving production processes owing to their advantages: no addition of chemical products, reduction in process energy consumption, and ease of scale up (Saxena et al., 2009). Downstream of purification/concentration, immobilization of the enzymes is a lucrative option to bypass the limitations of free enzymes by imparting the following properties: greater enzyme stability, operational ease of separation, and possibility of reuse. Immobilization may be achieved by encapsulation, by physical or chemical interactions with a particular media. In the process, following factors should be considered: economic, operational, experimental conditions employed in synthesis, ease of production, duration of the process, and the amount of enzyme incorporated into the support (Ficanha et al., 2015). In this context, we aimed at optimizing the production of amylolytic enzymes from Aspergillus niger using different fermentation processes, followed by their purification, immobilization, and application in the hydrolysis of Spirulina platensis biomass.
2. Material and Methods 2.1. Microorganisms, inoculum preparation, and enzyme production bioprocesses
The enzymes were produced using Aspergillus niger. The isolation and identification of this microorganism were well described in a previous work of our research group. Through the comparison of the partial area of the gene β-Tubulin of the isolated fragments or fungus with other sequences deposited in Genbank was obtained 98.2% of similarity of isolated fungus with Aspergillus niger, enough similarity to consider as the same species for fungus Treichel et al. (2016). The inoculum preparation was carried out by inoculating the microorganism in 1-L flasks, containing 100 mL of PDA medium (Potato Dextrose Agar), followed by incubation at 30 °C for 7 d. Wheat bran was used as a substrate for the bioprocess. In solid-state fermentation (SSF), moisture of the bran was adjusted to 60% with a sterile nutrient solution. For submerged fermentation (SF), the medium was prepared from the baking of 10% (w/v), at 100 °C for 30 min. After the cooking, filtration was carried out through a cotton filter to remove solids. In both culture media, 10% (v/v) of solution was added, which was composed of the following ingredients: 2 g.L-1 KH2PO4, 1 g.L-1 MgSO4, 0.225 g CaCl2, and 10 mL.L1
of trace solution (0.63 mg.L-1 FeSO4.7H2O, 0.01 mg.L-1 MnSO4 and 0.62 mg.L-1
ZnSO4 and distilled water to a volume of 1 L). The medium were sterilized at 121 °C, 101 kPa for 20 min, and the inoculation was performed after cooling the medium to room temperature. Solid-state fermentation experiments were conducted in 1-L beakers, with 100 g of bran inoculated with 107 spores/gmedium, and subsequent incubation at 30 °C for 144 h. The submerged cultivations were carried out in a 5-L bioreactor (Autoclavable
Fermentor Bioreactor SL - 135, SOLAB), containing 3 L of growth medium, inoculated with 4.106 spores/mLmedium, and incubated at 30 °C with shaking at 120 rpm for 72 h. The pH of the medium was around 5.8 and 6.0 at the beginning of fermentation processes in solid-state and submerged cultivation, respectively. 2.2. Purification of enzymes After the solid-state fermentation, the enzymes were extracted using a mediabuffer proportion of 1:17 (g fermented medium:mL phosphate buffer pH 7.0) in an orbital shaker at 35 °C for 60 min and 100 rpm (PARIS et al., 2012). The suspension was filtrated through cotton and centrifuged (Cientec Model CT - 5000) at 910 g for 20 min aiming at removing particulate solids. In this obtained crude extract, ammonium sulfate was added slowly and under stirring on ice bath, until 80% saturation (56.5 g ammonium sulfate in 100 mL of crude extract). After addition of salt, the mixture was kept standing overnight at 4 °C, and centrifuged at 1310 g for 30 min. The pellet was resuspended in distilled water, and the obtained enzyme solution was used in the analytical determinations following the purification process. The extract was called purified extract with ammonium sulfate. The medium of the submerged fermentation culture was filtered through cotton to separate the hyphae, followed by subsequent centrifugation at 910 g for 20 min to remove the particulate solids to yield the crude extract of submerged fermentation. The crude enzyme extract of the submerged fermentation and the purified extract with ammonium sulfate from solid-state fermentation were subjected to microfiltration to separate enzymes of different molecular weights from other low molecular weight compounds from the fermentation. The microfiltration system consisted of an acrylic module, a peristaltic pump, a ball valve to control the flow, pressure gauge, and hose
connector. System operation was at 200 kPa, using a flat microfiltration membrane consisting of polyvinylidene difluoride with a pore diameter of 0.4 µm (GE Water and Process Technologies). The enzymatic activity and the specific enzymatic activity of the purified extracts were determined according to the method described below (2.2.1.). 2.2.1. Determination of enzymatic activity of the extracts The amylolytic activity of the enzyme extracts was determined using starch as a substrate for the saccharification method, which is based on production of reducing sugars (Miller, 1959). One enzyme activity unit (U) was defined as the amount of enzyme capable of releasing 1 µmol of glucose per minute under the conditions of the proposed method. The quantification of proteins in the extracts was performed by the Kjeldahl method (A.O.A.C., 2005), and used for the calculation of specific activities (U.mgprotein-1).
2.2.2. Temperature stability of enzymes purified by microfiltration The thermal stability of amylases obtained from submerged and solid-state fermentation processes was measured by incubating the purified enzyme extracts from 40 ºC to 80 ºC. Aliquots were periodically taken to measure amylolytic activity (according to item 2.2.1.). All experiments were carried out in duplicate. The Arrhenius thermal deactivation and activation energy for thermal destruction constants were calculated according to previously reported methods by Colla et al. (2015), using the Equations 1 to 3 for the calculus of the thermal deactivation constant (Kd), half-lives (t1/2) and activation energy (Ea). The half-lives (t1/2) correspond to the time required, at the temperature tested, so that 50% of the initial enzyme concentration is inactivated. ln = − . ∆
(1)
/ =
,
(2)
= −
(3)
Where: RA = Residual activity of the enzyme; t = Time (min); Kd = Thermal deactivation constant; A = Arrhenius factor (depending, among other things, on the contact area); Ea = Activation energy; −1
−1
R = Ideal gases constant (8.314 J.mol .K ); T = Absolute temperature (K). 2.3. Immobilization of purified enzymes The retained fractions from microfiltered extracts of solid-state and submerged fermentations were used for immobilization in polyurethane. The monomers used for the synthesis of polyurethane are polyol polyether and toluene isocyanate, assigned by Mannes company (Brazil). The additives stannous octoate II, responsible for catalysis of the polymerization reaction, and the triethylenediamine, responsible for catalyzing the reaction between isocyanate and water, in amounts of 1.0% and 0.9% (m/m), respectively, were previously aggregated at polyether polyol. The experimental procedure for immobilization of amylolytic enzymes in polyurethane foam was accomplished in situ rigid polyurethane foam by the polymerization reaction of the monomers, using the volume ratio polyol polyether and toluene isocyanate of 3:3 (v/v), respectively. Previously, an optimization step where different proportions and volumes of monomers were tested (data not show).
The in situ experimental procedure for immobilization of amylolytic enzymes in polyurethane foam involved adding the enzyme prior to the polymerization step. Different volumes of enzyme extracts were used for the immobilization (1 mL, 1.5 mL, or 2 mL). The experiments were codified as E1-SSF, E1.5-SSF, E2-SSF, E1SF, E1.5-SF, E2-SF, corresponding to the volume of microfiltered enzyme extract added in the support. Microfiltrated enzyme extracts were added to polyol polyether and mixed until they were completely solubilized. Following the homogenization step, toluene isocyanate was added to the mixture, held for 30 seconds using a glass rod, and the polymerization reaction was initiated with expansion of the foam. This step was performed in an ice bath to prevent excessive increase in temperature due to the exothermic reaction generated by the mixture of monomers. The maximum temperatures reached by using the monomeric proportions employed in this study (3mL: 3mL) were 57 °C (Treichel et al., 2013.) and 45 °C for tests conducted in the absence and the ice bath presence, respectively. The polyurethane containing the enzyme (immobilized) was maintained for 24 h in a desiccator to minimize the moisture content and ensure complete solidification (Bustamante-Vargas et al., 2016). Thereafter, the support containing the immobilized enzyme was ground and characterized for enzyme activity. The yield of immobilized enzyme was calculated considering the overall activity of the free enzyme in solution (which considers the amount of enzyme extract purified by microfiltration employed in immobilization test) and the overall activity of the immobilized enzyme (which includes the total mass of immobilized produced), according to Equation 7 (Ficanha et al., 2015). % =
!"#$%&%'( )
. 100
(7)
Where: R = Immobilization yield (%); UImobilized = Enzymatic activity in immobilized fraction; U0 = Enzymatic activity of purified extract offered for the immobilization. 2.3.1. Evaluation of reaction recycling capacity of the immobilized amylase The reaction recycling capacity of amylase was tested in starch hydrolysis, considering that the greatest interest in the immobilization of enzymes is the need to reuse with cost reduction purposes. In 125 mL Erlenmeyer flasks, 0.1 g of immobilized enzyme was added to 1 mL of 1% starch solution (prepared in 0.5 M phosphate buffer, pH 7.0). The mixture was incubated for 30 min at 37 °C. After the reaction, the tests were filtered through filter paper to separate the immobilized enzyme, which was washed with distilled water and again subjected to the starch hydrolysis process. Reducing sugars were determined in the filtrate by the colorimetric method, using 3.5 dinitrosalicylic acid (DNS) (Miller, 1959). 2.4. Enzymatic hydrolysis of dry biomass of Spirulina platensis To perform the hydrolysis test, the biomass of Spirulina platensis LEB 52 containing 40% carbohydrate was used, which was cultured in a medium proposed by Margarites (2014). The hydrolysis was performed using concentrated (microfiltration) and immobilized enzymes. The Spirulina biomass used was characterized with respect to the following physicochemical parameters: carbohydrate concentration (Dubois et al., 1956), protein concentration determined by the Kjeldahl method, using a conversion factor of 5.22 that is specific for cyanobacteria (Lourenço et al., 2004), concentration of lipids by a method reported by Colla et al. (2004), humidity and ashes (A.O.A.C., 2005).
The biomass was subjected to thermal pretreatment at 121 °C, 101 kPa, for 30 min to facilitate ease of hydrolysis by disruption of the cell walls of microalgae. The enzymatic hydrolysis process (Table 2) was performed by the addition of 10% (v/v) of enzymes purified by microfiltration (EPM) or 5% (w/v) of enzymes purified by microfiltration and immobilized (EPMI) to a suspension of biomass of Spirulina platensis LEB 52 (100 g.L-1), pH fixed in 5.5, in Erlenmeyer flasks of 125 mL. The temperature was 40 °C for enzymes from solid-state fermentation and 50 °C for enzymes from submerged fermentation, chosen according to the results of thermal stability of the free enzymes, described in section 2.2.1. The enzymatic hydrolysis was carried out for 72 h, samples being collected every 24 h. The samples were then centrifuged at 4400 g for 30 min, followed by determination of reducing sugars using the 3.5 dinitrosalicylic based previously reported method (Miller, 1959).
3. Results and discussion 3.1. Production of amylolytic enzymes In the step of amylase production by solid-state and submerged fermentations, amylolytic activity was 24.63 U.gfermented bran-1 (corresponding to 1.63 U.mL-1 after extraction) after 72 h of fermentation and 0.79 U.mL-1 after 24 h of fermentation. These extracts were used in the purification process and enzyme immobilization. 3.2. Purification of amylolytic enzymes and evaluation of thermal stability of the purified enzyme extract Precipitation by salting out is one of the most widely applied methods for purification of enzymes, separating the proteins from other interfering compounds of the environment and harmless to the enzyme (Amid and Manap, 2014). The enzymatic
extracts from solid-state fermentation, precipitated with ammonium sulfate (EPSA) (Table 1), showed enzymatic activities similar to those from the unpurified crude extract (E), but the specific activity (SA) was higher, indicating that non-protein interfering agents from the crude extract were eliminated (Ugur et al., 2014). Table 1 After microfiltration, the solid-state fermentation extract purified with ammonium sulfate showed an increase in the enzyme activity of the retentate (RM), which can be explained by retention of a fraction of amylolytic enzymes with lower specificity in microfiltration membrane, with an increase by about 38% in the enzymatic activity. According to Fernández et al. (2013), low molecular-weight compounds that inhibited enzyme activity may have been eliminated in the fraction of the permeate. With respect to the specific enzymatic activities of the submerged-fermentation extracts purified by microfiltration, the results obtained for both the permeate extract (79.33±0.99 U.mgprotein-1) and the retentate (103.06±0.71 U.mgprotein-1), were superior to those observed for the crude extract (55.25±0.51 U.mgprotein-1). This shows that the membrane filtration process eliminated enzyme activity inhibitors such as ions or other low molecular-weight compounds that could be affecting the enzymatic activity (Mulder, 1997). Considering the higher specific activity, as compared with to those of the other extracts, the retentate fractions of enzymes from solid-state and submerged fermentations, obtained by microfiltration, were used for the immobilization process. 3.2.1. Evaluation of thermo stability of the purified extracts after purification by membrane filtration technology
Figures 1 and 2 show graphs of thermal stability of retentate extracts of solidstate and submerged fermentations determined at temperatures of 40 °C to 80 °C after microfiltration. Figure 1 Figure 2 Table 2 shows the values of thermal deactivation constants (Kd) and half-life time (t1/2) obtained from each condition. Table 2 Enzyme extracts from solid-state fermentation showed greater thermal stability at 40 °C and 50 °C (Table 2), which can be observed from the high half-life time (t1/2) at a temperature 50 °C (346.5 min) and the stable residual activity when the enzymes were subjected to a temperature of 40 °C (residual activity above 95%). In temperatures above 50 °C, the half-life time of the enzyme decreased considerably. At 60 °C, the enzymatic extract showed 42% residual activity after 10 min, with complete inactivation after 65 min incubation. After 1 min of exposure to temperatures of 80 °C and 70 °C, residual activity of these enzymes was less than 25%. Similarly, amylases obtained by submerged fermentation exhibited greater stability at lower temperatures. However, they showed more stability than enzymes obtained by solid-state fermentation at a temperature of 60 °C, with a half-life time (t1/2) of 330 min, compared to t1/2 of 12.55 min for enzymes in solid-state fermentation at this temperature. The enzymes of the submerged fermentation showed a residual activity of 50% at 80 °C after 20 min of incubation, and a residual activity of 50% at 70 °C after 60 min
of incubation. The enzymes were completely inactivated after 70 min and 240 min at 80 °C and 70 °C, respectively. The purified enzyme extracts from submerged fermentation showed greater thermal deactivation energy (195.02 kJ.mol-1), than those from solid-state fermentation (135.70 kJ.mol-1). The enzymes obtained from solid-state fermentation (SSF) were more unstable than the enzymes from submerged fermentation (SF), owing to the smaller values of thermal deactivation energy. The amylase produced in this study had higher energy thermal deactivation than that obtained by Aguilar et al. (2002), which was 32.60 kJ.mol-1 using Lactobacillus manihotivorans LMG 18010. Pires, Veiga and Finardi Filho (2002) showed that amylases produced by Arracacia xanthorrhiza presented thermal deactivation energies of 171.98 kJ.mol-1. The activation energy reflects the dependence of thermal deactivation constant with respect to temperature (Atkins and Jones, 2001). The enzymes produced in this work present interesting perspectives of application in processes that require temperatures around 40 °C to 50 °C, such as simultaneous saccharification and fermentation processes for the production of bioethanol. An optimum temperature for carrying out the two steps is necessary, since the action of the enzyme may occur at temperatures around 50 °C, and fermentative microorganisms grow between 30 °C and 37 °C. However, the energy consumption of the process is lower than that consumed by the steps conducted separately (Ishola et al., 2013; Binod et al., 2010). 3.3. Immobilization of amylolytic enzymes Table 3 shows the amylolytic activity (AA) of the purified enzyme extracts from both types of fermentation, used for immobilization, as well as the activity of enzymes
immobilized on the support per gram (AS), total amylolytic activity of the support (AT), and the immobilization yields (R) obtained. Table 3 In the E1-SSF test, an increase of 205.99% in the immobilized enzyme activity relative to the free enzyme (purified enzyme extract by microfiltration) was observed. The E1.5-SSF and E2-SSF trials had lower yields than the E1-SSF trial owing to higher amount of enzyme molecules immobilized per gram of support causing limiting effect of diffusion on the support. These values were higher than those reported by Bayramoglu et al. (2004) and Lim et al. (2003), who reported immobilization of less than 80% of amylases in sodium alginate. Day et al. (2003) achieved a α-amylase immobilization yield of about 75% in sodium alginate, using the enzymes produced by a GRS313 strain of Bacillus circulans. Of the immobilized amylases obtained from submerged fermentation, the E1-SF test also showed the highest yield (332.60±26.20%), compared to E1.5-SF (208.70±6.00%) and E2-SF (187.50±12.01%) tests. The increase in the enzymatic activity of amylase on binding to solid supports can be explained by favoring substrate binding to the enzyme due to conformational changes caused by binding of the enzyme to the support. The increase in enzyme stability in the solution reaction medium can also be an additional factor contributing to the same. The results for the tests of E1-SSF and E1-SF, which used 1 mL of the enzyme extract, showed a positive effect of immobilization on enzyme activity. This positive effect, according to Zhang et al. (2008), may be related to factors, such as availability and access to new active sites, and the possibility of reuse of immobilized biocatalyst. These results suggest that the polyurethane support provides protection to the enzyme
from the exterior environment, resulting in an increased amylolytic activity. In immobilized enzymes that can be linked to improvements in the microenvironment created within the support, which minimizes the influence of temperature and mimics the effects of crowding and confinement in a living cell. The use of a hydrophobic support, such as polyurethane for the conservation of the catalytic ability of immobilized enzymes is advantageous, since it simulates the natural environment of the enzyme, and can often promote hyper activation of catalytic activity. It also results in increased enzyme stability, because of the decrease in water retention due to the hydrophobicity of the support, possibly minimizing the denaturing processes that are related to the percentage of hydrating enzymes (Romaškevič et al., 2006). The possibility of reuse of immobilized amylases from Aspergillus niger was determined using the starch hydrolysis reaction as a model. The behavior of residual activity was obtained with reference to the activity of the first reaction (Figure 3). The reuse of enzymes in more than one reaction cycle is one of the main goals of immobilization. This fact is important for enzymes, as the high cost is a major problem in their industrial application. Generally, an enzyme could be reused until its activity is greater than or equal to 50% of the initial activity (Ficanha et al., 2015). Figure 3 From the results obtained for solid-state fermentation (Figure 3a), it is observed that E1-SSF assay permitted 11 recycles, and E1.5-SSF tests and E2-SSF permitted 10 recycles, until they reached a residual activity of about 50%. These immobilized enzymes could be used about 11 times in the same reaction process of enzymatic hydrolysis, thus reducing the process cost, because according to Wang, Liu and Wang
(2011) the cost involved in the immobilization of enzymes together with the use of cycles should be considered in the overall economy of an industrial process. The tests using 1 mL, 1.5 mL and 2 mL of immobilized enzymes from submerged fermentation (Figure 3b) also allowed 11 recycles before reaching a residual activity of about 50%. Thus, both immobilized purified extracts from solid-state fermentation and from submerged fermentation showed similar operational stability with 11 possible recycles. In this work, we demonstrated techniques for obtaining enzymes with high efficiency and using a reduced number of low cost purification steps in comparison with conventional chromatographic techniques. 3.4. Enzymatic hydrolysis of dry biomass of Spirulina platensis LEB 52, using free and immobilized enzymes To evaluate the influence of immobilization process on the catalytic efficiency of amylases produced, we carried out the enzymatic hydrolysis process for Spirulina platensis LEB 52 biomass. Table 4 shows the results of reducing sugars generated during the enzymatic hydrolysis of Spirulina platensis LEB 52 biomass. An increase in the amounts of reducing sugars was observed on comparing the results of initial and final time of hydrolysis. The higher amount of reducing sugar generated was 19.3 g.L-1, from enzymes produced from submerged fermentation and purified by microfiltration. Fermentable sugars around 2 g.L-1 were obtained from acid hydrolysis of Chlorella vulgaris by Klein (2013). However, acid hydrolysis can display effects such as the formation of certain chemical species from the degradation of sugars and other undesirable organic compounds, since they are detrimental to the fermenting agents in bioethanol synthesis. Table 4
Different amounts of enzymes (10% v/v for free enzymes and 5% w/v for immobilized enzymes) were added, as in the hydrolysis process. We calculated the amount of reducing sugars generated by the quantity of enzymes (assessed from the amount of protein in the extract purified from the microfiltration or amount of protein immobilized on polyurethane support), and these results are shown in Figure 4. The values shown above the vertical bars represent the increased amount of reducing sugars released per mg of protein, on comparing free and immobilized enzymes. We found that the enzymes produced via submerged fermentation showed higher reducing sugar released by the amount of protein present. When the enzymes were immobilized, the yields were 11.9 to 18.5 times higher than those obtained using free enzymes. Increased yields of 17.5 to 21.3 fold were observed on comparing the immobilized and free enzymes from solid-state fermentation. The best results of biomass saccharification obtained by submerged fermentation enzymes can be related to higher thermal stability and to higher immobilization yields. Figure 4 The conversion of carbohydrates obtained in the saccharification processes were 3.08% and 3.44 to the enzymatic extracts purified by microfiltration and immobilization, obtained from solid state and submerged fermentations respectively; and 3.49% and 5.33% to the enzymatic extracts purified by microfiltration, obtained from solid state and submerged fermentations, respectively. These conversions are considered low when compared to those reported previously by Choi et al. (2010), who obtained conversion factors of 23.5% using Chlamydomonas reinhardtii UTEX 90. Hernández et al. (2015) showed conversion factors of 18%, 15%, and 11%, using Chlorella sorokiniana gaditana, Nannochloropsis,
and Scenedesmus almeriense, respectively. The results obtained in the present study can be optimized with the study of fermentation medium for increasing the concentration of enzymes produced during the bioprocess, but that was not the main objective of this study. This work contributes with the research area of enzymes and biofuels in many ways. The use of wheat bran as a substrate in the production of amylases contributed to adding economic value to by-products of agribusiness. These enzymes have potential application in simultaneous saccharification and fermentation processes because of their stability at temperatures in the range of 50 °C. Immobilization using polyurethane as support showed a yield of 332% from enzymes in submerged fermentation and 205% from solid-state fermentation. Regarding operational stability, results showed the possible reuse of the immobilized amylases up to 11 times. Enzyme recycling will significantly lower the process cost; one of the main advantages of the purified enzyme extract. The concentrations of reducing sugars generated during the enzymatic hydrolysis of polysaccharides from Spirulina platensis LEB 52, using free (purified enzyme extract by microfiltration) and immobilized enzymes, were increased in all experiments after 72 h.
4. Conclusion The enzymes produced in submerged and solid state fermentations have potential application in simultaneous saccharification and fermentation processes. The use of microalgae technology on an industrial scale for the production of biofuels is still at an early stage. This work presented an alternative to the viability of this process. The purification with microfiltration and immobilization in polyurethane
can be and useful strategy to improve the saccharification of microalgae biomass. The inclusion of immobilization step improved the yields in sugars of 11.9 to 18.5 and of 17.5 to 21.3 times to the purified enzymes of submerged and solid state fermentation, respectively.
5 Acknowledgments The authors are pleased to acknowledge the National Council for Scientific and Technological Development (CNPq) and Coordination of Improvement of Higher Education Personnel (CAPES) for the financial support.
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Figure captions
Figure 1 - Thermal stability of the extracts purified by microfiltration, at temperatures of 60 °C to 80 °C for the extracts obtained from fermentation in solid state, and at 70 °C to 80 °C for the extracts obtained from submerged fermentation.
Figure 2 - Thermal stability of the extracts purified by microfiltration, at temperatures of 40 °C to 50 °C for the extracts obtained from fermentation in solid state, and at 40 °C to 60 °C for the extracts obtained from submerged fermentation.
Figure 3 - Residual amylolytic activity of the operational stability for supports of SSF and SF. (a) SSF, ♦ Free enzyme (purified enzyme extract by microfiltration), ▲ E1, ■ E1.5, × E2; (b) SF, ♦ Free enzyme (purified enzyme extract by microfiltration), ▲ E1, ■ E1.5, × E2.
Figure 4 - Reducing sugars (RS) generated per mg of protein present in the enzymatic extract purified by microfiltration (EPM) or by microfiltration and immobilization (EPMI), produced in submerged (SF) or solid state fermentation (SSF).
Figure 1
Figure 2
Figure 3a
Figure 3b
Figure 4
Table 1 - Amylolytic activities for concentrated/purified enzyme extracts obtained by SSF and SF.
SSF
SF
Determinations
E
EPSA
RM
PM
AA (U.mL-1)
1.63±0.01
1.64±0.03
2.26±0.01
1.03±0.01
PTN (mg.mL-1)
0.039±0.01
0.037±0.01
0.044±0.01
0.030±0.01
SA (U.mg-1)
41.82±0.13
44.09±0.75
50.77±0.19
34.41±0.36
AA (U.mL-1)
0.79±0.01
-
1.13±0.01
1.19±0.01
PTN (mg.mL-1)
0.014±0.01
-
0.011±0.01
0.015±0.01
AE (U.mgprotein-1)
55.25±0.51
-
103.06±0.71
79.33±0.99
AA: Amylolitic activity; PTN: Concentration of proteins in the enzymatic extract; SA: Specific activity; E: Enzymatic extract non purified; EPSA: Enzymatic extract after purification with ammonium sulfate (only to SSF); RM: Retentate of microfiltration process; PM: Permeate of microfiltration process. *Mean ± standard deviation.
Table 2 - Constant thermal deactivation (Kd), the coefficients of determination (R²) and half-life time (t1/2) of the enzymatic extracts obtained by SSF (solid state fermentation) and SF (submerged fermentation) after the concentration/purification by microfiltration. SSF
SF
Temperature (ºC)
Kd (min-1)
R²
t1/2 (min)
Kd (min-1)
R²
t1/2 (min)
50
0.0020
0.9502
346.5
-
-
-
60
0.0552
0.9846
12.55
0.0021
0.9893
330.00
70
0.7133
0.9682
0.97
0.0150
0.9978
46.20
80
0.7632
0.9679
0.90
0.0334
0.9899
20.74
Table 3 - Amylase enzyme immobilization yield purified obtained by SSF and SF polyurethane. E1-SSF
E1,5-SSF
E2-SSF
E1-SF
E1,5-SF
E2-SF
AA (U)*
2.26±0.01
3.39±0.01
4.52±0.01
1.13±0.01
1.69±0.01
2.26±0.01
AS (U)*
0.39±0.01
0.39±0.01
0.39±0.01
0.36±0.02
0.33±0.01
0.39±0.02
AT (U)*
4.67±0.15
3.42±0.15
4.62±0.12
3.78±0.29
3.56±0.10
4.26±0.28
R (%)*
205.99±6.87 103.30±4.60 92.60±2.50 332.60±26.20 208.70±6.00 187.50±12.01
AA: Amylolytic activity present in the mass in the purified enzyme extract by microfiltration added immobilization; AS: Amylolytic activity per gram of support; AT: Total activity in support; SSF: Solid state fermentation; SF: Submerged fermentation. E1; E1,5; E2: volumes of extracts used in the immobilization. *Mean ± standard deviation.
Table 4 - Reducing sugars generated during the enzymatic hydrolysis process of intracellular polysaccharides of microalgae biomass. Reducing sugars (g.L-1)* Experiment 0h
24 h
48 h
72 h
(1) EPM SSF + Sp. LEB 52 1.51±0.13ª 1.53±0.01b 1.71±0.02b 2.09±0.04b (2) EPMI SSF + Sp. LEB 52 1.51±0.13ª 1.26±0.03ª 1.44±0.03ª 1.84±0.06ª (3) EPM SF + Sp. LEB 52
1.51±0.13a 1.87±0.07d 2.71±0.02d 3.19±0.03c
(4) EPMI SF + Sp. LEB 52
1.51±0.13a 1.49±0.07c 1.89±0.03c 2.06±0.05b
EPM: Enzymatic extract purified by microfiltration; EPMI: Enzymatic extract purified by microfiltration and immobilization; SSF: Solid state fermentation; SF: Submerged fermentation; Sp.: Spirulina platensis. *Mean ± standard deviation.
Production and purification of amylolytic enzymes for saccharification of microalgal biomass
Highlights: - Production of enzymes using agro industrial residuals - Proposition of a combined strategy of purification, applying membrane technology and immobilization - The enzymes presented thermal stability at 50 ºC, allowing their use in procedures of simultaneous saccharification and fermentation of microalgal biomass. - We demonstrate the use of these enzymes in the saccharification of lab-produced, carbohydrate-rich biomass.