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An esterase from Penicillium decumbens P6 involved in lignite depolymerization
Fuel 214 (2018) 416–422
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Full Length Article
An esterase from Penicillium decumbens P6 involved in lignite depolymerization Yi Yanga, Jinshui Yanga, Baozhen Lia, Entao Wangb, Hongli Yuana, a b
T
⁎
State Key Laboratory of Agrobiotechnology, College of Biological Sciences, China Agricultural University, Beijing 100193, China Departamento de Microbiología, Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional, Mexico, DF, Mexico
A R T I C L E I N F O
A B S T R A C T
Keywords: Penicillium decumbens Lignite Depolymerization Esterase Humic acid
In this study, lignite was degraded using a purified esterase from Penicillium decumbens P6 for the first time. The esterase was purified using ammonium sulfate precipitation, anion exchange and gel filtration chromatography. The recovery and purification yield of the enzyme were 15% and 83 folds, respectively. The molecular weight of the purified enzyme was about 45 kDa. Both crude and purified esterases were studied for lignite depolymerization. The tendency of depolymerization by crude enzyme was consistent with the enzyme secreted in the medium. Along with the increased purified esterase concentration from 8 to 50 mg/ml, A450 value increased from 0.38 to 2.08. The contribution of esterase to the depolymerization was about 40% in the crude supernatant. Compared with aHA (crude lignite humic acid), bHA (esterase degraded lignite humic acid) has a lower percentage of aromatic carbon and ester groups, but a higher percentage of aliphatic carbon. bHA can promote the growth of asparagus lettuce. The results demonstrated that lignite was depolymerized by the purified esterase and evidenced the potential of esterase application in conversion of lignite into compounds with high bioactivities.
1. Introduction
organic matter, corresponding to one to two ester bonds per 100 lignite carbon atoms [9]. A direct correlation between lignite solubilization and enzymatic hydrolytic activity was evidenced in the deuteromycete Trichoderma atroviride [10]. Hölker et al. [11] reported that T. atroviride could cleave both the carboxylic esters and the phenolic ether bonds during the solubilization of 14C-labelled lignite. However, unlike the peroxidases, no low molecular weight co-factors and mediators were known in the activation of esterases, and the steric hindrance might be an obstacle for esterases to permeate the macromolecular coal network and to depolymerize the coal [12]. Therefore, more research is needed to determine the actual role and the mechanism of esterases in coal degradation. In our previous work, the fungal strain Penicillium decumbens P6 capable of degrading lignite effectively presented inducible esterase activity in the lignite degradation [3], that implied a possible role of esterase in the depolymerization of lignite. In addition, the contents of humic acids and water-soluble humic materials greatly increased and the molecular mass of humic acid decreased in the fungal depolymerized lignite [13]. Moreover, the fungal transformed lignite humic acids have a better bioactivity than the crude lignite humic acids [14,15]. It is well known that humic acids contribute substantially to global soil
With a lignin-like structure, lignite is defined as a low rank coal with low calorific value, high moisture and high sulfur content, which limits its usage and conversion [1]. The direct combustion of lignite results in low thermal efficiency and low industrial profit. In addition, the piling up of lignite in the open air for a long time causes energy waste and environmental pollution [2]. The high content of humic acid and fulvic acid and the great reserve (130 million tons) of lignite in China make it a potential natural resource [3]. Hence, research on conversion of lignite into industrial and chemical products with high additional value or into liquid fuel is rationally developed [4]. Enzymatic conversion has been considered as an economic and environment-friendly way for transforming macromolecular coal into simpler and low molecular weight products [5]. Lignin peroxidases, manganese peroxidases and laccases are the most widely studied oxidative enzymes participating in the coal solubilization [6,7], and they require low molecular weight cofactors and mediators to depolymerize the lignin [8]. In addition to the oxidative enzymes, hydrolytic enzymes also showed a great potential to depolymerize lignite. In lignite, carboxylic ester groups amount to approximately 2.5–5% of the total
Abbreviations: CP/MAS 13C NMR, 13C nuclear magnetic resonance with cross-polarization/magic angle spinning; aHA, lignite humic acid; bHA, esterase degraded lignite humic acid; SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis; DEAE, Diethyl-Aminoethanol; SDS, Sodium Dodecyl Sulfonate; EDTA, Ethylene Diamine Tetraacetic Acid ⁎ Corresponding author at: Center of Life Science, China Agriculture University, No.2 Yuanmingyuan West Road, Haidian District, Beijing 100193, China. E-mail addresses: [email protected] (Y. Yang), [email protected] (J. Yang), [email protected] (B. Li), [email protected] (H. Yuan). https://doi.org/10.1016/j.fuel.2017.11.035 Received 16 August 2017; Received in revised form 26 October 2017; Accepted 11 November 2017 0016-2361/ © 2017 Elsevier Ltd. All rights reserved.
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fertility and have been widely utilized in agriculture, for example, they can promote plant growth and metabolism [16] and reduce some plant diseases [17]. To prove the roles of esterase in the enzymatic attack on lignite, we performed the present study using purified esterase from P. decumbens P6 culture. The crude lignite humic acids (aHA) and esterase degraded lignite humic acids (bHA) were quantified and characterized using 13C nuclear magnetic resonance with cross-polarization/magic angle spinning (CP/MAS 13C NMR) and Fourier transformed infrared spectrophotometry (FT-IR). The humic acids produced by esterase depolymerization presented a significantly positive effect on the growth of asparagus lettuce. These results evidenced the key role of esterase for P. decumbens P6 in hydrolyzing carboxyl within the lignite structure.
The dried samples were dissolved in 1 ml of 50 mM phosphate buffer (pH 7.0) and were loaded onto a Sephadex G-75 gel filtration column (GE) that had been pre-equilibrated with the same phosphate buffer. The column was eluted with 50 mM phosphate buffer (pH 7.0) at a flow rate of 1.5 ml/min. The elute was collected into 1 ml fractions for esterase activity detection as mentioned previously [18]. The active fractions were loaded onto the Sephadex G-75 gel filtration column again for further purification. The molecular weight of the purified esterase was determined by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).
2. Materials and methods
Protein sequencing was performed according to the method [19]. The acquired amino acid sequence was used in BLAST searching (https://blast.ncbi.nlm.nih.gov/Blast.cgi) for extracting the related sequences in the database. The sequence was aligned together with the extracted reference sequences by MEGA 6.06 software [20].
2.3. Protein sequencing
2.1. Lignite samples and fungal strain Lignite sample was collected from the Huolingele Minerals Administration Coalmine, Inner Mongolia of China. Air-dried lignite sample was pulverized and sieved with a 70 mesh screen. The sample preparation and elemental analysis of the crude lignite humic acid were performed according to the previous reported methods [13]. The fungal strain P. decumbens P6 (CGMCC 0866) used throughout the study was maintained at 4 °C on the yeast extract – malt agar (YMPG) slants, consisting of: glucose, 10 g; malt extract, 10 g; peptone, 2 g; yeast extract, 2 g; asparagines, 1 g; KH2PO4·3H2O, 2 g and MgSO4·7H2O, 1 g in 1 l distilled water. All of the chemicals used in this study were reagent grade purchased from the Beijing Chemicals and Reagent Corp., China, unless otherwise stated. The fungus was cultured on YMPG agar plates at 28 °C for 1 week, then the spores were washed and suspended in 0.9% of sodium chloride solution. Aliquots of 2 ml of the spore suspension (106 spores/ml) were inoculated into 500 ml flasks containing 200 ml liquid medium (10 g wheat bran; 5 g urea; 2 g KH2PO4·3H2O; 1 g MgSO4·7H2O and 1 g asparagine in 1 l of distilled water) which had been optimized for esterase production. Powdered lignite was added into the medium at the ratio of 1% (w/v) as an inducer for enzyme production. The inoculated flasks were cultivated by shaking at 150 rpm and 28 °C for 10 days. Culture samples in aliquots of 1 ml were taken every day, and centrifuged at 4 °C, 8000 rpm for 10 min. Supernatants were used for analyzing the esterase activity with the method [18]. When the esterase activity reached the maximum level (after 10 days of incubation), whole cultures were harvested for purification studies as mentioned subsequently.
2.4. Enzyme properties The esterase activity was determined by the spectrophotometric method [18]. Briefly, 50 μL supernatant of the culture was mixed with 950 μL 4-nitrophenyl acetate in 50 mM phosphate buffer (pH 7.0). The reaction was performed at 37 °C for 15 min. A400 (absorbance at 400 nm) was determined with a UV-1800 spectrophotometer (Shimadzu, Japan) to estimate the released p-nitrophenol. In control, 50 μL of the supernatant boiled for 30 min were used. One unit of esterase activity was defined as the amount of proteins liberating 1 µmol p-nitrophenol per min under the defined conditions. The optimum pH for esterase was tested at 37 °C in 50 mM citric acid buffer (pH 3.0, 4.0 and 5.0) and 50 mM phosphate buffer (pH 6.0, 7.0, 8.0 and 9.0). The optimum temperature for esterase was determined by standard assay ranging from 30 to 80 °C in the 50 mM phosphate buffer (pH 7.0). The results were expressed as relative activity to the value obtained at either optimal pH assay or optimal temperature assay. The pH stability of esterase was determined by measuring the residual activity after incubating the enzyme at 37 °C for 1 h in the aforementioned buffer (pH 3.0–9.0). In the thermostability assay, the enzyme solution was incubated in 50 mM phosphate buffer (pH 7.0) at temperatures ranging from 30 to 80 °C for 1 h, and the remaining activity was measured under the optimum conditions. The esterase activity obtained under the optimum conditions was defined as 100%. The Michaelis-Menten constant (Km) and maximal velocity (Vmax) of the purified esterase were determined by measuring the esterase activity at various 4-nitrophenyl acetate concentrations (0.2 mM, 0.4 mM, 0.8 mM, 1.6 mM and 3.2 mM). The Km and Vmax of the enzyme were calculated using the Lineweaver-Burk plot constructed by plotting the reciprocal of the substrate concentration on the X-axis and the reciprocal of the esterase reaction velocity on the Y-axis. All determinations were conducted in triplicate. The effects of metal ions and chemical reagents on the esterase activity were determined. Different metal ions and chemical reagents such as NH4+, Al3+, Ba2+, Ca2+, Mn2+, Mg2+, Ag+, Cu2+, Sn2+, Fe2+, Co2+, Pb2+, Zn2+, Fe3+, urea, SDS (Sodium Dodecyl Sulfonate) and EDTA (Ethylene Diamine Tetraacetic Acid) at 10 mM and 1 mM were added to the purified enzyme solution and then incubated at 4 °C for 2 h. The residual activities were determined under the standard assay conditions. The esterase activity of the enzyme in culture without addition of metal ions or chemical reagents was defined as 100%. Protein concentration was determined using the Bradford Protein Assay Kit (GenStar, Beijing, China) according to the manufacturer’s instructions.
2.2. Purification of esterase and determination of molecular weight After 10 days of cultivation, the supernatant was separated from the mycelium by centrifugation of the culture at 8000 rpm for 20 min at 4 °C, and then filtration with Whatman No. 1 filter paper. Proteins in the cell-free extract were precipitated at 4 °C with ammonium sulfate (80%) for about 2 h following by centrifugation at 8000 rpm for 20 min under 4 °C. The sediment was resuspended in 20 ml of 20 mM Tris-HCl buffer (pH 8.0) and dialyzed against 2 l of the same buffer for 24 h to remove (NH4)2SO4. Then, the protein solution was loaded on the columns listed bellow to separate and purify acetyl xylan esterase. Firstly, the protein solution was loaded on a DEAE (DiethylAminoethanol) Sepharose anion exchange column (GE) pre-equilibrated with 20 mM Tris-HCl buffer (pH 8.0). The column was washed with the same buffer until A280 (absorbance at 280 nm) of effluent became zero and then eluted with a linear NaCl gradient (0–1 M) at a flow rate of 1.5 ml/min. The elute was collected into fractions of 1 ml. Esterase activity in the fractions was detected with the previously described method [18] and fractions presenting esterase activity were pooled and lyophilized for the next step of purification. 417
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2.5. Lignite depolymerization Crude enzyme was obtained from the daily sample during the incubation by centrifugation of the fermentation medium at 8000 rpm for 10 min at 4 °C. The depolymerization assay was carried out at 5% (w/v) sterile lignite loading in 0.1 M phosphate buffer (pH 7.0) in a final volume of 1 ml. Depolymerization assays were performed at a total esterase loading of 10 U/g lignite in an orbital shaker incubator at 28 °C for 24 h. Control assays including esterase sample without lignite and lignite without esterase sample were performed under the same conditions. The supernatants were collected by centrifugation at 8000 rpm for 10 min at 4 °C and then A450 (absorbance at 450 nm) was measured to represent the lignite depolymerization, since the humic and fulvic acids released from lignite degradation imparted a dark-brown color with a maximum absorbance at 450 nm [21]. The A450 value of crude sample was defined as 100%. The relative depolymerization degree was calculated from the ratio of A450 value at the purification step and that of the crude sample. All depolymerization assays were conducted in triplicate and mean values and standard deviations were calculated. To confirm the role of esterase in lignite degradation and estimate the purification efficiency, equivalent enzyme samples (0.5 mg protein) containing esterase from different purification steps were tested for lignite depolymerization with the same method mentioned above. Effects on lignite depolymerization by the esterases with different concentrations but from the same purity were also studied,
Fig. 1. The relationship between esterase activity and lignite solubilization. A400 represented the esterase activity; A450 represented the lignite depolymerization. Values are the means and standard deviations of triplicate experiments.
in culture filtrate to 894.7 U/mg in the final purified esterase (Table 1), with an 83-fold purification and a 15% yield finally. The purified esterase showed a single band by SDS-PAGE with the molecular weight of 45 kDa (Supplementary Fig. S1). The LC-MS/MS analyses correspond to 20% protein sequence coverage in the esterase band (Fig. 2a) that exhibited 86% and 85% identity with the esterases of Penicillium expansum and Penicillium italicum reported in the NCBI database, respectively (Fig. 2b).
2.6. Analysis of depolymerization products Samples of aHA and bHA were extracted by the method described earlier [13]. The solid-state CP/MAS 13C NMR spectra of the two samples were obtained with a Bruker av-300 spectrometer (Switzerland) at a frequency of 75.47 MHz with magic angle spinning at 4 kHz with a contact time of 3 ms and a pulse delay of 5 s. Approximately 2290 scans were performed for each spectrum. Micro-FT-IR spectra of local areas of sliced specimen were measured using Nicolet Magna-IR 750 spectrophotometer (USA), connected to a Nicolet NicPlan IR microscope and a MCT detector. The resolution was 4 cm−1 and the spectral range was 4000–650 cm−1.
3.2. Enzyme properties of the esterase The final purified esterase showed the highest acetyl xylan esterase activity at pH 7.0 (Fig. 3a) and had a strong pH stability that remained 100% enzyme activity at pH 7.0 after 1 h and more than 80% activity at pH 4.0–9.0 after 1 h (Fig. 3b). The optimal temperature for the esterase was 50 °C and the relative activity was more than 90% at 40 °C (Fig. 3c). At 30 °C, it remained 100% activity after 1 h and more than 80% activity was remained at 40 °C after 1 h. But the residual activity decreased rapidly when the temperature is higher than 40 °C and only 37% of the activity was left at 50 °C after 1 h (Fig. 3d). Therefore, it was better to select 40 °C for usage, which had higher stability and enzyme activity. The purified esterase presented Km and Vmax values 0.33 mM and 4.33 μmol·(L·min)−1, respectively. The esterase activity was enhanced more than 80% by the ions NH4+, Cu2+ and Fe2+ at the concentration of 10 mM (Table 2). It was greatly inhibited by the metal ions of Ba2+, Mg2+, Fe3+ and Zn2+ (10 mM), while partially inhibited by Ca2+ and Mn2+. Besides, the esterase activity was completely inhibited by the ions Al3+ and Pb2+.
2.7. Effect of humic acid on the growth of asparagus lettuce Aliquot of 5 ml of sterile humic acid solution (400 ppm) was added in the sterile Petri dishes (Φ = 9 cm) with two pieces of filter paper, then the asparagus lettuce seeds were put onto the filter paper. Sterile water instead of humic acid was added as control. The dishes with seeds were inoculated at 37 °C for 7 days. Then the roots, stems and total length of 30 seedlings in each treatment were measured to explore the effect of humic acids on the growth of asparagus lettuce. The data sets were tested for statistical significance using ANOVA. The p values < .05 were deemed significant.
3.3. Enzymatic depolymerization of lignite
3. Results
As was shown in Fig. 4a, with the increase of purity, the depolymerization ability of samples after DEAE Sephadex A-50 and Sephadex G-75 gradually increased. The value of A450 increased from 0.091 (crude sample) to 0.44 (Sephadex G-75 purfied sample). Results in Fig. 4b and c showed the increase of A450 values along with the protein concentration increase that revealed a positive relation between the lignite depolymerization and the esterases concentrations. After the DEAE Sephadex A-50 purification, A450 increased from 0.19 to 0.95 when the concentration of protein increased from 10 to 50 mg/ml and after the Sephadex G-75 purification, A450 increased from 0.38 to 2.08 when the concentration of protein increased from 8 to 50 mg/ml. In estimation of contribution that the esterase accounted for lignite depolymerization, the relative depolymerization degree (A450) was 69% after (NH4)2SO4 precipitation and was about 40% for the final purified esterase, compared with the crude sample. These results evidenced that
3.1. Enzyme purification, molecular weight determination and amino acid sequencing In the cultures supplied with lignite, A450 increased during the first 13 days and then gradually decreased, which represents the depolymerization kinetics of lignite and was in line with the trend of esterase activity (Fig. 1). These results showed that lignite depolymerization process was correlated with esterase production by P. decumbens P6 and that the esterase played a role in the depolymerization of lignite. The esterase activity reached almost the maximum on the tenth day and the activity peak (1927 U/L) on the thirteenth day. Hence, the culture broth was collected on the tenth day for subsequent purification. After the purification procedure, the specific activity increased from 10.78 U/mg 418
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Table 1 Purification of esterase from P. decumbens P6. Purification step
Total protein (mg)
Total activity (Unit)
Specific activity (Unit/mg)
Yield (%)
Purity (fold)
Culture filtrate Ammonium sulfate DEAE Sephadex A-50 Sephadex G-75
294.0 ± 2.1 252.0 ± 1.7 7.4 ± 0.1 0.5 ± 0.0
3170.0 ± 13.8 2030.0 ± 7.4 634.0 ± 3.7 475.5 ± 5.4
10.8 ± 0.1 8.1 ± 0.1 86.2 ± 0.5 894.7 ± 10.8
100.0 ± 0.0 64.0 ± 0.2 20.0 ± 0.1 15.0 ± 0.2
1.0 ± 0.0 0.8 ± 0.0 8.0 ± 0.1 83.0 ± 1.0
The mean values of three replicates and standard deviations are presented.
4. Discussion
esterase did play a vital contribution to the lignite depolymerization.
Lignite accounts for about 13% of the total coal reserves in China [1] and its utilization is greatly restricted due to the high oxygen content, high water content and low calorific value [22]. Therefore, upgrading lignite or converting it into high value-added products is of great importance. Bioconversion of lignite by microorganisms has been considered as an eco friendly way to converting lignite into clean fuels or value-added products [23]. There are three principal mechanisms of lignite bioconversion including direct utilization, solubilization and depolymerization [6]. Solubilization refers to nonenzymatic dissolution of lignite and is mediated by alkaline compounds, chelating agents and surfactants at alkaline pH [24,25]. Depolymerization of lignite is mediated by enzymes functioned at acidic conditions (pH < 6) [6]. The involvement of oxidizing enzymes, including lignin peroxidases, manganese peroxidases and laccases in depolymerization has been investigated for a long time [26]. Aside from the peroxidases, hydrolases also contributed to coal degradation. But the participation of esterase in coal degradation has been rarely studied previously [12,27]. It was postulated that an efficient participation of a mediator was necessary
3.4. Analysis of depolymerization product In 13C NMR for the humic acids, seven regions were divided in the spectra as shown in Table 3. When lignite was tackled with purified esterase, the relative intensity of carboxyl/carbonyl carbon at 165–220 ppm decreased from 12.5 to 5.81, mainly due to a decrease in the intensity of carboxyl carbons at 176 ppm. Compared with aHA, bHA has a lower percentage of aromatic carbon and a higher percentage of aliphatic carbon. The range of 165–185 ppm represented chemical shifts of carboxyl, ester and quinine, whose changes showed the depolymerization of ester bonds in lignite. 3.5. Effect of humic acid on asparagus lettuce growth The results summarized in Table 4 demonstrated that the length of root, stem and total plant were significantly increased (from 2.77 cm, 3.38 cm and 6.15 cm to 3.44 cm, 3.58 cm and 7.01 cm) with the treatment of humic acids at 400 ppm.
Fig. 2. Mass spectrometric analysis of the purified esterase band. a, Sequence of the purified esterase with amino acids observed by LC-MS/MS in the esterase band highlighted in red; b, The observed amino acid sequence alignment of the esterase and other esterases.
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Fig. 3. Effects of pH and temperature on the purified esterase. a, The effect of pH on the esterase activity; b, pH stability of the esterase after 1 h of incubation; c, The effect of temperature on the esterase activity; d, Thermostability of the esterase after 1 h of incubation. Values are the means and standard deviations of triplicate experiments.
The fungus P. decumbens P6 could degrade a certain amount of lignite completely within 7 days [15]. With the induction of lignite, the activity of esterase increased in the culture, indicating that esterase might play a role in depolymerization of lignite [3]. In the present study, the increased degree of lignite depolymerization along with the increase of purification fold of the esterase and along with the increased concentration of esterase proved the contribution of esterase to lignite depolymerization (Fig. 4). The relative depolymerization degree by the final purified esterase was about 40% compared with that by the crude enzyme sample. A direct correlation between the esterase activity and the process of lignite depolymerization has been elaborated for the first time in this study. It could be concluded that esterase did play a key role (at least 40%) in the lignite depolymerization by P. decumbens P6 during the whole process. In addition, after the purification, the small compounds should be eliminated and the lignite depolymerization by the purified esterase implied that no low molecular weight co-factors or mediators were necessary for the esterase activity in lignite depolymerization. In our present study, the 86% and 85% identity of amino acid sequence of the esterase purified from the fungus P. decumbens P6 culture with the esterase from Penicillium expansum and Penicillium italicum, respectively (Fig. 2), demonstrated that our purified enzyme was similar, but differed from the esterases produced by the related fungi. The maintenance of more than 80% of residual esterase activity at pH 4.0 through 9.0 for 1 h indicated its strong pH stability (Fig. 3). The pH stability in alkaline condition was important for the esterase of P. decumbens P6, because lignite solubilization occurs primarily at alkaline condition [29] and some alkaline materials were found to be effective in the supernatant and the pH of the medium increased in the procedure of lignite degradation by this fungus [3]. In addition, the mechanism for enhance of esterase activity the ions NH4+, Cu2+ and Fe2+, but inhibition by Ba2+, Mg2+, Fe3+ and Zn2+, Ca2+, Mn2+, Al3+ and Pb2+ (Table 2) demonstrated in this case was not clear. However, it has been reported that NH4+ was able to help the depolymerization of some kinds polymers [30]. The Cu2+ and Fe2+ stimulation on the esterase was similar to the previous report on Drosophila repleta esterase [31]. The 13C NMR spectra have been extensively applied to the structural studies of low-rank coal. Compared with aHA, bHA has a lower
Table 2 Effect of different ions and chemicals on the esterase activity. Metal ions
NH4+ Al3+ Ba2+ Ca2+ Mn2+ Mg2+ Ag+ Cu2+ Sn2+ Fe2+ Fe3+ Co2+ Pb2+ Zn2+ SDS EDTA Urea
Relative activity (%) 1 mM
10 mM
NDa ND ND ND ND ND ND ND ND 68.3 ± 0.6 145.87 ± 2.8 ND ND ND ND ND ND
275.5 ± 4.3 ND 34.3 ± 1.2 82.3 ± 1.3 93.1 ± 0.9 78.4 ± 1.6 120.6 ± 2.5 197.0 ± 1.9 119.2 ± 3.1 284.3 ± 1.7 3.9 ± 0.2 105.8 ± 1.4 ND 63.7 ± 1.6 ND ND ND
Values represent the mean ± SD (n = 3) relative to untreated control samples. a Means no activity was detected.
for the enzymes in depolymerization of lignite [12,27], but this property was not featured for the hydrolytic enzymes. The involvement of hydrolytic enzymes in lignite depolymerization was once a matter of dispute [11]. The study on possible targets for hydrolytic enzymes in lignite structure revealed that 1–2% of the total carbon in lignite can be ascribed to carboxylic esters and ester bonds play an important role in stabilizing the structure of lignite [9]. Supaluknari et al. [28] has confirmed the presence of ester groups in Australian lignite by IR spectra. The participation of extracellular esterases in lignite depolymerization was assessed by measuring the release of 14C radioactivity from selectively alkylated carboxylic groups [11]. However, up to date the actual role of esterases in coal degradation and its mechanism are unclear [12].
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Fig. 4. Lignite depolymerization by P. decumbens P6 esterase. a, Lignite depolymerization by the esterases from different purification steps; enzyme samples (0.5 mg protein) containing esterase from different purification steps were used to depolymerize lignite; b, Lignite depolymerization by the esterase purified after DEAE Sephadex A-50; c, Lignite depolymerization by the esterase purified after Sephadex G-75; d, The contribution by the purified esterases from different purification steps to lignite depolymerization. The A450 value of crude sample was defined as 100%. The relative depolymerization degree was calculated the ratio of the A450 value of different purification steps and that of crude sample. Values are the means and standard deviations of triplicate experiments.
Table 3 Relative intensity distribution in solid-state Sample
aHA bHA
Carbonyl/carboxyl C
13
C NMR spectra of lignite. C-substituted and protonated aromatic C 135–120 ppm
Protonated aromatic C 120–90 ppm
Oxygenated aliphatic C 90–60 ppm
Methoxyl C
Aliphatic C
220–165 ppm
O-substituted aromatic C 165–135 ppm
60–50 ppm
50–0 ppm
12.5 ± 0.2 5.81 ± 0.11
4.75 ± 0.15 4.44 ± 0.12
6.77 ± 0.31 6.59 ± 0.20
4.94 ± 0.11 3.92 ± 0.09
2.2 ± 0.1 3.87 ± 0.12
1.11 ± 0.06 1.5 ± 0.1
4.97 ± 0.14 5.81 ± 0.17
The mean values of three replicates and standard deviations are presented.
acids [34,35]. In this study, the humic acid transformed by esterase showed a significantly positive effect on root, stem and total length. This result confirmed that the humic acids obtained by fungal degradation of lignite have effective bioactivity.
percentage of aromatic carbon and a higher percentage of aliphatic carbon. This may be why the smaller humic acids contained more functional groups than the larger ones [32]. The range of 165–185 ppm represented chemical shifts of carboxyl, ester and quinine, whose decreases showed the depolymerization of ester bonds in lignite. The analysis in the present study (Table 3) demonstrated the depolymerization of ester bonds in lignite and further confirmed the participation of esterase in lignite depolymerazation. Humic acids influence the growth and metabolism of plants, especially on root development [33]. After transformed by P. decumbens P6, the molecular mass of the humic acid decreased, while the oxygen and nitrogen content increased, having a better bioactivity than the crude lignite humic acids [13]. Humic acids of low molecular size are generally found to have higher bioactivity than the larger-sized humic
5. Conclusions In conclusion, the fungus P. decumbens P6 produced an esterase with high pH stability and favorable capacity of lignite depolymerization. The purified esterase could depolymerize about 40% of lignite. A direct correlation between the esterase activity and the process of lignite depolymerization was elaborated for the first time in this study, which made an important contribution to our understanding of lignite depolymerization by hydrolytic enzymes. The depolymerization product of
Table 4 Effect of humic acids on the growth of asparagus lettuce. Concentration (ppm) 0 400
Root (cm)
Stem (cm) b
2.77 ± 0.18 3.44 ± 0.11a
Total (cm) b
3.38 ± 0.10 3.58 ± 0.08a
Root/Stem b
6.15 ± 0.14 7.01 ± 0.16a
0.833 ± 0.037b 0.966 ± 0.03a
Values given are mean ± standard deviation of 30 samples; Values with different alphabets in the same column are significantly (p < .05) different from each other, according to ANOVA.
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lignite had a significantly positive effect on the growth of asparagus lettuce. Therefore, the esterase could be a potential biocatalyst in lignite depolymerization. Declarations None. Conflict of interest None. Funding This work was sponsored by the fund for Shanxi “1331 Project” Collaborative Innovation Center and special fund for Agro-scientific Research in the Public Interest (201403048-2). Acknowledgements We thank the fund for Shanxi “1331 Project” Collaborative Innovation Center and the special fund for Agro-scientific Research in the Public Interest (201403048-2). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.fuel.2017.11.035. References [1] Feng X, Zhang C, Tan P, Zhang X, Fang Q, Chen G. Experimental study of the physicochemical structure and moisture readsorption characteristics of Zhaotong lignite after hydrothermal and thermal upgrading. Fuel 2016;185:112–21. [2] Tahmasebi A, Yu J, Han Y, Zhao H, Bhattacharya S. A kinetic study of microwave and fluidized-bed drying of a Chinese lignite. Chem Eng Res Des 2014;92(1):54–65. [3] Yuan HL, Yang JS, Chen WX. Production of alkaline materials, surfactants and enzymes by Penicillium decumbens strain P6 in association with lignite degradation/ solubilization. Fuel 2006;85(10–11):1378–82. [4] Cohen MS, Gabriele PD. Degradation of coal by the fungi Polyporus Versicolor and Poria Monticola. Appl Environ Microbiol 1982;44(1):23–7. [5] Ghani MJ, Rajoka MI, Akhtar K. Investigations in fungal solubilization of coal: mechanisms and significance. Biotechnol Bioprocess Eng 2015;20(4):634–42. [6] Romanowska I, Strzelecki B, Bielecki S. Biosolubilization of polish brown coal by Gordonia alkanivorans S7 and Bacillus mycoides NS1020. Fuel Process Technol 2015;131:430–6. [7] Bugg TD, Ahmad M, Hardiman EM, Singh R. The emerging role for bacteria in lignin degradation and bio-product formation. Curr Opin Biotechnol 2011;22(3):394–400. [8] Conesa A, Punt PJ, van den Hondel CAMJJ. Fungal peroxidases: molecular aspects and applications. J Biotechnol 2002;93:143–58. [9] Groβe SA. Untersuchungen zum Chemismus der Biokonversion von Braunkohle durch kohledegradierende Deuteromyceten; 2000. [10] Hölker U, Ludwig S, Scheel T, Höfer M. Mechanisms of coal solubilization by the deuteromycetes Trichoderma atroviride and Fusarium oxysporum. Appl Microbiol Biotechnol 1999;52:57–9. [11] Hölker U, Schmiers H, Grosse S. Solubilization of low-rank coal by Trichoderma atroviride: evidence for the involvement of hydrolytic and oxidative enzymes by
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Full Length Article
An esterase from Penicillium decumbens P6 involved in lignite depolymerization Yi Yanga, Jinshui Yanga, Baozhen Lia, Entao Wangb, Hongli Yuana, a b
T
⁎
State Key Laboratory of Agrobiotechnology, College of Biological Sciences, China Agricultural University, Beijing 100193, China Departamento de Microbiología, Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional, Mexico, DF, Mexico
A R T I C L E I N F O
A B S T R A C T
Keywords: Penicillium decumbens Lignite Depolymerization Esterase Humic acid
In this study, lignite was degraded using a purified esterase from Penicillium decumbens P6 for the first time. The esterase was purified using ammonium sulfate precipitation, anion exchange and gel filtration chromatography. The recovery and purification yield of the enzyme were 15% and 83 folds, respectively. The molecular weight of the purified enzyme was about 45 kDa. Both crude and purified esterases were studied for lignite depolymerization. The tendency of depolymerization by crude enzyme was consistent with the enzyme secreted in the medium. Along with the increased purified esterase concentration from 8 to 50 mg/ml, A450 value increased from 0.38 to 2.08. The contribution of esterase to the depolymerization was about 40% in the crude supernatant. Compared with aHA (crude lignite humic acid), bHA (esterase degraded lignite humic acid) has a lower percentage of aromatic carbon and ester groups, but a higher percentage of aliphatic carbon. bHA can promote the growth of asparagus lettuce. The results demonstrated that lignite was depolymerized by the purified esterase and evidenced the potential of esterase application in conversion of lignite into compounds with high bioactivities.
1. Introduction
organic matter, corresponding to one to two ester bonds per 100 lignite carbon atoms [9]. A direct correlation between lignite solubilization and enzymatic hydrolytic activity was evidenced in the deuteromycete Trichoderma atroviride [10]. Hölker et al. [11] reported that T. atroviride could cleave both the carboxylic esters and the phenolic ether bonds during the solubilization of 14C-labelled lignite. However, unlike the peroxidases, no low molecular weight co-factors and mediators were known in the activation of esterases, and the steric hindrance might be an obstacle for esterases to permeate the macromolecular coal network and to depolymerize the coal [12]. Therefore, more research is needed to determine the actual role and the mechanism of esterases in coal degradation. In our previous work, the fungal strain Penicillium decumbens P6 capable of degrading lignite effectively presented inducible esterase activity in the lignite degradation [3], that implied a possible role of esterase in the depolymerization of lignite. In addition, the contents of humic acids and water-soluble humic materials greatly increased and the molecular mass of humic acid decreased in the fungal depolymerized lignite [13]. Moreover, the fungal transformed lignite humic acids have a better bioactivity than the crude lignite humic acids [14,15]. It is well known that humic acids contribute substantially to global soil
With a lignin-like structure, lignite is defined as a low rank coal with low calorific value, high moisture and high sulfur content, which limits its usage and conversion [1]. The direct combustion of lignite results in low thermal efficiency and low industrial profit. In addition, the piling up of lignite in the open air for a long time causes energy waste and environmental pollution [2]. The high content of humic acid and fulvic acid and the great reserve (130 million tons) of lignite in China make it a potential natural resource [3]. Hence, research on conversion of lignite into industrial and chemical products with high additional value or into liquid fuel is rationally developed [4]. Enzymatic conversion has been considered as an economic and environment-friendly way for transforming macromolecular coal into simpler and low molecular weight products [5]. Lignin peroxidases, manganese peroxidases and laccases are the most widely studied oxidative enzymes participating in the coal solubilization [6,7], and they require low molecular weight cofactors and mediators to depolymerize the lignin [8]. In addition to the oxidative enzymes, hydrolytic enzymes also showed a great potential to depolymerize lignite. In lignite, carboxylic ester groups amount to approximately 2.5–5% of the total
Abbreviations: CP/MAS 13C NMR, 13C nuclear magnetic resonance with cross-polarization/magic angle spinning; aHA, lignite humic acid; bHA, esterase degraded lignite humic acid; SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis; DEAE, Diethyl-Aminoethanol; SDS, Sodium Dodecyl Sulfonate; EDTA, Ethylene Diamine Tetraacetic Acid ⁎ Corresponding author at: Center of Life Science, China Agriculture University, No.2 Yuanmingyuan West Road, Haidian District, Beijing 100193, China. E-mail addresses: [email protected] (Y. Yang), [email protected] (J. Yang), [email protected] (B. Li), [email protected] (H. Yuan). https://doi.org/10.1016/j.fuel.2017.11.035 Received 16 August 2017; Received in revised form 26 October 2017; Accepted 11 November 2017 0016-2361/ © 2017 Elsevier Ltd. All rights reserved.
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fertility and have been widely utilized in agriculture, for example, they can promote plant growth and metabolism [16] and reduce some plant diseases [17]. To prove the roles of esterase in the enzymatic attack on lignite, we performed the present study using purified esterase from P. decumbens P6 culture. The crude lignite humic acids (aHA) and esterase degraded lignite humic acids (bHA) were quantified and characterized using 13C nuclear magnetic resonance with cross-polarization/magic angle spinning (CP/MAS 13C NMR) and Fourier transformed infrared spectrophotometry (FT-IR). The humic acids produced by esterase depolymerization presented a significantly positive effect on the growth of asparagus lettuce. These results evidenced the key role of esterase for P. decumbens P6 in hydrolyzing carboxyl within the lignite structure.
The dried samples were dissolved in 1 ml of 50 mM phosphate buffer (pH 7.0) and were loaded onto a Sephadex G-75 gel filtration column (GE) that had been pre-equilibrated with the same phosphate buffer. The column was eluted with 50 mM phosphate buffer (pH 7.0) at a flow rate of 1.5 ml/min. The elute was collected into 1 ml fractions for esterase activity detection as mentioned previously [18]. The active fractions were loaded onto the Sephadex G-75 gel filtration column again for further purification. The molecular weight of the purified esterase was determined by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).
2. Materials and methods
Protein sequencing was performed according to the method [19]. The acquired amino acid sequence was used in BLAST searching (https://blast.ncbi.nlm.nih.gov/Blast.cgi) for extracting the related sequences in the database. The sequence was aligned together with the extracted reference sequences by MEGA 6.06 software [20].
2.3. Protein sequencing
2.1. Lignite samples and fungal strain Lignite sample was collected from the Huolingele Minerals Administration Coalmine, Inner Mongolia of China. Air-dried lignite sample was pulverized and sieved with a 70 mesh screen. The sample preparation and elemental analysis of the crude lignite humic acid were performed according to the previous reported methods [13]. The fungal strain P. decumbens P6 (CGMCC 0866) used throughout the study was maintained at 4 °C on the yeast extract – malt agar (YMPG) slants, consisting of: glucose, 10 g; malt extract, 10 g; peptone, 2 g; yeast extract, 2 g; asparagines, 1 g; KH2PO4·3H2O, 2 g and MgSO4·7H2O, 1 g in 1 l distilled water. All of the chemicals used in this study were reagent grade purchased from the Beijing Chemicals and Reagent Corp., China, unless otherwise stated. The fungus was cultured on YMPG agar plates at 28 °C for 1 week, then the spores were washed and suspended in 0.9% of sodium chloride solution. Aliquots of 2 ml of the spore suspension (106 spores/ml) were inoculated into 500 ml flasks containing 200 ml liquid medium (10 g wheat bran; 5 g urea; 2 g KH2PO4·3H2O; 1 g MgSO4·7H2O and 1 g asparagine in 1 l of distilled water) which had been optimized for esterase production. Powdered lignite was added into the medium at the ratio of 1% (w/v) as an inducer for enzyme production. The inoculated flasks were cultivated by shaking at 150 rpm and 28 °C for 10 days. Culture samples in aliquots of 1 ml were taken every day, and centrifuged at 4 °C, 8000 rpm for 10 min. Supernatants were used for analyzing the esterase activity with the method [18]. When the esterase activity reached the maximum level (after 10 days of incubation), whole cultures were harvested for purification studies as mentioned subsequently.
2.4. Enzyme properties The esterase activity was determined by the spectrophotometric method [18]. Briefly, 50 μL supernatant of the culture was mixed with 950 μL 4-nitrophenyl acetate in 50 mM phosphate buffer (pH 7.0). The reaction was performed at 37 °C for 15 min. A400 (absorbance at 400 nm) was determined with a UV-1800 spectrophotometer (Shimadzu, Japan) to estimate the released p-nitrophenol. In control, 50 μL of the supernatant boiled for 30 min were used. One unit of esterase activity was defined as the amount of proteins liberating 1 µmol p-nitrophenol per min under the defined conditions. The optimum pH for esterase was tested at 37 °C in 50 mM citric acid buffer (pH 3.0, 4.0 and 5.0) and 50 mM phosphate buffer (pH 6.0, 7.0, 8.0 and 9.0). The optimum temperature for esterase was determined by standard assay ranging from 30 to 80 °C in the 50 mM phosphate buffer (pH 7.0). The results were expressed as relative activity to the value obtained at either optimal pH assay or optimal temperature assay. The pH stability of esterase was determined by measuring the residual activity after incubating the enzyme at 37 °C for 1 h in the aforementioned buffer (pH 3.0–9.0). In the thermostability assay, the enzyme solution was incubated in 50 mM phosphate buffer (pH 7.0) at temperatures ranging from 30 to 80 °C for 1 h, and the remaining activity was measured under the optimum conditions. The esterase activity obtained under the optimum conditions was defined as 100%. The Michaelis-Menten constant (Km) and maximal velocity (Vmax) of the purified esterase were determined by measuring the esterase activity at various 4-nitrophenyl acetate concentrations (0.2 mM, 0.4 mM, 0.8 mM, 1.6 mM and 3.2 mM). The Km and Vmax of the enzyme were calculated using the Lineweaver-Burk plot constructed by plotting the reciprocal of the substrate concentration on the X-axis and the reciprocal of the esterase reaction velocity on the Y-axis. All determinations were conducted in triplicate. The effects of metal ions and chemical reagents on the esterase activity were determined. Different metal ions and chemical reagents such as NH4+, Al3+, Ba2+, Ca2+, Mn2+, Mg2+, Ag+, Cu2+, Sn2+, Fe2+, Co2+, Pb2+, Zn2+, Fe3+, urea, SDS (Sodium Dodecyl Sulfonate) and EDTA (Ethylene Diamine Tetraacetic Acid) at 10 mM and 1 mM were added to the purified enzyme solution and then incubated at 4 °C for 2 h. The residual activities were determined under the standard assay conditions. The esterase activity of the enzyme in culture without addition of metal ions or chemical reagents was defined as 100%. Protein concentration was determined using the Bradford Protein Assay Kit (GenStar, Beijing, China) according to the manufacturer’s instructions.
2.2. Purification of esterase and determination of molecular weight After 10 days of cultivation, the supernatant was separated from the mycelium by centrifugation of the culture at 8000 rpm for 20 min at 4 °C, and then filtration with Whatman No. 1 filter paper. Proteins in the cell-free extract were precipitated at 4 °C with ammonium sulfate (80%) for about 2 h following by centrifugation at 8000 rpm for 20 min under 4 °C. The sediment was resuspended in 20 ml of 20 mM Tris-HCl buffer (pH 8.0) and dialyzed against 2 l of the same buffer for 24 h to remove (NH4)2SO4. Then, the protein solution was loaded on the columns listed bellow to separate and purify acetyl xylan esterase. Firstly, the protein solution was loaded on a DEAE (DiethylAminoethanol) Sepharose anion exchange column (GE) pre-equilibrated with 20 mM Tris-HCl buffer (pH 8.0). The column was washed with the same buffer until A280 (absorbance at 280 nm) of effluent became zero and then eluted with a linear NaCl gradient (0–1 M) at a flow rate of 1.5 ml/min. The elute was collected into fractions of 1 ml. Esterase activity in the fractions was detected with the previously described method [18] and fractions presenting esterase activity were pooled and lyophilized for the next step of purification. 417
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2.5. Lignite depolymerization Crude enzyme was obtained from the daily sample during the incubation by centrifugation of the fermentation medium at 8000 rpm for 10 min at 4 °C. The depolymerization assay was carried out at 5% (w/v) sterile lignite loading in 0.1 M phosphate buffer (pH 7.0) in a final volume of 1 ml. Depolymerization assays were performed at a total esterase loading of 10 U/g lignite in an orbital shaker incubator at 28 °C for 24 h. Control assays including esterase sample without lignite and lignite without esterase sample were performed under the same conditions. The supernatants were collected by centrifugation at 8000 rpm for 10 min at 4 °C and then A450 (absorbance at 450 nm) was measured to represent the lignite depolymerization, since the humic and fulvic acids released from lignite degradation imparted a dark-brown color with a maximum absorbance at 450 nm [21]. The A450 value of crude sample was defined as 100%. The relative depolymerization degree was calculated from the ratio of A450 value at the purification step and that of the crude sample. All depolymerization assays were conducted in triplicate and mean values and standard deviations were calculated. To confirm the role of esterase in lignite degradation and estimate the purification efficiency, equivalent enzyme samples (0.5 mg protein) containing esterase from different purification steps were tested for lignite depolymerization with the same method mentioned above. Effects on lignite depolymerization by the esterases with different concentrations but from the same purity were also studied,
Fig. 1. The relationship between esterase activity and lignite solubilization. A400 represented the esterase activity; A450 represented the lignite depolymerization. Values are the means and standard deviations of triplicate experiments.
in culture filtrate to 894.7 U/mg in the final purified esterase (Table 1), with an 83-fold purification and a 15% yield finally. The purified esterase showed a single band by SDS-PAGE with the molecular weight of 45 kDa (Supplementary Fig. S1). The LC-MS/MS analyses correspond to 20% protein sequence coverage in the esterase band (Fig. 2a) that exhibited 86% and 85% identity with the esterases of Penicillium expansum and Penicillium italicum reported in the NCBI database, respectively (Fig. 2b).
2.6. Analysis of depolymerization products Samples of aHA and bHA were extracted by the method described earlier [13]. The solid-state CP/MAS 13C NMR spectra of the two samples were obtained with a Bruker av-300 spectrometer (Switzerland) at a frequency of 75.47 MHz with magic angle spinning at 4 kHz with a contact time of 3 ms and a pulse delay of 5 s. Approximately 2290 scans were performed for each spectrum. Micro-FT-IR spectra of local areas of sliced specimen were measured using Nicolet Magna-IR 750 spectrophotometer (USA), connected to a Nicolet NicPlan IR microscope and a MCT detector. The resolution was 4 cm−1 and the spectral range was 4000–650 cm−1.
3.2. Enzyme properties of the esterase The final purified esterase showed the highest acetyl xylan esterase activity at pH 7.0 (Fig. 3a) and had a strong pH stability that remained 100% enzyme activity at pH 7.0 after 1 h and more than 80% activity at pH 4.0–9.0 after 1 h (Fig. 3b). The optimal temperature for the esterase was 50 °C and the relative activity was more than 90% at 40 °C (Fig. 3c). At 30 °C, it remained 100% activity after 1 h and more than 80% activity was remained at 40 °C after 1 h. But the residual activity decreased rapidly when the temperature is higher than 40 °C and only 37% of the activity was left at 50 °C after 1 h (Fig. 3d). Therefore, it was better to select 40 °C for usage, which had higher stability and enzyme activity. The purified esterase presented Km and Vmax values 0.33 mM and 4.33 μmol·(L·min)−1, respectively. The esterase activity was enhanced more than 80% by the ions NH4+, Cu2+ and Fe2+ at the concentration of 10 mM (Table 2). It was greatly inhibited by the metal ions of Ba2+, Mg2+, Fe3+ and Zn2+ (10 mM), while partially inhibited by Ca2+ and Mn2+. Besides, the esterase activity was completely inhibited by the ions Al3+ and Pb2+.
2.7. Effect of humic acid on the growth of asparagus lettuce Aliquot of 5 ml of sterile humic acid solution (400 ppm) was added in the sterile Petri dishes (Φ = 9 cm) with two pieces of filter paper, then the asparagus lettuce seeds were put onto the filter paper. Sterile water instead of humic acid was added as control. The dishes with seeds were inoculated at 37 °C for 7 days. Then the roots, stems and total length of 30 seedlings in each treatment were measured to explore the effect of humic acids on the growth of asparagus lettuce. The data sets were tested for statistical significance using ANOVA. The p values < .05 were deemed significant.
3.3. Enzymatic depolymerization of lignite
3. Results
As was shown in Fig. 4a, with the increase of purity, the depolymerization ability of samples after DEAE Sephadex A-50 and Sephadex G-75 gradually increased. The value of A450 increased from 0.091 (crude sample) to 0.44 (Sephadex G-75 purfied sample). Results in Fig. 4b and c showed the increase of A450 values along with the protein concentration increase that revealed a positive relation between the lignite depolymerization and the esterases concentrations. After the DEAE Sephadex A-50 purification, A450 increased from 0.19 to 0.95 when the concentration of protein increased from 10 to 50 mg/ml and after the Sephadex G-75 purification, A450 increased from 0.38 to 2.08 when the concentration of protein increased from 8 to 50 mg/ml. In estimation of contribution that the esterase accounted for lignite depolymerization, the relative depolymerization degree (A450) was 69% after (NH4)2SO4 precipitation and was about 40% for the final purified esterase, compared with the crude sample. These results evidenced that
3.1. Enzyme purification, molecular weight determination and amino acid sequencing In the cultures supplied with lignite, A450 increased during the first 13 days and then gradually decreased, which represents the depolymerization kinetics of lignite and was in line with the trend of esterase activity (Fig. 1). These results showed that lignite depolymerization process was correlated with esterase production by P. decumbens P6 and that the esterase played a role in the depolymerization of lignite. The esterase activity reached almost the maximum on the tenth day and the activity peak (1927 U/L) on the thirteenth day. Hence, the culture broth was collected on the tenth day for subsequent purification. After the purification procedure, the specific activity increased from 10.78 U/mg 418
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Table 1 Purification of esterase from P. decumbens P6. Purification step
Total protein (mg)
Total activity (Unit)
Specific activity (Unit/mg)
Yield (%)
Purity (fold)
Culture filtrate Ammonium sulfate DEAE Sephadex A-50 Sephadex G-75
294.0 ± 2.1 252.0 ± 1.7 7.4 ± 0.1 0.5 ± 0.0
3170.0 ± 13.8 2030.0 ± 7.4 634.0 ± 3.7 475.5 ± 5.4
10.8 ± 0.1 8.1 ± 0.1 86.2 ± 0.5 894.7 ± 10.8
100.0 ± 0.0 64.0 ± 0.2 20.0 ± 0.1 15.0 ± 0.2
1.0 ± 0.0 0.8 ± 0.0 8.0 ± 0.1 83.0 ± 1.0
The mean values of three replicates and standard deviations are presented.
4. Discussion
esterase did play a vital contribution to the lignite depolymerization.
Lignite accounts for about 13% of the total coal reserves in China [1] and its utilization is greatly restricted due to the high oxygen content, high water content and low calorific value [22]. Therefore, upgrading lignite or converting it into high value-added products is of great importance. Bioconversion of lignite by microorganisms has been considered as an eco friendly way to converting lignite into clean fuels or value-added products [23]. There are three principal mechanisms of lignite bioconversion including direct utilization, solubilization and depolymerization [6]. Solubilization refers to nonenzymatic dissolution of lignite and is mediated by alkaline compounds, chelating agents and surfactants at alkaline pH [24,25]. Depolymerization of lignite is mediated by enzymes functioned at acidic conditions (pH < 6) [6]. The involvement of oxidizing enzymes, including lignin peroxidases, manganese peroxidases and laccases in depolymerization has been investigated for a long time [26]. Aside from the peroxidases, hydrolases also contributed to coal degradation. But the participation of esterase in coal degradation has been rarely studied previously [12,27]. It was postulated that an efficient participation of a mediator was necessary
3.4. Analysis of depolymerization product In 13C NMR for the humic acids, seven regions were divided in the spectra as shown in Table 3. When lignite was tackled with purified esterase, the relative intensity of carboxyl/carbonyl carbon at 165–220 ppm decreased from 12.5 to 5.81, mainly due to a decrease in the intensity of carboxyl carbons at 176 ppm. Compared with aHA, bHA has a lower percentage of aromatic carbon and a higher percentage of aliphatic carbon. The range of 165–185 ppm represented chemical shifts of carboxyl, ester and quinine, whose changes showed the depolymerization of ester bonds in lignite. 3.5. Effect of humic acid on asparagus lettuce growth The results summarized in Table 4 demonstrated that the length of root, stem and total plant were significantly increased (from 2.77 cm, 3.38 cm and 6.15 cm to 3.44 cm, 3.58 cm and 7.01 cm) with the treatment of humic acids at 400 ppm.
Fig. 2. Mass spectrometric analysis of the purified esterase band. a, Sequence of the purified esterase with amino acids observed by LC-MS/MS in the esterase band highlighted in red; b, The observed amino acid sequence alignment of the esterase and other esterases.
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Fig. 3. Effects of pH and temperature on the purified esterase. a, The effect of pH on the esterase activity; b, pH stability of the esterase after 1 h of incubation; c, The effect of temperature on the esterase activity; d, Thermostability of the esterase after 1 h of incubation. Values are the means and standard deviations of triplicate experiments.
The fungus P. decumbens P6 could degrade a certain amount of lignite completely within 7 days [15]. With the induction of lignite, the activity of esterase increased in the culture, indicating that esterase might play a role in depolymerization of lignite [3]. In the present study, the increased degree of lignite depolymerization along with the increase of purification fold of the esterase and along with the increased concentration of esterase proved the contribution of esterase to lignite depolymerization (Fig. 4). The relative depolymerization degree by the final purified esterase was about 40% compared with that by the crude enzyme sample. A direct correlation between the esterase activity and the process of lignite depolymerization has been elaborated for the first time in this study. It could be concluded that esterase did play a key role (at least 40%) in the lignite depolymerization by P. decumbens P6 during the whole process. In addition, after the purification, the small compounds should be eliminated and the lignite depolymerization by the purified esterase implied that no low molecular weight co-factors or mediators were necessary for the esterase activity in lignite depolymerization. In our present study, the 86% and 85% identity of amino acid sequence of the esterase purified from the fungus P. decumbens P6 culture with the esterase from Penicillium expansum and Penicillium italicum, respectively (Fig. 2), demonstrated that our purified enzyme was similar, but differed from the esterases produced by the related fungi. The maintenance of more than 80% of residual esterase activity at pH 4.0 through 9.0 for 1 h indicated its strong pH stability (Fig. 3). The pH stability in alkaline condition was important for the esterase of P. decumbens P6, because lignite solubilization occurs primarily at alkaline condition [29] and some alkaline materials were found to be effective in the supernatant and the pH of the medium increased in the procedure of lignite degradation by this fungus [3]. In addition, the mechanism for enhance of esterase activity the ions NH4+, Cu2+ and Fe2+, but inhibition by Ba2+, Mg2+, Fe3+ and Zn2+, Ca2+, Mn2+, Al3+ and Pb2+ (Table 2) demonstrated in this case was not clear. However, it has been reported that NH4+ was able to help the depolymerization of some kinds polymers [30]. The Cu2+ and Fe2+ stimulation on the esterase was similar to the previous report on Drosophila repleta esterase [31]. The 13C NMR spectra have been extensively applied to the structural studies of low-rank coal. Compared with aHA, bHA has a lower
Table 2 Effect of different ions and chemicals on the esterase activity. Metal ions
NH4+ Al3+ Ba2+ Ca2+ Mn2+ Mg2+ Ag+ Cu2+ Sn2+ Fe2+ Fe3+ Co2+ Pb2+ Zn2+ SDS EDTA Urea
Relative activity (%) 1 mM
10 mM
NDa ND ND ND ND ND ND ND ND 68.3 ± 0.6 145.87 ± 2.8 ND ND ND ND ND ND
275.5 ± 4.3 ND 34.3 ± 1.2 82.3 ± 1.3 93.1 ± 0.9 78.4 ± 1.6 120.6 ± 2.5 197.0 ± 1.9 119.2 ± 3.1 284.3 ± 1.7 3.9 ± 0.2 105.8 ± 1.4 ND 63.7 ± 1.6 ND ND ND
Values represent the mean ± SD (n = 3) relative to untreated control samples. a Means no activity was detected.
for the enzymes in depolymerization of lignite [12,27], but this property was not featured for the hydrolytic enzymes. The involvement of hydrolytic enzymes in lignite depolymerization was once a matter of dispute [11]. The study on possible targets for hydrolytic enzymes in lignite structure revealed that 1–2% of the total carbon in lignite can be ascribed to carboxylic esters and ester bonds play an important role in stabilizing the structure of lignite [9]. Supaluknari et al. [28] has confirmed the presence of ester groups in Australian lignite by IR spectra. The participation of extracellular esterases in lignite depolymerization was assessed by measuring the release of 14C radioactivity from selectively alkylated carboxylic groups [11]. However, up to date the actual role of esterases in coal degradation and its mechanism are unclear [12].
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Fig. 4. Lignite depolymerization by P. decumbens P6 esterase. a, Lignite depolymerization by the esterases from different purification steps; enzyme samples (0.5 mg protein) containing esterase from different purification steps were used to depolymerize lignite; b, Lignite depolymerization by the esterase purified after DEAE Sephadex A-50; c, Lignite depolymerization by the esterase purified after Sephadex G-75; d, The contribution by the purified esterases from different purification steps to lignite depolymerization. The A450 value of crude sample was defined as 100%. The relative depolymerization degree was calculated the ratio of the A450 value of different purification steps and that of crude sample. Values are the means and standard deviations of triplicate experiments.
Table 3 Relative intensity distribution in solid-state Sample
aHA bHA
Carbonyl/carboxyl C
13
C NMR spectra of lignite. C-substituted and protonated aromatic C 135–120 ppm
Protonated aromatic C 120–90 ppm
Oxygenated aliphatic C 90–60 ppm
Methoxyl C
Aliphatic C
220–165 ppm
O-substituted aromatic C 165–135 ppm
60–50 ppm
50–0 ppm
12.5 ± 0.2 5.81 ± 0.11
4.75 ± 0.15 4.44 ± 0.12
6.77 ± 0.31 6.59 ± 0.20
4.94 ± 0.11 3.92 ± 0.09
2.2 ± 0.1 3.87 ± 0.12
1.11 ± 0.06 1.5 ± 0.1
4.97 ± 0.14 5.81 ± 0.17
The mean values of three replicates and standard deviations are presented.
acids [34,35]. In this study, the humic acid transformed by esterase showed a significantly positive effect on root, stem and total length. This result confirmed that the humic acids obtained by fungal degradation of lignite have effective bioactivity.
percentage of aromatic carbon and a higher percentage of aliphatic carbon. This may be why the smaller humic acids contained more functional groups than the larger ones [32]. The range of 165–185 ppm represented chemical shifts of carboxyl, ester and quinine, whose decreases showed the depolymerization of ester bonds in lignite. The analysis in the present study (Table 3) demonstrated the depolymerization of ester bonds in lignite and further confirmed the participation of esterase in lignite depolymerazation. Humic acids influence the growth and metabolism of plants, especially on root development [33]. After transformed by P. decumbens P6, the molecular mass of the humic acid decreased, while the oxygen and nitrogen content increased, having a better bioactivity than the crude lignite humic acids [13]. Humic acids of low molecular size are generally found to have higher bioactivity than the larger-sized humic
5. Conclusions In conclusion, the fungus P. decumbens P6 produced an esterase with high pH stability and favorable capacity of lignite depolymerization. The purified esterase could depolymerize about 40% of lignite. A direct correlation between the esterase activity and the process of lignite depolymerization was elaborated for the first time in this study, which made an important contribution to our understanding of lignite depolymerization by hydrolytic enzymes. The depolymerization product of
Table 4 Effect of humic acids on the growth of asparagus lettuce. Concentration (ppm) 0 400
Root (cm)
Stem (cm) b
2.77 ± 0.18 3.44 ± 0.11a
Total (cm) b
3.38 ± 0.10 3.58 ± 0.08a
Root/Stem b
6.15 ± 0.14 7.01 ± 0.16a
0.833 ± 0.037b 0.966 ± 0.03a
Values given are mean ± standard deviation of 30 samples; Values with different alphabets in the same column are significantly (p < .05) different from each other, according to ANOVA.
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lignite had a significantly positive effect on the growth of asparagus lettuce. Therefore, the esterase could be a potential biocatalyst in lignite depolymerization. Declarations None. Conflict of interest None. Funding This work was sponsored by the fund for Shanxi “1331 Project” Collaborative Innovation Center and special fund for Agro-scientific Research in the Public Interest (201403048-2). Acknowledgements We thank the fund for Shanxi “1331 Project” Collaborative Innovation Center and the special fund for Agro-scientific Research in the Public Interest (201403048-2). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.fuel.2017.11.035. References [1] Feng X, Zhang C, Tan P, Zhang X, Fang Q, Chen G. Experimental study of the physicochemical structure and moisture readsorption characteristics of Zhaotong lignite after hydrothermal and thermal upgrading. Fuel 2016;185:112–21. [2] Tahmasebi A, Yu J, Han Y, Zhao H, Bhattacharya S. A kinetic study of microwave and fluidized-bed drying of a Chinese lignite. Chem Eng Res Des 2014;92(1):54–65. [3] Yuan HL, Yang JS, Chen WX. Production of alkaline materials, surfactants and enzymes by Penicillium decumbens strain P6 in association with lignite degradation/ solubilization. Fuel 2006;85(10–11):1378–82. [4] Cohen MS, Gabriele PD. Degradation of coal by the fungi Polyporus Versicolor and Poria Monticola. Appl Environ Microbiol 1982;44(1):23–7. [5] Ghani MJ, Rajoka MI, Akhtar K. Investigations in fungal solubilization of coal: mechanisms and significance. Biotechnol Bioprocess Eng 2015;20(4):634–42. [6] Romanowska I, Strzelecki B, Bielecki S. Biosolubilization of polish brown coal by Gordonia alkanivorans S7 and Bacillus mycoides NS1020. Fuel Process Technol 2015;131:430–6. [7] Bugg TD, Ahmad M, Hardiman EM, Singh R. The emerging role for bacteria in lignin degradation and bio-product formation. Curr Opin Biotechnol 2011;22(3):394–400. [8] Conesa A, Punt PJ, van den Hondel CAMJJ. Fungal peroxidases: molecular aspects and applications. J Biotechnol 2002;93:143–58. [9] Groβe SA. Untersuchungen zum Chemismus der Biokonversion von Braunkohle durch kohledegradierende Deuteromyceten; 2000. [10] Hölker U, Ludwig S, Scheel T, Höfer M. Mechanisms of coal solubilization by the deuteromycetes Trichoderma atroviride and Fusarium oxysporum. Appl Microbiol Biotechnol 1999;52:57–9. [11] Hölker U, Schmiers H, Grosse S. Solubilization of low-rank coal by Trichoderma atroviride: evidence for the involvement of hydrolytic and oxidative enzymes by
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