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Purification and biochemical characteristics of a novel fructosyltransferase with a high FOS transfructosylation activity from Aspergillus oryzae S719
Protein Expression and Purification 167 (2020) 105549
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Purification and biochemical characteristics of a novel fructosyltransferase with a high FOS transfructosylation activity from Aspergillus oryzae S719
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Susu Hana,b, Tong Yea,b, Shuo Lenga,b, Lixia Panc, Wei Zenga,b, Guiguang Chena,b,∗∗, Zhiqun Lianga,b,∗ a
State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources, Guangxi Microorganism and Enzyme Research Center of Engineering Technology, China b College of Life Science and Technology, Guangxi University, Nanning, 530004, Guangxi, China c National Engineering Research Center for Non-Food Biorefinery, State Key Laboratory of Non-Food Biomass and Enzyme Technology, Guangxi Key Laboratory of Biorefinery, Guangxi Biomass, Engineering Technology Research Center, Guangxi Academy of Sciences, 98 Daling Road, Nanning, 530007, China
ARTICLE INFO
ABSTRACT
Keywords: Fructosyltransferase Aspergillus oryzae Enzyme purification Fructooligosaccharides production
Fructooligosaccharides (FOS) have widely used for the manufacture of low-calorie and functional foods, because they can inhibit intestinal pathogenic microorganism growth and increase the absorption of Ca2+ and Mg2+. In this study, the novel fructosyltransferase (FTase) from Aspergillus oryzae strain S719 was successfully purified and characterized. The specific activity of the final purified material was 4200 mg−1 with purification ratio of 66 times and yield of 26%. The molecular weight of FTase of A. oryzae S719 was around 95 kDa by SDS-PAGE, which was identified as a type of FTase by Mass Spectrometry (MS). The purified FTase had optimum temperature and pH of 55 °C and 6.0, respectively. The FTase showed to be stable with more than 80% of its original activity at room temperature after 12 h and maintaining activity above 90% at pH 4.0–11.0. The Km and kcat values of the FTase were 310 mmol L−1 and 2.0 × 103 min−1, respectively. The FTase was activated by 5 mmol L−1 Mg2+ and 10 mmol L−1 Na+ (relative activity of 116 and 114%, respectively), indicating that the enzyme was Mg2+ and Na+ dependent. About 64% of FOS was obtained by the purified FTase under 500 g L−1 sucrose within 4 h of reaction time, which was the shortest reaction time to be reported regarding the purified enzyme production of FOS. Together, these results indicated that the FTase of A. oryzae S719 is an excellent candidate for the industrial production of FOS.
1. Introduction Fructooligosaccharides (FOS) are widely used as a bioactive ingredient in functional foods because it represents a major class of prebiotics [1]. FOS are a short-chain fructosyl unit (F) that is primarily linked to the terminal sucrose molecule (GF) at the reducing end. Commercial FOS formulations exhibit a low degree of polymerization, consisting primarily of 1-kestose (GF2), 1-nystose (GF3) and 1-fructofuranosylnystose (GF4). In recent years, they have been receiving increasing attention because they are stable in the range of pH 4.0–11.0, and stored for a long time at low temperatures, and their sweetness is only one-third of sucrose. In addition, their thermal stability and viscosity are higher than sucrose [2–4]. Most importantly, they have many
physiological functions such as inhibiting the growth of intestinal pathogenic microorganisms and increasing adsorption of Ca2+ and Mg2+ [5,6]. So far, commercially, the production of FOS has been primarily carried out from sucrose by the transfructosylation activity of fructofuranosidases (EC 2.4.1.9)(FTase). Based on overall amino acid sequence similarities, FTase belongs to the glycosyl hydrolase family 32 (GH32) and shares a common three-dimensional (3-D) structure with other GH32 members [7,8]. The enzyme reaction transfructosylation mechanism (Fig. 1) can be expressed as (Eq. (1)): F−R + E ↔ F−E + R F−E + acceptor → F−acceptor + E
(1)
Where F is fructose, E is transfructosylation and R represents a
∗ Corresponding author. State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources; Guangxi Microorganism and Enzyme Research Center of Engineering Technology; College of Life Science and Technology; Guangxi University, 100 Daxue Road, Nanning 530004, Guangxi, China. ∗∗ Corresponding author. State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources; Guangxi Microorganism and Enzyme Research Center of Engineering Technology; College of Life Science and Technology; Guangxi University, 100 Daxue Road, Nanning, 530004, Guangxi, China. E-mail addresses: [email protected] (G. Chen), [email protected] (Z. Liang).
https://doi.org/10.1016/j.pep.2019.105549 Received 24 July 2019; Received in revised form 28 November 2019; Accepted 28 November 2019 Available online 02 December 2019 1046-5928/ © 2019 Elsevier Inc. All rights reserved.
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steps towards understanding some of its properties. In this study, the purification and characterization of a fructofuranosidases (FTase) from A. oryzae S719 were described. In addition to the broad enzymatic properties, this enzyme displayed a high FOS synthesis efficiency with a FOS yield of approximately 64% within 4 h of reaction, which demonstrated that this enzyme might be a potential biocatalyst for the synthesis of FOS in industry. 2. Materials and methods 2.1. Strains and materials A. oryzae ZT65 (CCTCCM) was stored in the Culture and Information Center of Industrial Microorganism of China Universities. Mycelium was grown at 37 °C for 48 h on solid medium (sucrose 20 g L−1, agar 20 g L−1 and NaCl 7 g L−1, pH 5.5). Its spores were collected with 0.85% sterile saline and prepared as 10−8 spores ml−1 suspension. The A. oryzae ZT65 spore solution was applied to the plate medium, cultured in a 37 °C incubator for 4 h, and continuously irradiated for 50 s using an ultraviolet lamp (15 W of power, effective wavelength 254 nm, distance 30 cm). It was induced by UV to a positive mutant strain A. oryzae S719 with significantly improved enzyme activity, which was identified as the target strain (supplementary material 1). They were stored on solid medium composed of 250 g L−1 sucrose, 30 g L−1 maceration extract of bran, 5 g L−1 LiCl, and 20 g L−1 agar. FOS standards 1-kestose (GF2), 1-nystose (GF3) and 1-fructofuranosylnystose (GF4) were of biochemical grade obtained from Wako Pure Chemical Industries (Chuo-Ku, Osaka, Japan). Acetonitrile as mobile phase used for HPLC was obtained from Merck (Germany). Wheat bran was purchased from Guangdong baiyan grain and oil industry Co. (Guangdong, China). Corn steep powder was obtained from Shandong Kangyuan Bio-Tech Co., Ltd. (Yuncheng, China). All other chemicals were of analytical grade.
Fig. 1. Network of the reaction mechanism for the production of FOS Network of the reaction mechanism for the production of FOS from sucrose catalyzed by fructosyltransferase: G, GF, GF2, GF3, and GF4, means glucose, sucrose, I-kestose, nystose, and lF- fructofuranosyl nystose, respectively. Where F is fructose, E is fructosyltransferase and R represents sucrose.
carbonyl of an aldose. According to this mechanism, one molecule of sucrose serves as a donor and another acts as an acceptor for GF2 synthesis, releasing one molecule of glucose, for the production of GF3, the GF2 acts as an acceptor [7]. According to the report of Trollope et al. [9], FTase can be sub-classified into the two groups with low and high FOS synthesis activity, possessing either predominantly hydrolytic or high FTase activity. The theoretical yield of FOS from sucrose is 75% if 1-kestose is the only FOS produced. However, the actual FOS yields were considerably lower than 60% with A. niger ATCC 20611 [10] and 53–59% with Aureobasidium sp. [11–13]. One explanation for this is that the enzymes involved in FOS production are inhibited by the liberated glucose that accumulates as a by-product [14]. It is well known that the low activity of the wild-type FTase makes them unsuitable for large-scale industrial application in FOS production. For this reason, many studies have attempted to produce FTase and FOS using different recombinant strains with high levels of FOS synthesis, such as Pichia pastoris GS115 [15]. However, the fungus stable system of Aspergillus is often used to produce FOS industrially, especially the FopA enzyme from A. niger ATCC 20611 [16]. It can be seen that the stability and practicability of the fungal system still exerts great potential in FOS production. In the past two decades, a lot of studies have been carried out to improve the efficiency of FTase production, but little has been known about FTase studies with high-level FOS synthesis from nonrecombinant strains. In nature, there are wide range of FTase resources from microorganisms, but a small part of which can be used in industry. Common enzyme preparations are vulnerable to extreme environments in industrial production, such as strong acid, alkali, high pressure and high temperature conditions. The industrial application of FTase in the production of FOS is highly limited due to inherent properties of FTase, such as low FTase activity (most important), low transfructosylation activity, low catalytic efficiency and poor stability. Several microorganisms possessing FTase activity including Aspergillus niger, Aspergillus oryzae, Aspergillus japonicus, Neurospora crassa, Aureobasidium sp., Fusariumoxys porum, Arthrobacter sp. and yeast have been reported [17–20]. Simultaneously, purification and characterization of FTase from various sources have been studied [21–24]. However, this accumulated knowledge is confusing [21–25]. The information differs from one source to another, from one microorganism to another, even from one strain to another, thereby making it imperative to purify the enzyme from each source. The purification and partial characterization of the enzyme were undertaken as necessary
2.2. Fermentation of A. oryzae S719 The inoculum was developed by transferring a loopful of mycelia from a 3-day-old slant of A. oryzae S719 stored on a bevel into the inoculum medium (10 g bran plus 5 ml water). The 250 ml flasks were incubated for 60 h at 37 ± 1 °C. After that, the spores were washed with 120 ml of sterile water, the bran was filtered off, and the spores were shaken at 200 rpm for 10 min to reach the spore suspensions (108 spores). The inoculum was transferred into 50 ml of fermentation medium consisting of the following (% w/v): sucrose: 18; corn steep powder 4; ZnSO4·7H20 0.2, (initial pH of 5.5). The culture was incubated for 40 h at 34 ± 1 °C on a rotary shaker at 240 rpm. 2.3. Assays of enzyme activity (standard activity assay) and protein concentration The supernatant were used as the crude enzyme for catalyzing the sucrose reaction. Enzymatic synthesis of FOS crude enzymes was added to a 500 g L−1 sucrose dissolved in phosphate buffer (pH 5.5), incubated at 120 rpm for 30 min at 55 °C and terminated by heating at 100 °C for 15 min. The analysis was performed by an HPLC system (LC20AB, Shimadzu, Japan, equipped with a Kromasil NH2 column (5 μm, 4.6 mm × 250 mm)) under the conditions of mobile phase of acetonitrile/water (75:25, v/v) and flow rate of 1.0 ml min−1. One unit of enzyme activity was defined as the amount of enzyme required to generate 1 μmol 1-kestose per minute under experimental conditions. Under the same conditions as above, the FOS content in the fermentation broth was determined by HPLC. Protein concentration was determined according to the method of Bradford [26], using Bovine albumin (BSA) as a standard and measuring the absorbance at 595 nm. 2
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2.4. Cloning of A. oryzae S719 FTase
considered to be 100%. The buffers (0.2 M) used were: citrate (pH 3.0 and 4.0), acetate (pH 4.0–5.5), phosphate (pH 5.5–7.5), and Tris-HCl (pH 7.5–9.0) and three independent experiments were performed. To determine the optimum temperature of the enzyme, the enzyme was measured at different temperatures between 20 and 80 °C and pH 5.5. The relative activity of the purified FTase at optimum temperature was considered to be 100%. To investigate the heat stability of the enzyme, the residual activity was measured under optimum conditions after incubating the enzyme for 2, 12 and 24 h at various temperatures between 20 and 80 °C. The activity measured was 100% when incubated at 25 °C for 2 h. Three independent experiments were performed. The effects of various metals on FTase activity of A. oryzae S719 were detected after pre-incubation of the enzyme and with specific metal ions at 40 °C for 30 min. The activity of A. oryzae S719 activity without any additions was set as 100%. Three independent experiments were performed.
Fungal biomass was harvested by filtration through a filter paper and frozen in liquid nitrogen and homogenized by grinding in a mortar. Total RNA was isolated using SV total RNA isolated system (Promega). First strand cDNA was synthesized using the GoScript reverse transcript system (Promega) following manufacturer's instructions. The A. oryzae S719 FTase cDNA, without its introns, was synthesized by SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen, Carlsbad, CA, USA). The cDNA obtained by RT-PCR was used as a template for PCR. The sequences of the primers were as follows: Forward primer, 5′-ATGAGGCTCTCAACCGC-3’; Reverse primer, 5′-TTAGCCAATGCCT TGT-3’. The PCR conditions were as follows: initial denaturation step at 95 °C for 5 min; 30 cycles of denaturation (95°Cfor 30 s), annealing (54°Cfor 30 s), and extension (72 °C for 2 min); and a final extension step at 72 °C for 10 min. 2.5. Purification of enzyme and SDS-PAGE analysis
2.8. Determination of kinetic parameters
All procedures were conducted at 4 °C. First of all, the extracellular fluid (centrifuging the fermentation broth) was added with ammonium sulphate to a saturation of 60%. Following removal of the impurities by centrifugation, the crude enzyme was step-wisely precipitated by addition of ammonium sulphate to 95% saturation and left for 12 h. Then, the precipitates were dissolved in 50 mM phosphate buffer with an adjustment of pH to 5.5 and allowed to stand for 12 h, followed by centrifugation at 10,000 rpm for 10 min to obtain the supernatant which was subsequently dialyzed against phosphate buffer (pH 7.5). The acquired dialysate was filtrated in a tangential flow membrane filtration system [GE (Amersham), USA] and concentrated. Later on, the concentrated suspension was applied to a DEAE-Sepharose (Pharmacia, USA) pre-equilibrated with 50 mM phosphate buffer (pH 7.5) and then eluted with the same buffer containing 1.2 M sodium chloride. Next, the eluates were loaded onto a Sephacryl S-200 HR column [GE (Amersham), USA] and eluted with 50 mM phosphate buffer (pH 7.5) containing 0.15 M sodium chloride. At last, the active fractions were pooled, concentrated, and used as a purified FTase enzyme. The molecular weight of purified enzyme was determined by sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDSPAGE). Protein molecular weight marker was obtained from TaKaRa (Dalian, China). SDS-PAGE was carried out on a Mini-PROTEAN 3 Cell (Bio-Rad, USA) using 12% resolving and 4% stacking polyacrylamide gel. After electrophoresis, the gel was stained with Coomassie Brilliant Blue (CBB) R250.
Determination of kinetic parameters the kinetic parameters (Km, Vmax, kcat, and kcat/Km) of the purified enzyme were measured at pH 5.5 (phosphate buffer, 200 mM) at 50 °C. Substrate concentrations were in the range of 10–500 g L−1. The enzyme activity in the reaction mixtures were determined as described in Subsection 2.4. Km and Vmax values were calculated from a Line weaver–Burk plot, and the values were expressed as the mean of the triplicate experiments. Three independent experiments were performed. 2.9. Synthesis of FOS FOS synthesis was carried out for 24 h at 55 °C and pH 6.0 (phosphate buffer, 200 mM). The high concentration of sucrose (500 g L−1, w v−1) as the initial substrate was used to analyze the FOS synthesis ability of this FTase during FOS industrial production [26,27]. The amount of FTase used was 12 U g−1 (sucrose) during the FOS synthesis reaction. Aliquots were taken at intervals. The enzyme was inactivated by boiling for 20 min in water. Then the FOS was analyzed quantitatively by HPLC. 3. Results and discussion 3.1. Sequences analysis The cDNA of mature FTase without its introns and signal peptide was obtained by RT-PCR from the total RNA of A. oryzae S719. Nucleotide sequence analysis revealed that the gene was 1782 bp long (Supplementary material 2) and that the protein encoded consisted of 594 amino acids. The nucleotide sequence has been submitted to GenBank (MN555333). The cDNA sequence of FTase from A. oryzae S719 was compared with that of A. oryzae RIB40 unnamed protein product (GenBank: XM_001824873.1), which shared 99% homology. Fig. 2a showed that the two amino acids are different. The above results indicate that we have isolated a novel gene encoding FTase.
2.6. Protein identification by mass spectrometric peptide mapping For mass spectrometry, Coomassie-stained protein bands were manually excised from gels. In-gel digestion was conducted using trypsin and chymotrypsin. Digested peptides were analyzed by Q Exactive LC-MS/MS mass spectrometer coupled with 2-D nano LC system (Thermo Fisher, America), mass spectral data were searched against the NCBI database, and protein was identified using the Thermo Proteome Discoverer 1.3.0.339 program.
3.2. FTase purification and protein identification
2.7. Effect of pH, temperature effect, and metal ions specificity
A. oryzae has been widely used in the food industry for production of FOS. However, the A. oryzae FTase have been mainly reported at the gene level. Thus, further characterization of these enzymes is necessary. In this study, purification of the FTase from A. oryzae S719 was achieved by a combination of several steps listed in Table 1 and each purification step are listed shown in the supplementary material 3. As summarized in Table 1, the enzyme was purified 66-fold with a final yield of 26% compared to the crude extract and the specific activity of the final purified enzyme was estimated to be 4200 U mg−1 protein.
To determine the optimum pH of the enzyme, the reaction was performed in solution buffered between pH 3.0 and 8.0. The relative activity of the purified FTase at optimum pH was considered to be 100%. To evaluate pH stability of the enzyme, the residual enzymatic activity was determined under optimum reaction conditions after the enzyme was kept in buffer with pH range of 4.0–8.0 for 2 h. The residual FTase activity was measured immediately as described in Subsection 2.3. The relative activity of the purified FTase at pH 6 was 3
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(caption on next page)
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Fig. 2. Sequences analysis and protein identification The unique fragment is a typical protein fragment in which a protein is distinguished from other proteins. (a) Comparison of the amino acid sequences deduced from the cloned β-fructofuranosidase gene of A.orayze S719 and the fructosyltransferase (NO. Q2U3S3) of A. oryzae RIB40. Differential amino acids are represented by a red background. Those of the 12 unique fragments of purified FTase determined by Q Exactive MS are showed in the white box. N-glycosylation potential sites are represented by asterisks. N-glycosylation potential sites are represented by blue font. (b) The fragment Ion Peak of Secondary Mass Spectrometry of one unique peptide (EAGLVPFQVSPTTK) of FTase (b).
This purified enzyme had a high specific activity. The specific activity of the FTase from A. oryzae ZZ-01 was only 1 U mg−1 [28], and that of the FTase from onion bulbs was 27 U mg−1 [29]. The SDS–PAGE result indicated approximate relative masses of 95 kDa for the purified protease with FTase activity (Fig. 3 line 1), which was similar to the FTase from A. oryzae FS4 [30]and Aureobasidium melanogenum 11–1 [31]. Extracellular FTase was mostly monomer, and its molecular weight was generally 50–90 kDa [31–33]. Intracellular enzymes were mostly dimers or trimers with molecular weights between 200 and 300 kDa [30,34], and a few are tetramers or even hexamers. The cloned cDNA encodes a protein with 594 amino acids with a deduced molecular mass of 64 kDa. The purified FTase also showed a molecular mass of approximately 95 kDa, which is approximately 30 kDa higher than the predicted mature protein molecular weight. Using software (http:// www.cbs.dtu.dk/services/NetCGlyc/), 9 potential N-glycosylation sites (Asn-X-Ser/Thr) were identified, and the glycosylation condition of the purified enzymes was confirmed (Fig. 2a). Due to glycosylation, the apparent molecular mass of the protein by SDS-PAGE may be much higher than that calculated for the ORF. According to the above analysis, the high apparent molecular mass of the mature fructosyltransferase protein by SDS-PAGE probably resulted from post-translational modification. Li et al. [30,34]had purified a glycosylated fructosyltransferase, and the total percentage of carbohydrate was determined as approximately 31% of the total mass of the protein. Aung et al. [31] proved that fructofuranosidase from the transformant 33 was a glycosylated protein by using the Endo-H to treat and reduce the molecular weight of the purified fructofuranosidase. The protein was identified by Mass Spectrometry in order to determine whether the purified protein is known or related to a known protein. Fig. 2a showed that the 12 peptides of the purified FTase protein identified by MS matched the deposited uncharacterized protein sequence of A. oryzae RIB40 (UnitProtKB accession NO. Q2U3S3), accounting for 23% of the entire amino acid sequence, and the 11 peptides detected were unique to the protein of the GH32 family [5,35] (Fig. 2b just shown one unique peptide information). Combined the nucleotide sequence analysis of A. oryzae S719 fructosyltransferase, it indicated that a purified novel enzyme has been identified. To characterize the purified protein, the BLAST programs at the National Center for Biotechnology Information (NCBI) were used for the nucleotide sequence analysis and database searches. The amino acid sequence of uncharacterized protein from A. oryzae RIB40 shared 100% identity with that of A. oryzae FTase (GenBank: EU118292.1). These indicated that purified enzyme was successfully identified as a type of FTase in this work. Li et al. [30] reported that the purified fructosyltransferase were identified by MS, and the protein fingerprint mapping matched the deposited FTase protein (UnitProtKB accession No. Q27J21).
Fig. 3. Determination of molecular weight of the enzyme by SDS-PAGE Lane M, molecular mass markers; lane 1, the active fraction from a Sephacryl S-200 HR column; lane 2, the active fraction from a DEAE-Sepharose column.
3.3. Effect of pH on enzyme activity and stability The effects of pH and temperature on the activity of the purified FTase were shown in Fig. 4. The effect of pH on enzyme activity was examined at various pH values ranging from 3.0 to 12.0. The enzyme was most active at the approximate pH of 6.0 (Fig. 4a), which was relatively conservative. Above or below pH 6.0 the relative enzyme activity decreased rapidly. It indicated that the enzyme exhibits different dissociation states under different pH conditions, but only one dissociation state may be beneficial to the binding of the substrate. In addition, the “dissociation state” could be also important to catalysis of the structure of the enzyme. The optimum reaction pH of FTase was 5.0–6.0, which was consistent with the research reported [36,37]. It was stable at wide pH ranges and retained more than 90% of its activity at pH 4.0–11, but below pH 4.0, the enzyme activity reduced abruptly (Fig. 4b). These results are consistent with those of extracellular invertase from A. niger AS0023 [36]. This stability in the broad pH range of the enzyme enhances the adaptability of the enzyme to the food industry. 3.4. Effect of temperature on enzyme activity and stability The optimum temperature for the activity of the enzyme was found to be 55 °C (Fig. 4c). The purified FTase had a high relative activity
Table 1 Purification procedures of b-fructosidase from A. oryzae S719. Purification steps
Total protein (mg)
Total protease activity (U)
Specific activity (U mg−1)
Purification fold
Yield (%)
Crude extract Ammonium sulphate precipitation Membrane filtration DEAE-Sepharose Sephacryl S-200 HR
580 202 140 11 2.3
37000 31000 28000 18000 9700
64 156 196 1700 4200
1.000 2.4 3.1 26 66
100 85 76 49 26
Values given are the average of three replicates. 5
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Fig. 4. Effects of temperature and PH on activity and stability of FTase. Values are means of three independent experiments (a) Effects of PH on activity of FTase. (b) Effects of PH on stability of FTase. (c) Effects of temperature on activity of FTase. (d) Effects of temperature on stability of FTase. The relative enzyme activity is obtained by comparing the enzyme activity measured at different conditions with the maximum value between them.
(> 90% of maximum) at temperatures ranging from 55 to 65 °C. The optimum temperature of FTase was consistent with that (55 °C) of the FTase from P. pastoris GS115 [38]. For the industrial application of the enzyme, it is desirable to operate at temperature above 60 °C in order to avoid microbial contamination. Fig. 4d showed that the FTase retained approximately 89% of its activity after incubation at 60 °C for 2 h. However, its activity declined rapidly at temperatures above 65 °C. The FTase was stable at 25 °C for 12 h, and retained approximately 65% of its activity after incubation for 24 h. The FTase activity began to decline due to thermal denaturation of proteins, and most research described an upper temperature limit of 40 °C [39–41] to 50 °C [32,36]. Significantly, The FTase of A. orayze S719 could produce FOS under high temperature of 60 °C. It is a known fact that high temperature fermentation not only decreased the sterilization cost, but also reduced the contamination possibilities. Compared with the conventional fermentation, the consumption of cooling water could also greatly reduced in high temperature fermentation, which resulted in lower energy consumption and production cost for large-scale industrial fermentation especially in tropical or subtropical areas. In addition, the enzyme was capable of maintaining high activity at room temperature (25 °C), which was advantageous for the application of this enzyme in the largeterm transportation.
Fig. 5. Lineweaver–Burk plots for using sucrose as substrate by FTase. Values are means of three independent experiments.
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Table 2 Effect of metal ions on the activity of FTase. Metal ions
2+
Ca Mg2+ Zn2+ Cu2+ Pb2+ Fe2+ Mn2+ EDTA Na+ CKb
Concentration relative activity (%)a 5 mM
10 mM
97 ± 2.3 116 ± 1.3 19 ± 1.5 2.4 ± 2.7 1.9 ± 2.4 0 70 ± 2.2 96 ± 1.4 91 ± 1.9 100
91 ± 2.1 89 ± 2.4 10 ± 1.8 2.3 ± 1.6 0 ± 1.0 0 80 ± 1.3 91 ± 1.2 114 ± 2.1 100
(a)The relative activity with different metal ions added was determined and compared with the activity measured in Phosphate buffer (pH 5.5, 0.2 M) without the addition of any ions. (b) The relative activity measured in Phosphate buffer (pH 5.5, 0.2 M) without the addition of any ions. The experiments were independently performed at least three times. The errors are shown as standard deviation (SD).
Fig. 6. FOS production by FTase G: glucose, GF: sucrose, GF2: 1-kestose, GF3: nystose, GF4: fructosylnystose, FOS: fructooligosaccharides. The synthesis of FOS was carried out for 24 h by using sucrose as initial substrate (500 g L−1) at 50 °C and pH 5.5. Values are means of three independent experiments.
3.5. Kinetic studies The kinetics of FTase was determined using sucrose as a substrate. The Km and Vmax of FTase were 310 mmol L−1 and 1.4 mmol L−1 min−1, respectively (Fig. 5). The kcat and kcat/Km of FTase were 2.0 × 103 min−1 and 6.4 L mmol−1 min−1, respectively. These results indicated that FTase has high catalytic efficiency. The kinetic constant of FTase from Rhodotorula sp was determined. The Km value of sucrose was 198 g L−1 [42]. Jung et al. [43]determined the properties of the FTase from Aureobasidium pullulans, and found that the Km value and Vmax value for sucrose were 330 g L−1 and 130 g (L h)−1. It was indicated that a higher substrate affinity of these FTase from A. pullulans and Rhodotorula sp. was lower than that of FTase heterologously expressed in a A. oryaze S719 in this work. The lower Km value of the purified A. oryaze S719 FTase found here indicated that it had a greater affinity for the substrate.
HPLC demonstrated that within 4 h of the reaction, the yield of FOS was over 0.64 g of FOS/g of sucrose and percentages of GF2, GF3 and GF4 were 67%, 32% and 0.06% (data not shown) whereas within 6 h of the reaction, the yield of FOS was 0.64 g of FOS/g of sucrose and percentages of GF2, GF3 and GF4 were 56%, 42% and 1.4%. These results demonstrated that the FTase has high transfructosylation efficiency towards the synthesis of FOS from fructosyl groups and sucrose. The result comparing A. oryaze S719 production of FOS with other strains were shown in Table 3. The purification of the FTase from A. orayze S719 obtained a yield of 64% of FOS within 4 h of reaction, indicating a considerable reduction in the reaction time of 8–24 h reported in the literature. It also was the shortest reaction time to be reported regarding the purified enzyme production of FOS. This further indicated that the use of A. oryaze S719 to produce FOS could save more time compared to other strains. In the industry, it is known that shortening production time would not only speed up the production process, but also reduce waste of resources and reduce energy consumption. According to the above results, the A. oryaze S719 strain with high transfer efficiency would have the potential to lay a foundation for its application in FOS production. In addition, to the best of our knowledge, A. oryzae S719 strain therefore represents the first reported non-recombinant fructosyltransferase that have shortest reaction time for the production of FOS. According to the above results, the A. oryaze S719 strain with [8] high transfructosylation efficiency has the potential to replace the recombinant strain producing FOS.
3.6. Effect of metal ions on FTase activity The effect of different metal ions on FTase activity was determined (Table 2). FTase was significantly activated by 5 mmol L−1 Mg2+ and 10 mmol L−1 Na+, respectively, which indicated that the FTase was Mg2+ and Na+ dependent, with relative activities of 116 and 114%, respectively. The Na+ (5 mmol L−1), Ca2+ (5 mmol L−1), and EDTA (5 mmol L−1) had no effect on FTase activity. The enzyme was not inhibited by 5 mM EDTA indicating that it was not a metalloprotein. Other metal ions inhibited FTase activity, but the precise mechanisms by which heavy metal ions inhibit the FTase activity were unknown [44]. Yang et al. [44] analyzed effect of metal ions on the activity of the FTase from Pichia pastoris GS115 and found that Mg2+ and Ni+ could enhance its initial activity.
4. Conclusion We have isolated and purified a novel fructosyltransferase form A. oryzae S719, which was identified as a type of FTase by MS. The purified FTase from A. orayze S719 obtained a yield of 64% of FOS within 4 h of reaction, which was the shortest reaction time to be reported regarding the purified enzyme production of FOS. The molecular weight of the purified FTase was around 95 kDa, and the specific activity was 4200 U mg−1. This FTase had a high relative activity at temperatures ranging from 25 to 65 °C, and the FTase was stable at wide pH ranges and retained more than 80% of its activity at pH 4.0–11. The FTase had high catalytic efficiency, the kcat of which was 2.0 × 103 min−1. The FTase was Mg2+ and Na+ dependent. This study provided a novel FTase with high transfer efficiency and would have the potential to lay a foundation for its application in FOS production.
3.7. FOS production According to the results of 3.3 and 3.4, it was found that 12 U of FTase activity/g of sucrose, 500.0 mg sucrose mL−1, pH 6.0 in 0.1 M sodium acetate buffer and reaction temperature of 55 °C were the most suitable for the transfructosylation reaction. As shown in Fig. 6, several important time process parameters were monitored during the transglucosidic assay including FOS yield, yield of each oligomer, and residual sucrose. As the reaction time increased, the amount of sucrose was decreased and the amounts of glucose and FOS were increased, but the released fructose could not be detected. Analysis of the FOS generated, glucose (G), fructose (F) and sucrose (GF) standards by using 7
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Table 3 Comparison of some parameters obtained for different strains in the present study and the literature. Strains A.oryzae S719 A.tubingensis XG21 PichiapastorisGS115 Aureobasidium melanogenum 11-1 A. pullulans CFR77
Enzyme conc −1
12 U g 6 U g−1 8 U g−1 117 U g−1 NS
Carbon source Sucrose Sucrose Sucrose Sucrose Sucrose
(400 (200 (500 (300 (600
g g g g g
−1
L ) L−1) L−1) L−1) L−1)
Cultivation time
FOS yield (%)
References
4 8 18 7 9
63% 57% 64% 66% 59%
Present study Xie et al. [45]. Guo et al. [15]. Jiang et al. [31]. Agbaje et al. [34].
NS: not stated.
Decalaration of competing interest [17]
The authors declare that they have no conflict of interest.
[18]
Acknowledgments
[19]
This work was financially supported by the Natural Science Foundation of Guangxi Province (Grant No. 2016GXNSFAA380130, 2017GXNSFAA198010, 2018GXNSFAA281019 and 2018GXNSFAA 138024) and the National Natural Science Foundation of China (Grant No. 31560448, 31660251 and 31860245) and the central government directs special funds for local science and technology development projects (ZY1949015).
[20] [21] [22] [23]
Appendix A. Supplementary data
[24]
Supplementary data to this article can be found online at https:// doi.org/10.1016/j.pep.2019.105549.
[25]
References
[26]
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improvement of the industrial fungus Aspergillus niger ATCC 20611, J. Biotechnol. 249 (2017) 25–33. J.W. Yun, Fructooligosaccharides—occurrence, preparation, and application, Enzym. Microb. Technol. 19 (2) (1996) 107–117. M.A. Rodríguez, O.F. Sánchez, C.J. Alméciga-Díaz, Gene cloning and enzyme structure modeling of the Aspergillus oryzae N74 fructosyltransferase, Mol. Biol. Rep. 38 (2) (2011) 1151–1161. P.T. Sangeetha, M.N. Ramesh, S.G. Prapulla, Recent trends in the microbial production, analysis and application of Fructooligosaccharides, Trends Food Sci. Technol. 16 (10) (2005) 442–457. P. Chuankhayan, et al., Crystal structures of Aspergillus japonicus fructosyltransferase complex with donor/acceptor substrates reveal complete subsites in the active site for catalysis, J. Biol. Chem. 285 (30) (2010) 23251–23264. S. Hayashi, et al., Purification and properties of beta-fructofuranosidase from Aspergillus japonicus, World J. Microbiol. Biotechnol. 8 (3) (1992) 276–279. C.T. Chang, et al., Purification and properties of beta-fructofuranosidase from Aspergillus oryzae ATCC 76080, Biochem. Mol. Biol. Int. 32 (2) (1994) 269–277. J.S. Chen, et al., Purification and partial characterization of the high and low molecular weight form (S- and F-form) of invertase secreted by Aspergillus nidulans, Biochim. Biophys. Acta 1296 (2) (1996) 207. Q.D. Nguyen, et al., Production, purification and identification of fructooligosaccharides produced by β-fructofuranosidase from Aspergillus niger IMI 303386, Biotechnol. Lett. 21 (3) (1999) 183–186. L. L Hocine, et al., Purification and partial characterization of fructosyltransferase and invertase from Aspergillus niger AS0023, J. Biotechnol. 81 (1) (2000) 73–84. I. Ghazi, et al., Immobilisation of fructosyltransferase from Aspergillus aculeatus on epoxy-activated Sepabeads EC for the synthesis of fructo-oligosaccharides, J. Mol. Catal. B Enzym. 35 (1) (2005) 19–27. L.M. Boddy, Purification and characterisation of an Aspergillus niger invertase and its DNA sequence, Curr. Genet. 24 (1–2) (1993) 60–66. T. Wei, et al., Purification and evaluation of the enzymatic properties of a novel fructosyltransferase from Aspergillus oryzae: a potential biocatalyst for the synthesis of sucrose 6-acetate, Biotechnol. Lett. 36 (5) (2014) 1015–1020. G. Notton, P. Poggi, C. Cristofari, Purification and characterization of a fructosyltransferase from onion bulbs and its key role in the synthesis of fructo-oligosaccharides in vivo, New Phytol. 165 (2) (2010) 513–524. X. Li, et al., Purification, cloning, characterization, and N-glycosylation analysis of a novel β-fructosidase from Aspergillus oryzae FS4 synthesizing levan- and neolevantype fructooligosaccharides, PLoS One 9 (12) (2014) e114793. T. Aung, et al., Overproduction of a β-fructofuranosidase1 with a high FOS synthesis activity for efficient biosynthesis of fructooligosaccharides, Int. J. Biol. Macromol..130 (2019) 988–996. T. Wei, et al., Purification and evaluation of the enzymatic properties of a novel fructosyltransferase from Aspergillus oryzae: a potential biocatalyst for the synthesis of sucrose 6-acetate, Biotechnol. Lett. 36 (5) (2014) 1015–1020. G. Lihong, et al., Expression and activity analysis of fructosyltransferase from Aspergillus oryzae, Protein J. 36 (6) (2017) 1–9. A. Lateef, J.K. Oloke, S.G. Prapulla, Purification and partial characterization of intracellular fructosyltransferase from a novel strain of Aureobasidium pullulans, Turkish J. Biol. 31 (1) (2007) 147. M.L. Velázquez-Hernández, et al., Microbial fructosyltransferases and the role of fructans, J. Appl. Microbiol. 106 (6) (2009) 1763–1778. L. L Hocine, et al., Purification and partial characterization of fructosyltransferase and invertase from Aspergillus niger AS0023, J. Biotechnol. 81 (1) (2000) 73–84. J.P. Park, T. Oh, J. Yun, Purification and characterization of a novel transfructosylating enzyme from Bacillus macerans EG-6, Process Biochem. 37 (5) (2001) 471–476. H. Yang, et al., Heterologous expression and enzymatic characterization of fructosyltransferase from Aspergillus niger in Pichia pastoris, N. Biotech. 33 (1) (2016) 164. X.D. Wang, S.K. Rakshit, Iso-oligosaccharide production by multiple forms of transferase enzymes from Aspergillus foetidus, Process Biochem. 35 (8) (2000) 771–775. K.B. Song, et al., Characteristics of levan fructotransferase from Arthrobacter ureafaciens K2032 and difructose anhydride IV formation from levan, Enzym. Microb. Technol. 27 (3) (2000) 212–218.
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Abbreviations
[41] J. Cha, et al., Molecular and enzymatic characterization of a levan fructotransferase from Microbacterium sp. AL-210, J. Biotechnol. 91 (1) (2001) 49–61. [42] M. Alvarado, F. Maugeri, Kinetic modelling of fructooligosaccharide synthesis using a Rhodotorula sp. fructosyltransferase, J. Biotechnol. 131 (2) (2007) S91–S92. [43] K.H. Jung, et al., Mathematical model for enzymatic production of fructo-oligosaccharides from sucrose, Enzym. Microb. Technol. 11 (8) (1989) 491–494. [44] K.H. Jung, et al., Mathematical model for enzymatic production of fructo-oligosaccharides from sucrose, Enzym. Microb. Technol. 11 (8) (1989) 491–494. [45] Y. Xie, et al., A molasses habitat-derived fungus Aspergillus tubingensis XG21 with high β-fructofuranosidase activity and its potential use for fructooligosaccharides production, Amb. Express 7 (1) (2017) 128.
FOS: Fructooligosaccharides FTase: β-fructofuranosidases SDS-PAGE: sodium dodecyl sulphate polyacrylamide gel electrophoresis GF2: 1-kestose GF3: 1-nystose GF4: 1-fructofuranosylnystose UV: ultraviolet BSA: Bovine albumin NCBI: National Center for Biotechnology Information
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Protein Expression and Purification journal homepage: www.elsevier.com/locate/yprep
Purification and biochemical characteristics of a novel fructosyltransferase with a high FOS transfructosylation activity from Aspergillus oryzae S719
T
Susu Hana,b, Tong Yea,b, Shuo Lenga,b, Lixia Panc, Wei Zenga,b, Guiguang Chena,b,∗∗, Zhiqun Lianga,b,∗ a
State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources, Guangxi Microorganism and Enzyme Research Center of Engineering Technology, China b College of Life Science and Technology, Guangxi University, Nanning, 530004, Guangxi, China c National Engineering Research Center for Non-Food Biorefinery, State Key Laboratory of Non-Food Biomass and Enzyme Technology, Guangxi Key Laboratory of Biorefinery, Guangxi Biomass, Engineering Technology Research Center, Guangxi Academy of Sciences, 98 Daling Road, Nanning, 530007, China
ARTICLE INFO
ABSTRACT
Keywords: Fructosyltransferase Aspergillus oryzae Enzyme purification Fructooligosaccharides production
Fructooligosaccharides (FOS) have widely used for the manufacture of low-calorie and functional foods, because they can inhibit intestinal pathogenic microorganism growth and increase the absorption of Ca2+ and Mg2+. In this study, the novel fructosyltransferase (FTase) from Aspergillus oryzae strain S719 was successfully purified and characterized. The specific activity of the final purified material was 4200 mg−1 with purification ratio of 66 times and yield of 26%. The molecular weight of FTase of A. oryzae S719 was around 95 kDa by SDS-PAGE, which was identified as a type of FTase by Mass Spectrometry (MS). The purified FTase had optimum temperature and pH of 55 °C and 6.0, respectively. The FTase showed to be stable with more than 80% of its original activity at room temperature after 12 h and maintaining activity above 90% at pH 4.0–11.0. The Km and kcat values of the FTase were 310 mmol L−1 and 2.0 × 103 min−1, respectively. The FTase was activated by 5 mmol L−1 Mg2+ and 10 mmol L−1 Na+ (relative activity of 116 and 114%, respectively), indicating that the enzyme was Mg2+ and Na+ dependent. About 64% of FOS was obtained by the purified FTase under 500 g L−1 sucrose within 4 h of reaction time, which was the shortest reaction time to be reported regarding the purified enzyme production of FOS. Together, these results indicated that the FTase of A. oryzae S719 is an excellent candidate for the industrial production of FOS.
1. Introduction Fructooligosaccharides (FOS) are widely used as a bioactive ingredient in functional foods because it represents a major class of prebiotics [1]. FOS are a short-chain fructosyl unit (F) that is primarily linked to the terminal sucrose molecule (GF) at the reducing end. Commercial FOS formulations exhibit a low degree of polymerization, consisting primarily of 1-kestose (GF2), 1-nystose (GF3) and 1-fructofuranosylnystose (GF4). In recent years, they have been receiving increasing attention because they are stable in the range of pH 4.0–11.0, and stored for a long time at low temperatures, and their sweetness is only one-third of sucrose. In addition, their thermal stability and viscosity are higher than sucrose [2–4]. Most importantly, they have many
physiological functions such as inhibiting the growth of intestinal pathogenic microorganisms and increasing adsorption of Ca2+ and Mg2+ [5,6]. So far, commercially, the production of FOS has been primarily carried out from sucrose by the transfructosylation activity of fructofuranosidases (EC 2.4.1.9)(FTase). Based on overall amino acid sequence similarities, FTase belongs to the glycosyl hydrolase family 32 (GH32) and shares a common three-dimensional (3-D) structure with other GH32 members [7,8]. The enzyme reaction transfructosylation mechanism (Fig. 1) can be expressed as (Eq. (1)): F−R + E ↔ F−E + R F−E + acceptor → F−acceptor + E
(1)
Where F is fructose, E is transfructosylation and R represents a
∗ Corresponding author. State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources; Guangxi Microorganism and Enzyme Research Center of Engineering Technology; College of Life Science and Technology; Guangxi University, 100 Daxue Road, Nanning 530004, Guangxi, China. ∗∗ Corresponding author. State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources; Guangxi Microorganism and Enzyme Research Center of Engineering Technology; College of Life Science and Technology; Guangxi University, 100 Daxue Road, Nanning, 530004, Guangxi, China. E-mail addresses: [email protected] (G. Chen), [email protected] (Z. Liang).
https://doi.org/10.1016/j.pep.2019.105549 Received 24 July 2019; Received in revised form 28 November 2019; Accepted 28 November 2019 Available online 02 December 2019 1046-5928/ © 2019 Elsevier Inc. All rights reserved.
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steps towards understanding some of its properties. In this study, the purification and characterization of a fructofuranosidases (FTase) from A. oryzae S719 were described. In addition to the broad enzymatic properties, this enzyme displayed a high FOS synthesis efficiency with a FOS yield of approximately 64% within 4 h of reaction, which demonstrated that this enzyme might be a potential biocatalyst for the synthesis of FOS in industry. 2. Materials and methods 2.1. Strains and materials A. oryzae ZT65 (CCTCCM) was stored in the Culture and Information Center of Industrial Microorganism of China Universities. Mycelium was grown at 37 °C for 48 h on solid medium (sucrose 20 g L−1, agar 20 g L−1 and NaCl 7 g L−1, pH 5.5). Its spores were collected with 0.85% sterile saline and prepared as 10−8 spores ml−1 suspension. The A. oryzae ZT65 spore solution was applied to the plate medium, cultured in a 37 °C incubator for 4 h, and continuously irradiated for 50 s using an ultraviolet lamp (15 W of power, effective wavelength 254 nm, distance 30 cm). It was induced by UV to a positive mutant strain A. oryzae S719 with significantly improved enzyme activity, which was identified as the target strain (supplementary material 1). They were stored on solid medium composed of 250 g L−1 sucrose, 30 g L−1 maceration extract of bran, 5 g L−1 LiCl, and 20 g L−1 agar. FOS standards 1-kestose (GF2), 1-nystose (GF3) and 1-fructofuranosylnystose (GF4) were of biochemical grade obtained from Wako Pure Chemical Industries (Chuo-Ku, Osaka, Japan). Acetonitrile as mobile phase used for HPLC was obtained from Merck (Germany). Wheat bran was purchased from Guangdong baiyan grain and oil industry Co. (Guangdong, China). Corn steep powder was obtained from Shandong Kangyuan Bio-Tech Co., Ltd. (Yuncheng, China). All other chemicals were of analytical grade.
Fig. 1. Network of the reaction mechanism for the production of FOS Network of the reaction mechanism for the production of FOS from sucrose catalyzed by fructosyltransferase: G, GF, GF2, GF3, and GF4, means glucose, sucrose, I-kestose, nystose, and lF- fructofuranosyl nystose, respectively. Where F is fructose, E is fructosyltransferase and R represents sucrose.
carbonyl of an aldose. According to this mechanism, one molecule of sucrose serves as a donor and another acts as an acceptor for GF2 synthesis, releasing one molecule of glucose, for the production of GF3, the GF2 acts as an acceptor [7]. According to the report of Trollope et al. [9], FTase can be sub-classified into the two groups with low and high FOS synthesis activity, possessing either predominantly hydrolytic or high FTase activity. The theoretical yield of FOS from sucrose is 75% if 1-kestose is the only FOS produced. However, the actual FOS yields were considerably lower than 60% with A. niger ATCC 20611 [10] and 53–59% with Aureobasidium sp. [11–13]. One explanation for this is that the enzymes involved in FOS production are inhibited by the liberated glucose that accumulates as a by-product [14]. It is well known that the low activity of the wild-type FTase makes them unsuitable for large-scale industrial application in FOS production. For this reason, many studies have attempted to produce FTase and FOS using different recombinant strains with high levels of FOS synthesis, such as Pichia pastoris GS115 [15]. However, the fungus stable system of Aspergillus is often used to produce FOS industrially, especially the FopA enzyme from A. niger ATCC 20611 [16]. It can be seen that the stability and practicability of the fungal system still exerts great potential in FOS production. In the past two decades, a lot of studies have been carried out to improve the efficiency of FTase production, but little has been known about FTase studies with high-level FOS synthesis from nonrecombinant strains. In nature, there are wide range of FTase resources from microorganisms, but a small part of which can be used in industry. Common enzyme preparations are vulnerable to extreme environments in industrial production, such as strong acid, alkali, high pressure and high temperature conditions. The industrial application of FTase in the production of FOS is highly limited due to inherent properties of FTase, such as low FTase activity (most important), low transfructosylation activity, low catalytic efficiency and poor stability. Several microorganisms possessing FTase activity including Aspergillus niger, Aspergillus oryzae, Aspergillus japonicus, Neurospora crassa, Aureobasidium sp., Fusariumoxys porum, Arthrobacter sp. and yeast have been reported [17–20]. Simultaneously, purification and characterization of FTase from various sources have been studied [21–24]. However, this accumulated knowledge is confusing [21–25]. The information differs from one source to another, from one microorganism to another, even from one strain to another, thereby making it imperative to purify the enzyme from each source. The purification and partial characterization of the enzyme were undertaken as necessary
2.2. Fermentation of A. oryzae S719 The inoculum was developed by transferring a loopful of mycelia from a 3-day-old slant of A. oryzae S719 stored on a bevel into the inoculum medium (10 g bran plus 5 ml water). The 250 ml flasks were incubated for 60 h at 37 ± 1 °C. After that, the spores were washed with 120 ml of sterile water, the bran was filtered off, and the spores were shaken at 200 rpm for 10 min to reach the spore suspensions (108 spores). The inoculum was transferred into 50 ml of fermentation medium consisting of the following (% w/v): sucrose: 18; corn steep powder 4; ZnSO4·7H20 0.2, (initial pH of 5.5). The culture was incubated for 40 h at 34 ± 1 °C on a rotary shaker at 240 rpm. 2.3. Assays of enzyme activity (standard activity assay) and protein concentration The supernatant were used as the crude enzyme for catalyzing the sucrose reaction. Enzymatic synthesis of FOS crude enzymes was added to a 500 g L−1 sucrose dissolved in phosphate buffer (pH 5.5), incubated at 120 rpm for 30 min at 55 °C and terminated by heating at 100 °C for 15 min. The analysis was performed by an HPLC system (LC20AB, Shimadzu, Japan, equipped with a Kromasil NH2 column (5 μm, 4.6 mm × 250 mm)) under the conditions of mobile phase of acetonitrile/water (75:25, v/v) and flow rate of 1.0 ml min−1. One unit of enzyme activity was defined as the amount of enzyme required to generate 1 μmol 1-kestose per minute under experimental conditions. Under the same conditions as above, the FOS content in the fermentation broth was determined by HPLC. Protein concentration was determined according to the method of Bradford [26], using Bovine albumin (BSA) as a standard and measuring the absorbance at 595 nm. 2
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2.4. Cloning of A. oryzae S719 FTase
considered to be 100%. The buffers (0.2 M) used were: citrate (pH 3.0 and 4.0), acetate (pH 4.0–5.5), phosphate (pH 5.5–7.5), and Tris-HCl (pH 7.5–9.0) and three independent experiments were performed. To determine the optimum temperature of the enzyme, the enzyme was measured at different temperatures between 20 and 80 °C and pH 5.5. The relative activity of the purified FTase at optimum temperature was considered to be 100%. To investigate the heat stability of the enzyme, the residual activity was measured under optimum conditions after incubating the enzyme for 2, 12 and 24 h at various temperatures between 20 and 80 °C. The activity measured was 100% when incubated at 25 °C for 2 h. Three independent experiments were performed. The effects of various metals on FTase activity of A. oryzae S719 were detected after pre-incubation of the enzyme and with specific metal ions at 40 °C for 30 min. The activity of A. oryzae S719 activity without any additions was set as 100%. Three independent experiments were performed.
Fungal biomass was harvested by filtration through a filter paper and frozen in liquid nitrogen and homogenized by grinding in a mortar. Total RNA was isolated using SV total RNA isolated system (Promega). First strand cDNA was synthesized using the GoScript reverse transcript system (Promega) following manufacturer's instructions. The A. oryzae S719 FTase cDNA, without its introns, was synthesized by SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen, Carlsbad, CA, USA). The cDNA obtained by RT-PCR was used as a template for PCR. The sequences of the primers were as follows: Forward primer, 5′-ATGAGGCTCTCAACCGC-3’; Reverse primer, 5′-TTAGCCAATGCCT TGT-3’. The PCR conditions were as follows: initial denaturation step at 95 °C for 5 min; 30 cycles of denaturation (95°Cfor 30 s), annealing (54°Cfor 30 s), and extension (72 °C for 2 min); and a final extension step at 72 °C for 10 min. 2.5. Purification of enzyme and SDS-PAGE analysis
2.8. Determination of kinetic parameters
All procedures were conducted at 4 °C. First of all, the extracellular fluid (centrifuging the fermentation broth) was added with ammonium sulphate to a saturation of 60%. Following removal of the impurities by centrifugation, the crude enzyme was step-wisely precipitated by addition of ammonium sulphate to 95% saturation and left for 12 h. Then, the precipitates were dissolved in 50 mM phosphate buffer with an adjustment of pH to 5.5 and allowed to stand for 12 h, followed by centrifugation at 10,000 rpm for 10 min to obtain the supernatant which was subsequently dialyzed against phosphate buffer (pH 7.5). The acquired dialysate was filtrated in a tangential flow membrane filtration system [GE (Amersham), USA] and concentrated. Later on, the concentrated suspension was applied to a DEAE-Sepharose (Pharmacia, USA) pre-equilibrated with 50 mM phosphate buffer (pH 7.5) and then eluted with the same buffer containing 1.2 M sodium chloride. Next, the eluates were loaded onto a Sephacryl S-200 HR column [GE (Amersham), USA] and eluted with 50 mM phosphate buffer (pH 7.5) containing 0.15 M sodium chloride. At last, the active fractions were pooled, concentrated, and used as a purified FTase enzyme. The molecular weight of purified enzyme was determined by sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDSPAGE). Protein molecular weight marker was obtained from TaKaRa (Dalian, China). SDS-PAGE was carried out on a Mini-PROTEAN 3 Cell (Bio-Rad, USA) using 12% resolving and 4% stacking polyacrylamide gel. After electrophoresis, the gel was stained with Coomassie Brilliant Blue (CBB) R250.
Determination of kinetic parameters the kinetic parameters (Km, Vmax, kcat, and kcat/Km) of the purified enzyme were measured at pH 5.5 (phosphate buffer, 200 mM) at 50 °C. Substrate concentrations were in the range of 10–500 g L−1. The enzyme activity in the reaction mixtures were determined as described in Subsection 2.4. Km and Vmax values were calculated from a Line weaver–Burk plot, and the values were expressed as the mean of the triplicate experiments. Three independent experiments were performed. 2.9. Synthesis of FOS FOS synthesis was carried out for 24 h at 55 °C and pH 6.0 (phosphate buffer, 200 mM). The high concentration of sucrose (500 g L−1, w v−1) as the initial substrate was used to analyze the FOS synthesis ability of this FTase during FOS industrial production [26,27]. The amount of FTase used was 12 U g−1 (sucrose) during the FOS synthesis reaction. Aliquots were taken at intervals. The enzyme was inactivated by boiling for 20 min in water. Then the FOS was analyzed quantitatively by HPLC. 3. Results and discussion 3.1. Sequences analysis The cDNA of mature FTase without its introns and signal peptide was obtained by RT-PCR from the total RNA of A. oryzae S719. Nucleotide sequence analysis revealed that the gene was 1782 bp long (Supplementary material 2) and that the protein encoded consisted of 594 amino acids. The nucleotide sequence has been submitted to GenBank (MN555333). The cDNA sequence of FTase from A. oryzae S719 was compared with that of A. oryzae RIB40 unnamed protein product (GenBank: XM_001824873.1), which shared 99% homology. Fig. 2a showed that the two amino acids are different. The above results indicate that we have isolated a novel gene encoding FTase.
2.6. Protein identification by mass spectrometric peptide mapping For mass spectrometry, Coomassie-stained protein bands were manually excised from gels. In-gel digestion was conducted using trypsin and chymotrypsin. Digested peptides were analyzed by Q Exactive LC-MS/MS mass spectrometer coupled with 2-D nano LC system (Thermo Fisher, America), mass spectral data were searched against the NCBI database, and protein was identified using the Thermo Proteome Discoverer 1.3.0.339 program.
3.2. FTase purification and protein identification
2.7. Effect of pH, temperature effect, and metal ions specificity
A. oryzae has been widely used in the food industry for production of FOS. However, the A. oryzae FTase have been mainly reported at the gene level. Thus, further characterization of these enzymes is necessary. In this study, purification of the FTase from A. oryzae S719 was achieved by a combination of several steps listed in Table 1 and each purification step are listed shown in the supplementary material 3. As summarized in Table 1, the enzyme was purified 66-fold with a final yield of 26% compared to the crude extract and the specific activity of the final purified enzyme was estimated to be 4200 U mg−1 protein.
To determine the optimum pH of the enzyme, the reaction was performed in solution buffered between pH 3.0 and 8.0. The relative activity of the purified FTase at optimum pH was considered to be 100%. To evaluate pH stability of the enzyme, the residual enzymatic activity was determined under optimum reaction conditions after the enzyme was kept in buffer with pH range of 4.0–8.0 for 2 h. The residual FTase activity was measured immediately as described in Subsection 2.3. The relative activity of the purified FTase at pH 6 was 3
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(caption on next page)
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Fig. 2. Sequences analysis and protein identification The unique fragment is a typical protein fragment in which a protein is distinguished from other proteins. (a) Comparison of the amino acid sequences deduced from the cloned β-fructofuranosidase gene of A.orayze S719 and the fructosyltransferase (NO. Q2U3S3) of A. oryzae RIB40. Differential amino acids are represented by a red background. Those of the 12 unique fragments of purified FTase determined by Q Exactive MS are showed in the white box. N-glycosylation potential sites are represented by asterisks. N-glycosylation potential sites are represented by blue font. (b) The fragment Ion Peak of Secondary Mass Spectrometry of one unique peptide (EAGLVPFQVSPTTK) of FTase (b).
This purified enzyme had a high specific activity. The specific activity of the FTase from A. oryzae ZZ-01 was only 1 U mg−1 [28], and that of the FTase from onion bulbs was 27 U mg−1 [29]. The SDS–PAGE result indicated approximate relative masses of 95 kDa for the purified protease with FTase activity (Fig. 3 line 1), which was similar to the FTase from A. oryzae FS4 [30]and Aureobasidium melanogenum 11–1 [31]. Extracellular FTase was mostly monomer, and its molecular weight was generally 50–90 kDa [31–33]. Intracellular enzymes were mostly dimers or trimers with molecular weights between 200 and 300 kDa [30,34], and a few are tetramers or even hexamers. The cloned cDNA encodes a protein with 594 amino acids with a deduced molecular mass of 64 kDa. The purified FTase also showed a molecular mass of approximately 95 kDa, which is approximately 30 kDa higher than the predicted mature protein molecular weight. Using software (http:// www.cbs.dtu.dk/services/NetCGlyc/), 9 potential N-glycosylation sites (Asn-X-Ser/Thr) were identified, and the glycosylation condition of the purified enzymes was confirmed (Fig. 2a). Due to glycosylation, the apparent molecular mass of the protein by SDS-PAGE may be much higher than that calculated for the ORF. According to the above analysis, the high apparent molecular mass of the mature fructosyltransferase protein by SDS-PAGE probably resulted from post-translational modification. Li et al. [30,34]had purified a glycosylated fructosyltransferase, and the total percentage of carbohydrate was determined as approximately 31% of the total mass of the protein. Aung et al. [31] proved that fructofuranosidase from the transformant 33 was a glycosylated protein by using the Endo-H to treat and reduce the molecular weight of the purified fructofuranosidase. The protein was identified by Mass Spectrometry in order to determine whether the purified protein is known or related to a known protein. Fig. 2a showed that the 12 peptides of the purified FTase protein identified by MS matched the deposited uncharacterized protein sequence of A. oryzae RIB40 (UnitProtKB accession NO. Q2U3S3), accounting for 23% of the entire amino acid sequence, and the 11 peptides detected were unique to the protein of the GH32 family [5,35] (Fig. 2b just shown one unique peptide information). Combined the nucleotide sequence analysis of A. oryzae S719 fructosyltransferase, it indicated that a purified novel enzyme has been identified. To characterize the purified protein, the BLAST programs at the National Center for Biotechnology Information (NCBI) were used for the nucleotide sequence analysis and database searches. The amino acid sequence of uncharacterized protein from A. oryzae RIB40 shared 100% identity with that of A. oryzae FTase (GenBank: EU118292.1). These indicated that purified enzyme was successfully identified as a type of FTase in this work. Li et al. [30] reported that the purified fructosyltransferase were identified by MS, and the protein fingerprint mapping matched the deposited FTase protein (UnitProtKB accession No. Q27J21).
Fig. 3. Determination of molecular weight of the enzyme by SDS-PAGE Lane M, molecular mass markers; lane 1, the active fraction from a Sephacryl S-200 HR column; lane 2, the active fraction from a DEAE-Sepharose column.
3.3. Effect of pH on enzyme activity and stability The effects of pH and temperature on the activity of the purified FTase were shown in Fig. 4. The effect of pH on enzyme activity was examined at various pH values ranging from 3.0 to 12.0. The enzyme was most active at the approximate pH of 6.0 (Fig. 4a), which was relatively conservative. Above or below pH 6.0 the relative enzyme activity decreased rapidly. It indicated that the enzyme exhibits different dissociation states under different pH conditions, but only one dissociation state may be beneficial to the binding of the substrate. In addition, the “dissociation state” could be also important to catalysis of the structure of the enzyme. The optimum reaction pH of FTase was 5.0–6.0, which was consistent with the research reported [36,37]. It was stable at wide pH ranges and retained more than 90% of its activity at pH 4.0–11, but below pH 4.0, the enzyme activity reduced abruptly (Fig. 4b). These results are consistent with those of extracellular invertase from A. niger AS0023 [36]. This stability in the broad pH range of the enzyme enhances the adaptability of the enzyme to the food industry. 3.4. Effect of temperature on enzyme activity and stability The optimum temperature for the activity of the enzyme was found to be 55 °C (Fig. 4c). The purified FTase had a high relative activity
Table 1 Purification procedures of b-fructosidase from A. oryzae S719. Purification steps
Total protein (mg)
Total protease activity (U)
Specific activity (U mg−1)
Purification fold
Yield (%)
Crude extract Ammonium sulphate precipitation Membrane filtration DEAE-Sepharose Sephacryl S-200 HR
580 202 140 11 2.3
37000 31000 28000 18000 9700
64 156 196 1700 4200
1.000 2.4 3.1 26 66
100 85 76 49 26
Values given are the average of three replicates. 5
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Fig. 4. Effects of temperature and PH on activity and stability of FTase. Values are means of three independent experiments (a) Effects of PH on activity of FTase. (b) Effects of PH on stability of FTase. (c) Effects of temperature on activity of FTase. (d) Effects of temperature on stability of FTase. The relative enzyme activity is obtained by comparing the enzyme activity measured at different conditions with the maximum value between them.
(> 90% of maximum) at temperatures ranging from 55 to 65 °C. The optimum temperature of FTase was consistent with that (55 °C) of the FTase from P. pastoris GS115 [38]. For the industrial application of the enzyme, it is desirable to operate at temperature above 60 °C in order to avoid microbial contamination. Fig. 4d showed that the FTase retained approximately 89% of its activity after incubation at 60 °C for 2 h. However, its activity declined rapidly at temperatures above 65 °C. The FTase was stable at 25 °C for 12 h, and retained approximately 65% of its activity after incubation for 24 h. The FTase activity began to decline due to thermal denaturation of proteins, and most research described an upper temperature limit of 40 °C [39–41] to 50 °C [32,36]. Significantly, The FTase of A. orayze S719 could produce FOS under high temperature of 60 °C. It is a known fact that high temperature fermentation not only decreased the sterilization cost, but also reduced the contamination possibilities. Compared with the conventional fermentation, the consumption of cooling water could also greatly reduced in high temperature fermentation, which resulted in lower energy consumption and production cost for large-scale industrial fermentation especially in tropical or subtropical areas. In addition, the enzyme was capable of maintaining high activity at room temperature (25 °C), which was advantageous for the application of this enzyme in the largeterm transportation.
Fig. 5. Lineweaver–Burk plots for using sucrose as substrate by FTase. Values are means of three independent experiments.
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Table 2 Effect of metal ions on the activity of FTase. Metal ions
2+
Ca Mg2+ Zn2+ Cu2+ Pb2+ Fe2+ Mn2+ EDTA Na+ CKb
Concentration relative activity (%)a 5 mM
10 mM
97 ± 2.3 116 ± 1.3 19 ± 1.5 2.4 ± 2.7 1.9 ± 2.4 0 70 ± 2.2 96 ± 1.4 91 ± 1.9 100
91 ± 2.1 89 ± 2.4 10 ± 1.8 2.3 ± 1.6 0 ± 1.0 0 80 ± 1.3 91 ± 1.2 114 ± 2.1 100
(a)The relative activity with different metal ions added was determined and compared with the activity measured in Phosphate buffer (pH 5.5, 0.2 M) without the addition of any ions. (b) The relative activity measured in Phosphate buffer (pH 5.5, 0.2 M) without the addition of any ions. The experiments were independently performed at least three times. The errors are shown as standard deviation (SD).
Fig. 6. FOS production by FTase G: glucose, GF: sucrose, GF2: 1-kestose, GF3: nystose, GF4: fructosylnystose, FOS: fructooligosaccharides. The synthesis of FOS was carried out for 24 h by using sucrose as initial substrate (500 g L−1) at 50 °C and pH 5.5. Values are means of three independent experiments.
3.5. Kinetic studies The kinetics of FTase was determined using sucrose as a substrate. The Km and Vmax of FTase were 310 mmol L−1 and 1.4 mmol L−1 min−1, respectively (Fig. 5). The kcat and kcat/Km of FTase were 2.0 × 103 min−1 and 6.4 L mmol−1 min−1, respectively. These results indicated that FTase has high catalytic efficiency. The kinetic constant of FTase from Rhodotorula sp was determined. The Km value of sucrose was 198 g L−1 [42]. Jung et al. [43]determined the properties of the FTase from Aureobasidium pullulans, and found that the Km value and Vmax value for sucrose were 330 g L−1 and 130 g (L h)−1. It was indicated that a higher substrate affinity of these FTase from A. pullulans and Rhodotorula sp. was lower than that of FTase heterologously expressed in a A. oryaze S719 in this work. The lower Km value of the purified A. oryaze S719 FTase found here indicated that it had a greater affinity for the substrate.
HPLC demonstrated that within 4 h of the reaction, the yield of FOS was over 0.64 g of FOS/g of sucrose and percentages of GF2, GF3 and GF4 were 67%, 32% and 0.06% (data not shown) whereas within 6 h of the reaction, the yield of FOS was 0.64 g of FOS/g of sucrose and percentages of GF2, GF3 and GF4 were 56%, 42% and 1.4%. These results demonstrated that the FTase has high transfructosylation efficiency towards the synthesis of FOS from fructosyl groups and sucrose. The result comparing A. oryaze S719 production of FOS with other strains were shown in Table 3. The purification of the FTase from A. orayze S719 obtained a yield of 64% of FOS within 4 h of reaction, indicating a considerable reduction in the reaction time of 8–24 h reported in the literature. It also was the shortest reaction time to be reported regarding the purified enzyme production of FOS. This further indicated that the use of A. oryaze S719 to produce FOS could save more time compared to other strains. In the industry, it is known that shortening production time would not only speed up the production process, but also reduce waste of resources and reduce energy consumption. According to the above results, the A. oryaze S719 strain with high transfer efficiency would have the potential to lay a foundation for its application in FOS production. In addition, to the best of our knowledge, A. oryzae S719 strain therefore represents the first reported non-recombinant fructosyltransferase that have shortest reaction time for the production of FOS. According to the above results, the A. oryaze S719 strain with [8] high transfructosylation efficiency has the potential to replace the recombinant strain producing FOS.
3.6. Effect of metal ions on FTase activity The effect of different metal ions on FTase activity was determined (Table 2). FTase was significantly activated by 5 mmol L−1 Mg2+ and 10 mmol L−1 Na+, respectively, which indicated that the FTase was Mg2+ and Na+ dependent, with relative activities of 116 and 114%, respectively. The Na+ (5 mmol L−1), Ca2+ (5 mmol L−1), and EDTA (5 mmol L−1) had no effect on FTase activity. The enzyme was not inhibited by 5 mM EDTA indicating that it was not a metalloprotein. Other metal ions inhibited FTase activity, but the precise mechanisms by which heavy metal ions inhibit the FTase activity were unknown [44]. Yang et al. [44] analyzed effect of metal ions on the activity of the FTase from Pichia pastoris GS115 and found that Mg2+ and Ni+ could enhance its initial activity.
4. Conclusion We have isolated and purified a novel fructosyltransferase form A. oryzae S719, which was identified as a type of FTase by MS. The purified FTase from A. orayze S719 obtained a yield of 64% of FOS within 4 h of reaction, which was the shortest reaction time to be reported regarding the purified enzyme production of FOS. The molecular weight of the purified FTase was around 95 kDa, and the specific activity was 4200 U mg−1. This FTase had a high relative activity at temperatures ranging from 25 to 65 °C, and the FTase was stable at wide pH ranges and retained more than 80% of its activity at pH 4.0–11. The FTase had high catalytic efficiency, the kcat of which was 2.0 × 103 min−1. The FTase was Mg2+ and Na+ dependent. This study provided a novel FTase with high transfer efficiency and would have the potential to lay a foundation for its application in FOS production.
3.7. FOS production According to the results of 3.3 and 3.4, it was found that 12 U of FTase activity/g of sucrose, 500.0 mg sucrose mL−1, pH 6.0 in 0.1 M sodium acetate buffer and reaction temperature of 55 °C were the most suitable for the transfructosylation reaction. As shown in Fig. 6, several important time process parameters were monitored during the transglucosidic assay including FOS yield, yield of each oligomer, and residual sucrose. As the reaction time increased, the amount of sucrose was decreased and the amounts of glucose and FOS were increased, but the released fructose could not be detected. Analysis of the FOS generated, glucose (G), fructose (F) and sucrose (GF) standards by using 7
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Table 3 Comparison of some parameters obtained for different strains in the present study and the literature. Strains A.oryzae S719 A.tubingensis XG21 PichiapastorisGS115 Aureobasidium melanogenum 11-1 A. pullulans CFR77
Enzyme conc −1
12 U g 6 U g−1 8 U g−1 117 U g−1 NS
Carbon source Sucrose Sucrose Sucrose Sucrose Sucrose
(400 (200 (500 (300 (600
g g g g g
−1
L ) L−1) L−1) L−1) L−1)
Cultivation time
FOS yield (%)
References
4 8 18 7 9
63% 57% 64% 66% 59%
Present study Xie et al. [45]. Guo et al. [15]. Jiang et al. [31]. Agbaje et al. [34].
NS: not stated.
Decalaration of competing interest [17]
The authors declare that they have no conflict of interest.
[18]
Acknowledgments
[19]
This work was financially supported by the Natural Science Foundation of Guangxi Province (Grant No. 2016GXNSFAA380130, 2017GXNSFAA198010, 2018GXNSFAA281019 and 2018GXNSFAA 138024) and the National Natural Science Foundation of China (Grant No. 31560448, 31660251 and 31860245) and the central government directs special funds for local science and technology development projects (ZY1949015).
[20] [21] [22] [23]
Appendix A. Supplementary data
[24]
Supplementary data to this article can be found online at https:// doi.org/10.1016/j.pep.2019.105549.
[25]
References
[26]
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Abbreviations
[41] J. Cha, et al., Molecular and enzymatic characterization of a levan fructotransferase from Microbacterium sp. AL-210, J. Biotechnol. 91 (1) (2001) 49–61. [42] M. Alvarado, F. Maugeri, Kinetic modelling of fructooligosaccharide synthesis using a Rhodotorula sp. fructosyltransferase, J. Biotechnol. 131 (2) (2007) S91–S92. [43] K.H. Jung, et al., Mathematical model for enzymatic production of fructo-oligosaccharides from sucrose, Enzym. Microb. Technol. 11 (8) (1989) 491–494. [44] K.H. Jung, et al., Mathematical model for enzymatic production of fructo-oligosaccharides from sucrose, Enzym. Microb. Technol. 11 (8) (1989) 491–494. [45] Y. Xie, et al., A molasses habitat-derived fungus Aspergillus tubingensis XG21 with high β-fructofuranosidase activity and its potential use for fructooligosaccharides production, Amb. Express 7 (1) (2017) 128.
FOS: Fructooligosaccharides FTase: β-fructofuranosidases SDS-PAGE: sodium dodecyl sulphate polyacrylamide gel electrophoresis GF2: 1-kestose GF3: 1-nystose GF4: 1-fructofuranosylnystose UV: ultraviolet BSA: Bovine albumin NCBI: National Center for Biotechnology Information
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