Highly Efficient Synthesis of Fructooligosaccharides by Extracellular

Aug 5, 2016 - In this work, Aspergillus aculeatus M105 was obtained to produce high extracellular fructooligosaccharide-producing enzyme activity. The...
1 downloads 22 Views 798KB Size
Download PDF
Article pubs.acs.org/JAFC

Highly Efficient Synthesis of Fructooligosaccharides by Extracellular Fructooligosaccharide-Producing Enzymes and Immobilized Cells of Aspergillus aculeatus M105 and Purification and Biochemical Characterization of a Fructosyltransferase from the Fungus Mei-Ping Huang,∥ Min Wu,∥ Qiang-Sheng Xu, De-Jiao Mo, and Jia-Xun Feng* State Key Laboratory for Conservation and Utilization of Subtropical Agro-bioresources, Guangxi Key Laboratory of Subtropical Bioresources Conservation and Utilization, College of Life Science and Technology, Guangxi University, 100 Daxue Road, Nanning 530004, China S Supporting Information *

ABSTRACT: In this work, Aspergillus aculeatus M105 was obtained to produce high extracellular fructooligosaccharideproducing enzyme activity. The maximum yields of fructooligosaccharides produced by its extracellular enzymes and immobilized cells were 67.54 and 65.47% (w/w), respectively. A fructosyltransferase (FTase), AaFT32A, was purified from M105. The optimal pH and temperature of AaFT32A were pH 5.0−6.0 and 65 °C, respectively. The Km, Vmax, and kcat values for the transfructosylating activity of AaFT32A were 2267 mM, 1347 μmol/min/mg protein, and 1550.2 s−1, respectively, and those values for the hydrolytic activity of AaFT32A were 6.10 mM, 32.44 μmol/min/mg protein, and 37.3 s−1, respectively. The sequence of AaFT32A deduced from the cloned gene shared 99.4% identity with a FTase from Aspergillus japonicus CB05 and a fructofuranosidase from Aspergillus niger and 96.5% identity with a FTase (Aspacl_37092) from A. aculeatus ATCC 16872. The fungal strain and its FTase may have potential applications in the prebiotics industry. KEYWORDS: Aspergillus aculeatus, fructooligosaccharides, fructooligosaccharide-producing enzyme, fructosyltransferase, immobilized cells

■

INTRODUCTION The nutritional and health benefits of prebiotic oligosaccharides have received particular attention in recent decades. One such prebiotic, fructooligosaccharides (FOSs), can selectively stimulate the growth of intestinal microflora, especially Bifidobacterium, improve blood lipid composition in hyperlipidemia, decrease serum cholesterol levels, and promote calcium and magnesium absorption.1−4 In addition, they also have good physical properties, as they are low-calorie, noncariogenic, and nondigestible. Therefore, FOSs have been greatly explored in the food, feed, and pharmaceutical fields. Commercial FOSs mainly consist of 1-kestose (GF2), 1-nystose (GF3), and 1Ffructofuranosylnystose (GF4) in which one to three fructosyl residues are linked to sucrose via β-2,1 glycosidic bonds.4,5 FOSs are mainly synthesized from sucrose by various microorganisms via the action of extracellular/intracellular fructosyltransferases (FTases, EC 2.4.1.9) or β-fructofuranosidases (FFases, EC 3.2.1.26) with high transfructosylating activities.5,6 These two enzymes possess both transfructosylating and hydrolytic activities, but FTases almost exclusively exhibit transfructosylating activity when the sucrose concentration exceeds 100 mM.7 In industry, enzymes with high transfructosylating activities are likely to be used for the industrial production of FOSs. However, the main limiting factor in commercial FOS production is that the FOS yield ranges from 50 to 60% (w/w), and the product contains large amounts of reaction byproducts, such as glucose, fructose, and unreacted sucrose.4 © XXXX American Chemical Society

Recently, many studies have attempted to improve FOS production yields. Most of these studies have focused on identifying new microbial strains that produce high levels of FOSs. Several fungal strains, such as Aspergillus spp.8−11 and Aureobasidium spp.,12−14 have been reported to be potentially adequate for the industrial production of FOSs. Ganaie et al.15 identified a strain with high fructosyltransferase activity for transforming sucrose to FOSs by screening biocatalysts. In contrast, some studies improved FOS yields by promoting the use efficiency of enzymes or cells. Immobilization systems are common methods to reuse enzymes/cells to increase their efficiency and productivity, as well as to allow easier separation of the enzymes/cells from the fermentation broth.16,17 Compared with other methods, entrapment within insoluble calcium alginate beads has been shown to be the most effective approach because of its biocompatibility, low cost, and resistance to microbial contamination.18 FFases from Aspergillus japonicus19,20 and whole cells of Penicillium citrinum16 were immobilized by sodium alginate and exhibited good properties during FOS synthesis. Despite extensive studies of FTases and FFases from diverse microorganisms, it is still necessary to identify strains that produce extracellular FTases and FFases with high FOS yields and characterize their enzyme properties and coding genes. The Received: May 22, 2016 Revised: August 3, 2016 Accepted: August 5, 2016

A

DOI: 10.1021/acs.jafc.6b02115 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

a syringe (1D 1.0 mm). Beads were formed and kept at 4 °C for 24 h prior to use.22 FOSs Production from Sucrose. To investigate FOS production, crude extracellular enzyme from A. aculeatus strain M105 was added to sucrose solution. The final dosage of the enzyme and the sucrose concentration were 9.0 U/g and 600 g/L, respectively. Deionized water was added to a final volume of 30 mL. FOS-producing reactions were conducted at 40 °C. Immobilized cells formed by 1 g of mycelia in 3% alginate were added to 10 mL of sucrose at 600 g/L (pH 7.5), and the reaction was conducted at 45 °C for different periods of time. All of the products were analyzed by HPLC. The percentage of FOSs was calculated using the following formula:

main objective of this study was to find new microbial strains with high extracellular fructooligosaccharide-producing (FOSproducing) enzyme activity, to improve the strains’ production of enzymes responsible for the biotransformation of sucrose to FOSs, and to further purify and characterize the responsible enzymes.

■

MATERIALS AND METHODS

Chemicals. Fructose, glucose, sucrose, GF2, GF3, fluorescein-5thiosemicarbazide, and peptide-N-glycosidase F (PNGase F) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Standard GF4 was purchased from Megazyme (Wicklow, Ireland). Five-milliliter HiTrap QFF and HiPrep16/10 PhenylFF columns were purchased from GE Healthcare Life Sciences (Uppsala, Sweden). Molecular weight markers for sodium dodecyl sulfate−polyacrylamide gel electrophoresis (SDS-PAGE) were obtained from Thermo Fisher Scientific (Waltham, MA, USA). All other chemicals were of analytical grade and obtained from readily available commercial sources. Fungal Strains and Cultivation Conditions. Twenty-nine fungal strains, which were used to screen enzymes with extracellular FOSproducing enzyme activity, were obtained from the microbial culture collection of the authors’ laboratory. The fungal strains were cultivated on potato dextrose agar (PDA) plates at 28 °C for 6 days to collect spores, and a 1.5 mL of spore suspension (1 × 108 spores/mL) was inoculated into basal medium (150 mL in a 500 mL flask) consisting of (w/v) 3% sucrose, 0.15% yeast extract, 0.15% peptone, 0.3% NaNO3, 0.1% KH2PO4, 0.05% KCl, 0.05% MgSO4·7H2O, and 0.001% FeSO4·7H2O. To prepare crude enzymes and mycelia for immobilization, the culture was incubated at 28 °C with shaking at 180 rpm for 6 and 2 days, respectively. Identification and Mutagenesis of the Fungal Strain Producing High FOS-Producing Enzyme Activity. The fungal strain GXU15 was identified as Aspergillus aculeatus, and a mutant strain, M105, with higher FOS-producing enzyme activity, was selected by Co60 radiation of GXU15 (see the Supporting Information). Analytical Methods of FOSs. Quantitative Analysis of FOSs by High-Performance Liquid Chromatography (HPLC). Quantitative analysis of the carbohydrates was performed by HPLC (LC-10AT, Shimadzu, Kyoto, Japan) equipped with a refractive index detector. A 5-μm YMC-NH2 column (4.6 × 250 mm, YMC Co., Ltd. Japan) was used at 30 °C with acetonitrile/water (75:25, v/v) as the mobile phase at a flow rate of 1.0 mL/min.6 Enzyme Activity Assay. The FOS-producing enzyme activity of the fungal strains was determined using the DNS method21 with minor modifications. The reaction mixture for determining the enzyme activity consisted of 10% (w/v) sucrose (pH 6.0, 350 μL) as the substrate and the enzyme solution (50 μL). The reaction was performed at 65 °C for 30 min, stopped by adding 800 μL of DNS reagent, and then placed in boiling water for 5 min for color development. One unit of enzymatic activity was defined as the amount of enzyme required to produce 1 μmol of reducing sugars per minute under the above conditions. Unless stated otherwise, all determinations of FOS-producing enzyme activity used this method. Effects of the Sucrose Concentration and Enzyme Dosage on the Enzymatic Synthesis of FOSs. The effect of the sucrose concentration on the yield of FOSs was determined using the FOSproducing enzyme at a fixed activity dosage (4.5 U/g sucrose) during the synthesis of FOSs from different sucrose concentrations (400, 500, and 600 g/L). To study the effect of enzyme loading on the FOSs yield, a range of FOS-producing enzyme dosages (4.5, 6.0, 7.5, and 9.0 U/g sucrose) and a sucrose concentration of 600 g/L were used for the FOS synthesis. The total volume of each reaction system was 30 mL. The synthesis reactions were conducted at pH 6.0 and 40 °C for different periods of time. The products were analyzed using HPLC. Mycelia Immobilization. Immobilization of mycelia of fungal strain M105 was achieved by mixing 1 g of mycelia with sodium alginate (3%, w/v) at room temperature and with stirring to form a slurry. Then, drops were extruded into a CaCl2 solution (0.3 M) using

FOSs (%) =

GF2 + GF3 + GF4 (g) × 100 total sugars (g)

Purification of the Enzyme from A. aculeatus M105. The cell culture was filtered through multilayer gauze and centrifuged for 10 min at 7100g to obtain the crude enzyme (160 mL, 23.58 U/mL). Proteins in the solution were precipitated by slowly adding ethanol; the ratio of the volume of the protein solution to the volume ethanol solution was 1:0.75. After stirring on ice for 15 min, the liquor was centrifuged at 7100g at 4 °C for 15 min to obtain the supernatant. More ethanol was slowly added to the supernatant, until the ratio of the volume of the supernatant to the volume of ethanol solution was 1:1.5. After stirring on ice for 15 min, the precipitated proteins were collected by centrifugation at 7100g at 4 °C for 15 min and dissolved in 20 mM Tris-HCl buffer (pH 8.0). The enzyme solution was loaded onto an anion-exchange column (5-mL HiTrap Q FF) and eluted with Tris-HCl buffer (pH 8.0) using a linear gradient of 0−1 M NaCl. The active fractions were combined and concentrated. After the combined enzyme fractions were dialyzed against 50 mM phosphate buffer (pH 6.5) containing 2 M (NH4)2SO4, they were applied to a HiPrep 16/10 Phenyl FF (high sub) column (GE Healthcare Life Sciences), and the enzyme was eluted with a linear gradient from 2 to 0 M (NH4)2SO4 in 50 mM phosphate buffer (pH 6.5). The separated and purified active fractions were pooled and concentrated. The molecular weight of the enzyme was determined by 12% SDS-PAGE using a Mini-gel system (Bio-Rad, Hercules, CA, USA) as described previously by Laemmli.23 SDS-PAGE was performed with a 5% acrylamide stacking gel (pH 6.8) and a 12% separating gel (pH 8.8). The gel was stained with Coomassie brilliant blue R-250. Effects of pH and Temperature on Enzymatic Activity. The optimal pH of the purified enzyme was determined by measuring its enzymatic activity at different pH values using citrate−phosphate buffer (pH 3.5−6.5), phosphate buffer (pH 6.5−7.5), Tris-HCl buffer (pH 7.5−8.5), and glycine−NaOH buffer (pH 8.5−10.5) at 37 °C. To determine the effect of temperature on the enzymatic activity, the activity of the purified enzyme was measured at temperatures ranging from 30 to 80 °C at the optimal pH. The highest enzymatic activities obtained at the different pH values and temperatures were considered to be 100%. To determine the pH stability of the enzyme, the purified enzyme was incubated at pH values ranging from 3.5 to 10.5 at 4 °C for 24 h, and the residual activity was measured under the optimal condition. Thermal stability was investigated by pre-incubating the enzyme at 30−60 °C for 1−5 h, and the residual activity was determined under the optimal condition. The enzymatic activities prior to the incubations at different pH values and temperatures were measured and considered to be 100%. Effects of Different Metal Ions and Chemical Reagents on Enzymatic Activity. The effects of different metal ions and chemical reagents on enzymatic activity are provided in the Supporting Information. Determination of Kinetic Parameters. For the kinetic experiments, substrates at different concentrations (0.01−1.6 M) were prepared in phosphate buffer (pH 6.0) and mixed with the purified enzyme. The reactions were conducted under the optimal temperature B

DOI: 10.1021/acs.jafc.6b02115 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry (65 °C) for 30 min and terminated by heating the tubes in boiling water for 10 min. The amounts of fructose and glucose in the reaction products were analyzed by HPLC. The amount of free fructose detected is a direct determination of the hydrolyzing activity of the enzyme, whereas the amount of glucose minus the amount of free fructose represents the transferring activity of the enzyme. One unit of hydrolyzing activity was defined as the amount of enzyme needed to release 1 μmol of fructose per minute, whereas 1 unit of transferring activity was defined as the amount of enzyme required to catalyze the transfer of 1 μmol of fructose per minute.6 The Km, Vmax, and kcat values were calculated using GraphPad Prism5 software (La Jolla, CA, USA). Mass Spectrometric Analysis of the Purified Enzyme. The purified protein was separated on a 12% polyacrylamide gel. After the separation, the gel was stained by Coomassie brilliant blue R250. The target protein gel band was cut from the gel for in-gel digestion. Gel particles were washed three times with Milli-Q water and destained with an acetonitrile/water solution (1:1, v/v). For protein reduction and alkylation, gel particles were washed with 40 mM ammonium bicarbonate and incubated in 10 mM dithiothreitol and 55 mM iodoacetic acid at 55 °C and room temperature in the dark. Gel particles were dehydrated with 100% acrylonitrile, and the dry gel particles were saturated with trypsin and incubated overnight at 37 °C. The peptides were extracted by incubating the gel pieces twice in the extraction solution (1:1, v/v acetonitrile/water solution). The peptides were subjected to HPLC on a Nano-LC system (easy nLC-1000, Thermo Fisher Scientific) connected to a LTQ Orbitrap Elite mass spectrometer (Thermo Scientific, Bremen, Germany). The detailed procedures followed the methods reported by Zhang et al.24 Cloning of the Gene Encoding the Purified Enzyme. Genomic DNA was extracted using Horiuchi’s protocol,25 with some modifications. Total RNA was extracted using Trizol (Tiangen Biotech, Beijing, China) according to the manufacturer’s instructions. cDNA was obtained from the total RNA using the Reverse Transcription Kit (Tiangen Biotech). All of the protein MS detection results were analyzed using the Basic Local Align Search Tool (BLAST) at the National Center for Biotechnology Information (NCBI) Web site (http://www.ncbi.nlm. nih.gov/); the most similar FTase protein sequence was identified, and the corresponding nucleotide coding sequence was obtained. On the basis of the nucleotide sequence, forward 5′-ATGAAGCTCMCCACTACCAC-3′ and reverse 5′-TCACTTTCTCTCCGGCCAGGCGTT-3′ primers were designed to amplify the purified enzyme-encoding gene. The genomic DNA and cDNA from A. aculeatus strain M105 were used as templates. PCR amplification was conducted as follows: one cycle at 94 °C for 5 min, followed by 30 cycles at 94 °C for 30 s, 56 °C for 30 s, and 72 °C for 90 s, followed by one cycle at 72 °C for 10 min. The PCR products were purified, cloned into the pEASY-Blunt Cloning Vector (TransGen Biotech), and transformed into Escherichia coli Trans1-T1 phage-resistant competent cells. Then, the correct transformants were sequenced and analyzed. The upstream and downstream flanking sequences of the obtained sequence were amplified using the Genome Walking Kit (Takara, Shiga, Japan) according to the manufacturer’s instructions. Sequence analyses were conducted using the DNAStar software package, Vector NTI 11.0 software, NCBI-BLAST, CBS Prediction Servers (http://www.cbs.dtu.dk/services/), and the Simple Modular Architecture Research Tool (SMART) (http://smart.embl-heidelberg. de/). Nucleotide Sequence Accession Numbers. Sequences of the partial ITS, the β-tubulin gene in strain GXU15, and DNA and mRNA encoding the enzyme AaFT32A in strain M105 were deposited in the GenBank database under the accession numbers KU310905, KU310906, KX098014, and KU310907, respectively.

aculeatus M105 was reacted with 600 g/L sucrose at 40 °C for 10 h, GF2, GF3, and GF4 were produced (Figure 1), which

Figure 1. HPLC analysis of the reaction mixture catalyzed by crude FOS-producing enzyme from fungal strain M105: (A) standard samples; (B) the reaction mixture after 10 h. Peaks: 1, fructose; 2, glucose; 3, sucrose; 4, GF2; 5, GF3; 6, GF4.

confirmed that the enzyme exhibited extracellular FOSproducing enzyme activity. The optimal pH and temperature of the FOS-producing enzyme from strain M105 were pH 4.5− 5.5 and 65 °C, respectively. The enzyme was remarkably stable at pH 3.5−10.5 and below 40 °C (data not shown). Thus, the effects of the sucrose concentration and enzyme dosage on FOS synthesis were measured at pH 5.0 and 40 °C. Solutions of different sucrose concentrations (400, 500, and 600 g/L) were mixed with the crude enzyme at 4.5 U/g sucrose, and FOS production was measured. As shown in Figure 2A, total FOS production increased to the same extent (65−67%, w/w) by increasing the initial sucrose concentration, and no significant differences were observed in the FOS yields at sucrose concentrations ranging from 400 to 600 g/L. When different enzyme doses (4.5−9.0 U/g) were reacted with 600 g/L sucrose, the FOS synthesis by different enzyme dosages at 4.5, 6.0, 7.5, and 9.0 U/g were 233.83, 271.91, 312.06, and 328.56 g/L, respectively, at the first 3 h. It was clear that the more enzyme dosage added, the faster was product synthesis. However, it was noteworthy that the total FOS production increased to the same extent when the maximum FOS yields were reached (Figure 2B). Therefore, the increase in enzyme dosage only increased the reaction rate, but did not significantly affect the FOS yield. Similar results were found in previous studies.8,26 FOS Production by Crude Enzyme from A. aculeatus Strain M105. The reaction progress at an enzyme dosage of 9.0 U/g sucrose and 600 g/L sucrose at 40 °C is depicted in Figure 3. The FOS yield reached a maximum value of 394.42 g/ L after 12 h (205.41 g/L GF2, 176.45 g/L GF3, and 12.56 g/L GF4), which corresponded to 67.54% (w/w) of the total carbohydrates in the mixture. At the beginning of the reaction, GF2 was synthesized first and reached its maximum yield of 261.03 g/L at 3 h, and then the yield decreased slightly. GF3 increased continuously during the reaction, whereas GF4 was

■

RESULTS AND DISCUSSION Effects of the Sucrose Concentration and Enzyme Dosage on FOS Synthesis. When the supernatant of A. C

DOI: 10.1021/acs.jafc.6b02115 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

Figure 2. Enzymatic synthesis of FOSs by A. aculeatus strain M105: (A) effect of sucrose concentrations on the FOS yield (substrate concentrations were 400 (■), 500 (▲), and 600 g/L (◆); (B) effect of enzyme dosages on the FOS yield (reactions were conducted at pH 6.0 and 40 °C for 27 h; substrate concentration was 600 g/L). Data are the means ± standard deviation from three replicates. Different enzyme dosages were used in the syntheses: 4.5 (◆), 6.0 (■), 7.5 (▲), and 9.0 U/g (●).

Figure 4. Time course of FOS synthesis by immobilized cells from A. aculeatus strain M105: (◆) fructose; (■) glucose; (▲) sucrose; (×) GF2; (●) GF3; (+) GF4; (□) total FOSs. The reaction was conducted at 45 °C in a 600 g/L sucrose solution. The total volume of the reaction system was 10 mL.

Figure 3. Time course of FOS synthesis by crude enzyme from A. aculeatus strain M105: (◆) fructose; (■) glucose; (▲) sucrose; (×) GF2; (●) GF3; (+) GF4; (□) total FOSs. The reaction was conducted at 40 °C in a 600 g/L sucrose solution. The total volume of the reaction system was 30 mL.

w).10 Compared with the aforementioned yield of FOSs by the crude enzyme, the maximum yield of GF2 was just 28.65%, and the amount of GF2 was less than that of GF3 when the maximum yield of FOSs was reached. However, when the crude enzyme was reacted with sucrose, primarily GF2 accumulated first, and the maximum yield of GF2 achieved was 43.51%. As the maximum yield of FOSs was reached, the amount of GF2 was higher than that of GF3. It seemed that the accumulation of GF2 contributed to the higher FOSs synthesis. In addition, the immobilized cells could be reused to transform sucrose at least 5 times in buffer system and 15 times in nonbuffer system (data not shown), indicating that this process has better potential for FOS production. A previous study showed that the relative activities of dried alginateentrapped enzymes were still stable after they were recycled 13 times.30 For P. citrinum,22 immobilized cells could continue to produce neo-FOSs for 50 days. Ganaie et al.20 revealed that the recycling efficiency of alginate beads was better than that of chitosan beads. Although our study was preliminary, it still showed that immobilized cells from strain M105 have potential industrial-scale applications. Purification and Identification of a FOS-Producing Enzyme from A. aculeatus M105. To further understand the characteristics of the enzyme responsible for the high FOSproducing activity from A. aculeatus M105, it was purified in three steps: ethanol precipitation, ion-exchange chromatography, and hydrophobic interaction chromatography. The collected protein solutions of each purification step were

present until 12 h (Figure 3). These results indicated that the synthesis of FOSs was always sequential, in the order GF → GF2 → GF3 → GF4.19 Throughout the time course of the reaction, the remaining sucrose concentration was approximately 10%. The main reason for the remaining sucrose was that GF2 acts as a donor and acceptor to form GF3 and also because of glucose inhibition.27 After reaching the maximum FOS yield, FOSs were not hydrolyzed as the reaction proceeded, whereas during the entire reaction process, the level of fructose was negligible. This indicated that the crude enzyme produced by strain M105 had high transfructosylating activity. Crude FTases from A. flavus15 and Penicillium expansum28 achieved 63.40 and 58% FOS yield, respectively. Thus, the FOS yield of the crude enzyme from strain M105 was higher than those reported previously, indicating that the enzyme from strain M105 has a strong transfructosylating activity and great potential in FOS production. FOS Production by Immobilized Mycelia of Strain M105. The progress of the reaction using immobilized cells, in the presence of 600 g/L sucrose under the optimal reaction conditions, was also measured. The maximum FOS yield for immobilized cells reached 65.47% (w/w) (24.42% GF2, 34.34% GF3, and 6.71% GF4) after 9 h (Figure 4). This yield was lower than the yield of A. flavus (67.75%, w/w);20 however, it was higher than that of most strains, such as A. japonicas (61.3%, w/ w),19 A. aculeatus (61.5%, w/w),29 and A. oryzae (51.9%, w/ D

DOI: 10.1021/acs.jafc.6b02115 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry analyzed by SDS-PAGE analysis, which finally showed a single protein band (Figure 5A). The overall purification processes of the enzyme are summarized in Table 1.

(LC-MS/MS). The peptide sequences that matched the mass spectra of the peptides released from the purified enzyme of A. aculeatus strain M105 were LDQGPVIADHPFAVDVTAFR, NAPWYVAVSGGVHGVGPAQFLYR, VQTVENVVDNELVR, SQTSAAAPTNPGLDSFTESGK, VEFSPSMAGFLDWGFSAYAAAGK, GSETQSLAVAR, TLGITIAR, FALSTWAR, YVSFVWLTGDQYEQADGFPTAQQGWTGSLLLPR, and VLPASSAVSK, all of which were found in a putative FTase from A. japonicus CB05 (NCBI accession no. ADK46938.1) and a FFase from A. niger ATCC 20611 (NCBI accession no. BAB67771.1), which belong to the GH 32 family. Moreover, in catalyzing 100 mM sucrose, the amount of glucose detected was significantly higher than that of fructose (Figure 5C), indicating that the enzyme exhibited higher transfructosylating activity than hydrolytic activity. This phenomenon is in accordance with the fact that FTases almost exclusively exhibit transfructosylating activity at sucrose concentrations exceeding 100 mM.7 On the basis of the HPLC analysis, the oligosaccharide products were identified as GF2 and GF3, and the maximum FOS yield was 63.56% (w/w) (158.17 g/L GF2 and 102.01 g/L GF3). The final yield of FOSs (63.56%) was higher than that from A. aculeatus (60.7 and 61.4%).6,31 Together, the MS results confirmed that the protein purified from strain M105 was a FTase, which was named AaFT32A. Effects of pH and Temperature on the Activity and Stability of AaFT32A. The optimal pH for AaFT32A activity ranged from pH 5.0 to 6.0, and the relative activities declined significantly below pH 4.0 and above pH 7.0 (Figure 6A). The optimal pH values of FTases (AcFT1 and AcFT2) from A. aculeatus31 were 5.5 and 5.0. The optimal pH of the FTase from A. pullulans36 was 5.0, whereas those of FTases from A. oryzae32,37 and Streptococcus salivarius38 were 6.0 and those of FFases from P. oxalicum35 and A. niger39 were 5.5 and 5.8, respectively. The highest enzymatic activity was observed at 65 °C, and no significant activity was found at temperatures 80 °C (Figure 6B). The optimal temperature of FTases from A. aculeatus6,31 and A. oryzae37 was 60 °C, whereas the optimal temperature of a FTase from S. salivarius38 was 37 °C. AaFT32A was remarkably stable over a broad pH range (5.5−10.5) and at temperatures