Isolation of Fructosyl Transferase from Aspergillus niger Strain SS274 and Enhancing its Stability by Harnessing Magnetic Nanoparticles Assisted Enzyme Immobilization Strategy
Saba Iqbal1*, Zahoor Qadir Samra1 and Muhammad Imtiaz Shafiq2
1School of Biochemistry and Biotechnology, University of the Punjab, Lahore
2School of Chemistry, University of the Punjab, Lahore
ABSTRACT
Fructosyl trasferase is considered as an important industrial enzyme for production of prebiotics that hold numerous health benefits, is purified from Aspergillus niger strain SS274 using size exclusion chromatography Sephadex G-100 showed 91% of sample recovery, 3.57 purification fold and 393.32U/mg of specific activity. It was a glycoprotein dimer enzyme of approximately 75kDa molecular weight with 2 subunits of 45kDa and 30kDa which showed optimal activity from 50-60 ℃ and at pH-5 and its activity was increased up to 4% from normal activity in presence of Fe metal ions while Hg and Ag showed inhibitory effect on enzyme’s activity. Formation of prebiotics or fructooligosaccharides (FOS) was confirmed through HPLC analysis using sucrose as a substrate. Enzyme was further immobilized on magnetic nanoparticles using carbodiimide method to make it more industrial viable by increasing it shelf life and stability, its activity was observed in mobilize and immobilize state for period of 6 months stored at 4 ℃ and -20 ℃ and it showed maximum activity and better shelf life at -20 ℃ in immobilize form. Fructosyl transferase from A. niger have good potential for FOS production at industrial scale for health applications.
Article Information
Received 25 April 2024
Revised 25 June 2025
Accepted 03 July 2025
Available online 10 October 2025
(early access)
Published 04 April 2026
Authors’ Contribution
Conceptualization and design of study SI and ZQS, Acquisition of data SI, Analysis and interpretation of data SI and MIS, Drafting the article SI, Revising content SI, ZQS and MIS. Final approval of version SI and ZQS. All authors have read and agreed to the published version of the manuscript.
Key words
FTase, Fructooligosaccharides, Prebiotics, HPLC, Magnetic nanoparticles, FTIR
DOI: https://dx.doi.org/10.17582/journal.pjz/20240925110813
* Corresponding author: [email protected]
0030-9923/2026/0003-1311 $ 9.00/0
Copyright 2026 by the authors. Licensee Zoological Society of Pakistan.
This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/).
Introduction
Nowadays, foods that contributes to maintain good health have gained prominence due to their increased demand. People are getting more conscious about their food intake and possible effect of the ingredients present in food (Trujillo et al., 2019). Non-digestible ooligosaccharides are one among of them due to their health-related benefits among consumers (Mussatto and Mancilha, 2007). Fructooligosaccharides (FOS) belong to the class of oligosaccharides, that are low molecular weight carbohydrates, which basically contained 2 to 10 monosaccharides units linked through glycosidic linkages (Sabater-Molina et al., 2009). FOS being low caloric, mild sweet and non-carcinogenic are one of the important ooligosaccharides (Dhake and Patil, 2007; Pool-Zobel et al., 2002) and have proved prime in proliferating gut microflora of humans (Antošová and Polakovič, 2001; Campbell et al., 1997; Ozcan and Kurtuldu, 2014). They could easily be fermented by gut microflora bifidobacterial and lactobacilli to enhance their growth and function (Markowiak and Ślizewska, 2017; Trujillo et al., 2019). FOS due to their lower molecular weight show hygroscopic function and play important role in water retention, they also show good thermal stability due to their higher viscosity (Crittenden and Playne, 1996; Roberfroid, 2007). FOS possess many health benefits, cholesterol lowering effect, mineral absorption in body and cancer suppression has been studied throughout many years (Khanvilkar and Arya, 2015; Pool-Zobel et al., 2002; Rao, 2001).
FOS are represented by formula GFn, chain of fructose oligomers attached with terminal glucosyl (Michel et al., 2016; Sangeetha, 2003). 1-kestose (GF2), 1-nystose (GF3) and 1-β-fructofuranosylnystose (GF4) are frequent FOS used in functional foods (Borromei et al., 2009; Dake and Kumar, 2012).Chemical structure of FOS is given in Figure 1 Naturally FOS are present in many fruits and vegetables like banana, onion, garlic chicory, barley and oats, but in low concentrations that they cannot carry out their health-related functions properly (Sridevi et al., 2014; Yun, 1996).
FOS can be enzymatically produced by the action of Fructosyltransferases or (Ftase) enzymes [EC 2.4.1.9] which carry out the transfructsylation of sugars like sucrose, glucose, maltose and fructose (Sangeetha et al., 2004; Yang et al., 2016). Many microorganisms and plants could be source of Ftases (Arthee and Vijila, 2014; Cunha et al., 2019; Dake and Kumar, 2012; Fujishima et al., 2005; Heyer and Wendenburg, 2001; Itaya et al., 1999; Kingston Smith et al., 1999; Koops and Jonker, 1996; Luscher et al., 1993; Lasseur et al., 2009; Van Hijum et al., 2002). FTase catalyzes the chain of reaction employing disproportionate mechanism in which fructosyl moiety of donor sucrose is transferred to another sucrose molecule to release glucose and ooligosaccharides and so on, mechanism can be shown as follows:
GF + GFn G + GFn+1
GF + GFn+1 G + GFn+2
FOS being non-digestible fibers, play an important role in cancer suppression by increasing short chain fatty acids (SCFA) like propionic and butyric acid in gut gradually increasing the proliferation of gut microflora. Increasing SCFA in gut will ultimately decrease the fecal pH and bile acids that are main contributors of colorectal cancer (Bruno-Barcena and Azcarate-Peril, 2015; Dou et al., 2022; Grajek et al., 2005; Pool-Zobel et al., 2002; Rao, 2001).
This study aims to purify extracellular Ftase from Aspergillus niger SS274, its biochemical characterization for production of FOS and its immobilization on MNPs to enhance its shelf life.
Materials and Methods
Aspergillus niger SS274 pure culture was obtained from AMBER laboratory of School of Biochemistry and Biotechnology, University of the Punjab, Lahore used as an inoculum for pre-inoculum media. A loop full of stored inoculum was transferred to pre-inoculum media (yeast extract: 0.2%, sucrose: 2%, pH: 6) under sterile conditions and incubated for 24 h at 30℃ at 100 rpm.
Fermentation media
For extracellular enzyme production 500ml of fermentation media (Sucrose: 4%, Yeast extract: 10g/L, NaNO3: 1g/L, KH2PO4: 1g/L, K2HPO4: 0.5g/L, NaCl: 0.25g/L, MgSO4.7H2O: 0.05g/L, NH4Cl: 0.05g/L, pH-6) was prepared and inoculated with 24h old pre-inoculum media (20%). Fermentation media was incubated at 30℃ for 72 h at 200 rpm (Arthee and Vijila, 2014; Ojwach et al., 2020). Fermentation media without inoculum was used as a control and all reaction were run in duplicates.
Preparation of extracellular enzyme extract
After incubation fermentation media was filtered through Whatman No.1 filter paper and filtrate broth was centrifuged at 4℃ for 20 min at 8000 rpm, supernatant was separated and served as crude mycelial enzyme extract. Mycelial extract was subjected to different ammonium sulphate concentrations (20%, 30%, 40%. 50%, 60%,70% and 80%), respectively. The precipitate was obtained by centrifugation at 4℃ for 20 min at 13,000 rpm. Pellets were dissolved in 0.2M phosphate buffer saline pH-7.5 and dialyzed further by employing 15KDa cut off dialyzing tubing to remove any ammonium sulphate left behind. After dialysis extracellular FTase was further purified through size exclusion chromatography using Sephadex G-100 (1.5× 30cm) column, equilibrated twice with elution buffer (50mM Tris-Cl, 150mM NaCl and 10% Glycerol, pH: 8, using flow rate of 0.5ml/min. Each fraction was analyzed for FTase activity. Concentration of protein was determined by Bradford reagent. Extracellular FTase was used to perform enzyme assays and biochemical analysis.
Enzyme assay
Enzyme activity of extracellular Ftase is defined as µmol of glucose release during enzymatic reaction per minute under defined standard conditions. Enzyme activity was measured by DNS (Dinitrosalicylic) assay. For enzymatic reaction sucrose was used as a substrate, 20% of sucrose solution was prepared in 0.1M citrate buffer of pH-5. 1ml extracellular enzyme solution and 1ml of substrate solution were mixed together and placed in water bath at 55℃ for 60 min at 120 rpm. At the end of specified incubation time, reaction mixture was placed in boiling water bath for 5-10 min to stop enzymatic reaction. Reaction mixture was cooled at room temperature and centrifuged at 10,000 rpm for 5 min at 4℃. 2ml of DNS reagent was added in reaction mixture and boiled for 10 min. Absorbance of reaction mixture was measured at 540nm along control sample.
Optimal temperature, pH, metal ion and carbon source
Enzyme activity was accessed on different temperature ranges (30℃-90℃) to determine optimal and maximum activity of enzyme. Enzymatic reactions were incubated at different temperatures, pH-5.5 for 60 min, while using substrate concentration 580mM. Enzyme activity was accessed on different pH ranges. Different buffer systems (0.1M Citrate buffers pH 3-6, 0.1M Phosphate buffer pH 7-8 and 0.1M Tris-HCl buffer pH 9-12) were employed to determine optimal enzyme activity, while other reaction conditions (Temperature:55℃ and time 60 min) were same. Before enzymatic reactions, enzyme was first incubated with all three buffer systems for 20 min at room temperature.
Effect of metal ions on Ftase activity was analyzed. 0.5M solutions of following chemicals were prepared: NaCl, MgSO4. 7H2O, HgCl2, FeSO4, ZnSO4, KCl, CaCl2, CuSO4 and AgCl. Before enzymatic reactions, 500µl of enzyme solution was incubated with 500µl of metal ion solution for each reaction and enzyme activity was measured. Substrate solutions of different concentrations (10%: 0.29M, 20%: 0.58M, 30%: 0.88M, 40%: 1.17M, 50%: 1.46M, 60%: 1.75M, 70%: 2.04M and 80%: 2.33M) sucrose substrate solutions of different concentrations were prepared in 0.1M citrate buffer of pH 5.5. An enzyme activity was analyzed for each substrate concentration at temperature 55℃ for 60 min. Effect of carbon source was also analyzed on the production of enzyme by using different carbon source in fermentation media. Fermentation media was prepared in same way mentioned earlier but sucrose is replaced with other potential carbon source maltose, fructose and glucose. Incubation was carried at 30℃ for 72 h at 200 rpm.
HPLC analysis
To confirm formation of FOS, HPLC analysis of enzymatic reaction and of fermentation broth was performed using amino column. Isocratic analysis was carried at room temperature with water and acetonitrile (30:70) as a mobile phase. Fow rate was adjusted to 1ml/min with wavelength 195nm. The yield of FOS was measured as percentage of conversion yield from starting substrate concentration. Analysis was done on basis sequence of retention time of 1-Kestose (GF2), 1-Nystose (GF3) and 1-fructofuranosyl nystose (GF4), respectively.
Synthesis and characterization of MNPs
Magnetic nanoparticles were synthesized by employing co-precipitation method (Samra et al., 2012) , 0.25mM solutions of FeCl2 and FeCl3 were mixed in ratio 1:2 and pH was adjusted to 10 with NH4OH (30%). Resulting black solution was heated at 80℃ with continuous stirring until black precipitates were formed. Precipitates were firstly washed twice with deionized water secondly with acetone and later on dried under vacuum (70℃). 100mg of dried particles were suspended in 50ml deionized water and sonicated for 60 min, particles were filtered through 0.45µm filter. MNPs were characterized through FTIR (Fourier transform infrared), zeta potential, DLS (Dynamic light scattering) and SEM (Scanning electron microscope).
Conjugation of FTase with MNPs
Extracellular FTase was conjugated with MNPs through carbodiimide method. 50mg of MNPs were suspended in 1ml of deionized water and pH:8 was adjusted with NaOH (1M). 20mg of EDC (1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide) mixed in 1ml of deionized water and pH:6.4 was adjusted with HCL(1N). Both solutions were mixed at 120 rpm for 40 min, after 40 min 10mg/ml of EDC was added in solution. 0.2 mg of FTase was added in solution and incubated for 3 h in dark at 120 rpm. Particles were separated by centrifugation at 13,000 rpm for 15 min at 4℃ and washed twice with deionized water. Conjugation was confirmed by FTIR, particles were divided in different fractions and stored at both 4℃ and -20℃ for further analysis. Activity of extracellular FTase enzyme was analyzed in free and immobilized form for the period of six months stored at 4℃ and -20℃.
Results and Discussion
Aspergillus niger SS274 was analyzed for the production of Ftase during submerged fermentation. The crude enzyme extract was obtained after the end of incubation following filtration and centrifugation. Activity of extracellular FTase enzyme was analyzed through DNS method. The crude enzyme extract was subjected to different, from 20% to 80% ammonium sulphate precipitation. 30% of precipitate gave ideal concentration of ammonium sulphate for FTase precipitation showing FTase activity. Dialysis was performed to remove extra ammonium sulphate and gave 0.186mg/ml enzyme hence increasing the purity to 2.28-fold. Specific activity of FTase was increased from 88.69U/mg to 393.32U/mg after size exclusion purification. Detailed comparison of purification of FTase is given in Table I. Purified FTase is of 75kDa, 2 subunits of 45kDa and 30kDa are responsible for activity of FTase obtained after SEC shown in Figure 2.
Table I. Comparison of different purification steps on Specific Activity of extracellular FTase.
|
Steps |
Enzyme activity (µmol/min) |
Enzyme (mg) |
Specific activity U/mg |
Recovery of sample % |
Purification fold |
|
Enzyme extract |
67.76 |
0.764 |
88.69 |
100.00 |
1.00 |
|
Ammonium sulphate |
41.85 |
0.38 |
110.14 |
61.77 |
1.24 |
|
After dialysis |
37.54 |
0.186 |
201.80 |
55.40 |
2.28 |
|
Size exclusion |
38.15 |
0.097 |
393.32 |
91.16 |
3.57 |
Biochemical analysis of extracellular purified enzyme was performed to determine temperature, pH, substrate concentration and metal ion effect on FTase. Effect of temperature was studied to determine optimum temperature for enzyme’s activity, which was subjected to different temperature ranges 30℃ to 90℃ shown in Figure 3A.
FTase gave its maximum activity at 55℃, while enzyme was still showing 70% activity from 40℃ to 60℃. FTase retained its 40% activity at 30℃ and 65℃. Enzyme’s performance was recorded lowest only 5% at 80℃ and no activity was recorded at 90℃ at all. Effect of pH was recorded by subjecting FTase to different pH ranges (3-12) from acidic to basic using different buffer systems shown in Figure 3B. FTase showed its maximum activity at pH 5. FTase retained its 80% activity from pH 6-7 and maintained 50% activity at pH 4 and pH 8. Activity of FTase started to decline from pH 9 and almost no activity was recorded at pH 12. 0.5mM solution of different metal ions Na+, Ca2+, Mg2+, K+, Fe2+, Hg2+, Zn2+, Cu2+ and Ag2+ were prepared and FTase activity was recorded in the presence of these metal ions. Residual activity of FTase in presence of each metal ion is shown in Table II.
Table II. Effect of metal ions on residual activity of FTase.
|
Metal ions |
Concentration |
Residual activity % |
|
Control |
0mM |
100 |
|
Mg2+ |
0.5mM |
83.53 |
|
Na+ |
0.5mM |
90.12 |
|
K+ |
0.5mM |
93.41 |
|
Fe2+ |
0.5mM |
104.94 |
|
Ag2+ |
0.5mM |
15.35 |
|
Ca2+ |
0.5mM |
96.71 |
|
Zn2+ |
0.5mM |
95.39 |
|
Cu2+ |
0.5mM |
81.56 |
|
Hg2+ |
0.5mM |
13.70 |
Activity of FTase without any metal ion was taken as 100% or control. Out of all metal ions ferrous enhanced the enzyme residual activity to 4% when compared with its original activity. Ca2+, Zn2+, Na2+ and K+ had slightest effect on enzyme’s activity, decreasing its activity 5-7% while Mg2+ and Cu2+ caused decrease in FTase activity from 15-20%. Ag2+ and Hg2+ had decreased the enzyme activity up to 85% indicating these metal ions has shown inhibitory effect on activity of enzyme. Out of all Ferrous could be a better choice to enhance the activity of FTase for better production of FOS.
FTase production was also analyzed in the presence of different carbon sources (sucrose, glucose, maltose and fructose). Sucrose and glucose were proved better carbon source for FTase production shown in Figure 4.
FTase activity was calculated with increasing substrate concentration, with increasing substrate concentration activity of the enzyme gradually increase up to 70% of the substrate, after that enzyme activity ceased to increase showing enzyme is fully saturated. Km and Vmax of FTase was calculated by Michaelis Menten Kinetics equation by using enzyme activity and substrate relation. Km and Vmax were recorded 321mM and 64µM/min respectively. FOS production was confirmed through HPLC shown in Figure 5 and Table III.
To increase the stability and shelf life of FTase, it was immobilized on MNPs which were synthesized by co-precipitation method and characterized. Characterization of MNPs is shown in Figure 6, shows formation of amine functionalized MNPs of around 20-60nm in size were formed with circular morphology and stable while suspended in solution. Conjugated MNPs with FTase were characterized by FTIR, stretching peak at 1637.2cm-1 confirmed the formation of FTase immobilization with formation of amide bond. Schematic representation of MNPs synthesis and FTase conjugation is shown in Figure 7.
Table III. HPLC detail of products formed during FTase catalyzed reaction and in fermentation broth.
|
Products |
FTase reaction |
Fermentation broth |
||
|
Peak area |
Area % |
Peak area |
Area % |
|
|
Sucrose |
45074706 |
68.566 |
23311507 |
48.176 |
|
Glucose |
3454595 |
5.255 |
6943798 |
14.350 |
|
Fructose |
3242312 |
4.932 |
5119657 |
10.58 |
|
1-Kestose |
8324399 |
12.663 |
3121757 |
6.452 |
|
1-Nystose |
3076288 |
4.680 |
121177 |
0.250 |
|
1-Fructofuranosyl nystose |
1367211 |
2.080 |
23980 |
0.050 |
Immobilize and free state FTase were stored at two different temperatures (4℃ and -20℃) for period of 6 months to study better storage condition and shelf life of FTase by analyzing enzyme’s activity shown in Figure 8. FTase activity at first month was considered as 100% and compared with respective months accordingly. At 2nd month activity of FTase in free state stored at 4 ℃ decreased to 47.19% and at 3rd month it showed further decline of 12.86% and it continued to decrease to 1.41% in 4th month while no activity was observed at 5th and 6th month, respectively. Free state FTase activity stored at -20 ℃ was recorded 91.93% at 2nd month, 80.60% at 3rd month, 74.13% at 4th month, 54.73% at 5th month while 35.33% at the end of 6th month
FTase conjugated on MNPs were evaluated for its stability, at 2nd month MNP conjugated FTase stored at 4 ℃ showed 89.40% activity, it declined to 51.93% in
third month, 34.01% in 4th month, 14.43% in 5th month and to 4.65% in 6th month. In the same way activity of MNP conjugated FTase stored at -20 ℃ was recorded which showed 95.35% of activity at 2nd month, 89.43% at 3rd month, 81.44% at 4th month, 75.03% at 5th month and 65.42% at 6th month. These results showed FTase showed higher stability and better shelf life in immobilize form and -20 ℃ is optimum storage temperature for FTase to sustain its maximum activity. FTase from different sources and organisms exhibit different activity and catalytic properties which has increased importance of its purification and analysis from different sources for industrial production of prebiotics, this study showed purified FTase enzyme with excellent potential of prebiotic production on industrial scale especially when immobilized on MNPs for enhanced stability as well as activity.
Conclusion
Extracellular FTase was successfully isolated and purified from A. niger SS274 that can be employed for prebiotic production and its was successfully immobilized to MNPs in order to increase its stability as well as shelf life making it more suitable for industrial applications.
Declarations
Acknowledgments/ funding
This work was supported by School of Biochemistry and Biotechnology, University of the Punjab Lahore.
IRB approval
The punjab University Ethics Review Board reviewed and approved the study (No. D/171/FIMS, Sated: 12/10/2023).
Generative AI and AI-assisted technology statement
The authors have declared that no generative AI or AI-assisted technologies were used to create this manuscript.
Statement of conflict of interest
The authors have declared no conflict of interest.
References
Antošová, M. and Polakovič, M., 2001. Fructosyltransferases: The enzymes catalyzing production of fructooligosaccharides. Chem. Pap., 55: 350–358.
Arthee, R. and Vijila, K., 2014. Study on Fructosyltransferase enzyme from Aspergillus sp. in Fructooligosaccharides production. Res. J. Recent Sci., 3: 147–153.
Borromei, C., Cavazza, A., Merusi, C. and Corradini, C., 2009. Characterization and quantitation of short-chain fructooligosaccharides and inulooligosaccharides in fermented milks by high-performance anion-exchange chromatography with pulsed amperometric detection. J. Separat. Sci., 32: 3635–3642. https://doi.org/10.1002/jssc.200900322
Bruno-Barcena, J.M. and Azcarate-Peril, M.A., 2015. Galacto-oligosaccharides and colorectal cancer: Feeding our intestinal probiome. J. Funct. Fds., 12: 92–108. Elsevier Ltd. https://doi.org/10.1016/j.jff.2014.10.029
Campbell, J.M., Bauer, L.L., Fahey, G.C., Hogarth, A.J.C.L., Wolf, B.W. and Hunter, D.E., 1997. Selected fructooligosaccharide (1-Kestose, Nystose, and 1F-β-Fructofuranosylnystose) composition of foods and feeds. J. Agric. Fd. Chem., 45: 3076–3082. https://doi.org/10.1021/jf970087g
Crittenden, R.G. and Playne, M.J., 1996. Production, properties and applications of food-grade oligosaccharides. Trends Fd. Sci. Technol., 7: 353–361. https://doi.org/10.1016/S0924-2244(96)10038-8
Cunha, J.S., Ottoni, C.A., Morales, S.A.V., Silva, E.S., Maiorano, A.E. and Perna, R.F., 2019. Synthesis and characterization of fructosyltransferase from Aspergillus oryzae ipt-301 for high fructooligosaccharides production. Braz. J. Chem. Eng., 36: 657–668. https://doi.org/10.1590/0104-6632.20190362s20180572
Dake, M.S. and Kumar, G., 2012. Partial purification and characterization of fructosyltransferase from aureobasidium. Int. J. Sci. Environ., 1: 88–98.
Dhake, A.B. and Patil, M.B., 2007. Effect of substrate feeding on production of fructosyltransferase by Penicillium purpurogenum. Braz. J. Microbiol., 38: 194–199. https://doi.org/10.1590/S1517-83822007000200002
Dou, Y., Yu, X., Luo, Y., Chen, B., Ma, D. and Zhu, J., 2022. Effect of fructooligosaccharides supplementation on the gut microbiota in human: A systematic review and meta-analysis. Nutrients, 14: MDPI. https://doi.org/10.3390/nu14163298
Fujishima, M., Sakai, H., Ueno, K., Takahashi, N., Onodera, S., Benkeblia, N. and Shiomi, N., 2005. Purification and characterization of a fructosyltransferase from onion bulbs and its key role in the synthesis of fructo-oligosaccharides in vivo. New Phytol., 165: 513–524. https://doi.org/10.1111/j.1469-8137.2004.01231.x
Grajek, W., Olejnik, A. and Sip, A., 2005. Probiotics, prebiotics and antioxidants as functional foods. Acta Biochim. Polon., 52: 665–671. https://doi.org/10.18388/abp.2005_3428
Heyer, A.G. and Wendenburg, R., 2001. Gene cloning and functional characterization by heterologous expression of the fructosyltransferase of Aspergillus sydowi IAM 2544. Appl. environ. Microbiol., 67: 363–370. https://doi.org/10.1128/AEM.67.1.363-370.2001
Itaya, N.M., Figueiredo-Ribeiro, R.C.L. and Buckeridge, M.S., 1999. Synthesis of fructans by fructosyltransferase from the tuberous roots of Viguiera discolor (Asteraceae). Braz. J. med. biol. Res., 32: 435–442. https://doi.org/10.1590/S0100-879X1999000400010
Khanvilkar, S.S. and Arya, S.S., 2015. Fructooligosaccharides: Applications and health benefits: A review. Agro Fd. Ind. Hi-Tech, 26: 8–12.
Kingston, S.A.H., Walker, R.P. and Pollock, C.J., 1999. Invertase in leaves: Conundrum or control point? J. exp. Bot., 50: 735–743. https://doi.org/10.1093/jxb/50.335.735
Koops, A.J. and Jonker, H.H., 1996. Purification and characterization of the enzymes of fructan biosynthesis in tubers of Helianthus tuberosus Colombia: II. Purification of sucrose: Sucrose 1-fructosyltransferase and reconstitution of fructan synthesis in vitro with purified sucrose: Sucrose. Pl. Physiol., 110: 1167–1175. https://doi.org/10.1104/pp.110.4.1167
Lasseur, B., Schroeven, L., Lammens, W., Le Roy, K., Spangenberg, G., Manduzio, H., Vergauwen, R., Lothier, J., Prud’homme, M.P. and Van Den Ende, W., 2009. Transforming a fructan: Fructan 6G-fructosyltransferase from perennial ryegrass into a sucrose: Sucrose 1-fructosyltransferase. Pl. Physiol., 149: 327–339. https://doi.org/10.1104/pp.108.125559
Luscher, M., Frehner, M. and Nosberger, J., 1993. Purification and some properties of fructan: Fructan fructosyl transferase from dandelion (Taraxacum officinale Weber). New Phytol., 123: 437–442. https://doi.org/10.1111/j.1469-8137.1993.tb03755.x
Markowiak, P. and Ślizewska, K., 2017. Effects of probiotics, prebiotics, and synbiotics on human health. Nutrients, 9: https://doi.org/10.3390/nu9091021
Michel, M.R., Rodríguez-Jasso, R.M., Aguilar, C.N., Gonzalez-Herrera, S.M., Flores-Gallegos, A.C. and Rodríguez-Herrera, R., 2016. Fructosyltransferase sources, production, and applications for prebiotics production. In: Probiotics and prebiotics in human nutrition and health. https://doi.org/10.5772/62827
Mussatto, S.I. and Mancilha, I.M., 2007. Non-digestible oligosaccharides: A review. Carbohyd. Polym., 68: 587–597. https://doi.org/10.1016/j.carbpol.2006.12.011
Ojwach, J., Kumar, A., Mukaratirwa, S. and Mutanda, T., 2020. Purification and biochemical characterization of an extracellular fructosyltransferase enzyme from Aspergillus niger sp. XOBP48: Implication in fructooligosaccharide production. 3 Biotech, 10: 459. https://doi.org/10.1007/s13205-020-02440-w
Ozcan, T. and Kurtuldu, O., 2014. Influence of dietary fiber addition on the properties of probiotic yogurt. Int. J. Chem. Eng. Appl., 5: 397–401. https://doi.org/10.7763/IJCEA.2014.V5.417
Pool-Zobel, B., van Loo, J., Rowland, I. and Roberfroid, M.B., 2002. Experimental evidences on the potential of prebiotic fructans to reduce the risk of colon cancer. Br. J. Nutr., 87(S2): S273–S281. https://doi.org/10.1079/BJN/2002548
Rao, V. A., 2001. The prebiotic properties of oligofructose at low intake levels. Nutr. Res., 21: 843–848. https://doi.org/10.1016/S0271-5317(01)00284-6
Roberfroid, M.B., 2007. Inulin-type fructans: Functional. J. Nutr., 137: 2493–2502. https://doi.org/10.1093/jn/137.11.2493S
Sabater-Molina, M., Larque, E., Torrella, F. and Zamora, S., 2009. Dietary fructooligosaccharides. Ovid Med. J. Physiol. Biochem., 65: 315–328. https://doi.org/10.1007/BF03180584
Samra, Z.Q., Dar, N. and Athar, M.A., 2012. Biocatalytic activity of recombinant human β-mannosidase immobilized onto magnetic nanoparticles for bioprocess. Preparat. Biochem. Biotechnol., 42: 97–112. https://doi.org/10.1080/10826068.2011.575910
Sangeetha, P.T., 2003. Microbial production of fructooligosaccharides. Asian J. Microbiol. Biotechnol. environ. Sci., 5.
Sangeetha, P.T., Ramesh, M.N. and Prapulla, S.G., 2004. Production of fructosyl transferase by Aspergillus oryzae CFR 202 in solid-state fermentation using agricultural by-products. Appl. Microbiol. Biotechnol., 65: 530–537. https://doi.org/10.1007/s00253-004-1618-2
Sridevi, V., Sumathi, V., Guru, P.M. and Satish, K.M., 2014. Fructooligosaccharides type prebiotic: A review. J. Pharma. Res., 8: 321–330.
Trujillo, T.L.E., Martinez, G.D., Pérez, C.E., Rivera, I.L.M., Pérez, J.N. and Pais, C.J.M., 2019. Fructosyltransferases and invertases: Useful enzymes in the food and feed industries. In: Enzymes in food biotechnology, Elsevier Inc. pp. 451–469. https://doi.org/10.1016/B978-0-12-813280-7.00026-8
Van Hijum, S.A.F.T., Van Geel-Schutten, G.H., Rahaoui, H., Van der Maarel, M.J.E.C. and Dijkhuizen, L., 2002. Characterization of a novel fructosyltransferase from Lactobacillus reuteri that synthesizes high-molecular-weight inulin and inulin oligosaccharides. Appl. environ. Microbiol., 68: 4390–4398. https://doi.org/10.1128/AEM.68.9.4390-4398.2002
Yang, H., Wang, Y., Zhang, L. and Shen, W., 2016. Heterologous expression and enzymatic characterization of fructosyltransferase from Aspergillus niger in Pichia pastoris. New Biotechnol., 33: 164–170. https://doi.org/10.1016/j.nbt.2015.04.005
Yun, J.W., 1996. Fructooligosaccharides occurrence, preparation, and application. Enzyme Microbial Technol., 19: 107–117. https://doi.org/10.1016/0141-0229(95)00188-3