Enhanced fructooligosaccharides synthesis by engineered Trichoderma atroviride β-fructofuranosidase
Egle Narmontaite, Francisco J. Plou, María Fernández-Lobato
TL;DR
Researchers discovered a new enzyme from Trichoderma atroviride that efficiently produces fructooligosaccharides, which could be useful for prebiotic applications.
Contribution
The study introduces a novel β-fructofuranosidase from Trichoderma atroviride with high transfructosylation activity for FOS synthesis.
Findings
TaINV synthesized 252 g/L of total FOS, with 1-kestose as the major product.
Engineered variants achieved FOS yields of up to 62.7% of total sugars.
Structure-function analysis identified key residues for transfructosylation specificity.
Abstract
Here we report the first β-fructofuranosidase from the Trichoderma genus producing fructooligosaccharides (FOS). The novel enzyme from Trichoderma atroviride (TaINV) here characterized was heterologously expressed, purified, and biochemically analyzed. TaINV exhibited hydrolytic activity mainly toward sucrose and other substrates containing β-(2 → 1) linkages, with minor activity toward β-(2 → 6) bonds. In addition to hydrolysis, it catalyzed the synthesis of FOS of all three structural series (1F-FOS, 6F-FOS, and 6G-FOS). At the maximal production point, TaINV synthesized 252 g/L of total FOS, representing 50.3% (w/w) of the total sugars in the reaction mixture, with 1-kestose as the major product, representing ~ 85% of the total products synthesized. Structural analysis based on AlphaFold-predicted TaINV model and comparative superimposition with GH32-substrate complexes revealed…
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Figure 8- —Spanish Ministry of Science and Innovation
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Taxonomy
TopicsMicrobial Metabolites in Food Biotechnology · Diet, Metabolism, and Disease · Polysaccharides and Plant Cell Walls
Introduction
Nutraceuticals and foods are gaining importance due to increasing awareness of their health benefits (Peruzzolo et al. 2025). The global functional food and beverage market was valued at USD 364.2 billion in 2024 and is projected to reach USD 793.6 billion by 2032, with an annual growth rate (CAGR) of 10.3% (Fortune Business Insights 2024). Among bioactive components, oligosaccharides are increasingly valued as prebiotics in functional foods, often preferred over fibers, peptides, or fatty acids. Among them, fructooligosaccharides (FOS), galactooligosaccharides, and isomaltooligosaccharides are the most common ones (Narisetty et al. 2022; Vera et al., 2021). FOS exhibit numerous health benefits for humans, including the growth stimulation of beneficial bacteria in the gut and inhibition of the pathogenic ones, also known as a prebiotic effect (Bhadra et al. 2022; Rawat et al. 2024). They also improve the intestinal absorption of minerals and trace elements (Costa et al. 2021), decrease serum levels of cholesterol and triglycerides, show low carcinogenicity (Bhadra et al. 2022), reduce the risk of colon cancer (Alvandi et al. 2022; Bytautaitė et al. 2024), improve mental health (Taylor and Holscher 2020; Zhang et al. 2020), and even decrease neuroinflammation and promote neuroplasticity (Rodrigues de Paiva et al. 2023).
FOS consist of fructose oligomers with a glucose unit attached to the terminal fructose unit through a β-(2 → 1)-glycosidic bond. They are categorized into three primary series based on the linkage pattern: levan- or inulin-type, where fructose units are connected by β-(2 → 6) linkages (^6^F-FOS, as in the trisaccharide 6-kestose) or β-(2 → 1) bonds (^1^F-FOS, as in the trisaccharide 1-kestose), and neoseries fructan-type, where a β-(2 → 6) linkage joins a fructose unit to the terminal glucose unit (^6^G-FOS, as in the trisaccharide neokestose) (Jiménez-Ortega et al. 2022). Although FOS are naturally found in numerous fruits and vegetables, and in more than 36,000 plant species, they are produced only in small amounts, and their production levels may vary depending on the season and abiotic factors (Nobre et al. 2015, 2022). β-Fructofuranosidases (EC 3.2.1.26) are predominant biocatalysts to produce FOS. They belong to the glycosyl hydrolase family 32 (GH32) (CAZy; http://www.cazy.org), which also includes inulinases and fructosyltransferases, and together with the GH68 family (bacterial fructosyltransferases, levansucrases, and inulosucrases) form the GH-J clan. They share a common fivefold β-propeller catalytic domain, and GH32 enzymes have an additional C-terminal β-sandwich domain attached to their N-terminal catalytic domain (Jiménez-Ortega and Sanz-Aparicio 2024). Three key amino acid residues located in the active site, responsible for substrate binding and hydrolysis, are surrounded by conserved sequences among members of the GH32 family: WMNDPNG (D acting as a nucleophile), FRDP (D acting as a stabilizer of the transition state), and ECP (E acting as an acid–base catalyst) (Jiménez-Ortega et al. 2022).
Currently, fructosyltransferases and β-fructofuranosidases from fungi, such as Aspergillus sp. and Aureobasidium sp., are among the most widely used producers of FOS at the industrial scale, achieving production yields representing about 50–60% (w/w) of total products in the reaction mixtures (Nobre et al. 2022; Rawat et al. 2024). The commercialized enzymes used in FOS synthesis mostly synthesize inulin-type FOS with a degree of polymerization (DP) of about 3 units (mainly 1-kestose), and toward the end of the reaction, when the highest yield of FOS is reached, higher-DP FOS are also formed which have lower prebiotic potential (Karkeszová and Polakovič, 2023). Although several β-fructofuranosidases have been isolated from bacteria, plants, and fungi, and their ability to produce FOS has been studied (Jiménez-Ortega et al. 2022; Nadeem et al. 2015; Omori et al. 2010), to the best of our knowledge, no such enzymes have been reported in Trichoderma species. This fungal genus, within the family Hypocreaceae, is commonly known for its ubiquity in all soil types and for its high prevalence among culturable soil fungi. Many species in this genus are opportunistic avirulent plant symbionts, but others can even act as biocontrol agents against a wide range of economically important aerial and soilborne plant pathogens (Silva et al. 2014). The aim of this study was to identify and functionally characterize the first GH32 β-fructofuranosidase from the Trichoderma genus showing transfructosylation activity and analyze its biotechnological potential. In this work, a sucrose-hydrolyzing activity was detected in Trichoderma atroviride, and a gene encoding a putative GH32 β-fructofuranosidase was identified. The protein that it encoded (TaINV) was heterologously expressed, purified, biochemically characterized, and its potential to produce FOS analyzed. Comparison of the AlphaFold predicted TaINV model with GH32-substrate complex structures revealed conserved catalytic elements and positions associated with substrate recognition. Several TaINV variants were generated and evaluated for FOS production, with some achieving yields comparable to those reported for commercial β-fructofuranosidases from Aspergillus niger and Aureobasidium pullulans.
Materials and methods
Strains, plasmids, media, and growth conditions
Trichoderma atroviride IMI 206040 was used in this study for genomic DNA isolation. It was grown on potato dextrose agar (PDA 39 g/L) plates (Condalab, Madrid) at 25 °C for 7 days. Spores were harvested from PDA plates using 3 mL of sterile water and inoculated in 50 mL of Czapek liquid medium (30 g/L sucrose, 2 g/L sodium nitrate, 1 g/L dipotassium phosphate, 0.5 g/L magnesium sulfate, 0.5 g/L potassium chloride, 0.01 g/L ferrous sulfate) in a 250-mL flask and grown for 2–3 days. Mycelium was collected by filtration using a 0.22-μm filter from Merck Millipore Ltd (Tullagreen, Carrigtwohill, Co. Cork IRL) and approximately 300 mg of mycelium was transferred to a 1.5-mL tube and stored at − 70 °C.
Escherichia coli strains DH5α and BL21 (DE3) (Invitrogen, Carlsbad, CA, USA) were used in this study for the cloning and expression systems, respectively. The recombinant bacteria were maintained in Luria–Bertani (LB) plates (10 g/L tryptone, 5 g/L yeast extract, 10 g/L sodium chloride, 20 g/L agar) with kanamycin (30 μg/mL). The cellular growth was monitored spectrophotometrically at a wavelength of 600 nm.
YEP (S-BTB) plates (10 g/L yeast extract, 20 g/L peptone, 50 g/L sucrose, 20 g/L agar, 0.5 g/L BTB: bromothymol blue) were used for the sucrose hydrolytic activity assay as previously referred (Garcia-Gonzalez et al., 2019). BTB was used as a pH indicator: yellow, green, and blue in acidic, neutral, and basic solutions, respectively. In this assay, Komagataella phaffii GS115 (also known as Pichia pastoris GS115; Invitrogen, Carlsbad, CA, USA) and Rhodotorula dairenensis CECT1416 (also Rhodotorula glutinis var. dairenensis) were used as negative and positive controls for sucrose hydrolysis, respectively.
DNA amplification, cloning, and site-directed mutagenesis
Genomic* T*. atroviride IMI 206040 sequences were already available in databases (NCBI code: NW_014013638.1). To amplify the gene responsible for the invertase activity from this organism, genomic DNA was obtained as previously described (Vazquez-Angulo et al. 2012) and used as template in PCR reactions. The mRNA sequence codifying for a potential GH32 protein (uncharacterized protein TrAtP1_009821, NCBI: XM_014091134.1) was used for the primers TAINVF and TAINVR design (Table S1). In Vivo Assembly (IVA) cloning was used for the insertion of the amplified sequence (here named TaINV) into the expression vector pET28b(+) (Novagen, Darmstadt, Germany) as previously referred (García-Nafría et al. 2016). Shortly, in the first PCR: vector pET28b(+) was linearized by PCR using primers pET28b(+) openF and pET28b(+) openR. In the second PCR: the potential gene TaINV (predicted length of 1947 bp, including the stop codon) was amplified using the primers TAINVF and TAINVR that contained pET28B(+) recombination sequences. The two PCR products were mixed in a ratio 1:5 (vector:insert) and used to transform E. coli DH5α, where their assembly occurred. Conditions for the first PCR were as follows: 98 °C for 30 s; 35 cycles of 98 °C for 10 s, 61 °C for 30 s, 72 °C for 300 s; and a final elongation at 72 °C for 120 s. Conditions for the second PCR were that of the first but with 55 °C as annealing temperature and the final elongation reduced to 60 s. The TaINV sequence included in pET28B(+) was verified by sequencing (Macrogen, Madrid, Spain) and consisted of 1996 bp including one intron of 52 bp that was successfully removed applying transfer PCR using the primers IntF and IntR (Table S1) as previously described (Erijman et al. 2011). The PCR conditions were as follows: 98 °C for 30 s; 13 cycles of 98 °C for 10 s, 58 °C for 30 s, 72 °C for 15 s; 25 cycles of 98 °C for 10 s, 72 °C for 330 s; 72 °C for 120 s. The final construct pET28b(+)-TAINV contained the potential gene TaINV (1944 bp) flanked by the T7 lac promoter and sequences codifying a C-terminus 6-His-tag followed by the stop codon TGA. This plasmid was used as a template to generate all the protein TaINV variants produced in this work by site-directed mutagenesis (Liu and Naismith 2008) using the specific primers listed in Table S1. All mutations were verified by DNA sequencing (Macrogen, Madrid, Spain). In all PCR amplifications, Phusion High-Fidelity or Q5 High-Fidelity DNA Polymerases (both from NEB, Ipswich, UK) were used according to the manufacturer’s recommendations. Functionality of TaINV sequence was validated after analyzing the activity of the protein it encoded using E. coli BL21 (DE3) as heterologous expression system.
Protein expression and purification
E. coli BL21 (DE3) cells were transformed by electroporation using standard techniques, as previously described (Garcia-Gonzalez et al. 2020). Transformants carrying pET28b(+) empty vector were also obtained and used as controls. For protein expression, bacteria transformants were cultivated in 10 mL of LB media containing kanamycin (30 μg/mL) at 30 °C in an orbital shaker (200 rpm) for 16 h. Then the culture was used to inoculate 400 mL of LB media with kanamycin and cultivated until it reached 0.6–0.8 units at 600 nm. Protein expression was induced by adding 1 mM isopropyl-β-D-1-thiogalactopyranoside (IPTG) and the culture was incubated for 14–16 h at 18 °C. Cells were collected by centrifugation (5000 × g, 10 min) and resuspended in 35 mL of wash buffer (20 mM HEPES pH 8.0, 200 mM sodium chloride, 30 mM imidazole). The pH of HEPES buffers was adjusted using NaOH. Cell suspensions were disrupted by sonication using an MSE 150-Watt Ultrasonic Disintegrator Mk2 (MSE Scientific Instruments, Sussex, England), for 2 min (15-s on, 15-s off) followed by centrifugation (8000 × g, 20 min). The supernatant was filtered through 0.45-μm syringe PVDF filters (Merck Millipore, Burlington, MA, USA). The His-tagged proteins were purified by affinity chromatography using Ni–NTA agarose packed (Qiagen, Hilden, Germany) equilibrated with the same wash buffer at pH 8.0. Columns were washed with the same buffer and eluted with elution buffer (20 mM HEPES pH 8.0, 250 mM imidazole). The eluted fractions containing purified protein were dialyzed against 20 mM HEPES pH 7.0 at 4 °C for 16 h. If needed, the protein was further concentrated using 30,000 MWCO PES Amicon™ Ultra-0.5 centrifugal filters (Merck Millipore, Darmstadt, Germany). All protein variants generated in this work were produced in E. coli BL21 (DE3) and then purified as indicated above.
SDS-PAGE and zymogram analysis
For the protein purification analysis, sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE; 12%) was used. Gels were stained with BlueSafe (NzyTech, Lisbon, Portugal). Precision Plus Protein Standards Unstained 10–250 kDa (Bio-Rad, Hercules, CA, USA) were used as molecular weight markers. The concentration of the proteins was determined using a NanoDrop™. One/OneC Microvolume UV–Vis Spectrophotometer at 280 nm (ThermoFisher, Waltham, Massachusetts, USA), applying the extinction coefficient of 1 mg/mL = 1.725 calculated by the ExPASy ProtParam server (https://web.expasy.org/protparam/).
For determination of the sucrose-hydrolyzing activity, zymogram analysis was performed as previously described (Linde et al. 2012). Briefly, proteins were loaded onto 12% non-denaturing PAGE gels (without SDS) run for 2 h at 120 V and 4 °C. Then gels were washed twice with 50 mM sodium acetate pH 5.5 containing 1% Triton X-100 (v/v), and then incubated with 1 M sucrose, in the same washing buffer, for 30 min at 45 °C. After washing three times with distilled water, gels were stained with 1.0% 2,3,5-triphenyltetrazolium chloride in 0.25 M NaOH (previously heated), to reveal the reducing sugars. The reaction was stopped by adding 5.0% (v/v) acetic acid. Alternatively, gels were stained with colloidal Coomassie. Bovine Serum Albumin (BSA; 66 kDa) was used as a molecular weight control, which under non-denaturing conditions forms oligomers, and the invertase from Saccharomyces cerevisiae (Novozymes, Bagsvaerd, Denmark) as positive activity control.
Bioinformatic analysis and molecular modeling
The sequence of the protein TaINV from T. atroviride was obtained from the National Center for Biotechnology Information (NCBI; GeneBank accession number: XP_013946609) and was analyzed by using UniProt BLAST (https://www.uniprot.org/blast). Sequences were aligned using Clustal Omega (https://www.ebi.ac.uk/Tools/msa/clustalo/) (pairwise gap open penalty 10, gap extension penalty 0.5, multiple alignment gap open penalty 10, gap extension penalty 0.05), analyzed in SnapGene Viewer application and represented using ESPript 3.0 (https://espript.ibcp.fr/ESPript/ESPript/). The theoretical molecular weight was calculated on ExPASy ProtParam server (https://web.expasy.org/protparam/). The structural prediction of TaINV was performed using AlphaFold3 (Abramson et al. 2024), and experimentally solved GH32 structures used in this work were obtained from the Protein Data Bank (PDB) (https://www.rcsb.org/). 3D protein models were visualized, analyzed, and superimposed using UCSF ChimeraX version 1.9. All figures were prepared and visualized using GraphPad Prism version 8.0.2 (GraphPad Software, San Diego, CA, USA).
Enzyme characterization and kinetic analysis
The sucrose-hydrolyzing (also β-fructofuranosidase hydrolytic) activity was determined using the 3,5-dinitrosalicylic acid (DNS) assay adapted to a 96-well microplate as previously described (Gimeno-Pérez et al. 2021; Jiménez-Ortega et al. 2022). Glucose 0.0–3.0 g/L was used for the calibration curve. Reactions (50 μL) contained 5 μL of the enzymatic solutions (adequately diluted to fit the calibration curve) and 45 μL of 10 g/L sucrose, in 50 mM sodium acetate pH 5.5, were incubated at 45 °C for 10–20 min. One unit (U) of sucrose hydrolytic activity was defined as that catalyzing the formation of 1 μmol of reducing sugars per min in the mentioned conditions. The estimation of hydrolase activity under different pH (3.0–8.5) and temperature (30–65 °C) values was carried out under the mentioned conditions, using sucrose as substrate in sodium acetate (pH 3.0–5.5) or sodium phosphate (pH 5.5–8.5), all at 50 mM. For the pH stability studies, the enzyme TaINV was pre-incubated in ice for 1 h at different pH (4.0–8.0) values and for thermostability at different temperatures (30–65 °C) values for 15, 30, 45, 60, and 90 min. Inulin (BENEO, Barcelona, Spain), sucrose (Merck, MA, USA), raffinose, nystose, levan of bacterial origin (Sigma-Aldrich, MA, USA), and fructan-agave from Agave tequilana supplied by Dr. G. Sandoval (CIATEJ, Guadalajara, México); all at 1% (w/v) except levan at 0.1% (w/v; the highest concentration at which it was completely dissolved) were used as possible substrates, as referred above. All reactions were made in triplicate.
The kinetic analysis was performed in triplicate using sucrose (1–450 mM), raffinose (5–430 mM), inulin (1–205 mM), and nystose (2–1000 mM). Reactions were carried out for 10 min at 45 °C in 50 mM sodium acetate pH 5.5, for all enzyme variants. For inulin, an average molecular weight of 1640 g/mol was used for calculations. The plotting and analysis were carried out using the kinetic module in SigmaPlot (version 14.5). Kinetic parameters were calculated by fitting the initial rate values to the Michaelis–Menten equation.
Transferase activity, fructooligosaccharides production, and HPAEC-PAD analysis
The transferase activity was measured using 500 g/L sucrose in 50 mM sodium acetate pH 5.5, 5–10 U/mL of sucrose hydrolytic activity, and 45 °C in an orbital shaker (Vortem 56, Labnet International, Woodbridge, NJ, USA), as previously described (Jiménez-Ortega et al. 2022). Samples of 50 μL were withdrawn from the reaction mixture at different times (0–24 h), the enzyme was inactivated at 100 °C for 8 min, and samples stored at − 20 °C until further use. The samples were diluted 500–1000 times with H_2_O and filtered through 0.45-μm pore filter (Scharlau, S.L; Sentmenat, Spain). They were analyzed by high-performance anion-exchange chromatography with pulsed amperometry detection (HPAEC-PAD) on a Dionex ICS3000 system (Sunnyvale, CA, USA) and a CarboPack PA-1 column (4 × 250 mm) connected to a PA-1 guard column as previously referred (Míguez et al. 2018). Separation started with 100 mM NaOH at a flow rate of 1 mL/min for 8 min. Subsequently, a linear gradient was applied over 22 min, during which the proportion of 100 mM NaOH was decreased from 100 to 88%, while the proportion of 100 mM NaOH containing 600 mM sodium acetate was increased from 0 to 12% (corresponding to a final sodium acetate concentration of 72 mM). This eluent composition was maintained for 6 min, then changed to 50% 100 mM NaOH and 50% 100 mM NaOH/600 mM sodium acetate and maintained until the end of the chromatographic run. Eluents were degassed by flushing with helium and peaks were analyzed using Chromeleon software. Compounds were identified by comparison of their retention times with those of the standards and quantified based on peak areas using calibration curves. The standards employed were fructose, glucose, and sucrose (Merck, MA, USA), 1-kestose and nystose (TCI Europe, Zwijndrecht, Antwerp, Belgium), and fructosylnystose (Megazyme); blastose, neokestose, and 6-kestose were produced by the β-fructofuranosidases from R. dairenensis (and its protein variants) and Xanthophyllomyces dendrorhous and purified as previously described (Jiménez-Ortega et al. 2022; Linde et al. 2012; Zambelli et al. 2014). Peaks that could not be structurally identified were not included in quantitative calculations.
Results
Cloning and the heterologous expression of the GH32 protein from Trichoderma atroviride
Initially, the potential sucrose-hydrolyzing activity of Trichoderma atroviride IMI 206040 was evaluated using a rich solid medium containing sucrose and bromothymol blue as pH indicator (YEP (S-BTB)). The yeasts Rhodotorula dairenensis and Komagataella phaffii were used as positive and negative controls in this assay because of their proven or absent capacity to hydrolyze sucrose, respectively. As expected, all microbial species were able to grow in this yeast medium. R. dairenensis turned the plates color from green to yellow due to the acidic compounds formed after the sucrose utilization, whereas K. phaffii turned it into dark blue following the peptone utilization and the ammonium excretion. As a result of its sucrose-hydrolyzing activity, the medium in which T. atroviride grew also turned yellow (Fig. 1A).Fig. 1. Plate assay of sucrose-hydrolyzing activity and analysis of the protein TaINV expressed in E. coli. A Solid-rich medium including sucrose and BTB (YEP (S-BTB)) was used in sucrose-hydrolyzing activity assays. Non-inoculated (1; green) and inoculated plates with K. phaffii, as negative control (2; dark blue), T. atroviride showing sucrose hydrolase activity (3; dark yellow), and* R. dairenensis* as positive control (4; light yellow) are shown. B SDS-PAGE analysis of the protein TaINV expressed in E. coli and purified with Ni–NTA agarose packed columns (lane 1). C Zymogram analysis of the purified TaINV (lane 1). Commercial S. cerevisiae invertase was used as positive activity control (lane 2) and BSA as a molecular mass control. Numbers on the left and right of panels B and C, respectively, indicate the positions of molecular mass standards (lane M) in kDa
The nucleotide sequence located in the T. atroviride genome potentially encoding a GH32 protein resulted in a sequence of 648 amino acids (TaINV), which showed high similarity to other already annotated GH32 sequences. Thus, TaINV displayed the highest homology to the potential GH32 proteins from Trichoderma gamsii (PNP48467.1, query cover 99%, identity 92.3%) and Trichoderma harzianum (KKP03051.1, query cover 99%, identity 82.9%), none of which have been previously studied, nor their functionality demonstrated (Fig. S1).
The recombinant protein TaINV, including a C-terminal 6-His-tag, was expressed in E. coli cultures, and then purified by affinity chromatography. A major band of about 75 kDa was visualized by SDS-PAGE (Fig. 1B), which was consistent with the theoretical weight of TaINV (~ 74 kDa). The sucrose-hydrolyzing activity of the purified protein was confirmed by zymogram analysis (Fig. 1C), thus validating the TaINV sequence functionality. In the zymogram, TaINV migrated even above the 264 kDa BSA marker, suggesting that the protein does not behave as a monomer under these conditions (Fig. 1C).
Biochemical characterization and kinetic analysis of TaINV
The recombinant TaINV displayed the maximum sucrose-hydrolyzing activity at 40–45 °C and pH 5.5–6.5 (Fig. 2A, B). Regarding pH stability and thermostability, TaINV maintained more than 80% of its initial activity after 1 h in the pH range of 5.0–8.0, but only 50% (T_50_) after 30 min at 45 °C and was completely inactivated after 90 min at 65 °C (Fig. 2C, D).Fig. 2. Optimal temperature and pH, thermostability, and pH stability of the enzyme. The effect of temperature (A) and pH (B) on hydrolytic activity was evaluated using sucrose as substrate. For the optimal activity assays, pH 5.5 and 45 °C were used, with sodium acetate (blue) or sodium phosphate (red) buffers, respectively. In the thermostability study (C), the enzyme was incubated at 45 °C (blue circles), 50 °C (red squares), 55 °C (green triangles), 60 °C (purple inverted triangles), and 65 °C (orange diamonds), and residual activity was measured over time. For pH stability (D), the enzyme was incubated for 1 h at the indicated pH values, before activity determination
The hydrolytic activity of TaINV was evaluated using different substrates containing fructosyl β-(2 → 1), β-(2 → 6), α-(1 → 3), α-(1 → 5) or α-(1 → 6) linkages (Table 1). The enzyme liberated reducing sugars only from fructosyl β-(2 → 1) linked carbohydrates, such as sucrose, nystose, and to a lower extent, inulin, raffinose, fructan-agave, and levan. No activity was detected on sucrose isomers such as leucrose (α-D-Glc-(1 → 5)-D-Fru), palatinose (α-D-Glc-(1 → 6)-D-Fru), and turanose (α-D-Glc-(1 → 3)-D-Fru), indicating a clear preference for substrates including β-(2 → 1) linkages and to a lesser extent β-(2 → 6) as in levan or fructan-agave. The highest specific activity was obtained with sucrose (about 416 μmol/(min mg)), which decreased with increasing substrate polymerization degree, being reduced by about 10- and 30-fold with nystose and inulin, respectively. Table 1. TaINV-specific activity on the referred substratesSubstrate (main linkage-types)Specific activity (μmol/(min mg))Sucrose (β-(2 → 1))416.4 ± 0.6Nystose (β-(2 → 1))40.2 ± 1Inulin (β-(2 → 1))14.5 ± 0.1Raffinose (α-(1 → 6)/β-(2 → 1))13.8 ± 0.3Fructan-agave (β-(2 → 1)/β-(2 → 6))4.1 ± 0.1Levan (β-(2 → 6))0.8 ± 0.03Leucrose (α-(1 → 5))n.dPalatinose (α-(1 → 6))n.dTuranose (α-(1 → 3))n.dData are average of three independent experiments and standard errors (±) are indicated. n.d., not detected. Main linkage-types connecting the sugar units of each of the substrates are indicated
Enzyme kinetics for TaINV were examined with sucrose, nystose, inulin, and raffinose (Table 2, Fig. S2). As expected, TaINV showed the highest affinity for sucrose (Km of 8.7 mM), with nearly ninefold higher Km values for nystose. The enzyme catalytic efficiency, defined by the kcat/Km ratio, with sucrose was about 14–42-fold higher than with the other substrates tested. Table 2. TaINV kinetic parameters with different substratesSubstrateKm (mM)kcat (s^−1^)kcat/Km (s^−1^ mM^−1^)Sucrose8.7 ± 0.8479 ± 355 ± 4Raffinose51.2 ± 8195 ± 9.33.8 ± 0.2Inulin12.3 ± 1.435.4 ± 1.52.9 ± 0.2Nystose77.9 ± 8.4100 ± 31.3 ± 0.01Values represent an average of three experiments. The kcat values were calculated assuming a protein molecular mass of 74 kDa. The standard errors were calculated based on the curve fitting Michaelis–Menten model using SigmaPlot
Amino acid sequence, structural analysis, and catalytic residues identification
TaINV consists of 648 amino acids, a size comparable to other structurally resolved yeast and fungal GH32 enzymes, such as the invertase ScINV from Saccharomyces cerevisiae (533 aa; Sainz-Polo et al. 2013), the β-fructofuranosidases SoFfase from Schwanniomyces occidentalis (512 aa; Álvaro-Benito et al. 2010), XdINV from Xanthophyllomyces dendrorhous (665 aa; Ramírez-Escudero et al. 2016), and AkFfase from Aspergillus kawachii (634 aa; Nagaya et al. 2017), although it shows relatively low sequence identity with them. Generally, yeast invertases exhibit high sequence similarity among themselves, as do fungal ones, although the similarity between these two groups is markedly lower (Jiménez-Ortega et al. 2022; Nagaya et al. 2017). In line with the well-established structural conservation of the GH32 fold, the AlphaFold-derived model of TaINV adopts a structural fold similar to that of the fungal AkFfase (sequence coverage, 94%; identity, 26%) and the yeast XdINV (sequence coverage, 94%; identity, 24%) (Fig. 3A).Fig. 3. Preliminary structural model of TaINV. The protein model was obtained using AlphaFold3. A Structural superimposition of XdINV (PDB code: 5ANN) from Xanthophyllomyces dendrorhous (light blue) and AkFfase (PDB code: 5XH8) from Aspergillus kawachii (mint) onto TaINV model (orchid). Superposition with XdINV and AkFfase yielded Cα RMSD values of ~ 1.1 Å over approximately 320 pruned atom pairs, corresponding to the conserved GH32 catalytic core after exclusion of flexible peripheral regions and species-specific insertions. B The catalytic domain composed of blades I-V and the β-sandwich domain (blue) linked by α-helix (orange) are represented. C Close-up of the TaINV active-site region showing residues selected for site-directed mutagenesis. The sucrose molecule, taken from the crystal structure of XdINV-substrate complex (PDB: 5FIX), was positioned by structural superposition and is shown only as a reference for substrate-binding pocket location. Catalytic residues are shown in pink, and residues selected for this study in blue
The protein TaINV displayed the characteristic structural features of the GH32 family, composed of two domains. The β-propeller catalytic domain (residues 40–455) consists of five blades (I–V), each formed by four antiparallel β-strands (A–D) connected by loops in the classical W topology (Fig. 3B). The β-sandwich domain (residues 465–643) composed of twelve β-sheets arranged into two antiparallel six-stranded β-sheets, folded into a β-sandwich topology. Both domains are connected by a short α-helix (residues 456–464). Notably, the β-sandwich domain is the most variable region among GH32 enzymes and is proposed to contribute to substrate binding and specificity (Álvaro-Benito et al. 2010; Cuskin et al. 2012).
Figure 4 includes a sequence alignment of the catalytic domain of TaINV and some of the structurally resolved GH32 proteins from eukaryotic microorganisms that displayed the highest sequence similarity. All conserved motifs of the GH32 family were identified in the TaINV sequence, including WMNDPNG (WVNDPCG in TaINV), FRDP (WRDP in TaINV), and MWECPDF (NFEVTNF in TaINV). The catalytic triad residues (shown in bold) were identified as Asp63 (nucleophile), Asp201 (stabilizer of the transition-state), and Glu277 (acid–base catalyst). The relevance of these three amino acids for the hydrolytic activity was confirmed by site-directed mutagenesis. Thus, the protein variants, including substitutions D63A, D201A, and E277A, were obtained, expressed in E. coli, purified, and none of them showed any residual activity toward sucrose (data not shown). In addition, several other highly conserved amino acids in GH32 proteins were also identified in the TaINV sequence as potentially involved in sucrose hydrolysis based on sequence and structural correspondence with experimentally characterized GH32 enzymes (Fig. 4, Table S3). Residues corresponding to the −1 subsite, which accommodates the fructosyl moiety of sucrose, include Q81, F125, T126, R200, and W387, consistent with annotations reported for other GH32 enzymes (Kubota et al. 2022; Nagaya et al. 2017). Residues corresponding to the + 1 subsite, which interacts with the glucosyl unit of sucrose, can be identified as I154, H155, W156, E300, K328, and Y369 (Table S3).Fig. 4. Structural alignment of the catalytic domain sequences of TaINV with homologues. Catalytic domain of TaINV superimposed onto those of the β-fructofuranosidase from A. kawachii (AkFfase, PDB code 5XH8), β-fructofuranosidase from X. dendrorhous (XdINV, PDB code 5ANN), fructosyltransferase from Aspergillus japonicus (AjFfase, PDB code 3LDK), and invertase from S. cerevisiae (ScINV, PDB code 4EQV). Clustal Omega was used for the alignment and ESPript 3.0 for visualization. Conservative motifs are marked within black boxes and catalytic residues are marked with black asterisks
A series of TaINV variants was generated to examine the role of specific amino acids (highlighted in Fig. 3C) in the sucrose-hydrolyzing activity of the protein. Residue selection was based on conservation patterns within the GH32 family and on previous structure–function studies of homologous enzymes, including AkFfase, XdINV, AjFfase, and ScINV (Lafraya et al. 2011; Nagaya et al. 2017; Ramírez-Escudero et al. 2016; Trollope et al. 2015). Accordingly, TaINV variants carrying the substitutions W60Y, N62S, F125W, T126D, H151Y, E300R, and K328H/V were produced, purified, and evaluated for their activity toward sucrose (Table S2). All TaINV variants exhibited reduced specific activity on this substrate, ranging from a moderate twofold reduction with the substitution F125W to a 69-fold decrease with T126D. In contrast, variants including the substitutions E300R and K328H/V showed an almost complete loss of activity, with specific activity reduced by approximately 4000- to 8000-fold, respectively.
The kinetic analysis using sucrose as substrate revealed that substitutions W60Y and T126D did not significantly affect the TaINV Km values for sucrose, whereas H151Y practically doubled it, F125W quadrupled it, and N62S reduced it by half (Table 3, Figure S3). This suggests that, except for W60Y and T126D, these substitutions alter the affinity of the enzyme for sucrose (Table 3). In addition, only the variant including F125W showed an increased catalytic constant, while all the variants displayed a decreased catalytic efficiency (kcat/K _m_ ratio), with reductions ranging from 3- to 44-fold. These results highlight the importance of the substituted residues in the hydrolytic activity of TaINV. Table 3. Kinetic parameters of TaINV variants including the indicated substitutionsVariantKm (mM)kcat (s^−1^)kcat/Km (s⁻^1^ mM⁻^1^)WT8.7 ± 0.8479 ± 355 ± 4W60Y10.1 ± 176.1 ± 3.27.5 ± 0.3N62S3.6 ± 0.420.3 ± 1.55.6 ± 0.2F125W37.5 ± 5.6629.7 ± 1516.8 ± 2.7T126D7.2 ± 0.98.9 ± 0.11.2 ± 0.1H151Y22.1 ± 3.2144.3 ± 8.35.5 ± 0.3Sucrose was used as substrate, and values represent the average of three experiments. The kcat values were calculated assuming a protein molecular mass of 74 kDa. The low activity detected in TaINV variants including substitutions E300R and K328H/V (Table S2) was incompatible with their inclusion in this assay. The ± sign refers to the standard errors based on the curve fitting Michaelis–Menten model using SigmaPlot
Characterization of TaINV transfructosylation activity
The transfructosylating activity of TaINV was evaluated using 500 g/L sucrose as substrate, and most of the products detected in the reaction mixtures were identified and quantified by HPAEC-PAD (Fig. 5). Along with the expected glucose and fructose resulting from sucrose hydrolysis, at least nine peaks were detected in the reaction mixture. Six peaks (peaks 4–9 in Fig. 5A) were identified as 1-kestose, blastose, 6-kestose, neokestose, nystose, and fructosylnystose based on the commercially available or previously purified standards, while at least three additional peaks remained unidentified. Maximum FOS production was practically reached between 3 and 6 h of reaction (Fig. 5B). After 5 h, 88.3% of the sucrose was consumed, yielding 251.6 g/L of total FOS, which represented about 50.3% (w/w) of the total sugars in the reaction mixture. At the maximum FOS production point (~ 89% of initial sucrose consumed; Table S4), the main FOS produced by TaINV was 1-kestose (214.4 g/L representing 85% of total FOS), followed by the inulin-type FOS nystose (17.3 g/L), and fructosylnystose (2.3 g/L), the levan-type FOS 6-kestose (10.7 g/L) and with minor quantities of the neo-FOS neokestose (5.6 g/L) and blastose (1.3 g/L) (Fig. 5C, Table S4). Although TaINV predominantly produces 1-kestose, low levels of additional FOS, including levan-type and neo-FOS, were also detected. The formation of such minor products in addition to the primary product, 1-kestose, has also been reported for fungal extracts from Cladosporium cladosporioides and Penicillium sizovae (Zambelli et al. 2014).Fig. 5. Analysis of transferase reactions mediated by TaINV. A HPAEC-PAD chromatogram of a 5-h reaction performed with 10 U/mL of enzymatic activity and 500 g/L sucrose. Peaks: (1) glucose, (2) fructose, (3) sucrose, (4) 1-kestose, (5) blastose, (6) 6-kestose, (7) neokestose, (8) nystose, (9) fructosylnystose, and (*) unidentified products. B Time-course of quantified sugars in the reaction mixture: sucrose (pale blue triangles), fructose (green squares), glucose (red circles), and total FOS (orange inverted triangles). C Time-course of each of the FOS produced and identified, 1-kestose (pink circles) on the right y-axis, and the rest of the FOS individually on the left y-axis of which: neokestose (green inverted triangles), 6-kestose (blue triangles), blastose (coral squares), nystose (light blue rhombus), and fructosylnystose (purple circles). All reactions were performed in triplicate and standard errors are represented
Transfructosylation activity of TaINV variants
The ability of the five TaINV variants with the highest specific activity on sucrose (W60Y, N62S, F125W, T126D, and H151Y) to synthesize FOS was evaluated (Fig. 6 and Fig. 7). Among them, substitutions W60Y and N62S caused a significant increase in the production of total FOS, which at the maximum production point (~ 93–96% of initial sucrose consumed; Table S4) reached about 313 g/L (after 3 h) and 288 g/L (after 2 h), representing 62.7% and 57.5% (w/w) of the total sugars in the mixture, respectively (Fig. 6, Table S4). These increases were mainly due to enhanced 1-kestose production (Fig. 7). Indeed, W60Y and N62S variants produced approximately 258 g/L (83.2% of total FOS) and 232 g/L (80.3% of total FOS) of 1-kestose, corresponding to increases of ~ 20.3% and 8.2%, respectively, compared with the wild type, at maximal FOS-production points (Table S4). Both protein variants also promoted nystose formation, which increased up to 300% (103 g/L, 24 h) with W60Y and 122% (57 g/L, 4 h) with N62S. In the case of fructosylnystose, only the substitution W60Y increased production of this sugar (about 518% after 24 h; 23.7 g/L). W60Y and N62S variants showed decreased 6-kestose formation (3.6 g/L at 24 h and 8.6 g/L at 4 h, respectively). Regarding the neo-FOS production, substitutions W60Y and N62S increased blastose synthesis by 141% (10 g/L; 24 h) and 213% (13 g/L; 8 h), respectively, while both variants displayed reduced neokestose production.Fig. 6. Time course of total FOS production by the TaINV variants including the referred substitutions. Reactions included 5–10 U/mL of sucrose hydrolytic activity and 500 g/L sucrose in sodium acetate 50 mM pH 5.5 at 45 °C. The total FOS corresponds to the sum of all quantified FOS (1-kestose, nystose, fructosylnystose, 6-kestose, blastose, and neokestose)Fig. 7. Time course of the indicated FOS production mediated by the TaINV wild-type enzyme (WT) and variants carrying the indicated substitution. Reactions included 5–10 U/mL of sucrose hydrolytic activity and 500 g/L sucrose in sodium acetate 50 mM pH 5.5, and were incubated at 45 °C. Reactions were performed in triplicate and standard errors are represented. The sugar composition of the reaction mixture at the maximal FOS production point for each TaINV variant is presented in Table S4
In addition, substitutions F125W, T126D, and H151Y have also caused changes in total FOS and ^1^F-FOS production, at maximal FOS-production points (~ 88–90% of initial sucrose consumed; Table S4). The substitution F125W reduced the production of total FOS by 49%, mainly due to an equivalent decrease in 1-kestose synthesis, and the inability to produce nystose and fructosylnystose. In contrast, F125W also caused about 1.5-fold increase in blastose and neokestose production and reduced the 6-kestose synthesis by almost twofold. However, substitution T126D produced a similar total amount of FOS as the wild-type enzyme, but with notable differences in the FOS composition. Specifically, it led to a slight increase in 1-kestose (220 g/L; 85% of total FOS), 20% reductions in nystose, and 5.2-fold increase in blastose. Finally, the variant including H151Y, substitution located near the active site of TaINV (Fig. 3C), led to a modest reduction in total FOS and 1-kestose synthesis (by 10% and 11%, respectively; Table S4), whereas in 24 h reactions decreased production of nystose and 6-kestose by approximately twofold, neokestose by 1.5-fold and increased that of blastose by 1.7-fold (Fig. 7).
Discussion
The invertase activity of several Trichoderma species had been previously reported, although it is not universally present across the genus, and no FOS-producing enzymes had been described in any of them prior to this study (Delabona et al. 2012; Silva et al. 2014; Vargas et al. 2009). In this work, the availability of the T. atroviride genome sequence, together with the detection of sucrose-hydrolyzing activity in this organism, enabled the identification and characterization of a novel GH32 β-fructofuranosidase (TaINV) with transferase activity, thus expanding the diversity of fungal sources with potential for FOS production. Most GH32 proteins from fungi, bacteria, and plants (e.g., Aspergillus japonicus,* Bifidobacterium longum*,* Arabidopsis thaliana*) are monomeric, but oligomeric assemblies have also been described (Chuankhayan et al. 2010; Jiménez-Ortega and Sanz-Aparicio 2024). Among GH32 enzymes, most of the proteins studied from yeasts, including those from Sw. occidentalis, R. dairenensis, and X. dendrorhous as well as the enzyme from the fungus A. kawachii and the bacterium Thermotoga maritima, oligomerize as dimers (Álvaro-Benito et al. 2010; Jiménez-Ortega et al. 2022; Jiménez-Ortega and Sanz-Aparicio 2024; Nagaya et al. 2017; Ramírez-Escudero et al. 2016), whereas the invertase ScINV from S. cerevisiae predominantly assembles as octamers (Sainz-Polo et al. 2013). However, in the zymogram performed in this work, TaINV appears to have an apparent molecular mass that could be compatible with that of a multimeric form, a priori with a tetramer (Fig. 1C).
With respect to TaINV catalytic properties, both the thermal behavior and the pH dependence of this protein (optimum activity at 40–45 °C and pH 5.5–6.5) were very similar to those of other intracellular GH32 proteins (Nadeem et al. 2015), as well as the low TaINV thermostability. Thus, whereas the extracellular enzyme from X. dendrorhous displayed maximum activity at 60–75 °C and T_50_ values of 80–82 °C, the intracellular one was only relatively stable below 45 °C, showing a drastic reduction in activity above 50 °C (Chen et al. 2011; Gimeno-Pérez et al. 2015).
TaINV displayed a high capacity for FOS production, with 1-kestose as the predominant product (~ 85% of total FOS). In addition, unlike most commercial enzymes that predominantly produce inulin-type of FOS, with 1-kestose and nystose as the main products (Choukade and Kango 2021; Sánchez-Martínez et al. 2020), and an average products DP of 3.2–3.7 at ≥ 85% sucrose conversion (Karkeszová and Polakovič, 2023), TaINV mainly produces 1-kestose (Fig. 5), with an average products DP of about 3.1 even at 92% sucrose conversion (Table S5). Lower DP FOS are linked to enhanced prebiotic effects (Tochio et al. 2018), which would highlight the TaINV potential for functional food applications.
Site-directed mutagenesis confirmed the essential role of the TaINV predicted catalytic triad (Asp63, Asp201, Glu277) and the functional relevance of residues T126 and F125, at subsite −1. Substitution T126D, designed to mimic the equivalent position (Asp122) in the homologous AkFfase, where it has been reported to participate in hydrogen bonding with the fructose moiety of sucrose (Nagaya et al. 2017), drastically reduced the catalytic efficiency of TaINV (Table 3), confirming the critical role of T126 in substrate hydrolysis. However, this change did not substantially affect 1-kestose production (Fig. 7; Table S4). The F125W substitution, likely due to steric hindrance from the bulkier side chain, also reduced catalytic efficiency and caused the strongest decrease in FOS synthesis, with 1-kestose markedly diminished and longer-chain FOS undetected. Similar effects were observed with an equivalent substitution (W82F) in the S. cerevisiae invertase, which resulted in the absence of 1-kestose synthesis and reduced 6-kestose formation (Lafraya et al. 2011). K328 in TaINV corresponds to His343 in XdINV and His310 in AkFfase, both also mediating enzyme-glucosyl interactions, whereas E300 corresponds to E334 in XdINV and E296 in AkFfase, residues previously shown to be involved in sucrose binding (Nagaya et al. 2017; Ramírez-Escudero et al. 2016). Variants of TaINV including substitutions K328H/V and E300R drastically reduced the enzyme hydrolytic activity (Table S2), underscoring the conserved catalytic role of subsite + 1 residues among GH32 enzymes.
Differences in the FOS profiles produced by the TaINV variants provide insight into how subtle active-site changes shape transfructosylation specificity. Substitutions W60Y and N62S significantly enhanced total FOS production, mainly through increased 1-kestose and nystose synthesis, reflecting enhanced β-(2 → 1) transfructosylation activity. The rise in fructosylnystose observed only in W60Y may result from replacing Trp with a smaller hydrophobic residue, which facilitates access for larger acceptors in fructosylation. Interestingly, the equivalent substitutions in the S. cerevisiae ScINV (W19Y and N21S) enhanced 6-kestose synthesis, the major FOS produced by this enzyme (Lafraya et al. 2011), suggesting that these equivalent substitutions could improve the production of the predominant FOS generated by each enzyme. In addition, all generated variants except the one with T126D showed reduced 6-kestose synthesis (Fig. 7; Table S4), suggesting that none of the introduced substitutions could favor β-(2 → 6) bond formation between fructose units. Furthermore, the increased neokestose levels detected with substitutions F125W and T126D (the latter in the 24 h-reaction), both located in subsite −1, suggest a shift in fructosyl acceptor specificity that may impact neo-series FOS production. Finally, the change H151Y, equivalent to F140Y in the A. japonicus AjFfase, previously reported to enhance 1-kestose production (Trollope et al. 2015), did not show the same effect in TaINV, further highlighting the mechanistic differences among fungal GH32 enzymes.
In conclusion, this study described a novel intracellular β-fructofuranosidase, TaINV from T. atroviride, which exhibits high transferase activity and mainly synthesizes 1-kestose. Structural and mutagenic analysis identified key residues involved in hydrolysis and transfructosylation. Notably, the TaINV variant W60Y increased total FOS production to 62.6% (w/w) of total sugars, reaching yields comparable to those of commercial enzymes. This work provides new insights into the structure–function relationship of a fungal GH32 β-fructofuranosidase producing low-DP FOS and highlights TaINV as a promising candidate for biotechnological applications.
Supplementary Information
Below is the link to the electronic supplementary material. ESM1(PDF 695 KB)
The reference list from the paper itself. Each links out to its DOI / PubMed record.
- 1Fortune Business Insights (2024) Functional food and beverage market size, share & COVID-19 impact analysis, by type (functional dairy products, functional bakery products, functional cereals & grains, functional fats & oils, and others), and regional forecast, 2024–2032. https://www.fortunebusinessinsights.com/functional-foods-market-102269. Accessed 26 May 2025
- 2Jiménez-Ortega E, Sanz-Aparicio J (2024) Oligomeric structure of yeast and other invertases governs specificity. In: Harris JR, Marles-Wright J (eds) Macromolecular protein complexes V: structure and function. Springer, Cham, pp 503–530. 10.1007/978-3-031-58843-3_1910.1007/978-3-031-58843-3_1938963498 · doi ↗ · pubmed ↗