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3 March 2026

19 Pages

Application of Enzyme Engineering and Synthetic Biology for Modulated Transformation of Fructooligosaccharides (FOSs) to Elucidate the Catalytic Mechanism of Fructofuranosidases

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1
School of Chemistry and Chemical Engineering, Guangxi Minzu University, Nanning 530006, China
2
Guangxi Subtropical Crops Research Institute, Guangxi Academy of Agricultural Sciences, Nanning 530001, China
3
Guangxi Key Laboratory of Quality and Safety Control for Subtropical Fruits, Key Laboratory of Quality and Safety Control for Subtropical Fruit and Vegetable, Ministry of Agriculture and Rural Affairs, Nanning 530001, China
4
Guangxi Key Laboratory of Green Processing of Sugar Resources, College of Biological and Chemical Engineering, Guangxi University of Science and Technology, Liuzhou 545006, China
Foods2026, 15(5), 843;https://doi.org/10.3390/foods15050843
This article belongs to the Special Issue Advances in Food Biotechnology and Enzyme Engineering
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Abstract

Fructooligosaccharides (FOSs) are plant-based prebiotics widely utilized in the food and pharmaceutical industries. As a major sugar-producing region, Guangxi holds significant potential for enzymatic production of FOS from sucrose. This study engineered a mutant enzyme, 142P-242K, to address the low catalytic activity characteristic of wild-type enzymes. The mutation upregulated the FOS conversion efficiency from 29 to 52%, respectively. Optimal enzymatic activity was observed at 45 °C, pH 6.0, and in the presence of 1 mM Na+. Mechanistic investigations revealed that modifications to the catalytic domain pocket and shifts in substrate affinity were the primary factors driving enhanced FOS production. The accumulation of 1-Kestose (GF2) was attributed to the enhanced flexibility of the 142P-242K loop, which facilitates substrate access to the active site. However, the synthesis of nystose (GF3) from GF2 is hindered by the hydrophobic nature of the active site and strong hydrogen bonds binding GF2. Comparing the enzyme’s ability to produce FOS using sugarcane juice, sugarcane molasses, and adsorption-heating sugarcane molasses, it was determined that heat-adsorbed molasses yielded the highest FOS concentration (30.77%). This study offers a practical and cost-effective strategy for enzyme modification and efficient valorization of molasses.

1. Introduction

A primary benefit of functional foods lies in their capacity to promote health and well-being while mitigating the risk of various diseases [1]. Prebiotics have been extensively explored within the context of functional foods due to their diverse and substantial health benefits [2]. These compounds typically comprise short-chain carbohydrates, the most prominent group being fructooligosaccharides (FOSs), which are oligosaccharides formed by the attachment of one to three fructose residues to the fructose moiety of a sucrose molecule. FOS demonstrates significant efficacy in enhancing gastrointestinal health and modulating blood glucose levels [3,4]. Moreover, it serves technical functions, such as acting as a functional carrier [5]. Nowadays, consumer trends reflect a shift in purchasing behavior toward functional foods that positively impact health [6]. Given the rise of health-conscious consumers, food is increasingly perceived as a vehicle for essential nutrition and functional components. Consequently, the investigation of prebiotic dietary fibers is of critical importance.
Fructooligosaccharides are most abundant in chicory roots [7], onions [8], and jicama [9], which serve as the primary raw materials for industrial extraction. However, plant cultivation is subject to various environmental factors and limitations, which have hindered the widespread adoption of plant extraction methods at present. Research indicates that FOS is primarily synthesized through enzymatic catalysis of sucrose. These enzymes, including β-fructofuranosidases (FFases, EC 3.2.1.26) and β-D-fructosyltransferases (FTases, EC 2.4.1.9), are produced intracellularly or extracellularly by various fungal genera, such as Aureobasidium sp., Penicillium sp., and Aspergillus sp. [10]. Recently, research on fructosyltransferases has focused primarily on expanding resource libraries [11,12], achieving high-efficiency expression [13,14], enhancing catalytic activity, and optimizing fermentation processes [15,16]. Direct evolution is a robust and widely employed methodology in enzyme engineering that can effectively enhance catalytic performance. However, this approach generates an extensive library of mutants, rendering the screening process time-consuming and labor-intensive [17].
Protein engineering employing computer-aided rational design has emerged as a robust strategy for developing enzymes with properties tailored for industrial applications [18,19]. Recent structural analyses of FTases have revealed that, despite varying degrees of homology across different sources, these enzymes share highly conserved folds and characteristic domains [20,21]. Docking simulations further indicate that these enzymes possess a similar protein–ligand interaction profile. Previous observations indicated that the total FOS yield correlates with the summation of binding affinities for 1-kestose (GF2), nystose (GF3), and 1F-fructofuranosylnystose (GF4) [22]. Currently, dynamic equilibrium remains a critical bottleneck limiting the industrial-scale application of FTases. Consequently, computer-aided analysis facilitates a deeper understanding of the affinity, mass transfer, and dissociation kinetics between the substrate and the active site, thereby providing practical guidance for enhancing FOS accumulation [23].
As the largest sugarcane-producing region in China, Guangxi serves as a pivotal research and industrial hub for the sugarcane industry [24]. Although sucrose is currently utilized primarily for crystalline sugar production, the process generates molasses as a major byproduct. This molasses contains substantial residual sucrose and remains an ideal substrate for further enzymatic conversion [25]. The application of converting such agricultural and industrial byproducts into high-value products like FOS demonstrates the potential to minimize production costs and alleviate environmental pollution. Furthermore, these strategies align perfectly with the principles of a circular economy [26]. Based on these considerations, the objective of the present study is to construct a dual-mutant FTase to enhance FOS production. Furthermore, we aim to perform an in-depth structural analysis of the engineered enzyme to elucidate the molecular mechanisms underlying the improvement in catalytic yield. By synthesizing FOS from sugarcane molasses, this study successfully transformed an industrial byproduct into a functional product. These findings provide a theoretical basis for promoting high-quality development within the sugar industry of Guangxi, China.

2. Materials and Methods

2.1. Plasmids, Strains, and Culture Medium

All plasmids and strains used in the present study are summarized in Supplementary Table S1. E. coli DH5α (Sangon Biotech, Co., Ltd., Shanghai, China) was used to construct recombinant plasmids and grown at 37 °C in Luria–Bertani (LB) medium (10 g/L peptone, 5 g/L yeast extract, and 10 g/L NaCl, with or without 15 g/L agar) (Sangon Biotech, Co., Ltd., Shanghai, China). Additionally, zeocin (100 μg/mL) (Jino Biotechnology Co., Ltd., Nanning, China) was added appropriately to culture medium.

2.2. Genetic Constructions

The recombinant plasmid pPICzαA-142P was constructed in our previous study [23] and used as the template of mutagenesis. The E. coli strain DH5α and P. pastoris GS115 (Invitrogen, Carlsbad, CA, USA) were separately applied as the host for DNA manipulation and protein expression. Amino acids that were screened in the preliminary study were mutated at sites 154 and 242 to form hydrogen bonds (Table 1). Mutants and ligands were submitted to CB-DOCK2 Online Server (http://183.56.231.194:8001/cb-dock2/index.php) (accessed on 20 March 2025) for molecular docking to screen for mutants exhibiting increased hydrogen bond counts. Primers for the introduction of mutation sites are shown in Supplementary Table S2. Two gene fragments were amplified using the primer pairs F/R1 and F1/R, respectively, and then fused by overlap extension PCR to introduce the mutation site. The PCR products were ligated with the vector pPICzαA (Invitrogen, Carlsbad, CA, USA) to form the recombinant plasmid. The linearized recombinant plasmid was introduced into P. pastoris GS115 by electroporation to construct the mutant strain.
Table 1. Analytical results of the number of hydrogen bonds in mutants.

2.3. Enzyme Expression and Activity Assay

Seed solutions were inoculated in YPD (Sangon Biotech, Co., Ltd., Shanghai, China) medium for overnight incubation. After that, they were inoculated into BMGY medium at 10% inoculum for 24 h of preculture. Recombinant P. pastoris GS115 were collected, washed twice in sterile water, resuspended in BMMY and replenished with 1% methanol every 24 h. The target protein was purified by a Ni Sepharose 6 Fast Flow affinity column (Sangon Biotech, Co., Ltd., Shanghai, China) (the wild-type FTases and their mutants fused with a C-terminal 6-histidine tag). The enzyme activity of fructosyltransferase was determined according to Kubota [27] with slight modifications. An appropriate amount of enzyme solution and sucrose substrate (Sangon Biotech, Co., Ltd., Shanghai, China) (500 g/L) were mixed in 0.1 M phosphate buffer sucrose solution (pH 6.0) (Sangon Biotech, Co., Ltd., Shanghai, China). The reaction was terminated in a boiling water bath for 15 min after the reaction was carried out (45 °C, 10 min).
To monitor the changes in FOS composition during catalysis by different mutant enzymes, FOS production was carried out by adding purified enzyme samples (1.68 mg/mL) to sucrose (500 g/L, w/v) dissolved in 0.1 M phosphate buffer solution (pH 6.0) for 24 h at 45 °C. Samples were taken every 3 h for FOS yield detection. FOS yield is defined as the ratio of FOS produced in the reaction to the initial sucrose substrate. FOSs were detected through an XBridge BEH Amide Column (130 Å, 5 μm, 4.6 mm × 250 mm) (Infever Biotechnology Co., Ltd., Suzhou, China) by Liquid Chromatography Waters 2595with Water 2414 differential refractive detector parameters (Water, Milford, MA, USA), such as 35 °C column temperature. The retention times for fructose, glucose, sucrose, 1-kestose (GF2), and nystose (GF3) (Yuanye Bio-Technology Co., Ltd., Shanghai, China) were 7.166, 8.580, 11.307, 17.945, and 26.031 min. Here, 1 U fructosyltransferase is defined as the amount of enzyme used to catalyze the reaction of the substrate sucrose to produce 1 µmol of FOS in 1 min.

2.4. Analysis of Enzymatic Activities

The effect of temperature on enzyme activities were measured over 40–75 °C every 5 °C (pH 7.0), and the optimum enzyme activity was defined as 100% relative enzyme activity. To assess the effect of pH on the catalytic activity of variants, enzymes were catalyzed at different pH gradients (pH 3.0–9.0) with specific intervals of 1.0 for 10 min. The highest enzyme activity was defined as 100% relative enzyme activity. To obtain the optimum metal ions for enzymes, different metal ions of 1 mM concentration (Cu2+, Co2+, K+, Ca2+, Mg2+, Na+ and Ba2+) were added to the catalytic system. The control was treated with buffer solution in place of metal ions, and the enzyme activity of the control group was defined as 100% relative enzyme activity.

2.5. MD Simulation and Structural Analysis

Using fructosyltransferase (PDB ID: 3LF7) as a template, structural models of the 142P and its variants were obtained through the SWISS-MODEL online modeling server (https://swissmodel.expasy.org/interactive) (accessed on 20 March 2025). Enzyme solubility calculation involved importing the protein sequence into Chemistry of Health software (https://www-cohsoftware.ch.cam.ac.uk/index.php/camsolph) (accessed on 1 September 2025), setting the pH range, and submitting for computation. Enzyme–ligand complexes were obtained using CB-DOCK2 molecular docking. Molecular dynamics simulations of 100 ns were performed on the complexes obtained by molecular docking using Gromacs v2023.02 software [28].
The simulation system was constructed using GROMACS v2023.02 software, establishing a water box model. Parametrization was performed using the Amber99sb-ildn force field, with TIP3P water molecules implemented for system solvation. The minimum distance between protein atoms and the water box edges was set to at least 1.2 nm. The simulation system charge was neutralized by adding an appropriate number of Na+ and Cl- ions. The system was slowly heated from 0 to 300 K, then equilibrated in an isothermal–isobaric (NPT) ensemble at the temperature of 300 K and pressure of 1 bar (1 bar = 1 × 105 Pa) for an equilibration time of 100 ps. A 100 ns molecular dynamics simulation was performed on the complex. Based on the MD simulation results, the built-in tools of GROMACS software were used to calculate the complex’s root mean square deviation (RMSD), root mean square fluctuation (RMSF), radius of gyration (Rg), solution-accessible surface area (SASA), and number of hydrogen bonds (H-bonds).

2.6. Treatment of Different Raw Materials

Adsorption-heating treatment (H) was applied by adding 6% activated carbon (m/v), heating at 60 °C in a water bath for adsorption (30 min), then adding 3% siliceous earth (m/v) for adsorption (1 h). Finally, samples were centrifuged and filtered to remove impurities. Unprocessed sugarcane molasses was used as the control (CK). Sugarcane juice (ZZ) was centrifuged and filtered to remove impurities, and the supernatant was collected. The sucrose content in substrates from different sucrose sources was determined by HPLC, and substrates were diluted to ensure the same sucrose concentration for use in FOS synthesis. The method for synthesizing FOS was the same as the sucrose-based catalytic approach described in Section 2.3.

2.7. Statistical Analysis

All experiments were carried out independently thrice (n = 3) and data presented as mean ± SD. Statistical and numerical analyses were performed by GraphPad Prism 8.0.2 software (GraphPad Inc., Boston, MA, USA) and Origin version 9.1 (OriginLab, Northampton, MA, USA). Significant differences (p < 0.05) were determined by Fisher’s least significant difference (LSD) test.

3. Results and Discussion

3.1. Selection of Mutation Site Regions

Our preliminary research indicated that the substitution of serine with proline at position 142 enhances hydrophobic interactions and hydrogen bonding. This structural change ultimately alters the catalytic mechanism of the enzyme. Specifically, tryptophan at position 242 recognizes the oxygen atom within the fructose residue of sucrose. The enzyme then catalyzes 1-kestose (GF2) formation through sequence of hydrolysis, mass transfer, and covalent bonding with free fructose residue. Subsequently, the oxygen atom of the fructose residue in GF2 interacts with arginine at position 154 and covalently bonds with a new fructose residue to form nystose (GF3). The oxygen atom of the fructose residue in GF3 then re-associates with tryptophan at position 242 to facilitate the binding of subsequent fructose residues. Based on these observations, we hypothesized that arginine 154 and tryptophan 242 are critical residues for the FOS accumulation observed in the 142P mutant [23].
To validate this hypothesis, site-directed mutations were introduced at these two positions based on molecular dynamics simulations using GF2 as the docking ligand. Specifically, residues at positions 154 and 242 were substituted with various amino acids capable of forming enhanced hydrogen bond networks (Table 1). Eleven novel mutants were ultimately identified and selected for further investigation after comprehensive comparison of their hydrogen bond counts against the 142P control variant.

3.2. Enzyme Expression and Forward Mutation Screening

FTase initially hydrolyzes sucrose into glucose and a fructosyl moiety. Subsequently, the fructosyl group is transferred to another sucrose molecule to synthesize 1-kestose (GF2). A subsequent fructosyl group then covalently bonds to this trisaccharide to form nystose (GF3), a process that continues with the addition of further fructosyl groups to generate longer FOS chains [29]. In this study, mutant enzymes were incubated with the sucrose substrate for a 24 h catalytic reaction. The resulting FOS content was then quantified by using high-performance liquid chromatography (HPLC), with the total FOS yield serving as the primary indicator for assessing enzymatic performance (Figure 1). Among the tested variants, only the 142P-242K mutant exhibited enhanced FOS conversion relative to the 142P control group. Notably, mutations at position 154 resulted in a near-total loss of enzymatic activity.
Figure 1. Screening for FOS conversion ability. The mutant enzyme 142P was used as a control (CK). ****—p < 0.0001. Gray column—control, Green column—reduction, Pink column—increase.
To further investigate the underlying causes of activity loss, structural analyses of the relevant mutant enzymes were performed. The interactions between ligand GF2 and active-site residues are illustrated in Figure 2. Mutations at position 154 resulted in the disruption of critical interactions between this site and the ligand. In the 142P-154K mutant, site 154 still maintained hydrophobic interactions with GF2. Consequently, this mutant retained only marginal FOS conversion capacity. However, hydrophobic interactions alone were insufficient to improve FOS conversion. Table 1 illustrates that the hydrogen bonding between GF2 and mutant enzymes was strengthened. Despite this enhancement, the inability of site 154 to effectively recognize and bind GF2 prevented the catalytic reaction from proceeding. Based on the experimental data in Figure 1, mutations at position 154 led to significant loss of enzymatic activity. These findings indicate that position 154 is a highly conserved site essential for the catalytic function of the enzyme. Conserved substrate-binding sites are critical for maintaining catalytic performance, as substitutions at these residues can significantly compromise overall enzyme function [30]. Further structural analysis of mutants that retained activity revealed that the hydrogen bond interaction between site 154 and GF2 remained intact despite the mutations introduced at position 242 (Figure 2). This observation confirms that site 154 is a pivotal residue for ligand recognition by the enzyme. The loss of binding affinity at this position directly led to the complete abolition of catalytic activity. Based on the results presented in Figure 1, the mutant enzymes 142P-154K, 142P-242K, 142P-242D, and 142P-242Y were selected for subsequent detailed analysis.
Figure 2. Visualization analysis of molecular docking between GF2 and mutant enzymes of site 154. The mutant enzyme 142P was used as control (A), and mutant enzyme at site 154 (B1–B6). The red arrow and circle—the binding of the ligand to the 154th site of the receptor.

3.3. Analysis of Conversion Efficiency Regulation

The affinity of multistep reactions for various substrates determines the proportional distribution of mixed products [22]. The primary objective of this study was to enhance FOS accumulation by increasing the binding affinity for GF2. Consequently, the FOS conversion rates of the mutants identified in Section 3.2 were assessed (Figure 1). The FOS conversion rate of 142P-242K mutants increased (52%), representing 1.79-fold improvement over the control group. However, the other three mutants, specifically 142P-154K, 142P-242D, and 142P-242Y, exhibited varying degrees of reduction in conversion efficiency. Enzymatic activity is mediated by interactions between the active site and ligand through various non-covalent bonds, with a particular emphasis on the reaction transition state [31,32]. Internal factors, such as mutations, and external factors including pH, temperature, and ions determine enzymatic catalytic efficiency by influencing substrate binding, transition state stabilization, or product release [33,34].
Building upon the conclusions from Section 3.2 and integrating molecular docking results, we attributed the reduced conversion rates of 142P-242Y and 142P-242D to the loss of specific binding interaction between GF2 and site 154 (Figure 3). Although the total number of hydrogen bonds in these mutants exceeded those in the 142P variant, this increase did not translate into an improved FOS conversion rate. For the 142P-242K mutant, the number of hydrogen bonds was higher than that of 142P while its hydrophobic interactions were significantly lower. In most instances, enhanced hydrophobic interactions significantly upregulate the binding affinity between enzymes and substrates. A hydrophobic environment typically helps to reduce the dielectric constant of key chemical bonds to stabilize transition states characterized by charge separation [35]. Nevertheless, experimental results indicated that the enhanced conversion rate of the 142P-242K mutant occurred despite its reduced hydrophobicity (Figure 1). This observation suggests that hydrogen bonding plays a more dominant role than hydrophobic interactions in this catalytic process. Transition states typically possess greater hydrogen bonding potential than either substrates or products. Furthermore, enzymes significantly lower the activation energy of reactions by forming additional hydrogen bonds with the transition state [36]. In summary, we concluded that the observed variations in mutant enzyme activity result from the synergistic effects of hydrogen bonds and hydrophobic interactions.
Figure 3. Visualization analysis of molecular docking between GF2 and mutant enzymes of site 242. The mutant enzyme 142P was used as control (A), and mutant enzyme at site 242 (B–D). The red arrow and circle—the binding of the ligand to the 154th site of the receptor.
The sugar composition of the synthesized FOS was subsequently determined. As illustrated in Figure 4A, the proportions of GF2 and GF3 for the 142P-242K mutant were 47 and 5%, respectively. Compared to the control, the 142P-242K mutant exhibited increased GF2 content alongside the reduction in GF3. Based on the molecular docking results presented in Figure 5, the increased hydrogen bond interactions between 142P-242K and GF2, coupled with the reduced interactions with GF3, are hypothesized to be the primary cause of this shift in FOS composition. Furthermore, we evaluated the dynamic catalytic process of the mutant enzymes. Using sucrose (500 g/L) as the substrate, samples were collected every 4 h to observe the production of FOS. The FOS levels reached a plateau after 20 h of reaction (Figure 4B). Throughout the entire process, the 142P-154K, 142P-242Y, and 142P-242D mutants produced only GF2. In contrast, the 142P-242K mutant initiated GF3 synthesis after 16 h of reaction, and yield reached equilibrium by 24 h.
Figure 4. Yield of FOS in mutants. Total FOS production (A), such as GF2 (light green) and GF3 (bottle green). Dynamic monitoring of FOS production (B), the green curve: GF2; and the blue curve: GF3. ****—p < 0.0001.
Figure 5. Hydrogen bond analysis of molecular docking. The mutant enzyme 142P was used as a control (A, green), and blue labels indicate dual-mutant enzymes (B–E).

3.4. Biochemical Characterization of the Mutant 142P-242K

Temperature serves as a major factor in governing catalytic reactions [37]. It directly affects molecular motion within the reaction system and influences the frequency of contact between substrates and enzymatic active sites [38]. As illustrated in Figure 6A, the optimal catalytic temperature for the 142P-242K mutant was 45 °C. Enzymatic activity increased gradually within the temperature range of 30 to 45 °C. When the temperature exceeded 45 °C, the activity dropped sharply and approached complete inactivation beyond 55 °C. Compared with the control variant 142P, the optimal catalytic temperature reduced by 10 °C. The lower catalytic temperature offers significant advantages for industrial applications, including reduced energy consumption and facilitation of green manufacturing processes [39]. Theoretically, optimum temperatures accelerate molecular diffusion and upregulate the probability of substrate-active site encounters [40]. However, elevated temperatures enhance the fluidity of the reaction system while simultaneously destabilizing the enzyme structure [41,42]. This structural instability potentially leads to dissociative denaturation and subsequent inactivation. However, optimum temperatures are favorable for industrial-scale FOS production. Such conditions accelerate the conformational changes of the substrate and promote reaction equilibrium toward product synthesis. Furthermore, elevated temperatures increase sugar solubility and reduce the risk of microbial contamination during the process [43,44].
Figure 6. Enzymatic characterization of the mutant 142P-242K. Optimal temperature (A), optimal pH (green) and intrinsic solubility of the enzyme (red) (B), and optimal metal ions (C). For the investigation of temperature and pH, the maximum enzyme activity was defined as 100% relative enzyme activity. For the assessment of metal ions, the enzyme activity of the control group was defined as 100% relative enzyme activity. ns—non-significant, *—0.01 < p < 0.05, ****—p < 0.0001.
Similarly, pH serves as a critical factor influencing enzymatic activity. It can modify the spatial structure of enzymes and influence the binding affinity between substrates and active-site amino acid residues [45]. Consequently, identifying the appropriate pH value is essential for maximizing product accumulation. The 142P-242K mutant exhibited maximum activity at pH 6.0, whereas pH values below 4.5 resulted in enzymatic inactivation (Figure 6B). The shift of the optimal catalytic pH from 5.0 (142P) to 6.0 provides a milder reaction environment. In industrial applications, this adjustment reduces corrosion and pH-control costs, while supporting more environmentally friendly and sustainable biocatalytic processes [46]. These enzymes generally demonstrate a bias toward slightly acidic conditions. Stable folded proteins typically possess large insoluble regions from the hydrophobic core of the folded state [47]. Therefore, the intrinsic solubility of the enzyme across the range of pH values was calculated using computational analysis as shown by the red curve in Figure 6B. The observed reduction in solubility within the weakly acidic range further suggests that such conditions are more favorable for maintaining the hydrophobic structure of the enzymatic active sites and facilitating substrate conversion. Similar studies have also reported that the FTases typically exhibit a preference for acidic conditions and often require an acidic environment to maintain optimal catalytic activity [48,49].
Specific enzymes are classified as metal-dependent due to their multifunctional chemical reactivities, which include acidic, electrophilic, and nucleophilic properties [50]. The catalytic functions of these enzymes are typically enhanced when they are supplemented with specific metal ions to help facilitate some of the most complex reactions in nature [51]. Previous research confirmed that FTase functions as a metal-dependent enzyme [23]. Therefore, we investigated the effects of various metal ions on enzymatic activity to identify conditions for maximum performance. As illustrated in Figure 6C, all seven metal ions assessed promoted enzymatic activity to varying degrees. Among them, Na+ exhibited the most significant enhancement and increased the relative enzymatic activity by 61.33%. After engineering, the auxiliary metal ion was changed from Co2+ (142P) to Na+, reducing raw material costs and toxicity, avoiding issues related to heavy metal handling, and aligning with environmentally friendly and sustainable biocatalysis principles. FOS molecules carry a negative charge due to electrostatic repulsion between multiple groups bearing the same charge along the chain. This negatively charged substrate interacts with positively charged metal ions to form a transition state. This complex undergoes a nucleophilic reaction with the active-site amino acids to facilitate substrate conversion [52].

3.5. Enzyme Structure Analysis

The experimental results demonstrated the significant increase in the FOS conversion rate for the 142P-242K mutant, as discussed in Section 3.3. Molecular docking elucidated the mechanisms behind the enhanced FOS conversion in 142P-242K. Following the mutation at position 242, the active cavity underwent a slight expansion (Figure 7A,B). Enzyme structures typically undergo conformational modifications in response to their environment. Consequently, when different substrates enter the active site cavity, the distinct interactions between substrates and enzyme determine specific variations in the cavity structure [53,54]. For sucrose, the larger active pocket in the 142P variant allowed more flexible substrate entry/exit, explaining its higher hydrolytic activity compared to 142P-242K (Figure 7(A1,B1)).
Figure 7. Structural domain comparison analysis of 142P and 142P-242K, such as 142P (A), 142P-242K (B), 1–3, Molecular docking with sucrose, GF2 and GF3. The yellow box indicates the active center of the enzyme. The color variation within the active center represents structural flexibility, where blue denotes rigidity and red denotes flexibility.
However, for 1-kestose (GF2) synthesis, the active pocket of 142P-242K transitioned to a more open conformation, facilitating the accumulation of GF2 (Figure 7(A2,B2)). Similar research findings have also proposed that widening the substrate-binding pocket serves as an effective strategy to enhance enzymatic activity [55]. Conversely, when GF3 bound to 142P-242K, the pocket contracted due to altered intermolecular interactions (Figure 7(A3,B3)). FTase possesses hydrolytic and fructosyl-transferring activities, which allow the enzymatic reaction to progress continuously in the forward direction [56]. This restriction hindered the mobility of longer chains, inhibiting further fructosylation and leading to lower GF3 but notably higher GF2 levels compared to 142P.

3.6. Mechanism for Enhancement of Catalytic Activity by MD Simulations

To analyze the mechanisms underlying the enhancement of the conversion rate in depth, molecular dynamics (MD) simulations were conducted for various enzyme–substrate complexes. During the 100 ns simulation process, both 142P and 142P-242K exhibited a single energy well, which indicates that each maintained only one stable conformation with minimum energy (Figure 8(A1,A2,B1,B2) and Figure 9(A1,A2,B1,B2)). When the Gibbs free energy decreases, the system tends to undergo changes, such as the formation or breaking of chemical bonds [57]. The 142P-242K mutant exhibited higher values for the radius of gyration (Rg), RMSD, RMSF and SASA, indicating greater structural flexibility and surface exposure (Figure 8D,E1,F,G). Enzyme catalysis is the dynamic process where structural flexibility often correlates positively with catalytic activity [33,58]. As structural flexibility often correlates positively with catalytic activity, this enhanced flexibility facilitated easier substrate access to the catalytic center [59,60,61,62]. For the enzyme–GF2 complex, the stable hydrogen bond network and lower RMSD value in 142P-242K helped to maintain the stability of the bound substrate (Figure 8C,E2). However, for GF3, the mutant displayed lower Rg, RMSD, and SASA values compared to 142P, indicating higher stability that restricted GF3 transport (Figure 9C–G).
Figure 8. Comparison of molecular dynamics simulations for 142P and 142P-242K with GF2. Free energy landscape of 142P with two-dimensional and three-dimensional structures (A1,A2), the free energy landscape of 142P-242K with two-dimensional and three-dimensional structures (B1,B2), analysis of hydrogen bond changes (C), radius of gyration (D), root mean square deviation of complex and monomer (E1,E2), root mean square fluctuation (F), and solvent-accessible surface area (G).
Figure 9. Comparison of molecular dynamics simulations for 142P and 142P-242K with GF3. Free energy landscape of 142P with two-dimensional and three-dimensional structures (A1,A2), the free energy landscape of 142P-242K with two-dimensional and three-dimensional structures (B1,B2), hydrogen bond changes (C), radius of gyration (D), root mean square deviation of complex and monomer (E1,E2), root mean square fluctuation (F), and solvent-accessible surface area (G).
In summary, we concluded that the shift in the FOS conversion rate induced by the Lys mutation at position 242 resulted from the synergistic effects of multiple factors. This study identified a more spacious active binding domain within the 142P-242K mutant. This structural modification provided sufficient space for the efficient exchange of various substrates. Similar studies previously concluded that replacing key amino acids in the active site with Ala, which possesses a smaller side chain, develops a more spacious substrate-binding pocket. Such modifications promote structural flexibility and potentially increase the rate of entry and exit for substrates and their glycosides. Therefore, the specific residue composition and narrow shape of the channel determine the specificity of substrate recognition [63]. For example, the molecular dynamics simulation analysis of the T416C-I432C mutation demonstrated by Dodda et al. [64] revealed that the mutation widened the tunnel at the product exit site. This change facilitated substrate entry into the catalytic tunnel and allowed for the faster release of products compared to the wild-type enzyme.
Fructosyltransferase mediates multistep catalytic reactions, and its active domain is capable of recognizing multiple substrate molecules. The 142P-242K mutant exhibited a more flexible structure than the control group, which is a property critical for dynamic catalytic processes. Dotas et al. [65] proposed that conformational plasticity at enzymatic active sites controls substrate specificity. Generally, more flexible enzymes exhibit greater diversity and are capable of processing substrates with diverse chemical structures. As the simulation time increased, the magnitude of changes in hydrogen bonding for the 142P-242K mutant was smaller than that of the control group. Hydrogen bonds stabilize protein structures and are essential for catalysis, as the disruption of these bonds typically leads to protein denaturation [66]. Specifically, the more stable hydrogen bond network is essential when enzyme molecules are exposed to complex catalytic environments for extended periods. Although the FOS conversion capacity of the 142P-242K mutant increased, its GF3 content was significantly lower than that of the control group. We hypothesized that the extension of the substrate chain length exposed the hydrophobic environment during active-site binding, which subsequently weakened the binding affinity. Typically, active sites are located within hydrophobic cavities where substrates interact more effectively due to the surrounding non-polar environment [67].

3.7. Feasibility Evaluation of 142P-242K Mutant for FOS Synthesis from Different Raw Materials

Guangxi is a key region for the sugar industry, with abundant sugarcane resources, making it advantageous for FOS production [68]. This study compared the effects of sugarcane juice and sugarcane molasses on FOS production. The sucrose concentrations in these substrates were 272.13 (CK), 248.96 (H), and 135.67 g/L (ZZ), respectively (Figure 10A). The minimum loss of sucrose was observed after the adsorption-heating treatment compared to the untreated group. While maintaining a consistent initial sucrose concentration, we determined the effects of different treatments on the FOS conversion rate. As illustrated in Figure 10B, variations in FOS yields were observed across different raw materials, with measured values of 27.73, 30.77 and 5.61%, respectively. Molasses is a primary byproduct of the sugar industry. Due to its high sucrose content, it serves as a unique substrate for enzymatic conversion [25]. However, the complex sugar production process results in molasses that contains numerous impurities, such as colloids, ash, pigments, and metal ions [69].
Figure 10. Analysis of optimizing FOS synthesis from different sources. Different sources of sucrose content (A) and FOS production (B). Note: CK—control (untreated), ZZ—sugarcane juice, and H—adsorption heating. *—0.01 < p < 0.05, **—p < 0.01, and ****—p < 0.0001.
The H-treated method mainly removes most of the colloids [70]. The presence of colloids reduces the fluidity of the reaction system and hinders enzyme–substrate interaction. Furthermore, colloids can flocculate proteins, resulting in the loss of enzymatic activity. This serves as a primary factor contributing to the highest FOS yield observed after the adsorption-heating (H) treatment. Molasses possesses a strong natural pH buffering capacity, which mitigates pH fluctuations caused by acid production during side reactions and ensures that FTase remains within its optimal activity of pH 5.0–6.0 [71]. FTase is a metal-dependent enzyme, and an appropriate concentration of specific metal ions is essential to enhance its catalytic activity [50]. Residual metal ions in molasses may interact directly with the active site or allosteric sites of FTase to enhance its affinity for the sucrose substrate. This interaction potentially leads to higher FOS production within the same timeframe and explains why molasses exhibits a superior capacity for FOS synthesis compared to sugarcane juice.
Despite the significant improvements in FOS yield, this study has various limitations. Conceptually, while the 142P-242K mutant effectively enhances the accumulation of 1-kestose (GF2), its catalytic efficiency for longer-chain fructooligosaccharides, such as nystose (GF3), remains constrained due to the contraction of the active pocket and increased hydrophobic exposure. Methodologically, the engineered enzyme exhibits limited thermostability, with a sharp decline in activity beyond 45 °C, which may restrict its application in industrial processes requiring higher operating temperatures to minimize microbial contamination. Furthermore, although the adsorption-heating treatment successfully valorizes sugarcane molasses, the slight reduction of sucrose during this process suggests that the purification strategy could be further optimized to maximize substrate utilization. Future research should focus on multiobjective enzyme engineering to simultaneously enhance product specificity and thermal robustness.

4. Conclusions

Building upon our previous research, the present study employed molecular docking to identify the 142P-242K mutant, which increased FOS conversion efficiency from 29 to 52%, representing an approximately 1.79-fold enhancement. Enzymatic characterization revealed that the optimal conditions for its catalytic activity are 45 °C, pH 6.0, and the presence of Na+. Molecular dynamics simulations and structural visualization analyses were performed to elucidate the mechanisms underlying this enhanced conversion rate. The findings indicated that the altered catalytic domain pocket and shifts in substrate affinity following the Lys mutation at position 242 were key factors in the FOS enhancement. The resulting proportions of GF2 and GF3 in the FOS product were 47 and 5%. The accumulation of GF2 resulted from the enhanced flexibility of the 142P-242K mutant, which facilitates easier substrate access to the active site. However, the exposure of hydrophobic active sites and the formation of strongly bound hydrogen bonds contributed significantly to the inhibition of GF2 conversion into GF3. Finally, this study utilized molasses, a byproduct of the sugar industry, as a substrate for FOS production. Among the various impurity removal strategies, the adsorption application yielded the optimum FOS content. FOS yield from sugar molasses treated by adsorption heating was 30.77%. This study provides a significant strategy for modifying the substrate specificity of FTase and facilitates the efficient, comprehensive utilization of industrial molasses.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/foods15050843/s1, Table S1: Strains and plasmids used in this experiment. Table S2: Primers used in this experiment.

Author Contributions

Conceptualization, Data curation, Formal analysis, Methodology and Resources, G.-L.C. and J.C.; Resources and Software, L.-Z.Z., B.L., and K.K.V.; Writing—original draft, G.-L.C. and J.C.; Writing—review and editing, F.-J.Z., K.K.V., and L.-F.Y.; Project management, Funding acquisition, Supervision, Validation and Visualization, G.-L.C., F.-J.Z., and L.-F.Y. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Guangxi Science and Technology Program (GK-AA22117015), the Earmarked Fund for China Agriculture Research System Guangxi Innovation Team—Specialty Fruits (nycytxgxcxtd-2024-17), the Opening Project of Guangxi Key Laboratory of Green Processing of Sugar Resources (GXTZYKF202211), and the Guangxi Academy of Agricultural Sciences Basic Research Business Project (GNK2026YT164).

Institutional Review Board Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article/Supplementary Materials. Further inquiries can be directed to the corresponding authors.

Acknowledgments

The authors would like to thank the Guangxi Subtropical Crops Research Institute and the Guangxi Academy of Agricultural Sciences, Nanning, Guangxi, China, for providing the necessary facilities for this study.

Conflicts of Interest

The authors declare no conflicts of interest.

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