Rational Engineering of Cellobiose 2-Epimerase Through Flexible Loop Modulation and Structure-Guided Sequence Alignment for Enhanced Lactulose Synthesis

Abstract

Lactulose, a valuable functional disaccharide with pharmaceutical and food applications, is efficiently synthesized via enzymatic isomerization of lactose. This study developed an integrated strategy combining protein engineering of cellobiose 2-epimerase (CsCE) from Caldicellulosiruptor saccharolyticus and process optimization to enhance lactulose production. A dual-track engineering approach—incorporating flexible loop modulation (residues 161–193) and structure-guided sequence alignment with N-acetyl-D-glucosamine-2-epimerase—enabled the creation of two superior mutants, R17Q/L184S and R17Q/S142T. The R17Q/L184S variant exhibited a 37% increase in crude enzyme activity, improved thermostability (half-life of 200 min at 80 °C), and enhanced substrate affinity (K_m reduced by 23.2%). R17Q/S142T achieved a 21% higher specific activity (24.08 U/mg), the highest among all variants. Structural and molecular dynamics analyses revealed that L184S enriched hydrogen bonding and hydrophobic interactions, improving structural rigidity, while S142T introduced allosteric regulation that facilitated catalytic efficiency. Under optimized conditions (70 °C, pH 7.5, 40% lactose, 20 U/mL enzyme, 3 h), lactulose yield reached 75.6% with >95% purity. This work demonstrates the successful application of synergistic enzyme engineering and process intensification for high-efficiency lactulose biosynthesis, providing viable candidates and system solutions for industrial-scale production.

1. Introduction

Lactulose is a unique functional oligosaccharide with dual applications in both pharmaceutical and food industries [1,2,3]. Due to its distinctive physiological properties, it has been widely utilized in medications, functional foods, dietary supplements, and animal feed [4,5,6]. Recognized as an important therapeutic agent for digestive disorders, lactulose also shows promising potential for expansion into metabolic disease management [7,8,9,10]. With over 70 years of clinical validation, lactulose has been approved as an over-the-counter (OTC) drug in more than 100 countries, demonstrating an excellent safety profile. Notably, even at high doses, there have been no reported cases of toxicity, carcinogenicity, or teratogenicity associated with its use. Lactulose was first discovered in heat-treated milk, which naturally contains high levels of lactose [11]. Under high-temperature conditions, lactose undergoes an isomerization reaction, converting its glucose moiety into a fructose residue, thereby forming lactulose. However, the natural occurrence of lactulose is extremely limited, necessitating industrial synthesis for large-scale production [12].

The global utilization of lactose resources presents a dual challenge: approximately 70% of the world’s population suffers from lactose intolerance, while the dairy industry generates around 2 million metric tons of lactose byproducts annually, with only about 25% being effectively utilized [13]. In this context, cellobiose 2-epimerase (EC 5.1.3.11) emerges as a promising biocatalyst. This enzyme specifically catalyzes the epimerization of D-glucose residues at the reducing end of β-1,4-glycosidic bond-linked oligosaccharides (such as cellobiose and lactose) to either D-mannose or D-fructose residues [14].

Compared to β-galactosidases, cellobiose 2-epimerase exhibits superior specificity, utilizing lactose as the sole substrate to produce lactulose through isomerization while generating only epilactose (4-O-β-D-galactopyranosyl-D-mannose) as a byproduct [15]. Notably, Kim et al. achieved a remarkable 58% conversion rate when employing this enzyme to synthesize lactulose from 700 g/L lactose at 80 °C—significantly higher than conventional chemical synthesis methods [16]. The conversion efficiency could be further enhanced to 88% through the addition of borate at a 1:1 molar ratio to lactose [17]. Over the past decade, cellobiose 2-epimerases from various microbial sources have been extensively characterized for their lactulose-producing capabilities, establishing them as next-generation industrial enzymes with exceptional potential for large-scale lactulose production [18].

Extensive studies have revealed that native enzymes frequently exhibit suboptimal performance in terms of stability, selectivity, and tolerance, often failing to meet the stringent requirements of industrial or laboratory applications [19,20,21]. Protein engineering has consequently emerged as a pivotal strategy to enhance specific enzymatic properties through targeted molecular modifications, representing a critical focus in contemporary enzyme technology development [22,23,24,25,26].

To address the inherent limitations of wild-type cellobiose 2-epimerase (CE) for industrial-scale applications, researchers have implemented comprehensive molecular optimization strategies aimed at improving three key parameters: catalytic activity, thermostability, and substrate specificity [18]. Notably, Feng and Wang et al. (2020–2021) conducted pioneering work by investigating a flexible loop region (residues 149–181) in CE, demonstrating that its amino acid composition critically influences both isomerization activity and reaction equilibrium, while playing a fundamental role in substrate recognition. Their structure-guided engineering yielded mutant enzymes with simultaneously enhanced isomerization activity and thermal stability [27,28].

Furthermore, comparative sequence analysis between CE and N-acetyl-D-glucosamine 2-epimerase (a key enzyme in the AGE family) has identified critical amino acid variations at conserved positions. These structural insights provide valuable guidance for rational design of CE variants with potentially improved catalytic efficiency through targeted site-directed mutagenesis.

This study focuses on the protein engineering of cellobiose 2-epimerase from Caldicellulosiruptor saccharolyticus (CsCE) and its application in lactulose biosynthesis. An innovative “dual-track” protein engineering strategy was developed. Employing local conformation engineering to systematically modify the key flexible loop region through site-directed mutagenesis and truncation. Implementing cross-family functional element transplantation based on sequence alignment with N-acetyl-D-glucosamine 2-epimerase. Using these two complementary approaches, several CsCE variants with significantly improved catalytic performance were successfully obtained, which provides a solid foundation for developing efficient enzymatic lactulose production processes.

2. Materials and Methods

2.1. Materials

The gene sequence encoding CsCE was codon-optimized for Escherichia coli expression system and synthesized by GenScript Biotech Corporation (Suzhou, China). The optimized gene was subsequently cloned into the pET-28a(+) expression vector using BamHI and XhoI restriction sites. Competent cells including E. coli DH5α, E. coli BL21(DE3), E. coli DMT and Fast Mutagenesis System were purchased from TransGen Biotech (Beijing, China). All chemical reagents used in this study were obtained from Macklin Biochemical Technology Co., Ltd. (Shanghai, China).

2.2. Rational Design Strategy for Flexible Loop Engineering

Structural studies by Feng et al. revealed that the Loop 149–181 in CE occupies a strategic position at the entrance of the catalytic pocket [28]. This functionally critical loop exhibits distinct structural characteristics: (1) the N-terminal segment maintains high sequence conservation with limited conformational flexibility, while (2) the C-terminal portion demonstrates considerable structural plasticity that enables dynamic movement toward the active center during catalysis, facilitating novel substrate interactions that promote isomerization efficiency. Building upon these structural insights, we developed a systematic three-tiered engineering strategy targeting the corresponding flexible loop (Loop 161–193) in CsCE. Constructed a series of deletion mutants to investigate loop length effects: Δ182–183 (2-aa deletion), Δ184–185 (2-aa deletion), and Δ180–183 (4-aa deletion). Performed virtual saturation mutagenesis focusing on three critical positions: R182 (electrostatic modulation): R182D/E (charge reversal mutants); F183 (aromatic interactions): F183E/K/M (polarity-engineered variants); L184 (hydrophobic core): L184S/P (flexibility/rigidity mutants). Designed distal regulatory mutants based on dynamic correlation analysis: S185P (rigidity enhancement), A191S (hydrogen bonding potential), K193R (conserved positive charge maintenance).

2.3. Sequence Alignment-Based Molecular Engineering Strategy

Through NCBI BLAST (version 2.6.0, https://blast.ncbi.nlm.nih.gov/Blast.cgi (accessed on 8 July 2025)) screening, we identified four N-acetyl-D-glucosamine-2-epimerase (AGE) sequences with highest homology to CsCE. Comprehensive sequence alignment and UniProt conserved domain analysis (Figure 1) revealed six key variable regions (41–42, 90–94, 139–145, 197–201, 260–268, and Δ flexible loop). Initial screening of combinatorial mutants demonstrated that modifications in regions 90–94 and 139–145 exhibited promising functional improvements. Subsequent precision single-point mutagenesis included: F90D (hydrophobic-to-hydrophilic substitution enlarging substrate channel), A92G (reducing steric hindrance for improved substrate binding), E94A (charge neutralization optimizing local electrostatic environment), Y139H/E140R (salt bridge formation enhancing thermostability), S142T (hydrogen bond reinforcement decreasing catalytic center RMSF), and D144S/E145P (reducing binding free energy).

Figure 1.

Sequences alignment analysis of CsCE and AGEs. (HcAGE GeneBank: WP_066094377.1, RhAGE GenBank: MBC7707085.1, PbAGE GenBank: HXI59021.1, BbAGE GenBank: MBC7380411.1).

2.4. Mutant Construction Using Fast Mutagenesis System

The site-directed mutagenesis was performed using methylated pET28a(+)/CsCE plasmid as template with specifically designed overlapping primers (Table S1) containing desired mutation sites. PCR amplification was carried out in 20 μL reaction volumes using a Fast Mutagenesis System under optimized conditions: initial denaturation at 95 °C for 3 min followed by 30 cycles of denaturation (95 °C, 20 s), annealing (56 °C, 20 s) and extension (72 °C, 3 min), with final extension at 72 °C for 3.5 min. The PCR products were then treated with DMT enzyme—an engineered DpnI restriction enzyme—at 37 °C for 1.5 h to selectively digest methylated template plasmids before transforming into specialized DMT-competent E. coli cells capable of degrading residual methylated DNA. All mutant constructs were sequence-verified by Tsingke Biotechnology Co., Ltd. (Beijing, China) to confirm the introduced mutations.

2.5. Enzyme Expression and Activity Assay

The verified mutant plasmids were transformed into E. coli BL21(DE3) competent cells for protein expression. Single colonies were selected and pre-cultured in 50 mL LB medium supplemented with 50 μg/mL kanamycin at 37 °C overnight. The starter cultures (2% inoculum) were transferred into 1 L of optimized auto-induction medium containing: 10 g/L tryptone, 10 g/L KNO₃, 17.105 g/L Na₂HPO₄·12H₂O, 3 g/L KH₂PO₄, 1 g/L NH₄Cl, 0.1 mM MgSO₄·7H₂O, 5 g/L glycerol, and 5 g/L glucose. Cells were grown at 37 °C with shaking (220 rpm) until reaching OD600 of 1.8–2.2, followed by induction with 0.1 mM IPTG and continued cultivation at 25 °C for 18 h.

Cells were harvested by centrifugation (8000× g, 15 min, 4 °C) and resuspended in lysis buffer (20 mM acetic acid, 10 mM calcium acetate, pH 5.4). Cell disruption was performed by sonication (37 kHz, 10 min, pulse mode). The crude extract was heat-treated at 70 °C for 20 min to denature host proteins (Figure S1), immediately cooled on ice for 10 min, and clarified by centrifugation (12,000× g, 10 min). The supernatant containing the thermostable CsCE was collected for activity assays.

Enzyme activity was determined using lactose as substrate in a 3 mL reaction system containing: 0.3 g lactose, whose final concentration is 0.29 M (10% w/v final concentration), 1 mL appropriately diluted enzyme solution, and 10 mM KH₂PO₄-NaOH buffer (pH 7.5). Reactions were conducted at 80 °C for 20 min and terminated by adding 100 μL concentrated HCl. After centrifugation (8000× g, 5 min), 200 μL of the reaction mixture was mixed with 800 μL resorcinol-HCl reagent and heated at 100 °C for 7 min. The solution was cooled with 5 mL ice-cold water, and the absorbance at 480 nm was measured to quantify lactulose production.

One unit of enzyme activity (U) was defined as the amount of enzyme required to produce 1 μmol lactulose per minute under standard conditions (80 °C, pH 7.5, 10% lactose, that is 0.29 M lactulose). Protein concentration was determined by Bradford assay for specific activity calculation.

2.6. Quantitative Analysis and Optimized Purification of Lactulose

Lactulose quantification was performed by HPLC using an aminopropylsilane-bonded silica column with isocratic elution (acetonitrile:10 mM sodium phosphate buffer, pH 4.5 [84:16, v/v]) at 1.5 mL/min flow rate. The refractive index detector and column were maintained at 40 °C, with 20 μL injection volume. Samples were filtered through 0.22 μm nylon membranes and sonicated for 5 min prior to analysis.

The lactulose purification process was initiated by heat-inactivating the 50 mL enzymatic reaction mixture followed by centrifugation (8000× g, 15 min, 4 °C) to remove denatured enzymes. The supernatant was treated with 1% (w/v) activated carbon at 50 °C for 60 min, with subsequent removal of carbon particulates through sequential centrifugation and 0.22 μm membrane filtration. The clarified solution was concentrated to approximately 5 mL by rotary evaporation, then subjected to ethanol-assisted crystallization at 80 °C with continuous stirring and gradual addition of anhydrous ethanol until crystal formation, followed by overnight crystallization at 4 °C. For enhanced purity, the crystallized product was further purified using m-aminophenylboronic acid affinity chromatography, where the resin was pre-activated and loaded with the lactulose solution in alkaline conditions, washed with alkaline buffer to remove impurities, and finally eluted with 0.1 M HCl. This comprehensive protocol achieved >85% recovery yield and >95% purity, as verified by HPLC analysis, while maintaining scalability for potential industrial applications.

2.7. ¹H NMR Structural Characterization of Lactulose

For structural verification, purified sample (20 mg) was dissolved in 600 μL D₂O. ¹H NMR spectra were acquired at 300 K using a 600 MHz NMR spectrometer (Bruker Avance III HD) (Agilent Technologie Inc., Santa Clara, CA, USA) equipped with a TCI cryoprobe, employing 128 scans with a 16 ppm spectral width, 3.0 s acquisition time, and 2.0 s relaxation delay. As shown in Figure S2, chemical shifts (δ) were referenced to internal TMS (0 ppm) and revealed characteristic resonances:

¹H NMR (600 MHz, Deuterium Oxide) δ 4.40 (d, J = 7.8 Hz, 1H), 4.31 (d, J = 7.9 Hz, 0H), 4.18–4.08 (m, 1H), 4.07–4.02 (m, 1H), 3.97 (dd, J = 10.0, 3.3 Hz, 1H), 3.86 (dd, J = 13.0, 1.4 Hz, 1H), 3.80–3.74 (m, 3H), 3.68–3.49 (m, 10H), 3.47–3.44 (m, 1H), 3.44–3.42 (m, 1H), 3.42–3.40 (m, 1H).

3. Results and Discussion

3.1. Expression and Activity Assay of Mutant Enzymes

A comprehensive analysis of enzymatic activity and specific activity was conducted for the flexible loop mutants (Figure 2A). Compared to the parental R17Q strain (crude enzyme activity set as 100%), several mutants exhibited moderate improvements in catalytic performance: R182E (112%), F183E (110%), F183K (114%), F183M (115%), L184S (137%), A191S (110%), and Δ184–185 (109%), with L184S demonstrating the most substantial improvement. R182D (63%) showed the most significant decrease. The specific activity analysis after thermal purification revealed varying degrees of reduction across all mutants. The lowest value was observed in Δ184–185 (9.14 U/mg), while L184S maintained the highest specific activity (18.45 U/mg), representing only a minor decrease of approximately 1.5 U/mg compared to R17Q. Notably, L184S exhibited excellent reproducibility across five consecutive fermentation batches, consistently maintaining high levels of both crude enzyme activity and specific activity. Therefore, the double mutant R17Q/L184S, obtained through this flexible loop engineering strategy, combines high enzymatic activity with favorable specific activity, emerging as a promising candidate for industrial-scale applications.

Figure 2.

Analysis of enzyme activity and specific activity for (A) flexible loop mutants and (B) sequences alignment analysis mutants.

A comprehensive functional analysis was performed on mutants generated through sequence alignment (Figure 2B), with R17Q crude enzyme activity normalized to 100%. The results revealed distinct functional categories: Mutations in regions 41–42, 197–201, 260–268, and Δ flexible loop resulted in complete loss of enzymatic activity, indicating their essential roles in maintaining the catalytic core structure. The 90–94 region mutants exhibited moderate activity enhancement (up to 112%). The 139–145 region mutants showed variable effects, with relative activities ranging down to 78%. It is worth noting that F90D demonstrated the highest activity improvement in the 90–94 region (112%). S142T displayed exceptional specific activity (24.08 U/mg) with 103% relative activity, representing a 21% increase over R17Q and the highest specific activity observed among all characterized variants.

Through integrated molecular engineering strategies combining flexible loop optimization and structure-guided sequence alignment, we have successfully identified two superior mutants: R17Q/L184S and R17Q/S142T. Future investigations will systematically characterize the enzymatic properties of these optimized variants—including thermostability, substrate specificity, and kinetic parameters—to comprehensively evaluate their catalytic performance and application potential.

3.2. Enzymatic Characterization of Mutant Enzymes

The catalytic activities of the two mutant enzymes, R17Q/L184S and R17Q/S142T, were evaluated across a temperature gradient of 60–90 °C (Figure 3A). All variants exhibited similar parabolic activity-temperature profiles within this range. Notably, neither of the newly introduced mutations altered the optimal catalytic temperature, which remained at 80 °C. This observation suggests that the thermal stability of the core catalytic domain remains unaffected by these surface mutations, and that the modified residues are not involved in conformational rearrangements within temperature-sensitive regions. In the moderate temperature range (70–80 °C), R17Q/L184S demonstrated superior catalytic performance, maintaining >83% relative activity. Both R17Q and R17Q/S142T also retained substantial activity (>75%) in this range. However, under high-temperature conditions, R17Q/L184S showed reduced thermotolerance, with residual activity dropping to 36% at 90 °C. In contrast, R17Q and R17Q/S142T exhibited better thermal resilience, retaining approximately 50% activity at the same extreme temperature.

Figure 3.

Comparison of (A) optimal temperature characteristics, (B) the effect of pH on enzyme activity, and (C) temperature stability for double mutants R17Q/L184S and R17Q/S142T.

The enzymatic activities of R17Q/L184S and R17Q/S142T were assessed across a broad pH range (3.0–12.0). As illustrated in Figure 3B, both double mutants retained the same optimal pH (7.5) as the parental R17Q. Notably, under alkaline conditions (pH 9–11), R17Q/L184S and R17Q/S142T exhibited significantly higher residual activity compared to R17Q, suggesting that the L184S and S142T mutations may enhance structural stability through improved surface charge distribution. In contrast, all enzyme variants underwent rapid inactivation under acidic conditions (pH < 6.0). These findings indicate that while the optimal pH remains unchanged, the introduced mutations confer enhanced alkaline stability without compromising the intrinsic catalytic preference.

Real-time monitoring of enzymatic activity decay (Figure 3C) revealed distinct thermal stability profiles for the double mutants. The R17Q/L184S variant exhibited enhanced thermostability with a half-life of 200 min at 80 °C, representing a significant improvement over R17Q (180 min). In contrast, R17Q/S142T showed reduced thermal resistance, displaying a half-life of only 140 min at 80 °C—substantially shorter than that of R17Q. These findings highlight the complementary application potential of the two mutants: R17Q/L184S is better suited for continuous processes at elevated temperatures (70–80 °C), while R17Q/S142T, with its superior specific activity, offers distinct advantages for moderate-temperature applications (<70 °C). This differentiation provides a rational basis for selecting appropriate biocatalysts according to specific process requirements.

Enzyme kinetic studies (Table 1) revealed that the turnover numbers (kcat) of both double mutants, R17Q/L184S and R17Q/S142T, decreased by 9–22%, potentially due to altered transition-state stabilization. Notably, R17Q/L184S exhibited a reduced Michaelis constant (K_m = 195.63 mM), indicating a 23.2% enhancement in substrate affinity, whereas R17Q/S142T showed no significant change in K_m. This suggests that the L184S mutation may induce subtle conformational adjustments in the catalytic pocket entrance. Although the catalytic turnover decreased slightly, both mutants maintained high catalytic efficiency (k_cat/K_m = 0.048–0.069 mM⁻¹·min⁻¹). Obviously, the key outcome of the optimized hydrogen bond network is a balanced improvement in binding (Km) with minimal impact on catalytic rate (kcat), leading to a slightly more efficient enzyme. Combined with their distinct thermal stability profiles—R17Q/L184S demonstrated an extended half-life (200 min) at 80 °C compared to R17Q/S142T (140 min)—these kinetic properties enable precise enzyme selection for different process requirements: R17Q/L184S is preferable for continuous high-temperature (70–80 °C) operations, while R17Q/S142T offers advantages in moderate-temperature applications due to its superior specific activity.

Table 1.

Comparison of enzyme kinetic constants for double mutants R17Q/L184S and R17Q/S142T.

Mutant	Km (mM)	kcat (min−1)	kcat/Km (mM−1·min−1)
R17Q	241.00 ± 4.10	14.96 ± 0.25	0.062 ± 0.006
R17Q/L184S	195.63 ± 4.24	13.50 ± 0.29	0.069 ± 0.007
R17Q/S142T	241.26 ± 4.44	11.57 ± 0.21	0.048 ± 0.005

3.3. Structural Modeling and Molecular Dynamics Analysis of Mutants

The three-dimensional structures of the double mutants R17Q/L184S and R17Q/S142T were initially predicted using AlphaFold2 (https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb (accessed on 8 July 2025)), revealing that both mutants maintained a highly consistent overall structural framework with the parental R17Q protein, without significant conformational alterations. Subsequent analysis using Protein Tools (https://www.proteintools.uni-bayreuth.de/ (accessed on 8 July 2025)) for hydrogen bonding networks and salt bridge interactions demonstrated distinct molecular interaction patterns: the R17Q/L184S mutant exhibited a substantial enhancement in both salt bridge formation and hydrogen bonding networks, whereas the R17Q/S142T mutant primarily showed localized increases in hydrogen bonding while maintaining stable salt bridge interactions. These structural modifications, particularly the reinforced intermolecular interaction network in R17Q/L184S, are likely to critically influence enzymatic stability and catalytic properties.

Visualization of the hydrogen bond network surrounding the critical flexible loop (residues 161–193, Figure 4) revealed its essential role in modulating substrate recognition and reaction equilibrium, thereby significantly influencing catalytic efficiency. Specifically, the R17Q mutant formed only a limited set of hydrogen bonds localized within the loop region. It is noteworthy that the R17M mutant does not exhibit a more extensive hydrogen bond network in the same region. It can be inferred that the R17M mutation fails to enhance the hydrogen bond network within the flexible loop region at the entrance of the active site, which is consistent with the lack of improvement in its activity. In contrast, the R17Q/L184S double mutant not only established an extensive and intricate hydrogen bond network within the loop itself but also formed additional stabilizing interactions in distal regions near the active site entrance. This optimized hydrogen bond architecture enhances catalytic performance through several mechanisms: (1) facilitating optimal substrate orientation for entry into the active pocket, (2) strengthening binding affinity in the enzyme-substrate complex, and (3) stabilizing catalytic transition states to promote reaction progression. Consequently, exhibits marginally enhanced catalytic efficiency and reaction rates compared to R17Q. These structural advantages highlight the crucial regulatory role of flexible loop hydrogen bonding in enhancing catalytic efficiency, positioning R17Q/L184S as a superior biocatalyst for industrial applications.

Figure 4.

Analysis of hydrogen bond network in flexible loop for active pocket entrance ((A): R17Q, (B): R17Q/L184S, (C): R17M).

Simulated analysis of hydrophobic clusters using Protein Tools (Figure 5) revealed that the R17Q/L184S double mutant exhibits a more optimized hydrophobic core architecture compared to the R17Q single mutant. Quantitatively, R17Q/L184S contained 15 distinct hydrophobic clusters—a 25% increase over the 12 clusters identified in R17Q. Spatially, the four additional clusters were strategically positioned in key regions of the protein surface and around the active site pocket. These structural modifications are consistent with enhanced conformational stability, aligning with established literature demonstrating a positive correlation between hydrophobic core integrity and protein stability [29,30]. The optimized hydrophobic packing in R17Q/L184S likely contributes to its improved thermal resistance and catalytic robustness observed in previous experiments.

Figure 5.

Distribution characteristics of hydrophobic core for mutant proteins ((A): R17Q, (B): R17Q/L184S).

Structural-functional analysis of R17Q/S142T revealed no significant differences in hydrogen bond numbers within Loop 161–193 and the active pocket compared to R17Q, while hydrophobic interaction analysis showed essentially identical cluster counts. Detailed structural examination identified that the S142T substitution occurs at the junction between two critical α-helices. Further investigation demonstrated that the newly introduced threonine side chain enhances catalytic performance through three interconnected mechanisms: increased local steric hindrance, induced alterations in α-helical dihedral angles, and consequent optimization of the active site microenvironment. Although these structural modifications do not directly affect substrate recognition, they collectively improve specific activity by 21% through long-range effects, exemplifying the classic allosteric regulation paradigm of “distal mutation → local conformational adjustment → functional modulation” in enzyme systems.

Molecular dynamics simulations performed using Discovery Studio 2021 (Figure 6) demonstrated distinct dynamic properties among the mutants. Root-mean-square deviation (RMSD) analysis indicated that R17Q/S142T exhibited the largest overall backbone fluctuations, while R17Q/L184S maintained relatively lower fluctuations during the early simulation phase, though its mobility increased in later stages. Complementary root-mean-square fluctuation (RMSF) profiles revealed reduced flexibility in the 75–175 residue region of R17Q/L184S, consistent with the previously observed denser hydrogen-bonding network in this area. Conversely, R17Q/S142T displayed generally elevated RMSF values, particularly within the 50–200 residue region encompassing key catalytic residues, where enhanced flexibility was observed. This increased structural mobility correlates well with its experimentally determined decrease in thermal stability alongside improved specific activity. These computational findings provide mechanistic insights into how different mutation sites differentially modulate protein dynamics, ultimately governing distinct functional outcomes.

Figure 6.

Comparison of molecular dynamics characteristics for double mutants R17Q/L184S and R17Q/S142T ((A): RMSD, (B): RMSF).

3.4. Optimization of Lactulose Synthesis Process Conditions

Enzymatic reactions require an optimal temperature that balances high reaction rates with sustained enzyme activity. Building on the thermostability profile of the R17Q/L184S mutant (200 min half-life at 80 °C) established in Section 3.2, this study systematically evaluated the competitive effects of temperature on both the enzymatic reaction and Maillard side reactions.

As shown in Figure 7A, lactulose yield was highest at 70 °C (set as 100% baseline). The yield remained substantial at 65 °C (92.1%) and 85 °C (93.5%), indicating high catalytic efficiency across the 65–85 °C range. However, temperatures exceeding 70 °C led to a marked increase in solution browning and accumulation of Maillard byproducts, which suppressed enzymatic activity. Therefore, 70 °C was selected as the optimal temperature to maximize reaction rate while minimizing competitive side reactions.

Figure 7.

Optimization analysis of reaction for lactulose synthesis. (A) Temperature (20% substrate concentration, 15 U/mL enzyme dosage, 6 h reaction time); (B) substrate concentration (70 °C temperature, 15 U/mL enzyme dosage, 6 h reaction time); (C) enzyme dosage (40% substrate concentration, 70 °C temperature, 6 h reaction time); (D) reaction course (40% substrate concentration, 70 °C temperature, 20 U/mL enzyme dosage).

Substrate concentration plays a dual role in governing enzymatic reaction efficiency. At low concentrations, enzymes operate below saturation, leading to diffusion-limited kinetics and a thermodynamic equilibrium favoring reverse hydrolysis. Conversely, excessively high substrate levels—despite saturating active sites—can induce non-productive complex formation, reduce effective molecular collisions, and ultimately diminish both reaction rate and yield, while also raising material costs. Therefore, identifying a balanced substrate concentration is essential to maintain high catalytic rates, achieve elevated conversion within a practical timeframe, and maximize overall production efficiency.

The substrate lactose exhibits limited aqueous solubility that increases non-linearly with temperature, necessitating elevated temperatures to achieve complete dissolution at high concentrations. However, excessive concentrations of both lactose and enzyme can accelerate Maillard reactions, compromising both yield and product quality. To address these competing factors, we systematically evaluated concentration effects (Figure 7B). Maximum lactulose yield (set as 100%) was achieved 1.17 M (40%) lactose, where substrate dissolution equilibrium and reaction kinetics reached an optimal balance. Yields increased progressively within the 10–40% concentration range, but declined and became unstable beyond 50%, where phase separation occurred and excessive substrate impeded complete enzymatic conversion. Concurrently, Maillard reaction intensification led to noticeable browning at these higher concentrations. Therefore, 1.17 M (40%) lactose was identified as the optimal concentration, providing maximal and reproducible lactulose yield while maintaining practical dissolution characteristics and minimizing substrate waste.

Enzyme concentration is a critical parameter in enzymatic processes, significantly influencing both reaction rate and final yield. Insufficient enzyme levels lead to slow kinetics and prolonged reaction times, whereas excessive dosage increases production costs, wastes enzyme resources, and promotes undesirable Maillard side reactions. Thus, identifying an optimal enzyme concentration is essential for economically viable operation.

Systematic evaluation of enzyme dosage (Figure 7C) revealed a characteristic pattern of diminishing returns. With the yield at 20 U/mL set as 100%, increasing enzyme concentration consistently enhanced lactulose production, but with progressively smaller gains. When enzyme levels were low (5–10 U/mL), yield increased linearly (13.6% improvement from 5 to 10 U/mL), indicating unsaturated enzyme activity. As concentration rose to 15–20 U/mL, the growth rate slowed, reaching maximum yield at 20 U/mL with acceptable Maillard byproduct levels. Further increase to 25 U/mL provided negligible yield improvement (1.2%) but raised enzyme costs by 25% and intensified side reactions. Based on this cost–benefit analysis, 20 U/mL was selected as the optimal enzyme dosage, balancing high lactulose yield with operational economy. This parameter offers valuable guidance for industrial-scale implementation.

Systematic sampling and analysis of reaction components at different time intervals enabled precise monitoring of reaction progression and determination of the optimal termination point. As shown in Figure 7D, the reaction exhibited rapid product accumulation during the initial 0–3 h phase (production rate: 25.2%/h), followed by a plateau thereafter. Under optimized conditions (70 °C, pH 7.5, 40% substrate concentration, 20 U/mL enzyme dosage), lactulose yield reached its maximum at 3 h, achieving 75.6% substrate conversion with only 4.5% epilactose as byproduct. This time point represents the ideal balance between conversion efficiency and product specificity.

4. Conclusions

This study successfully established an integrated framework for efficient lactulose biosynthesis through synergistic protein engineering and process optimization. Structure-guided engineering of cellobiose 2-epimerase yielded two optimized variants, R17Q/L184S and R17Q/S142T, with complementary catalytic properties. R17Q/L184S demonstrated enhanced thermostability and substrate affinity, making it ideal for continuous high-temperature processes, while R17Q/S142T exhibited superior specific activity for moderate-temperature applications. Systematic optimization of reaction parameters achieved 75.6% lactulose yield with high purity, confirming the industrial feasibility of the developed process. These findings not only provide efficient biocatalysts for lactulose production but also illustrate a rational strategy for enzyme engineering that balances activity, stability, and efficiency—offering broad implications for the sustainable manufacturing of functional carbohydrates.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/biom16020206/s1, Table S1: Primer sequences of mutants; Figure S1: SDS-PAGE electrophoresis analysis of thermal purification process; Figure S2: ¹H NMR spectrum of product lactulose.

Author Contributions

Conceptualization, C.H. and J.Y.; methodology, C.H.; software, X.M.; validation, X.M.; formal analysis, X.M. and C.M.; investigation, C.H. and C.M.; resources, X.M.; data curation, C.H.; writing—original draft preparation, J.Y.; writing—review and editing, X.M. and J.Y.; visualization, X.M.; supervision, H.Z., X.H. and J.Y.; project administration, H.Z. and J.Y.; funding acquisition, H.Z. and J.Y. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data is contained within the article or Supplementary Material.

Conflicts of Interest

The authors declare no conflicts of interest.

Funding Statement

This research was supported by the National Natural Science Foundation of China (Grant No. 82473833), the Natural Science Foundation of Anhui Province (Grant No. 2408085MC051), the Key Science & Technology Project of Anhui Province (Grant No. 202423l10050064), and the Fundamental Research Funds for the Central Universities.