Preparation of Metal-Hybridized Magnetic Nanocellulose for ω-Transaminase Immobilization

Abstract

The enzyme ω-transaminase (ω-TA) has garnered significant attention due to its capacity to catalyze the synthesis of chiral amines with high efficiency. Nevertheless, the lack of stability of ω-TA and the difficulty of recycling and reuse are still challenges that limit its application. This study developed a novel magnetic nanocellulose composite carrier (NNC@Fe₃O₄@Ni), synthesized from microcrystalline cellulose via low-eutectic solvent treatment, amine modification, and metal hybridization. The NNC@Fe₃O₄@Ni was characterized by FTIR, XPS, XRD, BET, and VSM. Additionally, the performance and catalytic behavior of the immobilized enzyme were investigated. The results revealed that NNC@Fe₃O₄@Ni exhibited a high specific surface area, superparamagnetism, and dual-site functionality (amine/Ni²⁺). Response Surface Methodology (RSM) optimized the carrier-enzyme interaction parameters, yielding optimal immobilization conditions: a mass ratio of 50.8 mg g⁻¹, temperature of 12.5 °C, and duration of 58.6 min, achieving 82.91% enzyme activity recovery. Compared to free enzymes, the immobilized variant demonstrated enhanced catalytic stability, with expanded optimal pH (9.0) and temperature (30 °C). Thermal stability assessments showed 84.39% activity retention after 5 h at 30 °C and 90.30% residual activity post-120 h storage. The catalyst maintained >80% efficiency over 10 reuse cycles. These findings confirm the efficacy of magnetic nanocellulose carriers in enhancing ω-TA stability, reusability, and catalytic performance, offering a viable strategy for industrial biocatalytic processes.

1. Introduction

In biocatalysis, ω-transaminase (ω-TA) uses inexpensive amines as donor substrates to catalyze the reductive amination of prochiral ketones, efficiently producing large quantities of chiral amines [1,2]. These amines are key intermediates in pharmaceutical and fine chemical production. As a green catalyst, ω-TA is widely used in chiral amine biosynthesis due to its environmental benefits and high catalytic efficiency [3]. However, the industrial use of free ω-TA is limited by its low stability and challenges in enzyme recovery and reuse. Therefore, improving the stability of ω-TA for industrial applications is a critical problem to be addressed.

To enhance the stability and catalytic efficiency of ω-TA, various techniques have been developed, including protein engineering [4], solvent engineering [5,6], and enzyme immobilization [7,8]. Although protein engineering and solvent engineering have improved enzyme stability and catalytic performance, they face limitations such as high costs, complex control requirements, low biocompatibility, and challenges in recycling, which significantly restrict their applications in biocatalysis [9]. Enzyme immobilization involves anchoring or confining the enzyme to a specific region while preserving its catalytic activity, enabling recovery and recycling. This technique not only increases enzyme recyclability but also markedly improves its resistance to denaturation and durability [10,11]. Koszelewski et al. demonstrated the reuse of ω-TA by immobilizing it in a sol-gel matrix, maintaining high enzyme activity [12]. Furthermore, immobilization could enhance catalytic efficiency and stability through chemical modification and directed evolution.

The effectiveness of enzyme immobilization depends on selecting a robust and efficient carrier with superior properties, as its physicochemical characteristics directly influence enzyme stability, activity retention, mass transfer efficiency, and reusability [13]. These factors determine the overall performance and practical utility of the immobilized enzyme. Typical immobilized carriers include carbon nanotubes, nanosilica, agarose, and magnetic nanoparticles [14,15]. Notably, magnetic nanoparticles have been shown to facilitate rapid enzyme recovery. This is due to their unique magnetic properties, which enable efficient termination of enzymatic reactions and facilitate reuse of the enzymes [16]. However, conventional carriers often exhibit limited active sites for immobilization and poor biocompatibility, resulting in low immobilization efficiency or disruption of the spatial structure of the enzyme, which significantly reduces catalytic activity. Therefore, the development of new immobilization carriers is a key challenge in the field of enzyme immobilization.

Cellulose, the most abundant renewable resource on Earth, is known for its low cost, biodegradability, non-toxicity, and environmental sustainability, as well as its excellent chemical modifiability and biocompatibility [17,18,19]. It is important to note that nanocellulose displays a high specific surface area and a multilevel pore structure. The highly crystalline structure of nanocellulose provides superior mechanical strength and thermal stability [20]. It has been demonstrated that enzyme molecules can be immobilized and conformationally stabilized on carrier surfaces through strategies such as physical adsorption and chemical cross-linking. Currently, cellulose is widely employed as an immobilization carrier for enzymes such as lipase [21], amylase [22], and catalase [23]. However, the binding of modified cellulose to enzymes via physical adsorption or covalent bonding is often characterized by a lack of selectivity and stability, resulting in issues such as enzyme detachment and reduced activity. Therefore, the development of novel immobilization carriers and specific binding methods tailored for transaminases remains a pressing challenge in this field.

In this study, microcrystalline cellulose was used to prepare carboxylated nanocellulose (CNC) by the carboxylation of deep eutectic solvent (DES), followed by amino-modification and metal hybridization techniques, to form NNC@Fe₃O₄@Ni composites with excellent magnetic properties and binding capacity. The structural properties were characterized using various characterization techniques. Magnetic nanocellulose served as a carrier, with ω-TA immobilized via the specific interaction between Ni and the His-tag of ω-TA. By optimizing immobilization conditions, the physicochemical and catalytic properties of the immobilized enzyme, including its activity, storage stability, and reusability, were thoroughly analyzed.

2. Results and Discussion

2.1. Structural and Characteristics of Nanocomposite

As illustrated in Figure 1a, this study utilized microcrystalline cellulose as raw material to prepare carboxylated nanocellulose (CNC) through a deep eutectic solvent (DES). Subsequently, polyethyleneimine was introduced for amination, yielding aminated nanocellulose (NNC). Metal hybridization was then employed to deposit Fe₃O₄ and NiO uniformly onto the NNC surface, resulting in the successful synthesis of NNC@Fe₃O₄@Ni composites. These composites exhibited excellent magnetic properties and enzyme immobilization capacity, enabling ω-TA immobilization via the specific interaction between Ni and the His-tag of ω-TA. Figure 1b shows the microstructure of microcrystalline cellulose (cellulose), characterized by short, rod-like structures with diameters of 20–50 µm and diverse morphologies. In contrast, Figure 1c shows CNC obtained after DES treatment, revealing a fibrous network with diameters less than 100 nm and a more uniform size distribution. This indicates that the DES disrupted cellulose crystallinity, producing finer and shorter fibers. Consequently, nanocellulose offers a larger specific surface area, providing more sites for enzyme immobilization.

The infrared (IR) spectra of microcrystalline cellulose, CNC, and NNC are presented in Figure 2a. A peak at 3400 cm⁻¹ corresponds to the O–H stretching vibration. CNC exhibits a distinct peak at 1730 cm⁻¹, confirming carboxyl group formation due to DES treatment. For NNC, the carboxyl peak is absent, replaced by an N–H peak at 1572 cm⁻¹, indicating successful amination.

X-ray photoelectron spectroscopy (XPS) spectra, shown in Figure 2b, reveal elemental changes during preparation. All samples display C1s and O1s peaks. NNC shows an N1s peak at 398.4 eV, mainly originating from the amino group in polyethyleneimine by peak analysis (Figure S1). The NNC@Fe₃O₄@Ni composites show Fe2p (710.5 eV) and Ni2p (854.5 eV) peaks for Fe₃O₄ and NiO, confirming their successful deposition, whereas Ni2p is mainly present as Ni²⁺ attributed to NiO (Figure S2).

X-ray diffraction (XRD) patterns of cellulose, NNC, and NNC@Fe₃O₄@Ni are shown in Figure 2c. Cellulose exhibits a prominent diffraction peak at 2θ = 22.6°, indicative of its crystalline structure [24]. NNC displays reduced crystallinity, reflecting DES-induced disruption of hydrogen bonds, which disperses cellulose into smaller units. The NNC@Fe₃O₄@Ni composite shows distinct metal oxide peaks, matching standard PDF cards for Fe₃O₄ and NiO, further confirming successful functionalization.

Thermal stability, closely tied to material composition, is shown in Figure 2d, which illustrates mass changes with temperature for cellulose, NNC, NNC@Fe₃O₄, and NNC@Fe₃O₄@Ni. Cellulose remains stable up to 300 °C but decomposes rapidly at around 350 °C, leaving minimal residual carbon (5.5%). NNC exhibits reduced thermal stability due to its high specific surface area and low crystallinity, making it more sensitive to decomposition. However, its higher residual carbon content (26.6%) compared to cellulose likely results from amino groups altering pyrolysis pathways and releasing flame-retardant gases [25]. NNC@Fe₃O₄ and NNC@Fe₃O₄@Ni show improved thermal stability and higher residual carbon rates (>35%), attributed to metal oxides forming a protective barrier. The higher stability of NNC@Fe₃O₄@Ni with an increased residual carbon rate of 40% compared to NNC@Fe₃O₄@Ni suggests that nickel oxide contributes to the formation of a denser and stronger structure. Thermogravimetric (TG) analysis confirms the structural integrity and stability of NNC@Fe₃O₄@Ni.

2.2. Specific Surface Area and Magnetic Properties of NNC@Fe₃O₄@Ni

Nitrogen adsorption–desorption isotherms for cellulose and NNC@Fe₃O₄@Ni, shown in Figure 3a, were used to assess specific surface area and pore structure. NNC@Fe₃O₄@Ni exhibits a significantly higher adsorption volume than cellulose, indicating a larger specific surface area and pore volume. A sharp increase in adsorption at relative pressure (P/P₀) near 1 confirms a type IV isotherm, characteristic of a mesoporous structure. The specific surface area of cellulose is 2.7 m²/g, while that of NNC@Fe₃O₄@Ni is 63.2 m²/g, suggesting enhanced immobilization capacity. Magnetic nanoparticles facilitate rapid enzyme recovery and reuse. In this study, Fe₃O₄ was integrated with nanocellulose to improve separation efficiency. Hysteresis loops for NNC@Fe₃O₄ and NNC@Fe₃O₄@Ni, presented in Figure 3b, display typical S-shaped curves. Although NNC@Fe₃O₄@Ni has slightly lower magnetization than NNC@Fe₃O₄, both exhibit zero coercivity and remanence, indicating excellent superparamagnetism.

2.3. Optimization of Immobilization Reaction Conditions

To determine the optimal conditions for immobilizing ω-TA onto NNC@Fe₃O₄@Ni, the effects of enzyme-to-carrier mass ratio, immobilization time, pH, and temperature were investigated. As shown in Figure 4a, the enzyme-to-carrier mass ratio significantly influenced the relative activity and loading capacity of AtATA@NNC@Fe₃O₄@Ni. Peak relative enzyme activity was achieved at a mass ratio of 50 mg/g, with an enzyme loading of 16.81 mg/g. Further increases in the mass ratio led to a gradual decline in relative activity. This decline likely results from the saturation of active sites on the carrier surface, where excessive enzyme aggregation blocks or damages active centers, thereby reducing activity. Similarly, Figure 4b illustrates the impact of immobilization time, with both relative activity and enzyme loading (28.65 mg/g) peaking at 50 min. Prolonged immobilization times caused a subsequent decrease in activity. Figure 4c shows that immobilization at pH 8.5 maximized relative activity and enzyme loading (30.46 mg/g). In addition, Figure 4d shows that an immobilization temperature of 10 °C yielded the highest relative activity and enzyme loading (41.67 mg/g).

To further evaluate the influence of enzyme-to-carrier mass ratio, immobilization time, and temperature on AtATA@NNC@Fe₃O₄@Ni activity recovery, response surface methodology was employed. A second-order polynomial model was developed to correlate these variables with enzyme activity, expressed as

Y = 81.18 + 1.60 A + 0.0832 B + 1.76 C + 1.02 AB + 0.6389 AC − 2.82 BC − 5.52 A² − 4.22 B² − 2.27 C²

where Y represents enzyme activity recovery, A is the enzyme-to-carrier mass ratio, B is immobilization time, and C is immobilization temperature.

The model’s coefficient of determination (R² = 0.9795) indicates strong predictability, effectively capturing the relationships among variables. A p-value below 0.0001 confirms the model’s statistical significance, with predictor variables exerting a notable impact on the response. A signal-to-noise ratio of 16.0878, exceeding 4, indicates robust model performance. Three-dimensional response surface and contour plots (Figure S3) visually represent these relationships. Optimal conditions, determined through rigorous experimentation, were an enzyme-to-carrier mass ratio of 50.8 mg/g, immobilization time of 58.6 min, and temperature of 12.5 °C, predicting an activity recovery of 81.74%. Parallel validation experiments confirmed an actual recovery of 82.91% for AtATA@NNC@Fe₃O₄@Ni.

2.4. Enzymatic Properties of AtATA@NNC@Fe₃O₄@Ni

The catalytic activity and stability of immobilized enzymes often differ significantly from those of free enzymes. Figure 5a shows the catalytic performance of AtATA and AtATA@NNC@Fe₃O₄@Ni at various temperatures, with maximum relative activity achieved at 30 °C for both. Thermal stability at 30 °C, shown in Figure 5b, reveals that after 5 h of incubation, the immobilized enzyme retained 84.39% of its activity, 10.52% higher than the free enzyme, indicating greater resilience to temperature changes. This increased stability is likely due to immobilization reducing the conformational flexibility of the enzyme, making it less susceptible to thermal denaturation [15].

The pH environment significantly influences catalytic performance, as extreme pH values can alter enzyme structure, leading to irreversible inactivation. As shown in Figure 6, both AtATA and AtATA@NNC@Fe₃O₄@Ni exhibited maximum activity at pH 9.0, with similar activity trends across pH values. Notably, the immobilized enzyme demonstrated superior stability in strongly acidic conditions, likely due to reduced aggregation compared to the free enzyme, which is prone to precipitation at low pH. After 5 h at pH 9.0, the immobilized enzyme retained 91.25% activity, 6.09% higher than the free enzyme. At pH 4.0 for 50 min, the immobilized enzyme maintained 29.83% activity, compared to only 14.79% for the free enzyme, highlighting its enhanced acid resistance.

2.5. Recycling Capacity of AtATA@NNC@Fe₃O₄@Ni

Figure 7a shows that during continuous 50 mL catalytic reactions, AtATA@NNC@Fe₃O₄@Ni maintained a 1-(R)-PEA conversion rate above 80% for the first seven batches. As reactions progressed, the conversion rate gradually declined. Figure 7b indicates that after 10 batches, residual enzyme activity was 80.30%. Enzyme loss during recovery, possibly due to incomplete magnetic separation or operational errors, probably contributed to this decline [26].

2.6. Storage Stability of AtATA@NNC@Fe₃O₄@Ni

To assess storage stability, AtATA and AtATA@NNC@Fe₃O₄@Ni were incubated at 4 °C for 120 h, with residual activity measured periodically (Figure 8). The immobilized enzyme consistently exhibited high relative activity, retaining 90.30% after 120 h, demonstrating excellent storage stability. In contrast, the free enzyme, starting at 100% activity, declined to 81.7% over the same period, indicating significantly lower stability. Immobilization on the NNC@Fe₃O₄@Ni carrier enhances stability by protecting the enzyme from environmental factors, such as temperature fluctuations or solution-mediated inactivation [27].

2.7. Kinetics of Enzymatic Reactions of ω-TA

The kinetic parameters for AtATA and AtATA@NNC@Fe₃O₄@Ni were determined using Lineweaver–Burk plots based on the Michaelis–Menten model. As shown in

Table 1. Kinetics of enzymatic reactions of AtATA and AtATA@NNC@Fe₃O₄@Ni.

Name	kcatpyruvate (min−1)	Kmpyruvate (mM)	kcat/Kmpyruvate (L·(min.mmol)−1)	kcat1-(R)-PEA (min−1)	Km1-(R)-PEA (mM)	kcat/Km1-(R)-PEA (L·(min.mmol)−1)
AtATA	176.68 ± 59.93	2.70 ± 1.28	65.44 ± 38.14	60.25 ± 0.95	0.31 ± 0.11	194.35 ± 69.03
AtATA@NNC@Fe3O4@Ni	182.20 ± 57.18	1.67 ± 0.78	109.10 ± 61.10	169.86 ± 4.87	0.76 ± 0.05	223.5 ± 16.07

, the immobilized enzyme exhibited enhanced catalytic efficiency for pyruvate and 1-(R)-PEA substrates, with a particularly pronounced increase for 1-(R)-PEA. Immobilization increased enzyme affinity for pyruvate (lower K_m) but reduced affinity for 1-(R)-PEA (higher K_m). These changes make the immobilized enzyme particularly suitable for 1-(R)-PEA catalysis, together with the improved stability.

3. Materials and Methods

3.1. Chemicals and Reagents

The microcrystalline cellulose was extra pure, with an average diameter of 20–50 μm, and was purchased from Sinopharm Chemical Reagent Corporation (Shanghai, China); The NaCl, yeast powder, tryptone, acetophenone, and polyethyleneimine (Mw 10000) were 99% pure; pyridoxal 5-phosphate, Na₂HPO₄, and NaH₂PO₄ were BC grade, and the purity of imidazole, kanamycin sulfate, and isopropyl-β-D-thiogalactopyranoside (Imidazole, kanamycin sulfate, and isopropyl-β-D-thiogalactopyranoside (IPTG)) were of molecular biology grade and purchased from Shanghai Bioengineering Co., Ltd. (Shanghai, China); the anhydrous ethanol, formic acid, choline chloride, oxalic acid, NiO (<30 nm), Fe₃O₄ (50 nm), HCl, and dimethylsulfoxide were of analytical purity and were purchased from Shanghai McLean Biochemistry and Technology Co. (Shanghai, China).

3.2. Preparation of Carboxylated Nanocellulose (CNC)

The 3 g of microcrystalline cellulose was mixed with 30 g of deep eutectic solvent (DES, choline chloride: oxalic acid = 1:3, stirred at 80 °C until transparent) and heated in an oil bath at 100 °C with stirring at 400 rpm for 2 h. The reaction mixture was then frozen at −20 °C for 24 h, thawed at room temperature, and washed twice with anhydrous ethanol and three times with ultrapure water until colorless. Carboxylated nanocellulose (CNC) was obtained after freeze-drying.

3.3. Preparation of Aminated Nanocellulose (NNC)

The 2 g of CNC was dispersed in 100 mL of water, mixed with 4 g of polyethyleneimine, and adjusted to pH 1.5 using diluted hydrochloric acid. The mixture was stirred at room temperature for 1.5 h, washed three times, and freeze-dried for 48 h to obtain aminated nanocellulose (NNC).

3.4. Preparation of NNC@Fe₃O₄@Ni

The 2 g of NNC was dispersed in 160 mL of 50% ethanol solution, combined with 0.20 g of Fe₃O₄ and 0.30 g of NiO, ultrasonicated for 15 min, and stirred at room temperature for 24 h. The product was ultrasonically cleaned three times and separated using an external magnetic field, yielding the magnetic nanocellulose composite (NNC@Fe₃O₄@Ni).

3.5. Characterization

The structure and properties of the materials were comprehensively characterized using multiple analytical techniques. Morphological features were examined via scanning electron microscopy (SEM, Model SU1510, Hitachi, Tokyo, Japan) and transmission electron microscopy (TEM, Model JEM-2100, JEOL, Tokyo, Japan). Infrared spectra were obtained using a Fourier transform infrared spectrometer (FTIR, Vertex 70, Molecular Devices, San Jose, CA, USA) over a scanning range of 4000–400 cm⁻¹. Elemental composition and chemical states were analyzed with an X-ray photoelectron spectroscopy analyzer (XPS, EscaLab 250Xi, Thermo Fisher, Waltham, MA, USA). Structural properties were assessed using X-ray diffraction (XRD, Model XRD-7000, Sailiwei, Suzhou, China), with samples prepared as XRD slices, scanned from 10° to 80° at 5°/min. Thermal stability was evaluated via thermogravimetric analysis (TGA, Model TG209F3, NETZSCH, Selb, Germany) using 10 mg samples, tested from 50 K to 800 K at a heating rate of 10 K/min under a nitrogen atmosphere. Specific surface area and pore structure were determined using a BET analyzer (Model ASAP2460, Micromeritics, Norcross, GA, USA). Magnetic properties were assessed with a vibrating sample magnetometer (VSM, Type 4HF, ADE, Chicago, IL, USA) at 300 K, with magnetic field strengths ranging from ±10,000 Oe, and corresponding magnetization curves were plotted.

3.6. Activity Assay of Immobilized Enzyme

Preparation and purification of ω-TA and methods for the determination of protein and acetophenone are provided in the Supplementary Materials. In a 1.5 mL centrifuge tube, equal amounts of free and immobilized ω-TA were combined with a reaction substrate containing 2.5 mM pyruvate, 1-(R)-phenylethylamine (1-(R)-PEA), 0.1 mM PLP, and 50 mM PBS. The mixture was incubated with shaking at 800 rpm and 25 °C for 3 min in a metal bath. Subsequently, the supernatant was separated using an external magnetic field, heated in a boiling water bath for 10 min, and analyzed for acetophenone content via high-performance liquid chromatography (HPLC). Relative enzyme activity was calculated from the obtained data, with the highest activity of free and immobilized enzymes defined as 100%. The recovery of immobilized enzyme activity was determined using the provided Equation (1) [26].

$$\mathrm{Relative} \mathrm{activity}(\%)=({\displaystyle \frac{A_i}{A_f}}) \text{×} 100$$

(1)

where A_i represents the enzyme activity of the immobilized enzyme (U); A_f represents the enzymatic activity of the free enzyme with an equivalent amount of the immobilized enzyme solidified (U).

3.7. Response Surface Methodology for Optimization of Immobilized ω-TA Activity

First, four parameters of the immobilization conditions were optimized: enzyme-to-carrier mass ratio (35, 40, 45, 50, and 55 mg/g), immobilization time (10, 20, 30, 40, 50, 60, 70, and 80 min), immobilization temperature (5, 10, 15, 20, 25, and 30 °C), and immobilization pH (8.0, 8.5, 9.0, 9.5, and 10.0). Then, a response surface methodology was employed to analyze three factors influencing the activity of immobilized ω-TA: enzyme-to-carrier mass ratio, immobilization time, and immobilization temperature. Three-dimensional (3D) response surface plots and contour plots were generated to visualize the enzyme’s response under optimized conditions. A three-variable Box–Behnken design was used to evaluate the effects of these parameters on enzyme activity recovery, with a total of 17 experiments conducted. Through multiple regression analysis, experimental data were compared with mathematical model predictions to determine the optimal immobilization conditions for ω-TA.

3.8. Enzymatic Properties of Immobilized ω-TA

3.8.1. Effect of Temperature on the Activity of Immobilized ω-TA

Enzyme activity was measured at temperatures ranging from 20 to 60 °C at pH 8.0 to identify the optimal temperature. Thermal stability of free ω-TA (AtATA) and immobilized ω-TA (AtATA@NNC@Fe₃O₄@Ni) was assessed by measuring residual activity after incubation at 30 °C for varying durations, with the highest activity in each group defined as 100%.

3.8.2. Effect of pH on Immobilized ω-TA Activity

To determine the optimal pH for AtATA and AtATA@NNC@Fe₃O₄@Ni, enzyme immobilization was performed at 25 °C in 50 mM of the following buffers: citrate–phosphate (pH 4.0–6.0), PBS (pH 6.0–8.0), Tris-HCl (pH 8.0–9.0), and Na₂CO₃-NaHCO₃ (pH 10.0–12.0). Enzyme activity was measured, and pH stability was evaluated by incubating the enzymes at 25 °C for different durations and determining residual activity, with the highest activity in each group set as 100%.

3.8.3. Evaluation of Recycling Ability of Immobilized Enzyme

A 50 mL continuous reaction was conducted with 5.0 g of AtATA@NNC@Fe₃O₄@Ni, 20 mM 1-(R)-PEA, 20 mM pyruvate, and 10% (v/v) dimethyl sulfoxide at 25 °C and 800 rpm. Each batch lasted 9 h. After completion, the enzyme was magnetically separated, washed twice with 50 mM PBS (pH 8.0), and resuspended in a fresh reaction mixture for the next batch.

3.8.4. Storage Stability Assessment of Immobilized Enzymes

To assess storage stability, AtATA and AtATA@NNC@Fe₃O₄@Ni were stored in 50 mM PBS (pH 8.0) at 4 °C for 120 h. Samples were periodically analyzed to determine residual activities, with the initial activity of both free and immobilized enzymes defined as 100%.

3.9. Kinetic Analysis of ω-TA Enzymatic Reactions

As a dual-substrate enzyme, the kinetic parameters of ω-TA for 1-(R)-PEA and pyruvate were determined separately. Substrate solutions (0–2.5 mM 1-(R)-PEA and 0–2.5 mM pyruvate) were prepared, and their effects on enzyme activity were analyzed via HPLC. The relationship between reaction rate (V) and substrate concentration (S) at low concentrations was modeled using the provided Equation (2).

$$V={\displaystyle \frac{V_{\mathrm{m}\mathrm{a}\mathrm{x}}·\left[S\right]}{(K_{\mathrm{m}}+\left[S\right])}}$$

(2)

Nonlinear fitting was applied to calculate the Michaelis constant (K_m), maximum reaction rate (V_max), turnover number (k_cat), and catalytic efficiency (k_cat/K_m).

4. Conclusions

This study successfully synthesized NNC@Fe₃O₄@Ni composites by depositing Fe₃O₄ and NiO onto nanocellulose, achieving excellent immobilization and superparamagnetic properties. The ω-TA was immobilized on NNC@Fe₃O₄@Ni through specific binding of Ni to the His-tag of the enzyme molecule. Optimal immobilization conditions, determined through single-factor and response surface experiments, were an enzyme-to-carrier mass ratio of 50.8 mg/g, immobilization time of 58.6 min, and temperature of 12.5 °C, which resulted in maximum enzyme activity. The immobilized enzyme, AtATA@NNC@Fe₃O₄@Ni, exhibited optimal activity at pH 9.0 and 30 °C, with significantly enhanced thermal and pH stability compared to the free enzyme. In a 50 mL system, it sustained acetophenone conversion over 10 batches, demonstrating robust reusability. Furthermore, it retained 90.30% of its initial activity after 120 h of storage at room temperature (25 °C), indicating superior storage stability.