Biomacromolecule-Regulated Biomimetic Mineralization for Efficiently Immobilizing Cells to Enhance Thermal Stability

Abstract

The industrial application of free sucrose phosphorylase (SPase) is significantly limited due to cost, stability issues, and poor reusability. In this study, we employed organic–inorganic hybrid nanoflowers to achieve cell immobilization by co-assembling metal ions with cells. The surface of cells was coated with nanoflowers via chitosan-regulated biomimetic mineralization, thereby enhancing the activity of immobilized cells while providing a protective structure to improve stability. The relative activity of the immobilized cells was 30% higher than that of the free cells. After placing at 4 °C in 15 days, the relative activity of immobilized cells (80%) was substantially higher than that of free cells (40%). Moreover, the immobilized cells retained approximately 85% of their relative activity after 10 cycles. In summary, the novel biocatalysts developed in this study combine high catalytic performance with excellent reusability, demonstrating significant advantages in E. coli cell immobilization and providing a solid foundation for their application in industrial biocatalysis and related fields.

1. Introduction

Sucrose phosphorylase (SPase, EC 2.4.1.7), a member of glycoside hydrolase family 13 (GH13), is a versatile biocatalyst that catalyzes the reversible phosphorolysis of sucrose to yield glucose-1-phosphate and fructose [1]. Beyond this canonical activity, SPase catalyzes transglycosylation reactions using various monosaccharides, sugar alcohols, and phenolic or alcoholic compounds as acceptors. This versatility facilitates the production of high-value functional glycosides, exemplified by glucosyl glycerol (GG) [2] and 2-O-α-D-glucopyranosyl-L-ascorbic acid (AA-2G) [3]. Owing to its high regioselectivity, broad substrate range, and environmentally benign reaction conditions, SPase has attracted considerable attention for applications in cosmetics, food, and pharmaceutical industries [4]. However, free SPase suffers from poor thermostability, narrow pH tolerance, and low reusability, limiting its industrial application [5].

Traditional enzyme immobilization techniques, such as covalent binding [6] and cross-linking [7], could partially enhance enzyme stability but often lead to activity loss and involve complex procedures. In contrast, whole-cell immobilization presents notable advantages. This technique confines entire cells (engineered E. coil cells expressing SPase) within a defined space, creating reusable biocatalysts [8]. The primary goal is to preserve cellular physiological function or enzymatic activity, employing methods like adsorption and entrapment [9]. Immobilized cells circumvent the need for complex enzyme purification steps, thereby reducing activity loss and cost, while better maintaining the enzyme’s native conformation and stability [10]. Particularly for enzymes like SPase, which might require the high concentration of substrates in the industrial biosynthesis, immobilized cells could significantly enhance industrial operational stability, catalytic efficiency, and reusability. Consequently, developing efficient whole-cell immobilization strategies is crucial for advancing the practical application of SPase.

Magnetic nanoparticles (MNPs), including Fe₃O₄, Fe, Co, and Ni, have attracted considerable interest as immobilization carriers owing to their superparamagnetism, biocompatibility, and ease of magnetic recovery [11]. Fe₃O₄ nanoparticles, in particular, are widely employed because they can be readily functionalized with biomacromolecules, such as chitosan, and polyethyleneimine (PEI) groups to improve enzyme affinity [12,13,14]. However, MNPs alone provide limited structural protection, leaving covalently immobilized enzymes directly exposed to the reaction environment and thereby compromising their stability and reusability. In contrast, biomacromolecules alone offer excellent biocompatibility but generally lack mechanical rigidity and structural durability, resulting in poor long-term performance. These intrinsic limitations underscore the need for hybrid immobilization platforms that combine robustness with favorable biochemical compatibility.

In recent years, organic–inorganic hybrid nanoflowers have provided a novel biomimetic platform for enzyme and whole-cell immobilization [15]. This approach relies on the self-assembly of biomolecules and metal ions under mild aqueous conditions to form hierarchical flower-like architectures. Biomacromolecules, including proteins, chitosan, and peptides, regulate crystal nucleation and growth while preserving enzyme conformation and improving biocompatibility [16]. Extending this strategy to whole-cell immobilization enables nanoflowers to uniformly cover the E. coli surface, forming a protective matrix. Chitosan-regulated biomimetic mineralization provides a simple, low-cost knob to control nanoflower formation and cell loading, yielding a protective, flower-like scaffold that improves catalytic stability and recyclability for whole-cell biocatalysis. Nevertheless, hybrid nanoflowers have been extensively developed for enzyme immobilization, and their use for immobilizing cells remains limited.

In this study, we developed a biomolecule-mediated biomimetic mineralization strategy to immobilize host cells expressing SPase in hybrid nanoflowers. This system offers multiple advantages, including high surface area for enhanced enzyme loading, mild mineralization preserving enzyme activity, improved thermal and pH stability, and outstanding reusability. Immobilized cells were applied in the scale biosynthesis of AA-2G (Figure 1). The resulting hybrid catalyst exhibited superior performance to free SPase, providing a promising foundation for sustainable, large-scale production of AA-2G.

2. Results and Discussion

2.1. Screening of Biological Macromolecules and Inorganic Metal Ions

Biological macromolecules and inorganic metal ions represent critical constituents in the construction of organic–inorganic hybrid nanoflowers [13]. Because most enzymes exhibit high sensitivity toward their surrounding physicochemical environment, the influence of five representative metal ions (Co²⁺, Cu²⁺, Fe²⁺, Ca²⁺, and Fe³⁺) on intracellular enzymatic activity was systematically evaluated (Figure 2A). As shown in Figure 2A, Cu²⁺ markedly inhibited the catalytic activity of SPase, due to conformational perturbations and disruption of the enzyme’s active site. In contrast, Co²⁺, Ca²⁺, Fe²⁺, and Fe³⁺ have almost no effect on the enzyme activity. Comparing with Co²⁺, Fe²⁺, and Fe³⁺, Ca²⁺ provides better biocompatibility and a mature, physiologically relevant mineralization pathway, allowing mild, aqueous, tunable synthesis of nanoflowers [14].

Biological macromolecules containing abundant functional groups can act as molecular regulators during the nucleation and crystallization of hybrid materials [15,16]. To elucidate their influence on intracellular enzyme performance, three representative polymers, including chitosan, poly-L-lysine, and poly-L-glutamic acid, were comparatively investigated (Figure 2B). Both poly-L-glutamic acid and poly-L-lysine significantly caused the loss of catalytic activity, suggesting strong interactions between enzyme and polymers that impede substrate accessibility or alter protein conformation [17]. In contrast, chitosan maintained nearly unaltered catalytic activity. Owing to its minimal inhibitory effect, high biocompatibility, and capacity for coordination with metal ions [18,19], chitosan was identified as an optimal organic matrix for biomimetic mineralization during the formation of hybrid nanoflower.

After determining the metal ions and biomacromolecules, sodium tripolyphosphate (TPP), a biocompatible anionic crosslinker, further facilitates nanocomposite assembly with chitosan through electrostatic gelation [20]. Upon partial hydrolysis, TPP generates pyrophosphate ions that promote biomimetic mineralization with divalent metal ions, yielding hybrid organic–inorganic architectures. The addition of TPP to the solution of chitosan resulted in an immediate white precipitate. Metal ions and E. coli cells were then incorporated to initiate in situ mineralization. Based on the strong calcium-binding affinity and mineralization capability of chitosan, a novel immobilized cell system (CS–CaP@cells) was successfully constructed, providing a robust platform for enhanced catalytic activity and stability.

2.2. Effects of Immobilization Conditions on Relative Activity of CS-CaP@cells

During the immobilization of E. coli cells within organic–inorganic hybrid nanoflowers, the concentration of chitosan (CS), Ca²⁺, and TPP and the loading capacity of cell obviously influence the catalytic performance of the enzyme. As illustrated in Figure 3A, the highest relative activity was achieved at a chitosan concentration of 2 mg/mL, and relative activity was normalized to the catalytic performance obtained under the reference condition of 1.4 M L-AA, 0.8 M sucrose, and 45 °C for 48 h, yielding 156.7 g/L AA-2G, which corresponds to an L-AA conversion of approximately 40–45%. An increased concentration of chitosan led to excessively compact nanostructures that restricted substrate diffusion and consequently attenuated catalytic efficiency [21].

As shown in Figure 3B, the relative activity of SPase improved with the increase in cell loading, reaching its maximum at 600 μL. However, the further addition of cells might result in overcrowded microenvironments and aggravated mass-transfer resistance, ultimately diminishing enzymatic activity [22]. The influence of metal ion concentration on the SPase activity was further assessed. Figure 3C demonstrates that the optimal concentration of metal ions was 60 mM Ca²⁺. At higher Ca²⁺ concentrations (80–140 mM), the hybrid nanoflowers became excessively thick and dense, thereby impeding substrate accessibility and leading to a notable decline in catalytic output [23]. Similarly, Figure 3D revealed that the relative activity of SPase was the highest at a TPP concentration of 50 mg/mL. Collectively, owing to the excellent catalytic performance of SPase, the optimized immobilization conditions were determined as 2 mg/mL chitosan, 600 μL E. coli cells, 60 mM Ca²⁺, and 50 mg/mL TPP.

2.3. Characterization of Free Cells and CS-CaP@cells via SEM and EDS

After optimizing the immobilization conditions, the surface morphology, elemental composition, functional groups, and crystal structure of CS-CaP@cells were further characterized by SEM, EDS, FTIR, and XRD. The scanning electron microscopy (SEM) analysis was performed to elucidate the microstructural characteristics of the CS–CaP and CS–CaP@cells composites. As shown in Figure 4A,B, the CS–CaP material exhibited a distinctive flower-like hierarchical morphology. In Figure 4C, free cells are clearly observed to be encapsulated and uniformly distributed within the petal-like nanosheets of the CS–CaP matrix, confirming the effective immobilization of cells within the hybrid materials. Elemental mapping results (Figure 4D–G) revealed the well-defined distributions of calcium, phosphorus, carbon, nitrogen, and oxygen across the CS–CaP@cells structure. The distinct signals of carbon and nitrogen, characteristic elements of chitosan, confirm its successful integration into the composite architecture. Meanwhile, calcium and phosphorus were predominantly localized in the nanosheet regions, consistent with the formation of calcium phosphate nanoflowers. Furthermore, the energy-dispersive X-ray spectroscopy (EDS) analysis corroborated these findings, confirming that calcium phosphate and chitosan synergistically constitute the fundamental structural framework of the CS–CaP@cells composite. This hierarchical organization provides both mechanical robustness and biocompatibility, forming an ideal microenvironment for enzymatic catalysis.

2.4. Characterization of CS-CaP and CS-CaP@cells via FTIR and XRD

Fourier-transform infrared (FTIR) spectroscopy was employed to elucidate changes in chemical functionalities before and after E. coli immobilization in the CS–CaP nanocomposite. As depicted in Figure 5A, both CS–CaP and CS–CaP@cells exhibited absorption bands at 3454 cm⁻¹ and 2936 cm⁻¹, corresponding to the O–H and C–H stretching vibrations of chitosan. Distinct bands at 903 cm⁻¹, 758 cm⁻¹, and 698 cm⁻¹ were assigned to P–O vibrational modes, confirming the successful formation of calcium phosphate. Remarkably, an additional absorption band emerged at 1220 cm⁻¹ in CS–CaP@cells, attributable to –NH₂ groups originating from intracellular enzymes, thereby verifying effective cell immobilization in the hybrid materials.

X-ray diffraction (XRD) analysis further revealed the crystalline characteristics of the materials (Figure 5B). The CS–CaP composite displayed well-defined diffraction peaks at 2θ = 20.73°, 30.16°, 31.13°, 42.19°, 43.03°, and 51.00°, which are consistent with the reference pattern of Ca₂P₂O₇·4H₂O (PDF 44-0762). TPP undergoes partial hydrolysis to pyrophosphate, a process accelerated by acetic acid in the CS solution, and then resulting pyrophosphate ions adsorb onto the CS–TPP nanocomplexes. Subsequent introduction of Ca²⁺ triggers biomimetic mineralization of Ca₂P₂O₇·4H₂O on the nanocomplex network, templating the formation of hybrid nanoflowers [20]. The CS–CaP@cells sample exhibited identical diffraction peaks, indicating that the crystalline framework of calcium pyrophosphate was preserved during cell encapsulation. This attenuation in diffraction intensity suggests partial structural disorder introduced by biological components, further corroborating the successful biomimetic incorporation of cells within the inorganic matrix.

2.5. Optimal Reaction Conditions for Free Cells and CS-CaP@cells

The catalytic activity of free cells and CS-CaP@cells was systematically evaluated in diverse pH (4.5–7.5) and temperature (25–60 °C) conditions to assess their biochemical stability and functional adaptability. As shown in Figure 6A, free cells and CS-CaP@cells exhibited maximal catalytic activity at pH 5.2. Upon the immobilization, the relative activity of free cells maintained unchanged in the pH range of 4.5–7.5. The determination of optimal temperature (Figure 6B) revealed a consistent optimum temperature at 50 °C for free cells and CS-CaP@cells. The observed attenuation at elevated temperatures is likely due to the partial disintegration of the cellular envelope and the thermal denaturation of intracellular enzymes [22].

2.6. Thermal, pH, and Storage Stability and Reusability of Free Cells and CS-CaP@cells

The stability of free cells and CS-CaP@cells was systematically tested after incubating in 2 h across varying pH (4–9) and temperature (25–65 °C) conditions. As illustrated in Figure 7A, CS–CaP@cells maintained nearly constant catalytic activity (~95% retention) throughout the entire pH, markedly outperforming free cells. This exceptional pH stability can be ascribed to the protective flower-like CS–CaP architecture, which stabilizes enzyme conformation and prevents protonation-induced denaturation under extreme pH environments [24].

As shown in Figure 7B, CS-CaP@cells exhibited significantly enhanced thermal resistance, retaining over 50% of their initial activity at 65 °C, 2.13-fold higher than that of free cells. The improvement in thermal stability is attributed to the encapsulation of E. coli cells in chitosan-regulated hybrid nanoflowers, which mitigate thermal deactivation by restricting enzyme mobility and preserving tertiary structure integrity under heat stress.

As shown in Figure 7C, CS-CaP@cells retained approximately 85% of their initial activity after ten cycles, whereas free cells preserved only ~30%. The pronounced difference primarily arises from mechanical damage and enzyme leakage in free systems during repeated centrifugation, while nanoflowers encapsulation effectively shielded immobilized cells from shear-induced stress. Moreover, storage tests at 4 °C (Figure 7D) revealed that immobilized cells maintained approximately 80% of their initial activity after 13 days of preservation, nearly two-fold that of free cells (~40%). These findings collectively indicate that chitosan-mediated biomimetic mineralization establishes a robust protective microenvironment that substantially enhances the operational durability and storage longevity of intracellular enzymes, rendering the system promising for sustainable industrial-scale biocatalysis.

2.7. Organic Solvent and Byproduct Tolerance of Free Cells and CS-CaP@cells

SPase primarily accepts hydrophobic molecules such as glycosyl acceptors during transglycosylation reactions [25]. The inclusion of organic solvents in the reaction medium improves the solubility of these hydrophobic substrates, consequently enhancing catalytic performance [26]. However, organic solvents can permeate cellular membranes and disrupt the hydrophobic cores of enzymes, resulting in structural unfolding and loss of activity. To evaluate solvent tolerance, free cells and CS-CaP@cells were exposed to various organic solvents, including acetone, methanol, ethanol, acetonitrile, and dimethyl sulfoxide (DMSO). As illustrated in Figure 8A, CS-CaP@cells exhibited markedly superior tolerance to all tested solvents compared to free cells. The encapsulation of free cells within chitosan-regulated organic–inorganic hybrid nanoflowers effectively shielded intracellular enzymes from solvent-induced conformational disruption. This protective microenvironment endowed CS-CaP@cells with exceptional solvent resistance, underscoring their strong potential in the biosynthesis of glycosylated compounds under non-aqueous conditions.

Fructose, the principal by-product of SPase–catalyzed transglycosylation, acts as a potent competitive inhibitor through hydrogen-bonding interactions with protein surface residues. The accumulation of fructose significantly impairs enzymatic activity, thus constraining overall reaction efficiency. To assess the inhibition of by-product, enzyme activity was measured across fructose concentrations ranging from 0 to 2.4 M. As shown in Figure 8B, the catalytic activity of both systems declined progressively with increasing fructose concentration. Notably, CS-CaP@cells retained 60% of their initial activity at 2.4 M fructose, approximately four-fold higher than that of free cells (15%). This enhanced resistance arises from the CS–CaP composite framework, which provides a physical diffusion barrier that limits fructose accessibility to intracellular enzymes and mitigates inhibitory hydrogen bonding. To sum up, these findings highlight the superior structural resilience and operational stability of CS-CaP@cells under both solvent and product stress, validating their suitability for industrial-scale glycoside biocatalysis.

2.8. Comparing the Catalytic Performance of Free Cell and CS-CaP@cells

For comparing the industrial potential, free cell and CS-CaP@cells were added in a 1 L bioreactor to prepare the AA-2G, respectively. The AA-2G production of CS-CaP@cells was 156.7 g/L in 48 h (Figure 9), which was about 50 g/L higher than that of free cells. CS-CaP@cells demonstrated superior catalytic efficiency, providing a robust and scalable platform for high-yield glucoside production with strong industrial potential. (The yield of AA-2G was calculated using the standard curve provided in Appendix A).

3. Materials and Methods

3.1. Strains and Vector

The gene sequence corresponding to GenBank accession number AAO73025 was synthesized for this study (BGI Genomics, Shenzhen, China). The gene was subcloned into the pET-28a vector and ex-pressed in E. coli BL21(DE3). The E. coli strains DH5α and BL21(DE3) (TransGen Biotech, Beijing, China) and the expression vector pET-28a were obtained from our laboratory stock.

3.2. E. coli Culture

A 10 μL aliquot from the glycerol stock was inoculated into LB medium (Angel Yeast Co., Ltd., Yichang, China) containing 50 μg/mL kanamycin sulfate (Solarbio Life Sciences, Beijing, China) in a test tube, followed by incubation in a constant-temperature shaker (37 °C, 220 rpm) for 12 h. A 2 mL volume of the test tube culture was transferred as seed culture into LB medium (50 mL) supplemented with 50 μg/mL kanamycin sulfate and incubated under the same conditions until OD₆₀₀ reached 0.6–0.8. Protein expression was induced by adding isopropyl β-D-1-thiogalactopyranoside (IPTG) (Solarbio Life Sciences, Beijing, China) to a final concentration of 0.5 mM, and the culture was kept at 16 °C with shaking at 220 rpm for 20 h. Cells were harvested by centrifugation (6000 rpm, 4 °C, 10 min), washed three times with deionized water, and resuspended in an equal volume of deionized water. The cell suspension was stored at 4 °C until further use.

3.3. Screening of Biological Macromolecules and Metal Ions

Chitosan, poly-L-lysine hydrochloride, and poly-L-glutamic acid were each dis-solved at 2 mg/mL in 5 mL deionized water containing 1% (w/v) acetic acid. Separate 100 mM stock solutions of ZnSO₄, Co(NO₃)₂·6H₂O, CuSO₄·5H₂O, and CaCl₂ (Macklin Biochemical Co., Ltd., Shanghai, China) were prepared in deionized water. For each assay, 100 μL of one of the biological macromolecules or metal ion solutions (10%, v/v, final concentration) was added to a 1 mL whole-cell reaction mixture containing 1.2 mol/L ascorbic acid and 0.8 mol/L sucrose (Macklin Biochemical Co., Ltd., Shanghai, China). The reaction was carried out at 37 °C and 800 rpm in a thermostated metal bath for 6 h. Relative activity was calculated with the control reaction lacking any additive, which was defined as 100%.

3.4. Construction of CS-CaP@cells

An aqueous chitosan solution (Macklin Biochemical Co., Ltd., Shanghai, China, CS, 2 mg/mL, 500 μL) was mixed with a sodium tripolyphosphate solution (TPP, 50 mg/mL, 200 μL) and allowed to react at 20 °C and 800 rpm in a thermostated metal bath for 20 min. Subsequently, CaCl₂ (60 mM, 600 μL) and E. coli cells (OD600 = 40, 600 μL) were added, and the mixture was incubated for an additional 15 min under the same conditions. CS-CaP@cells were collected by centrifugation (8000 rpm, 5 min).

3.5. Characterization of Free Cells, CS-CaP, and CS-CaP@cells

The morphology and microstructure of E. coli, CS-CaP, and CS-CaP@cells were examined by scanning electron microscopy (SEM; HITACHI SU8600, Tokyo, Japan). Fourier-transform infrared (FTIR) spectroscopy (Nicolet IS5, Thermo Fisher Scientific, Waltham, MA, USA) was employed to analyze functional group compositions of immobilized cells within the 4000–400 cm⁻¹ range. Semi-quantitative elemental analysis and EDS mapping of CS-CaP@cells were determined using an energy-dispersive spectrometer (EDS; HITACHI, EMAX x-act, Tokyo, Japan) coupled to the SEM. The crystal structures of CS-CaP and CS-CaP@cells were characterized by X-ray diffraction (XRD; D8 Advance diffractometer, Bruker, Billerica, MA, USA).

3.6. SPase Activity Assay and Standard Curve Construction

An aqueous solution containing 1.2 mol/L ascorbic acid and 0.8 mol/L sucrose was prepared and adjusted to pH 5.2 with NaOH. Appropriate amounts of free cells or CS-CaP@cells were added to 1 mL of this solution and incubated at 37 °C and 800 rpm in a thermostated metal bath for 6 h. The reaction was terminated by heating the mixture at 100 °C for 10 min. Free cells or CS-CaP@cells were removed by centrifugation (8000 rpm, 5 min). The supernatant was filtered through a 0.22 μm microporous membrane, and 20 μL of the filtrate was injected into the HPLC system for analysis.

Separation was performed on a Diamonsil C18 column (4.6 × 250 mm, 5 μm, DIKMA Technologies, Beijing, China) thermostated at 30 °C. The mobile phase was an aqueous phosphate buffer (0.57 g/L K₂HPO₄·3H₂O, pH 2.0 adjusted with H₃PO₄) delivered at 1 mL/min flow rate. Detection employed a UV detector (Shimadzu UV-2600i Plus, Tokyo, Japan).

All experiments were performed in triplicate. Relative activity was calculated as follows:

Relative activity (%) = A_x/A₀ × 100,

where A₀ is the mean peak area of the control group, and A_x is the peak area of the test sample.

3.7. Optimal Reaction Conditions and Stability of Free Cells and CS-CaP@cells

Relative activity of free cells and CS-CaP@cells was determined at 25–60 °C. Free cells and CS-CaP@cells were incubated at 25–65 °C for 2 h, and residual activity was measured (maximum activity defined as 100%). Relative activity of free cells and CS-CaP@cells was assayed in various buffer (pH 4–9). Free cells and CS-CaP@cells were incubated in various buffers (20 mM, pH 4–9) for 2 h, followed by residual activity determination (maximum activity = 100%). Free cells and CS-CaP@cells stored at 4 °C for 15 days were assayed for relative activity (Day 1 activity = 100%). CS-CaP@cells were reacted under optimal conditions, washed with PBS buffer, and subjected to 15 reuse cycles (Cycle 1 activity = 100%).

4. Conclusions

In this study, a novel E. coli cell immobilization system (CS–CaP@cells) was successfully constructed through chitosan-regulated biomimetic mineralization. Comprehensive structural characterizations (SEM, XRD, FT–IR, and EDS) confirmed the effective encapsulation of cells within the calcium phosphate–chitosan hybrid nanomaterials. CS–CaP@cells showed approximately 30% higher relative activity than the initial activity of free cells under their optimal conditions. CS–CaP@cells maintained 85% relative activity after ten cycles. Moreover, the CS–CaP@cell system demonstrated enhanced tolerance to organic solvents and fructose inhibition. Overall, this biomimetic immobilization strategy could be used as a promising and scalable platform for cell immobilization to achieve the high-titer biosynthesis of glucosides.