Gas‐Shearing Microfluidic Fabrication of Alginate/Chitosan Hydrogel Microreactors for Genipin‐Mediated Dual‐Mode Lipase Immobilization

ABSTRACT

Enzyme immobilization is an effective strategy to improve catalyst recovery and operational stability, but conventional approaches are often limited by diffusion resistance, steric hindrance, and insufficient control over the enzyme microenvironment. Hydrogel‐based materials are attractive supports because their hydrated, tunable, and porous networks help preserve enzyme conformation while improving substrate diffusion and mass transfer. Here, a gas‐shearing microfluidic strategy was developed to fabricate alginate/chitosan hydrogel microreactors for dual‐mode lipase immobilization through entrapment and genipin‐mediated covalent binding. By tuning gas‐liquid shear, microspheres with controllable and narrow size distribution were generated without surfactants or UV‐initiated polymerization. Importantly, genipin was introduced as a mild and biocompatible cross‐linker to reinforce the alginate/chitosan network and create a stable interfacial microenvironment for enzyme immobilization. Compared with conventional cross‐linkers, the milder and more controllable reactivity of genipin is advantageous in preserving enzyme conformation and catalytic performance. The resulting microreactors exhibited enhanced catalytic activity, improved storage stability, and good reusability. After five consecutive cycles, the entrapped and covalently immobilized systems retained 70% and 81% of their initial activity, respectively. These results demonstrate that gas‐shearing microfluidics combined with genipin‐mediated network reinforcement provides a robust and biocompatible platform for constructing hydrogel‐based enzymatic microreactors with improved catalytic performance and operational durability.

A gas‐shearing microfluidic strategy was developed to fabricate alginate/chitosan hydrogel microreactors for dual‐mode lipase immobilization. By integrating enzyme entrapment with genipin‐mediated covalent binding, the microreactors provide a mild, biocompatible, and structurally reinforced microenvironment that improves catalytic performance, storage stability, and reusability. In addition, the immobilized systems retained high activity after repeated cycles, with 70% activity remaining for entrapment and 81% for covalent immobilization after five cycles.

1. Introduction

Although enzymes are highly efficient and selective biocatalysts with great potential for green biocatalysis and biomanufacturing, their poor stability, easy inactivation, and limited reusability under nonphysiological or industrial conditions constrain practical application, while conventional immobilization strategies, despite improving stability and recyclability, often fail to simultaneously optimize the enzyme microenvironment and mass transfer [1, 2, 3, 4].

Importantly, the efficiency of biochemical reaction networks depends not only on the intrinsic activity of individual enzymes but also, more critically, on spatiotemporal organization and microenvironmental regulation [5]. In cells, specific enzymes and substrates are frequently co‐localized within confined compartments to maintain metabolic order, increase local concentrations that facilitate thermodynamically unfavorable steps, and protect enzymes and labile intermediates from external stresses (e.g., proteases or toxic chemicals). Nowadays, materials based on three‐dimensional hydrogels have emerged as a significant research direction in the field of enzyme immobilization. Hydrogels possess tunable pore sizes, morphologies, composite structures, excellent specific surface areas, and mechanical strength. These characteristics ensure effective enzyme immobilization and provide pathways for directing substrate molecules towards active sites [6, 7, 8, 9, 10]. Three‐dimensional hydrogels retain substantial moisture, creating a hydrated environment that minimizes enzyme denaturation and preserves catalytic functionality [11].

Inspired by cellular compartmentalization, enzyme microreactors and bioinspired encapsulation systems have attracted increasing attention [12]. Among existing fabrication approaches, self‐assembly is simple and rapid but often yields polydisperse particles and limited batch‐to‐batch consistency, with constrained throughput or yield [13, 14]. By contrast, microfluidics enables precise control over size and morphology, producing more uniform microreactors [15, 16]. However, conventional microfluidic immobilization methods are often constrained by limited accessible surface area [17], steric hindrance, and the potential adverse effects of surfactants [18] or UV‐initiated polymerization [19] on biocompatibility and enzyme activity, underscoring the need for highly biocompatible miniaturized reactors with tunable dimensions and diverse morphologies. Therefore, this study combines three‐dimensional hydrogels with microfluidic technology to achieve enzyme encapsulation. Sol–gel immobilization encapsulates microorganisms or enzymes within a hydrogel network through a mild liquid‐to‐gel transition, enabling controlled incorporation while preserving their structural integrity and activity [11]. Owing to the tunable pore structure, functional groups, and surface reactivity of sol–gel materials, this strategy can improve enzyme thermal stability, solvent tolerance, mechanical resistance, and storage stability [20]. It has been widely applied as a simple and effective immobilization method for enzymes such as xylanase, horseradish peroxidase, oxidoreductase, and lipase.

Cross‐linking methods are frequently employed to immobilize enzymes. Glutaraldehyde is one of the most widely used catalysts in enzyme immobilization. It forms a covalent Schiff base (imide bond) via its aldehyde group, thereby stabilizing the enzyme's amino group [21, 22], however, it exhibits certain cytotoxic properties and poor biocompatibility [23, 24]. Currently, genipin is frequently employed as a cross‐linking agent. As a plant‐derived reagent, it can be extracted from the fruit of the Asian evergreen flowering plant Gardenia jasminoides J. Ellis, or obtained from the unripe fruit of Genipa americana L., which grows wild in the Amazonian lowland forests and other regions of Latin America. Within the food sector, genipin is commonly employed as a food colorant. Moreover, owing to its multifaceted properties—including antioxidant, anti‐inflammatory, anti‐thrombotic, and neuroprotective effects—it holds considerable promise in the field of pharmacology [25]. Genipin undergoes cross‐linking reactions with biopolymers containing primary amine groups. This cross‐linking alters porosity and adjusts microstructure [26, 27], whilst the relatively slow reaction rate of genipin cross‐linking allows precise control over cross‐linking speed and extent. Furthermore, genipin itself is colorless, but in the presence of oxygen, it reacts with primary amines and proteins to form a water‐soluble blue‐violet pigment [25]. This color change can be used to identify the extent of the cross‐linking reaction. Consequently, genipin is safer as a cross‐linking agent and exhibits excellent biocompatibility.

To the end, we developed a reliable and biocompatible gas‐shear microfluidic method to fabricate lipase microreactors. By tuning the gasflow, we generated microspheres with narrow size distributions and tunable morphologies without using surfactants or UV‐initiated polymerization [28, 29]. Moreover, genipin was employed as a biocompatible cross‐linker to reinforce a chitosan (CS)/alginate (Alg) entanglement network for stabilizing lipase immobilization [30]. Compared with glutaraldehyde, genipin provides milder and more controllable cross‐linking, which is beneficial for preserving enzyme conformation and catalytic performance. Catalytic performance was evaluated using a p‐nitrophenyl ester substrate system. Overall, this strategy offers a general platform for constructing high‐performance, scalable enzyme microreaction systems.

2. Experimental and materials

2.1. Materials

Sodium alginate (Alg, viscosity ≥0.02 (10 g/L, 20°C)/(Pa/s), CP) was purchased from Shanghai Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). Chitosan (CS, degree of deacetylation 80.0‐95.0%, BR) was obtained from Sinopharm Chemical Reagent Co., Ltd. (China). Lipase (30 U/mg, BR) was provided by Orileaf. Genipin (98%) was acquired from MACKLIN. Anhydrous calcium chloride (CaCl₂, AR) was purchased from Sinopharm Chemical Reagent Co. Ltd.

2.2. Fabrication of Alg@CS Microparticle Enzymatic Microreactors

The enzymatic microreactors were fabricated according to the method reported by Qu et al. [31]. Alg@CS microparticles were used as carriers to construct enzymatic microreactors. As shown in Scheme 1, the gas‐shearing microfluidic setup comprised five components: (i) a microsyringe pump to deliver the polymer solution, (ii) a collection bath to collect particles, (iii) a gas pump to supply the gas stream, (iv) a flow meter to monitor the gas flow rate, and (v) a spray ejector device (SED). The SED consisted of a concentric needle pair, with an inner needle delivering the polymer solution and an outer needle conveying gas. Gas‐liquid shear at the nozzle generated fine droplets. Subsequently, the droplets fell into a polymer‐containing collection bath and solidified into microspheres.

SCHEME 1.

Schematic diagram of microparticles fabricated by gas shearing microfluidic technology.

In this study, 2% (w/v) alginate solution was delivered through a coaxial needle assembly using a microsyringe pump at 1 mL/h. The inner needle (22G, 420 µm) carried the liquid phase, whereas the outer needle (17G, 1110 µm) delivered the gas phase. The collection bath consisted of 2% (w/v) CaCl₂ solution and chitosan (CS; 1% (w/v) in 1% (v/v) aqueous acetic acid), mixed at volume ratios of 3:7, 5:5, 7:3, and 9:1. The collection distance was set to 8 cm. Finally, the collected microspheres were rinsed thoroughly with deionized water to remove residual reagents, and the as‐prepared microspheres were stored at 4°C.

2.3. Enzyme Immobilization

2.3.1. Entrapment of Lipase

First, a 2% (w/v) alginate solution was mixed with 5 mL of lipase solution at 30, 90, 150 or 300 U/mL. Subsequently, the mixture was loaded into the spray ejector device and pumped at 1 mL/h, while being sheared by a gas stream at 1.0, 1.5, 2.0, 2.5, or 3.0 L/min. Meanwhile, a collection bath containing 1% (w/v) CS and 2% (w/v) CaCl₂ (v/v = 5:5) was used to collect the microspheres. Finally, the collected microspheres were washed three times with distilled water, resuspended in water, and stored at 4°C.

2.3.2. Covalent Binding of Lipase

The as‐prepared Alg@CS microspheres were used as carriers for covalent immobilization of lipase. First, about 1100 microspheres were activated in 2 mL of genipin solution (5, 0.5, 0.05, 0.005, or 0.0005 mg/mL) with shaking for 6 h at 37°C. Then the microspheres were then rinsed thoroughly with deionized water to remove unreacted genipin and other residuals. Next, lipase was covalently conjugated to the activated microspheres. A lipase solution (300 U/mL) was added to the activated microspheres and incubated with shaking (125 rpm, 37°C) for 0.5, 2, 4, 6, or 8 h to achieve adsorption/conjugation equilibrium. The lipase‐conjugated microspheres were washed with water to remove unbound enzyme and stored at 4°C.

2.4. Evaluation of Enzymatic Activity

For the free‐enzyme assay, a 90 U/mL lipase solution was reacted with a p‐nitrophenyl ester substrate, and the absorbance spectrum (350–550 nm) was recorded every 5 min.

For the immobilized‐enzyme assay, 1100 lipase‐immobilized microspheres (35 microspheres per min count by eyes) were placed in a 5 mL tube, followed by addition of the p‐nitrophenyl ester substrate. The absorbance spectrum (350‐550 nm) of the supernatant was measured every 5 min.

The P‐nitrophenyl ester substrate was converted to yellow under the catalytic action of the enzyme, with an absorption peak observed at 402 nm. When the absorption curve became nearly linear, the reaction was considered to have reached equilibrium. The reaction rate was determined from the slope of the absorbance‐time curve between 0 and 60 min.

2.5. Lipase Stability

For the reusability test, after completion of each reaction cycle, the microspheres were washed three times with deionized water. Fresh p‐nitrophenyl ester substrate was then added to the microspheres to initiate the next cycle. This procedure was repeated for a total of five cycles.

For the storage‐stability test, two types of microspheres were stored at 37°C, and enzyme activity was measured on days 0, 25.

3. Results and Discussion

3.1. Fabrication and Characterization of Alginate Microspheres

Alginate microspheres were prepared by gas‐shearing microfluidics (Figure 1a), and their size was tuned by varying the gas flow rate, which decreased with increasing gas flow rate (Figure 1b). Representative micrographs (Figure 1c and S1) showed a narrow size distribution and smooth particle surfaces. Particle diameters were measured from the micrographs using Image J, and 120 microparticles per group were used for size statistics. At gas flow rates of 1.0, 1.5, and 2.0 L/min, the mean diameters were 648.89 ± 1.48 µm, 468.55 ± 1.50 µm, and 464.83 ± 1.45 µm, respectively. It demonstrates that gas‐shearing microfluidics enables robust, tunable control of microsphere size and morphology. Scanning electron microscope (SEM) is used to observe the structure of blank microspher, enzyme‐encapsulated microsphere and enzyme‐crosslinked microsphere. Blank microsphere and enzyme‐encapsulated microsphere have a soft surface texture (Figures 1d,e). Enzyme‐crosslinked microsphere have a rough surface texture with some creases (Figure 1f) and its diameter increase by approximately 40 µm (Figure S2), demonstrates the successful cross‐linking of the microsphere surface and enzyme loading. Moreover, Energy‐dispersive X‐ray spectroscopy (EDS) elemental mapping observes the elements N, S, and Ca, further confirms successful fabrication (Figures 1d‐f).

FIGURE 1.

(a) Scheme of the fabrication process of the microspheres. (b) The size distributions of microspheres prepared with different gas flow rates. (c) The optical image and size distribution of the microspheres (added with purple dye) (n = 120). The SEM image of (d) blank microsphere (e) enzyme‐encapsulated microsphere (f) enzyme‐crosslinked microsphere with the corresponding EDS elemental mapping for N, S, and Ca, scale bar: 500 nm.

3.2. Activity of Free Enzyme and Enzyme‐Encapsulated Microsphere

Entrapment is a common strategy for enzyme immobilization (Figure 2a), and the effect of initial lipase concentration (30 U/mL, 90 U/mL, 150 U/mL) on immobilization performance was evaluated. As shown in Figure 2b and S3, the activity of entrapped lipase displayed a nonmonotonic dependence on enzyme concentration, reaching a maximum at 90 U/mL. The activity at 90 U/mL was 3.3 fold and 1.3 fold higher than those at 30 and 150 U/mL, respectively. At low enzyme concentration, the catalytic capacity is limited by insufficient enzyme loading. However, at excessively high concentration, the overly high local enzyme density within the confined matrix may induce molecular crowding, restrict substrate diffusion, and reduce active‐site accessibility, thereby lowering the apparent catalytic efficiency [32]. Moreover, free lipase at the same concentration showed 1.6 fold higher activity than entrapped lipase, suggesting that matrix confinement introduces additional diffusion resistance despite successful enzyme incorporation. Together, these results indicate that 90 U/mL is the optimal embedding concentration under the present conditions. In addition, to address the comparison with the covalent immobilization system, an entrapment experiment using a higher initial lipase concentration of 300 U/mL was also performed (Figure S4). The activity of the 300 U/mL entrapment system was lower than that of the 90 U/mL entrapment system, further confirming that excessive enzyme input was unfavorable for the entrapment mode under the present microfluidic preparation conditions.

FIGURE 2.

(a) Scheme of the fabrication process of the entrapped microspheres. (b) The change of absorbances at 402 nm of the reaction by free lipase, 30 U/mL, 90 U/mL, 150 U/mL embedding lipase.

3.3. Activity of Free Lipase and Covalently Immobilized Lipase

Compared with glutaraldehyde, a widely used cross‐linking agent, genipin is more biocompatible and typically functions under milder reaction conditions. These features may better preserve enzyme activity and provide improved control over the immobilization process, thereby facilitating a balance between catalytic stability and mass transfer. In addition, genipin crosslinking produces a characteristic blue coloration, which serves as a convenient visual indicator of activation (Figures 3d,e). As illustrated in the Figure 3a, genipin can crosslink with amino groups via two primary pathways. In the first pathway, the amino group directly attacks the carbonyl carbon of the ester group of genipin; subsequent nucleophilic substitution converts the ester into an amide bond, resulting in an amide‐type covalent linkage. In the second pathway, nucleophilic attack of the amino group at the C₃ position induces opening of the dihydrofuran ring, followed by further reactions via an aldehyde intermediate to form a heterocyclic amine‐type linkage [33]. Through these reactions, genipin‐mediated surface crosslinking/activation generates a more stable interface for subsequent lipase immobilization.

FIGURE 3.

(a) Scheme of the fabrication process of the covalent immobilized microspheres via genipin. (b) The change of absorbances at 402 nm of the reaction by free lipase, 100 U/mL, 300 U/mL, 500 U/mL covalent binding lipase. (c) The change of absorbances at 402 nm at different gas flow rate. The optical image of microspheres (d) before covalent immobilization and (e) after covalent immobilization.

As shown in Figure 3b and S5, the activated microspheres were incubated with lipase at 100, 300, and 500 U/mL to determine the optimal loading condition. The activity of immobilized lipase first increased and then decreased with increasing lipase concentration, reaching a maximum at 300 U/mL. Specifically, the activity at 300 U/mL was 6.5 fold and 5.0 fold higher than those at 100 and 500 U/mL, respectively. It suggests that an appropriate enzyme loading level is required to achieve optimal catalytic performance. At low lipase concentration, the limited amount of immobilized enzyme may be insufficient to provide high catalytic capacity. In contrast, excessive enzyme loading may increase local enzyme crowding at the microsphere interface, reduce substrate accessibility to active sites, and aggravate mass‐transfer limitations, thereby decreasing the apparent catalytic efficiency [34]. Moreover, the activity of immobilized lipase was 3.5 fold higher than that of free lipase at the same concentration, indicating that genipin‐mediated surface activation of the microspheres provided a favorable microenvironment for enzyme immobilization and catalysis [35]. As shown in Figure 3c and S6, microsphere activity decreased with increasing gas flow rate, suggesting that excessive gas shear is unfavorable for maintaining optimal catalytic performance. These results demonstrate that both enzyme loading and operating conditions play critical roles in determining the catalytic performance of the immobilized microsphere system.

To further clarify the comparison between the two immobilization strategies, an additional entrapment experiment was performed using the same initial lipase concentration as that used for the covalent binding system, namely 300 U/mL (Figure S4). Although 300 U/mL was the optimal initial enzyme concentration for covalent immobilization, the activity of the 300 U/mL entrapment system was lower than that of the 90 U/mL entrapment system. This result indicates that simply increasing the enzyme input did not improve the catalytic performance of the entrapment mode. The reduced activity at 300 U/mL may be attributed to excessive enzyme crowding within the hydrogel matrix, increased diffusion resistance, and decreased accessibility of substrate molecules to enzyme active sites. Moreover, the high‐concentration enzyme‐containing phase exhibited increased viscosity during microfluidic preparation and occasionally caused partial needle blockage, resulting in less stable droplet generation. Therefore, 90 U/mL was selected as the optimal and practically feasible enzyme concentration for the entrapment method, whereas 300 U/mL was optimal for the covalent binding method. These results indicate that the optimal enzyme input is strongly dependent on the immobilization strategy, and the comparison between the two modes should be interpreted by considering both catalytic performance and preparation feasibility.

A single‐factor optimization strategy was used to evaluate the effects of genipin concentration in the activation solution, CS/CaCl₂ volume ratio in the collection bath, and immobilization time on lipase activity (Figure 4). The initial lipase concentration was fixed at 300 U/mL. When optimizing the genipin concentration, the CS/CaCl₂ volume ratio and immobilization time were fixed at 5:5 (v/v) and 6 h, respectively. When optimizing the CS/CaCl₂ volume ratio, the genipin concentration and immobilization time were fixed at 0.05 mg/mL and 6 h, respectively. When optimizing the immobilization time, the genipin concentration and CS/CaCl₂ volume ratio were fixed at 0.05 mg/mL and 5:5 (v/v), respectively.

FIGURE 4.

The change of absorbances at 402 nm by covalently immobilized lipase under different optimization conditions: (a,d) genipin concentrations, with the CS/CaCl₂ volume ratio and immobilization time fixed at 5:5 (v/v) and 6 h, respectively; (b,e) CS/CaCl₂ volume ratios, with the genipin concentration and immobilization time fixed at 0.5 mg/mL and 6 h, respectively; and (c,f) immobilization times, with the genipin concentration and CS/CaCl₂ volume ratio fixed at 0.5 mg/mL and 5:5 (v/v), respectively.

As shown in Figures 4a,d, immobilized lipase activity exhibited a nonmonotonic dependence on genipin concentration (0.0005 to 5 mg/mL) at an activation time of 6 h, with an apparent optimum around 0.05 mg/mL. When genipin concentration was below 0.05 mg/mL, activity decreased with further reductions in genipin, suggesting that insufficient activation limits the number of available binding sites for lipase immobilization. In contrast, excessive genipin can reduce lipase activity for two main reasons. First, extensive multipoint covalent attachment may “lock” the enzyme in a constrained conformation, limiting the structural flexibility required for catalysis. Second, genipin may promote enzyme–enzyme crosslinking, forming dense crosslinked layers or aggregates that hinder substrate access to active sites and decrease apparent activity [36]. Accordingly, 0.05 mg/mL genipin was selected for subsequent experiments.

The effect of the CS/CaCl₂ ratio on the activity of immobilized lipase is shown in Figures 4b,e. Across the tested CS/CaCl₂ ratios (3:7 to 9:1,v/v), lipase activity increased initially and then decreased. At higher CS content, enhanced ionic interactions and covalent coupling involving hydroxyl and amino groups may promote the formation of a denser polyelectrolyte layer on the microsphere surface, which can hinder substrate diffusion and reduce apparent activity. Therefore, a CS/CaCl₂ ratio of 5:5 (v/v) in the collection bath was used in subsequent experiments.

The effect of immobilization time on lipase activity is shown in Figures 4c,f. Activity increased as the immobilization time was extended from 0.5 to 6 h; whereas beyond 6 h the change became decreased. Extending the immobilization duration can enhance enzyme loading by promoting sustained enzyme‐carrier interactions and reducing enzyme loss from surface and pore structures, thereby increasing activity up to an optimal duration. However, prolonged immobilization did not further improve activity under the tested conditions.

3.4. Storage Stability and Reusability

As biocatalytic platforms, microreactors are expected to maintain activity over extended periods and support repeated reaction cycles. As shown in Figure 5a, the activity of free lipase decreased from day 0 to day 25 at 37°C, whereas the apparent activity of the immobilized lipase increased over the same storage period. This phenomenon should not be interpreted as an intrinsic enhancement of lipase activity, but rather as an increase in apparent catalytic response associated with the hydrogel microenvironment. For freshly prepared Alg/CS hydrogel microreactors, the internal network may not have reached complete hydration and structural equilibrium at day 0. During storage, gradual water equilibration, network relaxation, and moderate swelling of the hydrogel matrix may improve substrate diffusion and increase the accessibility of lipase active sites. In addition, genipin‐mediated cross‐linking may further stabilize the local microenvironment around lipase, helping to reduce unfavorable conformational changes during storage [37, 38]. Therefore, the increased activity of immobilized lipase after storage may result from the combined effects of improved mass transfer, increased active‐site accessibility, and enhanced microenvironmental stabilization.

FIGURE 5.

(a) Storage stability of free lipase and microspheres. (b) The optical image of enzyme‐encapsulated microspheres and enzyme‐crosslinked microspheres before cycle and after 5 cycles. Reusing stability of (c) entrapped microspheres and (d) cross bonding microspheres.

Benefitting from the ease of separating the microspheres from the reaction system, their reusability was further tested. The immobilization microspheres prepared by entrapment and covalent cross‐linking could retain the high activity and structural integrity over five cycles (Figures 5b‐d). In Figure 5c, the reusability of entrapment maintained 70% after five consecutive cycles. In Figure 5d, the reusability of covalent binding maintained 81% after five consecutive cycles. This minor decrease may attributable to the cumulative effects of partial enzyme inactivation and enzyme loss during repeated reaction and washing processes. In addition, repeated use may alter the interfacial microenvironment of the microspheres and aggravate local mass‐transfer limitations, thereby reducing substrate accessibility to the active sites and lowering the apparent catalytic efficiency.

4. Conclusion

In this work, a gas‐shearing microfluidic strategy was combined with an Alg/CS hydrogel network and genipin‐mediated cross‐linking to construct lipase‐loaded hydrogel microreactors. The central hypothesis was that this mild and biocompatible microenvironment could improve enzyme immobilization while maintaining catalytic performance. Compared with conventional self‐assembly or harsh cross‐linking approaches, this strategy enabled controllable microsphere fabrication under surfactant‐free and UV‐free conditions, while genipin provided a milder alternative to glutaraldehyde for reinforcing the hydrogel network and stabilizing the enzyme microenvironment.

The results showed that enzyme input strongly affected the catalytic performance of different immobilization modes. The optimal initial lipase concentration was 90 U/mL for entrapment and 300 U/mL for covalent binding. Excessive enzyme input did not further improve the entrapment system, but instead reduced apparent activity and affected droplet formation due to increased viscosity and partial needle blockage. Both immobilization modes exhibited improved storage stability and reusability, suggesting that Alg/CS hydrogel microreactors have potential as recoverable enzymatic platforms.

Nevertheless, this study still has some limitations. First, the effects of pH and reaction temperature on immobilized lipase activity were not systematically investigated. Because the Alg/CS hydrogel network is sensitive to environmental changes, variations in pH or temperature may affect not only lipase conformation and catalytic activity, but also the swelling, deformation, or partial dissociation of the hydrogel matrix. Therefore, future studies should combine pH‐ and temperature‐dependent activity assays with particle morphology observation, swelling analysis, and enzyme leakage measurements. Second, the apparent increase in immobilized enzyme activity after storage requires further mechanistic verification. Time‐dependent structural characterization and mass‐transfer analysis will be necessary to clarify the relationship between hydrogel network relaxation, substrate diffusion, and enzyme active‐site accessibility.

Overall, this work demonstrates that gas‐shearing microfluidics combined with genipin‐mediated Alg/CS hydrogel reinforcement provides a robust, reproducible, and biocompatible strategy for constructing enzyme microreactors. Future work will focus on improving hydrogel structural robustness, expanding the system to other enzymes or multi‐enzyme cascades, and establishing a more comprehensive evaluation of operational stability under different pH, temperature, and reaction environments.

Author Contributions

Dong Pei initiated the project. Qingli Qu designed an experimental idea. Meng Wang and Chengfu Zhang conducted the experiments with assistance from Qingli Qu. Dong Pei, Puzhao Liu, Ningli Wang and Qingli Qu acquisite fundings. Dong Pei and Qingli Qu supervised the experimental work. Meng Wang, Hongzhen Jia and Qingli Qu prepared the original draft. Qingli Qu and Ningli Wang was in charge of revising. All the authors read and approved the final manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: mabi70238‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.