Fabrication of Yeast-Immobilized Porous Scaffolds Using a Water-in-Water Emulsion-Templating Strategy

Abstract

This study introduces an efficient, all-aqueous emulsion-templating strategy for fabricating highly tunable yeast immobilization carriers with superior biocatalytic performance. Utilizing cellulose nanocrystals (CNCs) to stabilize dextran/polyethylene glycol (Dex/PEG) water-in-water emulsions, an architecture-controlled void is obtained by crosslinking the PEG-rich phase with variable concentrations of polyethylene glycol diacrylate (PEGDA) (10–25 wt%). This approach successfully yielded macroporous networks, enabling precise tuning of void diameters from 10.4 to 6.6 μm and interconnected pores from 2.2 to 1.4 μm. The optimally designed carrier, synthesized with 15 wt% PEGDA, featured 9.6 μm voids and robust mechanical strength (0.82 MPa), and facilitated highly efficient yeast encapsulation (~100%). The immobilized yeast demonstrated exceptional fermentation activity, remarkable storage stability (maintaining > 95% productivity after 4 weeks), and high reusability (85% activity retention after seven cycles). These enhancements are attributed to the material’s excellent water retention capacity and the provision of a stable microenvironment. This green and straightforward method represents a significant advance in industrial cell immobilization, offering unparalleled operational stability, protection, and design flexibility.

1. Introduction

Cell immobilization refers to the confinement of animal, plant, or microbial cells within a defined carrier space through physical or chemical means, while maintaining their reproductive and metabolic capabilities [1,2]. This technology utilizes immobilized cells as biocatalysts, offering several advantages over free cells or immobilized enzymes: (1) elimination of enzyme extraction and purification steps, thereby preserving native enzyme activity; (2) retention of endogenous coenzyme regeneration systems, obviating the need for external cofactors while enabling multi-enzyme cascade catalysis; and (3) maintenance of enzyme activity and stability through the protective intracellular environment. The catalytic performance of immobilized cells is strongly influenced by carrier properties, with appropriate material selection enabling adaptation to diverse reaction conditions [3,4,5,6]. Currently, this technology finds widespread application in environmental remediation, food fermentation, energy production, and pharmaceutical manufacturing [1,2,3,4,7,8,9].

Immobilization methods can be broadly classified into chemical (covalent bonding and cross-linking) and physical (adsorption and embedding) approaches [2,10,11,12,13]. While chemical methods provide strong cell fixation, they often compromise viability due to harsh bonding conditions. In contrast, physical methods have gained increasing attention for their ability to maintain cell activity comparable to free cells. Physical immobilization techniques include: (a) surface adhesion/adsorption, (b) entrapment within porous matrices, (c) self-aggregation via flocculation, and (d) encapsulation behind semi-permeable barriers.

The choice of carrier material critically determines the success of cell immobilization by influencing microbial viability and, consequently, catalytic efficiency [1,8,10,11,13]. Recently, open-cell porous polymers, such as emulsion-templated porous polymers (polyHIPEs), have emerged as promising carriers due to their high cell-loading capacity, excellent mass transfer properties, and facile separation from reaction media [7,14,15,16,17,18,19,20,21,22,23,24]. Conventional polyHIPEs for cell immobilization are typically synthesized using oil-in-water (o/w) or water-in-oil (w/o) high internal phase emulsions (HIPEs), requiring post-fabricating cell infiltration (Cells@polyHIPE) to avoid exposure to harmful organic solvents and monomers [18,19,20,25,26,27]. However, this sequential approach suffers from several limitations: (1) time-consuming processing, (2) low cell-loading efficiency, (3) non-uniform cell distribution, and (4) cell leakage during operation.

This research is a continuation of our previous work [18], which was limited to post-preparation encapsulation. The key advancement presented here is the use of a water-in-water (w/w) emulsion-templating technique to achieve in situ yeast encapsulation during carrier synthesis. The w/w emulsion-templating method is a green, scalable process we developed for producing porous materials [28]. This technique utilizes an emulsion formed from two incompatible polymer aqueous solutions [29,30,31], entirely eliminating the need for organic solvents required by traditional emulsion-templating [28,32]. In this work, yeast cells are dispersed in a dextran aqueous solution (dispersed phase), which is emulsified in a polyethylene glycol diacrylate (PEGDA) aqueous solution (continuous phase). Subsequent polymerization of PEGDA and dextran removal yields porous PEG with uniformly distributed yeast cells (yeast@PPEG). This approach offers significant advantages: (1) complete avoidance of organic solvents, ensuring excellent cell viability; (2) single-step immobilization with near-quantitative cell incorporation (~100% efficiency); and (3) formation of a homogeneous cell distribution within the carrier. The entirely aqueous process represents an environmentally benign and cell-friendly immobilization platform with broad potential applications in biocatalysis, biomedicine, and food technology.

2. Results and Discussion

2.1. Preparation of w/w Emulsion-Template and PPEG

Unlike traditional oil-in-water (o/w) or water-in-oil (w/o) emulsions, water-in-water (w/w) emulsions consist of two aqueous phases with very low interfacial tension, making them difficult to stabilize using conventional surfactants. In this study, we employ nanocrystalline cellulose (CNC) particles as stabilizers to prepare w/w emulsions. Given the significant influence of polyethylene glycol diacrylate (PEGDA) concentration on the dextran (Dex)/PEG emulsion-templating method and the resulting porous material morphology, we prepared a series of w/w emulsions (EPEG-10, EPEG-15, EPEG-20, and EPEG-25) by varying PEGDA concentrations and investigated its effect on porous structure.

Due to the similar physical properties and extremely low interfacial tension between the two aqueous phases, conventional microscopy cannot clearly resolve the emulsion’s internal structure. Therefore, we fluorescently labeled the Dex phase and analyzed the emulsions using laser confocal microscopy. Figure 1 shows that in all four emulsions, the FITC-Dex-labeled phase (fluorescent green) forms the dispersed phase, while the unstained PEG phase (black) constitutes the continuous phase. This structure provides an effective template for subsequent polymerization of PEGDA in the continuous phase to form poly-PEGDA porous materials (PPEG).

Furthermore, Figure 1 reveals that as the PEG concentration increases, the droplet size (Dex phase) progressively decreases. The average diameters of the Dex droplets in EPEG-10, EPEG-15, EPEG-20, and EPEG-25 are 39.9 μm, 30.8 μm, 25.3 μm, and 18.0 μm, respectively. This trend arises because the w/w emulsion’s two aqueous phases undergo water diffusion and redistribution during emulsification due to osmotic imbalance, altering their volume fractions from the initial solution ratios. The actual phase volumes depend on the overall Dex/PEG concentration ratio in the emulsion. Thus, increasing PEG concentration while keeping Dex constant reduces the Dex/PEG ratio, leading to a smaller dispersed-phase volume fraction.

Given the temperature-sensitive stability of w/w emulsions, we employed a redox initiator system to rapidly polymerize PEGDA into PPEG. As shown in Figure 2, the resulting porous materials exhibit a networked, highly open-cell structure. The void size decreases with increasing PEGDA concentration, mirroring the emulsion droplet trend. However, statistical analysis reveals that the average void diameters of PPEG-10, PPEG-15, PPEG-20, and PPEG-25 (10.4 μm, 9.6 μm, 8.8 μm, and 6.6 μm, respectively) are notably smaller than their corresponding emulsion droplets. This discrepancy may stem from the polymerization process: PEGDA (Mn = 400) converts to high-molecular-weight PEG, increasing the continuous phase’s affinity for water. For water-soluble polymers of equal mass, higher molecular weight enhances water “adsorption,” thereby shrinking the Dex-phase droplets [33].

Given the critical role of mechanical properties in cell immobilization carriers, we performed compression tests on the PPEG materials. The stress–strain curves (Figure 3) reveal that both compressive strength and modulus increase with higher PEGDA content (

Table 1. Mechanical property data of different samples.

Sample	Average Void Size (µm)	Average Interconnected Pore (µm)	Compression Modulus (MPa)	Compressive Strength (MPa)	Crushing Compression Ratio (%)
PPEG-10	10.4	2.2	0.16	0.80	58.4
PPEG-15	9.6	1.8	0.45	0.82	55.2
PPEG-20	8.8	1.7	0.47	0.94	53.4
PPEG-25	6.6	1.4	0.94	1.04	51.1

). Specifically, PPEG-25 exhibits the highest compressive strength (1.04 MPa) and compression modulus (0.94 MPa), attributed to its greater material density compared to other PPEG variants. However, the crushing compression rate decreases with increasing PEGDA concentration—PPEG-10 demonstrates the highest rate (58.4%), indicating superior toughness. These results confirm that the synthesized PPEGs possess well-balanced mechanical properties suitable for cell carrier applications.

Additionally, since the target yeast cells in this study (Saccharomyces cerevisiae, 2–5 µm in size) [18,34] require sufficient space for mobility and retention during fermentation, we selected PPEG-15 as the immobilization carrier. Its average void size (9.6 µm) and interconnected pore diameter (1.8 µm) provide ample room for cell movement while preventing escape, ensuring optimal encapsulation efficiency.

2.2. Preparation of Yeast@PPEG

In this study, yeast cells were encapsulated within porous materials prepared via the w/w emulsion-templating method. Unlike conventional approaches requiring pre-fabricated scaffolds, this method enables direct in situ immobilization by simply adding yeast to the Dex phase prior to emulsification. Following polymerization, the yeast cells remain embedded within the porous matrix.

Due to the presence of yeast glucan in their cell walls, yeast exhibits a stronger affinity for the Dex solution than for PEG, preventing cell migration to the PEG phase during emulsification. To facilitate observation, both Dex and yeast cells were fluorescently labeled before emulsion formation and analyzed via laser confocal microscopy (Figure 4). The Dex phase appears fluorescent green, the yeast cells are blue, and the PEG phase remains unlabeled (black). Notably, the yeast cells were exclusively localized within Dex droplets, with no diffusion into the PEG phase, achieving 100% encapsulation efficiency in the dispersed phase. The sharp interfacial confinement further confirms the emulsion’s effectiveness in cell retention.

These findings demonstrate that w/w emulsions are a robust platform for fabricating porous materials with high-efficiency in situ cell immobilization, eliminating the need for post-synthesis loading. The PEGDA polymerization was initiated in the continuous phase of the Dex/PEG emulsion, yielding PPEG-15 embedded with yeast (yeast@PPEG-15). As illustrated in Figure 5, the yeast cells are distributed within the porous structure, exhibiting intact morphological integrity. The in situ encapsulation process successfully preserved cell viability and structure, with the muffle furnace thermogravimetric analysis confirming a near-100% encapsulation rate. These results demonstrate the w/w emulsion-templating method’s efficacy as a robust, green, and high-efficiency platform for yeast immobilization that eliminates the need for an organic solvent and post-synthesis loading.

2.3. Effect of Temperature and pH on Fermentation

Glucose serves as a key substrate in the modern green production of bioethanol through fermentation [35,36]. Given the significant influence of fermentation temperature and pH on yeast-catalyzed glucose-to-ethanol conversion, we evaluated the fermentation efficiency of yeast@PPEG-15 under varying conditions.

As shown in Figure 6a, fermentation efficiency initially increases with pH, peaking at pH 5.5, before declining at higher pH values (5.5–6.5). Similarly, Figure 6b reveals that ethanol yield follows a bell-shaped trend with temperature (24–36 °C), reaching its maximum at 33 °C. Thus, the optimal conditions for immobilized yeast fermentation in our w/w emulsion-templated porous materials are pH 5.5 and 33 °C.

Notably, under these optimized conditions (pH 5.0, 33 °C), the ethanol yield of yeast@PPEG-15 was 71.8%, significantly higher than Ca-alginate immobilized yeasts that are used for ethanol production, aiming for repeated batches (ethanol yield at 46% for the first batches, and 38% for the fifth batch) [37]. The ethanol yield of yeast@PPEG-15 was only 2.8% lower than that of free yeast cells (74.6%). This minor reduction compared to the free cells—attributable to both mass transfer limitations in immobilized systems and reduced cell activity caused by cell immobilization—is consistent with prior yeast immobilization studies [35,38]. Critically, our w/w emulsion-templating method achieves near-perfect encapsulation efficiency (~100%) with only a minor (<3%) loss in catalytic fermentation efficiency. These results demonstrate that this approach is a highly efficient and environmentally benign strategy for cell immobilization in bioethanol production. Building on recent studies demonstrating near-100% substrate conversion using recombinant yeast cells in Ca alginate [39], our future work will employ more efficient yeast strains to further enhance the fermentation efficiency of this immobilization system.

2.4. Reusability and Storability of Immobilized Yeast

Compared to free cells, immobilized cells offer superior reusability, a critical advantage for industrial applications. As demonstrated in Figure 7a, yeast@PPEG-15 exhibits excellent cyclic fermentation performance. During the first three cycles, the ethanol yields gradually decreased by a small margin, then stabilized after the fourth cycle. This trend can be attributed to the gradual detachment of loosely bound yeast cells from the material surface during initial cycles.

To enhance contact between the immobilized yeast and culture medium, we cut yeast@PPEG-15 into 5 mm cubes, exposing yeast cells in the void of the material’s cross-section. In early cycles, yeast near these exposed surfaces desorbed, leading to a minor decline in ethanol yield. However, after multiple cycles, the remaining yeast cells were firmly retained within the carrier, resulting in stable fermentation performance. This confirms the robust immobilization of yeast without cell escape during fermentation.

Remarkably, by the seventh cycle, the ethanol yield remained at ~85% of the initial value, significantly outperforming traditional o/w emulsion-templated materials (which typically retain only ~70% yield under similar conditions) [18]. These results highlight the superior continuous fermentation capability of our w/w emulsion-templated immobilization system.

The storage stability of immobilized yeast systems is a crucial performance parameter, as these biocatalysts are frequently required to remain viable and active during periods between production and application. As shown in Figure 7b, yeast@PPEG-15 demonstrates excellent storage performance. While fermentation yield gradually decreases with storage time, the decline is significantly smaller compared to free yeast cells.

This superior stability can be attributed to the material’s unique properties: although its dense pore structure slightly hinders mass transfer, it significantly enhances the water retention capacity. The material effectively absorbs and preserves the culture medium, providing more stable environmental conditions for yeast during storage. Consequently, the presence or absence of culture medium during storage has minimal impact on immobilized yeast performance.

Remarkably, after four weeks of storage, the immobilized system retained over 95% of its initial ethanol production capacity, with high yeast viability. This performance far surpasses that of free cells, which showed only 81% activity retention under identical conditions. These results underscore that our in situ immobilization method yields cell carriers with exceptional storage stability, making them ideally suited for practical applications involving delayed use.

3. Experimental Section

3.1. Materials

Polyethylene glycol diacrylate (PEGDA, M_n = 400), polyethylene glycol (PEG-20k, M_n = 20,000), and dextran (Dex, M_n = 70,000) were purchased from Aladdin (Shanghai, China). Fluorescein isothiocyanate dextran (FITC-dextran, M_w = 10,000) was obtained from Perfemiker Co., Ltd. (Shanghai, China). Nanocrystalline cellulose (CNC) was supplied by Shansi Technology Co., Ltd. (Suzhou, China). The following chemicals were acquired from Shanghai Titan Scientific Co., Ltd. (Shanghai, China): potassium persulfate (KPS, 99.5%), Tetramethylethylenediamine (TMEDA, 99%), YPD liquid medium (a nutrient-rich “workhorse” broth for growing yeast in a laboratory setting), D-Glucose anhydrous (99%), sodium chloride (NaCl, 99.5%), potassium chloride (KCl, 99.5%), disodium hydrogen phosphate dodecahydrate (Na₂HPO₄·12H₂O, 99%), potassium dihydrogen phosphate (KH₂PO₄, 99.5%), and hydrochloric acid (HCl, 36.0–38.0%). Dry yeast powder (A200) was obtained from Angel Yeast Co., Ltd. (Yichang, China). DAPI staining reagent was purchased from Sevilla Biotech Co., Ltd. (Wuhan, China). Deionized water was prepared by secondary distillation in the laboratory. KPS was purified by recrystallization prior to use.

3.2. Preparation of Porous PEG (PPEG)

To prepare PEG-based porous materials, a water-in-water (w/w) emulsion was prepared as a template [28]. PEGDA aqueous solutions (10–25% w/w) were first prepared by dissolving PEGDA into 6 mL deionized water containing 0.8 g PEG-20k and 0.05 g CNC. A 20% w/w dextran solution was separately prepared by dissolving 0.8 g Dex in 4 mL deionized water, which was then slowly added to the PEGDA solutions under 500 rpm magnetic stirring for 10 min to form Dex/PEG w/w emulsions, which were named as EPEG-x (where x = 10, 15, 20, or 25, representing the PEGDA concentration%).

For polymerization, KPS (6 wt% of PEGDA) was dissolved in the PEG solutions before emulsion formation, followed by addition of TMEDA (1 wt% of PEGDA) to the freshly prepared emulsions, which were immediately transferred to PTFE molds and polymerized at 35 °C in an oven for 12 h. The resulting porous hydrogels were demolded and washed in deionized water using an orbital shaker for 24 h, with the water changed every eight hours. These samples were designated as PPEG-x, where x = 10, 15, 20, or 25, representing the PEGDA concentration (%).

3.3. Preparation of Yeast@PPEG

The yeast immobilization was performed by first activating dry yeast in glucose solution, followed by centrifugation to collect the active yeast cells as described in the previous research [18,34]. Approximately 0.8 g of the harvested yeast cells were then dispersed in the prepared 20% (w/w) Dex aqueous solution of 4 mL. This yeast-containing Dex solution containing yeast of 0.2 g/mL served as the internal phase for preparing yeast-embedded porous PEG (yeast@PPEG) through the same w/w emulsion-templating method used for PPEG preparation.

3.4. Characterization of Emulsion-Template, PPEG, and Yeast@PPEG

The morphology of the w/w emulsion was analyzed using a Nikon A1R laser confocal microscope (Nikon, Japan). To visualize the Dex phase, FITC-dextran was dissolved in the Dex aqueous solution prior to emulsion preparation. The freshly prepared Dex/PEG emulsion was dropped onto a 20 mm glass-bottom confocal dish and immediately imaged. ImageJ software (Version 1.54g) was used to analyze the droplet size.

For yeast immobilization analysis, the active yeast cells were first stained with DAPI. The staining protocol involved (1) washing activated yeast cells three times with PBS buffer (5 min each wash); (2) incubating cells with ready-to-use DAPI staining reagent (sufficient to fully cover cells) in the dark at room temperature for 8 min; and (3) washing three times with PBS buffer (5 min each wash) to remove excess stain. The PBS buffer (0.01 M, pH 7.4) was prepared by dissolving 8.00 g NaCl, 0.20 g KCl, 3.63 g Na₂HPO₄·12H₂O, and 0.24 g KH₂PO₄ in 800 mL deionized water, adjusting the pH with HCl, and diluting to 1 L. The DAPI-stained yeast cells were dispersed in FITC-labeled Dex solution, and the emulsion was imaged via a Nikon A1R laser confocal microscope (Japan).

The porous hydrogels (PPEG and yeast@PPEG) were frozen for 24 h and lyophilized for 24 h to obtain dry samples. The dry samples were sectioned with a surgical knife, placed on a SEM stub with the cut face up, mounted on the stub with conductive tape, and sputter-coated with gold. The pore morphology and yeast distribution were examined with a Hitachi S4800 SEM (Japan). ImageJ was used to quantify the void size, with at least 100 voids measured according to the previous research [40,41].

The yeast mass on the yeast@PPEG was determined by combustion at 1000 °C in air using a muffle furnace. The resulting ash, constituting an inorganic component of the yeast cells (0.0315 g ash/g cells) [38], was weighed. The yeast content was then calculated by dividing the yeast mass (determined from the ash mass) by the total mass of the yeast@PPEG.

The compressive properties of hydrated porous materials (PPEG-10, PPEG-15, PPEG-20, and PPEG-25) were evaluated using a LABSANS LD23.104 universal testing machine (Shenzhen, China). Samples were tested immediately after polymerization and washing (without drying/rehydration). The compression tests were performed at a constant crosshead speed of 1 mm/min. Testing was terminated upon sample rupture, defined as the point of catastrophic structural failure where the stress–strain curve showed a sudden drop in load-bearing capacity. The maximum compressive strength was recorded at the point immediately preceding rupture. Each PPEG polyHIPE sample was tested five times to ensure the accuracy of the compression test results.

3.5. Fermentation Performance Evaluation

Yeast@PPEG-15 (containing 0.8 g yeast) was cut into cubes with a side length of 5 mm, and was then immersed in a 50 mL sterile YPD medium (20 g/L glucose) in sealed conical flasks. Fermentation was conducted in a shaking incubator at varied temperatures (24–36 °C), with different pH values (4.5–6.5), and at 120 rpm for 12 h. Each condition was tested in quintuplicate, with free yeast (0.8 g) serving as the control.

Ethanol production was quantified by gas chromatography (GC, 7890B + 7697A, American Agilent, Santa Clara, CA, USA) using the following protocol [18]: a standard curve was prepared by diluting anhydrous ethanol in deionized water to create solutions of 1, 2, 4, 6, and 10 mg/mL concentrations, which were analyzed using GC with headspace sampling (ethanol retention time: 3.049 min) (Table S1); the resulting peak areas were plotted against concentration to generate a linear calibration curve (Figure S1), which was then used to calculate ethanol concentrations in fermentation samples based on their GC peak areas, with all measurements performed in triplicate for accuracy.

To investigate the cycle fermentation performance, yeast@PPEG-15 underwent seven consecutive 12 h fermentation cycles in fresh YPD medium (pH 5.5, 33 °C, 120 rpm). Between cycles, samples were rinsed with deionized water. Ethanol production was monitored throughout all cycles.

To make the storage stability assessment, two sample groups (n = 4 each) of yeast@PPEG-15 were prepared: Group A: pre-fermented (6 h in YPD) before storage and Group B: direct storage. The samples were refrigerated and tested weekly for 4 weeks by measuring ethanol production after fermentation. Parallel tests with refrigerated free yeast served as controls.

4. Conclusions

We have developed an efficient water-in-water (w/w) emulsion-templating method for in situ yeast immobilization, producing materials with exceptional fermentation activity, storage stability (maintaining > 95% ethanol production after 4 weeks), and reusability (85% activity retention after 7 cycles). By employing cellulose nanocrystal (CNC) stabilizers in Dex/PEG emulsions and varying PEGDA concentration (10–25 wt%), we achieved pore size control, with void size tunable from 10.4 to 6.6 μm and interconnected pores from 2.2 to 1.4 μm. The optimized PPEG-15 carrier (9.6 μm voids, 1.8 μm interconnected pores) demonstrated robust mechanical properties (0.8–1.04 MPa compressive strength, >50% compressibility) ideal for yeast encapsulation (2–5 μm cells), as confirmed by laser confocal microscopy and SEM imaging. The immobilized yeast exhibited peak fermentation efficiency at pH 5.5 and 33 °C, combining the material’s excellent water retention with stable microenvironmental conditions. This simple, green immobilization strategy represents a significant advance in cell immobilization technology, offering tunable porosity, high encapsulation efficiency, and outstanding operational stability for industrial biocatalysis applications.