Algae oil contains polyunsaturated fatty acids (PUFAs) that are beneficial to human health and development. Enzymatic refining is a sustainable approach that can improve the quality and nutritional value of algae oil. However, this approach involves the use of specific enzymes to remove unwanted impurities such as phospholipids (PLs) and free fatty acids (FFAs). This study proposes a novel immobilized phospholipase, immobilized phospholipase A1 (PLA1@MCM-41-C8), which serves a dual purpose: it is effective for both degumming arachidonic acid (ARA)-enriched algal oil and its subsequent deacidification. The use of PLA1@MCM-41-C8 achieved a degumming rate of 95.9% for crude algal oil, with an ARA retention rate of up to 97.3%. The acidity of the degummed oil decreased from 14.5 to 1.1 mg KOH·g−1, while simultaneously producing 29.2% high-value diacylglycerol (DAG). By comparing enzymatically degummed and deacidified ARA-enriched algae oil (EDDO) with crude ARA-enriched algae oil (CO), commercialized refining algae oil (CRO), it was found that enzymatic refining had minimal impact on antioxidant stability, fatty acid composition, and oil content. In particular, the flavor characteristics of algal oil before and after enzymatic refining with PLA1@MCM-41-C8 remained unchanged and were dominated by the main volatile compounds: alcohols and aldehydes. Compared with free phospholipase A1 (PLA1), PLA1@MCM-41-C8 exhibited significantly improved thermostability, pH tolerance, solvent tolerance, and long-term reusability. These characteristics make it a promising candidate for complex oil refining in industrial applications.
Algae oil is derived from oleaginous microorganisms that rely on prokaryotic or eukaryotic photosynthesis and is minimally affected by seasonal or climatic changes [1], [2]. Notable sources of algae oil include species such as Schizochytrium sp., Ulkenia amoeboida, Crypthecodinium cohnii, and Mortierella alpina (M. alpina). Compared to plant- and animal-based lipids, algae oil has a much shorter production cycle and requires no agricultural land or freshwater. Algae oil is a rich source of polyunsaturated fatty acids (PUFAs), such as docosahexaenoic acid (DHA), arachidonic acid (ARA), and eicosapentaenoic acid (EPA), which have been shown to lower blood lipid levels, boost the immune system, and promote infant brain and visual development [3], [4]. Although fish oil and Antarctic krill oil are also sources of PUFAs, their production is increasingly affected by pollution, particularly heavy metal accumulation and microplastic contamination in the oceans [5], [6]. Hence, algae oil is increasingly recognized as a sustainable, economical, and functional lipid. It has gained significant attention in the food and healthcare industries due to its potential health benefits and versatile applications.
Currently, the algae-to-oil process involves multiple steps, including cultivation, harvesting/dewatering, processing, conversion, oil extraction, and refining [7]. During oil refining, degumming and deacidification are essential steps for removing phospholipids (PLs) and free fatty acids (FFAs), respectively, in order to preserve the nutritional value and quality of the oil. Traditional degumming can be classified into water and acid degumming, depending on the hydratability of the PLs [8]. Water degumming removes hydratable PLs, which absorb water to become hydrophilic, aggregate into a sticky gel-like mass (i.e., gums), and are then separated by centrifugation or by allowing the solids to settle under gravity [9]. Acid degumming involves adding phosphoric, citric, or malic acids at high temperatures to convert non-hydratable PLs into hydratable ones [10]. Both methods consume large amounts of freshwater and generate substantial wastewater, which can negatively impact the environment.
Traditional deacidification methods include alkali neutralization, physical distillation, and solvent extraction [11]. For instance, alkali refining uses sodium hydroxide or potassium hydroxide to saponify FFAs and reduce the acid value (AV) of the oil. However, the resulting soap micelles can absorb neutral oils and micronutrients such as phytosterols and vitamin E, leading to yield losses [12]. Physical distillation and solvent extraction are further constrained by challenges such as high energy consumption, toxic solvent residues, and environmental waste emissions [13].
Edible oil refining using biocatalytic enzymes is emerging as an alternative to chemical catalysis due to its high selectivity, mild reaction conditions, and carbon-neutral characteristics [14]. Enzymatic degumming involves the hydrolysis of PLs by phospholipases, while enzymatic deacidification uses lipases to convert FFAs into fatty acid esters [15], [16]. For example, Ye et al. [17] reported that enzymatic degumming using phospholipase C (PLC) reduced the phospholipid content in rapeseed oil (RO) from 690.0 to 7.3 mg·kg−1 in 1.5 h, compared to water degumming, which decreased the phosphorus content by only 54.2 mg·kg−1. Qu et al. [18] also found that phospholipase A2 reduced the phosphorus content in soybean oil (SO) from 142.4 to 9.8 mg·kg−1 within 5.0 h. Enzymatic deacidification involves lipase-catalyzed esterification between FFAs and acyl acceptors containing hydroxyl groups, such as phytosterols, monoglycerides (MAGs), and glycerol. This process results in the selective conversion of FFAs into phytosterol esters, diacylglycerols (DAGs), and other functional lipids.
However, the application of enzymatic degumming and deacidification in edible oil processing is hindered by the low catalytic stability, poor reusability, and narrow substrate specificity of enzymes. Although various enzyme immobilization techniques, such as physical adsorption, encapsulation, covalent bonding, and crosslinking—have been used to attach free enzymes to solid carriers like silica gel and microporous resins, challenges such as insufficient mechanical strength and low organic solvent tolerance remain unresolved [19]. To overcome these issues, several types of porous particles have been proposed, including metal-organic frameworks (MOFs), covalent organic frameworks (COFs), multi-shelled hollow spheres, and magnetic microspheres. However, the complex, time-consuming preparation procedures and low yields of these carriers limit their suitability for industrial applications [20], [21]. Mesoporous silica of Mobil Composition of Matter No. 41 (MCM-41), by contrast, has been produced on a large scale and is widely applied in petrochemical, pharmaceutical, and other industrial fields [22], [23]. With its hexagonally ordered pores (2-50 nm in diameter) and specific surface area of 800-1400 g·m−2, MCM-41 provides abundant sites for enzyme immobilization and subsequent catalytic reactions. This suggests that MCM-41 holds great potential for enzymatic applications [24]. Our previous study found that phospholipase A1 (PLA1) could not only catalyze the hydrolysis of PLs but also enhance the removal of FFAs, whereas traditional enzymatic degumming and deacidification typically require different types of enzymes. Hence, it was proposed that the immobilization of PLA1 on MCM-41 could facilitate both oil degumming and deacidification. This approach simplifies enzymatic refining procedures and reduces production costs.
In this study, a novel “one stone for two birds” strategy was proposed for the enzymatic degumming and deacidification of ARA-enriched algal oil using immobilized PLA1 based on hydrophobically functionalized MCM-41. Immobilized phospholipase A1 (PLA1@MCM-41-C8) was prepared, characterized, and applied in enzymatic refining. The composition, flavor characteristics, and antioxidant properties of algae oil before and after enzymatic treatment were compared. Additionally, the reaction kinetics of enzymatic deacidification using PLA1@MCM-41-C8 were investigated through molecular docking simulations. This work provides a green and efficient refining approach for ARA-enriched algal oil based on immobilized phospholipase, demonstrating both cost-effectiveness and time-saving potential for future oil processing.
2. Materials and methods
2.1. Materials
Crude ARA-enriched algae oil (CO), produced from M. alpina, was provided by CABIO Biotech Co., Ltd. (China). The initial phosphorus content was 321.1 mg·kg−1, and the original AV was 12.8 mg KOH·g−1. Tea seed oil (TSO, phosphorus content: 40.4 mg·kg−1; AV: 5.5 mg KOH·g−1), peanut oil (PO, phosphorus content of 277.6 mg·kg−1 and original AV of 1.9 mg KOH·g−1), SO (phosphorus content of 304.1 mg·kg−1 and original AV of 7.9 mg KOH·g−1), RO (phosphorus content of 300.6 mg·kg−1 and original AV of 5.7 mg KOH·g−1), rice bran oil (RBO, phosphorus content of 142.4 mg·kg−1 and original AV of 19.2 mg KOH·g−1), walnut oil (WO, phosphorus content of 155.9 mg·kg−1 and original AV of 1.9 mg KOH·g−1), and flaxseed oil (FO, phosphorus content of 16.5 mg·kg−1 and original AV of 4.8 mg KOH·g−1) were obtained from the Oil Crops Research Institute of the Chinese Academy of Agricultural Sciences. MCM-41 was purchased from Jiangyin Nanda Synthetic Chemical Co., Ltd. (China). PLA1 was obtained from Novozymes (Denmark). Phospholipase B (PLB) and phospholipase D (PLD) were purchased from Cangzhou Sunson Enzyme Biotechnology Co., Ltd. (China), and PLC was provided by Shanghai Macklin Biochemical Technology Co., Ltd. (China).
The bicinchoninic acid (BCA) Protein Assay Kit was purchased from Shanghai Yuanye Bio-Technology Co., Ltd. (China). Glycerol, oleic acid, isopropanol, ligroin (boiling point: 30-60 °C), n-hexane, phenolphthalein, citric acid, 95% ethanol, n-octyltrichlorosilane (C8), and other chemicals were obtained from Aladdin Reagent Co., Ltd. (China). All the reagents and solvents used in the experiments were of analytical, chromatographic, or biological grade. Ultrapure water was obtained using Milli-Q water and a Milli-Q apparatus (Millipore, USA).
2.2. Preparation of immobilized phospholipase
For phospholipase immobilization, mesoporous carriers (i.e., MCM-22, MCM-41, and MCM-49) were hydrophobically functionalized using the silane coupling agent C8, following the method described by Liu et al. [24]. The resulting functionalized carriers—MCM-22-C8, MCM-41-C8, and MCM-49-C8—were then used to immobilize free PLA1. Briefly, a specific amount of PLA1 was added to a phosphate buffer (50 mmol·L−1, pH 5-9) and mixed thoroughly. Separately, 0.1 g of each carrier was prewetted with 200 μL of pure ethanol and then added to 1-10 mL of the prepared PLA1 solution. The mixture was incubated in a shaking incubator (THZ-C; Taicang Qiangle Experimental Equipment Co., Ltd., China) at 30 °C and 220 r·min−1 for 40 min. After incubation, the suspension was centrifuged at 2000g for 10 min at 4 °C using a refrigerated centrifuge (Allegra X-12; Beckman, USA). The resulting supernatant was collected and analyzed for residual protein content using a BCA protein assay. The precipitates, corresponding to PLA1@MCM-41-C8, were collected and washed at least three times with phosphate buffer (50 mmol·L−1, pH 5-9), then freeze-dried using a vacuum freeze-drying machine (YTLG-18A; Shanghai Yetuo, China) at −70 °C for 24 h, to obtain the final PLA1@MCM-41-C8 product.
2.3. Characterization of MCM-41, MCM-41-C8, and PLA1@MCM-41-C8
Scanning electron microscopy (SEM; S-4800; Hitachi, Japan) and transmission electron microscopy (TEM; TECNAI G2 20S-TWIN; FEI, USA) were used to characterize the surface morphology and the internal microstructure of the solid samples. A computer-controlled nitrogen gas adsorption analyzer (ASAP 2010; Micromeritics, USA) was used to measure the surface area and total pore volume of MCM-41, MCM-41-C8, and PLA1@MCM-41-C8 within a relative pressure range of 0.05-1.00. Relative pressure was calculated as the ratio of the partial pressure of nitrogen to the saturated vapor pressure of liquid nitrogen at the adsorption temperature. The hydrophobicity of the samples was assessed using a contact angle goniometer (SDC-200S; SINDIN, China), the hydrophobicity of the samples was assessed by measuring surface water contact angles. Fourier-transform infrared (FT-IR) spectroscopy (Bruker, Germany) was used to obtain the FT-IR spectra of MCM-41, MCM-41-C8, PLA1@MCM-41-C8, and free PLA1. Additionally, the secondary structures of different phospholipase forms—including free PLA1, freshly prepared PLA1@MCM-41-C8, reused PLA1@MCM-41-C8, and deactivated PLA1@MCM-41-C8—were analyzed using FT-IR in the wavenumber range of 4000-400 cm−1.
Thermogravimetric (TG) analyses of free PLA1, MCM-41-C8, and PLA1@MCM-41-C8 were performed using a TG analyzer (Netzsch Thermo Gravimetric Analyzer, Germany).
The protein content of PLA1 before and after enzyme immobilization was determined using the BCA method [25]. Residual protein concentration in the phospholipase solution was calculated by:
where C0 and C are the protein concentrations (mg·mL−1) before and after PLA1 immobilization, respectively; V′ is the volume (mL) of the phospholipase solution; and M is the mass (g) of MCM-41-C8.
2.4. Enzyme activity of free PLA1 and PLA1@MCM-41-C8
The enzymatic activities of free PLA1 and PLA1@MCM-41-C8 were determined using the method reported by Si et al. [20]. Briefly, 50 g·L−1 of defatted soybean lecithin and a 4% (v/v) polyvinyl alcohol solution were emulsified at a volume ratio of 1:4 by conventional stirring at 200 r·min−1 for 10 min. Then, 4 mL of the prepared emulsion was mixed with 5 mL of 0.01 mol·L−1 citric acid buffer (pH 5.0) and 1 mL of a 100-fold diluted enzyme solution. After heating in a water bath at 37 °C for 10 min, 15 mL of 95% ethanol was added to terminate the reaction. The released fatty acids were titrated with 0.05 mol·L−1 sodium hydroxide.
The enzyme activity of PLA1 was calculated using:
where X represents the enzyme activity of the sample (U·g−1), V1 is the volume of NaOH standard solution consumed in the titration of samples (mL), V0 is the volume of NaOH standard solution consumed in the titration blank (mL), c is the concentration of NaOH standard solution (mol·L−1), t is the reaction time for enzyme activity determination (min) and n is the dilution multiple.
2.5. Enzymatic refining by PLA1@MCM-41-C8
Enzymatic refining of CO, comprising both degumming and deacidification, was conducted using the immobilized enzyme PLA1@MCM-41-C8. For degumming, 45.0 g of CO was first heated to 70 °C. Then, 0.2 mL of citric acid solution (45%, w/w) was added under stirring at 10 000 r·min−1 for 1 min. The mixture was then allowed to react further at 200 r·min−1 and 70 °C for 3 h before being cooled to 45 °C. Next, a 4% (w/w) NaOH solution was added to adjust the pH to 5.8. Following this, PLA1@MCM-41-C8 was added at concentrations ranging from 0.1% to 0.9% (w/w), and the mixture was maintained at a predetermined reaction temperature (15-55 °C) for 3 h for enzymatic degumming. During this period, 5-6 g of oil samples were collected at regular intervals for further analysis. Finally, the reaction mixture was centrifuged at 4863g for 10 min at 4 °C to recover both the immobilized enzyme and the supernatant (degummed oil) for further analysis. For enzymatic deacidification, the degummed algal oil was mixed with glycerol and a 4 Å molecular sieve at temperatures ranging from 25 to 75 °C. Glycerol served as the acyl receptor, while the molecular sieve adsorbed water generated during enzymatic esterification [24]. The immobilized PLA1@MCM-41-C8 recovered from the degumming step was then reintroduced into the preheated mixture to catalyze the esterification of FFAs with glycerol. Oil samples (1-2 g) were collected at regular intervals from each reaction for further analysis.
2.6. AV, peroxide value (PV), and phosphorus content determination
The AV and PV of the oils, both before and after enzymatic refining, were determined according to the official American Oil Chemists’ Society (AOCS) methods Cd 3d-63 and Cd 8-53, respectively [26]. The antioxidant capacities of different oils—including CO, commercialized refining algae oil (CRO), enzymatically degummed ARA-enriched algae oil (EDO), enzymatically degummed and deacidified ARA-enriched algae oil (EDDO), and alkali deacidified ARA-enriched algae oil (ADO)—were evaluated based on their oxidation induction times, measured using a Rancimat oxidation stability analyzer (Model 743; Herisau, Switzerland). Phosphorus content in the algal oil samples was determined according to the method reported by Xie and Dunford [27]. Detailed experimental procedures are provided in Appendix A.
2.7. Kinetic simulation of enzymatic deacidification
The Michaelis-Menten kinetic model was used to determine the kinetic parameters of the enzymatic deacidification of algal oil catalyzed by PLA1@MCM-41-C8 [28]. With glycerol concentration ranged from 0.05 to 0.6 mol·mL−1, enzymatic reactions were conducted using PLA1@MCM-41-C8. The Lineweaver-Burk double-reciprocal method was employed to plot the reciprocal values of reaction rates and substrate concentrations. The Michaelis constant (Km, mol·L−1) and the maximum reaction rate (Vm, mol·L−1·min−1) were subsequently calculated:
where V4 represents the reaction rate (mol·L−1·min−1) and S is the substrate concentration (mol·L−1).
The Arrhenius equation was applied to estimate the activation energy (Ea, kJ·mol−1) and the pre-exponential factor (A) of the esterification reaction over a temperature range of 15-35 °C [29]. The corresponding parameters were calculated as described in the following equation:
where k is the reaction rate constant (min−1), R is the universal gas constant (8.314 J·mol−1·K−1), and T is the absolute temperature (K). The relationship between the reaction rate and reactant concentration (A1) is shown in Appendix A.
2.8. Oil content, composition, and flavor characterization
The flavor compounds in the algal oil were analyzed using solid-phase microextraction (SPME) coupled with gas chromatography-mass spectrometry (GC-MS) [30]. Detailed procedures are provided in Appendix A. The contents and compositions of algal oil were analyzed using a gas chromatography system (GC-2030; Shimadzu, Japan), equipped with a fused silica capillary column (DB-5 HT, 15 m × 320 µm × 0.25 µm; Agilent Technologies, USA) and a flame ionization detector to determine the composition of the different oils [16]. Nitrogen was used as the carrier gas at a flow rate of 2 mL·min−1. The overall gas flow rate was 2 mL·min−1, with nitrogen serving as the carrier gas. Injector and detector temperatures were maintained at 320 and 350 °C, respectively. The oven temperature was programmed from an initial temperature of 170 °C (held initially), held for a specified duration, and then increased according to a defined temperature ramp. The split ratio was 50:1. Besides, the fatty acid composition was detected by the Agilent 7890A gas chromatography system (Agilent Technologies, USA) on a DB-FFAP column (30 m × 0.25 mm, 0.25 μm; Agilent Technologies) with the specific procedures described in Appendix A.
2.9. Molecular docking analysis
Molecular docking analysis was performed using AutoDock Vina to investigate the interaction mechanism between PLA1 and glycerol or oleic acid. The crystal structure of PLA1 (Protein Data Bank (PDB) ID: 4HYQ) was obtained from the protein database PDB.34 [31]. Three-dimensional (3D) structures of oleic acid and glycerol were obtained from the PubChem database. ChemDraw 20.0 was used for energy minimization of the small molecular structures. The Lamarckian genetic algorithm (LGA) was employed as the docking algorithm. The conformation with the lowest docking energy was selected as the most favorable binding configuration for both oleic acid and glycerol.
3. Results and discussion
3.1. Enzyme immobilization and optimization
To identify the optimal free enzyme and mesoporous carrier for enzyme immobilization, various phospholipases (PLA1, PLB, PLC, and PLD) and mesoporous molecular sieves (MCM-22, MCM-41, and MCM-49) were evaluated based on enzyme loading capacity, immobilization efficiency, and degumming performance of the corresponding immobilized enzymes. As shown in Fig. 1(a), PLA1 achieved the highest degumming rate, effectively removing 84.0% of PLs. In comparison, PLB, PLC, and PLD showed significantly lower degumming efficiencies of 71.7%, 47.2%, and 58.3%, respectively. All values were significantly lower than that achieved by PLA1 under the same conditions. These findings are consistent with the results reported by Jiang et al. [32], who found that PLA1 removed 60% of PLs from SO, compared to only 27.2% using PLC. The superior performance of PLA1 is attributed to its ability to hydrolyze the sn-1 ester bond of PLs, facilitating the conversion of non-hydratable PLs into hydratable forms [33]. In contrast, PLC targets the glycerophosphate bond at the C3 position of glyceroPLs, which may limit its degumming effectiveness. Therefore, because of its superior phospholipid hydrolysis ability, PLA1 was selected as the ideal biocatalyst for algal oil degumming in this study.
To further evaluate the effect of hydrophobic mesoporous carriers on enzyme immobilization, MCM-22-C8, MCM-41-C8, and MCM-49-C8 were assessed based on PLA1 loading capacity, immobilization efficiency, and enzymatic activity. As shown in Fig. 1(b), the PLA1 loading amounts on MCM-22-C8, MCM-41-C8, and MCM-49-C8 were 183.5, 189.5, and 155.7 mg·g−1, respectively. The corresponding immobilization efficiencies were 14.8%, 18.1%, and 13.0%, respectively.
As a result, the enzymatic activities of immobilized PLA1 reached 894.0 (MCM-22-C8), 1113.0 (MCM-41-C8), and 654.0 U·g−1 (MCM-49-C8). The superior PLA1 immobilization and catalytic activity observed with MCM-41-C8 can be attributed to its favorable internal structure and large carrier pore size [34]. These findings suggest that MCM-41-C8 is more suitable for PLA1 immobilization than MCM-22-C8 and MCM-49-C8, as further characterized and validated in Section 3.2.
To further optimize the catalytic efficiency (Vm/Km) of phospholipase immobilized using the selected combination of PLA1 and MCM-41-C8, two key factors were investigated: the MCM-41-C8/PLA1 ratio (g·mL−1) and the pH of the PLA1 solution. As shown in Fig. 1(c), increasing the MCM-41-C8/PLA1 ratio from 1:10 to 1:50 led to a steady increase in PLA1 loading, from 42.0 to 138.3 mg·g−1. This increase in the solid-liquid ratio promoted greater interaction between phospholipase and the carrier material, allowing more enzymes to enter the mesopores and thereby enhancing immobilization efficiency. However, the immobilization rate of PLA1 initially decreased from 16.0% to 8.6% as the solid-to-liquid ratio increased from 1:10 to 1:30, then fluctuated between 10.5% and 11.7% as the ratio increased further to 1:50. This trend is attributed to the saturation of available immobilization sites caused by the excess amount of enzyme solution [35]. Considering both PLA1 loading capacity and immobilization efficiency, an optimal MCM-41-C8/PLA1 ratio of 1:50 was selected for subsequent experiments.
The effect of the enzyme solution pH on PLA1 immobilization was also investigated. Fig. 1(d) shows that the PLA1 loading initially increased from 122.7 to 190.8 mg·g−1 as the solution pH rose from 5.0 to 7.0, followed by a continuous decline to 88.6 mg·g−1 as the pH increased further to 9.0. A similar trend was observed for both immobilization efficiency and enzymatic activity. Both reached their peaks at pH 7.0, with immobilization efficiency at 18.3% and enzyme activity at 1 230.0 U·g−1. These results indicate that neutral conditions (pH 7.0) enhance both PLA1 immobilization and enzymatic activity, whereas acidic or alkaline environments adversely affect the process [36]. Therefore, pH 7.0 was selected as the optimal condition for PLA1 immobilization.
3.2. Characterization of PLA1@MCM-41-C8
SEM and TEM were used to analyze the morphological and structural characteristics of MCM-41. As shown in Figs. 2(a) and (b), MCM-41 appeared as irregular and non-uniform spherical clusters. However, its tightly packed and evenly distributed hexagonal mesopores revealed a uniformly layered structure with fingerprint-like surface patterns [37]. Surface modification with C8 increased the water contact angle of MCM-41 from 21.6° to 117.4°, indicating successful hydrophobic functionalization that facilitated PLA1 immobilization (Fig. 2(c)). Thermogravimetric analysis (TGA) of MCM-41, free PLA1, and PLA1@MCM-41-C8 showed distinct weight loss patterns over the temperature range of 30-800 °C (Fig. 2(d)). MCM-41 exhibited a minor mass loss of 5.1%, indicating high thermal stability of the silica-based carrier. In contrast, free PLA1 showed significant weight loss—52.5% between 30 and 200 °C—primarily due to water evaporation, as PLA1 is a high-moisture enzyme that retains substantial water within its structure [38]. An additional 44.5% weight loss between 200 and 350 °C was attributed to the thermal decomposition of the organic phospholipase [39]. Compared to free PLA1, PLA1@MCM-41-C8 exhibited mass losses of 10.5%, 49.6%, and 14.4% in the temperature ranges of 30-200, 200-350, and 350-800 °C, respectively. These correspond to the loss of adsorbed water, dissociation of phospholipase, and removal of structural water at higher temperatures [40], [41]. The consistent weight loss observed between 200 and 800 °C confirms the successful immobilization of PLA1 and indicates that immobilization on MCM-41-C8 enhances the thermostability of the enzyme compared to its free form.
The nitrogen adsorption and desorption isotherms (Figs. 2(e) and (f)) reveal that both MCM-41 and MCM-41-C8 exhibit type-IV isotherms with H1 hysteresis loops—typical features of mesoporous materials [42]. Detailed textural parameters for MCM-41, MCM-41-C8, and PLA1@MCM-41-C8 obtained from Brunauer-Emmett-Teller (BET) analysis are presented in Table S1 in Appendix A. Notably, the specific surface area of MCM-41 decreased significantly from 1225.4 to 419.4 m2·g−1 after C8 modification, indicating successful surface grafting. Following enzyme immobilization, the surface area of PLA1@MCM-41-C8 further decreased to 64.8 m2·g−1, accompanied by a reduction in pore volume from 0.8 to 0.06 cm3·g−1, confirming the successful loading of PLA1 into the mesopores.
Additionally, FT-IR spectroscopy was used to identify the functional groups present in the carrier and immobilized enzyme. As shown in Fig. 2(g), the characteristic peaks at 2925 and 2854 cm−1 correspond to the asymmetric and symmetric stretching vibrations of −C-H bonds, respectively. A distinct peak at 1541 cm−1 observed in the spectrum of PLA1@MCM-41-C8 is attributed to −N-H bending vibrations, further confirming enzyme immobilization.
X-ray photoelectron spectroscopy (XPS) analysis supported these findings. Elemental peaks for C 1s, Si 2p, O 1s, and N 1s were observed at binding energies of 285.1, 102.1, 532.1, and 399.1 eV, respectively (Fig. 2(h)). Compared to MCM-41 which contained 14.4% of C 1s, MCM-41-C8 and PLA1@MCM-41-C8 showed significantly increased C 1s levels of 33.5% and 49.8%, respectively, owing to the incorporation of carbon-rich groups from C8 and the proteinaceous enzyme. Similarly, the N content increased from 1.1% to 4.0%, indicating the presence of amino groups in the immobilized phospholipase [43].
3.3. Enzymatic degumming
The schematic diagram in Fig. 3(a) illustrates the enzymatic degumming mechanism of PLA1@MCM-41-C8. PLA1 selectively hydrolyzes the acyl bonds of PLs, converting non-hydratable PLs into hydratable forms (i.e., hydratable PLs). These hydratable PLs were separated by centrifugation [17]. To evaluate the catalytic performance of PLA1@MCM-41-C8, the degumming rates of COs were compared with those of carrier-only controls using MCM-41 and the hydrophobic carrier MCM-41-C8. Fig. 3(b) shows that the degumming rates induced by MCM-41 and MCM-41-C8 were approximately 22.5%, which was attributed to the physical adsorption of PLs by the mesoporous silica [44]. By contrast, PLA1@MCM-41-C8 achieved a phospholipid removal rate of 94.3%, indicating that enzymatic degumming has a significantly higher phospholipid removal efficiency than physical degumming. Moreover, the retention of ARA in algal oils after physical and enzymatic degumming was 98.1% and 97.3%, respectively. Hence, enzymatic degumming via PLA1@MCM-41-C8 not only effectively removed PLs but also selectively preserved the functional components of ARA.
To further improve the degumming efficiency of the immobilized phospholipase, the effect of the PLA1@MCM-41-C8 amount was investigated over the range of 0.1%-0.9%. Fig. 3(c) shows that the degumming rate of algal oils increased rapidly from 46.4% to 55.8% and 94.9% as the PLA1@MCM-41-C8 content increased from 0.1% to 0.3% and 0.5%, respectively. This suggests that the self-made immobilized phospholipase was catalytically effective. However, further increase in PLA1@MCM-41-C8 from 0.5% to 0.9% only slightly improved the degumming rate to 96.1% because the enzymatic reaction could have reached a saturation state [45]. Hence, 0.5% of phospholipase was regarded as the optimal adding amount for enzymatic degumming of algae oil.
In addition to the amount of PLA1@MCM-41-C8, the effects of reaction temperature on enzymatic degumming were also studied. Fig. 3(d) shows that increasing the reaction temperature from 15 to 35 °C increased the phospholipid removal rate from 68.7% to 95.9%. However, further increasing the temperature to 45 and 55 °C caused the degumming rates to decrease to 67.1% and 18.6%, respectively.
The initial increase in temperature not only enhanced the catalytic activity of PLA1 but also reduced the reaction viscosity from 70.4 to 23.8 mPa·s (Table S2 in Appendix A), thereby improving the enzymatic degumming efficiency. However, temperatures above 35 °C negatively impacted the activity of PLA1. This is consistent with the findings of Thangaraju et al. [46], who also reported that PLA1 exhibited maximum catalytic performance around 35 °C. Therefore, a reaction temperature of 35 °C was selected for enzymatic degumming in subsequent experiments.
Finally, the oil quality of EDO with a 95% degumming rate was compared with CRO and CO based on their PVs, moisture content, and oxidation induction time. Fig. 3(e) shows that the commercialized algae oil produced through traditional refining had a low PV of 0.01 g·100 g−1, while the PVs for both CO and enzymatically degummed oil were 0.04 g·100 g−1. All values were within the edible oil standard of < 0.06 g·100 g−1. The lower PV observed in CRO can be attributed to the traditional degumming process, which completely removes trace components that may affect the stability of algae oil. However, this also results in the loss of beneficial micronutrients [47]. In contrast, enzymatic degumming selectively hydrolyzes PLs while retaining other components of the algal oil, which has minimal impact on oil stability, as reflected by the unchanged PV.
Similarly, the lowest moisture content (0.003%) was observed in CRO, likely due to the addition of citric acid and NaOH solutions during chemical degumming, which can absorb water from the oil. In comparison, the moisture content increased slightly from 0.01% in CO to 0.015% in EDO, owing to the enzymatic hydrolysis of PLs [48]. However, the oxidation induction time decreased from 2.6 h in CO to 2.1 h in EDO, indicating that the removal of PLs may accelerate the oxidation of algal oil. This is because PLs, due to their negatively charged groups, exhibit strong metal ion chelating activity, which helps inhibit lipid oxidation [49]. Geng et al. [50] further reported that free amino groups in PLs can react with lipid oxidation products through the Maillard reaction to produce antioxidants such as dihydropyrrole PLs, thereby enhancing the antioxidant capacity of oils.
Nevertheless, the oxidation induction time of enzymatically degummed algal oil still meets the requirements for edible oil.
3.4. Enzymatic deacidification
This section describes the further refinement of EDO by reducing its FFA content using PLA1@MCM-41-C8, glycerol, and a 4 Å molecular sieve, serving as the biocatalyst, acyl acceptor, and water absorbent, respectively (Fig. 4(a)).
To enhance the catalytic performance of PLA1@MCM-41-C8, the enzymatic deacidification conditions for EDO—including reaction temperature, substrate molar ratio of FFA to glycerol, and the amounts of PLA1@MCM-41-C8 and 4 Å molecular sieve—were systematically investigated.
Fig. 4(b) shows that increasing temperature from 25 to 35 °C reduced the final AV of EDO from 6.8 to 2.0 mg KOH·g−1. However, further increasing the temperature from 35 to 75 °C caused the final AV to gradually increase from 5.0 to 12.3 mg KOH·g−1. This trend is consistent with the optimal temperature identified in the enzymatic degumming study above, confirming that PLA1@MCM-41-C8 exhibits its highest catalytic activity at 35 °C. Glycerol was selected as the acyl acceptor because it can esterify with FFAs in algae oil, thereby reducing the AV while simultaneously lowering the levels of functional lipids such as MAGs and DAGs. The effect of the molar ratio of glycerol to FFA, ranging from 1:0.5 to 1:8, on EDO deacidification was also investigated.
Fig. 4(c) shows that the lowest AV of 1.2 mg KOH·g−1 was achieved when the molar ratio of FFA to glycerol was 1:6. However, further increasing the molar ratio to 1:8 resulted in no additional improvement in FFA removal. A moderate increase in glycerol enhances the lipase-substrate catalytic reaction; however, excessive glycerol increases system viscosity, which can hinder enzymatic activity [51]. Therefore, the optimal FFA-to-glycerol molar ratio for EDO deacidification was determined to be 1:6.
Based on the optimal reaction temperature of 35 °C and the FFA-to-glycerol molar ratio of 1:6, the effects of PLA1@MCM-41-C8 and the 4 Å molecular sieve on EDO deacidification were further investigated. Fig. 4(d) shows that increasing the amount of PLA1@MCM-41-C8 from 0.5% to 5% enhanced FFA removal, reducing the final AV from 8.8 to 1.2 mg KOH·g−1. This demonstrates the catalytic effectiveness of the immobilized phospholipase. However, increasing the amount to 6% did not yield any further decrease in AV, indicating that 5% PLA1@MCM-41-C8 was sufficient for the current reaction system. A similar trend was observed with the addition of the 4 Å molecular sieve, which achieved the lowest AV of 1.1 mg KOH·g−1 at a concentration of 4% (Fig. 4(e)). Lower amounts were insufficient to effectively remove the water generated by enzymatic esterification, while excessive amounts may hinder contact between PLs and substrates. Thurs, 4% of the 4 Å molecular sieve was determined to be the optimal for removing the water by-product in the enzymatic deacidification of EDO.
3.5. Kinetic model and molecular docking analysis of enzymatic deacidification
To better understand and predict the enzymatic deacidification of EDO, kinetic parameters—including Km, Vm, Vm/Km, and Ea—were compared between free PLA1 and immobilized PLA1 under the optimized conditions described above.
The Lineweaver-Burk double reciprocal plots (Fig. 4(f)) based on the Michaelis-Menten equation, indicate that the Km of PLA1@MCM-41-C8 (0.28 mol·L−1) was significantly lower than that of free PLA1 (3.22 mol·L−1) (Table S3 in Appendix A). This suggests that the immobilized PLs have a higher substrate affinity [52]. In contrast, the Vm of PLA1@MCM-41-C8 was 1.8 times higher than that of free PLA1, and its apparent Vm/Km was 20.6 times greater. These results confirm that the catalytic activity of PLA1 is substantially enhanced upon immobilization.
According to the activation energy results derived from the Arrhenius equation (Fig. 4(g)), the Ea of free PLA1 (77.8 kJ·mol−1) was reduced to 65.3 kJ·mol−1 after immobilization on MCM-41-C8. This further demonstrates the improved catalytic efficiency of PLA1@MCM-41-C8 compared to the free enzyme.
To gain insight into the catalytic mechanism of PLA1 during enzymatic deacidification, molecular docking simulations were used to investigate the binding affinity and interaction patterns between PLA1 and small molecules such as glycerol and oleic acid. The two-dimensional (2D) diagrams in Figs. 4(h) and (i) show that both oleic acid and glycerol formed stable interactions with the amino acid residues of PLA1 via hydrogen bonding and hydrophobic interactions.
Specifically, a hydrogen bond between oleic acid and PLA1 was formed with a single amino acid residue, Thr92(A), whereas the corresponding residues involved in glycerol binding were Asn83(A) and Ser11(A). The binding sites responsible for hydrophobic interactions differed between oleic acid and glycerol, as detailed in Table S4 in Appendix A. Overall, the binding energies of PLA1 to oleic acid and glycerol were -7.5 and -3.7 kcal·mol−1, respectively. A lower binding energy indicates stronger enzyme-substrate affinity, which is primarily influenced by the carbon chain length and molecular weight of the substrates [53].
3.6. Evaluation of oil quality and wide adaptability
Enzymatic deacidification of algal oil using glycerol as an acyl acceptor catalyzes esterification between FFAs and glycerol to produce MAG and DAG. The main components—triacylglycerol (TAG), DAG, MAG, and FFA—in ARA oils before and after enzymatic degumming and deacidification are compared in Fig. 5(a). CO initially consisted of 86.4% TAG, 5.2% DAG, 1.3% MAG, and 7.1% FFA. After enzymatic degumming with PLA1@MCM-41-C8, the composition changed slightly to 86.0% TAG, 5.8% DAG, 1.2% MAG, and 7.0% FFA, indicating that the main composition of the algal oil remained largely unchanged. This reflects the high specificity of PLA1@MCM-41-C8 for the hydrolysis of PLs without affecting other lipid components. However, subsequent enzymatic deacidification of EDDO resulted in a significant reduction in TAG and FFA contents to 61.7% and 0.5%, respectively. The reduction in FFA content was attributed to esterification with glycerol, while the decrease in TAG content was primarily due to glycolysis and simultaneous TAG hydrolysis. These reactions also promoted the formation of DAG and MAG, which increased to 34.4% and 3.4%, respectively. Therefore, enzymatic deacidification using PLA1@MCM-41-C8 not only effectively reduced the AV of ARA-enriched algal oil but also led to the production of functional lipids. The former improves oil quality, while the latter enhances its nutritional value.
The self-prepared immobilized PLA1 retained 90% of its initial catalytic capacity, maintaining the ability to reduce the oil AV to 3 mg KOH·g−1 after seven reuse cycles. This operational stability may be attributed to changes in the secondary structure of PLA1 upon immobilization (Figs. S1 and S2 in Appendix A). As shown in Table S5 in Appendix A, PLA1 immobilization increased the α-helix and β-turn contents from 31.3% and 0 to 35.4% and 29.6%, respectively, while decreasing the β-sheet and random coil contents from 27.6% and 41.1% to 0 and 35.0%, respectively. The enhanced structural stability of PLA1@MCM-41-C8 compared to free PLA1 was mainly attributed to the increased α-helix content, which contributes to enzyme rigidity through hydrogen bonding [54]. However, prolonged reuse of PLA1@MCM-41-C8 led to a decline in α-helix content to 24.5%, resulting in a corresponding decrease in catalytic activity.
The antioxidant stability of EDDO was compared with CRO and CO based on the PVs, moisture content, and oxidation induction time (Fig. 5(b)). Further enzymatic deacidification of degummed algae oil had minimal effect on the antioxidant properties, as the PV, moisture content, and oxidation induction time of EDDO remained nearly unchanged compared to those of CO. In contrast, CRO showed significantly lower PV and moisture content compared to CO—decreased by factors of 4 and 3.3, respectively. This suggests that conventional refining processes, which often involve high temperatures and chemical solvents, significantly alter oil quality. The broader applicability of PLA1@MCM-41-C8 was further evaluated through its use in enzymatic degumming and deacidification of various edible oils. Fig. 5(c) shows that all five tested vegetable oils—PO, SO, RO, RBO, and WO—achieved degumming rates above 91.7% through enzymatic hydrolysis of PLs by PLA1@MCM-41-C8. The highest degumming rate of 96.5% was observed in SO, where phospholipid content was decreased from 300.6 to 10.3 mg·kg−1.
Using the PLA1@MCM-41-C8 under optimal reaction conditions, the AVs of TSO, RO, RBO, SO, and FO decreased from 5.5, 5.7, 19.2, 7.9, and 4.8 mg KOH·g−1 to 1.6, 1.8, 1.0, 1.6, and 0.98 mg KOH·g−1, respectively (Fig. 5(d)). All enzymatically deacidified oils met the standard requirement for edible oils, which mandates an AV of less than 3 mg KOH·g−1. Notably, RBO and FO with final AVs below 1.5 mg KOH·g−1 met the first-grade standard for refined oils [55].
3.7. Fatty acid composition and contents
This section describes the content and composition of 11 fatty acids in different algae oils, as presented in Table 1. CO consisted of 13.10% saturated fatty acids (SFAs) and 83.18% unsaturated fatty acids (USFAs), with ARA (C20:4) accounting for 50.29%. These fatty acid profiles are largely consistent with those of CRO. Enzymatic degumming using PLA1@MCM-41-C8 led to slight reductions in SFA, USFA, and ARA contents, which decreased to 9.80%, 80.37%, and 49.89%, respectively. Further enzymatic deacidification resulted in a decrease in USFA to 76.46%, while SFA slightly increased to 11.09%. Notably, alkaline deacidification caused a more substantial reduction in USFA content—by 18.9%—with ARA and EPA (C20:5) significantly reduced to 46.43% and 1.44%, respectively. This indicates that alkaline deacidification leads to the degradation of USFAs, likely due to oxidation during exposure to alkaline catalysts and separation via rotary evaporation [56]. During enzymatic degumming, the total oil content decreases due to the removal of PLs. In contrast, enzymatic deacidification, through esterification of FFAs with added glycerol, generates MAG, DAG, and TAG, thereby increasing the total oil content. Consequently, changes in total oil content may also influence the observed fatty acid composition and should be considered when evaluating compositional shifts. Overall, these findings confirm that PLA1@MCM-41-C8 has a minimal impact on the fatty acid composition and content of algal oil during enzymatic refining.
3.8. Flavor characteristics of enzymatic-refined algae oils
Algae oils typically exhibit oceanic or fishy odors, which can be attributed to the origin of oleaginous microorganisms and the specific oil processing methods used [57]. This section explores the effects of enzymatic degumming and deacidification using PLA1@MCM-41-C8 on the aroma characteristics of different ARA-enriched algal oils. Fig. 6 presents a heatmap of the volatile flavor compounds identified in the various algal oil samples. A total of 74 volatile compounds were detected and classified into six categories: ketones (15 compounds), alcohols (26), acids (6), sulfur-containing compounds (2), aldehydes (17), and heterocyclic compounds (8). As summarized in Table S6 in Appendix A, 56 aroma-active compounds with a total concentration of 41 563.9 mg·kg−1 were identified in CRO. Among these, alcohols (21 512.0 mg·kg−1) and aldehydes (6 124.1 mg·kg−1) were the predominant contributors, responsible for the characteristic alcoholic, malty, meaty, and burnt-paste aroma profile [58]. Due to the higher content of PLs and FFAs, the total volatile flavor compound content in CO was reduced to 28 173.4 mg·kg−1. However, alcohols (15 615.7 mg·kg−1) and aldehydes (5 753.8 mg·kg−1) remained the dominant components in its aroma profile. Following enzymatic degumming with PLA1@MCM-41-C8, the total volatile compound content in EDO further decreased to 17 307.9 mg·kg−1 but increased to 25 675.3 mg·kg−1 after subsequent enzymatic deacidification. This increase was mainly attributed to the addition of glycerol, which led to a marked rise in alcohol content—from 10 199.2 mg·kg−1 in EDO to 15 025.1 mg·kg−1 in EDDO. In contrast, alkaline deacidification resulted in a significantly lower total volatile compound content of 12 696.0 mg·kg−1 in ADO, which is 2.2 times lower than that observed in CO. This reduction is attributed to the absorption of aroma compounds by residual soap formed during the alkaline deacidification process, which is subsequently separated from the algal oil. Thus, enzymatic refining exerts a milder impact on the volatile components of algae oil compared to conventional methods. Across all samples, alcohols and aldehydes remained the dominant contributors to the overall aroma profile.
3.9. Stability evaluation of PLA1@MCM-41-C8
Thermostability refers to the ability of an enzyme to maintain its structural integrity and catalytic activity at elevated temperatures. The thermostability of PLA1@MCM-41-C8 was evaluated based on its enzymatic activity following 2 h of incubation at temperatures ranging from 15 to 65 °C. As shown in Fig. 7(a), the enzyme activity of PLA1@MCM-41-C8 gradually increased from 1056.6 to 1205.7 U·g−1 as the incubation temperature increased from 15 to 35 °C. This suggests that moderately increasing the temperature can enhance the catalytic ability of phospholipase. However, further increasing the incubation temperature to 65 °C reduced the enzyme activity to 807.3 U·g−1. Free PLA1 is stable at around 450.0 U·g−1 at 15 to 35 °C but showed a sharp decline to 321.0 U·g−1 when the incubation temperature increased to 65 °C. Notably, PLA1@MCM-41-C8 consistently demonstrated significantly higher enzymatic activity than free PLA1 across all tested temperatures, particularly at elevated temperatures. These results suggest that enzyme immobilization on heat-resistant silica carriers effectively protects against structural degradation and loss of activity [59].
Fig. 7(b) presents a bar chart comparing enzyme activity across different pH values for immobilized and free phospholipase. The results indicate that immobilized phospholipase exhibited superior pH tolerance than free phospholipase. Fig. 7(c) shows enzyme activity versus different solvents. For immobilized phospholipase and free phospholipase, the results show that immobilized phospholipase exhibited better solvent tolerance than free phospholipase. Notably, since PLA1 exhibits an inherent optimal pH of approximately 5.5-5.8, both free and immobilized PLA1 showed maximum enzymatic activity at pH 5, reaching 503.3 and 1160.0 U·g−1, respectively. Both forms of PLA1 demonstrated low activity in polar solvents, such as methanol, ethanol, and acetonitrile but maintained high catalytic performance in acetone, ether, and petroleum ether. The highest enzyme activity was observed in petroleum ether, where PLA1@MCM-41-C8 and free PLA1 achieved 1 043.3 and 490.0 U·g−1, respectively. The long-term storage stability of PLA1@MCM-41-C8 was also evaluated over a 45-day period at 4 °C. Fig. 7(d) shows a bar-chart of enzyme activity versus storage days. Although enzyme activity gradually decreased, the immobilized PLA1 retained 75% of its initial activity by day 45. Thus, compared to free PLA1, the immobilization of PLA1 on MCM-41-C8 effectively improved thermostability, pH tolerance, solvent resistance, and storage stability of PLA1. This suggests strong potential for its application under harsh or complex industrial reaction conditions. Looking forward, to overcome limitations associated with batch reactions—such as high shear stress, limited mass transfer, and substrate inhibition—continuous-flow enzymatic reactors, including packed-bed and fluidized-bed reactors, should be explored.
4. Conclusions
In this study, an immobilized phospholipase, PLA1@MCM-41-C8, was developed and applied for the enzymatic degumming and deacidification of ARA-enriched algal oil. By optimizing both the PLA1 immobilization conditions and the subsequent enzymatic reaction parameters, a maximum degumming rate of 95.9% was achieved within 3 h. The degummed algae oil was then further deacidified using the same PLA1@MCM-41-C8; this reduced the AV from 14.5 to 1.1 mg KOH·g−1 within 6 h while simultaneously increasing DAG content from 5.2% to 34.4%, as analyzed through kinetic models and molecular docking simulations revealed the mechanism of immobilized phospholipase applied to deacidification. Both enzymatic degumming and deacidification had minimal effects on the oil quality, fatty acid composition, flavor characteristics, and ARA retention. Finally, the improved thermal stability, pH tolerance, solvent tolerance, and long-term reusability of PLA1@MCM-41-C8 demonstrate its strong potential for industrial applications under complex reaction conditions.
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