Enzymatic preparation and quality evaluation of diacylglycerol-rich Swida wilsoniana fruit oil

Abstract

Herein, diacylglycerol (DAG) was prepared from Swida wilsoniana fruit oil by a two-step enzymatic process, and the quality of crude Swida wilsoniana fruit oil (Crude oil), refined Swida wilsoniana fruit oil (Refined oil), DAG oil derived from crude oil (Crude oil-DAG), and DAG oil derived from refined oil (Refined oil-DAG) were compared and evaluated. The results showed that Lipozyme® RM IM enzyme exhibited the best hydrolysis effect in the hydrolysis reaction, while Novozym® 435 enzyme had a lower reaction energy barrier and a faster esterification catalytic rate in the esterification reaction, yielding a DAG content of up to 72.99%. Quality analysis showed that the physical and chemical indicators of the four oils all met the national standards, refining and enzymatic reactions can affect the tocopherol content, oxidative stability and flavor components of oils. The tocopherol content and oxidative stability in the crude oil were the best, and the volatile flavor substances of the crude oil and its DAG were richer. The research confirmed that the crude Swida wilsoniana fruit oil is an ideal raw material for the preparation of DAG, and the obtained DAG has excellent comprehensive quality.

Highlights

• •Optimization of the enzymatic preparation process of diglyceride-rich Swida wilsoniana fruit oil.
• •DAG 36:4 (C₃₉H₆₈O₅) and DAG 36:3 (C₃₉H₇₀O₅) account for 98% of the diglyceride content.
• •Crude oil and Crude oil-DAG are rich in tocopherol and have better oxidative stability.
• •Crude oil and Crude oil-DAG are rich in volatile flavor compounds (40 species).
• •Refining reduces the content of tocopherols and flavor compounds.

1. Introduction

Swida wilsoniana is a woody oil crop belonging to the Cornaceae family, primarily cultivated in Hunan, Hubei, and Jiangxi provinces of China (Wen et al., 2021). The average oil content of Swida Wilsoniana fruit is 33.95% ∼ 36% (Alessandra et al., 2022; Cross et al., 2021), of which the content of β-carotene is 4.7 mg/100 g. The fatty acids mainly include oleic acid, linoleic acid, linolenic acid, lauric acid, palmitic acid and stearic acid. Swida Wilsoniana fruit oil has been used as edible oil for more than 100 years (Xiao et al., 2022). Studies have shown that Swida Wilsoniana fruit oil has the effects of lowering blood lipids, preventing fatty liver and preventing atherosclerosis (Liu et al., 2020). Fu et al. (2012) conducted animal experiments on Swida Wilsoniana fruit oil and found that Swida Wilsoniana fruit oil can reduce serum triglycerides (TAG) and total cholesterol, and has a good lipid-lowering effect.

DAG is a structured lipid formed by replacing a fatty acid at a specific position in TAG with a hydroxyl group. Due to the different positions of the hydroxyl group, two isomers, 1,3-DAG and 1,2-DAG, are formed (deMan et al., 1992;Tada, 2004). As a component of natural oils, DAG is present in very low levels in most animal and plant oils. Even in palm oil, which has a high DAG content, it does not exceed 10% (St - Onge & Jones, 2002). Clinical studies have shown that when the DAG content in edible oil is greater than 40%, consuming 2.5 g of DAG edible oil per day can promote postprandial fat metabolism and inhibit fat accumulation (Ando et al., 2017). In the food industry, DAG is widely used as a green food ingredient in the production of dairy products, baked goods, chocolate, etc. (Butterbaugh & Sargent, 2001; Liu et al., 2019). In the pharmaceutical field, due to the lack of Sn-2-specific acyltransferase in the body, hydrolyzed DAG does not follow the 2-MAG re-esterification pathway and will not accumulate in the human body, thereby reducing the blood lipid levels. Therefore, DAG can be used for the prevention and treatment of fatty liver, hyperlipidemia and related cardiovascular and cerebrovascular diseases (Lee et al., 2020;Taguchi et al., 2000;Saito et al., 2006). In the chemical and biological industries, DAG can be used for the synthesis of a variety of compounds, such as resins, phospholipids, ester proteins, etc.. It can also be used in the production of enzyme activators and inhibitors in the bioindustry (Wright & Marangoni, 2002).

The main methods for preparing DAG include chemical and enzymatic methods. Although chemical methods are characterized by mature technology, low cost, and ease of large-scale production, their drawbacks include poor reaction selectivity, the tendency to produce mixtures of positional isomers, and the potential for side reactions and degradation of nutritional components at high temperatures. Enzymatic methods, on the other hand, offer mild reaction conditions, high positional selectivity, and better product purity. In particular, the use of 1,3-specific lipases for the targeted synthesis of 1,3-DAG aligns better with the principles of green chemistry (Lin et al., 2025; Shi et al., 2025).

As a new type of woody oil unique to my country, the Swida Wilsoniana fruit oil has a good market prospect. The unsaturated fatty acid content in the Swida Wilsoniana fruit oil is relatively high, ranging from 76.02% to 82.43%, of which the linoleic acid content is about 42%. The high content of unsaturated fatty acids and its unique fatty acid composition make it a specific raw material for DAG preparation. Meanwhile, the active substances contained in the Swida Wilsoniana fruit oil may also have beneficial effects on diglycerides, such as enhancing oxidative stability and increasing flavor components. However, systematic exploration is lacking in key parameters such as the enzymatic preparation of diglycerides from Swida Wilsoniana fruit oil, optimization of process parameters, catalyst selection based on its fatty acid specificity, and optimization of reaction pathways.

This study aimed to prepare diacylglycerol (DAG) from Swida Wilsoniana fruit oil using a two-step hydrolysis-esterification method with bio-enzymes as catalysts. The process conditions for hydrolysis-esterification were optimized, and the reaction mechanism of the enzyme-catalyzed process was thoroughly understood from kinetic and thermodynamic perspectives. Furthermore, the effects of processing (refining and enzyme catalysis) on key quality parameters (physicochemical properties, fatty acid composition, tocopherol determination, oxidative stability, and flavor) of Swida Wilsoniana fruit oil and its DAG were comprehensively evaluated. This research is of great significance for improving the value of Swida Wilsoniana fruit oil, enhancing human health, and promoting the industrial production of DAG.

2. Experimental section

2.1. Materials

Crude Swida wilsoniana fruit oil was extracted by the State Key Laboratory of Woody Oil Resources Utilization, Hunan Academy of Forestry, using low-temperature prepressing followed by subcritical n-butane extraction. Its acid value was 3.4 mg/g. After refining, the acid value of the Swida wilsoniana fruit oil was 1.2 mg/g. Immobilized lipases (Lipozyme® RM IM, Lipozyme® TL IM and Novozym® 435 with a nominal enzymatic activity of 275 IUN/g, 250 IUN/g and 8000 PLU/g, respectively) were purchased from Novozymes (China) Biotechnology Co., Ltd., analytical reagent (AR) grade. Free lipase F1 with a nominal enzymatic activity of 8000 u/g was supplied by Hunan Wanquan Yuxiang Biotechnology Co., Ltd. Petroleum ether(30–60 °C), isopropanol, glycerol (Propylene glycol), sodium hydroxide (NaOH), potassium iodide (KI), sodium bisulfate (NaHSO₄), soluble starch, chloroform (CHCl₃), glacial acetic acid, cyclohexane, Wei's reagent, potassium hydroxide-ethanol solution (KOH in EtOH), phenolphthalein and alkali Blue 6B were purchased from Sinopharm Chemical Reagent Co., Ltd., AR grade. Sodium thiosulfate (Na₂S₂O₃) and potassium hydroxide (KOH) were supplied by Tianjin Chemical Reagent Research Institute Co., Ltd. Activated clay and hydrochloric acid standard solution were provided by Macklin Biochemical Technology Co., Ltd., AR grade. Isooctane, isopropanol, anhydrous ethanol, n-hexane, methanol (MeOH), tetrahydrofuran (THF), Butylated hydroxytoluene (BHT) and tert-butyl methyl ether (MTBE) were purchased from MacLean Biochemical Materials Technology Co., Ltd. A 37-component fatty acid methyl ester (FAME) standard mix was supplied by TMRM Quality Inspection Technology Co., Ltd. Tocopherol reference standards (α-, β-, γ-, δ-tocopherol) were purchased from Sinopharm Chemical Reagent Co., Ltd., all of which were mass spectrometry (MAS) grade. Ultrapure water was used throughout the experimental procedures.

2.2. Methods

2.2.1. Hydrolysis

Crude Swida wilsoniana fruit oil was placed into the 250 mL iodine flasks, then ultrapure water was added with different water addition amounts (10%, 20%, 30%, 40%, 50%, 60% wt_water/wt_oil+water), and a certain amount of lipase (2%, 3%, 4%, 5%, 6% wt_lipase/wt_oil+water) was added subsequently. After sealing, the iodine flasks were placed in a constant temperature water bath shaker at a certain temperature (30 °C, 40 °C, 50 °C, 60 °C, 70 °C), and the shaking speed was selected to be 180 rpm. To determine the effect of time, separate but identical reaction flasks were set up for each predetermined time point (4 h, 8 h, 12 h, 16 h, 20 h, 24 h). This approach ensured that each data point was obtained from an independent reaction mixture quenched and processed at the exact target time, thereby accurately representing the hydrolysis state at that interval without the need for sampling from an ongoing reaction. After a certain reaction time, the reaction solution was poured into a 100 mL centrifuge tube and centrifuged at a speed of 5000 r/min for 30 min. The upper oil phase was aspirated to separate the oil phase from the lipase and the water phase after the centrifugation. The aspirated oil phase is the free fatty acid of the Swida wilsoniana fruit oil, referred to as S-FFA.

2.2.2. Esterification

S-FFA and glycerol were placed in a round-bottom flask at different substrate molar ratios (3:1, 2:1, 1:1, 1:2, 1:3), and a certain amount (1%, 2%, 3%, 4%, 5%) of lipase was added. The reaction was placed in a vacuum environment and stirred at 450 rpm at a certain temperature (30 °C, 40 °C, 50 °C, 60 °C, 70 °C). After a certain reaction time (2 h, 4 h, 6 h, 8 h, 10 h), the reaction solution was poured into a 50 mL centrifuge tube and centrifuged at 5000 r/min for 30 min. The upper oil phase was aspirated to separate it from lipase and glycerol after the centrifugation. The aspirated oil phase is S-DAG.

2.2.3. Determination of glyceride composition and calculation of esterification rate

Pretreatment: Weighed 10–20 mg of the sample into a 10 mL centrifuge tube, added 1 mL of methanol/chloroform (1:1) to dissolve, vortex mixed, taken 10 μL of the above solution to 800 mL of methanol/chloroform for dilution, then added 100 μL of full lipid internal standard (EquiSPLASH-330,731-1EA-REV, 10 μg/mL), vortex mixed, passed through a 0.22 μm organic filter membrane, and applied to the machine.

Chromatographic system: Shimadzu UPLC LC-30 A, chromatographic column: Phenomenex Kinete C18 column (100 × 2.1 mm, 2.6 μm); injection volume: 3 μL; flow rate: 0.4 mL/min; column temperature: 60 °C; sample chamber temperature: 4 °C. Phase A: water: methanol: acetonitrile (1:1:1) (containing 5 mM NH₄Ac); phase B: isopropanol: acetonitrile (5:1) (containing 5 mM NH₄Ac); gradient: 0.5 min, 20% B; 1.5 min, 40% B; 3 min, 60% B; 13 min, 95% B, hold 4 min; 20 min, 20% B; 25 min, 20% B.

Mass spectrometry system: AB Sciex Triple TOF®6600, ESI⁺ ion source, mass spectrometry scanning range: 100 m/z ∼ 1200 m/z; Curtain Gas: 35.00 psi; Ion Source Gas 1: 50.00; Ion Source Gas 2: 50.00; Ion Spray Voltage: 5500.00 V; Temperature: 600 °C.

Based on the peak area and retention time, the relative percentage contents of DAG, TAG and MAG of different fatty acid chains were calculated using (1), (2), (3).

$$\mathrm{MAG}\text{content}\left(\%\right)=\frac{\mathrm{MAG}\text{peak area}}{\text{Total glyceride peak area}}\times 100\%$$ (1)

$$\mathrm{DAG}\text{content}\left(\%\right)=\frac{\mathrm{DAG}\text{peak area}}{\text{Total glyceride peak area}}\times 100\%$$ (2)

$$\mathrm{TAG}\text{content}\left(\%\right)=\frac{\mathrm{TAG}\text{peak area}}{\text{Total glyceride peak area}}\times 100\%$$ (3)

Esterification efficiency was calculated using Eq. (4):

$$\text{Esterification efficiency}\left(\%\right)=\left(1-\frac{\text{Acid value of esterified oil}}{\text{Acid value of feedstock}\mathrm{FFA}}\right)\times 100\%$$ (4)

2.2.4. Thermodynamics and kinetics of esterification reaction

It is known that the esterification reaction between fatty acids and glycerol catalyzed by lipases follows a Ping-Pong Bi—Bi mechanism, accompanied by substrate inhibition, particularly inhibition of glycerol (Phuah et al., 2012). Therefore, we adopted this mechanism as the basis for the kinetic analysis. The esterification reaction rate is closely related to the reaction temperature. The reaction temperature will affect the rate constant and enzyme activity, thereby affecting the esterification effect and DAG content, but will not affect the reaction order. The relationship between the reaction rate and reaction temperature can be derived from the classical thermodynamic equation, as shown in formula (5):

$$v=k{c}_A^{\alpha }{c}_B^{\beta }$$ (5)

The reaction equilibrium constant (k) is primarily by the reaction temperature (T), which are expressed as eq. (6):

$$k=-\left(\frac{k_{B}T}{h}\right)\times \frac{e^{∆G\ne }}{\mathit{RT}}$$ (6)

where k is the reaction equilibrium constant, k_B is the Boltzmann constant 1.381 × 10⁻²³ J/K, h is the Planck constant 6.626 × 10⁻³⁴ J·s, ΔG^≠ is the change in reaction free energy (J/mol), T is the absolute temperature (K) and R is the gas constant 8.314 J/mol·K.

The Arrhenius equation is shown in eq. (7), which explains the relationship between reaction rate and reaction temperature.:

$$k=A{e}^{-E_a/\mathit{RT}}$$ (7)

where A is the Arrhenius constant, Ea is the activation energy (J/mol).

Assuming the reactant concentration remains approximately constant in the initial stage, substituting eq. (7) into eq. (5) yields eq. (8):

$$v=A^{\prime }{e}^{-E_a/\mathit{RT}}$$ (8)

where$A^{\prime }=A{c}_A^{\alpha }{c}_B^{\beta }$, taking the logarithm of both sides of eq. (8) yields eq. (9):

$$\ln v=\ln A^{\prime }-\frac{E_a}{\mathit{RT}}$$ (9)

Esterification was carried out under the optimal enzymatic hydrolysis conditions, and the initial reaction velocity corresponding to different substrate concentrations was calculated. According to eq. (9), a graph was drawn with ln v as the ordinate and 1/T as the abscissa. From the fitting slope = ($-\frac{E_a}{R}$), the activation energy of the reaction and the relationship between the initial reaction velocity and the reaction temperature can be obtained.

A ping-pong bi-bi mechanism was employed to evaluate initial reaction rates based on different mechanisms. By varying reaction conditions, the synthesis of diglycerides from fatty acids and glycerol under lipase catalysis was determined, identifying the primary reaction mechanism. The maximum reaction rate (v_m) and Michaelis constant (K_m) were calculated using the initial reaction rates obtained under different conditions. This ping-pong bi-bi model (eq. (10)) can calculate the kinetic parameters necessary to understand enzyme-substrate interactions, such as the inhibition constant.

$$v_0=v_m/\left(1+\frac{K_A}{\left[A\right]}+\frac{K_B\left(1+\frac{\left[A\right]}{K_{\mathit{IA}}}\right)}{\left[B\right]}\right)$$ (10)

where$v_0$ is the initial velocity, $v_m$ is the maximum reaction rate, [A] and [B] are the concentrations of glycerol and S-FFA, respectively, K_A and K_B are the kinetic parameters for glycerol and S-FFA, K_IA is the inhibition constant for glycerol.

When S-FFA is in excess in the reaction system, [B] can be regarded as a constant, and eq. (10) can be simplified to eq. (11):

$$v_0=\frac{v_m\left[S\right]}{K_m+\left[S\right]}$$ (11)

where $v_0$ is the initial velocity, $v_m$ is the maximum reaction rate, $K_m$ is the Michaelis constant.

According to the equation, a Lineweaver-Burk double reciprocal plot of $1/v$ corresponding to $1/\left[S\right]$ is made. From the vertical intercept of the fitting straight line = $1/v_m$, and the slope = $K_m/v_m$, the Michaelis constant ($K_m$) was calculated as: $K_{\mathrm{m}}=\frac{\text{Slope}}{\text{vertical intercept}}$.

2.2.5. Determination of fatty acids

According to GB5009.168–2016 National Food Safety Standard Determination of Fatty Acids in Foods, the oil samples were subjected to methyl esterification operation.

Gas phase conditions (Nicholson & Marangoni, 2021): Shimadzu GC-2010 Plus gas chromatograph, equipped with a flame ionization detector (FID). Capillary column: Dura Bond HP-88, polydicyanopropylsiloxane strong polar stationary phase, column length 100 m, inner diameter 0.25 mm, film thickness 0.2 μm. Operating temperature range 50 °C ∼ 250 °C. Injector temperature: 250 °C. Detector temperature: 280 °C. Program temperature rise: initial temperature 100 °C, lasting 13 min; 100 °C ∼ 180 °C, heating rate 10 °C/min, hold for 6 min; 180 °C ∼ 200 °C, heating rate 1 °C/min, hold for 20 min; 200 °C ∼ 230 °C, heating rate 4 °C/min, hold for 10.5 min. (85 min in total, column oven temperature 100 °C ∼ 230 °C). High-purity helium 25.00 mL/min as carrier gas, hydrogen: 30.00 mL/min, air: 300.00 mL/min. Split ratio: 100:1, injection volume: 1.00 μL. The oil sample was qualitatively analyzed with a standard mixture of 37 fatty acids, and then the fatty acids in the oil sample were qualitatively and quantitatively analyzed. The experiment was repeated three times and the average value was taken.

2.2.6. Determination of tocopherol content

Tocopherol was determined in accordance with GB/T 5009.82–2016 Vitamin A, D and E in food.

2.2.7. Oxidative stability determination

According to GB/T 21121–2007 Animal and vegetable fats and oils - Determination of oxidative stability (accelerated oxidation test) and reference (Ma et al., 2025):

The oil oxidation induction time (OSI) was measured by Rancimat 892 oil oxidation tester. The setting parameters were as follows: temperature 110 °C, oxygen flux 10 L/h, and 3 g of Swida wilsoniana fruit oil sample was taken for a single test to obtain the oxidation induction time.

2.2.8. Determination of volatile flavor compounds

According to the method of Fang et al. (2022), a GC-IMS instrument (Flavor Spec ® Flavor) equipped with a FSSE-54-CB-1 column was used as follows:

1 g of oil sample was added to a 20 mL headspace injection bottle. The sample was then incubated at 80 °C for 15 min at a speed of 500 r/min. 200 μL of incubated oil sample was automatically injected into the device through the injection needle. The column temperature was 60 °C, the carrier gas/drift gas was N₂, and the IMS temperature was 45 °C. Three parallel runs were made for each oil sample. The gas chromatography conditions were as follows: drift gas flow rate was 150 mL/min, initial carrier gas flow rate was 2 mL/min, 10 mL/min at 10 min, 100 mL/min at 15 min, and 150 mL/min at 20 min.

The spectra and data were analyzed using the analysis software supporting GC-IMS. The database was used to perform qualitative analysis on the substances, and quantitative analysis was performed after the standard curve was established. Reporter was used to obtain three-dimensional spectra, two-dimensional top views, and difference spectra. Gallery Plot was used to obtain fingerprint spectra. PCA was used to obtain principal component analysis and similarity analysis graphs.

3. Results and discussion

3.1. Factors affecting the hydrolysis process of swida wilsoniana fruit oil

The effect of time on the hydrolysis of swida wilsoniana fruit oil is shown in Fig. S1(A). With the increase of reaction time, the hydrolysis rate of the swida wilsoniana fruit oil increased. Lipozyme® RMIM had the best hydrolysis effect on the oil of the fruit of the safflower tree. The hydrolysis rate was the highest when the reaction time was 24 h. This is likely because Lipozyme® RMIM exhibits Sn-1,3 position specificity and selectivity for unsaturated fatty acids, enabling rapid hydrolysis of the 1,3-position fatty acids in oils, especially in swida wilsoniana fruit oil where unsaturated fatty acids account for over 80%.

The effect of reaction temperature on the hydrolysis rate of safflower oil is displayed in Fig. S1(B). The hydrolysis of swida wilsoniana fruit oil first increased with increasing temperature, reaching the highest hydrolysis rate at 60 °C, and then began to decrease after 60 °C. This may be because when the reaction temperature is low, the molecular motion is weak, the contact and collision between the enzyme and the substrate are reduced (Li et al., 2015), and the hydrolysis reaction proceeds slowly. As the temperature rises, the molecular motion becomes stronger, which is conducive to the hydrolysis reaction. When the temperature is too high, the protein structure in the enzyme is easily destroyed, the protein denatures, and the lipase gradually becomes inactive (Li & Zhang, 2020), resulting in a decrease in the hydrolysis rate. Therefore, it is more appropriate to maintain the hydrolysis temperature at around 60 °C.

The effect of water addition on the hydrolysis rate of swida wilsoniana fruit oil is listed in Fig. S1(C). The hydrolysis rate of swida wilsoniana fruit oil increased with the increase of water addition. The hydrolysis rate of lipozyme RMIM and Novozym® 435 on swida wilsoniana fruit oil tended to be flat after the water addition reached 50%. This may be because lipase can only effectively contact with the substrate when it is at the water-oil interface (Xiao et al., 2019). When the water addition is too high or too low, the water-oil interface is reduced, resulting in a smaller contact area between the enzyme and the substrate, which leads to a lower hydrolysis rate. Therefore, it is more appropriate to choose a water addition of 60%.

The effect of enzyme addition on the hydrolysis rate of safflower oil is shown in Fig. S1D. As the amount of enzyme added in the reaction system increased, the hydrolysis rate of swida wilsoniana fruit oil also increased overall. When the enzyme addition amount was 5%, the reaction had reached saturation. Too high an enzyme content will cause the reaction system to become thick, and the lipases will adhere to each other and cannot fully contact the substrate, thereby reducing the hydrolysis rate. Therefore, the optimal hydrolysis effect is achieved when the enzyme addition amount is 5%.

3.2. Factors affecting the esterification process of swida wilsoniana fruit oil fatty acids

The effects of enzyme types on the esterification effect were shown in Fig. S2 using lipase Lipozyme® RMIM, Lipozyme® TLIM, Novozym® 435 and free lipase F1 as catalysts. The results showed that the esterification effects of Lipozyme® RMIM and Novozym® 435 were good, with esterification rates of 94.53% and 90.55%, respectively. However, the esterification effects of Lipozyme® TLIM and F1 were poor, with esterification rates of 23.14% and 21.38%, respectively. Therefore, Lipozyme® RMIM and Novozym® 435 were selected for subsequent experiments to explore the effects of these two lipases as catalysts on the DAG content of the prepared products.

The effect of reaction time on the esterification effect of S-FFA were shown in Figs. 1A and 2A. As shown in Fig. 1A, with Lipozyme® RMIM enzyme as the catalyst, the esterification rate increased with the increase of reaction time. The esterification rate was the highest when the reaction time was 8 h (the content of S-DAG is 53.20%), and then began to decline, which may be because the content of S-FFA in the reaction system is low after 8 h of reaction, and the reaction begins to proceed in the reverse direction (hydrolysis reaction). The results in Fig. 2A showed that the content of S-DAG generated by esterification with Novozym® 435 enzyme as the catalyst is the highest at 8 h, which is 59.9%. When the reaction time exceeds 8 h, the content of S-DAG prepared begins to decline. In summary, when the reaction time was 8 h, the esterification effect of Novozym® 435 enzyme was better than that of Lipozyme® RMIM.

Fig. 1.

Effects of reaction time, substrate molar ratio, enzyme dosage and temperature on esterification efficiency and glycerol content when Lipozyme® RMIM is used as a catalyst.

Fig. 2.

Effects of reaction time, substrate molar ratio, enzyme dosage and temperature on esterification efficiency and glycerol content when Novozym® 435 is used as a catalyst.

The effect of substrate molar ratio on esterification effect was shown in Figs. 1B and 2B. When the substrate molar ratio is in the range of 3:1–1:1, the esterification rate of S-FFA catalyzed by Lipozyme® RMIM and Novozym® 435 increased with the decrease of glycerol addition. When the substrate molar ratio was 1:1, the esterification effect of Lipozyme® RMIM and Novozym® 435 enzymes was the best. When the substrate molar ratio was in the range of 1:1–1:3, the esterification rate decreased with the decrease of glycerol addition, showing a positive correlation. This may be because glycerol itself has a large density, when too much glycerol is added, the entire reaction system becomes viscous, reducing the contact frequency between the reactants, at the same time, glycerol will wrap the outer layer of lipase, reducing the contact area between lipase and FFA, resulting in a poor esterification effect. Compared with Lipozyme® RMIM enzyme, Novozym® 435 enzyme has stronger binding ability with substrate, and when the glycerol content is high, glycerol has less effect on the esterification effect of Novozym® 435 enzyme. Therefore, the reaction substrate molar ratio of 1:1 is more suitable, at which the esterification rate of S-FFA and the S-DAG content are higher.

The effect of enzyme addition on the esterification effect is shown in Figs. 1C and 2C. As shown in Fig. 1C, Lipozyme® RMIM was used as a catalyst, when the enzyme addition was less than 3%, the esterification rate increased with the increase of enzyme addition, but the change trend was slow; when the enzyme addition was greater than 3%, the esterification rate decreased with its increase. It may be because when the enzyme addition is less, the reaction rate has not reached saturation, and the esterification rate will increase with the increase of enzyme addition. However, when the enzyme is added too much, the lipases stick to each other, the viscosity of the reaction system increases, the contact area and contact frequency between the lipase and the reactant decrease, and the esterification rate decreases. The content change trend of S-DAG prepared by Lipozyme® RMIM esterification was consistent with the change trend of its esterification rate. When the enzyme addition is 3%, the S-DAG content is the highest, about 54.46%. The results in Fig. 2C revealed that when Novozym® 435 was used to catalyze esterification, the overall esterification rate was high, and the S-DAG content was the highest at 3% enzyme addition, was 55.30%. Therefore, it is better to choose a lipase addition of 3%.

The effect of reaction temperature on the esterification effect is shown in Figs. 1D and 2D. As shown in Fig. 1D, with Lipozyme® RMIM as the catalyst, the esterification of S-FFA first increased with increasing temperature, and the esterification rate was the highest at 50 °C, and then the esterification rate began to decrease, while the content of S-DAG had always been positively correlated with the reaction temperature. When the reaction temperature was 70 °C, the S-DAG content is the highest, about 66.48%. Fig. 2D shows that with Novozym® 435 enzyme as the catalyst, the esterification of S-FFA first increased with increasing temperature, and the change trend of S-DAG content slowed down between 50 °C and 60 °C. When the reaction temperature exceeded 60 °C, the esterification rate began to decline. The change trend of S-DAG content was basically consistent with the change trend of its esterification rate. When the reaction temperature was 50 °C, the S-DAG content was the highest, 72.99%, and when the temperature exceeded 50 °C, the S-DAG content gradually decreased. This may be because when the reaction temperature is low, the reactant molecules move slowly, as the temperature increases, the molecular movement speed increases, the collision frequency between molecules increases, and the esterification effect is enhanced. However, when the reaction temperature is too high, the protein structure in the lipase is destroyed, resulting in a decrease in lipase activity and a poor esterification effect. Therefore, considering all factors, it is more appropriate to select 50 °C as the reaction temperature.

3.3. Analysis of glyceride composition of S-DAG

S-DAG was prepared under the optimal conditions of pre-hydrolysis-esterification, and the glyceride composition was analyzed, as shown in Figs. 3.

Fig. 3.

Chromatographic characterization and compositional analysis of structured diacylglycerols (S-DAG). (A) Total ion current (TIC) chromatogram of the glycerides in the DAG oil. (B) Glyceride composition of DAG oil, expressed as the relative percentage (%, w/w) of DAG, MAG, and TAG based on TIC peak areas.

As shown in Fig. 3(A), MAG, DAG, and TAG showed good responses in the ESI⁺ mode and were ionized into [M + NH₄]⁺. The glyceride contents in S-DAG were added together to calculate the percentage of each type of glyceride, as shown in Fig. 3(B). A total of 138 glycerides were identified by UPLC-QTOF-MS, which were mainly divided into MAG, DAG, and TAG in terms of glyceride types. MAG has 16 lipid molecules, accounting for 19.8% of the total. TAG has 72 lipid molecules, but only 7.2% of the total. DAG is the main glyceride, with 50 lipid molecules, accounting for 73.0% of the total, among them, DAG 36:4 (C₃₉H₆₈O₅) has the highest content, which is 36.9%, followed by DAG 36:3 (C₃₉H₇₀O₅), accounting for 34.4% of the total DAG content, it is formed by the esterification of two linoleic acid molecules or one linoleic acid and one oleic acid molecule with one glycerol molecule and is commonly found in vegetable oils.

3.4. Esterification thermodynamics and kinetics simulation

The experimental data were linearly fitted with 1/T as the horizontal axis and lnv as the vertical axis (Bashiri & Pourbeiram, 2016), and the results were shown in Fig. 4. As the temperature increased, the initial reaction velocity continued to increase. According to formula (9), the activation energy E_a = k·R can be calculated. Given that R is 8.3145, it can be concluded that the E_a of the esterification reaction catalyzed by Lipozyme® RMIM is 10.58 KJ/mol (Fig. 4A), and the E_a of the esterification reaction catalyzed by Novozym® 435 is 10.33 KJ/mol (Fig. 4B). The E_a range of the esterification reaction catalyzed by most lipases is 0.97–34.5 KJ/mol, and the E_a of the esterification reactions of S-FFA catalyzed by the two enzymes are both within this range.

Fig. 4.

Relationship between reaction rate and reaction temperature esterification by different enzymes (A: Lipozyme® RMIM, B: Novozym® 435); Lineweaver-Burk double reciprocal diagram of esterified with different lipases. (C: Lipozyme® RMIM, D: Novozym® 435).

The Arrhenius equations are (12), (13):

$$\ln v=1.9480-\frac{1272}{T}$$ (12)

$$\ln v=1.8428-\frac{1243}{T}$$ (13)

The activation energy E_a is the minimum additional energy required for reactant molecules to reach the transition state in which a chemical reaction can occur. In the enzymatic esterification of oils, the activation energy E_a represents the energy barrier that the reaction system needs to overcome in the process of converting the substrate into the product. The E_a of the esterification reaction catalyzed by Lipozyme® RMIM is slightly greater than that of the esterification reaction catalyzed by Novozym® 435 enzyme, indicating that the energy barrier to be overcome for Lipozyme® RMIM catalyzed esterification is slightly greater than that for Novozym® 435 catalyzed esterification. Furthermore, the greater the activation energy of the reaction, the smaller the fraction of activated molecules, the fewer the number of activated molecules, and thus the smaller the reaction rate. Therefore, the rate of the esterification reaction catalyzed by Novozym® 435 enzyme is greater.

The data of the determination of the Michaelis equation constant were shown in Tables S1 and S2. Fig. 4C and D were Lineweaver-Burk double reciprocal plots of S-FFA esterification using different lipases as catalysts. The experimental data were linearly fitted with 1/[S] as the horizontal axis and 1/v as the vertical axis to obtain the Michaelis constant Km and the maximum reaction rate v_m of the reaction (Phuah et al., 2012). The results were shown in Table S3. The value of K_m represents the relationship between the affinity of lipase for a specific substrate and the reaction efficiency. As can be seen from Table S3, the K_m value of the S-FFA esterification reaction catalyzed by Novozym® 435 enzyme is the smallest. A lower K_m value signifies a higher apparent affinity of the enzyme for its substrate. This indicates that the substrate concentration at which the reaction rate in the Novozym® 435 enzymatic hydrolysis system reaches half of the maximum value is lower than that of the Lipozyme® RMIM enzyme. The Novozym® 435 enzyme has the strongest ability to bind to the substrate and has the best esterification effect, which is consistent with the experimental results. According to Table S3, when the reaction conditions remain unchanged, the esterification reaction rate of Novozym® 435 enzyme is greater than that of Lipozyme® RMIM. Based on the results of thermodynamics and kinetics, the esterification effect of Novozym® 435 enzyme is better.

3.5. Comparative evaluation of swida wilsoniana fruit oil and its DAG

3.5.1. Physical and chemical index determination

In order to explore the effects of refining treatment and enzyme-catalyzed reaction on the physical and chemical properties of Swida wilsoniana fruit oil, the color, transparency, acid value, saponification value and iodine value of crude Swida wilsoniana fruit oil (Crude oil), refined Swida wilsoniana fruit oil (Refined oil), diglyceride derived from crude Swida wilsoniana fruit oil (Crude oil-DAG) and diglyceride derived from refined Swida wilsoniana fruit oil (Refined oil-DAG) were tested respectively, and the results were shown in Table S2.

Table S4 showed that the transparency of the Swida wilsoniana fruit oil after refining is higher, the color is lighter, and the saponification value is lower, but the iodine value and acid value do not change much. Crude oil is obtained by low-temperature pre-pressing-subcritical n-butane method, so the acid value itself is low, and the acid value changes little after refining. After the crude oil was subjected to enzyme catalysis, the saponification value decreased slightly, the color became lighter, and the transparency increased. It should be that during the enzyme catalysis reaction, the lipase adsorbed certain impurities and pigments, and the centrifugal operation during the enzyme catalysis reaction removed impurities in the crude oil. At the same time, the iodine value difference between crude oil and S-DAG is small, indicating that the degree of unsaturation of fatty acids in the oil does not change much before and after the enzyme catalysis reaction. In addition, the physical and chemical indicators of Crude oil, Refined oil, Crude oil-DAG, and Refined oil-DAG all meet the requirements of “GB 2716-2018 National Food Safety Standard Vegetable Oil”.

3.5.2. Fatty acid composition analysis

The fatty acid compositions of the Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG were shown in Fig. 5A-5D and Table 1. The results showed that the main fatty acid composition of the Swida wilsoniana fruit oil was linoleic acid (C18:2), oleic acid (C18:1), palmitic acid (C16:0), linolenic acid (C18:3), stearic acid (C18:0) and palmitoleic acid (C16:1). The palmitic acid and oleic acid contents of crude oil were slightly higher than those of refined oil, and the linolenic acid content in S-DAG oil was slightly lower than that in crude oil. However, the fatty acid compositions of crude oil, refined oil, crude oil-DAG and refined oil-DAG were very similar. The fatty acid composition represents the physical and chemical properties of the oil itself (Ma et al., 2011). This indicates that the oil refining process and enzyme catalysis reaction have little effect on the fatty acid composition of the safflower oil itself.

Fig. 5.

Gas chromatograms, tocopherol composition (E) and oxidation induction times (F) of Crude oil (A), Refined oil (B), Crude oil-DAG (C) and Refined oil-DAG (D).

Table 1.

Fatty acid composition of Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG.

Fatty acid composition	Crude oil/%	Refined oil/%	Crude oil-DAG/%	Refined oil-DAG/%
C16:0	18.084	16.809	18.092	16.955
C16:1	1.113	1.175	1.119	1.157
C18:0	1.828	1.921	1.847	1.927
C18:1	33.431	30.317	33.91	30.275
C18:2	44.408	46.316	42.849	46.369
C18:3	2.262	3.091	2.183	2.965

3.5.3. Tocopherol content determination

The test results of the tocopherol content of the Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG were shown in Table 2 and Fig. 5E. The results in Table 2 and Fig. 5E showed that the δ-tocopherol content in the Swida wilsoniana fruit oil is the highest, followed by α-tocopherol and β-tocopherol, and γ-tocopherol is not detected. After refining, the total tocopherol content decreased from 5020.018 mg/100 g to 4511.616 mg/100 g, with a retention rate of 47.3046%. Among them, the α-tocopherol content was not detected after refining, and the retention rates of β-tocopherol and δ-tocopherol were 19.17% and 49.94%, respectively. The refining process involves deodorization, decolorization, deacidification, degumming and washing. The high temperatures, adsorbents, soap residue, and water involved in these steps all contribute to the decomposition and loss of tocopherol (Choe, 2013). After the enzyme-catalyzed reaction, the composition and content of tocopherol decreased to varying degrees. This may be due to the enzyme indirectly promoting oxidation, consuming tocopherol, and the loss of tocopherol during the washing process in the enzyme separation process. Therefore, to maximize the preservation of the bioactivity in oils, high temperatures and fewer processing steps should be avoided during processing, and enzymes with minimal impact on tocopherol should be selected.

Table 2.

The tocopherol content of Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG.

Composition	Crude oil (mg/100 g)	Refined oil (mg/100 g)	Crude oil-DAG (mg/100 g)	Refined oil-DAG (mg/100 g)
α- tocopherol	220.5388	–	200.5663	–
β- tocopherol	71.7125	13.75	60.2875	9.5
δ- tocopherol	4727.766	2360.95	4250.763	1915.6
Total amount	5020.018	2374.7	4511.616	1925.1

3.5.4. Oxidation stability analysis

The reaction temperature was set at 110 °C, and the oxidation stability of the Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG was shown in Fig. 5F. As can be seen from Fig. 5F, the oxidation induction time of the Swida wilsoniana fruit oil is greater than 24 h, and its oxidation stability is better than that of most oils and fats (Li et al., 2025). After refining and enzyme-catalyzed reaction, the oxidation induction time of the Swida wilsoniana fruit oil became shorter and its oxidation stability decreased. It may be that the high reaction temperature during the refining process and enzyme-catalyzed reaction causes the oxidation of the safflower oil (Yuan et al., 2018). At the same time, the content of MAG in S-DAG is higher than that in crude oil, MAG has only a single ester bond and is more easily destroyed than DAG and TAG under heating conditions. Therefore, crude oil has better oxidation stability, followed by crude oil-DGA.

3.5.5. Analysis of volatile flavor compounds

The volatile flavor compounds of Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG were analyzed using HS-GC-IMS, which can intuitively distinguish the flavor differences between different oil samples (Corona et al., 2023;Guo et al., 2018), the three-dimensional spectra and two-dimensional top-view images of the volatile components of the four oil samples were shown in Figs. 6 a and Fig. 7, respectively.

Fig. 6.

3D spectra (a), relative percentage of volatile compounds (b) and fingerprint of volatile flavor substances (c) for Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG.

Fig. 7.

2D top view and difference diagram of the four oils. (a is the 2D top view, b, c, d and e are the difference graphs with Crude oil, Refined oil, Crude oil-DAG and Refined oil-DAG as the main plots, respectively).

Each peak in the three-dimensional spectrum represents a different volatile compound (Geng et al., 2023). The results of Fig. 6 a displayed that there are significant differences in the types and contents of volatile compounds between crude oil and refined oil, the types of volatile substances between crude oil-DAG and refined oil-DAG are slightly different, and the main difference is in the content. A total of 40 volatile substances were identified in the GC-IMS library. The main volatile substance components and their relative contents in the four oil samples were shown in Fig. 7 and Table S5. These include 6 alkanes, 1 alcohol, 1 acid, 2 esters, 6 ketones, 6 benzenes, 4 aldehydes, 8 alkenes, 1 thiazole, 1 pyridine, and 2 others. The results of Fig. 6 b and Table S5 suggested that the main volatile compounds in the fruit oil of the light-skinned tree are alkenes, alkanes, ketones, and aldehydes. In Fig. 7, the brighter the color and the warmer the hue, the greater the concentration of the compound. In the difference graph, the dark blue background is the main graph, and the other three graphs are the difference comparison graphs of the main graph. The blue area represents that the compound represented by this point in this sample is lower than the content of the compound in the main graph, and the red area represents that the content of this compound is higher than that in the main graph. Fig. 7 and Table S5 demonstrated that crude oil has a higher content of volatile compounds and a stronger flavor, followed by crude oil-DAG. Refining treatment and enzyme-catalyzed reaction will reduce the volatile flavor compounds in the fruit oil of the light bark tree to varying degrees, which may be because the activated white clay in the bleaching process and the lipase in the enzyme-catalyzed reaction process will adsorb volatile flavor substances.

The results in Fig. 6 b revealed that the content of olefin compounds is the highest among the four oil samples, followed by benzene ring compounds, ketone compounds and ester compounds. Olefin compounds are associated with light herbaceous, or certain fruity aromas; benzene ring compounds often contribute aromatic, sweet, or nutty flavors; ketone compounds have a weaker contribution to flavor and may have subtle floral, fruity, or cheese-like notes; while ester compounds often possess prominent fruity aromas, such as the pleasant scents of apples and bananas. After enzyme catalysis, the content of olefin compounds and ester compounds in the light-peeled tree fruit oil increased, while the content of ketone compounds and alkane compounds decreased. This change may result in a purer flavor and enhanced fruity aroma of the oil. After refining, the content of olefin compounds and ester compounds in the light-peeled tree fruit oil decreased, while the content of ketone compounds and aldehyde compounds increased. Ketone compounds have a higher threshold and contribute less to flavor, aldehyde compounds also have a high threshold but often contribute grassy,  fatty, or citrus flavors to the oil. Oil oxidation during the refining process will lead to the production of ketone compounds and aldehyde compounds, which may bring a more pronounced oxidized flavor, astringency, or a faint fatty taste to the oil, while the original fruity characteristics are diminished.

Fig. 6 c presented the fingerprints of volatile substances in four oil samples: crude oil, refined oil, crude oil-DAG, and refined oil-DAG. Each row represents all the signal peaks selected from a sample, and the brightness and area of each color block in each column indicate the concentration of the compound it represents (Christmann et al., 2022). The fingerprint can more intuitively compare the differences in volatile flavor substances of different oil samples. The fingerprint is divided into eight parts: a, b, c, d, e, f, g, and h using red boxes.

The spectrum in zone a suggested that the volatile substances are higher in crude oil, crude oil-DAG, and refined oil-DAG, and lower in refined oil. They include Benzene, 1-Ethyl-2-methyl-benzene, 2,3-Butanedioldiacetate, which are benzene compounds and an acid. Acids are one of the products of oil oxidation. Benzene compounds have strong aromaticity, 1-Ethyl-2-methyl-benzene has a special aromatic smell and is highly volatile. The volatile substances in zone b are mainly 3-Iodobenzotrifluoride and Nonanal, which are benzene compounds and nonanal. Nonanal belongs to the alkanal class of compounds, is green and has a fatty smell (Xu et al., 2023), and is a product of fatty acid oxidation. Nonanal is detected in higher amounts in crude oil-DAG and refined oil-DAG, which may be caused by the oxidation of crude oil after enzyme catalysis.

Area c includes two alkenes, (E)-ocimene and cyclopentene, and one ketone, 2-butanone and 3,3-dimethyl. These compounds were present in high levels in Crude oil-DAG. Ketones mainly have aromatic and creamy flavors, but the threshold of ketones is high and has little effect on the flavor of oil. Ketones are mainly produced by the oxidation of unsaturated fatty acids, Maillard reaction and alcohol oxidation of oils. The alkenes are (E)-β-ocimene and cyclopentene. Cyclopentene mainly has an aromatic smell. (E)-β-ocimene was first isolated from basil oil and has a warm medicinal and citrus aroma (Farré - Armengol et al., 2017). Area d contains 2,3-dimethylbutane and an unidentified volatile compound, which is mainly detected in Refined oil-DAG.

The compounds in zone e are (E)-2-hexenal and 2-butanone. (E)-2-hexenal mainly has fruity and fatty aromas and is an oxidation product of oleic acid and linoleic acid. (E)-2-hexenal is present at higher levels in Refined oil-DAG and Crude oil-DAG. 2-Butanone was detected in all four oil samples, but its concentration was higher in Crude oil and Refined oil. Studies have shown (Luo et al., 2022) that 2-butanone is produced by fat oxidation and Maillard reaction, and has a creamy and mush roomy flavor. It is one of the main volatile compounds in cold-pressed peanut oil (Dun et al., 2019). Hexanal, β-Pinene, 2-methylthiazole, 2-Ethylpyridine and two alkanes, Norbornane and Methylene Chloride, can be observed in zone f. β-pinene mainly has an aromatic and lemony aroma, while 2-ethylpyridine has a caramel aroma (Zeng et al., 2024). 2-methylthiazole has a popcorn roasted flavor, which is produced by the Maillard reaction induced by glucose and lysine, arginine and histidine (Zheng et al., 2024). Hexanal has a fruity, fatty and beef tallow flavor, which is caused by the decomposition of linoleic acid and linolenic acid during oil oxidation (Kalua et al., 2007). Gulzar and Benjakul (2020) have shown that as the hexanal content increases, the sensory evaluation of the sample will decrease accordingly.

Zone g contains two olefin compounds, 2-heptene-3-methyl and 2-methyl-1-pentene, an ester compound, methyl pivalate, and a heterocyclic compound, 1,3-dioxolane-2,4-dimethyl. Among them, 2-methyl-1-pentene has a pleasant lemon aroma, and the heterocyclic compound 1,3-dioxolane-2,4-dimethyl is considered to be the product of Maillard reaction and Strecker degradation reaction, which can cause the food to have a roasted and nutty flavor. Methyl pivalate has a cheesy banana flavor. Ester compounds are relatively important aroma substances, and they play an important role in the flavor of the product even at low concentrations, and have a pleasant fruity aroma (Sun et al., 2021). It is worth noting that Jing et al. (2022) found that the concentration of methyl acetate in solvents and fruits is relatively high, while the concentration in pomace oil is relatively low.

There were many compounds in zone h, including three benzene rings: Benzene, 1-ethenyl-4-ethyl, 1-Ethyl-2-methyl-benzene and toluene, four ketone compounds: m-Menth-6-ene, 3-Pentanone, 2-heptanone and 2,3-Dimethyl-2-cyclopenten-1-one, three alkane compounds: isopropyl ethylphosphonofluoridate, 1,2,4,5-tetraoxacyclohexane and 3,3,6,6-tetramethyl-2,3-Dimethylbutane, one ester compound: methyl 3-(methylthio)propionate, one alcohol compound: 3-Methyl-3-buten-1-ol, and three olefin compounds: 1-Octene, cis-3-Methyl-2-pentene and 2-Methyl-2-pentene. m-Menth-6-ene has a minty aroma, 2-heptanone has a creamy aroma, heptanone is produced by oxidation of linoleic acid and has an oily, fruity, grassy and creamy flavor, 2,3-Dimethyl-2-cyclopenten-1-one has a caramel nut aroma, 3-Pentanone has a grassy aroma. Jia et al. (2024) used HS-GC-IMS to detect pentanone in microwave camellia oil, which is a low molecular weight aromatic compound. The olefin compounds are mainly 1-Octene, cis-3-Methyl-2-pentene, and 2-Methyl-2-pentene, which mainly have a fragrant and refreshing aroma. 2,3-Dimethylbutane mainly has a fruity aroma, methyl 3-(methylthio) propionate has a fresh floral and fruity aroma, and 3-Methyl-3-buten-1-ol has a nail polish and alcohol smell. Alcohol substances are mainly produced by a series of decomposition or enzymatic reactions of monohydroperoxides produced by air oxidation of vegetable oils (Koch et al., 2022). It is evident that woody plant oils, represented by the Swida wilsoniana fruit oil, not only have antioxidant and anti-inflammatory effects, but also contain alkenes, aldehydes, esters, ketones, and other flavor compounds that enhance their appeal and acceptability in the diet, encouraging consumers to incorporate them into their daily diet, replacing some traditional oils and achieving dietary diversification.

In order to further clarify the differences in volatile compounds among the four oil samples of Refined oil, Crude oil, Refined oil-DAG, and Crude oil-DAG, the Dynamic PCA plug-in was used to draw the PCA diagram, as shown in Fig. S3.

As shown in Fig. S3, the contribution rate of PC1 and PC2 is 60.9% and 30.1%, respectively, and the cumulative contribution rate reaches 91%, indicating that PC1 and PC2 can reflect most of the information of volatile flavor substances. In the PCA diagram, the closer the distance between samples, the smaller the difference within the sample group. The results revealed that there were significant differences in the volatile components among the four oil samples of Refined oil, Crude oil, Refined oil-DAG, and Crude oil-DAG. The distances between Refined oil and Crude oil, between Refined oil and Refined oil-DAG, and between Crude oil and Crude oil-DAG were far, indicating that there were large differences in their volatile flavor substances. The volatile flavor substances between Refined oil-DAG and Crude oil-DAG after enzyme-catalyzed hydrolysis and esterification were relatively close. Therefore, both the refining process and the enzyme-catalyzed process will have a certain impact on the volatile flavor substances of the Swida wilsoniana fruit oil.

4. Conclusion

This paper describes a two-step enzymatic hydrolysis-esterification method for preparing diglycerides (DAG) from the Swida wilsoniana fruit oil. Through enzyme screening, kinetic analysis, and systematic evaluation of oil samples, the paper not only verifies the feasibility of this technical route but also reveals the comprehensive impact of the processing on key quality indicators such as the nutritional content, stability and flavor of the oil.

For the first time, a clear process-quality correlation framework has been established for the enzymatic preparation of DAG from the Swida wilsoniana fruit oil from the perspective of kinetic mechanisms and overall quality evolution. The study shows that the combined use of Lipozyme® RM IM and Novozym® 435, with its specific catalytic properties (Sn-1, 3-position selectivity) and superior kinetic parameters (low activation energy E_a and low Michaelis constant K_m), can achieve highly efficient hydrolysis and esterification reactions, providing an efficient biocatalytic solution for the structural modification of similar oils. However, the study also clarified the key limitations of this process: both refining pretreatment and enzymatic catalysis irreversibly reduce the total tocopherol content and oxidative stability of the oil, and may significantly weaken its original flavor complexity through refining steps. This reveals that the current process, while pursuing high DAG yields, comes at the cost of sacrificing some natural active ingredients and flavor substances, indicating room for optimization in terms of nutritional and sensory quality preservation.

Based on the above conclusions, the following suggestions are made for future industrial practices: First, in terms of raw material selection, unrefined crude oil should be prioritized for DAG preparation to maximize the retention of natural antioxidants such as tocopherols and volatile flavor compounds, thereby producing DAG products with superior nutritional and sensory qualities. Second, in terms of process optimization, future research should focus on developing mild refining technologies or exploring direct enzymatic modification processes for unrefined raw materials to reduce the damage to active substances caused by pretreatment. Simultaneously, the addition of exogenous natural antioxidants to the reaction system or the use of an inert atmosphere can be investigated to mitigate the degradation and oxidation of active ingredients during enzymatic catalysis. Finally, it is recommended to establish a multi-dimensional quality evaluation system covering nutrition, stability, flavor, and functionality to more comprehensively assess and guide the development of functional oil products.

CRediT authorship contribution statement

Jingjing Xiao: Writing – original draft, Methodology, Investigation, Data curation. Yujie Xu: Writing – review & editing, Data curation. Yongjun Miao: Investigation, Data curation. Sisi Liu: Writing – review & editing, Supervision. Jia Tu: Writing – review & editing, Supervision. Baining Lin: Methodology, Investigation. Zengmin Kuang: Supervision, Methodology, Investigation. Zhihong Xiao: Supervision, Funding acquisition, Conceptualization. Rukuan Liu: Writing – review & editing, Supervision. Li Li: Writing – review & editing, Supervision. Changzhu Li: Supervision, Methodology. Aihua Zhang: Investigation, Data curation. Daliang Jiang: Writing – review & editing, Supervision.

Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Zhihong Xiao reports financial support was provided by the National Key Research and Development Program of China. Zhihong Xiao reports a relationship with the National Key Research and Development Program of China that includes: funding grants. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A. Supplementary data

Supplementary material

Data availability

Data will be made available on request.