Effects of Lactobacillus fermentation on the phenolic and flavor substances of coconut water

Effects of Lactobacillus fermentation on the phenolic and flavor substances of coconut water Hao-Ran Zhang1, Yu-Huan Wang1, Zi-Yi Peng1, Yun-Peng Hao1, Xiao-Peng Wang1 1 College of Food Science and Technology, Henan Agricultural University, 95 Wenhua Road, Zhengzhou, Henan Province 450002, China Corresponding author: Yun-Peng Hao (haoyp[email protected]m) Academic editor: Maria Manuela Silva♦Received 13 May 2025♦Accepted 2 October 2025♦Published 15 October 2025 Abstract In this study, Lactobacillus strains with beneficial biofunctions were screened and applied to ferment coconut water. The effects of Lactobacillus fermentation on the phenolic and flavor components of coconut water were investigated. The results showed that the strains Lactobacillus casei U2.1 and Lactobacillus paracasei M1.5 possessed excellent DPPH radical scavenging (30.91%) and α-amylase inhibitory activity (57.44%), and both of them survived over 50% in simulated gastrointestinal fluids. The co-fermentation of the two strains elevated the total phenolic content of coconut water by 19%, with a significant increase in free phenols and insoluble glycoside-type phenols (P < 0.05), indicating that lactobacilli fermentation promoted the release and conversion of phenolics. Through gas chromatography-mass spectrometry (GCMS) analysis, it was found that the variety of volatile flavor substances increased by 27 species after fermentation, and the generation of key flavor substances (phenylethanol and ethyl caprylate) enhanced the aroma quality of the coconut water. In addition, the accumulation of potential physiologically active components (palmitoleic acid and cis-5-dodecenoic acid) further enhanced the nutritional value of the product. In conclusion, the co-fermentation of the two strains improved the nutritional value and flavor of coconut water, which provided a theoretical basis for the development of fermented coconut water beverage. Keywords Coconut, Lactobacillus, phenolic, volatile flavor substances Introduction Coconut (Cocos nucifera L.) belongs to the Arecaceae family, and is an important cash crop, mainly distributed in Indonesia, Brazil, the Philippines and other regions. In mature coconuts, coconut water accounts for about 18% of the total weight. Coconut water is a clear nutritious liquid from the endosperm of the coconut, with a unique coconut flavor, rich in minerals, vitamins and electrolytes, making it a natural electrolyte drink. Meanwhile, coconut water is rich in phenols and flavonoids, which makes it a high-quality fermentation raw material. Coconut water has antioxidant, hypolipidemic antibacterial and antidiabetic properties (DebMandal et al. 2011). Coconut water can be categorized into immature and mature coconut water based on its maturity at the time of harvest. Raw coconut water is low in fat, contains sugar, amino acids, vitamins and minerals and can be consumed directly or processed into various beverages (Hahn 2012). The nutritional value of mature coconut water does not change significantly, but the increase in crude fat content leads to deterioration in flavor (Zhang et al. 2018a, 2018b). Moreover, coconut water is often discarded during the production of coconut milk or cream, resulting in a waste of resources. Therefore, new processing methods need to be explored to develop high-value coconut water products. Fermentation is a traditional method of food processing and preservation, and Lactobacillusfermented products representing a significant category. Lactobacilli are a group of active microorganisms that improve the host’s intestinal Copyright AuthorFamily, et al. This is an open access article distributed under the terms of the Creative Commons Attribution License (CC-BY 4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited. Emirates Journal of Food and Agriculture 37: 1–13 doi: 10.3897/ejfa.2025.158757 RESEARCH PAPER AuthorFamily, et al.: Lactobacillus in coconut water fermentation 2 Emirates Journal of Food and Agriculture environment and have positive effects on health, such as promoting nutrient absorption, alleviating lactose intolerance, and regulating intestinal health (Teng et al. 2020). Microorganisms of the genus Lactobacillus are mainly associated with the production of fermented foods (Plessas et al. 2017), altering the biochemical and organoleptic properties of fermented raw materials and providing abundant nutrients such as minerals, vitamins, amino acids and digestive enzymes (Ray et al. 2016; Du et al. 2024; Yang et al. 2024). In addition, lactobacilli are widely used as probiotics due to their high safety and diverse probiotic functions. Lactobacillus fermentation can change the composition of the substances in the plant-based substrate, so that the flavor substances in the product can be changed and the functionality can be enhanced. It has been shown that the fermentation of L. plantarum increased the total phenolics and antioxidant capacity in blueberry juice (Zhang et al. 2021). Furthermore, Kantachote (2017) and Giri (2018) found that coconut water fermented by different Lactobacillus had improved flavor and nutrients in the product as well as a better storage quality. In this study, Lactobacillus with good basic properties and potential physiological functions (antioxidant and hypoglycemic) were screened and applied to coconut water fermentation. And the effects of Lactobacillus fermentation on the content, composition and type of phenolics and volatile flavor substances in coconut water were further investigated. The result would provide a theoretical basis for the development of novel fermented coconut water products. Materials and methods Experimental reagents Coconut water: The coconuts produced in Wenchang, Hainan, were cracked open to extract the water, which was then filtered through four layers of gauze, mixed, and stored at 4 °C. Strain source: L. casei U2.1, L. paracasei O3.3, L. fermentum S1.5, L. paracasei M1.5 were isolated from traditional fermented foods (Chinese sauerkraut, kimchi) and preserved in College of Food Science and Technology, Henan Agricultural University, and L. rhamnosus LGG was isolated from live bacterial powder. All other reagents used in the test were analytically pure and purchased from general suppliers. Strain culture conditions The strains were streaked onto MRS solid medium using the streak plate method and subsequently incubated at 37 °C for a duration of 48 h. Subsequent to this period, a well-isolated single colony was transferred to MRS liquid medium and cultivated at the same temperature for an additional 24 h. The strains were subcultured twice in this manner. Preparation of Bacterial Suspension: After activating Lactobacillus for two generations, the bacterial cells were separated by centrifugation (10000 × g, 4 °C, 5 min). The concentration of viable bacteria was subsequently adjusted to 1 × 108 CFU/mL using phosphate-buffered saline (PBS, pH 7.0) based on the viable count. Basic properties of Lactobacillus strains Growth curves, pH and acidity measurement The activated bacterial solution was transferred to MRS liquid medium at an inoculum of 1% (v/v) and incubated at 37 °C for 24 h. Samples were taken at 0 h, 2 h, 4 h, 8 h, 12 h, 14 h, 16 h, 18 h, 20 h, 22 h, and 24 h to determine the OD600, pH values, and acidity. A pH meter was used to measure the pH, while the total acidity, expressed as lactic acid, was determined by the titration method. Simulated gastrointestinal fluid tolerance assay Preparation of simulated gastric and intestinal fluids was performed with reference to Son et al. (2018).1 mL of bacterial suspension was inoculated into 9 mL of simulated gastric and simulated intestinal fluids, respectively, and incubated with shaking and mixing at 37 °C for 3 h. Samples were taken at 0 and 3 h to calculate the number of viable bacteria in the sample by the plate count method. The results were calculated according to formulas (1) and (2). (1) Where NGas. indicates gastric fluid tolerance survival rate. (2) where NInt. indicates intestinal fluid tolerance survival rate. Measurement of the reducing power of the strains 1 mL of the bacterial suspension was mixed with 2.5 mL K3[Fe(CN)6] (1%) solution and 2.5 mL of PBS buffer (0.2 mol/L, pH 6.6). After a water bath at 50 °C, the suspension was cooled in ice water for 5 min. Subsequently, 2.5 mL trichloroacetic acid (TCA) solution (10%) was added to the cooled sample, followed by centrifugation at 4 °C and 3500 × g for 10 minutes. Following centrifugation, 2.5 mL of the supernatant was extracted, to which 2.5 mL of distilled water and 0.5 mL FeCl3 (0.1%) solution were added. The absorbance was then measured at 700 nm (Li et al. 2012). Measurement of free radical scavenging rate of ATBS The ABTS radical-scavenging activity was determined spectrophotometrically using the method described by Delgado et al. (2019). The ABTS working solution was prepared by diluting ABTS stock (7.68 g/L ABTS, 13.24 g/L K2S2O8 in pure water) with anhydrous ethanol to achieve a specific absorbance value of 0.7 ± 0.02 (A value). A sample of bacterial suspension (1.0 mL) was mixed with the Emir. J. Food Agric ⋅ Volume 37 ⋅ 2025 3 Emirates Journal of Food and Agriculture ABTS working solution (5 mL) and incubated in the dark at room temperature for 5 minutes before the absorbance was measured. The absorbance of the sample was measured at 734 nm as A0. The results were calculated using formula (3). (3) Where NABTS. indicates the ABTS free radical scavenging rate. Measurement of DPPH scavenging capacity The scavenging capacity of DPPH free radicals was determined spectrophotometrically (Oh et al. 2020). Briefly, 2.0 mL of bacterial suspension was mixed with 4.0 mL of DPPH solution (0.1 mmol/L). After incubation for 25 min at room temperature in the dark, the reaction mixture was centrifuged at 5000 × g for 5 min at 4 °C. Sample controls were prepared by mixing with an equal volume of anhydrous ethanol instead of the DPPH solution. For the control group, distilled water was used in place of the DPPH solution. The results were calculated using formula (4). (4) Where NDPPH. denotes DPPH clearance rate. ODS is sample group; ODSC is the sample control group; and ODC is the control group. Measurement of α-glucosidase inhibition rate α-glucosidase inhibition rate was performed according to Zhang et al. (2007). 100 μL of α-glucosidase (0.4 U/mL) was added to 50 μL of bacterial suspension and incubated for 10 minutes at 37 °C. Subsequently, 100 μL of PNPG (2.5 mmol/L) was added to each reaction tube, and incubation continued for 30 minutes at 37 °C. Finally, the reaction was terminated by the addition of 50 μL of Na2CO3 solution (1 mol/L).The absorbance at 405 nm was measured for the sample group, sample control group, control group and blank group. The results were calculated according to formula (5). (5) In this case, the sample control is a mixture of PNPG and sample without added enzyme. The control is a mixture of PNPG and enzyme without added sample. The blank group was PBS solution (0.1 mol/L, pH 6.8). Where NGlu. denotes the rate of glucosidase inhibition. ODS is the sample group; ODSC is the sample control group; ODC is the control group; ODB is the blank group. Measurement of α-amylase inhibition rate α-amylase inhibition was performed according to Meng et al. (2016). 0.25 mL of bacterial suspension was mixed with 0.25 mL of α-amylase (1.0 mg/mL) and incubated for 10 min at 37 °C. Subsequently, 0.5 mL of starch solution (1%) was added, and the incubation was continued for another 5 min at 37 °C. The reaction was terminated by adding 1 mL of DNS solution, and the mixture was then heated in a boiling water bath for 5 min before being rapidly cooled to room temperature. The reaction products were diluted using PBS. The absorbance at 540 nm was measured for the sample group, sample control group, control group and blank group. The results were calculated using formula (6). (6) In this case, the sample control is a mixture of starch solution and sample without added enzyme. The control is a mixture of starch solution and enzyme without added sample. The blank group was PBS solution (0.1 mol/L, pH 6.8). Where NAmy. denotes amylase inhibition rate. ODS is the sample group; ODSC is the sample control group; ODC is the control group; ODB is the blank group. Coconut water fermentation Preparation of fermented coconut water For fermentation, a mixture of L. paracasei M1.5 and L. casei U2.1, obtained from previous screening, was used as the fermenting agent at a mixing ratio of 1:1 (n:n). And the inoculum inoculated into the fermentation material was 4% (v/v) of the inoculum to the fermentation material. Following inoculation, the coconut water was fermented at a constant temperature of 35 °C for 22 h. At the conclusion of the fermentation process, the products were placed in a cold room at 4 °C for 24 h of post-ripening. Determination of phenolic content Polyphenol standard curve Standard curves for phenol were created using the Folin-Ciocalteu reagent (Managa et al. 2021). Standard solutions of varying concentrations were treated with the Folin-Ciocalteu reagent (10%, 2.5 mL) for 8 min, followed by the addition of sodium carbonate (7.5%, 2 mL). After incubation in the dark for 60 min, the absorbance was measured at 765 nm. The standard curve was then plotted with gallic acid concentration on the horizontal axis and absorbance values on the vertical axis. Preparation of free phenolics The extraction of phenol followed the method described by Ayaz et al. (2005) with slight modifications. 5 mL of fermented coconut water was taken and extracted with 25 mL of 80% (v/v) methanol solution. Subsequently, the mixture was centrifuged (10000 × g, 10 min, 4 °C). The filtrate was extracted twice by repeating the same procedure and the resulting supernatant was collected as soluble phenolic components. The soluble phenol obtained through centrifugation was concentrated to dryness using a rotary evaporator at 35 °C. Subsequently, the concentrate AuthorFamily, et al.: Lactobacillus in coconut water fermentation 4 Emirates Journal of Food and Agriculture was redissolved in 40 mL of distilled water. The mixture was then degreased using hexane, and the aqueous phase was separated. The aqueous phase was separated three times using ethyl acetate. The liquid from the combined extractions was concentrated under vacuum. The extract was redissolved in 10 mL of 80% (v/v) methanol to obtain the free phenol fraction. Preparation of soluble ester phenolics The aqueous phase from the separation in 2.4.2.2 was hydrolyzed by adding 40 mL of NaOH (2 mol/L) for 4 h (37 °C, 150 rpm) and then acidified with HCl (6 mol/L) to pH 2.0. The mixture was degreased, extracted, dried, and reconstituted according to section “Preparation of free phenolics”. The result of the re-solubilization was the soluble ester phenol fraction. Preparation of soluble glycoside phenolics The soluble glycosidic phenol fraction was extracted from the water phase following the ethyl acetate extraction of the soluble ester phenolic fraction obtained in section 2.4.2.3. The water phase was hydrolyzed using 10 mL of 6 mol/L HCl in a water bath at 85 °C for 30 min and then adjusted to pH 2.0 with 10 mol/L NaOH. The mixture was then defatted, extracted, dried, and reconstituted according to the protocol outlined in section “Preparation of free phenolics”. Preparation of insoluble ester phenolics The residue obtained in section 2.4.2.2 was initially subjected to digestion with 40 mL of 2 mol/L NaOH while being agitated (37 °C, 150 rpm) for 4 hours. The supernatant of the mixture, after centrifugation (11,960 × g, 10 min, 4 °C), was acidified to pH 2.0 with 6 mol/L HCl. The mixture was then defatted, extracted, dried, and redissolved following the procedures outlined in section “Preparation of free phenolics”. Preparation of insoluble glycoside phenolics The insoluble glycosidic phenol fraction was extracted from the residue obtained in section 2.4.2.5 (after centrifugation). The residue was hydrolyzed with 25 mL of 2 mol/L HCl in a water bath (at 85 °C for 60 minutes). The supernatant of the mixture, following centrifugation (at 11,960 × g for 10 minutes at 4 °C), was adjusted to pH 2.0 using 10 mol/L NaOH. Subsequently, the mixture underwent defatting, extraction, drying, and was redissolved as per section “Preparation of free phenolics”. Determination of phenolic content in different forms The method of Gong et al. (2017) was referred to with slight modification. The extracts (prepared in sections 2.4.2.2–2.4.2.6) were treated with a 10% (v/v) Folin-Ciocalteu reagent (2.5 mL) at 25 °C for 8 minutes. Subsequently, 7.5% (w/v) Na2CO3 (2 mL) was added to the mixture. The reaction was carried out in the dark for 60 minutes, and the absorbance of the mixture at 765 nm was measured. A standard curve was generated using gallic acid, and the results were expressed as Gallic acid equivalent (GAE) mg/g dry weight (DW). Determination of monomer phenolics content The determination was carried out using a 1260 Infinity II liquid chromatograph (Agilent, USA). The separation was performed on a Supersil ODS2-C18 column (4.6 mm × 250 mm, 5 μm). The column temperature was maintained at 35 °C. The mobile phase consisted of two solvents: (A) 1% formic acid and (B) methanol. Gradient elution was achieved by adjusting the volume ratio of solvent B in the mobile phase at a flow rate of 1 mL/min. The injection volume was 20 μL. All samples were analyzed using the diode array detector (DAD) at 280 nm. Components in phenol extracts were tentatively identified by comparing their retention time and peak area to those of standards under identical analysis conditions. Determination of volatile flavor substances A 7890A/5975C gas chromatography-mass spectrometer (Agilent, USA) was utilized for the analysis. A 10 mL sample was placed in a 15 mL vial, heated, and equilibrated to 80 °C for 30 minutes. Subsequently, the solid phase microextraction (SPME) probe was inserted into the vial and adsorbed in the headspace for 20 min with magnetic stirring at a speed of 100 × g. Separation was conducted on a capillary column DBWAX (30 mm × 0.25 mm, 0.25 μm). The inlet port temperature was set at 250 °C, and the SPME was inserted into the inlet well for desorption for 5 minutes. The temperature was programmed to increase: starting at 35 °C, held for 2 minutes, then ramped up to 230 °C at a rate of 5 °C/min, and held for 8 minutes. The carrier gas (He) flow rate was maintained at 1.0 mL/min without a shunt. Mass spectrometry conditions: ion source temperature set at 230 °C, quadrupole temperature at 150 °C, electron energy at 70 eV, and the mass scan range was 33–450 amu. Statistical analysis All data were expressed as mean ± standard deviation calculated from the results of 3 or more experiments. Data were analyzed and plotted using SPSS 26 (SPSS, Inc., Chicago, IL, USA) and ORIGIN 2021(Origin Lab Co., Northampton, MA, USA). Results were considered significant at P < 0.05 using one-way ANOVA. Results and discussion Basic properties of the strains Lactobacillus can utilize the monosaccharides present in coconut water to produce organic acids, and its growth and acid production capabilities significantly influence the fermentation process of coconut water. Consequently, Emir. J. Food Agric ⋅ Volume 37 ⋅ 2025 5 Emirates Journal of Food and Agriculture changes in pH and growth capacity can serve as reliable indicators to evaluate the suitability of Lactobacillus as a fermenting strain (Salmerón et al. 2015). The growth curves of Lactobacillus, along with the changes in pH and acidity, are illustrated in Fig. 1A–C. The lag phase for Lactobacillus incubated at 37 °C was observed to be 2 h. During the initial 18 h, no significant differences were noted in the growth rates and acid-producing capacities among L. paracasei M1.5, L. casei U2.1 and L. paracasei O3.3. However, the growth rate and acid-producing capacity of L. fermentum S1.5 were significantly lower than those of the other three Lactobacillus strains. L. casei U2.1 and L. fermentum S1.5 entered the stabilization phase at 18 hours, while L. paracasei M1.5 and L. paracasei O3.3 exhibited prolonged growth periods, entering the stabilization phase at 22 h. The pH of the culture broth began to stabilize after 14 h, with L. fermentum S1.5 displaying a significantly higher pH compared to the other three lactic acid bacteria strains. The results of the survival rate of lactic acid bacteria after incubation in simulated gastric and intestinal fluids are presented in Fig. 1D. The study by Megur et al. (2023) showed that excellent tolerance could be indicated when the number of surviving microorganisms was greater than 1 × 107 cfu/mL. Analysis of the survival rate data revealed that L. paracasei M1.5 demonstrated exceptional gastrointestinal tolerance. The viable counts of L. paracasei M1.5, L. casei U2.1, and L. fermentum S1.5 exceeded 1 × 107 cfu/ mL, indicating good tolerance. In summary, L. paracasei M1.5, L. casei U2.1, and L. fermentum S1.5 were classified as lactobacilli with gastrointestinal tolerance. In conclusion, L. paracasei M1.5 and L. casei U2.1 exhibited robust growth and acid production capabilities, significantly altering the acidity of the culture broth, making them suitable for use as fermenters in coconut water fermentation. Antioxidant capacity of strains The antioxidant function of food products is of increasing interest due to its potential positive impact on human health. The antioxidant function is mainly due to the bioactive substances they contain (Abountiolas et al. 2018). In this study, three indices such as total reducing power, DPPH and ABTS radical scavenging capacity were selected to evaluate the antioxidant capacity of lactic acid bacteria. The results, as shown in Fig. 2, showed that the DPPH radical scavenging rates of the four Lactobacillus strains were L. casei U2.1 (30.91% ± 0.012) > L. paracasei O3.3 (12.40% ± 0.009) > L. fermentum S1.5 (7.45% ± 0.006) > L. paracasei M1.5 (6.33%±0.008). The ABTS free radical scavenging activity of the four Lactobacillus strains was L. paracasei O3.3 (41.82% ± 0.012) > L. casei U2.1 (41.72% ± 0.017) > L. paracasei M1.5 (33.83% ± 0.007) > L. fermentum S1.5 (22.24% ± 0.008).The total reducing power of L. fermentum S1.5 and L. casei U2.1 was better. Figure 1. Basic properties of Lactobacillus. A. Growth curve; B. PH change curve; C. Acidity change curve; D. Gastrointestinal tolerance survival. AuthorFamily, et al.: Lactobacillus in coconut water fermentation 6 Emirates Journal of Food and Agriculture Xu et al. (2011) showed that the ability of Lactobacilli to scavenge DPPH radicals was positively correlated with the concentration of extracellular polysaccharides they produced and the activity of the extracellular polysaccharides. The scavenging of DPPH radicals is a process whereby the radical binds to a hydrogen atom, and DPPH forms stable compounds after acquiring hydrogen atoms (Berset et al. 1995; Berset et al. 1997; Tsimogiannis et al. 2006). L. casei U2.1 may produce more extracellular polysaccharides capable of providing hydrogen atoms to DPPH or other substances that achieve antioxidant activity by providing hydrogen atoms during growth. Unlike DPPH, scavenging of ABTS radicals is considered to be an electron-transfer reaction that achieves free radical scavenging mainly by obtaining electrons through a reaction with antioxidant substances (Vassalle et al. 2004). L. paracasei O3.3 had the strongest scavenging capacity of ABTS radicals. In scavenging both DPPH and ABTS radicals, L. casei U2.1 showed good scavenging ability. The scavenging capacity of two free radicals, ABTS and DPPH, was positively correlated with the strength of total reducing power, with some exceptions. Differences in the metabolism of lactic acid bacteria lead to differences in antioxidant capacity. L. fermentum S1.5 showed poorer scavenging capacity for two free radicals, DPPH and ABTS, but higher reducing power. L. casei U2.1 showed good positive correlation between the two and higher total reducing power. In conclusion, L. casei U2.1 was the superior strain for antioxidant function. In vitro assessment of the hypoglycemic capacity of the strains α-Glucosidase and α-amylase are important enzymes in the process of glucose metabolism, which can hydrolyze glycosidic bonds to produce glucose, and have an important effect on human blood glucose concentration. Therefore, the inhibition rate of α-glucosidase and α-amylase was chosen as an index to evaluate the hypoglycemic function of the strain in this study. The inhibition rates of α-glucosidase and α-amylase activity by lactic acid bacteria. The inhibition rate of α-glucosidase by five strains ranged from 3.52% to 40.94%, with the positive control strain L. rhamnosus LGG having an inhibition rate of 29.57. And the inhibition rates of L. casei U2.1 (37.5% ± 1.72) and L. paracasei O3.3 (40.94% ± 0.46) were significantly higher inhibition than L. rhamnosus LGG. The inhibition of α-amylase by lactic acid bacteria ranged from 10.77% to 57.44%. The inhibition rates of L. rhamnosus LGG was 21.53%, which was lower than that of L. paracasei M1.5 (57.44% ± 0.931) and L. casei U2.1 (48.21% ± 2.373). The hydrolysis of polysaccharides can be reduced by inhibiting the activities of α-glucosidase and α-amylase, which indirectly regulates the blood glucose level (Khan et al. 2023). Therefore, the inhibitory capacity of α-glucosidase and α-amylase can represent the hypoglycemic activity. L. casei U2.1 showed good inhibition of both indicator enzymes, suggesting the potential hypoglycemic capacity of L. casei U2.1. The inhibitory effects of Lactobacillus strains on α-glucosidase and α-amylase could be attributed to several mechanisms. Firstly, lactic acid bacteria are known to produce various metabolites during fermentation, including organic acids (e.g., lactic acid and acetic acid), exopolysaccharides, and bioactive peptides, which may interact with these enzymes and alter their catalytic activities (Wang et al. 2014; Zhang et al. 2021). Organic acids can lower the local pH or directly bind to the active sites of the enzymes, thereby reducing their efficiency. Secondly, certain components of the bacterial cell wall or extracellular polymeric substances might adsorb or complex with the enzymes, leading to steric hindrance or conformational changes that impede substrate access. Furthermore, the production of specific inhibitory compounds through microbial metabolism, or the ability of probiotics to modulate the gut microbiota, may indirectly influence host. Effect of Lactobacillus fermentation on the content and composition of polyphenolic substances in coconut water L. paracasei M1.5 and L. casei U2.1 were selected as the coconut water fermentation agent based on the results of the above basic performance experiments. After 24 h of fermentation with M1.5-U2.1, the total phenol content of coconut Figure 2. Antioxidant capacity of Lactobacillus.Figure 3. Hypoglycemic capacity of Lactobacillus. Emir. J. Food Agric ⋅ Volume 37 ⋅ 2025 7 Emirates Journal of Food and Agriculture water was increased by 19%, and the fermentation products contained significantly more soluble glycoside phenols, insoluble glycoside phenols, insoluble lipophilic phenols, and free phenols, and a significant decrease in fat-soluble phenols (Table 1, Fig. 4). In previous studies, Ghosh et al. (2015) and Freire et al. (2017) found that fermentation of cereal slurry by Lactobacillus led to a significant increase in phenolic content. The increase in free phenols may be attributed to several factors: (1) the production of reducing substances through Lactobacillus metabolism that protect free phenols from oxidation; (2) the generation of new free phenolics via glycosylation and methylation during metabolic processes; and (3) the enzymatic catabolism of macromolecules by Lactobacillus during fermentation (Jamal et al. 2011; Hur et al. 2014). Phenolics have a variety of functional properties and the increase in phenolic content in fermented coconut juice may indicate elevated levels of phenols with antioxidant and hypoglycemic functions. Coconut water contains phenolics such as epicatechin, p-hydroxybenzoic acid, butyric acid, o-coumaric acid, p-coumaric acid and caffeic acid (Mahayothee et al. 2016). Among them, epicatechin, p-hydroxybenzoic acid, butyric acid, and p-coumaric acid have been shown to be phenolics with hypoglycemic activity and antioxidant activity (Hii et al. 1985; Best et al. 2007; Youl et al. 2010; Kilic et al. 2013), and the fermentation of Lactobacillus may cause changes in their contents. Therefore, in this study, we chose to analyze the content of the above-mentioned monomer phenols in coconut water. The experimental results are shown in Table 2. Fermentation increased p-hydroxybenzoic acid by 75%, with no significant change in syringic acid content, while caused decrease in epicatechin, p-coumaric acid and caffeic acid content by 14%, 16.5% and 34%, respectively. The depletion of antioxidant phenols such as epicatechin, caffeic acid and p-coumaric acid may be related to the protective effect of antioxidants. Clark et al. (2008) reported that antioxidants in fermented wine produced a depletion of the protective effect on other oxidation-prone substances. Additionally, the content of p-coumaric acid was influenced by the hydrolysis of p-coumaroylated anthocyanins. Flavanol-type phenolic compounds such as epicatechin underwent polymerization and subsequent oxidation (Clark et al. 2010), all of which led to a decrease in monomeric phenol content. The increase in p-hydroxybenzoic acid content may be related to the action of aldehyde dehydrogenase (Payet et al. 2016), which converted p-hydroxybenzaldehyde analogs to p-hydroxybenzoic acid. In addition, p-hydroxybenzoic acid can be biosynthesized via the mangiferolic acid pathway. In this pathway, phosphoenolpyruvic acid produced by glycolysis was converted to chorismic acid and further produces p-hydroxybenzoic acid (Mir et al. 2015; Li et al. 2020). Figure 4. Effect of fermentation on phenolics in coconut water. Table 1. Effect of fermentation on phenolic contents. Sample Polyphenol content (μg/mL) Free phenol Insoluble ester phenol Soluble ester phenol Soluble glycosidic phenol Insoluble glycosidic phenol Total phenol Coconut water 17.62 ± 0.26Bc 9.56 ± 0.45Bd 21.13 ± 0.09Aa 19.60 ± 0.43Bb ND 67.91 ± 0.25B Fermentation broth 20.44 ± 0.51Aa 10.21 ± 0.40Ad 18.56 ± 0.31Bb 20.19 ± 0.23Aa 11.49 ± 0.37Ac 80.89 ± 0.46A Table 2. Effect of fermentation on the content of monomeric phenols. Sample Monomer phenol content (μg/mL) Epicatechin Syringic acid p-Hydroxybenzoic acid P-coumaric acid Caffeic acid Coconut water 4.15 ± 0.06A0.30 ± 0.16A0.12 ± 0.03B3.45 ± 0.07A0.50 ± 0.12A Fermentation broth 3.57 ± 0.17B0.30 ± 0.33A0.21 ± 0.02A2.88 ± 0.11B0.33 ± 0.03B AuthorFamily, et al.: Lactobacillus in coconut water fermentation 8 Emirates Journal of Food and Agriculture In conclusion, the fermentation of coconut water by lactic acid bacteria increased total phenolic content, with a notable rise in p-hydroxybenzoic acid among the five monophenols analyzed, a reduction in epicatechin, caffeic acid, and p-coumaric acid, and no significant alteration in syringic acid. Effect of Lactobacillus fermentation on volatile flavor substances of coconut water In the present work, the influence of lactic acid bacteria fermentation on the volatile flavor substances of coconut water was further studied. The volatile organic compound (VOC) composition of each sample is shown in Fig. 4A and 4B. After fermentation, there was a significant increase in the number of volatile organic compound (VOC) species, which increased by 27 species. And the number of alcohols, esters and ketones was also enhanced. In addition, there was a significant increase in the number of organic acid species in the fermented samples compared to the unfermented coconut water. Long-chain hydrocarbons and cycloalkanes underwent cracking, oxidation and other reactions to produce other flavor substances due to the fermentation of lactic acid bacteria, and the content of alkanes decreased (Maghrebi et al. 2021). These changes had an important effect on the flavor of coconut water after fermentation. During fermentation, changes in acids directly reflected the interaction between the metabolic activity of Lactobacillus and the raw material. Their dynamic balance determined the flavor, functionality, and stability of the product. Medium and long-chain fatty acids in coconut water underwent decomposition due to β-oxidation. The greatest increase in palmitoleic acid content among the organic acids was not detected in coconut water while it was produced in large quantities during the post-ripening stage (Fig. 4C, Table 3). Palmitoleic acid was capable of imparting specific flavors (e.g. Figure 5. Changes of volatile flavor substances. A. Changes in the number of volatile flavorant species in coconut water; B. Percentage change of volatile flavorant species in coconut water; C. Organic heat map. Emir. J. Food Agric ⋅ Volume 37 ⋅ 2025 9 Emirates Journal of Food and Agriculture Table 3. Effect of fermentation on the content of volatile components. Types Organic name Coconut water Sterilization liquid Fermentation liquid Post-ripening liquid acids Palmitic acid 15.02% 7.35% 6.04% 6.47% Acetic acid ND ND 1.71% 2.27% Nonanoic acid ND ND 2.49% 2.56% Suparic acid 3.93% 2.83% 2.38% 2.43% Hexanoic acid ND ND 2.30% 2.93% Lauric acid 6.47% 3.26% 3.70% 2.28% N-decanoic acid ND 2.69% 2.71% 2.03% Palmitoleic acid ND ND ND 9.99% Myristic acid 8.17% 3.90% 3.93% 3.54% Pentadecanoic acid 10.82% 5.40% ND 5.13% Cis-5-docosaenoic acid ND ND 2.92% 2.43% Alcohols Phenyl alcohol ND ND 2.11% 3.58% Ethanol ND 2.94% ND ND 2-ethylhexanol ND 1.72% ND 1.48% Neroli alcohol ND ND 1.85% ND Heptanol 1.70% 1.09% 1.04% ND 1-octanol 2.86% 2.07% 1.61% 1.99% Lauryl alcohol ND ND 2.14% ND Isoamyl alcohol ND 1.65% 1.12% ND 2,4,7,9-tetramethyl-5-decyne-4,7-diol 3.41% ND ND ND 1-nonanol ND ND 1.65% 1.72% 2-nonanol ND ND 1.54% 1.77% Cyclodecane-1-alcohol ND ND ND 2.29% 2-undecanol ND ND ND 1.70% 2-dodecenoaldol ND 2.53% ND ND 2-dodecanol ND ND 1.69% ND 3-cyclohexene-1-ethanol ND ND 2.06% 2.15% Esters Diisobutyl phthalate 4.98% 2.69% 2.75% 2.22% Dibutyl phthalate 6.56% 3.27% 2.96% 2.95% Ethyl octanoate 1.72% 1.04% 0.91% 1.05% Ethyl fetalate ND 2.50% ND ND Ethyl oleate ND ND 2.11% 1.87% Ethyl hexanoate ND 1.18% ND ND Ethyl myristate ester ND ND 1.91% 1.77% Dimethyl phthalate ND 2.54% ND ND Ethyl undecanoate ND ND 1.65% 2.45% Ethyl palmitate ND ND 1.89% 1.80% Butyric decanolide 4.16% ND 2.33% ND 2-chlornonyl acetate ND 2.27% ND ND Butyl 2-methylpropyl 1, 2-phthalate ND ND 2.52% ND Ethyl 9-bromononanoate ND 2.66% ND ND methyl-3-hydroxy-2,4,4-Trimethylpentyl propionate ND 2.84% 2.35% ND (1, 1-dimethylethyl)-2-methyl-1, 3-malondiyl ester ND ND 2.13% ND (3Z) -3-decene-1-ol acetate ND ND 1.82% 1.87% Disecobutyl phthalate ND 2.77% ND ND aldehyde Decyl aldehyde 2.95% ND 1.71% ND Nonenal aldehyde ND 1.76% ND ND 3, 5-dimethylbenz aldehyde ND ND 2.54% ND 3, 4-dimethylbenz aldehyde ND ND ND 2.39% Trans-2-dodecen aldehyde ND 2.72% ND ND alkanes Eicosane ND 3.19% ND 1.79% Cyclodecane ND ND ND 3.25% Cyclododecane 3.26% 3.44% ND 2.18% Cyclotetradecane 4.32% ND 2.17% ND n-hexadecane ND 2.41% ND ND pentadecane ND 1.41% ND ND n-heptacosane 3.01% ND ND ND 1-fifteen alkyne ND ND 1.91% ND 1, 11-docosadiene ND ND ND 1.70% (Z) -3-hexadecene ND 2.78% ND ND Nonylcyclopropane ND ND 2.15% ND