Green extraction approach for green microalga Tetraselmis tetrahele: Enhanced phenolic yield and bioactivity via Ultrasound-Assisted extraction

TL;DR

This study shows that using ultrasound-assisted extraction improves the yield and bioactivity of phenolic compounds from a green microalga, making it a promising method for producing high-value natural products.

Contribution

The study introduces ultrasound-assisted extraction as a green and scalable method for maximizing phenolic compound yield from Tetraselmis tetrahele.

Findings

Ultrasound-assisted extraction significantly increased phenolic and flavonoid content compared to maceration.

The UAE method enhanced anti-collagenase activity and photosynthesis pigment levels in microalgal extracts.

Extraction parameters like solvent composition and time were optimized for maximum bioactive yield.

Abstract

Microalgae are gaining attention as promising natural sources of bioactives due to their rich nutrient content. This study investigates the effects of ultrasound-assisted extraction (UAE) parameters (i.e., solvent composition, temperature, solid-to-solvent ratio, and time) on the extraction of phenolic compounds from Tetraselmis tetrahele. The UAE parameters were determined at a solid-to-solvent ratio of 1:75, a temperature of 25 °C, and an extraction time of 15 min with 75% ethanol as the solvent. Phenolic profile, total phenolic content, total flavonoid content, radical scavenging activity of DPPH and ABTS, photosynthesis pigment (chlorophyll a, b, and total carotenoid), anti-collagenase activity, and metabolic profile of Tetraselmis tetrahele extract obtained using UAE were determined. Tetraselmis tetrahele extracts obtained using the selected UAE method demonstrated significantly higher (p < 0.05) levels in gallic acid (1.81%), 4-hydroxybenzoic acid (7.0%), total phenolic content (84.0%), total flavonoid content (56.2%), ABTS (101.3%), chlorophyll a (119.4%), chlorophyll b (174.3%), total carotenoids (173.2%) and anti-collagenase activity (15.9%) compared to the maceration method. These findings provide UAE as a green and scalable method for maximizing the phenolic yield and support the application of Tetraselmis tetrahele in the development of high-value products across pharmaceutical, cosmetic, and functional food industries.

Figures

Absorption spectra in the UV–Vis region (200–700 nm) of the *Tetraselmis tetrahele* extracts with (A) UAE and (B) maceration. The peak at 337 nm suggests UV-absorbing compound, bands at 440 nm and 664 nm correspond to chlorophyll *a*, and the peak at 466 nm is associated with carotenoids. brief title: Absorption spectra of *Tetraselmis tetrahele* extracts.

Full text

Introduction

1

Microalgae are attracting global interest as a sustainable source of bioactive compounds for use in food and health applications due to their rich nutritional value and variety of bioactive compounds. Global microalgae production has achieved 1.02 million tonnes for use in bioenergy, feed, and pharmaceutical application [1]. The rich bioactive compounds in microalgae have demonstrated antioxidant, anti-inflammatory, antidiabetic, antiviral, and anticancer activities [2], [3]. Tetraselmis tetrahele, a member of the phylum Chlorophyta, is commonly used in aquaculture industry as an animal feed for aquatic organisms due to the presence of polyunsaturated fatty acids, polysaccharides, protein, enzymes, and carotenoids. In addition, Tetraselmis species was proved to reduce stress in fish and shellfish [4] due to their high antioxidant, anti-inflammatory, and antimicrobial properties [5]. Thus, Tetraselmis tetrahele has a great capacity to be used as a novel source of antioxidant compounds due to its ease of cultivation with fast growth rate, tolerance to a wide range of pH (3–10.5) and salinity (10–60 ppt), as well as high nutritional value, including protein, carbohydrates, fatty acids, and other bioactive compounds [6].

Phenolic compounds, the most abundant class of plant-derived secondary metabolites, play a pivotal role as potent exogenous antioxidants [7]. Amid growing safety concerns surrounding synthetic antioxidants, naturally derived alternatives such as phenolics, carotenoids, and astaxanthin have gained increasing scientific and industrial attention [8]. In particular, phenolic compounds from microalgae are emerging as promising candidates for applications in nutraceuticals, cosmeceuticals, and pharmaceuticals as a natural antioxidant. However, the exploration of phenolic compounds in microalgae remains limited, and the commercial scale production of these bioactives is hindered by high costs and inefficiencies in downstream processing, particularly extraction and purification steps [9]. Thus, the development of standardized, efficient, and scalable extraction protocols for microalgae species such as Tetraselmis tetrahele is essential to unlock their full biotechnological value.

Ultrasound-assisted extraction (UAE) offers an energy-efficient, rapid, cost-effective, safe, and environmentally friendly, compared with the traditional extraction methods [10], [11]. UAE can overcome the issues of high solvent consumption, maintenance costs, long extraction time, difficulties in usage, and energy consumption, making it a scalable and practical solution for industrial applications [12]. This technology can disrupt the cell wall of microalgae to facilitate the release of phenolic compounds to enhance the extraction yield. Moreover, it is appropriate for mass production of temperature-sensitive compounds such as phenolic compounds, which it is more advantageous than other modern extraction methods with heat requirements, such as microwave-assisted extraction and pressurized liquid extraction [10]. Although heat may also be generated during the ultrasound process, the overall extraction temperature of the UAE is generally mild and can be effectively controlled, especially for the ultrasonic bath equipped with temperature control. In addition, the relatively simple and low-cost of the equipment and operation of the UAE making it easier to scale-up for mass production, compared to other extraction methods, such as supercritical fluid extraction [10]. Fernandes et al. [13] investigated UAE as a green extraction approach for obtaining carotenoids and chlorophyll from microalgae. However, to date, there is limited information on the extraction of phenolic compounds from microalgae using UAE.

In this work, one of the challenges of using UAE is determining the extraction parameters for different bioactive compounds from various ingredient sources [12], as the thickness, chemical composition, cell wall structure, and phenolic compounds can vary significantly between microalgae species [14]. Despite the growing interest in microalgae, the phenolics profile of Tetraselmis sp. remains underexplored. Thus, this study aimed to investigate the effects of different UAE parameters on the extraction of phenolic compounds from Tetraselmis tetrahele by quantifying the individual phenolic compounds using ultra-high performance liquid chromatography (UHPLC). A single-factor experimental approach was adopted to investigate the influence of key UAE parameters, including solvent composition, temperature, solid-to-solvent ratio, and time. The efficiency of the selected UAE conditions was further validated by comparing the total phenolic content, total flavonoid content, antioxidant activities (DPPH and ABTS assays), photosynthetic pigment content (chlorophyll a, chlorophyll b, and total carotenoids), and anti-collagenase activity against extracts obtained via conventional maceration. These findings contribute to the development of an effective and environmentally friendly method for recovering phenolic compounds from microalgal biomass with promising antioxidant and bioactivity.

Materials and methods

2

Chemicals

2.1

Phenolic standards were purchased from Sigma-Aldrich Co. (St. Louis, MO, USA). 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid), 1,1-dipheny1-2-picrylhydrazyl, 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid, Folin-Ciocalteu reagent, and other chemicals were sourced from Sigma-Aldrich. HPLC-grade methanol and acetic acid were sourced from Fisher brand (New Hampshire, US).

Cultivation of microalgae

2.2

Tetraselmis tetrahele was cultivated in Conway medium [15], as detailed in Table 1, with micronutrients, trace metals, and silicate solution in 30 ppt seawater under the microalgae incubator (GZP-360 N, Senxin). Cultivation was carried out in a microalgae incubator (GZP-360 N, Senxin). A 7-day pre-inoculum was grown in test tubes, then scaled up to 200 mL in 300 mL flasks under LED light at an intensity of 60 µmol photons m^-2^s^-1^ with a 12:12 h light/dark cycle at room temperature. Constant aeration was provided using an air pump (ACO-328, Hailea, China). Cultures were further upscaled to 2 L in 3 L cylindrical flasks and the cells at stationary phase were harvested by centrifugation (4000 rpm, 5 min, 15 °C), washed twice, and freeze-dried (Cool Safe 110–4, LaboGene).Table 1. Composition of Conway medium solution.Name of ChemicalQuantitySolution A: Main mineral solutionNaNO_3_100.00 gEDTA-2Na(C_10_H_16_N_2_Na_2_O_8_)45.00 gH_3_BO_3_33.60 gNaH_2_PO_4_·2H_2_O20.00 gFeCL_3_·6H_2_O1.30 gMnCL_2_·4H_2_O0.36 gTrace metal solution (B)1.00 mLDissolved in deionised water and make up the volume to 1 LSolution B: Trace metal solutionZnCL_2_2.10 gCoCL_3_·6H_2_O2.00 g(NH_4_^+^)6_MO_7_O_2·4H_2_O0.90 gCuSO_4_·5H_2_O2.00 gDissolved in deionised water and make up the volume to 1 LSolution C: Vitamin solutionThiamine, B10.20 gCyanocobalamin, B120.01 gDissolved in deionised water and make up the volume to 100 mLSolution D: Silicate solutionNa_2_SiO_3_15.00 gDissolved in deionised water and make up the volume to 1 L

Microalgae extraction using UAE with different parameters

2.3

The freeze-dried microalgal biomass was used to extract phenolic compounds using UAE under various extraction parameters, including solvent composition, temperature, solid-to-solvent ratio, and time, based on single-factor experiments. After investigating one parameter, the selected parameter was applied to subsequent tests to determine the effect of the remaining parameters on phenolic compound yield.

Solvent composition

2.3.1

To determine the ideal solvent composition for phenolic compound extraction, 5 mL of different solvents (100% methanol, 25% ethanol, 50% ethanol, 75% ethanol, and ultrapure water) were added into corresponding centrifuge tubes (20 mg dried biomass). Then, all samples were extracted via UAE in an ultrasonication bath (KQ-300DE, Kunshan) with a frequency of 40 kHz (ultrasonic power: 300 W) at 25 °C for 15 min. Subsequently, the samples were centrifuged at 15 °C at 4500 rpm for 5 min. The above procedures were repeated three times, and the supernatants in each centrifugation of extraction were combined and collected. The selected solvent composition (75% ethanol) was determined based on the phenolic content quantified by UHPLC and was used in the following sections.

Temperature

2.3.2

To determine the ideal UAE temperature in extracting the phenolic compounds, 5 mL of the selected solvent composition (75% ethanol) was added into each corresponding centrifuge tube containing 20 mg of dried microalgal sample. Then, all samples were transferred to the ultrasonication bath at various temperatures (4 °C, 25 °C, and 45 °C) for 15 min. Subsequently, the extraction and collection steps were conducted as previously mentioned. The selected UAE temperature (25 °C) was determined based on the phenolic content quantified by UHPLC and used in the following sections.

Solid-to-solvent ratio

2.3.3

Twenty milligrams of dried microalgal biomass were mixed with different volumes (0.5 mL, 1 mL, 5 mL, and 10 mL) of selected solvent composition (75% ethanol), corresponding to solid-to-solvent ratios of 1:75 g/mL, 1:150 g/mL, 1:750 g/mL, and 1:1500 g/mL, respectively, after combining the supernatants from three repetitions of the process. The extraction and collection procedures were then conducted as previously mentioned. The selected solid-to-solvent ratio (1:75 g/mL) was determined based on the phenolic content quantified by UHPLC and used in the following sections.

Time

2.3.4

To determine the ideal ultrasonication time for extracting the phenolic compounds, 75% ethanol was added to the dried microalgal biomass (20 mg) based on the selected solid-to-solvent ratio (1:75 g/mL). Then, all samples were sonicated in an ultrasonication bath at 25 °C for different durations (5 min, 15 min, and 30 min). The extraction and collection procedures were then conducted as previously mentioned. The selected UAE time (15 min) was determined based on the phenolic content quantified by UHPLC.

Microalga extraction using maceration

2.4

The maceration method was carried out according to Dianursanti et al. [16] with slight modifications. Dried microalgal biomass (40 mg) was added with 75% ethanol at a solid-to-solvent ratio (1:75 g/mL) and stored in the dark for 48 h at room temperature. The sample was then centrifuged at 15 °C at 4500 rpm for 5 min. The supernatants were collected and subjected to the following analyses.

Determination of the phenolic compound by UHPLC-PDA

2.5

The collected supernatants were evaporated to dryness using a Hei-VAP Advantage vacuum evaporator (Heidolph, Germany), then redissolved in 2 mL of ultrapure water and filtered through a PVDF syringe filter (0.22 μm) prior to UHPLC analysis. The phenolic profile of microalgal extracts from UAE and maceration was quantified according to Lau et al. [17] using Waters Acquity Arc UHPLC (Waters Corporation, US) coupled with an Agilent InfinityLab Poroshell 120 EC-C18 column (4.6 x 250 mm, 4 μm) and a photodiode array (PDA) detector. The chromatographic separation employed a gradient elution by mobile phase A (methanol) and mobile phase B (1% acetic acid solution) from 0-4 min, 4% A; 4–24 min, 22% A; 24–30 min, 60% A; 30–34 min, 90% A; 34–40 min, 20% A, and 40–45 min, 4% A at a flow rate of 1 mL/min. The peaks were detected at 280, 290, 310, 330, and 360 nm and compared with the phenolic standards of 4-hydroxybenzoic acid, protocatechuic acid, p-coumaric acid, vanillic acid, gallic acid, caffeic acid, ferulic acid, catechin, kaempferol, quercetin, and chlorogenic acid. The concentration of the respective phenolics was expressed in μg/g dry weight of the microalgal biomass based on the reference standard curve.

Comparison between UAE and maceration

2.6

DPPH and ABTS radical scavenging activity assays

2.6.1

To compare the antioxidant activity of microalgal extracts from UAE and maceration, the DPPH and ABTS radical scavenging activity assays were conducted according to Chew and Nyam [18] with slight modifications. For the DPPH assay, 100 µL of sample was mixed with 2 mL of 0.15 mM DPPH solution and incubated in the dark for 30 min. For the ABTS assay, 50 µL of sample was mixed with 950 µL of diluted ABTS solution (absorbance 0.700 ± 0.050) and left undisturbed for 3 min. The radical scavenging activities were calculated based on the Trolox standard curve (0.02 to 0.20 mg/mL) with the calibration equation of y = 136.03 x – 0.2836 (R^2^ = 0.993) at 517 nm (for DPPH) and the calibration equation of y = 285.97 x  + 9.0324 (R^2^ = 0.998) at 734 nm (for ABTS). The results were expressed as mg Trolox equivalents (mg Teq/g biomass).

Total phenolic content (TPC)

2.6.2

TPC was measured by the Folin–Ciocalteu method according to Lau et al. [17]. A 100 µL sample was mixed with ultrapure water (600 µL) and Folin-Ciocalteu reagent (500 µL). Sodium carbonate solution (20%, 1.5 mL) was then added, followed by incubation for 1 h. TPC was calculated based on the standard curve of gallic acid (0.02 to 0.20 mg/mL) with a calibration equation of y = 3.0727 x – 0.0326 (R^2^ = 0.992) at 760 nm, and results were expressed as mg gallic acid equivalents (mg GAE/g biomass).

Total flavonoid content (TFC)

2.6.3

TFC was measured by the aluminium chloride colorimetric assay according to Lau et al. [17]. Sample (250 µL) was mixed with 10% AlCl_3_ solution (50 µL) and methanol (750 µL). Potassium acetate (50 µL, 1 M) and ultrapure water (1400 µL) were then added and the mixture was incubated for 30 min. The TFC was calculated from a quercetin standard (0.02 to 0.20 mg/mL) using the calibration equation of y = 3.2583 x – 0.0003 (R^2^ = 0.998) at 415 nm, and expressed as mg quercetin equivalents (mg QE/g biomass).

Total carotenoids and chlorophyll content

2.6.4

Total carotenoids and chlorophyll contents of microalgal extract (1 mg/mL) were determined according to Lichtenthaler and Buschmann [19] at 470, 648, and 664 nm using a UV–Vis spectrophotometer (V-770ST, Jasco). The concentration was calculated by the Lichtenthaler equations as follows, and expressed in the unit of mg/g biomass.

$$[eqn]$$ $$[eqn]$$ $$[eqn]$$

Uv–vis absorption spectra

2.6.5

The UV–Vis absorption spectra of the extracts were scanned using a UV–Vis spectrophotometer (V-770ST, Jasco) with a spectral window from 200 to 700 nm and resolution of 0.5 nm.

Collagenase inhibition assay

2.6.6

The collagenase inhibitory activity (IC_50_) was measured using the fluorometric Collagenase Inhibitor Screening Kit (Abcam, ab211108) according to the manufacturer’s instructions with the microalgal extract in the concentration of 0.67 to 10 µg/mL. In the assay, 1 µL of sample and diluted inhibitor (1,10-phenanthroline) were added in each well. Then, 5 µL collagenase (1:50 dilution) was added to the well, and the volume was adjusted to 50 µL with assay buffer. After 15 min of incubation, diluted collagenase substrate (gelatin) was added to the mixture, followed by measurement of the fluorescence signal in excitation/emission wavelengths of 490/520 nm using a fluorescence Spark^TM^ 10M multimode plate reader (Tecan Trading AG, Switzerland).

Liquid chromatography-quadrupole time of flight-mass spectrometry (LC-QTOF-MS)

2.7

Metabolic profiling of microalgal extract using selected UAE parameters was performed using an Agilent 1290 Infinity LC system coupled with an Agilent 6520 Accurate-Mass Quadrupole Time-of-Flight (QTOF) mass spectrometer (Agilent, Santa Clara, CA, USA). A qualitative untargeted profiling analyses of the metabolites was performed in positive and negative ionization modes. The chromatographic separation was performed using an Agilent Eclipse XDB-C18 column (150  mm × 2.1  mm, Agilent Technologies, USA). The mobile phase consisted of (A) 100% water and (B) 100% acetonitrile and a gradient elution was employed with 0–5 min, 95% A; 6–20 min, 0% A; and 21–25 min, 0% A; with a flow rate of 0.5 mLmin^−1^. The full-scan MS was recorded for 100–3200 m/z. Data were acquired using Agilent MassHunter qualitative analysis B.07.00 software. Metabolic profiling was performed using accurate mass and MS/MS fragmentation data with a mass tolerance of ± 5 ppm. Compounds annotation was performed by comparison with the METLIN database and only metabolites with high-confidence matches (score ≥ 80%) were considered for further interpretation.

Statistical analysis

2.8

All experiments were conducted in duplicate and each sample was measured two times. The statistical analysis was conducted with the statistical software MINITAB 21 (Minitab Inc, Pennsylvania, USA). The significant differences among the samples were determined based on 95% confidence level (p < 0.05). For comparing the parameters in UAE, one-way analysis of variance and Tukey's post-hoc test were conducted. For comparing UAE and maceration, 2-samples t-test was conducted.

Results and discussion

3

Effects of solvent composition on phenolic content

3.1

Table 2 presents the identified phenolic compounds extracted from Tetraselmis tetrahele using UAE with different solvent compositions. Seven phenolic compounds were identified in Tetraselmis tetrahele, including gallic acid, catechin, ferulic acid, quercetin, 4-hydroxybenzoic acid, p-coumaric acid, and protocatechuic acid. The total phenolic compounds ranged from 663.15 to 712.71 μg/g biomass, indicating that the solvent composition could significantly affect the extraction of the phenolic compound from microalga via UAE. The results showed that 75% ethanol was the most efficient extraction solvent, yielding significantly higher (p < 0.05) contents of ferulic acid (20.25 μg/g), 4-hydroxybenzoic acid (81.63 μg/g), and p-coumaric acid (54.26 μg/g), as well as the highest total phenolic compounds (712.71 ± 2.19 μg/g).Table 2. Identified phenolic compounds (μg/g biomass) at different UAE solvent composition.Content100% methanol25% ethanol50% ethanol75% ethanolwater4-Hydroxybenzoic acid64.68 ± 0.09^e^73.01 ± 1.10^c^76.20 ± 0.13^b^81.63 ± 2.48^a^67.74 ± 0.46^d^Protocatechuic acid12.80 ± 0.06^c^13.12 ± 1.00^b^12.80 ± 0.04^c^12.78 ± 0.06^c^13.85 ± 0.14^a^p-Coumaric acid54.26 ± 0.05^a^NDND54.26 ± 0.05^a^NDGallic acid171.57 ± 0.62^c^176.92 ± 1.17^b^177.29 ± 1.36^b^175.94 ± 0.60^b^183.19 ± 0.81^a^Ferulic acid19.89 ± 0.08^d^20.00 ± 0.09^cd^20.25 ± 0.12^a^20.21 ± 0.04^ab^20.07 ± 0.04^bc^Catechin272.20 ± 4.43^b^276.59 ± 1.78^b^275.97 ± 3.76^b^264.60 ± 1.60^c^300.46 ± 3.08^a^Quercetin104.12 ± 0.27^a^103.51 ± 0.08^b^103.34 ± 0.08^bc^103.20 ± 0.03^c^103.19 ± 0.03^c^Total identified phenolic compounds693.12 ± 11.02^b^663.15 ± 2.61^c^665.85 ± 3.33^c^712.71 ± 2.19^a^688.49 ± 4.24^b^ND: Not detected.^abcd^ Mean ± standard deviation with distinct superscript letters within the same row are significantly different (p < 0.05) based on Tukey’s test.

In the current findings, 75% ethanol yielded significantly higher (p < 0.05) total identified phenolic content than 100% methanol or water. This is likely due to the differences in polarity, viscosity, and surface tension. Polarity, in particular, plays a key role in extraction efficiency, as the formation and disruption of hydrogen bonds are essential during solubilization [20]. This is because the hydroxyl groups in these solvents can interact with oxygen atoms within phenolic structures, making polar protic solvents like ethanol and methanol more efficient for phenolic extraction. However, highly polar protic solvents like water can lower extraction efficiency because their strong solvation capacity limits transfer of mid-polarity phenolic compounds into the extraction phase [20], [21].

In terms of the solvent viscosity and surface tension, extraction solvent with a lower viscosity and surface tension enables more phenolic compounds dissolving into the extraction medium [22], while minimizing the cavitation resistance during UAE [12]. The current findings showed that water is effective in extracting highly polar phenolics, such as gallic acid and protocatechuic acid, but is less suitable for mid-polarity phenolics, such as ferulic acid and p-coumaric acid. This may be due to water’s high surface tension, which resulted from strong cohesive forces between its hydroxyl groups, creating greater inward pull [23].

At 25 °C, the surface tension of methanol (22.07 mN/m) and ethanol (22.32 mN/m) is substantially lower than that of water (72.75 mN/m) [24], facilitating solvent penetration and mass transfer during extraction. Although ethanol has slightly lower dynamic and kinematic viscosities than water, its superior surface property dominates extraction efficiency for nonpolar to mid-polar phenolic compounds. In this study, an interesting trend was observed in which the total identified phenolic compounds increased as the ethanol concentration increased from 25% to 75%. The higher ethanol composition in ethanol–water mixtures (e.g., at 25%, 50%, and 75% ethanol) helps in lowering surface tension, which in turn disrupts the hydrogen bond networks and enhances solvent penetration into cell tissues [25]. Consequently, a solvent with a higher ethanol composition provides higher extraction efficiency for phenolic compounds, which is in agreement with Bandyopadhyay et al. [23], Hossain et al. [22], and Do et al. [26]. Additionally, Monteiro et al. [27] reported that 80% ethanol was the optimal solvent for extracting TPC from Fucus vesiculosus extract, which further supports the current findings.

From a safety perspective, methanol is prohibited in products that come into contact with the skin or are intended for oral consumption. During production and processing, residual methanol may cause difficulties in the removal and purification of the final product. In contrast, water and ethanol are safer choices for industrial use. Compared to methanol, ethanol is safer, more environmentally friendly, and less toxic for human consumption [28]. Overall, 75% ethanol was identified as the appropriate solvent for extracting phenolic compounds by considering both safety and extraction efficiency.

Effects of temperature on phenolic content

3.2

Table 3 presents the individual phenolic compounds extracted at different ultrasonication temperatures (4 °C, 25 °C, and 45 °C). Gallic acid, catechin, ferulic acid, quercetin, 4-hydroxybenzoic acid, p-coumaric acid, protocatechuic acid, and kaempferol were identified. The total identified phenolics ranged from 784.67 to 799.71 μg/g biomass. At 4 °C, the extraction yielded a significantly lower amount (p < 0.05) of total identified phenolic compound (784.67 μg/g), which was 1.9% lower than 25 °C and 1.7% lower than 45 °C. This showed that increasing temperature would assist the extraction in different areas, including breaking the cell walls, enhancing the solubility and diffusivity of the phenolic compounds, and reducing the solvent viscosity [29].Table 3. Identified phenolic compounds (μg/g biomass) at different UAE temperatures.Content4 °C25 °C45 °C4-Hydroxybenzoic acid77.29 ± 0.76^b^84.20 ± 0.90^a^79.29 ± 1.45^b^Protocatechuic acid12.89 ± 0.08^b^13.14 ± 0.15^a^13.14 ± 0.11^a^p-Coumaric acid54.26 ± 0.13^b^54.56 ± 0.17^a^54.43 ± 0.11^ab^Ferulic acid20.21 ± 0.06^b^20.63 ± 0.09^a^20.80 ± 0.15^a^Gallic acid173.87 ± 0.29^b^175.78 ± 0.57^a^176.08 ± 0.62^a^Kaempferol73.58 ± 0.12^a^73.26 ± 0.07^b^72.93 ± 0.10^c^Catechin268.50 ± 3.10^b^273.44 ± 2.09^a^276.35 ± 1.90^a^Quercetin104.09 ± 0.36^b^104.71 ± 0.32^ab^105.07 ± 0.30^a^Total identified phenolic compounds784.67 ± 4.23^b^799.71 ± 3.64^a^798.08 ± 3.71^a^^abc^ Mean ± standard deviation with distinct superscript letters within the same row are significantly different (p < 0.05) based on Tukey’s test.

Under increasing temperature, the cavitation effect enhances extraction efficiency by disrupting cells. However, there was no significant difference (p > 0.05) in the total identified phenolic content between 25 °C (799.71 μg/g) and 45 °C (798.08 μg/g), revealed that comparable extraction efficiency of phenolic compounds at both temperatures. Temperatures above 45 °C were not selected in this study, as the evaporation process could reduce the solvent volume and affect the cavitation effect in UAE. In the extraction process, higher temperature in the ultrasonication would allow more solvent to evaporate, especially for the 75% ethanol used in this study. This would reduce the diffusion of the solutes to be extracted and the extraction efficiency [30]. Additionally, as temperature increases, the solvent vapor may fill the cavitation voids, leading to the disruption of cavitation bubbles and weakening the cavitation effect [12].

Previous studies have shown that higher temperatures increase the degradation rate of phenolics and diminish the diversity and the antioxidant capacity of these compounds [31]. Over 45 °C, phenolics are susceptible to hydrolysis, oxidation, and degradation, which significantly reduces the yield in UAE [32]. Since phenolic compounds are prone to oxidation and degradation, excessively high temperature may negatively impact extraction efficiency. This supports with the findings of the current study, in which increasing the temperature from 25 °C to 45 °C did not show a significant improvement. UAE at 25 °C was sufficient to effectively disrupt the cell walls and solubilize the phenolic compounds into the solvent. Thus, striking a balance between extraction efficiency and compound stability is essential to boost the extraction efficiency of the phenolic compounds while preserving the antioxidant activity. Overall, 25 °C was determined as the ideal extraction temperature, offering high phenolic yield while maintaining cost-effectiveness and operational simplicity.

Effects of solid-to-solvent ratio on phenolic content

3.3

The identified phenolic compounds at different solid-to-solvent ratios (1:75, 1:150, 1:750, and 1:1500 g/mL) are presented in Table 4. Eight phenolic compounds were identified in Tetraselmis tetrahele extract, including gallic acid, catechin, ferulic acid, quercetin, 4-hydroxybenzoic acid, p-coumaric acid, protocatechuic acid, and kaempferol. The total identified phenolic compound content ranged from 788.54 to 793.50 μg/g biomass. There was no significant difference (p > 0.05) in the total identified phenolic compounds across the different solid-to-solvent ratios. Even though the solid-to-solvent ratio of 1:1500 g/mL yielded the highest total phenolic compounds (793.50 μg/g), the lowest solid-to-solvent ratio (1:75 g/mL) would be better in balancing the extraction efficiency and cost-effectiveness, especially in the industry-scale production. These results are in agreement with Luque-Alcaraz et al. [33], which suggested that the variation of solid-to-solvent ratio did not affect significantly in the extraction of vine grape (Vitis vinifera L.) using UAE. This result may be attributed to the low solid-to-solvent ratio, such as 1:75, is sufficient for dissolving most phenolic compounds. In this context, the degree of cell disruption becomes a more critical factor than the solvent volume. Once equilibrium is achieved, further increasing the solvent volume does not proportionally enhance the extraction of bioactive compounds.Table 4. Identified phenolic compounds (μg/g biomass) at different solid-to-solvent ratios under UAE.Content1:75 g/mL1:150 g/mL1:750 g/mL1:1500 g/mL4-Hydroxybenzoic acid75.46 ± 3.65^bc^71.87 ± 0.72^c^77.26 ± 1.05^b^91.29 ± 1.75^a^Protocatechuic acid13.22 ± 0.19^ab^13.09 ± 0.22^ab^13.31 ± 0.31^a^12.78 ± 0.07^b^p-Coumaric acid53.93 ± 0.02^d^54.04 ± 0.06^c^54.29 ± 0.06^b^54.49 ± 0.07^a^Gallic acid174.85 ± 0.35^a^176.63 ± 1.62^a^175.86 ± 0.57^a^176.30 ± 1.59^a^Ferulic acid20.34 ± 0.06^a^20.40 ± 0.06^a^20.30 ± 0.05^a^19.98 ± 0.04^b^Kaempferol73.08 ± 0.16^a^72.84 ± 0.07^b^72.69 ± 0.04^bc^72.58 ± 0.04^c^Catechin276.22 ± 1.77^a^275.32 ± 2.05^a^273.84 ± 0.89^a^261.56 ± 0.21^b^Quercetin104.15 ± 0.24^b^104.36 ± 0.13^b^105.00 ± 0.26^a^104.52 ± 0.44^ab^Total identified phenolic compounds791.25 ± 2.00^a^788.54 ± 4.03^a^792.53 ± 2.49^a^793.50 ± 2.68^a^^abcd^Mean ± standard deviation with distinct superscript letters within the same row are significantly different (p < 0.05) based on Tukey’s test.

In the ultrasonication process with solid-to-solvent ratio higher than 1:75, more solvent would not significantly enhance the cell disruption rate and solubility. Conversely, excessive solvent may reduce the energy to propagate and hit the target biomass per unit. This may also counteract the benefit from the gradient and limit the effect of the high solid-to-solvent ratio. These results are consistent with Santos et al. [34], which mentioned that most species will have the optimal phenolic compounds and antioxidants using the biomass-to-solvent ratio of 1:75. Although a higher ratio, such as 1:1500 g/mL, may result in a marginally higher phenolic content, the difference is not statistically significant. Importantly, the 1:75 ratio uses 95% less solvent compared to 1:1500, leading to substantial cost savings, especially in large-scale industrial applications. Moreover, lower solvent volumes reduce the energy required in downstream processes such as evaporation and purification.

Effects of time on phenolic content

3.4

Table 5 shows five phenolic compounds detected at different time durations of UAE of Tetraselmis tetrahele, including gallic acid, catechin, ferulic acid, 4-hydroxybenzoic acid, and protocatechuic acid. The total identified phenolic compounds ranged from 277.20 to 280.00 μg/g biomass, with the highest yield achieved at 15 min of extraction time. This was significantly higher (p < 0.05) than the yield achieved at 45 and 90 min of extraction time. One unanticipated finding was that the identified individual and total phenolic compounds were different from the results presented in Table 2, Table 3, Table 4. This could be attributed to the physiological variation between different batches of Tetraselmis tetrahele biomass. Despite being cultivated in the same medium, environmental factors and storage conditions (e.g., light intensity, nutrient availability, and temperature fluctuations) could influence the cellular morphology and physiological state, which in turn affect metabolite composition and extraction outcomes [35].Table 5. Identified phenolic compounds (μg/g biomass) at different UAE times.Content15 min45 min90 min4-Hydroxybenzoic acid44.34 ± 0.65^a^42.54 ± 0.72^b^42.40 ± 0.38^b^Protocatechuic acid8.24 ± 0.12^a^7.96 ± 0.13^ab^8.08 ± 0.12^b^Gallic acid89.05 ± 0.23^a^88.37 ± 0.25^b^88.43 ± 0.10^b^Ferulic acid9.85 ± 0.037^a^9.81 ± 0.02^a^9.81 ± 0.05^a^Catechin128.54 ± 0.43^a^128.56 ± 0.05^a^128.47 ± 0.19^a^Total identified phenolic compound content280.00 ± 1.37^a^277.23 ± 1.12^b^277.20 ± 0.23^b^^ab^ Mean ± standard deviation with distinct superscript letters within the same row are significantly different (p < 0.05) based on Tukey’s test.

The current study found that 15 min of the ultrasonication time (with 3 repetitions of 5 min extraction) was the appropriate ultrasonication duration with the highest yield of major phenolics, such as 4-hydroxybenzoic acid, protocatechuic acid, gallic acid, and ferulic acid. This finding can be attributed to the prolonged ultrasonication may result in the decomposition of the phenolic compounds [36]. A high concentration of reactive species, such as hydroxyl radicals, can be generated through extended exposure to ultrasound treatment in a water–ethanol medium [37]. These radicals can decompose phenolic compounds, thereby reducing the total phenolic compounds content [38]. This observation aligns with Chimsook and Wannalangka [39] and Gao et al. [40], which reported that the optimal extraction of phenolics from Spirogyra sp. and Asparagopsis taxiformis was achieved at 15 min and 10–20 min, respectively. Overall, the selected ultrasonication time of 15 min yielded the highest content of phenolic compounds, which achieved effective yields while promoting sustainable practices in saving energy and production costs, especially in the aspect of mass production.

Comparison in phenolic compounds between UAE and maceration using UHPLC

3.5

Table 6 presents identified individual phenolic compounds from Tetraselmis tetrahele using UAE and maceration extraction methods, which revealed significant differences (p < 0.05) between UAE and maceration. The used biomass was obtained from different batches of cultivation, compared with the biomass used in the previous quantification of phenolic compounds. Thus, different phenolic compounds were observed in Table 6. Gallic acid, 4-hydroxybenzoic acid, protocatechuic acid, and vanillic acid were detected in Tetraselmis tetrahele extracts by UAE and maceration. The results showed that UAE (236.68 ± 0.61 μg/g) yielded significantly higher (p < 0.05) phenolic content compared to the maceration method (232.61 ± 1.53 μg/g), particularly in the levels of gallic acid and 4-hydroxybenzoic acid.Table 6. Comparison of identified phenolic compounds (μg/g biomass) between UAE and maceration methods.ContentUAEMaceration4-Hydroxybenzoic acid36.69 ± 0.26^a^34.28 ± 0.66^b^Protocatechuic acid6.97 ± 0.07^a^6.75 ± 0.25^a^Vanillic acid104.36 ± 0.07^a^104.49 ± 0.27^a^Gallic acid88.67 ± 0.32^a^87.09 ± 0.60^b^Total identified phenolic compounds236.68 ± 0.61^a^232.61 ± 1.53^b^^ab^ Mean ± standard deviation with distinct superscript letters within the same row are significantly different (p < 0.05) based on 2-sample t-test.

Overall, UAE proved to be more exceptional than the maceration in extracting phenolic compounds. This enhanced efficiency can be due to the intensified interaction between the solvent and biomass during ultrasound treatment. Ultrasonication has been shown to permeabilize cell walls and induce pore formation, facilitating the release of intracellular compounds. In the cavitation process, the cavitation bubbles will grow because the gas in the medium will diffuse into the bubbles with periodic pressure oscillation [41]. Because of the rapid change in pressure, the cavitation bubbles will grow to a critical size and collapse violently, producing localized shockwaves and extreme mechanical shear forces [42]. As a result, ultrasound-induced cavitation can disrupt the microalgal cell wall and cellular structure, promoting the distribution of compounds into the solvent. Alongside the cavitation effects, secondary phenomena would also be triggered and apply to the microalgal cells. For the biomass near the point of collapse, shear forces, immediate shock waves, interparticle collisions, and localized high temperatures can produce solvent microjets, which assist in the fragmentation or erosion of the boundary of the biomass. The reduction of the particle size would increase the surface area of the microalgal cells, contributing to a higher mass transfer, extraction rate, and yield. The cavitation process was proven to result in the fragmentation and erosion to enhance the extraction efficiency compared to the maceration. If the cavitation bubbles collapse in the liquid, macroscopic turbulence would be formed and mixed with the micro mixing, leading to an increased mixing effect and diffusion rate [43].

In addition, UAE can contribute to the ultrasonic capillary effect (UCE), which may increase the penetration speed and depth, leading to the rapid swelling and rehydration of the dry biomass. This would further enhance the desorption and diffusion of compounds from the microalgae biomass. Furthermore, the various physical effects generated during UAE (e.g., cavitation, shear forces, fragmentation and UCE) occur sequentially and synergistically, significantly degrading the microalgae cell wall and membrane, enhancing the penetration process and direct contact of the solvent and intracellular compound, improving both mass transfer and solute diffusion rates [43].

In contrast, maceration relies on prolonged soaking to soften and disrupt cell walls for compound release, making it a slower and less efficient method. Its disadvantages include extended extraction times, high solvent consumption, and limited scalability, rendering it unsuitable for industrial-scale production. Therefore, UAE presents a more efficient and sustainable alternative to maceration, offering enhanced extraction yields within a shorter duration.

Comparison in antioxidant activity and anti-collagenase activity between UAE and maceration

3.6

Table 7 shows the photosynthetic pigments contents (chlorophyll a, b, and total carotenoids) and the antioxidant activities (TFC, TPC, and ABTS) obtained from UAE were significantly higher (p < 0.05) than those using the maceration method, except for the DPPH, which showed no significant difference (p > 0.05). These results are consistent with the current findings that phenolic compounds in microalgae were better extracted using UAE. This could be attributed to UAE not only improves the extraction of phenolics but also facilitates the release of other non-phenolic antioxidants such as carotenoids, chlorophylls, and vitamin C [44], which collectively enhance antioxidant activity, as shown in Table 7. Farahin et al. [45] reported that Tetraselmis tetrahele extracts obtained via maceration with 100% methanol (1:20 ratio) for 4 days with stirring at 80 rpm exhibited a markedly high DPPH radical scavenging activity of 17.20 ± 1.81 mg Trolox/g biomass. In contrast, the ABTS radical scavenging activity (2.50 ± 0.43 mg Trolox/g biomass) and TPC value (2.04 ± 0.67 mg gallic acid/g biomass) were comparatively low relative to this study, suggesting limited extraction efficiency of certain bioactive compounds under maceration.Table 7. Comparison of the antioxidant activity and anti-collagenase activities of Tetraselmis tetrahele extracts obtained using UAE and maceration methods.AssayUAEMacerationChlorophyll a (mg/g biomass)8.60 ± 0.13^a^3.92 ± 0.02^b^Chlorophyll b (mg/g biomass)7.46 ± 0.44^a^2.72 ± 0.01^b^Total carotenoid content (mg/g biomass)6.72 ± 0.36^a^2.45 ± 0.02^b^TFC (mg QE/g biomass)16.07 ± 1.35^a^10.29 ± 0.90^b^TPC (mg GAE/g biomass)14.50 ± 2.89^a^7.88 ± 1.56^b^DPPH (mg Trolox/g biomass)1.85 ± 0.33^a^1.17 ± 0.09^a^ABTS (mg Trolox/g biomass)6.26 ± 0.18^a^3.11 ± 0.05^b^Anti-collagenase activity, IC_50_ (μg/mL)52.43 ± 0.62^a^63.17 ± 5.47^a^^ab^ Mean ± standard deviation followed by distinct superscript letters within the same row are significantly different (p < 0.05) based on 2-sample t-test.

Fig. 1 shows both UAE and maceration extracts displayed absorption peaks at the wavelengths of 337 nm, 440 nm, 466 nm, and 664 nm in the spectral profiles. The peaks at 440 nm and 664 nm correspond to chlorophyll a [46], indicating that chlorophyll a was the major pigment in the Tetraselmis tetrahele extract. The UAE extract presented significantly higher chlorophyll a content (8.60 ± 0.13 mg/g), which was twofold higher than the maceration extract (3.92 ± 0.02 mg/g). The linear relationship between absorbance and concentration was consistent with the Beer-Lambert Law, as indicated by the observed absorbance values. Besides that, the peak at 466 nm corresponds to the carotenoid pigment. Conlon et al. [47] reported the lutein (0.13 ± 0.02–0.39 ± 0.15 mg lutein/g) and β-carotene (3.81 ± 0.22–7.40 ± 0.29 mg β-carotene/g) were the main carotenoid pigments in Tetraselmis species. Moreover, the peak at 337 nm corresponds to the possible presence of UV-protective compounds, such as phenolics and mycosporine-like amino acids [48]. These findings recommend that Tetraselmis tetrahele extract has the capacity to offer UV protection in edible cosmetics or cosmeceutical formulations due to their high UV absorption efficiency.Fig. 1. Absorption spectra in the UV–Vis region (200–700 nm) of the Tetraselmis tetrahele extracts with (A) UAE and (B) maceration. The peak at 337 nm suggests UV-absorbing compound, bands at 440 nm and 664 nm correspond to chlorophyll a, and the peak at 466 nm is associated with carotenoids. brief title: Absorption spectra of Tetraselmis tetrahele extracts.

Regarding collagenase inhibition activity, the Tetraselmis tetrahele extract obtained through UAE (IC_50_: 52.9 μg/mL) exhibited 15.9% more effectively compared to the maceration extract (IC_50_: 62.9 μg/mL), although the difference was not statistically significant (p > 0.05). According to Findrianny et al. [49], IC_50_ values below 50 μg/mL indicate extremely strong activity, values between 50–100 μg/mL indicate strong activity, 101–150 μg/mL indicate moderate activity, and values above 150 μg/mL indicate weak activity. Thus, the Tetraselmis tetrathele showed strong anti-collagenase ability in both UAE and maceration. The observed anti-collagenase activity is likely attributed to the high levels of bioactive compounds, particularly phenolic compounds, extracted more efficiently through UAE, which can inhibit collagenase and thereby offer anti-aging potential. This indicates the potential of Tetraselmis tetrahele extract as an anti-aging agent in cosmeceutical formulations. Compared to previous studies, the collagenase inhibition activity of Tetraselmis tetrathele was higher than Chlorella emersonii (IC_50_: 2.50 ± 0.03 mg/mL) [50] and most of the red, brown, and green seaweed extracts (IC_50_: 77.14 − >100 µg/mL) [51]. However, the collagenase inhibition percentage of Tetraselmis tetrahele extracts (62.7 – 75.6%; 80 µg/mL) was slightly lower than 80 µg/mL of Spirulina platensis extract (86.21 ± 0.97%) and Chlorella vulgaris (90.52 ± 0.87%) extract [52]. Thus, Tetraselmis tetrathele extract by UAE could be a potential source in the food and pharmaceutical industries.

Metabolic profiling of Tetraselmis tetrahele extract using LC-QTOF-MS

3.7

The metabolic profiling of Tetraselmis tetrahele extract revealed a diverse metabolite profile consisting of amino-fatty acids, lipids, terpenoids, apocarotenoids, and xanthophyll derivatives, as shown in Table 8. DL-2-Aminooctanoic acid represents an amino-fatty acid-like molecule [53], which indicates a lipid and nitrogen metabolism in Tetraselmis tetrahele. 7E,10-undecadien-4-olide, a lactone related with lipid oxidation, and the short peptide Ala-Trp-Arg, were detected. These compounds showed an active lipid and protein metabolism in the microalgae, likely indicating physiological responses during growth and stress conditions [54]. Arginine (semi-essential amino acid) and tryptophan (essential amino acids) contribute to the antioxidant activity and anti-aging pathways to protect the cells against oxidative stress [55]. Peptides derived from microalgae can effectively scavenge free radicals and stimulate collagen synthesis, thereby helping to delay premature skin aging. In addition to direct anti-collagenase activity observed in this study, microalgae-derived peptides have been demonstrated to suppress collagen degradation, which contribute to long-term skin elasticity and firmness [56]. Moreover, the detection of membrane glycerophospholipids, including PE(20:5/0:0), PG(18:3/18:3), PG(22:6/18:3), and PG(20:5/18:4) indicate that Tetraselmis tetrahele produces polyunsaturated fatty acids, such as alpha-linolenic acid (C18:3), eicosapentanoic acid (EPA) (C20:5), and docosahexaenoic acid (DHA) (C22:6). This finding is consistent with previous study describing Tetraselmis tetrahele as being high in polyunsaturated fatty acids [57]. Polyunsaturated fatty acids have been proven to maintain the skin’s barrier functions, and omega-3 fatty acids can protect cells against UV-induced skin damage by minimizing oxidative stress and inflammation [56]. The detection of phytosphingosine, oleamide, and 13E-docosenamide may associate with lipid metabolism in microalgae [58].Table 8LC-ESI-QTOF-MS analysis of Tetraselmis tetrahele extract.Peak numberCompound nameMS fragment (m/z)Retention timeFormulaBase peakScoreChemical classPositive ionization mode1DL-2-Aminooctanoic acid159.12550.909C_8_H_17_NO_2_83.046997.33Amino fatty acid27E,10-undecadien-4-olide180.11428.398C_11_H_16_O_2_83.049383.89Lactone, lipid32′',6′'-Di-O-acetylononin514.148510.612C_26_H_26_O_11_201.054589.97Modified ononin, a natural isoflavone.4(3R,8E)-3-Hydroxy-5,8-megastigmadien-7-one208.146211.495C_13_H_20_O_2_83.081286.45Enone54,5-Dihydrovomifoliol226.156411.534C_13_H_22_O_3_81.067896.62Sesquiterpenoid (a type of terpene)6Phytosphingosine317.291812.394C_18_H_39_NO_3_318.299094.62Sphingoid base in complex lipids717-hydroxyandrostane-3-glucuronide484.266713.210C_25_H_40_O_9_507.255798.01Steroid glucuronide conjugate8Ala Trp Arg414.203814.005C_20_H_29_N_7_O_4_119.083598.12A short peptide (alanine, tryptophan, and arginine)9Estra-1,3,5(10)-triene-3,6alpha,17beta-triol triacetate431.228714.151C_24_H_30_O_6_119.083197.80Steroid ester10PE(20:5(5Z,8Z,11Z,14Z,17Z)/0:0)499.269514.293C_25_H_42_NO_7_P328.272697.83Phosphatidylethanolamine11Ganolucidic acid B502.328714.484C_30_H_46_O_6_520.363397.50Triterpenoid12Corchorosol A536.296315.030C_29_H_44_O_9_537.306788.69Cardenolide glycoside13Campesteryl ferulate576.417515.299C_38_H_56_O_4_221.148097.94Steroid ester14SLF524.290215.322C_30_H_40_N_2_O_6_104.102094.14Synthetic ligand15Verimol C300.136216.724C_18_H_20_O_4_834.260296.09Anisoles162′-Dehydroplectaniaxanthin566.414619.045C_40_H_54_O_2_121.101190.82Xanthophyll carotenoid17Oleamide281.271719.336C_18_H_35_NO284.328685.70Fatty acid amide1813E-Docosenamide337.335320.296C_22_H_43_NO69.069495.52Fatty acid amide19Tridodecylamine521.592220.467C_36_H_75_N522.598889.90Tertiary amine209Z-Hexacosene364.408420.758C_26_H_52_382.441693.19Alkene21PG(18:3(6Z,9Z,12Z)/18:3(6Z,9Z,12Z))766.477921.074C_42_H_71_O_10_P767.485088.70Phosphatidylglycerols2213-Methyl-1-tritriacontene476.532323.709C_34_H_68_494.566298.88AlkeneNegative ionization mode1Embelin294.182517.369C_17_H_26_O_4_96.960285.31Benzoquinone2PG(22:6(4Z,7Z,10Z,13Z,16Z,19Z)/18:3(9Z,12Z,15Z))816.494519.169C_46_H_73_O_10_P815.485391.24Phosphatidylglycerols3PG(20:5(5Z,8Z,11Z,14Z,17Z)/18:4(6Z,9Z,12Z,15Z))788.464120.487C_44_H_69_O_10_P275.196996.62Phosphatidylglycerols4Erythromycin E747.439221.094C_37_H_65_NO_14_61.988396.02Macrolide antibiotics5PI(18:3(9Z,12Z,15Z)/13:0)790.459821.236C_40_H_71_O_13_P277.205084.88Phosphatidylinositol

A range of apocarotenoids, such as (3R,8E)-3-hydroxy-5,8-megastigmadien-7-one, 4,5-dihydrovomifoliol, and the xanthophyll derivative, 2′-dehydroplectaniaxanthin, were also detected. Previous study reported Tetraselmis sp. presented high carotenoid content (8.48 ± 0.47 mg g^−1^ DW) with 3.17 ± 0.18 mg g^−1^ DW lutein and 3.21 ± 0.18 mg g^−1^ DW β-carotene. The high carotenoid content in microalgae responsible for the adaptation mechanisms to oxidative stress and photoprotection [59]. Thus, the presence of apocarotenoids and xanthophyll derivative indicates active carotenoid metabolism in Tetraselmis tetrahele. Furthermore, the improved oxidative defence mechanism by high carotenoid content in Tetraselmis tetrahele can contribute to skin health and fight against premature aging [56].

Ganolucidic acid B, corchorosol A, and verimol C represent terpenoid and triterpenoid families, while embelin and 2′',6′'-di-O-acetylononin represent the presence of phenolic secondary metabolites. Campesteryl ferulate, a phytosterol ester, further reflects the diversity of complex lipids in the extract [60]. These compounds are always associated with antioxidant, anti-inflammatory, antimicrobial, and cellular protection properties [61]. The wide array of bioactive compounds tentatively detected in Tetraselmis tetrahele suggests that the secondary metabolites production in microalgae encountering with different environmental conditions. These metabolites can enhance the antioxidant capacity of microalgae and fight against environmental damage and premature aging [56]. Overall, the diverse metabolite profile highlights the potential of Tetraselmis tetrahele as a valuable and sustainable source of bioactive compounds to use in aquaculture, nutraceutical, cosmeceutical, and pharmaceutical applications.

Conclusion

4

This study investigated the effects of solvent composition, temperature, solid-to-solvent ratio, and time in extracting phenolic compounds from microalga Tetraselmis tetrahele using UAE. The appropriate UAE parameters were determined as a biomass-to-solvent ratio of 1:75, a temperature of 25 °C, 75% ethanol, and an ultrasonication time of 15 min. The Tetraselmis tetrahele extract from the selected UAE exhibited higher phenolic compounds, antioxidant activities, and anti-collagenase activity than the conventional maceration method. The LC-QTOF-MS profiling of Tetraselmis tetrahele extract revealed a diverse metabolic profile constituting of amino-fatty acids, lipids, terpenoids, apocarotenoids, and xanthophyll derivatives. The metabolite diversity demonstrated the capacity of Tetraselmis tetrahele extract using UAE to serve as a sustainable and valuable source of bioactive compounds for applications.

Batch-to-batch variability in the phenolic profiles in Tetraselmis tetrahele represents a limitation in this study. This discrepancy may be attributed to physiological differences among different biomass batches. The time of harvesting, slight differences in cultivation stages, and small fluctuations in ambient temperature can affect phenolic biosynthesis and accumulation in the microalgae. Moreover, post-harvest handling and storage conditions would also affect the phenolic composition in the microalgae [35]. Such variability is expected in microalgal secondary metabolite production due to their sensitivity to the subtle environment and physiological changes. Multiple independent batches of microalgae cultivation and standardized environmental conditions are always expected to reduce the variability in the secondary metabolites production and bioactivity [62]. Besides that, scale-up study is required to enable industrial applications of UAE in obtaining Tetraselmis tetrahele extracts. In addition, future research should continue to explore additional bioactivities of Tetraselmis tetrahele extracts, including its anti-inflammatory, anti-cancer, and immunomodulatory activities, as well as comprehensive toxicological evaluations via in vitro and in vivo studies. This can provide comprehensive insights to the industry regarding the effects of these metabolites into the modulation of oxidative stress, enzyme inhibition, or signaling pathways involved in inflammation and tumor progression to expand its applications in pharmaceutical, food, and cosmetic industries.

CRediT authorship contribution statement

Tao Mai: Writing – original draft, Visualization, Validation, Software, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Norazira Abdu Rahman: Writing – review & editing, Supervision, Resources, Methodology, Investigation, Data curation. Swee Yun Pang: Writing – review & editing, Visualization, Validation, Supervision, Resources, Data curation. Jian Ping Tan: Writing – review & editing, Visualization, Validation, Resources. Sook Chin Chew: Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Methodology, Funding acquisition, Data curation, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.