Enhancing bioactivity of Chlorella vulgaris through enzymatic pretreatment and lactic acid fermentation

doi:10.21203/rs.3.rs-8249047/v1

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Enhancing bioactivity of Chlorella vulgaris through enzymatic pretreatment and lactic acid fermentation

https://doi.org/10.21203/rs.3.rs-8249047/v1

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This study explores an integrated bioprocessing approach to enhance the bioactivity of Chlorella vulgaris biomass through enzymatic pretreatment and lactic acid fermentation. Three pretreatment strategies—ultrasound-assisted hydrolysis, hydrothermal treatment, and enzymatic hydrolysis using Viscozyme, Alcalase, or their combination- were evaluated for their ability to release fermentable sugars and proteins. Enzymatic hydrolysis proved most effective, yielding up to 151.18 mg g DW^− 1 of reducing sugars and 14.4 mg g DW^− 1 of protein. Subsequent fermentation with Lactiplantibacillus plantarum and Levilactobacillus brevis demonstrated robust microbial growth (Δ Log CFU ≈ 4.0) and significant acidification (pH 3.6–4.2), accompanied by lactic acid production up to 17.35 g/L. Functional characterisation revealed that combined enzymatic pretreatment and fermentation improved antioxidant properties, as indicated by CUPRAC, ABTS, DPPH, and TPC assays, and selectively enhanced antimicrobial activity against Gram-positive bacteria. These findings highlight the synergistic effect of enzymatic pretreatment and microbial fermentation in enhancing the biofunctional value of C. vulgaris, offering a sustainable strategy for developing natural antioxidants and antimicrobial agents for food, nutraceutical, and cosmetic applications.

microalgae

Chlorella vulgaris

fermentation

lactic acid bacteria

enzyme pretreatment

antioxidant capacity

In recent years, with growing demand for sustainable and health-promoting food, there has been increased interest in microalgae as a renewable resource for food production. Beyond sustainability, microalgae, due to their nutrient-dense content and reported biofunctional properties, offer tremendous potential for applications in food, nutraceuticals, cosmetics, and pharmaceuticals (Sivaramakrishnan et al. 2025). Among microalgal species, Chlorella vulgaris is particularly attractive due to its high protein (≈ 43–58% DW) and carbohydrate (≈ 12–55% DW) content, as well as its role as a source of functional lipids, water-soluble vitamins and pigments (Wang et al. 2024). Its carbohydrate profile is rich in starch, specifically containing 30–40% of the linear α-glucan amylose. Beyond starch, Chlorella species have a rigid cell wall, composed primarily of polysaccharides containing galactose, uronic acids, and rhamnose (Ferreira et al. 2020). The latter features make C. vulgaris a promising substrate for microbial fermentation, particularly by lactic acid bacteria (LAB), which require fermentable sugars as primary carbon sources (Abedi and Hashemi 2020; Al-Hammadi and Güngörmüşler).

In fact, algal fermentation is increasingly recognised as a strategy that could enhance the nutritional and functional properties of algae, while improving their digestibility, bioaccessibility and sensory properties (Pérez-Alva et al. 2022). Particularly, lactic acid fermentation has been widely applied in food systems for thousands of years due to its capacity to improve safety, stability, and organoleptic characteristics while generating or releasing metabolites such as bacteriocins, peptides, and physically entrapped substances (Aguirre-Garcia et al. 2024). Therefore, the physicochemical transformations that occur during fermentation may enhance the health-promoting potential of microalgal biomass, thereby positioning it for use in functional foods and nutraceuticals (Garofalo et al. 2022).

However, the recalcitrant nature of C. vulgaris cell wall, consisting of resistant polysaccharides and glycoproteins, poses a barrier that limits the accessibility of intracellular nutrients, thereby hindering fermentation efficiency (Spain and Funk 2022). As a result, effective pretreatments that disrupt the cell wall and release fermentable sugars are a critical step in the downstream process before further valorisation of these substrates. Towards this, various studies have focused on different pretreatments, such as microwaves (Lee et al. 2013), bead milling (Postma et al. 2017), steam explosion (Lorente et al. 2015), ultrasonication, acid or alkali treatment, and enzymatic hydrolysis have been suggested to help break down its robust cell walls and facilitate the release of intracellular compounds. Nevertheless, recent reviews have highlighted the limitations of mechanical, thermal, and chemical cell disruption approaches, as economic inefficiency, sustainability concerns, and environmental impact often constrain them (Alhattab et al. 2019; Md Nadzir et al. 2023). As a result, biological approaches are increasingly favoured for their eco-friendly and cost-effective nature.

Despite growing interest in microalgal fermentation, few studies have systematically compared the effectiveness of various pretreatment techniques in enhancing fermentable sugar release from C. vulgaris, particularly in the context of LAB fermentation. In this study, we evaluated three pretreatment strategies: ultrasonication and autoclave with or without acid, and enzymatic hydrolysis using viscozyme, alcalase, or a combination of both, as strategies to enhance carbon availability from C. vulgaris for lactic acid fermentation. Among the various approaches, enzymatic hydrolysis yielded the best results. We then further assessed the fermentation performance using two LAB strains, Lactiplantibacillus plantarum DSM 24624 and Levilactobacillus brevis DSM 20556. Moreover, key fermentation parameters, including microbial growth, pH dynamics, and organic acid production, as well as functional properties such as antioxidant and antimicrobial activities, were monitored. Overall, his study provides a targeted evaluation of pretreatment strategies, followed by fermentation of enzyme-treated biomass variants, offering new insights into optimising microalgal substrates for functional food applications.

Chemicals and algal biomass

Spray-dried green algal biomass powder, C. vulgaris, was obtained by Phytobloom (Necton S.A., Portugal). Sulfuric acid (98%), phosphoric acid (85%), ethanol (99%) commercial enzyme complex, viscozyme (≥ 100 FBGU/g, Novozyme, Denmark), alcalase (2.964 U mL^− 1, Merck, Denmark) potassium sodium tartrate, sodium hydroxide, 3,5-dinitrosalicylic acid, D-glucose powder, bovine serum albumin, Coomassie brilliant blue G-250, 2,2′-azino-bis- (3-ethylbenzothiazolin-6-ammonium sulfonate) (ABTS), 6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox), potassium peroxodisulfate (K₂S₂O₈), sodium carbonate (Na2CO3), disodium phosphate, sodium chloride (NaCl), gallic acid (3,4,5-trihydroxybenzoic acid, 99%,), 2,2-diphenyl-1-picrylhydrazyl hydrate free radical (DPPH•, 95%), methanol, Folin–Ciocalteu reagent, copper (II) chloride solution (CuCl2.2H2O), potassium dihydrogen phosphate (KH₂PO₄), sodium hydrogen phosphate (Na₂HPO₄), neocuproine (C₁₄H₁₂N₂), ammonium acetate (NH₄CH₃CO₂), acetonitrile (C₂H₃N), formic acid (HCOOH), ammonium sulfate (NH₄)₂SO₄, and MRS broth medium (De Man, Rogosa, and Sharpe) were purchased from Sigma-Aldrich (St. Louis, MO, USA).

Proximate composition of C. vulgaris biomass

The protein content was measured using the Kjeldahl technique (AOAC 960.52). Moisture and ash levels were quantified through gravimetric analysis, following AOAC methods 925.10 and 900.02, respectively. Lipid content was evaluated by Soxhlet extraction with n-hexane, according to procedures previously established by our research team (Syrpas et al. 2018).

Pretreatment of C. vulgaris

Pretreatment methods were assessed using a 10% (w/v) spray-dried algal biomass solution. Specifically, 7.0 ± 0.01 g of biomass was combined with 70 mL of distilled water (control) or with the specified concentrations of sulfuric acid or enzymatic solutions. All experiments were performed in triplicate.

Ultrasound pretreatment

Ultrasound pretreatment was conducted using a Hielscher Ultrasonic Processor UP200Ht (Hielscher Ultrasonics, Teltow, Germany), as described previously with slight modifications (Syrpas et al. 2020). Specifically, the sonotrode operated at 90% amplitude (200 W, 26 kHz) and alternated between 10-second on and 10-second off cycles for a total duration of 30 minutes. Pretreatment was performed with sulfuric acid solutions at 1%, 2%, and 4% (v/v) or with distilled water as a control.

Autoclave pretreatment

For the autoclave pretreatment, C. vulgaris powder was mixed with sulfuric acid at 1, 2, and 4% (v/v) or with distilled water as a control. The mixture was autoclaved at 121°C and 1.1 atm for 30 minutes, then cooled to room temperature.

Enzymatic pretreatment

Enzymatic pretreatment was carried out following conditions previously described in the literature (Mahdy et al. 2014). Specifically, samples were treated with either Viscozyme (0.3 mL g DW^− 1), Alcalase (0.2 mL g DW^− 1), or a combination of both. The pH was adjusted to 5.5 for Viscozyme and 8.0 for Alcalase, and maintained as needed during hydrolysis. After enzyme addition, samples were incubated at 50°C in a Biosan Orbital Shaker Incubator (Latvia). Enzymes were then inactivated by heating at 75°C for 15 minutes, and the samples were stored at 4°C for further analysis.

Preparation of bacterial inoculum

L. plantarum DSM 24624 and L. brevis DSM 20556 were obtained from Biometrija (Kaunas, Lithuania). LAB were maintained on De Man–Rogosa–Sharpe (MRS) agar slants (REF 4017282, Liofilchem, Italy), at + 4°C in a Liebherr ProfiLine refrigerator. Inoculum preparation followed the method described before (Aboobacker et al. 2025). Briefly, a loopful of culture was transferred from the slant to 10 mL of MRS broth (REF 401729 REF 610037, Liofilchem, Italy) and incubated at 37°C for 16–18 hours. The MRS media was prepared by dissolving 14.04 g in 200 mL of distilled water, followed by autoclaving at 121 ⁰C for 15 minutes. The resulting overnight cultures were diluted 1:100 in sterile saline to prepare working suspensions of LAB before inoculation.

Fermentation of C. vulgaris with lactic acid bacteria

Prior to fermentation, 7 g of C. vulgaris biomass samples were pretreated with the enzymatic treatments as indicated above. The final volume of the corresponding solutions was 70 mL. Following this step, samples were inoculated with working suspensions of L. plantarum and L. brevis to achieve an initial LAB load of approximately 6 log CFU mL^− 1, under sterile conditions. Inoculated samples were then immediately placed in a Thermo Scientific Oxoid AnaeroJar (Thermo Fisher Scientific Inc., Waltham, MA, USA) anaerostats to maintain anaerobic conditions using Oxoid AnaeroGen Compact Sachets, and incubated at 37°C. After cell counting and pH measurements, the collected samples were centrifuged (Ortoalresa—Alvarez Redondo S.A., Madrid, Spain) at 6000 rpm for 10 minutes, then filtered through Whatman filter paper. Samples were stored at − 18°C in the freezer until subsequent analysis.

Microbial growth and pH analysis

Viable cell counts in the C. vulgaris fermentates were determined as described previously in the literature (Dave et al. 2025). At designated time points, samples were serially diluted (1:10) in 0.9% (w/v) saline and homogenised. Dilutions were plated on MRS agar (Scharlau Chemie, Spain) and incubated at 37°C for 36–48 hours. Results were expressed as log CFU mL^− 1.The pH of the fermented samples was monitored using a calibrated 32000P pH meter (Agilent Technologies Inc., Santa Clara, CA, USA). Measurements were taken immediately after aseptic sampling at each time point to monitor acidification during fermentation.

Determination of reducing sugar content

The concentration of reducing sugars released during pretreatment was determined using a modified 3,5-dinitrosalicylic acid (DNS) assay, based on the method by Miller (Miller 1959) with modifications by (Teixeira et al. 2012). Briefly, 1 mL of the sample was mixed with 1 mL of DNS reagent, prepared by dissolving 1 g of DNS and 30 g of sodium potassium tartrate in 100 mL of 0.4 M NaOH. Phenol was omitted from the reagent, as it can unnecessarily intensify colour. The mixture was boiled in a water bath for 5 minutes, then cooled to room temperature. The final volume was adjusted to 8 mL with distilled water, and absorbance was measured at 540 nm using a GENESYS 50 UV–Vis spectrophotometer (ThermoFisher Scientific, Waltham, United States). Reducing sugar concentrations were calculated using a glucose standard curve ranging from 0.2 to 1.0 mg mL^− 1.

Determination of protein content

Protein content in the treated samples was determined using a modified Bradford assay (Bradford 1976). In brief, 0.1 mL of the sample was mixed with 3 mL of filtered Bradford reagent, prepared by dissolving 100 mg of Coomassie Brilliant Blue G-250 in 50 mL of 95% ethanol, followed by the addition of 100 mL of 85% (w/v) phosphoric acid, and dilution to 1 L with distilled water. After vortexing, the samples were incubated at room temperature for 10 minutes, then centrifuged. Absorbance of optically clear supernatants was then measured at 595 nm using a GENESYS 50 UV–Vis spectrophotometer (ThermoFisher Scientific, Waltham, United States).

Determination of organic acids

Organic acids were analysed in a Shimadzu LC-30AD Nexera X2 HPLC system (Shimadzu Corporation, Kyoto, Japan). Chromatographic resolution was achieved on a Rezex ROA-organic acid H+ (8%) column (300 × 7.8 mm, Torrance, CA, USA), maintained at 60°C, using 0.005 M sulfuric acid as the mobile phase under isocratic conditions, at a flow rate of 0.5 mL min^− 1. Qualitative determination was performed using reference materials, whereas quantitative determination was performed using external calibration curves. The final results were expressed as mg mL^− 1 using dose-response curves for lactic acid (0.1-5 mg mL^− 1, y = 63740x − 6516.5, R² = 0.9979 ), acetic acid (0.1 -5 mg mL^− 1, y = 54906x − 748.81, R² = 0.9999)

Total phenolic content (TPC) by Folin-Ciocalteu’s assay

TPC of C. vulgaris supernatants was determined using the method of Singleton (Singleton et al. 1999) with modifications reported (Nagybákay et al. 2023). For each analysis, 150 µL of the fermented extract or distilled water (blank) was mixed with 750 µL of Folin–Ciocalteu reagent (diluted 1:9, v/v). After 3 minutes of reaction, 600 µL of a sodium carbonate solution (75 g/L) was added, and the mixture was vortexed for 15 seconds. The samples were then incubated in the dark for 2 hours. Absorbance was measured at 760 nm. TPC was expressed as milligrams of gallic acid equivalents (mg GAE/g extract or dry weight), based on a standard curve prepared with gallic acid (0–80 µg mL^− 1).

Cupric reducing antioxidant capacity assay

CUPRAC measurements were performed following the protocol of Apak (Apak et al. 2007). Shortly, 0.4 mL of Cu(II), neocuproine, and NH₄Ac buffer solutions were added to a test tube. Then, 0.4 mL of extract (or standard) was added to the initial mixture. The tubes were stoppered, and after 30 min, the absorbance was recorded at 450 nm. The CUPRAC antioxidant capacity was expressed as Trolox equivalents (either as mg TE g-1 extract or per g DW), employing a dose-response curve for Trolox (25–150 µmol L-1).

Determination of the ABTS•+ scavenging assay

The ABTS assay was carried out according to the method as described in the literature (Re et al. 1999). Firstly, phosphate-buffered saline (PBS) solution (75 mmol/L; pH 7.4) was prepared by dissolving 8.18 g of NaCl, 0.27 g of KH₂PO₄, 1.42 g of Na2HPO4, and 0.15 g of KCl in 1 L of ultra-pure water. The ABTS^•+ solution was prepared by mixing 50 mL of ABTS (2 mmol/L PBS) with 200 µL of K2S2O8 (70 mmol/L) and allowing the mixture to stand in the dark at room temperature for 15–16 hours prior to use. The working solution was prepared by diluting the ABTS•+ solution with PBS to achieve an absorbance of 0.700 ± 0.010 at 734 nm. To a 1500 µL working ABTS•+ solution, 25 µL of fermented extract or blank (methanol) was added. The mixtures were kept in the dark for 2 hours, and the absorbance was measured at 734 nm. The final results were expressed as Trolox equivalents (mg TE g-1 extract or DW), calculated employing dose-response curves for Trolox (0–1500 µmol L-1 methanol).

Determination of the DPPH scavenging assay

The DPPH radical scavenging assay was performed as previously described (Brand-Williams et al. 1995). To 500 µL of fermented extract or methanol (blank), 1000 µL of a ~ 89.7 µmol L-1 (absorbance adjusted to 0.800 ± 0.010 AU at 517 nm) DPPH• methanolic solution was added and vortexed for 15 s, and kept for 2 hours in the dark, and the absorbance of the optically clear supernatant was measured at 517 nm. Antioxidant activity was expressed as mg TE g-1 extract or DW, calculated using dose-response curves for Trolox (0–50 µmol L-1 methanol).

Determination of antimicrobial activity

The antibacterial activity of the samples was evaluated using the agar well diffusion method against reference strains: Escherichia coli ATCC 8739, Micrococcus luteus ATCC 9341, Staphylococcus aureus ATCC 25923, Bacillus subtilis ATCC 6633, and Pseudomonas aeruginosa. To prepare the bacterial inoculum, each strain was cultured on Plate Count Agar slants at 37°C for 18–20 hours. A loopful of the culture was transferred into sterile saline and adjusted to a 0.5 McFarland standard, corresponding to approximately 10⁸ CFU mL^− 1. Then, 20 mL of this suspension was mixed with 200 mL of molten Plate Count Agar, poured into Petri dishes, and allowed to solidify. Wells (9 mm diameter) were created using sterile pipette tips, and 25 µL of either unfermented control or 24-hour fermented samples was added to each well. Plates were incubated at 37°C for 24 hours, after which the diameters of inhibition zones were measured in millimetres using an electronic digital calliper.

Statistical Analysis

Mean values and standard deviations were calculated using Microsoft Excel. One-way ANOVA followed by Tukey’s post hoc test was conducted to compare means showing significant variation (p < 0.05), using GraphPad Prism software version 10.5 for Windows.

Compositional analysis of C. vulgaris

In the first part of this study, the proximate composition of C. vulgaris biomass was analysed, with the results presented in Fig. 1. As shown, proteins (~ 46%) and carbohydrates (~ 40%) were the primary components of the studied biomass. In comparison, lipid (~ 3% ) and ash contents (~ 6%) were comparatively lower (Fig. 1). These findings are consistent with a recent study, which reported ash, lipid, protein, and carbohydrate contents of approximately 9%, 0.5%, 40%, and 43%, respectively (Damayanti et al. 2025). Furthermore, previous reviews reported that the total protein content in mature C. vulgaris ranges from 42% to 58% of the biomass dry weight (Safi et al. 2014; Ijaola et al. 2024). Although carbohydrate levels are typically lower, Chlorella is known to accumulate high starch contents, particularly under nitrogen-limiting conditions, reaching a total carbohydrate content up to 55% of its dry weight (Brányiková et al. 2011). The observed lipid content in this study is lower than most reported values in the literature, ranging from 5% to 58% with higher content typically obtained under mixotrophic cultivation (Wang et al. 2024). Nevertheless, the biochemical and nutritional profile of C. vulgaris is highly dependent on cultivation conditions and the strain used (Hiltunen et al. 2026).

Evaluation of various pretreatment strategies

The first method used to disrupt the cell wall of C. vulgaris was ultrasound treatment, applied in distilled water or sulfuric acid solutions at varying concentrations (Fig. 2). Ultrasound-assisted extraction is a green technology used to recover bioactive compounds from algae with reduced solvent and energy input, while preserving molecular integrity. Ultrasound induces cavitation in liquid media, and the collapse of these microbubbles generates localised mechanical forces that disrupt the cell walls of microalgae, facilitating the release of intracellular compounds (Lee et al. 2017). As a technique, it has been widely applied to extract polysaccharides, pigments, and phenolics from various algal species (Carreira-Casais et al. 2021). The highest release of reducing sugars (~ 9 mg g DW^− 1) was achieved with 2% sulfuric acid (Fig. 2A), while the maximum protein content was observed with 4% acid. However, no statistically significant difference was observed compared with 2% acid (Fig. 2B). As shown, even at the highest yield achieved with this method, the carbohydrate conversion is relatively low, reaching 2.2%. This result closely agrees with a previous study utilising ultrasonic-assisted acid hydrolysis for Chlorella, which reported a carbohydrate conversion efficiency of 3.3%. The authors concluded that the application of ultrasonic treatments as the only method for releasing and depolymerising carbohydrates is insufficient (Nasirpour et al. 2023). Although ultrasound treatments are a useful pretreatment method in other applications, they may not achieve sufficient conversion of complex carbohydrates into simple sugars, which are essential for subsequent fermentation.

In the following step, hydrothermal hydrolysis was conducted at 121°C for 30 minutes using sulfuric acid concentrations of 1%, 2%, and 4% (v/v), with distilled water serving as the control. The results, summarised in Fig. 3, indicate that hydrolysis with 2% sulfuric acid yielded the highest amount of reducing sugars, reaching 102.4 mg g DW^− 1 (Fig. 3A). On the other hand, the lowest yield was observed for hydrothermal treatment with water as solvent, reaching a conversion efficiency of only 0.9%. A similar observation was also reported in a study on Chlorella, where the conversion efficiencies at 30 and 120 minutes of heating in the hydrothermal method were similarly low, at 0.5% and 1%, respectively (Nasirpour et al. 2023). Protein recovery was significantly higher in samples treated with 2% and 4% acid than in those treated with lower concentrations, although the difference between the two highest concentrations was not statistically significant (Fig. 3B).

Regardless of the technique used, the addition of sulfuric acid resulted in a substantially higher release of reducing sugars and proteins (Figs. 2 and 3). Dilute sulfuric acid is commonly employed for lignocellulosic biomass pretreatment due to its rapid reaction kinetics and enhanced cellulose hydrolysis efficiency (Esteghlalian et al. 1997; Sun and Cheng 2002). In practically all cases, a 2% sulfuric acid addition resulted in the highest recoveries. This observation aligns with a report by de Carvalho Silvello et al. (2023), who investigated the effect of 1%, 2.5%, and 4% sulfuric acid concentrations on the release of sugar from biomass. The authors indicated that 2.5% sulfuric acid yielded the highest sugar release, while increasing the concentration to 4% did not result in a significant improvement (de Carvalho Silvello et al. 2023). Moreover, in another study, the effects of 2% and 5.36% sulfuric acid at 120°C for 30 min on sugar release were compared, with the authors reporting that the highest yield (247 mg/ gDW) was achieved with the 2% acid concentration (Abou El-Souod et al. 2021). Nevertheless, it is worth noting that a previous study comparing five acids for C. vulgaris hydrolysis found that while hydrochloric, sulfuric, and nitric acids were more effective than peracetic and phosphoric acids, hydrochloric acid yielded the best overall results (Park et al. 2016). High recoveries (> 80%) have also been reported by other studies utilising hydrochloric acid for the pretreatment of Chlorella biomass towards bioethanol production (Kim et al. 2011; Zhou et al. 2011). Nonetheless, during acid hydrolysis, especially at elevated temperatures, carbohydrate degradation often leads to the formation of furanic compounds, such as hydroxymethylfurfural and furfural, which are known to inhibit microbial activity and reduce fermentation efficiency (Labbafi et al. 2025).

Considering these factors, the next step was to apply three enzymatic treatments to disrupt the cell wall matrix and enhance the release of intracellular compounds. Enzymatic hydrolysis has been proposed as an effective method to disrupt the cell wall of C. vulgaris, thereby facilitating the release of valuable intracellular components, including proteins, carbohydrates, and bioactive compounds (Rahman et al. 2022). Towards this, Viscozyme, a multi-enzyme complex with carbohydrase activity, was used to break down polysaccharides, while alcalase, a protease, targeted protein structures. Additionally, a combined treatment using both enzymes was performed to maximise cell wall degradation and improve nutrient availability for downstream processes. As shown in Fig. 4A, enzymatic hydrolysis, particularly with viscozyme alone and in combination, significantly increased the release of reducing sugars, yielding 139.61 and 151.18 mg g DW^− 1, respectively, compared to 4.34 mg g DW^− 1 in the control samples. These findings are in close agreement with a previous report, which reported that the highest soluble carbohydrate hydrolysis (160 mg g DW^− 1) was obtained with the carbohydrolase (V and V + A) after 2 h (Mahdy et al. 2014). On the other hand, alcalase treatment resulted in the highest protein release, reaching 14.4 mg g DW^− 1, corresponding to an extraction efficiency of 3.1%. Similar recovery values were reported in a previous study, where the enzymatic protein extraction process yielded an average efficiency of 3.80% (Yaghoubzadeh and Safari 2025).

Overall, among the various tested techniques, ultrasound resulted in the lowest recovery of fermentable sugars and protein, followed by the hydrothermal treatments. The low efficiency of ultrasound has also been reported in previous studies. For instance, de Carvalho et al. evaluated alkaline, acid, ultrasonic, and supercritical treatments and found that ultrasound in water (10 min, 200 W, 20 kHz) yielded less than 1 g/L of glucose, one of the lowest values among the tested methods (de Carvalho Silvello et al., 2023). Similarly, Paterson et al. applied ultrasound after freeze–thaw cycles, but microscopy revealed insufficient cell wall disruption, highlighting the need for more effective strategies (Paterson et al. 2024). Hydrothermal treatment also did not result in satisfactory results in this study. However, it should be noted that a previous report indicated that although the hydrothermal or ultrasonic treatment alone may be insufficient, their combination results in substantially higher recoveries of fermentable sugars (Nasirpour et al. 2023). The effectiveness of combinatorial approaches has also been suggested in other reports, where alkali and enzyme hydrolysis produced the highest reducing sugar yield (Effiom et al. 2023).

In this study, enzymatic treatments resulted in substantially higher recoveries of fermentable sugars, especially as compared to the other two methods. The efficiency of cellulases and alcalases in converting the complex carbohydrates of Chlorella into fermentable sugars has been demonstrated for biohydrogen (Liu et al. 2021; Sriyod et al. 2021) and bioethanol (Kim et al. 2014). In fact, a recent review concluded that compared to other pretreatment methods, enzymatic hydrolysis offers advantages for bioethanol production due to its low cost, minimal environmental impact, and higher ethanol yields, as it also operates under mild conditions, requiring less energy and no harsh chemicals, thereby avoiding the formation of fermentation inhibitors (Kusmiyati et al. 2023). Besides bioenergy production, enzymatic hydrolysis has also been used as a strategy to increase the functional properties of Chlorella hydrolysates (Yin et al. 2010; Gharehbeglou et al. 2024). An additional benefit of this approach is the absence of fermentation inhibitors. Considering all these factors, enzyme-assisted hydrolysis was selected as the optimal strategy prior to LAB fermentation, as discussed in the following sections.

Fermentation of C. vulgaris by lactic acid bacteria

Microbial growth and acification during LAB fermentation

In this part of the study, C. vulgaris biomass underwent enzymatic hydrolysis prior to fermentation by LAB. To evaluate the effectiveness of the fermentation process and the capacity of pretreated biomass to support microbial activity, the growth of inoculated LAB strains was monitored and expressed as log CFU mL⁻¹. Concurrently, pH was monitored to assess acidification of the fermentation medium. The results of these parts are summarised in Fig. 5. Before inoculation, the cell densities of L. plantarum and L. brevis were standardised to approximately 6 log CFU mL⁻¹ to ensure consistent initial microbial loads across all fermentation trials. As depicted in Fig. 5, both strains exhibited substantial growth after 24 hours of fermentation. In samples fermented with L. plantarum, the highest growth of 10.12 log CFU mL⁻¹, corresponding to a Δ Log CFU mL⁻¹ of 4.1, was observed for the sample treated with Viscozyme (FVPL, Fig. 5A). The other two treatments, FAPL and FVAPL, also showed a significant increase in cell density with Δ Log CFU mL⁻¹ of 3.6 and 4.0, respectively, thereby overall indicating robust proliferation under the given conditions (Fig. 5A).

Similar trends were observed in samples inoculated with L. Brevis, where all treatments supported effective growth. The Viscozyme-treated samples (FVBR) reached the highest cell density of 9.76 log CFU mL⁻¹, with Δ Log CFU mL⁻¹ of 4.1 (Fig. 5C). The other treatments also showed strong LAB proliferation, with increases of ~ 3.9 Log units.

In addition to microbial growth, acidification of the medium was used as an indicator of fermentation efficiency. As illustrated in Fig. 5B and 5D, both LAB strains demonstrated significant acidification capacity within 24 hours of fermentation. The lowest pH values were recorded in the Viscozyme-treated samples, reaching 3.6 for L. plantarum (Fig. 5B) and 4.2 for L. brevis (Fig. 5D). In contrast, the lowest acidification for both strains was observed in the alcalase-treated samples.

These findings align with previous studies that have reported the beneficial effects of Chlorella species on the viability and growth of probiotic strains such as Lactobacillus and Bifidobacterium. In most cases, Chlorella biomass was incorporated either into fermented semi-solid dairy matrices such as cheese and yoghurt (Kovaleski and Ventura 2025) or into freeze-dried probiotic formulations (Meireles Mafaldo et al. 2022; Fortuin et al. 2026). For instance, Ścieszka and Klewicka demonstrated that supplementing MRS medium with varying concentrations of C. vulgaris biomass accelerated the growth of four L. brevis strains, shortening their logarithmic growth phase (Ścieszka and Klewicka 2020). Moreover, another study reported that C. vulgaris increased the survival of Lactobacillus spp. in bile salts. They noted that the protective effect of algae at low pH is strain-dependent, concluding that the biomass of this microalga could be used in fermented dietary products (Sylwia and Elżbieta 2020).

However, only a limited number of studies have explored the use of C. vulgaris biomass as a sole carbon source for LAB fermentation. In a recent study, Niocolloti et al. investigated the fermentation of four C. vulgaris biomasses with varying carbohydrate content using 5 LAB strains, including L. plantarum 4932. Their results showed that all biomasses fermented with L. plantarum reached pH values below 4.5 after 72 hours of fermentation, although microbial growth was modest, reaching a maximum of 2.1 (Nicolotti et al. 2025). Moreover, another study reported co-fermentation of Aphanizomenon flos-aquae and C. vulgaris biomasses with L. plantarum and S. cerevisiae; however, the authors do not report either cell growth or pH values (Tomassi et al. 2025). Taken together, the results of the present study highlight the importance of enzymatic pretreatment in enhancing the fermentability of C. vulgaris biomass. Compared to previous reports, the observed microbial proliferation and acidification suggest that enzymatic hydrolysis significantly improves the bioavailability of fermentable substrates, thereby supporting more efficient LAB growth and metabolic activity.

Analysis of sugars and organic acids

In parallel with the growth experiments, the reducing sugar content and organic acid production were monitored, with results summarised in Fig. 6. Generally, L. plantarum exhibits facultative heterofermentative metabolism, meaning its fermentation products can differ depending on the environmental conditions (Siezen and van Hylckama Vlieg 2011). Most LAB thrive in anaerobic environments, where they convert pyruvate produced via the Embden-Meyerhof-Parnas pathway into lactate. On the other hand, L. brevis is an obligate heterofermentative LAB, as it consistently produces a mixture of lactic acid, carbon dioxide, and ethanol or acetic acid during fermentation, regardless of culture conditions. Unlike facultative fermenting bacteria, it relies on the phosphoketolase pathway to metabolise sugars. A significant reduction in reducing sugar concentration was observed after 24 h of fermentation (Supplementary Material Fig. S1). As compared to alcalase-treated samples, sugar consumption was higher in viscozyme-treated and combined-treated samples. Chromatographic analysis confirmed that the hydrolysates primarily contained glucose, fructose, and xylose, along with maltose (Fig. 6 and Supplementary Material Fig. S2). Due to chromatographic limitations, fructose and xylose were not fully resolved. These findings are consistent with previous reports on the carbohydrate composition and enzymatic hydrolysis of C. vulgaris, which typically release glucose, fructose, xylose, and maltose (Safi et al. 2014; Mahdy et al. 2014; El-Naggar et al. 2020). Both LAB strains exhibited similar overall sugar consumption, except in Alcalase-treated samples, where L. plantarum consumed more sugars than L. brevis. Xylose utilisation was greater in L. brevis, aligning with its metabolic capacity to ferment pentoses via the phosphoketolase pathway (Zhang et al. 2016). This pathway also explains the small amounts of acetic acid detected in samples fermented by L. brevis (Supplementary Fig. S3).

Correspondingly, the production of organic acid is presented in Fig. 7. The sample treated with combined enzyme and fermented with L. plantarum exhibited the highest lactic acid production among all tested conditions, at 17.35 g L-1, followed by the sample treated with viscozyme alone, at 12.20 g L^− 1. These findings align with the measured reducing sugar, as viscozyme- and combined-treated samples showed higher sugar release from the matrix, confirming the increased conversion to lactic acid. Besides, a small concentration of acetic acid production was observed in the samples, including those treated with alcalase and combined enzyme treatments, and fermented by L. brevis, with values of 1.03 and 1.10 g L^− 1, respectively (Supplementary Materials Fig. S3).

The enhanced fermentation performance observed in Viscozyme and combined enzyme treatments further highlights the importance of cell wall disruption in microalgae. Several reports on enzymatic pretreatment of Chlorella species for biofuel and bioproduct generation have shown similar benefits, including increased sugar release and improved microbial activity (Córdova et al. 2019; Yang et al. 2023). In a previous study, the authors explored fermentation strategies for lactic acid production with immobilised L. plantarum 23 using C. vulgaris hydrolysate as feedstock. They tested various aeration conditions to determine the oxygen requirements for this strain to be homolactic (Chen et al. 2020). The study concluded that although lactic acid’s yield and productivity were not substantially different under the different oxygen availability, the strain was homolactic only under anaerobic conditions (Chen et al. 2020).

Nevertheless, the ability of C. vulgaris to serve as a sole carbon source for LAB fermentation, especially when pretreated enzymatically, could open new avenues for sustainable bioprocessing. This strategy could also be considered particularly relevant for developing functional foods and probiotic formulations, as highlighted by recent work on engineered C. vulgaris strains with enhanced carbohydrate content (Saha et al. 2024).

Antioxidant activity of fermented C. Vulgaris

The antioxidant capacity of C. vulgaris biomass and its extracts is primarily associated with a diverse range of bioactive compounds, including chlorophyll, carotenoids (such as lutein and β-carotene), polysaccharides, and polyphenols (Mendes et al. 2024). Although fermentation has been proposed as a strategy to enhance the antioxidant activity of microalgae, data on its impact on C. vulgaris and its extracts remain limited. In this study, the in vitro antioxidant capacity of fermented C. vulgaris was assessed using four complementary assays: CUPRAC, total phenolic content (TPC), ABTS, and DPPH radical scavenging tests (Fig. 8 and Supplementary Material Fig. S4). Enzyme-treated but non-fermented samples served as controls (0h in all figures). Across all assays, combined enzyme treatment yielded the highest antioxidant capacity, with values of 8.64 mg Trolox g DW⁻¹ (CUPRAC), 9.67 mg GAE g DW⁻¹ in the TPC assay (Fig. 8). Similar observations with 19.46 mg Trolox g DW⁻¹ in the ABTS assay, and 1.45 mg Trolox g DW⁻¹ in the DPPH assay were recorded (Supplementary Material Fig. S4). However, most differences between fermented and enzyme-treated samples were not statistically significant.

When a single enzyme pretreatment was applied, the effects of fermentation became more evident, with fermented samples generally exhibiting higher antioxidant activity than unfermented ones. This observation aligns with recent findings by Tomassi et al., who reported that fermentation of C. vulgaris using LAB and yeast significantly improved its antioxidant profile. Specifically, TPC nearly doubled from 2.4 mg GAE g DW⁻¹ at the onset of fermentation to 4.8 mg GAE g DW⁻¹ after 24 hours, with similar trends observed for DPPH and ORAC assays (Tomassi et al. 2025).

The increase in antioxidant capacity observed in this study can be attributed to several mechanisms. Enzymatic pretreatment with cellulases (e.g., Viscozyme) facilitates the release of intracellular antioxidants, including phenolics, carotenoids, chlorophylls, and peptides (Sanjeewa et al. 2023; Badrunanto et al. 2025). Moreover, although proteases can contribute to cell wall disruption, the observed increase in antioxidant capacity is attributed to the generation of bioactive peptides with antioxidant properties. These observations are further supported by previous studies on other substrates that have shown similar effects with proteases and cellulases. For instance, Viscozyme was utilised to obtain a high-yield bilberry extract with enhanced antioxidant capacity (Syrpas et al. 2021). Alcalase was used to generate bioactive peptides from mesopelagic fish protein hydrolysates (Hayes et al. 2024). In a study involving lentil proteins, the highest content of phenolic compounds (3.8 mg GAE/g) was achieved through sequential hydrolysis using Alcalase and Flavourzyme (Rezvankhah et al. 2021) .

Additionally, studies suggest that fermentation enhances antioxidant capacity by breaking down complex or bound antioxidant compounds into more bioavailable and active forms, and by generating new bioactive molecules such as peptides and vitamins (Verni et al. 2019). These effects depend on the specific metabolic activities of the microbial strains used. Overall, these findings demonstrate that combining enzymatic pretreatment with microbial fermentation is an effective strategy for improving the biofunctional value of C. vulgaris. From an application perspective, such improvements expand the potential of microalgal biomass as a natural antioxidant source for food fortification, nutraceuticals, and cosmetic formulation, areas where stability and natural origin are increasingly prioritised.

Antimicrobial activity of fermented C. vulgaris

Alternative antimicrobial products have gained increasing attention as a response to rising drug resistance and environmental concerns. As a result, the antimicrobial potential of microalgae as a source of sustainable and effective bioactive compounds for diverse applications has been explored (Xia et al. 2021). In fact, extracts from C. vulgaris exhibit antimicrobial properties (Kitada et al. 2009; Fassi Fihri et al. 2024), making this species a promising candidate for the development of functional ingredients for food and pharmaceutical use (Yuan et al. 2020). To assess this potential, the antimicrobial activity of fermentation supernatants was evaluated against five bacterial strains (Table 1). Among the tested samples, only those treated with Viscozyme and fermented with L. plantarum displayed antimicrobial effects.

In contrast, neither the unfermented samples nor those fermented with L. brevis inhibited any strain. The strongest activity was observed in the Viscozyme-treated sample (FVPL), which inhibited all strains except P. aeruginosa, with zones ranging from 8.6 mm (S. aureus) to 12.4 mm (M. luteus). Moderate inhibition was noted against E. coli (Table 1). Enzymatic hydrolysis using alcalase did not enhance antimicrobial properties, as no inhibitory effects were detected in alcalase-treated samples.

Table 1

Antimicrobial activity of enzyme-treated and fermented *C. vulgaris* by *L. plantarum* and *L. brevis*
	Inhibition Zone, mm
	S. Aureus	B. Subtilis	M. Luteus	E. Coli	P. aeruginosa
UPL	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00
UBR	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00
VPL	8.64 ± 0.01	10.37 ± 0.06	12.38 ± 0.13	9.64 ± 0.03	0.00 ± 0.00
VAPL	7.79 ± 0.13	7.90 ± 0.05	9.54 ± 0.37	0.00 ± 0.00	0.00 ± 0.00

Results represent mean ± SD. (UPL and UBR: untreated biomass and fermented with L. plantarum and L. brevis, respectively; VPL: viscozyme treated and fermented by L. plantarum; VAPL: combined enzyme treated and fermented by L. plantarum).

These findings are consistent with previous work on macroalgae, in which alcalase treatment failed to confer antimicrobial activity across the species tested (K. Habeebullah et al. 2020). The activity observed in Viscozyme-treated samples likely reflects efficient carbohydrate release, supporting the higher metabolic activity of L. plantarum. Generally, the antimicrobial effects of LAB-fermented products are linked to organic acid production, peptide or proteinaceous bacteriocins, and other small molecules (Ibrahim et al. 2021). Among these, lactic acid is notable for lowering pH and permeabilising the membranes of Gram-negative bacteria, such as E. coli, thereby enhancing the action of other antimicrobial compounds (Alakomi et al. 2000). However, fermentate activity typically cannot be attributed to a single molecule (Figueroa et al. 2024). In fact, the absence of activity in L. brevis-fermented samples suggests that the antimicrobial effect of L. plantarum is likely due to bacteriocin production. Many L. plantarum strains synthesise plantaricins, bacteriocins of interest for their diverse applications, including food biopreservation, mitigation of irritable bowel syndrome symptoms, and protection against urinary tract infections (Abdulhussain Kareem and Razavi 2020). L. plantarum strains that produce plantaricin are valued for their gut-friendly characteristics, broad carbohydrate utilisation, pleasant sensory profile, and probiotic properties, making them highly suitable for applications in the food industry (Goel and Halami 2023).

This work demonstrates that enzymatic pretreatment combined with lactic acid fermentation is an effective and eco-friendly strategy to enhance the bioactivity of C.vulgaris biomass. Among the tested pretreatment methods, enzymatic hydrolysis, particularly the combined use of Viscozyme and Alcalase, significantly improved the release of fermentable sugars and proteins, thereby supporting efficient LAB growth and organic acid production. Fermentation further contributed to functional improvements, notably increased antioxidant capacity and selective antimicrobial activity. These results demonstrate the potential of integrated bioprocessing for valorising microalgal biomass into high-value biofunctional ingredients. Future research should focus on scaling up the process, optimising strain selection, and exploring metabolomic profiling to better understand the mechanisms underlying bioactivity enhancement.

Author contributions

Conceptualisation, M.S. and V.K.S.; methodology, M.S. and V.K.S.; investigation, H.B, S.A and A.S.; resources, M.S.; data curation, H.B., V.K.S. and M.S.; writing—original draft preparation, H.B.; writing—review and editing, A.S, A.S, V.K.S. and M.S.; visualisation, H.B.; V.K.S. and M.S. supervision, M.S.; project administration, M.S. and V.K.S.; funding acquisition, M.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Research Council of Lithuania (RCL), project no.: S-MIP-23-78, Projects carried out by researchers’ teams.

Ethics approval and consent to participate

Not applicable

Consent for publication

Not applicable. All authors have read and agreed to the published version of the manuscript.

Availability of data and materials

All data generated or analysed during this study are included in this published article [and its supplementary information files].

Competing interests

The authors declare no competing financial interests or personal relationships that could have influenced the work reported in this paper. The funders had no role in the study’s design, data collection, analysis, interpretation, manuscript writing, or decision to publish the results.

Abdulhussain Kareem R, Razavi SH (2020) Plantaricin bacteriocins: As safe alternative antimicrobial peptides in food preservation—A review. J Food Saf 40:e12735. https://doi.org/10.1111/jfs.12735
Abedi E, Hashemi SMB (2020) Lactic acid production – producing microorganisms and substrates sources-state of art. Heliyon 6:e04974. https://doi.org/10.1016/j.heliyon.2020.e04974
Aboobacker S, Kitrytė-Syrpa V, Šipailienė A, Rutkaitė R, Syrpas M (2025) Fermentation-induced nutritional and in vitro antioxidant capacity changes in Arthrospira platensis (spirulina). Food Bioscience 68:106747. https://doi.org/10.1016/j.fbio.2025.106747
Abou El-Souod GW, Morsy EM, Hassan LHS, El-Sheekh MM (2021) Efficient Saccharification of the Microalga Chlorella vulgaris and its Conversion into Ethanol by Fermentation. Iran J Sci Technol Trans Sci 45:767–774. https://doi.org/10.1007/s40995-021-01097-1
Aguirre-Garcia YL, Nery-Flores SD, Campos-Muzquiz LG, Flores-Gallegos AC, Palomo-Ligas L, Ascacio-Valdés JA, Sepúlveda-Torres L, Rodríguez-Herrera R (2024) Lactic Acid Fermentation in the Food Industry and Bio-Preservation of Food. Fermentation 10:168. https://doi.org/10.3390/fermentation10030168
Alakomi H-L, Skyttä E, Saarela M, Mattila-Sandholm T, Latva-Kala K, Helander IM (2000) Lactic Acid Permeabilizes Gram-Negative Bacteria by Disrupting the Outer Membrane. Appl Environ Microbiol 66:2001–2005. https://doi.org/10.1128/AEM.66.5.2001-2005.2000
Al-Hammadi M, Güngörmüşler M New insights into Chlorella vulgaris applications. https://doi.org/10.1002/bit.28666
Alhattab M, Kermanshahi-Pour A, Brooks MS-L (2019) Microalgae disruption techniques for product recovery: influence of cell wall composition. J Appl Phycol 31:61–88. https://doi.org/10.1007/s10811-018-1560-9
Apak R, Güçlü K, Demirata B, Özyürek M, Çelik SE, Bektaşoğlu B, Berker KI, Özyurt D (2007) Comparative Evaluation of Various Total Antioxidant Capacity Assays Applied to Phenolic Compounds with the CUPRAC Assay. Molecules 12:1496–1547. https://doi.org/10.3390/12071496
Badrunanto, Purba LDA, Wikantyasning ER, Maryati, Prayitno J, Hidhayati N, Agustini NWS, Munawaroh HSH, Susanti H, Kusumawaty D, Rahman DY, Apriastini M, Prasetyadi, Admirasari R (2025) Chlorella sp. as a promising natural antioxidant source: Research trends, bioactive components, and future perspectives. Algal Res 92:104341. https://doi.org/10.1016/j.algal.2025.104341
Bradford MM (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248–254. https://doi.org/10.1016/0003-2697(76)90527-3
Brand-Williams W, Cuvelier ME, Berset C (1995) Use of a free radical method to evaluate antioxidant activity. LWT - Food Sci Technol 28:25–30. https://doi.org/10.1016/S0023-6438(95)80008-5
Brányiková I, Maršálková B, Doucha J, Brányik T, Bišová K, Zachleder V, Vítová M (2011) Microalgae—novel highly efficient starch producers. Biotechnol Bioeng 108:766–776. https://doi.org/10.1002/bit.23016
Carreira-Casais A, Otero P, Garcia-Perez P, Garcia-Oliveira P, Pereira AG, Carpena M, Soria-Lopez A, Simal-Gandara J, Prieto MA (2021) Benefits and Drawbacks of Ultrasound-Assisted Extraction for the Recovery of Bioactive Compounds from Marine Algae. Int J Environ Res Public Health 18:9153. https://doi.org/10.3390/ijerph18179153
Chen P-T, Hong Z-S, Cheng C-L, Ng I-S, Lo Y-C, Nagarajan D, Chang J-S (2020) Exploring fermentation strategies for enhanced lactic acid production with polyvinyl alcohol-immobilized Lactobacillus plantarum 23 using microalgae as feedstock. Bioresour Technol 308:123266. https://doi.org/10.1016/j.biortech.2020.123266
Córdova O, Passos F, Chamy R (2019) Enzymatic Pretreatment of Microalgae: Cell Wall Disruption, Biomass Solubilisation and Methane Yield Increase. Appl Biochem Biotechnol 189:787–797. https://doi.org/10.1007/s12010-019-03044-8
Damayanti A, Megawati, Winaningsih I, Al Taghna E, Supriyadi M, Zahra SA, Khusniyyah R, Adonai Tandirogang N, Avril Ervian Then N (2025) Comparative study of pretreatment methods for the reverse enzymatic hydrolysis of Chlorella and Spirulina microalgae for bioethanol production. Biofuels 16:905–911. https://doi.org/10.1080/17597269.2025.2469384
Dave AD, Bilgin H, Kitrytė-Syrpa V, Syrpas M (2025) Optimisation of Pressurised Liquid Extraction and Subsequent Hydrolysate Fermentation by Lactiplantibacillus plantarum for Integrated Bioprocessing of Ulva sp. Mar Drugs 23:371. https://doi.org/10.3390/md23100371
de Carvalho Silvello MA, Gasparotto GA, Ferreira GF, Santos LO, Fregolente LV, Goldbeck R (2023) Nutrient Optimization Strategy to Increase the Carbohydrate Content of Chlorella vulgaris and Evaluation of Hydrolysis and Fermentation Performance. Bioenerg Res 16:2058–2067. https://doi.org/10.1007/s12155-023-10660-0
Effiom DI, Afegbua SL, Ado SA, Chia MA (2023) Effect of single and combined pretreatment strategies applied to Chlorella vulgaris biomass on bioethanol production. Biofuels 14:165–172. https://doi.org/10.1080/17597269.2022.2123938
El-Naggar NE-A, Hussein MH, Shaaban-Dessuuki SA, Dalal SR (2020) Production, extraction and characterization of Chlorella vulgaris soluble polysaccharides and their applications in AgNPs biosynthesis and biostimulation of plant growth. Sci Rep 10:3011. https://doi.org/10.1038/s41598-020-59945-w
Esteghlalian A, Hashimoto AG, Fenske JJ, Penner MH (1997) Modeling and optimization of the dilute-sulfuric-acid pretreatment of corn stover, poplar and switchgrass. Bioresour Technol 59:129–136. https://doi.org/10.1016/S0960-8524(97)81606-9
Fassi Fihri R, Ez-Zoubi A, Mbarkiou L, Amar A, Farah A, Bouchamma EO (2024) Antibacterial and antioxidant activities of Chlorella vulgaris and Scenedesmus incrassatulus using natural deep eutectic solvent under microwave assisted by ultrasound. Heliyon 10:e35071. https://doi.org/10.1016/j.heliyon.2024.e35071
Ferreira AS, Ferreira SS, Correia A, Vilanova M, Silva TH, Coimbra MA, Nunes C (2020) Reserve, structural and extracellular polysaccharides of Chlorella vulgaris: A holistic approach. Algal Res 45:101757. https://doi.org/10.1016/j.algal.2019.101757
Figueroa RHH, López-Malo A, Mani-López E (2024) Antimicrobial activity and applications of fermentates from lactic acid bacteria – a review. Sustainable Food Technol 2:292–306. https://doi.org/10.1039/D3FB00241A
Fortuin J, Leclercq CC, Silva RK, Shaplov AS, Contal S, Cambier S, Iken M, Fogliano V, Soukoulis C (2026) Chlorella vulgaris protein isolate effectively protects Lacticaseibacillus rhamnosus GG viability during processing, storage, and in vitro digestion. Food Hydrocolloids 172:111999. https://doi.org/10.1016/j.foodhyd.2025.111999
Garofalo C, Norici A, Mollo L, Osimani A, Aquilanti L (2022) Fermentation of Microalgal Biomass for Innovative Food Production. Microorganisms 10:2069. https://doi.org/10.3390/microorganisms10102069
Gharehbeglou P, Sarabandi K, Akbarbaglu Z (2024) Insights into enzymatic hydrolysis: Exploring effects on antioxidant and functional properties of bioactive peptides from Chlorella proteins. J Agric Food Res 16:101129. https://doi.org/10.1016/j.jafr.2024.101129
Goel A, Halami PM (2023) Structural and biosynthetic diversity of plantaricins from Lactiplantibacillus. Appl Microbiol Biotechnol 107:5635–5649. https://doi.org/10.1007/s00253-023-12692-0
Hayes M, Naik A, Mora L, Iñarra B, Ibarruri J, Bald C, Cariou T, Reid D, Gallagher M, Dragøy R, Galino J, Deyà A, Albrektsen S, Thoresen L, Solstad RG (2024) Generation, Characterisation and Identification of Bioactive Peptides from Mesopelagic Fish Protein Hydrolysates Using In Silico and In Vitro Approaches. Mar Drugs 22:297. https://doi.org/10.3390/md22070297
Hiltunen M, Kannangara R, Nakandalage B, Söderena O, Timilsina H, Pirhonen J, Pulkkinen K (2026) Growth and biomass composition of Chlorella vulgaris using nutrient-rich water and CO2 from a recirculating aquaculture system. Aquaculture 610:742956. https://doi.org/10.1016/j.aquaculture.2025.742956
Ibrahim SA, Ayivi RD, Zimmerman T, Siddiqui SA, Altemimi AB, Fidan H, Esatbeyoglu T, Bakhshayesh RV (2021) Lactic Acid Bacteria as Antimicrobial Agents: Food Safety and Microbial Food Spoilage Prevention. Foods 10:3131. https://doi.org/10.3390/foods10123131
Ijaola AO, Akamo DO, George TT, Sengul A, Adediji MY, Asmatulu E (2024) Algae as a potential source of protein: A review on cultivation, harvesting, extraction, and applications. Algal Res 77:103329. https://doi.org/10.1016/j.algal.2023.103329
Habeebullah K, Alagarsamy SF, Sattari S, Al-Haddad Z, Fakhraldeen S, Al-Ghunaim S, Al-Yamani A F (2020) Enzyme-assisted extraction of bioactive compounds from brown seaweeds and characterization. J Appl Phycol 32:615–629. https://doi.org/10.1007/s10811-019-01906-6
Kim KH, Choi IS, Kim HM, Wi SG, Bae H-J (2014) Bioethanol production from the nutrient stress-induced microalga Chlorella vulgaris by enzymatic hydrolysis and immobilized yeast fermentation. Bioresour Technol 153:47–54. https://doi.org/10.1016/j.biortech.2013.11.059
Kim N-J, Li H, Jung K, Chang HN, Lee PC (2011) Ethanol production from marine algal hydrolysates using Escherichia coli KO11. Bioresour Technol 102:7466–7469. https://doi.org/10.1016/j.biortech.2011.04.071
Kitada K, Machmudah S, Sasaki M, Goto M, Nakashima Y, Kumamoto S, Hasegawa T (2009) Antioxidant and Antibacterial Activity of Nutraceutical Compounds from Chlorella vulgaris Extracted in Hydrothermal Condition. Sep Sci Technol 44:1228–1239. https://doi.org/10.1080/01496390902729056
Kovaleski G, Ventura SPM (2025) Benefits, challenges and future of incorporation of algae into dairy products such as cheese and yogurt. J Food Compos Anal 147:108045. https://doi.org/10.1016/j.jfca.2025.108045
Kusmiyati K, Hadiyanto H, Fudholi A (2023) Treatment updates of microalgae biomass for bioethanol production: A comparative study. J Clean Prod 383:135236. https://doi.org/10.1016/j.jclepro.2022.135236
Labbafi S, Zahedi A, Kanani B (2025) Optimizing bioethanol production from Chlorella vulgaris and Dunaliella salina using integrated renewable energy systems. Biomass Bioenergy 202:108186. https://doi.org/10.1016/j.biombioe.2025.108186
Lee C-G, Kang D-H, Lee D-B, Lee H-Y (2013) Pretreatment for Simultaneous Production of Total Lipids and Fermentable Sugars from Marine Alga, Chlorella sp. Appl Biochem Biotechnol 171:1143–1158. https://doi.org/10.1007/s12010-013-0295-y
Lee SY, Show PL, Ling TC, Chang J-S (2017) Single-step disruption and protein recovery from Chlorella vulgaris using ultrasonication and ionic liquid buffer aqueous solutions as extractive solvents. Biochem Eng J 124:26–35. https://doi.org/10.1016/j.bej.2017.04.009
Liu H, Ru G, Zhang Z, Li Y, Xia C, Lu C, Zhang Q (2021) Experimental study on optimization of initial pH for photo-fermentation bio-hydrogen under different enzymatic hydrolysis of chlorella vulgaris. Bioresour Technol 338:125571. https://doi.org/10.1016/j.biortech.2021.125571
Lorente E, Farriol X, Salvadó J (2015) Steam explosion as a fractionation step in biofuel production from microalgae. Fuel Processing Technology 131:93–98. https://doi.org/10.1016/j.fuproc.2014.11.009
Mahdy A, Mendez L, Ballesteros M, González-Fernández C (2014) Enhanced methane production of Chlorella vulgaris and Chlamydomonas reinhardtii by hydrolytic enzymes addition. Energy Conv Manag 85:551–557. https://doi.org/10.1016/j.enconman.2014.04.097
Md Nadzir S, Yusof N, Nordin N, Kamari A, Yusoff MZM (2023) A review of microalgal cell wall composition and degradation to enhance the recovery of biomolecules for biofuel production. Biofuels 14:979–997. https://doi.org/10.1080/17597269.2023.2197730
Meireles Mafaldo Í, de Medeiros VPB, da Costa WKA, da Costa Sassi CF, da Costa Lima M, de Souza EL, Eduardo Barão C, Colombo Pimentel T, Magnani M (2022) Survival during long-term storage, membrane integrity, and ultrastructural aspects of Lactobacillus acidophilus 05 and Lacticaseibacillus casei 01 freeze-dried with freshwater microalgae biomasses. Food Res Int 159:111620. https://doi.org/10.1016/j.foodres.2022.111620
Mendes AR, Spínola MP, Lordelo M, Prates JAM (2024) Chemical Compounds, Bioactivities, and Applications of Chlorella vulgaris in Food, Feed and Medicine. Appl Sci 14:10810. https://doi.org/10.3390/app142310810
Miller GL (1959) Use of Dinitrosalicylic Acid Reagent for Determination of Reducing Sugar. Anal Chem 31:426–428. https://doi.org/10.1021/ac60147a030
Nagybákay NE, Sarapinaitė L, Syrpas M, Venskutonis PR, Kitrytė-Syrpa V (2023) Optimization of pressurized ethanol extraction for efficient recovery of hyperoside and other valuable polar antioxidant-rich extracts from Betula pendula Roth leaves. Ind Crops Prod 205:117565. https://doi.org/10.1016/j.indcrop.2023.117565
Nasirpour N, Ravanshad O, Mousavi SM (2023) Ultrasonic-assisted acid and ionic liquid hydrolysis of microalgae for bioethanol production. Biomass Conv Bioref 13:16001–16014. https://doi.org/10.1007/s13399-021-02274-3
Nicolotti C, Cirlini M, Del Vecchio L, Hadj Saadoun J, Bernini V, Gatti M, Bottari B, Martelli F (2025) Lactic Acid Fermentation of Chlorella vulgaris to Improve the Aroma of New Microalgae-Based Foods: Impact of Composition and Bacterial Growth on the Volatile Fraction. Foods 14:1511. https://doi.org/10.3390/foods14091511
Park C, Lee JH, Yang X, Yoo HY, Lee JH, Lee SK, Kim SW (2016) Enhancement of hydrolysis of Chlorella vulgaris by hydrochloric acid. Bioprocess Biosyst Eng 39:1015–1021. https://doi.org/10.1007/s00449-016-1570-4
Paterson S, Majchrzak M, Alexandru D, Di Bella S, Fernández-Tomé S, Arranz E, de la Fuente MA, Gómez-Cortés P, Hernández-Ledesma B (2024) Impact of the biomass pretreatment and simulated gastrointestinal digestion on the digestibility and antioxidant activity of microalgae Chlorella vulgaris and Tetraselmis chuii. Food Chem 453:139686. https://doi.org/10.1016/j.foodchem.2024.139686
Pérez-Alva A, MacIntosh AJ, Baigts-Allende DK, García-Torres R, Ramírez-Rodrigues MM (2022) Fermentation of algae to enhance their bioactive activity: A review. Algal Res 64:102684. https://doi.org/10.1016/j.algal.2022.102684
Postma PR, Suarez-Garcia E, Safi C, Yonathan K, Olivieri G, Barbosa MJ, Wijffels RH, Eppink MHM (2017) Energy efficient bead milling of microalgae: Effect of bead size on disintegration and release of proteins and carbohydrates. Bioresour Technol 224:670–679. https://doi.org/10.1016/j.biortech.2016.11.071
Rahman MM, Hosano N, Hosano H (2022) Recovering Microalgal Bioresources: A Review of Cell Disruption Methods and Extraction Technologies. Molecules 27:2786. https://doi.org/10.3390/molecules27092786
Re R, Pellegrini N, Proteggente A, Pannala A, Yang M, Rice-Evans C (1999) Antioxidant activity applying an improved ABTS radical cation decolorization assay. Free Radic Biol Med 26:1231–1237. https://doi.org/10.1016/S0891-5849(98)00315-3
Rezvankhah A, Yarmand MS, Ghanbarzadeh B, Mirzaee H (2021) Generation of bioactive peptides from lentil protein: degree of hydrolysis, antioxidant activity, phenol content, ACE-inhibitory activity, molecular weight, sensory, and functional properties. Food Measure 15:5021–5035. https://doi.org/10.1007/s11694-021-01077-4
Safi C, Zebib B, Merah O, Pontalier P-Y, Vaca-Garcia C (2014) Morphology, composition, production, processing and applications of Chlorella vulgaris: A review. Renew Sustain Energy Rev 35:265–278. https://doi.org/10.1016/j.rser.2014.04.007
Saha S, Maji S, Ghosh SK, Maiti MK (2024) Engineered Chlorella vulgaris improves bioethanol production and promises prebiotic application. World J Microbiol Biotechnol 40:271. https://doi.org/10.1007/s11274-024-04074-z
Sanjeewa KKA, Herath KHINM, Kim Y-S, Jeon Y-J, Kim S-K (2023) Enzyme-assisted extraction of bioactive compounds from seaweeds and microalgae. TRAC Trends Anal Chem 167:117266. https://doi.org/10.1016/j.trac.2023.117266
Ścieszka S, Klewicka E (2020) Influence of the Microalga Chlorella vulgaris on the Growth and Metabolic Activity of Lactobacillus spp. Bacteria Foods 9:959. https://doi.org/10.3390/foods9070959
Siezen RJ, van Hylckama Vlieg JE (2011) Genomic diversity and versatility of Lactobacillus plantarum, a natural metabolic engineer. Microb Cell Fact 10:S3. https://doi.org/10.1186/1475-2859-10-S1-S3
Singleton VL, Orthofer R, Lamuela-Raventós RM (1999) [14] Analysis of total phenols and other oxidation substrates and antioxidants by means of folin-ciocalteu reagent. In: Methods in Enzymology. Academic Press, pp 152–178
Sivaramakrishnan R, Kanwal S, Incharoensakdi A, Nirmal N, Srimongkol P (2025) Exploring the nutraceutical and functional food potential of microalgae: Implications for health and sustainability. J Agric Food Res 22:102148. https://doi.org/10.1016/j.jafr.2025.102148
Spain O, Funk C (2022) Detailed Characterization of the Cell Wall Structure and Composition of Nordic Green Microalgae. J Agric Food Chem 70:9711–9721. https://doi.org/10.1021/acs.jafc.2c02783
Sriyod K, Reungsang A, Plangklang P (2021) One-step multi enzyme pretreatment and biohydrogen production from Chlorella sp. biomass. Int J Hydrog Energy 46:39675–39687. https://doi.org/10.1016/j.ijhydene.2021.09.232
Sun Y, Cheng J (2002) Hydrolysis of lignocellulosic materials for ethanol production: a review. Bioresour Technol 83:1–11. https://doi.org/10.1016/S0960-8524(01)00212-7
Sylwia Ś, Elżbieta K (2020) Algae Chlorella vulgaris as a factor conditioning the survival of Lactobacillus spp. adverse Environ conditions LWT 133:109936. https://doi.org/10.1016/j.lwt.2020.109936
Syrpas M, Bukauskaite J, Paskauskas R, Basinskiene L, Venskutonis P (2018) Recovery of lipophilic products from wild cyanobacteria (Aphanizomenon flos-aquae) isolated from the Curonian Lagoon by means of supercritical carbon dioxide extraction. ALGAL RESEARCH-BIOMASS BIOFUELS Bioprod 35:10–21. https://doi.org/10.1016/j.algal.2018.08.006
Syrpas M, Bukauskaitė J, Ramanauskienė K, Karosienė J, Majienė D, Bašinskienė L, Venskutonis PR (2020) Ultrasound-Assisted Extraction and Assessment of Biological Activity of Phycobiliprotein-Rich Aqueous Extracts from Wild Cyanobacteria (Aphanizomenon flos-aquae). J Agric Food Chem 68:1896–1909. https://doi.org/10.1021/acs.jafc.9b05483
Syrpas M, Valanciene E, Augustiniene E, Malys N (2021) Valorization of Bilberry (Vaccinium myrtillus L.) Pomace by Enzyme-Assisted Extraction: Process Optimization and Comparison with Conventional Solid-Liquid Extraction. Antioxidants 10:773. https://doi.org/10.3390/antiox10050773
Teixeira RSS, da Silva AS, Ferreira-Leitão VS, Bon EP (2012) da S Amino acids interference on the quantification of reducing sugars by the 3,5-dinitrosalicylic acid assay mislead carbohydrase activity measurements. Carbohydrate Research 363:33–37. https://doi.org/10.1016/j.carres.2012.09.024
Tomassi E, Arouna N, Caruso MG, Girgenti A, Picone P, Nuzzo D, Pucci L (2025) Fermentation of Chlorella vulgaris and Aphanizomenon flos-aquae biomass improves the antioxidant profile. LWT 215:117183. https://doi.org/10.1016/j.lwt.2024.117183
Verni M, Verardo V, Rizzello CG (2019) How Fermentation Affects the Antioxidant Properties of Cereals and Legumes. Foods 8:362. https://doi.org/10.3390/foods8090362
Wang C, Onyeaka H, Miri T, Soltani F (2024) Chlorella vulgaris as a food substitute: Applications and benefits in the food industry. J Food Sci 89:8231–8247. https://doi.org/10.1111/1750-3841.17529
Xia D, Qiu W, Wang X, Liu J (2021) Recent Advancements and Future Perspectives of Microalgae-Derived Pharmaceuticals. Mar Drugs 19:703. https://doi.org/10.3390/md19120703
Yaghoubzadeh Z, Safari R (2025) Extraction of Bioactive Peptides from Chlorella vulgaris Using Enzymatic Hydrolysis: A Green Natural Antioxidant. Int J Pept Res Ther 31:33. https://doi.org/10.1007/s10989-025-10692-4
Yang J, Cai D, Liu X, Zhu L, Zhang C, Peng Q, Han Y, Liu G, Yang M (2023) Glucose Conversion for Biobutanol Production from Fresh Chlorella sorokiniana via Direct Enzymatic Hydrolysis. Fermentation 9:284. https://doi.org/10.3390/fermentation9030284
Yin L-J, Jiang S-T, Pon S-H, Lin H-H (2010) Hydrolysis of Chlorella by Cellulomonas sp. YJ5 Cellulases and Its Biofunctional Properties. J Food Sci 75:H317–H323. https://doi.org/10.1111/j.1750-3841.2010.01867.x
Yuan Q, Li H, Wei Z, Lv K, Gao C, Liu Y, Zhao L (2020) Isolation, structures and biological activities of polysaccharides from Chlorella: A review. Int J Biol Macromol 163:2199–2209. https://doi.org/10.1016/j.ijbiomac.2020.09.080
Zhang Y, Vadlani PV, Kumar A, Hardwidge PR, Govind R, Tanaka T, Kondo A (2016) Enhanced D-lactic acid production from renewable resources using engineered Lactobacillus plantarum. Appl Microbiol Biotechnol 100:279–288. https://doi.org/10.1007/s00253-015-7016-0
Zhou N, Zhang Y, Wu X, Gong X, Wang Q (2011) Hydrolysis of Chlorella biomass for fermentable sugars in the presence of HCl and MgCl2. Bioresour Technol 102:10158–10161. https://doi.org/10.1016/j.biortech.2011.08.051

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Enhancing bioactivity of Chlorella vulgaris through enzymatic pretreatment and lactic acid fermentation

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Abstract

Figures

Introduction

Materials and Methods

Chemicals and algal biomass

Ultrasound pretreatment

Autoclave pretreatment

Enzymatic pretreatment

Preparation of bacterial inoculum

Microbial growth and pH analysis

Determination of reducing sugar content

Determination of protein content

Determination of organic acids

Total phenolic content (TPC) by Folin-Ciocalteu’s assay

Cupric reducing antioxidant capacity assay

Determination of the ABTS•+ scavenging assay

Determination of the DPPH scavenging assay

Determination of antimicrobial activity

Statistical Analysis

Results and Discussion

Evaluation of various pretreatment strategies

Microbial growth and acification during LAB fermentation

Analysis of sugars and organic acids

Conclusions

Declarations

References

Supplementary Files

Status:

Version 1