Protective Effects on Keratinocytes by Extracts Enriched in Polysaccharides from Limnospira platensis Grown Under Autotrophic and Mixotrophic Conditions

TL;DR

This study shows that polysaccharide extracts from a cyanobacterium, grown under mixotrophic conditions, protect skin cells and may be useful in skincare or pharmaceuticals.

Contribution

The novel finding is that mixotrophically grown L. platensis extracts show comparable or better protective effects on keratinocytes than autotrophically grown ones.

Findings

Mixo−P extract showed better antioxidant activity in OH radical scavenging compared to Auto−P extract.

Both extracts protected HaCat cells against H2O2-induced damage and reduced inflammation markers like IL-1β and IL-6.

Mixo−P extract slightly outperformed Auto−P in preventing inflammatory effects on cell viability.

Abstract

Background/Objectives: Natural polysaccharides have many bio-pharmacological effects, which make them compounds with potential in healthcare. Limnospira platensis (Spirulina), a well-known blue–green cyanobacterium with relevance in the market of nutraceuticals, produces polysaccharides with recognized antioxidant and anti-inflammatory activities. Noteworthy, the growth of the cyanobacterium biomass may be obtained in a more sustainable manner under mixotrophic conditions. In the present study, we compared the antioxidant and anti-inflammatory effects of polysaccharide-enriched extracts from the cyanobacterium cultured under autotrophism (Auto−P extract) or mixotrophism (Mixo−P extract); this latter was realized using medium added with brewery wastewater (BWW). Methods and Results: Non-cellular investigation showed a better antioxidant profile for Mixo−P extract in the OH radical scavenging assay and a similar activity between the extracts in ABTS and ferrous chelation assays. The antioxidant protective activity of L. platensis extracts investigated on HaCat cells in the range of 0.3–10 μg/mL (not cytotoxic concentrations), against hydrogen peroxide (H2O2, 600 μM)-induced damage, revealed a similar activity by the two extracts. When tested against the inflammatory stimuli with lipopolysaccharide (LPS, 10 μg/mL) or tumor necrosis factor-α (TNF-α, 10 ng/mL), both Auto−P and Mixo−P showed an ability to prevent the effects of the inflammatory agents on cell viability and on interleukin-1β (IL-1β) and interleukin-6 (IL-6) release, with a slightly greater potency by Mixo−P extract. Conclusions: In conclusion, our data suggest the possible use of L. platensis polysaccharide-enriched extracts in biological-made pharmaceuticals for skin disorders or in cosmeceuticals. In addition, this study demonstrates that mixotrophic cultivation of L. platensis may be an alternative and sustainable way for biotechnological applications of the cyanobacterium biomass.

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1. Introduction

Natural polysaccharides, essential biopolymers synthesized by almost all living organisms, have a wide spectrum of bioactivities that suggest possible biotechnological applications in the pharmaceutical, nutraceutical, and cosmetic fields. Polysaccharides are present in many edible organisms (such as plants, algae, animals, and fungi), and, being poorly adsorbed after oral ingestion, exert their health effects mainly via the interaction with intestinal cells or with the microbiota [1]. However, after parenteral administration (i.e., subcutaneous and intravenous), which allows adequate bioavailability and then systemic effects. Or, when topically delivered, they may be promising candidates in different biomedical fields [2]. Indeed, natural polysaccharides exhibit wound healing, as well as antioxidant and anti-inflammatory properties, showcasing their potential in skincare and cutaneous disease treatment [3,4]. As a source of natural polysaccharides, marine organisms and, in particular, algae (seaweeds and microalgae) and cyanobacteria, received great attention [5,6,7]. Since they are a sustainable source of many value-added compounds with health impact, and they are easily cultivated under different growth conditions [8,9,10]. The search for alternative and more economically culture conditions that provide enriched metabolic profile for skin care is an exciting challenge. On this regard, an innovative opportunity is represented by the culture of microalgae/cyanobacteria in the presence of wastewater from the food industry or from fish aquaculture, addressing circular economy strategies [9,11].

Spirulina, formerly known as Arthrospira platensis, is a cyanobacterium (or blue–green microalga) with great commercial value. L. platensis has been widely studied and has been utilized historically for its nutrient rich composition as a dietary supplement [8]. Among macronutrients, proteins, lipids, and also polysaccharides were studied, looking also at new biotechnological approaches of cultivation to expand their sustainable production [9,10]. The polysaccharidic content, mainly consisting of sulfated polysaccharides, are synthesized by the cyanobacteria both as endometabolites and extracellular secreted molecules [12,13].

Today, the worldwide commercial production of L. platensis is linked to industrial approaches via two main types of culture systems, open ponds and bioreactors. Although open pond culture systems provide an economic advantage for the cultivation of L. platensis, it causes a higher level of contamination compared to bioreactor-based culture systems.

A third type of cultivation is a mix of the two methods above (hybrid systems) [14]. Recently, several alternative growth systems modulating environmental factors (light, pH, salt, temperature, wavelength) or media composition were shown to allow a more efficient and sustainable culture of the cyanobacteria [9,15]. In this context, L. platensis demonstrated recently the ability to grow under the supplementation of organic carbon source deriving from wastewaters [16]. It is worth noting that organic sources derived from food industry waste were used for the mixotrophic cultivation, with the aim to reduce environmental impact in line with the principles of circular bio-economy. In this study, the mixotrophic growth condition was optimized by exploiting the BWW as source of organic carbon. Under BWW addition to the medium, a great yield of biomass and changes in macronutrient composition were described by Russo et al. [17].

Although Spirulina-derived polysaccharides have been widely investigated for their antioxidant and anti-inflammatory activities, most studies have primarily relied on chemical radical-scavenging assays or systemic models, with limited attention to epidermal keratinocytes as a specific biological target. Furthermore, the majority of reports concern biomass obtained under conventional autotrophic cultivation, while the influence of alternative and sustainability-oriented growth conditions on polysaccharide composition and functional activity remains insufficiently explored. In particular, a comparative evaluation of polysaccharide-enriched extracts derived from autotrophic and BWW-based mixotrophic cultivation of L. platensis in skin-relevant in vitro models is currently lacking. Therefore, the relationship between cultivation strategy, compositional differences, and keratinocyte-protective efficacy has not yet been clearly established.

In light of the above considerations, the present study aims at investigating the different composition of aqueous extracts enriched in polysaccharides from L. platensis grown under autotrophic and mixotrophic conditions and their different protective activity against in vitro skin oxidative and inflammatory models. We expect to assess the potential of these extracts in the prevention or treatment of skin diseases and if the cultivation under mixotrophic condition may assure a valid alternative by the pharmacological point of view.

2. Materials and Methods

2.1. Materials

The cyanobacterial strain Limnospira platensis SAG 21.99 employed in this work was obtained in non-axenic form from the Algal Culture Collection of the University of Göttingen (Germany). All chemicals and reagents used throughout the study, including 0.5 N hydrochloric acid solution, 1-phenyl-3-methyl-5-pyrazolone (PMP), acetone, ammonium iron(II) sulfate, arabinose, ascorbic acid (VITC), barium chloride, chloroform, diethyl ether, dimethyl sulfoxide (DMSO), ethanol (95%), ferrous sulfate solution, fructose, fucose, galactose, galacturonic acid, glucose, glucuronic acid, hydrochloric acid (HCl), H_2_O_2_, Lowry protein assay kit, methanol, papain, potassium persulfate, rhamnose, ribose, salicylic acid, sodium hydroxide (NaOH), sodium sulfate, sulfuric acid (95.0–98.0%), trifluoroacetic acid (TFA), trichloroacetic acid, and xylose, were supplied by Merck Life Science S.r.l. (Milan, Italy). Solvents of chromatographic grade, namely acetonitrile, water, and formic acid for ultra-high-performance liquid chromatography (UHPLC) analyses, were purchased from Merck KGaA (Darmstadt, Germany). BWW was collected from the facility “Birrificio Emiliano”, located in Anzola (Bologna, Italy), and its principal physicochemical characteristics have been previously described by Russo et al. [17].

2.2. L. platensis Cultivation: Inoculums and Culture Media Preparation

Microalgal growth was carried out in a modified Jourdan Medium (JM), the full formulation of which has been previously reported by Cavallini et al. [18]. Cultivation was performed in 150 mL of Erlenmeyer flasks containing 50 mL of culture medium, which were inoculated with approximately 10 mL of algal culture and sealed using sterile cotton stoppers. The cultures were maintained at ambient temperature under continuous illumination provided by white fluorescent tubes (T8, 36 W, IP20; CMI, Frankfurt, Germany). Light intensity was set at a photon flux density of 50 µmol m^−2^ s^−1^, monitored using a calibrated luxmeter (HD 2302.0, Delta OHM, Padua, Italy). Prior to experimental use, the inoculum was expanded for about 7 days and harvested during the late exponential growth phase.

BWW samples were preserved at 4 °C until processing. Before being employed as a cultivation medium, the effluent was passed through glass microfiber filters (Whatman GF/CTM, 47 mm diameter; Incofar Srl, Modena, Italy) to remove particulate matter, followed by sterilization in an autoclave at 121 °C and 0.1 MPa for 20 min to guarantee aseptic conditions during microalgal growth. Although autoclaving effectively inactivates viable microorganisms, it is acknowledged that LPS from Gram-negative bacteria are relatively heat-stable. However, since L. platensis cultivation was conducted in JM under strongly alkaline conditions (pH > 9 during growth), which are intrinsically unfavorable for the survival and proliferation of most Gram-negative bacteria, the additional endotoxin generation during culture was limited. In addition, the physicochemical characteristics of the BWW used in this study, previously reported in [16], confirm that brewery effluents derived from food-grade raw materials are not typically associated with elevated heavy metal concentrations.

2.3. L. platensis Cultivation Conditions and Experimental Setup

Cultivation of L. platensis was carried out in cylindrical polyvinyl chloride (PVC) photobioreactors (PBRs) with a total volume of 600 L (60 cm external diameter and 2 m height). The reactors were supplied with filtered compressed air for aeration and operated under a 12 h light/12 h dark photoperiod. Illumination was provided by warm white LED sources (model 2835, 30 W, Aftertech, Montecavolo, Italy), delivering a photon flux density of 150 µmol m^−2^ s^−1^. Experimental runs were performed at a working volume of 500 L under continuous cultivation conditions, allowing for the evaluation of growth kinetics and biomass compositional changes using either conventional JM (CTRL) or a mixed JM–BWW formulation (JB). Medium replacement was carried out daily. Each experimental condition was maintained for 15 days and replicated three times, with cultures initiated at an initial biomass concentration of 0.3 g/L. Culture performance was monitored by measuring optical density, biomass concentration, and final dry weight over time. Additional information regarding PBR design and operational settings has been previously reported by Russo et al. [17].

2.4. L. platensis Cell Growth and Dry Weight Determination

The growth of L. platensis was followed by spectrophotometric analysis, recording the absorbance of the cultures at 680 nm with a UV–Vis spectrophotometer (ONDA V30 SCAN, ZetaLab, Padua, Italy). To convert optical density values into quantitative biomass data, a calibration curve was generated linking absorbance measurements to corresponding dry weight concentrations.

For direct determination of biomass by gravimetric analysis, aliquots of 10 mL were periodically collected from the PBRs and passed through pre-weighed glass microfiber filters (Whatman GF/C™, 55 mm diameter; Incofar Srl., Modena, Italy). Prior to use, the filters were conditioned by drying at 105 °C for 2 h in a forced-air drying oven (Model 30, Memmert GmbH, Schwabach, Germany), allowed to cool in a desiccator, and weighed to obtain the initial mass (W_1_). Following filtration, the biomass retained on the filters was dried overnight at 105 °C until a constant mass (W_2_) was achieved. The dry biomass concentration was subsequently calculated from the mass increase (W_2_ − W_1_) normalized to the volume of culture filtered (V).

2.5. Extraction, Purification, and Proximate Composition Analysis of Polysaccharide-Enriched Extracts Auto−P and Mixo−P

Biomass from L. platensis was processed through a combined ultrasound-assisted and hot-water extraction procedure, following the protocol previously described by Banti et al. [19]. For each experimental condition (autotrophic and mixotrophic growth), 1 g of dried biomass was suspended in 160 mL of distilled water and subjected to sonication in an ultrasonic bath (120 W; VWR^®^ USC 200T, Leicestershire, UK) for 1 h at 25 °C under continuous agitation. The suspension was subsequently stirred and heated at 80 °C for an additional 8 h. After centrifugation at 1220 RCF for 30 min, the supernatant was recovered and maintained at 80 °C until its volume was reduced to one-fifth of the initial volume. After that cold, 95% ethanol was added at a ratio of five volumes relative to the extract, followed by incubation at 4 °C overnight. The precipitate obtained after centrifugation (1220 RCF, 20 min) was dried and collected as a crude aqueous extract.

Each crude extract was then re-dissolved in distilled water at a ratio of 1:14 (w/v), and the pH was adjusted to 7.0 using hydrochloric acid. Proteolytic treatment was carried out by adding papain at 3% (w/w relative to crude extract) and incubating the mixture at 50 °C for 2.5 h. Enzymatic activity was subsequently terminated by boiling the solution. After that, 5% trichloroacetic acid was added, and the mixture was stored at 4 °C overnight. Following centrifugation, the supernatant was collected, and its pH was adjusted to 8.0 using concentrated ammonia. Oxidative treatment was then performed by adding H_2_O_2_ 30% to a final concentration of 5% and stirring the solution at 55 °C for 2 h. Polysaccharide-rich fractions were precipitated by the addition of five volumes of 95% ethanol and incubation at 4 °C overnight. The resulting precipitate was recovered by centrifugation (1220 RCF, 20 min), sequentially washed with absolute ethanol, acetone, and diethyl ether, and finally dried. Extraction yield was calculated as the ratio between the dry extract mass and the initial biomass weight, expressed as a percentage.

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For both Auto−P and Mixo−P extracts, total carbohydrate content was quantified using the phenol–sulfuric acid assay, while sulfate concentration was determined using the gelatin–barium method. Glucose and sodium sulfate were employed as calibration standards for carbohydrate and sulfate determinations, respectively, as previously reported [20,21].

The downstream extraction and purification procedure applied to obtain the polysaccharide-enriched fractions was designed to remove proteins, lipids, low-molecular-weight organic compounds, and potential endotoxin-associated impurities. These sequential purification steps significantly reduce the likelihood of wastewater-derived contaminants in the final preparations used for in vitro biological assays.

2.6. Monosaccharide Composition by UHPLC-DAD-HRMS/MS Analysis

2.6.1. Hydrolysis of Polysaccharide-Enriched Extracts Auto−P and Mixo−P and Derivatization Procedure

For monosaccharide analysis, 5 mg of the polysaccharide-rich fractions obtained from Auto−P and Mixo−P, were subjected to acid hydrolysis using 1 mL of 4 M TFA. The reaction mixture was incubated at 121 °C for 5 h to cleave the polysaccharide chains into their constituent monosaccharides. Following hydrolysis, methanol (1 mL) was added to facilitate removal of residual TFA, which was subsequently eliminated by vacuum evaporation at 60 °C. The resulting residue was reconstituted in 150 μL of ultrapure water, yielding a monosaccharide-containing solution suitable for subsequent derivatization and analysis.

Monosaccharide derivatization with PMP was performed following previously published protocols with minor modifications [22,23]. Briefly, 150 μL of either hydrolyzed polysaccharide samples or aqueous monosaccharide standards, comprising glucose, galactose, ribose, rhamnose, fucose, fructose, xylose, arabinose, glucuronic acid, and galacturonic acid at a concentration of 5 mmol L^−1^, were transferred into individual 2 mL centrifuge tubes. Each sample was combined with 150 μL of 0.6 M NaOH and 300 μL of 0.5 M PMP in methanol, and the reaction was carried out in a water bath at 70 °C for 100 min. After completion of derivatization, 300 μL of 0.3 M HCl were added to neutralize the reaction mixture, followed by extraction with an equal volume of chloroform to remove excess PMP. This extraction step was repeated at least three times. The aqueous phase was then centrifuged at 1220 RCF for 10 min. After that, the supernatant was collected, dried, and stored at −20 °C until chromatographic analysis.

2.6.2. LC-MS/MS Analysis

The monosaccharide profiles of polysaccharide fractions, following acid hydrolysis and derivatization with PMP, were determined by LC–MS/MS. High-resolution mass spectrometric analyses were carried out using a Q-ToF Agilent 6540 mass spectrometer (Agilent Technologies, Santa Clara, CA, USA) equipped with an electrospray ionization source operating in positive mode and coupled to an Agilent 1290 Infinity II ultra-high-performance liquid chromatography (UHPLC) system. Separation was achieved on a BEH C18 column (2.1 × 100 mm, 1.7 μm particle size; Acquity, Waters, Milford, CT, USA). The chromatographic system employed water containing 0.1% formic acid as mobile phase A and acetonitrile supplemented with 0.1% formic acid as mobile phase B, delivered at a flow rate of 0.3 mL min^−1^ with an injection volume of 5 μL. Analyses were performed under isocratic conditions, maintaining solvent B at 20% and solvent A at 80% for a total run time of 12 min.

The column oven temperature was fixed at 35 °C throughout the analysis. High-resolution mass spectra were collected using both full-scan acquisition (40.000 resolving power, maximum injection time of 220 ms) and data-dependent MS/MS experiments (17.500 resolving power, maximum injection time of 60 ms) over an m/z range of 200–1200 in positive electrospray ionization mode. Source conditions were optimized as follows: spray voltage set to 4000 V, capillary temperature maintained at 300 °C, nitrogen sheath gas at 24 arbitrary units, and auxiliary nitrogen gas at five arbitrary units. Fragmentation was achieved by higher-energy collisional dissociation (HCD) using a collision energy of 18 eV. Diode array detector (DAD) signals were acquired across the 200–600 nm spectral range, with 254 nm selected as the preferred wavelength for monitoring PMP-derivatized monosaccharides. Data acquisition and processing were performed using MassHunter software 10.0 (Agilent Technologies, Santa Clara, CA, USA).

2.7. Radical Scavenging Assays

2.7.1. ABTS Radical Scavenging Assay

The ABTS radical scavenging capacity of the polysaccharide fractions was evaluated following the protocol reported by Li et al. (2021) [24]. The ABTS radical stock solution was generated by combining 10 mL of an aqueous ABTS solution (7 mM) (Roche Diagnostics GmbH, Mannheim, Germany) with 163 μL of potassium persulfate (150 mM). The reaction mixture was allowed to stand for 16 h at 4 °C in the absence of light to ensure complete radical formation. Prior to use, the ABTS solution was diluted with ethanol until an absorbance value of 0.72 ± 0.01 at 734 nm was obtained.

For the assay, 180 μL of the ABTS radical solution were dispensed into each well of a 96-well microplate and mixed with 20 μL of aqueous extract solutions prepared in water/DMSO (4:1, v v^−1^) at concentrations ranging from 0.0078 to 1.0 mg mL^−1^. The reaction mixtures were incubated for 15 min at room temperature under dark conditions. The reduction of the ABTS radical, reflected by a decrease in absorbance, was measured at 734 nm using an EnSpire UV–Vis spectrophotometer (Thermo Fisher Scientific, Monza, Italy), with a water/DMSO (1:4, v v^−1^) mixture used as the blank. Control samples consisted of 180 μL of ABTS radical solution combined with 20 μL of water/DMSO (1:4), yielding absorbance values of approximately 0.720. Radical scavenging activity was expressed as percentage inhibition, calculated according to the equation reported below:

$$[eqn]$$

2.7.2. Hydroxyl Radical Scavenging Assay

Hydroxyl radical scavenging activity was assessed following the procedure described by Chen et al. [25], with minor modifications. In brief, 50 μL of an aqueous ferrous sulfate solution (6.0 mM) and 50 μL of a salicylic acid solution in ethanol (6.0 mM) were combined with 50 μL of extract solutions prepared in water/DMSO (4:1, v v^−1^) at concentrations ranging from 0.125 to 2.5 mg mL^−1^. The reaction was initiated by the addition of 50 μL of H_2_O_2_ (6.0 mM), after which the mixtures were incubated at 37 °C for 30 min.

Scavenging of hydroxyl radicals was evaluated by measuring the resulting absorbance at 510 nm using an EnSpire UV–Vis spectrophotometer, with a water/DMSO (1:4, v v^−1^) mixture employed as the blank. Control samples were prepared by replacing the extract solution with water/DMSO (1:4) while maintaining the same volumes of ferrous sulfate, salicylic acid, and hydrogen peroxide. The absorbance value of the control reaction was approximately 1.00. The percentage of radical inhibition was calculated using the equation previously reported. VITC was included as a reference antioxidant and used as a positive control in both radical scavenging assays.

2.7.3. Ferrous Ion Chelating Activity Assay

The iron-chelating activity of the aqueous extracts was evaluated following the protocol reported by Teixeira et al. [26], with minor modifications. The assay was carried out in ammonium acetate buffer adjusted to pH 6.7, using ammonium iron(II) sulfate dissolved in the same buffer as the source of ferrous ions. For each determination, 50 μL of extract solutions prepared in water/DMSO (4:1, v v^−1^) at concentrations ranging from 0.4 to 14 μg mL^−1^ were combined with ammonium iron(II) sulfate (final concentration 20 μM) in 96-well microplates. After incubation for 10 min, absorbance was recorded at 562 nm to account for any initial signal. Subsequently, a freshly prepared aqueous ferrozine solution (5 mM) was added to each well to reach a final concentration of 96 μM, and the reaction mixtures were further incubated at 37 °C for 10 min. The formation of the [Fe(ferrozine)3]^2+^ complex was then monitored by measuring absorbance at 562 nm. Blank samples were prepared by replacing the extract solutions with water/DMSO (4:1, v v^−1^). To eliminate any interference caused by the intrinsic absorbance of the extracts, the initial absorbance values recorded prior to ferrozine addition were subtracted from the final readings. Iron(II) chelation capacity was expressed as percentage inhibition, calculated using the equation provided below.

$$[eqn]$$

2.8. HaCaT Cell Culture and Viability Assay

The immortalized human keratinocyte cell line HaCaT (Cytion, Heidelberg, Germany) was maintained in Dulbecco’s Modified Eagle’s Medium (DMEM; Sigma-Aldrich, St. Louis, MO, USA) supplemented with 5% fetal bovine serum (FBS; Sigma-Aldrich; St. Louis, MO, USA) and a penicillin–streptomycin mixture (100 U mL^−1^ penicillin and 100 µg mL^−1^ streptomycin; Sigma-Aldrich; St. Louis, MO, USA). Cells were cultured at 37 °C in a humidified incubator with 5% CO_2_ and routinely passaged upon reaching 80–90% confluence. For assessment of cell viability, HaCaT cells were seeded into 96-well culture plates at a density of 5 × 10^3^ cells per well and allowed to attach overnight. Cells were then exposed to the polysaccharide-rich extracts obtained under autotrophic and mixotrophic conditions (Auto−P and Mixo−P) for 24 h at final concentrations of 0.3, 1, 3, 10, 30, and 100 µg mL^−1^. Cell viability was evaluated using the MTT assay [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] (Sigma-Aldrich; St. Louis, MO, USA). Following treatment, cells were incubated with MTT solution (0.5 mg mL^−1^) for 2 h at 37 °C. The culture medium was subsequently removed, and the resulting formazan crystals were solubilized in DMSO. Absorbance was measured at 540 nm, and cell viability was expressed as a percentage relative to untreated control cells.

2.9. Assessment of Antioxidant and Anti-Inflammatory Activities on HaCaT Cells

The protective potential of the extracts was investigated in HaCaT keratinocytes subjected to oxidative and inflammatory challenges using a defined treatment scheme. Cells were initially exposed to Auto−P or Mixo−P at concentrations ranging from 0.3 to 10 µg mL^−1^ for 24 h. Subsequently, the culture medium was replaced, and cells were challenged with the selected stress-inducing agents (described below) for an additional 16 h in the absence of the extracts, allowing evaluation of the pre-treatment effects. Oxidative stress was generated by the addition of H_2_O_2_ (600 µM). Inflammatory responses were triggered either by LPS derived from Escherichia coli O111:B4 (10 µg mL^−1^; Sigma-Aldrich; St. Louis, MO, USA) or by stimulation with the pro-inflammatory and mitogenic cytokine TNF-α (10 ng mL^−1^; Sigma-Aldrich; St. Louis, MO, USA).

2.9.1. DHE (Dihydroethidium) Staining for Intracellular ROS (Reactive Oxygen Species) Measurement in HaCaT Cells

Intracellular generation of the superoxide anion (O_2_•^−^) was assessed using the fluorescent probe DHE (Sigma-Aldrich; St. Louis, MO, USA). Following the H_2_O_2_ exposure described above, HaCaT cells were rinsed twice with Hank’s Balanced Salt Solution (HBSS; Sigma-Aldrich; St. Louis, MO, USA) and subsequently incubated in HBSS containing 5 µM of DHE for 30 min at 37 °C under light-protected conditions. After the staining period, cells were washed three additional times with HBSS, and fluorescence signals were quantified by spectrofluorimetric analysis using an excitation wavelength of 535 nm and an emission wavelength of 610 nm. The measured fluorescence intensity was taken as an index of ROS accumulation.

2.9.2. ELISA Assay for IL-1β and IL-6 Production

The secretion of the pro-inflammatory cytokines IL-1β and IL-6 by HaCaT cells was quantified in culture supernatants using commercially available human ELISA kits (IL-1β, IL-6, Thermo Fisher Scientific, Waltham, MA, USA) in accordance with the manufacturer’s protocols. HaCaT keratinocytes were first pre-treated with the polysaccharide-rich extracts Auto−P and Mixo−P for 24 h, followed by stimulation with either LPS or TNF-α for an additional 16 h. After completion of the treatments, cell culture supernatants were carefully harvested and subjected to cytokine quantification. Cytokine levels were measured by spectrophotometric detection at 450 nm and expressed as concentrations in picograms per milliliter (pg mL^−1^).

2.10. Data Analysis and Statistics

All experiments were conducted in triplicate and repeated independently at least three times. Each result is expressed as mean ± standard deviation (SD). Statistical analysis was performed through the software MetaboAnalysts 5.0 tuned by the McGill University, Montreal, Canada (for data regarding biomass cultivation) or by GraphPad Prism 10 (GraphPad Software, La Jolla, CA, USA). Differences among groups were evaluated by one-way analysis of variance (ANOVA) followed by Bonferroni’s post hoc test, with significance set at p < 0.05.

3. Results

3.1. L. platensis Biomass Production

The results illustrated in Figure 1 provide a comparative overview of L. platensis cultivation under photoautotrophic (JM) and mixotrophic conditions (JM + BWW), where BWW was added as an organic carbon source. Overall, the mixotrophic configuration exhibited higher final biomass concentration and specific growth rate, although the volumetric biomass productivity remained lower than JM, suggesting an incomplete optimization of process parameters. In terms of biomass concentration (Figure 1A), the JM + BWW system reached 0.377 ± 0.060 g/L, corresponding to an increase of approximately 82% compared to JM (0.207 ± 0.013 g/L). This enhancement indicates that the organic load of the BWW effectively stimulated biomass accumulation, likely due to the availability of easily assimilable carbon compounds and accessory nutrients such as amino acids, vitamins, and organic acids. Similar increases in final biomass concentration have been reported for mixotrophic cultures of Limnospira and Chlorella species using dairy or food-industry effluents with moderate organic loads [18,27,28]. Despite the higher final biomass, the average biomass productivity (Figure 1B) was lower in JM + BWW (8 ± 0.005 mg/L d) than in JM (53 ± 0.011 mg/L d). This apparent contradiction can be attributed to several interacting factors. The darker color and turbidity of the BWW may have reduced light penetration, limiting photosynthetic efficiency, particularly at higher cell densities. Moreover, the rapid consumption of the available organic carbon might have led to early saturation of growth or nutrient depletion, resulting in a lower average productivity over the cultivation period. Similar effects have been observed in mixotrophic systems where excessive organic carbon input or poor light transmission constrains the long-term productivity despite faster initial growth [29,30]. The specific growth rate (Figure 1C) revealed the most pronounced difference between systems, increasing from 0.060 ± 0.010 1/d in JM to 0.183 ± 0.035 1/d in JM + BWW. This nearly three-fold increase highlights a kinetic advantage during the exponential phase of the mixotrophic culture. Such acceleration can be explained by the co-utilization of inorganic (HCO_3_^−^/CO_2_) and organic carbon sources, providing enhanced ATP and reducing power for biosynthetic pathways and promoting rapid cell division. This trend is consistent with previous findings for mixotrophic cultures of L. platensis and Chlorella vulgaris cultivated with agro-industrial residues [27,28]. Taken together, the results suggest that the addition of BWW enhances the metabolic flexibility of L. platensis, supporting faster initial growth (higher µ) and greater final biomass (higher X), but that process conditions, with particular emphasis on light intensity, optical depth, and organic load, require optimization to translate these kinetic benefits into sustained volumetric productivity. Similar patterns have been reported in other mixotrophic systems, where insufficient dilution or excessive turbidity of the effluent limited long-term biomass productivity despite transient kinetic advantages. Future studies should therefore focus on optimizing BWW pre-treatment, dilution, and feeding strategy to fully exploit its potential as a sustainable feedstock for circular bioeconomy-based microalgal production.

3.2. L. platensis-Derived Extracts

3.2.1. Extraction and Chemical Analysis of Auto−P and Mixo−P Extracts

In the present study, the aqueous extracts enriched in polysaccharides, Auto−P and Mixo−P, were obtained by hot water extraction combined with ultrasound, followed by deproteinization and decolorization treatment. Based on the dried powder the yield for Auto−P was only slightly and not significantly higher than that for Mixo−P. On the contrary, the sulfate content was significantly greater in the Mixo−P extract (see values in Table 1).

3.2.2. Monosaccharide Composition

The identification of sugar components of the extracts Auto−P and Mixo−P was carried out by comparison of their retention time (tR), UV absorption, full mass spectra with the literature data [31], and data obtained by injection of monosaccharide standards after derivatization with PMP in the same conditions. A mass error <5 ppm was considered for compound annotation. The LC-MS/MS chromatogram recorded in positive ionization mode is shown in Figure 2, while chromatographic and spectral data in +ESI mode are reported in Table 2. The obtained data for both aqueous polysaccharide-enriched extracts, Auto−P and Mixo−P, allowed us to identify unambiguously the presence of ribose (peak 1), rhamnose (peak 1), glucuronic acid (peak 3), arabinose (peak 3), xylose (peak 4), and fucose, (peak 5), as confirmed by chromatographic and MS data of pure injected standards. The broad peak at 7.5 min and 7.6 min for Auto−P and Mixo−P, respectively, was attributed to hexose-PMP, but it was not possible to discriminate among glucose, galactose, and fructose due to the coelution of the three monosaccharide-PMP derivatives having very close t_R_ values and identical mass spectral data. Furthermore, the injection of galacturonic acid -PMP as a reference pure compound led to exclusion of the presence of this monosaccharide.

Based on these results, the monosaccharide components of Auto−P and Mixo−P were very similar for the qualitative profile, and xylose was the most abundant monosaccharide. Moreover, the presence of all the monosaccharides used as standards was confirmed, with the exception of galacturonic acid. Finally, some differences in the relative abundance between the monosaccharides in the two types of extracts were observed, as shown in Table 2 (last column).

3.3. Antioxidant Activities of L. platensis Extracts

3.3.1. Radical Scavenging Activity and Ferrous Ion Chelating Activity

The activity as antioxidants was studied by directly binding (scavenging effect) with free radicals, i.e., ABTS and hydroxyl [32,33], and by chelating transition metals, i.e., iron, which usually catalyze the synthesis of free radicals [34].

The radical scavenging activity of Auto−P and Mixo−P extracts is shown in Figure 3A,B. The ABTS scavenging ability of both extracts improved as the concentration increased, with EC_50_ values (concentration required to obtain a 50% radical scavenging effect) of 166.6 ± 0.98 μg/mL for Auto−P and 150.0 ± 1.32 μg/mL for Mixo−P. Similar to ABTS radical scavenging activity, the hydroxyl radical scavenging ability of both extracts improved with their concentration, with EC_50_ values of 685.6 ± 0.98 μg/mL for Auto−P and 428.2 ± 1.06 μg/mL for Mixo−P. In both cases, the radical scavenging activities of Auto−P and Mixo−P were significantly lower than those of VITC. The results demonstrated higher activity in the ABTS radical scavenging assay than in the hydroxyl one, according to the data reported in the literature analysing polysaccharides from L. platensis [35,36]. Furthermore, in both the ABTS and hydroxyl radical assay, Mixo−P showed a better radical scavenging abilities than Auto−P. This result could be due to the higher sulfate content in Mixo−P compared to that present in Auto−P (13.6 ± 1.1 vs. 6.5 ± 0.32, Table 1). In fact, as reported in the literature, the antioxidant activity of microalgae-derived polysaccharides is related not only to their sugar composition, molecular weight, and treatment process, but also to their degree of sulfation [34].

Regarding the ferrous ion chelating capacity, both Auto−P and Mixo−P extracts (Figure 3C) showed an activity that improved with their concentration, with EC_50_ values of 4.823 ± 0.12 µg/mL and 3.659 ± 0.25 µg/mL for Auto−P and Mixo−P, respectively. Similarly to that observed for the radical scavenging activity, Mixo−P showed better activity than Auto−P. As reported in the literature, the presence of uronic acid and sulfate groups appeared to be critical for the polysaccharide chelating activity [34]. Therefore, the better activity of Mixo−P compared to that of Auto−P could be due to the higher sulfate content of Mixo−P compared to that of Auto−P (Table 1). Instead, it was not possible to correlate the activity to the presence of glucuronic acid due to the coelution of this monosaccharide with arabinose (Table 2).

3.3.2. Antioxidant Effects of L. platensis Extracts in HaCaT Cells

The extracts Auto−P and Mixo−P were investigated for their antioxidant and anti-inflammatory activities on keratinocytes (HaCaT cells) in the concentration range of 0.3–10 µg/mL. This range was chosen after observing in a preliminary test, at 24 h of treatment, its absence of effects on HaCaT cell viability. A significant cytotoxicity was, in fact, evident only for concentrations above 10 µg/mL for both the extracts (Figure 4).

The H_2_O_2_ treatment at 600 µM was able to significantly decrease viability of HaCaT keratinocytes by about 25% (Figure 5A,B). As shown in Figure 5A,B, both extracts Auto−P and Mixo−P revealed a protective effect on viability at the highest concentrations used (3 and 10 µg/mL).

When intracellular ROS levels were investigated, using the DHE staining test, about 80% increase of the H_2_O_2_-induced oxidative condition was observed, as well as a significant decrease of ROS levels at 3 and 10 µg/mL, by both Auto−P and Mixo−P extracts (Figure 5C,D).

The effects of the extracts on ROS production were in agreement with the results from cell viability experiments, confirming a very similar protective effect between the two extracts.

3.4. Protective Effects of L. platensis Extracts Against LPS in HaCaT Cells

The potential anti-inflammatory effects of the extracts Auto−P and Mixo−P were firstly investigated against the classical pro-inflammatory agent derived from the gram-negative bacterial wall, the LPS (10 µg/mL). As shown in Figure 6A,B, HaCaT cells suffered a significant reduction in viability after exposure to LPS (by about 30%).

This reduction in viability was significantly prevented when cells were pre-treated for 24 h with Auto−P or Mixo−P prior to LPS exposure. Both extracts exerted a comparable protective effect, although Mixo−P was the only extract showing significant activity at the lowest tested concentration (0.3 µg/mL).

LPS stimulation markedly increased IL-1β and IL-6 release from basal levels of 11.57 ± 3.79 pg/mL and 10.03 ± 1.59 pg/mL to 32.65 ± 1.98 pg/mL and 77.95 ± 3.22 pg/mL, respectively (mean ± SD).

Pre-treatment with both extracts significantly attenuated cytokine production. At 10 µg/mL, Mixo−P reduced IL-1β levels to 17.34 ± 2.85 pg/mL compared with 19.25 ± 3.34 pg/mL for Auto−P, and decreased IL-6 to 23.22 ± 2.39 pg/mL versus 25.90 ± 2.37 pg/mL for Auto−P. A consistent trend was observed at 3 µg/mL, where Mixo−P further lowered IL-1β (22.48 ± 2.63 pg/mL vs. 23.54 ± 2.70 pg/mL for Auto−P) and IL-6 (33.25 ± 3.71 pg/mL vs. 35.98 ± 3.71 pg/mL for Auto−P). The complete quantitative dataset for all treatment conditions is reported in Table 3.

3.5. Protective Effects of L. platensis Extracts Against TNF-α in HaCaT Cells

When the endogenous inflammatory cytokine TNF-α was administered to HaCaT cells, a significant increase in cell viability (approximately 40%) was observed (Figure 7A,B), consistent with previous reports describing TNF-α-induced proliferation of HaCaT keratinocytes in psoriasis-related in vitro models [37].

Both extracts significantly prevented the TNF-α-induced increase in cell viability (Figure 7C,D). Mixo−P, also in this case, was slightly more potent than Auto−P, becoming significantly effective at 1 µg/mL.

TNF-α stimulation significantly increased IL-1β and IL-6 release from basal levels of 11.57 ± 3.79 pg/mL and 10.03 ± 0.80 pg/mL to 36.03 ± 2.43 pg/mL and 32.67 ± 2.10 pg/mL, respectively (mean ± SD). Pre-treatment with both extracts markedly attenuated cytokine production. At 10 µg/mL, Mixo−P reduced IL-1β levels to 20.26 ± 2.35 pg/mL compared with 22.90 ± 2.34 pg/mL for Auto−P, and decreased IL-6 to 12.32 ± 1.87 pg/mL versus 15.04 ± 1.83 pg/mL for Auto−P. A similar trend was observed at lower concentrations, with Mixo−P consistently demonstrating greater efficacy in reducing TNF-α-induced cytokine release. The complete quantitative data for all treatment conditions are summarized in Table 4.

4. Discussion

Compounds of natural origin with antioxidant and anti-inflammatory properties represent valuable candidates for dermatological and cosmetic applications targeting mechanisms involved in skin ageing or inflammageing conditions. Excessive production of ROS, together with the enhanced release of pro-inflammatory interleukins such as IL-1β and IL-6, plays a pivotal role in disrupting cellular balance and skin function. Natural bioactive molecules capable of modulating both oxidative stress and interleukin-driven inflammatory responses are, therefore, of significant interest for the development of protective and preventive skin strategies.

In this perspective, organisms such as cyanobacteria have shown great potential, offering the opportunity to use their metabolites as innovative ingredients in cosmeceuticals or as new pharmacological agents.

Among cyanobacteria, L. platensis is one of the most present on the market providing food supplements/nutraceuticals [38]. It has shown health benefits among which the antioxidant and the anti-inflammatory potentials are largely recognized from preclinical studies and a few clinical trials [32,39].

These properties, although prevalently correlated with C-phycocyanin, polyphenol, and polyunsaturated fatty acids (PUFA) omega-3 content, have been also attributed to the cyanobacterium polysaccharidic component [40,41], which represents the 15 to 20% in weight of the whole biomass and consists mostly of acidic heteropolysaccharides [10].

Recently, L. platensis was demonstrated to be cultivable under BWW as source of organic carbon (mixotrophic conditions), reaching a greater yield in terms of biomass, compared to the autotrophic cultivation, and some advantage in value-added compounds, such as an increase in C-phycocyanin and PUFA content [17]. In the same study, Russo et al. [17], as far as the total monosaccharides amount, did not observe a difference between the biomasses grown in the autotrophic or mixotrophic condition. A similar conclusion could be drawn for the polysaccharides content in our study, although some differences were evidenced in sulfated polysaccharides and in single or associated monosaccharides levels that can justify little differences in the biological effects (mainly in extract potency) investigated.

Natural polysaccharides are known to have antioxidant activity, since they can regulate ROS production, protect cells through free radical scavenging activity, and regulate the signal pathways and specific enzymes involved in antioxidation [42].

For the first time in our study, the hydrophilic polysaccharide extract from L. platensis, grown under BWW-mediated mixotrophic conditions, was observed to counteract the damage induced by oxidative stress in keratinocytes in the same way as the extract derived from the autotrophic growth condition.

The antioxidant effect was observed with the polysaccharide-enriched extracts at the greatest concentrations used. The antioxidant effects on the cellular assays confirmed previous results in our study, with non-cellular assays revealing the ability of both Auto−P and Mixo−P to exhibit scavenging activity against different radicals. Moreover, a better activity was observed with the Mixo−P extract in radical scavenging effects. This result might be due to a higher level of sulfated polysaccharides in Mixo−P when compared to Auto−P extract (see Table 1). In fact, the sulfate groups are able to increase the hydrogen release capacity of the anomeric carbon, increasing the hydrogen supply capacity of the polysaccharides and, therefore, enhancing their radical scavenging activity [35]. In addition, sulfate groups produce an acidic environment that improves the electrostatic trapping of free radicals [43]. This difference between the two extracts was not observed when their antioxidant effects were investigated on cells, studying either viability or ROS production directly. This may be the consequence of the different concentrations used in cell-based assays, chosen for avoiding cell toxicity by the extracts, although it may be due also to the biochemical complexity of cells possibly responsible for compensatory mechanisms able to mask the difference between the effects by Auto−P and Mixo−P extracts.

In a previous paper, we reported a greater antioxidant effect by another L. platensis polysaccharide-enriched extract on cochlear cells [19]. This greater effect may have some possible causes, which could be due to a different L. platensis commercial source and to different experimental conditions, i.e., the damaging condition (H_2_O_2_ vs. cisplatin as ROS enhancer) and different cells (keratinocytes vs. cochlear cells). The difference between the experimental settings in the two studies was observed also by the toxicity concentration threshold to L. platensis polysaccharides (10 µg/mL in keratinocytes vs. 80 µg/mL in cochlear cells).

In addition to the scavenging properties, L. platensis polysaccharides may protect skin through an upregulation of the enzyme SOD-2, via the Nrf2 pathway, as suggested very recently by Han et al. [44], who studied the effects of Spirulina polysaccharides against UVA/B-induced damage on mice skin. Nrf-2/HO-1/SOD2 is, indeed, a crucial pathway activated to defend cells against oxidative stress [45].

A restoration of mitochondrial activity and collagen production by L. platensis polysaccharides enhancing SOD-2 in fibroblasts has been previously reported by other authors [46]. In the present study, the low intensity in the antioxidant effect observed in HaCaT cells may be linked to a different response of keratinocytes in comparison with fibroblasts, suggesting these latter cells are likely more efficient in enhancing the expression of SOD-2.

Among the health-promoting activities described for natural polysaccharides, there is the anti-inflammatory one [47], described also in a few studies on L. platensis [40,44,48]. When we studied the anti-inflammatory protection of L. platensis polysaccharidic extracts, a greater activity by the two extracts in comparison with the antioxidant protection was observed, both when the inflammatory agent was LPS and when the cells were exposed to the action of TNF-α. In addition, a better potency by the Mixo−P extract in comparison with the Auto−P was evident. Structural characteristics and modifications of Spirulina polysaccharides may be responsible for different biological activities [41]. In addition to the molecular weight (Mw), which may interfere with the ability in membrane penetration and then in intracellular effects [49], also the monosaccharides composition and the sulfate content are a base for different activities that may be observed among polysaccharide-enriched preparations [41]. The extracts Auto−P and Mixo−P have differences both in sulfate content and in the relative percentual content of some monosaccharides, which could explain their similar but not identical potency.

In our study, protection against inflammation was revealed by the inhibition of both LPS and of TNF-α-induced changes in HaCaT cell viability and cytokine release. LPS is a standard inflammatory agent induced in skin as in other tissues, with several effects involving the activation of the Toll-like receptor (TLR) family members [50] and the promotion of the NF-kB pathway and cytokine release [51]. The inhibitory effects of natural polysaccharides, particularly sulfated polysaccharides, against LPS-induced tissue responses are well documented [52]. Moreover, in some papers, the interaction and blocking activity of polysaccharides and oligosaccharides with TLR-4 and other TLR receptors has been suggested [51,53].

Our data support the anti-LPS activity of L. platensis polysaccharides in this in vitro keratinocyte model, suggesting that polysaccharide-enriched extracts obtained under mixotrophic conditions may represent promising candidates for further investigation.

The protective potential of L. platensis polysaccharide-enriched extracts was evident also against TNF-α, at a concentration of this agent increasing HaCaT cell viability and releasing other inflammatory cytokines. TNF-α is known to promote cell proliferation participating in wound-healing processes and produce an inflammatory condition coupled to mitogenic effects in some pathological cutaneous processes [54]. For example, this molecule contributes to keratinocytes’ hyperproliferation in psoriasis, promoting the activation of the PI3K/AKT pathway [55,56]. So, the protection by L. platensis polysaccharides against TNF-α in the present study may be linked to their ability to inhibit the PI3K/AKT pathway, as already reported for other polysaccharides in cancer cells [57,58].

The present study presents some limitations that should be considered. The biological effects were assessed in a two-dimensional in vitro HaCaT model, which, although widely used, does not fully replicate the complexity of human skin. In addition, the molecular mechanisms underlying the observed effects were not directly explored. Further studies employing complementary experimental models would help to better define the mechanistic aspects and potential applicability of these findings.

5. Conclusions

In conclusion, the present study gives evidence for an important effect of crude polysaccharidic extracts from L. platensis in the control of pro-oxidant and inflammatory conditions of a cellular component of the skin epidermis, i.e., keratinocytes. These findings suggest the possible use of L. platensis polysaccharides in biological-made pharmaceuticals or in cosmeceutical products for prevention or treatment of skin disorders.

Moreover, our data show that a mixotrophic cultivation of the cyanobacterium may represent an alternative and sustainable way for the biotechnological application of L. platensis biomass in healthcare products.