Value-added co-products from biomass of the diatoms Staurosirella pinnata and Phaeodactylum tricornutum

Algal Research 47 (2020) 101830

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Value-added co-products from biomass of the diatoms Staurosirella pinnata and Phaeodactylum tricornutum

T

Saverio Savioa, Serena Farrottia, Debora Parisb, Esther Arnaìzc, Israel Díazc, Silvia Boladoc, ⁎ Raul Muñozc, Carlo Rodolfod, , Roberta Congestria Laboratory of Biology of Algae, Department of Biology, University of Rome ‘Tor Vergata’, Via Cracovia 1, 00133 Rome, Italy Institute of Biomolecular Chemistry (ICB) of National Research Council (CNR), Comprensorio Olivetti, Via Campi Flegrei 34, 80078 Pozzuoli, Italy c Institute of Sustainable Processes, Department of Chemical Engineering and Environmental Technology, University of Valladolid, Dr. Mergelina s/n, 47002 Valladolid, Spain d Laboratory of Cell Biology, Department of Biology, University of Rome Tor Vergata, Via della Ricerca Scientifica s.n.c., 00133 Rome, Italy a

b

A R T I C LE I N FO

A B S T R A C T

Keywords: Diatom biomass Multiple extractions Bioactivity Biomethane PUFAs

Microalgae bioprospecting indicates diatoms as a promising resource for biotechnology, with multiple extractions, and intermediate valorization protocols able to open new paths in biomedical, food and feed, and bioenergy fields. The biomass of two diatoms, Phaeodactylum tricornutum and Staurosirella pinnata, was sequentially extracted to obtain crude extracts, cellular lipids, and biomethane by means of compatible protocols. Hydrophilic fractions of crude extracts were characterized for their bioactivity on human melanoma (CHL-1) and keratinocytes (HaCaT) cell lines, with S. pinnata extract showing efficient anti-proliferative and cell death inducing activities only on CHL-1. Total lipids were extracted from residual biomass, and their chromatographic profiles evidenced percental amounts of eicosapentaenoic, hexadecenoic, and octadecanoic acid, exploitable in food and feed sectors. Finally, exhausted biomass was used for biomethane production, with P. tricornutum showing the highest rate. Cascade extraction from diatom biomass prospected potential to optimize the production of algal chemicals and their further biotechnological application.

1. Introduction In recent years, bioprospecting fast growing microalgae, able to produce high amounts of added value bio-products, raised special interest in the large-scale production of biomass, including research and development on novel strains, culture systems, and methods. Microalgae-based technologies include the production of novel food and feed, nutraceuticals and pharmaceuticals, and can improve the sustainability of existing industrial activities, as agriculture, wastewater treatment [1], and renewable fuel production [2]. These applications rely on both the exploitation of whole algal biomass and on the extraction of target products from wet or, more frequently, dried biomass. Microalgal biomass, as a whole, is mainly related to human nutrition, with Chlorella and Spirulina regarded as functional/health foods, as well as to animal feed production, also as ingredients, thanks to the biomass biochemical composition, and the

high protein, carbohydrate, and lipid contents [3]. Aquafeed is a primary sector of microalgal biomass exploitation, for the nutrition of larvae and early stages of molluscs, shrimps, and fishes in aquaculture systems [4]. A number of reports also exists on the beneficial effect of microalgal supplementation into the diet of cattle and domestic animals [5]. Novel agriculture operations can also benefit from microalgal biomass, due to the presence of interesting metabolites acting as biostimulants, growth enhancers, bio-fertilizer, and bio-pesticides [6,7]. In addition, whole microalgal biomass can be used for the production of liquid or gaseous bioenergy carriers. Finally, microalgal biomass represents a source of a wealth of biomolecules, spanning from bioactive pigments, as astaxanthin, beta-carotene, and phycobiliproteins, to polyunsaturated fatty acids (PUFAs), and essential amino acids and vitamins [8]. The recent development in downstream biomass procedures are focused on obtaining a variety of the above value-added products

⁎ Corresponding author at: Laboratory of Cell Biology, Department of Biology, University of Rome Tor Vergata, Via della Ricerca Scientifica s.n.c., 00133 Rome, Italy. E-mail addresses: [email protected] (S. Savio), [email protected] (S. Farrotti), [email protected] (D. Paris), [email protected] (E. Arnaìz), [email protected] (I. Díaz), [email protected] (S. Bolado), [email protected] (R. Muñoz), [email protected] (C. Rodolfo), [email protected] (R. Congestri).

https://doi.org/10.1016/j.algal.2020.101830 Received 17 June 2019; Received in revised form 3 February 2020; Accepted 4 February 2020 2211-9264/ © 2020 Elsevier B.V. All rights reserved.

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starting from the same biomass. This goal is reached by means of compatible protocols, with an increasing integration of ‘green’ methods for the extraction/conversion of raw material into multiple co-products, or commodities [9,10]. This approach, known as algal biorefinery, allows for the valorisation of wastes and intermediates of biomass conversion, thus ameliorating economic and environmental sustainability of these processes [11]. In this context, diatoms represent a promising group of microalgae, as they: thrive in every aquatic systems, both natural and artificial; are adaptable to a wide range of environmental conditions [12,13]; rapidly grow in laboratory, and in larger scale systems; their growth rates can be easily controlled, by modulating the availability of silicate [14]; the derived biomass is rich in valuable compounds, exploitable for different biotechnological practices. In fact, diatoms accumulate lipids, as carbon storage and for cell buoyancy, up to 25% (45% under nutrient depletion) of their dry weight. Their fatty acid profile revealed that eicosapentaenoic acid (EPA) is the most common PUFAs [15,16] and diatom oil represents a promising resource for human nutrition [5]. Furthermore, diatom dominant xanthophyll, fucoxanthin, active in light harvesting and energy transfer, showed important anti-proliferative properties against cancer [17,18], antioxidant [19], anti-diabetic and antiinflammatory potential [20]. Finally, diatoms produce frustules, siliceous rigid cell walls, finely perforated by nano-scale pores in highly ordered patterns, which confer interesting optical properties, and photonic crystal behavior. Frustules protect cells from grazing and pathogen attack, and, thanks to their photonic properties, manipulate incident light to protect cells from harmful light radiations [21,22]. As a whole, frustule properties showed potential for biotechnological exploitation in the field of random lasing, dye trapping [23,24], micromechanics device production [25], and biomedical application [26]. Application of a biorefinery approach to intensively cultivated diatom biomass is still scant, with reports of a stepwise microwave assisted extraction protocol, applied to the model diatom Phaeodactylum tricornutum, to obtain fucoxanthin, eicosapentaenoic acid (EPA), and chrysolaminarin [27]. The interest for the valorisation of the whole set of products obtainable from a single microalgal extract is increasing worldwide, as different fractions can be exploited for diverse applications and the hydro-soluble ones may contain molecules, peptides, and proteins with important properties such as antioxidant, anticancer, and antimicrobial activity [28]. In this work we develop a lab-scale cascade extraction procedure, to isolate different value-added co-products, starting from a single biomass of two intensively cultivated diatoms strains: the native fieldisolated diatom Staurosirella pinnata (Ehrenberg) D.M. Williams & Round, and the well-studied and ‘domesticated’ model organism Phaeodactylum tricornutum Bohlin. Our cascading extraction was developed to obtain different fractions starting from crude extract small metabolites, through biological molecules, such as lipids, and ending up with biomethane, from the exhausted biomass. The different co-products were characterized and tested for potential applications in the biomedical and bioenergy fields, showing the possibility of obtaining high value products from cascade extraction of the same biomass, and prospecting their potential as bio-actives in the nutraceutical and energy sectors.

Fig. 1. Cascade extraction protocol. Outline of the extraction steps and obtained product fractions.

was pneumatically stirred to optimize biomass production. Biomass accumulation was monitored every 48 h by measuring both the optical density (OD), of 1 ml culture samples, at 665 nm and 750 nm, with a Onda UV-30 Scan spectrophotometer, and the dry weight, after desiccation in a ventilated oven (Falc STF-N 80), of 5 ml culture samples. In order to evaluate the possible presence of contaminants, culture samples were analysed by light microscopy (Zeiss Axioskop, 40×), over the whole growth experiment. The biomass was harvested at the stationary phase, by settling and centrifugation (2.200 ×g, 5 min), freezedried (Edwards SO4), and stored at −80 °C. Biomass yields reached 3.00 ± 0.01 g/l, fresh weight and 0.210 ± 0.01 g/l, dry weight. The daily productivity was 14.00 ± 0.1 mg/l/day. Phaeodactylum tricornutum Bohlin biomass was purchased as lyophilized material from Phytobloom Necton (http://phytobloom.com/ index.php). 2.2. Biomass cascade extractions In the cascade extraction procedure, outlined in Fig. 1, the freezedried biomass of the two studied diatoms was first manually grinded and then extracted with methanol, in order to obtain a crude extract. Subsequently, the remaining biomass (residual biomass) was treated with chloroform and methanol, to extract total lipids, and finally, the “leftover” material (exhausted biomass), was anaerobically digested to evaluate the Biochemical Methane Potential (BMP). Each extraction step was also conducted on untreated material for yield comparisons. 2.2.1. Crude extract To obtain the crude extract 1 g of freeze-dried biomass was manually grinded, suspended in 20 ml methanol aqueous solution (20% v/v), and incubated for 2 h at 45 °C, in a thermostatic water bath. The solution was centrifuged (2.200 ×g, 10 min), the supernatant collected, and separated into hydrophilic and lipophilic fractions, by addition of 0.05 volumes of CHCl3, gentle mixing, and centrifuging (10.000 ×g, 10 min). Solvents were vacuum evaporated (Buchi Rotavapor WaterBath B-480), the two fractions lyophilized, and stored at −80 °C. The choice of this procedure was based on the current literature, reporting the extraction of very small metabolites from microalgae. We focused our attention on the hydrophilic fraction, which was characterized by means of NMR analysis and its bioactivity tested in cell assays.

2. Materials & methods 2.1. Biomass The strain of the colonial diatom VRUC 290 Staurosirella pinnata (Ehrenberg) D.M. Williams & Round was isolated from sediments of a Mediterranean lagoon (Cabras, Sardinia, Italy, [23]). The stock culture was maintained in Diatom Medium (DM) and was used as inoculum for the mass cultivation in an indoor 30 l photobioreactor (25 °C, irradiance of 80 μmol photons m−2 s−1 and 12:12 h Light/Dark cycle), the culture

2.2.2. Total lipid extract Total lipids extraction was performed on the residual biomass, resulting from the aqueous methanol extraction step, following the method of Bligh and Dyer [29]. Residual biomass was grinded manually for 10 min, then suspended in 2 ml of chloroform:methanol (2:1 v/v), 2

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iodide solution (50 μg/ml propidium iodide/0.1% Triton X-100/0.1% Na-citrate in PBS), and incubated overnight at 4 °C in the dark. Samples were then acquired by means of a FACSCalibur (BD) flow cytometer and collected data analysed with the FlowJo software (TreeStar). Statistical analysis was performed by means of two-way ANOVA, on GraphPad Prism (GraphPad Software, USA).

and centrifuged (2.200 ×g, 3 min). The supernatant was saved, and the pellet washed by repeated suspension in 1 ml of chloroform:methanol (2:1 v/v) and centrifuging (2.200 ×g, 3 min), till it turned whitish. The pooled supernatants were treated with 0.1 M HCl/0.5% MgCl2 and centrifuged (2.200 ×g, 3 min), in order to separate the proteins from the total lipids. The lower phase, containing lipids, was then recovered, dried by solvent evaporation, and the lipid content calculated as percentage of dry weight. For the fatty acid profile determination, 100 mg of the dried lipid extract was derivatized and qualitatively characterized using a Gas Chromatography equipped with a Flame Ionization Detector (GC-FID).

2.3.3. Profiling of lipid extracts The derivatization of lipid extracts was performed using a CH3OH:H2SO4 solution (15:1 v/v), at 60 °C, for 6 h, in order to obtain Fatty Acid Methyl Esters (FAMEs) by transesterification. FAMEs present in the extracts were identified by comparing the retention times of the peaks in the sample with those of an analytical commercial standard, a mixture of 37 FAMEs, purchased from Supelco (Sigma-Aldrich, Merck group, Germany). Qualitative characterization of FAMEs was performed on an Agilent 7890A (Agilent Technologies, California, USA) GC-FID instrument, equipped with a DB-Wax column (30 m × 0.25 mm, i.d. 0.25 μm). The carrier gas used was helium (1 ml/min), and the temperature program was as follows: the column was held at 50 °C for 1 min, ramped to 200 °C, at 25 °C/min, increased to 230 °C, at 3 °C/min, and then held for 18 min. FID detector temperature was fixed at 280 °C. Helium (99.999 purity) was purchased from Abello Linde S.A. (Barcelona, Spain).

2.3. Characterization of biomass multiple products 2.3.1. Characterization of hydrophilic fractions by nuclear magnetic resonance The metabolic content of the hydrophilic fractions was analysed by means of Nuclear Magnetic Resonance (NMR) spectroscopy. 630 μl of each hydrophilic fraction were supplemented with 70 μl of a D2O standard solution (1 mM sodium 3-trimethylsilyl (2,2,3,3-2H4) propionate (TSP)), providing a field frequency lock for the spectrometer, and rapidly transferred to an NMR tub, for metabolic profiling characterization. NMR spectra were recorded at 600.13 MHz on a Bruker Avance-600 spectrometer, equipped with a TCI CryoProbe™ fitted with a gradient along the Z-axis, at a probe temperature of 27 °C. For each sample, 1D and 2D homonuclear and heteronuclear NMR experiments were acquired. In particular, 1D proton spectra were recorded by using the excitation sculpting sequence [30] and TSP signal was used as an internal chemical shift reference for 1H nucleus. 2D homonuclear DIPSY experiment and heteronuclear HSQC and HMBC NMR experiments were acquired with water pre-saturation, in order to maximize metabolite peak intensities (see details in Supplementary Materials). The natural abundance 2D 1He13C HSQC and 1He13C HMBC spectra were recorded on the Avance-600 spectrometer operating at 150.90 MHz for 13 C. HSQC spectra were referred to the Lactate doublet (βCH3), resonating at 1.33 ppm for 1H and 20.08 ppm for 13C, while HMBC spectra were referred to the Alanine doublet (βCH3), resonating at δ = 1.48 ppm for 1H and δ = 51.10 ppm for 13C (relative to the αCH resonance). Typical proton spectrum of diatom extract is reported in the Supplementary Fig. 1. In order to unambiguously assign metabolites, when peaks overlap in 1D spectra, we resorted to homonuclear and heteronuclear 2D experiments such as DIPSY to identify 1He1H connectivity, 1He13C HSQC and 1He13C HMBC (example in Supplementary Figs. 2–4) for directly and long range bonded 1H and 13C nuclei, respectively. With these experiments, we were able to identify resonances by a comparison with literature data [31,32] and/or online database [33]; the 1H and 13C assignments are reported in Table. In order to better compare the metabolic profiles and to highlight the differences in the metabolic content of the two species, spectra were normalized on the total area with AMIX 3.9.15 software package (Bruker Biospin, Germany).

2.3.4. Biochemical methane potential of exhausted biomass In order to compare the methane production, after and before the cascade extraction procedure, we measured the Biochemical Methane Potential (BMP), of both exhausted and on non-treated biomass, as previously described [35], and following the method of Owen [36]. Tests were performed in 200 ml serum bottles, by adding 100 ml of anaerobic inoculum to the biomass, at a 0.5 g Organic Fractionsubstrate/g Organic Fractioninoculum, ratio, and using 100 ml of anaerobic inoculum as a blank. The organic fraction of the substrate (susceptible of anaerobic degradation) was calculated by determining the Volatile Solids (VS) content of dried samples according to Standard Methods [37]. In order to provide buffer capacity for anaerobic digestion, 5 g of NaHCO3/l were supplemented at the beginning of the test, bottles were closed with butyl septa, and helium was flushed for 10 min, to remove air from the residual headspace. Samples were incubated at 35 °C, with constant shaking at 120 rpm, and methane production monitored by measuring headspace pressure (PN 5007, IFM, Germany), and gas composition (GC-TCD) [38]. 3. Results and discussion In the last decade, the main strategies for the exploitation of microalgal biomass have been focused on single-product methods, and most of the attention has been paid at identifying strains able to hyperaccumulate the target product. Despite this, some bottlenecks, especially connected to downstream processes, still hampered major development in sustainability and economics of microalgae biotechnology. In this scenario, cascade extractions are able to recover multiple microalgal compounds and appear as promising strategies to both valorise the complete biomass, and to improve the economics of the value chain [39]. In this work, we use a cascade extraction procedure to obtain three different bioproducts, for potential biotechnological applications, starting from biomass of two intensively cultivated diatoms: one native strain of the chain forming species Staurosirella pinnata, and one commercial strain of the model diatom Phaedactylum tricornutum.

2.3.2. Bioactivity assay on crude extract hydrophilic fractions HaCaT, human immortalized keratinocytes, and CHL-1, human melanoma cell lines (American Type Culture Collection, ATCC), were cultured in Dulbecco Modified Eagle Medium (DMEM), supplemented with 10% heat-inactivated fetal bovine serum (FBS), penicillin (100 U/ ml), and streptomycin (100 μg/ml). Cell cultures were maintained at 37 °C with 5% CO2 in a humidified atmosphere. For the bioactivity assays, 5 × 104 cells were seeded in 12-well plates, and the next day incubated in presence of 0.2, 0.4, 0.8, 1.6, 3.2, 10.0 mg/ml of S. pinnata and P. tricornutum extracts, dissolved in DMEM medium at the concentration of 100 mg/ml, for another 24 h. At the end of the treatment, cell death and cell cycle analyses were performed by means of flow cytometry, as previously described [34]. Briefly, cells were detached with trypsin, washed twice in PBS, suspended in propidium

3.1. Cascade extraction Investigation of the potential bioactivity, in particular cytotoxic and anti-proliferative effects on human cancer cell lines, of diatom crude extracts, focused on strains of Cylindrotheca closterium, Pseudo-nitzschia 3

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concentration. In fact, we observed a progressive and significant dosedependent increase in the percentage of cells in the S phase at the lower doses, from 50.05% of control cells, up to 58.90% at 0.8 mg/ml (p < .0001), which then decreases to 33.15% at 3.2 mg/ml (p < .0001), and 31.64% at 10 mg/ml (p < .0001). This decrease was coupled with a significant increase in the percentage of cells in the G1 phase, which rises from 33.56% of the control cells up to 52.30% at 3.2 mg/ml (p < .0001), and 54.48% at 10 mg/ml (p < .0001). Interestingly, the CHL-1 cell line showed a completely different behaviour. In fact, we detected a progressive dose-dependent decrease in the percentage of cells in the G1 phase, from 48.87% of the control cells down to 25.20% at 3.2 mg/ml (p < .0001), which was coupled to an increase in the percentage of cells both in the S phase, from 29.72% of the control cells up to 42.10% at 3.2 mg/ml (p < .05), and in the G2/ M phase, from 20.27% of the control cells up to 30.02% at 3.2 mg/ml (p < .05). Fig. 3B, shows the effects of the P. tricornutum extract-induced cell cycle alterations. In the HaCaT cell line, we observed a response similar to that recorded for S. pinnata. In fact, we could detect a progressive and significant dose-dependent increase in the percentage of cells in the S phase, from 47.11% of the control cells, up to 53.48% at 1.6 mg/ml (p < .05), which then decreases down to 24.86% at 10.0 mg/ml (p < .0001). Also in this case, we observed a coupled decrease in the percentage of cells in the G1 phase, from 35.08% of the control cells, down to 28.35% at 0.4 mg/ml (p < .05), which then increase up to 52.11% at 3.2 mg/ml (p < .001). It is worth to note that, differently from the S. pinnata extract, the treatment with the highest dose (10.0 mg/ml) results in a strong decrease in the percentage of cells in the G1 phase, down to 20.17% (p < .0001), coupled to a significant increase of cells in the G2/M phase, from 15.76% of the control cells, up to 52.37% (p < .0001). On the contrary, the CHL-1 cell line showed only a slight increase in the percentage of cells in the G1 phase, from 59.10% of the control cells, up to 66.73% at 3.2 mg/ml (p < .0001), which was coupled to a decrease in the percentage of cells in the S phase, from 21.46% of the control cells, down to 13.53% at 3.2 mg/ml (p < .0001). Interestingly, for this cell line, the treatment with the highest dose (10 mg/ml), results in a drastic reduction of the percentage of cells in the G1 phase, from 59.10% of the control cells, down up to 35.53% (p < .0001), which was coupled to an increase in the percentage of cells both in the S phase, from 21.46% of the control cells, up to 26.15% (p < .05), and in the G2/M phase, from 19.40% of the control cells, up to 32.60% (p < .0001). The compared analysis of cell death and cell cycle data, suggested that extract-contained bioactive molecules could affect DNA integrity and/or the activity of the enzyme controlling the cell cycle checkpoints, thus leading to the observed increase in the percentage of cells in the S and G2/M phases of the cell cycle, as well as to the increased cell death rate. Of particular interest is the fact that the S. pinnata extract is able to induce significant levels of cell death in the melanoma cell line, already at 1,6 mg/ml, at which normal keratinocytes viability was not affected. These results suggest the presence, in the hydrophilic fraction of S. pinnata, of molecules able to affect different targets and/or cellular pathways in malignant vs. normal cells. Further and more detailed characterization of the small molecule contained in the hydrophilic fraction could provide us with new perspectives for future biomedical exploitation [47].

sp., Skeletonema sp., and Thalassiosira weissflogii [40], especially on the liposoluble fraction and on fucoxanthin, which represents the main commercial product of diatom crude extracts [18]. Following a biorefinery approach, we developed a cascade extraction protocol, in which the first fractionation step consists in a methanol aqueous solution (20% v/v) extraction of the grinded freeze-dried biomass. The resulting crude extract was further extracted with CHCl3, in order to separate the hydro-soluble intra-cellular components from the lipo-soluble ones. and the hydrophilic fraction was characterized in respect to its antioxidant and anti-proliferative potential. The residual biomass, from the first step, was subjected to a second extraction and the fatty acid composition was characterized. Finally, the exhausted biomass was tested for Biomethane production. 3.2. Characterization of the small metabolite pool of the hydrophilic fractions The characterization of small metabolites present in the S. pinnata and P. tricornutum hydrophilic extracts was performed by means of Nuclear Magnetic Resonance (NMR). This analysis allowed the identification of 52 metabolites, including amino acids (leucine, isoleucine, alanine, threonine, arginine, lysine, valine, glutamate, glutamine, aspartate, asparagine, glycine, tyrosine, phenylalanine and tryptophan), organic acids (fumarate, formate, succinate and acetate), sugars (α/β glucose), nucleosides (uridine and cytidine), and other compounds, such as trigonelline, inositol, and choline (Table 1). Peak intensities of the spectra provided additional information about the differences in metabolite contents in the two extracts (Table 2). These results are in agreement with the few existing reports on small metabolite pools in diatoms, e.g. Chaetoceros calcitrans [41], and in the aqueous extract of the haptophyte Isochrysis galbana [42]. In addition, the presence of compounds, such as gamma-aminobutyric acid (GABA) and trigonelline, suggested a potential bioactivity for these hydrophilic extracts. In fact, it has been reported that GABA, from the rhodophyte microalga Rhodosorus marinus, exhibited inhibitor effect on the release of neuroinflammation mediators, on human keratinocytes [43], and several studies showed that trigonelline exerts different bioactivities against diabetes and hyperlipidemia [44], cancer [45], and cardiovascular diseases [46]. 3.3. Determination of the antiproliferative potential on human cancer cell lines The antiproliferative potential of the two hydrophilic fractions was assessed by evaluating both cell death induction and cell cycle variations, in a 24 h dose-response assay, on HaCaT (human keratinocytes) and CHL-1 (human melanoma) cell lines. Fig. 2 shows that the S. pinnata extract had a strong, and dose-dependent cytotoxic effect on the CHL-1 melanoma cell line. In fact, we could detect significant cell death induction starting from 37,15% (1,6 mg/ml) up to 69.75% (10 mg/ml), at the highest concentration tested. Conversely, the same treatment administered to the human keratinocytes HaCaT cell line did not induce significant cell death levels, except for the highest dose used (26.75%, 10 mg/ml). We obtained quite different results following the administration of the P. tricornutum extract. In details, we did not observe great differences in the behaviour of the two cell lines nor a clear dose-dependent cytotoxic effect, with a significant level of cell death only at the highest dose used (10.0 mg/ml). We further asked ourselves whether the differences we observed in the cell death induction levels could be related to a different impact on the cell cycle. To answer this question, we quantified the percentage of cells in each phase of the cycle, in the same dose-response experiment, by means of propidium iodide staining and flow cytometry analysis. Fig. 3A, shows the S. pinnata extract-induced cell cycle alterations in HaCaT vs. CHL-1 cell lines. In details, for the HaCaT cell line, we could detect a dual modality effect on cell cycle, depending upon extract's

3.4. Determination of lipid yields and fatty acid profiles of residual biomass extracts The residual biomass, resulting from the first extraction step, was further subjected to a second chloroform/methanol extraction, and, after solvent evaporation, relative lipid yields were estimated by quantifying the relative lipids' percentage and by comparing these values with that obtained from un-treated biomass. Our results showed that the P. tricornutum extract has a higher lipid content, and that the CH3OH extraction step did not affect their availability (Supplementary 4

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Table 1 1 H and 13C chemical shift assignment (δ, ppm) of metabolites found in 1H-TOCSY, 1He13C-HSQC and 1He13C-HMBC-NMR spectra of diatom hydrophilic extracts. Entry

Metabolite

δ1H

δ13C

Group

Entry

Metabolite

δ1H

δ13C

Group

1

Saturated fatty acids

0.89 1.29

12.06

ωCH3 CH2-CH3

27

Myo-Inositol

73.11 71.60

2

Isoleucine

0.93 1.01 1.29 1.47 1.98

11.29 15.23

δCH3 γCH3 γCH γ′CH βCH

28

β-glucose

3

Leucine

0.95 0.97 1.72 3.72

22.29 21.97 40.30 61.81

δCH3 δCH3 βCH2 αCH

29

α-glucose

4

Valine

17.32 18.35 29.90

30

Scyllo-inositol

Propionate

31

Methanol

3.36

49.40

CH3

6

Threonine

32

Glycerol

7

Ethanol

33

Glycine

3.56 3.66 3.80 3.57

42.00

C1H C3H C2H αCH

8

Lactate

34

Urea

5.78

9

Lysine

35

Uracil

5.80 7.53

101.40 144.00

C5 ring C6 ring

10

Alanine

γCH3 γ′CH3 βCH αCH βCH3 αCH2 γCH3 αCH βCH βCH3 αCH2 βCH3 αCH γCH2 δCH2 βCH2 εCH2 αCH βCH3 αCH

96.40 74.60 76.22 70.70 76.80 61.40 92.40 72.20 73.30 70.03 71.61 62.37 74.20

C5H C3H C4, 6H C2H C1H C2H C3H C4H C5H C6H C1H C2H C3H C4H C5H C6H CH2

5

0.99 1.04 2.28 3.62 1.05 2.19 1.33 3.59 4.27 1.19 3.66 1.33 4.12 1.46 1.73 1.91 3.03 3.74 1.48 3.78

3.29 3.54 3.64 4.06 4.64 3.26 3.48 3.40 3.47 3.90 5.24 3.54 3.72 3.42 3.84 3.78 3.35

36

Uridine

11

Arginine

37

Cytidine

4.23 5.90 5.91 7.87 6.07 7.84

70.10 102.80 92.80 142.20 96.79 142.00

C4 ring C2 ring C1 ring C11 ring C10 ring C11 ring

12

Gaba

38

Inosine

13

Acetate

39

2-Deoxyuridine

14

Proline

40

2-Deoxy-guanosine

6.10 8.22 8.35 6.27 4.45 6.30 8.00

88.60 146.68 140.60 85.80 70.70 85.40 136.30

C2 ring C7 ring C12 ring C2 ring C4 ring C2 ring C7 ring

15

Glutamate

41

2-Deoxy-adenosine

6.50 8.22 8.33

84.90 145.80 140.80

C2 ring C12 ring C7 ring

16

Glutamine

42

Fumarate

6.52

136.30

α,β C=C

17

Methionine

43

p-Cresol

6.83 7.12

18

Succinate

44

4-hydroxyphenyl-Acetate

19

Aspartate

45

Tyrosine

6.87 7.16 6.90 7.20

20

Asparagine

46

τ-methylhisa

7.06 7.65

21

Cysteine

47

Phenylalanine

7.34 7.37 7.43

17.30 58.00 20.08 61.20 65.67 17.30 58.00 20.08 68.90 21.99 26.80 30.29 39.70 54.80 16.47 51.40

1.65 1.91 3.25 3.78 1.89 2.30 3.02 1.92

24.30 28.04

2.06 2.34 3.35 3.43 4.15 2.09 2.09 2.35 3.76 2.17 2.42 3.77 2.13 2.15 2.65 3.89 2.41

29.30 29.30 46.50

2.69 2.82 3.91 2.86 2.96 4.01 3.04 3.13 3.99

37.10

54.80 24.05 35.04 40.09 23.59

61.60 27.60 27.60 33.90 55.12 27.55 55.12

34.30

36.90

γCH2 βCH2 γCH3 αCH βCH2 αCH2 γCH2 βCH3 β′CH βCH δ′CH δCH αCH βCH γCH2 αCH βCH2 γCH2 αCH S-CH3 βCH2 γCH2 αCH α,βCH2 βCH β′CH αCH βCH β′CH αCH βCH β′CH αCH

(NH2)2

C2,6 ring C3,5 ring

115.90 117.60 116.30 131.10

C2,6 C3,5 C3,5 C2,6

ring ring ring ring

C4H ring C2H ring 129.02 138.03 127.00

C4 ring C2,6 ring C3.5 ring

(continued on next page) 5

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Table 1 (continued) Entry

a

22

Cr /PCr

23

24

25

26

δ1H

Metabolite a

δ13C

Group

Entry

Metabolite

3.06 3.96

39.40 54.18

NCH3 CH2

48

N-Phenyl-A-Gly

Choline

3.20 3.51 4.07

54.38

NCH3 CH2 CH2

49

Tryptophan

PCa

3.22 3.60 4.16 3.23 3.62 4.33 3.26 3.90

54.29

NCH3 CH2 CH2 NCH3 NCH2 CH2 NCH3 αCH2

50

Trigonelline

GPCa

Betaine

54.42

54.47 66.50

51

52

Nicotinate

Formate

a

δ1H

δ13C

Group

7.35 7.36 7.41 7.74 7.54 7.30 7.21 8.09 8.84 9.13 8.26 8.62 8.94 8.47

129.00 131.02 131.00 118.90 112.70 126.20 120.00

C4 ring C2,6 ring C3,5 ring C4 ring C7 ring C6 ring C5 ring C4 ring C3,5 ring C1 ring C4 ring C6 ring C2 ring HCOO−

146.10 146.40 150.70 149.20 171.50

a Abbreviations: Cr, Creatine; PCr, Phosphocreatine; PC, Phosphocholine; GPC, Glycerophosphocholine; τ-methylhis, τ-methylhistidine; N-Phenyl-A-Gly, NPhenylacetylglycine.

Table 2 Differences in the metabolites content of Staurosirella pinnata and Phaeodactylum tricornutum hydrophilic extracts. −, metabolite absence; +, different amount of metabolite. Metabolite

Staurosirella pinnata

Phaeodactylum tricornutum

Trigonelline Phenylalanine Formic acid Cytidine p-Cresol Uracil Short-chain fatty acids Ethanol Lysine Gaba Betaine Methionine Inosine Glutamate Scyllo-inositol Succinic acid Proline 4-hydroxyphenylacetate Cysteine τ-Methylhistidine

− ++ − ++ ++ − ++ + +++ +++ + ++ + + − + + + − +

+++ + ++ + − ++ + +++ + + +++ − ++ ++ ++ +++ +++ − + −

Fig. 3. Evaluation of cell cycle variations after administration of hydrophilic fraction extracts to human keratinocytes (HaCaT) and melanoma (CHL-1) cells lines. Cell lines were incubated, for 24 h, with the indicated amount of the S. pinnata (A) and P. tricornutum (B) hydrophilic fraction extract, and the percentage of cells in the G1, S, and G2/M phases quantified by flow cytometry.

hexadecanoic (C16:0, 29.79%), cis-9-hexadecenoic (C16:1, 27.41%), and eicosapentaenoic (EPA, C20:5, 12.59%) acids were the most abundant fatty acids in the S. pinnata extract, while eicosapentaenoic (EPA, C20:5, 20.59%), docosanoic (C22:0, 20.58%), cis-9-acid (C16:1, 17.27%), and hexadecanoic (16:0, 14.87%) acids, were the most abundant in the P. tricornutum extract. Our findings are consistent with data reported in literature, for the same species collected from natural environment or cultivated in photobioreactors, and not pre-extracted [23,48]. It is worth to note, that both extracts showed a relative abundance of eicosapentaenoic, hexadecenoic, and octadecanoic acids, all molecules with known biological activities and of interest for their potential biomedical application. In particular, EPA has been reported to play an important role in human metabolism [49], to protect cardiovascular system [50], and to be effective in cancer prevention, insulin resistance, tissue inflammation, and obesity [51], while hexadecanoic acid obtained from algal sources, has been shown to selectively inhibit topoisomerase I in human cancer cell lines [52]. In addition, the presence of PUFAs in the S. pinnata and P. tricornutum biomasses suggest a possible exploitation in feed industry, as supplements for larvae and early stages of fish, as well as shellfish in aquaculture. In fact, feed enriched with microalgal PUFAs improve the

Fig. 2. Evaluation of cell death induction after administration of hydrophilic fraction extracts to human keratinocytes (HaCaT) and melanoma (CHL-1) cells lines. Cell lines were incubated, for 24 h, with the indicated amount of S. pinnata and P. tricornutum hydrophilic fraction extract, and cell death was quantified as percentage of hypodiploid events. *, p < .05; **, p < .01; ****, p < .0001, respect to the HaCaT cell line.

Fig. 5). In details, the lipid content of the residual biomass was of 17.09% ± 1.50% for P. tricornutum and of 14.54% ± 1.61% for S. pinnata, with no significant difference as compared to non-treated biomass (17.29% ± 2.98% and 13.05% ± 2.54%, respectively). Qualitative analysis of the lipid fractions was performed by determining their fatty acid (FA) profile, by means of GC-FID. Table 3 shows that 6

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Table 3 Fatty acids composition of the lipid fraction extracted from Staurosirella pinnata and Phaeodactylum tricornutum residual biomass. The extraction was performed from the residual biomass and Fatty Acids (FA) profiles were qualitatively analysed using GC-FID (three replicates).

Fatty acid (FA) Butanoic acid (C4:0) Hexanoic acid (C6:0) Octanoic acid (C8:0) Decanoic acid (C10:0) Undecanoic acid (C11:0) Dodecanoic acid (C12:0) Tridecanoic acid (C13:0) Tetradecanoic acid (C14:0) Tetradecenoic acid (C14:1) Pentadecanoic acid (C15:0) Cis-10-Pentadecenoic acid (C15:1) Hexadecanoic acid (16:0) Cis-9-Hexadecenoic acid (C16:1) Heptadecanoic acid (C17:0) Cis-10-Heptadecenoic acid (C17:1) Octadecanoic acid (C18:0) (C18:1 c + t) (C18:2 c + t) (C18:3 n-6) (C18:3 n-3) (C20:3n6 + C21:0) (C20:3 n-3) Eicosapentaenoic acid (C20:5) Docosanoic acid (C22:0) Tetracosanoic acid (C24:0) (C22:6 + C24:1)

Staurosirella pinnata

Phaeodactylum tricornutum

% of FA 2.15 1.10 1.66 0.58 0.81 – 0.66 5.75 – 2.95 1.07 29.79 27.41 – 1.97 5.81 2.50 1.26 1.88 – – – 12.59 – – –

% of FA 0.58 0.19 0.09 0.09 0.12 0.15 0.35 6.80 0.30 0.32 – 14.87 17.27 3.01 5.92 1.92 0.74 1.65 0.19 0.40 0.30 0.70 20.59 20.58 1.15 1.64

Fig. 4. Biochemical methane potential rates in Staurosirella pinnata and Phaeodactylum tricornutum biomass. Non-treated and exhausted biomass were subjected to anaerobic digestion, and the Biochemical Methane Potential (BMP) rates were recorded over a period of 27 days.

biomethane (CH4) productivity rates, over a 27 days period, and by comparing the obtained values to that of non-treated diatom biomass. Fig. 4 shows that the CH4 production rates recorded for S. pinnata nontreated vs. exhausted biomass were similar, and the ultimate methane yields were 79.2 ± 5.9 and 68.6 ± 12.4 ml CH4 g−1 Organic Fraction, respectively. Conversely, production rates recorded for P. tricornutum revealed a different behaviour between non-treated and exhausted biomass. Indeed, CH4 production rates of non-treated biomass exponentially increased starting from day 3, with a final value of 239.4 ± 6.7 ml CH4 g−1 Organic Fraction, at day 27. By contrast, anaerobic digestion of the exhausted biomass results in an initial, from day 1 to day 9, inhibition of methane production, followed by a rapid increase of the production rates, from day 9 to day 23, which then stabilise until day 27, with an ultimate methane yield of 164.8 ± 10.3 ml CH4 g−1 Organic Fraction. The biomethane yield we obtained for P. tricornutum non-treated biomass, is comparable with the values reported in literature [54]. It is worth to note that the biomethane productivity of P. tricornutum exhausted biomass was comparable with that measured for Nannochloropsis salina biomass after lipid extraction, for the green algae Scenedesmus obliquus and Chlorella vulgaris untreated biomass, and for the cyanobacterium Arthrospira maxima [55]. On the other hand, biomethane yields recorded for both non-treated and exhausted biomass of S. pinnata were similar, thus not substantially altered by cascade extraction but significantly lower than those reported in literature for different microalgal species [55].

animal immune response, disease resistance, physiology, and, in general, quality and performance of the final product [53]. These findings highlight that microalgal oil, derived from both S. pinnata and P. tricornutum residual biomass, represents a second value added product with potential for large scale exploitation. 3.5. Determination of the biochemical methane potential of exhausted biomass As a final product characterization, we quantified the percentage of the Organic and Inorganic Fractions in the exhausted diatom biomass, as well as their potential exploitation for biomethane production, by means of anaerobic digestion in a Biochemical Methane Potential (BMP) assay. Table 4 shows that, the P. tricornutum exhausted biomass showed the highest Organic Fraction content (88.07 ± 0.50%), as compared to P. tricornutum non-treated biomass (81.66 ± 0.5%), and to both S. pinnata exhausted (45.81 ± 0.5%) and non-treated (49.88 ± 0.5%) ones. As for the Inorganic Fraction, the results were the opposite, with S. pinnata exhausted biomass showing the highest content (54.19 ± 0.5%), as compared to S. pinnata non-treated biomass (50.12 ± 0.5), and to both P. tricornutum exhausted (11.93 ± 0.5%) and non-treated biomass (18.34 ± 0.5%). The BMP of exhausted biomass was assessed by measuring

4. Conclusions In this work, we used a cascade extraction protocol to obtain multiple valued products from the same biomass of two intensively cultivated diatom species: S. pinnata and P. tricornutum. The hydro-soluble fraction of S. pinnata crude extract, which contains different small metabolites, exhibits an efficient cell death inducing potential, when tested on human melanoma cancer cells. The lipid fraction of both diatoms contains a high percentage of fatty acids, of interest for their potential exploitation in biomedical and feed industry. Finally, the exhausted biomass proved to be a suitable substrate for anaerobic digestion, thus being exploitable for biomethane production. These results represent a first step towards: the characterization of the compound(s) responsible for the bioactivity on the human cell lines; the evaluation of the bioactive potential of the lipidic fraction; the improvements of the extraction procedures, by using novel green extraction techniques, in order to reduce the use of solvents and to make the cascading process ecologically and economically sustainable.

Table 4 Quantification of the organic and inorganic fractions in non-treated and exhausted diatom biomasses, employed for the determination of the Biochemical Methane Potential.

Phaeodactylum tricornutum Staurosirella pinnata

Non-treated biomass Exhausted biomass Non-treated biomass Exhausted biomass

Organic (%)

Inorganic (%)

81.7 88.1 49.9 45.8

18.3 11.9 50.1 54.2

± ± ± ±

0.5 0.5 0.5 0.5

± ± ± ±

0.5 0.5 0.5 0.5

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Acknowledgements

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The authors acknowledge ES1408 Cost Action EUALGAE, for networking and knowledge transfer. This work was partially supported by “University of Rome Tor Vergata - Consolidate the Foundations” grant to C.R. Authors' contributions S.S., S.F. carried out the diatom cultivation and extraction protocol; D.P. performed the NMR analysis; E.A. performed the GC-FID; I.D. performed the BMP assay; I.D., S.B., R.M. elaborated the BMP data and revised the manuscript; S.S. and C.R. performed the cell assays; R.C., C.R. and S.S. conceived the research, wrote and revised the manuscript. Declaration of competing interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Appendix A. Supplementary data Supplementary data to this article can be found online at https:// doi.org/10.1016/j.algal.2020.101830. References [1] E. Posadas, C. Alcántara, P.A. García-Encina, L. Gouveia, B. Guieysse, Z. Norvill, F.G. Acién, G. Markou, R. Congestri, J. Koreiviene, R. Muñoz, Microalgae cultivation in wastewater, in: C. Gonzalez-Fernandez, R. Muñoz (Eds.), Microalgae-Based Biofuels and Bioproducts, Woodhead Publishing, 2017, pp. 67–91. [2] M.C. Deprá, A.M. dos Santos, I.A. Severo, A.B. Santos, L.Q. Zepka, E. Jacob-Lopes, Microalgal biorefineries for bioenergy production: can we move from concept to industrial reality? BioEnergy Research 11 (2018) 727–747, https://doi.org/10. 1007/s12155-018-9934-z. [3] P. Steinrucken, S.R. Erga, S.A. Mjos, H. Kleivdal, S.K. Prestegard, Bioprospecting North Atlantic microalgae with fast growth and high polyunsaturated fatty acid (PUFA) content for microalgae-based technologies, Algal Res. 26 (2017) 392–401, https://doi.org/10.1016/j.algal.2017.07.030. [4] M.S. Chauton, K.I. Reitan, N.H. Norsker, R. Tveterås, H.T. Kleivdal, A techno-economic analysis of industrial production of marine microalgae as a source of EPA and DHA-rich raw material for aquafeed: research challenges and possibilities, Aquaculture 436 (2015) 95–103, https://doi.org/10.1016/j.aquaculture.2014.10. 038. [5] E.W. Becker, Microalgae for human and animal nutrition, Handbook of Microalgal Culture, 2013. [6] J.A.V. Costa, B.C.B. Freitas, C.G. Cruz, J. Silveira, M.G. Morais, Potential of microalgae as biopesticides to contribute to sustainable agriculture and environmental development, J. Environ. Sci. Health B 54 (2019) 366–375, https://doi.org/10. 1080/03601234.2019.1571366. [7] C. Schreiber, H. Schiedung, L. Harrison, C. Briese, B. Ackermann, J. Kant, S.D. Schrey, D. Hofmann, D. Singh, O. Ebenhöh, W. Amelung, U. Schurr, T. MettlerAltmann, G. Huber, N.D. Jablonowski, L. Nedbal, Evaluating potential of green alga Chlorella vulgaris to accumulate phosphorus and to fertilize nutrient-poor soil substrates for crop plants, J. Appl. Phycol. 30 (2018) 2827–2836, https://doi.org/ 10.1007/s10811-018-1390-9. [8] M. Hayes, H. Skomedal, K. Skjånes, H. Mazur-Marzec, A. Toruńska-Sitarz, M. Catala, M. Isleten Hosoglu, M. García-Vaquero, Microalgal proteins for feed, food and health, in: C. Gonzalez-Fernandez, R. Muñoz (Eds.), Microalgae-Based Biofuels and Bioproducts, Woodhead Publishing, 2017, pp. 347–368. [9] B. Gilbert-Lopez, A. Barranco, M. Herrero, A. Cifuentes, E. Ibanez, Development of new green processes for the recovery of bioactives from Phaeodactylum tricornutum, Food Res. Int. 99 (2017) 1056–1065, https://doi.org/10.1016/j.foodres. 2016.04.022. [10] B. Gilbert-López, J.A. Mendiola, J. Fontecha, L.A.M. van den Broek, L. Sijtsma, A. Cifuentes, M. Herrero, E. Ibáñez, Downstream processing of Isochrysis galbana: a step towards microalgal biorefinery, Green Chem. 17 (2015) 4599–4609, https:// doi.org/10.1039/C5GC01256B. [11] L. Bastiaens, S. Van Roy, G. Thomassen, K. Elst, Biorefinery of Algae: Technical and Economic Considerations, (2017). [12] R. Congestri, E.J. Cox, P. Cavacini, P. Albertano, Diatoms (Bacillariophyta) in phototrophic biofilms colonising an Italian wastewater treatment plant, Diatom Research 20 (2005) 241–255, https://doi.org/10.1080/0269249X.2005.9705634. [13] M. Hildebrand, A.K. Davis, S.R. Smith, J.C. Traller, R. Abbriano, The place of diatoms in the biofuels industry, Biofuels 3 (2012) 221–240, https://doi.org/10.4155/ BFS.11.157.

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