Technologies for Polysaccharide Extraction from Marine Brown Algae

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This entry is adapted from the peer-reviewed paper 10.3390/md18030168

polysaccharides marine algae extraction fucoidan laminarin alginate

1. Introduction

The term macroalgae refers to aquatic photosynthetic organisms which are included in the Eukaryota domain as well as Plantae and Chromista kingdoms ^[1]. They differ according to several characteristics such as cell wall composition, presence or absence of flagella and ultrastructure of mitosis ^[2]. Their distribution, diversity and chemical composition are mainly limited by the environmental conditions, e.g., sunlight availability (chromatic adaptation) and water temperature. Based on their pigmentation and chemical composition, macroalgae can be classified into three groups: brown (Phaeophyceae), red (Rhodophyceae) and green (Chlorophyceae) ^[3]^[4].

Brown algae are a rich source of bioactive molecules such as proteins, amino acids, polysaccharides, fatty acids, vitamins, minerals, dietary fibre, sterols, pigments, polyphenols etc. which possess a broad spectrum of biological activities (anticoagulant, antithrombotic, anti-viral, anti-cancer, anti-inflammatory and antibacterial) ^[5]. These compounds therefore provide high potential for the application of brown algae extracts in the treatment of arteriosclerosis, rheumatic processes, hypertension, goitre, asthma, ulcers, menstrual disorders, syphilis, skin diseases etc. ^[4]^[6]^[7]. The biological potential of brown algae is significantly contributed by polysaccharides as one of the most common and most important groups of bioactive compounds. Over the last years, considerable interest has been raised about different types of polysaccharides in brown algae cell walls, including laminarins, alginates and fucoidans which have high potential for biological applications in functional foods, cosmeceutical and pharmaceutical products ^[8]. The structure and composition of algal polysaccharides (APS) is determined by algae species however it is also influenced by other factors causing inter-species variation, e.g., growth location and harvesting season ^[8]. Vast structural variation between the APS therefore presents a challenge in terms of pre-treatments application, extraction techniques and optimization, characterization of isolated fractions and determination of their biological properties.

Chemical structure and yield of APS isolated from marine macroalgae by conventional extraction (CE) techniques can be affected by various experimental conditions (pH, time, temperature, pressure, particle size, solvent, sample to solvent ratio, agitation speed etc.). In addition, different advanced techniques such as microwave assisted extraction (MAE), ultrasound assisted extraction (UAE), pressurized liquid extraction (PLE), enzyme-assisted extractions (EAE) are assessed and applied for APS extraction ^[9]^[10]^[11].

In general, the chemical structure of polysaccharides determines its physical, chemical and biochemical properties as well as its biological activities ^[12]. Several studies have reported that their biological activity is strongly associated with their chemical structure ^[9]. Due to very complex mechanisms that are affected by many factors, the correlation between polysaccharide structure and biological activity is still not sufficiently clarified.

In order to improve isolation of APS, pre-treatments are usually applied to the algal biomass prior to the extraction process with the two aims: (i) to prevent co-extraction of interfering bioactive compounds with similar solubility; and (ii) to disrupt cell walls and improve mass transfer of APS into extraction solvent. The first type of pre-treatments is therefore used to remove compounds which are highly bound to the APS such as proteins, phenols and lipids, as well as mannitol and chlorophyll ^[13]. For that purpose, the application of various pre-treatment solvents at different temperatures has been studied. A mixture of methanol, chloroform and water at 4:2:1 (v/v/v) has been successfully used for defatting, and acetone ^[14] as well as mixture of acetone and ethanol ^[15] or methanol ^[16] were used to remove lipids and pigments. The second type of pre-treatments which is carried out in order to disrupt cell wall material and enhance the mass transfer of the target compounds to the extraction solvent result in an improved extraction yield ^[13]. In addition, several mechanical and physical methods for cell disruption which include milling, high pressure extrusion, ultrasonication and microwave pre-treatment have been described in the literature ^[17].

2. The Chemical Structure and Bioactivity of Polysaccharides from Marine Brown Algae

Polysaccharides from marine macroalgae differ greatly from the ones present in terrestrial plants such as cellulose and starch ^[18]. Brown seaweed cell walls contain sulfated polysaccharides i.e., laminarin and alginate along with fucoidan, which is not present in any other type of seaweeds. These three types of APS have their own unique physical and chemical characteristics which are influenced by species, geographic location, season and population age ^[19].

2.1. Laminarin

Laminarin is a water-soluble linear polysaccharide that consists of β (1→3) and β (1→6) glucan in a 3:1 ratio ^[20] (Figure 1). Molecular weight (MW) of laminarin is around 5 kDa depending on the degree of polymerization ^[21]. In addition, in dependence on the type of sugar at the reducing end, there are M chains with terminal 1-O-substituted D-mannitol, and G chains with glucose. Laminarin is mainly isolated from the brown algae species Laminaria and Alaria. Based on the type of algae and harvest season ^[22] as well as the environmental conditions such as sea temperature, salinity, sea currents, depth and availability of nutrients ^[23], laminarin represents around 22–49% of algal dry matter. Apart from contributing to dietary fibre intake, studies have shown that laminarin and products of its enzymatic hydrolysis inhibit the production of melanoma cells and colon cancer ^[24] and also show anti-metastatic effects which makes them potentially useful in cancer treatment ^[25].

Figure 1. Structure of laminarin ^[26].

2.2. Alginates

Alginates are linear hetero-polysaccharides composed of β-D-manuronic acid (M) and α-L-guluronic acid (G) (Figure 2). These two monomers are linked in a 1→4 configuration and arranged as homogeneous MM, GG or alternatively MG blocks. The proportion of these three block types is responsible for the physical properties of alginates whereas alginates with high M blocks share have higher viscosity while alginates with high G blocks share have better gelling properties ^[27]. Alginates are obtained from cell walls of various brown algae that grow in colder seas such as Microcystis, Laminaria and Ascophyllum sp. ^[28]. In addition, alginates can be present in alginic acid form as well as in the form of its salt, which make about 40% dry matter of the algae biomass ^[29].

Figure 2. Structure of alginates ^[30].

Studies have shown that alginic acid has a positive effect on preventing the absorption of heavy metals in the body, reducing blood pressure and cholesterol as well as assisting in the absorption of cholesterol ^[31]^[32]. In addition, alginates with molecular mass larger than 50 kDa showed a positive effect in prevention of diabetes and adiposity ^[33]. Furthermore, they have anticarcinogenic properties ^[32] and inhibitory effect on microorganisms such as Staphylococcus, Listeria, Salmonella and Escherichia coli ^[34]. Due to their properties, alginates are widely used by the food industry as stabilizers, emulsifying agent or thickeners, as well as by the cosmetic and pharmaceutical industries ^[35].

2.3. Fucoidan

Fucoidan is sulfated polysaccharide found in fibrillar tissue of the cell wall and intercellular space of brown algae. It consists mainly of fucose interconnected by α−(1,3) glycoside bonds, alternating α−(1,3) and α−(1,4) bonds and rarely α−(1,2) bonds (Figure 3). Apart from fucose, it also contains other monosaccharides, including galactose, glucose, mannose, xylose, rhamnose, and uronic acids which contents vary depending on algal species and season ^[19]^[36]. The average relative MW of fucoidan varies from 7 kDa ^[37] to 2300 kDa ^[38]. Fucoidan is the most abundant in orders Laminariales and Fucales and, depending on the algae type, it represents approximately 5% to 10% of algae dry matter while its sulfate content varies between 5% ^[19] and 38% ^[39].

Figure 3. Structure of fucoidan from Fucus vesiculosus, with a backbone of alternating (1→3)-linked α-L-fucopyranose and (1→4)-linked α-L-fucopyranose residues and the presence of sulfate groups on both O-2 and O-3 ^[40].

Fucoidan is one of the most researched algae molecules and studies have found that it shows a wide range of positive effects such as antioxidant, anti-inflammatory and antitumor ^[10]^[19]^[41]. It has been increasingly studied not only for its potential applications as the heparin-like anticoagulant and antithrombotic agent but also, due to its non-toxicity and biodegradability, as a functional additive for developing novel drug delivery systems and functional foods. Biological activities of fucoidan are closely correlated to its physicochemical properties such as MW, types and ratios of constituent monosaccharides, features of glycosidic linkages, sulfation degree and the position of sulfate groups.

3. The Perspective of Advanced Technologies for Polysaccharide Extraction from Marine Brown Algae

In general, APS isolation process (Figure 4) includes several complex and time consuming steps such as seaweed preparation, pre-treatment, extraction (conventional or advanced) and purification. These procedures are usually followed by biological activity assays which enable the determination of APS potential industrial use.

Figure 4. Schematic overview of essential steps for extraction of brown algae polysaccharides.

Preparation of the seaweed includes washing the seaweed, preferably with distilled water to remove salt and impurities, as well as drying or freeze drying and milling in order to achieve the powder with higher surface-to-volume ratio. Removal of interfering algal compounds prior to polysaccharide extraction is useful for prevention of contamination of the target polysaccharide. Conventional polysaccharide extraction is usually performed by hot aqueous or acidic solutions at high temperatures for several hours. On the other hand, advanced technologies (e.g., MAE, UAE, PLE and EAE) have already demonstrated numerous advantages over CE especially in the terms of increased extraction efficiency, reduction of extraction time, energy and solvent usage as well as preservation of sensitive and unstable bioactive molecules (e.g., polyphenols). Regardless of which extraction technique is performed, the extraction parameters should be optimized as they may influence APS chemical structure, bioactivity and potential industrial use (Figure 5). Moreover, preservation of the APS structural integrity is crucial for obtaining the relevant structural characteristics required for their specific biological activities ^[10].

Figure 5. Schematic diagram of process parameters, chemical structure properties, biological activity and potential industrial uses of brown algae polysaccharides.

3.1. Pre-treatment of Marine Brown Algae

Prior to the APS extraction, it is beneficial to apply pre-treatment in order to remove lipids, pigments and low molecular weight compounds from the seaweed material. For that purpose, various solvents and solvent mixtures with different polarity, that do not cause any APS structural changes, have been used ^[13]. Lipids are traditionally removed by lower polarity solvents, such as chloroform, petroleum ether or dichloromethane; pigments with semipolar solvents, such as acetone, methanol or ethanol; while other high polarity molecules, such as monosaccharides, proteins and minerals are extracted in water ^[20]. Prior to laminarin extraction from Cystoseira barbata, Sellimi et al. ^[42] seaweed powder was sequentially treated twice with acetone-methanol (7:3) and twice with chloroform for 24 h at 30 °C with constant stirring (250 rpm). Similarly, January et al. ^[43] used the mixture of methanol-chloroform-water (4:2:1) as a pre-treatment prior to fucoidan extraction. However, as chloroform is toxic and classified as an extremely hazardous substance in the United States as it is defined in Section 302 of the U.S. Emergency Planning and Community Right-to-Know Act (42 U.S.C. 11002), new chloroform-free pre-treatment alternatives are being proposed. As an alternative solvent, relatively low cost and non-toxic petroleum ether has been used by Sahera et al. ^[44]. Furthermore, Cystoseira myrica powder was treated with petroleum ether and acetone in Soxhlet apparatus prior to polysaccharides extraction. The same procedure was applied by Abid et al. ^[45] for sodium alginate extraction from Dictyopteris membranaceae and Padina pavonica. However, in their work, algae powder was previously macerated three times by methanol and dichloromethane (1:1, v/v) for 48 h. In addition to acetone, 95% ethanol (v/v), 80% ethanol (v/v) and methanol were also used ^[15]^[16]^[46]. Compared to previously described procedures, where higher temperatures were not used because they might promote the extraction of fucoidan, with 80% ethanol (v/v) as a solvent, higher temperature is acceptable. Since fucoidan is not soluble in ethanol, even if it was unintentionally extracted, it would be precipitated and thus only impurities like pigments and lipids would be removed by filtration ^[19].

Prior to APS extraction, pre-treated seaweed is further exposed to the conventional, vacuum or freeze drying methods. In addition, as seaweed rigid cell walls are hard to break ^[47], a cell disruption pre-treatment is generally required to remove or weaken them, making the intracellular molecules more accessible to solvent in further extraction steps. For that purpose, various cell disruption pre-treatment methods such as mechanical, chemical, thermal, enzymatic or advanced techniques, like ultrasound and microwave, could be applied. Ultrasound and microwave pre-treatments are based on the energy waves that have an effect on the cell wall material causing its lysis and release of intracellular molecules. However, this approach is more often used in bioethanol production from algae where pre-treatment is intended for cell disruption and complex carbohydrates release. Complex carbohydrates are then broken down into their monosaccharide components (simple sugars) which are fermented into bioethanol and carbon dioxide ^[48]. To the best of our knowledge, the only research where cell disruption pre-treatment was applied prior to polysaccharide extraction from brown seaweed was made by Kadam et al. ^[49]. In their work, ultrasound pre-treatment for 10 min (20 Hz, amplitude 20–100%, 13 mm diameter probe) was applied followed by simple extraction in orbital shaker for 1 to 22 hours to extract fucose and uronic acid from Ascophyllum nodosum. Compared to control extraction, ultrasound pre-treatment increased fucose and uronic acid extraction yield, whereas ultrasound amplitude was the most significant factor contributing to the pre-treatment efficiency ^[49].

3.2. Extraction Techniques

Following pre-treatment, seaweed samples are subjected to various extraction techniques. General principal of these procedures is to extract target compounds with minimum co-extraction of other polysaccharide constituents, e.g., isolate fucoidan from alginate. If alginate is co-extracted, further steps are needed to remove alginate from fucoidan, thus increasing the purity of the extracted fucoidan ^[20].

3.2.1. Conventional Extraction Technique (CE)

Conventional extraction of APS is typically performed by treating the algal material with various solvents such as hot water, acidic or salt solutions at high temperatures for several hours. Table 1 summarizes the most frequently used parameters of CE for APS recovery from brown algae.

Table 1. The most frequent conventional extraction parameters used for the recovery of brown algae polysaccharides.

Algae	Polysaccharide	PRETREATMENT	EXTRACTION	Purification	Yield	References
Algae	Polysaccharide	Solvent; Time; Temperature	Solvent; Time; Temperature	Purification	Yield	References
S. henslowianum	fucoidan	95% EtOH; 2 × 12 h	H₂O; 3 × 2 h; reflux	EtOH precipitation; dialysis (12000 Da)	5.1%	^[46]
S. fusiforme	fucoidan	95% EtOH; 24 h; 30 °C	H₂O; 3 h; 80 °C	EtOH precipitation; dialysis (3,5 kDa)	3.94–11.24%	^[50]
			1.0M HCl; 6 h; 25 °C
			2% CaCl₂; 3 h; 50 °C
E. maxima L. pallida S. rugosum	fucoidan	/	H₂O; 24 h; 70 °C		/	^[43]
		/	0.15M HCl; 2 h; 65 °C	EtOH precipitation
		methanol-chloroform-H₂O (4:2:1); overnight; room temp.	2% CaCl₂; 5 h; 85 °C	10% CTAB
C. barbata	laminarin	acetone-methanol (7:3); 2 × 24 h; 30 °C chloroform; 2 × 24 h; 30 °C	0.1M HCl; 2 × 2 h; 60 °C	EtOH precipitation; ultrafiltration (50, 10 & 1 kDa)	7.27%	^[42]
Cystoseira compressa	sodium alginate	acetone; 2 × 24 h; 25 °C methanol; 2 × 24 h; 25 °C	0.1M HCl; 2 h; 60 °C 3% Na₂CO₃; 2 h; 60 °C	EtOH precipitation; dialysis (3,5 kDa)	fucoidan—5.2%	^[16]
Dictyopteris divaricata	polysaccharides	/	H₂O; 5–7 h; 80–100 °C; water to solid ratio 90–110 mL/g	EtOH precipitation	3.05%	^[51]
Sargassum latifolium	sodium alginate	/	2% citric acid; 2 h; room temperature 3% Na₂CO₃; 1–3 h; 25–45 °C	EtOH precipitation	18.89–40.43%	^[52]
Fucus serratus F. vesiculosus A. nodosum	fucoidan	85% EtOH; overnight; room temp.	0.1M HCl; 4 h; 80 °C 1% CaCl₂; overnight; 4 °C	EtOH precipitation	F. serratus—4.2–7.5% F. vesiculosus—8.1–12.2% A. nodosum—6.5–8.9%	^[53]
D. Membranaceae P. Pavonica	sodium alginate	methanol-dichloromethane (1:1); 3x48h; room temp. petroleum ether; soxhlet acetone; soxhlet	2% CaCl₂; 3 × 3h 1M Na₂CO₃; 2 h	EtOH precipitation	D. Membranaceae - 18.93% P. Pavonica—66.72%	^[45]
Cystoseira sedoides	fucoidan sodium alginate	acetone; 24 h; 25 °C 80% EtOH; 24 h; 25 °C 80% EtOH; 24 h; 78 °C	2% CaCl₂; 7 h; 70°C 2% Na₂CO₃; 70°C	dialysis (7 kDa)	fucoidan—4.2% alginate—11%	^[15]
C. myrica	polysaccharides	petroleum ether acetone	H₂O; 8 h; 80°C	EtOH precipitation; 10% CTAB; dialysis	5.3%	^[44]
Cystoseira crinite C. compressa C. sedoides	fucoidan	methanol-dichloromethane (1:1); 3 × 48 h; room temp.	2% CaCl₂; 3 × 3h	dialysis (30 kDa)	2.8–3.7%	^[54]

3.2.2. Advanced Extraction Techniques

Recently, advanced extraction techniques used for polysaccharide extraction from algae are microwave-assisted extraction (MAE), ultrasound assisted extraction (UAE), pressurized liquid extraction (PLE) and enzymatic assisted extraction (EAE). However, due to various extraction conditions, APS degradation could occur which may affect the extract viscosity, sulfate content, monosaccharide composition, MW and bioactivity. Therefore, extraction parameters such as temperature, time, power and sample to solvent ratio should be optimized.

3.3. Purification Procedure

In addition to having variable MWs, monosaccharides composition and sulfate content, extracted APS are usually contaminated with proteins and low molecular weight compounds which were also dissolved in water during extraction ^[3]. Therefore, they can be additionally purified using different purification methods such as ethanol precipitation, membrane separation, ion-exchange, size-exclusion and affinity chromatographic methods. As it depends on the purity requirements and further function, there are no standard protocols set for purification. In functional food industry ethanol precipitation is the most frequently used method while in scale-up testing membrane separation can be used ^[3].

Ethanol precipitation is often used as the first step in APS purification, which removes low molecular weight impurities from the polysaccharides. Numerous researchers conducted purification step by adding two or three volumes of ethanol to the extract and allowing the mixture to precipitate overnight at 4 °C ^[16]^[42]^[46]^[52]^[50]. Due to shielded oppositely charged groups APS are soluble in solvents with higher dielectric constants such as water. ^[13]. However, by using solvent with lower dielectric constant, like ethanol, polysaccharides sulfate groups and positive ions will form ionic bonds, which will result in precipitation. Precipitated polysaccharides can therefore be separated from the supernatant by centrifugation. Supernatant with other non-polysaccharide impurities is discarded while precipitate is then dissolved in water and sometimes treated with CaCl₂ to precipitate alginates. Alginates are then removed by centrifugation, while fucoidan is left in supernatant. Since fucoidan is a macromolecule, supernatant goes through dialysis against water to remove other low molecular weight impurities (laminarin, oligosaccharides and inorganic salts) prior to drying. To isolate fucoidan from the extract January et al. ^[43] and Sahera et al. ^[44] used cationic detergent hexadecyltrimethylammonium bromide (CETAVLON or CTAB). In addition, asfucoidan is a sulfated (SO₃^2-) polysaccharide, it is negatively charged (polyanion) and may form salts with cationic detergents. These salts are highly insoluble in water and hence they will precipitate. Laminarin and alginate are neutral APS and therefore do not react with cationic detergents and remain water soluble ^[55].

After isolation fucoidan needs to be dried with one of the various drying methods such as oven drying, vacuum drying, spray-drying and lyophilisation. Selection of drying method depends on the analysis requirements along with potential use of the extracted fucoidan ^[19]. Although it is quite slow and has relatively low capacity, lyophilisation is usually the preferred method due to its low effect on the fucoidan structure and bioactivity ^[19]. Furthermore, various combinations of chromatographic techniques can be used to achieve high throughput screening and percent purification. For validation purposes, numerous analytical equipment and instrumental techniques can be used to identify and quantify the active fractions of extracted compounds ^[56].

References

Cavalier-Smith, T. Evolution and relationships of algae: Major branches of the tree of life. In Unravelling the Algae the Past, Present, and Future of Algal Systematics; Juliet Brodie, J.L., Ed.; CRC Press: Boca Raton, FL, USA, 2007; pp. 21–55. ISBN 0-8493-7989-X.
Rindi, F. Diversity and Classification of Marine Benthic Algae. Available online: http://marinespecies.org/introduced/wiki/Diversity_and_classification_of_marine_benthic_algae#cite_note-Cavalier-1 (accessed on 24 November 2019).
Xu, S.Y.; Huang, X.; Cheong, K.L. Recent advances in marine algae polysaccharides: Isolation, structure, and activities. Mar. Drugs 2017, 15, 388.
Mišurcová, L.; Orsavová, J.; Ambrožová, J.V. Algal Polysaccharides and Health. In Polysaccharides; Springer: Berlin/Heidelberg, Germany, 2014; pp. 1–29.
Zhao, C.; Yang, C.; Liu, B.; Lin, L.; Sarker, S.D.; Nahar, L.; Yu, H.; Cao, H.; Xiao, J. Bioactive compounds from marine macroalgae and their hypoglycemic benefits. Trends Food Sci. Technol. 2018, 72, 1–12.
Costa, L.S.; Fidelis, G.P.; Cordeiro, S.L.; Oliveira, R.M.; Sabry, D.A.; Câmara, R.B.G.; Nobre, L.T.D.B.; Costa, M.S.S.P.; Almeida-Lima, J.; Farias, E.H.C.; et al. Biological activities of sulfated polysaccharides from tropical seaweeds. Biomed. Pharmacother. 2010, 64, 21–28.
Pereira, L. Seaweeds as source of bioactive substances and skin care therapy-Cosmeceuticals, algotheraphy, and thalassotherapy. Cosmetics 2018, 5, 68.
Jiao, G.; Yu, G.; Zhang, J.; Ewart, H.S. Chemical structures and bioactivities of sulfated polysaccharides from marine algae. Mar. Drugs 2011, 9, 196–233.
Garcia-Vaquero, M.; Rajauria, G.; O’Doherty, J.V.; Sweeney, T. Polysaccharides from macroalgae: Recent advances, innovative technologies and challenges in extraction and purification. Food Res. Int. 2017, 99, 1011–1020.
Ale, M.T.; Mikkelsen, J.D.; Meyer, A.S. Important determinants for fucoidan bioactivity: A critical review of structure-function relations and extraction methods for fucose-containing sulfated polysaccharides from brown seaweeds. Mar. Drugs 2011, 9, 2106–2130.
Praveen, M.A.; Parvathy, K.R.K.; Balasubramanian, P.; Jayabalan, R. An overview of extraction and purification techniques of seaweed dietary fibers for immunomodulation on gut microbiota. Trends Food Sci. Technol. 2019, 92, 46–64.
De Jesus Raposo, M.F.; De Morais, A.M.B.; De Morais, R.M.S.C. Marine polysaccharides from algae with potential biomedical applications. Mar. Drugs 2015, 13, 2967–3028.
Hahn, T.; Lang, S.; Ulber, R.; Muffler, K. Novel procedures for the extraction of fucoidan from brown algae. Process Biochem. 2012, 47, 1691–1698.
Dore, C.M.P.G.; Faustino Alves, M.G.D.C.; Pofírio Will, L.S.E.; Costa, T.G.; Sabry, D.A.; De Souza Rêgo, L.A.R.; Accardo, C.M.; Rocha, H.A.O.; Filgueira, L.G.A.; Leite, E.L. A sulfated polysaccharide, fucans, isolated from brown algae Sargassum vulgare with anticoagulant, antithrombotic, antioxidant and anti-inflammatory effects. Carbohydr. Polym. 2013, 91, 467–475.
Hadj Ammar, H.; Hafsa, J.; Le Cerf, D.; Bouraoui, A.; Majdoub, H. Antioxidant and gastroprotective activities of polysaccharides from the Tunisian brown algae (Cystoseira sedoides). J. Tunis. Chem. Soc. 2016, 18, 80–88.
Hentati, F.; Delattre, C.; Ursu, A.V.; Desbrières, J.; Le Cerf, D.; Gardarin, C.; Abdelkafi, S.; Michaud, P.; Pierre, G. Structural characterization and antioxidant activity of water-soluble polysaccharides from the Tunisian brown seaweed Cystoseira compressa. Carbohydr. Polym. 2018, 198, 589–600.
Mzibra, A.; Meftah Kadmiri, I.; El Arroussi, H. Enzymatic Technologies for Marine Polysaccharides; Trincone, A., Ed.; CRS PRESS: Boca Raton, FL, USA, 2019.
Chaminda Lakmal, H.H.; Lee, J.-H.; Jeon, Y.-J. Enzyme-assisted extraction of a marine algal polysaccharide, fucoidan and bioactivities. In Polysaccharides: Bioactivity and Biotechnology; Springer: Berlin/Heidelberg, Germany, 2015; pp. 1–2241. ISBN 9783319162980.
Lim, S.J.; Wan Aida, W.M. Extraction of sulfated polysaccharides (fucoidan) from brown seaweed. In Seaweed Polysaccharides; Elsevier: Amsterdam, The Netherlands, 2017; pp. 27–46. ISBN 9780128098172.
Nisizawa, K.; Yamaguchi, T.; Handa, N.; Maeda, M.; Yamazaki, H. Chemical nature of a uronic acid-containing polysaccharide in the peritrophic membrane of the silkworm. J. Biochem. 1963, 54, 419–426.
Kadam, S.U.; Tiwari, B.K.; O’Donnell, C.P. Extraction, structure and biofunctional activities of laminarin from brown algae. Int. J. Food Sci. Technol. 2015, 50, 24–31.
Quillet, M. Glucide metabolism of brown algae. Presence of small quantities of Laminarin in numerous new species, distributed over the entire group of Phaeophyceae. Comptes Rendus l’Académie Sci. 1958, 246, 812–815.
Chizhov, A.O.; Dell, A.; Morris, H.R.; Reason, A.J.; Haslam, S.M.; McDowell, R.A.; Chizhov, O.S.; Usov, A.I. Structural analysis of laminarans by MALDI and FAB mass spectrometry. Carbohydr. Res. 1998, 310, 203–210.
Menshova, R.V.; Ermakova, S.P.; Anastyuk, S.D.; Isakov, V.V.; Dubrovskaya, Y.V.; Kusaykin, M.I.; Um, B.H.; Zvyagintseva, T.N. Structure, enzymatic transformation and anticancer activity of branched high molecular weight laminaran from brown alga Eisenia bicyclis. Carbohydr. Polym. 2014, 99, 101–109.
Miao, H.Q.; Elkin, M.; Aingorn, E.; Ishai-Michaeli, R.; Stein, C.A.; Vlodavsky, I. Inhibition of heparanase activity and tumor metastasis by laminarin sulfate and synthetic phosphorothioate oligodeoxynucleotides. Int. J. Cancer 1999, 83, 424–431.
Bae, H.; Song, G.; Lee, J.; Hong, T.; Chang, M.; Lim, W. Laminarin-derived from brown algae suppresses the growth of ovarian cancer cells via mitochondrial dysfunction and ER stress. Mar. Drugs 2020, 18, 152.
Hernández-Carmona, G.; Freile-Pelegrín, Y.; Hernández-Garibay, E. Conventional and Alternative Technologies for the Extraction of Algal Polysaccharides; Woodhead Publishing: Shaston, UK, 2013; ISBN 9780857095121.
Bixler, H.J.; Porse, H. A decade of change in the seaweed hydrocolloids industry. J. Appl. Phycol. 2011, 23, 321–335.
Rasmussen, R.S.; Morrissey, M.T. Marine biotechnology for production of food ingredients. Adv. Food Nutr. Res. 2007, 52, 237–292.
Collado-González, M.; Cristina Ferreri, M.; Freitas, A.R.; Santos, A.C.; Ferreira, N.R.; Carissimi, G.; Sequeira, J.A.D.; Guillermo Díaz Baños, F.; Villora, G.; Veiga, F.; et al. Complex polysaccharide-based nanocomposites for oral insulin delivery. Mar. Drugs 2020, 18, 55.
Burtin, P. Nutritional value of seaweeds. Electron. J. Environ. Agric. Food Chem. 2003, 2, 498–503.
Murata, M.; Nakazoe, J. Production and use of marine algae in Japan. Jpn. Agric. Res. Q. 2001, 35, 281–290.
Kimura, Y.; Watanabe, K.; Okuda, H. Effects of soluble sodium alginate on cholesterol excretion and glucose tolerance in rats. J. Ethnopharmacol. 1996, 54, 47–54.
Kim, I.H.; Lee, J.H. Antimicrobial activities against methicillin-resistant Staphylococcus aureus from macroalgae. J. Ind. Eng. Chem. 2008, 14, 568–572.
Chapman, V.J.; Chapman, D.J. Seaweeds and Their Uses, 3rd ed.; Springer: Dordrecht, The Netherlands, 1980.
Zvyagintseva, T.N.; Shevchenko, N.M.; Chizhov, A.O.; Krupnova, T.N.; Sundukova, E.V.; Isakov, V.V. Water-soluble polysaccharides of some far-eastern brown seaweeds. Distribution, structure, and their dependence on the developmental conditions. J. Exp. Mar. Biol. Ecol. 2003, 294, 1–13.
Zvyagintseva, T.N.; Shevchenko, N.M.; Popivnich, I.B.; Isakov, V.V.; Scobun, A.S.; Sundukova, E.V.; Elyakova, L.A. A new procedure for the separation of water-soluble polysaccharides from brown seaweeds. Carbohydr. Res. 1999, 322, 32–39.
Rioux, L.E.; Turgeon, S.L.; Beaulieu, M. Characterization of polysaccharides extracted from brown seaweeds. Carbohydr. Polym. 2007, 69, 530–537.
Bilan, M.I.; Grachev, A.A.; Ustuzhanina, N.E.; Shashkov, A.S.; Nifantiev, N.E.; Usov, A.I. A highly regular fraction of a fucoidan from the brown seaweed Fucus distichus L. Carbohydr. Res. 2004, 339, 511–517.
Van Weelden, G.; Bobi, M.; Okła, K.; van Weelden, W.J.; Romano, A.; Pijnenborg, J.M.A. Fucoidan structure and activity in relation to anti-cancer mechanisms. Mar. Drugs 2019, 17, 32.
Li, B.; Lu, F.; Wei, X.; Zhao, R. Fucoidan: Structure and bioactivity. Molecules 2008, 13, 1671–1695.
Sellimi, S.; Maalej, H.; Rekik, D.M.; Benslima, A.; Ksouda, G.; Hamdi, M.; Sahnoun, Z.; Li, S.; Nasri, M.; Hajji, M. Antioxidant, antibacterial and in vivo wound healing properties of laminaran purified from Cystoseira barbata seaweed. Int. J. Biol. Macromol. 2018, 119, 633–644.
January, G.G.; Naidoo, R.K.; Kirby-McCullough, B.; Bauer, R. Assessing methodologies for fucoidan extraction from South African brown algae. Algal Res. 2019, 40, 101517.
Sahera, M.F.; Thani, S.M.; Salha, S.Y. Characterization of sulphated polysaccharide with antiviral activity from marine brown alga Cystoseira myrica collected from Jazan coasts, KSA. Int. J. PharmTech Res. 2015, 8, 198–203.
Deghrigue Abid, M.; Lajili, S.; Hadj Ammar, H.; Cherif, D.; Eltaief, N.; Majdoub, H.; Bouraoui, A. Chemical and biological properties of sodium alginates isolated from tow brown algae Dictyopteris Membranaceae and Padina Pavonica. Trends J. Sci. Res. 2019, 4, 62–67.
Sun, Q.L.; Li, Y.; Ni, L.Q.; Li, Y.X.; Cui, Y.S.; Jiang, S.L.; Xie, E.Y.; Du, J.; Deng, F.; Dong, C.X. Structural characterization and antiviral activity of two fucoidans from the brown algae Sargassum henslowianum. Carbohydr. Polym. 2020, 229, 115487.
Zhao, G.; Chen, X.; Wang, L.; Zhou, S.; Feng, H.; Chen, W.N.; Lau, R. Ultrasound assisted extraction of carbohydrates from microalgae as feedstock for yeast fermentation. Bioresour. Technol. 2013, 128, 337–344.
Harun, R.; Yip, J.W.S.; Thiruvenkadam, S.; Ghani, W.A.W.A.K.; Cherrington, T.; Danquah, M.K. Algal biomass conversion to bioethanol-a step-by-step assessment. Biotechnol. J. 2014, 9, 73–86.
Kadam, S.U.; Tiwari, B.K.; O’Connell, S.; O’Donnell, C.P. Effect of ultrasound pretreatment on the extraction kinetics of bioactives from brown seaweed (Ascophyllum nodosum). Sep. Sci. Technol. 2015, 50, 670–675.
Liu, J.; Wu, S.-Y.; Chen, L.; Li, Q.-J.; Shen, Y.-Z.; Jin, L.; Zhang, X.; Chen, P.-C.; Wu, M.-J.; Choi, J.; et al. Different extraction methods bring about distinct physicochemical properties and antioxidant activities of Sargassum fusiforme fucoidans. Int. J. Biol. Macromol. 2019.
Cui, Y.; Liu, X.; Li, S.; Hao, L.; Du, J.; Gao, D.H.; Kang, Q.; Lu, J. Extraction, characterization and biological activity of sulfated polysaccharides from seaweed Dictyopteris divaricata. Int. J. Biol. Macromol. 2018, 117, 256–263.
Fawzy, M.A.; Gomaa, M.; Hifney, A.F.; Abdel-Gawad, K.M. Optimization of alginate alkaline extraction technology from Sargassum latifolium and its potential antioxidant and emulsifying properties. Carbohydr. Polym. 2017, 157, 1903–1912.
Fletcher, H.R.; Biller, P.; Ross, A.B.; Adams, J.M.M. The seasonal variation of fucoidan within three species of brown macroalgae. Algal Res. 2017, 22, 79–86.
Yuan, Y.; Macquarrie, D. Microwave assisted extraction of sulfated polysaccharides (fucoidan) from Ascophyllum nodosum and its antioxidant activity. Carbohydr. Polym. 2015, 129, 101–107.
Scott, J.E. Fractionation by precipitation with quaternary ammonium salts. In Methods in Carbohydrate Chemistry; Whistler, R.L., BeMiller, J.N., Eds.; Academic Press: New York, NY, USA, 1965; pp. 38–44.
Sosa-Hernández, J.E.; Escobedo-Avellaneda, Z.; Iqbal, H.M.N.; Welti-Chanes, J. State-of-the-art extraction methodologies for bioactive compounds from algal biome to meet bio-economy challenges and opportunities. Molecules 2018, 23, 2953.

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