Open access peer-reviewed chapter

Antioxidant Activity of Sulfated Seaweeds Polysaccharides by Novel Assisted Extraction

Written By

Shao-Chi Wu

Submitted: December 11th, 2016 Reviewed: May 8th, 2017 Published: November 29th, 2017

DOI: 10.5772/intechopen.69633

Chapter metrics overview

2,274 Chapter Downloads

View Full Metrics


Seaweeds have an extremely numerous of species in the world and been able to be divided into several developmental systems. Broadly, three types of seaweeds can be defined according to their color: brown seaweeds, green seaweeds, and red seaweeds. Thousands of years ago, mankind used seaweeds as food and medicine. Seaweed extracts are gaining increasing attention due to their unique composition and the potential for widespread use in industry. A variety of novel (green) extraction techniques have been devised for converting seaweed biomass into seaweed extracts, such as enzyme-assisted extraction (EAE), microwave-assisted extraction (MAE), pressurized liquid extraction (PLE), supercritical fluid extraction (SFE), and ultrasound-assisted extraction (UAE), which are capable of extracting seaweeds’ biologically active compounds without causing degradation. Seaweed extracts contain compounds, such as carbohydrates, proteins, minerals, oils, fats, and polyunsaturated fatty acids and abundant bioactive compounds, such as antioxidants, pigments, and sulfated seaweed polysaccharides (SWP), as well as antibacterial, antifungal, anti-inflammatory, antioxidation, antitumor, antiviral, and hypolipidemic effect. The purpose of this article is to describe the antioxidant activity of SWP of brown seaweeds, green seaweeds, and red seaweeds by novel assisted extraction.


  • antioxidant activity
  • assisted extraction
  • sulfated seaweed polysaccharides

1. Introduction

Edible seaweeds are a good source of antioxidants, dietary fibers, essential amino acids, vitamins, phytochemicals, polyunsaturated fatty acids, and minerals [1]. Most seaweed polysaccharides (SWP) are derived from natural sources, such as agarose, alginates, carrageenan, fucoidan, porphyran, and ulvan. Seaweed polysaccharide is a very important biological macromolecule in marine life. Seaweed polysaccharides exhibit a wide range of structure and are still underexploited and should therefore be considered as a novel source of natural products for pharmaceutical discovery [2].

Seaweeds are plant-like organisms that generally live attached to rock or other hard substrata in coastal areas. They can be classified into three different groups, empirically distinguished since the mid-nineteenth century on the basis of thallus color: brown seaweeds (Phaeophyceae), green seaweeds (Chlorophyceae), and red seaweeds (Rhodophyceae) [3, 4]. Among marine resources, seaweeds, which are sometimes known as algae, are well-known natural sources of polysaccharides. SWP are well-known natural seaweed sources of polysaccharides which have considerable numerous bioactive compounds with significant biological features. SWP are most common in the seaweed cell walls, and their number and chemical structure are varying according to the specific seaweeds’ species [5]. Bioactive SWP extracted from seaweeds can be classified into three types. The major SWP found in brown seaweeds were fucan and fucoidan; green seaweeds were sulfated rhamnans and ulvan; red seaweeds were galactan and carrageenan [6]. It displays several physicochemical and biological features of potential interest for agricultural, chemical, food, and pharmaceutical applications [7].

The purpose of this chapter is to understand the potential applications of SWP in antioxidant activity of brown seaweeds, green seaweeds, and red seaweeds by enzyme-assisted extraction (EAE), microwave-assisted extraction (MAE), pressurized liquid extraction (PLE), supercritical fluid extraction (SFE), and ultrasound-assisted extraction (UAE), also named green-assisted extraction in recent literary.


2. Novel assisted extraction

Marine plant materials are increasingly being used to isolate and purify bioactive compounds; and then recent studies have reported on the antioxidant potential of SWP of seaweeds [8, 9]. Traditional techniques involve application of solid-liquid extraction (SLE) simply by means of solvent application and leaching. A domestic application of conventional solvent extraction (CSE) is quite familiar to everybody in daily life from making of coffee or tea at home. SLE encompasses conventional methods: Soxhlet extraction (SE), percolation, and maceration extraction (ME). These techniques have been utilized for more than a century for the separation of SWP. However, certain disadvantages pertaining to CSE render its application quite uneconomically due to excessive consumption of energy, polluting solvents, and time. These underlying drawbacks have triggered research that explores more cost-effective and greener techniques for the extraction of SWP from a wide range of seaweed matrices [10]. They include EAE, MAE, PLE, SFE, and UAE techniques [11]. The advantages and disadvantages of these novel assisted extraction methods are shown in Table 1.

Novel assisted extraction methods Advantages Disadvantages References
EAE 1. Non-consumed during reaction
2. High conversion yield
3. Nontoxic and biodegradable
4. High selectivity
5. High specificity
6. Utilization in soft conditions
7. Large-scale production
1. Stability
2. Cost
MAE 1. Decreased in extraction time
2. To avoid the loss of volatile substances during microwave
3. Less solvent is required because no evaporation occurs
4. No hazardous fumes during acid microwave since it is a closed vessel
1. High pressure used poses safety risks
2. The usual constituent material of the vessel does not allow high solution temperatures
3. Addition of reagents is impossible sine it is a single-step procedure
4. Vessel must be cooled down before it can be opened to prevent loss of volatile constituents
PLE 1. Better for increased operating temperature
2. Increase selectivity
3. Precision and reproducibility
4. Reduces oxidation risk
5. Relatively simple compared to SFE
6. Shorter extraction time
7. Reduced solvent consumption
8. Possibility for automation
1. Thermal degradation for thermolabile compounds is a cause for concern
2. Selectivity is mainly affected by varying the solvent type
3. Post-extraction cleanup step is still necessary
SFE 1. Enhanced extraction efficiency
2. Tunability of the solvent strength
3. Low organic solvent consumption
4. Preservation of bioactive properties
5. Organoleptic properties of the extracts
6. Inline integration with sample preparation and detection methods
1. High-capital investment
2. Large number of variables to optimize
3. Strong dependence on matrix analyte interactions
4. Difficulties in scale-up
5. Difficulties in technology transfer
6. Difficulty in implementing continuous extraction processes
7. Difficulty of extracting more polar compounds
UAE 1. Easy to use
2. Short time of extraction
3. Small amount of solvent
1. No good recoveries for most PCB congeners
2. Expensive system

Table 1.

Advantages and disadvantages of novel assisted extractions.

2.1. EAE

Enzymes can be derived from animal organs, bacteria, fruit extracts, fungi, or vegetable. All these known enzymes are classified according to six basic groups. These categories are organized according to how the enzyme works on a molecular level. These six types of enzymes are as follows: hydrolases, isomerases, ligases, lyases, oxidoreductases, and transferases. Hydrolases are the most common type, followed by oxidoreductases and transferases. Enzymes are ideal catalysts that can assist in the extraction of complex bioactive compounds of natural origin by degrading the plant cell walls and membranes. Consequently, they increase plant cell wall permeability, and thus higher extraction yields of bioactive compounds are achieved [12].

Seaweed cell walls are composed of a diverse array of fibrillar, matrix, and crystalline polymers, that is, sulfated and branched polysaccharides, interacting with proteins, various ions, and water. It is necessary to break down seaweed cell walls with enzymes that can be used to remove the cell wall specifically under optimal temperature and pH conditions and then get the desired bioactive compounds [11], depending on what organism you work with, which can be agarase [6], Alcalase [13], carragenanase [14], Celluclast [15], Kojizyme [16], Neutrase [17], Termamyl [18], Ultraflo [19], Umamizyme [20], Viscozyme [21], and xylanase [22].

2.2. MAE

MAE is a novel technique that has many advantages including high extraction efficiency, low solvent consumption, high-purity extracts, and shortened extraction time, which make it well suited for the extraction of bioactive compounds from plant materials [29]. The mechanism of microwave volumetric heating involves the inherent ionic conduction and dipolar relaxation inside a dielectric material. Microwave irradiation induces the rapid elevating temperature of solvent to accelerate the diffusion of pure solvent into plant matrix, as well as the dissolution of the targeted compound into solvent [30]. Microwave energy penetration causes quick elevation of temperature to build the internal pressure inside the cell of plant material. The high interior pressure may destroy the cell wall of plant material to easily release bioactive compounds into solvents [26]. High temperature would cause the dehydration of cellulose and reduce its mechanical strength in MAE, which promotes the solvent to penetrate into the cellular channels and subsequently increase the extraction yield [31].

Microwave is an electromagnetic radiation with wavelengths ranging from 1 m to 1 mm, with frequencies between 300 MHz (100 cm) and 300 GHz (0.1 cm), which can be transmitted as the wave [32]. When microwave passes through the seaweed medium, its energy may be absorbed and converted into thermal energy. Heating using microwave energy is based on two principles: (1) ionic conduction refers to the electrophoretic migration of the charge carriers (e.g. ions and electrons) under the influence of the electric field produced by microwave [33] and (2) dipole rotation happens when the dipolar molecules attempt to follow the electric field in the same alignment [34].

There are two main types of MAE systems available for industrial and commercial applications for natural product extraction: (1) the closed-vessel system and (2) the open-vessel system [10]. In the closed-vessel system, extraction is carried out under controlled conditions of temperature and pressure. This is generally employed for extractions under extremely high-temperature conditions. Diffused microwaves from a cavity magnetron radiate in all directions to interact with plant samples placed in extraction vessels in a closed-vessel chamber. Owing to the even dispersion of microwaves, this technique is also known as the multimode system [35]. In the open-vessel system, also known as the monomode system, the extraction vessel is partially exposed to microwave radiation (focused radiation). A circular metallic waveguide directs the focused microwaves toward the extraction vessel inside the microwave (monomode) cavity. This interaction promotes the initiation of mass transfer between the solute and extractant upon solvation [36].

2.3. PLE

PLE is another novel assisted extraction technique for natural product extraction from food and botanical sample matrices. The Dionex Corporation was the first to introduce the PLE technique as an accelerated solvent extraction (ASE) technology (ASER®) in 1995 [37]. PLE is called accelerated solvent extraction (ASE), enhanced solvent extraction (ESE), pressurized fluid extraction (PFE), or pressurized solvent extraction (PSE). When the solvent used is water, it is common to use other terms, such as high-temperature water extraction (HTWE), hot liquid water extraction (HLWE), hot water extraction (HWE), pressurized hot water extraction (PHWE), subcritical water extraction (SWE), or superheated water extraction (SWE) [38].

The principle of the PLE is based on using elevated temperatures (50–200°C) and pressures (50–150 atm) to extract analytes from solid or semisolid samples within short periods of time (5–15 min). PLE is similar to Soxhlet extraction, except that during the extraction process, the solvent condition inside the PLE cell approaches the supercritical region which results in more efficient extractions. Depending on the temperature at which the extraction is performed, PLE allows the sample to become more soluble and achieve a higher diffusion rate, while the elevated pressure keeps the solvent below its boiling point [39]. PLE permits high extraction efficiency with a low solvent volume (15–40 mL) and a short extraction time (15–20 min). PLE has used less solvent in a shorter period of time, and in oxygen and light-free environment, it has the potential to be a powerful tool in industry [4042].

2.4. SFE

SFE is the process of separating one component (the extractant) from another (the matrix) using supercritical fluids as the extracting solvent [43]. The supercritical fluid state occurs when a fluid is above its critical temperature (Tc) and critical pressure (Pc), when it is between the typical gas and liquid state. Manipulating the temperature and pressure of the fluid can solubilize the material of interest and selectively extract it. The unique physical properties of supercritical fluids have values for density, diffusivity, and viscosity between liquids and gases. Moreover, near-zero surface tension as well as low viscosities similar to gases allow supercritical fluids to easily penetrate into a microporous matrix material to extract desired compounds [44].

Carbon dioxide (CO2) is the most used supercritical fluid, sometimes modified by cosolvents, such as ethanol or methanol. Since the critical temperature and critical pressure of CO2 are only 31°C and 74 bar, extraction is done at temperatures that will not damage heat labile molecules, and the absence of oxygen minimizes oxidation. Thus, an extraction process can take 10–60 min with SFE, while it would take hours or even days with classical methods [45, 46]. On the other hand, it is to be noticed that no organic residue is found both in extract and solid residue and no thermal degradation appears, which results in very high-quality products [47].

2.5. UAE

Ultrasound is sound waves with frequencies higher than the upper audible limit of human hearing. This limit varies from person to person and is approximately 20 kHz (20,000 Hz) in healthy young adults. Ultrasound devices operate with frequencies from 20 kHz up to several GHz [48]. Sound waves produced by an ultrasonic probe allow greater penetration of solvent into the seaweeds, and ultrasonic power also produces high-energy cavitation bubbles containing solvent vapor. The bubbles implode near seaweed walls causing very high local temperatures, pressure increase, and seaweed cell wall destruction, which eases mass transfer from cell to solvent and enhances microstreaming [49]. This effect is much stronger at low frequencies (18–40 kHz) [50].

UAE has emerged as a promising technique that fulfills the required criteria as an inexpensive green extraction technique providing higher recovery of targeted compounds with lower solvent consumption and/or faster analysis and bioactivity properties. Notable UAE features include cost-effectiveness, eco-friendliness, rapidity, simplicity, safety, and versatility, due to the reduced consumption of time, energy, and expensive organic solvents, which is in contrast to traditional extraction techniques [10].


3. Sulfated seaweed polysaccharides (SWP)

Seaweeds are the most abundant source of nonanimal in nature. SWP from some seaweeds have become very important products in the food industry and also possess biological activity of potential medicinal value, such as anti-allergy, anticancer, anticoagulant, anti-inflammation, antioxidant, and antiviral [9, 5153]. SWP are commonly found in three major groups of seaweeds: brown seaweeds (Phaeophyta), green seaweeds (Chlorophyta), and red seaweeds (Rhodophyta). The major SWP of brown seaweeds are fucans, including fucoidan, sargassan, ascophyllan, and glucuronoxylofucan; and those of red seaweeds are galactans commercially known as agar and carrageenan. On the other hand, the major SWP of green seaweeds are usually sulfated heteropolysaccharides that contain galactose, xylose, arabinose, mannose, glucuronic acid, or glucose [54].

3.1. Brown seaweeds

There are about 1800 species of brown algae, and most are marine. In general, brown algae are larger and more species are found in colder waters. Brown seaweeds are usually grown or collected for food consumption and especially known for their health benefits and high nutritional value, such as kombu or haidai (Laminaria japonica), wakame or quandai-cai (Undaria pinnatifida), hijiki (Hizikia fusiforme), and mozuku (Cladosiphon okamuranus). The major fucoidan yielding brown seaweed genera are Fucus, Sargassum, Laminaria, and Undaria [55].

The term fucoidan is commonly applied for complex SWP, often isolated from seaweeds, mainly containing fucose residues but also many other monosaccharides [7]. Furthermore, fucoidan has a backbone built of (1→3)-linked α-l-fucopyranosyl or of alternating (1→3)- and (1→4)-linked α-l-fucopyranosyl residues, also including sulfated galactofucans with backbones built of (1→6)-β-d-galacto- and/or (1→2)-β-d-mannopyranosyl units with fucose or fuco-oligosaccharide branching and/or glucuronic acid, xylose, or glucose substitutions. There are at least two distinct forms of fucoidan: U-fucoidan, which is approximately 20% glucuronic acid, and F-fucoidan, which is >95% composed of sulfated esters of fucose [56].

Fucoidans with greater molecular masses and higher degrees of sulfatation form solutions of higher viscosity. Adding glycerol and diols also leads to a significant increase in viscosity [57]. Rheological characteristics of fucoidan from C. okamuranus showed shear thinning behavior below 1.5% (W/V) but plastic behavior at 2.0% (W/V). The dynamic viscoelasticity of the fucoidan increased linearly with an increase in concentration and decreased gradually with increase in temperature [58]. Fucoidan with covalent linkage of bovine serum albumin had emulsifying properties, high solubility after heating, and high melting temperature [59]. Crude fucoidan from Sargassum sp. demonstrated good emulsion-stabilizing capacities, especially with cedar wood oil and xylene [60].

3.2. Green seaweeds

Green seaweeds are usually grown or collected for food consumption and especially known for their health benefits and high nutritional value, such as aonori, hirohano-hitoegusa nori, or hitoegusa-nori (Monostroma spp.) or green laver (Enteromorpha spp.). SWP from green seaweeds can be found in Caulerpa (sulfated galacotans), Codium (sulfated arabinogalactans), Enteromorpha (ulvans), Monostroma (sulfated rhamnans), and Ulva (ulvans) [61].

The structural diversity of SWP found in seaweeds varies with species. Water-soluble extract polysaccharides from Caulerpa are mainly composed of glucans and SWP. Heteropolysaccharides (SWP) from Caulerpa consist of different monosaccharides, such as galactose, glucose, mannose, and xylose. Among them, galactose is the major sugar source, while glucose, mannose, and xylose are common components. The water-soluble fraction obtained from Caulerpa sertularioides with antimicrobial effects which grown under natural conditions contains sulfated galactans constituted of (1→3)-β-d-Gal and (1→6)-β-d-Gal units, and sulfation is observed to occur at the C-2 position of the residues [62]. A water extraction of SWP from Codium divaricatum with anticoagulant activity is a galactan which is highly sulfated and substituted with pyruvic acid ketals was mainly constituted of (1→3)-β-d-galactopyranose residues, branched by single (1→)-β-d-galactopyranose units, and the backbone of CP2-1 attached to the main chain at C-4 positions [63]. Monostroma nitidum extracted with boiling water could obtain rhamnan sulfate with antithrombin active that consists of α-1,3-linked l-rhamnose residues, some substituted with sulfate groups mainly at position O-2. Minor amounts were also exist internal 1,2-linked rhamnose and branched rhamnose linkages [64]. The structure of an ulvan with anticancer obtained by water extraction from Ulva lactuca consists of galactose, glucose, mannose, rhamnose, xylose, uronic acid, and sulfate content. This ulvan is mainly composed of disaccharide [→4)-β-d-GlcA-(1→4)-α-l-Rha3S-(1→] and other minor disaccharides β-GlcA-(1→2)-α-Xyl and β-GlcA-(→2)-α-Rha [65].

Dielectric properties of aqueous solutions from ulvan (U. meridionalis) and rhamnan sulfate (M. lattisimum) with H+-form hydrocolloids possess significant improvement in hydration function [66]. Ulvan of Ulva armoricana and Ulva rotundata showed that chemical structure, macromolecular characteristics, and rheological properties were affected by both species and seasons. The proportion of high-molecular-weight ulvan was the major factor positively correlated with the gelling properties [67]. Ulvan from Ulva fasciata in different ionic strengths (Na+ and Ca2+) had significant effects on the stability of o/w emulsions [68]. The rheological properties and zeta-potential of the emulsions appeared to be dependent on the ulvan concentration. The emulsifying and stabilizing mechanism of the ulvan may ascribe to its surface-active protein moiety, also to the hydrophobicity of the polysaccharide itself [69].

3.3. Red seaweeds

Red seaweeds are usually grown or collected for food consumption and especially known for their health benefits and high nutritional value, such as nori or purple lave (Porphyra spp.), ogo, ogonori, or sea moss (Gracilaria spp.) The red seaweeds Gracilaria and Gelidium are used in the manufacture of the agar, Kappaphycus and Betaphycus are now the most important sources of carrageenan, and Porphyra could extract porphyran for food and biotechnological applications [70].

Carrageenan are high-molecular-weight sulfated d-galactans composed of repeating disaccharide units with alternating 3-linked β-d-galactopyranose and 4-linked α-galactopyranose or 3,6-anhydro-α-galactopyranose [7]. There are at least 15 different carrageenan structures, including but not limited to κ-, ι-, λ-, μ-, θ-, β-, and ν-carrageenan [20]. Porphyran has hypolipidemic and antiallergic pharmacological property applications, is a complex SWP, and is consist of about 30% of agarose repeating units (1,3-linked β-D-galactopyranose followed by 1,4-linked 3,6-anhydro-α-l-galactopyranose), with the remaining residues being 3-linked β-d-galactopyranose and 4-linked α-l-galactopyranose-6-sulfate. The composition includes 6-O-sulfated l-galactose, 6-O-methylated d-galactose, l-galactose, 3,6-anhydro-l-galactose, 6-O-methyl d-galactose, and ester sulfate. Some of the ester is present as 1-4-linked l-galactose 6-sulfate [71].

Kappa carrageenan showed that shear stress-shear rate and viscosity curves clearly indicated sharp increases in viscosity and consistency coefficient (k), and addition of KCl was more effective in increasing viscosity [72]. Kappa-carrageenan film incorporation of plant oils increased the film thickness and plasticizing effect significantly. However, the moisture content, solubility, and tensile strength of films decreased significantly when plant oils were added [73]. Six formulations carrageenan/porphyran films from Pyropia columbina were homogeneous and flexible. Their moisture content, water solubility, and water vapor permeability significantly increased by glycerol content increasing could obtain more stretchable but less resistant films. When Ca+2 addition could mask glycerol effect in water solubility and water vapor permeability, it could stabilize the three-dimensional structure of carrageenans/porphyrans by interactions between sulfate groups, promote water retention, and open film structure [74]. Porphyran, alkali-treated porphyrin, and sulfated porphyran from Porphyra haitanensis demonstrate that the anticoagulant activities mainly depend on the position of sulfate and the antioxidant activities mainly depend on degree of sulfate substitution [75]. The rheological behavior of porphyran exist pseudoplastic behavior, which agrees with the Herschel-Bulkley model. The effect of temperature on the viscosity of porphyrans was increasing concentrations from 3 to 7% [76].


4. Antioxidant activity of SWP

Reactive oxygen species (ROS) is produced in the forms of H2O2, superoxide radical (O·̄2), hydroxyl radical (•OH) and nitric oxide (NO) in the organisms are produced by non-enzymatic and enzymatic reactions. Moreover, endogenous antioxidant enzymes and antioxidants in the body under synergy effect would been removed these ROS. But when the body is aging and experiences illness or fatigue, the body’s free radicals may be destroyed, and excess oxygen free radicals cause a series of oxidative damage to the body. Excessive free radicals can cause irreversible damage to the body. It can cause damage to the body at molecular level, cell level, and level of tissue and organs by attacking the life molecules and various kinds of cells [77]. Oxidative stress and disease have resulted in identification of important oxidative stress biomarkers—the products of oxidation of biological molecules: DNA, lipids, and proteins. It would lead to many health disorders including cancer, diabetes, cardiovascular, inflammatory, and neurodegenerative diseases [78]. SWP from a number of seaweeds has appreciable antioxidant capability. Antioxidant activity of SWP has been determined by various in vitro methods, such as ABTS radical scavenging [79], 1,1-diphenyl-2-picrylhydrazyl (DPPH) radical scavenging [80], ferric reducing antioxidant power (FRAP) [81], lipid peroxide inhibition [82], NO scavenging [83], and superoxide and hydroxyl radical scavenging assays [84, 85].

4.1. SWP by EAE

Enzymes are more action specific and operate at lower temperature during hydrolysis; thus, EAE is more amicable for degradation of biological materials without damaging the bioactive compounds. The extract of Ecklonia cava (brown seaweed) obtained using Celluclast was compared with extract obtained by organic solvents. Methanol extract, which gave the highest radical scavenging activities among the organic solvents investigated, was 20% less than the extract obtained by Celluclast [86, 87].

EAE was performed using Alcalase, cellulase, flavourzyme, and Viscozyme L applied on Sargassum muticum (brown seaweed), Codium tomentosum (green seaweed), and Osmundea pinnatifida (red seaweed). SWP with higher extraction yields were observed for Co. tomentosum EAE (for cellulase and Viscozyme L), followed by O. pinnatifida (except Alcalase) and S. muticum. A higher effect on hydroxyl-radical scavenging activity (35–50%) was observed for all SWP extracts [88].

The extracts of U. armoricana (green seaweed) were determined to assess the efficiency of endo-protease treatments which significantly increased the extraction yields compared to the control. The organic matter, neutral sugar, and protein contents were increased in all extracts compared to an extraction with water, up to 2.0-, 2.7-, and 1.75-fold, respectively. Free radical scavenging capacity at medium inhibition concentrations (IC50) of 1.8 and 12.5 mg/mL was shown for the extracts produced with endo-protease treatments and 6.0 mg/mL for the sample resulting from the extraction with the multiple mix of glycosyl hydrolases [89]. Enzyme-assisted extraction of Un. pinnatifida (green seaweed) was performed using five proteases (Alcalase, Flavourzyme, Neutrase, trypsin, and Protamex) and six carbohydrases (AMG, Dextrozyme, maltogenase, Promozyme, Viscozyme, and Celluclast). SWP of Un. pinnatifida exhibited strong radical scavenging activity on DPPH and hydroxyl radical, and activity increased with increasing concentration [90].

SWP were extracted from Pterocladia capillacea (red seaweed), and the produced fractions were hydrolyzed using different enzymes, such as Viscozyme L (a multienzyme complex containing arabanase, cellulase, β-glucanase, hemicellulase, and xylanase), β-glucanase from Trichoderma longibrachiatum, and β-galactosidase from Kluyveromyces lactis. The Viscozyme hydrolysate exhibited DPPH scavenging activity of about 92% with more than 50% increase over its mother fraction [27]. Three SWP were obtained from the Mastocarpus stellatus (red seaweed) using Alcalase hydrolysis at three different treatments. The Alcalase enzymatic hydrolysis at 50°C provided films with reducing power and radical scavenging capacity, which did not change as a result of subsequent heat treatment at 90°C. Their viscoelastic properties of the film-forming solutions showed improved gel-like behavior when the κ/ι-hybrid carrageenan extraction at 90°C was promoted [91].

4.2. SWP by MAE

SWP (fucoidan) from Ascophyllum nodosum (brown seaweed) were extracted by MAE technology. Different conditions of temperature (90–150°C) and extraction time (5–30 min) were evaluated, and optimal fucoidan yield was 16.08%, obtained from 120°C for 15 min of extraction. All SWP from A. nodosum exhibited antioxidant activities as measured by DPPH scavenging and reducing power, among which SWP extracted at 90°C was the highest [62]. SWP were extracted from Fucus vesiculosus (brown seaweed) seaweed by using MAE. The SWP obtained by MAE that contained 53.8 mole% of fucose, 35.3 mole% of xylose, and 10.8 mole% of galactose presented comparable values of antioxidant activity by the DPPH, ABTS+, and lipid oxidation inhibition methods [92].

Six representative (molecular weight 446.5, 247.0, 76.1, 19.0, 5.0, and 3.1 kDa) SWP from Enteromorpha prolifera (green seaweed) were extracted by MAE. All samples showed that great inhibitory effects on superoxide radical at a low concentration and high molecular weight exhibited higher inhibitory effects. Otherwise, samples with low molecular weight possessed stronger inhibitory effects on hydroxyl radical; IC50 of molecular weight 3.1 kDa was 0.39 mg/mL. The chelating effect of molecular weight 3.1 kDa was 77.3% at 5 mg/mL, which was twice more than initial polysaccharide [93].

Four SWP were extracted from Durvillaea antarctica (brown seaweed), Sarcodia ceylonensis (brown seaweed), U. lactuca L. (green seaweed), and Gracilaria lemaneiformis (red seaweed), respectively, by MAE. The average molecular weight of SWP of D. antarctica, S. ceylonensis, U. lactuca L., and G. lemaneiformis was 482, 466, 404, and 591 kDa, respectively. The in vitro antioxidant activity of all of the polysaccharides was evaluated using ABTS+, hydroxyl radical, nitrite scavenging capacity, and reducing power. SWP of U. lactuca L. presented the highest ABTS radical scavenging activity; SWP of U. lactuca L., S. ceylonensis, and D. antarctica also showed a strong effect on the ABTS radical scavenging activity. SWP of U. lactuca L. and S. ceylonensis exhibited excellent hydroxyl-radical scavenging activities, about 83.33% ± 2.31% and 80.07% ± 2.17%, respectively, at 4 mg/mL. The reducing power of SWP of D. Antarctica was relatively more pronounced than that of the three other polysaccharides. However, the nitrite scavenging activities of the four seaweed polysaccharides were weaker than other antioxidant activities (ABTS), hydroxyl radical scavenging capacities, and reducing powers [94].

SWP from Porphyra dentate (red seaweed) is adjusting the pH value of the ethanol solutions used as extraction solvents and then applying continuous or intermittent MAE to extract P. dentate solutions. The SWP content was significantly affected by ethanol concentration, pH value of the ethanol solution, and intermittency at a 1% significance level. It also showed high antioxidant activities by the DPPH and FRAP. Gelling property of the extracted SWP was not affected [95].

4.3. SWP by PLE

PLE was utilized to extract SWP from Saccharina japonica (brown seaweed). Various conditions of temperature (80–200°C), pressure (5–100 bar), and solvents (water, 0.1% sodium hydroxide, 0.1% formic acid, 70% ethanol, 50% ethanol, and 25% ethanol) were assessed; the best crude SWP yield was 8.23%, obtained from 140°C and 50 bar (sodium hydroxide). All crude SWP showed antioxidant activities as measured by DPPH radical and ABTS+ radical scavenging. Crude SWP demonstrates good emulsion-stabilizing capacities, especially with vegetable oils [96].

4.4. SWP by SFE

Gracilaria mammillaris (red seaweed) from the Colombian Caribbean coast were investigated as a source of extracts with antioxidant activity, by means of supercritical CO2 extraction with ethanol as cosolvent. A central composite design was used to investigate the effects of pressure (10, 20, and 30 MPa), temperature (40, 50, and 60°C), and cosolvent concentration (2, 5, and 8%) on antioxidant activity. The antioxidant activity of samples was evaluated by determining their capacity for protecting an edible oil against oxidation, upon accelerated oxidation trials. The extracts obtained at 30 MPa, 60°C, and 8% cosolvent showed the highest antioxidant activity, inhibiting 42.1% in the formation of TBARS after 6 days of accelerated oxidation [97].

4.5. SWP by UAE

UAE was carried out in a water bath ultrasonicator for 60 min (sonicate for 10 min and pause for 2 min) at 50°C applied on S. muticum (brown seaweed), Co. tomentosum (green seaweed), and O. pinnatifida (red seaweed). A higher effect on hydroxyl-radical scavenging activity (35–50%) was observed for all SWP [88]. The SWP from Gracilaria birdiae (red seaweed) were obtained using five different extraction conditions. Their infrared and electrophoresis analysis showed that all conditions extracted the same SWP. The total capacity antioxidant of the SWP was also affected by extraction condition, since GB2s (NaOH/sonication/proteolysis extraction) and GB1 (water extraction) showed lower activity in comparison to the other conditions. The data revealed that NaOH/sonication/proteolysis was the best condition to extract antioxidant SWP from G. birdiae [98].

In general, the antioxidant properties of SWP are influenced by chemical characteristics like molecular weight, degree of branching, type of monosaccharides, ratio of monosaccharides, intermolecular associations of polysaccharides, glycosidic branching, and modification of polysaccharides. It has been observed that crude polysaccharide has better antioxidant activity than purified polysaccharide components. Antioxidant activity of SWP depends on their structural features, such as degree of sulfating, molecular weight, type of the major sugar, and glycosidic branching. The rationale is that low-molecular-weight SWP may incorporate into the cells more efficiently and donate protons more effectively than high-molecular-weight SWP. Evidence suggests that SWP are higher useful candidates when searching for effective nontoxic substances with potential antioxidant activity among various naturally occurring compounds. Therefore, SWP could be used as a rich source of natural antioxidants with potential application in the food industry as well as in cosmetic and pharmaceutical areas [99].


5. Conclusions

Seaweeds are used in many maritime countries as a source of human food, for industrial applications, and as a fertilizer. The major utilization of these plants as functional food is in Asia, particularly China, Japan, and Korea. They have the potential to be used as a source of long- and short-chain chemicals with medicinal and industrial uses. Seaweeds are a rich source of amino acids, bioactive peptides, carotenoids, dietary fiber, minerals, omega-3 fatty acids, SWP, and vitamins. Among them, SWP have the potential to significantly improve extraction efficiency. SWP include a complex group of macromolecules with a wide range of important biological activities, such as antioxidant, anticoagulant, anticancer, antiviral, anti-allergy, and anti-inflammation. Future research priorities in this area should be concentrated on overcoming the challenges of employing these novel technologies on chemical characteristics of SWP.


  1. 1. Roohinejad S, Koubaa M, Barba, FJ, Saljoughian S, Amid M, Greiner R. Application of seaweeds to develop new food products with enhanced shelf-life, quality and health-related beneficial properties. Food Research International. 2016. (In Press) DOI: 10.1016/j.foodres.2016.08.016
  2. 2. Manivasagan P, Oh J. Marine polysaccharide-based nanomaterials as a novel source of nanobiotechnological applications. International Journal of Biological Macromolecules. 2016;82:315-327. DOI: 10.1016/j.ijbiomac.2015.10.081
  3. 3. Fernando IP, Nah, JW, Jeon, YJ. Potential anti-inflammatory natural products from marine algae. Environmental Toxicology and Pharmacology. 2016;48:22-30. DOI: 10.1016/j.etap.2016.09.023
  4. 4. McHugh DJ. A Guide to the Seaweed Industry; FAO Fisheries Technical Paper 441. Rome, Italy: Food and Agriculture Organization of the United Nations; 2003
  5. 5. Fimbres-Olivarría D, López-Elías JA, Carvajal-Millán E, Márquez-Escalante JA, Martínez-Córdova LR, Miranda-Baeza A, Enríquez-Ocaña F, Valdéz-Holguín JE, Brown-Bojórquez F. Navicula sp. sulfated polysaccharide gels induced by Fe(III): Rheology and microstructure. International Journal of Molecular Sciences. 2016;17:1-10. DOI: 10.3390/ijms17081238
  6. 6. Gurpilhares DB, Moreira TR, Bueno JL, Cinelli LP, Mazzola PG, Pessoa A, Sette LD. Algae’s sulfated polysaccharides modifications: Potential use of microbial enzymes. Process Biochemistry. 2016;51:989-998. DOI: 10.1016/j.procbio.2016.04.020
  7. 7. Cunha L, Grenha A. Sulfated seaweed polysaccharides as multifunctional materials in drug delivery applications. Marine Drugs. 2016;14:42. DOI: 10.3390/md14030042
  8. 8. Di T, Chen GJ, Sun Y, Ou SY, Zeng XX, Hong Y. Antioxidant and immunostimulating activities in vitro of sulfated polysaccharides isolated from Gracilaria rubra. Journal of Functional Foods. 2017;28:64-75. DOI: 10.1016/j.jff.2016.11.005
  9. 9. Wang JH, Zhang BW, Luo JP. Molecular weight, chain conformation and antioxidant activities of sulfated β-d-galactan derivatives from Dendrobium nobile Lindl. Current Topics in Nutraceutical Research. 2015;13:205-212
  10. 10. Ameer K, Shahbaz HM, Kwon JH. Green extraction methods for polyphenols from plant matrices and their byproducts: A review. Comprehensive Reviews in Food Science and Food Safety. 2017;16:295-315. DOI: 10.1111/1541-4337.12253
  11. 11. Kadam SU, Tiwari BK, O’Donnell CP. Application of novel extraction technologies for bioactives from marine algae. Journal of Agricultural and Food Chemistry. 2013;61:4667-4675. DOI: 10.1021/jf400819p
  12. 12. Cheng X, Bi LG, Zhao ZD, Chen YX. Advances in enzyme assisted extraction of natural products. In: 3rd International Conference on Material, Mechanical and Manufacturing Engineering (IC3ME 2015); 27-28 June, 2015; Amstelkade, Amsterdam, Netherlands. Atlantis Press; 2015. pp. 371-375
  13. 13. Wijesinghe WAJ, Jeon YJ. Enzyme-assistant extraction (EAE) of bioactive components: A useful approach for recovery of industrially important metabolites from seaweeds. Fitoterapia. 2012;83:6-12. DOI: 10.1016/j.fitote.2011.10.016
  14. 14. Jiao GL, Yu GL, Zhang JZ, Ewart HS. Chemical structures and bioactivities of sulfated polysaccharides from marine algae. Marine Drugs. 2011;9:196-223. DOI: 10.3390/md9020196
  15. 15. Michalak I, Chojnacka K. Algal extracts: Technology and advances. Engineering in Life Sciences. 2014;14:581-591. DOI: 10.1002/elsc.201400139
  16. 16. Lee SH, Athukorala Y, Lee JS, Jeon YJ. Simple separation of anticoagulant sulfated galactan from marine red algae. Journal of Applied Phycology. 2008;20:1053-1059. DOI: 10.1007/s10811-007-9306-0
  17. 17. Sila A, Bougatef A. Antioxidant peptides from marine by-products: Isolation, identification and application in food systems. A review. Journal of Functional Foods. 2016;21:10-26. DOI: 10.1016/j.jff.2015.11.007
  18. 18. Cao RA, Lee YJ, You SG. Water soluble sulfated-fucans with immune-enhancing properties from Ecklonia cava. International Journal of Biological Macromolecules. 2014;67:303-311. DOI: 10.1016/j.ijbiomac.2014.03.019
  19. 19. Hahn T, Lang S, Ulber R, Muffler K. Novel procedures for the extraction of fucoidan from brown algae. Process Biochemistry. 2012;47:1691-1698. DOI: 10.1016/j.procbio.2012.06.016
  20. 20. Hardouin K, Burlot AS, Umami A, Tanniou A, Stiger-Pouvreau V, Widowati I, Bedoux G, Bourgougnon N. Biochemical and antiviral activities of enzymatic hydrolysates from different invasive French seaweeds. Journal of Applied Phycology. 2014;26:1029-1042. DOI: 10.1007/s10811-013-0201-6
  21. 21. Fleita D, El-Sayed M, Rifaat D. Evaluation of the antioxidant activity of enzymatically-hydrolyzed sulfated polysaccharides extracted from red algae; Pterocladia capillacea. LWT - Food Science and Technology. 2015;63:1236-1244. DOI: 10.1016/j.lwt.2015.04.024
  22. 22. Chen Y, Yao FK, Ming K, Wang DY, Hu YL. Polysaccharides from traditional Chinese medicines: Extraction, purification, modification, and biological activity. Molecules. 2016;21:1-23. DOI: 10.3390/molecules21121705
  23. 23. Hardouin K, Bedoux G, Burlot AS, Navall-Collen P, Bourgougnon N. Enzymatic recovery of metabolites from seaweeds: Potential applications. In: Bourgougnon N, editor. Advances in Botanical Research. Vol. 71 Sea Plants. UK: Elsevier; 2014
  24. 24. Tatke P, Jaiswal Y. An overview of microwave assisted extraction and its applications in herbal drug research. Research Journal of Medicinal Plant. 2011;5:21-31. DOI: 10.3923/rjmp.2011.21.31
  25. 25. Delazar A, Nahar L, Hamedeyazdan S, Sarker SD. Microwave-assisted extraction in natural products isolation. Methods in Molecular Biology. 2012;864:89-115. DOI: 10.1007/978-1-61779-624-1_5
  26. 26. Chan CH, Yeoh HK, Yusoff R, Ngoh GC. A first-principles model for plant cell rupture in microwave-assisted extraction of bioactive compounds. Journal of Food Engineering. 2016;188:98-107. DOI: 10.1016/j.jfoodeng.2016.05.017
  27. 27. Chin CL. Study of Extraction Processes and their Impact on Bioactivity of Botanicals. A thesis submitted for the degree of doctor of philosophy department of pharmacy national university of Singapore; 2009
  28. 28. Halfadji A, Touabet A. Badjah-Hadj-Ahmed AY. Comparison of soxhlet extraction, microwave-assisted extraction and ultrasonic extraction for the determination of PCBs congeners in spiked soils by transformer oil (Askarel). International Journal of Advances in Engineering and Technology. 2013; 5: 63-75
  29. 29. Sookjitsumran W, Devahastin S, Mujumdar AS, Chiewchan N. Comparative evaluation of microwave-assisted extraction and preheated solvent extraction of bioactive compounds from a plant material: A case study with cabbages. International Journal of Food Science and Technology. 2016;51:2440-2449. DOI: 10.1111/ijfs.13225
  30. 30. Patil DM, Akamanchi KG. Microwave assisted process intensification and kinetic modelling: Extraction of camptothecin from Nothapodytes nimmoniana plant. Industrial Crops and Products. 2017;98:60-67. DOI: 10.1016/j.indcrop.2017.01.023
  31. 31. Sun Y, Xue HK, Liu CH, Liu C, Su XL, Zheng XZ. Comparison of microwave assisted extraction with hot reflux extraction in acquirement and degradation of anthocyanin from powdered blueberry. International Journal of Agricultural and Biological Engineering. 2016;9:186-199. DOI: 10.3965/j.ijabe.20160906.2724
  32. 32. Czarnecki S, During RA. Closed-vessel miniaturised microwave-assisted EDTA extraction to determine trace metals in plant materials. International Journal of Environmental Analytical Chemistry. 2014;94:801-811
  33. 33. Quoc LPT, van Muoi N. Microwave-assisted extraction of phenolic compounds from Polygonum multiflorum Thunb. roots. Acta Scientiarum Polonorum. Technologia Alimentaria. 2016;15:181-189. DOI: 10.17306/J.AFS.2016.2.18
  34. 34. Zhang HF, Yang XH, Wang Y. Microwave assisted extraction of secondary metabolites from plants: Current status and future directions. Trends in Food Science and Technology. 2011;22:672-688. DOI: 10.1016/j.tifs.2011.07.003
  35. 35. Yanik DK. Alternative to traditional olive pomace oil extraction systems: Microwave-assisted solvent extraction of oil from wet olive pomace. LWT - Food Science and Technology. 2017;77:45-51. DOI: 10.1016/j.lwt.2016.11.020
  36. 36. Welna M, Borkowska-Burnecka J, Popko M. Ultrasound- and microwave-assisted extractions followed by hydride generation inductively coupled plasma optical emission spectrometry for lead determination in geological samples. Talanta. 2015;144:953-959. DOI: 10.1016/j.talanta.2015.07.058
  37. 37. Benthin B, Danz H, Hamburger M. Pressurized liquid extraction of medicinal plants. Journal of Chromatography A. 1999;837:211-219. DOI: 10.1016/S0021-9673(99)00071-0
  38. 38. Osorio-Tobón JF, Meireles MAA. Recent applications of pressurized fluid extraction: Curcuminoids extraction with pressurized liquids. Food and Public Health. 2013;3:289-303. DOI: 10.5923/j.fph.20130306.05
  39. 39. Wu HZ, Wang JM, Yang H, Li GQ, Zeng YH, Xia W, Li ZG, Qian MR. Development and application of an in-cell cleanup pressurized liquid extraction with ultra-high-performance liquid chromatography-tandem mass spectrometry to detect prohibited antiviral agents sensitively in livestock and poultry feces. Journal of Chromatography A. 2017;1488:10-16. DOI: 10.1016/j.chroma.2017.01.070
  40. 40. Chen HP, Pan ML, Liu X, Lu CY. Evaluation of transfer rates of multiple pesticides from green tea into infusion using water as pressurized liquid extraction solvent and ultra-performance liquid chromatography tandem mass spectrometry. Food Chemistry. 2017;216:1-9. DOI: 10.1016/j.foodchem.2016.07.175
  41. 41. Klees M, Bogatzki C, Hiester E. Selective pressurized liquid extraction for the analysis of polychlorinated biphenyls, polychlorinated dibenzo-p-dioxins and dibenzofurans in soil. Journal of Chromatography A. 2016;1468:10-16. DOI: 10.1016/j.chroma.2016.09.029
  42. 42. Jiménez-Salcedo M, Tena MT. Determination of cinnamaldehyde, carvacrol and thymol in feedstuff additives by pressurized liquid extraction followed by gas chromatography-mass spectrometry. Journal of Chromatography A. 2017;1487:14-21. DOI: 10.1016/j.chroma.2017.01.042
  43. 43. Rai A, Mohanty B, Bhargava R. Fitting of broken and intact cell model to supercritical fluid extraction (SFE) of sunflower oil. Innovative Food Science and Emerging Technologies. 2016;38:32-40. DOI: 10.1016/j.ifset.2016.08.019
  44. 44. Salea R, Veriansyah B, Tjandrawinata RR. Optimization and scale-up process for supercritical fluids extraction of ginger oil from Zingiber officinale var. Amarum. The Journal of Supercritical Fluids. 2017;120:285-294. DOI: 10.1016/j.supflu.2016.05.035
  45. 45. Chen MH, Huang TC. Volatile and nonvolatile constituents and antioxidant capacity of oleoresins in three Taiwan citrus varieties as determined by supercritical fluid extraction. Molecules. 2016;21:1-12. DOI: 10.3390/molecules21121735
  46. 46. Osorio-Tobón JF, Carvalho PIN, Rostagno MA, Meireles MAA. Process integration for turmeric products extraction using supercritical fluids and pressurized liquids: Economic evaluation. Food and Bioproducts Processing: Transactions of the Institution of Chemical Engineers Part C. 2016;98:227-235. DOI: 10.1016/j.fbp.2016.02.001
  47. 47. Antunes-Ricardo M, Gutiérrez-Uribe JA, Guajardo-Flores D. Extraction of isorhamnetin conjugates from Opuntia ficus-indica (L.) Mill using supercritical fluids. The Journal of Supercritical Fluids. 2017;119:58-63. DOI: 10.1016/j.supflu.2016.09.003
  48. 48. Picó Y. Ultrasound-assisted extraction for food and environmental samples. Trends in Analytical Chemistry. 2013;43:84-99. DOI: 10.1016/j.trac.2012.12.005
  49. 49. Yilmaz T, Kumcuoglu S, Tavman S. Ultrasound-assisted extraction of lycopene and β-carotene from tomato-processing wastes. Italian Journal of Food Science. 2017;29:186-194
  50. 50. Zhang LJ, Wang MS. Polyethylene glycol-based ultrasound-assisted extraction and ultrafiltration separation of polysaccharides from Tremella fuciformis (snow fungus). Food and Bioproducts Processing: Transactions of the Institution of Chemical Engineers Part C. 2016;100:464-468. DOI: 10.1016/j.fbp.2016.09.007
  51. 51. Abdelhedi O, Nasri R, Souissi N, Nasri M, Jridi M. Sulfated polysaccharides from common smooth hound: Extraction and assessment of anti-ACE, antioxidant and antibacterial activities. Carbohydrate Polymers. 2016;152:605-614. DOI: 10.1016/j.carbpol.2016.07.048
  52. 52. Netanel Liberman G, Ochbaum G, (Malis) Arad S, Bitton R. The sulfated polysaccharide from a marine red microalga as a platform for the incorporation of zinc ions. Carbohydrate Polymers. 2016;152:658-664. DOI: 10.1016/j.carbpol.2016.07.025
  53. 53. Tang L, Chen YC, Jiang ZB, Zhong SP, Chen WZ, Zheng FC, Shi GG. Purification, partial characterization and bioactivity of sulfated polysaccharides from Grateloupia livida. International Journal of Biological Macromolecules. 2017;94:642-652. DOI: 10.1016/j.ijbiomac.2016.10.067
  54. 54. Ngo DH, Kim SK. Sulfated polysaccharides as bioactive agents from marine algae. International Journal of Biological Macromolecules. 2013;62:70-75. DOI: 10.1016/j.ijbiomac.2013.08.036
  55. 55. Lu KY, Li R, Hsu CH, Lin CW, Chou SC, Tsai ML, Mi FL. Development of a new type of multifunctional fucoidan-based nanoparticles for anticancer drug delivery. Carbohydrate Polymers. 2017;165:410-420. DOI: 10.1016/j.carbpol.2017.02.065
  56. 56. Fletcher HR, Biller P, Ross AB, Adams JMM. The seasonal variation of fucoidan within three species of brown macroalgae. Algal Research. 2017;22:79-86. DOI: 10.1016/j.algal.2016.10.015
  57. 57. Shchipunov YA, Mukhaneva OG, Zvyagintseva TN, Popivnich IB, Shevchenko NM. Rheological properties of aqueous fucoidan solutions. Vysokomolekulyarnye Soedineniya Seriya A and Seriya B. 2000;42:93-101
  58. 58. Tako M. Rheological characteristics of fucoidan isolated from commercially cultured Cladosiphon okamuranus. Botanica Marina. 2003;46:461-465
  59. 59. Kim DY, Shin WS. Functional improvements in bovine serum albumin–fucoidan conjugate through the Maillard reaction. Food Chemistry. 2016;190:974-981. DOI: 10.1016/j.foodchem.2015.06.046
  60. 60. Hifney AF, Fawzy MA, Abdel-Gawad KM, Gomaa M. Industrial optimization of fucoidan extraction from Sargassum sp. and its potential antioxidant and emulsifying activities. Food Hydrocolloids. 2016;54:77-88. DOI: 10.1016/j.foodhyd.2015.09.022
  61. 61. Wang LC, Wang XY, Wu H, Liu R. Overview on biological activities and molecular characteristics of sulfated polysaccharides from marine green algae in recent years. Marine Drugs. 2014;12:4984-5020. DOI: 10.3390/md12094984
  62. 62. Darah I, Tong WY, Nor-Afifah S, Nurul-Aili Z, Lim SH. Antimicrobial effects of Caulerpa sertularioides extract on foodborne diarrhea-caused bacteria. European Review for Medical and Pharmacological Sciences. 2014;18:171-178
  63. 63. Li N, Mao WJ, Yan MX, Liu X, Xia Z, Xia Z, Wang SY, Xiao B, Chen CL, Zhang LF, Cao SJ. Structural characterization and anticoagulant activity of a sulfated polysaccharide from the green alga Codium divaricatum. Carbohydrate Polymers. 2015;121:175-182. DOI: 10.1016/j.carbpol.2014.12.036
  64. 64. Harada N, Maeda M. Chemical structure of antithrombin-active rhamnan sulfate from Monostrom nitidum. Bioscience, Biotechnology, and Biochemistry. 1998;62:1647-1652
  65. 65. Thanh TTT, Quach TMT, Nguyen TN, Vu Luong D, Bui ML, Tran TTV. Structure and cytotoxic activity of ulvan extracted from green seaweed Ulva lactuca. International Journal of Biological Macromolecules. 2016;93:695-702. DOI: 10.1016/j.ijbiomac.2016.09.04
  66. 66. Tsubaki S, Hiraoka M, Hadano S, Nishimura H, Kashimura K. Functional group dependent dielectric properties of sulfated hydrocolloids extracted from green macroalgal biomass. Carbohydrate Polymer. 2014;107:192-197. DOI: 10.1016/j.carbpol.2014.03.002
  67. 67. Robic A, Sassi JF, Dion P, Lerat Y, Lahaye M. Seasonal variability of physicochemical and rheological properties of ulvan in two Ulva species (Chlorophyta) from the Brittany coast. Journal of Phycology. 2009;45:962-973
  68. 68. Shao P, Ma HL, Zhu JY, Qiu Q. Impact of ionic strength on physicochemical stability of o/w emulsions stabilized by Ulva fasciata polysaccharide. Food Hydrocolloids. 2017;69:202-209. DOI: 10.1016/j.foodhyd.2017.01.039
  69. 69. Shao P, Shao JM, Jiang YK, Sun PL. Influences of Ulva fasciata polysaccharide on the rheology and stabilization of cinnamaldehyde emulsions. Carbohydrate Polymers. 2016;135:27-34. DOI: 10.1016/j.carbpol.2015.08.075
  70. 70. Nishiguchi T, Cho K, Isaka S, Ueno M, Jin J-O, Yamaguchi K, Kim DK, Oda T. Protective effect of porphyran isolated from discolored nori (Porphyra yezoensis) on lipopolysaccharide-induced endotoxin shock in mice. International Journal of Biological Macromolecules. 2016;93:1273-1278. DOI: 10.1016/j.ijbiomac.2016.09.091
  71. 71. Bhatia S, Rathee P, Sharma K, Chaugule BB, Kar N, Bera T. Immuno-modulation effect of sulphated polysaccharide (porphyran) from Porphyra vietnamensis. International Journal of Biological Macromolecules. 2013;57:50-56. DOI: 10.1016/j.ijbiomac.2013.03.01
  72. 72. Azizi R, Farahnaky A. Ultrasound assisted-viscosifying of kappa carrageenan without heating. Food Hydrocolloids. 2016;61:85-91. DOI: 10.1016/j.foodhyd.2016.05.006
  73. 73. Nur Fatin Nazurah R, Nur Hanani ZA. Physicochemical characterization of kappa-carrageenan (Euchema cottoni) based films incorporated with various plant oils. Carbohydrate Polymers. 2017;157:1479-1487. DOI: 10.1016/j.carbpol.2016.11.026
  74. 74. Cian RE, Salgado PR, Drago SR, Mauri AN. Effect of glycerol and Ca+2 addition on physicochemical properties of edible carrageenan/porphyran-based films obtained from the red alga, Pyropia columbina. Journal of Applied Phycology. 2015;27:1699-1708
  75. 75. Zhang ZS, Zhang QB, Wang J, Song H, Zhang H, Niu XZ. Regioselective syntheses of sulfated porphyrans from Porphyra haitanensis and their antioxidant and anticoagulant activities in vitro. Carbohydrate Polymers. 2010;79:1124-1129. DOI: 10.1016/j.carbpol.2009.10.055
  76. 76. In SK, Koo JG. Chemical composition and rheological properties of enzymatic hydrolysate of porphyran isolated from Pyropia yezoensis. Korean Journal of Fisheries and Aquatic Sciences. 2015;48:58-63. DOI: 10.5657/KFAS.2015.0058
  77. 77. Shu GW, Zhang BW, Zhang Q, Wan HC, Li H. Effect of temperature, pH, enzyme to substrate ratio, substrate concentration and time on the antioxidative activity of hydrolysates from goat milk casein by alcalase. Acta Universitatis Cinbinesis, Series E: Food Technology. 2016;20:29-38. DOI: 10.1515/aucft-2016-0013
  78. 78. Dhingra N, Kar A, Sharma R, Bhasin S. In-vitro antioxidative potential of different fractions from Prunus dulcis seeds: Vis a vis antiproliferative and antibacterial activities of active compounds. South African Journal of Botany. 2017;108:184-192. DOI: 10.1016/j.sajb.2016.10.013
  79. 79. Gómez-Ordóñez E, Jiménez-Escrig A, Rupérez P. Bioactivity of sulfated polysaccharides from the edible red seaweed Mastocarpus stellatus. Bioactive Carbohydrates and Dietary Fibre. 2014;3:29-40. DOI: 10.1016/j.bcdf.2014.01.002
  80. 80. Seedevi P, Moovendhan M, Viramani S, Shanmugam A. Bioactive potential and structural characterization of sulfated polysaccharide from seaweed (Gracilaria corticata). Carbohydrate Polymers. 2017;155:516-524. DOI: 10.1016/j.carbpol.2016.09.0
  81. 81. Xie JH, Wang ZJ, Shen MY, Nie SP, Gong B, Li HS, Zhao Q, Li WJ, Xie MY. Sulfated modification, characterization and antioxidant activities of polysaccharide from Cyclocarya paliurus. Food Hydrocolloids. 2016;53:7-15. DOI: 10.1016/j.foodhyd.2015.02.018
  82. 82. Ben Gara A, Ben Abdallah Kolsi R, Chaaben R, Hammami N, Kammoun M, Paolo Patti F, El Feki A, Fki L, Belghith H, Belghith K. Inhibition of key digestive enzymes related to hyperlipidemia and protection of liver-kidney functions by Cystoseira crinita sulphated polysaccharide in high-fat diet-fed rats. Biomedicine and Pharmacotherapy. 2017;85:517-526. DOI: 10.1016/j.biopha.2016.11.059
  83. 83. Badrinathan S, Shiju TM, Christa ASS, Arya R, Pragasam V. Purification and structural characterization of sulfated polysaccharide from Sargassum myriocystum and its efficacy in scavenging free radicals. Indian Journal of Pharmaceutical Sciences. 2013;74:549-555
  84. 84. Abu R, Jiang Z, Ueno M, Okimura T, Yamaguchi K, Oda T. In vitro antioxidant activities of sulfated polysaccharide ascophyllan isolated from Ascophyllum nodosum. International Journal of Biological Macromolecules. 2013;59:305-312. DOI: 10.1016/j.ijbiomac.2013.04.035
  85. 85. Deng C, Xu JJ, Fu HT, Chen JH, Xu X. Characterization, antioxidant and cytotoxic activity of sulfated derivatives of a water-insoluble polysaccharides from Dictyophora indusiata. Molecular Medicine Reports. 2015;11:2991-2998. DOI: 10.3892/mmr.2014.3060
  86. 86. Heo SJ, Lee KW, Song CB, Jeon YJ. Antioxidant activity of enzymatic extracts from brown seaweeds. Bioresource Technology. 2005;96:1613-1623. DOI: 10.1016/j.biortech.2004.07.013
  87. 87. Heo SJ, Jeon YJ, Lee J, Kim HT, Lee KW. Antioxidant effect of enzymatic hydrolyzate from a Kelp, Ecklonia cava. Algae. 2003;18:341-347. DOI: 10.4490/ALGAE.2003.18.4.341
  88. 88. Rodrigues D, Sousa S, Silva A, Amorim M, Pereira L, Rocha-Santos TAP, Gomes AMP, Duarte AC, Freitas AC. Impact of enzyme- and ultrasound-assisted extraction methods on biological properties of red, brown, and green seaweeds from the central west coast of Portugal. Journal of Agricultural and Food Chemistry. 2015;63:3177-3318
  89. 89. Hardouin K, Bedoux G, Burlot AS, Donnay-Moreno C, Bergé JP, Nyvall-Collén P, Bourgougnon N. Enzyme-assisted extraction (EAE) for the production of antiviral and antioxidant extracts from the green seaweed Ulva armoricana (Ulvales, Ulvophyceae). Algal Research. 2016;16:233-239. DOI: 10.1016/j.algal.2016.03.013
  90. 90. Je JY, Park PJ, Kim EK, Park JS, Yoon HD, Kim KR, Ahn CB. Antioxidant activity of enzymatic extracts from the brown seaweed Undaria pinnatifida by electron spin resonance spectroscopy. LWT - Food Science and Technology. 2009;42:874-878. DOI: 10.1016/j.lwt.2008.10.012
  91. 91. Alemán A, Blanco-Pascual N, Montero MP, Gómez-Guillén MC. Simple and efficient hydrolysis procedure for full utilization of the seaweed Mastocarpus stellatus to produce antioxidant films. Food Hydrocolloids. 2016;56:277-284. DOI: 10.1016/j.foodhyd.2015.12.024
  92. 92. Rodriguez-Jasso R, Mussatto S, Pastrana L, Aguilar C, Teixeira J. Chemical composition and antioxidant activity of sulphated polysaccharides extracted from Fucus vesiculosus using different hydrothermal processes. Chemical Papers. 2014;68:203-209
  93. 93. Li B, Liu S, Xing RG, Li KC, Li RF, Qin YK, Wang XQ, Wei ZH, Li PC. Degradation of sulfated polysaccharides from Enteromorpha prolifera and their antioxidant activities. Carbohydrate Polymers. 2013;92:1991-1996. DOI: 10.1016/j.carbpol.2012.11.088
  94. 94. He JZ, Xu YY, Chen HB, Sun PL. Extraction, structural characterization, and potential antioxidant activity of the polysaccharides from four seaweeds. International Journal of Molecular Sciences. 2016;17:1-17. DOI: 10.3390/ijms17121988
  95. 95. Wu SC, Lin YP, King VAE. Optimization of intermittent microwave-assisted extraction of sulfated porphyran from Porphyra dentate. Transactions of the ASABE. 2014;57:103-110
  96. 96. Saravana PS, Cho YJ, Park YB, Woo HC, Chun BS. Structural, antioxidant, and emulsifying activities of fucoidan from Saccharina japonica using pressurized liquid extraction. Carbohydrate Polymers. 2016;153:518-525. DOI: 10.1016/j.carbpol.2016.08.014
  97. 97. Ospina M, Castro-Vargas HI, Parada-Alfonso F. Antioxidant capacity of Colombian seaweeds: 1. Extracts obtained from Gracilaria mammillaris by means of supercritical fluid extraction. The Journal of Supercritical Fluids. 2017. (In Press) DOI: 10.1016/j.supflu.2017.02.023
  98. 98. Fidelis GP, Gomes Camara RB, Queiroz MF, Santos Pereira Costa MS, Santos PC, Oliveira Rocha HA, Costa LS. Proteolysis, NaOH and ultrasound-enhanced extraction of anticoagulant and antioxidant sulfated polysaccharides from the edible seaweed, Gracilaria birdiae. Molecules. 2014;19:18511-18526
  99. 99. Li SQ, Dai SH, Shah NP. Sulfonation and antioxidative evaluation of polysaccharides from Pleurotus mushroom and Streptococcus thermophilus bacteria: A review. Comprehensive Reviews in Food Science and Food Safety. 2017;16:282-294. DOI: 10.1111/1541-4337.12252

Written By

Shao-Chi Wu

Submitted: December 11th, 2016 Reviewed: May 8th, 2017 Published: November 29th, 2017