Brown Algae-Derived Polysaccharides: From Sustainable Bioprocessing to Industrial Applications

Abstract

Brown seaweeds are marine bioresources rich in bioactive compounds such as carbohydrates, proteins, pigments, fatty acids, polyphenols, vitamins, and minerals. Among these substances, brown algae-derived polysaccharides (alginate, fucoidan, and laminarin) have promising industrial prospects owing to their distinctive structural features and diverse biological activities. Consequently, processing technologies have advanced substantially to address industrial requirements for biopolymer quality, cost-effectiveness, and sustainability. Over the years, significant progress has been made in developing various advanced methods for the sake of extracting, purifying, and structurally characterizing polysaccharides. Aside from that, numerous studies reported their broad spectrum of biological activities, such as antioxidant, anti-inflammatory, anticoagulant, and antimicrobial properties. Furthermore, these substances have various industrial, pharmaceutical, bioenergy, food, and other biotechnology applications. The present review systematically outlines the brown algae-derived polysaccharides treatment process, covering the entire value chain from seaweed harvesting to advanced extraction methods, while highlighting their biological activities and industrial potential as well.

1. Introduction

Phaeophyceae, commonly known as brown algae, are a large and diverse group of multicellular, photosynthetic organisms that belong to the kingdom Chromista [1]. Brown seaweed, comprising approximately 250 genera and 2000 species, is a significant part of intertidal zone biomass, notably from predominant genera such as Sargassum, Ascophyllum, Laminaria, and Macrocystis [2,3]. As a foundational group, brown algae species hold immense ecological value, providing essential habitats, nursery grounds, and shelter for numerous marine species while also playing a key role in carbon cycling [4,5]. Their name is derived from their distinctive greenish-brown color, which results from the fucoxanthin pigment in their chloroplasts and from specific tannins found in their cell walls that mask residual pigments [6,7].

For centuries, marine brown algae have been an integral part of oriental culture, especially in the Asian–Pacific region [8]. Their widespread use is owing to their dietary, medicinal, and functional food properties [9,10]. Today, seaweeds have become a globally important economic resource, especially in East and South Asia, where they are extensively cultivated [11]. Much attention has been paid to seaweeds as a valuable source of structurally novel and biologically active metabolites with a significant industrial potential, particularly polysaccharides [12,13,14]. As the principal structural polysaccharides in brown seaweed, fucoidan, alginate, and laminarin exhibit notable heterogeneity in their physicochemical properties and structural characteristics that directly influence their biological activities [15]. This diversity depends on several factors such as the species of algae, growth environment, harvesting season, and extraction process. Consequently, such variability poses substantial challenges in terms of pretreatment, extraction, characterization, and evaluation of bioactivity [16].

Although the discovery of new brown algal species is continually accelerating, a substantial number of these taxa is still without formal taxonomic classification or phylogenetic distinction [4]. Driven by accessible high-throughput sequencing, the development of comprehensive DNA barcode libraries has dramatically revolutionized specimen identification in taxonomic, ecological, and biogeographical research. DNA barcoding initiatives allow for the accurate identification of cryptic species, larval forms, and novel specimens previously unidentifiable after using traditional morphological methods [17,18,19]. The vision of a global and interconnected library of barcodes relied on standardized genetic markers exemplified by many initiatives targeting the ribosomal genes, ITS spacer, rbcL plastid gene, and the conserved mitochondrial genes as well [17,20].

The high quality of downstream extraction fundamentally depends on the preliminary treatment of raw seaweed material [21]. The production scheme involves various stages, including algal strain cultivation followed by biomass harvesting, which also involves the separation from carrier media and impurities (e.g., salts, heavy metals, and epiphytes), and ends in drying and milling into a usable form [22]. Thanks to great technological innovations, the processing of polysaccharides derived from brown algae has now become a highly sophisticated field [23]. Traditionally, solid-to-liquid extraction using acidic, alkaline, or aqueous solvents was the standard method for the extraction of polysaccharides. Still, innovative extraction systems have been developed in recent years to recover polysaccharides in a more efficient and environmentally friendly manner [24]. Current extraction technologies such as Ultrasound-Assisted Extraction (UAE), Microwave-Assisted Extraction (MAE), Enzymatic-Assisted Extraction (EAE), Pressurized Liquid Extraction (PLE), and Supercritical Fluid Extraction (SFE) have revolutionized brown algae processing. Indeed, these protocols have fewer drawbacks than conventional methods thanks to their considerable potential for enhancing extraction yields, reducing energy and solvent consumption, streamlining processing time, and providing safer working conditions through reduced chemical exposure. The crude extract typically contains impurities, which can lead to an overestimation of extraction yields and limit the biological applications of target polysaccharides [25]. However, promoting a high-purity product for advanced applications usually requires further purification through chromatographic and membrane-based methods [26,27]. These processes aim to decipher the molecular architecture such as the sugar composition, glycosidic linkage patterns, functional groups, molecular weight, and anomeric configuration [28]. For this purpose, a suite of sophisticated analytical techniques is required including chromatographic methods (e.g., HPAEC-PAD), spectroscopic analyses (e.g., NMR, FTIR), mass spectrometry (e.g., MS, GC-MS, LC-MS), and advanced microscopy [26,29].

As naturally derived biopolymers, brown algae polysaccharides have been extensively investigated for their broad spectrum of biological activities [30]. These biological effects are mediated via the direct initiation of complex reaction pathways [31]. This dual capacity accounts for the diverse biological characteristics observed including antioxidant, anti-inflammatory, anticoagulant, antimicrobial, and immunomodulatory activities [21]. Once their structure and functional properties are elucidated, these biologically active polysaccharides may serve as a source of fertile materials for various applications. Their innate biodegradability and low toxicity make them highly attractive candidates for diverse industries beyond traditional uses. Numerous studies highlight their importance in different sectors such as the food industry (as functional ingredients), pharmaceuticals (for drug delivery), cosmetics (in skincare formulations), agriculture (as biostimulants), and the environment (in bioremediation) [2,32].

In fact, many previous studies were made on specific biological activities or extraction techniques. Yet, a substantial gap still appears in providing a holistic system-level perspective that connects the entire valorization pipeline. Many studies predominantly focus on downstream aspects such as structure–activity relationships, often treating upstream biomass processing as a peripheral concern. To bridge this gap, this study is structured around the novel conceptual framework of an integrated value chain. By adopting this approach, the present study provides an extensive and updated overview of brown algae-derived polysaccharide processing, focusing on three core aspects essential for their research and application: (i) biomass pretreatment process, outlining key methods including washing, drying, size reduction, and taxonomic species delimitation; (ii) extraction and characterization techniques, from conventional to emerging methods, alongside the advanced analytical tools used to determine structural and functional properties; and (iii) bioactive properties associated with these polysaccharides. Through a critical analysis of these themes, this study seeks to offer valuable insights for the scientific community and industry partners, aiming to optimize the sustainable processing and application of brown algae polysaccharides across various fields.

2. Global Seaweed Production

Seaweed production originated approximately 1700 years ago in China for domestic purposes such as food, feed, and medicine prior to its adoption for industrial applications [33]. Over the years, the exponential growth of the global seaweed industry led to the cultivation and harvesting of a wide variety of seaweed species. This diversification is a direct response to market pressures seeking novel value-added products. However, the rapid expansion and transition from artisanal collection to large-scale commercial farming raise critical questions regarding its environmental and socio-economic sustainability, particularly for coastal countries in Asia [34].

According to statistics from the Food and Agriculture Organization [35], the global market share of seaweed farming production increased by approximately 203% between 2000 and 2019, reaching over 35 million tons per year. Recent studies (2020) have indicated that the algae industry, primarily dominated by Asia (97% of the total production), is expected to grow by 137% by 2027, increasing its value from 40 billion to 95 billion dollars. The leading Asian producers are China, Indonesia, Korea, and the Philippines, while recent growth has also been observed in Europe (0.8%, e.g., Ireland, Norway, and France) and the Americas (e.g., Chile, Canada, and Mexico) [36]. However, increasing production sustainably requires the optimization of cultivation systems [24]. The expansion of the seaweed sector is largely attributed to the increasing global demand for brown seaweed biomass and its bio-based derivatives. Statistical records indicate an average annual increase of around 11%, with production volumes expanding from 13,000 tons in 1950 to 17.6 million tons in 2021. Today, 96.5% of the total global production of brown seaweed comes from aquaculture [37]. This expanding biomass is crucial to respond to the market needs for valuable polysaccharides. For instance, the bioactivities of fucoidan have positioned it as a compound of notable commercial value, with a global market valuation of USD 30 million in 2022 [38]. Similarly, the alginate industry was valued at USD 728.4 million in 2020, with an expected annual growth rate of 5% until 2028 [39]. Microalgae can be farmed offshore in artificial ponds or wildly harvested from natural beds (Table 1) [11]. The choice of cultivation technique is largely determined by the biological requirements of the target species and its optimal growth environment [40].

Table 1. Seaweed cultivation techniques.

Cultivation Method	Advantages	Disadvantages	Examples	Species	Most-Used	References
Offshore seaweed farming	High environmental control (nutrients, temperature, light, pH, and pathogens) Stable productivity (yield, growth cycles) Lower physical risks (storms, currents)	Substantial operational costs (water, energy, and ponds) External input dependency (fertilizers, treatments) Limited scalability (available area, cost)	Bottom culture Pond culture Tank culture	Sargassum horneri Cystoseira barbata Himantothallus grandifolius	China Indonesia Philippines South Korea Netherlands USA	[41,42,43,44]
Onshore seaweed farming	Lower production costs Natural resources (nutrients) Unlimited space	High environmental risks (storms, waves, and grazing) Yield variability Logistical challenges (monitoring, harvesting, and maintenance)	Floating raft Open water Rope	Sargassum muticum Dictyota menstrualis Turbinaria ornata	Norway France Portugal Canada Japan Chile	[42,45,46]

Table 1. Seaweed cultivation techniques.

Cultivation Method	Advantages	Disadvantages	Examples	Species	Most-Used	References
Offshore seaweed farming	High environmental control (nutrients, temperature, light, pH, and pathogens) Stable productivity (yield, growth cycles) Lower physical risks (storms, currents)	Substantial operational costs (water, energy, and ponds) External input dependency (fertilizers, treatments) Limited scalability (available area, cost)	Bottom culture Pond culture Tank culture	Sargassum horneri Cystoseira barbata Himantothallus grandifolius	China Indonesia Philippines South Korea Netherlands USA	[41,42,43,44]
Onshore seaweed farming	Lower production costs Natural resources (nutrients) Unlimited space	High environmental risks (storms, waves, and grazing) Yield variability Logistical challenges (monitoring, harvesting, and maintenance)	Floating raft Open water Rope	Sargassum muticum Dictyota menstrualis Turbinaria ornata	Norway France Portugal Canada Japan Chile	[42,45,46]

3. Sample Processing

The postharvest handling of seaweed biomass is a critical factor influencing all subsequent valorization pathways. As illustrated in Figure 1, the standard processing chain involves key unit operations such as washing, drying, and size reduction. The choice of processing method primarily depends on the algal material, the intended end product, and the socio-economic objectives of production [47].

3.1. Washing

Unwashed algal biomass contains a wide range of natural contaminants including sediment (e.g., mud, sand), epibionts (e.g., small invertebrates), and associated macro algal species [31]. Washing with sea or tap waters is an important pretreatment step strongly recommended to reduce the salt content and eliminate adhering impurities [48]. The washing conditions for each macro algal species should be optimized to preserve color and texture regardless of whether freshwater or seawater was used [22].

3.2. Drying

Seaweeds, with a moisture content exceeding 90–95%, are considered highly perishable material [49]. Drying is a major operation in the seaweed industry, as it extends the shelf life and improves storage efficiency, transport, and distribution, as well [50]. Substantial progress has recently been made in drying technologies to satisfy the growing demands of the industry. Currently, two main drying technologies are commercially emerging: (i) natural sun drying, which is a relatively simple and low-cost technique, but its dependence on weather conditions affects both the quality and sanitary standards of the final product, hindering its application in some industrial sectors; and (ii) mechanical drying using industrial thermal drying systems, which, despite their fast processing rates, are constrained by limited throughput, high energy consumption, and degradation of key nutrients [51,52]. As a promising alternative, effective freezing ensures good preservation of the organoleptic and nutritional quality of seaweed, particularly for high-value applications that require fresh-like attributes [53].

3.3. Milling or Size Reduction

For extraction purposes, dried seaweeds are typically crushed or manually ground using a mechanical mill or blender to obtain a homogeneous mass and increase the surface-to-volume ratio [54]. Milling methods involve the production of particles with a wide range of sizes from millimeters to microns [55]. Finally, sieving is applied to separate algal particles into different size fractions and obtain the desired particle size [52]. However, the requirement for this mechanical pretreatment is highly species-specific. In taxa having fragile cell wall structures, the extraction medium led to effective cell disruption, thereby requiring less mechanical energy or potentially removing the necessity for prior milling [56].

4. Genus and Species Delimitation

Accurate algal species identification is crucial for ecological studies, biodiversity conservation, and sustainable management of marine resources, as well as for ensuring effective traceability and quality control in the commercialization of seaweed-based products [57]. The challenges associated with the morphological species concept revealed significant limitations. In algae, phenotypic plasticity, polymorphism, and convergent evolution are common phenomena, creating marked discrepancies between the morphologically defined species and those delimited by phylogenetic or reproductive isolation criteria [5,58]. Moreover, cryptic species are frequently observed in diverse lineages. In addition, morphological diagnostic keys may vary depending on the life stage or sex of the algae. Ultimately, the reliable application of morphological keys requires advanced taxonomic expertise for accurate identification [59].

To remove such constraints, integrative approaches that combine morphological characteristics with DNA data (nuclear, mitochondrial, and/or plastid) have been increasingly adopted in taxonomic and ecological research to complement classical morphological identification methods [57,60].

DNA-based tools, such as DNA barcoding, have proven to be a practical method for marine flora studies, making simple, rapid, and cost-effective identification of novel species [61]. As illustrated in Figure 2, this approach involves the sequencing of a target gene from accurately authenticated specimens to create a reference database and facilitate the identification of unknown species via a comparative sequence analysis with/of this library. With current advances in molecular technologies, multiple validated DNA barcodes are available in accessible sequence databases [17]. DNA barcoding markers are derived from distinct cellular compartments: (i) nuclear genomes, providing ribosomal RNA genes (18S/28S) and the ribosomal internal transcribed spacer region (ITS1-5.8SrDNA-ITS2); (ii) chloroplast genomes, featuring the widely adopted rbcL gene (ribulose-1,5-bisphosphate carboxylase/oxygenase large subunit) and psaA/psbA (photosystem I P700 chlorophyll Apo protein A1); and (iii) mitochondrial genomes, particularly the COI-5P (cytochrome c oxidase subunit I 5′ region) and the cox2–3 spacer (cytochrome c oxidase subunit II–III intergenic spacer) [17,62].

5. Marine Brown Algae Polysaccharides: Biodiversity and Chemical Structure

Brown seaweeds are characterized by a high content of polysaccharides that serve as structural cell wall constituents and energy-reserve storage compounds. Specifically, the brown algal intercellular matrix and cell walls are predominantly composed of alginate and fucoidan, which are both critical for thallus support in water. The primary storage of the polysaccharide laminarin is a direct product of photosynthesis and is sequestered within plastids as a carbon reserve for core metabolism [63]. As major components, brown seaweed polysaccharides, including sodium alginate, fucoidans, and laminarin, exhibit significant structural and chemical heterogeneity [64]. This heterogeneity is influenced by many factors such as species-specific traits, geographic location, harvesting season and extraction methods. The substantial variability among these polysaccharides presents great industrial challenges for valorization and innovation [65].

5.1. Alginate

Alginate, which accounts for approximately 40% of the dry weight, is the primary polysaccharide in the cell wall and intercellular matrix of brown algae [66]. It naturally occurs as an insoluble mixed salt of alginic acid complexed with various cations, mainly Ca²⁺, Na⁺, Mg²⁺, and K⁺, along with other ions normally existing in seawater. Structurally, alginate is a linear and unbranched polysaccharide with a molecular mass larger than 50 kDa [11]. Its structure consists of β−D−mannuronic acid (M) and α−L−guluronic acid (G), linked through (1,4)−glycosidic linkages [67,68]. These monomers are arranged in homogeneous (MM or GG) or heterogeneous (MG) blocks (Figure 3). The monomeric ratios (M/G) of sodium alginate vary in different species of brown algae, with reported values of 0.52 for Sargassum dentifolium, 0.77 for Cystoseira compressa, 1.12 for Laminaria digitata, and 1.5 for Padina gymnospora [69]. The relative proportions of M and G, as well as block distributions (MM, GG, MG, and GM), influence the physicochemical properties of alginate. For example, alginate with high-M blocks showed higher viscosity, whereas high-G alginate shares better gelling properties [66]. Alginate’s gelling ability is intrinsically linked to its anionic polyelectrolyte nature. The process is initiated by the exchange of sodium ions from guluronic acid residues with divalent cations, leading to the formation of cross-linked junctions described by the «egg-box» model [69]. As a direct result, its rheological behavior in an aqueous solution, including flow and viscosity, is highly dependent on pH and ionic strength [70].

5.2. Fucoidan

Fucoidan is a sulfated polysaccharide commonly found in brown seaweeds and some lower plants and typically yields up to 30% of dry weight [71]. As an alginate, fucoidan is essentially a structural component of the intercellular matrix where it plays a crucial role in maintaining cellular integrity and providing protection against environmental stresses. The relative molecular weight of fucoidan generally varies from about 7 kDa [72] to 2379 kDa [73]. The fucoidan backbone is highly variable among species primarily composed of fucose with additional monosaccharide residues (such as mannose, galactose, arabinose, glucose, xylose, and uronic acids). Generally, α−(1,3) and α−(1,4) glycosidic bonds constitute the main chain of the macromolecule (Figure 4) [21]. A major structural feature of fucoidan is its specific substitution pattern characterized by sulfate groups, particularly those attached at the C−2, C−4, and/or C−3 and the O-acetylation. These functionalities confer a strong polyanionic character and enhanced chain stiffness, which profoundly influence the molecule’s physicochemical behavior, particularly its conformation in aqueous media and its interactions with water and other polymers [74]. From a techno-functional standpoint, the polyanionic nature of fucoidan allows for interactions with counter-ions and cationic species (e.g., proteins, polysaccharides, or mineral ions), thereby modulating the rheology, stability, and texture of dispersions, emulsions, and gels, as well [75]. Rheologically, fucoidan solutions and blends typically exhibit non-Newtonian, shear-thinning (pseudoplastic) behavior, where the apparent viscosity decreases as the shear rate increases [76].

5.3. Laminarin

Laminarin, a water-soluble β−glucan, is another major storage polysaccharide found in brown algae, particularly in Laminaria, Saccharina, Ascophyllum, Fucus, and Alaria species [77]. Laminarin is mainly composed of β−(1,3)−β−D−glucan backbone with some 6-O-branching, glucose, and β−(1,6)−intrachain link. The ratio of β−(1,3) to β−(1,6) linkages vary depending on the type of algae [78]. Additionally, laminarin is structurally classified into two forms based on the type of sugar at the reducing end: M-chains, featuring a terminal D-mannitol (1−O-substituted), and G-chains, which end with glucose (Figure 5) [52]. The average molecular weight of laminarin is approximately 5 kDa, depending on the degree of polymerization. Furthermore, Laminarin represents around 22–49% of algal dry matter [30].

6. Polysaccharides Treatment Process

Processing polysaccharides from brown algae is a crucial technological pathway for accessing their functional and biological properties in various industrial applications [79]. Commercial-scale processing involves three interdependent critical stages (Figure 6): (i) pretreatment and primary extraction, (ii) successive purification, and (iii) complete analytical profiling [80].

6.1. Extraction Technologies

To obtain targeted brown algae hydrocolloids tailored to specific purposes and functions, a wide range of extraction methods, from the conventional to the more advanced ones, are documented (Figure 7). Although deep research has been made on extraction processes, few studies have systematically evaluated their efficiency, scalability, and suitability for industrial applications [9]. This lack of comprehensive assessment is a paramount concern, since effective recovery methods are required to achieve optimal yields of polysaccharides with high purity and strong bioactivity [80,81].

6.1.1. Hot Water Extraction

As the most commonly used extraction technique, hot water extraction (HWE) is adopted for its simplicity and selectivity towards water-soluble polysaccharide compounds [82,83]. This technique was implemented at temperatures above 80 °C for a controlled duration (1–4 h) using a solid-to-liquid ratio of 1:10 to 1:30 (w/v) to effectively disrupt the algal cell matrices and release selective polysaccharide fractions under mild conditions [80]. For instance, hydrothermal processing of Saccharina japonica under extreme conditions (180–420 °C, 13–520 bar) can achieve a yield of 98.91% of total organic carbon in dry seaweed [84].

6.1.2. Soxhlet-Assisted Extraction

The key advancement of the Soxhlet Extraction technique over traditional hot water extraction is the continuous flow of solvent through the sample matrix. This dynamic process enhances the extraction efficiency, typically completing a cycle within 2–3 h [85]. Despite the large-scale application, Soxhlet-Assisted Extraction is commonly applied as a preliminary step in macroalgae processing to remove non-targeted compounds (e.g., lipids, pigments) [86].

6.1.3. Maceration

Maceration is a flexible but cost-effective extraction technique that uses a combination of organic solvents (e.g., methanol, ethanol, and ethyl acetate) and/or acid or alkali solutions (e.g., HCl, H₂SO₄, Na₂CO₃, and NaOH) under controlled temperature and agitation [53,87]. It efficiently solubilizes and extracts functional polysaccharides from brown algal cell walls. For example, alginate extraction from Cystoseira compressa involves a sequential solvent-based approach using acidic pretreatment (0.1 M HCl) followed by alkali extraction (Na₂CO₃) at elevated temperatures (60 °C). The extraction protocol involves multiple maceration steps with acetone, methanol, and ethanol [29].

6.1.4. Ultrasound-Assisted Extraction

Ultrasound-Assisted Extraction (UAE) is an innovative non-thermal technique that uses high-frequency sound waves (16–100 kHz) to disrupt cellular structures through the cavitation phenomenon [88]. This process generates oscillating pressure waves in the liquid medium, creating microbubbles that mechanically damage cell walls and facilitate the release of bioactive compounds and polysaccharides from brown seaweeds [85]. According to the literature, water or acidic solutions are the most employed solvents for UAE. However, other organic solvents like methanol and ethanol are used for industrial-scale applications [86,89]. UAE combines enhanced yield, time efficiency, low solvent consumption, and low-temperature operation, all achievable through low-cost equipment [65].

6.1.5. Enzymatic-Assisted Extraction

Enzymatic-Assisted Extraction (EAE) is a promising green technology based on the use of selective enzymes as catalysts for cell wall degradation. The successful implementation of an EAE system depends on two interdependent factors: (i) screening target biocatalysts according to their specificity and selectivity profiles and (ii) adjustment of process parameters (temperature, pH, solvent polarity, and enzyme-to-substrate ratio) for the sake of maximizing extraction efficiency [90,91,92]. Various types of commercial enzymes are available, including proteases such as Alcalase and Flavourzymes, and carbohydrases like Celluclast, Viscozyme, Amyloglucosidase, Termamyl, and Ultraflo, as well [93,94]. By using biodegradable catalysts, EAE offers enhanced environmental sustainability, reduced energy consumption, and high extraction yields with improved specificity [95]. Yet, economic constraints often limit its industrial application on a large scale [80].

6.1.6. Microwave-Assisted Extraction

This energy-assisted technique is used to overcome the limitations of conventional procedures. Microwave-Assisted Extraction (MAE) is described as a process that employs microwave radiation to heat algal material, thereby accelerating the extraction process [92,96]. During microwave treatment, rapid heating occurs directly within the biological material, causing the evaporation of intracellular fluids [75]. This results in the disruption of the cell wall and the release of various bioactive compounds and polysaccharides into the solvent [97]. Generally, water and dilute acidic solutions (e.g., 0.01 M HCl) are used as solvents for this type of extraction [85]. This method is undoubtedly effective for brown algal polysaccharides, as it reduces processing time and energy and solvent consumption while allowing for integration with other extraction technologies [98]. Nevertheless, the main disadvantage lies in the potential alteration or degradation of thermo-labile compounds [86].

6.1.7. Pressurized Liquid Extraction

Pressurized Liquid Extraction (PLE) is also known as Pressurized Solvent Extraction (PSE), Accelerated Solvent Extraction (ASE), or Subcritical Water Extraction (SWE), depending on the used solvent [99]. The PLE technique is based on using water or organic solvents under high temperatures (50–200 °C) and pressures (35–200 bar) within an oxygen and light-free environment [94]. Higher temperatures enhance compound solubility, modify the physical properties of solvent, and accelerate diffusion rates, while elevated pressure maintains the solvent in its liquid state below its boiling point. The technique is known for its eco-efficiency and short extraction times [84]. Despite this efficiency, high-temperature operation may lead to undesired reactions or the degradation of compounds. Furthermore, industrial-scale implementation is still limited by high energy demand and infrastructure-related costs [86].

6.1.8. Supercritical Fluid Extraction

Another example of innovative techniques is the Supercritical Fluid Extraction (SFE) that is based on the application of supercritical fluids as solvents (e.g., CO₂) to extract polysaccharides under high pressure and high temperatures [9]. Each fluid is defined by its corresponding temperature and pressure, which determines its thermophysical behavior. Beyond its critical temperature, a supercritical fluid cannot be liquefied regardless of the applied pressure, but it retains a density similar to that of a liquid. According to various studies, carbon dioxide (CO₂) is the ideal solvent for Supercritical Fluid Extraction (SFE) due to its non-toxicity, safety, abundance, and ease of removal from the final product. Indeed, SFE is recognized as a highly selective, sustainable, and effective eco-friendly process for producing high-purity products [8].

The heterogeneity of experimental conditions, including variations in instrumental parameters, extraction duration, and solvent systems, together with the investigated broad spectrum of polysaccharides (Table 2), renders direct comparison among extraction techniques difficult.

Table 2. Overview of traditional and advanced methods for brown seaweed polysaccharide extraction.

Species	Polysaccharide	Extraction Conditions	Recovery	References
Soxhlet Extraction
Fucus vesiculosus	Fucoidan	CaCl2 (2%), 85 °C, 2 h	2.6%	[100]
Maceration Extraction
Ascophyllum nodosum	Fucoidan	HCl (0.01 M), 70 °C, 3 h	11.9%	[101]
Coccophora langsdorfii	Sodium alginate	Na2CO3, 60 °C, 24 h	13.3%	[102]
Ultrasound-Assisted Extraction
Sargassum muticum	Sodium alginate	H2O, 150 W/40 kHz, 25 °C, 30 min	15%	[76]
Sargassum binderi	Fucoidan	HCl (0.1 M), 60 min	22.40%	[103]
Laminaria hyperborea	Laminarin	HCl (0.1 M), 20 kHz, 15 min	6.24%	[52]
Enzymatic-Assisted Extraction
Padina gymnospora	Total polysaccharides	Alcalase (0.32% w/w), 60.5 °C, 1.95 h	87.45%	[104]
Ascophyllum nodosum	Fucoidan	Cellulase, pH 4.5, 50 °C, 24 h	3.89%	[101]
Nizamuddinia zanardinii	Fucoidan	Flavourzyme (5% v/v, pH 7), 50 °C, 24 h	4.36%	[105]
Fucus vesiculosus	Neutral sugars	β−glucosidase, Botrytis glucanase, protease (5% w/w), 50 °C, 17 h	60%	[106]
Microwave-Assisted Extraction
Sargassum siliquosum	Fucoidan	H2O, 750 W, 10 min	6.94%	[75]
Sargassum pallidum	Sulfated polysaccharides	EtOH (21%), (NH4)2SO4 (22%), 830 W, 95 °C, 15 min	7.56%	[107]
Ascophyllum nodosum	Fucoidan	HCl (0.1 M), 120 °C, 15 min	16.08%	[16]
Pressurized Liquid Extraction
Nizamuddinia zanardinii	Fucoidan	H2O, 7.5 bar, 150 °C, 29 min	25.98%	[108]
Saccharina japonica	Fucoidan	NaOH (0.1%), 80 bar, 127.01 °C, 11.98 min	13.56%	[84]

Table 2. Overview of traditional and advanced methods for brown seaweed polysaccharide extraction.

Species	Polysaccharide	Extraction Conditions	Recovery	References
Soxhlet Extraction
Fucus vesiculosus	Fucoidan	CaCl2 (2%), 85 °C, 2 h	2.6%	[100]
Maceration Extraction
Ascophyllum nodosum	Fucoidan	HCl (0.01 M), 70 °C, 3 h	11.9%	[101]
Coccophora langsdorfii	Sodium alginate	Na2CO3, 60 °C, 24 h	13.3%	[102]
Ultrasound-Assisted Extraction
Sargassum muticum	Sodium alginate	H2O, 150 W/40 kHz, 25 °C, 30 min	15%	[76]
Sargassum binderi	Fucoidan	HCl (0.1 M), 60 min	22.40%	[103]
Laminaria hyperborea	Laminarin	HCl (0.1 M), 20 kHz, 15 min	6.24%	[52]
Enzymatic-Assisted Extraction
Padina gymnospora	Total polysaccharides	Alcalase (0.32% w/w), 60.5 °C, 1.95 h	87.45%	[104]
Ascophyllum nodosum	Fucoidan	Cellulase, pH 4.5, 50 °C, 24 h	3.89%	[101]
Nizamuddinia zanardinii	Fucoidan	Flavourzyme (5% v/v, pH 7), 50 °C, 24 h	4.36%	[105]
Fucus vesiculosus	Neutral sugars	β−glucosidase, Botrytis glucanase, protease (5% w/w), 50 °C, 17 h	60%	[106]
Microwave-Assisted Extraction
Sargassum siliquosum	Fucoidan	H2O, 750 W, 10 min	6.94%	[75]
Sargassum pallidum	Sulfated polysaccharides	EtOH (21%), (NH4)2SO4 (22%), 830 W, 95 °C, 15 min	7.56%	[107]
Ascophyllum nodosum	Fucoidan	HCl (0.1 M), 120 °C, 15 min	16.08%	[16]
Pressurized Liquid Extraction
Nizamuddinia zanardinii	Fucoidan	H2O, 7.5 bar, 150 °C, 29 min	25.98%	[108]
Saccharina japonica	Fucoidan	NaOH (0.1%), 80 bar, 127.01 °C, 11.98 min	13.56%	[84]

6.2. Purification Procedure

The application of conventional extraction processes yields polysaccharide-rich fractions, such as fucoidan, alginate, and laminarin, which are invariably associated with coextracted compounds. Significant amounts of proteins, polyphenols, and flavonoids are often present as contaminants in the resulting mixture [109,110]. To obtain high-purity polysaccharides, various purification strategies can be applied, including ethanol precipitation, membrane separation, and chromatographic methods [91].

6.2.1. Ethanol Precipitation

Ethanol precipitation is the most commonly used method, often employed as the first step in polysaccharide purification. However, this approach not only effectively precipitates target polysaccharides but also removes low-molecular-weight impurities [111]. Standard precipitation is typically performed by adding two to three volumes of ethanol to the mixture and allowing it to precipitate overnight at 4° [112]. Ethanol’s low dielectric constant reduces ionic interactions between polysaccharide sulfate groups and positive ions, inducing aggregation. Target polysaccharides are then recovered as a pellet via centrifugation, while soluble non-polysaccharide impurities remain in the supernatant [94]. As a one-step purification strategy, ethanol precipitation delivers operational simplicity, cost-effectiveness, and industrial scalability. Several recent reports have highlighted the use of ethanol precipitation to purify polysaccharide fractions from brown algae species including Cystoseira compressa [29] and Halopteris scoparia [113].

6.2.2. Membrane-Based Methods

Membrane-based purification techniques, including ultrafiltration and dialysis, have emerged as size-selective approaches for the separation of heterogeneous mixtures [109]. In brief, the polysaccharide mixture containing low-molecular-weight contaminants such as salts, proteins, and phenolic compounds is placed inside a dialysis membrane to allow for selective and passive diffusion [114]. Membranes/filters with progressively graduated molecular-weight cut-off (MWCO) are available, providing diverse filtration modes including nanofiltration, microfiltration, ultrafiltration, and reverse osmosis [115,116]. Membrane-based systems offer extreme operational simplicity, cost-efficiency, and low chemical dependency, while allowing for the simultaneous removal of multi-contaminants and the processing of large volumes of solutions at a commercial scale [109,117]. Following fucoidan extraction from the brown seaweed Ascophyllum [27], a three-step purification process was applied involving a 10 kDa molecular-weight cut-off membrane, yielding 562.3 mg of pure fucoidan per g of dry extract.

6.2.3. Chromatographic Techniques

Brown algae polysaccharides are commonly purified using chromatography techniques such as ion exchange, size exclusion, and affinity chromatography, which separate molecules based on differences in their charge, size, and hydrophobicity.

Currently, ion exchange chromatography (IEC) is the most widely used tool for the separation and purification of charged biological polymers, such as proteins, DNA/RNA, and polysaccharides [118]. According to the principles of IEC, solute separation relies on three primary interactions: reversible ion exchange, electrostatic attraction, and adsorption phenomena, all involving charged groups from the column material, the biological sample, and eluent ions [119]. Briefly, charged solute molecules adsorb onto immobilized ion-exchange groups on the column matrix, followed by selective elution controlled by modulating the mobile phase buffer pH and/or ionic strength [120]. IEC separation modalities are charge-dependent: anion-exchange chromatography (AEC) is used for negatively charged targets, while cation-exchange chromatography (CEC) is used for cationic targets (pH < pI). Anion/cation exchangers are classified as “strong” or “weak” based on their pH-dependent charge variations. IEC’s prominence in modern chromatography derives from its superior resolution, high throughput, cost-effectiveness, and ease of automation. Recent literature revealed that modified Diethylaminoethyl (DEAE-Cellulose/Sepharose/Dextran) and quaternary ammonium (Q) groups are routinely adopted as AEC-ligands for polysaccharide separation [87].

As a physicochemical separation technique, Size-Exclusion Chromatography (SEC) resolves complex mixtures through differential elution based on the hydrodynamic volume and molecular weight of the target analytes [121,122]. Molecules are fractionated within a porous gel matrix, where larger molecules quickly pass through the column, while smaller molecules penetrate deeper and elute later. The resolution of polysaccharide separation depends mainly on the choice of gel filtration media relative to the target polysaccharide’s molecular weight. Current polysaccharide purification protocols predominantly rely on four common column types, namely, PL aquagel OH, Sepharose CL-6B, Sephacryl S-300, and Superdex 200 [93]. For sensitive biomolecules, SEC combines operational simplicity with high resolution while preserving their native molecular structures. Unlike binding-dependent techniques like IEC, SEC’s steric exclusion mechanism eliminates the influence of buffer chemistry on separation efficiency [80].

Affinity chromatography (AC) is a high-resolution liquid chromatography method that separates target compounds based on specific molecular interactions, such as antigen–antibody binding, enzyme–substrate recognition, and receptor–protein interactions. Molecular interactions are mediated through various chemical forces, including ionic, hydrophobic, hydrogen, and disulfide bonds. The selective binding between target molecules and the immobilized ligands on the chromatographic matrix enables selective retention, while nonspecific contaminants are eliminated during elution [116]. For sulfated polysaccharide extraction (e.g., fucoidans), conventional approaches leverage lectin proteins’ specific affinity for fucose residues, enabling selective separation from crude extracts. However, fucoidan’s sulfate groups disrupt lectin–fucose binding, reducing purification yields. As an alternative strategy, dye-based affinity chromatography using cationic dyes like toluidine blue and methylene blue can achieve selective purification. Cationic dyes form stable electrostatic complexes with sulfated polysaccharides via complementary charge interactions [123].

To understand the structure and activity relationships, a molecularly homogeneous preparation is required. Besides being a procedural step, purification critically influences or even defines the observed biological activity of these polysaccharides, often determining their expression or nature [124]. This process is necessary to remove interfering compounds (e.g., polyphenols, proteins) that may obscure the true effect of the polysaccharide itself and to precisely attribute bioactivity to specific structural features [100]. Yet, this technique is challenged by comparative research on Arctic brown seaweeds, where crude polysaccharide extracts exhibited consistently stronger antioxidant and anti-glycemic activities than their purified counterparts. This paradoxical finding reveals that the intrinsic synergistic interactions among compounds in the crude extract enhance bioactivity, or the purification removes essential co-factors needed for the full biological effect [125].

6.3. Quantitative and Qualitative Analyses of Polysaccharides

6.3.1. Colorimetric Assay

Colorimetric assays are analytical methods widely used for the quantitative determination of polysaccharides, relying on measurable color changes resulting from their reactions with specific chemical reagents [80]. Among the various strategies, cationic dye-based assays are noted for their high sensitivity, selectivity, and rapidity. However, their practical application is limited by critical drawbacks, including the requirement for stringently controlled conditions for dye–polyelectrolyte complex formation, as well as a pronounced vulnerability to matrix effects induced by common interferents like salts, acids, or bases [126]. Polysaccharide composition is algal-species-dependent, encompassing different proportions of neutral sugars, uronic acids, and sulfate groups, along with minor constituents such as proteins, polyphenols, and minerals [127]. Total carbohydrate content is typically quantified through two principal colorimetric methods: the phenol–sulfuric acid assay, using glucose as a universal calibration standard with detection at λ = 490 nm [128] and the anthrone method, which employs fucose as a standard for the specific analysis of fucose-rich polysaccharides at λ = 620 nm [129]. Neutral sugars are typically analyzed using the sulfuric resorcinol reaction (glucose standard, λ = 520 nm) [130], while uronic acids are determined via the m-hydroxydiphenyl method (glucuronic acid standard, λ = 525 nm) [131]. For evaluating the degree of sulfation in sulfated polysaccharides, two colorimetric approaches are available: turbidimetric BaCl₂/gelatin precipitation (potassium sulfate standard, λ = 500 nm), as described by Dodgson and Price [132], and Dimethylmethylene Blue assay (chondroitin sulfate standard, λ = 525 nm), according to Farndale et al. [133]. Proteins and polyphenolic compounds, as minority components, are assessed using either the Bradford assay [134] or Lowry method [135] for proteins with bovine serum albumin (BSA) as the standard (λ = 595 nm) and the Folin–Ciocalteu method (gallic acid standard, λ = 760 nm) for polyphenols, as described by Singleton et al. [136].

6.3.2. Methods for Structural and Physicochemical Characterization

The current literature reported that structural features of polysaccharides, such as molecular weight, branching degree, chain conformation, and functional groups, are the primary determinants of their bioactivity and functional performance [28]. However, a comprehensive understanding of each polysaccharide’s characteristics is essential prior to further application [137]. Such comprehensive profiling is achieved through a diverse and sophisticated array of analytical techniques (Figure 8).

Molecular-Weight Determination

High-performance Size-Exclusion Chromatography coupled with the Multiangle Laser Light Scattering and Refractive Index (HPSEC-MALLS-RI) is an analytical technique particularly valuable for the absolute determination of molecular weight and size distribution [28]. This system integrates an HPSEC column for hydrodynamic size-based separation, a MALLS detector for calibration-independent molecular-weight measurement (λ = 658 nm), and an RI detector for concentration quantification. Unlike conventional Size-Exclusion Chromatography (SEC), which relies on calibration standards for relative molecular-weight estimation, the MALLS-RI detection system provides calibration-free and absolute molecular-weight/size determination, allowing for the standardized quality control of polysaccharides [120,122]. Recent HPSEC-MALLS analyses have revealed a substantial variability in the molecular weights of polysaccharides, reporting values for fucoidan (Mw = 122 kDa) from Tubinaria decurrens [104], alginate (100 kDa) from Cystoseira compressa [29], and fucoidan (814 kDa) and alginate (252 kDa) from Halopteris scoparia [113].

Functional Groups Constitution

Fourier transform infrared spectroscopy (FTIR) is a highly effective technique for identifying polysaccharide functional groups. It relies on the analysis of vibrational absorption peaks at certain wave numbers within the 500–4000 cm⁻¹ spectral region [138]. Recently, the synergistic combination of conventional FTIR spectroscopy with Attenuated Total Reflectance (ATR) technology has significantly enhanced standard IR analytical capabilities. Therefore, functional groups of brown algal phycocolloids can be efficiently identified through rapid, direct, and non-destructive FTIR spectroscopy using minimal sample amounts [139]. Moreover, modern FTIR analysis software provides advanced tools for spectral data processing and interpretation. Seaweed-derived algal polysaccharides exhibit distinctive FTIR signatures, including O−H vibrations (3200–3400 cm⁻¹), C−H stretching (2927–2891 cm⁻¹), C=O stretching (1638–1597 cm⁻¹), sulfate ester vibrations (1247–1257 cm⁻¹), and carbohydrate backbone (C−H, 1134–1150 cm⁻¹) absorptions [85].

Chain Conformations Analysis

Marine brown algal polysaccharides display different monosaccharide profiles, featuring glucose (Glc), mannose (Man), galactose (Gal), fucose (Fuc), xylose (Xyl), arabinose (Ara), rhamnose (Rha), mannuronic acid (ManA), and glucuronic acid (GluA) [140], which exhibit varied chain conformations and spatial arrangements [141]. Therefore, the accurate determination of monosaccharide composition investigation is fundamental for elucidating structure–activity relationships. Standard quantitative monosaccharide analysis requires the complete acid hydrolysis of polysaccharides into their constituent monosaccharides, followed by precise quantification using chromatographic systems (Table 3) such as high-performance liquid chromatography (HPLC), high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD), gas chromatography (GC), and ion-exchange chromatography (IEC) [142].

Furthermore, the comprehensive structural analysis of polysaccharides requires the determination of glycosidic linkage positions (e.g., 1,3; 1,4), anomeric configurations (α/β), and branching patterns. Current analytical strategies combine spectroscopic, chemical, and enzymatic approaches. Spectroscopic approaches such as mass spectrometry (MS; e.g., MALDI-TOF, ESI-MS) provide robust information on monosaccharide composition, relative abundance, molecular-weight distributions, and branching topology [143,144], while Nuclear Magnetic Resonance (NMR) yields definitive data on anomeric configurations (α/β) and glycosidic linkage arrangements [145]. In contrast, methylation-assisted GC-MS specifically targets branch point mapping through controlled chemical derivatization [146]. Complementarily, enzyme-mediated hydrolysis targets specific glycosidic bonds, providing the experimental validation of proposed structural models [80].

Table 3. Comparative analysis of chromatographic methods for polysaccharide characterization.

Technique	Acronym	Carrier Gas	Stationary Phase	Separation	Reference
Gas chromatography	GC	Gas (He, H2, and N2)	Solid (silica-based column)	Polarity	[147]
High-performance liquid chromatography	HPLC	Liquid (acetonitrile/water)	Solid (C18, NH2, and HILIC)	Polarity	[148]
Ion chromatography	IC	Liquid (Na2CO3/NaHCO3)	Solid (ion-exchange resin)	Charge	[149]
High-performance anion-exchange chromatography with pulsed amperometric detection	HPAEC-PAD	Liquid (NaOH/KOH)	Solid (polymeric matrices)	Charge Affinity	[150]

Table 3. Comparative analysis of chromatographic methods for polysaccharide characterization.

Technique	Acronym	Carrier Gas	Stationary Phase	Separation	Reference
Gas chromatography	GC	Gas (He, H2, and N2)	Solid (silica-based column)	Polarity	[147]
High-performance liquid chromatography	HPLC	Liquid (acetonitrile/water)	Solid (C18, NH2, and HILIC)	Polarity	[148]
Ion chromatography	IC	Liquid (Na2CO3/NaHCO3)	Solid (ion-exchange resin)	Charge	[149]
High-performance anion-exchange chromatography with pulsed amperometric detection	HPAEC-PAD	Liquid (NaOH/KOH)	Solid (polymeric matrices)	Charge Affinity	[150]

Conformation Analysis

The spatial structure of phycocolloids can be described by a hierarchical model comprising four levels, namely, primary, secondary, tertiary, and quaternary structures [151]. The quaternary structure, defined as an aggregate formed through non-covalent interactions between polysaccharide chains, is a critical determinant of their physicochemical properties and biological functions. Owing to variations in monosaccharide composition, linkage stereochemistry, branching patterns, and inter-molecular forces, these molecules can adopt various conformations in a solution, defining their overall three-dimensional conformation [144]. Furthermore, the three-dimensional architecture dictates their interactions with solvents, ions, and other biomolecules, ultimately defining their functional roles in both natural and industrial contexts. The conformation of polysaccharide molecules is primarily governed by their molecular characteristics in dilute solutions. Key parameters of interest include molecular-weight distribution, chain size and flexibility, main-chain and substituent interactions, and morphological features of polysaccharide molecules. A multidisciplinary approach integrating data from Congo red assays, viscometry, scattering techniques (e.g., small-angle X-ray scattering, light scattering), chromatographic separation, fluorescence spectroscopy, and NMR spectroscopy is typically employed [152]. Furthermore, advanced microscopic techniques, such as Scanning Electron Microscopy (SEM), Atomic Force Microscopy (AFM), and Transmission Electron Microscopy (TEM), serve as a valuable complement to solution-based methods. Multiscale microscopy resolution (from the micro- to nano-level) includes (i) SEM scanning (1–10 nm) for 3D surface topology [153], (ii) AFM (0.1–5 nm) for nanomechanical mapping [154], and (iii) TEM (<0.5 nm) for internal ultrastructural analysis [155].

7. Biological Properties of Brown Algae Polysaccharides

Bioactive polysaccharides from brown algae, such as fucoidan, laminarin, and alginic acid derivatives, exhibit a wide range of biological properties, including anticoagulant, antioxidant, anti-inflammatory, antiviral, and immunostimulatory activities (Figure 9). These bioactivities are primarily attributed to their distinct structural features, such as sulfate group distribution and glycosidic bonding patterns, which enable complex interactions with biological systems [156]. This structure–function relationship makes brown seaweeds valuable sources for next-generation product innovation and functional ingredients design [90]. However, the transition from documented bioactivities into approved large-scale industrial products across pharmaceutical, energy, and environmental domains remains limited, highlighting a significant gap between experimental evidence and market-ready products [157].

7.1. Antioxidant Activity

Oxidative stress (OS) involves a cascade of reactions leading to a significant increase in oxidized cellular components. OS is a major factor in the aging process and can cause direct damage to the central nervous system. Antioxidants comprise a variety of protective components, including enzymatic antioxidants such as glutathione peroxidase, glutathione-S-transferase, catalase, and superoxide dismutase (SOD), as well as non-enzymatic antioxidants like carotenoids, microelements, vitamins, polysaccharides, and other antioxidative substances, which directly or indirectly contribute to the inhibition or suppression of oxidation processes [158]. Natural antioxidants play an important role in preventing ROS formation and eliminating ROS in organisms [159,160]. Numerous studies have demonstrated that brown algae polysaccharides, particularly sulfated ones like fucoidan, exhibit robust antioxidant properties in various in vitro models (Table 4). Their biological efficacy is largely determined by their distinctive structural characteristics—such as the degree of sulfation, glycosidic branching, and molecular weight—which synergistically enhance their electron-donating capacity, metal-ion chelation, and broad-spectrum free radical scavenging activity [28]. Common assays to evaluate the radical scavenging abilities of antioxidants in vitro include DPPH radical scavenging, nitric oxide scavenging, ABTS radical scavenging, hydroxyl radical scavenging, the beta-carotene–linoleic acid system, ferric reducing power, and superoxide radical scavenging [161,162]. Consequently, significant research efforts are directed toward natural antioxidants, particularly polysaccharides, as alternatives to synthetic antioxidants traditionally used in the food and pharmaceutical industries, such as tert-butyl hydroquinone (TBHQ), butylated hydroxyanisole (BHA), and butylated hydroxytoluene (BHT), due to their potential carcinogenicity and possible hepatotoxic effects [163].

Table 4. Antioxidant assays of sulfated polysaccharides from different brown seaweed species.

Species	Polysaccharide	Antioxidant Activity	Reference
Sargassum policystum	Fucoidan	FRAP: IC50 = 91.3 ppm	[164]
Dictyota dichotoma	Fucoidan	Total antioxidant capacity (TAC): 71.76 ± 3.23% DPPH scavenging: 51.13 ± 2.31% H2O2 scavenging: 40.45 ± 1.87% ABTS scavenging: 50.51 ± 2.27% NO scavenging: 38.13 ± 1.72% Ferrous ion chelating: 18.66 ± 0.84%	[162]
Cystoseira schiffneri	Fucoidan	DPPH scavenging: IC50 = 104 ± 5 µg/mL (Ref: BHA IC50 = 14 ± 0.2 µg/mL) Ferrous ion chelating: IC50 = 96 ± 3 µg/mL (Ref: EDTA IC50 = 10 ± 0.2 µg/mL) FRAP: IC50 = 63 ± 3 µg/mL (Ref: BHA IC50 = 25 ± 0.1 µg/mL)	[165]
Ericaria crinita	Fucoidan	DPPH scavenging: IC50 = 412 µg/mL FRAP: 118.72 µM Trolox Equivalent/g	[166]
Padina gymnospora	Fucoidan	DPPH scavenging: IC50 = 0.5256 ± 0.05 mg/mL (Ref: BHA IC50 = 0.31 ± 0.03 mg/mL) ABTS scavenging: IC50 = 2.565 ± 0.02 mg/mL (Ref: ascorbic acid IC50 = 0.2859 ± 0.02 mg/mL)	[167]
Padina boryana	Fucoidan	In vitro (reduced intracellular ROS levels, protection against apoptosis) In vivo, zebrafish model (reduced ROS levels, reduced lipid peroxidation)	[159]
Colpomenia sinuosa	Fucoidan	DPPH scavenging: IC50 = 46.2 ± 1.4 µg/mL SOD-like activity: IC50 = 23.7 ± 1.1 µg/mL	[152]
	Sodium alginate	DPPH scavenging: IC50 = 280 ± 1.2 µg/mL SOD-like activity: IC50 = 41.34 ± 1.07 µg/mL

Table 4. Antioxidant assays of sulfated polysaccharides from different brown seaweed species.

Species	Polysaccharide	Antioxidant Activity	Reference
Sargassum policystum	Fucoidan	FRAP: IC50 = 91.3 ppm	[164]
Dictyota dichotoma	Fucoidan	Total antioxidant capacity (TAC): 71.76 ± 3.23% DPPH scavenging: 51.13 ± 2.31% H2O2 scavenging: 40.45 ± 1.87% ABTS scavenging: 50.51 ± 2.27% NO scavenging: 38.13 ± 1.72% Ferrous ion chelating: 18.66 ± 0.84%	[162]
Cystoseira schiffneri	Fucoidan	DPPH scavenging: IC50 = 104 ± 5 µg/mL (Ref: BHA IC50 = 14 ± 0.2 µg/mL) Ferrous ion chelating: IC50 = 96 ± 3 µg/mL (Ref: EDTA IC50 = 10 ± 0.2 µg/mL) FRAP: IC50 = 63 ± 3 µg/mL (Ref: BHA IC50 = 25 ± 0.1 µg/mL)	[165]
Ericaria crinita	Fucoidan	DPPH scavenging: IC50 = 412 µg/mL FRAP: 118.72 µM Trolox Equivalent/g	[166]
Padina gymnospora	Fucoidan	DPPH scavenging: IC50 = 0.5256 ± 0.05 mg/mL (Ref: BHA IC50 = 0.31 ± 0.03 mg/mL) ABTS scavenging: IC50 = 2.565 ± 0.02 mg/mL (Ref: ascorbic acid IC50 = 0.2859 ± 0.02 mg/mL)	[167]
Padina boryana	Fucoidan	In vitro (reduced intracellular ROS levels, protection against apoptosis) In vivo, zebrafish model (reduced ROS levels, reduced lipid peroxidation)	[159]
Colpomenia sinuosa	Fucoidan	DPPH scavenging: IC50 = 46.2 ± 1.4 µg/mL SOD-like activity: IC50 = 23.7 ± 1.1 µg/mL	[152]
	Sodium alginate	DPPH scavenging: IC50 = 280 ± 1.2 µg/mL SOD-like activity: IC50 = 41.34 ± 1.07 µg/mL

7.2. Anti-Inflammatory Activity

Inflammation is the complex biological response of vascular tissues to harmful stimuli, such as pathogens, tissue damage, or deleterious agents, serving as a protective mechanism [168]. The inflammatory response involves precisely coordinated steps for effective immunity. Initially, immune cells recognize pathogens or damaged cells through pattern recognition receptors (PRRs), which identify specific molecular patterns. This recognition activates intracellular signaling pathways, such as NF-κB, MAPK, and JAK-STAT, leading to the production of inflammatory mediators. Activated immune cells then release these mediators, including cytokines and chemokines, which amplify the inflammatory response and recruit additional immune cells to the site of injury or infection. Once the harmful stimulus is eliminated, anti-inflammatory signals promote a resolution by inducing apoptosis of activated immune cells and initiating tissue repair mechanisms to restore homeostasis [169]. Studies across multiple brown algal species have consistently identified their polysaccharides as potent anti-inflammatory agents.

A sulfated heteropolysaccharide (PSCH) isolated from the brown alga Cystoseira humilis has been investigated for its anti-inflammatory potential. PSCH is mainly composed of fructose, xylose, galacturonic acid, arabinose, and glucuronic acid. In vivo experiments demonstrated pronounced anti-inflammatory activity, particularly in carrageenan-induced paw edema in rats, where treatment significantly reduced paw size. This response, which typically involves elevated leukocyte recruitment mediated by pro-inflammatory cytokines, was significantly attenuated by PSCH administration, suggesting the inhibition of cytokines as a likely mechanism. Additionally, platelet counts in treated rats remained comparable to healthy controls, suggesting that PSCH may modulate pro-inflammatory pathways involved in platelet synthesis. Overall, these findings highlight the potential of PSCH as a bioactive compound with cytokine-suppressive and platelet-modulatory effects contributing to its anti-inflammatory activity [170].

Another study reported by Wang et al. [171] investigated the mechanisms underlying the anti-inflammatory activity of two novel fucoidan fractions, ANP−6 and ANP−7, isolated from Ascophyllum nodosum, and further examined the relationship between their structural features and bioactivity. Both ANP−6 and ANP−7 were shown to modulate nitric oxide (NO) production and the mRNA expression of inflammatory mediators, such as iNOS, COX−2, TNF−α, IL−1β, IL−6, and IL−10 in lipopolysaccharide (LPS)-induced RAW 264.7 macrophages. Notably, the anti-inflammatory potential was closely associated with ANP−6 and ANP−7 structural features, which are also linked with the TLR/NF−κB signaling pathway, as evidenced by reduced expression levels of Toll-like receptors (TLR−2 and TLR−4). The lower molecular weight of ANP−6 correlated with the enhanced anti-inflammatory efficacy, underscoring the influence of molecular weight on fucoidan bioactivity [172].

Furthermore, the anti-inflammatory potential of SLCF5, a fucoidan fraction from Scytosiphon lomentaria, was demonstrated by its inhibition of LPS-induced NO production in macrophages. LPS stimulation induced prostaglandin E2, pro-inflammatory cytokines, and the phosphorylation of NF-κB and MAPK-related proteins, while SLCF5 treatment significantly downregulated their expression. Additionally, the in vivo results showed that SLCF5 increased the survival percentage while decreasing cell death, NO generation, and heart rate. In vivo experiments revealed that SLCF5 increased survival rates while decreasing cell death, NO generation, and heart rate [173].

7.3. Antibacterial Activity

The increasing prevalence of multidrug-resistant pathogens constitutes a significant challenge for public health worldwide, substantially reducing the effectiveness of standard antibiotic interventions against bacterial infections [174]. Moreover, the development of microbial biofilms represents a crucial survival strategy that enhances bacterial resistance against antimicrobial agents [175]. As a result, therapeutic options for such infections are increasingly constrained and frequently ineffective, leading to chronic and recalcitrant conditions. Therefore, the development of therapeutic alternatives with novel mechanisms of action constitutes a pressing need [176]. Accordingly, algal polysaccharides present a promising source of such novel agents, owing to their potent dual antibacterial and antibiofilm activities (Table 5). Their activity is mediated through multiple mechanisms, including membrane disruption, the inhibition of bacterial adhesion, and suppression of virulence factor expression [177]. Experimental evaluation commonly relies on standard in vitro methods, including Minimum Inhibitory Concentration (MIC) assays, biofilm biomass quantification (e.g., crystal violet staining), and confocal microscopy, while transcriptomic and proteomic analyses provide deeper insights into the molecular basis of their anti-virulence and antibiofilm properties [178].

Table 5. Antibacterial activity of brown algae polysaccharides.

Polysaccharide	Source	Mode of Action	Target Pathogens	Reference
Fucoidan	Sargassum spp.	Antibiofilm activity	Streptococcus mutan Porphyromonas gingivalis Fusobacterium nucleatum	[179]
Fucoidan	Sargassum polycystum	Antibacterial activity	Pseudomonas aeruginosa Staphylococcus aureus Escherichia coli Streptococcus mutans	[180]
Laminarin, Alginate	Alaria esculenta	Antibacterial activity Anti-yeast activity	Bacillus subtilis Escherichia coli Saccharomyces cerevisiae	[181]
Fucoidan	Sargassum wightii	OmpF porin inhibition	Salmonella typhi	[182]
Fucoidan	Fucus vesiculosus	Antibacterial activity	Escherichia coli Staphylococcus aureus Bacillus subtilis Pseudomonas aeruginosa	[183]

Table 5. Antibacterial activity of brown algae polysaccharides.

Polysaccharide	Source	Mode of Action	Target Pathogens	Reference
Fucoidan	Sargassum spp.	Antibiofilm activity	Streptococcus mutan Porphyromonas gingivalis Fusobacterium nucleatum	[179]
Fucoidan	Sargassum polycystum	Antibacterial activity	Pseudomonas aeruginosa Staphylococcus aureus Escherichia coli Streptococcus mutans	[180]
Laminarin, Alginate	Alaria esculenta	Antibacterial activity Anti-yeast activity	Bacillus subtilis Escherichia coli Saccharomyces cerevisiae	[181]
Fucoidan	Sargassum wightii	OmpF porin inhibition	Salmonella typhi	[182]
Fucoidan	Fucus vesiculosus	Antibacterial activity	Escherichia coli Staphylococcus aureus Bacillus subtilis Pseudomonas aeruginosa	[183]

7.4. Antidiabetic Activity

Diabetes is a chronic disease that arises either from insulin deficiency due to pancreatic beta-cell dysfunction or from impaired glucose homeostasis resulting from insulin deficiency. According to the World Health Organization (WHO), diabetes mellitus was responsible for an estimated 2 million deaths in 2021, making it one of the leading causes of premature mortality. Furthermore, hyperglycemia was associated with approximately 11% of cardiovascular-related deaths worldwide [184]. Brown algal bioactive polysaccharides have been reported to lower blood glucose levels through multiple mechanisms, including the modulation of glucose metabolism, stimulation of insulin secretion, and protection of pancreatic β−cells from postprandial hyperglycemia. Additionally, their ability to inhibit carbohydrate-hydrolyzing enzymes, including α-amylase and α-glucosidase, further contributes to reducing postprandial glucose levels, supporting an effective therapeutic approach for type 2 diabetes [30,185].

Recent investigations have identified a sulfated polysaccharide from Undaria pinnatifida, composed primarily of rhamnose, glucuronic acid, and fucose, which exhibited superior α-glucosidase inhibitory efficacy compared to acarbose, a standard antidiabetic drug, at a concentration of 100 μg/mL. In vivo evaluation using oral maltose and oral sucrose tolerance tests in C57BL/6J mice revealed significant anti-hyperglycemic effects. Specifically, during both the oral maltose and sucrose tolerance tests, which are commonly used to assess the inhibitory activity of compounds on α-glucosidase-mediated carbohydrate hydrolysis, the administration of the sulfated polysaccharide at a dose of 50 mg/kg resulted in a significant reduction in blood glucose levels within 30–60 min, exhibiting an effect comparable to acarbose [186].

Furthermore, the antidiabetic potential of alginate extracted from Laminaria digitata has been studied as an inhibitor of α-amylase. For this polysaccharide, the Km values increased due to the inhibitors’ competition with the substrates for a limited number of α-amylase active sites. In comparison to the control sample, the Vmax decreased when alginate was added. This demonstrates that the inhibitors are unable to attach to the enzyme’s active site. Instead, they bind allosterically to a different site on α-amylase, thereby affecting the enzyme–substrate complex and slowing the rate of reaction [187].

As noted by Daub et al. [188], fucoidan, isolated from Ecklonia maxima, has been identified as a potent mixed-type inhibitor of α-glucosidase compared to standard antidiabetic compounds. This sulfated polysaccharide may hold significant promise for the treatment of type 2 diabetes through the efficient regulation of postprandial hyperglycemia. The proposed inhibition mechanism involves direct interaction with the enzyme, independent of the enzyme–substrate complex, leading to the effective suppression of carbohydrate digestion. Notably, E. maxima fucoidan may avoid the gastrointestinal side effects commonly associated with acarbose therapy. These attributes make these bioactive molecules an ideal therapeutic candidate for the enzyme-targeted treatment of type 2 diabetes, particularly concerning action between starch and α-amylase.

7.5. Anticancer Activity

Cancer is a highly complex and multifactorial disease characterized by the uncontrolled growth and division of atypical cells. Unlike healthy cells, which follow a regulated cycle of growth, division, and programmed death (apoptosis), cancer cells evade these regulatory mechanisms, thereby enabling their uncontrolled proliferation. Recent research has highlighted the potential of polysaccharides derived from brown algae in cancer prevention and therapy. These bioactive compounds have been shown to inhibit the growth of cancer cells, induce cell death, and enhance immune system-mediated tumor suppression [189].

Fucoidan extracted from Spatoglossum vietnamense exhibited significant anticancer activity in vitro against human colon carcinoma cell lines HCT-116, HT-29, and DLD-1. These findings highlight a clear structure–activity relationship, showing that higher sulfate content increased colony formation inhibition from 2030% in low-sulfated fractions to 40–50% in highly sulfated fractions [190].

In vitro assays demonstrated that fucoidan from Dictyota dichotoma decreased the viability of Ehrlich ascites carcinoma cells in a concentration and time-dependent manner. At concentrations ranging from 0.05 to 1 mg/mL, fucoidan caused a significant decrease in cell viability over a 30 min exposure period. These findings emphasize the potential of fucoidan as an antitumor agent, particularly in the context of solid tumors [162].

Laminarin exhibited cytotoxic effects against human colon cancer cells (HT-29), predominantly through apoptosis. DNA fragmentation assays confirmed the induction of cell death in both cancer cell lines. Mechanistic investigations revealed that laminarin suppresses tumor cell proliferation and triggers apoptosis, reactive oxygen species (ROS) generation, DNA fragmentation, and activation of apoptotic signaling pathways. Additionally, laminarin promotes endoplasmic reticulum (ER) stress by disrupting calcium homeostasis and altering the ER–mitochondria axis, ultimately leading to mitochondrial dysfunction. These results highlight the potential of laminarin as a selective anticancer agent [191,192].

Alginate derived from Colpomenia sinuosa further demonstrated anticancer activity by reducing HCT-116 colon cancer cell viability in a dose- and time-dependent manner, mediated by increased ROS levels and apoptotic cell death. These findings highlight a clear structure–activity relationship, as higher sulfate content enhanced colony formation inhibition from 20–30% in low-sulfated fractions to 40–50% in highly sulfated fractions [152].

7.6. Others

Polysaccharides extracted from brown algae, such as fucoidan from Sargassum binderi and L-fucose-rich sulfated glycans from Scytosiphon lomentaria, demonstrated significant anti-obesity potential. In vivo studies in rats fed a high-fat diet showed that fucoidan treatment reduced body weight by 36% and visceral fat by 18% and also enhanced metabolic markers like insulin, leptin, and serum lipid profiles without inducing liver or kidney toxicity. Likewise, L-fucose-rich sulfated polysaccharides have been reported to suppress adipogenesis in vitro, modulating key regulators, including AMPK and PPARγ, while in zebrafish models, they increased energy expenditure and decreased lipid accumulation. Collectively, these findings highlight the potential of brown algae polysaccharides as natural functional agents for obesity management through their combined effects on fat reduction, metabolic regulation, and energy expenditure [75,174].

Furthermore, sulfated fucoidans from brown algae exhibit significant anticoagulant and antithrombotic effects, as demonstrated in recent studies. These compounds disrupt the coagulation cascade by inhibiting thrombin and factor Xa, modulating platelet aggregation, and improving fibrinolysis, illustrating a well-established structure–activity relationship where sulfation patterns and molecular size determine potency. Numerous studies highlight the diversity of marine polysaccharides that are efficient in producing these anticoagulant phenomena and provide experimental evidence, indicating that variations in fucoidan structures may improve or reduce these effects. Overall, these studies demonstrated that specific structural features of marine polysaccharides directly translate into measurable anticoagulant and antithrombotic activities, showing their potential as natural alternatives to conventional anticoagulants [193,194].

Polysaccharides derived from brown marine algae also exhibit a broad spectrum of neuroprotective properties including antioxidant activity that mitigates oxidative stress in neurons. They revealed anti-inflammatory effects through the downregulation of pro-inflammatory cytokines and anti-apoptotic actions that prevent programmed cell death in neuronal cells. In addition, these compounds display anti-amyloidogenic activities, inhibiting the aggregation of toxic amyloid−β proteins implicated in Alzheimer’s disease. In addition, they support mitochondrial function and protein clearance pathways, thereby maintaining neuronal integrity. Collectively, these mechanisms underscore a multi-targeted neuroprotective potential, enhancing neuronal survival, improving cognitive performance, and conferring resilience against neurotoxic stressors such as glutamate, hydrogen peroxide, amyloid−β, and ischemic injury [195].

8. Conclusions and Future Perspectives

Brown algae represent an approachable and abundant source of functional polysaccharides. Their biocompatible, biodegradable, and non-toxic properties have garnered significant industrial interest. Unlocking this potential requires the development of optimized and integrated processing value chains that simultaneously ensure product quality, cost-effectiveness, and environmental sustainability. This critical requirement has driven considerable innovation, resulting in advanced extraction, purification, and characterization techniques. Now, sustainable technologies (e.g., UAE, MAE, and SFE) and green solvents offer promising alternatives to conventional processing methods. Despite these advances, commercial-scale exploitation of brown algal polysaccharides faces several persistent technical and economic challenges. A primary obstacle is the lack of standardized processing protocols, leading to inconsistent outcomes due to methodological variations and source diversity. Additionally, while the bioactivities of these polysaccharides are well-established, their mechanistic pathways remain poorly elucidated. Further progress requires the integration of advanced molecular techniques, particularly proteomics, transcriptomics, and metabolomics, to identify key genes governing biosynthetic and regulatory pathways. Addressing these challenges through interdisciplinary collaboration represents a critical opportunity to unlock the full potential of brown algal polysaccharides for sustainable industrial development.