Fucoidan-Derived Functional Oligosaccharides: History
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Contributor:
Min Wang
,
Suresh Veeraperumal
,
Saiyi Zhong
,
Kit-Leong Cheong

Oligosaccharides derived from natural resources are attracting increasing attention as both food and nutraceutical products because of their beneficial health effects and lack of toxicity. New interest has emerged in fucoidan, partially hydrolysed into fuco-oligosaccharides (FOSs) or low-molecular weight fucoidan, owing to their superior solubility and biological activities compared with fucoidan. There is considerable interest in their development for use in the functional food, cosmetic, and pharmaceutical industries. 

  • fuco-oligosaccharides
  • brown algae
  • preparation
  • biological activities
  • prebiotics

1. Introduction

Fucoidan is a polysaccharide mainly composed of fucose, with a high sulphate content (more than 20%). This carbohydrate moiety is found in marine resources, generally in brown algae, sea cucumbers, and sea urchin eggs (as indicated in Table 1). The chemical structure of fucoidans consists of a backbone of α-(1→3)-L-fucopyranose residues, or alternating α-(1→3)-linked and α-(1→4)-linked L-fucopyranose or α-(1→2)-L-fucopyranose residues (Figure 1), with different substitutions, such as fucose, sulphate, acetate, and uronic acid [1][2]. Fucoidan presents a wide range of pharmaceutical activities, including antioxidant, anticancer, immunomodulatory, anticoagulant, and antimicrobial activities [3][4].
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Figure 1. The chemical structure of fucoidan. (A) backbone of α-(1→3)-L-fucopyranose residues; (B) backbone alternating α-(1→3)-linked and α-(1→4)-linked L-fucopyranose residues. R = OH or OSO3−.
Table 1. Sources of fucoidan derived from brown algae, sea cucumber, and sea urchin species.
Sources Species
Brown algae Adenocystis utricularis, Ascophyllum nodosum [5], Bifurcaria bifurcate [6], Cladosiphon okamuranus [6], Coccophora langsdorfii [7], Cystoseira sedoides [7], Desmarestia intermedia [6], Dictyosiphon foeniculaceus [6], Dictyota dichotoma [6], Ecklonia kurome [6], Eisenia bicyclis [6], Fucus vesiculosus [5], F. serratus [6], F. spiralis [6], Laminaria digitate [5], L. saccharina [5], Saundersella simplex [6], Sargassum siliquosum [6], Scytosiphon lomentaria [6], Undaria pinnatifida [6]
Sea cucumber Acaudina leucoprocta [2], A. molpadioides [2], Holothuria fuscopunctata [2], H. polii [2], Isostichopus badionotus [2], Ludwigothurea grisea [2], Stichopus horrens [2], Thelenota ananas [2]
Sea urchin Arbacia lixula [6], Lytechinus variegatus [6], Strongylocentrotus droebachiensis [8], S. franciscanus [6], S. intermedius [6], S. pallidus [8], S. purpuratus [6]
In fact, the preparation and structural characterisation of fucoidan seem to be well elucidated. However, haemorrhagic risk, high molecular weight (MW), high viscosity of fucoidan, and a poor dissolution rate in solution may limit their successful application in functional foods and pharmaceuticals [9][10]. Fucoidans often differ in terms of structural diversity and complexity; therefore, standardisation and quality assurance of fucoidans are challenging. Therefore, there has been a recent increase in interest in degrading fucoidan into fuco-oligosaccharides (FOSs) or low-MW fucoidan (LMWF) to broaden potential applications.
Oligosaccharides consist of a mixture of 2–20 monosaccharides, with a variable extent of polymerisation. They have attracted increasing interest owing to their nutritional and biological properties [11]. Many functional oligosaccharides, such as fructo-oligosaccharides, galacto-oligosaccharides, xylo-oligosaccharides, manno-oligosaccharides, and isomalto-oligosaccharides, have been widely used as prebiotics in food application [12][13][14]. Commercial prebiotics are derived from terrestrial plant oligosaccharides. However, research is underway to explore new prebiotics from natural resources, especially from renewable sources. Marine algae have unparalleled advantages as easily renewable and extremely abundant resources, as they grow in seawater and avoid the demand for freshwater. FOSs, also called LMWF, is an oligomer of fucoidan, which is normally obtained from brown algae. FOSs have a degree of polymerisation (DP) < 20 and an average MW of less than 10 kDa. FOSs also show a prebiotic ability and can be fermented by the gut microbiota, exert a range of physiological effects on body functions and improve human health [15]. Therefore, FOSs and other marine algae oligosaccharides are considered promising prebiotics.

2. Preparation of Fucoidan and/or Raw Materials for Fuco-Oligosaccharide Production

The advantages of FOSs include high water solubility, low viscosity, and good biological activity. However, FOSs are seldom found in natural resources. Therefore, the degradation of fucodain is necessary for the production of FOSs. Figure 2 shows a singular diagram of the process of preparing fucoidan from brown algae. The extraction processes are based on aqueous (water, acidic, or alkaline solution) extraction; the polysaccharides are precipitated by ethanol; proteins are removed by the Sevag or enzymatic method, and the low-molecular-weight impurity is discarded by diafiltration [16]. The Sevag method is widely used to discard protein. It was first exploited by Sevag et al., as they used an organic solvent, including chloroform and n-butanol, to precipitate protein from sample solution [17]. An acid or calcium chloride solution added to the polysaccharide mixture has been commonly used to separate alginic acid from fucoidan. Béress et al. precipitated the alginic acid by adding glacial acetic acid, and the process was completed after 12 h at 4 °C [18]. Furthermore, column chromatography, such as ion-exchange and size-exclusion methods, can be used to obtain fucoidan [19]. Currently, three major processes exist to obtain FOSs from fucoidan: degradation by mild acids, enzymes, and radical methods. The mild acids and radical degradation methods are considered to be random and non-specific in degrading fucosidic linkages, while enzymatic method recognizes the specific substrate to cleave fucosidic linkages.
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Figure 2. Schematic diagram of the extraction and preparation of fucoidan from brown algae.

3. Preparation of Fuco-Oligosaccharides

3.1. Mild Acid Hydrolysis

Mild acid hydrolysis usually uses dilute acid solutions, such as sulphuric acid, hydrochloric acid, and trifluoroacetic acid, at suitable heating temperatures and times. FOSs mixtures are obtained after hydrolysis and subsequent neutralisation. Mild acid hydrolysis seems to offer a simple, low-cost production, and an easy operation, especially to fulfil requirements in the laboratory. Therefore, the mild acid hydrolysis method is always adopted by laboratories to screen for potential biological activities of FOSs. An aliquot of 0.05 mol/L sulphuric acid solution was used to degrade Nemacystus decipiens fucoidan at 80 °C for 2 h to obtain FOSs with a DP ranging from 2 to 9, and the FOSs showed antithrombotic activity [20]. A high-MW fucoidan, derived from S. hemiphyllum, was degraded by 0.01 mol/L hydrochloric acid at 85 °C for 15 min to obtain FOSs with a MW of 800 Da. The result showed skin-protective effects against ultraviolet B damage [21].
However, the degradation method is non-specific and may produce a series of FOSs, which increases the challenge of purifying FOSs. Moreover, the sulphated group in fucoidan is acid-labile and easily hydrolysed during acid hydrolysis. Mild acid hydrolysis (parameters used 0.005–0.01 mol/L trifluoroacetic acid at 60–100 °C for 2–4 h) was conducted on fucan sulphate derived from S. herrmanni, to obtain a number of FOSs. They contained two series of FOSs: 2-desulphated and 2-sulphated residues at the reducing ends of FOSs [22]. Pomin et al. found that in the dilute acid hydrolysis of fucoidan, the 2-sulphate ester group is initially cleaved, subsequently degrading the glycosidic bonds between the non-sulphated residue and the 4-sulphate group [23]. The kinetic constant of mild acid hydrolysis on sulphated fucan derived from three sea cucumber species, I. badionotus, H. floridana, and L. variegatus, with core α (1→3)-linked fucose, degraded the sulphate group substitution at the 2-O and 4-O-positions. The mild acid hydrolysis progress was under 0.05 mol/L sulphuric acid at 60 °C for 9 h, and selective 2-desulphation in sulphated fucan and tetrasaccharides FOSs were obtained [24].
Mild acid hydrolysis is suitable for a large-scale production of FOSs. Future studies on the kinetics and mechanism of mild acid hydrolysis should focus on providing fundamental data for industrial application in the large-scale production of FOSs.

3.2. Enzymatic Hydrolysis

Several hydrolytic enzymes that catalyse the hydrolysis of polysaccharides have been characterised. Normally, the breakdown of fucoidans involves the degradation of glycosidic linkages by fucoidanases (EC 3.2.1.44) and α-L-fucosidases (EC 3.2.1.51, EC 3.2.1.63, EC 3.2.1.111, and EC 3.2.1.127), which converts fucoidans into FOSs [25]. The enzymes that produce FOSs are produced by marine microorganisms and include endo- and exo-enzymes [26]. The well characterized endo-enzymes for fucoidan hydrolysis include endo-fucanases FcnA, FFA1, FFA2, Fp273, Fp277, Fp279, FWf1, FWf2, Fhf1, Fhf2 of the GH107 family and endo-fucanase FunA of the GH168 family, while the α-L-fucosidase is exo-enzymes and described for the families of glycoside hydrolases (GH) 29, 95, and 141 [25][27][28]. Enzymatic depolymerisation is highly selective for specific substrates and efficient for producing the desired end-product under selective method conditions. Therefore, with the enzymatic method, unlike the mild acid hydrolysis method, it is possible to achieve hydrolysate of crude fucoidan yielding FOSs.
The extent of enzymatic action is strongly dependent on operating conditions, including temperature and pH. Fucoidan with a MW of approximately 400 kDa from F. distichus was incubated together with endo-α-(1,4)-fucoidanase from the marine bacterium Formosa haliotis (Fhf1Δ470), in Tris-HCl buffer (pH 8) at 37 °C for 1 d, yielding FOSs with a MW of 2 kDa [29]. The study demonstrated the use of the enzymatic technique to screen out the tailored fucoidan functional group (FOSs) to specifically improve bone regeneration [29]. Enzymatic depolymerisation of L. japonica fucoidan by fucoidanase—which is produced by Flavobacteriaceae RC2-3—yielded FOSs with a MW < 2 kDa (96.3%) after incubation at 37 °C for 10 h. The FOSs products showed an excellent radical scavenging activity and inhibition of tyrosinase activity, rendering them suitable for application as cosmetic skin-whitening additives [30]. Another advantage of the enzymatic method is that the sulphated groups in FOSs may be preserved during the depolymerisation process. A fucoidanase obtained from the digestive glands of mollusc Lambis sp., specifically, degraded (1→3) and (1→4)-α-L-fucans derived from F. evanescens and F. vesiculosus, yielding sulphated FOSs, without any effect on the sulphate group [31].
To overcome problems of using the enzyme, including those owing to its high cost and poor stability, the immobilised enzyme technique is preferred. The immobilised enzyme technique has the advantages of maintaining enzymatic activity, ease of recovery after coating monolithic substrates, and easy scaling up of processes for industrial application [32].
The enzymatic method is considered a good choice for producing FOSs for applications in the food, cosmetic, and pharmaceutical industries. Therefore, screening for new enzymes that can degrade fucoidan has become an increasingly important focus of scientists and industrial engineers; in particular, searching for new endo-fucoidanases and their subsequent kinetic analysis, molecular cloning, and immobilisation techniques to provide valuable fundamental knowledge to produce highly active FOSs.

3.3. Radical Depolymerisation

Radical depolymerisation is a process that uses radicals to attack glyosidic linkages of polysaccharides to form oligosaccharides. The radicals, including HOO−•, •O2−, and HO•, can be generated by hydrogen peroxide (H2O2) and a catalytic metal system. Depolymerisation by H2O2 involves radicals (HOO•, •O2−, and HO•) attacking the glycosidic linkages [33]. This method has the advantages of good reproducibility and controllability. The main parameters that influence the efficacy of depolymerisation include the temperature, concentration of H2O2, reaction time, and substrate concentration.
Radical depolymerisation of fucan sulphate from S. herrmanni was performed by adding H2O2 and copper (II) acetate monohydrate. On average, the MW of fucan sulfate was degraded from 790.8 kDa to 8.32–22.11 kDa. The resulting products exhibited good anticoagulant activity [34]. Moreover, a combination of ascorbic acid and H2O2 was used to degrade fucoidan at 25 °C for 16 h, to obtain a fraction with the MW of approximately 3 kDa, which exhibited anti-lung-cancer activity in vitro [35]. Qi et al. reported that radical depolymerisation can maintain the sulphate content or other functional units in the degradation process [36]. Qi et al. degraded U. pinnatifida fucoidan using a photocatalytic method, which processes the reaction in H2O2 and TiO2 under xenon light for 3 h. While the MW of 190 kDa was degraded to 3 kDa, the resulting product exhibited anticoagulant activity [36].
Moreover, photoirradiation techniques for the degradation of polysaccharides have been attracting increasing attention. These techniques provide efficient degradation methods for the production of FOSs from fucoidan, because photoirradiation can be achieved at room temperature and under normal pressure. Ultraviolet and gamma radiation are typically used to degrade fucoidan. They are an energetic form of electromagnetic radiation, with a short wavelength and high energy. Water radiolysis is the major effect of gamma irradiation of solutions. Higher gamma irradiation doses may result in a lower MW of fucoidan: the MW of F. vesiculosus fucoidan degraded from 210 kDa to 85 kDa and 7 kDa following irradiation doses of 8 kGy and 100 kGy, respectively, while the latter degraded product showed the highest antioxidant activity [37]. In addition, photoirradiation combined with radical depolymerisation increased the efficiency of FOSs production. The average MW of fucoidan decreased to 6.7 kDa at an irradiation dose of 20 kGy, while the degradation efficiency increased by 5% when a 10% H2O2 concentration was used in combination with gamma radiation. The hydroxyl radical scavenging abilities of degraded product of FOSs were more effective than fucoidan [38]. Park and Choi also used a combination of gamma radiation at a dose of 100 kGy and H2O2 to degrade U. pinnatifida fucoidan from MW 378 kDa to 6 kDa. Park and Choi found that degraded fucoidan has a higher inhibitory activity against tyrosinase and antioxidant activity than fucoidan [39].

4. Purification of Fuco-Oligosaccharides

The mixtures of FOSs produced by degradation methods include a variety of compounds, i.e., monosaccharides, FOSs ranges of different DPs, and large MW fucoidan. To obtain high-purity FOSs (for use in studies of the structure–function relationship and mechanism, and for applications in functional food or pharmaceutical areas), some suitable purification steps should be performed. Figure 3 shows a schematic diagram of the production and purification of FOSs.
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Figure 3. Schematic diagram of FOS degradation and purification processes.
Step-gradient ethanol precipitation is a convenient and time-saving method for the large-scale production of FOSs. Different MWs can be prepared by adding an ethanol gradient at a final concentration of 10–80%. However, this method yields a broad MW distribution of FOSs, and it is difficult to obtain high-purity FOSs. For example, Fernando et al. performed a step-gradient ethanol precipitation to exclude high-MW fucoidan, and a broad MW distribution of FOSs, ranging from 8 to 25 kDa, was achieved [40]. Thus, ethanol precipitation is useful as a preliminary step for discarding high-MW fucoidans or proteins.
Membrane filtration is another green technique for the purification of FOSs, and its performances are quite similar to gradient ethanol precipitation, which obtained a broad MW distribution of FOSs. Zhao et al. purified a degraded fucoidan mixture, using an ultrafiltration membrane (10 kDa and 2.5 kDa MW cut-off) to obtain the FOSs fraction, and the fraction range 2.5–10.0 kDa was collected [9]. The degraded fucoidan samples passed through a 30-kDa-MW cut-off membrane and subsequently through a 1-kDa membrane, obtaining an FOSs fraction MW range of 1–30 kDa [41]. Dialysis bags with different MW cut-off membranes have been used to obtain MW values of lower than 10 kDa fraction [42].
Column chromatography, including ion-exchange chromatography and size-exclusion chromatography, is widely used for fractionation oligosaccharides during purification. Size-exclusion chromatography has been used to fractionate oligosaccharides according to their MW, and this is suitable for the purification of FOSs and achieving high purity. Commercially available size-exclusion chromatography resins have been used to purify FOSs. The purified FOSs fractions are suitable for structural identification and pharmaceutical activity investigation. After mild acid hydrolysis the fucoidan hydrolysate sample was purified using a Bio-Gel P-2 column, and a high-purity fucotrioligosaccharide was obtained [43]. A mixture of oligosaccharides, produced from acid hydrolysis, was further purified using Bio-Gel P-4 and P-2, to obtain a high-purity yield of 6–9% of monosulphated fucobiose and monosulphated fucotriose [44]. Ion-exchange chromatography for FOSs purification proceeds according to the sulphate content of the FOSs, which leads to different ionic interactions with the resins. The FOSs mixture produced from fucoidan was degraded by the enzymatic method and further purified by ion-exchange chromatography on Q-Sepharose, to yield DP 4–10 FOSs with different sulphate contents [45].

This entry is adapted from the peer-reviewed paper 10.3390/foods12040878

References

  1. Ale, M.T.; Meyer, A.S. Fucoidans from brown seaweeds: An update on structures, extraction techniques and use of enzymes as tools for structural elucidation. RSC Adv. 2013, 3, 8131–8141.
  2. He, W.; Sun, H.; Su, L.; Zhou, D.; Zhang, X.; Shanggui, D.; Chen, Y. Structure and anticoagulant activity of a sulfated fucan from the sea cucumber Acaudina leucoprocta. Int. J. Biol. Macromol. 2020, 164, 87–94.
  3. Yao, Y.; Yim, E.K.F. Fucoidan for cardiovascular application and the factors mediating its activities. Carbohydr. Polym. 2021, 270, 118347.
  4. Lin, Z.; Tan, X.; Zhang, Y.; Li, F.; Luo, P.; Liu, H. Molecular targets and related biologic activities of fucoidan: A review. Mar. Drugs 2020, 18, 376.
  5. Chollet, L.; Saboural, P.; Chauvierre, C.; Villemin, J.-N.; Letourneur, D.; Chaubet, F. Fucoidans in Nanomedicine. Mar. Drugs 2016, 14, 145.
  6. Berteau, O.; Mulloy, B. Sulfated fucans, fresh perspectives: Structures, functions, and biological properties of sulfated fucans and an overview of enzymes active toward this class of polysaccharide. Glycobiology 2003, 13, 29R–40R.
  7. Xu, S.-Y.; Huang, X.; Cheong, K.-L. Recent advances in marine algae polysaccharides: Isolation, structure, and activities. Mar. Drugs 2017, 15, 388.
  8. Vilela-Silva, A.-C.E.; Castro, M.O.; Valente, A.-P.; Biermann, C.H.; Mourao, P.A. Sulfated fucans from the egg jellies of the closely related sea urchins Strongylocentrotus droebachiensis and Strongylocentrotus pallidus ensure species-specific fertilization. J. Biol. Chem. 2002, 277, 379–387.
  9. Zhao, X.; Guo, F.; Hu, J.; Zhang, L.; Xue, C.; Zhang, Z.; Li, B. Antithrombotic activity of oral administered low molecular weight fucoidan from Laminaria Japonica. Thromb. Res. 2016, 144, 46–52.
  10. Xie, X.-T.; Cheong, K.-L. Recent advances in marine algae oligosaccharides: Structure, analysis, and potential prebiotic activities. Crit. Rev. Food Sci. Nutr. 2022, 62, 7703–7717.
  11. Delzenne, N.M. Oligosaccharides: State of the art. Proc. Nutr. Soc. 2007, 62, 177–182.
  12. Nobre, C.; Teixeira, J.A.; Rodrigues, L.R. New trends and technological challenges in the industrial production and purification of fructo-oligosaccharides. Crit. Rev. Food Sci. Nutr. 2015, 55, 1444–1455.
  13. Vera, C.; Illanes, A.; Guerrero, C. Enzymatic production of prebiotic oligosaccharides. Curr. Opin. Food Sci. 2021, 37, 160–170.
  14. Davani-Davari, D.; Negahdaripour, M.; Karimzadeh, I.; Seifan, M.; Mohkam, M.; Masoumi, S.J.; Berenjian, A.; Ghasemi, Y. Prebiotics: Definition, types, sources, mechanisms, and clinical applications. Foods 2019, 8, 92.
  15. Zheng, L.-X.; Liu, Y.; Tang, S.; Zhang, W.; Cheong, K.-L. Preparation methods, biological activities, and potential applications of marine algae oligosaccharides: A review. Food Sci. Hum. Well. 2023, 12, 359–370.
  16. Zayed, A.; Ulber, R. Fucoidan production: Approval key challenges and opportunities. Carbohydr. Polym. 2019, 211, 289–297.
  17. Sevag, M.; Lackman, D.B.; Smolens, J. The isolation of the components of streptoeoeeal nueleoproteins in serologieally active form. J. Biol. Chem. 1938, 124, 425–436.
  18. Béress, A.; Wassermann, O.; Bruhn, T.; Béress, L.; Kraiselburd, E.N.; Gonzalez, L.V.; de Motta, G.E.; Chavez, P.I. A new procedure for the isolation of anti-HIV compounds (polysaccharides and polyphenols) from the marine alga Fucus vesiculosus. J. Nat. Prod. 1993, 56, 478–488.
  19. Zhang, R.; Zhang, X.; Tang, Y.; Mao, J. Composition, isolation, purification and biological activities of Sargassum fusiforme polysaccharides: A review. Carbohydr. Polym. 2020, 228, 115381.
  20. Cui, K.; Tai, W.; Shan, X.; Hao, J.; Li, G.; Yu, G. Structural characterization and anti-thrombotic properties of fucoidan from Nemacystus decipiens. Int. J. Biol. Macromol. 2018, 120, 1817–1822.
  21. Hwang, P.-A.; Yan, M.-D.; Kuo, K.-L.; Phan, N.N.; Lin, Y.-C. A mechanism of low molecular weight fucoidans degraded by enzymatic and acidic hydrolysis for the prevention of UVB damage. J. Appl. Phycol. 2017, 29, 521–529.
  22. Li, X.; Sun, H.; Ning, Z.; Yang, W.; Cai, Y.; Yin, R.; Zhao, J. Mild acid hydrolysis on Fucan sulfate from Stichopus herrmanni: Structures, depolymerization mechanism and anticoagulant activity. Food Chem. 2022, 395, 133559.
  23. Pomin, V.H.; Pereira, M.S.; Valente, A.-P.; Tollefsen, D.M.; Pavão, M.S.G.; Mourão, P.A.S. Selective cleavage and anticoagulant activity of a sulfated fucan: Stereospecific removal of a 2-sulfate ester from the polysaccharide by mild acid hydrolysis, preparation of oligosaccharides, and heparin cofactor II–dependent anticoagulant activity. Glycobiology 2004, 15, 369–381.
  24. Kim, S.B.; Farrag, M.; Mishra, S.K.; Misra, S.K.; Sharp, J.S.; Doerksen, R.J.; Pomin, V.H. Selective 2-desulfation of tetrasaccharide-repeating sulfated fucans during oligosaccharide production by mild acid hydrolysis. Carbohydr. Polym. 2023, 301, 120316.
  25. Silchenko, A.S.; Rubtsov, N.K.; Zueva, A.O.; Kusaykin, M.I.; Rasin, A.B.; Ermakova, S.P. Fucoidan-active α-L-fucosidases of the GH29 and GH95 families from a fucoidan degrading cluster of the marine bacterium Wenyingzhuangia fucanilytica. Arch. Biochem. Biophys. 2022, 728, 109373.
  26. Trincone, A. Update on marine carbohydrate hydrolyzing enzymes: Biotechnological applications. Molecules 2018, 23, 901.
  27. Vickers, C.; Liu, F.; Abe, K.; Salama-Alber, O.; Jenkins, M.; Springate, C.M.K.; Burke, J.E.; Withers, S.G.; Boraston, A.B. Endo-fucoidan hydrolases from glycoside hydrolase family 107 (GH107) display structural and mechanistic similarities to α-l-fucosidases from GH29. J. Biol. Chem. 2018, 293, 18296–18308.
  28. Silchenko, A.S.; Rasin, A.B.; Zueva, A.O.; Kusaykin, M.I.; Zvyagintseva, T.N.; Kalinovsky, A.I.; Kurilenko, V.V.; Ermakova, S.P. Fucoidan sulfatases from marine bacterium Wenyingzhuangia fucanilytica CZ1127T. Biomolecules 2018, 8, 98.
  29. Ohmes, J.; Mikkelsen, M.D.; Nguyen, T.T.; Tran, V.H.N.; Meier, S.; Nielsen, M.S.; Ding, M.; Seekamp, A.; Meyer, A.S.; Fuchs, S. Depolymerization of fucoidan with endo-fucoidanase changes bioactivity in processes relevant for bone regeneration. Carbohydr. Polym. 2022, 286, 119286.
  30. Chen, Q.; Kou, L.; Wang, F.; Wang, Y. Size-dependent whitening activity of enzyme-degraded fucoidan from Laminaria japonica. Carbohydr. Polym. 2019, 225, 115211.
  31. Silchenko, A.S.; Kusaykin, M.I.; Zakharenko, A.M.; Menshova, R.V.; Khanh, H.H.N.; Dmitrenok, P.S.; Isakov, V.V.; Zvyagintseva, T.N. Endo-1,4-fucoidanase from Vietnamese marine mollusk Lambis sp. which producing sulphated fucooligosaccharides. J. Mol. Catal. B: Enzym. 2014, 102, 154–160.
  32. Basso, A.; Serban, S. Industrial applications of immobilized enzymes—A review. Mol. Catal. 2019, 479, 110607.
  33. Chen, X.; Sun-Waterhouse, D.; Yao, W.; Li, X.; Zhao, M.; You, L. Free radical-mediated degradation of polysaccharides: Mechanism of free radical formation and degradation, influence factors and product properties. Food Chem. 2021, 365, 130524.
  34. Li, X.; Li, S.; Liu, J.; Lin, L.; Sun, H.; Yang, W.; Cai, Y.; Gao, N.; Zhou, L.; Qin, H.; et al. A regular fucan sulfate from Stichopus herrmanni and its peroxide depolymerization: Structure and anticoagulant activity. Carbohydr. Polym. 2021, 256, 117513.
  35. Wu, T.-C.; Hong, Y.-H.; Tsai, Y.-H.; Hsieh, S.-L.; Huang, R.-H.; Kuo, C.-H.; Huang, C.-Y. Degradation of Sargassum crassifolium fucoidan by ascorbic acid and hydrogen peroxide, and compositional, structural, and in vitro anti-lung cancer analyses of the degradation products. Mar. Drugs 2020, 18, 334.
  36. Qi, Y.; Wang, L.; You, Y.; Sun, X.; Wen, C.; Fu, Y.; Song, S. Preparation of low-molecular-weight fucoidan with anticoagulant activity by photocatalytic degradation method. Foods 2022, 11, 822.
  37. Lim, S.; Choi, J.-i.; Park, H. Antioxidant activities of fucoidan degraded by gamma irradiation and acidic hydrolysis. Radiat. Phys. Chem. 2015, 109, 23–26.
  38. Jeong, G.-W.; Choi, Y.-S. Physicochemical properties and antioxidant effects of fucoidans degraded by hydrogen peroxide under electron beam at various irradiation doses. Appl. Chem. Eng. 2022, 33, 322–327.
  39. Park, E.-J.; Choi, J.-i. Melanogenesis inhibitory effect of low molecular weight fucoidan from Undaria pinnatifida. J. Appl. Phycol. 2017, 29, 2213–2217.
  40. Fernando, I.P.S.; Dias, M.K.H.M.; Madusanka, D.M.D.; Han, E.J.; Kim, M.J.; Jeon, Y.-J.; Ahn, G. Step gradient alcohol precipitation for the purification of low molecular weight fucoidan from Sargassum siliquastrum and its UVB protective effects. Int. J. Biol. Macromol. 2020, 163, 26–35.
  41. Hwang, P.-A.; Hung, Y.-L.; Phan, N.N.; Hieu, B.-T.-N.; Chang, P.-M.; Li, K.-L.; Lin, Y.-C. The in vitro and in vivo effects of the low molecular weight fucoidan on the bone osteogenic differentiation properties. Cytotechnology 2016, 68, 1349–1359.
  42. Bi, D.; Yu, B.; Han, Q.; Lu, J.; White, W.L.; Lai, Q.; Cai, N.; Luo, W.; Gu, L.; Li, S.; et al. Immune activation of RAW264.7 macrophages by low molecular weight fucoidan extracted from New Zealand Undaria pinnatifida. J. Agric. Food. Chem. 2018, 66, 10721–10728.
  43. Cong, Q.; Chen, H.; Liao, W.; Xiao, F.; Wang, P.; Qin, Y.; Dong, Q.; Ding, K. Structural characterization and effect on anti-angiogenic activity of a fucoidan from Sargassum fusiforme. Carbohydr. Polym. 2016, 136, 899–907.
  44. Usoltseva, R.V.; Anastyuk, S.D.; Shevchenko, N.M.; Zvyagintseva, T.N.; Ermakova, S.P. The comparison of structure and anticancer activity in vitro of polysaccharides from brown algae Alaria marginata and A. angusta. Carbohydr. Polym. 2016, 153, 258–265.
  45. Silchenko, A.S.; Rasin, A.B.; Kusaykin, M.I.; Kalinovsky, A.I.; Miansong, Z.; Changheng, L.; Malyarenko, O.; Zueva, A.O.; Zvyagintseva, T.N.; Ermakova, S.P. Structure, enzymatic transformation, anticancer activity of fucoidan and sulphated fucooligosaccharides from Sargassum horneri. Carbohydr. Polym. 2017, 175, 654–660.
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