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Review

Sulfated Polysaccharides from Macroalgae—A Simple Roadmap for Chemical Characterization

by
Alice Martins
1,*,
Celso Alves
1,*,
Joana Silva
1,
Susete Pinteus
1,
Helena Gaspar
1,2 and
Rui Pedrosa
1,3
1
MARE—Marine and Environmental Sciences Centre/ARNET-Aquatic Research Network, Polytechnic of Leiria, 2520-630 Peniche, Portugal
2
BioISI-Biosystems and Integrative Sciences Institute, Faculty of Sciences, University of Lisbon, 1749-016 Lisboa, Portugal
3
MARE/ARNET/ESTM, Polytechnic of Leiria, 2520-614 Peniche, Portugal
*
Authors to whom correspondence should be addressed.
Polymers 2023, 15(2), 399; https://doi.org/10.3390/polym15020399
Submission received: 8 December 2022 / Revised: 4 January 2023 / Accepted: 9 January 2023 / Published: 12 January 2023
(This article belongs to the Special Issue Biopolymers Characterisation)
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Abstract

:
The marine environment presents itself as a treasure chest, full of a vast diversity of organisms yet to be explored. Among these organisms, macroalgae stand out as a major source of natural products due to their nature as primary producers and relevance in the sustainability of marine ecosystems. Sulfated polysaccharides (SPs) are a group of polymers biosynthesized by macroalgae, making up part of their cell wall composition. Such compounds are characterized by the presence of sulfate groups and a great structural diversity among the different classes of macroalgae, providing interesting biotechnological and therapeutical applications. However, due to the high complexity of these macromolecules, their chemical characterization is a huge challenge, driving the use of complementary physicochemical techniques to achieve an accurate structural elucidation. This review compiles the reports (2016–2021) of state-of-the-art methodologies used in the chemical characterization of macroalgae SPs aiming to provide, in a simple way, a key tool for researchers focused on the structural elucidation of these important marine macromolecules.

1. Introduction

Polysaccharides are condensate polymers of various sugars, which themselves are cyclic ethers that contain, typically, many hydroxy (–OH) substituents and, in some cases, other substituents such as amines and carboxylic acid groups. There are so many sugar monomers, and the diversity of polysaccharides is so broad, that it is not possible to write a single general structure as it is commonly done for proteins and nucleic acids.
The versatility of marine polysaccharides, e.g., their abundance, biodegradability, and biocompatibility, has been extensively investigated in the pharmaceutical and biomedical fields due to their wide range of therapeutic properties as antitumoral, anti-inflammatory, immunomodulatory, antimicrobial, and drug-release applications [1,2]. Additionally, these natural polymers are also reported for their cosmeceutical and nutraceutical potential [3], being increasingly explored by the cosmetic, food, and feed industries. Therefore, efforts focused on the elucidation of their accurate chemical structure are very important to establish a rational structure-bioactivity relationship.

2. Chemical Features of Macroalgae Sulfated Polysaccharides

As fully reported, macroalgae are known to be a good source of a variety of sulfated polysaccharides (SPs), with their bioactivities being influenced by their chemical structure [4,5,6]. However, a complete and unequivocal chemical characterization of SPs continues to be a challenge due to their structural complexity: type of polymer (homo/heteropolymer, linear/branched), molecular weight (MW), sugar composition, type of O-glycosidic linkage, sulfate pattern, and other substituents (e.g., acetate, pyruvate). These structural features strongly depend on a set of biotic and abiotic factors (Figure 1), such as macroalgae species, growth stage, harvest season, marine environment, climatic changes, geographical localization, and extraction/purification methodologies, which, taken together, also contribute to make SPs’ structural elucidation a very difficult task [7,8,9].
The extensive reviews reported in the literature [8,9,10,11,12,13,14,15,16,17,18,19] on the structural features of macroalgae SPs reveal that, despite their chemical structural variability, some similar backbones are characteristic of each seaweed phyllo. The most simple and representative structural backbones of the SPs biosynthesized by brown, red, and green macroalgae are depicted in Figure 2.
Fucoidans are the main SPs biosynthesized by brown algae. Besides fucose, the predominant sugar, other monomers such as glucose, galactose, xylose, mannose, and glucuronic acid also make up part of fucoidans’ structure. This group of SPs can be divided into two subgroups, one composed by alternating 1,3- and 1,4-linked α-l-fucopyranose residues and the other by α-1,3-l-fucopyranose, being sulfate groups linked to O-2 and/or O-3 and/or O-4 positions of fucose [4,13,16,17]. Fucoidans can be differentiated into several distinct groups according to the macroalgae species from which they are isolated, showing significant differences on their polydispersity behavior derived from a broad range of molecular weights, sugar, sulfate, and acetate contents, while enhanced bio-functional properties are achieved via structural modification of those SPs [9].
Carrageenans are the main characteristic SPs of red macroalgae and are conventionally categorized into six basic forms depending on their amount and position of sulfate groups, the number of 3,6-anhydrogalactose residues, source of extraction, and solubility, as: Kappa (κ)-, Iota (ɩ)-, Lambda (λ)-, Mu (μ)-, Nu (ν)-, and Theta (θ)-carrageenans. They are composed by alternating α-1,4-d-galactopyranose and β-1,3-d-galactopyranose (μ-, ν-, and λ-carrageenan) or by alternating β-1,3-d-galactopyranose and 3,6-anhydro-α-d-galactopyranose (κ-, ɩ-, and θ-carrageenan) [17,20]. Of these, κ, ɩ, and λ are of commercial importance due to their viscoelastic and gelling properties [10]. Due to their biocompatibility, emulsifying, thickening, gelling, and stabilizing abilities, they have several industrial applications, especially in the food, pharmaceutical, and cosmetic industries [21]. An example of a successful history is Carragelose®, an antiviral nasal spray that contains the linear SPs ɩ-carrageenan extracted from red edible seaweeds and is marketed as an over the counter (OTC) drug [22]. Due to the chemical properties of carrageenan-based hydrogels, these SPs are currently promising candidates for tissue engineering and regenerative medicine due to their similarity with native glycosaminoglycans [20].
Agar is a mixture of agarose and agaropectin consisting of d-galactose and 3,6-anhydro-α-l-galactose units joined by β-1,3- and α-1,4-glycosidic linkages. Sulfate and methoxyl groups, as well as pyruvic and d-guluronic acids, can be found in agar backbone [17]. Porphyrans and funorans, also known as agaroids, have a chemical structure very close to agars and are found in some species of red algae [16,23].
Ulvans and sulfated galactans are the main SPs found in green algae. Ulvans are water-soluble polyanionic heteropolysaccharides, with the ulvan backbone being frequently made of α- and β-(1,4)-linked monosaccharides (rhamnose, xylose, glucuronic, and iduronic acids) with characteristic repeating disaccharide units [16,17]. However, other monosaccharides are often reported in their composition, e.g., glucose, galactose, arabinose, and mannose [14]. Sulfated galactans are highly branched sulfated β-d-galactose molecules with (1,3) and (1,6) linkages, with sulfation mainly occurring at C-4 and C-6 positions [23].
Glycosaminoglycans (GAGs) are linear and heterogeneous sulfated glycans that can be found not only in green but also in red algae [13]. The skeletons of these polysaccharides are constituted by repeated building blocks of disaccharides composed of alternating uronic acid (UroA) or galactose (Gal) and hexosamine. The hexosamine may be glucosamine (GlcN) or N-acetylgalactosamine (GalNAc) and its differently substituted (mostly sulfated) derivatives. UroA can be either glucuronic acid (GlcA) or iduronic acid [13].
Some of these structural features are strictly linked with the selected extraction, depolymerization, and purification processes, which can be chosen according to the available technologies and therapeutic/industrial applications.

3. Extraction, Depolymerization, and Purification Processes

Different extraction/purification techniques employed to obtain polysaccharide-enriched products from macroalgae, and their pros and cons, were recently reviewed [6,14,17,24,25,26,27,28,29]. The chosen isolation procedure can strongly influence the molecular weight, monosaccharide composition, and sulfate content of SPs [28]. Although conventional extraction (CE) procedures (e.g., extraction with water in basic or acidic conditions at different temperatures) continue to be used, advanced extraction techniques such as subcritical water extraction (SWE), supercritical fluid extraction (SFE), microwave-assisted extraction (MAE), ultrasound-assisted extraction (UAE), pressurized liquid extraction (PLE), and enzymatic-assisted extraction (EAE) constitute efficient alternatives. Additionally, Matos et al. [29] reported the use of pulsed electric field (PEF) and ohmic heating (OH) as examples of promising and attractive electro-technologies to recover added-value compounds from macroalgae.
Since sulfated polysaccharides are complex macromolecules of high molecular weights, it is hard to achieve unequivocal structural characterization of intact polymers. Therefore, they need to be transformed into small oligomers and/or sugar monomers to facilitate further structural elucidation. Usually, the first step is the depolymerization, which can be achieved through acid (HCl, TFA, H2SO4), enzymatic (Celluclast, Viscozyme, Fucoidanase, etc.), or by high-pressure hydrolysis methods. In the following, the fractionation/purification steps of SPs’ hydrolysates can be performed with complementary methods: (i) physicochemical (precipitation, ultracentrifugation, complexation), (ii) membrane separation (dialysis, ultrafiltration), and (iii) chromatographic (ion-exchange chromatography (IEC) and size-exclusion chromatography (SEC), also referred to as gel permeation chromatography (GPC)). SPs are negatively charged molecules due to the presence of sulfate ions, and thus anion-exchange chromatography is very useful to eliminate neutral polymers, while size-exclusion chromatography allows measurements of total and molecular mass distributions. Therefore, the use of diethylaminoethyl anion-exchange (DEAE) chromatography, such as DEAE-Sepharose or DEAE-cellulose, is fully reported for SPs’ purification purposes and can be combined with SEC. More specific details regarding purification methodologies applied to polysaccharides from macroalgae and other natural sources were recently reviewed [6,30,31].

4. Chemical Characterization

The first approach aiming at the chemical characterization of macroalgae-derived SPs after extraction, fractionation, and/or purification procedures is the determination of the total content of carbohydrates, sulfates, and eventually other components, mostly proteins and phenolics, by using standard analytical methods.
The phenol-sulfuric acid method is the most used to estimate the concentration of total carbohydrates. The basic principle of the phenol-H2SO4 reaction established by Dubois et al. [32] is that carbohydrates, when dehydrated by reaction with concentrated sulfuric acid, produce furfural derivatives, which react with phenol, developing colored products [33]. d-glucose is widely used as a standard to obtain a calibration curve.
Sulfate content can be estimated by turbidimetric, colorimetric, and/or gravimetric methods. Turbidimetric methods, such as the gelatin-barium assay, quantify sulfate content on polysaccharide-enriched samples and are based on the reaction of the sulfate ion (SO42−) with the barium ion (Ba2+), originating barium sulfate (BaSO4), a water-insoluble precipitate at a low pH. The turbidity generated by the precipitate is commonly established by gelatin [34,35,36]. The quantification through colorimetric assays is preceded by the polysaccharide hydrolysis and can be accomplished by using Azure A dye, which is able to bind to sulfate groups [37]. Sodium sulfate is widely used as a standard. The method of precipitation and weighing of sulfate as BaSO4 according to AOAC [38] is a widely used gravimetric method to determine the sulfate content.
The presence of proteins on crude SPs’ fractions can be estimated by the methods developed by Bradford [39], Spector [40], and/or Lowry et al. [41], while the total phenolic content can be evaluated by the Folin-Ciocalteu method. For each determination, bovine serum albumin and gallic acid can be used as standards, respectively.
Besides the general component analysis usually performed on crude SPs (total carbohydrates, total protein, total phenolics, and total sulfate contents), more refined techniques need to be used to determine SPs’ chemical structural features. As reported by several authors [6,14,29], the elucidation of polysaccharides’ structure is a hard task due to the presence of multiple monosaccharide constituents, a variety of O-glycosidic linkages, high molecular weights, sugars’ branching, variable degrees of sulfation and substitution patterns, stereochemistry, as well as complex macromolecular properties as their aggregation modes. Effectively, to achieve a consistent structural characterization of these natural sugar polymers, it will be necessary to resort to several complementary analytical techniques to be applied to crude SPs and their derived hydrolysates. The most used techniques, and relevant information to be attained from each one, are summarized in Figure 3. Additionally, a set of chemical derivatization methods (methylation, periodate oxidation, etc.) coupled with those instrumental techniques can provide some insights into SPs’ chain structure.
Spectroscopy techniques such as Fourier transform infrared spectroscopy (FTIR), Fourier transform infrared spectroscopy-attenuated total reflection (FTIR-ATR), and Raman spectroscopy allow the detection of characteristic functional groups of SPs and can also provide some information regarding the type of glycosidic linkages. The anomeric configuration, sugar sequence, as well as the position of substituents, e.g., sulfate groups, can be determined by nuclear magnetic resonance (NMR) spectroscopy (1D and 2D experiments).
The determination of the average molecular weight (MW) and molecular weight distribution of SPs can be achieved through size-exclusion chromatography (SEC), while HPLC-SEC also offers high resolution and reproducibility and can simultaneously detect the homogeneity of polysaccharides. Refractive index (RI) and evaporative light scattering (ELSD) are the most common detectors coupled with SEC, but in some applications, multiangle laser light scattering (MALLS) is also used. The SEC-MALLS has the advantage to provide both molar mass and size independently of reference standards. Mass spectrometry techniques such as matrix-assisted laser desorption/ionization-time-of-flight mass spectrometry (MALDI-TOF-MS) and electrospray ionization tandem mass spectrometry (ESI-MS/MSn) are used to analyze macromolecules, including SPs, providing information not only about MW but also regarding monosaccharide type and substituents.
After hydrolysis, monosaccharides’ composition can be determined by gas chromatography coupled to mass spectrometry or to flame ionization detectors (GC-MS, GC-FID), high-performance liquid chromatography-refractive index detector (HPLC-RID), and high-performance anion-exchange chromatography combined with pulsed amperometric detection (HPAEC-PAD), as well as by high-performance capillary zone electrophoresis (HPCZE). GC analysis requires the conversion of sugars to volatile analogues such as alditol acetates, methyl, or trimethylsilyl derivatives, also providing information on the linkage positions and substitution patterns of constituent sugars.
Inductively coupled plasma-mass spectrometry (ICP-MS) or inductively coupled plasma-optical emission spectroscopy (ICP-OES) can be used to perform SPs’ elemental analysis. Other complementary techniques such as scanning electron microscopy (SEM), atomic force microscopy (AFM), X-ray diffraction (XRD), and circular dichroism (CD) can provide insights regarding the conformational analysis of SPs. More details about the above-outlined techniques were previously described [6,31]. Additionally, Table 1 compiles the methodologies used to attain the structural elucidation of SPs from brown, red, and green macroalgae over the last five years.
From the analysis of Table 1, it is evident that, besides the determination of total components (carbohydrates, sulfates, proteins, phenolics, glucuronic acid) and elemental (C, H, O, S) analysis, spectroscopic (FTIR, NMR) and chromatographic techniques (HPLC, GC, SEC, AEC) coupled to different detectors (MS, MALLS, RI, PAD, ELSD) are the most used techniques to attain the structural elucidation of SPs from macroalgae.
Examples of the application of several complementary techniques aiming at the structural elucidation of these marine macromolecules is evidenced by the work of Cao et al. [92] and Wahlström et al. [101]. Besides chemical modifications (acid hydrolysis, desulfation, methylation), Cao et al. [92] used HPAEC, HPGPC, FTIR, HILIC-FT-MS, GC-MS, and 1D- and 2D-NMR to perform the chemical characterization of SPs isolated from the green macroalgae Monostroma nitidum, while Wahlström et al. [101] have performed elemental analysis, FTIR, SEC, TGA, SEM, NMR, and HPAEC-PAD to characterize the SPs from Ulva spp.
A general roadmap of the main steps and techniques and/or methods currently used for extraction and chemical characterization of sulfated polysaccharides from macroalgae is summarized in Figure 4.

5. Conclusions and Further Directions

Over the last years, sulfated polysaccharides have aroused the interest of the research community due to their broad applications in biomedical, functional food, and technological areas. However, the widespread use of these macromolecules remains a challenge, mainly due to different factors that, directly and/or indirectly, affect their unequivocal chemical characterization, such as seasonality, macroalgae species, SPs’ structural and conformational variability, high molecular weights, etc., influencing their bioavailability and physicochemical behavior. Effectively, the diversity and chemical complexity of these natural polymers make their structural elucidation a hard task. Several strategies have been used to characterize SPs and it is very clear that only the integration of distinct methodologies/techniques will provide complementary information that will allow researchers to build on the puzzle of SPs’ structure. This work also evidenced the need for a set of highly costly equipment, many of them only available in a few research institutions. These constraints highlight the importance of strengthening and stimulating collaborative networks between scientists for the development of new advanced tools and strategies to reach the most accurate chemical characterization of SPs extracted from natural resources.

Author Contributions

Conceptualization, A.M., C.A., H.G., S.P. and J.S.; methodology, A.M., C.A. and H.G.; validation, A.M., C.A., H.G., S.P., J.S. and R.P.; formal analysis, C.A., A.M. and H.G.; investigation, A.M., C.A., H.G., S.P. and J.S.; resources, C.A., A.M., J.S., H.G. and R.P.; writing—original draft preparation, A.M., C.A., H.G. and S.P.; writing—review and editing, all authors; supervision, A.M., C.A., H.G. and R.P.; project administration, R.P.; funding acquisition, R.P. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Portuguese Foundation for Science and Technology (FCT) through the Strategic Projects granted to MARE—Marine and Environmental Sciences Centre (UIDP/04292/2020 and UIDB/04292/2020), Associate Laboratory ARNET (LA/P/0069/2020), and to BioISI—BioSystems and Integrative Sciences Institute (UIDP/Multi/04046/2020 and UIDB/04046/2020). FCT also funded this work through the project CROSS-ATLANTIC (PTDC/BIA-OUT/29250/2017), co-financed by the European Regional Development Fund (FEDER), through the Operational Programme for Competitiveness and Internationalization (COMPETE 2020; PO-CI-01-0145-FEDER-029250). This work was also supported by FCT and CAPES cooperation agreement through the project MArTics (FCT/DRI/CAPES 2019.00277.CBM).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data presented in this study are available on request from the corresponding author.

Acknowledgments

Authors are very grateful for the support of institutions/projects detailed in the Funding section.

Conflicts of Interest

The authors declare no conflict of interest.

Abbreviations

AASAtomic absorption spectroscopy
AECAnion-exchange chromatography
AGEAgarose gel electrophoresis
CDCircular dichroism
13C NMRCarbon-13 nuclear magnetic resonance
2D-NMRTwo-dimensional nuclear magnetic resonance spectroscopy
DEAE-CelluloseDiethylaminoethyl-Cellulose column chromatography
DEAE-SepharoseDiethylaminoethyl-Sepharose column chromatography
EDSEnergy-dispersive X-ray spectroscopy
FACEFluorophore-assisted carbohydrate electrophoresis
FTIRFourier transform infrared spectroscopy
FTIR-ATRFourier transform infrared spectroscopy-attenuated total reflectance
GC-FIDGas chromatography with flame ionization detection
GC-MSGas chromatography with mass spectrometry detection
GPCGel permeation chromatography
1H NMRProton nuclear magnetic resonance
HILIC-FT-MSHydrophilic interaction liquid chromatography-Fourier transform-mass spectrometry
HPAECHigh-performance anion-exchange chromatography
HPAEC-PADHigh-performance anion-exchange chromatography with pulsed amperometric detection
HPGPCHigh-performance gel-permeation chromatography
HPLC-ELSDHigh-performance liquid chromatography with evaporative light scattering detector
HPLC-RIDHigh-performance liquid chromatography with refractive index detection
HPSECHigh-performance size-exclusion chromatography
HPSEC-ELSDHigh-performance size-exclusion chromatography with evaporative light scattering detector
HPSEC-MALLSHigh-performance size-exclusion chromatography coupled with multi-angle laser light scattering
HPSEC-MALS-RIHigh-performance size-exclusion chromatography-multi-angle light scattering and refractive index detection
HPSEC-UV-MALLS-RIHigh-performance size-exclusion liquid chromatography with ultraviolet-multi-angle laser light scattering-refractive index detection
HPTLCHigh-performance thin-layer chromatography
ICP-MSInductively coupled plasma-mass spectrometry
ICP-OESInductively coupled plasma-optical emission spectrometry
IECIon-exchange chromatography
LC-ESI–MS/MSLiquid chromatography-electrospray ionization-tandem mass spectrometry
MALDI-TOF-MSMatrix-assisted laser desorption/ionization-time-of-flight mass spectrometry
MALLSMulti-angle laser light scattering detection
RP-HPLCReversed phase-high-performance liquid chromatography
SDS-PAGESodium dodecyl sulfate-polyacrylamide gel electrophoresis
SEC-MALLSSize-exclusion chromatography-multi-angle laser light scattering
SEMScanning electron microscopy
SEM-EDXScanning electron microscope-energy-dispersive X-ray analysis
SLS/DLSStatic and dynamic light scattering
TGAThermogravimetric analysis
TLCThin-layer chromatography
UV-VisUltraviolet-visible spectroscopy
XRDX-ray diffraction

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Polymers 15 00399 g001 550
Figure 1. Features related to macroalgae sulfated polysaccharides’ complexity.
Figure 1. Features related to macroalgae sulfated polysaccharides’ complexity.
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Polymers 15 00399 g002 550
Figure 2. Characteristic backbones of macroalgae sulfated polysaccharides.
Figure 2. Characteristic backbones of macroalgae sulfated polysaccharides.
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Polymers 15 00399 g003 550
Figure 3. Current techniques for macroalgae sulfated polysaccharides’ structural characterization.
Figure 3. Current techniques for macroalgae sulfated polysaccharides’ structural characterization.
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Figure 4. Roadmap of techniques/approaches for the chemical characterization of sulfated polysaccharides.
Figure 4. Roadmap of techniques/approaches for the chemical characterization of sulfated polysaccharides.
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Table
Table 1. Strategies for chemical characterization of sulfated polysaccharides isolated from macroalgae adopted in the last five years (2016–2021).
Table 1. Strategies for chemical characterization of sulfated polysaccharides isolated from macroalgae adopted in the last five years (2016–2021).
AlgaeSourceCompoundChemical CharacterizationReference
Ochrophyta (Brown Algae)
Chnoospora minimaSouthern coastal area of Sri LankaFucoidanComponent analysis
DEAE-Sepharose chromatography
FTIR
AGE
HPAE-PAD
NMR		[42]
Cladosiphon okamuranusIshigaki Island
(Okinawa, Japan)		FucoidanComponent analysis
IEC
GC-FID
GC-MS
Chemical modifications		[4]
Dictyota bartayesiana
Turbinaria decurrens		Mandapam Coastal region, Rameswaram, Tamil Nadu, IndiaFucoidanComponent analysis
FTIR
RP-HPLC
DEAE-Cellulose chromatography
Chemical modifications		[43]
Ecklonia maximaHIK-Abalone Farm, Hermanus, South AfricaFucoidanComponent analysis
Ultracentrifugation
FTIR
NMR
XRD		[44]
Fucus evanescens-FucoidanComponent analysis
SEC
IEC
NMR		[45]
Himanthalia elongataSpanish Atlantic
coasts (local supplier Porto-Muiños, A Coruña, Spain)		FucoidansFTIR
HPSEC
GC-FID		[46]
Hizikia fusiforme-FucoidanComponent analysis
HPGPC
HPAEC-PAD
FTIR
NMR		[47]
Kjellmaniella crassifoliaCoast of Dalian, ChinaFucoidansComponent analysis
HPLC
DEAE-Sepharose chromatography
SEC
FTIR
1D and 2D NMR 		[48]
Laminaria hyperboreaNortheast Atlantic Ocean, ScandinaviaSulfated fucansAEC
Raman spectroscopy
ICP-MS
HPSEC-MALLS		[49]
Laminaria japonicaPutian, Fujian, ChinaFucoidansDEAE-Cellulose chromatography
Ultrafiltration
Chemical modifications
GC		[50]
Laminaria japonicaCrude commercial fucoidan
(Rizhao Jiejing Ocean Biotechnology Development Co., Ltd., Rizhao, China)		FucoidanElemental analysis
Component analysis
Chemical modifications
1D and 2D NMR
GC-FID
GC-MS
SLS/DLS measurements
FTIR
AFM		[51]
Lessonia sp.Tekenika Bay, Southern ChileSulfated fucanFTIR
NMR
GC-FID
DEAE
HPLC		[52]
Nizamuddinia zanardiniiRocky beaches of Chabahr at Oman Sea, South of IranFucoidanComponent analysis
FTIR
GCMS
HPSEC-UV-MALLS-RI
SEM		[53]
Padina commersoniiCoast of Galle, Sri LankaFucoidanAEC
FTIR
NMR		[54]
Padina tetrastromaticaVizhinjam coast of Kerala, IndiaSPs1H NMR
DEAE-Cellulose chromatography		[55]
Padina tetrastromaticaVizhinjam
coast of Kerala, India		SPsElemental analysis
UV-Vis
GPC		[56]
Padina tetrastromaticaCoastal rocks of Mulloor, Vizhinjam, Thiruvananthapuram, Kerala, IndiaSulfated fucanHPTLC
LC-ESI-MS
DEAE-Cellulose chromatography		[57]
Saccharina japonicaGuemil-eup, Wando-gun, and Jeollanam-do, Republic of KoreaFucoidansComponent analysis
Elemental analysis
FTIR
UV-Vis
XRD
TGA
TLC
HPSEC-ELSD
HPLC-ELSD		[58]
Saccharina japonicaXiapu, Fujian province, ChinaFucoidanComponent analysis
HPSEC-MALLS-RID
FTIR
NMR
GC		[59]
Saccharina japonica
Sargassum fusiforme Sargassum hemiphyllum
Undaria pinnatifida		VariousSPsAEC
HPSEC-MALLS-Visc-RID
FTIR
HPLC		[60]
Sargassum binderiHikkaduwa southern coast of Sri Lanka-IEC
FTIR
NMR		[61]
Sargassum duplicatumNhatrang Bay (Socialist Republic of Vietnam)FucoidanComponent analysis
Chemical modifications
ESI-MS/MS
MALDI-TOF
NMR
HPSEC
DEAE-Cellulose chromatography
AGE		[62]
Sargassum duplicatum
Sargassum feldmannii		Nhatrang bay (Socialist Republic of Vietnam)FucoidanESI-MS/MS
NMR
HPSEC
Chemical modifications 		[63]
Sargassum horneri-Fucoidan and
sulfated fucooligosaccharides		NMR
IEC
PAGE		[64]
Sargassum muticumBuarcos Bay (Figueira da Foz, Portugal)FucoidansComponent analysis
ICP-OES
HPLC-UV
FTIR-ATR
1H NMR		[65]
Sargassum pallidumWeihai, Yellow Sea, ChinaFucoidansComponent analysis
HPGPC-FTIR
GC-FID		[66]
Sargassum swartziiCoast of Kanyakumari, IndiaSPsFTIR
NMR
UV-Vis
TLC
HPSEC
TGA		[67]
Sargassum wightiiTamil Nadu, India-Elemental analysis
Component analysis
FTIR
TGA		[68]
Turbinaria conoidesCoast of Mandapam, Rameswaram, Gulf of Mannar, Tamil Nadu, IndiaFucoidanGPC
HPLC
NMR
GC-MS
DEAE-Cellulose chromatography
Component analysis		[69]
Turbinaria ornataNhatrang Bay
(Socialist Republic of Vietnam)		FucoidanDEAE
ESI-MS/MS
GC-MS
NMR		[70]
Turbinaria turbinataMalaysian originSPsGC-FID
FTIR
HPSEC-MALS-RI
DEAE-Cellulose chromatography
NMR
TGA		[71]
Undaria pinnatifidaAuckland, New ZealandFucoidanComponent analysis
FTIR
2D-NMR
HPLC-RID		[72]
Rhodophyta (Red Algae)
Chondrus canaliculatusTunisian coasts, Sfax (“Sidi Mansour, Tabaroura”)Fractions of SPsComponent analysis
HPGPC
FTIR-ATR
HPLC-RID
Solid-state 13C NMR 		[73]
Gelidiella acerosaAtlantic coast, Brazil (Búzios Beach, Nísia Floresta—Rio Grande do Norte)SPsElemental analysis
Component analysis
FTIR
NMR
HPSEC		[74]
Gelidium crinaleNaozhou Island Sea, Zhanjiang City, Guangdong ProvinceSPsChemical modifications
Component analysis
FTIR
HPLC-UV
GPC		[75]
Gigartina pistillataCollected at Spanish Atlantic
coasts and obtained from a local supplier (Porto-Muiños, A Coruña, Spain)		CarrageenansComponent analysis
FTIR
HPSEC
GC-FID		[46]
Gracilaria caudataBrazilian Atlantic coast (Fleixeiras Beach, Trairí—Ceará)SPComponent analysis
GPC
ICP-OES
FTIR
NMR 		[76]
Gracilaria caudataNortheast Atlantic coast of Brazil (Fleixeira Beach, Trairi—CE,
Brazil)		SPsFTIR
NMR		[77]
Gracilaria gracilisWild Coast Abalone, East London, South AfricaSPsComponent analysis
SEM-EDX
FTIR
GC-MS		[78]
Gracilaria gracilisDakhala shoreline,
Morocco		AgarsFTIR
NMR		[79]
Gracilaria lemaneiformisNan’ao Island of ChinaSPsDEAE-Sephadex chromatography
HPLC-ELSD
FTIR
GC-FID
GC-MS		[80]
Laurencia obtusaCoastal region of Bizerte (Tunisia) in the Mediterranean SeaComplex SPsComponent analysis
DEAE-Sephadex chromatography
SEC-MALLS
FTIR
NMR 		[81]
Laurencia papillosaEast-Mediterranean coastal waters of Lattakia, SyriaCarrageenansComponent analysis
FTIR-ATR
NMR
GPC		[82]
Osmundea pinnatifidaBuarcos bay (Figueira da Foz, Portugal)AgaransComponent analysis
ICP-OES
HPLC
FTIR-ATR
NMR		[65]
Porphyra aitanensisPurchased from Pingtan Island, Fujian Province, ChinaSPsHPLC-SEC-MALLS-RI
UV
EDS
XRD		[83]
Solieria filiformisNortheast Atlantic coast of Brazil (Flexeiras Beach, Trairi—Ceará)SPsHPSEC
FTIR
NMR
Component analysis 		[84]
Chlorophyta (Green Algae)
Caulerpa cupressoides
var. flabellata		Nísia Floresta,
southern coast of Rio Grande do Norte, Brazil.		Sulfated galactansComponent analysis
GPC
NMR
IEC		[85]
Caulerpa lentilliferaCultivated, Dalian, Liaoning, ChinaSPsChemical modifications
NMR
GC-MS		[86]
Caulerpa lentilliferaTakalar, South of Sulawesi, IndonesiaSPsFTIR
HPLC
NMR		[87]
Caulerpa sertularioidesCoast of Rio Grande do Norte, BrazilSPsComponent analysis
HPLC-RID
GPC		[88]
Chaetomorpha gracilisIMTA system Cinvestav Marine Station, TelchacSPsComponent analysis
FTIR
NMR
XRD
TGA		[89]
Codium isthmocladumPirambuzios beach, Nisia Floresta, Rio Grande do Norte, BrazilSulfated homogalactansAEC
AGE
GPC
GC-MS
NMR		[90]
Gayralia brasiliensisBaía de Paranaguá, Paraná State, BrasilSulfated heterorhamnanComponent analysis
HPSEC-MALLS-RI
NMR		[91]
Monostroma nitidumCoast of Yantai, ChinaSPsComponent analysis
NMR
GC-MS
AEC
HILIC-FT-MS
FTIR
HPGPC		[92]
Monostroma nitidumYellow Sea of ChinaSulfated glucuronorhamnanComponent analysis
FTIR
NMR
HILIC-FT-MS
HPGPC
RP-HPLC		[93]
Ulva lactucaTaboulba and Sayada (Monastir—Tunisia)UlvansComponent analysis
GC-FID
HPSEC
FTIR
NMR		[94]
Ulva lactucaMediterranean Sea in Egypt (Alexandria in Abou Kir region)SPsComponent analysis
AEC
FTIR
HPLC-RID		[95]
Ulva lactucaWild Coast Abalone, East London, South AfricaSPsComponent analysis
SEM-EDX
FTIR
GC-MS		[78]
Ulva lactucaHo-Ping Island, Keelung, TaiwanUlvansComponent analysis
FTIR
HPSEC		[96]
Ulva lactuca L.Seashore of Nísia
Floresta, RN, Brazil		SPsComponent analysis
AGE
FACE
FT-Raman spectroscopy		[97]
Ulva linzaLebanese Mediterranean coastUlvansElemental analysis
Component analysis
SEC
HPLC
FTIR
NMR		[98]
Ulva pertusaChinaSPsDEAE-Cellulose chromatography
HPGPC
GC
FTIR
AAS		[99]
Ulva sp.Landrézac Beach, Sarzeau, Brittany, FranceUlvansComponent analysis
HPSEC
HPAEC
MALDI-TOF		[100]
Ulva spp.Swedish West coastUlvansElemental analysis
FTIR
SEC
TGA
SEM
NMR
HPAEC-PAD		[101]
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MDPI and ACS Style

Martins, A.; Alves, C.; Silva, J.; Pinteus, S.; Gaspar, H.; Pedrosa, R. Sulfated Polysaccharides from Macroalgae—A Simple Roadmap for Chemical Characterization. Polymers 2023, 15, 399. https://doi.org/10.3390/polym15020399

AMA Style

Martins A, Alves C, Silva J, Pinteus S, Gaspar H, Pedrosa R. Sulfated Polysaccharides from Macroalgae—A Simple Roadmap for Chemical Characterization. Polymers. 2023; 15(2):399. https://doi.org/10.3390/polym15020399

Chicago/Turabian Style

Martins, Alice, Celso Alves, Joana Silva, Susete Pinteus, Helena Gaspar, and Rui Pedrosa. 2023. "Sulfated Polysaccharides from Macroalgae—A Simple Roadmap for Chemical Characterization" Polymers 15, no. 2: 399. https://doi.org/10.3390/polym15020399

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