Animal-Derived Polysaccharides

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1		Tao Wu	--	3625	2024-01-10 13:34:48	\|
2	update references and layout	Rita Xu	Meta information modification	3625	2024-01-11 02:52:02	\|

This entry is adapted from the peer-reviewed paper 10.3390/foods13010173

Polysaccharides are biomolecules found in microorganisms, plants, and animals that constitute living organisms. Glycosaminoglycans, unique acidic polysaccharides in animal connective tissue, are often combined with proteins in the form of covalent bonds due to their potent biological activity, low toxicity, and minimal side effects, which have the potential to be utilized as nutrition healthcare and dietary supplements.

animal polysaccharides glycosaminoglycan extraction method

1. Introduction

Polysaccharides are carbohydrates formed via the dehydration and condensation of multiple monosaccharides linked together by glycosidic bonds ^[1]. The molecular weights can reach tens of thousands or even millions, and they are considered one of the fundamental substances that constitute life activities and maintain biological functions besides proteins and nucleic acids ^[2]. They exhibit diverse branching compositions, molecular weights, and conformations ^[3]. Glycosaminoglycans (GAGs), also known as mucopolysaccharides, are linear polyanionic compounds that belong to the family of animal polysaccharides, which are polymerized from hexuronic acid (except keratan sulfate) and hexosamine repeating units ^[4]. Proteoglycans (PGs) are glycosylated proteins whose structure consists of one or more GAGs covalently linked to a core protein ^[5]. Serving as indispensable constituents of PGs, GAGs are regally divided into five kinds: hyaluronic acid (HA), chondroitin sulfate (CS), dermatan sulfate (DS), keratan sulfate (KS), and heparan sulfate (HS) along with heparin (Hep) ^[6]. Cell growth and development, intercellular homeostasis, and repair and regeneration are all linked to GAGs in the extracellular matrix ^[7]. GAGs are predominantly sourced from aquatic organisms, including marine fish, shellfish, and mollusks ^[8]. These creatures live in hydrated buffer systems with small temperature differences and rich ionic species, and the polysaccharides synthesized in vivo have distinctive structural groups and physicochemical properties ^[9], which are found in a variety of aspects, such as drug carriers, cosmetics, and healthcare foods. Relationships between the conformation of polysaccharides and their activities have been reported. To elucidate these relationships, appropriate preparation and analytical techniques are required. A single method is not sufficient to characterize the structure of GAGs. Fourier-transform infrared (FTIR), nuclear magnetic resonance (NMR), high-performance liquid chromatography (HPLC), and other methods are usually used to orthogonally integrate the obtained data. The objective is to conduct a more in-depth analysis of animal polysaccharides.

2. Extraction Methods of Animal Polysaccharides

Animal polysaccharide extraction methods mainly include water extraction, acid–alkaline extraction, enzyme extraction, ultrasonic extraction, and combined extraction. Extraction serves as a vital step towards obtaining crude polysaccharides. The glycosidic linkage connection, monosaccharide type, and disaccharide composition of polysaccharides can be influenced by factors such as pH, temperature, and solvent used, which may interfere with subsequent structural analysis.

2.1. Water Extraction Method

Traditional water extraction is the most commonly used physical extraction method, which has the advantages of simple operation, ease of control, and low cost. The principle is that polysaccharides are extracted by submerging raw materials in boiling water for a period of time. This method utilizes the solubility of high-temperature water to accelerate the release of polysaccharides. According to polysaccharides, which are insoluble in organic solvents, ethanol-graded precipitation obtains crude polysaccharides. However, water molecules cannot fully destroy the cell membrane, resulting in the incomplete extraction and loss of polysaccharides and a low yield ^[10].

Zheng et al. compared four methods for extracting polysaccharides from lumpus cartilage using a fixed material–liquid ratio of 1:40. They found that the water extraction method heated at 80 °C for 4 h had the lowest extraction rate, at approximately 3.52% ^[11]. Luan extracted the crude polysaccharide of Solenidae using a boiling water bath. The material–liquid ratio was 1:30 for 2 h, and then the residue was boiled again with a material–liquid ratio of 1:20 for 1 h. The extraction rate of crude polysaccharide was 1.7% lower than that of the alkaline extraction method, but it maintained a higher total sugar content at 55.4% ^[12]. That is because the solvent was only water, and water-soluble polysaccharides were extracted through intermolecular forces to retain the original structure of polysaccharides as much as possible. Ticar et al. used 80 °C hot water to extract heads of silver-banded whiting twice, each time lasting for 1 h. However, the yield of GAGs only reached 0.8% ^[13]. Getachew et al. modified the hot water extraction method using subcritical water (SWE) at approximately 125 °C to extract polysaccharides from Pacific oysters for 15 min. The process was optimized, resulting in a significantly higher yield of approximately 18.42% ^[14]. Mohammadi et al. investigated the efficiency of extracting polysaccharides from abalone in a high-pressure autoclave extractor with subcritical water at different temperatures, and the polysaccharide content was found to be high in the range of 250–280 °C, with a peak at 250 °C ^[15].

2.2. Acid and Alkaline Extraction Method

Compared to water extraction, alkaline extraction more effectively breaks down glycopeptide chains between PGs and forms salts with acidic polysaccharides so that water-soluble polysaccharides can be easily released. Chen et al. discovered that the yield of polysaccharides from Andrias davidianus skin mucus increased by 2.45% when extracted using a 0.4 mol/L NaOH solution for 2 h at 45 °C with a material–liquid ratio of 1:25, compared to extraction using a 0.3 mol/L HCL solution under the same conditions ^[16]. On the other hand, Lu et al. obtained the opposite conclusion that by increasing the temperature to 50 °C and extending the extraction time to 3 h, the yield of polysaccharides in the coelomic fluid of Phasolosma esculenta extracted using NaOH with a mass concentration of 1.5% was only 0.92% ^[17].

The alkaline extraction method can enhance the yield of polysaccharides within a certain concentration range. However, the disadvantage is that some aminopolysaccharides may be degraded by strong alkaline, resulting in the Walden inversion and desulfurization ^[18], affecting polysaccharides’ structure analysis and activity verification. Therefore, low-concentration alkaline can be used for extraction.

Similar to the alkaline extraction method, the use of the acid extraction method can help break the bonds between polysaccharides and bound proteins, thereby improving the yield and purity of polysaccharides. Cheng et al. first used water to extract polysaccharides from Mytilus edulis. They then extracted the residual precipitate in a 0.2 mol/L HCL solution at 60 °C for 30 min and added 10% Na₂CO₃ for a further alkaline extraction of the residues. The acid extraction yield was the lowest at 3.5%, and the antioxidant capacity of the extracted polysaccharides was also the weakest ^[19]. It can be concluded that high concentrations of acid or base can cleave glycosidic bonds, irreversibly affecting the spatial structure of polysaccharides and reducing their biological activity. Therefore, neither laboratory nor industrial preparations are used very rarely.

2.3. Enzyme Extraction Method

Under mild conditions of enzyme action, each enzyme binds specifically to its substrate, allowing the reaction to be specific, which can selectively hydrolyze or degrade the cell membranes, reducing intermolecular resistance and allowing polysaccharides to be effectively released ^[20]. Enzymes function optimally at a specific pH and temperature, breaking down polysaccharides into small molecule fragments, thus improving the yield. Enzymes utilized in the preparation include alkaline protease, papain, neutral protease, etc.

Papain is the most commonly used enzyme for the extraction of animal polysaccharides. Maccari et al. selected papain for CS extraction from four species of bony fish by stirring overnight at 60 °C in a Na–acetate buffer solution at pH 5.5, and the highest yield among the four CS was only 0.34% ^[21], which is contrary to the findings of Zhang’s study ^[20]. The reason for this discrepancy may be because no screening of the optimum proteases was carried out prior to the extraction of polysaccharides. Bai et al. employed a trypsin–papain in a 0.1 mol/L Tris-HCL buffer at 55 °C to co-enzymatically extract the polysaccharides from the Lapemis curtus skin for 3 h with a crude extraction rate of 200 mg/g ^[22]. Wang also used both enzymes to extract GAGs from the Acipenser schrenckii cartilage, with a yield of 25% ^[23]. Wang et al. extracted CS from six marine animals using trypsin digestion at 37 °C for 4 h, followed by papain digestion at 60 °C for 4 h. The yield of CS was only 16.36% for Raja porosa, while the yield from the other five animals was less than 1%. This difference in yield may be due to the limitation that only Raja porosa is suitable for extraction using this method, and more suitable conditions need to be explored for the extraction of other animal polysaccharides ^[24]. Yuan et al. compared the extraction rate of Sinonovacula constricta polysaccharide (SCP) using a neutral protease extracted at 50 °C for 173 min and water extraction at 80 °C for 4 h. The enzyme extraction method resulted in a 12.26% higher yield than the water extraction method, but there was no significant difference in composition ^[25]. These findings suggest that enzyme extraction is a green and efficient method that can improve extraction rates while ensuring the integrity of the polysaccharide structure. Guo et al. extracted abalone viscera polysaccharide using an alkaline protease for enzymolysis at pH 9.5 for 19 h. They then selected a flavor enzyme at pH 7.0 from five enzymes for secondary enzymolysis. The extract had a high polysaccharide content of 51.75% and significantly reduced protein content ^[26]. The reason for this is the high efficiency of protein hydrolysis via alkaline protease and the fact that different binding sites of the flavor enzyme and alkaline enzyme facilitate further hydrolysis of polysaccharides and the removal of protein complexes.

Single enzyme extraction, complex enzyme extraction, and multiple enzyme segmented extraction methods are currently the most suitable for extracting animal polysaccharides.

2.4. Combined Extraction Method

In addition to the previously mentioned methods, several sustainable technologies have been steadily implemented, including microwave extraction, ultrasonic extraction, and supercritical fluid extraction. Each method possesses unique characteristics, depending on its specific mechanisms. Various methods were employed to obtain the most efficient extraction of animal polysaccharides, where enzymolysis, in conjunction with other methods, was mainly utilized. This method shortened the extraction time and increased the yield simultaneously.

Wang et al. compared the effects of enzyme, ultrasound, and ultrasound-assisted enzyme methods on the yield of polysaccharides from sea cucumber gonads. The results showed that the polysaccharide extracted via pH 7.5, 50 °C enzymolysis for 4 h followed by 400 Watt ultrasonic for 50 min had a higher sulfate acid group content, and the yield was better than the other two methods, about 6.09% ^[27]. Guo et al. conducted a study on the extraction of polysaccharides from various parts of Scophthalmus maximus using two methods: first, alkaline followed by enzyme, and second, enzyme followed by alkaline ^[28]. The results show that enzymes are used to hydrolyze the peptide bond first, which degrades the protein into small peptides and thus separates it from other substances. After that, the β-elimination reaction occurred under alkaline conditions, and the -O-glycopeptide chain was cleaved, which further released the polysaccharide from PG. Chen et al. optimized the specific parameters of the ultrasound-assisted enzymatic extraction of polysaccharides from thick-shelled mussels: acid protease was added to the solution, the extraction was performed at 64 °C for 36 min, and the ultrasonic power was 60 w ^[29]. The molecular weight of the polysaccharides obtained using this method was 1/10 of that obtained using hot water extraction, suggesting that ultrasonic treatment led to the degradation of polysaccharides. Although there is a decrease in molecular weight, no difference in monosaccharide composition was observed. Li et al. used a dual-phase saline extraction method papain, enzymatic extraction of polysaccharides from abalone guts while removing pigments and some heavy metal ions to shorten the time of purification and simplify experimental steps. The optimal extraction conditions were 30% ethanol and 12% Na₂CO₃. Under this condition, the yield of abalone visceral polysaccharides was 3.76% ^[30].

3. Structural Characterization of Animal Polysaccharides

The research and development of polysaccharides in terms of function, therapeutic properties, and toxicities are synchronized and continuous ^[31]. Without mastering the structure of isolated and purified polysaccharides, even if they have strong biological activity and safety, it is impossible to carry out pharmacological and toxicological studies, not to mention synthetic or structural modification, which hinders the development of new, high-quality drugs. Polysaccharides exhibit distinctive chain conformations, with structures ranging from spherical to helical, coiled, and rodlike shapes. Moreover, they contain (1→3), (1→2), (1→4), and (1→6) linkages and diverse α and β conformations ^[1]. The primary structures serve as the foundation for advanced structures. Currently, the identification of polysaccharide structures involves the relative molecular mass, the monosaccharide composition, the ratio of the number of substances, sugar ring conformations, anomeric carbon configurations, linkages between sugar residues, etc. Primary structures of polysaccharides are often analyzed using high-performance liquid chromatography (HPLC) ^[32], gas chromatography (GC), gas chromatography–mass spectrometry (GC-MS), Fourier-transform infrared spectroscopy (FTIR) ^[33], an atomic force microscope (AFM) ^[34], nuclear magnetic resonance spectroscopy (NMR) ^[35], and other methodologies.

3.1. Monosaccharide Composition

Oligosaccharides are highly water-soluble and more likely to enter the organism through multilayer membrane barriers to carry out their function. Acid hydrolysis, acetolysis, methanolysis, and other methods can degrade polysaccharides and explain their primary structure and active characteristics at a low molecular level ^[36]. HPLC is most suitable for glycan identification, especially for detecting the molecular weights of several thermosensitive glycans and polysaccharides. This technology is highly sensitive, requires only a small sample, has excellent reproducibility, can qualify components based on the retention time of spectra, and can quantify the peak area.

Wang isolated polysaccharides from various sea gonads and discovered that the monosaccharides consisted mainly of mannose (Man), glucosamine (GlcN), and glucose (Glc), with small amounts of rhamnose (Rha), glucuronic acid (GlcA), galactosamine (GalN), galactose (Gal), and fucose (Fuc) together ^[37]. Bai conducted a compositional analysis of CS in the sturgeon bone. The analysis revealed that the major constituents were Glu and GalN, with a proportion of 1.04:1 comprising 48.23% and 46.29% of the sample, respectively. A peak different from the monosaccharide standard was also captured, presumably a disaccharide produced via antacidolysis ^[38]. Yim et al. analyzed the monosaccharide composition of polysaccharides extracted from abalone and observed that the crude product had a higher fucose than other monosaccharides, which may be responsible for its antiviral activity ^[39].

3.2. Functional Groups and Chemical Bonds

After determining the compositions and molecular weights of the monosaccharides, FTIR was employed to detect the functional groups obtained in the polysaccharides. FTIR is applied for the qualitative analysis of unidentified substances, and spectra are based on the vibration or rotational excursion of atoms comprising chemical bonds or functional groups. According to the origin of the absorption peaks, they can be divided into the functional group region (2.5–7.7 μm, i.e., 4000–1330 cm⁻¹) and the fingerprint region (7.7–16.7 μm, i.e., 1330–400 cm⁻¹). The absorption peaks within the functional region arise from stretching vibrations of groups and are few, which are used to identify functional groups. The situation is different in the fingerprint region, where the peaks are numerous and complex, not typical, and are mainly generated via the stretching vibrations of some single bonds C-O, C-N, and C-X (halogen atoms) and bending vibrations of hydrogen-containing groups like C-H and O-H, as well as via vibrations of the C-C skeleton. When there are slight variations in the molecular structures, absorption peaks in this area will vary.

Gao et al. discovered that the purified polysaccharides from snail mucilage were pyranose, exhibiting distinct absorption peaks between 810 and 780 cm⁻¹. The absorption peak at 1426 cm⁻¹ is due to the stretching vibration of the C-O bond on the residue of iduronic acid, confirming the presence of -COO-. Furthermore, the presence of -COO- and CH₃CO- is evidenced by a strong characteristic absorption peak of the C=O bond at 1640 cm⁻¹. Taken together, these groups indicate that the polysaccharide is a typical GAG ^[40]. Similarly, Wang et al. also identified the snail mucilage polysaccharide as a β-pyranose by finding a characteristic β-glycosidic bond absorption peak at 891 cm⁻¹ ^[41]. Krichen et al. analyzed two purified fish skin sulfated polysaccharides, SHSP and GTSP, using IR, and the characteristic absorptions at 1382 cm⁻¹ and 1394 cm⁻¹ confirmed the presence of a C-O-S bond in the sulfate group. A pyranose unit is suggested via absorptions at 1152 and 1034 cm⁻¹ (SHSP) and 1156 and 1106 cm⁻¹ (GTSP) ^[42]. Song et al. conducted an analysis of the IR of polysaccharides from Patinopecten yessoensis viscera and attributed the broad peak at 3428.4 cm⁻¹ to the stretching vibration of the -OH bond, while the multiple peaks at 1100–1000 cm⁻¹ belong to the stretching vibration of the C=O and C-C bonds in the pyranoid ring, as well as the stretching vibration of the glycosidic bond C-O-C. The peak near 1247.6 cm⁻¹ is caused by the stretching vibration of the S=O bond of the sulfate group and is typical of sulfate polysaccharides ^[43].

3.3. Disaccharide Composition

NMR is used as an analytical chemistry technology to study the molecular structures and conformation of polysaccharides. Its principle involves identifying groups based on their chemical shifts, determining linkage and sequence of residues through numbers of coupled splitting peaks and coupling constants ^[44], and analyzing proton ratios of each group through H-peak integral areas. The study of polysaccharides is generally centered around Proton nuclear magnetic resonance (¹H NMR), Carbon-13 nuclear magnetic resonance (¹³C NMR), correlation spectroscopy (COSY) in a heteronuclear single quantum coherence (HSQC), as well as a heteronuclear multiple-bond correlation (HMBC).

Zuo analyzed the disaccharide composition of CS extracted from tilapia processing by-products. The findings indicate that CS-C are present in both fish heads and tails, with the CS in tails containing ∆Di-2S,6S, while disulfide disaccharides are absent from the CS in heads. The composition of CS in fish spines and fins was similar, but CS in spines does not contain ΔDi-2S,6S, and CS in fins does not contain ΔDi-4S,6S. Most of the absorption peaks of ¹H NMR appeared in three regions. The first region showed a characteristic peak of CS, while the middle region mainly consisted of sugar ring proton signals. The anomeric proton signal at 5.3 ppm in the last region was assigned to the C=O bond, thus confirming the existence of N-acetylglucosamine ^[45]. Signals elsewhere in the region were consistent with the analysis of disaccharide.¹³C NMR makes up for the shortcomings of ¹H NMR. It can directly determine the molecular skeleton, giving information on various carbon-containing functional groups. The oyster polysaccharides extracted using subcritical water by Getachew et al. were analyzed as follows: the absorption peak at 1150 cm⁻¹ was due to α-(1→4) glycosidic bonding. The stretching of the C-O bond appeared at 1020.16 cm⁻¹, while the 1077.04 cm⁻¹ peak was attributable to the deformation vibration of the -OH group. The presence of a characteristic peak at 848.52 cm⁻¹ indicated the presence of oyster polysaccharides in an α-configuration. In ¹³C NMR, there were no chemical shifts above 100 ppm, indicating the absence of acetyl and carboxyl groups. The absence of a furan ring C-1 resonance band within the 107–109 ppm range confirmed that the polysaccharide was pyranose. Additionally, the peaks detected in the 76.96–69.27 ppm and 60.50 ppm regions corresponded to an α-configuration of dextran’s C2-C5 and C6, respectively. Thus, based on the NMR spectrum, the polysaccharide sample could be confirmed to be in an α-configuration pyranose, corresponding to the FTIR analysis results ^[14]. Additionally, Seedevi et al. analyzed 2D NMR COSY spectra based on 1D NMR of GAG extracted from cuttlefish. The cross peaks between 4.50 and 4.75 ppm indicated the presence of unsubstituted amines H-1 and H-2 that contained N-acetylglucosamine residues within sulfate GAG, proving the presence of glucuronic acid. Moreover, the signals at 3.80, 3.87, and 3.29 ppm assigned to H-4, H-3, and H-2, respectively, identified the position of glucuronic acid residues in sulfate GAG ^[46].

3.4. Chain Conformation

SEM uses secondary electron signal imaging to observe the surface morphologies of samples, which can also be used for the qualitative and semi-quantitative analysis of components, with the advantages of a large imaging range and fast scanning speed. In contrast, the AFM avoids any special handling and preserves the physiological state and integrity of polysaccharide molecules as much as possible ^[27]. Its imaging technology reveals 3D information on molecular morphologies, sizes, and conformations of samples in a solution. Furthermore, it exploits the intermolecular force between atoms to emphasize sample properties, directly reflecting the sensitivity of conformation to the microenvironment.

Mou et al. extracted two sea cucumber polysaccharides, fCS-Hm and fCS-Aj, using an alkaline extraction–enzymolysis method. With a similar molecular weight but relatively low sulfate content, FCS-Hm demonstrated greater extended chain conformations than fCS-Aj. Since fCS contains complex sulfated fucose branches, in solution, it causes its chain to stretch outward, forming an extended linear chain conformation. fCS-Aj and fCS-Hm have a random coil and extended linear chain conformations, respectively ^[47]. Li et al. studied sea cucumber CS: fCS-Pg, fCS-Ib, fuc-Pg, and fuc-Ib from diverse origins, where Pg and Ib are two species, and fCS denotes fucosylated CS, whereas fuc stands for fucoidan sulfate. An analysis of AFM images revealed that all the CSs except for fuc-Ib exhibited random linear chains. Chain lengths ranged between 100 and 1000 nm with some spherical aggregates. Notably, fuc-Ib showed a completely spherical structure. The variation in CS morphologies could relate to its fucose branches ^[30]. Dong et al. visually observed CS from discarded codfish bones as a white flocculent with an irregular lamellar structure using SEM, and spherical particles were visible after magnification. An observation under an AFM was able to show that CS was an irregular spherical structure with uniform distribution. The height of the individual polysaccharides was generally between 0.1 and 1 nm, indicating that the samples were in an aggregated state ^[48]. Yang et al. observed the structure of purified SCVP-1 from sea cucumber viscera using SEM, which showed a loose and irregular lamellar structure when magnified 500 times using SEM and continued to magnify it up to 2000 times to observe a rough surface morphology, curling, and a few spherical aggregates. SCVP-1 was not completely uniform in size and aggregated to form surface protrusions, as observed using an AFM ^[49].

Most studies typically use a combination of SEM and AFM images to analyze the sample surface, using SEM for the initial exploration and an AFM for the higher-resolution images at the nm level to distinguish surface changes at the atomic level and calculate the roughness of the sample surface.

The aforementioned measures can complement each other and accurately define the real and clear structure of polysaccharides. Nevertheless, physicochemical properties and extraction procedures may impact morphologies, resulting in discrepancies in the polysaccharide analysis of the same species. A more rigorous system must be established in future research processes to standardize the extraction and purification of polysaccharides and to avoid any structural discrepancies caused by external factors.

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Bochun Yang

Conghao Yang

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Update Date: 11 Jan 2024

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