Next Article in Journal
Probiotics as a Possible Strategy for the Prevention and Treatment of Allergies. A Narrative Review
Next Article in Special Issue
Comparison of Alkaline/Oxidative and Hydrothermal Extraction of Wheat Bran Arabinoxylans
Previous Article in Journal
Valorization of Sicanaodorifera (Vell.) Naudin Epicarp as a Source of Bioactive Compounds: Chemical Characterization and Evaluation of Its Bioactive Properties
Previous Article in Special Issue
Morus alba L. Plant: Bioactive Compounds and Potential as a Functional Food Ingredient
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:

Hydrogen Peroxide Effects on Natural-Sourced Polysacchrides: Free Radical Formation/Production, Degradation Process, and Reaction Mechanism—A Critical Synopsis

Chigozie E. Ofoedu
Lijun You
Chijioke M. Osuji
Jude O. Iwouno
Ngozi O. Kabuo
Moses Ojukwu
Ijeoma M. Agunwah
James S. Chacha
Onyinye P. Muobike
Adedoyin O. Agunbiade
Giacomo Sardo
Gioacchino Bono
Charles Odilichukwu R. Okpala
7,* and
Małgorzata Korzeniowska
Department of Food Science and Technology, School of Engineering and Engineering Technology, Federal University of Technology, Owerri, 460114 Imo, Nigeria
School of Food Science and Engineering, South China University of Technology, Guangzhou 510640, China
Food Technology Division, School of Industrial Technology, Universiti Sains Malaysia, Minden 11800, Penang, Malaysia
Department of Food Technology, Nutrition and Consumer Sciences, Sokoine University of Agriculture, 3006 Morogoro, Tanzania
Department of Food Technology, University of Ibadan, 200284 Ibadan, Nigeria
Institute for Biological Resources and Marine Biotechnologies—IRBIM, National Research Council (CNR), Via Vaccara, 61, 91026 Mazara del Vallo, Italy
Department of Functional Food Products Development, Faculty of Biotechnology and Food Science, Wroclaw University of Environmental and Life Sciences, 51-630 Wroclaw, Poland
Authors to whom correspondence should be addressed.
Foods 2021, 10(4), 699;
Submission received: 18 February 2021 / Revised: 20 March 2021 / Accepted: 22 March 2021 / Published: 25 March 2021


Numerous reactive oxygen species (ROS) entities exist, and hydrogen peroxide (H2O2) is very key among them as it is well known to possess a stable but poor reactivity capable of generating free radicals. Considered among reactive atoms, molecules, and compounds with electron-rich sites, free radicals emerging from metabolic reactions during cellular respirations can induce oxidative stress and cause cellular structure damage, resulting in diverse life-threatening diseases when produced in excess. Therefore, an antioxidant is needed to curb the overproduction of free radicals especially in biological systems (in vivo and in vitro). Despite the inherent properties limiting its bioactivities, polysaccharides from natural sources increasingly gain research attention given their position as a functional ingredient. Improving the functionality and bioactivity of polysaccharides have been established through degradation of their molecular integrity. In this critical synopsis; we articulate the effects of H2O2 on the degradation of polysaccharides from natural sources. Specifically, the synopsis focused on free radical formation/production, polysaccharide degradation processes with H2O2, the effects of polysaccharide degradation on the structural characteristics; physicochemical properties; and bioactivities; in addition to the antioxidant capability. The degradation mechanisms involving polysaccharide’s antioxidative property; with some examples and their respective sources are briefly summarised.

1. Introduction

Free radicals are a group of reactive atoms, molecules, and compounds with electron-rich sites produced as a result of metabolic reactions during cellular respiration [1,2]. In biological/food systems, free radical reactions are predominantly present and play a key role in biochemical pathways [1]. Collectively, free radicals are classified in two groups, namely reactive oxygen species (ROS) and reactive nitrogen species (RNS). The ROS involves oxygen-based highly reactive free radicals whereas the RNS involves nitrogen moieties associated with oxygen. In other words, RNS is a subset of ROS. Since oxygen radicals were discovered as harmful molecules [3], the causative agent of oxidative stress [4], and among essential free radicals generated during cellular metabolism, research interest on ROS has increased impressively. The ROS remains in the group of reactive oxygen-containing molecules possessing one or more unpaired electrons capable of existing independently [5]. The ROS comprises radical species and non-radical species. The former involves chemically reactive oxygen radicals such as hydroxyl, superoxide, peroxy, and alkoxy radicals, whereas the latter involves non-radical derivatives of oxygen such as hydrogen peroxide (H2O2,), hypochlorous acid (HOCl), singlet oxygen (O2), ozone (O3), and peroxynitrite (ONOOH) [6]. Moreover, non-radical species though not free radicals can still easily result in free radical reactions in living organisms, once conditions are favourable [7,8]. The ROS can be seen as a natural by-product of aerobic cellular metabolism with a double role, in physiological regulation and a key biological messenger in cell signalling cascades [9,10,11] such that when in low concentration, could cause oxidative stress, and when in excess or high concentration, could significantly damage the cell structures. Oxidative stress has always been hypothesized to be linked to excess production of ROS and/or impaired antioxidant defense capacity [12] or has also been perceived to occur due to an imbalance between the production of ROS and the available antioxidant or radical scavengers [13]. Over time, the accumulation of oxidative stress determined by the action of free radicals can cause lipid, protein, and DNA damage or alterations [7,14,15], as well as many life-threatening diseases including cancer, cardiovascular, Alzheimer disease, and related neurodegenerative diseases [11,16] diabetes mellitus, rheumatoid arthritis, cataracts, respiratory disease and aging process [7].
Numerous ROS entities exist, and H2O2 is very key among them as it is well known to possess stable [7,17] but poor reactivity [1] capable of generating free radicals. In addition, H2O2 is a metabolite of superoxide metabolic reactions [17] and a substrate for the production of hydroxyl radicals during cellular respiration in living organisms. Since its discovery, there has been a substantial increase in research interest regarding in vivo and in vitro formation mechanisms of H2O2—a ubiquitous endogenous molecule considered toxic to human tissues [18], and how it functions as a biomarker (target) in directing therapeutic delivery for oxidative stress-related diseases [19]. H2O2 is generated via respiratory chain cascade, but also a by-product of cellular metabolism [17]. Chemically, in contrast to superoxide anion (O2•−) and hydroxyl (OH•) radical, H2O2 is not very reactive [1] especially in the absence of transition metal ions and can act as a mild reducing and oxidizing agent. Though H2O2 does not readily oxidize proteins, lipids, and DNA [20], how it gets involved in such numerous physiological processes as cell signal transduction, cell differentiation (apoptosis), growth, and maintenance, as well as mediation of immune responses [17] demonstrates its dual role potential. Like other ROS molecules, and besides the dual functionality of H2O2, there are reported levels of H2O2 being cytotoxic to cells when the concentration is greater than 50 µM and limited cell cytotoxicity when concentration is at or below about 20–50 µM [20]. The effect of H2O2 in a living system is dependent on the type of cell, its concentration, its physiological state, and duration of exposure [17,20,21,22]. Interestingly, H2O2 can be part of redox signalling but can induce oxidative destruction of organs and tissues, associated with other inflammatory responses when overexpressed in cells [11,23]. Note worthily, H2O2 can produce very reactive OH• free radicals either in the presence of redox metal ions, such as copper or iron (Fenton reaction) or in the presence of oxygen radical (Haber-Weiss reaction) [24]. Hence, the need for antioxidants to scavenge free radicals from ROS which are causative agents of oxidative stress cannot be over-emphasized.
Antioxidants are substances able to inhibit oxidation caused by free radicals and unstable molecules produced in a living organism. They are applied to foods to inhibit free radical formation due to lipid oxidation and reduce the in vivo free radical concentration after (food) digestion [25]. The oxidative processes of antioxidants are underpinned by free radicals, being propagated by chain reactions in foods. For the most part, lipid oxidation (LOx) is sure to occur via mechanisms that involve complex processes such as autoxidation, enzymatic/thermal oxidation, and photo-oxidation. These processes are already well explained elsewhere [26,27]. For emphasis, antioxidants have two major groups that include: natural and synthetic types. Natural antioxidants have two major groups, namely, enzymatic and non-enzymatic antioxidants. Enzymatic antioxidants have primary (catalase, glutathione peroxidase (GPx), and superoxide dismutase) and secondary (glucose-6-phosphate dehydrogenase and glutathione reductase) enzymes, whereas non-enzymatic antioxidants have carotenoids, cofactors, flavonoids, minerals, phenolic acids, and vitamins. On the other hand, synthetic antioxidants can include butylated hydroxytoluene (BHT), butylated hydroxyanisole (BHA), tert-butylhydroquinone (TBHQ), and esters of gallic acid (e.g., propyl gallate) [27]. Nowadays, nutritional awareness has significantly influenced consumer demand and acceptance for safe, organic, and natural antioxidant sources. The preference for organic (food) products and/or additives such as antioxidants over synthetic ones by most consumers has increased interest in the use of antioxidants from natural sources. In addition, the components that make the antioxidant functions/mechanism work include its hydroperoxide destroyers, metal chelators, oxygen quenchers, radical scavengers, as well as (antioxidant) regenerators [27].
Polysaccharides are among the most abundant biomaterials which have attracted serious research interest in the field of pharmacology and biochemistry [25,28,29]. In recent years, polysaccharides obtained from natural (plants, animals, and microorganisms) sources have shown promising biological activity, such as antioxidation, antitumor, immunity, hypoglycaemic, and anti-aging properties, as well as cardiovascular, liver, and kidney protective effects [30,31,32,33,34], which is indicative of their high potential, especially in the field of medicine [25,35]. However, the high molecular viscosity/weight of many polysaccharides is considered to limit their pharmaceutical application. Modifying the chemical aspects of polysaccharides can therefore help achieve promising pharmacological/therapeutic targets [36,37]. The biological activities of polysaccharides may depend on several structural parameters, such as the branch structures, glycosidic linkages, and their types, molecular weight, and monosaccharide compositions. For instance, the β-(1→3) and β-(1→6) glycosidic linkages present within the repeating units of the polysaccharides have been deemed vital for anticancer activity [38]. Natural polysaccharides’ bioactivities and application are significantly influenced by the molecular weight of the native polysaccharide. For instance, a polysaccharide with larger molecular weight, on the one hand, may not easily penetrate cell membrane barriers in organisms and this could result in impaired biological effects [34,39,40]. On the other hand, Wu et al. [41] have reported higher biological activity in low molecular weight natural polysaccharides. Thus, finding efficient methods for the degradation of natural polysaccharides is crucial, if the biological activity of polysaccharides were to be facilitated. The physical, chemical, enzymatic, and/or a combination of these modification methods could substantially enhance both characteristics and utilities of (natural) polysaccharides.

2. Justification of Synopsis and Objective Statement

Ranked third among the macromolecules after nucleic acids and proteins that carry essential biological information, polysaccharides and their associated (protein) complexes from natural sources (such as bacteria, fungi, algae, and plants) continue to display effectiveness as non-toxic substances, demonstrated by diverse biological functions [42]. It is important to reiterate that among the active polysaccharides from natural sources, those that are sulfated are considered acceptable and safe [43]. Various polysaccharide extraction methods are possible due to novel techniques like enzyme-assisted extraction (EAE), microwave-assisted extraction (MAE), as well as ultrasonic-assisted extraction (UAE). Additionally, polysaccharide extraction has largely focused on both effects as well as the yield of a single variable, not forgetting its activity (that is, polysaccharides) [44]. However, considering the existence of ROS such as H2O2, the presence of free radicals that induce oxidative stress, and the need for an antioxidant to curb its (free radicals) overproduction within biological systems (in vivo and in vitro), it is important to understand how the aforementioned (physical, chemical, and/or enzymatic modification) methods can extend the characteristics and utilities of (natural) polysaccharides.
The chemical method of polysaccharide modification (degradation) via oxidation has been of increasing research interest, given the contributions to this body of knowledge specific to the degradation of polysaccharides via oxidation method largely by the H2O2 method. Although studies focused on the degradation of polysaccharides by free radicals increasingly grows, there is a paucity of literature that has synthesized the polysaccharide degradation by free radicals from H2O2, to particularly demonstrate its potential use in the agro-food industry, together with it being a viable candidate for in vivo biological activity especially when incorporated within the food matrix/system. On this premise, this current critical synopsis articulates the H2O2 effects on the degradation of polysaccharides from natural sources. Specifically, the structure of this synopsis will give emphasis to free radical formation/production, the effects of polysaccharide degradation with H2O2 on the structural characteristics, physicochemical properties, and bioactivities, in addition to the antioxidant capability, and reaction mechanism.

3. Survey Methodology

To actualise this critical synopsis, the first step taken was to formulate the research questions, ensuring the contribution of this review is relevant to this field, and at the same time, considering the intended/target audience. In the next step, after having agreed with the research questions, we developed a search strategy. This allowed for the establishment of both inclusion and exclusion criteria, which determined the appropriate research articles, and considered relevant specific to degradation of polysaccharides by free radicals from H2O2. Based on this chosen topic, the search terms such as polysaccharides, natural antioxidants, free radicals, ROS, polysaccharide degradation, H2O2 (formation/sources and reaction mechanism), and natural polysaccharides were identified and selected.
Published papers and their references were collated from databases like EMBASE, Google Scholar, Web of Science, Interscience Online Library, MEDLINE, ScienceDirect, and PubMed. By way of inclusion and exclusion criteria, the research articles not addressing the research question(s) were expunged, considering the year of publication and the language of the article (publications in languages other than English were excluded). Importantly, the data from the literature of selected studies were analysed and relevant information incorporated helped to expatiate the discourse of this conducted critical synopsis.

4. Discussion of Findings

4.1. H2O2 Formation and Factors Influencing its Production

The H2O2 remains a key affiliate of the group of ROS next to superoxide anion (O2•−) and OH• radicals, which is produced through the respiratory chain cascade and a by-product of cellular metabolism [17,45,46]. As a member of ROS, its reactivity targets proteins, lipids, and DNA alteration or damage, which brings about oxidative stress [47,48,49]. H2O2, as a crucial second messenger in normal cellular signalling [11], has had its overproduction associated with various life-threatening diseases [2,6,11,20] since it is considered cytotoxic [1,20]. Besides being the simplest form of peroxide [5] having two oxygen atoms covalently bonded (single bond), it is endogenously produced in diverse cellular aspects of tissues or organs (peroxisomes, endoplasmic reticulum, and mitochondria) at high levels of oxygen utilisation [7].
However, many studies have reported that H2O2 is a metabolite of O2•− produced in the mitochondria during cellular respiration propelled by electron transport chain (ETC) [6,7,17], and a substrate for the production of OH• radical during cellular metabolism [7]. The O2•− (half-life of 10−6s), OH• radical (half-life of 10−10s), and H2O2 (stable half-life) are key members of ROS [7]. H2O2 as a non-radical is fairly reactive (in the absence of transition metal ions) probably due to its stable half-life, but its reactivity is enhanced when it reacts with transition metal ion to yield OH• radical (Fenton reaction), as well as when it reacts with O2•− to yield OH• radical (Haber-Weiss reaction) [24].
Fenton   reaction : Fe 3 + + O 2     Fe 2 + + O 2   ( auto-oxidation ) Fe 2 + + H 2 O 2     Fe 3 + + OH + OH
Haber-Weisss   reaction : H 2 O 2 + O 2     O 2 + OH + OH
In the first instance (Equation (1)), the Fenton reaction begins with the production of superoxide radical via cellular metabolism. During cellular respiration, some electrons that escaped the electron transport chain can interact with oxygen (O2) to yield (unstable) O2•−, a member of ROS. As a non-enzymatic product, the generation of superoxide anion (O2•−) increases with an increase in metabolic rate, which causes cellular components destruction. Consequently, low amounts of superoxide anion (O2•−) are maintained in the cells with the aid of a metalloprotein enzyme [2] called superoxide dismutase (SOD), which catabolizes superoxide anion (O2•−) to H2O2 and subsequently catabolized (detoxified) by the enzyme catalase (metalloenzyme) and glutathione peroxidase (GPx) [7] to water and oxygen.
As shown in Figure 1, there is the electron transport chain, which involves a series of electron transporters embedded within the inner mitochondrial membrane that shuttles the cascade of electrons from the NADH and FADH2 to molecular oxygen. It is at this process that protons are pumped from the mitochondrial matrix to the intermembrane space, and oxygen is reduced to form water. Specifically, O2•− levels may rise, at times, after the metabolism of alcohol by actions of alcohol dehydrogenase (ADH) enzyme, and NAD+ generates a lot of nicotinamide adenine dinucleotide (NADH) [50]. As a result, more H2O2 is produced which may not be either detoxified and or hydrolysed, by the limited amount of enzyme (catalase and glutathione peroxidase). Moreover, the H2O2 would react with the transition metal ions, such as iron (Fe2+) and copper (Cu+), respectively found in the cells as ferritin and ceruloplasmin [51], and in that way, able to generate OH• radical, which remains a highly unstable molecule [52,53].
On the other hand, the Haber-Weiss reaction (Equation (2)) mechanism, which generates OH• radical from H2O2 and O2•− catalysed by iron as described by Fritz Haber and his student Joseph Joshua Weiss [5] is considered as among the key foundation of free radical biochemistry [53]. Figure 2 shows the role of H2O2 in cell dysfunction and transformation. Notably, both the Fenton and the Haber-Weiss reactions have proven the role of H2O2 as an intermediate and substrate respectively, for the production of OH• radicals responsible for oxidative stress and cellular damages [5,53]. The OH• radical, as a neutral form of hydroxyl ion (OH), remains the most reactive member of the ROS because of its short-lived half-life, estimated to be about 10−10 s. The destructive action of this unstable molecule (OH•) is implicated in series of cell dysfunction, transformation, and even death [5,54,55].

4.1.1. Formation of H2O2

The formation of H2O2, a member of ROS, is the outcome of an unavoidable aerobic metabolism or reaction [56]. Many studies have understood H2O2 as poorly reactive yet somewhat stable. However, the destructive activity and cytotoxic effects of H2O2 have been owed to the formation of OH• radical, when it reacts with either superoxide anion(s) or transition metals. The formation/production of H2O2 in vitro and in vivo [57] can, respectively, take place in certain beverages and humans.
In vitro production of H2O2
The production of H2O2 by various food systems has been reported previously. For instance, studies have shown H2O2 production from thiol-rich proteins during barley (malt) beer production [58] and in the association of L-cysteine with Cu(II)-catalysts during malting and beer brewing [59]. Also, H2O2 can be generated in vitro in certain polyphenol-rich beverages such as green tea, black tea [60,61,62], red wine, and coffee [60,63]. Polyphenols are a large group of compounds naturally found in fruits, vegetables, and certain beverages, possessing antioxidant properties. The protective effect of polyphenols which is attributed to its antioxidative property is believed to be due to its ability to act as a free radical scavenger, quenching the activities of ROS such as O2•− radical, OH• radical, and H2O2 [62,64]. In contrast, the production of H2O2 by polyphenols understood from the (polyphenol) pro-oxidant action via autoxidation [60,65] has been associated with induced mutagenesis and promotion of cancer [62], as well as to promote oxidative damage to DNA, lipids, and proteins in the presence of transition metal ions under certain conditions in vitro [66].
The phenolic compounds that produce H2O2 through the autoxidation process include tea catechins (epicatechin [EC], epicatechin gallate [ECg], epigallocatechin [EGC], and epigallocatechin gallate [EGCg]) [61] (+)-catechin, and gallic acid [67]. Further, Grzesik et al. [63] reported that (black and green) teas and coffee but not cocoa were shown to produce considerable concentrations of H2O2. In such beverages (green tea, black tea, and coffee), the H2O2 generating property is poised to coincide with the content of phenolic compounds. This suggested that polyphenols are liable for H2O2 production in beverages. Results from previous researches indicated that the formation of H2O2 from the aforementioned phenolic compounds in an aqueous solution would depend on the content and composition of polyphenols, structure of the phenolic compound, oxygen, metal ions, temperature, pH, and incubation time [66]. Therefore, the combined effect of these conditions would result in the formation as well as diverse concentration(s) of in vitro H2O2 production.
In vivo production of H2O2
The H2O2 can be naturally produced in vivo for example, in humans, as a by-product of oxidative metabolism [5] by the dismutation of superoxide radical, both non-enzymatically and catalysed by superoxide dismutase (SOD) enzymes [68]. Besides the superoxide dismutase (SOD) enzyme, there are other oxidase enzymes such as glycollate and monoamine oxidases, directly linked to the production of H2O2 [68,69]. Additionally, the glutathione peroxidase and thioredoxin-linked peroxidase are enzymes contained in the mitochondria. However, the effectiveness of H2O2 removal by the enzymes remains unclear, given the rate at which it is generated in the mitochondria [70,71]. Although certain tissues may be exposed to higher H2O2 concentrations with the mitochondria being at the helm of activity in human cells, most cells may also be exposed to certain H2O2 levels.
The formation of H2O2 in the oral cavity, oesophagus, stomach, respiratory system, kidney, urinary tract and bladder, vascular endothelial and circulating blood vessel as well as ocular tissues are well documented. Lambert et al. [72] reported the in vivo generation of H2O2 in micromolar concentrations produced when green tea solution is held in the mouth. Halliwell et al. [68] reported that beverages like green tea, black tea, and instant coffee contain H2O2 at concentrations above 100 µM, probably because of the joint action of autoxidation of antioxidants in the beverages and oral bacteria. Production of H2O2 by oral bacteria has been previously reported, although its impact on oral tissues remains unknown. Moreover, H2O2 can be present in exhaled air of healthy humans although its concentration is higher in those with inflammatory lung diseases and in cigarette smokers [68]. This H2O2 in exhaled air can possibly be produced by oral bacteria since cells lining the respiratory system are exposed to about 21% oxygen. Besides H2O2 detected in freshly voided human urine [68], it could be at a higher concentration in the urine of coffee drinkers [63]. The presence of H2O2 in human urine may be due to the dismutation of superoxide radicals which may come from the diet via autoxidation. Adahi et al. [73] detected some traces of superoxide dismutase (SOD) in the urine. Additionally, the exhalation of H2O2 and its removal through urine could be part of the human excretory mechanism.
The in vivo generation of H2O2 in blood plasma by the action of xanthine oxidase enzyme is degraded in part by traces of catalase. Like in blood plasma, ocular fluids have been reported to generate H2O2 constantly which is usually rapidly removed [74,75]. Though the origin of ocular H2O2 is unknown, oxidation of ascorbate may likely be a contributory factor [76]. Importantly, it is expected that H2O2 would accumulate as a result of the joint action of in vivo dismutation of superoxide radical and in vitro autoxidation of phenolic compounds (antioxidants). However, there is no evidence that these beverages (teas and coffee) promote or induce mutagenesis and carcinogenesis in vivo. Possibly, any threat the in vitro generated H2O2 from polyphenols is likely to pose could be inhibited by phenolic compounds and other constituents of the beverages [60], or even catalase and glutathione peroxidases in the body, and decomposed by the reaction with biomolecules [62]. In a research on dietary antioxidants as a source of H2O2, Grzesik et al. [63] reported that 27 out of 52 antioxidants generated H2O2, suggesting the availability of other constituents capable of exerting antioxidant property in the beverage.

4.1.2. Factors That Influence the Production of H2O2

As was earlier mentioned, the in vivo and in vitro formation of H2O2 is dependent on some factors that could inhibit or enhance its production, and they include the composition and structure of phenolic compounds, oxygen, metal ions, pH, enzyme, and incubation condition (temperature and time).
Composition and structure of phenolic compounds
According to Akagawa et al. [62], only o- and p-phenolic compounds undergo autoxidation. Additionally, EGC, EGCg, and compounds with gallic acid moieties have been reported to produce H2O2 during oxidation in aqueous media [61,77,78,79] while Akagawa et al. [62] reported that the formation of H2O2 by tea catechins followed the order: epigallocatechin gallate > epicatechin gallate ≡ epigallocatechin > epicatechin > catechin.
Phenolic compounds with OH-group, substituted at the 2 and 4 positions of phenol, would generate H2O2 [80]. Such production would significantly increase by the insertion of a third OH-group. This, therefore, corroborates the relationship between the H2O2 generating property of phenolic compounds and their structures [61] (Figure 3). Akagawa et al. [62] reiterated this, in that the insertion of a third adjacent hydroxyl group, that is, the presence of gallate substituent (pyrogallol moiety) in the catechins, would increase the formation of H2O2. In practice, a significant amount of H2O2 can be produced by gallic acid. The H2O2 generating property in these beverages was found to coincide with the content of phenolic compounds, indicating that polyphenols are responsible for the production of H2O2 in beverages.
Oxygen and metal ions
Oxygen is essential for the viability and maintenance of cell metabolism but has been found to play a key role as a potential promoter of in vitro and in vivo free radical reactions [81,82]. Electronically it is an highly excited molecule that exists in two excited states upon activation. In the mitochondria, the formation of H2O2 is initiated by oxygen during the mitochondrial respiratory chain, where about 85% of oxygen is metabolized. The transition metal ions mainly considered by researchers are iron, manganese, copper, and sometimes zinc because these species are major players in free radical or electron transfer reactions [1]. The joint action of transition metal ions and oxygen in the production of H2O2 has been reported in many studies. According to Mochizuki et al. [83], tea polyphenols are easily oxidized to the corresponding quinones by oxygen (O2) in the presence of transition metal ions, incidentally resulting in the formation of O2•− and H2O2 through autoxidation (Figure 4). In addition, higher H2O2 formation has been detected in tea infused in tap water containing transition metal ions than a tea infused with deionised water [63], thus suggesting that the autoxidation of polyphenols would produce H2O2, catalysed by transition metal ions. The autoxidation of polyphenols by oxygen to yield semiquinone radical and O2•− in the absence of metal ions was reported by Akagwa et al. [62] though with a slow reaction rate due to the low redox potential of the O2/O2-. On the other hand, metal ion (Cu2+) catalysed autoxidation of polyphenols to yield semiquinone radical and Cu+ have also been reported. Subsequently, O2•− and quinone is produced when the generated semiquinone radical is rapidly oxidized by oxygen [83]. Hence, the generated O2•− helps to make the H2O2 spontaneously via dismutation [62], and also to H2O2 and semiquinone radical via oxidation of polyphenols.
Multiple studies have reported the correlation between pH and H2O2 formation from tea polyphenols. Mochizuki et al. [83] reported that the autoxidation of polyphenols depends on the pH of the medium (tea) and is accelerated by alkaline pH. Similarly, Grzesik et al. [63] reported that the rate of autoxidation of compounds increased with pH, which corroborated with findings of Rinaldi et al. [84] and Akagawa et al. [62] wherein the strong electron-donating ability of acid dissociated the phenolic compound, as polyphenols underwent autoxidation at a higher rate with increased pH. Nakayama et al. [61] detected H2O2 in polyphenol-rich beverages (tea and coffee) when the pH value is above 5.8 and its concentration increased with pH up to 8.0. A maximum formation of H2O2 at pH 7.0–9.0 for tea polyphenols has also been reported [62]. However, the addition of milk and lemon respectively decreased and prevented H2O2 formation in tea [60,63]. This was shown by the lower pH ranges in milk (pH 6.5–6.7) and lemon (pH 2–3), which suggested H2O2 formation as pH-dependent, where a decrease in pH reduces or prevents its (H2O2) formation and vice versa.
Incubation condition
The incubation condition of tea polyphenols plays a significant role in H2O2 formation. The action of temperature-time incubation conditions cannot be over emphasised. Studies have shown that an increase in incubation temperature and time increases the rate of H2O2 formation. In the same vein, Nakayama et al. [61] reported an increase in H2O2 formation when the incubation temperature of tea was increased from 37 °C to 77 °C. However, Nakayama et al. [66] reported that the concentration of H2O2 in tea increased concavely with temperature when increased from 32 °C to 82 °C. Also, the amount of H2O2 formation in tea has been reported to increase almost linearly with incubation time [66], suggesting that autoxidation of polyphenols is also time-dependent.
Enzymes have been found to participate in the production of H2O2. Some H2O2-producing enzymes include but are not limited to superoxide dismutase (SOD), and other oxidase enzymes such as glycollate and monoamine oxidase [68,69], glucose oxidase, galactose oxidase, and cholesterol oxidase [85]. Superoxide dismutase catabolizes O2•− to form H2O2 [7], while glucose oxidase catalyses the oxidation of β-D-glucose by oxygen to form H2O2 and gluconolactone (Figure 5) [85]. Also, cholesterol oxidase catalyses the oxidation of cholesterol by oxygen to yield H2O2 (Figure 6) while galactose oxidase catalyses the oxidation of primary alcohol and polysaccharides (for example, D-galactose) and primary alcohols by oxygen, resulting to corresponding aldehydes and H2O2 as shown in the equation below:
RCH2OH + O2 → RCHO + H2O2
Notably, some key situations associated with the formation of H2O2 which can combine together via series of chain reactions under both in vitro and in vivo conditions are briefly presented in Table 1. In the mitochondria (in vivo), oxygen combines with an escaped electron in the electron transport chain to form O2•−, which is subsequently oxidised to H2O2 by superoxide dismutase (SOD). This implies that the formation of H2O2 in the mitochondria is dependent on the O2•−, which may increase in concentration with an increasing amount of NADH. On the other hand, in beverages (in vitro) such as tea and coffee, autoxidation of polyphenols by oxygen, catalysed by transition metal ions to yield corresponding quinones (depending on the composition and structure of phenolic compound) subsequently generates H2O2, whose concentration is dependent on the beverage incubation condition (temperature and time).

4.2. Polysaccharide: The Degradation Process and Effects with H2O2

Polysaccharides are the most common biopolymers and copious organic material in existence [25,92,93]. For instance, the carbohydrate molecules of carbon and oxygen atoms differ in size and range from one to thousands of units. Simple sugars (monosaccharides) are the smallest unit of a polysaccharide molecule. Examples include glucose, fructose, mannose, and galactose. Polysaccharides are broadly classified as homo-polysaccharides, which can include, one type of monosaccharide units like glycogen, cellulose, and starch, and the hetero-polysaccharides, which can include two or more types of monosaccharide units like hyaluronic acid, heparin, etc. Furthermore, cellulose is a structural polysaccharide found in plants that forms the main structural component of plant cell wall [25] whereas glycogen and starch are storage polysaccharides found in animals (muscles and liver), and plants respectively. Besides the well-known animal/plant polysaccharides, there are other less popular polysaccharides that arise from sources like bacteria, fungi, and algae [25].
Ever since polysaccharides were discovered to possess some biological activity, there has been increasing research interest on this specific aspect in the field of biochemistry, life science, pharmacology, and medicine, together with its possible applications [25,94]. Garcia-Gonzalez et al. [95] highlighted the use of polysaccharides as key functional ingredients in the production of bio-based materials in food, cosmetics, medical devices, and pharmaceutics. However, polysaccharides obtained from plants, animals, and microorganisms have been shown to possess diverse properties not limited to antioxidative, anti-inflammatory, antimutagenic, antitumor, health-promoting, and therapeutic properties [40,94,96,97,98]. With the many bioactivities/properties exerted by polysaccharides, our focus in this current review was on the resurgent antioxidative effect of polysaccharides, which can be explored as a potential novel functional food additive in food formulation.

4.2.1. Polysaccharide Degradation

There is evidence that, despite exerting antioxidative properties, not all polysaccharides have bioactivity, which is the situation, for example, of untreated pachymaran [99]. Additionally, some polysaccharides might not exhibit any bioactivities given their structural and physicochemical properties [94]. Many studies have shown that the functional role of polysaccharides is dependent on their degree of polymerization and molecular weight [46]. This was reiterated by Li and Shah [100] wherein the bioactivities of most polysaccharides were influenced by molecular weight and structural characteristics such as monosaccharide compositions, linkage patterns, anomeric carbon configuration, branching features, degree of polymerization, and sequence of sugar units. Despite these factors, several studies have shown that polysaccharides’ bioactivities are significantly influenced by the molecular weight of the native polysaccharide because polysaccharides with too high (large) molecular weight may not easily penetrate cell membrane barriers in organisms due to their high viscosity and low solubility, and will therefore result to impaired biological effect [34,39,40,94,101]. Hence, the physicochemical properties and bioactivities of polysaccharides are significantly influenced by their structure; so the structure of polysaccharides must be modified via degradation to the desired molecular weights that will enhance the bioactivities of polysaccharides.

4.2.2. Methods of Polysaccharide Degradation

The growing realisation of the potential novel products with diverse applications which may be obtained from degradation (depolymerisation) of the structural integrity of polysaccharides gave rise to the development of different degradation methods [102]. Generally, the degradation of polysaccharides can be achieved via physical, chemical, and enzymatic methods [40,102,103], as well as biological means [94], to obtain a diversity of structural derivatives. Additionally, some other methods of polysaccharide degradation such as thermal, ultrasonic, mechanical, lyophilisation-mediated, metal-catalysed, free radical-catalysed [102], and irradiation [104] degradation have been previously described. Besides the advantages of primary degradation methods such as being simple to operate or easy to handle (chemical method), environmentally friendly (physical method), low cost, good safety, and mild reaction condition (biological method), each method also has its limitations. Multiple studies have reported environmental pollution and serious safety concerns (chemical method), utilization of specialized equipment which limits their industrial application (physical method), and difficulty in the screening of enzymes and microbes (biological method), as some of the challenges attributed to these treatment methods for polysaccharide degradation [105,106,107,108]. With the aforementioned methods of polysaccharide degradation, the chemical degradation through the oxidation method (either singly or combined) continues to attract researchers’ interest. For example, Zhao. [34] showed that the free radical-catalysed H2O2 method is frequently employed when they investigated polysaccharides from Tremella fuciformis. Furthermore, the fragmentation (degradation) of polysaccharides by ROS has been increasingly evidenced, as the bioactivities and overall functionality (of polysaccharides) gets altered [46]. Noteworthily, treating polysaccharides with H2O2 (only) does not generally degrade its high molecular weight, instead, the activation of H2O2 is necessary to produce highly reactive OH• radicals that can enhance its degradation property [106].
H2O2, as a member of ROS, during polysaccharide degradation operates under mild reaction conditions, with high efficiency, despite its inability to compromise the integrity of the main chain (polysaccharide) structure [40,98,109]. However, the various degradation methods employed are mediated by different reaction mechanisms, to yield potentially distinct products. Recently, desirable modified polysaccharides have been obtained with higher bioactivities by combining different degradation methods [94]. According to Wu et al. [104], the synergistic effect of combining different polysaccharide degradation methods significantly affects the depolymerisation kinetics of the process. Based on this, several combined polysaccharide degradation methods developed include H2O2-Vitamin C, H2O2-UV, H2O2-NaClO2, H2O2-HAc, Fe2+-H2O2, Fe2+-H2O2-vitamin C, Solution plasma process (SPP) irradiation, and SPP irradiation–H2O2, Ultrasonic irradiation-H2O2 [34,36,40,104,110,111,112], as well as a compound treatment of H2O2-vitamin C-microwave treatment [107]. In other words, H2O2 serves as an important source of reactive oxygen species (ROS) for these combined treatments, to assure adequate production of OH• radicals for significant degradation of polysaccharides.

4.2.3. Effect of Polysaccharide Degradation with H2O2

H2O2 continues to be a promising candidate for polysaccharide degradation. Polysaccharide degradation (depolymerisation) with H2O2 or in combination with another technique, by changing its functional role, would initiate its (polysaccharide) modification. Largely, the H2O2 is used in polysaccharides degradation given that it is easily available, easy to handle, and environmentally friendly since it decomposes to water and oxygen [106,113] as well as its ability to form reactive OH• radical through Fenton reaction [104]. In free radical-catalysed depolymerisation, the hydroxyl radicals via bond (glycosidic linkage) cleavage on the polysaccharides chain [102], would bring about molecular modification. Consequently, the modification of polysaccharides would lead to changes in physicochemical (relative molecular mass, viscosity, and solubility) and structural properties, as well as improved bioactivities. Li et al. [94] obtained enhanced biological properties in degraded/modified polysaccharides. Thus, both physicochemical and structural properties will be tersely discussed below:
Physicochemical Properties
Polysaccharide degradation can cause numerous changes in the physicochemical properties, from a decrease in its crystallinity as a result of structural rearrangements [114] to the improved tensile strength of its chitosan polysaccharide [94,115]. To understand the effect of polysaccharide degradation on the physical and chemical property, we attempt to examine aspects like its molecular weight, intrinsic viscosity, and water solubility, which are discussed briefly below.
Molecular weight
The molecular weight of polysaccharides is the basis for their bioactivities. Polysaccharides with high molecular weight are associated with low bioactivity, due to the compactness of their structures, low surface area, and stronger intramolecular hydrogen bonds [30]. Interestingly, the intrinsic viscosity and water solubility of polysaccharides depends significantly on their molecular weight, which could indicate a clear correlation between these properties. High molecular weight polysaccharides could lead to increased viscosity and decreased water solubility. On this premise, the degradation of high molecular weight polysaccharides could result in a modified (polysaccharide) type, able to easily permeate the cell membrane barriers of organisms, which could, in turn, exert improved bioactivity. Suggestively, the bioactivity of polysaccharides significantly would depend on their molecular weight.
The molecular weight of degraded or modified polysaccharides would decrease significantly, compared to the native (unmodified) polysaccharide. For example, Zhang et al. [34] reported a successful degradation of high molecular weight polysaccharides from 599,580 to 26,895 Da. In addition, the rate of polysaccharide degradation into lower molecular weight (degraded) polysaccharide significantly depends on the concentration of degradation reagent (H2O2). A high molecular weight Astragalus polysaccharide (11,033 Da) treated with different concentrations of H2O2 (4%, 6%, and 14%) was significantly modified (degraded) to lower molecular weight polysaccharides of 8376, 4716, and 2600 Da, respectively [30]. Therefore, whilst degradation of polysaccharides enhances their bioactivities, the rate of degradation into lower molecular weights is determined by the concentration of H2O2 which was utilized during the degradation. Notwithstanding that low molecular weight polysaccharides can exert added bioactivities, degraded polysaccharides with very low molecular weight, on the other hand, might bring about a reduction or loss in polysaccharide bioactivity as the inactive polymeric structure develops [101]. That is to say, the destruction of the polysaccharide ring structure by the H2O2 degradation system in very low molecular weight polysaccharide also breaks its intramolecular hydrogen bond, making the carbonyl group be in an open chain, thereby resulting in reduced bioactivity.
Intrinsic viscosity
Multiple studies have shown the correlation between molecular weight and intrinsic viscosity. Due to the high molecular weight of polysaccharides, the high intrinsic viscosity of polysaccharides limits their absorption and utilization in vivo [40,94]. Notably, the intrinsic viscosity of degraded or modified polysaccharides decreases significantly with a decrease in molecular weight compared to the native polysaccharide. Xu et al. [40] observed a decrease in intrinsic viscosity of degraded polysaccharides from 2.26 to 1.43 mPa.s. Similarly, Wu et al. [104] observed a reduction in intrinsic viscosity of chitosan polysaccharides by 82.19% and 70.04% when the chitosan polysaccharide was treated with SPP irradiation-H2O2 and SPP irradiation degradation method, respectively. This implies that the rate of reduction of intrinsic viscosity of polysaccharides is higher when two or more polysaccharide degradation methods are combined. Zhang et al. [34] observed a successful degradation with a 77.07% decrease in the intrinsic viscosity of polysaccharides, from 285.24 to 65.39 mL/g using Fe2+-H2O2-Vc degradation method. Similarly, Li et al. [94] observed a decrease in intrinsic viscosity of modified polysaccharide solution from 34.67 to 22. 92 mL/g which was believed to be attributed to a reduction of the hydrogen bonds. It is evident that the molecular weight of polysaccharides is concordant to their viscosity. This was consistent with Zhang et al. [36] who reported that the decrease in viscosity of polysaccharides is attributed to the reduced interaction between macromolecules, due to a slight decrease in their molecular weight.
Water solubility
Most bioactive polysaccharides have not found useful applications because of their poor water solubility, a result of their high molecular weight [40,94]. On the other hand, many studies have recorded a significant increase in water solubility of polysaccharides after degradation or depolymerisation,. Xu et al. [40] found that the solubility of degraded polysaccharides increased from 18.24 to 24.40 mg/mL with a decrease in molecular weight. Free radical-catalysed degradation offered by H2O2 has been found to increase polysaccharide solubility because of the inauguration or incorporation of carboxyl groups, ionic groups, or other hydrophilic groups into the polymer structure [94]. In addition to increased water solubility, degraded polysaccharides had a greater surface area which translates to higher reaction rates [34,98]. Indeed, the increase in water solubility of degraded polysaccharides would bring about a significant transformation of polysaccharides especially to their application, to exert improved bioactivities.
Structural Properties
The structure of polysaccharides influences their biological activity [30]. Polysaccharide degradation can produce different types of structures with varying bioactivities which lays a solid foundation for the functionality of the degraded or modified polysaccharide [94]. Free radical-catalysed degradation of polysaccharides offered by H2O2 reduces the association of amylose molecules by introducing small amounts of carbonyl and carboxylate groups [116] into the polymer structure which can improve the hydrophilic properties of the modified or degraded polysaccharides [94,117]. However, the surface area of the fragmented structure of degraded or modified polysaccharides could decrease significantly compared to its native polysaccharides, specifically due to the destruction of the aggregates or original interconnection in the polysaccharides by free radical-catalysed treatment offered by H2O2 [40,44]. Generally, by using only H2O2 or in combination with other degradation techniques, the main chain structure in the degraded or modified polysaccharides may not alter, but the side chain structure can change [30]. The degradation of chitosan structure, according to Tian et al. [118], only alters the side groups, but no significant change took place within the main chain structure. Some enhancement can take place at the side groups on the main chain polysaccharide structure, such as hydroxyl (–OH) and carboxyl (-COOH) groups, especially after the polysaccharide degradation. This can happen owing to the exposure of its effective active sites (groups), which thereby increases its content. Moreover, Wang et al. [30] detected a slight increase in degraded polysaccharide after the polysaccharide chain was broken by the OH• radical produced by the H2O2 degradation system, thus exposing the –COOH groups, whereas Zhao et al. [119] detected high levels of –COOH group in degraded polysaccharides. Similarly, Chang et al. [120] showed that a polysaccharide with a molecular weight of 129 KDa, degraded to lower molecular weights of 60 and 52 KDa, recorded an increase in their uronic acid content from 70.8% before degradation to 79.5% and 86.2% in the degraded polysaccharide, respectively. This was also consistent with the findings of Xu et al. [40] and Wu et al. [104]. Additionally, the helical structure of the main chain and hydrophilic group (hydroxyl group) influences the polysaccharides’ bioactivities. Also, polysaccharides with compact structures tend to possess stronger intramolecular hydrogen bonds, leading to less accessibility or exposure of its effective active sites (groups). To shed more understanding on this, we tersely discussed polysaccharides’ bioactivities and their workings.
There is increasing evidence that the functionality of polysaccharides such as their bioactivities can be enhanced only when their solubility in water increases, and this can be achieved via degradation or modification of polysaccharides [121]. In particular, the increase in hydrophilic properties of degraded polysaccharides, which has been attributed to their decreased molecular weight, lowered intrinsic viscosity, increased surface area, and structural changes (incorporation of carboxyl groups into the polymer structure), makes these degraded polysaccharides soluble in water, and as a result, exerts better bioactivities. Despite the many bioactivities, which include but are not limited to, antioxidant activity, antitumor activity, anti-inflammatory activity, anti-HIV activity, anticoagulant activity, antibacterial activity, antiviral activity, and immunomodulatory activity [94,121], our focus in this current literature synthesis is on the antioxidant activity of polysaccharides. Some researchers, however, have demonstrated that improved antioxidant activity of polysaccharides would associate with its degree of water-solubility [34,37]. To understand the workings of bioactivities, we shed some light on polysaccharides’ antioxidant activity in terms of their radical scavenging capacity, reducing power, and metal ion chelating ability.
Antioxidant activity
Antioxidants are substances or compounds that inhibit oxidation caused by free radicals and unstable molecules produced in food or living organism. Antioxidants are specific in their action by performing their primary function of breaking chain reactions through free radical scavenging and also by performing their secondary function (preventive role) through chelation of metal ions that catalyse a free radical reaction. The antioxidant activity of substances or compounds is displayed through series of reaction mechanisms which include transition metals chelating ability, hydrogen electron transfer, and single electron transfer [81]. However, much research has shown that various polysaccharides obtained from plants, animals, fungi, bacteria, and algae possess antioxidant properties [94], especially when such a polysaccharide is the degraded or modified type. Specifically, polysaccharides are compounds with large molecular weights and as a result would have lesser non-reducing end contents, hence, weaker antioxidant activity [30]. Based on this fact, the degradation of polysaccharides can effectively better their antioxidant activity. Furthermore, series of studies on free radical-catalysed polysaccharide degradation have shown that the antioxidant activity of degraded polysaccharides was ascertained by determining their radical (OH•, O2•−, DPPH, and ABTS) scavenging property, reducing power and metal ion chelating ability.
  • Radical scavenging capacity
The scavenging capacity of free radicals (OH•, O2•−, DPPH, and ABTS) by degraded polysaccharides has been documented in many studies. The OH• radical is one of the most unstable (half-life of 10−10s) and strongest oxidants among the ROS, randomly able to attack biomolecules, and cause oxidative damage to certain proteins, lipids, and nucleic acids [109,122]. Degraded polysaccharides have been found to show higher OH• radical scavenging capacity compared to the native (undegraded) polysaccharides. The OH• radical scavenging rate would be dependent on the concentration of degraded polysaccharides [109,122]. Wang et al. [30] reported an increase in radical scavenging rate when the concentration of degraded polysaccharide was increased from 0.15 mg/mL to 3 mg/mL. Similarly, Xu et al. [40] found that degraded polysaccharides of varying molecular weights exhibited different degrees of scavenging effects on the hydroxyl group, and this effect increased with decreasing molecular weight of polysaccharides. In addition, a polysaccharide with a molecular weight of 3.26 × 104 KDa, degraded to 1.30 × 104 and 9.62 × 103 KDa was found to show an increased OH• radical scavenging rates from 66.89% before H2O2 degradation to 92.58% and 93.21% in the degraded polysaccharide respectively [40].
The O2•− radical (half-life 10−6 s) is another key member of ROS, produced continuously in biological systems, and could either cause harm to cell components or serve as a precursor to other unstable reactive free radical species [40,100]. Just like in OH• radical, degraded polysaccharide showed higher O2•− radical scavenging capacity than the native polysaccharide, and the scavenging capacity was also enhanced with increasing concentration of degraded polysaccharide. The O2•− radical scavenging capacity of native polysaccharide (21%) and degraded polysaccharide (39%) at a concentration of 0.2 mg/mL increased significantly to 53.41% (native polysaccharide) and 62.12% (degraded polysaccharide) when the concentration was raised to 1.2 mg/mL [40]. Similarly, Wu et al. [104] also reported a significant increase in O2•− radical scavenging capacity (rate), when the concentration was 0.2 mg/mL with a scavenging rate of 12% and 32% for native polysaccharide and degraded polysaccharide was increased to 1.2 mg/mL, resulting in an increased scavenging capacity of 20% and 66% for native polysaccharide and degraded polysaccharide, respectively.
Furthermore, DPPH and ABTS radicals are (stable) free radicals frequently employed in radical scavenging assays for evaluating antioxidant capacity (of relevant compound(s)). Similarly, studies have shown that degraded polysaccharides showed significant scavenging capacity on DPPH and ABTS radicals, over the native polysaccharide [25,40,104,123] and the degree of the radical scavenging capacity of degraded polysaccharide on DPPH and ABTS radical were concentration-dependent. This implies that the degraded polysaccharide can easily react with free radicals to terminate their radical chain reaction or donate electrons from the degraded polysaccharide’s free hydroxyl group to reduce the radical to a more stable form [124]. Studies have also shown that degraded polysaccharides have more hydroxyl groups than their native polysaccharide which is an indication of a more available reductive hydroxyl group, which could accept and eliminate the free radicals [40,125]. Importantly, an increase in concentration and lower molecular weight of degraded polysaccharides play a significant role in the radical scavenging capacity [104], thus higher antioxidant capacity.
  • Reducing power
The reducing power of a compound is a potential indicator of its antioxidant activity [40,126]. Degraded polysaccharides with low molecular weight have strong reducing power. The reducing power of a blackcurrant polysaccharide with an initial molecular weight of 3.26 × 104 KDa degraded to a molecular weight of 9.62 × 103 KDa was found to increase from 0.183 before degradation to 0.414 in the degraded polysaccharide [40]. Additionally, a positive correlation exists between the concentration of polysaccharides and reducing power. The reducing power of native polysaccharide (0.02) and degraded polysaccharide (0.3) at a concentration of 0.2 mg/mL increased strongly to 0.11 in native polysaccharide and 0.87 in degraded polysaccharide when the concentration was increased to 1.2 mg/mL. This implies that reducing power increases with a decrease in molecular weight and increasing concentration of polysaccharides. The reducing power of degraded polysaccharides comes from their ability to donate a hydrogen atom to destroy free radical chain reactions, thus exerting its antioxidant activity [40,104]. The increased reducing power in low molecular weight degraded polysaccharides is attributed to the higher number of reducing and non-reducing ends, compared to their native polysaccharide [109].
  • Metal ion chelating ability
Transition metals such as iron and copper are well known to enhance the production of free radicals which are precursors of oxidative stress [40]. They are regarded as stimulants/facilitators of lipid peroxidation (Fenton reaction) consequently yielding more reactive alkoxyl and peroxyl radicals [5]. Amongst the transition metals, iron is the most important pro-oxidant due to its high reactivity [5,127]. Besides being an essential mineral for normal body physiology, the overproduction (excess) of iron can result in cellular damage from oxidative stress induced by free radicals. As a result, reduction in the concentration of the catalyzing transition metal-mediated autoxidation through metal ion chelation is commonly considered as an antioxidant mechanism [40,128]. Polysaccharides can inhibit free radical production by chelating metal ions instead of scavenging them directly [123]. Further, Chun-hui et al. [129] isolated two polysaccharide fractions (GAPS-1 and SAPS-1) from A. barbadensis Miller that showed good metal ion chelating ability against ferrous ion. Similarly, a polysaccharide from the fern Lygodium japonicum was reported to also show strong metal ion chelating potential [130,131]. Polysaccharide’s metal ion chelating ability has been shown to serve as a secondary (preventive) antioxidant because of their redox potential reduction ability which stabilizes the oxidized form of the metal ions [132].
Furthermore, studies have shown that degraded polysaccharides exhibited significantly higher metal ion chelating ability on Fe2+ over the native polysaccharide [40], as the metal-chelating effects of the degraded polysaccharides depend on concentration. Polysaccharide with a molecular weight of 3.26 × 104 KDa degraded to 1.30 × 104 and 9.62 × 103 KDa was found to show an increased metal chelating ability from 5.0% before H2O2 degradation to 22.28% and 23.32 in the degraded polysaccharide respectively when concentration was increased from 0.2–1.2 mg/mL [40]. Specifically, the efficiency of polysaccharides in chelating metal ions is dependent on its structural configuration especially for functional groups of –OH, –SH, –COOH, –PO3H2, –C=O, –NR2, –S–, and –O– [123,133]. This was corroborated by Fan et al. [134] who confirmed that the chelating effect of degraded polysaccharides from leaves of Ilex latifolia Thunb. was partly due to the presence of functional groups such as the carboxyl group in the polysaccharide structure. Similarly, the presence of hydroxyl groups and the number of hydroxyl group on a polysaccharide structure have been found to influence the chelating affinity for the ferrous ion [123,135]. In line with polysaccharide’s ability to act as a metal ion chelator, its chelation mechanism is owed to complexes formed between the metal ion and aforementioned functional groups on the polysaccharide structure [136]. Polysaccharides having exclusively hydroxyl groups in equatorial position (for example, glucose molecule) do not seem to form complexes with metal ions, rather a solely favorable arrangement or configuration of two or three hydroxyl groups allows for the formation of metal ion complexes [137]. This implies that structural configuration and the arrangement of functional group (OH group) on a polysaccharide influences the formation of metal ion complex, which is the stable form of the oxidized metal ion.

4.3. The Reaction Mechanism Involving Polysaccharides’ Antioxidant Capacity on Free Radicals: A Primer

Polysaccharide has two mechanisms of degrading the OH• radical: (a) Removal of already generated OH• radicals through hydrogen electron transfer or single electron transfer [30,81]; and (b) inhibition of OH• radical formation [30,138]. Figure 7 shows the proposed reaction mechanism of OH• on a polysaccharide chain. Firstly, OH• radicals attack polysaccharides at different carbon positions and only a few of the polysaccharide backbones are influenced by these attacks, while oxo groups are simultaneously introduced into the polysaccharide chain, without altering the main chain. An OH• radical can abstract a hydrogen atom of the C-H chain of polysaccharides to form water, while on the other hand, a carbon-centred radical (carbon-free radical) is formed. In aerobic conditions, the carbon-free radical is oxidized to peroxy radicals and subsequently, reacts with hydroperoxyl radical (HO2•, often referred to as superoxide anion radical, O2•− in its ionized state) to produce ketone, lactone, or aldehyde [139,140].
The introduction of oxo group (reactions c4 and d4) is a result of polysaccharide chain scission caused by the elimination of hydrogen from C-1 or C-4 position on a pyranose ring. Conversely, when hydrogen is abstracted from the C-5 position, there is no occurrence of polysaccharide chain scission, rather a more labile ester is formed (reaction a4). Additionally, when hydrogen is abstracted from the C-3, C-2, or C-6 positions, there is also no scission of polysaccharide chain but a relatively stable oxo group creating a glycosulose residue would be formed (reaction b3) [140]. Another mechanism of polysaccharides to exhibit their antioxidant capacity is the ability to inhibit free radical formation by chelating transition metal ions, especially Fe2+ that catalyse the formation of highly reactive OH• radical from O2•− and H2O2 (Fenton reaction) [25,135,140].

4.4. Some Examples of Antioxidative Polysaccharides and Their Sources

Table 1 shows some examples of antioxidative polysaccharides and their sources. Out of 46 examples of antioxidative polysaccharides total presented herein, two (2) that include Hyriopsis cumingii and Misgurnus anguillicaudatus were of animal [25,141,142], four (4) that include Bacillus coagulans, Edwardsiella tarda, Paenibacillus polymyxa, and Streptomyces virginia were of bacterial [25,143,144,145,146,147,148,149,150,151], seven (7) that include Bifurcaria bifurcata, Enteromorpha linza, Fucus vesiculosus, Laminaria angustata, L. japonica, Padina gymnospora, and Phaeophyceae spp were of algae [25,36,152,153,154,155,156,157,158,159,160], fifteen (15) that include Astragalus mongholicus, Auricularia auricula, Azadirachta indica leaves, Chuanminshen violaceum, Dendrobium huoshanense, Litchi chinensis Sonn., Longan, Lycium barbarum, Persimmon, Portulaca oleracea L., pumpkin, Astragali radix, Ribes nigrum L. (blackcurrant fruits), Scutellaria barbata D. Don, and Tremella fuciformis were of plant [34,42,43,123,161,162,163,164,165,166,167,168,169,170,171,172,173,174,175,176,177,178,179,180,181,182,183,184], as well as eighteen (18) that include Athyrium multidentatum, Calocybe indica var, Cordyceps gunni, Coriolus versicolor, Dictyophora indusiata, Flammulina spp, Ganoderma atrum, G. lucidum, Grifola frondosa, Lachnum, Lentinus edodes, Lentinus polychrous, Pleurotus abalonus, P. cornucopiae, P. eryngii, P. florida, P. nebrodensis, and P. ostreatus were of fungal [15,25,38,44,117,131,185,186,187,188,189,190,191,192,193,194,195,196,197,198,199,200,201,202,203] origin.
From Table 2, we can see that the various sources, from the plant, animal, to bacteria and algae, gave various polysaccharide specificities. We can also see that within the same sources, aspects of chain-linkages here and there may be similar. Additionally, there could be some polysaccharide types in common. Nonetheless, several antioxidative polysaccharides discovered from plants, animals, fungi, bacteria, and algae sources [25] have been reported to exhibit antioxidant properties in in vitro and in vivo systems [144,204,205,206,207,208]. The antioxidative polysaccharides in these studies were found to possess reducing power, radical scavenging ability, and metal ion chelating property [25]. It is important to reiterate here that there are some other natural polysaccharides reported to exert biological and antioxidant effect that include, but are not limited to, white jelly mushroom (Tremella fuciformis) which is also known as ‘snow fungus’ [34], oyster mushroom (Pleurotus spp.) [209], blackcurrant (Ribes nigrum L.) [40], and polysaccharides from Auricularia auricular, Radix astragali, Ganoderma lucidum [34], Athyrium multidentatum, Scutellaria barbata D. Don, Astragalus mongholicus, Litchi chinensis Sonn., Lycium barbarum, Lentinus edodes and Cordyceps sinensis [25].

5. Concluding Remarks and Future Prospects

Polysaccharide degradation by the free radical-catalysed system offered by H2O2 is indeed effective in improving the bioactivity, especially the antioxidant activity. The current review has highlighted how H2O2 singly or in combination with other techniques can degrade natural polysaccharides. How the effects of polysaccharide degradation would influence the structural characteristics, physicochemical properties, and bioactivities (antioxidant capacity) of degraded polysaccharides has also been enumerated in this review. The free radical-catalysed polysaccharide degradation method offered by H2O2 is frequently employed because of its mild reaction conditions, high efficiency, inability to alter the integrity of the main chain structure of polysaccharides, and ease of OH• radical formation. Overall, the antioxidant capacity of polysaccharides is not dependent on a single factor but rather a combination of several related factors. Compared with the native polysaccharide, degraded polysaccharides exerts their antioxidant activities due to their low molecular weight, increased water solubility, decreased intrinsic viscosity, and modified structural characteristics.
Polysaccharide degradation comes with free radical scavenging capacity on OH• and O2•− radicals, transition metal chelating ability, and reducing power through their hydrogen atom donating ability. However, there is a need to further understand what happens before the degradation, specific to the level of purity of polysaccharide with antioxidant properties, and this should be the direction of future literature syntheses/experimental programs to supplement existing information, given that other antioxidant compounds such as polyphenols may form part of the coextractives retained in the polysaccharide. Degraded polysaccharides may find wide application in the food and beverage industries as novel functional ingredients. The direction of future studies should focus on how polysaccharides and other food components interact within the food or beverage matrix, as this may either change, enhance or inhibit the antioxidant potential of the polysaccharide. The information this overview provides should prove useful, not only to biotechnologists, food scientists, and technologists, but also processors, as well as product developers of novel and functional food formulations.

Author Contributions

Conceptualization, C.E.O., L.Y., A.O.A., G.B. and C.O.R.O.; methodology, C.M.O., J.O.I., N.O.K., M.O., J.S.C.., O.P.M., and C.E.O.; validation, J.O.I., N.O.K., M.K., J.S.C., O.P.M., A.O.A., and C.O.R.O.; formal analysis, N.O.K., G.S., I.M.A. and O.P.M..; investigation, C.E.O., L.Y., G.S., I.M.A., G.B., C.M.O., and M.O.; resources, C.O.R.O. and M.K.; data curation, C.M.O., C.E.O., M.K., J.O.I., J.S.C., and M.O.; visualization, L.Y., I.M.A., G.S., A.O.A., G.B., C.O.R.O. and M.K.; funding acquisition, C.O.R.O. and M.K.; writing—original draft preparation, C.E.O., L.Y., A.O.A., I.M.A., M.O., and J.S.C.; writing—review and editing, C.M.O., J.O.I., N.O.K., G.B., O.P.M., G.S.,C.O.R.O., and M.K. All authors have read and agreed to the published version of the manuscript.


Publication financed by the project UPWR 2.0: international and interdisciplinary programme of development of Wrocław University of Environmental and Life Sciences, co-financed by the European Social Fund under the Operational Program Knowledge Education Development, under contract No. POWR.03.05.00-00-Z062/18 of 4 June 2019.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data sharing is not applicable.


C.E.O.: J.C., A.O.A. and L.Y. acknowledge financial support from South China University of Technology, Guangdong, China. C.O.R.O. and M.K. acknowledge financial support from Wroclaw University of Environmental and Life Sciences, Poland.

Conflicts of Interest

All authors declare that they have no conflicts of interest involving this work.


  1. Morello, M.J.; Shahidi, F.; Ho, C.T. Free Radicals in Foods: Chemistry, Nutrition and Health Effects; ACS Symposium Series; American Chemical Society: Washington, DC, USA, 2002; pp. 1–9. [Google Scholar]
  2. Maddu, N. Diseases Related to Types of Free Radicals; Intech Open: London, UK, 2019; pp. 1–8. [Google Scholar]
  3. Turkan, I. ROS and RNS key signalling molecules in plants. J. Exp. Bot. 2018, 69, 3313–3315. [Google Scholar] [CrossRef] [Green Version]
  4. Messner, K.R.; Imlay, J.A. Mechanism of Superoxide and Hydrogen Peroxide Formation by Fumarate Reductase, Succinate Dehydrogenase, and Aspartate Oxidase; American Society for Biochemistry and Molecular Biology Inc.: Rockville, MD, USA, 2002. [Google Scholar]
  5. Gulcin, I. Antioxidants and Antioxidant Methods: An Updated Review. Archives of Toxicology; Springer: Berlin/Heidelberg, Germany, 2020. [Google Scholar]
  6. Patel, R.; Rinker, L.; Peng, J.; Chilian, W.M. Reactive Oxygen Species: The Good and the Bad. React. Oxyg. Species (ROS) Living Cells 2018, 7, 7–20. [Google Scholar]
  7. Phaniendra, A.; Jestadi, D.B.; Periyasamy, L. Free Radicals: Properties, Sources, Targets, and Their Implication in Various Diseases. Indian J. Clin. Biochem. 2015, 30, 11–26. [Google Scholar] [CrossRef] [Green Version]
  8. Genestra, M. Oxyl radicals, redox-sensitive signalling cascades and antioxidants. Cell. Signal. 2007, 19, 1807–1819. [Google Scholar] [CrossRef]
  9. Lee, J.Y.; Jang, Y.W.; Kang, H.S.; Moon, H.; Sim, S.S.; Kim, C.J. Anti-inflammatory action of phenolic compounds from Gastrodia elata root. Arch. Pharmacal Res. 2006, 29, 849–858. [Google Scholar] [CrossRef] [PubMed]
  10. Miller, E.W.; Albers, A.E.; Pralle, A.; Isacoff, E.Y.; Chang, C.J. Boronate-Based Fluorescent Probes for Imaging Cellular Hydrogen Peroxide. J. Am. Chem. Soc. 2005, 127, 16652–16659. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  11. Lee, L.; Hwang, O.; Yoo, D.; Khang, G.; Lee, D. Detection of hydrogen peroxide in vitro and in vivo using peroxalate chemilumiescent micelles. Bull. Korean Chem. Soc. 2011, 33, 2187–2192. [Google Scholar] [CrossRef] [Green Version]
  12. Elmarakby, A.A.; Pollock, D.M.; Imig, J.D. Renal dysfunction in hypertension and obesity. In Comprehensive Medicinal Chemistry II; Taylor, J.B., Triggle, D.J., Eds.; Elsevier Ltd.: Amsterdam, The Netherlands, 2007; Volume 6, pp. 575–595. [Google Scholar]
  13. Chatterjee, S. Oxidative Stress, Inflammation, and Disease. In Oxidative Stress and Biomaterials; Dziuba, T., Butterfield, A.D., Eds.; Elsevier Academic Press: Amsterdam, The Netherlands, 2016; pp. 35–58. ISBN 9780128032695. [Google Scholar]
  14. Lobo, V.; Patil, A.; Phatak, A.; Chandra, N. Free radicals, antioxidants and functional foods: Impact on human health. Pharmacogn. Rev. 2010, 4, 118–126. [Google Scholar] [CrossRef] [Green Version]
  15. Vamanu, E. Antioxidant properties of polysaccharides obtained by batch cultivation of Pleurotus ostreatus mycelium. Nat. Prod. Res. 2012, 27, 1115–1118. [Google Scholar] [CrossRef]
  16. Singh, A.; Kukreti, R.; Saso, L.; Kukreti, S. Oxidative Stress: A Key Modulator in Neurodegenerative Diseases. Molecules 2019, 24, 1583. [Google Scholar] [CrossRef] [Green Version]
  17. Lennicke, C.; Rahn, J.; Lichtenfels, R.; Wessjohann, J.A.; Seliger, B. Hydrogen peroxide—production, fate and role in redox signalling of tumour cells. Cell Commun. Signal. 2015, 13, 1–19. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  18. Armogida, M.; Nisticò, R.; Mercuri, N.B. Therapeutic potential of targeting hydrogen peroxide metabolism in the treatment of brain ischaemia. Br. J. Pharmacol. 2012, 166, 1211–1224. [Google Scholar] [CrossRef] [Green Version]
  19. Weinstain, R.; Savariar, E.N.; Felsen, C.N.; Tsien, R.Y. In Vivo Targeting of Hydrogen Peroxide by Activatable Cell-Penetrating Peptides. J. Am. Chem. Soc. 2014, 136, 874–877. [Google Scholar] [CrossRef] [PubMed]
  20. Halliwell, B.; Clement, M.V.; Long, L.H. Hydrogen peroxide in the human body. FEBS Lett. 2000, 486, 10–13. [Google Scholar] [CrossRef] [Green Version]
  21. Schieber, M.; Chandel, N.S. ROS Function in Redox Signaling and Oxidative Stress. Curr. Biol. 2014, 24, R453–R462. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  22. Holmström, K.M.; Finkel, T. Cellular mechanisms and physiological consequences of redox-dependent signalling. Nat. Rev. Mol. Cell Biol. 2014, 15, 411–421. [Google Scholar] [CrossRef]
  23. Park, H.; Kim, S.; Song, Y.; Seung, K.; Hong, D.; Khang, G.; Lee, D. Antioxidant and anti-inflammatory activities of hydroxybenzyl alcohol releasing biodegradable polyoxalate nanoparticles. Biomacromolecules 2010, 11, 2103–2108. [Google Scholar] [CrossRef] [PubMed]
  24. Collin, F. Chemical Basis of Reactive Oxygen Species Reactivity and Involvement in Neurodegenerative Diseases. Int. J. Mol. Sci. 2019, 20, 2407. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  25. Wang, H.; Lui, Y.M.; Qi, Z.M.; Wang, S.Y.; Liu, S.X.; Li, X.; Wang, H.J.; Xia, X.C. An overview on natural polysaccharides with antioxidant properties. Curr. Med. Chem. 2013, 20, 2899–2913. [Google Scholar] [CrossRef]
  26. Erickson, M.C. Lipid oxidation in muscle foods. In Food Lipids Chemistry, Nutrition and Biotechnology; Akoh, C.C., Min, D.B., Eds.; CRC Press: New York, NY, USA, 2003; pp. 365–411. [Google Scholar]
  27. Bensid, A.; El Abed, N.; Houicher, A.; Regenstein, J.M.; Özogul, F. Antioxidant and antimicrobial preservatives: Properties, mechanism of action and applications in food—A review. Crit. Rev. Food Sci. Nutr. 2020, 1–17. [Google Scholar] [CrossRef] [PubMed]
  28. Jamila, V.; Vavrikova, E. Chitosan derivatives with antimicrobial, antitumor and anti-oxidant activities: A review. Curr. Pharm. Design. 2011, 17, 3596–3607. [Google Scholar]
  29. Yang, L.; Zhang, L.M. Chemical structural and chain conformational characterization of some bioactive polysaccharides isolated from natural sources. Carbohydr. Polym. 2009, 76, 349–361. [Google Scholar] [CrossRef]
  30. Wang, N.; Zhang, X.; Wang, S.; Guo, Q.; Li, Z.; Liu, H.; Wang, C. Structural characterisation and immunomodulatory activity of polysaccharides from white asparagus skin. Carbohydr. Polym. 2020, 227, 115314. [Google Scholar] [CrossRef] [PubMed]
  31. Wang, H.; Gao, X.D.; Zhou, G.C.; Cai, L.; Yao, W.B. In vitro and in vivo antioxidant activity of aqueous extract from Choerospondias axillaris fruit. Food Chem. 2008, 106, 888–895. [Google Scholar] [CrossRef]
  32. Zhang, Y.; Dai, L.; Kong, X.; Chen, L. Characterization and in vitro antioxidant activities of polysaccharides from Pleurotus ostreatus. Int. J. Biol. Macromol. 2012, 51, 259–265. [Google Scholar] [CrossRef]
  33. Fan, L.; Ding, S.; Ai, L.; Deng, K. Antitumor and immunomodulatory activity of water-soluble polysaccharide from Inonotus obliquus. Carbohydr. Polym. 2012, 90, 870–874. [Google Scholar] [CrossRef]
  34. Zhang, Z.; Wang, X.; Zhao, M.; Qi, H. Free-radical degradation by Fe2+/Vc/H2O2 and antioxidant activity of polysaccharide from Tremella fuciformis. Carbohydr. Polym. 2014, 112, 578–582. [Google Scholar] [CrossRef] [PubMed]
  35. Jhamandas, J.H.; Wie, M.B.; Harris, K.; MacTavish, D.; Kar, S. Fucoidaninhibits cellular and neurotoxic effects of β-amyloid (Aβ) in rat cholinergic basalforebrain neurons. Eur. J. Neurosci. 2005, 21, 2649–2659. [Google Scholar] [CrossRef]
  36. Zhang, N.; Chen, H.; Ma, L.; Zhang, Y. Physical modifications of polysaccharide from Inonotus obliquus and the antioxidant properties. Int. J. Biol. Macromol. 2013, 54, 209–215. [Google Scholar] [CrossRef]
  37. Zhang, Z.S.; Wang, X.M.; Mo, X.F.; Qi, H.M. Degradation and the antioxidant activity of polysaccharide from Enteromorpha linza. Carbohydr. Polym. 2013, 92, 2084–2087. [Google Scholar] [CrossRef]
  38. Jing, L.; Sheng, J.; Jiang, J.; Wang, Y.; Shen, X.; Liu, D.; Zhang, W.; Mao, S. Chemical characteristics and cytoprotective activities of polysaccharide fractions from Athyrium Multidentatum (Doll.) Ching. Int. J. Biol. Macromol. 2020, 158, 1227–1238. [Google Scholar] [CrossRef]
  39. Sun, L.; Wang, L.; Li, J.; Liu, H. Characterization and antioxidant activities of degraded polysaccharides from two marine Chrysophyta. Food Chem. 2014, 160, 1–7. [Google Scholar] [CrossRef]
  40. Xu, Y.; Niu, X.; Liu, N.; Gao, Y.; Wang, L.; Xu, G.; Li, X.; Yang, Y. Characterisation, antioxidant and hypoglycemic activities of degraded polysaccharides from black currant (Ribes nigrum L.) fruits. Food Chem. 2017, 243, 26–35. [Google Scholar] [CrossRef] [PubMed]
  41. Wu, H.; Zhu, H.; Li, X.; Liu, Z.; Zheng, W.; Chen, T.; Yu, B.; Wong, K.-H. Induction of Apoptosis and cell cycle arrest in A549 Human Lung Adernocarcinoma cells by Surface-Capping Selenium Nanoparticles: An Effect enhanced by Polysachharide-Protein Complexes from Polyporus rhinoceros. J. Agric. Food Chem. 2013, 61, 9859–9866. [Google Scholar] [CrossRef]
  42. Shen, H.; Tang, G.; Zeng, G.; Yang, Y.; Cai, X.; Li, D.; Liu, H.; Zhou, N. Purification and characterisation of an antitumor polysaccharide from Portulaca oleracea L. Carbohydr. Polym. 2013, 93, 395–400. [Google Scholar] [CrossRef]
  43. Saha, S.; Galhardi, L.C.; Yamamoto, K.A.; Linhares, R.E.C.; Bandyopadhyay, S.S.; Sinha, S.; Nozawa, C.; Ray, B. Water-extracted polysaccharides from Azadirachta indica leaves: Structural features, chemical modification and anti-bovine herpesvirus type 1 (BoHV-1) activity. Int. J. Biol. Macromol. 2010, 47, 640–645. [Google Scholar] [CrossRef]
  44. Zhu, Z.-Y.; Dong, F.; Liu, X.; Lv, Q.; Yang, Y.; Liu, F.; Chen, L.; Wang, T.; Wang, Z.; Zhang, Y. Effects of extraction methods on the yield, chemical structure and anti-tumor activity of polysaccharides from Cordyceps gunnii mycelia. Carbohydr. Polym. 2016, 140, 461–471. [Google Scholar] [CrossRef] [PubMed]
  45. Sies, H. Hydrogen peroxide as a central redox signaling molecule in physiological oxidative stress: Oxidative eustress. Redox Biol. 2017, 11, 613–619. [Google Scholar] [CrossRef] [PubMed]
  46. Duan, J.; Kasper, D.L. Oxidative depolymerisation of polysaccharides by reactive oxygen/nitrogen species. Glycobiology 2011, 21, 401–409. [Google Scholar] [CrossRef]
  47. Pagano, G.; Talamanca, A.A.; Castello, G.; Cordero, M.D.; Dischia, M.; Gadaleta, M.N. Oxidative stress and mitochondrial dysfunction across broad-ranging pathologies: Toward mitochondria-targeted clinical strategies. Oxidative Med. Cell. Longev. 2014, 54, 12–30. [Google Scholar] [CrossRef] [PubMed]
  48. Lv, W.; Booz, G.W.; Fan, F.; Wang, Y.; Roman, R.J. Oxidative stress and renal fibrosis: Recent insights for the development of novel therapeutic strategies. Front. Physiol. 2018, 9, 105. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  49. Zhao, J.; Wang, S.; Zhong, W.; Yang, B.; Sun, L.; Zheng, Y. Oxidative stress in the trabecular meshwork (Review). Int. J. Mol. Med. 2016, 38, 995–1002. [Google Scholar] [CrossRef] [Green Version]
  50. Electron Transport Chain, Lumen, Biology for Majors. Available online: (accessed on 23 December 2020).
  51. Fenton, H.J.H. Oxidation of tartaric acid in the presence of iron. J. Chem. Soc. Trans. 1894, 65, 899–910. [Google Scholar] [CrossRef] [Green Version]
  52. Halliwell, B.; Gutteridge, J.M.C. Free Radicals in Biology and Medicine, 2nd ed.; Clarendon Press: Oxford, UK, 1989; pp. 1–3. [Google Scholar]
  53. Kehrer, J.P. The Haber-Weiss reaction and mechanisms of toxicity. Toxicology 2000, 149, 43–50. [Google Scholar] [CrossRef]
  54. Anraku, M.; Gebicki, J.M.; Iohara, D.; Tomida, H.; Uekama, K.; Maruyama, T.; Hirayama, F.; Otagiri, M. Antioxidant activities of chitosans and its derivatives in in vitro and in vivo studies. Carbohydr. Polym. 2018, 199, 141–149. [Google Scholar] [CrossRef] [PubMed]
  55. Sindhi, V.; Gupta, V.; Sharma, K.; Bhatnagar, S.; Kumari, R.; Dhaka, N. Potential applications of antioxidants—A review. J. Pharm. Res. 2013, 7, 828–835. [Google Scholar] [CrossRef]
  56. Sharma, P.; Jha, A.B.; Dubey, R.S.; Pessarakli, M. Reactive Oxygen Species, Oxidative Damage, and Antioxidative Defense Mechanism in Plants under Stressful Conditions. J. Bot. 2012, 2012, 217037. [Google Scholar] [CrossRef] [Green Version]
  57. Nakagawa, H.; Wachi, M.; Woo, J.-T.; Kato, M.; Kasai, S.; Takahashi, F.; Lee, I.-S.; Nagai, K. Fenton Reaction Is Primarily Involved in a Mechanism of (−)-Epigallocatechin-3-gallate to Induce Osteoclastic Cell Death. Biochem. Biophys. Res. Commun. 2002, 292, 94–101. [Google Scholar] [CrossRef]
  58. Muller, R. The formation of hydrogen peroxide during oxidation of thiol-containing proteins. J. Inst. Brew. 1997, 103, 307–310. [Google Scholar] [CrossRef]
  59. Bopitiya, D.; Christensen, D.; Martin, M.; Zhang, J.; Bennett, L.E. Production of hydrogen peroxide in formulated beverages is associated with the presence of ascorbic acid combined with selected redox-active functional ingredients. Food Chem. 2021, 338, 127947. [Google Scholar] [CrossRef]
  60. Long, L.L.; Lan, A.N.B.; Hsuan, F.L.Y.; Halliwell, B. Generation of hydrogen peroxide by “Antioxidant” beverages and the effect of milk addition. Is cocoa the best beverage? Free Radic. Res. 1999, 31, 67–71. [Google Scholar] [CrossRef] [PubMed]
  61. Nakayama, T.; Ichiba, M.; Kuwabara, M.; Kujiya, K.; Kumazawa, S. Mechanisms and structural specificity of hydrogen peroxide formation during oxidation of catechins. Food Sci. Technol. Res. 2002, 8, 261–267. [Google Scholar] [CrossRef] [Green Version]
  62. Akagawa, M.; Shigemitsu, T.; Suyama, K. Production of Hydrogen Peroxide by Polyphenols and Polyphenol-rich Beverages under Quasi-physiological Conditions. Biosci. Biotechnol. Biochem. 2003, 67, 2632–2640. [Google Scholar] [CrossRef]
  63. Grzesik, M.; Bartosz, G.; Stefaniuk, I.; Pichla, M.; Namieśnik, J.; Sadowska-Bartosz, I. Dietary antioxidants as a source of hydrogen peroxide. Food Chem. 2019, 278, 692–699. [Google Scholar] [CrossRef]
  64. Nakagawa, T.; Yokozawa, T. Direct scavenging of nitric oxide and superoxide by green tea. Food Chem. Toxicol. 2002, 40, 1745–1750. [Google Scholar] [CrossRef]
  65. Kim, H.-S.; Quon, M.J.; Kim, J.-A. New insights into the mechanisms of polyphenols beyond antioxidant properties; lessons from the green tea polyphenol, epigallocatechin 3-gallate. Redox Biol. 2014, 2, 187–195. [Google Scholar] [CrossRef] [Green Version]
  66. Nakayama, T.; Enoki, Y.; Hashimoto, K. Hydrogen Peroxide Formation during Catechin Oxidation Is Inhibited by Superoxide Dismutase. Food Sci. Technol. Int. Tokyo 1995, 1, 65–69. [Google Scholar] [CrossRef] [Green Version]
  67. Wee, L.M.; Long, L.H.; Whiteman, M.; Halliwell, B. Factors affecting the ascorbate- and phenolic-dependent generation of hydrogen peroxide in Dulbecco’s Modified Eagles Medium. Free Radic. Res. 2003, 37, 1123–1130. [Google Scholar] [CrossRef] [PubMed]
  68. Halliwell, B.; Clement, M.V.; Ramalingam, J.; Long, L.H. Hydrogen Peroxide. Ubiquitous in cell culture and in vivo? Life 2000, 50, 251–257. [Google Scholar]
  69. Halliwell, B.; Gutteridge, J. Free Radicals in Biology and Medicine, 3rd ed.; Oxford University Press: Oxford, UK, 1999; p. 936. [Google Scholar]
  70. Kwong, L.K.; Sohal, R.S. Substrate and site specificity of hydrogen peroxide generation in mouse mitochondria. Arch. Biochem. Biophys. 1998, 350, 118–126. [Google Scholar] [CrossRef]
  71. Braidot, E.; Petrissa, E.; Vicunello, A.; Macri, F. Hydrogen peroxide generation by higher plant mitochondria oxidizing complex I or complex II substrates. FEBS Lett. 1999, 451, 347–350. [Google Scholar] [CrossRef] [Green Version]
  72. Lambert, J.D.; Kwon, S.J.; Hong, J.; Yang, C.S. Salivary hydrogen peroxide produced by holding or chewing green tea in the oral cavity. Free Radic. Res. 2007, 41, 850–853. [Google Scholar] [CrossRef]
  73. Adahi, T.; Yamada, H.; Hara, H.; Futenma, A.; Kakumu, S. Increase of urinary EC-SOD level correlated with cAMP. FEBS Lett. 1999, 458, 370–374. [Google Scholar] [CrossRef] [Green Version]
  74. Spector, A.; Ma, W.; Wang, R.R. The aqueous humor is capable of generating and degrading H2O2. Investig. Ophthalmol. Vis. Sci. 1998, 39, 1188–1197. [Google Scholar]
  75. Chen, Y.; Mehta, G.; Vasiliou, V. Antioxidant defences in the ocular surface. Ocul. Surf. 2009, 7, 176–185. [Google Scholar] [CrossRef] [Green Version]
  76. Wielgus, A.R.; Sarna, T. Ascorbate enhances photogeneration of hydrogen peroxide mediated by the Iris melanin. Photochem. Photobiol. 2008, 84, 683–691. [Google Scholar] [CrossRef]
  77. Long, L.H.; Clément, M.V.; Halliwell, B. Artifacts in Cell Culture: Rapid Generation of Hydrogen Peroxide on Addition of (−)-Epigallocatechin, (−)-Epigallocatechin Gallate, (+)-Catechin, and Quercetin to Commonly Used Cell Culture Media. Biochem. Biophys. Res. Commun. 2000, 273, 50–53. [Google Scholar] [CrossRef] [PubMed]
  78. Akagawa, M.; Shigemitsu, T.; Suyama, K. Oxidative deamination of Benzylamine and Lysine residue in bovine serum albumin by green tea, black tea, and coffee. J. Agric. Food Chem. 2005, 53, 8019–8024. [Google Scholar] [CrossRef]
  79. Hoshino, N.; Kimura, T.; Yamaji, A.; Ando, T. Damage to the cytoplasmic membrane of Escherichia coli by catechin-copper (II) complexes. Free Radic. Biol. Med. 1999, 27, 1245–1250. [Google Scholar] [CrossRef]
  80. Hiramoto, K.; Kida, T.; Kikugawa, K. Increased urinary hydrogen peroxide levels caused by coŠee drinking. Biol. Pharm. Bull. 2002, 25, 1467–1471. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  81. Santos-Sánchez, N.F.; Salas-Coronado, R.; Villanueva-Cañongo, C.; Hernández-Carlos, B. Antioxidant Compounds and Their Antioxidant Mechanism. In Antioxidants; Shalaby, E., Ed.; IntechOpen: London, UK, 2019; pp. 1–28. [Google Scholar]
  82. Davies, K.J. Oxidative stress: The paradox of aerobic life. Biochem. Soc. Symp. 1995, 61, 1–31. [Google Scholar] [CrossRef]
  83. Mochizuki, M.; Yamazaki, S.; Kano, K.; Ikeda, T. Kinetic analysis and mechanistic aspects of autoxidation of catechins. Biochim. Biophys. Acta 2002, 1569, 35–44. [Google Scholar] [CrossRef]
  84. Rinaldi, A.; Porcu, M.C.G.; Curreli, N.; Rescigno, A.; Finazziagro, A.; Pedersen, J.; Rinaldi, A.; Sanjust, E. Autoxidation of 4-Methylcatechol: A Model for the Study of the Biosynthesis of Copper Amine Oxidases Quinonoid Cofactor. Biochem. Biophys. Res. Commun. 1995, 214, 559–567. [Google Scholar] [CrossRef] [PubMed]
  85. Adányi, N.; Barna, T.; Emri, T.; Miseki, M.; Pocsi, I. Hydrogen peroxide producing and decomposing enzymes: Their use in biosensors and other applications. In Industrial Enzymes; Polaina, J., MacCabe, A.P., Eds.; Springer: Dordrecht, The Netherlands, 2007; pp. 441–459. ISBN 978-1-4020-5376-4. [Google Scholar]
  86. Baveris, A.; Cadenas, E. Mitochondrial production of hydrogen peroxide regulation by nitric oxide and the role of ubisemiquinone. IUBMB Life 2000, 50, 245–250. [Google Scholar] [CrossRef]
  87. Wong, H.-S.; Benoit, B.; Brand, M.D. Mitochondrial and cytosolic sources of hydrogen peroxide in resting C2C12 myoblasts. Free Radic. Biol. Med. 2019, 130, 140–150. [Google Scholar] [CrossRef]
  88. Treberg, J.R.; Braun, K.; Selseleh, P. Mitochondria can act as energy-sensing regulators of hydrogen peroxide availability. Redox Biol. 2019, 20, 484–488. [Google Scholar] [CrossRef]
  89. Druck, T.; Cheung, D.G.; Park, D.; Trapasso, F.; Pichiorri, F.; Gaspari, M.; Palumbo, T.; Aqeilan, R.I.; Gaudio, E.; Okumura, H.; et al. Fhit–Fdxr interaction in the mitochondria: Modulation of reactive oxygen species generation and apoptosis in cancer cells. Cell Death Dis. 2019, 10, 1–10. [Google Scholar] [CrossRef] [PubMed]
  90. Bottje, W.G. Oxidative metabolism and efficiceny; the delicate balancing act of mitochondria. Poult. Sci. 2019, 98, 4223–4230. [Google Scholar] [CrossRef]
  91. Fang, J.; Wong, H.-S.; Brand, M.D. Production of superoxide and hydrogen peroxide in the mitochondrial matrix is dominated by site IQ of complex I in diverse cell lines. Redox Biol. 2020, 37, 101722. [Google Scholar] [CrossRef]
  92. Theng, B.K.G. Polysaccharides. In Formation and Properties of Clay-Polymer Complexes; Theng, B.K.G., Ed.; Elsevier Science: Amsterdam, The Netherlands, 2012; Volume 4, pp. 351–390. [Google Scholar]
  93. Udayan, A.; Arumugam, M.; Pandey, A. Nutraceuticals from algae and cynobacteria. In Algal Green Chemistry—Recent Progress in Biotechnology; Rastogi, R.P., Madamwav, D., Pandey, A., Eds.; Elsevier: Amsterdam, The Netherlands, 2017; pp. 65–89. [Google Scholar]
  94. Li, S.; Xiong, Q.; Lai, X.; Li, X.; Wan, M.; Zhang, J.; Yan, Y.; Cao, M.; Lu, L.; Guan, J.; et al. Molecular Modification of Polysaccharides and Resulting Bioactivities. Compr. Rev. Food Sci. Food Saf. 2015, 15, 237–250. [Google Scholar] [CrossRef] [Green Version]
  95. Garcia-Gonzalez, C.A.; Alnaief, M.; Smirnova, I. Polysaccharide-based aerogels—Promising biodegradable carriers for drug delivery systems-Review Article. Carbohydr. Polym. 2011, 86, 1425–1438. [Google Scholar] [CrossRef]
  96. Huang, D.; Li, Y.; Cui, F.; Chen, J.; Sun, J. Purification and characterization of a novel polysaccharide-peptide complex from Clinacanthus nutans Lindau leaves. Carbohydr. Polym. 2016, 137, 701–708. [Google Scholar] [CrossRef]
  97. Tian, F.; Liu, Y.; Hu, K.; Zhao, B. The depolymerisation mechanism of chitosan by hydrogen peroxide. J. Mater. Sci. 2003, 38, 4709–4712. [Google Scholar] [CrossRef]
  98. Zhang, Z.S.; Wang, X.M.; Liu, C.B.; Li, J.F. The degradation, antioxidant and antimutagenic activity of the mucilage polysaccharide from Dioscorea opposite. Carbohydr. Polym. 2016, 150, 227–231. [Google Scholar] [CrossRef]
  99. Gao, G.; Jiao, Q.; Li, X. Structural modification of pachymaran and the mechanism interaction between Azur A. Acta Laser. Biol. Sin. 2005, 14, 228–232. [Google Scholar]
  100. Li, S.Q.; Shah, N.P. Antioxidant and antibacterial activities of sulphated polysaccharides from Pleurotus eryngii and Streptococcus thermophilus ASCC 1275. Food Chem. 2014, 165, 262–270. [Google Scholar] [CrossRef] [PubMed]
  101. Alban, S.; Franz, G. Characterization of the anticoagulant actions of a semisynthetic curdlan sulfate. Thromb. Res. 2000, 99, 377–388. [Google Scholar] [CrossRef]
  102. Yalpani, M. Chemistry of polysaccharide modification and degradation. In Carbohydrates: Structures, Syntheses and Dynamics; Finch, P., Ed.; Springer: Dordrecht, The Netherlands, 1999; pp. 294–318. [Google Scholar]
  103. Cardoso, M.J.; Caridade, S.G.; Costa, R.R.; Mano, J.F. Enzymatic degradation of polysaccharide-based layer-by-layer structures. Biomacromolecules 2016, 17, 1347–1357. [Google Scholar] [CrossRef] [PubMed]
  104. Wu, J.; Li, P.; Zhao, H.; Sun, R.; Ma, F.; Zhang, B. Effect of solution plasma process with hydrogen peroxide on the degradation and antioxidant activity of polysaccharides from Auricularia acuricula. Int. J. Biol. Macromol. 2018, 117, 1299–1304. [Google Scholar] [CrossRef]
  105. Ren, Y.L.; Zheng, G.Q.; You, L.J.; Wen, L.R.; Li, C.; Fu, X. Structural characterization and macrophage immunomodulatory activity of a polysaccharide isolated from Gracilaria lemaneiformis. J. Funct. Foods 2017, 33, 286–296. [Google Scholar] [CrossRef]
  106. Chen, X.; Zhange, R.; Li, Y.; You, L.; Kulikouskaya, V.; Hileuskaya, K. Degradation of polysaccharides from Sargassum fusiforme using UV/H2O2 and its effects on structural characteristics. Carbohydr. Polym. 2019. [Google Scholar] [CrossRef] [PubMed]
  107. Chen, S.; Liu, H.; Yang, X.; Li, L.; Qi, B.; Hu, X.; Ma, H.; Li, C.; Pan, C. Degradation of sulphated polysaccharides from Grateloupia livida and antioxidant activity of the degraded components. Int. J. Biol. Macromol. 2020, 156, 660–668. [Google Scholar] [CrossRef]
  108. Yang, H.; Bai, J.; Ma, C.; Wang, L.; Li, X.; Zhang, Y.; Xu, Y.; Yang, Y. Degradation models, structure, rheological properties and protective effects on erythrocyte hemolysis of the polysaccharides from Ribes nigrum L. Int. J. Biol. Macromol. 2020, 165, 738–746. [Google Scholar] [CrossRef]
  109. Hou, Y.; Wang, J.; Jin, W.H.; Zhang, H.; Zhang, Q.B. Degradation of Laminaria japonica fucoidan by hydrogen peroxide and antioxidant activities of the degradation products of different molecular weights. Carbohydr. Polym. 2012, 87, 153–159. [Google Scholar] [CrossRef]
  110. Huang, Q.Z.; Wang, S.M.; Wang, Q.S.; Huang, J.F. Study on degradation of chitosan with UV/H2O2. J. Fujian Norm. Univ. 2004, 20, 63–67. [Google Scholar]
  111. Qin, C.Q.; Du, Y.M.; Liu, Y.; Xiao, L. Antibacterial activity of degraded chitosan by H2O2/HAc. J. Xiaogan Univ. 2001, 21, 9–13. [Google Scholar]
  112. Li, M.; Ma, F.; Li, R.; Ren, G.; Yan, D.; Zhang, H.; Zhu, X.; Wu, R.; Wu, J. Degradation of Tremella fuciformis polysaccharide by a combined ultrasound and hydrogen peroxide treatment: Process paramters, structural characteristics, and antioxidant activities. Int. J. Biol. Macromol. 2020, 160, 979–990. [Google Scholar] [CrossRef]
  113. Ayanda, O.S.; Nelana, S.M.; Naidoo, E.B. Ultrasonic degradation of aqueous phenolsulfonphthalein (PSP) in the presence of nano-Fe/H2O2. Ultrason. Sonochem. 2018, 47, 29–35. [Google Scholar] [CrossRef]
  114. Carbinatto, F.M.; De Castro, A.D.; Cury, B.S.; Magalhães, A.; Evangelista, R.C. Physical properties of pectin–high amylose starch mixtures cross-linked with sodium trimetaphosphate. Int. J. Pharm. 2012, 423, 281–288. [Google Scholar] [CrossRef] [Green Version]
  115. Tuhin, M.O.; Rahman, N.; Haque, M.E.; Khan, R.A.; Dafader, N.C.; Islam, R.; Nurnabi, M.; Tonny, W. Modification of mechanical and thermal property of chitosan-starch blend films. Radiat. Phys. Chem. 2012, 81, 1659–1668. [Google Scholar] [CrossRef]
  116. BeMiller, J.N. Modified Starch. In Encyclopedia of Food Science and Nutrition, 2nd ed.; Caballero, B., Trugo, L.C., Finglas, P.M., Eds.; Elsevier Academic Press: Amsterdam, The Netherlands, 2003; Volume 9, pp. 5576–5579. [Google Scholar]
  117. Bae, I.Y.; Kim, H.Y.; Lee, S.; Lee, H.G. Effect of the degree of oxidation on the physicochemical and biological properties of Grifola frondosa polysaccharides. Carbohydr. Polym. 2011, 83, 1298–1302. [Google Scholar] [CrossRef]
  118. Tian, Y.; Zeng, H.; Xu, Z.; Zheng, B.; Lin, Y.; Gan, C.; Lo, Y.M. Ultrasonic-assisted extraction and antioxidant activity of polysaccharides recovered from white button mushroom (Agaricus bisporus). Carbohydr. Polym. 2012, 2, 522–529. [Google Scholar] [CrossRef]
  119. Zhao, Z.; Li, J.; Wu, X.; Dai, H.; Gao, X.; Liu, M.; Tu, P. Structures and immunological activities of two pectic polysaccharides from the fruits of Ziziphus jujuba Mill. cv. jinsixiaozao Hort. Food Res. Int. 2006, 39, 917–923. [Google Scholar] [CrossRef]
  120. Chang, S.; Hsu, B.; Chen, B. Structural characterization of polysaccharides from Zizyphus jujuba and evaluation of antioxidant activity. Int. J. Biol. Macromol. 2010, 47, 445–453. [Google Scholar] [CrossRef] [PubMed]
  121. Li, C.; Huang, Q.; Fu, X.; Yue, X.-J.; Liu, R.H.; You, L. Characterization, antioxidant and immunomodulatory activities of polysaccharides from Prunella vulgaris Linn. Int. J. Biol. Macromol. 2015, 75, 298–305. [Google Scholar] [CrossRef]
  122. Hecht, F.; Pessoa, C.F.; Gentile, L.B.; Rosenthal, D.; Carvalho, D.P.; Fortunato, R.S. The role of oxidative stress on breast cancer development and therapy. Tumor Biol. 2016, 37, 4281–4291. [Google Scholar] [CrossRef]
  123. Wang, X.; Zhang, Z.; Zhao, M. Carboxymethylation of polysaccharides from Tremella fuciformis for antioxidant and moisture-preserving activities. Int. J. Biol. Macromol. 2015, 72, 526–530. [Google Scholar] [CrossRef]
  124. Lindsay, R.C. Food additives. In Food Chemistry; Fennema, O.R., Ed.; Marcel Dekker Inc.: New York, NY, USA, 1996; pp. 778–780. [Google Scholar]
  125. Yan, J.K.; Li, L.; Weng, Z.M.; Leung, P.H.; Wang, W.Q.; Wu, J.Y. Acidic degradation and enhanced antioxidant activities of exopolysaccharides from Cordyceps sinensis mycelial culture. Food Chem. 2009, 117, 641–646. [Google Scholar] [CrossRef]
  126. You, Q.H.; Yin, X.L.; Zhang, S.N.; Jiang, Z.H. Extraction, purification, and antioxidant activities of polysaccharides from Tricholoma mongolicum Imai. Carbohydr. Polym. 2014, 99, 1–10. [Google Scholar] [CrossRef]
  127. Gulcin, I. Antioxidant activity of food constituents: An overview. Arch. Toxicol. 2012, 86, 345–391. [Google Scholar] [CrossRef]
  128. Chen, H.; Ju, Y.; Li, J.; Yu, M. Antioxidant activities of polysaccharides from Lentinus edodes and their significance for disease prevention. Int. J. Biol. Macromol. 2012, 50, 214–218. [Google Scholar] [CrossRef] [PubMed]
  129. Chun-hui, L.; Chang-hai, W.; Xu, Z.-L.; Wang, Y. Isolation, chemical characterization and antioxidant activities of two polysaccharides from the gel and the skin of Aloe barbadensis Miller irrigated with sea water. Process Biochem. 2007, 42, 961–970. [Google Scholar] [CrossRef]
  130. Li, X.; Zhou, A.; Han, Y. Anti-oxidation and anti-microorganism activities of purification polysaccharide from Lygodium japonicum in vitro. Carbohydr. Polym. 2006, 66, 34–42. [Google Scholar] [CrossRef]
  131. He, X.; Wang, X.; Fang, J.; Chang, Y.; Ning, N.; Guo, H.; Huang, L.; Huang, X.; Zhao, Z. Polysaccharides in Grifola frondosa mushroom and their health promoting properties: A review. Int. J. Biol. Macromol. 2017, 101, 910–921. [Google Scholar] [CrossRef] [PubMed]
  132. Subhapradha, N.; Ramasamy, P.; Seedevi, P.; Shanmugam, V.; Srinivasan, A.; Shamugam, A. Extraction, characterization and its antioxidant efficacy of polysaccharides from Sepia aculeate (Orbigny, 1848) cuttlebone. Afr. J. Biotechnol. 2013, 13, 138–144. [Google Scholar]
  133. Gulcin, I. Antioxidant and antiradical activities of L-carnitine. Life Sci. 2006, 78, 803–811. [Google Scholar] [CrossRef]
  134. Fan, J.; Wu, Z.; Zhao, T.; Sun, Y.; Ye, H.; Xu, R.; Zeng, X. Characterization, antioxidant and hepatoprotective activities of polysaccharides from Ilex latifolia Thunb. Carbohydr. Polym. 2014, 101, 990–997. [Google Scholar] [CrossRef] [PubMed]
  135. Wang, J.; Zhang, Q.; Zhang, Z.; Li, Z. Antioxidant activity of sulfated polysaccharide fractions extracted from Laminaria japonica. Int. J. Biol. Macromol. 2008, 42, 127–132. [Google Scholar] [CrossRef]
  136. Ghosh, N.; Chakraborty, T.; Mallick, S.; Mana, S.; Singha, D.; Ghosh, B.; Roy, S. Synthesis, characterization and study of antioxidant activity of quercetin–magnesium complex. Spectrochim. Acta Part A Mol. Biomol. Spectrosc. 2015, 151, 807–813. [Google Scholar] [CrossRef]
  137. Joly, N.; Ghemati, D.; Aliouche, D.; Martin, P. Interaction of metal ions with mono- and polysaccharides for wastewater treatment: A Review. Nat. Prod. Chem. Res. 2020, 8, 1–17. [Google Scholar]
  138. Li, B.; Liu, S.; Xing, R.; Li, K.; Li, R.; Qin, Y.; Wang, X.; Wei, Z.; Li, P. Degradation of sulfated polysaccharides from Enteromorpha prolifera and their antioxidant activities. Carbohydr. Polym. 2013, 92, 1991–1996. [Google Scholar] [CrossRef] [PubMed]
  139. Lindsay, S.E.; Fry, S.C. Redox and wall-restructuring. In The Expanding Cell; Verbelen, J.-P., Vissenberg, K., Eds.; Springer: Berlin, Germany, 2007; pp. 159–190. [Google Scholar]
  140. Vreeburg, R.A.M.; Aririanah, O.B.; Fry, S.C. Fingerprinting of hydroxyl radical-attacked polysaccharides by N-isopropyl-2-aminoacridone labelling. Biochem. J. 2014, 463, 225–237. [Google Scholar] [CrossRef] [Green Version]
  141. Qiao, D.; Ke, C.; Hu, B.; Luo, J.; Ye, H.; Sun, Y.; Yan, X.; Zeng, X. Antioxidant activities of polysaccharides from Hyriopsis cumingii. Carbohydr. Polym. 2010, 78, 199–204. [Google Scholar] [CrossRef]
  142. Qin, C.; Huang, K.; Xu, H. Isolation and characterisation of a novel polysaccharide from the mucus of the loach, Misgurnus anguillicaudatus. Carbohydr. Polym. 2002, 49, 367–371. [Google Scholar] [CrossRef]
  143. Daud, N.S.; Din, A.R.J.M.; Rosli, M.A.; Azam, Z.M.; Othman, N.Z.; Sarmidi, M.R. Paenibacillus polymyxa bioactive compounds for agricultural and biotechnological applications. Biocatal. Agric. Biotechnol. 2019, 18, 101092. [Google Scholar] [CrossRef]
  144. Guo, S.; Mao, W.; Han, Y.; Zhang, X.; Yang, C.; Chen, Y.; Chen, Y.; Xu, J.; Li, H.; Qi, X.; et al. Structural characteristics and antioxidant activities of the extracellular polysaccharides produced by marine bacterium Edwardsiella tarda. Bioresour. Technol. 2010, 101, 4729–4732. [Google Scholar] [CrossRef]
  145. He, F.; Yang, Y.; Yang, G.; Yu, L. Components and antioxidant activity of the polysaccharide from Streptomyces virginia H03. Zeitschrift Naturforschung C 2008, 63, 181–188. [Google Scholar] [CrossRef] [Green Version]
  146. Qiao, D.; Luo, J.; Ke, C.; Sun, Y.; Ye, H.; Zeng, X. Immunostimulatory activity of the polysaccharides from Hyriopsis cumingii. Int. J. Biol. Macromol. 2010, 47, 676–680. [Google Scholar] [CrossRef] [PubMed]
  147. Katzenellenbogen, E.; Kocharova, N.A.; Shashkov, A.S.; Górska-Frączek, S.; Bogulska, M.; Gamian, A.; Knirel, Y.A. Structure of the O-polysaccharide of Edwardsiella tarda PCM 1150 containing an amide of d-glucuronic acid with l-alanine. Carbohydr. Res. 2013, 368, 84–88. [Google Scholar] [CrossRef]
  148. Katzenellenbogen, E.; Kocharova, N.A.; Shashkov, A.S.; Górska-Frączek, S.; Gamian, A.; Knirel, Y.A. Structure of the O-polysaccharide of Edwardsiella tarda PCM 1156. Carbohydr. Res. 2013, 374, 45–48. [Google Scholar] [CrossRef] [PubMed]
  149. Kodali, V.P.; Das, S.; Sen, R. An exopolysaccharide from a probiotic: Biosynthesis dyanmics, composition and emulsifying activity. Food Res. Int. 2009, 42, 695–699. [Google Scholar] [CrossRef]
  150. Ramanathan, T.; Ahmad, A.; Ahmad, A.S.; Kalimutho, M. Taxonomical identity and polysaccharide produced by Bacillus species isolated from old aged medicinal decoctions. J. Sustain. Sci. Manag. 2011, 6, 2–9. [Google Scholar]
  151. Raza, W.; Makeen, K.; Wang, Y.; Xu, Y.; Qirong, S. Optimization, purification, characterization and antioxidant activity of an extracellular polysaccharide produced by Paenibacillus polymyxa SQR-21. Bioresour. Technol. 2011, 102, 6095–6103. [Google Scholar] [CrossRef]
  152. Mian, J.A.; Percival, E. Carbohydrates of the brown seaweeds Himanthalia lorea, Bifurcaria bifurcata, and Padina pavonia: Part 1. Extraction and fractionation. Carbohydr. Res. 1973, 26, 133–146. [Google Scholar] [CrossRef]
  153. Rodríguez-Jasso, R.M.; Mussatto, S.I.; Pastrana, L.; Aguilar, C.N.; Teixeira, J.A. Chemical composition and antioxidant activity of sulphated polysaccharides extracted from Fucus vesiculosus using different hydrothermal processes. Chem. Pap. 2014, 68, 203–209. [Google Scholar] [CrossRef] [Green Version]
  154. Rupérez, P.; Ahrazem, O.; Leal, J.A. Potential Antioxidant Capacity of Sulfated Polysaccharides from the Edible Marine Brown SeaweedFucus vesiculosus. J. Agric. Food Chem. 2002, 50, 840–845. [Google Scholar] [CrossRef]
  155. Saha, S.; Navid, M.H.; Bandyopadhyay, S.S.; Schnitzler, P.; Ray, B. Sulfated polysaccharides from Laminaria angustata: Structural features and in vitro antiviral activities. Carbohydr. Polym. 2012, 87, 123–130. [Google Scholar] [CrossRef]
  156. Soukaina, B.; Zainab, E.A.-T.; Guillaume, P.; Halima, R.; Philippe, M.; Cherkaoui, E.M.; Cedric, D. Radical depolymerisation of Alginate extracted from Moroccan Brown Seaweed Bifiucaria bifurcate. Appl. Sci. 2020, 10, 4166. [Google Scholar] [CrossRef]
  157. Teruya, T.; Takeda, S.; Tamaki, Y.; Tako, M. Fucoidan Isolated from Laminaria angustata var. longissima Induced Macrophage Activation. Biosci. Biotechnol. Biochem. 2010, 74, 1960–1962. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  158. Vasantharaja, R.; Abraham, L.S.; Gopinath, V.; Hariharan, D.; Smita, K.M. Attenuation of oxidative stress induced mitochondrial dysfunction and cytoxicity in fibroblast cells by sulphated polysaccharide from Padina gymnospora. Int. J. Biol. Macromol. 2019, 124, 50–59. [Google Scholar] [CrossRef]
  159. Wijesekara, I.; Pangestuti, R.; Kim, S.-K. Biological activities and potential health benefits of sulphated polysaccharides derived from marine algae. Carbohydr. Polym. 2011, 84, 14–21. [Google Scholar] [CrossRef]
  160. Xu, C.H.; Fang, Y.; Lin, H.; Chen, L.; Li, Z.J.; Deng, D.; Lu, C.X. Chemical characters and anti oxidative properties of sulphated polysaccharides from Lamina japonica. J. Appl. Phycol. 2001, 13, 67–70. [Google Scholar] [CrossRef]
  161. Chen, T.; Li, B.; Li, Y.; Zhao, C.; Shen, J.; Zhang, H. Catalytic synthesis and antitumor activities of sulfated polysaccharide from Gynostemma pentaphyllum Makino. Carbohydr. Polym. 2011, 83, 554–560. [Google Scholar] [CrossRef]
  162. Chen, L.; Huang, G. Extraction, characterization and antioxidant activities of pumpkin polysaccharide. Int. J. Biol. Macromol. 2018, 118, 770–774. [Google Scholar] [CrossRef] [PubMed]
  163. Deng, Y.; Chen, L.-X.; Wu, D.-T.; Cheong, K.-L.; Chen, N.-F.; Zhao, J.; Li, S.-P. Qualitative and quantitative analysis of specific polysaccharides in Dendrobium huoshanense by using saccharide mapping and chromatographic methods. J. Pharm. Biomed. Anal. 2016, 129, 163–171. [Google Scholar] [CrossRef] [PubMed]
  164. Dong, H.; Zhang, Q.; Li, Y.; Li, L.; Lan, W.; He, J.; Li, H.; Xiong, Y.; Qin, W. Extraction, characterisation and antioxidant activities of polysaccharides of Chuanminshen violaceum. Int. J. Biol. Macromol. 2016, 86, 224–232. [Google Scholar] [CrossRef]
  165. Jiang, J.; Meng, F.Y.; He, Z.; Ning, Y.L.; Li, X.H.; Song, H.; Wang, J.; Zhou, R. Sulfated modification of longan polysaccharide and its immunomodulatory and antitumor activity in vitro. Int. J. Biol. Macromol. 2014, 67, 323–329. [Google Scholar] [CrossRef]
  166. Jing, Y.; Huang, L.; Lv, W.; Tong, H.; Song, L.; Hu, X.; Yu, R. Structural Characterization of a Novel Polysaccharide from Pulp Tissues of Litchi chinensis and Its Immunomodulatory Activity. J. Agric. Food Chem. 2014, 62, 902–911. [Google Scholar] [CrossRef] [PubMed]
  167. Khaskheli, S.G.; Zheng, W.; Sheikh, S.A.; Khaskheli, A.A.; Liu, Y.; Soomro, A.H.; Feng, X.; Sauer, M.B.; Wang, Y.F.; Huang, W. Characterisation of Auricularia auricula polysaccharides and its antioxidant properties in fresh and pickled product. Int. J. Biol. Macromol. 2015, 81, 387–395. [Google Scholar] [CrossRef]
  168. Li, H.; Su, J.; Jiang, J.; Li, Y.; Gan, Z.; Ding, Y.; Li, Y.; Liu, J.; Wang, S.; Ke, Y. Characterization of polysaccharide from Scutellaria barbata and its antagonistic effect on the migration and invasion of HT-29 colorectal cancer cells induced by TGF-β1. Int. J. Biol. Macromol. 2019, 131, 886–895. [Google Scholar] [CrossRef]
  169. Liu, J.; Zhao, Z.Z.; Chen, H.-B. Review of Astragali radix. Chin. Herb. Med. 2011, 3, 90–105. [Google Scholar]
  170. Liu, J.; Meng, C.-G.; Yan, Y.-H.; Shan, Y.-N.; Kan, J.; Jin, C.-H. Structure, physical property and antioxidant activity of catechin grafted Tremella fuciformis polysaccharide. Int. J. Biol. Macromol. 2016, 82, 719–724. [Google Scholar] [CrossRef]
  171. Lu, X.; Mo, X.; Guo, H.; Zhang, Y. Sulfation modification and anticoagulant activity of the polysaccharides obtained from persimmon (Diospyros kaki L.) fruits. Int. J. Biol. Macromol. 2012, 51, 1189–1195. [Google Scholar] [CrossRef] [PubMed]
  172. Luo, Q.; Cai, Y.; Yan, J.; Sun, M.; Corke, H. Hypoglycemic and hypolipidemic effects and antioxidant activity of fruit extracts from Lycium barbarum. Life Sci. 2004, 76, 137–149. [Google Scholar] [CrossRef] [PubMed]
  173. Mckenna, D.J.; Hughes, K.; Jones, K. Astragalus. Altern. Ther. Health Med. 2002, 8, 34–40. [Google Scholar]
  174. Nguyen, T.L.; Chen, J.; Hu, Y.; Wang, D.; Fan, Y.; Wang, J.; Abula, S.; Zhang, J.; Qin, T.; Chen, X. In vitro antiviral activity of sulfated Auricularia auricular polysaccharides. Carbohydr. Polym. 2012, 90, 1254–1258. [Google Scholar] [CrossRef]
  175. Qian, X.P.; Zha, X.Q.; Xiao, J.J.; Zhang, H.L.; Pan, L.H.; Luo, J.P. Sulfated modification can enhance antiglycation abilities of polysaccharides from Dendrobium huoshanense. Carbohydr. Polym. 2014, 101, 982–989. [Google Scholar] [CrossRef]
  176. Song, X.; Yin, Z.; Zhao, X.; Cheng, A.; Jia, R.; Yuan, G.; Xu, J.; Fan, Q.; Dai, S.; Lu, H. Antiviral activity of sulfated Chuanmingshen violaceum polysaccharide against Newcastle disease virus. J. Gen. Virol. 2013, 94, 2164–2174. [Google Scholar] [CrossRef] [Green Version]
  177. Song, X.; Xu, J.; Yin, Z.; Jia, R.; Cheng, A.; Deng, Y.; Lu, C.; Liang, X.; Wang, Y.; Yao, X.; et al. Effects of Polysaccharide from Chuanminshen violaceum on Immune Response of Newcastle Disease Vaccine in Chicken. Acta Sci. Vet. 2013, 41, 1104. [Google Scholar]
  178. Song, Y.; Yang, Y.; Zhang, Y.; Duan, L.; Zhou, C.; Ni, Y.; Liao, X.; Li, Q.; Hu, X. Effect of acetylation on antioxidant and cytoprotective activity of polysaccharides isolated from pumpkin (Cucurbita pepo, lady godiva). Carbohydr. Polym. 2013, 98, 686–691. [Google Scholar] [CrossRef] [PubMed]
  179. Xu, Y.; Jiang, H.; Sun, C.; Adu-Frimpong, M.; Deng, W.; Yu, J.; Xu, X. Antioxidant and hepatoprotective effects of purified Rhodiola rosea polysaccharides. Int. J. Biol. Macromol. 2018, 117, 167–178. [Google Scholar] [CrossRef] [PubMed]
  180. Yan, J.-K.; Wang, Y.-Y.; Ma, H.-L.; Wang, Z.-B. Ultrasonic effects on the degradation kinetics, preliminary characterization and antioxidant activities of polysaccharides from Phellinus linteus mycelia. Ultrason. Sonochem. 2015, 29, 251–257. [Google Scholar] [CrossRef]
  181. Yang, C.; He, N.; Ling, X.; Ye, M.; Zhang, C.; Shah, W.; Yao, C.; Wang, Z.; Li, Q. The isolation and characterisation of polysaccharides from longan pulp. Sep. Purif. Technol. 2008, 63, 226–230. [Google Scholar] [CrossRef]
  182. Yin, J.Y.; Chan, B.C.L.; Yu, H.; Lau, I.Y.K.; Han, X.Q.; Cheng, S.W. Separation, structure characterization, conformation and immunomodulating effect of a hyperbranched heteroglycan from Radix astragali. Carbohydr. Polym. 2012, 87, 667–675. [Google Scholar] [CrossRef]
  183. Zhang, L.; Yang, L.; Ding, Q.; Chen, X. Studies on molecular weights of polysaccharides of Auricularia auricula-judae. Carbohydr. Res. 1995, 270, 1–10. [Google Scholar] [CrossRef]
  184. Zhang, Y.; Lu, X.; Fu, Z.; Wang, Z.; Zhang, J. Sulphated modification of a polysaccharide obtained from fresh persimmon (Diospyros kaki L.) fruit and antioxidant activities of the sulphated derivatives. Food Chem. 2011, 127, 1084–1090. [Google Scholar] [CrossRef] [PubMed]
  185. Ayimbila, F.; Keawsompong, S. Functional composition and antioxidant property of crude polysaccharides from the fruiting bodies of Lentinus squarrosulus. 3 Biotech. 2021, 11, 1–13. [Google Scholar] [CrossRef] [PubMed]
  186. Chen, C.; Wu, W.H.; Xu, X.J.; Zhang, L.N.; Liu, Y.; Wang, K.P. Chain conformation and anti-tumor activity of derivatives of polysaccharide from Rhizoma Panacis Japonici. Carbohydr. Polym. 2014, 105, 308–316. [Google Scholar] [CrossRef] [PubMed]
  187. Deng, Y.Y.; Yi, Y.; Zhang, L.F.; Zhang, R.F.; Zhang, Y.; Wei, Z.C.; Tang, X.J.; Zhang, M.W. Immunomodulatory activity and partial characterization of polysaccharides from Momordica charantia. Molecules 2014, 19, 13432–13447. [Google Scholar] [CrossRef] [Green Version]
  188. Deng, C.; Fu, H.; Xu, J.; Shang, J.; Cheng, Y. Physiochemical and biological properties of phosphorylated polysaccharides from Dictyophora indusiata. Int. J. Biol. Macromol. 2015, 72, 894–899. [Google Scholar] [CrossRef]
  189. Jung, H.Y.; Bae, I.Y.; Lee, S.; Lee, H.G. Effect of the degree of sulfation on the physicochemical and biological properties of Pleurotus eryngii polysaccharides. Food Hydrocoll. 2011, 25, 1291–1295. [Google Scholar] [CrossRef]
  190. Juyi, G.; Jiantin, Z.; Yihong, H. Isolation, Purification and Characterization of Polysaccharide from the Fruit Bodies of Pleurotus cornucopiae (Paul:Pers.) Roll. 1999. Available online: (accessed on 19 January 2021).
  191. Ren, D.; Jiao, Y.; Yang, X.; Yuan, L.; Guo, J.; Zhao, Y. Antioxidant and antitumor effects of polysaccharides from the fungus Pleurotus abalonus. Chem. Interact. 2015, 237, 166–174. [Google Scholar] [CrossRef] [PubMed]
  192. Rout, D.; Mondal, S.; Chakraborty, I.; Islam, S.S. The structure of a polysaccharide from Fraction-II of an edible mushroom, Pleurotus florida. Carbohydr. Res. 2006, 341, 995–1002. [Google Scholar] [CrossRef] [PubMed]
  193. Villares, A. Polysaccharides from the edible mushroom Calocybe gambosa: Structure and chain conformation of a (1→4),(1→6)-linked glucan. Carbohydr. Res. 2013, 375, 153–157. [Google Scholar] [CrossRef] [PubMed]
  194. Wang, F.; Hao, L.; Jia, S.; Wang, Q.; Zhang, X.; Niu, S. A Review of Research on Polysaccharide from Coriolus versicolor. In Proceedings of the 2012 International Conference on Applied Biotechnology (ICAB 2012); Lecture Notes in Electrical Engineering; Zhang, T.C., Ouyang, P., Kaplan, S., Skarnes, B., Eds.; Springer: Berlin/Heidelberg, Germany, 2014; Volume 249. [Google Scholar]
  195. Wu, Y.; Ye, M.; Du, Z.; Jing, L.; Surahio, M.; Yang, L. Carboxymethylation of an exopolysaccharide from Lachnum and effect of its derivatives on experimental chronic renal failure. Carbohydr. Polym. 2014, 114, 190–195. [Google Scholar] [CrossRef]
  196. Xin, X.; Zheng, K.; Niu, Y.; Song, M.; Kang, W. Effect of Flammulina velutipes (golden needle mushroom, eno-kitake) polysaccharides on constipation. Open Chem. 2018, 16, 155–162. [Google Scholar] [CrossRef]
  197. Xu, W.; Zhang, F.; Luo, Y.; Ma, L.; Kou, X.; Huang, K. Antioxidant activity of a water-soluble polysaccharide purified from Pteridium aquilinum. Carbohydr. Res. 2009, 344, 217–222. [Google Scholar] [CrossRef] [PubMed]
  198. Xu, X.; Yan, H.; Tang, J.; Chen, J.; Zhang, X. Polysaccharides inLentinus edodes: Isolation, Structure, Immunomodulating Activity and Future Prospective. Crit. Rev. Food Sci. Nutr. 2014, 54, 474–487. [Google Scholar] [CrossRef] [PubMed]
  199. Ye, M.; Chen, W.-X.; Qiu, T.; Yuan, R.-Y.; Ye, Y.-W.; Cai, J.-M. Structural characterisation and anti-ageing activity of extracellular polysaccharide from a strain of Lachnum sp. Food Chem. 2012, 132, 338–343. [Google Scholar] [CrossRef]
  200. Zhang, H.; Li, W.-J.; Nie, S.-P.; Chen, Y.; Wang, Y.-X.; Xie, M.-Y. Structural characterisation of a novel bioactive polysaccharide from Ganoderma atrum. Carbohydr. Polym. 2012, 88, 1047–1054. [Google Scholar] [CrossRef]
  201. Zhao, L.; Dong, Y.; Chen, G.; Hu, Q. Extraction, purification, characterization and antitumor activity of polysaccharides from Ganoderma lucidum. Carbohydr. Polym. 2010, 80, 783–789. [Google Scholar] [CrossRef]
  202. Zhu, Z.Y.; Pang, W.; Li, Y.Y.; Ge, X.R.; Chen, L.J.; Liu, X.C.; Lv, Q.; Dong, G.L.; Liu, A.J.; Zhang, Y. Effect of ultrasonic treatment on structure and antitumor activity of mycelial polysaccharides from Cordyceps gunnii. Carbohydr. Polym. 2014, 114, 12–20. [Google Scholar] [CrossRef] [PubMed]
  203. Gao, Y.; Guo, Q.; Zhang, K.; Wang, N.; Li, C.; Li, Z.; Zhang, A.; Wang, C. Polysaccharide from Pleurotus nebrodensis: Physicochemical, structural characterization and in vitro fermentation characteristics. Int. J. Biol. Macromol. 2020, 165, 1960–1969. [Google Scholar] [CrossRef] [PubMed]
  204. Ng, T.B.; Pi, Z.F.; Fu, M.; Li, L.; Hou, J.; Shi, L.S.; Chen, R.R.; Jiang, Y.; Liu, F.; Yue, H.; et al. A polysaccharopeptide complex and a condensed tannin with antioxidant activity from dried rose (Rosa rugosa) flowers. J. Pharm. Pharmacol. 2006, 58, 529–534. [Google Scholar] [CrossRef]
  205. Chen, H.; Zhang, M.; Xie, B. Components and antioxidant activity of polysaccharide conjugate from green tea. Food Chem. 2005, 90, 17–21. [Google Scholar] [CrossRef]
  206. Tannin-Spitz, T.; Bergman, M.; Van-Moppes, D.; Grossman, S.; Arad, S. (Malis) Antioxidant activity of the polysaccharide of the red microalga Porphyridium sp. Environ. Boil. Fishes 2005, 17, 215–222. [Google Scholar] [CrossRef]
  207. Zhang, Z.; Wang, X.; Zhao, M.; Yu, S.; Qi, H. The immunological and antioxidant activities of polysaccharides extracted from Enteromorpha linza. Int. J. Biol. Macromol. 2013, 57, 45–49. [Google Scholar] [CrossRef]
  208. Kodali, V.P.; Sen, R. Antioxidant and free radical scavenging activities of an exopolysaccharide from a probiotic bacterium. Biotechnol. J. 2008, 3, 245–251. [Google Scholar] [CrossRef]
  209. Okafor, D.C.; Onuegbu, N.C.; Odimegwu, N.E.; Ibeabuchi, J.C.; Njoku, N.E.; Agunwa, M.; Ofoedu, C.E.; Njoku, C.C. Antioxidant and Antimicrobial Activities of Oyster Mushroom. Am. J. Food Sci. Technol. 2017, 5, 64–69. [Google Scholar] [CrossRef]
  210. Zhu, Z.-Y.; Liu, R.-Q.; Si, C.-L.; Zhou, F.; Wang, Y.-X.; Ding, L.-N.; Jing, C.; Liu, A.-J.; Zhang, Y.-M. Structural analysis and anti-tumor activity comparison of polysaccharides from Astragalus. Carbohydr. Polym. 2011, 854, 895–902. [Google Scholar] [CrossRef]
Figure 1. The electron transport chain is a series of electron transporters embedded in the inner mitochondrial membrane that shuttles electrons from NADH and FADH2 to molecular oxygen. In the process, protons are pumped from the mitochondrial matrix to the intermembrane space, and oxygen is reduced to form water (Source: Electron Transport Chain [50]).
Figure 1. The electron transport chain is a series of electron transporters embedded in the inner mitochondrial membrane that shuttles electrons from NADH and FADH2 to molecular oxygen. In the process, protons are pumped from the mitochondrial matrix to the intermembrane space, and oxygen is reduced to form water (Source: Electron Transport Chain [50]).
Foods 10 00699 g001
Figure 2. The role of hydrogen peroxide (H2O2) in cell dysfunction and transformation.
Figure 2. The role of hydrogen peroxide (H2O2) in cell dysfunction and transformation.
Foods 10 00699 g002
Figure 3. Proposed mechanism of oxidation of gallocatechin (Source: Nakayama et al. [61]).
Figure 3. Proposed mechanism of oxidation of gallocatechin (Source: Nakayama et al. [61]).
Foods 10 00699 g003
Figure 4. Production of hydrogen peroxide (H2O2) by polyphenols via autoxidation (Source: Akagawa et al. [62]).
Figure 4. Production of hydrogen peroxide (H2O2) by polyphenols via autoxidation (Source: Akagawa et al. [62]).
Foods 10 00699 g004
Figure 5. Schematic diagram for the oxidation of β-D-glucose, as catalysed by glucose oxidase (Source: Adányi et al. [85]).
Figure 5. Schematic diagram for the oxidation of β-D-glucose, as catalysed by glucose oxidase (Source: Adányi et al. [85]).
Foods 10 00699 g005
Figure 6. Schematic diagram of cholesterol oxidation and isomerization catalysed by cholesterol oxidase (Source: Adányi et al. [85]).
Figure 6. Schematic diagram of cholesterol oxidation and isomerization catalysed by cholesterol oxidase (Source: Adányi et al. [85]).
Foods 10 00699 g006
Figure 7. Proposed reaction mechanism of OH• on a polysaccharide chain. The diagram shows some of the proposed reaction mechanism of polysaccharide (homogalacturonan) exerting its antioxidant capacity on OH•, where the short lines (- - -) depicts the polysaccharide chain continuation. This figure presents the predicted reactions occurring after •OH abstracts a hydrogen atom from C-1 (reactions c1–c4), C-3 (reactions b1–b3), C-4 (reactions d1–d4) or C-5 (reactions a1–a4). Abstraction of hydrogen from C-2 (not shown) is expected to give products directly comparable with those shown for C-3. (Source: Vreeburg et al. [140]).
Figure 7. Proposed reaction mechanism of OH• on a polysaccharide chain. The diagram shows some of the proposed reaction mechanism of polysaccharide (homogalacturonan) exerting its antioxidant capacity on OH•, where the short lines (- - -) depicts the polysaccharide chain continuation. This figure presents the predicted reactions occurring after •OH abstracts a hydrogen atom from C-1 (reactions c1–c4), C-3 (reactions b1–b3), C-4 (reactions d1–d4) or C-5 (reactions a1–a4). Abstraction of hydrogen from C-2 (not shown) is expected to give products directly comparable with those shown for C-3. (Source: Vreeburg et al. [140]).
Foods 10 00699 g007
Table 1. Some key situations associated with the formation of H2O2.
Table 1. Some key situations associated with the formation of H2O2.
LocationIn VitroReference(s)In VivoReference(s)
Source/siteBeverages (e.g., green tea, black tea, wine, beer, coffee)Grzesik et al. [63]MitochondriaLennicke et al. [15];
Baveris and Cadenas [86];
Wong et al. [87];
Treberg et al. [88]
Trigger agentCertain phenolic compounds (e.g., tea catechins, (+)-catechin, and gallic acid)Wee et al. [67];
Grzesik et al. [63]
Escaped electron (e) in the electron transport chainPhaniendra et al. [7];
Druck et al. [89];
Maddu [2]
Production processAuto-oxidationErickson [26];
Kim et al. [65];
Bensid et al. [27]
Cellular oxidative metabolismPatel et al. [6];
Bottje [90];
Fang et al. [91]
Factors responsible for H2O2 formationContent and composition of phenolic compounds;
Structure of phenolic compounds;
Metal ions;
Incubation time.
Akagawa et al. [62];
Grzesik et al. [63];
Bopitiya et al. [59]
The action of certain enzymes such as superoxide dismutase (SOD), xanthine oxidase, monoamine oxidase, and glycollate oxidase;
Type of ingested beverage such as green tea, black tea, and/or coffee;
Available oxygen;
The joint action of in vivo dismutation of superoxide radical and in vitro auto-oxidation of phenolic compounds
Fang et al. [91];
Grzesik et al. [63];
Bopitiya et al. [59]
Table 2. Antioxidative polysaccharides, with corresponding sources, and polysaccharides specifics.
Table 2. Antioxidative polysaccharides, with corresponding sources, and polysaccharides specifics.
Antioxidative PolysaccharideSourcesPolysaccharides SpecificsReferences
Astragali radixPlant From Astragali radix, isolated two glacans (AG-1, identified as as α-(1,4) and α-(1,6) glucan (5:2); and AG-2 identified as α-(1,4) glucan) and two heterosaccharides (AH-1, identified as acidic polysaccharide, composed of hexouronic acid, glucose, rhamnose, and arabinose in the ratio of 1:0.04:0.02:0.0; and AH-2, identified as glucose and arabinose in the ratio of 1:0.15) from an Astragali radix polysaccharidesLiu et al. [169];
Mckenna et al. [173]
Astragalus mongholicusPlantPolysaccharides from A. mongholicus, named as APS-I and APS-II, comprised backbone (1,3)-β-d-glucopyranosyl (Glcp) residues, with bioactivities closely related to their chemical composition, configuration, molecular weight, and physical propertiesYin et al. [182];
Zhu et al. [210]
Litchi chinensis Sonn.PlantA novel polysaccharide (LCP50W) from pulp tissues of Litchi chinensis had main chain consist of (1→3)-linked β-l-rhamnopyranosyl, (1→6)-linked α-d-glucopyranosyl, and (1→2,6)-linked α-d-glucopyranosyl residues, which branched at O-6. The three branches consisted of (1→2)-linked α-l-rhamnopyranosyl, (1→3)-linked α-d-galactopyranosyl, and (1→3)-linked α-l-mannopyranosyl residues, terminated with (1→)-linked α-l-arabinopyranosyl residues, respectivelyJing et al. [166];
Yin et al. [182]
Lycium barbarumPlantRaw and Purified Lycium barbarum polysaccharides (LBP) would contain sugars such as Rha, Gal, Glc, Ara, Man, Xyl, with molar ratio of 4.22, 2.43, 1.38, 1, 0.95, and 0.38, respectively Luo et al. [172];
Yin et al. [182]
Scutellaria barbata D. DonPlantFrom the whole plant of Scutellaria barbata D. Don, polysaccharide, SBPW3, showed a molecular weight of 10.2 kDa and composed of arabinose (25.68%), galactose (27.72%), glucose (20.59%), mannose (12.56%), rhamnose (2.51%), and xylose (10.94%)Li et al. [168];
Yin et al. [182]
Ribes nigrum L.PlantPolysaccharides from blackcurrant (Ribes nigrum L.) (BCP), showed two low-molecular-weight polysaccharides (DBCP-1, DBCP-2) with same monosaccharide units with resembling glycosidic linkage patternsXu et al. [179];
Yin et al. [182]
Tremella fuciformisPlantTremella fuciformis polysaccharide (catechin-g-TPS) with molecular weight of 5.82 × 105 Da, given by polysaccharide fractions (F-1 and F-2)Liu et al. [170];
Wang et al. [123];
Zhang et al. [34]
Auricularia auriculaPlant- Fruit bodies of Auricularia auricula were comprised of Glucans A, C, and E, mainly backbone chain of Beta (1-3)-d-glucose residues, with various branched groups. Specifically, polysaccharides D and B would have residues of d-xylose, d-mannose, d-galactose, d-glucose, and d-glucuronic acid.
- Two polysaccharide fractions (AAPF, AAPP) purified from the fruiting body of A. auricula, showed AAPF with five monosaccharides, including glucose, rhamnose, arabinose, mannose, and galactose with a molar ratio of 16.74:1.0:1.18:1.0:1.0; and AAPP with four monosaccharides, namely arabinose, mannose, galactose, and xylose with the molar ratio of 15.59:1.52:4.76:1.0
Zhang et al. [183];
Khaskheli et al. [167];
Nguyen et al. [174];
Zhang et al. [34]
Chuanminshen violaceumPlant- Chuanminshen violaceum polysaccharides (CVPS) comprise d-carubinose and d-glucose, in the ratio of 1: 16.2. The average molecular and number-average molecular weight CVPS of 9.7632 × 105 Da and 5.2270 × 104 Da respectively.
- Another study showed Chuanminshen violaceum polysaccharides (CVPs) with neutral polysaccharides, mainly composed of glucose and galactose, with molecular weights ranging between 233.69 and 11.02 kDa.
Dong et al. [164];
Song et al. [176,178]
Dendrobium huishanensePlantWith molecular weights range 1.16 × 105 to 2.17 × 105 Da, with main monosaccharide compositions that include Man and Glc, with glycosidic linkages β-1,4-Manp and β-1,4-Glcp, and substituted with acetyl groups at O-2 and O-3 of 1,4-linked ManpDeng et al. [163];
Qian et al. [175]
PersimmonPlant Sulphated modification, polysaccharide was obtained from fresh (Diospyros kaki L.) persimmon fruit (PFP), Three sulphated derivatives of PFP, known as PFP-SI, PFP-SII and PFP-SIII, were obtained, with average molecular weights of 53, 51 and 48 kDa, respectively,Lu et al. [171];
Zhang et al. [184]
LonganPlant Longan polysaccharides (LPSs), extracted from (logan) pulp, showed β-type acidic heteropolysaccharides with pyran group, wherein LPS-N had glucose/xylose molar ratio of 1.9:1, with LPS-A1 having rhamnose, xylose, arabinose and galactose molar ratio of 1:1.64:4.33:2.28, and LPS-A2 having only rhamnoseJiang et al. [165];
Yang et al. [181]
Azadirachta indica leavesPlant A native polysaccharide of Azadirachta indica (leaves) obtained an apparent molecular mass of 80 kDa, constituting (1→5)-/(1→3,5)-linked α-l-arabinosyl, (1→3)-/(1→6)-/(1→3,6)-linked β-d-galactosyl, and terminal-rhamnosyl and α-l-arabinosyl residuesSaha et al. [42]
Portulaca oleracea L. Plant A yield of 6.45% of dried raw material, P. oleracea showed water-soluble crude polysaccharides (DCPOP), 92% carbohydrate, free of proteins and with average molecular weight of 24.6 kDaChen et al. [161];
Shen et al. [42]
PumpkinPlant Potentially containing acetyl groups, the pumpkin polysaccharide showed α/β-glycosidic bond linkage. It is considered a heteropolysaccharide, composed of six monosaccharides, namely arabinose, galactose, glucose, glucuronic acid, rhamnose, and xylose Chen and Huang [162];
Song et al. [177]
Ganoderma lucidumFungiPolysaccharides of Ganoderma lucidum consisted primarily of glucose and galactose, small amounts of mannose, rhamnose and fucose, and minor amount of glucuronic acidWang et al. [25];
Xu et al. [197];
Zhao et al. [201]
Athyrium multidentatumFungiFive polysaccharide fractions (PS-1, PS-2, PS-3, PS-4, and PS-5) isolated from Athyrium multidentatum (Doll.) Ching, was heteropolysaccharides with different molecular weights and monosaccharide compositionsJing et al. [38];
Wang et al. [25]
Flammulina sppFungiFlammulina polysaccharide comprise glucan, which could have a mix of some other fractions, like fucosan, galactose, glycan, mannan, and xylanWang et al. [25];
Xin et al. [196]
Coriolus versicolorFungiPolysaccharide of Coriolus versicolor contains extracellular polysaccharide (EPS) mainly β-1, 3/β-1, 6-linked d-glucose molecules, predominantly comprising glucose and small amounts of arabinose, galactose, mannose, and xyloseWang et al. [194];
Wang et al. [25]
Pleurotus nebrodensisFungiPolysaccharide (PNPS) from the fruiting body of Pleurotus nebrodensis contained mainly glucose, with a small percentage of galactose and mannose Gao et al. [203];
Wang et al. [25]
Pleurotus abalunesFungiPolysaccharides of Pleurotus abalonus is heteropolysaccharide, comprise d-mannose, d-ribose, l-rhamnose, d-glucuronic acid, d-glucose and d-galactose, having corresponding mole percentages 3.4%, 1.1%, 1.9%, 1.4%, 87.9% and 4.4%, respectivelyRen et al. [191];
Wang et al. [25]
Pleurotus eryngiiFungiPolysaccharides from P. eryngii comprise mainly of β-(1→3)-glucans with β-(1→6) branchesJung et al. [189];
Wang et al. [25]
Pleurotus corncopiaeFungiPolysaccharides isolated from Pleurotus cornucopiae composed of arabinose, glucose, mannose and xyloseJuyi et al. [190];
Wang et al. [25]
Pleurotus floridaFungiP. florida consists of three different polysaccharides, which include (1→3)-, (1→6)-branched glucan, and (1→6)-α-glucanRout et al. [192];
Wang et al. [25]
Calocybe gambosaFungi-Three polysaccharides were isolated from the fruiting bodies Calocybe gambosa mushroom, composed of glucose, and methylation analysis, showed the units were (1→4),(1→6)-linked with a degree of branching (DB) of 4%Villares [193];
Wang et al. [25]
Lentinus edodesFungiL. edodes can be divided into three layers: (1) the outside layer has heteropolysaccharide and β-(1→3)-glucan with β-(1→6) branches; (2) middle layer has mainly β-(1→6)-glucan with a small number of β-(1→3) branches; (3) inner layer being complex chitin, β-glucan, with small amount of acid polymerXu et al. [198];
Wang et al. [25]
Lentinus polyschrous Lev.FungiThe major monosaccharides of Lentinus polychrous include d-galactose, d-glucose, and d-mannose at different proportions, fucose, and xylose Ayimbila and Keawsompong [185];
Wang et al. [25]
Pleurotus ostreatusFungiPolysaccharide isolated from Pleurotus ostreatus is heteropolysaccharide, composed of galactose, glucose, mannose, rhamnose, and xyloseVamanu [15];
Wang et al. [25];
Zhang et al. [200]
Cordyceps gunniFungiThe polysaccharide of Cordyceps gunnii was the α-polysaccharide type, with a glycosidic bond, mainly composed of d-mannose, d-glucose, and d-galactoseZhu et al. [202];
Zhu et al. [45]
Grifola frondosaFungiG. frondosa polysaccharides contain glucose, galactose, mannose, fucose, and ribose, with high amounts of (1→3,1→6)-β-d-glucans, which would account for 13.2% of water-soluble polysaccharidesBae et al. [117];
He et al. [131]
LachnumFungi Polysaccharide of Lachnum YM261(LEPS-1) is considered homogenous with a molecular weight of 21,670 Da, with its glucan linked by β-(1→3)-d-pyran glycosidic bondWu et al. [195];
Ye et al. [199]
Dictyophora indusiataFungiPolysaccharide (DIP) extracted from Dictyophora indusiata indicated a specific polysaccharide of (1→3)- β-d-glucan with (1→6)-β-glucopyranoside side chainsDeng et al. [187];
Deng et al. [188]
Ganoderma atrumFungi Polysaccharide fraction from Ganoderma atrum composed of glucose (Glc), mannose (Man), galactose (Gal) and galacturonic acid (GalA) in molar ratio of 4.91:1:1.28:0.71Chen et al. [186];
Zhang et al. [200]
Paenibacillus polymyxaBacteria- Paenibacillus polymyxa bioactive compounds like exopolysaccharides, for example, d-glucuronic acid, and polysaccharide beads
- Paenibacillus polymyxa was shown to comprise mannose, galactose, and glucose in a ratio of 1.23:1.14:1
Daud et al. [143];
Raza et al. [151];
Wang et al. [25]
Bacillus coagulansBacteria- Bacillus coagulans polysaccharide could have with monosaccharides like galactose, glucose, lactose, mannose, raffinose, rhamnose, and xylose
- Exopolysaccharide (EPS) from Bacillus coagulans, comprise galactose, mannose, fucose, glucose, and glucosamine
Kodali et al. [149];
Ramanathan et al. [150];
Wang et al. [25];
Edwardsiella tardaBacteria- E. tarda polysaccharide comprise glucosamine, a fucose, a mannose and a galactosamine unit;
- Another E. tarda polysaccharide comprise d-glucuronic acid with l-alanine, branched hexasaccharide repeating unit.
- branch points situated at C-2 and C-6 positions of the (1→3)-linked mannose residues, whereas the side chains composed of (1→2)-linked and (1→)-linked mannose residues
Guo et al. [144];
Katzenellenbogen et al. [147,148];
Wang et al. [25]
Streptomyces virginiaBacteria-A purified polysaccharide Streptomyces virginia (HO3 type) comprised mannose, glucose and galactose, in a 2:1:1 proportion, with average apparent molecular weight of 3.76 × 104 DaHe et al. [145]
Hyriopsis cumingiiAnimal Polysaccharides of Hyriopsis cumingii have pyranose rings, with glycosyl residues, either linked by α- and β-configuration glycosidic bonds Qiao et al. [141];
Wang et al. [25]
Misgurnus anguillicaudatusAnimalMisgurnus anguillicaudatus, polysaccharide (MAP) or misgurnan, comprise the major structural monomers identified included d-galactose, l-fucose and d-mannose, and monoses that link each other by α-1,3 bondsQin et al. [142];
Wang et al. [25]
Laminaria japonicaAlgaesulfated polysaccharide (L-A) has high galactose content but low fucose residues proportionWang et al. [25];
Xu et al. [160]
Enteromorpha linzaAlgaeEnteromorpha linza polysaccharides contain large amount of sulfated polysaccharides, composed mainly of galactose, mannose, glucuronic acid, glucose, rhamnose, and xyloseWang et al. [25];
Zhang et al. [36]
Bifurcaria bifurcataAlgaeGlucose and myo-inositol have been detected in Bifurcaria bifurcataMian and Percival [152];
Soukaina et al. [156]
Phaeophyceae sppAlgaeSulfated polysaccharides (SPs) found in Phaeophyceae spp can include laminarans, which comprise repeating dimeric units of λ-carrageenanWang et al. [25];
Wijesekara et al. [159]
Fucus vesiculosus LinnaeusAlgae- The main neutral sugars of Fucus vesiculosus include fucose, glucose, galactose, and xylose
- Sulphated polysaccharides of Fucus vesiculosus has a heterogeneous structure
Rodriguez-Jasso et al. [153];
Rupérez, Ahrazem, and Antonio Leal [154];
Wang et al. [25]
Padina gymnospora SonderAlgaeSulfated polysaccharide (SP) isolated from Padina gymnospora contains 29.4 ± 0.35% of sulfate, 27 ± 0.11% of fucose, 0.05 ± 0.12% of proteinVasantharaja et al. [158];
Wang et al. [25]
Laminaria angustataAlgae Polysaccharides from Laminaria angustata obtained via fucoidan -led macrophage activation produced chemical composition ratio of l-fucose:d-galactose:d-glucose:d-xylose:uronic acid:sulfate = 1.00:0.54:0.08:0.08:0.64:0.87Saha et al. [155];
Teruya et al. [157]
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Ofoedu, C.E.; You, L.; Osuji, C.M.; Iwouno, J.O.; Kabuo, N.O.; Ojukwu, M.; Agunwah, I.M.; Chacha, J.S.; Muobike, O.P.; Agunbiade, A.O.; et al. Hydrogen Peroxide Effects on Natural-Sourced Polysacchrides: Free Radical Formation/Production, Degradation Process, and Reaction Mechanism—A Critical Synopsis. Foods 2021, 10, 699.

AMA Style

Ofoedu CE, You L, Osuji CM, Iwouno JO, Kabuo NO, Ojukwu M, Agunwah IM, Chacha JS, Muobike OP, Agunbiade AO, et al. Hydrogen Peroxide Effects on Natural-Sourced Polysacchrides: Free Radical Formation/Production, Degradation Process, and Reaction Mechanism—A Critical Synopsis. Foods. 2021; 10(4):699.

Chicago/Turabian Style

Ofoedu, Chigozie E., Lijun You, Chijioke M. Osuji, Jude O. Iwouno, Ngozi O. Kabuo, Moses Ojukwu, Ijeoma M. Agunwah, James S. Chacha, Onyinye P. Muobike, Adedoyin O. Agunbiade, and et al. 2021. "Hydrogen Peroxide Effects on Natural-Sourced Polysacchrides: Free Radical Formation/Production, Degradation Process, and Reaction Mechanism—A Critical Synopsis" Foods 10, no. 4: 699.

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop