Next Article in Journal
Nutrient Deprivation-Associated Changes in Green Microalga Coelastrum sp. TISTR 9501RE Enhanced Potent Antioxidant Carotenoids
Next Article in Special Issue
Padina pavonica Extract Promotes In Vitro Differentiation and Functionality of Human Primary Osteoblasts
Previous Article in Journal
Sea Anemone Toxins: A Structural Overview
Previous Article in Special Issue
Integral Utilization of Red Seaweed for Bioactive Production
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Marine Drugs
Volume 17
Issue 6
10.3390/md17060327
marinedrugs-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Prebiotics from Seaweeds: An Ocean of Opportunity?

by
Paul Cherry
1,2,3,
Supriya Yadav
2,
Conall R. Strain
2,3,
Philip J. Allsopp
1,
Emeir M. McSorley
1,
R. Paul Ross
3,4 and
Catherine Stanton
2,3,*
1
Nutrition Innovation Centre for Food and Health, Ulster University, Cromore Road, Coleraine, Co. Londonderry BT52 1SA, UK
2
Teagasc Food Research Centre, Moorepark, Fermoy, P61 C996 Co. Cork, Ireland
3
APC Microbiome Ireland, University College Cork, T12 YT20 Cork, Ireland
4
College of Science, Engineering and Food Science, University College Cork, T12 K8AF Cork, Ireland
*
Author to whom correspondence should be addressed.
Mar. Drugs 2019, 17(6), 327; https://doi.org/10.3390/md17060327
Submission received: 1 May 2019 / Revised: 27 May 2019 / Accepted: 29 May 2019 / Published: 1 June 2019
(This article belongs to the Special Issue Seaweeds Secondary Metabolites: Successes in and/or Probable Therapeutic Applications)
Versions Notes

Abstract

:
Seaweeds are an underexploited and potentially sustainable crop which offer a rich source of bioactive compounds, including novel complex polysaccharides, polyphenols, fatty acids, and carotenoids. The purported efficacies of these phytochemicals have led to potential functional food and nutraceutical applications which aim to protect against cardiometabolic and inflammatory risk factors associated with non-communicable diseases, such as obesity, type 2 diabetes, metabolic syndrome, cardiovascular disease, inflammatory bowel disease, and some cancers. Concurrent understanding that perturbations of gut microbial composition and metabolic function manifest throughout health and disease has led to dietary strategies, such as prebiotics, which exploit the diet-host-microbe paradigm to modulate the gut microbiota, such that host health is maintained or improved. The prebiotic definition was recently updated to “a substrate that is selectively utilised by host microorganisms conferring a health benefit”, which, given that previous discussion regarding seaweed prebiotics has focused upon saccharolytic fermentation, an opportunity is presented to explore how non-complex polysaccharide components from seaweeds may be metabolised by host microbial populations to benefit host health. Thus, this review provides an innovative approach to consider how the gut microbiota may utilise seaweed phytochemicals, such as polyphenols, polyunsaturated fatty acids, and carotenoids, and provides an updated discussion regarding the catabolism of seaweed-derived complex polysaccharides with potential prebiotic activity. Additional in vitro screening studies and in vivo animal studies are needed to identify potential prebiotics from seaweeds, alongside untargeted metabolomics to decipher microbial-derived metabolites from seaweeds. Furthermore, controlled human intervention studies with health-related end points to elucidate prebiotic efficacy are required.

1. Introduction

Seaweeds are an underexploited and sustainable crop which offer a rich source of bioactive compounds, including novel dietary fibres, polyphenols, fatty acids, and carotenoids [1,2]. Epidemiological evidence comparing Japanese and Western diets have correlated seaweed consumption (5.3 g/day in Japan) with decreased incidence of chronic disease [3], while the purported efficacies of seaweed phytochemicals have led to potential functional food and nutraceutical applications which aim to protect against cardiometabolic and inflammatory risk factors associated with non-communicable diseases, such as obesity, type two diabetes, metabolic syndrome, cardiovascular disease, inflammatory bowel disease, and some cancers [1].
Current understanding of mutualistic diet-host-microbe interactions has generated efforts to exploit diet to maintain health status, and to prevent or overcome non-communicable diseases, where an imbalance of gut microbiota composition and metabolic function manifests during the onset and pathophysiology of gastrointestinal, neurological, and cardio-metabolic diseases, often congruent with intestinal inflammation and compromised gut barrier function [4,5]. As such, it has become pertinent to explore dietary strategies which modulate gut microbial composition and function to improve host health. This includes the use of prebiotics as fermentable substrates to enable selective gut commensal metabolism.
The prebiotic definition was recently updated to “a substrate that is selectively utilised by host microorganisms conferring a health benefit” [6], which includes the inhibition of pathogens, immune system activation, and vitamin synthesis and provides opportunity to explore the prebiotic efficacy of non-complex polysaccharide components such polyphenols, phytochemicals, and polyunsaturated fatty acids (PUFAs) [6]. It is also recognised that other microbial species have the potential to catabolise prebiotics, besides the classical examples of Bifidobacterium and Lactobacillus [6], courtesy of culture-independent techniques, such as 16S rRNA next generation sequencing and whole genome shotgun metagenomic sequencing which have provided taxonomic classification to identify microbial abundance/diversity and inferred or identified metabolic function [7].
Given that previous discussion regarding the prebiotic potential of seaweed components has focused solely upon the saccharolytic fermentation of complex polysaccharides and the physiological effects of short chain fatty acid metabolites (SCFAs) [3,8,9], scope exists to explore the prebiotic potential of other phytochemical components derived from seaweeds, namely polyphenols, carotenoids, and PUFAs, applicable to both human and animal health.
This review aims to provide an updated discussion regarding the fermentation and potential prebiotic effect of seaweed polysaccharides and oligosaccharides, based on recent evidence from in vitro fermentation studies and in vivo animal models, and to postulate how other seaweed phytochemicals, such as polyphenols, PUFAs, and carotenoids, may interact with the gut microbiota to manipulate microbial composition and/or function to elicit bioactivities pertained to a prebiotic. The latter provides new opportunities to complete prebiotic screening studies using in vitro techniques and pre-clinical animal models to understand how parent compound biotransformation into endogenously-derived or gut microbiota-derived metabolites impact bioaccessibility and bioavailability to influence gut microbial community structure and function, conducive to a prebiotic effect. Evidence from clinical trials with health-related end-points and mechanistic insight is imperative to substantiate health claims associated with a prebiotic effect.

2. Complex Polysaccharides

Seaweeds contain 2.97–71.4% complex polysaccharides [2,3], which include alginate, fucoidan, and laminarin in brown seaweeds; xylan and sulphated galactans, such as agar, carrageenan, and porphyran in red seaweeds; whilst ulvan and xylan are found in green seaweeds. The monosaccharide composition of the major brown, red, and green seaweed glycans are presented in Table 1, Table 2, Table 3, respectively. Whilst no human study to date has explored prebiotic sources from seaweeds, several in vitro studies [10,11,12,13], and in vivo animal studies [14,15], have explored the prebiotic potential seaweeds and their polysaccharide components.
Seaweed polysaccharides are atypical in structure to terrestrial glycans, and are understood to resist gastric acidity, host digestive enzymes, and gastrointestinal absorption [8]. Seaweed glycans may, therefore, serve as fermentation substrates for specific gut microbial populations or facilitate substrate cross-feeding of partially broken-down intermediates, such as oligosaccharides and metabolic cross-feeding of SCFAs to cause indirect proliferation of specific bacteria [16,17,18,19,20]. The physiological effects of SCFAs, primarily acetate, propionate, and butyrate, include the reduction of luminal pH to inhibit pathogens, the provision of energy sources to colonocytes, and the activation of free fatty acid receptors; where acetate and propionate are ligands for anorexigenic pathways in appetite regulation and can inhibit the rate limiting step of hepatic cholesterol synthesis via 3-hydroxy-3-methylglutaryl CoA reductase inhibition [21,22,23].
To facilitate saccharolytic fermentation in the colon, the gut microbiota must express functional carbohydrate active enzymes (CAZymes) to catabolise seaweed glycans as carbon sources within the colonic digesta. The repertoire of CAZymes expressed by the human gut microbiota includes glycoside hydrolase and polysaccharide lyase families to facilitate degradation via hydrolysis and elimination reactions, respectively [24,25,26]. Whole genome sequencing has previously identified gene clusters which encode the catabolic machinery responsible for the breakdown of prebiotics, which includes the CAZyme families responsible for the catabolism of inulin, lactulose, fructo-oligosaccharides, xylo-oligosaccharides, and galacto-oligosaccharides by human gut commensal strains, including Bifidobacterium longum NCC2705, Bifidobacterium adolescentis ATCC 15703, Streptococcus thermophilus LMD9, Eubacterium rectale ATCC 33656, Bacteroides vulgatus ATCC 8482, and Fecalibacterium prausnitzii KLE1255 [27].
Based on open source data from the Carbohydrate-Active enZYmes Database [28], Table 1, Table 2 and Table 3 detail the CAZyme families which may exert specificity for seaweed glycans and highlights the gut bacterial populations which have demonstrable evidence for seaweed glycan utilisation. This is dominated by Bacteroides, which have extensive glycolytic versatility [26,29]. This may explain why in vitro batch culture fermentation data of seaweeds and seaweed glycans indicate the proliferation of Bacteroides; whilst the degradation of complex seaweed glycans by Bacteroides could also facilitate the cross-feeding of oligosaccharides, monosaccharides, and SCFAs for gut commensals deemed beneficial to health, including Bifidobacterium.
In vitro fermentation studies are frequently used as screening tools to model colonic fermentation and determine substrate utilisation by an ex vivo faecal inoculum, with seaweed as a sole carbon source. An overview of recent in vitro fermentation studies which have evaluated the fermentation of whole seaweeds or extracted complex polysaccharide components by the human gut microbiota is presented in Table 4 (brown seaweeds), Table 5 (red seaweeds), and Table 6 (green seaweeds). These tables include differences in study methodologies, for example, test substrate dosage; the use of an in vitro digestion before the fermentation experiment (declared within the methods section of the cited research paper); how the inoculum was prepared; duration of the faecal fermentation experiment; microbial enumeration method; and the analytical technique used to ascertain metabolite changes during the fermentation. The use of an in vitro digestion before in vitro fermentation is often used to determine whether a substrate is resistant to endogenous digestive enzymes and small intestinal absorption, and to provide the fraction of a dietary component which is bioaccessible in the colon [30]. The lack of an in vitro digestion before fermentation experiments may cause false positive results, given that low molecular weight components present in seaweed extracts, normally absorbed in the small intestine, are used as fermentation substrates for the ex vivo microbiota. Table 7 highlights data from in vivo rodent studies which have evaluated the potential prebiotic effect of seaweeds and seaweed glycans.

2.1. Brown Seaweed Polysaccharides

Brown seaweeds are commonly used as food ingredients owing to their commercial abundance [53]. The anti-obesogenic effects of brown seaweeds are reported in mice, where supplementation of 5% (w/w) Saccorhiza polyschides extract, containing 12% dietary fibre, reduced body weight gain and fat mass of mice with diet-induced obesity [54]. The anti-obesogenic effect was attributed to the fermentation of alginate and fucoidan complex polysaccharide components, owing to reduced microbial bile salt hydrolase activity; however, no gut microbial compositional data were provided. Elsewhere, the in vitro evidence (Table 4) indicates that whole brown seaweeds and their extracted complex polysaccharide components are fermented by the ex vivo faecal microbiota, with increased production of acetate, propionate, butyrate, and total SCFAs reported during fermentation experiments. A corresponding increase in populations, such as Bifidobacterium, Bacteroides, Lactobacillus, Roseburia, Parasutterella, Fusicatenibacter, Coprococcus, Fecalibacterium is also reported [55,56,57].

2.1.1. Alginate

Alginates are composed of 1,4-linked α-l-guluronic (G) and β-d-mannuronic acid (M) residues to form GM, GG and MM blocks, and represent 17−45% dry weight of brown seaweeds [58]. The colloidal properties of alginates have wide application in food processing, biotechnology, medicine and pharmaceutical industries [59], while degraded sodium alginate is an approved item of “foods with specified uses”, under the categories of “Foods that act on cholesterol plus gastrointestinal conditions” and “Foods that act on blood cholesterol levels” in Japan [60]. The presence of water soluble alginate oligosaccharides in the faeces of pigs fed alginate is indicative of alginate lyase activity by the luminal or mucus adherent gut microbiota [61], although an adaptation period of > 39 days is reported for the degradation of G blocks by the porcine microbiota whilst M blocks are readily degraded [62].
The capacity for alginate to modulate the gut microbiota of Japanese individuals was highlighted over 20 years ago [63], where alginate supplementation (30 kDa, 10 g/day, n = 8) significantly increased faecal Bifidobacterium populations in healthy male volunteers after both one and two weeks, alongside significantly increased acetic and propionic acids after two weeks. Deleterious metabolites, including faecal sulphide, phenol, p-cresol, indole, ammonia and skatole were significantly reduced compared to the control (free living) diet. Notably, faecal Bifidobacterium counts and SCFA concentrations returned to baseline in the week after alginate diet cessation, which highlights the transient nature of the gut microbiota and the need for greater powered long-term human intervention studies.
Subsequent in vitro fermentation studies have indicated that alginate is fermented by the human gut microbiota, for example, a 24 h in vitro fermentation of a 212 kDa alginate increased total bacterial populations, although no statistical increase in individual Bifidobacterium, Bacteroides/Prevotella, Lactobacillus/Enterococcus, Eubacterium rectale/Clostridium coccoides, or Clostridium histolyticum populations were observed [11]. Acetic acid, propionic acid and total SCFAs were significantly increased after 24 h fermentation with the 212 kDa alginate, while alginate of 97 kDa increased total SCFA and acetate production after 10 h of fermentation. Alginates of 38 kDa, and 97 kDa did not change microbial abundance, although the authors could not correlate molecular weight with fermentation patterns.
Alginate oligosaccharides (AOS) (~3.5 kDa) can be obtained via acidic or enzymatic hydrolysis of alginate polysaccharides [58], and enzymatically derived AOS has promoted the growth of Bifidobacterium bifidum ATCC 29521, Bifidobacterium longum SMU 27001 and Lactobacilli, in vitro [64,65]. Supplementation of 2.5% AOS for two weeks significantly increased faecal Bifidobacterium. in rats compared to control and 5% FOS supplemented diets (13-fold and 4.7-fold increase, respectively), while faecal Lactobacillus were 5-fold greater in rats who consumed AOS compared to FOS. Enterobacteriaceae and Enterococcus populations were significantly decreased following AOS supplementation. Elsewhere, the hydrolysis of alginate, mannuronic acid oligosaccharides (MO) and guluronic oligosaccharides (GO) during a 48 h batch culture fermentation with the faecal microbiota of Chinese individuals demonstrated increased production of acetate, propionate, butyrate, and total SCFAs compared to the substrate-free control, where GO generated the greatest increase [36]. Subsequent strain isolation from the stools of individuals who demonstrated alginate degradation during fermentation identified Bacteroides xylanisovlens G25, Bacteroides thetaiotomicron A12, Bacteroides ovatus A9, and Bacteroides ovatus G19 as strains capable of hydrolysing alginate and AOS, where Bacteroides ovatus G19 expressed α-1,4-guluronanlyase and β-1,4-mannuronanlyase CAZymes [34].
A Bacteroides xylanisolvens strain with 99% similarity to Bacteroides xylanisolvens XB1A was recently isolated from the stool of a Chinese individual and the alginate lyase gene expressed was 100% homologous to the alginate lyase of Bacteroides ovatus strain ATCC 8483 [33]. The preceding in vitro fermentation study demonstrated increased production of acetate, propionate, butyrate, and total SCFAs compared to the soluble starch control vessel following a 72 h fermentation of alginate.
Alginate lyase depolymerises alginate polysaccharides to lower molecular weight oligosaccharides via β-elimination, and is most commonly expressed by marine bacteria, including Flammeovirga, Vibrio, Pseudoalteromonas, Glaciecola chathamensis S18K6, and Zobellia galactanivorans [65,66,67,68,69], while the terrestrial bacteria Paenibacillus sp. Strain MY03 was recently reported to possess genes encoding alginate lyase and agarase enzymes [70]. The acquisition of genes encoding alginate lyase enzymes by human gut Bacteroides is a suggested consequence of horizontal gene transfer from the marine environment [31,71], where seaweed consumption may have provided a vector to exert a selective pressure to induce diet-driven adaptations of the gut microbiota [35,72,73,74,75,76]. Recent work by Matthieu et al. [35] suggests that an alginate degradation system within the genome of human gut Bacteroides was a result of ancient acquisition, where the polysaccharide utilisation loci encodes PL6 and PL17 alginate lyase enzymes and hypothetical proteins responsible for alginate recognition, internalisation, and catabolism, including bacterial ABC transporter proteins to facilitate alginate uptake across the bacterial membrane [77]. Nevertheless, in vivo rodent studies have demonstrated that seaweed glycans are fermented even though animals have never been exposed to dietary seaweeds before the intervention, which suggests that the gut microbiome contains genes for CAZymes which can degrade seaweed glycans when expressed.

2.1.2. Laminarin

Laminarin is a water-soluble storage polysaccharide consisting of 1,3- or 1,6-β-glucose with an average molecular weight of 5 kDa [78] and accounts for 10–35% of the dry weight of brown seaweeds [58]. One in vitro batch culture fermentation of laminarin demonstrated increased Bifidobacteria and Bacteroides after 24 h [79], while another demonstrated increased propionate and butyrate production after 24 h [14]. A subsequent in vivo rat study (143 mg laminarin per kg body weight per day for 14 days) indicated that laminarin was not selectively fermented by Lactobacillus and Bifidobacterium, but could modify jejunal, ileal, caecal and colonic mucus composition, secretion, and metabolism to protect against bacterial translocation. The authors suggest that increased luminal acidity and/or catabolism of laminarin by mucolytic commensals could elicit such effects, which corroborates the evidence that a complex polysaccharide-rich diet maintains mucus layer integrity to promote gut barrier function [80,81]. Future studies regarding intestinal mucus modulation by laminarin may wish to characterise gut microbiota compositional and functional changes following laminarin ingestion, to detect the abundance and metabolic activity of glycan degraders, such as Bacteroides [82,83] or mucolytic species associated with health, such as Akkermansia muciniphila or Ruminococcus [84,85]. Elsewhere, laminarin increased L-cell GLP-1 secretion to attenuate diet-induced obesity in mice, and improved glucose homeostasis and insulin sensitivity [86]. The authors suggested that the observed cytosolic Ca2+ cascade caused GLP-1 secretion, which is in agreement with GPR41/43 receptor activation by SCFAs produced by gut microbial fermentation [87,88], however, data obtained to assess laminarin-induced changes to gut microbiota composition and metabolic output is needed to ascribe a prebiotic effect in this study.
The abundance of glycoside hydrolase and β-glucosidase enzymes expressed by the human gut microbiota may have the capacity to catabolise laminarin [24,89,90,91], for example, a Bacteroides cellulosyliticus WH2 human gut isolate was able to grow on laminarin-supplemented minimal media in vitro, (incidentally it did not grow on alginate, carrageenan, or porphyran) [92]; however, the molecular mechanisms by which human gut Bacteroides breakdown laminarin are likely distinct from those responsible for the degradation of mix linked β 1,3- 1,4- glucans, such as those found in cereals (e.g., by BoGH16MLG) [93].

2.1.3. Fucoidan

Fucoidans are water soluble polysaccharides composed of sulphated 1,2- or 1,3- or 1,4-α-l-fucose which exist as structural polysaccharides in brown seaweeds and occupy 5–20% of algal dry weight [58,94]. The structural heterogeneity of fucoidan encompasses varying degrees of branching, sulphate content, polydispersity, and irregular monomer patterns, which can include fucose, uronic acid, galactose, xylose, arabinose, mannose, and glucose residues [9,59,95].
A recent in vitro fermentation study of fucoidan (<30 kDa) extracted from Laminaria japonica demonstrated a greater increase in Bifidobacterium and Lactobacillus following 24 h and 48 h fermentation relative to >30 kDa fucoidan [12], while fucoidan from Ascophyllum nodosum (1330 kDa) and Laminaria japonica (310 kDa) were shown to increase Lactobacillus and Ruminococcaceae, respectively, in the caecal microbiota of mice gavaged with 100 mg/kg/day [96]. Fucoidan also reduced serum LPS-binding protein levels in this study—indicative of a reduced antigen load and reduced inflammatory response. In contrast, fucoidan with a fucose-rich and highly sulphated fucoidan extracted from Cladosiphon okamuranus was not fermented by the rat gut microbiota [97].
While the purported bioactivities of fucoidan include anti-obesogenic, anti-diabetic, anti-microbial, and anti-cancer properties [98], there is limited evidence to implicate a role for the gut microbiota with such bioactivities, and studies are needed to evaluate the structure-dependent fermentation of fucoidan to ascribe a prebiotic effect. For the latter, this is surprising given the myriad of α-fucosidase enzymes present in the human gut bacterial glycobiome.

2.2. Red Seaweed Polysaccharides

2.2.1. Galactans (Carrageenan, Agar, and Porphyran)

Red seaweeds, such as Gelidium spp. and Gracilaria spp., are used in the commercial production of agar and carrageenan food additives, including thickening, stabilizing and encapsulation agents [53]. A summary of evidence from recent in vitro fermentation experiments using red seaweed-derived substrates are presented in Table 5.
Carrageenans are composed of sulphated 1,4-β-d-galactose, 1,3-α-d-galactose, and 3,6-anhydro-d-galactose [43], and constitutes 30−75% dry weight of red seaweeds [58]. In rats fed 2.5% Chondrus crispus, of which carrageenan is a major polysaccharide component, faecal Bifidobacterium breve, and acetate, propionate, and butyrate SCFAs were significantly increased alongside a significant decrease in the pathogens Clostridium septicum and Streptococcus pneumonia, as compared to the basal diet [15]. Furthermore, a 1:1 mixture of polysaccharide extracts from Kappaphycus alvarezii (containing carrageenan) and Sargassum polycystum (brown seaweed) has lowered serum lipids in rats [39]. In a study by Li et al. [34], β-carrageenase activity in a Bacteroides uniforms 38F6 isolate complex of Bacteroides xylanisolvens and Escherichia coli hydrolysed κ-carrageenan oligosaccharides into 4-O-sulfate-d-galactose, κ-carratriose, κ-carrapentaose, and κ-carraheptaose, which could facilitate cross-feeding to promote the growth of Bifidobacterium populations.
Agar is composed of sulphated 1,3-β-d-galactose and 1,4- 3,6-anhydro-α-l-galactose [40] and can be fractionated into agarose and agaropectin [8]. Low molecular weight agar of 64.64 kDa has demonstrated a bifidogenic effect alongside increased acetate and propionate SCFA concentrations after 24 h in vitro fermentation with human stool inoculum [11], while mice fed with 2.5% (w/v) neoagarose oligosaccharides for 7 days demonstrated increased caecal and faecal Lactobacillus and Bifidobacterium [102]. The utilisation of agaro-oligosaccharides was noted in vitro by Bacteroides uniforms L8, isolated from Chinese individuals, which secreted a β-agarase CAZyme to breakdown agarooligosaccharides into agarotriose and subsequently facilitated the growth of Bifidobacterium infantis and Bifidobacterium adolescentis via the cross feeding of agarotriose [103].
Porphyran is made up of sulphated 1,3-β-d-galactose, 1,4-α-l-galactose-6-sulfate and 3,6-anhydro-α-l-galactose [41,104,105]. An in vitro faecal fermentation study indicated that porphyran did not significantly increase SCFAs, but stimulated Lactobacillus and Bacteroides populations [79]. While pure cultures of Bifidobacterium breve, Bifidobacterium longum, Bifidobacterium infantis, Bifidobacterium adolescentis, but not Bifidobacterium bifidum, were able to ferment dried Porphyra yezoensis (Nori), containing a low protein content (25%), whereas Nori with a high protein content (41%) was not fermented [105]. It is likely that carbohydrate content was highest in the low protein Nori, thus seasonal- and species-variation and in seaweed macronutrient content should be considered a determinant factor for the fermentability of whole seaweeds [106,107,108,109].
Evidence for the horizontal transfer of genes for porphyranase and agarase CAZymes from the marine bacteria, Zobellia galactanivorans, to Bacteroides plebeius of Japanese individuals is indicative of diet-driven adaptations of the human gut microbiome [41,44]; however, the North American counterparts in this study did not consume seaweeds and the gut microbiota of these individuals did not express such CAZymes. This may mean that the fermentation of seaweed polysaccharides, such as porphyran and agar, requires exposure to, and acquisition of, specific CAZymes usually present in the marine environment [73]. Red seaweed galactans are emerging prebiotic candidates given the commercial availability of red seaweed hydrocolloids and the potential gut modulatory effects of oligosaccharides obtained from red seaweeds. Nevertheless, further in vivo evidence is needed, given the purported pro-inflammatory effects of low molecular weight carrageenan [110,111,112].

2.2.2. Xylan

Xylan, composed of 1,3-1,4-β-D-xylose, is a major constituent of red seaweeds, such as Palmaria palmata [46]. A previous in vitro faecal fermentation study of xylan derived from P. palmata, reported that xylose was fermented after six h alongside a 58:28:14 ratio of acetate, propionate, and butyrate SCFAs (total SCFAs were 107 mM/L) [113]. This study did not ascertain bacterial compositional data, and thus a knowledge gap is presented given that xylans and xylooligosaccharides (XOS) extracted from terrestrial plants, such as wheat husks and maize, are mooted as potential prebiotics owing to evidence of bifidogenesis, improved plasma lipid profile, and positive modulation of immune function markers in healthy adults [114,115]. Given that human gut Bacteroides express a repertoire of xylanase and xylosidase CAZymes [116], investigations regarding the capacity of the human gut microbiota to catabolise red seaweed xylans and XOS are suggested.

2.3. Green Seaweed Polysaccharides

Ulvan

Ulvans are water-soluble cell wall polysaccharides that account for 8–29% dry weight of green seaweeds, and are composed of sulphated 1,3-α-l-rhamnose, 1,4-β-d-glucuronic acid, and 1,4-β-d-xyloglucan [51]. Previous reports indicate that Ulva lactuca and ulvan polysaccharides are poorly fermented by the human gut microbiota [8,95,118], while an in vitro fermentation study of Enteromorpha spp. With a human faecal inoculum reported no difference in Enterococcus, Lactobacillus, and Bifidobacterium populations compared to the control; only an increase in Enterobacter after 24 h and 48 h of fermentation (Table 6) [12]. In contrast, a recent in vitro faecal fermentation study indicated that Ulvan stimulated the growth of Bifidobacterium and Lactobacillus populations and promoted the production of lactate and acetate [79]. Further, a murine study showed that Enteromorpha (EP) and Enteromorpha polysaccharides (PEP) ameliorated inflammation associated with Loperamide-induced constipation in mice [119], where alpha diversity, Firmicutes, and Actinobacteria were increased in the faecal microbiota of seaweed-supplemented mice compared to the constipated control. Bacteroidetes and Proteobacteria were decreased, while Bacteroidales family S24-7 and Prevotellaceae were increased in EP and PEP, respectively. Current evidence for the fermentation of green seaweeds and their polysaccharides is limited and fermentation may require specific α-l-rhamnosidase activity by gut commensals [50]. More experimental evidence is needed to understand the impact of ulvans and ulvan-oligosaccharides in the human and animal diet.

2.4. Future Prospective–Obtaining Oligosaccharides

Enzyme technologies are reported to increase the extraction yield and reduce the molecular weight of bioactive components obtained from seaweeds, with examples of enhanced prebiotic activity when commercially available cellulases or seaweed-specific enzymes were used to hydrolyse polysaccharides [99,120]. Despite limited commercial availability of seaweed-specific enzymes, an avenue for functional oligosaccharide production is presented if efforts to develop commercially viable saccharolytic enzymes from microorganisms (primarily marine). Examples of such glycoside hydrolases include fucoidanase from Sphingomonas paucimobilis PF-1 [121]; ulvan lyase from Alteromonas spp. [122] and the family Flavobacteriaceae [123]; β-agarase from Cellulophaga omnivescoria W5C [124] and Cellvibrio PR1 [125]; alginate lyase from Flammeovirga [126], and Paenibacillus [70]; and laminarinase from Clostridiium thermocellum [127]. Factors which influence the stability and efficacy of such hydrolytic enzymes may include metal ion interaction, or thermostability at the high temperatures needed to prevent gelling of polysaccharides. Recent insight into the production of agarose oligosaccharides and neoagarose oligosaccharides from agar exemplify this [128].

3. Polyphenols

Seaweeds are rich in polyphenols, such as catechins, flavonols, and phlorotannins. Red and green seaweeds are a source of bromophenols, phenolic acids, and flavonoids [132], while phlorotannins are the most abundant polyphenol in brown seaweeds. Most research to date concerns the bioactivity of phlorotannins, a class of polyphenol unique to brown algae comprised of phloroglucinol monomers and categorised as eckols, fucols, fuhalols, ishofuhalols, phloroethols, or fucophloroethols [132]. The purported bioactivities of seaweed polyphenols are associated with the mitigation of risk factors pertained to type 2 diabetes and cardiovascular disease, including hyperglycemia, hyperlipidemia, inflammation and oxidative stress [133,134,135,136,137], and also anti-microbial activity [138]. Owing to heterogeneity in both molecular weight and the level of isomerisation, characterisation of polyphenols is difficult [139,140,141], and a paucity of information exists regarding the endogenous digestion and microbial catabolism of seaweed polyphenols, with a scarce mechanistic understanding of how they may exert health benefits via the gut microbiota.
Most polyphenols of plant origin must undergo intestinal biotransformation by endogenous enzymes and the gut microbiota prior to absorption across enterocytes. These enzymatic transformations include the elimination of glycosidic bonds, for example, flavonoids are converted to glycones (sugars) and aglycones (non-sugars–polyphenols) by endogenous β-glucosidases in the small intestine [142]. The transport of aglycones to the liver via the portal vein results in phase II biotransformation (coupling reactions, chiefly hepatic conjugation to O-glucuronides and O-sulfates) to facilitate urinary and biliary elimination. Phase II metabolites are absorbed into the systemic circulation, or excreted in bile and re-enter the duodenum (hepatic recycling), where subsequent glucuronidase, glycosidase, or sulphatase-mediated deconjugation by the colonic microbiota may favour aglycone reabsorption [143].
Approximately 90–95% of dietary polyphenols reach the colon intact [144], where biotransformation and metabolism by the gut microbiota occurs via hydrolysis, reduction, decarboxylation, demethylation, dehydroxylation, isomerisation, and fission [145], to produce low-molecular weight compounds with less chemical heterogeneity than the polyphenol parent compound [142]. It is suggested that a complex network of gut microbial species is necessary for full biotransformation of polyphenols, whereas simple reactions, such as deglycosylation, can be achieved by individual gut strains. Furthermore, the bioactivities associated with dietary polyphenol intake may be dependent on the catabolic capacity and composition of the gut microbiota, owing to the biological activity of metabolites rather than the parent polyphenol compound present in food [146,147], while a synergistic effect between prebiotic polyphenols and probiotic bacteria may occur [6].
The identification of bacteria which possess the metabolic capabilities to utilise polyphenols was previously identified in Eubacterium oxidoreducens, which could catabolise gallate, pyrogallol, phloroglucinol and quercetin [148]. Quercetin biotransformation by Eubacterium ramulus has also been identified [149], and multiple human gut microbes which possess phenolic enzymes capable of breaking down glycosides, glucuronides, sulphates, esters, and lactones was summarised by Selma et al. [145]. Such microorganisms included E. coli with β-glucuronidase activity; Eubacterium, Bacteroides, and Clostridium with β-glucosidase activity; Lactobacillus, Eubacterium, Clostridium , Butyrbacterium, Streptococcus, and Methylotrophicum with demethylase activity; and E. coli, Bifidobacterium, Lactobacillus, Bacteroides, Streptococcus , Ruminococcus, and Enterococcus with esterase activity. There is also evidence for α-l-Rhamnosidase mediated hydrolysis of rutinose, present on glycosylated polyphenols (rhamnoglycosides), to produce aglycones, by species, such as Bacteroides thetaiotaomicron [50], Bifidobacterium dentium [150], Bifidobacterium catenulatum [151], Bifidobacterium pseudocatenulatum [151], and Lactobacillus plantarum [152].
Current knowledge regarding the fate of seaweed polyphenols in the human gastrointestinal tract is scarce; however, it is understood that the limited absorption of Ascophyllum nodosum polyphenols from small intestinal enterocytes to the portal vein may facilitate the conjugation of polyphenols to methylated, glucuronidated, or sulphated forms rather than hydrolysis to aglycones [153,154]. Subsequently, unabsorbed conjugated polyphenols are available for biotransformation by the colonic microbiota, then potentially absorbed across the colonocytes. Indeed, Corona et al. [154], observed a reduction of total polyphenol contents of an Ascophyllum nodosum polyphenol extract, high molecular weight fraction (>10 kDa), and low molecular weight fraction (1–10 kDa) following in vitro digestion and batch culture fermentation; although anti-genotoxic activity against H2O2 induced DNA damage of HT-29 cells was increased (to a greater extent by the high molecular weight fraction). This study did not assess the microbiota composition, however, elsewhere, an in vitro fermentation of an Ecklonia radiata phlorotannin extract significantly increased Bacteroidetes, Clostridium coccoides, E. coli, and Fecalibacterium prausnitzii, but decreased Bifidobacterium and Lactobacillus populations after 24 h fermentation [56]. More in vitro digestion studies would be useful to understand the stability of seaweed polyphenols as extracts or within the seaweed matrix [155,156]. These studies may be complemented by studies which use ileostomy patient cohorts to determine structural changes to seaweed polyphenols following upper GI digestion in vivo to indicate polyphenol bioaccessibility in the colon [157].
A recent review highlighted the potential for dietary polyphenols to modulate the gut microbiota by increasing Bifidobacterium, Lactobacillus, Bacteroides, Enterococcus, Akkermansia muciniphila, and Fecalibacterium prausnitzii populations [158]. This review did not include any studies which assessed modulation of the gut microbiota by seaweed polyphenols and, therefore, a research opportunity is presented. Inter-individual variation of gut microbiota composition and function is the key determinant for gut microbiota-mediated biotransformation of phenolic compounds to bioactive metabolites [159,160]. Therefore, identification of bacterial species or strains with the ability to catabolise seaweed polyphenols and their respective catabolic machinery is needed to understand if seaweed polyphenols could be prebiotic [161,162]. Moreover, considering that gut microbiota-derived secondary metabolites reach a peak plasma concentration much later than the original aglycone or hepatic conjugates, controlled nutrikinetic studies could elude how dietary polyphenols from seaweeds interact with host-microbiota metabolism [163,164]. Identification of faecal, urinary, serum, or tissue biomarkers via untargeted and targeted metabolomics approaches, and the use of stable isotope studies, may also indicate variation in synthesis, bioavailability, metabolism, and excretion of polyphenols and associated metabolites. While integration of dose response studies alongside metagenomics and metabolomics analyses, akin to those conducted for berry and wine polyphenols, could elude how much seaweed polyphenol is required to have an impact, if any, on gut microbial composition, metabolic function, and host health [165,166].

4. Other Seaweed Phytochemicals

4.1. Carotenoids

Carotenoids are lipid soluble compounds which function within the photosynthetic machinery of seaweeds to produce pigments. Fucoxanthin is the predominant carotenoid in brown seaweeds [167], while lutein, β-carotene, astaxanthin, echinenone, violaxanthin, neoxanthin, and zeaxanthin are found in red and green seaweeds. Carotenoids are used as food colouring additives, while the application of fucoxanthin as functional food ingredients is suggested, owing to putative anti-oxidant, anti-inflammatory, anti-cancer, anti-obesity, and anti-diabetic bioactivities [168,169,170,171,172,173].
While some carotenoids are absorbed by enterocytes and converted into vitamin A and retinoid derivatives by endogenous beta-carotene oxygenase 1 (BCO1) and beta-carotene dioxygenase 2 (BCO2) enzymes [174,175], the bioavailability of carotenoids in the blood is reported as 10−40% [176], which has led to suggestions that carotenoids could be fermented by the gut microbiota [174,177]. The only evidence to date has demonstrated that male C57BL/6J mice supplemented with 0.04% (w/w) astaxanthin during an eight-week pilot study had increased abundance of caecal Bifidobacterium [178], whereas Proteobacteria and Bacteroides were significantly increased in the caecum of BCO2 knockout mice; however, analysis of health biomarkers was not reported. Given the differences in microbiota composition between wild type and BCO2 knockout mice in this study, there is scope to investigate how carotenoids and their endogenous derivatives interact with the gut microbiota. Looking ahead, the use of in vitro models of gastrointestinal digestion and colonic fermentation would be useful to assess whether there is a direct substrate to microbiota effect or a host–microbe effect [179].

4.2. Polyunsaturated Fatty Acids (PUFAs)

The lipid content of seaweed ranges from 1–5% dry weight, which includes n-3 PUFAs, such as eicosapentanoic acid (EPA) and docosahexaenoic acid (DHA) [180,181]. The n-3 PUFA are associated with the anti-inflammatory activity to reduce cardiovascular disease risk and may also exert beneficial effects on brain function and behaviour, as mediated by the microbiota-gut-brain axis [182]. Dietary EPA and DHA intake are reported to improve microbial diversity, reduce the Firmicutes/Bacteroidetes ratio, reduce LPS-producing bacteria, and increase populations of Bifidobacterium, Lachnospiraceae, and lipopolysaccharide (LPS)-suppressing bacteria in both humans and animal models [182,183,184]. Although the evidence to date has focused on fish-derived n-3 PUFA, great scope exists to evaluate the prebiotic effect of n-3 PUFA obtained from seaweeds.

5. Fermented Foods

Fermented foods are understood to have improved nutritional and functional properties owing to bioactive or bioavailable components [185]. Seaweeds (mainly kelp) are a common vegetable ingredient in the fermented food, Kimchi. The microbial content of kimchi provides a source of probiotics, nutrients, and bioactive metabolites, which are reported to have anti-microbial, anti-oxidant, and anti-obesogenic activities [186,187,188]. One randomised controlled trial (RCT) observed that consumption of a seaweed Kimchi made from L. japonica for four weeks promoted the growth and survival of gut microbial lactic acid bacteria in humans [189], whilst another RCT concluded that consumption of 1.5 g/day fermented L. japonica containing 5.56% γ-aminobutyric acid (GABA) (Lactobacillus brevis BJ2 culture) was associated with a reduction in oxidative stress in healthy adults over four weeks, indicated by decreased serum γ-glutamyltransferse (GGT) and malondialdehyde, and increased antioxidant activity of superoxide dismutase and catalase compared to the placebo [190]. The latter study indicates that foods containing fermented brown seaweeds, such as L. japonica, may offer a novel source of GABA enriched ingredients, which are associated with hypotensive and anti-inflammatory effects [188]. Anti-oxidant, anti-diabetic, and anti-hypertensive efficacies are also reported for Korean rice wine fermented with L. japonica [191], while Sargassum fermented with a starter culture of Enterococcus faecium was reported to contain higher soluble polyphenol and mannuronic acid-rich alginate contents [192], which may increase the provision of microbiota accessible components for colonic fermentation.
Reports of the functional properties of fermented foods containing red seaweeds are scarce; however, examples of red seaweed fermented foods include a fermented Porphyra yezoensis seaweed sauce, which used the marine halophilic lactic acid bacteria, Tetragenococcus halophilus, as a starter culture [193]; a Gracilaria domingensis aqueous extract applied as a texture modifier in fermented milks as a non-animal alternative to gelatin [194]; and carrageenan as a salt replacer in the production of fat-free cheese [195].
Given the availability of red, brown, and green seaweeds both commercially and locally [196], the production of seaweed-containing fermented foods could be a cost-effective alternative to bioactive component extraction. Nevertheless, an understanding of how live bacteria and bacterial metabolites present in fermented foods contribute towards health is required [185].

6. Seaweeds and Animal Health

Seaweeds also have a historical use as animal feed ingredients [197]. The capacity for seaweeds to modulate the gut microbiota of monogastrics, such as pigs and hens, is presented in Table 8 and Table 9, respectively, which complements the recent evidence for the application of seaweed bioactives in monogastric animal feed [198]. Table 8 shows limited evidence that the β-glucan, laminarin, may increase Lactobacillus populations but not Bifidobacterium populations. While there is scarce evidence for the selective stimulation of health-associated bacteria in pigs by the sulphated fucose, fucoidan. Only one recent study has evaluated the effect of dietary alginate on the porcine microbiota, where the genera Ruminococcus, Roseburia and Lachnospira, and an unclassified bacterium of the F16 family were increased, alongside a significant decrease in the genus Blautia, the family Clostridiaceae, and an unclassified bacterium of RF39 family [199]. In Table 9, recent evidence indicates that hens fed Chondrus crispus and Sarcodiotheca gaudichaudii red seaweeds may increase ceacal SCFA concentrations and modulate populations of Bifidobacterium longum, Lactobacillus acidophilus, Streptococcus salivarius, and Clostridium perfringens [200,201]; however, a bidirectional change in microbial composition was dose dependent and has only been assessed in two studies to date. Given the use of pigs as an animal model of humans [202], data from in vivo monogastric studies which are designed to evaluate the prebiotic potential of dietary seaweeds and seaweed-derived components could provide insight into the potential for human applications.
Table 10 and Table 11 summarise recent studies which have examined the impact of seaweed diets on the ruminant microbiota of cows and sheep, respectively, with the potential application of reducing methane production. Despite demonstrating decreased methane production, the cow rumen in vitro fermentation studies presented in Table 10 did not assess microbiota compositional changes, thus a knowledge gap is presented to understand which bacteria (if any), are increased or decreased, and are associated with a reduction in methane production. Table 11 shows that methanogenic bacteria and methane production were significantly decreased compared to the basal grass substrate control following the in vitro fermentation of sheep rumen with the red seaweed Asparagopsis taxiformis [203]. While sheep given an ad libitum diet of Ascophyllum nodosum brown seaweed (1%, 3%, or 5% w/w) for 21 days demonstrated a dose-dependent decrease in propionate and butyrate SCFAs and a dose-dependent increase in acetate [204], while several bacteria were significantly decreased, including Prevotella copri, Roseburia, and Coprococcus, while Blautia producta and the family Veillonellaceae were significantly increased compared to the basal diet. Moreover, the specific case of seaweed-fed North Ronaldsay sheep highlights how isolated organisms of the ruminant microbiome, such as Prevotella, Clostridium butyricum, Butyrivibrio fibrisolvens, and Spirochaetes have adapted to hydrolyse alginate laminarin, and fucoidan [205,206]. However, there is a paucity of evidence to implicate any health benefits attributed to a seaweed diet in these animals.

7. Conclusions

Current evidence regarding the prebiotic effects of seaweeds is dominated by complex polysaccharide components. This is because prebiotic research was previously focused on saccharolytic fermentation by the gut microbiota. Accumulating evidence from in vitro and in vivo animal studies provides encouraging data regarding the utilisation of red seaweed galactans and brown seaweed glycans, such as alginates and laminarins, with minor evidence for fucoidan and the green seaweed polysaccharide, ulvan.
Given that the most recent definition of prebiotic places non-complex polysaccharide components in vogue, an opportunity is presented to explore how other seaweed phytochemicals, including polyphenols, carotenoids, and PUFAs, are metabolised by host microbial populations to benefit host health. Future investigations should consider the use of in vitro screening studies and in vivo animal studies to identify putative prebiotic compounds from seaweeds via the identification of host organisms which utilise seaweed components and the bioactive metabolites produced (via untargeted metabolomics). Furthermore, controlled human intervention studies with health-related end points to elucidate prebiotic efficacy are required.

Author Contributions

Conceptualization, P.C., S.Y., C.R.S., C.S, P.J.A., E.M.M., R.P.R.; writing—original draft preparation, P.C., S.Y., and C.R.S X.X.; writing—review and editing, P.C., S.Y., C.R.S., C.S, P.J.A., E.M.M., and R.P.R.; funding acquisition, C.S, P.J.A., E.M.M., and R.P.R.

Funding

This research was funded by The Department of Agriculture Food and the Marine (FIRM) under the National Development Plan 2007–2013, Project number 13F511 (PREMARA); and Science Foundation of Ireland—funded Centre for Science, Engineering and Technology, and APC Microbiome Ireland. Paul Cherry is in receipt of a Department for Employment and Learning Northern Ireland Postgraduate studentship.

Acknowledgments

Please see the above funding section.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Brown, E.S.; Allsopp, P.J.; Magee, P.J.; Gill, C.I.; Nitecki, S.; Strain, C.R.; McSorley, E.M. Seaweed and human health. Nutr. Rev. 2014, 72, 205–216. [Google Scholar] [CrossRef] [PubMed]
  2. Cherry, P.; O’Hara, C.; Magee, P.J.; McSorley, E.M.; Allsopp, P.J. Risks and benefits of consuming edible seaweeds. Nutr. Rev. 2019. [Google Scholar] [CrossRef] [PubMed]
  3. De Jesus Raposo, M.F.; de Morais, A.M.; de Morais, R.M. Emergent sources of prebiotics: Seaweeds and microalgae. Mar. Drugs 2016, 14, 27. [Google Scholar] [CrossRef] [PubMed]
  4. Thursby, E.; Juge, N. Introduction to the human gut microbiota. Biochem. J. 2017, 474, 1823–1836. [Google Scholar] [CrossRef] [PubMed]
  5. Schippa, S.; Conte, M.P. Dysbiotic events in gut microbiota: Impact on human health. Nutrients 2014, 6, 5786–5805. [Google Scholar] [CrossRef] [PubMed]
  6. Gibson, G.R.; Hutkins, R.; Sanders, M.E.; Prescott, S.L.; Reimer, R.A.; Salminen, S.J.; Scott, K.; Stanton, C.; Swanson, K.S.; Cani, P.D.; et al. Expert consensus document: The International Scientific Association for Probiotics and Prebiotics (ISAPP) consensus statement on the definition and scope of prebiotics. Nat. Rev. Gastroenterol. Hepatol. 2017, 14, 491–502. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  7. Arnold, J.W.; Roach, J.; Azcarate-Peril, M.A. Emerging technologies for gut microbiome research. Trends Microbiol. 2016, 24, 887–901. [Google Scholar] [CrossRef]
  8. O’Sullivan, L.; Murphy, B.; McLoughlin, P.; Duggan, P.; Lawlor, P.G.; Hughes, H.; Gardiner, G.E. Prebiotics from marine macroalgae for human and animal health applications. Mar. Drugs 2010, 8, 2038–2064. [Google Scholar] [CrossRef]
  9. Zaporozhets, T.S.; Besednova, N.N.; Kuznetsova, T.A.; Zvyagintseva, T.N.; Makarenkova, I.D.; Kryzhanovsky, S.P.; Melnikov, V.G. The prebiotic potential of polysaccharides and extracts of seaweeds. Russ. J. Mar. Biol. 2014, 40, 1–9. [Google Scholar] [CrossRef]
  10. Devillé, C.; Damas, J.; Forget, P.; Dandrifosse, G.; Peulen, O. Laminarin in the dietary fibre concept. J. Sci. Food Agric. 2004, 84, 1030–1038. [Google Scholar] [CrossRef]
  11. Ramnani, P.; Chitarrari, R.; Tuohy, K.; Grant, J.; Hotchkiss, S.; Philp, K.; Campbell, R.; Gill, C.; Rowland, I. In vitro fermentation and prebiotic potential of novel low molecular weight polysaccharides derived from agar and alginate seaweeds. Anaerobe 2012, 18, 1–6. [Google Scholar] [CrossRef] [PubMed]
  12. Kong, Q.; Dong, S.; Gao, J.; Jiang, C. In vitro fermentation of sulfated polysaccharides from E. prolifera and L. japonica by human fecal microbiota. Int. J. Biol. Macromol. 2016, 91, 867–871. [Google Scholar] [CrossRef] [PubMed]
  13. Bajury, D.M.; Rawi, M.H.; Sazali, I.H.; Abdullah, A.; Sarbini, S.R. Prebiotic evaluation of red seaweed (Kappaphycus alvarezii) using in vitro colon model. Int. J. Food Sci. Nutr. 2017, 68, 821–828. [Google Scholar] [CrossRef] [PubMed]
  14. Devillé, C.; Gharbi, M.; Dandrifosse, G.; Peulen, O. Study on the effects of laminarin, a polysaccharide from seaweed, on gut characteristics. J. Sci. Food Agric. 2007, 87, 1717–1725. [Google Scholar] [CrossRef]
  15. Liu, J.; Kandasamy, S.; Zhang, J.; Kirby, C.W.; Karakach, T.; Hafting, J.; Critchley, A.T.; Evans, F.; Prithiviraj, B.J.B.C.; Medicine, A. Prebiotic effects of diet supplemented with the cultivated red seaweed Chondrus crispus or with fructo-oligo-saccharide on host immunity, colonic microbiota and gut microbial metabolites. BMC Complement. Altern. Med. 2015, 15, 279. [Google Scholar] [CrossRef] [PubMed]
  16. Rose, D.J.; Keshavarzian, A.; Patterson, J.A.; Venkatachalam, M.; Gillevet, P.; Hamaker, B.R. Starch-entrapped microspheres extend in vitro fecal fermentation, increase butyrate production, and influence microbiota pattern. Mol. Nutr. Food Res. 2009, 53, S121–S130. [Google Scholar] [CrossRef] [PubMed]
  17. Timm, D.A.; Stewart, M.L.; Hospattankar, A.; Slavin, J.L. Wheat dextrin, psyllium, and inulin produce distinct fermentation patterns, gas volumes, and short-chain fatty acid profiles in vitro. J. Med. Food 2010, 13, 961–966. [Google Scholar] [CrossRef]
  18. Belenguer, A.; Duncan, S.H.; Calder, A.G.; Holtrop, G.; Louis, P.; Lobley, G.E.; Flint, H.J. Two routes of metabolic cross-feeding between Bifidobacterium adolescentis and butyrate-producing anaerobes from the human gut. Appl. Environ. Microbiol. 2006, 72, 3593–3599. [Google Scholar] [CrossRef]
  19. Macfarlane, G.T.; Macfarlane, S. Bacteria, colonic fermentation, and gastrointestinal health. J. AOAC Int. 2012, 95, 50–60. [Google Scholar] [CrossRef]
  20. Ríos-Covián, D.; Ruas-Madiedo, P.; Margolles, A.; Gueimonde, M.; de los Reyes-Gavilán, C.G.; Salazar, N. Intestinal short chain fatty acids and their link with diet and human health. Front. Microbiol. 2016, 7, 185. [Google Scholar] [CrossRef]
  21. Byrne, C.S.; Chambers, E.S.; Morrison, D.J.; Frost, G. The role of short chain fatty acids in appetite regulation and energy homeostasis. Int. J. Obes. (2005) 2015, 39, 1331–1338. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  22. Gunness, P.; Gidley, M.J. Mechanisms underlying the cholesterol-lowering properties of soluble dietary fibre polysaccharides. Food Funct. 2010, 1, 149–155. [Google Scholar] [CrossRef] [PubMed]
  23. Den Besten, G.; van Eunen, K.; Groen, A.K.; Venema, K.; Reijngoud, D.-J.; Bakker, B.M. The role of short-chain fatty acids in the interplay between diet, gut microbiota, and host energy metabolism. J. Lipid Res. 2013, 54, 2325–2340. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  24. Tasse, L.; Bercovici, J.; Pizzut-Serin, S.; Robe, P.; Tap, J.; Klopp, C.; Cantarel, B.L.; Coutinho, P.M.; Henrissat, B.; Leclerc, M.; et al. Functional metagenomics to mine the human gut microbiome for dietary fiber catabolic enzymes. Genome Res. 2010, 20, 1605–1612. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  25. El Kaoutari, A.; Armougom, F.; Leroy, Q.; Vialettes, B.; Million, M.; Raoult, D.; Henrissat, B. Development and validation of a microarray for the investigation of the CAZymes encoded by the human gut microbiome. PLoS ONE 2013, 8, e84033. [Google Scholar] [CrossRef] [PubMed]
  26. Ndeh, D.; Gilbert, H.J. Biochemistry of complex glycan depolymerisation by the human gut microbiota. FEMS Microbiol. Rev. 2018, 42, 146–164. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  27. Cecchini, D.A.; Laville, E.; Laguerre, S.; Robe, P.; Leclerc, M.; Doré, J.; Henrissat, B.; Remaud-Siméon, M.; Monsan, P.; Potocki-Véronèse, G. Functional metagenomics reveals novel pathways of prebiotic breakdown by human gut bacteria. PLoS ONE 2013, 8, e72766. [Google Scholar] [CrossRef] [PubMed]
  28. Cantarel, B.L.; Coutinho, P.M.; Rancurel, C.; Bernard, T.; Lombard, V.; Henrissat, B. The Carbohydrate-Active EnZymes database (CAZy): An expert resource for Glycogenomics. Nucleic Acids Res. 2009, 37, D233–D238. [Google Scholar] [CrossRef]
  29. Benitez-Paez, A.; Gomez Del Pulgar, E.M.; Sanz, Y. the glycolytic versatility of bacteroides uniformis CECT 7771 and its genome response to oligo and polysaccharides. Front. Cell. Infect. Microbiol. 2017, 7, 383. [Google Scholar] [CrossRef]
  30. Brodkorb, A.; Egger, L.; Alminger, M.; Alvito, P.; Assunção, R.; Ballance, S.; Bohn, T.; Bourlieu-Lacanal, C.; Boutrou, R.; Carrière, F.; et al. INFOGEST static in vitro simulation of gastrointestinal food digestion. Nat. Protoc. 2019, 14, 991–1014. [Google Scholar] [CrossRef]
  31. Thomas, F.; Barbeyron, T.; Tonon, T.; Genicot, S.; Czjzek, M.; Michel, G. Characterization of the first alginolytic operons in a marine bacterium: From their emergence in marine Flavobacteriia to their independent transfers to marine Proteobacteria and human gut Bacteroides. Environ. Microbiol. 2012, 14, 2379–2394. [Google Scholar] [CrossRef] [PubMed]
  32. Brownlee, I.A.; Allen, A.; Pearson, J.P.; Dettmar, P.W.; Havler, M.E.; Atherton, M.R.; Onsoyen, E. Alginate as a source of dietary fiber. Crit. Rev. Food Sci. Nutr. 2005, 45, 497–510. [Google Scholar] [CrossRef] [PubMed]
  33. Bai, S.; Chen, H.; Zhu, L.; Liu, W.; Yu, H.D.; Wang, X.; Yin, Y. Comparative study on the in vitro effects of Pseudomonas aeruginosa and seaweed alginates on human gut microbiota. PLoS ONE 2017, 12, e0171576. [Google Scholar] [CrossRef] [PubMed]
  34. Li, M.; Shang, Q.; Li, G.; Wang, X.; Yu, G. Degradation of marine algae-derived carbohydrates by bacteroidetes isolated from human gut microbiota. Mar. Drugs 2017, 15, 92. [Google Scholar] [CrossRef] [PubMed]
  35. Mathieu, S.; Touvrey-Loiodice, M.; Poulet, L.; Drouillard, S.; Vincentelli, R.; Henrissat, B.; Skjåk-Bræk, G.; Helbert, W. Ancient acquisition of “alginate utilization loci” by human gut microbiota. Sci. Rep. 2018, 8, 8075. [Google Scholar] [CrossRef] [PubMed]
  36. Li, M.; Li, G.; Shang, Q.; Chen, X.; Liu, W.; Pi, X.; Zhu, L.; Yin, Y.; Yu, G.; Wang, X. In vitro fermentation of alginate and its derivatives by human gut microbiota. Anaerobe 2016, 39, 19–25. [Google Scholar] [CrossRef] [PubMed]
  37. Zhang, J.J.; Zhang, Q.B.; Wang, J.; Shi, X.L.; Zhang, Z.S. Analysis of the monosaccharide composition of fucoidan by precolumn derivation HPLC. Chin. J. Oceanol. Limnol. 2009, 27, 578–582. [Google Scholar] [CrossRef]
  38. Salyers, A.A.; Palmer, J.K.; Wilkins, T.D. Laminarinase (beta-glucanase) activity in Bacteroides from the human colon. Appl. Environ. Microbiol. 1977, 33, 1118–1124. [Google Scholar] [Green Version]
  39. Dousip, A.; Matanjun, P.; Sulaiman, M.R.; Tan, T.S.; Ooi, Y.B.H.; Lim, T.P.J.J.o.A.P. Effect of seaweed mixture intake on plasma lipid and antioxidant profile of hyperholesterolaemic rats. J. Appl. Phycol. 2014, 26, 999–1008. [Google Scholar] [CrossRef]
  40. Lahaye, M.; Rochas, C. Chemical-structure and physicochemical properties of agar. Hydrobiologia 1991, 221, 137–148. [Google Scholar] [CrossRef]
  41. Hehemann, J.H.; Correc, G.; Barbeyron, T.; Helbert, W.; Czjzek, M.; Michel, G. Transfer of carbohydrate-active enzymes from marine bacteria to Japanese gut microbiota. Nature 2010, 464, 908–912. [Google Scholar] [CrossRef] [PubMed]
  42. Rebuffet, E.; Groisillier, A.; Thompson, A.; Jeudy, A.; Barbeyron, T.; Czjzek, M.; Michel, G. Discovery and structural characterization of a novel glycosidase family of marine origin. Environ. Microbiol. 2011, 13, 1253–1270. [Google Scholar] [CrossRef] [PubMed]
  43. Weiner, M.L. Food additive carrageenan: Part II: A critical review of carrageenan in vivo safety studies. Crit. Rev. Toxicol. 2014, 44, 244–269. [Google Scholar] [CrossRef] [PubMed]
  44. Hehemann, J.H.; Kelly, A.G.; Pudlo, N.A.; Martens, E.C.; Boraston, A.B. Bacteria of the human gut microbiome catabolize red seaweed glycans with carbohydrate-active enzyme updates from extrinsic microbes. Proc. Natl. Acad. Sci. USA 2012, 109, 19786–19791. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Zhang, Q.B.; Qi, H.M.; Zhao, T.T.; Deslandes, E.; Ismaeli, N.M.; Molloy, F.; Critchley, A.T. Chemical characteristics of a polysaccharide from Porphyra capensis (Rhodophyta). Carbohydr. Res. 2005, 340, 2447–2450. [Google Scholar] [CrossRef] [PubMed]
  46. Usov, A.I. Polysaccharides of the Red Algae. Adv. Carbohydr. Chem. Biochem. 2011, 65, 115–217. [Google Scholar] [CrossRef] [PubMed]
  47. Mirande, C.; Kadlecikova, E.; Matulova, M.; Capek, P.; Bernalier-Donadille, A.; Forano, E.; Bera-Maillet, C. Dietary fibre degradation and fermentation by two xylanolytic bacteria Bacteroides xylanisolvens XB1AT and Roseburia intestinalis XB6B4 from the human intestine. J. Appl. Microbiol. 2010, 109, 451–460. [Google Scholar] [CrossRef]
  48. Hong, P.Y.; Iakiviak, M.; Dodd, D.; Zhang, M.L.; Mackie, R.I.; Cann, I. Two new xylanases with different substrate specificities from the human gut bacterium bacteroides intestinalis DSM 17393. Appl. Environ. Microbiol. 2014, 80, 2084–2093. [Google Scholar] [CrossRef]
  49. Despres, J.; Forano, E.; Lepercq, P.; Comtet-Marre, S.; Jubelin, G.; Chambon, C.; Yeoman, C.J.; Miller, M.E.B.; Fields, C.J.; Martens, E.; et al. Xylan degradation by the human gut Bacteroides xylanisolvens XB1A(T) involves two distinct gene clusters that are linked at the transcriptional level. BMC Genom. 2016, 17. [Google Scholar] [CrossRef]
  50. Munoz-Munoz, J.; Cartmell, A.; Terrapon, N.; Henrissat, B.; Gilbert, H.J. Unusual active site location and catalytic apparatus in a glycoside hydrolase family. Proc. Natl. Acad. Sci. USA 2017, 114, 4936–4941. [Google Scholar] [CrossRef] [Green Version]
  51. Lahaye, M.; Robic, A. Structure and functional properties of ulvan, a polysaccharide from green seaweeds. Biomacromolecules 2007, 8, 1765–1774. [Google Scholar] [CrossRef] [PubMed]
  52. Liang, W.-S.; Liu, T.C.; Chang, C.-J.; Pan, C.-L. Bioactivity of β-1,3-xylan Extracted from Caulerpa lentillifera by Using Escherichia coli ClearColi BL21(DE3)-β-1,3-xylanase XYLII. J. Food Nutr. Res. 2015, 3, 437–444. [Google Scholar] [CrossRef]
  53. Usman, A.; Khalid, S.; Usman, A.; Hussain, Z.; Wang, Y. Chapter 5—algal polysaccharides, novel application, and outlook. In Algae Based Polymers, Blends, and Composites, 1st ed.; Zia, K.M., Zuber, M., Ali, M., Eds.; Elsevier: Amsterdam, The Netherlands, 2017; Volume 1, pp. 115–153. [Google Scholar] [CrossRef]
  54. Huebbe, P.; Nikolai, S.; Schloesser, A.; Herebian, D.; Campbell, G.; Glüer, C.-C.; Zeyner, A.; Demetrowitsch, T.; Schwarz, K.; Metges, C.C.; et al. An extract from the Atlantic brown algae Saccorhiza polyschides counteracts diet-induced obesity in mice via a gut related multi-factorial mechanisms. Oncotarget 2017, 8, 73501–73515. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Fu, X.; Cao, C.; Ren, B.; Zhang, B.; Huang, Q.; Li, C. Structural characterization and in vitro fermentation of a novel polysaccharide from Sargassum thunbergii and its impact on gut microbiota. Carbohydr. Polym. 2018, 183, 230–239. [Google Scholar] [CrossRef] [PubMed]
  56. Charoensiddhi, S.; Conlon, M.A.; Vuaran, M.S.; Franco, C.M.M.; Zhang, W. Polysaccharide and phlorotannin-enriched extracts of the brown seaweed Ecklonia radiata influence human gut microbiota and fermentation in vitro. J. Appl. Phycol. 2017, 29, 2407–2416. [Google Scholar] [CrossRef]
  57. Charoensiddhi, S.; Conlon, M.A.; Vuaran, M.S.; Franco, C.M.M.; Zhang, W. Impact of extraction processes on prebiotic potential of the brown seaweed Ecklonia radiata by in vitro human gut bacteria fermentation. J. Funct. Foods 2016, 24, 221–230. [Google Scholar] [CrossRef]
  58. Vera, J.; Castro, J.; Gonzalez, A.; Moenne, A. Seaweed polysaccharides and derived oligosaccharides stimulate defense responses and protection against pathogens in plants. Mar. Drugs 2011, 9, 2514–2525. [Google Scholar] [CrossRef] [PubMed]
  59. García-Ríos, V.; Ríos-Leal, E.; Robledo, D.; Freile-Pelegrin, Y. Polysaccharides composition from tropical brown seaweeds. Phycol. Res. 2012, 60, 305–315. [Google Scholar] [CrossRef]
  60. Maeda-Yamamoto, M. Development of functional agricultural products and use of a new health claim system in Japan. Trends Food Sci. Technol. 2017, 69, 324–332. [Google Scholar] [CrossRef]
  61. Jonathan, M.C.; Bosch, G.; Schols, H.A.; Gruppen, H. Separation and identification of individual alginate oligosaccharides in the feces of alginate-fed pigs. J. Agric. Food Chem. 2013, 61, 553–560. [Google Scholar] [CrossRef]
  62. Jonathan, M.; Souza da Silva, C.; Bosch, G.; Schols, H.; Gruppen, H. In vivo degradation of alginate in the presence and in the absence of resistant starch. Food Chem. 2015, 172, 117–120. [Google Scholar] [CrossRef] [PubMed]
  63. Terada, A.; Hara, H.; Mitsuoka, T. Effect of dietary alginate on the fecal microbiota and fecal metabolic activity in humans. Microb. Ecol. Health Dis. 1995, 8, 259–266. [Google Scholar] [CrossRef]
  64. Wang, Y.; Han, F.; Hu, B.; Li, J.; Yu, W. In vivo prebiotic properties of alginate oligosaccharides prepared through enzymatic hydrolysis of alginate. Nutr. Res. 2006, 26, 597–603. [Google Scholar] [CrossRef]
  65. Han, W.; Gu, J.; Cheng, Y.; Liu, H.; Li, Y.; Li, F. Novel Alginate Lyase (Aly5) from a Polysaccharide-degrading marine bacterium, Flammeovirga sp. Strain MY04: Effects of module truncation on biochemical characteristics, alginate degradation patterns, and oligosaccharide-yielding properties. Appl. Environ. Microbiol. 2016, 82, 364–374. [Google Scholar] [CrossRef] [PubMed]
  66. Chen, X.-L.; Dong, S.; Xu, F.; Dong, F.; Li, P.-Y.; Zhang, X.-Y.; Zhou, B.-C.; Zhang, Y.-Z.; Xie, B.-B. Characterization of a New Cold-Adapted and Salt-Activated Polysaccharide Lyase Family 7 Alginate Lyase from Pseudoalteromonas sp. SM0524. Front. Microbiol. 2016, 7, 1120. [Google Scholar] [CrossRef]
  67. Dong, Q.; Ruan, L.; Shi, H. Genome sequence of a high agarase-producing strain Flammeovirga sp. SJP92. Stand. Genom. Sci. 2017, 12, 13. [Google Scholar] [CrossRef] [Green Version]
  68. Xu, F.; Dong, F.; Wang, P.; Cao, H.Y.; Li, C.Y.; Li, P.Y.; Pang, X.H.; Zhang, Y.Z.; Chen, X.L. Novel molecular insights into the catalytic mechanism of marine bacterial alginate lyase AlyGC from polysaccharide lyase family 6. J. Biol. Chem. 2017, 292, 4457–4468. [Google Scholar] [CrossRef]
  69. Zhu, B.; Sun, Y.; Ni, F.; Ning, L.; Yao, Z. Characterization of a new endo-type alginate lyase from Vibrio sp. NJU-03. Int. J. Biol. Macromol. 2018, 108, 1140–1147. [Google Scholar] [CrossRef]
  70. Liu, H.; Cheng, Y.; Gu, J.; Wang, Y.; Li, J.; Li, F.; Han, W. Draft genome sequence of Paenibacillus sp. Strain MY03, a terrestrial bacterium capable of degrading multiple marine-derived polysaccharides. Genome Announc. 2017, 5. [Google Scholar] [CrossRef]
  71. Mathieu, S.; Henrissat, B.; Labre, F.; Skjåk-Bræk, G.; Helbert, W. Functional exploration of the polysaccharide lyase family PL6. PLoS ONE 2016, 11, e0159415. [Google Scholar] [CrossRef]
  72. Cantarel, B.L.; Lombard, V.; Henrissat, B. Complex carbohydrate utilization by the healthy human microbiome. PLoS ONE 2012, 7, e28742. [Google Scholar] [CrossRef] [PubMed]
  73. Hehemann, J.-H.; Boraston, A.B.; Czjzek, M. A sweet new wave: Structures and mechanisms of enzymes that digest polysaccharides from marine algae. Curr. Opin. Struct. Biol. 2014, 28, 77–86. [Google Scholar] [CrossRef] [PubMed]
  74. Martin, M.; Barbeyron, T.; Martin, R.; Portetelle, D.; Michel, G.; Vandenbol, M. The cultivable surface microbiota of the brown alga Ascophyllum nodosum is enriched in macroalgal-polysaccharide-degrading bacteria. Front Microbiol 2015, 6, 1487. [Google Scholar] [CrossRef] [PubMed]
  75. Singh, R.P.; Reddy, C.R.K. Unraveling the functions of the macroalgal microbiome. Front. Microbiol. 2015, 6, 1488. [Google Scholar] [CrossRef] [PubMed]
  76. Bhattacharya, T.; Ghosh, T.S.; Mande, S.S. global profiling of carbohydrate active enzymes in human gut microbiome. PLoS ONE 2015, 10, e0142038. [Google Scholar] [CrossRef] [PubMed]
  77. Maruyama, Y.; Itoh, T.; Kaneko, A.; Nishitani, Y.; Mikami, B.; Hashimoto, W.; Murata, K. Structure of a Bacterial ABC Transporter Involved in the Import of an Acidic Polysaccharide Alginate. Structure 2015, 23, 1643–1654. [Google Scholar] [CrossRef] [Green Version]
  78. Kadam, S.U.; O’Donnell, C.P.; Rai, D.K.; Hossain, M.B.; Burgess, C.M.; Walsh, D.; Tiwari, B.K. Laminarin from Irish Brown Seaweeds Ascophyllum nodosum and Laminaria hyperborea: Ultrasound Assisted Extraction, Characterization and Bioactivity. Mar. Drugs 2015, 13, 4270–4280. [Google Scholar] [CrossRef]
  79. Seong, H.; Bae, J.-H.; Seo, J.S.; Kim, S.-A.; Kim, T.-J.; Han, N.S. Comparative analysis of prebiotic effects of seaweed polysaccharides laminaran, porphyran, and ulvan using in vitro human fecal fermentation. J. Funct. Foods 2019, 57, 408–416. [Google Scholar] [CrossRef]
  80. Brownlee, I.A.; Havler, M.E.; Dettmar, P.W.; Allen, A.; Pearson, J.P. Colonic mucus: Secretion and turnover in relation to dietary fibre intake. Proc. Nutr. Soc. 2003, 62, 245–249. [Google Scholar] [CrossRef]
  81. Desai, M.S.; Seekatz, A.M.; Koropatkin, N.M.; Kamada, N.; Hickey, C.A.; Wolter, M.; Pudlo, N.A.; Kitamoto, S.; Terrapon, N.; Muller, A.; et al. A dietary fiber-deprived gut microbiota degrades the colonic mucus barrier and enhances pathogen susceptibility. Cell 2016, 167, 1339–1353. [Google Scholar] [CrossRef]
  82. Salyers, A.A.; Vercellotti, J.R.; West, S.E.; Wilkins, T.D. Fermentation of mucin and plant polysaccharides by strains of Bacteroides from the human colon. Appl. Environ. Microbiol. 1977, 33, 319–322. [Google Scholar] [Green Version]
  83. Salyers, A.A.; West, S.E.; Vercellotti, J.R.; Wilkins, T.D. Fermentation of mucins and plant polysaccharides by anaerobic bacteria from the human colon. Appl. Environ. Microbiol. 1977, 34, 529–533. [Google Scholar] [PubMed]
  84. Tailford, L.E.; Crost, E.H.; Kavanaugh, D.; Juge, N. Mucin glycan foraging in the human gut microbiome. Front. Genet. 2015, 6, 81. [Google Scholar] [CrossRef] [PubMed]
  85. Dao, M.C.; Everard, A.; Aron-Wisnewsky, J.; Sokolovska, N.; Prifti, E.; Verger, E.O.; Kayser, B.D.; Levenez, F.; Chilloux, J.; Hoyles, L.; et al. Akkermansia muciniphila and improved metabolic health during a dietary intervention in obesity: Relationship with gut microbiome richness and ecology. Gut 2016, 65, 426. [Google Scholar] [CrossRef] [PubMed]
  86. Yang, L.; Wang, L.; Zhu, C.; Wu, J.; Yuan, Y.; Yu, L.; Xu, Y.; Xu, J.; Wang, T.; Liao, Z.; et al. Laminarin counteracts diet-induced obesity associated with glucagon-like peptide-1 secretion. Oncotarget 2017, 8, 99470–99481. [Google Scholar] [CrossRef]
  87. Tolhurst, G.; Heffron, H.; Lam, Y.S.; Parker, H.E.; Habib, A.M.; Diakogiannaki, E.; Cameron, J.; Grosse, J.; Reimann, F.; Gribble, F.M. Short-chain fatty acids stimulate glucagon-like peptide-1 secretion via the G-protein-coupled receptor FFAR2. Diabetes 2012, 61, 364–371. [Google Scholar] [CrossRef]
  88. Everard, A.; Cani, P.D. Gut microbiota and GLP-1. Rev. Endocr. Metab. Disord. 2014, 15, 189–196. [Google Scholar] [CrossRef]
  89. Dabek, M.; McCrae, S.I.; Stevens, V.J.; Duncan, S.H.; Louis, P. Distribution of beta-glucosidase and beta-glucuronidase activity and of beta-glucuronidase gene gus in human colonic bacteria. FEMS Microbiol. Ecol. 2008, 66, 487–495. [Google Scholar] [CrossRef]
  90. Gloux, K.; Berteau, O.; El oumami, H.; Béguet, F.; Leclerc, M.; Doré, J. A metagenomic β-glucuronidase uncovers a core adaptive function of the human intestinal microbiome. Proc. Natl. Acad. Sci. USA 2011, 108, 4539–4546. [Google Scholar] [CrossRef]
  91. Michalska, K.; Tan, K.; Li, H.; Hatzos-Skintges, C.; Bearden, J.; Babnigg, G.; Joachimiak, A. GH1-family 6-P-β-glucosidases from human microbiome lactic acid bacteria. Acta Crystallogr. Sect. D Biol. Crystallogr. 2013, 69, 451–463. [Google Scholar] [CrossRef]
  92. McNulty, N.P.; Wu, M.; Erickson, A.R.; Pan, C.; Erickson, B.K.; Martens, E.C.; Pudlo, N.A.; Muegge, B.D.; Henrissat, B.; Hettich, R.L.; et al. Effects of diet on resource utilization by a model human gut microbiota containing Bacteroides cellulosilyticus WH2, a symbiont with an extensive glycobiome. PLoS Biol. 2013, 11, e1001637. [Google Scholar] [CrossRef] [PubMed]
  93. Tamura, K.; Hemsworth, G.R.; Déjean, G.; Rogers, T.E.; Pudlo, N.A.; Urs, K.; Jain, N.; Davies, G.J.; Martens, E.C.; Brumer, H. Molecular mechanism by which prominent human gut bacteroidetes utilize mixed-linkage beta-glucans, major health-promoting cereal polysaccharides. Cell Rep. 2017, 21, 417–430. [Google Scholar] [CrossRef] [PubMed]
  94. Li, B.; Lu, F.; Wei, X.; Zhao, R. Fucoidan: Structure and Bioactivity. Molecules 2008, 13, 1671–1695. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Jiao, G.; Yu, G.; Zhang, J.; Ewart, H.S. Chemical structures and bioactivities of sulfated polysaccharides from marine algae. Mar. Drugs 2011, 9, 196–223. [Google Scholar] [CrossRef] [PubMed]
  96. Shang, Q.; Shan, X.; Cai, C.; Hao, J.; Li, G.; Yu, G. Dietary fucoidan modulates the gut microbiota in mice by increasing the abundance of Lactobacillus and Ruminococcaceae. Food Funct. 2016, 7, 3224–3232. [Google Scholar] [CrossRef]
  97. An, C.; Yazaki, T.; Takahashi, H.; Kuda, T.; Kimura, B. Diet-induced changes in alginate- and laminaran-fermenting bacterial levels in the caecal contents of rats. J. Funct. Foods 2013, 5, 389–394. [Google Scholar] [CrossRef]
  98. Collins, K.G.; Fitzgerald, G.F.; Stanton, C.; Ross, R.P. Looking beyond the terrestrial: The potential of seaweed derived bioactives to treat non-communicable diseases. Mar. Drugs 2016, 14. [Google Scholar] [CrossRef]
  99. Rodrigues, D.; Walton, G.; Sousa, S.; Rocha-Santos, T.A.P.; Duarte, A.C.; Freitas, A.C.; Gomes, A.M.P. In vitro fermentation and prebiotic potential of selected extracts from seaweeds and mushrooms. LWT Food Sci. Technol. 2016, 73, 131–139. [Google Scholar] [CrossRef]
  100. Chen, L.; Xu, W.; Chen, D.; Chen, G.; Liu, J.; Zeng, X.; Shao, R.; Zhu, H. Digestibility of sulfated polysaccharide from the brown seaweed Ascophyllum nodosum and its effect on the human gut microbiota in vitro. Int. J. Biol. Macromol. 2018, 112, 1055–1061. [Google Scholar] [CrossRef]
  101. Strain, C.R.; Collins, K.C.; Naughton, V.; McSorley, E.M.; Stanton, C.; Smyth, T.J.; Soler-Vila, A.; Rea, M.C.; Ross, P.R.; Cherry, P.; et al. Effects of a polysaccharide-rich extract derived from Irish-sourced Laminaria digitata on the composition and metabolic activity of the human gut microbiota using an in vitro colonic model. Eur. J. Nutr. 2019. [Google Scholar] [CrossRef]
  102. Hu, B.; Gong, Q.; Wang, Y.; Ma, Y.; Li, J.; Yu, W. Prebiotic effects of neoagaro-oligosaccharides prepared by enzymatic hydrolysis of agarose. Anaerobe 2006, 12, 260–266. [Google Scholar] [CrossRef] [PubMed]
  103. Li, M.; Li, G.; Zhu, L.; Yin, Y.; Zhao, X.; Xiang, C.; Yu, G.; Wang, X. Isolation and characterization of an agaro-oligosaccharide (AO)-hydrolyzing bacterium from the gut microflora of Chinese individuals. PLoS ONE 2014, 9, e91106. [Google Scholar] [CrossRef] [PubMed]
  104. Zhang, Q.; Li, N.; Liu, X.; Zhao, Z.; Li, Z.; Xu, Z. The structure of a sulfated galactan from Porphyra haitanensis and its in vivo antioxidant activity. Carbohydr. Res. 2004, 339, 105–111. [Google Scholar] [CrossRef] [PubMed]
  105. Muraoka, T.; Ishihara, K.; Oyamada, C.; Kunitake, H.; Hirayama, I.; Kimura, T. Fermentation Properties of Low-Quality Red Alga Susabinori Porphyra yezoensis by Intestinal Bacteria. Biosci. Biotechnol. Biochem. 2008, 72, 1731–1739. [Google Scholar] [CrossRef] [PubMed]
  106. Rioux, L.-E.; Turgeon, S.L.; Beaulieu, M. Effect of season on the composition of bioactive polysaccharides from the brown seaweed Saccharina longicruris. Phytochemistry 2009, 70, 1069–1075. [Google Scholar] [CrossRef] [PubMed]
  107. Kravchenko, A.O.; Byankina Barabanova, A.O.; Glazunov, V.P.; Yakovleva, I.M.; Yermak, I.M. Seasonal variations in a polysaccharide composition of Far Eastern red seaweed Ahnfeltiopsis flabelliformis (Phyllophoraceae). J. Appl. Phycol. 2018, 30, 535–545. [Google Scholar] [CrossRef]
  108. Skriptsova, A.V. Seasonal variations in the fucoidan content of brown algae from Peter the Great Bay, Sea of Japan. Russ. J. Mar. Biol. 2016, 42, 351–356. [Google Scholar] [CrossRef]
  109. Medcalf, D.G.; Lionel, T.; Brannon, J.H.; Scott, J.R. Seasonal variation in the mucilaginous polysaccharides from Ulva lactuca. Bot. Mar. 1975, 18, 67–70. [Google Scholar] [CrossRef]
  110. Shang, Q.; Sun, W.; Shan, X.; Jiang, H.; Cai, C.; Hao, J.; Li, G.; Yu, G. Carrageenan-induced colitis is associated with decreased population of anti-inflammatory bacterium, Akkermansia muciniphila, in the gut microbiota of C57BL/6J mice. Toxicol. Lett. 2017, 279, 87–95. [Google Scholar] [CrossRef]
  111. Bhattacharyya, S.; Liu, H.; Zhang, Z.; Jam, M.; Dudeja, P.K.; Michel, G.; Linhardt, R.J.; Tobacman, J.K. Carrageenan-induced innate immune response is modified by enzymes that hydrolyze distinct galactosidic bonds. J. Nutr. Biochem. 2010, 21, 906–913. [Google Scholar] [CrossRef] [Green Version]
  112. Younes, M.; Aggett, P.; Aguilar, F.; Crebelli, R.; Filipič, M.; Frutos, M.J.; Galtier, P.; Gott, D.; Gundert-Remy, U.; Kuhnle, K.K.; et al. Re-evaluation of carrageenan (E 407) and processed Eucheuma seaweed (E 407a) as food additives. EFSA J. 2018, 16, e05238. [Google Scholar] [CrossRef]
  113. Lahaye, M.; Michel, C.; Barry, J.L. Chemical, physicochemical and in-vitro fermentation characteristics of dietary fibres from Palmaria palmata (L.) Kuntze. Food Chem. 1993, 47, 29–36. [Google Scholar] [CrossRef]
  114. Childs, C.E.; Roytio, H.; Alhoniemi, E.; Fekete, A.A.; Forssten, S.D.; Hudjec, N.; Lim, Y.N.; Steger, C.J.; Yaqoob, P.; Tuohy, K.M.; et al. Xylo-oligosaccharides alone or in synbiotic combination with Bifidobacterium animalis subsp. lactis induce bifidogenesis and modulate markers of immune function in healthy adults: A double-blind, placebo-controlled, randomised, factorial cross-over study. Br. J. Nutr. 2014, 111, 1945–1956. [Google Scholar] [CrossRef] [PubMed]
  115. Lecerf, J.M.; Depeint, F.; Clerc, E.; Dugenet, Y.; Niamba, C.N.; Rhazi, L.; Cayzeele, A.; Abdelnour, G.; Jaruga, A.; Younes, H.; et al. Xylo-oligosaccharide (XOS) in combination with inulin modulates both the intestinal environment and immune status in healthy subjects, while XOS alone only shows prebiotic properties. Br. J. Nutr. 2012, 108, 1847–1858. [Google Scholar] [CrossRef] [PubMed]
  116. Mirande, C.; Mosoni, P.; Bera-Maillet, C.; Bernalier-Donadille, A.; Forano, E. Characterization of Xyn10A, a highly active xylanase from the human gut bacterium Bacteroides xylanisolvens XB1A. Appl. Microbiol. Biotechnol. 2010, 87, 2097–2105. [Google Scholar] [CrossRef] [PubMed]
  117. Di, T.; Chen, G.; Sun, Y.; Ou, S.; Zeng, X.; Ye, H. In vitro digestion by saliva, simulated gastric and small intestinal juices and fermentation by human fecal microbiota of sulfated polysaccharides from Gracilaria rubra. J. Funct. Foods 2018, 40, 18–27. [Google Scholar] [CrossRef]
  118. Andrieux, C.; Hibert, A.; Houari, A.-M.; Bensaada, M.; Popot, F.; Szylit, O. Ulva lactuca is poorly fermented but alters bacterial metabolism in rats inoculated with human fecal flora from methane and non-methane producers. J. Sci. Food Agric. 1998, 77, 25–30. [Google Scholar] [CrossRef]
  119. Ren, X.; Liu, L.; Gamallat, Y.; Zhang, B.; Xin, Y. Enteromorpha and polysaccharides from enteromorpha ameliorate loperamide-induced constipation in mice. Biomed. Pharmacother. 2017, 96, 1075–1081. [Google Scholar] [CrossRef]
  120. Charoensiddhi, S.; Conlon, M.A.; Franco, C.M.M.; Zhang, W. The development of seaweed-derived bioactive compounds for use as using enzyme technologies. Trends Food Sci. Technol. 2017, 70, 20–33. [Google Scholar] [CrossRef]
  121. Kim, W.J.; Park, J.W.; Park, J.K.; Choi, D.J.; Park, Y.I. Purification and Characterization of a Fucoidanase (FNase S) from a Marine Bacterium Sphingomonas paucimobilis PF-1. Mar. Drugs 2015, 13, 4398–4417. [Google Scholar] [CrossRef] [Green Version]
  122. Coste, O.; Malta, E.-j.; López, J.C.; Fernández-Díaz, C. Production of sulfated oligosaccharides from the seaweed Ulva sp. using a new ulvan-degrading enzymatic bacterial crude extract. Algal Res. 2015, 10, 224–231. [Google Scholar] [CrossRef]
  123. Barbeyron, T.; Lerat, Y.; Sassi, J.F.; Le Panse, S.; Helbert, W.; Collen, P.N. Persicivirga ulvanivorans sp. nov., a marine member of the family Flavobacteriaceae that degrades ulvan from green algae. Int. J. Syst. Evol. Microbiol. 2011, 61, 1899–1905. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  124. Ramos, K.R.M.; Valdehuesa, K.N.G.; Nisola, G.M.; Lee, W.K.; Chung, W.J. Identification and characterization of a thermostable endolytic beta-agarase Aga2 from a newly isolated marine agarolytic bacteria Cellulophaga omnivescoria W5C. New Biotechnol. 2018, 40, 261–267. [Google Scholar] [CrossRef] [PubMed]
  125. Xie, Z.; Lin, W.; Luo, J. Comparative Phenotype and Genome Analysis of Cellvibrio sp. PR1, a Xylanolytic and Agarolytic Bacterium from the Pearl River. BioMed Res. Int. 2017, 2017, 6304248. [Google Scholar] [CrossRef] [PubMed]
  126. Cheng, Y.; Wang, D.; Gu, J.; Li, J.; Liu, H.; Li, F.; Han, W. Biochemical characteristics and variable alginate-degrading modes of a novel bifunctional endolytic alginate lyase. Appl. Environ. Microbiol. 2017, 83. [Google Scholar] [CrossRef] [PubMed]
  127. Kislitsyn, Y.A.; Samygina, V.R.; Dvortsov, I.A.; Lunina, N.A.; Kuranova, I.P.; Velikodvorskaya, G.A. Crystallization and preliminary X-ray diffraction studies of the family 54 carbohydrate-binding module from laminarinase (beta-1,3-glucanase) Lic16A of Clostridium thermocellum. Acta Crystallogr. Sect. F Struct. Biol. Commun. 2015, 71, 217–220. [Google Scholar] [CrossRef]
  128. Xu, X.Q.; Su, B.M.; Xie, J.S.; Li, R.K.; Yang, J.; Lin, J.; Ye, X.Y. Preparation of bioactive neoagaroligosaccharides through hydrolysis of Gracilaria lemaneiformis agar: A comparative study. Food Chem. 2018, 240, 330–337. [Google Scholar] [CrossRef]
  129. Charoensiddhi, S.; Conlon, M.A.; Methacanon, P.; Franco, C.M.M.; Su, P.; Zhang, W. Gut health benefits of brown seaweed Ecklonia radiata and its polysaccharides demonstrated in vivo in a rat model. J. Funct. Foods 2017, 37, 676–684. [Google Scholar] [CrossRef]
  130. Nakata, T.; Kyoui, D.; Takahashi, H.; Kimura, B.; Kuda, T. Inhibitory effects of laminaran and alginate on production of putrefactive compounds from soy protein by intestinal microbiota in vitro and in rats. Carbohydr. Polym. 2016, 143, 61–69. [Google Scholar] [CrossRef]
  131. Kaewmanee, W.; Suwannaporn, P.; Huang, T.C.; Al-Ghazzewi, F.; Tester, R.F. In vivo prebiotic properties of Ascophyllum nodosum polysaccharide hydrolysates from lactic acid fermentation. J. Appl. Phycol. 2019. [Google Scholar] [CrossRef]
  132. Gomez-Guzman, M.; Rodriguez-Nogales, A.; Algieri, F.; Galvez, J. potential role of seaweed polyphenols in cardiovascular-associated disorders. Mar. Drugs 2018, 16, 250. [Google Scholar] [CrossRef] [PubMed]
  133. Lee, D.H.; Park, M.Y.; Shim, B.J.; Youn, H.J.; Hwang, H.J.; Shin, H.C.; Jeon, H.K. Effects of Ecklonia cava polyphenol in individuals with hypercholesterolemia: A pilot study. J. Med. Food 2012, 15, 1038–1044. [Google Scholar] [CrossRef] [PubMed]
  134. Lopes, G.; Andrade, P.B.; Valentao, P. Phlorotannins: Towards new pharmacological interventions for diabetes mellitus type 2. Molecules 2016, 22, 56. [Google Scholar] [CrossRef] [PubMed]
  135. Murray, M.; Dordevic, A.L.; Ryan, L.; Bonham, M.P. An emerging trend in functional foods for the prevention of cardiovascular disease and diabetes: Marine algal polyphenols. Crit. Rev. Food Sci. Nutr. 2018, 58, 1342–1358. [Google Scholar] [CrossRef] [PubMed]
  136. Murugan, A.C.; Karim, M.R.; Yusoff, M.B.; Tan, S.H.; Asras, M.F.; Rashid, S.S. New insights into seaweed polyphenols on glucose homeostasis. Pharm. Biol. 2015, 53, 1087–1097. [Google Scholar] [CrossRef] [PubMed]
  137. Fernando, I.P.; Kim, M.; Son, K.T.; Jeong, Y.; Jeon, Y.J. Antioxidant activity of marine algal polyphenolic compounds: A mechanistic approach. J. Med. Food 2016, 19, 615–628. [Google Scholar] [CrossRef] [PubMed]
  138. Eom, S.H.; Kim, Y.M.; Kim, S.K. Antimicrobial effect of phlorotannins from marine brown algae. Food Chem. Toxicol. Int. J. Publ. Br. Ind. Biol. Res. Assoc. 2012, 50, 3251–3255. [Google Scholar] [CrossRef] [PubMed]
  139. Heffernan, N.; Brunton, N.P.; FitzGerald, R.J.; Smyth, T.J. Profiling of the molecular weight and structural isomer abundance of macroalgae-derived phlorotannins. Mar. Drugs 2015, 13, 509–528. [Google Scholar] [CrossRef]
  140. Melanson, J.E.; MacKinnon, S.L. Characterization of Phlorotannins from Brown Algae by LC-HRMS. Methods Mol. Biol. 2015, 1308, 253–266. [Google Scholar] [CrossRef]
  141. Montero, L.; Sanchez-Camargo, A.P.; Garcia-Canas, V.; Tanniou, A.; Stiger-Pouvreau, V.; Russo, M.; Rastrelli, L.; Cifuentes, A.; Herrero, M.; Ibanez, E. Anti-proliferative activity and chemical characterization by comprehensive two-dimensional liquid chromatography coupled to mass spectrometry of phlorotannins from the brown macroalga Sargassum muticum collected on North-Atlantic coasts. J. Chromatogr. A 2016, 1428, 115–125. [Google Scholar] [CrossRef]
  142. Lewandowska, U.; Szewczyk, K.; Hrabec, E.; Janecka, A.; Gorlach, S. Overview of metabolism and bioavailability enhancement of polyphenols. J. Agric. Food Chem. 2013, 61, 12183–12199. [Google Scholar] [CrossRef] [PubMed]
  143. Opara, E.I.; Chohan, M. culinary herbs and spices: Their bioactive properties, the contribution of polyphenols and the challenges in deducing their true health benefits. Int. J. Mol. Sci. 2014, 15, 19183–19202. [Google Scholar] [CrossRef] [PubMed]
  144. Clifford, M.N. Diet-derived phenols in plasma and tissues and their implications for health. Planta Med. 2004, 70, 1103–1114. [Google Scholar] [CrossRef] [PubMed]
  145. Selma, M.V.; Espín, J.C.; Tomás-Barberán, F.A. Interaction between phenolics and gut microbiota: Role in human health. J. Agric. Food Chem. 2009, 57, 6485–6501. [Google Scholar] [CrossRef] [PubMed]
  146. Williamson, G.; Clifford, M.N. Role of the small intestine, colon and microbiota in determining the metabolic fate of polyphenols. Biochem. Pharmacol. 2017, 139, 24–39. [Google Scholar] [CrossRef] [PubMed]
  147. Espin, J.C.; Gonzalez-Sarrias, A.; Tomas-Barberan, F.A. The gut microbiota: A key factor in the therapeutic effects of (poly)phenols. Biochem. Pharmacol. 2017, 139, 82–93. [Google Scholar] [CrossRef] [PubMed]
  148. Krumholz, L.R.; Bryant, M.P. Eubacterium oxidoreducens sp. nov. requiring H2 or formate to degrade gallate, pyrogallol, phloroglucinol and quercetin. Arch. Microbiol. 1986, 144, 8–14. [Google Scholar] [CrossRef]
  149. Schneider, H.; Blaut, M. Anaerobic degradation of flavonoids by Eubacterium ramulus. Arch. Microbiol. 2000, 173, 71–75. [Google Scholar] [CrossRef]
  150. Bang, S.H.; Hyun, Y.J.; Shim, J.; Hong, S.W.; Kim, D.H. Metabolism of rutin and poncirin by human intestinal microbiota and cloning of their metabolizing alpha-l-rhamnosidase from Bifidobacterium dentium. J. Microbiol. Biotechnol. 2015, 25, 18–25. [Google Scholar] [CrossRef]
  151. Amaretti, A.; Raimondi, S.; Leonardi, A.; Quartieri, A.; Rossi, M. Hydrolysis of the rutinose-conjugates flavonoids rutin and hesperidin by the gut microbiota and bifidobacteria. Nutrients 2015, 7, 2788–2800. [Google Scholar] [CrossRef]
  152. Delgado, S.; Florez, A.B.; Guadamuro, L.; Mayo, B. Genetic and biochemical characterization of an oligo-alpha-1,6-glucosidase from Lactobacillus plantarum. Int. J. Food Microbiol. 2017, 246, 32–39. [Google Scholar] [CrossRef] [PubMed]
  153. Corona, G.; Ji, Y.; Anegboonlap, P.; Hotchkiss, S.; Gill, C.; Yaqoob, P.; Spencer, J.P.E.; Rowland, I. Gastrointestinal modifications and bioavailability of brown seaweed phlorotannins and effects on inflammatory markers. Br. J. Nutr. 2016, 115, 1240–1253. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  154. Corona, G.; Coman, M.M.; Guo, Y.; Hotchkiss, S.; Gill, C.; Yaqoob, P.; Spencer, J.P.E.; Rowland, I. Effect of simulated gastrointestinal digestion and fermentation on polyphenolic content and bioactivity of brown seaweed phlorotannin-rich extracts. Mol. Nutr. Food Res. 2017, 61. [Google Scholar] [CrossRef] [PubMed]
  155. Sadeghi Ekbatan, S.; Sleno, L.; Sabally, K.; Khairallah, J.; Azadi, B.; Rodes, L.; Prakash, S.; Donnelly, D.J.; Kubow, S. Biotransformation of polyphenols in a dynamic multistage gastrointestinal model. Food Chem. 2016, 204, 453–462. [Google Scholar] [CrossRef] [PubMed]
  156. Oliveira, A.; Pintado, M. Stability of polyphenols and carotenoids in strawberry and peach yoghurt throughout in vitro gastrointestinal digestion. Food Funct. 2015, 6, 1611–1619. [Google Scholar] [CrossRef] [PubMed]
  157. Brown, E.M.; Nitecki, S.; Pereira-Caro, G.; McDougall, G.J.; Stewart, D.; Rowland, I.; Crozier, A.; Gill, C.I. Comparison of in vivo and in vitro digestion on polyphenol composition in lingonberries: Potential impact on colonic health. BioFactors 2014, 40, 611–623. [Google Scholar] [CrossRef]
  158. Dueñas, M.; Muñoz-González, I.; Cueva, C.; Jiménez-Girón, A.; Sánchez-Patán, F.; Santos-Buelga, C.; Moreno-Arribas, M.V.; Bartolomé, B. A survey of modulation of gut microbiota by dietary polyphenols. BioMed Res. Int. 2015, 2015, 850902. [Google Scholar] [CrossRef]
  159. Tomas-Barberan, F.A.; Selma, M.V.; Espin, J.C. Interactions of gut microbiota with dietary polyphenols and consequences to human health. Curr. Opin. Clin. Nutr. Metab. Care 2016. [Google Scholar] [CrossRef]
  160. Duda-Chodak, A.; Tarko, T.; Satora, P.; Sroka, P. Interaction of dietary compounds, especially polyphenols, with the intestinal microbiota: A review. Eur. J. Nutr. 2015, 54, 325–341. [Google Scholar] [CrossRef]
  161. Nunez-Sanchez, M.A.; Gonzalez-Sarrias, A.; Romo-Vaquero, M.; Garcia-Villalba, R.; Selma, M.V.; Tomas-Barberan, F.A.; Garcia-Conesa, M.T.; Espin, J.C. Dietary phenolics against colorectal cancer--From promising preclinical results to poor translation into clinical trials: Pitfalls and future needs. Mol. Nutr. Food Res. 2015, 59, 1274–1291. [Google Scholar] [CrossRef]
  162. Ozdal, T.; Sela, D.A.; Xiao, J.; Boyacioglu, D.; Chen, F.; Capanoglu, E. The Reciprocal Interactions between Polyphenols and Gut Microbiota and Effects on Bioaccessibility. Nutrients 2016, 8, 78. [Google Scholar] [CrossRef] [PubMed]
  163. Van Duynhoven, J.; Vaughan, E.E.; Jacobs, D.M.; Kemperman, R.A.; van Velzen, E.J.; Gross, G.; Roger, L.C.; Possemiers, S.; Smilde, A.K.; Dore, J.; et al. Metabolic fate of polyphenols in the human superorganism. Proc. Natl. Acad. Sci. USA 2011, 108, 4531–4538. [Google Scholar] [CrossRef] [PubMed]
  164. Rubio, L.; Macia, A.; Motilva, M.J. Impact of various factors on pharmacokinetics of bioactive polyphenols: An overview. Curr. Drug Metab. 2014, 15, 62–76. [Google Scholar] [CrossRef] [PubMed]
  165. Feliciano, R.P.; Mills, C.E.; Istas, G.; Heiss, C.; Rodriguez-Mateos, A. Absorption, metabolism and excretion of cranberry (Poly)phenols in humans: A dose response study and assessment of inter-individual variability. Nutrients 2017, 9, 268. [Google Scholar] [CrossRef] [PubMed]
  166. Dueñas, M.; Cueva, C.; Muñoz-González, I.; Jiménez-Girón, A.; Sánchez-Patán, F.; Santos-Buelga, C.; Moreno-Arribas, M.V.; Bartolomé, B. Studies on modulation of gut microbiota by wine polyphenols: From isolated cultures to omic approaches. Antioxidants 2015, 4, 1–21. [Google Scholar] [CrossRef]
  167. Rajauria, G.; Foley, B.; Abu-Ghannam, N. Characterization of dietary fucoxanthin from Himanthalia elongata brown seaweed. Food Res. Int. 2017, 99, 995–1001. [Google Scholar] [CrossRef]
  168. Christaki, E.; Bonos, E.; Giannenas, I.; Florou-Paneri, P. Functional properties of carotenoids originating from algae. J. Sci. Food Agric. 2013, 93, 5–11. [Google Scholar] [CrossRef]
  169. Mikami, K.; Hosokawa, M. Biosynthetic pathway and health benefits of fucoxanthin, an algae-specific xanthophyll in brown seaweeds. Int. J. Mol. Sci. 2013, 14, 13763–13781. [Google Scholar] [CrossRef]
  170. Lopes-Costa, E.; Abreu, M.; Gargiulo, D.; Rocha, E.; Ramos, A.A. Anticancer effects of seaweed compounds fucoxanthin and phloroglucinol, alone and in combination with 5-fluorouracil in colon cells. J. Toxicol. Environ. Health 2017, 80, 776–787. [Google Scholar] [CrossRef]
  171. Maeda, H.; Tsukui, T.; Sashima, T.; Hosokawa, M.; Miyashita, K. Seaweed carotenoid, fucoxanthin, as a multi-functional nutrient. Asia Pac. J. Clin. Nutr. 2008, 17, 196–199. [Google Scholar] [CrossRef]
  172. Woo, M.N.; Jeon, S.M.; Kim, H.J.; Lee, M.K.; Shin, S.K.; Shin, Y.C.; Park, Y.B.; Choi, M.S. Fucoxanthin supplementation improves plasma and hepatic lipid metabolism and blood glucose concentration in high-fat fed C57BL/6N mice. Chem. Biol. Interact. 2010, 186, 316–322. [Google Scholar] [CrossRef] [PubMed]
  173. Kulczyński, B.; Gramza-Michałowska, A.; Kobus-Cisowska, J.; Kmiecik, D. The role of carotenoids in the prevention and treatment of cardiovascular disease–Current state of knowledge. J. Funct. Foods 2017, 38, 45–65. [Google Scholar] [CrossRef]
  174. Bohn, T.; McDougall, G.J.; Alegria, A.; Alminger, M.; Arrigoni, E.; Aura, A.M.; Brito, C.; Cilla, A.; El, S.N.; Karakaya, S.; et al. Mind the gap-deficits in our knowledge of aspects impacting the bioavailability of phytochemicals and their metabolites—A position paper focusing on carotenoids and polyphenols. Mol. Nutr. Food Res. 2015, 59, 1307–1323. [Google Scholar] [CrossRef]
  175. Widjaja-Adhi, M.A.K.; Lobo, G.P.; Golczak, M.; Von Lintig, J. A genetic dissection of intestinal fat-soluble vitamin and carotenoid absorption. Hum. Mol. Genet. 2015, 24, 3206–3219. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  176. Rein, M.J.; Renouf, M.; Cruz-Hernandez, C.; Actis-Goretta, L.; Thakkar, S.K.; da Silva Pinto, M. Bioavailability of bioactive food compounds: A challenging journey to bioefficacy. Br. J. Clin. Pharmacol. 2013, 75, 588–602. [Google Scholar] [CrossRef] [PubMed]
  177. Bohn, T. Chapter 9 Metabolic Fate of Bioaccessible and Non-bioaccessible Carotenoids. In Non-Extractable Polyphenols and Carotenoids: Importance in Human Nutrition and Health, 1st ed.; Saura-Calixto, F., Pérez-Jiménez, J., Eds.; The Royal Society of Chemistry: London, UK, 2018; pp. 165–200. [Google Scholar] [CrossRef]
  178. Lyu, Y.; Wu, L.; Wang, F.; Shen, X.; Lin, D. Carotenoid supplementation and retinoic acid in immunoglobulin A regulation of the gut microbiota dysbiosis. Exp. Biol. Med. 2018, 243, 613–620. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  179. Kamiloglu, S.; Capanoglu, E. Chapter 10 models for studying polyphenols and carotenoids digestion, bioaccessibility and colonic fermentation. In Non-Extractable Polyphenols and Carotenoids: Importance in Human Nutrition and Health, 1st ed.; Saura-Calixto, F., Pérez-Jiménez, J., Eds.; The Royal Society of Chemistry: London, UK, 2018; pp. 201–219. [Google Scholar] [CrossRef]
  180. Van Ginneken, V.J.T.; Helsper, J.P.F.G.; de Visser, W.; van Keulen, H.; Brandenburg, W.A. Polyunsaturated fatty acids in various macroalgal species from north Atlantic and tropical seas. Lipids Health Dis. 2011, 10, 104. [Google Scholar] [CrossRef] [PubMed]
  181. Robertson, R.C.; Guihéneuf, F.; Bahar, B.; Schmid, M.; Stengel, D.B.; Fitzgerald, G.F.; Ross, R.P.; Stanton, C. The Anti-Inflammatory Effect of Algae-Derived Lipid Extracts on Lipopolysaccharide (LPS)-Stimulated Human THP-1 Macrophages. Mar. Drugs 2015, 13, 5402–5424. [Google Scholar] [CrossRef] [Green Version]
  182. Costantini, L.; Molinari, R.; Farinon, B.; Merendino, N. Impact of Omega-3 fatty acids on the gut microbiota. Int. J. Mol. Sci. 2017, 18, 2645. [Google Scholar] [CrossRef]
  183. Menni, C.; Zierer, J.; Pallister, T.; Jackson, M.A.; Long, T.; Mohney, R.P.; Steves, C.J.; Spector, T.D.; Valdes, A.M. Omega-3 fatty acids correlate with gut microbiome diversity and production of N-carbamylglutamate in middle aged and elderly women. Sci. Rep. 2017, 7, 11079. [Google Scholar] [CrossRef]
  184. Robertson, R.C.; Kaliannan, K.; Strain, C.R.; Ross, R.P.; Stanton, C.; Kang, J.X. Maternal omega-3 fatty acids regulate offspring obesity through persistent modulation of gut microbiota. Microbiome 2018, 6, 95. [Google Scholar] [CrossRef] [PubMed]
  185. Marco, M.L.; Heeney, D.; Binda, S.; Cifelli, C.J.; Cotter, P.D.; Foligne, B.; Ganzle, M.; Kort, R.; Pasin, G.; Pihlanto, A.; et al. Health benefits of fermented foods: Microbiota and beyond. Curr. Opin. Biotechnol. 2017, 44, 94–102. [Google Scholar] [CrossRef] [PubMed]
  186. Chilton, S.N.; Burton, J.P.; Reid, G. Inclusion of fermented foods in food guides around the world. Nutrients 2015, 7, 390–404. [Google Scholar] [CrossRef] [PubMed]
  187. Tamang, J.P.; Shin, D.H.; Jung, S.J.; Chae, S.W. Functional properties of microorganisms in fermented foods. Front. Microbiol. 2016, 7, 578. [Google Scholar] [CrossRef] [PubMed]
  188. Wilburn, J.R.; Ryan, E.P. Chapter 1—Fermented foods in health promotion and disease prevention: An overview. In Fermented Foods in Health and Disease Prevention; Frias, J., Martinez-Villaluenga, C., Peñas, E., Eds.; Academic Press: Boston, MA, USA, 2017; pp. 3–19. [Google Scholar] [CrossRef]
  189. Ko, S.J.; Kim, J.; Han, G.; Kim, S.K.; Kim, H.G.; Yeo, I.; Ryu, B.; Park, J.W. Laminaria japonica combined with probiotics improves intestinal microbiota: A randomized clinical trial. J. Med. Food 2014, 17, 76–82. [Google Scholar] [CrossRef] [PubMed]
  190. Kang, Y.M.; Lee, B.J.; Kim, J.I.; Nam, B.H.; Cha, J.Y.; Kim, Y.M.; Ahn, C.B.; Choi, J.S.; Choi, I.S.; Je, J.Y. Antioxidant effects of fermented sea tangle (Laminaria japonica) by Lactobacillus brevis BJ20 in individuals with high level of gamma-GT: A randomized, double-blind, and placebo-controlled clinical study. Food Chem. Toxicol. Int. J. Publ. Br. Ind. Biol. Res. Assoc. 2012, 50, 1166–1169. [Google Scholar] [CrossRef] [PubMed]
  191. Choi, J.S.; Seo, H.J.; Lee, Y.R.; Kwon, S.J.; Moon, S.H.; Park, S.M.; Sohn, J.H. Characteristics and in vitro Anti-diabetic Properties of the Korean Rice Wine, Makgeolli Fermented with Laminaria japonica. Prev. Nutr. Food Sci. 2014, 19, 98–107. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  192. Shobharani, P.; Nanishankar, V.H.; Halami, P.M.; Sachindra, N.M. Antioxidant and anticoagulant activity of polyphenol and polysaccharides from fermented Sargassum sp. Int. J. Biol. Macromol. 2014, 65, 542–548. [Google Scholar] [CrossRef] [PubMed]
  193. Uchida, M.; Miyoshi, T.; Yoshida, G.; Niwa, K.; Mori, M.; Wakabayashi, H. Isolation and characterization of halophilic lactic acid bacteria acting as a starter culture for sauce fermentation of the red alga Nori (Porphyra yezoensis). J. Appl. Microbiol. 2014, 116, 1506–1520. [Google Scholar] [CrossRef]
  194. Tavares Estevam, A.C.; Alonso Buriti, F.C.; de Oliveira, T.A.; Pereira, E.V.; Florentino, E.R.; Porto, A.L. Effect of Aqueous Extract of the Seaweed Gracilaria domingensis on the Physicochemical, Microbiological, and Textural Features of Fermented Milks. J. Food Sci. 2016, 81, C874–C880. [Google Scholar] [CrossRef]
  195. Blaszak, B.B.; Gozdecka, G.; Shyichuk, A. Carrageenan as a functional additive in the production of cheese and cheese-like products. Acta Sci. Polonorum. Technol. Aliment. 2018, 17, 107–116. [Google Scholar] [CrossRef]
  196. Bixler, H.J.; Porse, H.J. A decade of change in the seaweed hydrocolloids industry. J. Appl. Phycol. 2011, 23, 321–335. [Google Scholar] [CrossRef]
  197. Makkar, H.P.S.; Tran, G.; Heuzé, V.; Giger-Reverdin, S.; Lessire, M.; Lebas, F.; Ankers, P. Seaweeds for livestock diets: A review. Anim. Feed Sci. Technol. 2016, 212, 1–17. [Google Scholar] [CrossRef]
  198. Overland, M.; Mydland, L.T.; Skrede, A. Marine macroalgae as sources of protein and bioactive compounds in feed for monogastric animals. J. Sci. Food Agric. 2018, 99, 13–24. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  199. Umu, O.C.; Frank, J.A.; Fangel, J.U.; Oostindjer, M.; da Silva, C.S.; Bolhuis, E.J.; Bosch, G.; Willats, W.G.; Pope, P.B.; Diep, D.B. Resistant starch diet induces change in the swine microbiome and a predominance of beneficial bacterial populations. Microbiome 2015, 3, 16. [Google Scholar] [CrossRef] [PubMed]
  200. Kulshreshtha, G.; Rathgeber, B.; Stratton, G.; Thomas, N.; Evans, F.; Critchley, A.; Hafting, J.; Prithiviraj, B. Feed supplementation with red seaweeds, Chondrus crispus and Sarcodiotheca gaudichaudii, affects performance, egg quality, and gut microbiota of layer hens. Poult. Sci. 2014, 93, 2991–3001. [Google Scholar] [CrossRef] [PubMed]
  201. Kulshreshtha, G.; Rathgeber, B.; MacIsaac, J.; Boulianne, M.; Brigitte, L.; Stratton, G.; Thomas, N.A.; Critchley, A.T.; Hafting, J.; Prithiviraj, B. Feed Supplementation with Red Seaweeds, Chondrus crispus and Sarcodiotheca gaudichaudii, Reduce Salmonella Enteritidis in Laying Hens. Front. Microbiol. 2017, 8, 567. [Google Scholar] [CrossRef] [Green Version]
  202. Litten-Brown, J.C.; Corson, A.M.; Clarke, L. Porcine models for the metabolic syndrome, digestive and bone disorders: A general overview. Animal 2010, 4, 899–920. [Google Scholar] [CrossRef]
  203. Machado, L.; Tomkins, N.; Magnusson, M.; Midgley, D.J.; de Nys, R.; Rosewarne, C.P. In Vitro Response of Rumen Microbiota to the Antimethanogenic Red Macroalga Asparagopsis taxiformis. Microb. Ecol. 2018, 75, 811–818. [Google Scholar] [CrossRef]
  204. Zhou, M.; Hünerberg, M.; Chen, Y.; Reuter, T.; McAllister, T.A.; Evans, F.; Critchley, A.T.; Guan, L.L. Air-Dried Brown Seaweed, Ascophyllum nodosum, Alters the Rumen Microbiome in a Manner That Changes Rumen Fermentation Profiles and Lowers the Prevalence of Foodborne Pathogens. mSphere 2018, 3, e00017-18. [Google Scholar] [CrossRef]
  205. Orpin, C.G.; Greenwood, Y.; Hall, F.J.; Paterson, I.W. The rumen microbiology of seaweed digestion in Orkney sheep. J. Appl. Bacteriol. 1985, 58, 585–596. [Google Scholar] [CrossRef] [PubMed]
  206. Williams, A.G.; Withers, S.; Sutherland, A.D. The potential of bacteria isolated from ruminal contents of seaweed-eating North Ronaldsay sheep to hydrolyse seaweed components and produce methane by anaerobic digestion in vitro. Microb. Biotechnol. 2013, 6, 45–52. [Google Scholar] [CrossRef] [PubMed]
  207. Heim, G.; O’Doherty, J.V.; O’Shea, C.J.; Doyle, D.N.; Egan, A.M.; Thornton, K.; Sweeney, T. Maternal supplementation of seaweed-derived polysaccharides improves intestinal health and immune status of suckling piglets. J. Nutr. Sci. 2015, 4, e27. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  208. Choi, Y.; Hosseindoust, A.; Goel, A.; Lee, S.; Jha, P.K.; Kwon, I.K.; Chae, B.-J. Effects of Ecklonia cava as fucoidan-rich algae on growth performance, nutrient digestibility, intestinal morphology and caecal microflora in weanling pigs. Asian Australas. J. Anim. Sci. 2017, 30, 64–70. [Google Scholar] [CrossRef] [PubMed]
  209. Murphy, P.; Dal Bello, F.; O’Doherty, J.; Arendt, E.K.; Sweeney, T.; Coffey, A. The effects of liquid versus spray-dried Laminaria digitata extract on selected bacterial groups in the piglet gastrointestinal tract (GIT) microbiota. Anaerobe 2013, 21, 1–8. [Google Scholar] [CrossRef]
  210. Heim, G.; Walsh, A.M.; Sweeney, T.; Doyle, D.N.; O’Shea, C.J.; Ryan, M.T.; O’Doherty, J.V. Effect of seaweed-derived laminarin and fucoidan and zinc oxide on gut morphology, nutrient transporters, nutrient digestibility, growth performance and selected microbial populations in weaned pigs. Br. J. Nutr. 2014, 111, 1577–1585. [Google Scholar] [CrossRef] [PubMed]
  211. Murphy, P.; Dal Bello, F.; O’Doherty, J.; Arendt, E.K.; Sweeney, T.; Coffey, A. Analysis of bacterial community shifts in the gastrointestinal tract of pigs fed diets supplemented with β-glucan from Laminaria digitata, Laminaria hyperborea and Saccharomyces cerevisiae. Animal 2013, 7, 1079–1087. [Google Scholar] [CrossRef]
  212. Walsh, A.M.; Sweeney, T.; O’Shea, C.J.; Doyle, D.N.; ’Doherty, J.V.O. Effect of supplementing varying inclusion levels of laminarin and fucoidan on growth performance, digestibility of diet components, selected fecal microbial populations and volatile fatty acid concentrations in weaned pigs. Anim. Feed Sci. Technol. 2013, 183, 151–159. [Google Scholar] [CrossRef]
  213. Belanche, A.; Ramos-Morales, E.; Newbold, C.J. In vitro screening of natural feed additives from crustaceans, diatoms, seaweeds and plant extracts to manipulate rumen fermentation. J. Sci. Food Agric. 2016, 96, 3069–3078. [Google Scholar] [CrossRef]
  214. Kinley, R.D.; de Nys, R.; Vucko, M.J.; Machado, L.; Tomkins, N.W. The red macroalgae Asparagopsis taxiformis is a potent natural antimethanogenic that reduces methane production during in vitro fermentation with rumen fluid. J. Anim. Prod. Sci. 2016, 56, 282–289. [Google Scholar] [CrossRef]
  215. Maia, M.R.; Fonseca, A.J.; Oliveira, H.M.; Mendonca, C.; Cabrita, A.R. The potential role of seaweeds in the natural manipulation of rumen fermentation and methane production. Sci. Rep. 2016, 6, 32321. [Google Scholar] [CrossRef] [PubMed]
  216. Hong, Z.S.; Kim, E.J.; Jin, Y.C.; Lee, J.S.; Choi, Y.J.; Lee, H.G. Effects of supplementing brown seaweed by-products in the diet of Holstein cows during transition on ruminal fermentation, growth performance and endocrine responses. Asian Australas. J. Anim. Sci. 2015, 28, 1296–1302. [Google Scholar] [CrossRef] [PubMed]
Table
Table 1. Potential degradation of brown seaweed glycans by the human gut microbiota.
Table 1. Potential degradation of brown seaweed glycans by the human gut microbiota.
CarbohydrateCarbohydrate-Active Enzyme (CAZyme)Evidenced Glycolytic BacteriaReference
Alginate1,4-β-d-mannuronic acid
α-l-guluronic acid		PL6 Alginate lyase
PL6 MG-specific alginate lyase		Bacteroides clarus
Bacteroides eggerthii
PL15 Alginate lyase
PL15 Oligoalginate lyase		Bacteroides ovatus
Bacteroides thetaiotaomicron
Bacteroides xylanisolvens		[31,32,33,34,35,36]
PL17 Alginate lyase
PL17 Oligoalginate lyase		Bacteroides clarus
Bacteroides eggerthii
FucoidanSulphated 1,2-1,3-1,4-α-l-fucoseGH29 α-l-fucosidase
GH29 α-1,3/1,4-l-fucosidase		Not determined[37]
GH95 α-l-fucosidase
GH95 α-1,2-l-fucosidase
Laminarin1,3-1,6-β-glucoseGH16 β-glucanase
GH16 β-1,3-1,4-glucanase
GH16 endo-1,3-β-glucanase		Bacteroides distasonis
Bacteroides fragilis
Bacteroides thetaiotaomicron		[10,38]
PL, Polysaccharide Lyase family; GH, Glycoside Hydrolase family. Potential glycolytic bacteria were identified using the Carbohydrate-Active enZYmes Database [28].
Table
Table 2. Potential degradation of red seaweed glycans by the human gut microbiota.
Table 2. Potential degradation of red seaweed glycans by the human gut microbiota.
CarbohydrateCarbohydrate-Active Enzyme (CAZyme)Evidenced Glycolytic BacteriaReference
Agar (Galactan)1,3-β-d-galactose
1,4-3,6-anhydro-α-l-galactose		GH2 β-galactosidaseBacteroidetes plebeius[39,40,41,42]
GH16 β-agarase
GH86 β-agarase
GH117 1,3-α-3,6-anhydro-l-galactosidase
Carrageenan (Galactan)1,4-β-d-galactose
1,3-α-d-galactose
3,6-anhydro-d-galactose		GH2 β-galactosidaseBacteroides plebeius[41,43]
GH117 1,3-α-3,6-anhydro-l-galactosidase
Porphyran (Galactan)Sulphated 1,3-β-d-galactose
1,4-α-l-galactose-6-sulfate
3,6-anhydro-α-l-galactose		GH16 β-porphyranase
GH86 β-porphyranase		Bacteroides plebeius[41,44,45]
Xylan1,3-1,4-β-d-xyloseGH3 xylan 1,4-β-xylosidaseNot determined[46,47,48,49]
GH5 endo-1,4-β-xylanase
GH10 endo-1,4-β-xylanase
GH10 endo-1,3-β-xylanase
GH11 endo-β-1,4-xylanase
GH11 endo-β-1,3-xylanase
GH43 β-xylosidase
GH43 xylanase
GH43 β-1,3-xylosidase
GH67 xylan α-1,2-glucuronidase
GH115 xylan α-1,2-glucuronidase
CE1−CE7 and CE12 acetyl xylanesterases
PL, Polysaccharide Lyase family; GH, Glycoside Hydrolase family. Potential glycolytic bacteria were identified using the Carbohydrate-Active enZYmes Database [28].
Table
Table 3. Potential degradation of green seaweed glycans by the human gut microbiota.
Table 3. Potential degradation of green seaweed glycans by the human gut microbiota.
CarbohydrateCarbohydrate-Active Enzyme (CAZyme)Evidenced Glycolytic BacteriaReference
UlvanSulphated 1,4-β-d-Glucuronic acid
α-l-Rhamnose
1,4-β-d-xyloglucan		GH78 α-l-rhamnosidaseNot determined[50,51]
GH145 α-l-rhamnosidase
Xylan1,3-β-d-xyloseGH10 endo-1,3-β-xylanase,Not determined[52]
GH11 endo-β-1,3-xylanase
GH43 β-1,3-xylosidase
PL, Polysaccharide Lyase family; GH, Glycoside Hydrolase family. Potential glycolytic bacteria were identified using the Carbohydrate-Active enZYmes Database [28].
Table
Table 4. In vitro fermentation of brown seaweeds with human faecal inoculum.
Table 4. In vitro fermentation of brown seaweeds with human faecal inoculum.
SeaweedSubstrateDoseUse of a Simulated In Vitro Digestion Before Fermentation?Experimental ParametersMicrobial EnumerationMicrobial ChangesMetabolomics Analysis TechniqueMetabolite ChangesReference
Ecklonia radiataCrude fraction (CF)
Phlorotannin-enriched fraction (PF)
Low-molecular weight polysaccharide fraction (LPF)
High-molecular weight polysaccharide fraction (HPF)		1.5% (w/v)Yes
CF = 71.5% digestible
PF = 87.3% digestible
LPF = 86.1% digestible
HPF = non-digestible		10% (w/v) pooled inoculum
(n = 3)
24 h		qPCR↑Bifidobacterium↑Lactobacillus
(LPF)
↑F. prausnitzii↑C. coccoides
↑Firmicutes
(CF, LPF)
↑Bacteroidetes↑E. coli
(CF, PF, LPF, HPF)
↓Enterococcus
(CF, PF)		GC-FID↑Acetate
(CF)
↑Propionate
(CF, LPF, HPF)
↑ Butyrate
(CF, LPF, HPF)
↑ Total SCFA
(CF, LPF, HPF)		[56]
Ecklonia radiataWater extract (WE)
Acid extract (AE)
Celluclast enzyme extract
(CEE)
Alcalase enzyme extract (AEE)
Free sugar fraction (FF)
Polysaccharide fraction (PF)
Seaweed residue (SR)
Seaweed powder (SP)		1.5% (w/v)No–digestibility unknown10% (w/v) pooled inoculum
(n = 3)
24 h		qPCR= F. prausnitzii = C. leptum
= R. bromii ↑ Total bacteria
(CEE, AEE, WE, FF)
↑ Bifidobacterium ↑ Bacteroidetes
↑ Lactobacillus ↑ C. coccoides
(CEE)
↑ E. coli ↑ Enterocccus
(WE, AE, CEE, AEE, FF, PF, SP)		GC-FID↑ Acetate
↑ Propionate
↑ Butyrate
(WE, AE, CEE, AEE, FF, PF, SP)
↑ Total SCFA		[57]
Sargassum muticumSargassum muticum Alcalase enzyme extract (SAE)1% (w/v)Yes–non-digestible (% digestible undisclosed)10% (w/v) single inoculum
24 h		FISH= Bifidobacterium
= Lactobacillus
= Clostridium histolticum
↑ Bacteroides/Prevotella
↓ C.coccoides/E.rectale		HPLC↑ Total SCFA[99]
Sargassum thunbergiiPolysaccharide extract0.3% (w/v)No–digestibility unknown20% (w/v) pooled inoculum
(n = 3)
24 h		16S rRNA
NGS		↑ Bacteroidetes
↑Bacteroidetes:Firmicutes ratio
↑ Bifidobacterium
↑ Roseburia
↑ Parasutterella
↑ Fusicatenibacter
↑ Coprococcus
↑ Fecalibacterium 		GC-MS↑ Acetate
↑ Propionate
↑ Butyrate
↑ Valerate
↑ Total SCFA		[55]
-Alginate5% (w/v)No–digestibility unknown10% (w/v) single inoculum
72 h		16S rRNA
DGGE
16S rRNA
NGS		↑BacteroidesGC-FID↑ Propionate
↑ Butyrate
↑ Total SCFA		[33]
-Alginate (A)
Mannuronic acid oligosaccharides (MO)
Guluronic acid oligosaccharides (GO) Propylene glycol alginate sodium sulphate (PSS)		5 g/L (A)
8 g/L (MO, GO, PSS)		No–digestibility unknown10% (w/v) single inoculum
48 h		16S rRNA DGGEDetection of Bacteroides xylanisolvens, Clostridium clostridioforme/Clostridium symbiosum, Bacteroides finegoldii, Shigella flexneri/E.coli, E.fergusonii, and Bacteroides ovatusHPLCA, MO, GO:
↑ Acetate
↑ Propionate
↑ Butyrate
↑ Total SCFA		[36]
Ascophyllum nodosumSulphated polysaccharide extract9 mg/mLYes–non-digestible (% digestible undisclosed)10% (w/v)
pooled inoculum
(n = 4)
24 h		16S rRNA NGS↑Bacteroides
↑Phascolarctobacterium
↑Oscillospira
↑Fecalibacterium		GC-FID↑ Acetate
↑ Propionate
↑ Butyrate
↑ Total SCFA		[100]
Laminaria digitataCrude polysaccharide extract (CE)
Depolymerised crude polysaccharide extract (DE)		1% (w/v)Yes–non-digestible (% digestible undisclosed)20% (w/v) pooled inoculum
(n = 3)
48 h		16S rRNA NGS↑Parabacteroides (CE, DE)
↑ Fibrobacter (CE)
↓ Streptococcus
↓ Ruminococcus
↑ Lachnospiraceae UC (DE)
↓ Peptostreptococcaceae IS (DE)
↑ Dialister (CE, DE)
↑ γ B38UC (CE)		GC-FID↑ Acetate (CE, DE)
↑ Propionate (CE, DE)
↑ Butyrate (CE, DE)
↑ Total SCFA (CE, DE)		[101]
-Laminarin1% (w/v)No–digestibility unknown10% (w/v) pooled inoculum
(n = 5)
24 h		qPCR↑Bifidobacterium
↑Bacteroides		HPLC↑ Acetate
↑ Propionate
↑ Total SCFA		[79]
qPCR, Quantitative PCR; GC-FID, Gas Chromatography with Flame Ionisation Detector; FISH, Flourescence in situ Hybridisation; 16S rRNA NGS, 16S rRNA Next Generation Sequencing; HPLC, High Performance Liquid Chromaography; GC-MS, Gas Chromatography-Mass Spectrometry; 16S rRNA DGGE, 16S rRNA Denaturing Gradient Gel Electrophoresis; 16S rRNA NGS, 16S rRNA Next Generation Sequencing; GC-FID, Gas Chromatography with Flame Ionisation Detector; HPLC, High Performance Liquid Chromaography; 16S rRNA NGS, 16S rRNA Next Generation Sequencing; qPCR, Quantitative PCR; GC-FID, Gas Chromatography with Flame Ionisation Detector; HPLC, High Performance Liquid Chromatography; SCFA, Short Chain Fatty Acid; =, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 5. In vitro fermentation of red seaweeds with human faecal inoculum.
Table 5. In vitro fermentation of red seaweeds with human faecal inoculum.
SeaweedSubstrateDoseUse of a Simulated in vitro Digestion Before Fermentation?Experimental ParametersMicrobial EnumerationMicrobial ChangesMetabolomics Analysis TechniqueMetabolite ChangesReference
Kappaphycus alvareziiWhole Seaweed (WS)1% (w/v)Yes–non-digestible
(% digestible undisclosed)		10% (w/v) single inoculum
24 h		FISH↑Bifidobacterium
↓Clostridium coccoides/
Eubacterium rectale		HPLC↑ Total SCFA[13]
Osmundea pinnatifidaOsmundea pinnatifida
Viscozyme extract (OVE)		1% (w/v)Yes–non-digestible
(% digestible undisclosed)		10% (w/v) single inoculum
24 h		FISH= Bifidobacterium
= Lactobacillus
= Clostridium histolticum		HPLC↑Total SCFA[99]
Gracilaria rubraPolysaccharide extract (PE)1%
(w/v)		Yes–non-digestible
(% digestible undisclosed)		10% (w/v) pooled
inoculum (n = 4)
24 h		16S rRNA
NGS		↑Bacteroides
↑Prevotella
↑Phascolarctobacterium
↓Firmicutes:Bacteroidetes		GC-FID↑ Acetate
↑ Propionate
↑ Isobutyrate
↑ Total SCFA		[117]
-Porphyran1% (w/v)No–digestibility unknown10% (w/v) pooled inoculum (n = 5)
24 h		qPCR↑Bifidobacterium
↑Bacteroides		HPLC= Acetate
= Propionate
= Butyrate
= Total SCFA		[79]
FISH, Flourescence in situ Hybridisation; 16S rRNA NGS, 16S rRNA Next Generation Sequencing; qPCR, Quantitative PCR; GC-FID, Gas Chromatography with Flame Ionisation Detector; HPLC, High Performance Liquid Chromatography; SCFA, Short Chain Fatty Acid; =, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 6. In vitro fermentation of green seaweeds with human faecal inoculum.
Table 6. In vitro fermentation of green seaweeds with human faecal inoculum.
SeaweedSubstrateDoseUse of a Simulated in vitro Digestion Before Fermentation?Experimental ParametersMicrobial EnumerationMicrobial ChangesMetabolomics Analysis TechniqueMetabolite ChangesReference
Enteromorpha proliferaPolysaccharide extract (PE)0.2 g in 9.5 mL
0.8 g in 9.5mL		Yes-non-digestible
(% digestible undisclosed)		10.5% (w/v) pooled inoculum (n = 3)
12, 24, and 48 h		Microbial culture↑Enterobacter (0.2 PE and 0.8 PE at 24 h and 48 h)
= Enterococcus
= Lactobacillus
= Bifidobacterium 		GC-FID= Acetate
= Butyrate
= Lactate		[12]
-Ulvan1% (w/v)No-digestibility unknown10% (w/v) pooled inoculum (n = 5)
24 h		qPCR↑Bifidobacterium
↑Lactobacillus		HPLC↑ Acetate
↑ Lactate		[79]
qPCR, Quantitative PCR; GC-FID, Gas Chromatography with Flame Ionisation Detector; HPLC, High Performance Liquid Chromatography; =, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 7. Impact of seaweeds on the rodent gut microbiota.
Table 7. Impact of seaweeds on the rodent gut microbiota.
AnimalSubstrateDoseDurationBiological SampleMicrobial ChangesMetabolite ChangesReference
30 Male Sprague-Dawley RatsChondrus crispus
Whole Seaweed (WS)		0.5% (w/w)
2.5% (w/w) 		21 daysFaeces↑Bifidobacterium↑Legionella↑Sutterella
↑Blautia↑Holdemania
↑Shewanella↑Agarivorans
↓Streptococcus
↑Bifidobacterium breve (2.5% WS)		↑ Acetate
↑ Propionate (2.5% WS)
↑ Butyrate
↑ Total SCFA		[15]
24 Male Sprague-Dawley RatsEcklonia radiata Whole Seaweed (WS)
Ecklonia radiata Polysaccharide Fraction (PF)		5% (w/w) WS
5% (w/w) PF		7 daysCaecum↑F. prausnitzii↑E. coli (PF)
↓ Enterococcus (WS)
↓Lactobacillus ↓Bifidobacterium
↓ Firmicutes:Bacteroidetes 		↑ Acetate
↑ Propionate
↑ Butyrate (PF)
↓ Valerate
↓ Hexanoate
↑ Total SCFA
↓ i-Butyrate
↓ i-Valerate
↓ phenol
↓ p-cresol		[129]
18 Male Wistar RatsAlginate (A)
Laminarin (L)
Fucoidan (F)		2% (w/w)14 daysCaecum↑Bacteroides (Bacteroides capillosus)
Presence of Enterorhabdus (A)
↑ Proteobacteria. Presence of Lachnospiracea, Parabacteroides (Parabacteroides distasonis) and Parasutterella (L)
Not fermented (F)		↑ Propionate (L)
↑ Total SCFA (A, L)		[97]
16 Male
C57 BL/6 Mice		Saccorhiza polyschides extract (BAE)High fat diet + 5% (w/w) BAE8 monthsFaeces↓ Faecal bile salt hydrolase activity↓ Secondary bile acids[54]
18 Male Wistar RatsAlginate (A)
Laminarin (L)		2% (w/w) 14 daysCaecum↑Lactobacillus↑Porphyromonas↑Coprobacillus
↑Oscillibacter valencigenes
↓Parabacteroides (L)
↑ Catabacter honkongensis
↑ Stomatobaculum longum
↓ Adlercreuzia (A)
↓ Helicobacter (A, L)		↑ Lactic acid (L)
= Acetate
= Propionate
= Butyrate
= Total SCFA
↓ Indole		[130]
18 Male C57BL/6 miceAscophyllum nodosum Fucoidan (FuA)
Laminaria
japonica Fucoidan (FuL)		100 mg/kg/day6 weeksCaecum↑Lactobacillus↑Anaeroplasma
↑Thalassospira (FuA)
↑Ruminococcaceae ↓ Alistipes
↓ Clostridiales
↓ Akkermansia (FuL)
↓ Candidatus ↓ Arthromitus
↓ Peptococcus
↓ Lachnospiraceae Incertae Sedis (FuA, FuL) 		-[96]
15 Male Wistar ratsAscophyllum nodosum seaweed crude polysaccharide (SCP)
SCP Lactobacillus plantarum hydrolysate (SCPH Lp)
SCP Enterococcus fecalis hydrolysate (SCPH Ef)
Alginate (A)
Hydrolysed Alginate (HA)		0.2 g per 180 –200 g rat weight4 daysFaeces-↑ Acetate (HA > A > SCPH Lp > SCPH Ef)
↑ Propionate (HA = A = SCPH Lp = SCPH Ef)
↑ Butyrate (HA = A = SCPH Lp = SCPH Ef)
(relative to day zero)		[131]
32 Female Kunming miceEnteromorpha prolifera (EP)
Enteromorpha polysaccharide extract (PEP)		1:5 (w/w)7 daysFaeces↑Alpha diversity (EP)
↑Bacteroidales S24-7 (EP)
↑ Prevotellaceae (PEP)
↑ Firmicutes ↑Actinobacteria (EP, PEP)
↓ Bacteroidetes ↓ Proteobacteria (EP, PEP)		-[119]
SCFA, Short Chain Fatty Acid; =, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 8. Impact of seaweeds on the porcine gut microbiota.
Table 8. Impact of seaweeds on the porcine gut microbiota.
AnimalSeaweed ComponentDoseDurationBiological SampleMicrobial ChangesMetabolite ChangesReference
20 pregnant gilts and 48 pigletsLaminarin/Fucoidan Extract10 g/day Gestation (day 83) to weaning (day 28)Faeces (Sow)
Colonic digesta (Piglet)		Sows (parturition): ↓ Enterobacteriaceae
= Lactobacilli
Piglets (birth, 48h after birth, weaning):
= Enterobacteriaceae = Lactobacilli		-[207]
200 pigsEcklonia cava Whole Seaweed0.05% (w/w)
0.1% (w/w)
0.15% (w/w) 		28 daysCaecum↑Lactobacillus
↓E. coli
= Total Anaerobes		-[208]
24 pigsLaminarin/Fucoidan
Extract (SD)
Laminarin/Fucoidan Wet Seaweed (WS)		5.37 Kg/tonne SD
26.3 Kg/tonne WS		21 daysIleum
Caecum
Colon		= Bifidobacteria
= Lactobacillus
= Enterobacterium (SD, WS)
↑Lactobacillus agilis (colon)		-[209]
48 pigsLaminarin Extract300 ppm 32 daysFaeces↑Lactobacillus
= Bifidobacteria		= Acetate
↓ Propionate
= Butyrate
= Valerate
= i-Butyrate
= i-Valerate		[210]
48 pigsβ-glucan250 g/tonne
150 g/tonne		29 daysIleum
Caecum
Proximal Colon Distal Colon		= Lactobacilli
= Bifidobacteria.
↑ Lactobacillus diversity		-[211]
168 pigsLaminarin (L)
Fucoidan (F)		240 mg/kg F
150 mg/kg L
300 mg/kg L
150 mg/kg L and 240 mg/kg F
300 mg/kg L and 240 mg/kg F		35 daysFaeces= E. coli
= Bifidobacteria
↑ Lactobacilli		= Acetate
= Propionate
= Butyrate
= Valerate
= i-Butyrate
= i-Valerate
= Total SCFA		[212]
9 pigsAlginate5.14% (w/w) 84 daysFaeces= Diversity
↑ Unclassified F16 family
↓ Clostridiaceae
↓ Unclassified RF39 (Mollicutes)
↑ Ruminococcus
↑ Roseburia
↑ unclassified F16 genus (TM7)
↑ Lachnospira
↓ Blautia 		-[199]
=, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 9. Impact of seaweeds on the hen gut microbiota.
Table 9. Impact of seaweeds on the hen gut microbiota.
AnimalSeaweed ComponentDoseDurationBiological SampleMicrobial ChangesMetabolite ChangesReference
160 laying hensChondrus crispus
Whole Seaweed (CC)
Sarcodiotheca gaudichaudii
Whole Seaweed (SG)		0.5% (w/w)
1% (w/w)
2% (w/w) 		30 daysIleum
Caecal digesta		↑Bifidobacterium longum (CC2, SG1, SG2)
↑Streptococcus salivarius (CC1, CC2, SG2)
↓Clostridium perfringens (CC1, CC2, SG1, SG2)
↓Lactobacillus acidophilus (CC1, CC2)		↑ Acetate (CC1, SG1)
↑ Propionate (CC2)
↑ Butyrate (SC2)		[200]
96 laying hensChondrus crispus
Whole Seaweed (CC)
Sarcodiotheca gaudichaudii
Whole Seaweed (SG)		Control diet + 2% (w/w) seaweed
Control diet + 4% (w/w) seaweed		28 daysCaecum↑Lactobacillus acidophilus (CC4)
↓Bifidobacterium longum (SG2, SG4, CC4)
↓ Streptococcus salivarius (SG2, SG4, CC2, CC4)
↑ Bacteroidetes (SG4, CC2, CC4)		↑ Propionate (CC4)[201]
=, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 10. In vitro fermentation of seaweeds with cow rumen inoculum.
Table 10. In vitro fermentation of seaweeds with cow rumen inoculum.
SeaweedSubstrateExperimental ParametersDose (w/v)Microbial EnumerationMicrobial ChangesMetabolomics Analysis TechniqueMetabolite ChangesReference
Ascophyllum nodosum (AN)
Laminaria digitata (LD) 		Whole Seaweed50% pooled inoculum
(n = 4)
24 h		0.5 g/L
1 g/L
2 g/L		- -GC-FID↑Propionate
↑Butyrate (LD)
↓ BCFA
↓Methane		[213]
Asparagopsis taxiformisWhole Seaweed20% pooled inoculum
(n = 4)
72 h		0.5%
1%
2%
5%
10%		- -GC-FID↓ Total gas production
↓ Methane
↓ Acetate
↑ Propionate
↑ Butyrate (2%, 10%)
↓ Total SCFA (5%, 10%)		[214]
Ulva sp.
Laminaria ochroleuca
Saccharina latissima
Gigartina sp.
Gracilaria vermiculophylla		Whole Seaweed20% pooled inoculum
(n = 2)
24 h		25%--GC-FID↓ Methane[215]
Brown seaweed by-products (BSB)-50% (v/v)
single inoculum
0, 3, 6, 9, 12, and 24 h		2%
4%		--GC-FID↓ Ammonia (3, 9, 12 and 24 h)
↓ Total SCFA (24 h)		[216]
GC-FID, Gas Chromatography; =, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.
Table
Table 11. Impact of seaweeds on the sheep rumen microbiota.
Table 11. Impact of seaweeds on the sheep rumen microbiota.
SeaweedDoseExperimental ParametersMicrobial EnumerationMicrobial ChangesMetabolomics Analysis TechniqueMetabolite ChangesReference
Asparagopsis taxiformis
Whole Seaweed		2%in vitro batch culture fermentation
20% (v/v) pooled sheep rumen fluid inoculum (n = 4)
48 and 72 h		16S rRNA NGS
qPCR		↓Methanogens
↓Bacteroidetes/Firmicutes ratio
↓ mcrA gene expression		GC-MS↓ Total Gas
↓ Methane
↑ Hydrogen		[203]
Ascophyllum nodosum
Whole Seaweed		1%
3%
5%		Rams (n = 8)
21 days ad libitum		16S rRNA NGS↓ undefined TM7-1
↓undefined Coriobacteriaceae ↓Roseburia
↓Coprococcus
↓Prevotella copri
↑Blautia producta
↑ Entodinium species 1
↑Veillonellaceae		GC-FIDDose dependent:
↑ Acetate
↓ Propionate
↓ Butyrate
PICRUSt:
↑Butanoate metabolism
↑ Fatty acid metabolism
↓Glycerophospholipid metabolism		[204]
16S rRNA NGS, 16S rRNA Next Generation Sequencing; qPCR, Quantitative PCR; GC-FID, Gas Chromatography; =, no statistical difference compared to the control; ↑, significant increase compared to the control; ↓ significant decrease compared to the control. Microbial and metabolite changes with abbreviations in parenthesis indicate the substrate(s) which exerted the effect. If no abbreviations in parenthesis are presented, then all of the seaweed substrates tested exerted the effect.

Share and Cite

MDPI and ACS Style

Cherry, P.; Yadav, S.; Strain, C.R.; Allsopp, P.J.; McSorley, E.M.; Ross, R.P.; Stanton, C. Prebiotics from Seaweeds: An Ocean of Opportunity? Mar. Drugs 2019, 17, 327. https://doi.org/10.3390/md17060327

AMA Style

Cherry P, Yadav S, Strain CR, Allsopp PJ, McSorley EM, Ross RP, Stanton C. Prebiotics from Seaweeds: An Ocean of Opportunity? Marine Drugs. 2019; 17(6):327. https://doi.org/10.3390/md17060327

Chicago/Turabian Style

Cherry, Paul, Supriya Yadav, Conall R. Strain, Philip J. Allsopp, Emeir M. McSorley, R. Paul Ross, and Catherine Stanton. 2019. "Prebiotics from Seaweeds: An Ocean of Opportunity?" Marine Drugs 17, no. 6: 327. https://doi.org/10.3390/md17060327

APA Style

Cherry, P., Yadav, S., Strain, C. R., Allsopp, P. J., McSorley, E. M., Ross, R. P., & Stanton, C. (2019). Prebiotics from Seaweeds: An Ocean of Opportunity? Marine Drugs, 17(6), 327. https://doi.org/10.3390/md17060327

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