Open access peer-reviewed chapter

Beneficial Effects of Proanthocyanidins on Intestinal Permeability and Its Relationship with Inflammation

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Carlos González-Quilen, Esther Rodríguez-Gallego, Raúl Beltrán-Debón, Montserrat Pinent, Anna Ardévol, Maria Teresa Blay and Ximena Terra

Submitted: November 22nd, 2019 Reviewed: January 15th, 2020 Published: February 25th, 2020

DOI: 10.5772/intechopen.91212

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Abstract

The intestinal barrier is constantly exposed to potentially harmful environmental factors including food components and bacterial endotoxins. When the intestinal barrier function and immune homeostasis are compromised, inflammatory conditions may be developed and impact overall health. Evidence from experimental animal and cell-culture studies suggests that exposure of intestinal mucosa to proanthocyanidin-rich plant products may contribute to maintain the barrier function and to ameliorate the inflammation present in prevalent pathologies such as diet-induced obesity and inflammatory bowel disease. In this review, we aim to update the current knowledge on the bioactivity of PACs in experimental models of altered intestinal permeability and in humans, emphasizing the beneficial effects of grape-seed proanthocyanidin extracts in intestinal health and giving insights into the subjacent biochemical and molecular mechanism.

Keywords

  • gut
  • permeability
  • inflammation
  • metabolic endotoxemia
  • obesity
  • IBD
  • flavonoid
  • flavan-3-ol
  • condensed tannin
  • procyanidin

1. Introduction

The primary function of the intestinal tract is to digest food components and absorb nutrients and water from the lumen to the systemic circulation. The intestine is also a physical barrier that is in contact with the environment. As a result, the intestinal epithelium is constantly exposed to potentially pathogenic microorganisms, toxins, and harmful components of the diet. When there are disturbances in the barrier function and mucosal immune homeostasis, the influx of intestine luminal content triggers barrier dysfunction and an exaggerated mucosal immune response [1]. Ultimately, chronic exposition to these detrimental environmental stimuli may lead to the development of local and systemic inflammatory conditions [2, 3] that contribute to barrier dysfunction.

Natural products have been recognized as a source of therapeutic agents for many years [4]. Some plant-derived phenolic compounds show promising anti-inflammatory effects and have been associated with the prevention of certain chronic diseases [5]. Proanthocyanidins (PACs), also known as condensed tannins, are oligo- and polymeric end products of the flavonoid biosynthesis pathway in plants [6]. There has been extensive laboratory research into the effects of both pure PAC molecules and PAC-rich extracts on overall health. These phytochemicals show a wide range of physiological activities [7], including anti-inflammatory and barrier-protective effects in the intestine [8, 9, 10], which may be interesting in the context of diet-induced obesity and inflammatory bowel disease (IBD).

We have reported previously that grape-seed PACs and other flavonoids have beneficial effects on inflammation [11, 12, 13] and protect the intestine against alterations associated with diet-induced obesity in rats [8, 9, 14, 15]. In addition, research conducted during the last decade with cell culture and animal models has made significant progress in determining the underlying mechanism of the health-promoting properties of PACs in the gastrointestinal tract and peripheral tissues.

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2. Altered networks in intestinal dysfunction: barrier integrity and inflammatory response

The intestinal epithelium is a single cell-layer responsible for separating underlying mucosal tissues from the environment and is the largest exposed surface area in the body [16]. As there is a prolific commensal microbial community in the intestinal lumen (intestinal microbiota), epithelial integrity plays a pivotal role in maintaining overall health [16, 17]. The intestinal epithelium is integrated by several cell types with specialized functions. The enterocytes are responsible for the absorptive function and constitute the most abundant epithelial cell lineage. The goblet cells are implicated in the synthesis of secretory mucin glycoproteins that form the mucus layer [18]. Other cellular types integrating the epithelium, microfold (M) [19], Paneth and enteroendocrine cells are specialized in antigen sampling and presentation to dendritic cells, synthesis of antimicrobial peptides, and secretion of hormones, respectively.

The first strategy the host tissue has to maintain its homeostatic relationship with the intestinal microbiota is to minimize the physical interaction with microorganisms, thus limiting microbial translocation and physiological inflammation [20, 21]. The thick mucus layer secreted by goblet cells represents a primary defense line against environmental insults [18]. In addition, the enterocytes are joined together forming an intricately and well-regulated barrier sustained by intercellular junctions linked to the cell cytoskeleton, such as tight junctions (TJs), desmosomes, and adherent junctions. TJs partially seal the paracellular space and prevent passive transport of large molecules, including microbial components and other potentially harmful agents [1, 22].

The paracellular and transcellular pathways are the two major pathways mediating transmembrane transfer of intestinal bacterial substances. Both mechanisms may be involved in intestinal mucosal barrier damage and bacterial translocation. The paracellular pathway is integrated by tight junctions (TJs), consisting of zonulin/zonula occludens (ZO)-1, occludin, claudins, junction adhesion molecules (JAMs), and actin-myosin cytoskeletal proteins. Previous studies have shown that inflammatory cytokines and bacterial antigens can affect the expression level and assembly of these elements, thereby exerting an influence on TJ functions [23]. Immune cells, including neutrophils, dendritic cells, and monocytes, have also been directly implicated in inducing disturbances in TJ barrier function. It has been postulated that pro-inflammatory cytokine-induced opening of the intestinal TJ barrier is an important mechanism contributing to the TJ barrier defects present in various inflammatory conditions of the gut [24]. Previous studies [25, 26, 27, 28] have shown that myosin light chain kinase (MLCK) plays a central role in the regulation of intestinal TJ permeability. The activation of MLCK catalyzes the phosphorylation of myosin light chain (MLC), inducing contraction of the peri-junctional actin-myosin filaments and the opening of the TJ barrier. In contrast, inhibition of MLCK activation prevents this effect [27]. It has been suggested that the cytokine-mediated barrier dysfunction could be mediated by an increase in Nuclear Factor (NF)-kB, which, in turn, activates MLCK gene and protein expression [29] (Figure 1).

Figure 1.

Protective properties of PACs in the intestinal barrier function. (A) Chronic exposition to detrimental environmental stimuli may lead to dysbiosis, breakdown of the intestinal barrier, influx of bacterial endotoxins and mucosal inflammation. (B) PACs ameliorate loss of barrier function blocking the activation of MLCK mediated by NF-κB and MAPK signaling. See text for details.

Once intestinal bacteria and endotoxins enter the portal vein and/or lymphatic system, they can reach other tissues and organs, leading to a cascade response modulated by inflammatory mediators. This situation can induce a systemic inflammatory response, which further damages the function of the intestinal barrier [30]. The endotoxin-signaling pathway includes the binding of LPS to LPS-binding protein (LBP) and its subsequent transfer to the CD14 receptor. LBP-bound LPS initiates inflammation via TLRs associated with membrane-anchored CD14 [31]. TLRs are a family of pattern-recognition receptors that play a key role in the innate immune system. Among all, the TLR4 is expressed at high levels in the intestinal tract, and given that LPS is its specific ligand, TLR4 could be considered the first barrier for recognition of bacterial presence in the gastrointestinal tract. NF-kB is the final effector transcription factor of the TLR4 signaling pathway. It promotes the development of many intestinal diseases and also plays a pivotal role in the translation and transcription of inflammatory mediators [30].

In mammals, the NF-kB family comprises five proteins, including p65 (RelA), RelB, c-Rel, p105/p50 (NF-kB1), and p100/p52 (NF-kB2), which associate with each other to form transcriptionally distinct homo- and heterodimeric complexes; the p65:p50 heterodimer is the most abundant and the most relevant for inflammation [32]. In resting cells, the p65:p50 NF-kB heterodimer is sequestered in the cytoplasm by binding to its inhibitory protein, IkappaB (IkB). In response to an inflammatory stimulus, such as LPS, the classical NF-kB activation pathway leads to the activation of the IkB kinase (IkkB), a member of the IKK complex, triggering IkB-a phosphorylation (pIkB-a). Then, pIkB-a is recognized by the ubiquitin ligase machinery, resulting in its polyubiquitination and subsequent proteasomal degradation. After pIkB-a degradation, the p65:p50 heterodimers are able to translocate to the nucleus, where they bind to the kB motif found in the promoter or enhancer regions of numerous pro-inflammatory genes to induce their expression [33].

NF-kB target genes include cytokines (e.g., tumor necrosis factor (TNF)-α and interleukins), adhesion molecules, acute phase proteins, and inducible enzymes (inducible nitric oxide synthase (iNOS) and cyclooxygenase 2 (COX2)), among others [11]. All of these genes contain verified NF-kB binding sites in their sequences, providing strong experimental evidence for their direct control by NF-kB [34]. Among all of these genes, the expression of iNOS and COX2 has been widely studied in relation to intestinal inflammation. In this regard, sustained high nitric oxide (NO) production by iNOS plays a role in the pathology of chronic inflammatory bowel disease [35, 36]. During the last decade, it has become increasingly clear that NO overproduction by iNOS is deleterious to intestinal function [37], thus contributing significantly to gastrointestinal immunopathology. Cyclooxygenases are enzymes that are responsible for the metabolism of arachidonic acid, converting it into prostaglandins. These products influence a wide variety of biological processes, ranging from homeostasis to inflammation [38]. There are two cyclooxygenase isoforms: the constitutive COX1 isoform and the inducible COX2 isoform [38, 39]. As a result of COX2 induction, prostaglandin E2 levels increase at the site of inflammation and can also be detected systemically.

Multiple environmental factors have been identified as potential triggers of intestinal inflammatory conditions, including Western dietary habits [40]. It has been described that saturated fats play a direct role in inflammatory signaling. Saturated fatty acids (SFA) such as lauric (C12:0) and palmitic (C16:0) directly induce NF-κB activation, acting as non-microbial TLR2 and TLR4 agonists in macrophages [41]. Data suggest that activation of TLRs by SFA is mediated by TLR dimerization and recruitment into lipid rafts [42]. We have reported mild intestinal inflammation and increased permeability in rats feeding on a cafeteria diet consisting of high-saturated fat/high-refined sugar food products [43]. This enhanced permeability has been shown to favor bacterial LPS and other potentially pro-inflammatory molecules entering the systemic circulation, which is known as metabolic endotoxemia [15].

Taken together, these data suggest that HF diet-induced changes in the intestinal microbiota could be responsible for metabolic endotoxemia and for the onset of the corresponding diseases. The causative link between changes in intestinal bacteria populations, endotoxemia, and metabolic disease needs further assessment [44], but the mechanisms likely include altered epithelial permeability, translocation of bacterial products, and upregulation of pro-inflammatory cytokines and hormones produced by gut endocrine cells, mechanisms which might be modulated by PACs.

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3. PACs: chemical structure, occurrence, and intake

PACs consist of flavan-3-ol subunits with a degree of polymerization (DP) equal to or greater than 2, mainly linked by (4 → 8) or (4 → 6) carbon-carbon bonds (B-type PACs) [45]. In some botanical sources an additional (2 → 7) ether-linkage also occurs (A-type PACs) [46] (Figure 2). Depending on the type of monomers, PACs can be classified into procyanidins, prodelphinidins, and propelargonidins. The most abundant group, procyanidins, consists exclusively of (+)- catechin and (−)- epicatechin monomers [47]. Prodelphinidins and propelargonidins are composed of (−)- gallocatechin/(−)- epigallocatechin and (+)- afzelechin/(−)- epiafzelechin monomers, respectively [45], and have a more limited distribution

Figure 2.

Chemical structures of PACs. Flavan-3-ol monomers differ based on the hydroxylation pattern and their cis- or trans-configuration. Dimers A1/A2 and B1/B2 are shown as example of A- and B-type PACs, respectively.

Dietary assessment studies have shown that PACs, especially procyanidins are among the most abundant polyphenols in the human diet [6], as they are present in a variety of botanical sources and plant food products such as tea, fruits, nuts, cacao products, legumes, and cereal grains [1, 2]. However, PAC intake varies widely between geographical regions and cultures and is greatly dependent on eating habits, lifestyle behaviors, and socioeconomic status [48]. The daily PAC (dimers to polymers) intakes in adult populations from Korea, the U.S., Mexico, and EU were estimated as 71 [49], 73 [48], 103 [50], and 123–180 mg [51, 52], respectively, but intakes up to 230 mg d−1 have been reported in some regions of Spain and Norway [53].

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4. The fate of PACs after ingestion

Flavan-3-ols are remarkably stable during gastric transit in humans [54]. Monomers such as (+)- catechin and (−)- epicatechin are readily absorbed in the upper sections of the small intestine [55, 56], recognized as xenobiotics and then subjected to an extensive phase II metabolism that generates glucuronidated, sulfated, and methylated conjugates [57]. Flavan-3-ol monomers and their conjugated metabolites reach peak plasma concentration 1–4 h after flavan-3-ol-rich food consumption [58, 59, 60]. Studies conducted in cultivated epithelial monolayers [61, 62, 63], rats [64, 65], and humans [60, 66] indicate that PAC absorption is conversely more limited and is highly dependent on DP, and that the permeation of larger oligomers (DP > 5) and polymers is negligible. No PAC transporter has been identified in the enterocyte membrane in the small intestine. Thus, dimers to tetramers are passively transported across the intestinal epithelium essentially by paracellular diffusion. Although transcellular passive diffusion is not likely to occur due to the hydrophilic nature of PACs conferred by the multiple hydroxyl groups, uptake might be possible by endocytic mechanisms [62].

In humans, a study assessed the contribution of the ingested cocoa flavan-3-ols and procyanidins to the systemic pool, and found that the plasma (−)- epicatechin came from the orally administered cocoa (−)- epicatechin and not from their oligomers or polymers [67]. This is in agreement with the evidence obtained with rats that suggests that PACs from different sources do not depolymerize to monomers after ingestion [68, 69]. Stalmach et al. [56] conducted a study with ileostomized patients who were administered green tea, and found 70% of the ingested flavan-3-ol in the ileal fluid after 24 h. Altogether, these findings suggest that substantial amounts of ingested flavan-3-ol monomers and PACs remain unabsorbed in the small intestine and reach the colon. There, they are efficiently transformed by the colonic microbiota into low molecular weight phenolic compounds that can be absorbed by colonocytes [57].

In vitro fermentation of purified procyanidin dimers with human fecal microbiota has shown to produce mainly 2-(3′,4′-dihydroxyphenyl) acetic acid and 5-(3′,4′-dihydroxyphenyl)-γ-valerolactone [70]. In agreement with this, a randomized cross-over study in healthy humans found that a great portion of the ingested (−)- epicatechin and procyanidin B1 was metabolized by the colonic microbiota to produce phenyl-γ-valerolactones as the major microbial metabolites [60]. In this study, microbial degradation of larger procyanidins was substantially lower, possibly to the inhibition of digesting enzymes or to the antibacterial properties exhibited by these compounds. Other human studies analyzing the bioavailability of flavan-3-ols, reported high levels of phenyl-γ-valerolactones in the circulation and urinary excretion after ingestion of a red grape pomace drink [71] and apple juice [72]. In the colonocytes and hepatocytes, these microbial products undergo further metabolism by phase II enzymes to produce conjugated derivatives. Margalef et al. [73] analyzed the tissue distribution of metabolites derived from a grape-seed proanthocyanidin extract (GSPE) 2 h after ingestion by rats. These authors detected a few microbial metabolites (methyl conjugated phenols) at low concentrations in the colon tissue, while most phase II metabolites (glucuronidated and methyl-glucuronidated forms) were found in the kidneys and liver. In humans, the major contributors to the excretion of phenyl-γ-valerolactones after ingestion of a red grape pomace drink, are sulfated and glucuronidated conjugates of 5-(3′,4′-dihydroxyphenyl)-γ-valerolactone [71].

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5. In vitro, in vivo and ex vivo studies on the benefits of PACs for intestinal dysfunction

During the last decade, the beneficial properties of PACs for intestinal function have been reported in several studies performed with cell-culture models and experimental animals (Tables 1 and 2). This experimental data indicate that PACs contribute to maintaining the intestinal barrier and improving mucosal inflammation induced by environmental insults. However, there are few studies on the effect of PACs on human intestinal health, although epidemiological studies connect PAC-rich food consumption with a lower risk of colorectal cancer [88].

Table 1.

Interaction of PACs with intestinal permeability markers in cell culture and animal models [10, 76, 82, 83, 84, 85, 86, 87].

Table 2.

Interaction of PACs with permeability and inflammatory markers in animal models of intestinal dysfunction [8, 13, 14, 15, 88, 89, 90, 91, 92, 93, 94].

In vitro models of inflammation have been fundamental in the comprehension of cellular mechanisms driving physiological effects of bioactive molecules. Studies on intestinal dysfunction have employed human colon carcinoma cell lines, being Caco-2 the most well-established and widely used model of the human intestine barrier ([89] and Table 1). Mucus producer [79], macrophages [90], and B cell lines [91] have been employed in co-culture systems to explore the interaction between cell populations. Although there is a strong trend in the industry toward replacing animal experiments with human cell-culture based models [92, 93], there are no in vitro models of the human intestine that replicate the complex interplay between cell types and the regulation of the barrier function by the mucosal innate and adaptive immunity. Therefore, most physiologically relevant data on intestinal dysfunction comes from the animal model. Most in vivo studies testing the effect of PAC supplementation on intestinal health have been performed in diet-induced obesity models and chemical-induced colitis models. The first resemble intestinal alterations seen in humans with metabolic syndrome [43]. The latter closely mimic histopathological features of human colitis and are frequently used to study the pathophysiology of IBD and the effectiveness of novel therapeutic drugs [94]. Notably, PAC-rich grape-seed extracts (GSPE) are among the most studied botanical extracts, mainly by in vivo approaches in rodents (Table 2).

5.1 In vitro studies of barrier integrity

The data available on the interaction between PACs and permeability and inflammation markers in cell models of intestinal dysfunction are summarized in Table 1. Caco-2-based models have shown to be responsive to pro-inflammatory stimulation, producing a wide range of inflammatory mediators and increasing the paracellular permeability. Pro-inflammatory agents such as LPS, phorbol 12-myristate 13-acetate (PMA), and cytokines (TNF-α and IL-1β) have been used in multiple studies testing the effect of PAC molecules and PAC-rich botanical extracts on Caco-2 cells [10, 74, 77, 78]. Stimulated-Caco-2 cell monolayers incubated with PACs generally show a reduction in gene expression and secretion of TNFα, IL-6, and IL-8 [10, 74, 75, 77], which is often linked to the downregulation of NF-κB signaling at different levels [10, 76, 77]. An increased expression of antioxidant enzymes, such as glutathione peroxidase (GPx), superoxidase dismutase (SOD), and hemeoxygenase 1 (HO-1), has also been reported [10].

When permeable support systems such as transwell or Ussing chamber (UCh) are used, alterations in barrier integrity and paracellular permeability of epithelial cell monolayers are evaluated by transepithelial electrical resistance (TEER), an electrophysiological parameter that measures ion conductance across the monolayer, and by the transepithelial transport of molecular markers such as Lucifer yellow (LY) and fluorescently labeled dextrans (FD) [95, 96]. Some in vitro studies have associated PACs with increased TEER and decreased transport of permeability markers in the context of barrier dysfunction [77, 78, 80]. The expression levels of TJ proteins (claudins, occludins, and ZOs) often correlate, but not always [79], with intestinal permeability and are also considered markers of epithelial integrity. Bitzer et al. [78] found that the dextran sodium sulfate (DSS)-induced loss of barrier function in Caco-2 cells was significantly inhibited by polymeric PACs of cocoa but not by oligomers. Moreover, a higher barrier-protective activity was determined in PACs with DP ≥ 7, which were able to reduce the detrimental effect of DSS in a dose-dependent fashion [78]. Effectiveness of procyanidin B2 ameliorating dextran sodium sulfate (DSS)-induced permeability alterations was examined using a Caco-2/HT29-MTX co-culture model [79]. Although procyanidin B2-incubated cells showed increased levels of the TJ proteins claudin-7, occludin, and ZO-1, these changes did not reduce TEER loss. Altogether, these results suggest that the ability of PACs to strengthen the intestinal barrier integrity depends on the degree of polymerization (DP).

5.2 In vivo studies of diet-induced intestinal permeability

The cafeteria (CAF) diet is a self-selected high-saturated fat/high-refined sugar diet that stimulates hyperphagia and a rapid weight gain in experimental animals [97, 98]. In this feeding regime, highly palatable and energy dense foods commercially available, such as muffins, biscuits, bacon, sausages, and sugared milk, are offered ad libitum[15, 99]. A long-term CAF diet (62% carbohydrate (mostly sugar), 23% lipid, and 13% protein) has negative effects on intestinal function in rodents, increasing intestinal permeability, and inducing mucosal inflammation [43]. We have described the beneficial effects of administering GSPE against the intestinal dysfunction induced by a long-term CAF diet (17–18 weeks) in Wistar rats [8, 14, 15]. The composition of the GSPE used in these studies has been analyzed in detail [100]. Both nutritional (5–50 mg kg−1 [14]) and pharmacological (100–500 mg kg−1 [8, 15]) doses of GSPE administered orally as a preventive [8] or counteractive treatment [14, 15], tended to reduce intestinal inflammatory markers such as TNF-α release or myeloperoxidase (MPO) activity (an indicator of neutrophil infiltration in tissues). The reduction of plasma ovalbumin (OVA), an in vivo marker of intestinal permeability, was supported by (1) the increase in TEER in small and large intestine segments. This parameter is determined ex vivoby UCh-based protocols [8, 15]; and (2) by the upregulation of TJ proteins such as ZO-1 [14] and claudin-1 [8, 15]. Notably, the protective effect of GSPE in the intestinal barrier function was linked to the amelioration of metabolic endotoxemia (reduction of plasma LPS) and systemic inflammation (reduction of plasma TNF-α) in obese rats [15, 101]. Other authors have also reported the upregulation of ZO-1 and claudin-1 TJ proteins in high-fat fed rats supplemented with other PAC-rich extracts [81].

5.3 In vivo studies of chemical-induced intestinal dysfunction

Chemical agents administered orally to induce colitis in rodents include trinitrobenzene sulfonic acid (TNBS) and DSS. These agents erode the colonic mucosal lining and produce the loss of the intestinal barrier function and colonic inflammation. In these models, the severity of outcomes depends on the dose of the chemical agent and the frequency of administration. Li et al. [102] found that intragastric administration of GSPE in rats at pharmacological doses (100–400 mg kg−1 d−1) prior to TNBS-induced recurrent colitis, reduced weight loss, and attenuated macro- and microscopic tissue damage scores in the colon. This protective effect was accompanied by a reduction in oxidative stress (malondialdehyde; MDA), inflammation (IL-1β), and neutrophil infiltration (MPO activity) in colonic tissues. Remarkably, the beneficial effects of low to high doses of GSPE were comparable to those of sulfasalazine (200 mg kg−1 d−1), a potent inhibitor of NF-κB. Subsequent studies carried out by these authors with the same model, confirmed the role of the GSPE down-regulating NF-κB response [83, 84]. A preventive effect of procyanidin B2 was also evidenced in a mouse model of DSS-induced colitis [85]. Administration of procyanidin B2 (10–40 mg kg−1 d−1) attenuated the severity of tissue damage in the colon and reduced the levels of matrix metalloproteinase-9 (MMP-9), a marker of macrophage infiltration. In addition, inhibition of the NF-κB signaling and of NLRP3 inflammasome activation was observed, with a concomitant reduction in the gene expression of pro-inflammatory cytokines. Overall, the benefits of procyanidin B2 administration, especially at the highest dosage (40 mg kg−1), were comparable to those of mesalazine (200 mg kg−1), a COX inhibitor. The authors suggest that these effects were largely due to the reduction in activated macrophages infiltrating colonic tissues, probably driven by ROS clearance.

5.4 Other in vivo studies with animal models

The IL-10 deficient mouse is a classic knockout model that develops spontaneous colitis under pathogen-free conditions. Some authors have explored the influence of GSPE in this model, supplementing colitic animals with 0.1–1 g 100 g−1 of dry feed weight for 12–16 days [86, 87]. These studies evidenced a reduction of multiple inflammatory markers in the jejunum and colon, such as TNF-α, IL-1β, IL-6, and IFN-γ gene expressions, as well as MPO activity. This anti-inflammatory effect was associated with the inhibition of the NF-κB signaling. Interestingly, GSPE supplementation also increased the density of goblet cells in the jejunum of treated animals, suggesting that there is an alternative mechanism by which inflammation is attenuated.

Cardoso et al. [13] recently tested both dietary (75 mg kg−1) and pharmacological doses of GSPE (375 mg kg−1) in a rat model of mild intestinal dysfunction induced by intraperitoneal injection of LPS. GSPE was administered daily by oral gavage for 15 days prior to LPS-induced intestinal dysfunction. LPS enhanced intestinal permeability and induced both oxidative stress and inflammation. GSPE-treated animals reduced OVA permeation to the circulation, MPO activity and COX-2 in the small intestine tissues, and reactive oxygen species (ROS) levels in the colon. Furthermore, a gene expression analysis with a low-density microarray technique revealed that unlike the dietary dose of GSPE, the pharmacological dose had a striking effect on the LPS-gene expression profile, showing a strong modulation of multiple genes associated with chemokines and ILs, including upregulation of the anti-inflammatory cytokine IL-13.

5.5 Human ex vivo studies

Although the use of animal models is the predominate approximation at preclinical stages for testing novel therapies in intestinal permeability, there is a strong trend in the industry towards replacing animal experiments with human cell-culture based models [92, 93]. Nevertheless, advantages related to the usefulness of in vitro models for screening of bioactives and exploring action mechanisms, are offset by limitations regarding the mimicking of the in vivo situation and translation to the human [103]. Thus, some human ex vivo models have been proposed to test immunomodulatory properties of drug candidates in intestinal explants from IBD patients [104, 105]. Intestinal function can also be studied with UCh-based protocols. The UCh system consists of two halves with an opening between them, where mucosal tissue is adapted, thus isolating the apical and basolateral sides of the tissue. This technique has been applied for studying drug absorption [106] and secretion of enterohormones [107] in human endoscopic biopsies. An advantage of UCh models over explant-based models is that UCh models make it possible to measure the electrophysiological parameters, including TEER [106]. All these set-ups permit analyzing the cytokine profiling of intestinal explants or biopsies retaining their in situ conditioning in a polarized fashion [105, 108]. We have employed the UCh to determine TEER and cytokine release (TNF-α) in intestinal tissues from cafeteria diet-induced obese rats treated with GSPE [8, 15]. It could also be useful for testing the effect of bioactives on dysfunctional human intestine. A feature of ex vivo models is that screening of drug effects does not compromise the patients by exposing them to unknown outcomes.

5.6 Clinical trials

Translation of doses of PAC-rich extracts used in rodent models of intestinal dysfunction to human equivalent doses (HED) indicates that pharmacological doses (up to 5 g d−1 for a 60 kg person) could be required to achieve beneficial effects in clinical trials [14, 15]. Thus, the first uncertainty involved in assessing the use of PACs as therapy agents in humans, is safety. Grape seed and skin proanthocyanidin-rich extracts have been subjected to toxicological tests in rats to determine their safety for use in functional foods [109, 110, 111]. In these studies, the median lethal dose (LD50) was found to be greater than 5000 mg kg−1 bw (HED of ≈50 g) when administered once by oral gavage, and 1400–2000 mg kg−1 d−1 (HED of ≈14–20 g d−1) was found to be the no-observed-adverse-effect level (NOAEL) for systemic toxicity in sub-chronic administration. A recent study evaluated the safety and tolerability of GSPE intake (up to 2.5 g d−1) in a small number of healthy adults for a 4-week period and found a good tolerability without adverse effects on hematological or biochemical parameters [47].

To date, there are few clinical studies that evaluate the influence of PACs on intestinal inflammatory conditions. A clinical study revealed that the postprandial increase of plasma LPS associated with the intake of a high-fat meal was significantly reduced in obese subjects who consumed 1 g of GSPE [112]. As translocation of LPS to the circulation is considered an indicator of intestinal permeability and a critical factor in the appearance of systemic low-grade inflammation in patients with metabolic syndrome [113], reduction of postprandial endotoxemia could be particularly interesting from a therapeutic perspective. Large double-blind clinical studies need to be conducted to provide more information on PAC clinical efficacy in intestinal dysfunction so that these phytochemicals can be used therapeutically to improve intestinal health in obese and IBD individuals.

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6. Biochemical and molecular mechanisms underlying the barrier-protective and anti-inflammatory properties of PAC in the intestine

PACs were often considered to be nutritionally undesirable due to their ability to form complexes with macronutrients and reduce the activity of virtually any enzyme implicated in digestion [114, 115]. Nevertheless, based on the anti-cancerous, anti-mutagenic, and anti-microbial activities these phytochemicals elicited in laboratory experiments, a role in the modulation of the metabolism and immune system was suggested [115]. The ability of PACs to form cross-links with biomolecules can be attributed to the hydroxyl groups and aromatic rings in their structure that can establish hydrogen bonds and hydrophobic interactions [116]. PACs have a significant affinity for proline-rich proteins and peptides [117]. In general, binding to proteins seems to increase with the DP as larger PAC molecules have more potential binding sites for the associations with proline residues [117]. The interaction results in effects determined by the biological function of the target protein. Thus, PACs not only alter enzymatic activity, but they may also prevent ligand-receptor interactions and the binding of transcription factors to their specific sites in DNA. In addition, some PAC molecules can be adsorbed non-specifically onto biomembrane surfaces [118], affecting their physical characteristics, such as fluidity and density, and potentially altering membrane-dependent processes, including protein receptor activity [119]. Altogether, these effects lead ultimately to the alteration of cell signaling pathways and the modulation of gene expression.

6.1 Modulation of TJ integrity

The precise mechanisms underlying the improvement in intestine paracellular permeability due to PACs in inflammation are not yet completely elucidated; however, it is known that they lead ultimately to the upregulation (e.g., ZO-1 and claudin-1 [8, 13]) or downregulation (e.g., claudin-2 [86]) of TJ protein expression. Loss of TJ integrity in the pro-inflammatory state is mediated by the NF-κB signaling pathway and by the activation of protein kinases MAPKs, PI3Ks, AMPK, and MLCK [120]. MLCK is particularly crucial in actomyosin-based cytoskeletal functions and multiple studies highlight its important role in intestinal TJ remodeling [121, 122]. PACs reduce the production of pro-inflammatory mediators (e.g., TNF-α) and reactive oxygen species (i.e., iNOS activity) associated with enhancing intestinal permeability by antagonizing the NF-κB signaling pathway. In addition, PACs are potent inhibitors of kinases including MLCK [120, 123]. Contreras et al. [124] also suggested that there is an upstream novel mechanism associated with flavan-3-ols that leads to the prevention of TNF-α-induced intestinal permeability. In this study, TNF-α-stimulated Caco-2 monolayers incubated with (−)- epicatechin showed a reduction of NOX activity, an enzyme that also facilitates activation of TNF-α signaling. This effect was directly associated with the inhibition of ERK1/2 MAPK activity of IκB phosphorylation and of MLCK activation.

6.2 Interaction with bacterial endotoxins

Delehanty et al. [125] demonstrated that naturally occurring A- and B-type cranberry PACs were able to bind the lipid A moiety of LPS, exhibiting an affinity similar to that of polymyxin B, a potent LPS-binding molecule. In this study, PACs efficiently blocked endocytosis of bacterial LPS in a dose-dependent manner in HEK 293 (human embryonic kidney cells) that expressed receptors TLR4/MD-2 and CD14, thus preventing the induction of the NF-κB signaling pathway without any interaction with cellular components. However, other authors reported that PACs isolated from cocoa beans did not abrogate the binding of LPS to TLR4 in cultivated human dendritic cells [126]. PAC-LPS binding has been linked to the reduction of the post prandial increase in blood LPS associated with the ingestion of a high-fat meal in obese subjects ingesting an oral dose of GSPE [112].

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7. PACs modulation of intestinal microbiota

Diet plays an important role in the composition of intestine microbiota, promoting or inhibiting growth of microorganisms [127]. Alterations in the composition and metabolism of the intestinal microbiota (dysbiosis) have also been associated with the consumption of high-saturated fat diets in rodents and humans [128, 129]. In fact, metagenomic analysis of the intestinal microbiome in Western populations has shown a reduction not only of microbial diversity, but also of functional potential [130]. Dysbiosis is linked to obesity-associated intestinal inflammation, although the “egg or hen” question related to the cause-effect relationship is not answered yet [131]. High-fat intake in rodents often decreases overall diversity of microbiota and the abundance of Bacteroidetes, and increases the relative abundance of Firmicutes [132, 133]. Several human studies have described similar associations [134, 135], but the importance of the ratio Firmicutes to Bacteroidetes remains controversial [136, 137], and some authors state that the experimental results are not sufficiently consistent [138]. Interestingly, the existence of a colitogenic microbiota was demonstrated in T-bet−/− × RAG2−/− deficient mice whose spontaneous ulcerative colitis was horizontally transmissible to wild-type individuals when co-housed [139]. Although mechanisms by which dysbiosis trigger intestinal dysfunction are not fully understood, it is known that they involve the loss of immune tolerance due to local immune homeostasis disruption and continuous abnormal activation of TLRs [140].

Several authors have suggested that both dietary PACs, which are the substrates of intestinal bacteria, and the metabolites produced during PACs degradation in the colon may modulate and induce oscillations in the composition of the microbiota populations by means of prebiotic and antimicrobial effects against gut pathogenic microorganisms [141, 142, 143, 144]. Dietary PACs, specifically longer polymers, reach the distal intestine nearly intact, where they become fermentable substrates for the commensal microbiota [145]. PACs have been associated with prebiotic properties, boosting the composition of several kinds of probiotics such as Bifidobacteriumspp., Lactobacillusspp. [146] and the stimulator of mucus production Akkermansia muciniphila[147, 148]. Nevertheless, current evidence is somewhat controversial as effects described in different in vivo studies mainly performed with rodents, do not always agree. This suggests that interactions between PACs and microbiota depend largely on the botanical source, the types of molecules present in the extracts tested and the animal model [149].

A recent study by Casanova-Marti et al. [150] found that oral administration of GSPE in Wistar rats for 8 days resulted in profound changes in the cecal microbiota composition, reducing diversity indices and the ratio of Firmicutesto Bacteroidetes. Similar results were found in diet-induced obese Sprague Dawley rats supplemented with a PAC-rich extract of the Pyracantha fortuneanafruit, although in this study an increase in microbiota diversity was also reported [81]. GSPE supplementation in IL-10 deficient mice resulted in an increased abundance of Bacteroidesand Lactobacilli[86]. Xing et al. [148] reported that the administration of procyanidin B2 in rabbits feeding a high-fat-cholesterol diet, promoted an increase in the relative abundance of Akkermansia. These authors proposed that the reduction of metabolic endotoxemia found in animals treated with procyanidin B2 was attributed to the ability of Akkermansiato retain the thickness of the intestinal mucus layer, thus reducing intestinal permeability and the leakage of LPS into the circulation [151].

Cueva et al. [146] found that in vitro fermentation of grape-seed monomers and PACs in human feces resulted in a reduced abundance of Clostridium histolyticum. Inhibition of the growth of some infectious microorganisms, such as the mentioned C. histolyticumin the intestine and Helicobacter pyloriin the stomach [152], may be related to the anti-adherence activity that PACs have demonstrated in in vitro studies [153], as adherence to the epithelium is a prerequisite for colonization and infection of the intestinal gastrointestinal mucosa.

Finally, phenolic acids and phenyl-γ-valerolactones resulting from the colonic fermentation of PACs also exhibit a significant bioactivity in cell models and experimental animals [154]. They therefore may partially account for the beneficial anti-inflammatory effects reported in intestinal and peripheral tissues in vivo. Further research is needed to clarify the importance of these microbial products in health-promoting properties associated with the intake of PACs.

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8. Conclusions and future perspectives

The health-promoting properties of PACs in the intestine are attributed not only to the antioxidant activity inherent to phenolic compounds, but also to the capacity of these phytochemicals to interact with multiple biomolecules, including proteins, biomembrane lipids, and endotoxins. Bioactivity of PACs is highly structure-dependent and enriched botanical extracts composed by a large variety of molecular structures exert a wide range of unrelated physiological effects. In this way, PAC-rich extracts can modulate kinase activity, several signal transduction pathways implicated in the inflammatory response and the remodeling of TJs. Flavan-3-ol monomers and short PAC oligomers are absorbed by enterocytes and immune cells and exert a direct action on kinases and transcription factors. Bioactivity of larger oligomers and polymeric PACs do not require direct intestinal absorption and are able to bind protein receptors on the enterocyte and immune cell surfaces as well as luminal bacterial endotoxins, thus inhibiting pro-inflammatory signaling and improving barrier integrity. Due to the negligible absorption of large PAC molecules in the small intestine, phenyl-γ-valerolactones and phenolic acids produced by the microbiota metabolism in the colon are thought to play an important role in these health-promoting effects, and thus need to be further researched.

The barrier-protective properties of PACs are emerging as a potential adjunctive support to current therapies for managing obesity related intestinal dysfunction and IBD. However, there have been no large, well-designed clinical trials establishing the efficacy of these phytochemicals in chronic conditions. At preclinical stages, the use of animal models is the predominant approach for testing novel therapies for intestinal dysfunction, although several strategies for replacing animal experiments have been proposed. As there are still no studies on the impact of PACs on human intestinal health, ex vivo models of the human intestine could be a more physiologically reliable alternative to human cell lines and an alternative to animal experimentation in preclinical development.

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Acknowledgments

C. González-Quilen has received financial support through a FI-AGAUR grant from the Generalitat de Catalunya. M. Pinent and X. Terra are Serra-Húnter fellows at the Universitat Rovira i Virgili, Tarragona, Spain.

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Conflicts of interest

The authors declare no conflict of interest.

Funding

This work was funded by the Spanish Ministerio de Economía y Competitividad (Grant: AGL2017-83477-R).

References

  1. 1. Chelakkot C, Ghim J, Ryu SH. Mechanisms regulating intestinal barrier integrity and its pathological implications. Experimental & Molecular Medicine. 2018;50:103
  2. 2. Fukui H. Increased intestinal permeability and decreased barrier function: Does it really influence the risk of inflammation? Inflammatory Intestinal Diseases. 2016;1:135-145. DOI: 10.1159/000447252
  3. 3. Clemente-Postigo M, Oliva-Olivera W, Coin-Aragüez L, Ramos-Molina B, Giraldez-Perez RM, Lhamyani S, et al. Metabolic endotoxemia promotes adipose dysfunction and inflammation in human obesity. American Journal of Physiology-Endocrinology and Metabolism. 2019;316:E319-E332. DOI: 10.1152/ajpendo.00277.2018
  4. 4. Harvey A. Strategies for discovering drugs from previously unexplored natural products. Drug Discovery Today. 2000;5:294-300
  5. 5. Costa C, Tsatsakis A, Mamoulakis C, Teodoro M, Briguglio G, Caruso E, et al. Current evidence on the effect of dietary polyphenols intake on chronic diseases. Food and Chemical Toxicology. 2017;110:286-299
  6. 6. Smeriglio A, Barreca D, Bellocco E, Trombetta D. Proanthocyanidins and hydrolysable tannins: Occurrence, dietary intake and pharmacological effects. British Journal of Pharmacology. 2017;174:1244-1262. DOI: 10.1111/bph.13630
  7. 7. Bladé C, Aragonès G, Arola-Arnal A, Muguerza B, Bravo FI, Salvadó MJ, et al. Proanthocyanidins in health and disease. BioFactors. 2016;42:5-12. DOI: 10.1002/biof.1249
  8. 8. Gil-Cardoso K, Ginés I, Pinent M, Ardévol A, Blay M, Terra X. The co-administration of proanthocyanidins and an obesogenic diet prevents the increase in intestinal permeability and metabolic endotoxemia derived to the diet. The Journal of Nutritional Biochemistry. 2018;62:35-42. DOI: 10.1016/J.JNUTBIO.2018.07.012
  9. 9. Gil-Cardoso K, Ginés I, Pinent M, Ardévol A, Blay M, Terra X. Effects of flavonoids on intestinal inflammation, barrier integrity and changes in gut microbiota during diet-induced obesity. Nutrition Research Reviews. 2016;29:1-15. DOI: 10.1017/S0954422416000159
  10. 10. Wu H, Luo T, Li YM, Gao ZP, Zhang KQ, Song JY, et al. Granny Smith apple procyanidin extract upregulates tight junction protein expression and modulates oxidative stress and inflammation in lipopolysaccharide-induced Caco-2 cells. Food & Function. 2018;9:3321-3329. DOI: 10.1039/c8fo00525g
  11. 11. Terra X, Valls J, Vitrac X, Mérrillon J-M, Arola L, Ardèvol A, et al. Grape-seed procyanidins act as antiinflammatory agents in endotoxin-stimulated RAW 264.7 macrophages by inhibiting NFkB signaling pathway. Journal of Agricultural and Food Chemistry. 2007;55:4357-4365. DOI: 10.1021/jf0633185
  12. 12. Martinez-Micaelo N, González-Abuín N, Pinent M, Ardévol A, Blay M. Procyanidin B2 inhibits inflammasome-mediated IL-1β production in lipopolysaccharide-stimulated macrophages. Molecular Nutrition & Food Research. 2015;59:262-269. DOI: 10.1002/mnfr.201400370
  13. 13. Gil-Cardoso K, Comitato R, Ginés I, Ardévol A, Pinent M, Virgili F, et al. Protective effect of proanthocyanidins in a rat model of mild intestinal inflammation and impaired intestinal permeability induced by LPS. Molecular Nutrition & Food Research. 2019;63:1800720. DOI: 10.1002/mnfr.201800720
  14. 14. Gil-Cardoso K, Ginés I, Pinent M, Ardévol A, Arola L, Blay M, et al. Chronic supplementation with dietary proanthocyanidins protects from diet-induced intestinal alterations in obese rats. Molecular Nutrition & Food Research. 2017;61:1601039. DOI: 10.1002/mnfr.201601039
  15. 15. González-Quilen C, Gil-Cardoso K, Ginés I, Beltrán-Debón R, Pinent M, Ardévol A, et al. Grape-seed proanthocyanidins are able to reverse intestinal dysfunction and metabolic endotoxemia induced by a cafeteria diet in wistar rats. Nutrients. 2019;11:pii: E979. DOI: 10.3390/nu11050979
  16. 16. Allaire JM, Crowley SM, Law HT, Chang SY, Ko HJ, Vallance BA. The intestinal epithelium: Central coordinator of mucosal immunity. Trends in Immunology. 2018;39:677-696
  17. 17. Bischoff SC, Barbara G, Buurman W, Ockhuizen T, Schulzke J-D, Serino M, et al. Intestinal permeability—A new target for disease prevention and therapy. BMC Gastroenterology. 2014;14:189. DOI: 10.1186/s12876-014-0189-7
  18. 18. Birchenough GMH, Johansson MEV, Gustafsson JK, Bergström JH, Hansson GC. New developments in goblet cell mucus secretion and function. Mucosal Immunology. 2015;8:712-719. DOI: 10.1038/mi.2015.32
  19. 19. Nakamura Y, Kimura S, Hase K. M cell-dependent antigen uptake on follicle-associated epithelium for mucosal immune surveillance. Inflammation and Regeneration. 2018;38:15. DOI: 10.1186/s41232-018-0072-y
  20. 20. Fiocchi C. What is “physiological” intestinal inflammation and how does it differ from “pathological” inflammation? Inflammatory Bowel Diseases. 2008;14:S77-S78. DOI: 10.1097/00054725-200810001-00040
  21. 21. Belkaid Y, Harrison OJ. Homeostatic immunity and the microbiota. Immunity. 2017;46:562-576
  22. 22. Ulluwishewa D, Anderson RC, McNabb WC, Moughan PJ, Wells JM, Roy NC. Regulation of tight junction permeability by intestinal bacteria and dietary components. The Journal of Nutrition. 2011;141:769-776. DOI: 10.3945/jn.110.135657
  23. 23. Luo H, Guo P, Zhou Q. Role of TLR4/NF-κB in damage to intestinal mucosa barrier function and bacterial translocation in rats exposed to hypoxia. PLoS One. 2012;7:e46291. DOI: 10.1371/journal.pone.0046291
  24. 24. Suzuki T. Regulation of intestinal epithelial permeability by tight junctions. Cellular and Molecular Life Sciences. 2013;70:631-659. DOI: 10.1007/s00018-012-1070-x
  25. 25. Suzuki M, Nagaishi T, Yamazaki M, Onizawa M, Watabe T, Sakamaki Y, et al. Myosin light chain kinase expression induced via tumor necrosis factor receptor 2 signaling in the epithelial cells regulates the development of colitis-associated carcinogenesis. PLoS One. 2014;9:e88369. DOI: 10.1371/journal.pone.0088369
  26. 26. Barreau F, Hugot JP. Intestinal barrier dysfunction triggered by invasive bacteria. Current Opinion in Microbiology. 2014;17:91-98. DOI: 10.1016/j.mib.2013.12.003
  27. 27. Al-Sadi R, Ye D, Dokladny K, Ma TY. Mechanism of IL-1beta-induced increase in intestinal epithelial tight junction permeability. Journal of Immunology. 2008;180:5653-5661. DOI: 180/8/5653 [pii]
  28. 28. Al-Sadi R, Guo S, Ye D, Dokladny K, Alhmoud T, Ereifej L, et al. Mechanism of IL-1β modulation of intestinal epithelial barrier involves p38 kinase and activating transcription factor-2 activation. Journal of Immunology. 2013;190:6596-6606. DOI: 10.4049/jimmunol.1201876
  29. 29. Ye D, Ma TY. Cellular and molecular mechanisms that mediate basal and tumour necrosis factor-alpha-induced regulation of myosin light chain kinase gene activity. Journal of Cellular and Molecular Medicine. 2008;12:1331-1346. DOI: 10.1111/j.1582-4934.2008.00302.x
  30. 30. Luo H, Guo P, Zhou Q. Role of TLR4/NF-kappaB in damage to intestinal mucosa barrier function and bacterial translocation in rats exposed to hypoxia. PLoS One. 2012;7:e46291. DOI: 10.1371/journal.pone.0046291
  31. 31. Laugerette F, Furet J-P, Debard C, Daira P, Loizon E, Geloen a, et al. Oil composition of high-fat diet affects metabolic inflammation differently in connection with endotoxin receptors in mice. American Journal of Physiology-Endocrinology and Metabolism. 2012;302:E374-E386. DOI: 10.1152/ajpendo.00314.2011
  32. 32. Li Q, Verma IM. NF-kappaB regulation in the immune system. Nature Reviews. Immunology. 2002;2:725-734. DOI: 10.1038/nri910
  33. 33. Hayden MS, West AP, Ghosh S. NF-kappaB and the immune response. Oncogene. 2006;25:6758-6780. DOI: 10.1038/sj.onc.1209943
  34. 34. Pahl HL. Activators and target genes of Rel/NF-kappaB transcription factors. Oncogene. 1999;18:6853-6866. DOI: 10.1038/sj.onc.1203239
  35. 35. Toumi R, Soufli I, Rafa H, Belkhelfa M, Biad A, Touil-Boukoffa C. Probiotic bacteriaLactobacillusandBifidobacteriumattenuate inflammation in dextran sulfate sodium-induced experimental colitis in mice. International Journal of Immunopathology and Pharmacology. 2014;27:615-627
  36. 36. Wang W, Xia T, Yu X. Wogonin suppresses inflammatory response and maintains intestinal barrier function via TLR4-MyD88-TAK1-mediated NF-κB pathway in vitro. Inflammation Research. 2015;64:423-431. DOI: 10.1007/s00011-015-0822-0
  37. 37. Kolios G, Valatas V, Ward SG. Nitric oxide in inflammatory bowel disease: A universal messenger in an unsolved puzzle. Immunology. 2004;113:427-437. DOI: 10.1111/j.1365-2567.2004.01984.x
  38. 38. Martinez-Micaelo N, Gonzalez-Abuin N, Terra X, Richart C, Ardevol A, Pinent M, et al. Omega-3 docosahexaenoic acid and procyanidins inhibit cyclo-oxygenase activity and attenuate NF-kappaB activation through a p105/p50 regulatory mechanism in macrophage inflammation. The Biochemical Journal. 2012;441:653-663. DOI: 10.1042/BJ20110967
  39. 39. Du CYQ, Choi RCY, Dong TTX, Lau DTW, Tsim KWK. Yu Ping Feng San, an ancient Chinese herbal decoction, regulates the expression of inducible nitric oxide synthase and cyclooxygenase-2 and the activity of intestinal alkaline phosphatase in cultures. PLoS One. 2014;9:e100382. DOI: 10.1371/journal.pone.0100382
  40. 40. Ananthakrishnan AN, Bernstein CN, Iliopoulos D, Macpherson A, Neurath MF, Ali RAR, et al. Environmental triggers in IBD: A review of progress and evidence. Nature Reviews. Gastroenterology & Hepatology. 2018;15:39-49
  41. 41. Huang S, Rutkowsky JM, Snodgrass RG, Ono-Moore KD, Schneider DA, Newman JW, et al. Saturated fatty acids activate TLR-mediated proinflammatory signaling pathways. Journal of Lipid Research. 2012;53:2002-2013. DOI: 10.1194/jlr.D029546
  42. 42. Wong SW, Kwon MJ, Choi AMK, Kim HP, Nakahira K, Hwang DH. Fatty acids modulate toll-like receptor 4 activation through regulation of receptor dimerization and recruitment into lipid rafts in a reactive oxygen species-dependent manner. The Journal of Biological Chemistry. 2009;284:27384-27392. DOI: 10.1074/jbc.M109.044065
  43. 43. Gil-Cardoso K, Ginés I, Pinent M, Ardévol A, Terra X, Blay M. A cafeteria diet triggers intestinal inflammation and oxidative stress in obese rats. The British Journal of Nutrition. 2017;117:218. DOI: 10.1017/S0007114516004608
  44. 44. Cani PD, Bibiloni R, Knauf C, Waget A, Neyrinck AM, Delzenne NM, et al. Changes in gut microbiota control metabolic endotoxemia-induced inflammation in high-fat diet-induced obesity and diabetes in mice. Diabetes. 2008;57:1470-1481. DOI: 10.2337/db07-1403
  45. 45. Serrano J, Puupponen-Pimiä R, Dauer A, Aura AM, Saura-Calixto F. Tannins: Current knowledge of food sources, intake, bioavailability and biological effects. Molecular Nutrition & Food Research. 2009;53:310-329. DOI: 10.1002/mnfr.200900039
  46. 46. Bittner K, Rzeppa S, Humpf H-U. Distribution and quantification of flavan-3-ols and procyanidins with low degree of polymerization in nuts, cereals, and legumes. Journal of Agricultural and Food Chemistry. 2013;61:9148-9154. DOI: 10.1021/jf4024728
  47. 47. Rue EA, Rush MD, van Breemen RB. Procyanidins: A comprehensive review encompassing structure elucidation via mass spectrometry. Phytochemistry Reviews. 2018;17:1-16. DOI: 10.1007/s11101-017-9507-3
  48. 48. Wang Y, Chung S-J, Song WO, Chun OK. Estimation of daily proanthocyanidin intake and major food sources in the U.S. diet. The Journal of Nutrition. 2011;141:447-452. DOI: 10.3945/jn.110.133900
  49. 49. Jun S, Shin S, Joung H. Estimation of dietary flavonoid intake and major food sources of Korean adults. The British Journal of Nutrition. 2016;115:480-489. DOI: 10.1017/S0007114515004006
  50. 50. Zamora-Ros R, Biessy C, Rothwell JA, Monge A, Lajous M, Scalbert A, et al. Dietary polyphenol intake and their major food sources in the Mexican Teachers’ Cohort. The British Journal of Nutrition. 2018;120:353-360. DOI: 10.1017/S0007114518001381
  51. 51. Vogiatzoglou A, Mulligan AA, Luben RN, Lentjes MAH, Heiss C, Kelm M, et al. Assessment of the dietary intake of total flavan-3-ols, monomeric flavan-3-ols, proanthocyanidins and theaflavins in the European Union. The British Journal of Nutrition. 2014;111:1463-1473. DOI: 10.1017/S0007114513003930
  52. 52. Zamora-Ros R, Knaze V, Rothwell JA, Hémon B, Moskal A, Overvad K, et al. Dietary polyphenol intake in Europe: The European Prospective Investigation into Cancer and Nutrition (EPIC) Study. European Journal of Nutrition. 2016;55:1359-1375. DOI: 10.1007/s00394-015-0950-x
  53. 53. Knaze V, Zamora-Ros R, Luján-Barroso L, Romieu I, Scalbert A, Slimani N, et al. Intake estimation of total and individual flavan-3-ols, proanthocyanidins and theaflavins, their food sources and determinants in the European Prospective Investigation into Cancer and Nutrition (EPIC) study. The British Journal of Nutrition. 2012;108:1095-1108. DOI: 10.1017/S0007114511006386
  54. 54. Rios LY, Bennett RN, Lazarus SA, Remesy C, Scalbert A, Williamson G. Cocoa procyanidins are stable during gastric transit in humans. The American Journal of Clinical Nutrition. 2002;76:1106-1110
  55. 55. Donovan JL, Crespy V, Manach C, Morand C, Besson C, Scalbert A, et al. Catechin is metabolized by both the small intestine and liver of rats. The Journal of Nutrition. 2001;131:1753-1757. DOI: 10.1093/jn/131.6.1753
  56. 56. Stalmach A, Mullen W, Steiling H, Williamson G, Lean MEJ, Crozier A. Absorption, metabolism, and excretion of green tea flavan-3-ols in humans with an ileostomy. Molecular Nutrition & Food Research. 2010;54:323-334. DOI: 10.1002/mnfr.200900194
  57. 57. Thilakarathna SH, Vasantha Rupasinghe HP. Flavonoid bioavailability and attempts for bioavailability enhancement. Nutrients. 2013;5:3367-3387. DOI: 10.3390/nu5093367
  58. 58. Rein D, Lotito S, Holt RR, Keen CL, Schmitz HH, Fraga CG. Epicatechin in human plasma: In vivo determination and effect of chocolate consumption on plasma oxidation status. The Journal of Nutrition. 2000;130:2109S-2114S. DOI: 10.1093/jn/130.8.2109S
  59. 59. Ullmann U, Haller J, Decourt JP, Girault N, Girault J, Richard-Caudron AS, et al. A single ascending dose study of epigallocatechin gallate in healthy volunteers. The Journal of International Medical Research. 2003;31:88-101. DOI: 10.1177/147323000303100205
  60. 60. Wiese S, Esatbeyoglu T, Winterhalter P, Kruse HP, Winkler S, Bub A, et al. Comparative biokinetics and metabolism of pure monomeric, dimeric, and polymeric flavan-3-ols: A randomized cross-over study in humans. Molecular Nutrition & Food Research. 2015;59:610-621. DOI: 10.1002/mnfr.201400422
  61. 61. Deprez S, Mila I, Huneau J-F, Tome D, Scalbert A. Transport of proanthocyanidin dimer, trimer, and polymer across monolayers of human intestinal epithelial Caco-2 cells. Antioxidants & Redox Signaling. 2001;3:957-967. DOI: 10.1089/152308601317203503
  62. 62. Zumdick S, Deters A, Hensel A. In vitro intestinal transport of oligomeric procyanidins (DP 2 to 4) across monolayers of Caco-2 cells. Fitoterapia. 2012;83:1210-1217. DOI: 10.1016/j.fitote.2012.06.013
  63. 63. Mendoza-Wilson AM, Carmelo-Luna FJ, Astiazarán-García H, Mata-Haro V, Espinosa-Plascencia A, del Carmen Bermúdez-Almada M, et al. Absorption of dimers, trimers and tetramers of procyanidins present in apple skin by IEC-18 cell monolayers. Journal of Functional Foods. 2016;27:386-391. DOI: 10.1016/J.JFF.2016.09.020
  64. 64. Shoji T, Masumoto S, Moriichi N, Akiyama H, Kanda T, Ohtake Y, et al. Apple procyanidin oligomers absorption in rats after oral administration: Analysis of procyanidins in plasma using the porter method and high-performance liquid chromatography/tandem mass spectrometry. Journal of Agricultural and Food Chemistry. 2006;54:884-892. DOI: 10.1021/jf052260b
  65. 65. Serra A, MacI A, Romero MP, Valls J, Bladé C, Arola L, et al. Bioavailability of procyanidin dimers and trimers and matrix food effects in in vitro and in vivo models. The British Journal of Nutrition. 2010;103:944-952. DOI: 10.1017/S0007114509992741
  66. 66. Sano A, Yamakoshi J, Tokutake S, Tobe K, Kubota Y, Kikuchi M. Procyanidin B1 is detected in human serum after intake of proanthocyanidin-rich grape seed extract. Bioscience, Biotechnology, and Biochemistry. 2003;67:1140-1143
  67. 67. Ottaviani JI, Kwik-Uribe C, Keen CL, Schroeter H. Intake of dietary procyanidins does not contribute to the pool of circulating flavanols in humans. The American Journal of Clinical Nutrition. 2012;95:851-858. DOI: 10.3945/ajcn.111.028340
  68. 68. Tsang C, Auger C, Mullen W, Bornet A, Rouanet J-M, Crozier A, et al. The absorption, metabolism and excretion of flavan-3-ols and procyanidins following the ingestion of a grape seed extract by rats. The British Journal of Nutrition. 2005;94:170-181
  69. 69. Gu L, House SE, Rooney L, Prior RL. Sorghum bran in the diet dose dependently increased the excretion of catechins and microbial-derived phenolic acids in female rats. Journal of Agricultural and Food Chemistry. 2007;55:5326-5334. DOI: 10.1021/jf070100p
  70. 70. Appeldoorn MM, Vincken JP, Aura AM, Hollman PCH, Gruppen H. Procyanidin dimers are metabolized by human microbiota with 2-(3,4-dihydroxyphenyl)acetic acid and 5-(3,4-dihydroxyphenyl)-γ-valerolactone as the major metabolites. Journal of Agricultural and Food Chemistry. 2009;57:1084-1092. DOI: 10.1021/jf803059z
  71. 71. Castello F, Costabile G, Bresciani L, Tassotti M, Naviglio D, Luongo D, et al. Bioavailability and pharmacokinetic profile of grape pomace phenolic compounds in humans. Archives of Biochemistry and Biophysics. 2018;646:1-9. DOI: 10.1016/j.abb.2018.03.021
  72. 72. Trošt K, Ulaszewska MM, Stanstrup J, Albanese D, De Filippo C, Tuohy KM, et al. Host: Microbiome co-metabolic processing of dietary polyphenols—An acute, single blinded, cross-over study with different doses of apple polyphenols in healthy subjects. Food Research International. 2018;112:108-128. DOI: 10.1016/j.foodres.2018.06.016
  73. 73. Margalef M, Pons Z, Bravo FI, Muguerza B, Arola-Arnal A. Tissue distribution of rat flavanol metabolites at different doses. The Journal of Nutritional Biochemistry. 2015;26:987-995. DOI: 10.1016/j.jnutbio.2015.04.006
  74. 74. Yoshioka Y, Akiyama H, Nakano M, Shoji T, Kanda T, Ohtake Y, et al. Orally administered apple procyanidins protect against experimental inflammatory bowel disease in mice. International Immunopharmacology. 2008;8:1802-1807. DOI: 10.1016/j.intimp.2008.08.021
  75. 75. Denis M-C, Desjardins Y, Furtos A, Marcil V, Dudonné S, Montoudis A, et al. Prevention of oxidative stress, inflammation and mitochondrial dysfunction in the intestine by different cranberry phenolic fractions. Clinical Science. 2015;128:197-212. DOI: 10.1042/CS20140210
  76. 76. Erlejman AG, Jaggers G, Fraga CG, Oteiza PI. TNFα-induced NF-κB activation and cell oxidant production are modulated by hexameric procyanidins in Caco-2 cells. Archives of Biochemistry and Biophysics. 2008;476:186-195. DOI: 10.1016/j.abb.2008.01.024
  77. 77. Gentile C, Perrone A, Attanzio A, Tesoriere L, Livrea MA. Sicilian pistachio (Pistacia veraL.) nut inhibits expression and release of inflammatory mediators and reverts the increase of paracellular permeability in IL-1β-exposed human intestinal epithelial cells. European Journal of Nutrition. 2015;54:811-821. DOI: 10.1007/s00394-014-0760-6
  78. 78. Bitzer ZT, Glisan SL, Dorenkott MR, Goodrich KM, Ye L, O’Keefe SF, et al. Cocoa procyanidins with different degrees of polymerization possess distinct activities in models of colonic inflammation. The Journal of Nutritional Biochemistry. 2015;26:827-831. DOI: 10.1016/j.jnutbio.2015.02.007
  79. 79. Bianchi MG, Chiu M, Taurino G, Brighenti F, Del Rio D, Mena P, et al. Catechin and Procyanidin B2 modulate the expression of tight junction proteins but do not protect from inflammation-induced changes in permeability in human intestinal cell monolayers. Nutrients. 2019;11:2271. DOI: 10.3390/nu11102271
  80. 80. Wong X, Carrasco-Pozo C, Escobar E, Navarrete P, Blachier F, Andriamihaja M, et al. Deleterious effect of p-cresol on human colonic epithelial cells prevented by proanthocyanidin-containing polyphenol extracts from fruits and proanthocyanidin bacterial metabolites. Journal of Agricultural and Food Chemistry. 2016;64:3574-3583. DOI: 10.1021/acs.jafc.6b00656
  81. 81. Xu H, Zhao C, Li Y, Liu R, Ao M, Li F, et al. The ameliorative effect of the:Pyracantha fortuneana(Maxim.) H. L. Li extract on intestinal barrier dysfunction through modulating glycolipid digestion and gut microbiota in high fat diet-fed rats. Food & Function. 2019;10:6517-6532. DOI: 10.1039/c9fo01599j
  82. 82. Wang Y-H, Yang X-L, Wang L, Cui M-X, Cai Y-Q, Li X, et al. Effects of proanthocyanidins from grape seed on treatment of recurrent ulcerative colitis in rats. Canadian Journal of Physiology and Pharmacology. 2010;898:888-898. DOI: 10.1139/Y10-071
  83. 83. Wang Y-H, Ge B, Yang X-L, Zhai J, Yang L-N, Wang X-X, et al. Proanthocyanidins from grape seeds modulates the nuclear factor-kappa B signal transduction pathways in rats with TNBS-induced recurrent ulcerative colitis. International Immunopharmacology. 2011;11:1620-1627. DOI: 10.1016/j.intimp.2011.05.024
  84. 84. Li X, Yang X, Cai Y, Qin H, Wang L, Wang Y, et al. Proanthocyanidins from grape seeds modulate the NF-κB signal transduction pathways in rats with TNBS-induced ulcerative colitis. Molecules. 2011;16:6721-6731. DOI: 10.3390/molecules16086721
  85. 85. Chen L, You Q, Hu L, Gao J, Meng Q, Liu W, et al. The antioxidant procyanidin reduces reactive oxygen species signaling in macrophages and ameliorates experimental colitis in mice. Frontiers in Immunology. 2018;8:1910. DOI: 10.3389/fimmu.2017.01910
  86. 86. Wang H, Xue Y, Zhang H, Huang Y, Yang G, Du M, et al. Dietary grape seed extract ameliorates symptoms of inflammatory bowel disease in IL10-deficient mice. Molecular Nutrition & Food Research. 2013;57:2253-2257. DOI: 10.1002/mnfr.201300146
  87. 87. Bibi S, Kang Y, Yang G, Zhu M-J. Grape seed extract improves small intestinal health through suppressing inflammation and regulating alkaline phosphatase in IL-10-deficient mice. Journal of Functional Foods. 2016;20:245-252. DOI: 10.1016/J.JFF.2015.10.021
  88. 88. He X, Sun LM. Dietary intake of flavonoid subclasses and risk of colorectal cancer: Evidence from population studies. Oncotarget. 2016;7:26617-26627. DOI: 10.18632/oncotarget.8562
  89. 89. Sambuy Y, De Angelis I, Ranaldi G, Scarino ML, Stammati A, Zucco F. The Caco-2 cell line as a model of the intestinal barrier: Influence of cell and culture-related factors on Caco-2 cell functional characteristics. Cell Biology and Toxicology. 2005;21:1-26
  90. 90. Kämpfer AAM, Urbán P, Gioria S, Kanase N, Stone V, Kinsner-Ovaskainen A. Development of an in vitro co-culture model to mimic the human intestine in healthy and diseased state. Toxicology in Vitro. 2017;45:31-43. DOI: 10.1016/j.tiv.2017.08.011
  91. 91. Chassaing B, Rolhion N, De Vallée A, Salim SY, Prorok-Hamon M, Neut C, et al. Crohn disease-associated adherent-invasiveE. colibacteria target mouse and human Peyer’s patches via long polar fimbriae. The Journal of Clinical Investigation. 2011;121:966-975. DOI: 10.1172/JCI44632
  92. 92. Goh JY, Weaver RJ, Dixon L, Platt NJ, Roberts RA. Development and use of in vitro alternatives to animal testing by the pharmaceutical industry 1980–2013. Toxicology Research (Cambridge). 2015;4:1297-1307. DOI: 10.1039/c5tx00123d
  93. 93. Kang TH, Kim HJ. Farewell to animal testing: Innovations on human intestinal microphysiological systems. Micromachines. 2016;7:107
  94. 94. Randhawa PK, Singh K, Singh N, Jaggi AS. A review on chemical-induced inflammatory bowel disease models in rodents. The Korean Journal of Physiology & Pharmacology. 2014;18:279-288. DOI: 10.4196/kjpp.2014.18.4.279
  95. 95. Hidalgo IJ, Raub TJ, Borchardt RT. Characterization of the human colon carcinoma cell line (Caco-2) as a model system for intestinal epithelial permeability. Gastroenterology. 1989;96:736-749. DOI: 10.1016/S0016-5085(89)80072-1
  96. 96. Srinivasan B, Kolli AR, Esch MB, Abaci HE, Shuler ML, Hickman JJ. TEER measurement techniques for in vitro barrier model systems. Journal of Laboratory Automation. 2015;20:107-126. DOI: 10.1177/2211068214561025
  97. 97. Sampey BP, Vanhoose AM, Winfield HM, Freemerman AJ, Muehlbauer MJ, Fueger PT, et al. Cafeteria diet is a robust model of human metabolic syndrome with liver and adipose inflammation: Comparison to high-fat diet. Obesity (Silver Spring). 2011;19:1109-1117. DOI: 10.1038/oby.2011.18
  98. 98. Ginés I, Gil-Cardoso K, Serrano J, Casanova-Martí À, Blay M, Pinent M, et al. Effects of an intermittent grape-seed proanthocyanidin (GSPE) treatment on a cafeteria diet obesogenic challenge in rats. Nutrients. 2018;10:315. DOI: 10.3390/nu10030315
  99. 99. Baselga-Escudero L, Pascual-Serrano A, Ribas-Latre A, Casanova E, Salvadó MJ, Arola L, et al. Long-term supplementation with a low dose of proanthocyanidins normalized liver miR-33a and miR-122 levels in high-fat diet-induced obese rats. Nutrition Research. 2015;35:337-345. DOI: 10.1016/j.nutres.2015.02.008
  100. 100. Margalef M, Pons Z, Iglesias-Carres L, Arola L, Muguerza B, Arola-Arnal A. Gender-related similarities and differences in the body distribution of grape seed flavanols in rats. Molecular Nutrition & Food Research. 2016;60:760-772. DOI: 10.1002/mnfr.201500717
  101. 101. Terra X, Pallarés V, Ardèvol A, Bladé C, Fernández-Larrea J, Pujadas G, et al. Modulatory effect of grape-seed procyanidins on local and systemic inflammation in diet-induced obesity rats. The Journal of Nutritional Biochemistry. 2011;22:380-387. DOI: 10.1016/j.jnutbio.2010.03.006
  102. 102. Li X-L, Cai Y-Q, Qin H, Wu Y-J. Therapeutic effect and mechanism of proanthocyanidins from grape seeds in rats with TNBS-induced ulcerative colitis. Canadian Journal of Physiology and Pharmacology. 2008;86:841-849. DOI: 10.1139/Y08-089
  103. 103. Hollman PCH, Cassidy A, Comte B, Heinonen M, Richelle M, Richling E, et al. The biological relevance of direct antioxidant effects of polyphenols for cardiovascular health in humans is not established. The Journal of Nutrition. 2011;141:989S-1009S. DOI: 10.3945/jn.110.131490
  104. 104. Tsilingiri K, Sonzogni A, Caprioli F, Rescigno M. A novel method for the culture and polarized stimulation of human intestinal mucosa explants. Journal of Visualized Experiments. 2013:e4368. DOI: 10.3791/4368
  105. 105. Vadstrup K, Galsgaard ED, Gerwien J, Vester-Andersen MK, Pedersen JS, Rasmussen J, et al. Validation and optimization of an ex vivo assay of intestinal mucosal biopsies in Crohn’s disease: Reflects inflammation and drug effects. PLoS One. 2016;11:e0155335. DOI: 10.1371/journal.pone.0155335
  106. 106. Sjöberg Å, Lutz M, Tannergren C, Wingolf C, Borde A, Ungell A-L. Comprehensive study on regional human intestinal permeability and prediction of fraction absorbed of drugs using the Ussing chamber technique. European Journal of Pharmaceutical Sciences. 2013;48:166-180. DOI: 10.1016/J.EJPS.2012.10.007
  107. 107. Geraedts MCP, Troost FJ, De Ridder RJ, Bodelier AGL, Masclee AAM, Saris WHM. Validation of Ussing chamber technology to study satiety hormone release from human duodenal specimens. Obesity. 2012;20:678-682. DOI: 10.1038/oby.2011.104
  108. 108. Wood MW, Breitschwerdt EB, Nordone SK, Linder KE, Gookin JL. UropathogenicE. colipromote a paracellular urothelial barrier defect characterized by altered tight junction integrity, epithelial cell sloughing and cytokine release. Journal of Comparative Pathology. 2012;147:11-19. DOI: 10.1016/j.jcpa.2011.09.005
  109. 109. Ray S, Bagchi D, Lim PM, Bagchi M, Gross SM, Kothari SC, et al. Acute and long-term safety evaluation of a novel IH636 grape seed proanthocyanidin extract. Research Communications in Molecular Pathology and Pharmacology. 2001;109:165-197
  110. 110. Yamakoshi J, Saito M, Kataoka S, Kikuchi M. Safety evaluation of proanthocyanidin-rich extract from grape seeds. Food and Chemical Toxicology. 2002;40:599-607. DOI: 10.1016/s0278-6915(02)00006-6
  111. 111. Lluís L, Muñoz M, Rosa Nogués M, Sánchez-Martos V, Romeu M, Giralt M, et al. Toxicology evaluation of a procyanidin-rich extract from grape skins and seeds. Food and Chemical Toxicology. 2011;49:1450-1454. DOI: 10.1016/j.fct.2011.03.042
  112. 112. Wong X, Madrid AM, Tralma K, Castillo R, Carrasco-Pozo C, Navarrete P, et al. Polyphenol extracts interfere with bacterial lipopolysaccharide in vitro and decrease postprandial endotoxemia in human volunteers. Journal of Functional Foods. 2016;26:406-417. DOI: 10.1016/J.JFF.2016.08.011
  113. 113. Cani PD, Osto M, Geurts L, Everard A. Involvement of gut microbiota in the development of low-grade inflammation and type 2 diabetes associated with obesity. Gut Microbes. 2012;3:279-288
  114. 114. Butler LG. Effects of condensed tannin on animal nutrition. In: Chemistry and Significance of Condensed Tannins. US: Springer; 1989. pp. 391-402
  115. 115. Chung KT, Wong TY, Wei CI, Huang YW, Lin Y. Tannins and human health: A review. Critical Reviews in Food Science and Nutrition. 1998;38:421-464. DOI: 10.1080/10408699891274273
  116. 116. Brás NF, Gonçalves R, Fernandes PA, Mateus N, Ramos MJ, De Freitas V. Understanding the binding of procyanidins to pancreatic elastase by experimental and computational methods. Biochemistry. 2010;49:5097-5108. DOI: 10.1021/bi100410q
  117. 117. De Freitas V, Mateus N. Structural features of procyanidin interactions with salivary proteins. Journal of Agricultural and Food Chemistry. 2001;49:940-945. DOI: 10.1021/jf000981z
  118. 118. Zhu W, Xiong L, Peng J, Deng X, Gao J, Li CM. Molecular insight into affinities of gallated and nongallated proanthocyanidins dimers to lipid bilayers. Scientific Reports. 2016;6:37680. DOI: 10.1038/srep37680
  119. 119. Verstraeten SV, Jaggers GK, Fraga CG, Oteiza PI. Procyanidins can interact with Caco-2 cell membrane lipid rafts: Involvement of cholesterol. Biochimica et Biophysica Acta, Biomembranes. 2013;1828:2646-2653. DOI: 10.1016/j.bbamem.2013.07.023
  120. 120. Yang G, Bibi S, Du M, Suzuki T, Zhu MJ. Regulation of the intestinal tight junction by natural polyphenols: A mechanistic perspective. Critical Reviews in Food Science and Nutrition. 2017;57:3830-3839. DOI: 10.1080/10408398.2016.1152230
  121. 121. Cunningham KE, Turner JR. Myosin light chain kinase: Pulling the strings of epithelial tight junction function. Annals of the New York Academy of Sciences. 2012;1258:34-42. DOI: 10.1111/j.1749-6632.2012.06526.x
  122. 122. Marchiando AM, Shen L, Graham WV, Edelblum KL, Duckworth CA, Guan Y, et al. The epithelial barrier is maintained by in vivo tight junction expansion during pathologic intestinal epithelial shedding. Gastroenterology. 2011;140:1208.e2-1218.e2. DOI: 10.1053/j.gastro.2011.01.004
  123. 123. Wang BH, Lai Yeap F, Polya GM. Differential inhibition of eukaryote protein kinases by condensed tannins. Phytochemistry. 1996;43:359-365. DOI: 10.1016/0031-9422(96)00259-2
  124. 124. Contreras TC, Ricciardi E, Cremonini E, Oteiza PI. (−)-Epicatechin in the prevention of tumor necrosis alpha-induced loss of Caco-2 cell barrier integrity. Archives of Biochemistry and Biophysics. 2015;573:84-91. DOI: 10.1016/j.abb.2015.01.024
  125. 125. Delehanty JB, Johnson BJ, Hickey TE, Pons T, Ligler FS. Binding and neutralization of lipopolysaccharides by plant proanthocyanidins. Journal of Natural Products. 2007;70:1718-1724. DOI: 10.1021/np0703601
  126. 126. Williams AR, Klaver EJ, Laan LC, Ramsay A, Fryganas C, Difborg R, et al. Co-operative suppression of inflammatory responses in human dendritic cells by plant proanthocyanidins and products from the parasitic nematodeTrichuris suis. Immunology. 2017;150:312-328. DOI: 10.1111/imm.12687
  127. 127. Zmora N, Suez J, Elinav E. You are what you eat: Diet, health and the gut microbiota. Nature Reviews. Gastroenterology & Hepatology. 2019;16:35-56
  128. 128. Guirro M, Costa A, Gual-Grau A, Herrero P, Torrell H, Canela N, et al. Effects from diet-induced gut microbiota dysbiosis and obesity can be ameliorated by fecal microbiota transplantation: A multiomics approach. PLoS One. 2019;14:e0218143. DOI: 10.1371/journal.pone.0218143
  129. 129. Wan Y, Wang F, Yuan J, Li J, Jiang D, Zhang J, et al. Effects of dietary fat on gut microbiota and faecal metabolites, and their relationship with cardiometabolic risk factors: A 6-month randomised controlled-feeding trial. Gut. 2019;68:1417-1429. DOI: 10.1136/gutjnl-2018-317609
  130. 130. Segata N. Gut microbiome: Westernization and the disappearance of intestinal diversity. Current Biology. 2015;25:R611-R613
  131. 131. Buttó LF, Haller D. Dysbiosis in intestinal inflammation: Cause or consequence. International Journal of Medical Microbiology. 2016;306:302-309
  132. 132. Ley RE, Bäckhed F, Turnbaugh P, Lozupone CA, Knight RD, Gordon JI. Obesity alters gut microbial ecology. Proceedings of the National Academy of Sciences of the United States of America. 2005;102:11070-11075. DOI: 10.1073/pnas.0504978102
  133. 133. Lee SM, Kim N, Yoon H, Nam RH, Lee DH. Microbial changes and host response in F344 rat colon depending on sex and age following a high-fat diet. Frontiers in Microbiology. 2018;9:2236. DOI: 10.3389/fmicb.2018.02236
  134. 134. Ley RRE, Turnbaugh PJP, Klein S, Gordon JIJ. Microbial ecology: Human gut microbes associated with obesity. Nature. 2006;444:1022-1023. DOI: 10.1038/4441022a
  135. 135. Koliada A, Syzenko G, Moseiko V, Budovska L, Puchkov K, Perederiy V, et al. Association between body mass index andFirmicutes/Bacteroidetesratio in an adult Ukrainian population. BMC Microbiology. 2017;17:120. DOI: 10.1186/s12866-017-1027-1
  136. 136. Duncan SH, Lobley GE, Holtrop G, Ince J, Johnstone AM, Louis P, et al. Human colonic microbiota associated with diet, obesity and weight loss. International Journal of Obesity. 2008;32:1720-1724. DOI: 10.1038/ijo.2008.155
  137. 137. Schwiertz A, Taras D, Schäfer K, Beijer S, Bos NA, Donus C, et al. Microbiota and SCFA in lean and overweight healthy subjects. Obesity. 2010;18:190-195. DOI: 10.1038/oby.2009.167
  138. 138. Castaner O, Goday A, Park YM, Lee SH, Magkos F, Shiow SATE, et al. The gut microbiome profile in obesity: A systematic review. International Journal of Endocrinology. 2018;2018:1-9. DOI: 10.1155/2018/4095789
  139. 139. Garrett WS, Lord GM, Punit S, Lugo-Villarino G, Mazmanian SKK, Ito S, et al. Communicable ulcerative colitis induced by T-bet deficiency in the innate immune system. Cell. 2007;131:33-45. DOI: 10.1016/j.cell.2007.08.017
  140. 140. Karczewski J, Poniedziałek B, Adamski Z, Rzymski P. The effects of the microbiota on the host immune system. Autoimmunity. 2014;47:494-504
  141. 141. Etxeberria U, Fernández-Quintela A, Milagro FI, Aguirre L, Martínez JA, Portillo MP. Impact of polyphenols and polyphenol-rich dietary sources on gut microbiota composition. Journal of Agricultural and Food Chemistry. 2013;61:9517-9533. DOI: 10.1021/jf402506c
  142. 142. Tzounis X, Rodriguez-Mateos A, Vulevic J, Gibson GR, Kwik-Uribe C, Spencer JPE. Prebiotic evaluation of cocoa-derived flavanols in healthy humans by using a randomized, controlled, double-blind, crossover intervention study. The American Journal of Clinical Nutrition. 2011;93:62-72. DOI: 10.3945/ajcn.110.000075.Diet
  143. 143. Queipo-Ortuño MI, Boto-Ordóñez M, Murri M, Gomez-Zumaquero JM, Clemente-Postigo M, Estruch R, et al. Influence of red wine polyphenols and ethanol on the gut microbiota ecology and biochemical biomarkers. The American Journal of Clinical Nutrition. 2012;95:1323-1334. DOI: 10.3945/ajcn.111.027847
  144. 144. Hervert-Hernández D, Goñi I. Dietary polyphenols and human gut microbiota: A review. Food Review International. 2011;27:154-169. DOI: 10.1080/87559129.2010.535233
  145. 145. Cardona F, Andrés-Lacueva C, Tulipani S, Tinahones FJ, Queipo-Ortuño MI. Benefits of polyphenols on gut microbiota and implications in human health. The Journal of Nutritional Biochemistry. 2013;24:1415-1422. DOI: 10.1016/j.jnutbio.2013.05.001
  146. 146. Cueva C, Sánchez-Patán F, Monagas M, Walton GE, Gibson GR, Martín-Álvarez PJ, et al. In vitro fermentation of grape seed flavan-3-ol fractions by human faecal microbiota: Changes in microbial groups and phenolic metabolites. FEMS Microbiology Ecology. 2013;83:792-805. DOI: 10.1111/1574-6941.12037
  147. 147. Anhê FF, Pilon G, Roy D, Desjardins Y, Levy E, Marette A. Triggering Akkermansia with dietary polyphenols: A new weapon to combat the metabolic syndrome? Gut Microbes. 2016;7:146-153. DOI: 10.1080/19490976.2016.1142036
  148. 148. Xing Y-W, Lei G-T, Wu Q-H, Jiang Y, Huang M-X. Procyanidin B2 protects against diet-induced obesity and non-alcoholic fatty liver disease via the modulation of the gut microbiota in rabbits. World Journal of Gastroenterology. 2019;25:955-966. DOI: 10.3748/wjg.v25.i8.955
  149. 149. Tao W, Zhang Y, Shen X, Cao Y, Shi J, Ye X, et al. Rethinking the mechanism of the health benefits of proanthocyanidins: Absorption, metabolism, and interaction with gut microbiota. Comprehensive Reviews in Food Science and Food Safety. 2019;18:971-985. DOI: 10.1111/1541-4337.12444
  150. 150. Casanova-Martí À, Serrano J, Portune KJ, Sanz Y, Blay MT, Terra X, et al. Grape seed proanthocyanidins influence gut microbiota and enteroendocrine secretions in female rats. Food & Function. 2018;9:1672-1682. DOI: 10.1039/c7fo02028g
  151. 151. Everard A, Belzer C, Geurts L, Ouwerkerk JP, Druart C, Bindels LB, et al. Cross-talk betweenAkkermansia muciniphilaand intestinal epithelium controls diet-induced obesity. Proceedings of the National Academy of Sciences of the United States of America. 2013;110:9066-9071. DOI: 10.1073/pnas.1219451110
  152. 152. Gotteland M, Andrews M, Toledo M, Muñoz L, Caceres P, Anziani A, et al. Modulation ofHelicobacter pyloricolonization with cranberry juice andLactobacillus johnsoniiLa1 in children. Nutrition. 2008;24:421-426. DOI: 10.1016/j.nut.2008.01.007
  153. 153. Pastene E, Parada V, Avello M, Ruiz A, García A. Catechin-based procyanidins fromPeumus boldusMol. aqueous extract inhibitHelicobacter pyloriurease and adherence to adenocarcinoma gastric cells. Phytotherapy Research. 2014;28:1637-1645. DOI: 10.1002/ptr.5176
  154. 154. Mena P, Bresciani L, Brindani N, Ludwig IA, Pereira-Caro G, Angelino D, et al. Phenyl-γ-valerolactones and phenylvaleric acids, the main colonic metabolites of flavan-3-ols: Synthesis, analysis, bioavailability, and bioactivity. Natural Product Reports. 2019;36:714-752

Written By

Carlos González-Quilen, Esther Rodríguez-Gallego, Raúl Beltrán-Debón, Montserrat Pinent, Anna Ardévol, Maria Teresa Blay and Ximena Terra

Submitted: November 22nd, 2019 Reviewed: January 15th, 2020 Published: February 25th, 2020