Next Article in Journal
Exogenous Brassinosteroid Facilitates Xylem Development in Pinus massoniana Seedlings
Next Article in Special Issue
The Role of Leaky Gut in Nonalcoholic Fatty Liver Disease: A Novel Therapeutic Target
Previous Article in Journal
Chromatin Profiling Techniques: Exploring the Chromatin Environment and Its Contributions to Complex Traits
Previous Article in Special Issue
Increase in Epithelial Permeability and Cell Metabolism by High Mobility Group Box 1, Inflammatory Cytokines and TPEN in Caco-2 Cells as a Novel Model of Inflammatory Bowel Disease
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
IJMS
Volume 22
Issue 14
10.3390/ijms22147613
ijms-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

Leaky Gut: Effect of Dietary Fiber and Fats on Microbiome and Intestinal Barrier

by
Haruki Usuda
*,
Takayuki Okamoto
* and
Koichiro Wada
Department of Pharmacology, Faculty of Medicine, Shimane University, 89-1 Enyachō, Izumo 693-8501, Japan
*
Authors to whom correspondence should be addressed.
Int. J. Mol. Sci. 2021, 22(14), 7613; https://doi.org/10.3390/ijms22147613
Submission received: 7 May 2021 / Revised: 10 July 2021 / Accepted: 13 July 2021 / Published: 16 July 2021
(This article belongs to the Special Issue Elucidation of Leaky Gut Syndrome and the Discovery of Therapeutic Targets)
Browse Figures
Versions Notes

Abstract

:
Intestinal tract is the boundary that prevents harmful molecules from invading into the mucosal tissue, followed by systemic circulation. Intestinal permeability is an index for intestinal barrier integrity. Intestinal permeability has been shown to increase in various diseases—not only intestinal inflammatory diseases, but also systemic diseases, including diabetes, chronic kidney dysfunction, cancer, and cardiovascular diseases. Chronic increase of intestinal permeability is termed ‘leaky gut’ which is observed in the patients and animal models of these diseases. This state often correlates with the disease state. In addition, recent studies have revealed that gut microbiota affects intestinal and systemic heath conditions via their metabolite, especially short-chain fatty acids and lipopolysaccharides, which can trigger leaky gut. The etiology of leaky gut is still unknown; however, recent studies have uncovered exogenous factors that can modulate intestinal permeability. Nutrients are closely related to intestinal health and permeability that are actively investigated as a hot topic of scientific research. Here, we will review the effect of nutrients on intestinal permeability and microbiome for a better understanding of leaky gut and a possible mechanism of increase in intestinal permeability.

1. Introduction

Leaky gut refers to the dysfunction of the intestinal barrier and often leads to the generation of leaky gut syndrome (LGS) under chronic states. As the name indicates, the pathological manifestation of leaky gut is increased intestinal permeability, which is induced by various causative factors. The term ‘intestinal permeability’ is simple; however, the definition of intestinal permeability is ambiguous, since it potentially has two meanings—epithelium permeability and permeability of capillary vessels in villi. It is difficult to evaluate the latter in vivo. So, the term ‘leaky gut’ is used to indicate abnormal translocation of big-size molecules from lumen to villi or excessive absorption of such molecules from lumen into systemic circulation, which, in turn, induces various organ disorders. Historically, the concept of leaky gut began to emerge as epithelium permeability. Classical LGS has been observed concomitantly with gut inflammation, including inflammatory bowel diseases (IBD) or coeliac disease, and NSAIDs-induced ulceration that was studied from 1970 to 1990 extensively [1,2,3]. Hence, leaky gut was recognized as a typical manifestation of gut inflammation during this period and ceased to be focused on. Leaky gut has again come under the scientific spotlight due to the attention gained by the gut microbiome. Genome-sequencing methods have dramatically evolved within the last two decades, which now enables analysis of the whole intestinal bacterial genome, or identification of bacteria in stool sample by sequencing conserved marker genes such as the 16srRNA gene of bacteria [4]. Now we know the entire genetic content of human gut microbiota, the amount of which is calculated to be 100-fold or higher compared to human genomic content [5]. These findings have led to the emergence of the concept of intestinal enterotypes. Two major phyla, Bacteroidetes and Firmicutes, dominate among the microbiota of humans, and many studies have shown the considerable inter- and intrapersonal variability at the genus level and above [6]. Firmicutes phylum contains various types of bacteria including facultative, anaerobic, cocci, and bacilli bacteria. Gram-positive bacteria are major in this phylum and have relatively low guanine and cytosine contents [7,8]. On the contrary to Firmicutes, gram-negative bacteria are dominant in Bacteroidetes phylum and human gut have Bacteroides, Alistipes, Parabacteroides, and Prevotella genus primarily [9]. Bacteria interacts with a host directly or indirectly via physiological active molecules secreted from bacteria including short-chain fatty acids (SCFAs), p-cresol, p-cresyl-glucuronide (pCG), indoxyl sulphate (IS), indole-3 acetic acid (IAA), and H2S and trimethylamine N-oxide (TMAO) that may result in protective or harmful effects on the intestinal barrier and various organs distant from the intestine including the liver, kidney, and brain [10,11]. In fact, dysbiosis, an unhealthy alternation of gut microbiota, correlates with several types of diseases, including IBD [12], cancer [13], neuropsychiatric disorders [14], chronic kidney diseases (CKD) [15,16], and cardiometabolic diseases including obesity, type 2 diabetes (T2D), and cardiovascular disease [17], along with increased intestinal permeability. Dysbiosis results in an increased population of pathogenic bacteria which are likely to produce higher levels of lipopolysaccharides (LPS) and induce damage to epithelial cells which is one of the possible mechanisms of increased intestinal permeability observed with dysbiosis [18,19,20]. Furthermore, disruptions of intercellular connections let harmful molecules, including LPS, invade the intestinal tissue, and consequently access the blood stream, which provokes or worsens not only IBD [21] but also systemic diseases [18,19,20]. Hence, the concept of LGS shifted from a mere inflammatory phenotype to an exacerbation factor of systemic diseases, although its etiology is still unclear. In addition, bidirectional translocation of molecules, metabolites, and toxins derived from systemic circulation to gut lumen also can be recognized as LGS although the evidence is poor.
Dietary components come into contact with intestinal lumen for a long time and are likely to regulate gut microbiota and intestinal permeability. However, many studies which are now focusing on gut microbiota recognize alteration of intestinal permeability as an ancillary symptom. In this review, we provide an overview of recent studies that investigate the effect of dietary components on intestinal permeability and microbiome and hypothesize a possible correlation among them (Figure 1). Especially, the effects of dietary fiber and a high fat diet (HFD) on microbiome and intestinal permeability have been attracting attention globally as targets of research and are investigated most actively. Hence, we will focus on these nutrients.

2. Intestinal Barrier

Luminal components including unstirred water, glycocalyx, and mucus as well as antibacterial molecules including defensins, lysosome, and IgA provide the first line of defense before harmful bacteria come in contact with the epithelium. In addition, microclimate and secretions from the stomach, pancreas, and pancreatic acids destroy bacteria and antigens in the lumen [22].
Mucus layers and epithelium are the most important and major structures of intestinal barrier. A mucus layer exists on the outer surface of the large intestine. This layer is composed of two sub-layers. The outer layer is thick and loose, where bacteria and bacteria-derived molecules are abundant. Many species of commensal bacteria grow and form colonies on the outer layer, such that under healthy conditions, pathogenic bacteria cannot overgrow or invade this territory of commensal bacteria. The inner layer is firm, adherent, and harbors quite small numbers of bacteria. This layer acts as a boundary between bacteria and epithelium [23]. In the case of the small intestine, the mucus layer is single and fluid and contains abundant antimicrobial substances. Studies using experimental animals have shown that the disruption of mucus production can lead to intestinal damage and inflammation [24].
The epithelium is located just below inner layers and is composed of normal epithelial cells and several types of cells possessing specific functions, including Paneth cells, goblet cells, etc. Paneth cells secrete antibacterial peptides such as lysozyme and defensins and prevent colonization of harmful bacteria [25], while enterocytes produce chloride in response to noxious stimuli [26]. Goblet cells contribute to maintaining a mucosal layer by secreting mucin [27]. Epithelial cells are linked by the apical junctional complex which consists of the tight junction (TJ) and adherence junction. TJ is further composed of claudin, occludin, and junctional adhesion protein molecule-A (JAM-A) as well as intracellular plaque proteins such as zonula occludens (ZOs) and cingulin [28] (Figure 2). Twenty-seven human claudin genes have been identified, though the protein expression of some of them is not confirmed [29]. Of note, claudin-13 is expressed in rodents but is absent in humans [30]. Claudin and occludin function cooperatively. Nevertheless, embryonic stem cells that lack occludin differentiate into polarized epithelia with functional TJ [31]. In addition, occludin knockout leads to the normal barrier function of intestinal epithelium [32]. Thus, occluding plays a supportive role in intestinal permeability.
Molecules can pass through the epithelium passively via transcellular route [26,33] or paracellular route [26,34] (Figure 3). Soluble lipids, small hydrophilic compounds, ions, and water molecules pass through the transcellular route. The paracellular route allows the passage of bigger molecules, although the size is limited at up to 600 Da in in vivo and 10 kDa in in vitro [35,36] via apical junctional complex. The latter route is further divided into two types of pathways and regulated by IL-13 and tumor necrosis factor (TNF). IL-13 specifically increases flux across the small molecules, including ions and water, through the pore pathway [37]. IL-13 causes barrier loss by inducing claudin-2 expression as well as by increasing apoptosis and inhibiting wound healing both in vitro and in vivo, whereas lower doses of IL-13 induce claudin-2 upregulation and claudin-2-dependent pore-pathway activation in response to IL-13 exposure without increase in leaks or unrestricted pathway flux [37,38]. TNF opens the leak pathway via myosin light chain kinase (MLCK) and lets bigger molecules pass through the intracellular space [37]. Hence, the leak pathway is likely to be associated with inflammation, which would permit the passage of macromolecules, bacterial products, and food antigens. TNFα has also been shown to regulate TJ and the clinically relevant role of TNF in IBD pathogenesis is clearly demonstrated by the efficacy of anti-TNF antibodies in IBD, which reduces disease severity and restores intestinal barrier function [39]. Restoration of epithelial barrier function by anti-TNF therapy may reflect mucosal healing in the setting of a dampened immune system; however, pre-clinical studies have shown that TNF signaling also modulates TJ. Many studies have shown decreased levels of intestinal TJ-consisting proteins in animal models of IBD [40], obesity, and T2D [41,42], with increased permeability, suggesting that a leaky pathway is responsible for LGS.
Epithelial cells bind each other with tight junction, adherens junction, and desmosome. Tight junction consists of claudin, occluding, and junctional adhesion protein molecule-A (JAM) recognized as a key construction which regulate the absorption of molecules via the paracellular route. These molecules, associated with zonula occluden-1 (ZO-1), which contribute to the formation of tight junction as a scaffold protein. Myosin-actin complex which is associated with myosin light-chain kinase (MLCK) is also interacting with ZO-1 in a steady state. Adherens junction is comprised by α-catenin, β-catenin, and E-cadherin. Desmosome consists of three kinds of molecules with binding to keratin.
Molecules are absorbed from intestinal lumen into the tissue via three pathways. Lipid soluble molecules, small hydrophilic molecules, ions, and water pass through the cell body. Ions and water are also absorbed intracellularly which is called the pore pathway. Bigger molecules (>600 Da) can be absorbed when the tight junction becomes loose or collapses by inflammation or other harmful stimuli. The modes of action for opening pore pathway by interleukin-13 (IL-13) and leak pathway by tumor necrosis factor (TNF-α) are relatively well known. Signaling from IL-13 receptor, activate casein kinase 2 and phosphorylate occluding successively which allows the interaction of claudin-2 and occluding linking with zonula occluden-1 (ZO-1) as shown in Figure 3. In the case of TNF-α, myosin light-chain kinase (MLCK) is activated and endocytosis of occluding is promoted which results in the collapse of tight junction.

3. How to Evaluate Intestinal Barrier Function

Oral administration of evaluating-reagents is a standard method and carried out most frequently in in vivo study. Especially, the administration of two non-metabolized sugars, lactulose and mannitol, is used as a golden standard method for human-targeted studies [43]. The absorption of lactulose is increased when the paracellular epithelial barrier is compromised whereas the smaller sized mannitol is constantly absorbed regardless of barrier function, which reflects the basal ability of intestinal absorption [43]. Hence, the value of urine lactulose/mannitol ratio is used as an index of disruption of the intestinal barrier [43]. However, these sugars can detect the status of barrier function in the small intestine only since these are degraded by bacteria [43]. Sucralose and 51Cr-ethylenediaminetetraacetic acid (EDTA) can evaluate whole intestinal permeability since these molecules are not metabolized by bacteria and used widely from in vitro studies to clinical studies [44].
The size of these sugars and 51Cr-(EDTA is small so that more big-sized molecules which are not non-metabolized by human enzymes and bacteria are also used in animal studies, in vitro and ex vivo, to evaluate absorption of molecules from the leak pathway. Ovalbumin (OVA) is frequently used to measure antigen uptake following oral gavage [43]. Polyethylene glycol and fluorescein isothiocyanate (FITC)-dextran are convenient to estimate the size of the molecule that leaks from the lumen since various molecular sizes of them are available. The molecular weight of these reagents which are most frequently used is around 4000 [45].
There are several other methods to evaluate permeability. Calculating the electrical resistance of the cell membrane, which reflects paracellular ion permeability, is specifically applicable to in vitro and ex vivo study [46]. In addition, various biomarkers relating to epithelial damage including citrulline, fatty acid-binding proteins (FABP) and LPS have been used as an indirect index of decreased intestinal barrier as shown elsewhere in detail [45].

4. Dietary Fiber

4.1. Dietary Fiber and Intestinal Barrier

4.1.1. Fermentation of Dietary Fiber and SCFA

Dietary fibers are largely metabolized by gut bacteria. This is because they can break down various types of carbohydrates since they encode over 260 glycoside hydrolases for the degradation of carbohydrates whereas a human has only 17 enzymes for digestion [47], meaning that human themselves are poor at digesting a variety of dietary fiber. In addition, more than 100 trillion bacteria can be engaged in consuming carbohydrates derived from dietary fiber [39] that would help effective digestion of dietary fiber. Hence, dietary fibers resistant to digestion by the host are termed as microbiota-accessible carbohydrates (MACs) [48]. Dietary fibers are divided into soluble and insoluble fibers. Insoluble fiber includes cellulose, some hemicellulose, and lignin [49]. Soluble fiber encompasses wheat dextrin, pectin, gums, β-glucan, psyllium, and fructans, as well as some hemicellulose [49]. These fibers are derived from grains, fruits, vegetables, and legumes [50]. Generally, insoluble fibers are poorly fermented by gut microbes, but they likely promote the gut transit rate and thus reduce the amount of time available for colonic bacterial fermentation of non-digested foodstuff [51]. Soluble fibers can be further processed by bacteria into SCFAs as metabolites [52] although some of them are not fermentable including psyllium and gums.
Different types of bacteria produce different types of SCFAs [53,54,55,56,57]. The most abundant SCFAs in the human colon are acetate, propionate, and butyrate, with a molar ratio of 60:20:20, approximately [58]. Colonocytes absorb SCFAs via transporters or simple diffusion. SCFA are transported to various organs via solute carrier family 16 member 1 (SLC16a1) and SLC5a8, transporters for SCFAs [59]. A majority of the acetate bypasses the splanchnic circulation to be converted into acetyl-CoA for lipogenesis or oxidation in peripheral muscles. The remaining acetate is converted to butyrate and is used by colonocytes [60,61]. Propionate primarily contributes to gluconeogenesis in liver. Some bacteria species intake lactate and succinate and convert them into propionates. These physiological functions are regulated by SCFAs via G-protein coupled receptors including GPR40, GPR41, GPR43, and GPR120, which are distributed across various types of cells. GPR41 and GPR43 are highly expressed in the intestine [62]. It has been reported that, in humans, the fecal SCFA level increases after intaking an MACs-rich diet [63].

4.1.2. Contribution of SCFA to Intestinal Barrier

Butyrate is a well-documented beneficial factor for maintaining colonocyte health by providing energy to intestinal epithelial cells, which likely contributes to intestinal epithelial integrity [64]. Butyrate suppresses cytokine-induced barrier dysfunction by modifying claudin-2 levels in vitro [65]. Animal studies have also shown similar protective effects on not only the intestinal barrier [66,67] but also the airway epithelial barrier [68]. Fecal calprotectin, a marker of intestinal inflammation, is diminished by administration of butyrate to UC patients [69]. In addition to these effects, butyrate seems to maintain intestinal integrity by inducing colonic mucin expression as shown in an in vitro study [70]. Besides, animal studies show that acetate directly activates nucleotide-binding oligomerization domain 3 (NLRP3) inflammasome in gut epithelial cells which results in the release of IL-18 [71], which, in turn, promotes intestinal barrier integrity via activation of IL-18 receptor on the mice epithelial cells [72]. However, genetical knocking out of IL-18 in the whole body or blocking of IL-18 receptor enhances the sensitivity to chemicals which induces colitis in mice [73,74], suggesting that acetate contributes to maintaining the epithelial barrier via NLRP3 signaling, followed by IL-18 production. Propionate also diminishes hyper intestinal permeability induced by dextran sulfate sodium (DSS) via amelioration of downregulation of ZO-1, occludin, and E-cadherin expressions in the colonic tissue in mice [75]. Recently, acetate, propionate, and butyrate have been shown to synergistically promote intracellular permeability by modifying TJ expression or distribution, including ZO-1 [76] in vitro. Together with these findings, SFCAs are recognized as a key factor for maintaining the intestinal barrier (Figure 4).

4.1.3. Contribution of Dietary Fiber to Intestinal Barrier Integrity

The impact of MACs on intestinal permeability has been investigated through two kinds of approaches using a fiber-deprived or high fiber diet. Mice fed on an MACs-deprived diet exhibited severe colitis and increased intestinal permeability induced by DSS along with less serum IL-18 levels in mice [71]. A recent animal study targeting mice demonstrated that the removal of MACs affects the secretions of glucagon-like peptide-1 (GLP-1) and GLP-2, which synergistically ameliorate intestinal injury and improve intestinal healing [77,78]: they are downregulated in ileum and colon, along with increased intestinal permeability [79]. These results indicate that MACs contribute to the intestinal barrier via regulating the intestinal immune system and regulation of secretion of peptide hormones from colonocytes (Figure 4). Besides, the supplementation of fructo-oligosaccharides that are produced by degradation of inulin, a representative water-soluble dietary fiber, induces IgA production in rat cecum and suppresses a MACs-deficient diet-induced decrease in intestinal permeability [80]. In addition, the severity of DSS-induced colitis is diminished after administration of an MACs-containing diet, along with a marked increase in fecal butyrate [81]. These results also suggest that MACs have beneficial effects on the intestinal barrier. Nevertheless, whether MAC supplementation is really recommended to maintain intestinal health including barrier function is not confirmed yet since some clinical studies show beneficial effects of fructo-oligosaccharides and polydextrose on intestinal barrier function in healthy subjects or patients of pancreatitis [82,83], but administration of oat β-glucan, arabinoxylan (soluble hemicellulose) do not show significant effects on acute indomethacin-induced intestinal hyperpermeability [84,85]. Oligofructose-enriched inulin also does not improve intestinal barrier function effective in the patients of celiac disease [84,85].

4.2. MACs-Induced Alteration of Microbiota and Intestinal Barrier Integrity

Several studies have shown alteration of microbiota after MAC administration. The fecal microbiota composition of HFD-fed mice is modified by inulin administration in a dose-dependent manner. In this study, populations of Roseburia, Clostridium I, IV, and XIV spp. decreased, whereas the levels of Bifidobacterium spp. and Bacteroidetes increased [86], along with decreased caloric intake. Another study targeting humans with mild constipation also demonstrated that inulin induces an increase in the populations of Anaerostipes, Bilophila, and Bifidobacterium genus [87]. Especially, Bilophila spp. have been reported to be associated with softer stool and a favorable change in constipation-specific quality-of-life measures [87]. In addition, T2D is associated with reduced abundance of fiber-degrading bacteria in humans [88,89]. On the other hand, prolonged low-fiber feeding aggravated allergic airway disease in mice, which could be corrected by administration of the SCFA propionate [90], implying alternation of microbiota. These results suggest that MACs exhibit a favorable effect on microbiota and intestinal health that would affect the state of various systemic diseases, although the precise mechanism remains unknown. Besides, an MAC-deprived diet leads to an increase in levels of Bacteroides thetaiotaomicron (B. thetaiotaomicron), which intakes intestinal mucus glycans in mice [48]. Of note, microbiota transplant to germ-free mice showed that the population of mucin-degrading bacteria including B. thetaiotaomicron and Akkermansia muciniphila (A. muciniphila) increased in the intestine under deficiency of dietary MACs [91,92]. Interestingly, animal study shows that A. muciniphila is a promising probiotic bacteria despite its mucin-degrading feature [93]. A. muciniphila accounts for 1–4% of human gut microbiota starting from early life [94]. Decreased population of A. muciniphila has been reported in patients with IBD [95,96]. Conversely, the population of intestinal A. muciniphila increased in accordance with DSS-induced colitis in mice [97,98]. However, intervention studies indicate protective effects of A. muciniphila on intestinal integrity or inflammatory states in an animal colitis model. Administration of live A. muciniphila ameliorates DSS-induced colitis along with suppression of increase in intestinal permeability in mice [99]. Another animal study demonstrated that spleen weight, colon inflammation index, and colon histological score as well as the expression of the pro-inflammatory cytokines including TNF-α and IFN-γ in the colon are decreased by the administration of A. muciniphila [100]. Furthermore, outer membrane protein derived from A. muciniphila also shows similar protective effects in a mice colitis model [101]. This protein has been shown to enhance intracellular permeability, activate signaling pathway through Toll-like receptor 2 (TLR2) and TLR4, and modulate cytokine production from peripheral blood mononuclear cells in vitro [102]. The mode of action of A. muciniphila in serving these effects is still unclear. Further investigation might disclose this mechanism in the future.

5. HFD and Bile Acids

5.1. HFD and Intestinal Barrier

5.1.1. Effects of Fatty Acid and HFD on Intestinal Barrier

Dietary fats-derived fatty acids are broadly categorized into saturated or unsaturated fatty acids. In parallel, there is another classification based on the length of fatty acids: SFCAs, middle-chain fatty acids (MCFAs), and long-chain fatty acid (LCFAs). They have the ability to affect paracellular permeability as demonstrated in previous in vitro studies [103,104]. Eicosapentaenoic acid (EPA), docosahexaenoic acid (DHA), and γ-linolenic acid, which belong to unsaturated LCFAs increase TJ permeability under normal physiological state without any disruptive stimuli to the intestinal barrier in Caco-2 cells [103,104]. However, these LCFAs decrease the TJ permeability in T84 cells, a cell line similar to Caco-2 cells. In addition, EPA and DHA reduce IL-4-mediated increase in paracellular permeability in T84 cells [105]. Both capric acid and lauric acid, known as saturated MCFAs, enhance the increase in paracellular permeability through activation of MLCK in Caco-2 cells [106]. Additionally, capric acid induces conformational alteration of TJ proteins including occludin and ZO-1 [107], whereas lauric acid does not [106]. These results indicate that dietary fats can alter intestinal permeability through the TJ-associated paracellular route by interacting with epithelial cells directly (Figure 5).
Etiological studies show the association of LCFAs with IBD progression, which implies that LCFAs alter the intestinal barrier function in humans. A large prospective cohort study demonstrated a relationship between higher consumption of DHA and a decrease in the incidence of ulcerative colitis (UC) [108]. Association of n-6 and n-3 polyunsaturated fatty acids (PUFAs) with IBD state has also been reported. Generally, n-6 PUFAs are suggested to trigger or enhance inflammatory signaling pathways, whereas n-3 PUFAs exhibit anti-inflammatory effects [109,110,111]. Higher intake of n-3 PUFA is associated with an increase in n-3/n-6 ratio in the erythrocyte membrane of the IBD patients who are in a remission state compared to those who had relapsed [112]. Serum n-3 fatty acids and EPA levels have been reported to be positively correlated with pro-inflammatory cytokine levels and disease activity, whereas serum n-6 fatty acids are inversely correlated with these indexes [113].
HFD can induce an increase in intestinal permeability in mice or rats with decreased mRNA or protein expression of TJ including claudins-1, claudins-2, claudins-3, and ZO-1 [114,115,116,117]. More specifically, in IL-10 knockout mice with IBD, IBD-like colitis is spontaneously triggered with an increase in intestinal permeability [118]. A high saturated fat diet was observed to promote Th1 immune response and increase the incidence of colitis [119]. These effects possibly arise due to an increased population of Bilophila wadsworthi, a sulfite-reducing pathobiont, through taurine-conjugation of hepatic bile acids. Mice fed a high-fat and high-sugar combination diet showed increased fecal inflammatory markers with increased levels of proteobacteria in the stool [120]. In accordance with these changes, these mice are more susceptible to DSS-induced colitis. HFD also affects IgA secretion. The level of secretory IgA coating the gut microbiota is elevated both in normal diet and HFD fed mice. However, this increase is diminished in HFD fed mice [121]. Hence, HFD diet induces an increase in intestinal permeability via downregulation of TJ and alters the immune response mediated by T cells and IgA-producing plasma cells in vivo.

5.1.2. HFD-Induced LPS Absorption and Intestinal Barrier

Increased LPS absorption from leaky gut has been suggested by several studies on HFD-fed mice showing increased permeability with elevated serum LPS levels [114,122]. On the other hand, mice injected with LPS exhibited a reduction in plasma HDL cholesterol and elevations in plasma triglycerides [123]. A human cohort study also demonstrated that high serum LPS level in type 1 diabetic patients is associated with high serum triglyceride levels and increased diastolic blood pressure [124]. These results suggest that leaky gut affects dyslipidemia via absorption of LPS (Figure 5). In fact, mouse models have shown the importance of TLR4, a receptor for LPS, and its signaling in diet-induced insulin resistance and atherosclerosis [125]. TLR4 knockout mice are resistant to HFD-induced glucose intolerance [126,127] as well as HFD-induced atherosclerosis [128]. Similarly, human studies have shown that high fat and high carbohydrate intake modulates TLR2 and TLR4 expression in mononuclear cells with increased LPS level in the serum [129,130]. Besides, LPS stimulates T helper cell (Th) 17 differentiation though TLR4 in response to LPS [131] that may contribute to an LPS-induced inflammatory reaction, since IL-17A decreases the expression of peroxisome proliferator-activated receptor-α, which is anti-inflammatory and inhibits fatty-acid oxidation [132]. An advanced study has shown that leukocyte infiltration to the liver and mRNA expression of inflammatory cytokines TLR4 and TLR9 were increased in HFD-fed mice, in which the intestinal barrier was disrupted by co-administration of DSS with HFD, which downregulated ZO-1 and Claudin-1 expression in the colon, suggesting that leaky gut and the following attack to liver by LPS are involved in pathogenesis of liver dysfunction [133].

5.2. Alteration of Microbiome by HFD and Intestinal Permeability

Several studies have shown alteration of the microbiome by HFD both in animals [134,135,136] and humans [137,138]. A recent study demonstrated the possibility that HFD-induced alteration of the microbiome indirectly modulates the expression of TJ. Constant light exposure with HFD induces glucose abnormalities, insulin resistance, inflammation, and liver steatohepatitis in mice, which is associated with less abundance of Butyricicoccus, Clostridium, and Turicibacter, also known as butyrate producers. These changes correlate with decreased butyrate levels in colon contests, decreased colon expressions of occludin-1 and ZO-1, and increased serum LPS and mRNA expression of liver LPS-binding protein [139]. As described earlier, A. muciniphila exhibits protective effects on intestinal permeability. The effect of these bacteria on intestinal barrier dysfunction is induced by HFD and associated systemic disorders, including obesity and T2D. An abundance of A. muciniphila is negatively correlated with fasting blood glucose, waist-to-hip ratio, and subcutaneous fat cell diameter in obese subjects and an increased abundance of A. muciniphila positively correlates with improvement in insulin sensitivity markers and other clinical parameters after calorie restriction [140]. Compared to healthy subjects, T2D patients harbor less fecal A. muciniphila extracellular vehicles (AmEVs) [141], which suppresses HFD-induced weight gain, increased plasma cholesterol, triglycerides, and glucose, along with downregulation of intestinal TLR4 and TJ mRNAs in mice [142]. In addition, the membrane protein derived from A. muciniphila reduces intolerance to glucose and plasma LPS levels along with upregulation of the molecules related to insulin signaling and claudin-3 [143]. These results imply that A. muciniphila exhibits a beneficial effect on HFD-induced disorders and intestinal barrier integrity might be involved in this mechanism (Figure 5).

5.3. Bile Acids and Intestinal Permeability

Bile acids (BAs) secreted from duodenum help in digestion of lipids as well as cholesterol and fat-soluble vitamins. HFD induce enhanced BA discharge, resulting in increased colonic concentrations of primary BAs, compared with low or normal fat diets [144]. However, 5% to 10% of BAs are not reabsorbed but are converted to secondary bile acids by bacteria in the large intestine. Secondary bile acids are harmful and are suggested to promote colon carcinogenesis [145]. In addition to their role in dietary lipid absorption and cholesterol homeostasis, bile acids act as signaling molecules via two major signaling pathways. One is G protein-coupled bile acid receptor (GPBAR1, also known as TGR5) and another comprises members of the nuclear hormone receptor superfamily including the farnesoid X receptor (FXR) [146]. Animal studies indicate that both GPBAR1 and FXR contribute to the integrity of the intestinal barrier. GPBAR1 knock out mice show abnormal molecular architecture of epithelial TJ with increased expression and abnormal subcellular distribution of zonulin-1 with increased intestinal permeability. In addition, these mice are more sensitive to DSS stimuli and exhibit severe colitis [147]. Activation of GPBAR1 with synthetic agonist reversed intestinal inflammation in chemically induced colitis models. The possible mechanisms include reduction in the trafficking of monocytes from blood to intestinal mucosa and modulation of the activation state of macrophages which results in decreased expression of inflammatory genes, including TNF-a, IFN-g, IL-1b, IL-6, and CCL2 [148]. Intestinal permeability is increased in FXR knockout mice [149] and FXR-agonist exhibits a protective effect on chemically induced colitis with reduced epithelial permeability and several inflammatory responses [150]. A recent study demonstrated that gut-specific FXR-deficient mice exhibit increased intestinal permeability, possibly due to reduced mucosal integrity, associated with decreased secretion of mucin 2 protein and lower levels of E-cadherin protein [151]. Taken together, these results indicate that BAs maintain the intestinal barrier through the BA sensing receptor at least partially (Figure 5).

6. Concluding Remarks

We reviewed the effect of dietary fiber and fats on intestinal permeability and microbiome, and the correlations between them. Dietary fiber is recognized as a protective nutrient for the intestinal barrier and contributes to maintaining the microbiota in a healthy state. These beneficial effects are mediated by SCFAs, and epithelial IL-18 seems to be involved in a molecular mechanism of barrier-regulation. In contrast, fats likely promote an increase in epithelial permeability. In vitro studies show that fatty acids directly impair the epithelial barrier. Besides, fat-induced increased plasma LPS levels are able to disrupt the intestinal barrier. These findings indicate that a westernized diet contains high fats and poor fibers likely to induce or exacerbate various systemic diseases due to bad intestinal health including impaired intestinal barrier function. In addition, both dietary fiber and high fat can affect the relative population of A. muciniphila which imparts a protective effect on intestinal barrier integrity and disease state, suggesting possible causative relationships between dietary fibers and fats, microbiome, LGS, and systemic diseases. Nevertheless, these relationships have a missing link since the molecules which are responsible for LGS and the successive dysfunction of various organs are not identified. To clarify this point, we should identify the precise region of LGS which conventional LGS-evaluating reagents cannot do [45]. This is the bottleneck to evaluate whether the subjects are LGS or not in animals and clinical studies. That is why leaky gut is not something that can be concluded as easily so far, especially in humans. Improvement of LGS-evaluating methods or development of an ex vivo organ like an organoid would connect the missing link and provide more information about the relationship between nutrients and LGS.

Author Contributions

Conceptualization, H.U.; writing—original draft preparation, H.U.; writing—review and editing, H.U., T.O. and K.W. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Chadwick, V.S.; Phillips, S.F.; Hofmann, A.F. Measurements of intestinal permeability using low molecular weight polyethylene glycols (PEG 400). II. Application to normal and abnormal permeability states in man and animals. Gastroenterology 1977, 73, 247–251. [Google Scholar] [CrossRef]
  2. Bjarnason, I.; Peters, T.J.; Veall, N. A persistent defect in intestinal permeability in coeliac disease demonstrated by a 51Cr-labelled EDTA absorption test. Lancet 1983, 1, 323–325. [Google Scholar] [CrossRef]
  3. Hollander, D.; Vadheim, C.M.; Brettholz, E.; Petersen, G.M.; Delahunty, T.; Rotter, J.I. Increased intestinal permeability in patients with Crohn’s disease and their relatives. A possible etiologic factor. Ann. Intern. Med. 1986, 105, 883–885. [Google Scholar] [CrossRef]
  4. Oulas, A.; Pavloudi, C.; Polymenakou, P.; Pavlopoulos, G.A.; Papanikolaou, N.; Kotoulas, G.; Arvanitidis, C.; Iliopoulos, I. Metagenomics: Tools and insights for analyzing next-generation sequencing data derived from biodiversity studies. Bioinform. Biol. Insights 2015, 9, 75–88. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  5. Bäckhed, F.; Ley, R.E.; Sonnenburg, J.L.; Peterson, D.A.; Gordon, J.I. Host-bacterial mutualism in the human intestine. Science 2005, 307, 1915–1920. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  6. Hillman, E.T.; Lu, H.; Yao, T.; Nakatsu, C.H. Microbial Ecology along the Gastrointestinal Tract. Microbes Environ. 2017, 32, 300–313. [Google Scholar] [CrossRef] [Green Version]
  7. Ciccarelli, F.D.; Doerks, T.; von Mering, C.; Creevey, C.J.; Snel, B.; Bork, P. Toward automatic reconstruction of a highly resolved tree of life. Science 2006, 311, 1283–1287. [Google Scholar] [CrossRef] [Green Version]
  8. Tocheva, E.I.; Matson, E.G.; Morris, D.M.; Moussavi, F.; Leadbetter, J.R.; Jensen, G.J. Peptidoglycan remodeling and conversion of an inner membrane into an outer membrane during sporulation. Cell 2011, 146, 799–812. [Google Scholar] [CrossRef] [Green Version]
  9. Human Microbiome Project Consortium. Structure, function and diversity of the healthy human microbiome. Nature 2012, 486, 207–214. [Google Scholar] [CrossRef] [Green Version]
  10. Guarner, F.; Malagelada, J.R. Gut flora in health and disease. Lancet 2003, 361, 512–519. [Google Scholar] [CrossRef]
  11. Stecher, B.; Hardt, W.D. The role of microbiota in infectious disease. Trends Microbiol. 2008, 16, 107–114. [Google Scholar] [CrossRef]
  12. Hold, G.L.; Smith, M.; Grange, C.; Watt, E.R.; El-Omar, E.M.; Mukhopadhya, I. Role of the gut microbiota in inflammatory bowel disease pathogenesis: What have we learnt in the past 10 years? World J. Gastroenterol. 2014, 20, 1192–1210. [Google Scholar] [CrossRef]
  13. Tilg, H.; Adolph, T.E.; Gerner, R.R.; Moschen, A.R. The Intestinal Microbiota in Colorectal Cancer. Cancer Cell 2018, 33, 954–964. [Google Scholar] [CrossRef] [Green Version]
  14. Grochowska, M.; Wojnar, M.; Radkowski, M. The gut microbiota in neuropsychiatric disorders. Acta Neurobiol. Exp. Wars 2018, 78, 69–81. [Google Scholar] [CrossRef] [Green Version]
  15. Castillo-Rodriguez, E.; Fernandez-Prado, R.; Esteras, R.; Perez-Gomez, M.V.; Gracia-Iguacel, C.; Fernandez-Fernandez, B.; Kanbay, M.; Tejedor, A.; Lazaro, A.; Ruiz-Ortega, M.; et al. Impact of Altered Intestinal Microbiota on Chronic Kidney Disease Progression. Toxins 2018, 10, 300. [Google Scholar] [CrossRef] [Green Version]
  16. Glorieux, G.; Gryp, T.; Perna, A. Gut-Derived Metabolites and Their Role in Immune Dysfunction in Chronic Kidney Disease. Toxins 2020, 12, 245. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  17. Hansen, T.H.; Gøbel, R.J.; Hansen, T.; Pedersen, O. The gut microbiome in cardio-metabolic health. Genome Med. 2015, 7, 33. [Google Scholar] [CrossRef] [Green Version]
  18. Salguero, M.V.; Al-Obaide, M.A.I.; Singh, R.; Siepmann, T.; Vasylyeva, T.L. Dysbiosis of Gram-negative gut microbiota and the associated serum lipopolysaccharide exacerbates inflammation in type 2 diabetic patients with chronic kidney disease. Exp. Ther. Med. 2019, 18, 3461–3469. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  19. Xi, Y.; Yan, J.; Li, M.; Ying, S.; Shi, Z. Gut microbiota dysbiosis increases the risk of visceral gout in goslings through translocation of gut-derived lipopolysaccharide. Poult. Sci. 2019, 98, 5361–5373. [Google Scholar] [CrossRef]
  20. Belančić, A. Gut microbiome dysbiosis and endotoxemia—Additional pathophysiological explanation for increased COVID-19 severity in obesity. Obes. Med. 2020, 20, 100302. [Google Scholar] [CrossRef] [PubMed]
  21. Pastor Rojo, O.; López San Román, A.; Albéniz Arbizu, E.; de la Hera Martínez, A.; Ripoll Sevillano, E.; Albillos Martínez, A. Serum lipopolysaccharide-binding protein in endotoxemic patients with inflammatory bowel disease. Inflamm. Bowel Dis. 2007, 13, 269–277. [Google Scholar] [CrossRef]
  22. Keita, A.V.; Söderholm, J.D. The intestinal barrier and its regulation by neuroimmune factors. Neurogastroenterol. Motil. 2010, 22, 718–733. [Google Scholar] [CrossRef]
  23. Quigley, E.M. Leaky gut—Concept or clinical entity? Curr. Opin. Gastroenterol. 2016, 32, 74–79. [Google Scholar] [CrossRef]
  24. Van der Sluis, M.; De Koning, B.A.; De Bruijn, A.C.; Velcich, A.; Meijerink, J.P.; Van Goudoever, J.B.; Büller, H.A.; Dekker, J.; Van Seuningen, I.; Renes, I.B.; et al. Muc2-deficient mice spontaneously develop colitis, indicating that MUC2 is critical for colonic protection. Gastroenterology 2006, 131, 117–129. [Google Scholar] [CrossRef]
  25. Wang, S.L.; Shao, B.Z.; Zhao, S.B.; Fang, J.; Gu, L.; Miao, C.Y.; Li, Z.S.; Bai, Y. Impact of Paneth Cell Autophagy on Inflammatory Bowel Disease. Front. Immunol. 2018, 9, 693. [Google Scholar] [CrossRef] [Green Version]
  26. Schoultz, I.; Keita, Å.V. The Intestinal Barrier and Current Techniques for the Assessment of Gut Permeability. Cells 2020, 9, 1909. [Google Scholar] [CrossRef] [PubMed]
  27. Camilleri, M. Leaky gut: Mechanisms, measurement and clinical implications in humans. Gut 2019, 68, 1516–1526. [Google Scholar] [CrossRef] [PubMed]
  28. Buckley, A.; Turner, J.R. Cell Biology of Tight Junction Barrier Regulation and Mucosal Disease. Cold Spring Harb. Perspect. Biol. 2018, 10, a029314. [Google Scholar] [CrossRef] [PubMed]
  29. Liu, F.; Koval, M.; Ranganathan, S.; Fanayan, S.; Hancock, W.S.; Lundberg, E.K.; Beavis, R.C.; Lane, L.; Duek, P.; McQuade, L.; et al. Systems Proteomics View of the Endogenous Human Claudin Protein Family. J. Proteome Res. 2016, 15, 339–359. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  30. Holmes, J.L.; Van Itallie, C.M.; Rasmussen, J.E.; Anderson, J.M. Claudin profiling in the mouse during postnatal intestinal development and along the gastrointestinal tract reveals complex expression patterns. Gene Expr. Patterns 2006, 6, 581–588. [Google Scholar] [CrossRef] [PubMed]
  31. Saitou, M.; Fujimoto, K.; Doi, Y.; Itoh, M.; Fujimoto, T.; Furuse, M.; Takano, H.; Noda, T.; Tsukita, S. Occludin-deficient embryonic stem cells can differentiate into polarized epithelial cells bearing tight junctions. J. Cell Biol. 1998, 141, 397–408. [Google Scholar] [CrossRef]
  32. Saitou, M.; Furuse, M.; Sasaki, H.; Schulzke, J.D.; Fromm, M.; Takano, H.; Noda, T.; Tsukita, S. Complex phenotype of mice lacking occludin, a component of tight junction strands. Mol. Biol. Cell 2000, 11, 4131–4142. [Google Scholar] [CrossRef] [Green Version]
  33. Sandvig, K.; Kavaliauskiene, S.; Skotland, T. Clathrin-independent endocytosis: An increasing degree of complexity. Histochem. Cell Biol. 2018, 150, 107–118. [Google Scholar] [CrossRef] [Green Version]
  34. Hollander, D.; Kaunitz, J.D. The "Leaky Gut": Tight Junctions but Loose Associations? Dig. Dis. Sci. 2020, 65, 1277–1287. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Farquhar, M.G.; Palade, G.E. Junctional complexes in various epithelia. J. Cell Biol. 1963, 17, 375–412. [Google Scholar] [CrossRef] [Green Version]
  36. Fihn, B.M.; Sjöqvist, A.; Jodal, M. Permeability of the rat small intestinal epithelium along the villus-crypt axis: Effects of glucose transport. Gastroenterology 2000, 119, 1029–1036. [Google Scholar] [CrossRef] [PubMed]
  37. Weber, C.R.; Raleigh, D.R.; Su, L.; Shen, L.; Sullivan, E.A.; Wang, Y.; Turner, J.R. Epithelial myosin light chain kinase activation induces mucosal interleukin-13 expression to alter tight junction ion selectivity. J. Biol. Chem. 2010, 285, 12037–12046. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  38. Heller, F.; Florian, P.; Bojarski, C.; Richter, J.; Christ, M.; Hillenbrand, B.; Mankertz, J.; Gitter, A.H.; Bürgel, N.; Fromm, M.; et al. Interleukin-13 is the key effector Th2 cytokine in ulcerative colitis that affects epithelial tight junctions, apoptosis, and cell restitution. Gastroenterology 2005, 129, 550–564. [Google Scholar] [CrossRef] [PubMed]
  39. Suenaert, P.; Bulteel, V.; Lemmens, L.; Noman, M.; Geypens, B.; Van Assche, G.; Geboes, K.; Ceuppens, J.L.; Rutgeerts, P. Anti-tumor necrosis factor treatment restores the gut barrier in Crohn’s disease. Am. J. Gastroenterol. 2002, 97, 2000–2004. [Google Scholar] [CrossRef] [PubMed]
  40. Nighot, M.; Ganapathy, A.S.; Saha, K.; Suchanec, E.; Castillo, E.; Gregory, A.; Shapiro, S.; Ma, T.; Nighot, P. Matrix Metalloproteinase MMP-12 promotes macrophage transmigration across intestinal epithelial tight junctions and increases severity of experimental colitis. J. Crohns Colitis 2021, jjab064. [Google Scholar] [CrossRef] [PubMed]
  41. Zhang, B.; Yue, R.; Chen, Y.; Huang, X.; Yang, M.; Shui, J.; Peng, Y. The Herbal Medicine Scutellaria-Coptis Alleviates Intestinal Mucosal Barrier Damage in Diabetic Rats by Inhibiting Inflammation and Modulating the Gut Microbiota. Evid. Based Complement. Alternat. Med. 2020, 2020, 4568629. [Google Scholar] [CrossRef]
  42. Ahmad, R.; Rah, B.; Bastola, D.; Dhawan, P.; Singh, A.B. Obesity-induces Organ and Tissue Specific Tight Junction Restructuring and Barrier Deregulation by Claudin Switching. Sci. Rep. 2017, 7, 5125. [Google Scholar] [CrossRef] [Green Version]
  43. Menard, S.; Cerf-Bensussan, N.; Heyman, M. Multiple facets of intestinal permeability and epithelial handling of dietary antigens. Mucosal Immunol. 2010, 3, 247–259. [Google Scholar] [CrossRef] [PubMed]
  44. Bischoff, S.C.; Barbara, G.; Buurman, W.; Ockhuizen, T.; Schulzke, J.D.; Serino, M.; Tilg, H.; Watson, A.; Wells, J.M. Intestinal permeability--a new target for disease prevention and therapy. BMC Gastroenterol. 2014, 14, 189. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Galipeau, H.J.; Verdu, E.F. The complex task of measuring intestinal permeability in basic and clinical science. Neurogastroenterol. Motil. 2016, 28, 957–965. [Google Scholar] [CrossRef] [PubMed]
  46. Clarke, L.L. A guide to Ussing chamber studies of mouse intestine. Am. J. Physiol. Gastrointest. Liver Physiol. 2009, 296, G1151–G1166. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  47. Cantarel, B.L.; Lombard, V.; Henrissat, B. Complex carbohydrate utilization by the healthy human microbiome. PLoS ONE 2012, 7, e28742. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  48. Sonnenburg, E.D.; Smits, S.A.; Tikhonov, M.; Higginbottom, S.K.; Wingreen, N.S.; Sonnenburg, J.L. Diet-induced extinctions in the gut microbiota compound over generations. Nature 2016, 529, 212–215. [Google Scholar] [CrossRef] [Green Version]
  49. Soliman, G.A. Dietary Fiber, Atherosclerosis, and Cardiovascular Disease. Nutrients 2019, 11, 1155. [Google Scholar] [CrossRef] [Green Version]
  50. Swann, O.G.; Kilpatrick, M.; Breslin, M.; Oddy, W.H. Dietary fiber and its associations with depression and inflammation. Nutr. Rev. 2020, 78, 394–411. [Google Scholar] [CrossRef]
  51. Titgemeyer, E.C.; Bourquin, L.D.; Fahey, G.C., Jr.; Garleb, K.A. Fermentability of various fiber sources by human fecal bacteria in vitro. Am. J. Clin. Nutr. 1991, 53, 1418–1424. [Google Scholar] [CrossRef]
  52. Kim, C.H. Immune regulation by microbiome metabolites. Immunology 2018, 154, 220–229. [Google Scholar] [CrossRef]
  53. Mahowald, M.A.; Rey, F.E.; Seedorf, H.; Turnbaugh, P.J.; Fulton, R.S.; Wollam, A.; Shah, N.; Wang, C.; Magrini, V.; Wilson, R.K.; et al. Characterizing a model human gut microbiota composed of members of its two dominant bacterial phyla. Proc. Natl. Acad. Sci. USA 2009, 106, 5859–5864. [Google Scholar] [CrossRef] [Green Version]
  54. Rios-Covian, D.; Gueimonde, M.; Duncan, S.H.; Flint, H.J.; de los Reyes-Gavilan, C.G. Enhanced butyrate formation by cross-feeding between Faecalibacterium prausnitzii and Bifidobacterium adolescentis. FEMS Microbiol. Lett. 2015, 362, fnv176. [Google Scholar] [CrossRef] [Green Version]
  55. Morrison, D.J.; Preston, T. Formation of short chain fatty acids by the gut microbiota and their impact on human metabolism. Gut Microbes 2016, 7, 189–200. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  56. Rios-Covian, D.; Ruas-Madiedo, P.; Margolles, A.; Gueimonde, M.; de Los Reyes-Gavilan, 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] [PubMed] [Green Version]
  57. Schirmer, M.; Smeekens, S.P.; Vlamakis, H.; Jaeger, M.; Oosting, M.; Franzosa, E.A.; Ter Horst, R.; Jansen, T.; Jacobs, L.; Bonder, M.J.; et al. Linking the Human Gut Microbiome to Inflammatory Cytokine Production Capacity. Cell 2016, 167, 1125–1136.e8. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  58. Canfora, E.E.; Jocken, J.W.; Blaak, E.E. Short-chain fatty acids in control of body weight and insulin sensitivity. Nat. Rev. Endocrinol. 2015, 11, 577–591. [Google Scholar] [CrossRef] [PubMed]
  59. Kim, C.H.; Park, J.; Kim, M. Gut microbiota-derived short-chain Fatty acids, T cells, and inflammation. Immune Netw. 2014, 14, 277–288. [Google Scholar] [CrossRef] [Green Version]
  60. Chambers, E.S.; Viardot, A.; Psichas, A.; Morrison, D.J.; Murphy, K.G.; Zac-Varghese, S.E.; MacDougall, K.; Preston, T.; Tedford, C.; Finlayson, G.S.; et al. Effects of targeted delivery of propionate to the human colon on appetite regulation, body weight maintenance and adiposity in overweight adults. Gut 2015, 64, 1744–1754. [Google Scholar] [CrossRef] [Green Version]
  61. Zambell, K.L.; Fitch, M.D.; Fleming, S.E. Acetate and butyrate are the major substrates for de novo lipogenesis in rat colonic epithelial cells. J. Nutr. 2003, 133, 3509–3515. [Google Scholar] [CrossRef]
  62. Kim, M.H.; Kang, S.G.; Park, J.H.; Yanagisawa, M.; Kim, C.H. Short-chain fatty acids activate GPR41 and GPR43 on intestinal epithelial cells to promote inflammatory responses in mice. Gastroenterology 2013, 145, e1–e10. [Google Scholar] [CrossRef] [PubMed]
  63. Pham, V.T.; Calatayud, M.; Rotsaert, C.; Seifert, N.; Richard, N.; Van den Abbeele, P.; Marzorati, M.; Steinert, R.E. Antioxidant Vitamins and Prebiotic FOS and XOS Differentially Shift Microbiota Composition and Function and Improve Intestinal Epithelial Barrier In Vitro. Nutrients 2021, 13, 1125. [Google Scholar] [CrossRef] [PubMed]
  64. Mathewson, N.D.; Jenq, R.; Mathew, A.V.; Koenigsknecht, M.; Hanash, A.; Toubai, T.; Oravecz-Wilson, K.; Wu, S.R.; Sun, Y.; Rossi, C.; et al. Gut microbiome-derived metabolites modulate intestinal epithelial cell damage and mitigate graft-versus-host disease. Nat. Immunol. 2016, 17, 505–513. [Google Scholar] [CrossRef] [PubMed]
  65. Huang, X.; Oshima, T.; Tomita, T.; Fukui, H.; Miwa, H. Butyrate Alleviates Cytokine-Induced Barrier Dysfunction by Modifying Claudin-2 Levels. Biology 2021, 10, 205. [Google Scholar] [CrossRef]
  66. Chen, G.; Ran, X.; Li, B.; Li, Y.; He, D.; Huang, B.; Fu, S.; Liu, J.; Wang, W. Sodium Butyrate Inhibits Inflammation and Maintains Epithelium Barrier Integrity in a TNBS-induced Inflammatory Bowel Disease Mice Model. EBioMedicine 2018, 30, 317–325. [Google Scholar] [CrossRef] [Green Version]
  67. Zhao, H.B.; Jia, L.; Yan, Q.Q.; Deng, Q.; Wei, B. Effect of Clostridium butyricum and Butyrate on Intestinal Barrier Functions: Study of a Rat Model of Severe Acute Pancreatitis with Intra-Abdominal Hypertension. Front. Physiol. 2020, 11, 561061. [Google Scholar] [CrossRef]
  68. Richards, L.B.; Li, M.; Folkerts, G.; Henricks, P.A.J.; Garssen, J.; van Esch, B. Butyrate and Propionate Restore the Cytokine and House Dust Mite Compromised Barrier Function of Human Bronchial Airway Epithelial Cells. Int. J. Mol. Sci. 2020, 22, 65. [Google Scholar] [CrossRef]
  69. Vernero, M.; De Blasio, F.; Ribaldone, D.G.; Bugianesi, E.; Pellicano, R.; Saracco, G.M.; Astegiano, M.; Caviglia, G.P. The Usefulness of Microencapsulated Sodium Butyrate Add-On Therapy in Maintaining Remission in Patients with Ulcerative Colitis: A Prospective Observational Study. J. Clin. Med. 2020, 9, 3941. [Google Scholar] [CrossRef]
  70. Gaudier, E.; Jarry, A.; Blottière, H.M.; de Coppet, P.; Buisine, M.P.; Aubert, J.P.; Laboisse, C.; Cherbut, C.; Hoebler, C. Butyrate specifically modulates MUC gene expression in intestinal epithelial goblet cells deprived of glucose. Am. J. Physiol. Gastrointest. Liver Physiol. 2004, 287, G1168–G1174. [Google Scholar] [CrossRef] [Green Version]
  71. Macia, L.; Tan, J.; Vieira, A.T.; Leach, K.; Stanley, D.; Luong, S.; Maruya, M.; Ian McKenzie, C.; Hijikata, A.; Wong, C.; et al. Metabolite-sensing receptors GPR43 and GPR109A facilitate dietary fibre-induced gut homeostasis through regulation of the inflammasome. Nat. Commun. 2015, 6, 6734. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  72. Nowarski, R.; Jackson, R.; Gagliani, N.; de Zoete, M.R.; Palm, N.W.; Bailis, W.; Low, J.S.; Harman, C.C.; Graham, M.; Elinav, E.; et al. Epithelial IL-18 Equilibrium Controls Barrier Function in Colitis. Cell 2015, 163, 1444–1456. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Impellizzeri, D.; Siracusa, R.; Cordaro, M.; Peritore, A.F.; Gugliandolo, E.; Mancuso, G.; Midiri, A.; Di Paola, R.; Cuzzocrea, S. Therapeutic potential of dinitrobenzene sulfonic acid (DNBS)-induced colitis in mice by targeting IL-1β and IL-18. Biochem. Pharmacol. 2018, 155, 150–161. [Google Scholar] [CrossRef]
  74. Sivakumar, P.V.; Westrich, G.M.; Kanaly, S.; Garka, K.; Born, T.L.; Derry, J.M.; Viney, J.L. Interleukin 18 is a primary mediator of the inflammation associated with dextran sulphate sodium induced colitis: Blocking interleukin 18 attenuates intestinal damage. Gut 2002, 50, 812–820. [Google Scholar] [CrossRef] [Green Version]
  75. Tong, L.C.; Wang, Y.; Wang, Z.B.; Liu, W.Y.; Sun, S.; Li, L.; Su, D.F.; Zhang, L.C. Propionate Ameliorates Dextran Sodium Sulfate-Induced Colitis by Improving Intestinal Barrier Function and Reducing Inflammation and Oxidative Stress. Front. Pharmacol. 2016, 7, 253. [Google Scholar] [CrossRef]
  76. Feng, Y.; Wang, Y.; Wang, P.; Huang, Y.; Wang, F. Short-Chain Fatty Acids Manifest Stimulative and Protective Effects on Intestinal Barrier Function Through the Inhibition of NLRP3 Inflammasome and Autophagy. Cell Physiol. Biochem. 2018, 49, 190–205. [Google Scholar] [CrossRef]
  77. Hytting-Andreasen, R.; Balk-Møller, E.; Hartmann, B.; Pedersen, J.; Windeløv, J.A.; Holst, J.J.; Kissow, H. Endogenous glucagon-like peptide- 1 and 2 are essential for regeneration after acute intestinal injury in mice. PLoS ONE 2018, 13, e0198046. [Google Scholar] [CrossRef]
  78. Maruta, K.; Takajo, T.; Akiba, Y.; Said, H.; Irie, E.; Kato, I.; Kuwahara, A.; Kaunitz, J.D. GLP-2 Acutely Prevents Endotoxin-Related Increased Intestinal Paracellular Permeability in Rats. Dig. Dis. Sci. 2020, 65, 2605–2618. [Google Scholar] [CrossRef]
  79. Hunt, J.E.; Hartmann, B.; Schoonjans, K.; Holst, J.J.; Kissow, H. Dietary Fiber Is Essential to Maintain Intestinal Size, L-Cell Secretion, and Intestinal Integrity in Mice. Front. Endocrinol. 2021, 12, 640602. [Google Scholar] [CrossRef] [PubMed]
  80. Genda, T.; Sasaki, Y.; Kondo, T.; Hino, S.; Nishimura, N.; Tsukahara, T.; Sonoyama, K.; Morita, T. Fructo-oligosaccharide-Induced Transient Increases in Cecal Immunoglobulin A Concentrations in Rats Are Associated with Mucosal Inflammation in Response to Increased Gut Permeability. J. Nutr. 2017, 147, 1900–1908. [Google Scholar] [CrossRef] [PubMed]
  81. Wang, S.; Zhang, S.; Huang, S.; Wu, Z.; Pang, J.; Wu, Y.; Wang, J.; Han, D. Resistant Maltodextrin Alleviates Dextran Sulfate Sodium-Induced Intestinal Inflammatory Injury by Increasing Butyric Acid to Inhibit Proinflammatory Cytokine Levels. Biomed. Res. Int. 2020, 2020, 7694734. [Google Scholar] [CrossRef]
  82. Wilms, E.; Gerritsen, J.; Smidt, H.; Besseling-van der Vaart, I.; Rijkers, G.T.; Garcia Fuentes, A.R.; Masclee, A.A.; Troost, F.J. Effects of Supplementation of the Synbiotic Ecologic® 825/FOS P6 on Intestinal Barrier Function in Healthy Humans: A Randomized Controlled Trial. PLoS ONE 2016, 11, e0167775. [Google Scholar] [CrossRef] [Green Version]
  83. Chen, T.; Ma, Y.; Xu, L.; Sun, C.; Xu, H.; Zhu, J. Soluble Dietary Fiber Reduces Feeding Intolerance in Severe Acute Pancreatitis: A Randomized Study. JPEN J. Parenter. Enteral Nutr. 2021, 45, 125–135. [Google Scholar] [CrossRef] [PubMed]
  84. Drabińska, N.; Krupa-Kozak, U.; Jarocka-Cyrta, E. Intestinal Permeability in Children with Celiac Disease after the Administration of Oligofructose-Enriched Inulin into a Gluten-Free Diet-Results of a Randomized, Placebo-Controlled, Pilot Trial. Nutrients 2020, 12, 1736. [Google Scholar] [CrossRef]
  85. Ganda Mall, J.P.; Fart, F.; Sabet, J.A.; Lindqvist, C.M.; Nestestog, R.; Hegge, F.T.; Keita, Å.V.; Brummer, R.J.; Schoultz, I. Effects of Dietary Fibres on Acute Indomethacin-Induced Intestinal Hyperpermeability in the Elderly: A Randomised Placebo Controlled Parallel Clinical Trial. Nutrients 2020, 12, 1954. [Google Scholar] [CrossRef]
  86. Singh, A.; Zapata, R.C.; Pezeshki, A.; Reidelberger, R.D.; Chelikani, P.K. Inulin fiber dose-dependently modulates energy balance, glucose tolerance, gut microbiota, hormones and diet preference in high-fat-fed male rats. J. Nutr. Biochem. 2018, 59, 142–152. [Google Scholar] [CrossRef] [PubMed]
  87. Vandeputte, D.; Falony, G.; Vieira-Silva, S.; Wang, J.; Sailer, M.; Theis, S.; Verbeke, K.; Raes, J. Prebiotic inulin-type fructans induce specific changes in the human gut microbiota. Gut 2017, 66, 1968–1974. [Google Scholar] [CrossRef] [PubMed]
  88. Karlsson, F.H.; Tremaroli, V.; Nookaew, I.; Bergstrom, G.; Behre, C.J.; Fagerberg, B.; Nielsen, J.; Backhed, F. Gut metagenome in European women with normal, impaired and diabetic glucose control. Nature 2013, 498, 99–103. [Google Scholar] [CrossRef]
  89. Qin, J.; Li, Y.; Cai, Z.; Li, S.; Zhu, J.; Zhang, F.; Liang, S.; Zhang, W.; Guan, Y.; Shen, D.; et al. A metagenome-wide association study of gut microbiota in type 2 diabetes. Nature 2012, 490, 55–60. [Google Scholar] [CrossRef] [PubMed]
  90. Trompette, A.; Gollwitzer, E.S.; Yadava, K.; Sichelstiel, A.K.; Sprenger, N.; Ngom-Bru, C.; Blanchard, C.; Junt, T.; Nicod, L.P.; Harris, N.L.; et al. Gut microbiota metabolism of dietary fiber influences allergic airway disease and hematopoiesis. Nat. Med. 2014, 20, 159–166. [Google Scholar] [CrossRef]
  91. Sonnenburg, J.L.; Xu, J.; Leip, D.D.; Chen, C.H.; Westover, B.P.; Weatherford, J.; Buhler, J.D.; Gordon, J.I. Glycan foraging in vivo by an intestine-adapted bacterial symbiont. Science 2005, 307, 1955–1959. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  92. 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.e21. [Google Scholar] [CrossRef] [Green Version]
  93. Everard, A.; Belzer, C.; Geurts, L.; Ouwerkerk, J.P.; Druart, C.; Bindels, L.B.; Guiot, Y.; Derrien, M.; Muccioli, G.G.; Delzenne, N.M.; et al. Cross-talk between Akkermansia muciniphila and intestinal epithelium controls diet-induced obesity. Proc. Natl. Acad. Sci. USA 2013, 110, 9066–9071. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Derrien, M.; Collado, M.C.; Ben-Amor, K.; Salminen, S.; de Vos, W.M. The Mucin degrader Akkermansia muciniphila is an abundant resident of the human intestinal tract. Appl. Environ. Microbiol. 2008, 74, 1646–1648. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Zhang, T.; Li, P.; Wu, X.; Lu, G.; Marcella, C.; Ji, X.; Ji, G.; Zhang, F. Alterations of Akkermansia muciniphila in the inflammatory bowel disease patients with washed microbiota transplantation. Appl. Microbiol. Biotechnol. 2020, 104, 10203–10215. [Google Scholar] [CrossRef] [PubMed]
  96. Bajer, L.; Kverka, M.; Kostovcik, M.; Macinga, P.; Dvorak, J.; Stehlikova, Z.; Brezina, J.; Wohl, P.; Spicak, J.; Drastich, P. Distinct gut microbiota profiles in patients with primary sclerosing cholangitis and ulcerative colitis. World J. Gastroenterol. 2017, 23, 4548–4558. [Google Scholar] [CrossRef] [PubMed]
  97. Shang, L.; Yu, H.; Liu, H.; Chen, M.; Zeng, X.; Qiao, S. Recombinant antimicrobial peptide microcin J25 alleviates DSS-induced colitis via regulating intestinal barrier function and modifying gut microbiota. Biomed. Pharmacother. 2021, 139, 111127. [Google Scholar] [CrossRef]
  98. Håkansson, Å.; Tormo-Badia, N.; Baridi, A.; Xu, J.; Molin, G.; Hagslätt, M.L.; Karlsson, C.; Jeppsson, B.; Cilio, C.M.; Ahrné, S. Immunological alteration and changes of gut microbiota after dextran sulfate sodium (DSS) administration in mice. Clin. Exp. Med. 2015, 15, 107–120. [Google Scholar] [CrossRef] [Green Version]
  99. Bian, X.; Wu, W.; Yang, L.; Lv, L.; Wang, Q.; Li, Y.; Ye, J.; Fang, D.; Wu, J.; Jiang, X.; et al. Administration of Akkermansia muciniphila Ameliorates Dextran Sulfate Sodium-Induced Ulcerative Colitis in Mice. Front. Microbiol. 2019, 10, 2259. [Google Scholar] [CrossRef] [Green Version]
  100. Zhai, R.; Xue, X.; Zhang, L.; Yang, X.; Zhao, L.; Zhang, C. Strain-Specific Anti-inflammatory Properties of Two Akkermansia muciniphila Strains on Chronic Colitis in Mice. Front. Cell Infect. Microbiol. 2019, 9, 239. [Google Scholar] [CrossRef]
  101. Wang, L.; Tang, L.; Feng, Y.; Zhao, S.; Han, M.; Zhang, C.; Yuan, G.; Zhu, J.; Cao, S.; Wu, Q.; et al. A purified membrane protein from Akkermansia muciniphila or the pasteurised bacterium blunts colitis associated tumourigenesis by modulation of CD8(+) T cells in mice. Gut 2020, 69, 1988–1997. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  102. Ottman, N.; Reunanen, J.; Meijerink, M.; Pietilä, T.E.; Kainulainen, V.; Klievink, J.; Huuskonen, L.; Aalvink, S.; Skurnik, M.; Boeren, S.; et al. Pili-like proteins of Akkermansia muciniphila modulate host immune responses and gut barrier function. PLoS ONE 2017, 12, e0173004. [Google Scholar] [CrossRef]
  103. Usami, M.; Muraki, K.; Iwamoto, M.; Ohata, A.; Matsushita, E.; Miki, A. Effect of eicosapentaenoic acid (EPA) on tight junction permeability in intestinal monolayer cells. Clin. Nutr. 2001, 20, 351–359. [Google Scholar] [CrossRef] [PubMed]
  104. Usami, M.; Komurasaki, T.; Hanada, A.; Kinoshita, K.; Ohata, A. Effect of gamma-linolenic acid or docosahexaenoic acid on tight junction permeability in intestinal monolayer cells and their mechanism by protein kinase C activation and/or eicosanoid formation. Nutrition 2003, 19, 150–156. [Google Scholar] [CrossRef]
  105. Willemsen, L.E.; Koetsier, M.A.; Balvers, M.; Beermann, C.; Stahl, B.; van Tol, E.A. Polyunsaturated fatty acids support epithelial barrier integrity and reduce IL-4 mediated permeability in vitro. Eur. J. Nutr. 2008, 47, 183–191. [Google Scholar] [CrossRef] [PubMed]
  106. Lindmark, T.; Nikkilä, T.; Artursson, P. Mechanisms of absorption enhancement by medium chain fatty acids in intestinal epithelial Caco-2 cell monolayers. J. Pharmacol. Exp. Ther. 1995, 275, 958–964. [Google Scholar]
  107. Anderberg, E.K.; Lindmark, T.; Artursson, P. Sodium caprate elicits dilatations in human intestinal tight junctions and enhances drug absorption by the paracellular route. Pharm. Res. 1993, 10, 857–864. [Google Scholar] [CrossRef]
  108. John, S.; Luben, R.; Shrestha, S.S.; Welch, A.; Khaw, K.T.; Hart, A.R. Dietary n-3 polyunsaturated fatty acids and the aetiology of ulcerative colitis: A UK prospective cohort study. Eur. J. Gastroenterol. Hepatol. 2010, 22, 602–606. [Google Scholar] [CrossRef]
  109. Chapkin, R.S.; Davidson, L.A.; Ly, L.; Weeks, B.R.; Lupton, J.R.; McMurray, D.N. Immunomodulatory effects of (n-3) fatty acids: Putative link to inflammation and colon cancer. J. Nutr. 2007, 137 (Suppl. 1), 200s–204s. [Google Scholar] [CrossRef] [Green Version]
  110. Patterson, E.; Wall, R.; Fitzgerald, G.F.; Ross, R.P.; Stanton, C. Health implications of high dietary omega-6 polyunsaturated Fatty acids. J. Nutr. Metab. 2012, 2012, 539426. [Google Scholar] [CrossRef]
  111. Schreiner, P.; Martinho-Grueber, M.; Studerus, D.; Vavricka, S.R.; Tilg, H.; Biedermann, L. Nutrition in Inflammatory Bowel Disease. Digestion 2020, 101 (Suppl. 1), 120–135. [Google Scholar] [CrossRef] [PubMed]
  112. Uchiyama, K.; Nakamura, M.; Odahara, S.; Koido, S.; Katahira, K.; Shiraishi, H.; Ohkusa, T.; Fujise, K.; Tajiri, H. N-3 polyunsaturated fatty acid diet therapy for patients with inflammatory bowel disease. Inflamm. Bowel Dis. 2010, 16, 1696–1707. [Google Scholar] [CrossRef]
  113. Scoville, E.A.; Allaman, M.M.; Adams, D.W.; Motley, A.K.; Peyton, S.C.; Ferguson, S.L.; Horst, S.N.; Williams, C.S.; Beaulieu, D.B.; Schwartz, D.A.; et al. Serum Polyunsaturated Fatty Acids Correlate with Serum Cytokines and Clinical Disease Activity in Crohn’s Disease. Sci. Rep. 2019, 9, 2882. [Google Scholar] [CrossRef] [PubMed]
  114. Nascimento, J.C.; Matheus, V.A.; Oliveira, R.B.; Tada, S.F.S.; Collares-Buzato, C.B. High-Fat Diet Induces Disruption of the Tight Junction-Mediated Paracellular Barrier in the Proximal Small Intestine Before the Onset of Type 2 Diabetes and Endotoxemia. Dig. Dis. Sci. 2020. [CrossRef] [PubMed]
  115. Mujawdiya, P.K.; Sharma, P.; Sharad, S.; Kapur, S. Reversal of Increase in Intestinal Permeability by Mangifera indica Seed Kernel Extract in High-Fat Diet-Induced Obese Mice. Pharmaceuticals 2020, 13, 190. [Google Scholar] [CrossRef]
  116. Tian, B.; Zhao, J.; Zhang, M.; Chen, Z.; Ma, Q.; Liu, H.; Nie, C.; Zhang, Z.; An, W.; Li, J. Lycium ruthenicum Anthocyanins Attenuate High-Fat Diet-Induced Colonic Barrier Dysfunction and Inflammation in Mice by Modulating the Gut Microbiota. Mol. Nutr. Food Res. 2021, 65, e2000745. [Google Scholar] [CrossRef]
  117. de La Serre, C.B.; Ellis, C.L.; Lee, J.; Hartman, A.L.; Rutledge, J.C.; Raybould, H.E. Propensity to high-fat diet-induced obesity in rats is associated with changes in the gut microbiota and gut inflammation. Am. J. Physiol. Gastrointest. Liver Physiol. 2010, 299, G440–G448. [Google Scholar] [CrossRef]
  118. Zhao, J.; Wang, H.; Yang, H.; Zhou, Y.; Tang, L. Autophagy induction by rapamycin ameliorates experimental colitis and improves intestinal epithelial barrier function in IL-10 knockout mice. Int. Immunopharmacol. 2020, 81, 105977. [Google Scholar] [CrossRef]
  119. Devkota, S.; Wang, Y.; Musch, M.W.; Leone, V.; Fehlner-Peach, H.; Nadimpalli, A.; Antonopoulos, D.A.; Jabri, B.; Chang, E.B. Dietary-fat-induced taurocholic acid promotes pathobiont expansion and colitis in Il10-/- mice. Nature 2012, 487, 104–108. [Google Scholar] [CrossRef] [Green Version]
  120. Agus, A.; Denizot, J.; Thévenot, J.; Martinez-Medina, M.; Massier, S.; Sauvanet, P.; Bernalier-Donadille, A.; Denis, S.; Hofman, P.; Bonnet, R.; et al. Western diet induces a shift in microbiota composition enhancing susceptibility to Adherent-Invasive E. coli infection and intestinal inflammation. Sci. Rep. 2016, 6, 19032. [Google Scholar] [CrossRef] [Green Version]
  121. Muhomah, T.A.; Nishino, N.; Katsumata, E.; Haoming, W.; Tsuruta, T. High-fat diet reduces the level of secretory immunoglobulin A coating of commensal gut microbiota. Biosci. Microbiota Food Health 2019, 38, 55–64. [Google Scholar] [CrossRef] [Green Version]
  122. Amar, J.; Burcelin, R.; Ruidavets, J.B.; Cani, P.D.; Fauvel, J.; Alessi, M.C.; Chamontin, B.; Ferriéres, J. Energy intake is associated with endotoxemia in apparently healthy men. Am. J. Clin. Nutr. 2008, 87, 1219–1223. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  123. Yang, Y.; Zhong, Z.; Wang, B.; Xia, X.; Yao, W.; Huang, L.; Wang, Y.; Ding, W. Early-life high-fat diet-induced obesity programs hippocampal development and cognitive functions via regulation of gut commensal Akkermansia muciniphila. Neuropsychopharmacology 2019, 44, 2054–2064. [Google Scholar] [CrossRef] [PubMed]
  124. Lassenius, M.I.; Pietiläinen, K.H.; Kaartinen, K.; Pussinen, P.J.; Syrjänen, J.; Forsblom, C.; Pörsti, I.; Rissanen, A.; Kaprio, J.; Mustonen, J.; et al. Bacterial endotoxin activity in human serum is associated with dyslipidemia, insulin resistance, obesity, and chronic inflammation. Diabetes Care 2011, 34, 1809–1815. [Google Scholar] [CrossRef] [Green Version]
  125. Shi, H.; Kokoeva, M.V.; Inouye, K.; Tzameli, I.; Yin, H.; Flier, J.S. TLR4 links innate immunity and fatty acid-induced insulin resistance. J. Clin. Investig. 2006, 116, 3015–3025. [Google Scholar] [CrossRef]
  126. Lee, J.J.; Wang, P.W.; Yang, I.H.; Huang, H.M.; Chang, C.S.; Wu, C.L.; Chuang, J.H. High-fat diet induces toll-like receptor 4-dependent macrophage/microglial cell activation and retinal impairment. Invest. Ophthalmol. Vis. Sci. 2015, 56, 3041–3050. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  127. Jia, L.; Vianna, C.R.; Fukuda, M.; Berglund, E.D.; Liu, C.; Tao, C.; Sun, K.; Liu, T.; Harper, M.J.; Lee, C.E.; et al. Hepatocyte Toll-like receptor 4 regulates obesity-induced inflammation and insulin resistance. Nat. Commun. 2014, 5, 3878. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  128. Ding, Y.; Subramanian, S.; Montes, V.N.; Goodspeed, L.; Wang, S.; Han, C.; Teresa, A.S., 3rd; Kim, J.; O’Brien, K.D.; Chait, A. Toll-like receptor 4 deficiency decreases atherosclerosis but does not protect against inflammation in obese low-density lipoprotein receptor-deficient mice. Arterioscler. Thromb. Vasc. Biol. 2012, 32, 1596–1604. [Google Scholar] [CrossRef] [Green Version]
  129. Ghanim, H.; Abuaysheh, S.; Sia, C.L.; Korzeniewski, K.; Chaudhuri, A.; Fernandez-Real, J.M.; Dandona, P. Increase in plasma endotoxin concentrations and the expression of Toll-like receptors and suppressor of cytokine signaling-3 in mononuclear cells after a high-fat, high-carbohydrate meal: Implications for insulin resistance. Diabetes Care 2009, 32, 2281–2287. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  130. Pendyala, S.; Walker, J.M.; Holt, P.R. A high-fat diet is associated with endotoxemia that originates from the gut. Gastroenterology 2012, 142, 1100–1101.e2. [Google Scholar] [CrossRef] [Green Version]
  131. Park, J.H.; Jeong, S.Y.; Choi, A.J.; Kim, S.J. Lipopolysaccharide directly stimulates Th17 differentiation in vitro modulating phosphorylation of RelB and NF-κB1. Immunol. Lett. 2015, 165, 10–19. [Google Scholar] [CrossRef]
  132. Shen, T.; Chen, X.; Li, Y.; Tang, X.; Jiang, X.; Yu, C.; Zheng, Y.; Guo, H.; Ling, W. Interleukin-17A exacerbates high-fat diet-induced hepatic steatosis by inhibiting fatty acid β-oxidation. Biochim. Biophys. Acta Mol. Basis Dis. 2017, 1863, 1510–1518. [Google Scholar] [CrossRef]
  133. Cheng, C.; Tan, J.; Qian, W.; Zhang, L.; Hou, X. Gut inflammation exacerbates hepatic injury in the high-fat diet induced NAFLD mouse: Attention to the gut-vascular barrier dysfunction. Life Sci. 2018, 209, 157–166. [Google Scholar] [CrossRef]
  134. Fujisaka, S.; Avila-Pacheco, J.; Soto, M.; Kostic, A.; Dreyfuss, J.M.; Pan, H.; Ussar, S.; Altindis, E.; Li, N.; Bry, L.; et al. Diet, Genetics, and the Gut Microbiome Drive Dynamic Changes in Plasma Metabolites. Cell Rep. 2018, 22, 3072–3086. [Google Scholar] [CrossRef] [Green Version]
  135. Münch, N.S.; Fang, H.Y.; Ingermann, J.; Maurer, H.C.; Anand, A.; Kellner, V.; Sahm, V.; Wiethaler, M.; Baumeister, T.; Wein, F.; et al. High-Fat Diet Accelerates Carcinogenesis in a Mouse Model of Barrett’s Esophagus via Interleukin 8 and Alterations to the Gut Microbiome. Gastroenterology 2019, 157, 492–506.e2. [Google Scholar] [CrossRef] [Green Version]
  136. Hassan, A.M.; Mancano, G.; Kashofer, K.; Fröhlich, E.E.; Matak, A.; Mayerhofer, R.; Reichmann, F.; Olivares, M.; Neyrinck, A.M.; Delzenne, N.M.; et al. High-fat diet induces depression-like behaviour in mice associated with changes in microbiome, neuropeptide Y, and brain metabolome. Nutr. Neurosci. 2019, 22, 877–893. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  137. Bisanz, J.E.; Upadhyay, V.; Turnbaugh, J.A.; Ly, K.; Turnbaugh, P.J. Meta-Analysis Reveals Reproducible Gut Microbiome Alterations in Response to a High-Fat Diet. Cell Host Microbe 2019, 26, 265–272.e4. [Google Scholar] [CrossRef] [PubMed]
  138. Sun, L.; Xie, C.; Wang, G.; Wu, Y.; Wu, Q.; Wang, X.; Liu, J.; Deng, Y.; Xia, J.; Chen, B.; et al. Gut microbiota and intestinal FXR mediate the clinical benefits of metformin. Nat. Med. 2018, 24, 1919–1929. [Google Scholar] [CrossRef] [PubMed]
  139. Wei, L.; Yue, F.; Xing, L.; Wu, S.; Shi, Y.; Li, J.; Xiang, X.; Lam, S.M.; Shui, G.; Russell, R.; et al. Constant Light Exposure Alters Gut Microbiota and Promotes the Progression of Steatohepatitis in High Fat Diet Rats. Front. Microbiol. 2020, 11, 1975. [Google Scholar] [CrossRef]
  140. 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, 426–436. [Google Scholar] [CrossRef] [Green Version]
  141. Chelakkot, C.; Choi, Y.; Kim, D.K.; Park, H.T.; Ghim, J.; Kwon, Y.; Jeon, J.; Kim, M.S.; Jee, Y.K.; Gho, Y.S.; et al. Akkermansia muciniphila-derived extracellular vesicles influence gut permeability through the regulation of tight junctions. Exp. Mol. Med. 2018, 50, e450. [Google Scholar] [CrossRef]
  142. Ashrafian, F.; Shahriary, A.; Behrouzi, A.; Moradi, H.R.; Keshavarz Azizi Raftar, S.; Lari, A.; Hadifar, S.; Yaghoubfar, R.; Ahmadi Badi, S.; Khatami, S.; et al. Akkermansia muciniphila-Derived Extracellular Vesicles as a Mucosal Delivery Vector for Amelioration of Obesity in Mice. Front. Microbiol. 2019, 10, 2155. [Google Scholar] [CrossRef]
  143. Plovier, H.; Everard, A.; Druart, C.; Depommier, C.; Van Hul, M.; Geurts, L.; Chilloux, J.; Ottman, N.; Duparc, T.; Lichtenstein, L.; et al. A purified membrane protein from Akkermansia muciniphila or the pasteurized bacterium improves metabolism in obese and diabetic mice. Nat. Med. 2017, 23, 107–113. [Google Scholar] [CrossRef] [Green Version]
  144. Murakami, Y.; Tanabe, S.; Suzuki, T. High-fat Diet-induced Intestinal Hyperpermeability is Associated with Increased Bile Acids in the Large Intestine of Mice. J. Food Sci. 2016, 81, H216–H222. [Google Scholar] [CrossRef] [PubMed]
  145. Zeng, H.; Umar, S.; Rust, B.; Lazarova, D.; Bordonaro, M. Secondary Bile Acids and Short Chain Fatty Acids in the Colon: A Focus on Colonic Microbiome, Cell Proliferation, Inflammation, and Cancer. Int. J. Mol. Sci. 2019, 20, 1214. [Google Scholar] [CrossRef] [Green Version]
  146. Chiang, J.Y.; Pathak, P.; Liu, H.; Donepudi, A.; Ferrell, J.; Boehme, S. Intestinal Farnesoid X Receptor and Takeda G Protein Couple Receptor 5 Signaling in Metabolic Regulation. Dig. Dis. 2017, 35, 241–245. [Google Scholar] [CrossRef] [PubMed]
  147. Cipriani, S.; Mencarelli, A.; Chini, M.G.; Distrutti, E.; Renga, B.; Bifulco, G.; Baldelli, F.; Donini, A.; Fiorucci, S. The bile acid receptor GPBAR-1 (TGR5) modulates integrity of intestinal barrier and immune response to experimental colitis. PLoS ONE 2011, 6, e25637. [Google Scholar] [CrossRef] [PubMed]
  148. Biagioli, M.; Carino, A.; Cipriani, S.; Francisci, D.; Marchianò, S.; Scarpelli, P.; Sorcini, D.; Zampella, A.; Fiorucci, S. The Bile Acid Receptor GPBAR1 Regulates the M1/M2 Phenotype of Intestinal Macrophages and Activation of GPBAR1 Rescues Mice from Murine Colitis. J. Immunol. 2017, 199, 718–733. [Google Scholar] [CrossRef] [Green Version]
  149. Inagaki, T.; Moschetta, A.; Lee, Y.K.; Peng, L.; Zhao, G.; Downes, M.; Yu, R.T.; Shelton, J.M.; Richardson, J.A.; Repa, J.J.; et al. Regulation of antibacterial defense in the small intestine by the nuclear bile acid receptor. Proc. Natl. Acad. Sci. USA 2006, 103, 3920–3925. [Google Scholar] [CrossRef] [Green Version]
  150. Gadaleta, R.M.; van Erpecum, K.J.; Oldenburg, B.; Willemsen, E.C.; Renooij, W.; Murzilli, S.; Klomp, L.W.; Siersema, P.D.; Schipper, M.E.; Danese, S.; et al. Farnesoid X receptor activation inhibits inflammation and preserves the intestinal barrier in inflammatory bowel disease. Gut 2011, 60, 463–472. [Google Scholar] [CrossRef]
  151. Huang, M.; Kong, B.; Zhang, M.; Rizzolo, D.; Armstrong, L.E.; Schumacher, J.D.; Chow, M.D.; Lee, Y.H.; Joseph, L.B.; Stofan, M.; et al. Enhanced alcoholic liver disease in mice with intestine-specific farnesoid X receptor deficiency. Lab. Investig. 2020, 100, 1158–1168. [Google Scholar] [CrossRef] [PubMed]
Ijms 22 07613 g001 550
Figure 1. Relationships between nutrients, gut microbiome, and intestinal barrier. LPS: lipopolysaccharides, SCFAs: short-chain fatty acids.
Figure 1. Relationships between nutrients, gut microbiome, and intestinal barrier. LPS: lipopolysaccharides, SCFAs: short-chain fatty acids.
Ijms 22 07613 g001
Ijms 22 07613 g002 550
Figure 2. Construction of intestinal barrier.
Figure 2. Construction of intestinal barrier.
Ijms 22 07613 g002
Ijms 22 07613 g003 550
Figure 3. The mode of action for opening pore and leak pathways.
Figure 3. The mode of action for opening pore and leak pathways.
Ijms 22 07613 g003
Ijms 22 07613 g004 550
Figure 4. Effects of microbiota-accessible carbohydrates (MACs) on tight junction (TJ). Intestinal bacteria digest fermentable MACs and produce short-chain fatty acids (SFCA) which are absorbed into epithelial cells via solute carrier family (SLC) transporter or simple diffusion. Acetate, butyrate, and propionate are converted to acetyl-CoA or propynyl-CoA via acetyl-CoA carboxylases (ACSSs) or β-oxidation to produce ATP which maintain cell homeostasis including the function of TJ. Acetate activates nucleotide-binding oligomerization domain 3 (NLRP3) and promotes the secretion of IL-18 from epithelial cells, which contributes to TJ function. Other SCFA also have a protective effect on TJ. MACs also may contribute to TJ function via regulating the growth of Akkermansia muciniphilla (A. muciniphilla) or the production of Glucagon-Like Peptide (GLP) 1 and GLP-2.
Figure 4. Effects of microbiota-accessible carbohydrates (MACs) on tight junction (TJ). Intestinal bacteria digest fermentable MACs and produce short-chain fatty acids (SFCA) which are absorbed into epithelial cells via solute carrier family (SLC) transporter or simple diffusion. Acetate, butyrate, and propionate are converted to acetyl-CoA or propynyl-CoA via acetyl-CoA carboxylases (ACSSs) or β-oxidation to produce ATP which maintain cell homeostasis including the function of TJ. Acetate activates nucleotide-binding oligomerization domain 3 (NLRP3) and promotes the secretion of IL-18 from epithelial cells, which contributes to TJ function. Other SCFA also have a protective effect on TJ. MACs also may contribute to TJ function via regulating the growth of Akkermansia muciniphilla (A. muciniphilla) or the production of Glucagon-Like Peptide (GLP) 1 and GLP-2.
Ijms 22 07613 g004
Ijms 22 07613 g005 550
Figure 5. Effects of high-fat diet (HFD) and bile acids (BAs) on tight junction (TJ). Fatty acids impair TJ function directly and open the leak pathway. In addition, HFD affects the pollution of bacteria and exacerbates LPS production. Dysfunction of TJ lets LPS invade into epithelial cells where it becomes the target of immune cells, which provokes further damage to TJ. HFD also induces secretion of BAs which are converted to cytotoxic secondary BAs (SBAs) by bacteria. Nevertheless, SBAs show a protective effect on TJ function via G protein-coupled bile acid receptor (GPBAR1) and farnesoid X receptor (FXR).
Figure 5. Effects of high-fat diet (HFD) and bile acids (BAs) on tight junction (TJ). Fatty acids impair TJ function directly and open the leak pathway. In addition, HFD affects the pollution of bacteria and exacerbates LPS production. Dysfunction of TJ lets LPS invade into epithelial cells where it becomes the target of immune cells, which provokes further damage to TJ. HFD also induces secretion of BAs which are converted to cytotoxic secondary BAs (SBAs) by bacteria. Nevertheless, SBAs show a protective effect on TJ function via G protein-coupled bile acid receptor (GPBAR1) and farnesoid X receptor (FXR).
Ijms 22 07613 g005
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Usuda, H.; Okamoto, T.; Wada, K. Leaky Gut: Effect of Dietary Fiber and Fats on Microbiome and Intestinal Barrier. Int. J. Mol. Sci. 2021, 22, 7613. https://doi.org/10.3390/ijms22147613

AMA Style

Usuda H, Okamoto T, Wada K. Leaky Gut: Effect of Dietary Fiber and Fats on Microbiome and Intestinal Barrier. International Journal of Molecular Sciences. 2021; 22(14):7613. https://doi.org/10.3390/ijms22147613

Chicago/Turabian Style

Usuda, Haruki, Takayuki Okamoto, and Koichiro Wada. 2021. "Leaky Gut: Effect of Dietary Fiber and Fats on Microbiome and Intestinal Barrier" International Journal of Molecular Sciences 22, no. 14: 7613. https://doi.org/10.3390/ijms22147613

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