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Review

What We Know So Far about the Metabolite-Mediated Microbiota-Intestinal Immunity Dialogue and How to Hear the Sound of This Crosstalk

1
Faculty of Medicine, CNRS, Grenoble INP, CHU Grenoble-Alpes, University Grenoble Alpes, TIMC (UMR5525), 38000 Grenoble, France
2
Service de Biochimie Biologie Moléculaire Toxicologie Environnementale, UM Biochimie des Enzymes et des Protéines, Institut de Biologie et Pathologie, CHU Grenoble-Alpes, 38000 Grenoble, France
3
Plateforme de Métabolomique GEMELI-GExiM, Institut de Biologie et Pathologie, CHU Grenoble-Alpes, 38000 Grenoble, France
4
Department of Infectiology-Pneumology, CHU Grenoble-Alpes, 38000 Grenoble, France
*
Authors to whom correspondence should be addressed.
Co-first authors.
Co-last authors.
Metabolites 2021, 11(6), 406; https://doi.org/10.3390/metabo11060406
Received: 17 May 2021 / Revised: 15 June 2021 / Accepted: 16 June 2021 / Published: 21 June 2021
(This article belongs to the Special Issue Reviews and Advances in Microbial Metabolomics)

Abstract

:
Trillions of microorganisms, termed the “microbiota”, reside in the mammalian gastrointestinal tract, and collectively participate in regulating the host phenotype. It is now clear that the gut microbiota, metabolites, and intestinal immune function are correlated, and that alterations of the complex and dynamic host-microbiota interactions can have deep consequences for host health. However, the mechanisms by which the immune system regulates the microbiota and by which the microbiota shapes host immunity are still not fully understood. This article discusses the contribution of metabolites in the crosstalk between gut microbiota and immune cells. The identification of key metabolites having a causal effect on immune responses and of the mechanisms involved can contribute to a deeper insight into host-microorganism relationships. This will allow a better understanding of the correlation between dysbiosis, microbial-based dysmetabolism, and pathogenesis, thus creating opportunities to develop microbiota-based therapeutics to improve human health. In particular, we systematically review the role of soluble and membrane-bound microbial metabolites in modulating host immunity in the gut, and of immune cells-derived metabolites affecting the microbiota, while discussing evidence of the bidirectional impact of this crosstalk. Furthermore, we discuss the potential strategies to hear the sound of such metabolite-mediated crosstalk.

Graphical Abstract

1. Introduction

The human body is in a complex equilibrium with its microbial flora, and their synergistic interactions have been and still are the object of intense investigation. In 1991, Dr. Lynn Margulis introduced the concept of “holobiont” in her book” Symbiosis as a Source of Evolutionary Innovation”; according to this definition, the human body can be described as an assemblage of the host and the many species living in or around it [1]. The components of a holobiont are individual species or bionts, while the genomes of these bionts, including the human genome, the virome, the mycobiome, the microbiome, together define the hologenome. For decades, the development of genomic approaches, including phylogenetic marker-based microbiome profiling and shotgun metagenomics, has enabled a descriptive characterization of the microbiota composition and numerous links between its composition and diseases [2,3]. In adulthood, more than 1000 bacterial species have been identified, which shows the complexity of this large microbial community. Metagenomics gives access to the characterization of the microbiota at the taxonomic level, and at the level of the putative functions encoded by the numerous microbial genes. The recent advent of systems biology makes it possible to consider functional analysis (also simply called function) of the microbial community. This means, in particular, to quantify the metabolic activity, thanks to the measurement of RNA, by metatranscriptomics [4], of proteins, by metaproteomics [5], and finally, more recently, of metabolites, by metabolomics. Although some of the tools necessary for these integrative studies are still under development, they are essential to better understand the molecular mechanisms involved in symbiosis, but also related to the emergence of disease in the case of dysbiosis.
The human immune system plays an essential role in the development and education of innate and adaptive immunity after birth [6], and in maintaining homeostasis between humans and their resident microbial communities, thereby ensuring that the symbiotic nature of the host-microbe relationship is maintained. At the same time, commensal bacteria profoundly shape immunity [7,8,9]. The functions of the immune system could even include promoting the growth of beneficial microbes, as well as limiting the growth of harmful microbes, since the same microbe could be harmful or beneficial depending on the context (body sites, physiological state of the host, etc.). The crosstalk between immune cells and the microbiota engages a bidirectional communication, which has been studied for many years. Pattern Recognition receptors, especially Toll-like receptors (TLRs), initially described to recognize microbial signals during infection, are now known to be important players in shaping the gut microbiota and triggering a situation-specific immune response [9]. Recently, researchers have proposed that the interaction between gut microbiota and immunity is highly diverse and dynamic, and have demonstrated the cornerstone role that metabolites play in the evolutionary connection between symbiotic microbes and their hosts [10]. It is therefore important to study the content of gut metabolites and try to better characterize this dialogue (who are the players involved in the production of these metabolites, under what circumstances, and with which effects). According to Goodacre, “Metabolomics experiments aim to quantify all metabolites in a cellular system (cell or tissue) in defined states and at different times, so that the dynamics of any biotic, abiotic or genetic perturbation can be accurately assessed” and he positions metabolomics as an important systems biology approach to study the concepts of supraorganism or holobiont [11]. Despite the difficulty, metabolomics studies have clarified the metabolic profiles in the gut lumen and proved that the gut microbiome strongly influences the metabolome of the colonic lumen [12]. Microbiota-derived metabolites have been identified as components of the human metabolome [13]; some of these are recognized by immune cells via membrane or nuclear receptors and trigger a cascade of events leading to the maintenance of tolerance in the gut or the activation of antimicrobial strategies. These metabolites have a local effect in the gut on intestinal epithelial cells (IEC) or immune cells after diffusion, or are released systemically, impacting many host biological functions [13].
In a healthy state, the gut is a compartmentalized system that separates microbes from the host. Control of the microbiota biomass in the gut lumen relies on the integrity of the epithelial barrier and the effectiveness of the immune system in eliminating incoming pathogens penetrating the lamina propria. However, in this apparently hermetic system, there is constant communication between each of the players. A defective interaction between them is implicated in the pathogenesis, development, or severity of several diseases.
The present contribution attempts to provide a state-of-the-art description of the bidirectional interactions between the gut microbiota and the immune system and focuses on metabolites, defined as small organic molecules, intermediates, and products of an organism’s metabolism (<1.5 kDa). First, we review the most compelling microbiota-related metabolites associated with local or systemic immune modulations and the immune pathways that are triggered in the gut. Next, we present the traditional strategies used by the immune system to control the biomass and composition of the microbiota. Finally, we discuss the challenges and perspectives of metabolomics strategies and integrative tools in studying the crosstalk between immune cells and microbiota.

2. Microbiota-Derived Metabolites That Modulate Host Immunity in the Gut

The number of metabolites in the gut lumen is currently unknown and includes molecules of dietary, host, and microbial origin. Microbiota-derived metabolites that stimulate or suppress the immune response are either soluble bioactive molecules or membrane-bound compounds. It is already well-known that “immunometabolites”, such as succinate, itaconate, acetyl-CoA, and 2-hydroxyglutarate, to name a few, serve as signal transducers to regulate the immune cell function and disease outcome. Here, we discuss the role of the main gut immunometabolites currently known, and we provide a brief description of the classification, function, and mechanism by which they are involved in immune dysregulation (see the Supplementary Table S1).

2.1. Soluble Microbially-Derived Metabolites Affecting the Immune System

2.1.1. Short Chain Fatty Acids (Propionate, Butyrate, and Acetate)

The most well-known microbiota-related metabolites with immunomodulatory properties are the so-called “short-chain fatty acids” (SCFA), including propionate, butyrate, and acetate. Their production is the result of anaerobic bacterial fermentation of dietary fibers within the intestine and especially within the colon [14]. SCFAs are mainly produced by Bacteroidetes and Firmicutes phyla, in particular, acetate and propionate from the former and butyrate from the latter [14,15]. SCFAs average concentrations are 70–140 mM in the proximal colon versus 20–70 mM in the distal colon, and vary according to the presence of infection or inflammation [14,15,16] (Figure 1). These variations highlight the significant and rapid absorption of SCFA by colonocytes after their production via passive diffusion or via some transporters, such as MCT (Monocarboxylate Transporter) and SMCT1 (Sodium-coupled Monocarboxylate Transporter 1) [17]. Their primary role after absorption is to serve as an energy substrate for ATP generation in the tricarboxylic acid cycle (TCA), especially for the colonocytes [18]. Hepatocytes are the second main consumers, and only a small part of SCFA reach the systemic circulation, especially for propionate and butyrate [19]. Indeed, the blood concentrations are 25–250 µM for acetate, 1.4–13.4 µM for propionate, and 0.5–14.2 µM for butyrate [20]. At a cellular level, they exert a major role on gene expression through HDAC (Histone DeAcetylase) inhibition and HAT activation (Histone acetyltransferase) [21,22]. SCFA signaling through GPCRs (G protein-coupled receptors), especially three types of GPCRs: FFAR2 (GPR43), FFAR3 (GPR41), and the niacin/butyrate Receptor (GPR109A), which affect chemotaxis, apoptosis, proliferation, and cell differentiation [23,24,25,26,27]. SCFAs are necessary for intestinal homeostasis and for IECs growth [28]. They are also critical for maintaining intestinal barrier integrity through hypoxia-inducible factor 1 alpha (Hif1-α) stabilization [29]. SCFAs can lead to opposite effects on intestinal stem cell growth, depending on their concentration. Lower butyrate concentration in intestinal villi crypts compared to apex appears to inhibit the proliferation of intestinal stem cells through the activation of the transcription factor: FOXO3 [28].
Among SCFAs, butyrate is the main immunomodulatory metabolite, displaying immunosuppressive properties [30]. At the level of adaptive immunity, butyrate promotes regulatory T-cells (Tregs) and IL-22 production through several mechanisms [31,32]. First, by acting as an HDAC inhibitor, butyrate increases FoxP3 expression [33,34]. It also activates indoleamine 2,3 dioxygenase 1 (IDO-1) (that leads to tryptophan deprivation and promotes the production of the immunosuppressive metabolites kynurenine, among others), and aldehyde dehydrogenase (ALDH)1A2 (which is involved in acid retinoic metabolism), two enzymes promoting the conversion of naïve T-cells into FoxP3+ Tregs [33]. Finally, it suppresses the conversion of naïve T-cells into proinflammatory interferon-γ (IFN-γ)-producing cells [33]. It also inhibits Dendritic Cell (DCs) activation/maturation via GPR109A. Butyrate-induced tolerogenic DCs further contribute to immunosuppression through IL-10 production, generation of Tregs, and decrease in T helper (Th)17 [35,36,37]. At the level of innate immunity, butyrate promotes macrophage differentiation, and antimicrobial activity through HDAC3 inhibition that leads to increased ROS production. Hence, butyrate promotes through macrophages the reduction of bacterial translocation, contributing to prevent gut inflammation [38,39]. Due to their antiinflammatory and HDAC inhibitory properties, SCFAs are critical for health. For instance, their properties are implicated in Inflammatory Bowel Diseases, especially in Colitis [40]. Finally, in certain pathophysiological conditions, SCFA mediated immunomodulation is not restricted only to antiinflammatory properties, but it also depends on their concentration, cell type, and metabolic state. Several studies have described the immunostimulatory properties of SCFAs [41]. SCFAs have been shown to increase IL-18 production by IECs, as well as proinflammatory cytokines and chemokines, such as TNF-α (Tumor Necrosis Factor α), IL-6, CXCL1 (C-X-C Motif Chemokine Ligand 1), and CXCL10 by colon epithelial cells in vitro [42,43] (Figure 1). SCFAs improve even the generation of Th1 and Th17 T lymphocytes in an infectious context [44].

2.1.2. Lactate

In addition to the common SCFAs, lactate, which is a hydroxy-carboxylic acid, is derived from food and easily produced by intestinal lactic acid bacteria (LAB), bifidobacteria, and other anaerobes [45,46,47] (Figure 1). In order to stabilize the pH and to preserve the balance between the different microbial populations and metabolism of the colonic microbiota, the concentration of lactic acid is closely regulated in the gastrointestinal tract by two mechanisms. The first one is the cycle of production and consumption of lactate, which is used as an energy substrate by other microbiota bacteria. The second one involves the metabolization of lactic acid into SCFAs, especially propionate and butyrate, which contributes to decrease its concentration in the intestinal lumen. Lactate concentration is ranging from 5 to 10 mM in the intestinal lumen and increases in patients suffering from gastrointestinal disorders [48,49]. Microbial communities with low numbers of lactate-utilizing bacteria are inherently less stable, and therefore, more prone to lactate-induced perturbations [50]. Interestingly, on innate immunity, lactate acts as a signaling molecule on GPR81, a cell-surface G-protein-coupled receptor. The GPCR-mediated signaling of metabolites is not covered in the present review, as it has already been extensively reviewed in [51]. GPR81-mediated signaling in colonic DCs and macrophages plays an important role in suppressing colonic inflammation [52]. Lactate also induces downregulation of myeloid cell activation mediated by Lipopolysaccharide (LPS) and leads to a decrease in the production of TNF-α and IL-6 [53]. Moreover, it promotes M2 polarization of macrophages producing a high level of IL-10 and a decrease in the production of IL-12 [54]. Furthermore, Lactate is internalized into the cell by transmembrane proteins, such as MCT-1/4, SLC5A12, and SCL5A8 (subtypes of SMCT channel), and it modulates the gene expression induced through the inhibition of HDAC activity (Figure 1). Lactate is also responsible for metabolic reprogramming, inducing inhibition of glycolysis [55]. In addition to promoting an immunopermissive environment by targeting immune cells present in lamina propria, lactate plays a critical role in regulating IECs inflammatory activity. It inhibits the activation of IECs depending on TLRs and IL-1β. On adaptive immunity, lactate has been shown to decrease the motility of both CD4+ and CD8+ T-cells and the cytolytic activity of CD8+ T-cells [56]. Lactic acid also decreases the proliferation and effective functions of effector T (Teff) cells, and at the same time, increases the conversion of naive to Tregs [57]. Moreover, lactic acid has been implicated in the fight against the development and the aggravation of some pathologies. Lactic acid produced by lactobacilli has shown to be essential in regulating inflammation-induced during injury of the small intestine by indomethacin. A metabolomics study has been shown a difference in the concentration of some metabolites in the blood of patients suffering from celiac disease, and notably an increase of the concentration of lactate, which highlights an alteration of the energy metabolism [58]. Lactate may rise to high concentrations (90 mM) in the colonic lumen of patients suffering from Ulcerative Colitis (UC) [48]. In line with this, it has been shown that the presence of lactate triggers Rorc and Il17 expression by naïve and polarized CD4+ T-cells upon in vitro activation. These data suggest that high lactate concentration might promote IL-17-driven inflammation in those diseases.

2.1.3. Succinate

Similar to lactate, succinate, a dicarboxylic acid, is produced in large amounts during bacterial fermentation of dietary fibers [15]. Succinate is a metabolic intermediate of the tricarboxylic acid (TCA) cycle; therefore, it is produced by both bacteria and immune cells. However, as Germ-free (GF) mice have a low amount of succinate in the gut lumen, microbes seem to be the principal producers within the gut [59]. Succinate concentration is very low, ranging from 1 to 3 mM in the intestinal lumen and feces [60]. This is mainly explained by the succinate metabolization into SCFAs, particularly for the production of propionate. Several studies have highlighted the role of succinate as a proinflammatory mediator. In particular, the expression of succinate receptor 1 (SUCNR1, also known as GPR91) on macrophages is associated with an M1 phenotype and a type I response upon stimulation [61]. Upon activation, macrophages change their metabolism to produce and secrete a significant amount of succinate. They simultaneously upregulate SUCNR1, which acts as an autocrine or paracrine sensor for extracellular succinate to subsequently enhance IL-1β production via Hif-1α activation [62]. Hif-1α is a transcription factor that senses and responds to the metabolic state of the cell, acting as a key mediator in mounting both innate and adaptive immune responses [63]. Extracellular succinate also triggers the SUCNR1 receptor expression on DCs, acts as a chemotactic factor, and participates in their activation via TLR3 and TLR7 [64,65]. Finally, it is also reported that succinate is a potent activator of intestinal tuft cells and leads to the activation of type 2 innate lymphoid cells in the lamina propria and the remodeling of the small intestine [66,67,68]. A dysbiosis, including the reduction of succinate-consuming bacteria, can lead to both succinate accumulation and lower SCFAs production within the gut lumen, as observed mainly in patients with a high-fat diet or after antibiotic treatment [69]. Succinate accumulation seems to play a role in acute gut inflammation and may participate in the maintenance of inflammatory bowel disease (IBD) [61,70]. A recent review gathered evidence suggesting that dysbiosis and the imbalance in the SCFA/succinate ratio is one of the causes of IBD [71].

2.1.4. B Vitamins

B vitamins (VB) form a group of hydrosoluble micronutrients counting eight members. Interestingly, though dietary intake is the principal source, the microbiota plays a large part in the daily intake of VB, either by modifying dietary precursors or by de novo synthesis. Most of these modifications cannot be performed by the mammalian host. Genes related to enzymes involved in VB synthesis are scattered across bacterial and yeast genomes [72]. Early genomic research for these pathways in the digestive tract found that half of the bacteria lacks at least one gene for their synthesis [73]. Producing an accessible and active form of VB involves multiple actors among the microbiota, to finally produce the VB benefiting the host [74]. Thus, VB represents an explicit example of a symbiosis both within a polymicrobial community [75], and between the microbiota and the host. As the links between microbes and VB become clearer, the synthesis of each vitamin can now be related to specific bacteria. VB-producing strains evolve through life; for instance, in early childhood, VB2, VB6, and VB9 producing bacteria strains predominate, whereas VB1 and VB5 producing strains are more prevalent in older ages [76]. The discovery of conserved microbial phenotypes in human populations, known as enterotypes, has also led to the findings that enterotypes are specialized in the production of some VB rather than others, without further mechanistic explanation [77,78]. All VB are enzyme cofactors involved in various pathways, ranging from fatty acid metabolism to DNA repair, and their deficiency leads to serious systemic manifestations, which, for most of them, led to their discovery. The indisputable influence of such vitamins on immunity has been extensively reviewed elsewhere [79], but we briefly summarize here the main described VB effects on immunity reviewing the most up-to-date articles (Figure 2).

Vitamin B1

Thiamine (VB1) is one of the first VB discovered, due to the particularly notable symptoms of its deficiency, such as Beriberi disease or organ-related diseases, mostly in the central nervous system, as seen in Wernicke’s encephalopathy or optic nerve disorders. VB1 participates in aerobic cellular energy production through amino-acids and alpha keto-acids catabolism, and its derivative, thiamine pyrophosphate (TPP), acts as a cofactor in pyruvate dehydrogenase and the TCA cycle. VB1 comes from diet, and microflora in the large intestine is easily internalized thanks to high-affinity thiamine transporter-1 and -2 (SLC19A2 and SLC19A3) [80]. The TPP, the active form of VB1, which is transformed by the gut bacteria, is an important cofactor for enzymes, such as pyruvate dehydrogenase, alpha-ketoglutarate dehydrogenase, and transketolase [81]. Finally, many factors influence thiamine absorption, such as alcoholism and digestive pathogens, but also multiple other factors independent of the microbiota [82,83]. From the immunological standpoint, VB1 plasmatic concentrations seem to be inversely linked to immune cell counts (Figure 2). Malignant lymphocytic proliferations have been correlated to a higher risk of VB1 deficiency, probably due to intense VB1 consumption [84]. Inversely, patients with low CD4+ T-cells in HIV seem to have higher VB1 blood concentrations, even in the early stages of the disease [85], suggesting VB1 consumption requirement for lymphocyte production, activation, or survival, as demonstrated in other animals [86].

Vitamin B2

Riboflavin (VB2) is the precursor for two coenzymes, the flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD), that are required, among others, in the metabolism of niacin (VB3) and pyridoxine (VB6), respectively. VB2 is naturally accessible from a wide variety of foods (e.g., eggs, organ meats, lean meats, and milk are particularly rich in riboflavin). Its production, however, is enhanced by LAB residing in the large intestine, where the VB2 is efficiently absorbed by colonocytes. Riboflavin blood levels correlate with LAB quantity [87], with microbiota diversity and its enterotype [88], as well as the host’s nutritional status [89]. It was shown that VB2 supplementation in mice led to a decrease in proinflammatory cytokines under septic conditions and obesity-associated inflammatory states [90], notably by decreasing TNF-α production by macrophages (Figure 2). VB2 also enhances the macrophage’s production of reactive oxygen species (ROS) and their phagocytic activity [91]. These effects could be a result of NF-κB pathway tampering, or secondary to the proteasome inhibitory properties of VB2 [92]. In addition, human mucosal-associated invariant T (MAIT) cell receptors (TCRs) recognize bacterial riboflavin pathway metabolites through the MHC class 1-related molecule MR1 [93]. MAIT cells kill host cells infected with bacteria and yeast, and secrete soluble mediators, such as TNF-α, IFN-γ, IL-17, etc. The fluctuations of MAIT, both in blood samples and intestinal lumen, are actively studied in the field of IBD [94]. In human diseases, a three week riboflavin supplementation in Crohn patients is beneficial on clinical symptoms, systemic inflammation, and improved the composition of the gut microbiota, underlying its therapeutic potential [95,96].

Vitamin B3

Niacin (VB3) is the precursor of nicotinamide adenine dinucleotide (NAD), a crucial electron transporter in central metabolism, oxidative stress control, mitochondrial homeostasis, and senescence regulation. Interestingly, NAD is metabolized through different possible pathways, which often imply transkingdomal cooperation [97], sometimes relying on VB3 intake and sometimes on the kynurenine pathway [98,99]. VB3 directly activates GPCR109A, which also recognizes butyrate, leading to similar effects, notably on lipid regulation. Reintroduction of Akkermansia muciniphila in GF mice partially restored the lack of nicotinamide-producing species [100]. Another study suggests the involvement of Methylobacteriaceae in VB3 synthesis [101]. VB3 and derivatives are known to participate in macrophage polarization, towards proinflammatory or unconventional phenotypes depending on the stimuli [102]. VB3 also tampers the reactive oxygenated species production both by macrophages and neutrophils, increasing survival in LPS induced sepsis models in mice [103], along with the neutrophils’ myeloperoxidase production and recruitment [104,105]. In preclinical models of colonic inflammation, activation of GPCR109A seems to correlate with the abundance of Treg in the gut lamina propria and the production of IL-10 [37], and with intestinal innate lymphoid cells polarization [106] (Figure 2).

Vitamin B5

When considering pantothenic acid (VB5), evidence is scarcer. Because of its wide distribution in nutritional sources, deficiency in pantothenic acid is very uncommon. However, its relevance in cell metabolism is indisputable, as a precursor of the coenzyme CoA and as a catalyzer for TCA mediated oxidation reactions in fatty acids metabolism. Pantothenic acid has been linked to inflammatory homeostasis. Firstly, metabolic pathways allowing this vitamin to be synthesized into coenzyme CoA are strongly expressed in white blood cells, and the tampering of these pathways leads to unsolicited inflammatory reactions. Panthotein metabolism is relevant to cellular adhesion and polynuclear efficacy, and oxidative stress control, among others [107]. In both isolated macrophages and infected mice, supplementation of VB5 helped clearance of Mycobacterium tuberculosis and promoted antibacterial cytokine production [108]. Finally, Ghosal et al. developed a murine model with a knockout (KO) for the gene coding for the intestinal transporter of VB5 and biotin (VB8) (SLC5A6) and shown histopathological alterations associated with a high mucosal inflammation of the colon of those KO mice [109]. This intestinal inflammation was reduced by supplementing the lacking vitamins to the mice’s diets [110].

Vitamin B6

Pyridoxine (VB6) and its active derivatives, pyridoxal, and pyridoxamine, assist as cofactors to various reactions (e.g., pyridoxal 5′phosphate in amino acid metabolism, but also carbohydrate and lipid metabolic pathways, kynurenine metabolism, and to VB3 synthesis). Isolated VB6 deficiency is rare, frequently associated with other deficiencies, and it manifests itself by a large array of nonspecific symptoms [111]. As with other VB, the different metabolic pathways required to synthesize the bioactive forms of VB6 are shared by the microbiota [112]. The most notable pathways can be, for example, found between Bacillus subtilis and in Escherichia coli [113]. Studies on VB6 supplementation in human patients without underlying deficiency failed to strongly link the flora and the VB6 uptake [114]. In carps, a VB6 depleted diet accentuated the transcription of proinflammatory cytokines in all segments of the digestive tract [115]. In patients with rheumatoid arthritis (RA), inflammatory biomarkers, such as TNF-α or C-Reactive Protein, were inversely correlated to VB6 level in sera [116], but again VB6 medication in non-deficient subjects showed no beneficial effect [117]. A similar observation was made in patients with critical illness [118,119]. Curiously, in murine models of IBD with a KO in IL-10, both deficiency and supplementation of VB6 had beneficial results on gut inflammation [120]. VB6 is likely to be involved in regulating the NF-κB pathway [121], and lymphocyte polarization in favor of a Th1 mediated immune response, rather than Th2 [122,123,124]. However, the participation of VB6 derivatives in the kynurenine and sphingosine pathways complicates the understanding of its genuine implications in human health and microbiota. At this time, it seems possible that VB6 is required for a stable, non-inflammatory baseline state, and perhaps to limit the cytokine storm in COVID-19 [125], but further evidence is needed.

Vitamin B8

Biotin (VB8) is the vitamin of the B group with the most admitted relevance in cellular health. It catalyzes up to six vital carboxylases involved in numerous roles, ranging from fatty acids to amino-acid metabolism [126]. The understanding of the role of VB8 on immune health, and by extension, the consequences of VB8 deficiency or supplementation, is yet still beginning and has been reviewed elsewhere [127]. Notably, its possible involvement in the NF-κB pathway regulation holds high expectations in a potential antiinflammatory process. Recently, this hypothesis has been strengthened by Skupsky et al., which showed a lowered NF-κB activity and histological signs of mucosal inflammation in the colon of biotin-supplemented mice [128]. This observation concurs with a previously mentioned study, exploring the supplementation of VB5 and VB8 [110]. Another team focused on the properties of VB8 on human T-cell population, and showed that VB8 deficient growth media promoted higher titers of proinflammatory cytokines, such as IFN-γ and IL-17, enhanced the T-cell polarization towards Th1 and Th17 proinflammatory profiles, and reduced the proportion of Treg-cells through the rapamycin-mTOR pathway [129].

Vitamin B9 and B12

Folate (VB9) and cyanocobalamin (VB12) are often studied together. In a human randomized trial, dietary uptake of probiotic species (an association of Bifidobacter spp. and lactobacillus spp.) was associated with higher blood titers of VB9 and VB12, without any additional supplementation of these vitamins [130]. Folate is an essential cofactor for synthesizing amino acids, nitrogenous bases, and ribonucleoside, which is why antifolates are commonly used in cytotoxic chemotherapies and antiinfectious treatments. It is also involved in mitochondrial RNA metabolism [131]. VB12 also catalyzes the synthesis of amino acids, both from the cell and the mitochondria [132]. Evidence of bacterial involvement in their synthesis and absorption is indisputable since their full metabolic pathways have been elucidated [133,134,135,136,137]. In immune homeostasis, though both vitamins are required for hematopoiesis, the role of VB9 is better understood. Lower titers of VB12 seem to correlate with higher inflammation levels in critically ill patients [138], and were associated with higher mortality in hospitalized patients [139], but the vitamin’s direct involvement remains unclear. VB9 titers, on the other hand, are strongly correlated with leukocyte counts [140]. VB9 deficiency induces DNA damage in lymphocyte precursors, similarly to those observed in radiation-induced injuries [141]. In allergic children, excessive levels of VB9 are also proportional to allergic symptoms, and inversely proportionate to circulating regulatory T lymphocytes levels [142]. Folate-associated receptors are found on mature immune cells, e.g., naive T lymphocytes, and might be involved in their activation or regulation [143]. Similar receptors on the surface of macrophages are overexpressed under certain conditions, such as hyperlipidemia, and lead to activation of macrophages in a proinflammatory profile, promoting, for example, atherosclerosis [144]. These receptors are targets for potential antiinflammatory drugs, notably for the control of chronic inflammatory states [145] and chronic IBD [146].

2.1.5. Amino Acids (AA) and AA-Derived Metabolites

Tryptophan

Tryptophan (Trp) is an essential amino acid mainly supplied by the diet. The majority of Trp is absorbed in the small intestine. A small portion can reach the colon, where it can be used by commensal bacteria. In humans, three Trp pathways are generally described: The kynurenine pathway, the indole pathway, and the serotonin pathway. Serotonin (also known as hydroxytryptamine or 5-HT) can modulate the immune response, and hence, potentially influence intestinal inflammation [147]. It has been demonstrated that GF mice increased plasma serotonin concentrations, which can be normalized following colonization of the mice immediately post-weaning [148]. Several of the 5-HT receptors have been associated with immune cells, such as lymphocytes, monocytes, macrophages, and DCs, which indicates that 5-HT plays an immune-modulatory role. Tryptamine concentrations increase nearly 200 fold in feces following colonization of GF mice with human gut microbiota, suggesting that bacterial metabolism of Trp generates luminal tryptamine [149]. The enzyme tryptophan decarboxylase (TDC1) is responsible for converting Trp to tryptamine, which is converted to serotonin by tryptamine 5-hydroxylase. The genes encoding the homologs of this TDC1 were found in at least 10% of the representative human gut microbiota [150]. Two comprehensive reviews have already discussed the link between the gut microbiota regulation of Trp and its impact on health and disease [151,152]. In addition to tryptamine, other metabolites, like indole derivatives or skatoles, are derived from the catabolism of Trp by the intestinal microbiota. Those are described in the review [153]. They have been intensively studied for their role in immunity, mainly through the activation of the aryl hydrocarbon receptor (AhR). In addition to this activity, indole is now studied for its ability to influence the gene transcription in some immune cell subsets and epithelial cells. Indeed, Microbiota-derived AhR ligands play a critical role in gut homeostasis; for instance, they have been associated with the increase of Il22 transcript in type 3 innate lymphoid cells or ILC3 [154], and subsequent mucosal integrity. Trp-derived indoles also activate the pregnane X receptor (PXR) to support the barrier function in colitis mouse models [155]. One example is indole 3-propionic acid (IPA) produced by Clostridium sporogenes. This metabolite is absent in GF mice. The ability of C. sporogenes to produce IPA from Trp was recently described and is limited to a small group of organisms sharing the proper metabolic pathway [156]. GF-mice colonization with an IPA-producing strain like C. sporogenes restores the IPA serum level [13]. It was shown to activate the AhR and the PXR pathways leading to the reduction of the symptoms in a mice-dextran sodium sulfate (DSS) model of colitis [157,158]. Thus, IPA might be an interesting candidate for the treatment of UC. Other indoles, such as Indole Acrylic Acid (IA) [159] and Indole-3-aldehyde [154], are described in the literature for their effects on immune cells (Figure 3). Microbial metabolism of Trp resulting in indole derivatives, such as indole acetic acid and IPA, has been recently investigated in several studies for their contributions to host physiology. In the future, Trp-derived metabolites should be considered as a promising strategy that can be used to treat human diseases [160].

Taurine

Taurine is a sulfur-containing amino acid that can be sourced from the diet or synthesized by the pancreas via the cysteine sulfinic acid pathway. Taurine is not per se a metabolite produced by bacteria. However, as described below, taurine reaches the colon in a conjugated form with bile acids (Bas). Its release in the colon depends directly on microbiota activity that can result in taurine deconjugation via bile salt hydrolase (BSH) activity. Thus, it can be considered as an indirect product of the colonic microbes. Taurine is known to be involved in many physiological events, such as osmoregulation, membrane stabilization, calcium mobilization, neurotransmission, reproduction, and detoxification [161,162,163], along with providing antiinflammatory effects and protection to cells from cytotoxic effects of inflammation. During Bas synthesis, glycine is more often used than taurine (ratio of 3:1) [164]. However, this ratio is subject to variation since the availability of taurine is diet-dependent (a western diet will favorize a taurine conjugation when vegetarians are using glycine [165,166]. In the Dexamethasone-induced immunosuppressive mice model, taurine was involved in the increase of lymphocytes in Peyer’s patches, a well-known lymphoid follicle in the intestine classified as a gut-associated lymphoid tissue [167]. Similarly, changes in the microbiota were observed after treating the mice with taurine. These groups showed a modification of the pattern, suggesting that some beneficial bacteria, such as Lachnospiraceae and Ruminococcaceae, groups were significantly re-increased after treatment with taurine. The role of taurine on gut epithelial cells was also presented by Levy et al. and reported as an activator of the NLRP6 inflammasome leading to the activation of IL-18 and the production of antimicrobial peptides (AMPs) [168].

p-Cresol and Its Derivatives

p-Cresol is a methyl phenol produced by colonic fermentation of tyrosine and phenylalanine. After its production, most of the p-cresol (approximately 80%) in the intestinal wall is conjugated by the intestinal flora into p-cresyl sulfate (pCS) or p-cresyl glucuronidate. The association between the microbiota and p-cresol or its derivatives has been proven, for example, in GF-mice models, in which p-cresol or its derivatives were not found in the blood [13]. In humans, pCS is considered a uremic solute, and patients with chronic kidney failure display high blood pCS levels. However, among these patients, those who are also colonectomized normal display levels of pCS, demonstrating the colonic origin of pCS synthesis [169]. Culture-based tests have identified several intestinal bacteria as producers of p-cresol, such as those belonging to the Coriobacteriaceae and Clostridium clusters XI and XIVa [170]. A comprehensive overview of bacterial species able to produce phenolic compounds has been reviewed elsewhere [171]. Derivatives of p-cresol are considered as uremic toxins, involved in uremic syndromes developed during renal failure progression. The immunomodulatory effect of p-cresol and derivatives has been highlighted in chronic kidney disease (CKD) patients that are highly susceptible to infectious diseases [172]. The most studied p-cresol derivative is pCS, for its role on both innate and adaptive immunity. In an in vitro study, it was found to induce ROS production and phagocytosis at lower concentrations, while at higher concentrations, it was shown to interfere with antigen processing in human monocyte-derived macrophages [173]. Moreover, in a mouse model of adenine-induced renal dysfunction, pCS was found to be involved in immune dysfunction in CKD. It decreased peripheral B lymphocytes number by inhibiting the proliferation of CD43+ B-cell progenitor [174]. The role of pCS on Th1-type immune responses has been studied through a tyrosine-enriched diet mouse model. pCS was negatively correlated with the percentage of IFN-γ-producing Th1 cells during a 2, 4-dinitrofluorobenzen-induced contact hypersensitivity. In vitro assays on splenocytes exposed to a variable concentration of pCS suggested that intestinal-derived pCS suppresses the percentage of IFN-γ-producing Th1 cells and favors a Th2 response [175]. Finally, while the role of p-cresol derivatives has been increasingly characterized, the exact effect of p-cresol itself is still controversial [176]. Even if CKD is a good study model, it may be interesting to understand if pCS can influence the immune system in a healthy state, as p-cresol derivatives are constantly eliminated in urine through tubular secretion.

Histidine and Derivatives

Gut bacteria can convert amino acids, such as L-histidine, into biogenic amines (i.e., molecules containing one or more amino groups), such as histamine, by the action of histamine decarboxylase (HDC) [177]. Numerous immune cells produce histamine, mainly basophilic cells, and mast cells, and to a lesser extent, monocytes, DC, and lymphocytes [178]. Histamine production by basophils and mast cells, is modulated by cytokines, such as IL-3, IL-12, IL-18, TNF-α [179]. Histamine is mainly produced within the caecum, and found at lower levels in the wall of the small intestine in GF mice, compared to conventional mice [180]. Histamine, in addition to its role as a neuromediator and regulation of gastric acidity, is mainly described as a cytokine and an inflammatory mediator in acute inflammation and hypersensitivity. Recently, it has been shown that histamine produced by microbiota-associated bacteria impacts both intestinal epithelial cells and immune cells. In humans, Barcik et al. have performed a PCR analysis of bacterial HDC expression on fecal samples from 74 healthy donors, and identified E. coli, Lactobacillus vaginalis, and Morganella morganii as histamine-secreting bacteria. Interestingly, they also collected samples from 74 asthma patients, highlighting an increased bacterial HDC copy number compared to healthy donors. Furthermore, Morganella morganii relative abundance was correlated with disease severity, suggesting a possible involvement in asthma pathogenesis [181]. Histamine action is mediated by four types of receptors noted H1R to H4R, and leads to different effects, depending on the receptor subtype, its expression level, as well as the targeted cell. DCs express H1R, H2R, and H4R, receptor subtype. H1R engagement is involved in immediate hypersensitivity reaction, inducing vasodilatation, and promoting Th1 responses, while H2R activation leads to an inhibition of Th1 and Th2 responses and promotes Treg by increasing the production of IL-10 [182,183,184]. On adaptive immunity, in the same way, H1R promotes IFN-γ, the production, and the proliferation of Th1 cells, while H2R inhibits Th2 response by blocking the synthesis of IL-4 and IL-13 [178,185]. Thus, histamine affects the balance between Th1 and Th2 responses [178]. Levy et al. analyzed the immunomodulatory effects of some microbiota-associated metabolites and showed that histamine intake in drinking water led to a strong reduction of IL-18 proinflammatory cytokine production by inhibiting NLRP6 inflammasome assembly [168]. Of note, the therapeutic potential of a bioengineered histamine-overexpressing bacteria has also been investigated in an in vivo asthma model. When orally administrated, it led to a decrease of Th2 cytokines secretion by lung-derived cells, as well as proinflammatory cytokines and IL-10. These results suggest a global antiinflammatory effect, not only mediated by an inhibition of Th2 response. Thus, these data indicate that histamine-producing bacteria within the gut can potently modulate systemic host immunity [186].

Polyamines

Among biogenic amines, polyamines (Pas) are organic compounds having more than two amino groups. They have been studied for their ability to modulate cellular functions, including gene regulation, stress resistance, cell proliferation, and differentiation [187]. Spermidine (N3) and spermine (N4), two putrescine-derivatives, are the major examples of Pas in human cells. These two metabolites can be found in food or directly produced by gut microbiota or any eukaryotic cells. While the ingested food is the major source of Pas in the small intestine and is a crucial source for the stability and the maintenance of gut tissues [188], the microbiota appears to be a major player in Pas production in the lower part of the intestine, i.e., the colon [189,190,191]. Pas are extensively studied for their interaction with epithelial cells and the immune system. Interestingly, Levy et al. reported that exogenous spermine significantly reduces the activation of NLRP6 inflammasome in IECs and subsequent release of IL-18, a cytokine that promotes the production of antimicrobial peptides (AMPs), an important element involved in gut microbiota modulation [168]. Spermine also exerts antiinflammatory effects directly on immune cells by inhibiting LPS-induced expression of proinflammatory cytokines, including IL-6, TNF-α, IL-1, MIP-1α, MIP-1β, by monocytes and macrophages [192,193]. This was confirmed in an in vivo sepsis mouse model where a systemic antiinflammatory effect was observed [194]. In addition to spermine, spermidine is also of interest for its effect in age-associated diseases [195,196,197], cardioprotection [198], tumor suppression, immune modulation, neuroprotection, metabolic syndromes, and stem cell function. All of these aspects have been extensively reviewed elsewhere [199]. Recently, spermidine was shown to play a major role in regulating T-cell differentiation and function. Spermidine exposition potentiates the in vitro Foxp3 + T-cell differentiation from both naïve and Th17 CD4+T-cells, in an autophagy-dependent manner [200], and dampens the IFN-γ-mediated monocyte response [201]. Moreover, the proTreg effect of spermidine has been confirmed in an in vivo colitis mouse model [200]. A considerable amount of studies suggested that Pas have real health benefits, making them potential therapeutic candidates [202,203].

D-Amino Acids

While mammalian cells can produce only two D-amino acids (D-AA), namely, D-serine and D-aspartate [204], pathogenic and commensal bacteria can produce several D-amino acids, which play an essential role in multiple biological processes [205]. Sasabe et al. reported that the mouse intestine is rich in free D-AA produced by gut microbiota [206]. They have found the D-amino acid oxidase (DAO), which is associated with D-AA, expressed in the villous epithelium of the small intestine. The oxidative deamination of the intestinal D-AA by DAO results in the generation of hydrogen peroxide, H2O2, a strong oxidizing agent with antimicrobial activity, and leads to the modification of the human microbiota, protecting the mucosal surface of the small intestine against patens. Thus, they proposed bacterial D-AA acids and DAO as new examples of inter-kingdom communication. Beyond bacterial communication, D-amino acids display antimicrobial properties, such as ROS production, and bacteriostatic activity. They can regulate neutrophil chemotaxis and modulate immune tolerance. These properties have been comprehensively reviewed elsewhere [207].

Gamma-Aminobutyric Acid

The gut microbiota has a critical role in the production of gamma-aminobutyric acid (GABA). It is mainly produced through a one-step reaction from glutamate, catalyzed by glutamic acid decarboxylase [208]. It can also be produced by some microorganisms from putrescine [209] (see section polyamines). GABA production was described for several Lactobacillus and Bifidobacterium strains [210], and it was reported that the introduction of a GABA-producing Bifidobacterium strain was sufficient to modulate GABA levels in the gut [211]. Among them, Lact. Brevis and Bif. Dentium was those producing the highest amounts of GABA [210]. GABA produced by Lactobacillus rhamnosus JB-1 in the gut has been shown to have effects on the brain (the microbiota-gut-brain axis), and, in particular, resulting in a reduction of stress-induced corticosterone, and in depression- and anxiety-related behavior. In the context of enterotoxigenic E. coli (ETEC) infection, ETEC induces dysbiosis, increasing the GABA-producing Lactobacillus lactis subsp. lactis. The increased content of GABA in the jejunum of mice promotes intestinal IL-17 expression through a GABA-mTORC1 signaling pathway [212]. The effects of GABA on intestinal homeostasis have been described through its action on enterocytes. Firstly, it has been described as a selective stimulator of mucin-1 expression in epithelial cells [213]. Then, exposure of GABA to epithelial cells resulted in a decrease in IL-1β-mediated inflammation and an increase in tight junctions and transforming growth factor beta (TGF-β) expression, thus providing a protective effect against the disruption of the intestinal barrier [214].

Quorum Sensing Molecules

Quorum sensing molecules (QSMs) consist of signaling molecules, sensing molecules, and downstream regulatory proteins produced and used by prokaryotes to monitor population density and assist a number of biological functions, such as biofilm formation and production of virulence factors. The gut microbiota has shown to have a very stable structure despite challenges in the gut, such as stomach acid and intestinal bile; its stability depends on QSMs [215]. Generally, in Gram-Negative bacteria, quorum sensing (QS) is composed of two components: A small soluble signaling molecule (predominantly N-acyl homoserine lactone (AHL) molecules) and a transcriptional regulatory protein (R protein). In gram-positive bacteria, QS systems are generally composed of three components, a signal peptide and a two-component regulatory system (TCS) itself composed of a membrane-bound histidine kinase (HK) sensor and an intracellular response regulator (RR) [216,217]. The main QSMs are AHLs, diketopiperazines (DKPs), 4-hydroxy-2-alkylquinolines (HAQs), diffusible signal factors (DSFs), autoinducer-2 (AI-2). QSMs are products of nutrients utilization, and many of them are synthesized from metabolites present in the environment. For instance, AHLs and AI-2 are both produced from S-adenosylmethionine, which is part of the methionine metabolism. AHLs are characterized by carbon acyl chains, some of which are derived from intermediate molecules of the fatty acid biosynthesis in the host [216]. Surprisingly, some organisms use more than one QS signaling system, which means that QS is not only involved in intra-species communication, but also involved in inter-species communications. It is the case of AI-2, which is used both by Gram-positive and Gram-negative bacteria, and which is the only major type of QSM known to promote inter-species bacterial communication across distantly related bacterial species [218]. AI-2 has been shown to influence the composition of antibiotic-treated microbiota and modulate the abundance of certain phyla [219]. QSMs are also detected by immune cells and regulate the activity of both innate and adaptive immunity. For instance, bacterial autoinducers (Ais) elicit proinflammatory effects and modulate the activities of gut-associated T lymphocytes, macrophages, DCs, and neutrophils [220]. Moreover, Mast Cell activation through the mast-cell-specific GPCR, MRGPRX2, which are Gram-positive QSMs receptors [221] leads to degranulation and release of ROS, TNF-α, and Prostaglandin D2, thereby triggering a local inflammatory response. In 2019, the association between gut microbiota-derived AI-2 and the progression of colorectal cancer (CRC) has been highlighted [222]. This association was mediated by tumor-associated immune cells, such as macrophages. AI-2 was proposed as a novel biomarker for human CRC as its concentration was higher in colorectal tissue and stool of CRC groups compared with normal colonic mucosa and colorectal adenoma. Moreover, AI-2 levels increased depending on CRC staging in both tumor tissues and stool samples. A positive correlation was also found between AI-2 and Tumor necrosis factor ligand superfamily member 9 (TNFSF9), a costimulator of T-cell proliferation belonging to the TNF receptor family. This association was confirmed experimentally through the stimulation of macrophages with AI-2 extracts from F. nucleatum that showed a significant increase in TNFSF9 expression, thus confirming the link between microbial QS, inflammation, and CRC.

2.1.6. Catecholamines

Catecholamines (Cas), which include epinephrine (E, also known as adrenaline), norepinephrine (NE, also called noradrenaline), and dopamine, are a class of organic compounds characterized by a catechol structure, i.e., a benzene ring bearing two adjacent hydroxyl groups, and a side-chain amine, which contributes to receptor specificity [223]. Cas synthesis starts with dietary L-dopa as substrate, which is then enzymatically converted into dopamine, NE, and finally, E. Cas, especially NE and dopamine, are abundantly present within the human gastrointestinal tract [224]; in particular, approximately half of the NE amount in the mammalian body is produced within the pre-vertebral ganglia innervating the gut mucosa, while dopamine is synthesized in non-sympathetic enteric neurons within the intestinal wall [225,226]. It is unlikely instead to normally find E in the gut at a significant level, since the enzymes required for its conversion from NE are not expressed in the intestinal mucosa [227]. In addition to this, there is increasing evidence that the intestinal lumen Cas levels are gut microbiota-dependent [228]. While data on the magnitude of the microbial contribution to Cas levels in the gut are still scarce, because the normal NE and dopamine luminal concentrations are difficult to determine, it is instead clear that the presence of bacterially derived NE and dopamine contributes to make the gut a Cas-rich environment, together with dietary intake and enteric nervous system sources [228]. Among others [229,230], Asano and coworkers demonstrated that the gut microbiota plays a crucial role in producing biologically active free Cas in the gut lumen [231]. They showed that levels of NE and dopamine increased in specific pathogen-free mice with normal gut microbiota, when bacteria were present. The role of Cas as chemical neurotransmitters in the central and peripheral nervous systems, with a key position in regulating various physiological processes and functions, such as cognitive abilities and intestinal motility, is well-recognized [232]. In addition to this, Cas have shown to be potential inter-kingdom signaling molecules in the gut: In the 1990s, the pioneering work of Lyte et al. demonstrated that some pathogenic species could recognize exogenous Cas in vitro, leading to an increase in bacterial growth [233,234]. Later, Sperandio et al. showed for E. coli O157:H7 that the two-component regulator sensor kinase QseC is a receptor for the host E and NE [235], supporting the observation that Cas could act as inter-kingdom signals [236]. In Salmonella Typhimurium, NE has been shown to trigger the expression of virulence-associated factors, including flagella-mediated motility and Type III protein secretion [237,238,239]. Signaling between host cells and microbes through Cas continues to attract interest among the scientific community, especially as it might play a key role in microbial dysbiosis and increase susceptibility to infection by altering the growth and virulence of human pathogens [228]. However, further investigation is needed to fully elucidate the luminal CA-related functions and their mechanisms. Finally, there are many studies indicating that Cas is also important immunomodulators during health and disease [240,241,242,243]. These functions were first observed in 1904, when pronounced leukocytosis was reported following subcutaneous administration of E in humans. It is now clear that lymphocytes and phagocytes are also Cas-producing cells [244], and there is evidence that Cas exert autoregulatory functions on immune cells by means of intracellular oxidative mechanisms [245]. To conclude, the consequences of lymphocyte/phagocyte-derived or exogenously administered Cas during shock and trauma are becoming increasingly clear [244]. During severe tissue trauma, the body experiences the destruction of noradrenergic nerve cell innervation, with a consequent release of NE into the systemic circulation. Lyte and Bailey reported that this led to increased proliferation of bacteria within the gastrointestinal system of an experimental murine model, probably contributing to the high incidence of systemic bacterial inflammation and sepsis following trauma hemorrhage [246]. Metabolites shared between different cell types and the gut microbiota, such as Cas, are particularly difficult to study. Further research is needed to understand the dynamics of Cas in the gut lumen and its impact on mucosal immunity. Mono-colonization with CA-producing microbial species and well-defined in vivo models will certainly help to decipher the role of each player.

2.1.7. Cyclic-Dinucleotides (CDNs) and Cyclic-Trinucleotides (CTNs)

CDNs are small nucleic acids produced by bacteria (and viruses) that act as essential secondary messengers that help them to coordinate a new response and shape their behavior after a change in their environment. For example, cyclic di-adenosine monophosphate (cyclic di-AMP) is involved in bacterial growth, biofilm formation, stress response, and antibiotic resistance [247]. When CDNs are sensed by immune cells, this results in proinflammatory responses. Depending on CDNs types, they can activate either STING (Stimulator of interferon genes) pathway, or the oxidoreductase RECON (NF-kB controlling reductase). CDNs with two purine bases (such as c-di-GMP, 3′,3′ cGAMP, and c-di-AMP) triggers STING-mediated NF-κB activation and type-I interferon [248]. In addition, the oxidoreductase RECON can specifically bind to cGAMP, and c-di-AMP, cUMP-AMP, resulting in the inhibition of its enzymatic activity and promoting a proinflammatory antibacterial state [249]. The RECON specificity is not limited to purine bases. It can also sense cyclic-trinucleotides (CTNs), such as cAMP-AMP-GMP produced by bacteria. Some of these CTNs are only recognized by the RECON sensor, but not by STING, leading to the inhibition of the NF-κB responses [250]. In a pathological context where intracellular bacteria are involved, the ability of these bacteria to increase the diversity of their CDNs/CTNs can be considered as an evolutionary adaptation mechanism to evade host innate immunity and metabolic pathways. This area of research is still emerging, and we are certainly at the beginning of a concept.

2.1.8. Inosine

Inosine is a nucleoside formed from ribofuranose and hypoxanthine. Recently, it was shown that inosine produced by Bifidobacterium pseudolongum is able to change the immune response after an immune checkpoint blockade (ICB) therapy [251]. In their work, Mager et al. studied the difference in microbiota between CRC-positive mice challenged with conventional immunotherapy versus CRC-positive mice with no treatment. Seven bacterial species were found to be specific to ICB-treated tumors. A mono-colonization assay with five of the seven species in a heterotopic model of CRC showed that only B. pseudolongum had the most important effect on immunotherapy, and this effect was mediated by inosine, which is found in high amounts in the sera of mice mono-colonized by this bacteria. Inosine can activate the adenosine 2A receptor in immune T-cells, and drive the Th1 differentiation of costimulated T lymphocytes.

2.1.9. Secondary Bile Acids

Bile acids are hydroxylated steroids involved in digestion and lipids absorption; they are considered hormones. Currently, it is known that primary bile acids, such as cholic acid and chenodeoxycholic acid, are synthesized in the liver and secreted in the intestine in a conjugated form with glycine or taurine. 95% of them are finally reabsorbed in the ileum and recycled by the liver; this cycle is known as enterohepatic circulation. The remaining 5% activates the FXR in the liver and intestine tissues. This leads to the repression of both the CYP7A1 gene, encoding for a rate-limiting enzyme in the classic bile acid synthesis pathway, and the CYP8B1 gene, required for cholic acid synthesis [252]. The microbiota interacts with the metabolism of bile acids at different stages. First, primary bile acids are deconjugated by bacterial communities via BSH. Then, they can also be transformed by the microbiota to produce more than 20 secondary bile acids (e.g., deoxycholic acid (DCA) and lithocholic acid (LCA)). Secondary bile acids can act as soluble mediators signaling through bile acid receptors (BARs) expressed by many types of cells, such as epithelial cells and some immune cell subsets. BARs encompassed a large family of GPCRs (e.g., GPBAR1) and nuclear (e.g., FXR) receptors. BARs can recognize both primary and secondary bile acids. A given BAR can recognize several bile acids that can either trigger an activating or an inhibitory signal, depending on the bile acid and BAR involved (Reviewed in [253]). They are viewed as a negative regulator of macrophages [254], DCs, and NKT [255] functions, supporting the idea that they contribute to the maintenance of a tolerogenic environment in the liver and intestine [256]. Recently, Song et al. demonstrated that deconjugated bile acids can promote the generation of colonic RORγt-expressing FoxP3+ Tregs through Vitamin D receptor (VDR) [257]. They demonstrated that genetic abolitions of bile acids metabolic pathways (required for secondary bile acids production) in individual gut symbiont (Bacteroides thetaiotaomicron and Bacteroides fragilis) decreases this T-cell population. This effect was cell-type- and tissue-type- specific, as no similar effects were observed either on colonic Th17, nor on Tregs and Th17 from the spleen, the mesenteric lymph node, or the small intestine, respectively. Importantly, these cells have been shown to display superior suppressive capacities in T-cell-mediated intestinal inflammation [258] than conventional FoxP3+ Tregs. Recently, Campbell et al. screened several secondary bile acids and identified the 3β-hydroxydeoxycholic acid (isoDCA) as an immunosuppressive bile acid. IsoDCA alters DC-mediated T-cell stimulation, by decreasing DC costimulatory molecules expression and proinflammatory cytokine production that in turn promotes FoxP3+ Tregs generation [259].

2.2. Microbial Membrane Metabolites Affecting the Immune System

Microbes not only modulate host immunity through soluble metabolite secretion, but can also use membrane-bound metabolites. Unlike soluble metabolites, membrane-bound metabolites require more complex mechanisms to cross the epithelial gut barrier. It has been proposed that these strategies are based on outer membrane vesicle (OMV) formation or dead bacteria-derived membrane fragments. Presumably, followed by endocytosis or passive diffusion through epithelial cells, this class of metabolites then interacts with the gut-associated immune system. The entire mechanism needs to be further investigated to clearly understand how these metabolites move from the microbial membrane to immune cells (Figure 4).

2.2.1. Sphingolipids

Sphingolipids are a class of plasma-membrane-associated lipids containing a backbone of sphingosine. They are produced by both the host and specific bacteria. Host sphingolipids participate in a large and diverse range of physiologic cellular functions, as they are involved in specific signaling pathways, either acting themselves as signaling molecules or regulating downstream signaling molecule functions [260]. Within the gut, host-sphingolipid complexes, such as sphingomyelin (SM) and glycosphingolipids (GSLs), are essential components of epithelial cells, playing a role in lipid absorption, providing protection, and integrity of the mucosa. Host sphingolipids (notably ceramide-1-phosphate (C1P), sphingosine-1-phosphate (S1P), and ceramide (Cer)) have been described alternatively as anti or proinflammatory and act directly or indirectly as inflammatory mediators [261]. Along the intestine, the ratio of sphingolipids (pro/antiinflammatory) is tightly controlled, and dysregulation of this balanced ratio is observed in diseases, and notably in IBD. It has been shown that sphingolipid levels are significantly different between inflamed and non-inflamed intestinal tissues and may affect the immune gut ecosystem [262]. The production of SM-like sphingophospholipids is described in the gut from members of the Bacteroidetes phylum (e.g., Bacteroides ([263] and Prevotella [264]). Sphingolipids in sphingolipids-producer bacterial strains participate in stress resistance by preserving membrane integrity [265]. It is believed that Bacteroides-derived sphingolipids may cross the epithelium barrier and reach immune cells via OMV formation [266]. Sphingolipids are presented by antigen-presenting cells (APCs) through the CD1d receptor, an MHC class I-like receptor to CD1d-restricted innate-like lymphocytes called invariant Natural killer T (iNKT). The best known CD1d-restricted iNKT ligand is aGalCer produced by Sphingomonas spp [267]. Recently, Kinjo et al. described that Bacteroides fragilis produced natural aGalCer analogs [267]. Early exposition to microbiota-derived sphingolipids, and notably Bacteroides fragilis-derived α-Galactosylceramide GSL-Bf717, prevents excessive colonic iNKT accumulation [268], and subsequent iNKT-related inflammatory disorders in adulthood [269]. The antiinflammatory properties of Bacteroides-derived sphingolipids were revealed by Brown et al., who showed that patients with Crohn’s disease and UC had elevated levels of host-derived sphingolipids, while the abundance of Bacteroides was decreased [270]. To better understand the exact role of Bacteroides-derived sphingolipids, a GF mice mono-colonization experiment was performed with sphingolipid-deficient B. thetaiotaomicron strain and compared to the wild-type strain. The sphingolipid-deficient strain triggered intestinal inflammation, as revealed by histopathological inflammatory signatures, also characterized by an increase of IL-6 and MCP-1 proinflammatory cytokines within the colon [270]. Interestingly, Ultra-High Performance Liquid Chromatography (U-HLPC) coupled to an Exactive Plus orbitrap mass spectrometer lipidomics analysis revealed 35 unique sphingolipids derived from Bacteroides, highlighting the diversity of these molecules [270]. In conclusion, it is now clear that sphingolipids can modulate host immunity. However, depending on the producing organism, sphingolipids can differentially impact the immunity either toward inflammation or tolerance. Thus, microbial-derived sphingolipids have to be identified, isolated, and screened for their immune properties.

2.2.2. Lipoteichoic Acids

Lipoteichoic acids (LTA, from the Greek “teichos” meaning wall) are amphipathic polymeric molecules anchored onto the Gram-positive bacteria wall, which regulate several autolytic wall enzymes (muramidases) [271]. They are composed of a repeating unit of glycerophosphate or ribitolphosphate bound to a glycolipid anchor [272]. Another form of bacterial cell wall polymers are wall teichoic acids (WTA), which are instead covalently bound onto the peptidoglycan and that have been reviewed elsewhere [273]. For Gram-positive bacteria, TLR2 ligands are the primary host Toll-like receptors (TLRs) involved in the innate immune response [274]. Even if their nature is still controversial, several studies have suggested that LTA are their main activator [275,276,277]. It appears that LTA can play an immunostimulatory role, by activating in vitro the NF-κB pathways in monocytes and macrophages, in a TLR2-dependent manner [276,278,279]. Other pattern recognition receptors (PRRs), such as C-lectins or Ig-receptors and CD14, have been shown to be involved in the immune response triggered by LTA during bacterial infections [280,281]. The activation of phagocytes promotes the production of cytokines, participating in appropriate antibacterial immune response in Gram-positive bacterial infection models [282,283,284]. However, some of the most compelling evidence on probiotics revolves around the LAB. The LTA of some LAB seems to exert a particular immuno-modulatory effect [285]. In murine models, dietary Lactobacillus rhamnosus allowed better healing from intestinal radiation, and in vitro studies confirmed the involvement of its specific LTA in the protective effect [286]. Other teams have shown a diminution in proinflammatory cytokine production from macrophages when exposed to the same LAB [287]. Animal experimentations with heat-killed Lactobacillus paracasei suggest that the exposition alone to the cell wall is enough to trigger the immunomodulatory effect, along with other beneficial effects of LTA, such as an increase in mucosal production via MUC2 production [288]. Grangette et al. demonstrated the possibility to enhance its antiinflammatory capacity by modifying the composition of an already beneficial type of LTA from Lactobacillus plantarum, therefore emphasizing the role of structural variations of LTA on the modulation of the host immune response [289]. The interactions between LTA and the immune system are not yet fully understood. Different levels of expression for TLR2 as a protective mechanism or a risk factor for inappropriate inflammation, as suggested by Melmed et al. [290], and other complex crosstalks should be furthermore examined to get a grasp on the intricated interactions that developed between bacterial LTA and human immune system during our long coevolution. The structural diversity of LTA and their effects, combined with the complexity of microbiota composition, could participate in several reactions in the immune response, and thus, should be explored in appropriate samples whenever the question is relevant.

3. Metabolites Derived from Immune Cells Affecting the Microbiota

Although interactions between microbiota and host are bidirectional, most studies have mainly focused on the impact of bacteria strains and derived-metabolites onto immunity. To our knowledge, no study has yet described immune-derived “bona fide” metabolites impacting gut microbiota composition and functions. However, some studies highlighted that host immune activation triggers rapid microbiota modulation, at least at the transcriptional (and thus, functional) level. Moreover, it is now clear that the immune system acts on microbiota through several effector molecules, ranging from antimicrobial peptides to soluble Immunoglobulin A (sIgA). In order to emphasize the importance of this bidirectional dialogue, we briefly describe in this part the impact of the host immune system on gut microbiota modulation.

3.1. Gut Microbiota Modulation upon Acute and Chronic Host Immune System Activation

Most chronic inflammatory diseases are associated with dysbiotic gut microbiota. Many efforts have been made to determine whether dysbiosis was a cause or consequence of chronic inflammation. In IBD, for instance, patients display a dysbiotic profile [291,292,293]. Mice experiments revealed that in genetically identical hosts, the susceptibility to colitis relies on microbiota composition [294]. Recent studies have demonstrated that chronic inflammation also impacts microbiota at the meta-transcriptome level, in both IBD patients [295], and mouse colitis models [296,297]. Very recently, Becattini et al. assessed if gut microbiota also responds to acute (thus transient) host immune activation. In order to address this question, they used a simplified mouse model where GF mice were reconstituted with four anaerobic bacterial strains able to stably colonize the intestine [298]. Data indicates that following systemic immune activation via intraperitoneal flagellin administration (TLR5 activation), gut microbiota transcriptional and metabolic profiles rapidly change within 6 h, while the relative proportions of each strain remain stable. At 24h post immune activation, transcriptional profiles largely return to baseline. Flagellin administration triggers transcriptional modifications of hundreds of genes, upregulating genes involved in protein folding factors (chaperones), oxidative-species-scavenging molecules, and stress-response mediators, while downregulating genes mostly encoding metabolic enzymes, in particular, those involved in sugar catabolism and amino acid synthesis. Interestingly the use of another stimulus, e.g., anti-CD3 (T-cell activation), also triggers rapid and transient transcriptional modifications. Most of them are shared with flagellin conditions, but some differ, suggesting a gut microbiota adaption, depending on the stimulus and subsequent immune activation. Of note, when mice were reconstituted with three strains instead of four, the transcriptional profiles changes upon acute immune activation were different, suggesting that microbiota response to immunity also depends on its composition and on intra-microbes crosstalks. Altogether, those studies clearly indicate that immunity also deeply impact gut microbiota composition and function, depending on nature and lasting of the immune activation (i.e., acute versus chronic). The challenge is now to identify effector molecules involved in such dialogue, including immune system-derived metabolites.

3.2. The Antimicrobial Peptides

Antimicrobial peptides (AMPs) are key components of innate immunity against invading pathogens and represent one of the oldest innate effector systems. They act by disrupting bacterial cell membranes, modulating the immune response, and regulating inflammation. Being completely accurate, AMPs are not metabolites: They are small cationic molecules of 2–6 kDa that play an important role. The majority of intestinal AMPs is produced by specialized secretory cells of the small intestinal mucosa, namely, the Paneth cells. The expression of AMPs is tightly regulated by the presence of microorganisms via different mechanisms, mainly implicating the activation of PRRs (TLRs and NLRs) in IECs [299]. Of note, IL-22 further promotes their production [300]. Among the various AMPs produced in the GI tract, defensins and cathelicidins constitute the two major classes. It has been shown that GF mice display a reduced AMPs expression [301], and that Paneth cell alteration can lead to dysbiosis [302], underlining the importance of microbiota and Paneth cells for optimal AMPs production and subsequent homeostatic control of intestinal-associated microorganisms, respectively. It is noteworthy that some AMPs can be further metabolized into bioactive metabolites that can shape the microbiota. Ehmann et al. showed that the Human Defensin 5 can be further degraded by proteases into peptides displaying antimicrobial properties. Among them, one peptide fragment promoted Akkermansia sp. growth without decreasing microbial diversity in mice [303]. Thus, AMPs are molecules belonging to innate immunity that can profoundly shape microbiota. Further investigations are required to better characterize the in vivo impact of AMPs and derived bioactive products, to design new microbiota modulation strategies.

3.3. Immune Cells-Derived Metabolites and Metabolite Mimicry

Upon activation, effector immune cells display specific metabolic states (immunometabolism). Indeed, Glycolysis or OXPHOS are preferentially initiated by Teff- and Treg-cells, respectively [304]. Thus, immune cells both consume and produce metabolites that can presumably modify the micro-environment milieu, and thus, influence immune cells and bacterial functions. For instance, Treg-cells can consume extracellular ATP via CD39/CD73 ecto-ATPase activity, leading to Teff-cell dysfunction [305]. Another example, is the production of retinoic acid (RA), a 300 Da metabolite produced by epithelial cells and dendritic cells from retinol. Retinol and RA are derivatives of vitamin A, an essential fat-soluble micronutrient. Their production in the gut relies on their nutritional intake. RA, the bioactive form of vitamin A derivate, is known to modulate the immune system. Indeed, it is involved in T helper cell differentiation [306,307,308], it is a modulator of the innate immune cells [309,310], and a modulator of epithelial tight junction proteins [311,312]. The bioavailability of epithelium-derived RA depends on the microbiota. Intestinal bacteria regulate the expression of the Rdh7 gene of epithelial cells, coding for a retinol dehydrogenase which is involved in RA synthesis [313]. This example demonstrates that a metabolite produced by the intestinal epithelium following a stimulation of the microbiota can impact the immune response. Thus, the effect of RA on gut microbial communities relies on the indirect effect of RA regulating the immune system and mucosal barrier. It is thus conceivable that gut-associated immunity might trigger such metabolic-mediated effects on microbiota. Moreover, it has been shown that mammalian cells can produce molecules resembling bacterial metabolites. Ismail et al. described that colonic cells could produce an AutoInducer (AI)-2 mimic metabolite following tight-junctions disruption, or when submitted to a bacteria-derived secreted molecule [314]. AI-2 is proposed to promote interspecies bacterial communication [219], including in the mammalian gut. Interestingly, AI-2 mimic metabolite production by eukaryotic cells, and subsequent release within the gut lumen, modulates bacterial gene expression through AI-2 receptor signaling and activates quorum sensing system, including in pathogenic strains such as Salmonella typhimurium [314]. One can imagine that several metabolite mimics, able to modulate bacterial activity, might be produced by eukaryotic cells, including immune cells. It becomes now urgent to identify such immune-derived metabolites and metabolites mimics to better understand microbiota/host dialogue and design relevant strategies for manipulating these interactions.

4. Conclusions

This review provided a state-of-the-art description of the metabolites (produced from the microbiota, from the host, or from both) playing a major role in the crosstalk between gut microbiota and immune system.
Forty key metabolites are discussed, together with the modulations and immune pathways triggered in the gut. Evidence of the bidirectional impact of this crosstalk has been reviewed to contribute to a deeper understanding of the host-microorganism relationships, with a specific focus on the correlation between dysbiosis, microbial-based dysmetabolism, and pathogenesis.
To conclude, however extensive this review is, we should consider that 117 known microbial metabolites detected in fecal samples are present on the HMDB website. This suggests that we are at the dawn of the discovery of molecules that can have an impact on human health: Being part of this story is both exciting and challenging.

5. Challenges and Perspectives: How to Hear the Sound from Metabolite-Mediated Microbiota-Host Immunity Crosstalk

Research on metabolites in the gut has evolved extremely rapidly and enhanced our understandings of the crosstalk between host and microbiota in homeostasis and disease. Exploration of their richness and diversity requires special attention; in particular, the following five points should be carefully considered:
  • Improve the analytical tools to detect metabolites in the gut (depth of data acquisition, detection of chemicals of various nature, dynamic range of detection with a large order of magnitude of detection). Currently, ultra-high performance liquid chromatography-high resolution mass spectrometry (UHPLC-HRMS) variants represent the best methods to address these challenges, and, in particular, to increase metabolite detection coverage [315]. The main limitations of LC-MS are the limited structural information and the complexity of obtaining absolute quantification, especially in non-targeted approaches.
  • Correctly assign the identity of a metabolite. The vast majority of information collected by metabolomics is “dark matter”, i.e., chemical signatures that remain uncharacterized. Therefore, new computational solutions to illuminating dark matter are needed [316]. Significant progress has been reported in exploring public data-rich libraries, finding chemicals and associated metadata, and applying molecular networking strategies to accelerate metabolite annotation, especially in the last few years [317,318,319,320,321]. Moreover, the development and maintenance of MS-based spectral databases, together with the increasing practice of sharing metabolomics data through these resources, play a key role in translating dark matter into biological knowledge [322]. Authentic chemical standards should be used to acquire both positive and negative mode MS/MS and MSn spectra to supplement spectral databases such as HMBD [323], NIST20 [324], METLIN [325], MZcloud [326], MassBank [327], ReSpect [328], and GNPS [329].
  • Establish an extensive reference catalog of microbial genomes present in the human gut, as several big projects are already doing, such as the MetaHIT (METAgenomics of the Human Intestinal Tract) project; the Human Microbiome Project, or others [2,3].
  • Integrate multi-omics datasets to recover microbe-metabolite relationships by using statistical analysis. Linear or neural network methods estimate the conditional probability that each molecule is present, given the presence of a specific microorganism [330,331,332]. The mmmvec tool (https://github.com/biocore/mmvec; accessed on 15 January 2021), for instance, can reliably identify all of the experimentally determined P. aeruginosa-produced molecules of interest in the lung of cystic fibrosis patient chronically infected with P. aeruginosa [331].
  • Use experimental models relevant to the concept of molecular dialogue to unveil the respective role of microbiota and/or immune cells in the production of specific metabolites of interest. In a context where the microbial and host metabolite relationships are not elucidated, this first work consists of a detailed experimental design, followed by a fundamental study on metabolites using specific pathological models (KO mice, mono-colonization of the microbiota with certain strains, synthetic biology strategy, GF and specific pathogen-free model, wilding mice [333], culturomic, metabolomic model with minimal microbiota, (reviewed in [334]) and in vitro models [335,336].
Non-targeted metabolomics approaches can stratify the metabolome and to reveal notable differences from a “pathological” metabolome to a “physiological” metabolome. The detection and identification of metabotypes associated with the desired context, thus refining the reading of the metabolome and then encourage to associate its knowledge with previously observed cellular and molecular mechanisms or to set up experiments to define them.
The concept of exploiting metabolomics to perform activity screens to identify biologically active metabolites—which we term activity metabolomics—has been proposed recently by G. Siuzdak [337], and is already having a broad impact on biology. Finally, metabolomics investigation has shown to be a promising tool for the targeting and understanding of the microbiota-host crosstalk; however, this field is still in its infancy and more research and integrative tools are still needed.

Supplementary Materials

The following are available online at https://www.mdpi.com/article/10.3390/metabo11060406/s1, Table S1: Summarizing table of Microbiota-derived metabolites that modulate host immunity in the gut.

Author Contributions

All the authors participated extensively to the work presented in this review. Conceptualization, C.C., G.M. and A.L.G. Writing—original draft preparation, C.P., F.F., C.C., G.M. and A.L.G.; writing—review and editing, B.T., D.H., S.T., A.-R.T., C.P., F.F., C.C., G.M. and A.L.G.; visualization, C.P., G.M., C.C. and A.L.G.; supervision and administration, D.H. and A.L.G.; funding acquisition, D.H., A.L.G. and B.T. All authors have read and agreed to the published version of the manuscript.

Funding

D.H. is supported by GEFLUC Dauphiné-Savoie, Ligue contre le Cancer Comité Isère, Université Grenoble Alpes IDEX Initiatives de Recherche Stratégiques. A.L.G. and B.T. were supported by “Vaincre la mucoviscidose” (VLM) and “Association Grégory Lemarchal” (AGL) (Grant number RF20180502189 and RF20190502416/1/1/68), ANR-15-IDEX-02, SATT Linksium and Fondation Université Grenoble Alpes.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Cavalier-Smith, T. Symbiosis as a Source of Evolutionary Innovation: Speciation and Morphogenesis; The MIT Press: Cambridge, MA, USA, 1992; Volume 7, ISBN 0262519909. [Google Scholar]
  2. Qin, J.; Li, R.; Raes, J.; Arumugam, M.; Burgdorf, K.S.; Manichanh, C.; Nielsen, T.; Pons, N.; Levenez, F.; Yamada, T.; et al. A human gut microbial gene catalogue established by metagenomic sequencing. Nature 2010, 464, 59–65. [Google Scholar] [CrossRef] [PubMed][Green Version]
  3. Huttenhower, C.; Gevers, D.; Knight, R.; Abubucker, S.; Badger, J.H.; Chinwalla, A.T.; Creasy, H.H.; Earl, A.M.; Fitzgerald, M.G.; Fulton, R.S.; et al. Structure, function and diversity of the healthy human microbiome. Nature 2012, 486, 207–214. [Google Scholar] [CrossRef][Green Version]
  4. Shi, Y.; Tyson, G.W.; Delong, E.F. Metatranscriptomics reveals unique microbial small RNAs in the oceans water column. Nature 2009, 459, 266–269. [Google Scholar] [CrossRef]
  5. Maron, P.A.; Ranjard, L.; Mougel, C.; Lemanceau, P. Metaproteomics: A new approach for studying functional microbial ecology. Microb. Ecol. 2007, 53, 486–493. [Google Scholar] [CrossRef]
  6. Francino, M.P. Early development of the gut microbiota and immune health. Pathogens 2014, 3, 769–790. [Google Scholar] [CrossRef][Green Version]
  7. Hooper, L.V.; Littman, D.R.; Macpherson, A.J. Interactions between the microbiota and the immune system. Science 2012, 336, 1268–1273. [Google Scholar] [CrossRef] [PubMed][Green Version]
  8. Belkaid, Y.; Hand, T.W. Role of the microbiota in immunity and inflammation. Cell 2014, 157, 121–141. [Google Scholar] [CrossRef][Green Version]
  9. Zheng, D.; Liwinski, T.; Elinav, E. Interaction between microbiota and immunity in health and disease. Cell Res. 2020, 30, 492–506. [Google Scholar] [CrossRef] [PubMed]
  10. Dorrestein, P.C.; Mazmanian, S.K.; Knight, R. Finding the Missing Links among Metabolites, Microbes, and the Host. Immunity 2014, 40, 824–832. [Google Scholar] [CrossRef][Green Version]
  11. Goodacre, R. Metabolomics of a superorganism. J. Nutr. 2007, 137, 259S–266S. [Google Scholar] [CrossRef] [PubMed][Green Version]
  12. Matsumoto, M.; Kibe, R.; Ooga, T.; Aiba, Y.; Kurihara, S.; Sawaki, E.; Koga, Y.; Benno, Y. Impact of intestinal microbiota on intestinal luminal metabolome. Sci. Rep. 2012, 2, 1–10. [Google Scholar] [CrossRef] [PubMed][Green Version]
  13. Wikoff, W.R.; Anfora, A.T.; Liu, J.; Schultz, P.G.; Lesley, S.A.; Peters, E.C.; Siuzdak, G. Metabolomics analysis reveals large effects of gut microflora on mammalian blood metabolites. Proc. Natl. Acad. Sci. USA 2009, 106, 3698–3703. [Google Scholar] [CrossRef][Green Version]
  14. Macfarlane, S.; Macfarlane, G.T. Regulation of short-chain fatty acid production. Proc. Nutr. Soc. 2003, 62, 67–72. [Google Scholar] [CrossRef] [PubMed]
  15. Louis, P.; Flint, H.J. Formation of propionate and butyrate by the human colonic microbiota. Environ. Microbiol. 2017, 19, 29–41. [Google Scholar] [CrossRef] [PubMed][Green Version]
  16. Cummings, J.H.; Pomare, E.W.; Branch, H.W.J.; Naylor, C.P.E.; MacFarlane, G.T. Short chain fatty acids in human large intestine, portal, hepatic and venous blood. Gut 1987, 28, 1221–1227. [Google Scholar] [CrossRef][Green Version]
  17. Sivaprakasam, S.; Bhutia, Y.D.; Yang, S.; Ganapathy, V. Short-chain fatty acid transporters: Role in colonic homeostasis. In Comprehensive Physiology; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2018; Volume 8, pp. 299–314. [Google Scholar]
  18. Donohoe, D.R.; Garge, N.; Zhang, X.; Sun, W.; O’Connell, T.M.; Bunger, M.K.; Bultman, S.J. The microbiome and butyrate regulate energy metabolism and autophagy in the mammalian colon. Cell Metab. 2011, 13, 517–526. [Google Scholar] [CrossRef][Green Version]
  19. Schönfeld, P.; Wojtczak, L. Short- and medium-chain fatty acids in energy metabolism: The cellular perspective. J. Lipid Res. 2016, 57, 943–954. [Google Scholar] [CrossRef][Green Version]
  20. Dalile, B.; Van Oudenhove, L.; Vervliet, B.; Verbeke, K. The role of short-chain fatty acids in microbiota–gut–brain communication. Nat. Rev. Gastroenterol. Hepatol. 2019, 16, 461–478. [Google Scholar] [CrossRef]
  21. Donohoe, D.R.; Collins, L.B.; Wali, A.; Bigler, R.; Sun, W.; Bultman, S.J. The Warburg Effect Dictates the Mechanism of Butyrate-Mediated Histone Acetylation and Cell Proliferation. Mol. Cell 2012, 48, 612–626. [Google Scholar] [CrossRef][Green Version]
  22. Gerhauser, C. Impact of dietary gut microbial metabolites on the epigenome. Philos. Trans. R. Soc. B Biol. Sci. 2018, 373, 20170359. [Google Scholar] [CrossRef][Green Version]
  23. Brown, A.J.; Goldsworthy, S.M.; Barnes, A.A.; Eilert, M.M.; Tcheang, L.; Daniels, D.; Muir, A.I.; Wigglesworth, M.J.; Kinghorn, I.; Fraser, N.J.; et al. The orphan G protein-coupled receptors GPR41 and GPR43 are activated by propionate and other short chain carboxylic acids. J. Biol. Chem. 2003, 278, 11312–11319. [Google Scholar] [CrossRef][Green Version]
  24. Le Poul, E.; Loison, C.; Struyf, S.; Springael, J.Y.; Lannoy, V.; Decobecq, M.E.; Brezillon, S.; Dupriez, V.; Vassart, G.; Van Damme, J.; et al. Functional characterization of human receptors for short chain fatty acids and their role in polymorphonuclear cell activation. J. Biol. Chem. 2003, 278, 25481–25489. [Google Scholar] [CrossRef] [PubMed][Green Version]
  25. Thangaraju, M.; Cresci, G.A.; Liu, K.; Ananth, S.; Gnanaprakasam, J.P.; Browning, D.D.; Mellinger, J.D.; Smith, S.B.; Digby, G.J.; Lambert, N.A.; et al. GPFM 09A is a G-protein-coupled receptor for the bacterial fermentation product butyrate and functions as a tumor suppressor in colon. Cancer Res. 2009, 69, 2826–2832. [Google Scholar] [CrossRef][Green Version]
  26. Corrêa-Oliveira, R.; Fachi, J.L.; Vieira, A.; Sato, F.T.; Vinolo, M.A.R. Regulation of immune cell function by short-chain fatty acids. Clin. Transl. Immunol. 2016, 5, e73. [Google Scholar] [CrossRef]
  27. Sina, C.; Gavrilova, O.; Förster, M.; Till, A.; Derer, S.; Hildebrand, F.; Raabe, B.; Chalaris, A.; Scheller, J.; Rehmann, A.; et al. G Protein-Coupled Receptor 43 Is Essential for Neutrophil Recruitment during Intestinal Inflammation. J. Immunol. 2009, 183, 7514–7522. [Google Scholar] [CrossRef]
  28. Kaiko, G.E.; Ryu, S.H.; Koues, O.I.; Collins, P.L.; Solnica-Krezel, L.; Pearce, E.J.; Pearce, E.L.; Oltz, E.M.; Stappenbeck, T.S. The Colonic Crypt Protects Stem Cells from Microbiota-Derived Metabolites. Cell 2016, 165, 1708–1720. [Google Scholar] [CrossRef][Green Version]
  29. Kelly, C.J.; Zheng, L.; Campbell, E.L.; Saeedi, B.; Scholz, C.C.; Bayless, A.J.; Wilson, K.E.; Glover, L.E.; Kominsky, D.J.; Magnuson, A.; et al. Crosstalk between microbiota-derived short-chain fatty acids and intestinal epithelial HIF augments tissue barrier function. Cell Host Microbe 2015, 17, 662–671. [Google Scholar] [CrossRef][Green Version]
  30. Tedelind, S.; Westberg, F.; Kjerrulf, M.; Vidal, A. Anti-inflammatory properties of the short-chain fatty acids acetate and propionate: A study with relevance to inflammatory bowel disease. World J. Gastroenterol. 2007, 13, 2826–2832. [Google Scholar] [CrossRef] [PubMed]
  31. Arpaia, N.; Campbell, C.; Fan, X.; Dikiy, S.; Van Der Veeken, J.; Deroos, P.; Liu, H.; Cross, J.R.; Pfeffer, K.; Coffer, P.J.; et al. Metabolites produced by commensal bacteria promote peripheral regulatory T-cell generation. Nature 2013, 504, 451–455. [Google Scholar] [CrossRef]
  32. Yang, W.; Yu, T.; Huang, X.; Bilotta, A.J.; Xu, L.; Lu, Y.; Sun, J.; Pan, F.; Zhou, J.; Zhang, W.; et al. Intestinal microbiota-derived short-chain fatty acids regulation of immune cell IL-22 production and gut immunity. Nat. Commun. 2020, 11, 1–18. [Google Scholar] [CrossRef] [PubMed]
  33. Gurav, A.; Sivaprakasam, S.; Bhutia, Y.D.; Boettger, T.; Singh, N.; Ganapathy, V. Slc5a8, a Na+-coupled high-affinity transporter for short-chain fatty acids, is a conditional tumour suppressor in colon that protects against colitis and colon cancer under low-fibre dietary conditions. Biochem. J. 2015, 469, 267–278. [Google Scholar] [CrossRef]
  34. Hui, W.; Yu, D.; Cao, Z.; Zhao, X. Butyrate inhibit collagen-induced arthritis via Treg/IL-10/Th17 axis. Int. Immunopharmacol. 2019, 68, 226–233. [Google Scholar] [CrossRef] [PubMed]
  35. Liu, L.; Li, L.; Min, J.; Wang, J.; Wu, H.; Zeng, Y.; Chen, S.; Chu, Z. Butyrate interferes with the differentiation and function of human monocyte-derived dendritic cells. Cell. Immunol. 2012, 277, 66–73. [Google Scholar] [CrossRef]
  36. Singh, N.; Thangaraju, M.; Prasad, P.D.; Martin, P.M.; Lambert, N.A.; Boettger, T.; Offermanns, S.; Ganapathy, V. Blockade of dendritic cell development by bacterial fermentation products butyrate and propionate through a transporter (Slc5a8)-dependent inhibition of histone deacetylases. J. Biol. Chem. 2010, 285, 27601–27608. [Google Scholar] [CrossRef] [PubMed][Green Version]
  37. Singh, N.; Gurav, A.; Sivaprakasam, S.; Brady, E.; Padia, R.; Shi, H.; Thangaraju, M.; Prasad, P.D.; Manicassamy, S.; Munn, D.H.; et al. Activation of Gpr109a, receptor for niacin and the commensal metabolite butyrate, suppresses colonic inflammation and carcinogenesis. Immunity 2014, 40, 128–139. [Google Scholar] [CrossRef][Green Version]
  38. Schulthess, J.; Pandey, S.; Capitani, M.; Rue-Albrecht, K.C.; Arnold, I.; Franchini, F.; Chomka, A.; Ilott, N.E.; Johnston, D.G.W.; Pires, E.; et al. The Short Chain Fatty Acid Butyrate Imprints an Antimicrobial Program in Macrophages. Immunity 2019, 50, 432–445.e7. [Google Scholar] [CrossRef] [PubMed][Green Version]
  39. Lewis, K.; Lutgendorff, F.; Phan, V.; Söderholm, J.D.; Sherman, P.M.; McKay, D.M. Enhanced translocation of bacteria across metabolically stressed epithelia is reduced by butyrate. Inflamm. Bowel Dis. 2010, 16, 1138–1148. [Google Scholar] [CrossRef]
  40. Venegas, D.P.; De La Fuente, M.K.; Landskron, G.; González, M.J.; Quera, R.; Dijkstra, G.; Harmsen, H.J.M.; Faber, K.N.; Hermoso, M.A. Short chain fatty acids (SCFAs)mediated gut epithelial and immune regulation and its relevance for inflammatory bowel diseases. Front. Immunol. 2019, 10, 277. [Google Scholar] [CrossRef][Green Version]
  41. Mirmonsef, P.; Zariffard, M.R.; Gilbert, D.; Makinde, H.; Landay, A.L.; Spear, G.T. Short-Chain Fatty Acids Induce Pro-Inflammatory Cytokine Production Alone and in Combination with Toll-Like Receptor Ligands. Am. J. Reprod. Immunol. 2012, 67, 391–400. [Google Scholar] [CrossRef][Green Version]
  42. 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, 1–15. [Google Scholar] [CrossRef][Green Version]
  43. 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, 396–406.e10. [Google Scholar] [CrossRef]
  44. Park, J.; Kim, M.; Kang, S.G.; Jannasch, A.H.; Cooper, B.; Patterson, J.; Kim, C.H. Short-chain fatty acids induce both effector and regulatory T cells by suppression of histone deacetylases and regulation of the mTOR-S6K pathway. Mucosal Immunol. 2015, 8, 80–93. [Google Scholar] [CrossRef][Green Version]
  45. Belenguer, A.; Duncan, S.H.; Holtrop, G.; Anderson, S.E.; Lobley, G.E.; Flint, H.J. Impact of pH on lactate formation and utilization by human fecal microbial communities. Appl. Environ. Microbiol. 2007, 73, 6526–6533. [Google Scholar] [CrossRef][Green Version]
  46. Belenguer, A.; Holtrop, G.; Duncan, S.H.; Anderson, S.E.; Calder, A.G.; Flint, H.J.; Lobley, G.E. Rates of productionand utilization of lactate by microbial communities fromthe human colon. FEMS Microbiol. Ecol. 2011, 77, 107–119. [Google Scholar] [CrossRef][Green Version]
  47. Iraporda, C.; Errea, A.; Romanin, D.E.; Cayet, D.; Pereyra, E.; Pignataro, O.; Sirard, J.C.; Garrote, G.L.; Abraham, A.G.; Rumbo, M. Lactate and short chain fatty acids produced by microbial fermentation downregulate proinflammatory responses in intestinal epithelial cells and myeloid cells. Immunobiology 2015, 220, 1161–1169. [Google Scholar] [CrossRef]
  48. Hove, H.; Nordgaard-Andersen, I.; Mortensen, P.B. Faecal DL-lactate concentration in 100 gastrointestinal patients. Scand. J. Gastroenterol. 1994, 29, 255–259. [Google Scholar] [CrossRef]
  49. Vernia, P.; Caprilli, R.; Latella, G.; Barbetti, F.; Magliocca, F.M.; Cittadini, M. Fecal Lactate and Ulcerative Colitis. Gastroenterology 1988, 95, 1564–1568. [Google Scholar] [CrossRef]
  50. Wang, S.P.; Rubio, L.A.; Duncan, S.H.; Donachie, G.E.; Holtrop, G.; Lo, G.; Farquharson, F.M.; Wagner, J.; Parkhill, J.; Louis, P.; et al. Pivotal Roles for pH, Lactate, and Lactate-Utilizing Bacteria in the Stability of a Human Colonic Microbial Ecosystem. mSystems 2020, 5, 645–665. [Google Scholar] [CrossRef]
  51. Krautkramer, K.A.; Fan, J.; Bäckhed, F. Gut microbial metabolites as multi-kingdom intermediates. Nat. Rev. Microbiol. 2021, 19, 77–94. [Google Scholar] [CrossRef] [PubMed]
  52. Ranganathan, P.; Shanmugam, A.; Swafford, D.; Suryawanshi, A.; Bhattacharjee, P.; Hussein, M.S.; Koni, P.A.; Prasad, P.D.; Kurago, Z.B.; Thangaraju, M.; et al. GPR81, a Cell-Surface Receptor for Lactate, Regulates Intestinal Homeostasis and Protects Mice from Experimental Colitis. J. Immunol. 2018, ji1700604. [Google Scholar] [CrossRef] [PubMed]
  53. Errea, A.; Cayet, D.; Marchetti, P.; Tang, C.; Kluza, J.; Offermanns, S.; Sirard, J.-C.; Rumbo, M. Lactate Inhibits the Pro-Inflammatory Response and Metabolic Reprogramming in Murine Macrophages in a GPR81-Independent Manner. PLoS ONE 2016, 11, e0163694. [Google Scholar] [CrossRef] [PubMed][Green Version]
  54. SHAN, T.; CHEN, S.; CHEN, X.; WU, T.; YANG, Y.; LI, S.; MA, J.; ZHAO, J.; LIN, W.; LI, W.; et al. M2-TAM subsets altered by lactic acid promote T-cell apoptosis through the PD-L1/PD-1 pathway. Oncol. Rep. 2020, 44, 1885–1894. [Google Scholar] [CrossRef] [PubMed]
  55. Dietl, K.; Renner, K.; Dettmer, K.; Timischl, B.; Eberhart, K.; Dorn, C.; Hellerbrand, C.; Kastenberger, M.; Kunz-Schughart, L.A.; Oefner, P.J.; et al. Lactic Acid and Acidification Inhibit TNF Secretion and Glycolysis of Human Monocytes. J. Immunol. 2010, 184, 1200–1209. [Google Scholar] [CrossRef] [PubMed]
  56. Haas, R.; Smith, J.; Rocher-Ros, V.; Nadkarni, S.; Montero-Melendez, T.; D’Acquisto, F.; Bland, E.J.; Bombardieri, M.; Pitzalis, C.; Perretti, M.; et al. Lactate regulates metabolic and proinflammatory circuits in control of T cell migration and effector functions. PLoS Biol. 2015, 13, e1002202. [Google Scholar] [CrossRef] [PubMed]
  57. Angelin, A.; Gil-de-Gómez, L.; Dahiya, S.; Jiao, J.; Guo, L.; Levine, M.H.; Wang, Z.; Quinn, W.J.; Kopinski, P.K.; Wang, L.; et al. Foxp3 Reprograms T Cell Metabolism to Function in Low-Glucose, High-Lactate Environments. Cell Metab. 2017, 25, 1282–1293.e7. [Google Scholar] [CrossRef][Green Version]
  58. Upadhyay, D.; Singh, A.; Das, P.; Mehtab, J.; Dattagupta, S.; Ahuja, V.; Makharia, G.K.; Jagannathan, N.R.; Sharma, U. Abnormalities in metabolic pathways in celiac disease investigated by the metabolic profiling of small intestinal mucosa, blood plasma and urine by NMR spectroscopy. NMR Biomed. 2020, 33, e4305. [Google Scholar] [CrossRef]
  59. Faith, J.J.; Ahern, P.P.; Ridaura, V.K.; Cheng, J.; Gordon, J.I. Identifying gut microbe-host phenotype relationships using combinatorial communities in gnotobiotic mice. Sci. Transl. Med. 2014, 6, 220ra11. [Google Scholar] [CrossRef][Green Version]
  60. Meijer-Severs, G.J.; Van Santen, E. Short-chain fatty acids and succinate in feces of healthy human volunteers and their correlation with anaerobe cultural counts. Scand. J. Gastroenterol. 1987, 22, 672–676. [Google Scholar] [CrossRef]
  61. Macias-Ceja, D.C.; Ortiz-Masiá, D.; Salvador, P.; Gisbert-Ferrándiz, L.; Hernández, C.; Hausmann, M.; Rogler, G.; Esplugues, J.V.; Hinojosa, J.; Alós, R.; et al. Succinate receptor mediates intestinal inflammation and fibrosis. Mucosal Immunol. 2019, 12, 178–187. [Google Scholar] [CrossRef][Green Version]
  62. Littlewood-Evans, A.; Sarret, S.; Apfel, V.; Loesle, P.; Dawson, J.; Zhang, J.; Muller, A.; Tigani, B.; Kneuer, R.; Patel, S.; et al. GPR91 senses extracellular succinate released from inflammatory macrophages and exacerbates rheumatoid arthritis. J. Exp. Med. 2016, 213, 1655–1662. [Google Scholar] [CrossRef]
  63. Pålsson-McDermott, E.M.; O’Neill, L.A.J. Targeting immunometabolism as an anti-inflammatory strategy. Cell Res. 2020, 30, 300–314. [Google Scholar] [CrossRef][Green Version]
  64. Rubic, T.; Lametschwandtner, G.; Jost, S.; Hinteregger, S.; Kund, J.; Carballido-Perrig, N.; Schwärzler, C.; Junt, T.; Voshol, H.; Meingassner, J.G.; et al. Triggering the succinate receptor GPR91 on dendritic cells enhances immunity. Nat. Immunol. 2008, 9, 1261–1269. [Google Scholar] [CrossRef]
  65. Saraiva, A.L.; Veras, F.P.; Peres, R.S.; Talbot, J.; De Lima, K.A.; Luiz, J.P.; Carballido, J.M.; Cunha, T.M.; Cunha, F.Q.; Ryffel, B.; et al. Succinate receptor deficiency attenuates arthritis by reducing dendritic cell traffic and expansion of Th17 cells in the lymph nodes. FASEB J. 2018, 32, 6550–6558. [Google Scholar] [CrossRef]
  66. Nadjsombati, M.S.; McGinty, J.W.; Lyons-Cohen, M.R.; Jaffe, J.B.; DiPeso, L.; Schneider, C.; Miller, C.N.; Pollack, J.L.; Nagana Gowda, G.A.; Fontana, M.F.; et al. Detection of Succinate by Intestinal Tuft Cells Triggers a Type 2 Innate Immune Circuit. Immunity 2018, 49, 33–41.e7. [Google Scholar] [CrossRef][Green Version]
  67. Schneider, C.; O’Leary, C.E.; von Moltke, J.; Liang, H.E.; Ang, Q.Y.; Turnbaugh, P.J.; Radhakrishnan, S.; Pellizzon, M.; Ma, A.; Locksley, R.M. A Metabolite-Triggered Tuft Cell-ILC2 Circuit Drives Small Intestinal Remodeling. Cell 2018, 174, 271–284.e14. [Google Scholar] [CrossRef] [PubMed][Green Version]
  68. Lei, W.; Ren, W.; Ohmoto, M.; Urban, J.F.; Matsumoto, I.; Margolskee, R.F.; Jiang, P. Activation of intestinal tuft cell-expressed sucnr1 triggers type 2 immunity in the mouse small intestine. Proc. Natl. Acad. Sci. USA 2018, 115, 5552–5557. [Google Scholar] [CrossRef][Green Version]
  69. Jakobsdottir, G.; Xu, J.; Molin, G.; Ahrné, S.; Nyman, M. High-fat diet reduces the formation of butyrate, but increases succinate, inflammation, liver fat and cholesterol in rats, while dietary fibre counteracts these effects. PLoS ONE 2013, 8, e80476. [Google Scholar] [CrossRef] [PubMed][Green Version]
  70. Ooi, M.; Nishiumi, S.; Yoshie, T.; Shiomi, Y.; Kohashi, M.; Fukunaga, K.; Nakamura, S.; Matsumoto, T.; Hatano, N.; Shinohara, M.; et al. GC/MS-based profiling of amino acids and TCA cycle-related molecules in ulcerative colitis. Inflamm. Res. 2011, 60, 831–840. [Google Scholar] [CrossRef]
  71. Connors, J.; Dawe, N.; Van Limbergen, J. The role of succinate in the regulation of intestinal inflammation. Nutrients 2019, 11, 25. [Google Scholar] [CrossRef] [PubMed][Green Version]
  72. Wolak, N.; Zawrotniak, M.; Gogol, M.; Kozik, A.; Rapala-Kozik, M. Vitamins B1, B2, B3 and B9—Occurrence, Biosynthesis Pathways and Functions in Human Nutrition. Mini-Rev. Med. Chem. 2016, 17, 1075–1111. [Google Scholar] [CrossRef]
  73. Magnúsdóttir, S.; Ravcheev, D.; De Crécy-Lagard, V.; Thiele, I. Systematic genome assessment of B-vitamin biosynthesis suggests cooperation among gut microbes. Front. Genet. 2015, 6, 148. [Google Scholar] [CrossRef] [PubMed][Green Version]
  74. Putnam, E.E.; Goodman, A.L. B vitamin acquisition by gut commensal bacteria. PLoS Pathog. 2020, 16, e1008208. [Google Scholar] [CrossRef][Green Version]
  75. Rodionov, D.A.; Arzamasov, A.A.; Khoroshkin, M.S.; Iablokov, S.N.; Leyn, S.A.; Peterson, S.N.; Novichkov, P.S.; Osterman, A.L. Micronutrient requirements and sharing capabilities of the human gut microbiome. Front. Microbiol. 2019, 10, 1316. [Google Scholar] [CrossRef][Green Version]
  76. Radjabzadeh, D.; Boer, C.G.; Beth, S.A.; van der Wal, P.; Kiefte-De Jong, J.C.; Jansen, M.A.E.; Konstantinov, S.R.; Peppelenbosch, M.P.; Hays, J.P.; Jaddoe, V.W.V.; et al. Diversity, compositional and functional differences between gut microbiota of children and adults. Sci. Rep. 2020, 10, 1–13. [Google Scholar] [CrossRef] [PubMed][Green Version]
  77. Arumugam, M.; Raes, J.; Pelletier, E.; Le Paslier, D.; Yamada, T.; Mende, D.R.; Fernandes, G.R.; Tap, J.; Bruls, T.; Batto, J.M.; et al. Enterotypes of the human gut microbiome. Nature 2011, 473, 174–180. [Google Scholar] [CrossRef]
  78. Costea, P.I.; Hildebrand, F.; Manimozhiyan, A.; Bäckhed, F.; Blaser, M.J.; Bushman, F.D.; De Vos, W.M.; Ehrlich, S.D.; Fraser, C.M.; Hattori, M.; et al. Enterotypes in the landscape of gut microbial community composition. Nat. Microbiol. 2017, 3, 8–16. [Google Scholar] [CrossRef] [PubMed][Green Version]
  79. Yoshii, K.; Hosomi, K.; Sawane, K.; Kunisawa, J. Metabolism of dietary and microbial vitamin b family in the regulation of host immunity. Front. Nutr. 2019, 6, 48. [Google Scholar] [CrossRef] [PubMed][Green Version]
  80. Nabokina, S.M.; Said, H.M. A high-affinity and specific carrier-mediated mechanism for uptake of thiamine pyrophosphate by human colonic epithelial cells. Am. J. Physiol.-Gastrointest. Liver Physiol. 2012, 303, G389–G395. [Google Scholar] [CrossRef] [PubMed][Green Version]
  81. Frank, R.A.W.; Leeper, F.J.; Luisi, B.F. Structure, mechanism and catalytic duality of thiamine-dependent enzymes. Cell. Mol. Life Sci. 2007, 64, 892–905. [Google Scholar] [CrossRef]
  82. Ashokkumar, B.; Kumar, J.S.; Hecht, G.A.; Said, H.M. Enteropathogenic Escherichia coli inhibits intestinal vitamin B1 (thiamin) uptake: Studies with human-derived intestinal epithelial Caco-2 cells. Am. J. Physiol.-Gastrointest. Liver Physiol. 2009, 297, 825–833. [Google Scholar] [CrossRef]
  83. Subramanya, S.B.; Subramanian, V.S.; Said, H.M. Chronic alcohol consumption and intestinal thiamin absorption: Effects on physiological and molecular parameters of the uptake process. Am. J. Physiol.-Gastrointest. Liver Physiol. 2010, 299, G23–G31. [Google Scholar] [CrossRef] [PubMed][Green Version]
  84. Seligmann, H.; Levi, R.; Konijn, A.M.; Prokocimer, M. Thiamine deficiency in patients with B-chronic lymphocytic leukaemia: A pilot study. Postgrad. Med. J. 2001, 77, 582–585. [Google Scholar] [CrossRef][Green Version]
  85. Müri, R.M.; Von Overbeck, J.; Furrer, J.; Ballmer, P.E. Thiamin deficiency in HIV-positive patients: Evaluation by erythrocyte transketolase activity and thiamin pyrophosphate effect. Clin. Nutr. 1999, 18, 375–378. [Google Scholar] [CrossRef]
  86. Ottinger, C.A.; Honeyfield, D.C.; Densmore, C.L.; Iwanowicz, L.R. Impact of thiamine deficiency on T-cell dependent and T-cell independent antibody production in lake trout. J. Aquat. Anim. Health 2012, 24, 258–273. [Google Scholar] [CrossRef]
  87. de Moreno de LeBlanc, A.; Levit, R.; de Giori, G.S.; LeBlanc, J.G. Vitamin Producing Lactic Acid Bacteria as Complementary Treatments for Intestinal Inflammation. Antiinflamm. Antiallergy Agents Med. Chem. 2018, 17, 50–56. [Google Scholar] [CrossRef][Green Version]
  88. Li, L.; Krause, L.; Somerset, S. Associations between micronutrient intakes and gut microbiota in a group of adults with cystic fibrosis. Clin. Nutr. 2017, 36, 1097–1104. [Google Scholar] [CrossRef]
  89. Ghashut, R.A.; McMillan, D.C.; Kinsella, J.; Talwar, D. Erythrocyte concentrations of B1, B2, B6 but not plasma C and E are reliable indicators of nutrition status in the presence of systemic inflammation. Clin. Nutr. ESPEN 2017, 17, 54–62. [Google Scholar] [CrossRef] [PubMed][Green Version]
  90. Dey, S.; Bishayi, B. Riboflavin along with antibiotics balances reactive oxygen species and inflammatory cytokines and controls Staphylococcus aureus infection by boosting murine macrophage function and regulates inflammation. J. Inflamm. 2016, 13, 1–21. [Google Scholar] [CrossRef][Green Version]
  91. Mazur-Bialy, A.I.; Pocheć, E. Riboflavin reduces pro-inflammatory activation of adipocyte-macrophage co-culture. Potential application of vitamin B2 enrichment for attenuation of insulin resistance and metabolic syndrome development. Molecules 2016, 21, 1724. [Google Scholar] [CrossRef][Green Version]
  92. Qureshi, A.A.; Tan, X.; Reis, J.C.; Badr, M.Z.; Papasian, C.J.; Morrison, D.C.; Qureshi, N. Suppression of nitric oxide induction and pro-inflammatory cytokines by novel proteasome inhibitors in various experimental models. Lipids Health Dis. 2011, 10, 1–25. [Google Scholar] [CrossRef][Green Version]
  93. Tastan, C.; Karhan, E.; Zhou, W.; Fleming, E.; Voigt, A.Y.; Yao, X.; Wang, L.; Horne, M.; Placek, L.; Kozhaya, L.; et al. Tuning of human MAIT cell activation by commensal bacteria species and MR1-dependent T-cell presentation. Mucosal Immunol. 2018, 11, 1591–1605. [Google Scholar] [CrossRef][Green Version]
  94. Kumar, V.; Ahmad, A. Role of MAIT cells in the immunopathogenesis of inflammatory diseases: New players in old game. Int. Rev. Immunol. 2018, 37, 90–110. [Google Scholar] [CrossRef]
  95. Von Martels, J.Z.H.; Bourgonje, A.R.; Klaassen, M.A.Y.; Alkhalifah, H.A.A.; Sadaghian Sadabad, M.; Vich Vila, A.; Gacesa, R.; Gabriëls, R.Y.; Steinert, R.E.; Jansen, B.H.; et al. Riboflavin Supplementation in Patients with Crohn’s Disease [the RISE-UP study]. J. Crohn’s Colitis 2020, 14, 595–607. [Google Scholar] [CrossRef]
  96. Mohedano, M.L.; Hernández-Recio, S.; Yépez, A.; Requena, T.; Martínez-Cuesta, M.C.; Peláez, C.; Russo, P.; LeBlanc, J.G.; Spano, G.; Aznar, R.; et al. Real-time detection of riboflavin production by Lactobacillus plantarum strains and tracking of their gastrointestinal survival and functionality in vitro and in vivo using mCherry labeling. Front. Microbiol. 2019, 10, 1748. [Google Scholar] [CrossRef][Green Version]
  97. Shats, I.; Williams, J.G.; Liu, J.; Makarov, M.V.; Wu, X.; Lih, F.B.; Deterding, L.J.; Lim, C.; Xu, X.; Randall, T.A.; et al. Bacteria Boost Mammalian Host NAD Metabolism by Engaging the Deamidated Biosynthesis Pathway. Cell Metab. 2020, 31, 564–579.e7. [Google Scholar] [CrossRef]
  98. Cetina Biefer, H.R.; Vasudevan, A.; Elkhal, A. Aspects of tryptophan and nicotinamide adenine dinucleotide in immunity: A new twist in an old tale. Int. J. Tryptophan Res. 2017, 10, 117864691771349. [Google Scholar] [CrossRef][Green Version]
  99. Rajman, L.; Chwalek, K.; Sinclair, D.A. Therapeutic Potential of NAD-Boosting Molecules: The In Vivo Evidence. Cell Metab. 2018, 27, 529–547. [Google Scholar] [CrossRef][Green Version]
  100. Blacher, E.; Bashiardes, S.; Shapiro, H.; Rothschild, D.; Mor, U.; Dori-Bachash, M.; Kleimeyer, C.; Moresi, C.; Harnik, Y.; Zur, M.; et al. Potential roles of gut microbiome and metabolites in modulating ALS in mice. Nature 2019, 572, 474–480. [Google Scholar] [CrossRef] [PubMed]
  101. Yamamoto, Y.; Nakanishi, Y.; Murakami, S.; Aw, W.; Tsukimi, T.; Nozu, R.; Ueno, M.; Hioki, K.; Nakahigashi, K.; Hirayama, A.; et al. A Metabolomic-Based Evaluation of the Role of Commensal Microbiota throughout the Gastrointestinal Tract in Mice. Microorganisms 2018, 6, 101. [Google Scholar] [CrossRef][Green Version]
  102. Montserrat- de la Paz, S.; Naranjo, M.C.; Lopez, S.; Abia, R.; Muriana, F.J.G.; Bermudez, B. Niacin and its metabolites as master regulators of macrophage activation. J. Nutr. Biochem. 2017, 39, 40–47. [Google Scholar] [CrossRef] [PubMed]
  103. Hong, G.; Zheng, D.; Zhang, L.; Ni, R.; Wang, G.; Fan, G.C.; Lu, Z.; Peng, T. Administration of nicotinamide riboside prevents oxidative stress and organ injury in sepsis. Free Radic. Biol. Med. 2018, 123, 125–137. [Google Scholar] [CrossRef]
  104. Ganji, S.H.; Kamanna, V.S.; Kashyap, M.L. Niacin decreases leukocyte myeloperoxidase: Mechanistic role of redox agents and Src/p38MAP kinase. Atherosclerosis 2014, 235, 554–561. [Google Scholar] [CrossRef]
  105. Ferreira, R.G.; Matsui, T.C.; Godin, A.M.; Gomides, L.F.; Pereira-Silva, P.E.M.; Duarte, I.D.G.; Menezes, G.; Coelho, M.M.; Klein, A. Neutrophil recruitment is inhibited by nicotinamide in experimental pleurisy in mice. Eur. J. Pharmacol. 2012, 685, 198–204. [Google Scholar] [CrossRef][Green Version]
  106. Bhatt, B.; Zeng, P.; Zhu, H.; Sivaprakasam, S.; Li, S.; Xiao, H.; Dong, L.; Shiao, P.; Kolhe, R.; Patel, N.; et al. Gpr109a Limits Microbiota-Induced IL-23 Production To Constrain ILC3-Mediated Colonic Inflammation. J. Immunol. 2018, 200, 2905–2914. [Google Scholar] [CrossRef][Green Version]
  107. Nitto, T.; Onodera, K. The linkage between coenzyme A metabolism and inflammation: Roles of pantetheinase. J. Pharmacol. Sci. 2013, 123, 1–8. [Google Scholar] [CrossRef][Green Version]
  108. He, W.; Hu, S.; Du, X.; Wen, Q.; Zhong, X.P.; Zhou, X.; Zhou, C.; Xiong, W.; Gao, Y.; Zhang, S.; et al. Vitamin B5 reduces bacterial growth via regulating innate immunity and adaptive immunity in mice infected with Mycobacterium tuberculosis. Front. Immunol. 2018, 9, 365. [Google Scholar] [CrossRef][Green Version]
  109. Ghosal, A.; Lambrecht, N.; Subramanya, S.B.; Kapadia, R.; Said, H.M. Conditional knockout of the Slc5a6 gene in mouse intestine impairs biotin absorption. Am. J. Physiol.-Gastrointest. Liver Physiol. 2013, 304, G64–G71. [Google Scholar] [CrossRef][Green Version]
  110. Sabui, S.; Kapadia, R.; Ghosal, A.; Schneider, M.; Lambrecht, N.W.G.; Said, H.M. Biotin and pantothenic acid oversupplementation to conditional SLC5A6 KO mice prevents the development of intestinal mucosal abnormalities and growth defects. Am. J. Physiol.-Cell Physiol. 2018, 315, C73–C79. [Google Scholar] [CrossRef]
  111. Spady, M. Vitamin B6 Deficiency. AJN, Am. J. Nurs. 1960, 60, 83. [Google Scholar] [CrossRef]
  112. Rosenberg, J.; Ischebeck, T.; Commichau, F.M. Vitamin B6 metabolism in microbes and approaches for fermentative production. Biotechnol. Adv. 2017, 35, 31–40. [Google Scholar] [CrossRef]
  113. Rosenberg, J.; Yeak, K.Y.C.; Commichau, F.M. A two-step evolutionary process establishes a non-native vitamin B6 pathway in Bacillus subtilis. Environ. Microbiol. 2018, 20, 156–168. [Google Scholar] [CrossRef]
  114. Nirmagustina, D.E.; Yang, Y.; Kumrungsee, T.; Yanaka, N.; Kato, N. Gender difference and dietary supplemental vitamin B6: Impact on colon luminal environment. J. Nutr. Sci. Vitaminol. 2018, 64, 116–128. [Google Scholar] [CrossRef][Green Version]
  115. Zheng, X.; Feng, L.; Jiang, W.D.; Wu, P.; Liu, Y.; Jiang, J.; Kuang, S.Y.; Tang, L.; Tang, W.N.; Zhang, Y.A.; et al. Dietary pyridoxine deficiency reduced growth performance and impaired intestinal immune function associated with TOR and NF-κB signalling of young grass carp (Ctenopharyngodon idella). Fish Shellfish. Immunol. 2017, 70, 682–700. [Google Scholar] [CrossRef]
  116. Roubenoff, R.; Roubenoff, R.A.; Selhub, J.; Nadeau, M.R.; Cannon, J.G.; Freeman, L.M.; Dinarello, C.A.; Rosenberg, I.H. Abnormal vitamin b6 status in rheumatoid cachexia association with spontaneous tumor necrosis factor α production and markers of inflammation. Arthritis Rheum. 1995, 38, 105–109. [Google Scholar] [CrossRef]
  117. Chiang, E.P.I.; Selhub, J.; Bagley, P.J.; Dallal, G.; Roubenoff, R. Pyridoxine supplementation corrects vitamin B6 deficiency but does not improve inflammation in patients with rheumatoid arthritis. Arthritis Res. Ther. 2005, 7, R1404. [Google Scholar] [CrossRef][Green Version]
  118. Molina-López, J.; Florea, D.; Quintero-Osso, B.; de la Cruz, A.P.; Rodríguez-Elvira, M.; del Pozo, E.P. Pyridoxal-5′-phosphate deficiency is associated with hyperhomocysteinemia regardless of antioxidant, thiamine, riboflavin, cobalamine, and folate status in critically ill patients. Clin. Nutr. 2016, 35, 706–712. [Google Scholar] [CrossRef] [PubMed]
  119. Dusitanond, P.; Eikelboom, J.W.; Hankey, G.J.; Thom, J.; Gilmore, G.; Loh, K.; Yi, Q.; Klijn, C.J.M.; Langton, P.; Van Bockxmeer, F.M.; et al. Homocysteine-lowering treatment with folic acid, cobalamin, and pyridoxine does not reduce blood markers of inflammation, endothelial dysfunction, or hypercoagulability in patients with previous transient ischemic attack or stroke: A randomized substudy of the Vitatops. Stroke 2005, 36, 144–146. [Google Scholar] [CrossRef]
  120. Selhub, J.; Byun, A.; Liu, Z.; Mason, J.B.; Bronson, R.T.; Crott, J.W. Dietary vitamin B6 intake modulates colonic inflammation in the IL10-/- model of inflammatory bowel disease. J. Nutr. Biochem. 2013, 24, 2138–2143. [Google Scholar] [CrossRef][Green Version]
  121. Zhang, P.; Tsuchiya, K.; Kinoshita, T.; Kushiyama, H.; Suidasari, S.; Hatakeyama, M.; Imura, H.; Kat, N.; Suda, T. Vitamin B6 Prevents IL-1β Protein Production by Inhibiting NLRP3 Inflammasome Activation. J. Biol. Chem. 2016, 291, 24517–24527. [Google Scholar] [CrossRef][Green Version]
  122. Qian, B.; Shen, S.; Zhang, J.; Jing, P. Effects of Vitamin B6 Deficiency on the Composition and Functional Potential of T Cell Populations. J. Immunol. Res. 2017, 2017. [Google Scholar] [CrossRef] [PubMed][Green Version]
  123. Kobayashi, C.; Kurohane, K.; Imai, Y. High dose dietary pyridoxine induces T-helper type 1 polarization and decreases contact hypersensitivity response to fluorescein isothiocyanate in mice. Biol. Pharm. Bull. 2012, 35, 532–538. [Google Scholar] [CrossRef][Green Version]
  124. Katunuma, N.; Matsui, A.; Endo, K.; Hanba, J.; Sato, A.; Nakano, M.; Yuto, Y.; Tada, Y.; Asao, T.; Himeno, K.; et al. Inhibition of intracellular cathepsin activities and suppression of immune responses mediated by helper T lymphocyte type-2 by peroral or intraperitoneal administration of vitamin B6. Biochem. Biophys. Res. Commun. 2000, 272, 151–155. [Google Scholar] [CrossRef] [PubMed]
  125. Kumrungsee, T.; Zhang, P.; Chartkul, M.; Yanaka, N.; Kato, N. Potential Role of Vitamin B6 in Ameliorating the Severity of COVID-19 and Its Complications. Front. Nutr. 2020, 7, 562051. [Google Scholar] [CrossRef]
  126. León-Del-Río, A. Biotin in metabolism, gene expression, and human disease. J. Inherit. Metab. Dis. 2019, 42, 647–654. [Google Scholar] [CrossRef] [PubMed]
  127. Kuroishi, T. Regulation of immunological and inflammatory functions by biotin. Can. J. Physiol. Pharmacol. 2015, 93, 1091–1096. [Google Scholar] [CrossRef]
  128. Skupsky, J.; Sabui, S.; Hwang, M.; Nakasaki, M.; Cahalan, M.D.; Said, H.M. Biotin Supplementation Ameliorates Murine Colitis by Preventing NF-κB Activation. Cell. Mol. Gastroenterol. Hepatol. 2020, 9, 557–567. [Google Scholar] [CrossRef][Green Version]
  129. Elahi, A.; Sabui, S.; Narasappa, N.N.; Agrawal, S.; Lambrecht, N.W.; Agrawal, A.; Said, H.M. Biotin Deficiency Induces Th1- and Th17-Mediated Proinflammatory Responses in Human CD4 + T Lymphocytes via Activation of the mTOR Signaling Pathway. J. Immunol. 2018, 200, 2563–2570. [Google Scholar] [CrossRef] [PubMed][Green Version]
  130. Valentini, L.; Pinto, A.; Bourdel-Marchasson, I.; Ostan, R.; Brigidi, P.; Turroni, S.; Hrelia, S.; Hrelia, P.; Bereswill, S.; Fischer, A.; et al. Impact of personalized diet and probiotic supplementation on inflammation, nutritional parameters and intestinal microbiota—The “RISTOMED project”: Randomized controlled trial in healthy older people. Clin. Nutr. 2015, 34, 593–602. [Google Scholar] [CrossRef][Green Version]
  131. Morscher, R.J.; Ducker, G.S.; Li, S.H.J.; Mayer, J.A.; Gitai, Z.; Sperl, W.; Rabinowitz, J.D. Mitochondrial translation requires folate-dependent tRNA methylation. Nature 2018, 554, 128–132. [Google Scholar] [CrossRef]
  132. Kapadia, C.R. Vitamin B12 in health and disease. Part I—Inherited disorders of function, absorption, and transport. Gastroenterologist 1995, 3, 329–344. [Google Scholar]
  133. Kok, D.E.; Steegenga, W.T.; McKay, J.A. Folate and epigenetics: Why we should not forget bacterial biosynthesis. Epigenomics 2018, 10, 1147–1150. [Google Scholar] [CrossRef][Green Version]
  134. Bermingham, A.; Derrick, J.P. The folic acid biosynthesis pathway in bacteria: Evaluation of potential for antibacterial drug discovery. BioEssays 2002, 24, 637–648. [Google Scholar] [CrossRef] [PubMed]
  135. Martens, J.H.; Barg, H.; Warren, M.; Jahn, D. Microbial production of vitamin B12. Appl. Microbiol. Biotechnol. 2002, 58, 275–285. [Google Scholar] [CrossRef] [PubMed]
  136. LeBlanc, J.G.; Milani, C.; de Giori, G.S.; Sesma, F.; van Sinderen, D.; Ventura, M. Bacteria as vitamin suppliers to their host: A gut microbiota perspective. Curr. Opin. Biotechnol. 2013, 24, 160–168. [Google Scholar] [CrossRef] [PubMed]
  137. Fang, H.; Kang, J.; Zhang, D. Microbial production of vitamin B12: A review and future perspectives. Microb. Cell Fact. 2017, 16, 15. [Google Scholar] [CrossRef] [PubMed][Green Version]
  138. Corcoran, T.B.; O’Neill, M.P.; Webb, S.A.R.; Ho, K.M. Inflammation, vitamin deficiencies and organ failure in critically ill patients. Anaesth. Intensive Care 2009, 37, 740–747. [Google Scholar] [CrossRef][Green Version]
  139. Tal, S.; Shavit, Y.; Stern, F.; Malnick, S. Association between vitamin B12 levels and mortality in hospitalized older adults. J. Am. Geriatr. Soc. 2010, 58, 523–526. [Google Scholar] [CrossRef]
  140. Abe, I.; Shirato, K.; Hashizume, Y.; Mitsuhashi, R.; Kobayashi, A.; Shiono, C.; Sato, S.; Tachiyashiki, K.; Imaizumi, K. Folate-deficiency induced cell-specific changes in the distribution of lymphocytes and granulocytes in rats. Environ. Health Prev. Med. 2013, 18, 78–84. [Google Scholar] [CrossRef][Green Version]
  141. Courtemanche, C.; Huang, A.C.; Elson-Schwab, I.; Kerry, N.; Ng, B.Y.; Ames, B.N. Folate deficiency and ionizing radiation cause DNA breaks in primary human lymphocytes: A comparison. FASEB J. 2004, 18, 209–211. [Google Scholar] [CrossRef][Green Version]
  142. Socha-Banasiak, A.; Kamer, B.; Gach, A.; Wysocka, U.; Jakubowski, L.; Głowacka, E.; Czkwianianc, E. Folate status, regulatory T cells and MTHFR C677T polymorphism study in allergic children. Adv. Med. Sci. 2016, 61, 300–305. [Google Scholar] [CrossRef]
  143. Kunisawa, J.; Hashimoto, E.; Ishikawa, I.; Kiyono, H. A pivotal role of vitamin B9 in the maintenance of regulatory T cells in vitro and in vivo. PLoS ONE 2012, 7, e32094. [Google Scholar] [CrossRef] [PubMed][Green Version]
  144. Xia, W.; Hilgenbrink, A.R.; Matteson, E.L.; Lockwood, M.B.; Cheng, J.X.; Low, P.S. A functional folate receptor is induced during macrophage activation and can be used to target drugs to activated macrophages. Blood 2009, 113, 438–446. [Google Scholar] [CrossRef][Green Version]
  145. Hilgendorf, I.; Swirski, F.K. Folate Receptor: A Macrophage “Achilles’ Heel”? J. Am. Heart Assoc. 2012, 1, e004036. [Google Scholar] [CrossRef][Green Version]
  146. Poh, S.; Putt, K.S.; Low, P.S. Folate-Targeted Dendrimers Selectively Accumulate at Sites of Inflammation in Mouse Models of Ulcerative Colitis and Atherosclerosis. Biomacromolecules 2017, 18, 3082–3088. [Google Scholar] [CrossRef] [PubMed]
  147. Spiller, R. Serotonin, inflammation, and IBS: Fitting the jigsaw together? J. Pediatric Gastroenterol. Nutr. 2007, 45, S115–S119. [Google Scholar] [CrossRef] [PubMed]
  148. Heijtz, R.D.; Wang, S.; Anuar, F.; Qian, Y.; Björkholm, B.; Samuelsson, A.; Hibberd, M.L.; Forssberg, H.; Pettersson, S. Normal gut microbiota modulates brain development and behavior. Proc. Natl. Acad. Sci. USA 2011, 108, 3047–3052. [Google Scholar] [CrossRef][Green Version]
  149. Marcobal, A.; Kashyap, P.C.; Nelson, T.A.; Aronov, P.A.; Donia, M.S.; Spormann, A.; Fischbach, M.A.; Sonnenburg, J.L. A metabolomic view of how the human gut microbiota impacts the host metabolome using humanized and gnotobiotic mice. ISME J. 2013, 7, 1933–1943. [Google Scholar] [CrossRef][Green Version]
  150. Williams, B.B.; Van Benschoten, A.H.; Cimermancic, P.; Donia, M.S.; Zimmermann, M.; Taketani, M.; Ishihara, A.; Kashyap, P.C.; Fraser, J.S.; Fischbach, M.A. Discovery and characterization of gut microbiota decarboxylases that can produce the neurotransmitter tryptamine. Cell Host Microbe 2014, 16, 495–503. [Google Scholar] [CrossRef][Green Version]
  151. Agus, A.; Planchais, J.; Sokol, H. Gut Microbiota Regulation of Tryptophan Metabolism in Health and Disease. Cell Host Microbe 2018, 23, 716–724. [Google Scholar] [CrossRef][Green Version]
  152. Roager, H.M.; Licht, T.R. Microbial tryptophan catabolites in health and disease. Nat. Commun. 2018, 9, 1–10. [Google Scholar] [CrossRef] [PubMed][Green Version]
  153. Gao, J.; Xu, K.; Liu, H.; Liu, G.; Bai, M.; Peng, C.; Li, T.; Yin, Y. Impact of the gut microbiota on intestinal immunity mediated by tryptophan metabolism. Front. Cell. Infect. Microbiol. 2018, 8, 13. [Google Scholar] [CrossRef][Green Version]
  154. Zelante, T.; Iannitti, R.G.; Cunha, C.; DeLuca, A.; Giovannini, G.; Pieraccini, G.; Zecchi, R.; D’Angelo, C.; Massi-Benedetti, C.; Fallarino, F.; et al. Tryptophan catabolites from microbiota engage aryl hydrocarbon receptor and balance mucosal reactivity via interleukin-22. Immunity 2013, 39, 372–385. [Google Scholar] [CrossRef][Green Version]
  155. Venkatesh, M.; Mukherjee, S.; Wang, H.; Li, H.; Sun, K.; Benechet, A.P.; Qiu, Z.; Maher, L.; Redinbo, M.R.; Phillips, R.S.; et al. Symbiotic bacterial metabolites regulate gastrointestinal barrier function via the xenobiotic sensor PXR and toll-like receptor 4. Immunity 2014, 41, 296–310. [Google Scholar] [CrossRef] [PubMed][Green Version]
  156. Dodd, D.; Spitzer, M.H.; Van Treuren, W.; Merrill, B.D.; Hryckowian, A.J.; Higginbottom, S.K.; Le, A.; Cowan, T.M.; Nolan, G.P.; Fischbach, M.A.; et al. A gut bacterial pathway metabolizes aromatic amino acids into nine circulating metabolites. Nature 2017, 551, 648–652. [Google Scholar] [CrossRef] [PubMed]
  157. Alexeev, E.E.; Lanis, J.M.; Kao, D.J.; Campbell, E.L.; Kelly, C.J.; Battista, K.D.; Gerich, M.E.; Jenkins, B.R.; Walk, S.T.; Kominsky, D.J.; et al. Microbiota-Derived Indole Metabolites Promote Human and Murine Intestinal Homeostasis through Regulation of Interleukin-10 Receptor. Am. J. Pathol. 2018, 188, 1183–1194. [Google Scholar] [CrossRef] [PubMed][Green Version]
  158. Li, J.; Zhang, L.; Wu, T.; Li, Y.; Zhou, X.; Ruan, Z. Indole-3-propionic Acid Improved the Intestinal Barrier by Enhancing Epithelial Barrier and Mucus Barrier. J. Agric. Food Chem. 2021, 69, 1487–1495. [Google Scholar] [CrossRef]
  159. Wlodarska, M.; Luo, C.; Kolde, R.; d’Hennezel, E.; Annand, J.W.; Heim, C.E.; Krastel, P.; Schmitt, E.K.; Omar, A.S.; Creasey, E.A.; et al. Indoleacrylic Acid Produced by Commensal Peptostreptococcus Species Suppresses Inflammation. Cell Host Microbe 2017, 22, 25–37.e6. [Google Scholar] [CrossRef] [PubMed][Green Version]
  160. Marsland, B.J. Regulating inflammation with microbial metabolites. Nat. Med. 2016, 22, 581–583. [Google Scholar] [CrossRef]
  161. Huxtable, R.J. Physiological actions of taurine. Physiol. Rev. 1992, 72, 101–164. [Google Scholar] [CrossRef] [PubMed][Green Version]
  162. Sturman, J.A. Taurine in development. Physiol. Rev. 1993, 73, 119–148. [Google Scholar] [CrossRef]
  163. Trachtman, H.; Barbour, R.; Sturman, J.A.; Finberg, L. Taurine and osmoregulation: Taurine is a cerebral osmoprotective molecule in chronic hypernatremic dehydration. Pediatr. Res. 1988, 23, 35–39. [Google Scholar] [CrossRef] [PubMed][Green Version]
  164. Smith, B.F. Hepatology: A Textbook of Liver Disease. Arch. Intern. Med. 1983, 143, 2238. [Google Scholar] [CrossRef]
  165. Hardison, W.G.M. Hepatic Taurine Concentration and Dietary Taurine as Regulators of Bile Acid Conjugation with Taurine. Gastroenterology 1978, 75, 71–75. [Google Scholar] [CrossRef]
  166. Sjövall, J. Dietary Glycine and Taurine on Bile Acid Conjugation in Man. Bile Acids and Steroids 75. Proc. Soc. Exp. Biol. Med. 1959, 100, 676–678. [Google Scholar] [CrossRef]
  167. Fang, H.; Meng, F.; Piao, F.; Jin, B.; Li, M.; Li, W. Effect of Taurine on Intestinal Microbiota and Immune Cells in Peyer’s Patches of Immunosuppressive Mice. In Advances in Experimental Medicine and Biology; Springer: New York, NY, USA, 2019; Volume 1155, pp. 13–24. [Google Scholar]
  168. Levy, M.; Thaiss, C.A.; Zeevi, D.; Dohnalová, L.; Zilberman-Schapira, G.; Mahdi, J.A.; David, E.; Savidor, A.; Korem, T.; Herzig, Y.; et al. Microbiota-Modulated Metabolites Shape the Intestinal Microenvironment by Regulating NLRP6 Inflammasome Signaling. Cell 2015, 163, 1428–1443. [Google Scholar] [CrossRef] [PubMed][Green Version]
  169. Aronov, P.A.; Luo, F.J.G.; Plummer, N.S.; Quan, Z.; Holmes, S.; Hostetter, T.H.; Meyer, T.W. Colonic contribution to uremic solutes. J. Am. Soc. Nephrol. 2011, 22, 1769–1776. [Google Scholar] [CrossRef][Green Version]
  170. Saito, Y.; Sato, T.; Nomoto, K.; Tsuji, H. Identification of phenol- and p-cresol-producing intestinal bacteria by using media supplemented with tyrosine and its metabolites. FEMS Microbiol. Ecol. 2018, 94, 125. [Google Scholar] [CrossRef]
  171. Gryp, T.; Vanholder, R.; Vaneechoutte, M.; Glorieux, G. P-Cresyl Sulfate. Toxins 2017, 9, 52. [Google Scholar] [CrossRef][Green Version]
  172. Dalrymple, L.S.; Go, A.S. Epidemiology of acute infections among patients with chronic kidney disease. Clin. J. Am. Soc. Nephrol. 2008, 3, 1487–1493. [Google Scholar] [CrossRef][Green Version]
  173. Azevedo, M.L.V.; Bonan, N.B.; Dias, G.; Brehm, F.; Steiner, T.M.; Souza, W.M.; Stinghen, A.E.M.; Barreto, F.C.; Elifio-Esposito, S.; Pecoits-Filho, R.; et al. p-Cresyl sulfate affects the oxidative burst, phagocytosis process, and antigen presentation of monocyte-derived macrophages. Toxicol. Lett. 2016, 263, 1–5. [Google Scholar] [CrossRef]
  174. Shiba, T.; Makino, I.; Sasaki, T.; Fukuhara, Y.; Kawakami, K.; Kato, I.; Kobayashi, T. p-Cresyl sulfate decreases peripheral B cells in mice with adenine-induced renal dysfunction. Toxicol. Appl. Pharmacol. 2018, 342, 50–59. [Google Scholar] [CrossRef]
  175. Shiba, T.; Kawakami, K.; Sasaki, T.; Makino, I.; Kato, I.; Kobayashi, T.; Uchida, K.; Kaneko, K. Effects of intestinal bacteria-derived p-cresyl sulfate on Th1-type immune response in vivo and in vitro. Toxicol. Appl. Pharmacol. 2014, 274, 191–199. [Google Scholar] [CrossRef]
  176. Vanholder, R.; Bammens, B.; De Loor, H.; Glorieux, G.; Meijers, B.; Schepers, E.; Massy, Z.; Evenepoel, P. Warning: The unfortunate end of p-cresol as a uraemic toxin. Nephrol. Dial. Transplant. 2011, 26, 1464–1467. [Google Scholar] [CrossRef] [PubMed][Green Version]
  177. Sudo, N. Biogenic Amines: Signals Between Commensal Microbiota and Gut Physiology. Front. Endocrinol. 2019, 10, 504. [Google Scholar] [CrossRef] [PubMed][Green Version]
  178. Jutel, M.; Watanabe, T.; Klunker, S.; Akdis, M.; Thomet, O.A.R.; Malolepszy, J.; Zak-Nejmark, T.; Koga, R.; Kobayashi, T.; Blaser, K.; et al. Histamine regulates T-cell and antibody responses by differential expression of H1 and H2 receptors. Nature 2001, 413, 420–425. [Google Scholar] [CrossRef][Green Version]
  179. Yoshimoto, T.; Tsutsui, H.; Tominaga, K.; Hoshino, K.; Okamura, H.; Akira, S.; Paul, W.E.; Nakanishi, K. IL-18, although antiallergic when administered with IL-12, stimulates IL-4 and histamine release by basophils. Proc. Natl. Acad. Sci. USA 1999, 96, 13962–13966. [Google Scholar] [CrossRef][Green Version]
  180. BEAVER, M.H.; WOSTMANN, B.S. Histamine and 5-Hydroxytryptamine in the Intestinal Tract of Germ-Free Animals, Animals Harbouring One Microbial Species and Conventional Animals. Br. J. Pharmacol. Chemother. 1962, 19, 385–393. [Google Scholar] [CrossRef][Green Version]
  181. Barcik, W.; Pugin, B.; Westermann, P.; Perez, N.R.; Ferstl, R.; Wawrzyniak, M.; Smolinska, S.; Jutel, M.; Hessel, E.M.; Michalovich, D.; et al. Histamine-secreting microbes are increased in the gut of adult asthma patients. J. Allergy Clin. Immunol. 2016, 138, 1491–1494.e7. [Google Scholar] [CrossRef] [PubMed][Green Version]
  182. Caron, G.; Delneste, Y.; Roelandts, E.; Duez, C.; Herbault, N.; Magistrelli, G.; Bonnefoy, J.-Y.; Pestel, J.; Jeannin, P. Histamine Induces CD86 Expression and Chemokine Production by Human Immature Dendritic Cells. J. Immunol. 2001, 166, 6000–6006. [Google Scholar] [CrossRef] [PubMed]
  183. Mazzoni, A.; Young, H.A.; Spitzer, J.H.; Visintin, A.; Segal, D.M. Histamine regulates cytokine production in maturing dendritic cells, resulting in altered T cell polarization. J. Clin. Investig. 2001, 108, 1865–1873. [Google Scholar] [CrossRef] [PubMed]
  184. Van Der Pouw Kraan, T.C.T.M.; Snijders, A.; Boeije, L.C.M.; De Groot, E.R.; Alewijnse, A.E.; Leurs, R.; Aarden, L.A. Histamine inhibits the production of interleukin-12 through interaction with H2 receptors. J. Clin. Investig. 1998, 102, 1866–1873. [Google Scholar] [CrossRef][Green Version]
  185. Noubade, R.; Milligan, G.; Zachary, J.F.; Blankenhorn, E.P.; Del Rio, R.; Rincon, M.; Teuscher, C. Histamine receptor H1 is required for TCR-mediated p38 MAPK activation and optimal IFN-γ production in mice. J. Clin. Investig. 2007, 117, 3507–3518. [Google Scholar] [CrossRef][Green Version]
  186. Barcik, W.; Pugin, B.; Brescó, M.S.; Westermann, P.; Rinaldi, A.; Groeger, D.; Van Elst, D.; Sokolowska, M.; Krawczyk, K.; Frei, R.; et al. Bacterial secretion of histamine within the gut influences immune responses within the lung. Allergy Eur. J. Allergy Clin. Immunol. 2019, 74, 899–909. [Google Scholar] [CrossRef] [PubMed]
  187. Igarashi, K.; Kashiwagi, K. Modulation of cellular function by polyamines. Int. J. Biochem. Cell Biol. 2010, 42, 39–51. [Google Scholar] [CrossRef] [PubMed]
  188. Milovic, V. Polyamines in the gut lumen: Bioavailability and biodistribution. Eur. J. Gastroenterol. Hepatol. 2001, 13, 1021–1025. [Google Scholar] [CrossRef] [PubMed]
  189. Ramos-Molina, B.; Queipo-Ortuño, M.I.; Lambertos, A.; Tinahones, F.J.; Peñafiel, R. Dietary and gut microbiota polyamines in obesity- And age-related diseases. Front. Nutr. 2019, 6, 1–15. [Google Scholar] [CrossRef][Green Version]
  190. Nakamura, A.; Ooga, T.; Matsumoto, M. Intestinal luminal putrescine is produced by collective biosynthetic pathways of the commensal microbiome. Gut Microbes 2019, 10, 159–171. [Google Scholar] [CrossRef][Green Version]
  191. Matsumoto, M.; Benno, Y. The relationship between microbiota and polyamine concentration in the human intestine: A pilot study. Microbiol. Immunol. 2007, 51, 25–35. [Google Scholar] [CrossRef] [PubMed][Green Version]
  192. Zhang, M.; Borovikova, L.V.; Wang, H.; Metz, C.; Tracey, K.J. Spermine inhibition of monocyte activation and inflammation. Mol. Med. 1999, 5, 595–605. [Google Scholar] [CrossRef]
  193. Zhang, M.; Caragine, T.; Wang, H.; Cohen, P.S.; Botchkina, G.; Soda, K.; Bianchi, M.; Ulrich, P.; Cerami, A.; Sherry, B.; et al. Spermine inhibits proinflammatory cytokine synthesis in human mononuclear cells: A counterregulatory mechanism that restrains the immune response. J. Exp. Med. 1997, 185, 1759–1768. [Google Scholar] [CrossRef][Green Version]
  194. Zhu, S.; Ashok, M.; Li, J.; Li, W.; Yang, H.; Wang, P.; Tracey, K.J.; Sama, A.E.; Wang, H. Spermine protects mice against lethal sepsis partly by attenuating surrogate inflammatory markers. Mol. Med. 2009, 15, 275–282. [Google Scholar] [CrossRef] [PubMed]
  195. Soda, K.; Dobashi, Y.; Kano, Y.; Tsujinaka, S.; Konishi, F. Polyamine-rich food decreases age-associated pathology and mortality in aged mice. Exp. Gerontol. 2009, 44, 727–732. [Google Scholar] [CrossRef] [PubMed]
  196. Eisenberg, T.; Knauer, H.; Schauer, A.; Büttner, S.; Ruckenstuhl, C.; Carmona-Gutierrez, D.; Ring, J.; Schroeder, S.; Magnes, C.; Antonacci, L.; et al. Induction of autophagy by spermidine promotes longevity. Nat. Cell Biol. 2009, 11, 1305–1314. [Google Scholar] [CrossRef] [PubMed]
  197. Kibe, R.; Kurihara, S.; Sakai, Y.; Suzuki, H.; Ooga, T.; Sawaki, E.; Muramatsu, K.; Nakamura, A.; Yamashita, A.; Kitada, Y.; et al. Upregulation of colonic luminal polyamines produced by intestinal microbiota delays senescence in mice. Sci. Rep. 2014, 4, 1–11. [Google Scholar] [CrossRef] [PubMed]
  198. Eisenberg, T.; Abdellatif, M.; Schroeder, S.; Primessnig, U.; Stekovic, S.; Pendl, T.; Harger, A.; Schipke, J.; Zimmermann, A.; Schmidt, A.; et al. Cardioprotection and lifespan extension by the natural polyamine spermidine. Nat. Med. 2016, 22, 1428–1438. [Google Scholar] [CrossRef]
  199. Madeo, F.; Eisenberg, T.; Pietrocola, F.; Kroemer, G. Spermidine in health and disease. Science 2018, 359, 6374. [Google Scholar] [CrossRef][Green Version]
  200. Carriche, G.M.; Almeida, L.; Stüve, P.; Velasquez, L.; Dhillon-LaBrooy, A.; Roy, U.; Lindenberg, M.; Strowig, T.; Plaza-Sirvent, C.; Schmitz, I.; et al. Regulating T-cell differentiation through the polyamine spermidine. J. Allergy Clin. Immunol. 2021, 147, 335–348.e11. [Google Scholar] [CrossRef]
  201. Morón, B.; Spalinger, M.; Kasper, S.; Atrott, K.; Frey-Wagner, I.; Fried, M.; McCole, D.F.; Rogler, G.; Scharl, M. Activation of Protein Tyrosine Phosphatase Non-Receptor Type 2 by Spermidine Exerts Anti-Inflammatory Effects in Human THP-1 Monocytes and in a Mouse Model of Acute Colitis. PLoS ONE 2013, 8, e73703. [Google Scholar] [CrossRef][Green Version]
  202. Madeo, F.; Bauer, M.A.; Carmona-Gutierrez, D.; Kroemer, G. Spermidine: A physiological autophagy inducer acting as an anti-aging vitamin in humans? Autophagy 2019, 15, 165–168. [Google Scholar] [CrossRef]
  203. Wirth, M.; Schwarz, C.; Benson, G.; Horn, N.; Buchert, R.; Lange, C.; Köbe, T.; Hetzer, S.; Maglione, M.; Michael, E.; et al. Effects of spermidine supplementation on cognition and biomarkers in older adults with subjective cognitive decline (SmartAge)—Study protocol for a randomized controlled trial. Alzheimer’s Res. Ther. 2019, 11, 36. [Google Scholar] [CrossRef]
  204. Wolosker, H.; Dumin, E.; Balan, L.; Foltyn, V.N. D-amino acids in the brain: D-serine in neurotransmission and neurodegeneration. FEBS J. 2008, 275, 3514–3526. [Google Scholar] [CrossRef]
  205. Cava, F.; Lam, H.; De Pedro, M.A.; Waldor, M.K. Emerging knowledge of regulatory roles of d-amino acids in bacteria. Cell. Mol. Life Sci. 2011, 68, 817–831. [Google Scholar] [CrossRef][Green Version]
  206. Sasabe, J.; Miyoshi, Y.; Rakoff-Nahoum, S.; Zhang, T.; Mita, M.; Davis, B.M.; Hamase, K.; Waldor, M.K. Interplay between microbial d-amino acids and host d-amino acid oxidase modifies murine mucosal defence and gut microbiota. Nat. Microbiol. 2016, 1, 1–7. [Google Scholar] [CrossRef]
  207. Sasabe, J.; Suzuki, M. Emerging role of D-Amino acid metabolism in the innate defense. Front. Microbiol. 2018, 9, 933. [Google Scholar] [CrossRef][Green Version]
  208. Yunes, R.A.; Poluektova, E.U.; Dyachkova, M.S.; Klimina, K.M.; Kovtun, A.S.; Averina, O.V.; Orlova, V.S.; Danilenko, V.N. GABA production and structure of gadB/gadC genes in Lactobacillus and Bifidobacterium strains from human microbiota. Anaerobe 2016, 42, 197–204. [Google Scholar] [CrossRef] [PubMed]
  209. Kurihara, S.; Oda, S.; Tsuboi, Y.; Hyeon, G.K.; Oshida, M.; Kumagai, H.; Suzuki, H. γ-glutamylputrescine synthetase in the putrescine utilization pathway of Escherichia coli K-12. J. Biol. Chem. 2008, 283, 19981–19990. [Google Scholar] [CrossRef] [PubMed][Green Version]
  210. Barrett, E.; Ross, R.P.; O’Toole, P.W.; Fitzgerald, G.F.; Stanton, C. γ-Aminobutyric acid production by culturable bacteria from the human intestine. J. Appl. Microbiol. 2012, 113, 411–417. [Google Scholar] [CrossRef] [PubMed]
  211. Matsumoto, M.; Ooga, T.; Kibe, R.; Aiba, Y.; Koga, Y.; Benno, Y. Colonic absorption of low-molecular-weight metabolites influenced by the intestinal microbiome: A pilot study. PLoS ONE 2017, 12, e0169207. [Google Scholar] [CrossRef]
  212. Ren, W.; Yin, J.; Xiao, H.; Chen, S.; Liu, G.; Tan, B.; Li, N.; Peng, Y.; Li, T.; Zeng, B.; et al. Intestinal microbiota-derived GABA mediates interleukin-17 expression during enterotoxigenic Escherichia coli infection. Front. Immunol. 2017, 7, 685. [Google Scholar] [CrossRef][Green Version]
  213. Braun, H.S.; Sponder, G.; Pieper, R.; Aschenbach, J.R.; Deiner, C. GABA selectively increases mucin-1 expression in isolated pig jejunum. Genes Nutr. 2015, 10, 1–8. [Google Scholar] [CrossRef]
  214. Bajic, S.S.; Djokic, J.; Dinic, M.; Veljovic, K.; Golic, N.; Mihajlovic, S.; Tolinacki, M. GABA-producing natural dairy isolate from artisanal zlatar cheese attenuates gut inflammation and strengthens gut epithelial barrier in vitro. Front. Microbiol. 2019, 10, 527. [Google Scholar] [CrossRef]
  215. Thompson, J.A.; Oliveira, R.A.; Xavier, K.B. Chemical conversations in the gut microbiota. Gut Microbes 2016, 7, 163–170. [Google Scholar] [CrossRef] [PubMed][Green Version]
  216. Papenfort, K.; Bassler, B.L. Quorum sensing signal-response systems in Gram-negative bacteria. Nat. Rev. Microbiol. 2016, 14, 576–588. [Google Scholar] [CrossRef]
  217. Bhatt, V.S. Quorum sensing mechanisms in gram positive bacteria. In Implication of Quorum Sensing System in Biofilm Formation and Virulence; Springer: Singapore, 2019; pp. 297–311. ISBN 9789811324291. [Google Scholar]
  218. Xavier, K.B.; Bassler, B.L. Interference with AI-2-mediated bacterial cell-cell communication. Nature 2005, 437, 750–753. [Google Scholar] [CrossRef] [PubMed][Green Version]
  219. Thompson, J.A.; Oliveira, R.A.; Djukovic, A.; Ubeda, C.; Xavier, K.B. Manipulation of the quorum sensing signal AI-2 affects the antibiotic-treated gut microbiota. Cell Rep. 2015, 10, 1861–1871. [Google Scholar] [CrossRef]
  220. Li, Q.; Ren, Y.; Fu, X. Inter-kingdom signaling between gut microbiota and their host. Cell. Mol. Life Sci. 2019, 76, 2383–2389. [Google Scholar] [CrossRef] [PubMed]
  221. Pundir, P.; Liu, R.; Vasavda, C.; Serhan, N.; Limjunyawong, N.; Yee, R.; Zhan, Y.; Dong, X.; Wu, X.; Zhang, Y.; et al. A Connective Tissue Mast-Cell-Specific Receptor Detects Bacterial Quorum-Sensing Molecules and Mediates Antibacterial Immunity. Cell Host Microbe 2019, 26, 114–122.e8. [Google Scholar] [CrossRef]
  222. Li, Q.; Peng, W.; Wu, J.; Wang, X.; Ren, Y.; Li, H.; Peng, Y.; Tang, X.; Fu, X. Autoinducer-2 of gut microbiota, a potential novel marker for human colorectal cancer, is associated with the activation of TNFSF9 signaling in macrophages. Oncoimmunology 2019, 8, e1626192. [Google Scholar] [CrossRef]
  223. Goldstein, D.S.; Eisenhofer, G.; Kopin, I.J. Sources and significance of plasma levels of catechols and their metabolites in humans. J. Pharmacol. Exp. Ther. 2003, 305, 800–811. [Google Scholar] [CrossRef][Green Version]
  224. Flierl, M.A.; Rittirsch, D.; Nadeau, B.A.; Chen, A.J.; Sarma, J.V.; Zetoune, F.S.; McGuire, S.R.; List, R.P.; Day, D.E.; Hoesel, L.M.; et al. Phagocyte-derived catecholamines enhance acute inflammatory injury. Nature 2007, 449, 721–725. [Google Scholar] [CrossRef][Green Version]
  225. Costa, M.; Brookes, S.J.H.; Hennig, G.W. Anatomy and physiology of the enteric nervous system. Gut 2000, 47, iv15–iv19. [Google Scholar] [CrossRef][Green Version]
  226. Eisenhofer, G.; Aneman, Å.; Friberg, P.; Hooper, D.; Fåndriks, L.; Lonroth, H.; Hunyady, B.; Mezey, E. Substantial production of Dopamine in the human gastrointestinal tract. J. Clin. Endocrinol. Metab. 1997, 82, 3864–3871. [Google Scholar] [CrossRef] [PubMed]
  227. Furness, J.B. The Enteric Nervous System. Available online: https://www.wiley.com/en-us/The+Enteric+Nervous+System-p-9781405133760 (accessed on 14 May 2021).
  228. Sandrini, S.; Aldriwesh, M.; Alruways, M.; Freestone, P. Microbial endocrinology: Host-bacteria communication within the gut microbiome. J. Endocrinol. 2015, 225, R21–R34. [Google Scholar] [CrossRef][Green Version]
  229. Patel, P.; Nankova, B.B.; LaGamma, E.F. Butyrate, a gut-derived environmental signal, regulates tyrosine hydroxylase gene expression via a novel promoter element. Dev. Brain Res. 2005, 160, 53–62. [Google Scholar] [CrossRef]
  230. Tsavkelova, E.A.; Klimova, S.Y.; Cherdyntseva, T.A.; Netrusov, A.I. Hormones and hormone-like substances of microorganisms: A review. Appl. Biochem. Microbiol. 2006, 42, 229–235. [Google Scholar] [CrossRef]
  231. Asano, Y.; Hiramoto, T.; Nishino, R.; Aiba, Y.; Kimura, T.; Yoshihara, K.; Koga, Y.; Sudo, N. Critical role of gut microbiota in the production of biologically active, free catecholamines in the gut lumen of mice. Am. J. Physiol.-Gastrointest. Liver Physiol. 2012, 303, 1288–1295. [Google Scholar] [CrossRef][Green Version]
  232. Eisenhofer, G.; Kopin, I.J.; Goldstein, D.S. Catecholamine metabolism: A contemporary view with implications for physiology and medicine. Pharmacol. Rev. 2004, 56, 331–349. [Google Scholar] [CrossRef] [PubMed]
  233. Lyte, M.; Ernst, S. Alpha and beta adrenergic receptor involvement in catecholamine-induced growth of gram-negative bacteria. Biochem. Biophys. Res. Commun. 1993, 190, 447–452. [Google Scholar] [CrossRef]
  234. Lyte, M.; Ernst, S. Catecholamine induced growth of gram negative bacteria. Life Sci. 1992, 50, 203–212. [Google Scholar] [CrossRef]
  235. Sperandio, V.; Torres, A.G.; Jarvis, B.; Nataro, J.P.; Kaper, J.B. Bacteria-host communication: The language of hormones. Proc. Natl. Acad. Sci. USA 2003, 100, 8951–8956. [Google Scholar] [CrossRef] [PubMed][Green Version]
  236. Clarke, M.B.; Hughes, D.T.; Zhu, C.; Boedeker, E.C.; Sperandio, V. The QseC sensor kinase: A bacterial adrenergic receptor. Proc. Natl. Acad. Sci. USA 2006, 103, 10420–10425. [Google Scholar] [CrossRef][Green Version]
  237. Bearson, B.L.; Bearson, S.M.D. The role of the QseC quorum-sensing sensor kinase in colonization and norepinephrine-enhanced motility of Salmonella enterica serovar Typhimurium. Microb. Pathog. 2008, 44, 271–278. [Google Scholar] [CrossRef] [PubMed]
  238. Moreira, C.G.; Weinshenker, D.; Sperandio, V. QseC mediates Salmonella enterica serovar typhimurium virulence in vitro and in vivo. Infect. Immun. 2010, 78, 914–926. [Google Scholar] [CrossRef][Green Version]
  239. Gart, E.V.; Suchodolski, J.S.; Welsh, T.H.; Alaniz, R.C.; Randel, R.D.; Lawhon, S.D. Salmonella typhimurium and multidirectional communication in the gut. Front. Microbiol. 2016, 7, 1827. [Google Scholar] [CrossRef] [PubMed][Green Version]
  240. Madden, K.S.; Sanders, V.M.; Felten, D.L. Catecholamine influences and sympathetic neural modulation of immune responsiveness. Annu. Rev. Pharmacol. Toxicol. 1995, 35, 417–448. [Google Scholar] [CrossRef] [PubMed]
  241. Sanders, V.M.; Straub, R.H. Norepinephrine, the β-adrenergic receptor, and immunity. Brain. Behav. Immun. 2002, 16, 290–332. [Google Scholar] [CrossRef] [PubMed]
  242. Ottaway, C.A.; Husband, A.J. The influence of neuroendocrine pathways on lymphocyte migration. Immunol. Today 1994, 15, 511–517. [Google Scholar] [CrossRef]
  243. Elenkov, I.J.; Wilder, R.L.; Chrousos, G.P.; Vizi, E.S. The sympathetic nerve—An integrative interface between two supersystems: The brain and the immune system. Pharmacol. Rev. 2000, 52, 595–638. [Google Scholar]
  244. Flierl, M.A.; Rittirsch, D.; Huber-Lang, M.; Vidya Sarma, J.; Award, P. Catecholamines—Crafty weapons in the inflammatory arsenal of immune/inflammatory cells or opening Pandora’s box§? Mol. Med. 2008, 14, 195–204. [Google Scholar] [CrossRef] [PubMed]
  245. Brown, S.W.; Meyers, R.T.; Brennan, K.M.; Rumble, J.M.; Narasimhachari, N.; Perozzi, E.F.; Ryan, J.J.; Stewart, J.K.; Fischer-Stenger, K. Catecholamines in a macrophage cell line. J. Neuroimmunol. 2003, 135, 47–55. [Google Scholar] [CrossRef]
  246. Lyte, M.; Bailey, M.T. Neuroendocrine-bacterial interactions in a neurotoxin-induced model of trauma. J. Surg. Res. 1997, 70, 195–201. [Google Scholar] [CrossRef]
  247. Corrigan, R.M.; Gründling, A. Cyclic di-AMP: Another second messenger enters the fray. Nat. Rev. Microbiol. 2013, 11, 513–524. [Google Scholar] [CrossRef]
  248. Woodward, J.J.; Lavarone, A.T.; Portnoy, D.A. C-di-AMP secreted by intracellular Listeria monocytogenes activates a host type I interferon response. Science 2010, 328, 1703–1705. [Google Scholar] [CrossRef] [PubMed][Green Version]
  249. McFarland, A.P.; Luo, S.; Ahmed-Qadri, F.; Zuck, M.; Thayer, E.F.; Goo, Y.A.; Hybiske, K.; Tong, L.; Woodward, J.J. Sensing of Bacterial Cyclic Dinucleotides by the Oxidoreductase RECON Promotes NF-κB Activation and Shapes a Proinflammatory Antibacterial State. Immunity 2017, 46, 433–445. [Google Scholar] [CrossRef][Green Version]
  250. Whiteley, A.T.; Eaglesham, J.B.; de Oliveira Mann, C.C.; Morehouse, B.R.; Lowey, B.; Nieminen, E.A.; Danilchanka, O.; King, D.S.; Lee, A.S.Y.; Mekalanos, J.J.; et al. Bacterial cGAS-like enzymes synthesize diverse nucleotide signals. Nature 2019, 567, 194–199. [Google Scholar] [CrossRef]
  251. Mager, L.F.; Burkhard, R.; Pett, N.; Cooke, N.C.A.; Brown, K.; Ramay, H.; Paik, S.; Stagg, J.; Groves, R.A.; Gallo, M.; et al. Microbiome-derived inosine modulates response to checkpoint inhibitor immunotherapy. Science 2020, 369, 1481–1489. [Google Scholar] [CrossRef] [PubMed]
  252. Kim, I.; Ahn, S.H.; Inagaki, T.; Choi, M.; Ito, S.; Guo, G.L.; Kliewer, S.A.; Gonzalez, F.J. Differential regulation of bile acid homeostasis by the farnesoid X receptor in liver and intestine. J. Lipid Res. 2007, 48, 2664–2672. [Google Scholar] [CrossRef][Green Version]
  253. Fiorucci, S.; Biagioli, M.; Zampella, A.; Distrutti, E. Bile acids activated receptors regulate innate immunity. Front. Immunol. 2018, 9, 1. [Google Scholar] [CrossRef] [PubMed][Green Version]
  254. 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]
  255. Mencarelli, A.; Renga, B.; Migliorati, M.; Cipriani, S.; Distrutti, E.; Santucci, L.; Fiorucci, S. The Bile Acid Sensor Farnesoid X Receptor Is a Modulator of Liver Immunity in a Rodent Model of Acute Hepatitis. J. Immunol. 2009, 183, 6657–6666. [Google Scholar] [CrossRef] [PubMed][Green Version]
  256. Gadaleta, R.M.; Van Erpecum, K.J.; Oldenburg, B.; Willemsen, E.C.L.; Renooij, W.; Murzilli, S.; Klomp, L.W.J.; Siersema, P.D.; Schipper, M.E.I.; 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]
  257. Song, X.; Sun, X.; Oh, S.F.; Wu, M.; Zhang, Y.; Zheng, W.; Geva-Zatorsky, N.; Jupp, R.; Mathis, D.; Benoist, C.; et al. Microbial bile acid metabolites modulate gut RORγ+ regulatory T cell homeostasis. Nature 2020, 577, 410–415. [Google Scholar] [CrossRef]
  258. Yang, B.H.; Hagemann, S.; Mamareli, P.; Lauer, U.; Hoffmann, U.; Beckstette, M.; Föhse, L.; Prinz, I.; Pezoldt, J.; Suerbaum, S.; et al. Foxp3+ T cells expressing RORγt represent a stable regulatory T-cell effector lineage with enhanced suppressive capacity during intestinal inflammation. Mucosal Immunol. 2016, 9, 444–457. [Google Scholar] [CrossRef]
  259. Campbell, C.; McKenney, P.T.; Konstantinovsky, D.; Isaeva, O.I.; Schizas, M.; Verter, J.; Mai, C.; Jin, W.B.; Guo, C.J.; Violante, S.; et al. Bacterial metabolism of bile acids promotes generation of peripheral regulatory T cells. Nature 2020, 581, 475–479. [Google Scholar] [CrossRef] [PubMed]
  260. Hannun, Y.A.; Obeid, L.M. Sphingolipids and their metabolism in physiology and disease. Nat. Rev. Mol. Cell Biol. 2018, 19, 175–191. [Google Scholar] [CrossRef] [PubMed]
  261. MacEyka, M.; Spiegel, S. Sphingolipid metabolites in inflammatory disease. Nature 2014, 510, 58–67. [Google Scholar] [CrossRef][Green Version]
  262. Abdel Hadi, L.; Di Vito, C.; Riboni, L. Fostering Inflammatory Bowel Disease: Sphingolipid Strategies to Join Forces. Mediat. Inflamm. 2016, 2016. [Google Scholar] [CrossRef][Green Version]
  263. Stoffel, W.; Dittmar, K.; Wilmes, R. Sphingolipid Metabolism in Bacteroideaceae. Hoppe Seylers Z. Physiol. Chem. 1975, 356, 715–726. [Google Scholar] [CrossRef]
  264. LaBach, J.P.; White, D.C. Identification of ceramide phosphorylethanolamine and ceramide phosphorylglycerol in the lipids of an anaerobic bacterium. J. Lipid Res. 1969, 10, 528–534. [Google Scholar] [CrossRef]
  265. Ana, D.; Na, C.; Bielawski, J.; Hannun, Y.A.; Kasper, D.L. Membrane sphingolipids as essential molecular signals for Bacteroides survival in the intestine. Proc. Natl. Acad. Sci. USA 2011, 108, 4666–4671. [Google Scholar] [CrossRef][Green Version]
  266. Hickey, C.A.; Kuhn, K.A.; Donermeyer, D.L.; Porter, N.T.; Jin, C.; Cameron, E.A.; Jung, H.; Kaiko, G.E.; Wegorzewska, M.; Malvin, N.P.; et al. Colitogenic Bacteroides thetaiotaomicron antigens access host immune cells in a sulfatase-dependent manner via outer membrane vesicles. Cell Host Microbe 2015, 17, 672–680. [Google Scholar] [CrossRef][Green Version]
  267. Kinjo, Y.; Wu, D.; Kim, G.; Xing, G.W.; Poles, M.A.; Ho, D.D.; Tsuji, M.; Kawahara, K.; Wong, C.H.; Kronenberg, M. Recognition of bacterial glycosphingolipids by natural killer T cells. Nature 2005, 434, 520–525. [Google Scholar] [CrossRef]
  268. An, D.; Oh, S.F.; Olszak, T.; Neves, J.F.; Avci, F.Y.; Erturk-Hasdemir, D.; Lu, X.; Zeissig, S.; Blumberg, R.S.; Kasper, D.L. Sphingolipids from a symbiotic microbe regulate homeostasis of host intestinal natural killer T cells. Cell 2014, 156, 123–133. [Google Scholar] [CrossRef][Green Version]
  269. Olszak, T.; An, D.; Zeissig, S.; Vera, M.P.; Richter, J.; Franke, A.; Glickman, J.N.; Siebert, R.; Baron, R.M.; Kasper, D.L.; et al. Microbial exposure during early life has persistent effects on natural killer T cell function. Science 2012, 336, 489–493. [Google Scholar] [CrossRef] [PubMed][Green Version]
  270. Brown, E.M.; Ke, X.; Hitchcock, D.; Jeanfavre, S.; Avila-Pacheco, J.; Nakata, T.; Arthur, T.D.; Fornelos, N.; Heim, C.; Franzosa, E.A.; et al. Bacteroides-Derived Sphingolipids Are Critical for Maintaining Intestinal Homeostasis and Symbiosis. Cell Host Microbe 2019, 25, 668–680. [Google Scholar] [CrossRef] [PubMed]
  271. Cleveland, R.F.; Holtje, J.V.; Wicken, A.J.; Tomasz, A.; Daneo-Moore, L.; Shockman, G.D. Inhibition of bacterial wall lysins by lipoteichoic acids and related compounds. Biochem. Biophys. Res. Commun. 1975, 67, 1128–1135. [Google Scholar] [CrossRef]
  272. Fischer, W. Bacterial Phosphoglycolipids and Lipoteichoic Acids. In Glycolipids, Phosphoglycolipids, and Sulfoglycolipids; Springer: New York, NY, USA, 1990; pp. 123–234. [Google Scholar]
  273. Brown, S.; Santa Maria, J.P.; Walker, S. Wall teichoic acids of gram-positive bacteria. Annu. Rev. Microbiol. 2013, 67, 313–336. [Google Scholar] [CrossRef] [PubMed][Green Version]
  274. Vasselon, T.; Detmers, P.A. Toll receptors: A central element in innate immune responses. Infect. Immun. 2002, 70, 1033–1041. [Google Scholar] [CrossRef] [PubMed][Green Version]
  275. Draing, C.; Pfitzenmaier, M.; Zummo, S.; Mancuso, G.; Geyer, A.; Hartung, T.; Von Aulock, S. Comparison of lipoteichoic acid from different serotypes of Streptococcus pneumoniae. J. Biol. Chem. 2006, 281, 33849–33859. [Google Scholar] [CrossRef][Green Version]
  276. Schröder, N.W.J.; Morath, S.; Alexander, C.; Hamann, L.; Hartung, T.; Zähringer, U.; Göbel, U.B.; Weber, J.R.; Schumann, R.R. Lipoteichoic acid (LTA) of Streptococcus pneumoniae and Staphylococcus aureus activates immune cells via Toll-like receptor (TLR)-2, lipopolysaccharide-binding protein (LBP), and CD14, whereas TLR-4 and MD-2 are not involved. J. Biol. Chem. 2003, 278, 15587–15594. [Google Scholar] [CrossRef][Green Version]
  277. Trianiafilou, M.; Manukyan, M.; Mackie, A.; Morath, S.; Hartung, T.; Heine, H.; Triantafilou, K. Lipoteichoic acid and Toll-like receptor 2 internalization and targeting to the Golgi are lipid raft-dependent. J. Biol. Chem. 2004, 279, 40882–40889. [Google Scholar] [CrossRef][Green Version]
  278. Michelsen, K.S.; Aicher, A.; Mohaupt, M.; Hartung, T.; Dimmeler, S.; Kirschning, C.J.; Schumann, R.R. The role of toll-like receptors (TLRs) in bacteria-induced maturation of murine dendritic cells (DCs): Peptidoglycan and lipoteichoic acid are inducers of DC maturation and require TLR2. J. Biol. Chem. 2001, 276, 25680–25686. [Google Scholar] [CrossRef] [PubMed][Green Version]
  279. Hong, S.W.; Baik, J.E.; Kang, S.S.; Yun, C.H.; Seo, D.G.; Han, S.H. Lipoteichoic acid of Streptococcus mutans interacts with Toll-like receptor 2 through the lipid moiety for induction of inflammatory mediators in murine macrophages. Mol. Immunol. 2014, 57, 284–291. [Google Scholar] [CrossRef]
  280. Dunne, D.W.; Resnick, D.; Greenberg, J.; Krieger, M.; Joiner, K.A. The type I macrophage scavenger receptor binds to Gram-positive bacteria and recognizes lipoteichoic acid. Proc. Natl. Acad. Sci. USA 1994, 91, 1863–1867. [Google Scholar] [CrossRef] [PubMed][Green Version]
  281. Ellingsen, E.; Morath, S.; Flo, T.; Schromm, A.; Hartung, T.; Thiemermann, C.; Espevik, T.; Golenbock, D.; Foster, D.; Solberg, R.; et al. Induction of Cytokine Production in Human T Cells and Monocytes by Highly Purified Lipoteichoic Acid: Involvement of Toll-Like Receptors and CD14. Available online: https://www.medscimonit.com/download/index/idArt/420854 (accessed on 15 May 2021).
  282. von Aulock, S.; Morath, S.; Hareng, L.; Knapp, S.; van Kessel, K.P.M.; van Strijp, J.A.G.; Hartung, T. Lipoteichoic acid from Staphylococcus aureus is a potent stimulus for neutrophil recruitment. Immunobiology 2003, 208, 413–422. [Google Scholar] [CrossRef] [PubMed][Green Version]
  283. Wang, J.E.; Dahle, M.K.; McDonald, M.; Foster, S.J.; Aasen, A.O.; Thiemermann, C. Peptidoglycan and lipoteichoic acid in gram-positive bacterial sepsis: Receptors, signal transduction, biological effects, and synergism. Shock 2003, 20, 402–414. [Google Scholar] [CrossRef][Green Version]
  284. Hara, H.; Seregin, S.S.; Yang, D.; Fukase, K.; Chamaillard, M.; Alnemri, E.S.; Inohara, N.; Chen, G.Y.; Núñez, G. The NLRP6 Inflammasome Recognizes Lipoteichoic Acid and Regulates Gram-Positive Pathogen Infection. Cell 2018, 175, 1651–1664.e14. [Google Scholar] [CrossRef][Green Version]
  285. Delgado, S.; Sánchez, B.; Margolles, A.; Ruas-Madiedo, P.; Ruiz, L. Molecules produced by probiotics and intestinal microorganisms with immunomodulatory activity. Nutrients 2020, 12, 391. [Google Scholar] [CrossRef][Green Version]
  286. Riehl, T.E.; Alvarado, D.; Ee, X.; Zuckerman, A.; Foster, L.; Kapoor, V.; Thotala, D.; Ciorba, M.A.; Stenson, W.F. Lactobacillus rhamnosus GG protects the intestinal epithelium from radiation injury through release of lipoteichoic acid, macrophage activation and the migration of mesenchymal stem cells. Gut 2019, 68, 1003–1013. [Google Scholar] [CrossRef]
  287. Peña, J.A.; Versalovic, J. Lactobacillus rhamnosus GG decreases TNF-α production in lipopolysaccharide-activated murine macrophages by a contact-independent mechanism. Cell. Microbiol. 2003, 5, 277–285. [Google Scholar] [CrossRef] [PubMed]
  288. Wang, S.; Ahmadi, S.; Nagpal, R.; Jain, S.; Mishra, S.P.; Kavanagh, K.; Zhu, X.; Wang, Z.; McClain, D.A.; Kritchevsky, S.B.; et al. Lipoteichoic acid from the cell wall of a heat killed Lactobacillus paracasei D3-5 ameliorates aging-related leaky gut, inflammation and improves physical and cognitive functions: From C. elegans to mice. GeroScience 2020, 42, 333–352. [Google Scholar] [CrossRef]
  289. Grangette, C.; Nutten, S.; Palumbo, E.; Morath, S.; Hermann, C.; Dewulf, J.; Pot, B.; Hartung, T.; Hols, P.; Mercenier, A. Enhanced antiinflammatory capacity of a Lactobacillus plantarum mutant synthesizing modified teichoic acids. Proc. Natl. Acad. Sci. USA 2005, 102, 10321–10326. [Google Scholar] [CrossRef] [PubMed][Green Version]
  290. Melmed, G.; Thomas, L.S.; Lee, N.; Tesfay, S.Y.; Lukasek, K.; Michelsen, K.S.; Zhou, Y.; Hu, B.; Arditi, M.; Abreu, M.T. Human Intestinal Epithelial Cells Are Broadly Unresponsive to Toll-Like Receptor 2-Dependent Bacterial Ligands: Implications for Host-Microbial Interactions in the Gut. J. Immunol. 2003, 170, 1406–1415. [Google Scholar] [CrossRef][Green Version]
  291. Ott, S.J.; Musfeldt, M.; Wenderoth, D.F.; Hampe, J.; Brant, O.; Fölsch, U.R.; Timmis, K.N.; Schreiber, S. Reduction in diversity of the colonic mucosa associated bacterial microflora in patients with active inflammatory bowel disease. Gut 2004, 53, 685–693. [Google Scholar] [CrossRef] [PubMed][Green Version]
  292. Manichanh, C.; Rigottier-Gois, L.; Bonnaud, E.; Gloux, K.; Pelletier, E.; Frangeul, L.; Nalin, R.; Jarrin, C.; Chardon, P.; Marteau, P.; et al. Reduced diversity of faecal microbiota in Crohn’s disease revealed by a metagenomic approach. Gut 2006, 55, 205–211. [Google Scholar] [CrossRef][Green Version]
  293. Sokol, H.; Seksik, P.; Rigottier-Gois, L.; Lay, C.; Lepage, P.; Podglajen, I.; Marteau, P.; Doré, J. Specificities of the fecal microbiota in inflammatory bowel disease. Inflamm. Bowel Dis. 2006, 12, 106–111. [Google Scholar] [CrossRef]
  294. Roy, U.; Gálvez, E.J.C.; Iljazovic, A.; Lesker, T.R.; Błażejewski, A.J.; Pils, M.C.; Heise, U.; Huber, S.; Flavell, R.A.; Strowig, T. Distinct Microbial Communities Trigger Colitis Development upon Intestinal Barrier Damage via Innate or Adaptive Immune Cells. Cell Rep. 2017, 21, 994–1008. [Google Scholar] [CrossRef][Green Version]
  295. Schirmer, M.; Franzosa, E.A.; Lloyd-Price, J.; McIver, L.J.; Schwager, R.; Poon, T.W.; Ananthakrishnan, A.N.; Andrews, E.; Barron, G.; Lake, K.; et al. Dynamics of metatranscription in the inflammatory bowel disease gut microbiome. Nat. Microbiol. 2018, 3, 337–346. [Google Scholar] [CrossRef] [PubMed]
  296. Patwa, L.G.; Fan, T.J.; Tchaptchet, S.; Liu, Y.; Lussier, Y.A.; Sartor, R.B.; Hansen, J.J. Chronic intestinal inflammation induces stress-response genes in commensal Escherichia coli. Gastroenterology 2011, 141, 1842–1851. [Google Scholar] [CrossRef][Green Version]
  297. Ilott, N.E.; Bollrath, J.; Danne, C.; Schiering, C.; Shale, M.; Adelmann, K.; Krausgruber, T.; Heger, A.; Sims, D.; Powrie, F. Defining the microbial transcriptional response to colitis through integrated host and microbiome profiling. ISME J. 2016, 10, 2389–2404. [Google Scholar] [CrossRef]
  298. Becattini, S.; Sorbara, M.T.; Kim, S.G.; Littmann, E.L.; Dong, Q.; Walsh, G.; Wright, R.; Amoretti, L.; Fontana, E.; Hohl, T.M.; et al. Rapid transcriptional and metabolic adaptation of intestinal microbes to host immune activation. Cell Host Microbe 2021, 29, 378–393.e5. [Google Scholar] [CrossRef]
  299. Cash, H.L.; Whitham, C.V.; Behrendt, C.L.; Hooper, L.V. Symbiotic bacteria direct expression of an intestinal bactericidal lectin. Science 2006, 313, 1126–1130. [Google Scholar] [CrossRef][Green Version]
  300. Rubino, S.J.; Geddes, K.; Girardin, S.E. Innate IL-17 and IL-22 responses to enteric bacterial pathogens. Trends Immunol. 2012, 33, 112–118. [Google Scholar] [CrossRef]
  301. Bouskra, D.; Brézillon, C.; Bérard, M.; Werts, C.; Varona, R.; Boneca, I.G.; Eberl, G. Lymphoid tissue genesis induced by commensals through NOD1 regulates intestinal homeostasis. Nature 2008, 456, 507–510. [Google Scholar] [CrossRef] [PubMed]
  302. Salzman, N.H.; Bevins, C.L. Dysbiosis-A consequence of Paneth cell dysfunction. Semin. Immunol. 2013, 25, 334–341. [Google Scholar] [CrossRef]
  303. Ehmann, D.; Wendler, J.; Koeninger, L.; Larsen, I.S.; Klag, T.; Berger, J.; Marette, A.; Schaller, M.; Stange, E.F.; Malek, N.P.; et al. Paneth cell α-defensins HD-5 and HD-6 display differential degradation into active antimicrobial fragments. Proc. Natl. Acad. Sci. USA 2019, 116, 3746–3751. [Google Scholar] [CrossRef][Green Version]
  304. Bantug, G.R.; Galluzzi, L.; Kroemer, G.; Hess, C. The spectrum of T cell metabolism in health and disease. Nat. Rev. Immunol. 2018, 18, 19–34. [Google Scholar] [CrossRef] [PubMed]
  305. Deaglio, S.; Dwyer, K.M.; Gao, W.; Friedman, D.; Usheva, A.; Erat, A.; Chen, J.F.; Enjyoji, K.; Linden, J.; Oukka, M.; et al. Adenosine generation catalyzed by CD39 and CD73 expressed on regulatory T cells mediates immune suppression. J. Exp. Med. 2007, 204, 1257–1265. [Google Scholar] [CrossRef] [PubMed][Green Version]
  306. Hill, J.A.; Hall, J.A.; Sun, C.M.; Cai, Q.; Ghyselinck, N.; Chambon, P.; Belkaid, Y.; Mathis, D.; Benoist, C. Retinoic Acid Enhances Foxp3 Induction Indirectly by Relieving Inhibition from CD4+CD44hi Cells. Immunity 2008, 29, 758–770. [Google Scholar] [CrossRef][Green Version]
  307. Goverse, G.; Labao-Almeida, C.; Ferreira, M.; Molenaar, R.; Wahlen, S.; Konijn, T.; Koning, J.; Veiga-Fernandes, H.; Mebius, R.E. Vitamin A Controls the Presence of RORγ + Innate Lymphoid Cells and Lymphoid Tissue in the Small Intestine. J. Immunol. 2016, 196, 5148–5155. [Google Scholar] [CrossRef][Green Version]
  308. Mucida, D.; Park, Y.; Kim, G.; Turovskaya, O.; Scott, I.; Kronenberg, M.; Cheroutre, H. Reciprocal TH17 and regulatory T cell differentiation mediated by retinoic acid. Science 2007, 317, 256–260. [Google Scholar] [CrossRef] [PubMed][Green Version]
  309. Jaensson-Gyllenbäck, E.; Kotarsky, K.; Zapata, F.; Persson, E.K.; Gundersen, T.E.; Blomhoff, R.; Agace, W.W. Bile retinoids imprint intestinal CD103+ dendritic cells with the ability to generate gut-tropic T cells. Mucosal Immunol. 2011, 4, 438–447. [Google Scholar] [CrossRef][Green Version]
  310. Mielke, L.A.; Jones, S.A.; Raverdeau, M.; Higgs, R.; Stefanska, A.; Groom, J.R.; Misiak, A.; Dungan, L.S.; Sutton, C.E.; Streubel, G.; et al. Retinoic acid expression associates with enhanced IL-22 production by γδ T cells and innate lymphoid cells and attenuation of intestinal inflammation. J. Exp. Med. 2013, 210, 1117–1124. [Google Scholar] [CrossRef] [PubMed]
  311. Osanai, M.; Nishikiori, N.; Murata, M.; Chiba, H.; Kojima, T.; Sawada, N. Cellular retinoic acid bioavailability determines epithelial integrity: Role of retinoic acid receptor α agonists in colitis. Mol. Pharmacol. 2007, 71, 250–258. [Google Scholar] [CrossRef][Green Version]
  312. Kubota, H.; Chiba, H.; Takakuwa, Y.; Osanai, M.; Tobioka, H.; Kohama, G.I.; Mori, M.; Sawada, N. Retinoid X receptor α and retinoic acid receptor γ mediate expression of genes encoding tight-junction proteins and barrier function in F9 cells during visceral endodermal differentiation. Exp. Cell Res. 2001, 263, 163–172. [Google Scholar] [CrossRef]
  313. Grizotte-Lake, M.; Zhong, G.; Duncan, K.; Kirkwood, J.; Iyer, N.; Smolenski, I.; Isoherranen, N.; Vaishnava, S. Commensals Suppress Intestinal Epithelial Cell Retinoic Acid Synthesis to Regulate Interleukin-22 Activity and Prevent Microbial Dysbiosis. Immunity 2018, 49, 1103–1115.e6. [Google Scholar] [CrossRef] [PubMed][Green Version]
  314. Ismail, A.S.; Valastyan, J.S.; Bassler, B.L. A Host-Produced Autoinducer-2 Mimic Activates Bacterial Quorum Sensing. Cell Host Microbe 2016, 19, 470–480. [Google Scholar] [CrossRef] [PubMed][Green Version]
  315. Perez de Souza, L.; Alseekh, S.; Scossa, F.; Fernie, A.R. Ultra-high-performance liquid chromatography high-resolution mass spectrometry variants for metabolomics research. Nat. Methods 2021, 1–14. [Google Scholar] [CrossRef]
  316. Da Silva, R.R.; Dorrestein, P.C.; Quinn, R.A. Illuminating the dark matter in metabolomics. Proc. Natl. Acad. Sci. USA 2015, 112, 12549–12550. [Google Scholar] [CrossRef][Green Version]
  317. Jarmusch, A.K.; Wang, M.; Aceves, C.M.; Advani, R.S.; Aguirre, S.; Aksenov, A.A.; Aleti, G.; Aron, A.T.; Bauermeister, A.; Bolleddu, S.; et al. ReDU: A framework to find and reanalyze public mass spectrometry data. Nat. Methods 2020, 17, 901–904. [Google Scholar] [CrossRef]
  318. Nothias, L.F.; Petras, D.; Schmid, R.; Dührkop, K.; Rainer, J.; Sarvepalli, A.; Protsyuk, I.; Ernst, M.; Tsugawa, H.; Fleischauer, M.; et al. Feature-based molecular networking in the GNPS analysis environment. Nat. Methods 2020, 17, 905–908. [Google Scholar] [CrossRef]
  319. Dührkop, K.; Fleischauer, M.; Ludwig, M.; Aksenov, A.A.; Melnik, A.V.; Meusel, M.; Dorrestein, P.C.; Rousu, J.; Böcker, S. SIRIUS 4: A rapid tool for turning tandem mass spectra into metabolite structure information. Nat. Methods 2019, 16, 299–302. [Google Scholar] [CrossRef] [PubMed][Green Version]
  320. Schmid, R.; Petras, D.; Nothias, L.F.; Wang, M.; Aron, A.T.; Jagels, A.; Tsugawa, H.; Rainer, J.; Garcia-Aloy, M.; Dührkop, K.; et al. Ion identity molecular networking in the GNPS environment. bioRxiv 2020. [Google Scholar] [CrossRef]
  321. Shahaf, N.; Rogachev, I.; Heinig, U.; Meir, S.; Malitsky, S.; Battat, M.; Wyner, H.; Zheng, S.; Wehrens, R.; Aharoni, A. The WEIZMASS spectral library for high-confidence metabolite identification. Nat. Commun. 2016, 7, 1–13. [Google Scholar] [CrossRef][Green Version]
  322. Vinaixa, M.; Schymanski, E.L.; Neumann, S.; Navarro, M.; Salek, R.M.; Yanes, O. Mass spectral databases for LC/MS- and GC/MS-based metabolomics: State of the field and future prospects. TrAC-Trends Anal. Chem. 2016, 78, 23–35. [Google Scholar] [CrossRef][Green Version]
  323. Wishart, D.S.; Jewison, T.; Guo, A.C.; Wilson, M.; Knox, C.; Liu, Y.; Djoumbou, Y.; Mandal, R.; Aziat, F.; Dong, E.; et al. HMDB 3.0-The Human Metabolome Database in 2013. Nucleic Acids Res. 2013, 41. [Google Scholar] [CrossRef]
  324. NIST20: Updates to the NIST Tandem and Electron Ionization Spectral Libraries|NIST. Available online: https://www.nist.gov/programs-projects/nist20-updates-nist-tandem-and-electron-ionization-spectral-libraries (accessed on 14 May 2021).
  325. Smith, C.A.; O’Maille, G.; Want, E.J.; Qin, C.; Trauger, S.A.; Brandon, T.R.; Custodio, D.E.; Abagyan, R.; Siuzdak, G. METLIN: A metabolite mass spectral database. Ther. Drug Monit. 2005, 27, 747–751. [Google Scholar] [CrossRef] [PubMed]
  326. Advanced Mass Spectral Database—mzCloud. Available online: https://www.mzcloud.org/ (accessed on 14 May 2021).
  327. Horai, H.; Arita, M.; Kanaya, S.; Nihei, Y.; Ikeda, T.; Suwa, K.; Ojima, Y.; Tanaka, K.; Tanaka, S.; Aoshima, K.; et al. MassBank: A public repository for sharing mass spectral data for life sciences. J. Mass Spectrom. 2010, 45, 703–714. [Google Scholar] [CrossRef]
  328. Sawada, Y.; Nakabayashi, R.; Yamada, Y.; Suzuki, M.; Sato, M.; Sakata, A.; Akiyama, K.; Sakurai, T.; Matsuda, F.; Aoki, T.; et al. RIKEN tandem mass spectral database (ReSpect) for phytochemicals: A plant-specific MS/MS-based data resource and database. Phytochemistry 2012, 82, 38–45. [Google Scholar] [CrossRef][Green Version]
  329. Wang, M.; Carver, J.J.; Phelan, V.V.; Sanchez, L.M.; Garg, N.; Peng, Y.; Nguyen, D.D.; Watrous, J.; Kapono, C.A.; Luzzatto-Knaan, T.; et al. Sharing and community curation of mass spectrometry data with Global Natural Products Social Molecular Networking. Nat. Biotechnol. 2016, 34, 828–837. [Google Scholar] [CrossRef][Green Version]
  330. Quinn, T.P.; Erb, I. Examining microbe–metabolite correlations by linear methods. Nat. Methods 2021, 18, 37–39. [Google Scholar] [CrossRef] [PubMed]
  331. Morton, J.T.; Aksenov, A.A.; Nothias, L.F.; Foulds, J.R.; Quinn, R.A.; Badri, M.H.; Swenson, T.L.; Van Goethem, M.W.; Northen, T.R.; Vazquez-Baeza, Y.; et al. Learning representations of microbe–metabolite interactions. Nat. Methods 2019, 16, 1306–1314. [Google Scholar] [CrossRef] [PubMed][Green Version]
  332. Morton, J.T.; McDonald, D.; Aksenov, A.A.; Nothias, L.F.; Foulds, J.R.; Quinn, R.A.; Badri, M.H.; Swenson, T.L.; Van Goethem, M.W.; Northen, T.R.; et al. Reply to: Examining microbe–metabolite correlations by linear methods. Nat. Methods 2021, 18, 40–41. [Google Scholar] [CrossRef] [PubMed]
  333. Rosshart, S.P.; Herz, J.; Vassallo, B.G.; Hunter, A.; Wall, M.K.; Badger, J.H.; McCulloch, J.A.; Anastasakis, D.G.; Sarshad, A.A.; Leonardi, I.; et al. Laboratory mice born to wild mice have natural microbiota and model human immune responses. Science 2019, 365. [Google Scholar] [CrossRef]
  334. Cassotta, M.; Forbes-Hernández, T.Y.; Iglesias, R.C.; Ruiz, R.; Zabaleta, M.E.; Giampieri, F.; Battino, M. Links between nutrition, infectious diseases, and microbiota: Emerging technologies and opportunities for human-focused research. Nutrients 2020, 12, 1827. [Google Scholar] [CrossRef]
  335. Venema, K.; Van Den Abbeele, P. Experimental models of the gut microbiome. Best Pract. Res. Clin. Gastroenterol. 2013, 27, 115–126. [Google Scholar] [CrossRef]
  336. Marzorati, M.; Van De Wiele, T. An Advanced in Vitro Technology Platform to Study the Mechanism of Action of Prebiotics and Probiotics in the Gastrointestinal Tract. J. Clin. Gastroenterol. 2016, 50, S124–S125. [Google Scholar] [CrossRef]
  337. Rinschen, M.M.; Ivanisevic, J.; Giera, M.; Siuzdak, G. Identification of bioactive metabolites using activity metabolomics. Nat. Rev. Mol. Cell Biol. 2019, 20, 353–367. [Google Scholar] [CrossRef]
Figure 1. Role of short chain fatty acids (SCFAs) and lactate in immune regulation. SCFAs are produced from bacteria anaerobic fermentation of carbohydrates and dietary fibers. SCFAs have different modes of action on both local and systemic immunity. In physiological conditions, they regulate the intestinal barrier function by upregulation of the expression of the tight junction. Then, they play an important role in T-cell functioning, via regulation of G-protein-coupled receptors (GPR43, GPR41, GPR109A) and inhibition of HDAC (i.e., histone deacetylase). SCFAs regulate dendritic cells (DC) in the differentiation of T-cells, and Treg, Th1, and Th17 generation in different cytokine environments. Moreover, they inhibit the production of proinflammatory cytokines (e.g., TNF-α, IL-6, IL-1β) from intestinal macrophages and induce the production of interleukin-18 (IL18) from intestinal epithelial cells (IEC). In pathological conditions, SCFAs activate the production of proinflammatory cytokines and chemokines, such as Tumor Necrosis Factor α (TNF-α), Interleukin-6 (IL-6), CXCL1, and CXCL10, as well as increasing the production of Th1 and Th17 T lymphocytes. Lactate is a signal molecule of GPR81 and plays an important role in suppressing colonic inflammation.
Figure 1. Role of short chain fatty acids (SCFAs) and lactate in immune regulation. SCFAs are produced from bacteria anaerobic fermentation of carbohydrates and dietary fibers. SCFAs have different modes of action on both local and systemic immunity. In physiological conditions, they regulate the intestinal barrier function by upregulation of the expression of the tight junction. Then, they play an important role in T-cell functioning, via regulation of G-protein-coupled receptors (GPR43, GPR41, GPR109A) and inhibition of HDAC (i.e., histone deacetylase). SCFAs regulate dendritic cells (DC) in the differentiation of T-cells, and Treg, Th1, and Th17 generation in different cytokine environments. Moreover, they inhibit the production of proinflammatory cytokines (e.g., TNF-α, IL-6, IL-1β) from intestinal macrophages and induce the production of interleukin-18 (IL18) from intestinal epithelial cells (IEC). In pathological conditions, SCFAs activate the production of proinflammatory cytokines and chemokines, such as Tumor Necrosis Factor α (TNF-α), Interleukin-6 (IL-6), CXCL1, and CXCL10, as well as increasing the production of Th1 and Th17 T lymphocytes. Lactate is a signal molecule of GPR81 and plays an important role in suppressing colonic inflammation.
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Figure 2. Overview of the impact of the different B vitamins on immunity. The main source of VB is the dietary intake, together with modification of dietary precursors or de novo synthesis carried out by the microbiota. As VB plays a role as precursors or as enzyme cofactors in numerous metabolic reactions, they impact the regulation of immune homeostasis through the metabolism, which is summarized here for each VB. MAIT, mucosa-associated invariant T lymphocytes; IBD, inflammatory bowel disease; ROS, reactive oxygen species).
Figure 2. Overview of the impact of the different B vitamins on immunity. The main source of VB is the dietary intake, together with modification of dietary precursors or de novo synthesis carried out by the microbiota. As VB plays a role as precursors or as enzyme cofactors in numerous metabolic reactions, they impact the regulation of immune homeostasis through the metabolism, which is summarized here for each VB. MAIT, mucosa-associated invariant T lymphocytes; IBD, inflammatory bowel disease; ROS, reactive oxygen species).
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Figure 3. Summary of the immunomodulatory properties of the main described soluble metabolites on epithelial cells, as well as on innate and adaptive immune cells. D-AA, Stereoisomer D-amino acids; QSM, Quorum Sensing Molecules; CDN, Cyclic-DiNucleotides; CTN, Cyclic TriNucleotides.
Figure 3. Summary of the immunomodulatory properties of the main described soluble metabolites on epithelial cells, as well as on innate and adaptive immune cells. D-AA, Stereoisomer D-amino acids; QSM, Quorum Sensing Molecules; CDN, Cyclic-DiNucleotides; CTN, Cyclic TriNucleotides.
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Figure 4. Summary of membrane-bound metabolites and their impact on immune cells. iNKT invariant Natural killer T, LTA, lipoteichoic acids; WTA, wall teichoic acids; PGE2, Prostaglandin E2.
Figure 4. Summary of membrane-bound metabolites and their impact on immune cells. iNKT invariant Natural killer T, LTA, lipoteichoic acids; WTA, wall teichoic acids; PGE2, Prostaglandin E2.
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Caffaratti, C.; Plazy, C.; Mery, G.; Tidjani, A.-R.; Fiorini, F.; Thiroux, S.; Toussaint, B.; Hannani, D.; Le Gouellec, A. What We Know So Far about the Metabolite-Mediated Microbiota-Intestinal Immunity Dialogue and How to Hear the Sound of This Crosstalk. Metabolites 2021, 11, 406. https://doi.org/10.3390/metabo11060406

AMA Style

Caffaratti C, Plazy C, Mery G, Tidjani A-R, Fiorini F, Thiroux S, Toussaint B, Hannani D, Le Gouellec A. What We Know So Far about the Metabolite-Mediated Microbiota-Intestinal Immunity Dialogue and How to Hear the Sound of This Crosstalk. Metabolites. 2021; 11(6):406. https://doi.org/10.3390/metabo11060406

Chicago/Turabian Style

Caffaratti, Clément, Caroline Plazy, Geoffroy Mery, Abdoul-Razak Tidjani, Federica Fiorini, Sarah Thiroux, Bertrand Toussaint, Dalil Hannani, and Audrey Le Gouellec. 2021. "What We Know So Far about the Metabolite-Mediated Microbiota-Intestinal Immunity Dialogue and How to Hear the Sound of This Crosstalk" Metabolites 11, no. 6: 406. https://doi.org/10.3390/metabo11060406

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