Redox signaling in the gastrointestinal tract

Author’s Accepted Manuscript Redox Signaling In The Gastrointestinal Tract Salvador Pérez, Raquel Taléns-Visconti, Sergio Rius-Pérez, Isabela Finamor, Juan Sastre

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To appear in: Free Radical Biology and Medicine Received date: 2 July 2016 Revised date: 20 December 2016 Accepted date: 31 December 2016 Cite this article as: Salvador Pérez, Raquel Taléns-Visconti, Sergio Rius-Pérez, Isabela Finamor and Juan Sastre, Redox Signaling In The Gastrointestinal Tract, Free Radical Biology and Medicine, http://dx.doi.org/10.1016/j.freeradbiomed.2016.12.048 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

REDOX SIGNALING IN THE GASTROINTESTINAL TRACT Salvador Pérez, Raquel Taléns-Visconti2, Sergio Rius-Pérez, Isabela Finamor, and Juan Sastre Department of Physiology, Faculty of Pharmacy. University of Valencia, 46100 Burjasot, Valencia, Spain 2

Department of Pharmacy and Pharmaceutical Technology and Parasitology, Faculty of

Pharmacy University of Valencia, 46100 Burjasot, Valencia, Spain *

CORRESPONDING AUTHOR: Prof. Juan Sastre, Department of Physiology,

Faculty of Pharmacy, University of Valencia Avda. Vicent Andres Estellés s/n, 46100 Burjassot, Valencia, Spain. Tel. 0034 96 354 38 15; Fax: 0034 96 354 33 95. [email protected] ABSTRACT Redox signaling regulates physiological self-renewal, proliferation, migration and differentiation in gastrointestinal epithelium by modulating Wnt/β-catenin and Notch signaling pathways mainly through NADPH oxidases (NOXs). In the intestine, intracellular and extracellular thiol redox status modulates the proliferative potential of epithelial cells. Furthermore, commensal bacteria contribute to intestine epithelial homeostasis through NOX1- and dual oxidase 2-derived reactive oxygen species (ROS). The loss of redox homeostasis is involved in the pathogenesis and development of a wide diversity of gastrointestinal disorders, such as Barrett’s esophagus, esophageal adenocarcinoma, peptic ulcer, gastric cancer, ischemic intestinal injury, celiac disease, inflammatory bowel disease and colorectal cancer. The overproduction of superoxide anion together with inactivation of superoxide dismutase are involved in the pathogenesis of Barrett’s esophagus and its transformation to adenocarcinoma. In Helicobacter pylori-induced peptic ulcer, oxidative stress derived from the leukocyte

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infiltrate and NOX1 aggravates mucosal damage, especially in HspB+ strains that downregulate Nrf2. In celiac disease, oxidative stress mediates most of the cytotoxic effects induced by gluten peptides and increases transglutaminase levels, whereas nitrosative stress contributes to the impairment of tight junctions. Progression of inflammatory bowel disease relies on the balance between pro-inflammatory redoxsensitive pathways, such as NLRP3 inflammasome and NF-κB, and the adaptive upregulation of Mn superoxide dismutase and glutathione peroxidase 2. In colorectal cancer, redox signaling exhibits two Janus faces: On the one hand, NOX1 up-regulation and derived hydrogen peroxide enhance Wnt/β-catenin and Notch proliferating pathways; on the other hand, ROS may disrupt tumor progression through different proapoptotic mechanisms. In conclusion, redox signaling plays a critical role in the physiology and pathophysiology of gastrointestinal tract. ABBREVIATIONS HNE, 4-hydroxynonenal; 8-OHdG, 8-oxo-7,8-dihydro-2’-deoxyguanosine; AP-1, Activator protein 1; APC, Adenomatous polyposis coli; APE-1, Apurinic/apyrimidinic endonuclease 1; Math1, Atonal homolog 1; Bax, BCL2-associated X protein; β-TrCP, Beta-transducin repeat containing protein; BMP, Bone morphogenic protein; CD, Celiac disease; CREB, cAMP response element binding protein; JNK, c-Jun N-terminal kinase; CXCR2, C-X-C chemokine receptor type 2; COX, Cyclooxygenase; CagA, Cytotoxin-associated protein gene A; DCC, Deleted in Colorectal Carcinoma; DPC4, Deleted in Pancreatic Cancer locus 4; Dvl, Dishevelled; DUOX, Dual oxidase; eNOS, Endothelial nitric oxide synthase; EGFR, Epidermal growth factor receptor; ERBB-2, Erythroblastic leukemia associated viral oncogene B-2; Eph, Erythropoietin-producing hepatocyte; ERK, Extracellular signal–regulated kinase; FPR, Formyl peptide receptor; GWAS, Genome-wide association studies; GPx, Glutathione peroxidase; GST, Glutathione S-transferase; GSK-3 β, Glycogen synthase kinase-3β; Hes, Hairy/Enhancer of Slipt; HspB, Heat-shock protein B; HO-1, Heme oxygenase-1; HNF4α, Hepatocyte nuclear factor-4α; HDAC-1, Histone deacetylase 1; HLA, Human leukocyte antigen; HIF-1α, Hypoxia inducible factor-1 alpha; IER3, Immediate early response-3; iNOS, Inducible nitric oxide synthase; IL, Interleukin; Keap 1, Kelch-like ECH-associated protein 1; MDA, Malondialdehyde; MMP, Matrix metalloproteinase; MAPK, Mitogen-activated protein kinase; NH2Cl, Monochloramine; MUC, Mucin;

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MPO, Myeloperoxidase; NOX, NADPH oxidase; NOXA1, NADPH oxidase activator 1; NOXO1, NADPH oxidase organizer 1; NQO1, NADPH quinone oxidoreductase-1; NADPH, Nicotinamide adenine dinucleotide phosphate; NO, Nitric oxide; NOS, Nitric oxide synthase; NSAIDs, Nonsteroidal anti-inflammatory drugs; NICD, Notch intracellular domain; Nrf2, Nuclear factor erythroid 2-related factor-2; NF-κB, Nuclear factor kappa B; NOD, Nucleotide-binding oligomerization domain; GSSG, Oxidized glutathione; PUMA, p53-upregulated modulator of apoptosis; PRR, Pattern recognition receptor; PTEN, Phosphatase and tensin homolog deleted on chromosome 10; PI3K, Phosphatidylinositol 3-kinase; Pdgf-A, Platelet-derived growth factor-A; PG, Prostaglandin; RAC1, Ras-related C3 botulinum toxin substrate 1; RNS, Reactive nitrogen species; ROS, Reactive oxygen species; RBPJ, Recombining binding protein suppressor of hairless; GSH, Reduced glutathione; STAT, Signal transducers and activators of transcription; SOD, Superoxide dismutase; SMAD, Suppressor of mothers against decapentaplegic; TCF, T-cell factor; Th, T-helper; TG, Tissue transglutaminase; TLR, Toll-like receptor; TIGAR, Tp53-inducible glycolysis and apoptosis regulator; TGF-β, Transforming growth factor-beta; TFFs, Trefoil factors; TNF-α, Tumor necrosis factor-alpha; VEGF, Vascular endothelial growth factor

KEYWORDS Intestinal epithelium; NADPH oxidase 1; dual oxidase 2; Nrf2; APE1/Ref1; MnSOD; GPx2; Barrett’s esophagus; peptic ulcer; gastric cancer; celiac disease; inflammatory bowel disease; colorectal cancer

This review is divided into two parts, the first one devoted to gastrointestinal physiological redox signaling, and the second one deals with gastroeintestinal disorders. Indeed, the first part of this review encompasses a critical and updated overview on redox biology in the stomach and intestine, explaining how redox signaling regulates physiological

self-renewal,

proliferation,

migration

and

differentiation

in

gastrointestinal epithelium. The important interplay between the intestinal microbiota and physiological redox signaling is also covered.

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The second part of this review focuses on the role of redox signaling in the pathogenesis and development of the most prevalent gastrointestinal disorders, namely Barrett’s esophagus, esophageal adenocarcinoma, peptic ulcer, gastric cancer, ischemic intestinal injury, celiac disease, inflammatory bowel disease and colorectal cancer.

1.- PHYSIOLOGICAL REDOX SIGNALING IN THE GASTROINTESTINAL TRACT 1.1.- Redox biology in the stomach: A physiological nitrosative stress The gastric mucosa contains different mucosal specialized glands along the stomach. Mucosal glands of the body and fundus of the stomach contain parietal (oxyntic) cells specialized in secretion of hydrochloric acid and intrinsic factor, together with chief cells that secrete pepsinogen and mast cells that secrete histamine. Pepsinogen may be also secreted by cells of pyloric glands. Antral pyloric glands contain G (gastrin) cells that secrete gastrin, the most potent stimulator of acid secretion. Gastric mucous cells secrete mucus rich in a large glycoprotein composed by four subunits bound by disulfide bridges, which are required to form a viscous protective gel. The gastric mucosal barrier and the gastric blood flow maintain the stomach integrity against the gastric juice and dietary irritants [1]. Reduced glutathione (GSH) levels are higher in gastric mucosa (7-8 mM) in comparison with other parts of the gastrointestinal tract, providing additional protection against gastric acid [2,3]. Gastric blood flow is regulated by nitric oxide (NO), which can be generated via nitric oxide synthase (NOS) in the vascular endothelium, epithelial cells and sensory nerve endings [4]. Nevertheless, NO may be also formed in the stomach independently of NOS.

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Dietary nitrate is readily absorbed in the gastrointestinal tract and around 25% is taken by salivary glands, which greatly concentrate nitrate in saliva (up to 20 fold) [5] leading to the entero-salivary circulation of nitrate [6]. Nitrate reductase from commensal bacteria reduces salivary nitrate to nitrite in the oral cavity [7] and in fact human saliva contains high nitrite levels (50-250 µM) [8]. When the nitrite reaches the acidic gastric pH it is rapidly protonated to nitrous acid (HNO2), which decomposes to generate nitric oxide (.NO) and other nitrogen oxides, such as nitrogen dioxide (.NO2), dinitrogen trioxide (N2O3), and peroxynitrite [9–11]. Hence, in addition to the classical pathway to generate NO in vivo through NOS, it can be also formed through the reduction of nitrite in the stomach yielding very high NO levels [6,12]. This pathway requires oral commensal bacteria because NO formation is almost abrogated in germ-free sterile rats after a dietary load of nitrate [13]. It is worth noting that dietary vitamin C and polyphenols largely increase nitrite-derived NO in the stomach [12,14,15]. The NOS-independent generation of NO plays a key role in the biology of the gastric mucosa because NO-derived from salivary nitrite increases gastric mucosal blood flow and mucus thickness [8] and triggers nitration of pepsinogen and pepsin, which decreases its proteolytic activity protecting against peptic ulcer [16,17]. NO may also induce smooth muscle relaxation modulating gastric emptying [12,18]. In addition, gastric NO and other nitrogen oxides derived from nitrite significantly helps to host defense by their potent microbicidal activity against pathogenic bacteria such as Helicobacter pylori, Pseudomonas aeruguinosa, Salmonella, Yersinia, and Shigella species [6,9,19]. Furthermore, an anti-inflammatory role of lipid nitration in gastric pathophysiology has been proposed [11], which deserves further investigation. Consequently, a physiological nitrosative stress may be present in the stomach after a meal. Nevertheless, it should be taken into account that an excess of nitrate in the diet

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might generate nitrite-derived nitrosamines with potential carcinogenic effects and therefore epidemiological studies in this regard should be performed.

1.2.- The self-renewing intestinal epithelium The intestinal mucosa is a complex and dynamic tissue that comprises the surface monolayer of self-renewing epithelial cells and the underlying lamina propria with its immune, vascular and structural components [20]. The intestinal epithelium is composed of a single and heterogeneous layer of cells that are completely renewed every 2-7 days [21]. In the small intestine there are invaginations into the mucosa called crypts of Lieberkühn that contain proliferative stem cells and prominences into the lumen called villi with terminally differentiated cells (Figure 1). The colon epithelium lacks villi. Two types of stem cells have been identified in the crypts, the radiation-sensitive pluripotent stem cells at position +4 relative to the crypt base, and the small and cycling crypt base columnar cells (Figure 1) [22]. The crypts also contain transit-amplifying cells that follow upward migration, and differentiated Paneth cells that are kept at the bottom and secrete bactericidal cryptdins/defensins and lysozymes [23]. Paneth cells are the only differentiated cells in the crypts, being generated at positions 5-7 of the crypt and then migrate downward to occupy the first three positions at the crypt bottom, and function not only in innate immunity and antibacterial defense, but also provide essential niche signals to intestinal stem cells [22,23]. They do not follow the typical rapid renewal of differentiated intestinal cells because their lifespan is at least three weeks [24]. Although Paneth cells are not present in the colon, CD24+ cells located in colon crypts between intestinal stem cells may represent the Paneth cell equivalents

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[23]. Each stem cell usually divides asymmetrically yielding one daughter stem cell and one transit-amplifying daughter that gives rise to one of the five types of differentiated epithelial cells, Paneth cells in crypts and four types of differenciated cells in villi (Figure 1) [25,26]. When there is intestinal damage, stem cells divide symmetrically yielding two daughter stem cells in order to replace damaged cells [27]. The four types of differentiated cells in villi are the majority absorptive enterocytes (>80% of all epithelial cells), and the rest secretory cells, mainly globet cells and a minority of enteroendocrine cells and tuft cells [21,22]. The highly polarized enterocytes exhibit an apical brush border specialized in nutrient and water absorption. Globet cells secrete mucus containing mucins and trefoil proteins that protect the mucosa against chemical aggression and shear stress, favor movement of the intestinal content, and also provide a supportive environment for the intestinal microbiota [22,28]. The percentage of globet cells rises along the intestine, being around 4% in the duodenum and around 16-50% in the colon [28]. Although enteroendocrine cells constitute only a minority (1% of all epithelial cells), they play a key role in digestive physiology through secretion of gastrointestinal hormones, such as secretin and cholecystokinin. Tuft cells are the only ones that release intestinal opioids and prostanoides [21]. All these intestinal differentiated cells are generated at the crypts and mature and differentiate after cell cycle arrest during their upward migration towards the villi [26,29]. Differentiated cells of the villi exhibit short lifespan because they suffer spontaneous apoptosis and shedding when they reach the tip of the villi after three days of their terminal differentiation [22].

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The intestinal immune defense relies on the gut-associated lymphoid tissue, which is composed by Peyer’s patches scattered along the intestinal wall and surrounded by a follicle-associate epithelium containing invaginated microfold cells (M cells) that present antigens to lymphocytes and dendritic cells [28].

1.3.- Redox biology in the intestine The dynamics of the intestinal epithelium imply changes in the intracellular redox potential of glutathione [GSH/oxidized glutathione (GSSG)] from cell proliferation (260 to -240 mV), through differentiation (-220 to -200 mV), and finally till apoptosis (170 to -150 mV) [30–32]. GSH levels in the intestinal epithelium are within the millimolar range similarly to other cell types [28,33,34]. In addition to the absorption of its constituent amino acids and subsequent intracellular GSH synthesis, trans-epithelial transport of GSH across the apical membrane exists, which can be sodium-dependent or independent [35]. Generally extracellular GSH levels are very low, except in the intestinal lumen (60-300 µM) where GSH derives from the dietary intake and mainly from the biliary output [33], as GSH concentration is high in the bile (1-2 mM in rat bile) [36]. Luminal GSH promotes reduction of dietary disulfides, metabolism of peroxidized lipids, xenobiotic detoxification, and the assembly of mucin oligomers to maintain mucus fluidity [28,33,34]. Luminal and intracellular GSH greatly protects against the lipid peroxides often present in the diet [37], in fact the estimated daily intake of lipid peroxides is around 1.4 mmol associated with 84 g of fat [38]. Accordingly, GSH supplementation restores mucosal proliferation that was suppressed in the rat intestine by chronic administration of lipid peroxides [39]. Nevertheless, the effect of lipid hydroperoxides

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on cell proliferation in the gut is dose-dependent because they promote cell proliferation at low levels (1-5 µM), but growth arrest and apoptosis at high levels (10-50 µM) [40,41]. The luminal thiol redox status is regulated mainly by the cysteine/cystine shuttle that involves cystine uptake by epithelial cells, intracellular cystine reduction, and subsequent release of cysteine to the lumen that joins the cysteine pool generated from luminal GSH hydrolysis [28,34]. Furthermore, the extracellular cysteine-to-cystine ratio modulates the proliferative potential of intestinal epithelial cells [42]. Thus, extracellular cysteine and cystine promote proliferation of CaCo-2 cells, independently of intracellular GSH levels, through phosphorylation of the epidermal growth factor receptor (EGFR) and subsequently of extracellular signal–regulated kinase (ERK) (p44/p42) mitogen-activated protein kinase (MAPK), which were mediated by a redoxsensitive metalloproteinase [43,44]. Moreover, intestinal CaCo-2 cells are able to modulate the extracellular cysteine-to-cystine ratio by stimulating their transport through the sodium-dependent transport system y+L and the sodium independent system xc- [44,45]. Nevertheless, all the experiments on modulation of the proliferative potential of intestinal epithelial cells by intracellular and extracellular thiol redox status have been performed in vitro in CaCo-2 cells and would require confirmation in vivo taking into account the complex and dynamic self-renewing intestinal epithelium.

1.4.- The Wnt, Notch, and BMP pathways drive the intestinal epithelium dynamics Stem cell self-renewal, cell proliferation, migration and differentiation are strictly controlled mainly by three signaling pathways, the Wnt, Notch and bone morphogenetic

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proteins (BMP) (Figure 2). The Wnt and Notch pathways are redox-sensitive since they may be modulated by NADPH oxidase (NOX) as explained later in item 1.5. The Wnt pathway (Figure 2) drives the proliferation of stem cells and progenitor epithelial cells in the crypts and its activity follows a gradient with its maximum at the crypt bottom [22,26,46]. The Wnt signaling has been also involved in the migration of epithelial cells towards the villi and in the terminal differentiation of Paneth cells [26]. Wnt signaling may proceed through canonical (β-catenin/ T-cell factor (TCF) dependent) or non-canonical (β-catenin/TCF independent) pathways, but only the former has been described so far in the intestine [26]. The canonical Wnt pathway requires β-catenin. The lack of Wnt activity triggers N-terminal phosphorylation of βcatenin and its ubiquitination and degradation by the proteasome beta-transducin repeat containing protein (β-TrCP), an F-box containing E3 ubiquitin ligase [47,48]. The phosphorylation status of β-catenin is controlled by a destruction complex composed of protein scaffolds and the tumor suppressors axin and adenomatous polyposis coli (APC) together with the kinases glycogen synthase kinase-3β (GSK-3 β) and casein kinase I [22,26]. This degradation complex is inactivated upon binding of Wnt ligands to the Frizzled and low-density lipoprotein receptor-related protein (LRP) receptors triggering accumulation of β-catenin and its translocation to the nucleus where it forms an active transcriptional complex with transcription factors of TCF/lymphocyte enhancer factor (TCF/LEF) to upregulate target genes, such as c-MYC, CYCLIN D1, and erythropoietinproducing hepatocyte B (EPHB) [22,26]. The ligands Wnt3, Wnt6 and Wnt9B are considered responsible for canonical Wnt signaling in the small intestine [46]. In absence of β-catenin, transcriptional repressors, such as Groucho, bind TCF/LEF transcription factors silencing signaling [49].

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Wnt signaling is essential for proliferative activity in crypts and accordingly intestinespecific Wnt target genes are up-regulated in proliferative progenitors [22]. Indeed, lack of Tcf-4 transcription factor [50], blockade of Wnt signaling by the soluble Dickkopf-1 inhibitor [51,52], or depletion of β-catenin in the intestinal epithelium [53,54] all lead to marked reduction in the intestinal proliferative activity. Conversely, the Wnt agonist RSpondin 1 caused intense hyperproliferation of intestinal crypts [55]. EphB receptors are Wnt target genes and together with Ephrin B ligands seem to regulate the migration, maturation and differentiation of intestinal epithelial cells along the crypt-villus axis [26]. In fact, mutant APC mice exhibit blockade of epithelial cell migration towards the villi [56]. The expression of EphB receptors parallels that of the Wnt signaling following the same gradient along the crypt and villi. Thus, EphB2 is mainly expressed on crypt base columnar cells, but not in Paneth cells, and reduces its expression from just above the Paneth cells towards the crypt top. Particularly, EphB3 is expressed only in crypt base columnar cells and in Paneth cells. In contrast to the EphB expression, Ephrin B1 and B3 are mostly expressed in differentiated cells in the villi and their expression diminishes along the way towards the crypt bottom, with absence in Paneth cells [26]. It is believed that the EphB-Ephrin B interaction exhibits repellant effects that might explain why Paneth cells, which exhibit EphB3 but lack Ephrin B, avoid upward migration and remain at the crypt bottom [26]. The Wnt pathway also regulates the appropriate maturation and differentiation of secretory epithelial cells. Thus, strikingly enteroendocrine cells are absent in mice deficient in Tcf-4 [57]. Furthermore, constitutive activation of the Wnt pathway due to APC mutation leads to marked reduction in differentiated cells in the villi [56].

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The Notch pathway (Figure 2) contributes decisively to keep proliferative and undifferentiated cells in the crypts and additionally controls the secretory fate decisions in the intestinal epithelium [22]. The Notch signaling is activated when one of the Delta or Jagged Notch transmembrane receptors interacts with one of the five Notch ligands triggering proteolytic cleavage of the receptor by the γ-secretase protease complex [22,26]. This cleavage releases the free Notch 1 intracellular domain (NICD) that translocates into the nucleus where it binds the transcription factor recombination signal binding protein J-κ (RBPJ-k) (CSL or CBF1) to upregulate target genes, mainly genes of the Hairy/Enhancer of Slipt (Hes) class that encode transcriptional repressors, such as Hes1 and Hes5 in the intestine. Hes repressors down-regulate basic-loop-helix transcription factors, especially Math1 in a Hes1 dependent manner in the intestinal epithelium, leading to differentiation towards specific lineages, particularly triggering absorptive determination [22,58]. It has been proposed that Notch signaling controls a binary decision in the intestine between absorptive and secretory cell fates [29,59,60]. Hes1 seems to be mainly involved into the differentiation of precursor cells into enterocytes, while Math1 and neurogenin 3 regulates the differentiation of secretory precursors into globet cells or enteroendocrine cells, respectively [58,61,62]. Blockade of the Notch pathway using γ-secretase inhibitors or through deletion of the Csl gene leads to conversion of all intestinal epithelial cells into globet cells [22,63]. The intestine from Hes1 deficient mice exhibit abundance of globet cells as well as enteroendocrine and Paneth cells, but a reduced number of enterocytes [61,64]. Conversely, deficiency in Math1 leads to an intestinal epithelium formed only by enterocytes [58]. Notch1 proteolysis is active in crypt epithelial cells, especially in intestinal stem cells, whereas Notch2 expression is weak in the crypts [65]. Constitutive

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overexpression of Notch1 receptor enhances proliferation reducing the differentiation of enteroendocrine and Paneth cells, thus decreasing their numbers [59,66]. The normal and complex homeostasis of the intestinal epithelium requires epitheliummesenchyme interactions in both directions, which originate the intestinal stem cell niche [67]. In fact, subepithelial myofibroblasts secrete putative growth factors and cytokines that stimulate epithelial proliferation [68]. Furthermore, BMP signaling (Figure 2) is located in the mesenchyme and epithelium of villi but not in the crypts and negatively regulates them [22,69,70]. BMPs belong to the transforming growth factorbeta (TGF-β) family and BMP ligands bind to their receptors triggering phosphorylation of mothers against decapentaplegic homolog (SMAD) transcription factors 1, 5, or 8, which associates with SMAD 4 and subsequently the heterodimeric complex translocates into the nucleus to up-regulate target genes [26,71]. It is noteworthy that overexpression of the BMP inhibitor Noggin or deficiency in BMP receptor 1A leads to hyperproliferative crypts because BMP signals suppress Wnt/β-catenin signaling [69,70]. The interaction between the epithelium and the mesenchyme also takes place through the hedgehog pathway and platelet-derived growth factor-A (Pdgf-A) [22]. Epithelial cells express hedgehog ligands whereas hedgehog patched receptors are expressed in the mesenchyme, and overexpression of a pan-hedgehog inhibitor leads to a hyperproliferative epithelium with reduced villi [72]. Similarly, Pdgf-A is expressed only in the epithelium whereas its receptor is expressed only in the mesenchyme, and deficiency in any of them also reduces villus morphogenesis [73].

1.5.- RAC1-NOX vs TIGAR in the control of ROS-dependent Wnt-driven proliferation

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Tp53-inducible glycolysis and apoptosis regulator (TIGAR) is a protein that modulates GSH levels by controlling glucose metabolism and NADPH levels [74,75], whereas Ras-related C3 botulinum toxin substrate 1 (RAC1) is a GTPase member of the NOX complex that generates superoxide radicals (O2-∙) [76]. Paradoxically, the loss of TIGAR enhances reactive oxygen species (ROS) but reduces Wnt-driven proliferation in the intestine, whereas the loss of RAC1 decreases ROS and also diminishes Wnt-dependent proliferation [75]. Accordingly, too much ROS hinders proliferation in the intestine whereas too little ROS may also inhibit proliferation. The identification of two different intracellular ROS pools recently proposed (TIGAR-controlled damaging ROS vs RAC1-triggered signaling ROS) [75] requires further confirmation. RAC1-mediated signaling should be ascribed to hydrogen peroxide (H2O2) following the FRBM guideline [77]. Redox signaling occurs in the colon epithelium through NOX1 under the delicate balance between RAC1 as enhancer and TIGAR as inhibitor, but it is still unkown which precise NOX isoform regulates epithelium dynamics under normal conditions in the small intestine. NOX1 is expressed at high levels in intestinal epithelial cells only in the colon [78], especially in the distal colon [79], but not in the normal small intestine where its expression is very low [80,81]. NOX1 is critically involved in the regulation of both Wnt/β-catenin and Notch signaling in the colon [29,82]. Indeed, the colon of Nox1 deficient mice exhibits simultaneous repression of both the Wnt/β-catenin and the Notch signaling causing marked decrease in proliferative cells, cell cycle arrest in progenitor cells, and conversion of all progenitor cells into globet cells, which were also expanded within crypts [29].

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NOX1 regulates the Wnt/β-catenin pathway through three different mechanisms (Figure 2): directly as an essential part of Wnt signaling, and indirectly by enhancing this pathway through oxidation of nucleoredoxin or phosphatase and tensin homolog deleted on chromosome 10 (PTEN). NOX1-derived O2-∙ generation is required to trigger the dephosphorylation and accumulation of β-catenin induced by Wnt3a, the up-regulation of its targets cyclin D1 and C-myc as well as Wnt-induced cell proliferation [82]. Wnt3a activates the RAC1 GTPase, which then translocates to the plasma membrane to assemble with the regulatory subunits NADPH oxidase organizer 1 (NOXO1) and NADPH oxidase activator 1 (NOXA1) as well as with NOX1 and p22Phox to form the active enzyme complex [83,84]. NOX1 also stimulates Wnt/β-catenin signaling by modulating the redox-sensitive interaction between the thioredoxin family protein nucleoredoxin and the Axin-binding protein Dishevelled (Dvl) [82]. Nucleoredoxin selectively inhibits the Wnt/β-catenin signaling because it activates the destruction complex and enhances β-catenin degradation upon binding to Dvl, an inhibitor of the destruction complex [85]. NOX1derived H2O2 triggers cysteine oxidation in nucleoredoxin, which inhibits its association with Dvl leading to activation of the Wnt/β-catenin pathway [82,85]. In addition, the lack of O2-∙ derived from NOX1 activates PTEN by lowering oxidation of its active-site Cys124 residue and its phosphorylation at Ser380, which reduces Akt activity and β-catenin transcriptional activity towards c-Myc and cyclin D1 as target genes [29]. On the other hand, NOX1 also stimulates the Notch pathway. Thus, absence of NOX1derived O2-∙ in Nox1 knock-out mice down-regulates the Notch pathway through decreased expression of γ-secretase and NICD, leading to lower Hes1 expression and

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Math1 up-regulation [29]. Nervertheless, the precise mechanism responsible for the O2-∙ -dependent up-regulation of γ-secretase and NICD remains to be elucidated. Hence, NOX1 is essential to maintain the proliferative state of crypt progenitors in colon and regulates cell fate decision of transit amplifying precursor cells by controlling both Wnt and Notch signaling [29]. Nevertheless, ROS derived from other sources may also be involved in this regulation. Thus, the loss of glutathione peroxidase 3 (GPx3) enhances H2O2 levels and stimulates Wnt-driven proliferation [86], and moreover TIGAR as well as redox-sensitive nucleoredoxin and PTEN open multiple alternative regulatory mechanisms. In addition, future research is required to explore potential redox BMP signaling in the intestinal mesenchyme or epithelium of villi.

1.6.- Antioxidant enzymes critically protect the intestinal epithelium Glutathione S-transferases (GSTs), superoxide dismutases, catalase, and particularly glutathione peroxidases exert a tremendous protective role in the intestinal epithelium against injury and inflammation. Crypt epithelial cells seems to be better protected against oxidative damage than the villi, since they exhibit 2- to 3-fold increase in glutathione peroxidase activity and 3- to 5-fold increase in catalase activity versus the villus [87]. Superoxide dismutase prevents mucosal injury [2]. GPx1 as well as intestine specific GPx2 and extracellular GPx3 all provide a very important protection in the intestine against inflammation and cancer, as it is explained later in the item on inflammatory bowel disease. GPx1 homogeneously expressed along the intestinal epithelium, whereas GPx2 expression is higher in crypts than in villi and exhibits its highest expression in ileum and cecum [87,88]. GPx3 is expressed in the

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gastrointestinal tract and additionally plasma GPx3 binds to the basement membranes of intestinal epithelial cells [86]. GSTs also play an important role in the intestine through the detoxification of electrophilic compounds present in dietary intake or drug treatment [36]. GST activity is high in villi and along the intestinal epithelium, except in crypts that show lower activity [87]. The presence of active GST in the extracellular mucus layer also allows detoxification of electrophilic agents without being absorbed [36]. Since GST activity is largely (100 –fold) higher than the rate of GSH synthesis and uptake, GSH concentration in enterocytes is considered the limiting factor for conjugation and elimination of electrophilic compounds [36]. The thioredoxin-thioredoxin reductase system contributes to the intestinal immune defense as thioredoxin is highly expressed in lamina propria T-lymphocytes, where it enhances cytokine expression, and inhibition of thioredoxin reductase abrogates cytokine expression [89]. Glutaredoxin 2 as well as peroxiredoxins 4 and 5 are also highly expressed in enterocytes [90], but their functional relevance in intestinal physiology requires further studies.

1.7.- Redox regulation of innate immunity in the gastrointestinal tract Innate immunity in the gastrointestinal tract is critical against pathogenic bacteria and involves a complex circuitry related to interleukin 23 (IL-23), IL-22, and toll-like receptors (TLRs), respiratory bursts mediated mainly by dual oxidase 2 (DUOX2) in intestinal epithelial cells in addition to the classical NOX2 in macrophages, and microbicidal defensins, all redox sensitive processes.

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DUOX2 belongs to the subfamily of epithelial-specific NADPH oxidases that contain an additional peroxidase homology domain generating H2O2 in a Ca2+/NADPHdependent manner and it is the predominant DUOX isoenzyme in the gut [84,91]. DUOX2 protein is located at the apical membrane of enterocytes with maximal expression in the tip epithelium of ileum, colon, and cecum [92]. DUOX2 mediates the circuitry driven by IL-23 and IL-22 that coordinates the innate defense in the gut in mammals [93]. IL-23 is released from lamina propria macrophages upon contact with bacteria or bacterial products and it regulates IL-22, which is released from T-helper 17 (Th17) cells and lymphoid cells of the lamina propria, Peyer’s patches and mesenteric lymph nodes, markedly activating DUOX2 [94] and enhancing the expression of antimicrobial peptides [93]. DUOX may trigger an oxidative burst to limit proliferation of pathogenic bacteria in the gut, and it is considered indispensable to cope with microbial gut infection in Drosophila melanogaster [91]. In mice, H2O2 generated by DUOX2 prevent the infection and inflammation of the gastric mucosa by Helicobacter felis [95]. The p38α MAPK pathway is responsible for Duox up-regulation in mammals as soluble microbial extract triggers p38α MAPK activation and DUOX2-dependent H2O2 generation in human CaCo-2 cells [96]. NOX1 may also contribute to innate immunity in the gastrointestinal tract, as it mediates the TLR5 pathway [80], and it is up-regulated by interferon gamma [78], lipopolysaccharide [80] or flagelin [97]. Nevertheless, the major role in redox regulation of innate immunity in the gut seems to be ascribed to DUOX2. The formation of active microbicidal defensins in Paneth cells requires thiol oxidation and additionally their activity is modulated by the redox status. Thus, the redox-

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sensitive matrix metalloproteinase-7 (MMP-7) (matrilysin) is required to convert cryptidin precursors into active microbicidal defensin peptides in Paneth cells and ProMMP-7 may be activated through oxidation of the thiol residue of the cysteine switch domain to sulfinic acid [98]. Moreover, MMP-7 activity is modulated by intestinespecific GPx2 [88,99]. Defensins generally harbor three intramolecular disulphide bridges that generally provide them with maximal microbicidal activity, with the exception of β-defensin 1 that exhibits potent microbicidal activity upon reduction of its thiols by thioredoxin, which is highly expressed in the intestinal mucosa and colocalizes with reduced β-defensin 1 [90,100]. Progress in the pathogenesis and pathophysiology of gastrointestinal infection requires indentifying how pathogenic bacteria counteract the multiple pro-oxidant mechanisms associated with these powerful innate immune processes becoming inefficient to cope with the bacterial aggression. On the other hand, excessive chronic stimulation of DUOX2 might contribute to inflammatory bowel disease and this would deserve further investigation.

1.8- Redox regulation of mucins and wound healing in the gastrointestinal tract The gastrointestinal mucosal barrier is protected from chemical attack and bacterial invasion by layers of mucins, whose synthesis is stimulated though specific pro-oxidant mechanisms. Mucins are high molecular weight, highly glycosylated proteins expressed in Globet cells that constitute the major components of gastrointestinal mucus [101]. They can be either secretory forming gel-like highly hydropholic polymers rich of glycans or membrane-bound proteins anchored to the apical membrane of gastrointestinal epithelial cells. DUOX2-derived O2-∙ is required for the up-regulation of

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some of the most important mucins, such as secretory mucin 3 (MUC3) and membranebound MUC5AC, induced by growth factors [101]. Appart from mucins, wound healing capacity is required for maintenance of the integrity of the gastrointestinal mucosal barrier and it relies on redox signaling through NOX1 [102]. Formyl peptide receptors (FPRs), a class of pattern recognition receptors (PRRs) found in intestinal crypts, mediate mucosal wound healing through NOX1 [103]. FPRs may be stimulated in intestinal epithelial cells not only by exogenous ligands, such as microbial formyl peptides, but also by endogenous anti-inflammatory and pro-resolution mediators, such as annexin A1, lipoxin A4, and resolving D1 [103,104]. Importantly, annexin A1 induces NOX1-derived O2-∙ via FPR1 to oxidize and inactivate redox-sensitive phosphatases PTP-PEST and PTEN, stimulating phosphorylation and activation of focal adhesion kinase (FAK) and paxillin that subsequently enhance cell migration and mucosal wound repair [105].

1.9.- Redox signaling triggered by commensal bacteria is mediated by NOX1 and DUOX2 The human gut microbiota is made up of 10-100 trillion microbes mostly distributed into two phyla: Bacterioidetes and Firmicutes [20,106–108]. Initially the gut is sterile in utero starting its colonization after birth, and yielding eventually a huge population of microbes especially abundant in the colon (≈1011-12 cells/g) and distal ileum (≈107-8 cells/g) [109,110]. Numerous beneficial functions have been reported for the symbiotic gut-microbe interaction including nutrition, host immunity, competitive exclusion of pathogenic microorganisms, metabolic regulation, as well as epithelial homeostasis related to

cell proliferation, migration, differentiation and barrier function

- 20 -

[106,109,111–113]. At least some of these important functions are related to the redoxbased response of intestinal epithelial cells to bacteria (see Table 1) [20]. Changes in oxygen partial pressure determine variation of redox potential along the gut. At colonic mucosa, oxygen partial pressure is below 25% of airborne oxygen content, moreover microbial metabolism causes reduction to a low redox potential (from 2200 mV to –300 mV) in the colon [114]. Accordingly, in the gut the fraction of strict anaerobes increases from proximal to distal, reaching 99% of bacterial species in the colon [115]. Intestinal epithelial cells exhibit TLRs and related nucleotide-binding oligomerization domain (NOD) proteins as well as FPRs to perceive “microbe associated molecular patterns” and signaling through these receptors contributes to the normal intestinal epithelial homeostasis without causing overt inflammation [102,116,117]. In intestinal epithelial cells NOX1 and DUOX2 are considered responsible for O2-∙ and H2O2 generation upon bacterial contact [102,118,119]. Nevertheless, ROS generation in intestinal epithelial cells largely varies upon exposure to different strains of commensal bacteria, being Lactobacilli the most potent inducers [20]. Indeed, although Lactobacilli rapidly generate ROS in CaCo-2 cells, neither Escherichia coli nor Bacteroides thetaiotaomicron produce ROS in vitro in these cells [120]. Commensal Lactobacilli stimulate FPR1 and rapidly generate physiological levels of ROS through NOX1 in the distal small intestine and colon in mice and stimulate ROSdependent cell proliferation in the colon [102,120]. Importantly, germ-free mice exhibit lack of ROS production and suppression of epithelial growth [120]. FPR1/NOX1dependent H2O2 induced by Lactobacilli trigger oxidation and inactivation of dual

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specificity protein phosphatase 3 (DUSP3), which stimulates ERK-mediated proliferation in intestinal epithelial cells [102,121,122]. Commensal bacteria control the ubiquitin-proteasome system in the gut through NOXderived H2O2, which provides an additional mechanism for stimulation of Wnt-driven proliferation [123]. Indeed, H2O2 generated through the interaction of Lactobacilli with intestinal epithelial cells inactivate the redox-sensitive NEDD8 ligase Ub‐like conjugating enzymes (Ubc) 12 by oxidizing its catalytic cysteine [123]. Ubc12 oxidation triggers rapid and reversible loss of the covalent neddylation of cullin-1, which is required for the function of the SCFβ-TrCP ubiquitin ligase formed by S-phase kinase-associated protein 1 (Skp1), cell division control protein 53 (Cdc53)/Cullin and F box receptor [124–126], blocking inhibitor kappa B (IκB) ubiquitination and consequently nuclear factor kappa B (NF-κB) activation [125]. The redox modulation of SCFβ-TrCP ubiquitin ligase by commensal bacteria also stimulates the Wnt/β-catenin pathway in vitro [123], but there is no experimental evidence in vivo in this regard yet. Recently it has been reported that the canonical Wnt ligand Wnt3A increases nuclear factor erythroid 2-related factor-2 (Nrf2) levels and its transcriptional activity in hepatocytes through disruption of its complex with Axin1/GSK-3/β-TrCP, abrogating GSK-3β-dependent Nrf2 phosphorylation and subsequent proteasomal degradation [127]. The role of Wnt signaling in the regulation of Nrf2 transcriptional activity and antioxidant defense in intestinal cells has not been explored yet and deserves further research. In any case, the link between NOX1 and Nrf2 at least exists in the gut because NOX1-derived ROS induced by Lactobacilli activate Nrf2 and up-regulate Nrf2dependent cytoprotective genes (Cyp2c65, Cyp2c55, Cyp4b1) in mouse colon [128]. Hence, Wnt and Nrf2 could be stimulated in parallel by NOX1 in the colon.

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Commensal bacteria are clearly required for normal epithelial dynamics and should modulate Wnt- and Notch- driven proliferation because it is enhanced by NOX1dependent O2-∙ generation. Hence, it would be worth assessing how redox signaling by different commensal bacteria affects the renewal rate of intestinal epithelial cells and the whole epithelium dynamics, particularly focusing on the delicate balance between NOX1, RAC1 and TIGAR described before. NOX1-derived H2O2 induced by certain commensal bacteria, such as Lactobacilli, may also contribute to mucosal would healing through focal adhesions. Indeed, FPR1/NOX1-derived H2O2 induced by Lactobacilli inactivate redox sensitive tyrosine phosphatases low molecular weight phosphotyrosine protein phosphatase (LMWPTPase) and SHP2 promoting phosphorylation and activation of focal adhesion kinase that increases the number of focal adhesions, enhancing epithelial cell migration and restitution after injury, which promotes mucosal wound healing [102,121,122]. DUOX2 is presently considered a key modulator of the symbiotic gut-microbiota interactions required to maintain intestinal immune homeostasis [94]. DUOX-derived H2O2 are required for control of intestinal microbiota in Drosophila [91]. In mammals our knowledge about how intestinal epithelial cells respond to non-pathogenic bacteria is still much limited [20]. In mice, Duox2 is markedly up-regulated in the ileum and colon epithelium in response to colonization by normal microbiota [92] or monocolonization by epithelial-attaching commensal segmented filamentous bacteria [94]. It has been proposed that direct epithelial contact would be required for Duox2 induction [94]. However, Duox2 expression is not induced in germ-free mice after colonization with different single commensal bacteria, such as Escherichia coli, Bifidobacterium longum, or Bacteroides thetaiotaomicron, and hence a single microbe-associated molecular pattern from any of these strains doesn’t seem to be the stimulus [92]. The mechanism

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involved in Duox2 up-regulation differs between the ileum and the colon [92], being mediated by NF-κB signaling and TIR-domain-containing adaptor protein including interferon-β (TRIF) in the ileum upon stimulation with IL-1β, but through MyD88 and p38 MAPK in the colon upon stimulation either with IL-1β or lipopolysaccharide [92]. Since DUOX2 may trigger a respiratory burst on intestinal epithelial cells it would be important to elucidate how commensal bacteria handle DUOX2 activation avoiding overt inflammation to maintain appropriate intestine homeostasis. Could DUOX2derived H2O2 up-regulate TIGAR or Nrf-2 in intestinal epithelial cells? Could certain commensal bacteria cause overexpression of DUOX2 and oxidative stress in the gut? Which is the impact of DUOX2 activation by commensal bacteria on intestinal epithelium dynamics? These questions remain to be answered and would help to identify potential pro-inflammatory conditions and niches with abnormal epithelium renewal that eventually may lead to loss of intestinal homeostasis.

2.- REDOX SIGNALING IN INFLAMMATION AND CANCER IN THE GASTROINTESTINAL TRACT 2.1.- Barrett’s esophagus and esophageal adenocarcinoma Patients with chronic gastroesophageal reflux exhibit esophagitis, and in around 15% of these patients the normal squamous esophageal epithelium is transformed into an intestinal-type columnar epithelium called Barrett’s esophagus, which greatly increases (125-fold) the risk of esophageal adenocarcinoma [129,130]. Indeed Barrett’s esophagus frequently progresses towards dysplasia and invasive carcinoma, whose incidence has risen markedly during the last decades in Western countries and exhibits high mortality -around 15.000 deaths each year in USA- [131]. Risk factors for Barrett’s

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esophagus include not only symptomatic gastroesophageal reflux, but also abdominal adiposity, and smoking [132]. Bile reflux is relatively common in patients with gastroesophageal reflux and it may also promote Barrett’s esophagus in patients with gastrectomy [133]. The development of Barrett’s esophagus and intestinal metaplasia towards adenocarcinoma is associated with chromosome aberrations, microsatellite instability, and epigenetic changes [134]. Genome-wide association studies (GWAS) have found increased adenocarcinoma risk in genetic loci related to DNA-repair capacity, telomere length, and mitochondrial DNA copy number [134]. A recent GWAS has identified increased risk for Barrett’s esophagus in single-nucleotide polymorphisms of the cyclooxygenase pathway and microsomal glutathione-S-transferase 1 [132]. Cyclooxygenase-2 (COX-2), the rate-limiting enzyme for the formation of prostaglandins and thromboxanes from arachidonic acid, is overexpressed in Barrett’s esophagus and esophageal adenocarcinoma in comparison with normal esophageal mucosa [129,133]. COX-2 is also highly expressed in gastric cancer and colorectal carcinoma and exhibits mutagenic and tumorigenic activity [129]. Hence COX-2 seems to be involved in the pathogenesis of cancer in the gastrointestinal tract. Interestingly, unconjugated bile acids potently induce COX-2 in esophageal adenocarcinoma cells via cAMP response element binding protein (CREB) through ROS-mediated activation of phosphatidylinositol 3-kinase (PI3K)/AKT and ERK-1/2 [133]. In fact, O2-∙ generation and lipid peroxidation levels are increased in esophagitis and Barrett’s epithelium, whereas superoxide dismutase activity is markedly diminished in these esophageal disorders [135–137]. Paradoxically the protein steady state levels of both Cu/Zn superoxide dismutase (Cu/ZnSOD) and Mn superoxide dismutase (MnSOD) are up-regulated in esophagitis and Barrett’s esophagus [137]. The increased

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nitration of MnSOD may explain the decrease in its activity [137] as Tyr34, present in the active site of human MnSOD, is the most susceptible for nitration. Nevertheless, oxidation of critical tyrosine residues of MnSOD in addition to nitration is required for complete inactivation of the enzyme. The nitration or oxidation of tryptophan 32 in Cu/ZnSOD may also lead to its partial inactivation. The critical role of O2-∙ overproduction in the pathogenesis of esophagitis and esophageal adenocarcinoma is supported by the efficient treatment with SOD, which prevented esophagitis induced by duodenogastroesophageal reflux in rats [138] and markedly reduced the risk of developing esophageal adenocarcinoma [139]. Other antioxidants, such as resveratrol, protect the esophageal epithelium restraining reflux esophagitis and its progression to Barrett’s esophagus and esophageal adenocarcinoma in animals [2,140]. Among the different NOXs, NOX5-S -one of the NOX5 isoforms- seems to play a major role in the enhanced O2-∙ production in Barrett’s esophagus as it is overexpressed in Barrett’s mucosa [141]. This overexpression of NOX5-S is mediated by the calciumdependent activation of Rho kinase ROCK2 [142]. Moreover, NOX5-S triggers the acid-induced generation of H2O2 and DNA damage in Barrett’s cells and esophageal adenocarcinoma cells [143]. H2O2 generation also seems to be enhanced in Barrett’ esophagus through downregulation of GPx7 by promoter DNA methylation [144]. Indeed, GPx7 decreases bile acid-induced H2O2 generation, oxidative DNA damage and double-strand breaks in esophageal cells, whereas its silencing enhances all these H2O2-dependent processes [145]. GPx7 silencing also promotes tumor necrosis factor-alpha (TNF-α)-induced NFκB activation as GPx7 abrogates NF-B signaling upstream via degradation of TNF receptor 1 (TNFR1) and TNF receptor-associated factor 2 (TRAF2) [146]. Importantly,

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GPx7 exhibits tumor suppressor functions in vivo by promoting cell senescence and blockade of G1/S progression [144]. A decrease in autophagy might also contribute to the enhanced ROS generation associated with the pathogenesis of Barrett’s esophagus, since inhibition of autophagy upon exposure to acid leads to increased ROS generation and cell death in human Barrett’s esophagus cell lines [131], and autophagic vesicles are markedly reduced in esophageal adenocarcinoma [131]. Paradoxically, the increased ROS generation in chronic esophagitis may be accompanied by clear anti-apoptotic mechanisms, such as increased glutathione levels and over-expression of

apurinic/apyrimidinic endonuclease-1

(APE-1). Thus,

experimental acute esophagitis in rats triggers a transient decrease in GSH levels followed by an increase in its levels [138,147]. GSH levels are high in esophageal mucosa of patients with chronic esophagitis [137]. Furthermore, many cases of esophageal adenocarcinoma (> 50%) exhibit over-expression of APE-1, which protects against acidic bile salts-induced oxidative DNA damage, DNA breaks, and apoptosis through down-regulation of c-Jun N-terminal kinase (JNK) and p38 MAPKs [148]. Chronic esophagitis and Barrett’s esophagus should be examples of translational research in which antioxidant therapy could provide clear beneficial effects to hinder their progression to esophageal adenocarcinoma. Which are the best antioxidants and at which doses still needs to be established in patients. Regarding redox signaling, the increase in GSH levels upon chronic esophagitis suggests that Nrf2 may be activated and might contribute to the anti-apoptotic mechanisms involved in the progression towards esophageal adenocarcinoma, which deserves further research. In addition, new insights would be underway regarding the investigation of the potential relationship between O2-∙ and H2O2 overproduction, activation of Wnt/Notch signaling, and

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epigenetic changes in proto-oncogenes and tumor suppressor genes in the pathogenesis of esophageal adenocarcinoma.

2.2.- Peptic ulcer Peptic ulcer disease includes both gastric and duodenal ulcers [149]. This condition affects more than 4 million people each year involving high costs for the health system [150]. The main symptoms of peptic ulcer are epigastric pain and dyspeptic symptoms such as fullness, bloating, early satiety, and nausea, although chronic ulcers can be asymptomatic [149]. Complications for the peptic ulcer disease include perforation, bleeding and obstruction. In general, peptic ulcer disease is caused by an imbalance between ulcerative factors and protective factors of gastric mucosa. Environmental and endogenous factors may contribute to the appearance of ulcers, increasing gastric acid and pepsin production or weakening the gastric mucosa barrier [149,151]. Currently, the two major etiological factors of peptic ulcers are Helicobacter pylori infection and excessive use of nonsteroidal anti-inflammatory drugs (NSAIDs). Risk factors for peptic ulcers are smoking, alcohol consumption and psychological stress [149,152]. 2.2.1.- Helicobacter pylori-induced ulcer Nowadays there is no doubt about the causal relationship between H. pylori infection and the development of gastric and duodenal ulcers [153,154]. In fact, it has been shown that H. pylori eradication is an adequate therapy for treatment of peptic ulcer [155,156]. Therefore, since Marshall and Warren discovered Gram-negative bacterium H. pylori in 1984 in patients with gastritis and peptic ulcer disease [157], the pathogenesis of this bacteria has been studied thoroughly [158]. H. pylori colonizes the gastric epithelium and modifies the pH of the gastric mucosa. This is due to H. pylori

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urease activity that produces ammonia from urea, which neutralizes HCl on the gastric mucosa. As a result, an inadequate amount of somatostatin is released increasing gastrin release and acid production by gastric mucosa [159,160]. Furthermore, H. pylori releases chemotactic factors, such as N-formyl-methionyl-leucyl phenylalanine peptide, and causes an inflammatory response in gastric mucosa, promoting IL-8 and IL-1β release from gastric epithelial cells [161,162]. This promotes infiltration of neutrophils and macrophages to the gastric mucosa, enhancing the inflammatory response and ROS production [2,163,164]. Additional pathogenic factors in type I H. pylori bacterial strains are vacuolating cytotoxin A (VacA), and a pro-apoptotic cytotoxin-associated protein gene A (CagA) [165,166] (Figure 3). H. pylori stimulates production of O2-∙ and H2O2 mainly through the associated inflammatory infiltrate [167,168]. Indeed neutrophils infiltrated in gastric mucosa, as a result of the inflammatory process triggered by H. pylori infection, seem the most important source of O2-∙ and H2O2 [163,164]. The oxidation of chloride with H2O2 catalyzed by neutrophil myeloperoxidase (MPO) activity leads to formation of OCl-, which

can

react

with

ammonia

produced

by bacterial

urease

generating

monochloramine (NH2Cl), a potent lipophilic oxidizing compound that crosses biological membranes causing oxidation of intracellular components of gastric cells [169]. H. pylori may also trigger ROS generation through NOXs in non-phagocytic cells. Thus, H. pylori membrane lipid A activates TLR4 receptor to induce expression of NOX components and O2-∙ production in gastric mucosal cells [170]. Indeed, H. pylori triggers the production of O2-∙ and activates the transcription of Nox1 and its organizer Noxo1 through Rac1 in gastric mucosal cells [171].

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Oxidative stress contributes to the inflammatory response during H. pylori infection through multiple redox signaling pathways. In fact, ROS induce activation of NF-κB and activator protein 1 (AP-1), which both enhance IL-8 transcription in gastric epithelial cells promoting the inflammatory response in gastric mucosa [172,173]. Nevertheless, ROS production may also have a protective role as gastric microbicide during H. pylori infection. The role of NOX2 in the immune defense against H. pylori was demonstrated in mice with chronic granulomatous disease caused by a genetic defect in NOX2 that prevents the production of oxidizing species by this enzyme. These mice exhibited enhanced tissue damage upon infection by H. pylori, which correlates with absence of oxidants derived from neutrophil infiltrates [174]. Although H. pylori infection may increase the activities of some antioxidant enzymes, such as catalase, glutathione peroxidase and specially MnSOD [175–177] as adaptive mechanism, glutathione depletion [178,179] as well as oxidative inactivation of gastric mucosa peroxidase [180] lead to increased lipid peroxidation and membrane damage exacerbating peptic ulcer disease [176,181]. Furthermore, those strains of H. pylori that express heat-shock protein B (HspB) may interfere with the antioxidant response regulated by Nrf2/Kelch-like ECH-associated protein 1 (Keap1) (Figure 3). In fact, HspB increases KEAP1 expression but decreases expression of NRF2 and target antioxidant enzymes, enhancing the inflammatory response [182]. Interestingly, it has been reported that chronic infection with H. pylori HspB+ strains increases the risk of gastric carcinoma [183]. The intracellular antioxidant defense should restrain the inflammatory response, but it is impaired by H. pylori infection. H. pylori γ-glutamyl transpeptidase (GGT) is a virulence factor that can induce apoptosis and enhance the inflammatory response in

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gastric epithelial cells. Bacterial GGT uses gastric extracellular glutamine and GSH for amino acid bacterial metabolism, which causes gastric glutamine and GSH depletion leading to DNA damage and enhanced NF-κB activation with the corresponding IL-8 release [184]. In addition, cysteine rich protein metallothionein protects against NF-κB activation as metallothionein knock-out mice infected with H. pylori exhibit enhanced NF-κB-driven inflammatory response and injury in gastric mucosa [185]. Gastric homeostasis depends on the balance between proliferation and cell death of gastric epithelial cells. This balance may be disrupted due to the injury and chronic oxidative stress triggered by H. pylori infection. Oxidative stress associated with H. pylori infection induces cell apoptosis through the mitochondrial pathway in human gastric epithelial cells [186,187]. However, in gastric mucosa the situation is more complex because adaptative anti-apoptotic mechanism may be triggered. Thus, APE1/redox factor 1 (Ref-1) is markedly up-regulated by H2O2 and also by ROS induced by H. pylori infection in human gastric epithelial cells [188]. APE-1 is a negative regulator of NOX1-driven O2-∙ generation induced by H. pylori [189]. Thus, APE-1 interacts with Rac1 abrogating NOX1 up-regulation and the corresponding O2-∙ generation induced by H. pylori infection [189] (Figure 3). Hence, APE-1 up-regulation restrains the oxidative stress associated with a chronic H. pylori infection. Unlike ROS, nitrogen free radicals seem to play a protective role in gastric mucosa against infection of H. pylori [2]. H. pylori infection increases the expression of inducible nitric oxide synthase (iNOS) and COX-2 in gastric mucosa to promote NO production through NF-κB activation, which decreases apoptosis in gastric epithelial cells [190–193]. Nevertheless, H. pylori expresses arginase that degrades arginine, the substrate of iNOS for NO production, which is a defense strategy against the immune

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response mediated by antimicrobial NO [194]. Gastric NO can react with O2-∙ generated during H. pylori infection producing peroxynitrite that could affect the survival of H. pylori in gastric mucosa [195]. However, the urease activity of H. pylori produces NH3 and CO2, which facilitates the peroxynitrite reaction with CO2 to produce NO3-, reducing the oxidizing ability of peroxynitrite [196]. 2.2.2.- NSAIDs-induced ulcer Excessive consumption of NSAIDs is another principal etiological factor of peptic ulcer. In the pathogenesis of NSAIDs-inducing ulceration, prostaglandin-dependent mechanisms and prostaglandin-independent mechanisms are involved [149,152]. The integrity of gastric mucosa depends on the synthesis of prostaglandins by cyclooxygenase. Continuous production of prostaglandins, such as PGE2 and PGI2, in gastric mucosa is a critical factor to maintain mucosal integrity protecting against injuring factors. Prostaglandins reduce acid release, stimulate mucus, bicarbonate and phospholipid production, increase mucosal blood flow, and inhibit inflammatory response [197]. The main deleterious effects mediated by NSAIDs occur through inhibition of prostaglandin production by COX-1 [149,152] Regarding the prostaglandin-independent NSAID-induced ulceration, the inflammatory response and excessive ROS production are the most important pathogenic factors. Excessive consumption of NSAIDs promotes neutrophil infiltration and release of neutrophil-derived factors to the gastric mucosa [198]. The increased level of TNF-α is the primary initiating event and induces gastric ulceration [199]. Indomethacin-induced gastric mucosa damage is mediated, at least in part, by increased production of hydroxyl radical and glutathione depletion [200–202] as well as by decreased activities of glutathione peroxidase [200] and gastric peroxidase [202]. Oxidative stress associated

- 32 -

with NSAID consumption leads to cell apoptosis through the mitochondrial pathway. Indomethacin disturbs the morphology and functionality of mitochondria and causes mitochondrial oxidative stress increasing mitochondrial lipid peroxidation, inducing intra-mitochondrial free iron accumulation and the generation of O2-∙, H2O2 and hydroxyl radical [203]. However, multiple intracellular antioxidant mechanisms related to thioredoxin and Nrf2/ heme oxygenase-1 (HO-1) have an important protective role inhibiting NSAIDinduced apoptotic signals and gastric injury progression. Thus, the antioxidant action of thioredoxin prevents gastric damage induced by indomethacin inhibiting the upregulation of apoptotic signals [204]. Up-regulation of HO-1 also protects gastric mucosa during indomethacin-induced gastric ulceration by inhibiting apoptosis [205]. Indomethacin stimulates p38 MAPK-dependent activation of Nrf2, which binds to the HO-1 promoter up-regulating its transcription in gastric mucosal cells, although it is still unclear how it activates p38 MAPK [205]. Additionally, Nrf-2-dependent HO-1 upregulation is triggered by the product of lipid peroxidation 4-hydroxynonenal (HNE) in gastric macrophages. Indeed, activation of the HNE-Nrf2-HO-1 pathway in macrophages, located in the periphery of the gastric mucosa during NSAID-induced gastric ulceration, leads to the activation of the adaptive response through up-regulation of Ho-1 [206]. HO-1 catalyzes the conversion of heme into Fe2+, carbon monoxide and biliverdin, being the latter reduced to bilirubin in macrophages. Biliverdin and bilirubin have antioxidant properties and carbon monoxide acts as anti-apoptotic signal molecule [205]. On the other hand, HO-1 also acts as an anti-apoptotic molecule by preventing mitochondrial oxidative stress. During indomethacin-induced gastric ulceration,

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mitochondrial accumulation of heme occurs probably due to erythrocyte lysate generated from bleeding or erosions during gastric ulceration. When heme accumulates into the mitochondria, increased ROS generation acts as a signal for Nrf2 activation inducing HO-1 expression. This leads to the detoxification of intra-mitochondrial heme and thus to diminished mitochondrial generation of ROS, preventing mitochondrial pathology and cellular apoptosis. Although it is true that heme metabolism by HO-1 generates iron, which could enhance ROS generation by the Fenton reaction, increased mitochondrial ferritin becomes an additional mechanism to rescue mitochondria from oxidative damage induced by indomethacin treatment [207]. Moreover, the activity of HO-1 may prevent induction of inflammatory response abrogating NF-κB activation [208,209]. MMPs also play an important role in indomethacin-induced gastric ulceration and their levels are redox regulated. MMPs are involved in degradation and remodeling of the extracellular matrix and ROS production modulates the expression of MMP genes. During indomethacin-induced gastric ulceration, H2O2 production leads to suppression of MMP-2 transcription by inhibiting the ERK/SP1 pathway [210–212]. However, the effect of NSAIDs-derived ROS towards promoting MMP-9 transcription is different and seems to be mediated by NF-κB and AP-1 [211,213]. Accordingly, MMP-9 but not MMP-2 promoters contain several putative NF-κB and AP-1 binding sites [214]. Interestingly, the activity ratio MMP-9/MMP-2 increases in parallel with the ulcer index [211]. Probably MMP-2 participates in gastric mucosa homeostasis, whereas MMP-9 would be involved in degradation of the extracellular matrix associated with ulceration [211,214].

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Consequently, in H. pylori-induced ulcer the oxidative stress associated with gastric ulcerogenesis is mediated mainly by NOX2 and MPO from infiltrated leukocytes as well as by chloramines produced during the infection and by cytotoxins released by certain bacterial strains, whereas in NSAIDs-induced ulcer TNF-alpha together with infiltrated neutrophils would be responsible for the associated oxidative stress. It remains to be elucidated the role of NOX1 from gastric epithelial cells in NSAIDsinduced ulcer and also the potential pathogenic role of ROS-dependent MMP-9 upregulation requires confirmation. In addition, it should be further investigated the controversial role of anti-apoptotic mechanisms, such as up-regulation of APE-1 or Nrf2/heme oxygenase, that protect against inflammation, mucosal damage and ulcer development, but might promote the progression of gastritis towards gastric cancer.

2.3.- Gastric cancer There are two clinical types of gastric carcinoma, the intestinal and the diffuse one [215]. In chronic atrophic gastritis, gastric glands are replaced by intestinal-type epithelium that eventually may evolve into metaplastic glands, dysplasia and carcinoma of the intestinal type [215,216]. The precancerous phase of intestinal-type gastric carcinoma is characterized by inflammation (chronic gastritis), multifocal atrophy (gland loss), and eventually intestinal metaplasia, dysplasia, and anaplasia [216]. This precancerous stage does not occur in diffuse gastric carcinoma. Infection by H. pylori is the major cause of gastric cancer, which is the fourth most common cancer and the second major cause of cancer death [2,3,215,217]. Nevertheless, only around 3% of H. pylori-infected patients develop gastric cancer within 8 years [218], because bacterial, host, and population-related factors contribute to the development of gastric carcinoma [3,217]. Infection by H. pylori triggers infiltration

- 35 -

of leukocytes in the gastric mucosa, but generally neutrophils, macrophages and lymphocytes cannot cope with the bacteria and chronic inflammation develops [3]. H. pylori may also activate the ubiquitous and latent Epstein-Barr virus present in the gastric mucosa promoting gastric carcinoma [219]. Indeed, the potent oxidizing NH2Cl generated from the reaction of NH3 with the hypochlorous acid (HOCl) produced by neutrophil MPO [3] triggers the viral lytic of the Epstein-Barr virus aggravating mucosal injury [219]. Strikingly, H. pylori exhibits an injection system able to cross the membrane of gastric epithelial cells to inject into the cytosol bacterial proteins, such as CagA [3]. In Western countries, patients infected with CagA-positive strains exhibit higher risk for atrophic gastritis, peptic ulcer, and gastric cancer [3,220,221]. The CagE protein seems critical for H. pylori-induced gastritis and associated cancer since cagE deficient animals do not develop gastritis nor metaplasia in gastric mucosa [222]. Levels of O2-∙ and H2O2 detected by chemiluminiscence were elevated in gastric mucosa from infected patients, being even higher upon infection with H. pylori strains harboring the cytotoxin Cag pathogenicity island [169,186]. CagA-positive strains trigger an intense ROS production associated with an oxidative burst in neutrophils [3]. In addition, H. pylori activates non-phagocytic NOX in gastric pit cells [223]. Although malondialdehyde (MDA) levels were high in tumor-free mucosa of patients with gastric adenocarcinoma, the progression towards adenocarcinoma seems to change the redox phenotype favoring cell proliferation as adenocarcinoma specimens exhibited less MDA levels and high GSH levels, as well as low NOX activity due to marked downregulation of NOX2 and 4 [224]. Interestingly, NOX1 was over-expressed in gastric adenocarcinoma [224]. The up-regulation of NOX1, but not of other isoforms of NOXs,

- 36 -

suggests that compartimentation of O2-∙ generation might occur in oncogenesis, particularly in the outer membrane acting on receptors that trigger cell proliferation. H2O2 production is also enhanced in gastric epithelial cells by H. pylori through upregulation of spermine oxidase, which produces H2O2 in the conversion of polyamine spermine into spermidine leading to oxidative DNA damage and apoptosis [225]. Spermine oxidase expression increases in gastritis and gastric carcinoma, with its highest levels upon infection with CagA positive strains and in intestinal metaplasia [225]. EGFR mediates the up-regulation of pro-apoptotic spermine oxidase in gastric epithelial cells, but paradoxically some cells exhibiting high phosphorylated EGFR avoid apoptosis via erythroblastic leukemia associated viral oncogene B-2 (ERBB-2) [225], which additionally may promote oncogenesis and tumor progression through tyrosine kinases. This mechanism might mediate gastric carcinogenesis as phosphoEGFR and phosho-ERBB-2 levels are elevated in gastric mucosa from patients who eventually develop gastric intestinal metaplasia or dysplasia [225]. Although H. pylori triggers apoptosis in vitro in gastric epithelial cells and the gastric epithelium of infected patients exhibits higher number of apoptotic cells than that from non-infected subjects, increased cell proliferation is found in vivo upon H. pylori infection [186]. The mechanism for escaping apoptosis may be related to the APE-1 upregulation found in H. pylori-infected gastritis and gastric carcinoma [226]. Indeed, suppression of APE-1 results in increased apoptosis in gastric epithelial cells after H. pylori infection [227] and the DNA-repair activity of APE-1 together with the transcriptional activation induced by APE-1/Ref-1 can inhibit both the mitochondrialdependent intrinsic and caspase 8-dependent extrinsic pathways of apoptosis in gastric epithelial cells [227]. In addition, H. pylori triggers p300-driven acetylation of nuclear APE-1, which forms a repressor complex with histone deacetylase-1 (HDAC-1) that

- 37 -

binds to the BCL2-associated X protein (Bax) promoter inhibiting its expression [228] (Figure 3). Thus, H. pylori may suppress cell apoptosis by up-regulating APE1 and inducing acetylation of APE-1 preventing the expression of Bax. Therefore, APE-1 enhances the anti-apoptotic mechanisms that can act as a double edge sword because they may promote the development of gastric cancer as a result of chronic H. pylori infection. In addition, APE-1 is involved in hypoxia inducible factor-1 alpha (HIF-1α) stabilization induced by H. pylori-generated ROS [229]. In fact, the induction of APE-1 by H. pylori infection increases the expression of HIF-1α in gastric epithelial cells, which is associated with the progression of gastric cancer [229]. APE-1 also modulates the reduction of cysteine residues in redox-sensitive transcription factors, such as AP-1 and NF-B, promoting their DNA binding activity and the up-regulation of IL-8 in gastric epithelial cells [173], which would enhance neutrophil infiltration. Trefoil factors (TFFs) are conserved small proteins synthesized and secreted by mucussecreting cells as components of the protective mucus layer, which may act as tumor suppressors. TFF1, secreted by pit cells in gastric epithelium, is required for normal differentiation of the gastric mucosa [230] and inhibits the growth of a human adenocarcinoma cell line [231]. Hence, TFF1 is considered a tumor suppressor gene. TFF1 also suppresses TNF-α-induced NF-B activation and cell growth [232]. As TFF1 expression is lost in most specimens of gastric carcinoma, the lack of TFF1 may contribute to the aberrant NF-B activation associated with gastric carcinoma [232]. The impact of the increased generation of NOX1-derived O2-∙ and spermine oxidase derived-H2O2 on Wnt and Notch signaling in the gastric mucosa still remains to be investigated. The H2O2-dependent stimulation of cell proliferation in the gastric epithelium is likely to contribute decisively to oncogenesis in the stomach.

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2.4.- Ischemic intestinal injury Ischemic intestinal injury casued by intestinal ischemia-reperfusion may occur in patients with abdominal trauma, major surgery, strangulated hernias, severe burns as well as septic and hypovolemic shock [233,234]. Mucosal injury and hemorrhage after intestinal ischemia are mainly associated with the reperfusion phase, which largely contributes to O2-∙ and H2O2 generation from infiltrated neutrophils and xanthine oxidase [2,235,236]. Ischemia-reperfusion in the gut may trigger translocation of bacteria and endotoxin that activates TLRs causing neutrophil recruitment and tissue damage not only in the intestine but also in lungs [237]. Intestinal ischemia-reperfusion leads to enhanced levels of MDA and protein carbonyls, but diminished activities of SOD and GPx [234]. All these changes as well as intestinal injury were restrained by antioxidant treatment with erdosteine and ebselen [234]. The major form of ischemic intestinal injury in elderly patients is ischemic colitis, which may be occlusive or non-occlusive [2]. The former is caused by strangulated hernias, volvulus or thrombus that blockade the blood flow in the colon, whereas the latter may be triggered by low intestinal blood flow induced by hypotension or mesenteric atherosclerosis. Mild ischemic colitis is associated with necrosis or ulceration, but patients with severe ischemic colitis also exhibit sepsis or fatal infarctions complicated with gangrene [2]. Ischemic colitis may eventually arise in vascular surgery, vascular thromboembolism, and treatment with vasoconstrictors and is associated with high morbidity and mortality [238]. Ischemic colitis leads to increased lipid peroxidation and oxidative stress-associated NF-kB activation that triggers TNF-α production from intestinal myocytes [233].

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Antioxidants, such as allopurinol and SOD, greatly ameliorate the inflammatory response and tissue damage in experimental reperfusion injury [2,235]. The redox modulation of the acute inflammatory cascade in intestinal ischemia and reperfusion is common to other inflammatory disorders and it would be worthwhile to elucidate the role of redox-sensitive phosphatases and redox signaling in epigenetic regulation of pro-inflammatory cytokines under this condition and during the resolution process. In addition, ischemic intestinal injury is another target of translational research in redox Medicine involving antioxidant therapy. The acute local and systemic inflammatory response associated with this condition may be ameliorated by certain antioxidants and the efficient doses should be clearly established.

2.5.- Celiac disease 2.5.1.- Clinical aspects and pathogenesis of celiac disease Celiac disease (CD) is an autoimmune heritable condition characterized by chronic inflammation of the small intestine due to permanent intolerance to gluten proteins. In fact, there is a massive, pro-inflammatory and pathogenic immune response towards certain parts of gluten and also towards the intestinal tissue itself resulting in structural changes. Gluten is a constituent derived from cereal grains of wheat, barley and rye, which is present in the common daily diet in the Western world [239]. Over the last decades, the prevalence of CD has increased dramatically and represents the most chronic gastrointestinal disease in Europe, affecting around 1% of the white ethnic population in USA and more than 10% of type 1 diabetic patients [240,241]. Pediatric CD patients often exhibit distinct clinical symptoms such as diarrhea,

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malnutrition, weight loss, weakness and failure to thrive, although the classical presentation is characterized by small-intestinal villous atrophy and crypt cell hyperplasia [242,243]. In adults, a wide variety of symptoms can occur, but many are often diffuse and not readily associated with an intestinal pathology [242,243]. Furthermore, some individuals may have serological and histological findings compatible with the disease, but with no clinical symptoms. Removal of gluten from the diet results in a clinical improvement in CD patients as it is required not only for the disease to precipitate, but also to drive inflammation towards resolution. Accordingly, the morphology of the intestine normalizes and clinical symptoms are reduced in most patients (> 80%) upon a gluten-free diet [244]. Despite decades of research, the mechanisms behind CD etiology are still not fully understood, although it is clear that a complex interplay between genetic and environmental factors is involved [245,246]. The genetic predisposition for development of CD has been broadly studied by CD GWAS, showing risk variants in the human leukocyte antigen (HLA) region, specially HLA-DQ2.5, which is present in most of patients (90%) [247,248]. Nevertheless, HLA-DQ2.5, HLA-DQ2.2, or HLADQ8 risk alleles are necessary but not sufficient for celiac disease development, as they are highly prevalent in the general population. HLA-DQ molecules predispose to disease by preferential presentation of gluten antigens to CD4+ T cells to induce an adaptive immune response [249]. The environmental stimuli responsible for development of intestinal damage associated with CD are prolamin fractions in cereal grains (gliadin in wheat, hordeins in barley, and secalins in rye) [246,250]. The gliadin sequence contains digestion-resistant regions rich in glutamine and proline, which play a special role in CD pathogenesis: some

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undigested peptides, mainly the 33-mer (P55-P87) and the 25-mer (P31-P55), exert a cytotoxic activity or immune-modulatory activity [251,252]. Tissue transglutaminase (TG) 2 deamidates the gliadin 33-mer increasing its immunogenicity as it binds to HLA-DQ2.5, HLA-DQ2.2, or HLA-DQ8 to trigger a potent and pathogenic adaptive Th1 CD4+ T-cell response [245,253]. The gliadin 25mer gives rise to the P31-P43 peptide that triggers a stress response with up-regulation of heat shock proteins [254], and an innate immune response mainly through IL-15 [255,256] causing injury to the intestinal mucosa [250,254]. Moreover, gliadin peptides may induce endoplasmic reticulum-stress that can be relevant in CD pathogenesis [257]. The stress responses in the intestinal epithelium are considered essential for the development of CD and its associated intestinal mucosa damage [258]. Therefore, the adaptive immune response to gluten appears to act in synergy with epithelial stress to allow intraepithelial cytotoxic T cells to kill epithelial cells and induce villous atrophy in patients with CD [254]. A hallmark of CD is the gluten dependent production of IgA autoantibodies toward the self-protein TG2. Hence, inflammation of the intestinal mucosa that occurs in genetically predisposed individuals is due to a complex interplay between innate and adaptive immune responses to ingested gluten [250,259]. 2.5.2.- Oxidative stress in celiac disease The relationship between oxidative stress and CD is supported by many studies on intestinal biopsies, plasma and blood cells of patients. An increase in lipid peroxidation (thiobarbituric acid-reactive substances and lipid hydroperoxides), protein carbonyls, and oxidative DNA damage 8-oxo-7,8-dihydro-2’-deoxyguanosine (8-OHdG) was demonstrated in intestinal cells and biological fluids of celiac patients with respect to controls [260–262]. Oxidative stress mediates most of the cytotoxic effects, such as

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apoptosis and impaired cell differentiation, induced by gluten peptides in intestinal epithelial cells. In fact, accumulation of some gliadin peptides, such as P31–43, in lysosomes leads to increased levels of O2-∙, H2O2, peroxynitrite, and lipid peroxides [263]. Different in vitro cell models have shown that gliadin increases the levels of lipid peroxidation products, such as HNE, as well as glutathione oxidation, but decreases protein thiols [264,265]. Besides, there is a close relationship between the redox state of enterocytes and their susceptibility to gliadins [264]. The activities of glutathione peroxidase and reductase as well as GSH levels were decreased in small intestine from celiac disease patients, although Cu/ZnSOD activity was increased in patients with active disease [266,267]. The decrease in glutathione peroxidase activity may be explained, at least in part, by the selenium deficiency associated with chronic inflammatory damage of the small intestine [268]. Additionally, the absorption of vitamin E as well as other vitamins is reduced in celiac disease patients. A decreased mRNA expression of the antioxidant and anti-inflammatory paraoxonase enzymes PON1 and PON3 was reported in intestinal biopsies of celiac patients [269]. Consequently, the inflamed mucosa would be more susceptible to oxidative tissue damage due to the general decrease in antioxidant defense, hampering the recovery of the mucosa and its epithelial integrity. Hence, oxidative stress has been considered as one of the major mechanisms involved in celiac disease pathogenesis and progression [270], specially in gliadin cytotoxicity, affecting cell morphology, proliferation, and viability [264,271].

2.5.3.- Redox signaling in celiac disease

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The redox imbalance induced by gliadin peptides in intestinal epithelial cells contributes to NF-κB activation and subsequent up-regulation of pro-inflammatory cytokines and adhesion molecules [272–274]. NF-κB and AP-1 transcriptional activities are dependent on the intracellular redox state, which can promote synergistically transcriptional activity of pro-inflammatory genes [273]. NF-κB activation increases the expression of COX-2 and iNOS with consequent higher production of prostaglandins and NO metabolites, contributing to inflammation and nitrosative stress [272,275]. Uncontrolled activation of the ROS-TG axis leads to down-regulation of peroxisome proliferatoractivated receptor gamma (PPARγ), lowering its “trans-repressing” activity towards NF-κB and the inflammatory response [265]. Interestingly, ROS generated by the gliadin peptide p31-43 inhibits tissue TG2 ubiquitination reducing its degradation by the ubiquitin-proteasome system, thus leading to increased TG2 protein levels [265]. Nitrosative stress also seems to be involved in CD pathogenesis. High levels of NO are present in serum [276,277] and urine [278] of celiac children and correlate with increased expression of iNOS in the small intestine [279]. iNOS is constitutively expressed in human duodenal enterocytes and its activity is increased in patients with untreated CD [280], but this is partially corrected when patients follow a gluten-free diet [281]. On the other hand, nitrosative stress may contribute to the loss of barrier function in the small intestine of celiac disease patients. Indeed, nitric oxide may decrease epithelial barrier function in CaCo-2 monolayers by down-regulating the expression of zona ocludens-1 [282] and intestinal biopsies of these patients exhibit reduced levels of zona ocludens-1 [283,284]. Hence, ROS not only mediate the cytotoxic effects of gliadine peptides but also enhance the inflammatory cascade via NF-B, and the pathogenic role of tissue

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transglutaminase. In addition, nitrosative stress promotes the impairment of tight junctions in celiac disease. Further research should be focused on the potential role of NOXes in gluten-derived production of O2-∙ and H2O2 in intestinal epithelial cells and the possible lack of Nrf2 activation that would explain glutathione depletion in this condition of chronic mucosal inflammation.

2.6.- Inflammatory bowel disease Inflammatory bowel disease accounts for idiopathic chronic and relapsing inflammatory disorders of the gut, which comprise mainly Crohn’s disease and ulcerative colitis and affect millions of people throughout the world [285]. The prevalence is higher for ulcerative colitis than for Crohn’s disease, with the highest rates of ulcerative colitis found in North America and northern Europe (156-291/100.000 persons) [286]. Interestingly, developed countries exhibit higher incidence of ulcerative colitis than developing countries, and accordingly urban areas show higher incidence than rural areas [286]. Inflammation in Crohn’s disease is typically segmental and transmural, commonly affecting the terminal ileum [287]. The first lesions usually arise in Peyer’s patches, and granulomas containing macrophages are considered features of Crohn’s disease [287]. However, mucosal inflammation in ulcerative colitis starts in the rectum and extends proximally in a diffuse and continuous manner through the colon [287]. Instead of granulomas, micro-abscesses containing neutrophils develop in crypts and lamina propria in ulcerative colitis [287]. Nevertheless, common mechanisms in both Crohn’s disease and ulcerative colitis are the decrease in mucin secretion that favors the extensive superficial ulceration of the mucosa in ulcerative colitis and mucosal lesions

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in Crohn’s disease, as well as the impairment of tight junctions that leads to loss of epithelial barrier and increased permeability to bacterial antigens [286,288]. GWAS have identified a large number of gene polymorphisms associated with inflammatory bowel disease (71 associated with Crohn’s disease and 47 to ulcerative colitis, being 17 in common), most of them related to NOD2, TLRs, and autophagy in the case of Crohn’s disease, and to HLA class II genes and epithelial defense function (cadherin, laminin β1, and MUC19) in ulcerative colitis [286,288–291]. Hence, it is noteworthy that gene variants of pattern recognition receptors, which recognize specific bacterial molecular patterns, and HLA II that act in the interplay between host cells and pathogens confer risk for inflammatory bowel disease [290]. Therefore, commensal bacteria that behave as pathogens or pathogenic microbes may also contribute to chronic inflammation in these patients [290]. Both innate and adaptive immunity are affected in inflammatory bowel disease [287]. Nevertheless, the adaptive immune system seems to maintain gut inflammation in Crohn’s disease, but not to initiate it [288]. A key pathophysiological mechanism is the imbalance between Th cells and regulatory T cells and particularly the loss of tolerogenic regulatory T cells. In Crohn’s disease the excessive immune response seems to occur via Th1 and Th17 cells, whereas ulcerative colitis would be mediated by Th2 cells [287,288,292]. Microbes stimulate the Th1 response that produces interferon-γ and TNF-α, which largely promotes intestinal damage in Crohn’s disease [290]. Accordingly, in patients with this disease NF-κB is activated in the inflamed mucosa, not only in leukocytes but also in epithelial cells [293]. Pathogenesis in ulcerative colitis is mediated by natural killer T cells that produce Th2 cytokines, particularly IL-13 that causes apoptosis in epithelial cells and triggers a positive feedback on natural killer T cells [286,294].

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2.6.1.- Oxidative stress in inflammatory bowel disease Marked glutathione depletion and oxidation have been reported in the inflamed mucosa of patients with Crohn’s disease; even GSH levels were slightly decreased in noninflamed mucosa [295]. A deficit in GSH synthesis mediated by diminished activity of glutamate cysteine ligase would account for GSH depletion, whereas the overproduction of H2O2 associated with chronic inflammation would explain glutathione oxidation [290,295,296]. GSH depletion contributes to mucosal injury as administration of Nacetyl cysteine or GSH in experimental models of colitis in rats and mice diminished damage of the intestinal mucosa [297–299]. High HNE levels and increased MDA levels have been found in intestinal mucosa from patients with Crohn’s disease [300– 302]. DNA is also a target of oxidative and nitrosative stress in inflammatory bowel disease and DNA damage may certainly contribute to inflammation-associated carcinogenesis. 8-OHdG levels, a marker of oxidative DNA damage, increases in the mucosa of rodents with colitis and in patients with inflammatory bowel disease, and it may promote colon carcinogenesis [303–306]. 8-nitroguanine levels, and indicator of nitrative DNA damage, also increased in a model of inflammatory bowel disease in mice [305]. A harmful “oxy-radical overload” severely affects the intestinal mucosa and correlates with the clinical outcome in inflammatory bowel disease [290,307–309]. Nowadays, oxidative stress is considered a potential etiological factor and also a critical factor in the progression and severity of the disease rather than a concomitant consequence of chronic intestinal inflammation [310–312]. Thus, H2O2 may oxidize actin and tubulin causing disassembly of F-actin and tubulin, which leads to disruption of cytoskeleton and intestinal barrier dysfunction [313,314]. Accordingly, carbonylation of actin and

- 47 -

tubulin as well as nitration of actin increase in the inflamed mucosa of patients with active ulcerative colitis leading to disruption of the actin cytoskeleton in epithelial cells [315]. Mitochondrial O2-∙ also contributes to epithelial barrier dysfunction in experimental colitis [316]. Macrophages and neutrophils are considered the major source of ROS and RNS during intestinal inflammation and play a key role not only in the initiation but also in the maintenance of inflammation in colitis [311,317]. In experimental models of colitis, selective depletion of neutrophils or macrophages diminished ROS/RNS generation and decreased intestinal inflammation and injury as well as pro-inflammatory cytokine expression [311,318]. In fact, high levels of 3-chlorotyrosine and the DNA chlorination product 5-chloro-2’-deoxycytidine have been found in colon samples from patients with inflammatory bowel disease [317]. 2.6.2.- Nitrosative stress in inflammatory bowel disease Protein nitration markedly increases in the inflamed mucosa of patients with ulcerative colitis, exhibiting higher levels than in patients with Crohn’s disease [302]. Interestingly, protein nitration was associated with neutrophilic MPO but not with iNOS [302], suggesting its formation through a peroxynitrite-independent pathway that is likely to be via MPO-derived NO2∙ [311]. Importantly, administration of peroxynitrite in rectum triggered colitis in rats [319]. Nevertheless, nitrosative stress is less intense in humans in comparison with rodent colitis [317], and NO seems to play a dual role in inflammatory bowel disease, being beneficial when present in intestinal endothelial cells but detrimental when derived from leukocytes [290]. NO derived from endothelial nitric oxide synthase (eNOS) protects the microvasculature though relaxation of smooth muscle cells and prevention of thrombosis and leukocyte adhesion [320–322]. Accordingly, disease severity was dramatically increased in eNos deficient mice, which

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exhibited markedly increased inflammatory infiltrate and intestinal injury [323]. The increase in iNOS expression and peroxynitrite levels found in chronically inflamed intestinal mucosa should be ascribed to leukocyte infiltration, but iNOS expression decreased in endothelial cells of the intestinal mucosa of patients with Crohn’s disease or ulcerative colitis [320,324]. In contrast to the findings in eNos knock-out mice, the inflammatory infiltrate in experimental colitis was much less intense in iNos knock-out mice than in wild type [325]. Other studies have also reported protective effects of iNOS deficiency or inhibition against intestinal inflammation and damage in experimental colitis in mice, at least in part via TLR4/ NF-κB signaling [326–328]. 2.6.3.- Redox signaling in the pathogenesis and progression of inflammatory bowel disease Although the etiology of inflammatory bowel disease still remains unknown, ROS have been involved in the potential pathogenesis of this disease through three different proposed stimuli and mechanisms, which are the increase in serotonine levels, inflammasome activation induced by titanium dioxide (TiO2) nanoparticles, and certain NOX2 polymorphisms (Figure 4, Tables 2-3). These mechanisms might cause the loss of a delicate balance of controlled ROS production that should be maintained within a relatively narrow range. Serotonin (5-hydroxytryptamine), which is mostly synthesized and localized in the gut, has been linked with the pathogenesis of inflammatory bowel disease, because its synthesis is increased in these patients [329] and it markedly enhances colon inflammation through NOX2-derived O2-∙ in an experimental model of colitis in rats [330]. It has been recently reported that titanium dioxide nanoparticles, which are commonly used as food additives (E171) and in pharmaceutical formulations, aggravate

- 49 -

experimental colitis via enhanced generation of ROS and activation of nucleotidebinding oligomerisation domain receptor, pyrin domain containing (NLRP) 3 inflammasome [331]. It is worth noting that titanium levels are increased in patients with inflammatory bowel disease [331]. However, NOX2 polymorphisms that exhibit diminished O2-∙ production have been found in patients with very early onset inflammatory bowel disease, which is a variant with disease onset before 6 years of age [332]. In addition, a polymorphism of NCF4 (p40phox), a member of the NOX family, confers risk for development of inflammatory bowel disease [333]. Importantly, HNE causes apoptosis in colon cells via JNK, reacts with DNA producing etheno-modified DNA bases, which increase in colon of patients with Crohn’s disease or ulcerative colitis [301,334], and also reacts with IgA reducing its intestinal bactericidal activity in rats [335]. Additionally cholesterol oxidation products named oxysterols, particularly 7-ketocholesterol and 7β-hydroxycholesterol, cause apoptosis driven by NOX1-derived O2-∙ in differentiated CaCo-2 cells and also induce the expression of cytokines [IL-1β, IL-6, IL-8, IL-23, and monocyte chemotactic protein-1 (MCP-1)] as well as receptors TLR2 and TLR9, at least in part through the activation of NOX1 [336,337]. Hence, oxysterols may contribute to the impairment of the intestinal epithelial barrier found in inflammatory bowel disease [336]. MnSOD protein levels and activity strongly increase in inflamed mucosa of patients with Crohn’s disease and ulcerative colitis, and other antioxidant enzymes such as Cu/ZnSOD, glutathione peroxidase (particularly intestinal GPx2), and catalase also increase [338–340]. The up-regulation of Gpx2 in mice during colitis is mediated by signal transducers and activators of transcription (STAT) transcription factors, but not by Nrf2 [341]. Cu/ZnSod overexpression in mice increased survival in an experimental

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model of severe acute colitis, although it did not protect against chronic intestinal inflammation [342]. Mice with double deficiency in glutathione peroxidases Gpx1 and Gpx2 develop spontaneous ileocolitis [343,344], which is mainly due to Nox1 up-regulation and its associated O2-∙ and H2O2 generation [345]. Indeed, NOX1 triggers marked TNF-upregulation, apoptosis and ileocolitis in mice [345]. It is worth noting that GPx2 seems to exert a potent anti-inflammatory function in the intestine by restraining COX-2 expression, as GPX2 silencing in HT-29 cells triggers up-regulation of COX-2 and membrane-associated PGE synthase-1 (mPGES-1) and induces the release of prostaglandin PGE2 [346]. Thioredoxin activity increases in intestinal epithelial cells exposed to low concentration of H2O2 [347]. Furthermore, the mRNA expression of thioredoxin interacting protein, a thioredoxin negative regulator, is decreased in the inflamed mucosa of patients with ulcerative colitis [348]. These findings would support an increase of thioredoxin activity in the inflamed mucosa of patients with inflammatory bowel disease [290]. NF-κB is a downstream transcription factor for microbial-activated TLRs and TNF-α in intestinal epithelial cells, in the latter case through activation of NADPH oxidases [290,349]. Redox-sensitive NF-κB is chronically over-activated in the intestinal mucosa of patients with Crohn´s disease and ulcerative colitis [293,350,351], and thus it also seems to contribute to chronic inflammation and injury in the gut of these patients. Indeed, treatment with the NF-κB essential modulator (NEMO)-binding peptide, which avoids the interaction between NEMO and inhibitory kappa B kinase (IKKβ), abrogated IKK activation and markedly diminished intestinal inflammation and injury in experimental models of colitis [352]. Furthermore, NF-κB contributes to the development of ulcerative colitis by inducing apoptosis of intestinal epithelial cells

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through up-regulation

of

p53-upregulated modulator

of apoptosis

(PUMA),

paradoxically in a p53 independent manner [353]. TNF-α-triggered loss of intestinal barrier is also mediated by NF-κB via induction of metalloproteinases and myosin light chain kinase [354,355]. Importantly, TNF-α is a very powerful activator of NOX1derived O2-∙ generation in colonic cells through induction of NOX1 and NOXO1 [356]. Nrf2 activation restrains the inflammatory response and mucosal injury in inflammatory bowel disease. Indeed, in experimental models of colitis Nrf2 knock-out mice exhibited increased loss of crypts and severe inflammatory infiltrate as well as enhanced upregulation of pro-inflammatory Il-1β, Il-6, Il-8, Tnf-α, iNos and Cox-2, and downregulation of target genes (Ho-1, NADPH quinone oxidoreductase-1 (Nqo1), and Gst Mu-1) [290,357–359]. Nrf2 activation in colonic cells is negatively regulated by the stress-inducible gene immediate early response-3 (IER3) through the PI3K/Akt-Fyn pathway [360]. Thus, Ier3 deficiency triggers activation of both Akt and Nrf2 leading to up-regulation of its target genes, diminished ROS levels and reduced apoptosis in experimental colitis [360]. Hence Nrf2 activity might be kept low in inflammatory bowel disease due to the high level expression of IER3 in the inflamed mucosa [360]. The gut microbiota should be taken also into account because it may contribute to inflammatory bowel disease or conversely restrain the intestinal chronic inflammation by modulating ROS production and antioxidant defense (see Table 2). Indeed, Enteroccocus fecalis generates high levels of H2O2 aggravating intestinal damage in inflammatory bowel disease [361,362], whereas Lactobacillus up-regulates antioxidant enzymes improving the evolution of the disease [363,364]. In addition, the pathogenic adherent-invasive strain of Escherichia coli LF82, obtained from the ileum of patients with Crohn’s disease, enhanced ROS generation in vitro in epithelial cells via NOX1

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and NOXO1 up-regulation leading to reduced mucin expression and increased IL-8 expression [365]. 2.6.4.- Redox signaling in the therapy of inflammatory bowel disease The host-microbiota interaction represents a great therapeutic tool because it may attenuate chronic intestinal inflammation by triggering the release of bacterial antioxidant enzymes on the inflamed mucosa. Thus, a strain of Lactococcus lactis that encodes cytoplasmic superoxide dismutase A (SodA) diminished oxidative stress in the colon and reduced colitis in different experimental models by releasing the enzyme at inflamed sites after host lysozyme-mediated lysis [366]. It is worth noting that the beneficial effects of this lactococcal strain were completely dependent on SodA [366]. On the other hand, protein intake may modulate intestinal inflammation through autophagy, ROS and inflammasome activation. Thus, general controlled non-repressed 2 (GCN2) kinase, a sensor of protein starvation that modulates proteins synthesis and the stress response, markedly restrains intestinal inflammation by stimulating autophagy that leads to low ROS-dependent inflammasome activation and Th17 cell responses [367]. Indeed, Gcn2 deficient mice exhibited enhanced IL-1β production and Th17 cell responses due to reduced autophagy, which led to increased mitochondrial O2-∙ levels in colonic cells and subsequently to enhaced inflammasome activation [367]. The loss of the delicate balance between NOX1-derived O2-∙ generation, Mn and Cu/ZnSOD, GPx1/2, and Nrf2/IER3 contributes to the pathogenesis and progression of chronic inflammation in the gut during Crohn’s disease and ulcerative colitis (see table 3). The up-regulation of MnSOD, Cu/ZnSOD, GPx2, and catalase that occurs in intestinal mucosa during inflammatory bowel disease cannot cope with the O2-∙ and H2O2 generated by TNF-α-induced NOX1 together with NOX2 from activated leukocytes. It remains to be investigated the mechanism for the reduction in mucin

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secretion induced by O2-∙ and H2O2 as well as the role of Nrf2 and AP-1 pathways in Crohn’s disease and ulcerative colitis, particularly the Nrf2 regulation by IER3. It would be also worth elucidating the potential role of ROS generated from pathogenic microbiota specifically in the rectum concerning the initiation of ulcerative colitis. Additionally, future research should focus on therapeutic approaches using the microbiota or stimulation of autophagy to restrain the inflammatory response by decreasing O2-∙, H2O2 and oxidative stress in the intestinal mucosa.

2.7.- Colorectal cancer Colorectal cancer is the second most common cancer in women and the third in men, with 1.36 million cases and almost 700.000 deaths in 2012 [368]. Its highest rates have been reported in Australia/New Zealand, Oceania, Europe, and Northern America, whereas the lowest rates have been found in Western Africa [368]. The incidence of colorectal cancer increases with age and it is estimated that half of the Western population develops adenomatous polyps when reaching 70 years of age, which may progress to cancer [369]. Indeed, the classical tumor progression from adenoma to carcinoma [370] is still the basic model to explain most cases of the disease, although the required multiple gene mutations affect the physiological regulation of the epithelium instead of involving simultaneous mutations of generic oncogenes and tumor-suppressor genes [369]. In any case, colon tumors contain tumor cells at different differentiation status, being undifferentiated cancer stem cells or tumor-initiating cells those with clonogenic and tumor initiating potential [371,372]. The three major forms of colorectal cancer are colitis associated, hereditary, and sporadic colorectal cancer [373]. Importantly, patients with inflammatory bowel

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disease, particularly with pan-colitis, are at risk of developing colorectal cancer [285,374,375]. Indeed, chronic inflammation contributes to the initiation and progression of colorectal carcinoma [309,311]. Tumor progression from multifocal dysplasia to carcinoma is typical in these patients [309]. Thus, more than 1/5 of patients with inflammatory bowel disease suffer colorectal carcinoma during the first 30 years of the disease, and more than half of them die due to this cancer [376]. The duration of colitis and the extent of colitis are risk factors for colorectal cancer [309]. Both tumor associated macrophages, which exhibit the protumoral M2 phenotype, and tumor associated neutrophils with protumoral N2 phenotype have been involved in colorectal carcinogenesis [377]. M2 macrophages promote cell proliferation and angiogenesis,

restrain

the

adaptive

immune

response,

and

release

matrix

metaloproteinases that facilitate tumor development and invasiveness [377,378]. Increased expression of COX-2, TNF-α, IL-6, IL-8, CXCR2 as well as IL-17 and IL-23 all contribute to progression of colorectal cancer [309,373,378–380]. In addition, persistent activation of NF-κB and STAT3 occurs in colorectal carcinoma [381,382]. Furthermore, colorectal cancer may be induced in the Apc knock-out mice by commensal Bacteroides enterotoxigenic fragilis through the Th17/STAT3 pathway (see Table 2) [383]. It is worth noting that in addition to chronic inflammation, microbiotaderived genotoxins seem to be required for initiation and progression of colitisassociated colorectal cancer [311,384]. Indeed, mice deficient in IL-10 suffering from colitis developed colon tumors upon colonization by a commensal E. coli strain that expresses genotoxin colibactin, which is carcinogenic [384]. A deficiency in plasma glutathione peroxidase GPx3 may also contribute to colitisassociated cancer. Thus, GPx3 is often downregulated in colon cancer due to

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hypermethylation, and GPx3-deficient mice exhibited enhanced colitis-associated tumorigenesis [85]. Indeed, GPx3 deficiency led to increased number of tumors, oxidative DNA damage, and overactivation of the Wnt pathway in inflammatory carcinogenesis induced in mice by azoxymethane/dextran sodium sulfate [85]. Hence, GPx3 is considered tumor suppressor in colitis-associated cancer. The molecular mechanisms responsible for colorectal cancer have been mostly identified in studies of hereditary colorectal cancer, mainly familial adenomatous polyposis, hereditary non-polyposis colorectal cancer, and juvenile polyposis syndrome [369]. Familial adenomatous polyposis is mainly caused by a germ-line inactivating mutation of the tumor suppressor gene APC [385–387] and these patients exhibit colorectal cancer when reaching 40 years of age [369]. Mutyh-associated polyposis is another form of hereditary and familial polyposis characterized by multiple colorectal adenomas and cancer, which is due to biallelic germline mutations of MUTYH that causes deficiency in base-excision repair [388,389]. A somatic inactivating mutation of APC is also present in most cases (85%) of sporadic colorectal carcinoma [387]. Hence, activating mutations of Wnt signaling are the critical first step in most cases of colorectal cancer [369,390]. The interplay between Wnt signaling and Notch signaling should be taken also into account, because γ-secretase inhibitors that modulate the expression of Hes1 abrogate the proliferation of adenomas transforming proliferative cells into differentiated globet cells [391]. Hereditary non-polyposis colorectal cancer account for 2-7% of all cases of colorectal cancer and is characterized by instability of DNA microsatellites [392], which leads to inherited mutations in genes for DNA mismatch repair such as MLH1, MSH2 and MSH6 [369,373,393,394]. On the other hand, a majority of hamartomatous polyposis syndromes, such as juvenile polyposis syndrome, is caused by inactivation of the BMP - 56 -

pathway, particularly mutations in SMAD4 [395] or BMP receptor type 1A [396]. Accordingly, conditional inactivation of the BMP pathway in the intestine in mice leads to the generation of hamartomatous polyps [397]. Furthermore, even impairment of BMP signaling in non-epithelial cells, such as SMAD4 mutation in T lymphocytes, may lead to intestinal polyposis [398]. Inactivation of TGF-β signaling promotes tumorigenesis, and accordingly SMAD2, SMAD3, SMAD4 and transforming growth factor-beta receptor 2 (TGFBR2) somatic mutations have been reported in patients with sporadic colorectal cancer [399]. The molecular pathogenesis of colitis-associated colon cancer and sporadic colon cancer involve mutations in APC, p53, deleted in colorectal carcinoma/deleted in pancreatic cancer locus 4 (DCC/DPC4), and K-RAS as well as aneuploidy, microsatellite instability, and DNA methylation [309]. Importantly, mutations must occur in crypt base stem cells, but not in differentiated epithelial cells or transit-amplifying cells, to origin intestinal cancer [400]. 2.7.1.- Oxidative stress in colorectal cancer Oxidative stress has been associated with human colon cancer as MDA, DNA hydroxylation, protein carbonyls, and cysteine oxidation increase in colorectal tumors [401–406]. Interestingly, colorectal adenocarcinoma, but not adenoma, exhibits enhanced cell proliferation together with persistent oxidative stress, evidenced by increased levels of 8-hydroxy-2’-deoxyguanosine, 4-hydroxy-2-nonenal-histidine adducts, and 3-nitro-L-tyrosine [407]. These findings suggest that colorectal adenocarcinoma cells can cope with maintained oxidative stress, which might stimulates cell proliferation without inducing apoptosis [407]. Hence, reactive oxygen and nitrogen species, mainly released by activated leukocytes during chronic inflammation, may act

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not only as tumor promoters in colorectal cancer through enhancing cell proliferation, but also as tumor initiators by inducing DNA mutations (Figure 5) [309,378,408]. The high levels of DNA oxidation present in colorectal tumors [409,410], together with the inactivation of the DNA mismatch repair system by oxidative stress [411], may lead to genomic mutations and microsatellite instability that initiate and promote colitisassociated carcinogenesis (Figure 5) [311]. On the other hand, deficiency in selenoproteins is also associated with increased susceptibility to oxidative stress and may contribute to colitis-induced tumorigenesis [412]. The major selenoprotein P (SEPP1) transports selenium from the liver to other tissues and exhibits antioxidant properties due to its ten selenocysteines, and its deficiency or the loss of its redox-active site promotes colonical epithelial stem cell features via stimulation of Wnt/β-catenin signaling, triggers genetic instability, and enhances

oxidative

stress-induced

DNA

damage

and

inflammatory

colonic

tumorigenesis [412]. Importantly, specimens of colorectal cancer exhibit low SEPP1 expression, and certain polymorphisms of the gene increased the risk of adenoma [413,414]. 2.7.2.- The Janus faces of redox signaling in colorectal cancer Simultaneous deficiency in both glutathione peroxidases 1 and 2 in mice triggers ileocolitis and also promotes mutation accumulation and intestinal cancer (Figure 5) [343,408,415]. Nevertheless, the genetic background remarkably affects the severity of ileocolitis and the incidence of ileocolonic tumors [408]. Indeed, B6.129 Gpx1/2-double knock-out mice were prone to severe ileocolitis and 25% of them developed intestinal tumors, whereas B6 mice exhibited mild ileocolitis and less inflammatory infiltrate as well as low incidence of intestinal tumors (2.5%) [408]. Restoration of GPx2, but not

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GPx1, avoids ileocolitis and intestinal cancer [416]. Accordingly, Gpx2 knock-out mice exhibited severe inflammation and more number of tumors in a model of inflammationassociated colon cancer [417]. Importantly, GPx2 may lower inflammation by downregulating COX-2 expression in human colon cancer cells [418]. The loss of tumor suppressor APC triggers generation of O2-∙ and H2O2, which is required for proliferation of intestinal stem cells and tumor initiation (Figure 5) [76]. Apc deficient mice exhibited markedly less number of intestinal adenomas when made deficient also in iNos [419]. Increased expression of NOX1, NOXO1 and NOXA1 was found in malignant colorectal tissue [420,421], which could promote colorectal tumorigenesis and vascularization through H2O2-driven up-regulation of the Wnt and Notch cascades [29] and induction of angiogenic factors (Figure 5) [422]. Interestingly, NOX1 activity is maintained low under basal conditions in colon cancer cells by ubiquitination and proteasomal degradation of NOXO1 mediated by growth receptor bound protein 1 (Grb2) and casitas B-lineage lymphoma (Cbl) E3 ligase, but upon stimulation by EGF serine 154 phosphorylation of NOXO1 blocks its degradation increasing NOX1 activity [421]. However, redox signaling exhibits two Janus faces in colon cancer because O2-∙ and H2O2 may also paradoxically protect against tumor progression and dissemination (Figure 5). Thus, ROS are required for apoptosis of colon cancer cells. p53-dependent apoptosis is mediated, at least in part, by the induction of mitochondrial proline oxidase and subsequently proline-dependent ROS generation in human colon cancer cells [423]. Proline oxidase converts proline into pyrroline-5-carboxylate leading to electron transfer to cytochrome c [423]. PUMA can also mediate apoptosis induced by p53 or DNA-damaging agents, as it induces Bax- and mitochondrial-dependent apoptosis in

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colorectal cancer cells through the generation of O2-∙ and H2O2 that causes proteasomemediated degradation of stathmin and disruption of cellular microtubules [424]. In addition, mitochondrial O2-∙ generated by oxidative phosphorylation uncoupling markedly enhance tumor necrosis factor-related apoptosis-inducing ligand (TRAIL)induced release of Smac/DIABLO from mitochondria and apoptosis in TRAIL-resistant human colon carcinoma cells [425]. HNE, one of the major end products of lipid peroxidation, induces expression and synthesis of TGF-β1 [426], which triggers growth inhibition. Strikingly, HNE levels as well as TGF-β1 protein were decreased in specimens of colon adenocarcinoma [300]. Furthermore, TGF-β1 receptors 1 and 2 were also generally down-regulated in colon cancer [300,426]. Hence, the low lipid peroxidation associated with colon cancer may favor colon tumor cells to escape from TGF-β1-mediated apoptosis and growth inhibition thus promoting tumor progression [300]. ROS also mediate the inhibition of AP-1 activity and the decrease in COX-2 and vascular endothelial growth factor (VEGF) expression exerted by the anti-inflammatory prostanoid 15-deoxy-∆12,14-prostaglandin J2 (15d-PGJ2) in colon carcinoma cells [427]. Importantly,

COX-2

gene

expression

is

up-regulated

in

human

colorectal

adenocarcinoma [428] and COX-2 together with VEGF contribute to tumor growth, metastasis, and angiogenesis [429,430]. Colon cancer cells exhibit high Nrf2 activity, which confers resistance to death ligands and anticancer treatment [431]. Nrf2 up-regulation may be due to the loss of function of its negative regulator IER3 that occurs in colorectal cancer [360]. Nrf2 seems to promote colon cancer at least in part by enhancing tumor angiogenesis as Nrf2 knockout reduced tumor growth, VEGF expression and vascular formation in tumor

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xenografts developed from human colon cancer cells in mice [432]. Nrf2 silencing inhibited HIF-1α-VEGF signaling in human colon cancer cells [432]. Accordingly, levels of MnSOD levels -a target of Nrf2- are elevated in primary colon cancer and further increase in metastasis, suggesting that MnSOD might promote tumor development and dissemination [433]. In addition, HO-1 is up-regulated in colorectal carcinoma and it stimulates angiogenesis contributing to tumor vascularization and progression and also confers resistance against anticancer treatment [434]. Interestingly in the short term of experimental colitis, Nrf2-deficient mice exhibited much more preinvasive aberrant crypt foci in the colon, and even in the long term most of them developed tumors (92% vs 53% in wild type mice), but paradoxically in Nrf2 knock-out mice the incidence of adenocarcinoma was much higher than that of adenoma (80% vs 20%), whereas wild type mice exhibited 80% adenomas and only 20% adenocarcinomas [357,358]. GPx2 protects against colon tumor initiation by restraining inflammation, but paradoxically stimulates tumor growth and metastasis [435]. Indeed, colon tumors with high GPx2 expression exhibit high Wnt signaling and proliferation, and GPx2 is required for the formation of metastasis by the aggressive colon cancer subtype 3 tumor cells [435]. The loss of hepatocyte nuclear factor-4α (HNF-4α) stimulates the Wnt/β-catenin pathway in crypt cells that might promote tumorigenesis, but unexpectedly HNF-4α deficiency protects against colon cancer by down-regulating expression of oxidoreductase genes, such as CYPB6, CYPD6, GSTA4, GSTK1 and NQO1, leading to increased ROS, lipid peroxidation and apoptosis [436]. Accordingly, increased levels of

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HNF-4α together with up-regulation of its target ROS-neutralizing genes were found in human samples of colorectal cancer [436]. The enzyme TIGAR lowers oxidative stress and supports glutathione-dependent antioxidant defense, but it contributes to intestinal adenoma formation and tumor development in mice and is increased in human colon cancer [74]. TIGAR functions as a fructose 6-biphosphatase that lowers the glycolytic flux and instead enhances the production of NADPH available for antioxidant defense and nucleotide synthesis[74]. Finally, the administration of antioxidants is not recommended as a preventive therapy against colorectal cancer since a meta-analysis of intervention studies stated that oral administration of antioxidants did not prevent colorectal neoplasia in the general population [437]. The two Janus faces of redox signaling in colorectal cancer creates a controversy on the precise role of ROS in the pathogenesis and progression of this neoplasia. On the one hand, persistent oxidative stress is associated with colorectal cancer cells and the upregulation of NOX1 in malignant colorectal tissue may stimulate cell proliferation through activation of Wnt and Notch signaling and would lead to Nrf2 activation, which promotes anti-apoptotic mechanisms and angiogenesis. However, on the other hand ROS are required for p53- and PUMA-induced apoptosis and also mediate COX-2 and VEGF down-regulation triggered by prostanoid 15d-PGJ2. Hence, further research is needed to clarify which mechanism is predominant and critical specifically in crypt base stem cells, those cells that harbor the mutation eventually responsible for the pathogenesis of colorectal cancer. Additionally, those redox-sensitive pathways that promote or restrain tumor progression in vivo should be clearly identified and further elucidated. In any case, it should be taken into account that the role of redox signaling

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may substantially change among the three major forms of colorectal cancer –colitis associated, hereditary, and sporadic-.

3.- CONCLUDING REMARKS The present broad and extensive overview highlights the great impact of redox signaling, oxidative stress, and nitrosative stress in the physiology and pathophysiology of inflammation and cancer of the gastrointestinal tract. A physiological nitrosative stress is common in the lumen of the stomach as high levels of nitric oxide are formed at the acidic gastric pH from salivary nitrite independently of nitric oxide synthase. Under physiological conditions, the normal and dynamic homeostasis of the gastrointestinal epithelium requires an appropriate thiol redox status and redox signaling through NOX1 and DUOX-2. In fact, intracellular glutathione redox status and extracellular cysteine redox status modulate the proliferative potential of intestinal epithelial cells. Furthermore, physiological self-renewal, proliferation, migration and differentiation of the intestinal epithelium by Wnt/β-catenin and Notch signaling pathways occur through a delicate balance between RAC1-NOX1-derived ROS and TIGER. Commensal bacteria, particularly Lactobacilli, decisively contribute to intestinal epithelial homeostasis through generation of O2-∙ and H2O2 by both NOX1and DUOX-2. Strikingly, the loss of redox homeostasis leads to the pathogenesis and progression of a wide diversity of gastrointestinal disorders, such as Barrett’s esophagus, esophageal adenocarcinoma, peptic ulcer, gastric cancer, ischemic intestinal injury, celiac disease, inflammatory bowel disease, and colorectal cancer. The overproduction of O2-∙ anion together with inactivation of superoxide dismutase are involved in the pathogenesis of - 63 -

Barrett’s esophagus and its transformation to adenocarcinoma. In Helicobacter pyloriinduced peptic ulcer, oxidative stress derived from the leukocyte infiltrate and NOX1 aggravates mucosal damage, especially in HspB+ strains that down-regulate Nrf2. The oncogenesis of gastric carcinoma is associated with up-regulation of NOX-1 and spermine oxidase. In celiac disease, oxidative stress mediates most of the cytotoxic effects induced by gluten peptides and increases transglutaminase levels, whereas nitrosative stress contributes to the impairment of tight junctions. Progression of inflammatory bowel disease relies on the balance between pro-inflammatory redoxsensitive pathways, such as NLRP3 inflammasome and NF-κB, and the adaptive upregulation of Mn superoxide dismutase and glutathione peroxidase 2. In colorectal cancer, redox signaling exhibits two Janus faces: On the one hand, NOX1 up-regulation and derived H2O2 enhance Wnt/β-catenin and Notch proliferating pathways; on the other hand, O2-∙ and H2O2 may disrupt tumor progression through different pro-apoptotic mechanisms. We propose a ROS-dependent pathogenic triad that would explain the critical interplay between O2-∙ and H2O2 as initiator factors, chronic inflammation and oncogenesis in the gastrointestinal tract (see Figure 6). Chronic inflammation may trigger overproduction of O2-∙ radicals and/or H2O2 through different mechanisms that overwhelm the antioxidant defense due to loss of superoxide dismutase activity or glutathione peroxidases. The increased generation of O2-∙ and H2O2 would be triggered by NOX-1 or DUOX-2 up-regulation that causes oxidative inactivation of superoxide dismutases and renders inefficient the glutathione-dependent antioxidant system. The enhanced production of H2O2 may be mediated by up-regulation of spermine oxidase and/or deficiency in glutathione peroxidases 1, 2, 3 or 7. In order to promote oncogenesis, the

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overwhelming ROS would trigger simultaneously two parallel mechanisms within the same cell, firstly one anti-apoptotic mechanism -such as APE1 up-regulation and acetylation, NRF-2 activation, or EGFR activation-, and secondly another promoter factor that causes uncontrolled cell proliferation related to Wnt, Notch, p53 or K-ras, which would be altered by mutation. The triad would close with the stimulation of Wnt, Notch-, or K-ras-dependent cell proliferation promoted by the anti-apoptotic mechanisms. In conclusion, overproduction of O2-∙ and H2O2, and redox signaling play a critical role in the physiology and pathophysiology of the gastrointestinal tract, particularly in inflammation and its progression towards cancer.

ACKNOWLEDGMENTS The authors acknowledge the support by Grant SAF2015-71208-R with FEDER funds from the Spanish Ministry of Economy and Competitiveness to J.S. I.F. was recipient of a fellowship from "Programa de Pós-Doutorado no Exterior (PDE)” that belongs to the “Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq)”.

REFERENCES

- 65 -

[1]

L. Laine, K. Takeuchi, A. Tarnawski, Gastric mucosal defense and cytoprotection: bench to bedside, Gastroenterology. 135 (2008) 41–60. doi:10.1053/j.gastro.2008.05.030.

[2]

A. Bhattacharyya, R. Chattopadhyay, S. Mitra, S.E. Crowe, Oxidative stress: an essential factor in the pathogenesis of gastrointestinal mucosal diseases, Physiol.Rev. 94 (2014) 329–354. doi:10.1152/physrev.00040.2012.

[3]

Y. Naito, T. Yoshikawa, Molecular and cellular mechanisms involved in Helicobacter pylori-induced inflammation and oxidative stress, Free Radic. Biol. Med. 33 (2002) 323–336.

[4]

S.

Moncada,

R.M.

Palmer,

E.A.

Higgs,

Nitric

oxide:

physiology,

pathophysiology, and pharmacology, Pharmacol. Rev. 43 (1991) 109–142. [5]

J.O. Lundberg, M. Govoni, Inorganic nitrate is a possible source for systemic generation of nitric oxide, Free Radic. Biol. Med. 37 (2004) 395–400. doi:10.1016/j.freeradbiomed.2004.04.027.

[6]

J.O. Lundberg, E. Weitzberg, M.T. Gladwin, The nitrate-nitrite-nitric oxide pathway in physiology and therapeutics, Nat Rev Drug Discov. 7 (2008) 156– 167. doi:10.1038/nrd2466.

[7]

C. Duncan, H. Dougall, P. Johnston, S. Green, R. Brogan, C. Leifert, L. Smith, M. Golden, N. Benjamin, Chemical generation of nitric oxide in the mouth from the enterosalivary circulation of dietary nitrate, Nat. Med. 1 (1995) 546–551.

[8]

H. Björne H, J. Petersson, M. Phillipson, E. Weitzberg, L. Holm, J.O. Lundberg, Nitrite in saliva increases gastric mucosal blood flow and mucus thickness, J. Clin. Invest. 113 (2004) 106–114. doi:10.1172/JCI19019.

- 66 -

[9]

N. Benjamin, F. O’Driscoll, H. Dougall, C. Duncan, L. Smith, M. Golden, H. McKenzie,

Stomach

NO

synthesis,

Nature.

368

(1994)

502.

doi:10.1038/368502a0. [10] J.O. Lundberg, E. Weitzberg, J.M. Lundberg, K. Alving, Intragastric nitric oxide production in humans: measurements in expelled air, Gut. 35 (1994) 1543–1546. [11] B.S. Rocha, B. Gago, R.M. Barbosa, J.O. Lundberg, R. Radi, J. Laranjinha, Intragastric nitration by dietary nitrite: implications for modulation of protein and lipid

signaling,

Free

Radic.

Biol.

Med.

52

(2012)

693–698.

doi:10.1016/j.freeradbiomed.2011.11.011. [12] C. Pereira, N.R. Ferreira, B.S. Rocha, R.M. Barbosa, J. Laranjinha, The redox interplay between nitrite and nitric oxide: From the gut to the brain, Redox Biol. 1 (2013) 276–284. doi:10.1016/j.redox.2013.04.004. [13] T. Sobko, C. Reinders, E. Norin, T. Midtvedt, L.E. Gustafsson, J.O. Lundberg, Gastrointestinal nitric oxide generation in germ-free and conventional rats, Am. J. Physiol.

Gastrointest.

Liver

Physiol.

287

(2004)

G993-997.

doi:10.1152/ajpgi.00203.2004. [14] L. Peri, D. Pietraforte, G. Scorza, A. Napolitano, V. Fogliano, M. Minetti, Apples increase nitric oxide production by human saliva at the acidic pH of the stomach: a new biological function for polyphenols with a catechol group?, Free Radic. Biol. Med. 39 (2005) 668–681. doi:10.1016/j.freeradbiomed.2005.04.021. [15] B. Gago, J.O. Lundberg, R.M. Barbosa, J. Laranjinha, Red wine-dependent reduction of nitrite to nitric oxide in the stomach, Free Radic. Biol. Med. 43 (2007) 1233–1242. doi:10.1016/j.freeradbiomed.2007.06.007. [16] B.S. Rocha, B. Gago, R.M. Barbosa, J.O. Lundberg, G.E. Mann, R. Radi, J. Laranjinha, Pepsin is nitrated in the rat stomach, acquiring antiulcerogenic

- 67 -

activity: a novel interaction between dietary nitrate and gut proteins, Free Radic. Biol. Med. 58 (2013) 26–34. doi:10.1016/j.freeradbiomed.2012.12.017. [17] B.S. Rocha, J.O. Lundberg, R. Radi, J. Laranjinha, Role of nitrite, urate and pepsin in the gastroprotective effects of saliva, Redox Biol. 8 (2016) 407–414. doi:10.1016/j.redox.2016.04.002. [18] E.A. Jansson, J. Petersson, C. Reinders, T. Sobko, H. Björne, M. Phillipson, E. Weitzberg, L. Holm, J.O. Lundberg, Protection from nonsteroidal antiinflammatory drug (NSAID)-induced gastric ulcers by dietary nitrate, Free Radic. Biol. Med. 42 (2007) 510–518. doi:10.1016/j.freeradbiomed.2006.11.018. [19] R.S. Dykhuizen, A. Fraser, H. McKenzie, M. Golden, C. Leifert, N. Benjamin, Helicobacter pylori is killed by nitrite under acidic conditions, Gut. 42 (1998) 334–337. [20] A.S. Neish, Redox signaling mediated by the gut microbiota, Free Radic. Res. 47 (2013) 950–957. doi:10.3109/10715762.2013.833331. [21] F. Gerbe, J.H. van Es, L. Makrini, B. Brulin, G. Mellitzer, S. Robine, B. Romagnolo, N.F. Shroyer, J.-F. Bourgaux, C. Pignodel, H. Clevers, P. Jay, Distinct ATOH1 and Neurog3 requirements define tuft cells as a new secretory cell type in the intestinal epithelium, J. Cell Biol. 192 (2011) 767–780. doi:10.1083/jcb.201010127. [22] L.G. van der Flier, H. Clevers, Stem cells, self-renewal, and differentiation in the intestinal

epithelium,

Annu.

Rev.

Physiol.

71

(2009)

241–260.

doi:10.1146/annurev.physiol.010908.163145. [23] T. Sato, J.H. van Es, H.J. Snippert, D.E. Stange, R.G. Vries, M. van den Born, N. Barker, N.F. Shroyer, M. van de Wetering, H. Clevers, Paneth cells constitute the

- 68 -

niche for Lgr5 stem cells in intestinal crypts, Nature. 469 (2011) 415–418. doi:10.1038/nature09637. [24] M. Bjerknes, H. Cheng, The stem-cell zone of the small intestinal epithelium. II. Evidence from paneth cells in the newborn mouse, Am. J. Anat. 160 (1981) 65– 75. doi:10.1002/aja.1001600106. [25] Y. Yatabe, S. Tavaré, D. Shibata, Investigating stem cells in human colon by using methylation patterns, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 10839– 10844. doi:10.1073/pnas.191225998. [26] D.H. Scoville, T. Sato, X.C. He, L. Li, Current view: intestinal stem cells and signaling,

Gastroenterology.

134

(2008)

849–864.

doi:10.1053/j.gastro.2008.01.079. [27] C. Booth, C.S. Potten, Gut instincts: thoughts on intestinal epithelial stem cells, J. Clin. Invest. 105 (2000) 1493–1499. doi:10.1172/JCI10229. [28] M.L. Circu, T.Y. Aw, Intestinal redox biology and oxidative stress, Semin. Cell Dev. Biol. 23 (2012) 729–737. doi:10.1016/j.semcdb.2012.03.014. [29] N. Coant, S. Ben Mkaddem, E. Pedruzzi, C. Guichard, X. Tréton, R. Ducroc, J.N. Freund, D. Cazals-Hatem, Y. Bouhnik, P.-L. Woerther, D. Skurnik, A. Grodet, M. Fay, D. Biard, T. Lesuffleur, C. Deffert, R. Moreau, A. Groyer, K.-H. Krause, F. Daniel, E. Ogier-Denis, NADPH oxidase 1 modulates WNT and NOTCH1 signaling to control the fate of proliferative progenitor cells in the colon, Mol. Cell. Biol. 30 (2010) 2636–2650. doi:10.1128/MCB.01194-09. [30] D.P. Jones, Redox potential of GSH/GSSG couple: assay and biological significance, Meth. Enzymol. 348 (2002) 93–112. [31] D.P. Jones, Redefining oxidative stress, Antioxid. Redox Signal. 8 (2006) 1865– 1879. doi:10.1089/ars.2006.8.1865.

- 69 -

[32] Y.S. Nkabyo, T.R. Ziegler, L.H. Gu, W.H. Watson, D.P. Jones, Glutathione and thioredoxin redox during differentiation in human colon epithelial (Caco-2) cells, Am. J. Physiol. Gastrointest. Liver Physiol. 283 (2002) G1352-1359. doi:10.1152/ajpgi.00183.2002. [33] T.Y. Aw, Biliary glutathione promotes the mucosal metabolism of luminal peroxidized lipids by rat small intestine in vivo, J. Clin. Invest. 94 (1994) 1218– 1225. doi:10.1172/JCI117439. [34] M.L. Circu, T.Y. Aw, Redox biology of the intestine, Free Radic. Res. 45 (2011) 1245–1266. doi:10.3109/10715762.2011.611509. [35] T.Y. Aw, G. Wierzbicka, D.P. Jones, Oral glutathione increases tissue glutathione in vivo, Chem. Biol. Interact. 80 (1991) 89–97. [36] P.S. Samiec, L.J. Dahm, D.P. Jones, Glutathione S-transferase in mucus of rat small intestine, Toxicol. Sci. 54 (2000) 52–59. [37] T.Y. Aw, Intestinal glutathione: determinant of mucosal peroxide transport, metabolism, and oxidative susceptibility, Toxicol. Appl. Pharmacol. 204 (2005) 320–328. doi:10.1016/j.taap.2004.11.016. [38] S.P. Wolff, J. Nourooz-Zadeh, Hypothesis: UK consumption of dietary lipid hydroperoxides--a possible contributory factor to atherosclerosis, Atherosclerosis. 119 (1996) 261–263. [39] S. Tsunada, R. Iwakiri, T. Noda, K. Fujimoto, J. Fuseler, C.A. Rhoads, T.Y. Aw, Chronic exposure to subtoxic levels of peroxidized lipids suppresses mucosal cell turnover in rat small intestine and reversal by glutathione, Dig. Dis. Sci. 48 (2003) 210–222.

- 70 -

[40] T. Noda, R. Iwakiri, K. Fujimoto, T.Y. Aw, Induction of mild intracellular redox imbalance inhibits proliferation of CaCo-2 cells, FASEB J. 15 (2001) 2131–2139. doi:10.1096/fj.01-0131com. [41] Y. Gotoh, T. Noda, R. Iwakiri, K. Fujimoto, C.A. Rhoads, T.Y. Aw, Lipid peroxide-induced

redox

imbalance

differentially mediates

CaCo-2

cell

proliferation and growth arrest, Cell Prolif. 35 (2002) 221–235. [42] C.R. Jonas, T.R. Ziegler, L.H. Gu, D.P. Jones, Extracellular thiol/disulfide redox state affects proliferation rate in a human colon carcinoma (Caco2) cell line, Free Radic. Biol. Med. 33 (2002) 1499–1506. [43] Y.S. Nkabyo, Y.-M. Go, T.R. Ziegler, D.P. Jones, Extracellular cysteine/cystine redox regulates the p44/p42 MAPK pathway by metalloproteinase-dependent epidermal growth factor receptor signaling, Am. J. Physiol. Gastrointest. Liver Physiol. 289 (2005) G70-78. doi:10.1152/ajpgi.00280.2004. [44] C.L. Anderson, S.S. Iyer, T.R. Ziegler, D.P. Jones, Control of extracellular cysteine/cystine redox state by HT-29 cells is independent of cellular glutathione, Am. J. Physiol. Regul. Integr. Comp. Physiol. 293 (2007) R1069-1075. doi:10.1152/ajpregu.00195.2007. [45] Y.O. Mannery, T.R. Ziegler, L. Hao, Y. Shyntum, D.P. Jones, Characterization of apical and basal thiol-disulfide redox regulation in human colonic epithelial cells, Am.

J.

Physiol.

Gastrointest.

Liver

Physiol.

299

(2010)

G523-530.

doi:10.1152/ajpgi.00359.2009. [46] A. Gregorieff, D. Pinto, H. Begthel, O. Destrée, M. Kielman, H. Clevers, Expression pattern of Wnt signaling components in the adult intestine, Gastroenterology. 129 (2005) 626–638. doi:10.1016/j.gastro.2005.06.007.

- 71 -

[47] B. Rubinfeld, I. Albert, E. Porfiri, C. Fiol, S. Munemitsu, P. Polakis, Binding of GSK3beta to the APC-beta-catenin complex and regulation of complex assembly, Science. 272 (1996) 1023–1026. [48] S. Amit, A. Hatzubai, Y. Birman, J.S. Andersen, E. Ben-Shushan, M. Mann, Y. Ben-Neriah, I. Alkalay, Axin-mediated CKI phosphorylation of beta-catenin at Ser 45: a molecular switch for the Wnt pathway, Genes Dev. 16 (2002) 1066– 1076. doi:10.1101/gad.230302. [49] R.A. Cavallo, R.T. Cox, M.M. Moline, J. Roose, G.A. Polevoy, H. Clevers, M. Peifer, A. Bejsovec, Drosophila Tcf and Groucho interact to repress Wingless signalling activity, Nature. 395 (1998) 604–608. doi:10.1038/26982. [50] V. Korinek, N. Barker, P. Moerer, E. van Donselaar, G. Huls, P.J. Peters, H. Clevers, Depletion of epithelial stem-cell compartments in the small intestine of mice lacking Tcf-4, Nat. Genet. 19 (1998) 379–383. doi:10.1038/1270. [51] D. Pinto, A. Gregorieff, H. Begthel, H. Clevers, Canonical Wnt signals are essential for homeostasis of the intestinal epithelium, Genes Dev. 17 (2003) 1709–1713. doi:10.1101/gad.267103. [52] F. Kuhnert, C.R. Davis, H.-T. Wang, P. Chu, M. Lee, J. Yuan, R. Nusse, C.J. Kuo, Essential requirement for Wnt signaling in proliferation of adult small intestine and colon revealed by adenoviral expression of Dickkopf-1, Proc. Natl. Acad. Sci. U.S.A. 101 (2004) 266–271. doi:10.1073/pnas.2536800100. [53] H. Ireland, R. Kemp, C. Houghton, L. Howard, A.R. Clarke, O.J. Sansom, D.J. Winton, Inducible Cre-mediated control of gene expression in the murine gastrointestinal tract: effect of loss of beta-catenin, Gastroenterology. 126 (2004) 1236–1246.

- 72 -

[54] T. Fevr, S. Robine, D. Louvard, J. Huelsken, Wnt/beta-catenin is essential for intestinal homeostasis and maintenance of intestinal stem cells, Mol. Cell. Biol. 27 (2007) 7551–7559. doi:10.1128/MCB.01034-07. [55] K.-A. Kim, M. Kakitani, J. Zhao, T. Oshima, T. Tang, M. Binnerts, Y. Liu, B. Boyle, E. Park, P. Emtage, W.D. Funk, K. Tomizuka, Mitogenic influence of human R-spondin1 on the intestinal epithelium, Science. 309 (2005) 1256–1259. doi:10.1126/science.1112521. [56] O.J. Sansom, K.R. Reed, A.J. Hayes, H. Ireland, H. Brinkmann, I.P. Newton, E. Batlle, P. Simon-Assmann, H. Clevers, I.S. Nathke, A.R. Clarke, D.J. Winton, Loss of Apc in vivo immediately perturbs Wnt signaling, differentiation, and migration, Genes Dev. 18 (2004) 1385–1390. doi:10.1101/gad.287404. [57] V. Korinek, N. Barker, P. Moerer, E. van Donselaar, G. Huls, P.J. Peters, H. Clevers, Depletion of epithelial stem-cell compartments in the small intestine of mice lacking Tcf-4, Nat. Genet. 19 (1998) 379–383. doi:10.1038/1270. [58] Q. Yang, N.A. Bermingham, M.J. Finegold, H.Y. Zoghbi, Requirement of Math1 for secretory cell lineage commitment in the mouse intestine, Science. 294 (2001) 2155–2158. doi:10.1126/science.1065718. [59] S. Fre, M. Huyghe, P. Mourikis, S. Robine, D. Louvard, S. Artavanis-Tsakonas, Notch signals control the fate of immature progenitor cells in the intestine, Nature. 435 (2005) 964–968. doi:10.1038/nature03589. [60] J.H. van Es, M.E. van Gijn, O. Riccio, M. van den Born, M. Vooijs, H. Begthel, M. Cozijnsen, S. Robine, D.J. Winton, F. Radtke, H. Clevers, Notch/gammasecretase inhibition turns proliferative cells in intestinal crypts and adenomas into goblet cells, Nature. 435 (2005) 959–963. doi:10.1038/nature03659.

- 73 -

[61] J. Jensen, E.E. Pedersen, P. Galante, J. Hald, R.S. Heller, M. Ishibashi, R. Kageyama, F. Guillemot, P. Serup, O.D. Madsen, Control of endodermal endocrine

development

by

Hes-1,

Nat.

Genet.

24

(2000)

36–44.

doi:10.1038/71657. [62] M. Jenny, C. Uhl, C. Roche, I. Duluc, V. Guillermin, F. Guillemot, J. Jensen, M. Kedinger, G. Gradwohl, Neurogenin3 is differentially required for endocrine cell fate specification in the intestinal and gastric epithelium, EMBO J. 21 (2002) 6338–6347. [63] G.T. Wong, D. Manfra, F.M. Poulet, Q. Zhang, H. Josien, T. Bara, L. Engstrom, M. Pinzon-Ortiz, J.S. Fine, H.-J.J. Lee, L. Zhang, G.A. Higgins, E.M. Parker, Chronic treatment with the gamma-secretase inhibitor LY-411,575 inhibits betaamyloid peptide production and alters lymphopoiesis and intestinal cell differentiation,

J.

Biol.

Chem.

279

(2004)

12876–12882.

doi:10.1074/jbc.M311652200. [64] K. Suzuki, H. Fukui, T. Kayahara, M. Sawada, H. Seno, H. Hiai, R. Kageyama, H. Okano, T. Chiba, Hes1-deficient mice show precocious differentiation of Paneth cells in the small intestine, Biochem. Biophys. Res. Commun. 328 (2005) 348–352. doi:10.1016/j.bbrc.2004.12.174. [65] M. Vooijs, C.-T. Ong, B. Hadland, S. Huppert, Z. Liu, J. Korving, M. van den Born, T. Stappenbeck, Y. Wu, H. Clevers, R. Kopan, Mapping the consequence of Notch1 proteolysis in vivo with NIP-CRE, Development. 134 (2007) 535–544. doi:10.1242/dev.02733. [66] B.Z. Stanger, R. Datar, L.C. Murtaugh, D.A. Melton, Direct regulation of intestinal fate by Notch, Proc. Natl. Acad. Sci. U.S.A. 102 (2005) 12443–12448. doi:10.1073/pnas.0505690102.

- 74 -

[67] J.C. Mills, J.I. Gordon, The intestinal stem cell niche: there grows the neighborhood, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 12334–12336. doi:10.1073/pnas.231487198. [68] D.W. Powell, R.C. Mifflin, J.D. Valentich, S.E. Crowe, J.I. Saada, A.B. West, Myofibroblasts. II. Intestinal subepithelial myofibroblasts, Am. J. Physiol. 277 (1999) C183-201. [69] A.-P.G. Haramis, H. Begthel, M. van den Born, J. van Es, S. Jonkheer, G.J.A. Offerhaus, H. Clevers, De novo crypt formation and juvenile polyposis on BMP inhibition

in

mouse

intestine,

Science.

303

(2004)

1684–1686.

doi:10.1126/science.1093587. [70] X.C. He, J. Zhang, W.-G. Tong, O. Tawfik, J. Ross, D.H. Scoville, Q. Tian, X. Zeng, X. He, L.M. Wiedemann, Y. Mishina, L. Li, BMP signaling inhibits intestinal stem cell self-renewal through suppression of Wnt-beta-catenin signaling, Nat. Genet. 36 (2004) 1117–1121. doi:10.1038/ng1430. [71] J. Massagué, J. Seoane, D. Wotton, Smad transcription factors, Genes Dev. 19 (2005) 2783–2810. doi:10.1101/gad.1350705. [72] B.B. Madison, K. Braunstein, E. Kuizon, K. Portman, X.T. Qiao, D.L. Gumucio, Epithelial hedgehog signals pattern the intestinal crypt-villus axis, Development. 132 (2005) 279–289. doi:10.1242/dev.01576. [73] L. Karlsson, P. Lindahl, J.K. Heath, C. Betsholtz, Abnormal gastrointestinal development in PDGF-A and PDGFR-(alpha) deficient mice implicates a novel mesenchymal

structure

with

putative

instructive

properties

in

villus

morphogenesis, Development. 127 (2000) 3457–3466. [74] E.C. Cheung, D. Athineos, P. Lee, R.A. Ridgway, W. Lambie, C. Nixon, D. Strathdee, K. Blyth, O.J. Sansom, K.H. Vousden, TIGAR is required for efficient

- 75 -

intestinal regeneration and tumorigenesis, Dev. Cell. 25 (2013) 463–477. doi:10.1016/j.devcel.2013.05.001. [75] E.C. Cheung, P. Lee, F. Ceteci, C. Nixon, K. Blyth, O.J. Sansom, K.H. Vousden, Opposing effects of TIGAR- and RAC1-derived ROS on Wnt-driven proliferation in the mouse intestine, Genes Dev. 30 (2016) 52–63. doi:10.1101/gad.271130.115. [76] K.B. Myant, P. Cammareri, E.J. McGhee, R.A. Ridgway, D.J. Huels, J.B. Cordero, S. Schwitalla, G. Kalna, E.-L. Ogg, D. Athineos, P. Timpson, M. Vidal, G.I. Murray, F.R. Greten, K.I. Anderson, O.J. Sansom, ROS production and NFκB activation triggered by RAC1 facilitate WNT-driven intestinal stem cell proliferation and colorectal cancer initiation, Cell Stem Cell. 12 (2013) 761–773. doi:10.1016/j.stem.2013.04.006. [77] H.J. Forman, O. Augusto, R. Brigelius-Flohe, P.A. Dennery, B. Kalyanaraman, H. Ischiropoulos, G.E. Mann, R. Radi, L.J. Roberts, J. Vina, K.J.A. Davies, Even free radicals should follow some rules: a guide to free radical research terminology and methodology, Free Radic. Biol. Med. 78 (2015) 233–235. doi:10.1016/j.freeradbiomed.2014.10.504. [78] M. Geiszt, K. Lekstrom, S. Brenner, S.M. Hewitt, R. Dana, H.L. Malech, T.L. Leto, NAD(P)H oxidase 1, a product of differentiated colon epithelial cells, can partially replace glycoprotein 91phox in the regulated production of superoxide by phagocytes, J. Immunol. 171 (2003) 299–306. [79] A.J. Valente, Q. Zhou, Z. Lu, W. He, M. Qiang, W. Ma, G. Li, L. Wang, B. Banfi, K. Steger, K.-H. Krause, R.A. Clark, S. Li, Regulation of NOX1 expression by GATA, HNF-1alpha, and Cdx transcription factors, Free Radic. Biol. Med. 44 (2008) 430–443. doi:10.1016/j.freeradbiomed.2007.10.035.

- 76 -

[80] T. Kawahara, Y. Kuwano, S. Teshima-Kondo, R. Takeya, H. Sumimoto, K. Kishi, S. Tsunawaki, T. Hirayama, K. Rokutan, Role of nicotinamide adenine dinucleotide phosphate oxidase 1 in oxidative burst response to Toll-like receptor 5 signaling in large intestinal epithelial cells, J. Immunol. 172 (2004) 3051–3058. [81] E. Laurent, J.W. McCoy, R.A. Macina, W. Liu, G. Cheng, S. Robine, J. Papkoff, J.D. Lambeth, Nox1 is over-expressed in human colon cancers and correlates with activating mutations in K-Ras, Int. J. Cancer. 123 (2008) 100–107. doi:10.1002/ijc.23423. [82] S. Kajla, A.S. Mondol, A. Nagasawa, Y. Zhang, M. Kato, K. Matsuno, C. YabeNishimura, T. Kamata, A crucial role for Nox 1 in redox-dependent regulation of Wnt-β-catenin signaling, FASEB J. 26 (2012) 2049–2059. doi:10.1096/fj.11196360. [83] J.D. Lambeth, Nox enzymes, ROS, and chronic disease: an example of antagonistic pleiotropy, Free Radic. Biol. Med. 43 (2007) 332–347. doi:10.1016/j.freeradbiomed.2007.03.027. [84] J.D. Lambeth, T. Kawahara, B. Diebold, Regulation of Nox and Duox enzymatic activity and expression, Free Radic. Biol. Med. 43 (2007) 319–331. doi:10.1016/j.freeradbiomed.2007.03.028. [85] Y. Funato, T. Michiue, M. Asashima, H. Miki, The thioredoxin-related redoxregulating protein nucleoredoxin inhibits Wnt-beta-catenin signalling through dishevelled, Nat. Cell Biol. 8 (2006) 501–508. doi:10.1038/ncb1405. [86] C.W. Barrett, W. Ning, X. Chen, J.J. Smith, M.K. Washington, K.E. Hill, L.A. Coburn, R.M. Peek, R. Chaturvedi, K.T. Wilson, R.F. Burk, C.S. Williams, Tumor suppressor function of the plasma glutathione peroxidase gpx3 in colitis-

- 77 -

associated carcinoma, Cancer Res. 73 (2013) 1245–1255. doi:10.1158/00085472.CAN-12-3150. [87] F.F. Chu, R.S. Esworthy, The expression of an intestinal form of glutathione peroxidase (GSHPx-GI) in rat intestinal epithelium, Arch. Biochem. Biophys. 323 (1995) 288–294. [88] F.-F. Chu, R.S. Esworthy, J.H. Doroshow, Role of Se-dependent glutathione peroxidases in gastrointestinal inflammation and cancer, Free Radic. Biol. Med. 36 (2004) 1481–1495. doi:10.1016/j.freeradbiomed.2004.04.010. [89] B. Sido, T. Giese, F. Autschbach, F. Lasitschka, J. Braunstein, S.C. Meuer, Potential role of thioredoxin in immune responses in intestinal lamina propria T lymphocytes, Eur. J. Immunol. 35 (2005) 408–417. doi:10.1002/eji.200424500. [90] J.R. Godoy, M. Funke, W. Ackermann, P. Haunhorst, S. Oesteritz, F. Capani, H.P. Elsässer, C.H. Lillig, Redox atlas of the mouse. Immunohistochemical detection of glutaredoxin-, peroxiredoxin-, and thioredoxin-family proteins in various tissues of the laboratory mouse, Biochim. Biophys. Acta. 1810 (2011) 2– 92. doi:10.1016/j.bbagen.2010.05.006. [91] E.-M. Ha, C.-T. Oh, Y.S. Bae, W.-J. Lee, A direct role for dual oxidase in Drosophila

gut

immunity,

Science.

310

(2005)

847–850.

doi:10.1126/science.1117311. [92] F. Sommer, F. Bäckhed, The gut microbiota engages different signaling pathways to induce Duox2 expression in the ileum and colon epithelium, Mucosal Immunol. 8 (2015) 372–379. doi:10.1038/mi.2014.74. [93] S.C. Liang, X.-Y. Tan, D.P. Luxenberg, R. Karim, K. Dunussi-Joannopoulos, M. Collins, L.A. Fouser, Interleukin (IL)-22 and IL-17 are coexpressed by Th17 cells

- 78 -

and cooperatively enhance expression of antimicrobial peptides, J. Exp. Med. 203 (2006) 2271–2279. doi:10.1084/jem.20061308. [94] H. Grasberger, J. Gao, H. Nagao-Kitamoto, S. Kitamoto, M. Zhang, N. Kamada, K.A. Eaton, M. El-Zaatari, A.B. Shreiner, J.L. Merchant, C. Owyang, J.Y. Kao, Increased Expression of DUOX2 Is an Epithelial Response to Mucosal Dysbiosis Required for Immune Homeostasis in Mouse Intestine, Gastroenterology. 149 (2015) 1849–1859. doi:10.1053/j.gastro.2015.07.062. [95] H. Grasberger, M. El-Zaatari, D.T. Dang, J.L. Merchant, Dual oxidases control release of hydrogen peroxide by the gastric epithelium to prevent Helicobacter felis infection and inflammation in mice, Gastroenterology. 145 (2013) 1045– 1054. doi:10.1053/j.gastro.2013.07.011. [96] E.-M. Ha, K.-A. Lee, Y.Y. Seo, S.-H. Kim, J.-H. Lim, B.-H. Oh, J. Kim, W.-J. Lee, Coordination of multiple dual oxidase-regulatory pathways in responses to commensal and infectious microbes in drosophila gut, Nat. Immunol. 10 (2009) 949–957. doi:10.1038/ni.1765. [97] K. Rokutan, T. Kawahara, Y. Kuwano, K. Tominaga, A. Sekiyama, S. TeshimaKondo, NADPH oxidases in the gastrointestinal tract: a potential role of Nox1 in innate immune response and carcinogenesis, Antioxid. Redox Signal. 8 (2006) 1573–1582. doi:10.1089/ars.2006.8.1573. [98] X. Fu, S.Y. Kassim, W.C. Parks, J.W. Heinecke, Hypochlorous acid oxygenates the cysteine switch domain of pro-matrilysin (MMP-7). A mechanism for matrix metalloproteinase myeloperoxidase,

activation J.

Biol.

and

atherosclerotic

Chem.

doi:10.1074/jbc.M106958200.

- 79 -

276

plaque (2001)

rupture

by

41279–41287.

[99] C.L. Wilson, A.J. Ouellette, D.P. Satchell, T. Ayabe, Y.S. López-Boado, J.L. Stratman, S.J. Hultgren, L.M. Matrisian, W.C. Parks, Regulation of intestinal alpha-defensin activation by the metalloproteinase matrilysin in innate host defense, Science. 286 (1999) 113–117. [100] B.O. Schroeder, Z. Wu, S. Nuding, S. Groscurth, M. Marcinowski, J. Beisner, J. Buchner, M. Schaller, E.F. Stange, J. Wehkamp, Reduction of disulphide bonds unmasks potent antimicrobial activity of human β-defensin 1, Nature. 469 (2011) 419–423. doi:10.1038/nature09674. [101] S. Damiano, A. Morano, V. Ucci, R. Accetta, P. Mondola, R. Paternò, V.E. Avvedimento, M. Santillo, Dual oxidase 2 generated reactive oxygen species selectively mediate the induction of mucins by epidermal growth factor in enterocytes,

Int.

J.

Biochem.

Cell

Biol.

60

(2015)

8–18.

doi:10.1016/j.biocel.2014.12.014. [102] A. Alam, G. Leoni, C.C. Wentworth, J.M. Kwal, H. Wu, C.S. Ardita, P.A. Swanson, J.D. Lambeth, R.M. Jones, A. Nusrat, A.S. Neish, Redox signaling regulates commensal-mediated mucosal homeostasis and restitution and requires formyl

peptide

receptor

1,

Mucosal

Immunol.

7

(2014)

645–655.

doi:10.1038/mi.2013.84. [103] B.A. Babbin, A.J. Jesaitis, A.I. Ivanov, D. Kelly, M. Laukoetter, P. Nava, C.A. Parkos, A. Nusrat, Formyl peptide receptor-1 activation enhances intestinal epithelial cell restitution through phosphatidylinositol 3-kinase-dependent activation of Rac1 and Cdc42, J. Immunol. 179 (2007) 8112–8121. [104] B.A. Babbin, W.Y. Lee, C.A. Parkos, L.M. Winfree, A. Akyildiz, M. Perretti, A. Nusrat, Annexin I regulates SKCO-15 cell invasion by signaling through formyl

- 80 -

peptide

receptors,

J.

Biol.

Chem.

281

(2006)

19588–19599.

doi:10.1074/jbc.M513025200. [105] G. Leoni, A. Alam, P.-A. Neumann, J.D. Lambeth, G. Cheng, J. McCoy, R.S. Hilgarth, K. Kundu, N. Murthy, D. Kusters, C. Reutelingsperger, M. Perretti, C.A. Parkos, A.S. Neish, A. Nusrat, Annexin A1, formyl peptide receptor, and NOX1 orchestrate epithelial repair, J. Clin. Invest. 123 (2013) 443–454. doi:10.1172/JCI65831. [106] F. Bäckhed, R.E. Ley, J.L. Sonnenburg, D.A. Peterson, J.I. Gordon, Hostbacterial mutualism in the human intestine, Science. 307 (2005) 1915–1920. doi:10.1126/science.1104816. [107] S.R. Gill, M. Pop, R.T. Deboy, P.B. Eckburg, P.J. Turnbaugh, B.S. Samuel, J.I. Gordon, D.A. Relman, C.M. Fraser-Liggett, K.E. Nelson, Metagenomic analysis of the human distal gut microbiome, Science. 312 (2006) 1355–1359. doi:10.1126/science.1124234. [108] L. Dethlefsen, M. McFall-Ngai, D.A. Relman, An ecological and evolutionary perspective on human-microbe mutualism and disease, Nature. 449 (2007) 811– 818. doi:10.1038/nature06245. [109] P.B. Eckburg, E.M. Bik, C.N. Bernstein, E. Purdom, L. Dethlefsen, M. Sargent, S.R. Gill, K.E. Nelson, D.A. Relman, Diversity of the human intestinal microbial flora, Science. 308 (2005) 1635–1638. doi:10.1126/science.1110591. [110] J. Xu, M.A. Mahowald, R.E. Ley, C.A. Lozupone, M. Hamady, E.C. Martens, B. Henrissat, P.M. Coutinho, P. Minx, P. Latreille, H. Cordum, A. Van Brunt, K. Kim, R.S. Fulton, L.A. Fulton, S.W. Clifton, R.K. Wilson, R.D. Knight, J.I. Gordon, Evolution of symbiotic bacteria in the distal human intestine, PLoS Biol. 5 (2007) e156. doi:10.1371/journal.pbio.0050156.

- 81 -

[111] L.V. Hooper, J.I. Gordon, Commensal host-bacterial relationships in the gut, Science. 292 (2001) 1115–1118. [112] K. Madsen, A. Cornish, P. Soper, C. McKaigney, H. Jijon, C. Yachimec, J. Doyle, L. Jewell, C. De Simone, Probiotic bacteria enhance murine and human intestinal epithelial barrier function, Gastroenterology. 121 (2001) 580–591. [113] A.S. Neish, Microbes in gastrointestinal health and disease, Gastroenterology. 136 (2009) 65–80. doi:10.1053/j.gastro.2008.10.080. [114] M. Wilson, The gastrointestinal tract and its indigenous microbiota, in: Microbial Inhabitants of Humans, 1 edition, Cambridge University Press, New York, 2004: pp. 251–317. [115] L.V. Hooper, M.H. Wong, A. Thelin, L. Hansson, P.G. Falk, J.I. Gordon, Molecular analysis of commensal host-microbial relationships in the intestine, Science. 291 (2001) 881–884. doi:10.1126/science.291.5505.881. [116] S. Rakoff-Nahoum, J. Paglino, F. Eslami-Varzaneh, S. Edberg, R. Medzhitov, Recognition of commensal microflora by toll-like receptors is required for intestinal homeostasis, Cell. 118 (2004) 229–241. doi:10.1016/j.cell.2004.07.002. [117] J. Lee, J.-H. Mo, K. Katakura, I. Alkalay, A.N. Rucker, Y.-T. Liu, H.-K. Lee, C. Shen, G. Cojocaru, S. Shenouda, M. Kagnoff, L. Eckmann, Y. Ben-Neriah, E. Raz, Maintenance of colonic homeostasis by distinctive apical TLR9 signalling in intestinal

epithelial

cells,

Nat.

Cell

Biol.

8

(2006)

1327–1336.

doi:10.1038/ncb1500. [118] J.D. Lambeth, NOX enzymes and the biology of reactive oxygen, Nat. Rev. Immunol. 4 (2004) 181–189. doi:10.1038/nri1312.

- 82 -

[119] K. Bedard, K.-H. Krause, The NOX family of ROS-generating NADPH oxidases: physiology

and

pathophysiology,

Physiol.

Rev.

87

(2007)

245–313.

doi:10.1152/physrev.00044.2005. [120] R.M. Jones, L. Luo, C.S. Ardita, A.N. Richardson, Y.M. Kwon, J.W. Mercante, A. Alam, C.L. Gates, H. Wu, P.A. Swanson, J.D. Lambeth, P.W. Denning, A.S. Neish, Symbiotic lactobacilli stimulate gut epithelial proliferation via Noxmediated generation of reactive oxygen species, EMBO J. 32 (2013) 3017–3028. doi:10.1038/emboj.2013.224. [121] P.A. Swanson, A. Kumar, S. Samarin, M. Vijay-Kumar, K. Kundu, N. Murthy, J. Hansen, A. Nusrat, A.S. Neish, Enteric commensal bacteria potentiate epithelial restitution via reactive oxygen species-mediated inactivation of focal adhesion kinase phosphatases, Proc. Natl. Acad. Sci. U.S.A. 108 (2011) 8803–8808. doi:10.1073/pnas.1010042108. [122] C.C. Wentworth, A. Alam, R.M. Jones, A. Nusrat, A.S. Neish, Enteric commensal bacteria induce extracellular signal-regulated kinase pathway signaling via formyl peptide receptor-dependent redox modulation of dual specific

phosphatase

3,

J.

Biol.

Chem.

286

(2011)

38448–38455.

doi:10.1074/jbc.M111.268938. [123] A. Kumar, H. Wu, L.S. Collier-Hyams, J.M. Hansen, T. Li, K. Yamoah, Z.-Q. Pan, D.P. Jones, A.S. Neish, Commensal bacteria modulate cullin-dependent signaling via generation of reactive oxygen species, EMBO J. 26 (2007) 4457– 4466. doi:10.1038/sj.emboj.7601867. [124] A.S. Neish, A.T. Gewirtz, H. Zeng, A.N. Young, M.E. Hobert, V. Karmali, A.S. Rao, J.L. Madara, Prokaryotic regulation of epithelial responses by inhibition of IkappaB-alpha ubiquitination, Science. 289 (2000) 1560–1563.

- 83 -

[125] L.S. Collier-Hyams, V. Sloane, B.C. Batten, A.S. Neish, Cutting edge: bacterial modulation of epithelial signaling via changes in neddylation of cullin-1, J. Immunol. 175 (2005) 4194–4198. [126] W.-J. Lee, Bacterial-modulated signaling pathways in gut homeostasis, Sci Signal. 1 (2008) pe24. doi:10.1126/stke.121pe24. [127] P. Rada, A.I. Rojo, A. Offergeld, G.J. Feng, J.P. Velasco-Martín, J.M. GonzálezSancho, Á.M. Valverde, T. Dale, J. Regadera, A. Cuadrado, WNT-3A regulates an Axin1/NRF2 complex that regulates antioxidant metabolism in hepatocytes, Antioxid. Redox Signal. 22 (2015) 555–571. doi:10.1089/ars.2014.6040. [128] R.M. Jones, C. Desai, T.M. Darby, L. Luo, A.A. Wolfarth, C.D. Scharer, C.S. Ardita, A.R. Reedy, E.S. Keebaugh, A.S. Neish, Lactobacilli Modulate Epithelial Cytoprotection through the Nrf2 Pathway, Cell Rep. 12 (2015) 1217–1225. doi:10.1016/j.celrep.2015.07.042. [129] K.T. Wilson, S. Fu, K.S. Ramanujam, S.J. Meltzer, Increased expression of inducible nitric oxide synthase and cyclooxygenase-2 in Barrett’s esophagus and associated adenocarcinomas, Cancer Res. 58 (1998) 2929–2934. [130] P. Sharma, Clinical practice. Barrett’s esophagus, N. Engl. J. Med. 361 (2009) 2548–2556. doi:10.1056/NEJMcp0902173. [131] J. Kong, K.A. Whelan, D. Laczkó, B. Dang, A. Caro Monroig, A. Soroush, J. Falcone, R.K. Amaravadi, A.K. Rustgi, G.G. Ginsberg, G.W. Falk, H. Nakagawa, J.P. Lynch, Autophagy levels are elevated in barrett’s esophagus and promote cell survival

from

acid

and

oxidative

stress,

Mol.

Carcinog.

(2015).

doi:10.1002/mc.22406. [132] M.F. Buas, Q. He, L.G. Johnson, L. Onstad, D.M. Levine, A.P. Thrift, P. Gharahkhani, C. Palles, J. Lagergren, R.C. Fitzgerald, W. Ye, C. Caldas, N.C.

- 84 -

Bird, N.J. Shaheen, L. Bernstein, M.D. Gammon, A.H. Wu, L.J. Hardie, P.D. Pharoah, G. Liu, P. Iyer, D.A. Corley, H.A. Risch, W.-H. Chow, H. Prenen, L. Chegwidden, S. Love, S. Attwood, P. Moayyedi, D. MacDonald, R. Harrison, P. Watson, H. Barr, J. deCaestecker, I. Tomlinson, J. Jankowski, D.C. Whiteman, S. MacGregor,

T.L.

Vaughan,

M.M.

Madeleine,

Germline

variation

in

inflammation-related pathways and risk of Barrett’s oesophagus and oesophageal adenocarcinoma, Gut. (2016). doi:10.1136/gutjnl-2016-311622. [133] S. Song, S. Guha, K. Liu, N.S. Buttar, R.S. Bresalier, COX-2 induction by unconjugated bile acids involves reactive oxygen species-mediated signalling pathways in Barrett’s oesophagus and oesophageal adenocarcinoma, Gut. 56 (2007) 1512–1521. doi:10.1136/gut.2007.121244. [134] A. Baruah, N. Buttar, R. Chandra, X. Chen, N.J. Clemons, D. Compare, W. ElRifai, J. Gu, C.W. Houchen, S.Y. Koh, W. Li, G. Nardone, W.A. Phillips, A. Sharma, I. Singh, M.P. Upton, K.J. Vega, X. Wu, Translational research on Barrett’s esophagus, Ann. N. Y. Acad. Sci. 1325 (2014) 170–186. doi:10.1111/nyas.12531. [135] G.J. Wetscher, R.A. Hinder, D. Bagchi, P.R. Hinder, M. Bagchi, G. Perdikis, T. McGinn, Reflux esophagitis in humans is mediated by oxygen-derived free radicals, Am. J. Surg. 170 (1995) 552-556-557. [136] G.J. Wetscher, R.A. Hinder, P. Klingler, M. Gadenstätter, G. Perdikis, P.R. Hinder, Reflux esophagitis in humans is a free radical event, Dis. Esophagus. 10 (1997) 29–32; discussion 33. [137] P. Jiménez, E. Piazuelo, M.T. Sánchez, J. Ortego, F. Soteras, A. Lanas, Free radicals and antioxidant systems in reflux esophagitis and Barrett’s esophagus, World J. Gastroenterol. 11 (2005) 2697–2703.

- 85 -

[138] G.J. Wetscher, G. Perdikis, D.H. Kretchmar, R.G. Stinson, D. Bagchi, E.J. Redmond, T.E. Adrian, R.A. Hinder, Esophagitis in Sprague-Dawley rats is mediated by free radicals, Dig. Dis. Sci. 40 (1995) 1297–1305. [139] E. Piazuelo, C. Cebrián, A. Escartín, P. Jiménez, F. Soteras, J. Ortego, A. Lanas, Superoxide dismutase prevents development of adenocarcinoma in a rat model of Barrett’s esophagus, World J. Gastroenterol. 11 (2005) 7436–7443. [140] J.H. Song, Y.-M. Han, W.H. Kim, J.-M. Park, M. Jeong, E.J. Go, S.P. Hong, K.B. Hahm, Oxidative stress from reflux esophagitis to esophageal cancer: the alleviation

with

antioxidants,

Free

Radic.

Res.

(2016)

1–12.

doi:10.1080/10715762.2016.1181262. [141] X. Fu, D.G. Beer, J. Behar, J. Wands, D. Lambeth, W. Cao, cAMP-response element-binding protein mediates acid-induced NADPH oxidase NOX5-S expression in Barrett esophageal adenocarcinoma cells, J. Biol. Chem. 281 (2006) 20368–20382. doi:10.1074/jbc.M603353200. [142] J. Hong, D. Li, W. Cao, Rho Kinase ROCK2 Mediates Acid-Induced NADPH Oxidase NOX5-S Expression in Human Esophageal Adenocarcinoma Cells, PLoS ONE. 11 (2016) e0149735. doi:10.1371/journal.pone.0149735. [143] D. Li, W. Cao, Role of intracellular calcium and NADPH oxidase NOX5-S in acid-induced DNA damage in Barrett’s cells and Barrett’s esophageal adenocarcinoma cells, Am. J. Physiol. Gastrointest. Liver Physiol. 306 (2014) G863-872. doi:10.1152/ajpgi.00321.2013. [144] D. Peng, T. Hu, M. Soutto, A. Belkhiri, A. Zaika, W. El-Rifai, Glutathione peroxidase 7 has potential tumour suppressor functions that are silenced by location-specific methylation in oesophageal adenocarcinoma, Gut. 63 (2014) 540–551. doi:10.1136/gutjnl-2013-304612.

- 86 -

[145] D. Peng, A. Belkhiri, T. Hu, R. Chaturvedi, M. Asim, K.T. Wilson, A. Zaika, W. El-Rifai, Glutathione peroxidase 7 protects against oxidative DNA damage in oesophageal cells, Gut. 61 (2012) 1250–1260. doi:10.1136/gutjnl-2011-301078. [146] D.-F. Peng, T.-L. Hu, M. Soutto, A. Belkhiri, W. El-Rifai, Loss of glutathione peroxidase 7 promotes carcinogenesis,

TNF-α-induced

Carcinogenesis.

NF-κB 35

activation in (2014)

Barrett’s

1620–1628.

doi:10.1093/carcin/bgu083. [147] G.J. Wetscher, P.R. Hinder, D. Bagchi, G. Perdikis, E.J. Redmond, K. Glaser, T.E. Adrian, R.A. Hinder, Free radical scavengers prevent reflux esophagitis in rats, Dig. Dis. Sci. 40 (1995) 1292–1296. [148] J. Hong, Z. Chen, D. Peng, A. Zaika, F. Revetta, M.K. Washington, A. Belkhiri, W. El-Rifai, APE1-mediated DNA damage repair provides survival advantage for esophageal adenocarcinoma cells in response to acidic bile salts, Oncotarget. 7 (2016) 16688–16702. doi:10.18632/oncotarget.7696. [149] P. Malfertheiner, F.K. Chan, K.E. McColl, Peptic ulcer disease, Lancet. 374 (2009) 1449–1461. doi:10.1016/S0140-6736(09)60938-7. [150] R.S. Sandler, J.E. Everhart, M. Donowitz, E. Adams, K. Cronin, C. Goodman, E. Gemmen, S. Shah, A. Avdic, R. Rubin, The burden of selected digestive diseases in the United States, Gastroenterology. 122 (2002) 1500–1511. [151] K. Soreide, K. Thorsen, E.M. Harrison, J. Bingener, M.H. Moller, M. OheneYeboah, J.A. Soreide, Perforated peptic ulcer, Lancet. 386 (2015) 1288–1298. doi:10.1016/S0140-6736(15)00276-7. [152] Y. Yuan, I.T. Padol, R.H. Hunt, Peptic ulcer disease today, Nat Clin Pract Gastroenterol Hepatol. 3 (2006) 80–89. doi:10.1038/ncpgasthep0393.

- 87 -

[153] A. Nomura, G.N. Stemmermann, P.H. Chyou, G.I. Perez-Perez, M.J. Blaser, Helicobacter pylori infection and the risk for duodenal and gastric ulceration, Ann.Intern.Med. 120 (1994) 977–981. [154] E.J. Kuipers, J.C. Thijs, H.P. Festen, The prevalence of Helicobacter pylori in peptic ulcer disease, Aliment.Pharmacol.Ther. 9 Suppl 2 (1995) 59–69. [155] A. Ford, B. Delaney, D. Forman, P. Moayyedi, Eradication therapy for peptic ulcer disease in Helicobacter pylori positive patients, Cochrane Database Syst.Rev. (4) (2004) CD003840. doi:10.1002/14651858.CD003840.pub2. [156] A.C. Ford, B.C. Delaney, D. Forman, P. Moayyedi, Eradication therapy for peptic ulcer disease in Helicobacter pylori positive patients, Cochrane Database Syst.Rev. (2) (2006) CD003840. doi:10.1002/14651858.CD003840.pub4. [157] B.J. Marshall, J.R. Warren, Unidentified curved bacilli in the stomach of patients with gastritis and peptic ulceration, Lancet. 1 (1984) 1311–1315. [158] J.G. Kusters, A.H.M. van Vliet, E.J. Kuipers, Pathogenesis of Helicobacter pylori infection, Clin. Microbiol. Rev. 19 (2006) 449–490. doi:10.1128/CMR.00054-05. [159] S.F. Moss, S. Legon, A.E. Bishop, J.M. Polak, J. Calam, Effect of Helicobacter pylori on gastric somatostatin in duodenal ulcer disease, Lancet. 340 (1992) 930– 932. [160] E.M. el-Omar, I.D. Penman, J.E. Ardill, R.S. Chittajallu, C. Howie, K.E. McColl, Helicobacter pylori infection and abnormalities of acid secretion in patients with duodenal ulcer disease, Gastroenterology. 109 (1995) 681–691. [161] C. Mooney, J. Keenan, D. Munster, I. Wilson, R. Allardyce, P. Bagshaw, B. Chapman, V. Chadwick, Neutrophil activation by Helicobacter pylori, Gut. 32 (1991) 853–857.

- 88 -

[162] S.E. Crowe, L. Alvarez, M. Dytoc, R.H. Hunt, M. Muller, P. Sherman, J. Patel, Y. Jin, P.B. Ernst, Expression of interleukin 8 and CD54 by human gastric epithelium after Helicobacter pylori infection in vitro, Gastroenterology. 108 (1995) 65–74. [163] Q.B. Zhang, I.M. Nakashabendi, M.S. Mokhashi, J.B. Dawodu, C.G. Gemmell, R.I. Russell, Association of cytotoxin production and neutrophil activation by strains of Helicobacter pylori isolated from patients with peptic ulceration and chronic gastritis, Gut. 38 (1996) 841–845. [164] Q.B. Zhang, J.B. Dawodu, A. Husain, G. Etolhi, C.G. Gemmell, R.I. Russell, Association of antral mucosal levels of interleukin 8 and reactive oxygen radicals in patients infected with Helicobacter pylori, Clin.Sci.(Lond). 92 (1997) 69–73. [165] M.J. Blaser, G.I. Perez-Perez, H. Kleanthous, T.L. Cover, R.M. Peek, P.H. Chyou, G.N. Stemmermann, A. Nomura, Infection with Helicobacter pylori strains possessing cagA is associated with an increased risk of developing adenocarcinoma of the stomach, Cancer Res. 55 (1995) 2111–2115. [166] L. Manente, A. Perna, E. Buommino, L. Altucci, A. Lucariello, G. Citro, A. Baldi, G. Iaquinto, M.A. Tufano, A. De Luca, The Helicobacter pylori’s protein VacA has direct effects on the regulation of cell cycle and apoptosis in gastric epithelial cells, J.Cell.Physiol. 214 (2008) 582–587. doi:10.1002/jcp.21242. [167] G.R. Davies, N.J. Simmonds, T.R. Stevens, A. Grandison, D.R. Blake, D.S. Rampton, Mucosal reactive oxygen metabolite production in duodenal ulcer disease, Gut. 33 (1992) 1467–1472. [168] G.R. Davies, N.J. Simmonds, T.R. Stevens, M.T. Sheaff, N. Banatvala, I.F. Laurenson, D.R. Blake, D.S. Rampton, Helicobacter pylori stimulates antral mucosal reactive oxygen metabolite production in vivo, Gut. 35 (1994) 179–185.

- 89 -

[169] H. Suzuki, S. Miura, H. Imaeda, M. Suzuki, J.Y. Han, M. Mori, D. Fukumura, M. Tsuchiya, H. Ishii, Enhanced levels of chemiluminescence and platelet activating factor in urease-positive gastric ulcers, Free Radic. Biol. Med. 20 (1996) 449– 454. [170] T. Kawahara, S. Teshima, A. Oka, T. Sugiyama, K. Kishi, K. Rokutan, Type I Helicobacter pylori lipopolysaccharide stimulates toll-like receptor 4 and activates mitogen oxidase 1 in gastric pit cells, Infect.Immun. 69 (2001) 4382– 4389. doi:10.1128/IAI.69.7.4382-4389.2001. [171] T. Kawahara, M. Kohjima, Y. Kuwano, H. Mino, S. Teshima-Kondo, R. Takeya, S. Tsunawaki, A. Wada, H. Sumimoto, K. Rokutan, Helicobacter pylori lipopolysaccharide activates Rac1 and transcription of NADPH oxidase Nox1 and its organizer NOXO1 in guinea pig gastric mucosal cells, Am. J. Physiol., Cell Physiol. 288 (2005) C450-457. doi:10.1152/ajpcell.00319.2004. [172] S.H. Chu, H. Kim, J.Y. Seo, J.W. Lim, N. Mukaida, K.H. Kim, Role of NFkappaB and AP-1 on Helicobater pylori-induced IL-8 expression in AGS cells, Dig.Dis.Sci. 48 (2003) 257–265. [173] A.M. O’Hara, A. Bhattacharyya, R.C. Mifflin, M.F. Smith, K.A. Ryan, K.G.-E. Scott, M. Naganuma, A. Casola, T. Izumi, S. Mitra, P.B. Ernst, S.E. Crowe, Interleukin-8 induction by Helicobacter pylori in gastric epithelial cells is dependent on apurinic/apyrimidinic endonuclease-1/redox factor-1, J. Immunol. 177 (2006) 7990–7999. [174] J.I. Keenan, R.A. Peterson, M.B. Hampton, NADPH oxidase involvement in the pathology of Helicobacter pylori infection, Free Radic. Biol. Med. 38 (2005) 1188–1196. doi:10.1016/j.freeradbiomed.2004.12.025.

- 90 -

[175] J.M. Götz, J.L. Thio, H.W. Verspaget, G.J. Offerhaus, I. Biemond, C.B. Lamers, R.A. Veenendaal, Treatment of Helicobacter pylori infection favourably affects gastric mucosal superoxide dismutases, Gut. 40 (1997) 591–596. [176] H. Suzuki, M. Mori, K. Seto, A. Kai, C. Kawaguchi, M. Suzuki, M. Suematsu, T. Yoneta, S. Miura, H. Ishii, Helicobacter pylori-associated gastric pro- and antioxidant formation in Mongolian gerbils, Free Radic. Biol. Med. 26 (1999) 679–684. [177] D.T. Smoot, T.B. Elliott, H.W. Verspaget, D. Jones, C.R. Allen, K.G. Vernon, T. Bremner, L.C. Kidd, K.S. Kim, J.D. Groupman, H. Ashktorab, Influence of Helicobacter pylori on reactive oxygen-induced gastric epithelial cell injury, Carcinogenesis. 21 (2000) 2091–2095. [178] M. Kimura, S. Goto, Y. Ihara, A. Wada, K. Yahiro, T. Niidome, H. Aoyagi, T. Hirayama, T. Kondo, Impairment of glutathione metabolism in human gastric epithelial cells treated with vacuolating cytotoxin from Helicobacter pylori, Microb.Pathog. 31 (2001) 29–36. doi:10.1006/mpat.2001.0446. [179] H. Shirin, J.T. Pinto, L.U. Liu, M. Merzianu, E.M. Sordillo, S.F. Moss, Helicobacter pylori decreases gastric mucosal glutathione, Cancer Lett. 164 (2001) 127–133. [180] M. Bhattacharjee, S. Bhattacharjee, A. Gupta, R.K. Banerjee, Critical role of an endogenous gastric peroxidase in controlling oxidative damage in H. pylorimediated and nonmediated gastric ulcer, Free Radic. Biol. Med. 32 (2002) 731– 743. [181] I.M. Drake, N.P. Mapstone, C.J. Schorah, K.L. White, D.M. Chalmers, M.F. Dixon, A.T. Axon, Reactive oxygen species activity and lipid peroxidation in

- 91 -

Helicobacter pylori associated gastritis: relation to gastric mucosal ascorbic acid concentrations and effect of H pylori eradication, Gut. 42 (1998) 768–771. [182] E. Buommino, G. Donnarumma, L. Manente, A. De Filippis, F. Silvestri, S. Iaquinto, M.A. Tufano, A. De Luca, The Helicobacter pylori protein HspB interferes with Nrf2/Keap1 pathway altering the antioxidant response of Ags cells, Helicobacter. 17 (2012) 417–425. doi:10.1111/j.1523-5378.2012.00973.x. [183] G. Iaquinto, A. Todisco, N. Giardullo, V. D’Onofrio, L. Pasquale, A. De Luca, A. Andriulli, F. Perri, C. Rega, G. De Chiara, M. Landi, W. Taccone, G. Leandro, N. Figura, Antibody response to Helicobacter pylori CagA and heat-shock proteins in determining the risk of gastric cancer development, Dig Liver Dis. 32 (2000) 378–383. [184] M. Gong, S.S. Ling, S.Y. Lui, K.G. Yeoh, B. Ho, Helicobacter pylori gammaglutamyl transpeptidase is a pathogenic factor in the development of peptic ulcer disease,

Gastroenterology.

139

(2010)

564–573.

doi:10.1053/j.gastro.2010.03.050. [185] M. Mita, M. Satoh, A. Shimada, M. Okajima, S. Azuma, J.S. Suzuki, K. Sakabe, S. Hara, S. Himeno, Metallothionein is a crucial protective factor against Helicobacter pylori-induced gastric erosive lesions in a mouse model, Am.J.Physiol.Gastrointest.Liver

Physiol.

294

(2008)

G877-84.

doi:10.1152/ajpgi.00251.2007. [186] S.-Z. Ding, Y. Minohara, X.J. Fan, J. Wang, V.E. Reyes, J. Patel, B. DirdenKramer, I. Boldogh, P.B. Ernst, S.E. Crowe, Helicobacter pylori infection induces oxidative stress and programmed cell death in human gastric epithelial cells, Infect. Immun. 75 (2007) 4030–4039. doi:10.1128/IAI.00172-07.

- 92 -

[187] M. Calvino-Fernandez, S. Benito-Martinez, T. Parra-Cid, Oxidative stress by Helicobacter pylori causes apoptosis through mitochondrial pathway in gastric epithelial cells, Apoptosis. 13 (2008) 1267–1280. doi:10.1007/s10495-008-02550. [188] S.-Z. Ding, A.M. O’Hara, T.L. Denning, B. Dirden-Kramer, R.C. Mifflin, V.E. Reyes, K.A. Ryan, S.N. Elliott, T. Izumi, I. Boldogh, S. Mitra, P.B. Ernst, S.E. Crowe, Helicobacter pylori and H2O2 increase AP endonuclease-1/redox factor-1 expression in human gastric epithelial cells, Gastroenterology. 127 (2004) 845– 858. [189] G. den Hartog, R. Chattopadhyay, A. Ablack, E.H. Hall, L.D. Butcher, A. Bhattacharyya, L. Eckmann, P.R. Harris, S. Das, P.B. Ernst, S.E. Crowe, Regulation of Rac1 and Reactive Oxygen Species Production in Response to Infection of Gastrointestinal Epithelia, PLoS Pathog. 12 (2016) e1005382. doi:10.1371/journal.ppat.1005382. [190] S. Fu, K.S. Ramanujam, A. Wong, G.T. Fantry, C.B. Drachenberg, S.P. James, S.J. Meltzer, K.T. Wilson, Increased expression and cellular localization of inducible nitric oxide synthase and cyclooxygenase 2 in Helicobacter pylori gastritis, Gastroenterology. 116 (1999) 1319–1329. [191] X. Zhang, B. Ruiz, P. Correa, M.J. Miller, Cellular dissociation of NF-kappaB and inducible nitric oxide synthase in Helicobacter pylori infection, Free Radic. Biol. Med. 29 (2000) 730–735. [192] J.W. Lim, H. Kim, K.H. Kim, NF-kappaB, inducible nitric oxide synthase and apoptosis by Helicobacter pylori infection, Free Radic. Biol. Med. 31 (2001) 355–366.

- 93 -

[193] D.Y. Lu, C.H. Tang, C.H. Chang, M.C. Maa, S.H. Fang, Y.M. Hsu, Y.H. Lin, C.J. Lin, W.C. Lee, H.J. Lin, C.H. Lee, C.H. Lai, Helicobacter pylori attenuates lipopolysaccharide-induced nitric oxide production by murine macrophages, Innate Immun. 18 (2012) 406–417. doi:10.1177/1753425911413164. [194] A.P. Gobert, D.J. McGee, M. Akhtar, G.L. Mendz, J.C. Newton, Y. Cheng, H.L. Mobley, K.T. Wilson, Helicobacter pylori arginase inhibits nitric oxide production

by

eukaryotic

cells:

Proc.Natl.Acad.Sci.U.S.A.

98

a

strategy

for

(2001)

bacterial

survival,

13844–13849.

doi:10.1073/pnas.241443798. [195] K. Nagata, H. Yu, M. Nishikawa, M. Kashiba, A. Nakamura, E.F. Sato, T. Tamura, M. Inoue, Helicobacter pylori generates superoxide radicals and modulates nitric oxide metabolism, J.Biol.Chem. 273 (1998) 14071–14073. [196] A. Elfvin, A. Edebo, P. Hallersund, A. Casselbrant, L. Fandriks, Oxidative and nitrosative stress enzymes in relation to nitrotyrosine in Helicobacter pyloriinfected humans, World J.Gastrointest.Pathophysiol. 5 (2014) 373–379. doi:10.4291/wjgp.v5.i3.373. [197] T. Brzozowski, P.C. Konturek, S.J. Konturek, I. Brzozowska, T. Pawlik, Role of prostaglandins in gastroprotection and gastric adaptation, J.Physiol.Pharmacol. 56 Suppl 5 (2005) 33–55. [198] J.L. Wallace, C.M. Keenan, D.N. Granger, Gastric ulceration induced by nonsteroidal anti-inflammatory drugs is a neutrophil-dependent process, Am.J.Physiol. 259 (1990) G462-7. [199] S.K. Yadav, B. Adhikary, S. Chand, B. Maity, S.K. Bandyopadhyay, S. Chattopadhyay, Molecular mechanism of indomethacin-induced gastropathy,

- 94 -

Free

Radic.Biol.Med.

52

(2012)

1175–1187.

doi:10.1016/j.freeradbiomed.2011.12.023. [200] T. Yoshikawa, Y. Naito, A. Kishi, T. Tomii, T. Kaneko, S. Iinuma, H. Ichikawa, M. Yasuda, S. Takahashi, M. Kondo, Role of active oxygen, lipid peroxidation, and antioxidants in the pathogenesis of gastric mucosal injury induced by indomethacin in rats, Gut. 34 (1993) 732–737. [201] K. Biswas, U. Bandyopadhyay, I. Chattopadhyay, A. Varadaraj, E. Ali, R.K. Banerjee, A novel antioxidant and antiapoptotic role of omeprazole to block gastric ulcer through scavenging of hydroxyl radical, J.Biol.Chem. 278 (2003) 10993–11001. doi:10.1074/jbc.M210328200. [202] I. Chattopadhyay, U. Bandyopadhyay, K. Biswas, P. Maity, R.K. Banerjee, Indomethacin inactivates gastric peroxidase to induce reactive-oxygen-mediated gastric mucosal injury and curcumin protects it by preventing peroxidase inactivation and scavenging reactive oxygen, Free Radic. Biol. Med. 40 (2006) 1397–1408. doi:10.1016/j.freeradbiomed.2005.12.016. [203] P. Maity, S. Bindu, S. Dey, M. Goyal, A. Alam, C. Pal, K. Mitra, U. Bandyopadhyay, Indomethacin, a non-steroidal anti-inflammatory drug, develops gastropathy by inducing reactive oxygen species-mediated mitochondrial pathology and associated apoptosis in gastric mucosa: a novel role of mitochondrial aconitase oxidation, J.Biol.Chem. 284 (2009) 3058–3068. doi:10.1074/jbc.M805329200. [204] A. Tan, H. Nakamura, N. Kondo, M. Tanito, Y.-W. Kwon, M.K. Ahsan, H. Matsui, M. Narita, J. Yodoi, Thioredoxin-1 attenuates indomethacin-induced gastric mucosal injury in mice, Free Radic. Res. 41 (2007) 861–869. doi:10.1080/10715760701199618.

- 95 -

[205] M. Aburaya, K.-I. Tanaka, T. Hoshino, S. Tsutsumi, K. Suzuki, M. Makise, R. Akagi, T. Mizushima, Heme oxygenase-1 protects gastric mucosal cells against non-steroidal anti-inflammatory drugs, J. Biol. Chem. 281 (2006) 33422–33432. doi:10.1074/jbc.M602074200. [206] K. Ueda, T. Ueyama, K. Yoshida, H. Kimura, T. Ito, Y. Shimizu, M. Oka, Y. Tsuruo, M. Ichinose, Adaptive HNE-Nrf2-HO-1 pathway against oxidative stress is associated with acute gastric mucosal lesions, Am.J.Physiol.Gastrointest.Liver Physiol. 295 (2008) G460-9. doi:10.1152/ajpgi.00204.2007. [207] S. Bindu, C. Pal, S. Dey, M. Goyal, A. Alam, M.S. Iqbal, S. Dutta, S. Sarkar, R. Kumar, P. Maity, U. Bandyopadhyay, Translocation of heme oxygenase-1 to mitochondria is a novel cytoprotective mechanism against non-steroidal antiinflammatory drug-induced mitochondrial oxidative stress, apoptosis, and gastric mucosal

injury,

J.Biol.Chem.

286

(2011)

39387–39402.

doi:10.1074/jbc.M111.279893. [208] S. Bindu, S. Mazumder, S. Dey, C. Pal, M. Goyal, A. Alam, M.S. Iqbal, S. Sarkar, A. Azhar Siddiqui, C. Banerjee, U. Bandyopadhyay, Nonsteroidal antiinflammatory drug induces proinflammatory damage in gastric mucosa through NF-kappaB activation and neutrophil infiltration: anti-inflammatory role of heme oxygenase-1 against nonsteroidal anti-inflammatory drug, Free Radic.Biol.Med. 65 (2013) 456–467. doi:10.1016/j.freeradbiomed.2013.07.027. [209] S.K. Yadav, B. Adhikary, S.K. Bandyopadhyay, S. Chattopadhyay, Inhibition of TNF-alpha, and NF-kappaB and JNK pathways accounts for the prophylactic action of the natural phenolic, allylpyrocatechol against indomethacin gastropathy,

Biochim.Biophys.Acta.

doi:10.1016/j.bbagen.2013.03.013.

- 96 -

1830

(2013)

3776–3786.

[210] M.R. Pan, W.C. Hung, Nonsteroidal anti-inflammatory drugs inhibit matrix metalloproteinase-2 via suppression of the ERK/Sp1-mediated transcription, J.Biol.Chem. 277 (2002) 32775–32780. doi:10.1074/jbc.M202334200. [211] S. Swarnakar, K. Ganguly, P. Kundu, A. Banerjee, P. Maity, A.V. Sharma, Curcumin regulates expression and activity of matrix metalloproteinases 9 and 2 during prevention and healing of indomethacin-induced gastric ulcer, J. Biol. Chem. 280 (2005) 9409–9415. doi:10.1074/jbc.M413398200. [212] K. Ganguly, P. Kundu, A. Banerjee, R.J. Reiter, S. Swarnakar, Hydrogen peroxide-mediated downregulation of matrix metalloprotease-2 in indomethacininduced acute gastric ulceration is blocked by melatonin and other antioxidants, Free

Radic.

Biol.

Med.

41

(2006)

911–925.

doi:10.1016/j.freeradbiomed.2006.04.022. [213] K.K. Nelson, J.A. Melendez, Mitochondrial redox control of matrix metalloproteinases,

Free

Radic.Biol.Med.

37

(2004)

768–784.

doi:10.1016/j.freeradbiomed.2004.06.008. [214] W. Eberhardt, M. Schulze, C. Engels, E. Klasmeier, J. Pfeilschifter, Glucocorticoid-mediated

suppression

of

cytokine-induced

matrix

metalloproteinase-9 expression in rat mesangial cells: involvement of nuclear factor-kappaB and Ets transcription factors, Mol.Endocrinol. 16 (2002) 1752– 1766. doi:10.1210/me.2001-0278. [215] P. Correa, J. Houghton, Carcinogenesis of Helicobacter pylori, Gastroenterology. 133 (2007) 659–672. doi:10.1053/j.gastro.2007.06.026. [216] P. Correa, A human model of gastric carcinogenesis, Cancer Res. 48 (1988) 3554–3560.

- 97 -

[217] S.E. Crowe, Helicobacter infection, chronic inflammation, and the development of malignancy, Curr. Opin. Gastroenterol. 21 (2005) 32–38. [218] N. Uemura, S. Okamoto, S. Yamamoto, N. Matsumura, S. Yamaguchi, M. Yamakido, K. Taniyama, N. Sasaki, R.J. Schlemper, Helicobacter pylori infection and the development of gastric cancer, N. Engl. J. Med. 345 (2001) 784–789. doi:10.1056/NEJMoa001999. [219] J. Minoura-Etoh, K. Gotoh, R. Sato, M. Ogata, N. Kaku, T. Fujioka, A. Nishizono, Helicobacter pylori-associated oxidant monochloramine induces reactivation of Epstein-Barr virus (EBV) in gastric epithelial cells latently infected

with

EBV,

J.

Med.

Microbiol.

55

(2006)

905–911.

doi:10.1099/jmm.0.46580-0. [220] J. Parsonnet, G.D. Friedman, N. Orentreich, H. Vogelman, Risk for gastric cancer in people with CagA positive or CagA negative Helicobacter pylori infection, Gut. 40 (1997) 297–301. [221] J.Q. Huang, G.F. Zheng, K. Sumanac, E.J. Irvine, R.H. Hunt, Meta-analysis of the

relationship

between

cagA

seropositivity

and

gastric

cancer,

Gastroenterology. 125 (2003) 1636–1644. [222] K. Ogura, S. Maeda, M. Nakao, T. Watanabe, M. Tada, T. Kyutoku, H. Yoshida, Y. Shiratori, M. Omata, Virulence factors of Helicobacter pylori responsible for gastric diseases in Mongolian gerbil, J. Exp. Med. 192 (2000) 1601–1610. [223] S. Teshima, S. Tsunawaki, K. Rokutan, Helicobacter pylori lipopolysaccharide enhances the expression of NADPH oxidase components in cultured guinea pig gastric mucosal cells, FEBS Lett. 452 (1999) 243–246. [224] E.E. Montalvo-Javé, M. Olguín-Martínez, D.R. Hernández-Espinosa, L. SánchezSevilla, E. Mendieta-Condado, M.L. Contreras-Zentella, L.F. Oñate-Ocaña, T.

- 98 -

Escalante-Tatersfield, A. Echegaray-Donde, J.M. Ruiz-Molina, M.F. Herrera, J. Morán, R. Hernández-Muñoz, Role of NADPH oxidases in inducing a selective increase of oxidant stress and cyclin D1 and checkpoint 1 over-expression during progression to human gastric adenocarcinoma, Eur. J. Cancer. 57 (2016) 50–57. doi:10.1016/j.ejca.2015.11.027. [225] R. Chaturvedi, M. Asim, M.B. Piazuelo, F. Yan, D.P. Barry, J.C. Sierra, A.G. Delgado, S. Hill, R.A. Casero, L.E. Bravo, R.L. Dominguez, P. Correa, D.B. Polk, M.K. Washington, K.L. Rose, K.L. Schey, D.R. Morgan, R.M. Peek, K.T. Wilson, Activation of EGFR and ERBB2 by Helicobacter pylori results in survival of gastric epithelial cells with DNA damage, Gastroenterology. 146 (2014) 1739–1751.e14. doi:10.1053/j.gastro.2014.02.005. [226] S. Futagami, T. Hiratsuka, T. Shindo, A. Horie, T. Hamamoto, K. Suzuki, M. Kusunoki, K. Miyake, K. Gudis, S.E. Crowe, T. Tsukui, C. Sakamoto, Expression of apurinic/apyrimidinic endonuclease-1 (APE-1) in H. pyloriassociated gastritis, gastric adenoma, and gastric cancer, Helicobacter. 13 (2008) 209–218. doi:10.1111/j.1523-5378.2008.00605.x. [227] R. Chattopadhyay, A. Bhattacharyya, S.E. Crowe, Dual regulation by apurinic/apyrimidinic endonuclease-1 inhibits gastric epithelial cell apoptosis during Helicobacter pylori infection, Cancer Res. 70 (2010) 2799–2808. doi:10.1158/0008-5472.CAN-09-4136. [228] A. Bhattacharyya, R. Chattopadhyay, B.R. Burnette, J.V. Cross, S. Mitra, P.B. Ernst, K.K. Bhakat, S.E. Crowe, Acetylation of apurinic/apyrimidinic endonuclease-1 regulates Helicobacter pylori-mediated gastric epithelial cell apoptosis,

Gastroenterology.

doi:10.1053/j.gastro.2009.02.014.

- 99 -

136

(2009)

2258–2269.

[229] A. Bhattacharyya, R. Chattopadhyay, E.H. Hall, S.T. Mebrahtu, P.B. Ernst, S.E. Crowe, Mechanism of hypoxia-inducible factor 1 alpha-mediated Mcl1 regulation

in

Helicobacter

Am.J.Physiol.Gastrointest.Liver

pylori-infected Physiol.

human 299

gastric

epithelium,

(2010)

G1177-86.

doi:10.1152/ajpgi.00372.2010. [230] O. Lefebvre, M.P. Chenard, R. Masson, J. Linares, A. Dierich, M. LeMeur, C. Wendling, C. Tomasetto, P. Chambon, M.C. Rio, Gastric mucosa abnormalities and tumorigenesis in mice lacking the pS2 trefoil protein, Science. 274 (1996) 259–262. [231] D.P. Calnan, B.R. Westley, F.E. May, D.N. Floyd, T. Marchbank, R.J. Playford, The trefoil peptide TFF1 inhibits the growth of the human gastric adenocarcinoma

cell

line

AGS,

J.

Pathol.

188

(1999)

312–317.

doi:10.1002/(SICI)1096-9896(199907)188:3<312::AID-PATH360>3.0.CO;2-P. [232] M. Soutto, A. Belkhiri, M.B. Piazuelo, B.G. Schneider, D. Peng, A. Jiang, M.K. Washington, Y. Kokoye, S.E. Crowe, A. Zaika, P. Correa, R.M. Peek, W. ElRifai, Loss of TFF1 is associated with activation of NF-κB-mediated inflammation and gastric neoplasia in mice and humans, J. Clin. Invest. 121 (2011) 1753–1767. doi:10.1172/JCI43922. [233] A. Jarry, K. Bach-Ngohou, D. Masson, T. Dejoie, P.-A. Lehur, J.-F. Mosnier, M.G. Denis, C.L. Laboisse, Human colonic myocytes are involved in postischemic inflammation through ADAM17-dependent TNFalpha production, Br. J. Pharmacol. 147 (2006) 64–72. doi:10.1038/sj.bjp.0706449. [234] T. Tunc, B. Uysal, C. Atabek, V. Kesik, B. Caliskan, E. Oztas, N. Ersoz, S. Oter, A.

Guven,

Erdosteine

and

ebselen

- 100 -

as

useful

agents

in

intestinal

ischemia/reperfusion

injury,

J.

Surg.

Res.

155

(2009)

210–216.

doi:10.1016/j.jss.2008.06.017. [235] D.A. Parks, G.B. Bulkley, D.N. Granger, S.R. Hamilton, J.M. McCord, Ischemic injury in the cat small intestine: role of superoxide radicals, Gastroenterology. 82 (1982) 9–15. [236] D.A. Parks, D.N. Granger, Contributions of ischemia and reperfusion to mucosal lesion formation, Am. J. Physiol. 250 (1986) G749-753. [237] T. Victoni, F.R. Coelho, A.L. Soares, A. de Freitas, T. Secher, R. Guabiraba, F. Erard, R.M. de Oliveira-Filho, B.B. Vargaftig, G. Lauvaux, M.A. Kamal, B. Ryffel, R. Moser, W. Tavares-de-Lima, Local and remote tissue injury upon intestinal ischemia and reperfusion depends on the TLR/MyD88 signaling pathway, Med. Microbiol. Immunol. 199 (2010) 35–42. doi:10.1007/s00430-0090134-5. [238] P.D.R. Higgins, K.J. Davis, L. Laine, Systematic review: the epidemiology of ischaemic

colitis,

Aliment.

Pharmacol.

Ther.

19

(2004)

729–738.

doi:10.1111/j.1365-2036.2004.01903.x. [239] M.N. Marsh, Clinical and pathological spectrum of coeliac disease, Gut. 34 (1993) 1740; author reply 1741. [240] M. Rewers, G.S. Eisenbarth, Autoimmunity: Celiac disease in T1DM-the need to look long term, Nat Rev Endocrinol. 8 (2012) 7–8. doi:10.1038/nrendo.2011.193. [241] A. Rubio-Tapia, J.F. Ludvigsson, T.L. Brantner, J.A. Murray, J.E. Everhart, The prevalence of celiac disease in the United States, Am. J. Gastroenterol. 107 (2012) 1538-1544, 1545. doi:10.1038/ajg.2012.219. [242] P.H.R. Green, C. Cellier, Celiac disease, N. Engl. J. Med. 357 (2007) 1731–1743. doi:10.1056/NEJMra071600.

- 101 -

[243] S.A. Scanlon, J.A. Murray, Update on celiac disease - etiology, differential diagnosis, drug targets, and management advances, Clin Exp Gastroenterol. 4 (2011) 297–311. doi:10.2147/CEG.S8315. [244] D.A. Leffler, M. Dennis, B. Hyett, E. Kelly, D. Schuppan, C.P. Kelly, Etiologies and predictors of diagnosis in nonresponsive celiac disease, Clin. Gastroenterol. Hepatol. 5 (2007) 445–450. doi:10.1016/j.cgh.2006.12.006. [245] L.M. Sollid, Molecular basis of celiac disease, Annu. Rev. Immunol. 18 (2000) 53–81. doi:10.1146/annurev.immunol.18.1.53. [246] D.A. van Heel, J. West, Recent advances in coeliac disease, Gut. 55 (2006) 1037– 1046. doi:10.1136/gut.2005.075119. [247] W. Vader, D. Stepniak, Y. Kooy, L. Mearin, A. Thompson, J.J. van Rood, L. Spaenij, F. Koning, The HLA-DQ2 gene dose effect in celiac disease is directly related to the magnitude and breadth of gluten-specific T cell responses, Proc. Natl. Acad. Sci. U.S.A. 100 (2003) 12390–12395. doi:10.1073/pnas.2135229100. [248] D.A. van Heel, L. Franke, K.A. Hunt, R. Gwilliam, A. Zhernakova, M. Inouye, M.C. Wapenaar, M.C.N.M. Barnardo, G. Bethel, G.K.T. Holmes, C. Feighery, D. Jewell, D. Kelleher, P. Kumar, S. Travis, J.R.F. Walters, D.S. Sanders, P. Howdle, J. Swift, R.J. Playford, W.M. McLaren, M.L. Mearin, C.J. Mulder, R. McManus, R. McGinnis, L.R. Cardon, P. Deloukas, C. Wijmenga, A genomewide association study for celiac disease identifies risk variants in the region harboring IL2 and IL21, Nat. Genet. 39 (2007) 827–829. doi:10.1038/ng2058. [249] M. Bodd, C.-Y. Kim, K.E.A. Lundin, L.M. Sollid, T-cell response to gluten in patients with HLA-DQ2.2 reveals requirement of peptide-MHC stability in celiac disease,

Gastroenterology.

doi:10.1053/j.gastro.2011.11.021.

- 102 -

142

(2012)

552–561.

[250] M.V. Barone, R. Troncone, S. Auricchio, Gliadin peptides as triggers of the proliferative and stress/innate immune response of the celiac small intestinal mucosa, Int J Mol Sci. 15 (2014) 20518–20537. doi:10.3390/ijms151120518. [251] L. Shan, Ø. Molberg, I. Parrot, F. Hausch, F. Filiz, G.M. Gray, L.M. Sollid, C. Khosla, Structural basis for gluten intolerance in celiac sprue, Science. 297 (2002) 2275–2279. doi:10.1126/science.1074129. [252] R. Ciccocioppo, A. Di Sabatino, G.R. Corazza, The immune recognition of gluten in coeliac disease, Clin. Exp. Immunol. 140 (2005) 408–416. doi:10.1111/j.13652249.2005.02783.x. [253] O. Molberg, S.N. Mcadam, R. Körner, H. Quarsten, C. Kristiansen, L. Madsen, L. Fugger, H. Scott, O. Norén, P. Roepstorff, K.E. Lundin, H. Sjöström, L.M. Sollid, Tissue transglutaminase selectively modifies gliadin peptides that are recognized by gut-derived T cells in celiac disease, Nat. Med. 4 (1998) 713–717. [254] M. Setty, V. Discepolo, V. Abadie, S. Kamhawi, T. Mayassi, A. Kent, C. Ciszewski, M. Maglio, E. Kistner, G. Bhagat, C. Semrad, S.S. Kupfer, P.H. Green, S. Guandalini, R. Troncone, J.A. Murray, J.R. Turner, B. Jabri, Distinct and Synergistic Contributions of Epithelial Stress and Adaptive Immunity to Functions

of

Intraepithelial

Killer Cells

and Active Celiac

Disease,

Gastroenterology. 149 (2015) 681–691.e10. doi:10.1053/j.gastro.2015.05.013. [255] L. Maiuri, C. Ciacci, I. Ricciardelli, L. Vacca, V. Raia, S. Auricchio, J. Picard, M. Osman, S. Quaratino, M. Londei, Association between innate response to gliadin and activation of pathogenic T cells in coeliac disease, Lancet. 362 (2003) 30–37. doi:10.1016/S0140-6736(03)13803-2. [256] R.W. DePaolo, V. Abadie, F. Tang, H. Fehlner-Peach, J.A. Hall, W. Wang, E.V. Marietta, D.D. Kasarda, T.A. Waldmann, J.A. Murray, C. Semrad, S.S. Kupfer,

- 103 -

Y. Belkaid, S. Guandalini, B. Jabri, Co-adjuvant effects of retinoic acid and IL-15 induce inflammatory immunity to dietary antigens, Nature. 471 (2011) 220–224. doi:10.1038/nature09849. [257] I. Caputo, A. Secondo, M. Lepretti, G. Paolella, S. Auricchio, M.V. Barone, C. Esposito, Gliadin peptides induce tissue transglutaminase activation and ERstress through Ca2+ mobilization in Caco-2 cells, PLoS ONE. 7 (2012) e45209. doi:10.1371/journal.pone.0045209. [258] Y.L. Allegretti, C. Bondar, L. Guzman, E. Cueto Rua, N. Chopita, M. Fuertes, N.W. Zwirner, F.G. Chirdo, Broad MICA/B expression in the small bowel mucosa: a link between cellular stress and celiac disease, PLoS ONE. 8 (2013) e73658. doi:10.1371/journal.pone.0073658. [259] B. Jabri, D.D. Kasarda, P.H.R. Green, Innate and adaptive immunity: the yin and yang of celiac disease, Immunol. Rev. 206 (2005) 219–231. doi:10.1111/j.01052896.2005.00294.x. [260] P. Odetti, S. Valentini, I. Aragno, S. Garibaldi, M.A. Pronzato, E. Rolandi, T. Barreca, Oxidative stress in subjects affected by celiac disease, Free Radic. Res. 29 (1998) 17–24. [261] V. Stojiljković, A. Todorović, N. Radlović, S. Pejić, M. Mladenović, J. Kasapović, S.B. Pajović, Antioxidant enzymes, glutathione and lipid peroxidation in peripheral blood of children affected by coeliac disease, Ann. Clin. Biochem. 44 (2007) 537–543. doi:10.1258/000456307782268075. [262] A. Szaflarska-Poplawska, A. Siomek, M. Czerwionka-Szaflarska, D. Gackowski, R. Rózalski, J. Guz, A. Szpila, E. Zarakowska, R. Olinski, Oxidatively damaged DNA/oxidative stress in children with celiac disease, Cancer Epidemiol. Biomarkers Prev. 19 (2010) 1960–1965. doi:10.1158/1055-9965.EPI-10-0295.

- 104 -

[263] K.-P. Zimmer, I. Fischer, T. Mothes, G. Weissen-Plenz, M. Schmitz, H. Wieser, J. Büning, M.M. Lerch, P.C. Ciclitira, P. Weber, H.Y. Naim, Endocytotic segregation of gliadin peptide 31-49 in enterocytes, Gut. 59 (2010) 300–310. doi:10.1136/gut.2008.169656. [264] R. Rivabene, E. Mancini, M. De Vincenzi, In vitro cytotoxic effect of wheat gliadin-derived peptides on the Caco-2 intestinal cell line is associated with intracellular oxidative imbalance: implications for coeliac disease, Biochim. Biophys. Acta. 1453 (1999) 152–160. [265] A. Luciani, V.R. Villella, A. Vasaturo, I. Giardino, M. Pettoello-Mantovani, S. Guido, O.N. Cexus, N. Peake, M. Londei, S. Quaratino, L. Maiuri, Lysosomal accumulation of gliadin p31-43 peptide induces oxidative stress and tissue transglutaminase-mediated PPARgamma downregulation in intestinal epithelial cells and coeliac mucosa, Gut. 59 (2010) 311–319. doi:10.1136/gut.2009.183608. [266] E. Dolfini, L. Elli, T. Dasdia, B. Bufardeci, M.P. Colleoni, B. Costa, I. Floriani, M.L. Falini, N. Guerrieri, F. Forlani, M.T. Bardella, In vitro cytotoxic effect of bread wheat gliadin on the LoVo human adenocarcinoma cell line, Toxicol In Vitro. 16 (2002) 331–337. [267] V. Stojiljković, A. Todorović, S. Pejić, J. Kasapović, Z.S. Saicić, N. Radlović, S.B. Pajović, Antioxidant status and lipid peroxidation in small intestinal mucosa of children with celiac disease, Clin. Biochem. 42 (2009) 1431–1437. doi:10.1016/j.clinbiochem.2009.06.009. [268] A.V. Stazi, B. Trinti, [Selenium deficiency in celiac disease: risk of autoimmune thyroid diseases], Minerva Med. 99 (2008) 643–653. [269] L. Rothem, C. Hartman, A. Dahan, J. Lachter, R. Eliakim, R. Shamir, Paraoxonases are associated with intestinal inflammatory diseases and

- 105 -

intracellularly localized to the endoplasmic reticulum, Free Radic. Biol. Med. 43 (2007) 730–739. doi:10.1016/j.freeradbiomed.2007.05.003. [270] B. Diosdado, E. van Oort, C. Wijmenga, “Coelionomics”: towards understanding the molecular pathology of coeliac disease, Clin. Chem. Lab. Med. 43 (2005) 685–695. doi:10.1515/CCLM.2005.117. [271] A. Di Sabatino, R. Ciccocioppo, S. D’Alò, R. Parroni, D. Millimaggi, M.G. Cifone, G.R. Corazza, Intraepithelial and lamina propria lymphocytes show distinct patterns of apoptosis whereas both populations are active in Fas based cytotoxicity in coeliac disease, Gut. 49 (2001) 380–386. [272] M.C. Maiuri, D. De Stefano, G. Mele, B. Iovine, M.A. Bevilacqua, L. Greco, S. Auricchio, R. Carnuccio, Gliadin increases iNOS gene expression in interferongamma-stimulated RAW 264.7 cells through a mechanism involving NF-kappa B,

Naunyn

Schmiedebergs

Arch.

Pharmacol.

368

(2003)

63–71.

doi:10.1007/s00210-003-0771-y. [273] M.C. Maiuri, D. De Stefano, G. Mele, S. Fecarotta, L. Greco, R. Troncone, R. Carnuccio, Nuclear factor kappa B is activated in small intestinal mucosa of celiac patients, J. Mol. Med. 81 (2003) 373–379. doi:10.1007/s00109-003-04400. [274] V. De Re, M.P. Simula, A. Notarpietro, V. Canzonieri, R. Cannizzaro, G. Toffoli, Do gliadin and tissue transglutaminase mediate PPAR downregulation in intestinal cells of patients with coeliac disease?, Gut. 59 (2010) 1730–1731. doi:10.1136/gut.2010.209395. [275] L. Flohé, R. Brigelius-Flohé, C. Saliou, M.G. Traber, L. Packer, Redox regulation of NF-kappa B activation, Free Radic. Biol. Med. 22 (1997) 1115–1126.

- 106 -

[276] J.C. ter Steege, L. Koster-Kamphuis, E.A. van Straaten, P.P. Forget, W.A. Buurman, Nitrotyrosine in plasma of celiac disease patients as detected by a new sandwich ELISA, Free Radic. Biol. Med. 25 (1998) 953–963. [277] V. Ertekin, M.A. Selimoğlu, Y. Türkan, F. Akçay, Serum nitric oxide levels in children with celiac disease, J. Clin. Gastroenterol. 39 (2005) 782–785. [278] E.A. van Straaten, L. Koster-Kamphuis, I.M. Bovee-Oudenhoven, R. van der Meer, P.P. Forget, Increased urinary nitric oxide oxidation products in children with active coeliac disease, Acta Paediatr. 88 (1999) 528–531. [279] I.A. Murray, I. Daniels, K. Coupland, J.A. Smith, R.G. Long, Increased activity and expression of iNOS in human duodenal enterocytes from patients with celiac disease, Am. J. Physiol. Gastrointest. Liver Physiol. 283 (2002) G319-326. doi:10.1152/ajpgi.00324.2001. [280] I. Daniels, D. Cavill, I.A. Murray, R.G. Long, Elevated expression of iNOS mRNA and protein in coeliac disease, Clin. Chim. Acta. 356 (2005) 134–142. doi:10.1016/j.cccn.2005.01.029. [281] C.G. Beckett, D. Dell’Olio, H.J. Ellis, S. Rosen-Bronson, P.J. Ciclitira, The detection and localization of inducible nitric oxide synthase production in the small intestine of patients with coeliac disease, Eur J Gastroenterol Hepatol. 10 (1998) 641–647. [282] X. Han, M.P. Fink, R.L. Delude, Proinflammatory cytokines cause NO*dependent and -independent changes in expression and localization of tight junction proteins in intestinal epithelial cells, Shock. 19 (2003) 229–237. [283] M. Montalto, L. Cuoco, R. Ricci, N. Maggiano, F.M. Vecchio, G. Gasbarrini, Immunohistochemical analysis of ZO-1 in the duodenal mucosa of patients with untreated and treated celiac disease, Digestion. 65 (2002) 227–233. doi:63817.

- 107 -

[284] R. Ciccocioppo, A. Finamore, C. Ara, A. Di Sabatino, E. Mengheri, G.R. Corazza, Altered expression, localization, and phosphorylation of epithelial junctional proteins in celiac disease, Am. J. Clin. Pathol. 125 (2006) 502–511. doi:10.1309/DTYR-A91G-8R0K-TM8M. [285] C. Abraham, J.H. Cho, Inflammatory bowel disease, N. Engl. J. Med. 361 (2009) 2066–2078. doi:10.1056/NEJMra0804647. [286] I. Ordás, L. Eckmann, M. Talamini, D.C. Baumgart, W.J. Sandborn, Ulcerative colitis, Lancet. 380 (2012) 1606–1619. doi:10.1016/S0140-6736(12)60150-0. [287] R.J. Xavier, D.K. Podolsky, Unravelling the pathogenesis of inflammatory bowel disease, Nature. 448 (2007) 427–434. doi:10.1038/nature06005. [288] D.C. Baumgart, W.J. Sandborn, Crohn’s disease, Lancet. 380 (2012) 1590–1605. doi:10.1016/S0140-6736(12)60026-9. [289] D.P.B. McGovern, A. Gardet, L. Törkvist, P. Goyette, J. Essers, K.D. Taylor, B.M. Neale, R.T.H. Ong, C. Lagacé, C. Li, T. Green, C.R. Stevens, C. Beauchamp, P.R. Fleshner, M. Carlson, M. D’Amato, J. Halfvarson, M.L. Hibberd, M. Lördal, L. Padyukov, A. Andriulli, E. Colombo, A. Latiano, O. Palmieri, E.-J. Bernard, C. Deslandres, D.W. Hommes, D.J. de Jong, P.C. Stokkers, R.K. Weersma, NIDDK IBD Genetics Consortium, Y. Sharma, M.S. Silverberg, J.H. Cho, J. Wu, K. Roeder, S.R. Brant, L.P. Schumm, R.H. Duerr, M.C. Dubinsky, N.L. Glazer, T. Haritunians, A. Ippoliti, G.Y. Melmed, D.S. Siscovick, E.A. Vasiliauskas, S.R. Targan, V. Annese, C. Wijmenga, S. Pettersson, J.I. Rotter, R.J. Xavier, M.J. Daly, J.D. Rioux, M. Seielstad, Genomewide association identifies multiple ulcerative colitis susceptibility loci, Nat. Genet. 42 (2010) 332–337. doi:10.1038/ng.549.

- 108 -

[290] F. Biasi, G. Leonarduzzi, P.I. Oteiza, G. Poli, Inflammatory bowel disease: mechanisms, redox considerations, and therapeutic targets, Antioxid. Redox Signal. 19 (2013) 1711–1747. doi:10.1089/ars.2012.4530. [291] C.A. Anderson, G. Boucher, C.W. Lees, A. Franke, M. D’Amato, K.D. Taylor, J.C. Lee, P. Goyette, M. Imielinski, A. Latiano, C. Lagacé, R. Scott, L. Amininejad, S. Bumpstead, L. Baidoo, R.N. Baldassano, M. Barclay, T.M. Bayless, S. Brand, C. Büning, J.-F. Colombel, L.A. Denson, M. De Vos, M. Dubinsky, C. Edwards, D. Ellinghaus, R.S.N. Fehrmann, J.A.B. Floyd, T. Florin, D. Franchimont, L. Franke, M. Georges, J. Glas, N.L. Glazer, S.L. Guthery, T. Haritunians, N.K. Hayward, J.-P. Hugot, G. Jobin, D. Laukens, I. Lawrance, M. Lémann, A. Levine, C. Libioulle, E. Louis, D.P. McGovern, M. Milla, G.W. Montgomery, K.I. Morley, C. Mowat, A. Ng, W. Newman, R.A. Ophoff, L. Papi, O. Palmieri, L. Peyrin-Biroulet, J. Panés, A. Phillips, N.J. Prescott, D.D. Proctor, R. Roberts, R. Russell, P. Rutgeerts, J. Sanderson, M. Sans, P. Schumm, F. Seibold, Y. Sharma, L.A. Simms, M. Seielstad, A.H. Steinhart, S.R. Targan, L.H. van den Berg, M. Vatn, H. Verspaget, T. Walters, C. Wijmenga, D.C. Wilson, H.J. Westra, R.J. Xavier, Z.Z. Zhao, C.Y. Ponsioen, V. Andersen, L. Torkvist, M. Gazouli, N.P. Anagnou, T.H. Karlsen, L. Kupcinskas, J. Sventoraityte, J.C. Mansfield, S. Kugathasan, M.S. Silverberg, J. Halfvarson, J.I. Rotter, C.G. Mathew, A.M. Griffiths, R. Gearry, T. Ahmad, S.R. Brant, M. Chamaillard, J. Satsangi, J.H. Cho, S. Schreiber, M.J. Daly, J.C. Barrett, M. Parkes, V. Annese, H. Hakonarson, G. Radford-Smith, R.H. Duerr, S. Vermeire, R.K. Weersma, J.D. Rioux, Meta-analysis identifies 29 additional ulcerative colitis risk loci, increasing the number of confirmed associations to 47, Nat. Genet. 43 (2011) 246–252. doi:10.1038/ng.764.

- 109 -

[292] R.K.W. Mailer, A.-L. Joly, S. Liu, S. Elias, J. Tegner, J. Andersson, IL-1β promotes Th17 differentiation by inducing alternative splicing of FOXP3, Sci Rep. 5 (2015) 14674. doi:10.1038/srep14674. [293] G. Rogler, K. Brand, D. Vogl, S. Page, R. Hofmeister, T. Andus, R. Knuechel, P.A. Baeuerle, J. Schölmerich, V. Gross, Nuclear factor kappaB is activated in macrophages and epithelial cells of inflamed intestinal mucosa, Gastroenterology. 115 (1998) 357–369. [294] F. Heller, P. Florian, C. Bojarski, J. Richter, M. Christ, B. Hillenbrand, J. Mankertz, A.H. Gitter, N. Bürgel, M. Fromm, M. Zeitz, I. Fuss, W. Strober, J.D. Schulzke, Interleukin-13 is the key effector Th2 cytokine in ulcerative colitis that affects epithelial tight junctions, apoptosis, and cell restitution, Gastroenterology. 129 (2005) 550–564. doi:10.1016/j.gastro.2005.05.002. [295] B. Sido, V. Hack, A. Hochlehnert, H. Lipps, C. Herfarth, W. Dröge, Impairment of intestinal glutathione synthesis in patients with inflammatory bowel disease, Gut. 42 (1998) 485–492. [296] W. Cao, M.D. Vrees, M.T. Kirber, C. Fiocchi, V.E. Pricolo, Hydrogen peroxide contributes to motor dysfunction in ulcerative colitis, Am. J. Physiol. Gastrointest. Liver Physiol. 286 (2004) G833-843. doi:10.1152/ajpgi.00414.2003. [297] E. Ardite, M. Sans, J. Panés, F.J. Romero, J.M. Piqué, J.C. Fernández-Checa, Replenishment of glutathione levels improves mucosal function in experimental acute colitis, Lab. Invest. 80 (2000) 735–744. [298] C. Loguercio, G. D’Argenio, M. Delle Cave, V. Cosenza, N. Della Valle, G. Mazzacca, C. Del Vecchio Blanco, Glutathione supplementation improves oxidative damage in experimental colitis, Dig Liver Dis. 35 (2003) 635–641.

- 110 -

[299] Y. You, J.-J. Fu, J. Meng, G.-D. Huang, Y.-H. Liu, Effect of N-acetylcysteine on the murine model of colitis induced by dextran sodium sulfate through upregulating

PON1

activity,

Dig.

Dis.

Sci.

54

(2009)

1643–1650.

doi:10.1007/s10620-008-0563-9. [300] F. Biasi, L. Tessitore, D. Zanetti, J.C. Cutrin, B. Zingaro, E. Chiarpotto, N. Zarkovic, G. Serviddio, G. Poli, Associated changes of lipid peroxidation and transforming growth factor beta1 levels in human colon cancer during tumour progression, Gut. 50 (2002) 361–367. [301] F. Biasi, B. Vizio, C. Mascia, E. Gaia, N. Zarkovic, E. Chiarpotto, G. Leonarduzzi, G. Poli, c-Jun N-terminal kinase upregulation as a key event in the proapoptotic interaction between transforming growth factor-beta1 and 4hydroxynonenal in colon mucosa, Free Radic. Biol. Med. 41 (2006) 443–454. doi:10.1016/j.freeradbiomed.2006.04.005. [302] L. Kruidenier, I. Kuiper, C.B.H.W. Lamers, H.W. Verspaget, Intestinal oxidative damage in inflammatory bowel disease: semi-quantification, localization, and association with mucosal antioxidants, J. Pathol. 201 (2003) 28–36. doi:10.1002/path.1409. [303] D. Tardieu, J.P. Jaeg, A. Deloly, D.E. Corpet, J. Cadet, C.R. Petit, The COX-2 inhibitor nimesulide suppresses superoxide and 8-hydroxy-deoxyguanosine formation, and stimulates apoptosis in mucosa during early colonic inflammation in rats, Carcinogenesis. 21 (2000) 973–976. [304] R. D’Incà, R. Cardin, L. Benazzato, I. Angriman, D. Martines, G.C. Sturniolo, Oxidative DNA damage in the mucosa of ulcerative colitis increases with disease duration and dysplasia, Inflamm. Bowel Dis. 10 (2004) 23–27.

- 111 -

[305] X. Ding, Y. Hiraku, N. Ma, T. Kato, K. Saito, M. Nagahama, R. Semba, K. Kuribayashi, S. Kawanishi, Inducible nitric oxide synthase-dependent DNA damage in mouse model of inflammatory bowel disease, Cancer Sci. 96 (2005) 157–163. doi:10.1111/j.1349-7006.2005.00024.x. [306] L.B. Meira, J.M. Bugni, S.L. Green, C.-W. Lee, B. Pang, D. Borenshtein, B.H. Rickman, A.B. Rogers, C.A. Moroski-Erkul, J.L. McFaline, D.B. Schauer, P.C. Dedon, J.G. Fox, L.D. Samson, DNA damage induced by chronic inflammation contributes to colon carcinogenesis in mice, J. Clin. Invest. 118 (2008) 2516– 2525. doi:10.1172/JCI35073. [307] S.P. Hussain, L.J. Hofseth, C.C. Harris, Radical causes of cancer, Nat. Rev. Cancer. 3 (2003) 276–285. doi:10.1038/nrc1046. [308] Y. Naito, T. Takagi, T. Yoshikawa, Molecular fingerprints of neutrophildependent oxidative stress in inflammatory bowel disease, J. Gastroenterol. 42 (2007) 787–798. doi:10.1007/s00535-007-2096-y. [309] T.A.

Ullman,

S.H.

Itzkowitz,

Intestinal

inflammation

and

cancer,

Gastroenterology. 140 (2011) 1807–1816. doi:10.1053/j.gastro.2011.01.057. [310] H. Zhu, Y.R. Li, Oxidative stress and redox signaling mechanisms of inflammatory bowel disease: updated experimental and clinical evidence, Exp. Biol. Med. (Maywood). 237 (2012) 474–480. doi:10.1258/ebm.2011.011358. [311] A. Mangerich, P.C. Dedon, J.G. Fox, S.R. Tannenbaum, G.N. Wogan, Chemistry meets biology in colitis-associated carcinogenesis, Free Radic. Res. 47 (2013) 958–986. doi:10.3109/10715762.2013.832239. [312] I. Moret-Tatay, M. Iborra, E. Cerrillo, L. Tortosa, P. Nos, B. Beltrán, Possible Biomarkers in Blood for Crohn’s Disease: Oxidative Stress and MicroRNAs-

- 112 -

Current Evidences and Further Aspects to Unravel, Oxid Med Cell Longev. 2016 (2016) 2325162. doi:10.1155/2016/2325162. [313] A. Banan, Y. Zhang, J. Losurdo, A. Keshavarzian, Carbonylation and disassembly of the F-actin cytoskeleton in oxidant induced barrier dysfunction and its prevention by epidermal growth factor and transforming growth factor alpha in a human colonic cell line, Gut. 46 (2000) 830–837. [314] A. Banan, G.S. Smith, E.R. Kokoska, T.A. Miller, Role of actin cytoskeleton in prostaglandin-induced protection against ethanol in an intestinal epithelial cell line, J. Surg. Res. 88 (2000) 104–113. doi:10.1006/jsre.1999.5786. [315] A. Keshavarzian, A. Banan, A. Farhadi, S. Komanduri, E. Mutlu, Y. Zhang, J.Z. Fields, Increases in free radicals and cytoskeletal protein oxidation and nitration in the colon of patients with inflammatory bowel disease, Gut. 52 (2003) 720– 728. [316] A. Wang, Å.V. Keita, V. Phan, C.M. McKay, I. Schoultz, J. Lee, M.P. Murphy, M. Fernando, N. Ronaghan, D. Balce, R. Yates, M. Dicay, P.L. Beck, W.K. MacNaughton, J.D. Söderholm, D.M. McKay, Targeting mitochondria-derived reactive oxygen species to reduce epithelial barrier dysfunction and colitis, Am. J. Pathol. 184 (2014) 2516–2527. doi:10.1016/j.ajpath.2014.05.019. [317] C.G. Knutson, A. Mangerich, Y. Zeng, A.R. Raczynski, R.G. Liberman, P. Kang, W. Ye, E.G. Prestwich, K. Lu, J.S. Wishnok, J.R. Korzenik, G.N. Wogan, J.G. Fox, P.C. Dedon, S.R. Tannenbaum, Chemical and cytokine features of innate immunity characterize serum and tissue profiles in inflammatory bowel disease, Proc.

Natl.

Acad.

Sci.

U.S.A.

doi:10.1073/pnas.1222669110.

- 113 -

110

(2013)

E2332-2341.

[318] S. Muthupalani, Z. Ge, Y. Feng, B. Rickman, M. Mobley, A. McCabe, N. Van Rooijen, J.G. Fox, Systemic macrophage depletion inhibits Helicobacter bilisinduced proinflammatory cytokine-mediated typhlocolitis and impairs bacterial colonization dynamics in a BALB/c Rag2-/- mouse model of inflammatory bowel disease, Infect. Immun. 80 (2012) 4388–4397. doi:10.1128/IAI.00530-12. [319] D. Rachmilewitz, J.S. Stamler, F. Karmeli, M.E. Mullins, D.J. Singel, J. Loscalzo, R.J. Xavier, D.K. Podolsky, Peroxynitrite-induced rat colitis--a new model of colonic inflammation, Gastroenterology. 105 (1993) 1681–1688. [320] D.G. Binion, P. Rafiee, K.S. Ramanujam, S. Fu, P.J. Fisher, M.T. Rivera, C.P. Johnson, M.F. Otterson, G.L. Telford, K.T. Wilson, Deficient iNOS in inflammatory bowel disease intestinal microvascular endothelial cells results in increased leukocyte adhesion, Free Radic. Biol. Med. 29 (2000) 881–888. [321] C. Bogdan, Nitric oxide and the immune response, Nat. Immunol. 2 (2001) 907– 916. doi:10.1038/ni1001-907. [322] R.K. Cross, K.T. Wilson, Nitric oxide in inflammatory bowel disease, Inflamm. Bowel Dis. 9 (2003) 179–189. [323] M. Sasaki, S. Bharwani, P. Jordan, J.W. Elrod, M.B. Grisham, T.H. Jackson, D.J. Lefer, J.S. Alexander, Increased disease activity in eNOS-deficient mice in experimental colitis, Free Radic. Biol. Med. 35 (2003) 1679–1687. [324] S. Kawanishi, Y. Hiraku, Oxidative and nitrative DNA damage as biomarker for carcinogenesis with special reference to inflammation, Antioxid. Redox Signal. 8 (2006) 1047–1058. doi:10.1089/ars.2006.8.1047. [325] C.F. Krieglstein, C. Anthoni, W.H. Cerwinka, K.Y. Stokes, J. Russell, M.B. Grisham, D.N. Granger, Role of blood- and tissue-associated inducible nitric-

- 114 -

oxide synthase in colonic inflammation, Am. J. Pathol. 170 (2007) 490–496. doi:10.2353/ajpath.2007.060594. [326] R. Hokari, S. Kato, K. Matsuzaki, M. Kuroki, A. Iwai, A. Kawaguchi, S. Nagao, T. Miyahara, K. Itoh, E. Sekizuka, H. Nagata, H. Ishii, S. Miura, Reduced sensitivity of inducible nitric oxide synthase-deficient mice to chronic colitis, Free Radic. Biol. Med. 31 (2001) 153–163. [327] P.L. Beck, R. Xavier, J. Wong, I. Ezedi, H. Mashimo, A. Mizoguchi, E. Mizoguchi, A.K. Bhan, D.K. Podolsky, Paradoxical roles of different nitric oxide synthase isoforms in colonic injury, Am. J. Physiol. Gastrointest. Liver Physiol. 286 (2004) G137-147. doi:10.1152/ajpgi.00309.2003. [328] X. Tun, K. Yasukawa, K. Yamada, Involvement of nitric oxide with activation of Toll-like receptor 4 signaling in mice with dextran sodium sulfate-induced colitis, Free

Radic.

Biol.

Med.

74

(2014)

108–117.

doi:10.1016/j.freeradbiomed.2014.06.020. [329] I.M. Minderhoud, B. Oldenburg, M.E.I. Schipper, J.J.M. ter Linde, M. Samsom, Serotonin synthesis and uptake in symptomatic patients with Crohn’s disease in remission,

Clin.

Gastroenterol.

Hepatol.

5

(2007)

714–720.

doi:10.1016/j.cgh.2007.02.013. [330] S.C. Regmi, S.-Y. Park, S.K. Ku, J.-A. Kim, Serotonin regulates innate immune responses of colon epithelial cells through Nox2-derived reactive oxygen species, Free

Radic.

Biol.

Med.

69

(2014)

377–389.

doi:10.1016/j.freeradbiomed.2014.02.003. [331] P.A. Ruiz, B. Morón, H.M. Becker, S. Lang, K. Atrott, M.R. Spalinger, M. Scharl, K.A. Wojtal, A. Fischbeck-Terhalle, I. Frey-Wagner, M. Hausmann, T. Kraemer, G. Rogler, Titanium dioxide nanoparticles exacerbate DSS-induced

- 115 -

colitis: role of the NLRP3 inflammasome, Gut. (2016). doi:10.1136/gutjnl-2015310297. [332] S.S. Dhillon, R. Fattouh, A. Elkadri, W. Xu, R. Murchie, T. Walters, C. Guo, D. Mack, H.Q. Huynh, S. Baksh, M.S. Silverberg, A.M. Griffiths, S.B. Snapper, J.H. Brumell, A.M. Muise, Variants in nicotinamide adenine dinucleotide phosphate oxidase complex components determine susceptibility to very early onset inflammatory bowel disease, Gastroenterology. 147 (2014) 680–689.e2. doi:10.1053/j.gastro.2014.06.005. [333] J.D. Rioux, R.J. Xavier, K.D. Taylor, M.S. Silverberg, P. Goyette, A. Huett, T. Green, P. Kuballa, M.M. Barmada, L.W. Datta, Y.Y. Shugart, A.M. Griffiths, S.R. Targan, A.F. Ippoliti, E.-J. Bernard, L. Mei, D.L. Nicolae, M. Regueiro, L.P. Schumm, A.H. Steinhart, J.I. Rotter, R.H. Duerr, J.H. Cho, M.J. Daly, S.R. Brant, Genome-wide association study identifies new susceptibility loci for Crohn disease and implicates autophagy in disease pathogenesis, Nat. Genet. 39 (2007) 596–604. doi:10.1038/ng2032. [334] J. Nair, F. Gansauge, H. Beger, P. Dolara, G. Winde, H. Bartsch, Increased etheno-DNA adducts in affected tissues of patients suffering from Crohn’s disease, ulcerative colitis, and chronic pancreatitis, Antioxid. Redox Signal. 8 (2006) 1003–1010. doi:10.1089/ars.2006.8.1003. [335] H. Kimura, M. Mukaida, K. Kuwabara, T. Ito, K. Hashino, K. Uchida, K. Matsumoto, K. Yoshida, 4-Hydroxynonenal modifies IgA in rat intestine after lipopolysaccharide injection, Free Radic. Biol. Med. 41 (2006) 973–978. doi:10.1016/j.freeradbiomed.2006.06.005. [336] F. Biasi, C. Mascia, M. Astegiano, E. Chiarpotto, M. Nano, B. Vizio, G. Leonarduzzi, G. Poli, Pro-oxidant and proapoptotic effects of cholesterol

- 116 -

oxidation products on human colonic epithelial cells: a potential mechanism of inflammatory bowel disease progression, Free Radic. Biol. Med. 47 (2009) 1731– 1741. doi:10.1016/j.freeradbiomed.2009.09.020. [337] C. Mascia, M. Maina, E. Chiarpotto, G. Leonarduzzi, G. Poli, F. Biasi, Proinflammatory effect of cholesterol and its oxidation products on CaCo-2 human enterocyte-like cells: effective protection by epigallocatechin-3-gallate, Free

Radic.

Biol.

Med.

49

(2010)

2049–2057.

doi:10.1016/j.freeradbiomed.2010.09.033. [338] L. Kruidenier, I. Kuiper, W. van Duijn, S.L. Marklund, R.A. van Hogezand, C.B.H.W. Lamers, H.W. Verspaget, Differential mucosal expression of three superoxide dismutase isoforms in inflammatory bowel disease, J. Pathol. 201 (2003) 7–16. doi:10.1002/path.1407. [339] L. Kruidenier, I. Kuiper, W. Van Duijn, M.A.C. Mieremet-Ooms, R.A. van Hogezand, C.B.H.W. Lamers, H.W. Verspaget, Imbalanced secondary mucosal antioxidant response in inflammatory bowel disease, J. Pathol. 201 (2003) 17–27. doi:10.1002/path.1408. [340] A.A. Te Velde, I. Pronk, F. de Kort, P.C.F. Stokkers, Glutathione peroxidase 2 and aquaporin 8 as new markers for colonic inflammation in experimental colitis and inflammatory bowel diseases: an important role for H2O2?, Eur J Gastroenterol

Hepatol.

20

(2008)

555–560.

doi:10.1097/MEG.0b013e3282f45751. [341] F. Hiller, K. Besselt, S. Deubel, R. Brigelius-Flohé, A.P. Kipp, GPx2 Induction Is Mediated Through STAT Transcription Factors During Acute Colitis, Inflamm. Bowel Dis. 21 (2015) 2078–2089. doi:10.1097/MIB.0000000000000464.

- 117 -

[342] L. Kruidenier, M.E. van Meeteren, I. Kuiper, D. Jaarsma, C.B.H.W. Lamers, F.J. Zijlstra, H.W. Verspaget, Attenuated mild colonic inflammation and improved survival from severe DSS-colitis of transgenic Cu/Zn-SOD mice, Free Radic. Biol. Med. 34 (2003) 753–765. [343] R.S. Esworthy, R. Aranda, M.G. Martín, J.H. Doroshow, S.W. Binder, F.F. Chu, Mice with combined disruption of Gpx1 and Gpx2 genes have colitis, Am. J. Physiol. Gastrointest. Liver Physiol. 281 (2001) G848-855. [344] F.-F. Chu, R.S. Esworthy, P.G. Chu, J.A. Longmate, M.M. Huycke, S. Wilczynski, J.H. Doroshow, Bacteria-induced intestinal cancer in mice with disrupted Gpx1 and Gpx2 genes, Cancer Res. 64 (2004) 962–968. [345] R.S. Esworthy, B.-W. Kim, J. Chow, B. Shen, J.H. Doroshow, F.-F. Chu, Nox1 causes ileocolitis in mice deficient in glutathione peroxidase-1 and -2, Free Radic. Biol. Med. 68 (2014) 315–325. doi:10.1016/j.freeradbiomed.2013.12.018. [346] A. Banning, S. Florian, S. Deubel, S. Thalmann, K. Müller-Schmehl, G. Jacobasch, R. Brigelius-Flohé, GPx2 counteracts PGE2 production by dampening COX-2 and mPGES-1 expression in human colon cancer cells, Antioxid. Redox Signal. 10 (2008) 1491–1500. doi:10.1089/ars.2008.2047. [347] A. Higashikubo, N. Tanaka, N. Noda, I. Maeda, K. Yagi, T. Mizoguchi, H. Nanri, Increase in thioredoxin activity of intestinal epithelial cells mediated by oxidative stress, Biol. Pharm. Bull. 22 (1999) 900–903. [348] Y. Takahashi, H. Masuda, Y. Ishii, Y. Nishida, M. Kobayashi, S. Asai, Decreased expression of thioredoxin interacting protein mRNA in inflamed colonic mucosa in patients with ulcerative colitis, Oncol. Rep. 18 (2007) 531–535. [349] W. Jeong, Y. Jung, H. Kim, S.J. Park, S.G. Rhee, Thioredoxin-related protein 14, a new member of the thioredoxin family with disulfide reductase activity:

- 118 -

implication in the redox regulation of TNF-alpha signaling, Free Radic. Biol. Med. 47 (2009) 1294–1303. doi:10.1016/j.freeradbiomed.2009.07.021. [350] L. Andresen, V.L. Jørgensen, A. Perner, A. Hansen, J. Eugen-Olsen, J. RaskMadsen, Activation of nuclear factor kappaB in colonic mucosa from patients with

collagenous

and

ulcerative

colitis,

Gut.

54

(2005)

503–509.

doi:10.1136/gut.2003.034165. [351] A. Visekruna, T. Joeris, D. Seidel, A. Kroesen, C. Loddenkemper, M. Zeitz, S.H.E. Kaufmann, R. Schmidt-Ullrich, U. Steinhoff, Proteasome-mediated degradation of IkappaBalpha and processing of p105 in Crohn disease and ulcerative colitis, J. Clin. Invest. 116 (2006) 3195–3203. doi:10.1172/JCI28804. [352] W. Shibata, S. Maeda, Y. Hikiba, A. Yanai, T. Ohmae, K. Sakamoto, H. Nakagawa, K. Ogura, M. Omata, Cutting edge: The IkappaB kinase (IKK) inhibitor, NEMO-binding domain peptide, blocks inflammatory injury in murine colitis, J. Immunol. 179 (2007) 2681–2685. [353] W. Qiu, B. Wu, X. Wang, M.E. Buchanan, M.D. Regueiro, D.J. Hartman, R.E. Schoen, J. Yu, L. Zhang, PUMA-mediated intestinal epithelial apoptosis contributes to ulcerative colitis in humans and mice, J. Clin. Invest. 121 (2011) 1722–1732. doi:10.1172/JCI42917. [354] A. Ravi, P. Garg, S.V. Sitaraman, Matrix metalloproteinases in inflammatory bowel disease: boon or a bane?, Inflamm. Bowel Dis. 13 (2007) 97–107. doi:10.1002/ibd.20011. [355] D. Ye, T.Y. Ma, Cellular and molecular mechanisms that mediate basal and tumour necrosis factor-alpha-induced regulation of myosin light chain kinase gene activity, J. Cell. Mol. Med. 12 (2008) 1331–1346. doi:10.1111/j.15824934.2008.00302.x.

- 119 -

[356] Y. Kuwano, K. Tominaga, T. Kawahara, H. Sasaki, K. Takeo, K. Nishida, K. Masuda, T. Kawai, S. Teshima-Kondo, K. Rokutan, Tumor necrosis factor alpha activates transcription of the NADPH oxidase organizer 1 (NOXO1) gene and upregulates superoxide production in colon epithelial cells, Free Radic. Biol. Med. 45 (2008) 1642–1652. doi:10.1016/j.freeradbiomed.2008.08.033. [357] T.O. Khor, M.-T. Huang, K.H. Kwon, J.Y. Chan, B.S. Reddy, A.-N. Kong, Nrf2deficient mice have an increased susceptibility to dextran sulfate sodium-induced colitis, Cancer Res. 66 (2006) 11580–11584. doi:10.1158/0008-5472.CAN-063562. [358] W.O. Osburn, B. Karim, P.M. Dolan, G. Liu, M. Yamamoto, D.L. Huso, T.W. Kensler, Increased colonic inflammatory injury and formation of aberrant crypt foci in Nrf2-deficient mice upon dextran sulfate treatment, Int. J. Cancer. 121 (2007) 1883–1891. doi:10.1002/ijc.22943. [359] T.O. Khor, M.-T. Huang, A. Prawan, Y. Liu, X. Hao, S. Yu, W.K.L. Cheung, J.Y. Chan, B.S. Reddy, C.S. Yang, A.-N. Kong, Increased susceptibility of Nrf2 knockout mice to colitis-associated colorectal cancer, Cancer Prev Res (Phila). 1 (2008) 187–191. doi:10.1158/1940-6207.CAPR-08-0028. [360] I. Stachel, C. Geismann, K. Aden, F. Deisinger, P. Rosenstiel, S. Schreiber, S. Sebens, A. Arlt, H. Schäfer, Modulation of nuclear factor E2-related factor-2 (Nrf2) activation by the stress response gene immediate early response-3 (IER3) in colonic epithelial cells: a novel mechanism of cellular adaption to inflammatory

stress,

J.

Biol.

Chem.

289

(2014)

1917–1929.

doi:10.1074/jbc.M113.490920. [361] J. Marcinkiewicz, M. Ciszek, M. Bobek, M. Strus, P.B. Heczko, M. Kurnyta, R. Biedroń, A. Chmielarczyk, Differential inflammatory mediator response in vitro

- 120 -

from murine macrophages to lactobacilli and pathogenic intestinal bacteria, Int J Exp Pathol. 88 (2007) 155–164. doi:10.1111/j.1365-2613.2007.00530.x. [362] M.M. Huycke, V. Abrams, D.R. Moore, Enterococcus faecalis produces extracellular superoxide and hydrogen peroxide that damages colonic epithelial cell DNA, Carcinogenesis. 23 (2002) 529–536. [363] I.M. Carroll, J.M. Andrus, J.M. Bruno-Bárcena, T.R. Klaenhammer, H.M. Hassan, D.S. Threadgill, Anti-inflammatory properties of Lactobacillus gasseri expressing manganese superoxide dismutase using the interleukin 10-deficient mouse model of colitis, Am. J. Physiol. Gastrointest. Liver Physiol. 293 (2007) G729-738. doi:10.1152/ajpgi.00132.2007. [364] P.W. Lin, L.E.S. Myers, L. Ray, S.-C. Song, T.R. Nasr, A.J. Berardinelli, K. Kundu, N. Murthy, J.M. Hansen, A.S. Neish, Lactobacillus rhamnosus blocks inflammatory signaling in vivo via reactive oxygen species generation, Free Radic.

Biol.

Med.

47

(2009)

1205–1211.

doi:10.1016/j.freeradbiomed.2009.07.033. [365] I. Elatrech, V. Marzaioli, H. Boukemara, O. Bournier, C. Neut, A. DarfeuilleMichaud, J. Luis, L. Dubuquoy, J. El-Benna, P. My-Chan Dang, J.-C. Marie, Escherichia coli LF82 differentially regulates ROS production and mucin expression in intestinal epithelial T84 cells: implication of NOX1, Inflamm. Bowel Dis. 21 (2015) 1018–1026. doi:10.1097/MIB.0000000000000365. [366] S.A. Ballal, P. Veiga, K. Fenn, M. Michaud, J.H. Kim, C.A. Gallini, J.N. Glickman, G. Quéré, P. Garault, C. Béal, M. Derrien, P. Courtin, S. Kulakauskas, M.-P. Chapot-Chartier, J. van Hylckama Vlieg, W.S. Garrett, Host lysozymemediated lysis of Lactococcus lactis facilitates delivery of colitis-attenuating

- 121 -

superoxide dismutase to inflamed colons, Proc. Natl. Acad. Sci. U.S.A. 112 (2015) 7803–7808. doi:10.1073/pnas.1501897112. [367] R. Ravindran, J. Loebbermann, H.I. Nakaya, N. Khan, H. Ma, L. Gama, D.K. Machiah, B. Lawson, P. Hakimpour, Y. Wang, S. Li, P. Sharma, R.J. Kaufman, J. Martinez, B. Pulendran, The amino acid sensor GCN2 controls gut inflammation by inhibiting inflammasome activation, Nature. 531 (2016) 523–527. doi:10.1038/nature17186. [368] J. Ferlay, I. Soerjomataram, R. Dikshit, S. Eser, C. Mathers, M. Rebelo, D.M. Parkin, D. Forman, F. Bray, Cancer incidence and mortality worldwide: sources, methods and major patterns in GLOBOCAN 2012, Int. J. Cancer. 136 (2015) E359-386. doi:10.1002/ijc.29210. [369] F. Radtke, H. Clevers, Self-renewal and cancer of the gut: two sides of a coin, Science. 307 (2005) 1904–1909. doi:10.1126/science.1104815. [370] E.R. Fearon, B. Vogelstein, A genetic model for colorectal tumorigenesis, Cell. 61 (1990) 759–767. [371] C.A. O’Brien, A. Pollett, S. Gallinger, J.E. Dick, A human colon cancer cell capable of initiating tumour growth in immunodeficient mice, Nature. 445 (2007) 106–110. doi:10.1038/nature05372. [372] L. Ricci-Vitiani, D.G. Lombardi, E. Pilozzi, M. Biffoni, M. Todaro, C. Peschle, R. De Maria, Identification and expansion of human colon-cancer-initiating cells, Nature. 445 (2007) 111–115. doi:10.1038/nature05384. [373] D. Wang, R.N. Dubois, The role of COX-2 in intestinal inflammation and colorectal cancer, Oncogene. 29 (2010) 781–788. doi:10.1038/onc.2009.421.

- 122 -

[374] J.D. Lewis, J.J. Deren, G.R. Lichtenstein, Cancer risk in patients with inflammatory bowel disease, Gastroenterol. Clin. North Am. 28 (1999) 459–477, x. [375] A.A. Izzo, M. Camilleri, Emerging role of cannabinoids in gastrointestinal and liver diseases: basic and clinical aspects, Gut. 57 (2008) 1140–1155. doi:10.1136/gut.2008.148791. [376] B.D. Lovasz, L. Lakatos, A. Horvath, I. Szita, T. Pandur, M. Mandel, Z. Vegh, P.A. Golovics, G. Mester, M. Balogh, C. Molnar, E. Komaromi, L.S. Kiss, P.L. Lakatos, Evolution of disease phenotype in adult and pediatric onset Crohn’s disease in a population-based cohort, World J. Gastroenterol. 19 (2013) 2217– 2226. doi:10.3748/wjg.v19.i14.2217. [377] S. Edin, M.L. Wikberg, A.M. Dahlin, J. Rutegård, Å. Öberg, P.-A. Oldenborg, R. Palmqvist, The distribution of macrophages with a M1 or M2 phenotype in relation to prognosis and the molecular characteristics of colorectal cancer, PLoS ONE. 7 (2012) e47045. doi:10.1371/journal.pone.0047045. [378] T. Guina, F. Biasi, S. Calfapietra, M. Nano, G. Poli, Inflammatory and redox reactions in colorectal carcinogenesis, Ann. N. Y. Acad. Sci. 1340 (2015) 95– 103. doi:10.1111/nyas.12734. [379] W.-W. Lin, M. Karin, A cytokine-mediated link between innate immunity, inflammation,

and

cancer,

J.

Clin.

Invest.

117

(2007)

1175–1183.

doi:10.1172/JCI31537. [380] V. De Simone, E. Franzè, G. Ronchetti, A. Colantoni, M.C. Fantini, D. Di Fusco, G.S. Sica, P. Sileri, T.T. MacDonald, F. Pallone, G. Monteleone, C. Stolfi, Th17type cytokines, IL-6 and TNF-α synergistically activate STAT3 and NF-kB to

- 123 -

promote colorectal cancer cell growth, Oncogene. 34 (2015) 3493–3503. doi:10.1038/onc.2014.286. [381] Z. He, X. He, Z. Chen, J. Ke, X. He, R. Yuan, Z. Cai, X. Chen, X. Wu, P. Lan, Activation of the mTORC1 and STAT3 pathways promotes the malignant transformation of colitis in mice, Oncol. Rep. 32 (2014) 1873–1880. doi:10.3892/or.2014.3421. [382] M. Karin, Nuclear factor-kappaB in cancer development and progression, Nature. 441 (2006) 431–436. doi:10.1038/nature04870. [383] S. Wu, K.-J. Rhee, E. Albesiano, S. Rabizadeh, X. Wu, H.-R. Yen, D.L. Huso, F.L. Brancati, E. Wick, F. McAllister, F. Housseau, D.M. Pardoll, C.L. Sears, A human colonic commensal promotes colon tumorigenesis via activation of T helper type 17 T cell responses, Nat. Med. 15 (2009) 1016–1022. doi:10.1038/nm.2015. [384] J.C. Arthur, E. Perez-Chanona, M. Mühlbauer, S. Tomkovich, J.M. Uronis, T.-J. Fan, B.J. Campbell, T. Abujamel, B. Dogan, A.B. Rogers, J.M. Rhodes, A. Stintzi, K.W. Simpson, J.J. Hansen, T.O. Keku, A.A. Fodor, C. Jobin, Intestinal inflammation targets cancer-inducing activity of the microbiota, Science. 338 (2012) 120–123. doi:10.1126/science.1224820. [385] J. Groden, A. Thliveris, W. Samowitz, M. Carlson, L. Gelbert, H. Albertsen, G. Joslyn, J. Stevens, L. Spirio, M. Robertson, Identification and characterization of the familial adenomatous polyposis coli gene, Cell. 66 (1991) 589–600. [386] K.W. Kinzler, M.C. Nilbert, L.K. Su, B. Vogelstein, T.M. Bryan, D.B. Levy, K.J. Smith, A.C. Preisinger, P. Hedge, D. McKechnie, Identification of FAP locus genes from chromosome 5q21, Science. 253 (1991) 661–665.

- 124 -

[387] S.M. Powell, N. Zilz, Y. Beazer-Barclay, T.M. Bryan, S.R. Hamilton, S.N. Thibodeau, B. Vogelstein, K.W. Kinzler, APC mutations occur early during colorectal tumorigenesis, Nature. 359 (1992) 235–237. doi:10.1038/359235a0. [388] E. Chow, C. Thirlwell, F. Macrae, L. Lipton, Colorectal cancer and inherited mutations in base-excision repair, Lancet Oncol. 5 (2004) 600–606. doi:10.1016/S1470-2045(04)01595-5. [389] K.W. Jasperson, T.M. Tuohy, D.W. Neklason, R.W. Burt, Hereditary and familial colon

cancer,

Gastroenterology.

138

(2010)

2044–2058.

doi:10.1053/j.gastro.2010.01.054. [390] R. Fodde, T. Brabletz, Wnt/beta-catenin signaling in cancer stemness and malignant

behavior,

Curr.

Opin.

Cell

Biol.

19

(2007)

150–158.

doi:10.1016/j.ceb.2007.02.007. [391] J.H. van Es, H. Clevers, Notch and Wnt inhibitors as potential new drugs for intestinal

neoplastic disease, Trends

Mol

Med. 11 (2005) 496–502.

doi:10.1016/j.molmed.2005.09.008. [392] S.N. Thibodeau, G. Bren, D. Schaid, Microsatellite instability in cancer of the proximal colon, Science. 260 (1993) 816–819. [393] F.S. Leach, N.C. Nicolaides, N. Papadopoulos, B. Liu, J. Jen, R. Parsons, P. Peltomäki, P. Sistonen, L.A. Aaltonen, M. Nyström-Lahti, Mutations of a mutS homolog in hereditary nonpolyposis colorectal cancer, Cell. 75 (1993) 1215– 1225. [394] C.E. Bronner, S.M. Baker, P.T. Morrison, G. Warren, L.G. Smith, M.K. Lescoe, M. Kane, C. Earabino, J. Lipford, A. Lindblom, Mutation in the DNA mismatch repair gene homologue hMLH1 is associated with hereditary non-polyposis colon cancer, Nature. 368 (1994) 258–261. doi:10.1038/368258a0.

- 125 -

[395] J.R. Howe, S. Roth, J.C. Ringold, R.W. Summers, H.J. Järvinen, P. Sistonen, I.P. Tomlinson, R.S. Houlston, S. Bevan, F.A. Mitros, E.M. Stone, L.A. Aaltonen, Mutations in the SMAD4/DPC4 gene in juvenile polyposis, Science. 280 (1998) 1086–1088. [396] J.R. Howe, J.L. Bair, M.G. Sayed, M.E. Anderson, F.A. Mitros, G.M. Petersen, V.E. Velculescu, G. Traverso, B. Vogelstein, Germline mutations of the gene encoding bone morphogenetic protein receptor 1A in juvenile polyposis, Nat. Genet. 28 (2001) 184–187. doi:10.1038/88919. [397] A.-P.G. Haramis, H. Begthel, M. van den Born, J. van Es, S. Jonkheer, G.J.A. Offerhaus, H. Clevers, De novo crypt formation and juvenile polyposis on BMP inhibition

in

mouse

intestine,

Science.

303

(2004)

1684–1686.

doi:10.1126/science.1093587. [398] B.-G. Kim, C. Li, W. Qiao, M. Mamura, B. Kasprzak, B. Kasperczak, M. Anver, L. Wolfraim, S. Hong, E. Mushinski, M. Potter, S.-J. Kim, X.-Y. Fu, C. Deng, J.J. Letterio, Smad4 signalling in T cells is required for suppression of gastrointestinal cancer, Nature. 441 (2006) 1015–1019. doi:10.1038/nature04846. [399] N.I. Fleming, R.N. Jorissen, D. Mouradov, M. Christie, A. Sakthianandeswaren, M. Palmieri, F. Day, S. Li, C. Tsui, L. Lipton, J. Desai, I.T. Jones, S. McLaughlin, R.L. Ward, N.J. Hawkins, A.R. Ruszkiewicz, J. Moore, H.-J. Zhu, J.M. Mariadason, A.W. Burgess, D. Busam, Q. Zhao, R.L. Strausberg, P. Gibbs, O.M. Sieber, SMAD2, SMAD3 and SMAD4 mutations in colorectal cancer, Cancer Res. 73 (2013) 725–735. doi:10.1158/0008-5472.CAN-12-2706. [400] N. Barker, R.A. Ridgway, J.H. van Es, M. van de Wetering, H. Begthel, M. van den Born, E. Danenberg, A.R. Clarke, O.J. Sansom, H. Clevers, Crypt stem cells

- 126 -

as the cells-of-origin of intestinal cancer, Nature. 457 (2009) 608–611. doi:10.1038/nature07602. [401] C.W. Hendrickse, R.W. Kelly, S. Radley, I.A. Donovan, M.R. Keighley, J.P. Neoptolemos, Lipid peroxidation and prostaglandins in colorectal cancer, Br J Surg. 81 (1994) 1219–1223. [402] H. Wiseman, B. Halliwell, Damage to DNA by reactive oxygen and nitrogen species: role in inflammatory disease and progression to cancer, Biochem. J. 313 ( Pt 1) (1996) 17–29. [403] E. Skrzydlewska, A. Stankiewicz, M. Sulkowska, S. Sulkowski, I. Kasacka, Antioxidant status and lipid peroxidation in colorectal cancer, J. Toxicol. Environ. Health Part A. 64 (2001) 213–222. doi:10.1080/15287390152543690. [404] H.-Y. Yang, K.-O. Chay, J. Kwon, S.-O. Kwon, Y.-K. Park, T.-H. Lee, Comparative proteomic analysis of cysteine oxidation in colorectal cancer patients, Mol. Cells. 35 (2013) 533–542. doi:10.1007/s10059-013-0058-1. [405] S. Mehrabi, L. Wallace, S. Cohen, X. Yao, F.O. Aikhionbare, Differential Measurements of Oxidatively Modified Proteins in Colorectal Adenopolyps, Int J Clin Med. 6 (2015) 288–299. doi:10.4236/ijcm.2015.64037. [406] M. Perše, Oxidative stress in the pathogenesis of colorectal cancer: cause or consequence?, Biomed Res Int. 2013 (2013) 725710. doi:10.1155/2013/725710. [407] S. Kondo, S. Toyokuni, Y. Iwasa, T. Tanaka, H. Onodera, H. Hiai, M. Imamura, Persistent oxidative stress in human colorectal carcinoma, but not in adenoma, Free Radic. Biol. Med. 27 (1999) 401–410. [408] D.-H. Lee, R.S. Esworthy, C. Chu, G.P. Pfeifer, F.-F. Chu, Mutation accumulation in the intestine and colon of mice deficient in two intracellular

- 127 -

glutathione peroxidases, Cancer Res. 66 (2006) 9845–9851. doi:10.1158/00085472.CAN-06-0732. [409] M.R. Oliva, F. Ripoll, P. Muñiz, A. Iradi, R. Trullenque, V. Valls, E. Drehmer, G.T. Sáez, Genetic alterations and oxidative metabolism in sporadic colorectal tumors from a Spanish community, Mol. Carcinog. 18 (1997) 232–243. [410] Y. Murawaki, H. Tsuchiya, T. Kanbe, K. Harada, K. Yashima, K. Nozaka, O. Tanida, M. Kohno, T. Mukoyama, E. Nishimuki, H. Kojo, T. Matsura, K. Takahashi, M. Osaki, H. Ito, J. Yodoi, Y. Murawaki, G. Shiota, Aberrant expression of selenoproteins in the progression of colorectal cancer, Cancer Lett. 259 (2008) 218–230. doi:10.1016/j.canlet.2007.10.019. [411] C.L. Chang, G. Marra, D.P. Chauhan, H.T. Ha, D.K. Chang, L. Ricciardiello, A. Randolph, J.M. Carethers, C.R. Boland, Oxidative stress inactivates the human DNA mismatch repair system, Am. J. Physiol., Cell Physiol. 283 (2002) C148154. doi:10.1152/ajpcell.00422.2001. [412] C.W. Barrett, V.K. Reddy, S.P. Short, A.K. Motley, M.K. Lintel, A.M. Bradley, T. Freeman, J. Vallance, W. Ning, B. Parang, S.V. Poindexter, B. Fingleton, X. Chen, M.K. Washington, K.T. Wilson, N.F. Shroyer, K.E. Hill, R.F. Burk, C.S. Williams, Selenoprotein P influences colitis-induced tumorigenesis by mediating stemness and oxidative damage, J. Clin. Invest. 125 (2015) 2646–2660. doi:10.1172/JCI76099. [413] O.H. Al-Taie, N. Uceyler, U. Eubner, F. Jakob, H. Mörk, M. Scheurlen, R. Brigelius-Flohe, K. Schöttker, J. Abel, A. Thalheimer, T. Katzenberger, B. Illert, R. Melcher, J. Köhrle, Expression profiling and genetic alterations of the selenoproteins GI-GPx and SePP in colorectal carcinogenesis, Nutr Cancer. 48 (2004) 6–14. doi:10.1207/s15327914nc4801_2.

- 128 -

[414] U. Peters, N. Chatterjee, R.B. Hayes, R.E. Schoen, Y. Wang, S.J. Chanock, C.B. Foster, Variation in the selenoenzyme genes and risk of advanced distal colorectal adenoma, Cancer Epidemiol. Biomarkers Prev. 17 (2008) 1144–1154. doi:10.1158/1055-9965.EPI-07-2947. [415] F.-F. Chu, R.S. Esworthy, J.H. Doroshow, Role of Se-dependent glutathione peroxidases in gastrointestinal inflammation and cancer, Free Radic. Biol. Med. 36 (2004) 1481–1495. doi:10.1016/j.freeradbiomed.2004.04.010. [416] R.S. Esworthy, L. Yang, P.H. Frankel, F.-F. Chu, Epithelium-specific glutathione peroxidase, Gpx2, is involved in the prevention of intestinal inflammation in selenium-deficient mice, J. Nutr. 135 (2005) 740–745. [417] S. Krehl, M. Loewinger, S. Florian, A.P. Kipp, A. Banning, L.A. Wessjohann, M.N. Brauer, R. Iori, R.S. Esworthy, F.-F. Chu, R. Brigelius-Flohé, Glutathione peroxidase-2 and selenium decreased inflammation and tumors in a mouse model of inflammation-associated carcinogenesis whereas sulforaphane effects differed with

selenium

supply,

Carcinogenesis.

33

(2012)

620–628.

doi:10.1093/carcin/bgr288. [418] A. Banning, A. Kipp, S. Schmitmeier, M. Löwinger, S. Florian, S. Krehl, S. Thalmann, R. Thierbach, P. Steinberg, R. Brigelius-Flohé, Glutathione Peroxidase 2 Inhibits Cyclooxygenase-2-Mediated Migration and Invasion of HT-29 Adenocarcinoma Cells but Supports Their Growth as Tumors in Nude Mice, Cancer Res. 68 (2008) 9746–9753. doi:10.1158/0008-5472.CAN-08-1321. [419] B. Ahn, H. Ohshima, Suppression of intestinal polyposis in Apc(Min/+) mice by inhibiting nitric oxide production, Cancer Res. 61 (2001) 8357–8360. [420] K. Rokutan, T. Kawahara, Y. Kuwano, K. Tominaga, A. Sekiyama, S. TeshimaKondo, NADPH oxidases in the gastrointestinal tract: a potential role of Nox1 in

- 129 -

innate immune response and carcinogenesis, Antioxid. Redox Signal. 8 (2006) 1573–1582. doi:10.1089/ars.2006.8.1573. [421] J.H. Joo, H. Oh, M. Kim, E.J. An, R.-K. Kim, S.-Y. Lee, D.H. Kang, S.W. Kang, C. Keun Park, H. Kim, S.-J. Lee, D. Lee, J.H. Seol, Y.S. Bae, NADPH Oxidase 1 Activity and ROS Generation Are Regulated by Grb2/Cbl-Mediated Proteasomal Degradation of NoxO1 in Colon Cancer Cells, Cancer Res. 76 (2016) 855–865. doi:10.1158/0008-5472.CAN-15-1512. [422] J.L. Arbiser, J. Petros, R. Klafter, B. Govindajaran, E.R. McLaughlin, L.F. Brown, C. Cohen, M. Moses, S. Kilroy, R.S. Arnold, J.D. Lambeth, Reactive oxygen generated by Nox1 triggers the angiogenic switch, Proc. Natl. Acad. Sci. U.S.A. 99 (2002) 715–720. doi:10.1073/pnas.022630199. [423] S.P. Donald, X.Y. Sun, C.A. Hu, J. Yu, J.M. Mei, D. Valle, J.M. Phang, Proline oxidase, encoded by p53-induced gene-6, catalyzes the generation of prolinedependent reactive oxygen species, Cancer Res. 61 (2001) 1810–1815. [424] Z. Liu, H. Lu, H. Shi, Y. Du, J. Yu, S. Gu, X. Chen, K.J. Liu, C.-A.A. Hu, PUMA overexpression induces reactive oxygen species generation and proteasome-mediated stathmin degradation in colorectal cancer cells, Cancer Res. 65 (2005) 1647–1654. doi:10.1158/0008-5472.CAN-04-1754. [425] K. Izeradjene, L. Douglas, D.M. Tillman, A.B. Delaney, J.A. Houghton, Reactive oxygen species regulate caspase activation in tumor necrosis factor-related apoptosis-inducing ligand-resistant human colon carcinoma cell lines, Cancer Res. 65 (2005) 7436–7445. doi:10.1158/0008-5472.CAN-04-2628. [426] G. Leonarduzzi, A. Scavazza, F. Biasi, E. Chiarpotto, S. Camandola, S. Vogel, R. Dargel, G. Poli, The lipid peroxidation end product 4-hydroxy-2,3-nonenal up-

- 130 -

regulates transforming growth factor beta1 expression in the macrophage lineage: a link between oxidative injury and fibrosclerosis, FASEB J. 11 (1997) 851–857. [427] R. Grau, M.A. Iñiguez, M. Fresno, Inhibition of activator protein 1 activation, vascular endothelial growth factor, and cyclooxygenase-2 expression by 15deoxy-Delta12,14-prostaglandin J2 in colon carcinoma cells: evidence for a redox-sensitive peroxisome proliferator-activated receptor-gamma-independent mechanism, Cancer Res. 64 (2004) 5162–5171. doi:10.1158/0008-5472.CAN-040849. [428] C.E. Eberhart, R.J. Coffey, A. Radhika, F.M. Giardiello, S. Ferrenbach, R.N. DuBois, Up-regulation of cyclooxygenase 2 gene expression in human colorectal adenomas and adenocarcinomas, Gastroenterology. 107 (1994) 1183–1188. [429] K. Subbaramaiah, A.J. Dannenberg, Cyclooxygenase 2: a molecular target for cancer prevention and treatment, Trends Pharmacol. Sci. 24 (2003) 96–102. doi:10.1016/S0165-6147(02)00043-3. [430] M.A. Iñiguez, A. Rodríguez, O.V. Volpert, M. Fresno, J.M. Redondo, Cyclooxygenase-2: a therapeutic target in angiogenesis, Trends Mol Med. 9 (2003) 73–78. [431] A. Arlt, I. Bauer, C. Schafmayer, J. Tepel, S.S. Müerköster, M. Brosch, C. Röder, H. Kalthoff, J. Hampe, M.P. Moyer, U.R. Fölsch, H. Schäfer, Increased proteasome subunit protein expression and proteasome activity in colon cancer relate to an enhanced activation of nuclear factor E2-related factor 2 (Nrf2), Oncogene. 28 (2009) 3983–3996. doi:10.1038/onc.2009.264. [432] T.-H. Kim, E. Hur, S.-J. Kang, J.-A. Kim, D. Thapa, Y.M. Lee, S.K. Ku, Y. Jung, M.-K. Kwak, NRF2 blockade suppresses colon tumor angiogenesis by inhibiting

- 131 -

hypoxia-induced activation of HIF-1α, Cancer Res. 71 (2011) 2260–2275. doi:10.1158/0008-5472.CAN-10-3007. [433] A. Miar, D. Hevia, H. Muñoz-Cimadevilla, A. Astudillo, J. Velasco, R.M. Sainz, J.C. Mayo, Manganese superoxide dismutase (SOD2/MnSOD)/catalase and SOD2/GPx1 ratios as biomarkers for tumor progression and metastasis in prostate, colon, and lung cancer, Free Radic. Biol. Med. 85 (2015) 45–55. doi:10.1016/j.freeradbiomed.2015.04.001. [434] H. Was, J. Dulak, A. Jozkowicz, Heme oxygenase-1 in tumor biology and therapy, Curr Drug Targets. 11 (2010) 1551–1570. [435] B.L. Emmink, J. Laoukili, A.P. Kipp, J. Koster, K.M. Govaert, S. Fatrai, A. Verheem, E.J.A. Steller, R. Brigelius-Flohé, C.R. Jimenez, I.H.M. Borel Rinkes, O. Kranenburg, GPx2 suppression of H2O2 stress links the formation of differentiated tumor mass to metastatic capacity in colorectal cancer, Cancer Res. 74 (2014) 6717–6730. doi:10.1158/0008-5472.CAN-14-1645. [436] M. Darsigny, J.-P. Babeu, E.G. Seidman, F.-P. Gendron, E. Levy, J. Carrier, N. Perreault, F. Boudreau, Hepatocyte nuclear factor-4alpha promotes gut neoplasia in mice and protects against the production of reactive oxygen species, Cancer Res. 70 (2010) 9423–9433. doi:10.1158/0008-5472.CAN-10-1697. [437] D. Papaioannou, K.L. Cooper, C. Carroll, D. Hind, H. Squires, P. Tappenden, R.F. Logan, Antioxidants in the chemoprevention of colorectal cancer and colorectal adenomas in the general population: a systematic review and metaanalysis,

Colorectal

Dis.

13

(2011)

1318.2010.02289.x.

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1085–1099.

doi:10.1111/j.1463-

FIGURE LEGENDS Figure

1.-Schematic

representation

of

the

self-renewing

gastrointestinal

epithelium. A) Small intestine, B) Colon. In the small intestine (A) the crypts of Lieberkühn contain proliferative stem cells and prominences into the lumen called villi with terminally differentiated cells. The colon epithelium (B) lacks villi. The crypts contain two types of stem cells, the radiation-sensitive pluripotent stem cells at position +4 relative to the crypt base (shown in red), and the crypt base columnar stem cells, at position +1, +2, +3 (shown in dark green). The only differentiated cells in the crypts are Paneth cells (shown in yellow), generated at positions 5-7 and then migrate downward to the crypt bottom. Paneth cells are not present in the colon. Intestinal crypt stem cells divide yielding progenitor cells (shown in blue) that gives rise to one of the five types of differentiated epithelial cells: Paneth cells in crypts and enterocytes (shown in white), globet cells (shown in grey), enteroendocrine cells (shown in orange) and tuft cells in villi. These intestinal differentiated cells generated at the crypts -except Paneth cells- mature and differentiate during their upward migration towards the villi. When they reach the top of the villi after three days of their terminal differentiation they suffer spontaneous apoptosis. The Microfold cells (M cells) are shown in clear green. Stem cell self-renewal, cell proliferation, migration and differentiation are controlled by the Wnt, Notch and bone morphogenetic proteins (BMP). The Wnt pathway drives the proliferation of stem cells and progenitor epithelial cells and its activity follows a gradient with its maximum at the crypt bottom. The Notch pathway contributes to

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keep proliferative cells in crypts and also controls differentiation towards either globet cells or enterocytes. BMP signaling is located in the mesenchyme and epithelium of villi but not in the crypts and negatively regulates epithelial proliferation.

Figure 2.-

Redox signaling in the gastrointestinal tract. The three signaling pathways controlling the intestinal epithelium dynamics are Wnt, Notch and BMP. The Wnt and Notch pathways are redox-sensitive as they may be modulated by NOX1-derived hydrogen peroxide. Nucleoredoxin and PTEN mediate NOX1-dependent redox signaling via Wnt pathway, whereas NFB regulates redox signaling through Notch pathway. 3-phosphoinositide dependent protein kinase-1 (PDK1); Adenomatous polyposis coli (APC); Atonal homolog 1 (Math1); Bone morphogenic protein (BMP); BMP receptor (BMPR); Casein kinase 1 (CKI); Dishevelled protein (Dsh); Glycogen synthase kinase-3β (GSK-3β); Hairy/Enhancer of Slipt (Hes); Inositol phosphate (IP); Suppressor of mothers against decapentaplegic (Smad); NADPH oxidase 1 (NOX1); Notch intracellular domain (NICD); Nuclear factor kB (NF-kB); nucleoredoxin (NRX); Phosphatase and tensin homolog deleted on chromosome 10 (PTEN); Phosphatidylinositol 3-kinase (PI3K); T cell factor (TCF); Recombination signal binding protein J-κ (RBPJ-k) (CSL)

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Figure 3.- Redox signaling in Helicobacter pylori-induced ulcer. Helicobacter pylori colonizes the gastric epithelium enhancing ROS production and glutathione depletion. Additional pathogenic factors in H. pylori bacterial strains are VacA, and a pro-apoptotic CagA, and HspB, which may abrogate the antioxidant response regulated by Nrf2/Keap1. H. pylori activates the transcription of Nox1 and its organizer Noxo1 through the Rac1 GTPase in gastric epithelial cells. When Rac1 is activated it translocates to the plasma membrane to assemble with the regulatory subunits NOXO1 and NOXA1 with NOX1 and p22Phox to form the active enzyme complex. In gastric mucosa adaptive anti-apoptotic mechanism may be induced by H. pylori, as it triggers p300-driven acetylation of nuclear APE-1, which forms a repressor complex with HDAC-1 that binds to Bax promoter inhibiting its expression. On the other hand, APE-1 interacts with Rac1 abrogating NOX1 up-regulation and the corresponding hydrogen peroxide generation induced by H. pylori infection. Apurinic/apyrimidinic endonuclease 1 (APE-1); Acetylated Apurinic/apyrimidinic endonuclease 1 (Ac-APE1); BCL2associated X protein (bax); Cytotoxin-associated protein (CagA); Heatshock protein B (HspB); Histone deacetylase 1 (HDAC-1); Kelch-like ECHassociated protein 1 (Keap 1); NADPH oxidase 1 (NOX1); NADPH oxidase activator 1 (NOXA1); NADPH oxidase organizer 1 (NOXO1); Nuclear factor erythroid 2-related factor-2 (Nrf2); Poly [ADP-ribose] polymerase 1 (PARP-1); Ras-related C3 botulinum toxin substrate 1 GTPase (Rac1); Vacuolating cytotoxin A (VacA).

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Figure 4.- Redox signaling in the pathogenesis and progression of inflammatory bowel disease. ROS are involved through different stimuli and mechanisms in the pathogenesis of of inflammatory bowel disease. Macrophages and neutrophils are major sources of superoxide, hydrogen peroxide, and hypochlorous acid during intestinal inflammation. Pathogenic adherentinvasive bacteria enhance superoxide generation via NOX1 and NOXO1 upregulation. Increased protein nitration in inflamed mucosa was associated with neutrophilic MPO. Serotonin levels are increased and enhance intestinal inflammation through NOX2-derived superoxide. High HNE and oxysterols levels cause apoptosis driven by NOX1-derived ROS. MnSOD protein levels and activity and glutathione peroxidase 2 strongly increase in inflamed mucosa restraining the inflammatory response through COX2 and PGE2. NF-κB is over-activated contributing to chronic inflammation, inducing apoptosis through up-regulation of p53 and loss of epithelial barrier via induction of metalloproteinases and MLCK. ROS may also oxidize actin and tubulin, which leads to disruption of cytoskeleton and intestinal barrier dysfunction. Protein starvation stimulates autophagy via GCN2 that leads to low ROS-dependent inflammasome activation. 4hydroxynonenal (HNE); Cyclooxygenase (COX); General controlled nonrepressed 2 kinase (GCN2); Glutathione peroxidase (GPx); Mn superoxide dismutase (MnSOD); Myeloperoxidase (MPO); Myosin light-chain kinase (MLCK);

NADPH

oxidase

(NOX);

Nuclear

factor

kB

(NF-kB);

Prostaglandin E2 (PGE2); Tumor necrosis factor-alpha (TNF-α) Figure 5.- The Janus two-faces of the redox signaling in colorectal cancer. ROS may be either antitumoral as pro-apoptotic via p53/PUMA or TRAIL; or

- 136 -

protumoral when stimulators of cell proliferation via Wnt/Notch, and mutagenic via inactivation of DNA mismatch repair system. In addition, there are anti-tumoral redox-sensitive pathways, such as 15d-PGJ2 signaling, and also pro-tumoral redox sensitive pathways, such as Nrf2, HNE, TIGAR amd HNF4 signaling pathways. 4-hydroxynonenal (HNE); Activator protein 1 (AP-1); Adenomatous polyposis coli (APC); Cyclooxygenase (COX); Cytochrome P450 (CYP); Glutathione peroxidase 2 (GPx2); glutathione S-transferase alpha 4 (GSTA4); glutathione Stransferase kappa 1 (GSTK1); Hepatocyte Nuclear Factor 4 (HNF4); Mn superoxide dismutase (MnSOD); NADPH quinone oxidoreductase-1 (NQO1); NADPH oxidase 1 (NOX1); Nuclear factor erythroid 2-related factor-2 (Nrf2); p53 up-regulated modulator of apoptosis (PUMA); prostanoid 15-deoxy-∆12,14-prostaglandin J2 (15d-PGJ2); selenoprotein P 1 (SEPP1); Tp53-inducible glycolysis and apoptosis regulator (TIGAR); Transforming growth factor-beta (TGF-β); Tumor necrosis factor-related apoptosis-inducing ligand (TRAIL); Vascular endothelial growth factor (VEGF).

Figure 6.- ROS-dependent pathogenic triad in inflammation-induced cancer in the gastrointestinal tract. Superoxide anions and/or hydrogen peroxide would be initiators of the pathogenic triad. Superoxide overproduction may be generated by increased activity of NOX1, NOX2, DUOX or NOX5-S, associated in some cases with oxidative inactivation of superoxide dismutases. Hydrogen peroxide overproduction may be triggered by deficiency in glutathione peroxidases 2 or 7 and/or up-regulation of

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spermine oxidase. The excess of ROS would cause simultaneously two parallel mechanisms within the same cell: one anti-apoptotic mechanism such as APE1 up-regulation and acetylation, COX2 up-regulation, NRF-2 or EGFR activation; and another promoter factor induced by mutation of Wnt, Notch, p53 or K-ras leading to uncontrolled cell proliferation. The Wnt-, Notch-, or K-ras-dependent cell proliferation would be further stimulated by the anti-apoptotic mechanism. Apurinic/apyrimidinic endonuclease 1 (APE1);

Acetylated

Apurinic/apyrimidinic

endonuclease

1

(Ac-APE1);

Cyclooxygenase 2 (COX2); dual oxidase 2 (DUOX2); Epidermal growth factor receptor (EGFR); Glutathione peroxidase (GPx); Superoxide dismutase (SOD); NADPH oxidase (NOX); Nuclear factor erythroid 2related factor-2 (Nrf2)

Table 1.- Redox signaling in the intestine by Lactobacillus spp.

Pathway

ROS source

Effect

Reference

FPR1 stimulation

NOX1

Pro-proliferative and pro-

Alam et al. (2014)

migratory effects on human

[101]

intestinal epithelial cell line SKCO15 and repair of mucosal wounds in murine colon

Swanson et al. SHP-2 and LMW-PTP

NOX2

Increase phosphorylation of (2011) focal adhesion kinase (FAK) and

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inhibition

regulate cell migration in Caco-

[120]

2, T-84, and HeLa cells

Wentworth et al. DUSP3 inhibition/

FPR-

Cell growth, differentiation and

ERK activation

dependent

proliferation in human

(2011)

intestinal epithelial cell line SK-

[121]

CO15

Kumar et al. Oxidative

NOXs

Anti-inflammatory mechanism (2007)

inactivation of Ubc12

by NF-κB inhibition pathway in [122]

human (Caco-2, HeLa, T-84) epithelial cells

Jones et al. Nrf2-dependent

NOX1

Cytoprotective against external (2015)

cytoprotective genes

injury in murine colon

transcription

[127]

Table 2.- Redox signaling triggered by microbiota in the intestinal disorders

Bacterial spp.

Pathway

Effect

Lactobacillus spp.

Increased ROS

Anti-inflammatory

production in

cytokine synthesis in

murine

inflammatory bowel

Reference

Marcinkiewicz et al. (2007)

peritoneal

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[360]

macrophages

disease

ROS inhibition

Pro-inflammatory

in murine

cytokines synthesis in

peritoneal

inflammatory bowel

macrophages

disease

Increased of

DNA damage in

H2O2

inflammatory bowel

production in

disease

Marcinkiewicz et al. Escherichia coli

(2007)

[360]

Huycke et al. (2002) Enterococcus faecalis

[361]

CHO and HT-29 cells

Carrol et al. (2007) Lactobacillus gasseri

Increased

Anti-inflammatory

expression of

activity that reduces

MnSOD in IL-

the severity of colitis

[362]

10-deficient mouse model

Lin et al. (2009) Lactobacillus rhamnosus

ROS generation

NF-κB inhibition by

in human fetal

blocking neddylation

intestinal

of cullin-1

[363]

epithelial cells

Elatrech et al.(2015) Escherichia coli LF82

Increased

Inhibition of mucin

expression of

gene expression and

- 140 -

NOX1 and

increased expression

NOXO1 in

of IL-8 in

cultured

inflammatory bowel

epithelial T84

disease

[364]

cells

Wu et al. (2009) Bacteroides fragilis

Induction of

Colonic hyperplasia

STAT3 and Th17

and tumor formation

in colon of

in colorectal cancer

[382]

multiple intestinal neoplasia (Min) mice

Table 3.- Redox signaling pathway in inflammatory bowel disease

Initiator

Pathway

Effect

- 141 -

Reference

Biasi et al. (2006) HNE

JNK and SMAD4

Anti-proliferative and

activation

pro-apoptotic effect in

[300]

CaCo-2 human colon adenocarcinoma cells

Serotonin

NOX2 activation

Cytokine synthesis by

Regmi et al. (2014) [329]

activation of p38, ERK, CREB and NF-κB in colon epithelial cells (CCD 841, HT-29, Caco-2)

IL-1β cleavage and IL-8 TiO2

Ruíz et al. (2016)

NLRP3 inflammasome synthesis activation

[330] in THP-1, CaCo-2 and HT29 intestinal epithelial cells

Biasi et al. (2009) Oxysterols

NOX1 activation

Up-regulation of the mitochondrial pathway

[335]

of programmed death of differentiated CaCo-2 cells

Esworthy et al. (2014) GPx1 and GPx2

NOX1 activation

Pro-apoptotic and proinflammatory activity by

- 142 -

[344]

deficiency

TNF-α synthesis in GPx1/2-Nox1-triple KO mice

Quiu et al. (2011) NF-κB

PUMA induction

Apoptotic pathway through the TNF-α/NF-

[352]

κB/PUMA axis in murine and human colon

Stachel et al. (2014) IER3

NRF2 activation

deficiency

Tumor progression by Nrf2-dependent

[359]

resistance to death ligands and anticancer drugs in human NCM460 colonocytes

Ravindran et al. (2016) GCN2

Autophagy inhibition

deficiency

NLRP3 inflammasome induction and Th17 response in intestinal antigen presenting cells (APCs) and epithelial cells

HIGHLIGHTS

- 143 -

[366]



NOX1-derived hydrogen peroxide regulates intestinal epithelial homeostasis by Wnt and Notch signaling



O2-. overproduction and SOD inactivation contribute to Barrett’s esophagus and esophageal adenocarcinoma



In H. pylori-induced peptic ulcer, oxidative stress aggravates mucosal damage



Progression of inflammatory bowel disease relies on redox-sensitive NLRP3, NF-κB, Mn-SOD and GPx2



In colorectal cancer, redox signaling exhibits two Janus faces: proliferation vs apoptosis

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- 145 -

- 146 -

- 147 -

- 148 -

- 149 -