The intestinal epithelium serves the unique and critical function of harvesting dietary nutrients, while simultaneously acting as a cellular barrier separating tissues from the luminal environment and gut microbial ecosystem. Two salient features of the intestinal epithelium enable it to perform these complex functions. First, cells within the intestinal epithelium achieve a wide range of specialized identities, including different cell types and distinct anterior–posterior patterning along the intestine. Second, intestinal epithelial cells are sensitive and responsive to the dynamic milieu of dietary nutrients, xenobiotics and microorganisms encountered in the intestinal luminal environment. These diverse identities and responsiveness of intestinal epithelial cells are achieved in part through the differential transcription of genes encoded in their shared genome. Here, we review insights from mice and other vertebrate models into the transcriptional regulatory mechanisms underlying intestinal epithelial identity and microbial responsiveness, including DNA methylation, chromatin accessibility, histone modifications and transcription factors. These studies are revealing that most transcription factors involved in intestinal epithelial identity also respond to changes in the microbiota, raising both opportunities and challenges to discern the underlying integrative transcriptional regulatory networks.
Regional and cell identities in the intestinal epithelium of vertebrates are patterned through interactions between changes in the chromatin landscape and transcription factors.
DNA methylation and accessible chromatin in intestinal epithelial cells are relatively stable in response to the gut microbiota.
Histone modifications and transcription factor activity respond dynamically to microbial colonization.
Transcription factors often have dual roles, to various degrees, in specifying intestinal epithelial identity and microbial responsiveness.
Nuclear receptors seem to be key mediators of intestinal epithelial responses to the gut microbiota.
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Haber, A. L. et al. A single-cell survey of the small intestinal epithelium. Nature 551, 333–339 (2017). This single-cell analysis shows the full diversity of cell types in the small intestine, providing a framework resource for future cell-type specific studies of the small-intestinal epithelium, and provides an online portal tool for exploration.
Moor, A. E. et al. Spatial reconstruction of single enterocytes uncovers broad zonation along the intestinal villus axis. Cell 175, 1156–1167.e15 (2018). This paper reconstructs the pattern of gene expression along the crypt–villus axis using scRNA-seq with unprecedented resolution.
Larsson, E. et al. Analysis of gut microbial regulation of host gene expression along the length of the gut and regulation of gut microbial ecology through MyD88. Gut 61, 1124–1131 (2012).
El Aidy, S. et al. Temporal and spatial interplay of microbiota and intestinal mucosa drive establishment of immune homeostasis in conventionalized mice. Mucosal Immunol 5, 567–579 (2012). This paper highlights the temporal and spatial dynamics of genome-wide gene expression in the intestine following microbiota colonization of germ-free mice.
Smale, S. T. Selective transcription in response to an inflammatory stimulus. Cell 140, 833–844 (2010).
Howitt, M. R. et al. Tuft cells, taste-chemosensory cells, orchestrate parasite type 2 immunity in the gut. Science 351, 1329–1333 (2016).
Thompson, C. A., DeLaForest, A. & Battle, M. A. Patterning the gastrointestinal epithelium to confer regional-specific functions. Dev. Biol. 435, 97–108 (2018). This is a fundamental review on the development and patterning of the gatrointestinal tract.
Stevens, C. E. & Hume, I. D. Comparative Physiology of the Vertebrate Digestive System 2nd edn (Cambridge University Press, 2004).
Wallace, K. N., Akhter, S., Smith, E. M., Lorent, K. & Pack, M. Intestinal growth and differentiation in zebrafish. Mech. Dev. 122, 157–173 (2005).
Ng, A. N. et al. Formation of the digestive system in zebrafish: III. Intestinal epithelium morphogenesis. Dev. Biol. 286, 114–135 (2005).
Aghaallaei, N. et al. Identification, visualization and clonal analysis of intestinal stem cells in fish. Development 143, 3470–3480 (2016).
Lickwar, C. R. et al. Genomic dissection of conserved transcriptional regulation in intestinal epithelial cells. PLoS Biol. 15, e2002054 (2017).
Camp, J. G. et al. Microbiota modulate transcription in the intestinal epithelium without remodeling the accessible chromatin landscape. Genome Res. 24, 1504–1516 (2014). This paper demonstrates that microbial colonization did not lead to major changes in chromatin accessibility and that transcription factor binding, including by nuclear receptors, is likely to mediate much of the transcriptional response to the microbiota.
Graham, D. B. & Xavier, R. J. Pathway paradigms revealed from the genetics of inflammatory bowel disease. Nature 578, 527–539 (2020).
Gordon, J. I. & Hermiston, M. L. Differentiation and self-renewal in the mouse gastrointestinal epithelium. Curr. Opin. Cell Biol. 6, 795–803 (1994).
Moorefield, E. C. et al. Aging effects on intestinal homeostasis associated with expansion and dysfunction of intestinal epithelial stem cells. Aging 9, 1898–1915 (2017).
Williams, J. M. et al. Epithelial cell shedding and barrier function: a matter of life and death at the small intestinal villus tip. Vet. Pathol. 52, 445–455 (2015).
Uribe, A., Alam, M., Midtvedt, T., Smedfors, B. & Theodorsson, E. Endogenous prostaglandins and microflora modulate DNA synthesis and neuroendocrine peptides in the rat gastrointestinal tract. Scand. J. Gastroenterol. 32, 691–699 (1997).
Rawls, J. F., Samuel, B. S. & Gordon, J. I. Gnotobiotic zebrafish reveal evolutionarily conserved responses to the gut microbiota. Proc. Natl Acad. Sci. USA 101, 4596–4601 (2004).
Cheesman, S. E., Neal, J. T., Mittge, E., Seredick, B. M. & Guillemin, K. Epithelial cell proliferation in the developing zebrafish intestine is regulated by the Wnt pathway and microbial signaling via Myd88. Proc. Natl Acad. Sci. USA 108 (Suppl 1), 4570–4577 (2011).
Gehart, H. & Clevers, H. Tales from the crypt: new insights into intestinal stem cells. Nat. Rev. Gastroenterol. Hepatol. 16, 19–34 (2019). This paper is a current and thorough review of the intestinal stem cell field.
Tan, D. W. & Barker, N. Intestinal stem cells and their defining niche. Curr. Top. Dev. Biol. 107, 77–107 (2014).
Sato, T. et al. Long-term expansion of epithelial organoids from human colon, adenoma, adenocarcinoma, and Barrett’s epithelium. Gastroenterology 141, 1762–1772 (2011).
Zhang, Z. & Liu, Z. Paneth cells: the hub for sensing and regulating intestinal flora. Sci. China Life Sci. 59, 463–467 (2016).
Korinek, V. et al. Depletion of epithelial stem-cell compartments in the small intestine of mice lacking Tcf-4. Nat. Genet. 19, 379–383 (1998).
van der Flier, L. G. et al. Transcription factor achaete scute-like 2 controls intestinal stem cell fate. Cell 136, 903–912 (2009).
Schuijers, J. et al. Ascl2 acts as an R-spondin/Wnt-responsive switch to control stemness in intestinal crypts. Cell Stem Cell 16, 158–170 (2015).
Bankaitis, E. D., Ha, A., Kuo, C. J. & Magness, S. T. Reserve stem cells in intestinal homeostasis and injury. Gastroenterology 155, 1348–1361 (2018).
Snippert, H. J. et al. Intestinal crypt homeostasis results from neutral competition between symmetrically dividing Lgr5 stem cells. Cell 143, 134–144 (2010).
Yang, Q., Bermingham, N. A., Finegold, M. J. & Zoghbi, H. Y. Requirement of Math1 for secretory cell lineage commitment in the mouse intestine. Science 294, 2155–2158 (2001).
Shroyer, N. F. et al. Intestine-specific ablation of mouse atonal homolog 1 (Math1) reveals a role in cellular homeostasis. Gastroenterology 132, 2478–2488 (2007).
Zwick, R. K., Ohlstein, B. & Klein, O. D. Intestinal renewal across the animal kingdom: comparing stem cell activity in mouse and Drosophila. Am. J. Physiol. Gastrointest. Liver Physiol. 316, G313–G322 (2018).
Peterson, K. J., Cotton, J. A., Gehling, J. G. & Pisani, D. The Ediacaran emergence of bilaterians: congruence between the genetic and the geological fossil records. Philos. Trans. R. Soc. Lond. B Biol. Sci. 363, 1435–1443 (2008).
Dempsey, P. J., Bohin, N. & Samuelson, L. C. in Physiology of the Gastrointestinal Tract 6th edn (ed Said, H. M.) 141-183 (Academic Press, 2018).
Sancho, R., Cremona, C. A. & Behrens, A. Stem cell and progenitor fate in the mammalian intestine: Notch and lateral inhibition in homeostasis and disease. EMBO Rep. 16, 571–581 (2015).
Stamataki, D. et al. Delta1 expression, cell cycle exit, and commitment to a specific secretory fate coincide within a few hours in the mouse intestinal stem cell system. PLoS ONE 6, e24484 (2011).
Gregorieff, A. et al. The ets-domain transcription factor Spdef promotes maturation of goblet and paneth cells in the intestinal epithelium. Gastroenterology 137, 1333–1345.e1–3 (2009).
Lopez-Diaz, L. et al. Intestinal Neurogenin 3 directs differentiation of a bipotential secretory progenitor to endocrine cell rather than goblet cell fate. Dev. Biol. 309, 298–305 (2007).
Gerbe, F. et al. Intestinal epithelial tuft cells initiate type 2 mucosal immunity to helminth parasites. Nature 529, 226–230 (2016).
Kosinski, C. et al. Gene expression patterns of human colon tops and basal crypts and BMP antagonists as intestinal stem cell niche factors. Proc. Natl Acad. Sci. USA 104, 15418–15423 (2007).
Auclair, B. A., Benoit, Y. D., Rivard, N., Mishina, Y. & Perreault, N. Bone morphogenetic protein signaling is essential for terminal differentiation of the intestinal secretory cell lineage. Gastroenterology 133, 887–896 (2007).
Beumer, J. et al. Enteroendocrine cells switch hormone expression along the crypt-to-villus BMP signalling gradient. Nat. Cell Biol. 20, 909–916 (2018).
Chen, L. et al. A reinforcing HNF4-SMAD4 feed-forward module stabilizes enterocyte identity. Nat. Genet. 51, 777–785 (2019).
Gilmore, A. P. Anoikis. Cell Death Differ. 12 (Suppl 2), 1473–1477 (2005).
Banerjee, K. K. et al. Enhancer, transcriptional, and cell fate plasticity precedes intestinal determination during endoderm development. Genes Dev. 32, 1430–1442 (2018). This paper defines developmentally important enhancers and defines a developmental window during which Cdx2 activity is critical for establishing regional identities of the intestinal epithelium.
Kleinman, R. E. & Walker, W. A. Antigen processing and uptake from the intestinal tract. Clin. Rev. Allergy 2, 25–37 (1984).
Park, J. et al. Lysosome-rich enterocytes mediate protein absorption in the vertebrate gut. Dev. Cell 51, 7–20.e6 (2019). This paper describes an often over-looked cell type common in the gastrointestinal tract of vertebrates, but often restricted to early mammalian developmental stages.
Langille, R. M. & Youson, J. H. Morphology of the intestine of prefeeding and feeding adult lampreys, Petromyzon marinus (L): the mucosa of the posterior intestine and hindgut. J. Morphol. 182, 137–152 (1984).
van den Bogert, B., Leimena, M. M., de Vos, W. M., Zoetendal, E. G. & Kleerebezem, M. in Handbook of Molecular Microbial Ecology II: Metagenomics in Different Habitats (ed de Bruijn, F. J.) 175–190 (Wiley, 2011).
Hu, B. et al. Transgenic overexpression of cdx1b induces metaplastic changes of gene expression in zebrafish esophageal squamous epithelium. Zebrafish 10, 218–227 (2013).
Boyd, M. et al. Characterization of the enhancer and promoter landscape of inflammatory bowel disease from human colon biopsies. Nat. Commun. 9, 1661 (2018). This paper identifies non-coding promoter and enhancer regions, and the SNPs and transcription factor binding sites they contain, that are common and distinct among patients with ulcerative colitis, Crohn’s disease and controls.
Weiser, M. et al. Molecular classification of Crohn’s disease reveals two clinically relevant subtypes. Gut 67, 36–42 (2018). This paper finds two distinct transcriptional patterns in patients with Crohn’s disease that suggest molecular subtyping based on region-specific similarities to either normal ileal or colonic tissue.
Middendorp, S. et al. Adult stem cells in the small intestine are intrinsically programmed with their location-specific function. Stem Cell 32, 1083–1091 (2014).
Que, J. et al. Multiple dose-dependent roles for Sox2 in the patterning and differentiation of anterior foregut endoderm. Development 134, 2521–2531 (2007).
Beck, F., Erler, T., Russell, A. & James, R. Expression of Cdx-2 in the mouse embryo and placenta: possible role in patterning of the extra-embryonic membranes. Dev. Dyn. 204, 219–227 (1995).
James, R., Erler, T. & Kazenwadel, J. Structure of the murine homeobox gene cdx-2. Expression in embryonic and adult intestinal epithelium. J. Biol. Chem. 269, 15229–15237 (1994).
Chawengsaksophak, K., James, R., Hammond, V. E., Kontgen, F. & Beck, F. Homeosis and intestinal tumours in Cdx2 mutant mice. Nature 386, 84–87 (1997).
Gao, N., White, P. & Kaestner, K. H. Establishment of intestinal identity and epithelial-mesenchymal signaling by Cdx2. Dev. Cell 16, 588–599 (2009).
Grainger, S., Savory, J. G. & Lohnes, D. Cdx2 regulates patterning of the intestinal epithelium. Dev. Biol. 339, 155–165 (2010).
Stringer, E. J. et al. Cdx2 determines the fate of postnatal intestinal endoderm. Development 139, 465–474 (2012).
Mutoh, H. et al. Conversion of gastric mucosa to intestinal metaplasia in Cdx2-expressing transgenic mice. Biochem. Biophys. Res. Commun. 294, 470–479 (2002).
Silberg, D. G. et al. Cdx2 ectopic expression induces gastric intestinal metaplasia in transgenic mice. Gastroenterology 122, 689–696 (2002). This paper describes the dominant role of Cdx2 in specifying intestinal fate along the gastrointestinal tract in mice.
Jiang, M. et al. Transitional basal cells at the squamous-columnar junction generate Barrett’s oesophagus. Nature 550, 529–533 (2017).
Bosse, T. et al. Gata4 is essential for the maintenance of jejunal-ileal identities in the adult mouse small intestine. Mol. Cell. Biol. 26, 9060–9070 (2006).
Battle, M. A. et al. GATA4 is essential for jejunal function in mice. Gastroenterology 135, 1676–1686.e1 (2008).
Thompson, C. A. et al. GATA4 is sufficient to establish jejunal versus ileal identity in the small intestine. Cell Mol. Gastroenterol. Hepatol. 3, 422–446 (2017).
Walker, E. M., Thompson, C. A. & Battle, M. A. GATA4 and GATA6 regulate intestinal epithelial cytodifferentiation during development. Dev. Biol. 392, 283–294 (2014).
Lindeboom, R. G. et al. Integrative multi-omics analysis of intestinal organoid differentiation. Mol. Syst. Biol. 14, e8227 (2018).
Wilson, J. M. & Castro, L. F. C. in Fish Physiology Vol. 30 (eds Grosell, M., Farrell, A. P. & Brauner, C. J.) 1–55 (Academic Press, 2010).
Castro, L. F. et al. Recurrent gene loss correlates with the evolution of stomach phenotypes in gnathostome history. Proc. Biol. Sci. 281, 20132669 (2014).
Muncan, V. et al. T-cell factor 4 (Tcf7l2) maintains proliferative compartments in zebrafish intestine. EMBO Rep. 8, 966–973 (2007).
Creyghton, M. P. et al. Histone H3K27ac separates active from poised enhancers and predicts developmental state. Proc. Natl Acad. Sci. USA 107, 21931–21936 (2010).
Edwards, J. R., Yarychkivska, O., Boulard, M. & Bestor, T. H. DNA methylation and DNA methyltransferases. Epigenetics Chromatin 10, 23 (2017).
Sheaffer, K. L. et al. DNA methylation is required for the control of stem cell differentiation in the small intestine. Genes Dev. 28, 652–664 (2014). This is the first study to show that DNA methylation machinery is required for control of differentiation and proliferative processes along the crypt–villus axis of the mouse intestine, and reveals transcription factor binding sites that are enriched in differentially methylated regions in Dnmt1 mutants.
Elliott, E. N., Sheaffer, K. L., Schug, J., Stappenbeck, T. S. & Kaestner, K. H. Dnmt1 is essential to maintain progenitors in the perinatal intestinal epithelium. Development 142, 2163–2172 (2015).
Marjoram, L. et al. Epigenetic control of intestinal barrier function and inflammation in zebrafish. Proc. Natl Acad. Sci. USA 112, 2770–2775 (2015).
Yu, D. H. et al. Postnatal epigenetic regulation of intestinal stem cells requires DNA methylation and is guided by the microbiome. Genome Biol. 16, 211 (2015).
Kaaij, L. T. et al. DNA methylation dynamics during intestinal stem cell differentiation reveals enhancers driving gene expression in the villus. Genome Biol. 14, R50 (2013).
Kraiczy, J. et al. Assessing DNA methylation in the developing human intestinal epithelium: potential link to inflammatory bowel disease. Mucosal Immunol. 9, 647–658 (2016).
Howell, K. J. et al. DNA methylation and transcription patterns in intestinal epithelial cells from pediatric patients with inflammatory bowel diseases differentiate disease subtypes and associate with outcome. Gastroenterology 154, 585–598 (2018).
Kraiczy, J. et al. DNA methylation defines regional identity of human intestinal epithelial organoids and undergoes dynamic changes during development. Gut 68, 49–61 (2019). First study exploring the segment and development-specific DNA methylation and transcriptomic differences in human terminal ileum and sigmoid colon.
Kazakevych, J., Sayols, S., Messner, B., Krienke, C. & Soshnikova, N. Dynamic changes in chromatin states during specification and differentiation of adult intestinal stem cells. Nucleic Acids Res. 45, 5770–5784 (2017). This is the first study achieving chromatin and transcriptome profiling from embryonic to adult intestinal development, defining these differences between specification and differentiation of intestinal stem cells.
Pan, W. H. et al. Exposure to the gut microbiota drives distinct methylome and transcriptome changes in intestinal epithelial cells during postnatal development. Genome Med. 10, 27 (2018). This study reveals information about the contribution of an antibiotic treatment (and thus changes in microbiota composition) in pigs on DNA methylation, protein expression and physiology in the small intestine.
Elliott, E. N. & Kaestner, K. H. Epigenetic regulation of the intestinal epithelium. Cell. Mol. Life Sci. 72, 4139–4156 (2015).
Klemm, S. L., Shipony, Z. & Greenleaf, W. J. Chromatin accessibility and the regulatory epigenome. Nat. Rev. Genet. 20, 207–220 (2019).
Kim, T. H. et al. Broadly permissive intestinal chromatin underlies lateral inhibition and cell plasticity. Nature 506, 511–515 (2014). This study establishes a basic framework for accessible chromatin maps and dynamics in multiple intestinal cell types and lineages undergoing differentiation.
Jadhav, U. et al. Acquired tissue-specific promoter bivalency is a basis for PRC2 necessity in adult cells. Cell 165, 1389–1400 (2016).
Jadhav, U. et al. Dynamic reorganization of chromatin accessibility signatures during dedifferentiation of secretory precursors into Lgr5+ intestinal stem cells. Cell Stem Cell 21, 65–77.e5 (2017).
Troll, J. V. et al. Microbiota promote secretory cell determination in the intestinal epithelium by modulating host Notch signaling. Development 145, dev155317 (2018).
Davison, J. M. et al. Microbiota regulate intestinal epithelial gene expression by suppressing the transcription factor Hepatocyte nuclear factor 4 alpha. Genome Res. 27, 1195–1206 (2017). This study identifies microbially responsive regulatory regions genome wide and establishes the involvement of Hnf4a in regulating microbially responsive genes in the mouse and zebrafish intestine.
Richards, A. L. et al. Gut microbiota has a widespread and modifiable effect on host gene regulation. mSystems 4, e00323–18 (2019).
Shlyueva, D., Stampfel, G. & Stark, A. Transcriptional enhancers: from properties to genome-wide predictions. Nat. Rev. Genet. 15, 272–286 (2014).
Oittinen, M. et al. Polycomb repressive complex 2 enacts Wnt signaling in intestinal homeostasis and contributes to the instigation of stemness in diseases entailing epithelial hyperplasia or neoplasia. Stem Cell 35, 445–457 (2017).
Jadhav, U. et al. Extensive recovery of embryonic enhancer and gene memory stored in hypomethylated enhancer DNA. Mol. Cell 74, 542–554.e5 (2019). This paper describes how DNA methylation and histone methylation act as combinatorial signals to create an epigenetic memory in bivalent promoters.
Binda, O. & Fernandez-Zapico, M. E. (eds) Chromatin Signaling and Diseases. (Academic Press, 2016).
Turgeon, N. et al. HDAC1 and HDAC2 restrain the intestinal inflammatory response by regulating intestinal epithelial cell differentiation. PLoS ONE 8, e73785 (2013).
Zimberlin, C. D. et al. HDAC1 and HDAC2 collectively regulate intestinal stem cell homeostasis. FASEB J. 29, 2070–2080 (2015).
Alenghat, T. et al. Histone deacetylase 3 coordinates commensal-bacteria-dependent intestinal homeostasis. Nature 504, 153–157 (2013).
Mukherji, A., Kobiita, A., Ye, T. & Chambon, P. Homeostasis in intestinal epithelium is orchestrated by the circadian clock and microbiota cues transduced by TLRs. Cell 153, 812–827 (2013).
Kuang, Z. et al. The intestinal microbiota programs diurnal rhythms in host metabolism through histone deacetylase 3. Science 365, 1428–1434 (2019).
Morrison, D. J. & Preston, T. Formation of short chain fatty acids by the gut microbiota and their impact on human metabolism. Gut Microbes 7, 189–200 (2016).
Krautkramer, K. A. et al. Diet-microbiota interactions mediate global epigenetic programming in multiple host tissues. Mol. Cell 64, 982–992 (2016).
Yuille, S., Reichardt, N., Panda, S., Dunbar, H. & Mulder, I. E. Human gut bacteria as potent class I histone deacetylase inhibitors in vitro through production of butyric acid and valeric acid. PLoS ONE 13, e0201073 (2018).
Donohoe, D. R. et al. The Warburg effect dictates the mechanism of butyrate-mediated histone acetylation and cell proliferation. Mol. Cell 48, 612–626 (2012).
Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).
Chen, X. et al. Integration of external signaling pathways with the core transcriptional network in embryonic stem cells. Cell 133, 1106–1117 (2008).
Lee, T. I. & Young, R. A. Transcriptional regulation and its misregulation in disease. Cell 152, 1237–1251 (2013).
Iwafuchi-Doi, M. & Zaret, K. S. Pioneer transcription factors in cell reprogramming. Genes Dev. 28, 2679–2692 (2014).
Verzi, M. P. et al. Differentiation-specific histone modifications reveal dynamic chromatin interactions and partners for the intestinal transcription factor CDX2. Dev. Cell 19, 713–726 (2010).
Verzi, M. P., Shin, H., San Roman, A. K., Liu, X. S. & Shivdasani, R. A. Intestinal master transcription factor CDX2 controls chromatin access for partner transcription factor binding. Mol. Cell. Biol. 33, 281–292 (2013). This paper describes the role of Cdx2 as a lineage-specifying transcription factor, whose activities are a critical part of establishing the chromatin landscape in the intestinal epithelium.
San Roman, A. K., Tovaglieri, A., Breault, D. T. & Shivdasani, R. A. Distinct processes and transcriptional targets underlie CDX2 requirements in intestinal stem cells and differentiated villus cells. Stem Cell Rep. 5, 673–681 (2015).
Flores, M. V. et al. Intestinal differentiation in zebrafish requires Cdx1b, a functional equivalent of mammalian Cdx2. Gastroenterology 135, 1665–1675 (2008).
Chen, T. & Dent, S. Y. R. Chromatin modifiers and remodellers: regulators of cellular differentiation. Nat. Rev. Genet. 15, 93–106 (2014).
Kumar, N. et al. The lineage-specific transcription factor CDX2 navigates dynamic chromatin to control distinct stages of intestine development. Development 146, dev172189 (2019).
Hryniuk, A., Grainger, S., Savory, J. G. & Lohnes, D. Cdx function is required for maintenance of intestinal identity in the adult. Dev. Biol. 363, 426–437 (2012).
Kazumori, H., Ishihara, S., Rumi, M. A., Kadowaki, Y. & Kinoshita, Y. Bile acids directly augment caudal related homeobox gene Cdx2 expression in oesophageal keratinocytes in Barrett’s epithelium. Gut 55, 16–25 (2006).
Domon-Dell, C. et al. Stimulation of the intestinal Cdx2 homeobox gene by butyrate in colon cancer cells. Gut 50, 525–529 (2002).
Lee, S. H. et al. Burkholderia pseudomallei suppresses Caenorhabditis elegans immunity by specific degradation of a GATA transcription factor. Proc. Natl Acad. Sci. USA 110, 15067–15072 (2013).
Yang, W. et al. The inducible response of the nematode Caenorhabditis elegans to members of its natural microbiome across development and adult life. Front Microbiol. 10, 1793 (2019).
Yang, W., Dierking, K., Rosenstiel, P. C. & Schulenburg, H. GATA transcription factor as a likely key regulator of the Caenorhabditis elegans innate immune response against gut pathogens. Zoology 119, 244–253 (2016).
Kawasaki, T. & Kawai, T. Toll-like receptor signaling pathways. Front. Immunol. 5, 461 (2014).
Lei-Leston, A. C., Murphy, A. G. & Maloy, K. J. Epithelial cell inflammasomes in intestinal immunity and inflammation. Front. Immunol. 8, 1168 (2017).
Price, A. E. et al. A map of toll-like receptor expression in the intestinal epithelium reveals distinct spatial, cell type-specific, and temporal patterns. Immunity 49, 560–575.e6 (2018). The authors created transgenic reporter mouse strains that were used to visualize the localization of Toll-like receptors in the intestine along both the longitudinal and the crypt–villus axes, on specific cell types, and on the basolataral and/or apical faces of the epithelium.
Schmidt-Ullrich, R. et al. NF-kappaB activity in transgenic mice: developmental regulation and tissue specificity. Development 122, 2117–2128 (1996).
Karrasch, T., Kim, J. S., Muhlbauer, M., Magness, S. T. & Jobin, C. Gnotobiotic IL-10−/−;NF-κB(EGFP) mice reveal the critical role of TLR/NF-κB signaling in commensal bacteria-induced colitis. J. Immunol. 178, 6522–6532 (2007).
Kanther, M. et al. Microbial colonization induces dynamic temporal and spatial patterns of NF-κB activation in the zebrafish digestive tract. Gastroenterology 141, 197–207 (2011). These authors created and characterized a transgenic zebrafish enabling real-time, in vivo imaging of the dynamic patterns of NF-κB activation in response to microbiota colonization.
Vlantis, K. et al. NEMO prevents RIP kinase 1-mediated epithelial cell death and chronic intestinal inflammation by NF-κB-dependent and -independent functions. Immunity 44, 553–567 (2016).
Steinbrecher, K. A., Harmel-Laws, E., Sitcheran, R. & Baldwin, A. S. Loss of epithelial RelA results in deregulated intestinal proliferative/apoptotic homeostasis and susceptibility to inflammation. J. Immunol. 180, 2588–2599 (2008).
Cuiv, O. P. et al. Enterococcus faecalis AHG0090 is a genetically tractable bacterium and produces a secreted peptidic bioactive that suppresses nuclear factor kappa B activation in human gut epithelial cells. Front. Immunol. 9, 790 (2018).
Quevrain, E. et al. Identification of an anti-inflammatory protein from Faecalibacterium prausnitzii, a commensal bacterium deficient in Crohn’s disease. Gut 65, 415–425 (2016).
Nie, N. et al. Bifidobacterium plays a protective role in TNF-α-induced inflammatory response in Caco-2 cell through NF-κB and p38MAPK pathways. Mol. Cell. Biochem. 464, 83–91 (2020).
Zaidi, D. & Wine, E. Regulation of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κβ) in inflammatory bowel diseases. Front. Pediatr. 6, 317 (2018). This review covers a wide range of literature linking NF-κβ to inflammatory bowel disease, including recent papers on NF-κβ regulation in intestinal epithelial cells.
Kisseleva, T., Bhattacharya, S., Braunstein, J. & Schindler, C. W. Signaling through the JAK/STAT pathway, recent advances and future challenges. Gene 285, 1–24 (2002).
LaPorte, S. L. et al. Molecular and structural basis of cytokine receptor pleiotropy in the interleukin-4/13 system. Cell 132, 259–272 (2008).
De Oliveira, T. et al. Loss of Stat6 affects chromatin condensation in intestinal epithelial cells causing diverse outcome in murine models of inflammation-associated and sporadic colon carcinogenesis. Oncogene 38, 1787–1801 (2019).
Schubart, C. et al. Selective expression of constitutively activated STAT6 in intestinal epithelial cells promotes differentiation of secretory cells and protection against helminths. Mucosal Immunol. 12, 413–424 (2019).
Bollrath, J. et al. gp130-mediated Stat3 activation in enterocytes regulates cell survival and cell-cycle progression during colitis-associated tumorigenesis. Cancer Cell 15, 91–102 (2009).
Pickert, G. et al. STAT3 links IL-22 signaling in intestinal epithelial cells to mucosal wound healing. J. Exp. Med. 206, 1465–1472 (2009).
Sugimoto, K. et al. IL-22 ameliorates intestinal inflammation in a mouse model of ulcerative colitis. J. Clin. Invest. 118, 534–544 (2008).
Choi, S. M. et al. Innate Stat3-mediated induction of the antimicrobial protein Reg3γ is required for host defense against MRSA pneumonia. J. Exp. Med. 210, 551–561 (2013).
Lee, K. S. et al. Helicobacter pylori CagA triggers expression of the bactericidal lectin REG3γ via gastric STAT3 activation. PLoS ONE 7, e30786 (2012).
Hertz, R., Magenheim, J., Berman, I. & Bar-Tana, J. Fatty acyl-CoA thioesters are ligands of hepatic nuclear factor-4α. Nature 392, 512–516 (1998).
Yuan, X. et al. Identification of an endogenous ligand bound to a native orphan nuclear receptor. PLoS ONE 4, e5609 (2009).
Darsigny, M. et al. Loss of hepatocyte-nuclear-factor-4α affects colonic ion transport and causes chronic inflammation resembling inflammatory bowel disease in mice. PLoS ONE 4, e7609 (2009).
Gerdin, A. K. et al. Phenotypic screening of hepatocyte nuclear factor (HNF) 4-γ receptor knockout mice. Biochem. Biophys. Res. Commun. 349, 825–832 (2006).
Darsigny, M. et al. Hepatocyte nuclear factor-4α promotes gut neoplasia in mice and protects against the production of reactive oxygen species. Cancer Res. 70, 9423–9433 (2010).
Babeu, J. P., Darsigny, M., Lussier, C. R. & Boudreau, F. Hepatocyte nuclear factor 4α contributes to an intestinal epithelial phenotype in vitro and plays a partial role in mouse intestinal epithelium differentiation. Am. J. Physiol. Gastrointest. Liver Physiol. 297, G124–G134 (2009).
Frochot, V. et al. The transcription factor HNF-4α: a key factor of the intestinal uptake of fatty acids in mouse. Am. J. Physiol. Gastrointest. Liver Physiol. 302, G1253–G1263 (2012).
Cattin, A. L. et al. Hepatocyte nuclear factor 4α, a key factor for homeostasis, cell architecture, and barrier function of the adult intestinal epithelium. Mol. Cell. Biol. 29, 6294–6308 (2009).
Chahar, S. et al. Chromatin profiling reveals regulatory network shifts and a protective role for hepatocyte nuclear factor 4α during colitis. Mol. Cell. Biol. 34, 3291–3304 (2014).
Baraille, F. et al. Glucose tolerance is improved in mice invalidated for the nuclear receptor HNF-4γ: a critical role for enteroendocrine cell lineage. Diabetes 64, 2744–2756 (2015).
Shukla, S. et al. Aberrant activation of a gastrointestinal transcriptional circuit in prostate cancer mediates castration resistance. Cancer Cell 32, 792–806.e7 (2017).
Chen, L. et al. HNF4 factors control chromatin accessibility and are redundantly required for maturation of the fetal intestine. Development 146, dev179432 (2019). This is the first study to determine the genetic redundancy between Hnf4a and Hnf4g transcription factors during mouse intestinal development.
Chen, L. et al. HNF4 regulates fatty acid oxidation and is required for renewal of intestinal stem cells in mice. Gastroenterology 158, 985–999.e9 (2019). This work identifies a role for a ligand-regulated nuclear receptor transcription factor in intestinal stem cell renewal, which might implicate potential microbial regulation of this factor in influencing intestinal stem cell function.
Yamagata, K. et al. Mutations in the hepatocyte nuclear factor-4α gene in maturity-onset diabetes of the young (MODY1). Nature 384, 458–460 (1996).
Barrett, J. C. et al. Genome-wide association study of ulcerative colitis identifies three new susceptibility loci, including the HNF4A region. Nat. Genet. 41, 1330–1334 (2009).
Jostins, L. et al. Host-microbe interactions have shaped the genetic architecture of inflammatory bowel disease. Nature 491, 119–124 (2012).
Marcil, V. et al. Association between genetic variants in the HNF4A gene and childhood-onset Crohn’s disease. Genes Immun. 13, 556–565 (2012).
Ahn, S. H. et al. Hepatocyte nuclear factor 4α in the intestinal epithelial cells protects against inflammatory bowel disease. Inflamm. Bowel Dis. 14, 908–920 (2008).
Mokry, M. et al. Many inflammatory bowel disease risk loci include regions that regulate gene expression in immune cells and the intestinal epithelium. Gastroenterology 146, 1040–1047 (2014).
Meddens, C. A. et al. Systematic analysis of chromatin interactions at disease associated loci links novel candidate genes to inflammatory bowel disease. Genome Biol. 17, 247 (2016).
Franke, A. et al. Systematic association mapping identifies NELL1 as a novel IBD disease gene. PLoS ONE 2, e691 (2007).
Camp, J. G., Jazwa, A. L., Trent, C. M. & Rawls, J. F. Intronic cis-regulatory modules mediate tissue-specific and microbial control of angptl4/fiaf transcription. PLoS Genet. 8, e1002585 (2012).
Dhe-Paganon, S., Duda, K., Iwamoto, M., Chi, Y. I. & Shoelson, S. E. Crystal structure of the HNF4α ligand binding domain in complex with endogenous fatty acid ligand. J. Biol. Chem. 277, 37973–37976 (2002).
Wisely, G. B. et al. Hepatocyte nuclear factor 4 is a transcription factor that constitutively binds fatty acids. Structure 10, 1225–1234 (2002).
Palanker, L., Tennessen, J. M., Lam, G. & Thummel, C. S. Drosophila HNF4 regulates lipid mobilization and β-oxidation. Cell Metab. 9, 228–239 (2009).
Devillard, E., McIntosh, F. M., Duncan, S. H. & Wallace, R. J. Metabolism of linoleic acid by human gut bacteria: different routes for biosynthesis of conjugated linoleic acid. J. Bacteriol. 189, 2566–2570 (2007).
Druart, C. et al. Role of the lower and upper intestine in the production and absorption of gut microbiota-derived PUFA metabolites. PLoS ONE 9, e87560 (2014).
Makishima, M. et al. Identification of a nuclear receptor for bile acids. Science 284, 1362–1365 (1999).
Hwang, S. T., Urizar, N. L., Moore, D. D. & Henning, S. J. Bile acids regulate the ontogenic expression of ileal bile acid binding protein in the rat via the farnesoid X receptor. Gastroenterology 122, 1483–1492 (2002).
Zhu, Y., Li, F. & Guo, G. L. Tissue-specific function of farnesoid X receptor in liver and intestine. Pharmacol. Res. 63, 259–265 (2011).
Gonzalez, F. J., Jiang, C. & Patterson, A. D. An intestinal microbiota-farnesoid X receptor axis modulates metabolic disease. Gastroenterology 151, 845–859 (2016).
Devlin, A. S. & Fischbach, M. A. A biosynthetic pathway for a prominent class of microbiota-derived bile acids. Nat. Chem. Biol. 11, 685–690 (2015).
Sayin, S. I. et al. Gut microbiota regulates bile acid metabolism by reducing the levels of tauro-beta-muricholic acid, a naturally occurring FXR antagonist. Cell Metab. 17, 225–235 (2013).
Wahlstrom, A. et al. Induction of farnesoid X receptor signaling in germ-free mice colonized with a human microbiota. J. Lipid Res. 58, 412–419 (2017).
Li, F. et al. Microbiome remodelling leads to inhibition of intestinal farnesoid X receptor signalling and decreased obesity. Nat. Commun. 4, 2384 (2013).
Degirolamo, C., Rainaldi, S., Bovenga, F., Murzilli, S. & Moschetta, A. Microbiota modification with probiotics induces hepatic bile acid synthesis via downregulation of the Fxr-Fgf15 axis in mice. Cell Rep. 7, 12–18 (2014).
Parséus, A. et al. Microbiota-induced obesity requires farnesoid X receptor. Gut 66, 429–437 (2016). This study investigated the interaction between diet, the microbiota and FXR signalling, and shows that FXR is required for high-fat diet to induce weight gain and that the effects might be due to differences in the microbiota observed in Fxr −/− mice.
Xie, C. et al. An intestinal farnesoid X receptor-ceramide signaling axis modulates hepatic gluconeogenesis in mice. Diabetes 66, 613–626 (2017).
Gadaleta, R. M. et al. Farnesoid X receptor activation inhibits inflammation and preserves the intestinal barrier in inflammatory bowel disease. Gut 60, 463–472 (2011).
Evans, R. M. & Mangelsdorf, D. J. Nuclear receptors, RXR, and the big bang. Cell 157, 255–266 (2014).
Duszka, K. & Wahli, W. Enteric microbiota–gut–brain axis from the perspective of nuclear receptors. Int. J. Mol. Sci. 19, 2210 (2018).
Chinetti, G., Fruchart, J. C. & Staels, B. Peroxisome proliferator-activated receptors (PPARs): nuclear receptors at the crossroads between lipid metabolism and inflammation. Inflamm. Res. 49, 497–505 (2000).
Hasan, A. U., Rahman, A. & Kobori, H. Interactions between host PPARs and gut microbiota in health and disease. Int. J. Mol. Sci. 20, 387 (2019).
Byndloss, M. X. et al. Microbiota-activated PPAR-γ signaling inhibits dysbiotic Enterobacteriaceae expansion. Science 357, 570–575 (2017).
Peyrin-Biroulet, L. et al. Peroxisome proliferator-activated receptor gamma activation is required for maintenance of innate antimicrobial immunity in the colon. Proc. Natl Acad. Sci. USA 107, 8772–8777 (2010).
Su, C. G. et al. A novel therapy for colitis utilizing PPAR-γ ligands to inhibit the epithelial inflammatory response. J. Clin. Invest. 104, 383–389 (1999).
Celinski, K. et al. Comparison of the anti-inflammatory and therapeutic actions of PPAR-gamma agonists rosiglitazone and troglitazone in experimental colitis. J. Physiol. Pharmacol. 63, 631–640 (2012).
Dubuquoy, L. et al. Impaired expression of peroxisome proliferator-activated receptor γ in ulcerative colitis. Gastroenterology 124, 1265–1276 (2003).
Dou, X., Xiao, J., Jin, Z. & Zheng, P. Peroxisome proliferator-activated receptor-γ is downregulated in ulcerative colitis and is involved in experimental colitis-associated neoplasia. Oncol. Lett. 10, 1259–1266 (2015).
Lu, P. et al. Intestinal epithelial Toll-like receptor 4 prevents metabolic syndrome by regulating interactions between microbes and intestinal epithelial cells in mice. Mucosal Immunol. 11, 727–740 (2018).
Kelly, D. et al. Commensal anaerobic gut bacteria attenuate inflammation by regulating nuclear-cytoplasmic shuttling of PPAR-γ and RelA. Nat. Immunol. 5, 104–112 (2004).
Manoharan, I. et al. Homeostatic PPARα signaling limits inflammatory responses to commensal microbiota in the intestine. J. Immunol. 196, 4739–4749 (2016).
Tomas, J. et al. High-fat diet modifies the PPAR-γ pathway leading to disruption of microbial and physiological ecosystem in murine small intestine. Proc. Natl Acad. Sci. USA 113, E5934–E5943 (2016).
Prakash, C. et al. Nuclear receptors in drug metabolism, drug response and drug interactions. Nucl. Receptor Res. 2, 101173 (2015).
Wang, Y. M., Ong, S. S., Chai, S. C. & Chen, T. Role of CAR and PXR in xenobiotic sensing and metabolism. Expert. Opin. Drug Metab. Toxicol. 8, 803–817 (2012).
Hudson, G. M. et al. Constitutive androstane receptor regulates the intestinal mucosal response to injury. Br. J. Pharmacol. 174, 1857–1871 (2017).
Venkatesh, M. et al. Symbiotic bacterial metabolites regulate gastrointestinal barrier function via the xenobiotic sensor PXR and Toll-like receptor 4. Immunity 41, 296–310 (2014).
Terc, J., Hansen, A., Alston, L. & Hirota, S. A. Pregnane X receptor agonists enhance intestinal epithelial wound healing and repair of the intestinal barrier following the induction of experimental colitis. Eur. J. Pharm. Sci. 55, 12–19 (2014).
Garg, A. et al. Pregnane X receptor activation attenuates inflammation-associated intestinal epithelial barrier dysfunction by inhibiting cytokine-induced myosin light-chain kinase expression and c-Jun N-terminal kinase 1/2 activation. J. Pharmacol. Exp. Ther. 359, 91–101 (2016).
Shakhnovich, V. et al. Decreased pregnane X receptor expression in children with active Crohn’s disease. Drug Metab. Dispos. 44, 1066–1069 (2016).
Swanson, H. I. Drug metabolism by the host and gut microbiota: a partnership or rivalry? Drug Metab. Dispos. 43, 1499–1504 (2015).
Kawajiri, K. & Fujii-Kuriyama, Y. The aryl hydrocarbon receptor: a multifunctional chemical sensor for host defense and homeostatic maintenance. Exp. Anim. 66, 75–89 (2017).
Bock, K. W. Human and rodent aryl hydrocarbon receptor (AHR): from mediator of dioxin toxicity to physiologic AHR functions and therapeutic options. Biol. Chem. 398, 455–464 (2017).
Moura-Alves, P. et al. AhR sensing of bacterial pigments regulates antibacterial defence. Nature 512, 387–392 (2014).
Hubbard, T. D. et al. Adaptation of the human aryl hydrocarbon receptor to sense microbiota-derived indoles. Sci. Rep. 5, 12689 (2015).
Lamas, B. et al. CARD9 impacts colitis by altering gut microbiota metabolism of tryptophan into aryl hydrocarbon receptor ligands. Nat. Med. 22, 598–605 (2016).
Sun, M., Ma, N., He, T., Johnston, L. J. & Ma, X. Tryptophan (Trp) modulates gut homeostasis via aryl hydrocarbon receptor (AhR). Crit. Rev. Food Sci. Nutr. 60, 1760–1768 (2019).
Marinelli, L. et al. Identification of the novel role of butyrate as AhR ligand in human intestinal epithelial cells. Sci. Rep. 9, 643 (2019).
Fukumoto, S. et al. Identification of a probiotic bacteria-derived activator of the aryl hydrocarbon receptor that inhibits colitis. Immunol. Cell Biol. 92, 460–465 (2014).
Lamas, B., Natividad, J. M. & Sokol, H. Aryl hydrocarbon receptor and intestinal immunity. Mucosal Immunol. 11, 1024–1038 (2018).
Furumatsu, K. et al. A role of the aryl hydrocarbon receptor in attenuation of colitis. Dig. Dis. Sci. 56, 2532–2544 (2011).
Yu, M. et al. Aryl hydrocarbon receptor activation modulates intestinal epithelial barrier function by maintaining tight junction integrity. Int. J. Biol. Sci. 14, 69–77 (2018).
Qiu, J. et al. The aryl hydrocarbon receptor regulates gut immunity through modulation of innate lymphoid cells. Immunity 36, 92–104 (2012).
Monteleone, I. et al. Aryl hydrocarbon receptor-induced signals up-regulate IL-22 production and inhibit inflammation in the gastrointestinal tract. Gastroenterology 141, 237–248.e1 (2011).
Lanis, J. M. et al. Tryptophan metabolite activation of the aryl hydrocarbon receptor regulates IL-10 receptor expression on intestinal epithelia. Mucosal Immunol. 10, 1133–1144 (2017).
Kawai, S. et al. Indigo naturalis ameliorates murine dextran sodium sulfate-induced colitis via aryl hydrocarbon receptor activation. J. Gastroenterol. 52, 904–919 (2017).
Yoshimatsu, Y. et al. Development of an Indigo naturalis suppository for topical induction therapy in patients with ulcerative colitis. Digestion 101, 1–7 (2019).
Marafini, I. et al. NPD-0414-2 and NPD-0414-24, two chemical entities designed as aryl hydrocarbon receptor (AhR) ligands, inhibit gut inflammatory signals. Front. Pharmacol. 10, 380 (2019).
Kitajima, S., Morimoto, M., Sagara, E., Shimizu, C. & Ikeda, Y. Dextran sodium sulfate-induced colitis in germ-free IQI/Jic mice. Exp. Anim. 50, 387–395 (2001).
Maslowski, K. M. et al. Regulation of inflammatory responses by gut microbiota and chemoattractant receptor GPR43. Nature 461, 1282–1286 (2009).
Morgun, A. et al. Uncovering effects of antibiotics on the host and microbiota using transkingdom gene networks. Gut 64, 1732–1743 (2015).
The authors are grateful to members of the laboratory of J.F.R. for their helpful feedback on this manuscript. The authors were supported by a Pew Scholars Innovation Fund Award (J.F.R.) and NIH grants R01-DK081426, R24-DK110492, R01-DK093399, R01-DK113123, R24-OD016761, and P01-DK094779 (J.F.R.), F31-DK121392 (C.K.), and the UNC-CH Gastroenterology Research Training Program T32-DK07737 (J.K.H.).
The authors declare no competing interests.
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- Intestinal epithelial cells
(IECs). The cells that comprise the columnar epithelial layer that lines the lumen of the digestive tract from the anterior small intestine to the rectum, serving multiple functions including as a physical barrier and an absorptive tissue.
- Gut microbiota
The microorganisms that colonize the lumen and mucosal surfaces of the gastrointestinal tract.
- Intestinal cell type or lineage identity
The intestinal epithelium comprises many different specialized types of cells (for example, absorptive enterocytes, enteroendocrine cells, goblet cells) that are specified partly through distinct gene expression programmes.
The complex consisting of chromosomal DNA and associated protein and RNA within the nucleus.
- Inflammatory bowel disease
(IBD). Conditions characterized by chronic inflammation of the intestine, most commonly ulcerative colitis and Crohn’s disease.
- Transcription factors
Proteins that regulate transcription (gene expression), typically by binding to specific DNA sequences.
- Anterior–posterior axis of the intestine
Differences in the physiological function and underlying cellular composition and gene expression patterns along the anterior–posterior axis of the gastrointestinal tract. This regionality is typically categorized into the major regions of the gastrointestinal tract (for example, duodenum, jejunum, ileum, colon, etc.).
- Intestinal stem cells
(ISCs). Cells in the intestinal epithelium that undergo self-renewal and also give rise to all of the different intestinal epithelial cell types. They are stereotypically located at the base of intestinal crypts or rugae.
- Secretory cells
A major IEC lineage with diverse functions mediated partially through secretion of products into the lumen. The secretory cell lineage includes Paneth cells, tuft cells, goblet cells and enteroendocrine cells.
- Absorptive cell
A major IEC lineage that gives rise to absorptive enterocytes, the most abundant cell type in the intestine, which are also responsible for absorption of nutrients and other cargoes.
- Lysosome-rich enterocytes
(LREs). Specialized absorptive cell type found in the intestines characterized by a large lysosomal vacuole and involved in cellular digestion of macromolecular cargoes. These cells develop in diverse vertebrate species but are often overlooked in mammals because they are only present during the ‘suckling’ or perinatal developmental stages.
- Lineage-specific transcription factors
Transcription factors that function early on in the differentiation of a cell lineage or tissue that have a role in shaping the chromatin landscape, setting the stage for further transcription factor binding and lineage specification. Lineage-specifying transcription factors are functionally similar to pioneer transcription factors, but unlike pioneer transcription factors they have not definitively been shown to directly bind to condensed chromatin.
- DNA methylation
The addition of methyl groups on DNA bases that serves as an additional layer of genetic information that can impact the expression of genes. Typically, methylation of a gene promoter results in suppression of that gene’s transcription.
- Chromatin accessibility
The degree to which regions of chromatin DNA are available for transcription factor binding or other regulatory processes. Typically, chromatin accessibility is low due to nucleosome occupancy and can be modulated by post-translational modification of histone tails. This term is used interchangeably with chromatin openness.
- Histone modifications
Post-translational modifications on the tails of histone proteins that can influence regional gene expression and other physical and enzymatic genome utilization dynamics.
- Cis-regulatory DNA regions
(CRRs). Typically non-coding regions of the genome often involved in modulating the transcription of a nearby gene. This term is used interchangeably with cis-regulatory element (CRE).
Structural unit of DNA organization in the nucleus, consisting of DNA wrapped around a complex of eight histone proteins.
- Short-chain fatty acids
(SCFAs). Short-chain fatty acids are products of microbial fermentation of non-digestible dietary fibre and protein, and a main source of energy for epithelial cells in the colon.
- Bile acid
Cholesterol-derived acids that are secreted from the gallbladder into the small intestine. They assist in emulsification of dietary fats, act as ligands for multiple host receptors and transcription factors and can be chemically modified by the intestinal microbiota.
- Nuclear receptor
A family of ligand-binding transcription factors with important roles in intestinal epithelial cells that regulate the expression of genes associated with diverse processes from development to metabolism.
Chemicals that are foreign to the body.
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Heppert, J.K., Davison, J.M., Kelly, C. et al. Transcriptional programmes underlying cellular identity and microbial responsiveness in the intestinal epithelium. Nat Rev Gastroenterol Hepatol 18, 7–23 (2021). https://doi.org/10.1038/s41575-020-00357-6
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