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Distribution and Stimulus Secretion Coupling of Enteroendocrine Cells along the Intestinal Tract

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ABSTRACT

The enteroendocrine system of the gut acts both locally and peripherally, regulating gastrointestinal function as well as metabolism, energy expenditure, and central appetite control through the release of a variety of hormones. The chemosensing ability of enteroendocrine cells is integral to their role in eliciting physiological changes in response to fluctuations in the composition of the intestinal lumen. Regulation of enteroendocrine cell activity is complex, and requires that these cells can integrate signals deriving from dietary sources as well as the nervous and endocrine systems. Here, we provide an overview of enteroendocrine cell form and function, with a focus on new insights into their distribution throughout the intestine and the stimulus secretion coupling mechanisms underlying the activity of these important members of the gut‐brain axis. © 2018 American Physiological Society. Compr Physiol 8:1603‐1638, 2018.

Figure 1. Figure 1. Schematic diagram illustrating the microstructure of the intestine. Epithelial cell types are identified along the crypt‐villus axis.
Figure 2. Figure 2. Examples of the crypt‐villus axis. (A) Widefield image of the base of a villus and its accompanying crypts from enzymatically dispersed tissue. (B) Confocal image of a villus and attached crypt. Chromogranin‐A‐positive enteroendocrine cells are stained in red. E‐cadherin, staining adherin junctions between epithelial cells, is stained in green. (Images courtesy of Deborah Goldspink, Reimann/Gribble laboratories, with permission.)
Figure 3. Figure 3. Depiction of the variety of morphologies and signaling modalities adopted by enteroendocrine cells. An open‐type enteroendocrine cell (red) is being contacted by a basolateral extension from a neighboring enteroendocrine cell (blue), facilitating paracrine signaling. Capillaries within the lamina propria mediate endocrine signaling from gut hormones secreted into the paracellular space. A closed‐type enteroendocrine cell is illustrated in grey. Neuroendocrine communication is enabled by neuronal nerve fibers that project into the intestinal mucosa (black). Some enteroendocrine cells form “neuropods” from their basolateral surface, which synapse with neuronal termini (orange).
Figure 4. Figure 4. Key sites of release for prominent gut hormones expressed throughout the gastrointestinal tract. Details are discussed in the text. Abbreviations: CCK, cholecystokinin; GIP, glucose‐dependent insulinotropic polypeptide; GLP‐1, glucagon‐like peptide‐1; Insl5, Insulin‐like peptide 5; PYY, peptide YY; SST, somatostatin; 5‐HT, 5‐hydroxy‐tryptamine.
Figure 5. Figure 5. Coproduction of GLP‐1 and PYY in human colonic enteroendocrine cells. Human colonic epithelial cells were fixed and stained for GLP‐1 and PYY and analyzed by flow cytometry (A). Coproduction of PYY and GLP‐1 was confirmed via immunohistochemistry in cultured human colon epithelium. The scale bar represents 10 µm. Data were obtained and adapted, with permission, from Habib et al., Diabetologia, 2013 ().
Figure 6. Figure 6. Overlap of gut hormone production in enteroendocrine cells purified from the mouse intestine. (A) GLP‐1‐ (“L”) and GIP‐expressing (“K”) cells were sorted from upper/lower small intestinal (“up‐/low‐”) and colonic (“LC”) epithelium by flow cytometry from transgenic mice expressing a fluorescent reporter (venus) under the control of the proglucagon or GIP promoter, respectively. Nonfluorescent control samples were collected from the same regions. Expression of gut hormone transcripts was measured in both fluorescent (“+”) and nonfluorescent (“−”) samples by quantitative PCR and compared to Actb expression. Data are presented as means and SEM values. (B) Fluorescence‐assisted cell sorting (FACS) analysis of small intestinal cells from transgenic mice expressing venus under the control of the proglucagon promoter costained for GLP‐1, GIP, CCK, SST, and PYY. Percentages of venus‐positive cells costaining for gut peptides were calculated and presented in a histogram for cells isolated from the upper and lower small intestine (SI). Data represent the mean and SEM for the number of samples indicated above each bar. (C,D) The frequency of either GLP‐1‐(C) or PYY‐(D)‐producing cells along the intestinal tract was analyzed using FACS. Intestinal epithelial cells were isolated from mice expressing venus under the control of the proglucagon promoter and stained for either GLP‐1 or PYY. The y‐axis represents the cell frequency. Data were obtained and adapted, with permission, from Habib et al., Endocrinology 2012 ().
Figure 7. Figure 7. Diet‐induced changes in enteroendocrine cells. Mice expressing venus under the control of the proglucagon promoter were subjected to either a high fat (HFD) or control chow diet for two weeks. The number of GLP‐1‐expressing cells in colonic epithelium was determined by flow cytometry. HFD mice exhibited a significant decrease in GLP‐1‐expressing cells compared to controls. Differences in enteroendocrine cell numbers between chow and HFD samples were compared using a Student's t‐test: *P < 0.05 (A). Colonic epithelial cells isolated from mice expressing venus under the control of the proglucagon promoter fed a HFD or control diet for 16 weeks were isolated via flow cytometry. The expression of gut hormones (B), prohormone processing enzymes (C), granins (D), and fatty acid binding proteins (E) was assessed via Affymetrix ST1.0 microarray. Venus‐negative cells are represented by striped bars (chow, horizontal; HFD, diagonal). Venus‐positive cells are represented by solid bars (chow, white; HFD, black). Data are represented as mean RMA intensity values and SEM. Expression levels between chow and HFD samples were compared using a Student's t‐test: *P < 0.05, **P < 0.01, ***P < 0.001. Data were obtained and adapted, with permission, from Richards et al., Peptides, 2016 ().
Figure 8. Figure 8. Proposed mechanisms for glucose sensing in enteroendocrine cells. The electrochemical gradient of sodium drives uptake of glucose from the intestinal lumen via the sodium‐glucose cotransporter, SGLT1. SGLT1 activity evokes a small depolarization of the plasma membrane, which activates voltage‐gated calcium channels (VGCCs). The resultant calcium influx further depolarizes the enteroendocrine cell plasma membrane and leads to secretory granule release. Glucose transported into the cell by either SGLT1 or facilitative transport is metabolized, raising cytosolic ATP levels, which closes KATP channels at the cell surface. Inhibition of potassium efflux further depolarizes the cell membrane, contributing to enteroendocrine cell excitation and subsequent gut hormone secretion.
Figure 9. Figure 9. Sensing of dietary protein in enteroendocrine cells. L‐Gln‐stimulated gut hormone release proceeds via a mechanism linked to sodium‐coupled transport as well as by the activation of a currently unidentified Gs‐coupled GPCR. Electrogenic L‐Gln uptake depolarizes the cell membrane, leading to the downstream activation of VGCCs. L‐Gln‐stimulated Gs signaling increases intracellular cAMP levels, thereby activating PKA‐ and Epac2‐dependent pathways leading to vesicle release. Oligopeptides are sensed by the sodium‐coupled transporter, PEPT‐1. Similar to L‐Gln sensing, proton‐dependent transport‐induced cell depolarization activates VGCCs fully depolarizing the cell and resulting in hormone release. Both oligopeptides and aromatic amino acids bind CasR on the surface of enteroendocrine cells, triggering the production of IP3 and the induction of calcium release from intracellular stores, leading to cell activation and vesicle fusion. CasR stimulation can also lead to calcium influx through VGCCs as well as transient receptor potential (TRP) channels. Though in this illustration CasR and the postulated L‐Gln sensitive Gs‐coupled GPCR are located at the apical surface, the localization of these receptors to either the basolateral or apical surface is yet to be characterized.
Figure 10. Figure 10. Sensing of products of fat digestion in enteroendocrine cells. Lipid sensing in enteroendocrine cells is largely mediated by GPCRs. Luminal fatty acids may bind Gq‐coupled FFA4 receptors at the apical surface of enteroendocrine cells, though the basolateral or apical localization of FFA4 has not been confirmed. Absorbed fat digestion products are more potent stimuli for gut hormone release. Luminal fat is absorbed by enterocytes, reesterified into triacylglycerides, packaged into chylomicrons, and released into the paracellular space where they may act locally or are taken up by lacteals from which they eventually reach the general circulation. Local lipases release monaclyglycerols and fatty acids from chylomicrons. These are bound by receptors expressed at the basolateral surface of enteroendocrine cells. FFA1 is a Gq‐coupled receptor that binds long‐chain fatty acids. Upon activation, gut hormone release is triggered following IP3‐mediated release of intracellular calcium stores. GPR119 is Gs coupled, and binds both oleoylethanolamide and 2‐oleoyglycerol. GPR119 binding elevates cAMP levels, stimulating PKA‐ and Epac2‐mediated vesicle release.
Figure 11. Figure 11. Bile acids trigger GLP‐1 release by accessing basolaterally located G protein‐coupled bile acid receptors. Epithelium isolated from murine Ileum was mounted in Ussing chambers (A). The concentration of GLP‐1 released from the basolateral surface was measured following either bidirectional (Bi), apical (Ap), or basolateral (Ba) application of the bile acids, taurodeoxycholate (TDCA) and taurolithocholate (TLCA), or the GPBAR1 agonist, GPBAR‐A. In some tissues, an inhibitor of ASBT (ASBT‐I) was applied 10 min before the addition of TDCA. Data represent means +/− SEM. The numbers of tissue samples analyzed is indicated above each bar. Statistical differences were determined on log10‐transformed data via one‐way ANOVA with a post hoc Bonferroni test. * denotes differences from basal conditions, denotes differences from experimental conditions (, P < 0.05; **/††, P < 0.01; ***, P < 0.001). (B‐D) Vascular GLP‐1 secretion was measured from perfused rat intestine. TDCA was applied to either the luminal (lum) or vascular (vasc) side. Bombesin (BBS) was used as a positive control (B,C). Data represent means +/− SEM GLP‐1 secretion from six to seven perfused samples. (D) Baseline and TDCA‐stimulated time‐averaged GLP‐1 release was calculated for experiments performed in (B) and (C). Statistical significance was calculated via one‐way ANOVA followed by a post hoc Bonferroni test (***, P < 0.001). Data were obtained and adapted, with permission, from Brighton et al., Endocrinology, 2015 ().
Figure 12. Figure 12. Sensing of bile acids in enteroendocrine cells. Bile acids from the lumen are absorbed by epithelial cells in the ileum via ASBT and released into the circulation, where they act as key signaling molecules. The GPCR, GPBAR1, binds bile acids circulated to the intestinal epithelium at the basolateral surface of enteroendocrine cells. Gs‐coupled GPBAR1 activation triggers elevations in cAMP levels, leading to PKA/Epac2‐mediated vesicle release. A currently unknown mechanism also triggers elevations in intracellular calcium levels in response to some bile acids, such as TDCA.
Figure 13. Figure 13. Sensing of microbial metabolites in enteroendocrine cells. Microbial fermentation products are stimuli for gut hormone release. Short chain fatty acids (SCFAs) derived from microbial fermentation of indigestible fiber bind the Gq‐coupled receptor FFAR2. Their binding elicits IP3‐mediated increases in intracellular calcium levels, leading to secretory vesicle release. Isovalerate binds Olfr558, increasing cAMP levels, which promotes calcium influx through VGCCs. Indole, a bacterial‐mediated degradation product of dietary tryptophan, exhibits bimodal regulation of gut hormone release. Short‐term exposure to indole reduces potassium efflux through voltage‐gated potassium channels (Kv), widening action potentials, and leading to enhanced intracellular calcium increase during electrical activity. Long‐term exposure, by contrast, deactivates enteroendocrine cells by uncoupling mitochondrial oxidative phosphorylation.
Figure 14. Figure 14. Neural and hormonal factors mediate water balance via regulating PYY release from enteroendocrine cells. AVP, released from the posterior pituitary, and angiotensin II, part of the renin‐angiotensin system, converge upon enteroendocrine cells in the colon to regulate anion secretion and subsequent water uptake. AVP released into the circulation binds the Gs/q‐coupled AVPR1B at the basolateral surface of PYY‐releasing cells. AVPR1B activation stimulates PKA/Epac2‐mediated granule release. Systemically (involving crosstalk of liver, kidneys, and lungs as shown in the figure) or locally produced angiotensin II binds to AT1R receptors at the basolateral surface of PYY‐producing cells. AT1R is Gq‐coupled, and its activation leads to the release of intracellular calcium stores and subsequent secretion of PYY. Secreted PYY may act in both a local paracrine manner, as well as in an endocrine manner after entering the circulation to inhibit chloride secretion from enterocytes by binding Gi‐coupled NPY1R receptors. Gi‐coupled signaling decreases cAMP levels, thereby inhibiting chloride transit through CFTR channels.


Figure 1. Schematic diagram illustrating the microstructure of the intestine. Epithelial cell types are identified along the crypt‐villus axis.


Figure 2. Examples of the crypt‐villus axis. (A) Widefield image of the base of a villus and its accompanying crypts from enzymatically dispersed tissue. (B) Confocal image of a villus and attached crypt. Chromogranin‐A‐positive enteroendocrine cells are stained in red. E‐cadherin, staining adherin junctions between epithelial cells, is stained in green. (Images courtesy of Deborah Goldspink, Reimann/Gribble laboratories, with permission.)


Figure 3. Depiction of the variety of morphologies and signaling modalities adopted by enteroendocrine cells. An open‐type enteroendocrine cell (red) is being contacted by a basolateral extension from a neighboring enteroendocrine cell (blue), facilitating paracrine signaling. Capillaries within the lamina propria mediate endocrine signaling from gut hormones secreted into the paracellular space. A closed‐type enteroendocrine cell is illustrated in grey. Neuroendocrine communication is enabled by neuronal nerve fibers that project into the intestinal mucosa (black). Some enteroendocrine cells form “neuropods” from their basolateral surface, which synapse with neuronal termini (orange).


Figure 4. Key sites of release for prominent gut hormones expressed throughout the gastrointestinal tract. Details are discussed in the text. Abbreviations: CCK, cholecystokinin; GIP, glucose‐dependent insulinotropic polypeptide; GLP‐1, glucagon‐like peptide‐1; Insl5, Insulin‐like peptide 5; PYY, peptide YY; SST, somatostatin; 5‐HT, 5‐hydroxy‐tryptamine.


Figure 5. Coproduction of GLP‐1 and PYY in human colonic enteroendocrine cells. Human colonic epithelial cells were fixed and stained for GLP‐1 and PYY and analyzed by flow cytometry (A). Coproduction of PYY and GLP‐1 was confirmed via immunohistochemistry in cultured human colon epithelium. The scale bar represents 10 µm. Data were obtained and adapted, with permission, from Habib et al., Diabetologia, 2013 ().


Figure 6. Overlap of gut hormone production in enteroendocrine cells purified from the mouse intestine. (A) GLP‐1‐ (“L”) and GIP‐expressing (“K”) cells were sorted from upper/lower small intestinal (“up‐/low‐”) and colonic (“LC”) epithelium by flow cytometry from transgenic mice expressing a fluorescent reporter (venus) under the control of the proglucagon or GIP promoter, respectively. Nonfluorescent control samples were collected from the same regions. Expression of gut hormone transcripts was measured in both fluorescent (“+”) and nonfluorescent (“−”) samples by quantitative PCR and compared to Actb expression. Data are presented as means and SEM values. (B) Fluorescence‐assisted cell sorting (FACS) analysis of small intestinal cells from transgenic mice expressing venus under the control of the proglucagon promoter costained for GLP‐1, GIP, CCK, SST, and PYY. Percentages of venus‐positive cells costaining for gut peptides were calculated and presented in a histogram for cells isolated from the upper and lower small intestine (SI). Data represent the mean and SEM for the number of samples indicated above each bar. (C,D) The frequency of either GLP‐1‐(C) or PYY‐(D)‐producing cells along the intestinal tract was analyzed using FACS. Intestinal epithelial cells were isolated from mice expressing venus under the control of the proglucagon promoter and stained for either GLP‐1 or PYY. The y‐axis represents the cell frequency. Data were obtained and adapted, with permission, from Habib et al., Endocrinology 2012 ().


Figure 7. Diet‐induced changes in enteroendocrine cells. Mice expressing venus under the control of the proglucagon promoter were subjected to either a high fat (HFD) or control chow diet for two weeks. The number of GLP‐1‐expressing cells in colonic epithelium was determined by flow cytometry. HFD mice exhibited a significant decrease in GLP‐1‐expressing cells compared to controls. Differences in enteroendocrine cell numbers between chow and HFD samples were compared using a Student's t‐test: *P < 0.05 (A). Colonic epithelial cells isolated from mice expressing venus under the control of the proglucagon promoter fed a HFD or control diet for 16 weeks were isolated via flow cytometry. The expression of gut hormones (B), prohormone processing enzymes (C), granins (D), and fatty acid binding proteins (E) was assessed via Affymetrix ST1.0 microarray. Venus‐negative cells are represented by striped bars (chow, horizontal; HFD, diagonal). Venus‐positive cells are represented by solid bars (chow, white; HFD, black). Data are represented as mean RMA intensity values and SEM. Expression levels between chow and HFD samples were compared using a Student's t‐test: *P < 0.05, **P < 0.01, ***P < 0.001. Data were obtained and adapted, with permission, from Richards et al., Peptides, 2016 ().


Figure 8. Proposed mechanisms for glucose sensing in enteroendocrine cells. The electrochemical gradient of sodium drives uptake of glucose from the intestinal lumen via the sodium‐glucose cotransporter, SGLT1. SGLT1 activity evokes a small depolarization of the plasma membrane, which activates voltage‐gated calcium channels (VGCCs). The resultant calcium influx further depolarizes the enteroendocrine cell plasma membrane and leads to secretory granule release. Glucose transported into the cell by either SGLT1 or facilitative transport is metabolized, raising cytosolic ATP levels, which closes KATP channels at the cell surface. Inhibition of potassium efflux further depolarizes the cell membrane, contributing to enteroendocrine cell excitation and subsequent gut hormone secretion.


Figure 9. Sensing of dietary protein in enteroendocrine cells. L‐Gln‐stimulated gut hormone release proceeds via a mechanism linked to sodium‐coupled transport as well as by the activation of a currently unidentified Gs‐coupled GPCR. Electrogenic L‐Gln uptake depolarizes the cell membrane, leading to the downstream activation of VGCCs. L‐Gln‐stimulated Gs signaling increases intracellular cAMP levels, thereby activating PKA‐ and Epac2‐dependent pathways leading to vesicle release. Oligopeptides are sensed by the sodium‐coupled transporter, PEPT‐1. Similar to L‐Gln sensing, proton‐dependent transport‐induced cell depolarization activates VGCCs fully depolarizing the cell and resulting in hormone release. Both oligopeptides and aromatic amino acids bind CasR on the surface of enteroendocrine cells, triggering the production of IP3 and the induction of calcium release from intracellular stores, leading to cell activation and vesicle fusion. CasR stimulation can also lead to calcium influx through VGCCs as well as transient receptor potential (TRP) channels. Though in this illustration CasR and the postulated L‐Gln sensitive Gs‐coupled GPCR are located at the apical surface, the localization of these receptors to either the basolateral or apical surface is yet to be characterized.


Figure 10. Sensing of products of fat digestion in enteroendocrine cells. Lipid sensing in enteroendocrine cells is largely mediated by GPCRs. Luminal fatty acids may bind Gq‐coupled FFA4 receptors at the apical surface of enteroendocrine cells, though the basolateral or apical localization of FFA4 has not been confirmed. Absorbed fat digestion products are more potent stimuli for gut hormone release. Luminal fat is absorbed by enterocytes, reesterified into triacylglycerides, packaged into chylomicrons, and released into the paracellular space where they may act locally or are taken up by lacteals from which they eventually reach the general circulation. Local lipases release monaclyglycerols and fatty acids from chylomicrons. These are bound by receptors expressed at the basolateral surface of enteroendocrine cells. FFA1 is a Gq‐coupled receptor that binds long‐chain fatty acids. Upon activation, gut hormone release is triggered following IP3‐mediated release of intracellular calcium stores. GPR119 is Gs coupled, and binds both oleoylethanolamide and 2‐oleoyglycerol. GPR119 binding elevates cAMP levels, stimulating PKA‐ and Epac2‐mediated vesicle release.


Figure 11. Bile acids trigger GLP‐1 release by accessing basolaterally located G protein‐coupled bile acid receptors. Epithelium isolated from murine Ileum was mounted in Ussing chambers (A). The concentration of GLP‐1 released from the basolateral surface was measured following either bidirectional (Bi), apical (Ap), or basolateral (Ba) application of the bile acids, taurodeoxycholate (TDCA) and taurolithocholate (TLCA), or the GPBAR1 agonist, GPBAR‐A. In some tissues, an inhibitor of ASBT (ASBT‐I) was applied 10 min before the addition of TDCA. Data represent means +/− SEM. The numbers of tissue samples analyzed is indicated above each bar. Statistical differences were determined on log10‐transformed data via one‐way ANOVA with a post hoc Bonferroni test. * denotes differences from basal conditions, denotes differences from experimental conditions (, P < 0.05; **/††, P < 0.01; ***, P < 0.001). (B‐D) Vascular GLP‐1 secretion was measured from perfused rat intestine. TDCA was applied to either the luminal (lum) or vascular (vasc) side. Bombesin (BBS) was used as a positive control (B,C). Data represent means +/− SEM GLP‐1 secretion from six to seven perfused samples. (D) Baseline and TDCA‐stimulated time‐averaged GLP‐1 release was calculated for experiments performed in (B) and (C). Statistical significance was calculated via one‐way ANOVA followed by a post hoc Bonferroni test (***, P < 0.001). Data were obtained and adapted, with permission, from Brighton et al., Endocrinology, 2015 ().


Figure 12. Sensing of bile acids in enteroendocrine cells. Bile acids from the lumen are absorbed by epithelial cells in the ileum via ASBT and released into the circulation, where they act as key signaling molecules. The GPCR, GPBAR1, binds bile acids circulated to the intestinal epithelium at the basolateral surface of enteroendocrine cells. Gs‐coupled GPBAR1 activation triggers elevations in cAMP levels, leading to PKA/Epac2‐mediated vesicle release. A currently unknown mechanism also triggers elevations in intracellular calcium levels in response to some bile acids, such as TDCA.


Figure 13. Sensing of microbial metabolites in enteroendocrine cells. Microbial fermentation products are stimuli for gut hormone release. Short chain fatty acids (SCFAs) derived from microbial fermentation of indigestible fiber bind the Gq‐coupled receptor FFAR2. Their binding elicits IP3‐mediated increases in intracellular calcium levels, leading to secretory vesicle release. Isovalerate binds Olfr558, increasing cAMP levels, which promotes calcium influx through VGCCs. Indole, a bacterial‐mediated degradation product of dietary tryptophan, exhibits bimodal regulation of gut hormone release. Short‐term exposure to indole reduces potassium efflux through voltage‐gated potassium channels (Kv), widening action potentials, and leading to enhanced intracellular calcium increase during electrical activity. Long‐term exposure, by contrast, deactivates enteroendocrine cells by uncoupling mitochondrial oxidative phosphorylation.


Figure 14. Neural and hormonal factors mediate water balance via regulating PYY release from enteroendocrine cells. AVP, released from the posterior pituitary, and angiotensin II, part of the renin‐angiotensin system, converge upon enteroendocrine cells in the colon to regulate anion secretion and subsequent water uptake. AVP released into the circulation binds the Gs/q‐coupled AVPR1B at the basolateral surface of PYY‐releasing cells. AVPR1B activation stimulates PKA/Epac2‐mediated granule release. Systemically (involving crosstalk of liver, kidneys, and lungs as shown in the figure) or locally produced angiotensin II binds to AT1R receptors at the basolateral surface of PYY‐producing cells. AT1R is Gq‐coupled, and its activation leads to the release of intracellular calcium stores and subsequent secretion of PYY. Secreted PYY may act in both a local paracrine manner, as well as in an endocrine manner after entering the circulation to inhibit chloride secretion from enterocytes by binding Gi‐coupled NPY1R receptors. Gi‐coupled signaling decreases cAMP levels, thereby inhibiting chloride transit through CFTR channels.

 

Teaching Material

A. E. Adriaenssens, F. Reimann, F. M. Gribble. Distribution and Stimulus Secretion Coupling of Enteroendocrine Cells along the Intestinal Tract. Compr Physiol 8: 2018, 1603-1638.

Didactic Synopsis

Major Teaching Points:

  • The intestinal epithelium provides a critical interface between the outside world and the body that relays information about an organism's nutritive status to other organ systems using an array of endocrine signals and neural networks.
  • Enteroendocrine cells collectively form the body's largest endocrine organ, and their hormonal signals regulate key physiological processes ranging from local gut motility to central appetite control.
  • Many individual enteroendocrine cells are capable of releasing several gut hormones.
  • The enteroendocrine cell population is plastic, and relative numbers of enteroendocrine cells and their expression of gut hormones appear to be influenced by changes in diet, pathology, and gastric bypass surgery.
  • Enteroendocrine cells sense and respond to changes in their environment via G protein-coupled receptors, ligand-gated ion channels, and electrogenic nutrient transporters that initiate signaling cascades mediating gut hormone release.

Didactic Legends

The figures—in a freely downloadable PowerPoint format—can be found on the Images tab along with the formal legends published in the article. The following legends to the same figures are written to be useful for teaching.

Figure 1 Teaching points: The intestinal mucosa, comprising the epithelium and lamina propria, is encased by connective tissue (the submucosa) and layers of muscle. Blood vessels and nerve endings are embedded within the mucosa, allowing for neuroendocrine exchange between the gut and the rest of the body. The absorptive surface of the intestine is greatly increased by villi, which decrease in density along the cephalocaudal axis. Crypts at the base of each villus house stem cells that give rise to both the absorptive and secreting cells that populate the epithelium. Absorptive cells include enterocytes and M cells. Secreting cells include progenitor, Paneth, goblet, tuft, and enteroendocrine cells.

Figure 2 Teaching points: Crypts lie at the base of the crypt-villus axis, and their progenitor cells give rise to absorptive and secreting cells that populate the villus. Enteroendocrine cells can be found in both the crypt and villus, as shown here in red (B).

Figure 3 Teaching points: Enteroendocrine cells exhibit a wide array of morphologies. Many enteroendocrine cells are “open-type” with apical extensions that extend into the gut lumen. Some enteroendocrine cells lack these apical extensions, and are termed “closed-type.” The secretion of gut hormones from enteroendocrine cells proceeds via several pathways. Once triggered, enteroendocrine cells release their secretory cargo into the paracellular space. Here, released gut hormones are either taken up from the interstitium by capillaries—constituting endocrine signaling—or act directly on their cognate receptors expressed at the surface of neighboring cells, thereby providing paracrine regulation. Enteroendocrine cells can interdigitate with neighboring cells via cytoplasmic extensions from their basolateral surface, facilitating paracrine crosstalk. Released gut hormones can also interact with local afferent nerve fibers, relaying their information centrally. In some instances, nerve termini synapse with enteroendocrine cells using structures called “neuropods” at the basolateral surface of the cell. These structures are identified in the aforementioned formal figure legend.

Figure 4 Teaching points: Gut hormones exhibit unique secretion profiles, which are dependent on the distribution of enteroendocrine cells producing a particular gut hormone along the cephalocaudal axis as well as meal patterns and nutrient absorption. Most gut hormones are predominantly produced in one area of the gut, as outlined in the diagram. Gut peptide abbreviations are decoded in the formal figure legend.

Figure 5 Teaching points: (A) this figure demonstrates that the majority of GLP-1-producing cells in the human colon also produce PYY, and vice versa. Part (B) confirms at the cellular level that GLP-1 and PYY are coexpressed in enteroendocrine cells.

Figure 6 Teaching points: This figure illustrates the multihormonal expression pattern of enteroendocrine cells. GLP-1- and GIP-expressing enteroendocrine cells were purified from transgenic mice. These cell populations were tested for the co-production of other gut hormones at the mRNA (A) and protein (B) levels. In both GLP-1- and GIP-positive cells, a number of gut hormones were coexpressed at the transcript level. In isolated GLP-1-expressing cells, the coproduction of GIP, CCK, and PYY protein was confirmed using immunoprobes followed by FACS. Together these data confirm that enteroendocrine cells produce more than one gut peptide. In parts C and D, the frequency of either GLP-1 or PYY positive cells was assessed along the cephaocaudal axis of the intestine. The number of GLP-1-expressing cells co-staining for PYY increased in increasingly distal regions, indicating regional-specificity to gut hormone overlap.

Figure 7 Teaching points: These data demonstrate plasticity of the enteroendocrine cell population. In this study, the effect of diet on GLP-1-expressing cells was assessed. The graph in part (A) indicates that mice kept on a high-fat diet (HFD) exhibit altered numbers of enteroendocrine cells in the colon. Gene expression studies on GLP-1-expressing cells from the intestinal epithelium showed that cells isolated from mice kept on a HFD express lower levels of enteroendocrine-enriched genes, many with proposed functions within secretory vesicles, indicating that the function of these enteroendocrine cells could be impaired.

Figure 8 Teaching points: Glucose uptake from the intestinal lumen results in the electrical activation of enteroendocrine cells. This electrical excitation is primarily mediated by the electrogenic activity of SGLT-1—the primary transporter mediating glucose absorption from the gut lumen. Metabolism-dependent electrical excitation through the closure of ATP-sensitive potassium channels can also contribute to enteroendocrine cell activation and hormone release.

Figure 9 Teaching points: Products from protein digestion stimulate a variety of cell signaling pathways leading to gut hormone release from enteroendocrine cells. Similar to glucose sensing in enteroendocrine cells, the electrogenic activity of transporters for L-Gln and oligopeptides excites the enteroendocrine cell, which triggers downstream ion channels that further depolarize the cell membrane, leading to vesicle release. GPCR signaling also plays a role in sensing protein digestion products. CasR, a Gq-coupled receptor that also activates voltage gated ion channels, binds small oligopeptides as well as single aromatic amino acids. A yet unidentified GPCR coupled to Gs G proteins underlies the sensing of L-Gln.

Figure 10 Teaching points: The absorption of dietary fat is an important step for lipid-induced gut hormone release.

Figure 11 Teaching points: These data were obtained using techniques that allow for the evaluation of the potency of apically versus basolaterally applied stimuli in triggering gut hormone release. From this data we can see that basolaterally (vascularly) applied bile acids (TDCA and TLCA) are more effective stimuli for GLP-1 release from intact ileal epithelium than when they are applied to the apical (luminal) surface. The blockade of bile acid transport from the luminal side significantly inhibits the GLP-1 response to TDCA, indicating that bile acids stimulate GLP-1 release predominantly postabsorption from the basolateral surface.

Figure 12 Teaching points: Bile acid-stimulated gut hormone release is mediated by absorbed bile acids that are circulated to the gut epithelium. Here they bind the Gs-coupled GPCR, GPBAR1, which stimulates cAMP elevation and downstream PKA/Epac2 pathway activation. Another mechanism underlying bile acid-dependent gut hormone release proceeds by a currently uncharacterized mechanism stimulating extracellular calcium influx.

Figure 13 Teaching points: SCFAs are products of microbial fermentation, and include propionate, acetate, and butyrate and isovalerate. Enteroendocrine cells express GPCRs, which bind SCFAs. SCFA-stimulated signaling mechanisms are outlined in the formal figure legend. Other products of microbial metabolism include indole, a derivative of dietary tryptophan. Indole elicits both stimulatory and inhibitory effects on gut hormone release, depending on the length of time enteroendocrine cells are exposed to indole, as outlined earlier.

Figure 14 Teaching points: Enteroendocrine cells integrate signals from the brain as well as the periphery to regulate water balance in the body. The colon is a key site for water absorption. Both centrally derived AVP, and angiotensin II from the renin-angiotensin system inhibit anion secretion from the colon by stimulating PYY release from enteroendocrine cells. This is achieved through the activation of their cognate receptors expressed at the basolateral surface of PYY-producing enteroendocrine cells, as outlined earlier in the formal figure legend.

 


Related Articles:

Distribution of Gut Peptides
Endocrine Cells of the Gut
Teaching Material

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How to Cite

Alice E. Adriaenssens, Frank Reimann, Fiona M. Gribble. Distribution and Stimulus Secretion Coupling of Enteroendocrine Cells along the Intestinal Tract. Compr Physiol 2018, 8: 1603-1638. doi: 10.1002/cphy.c170047