Open access peer-reviewed chapter
Abstract
Fruit fly, Drosophila melanogaster is a most powerful animal model for exploring fundamental biological processes and modeling molecular and cellular aspects of human diseases. It provides the flexibility and tool box with which scientists can experimentally manipulate and study behavior as well as gene expression in specific, defined population of cells in their normal tissue contexts. The utility and increasing value of a sophisticated genetic system of flies, the tool box available for studying physiological function, functional imaging, neural circuitry from gut to brain, taste receptors expression and controlling gene expression by determining the specific cells in the intestine, makes fly gut the most useful tissue for studying the regulation of feeding behavior under changing internal state. To understand the intestine and its connectivity with the brain, Drosophila has proved an ideal model organism for studying gut brain axis aspects of human metabolic diseases. Various markers and fly lines are available to characterize the expression of transgenes in the intestine. The newly generated genetic tools aim to streamline the design of experiments to target specific cells in intestine for genetic manipulations based on their type and location within physiologically specialized intestinal regions. This chapter will be useful for understanding post-ingestive sensing system that mediate food preferences and to investigate fundamental biological processes and model human diseases at the level of single cells in the fly gut. Furthermore, the utility of adult fly gut can be extended to the study of dietary and environmental factors relevant to health and disease by screening for cells and micro circuits stimulated by internal state or the consumption of various nutrients.
Keywords
- gut brain axis
- Drosophila
- taste receptors
- enterocytes
- enteroendocrine cells
1. Introduction
Mammalian central nervous system (CNS) consists of brain and spinal cord. The neuronal network extends from the brain to all over the body and various neurotransmitters help transmit the message to target cells in different tissues. The part of the nervous system located in our gut is called the enteric nervous system (ENS). In all animals, gut brain axis is a lesser known nervous system so far. Our gut or “second brain” (ENS) chemically connects with the brain through neurons, secreted chemicals like hormones and neurotransmitters that send messages to the brain. The enteric nervous system’s network of neurons and neurotransmitters extends along the entire digestive tract – it starts from the esophagus to the stomach and intestines, and down to the anus. The “gut microbiome” comprises of microorganisms living in the gut (bacteria, viruses, and fungi) that can affect the chemical messages that pass between the gut and the brain. Microorganisms residing in the gut help regulate the body’s immune response. Since, the brain and the gastrointestinal (GI) system are intimately connected, therefore, they play key roles in certain diseases and to maintain our overall health to regulate cognitive and digestive behavior. The bidirectional communication between the brain and digestive system hence, are opening up avenues to think about diseases considering this angle.
Gut has recently become a subject of research in medical sciences wherein subjects with depressive symptoms, Parkinson’s and Alzheimer’s disease, autism, amyotrophic lateral sclerosis, multiple sclerosis, pain, anxiety and other neurodegenerative conditions are beginning to be looked to see what is going on in the gut. Effects of conditions like ulcers, constipation, and other GI problems have been a focus of research on aspects of brain functioning. The enteric microbiota impacts the gut brain axis (GBA), interacts locally with the intestinal cells and ENS as well as with the CNS through neuroendocrine and metabolic pathways. These studies suggest that GBA plays a vital role in maintaining mental health and can affect the feeding behavior when nutrient detection or absorption does not function properly as in case of metabolic conditions.
To ensure stability in the internal environment of body and drive adaptive changes, control mechanisms are key to animal’s survival. Recently GI has been recognized as a major source of signals modulating feeding behaviors, food intake, metabolism, insulin secretion and energy balance. Through its interaction with microbiota, it can shape our physiology and behavior in complex and sometimes unexpected ways. A growing scientific community has exploited the genetic amenability of
2. Drosophila as an emerging model system for studying gut: comparison of human and fly gut
Easy genetic manipulation and effortless genetic tools make flies an insect of choice to study inter organ neuronal signaling including gut-brain axis neuronal connectivity. To understand human metabolic diseases and how a GI play a key role there is a recent focus of research. Some progressive studies also draw a link between neurodegenerative disorders and gut microbiota of humans and other insects.
Fruit flies have a simple and similar to humans- gut system (Figure 1). In mammals including humans, the esophagus (
Figure 1.
Comparison between human and
The adult
Figure 2.
Fly gut anatomy. (A) the
The fly midgut has been segmented into the anterior, middle and the posterior midgut (Figures 1,2A and B). It has been further subdivided morphologically and molecularly into 10–14 regions (Figure 2B) [11, 12, 13]. Midgut regionalization has been seen in the muscles, trachea and neurons that surround it [12, 13, 14, 15]. Physical properties (
The Malpighian tubules release at the junction between the midgut and hindgut (Figures 1,2A and B). The water/ion exchange occurs in the hindgut which consist of pylorus (a second valve-like structure), ileum, and rectum [7, 8]. The muscles that surround the epithelium in flies are striated, as opposed to the smooth muscles found in mammalian intestines [18]. An outer layer of longitudinal muscles found surrounding the midgut. Circular muscles are found to be present throughout the fly tract. Physiology of the intestine is maintained and regulated by autonomic innervation and by hormones. The tracheal system forms a branched structure surrounding the gut during development [15] and may influence epithelial regeneration in the adult. Owning to similarity of flies and human gut, we will be discussing further how gut of the flies is handled and controlled to understand about neural circuitry drawing it closer to the brain and diseases related to intestinal illnesses.
2.1 Intestinal anatomy of Drosophila and human
Both humans and fly intestines share similar tissue, anatomy and physiological function [19, 20]. Their gut are of endothelial origin in nature [21, 22] and comprise of an epithelial monolayer of columnar or cuboidal ECs. A series of sequential depressions called the crypts of Lieberkühn, along the small and large intestine, and protruding villi along the internal surface of the small intestine in mammalian intestinal epithelium maximize its surface area [23]. Extensive folding has not been reported in the
In both flies and mammals, the epithelial monolayer is associated on its basal side on an extracellular collagenous matrix (known as basement membrane) [29]. A checkerboard of innervated and trachea-oxygenated longitudinal and circular muscles tissue underneath the basement membrane in flies drive the peristaltic movements [30] (Figure 2C). Mammalian intestine has a similar organization of intestinal external musculature in the outer layers where musculature is also innervated and oxygenated by a plexus of vasculature [31, 32]. Layers including, the submucosa, (a dense layer of connective tissue containing nerves and lymphatic and blood vessels); muscularis mucosae (an additional muscle layer); and the lamina propria underling the intestinal epithelium and contains connective tissue, lymph nodes (Peyer’s patches), immune cells (leukocytes, and dendritic and mast cells), vessels and myofibroblasts [33], fill the space between the outer musculature and the basement membrane in mammals.
2.2 ISCs, ECs, and EE cells of fly gut
About 65% of human-disease causing genes are shared as a functional homolog in fruit flies. This shows conservation of genes and function at an evolutionary level. Fundamental processes such as digestion is also conserved from flies to humans.
The lineage of fly posterior midgut with only one type of mature absorptive cell and one main type of secretory cell is very simple. Although asymmetric ISC divisions in the fly midgut produce transient cells (EBs), these cells do not undergo further cell division and remain close to the ISCs before maturation. Fly midgut ISCs are situated basally and are broadly dispersed in the intestinal epithelium. The cellular composition and regeneration in
2.3 The fly intestine functions
Like the regional specialization of digestive functions, the expression of digestive enzymes has also been found to confined to specific segments of the digestive tract in flies [12, 46, 47]. In addition to its roles in nutrient extraction and utilization, the digestive tract responds to the food and bacteria in its lumen. Digestion takes place in fly midgut [48] which can be further modulated by various factors like temperature, redox potential, pH, and intestinal transit [8, 48]. It has been shown that the expression and activity of digestive enzymes are tightly regulated in many insects like enzymes involved in the breakdown of sugars in flies are enriched in anterior (R1/R3) portions of the adult midgut and Peptidase genes express more posteriorly [47].
The enzymatic activity of the intestine is a key factor determining availability of certain nutrients. A substantial reduction of intestinal digestive enzyme activities (trypsin, chymotrypsin, aminopeptidase, and acetate esterase) has been reported in flies lacking EE cells [49]. Though not extensively investigated in
2.3.1 Role in nutrients absorption
Changes in the expression of p38 kinase or the Atf3 and Foxo transcription factors cause accumulation of neutral lipid in ECs [58, 84]. It has been shown that neutral lipid increase following depletion of the EE hormone Tk [85], or in sterile female flies after mating [86]. It has been suggested that activation of intestinal lipogenesis is key to survival in diet-restricted flies. Indeed, nutrient scarcity induces expression of the sugar sensor transcription factor
2.3.2 Intestinal pH
Many animals generate localized regions of low pH inside the intestinal lumen to facilitate protein breakdown, absorption of minerals and metals, and limit the survival of ingested microbes. While mammalian digestion takes place in acidic conditions, insect digestion occurs at neutral or basic pH including
The contribution of five ion transporters enriched in the acidic region have been studied [88]. These include: the potassium/chloride symporter Kazachoc (Kcc), a member the Slc12 family of electroneutral cation-chloride transporters (express in intestine) [91, 92]; the Slowpoke pore-forming subunit of a calcium-activated K+ channel (express in neurons, muscles, tracheal cells, and two types of midgut ECs in the copper and iron cell regions) [93]; the ligand-gated chloride channel pHCL-2 which, in addition to regulating fluid secretion in malpighian tubules (express in the copper cell, iron, and large flat cell regions of the midgut) [94, 95]; the carbonic anhydrase CAH1; and the bicarbonate/chloride exchanger CG8177, belonging to the Slc4a1–3 subfamily of anion exchangers (express in a specific midgut pattern similar to that of pHCl-2) [96]. Collectively, these findings suggest that the transport of H+, Cl−, K+, and HCO3− contributes to acid generation in the
2.3.3 Water and osmolytes
Flies extract water from their diet to maintain hydration and ionic balance. This compensates for substantial water loss resulting from metabolic and physiological processes. Although malpighian tubules are important for this process, but intestine also contributes. Water absorption from the food occurs in the insect midgut and in rectal pads of rectum [8]. The rectal pads are also the crucial site for reabsorption of ions. Ions and water can cross the intestinal epithelium through or between cells and their transport play an important role in the maintenance of ion gradients that sustain active transport in the intestinal epithelium. The scanning ion-selective electrode technique (SIET) provides a way to probe intestinal gradients for ions such as K+, Na+, H+, or Cl− [24, 97]. K+ and Na+ absorption occur largely in the large flat cell and posterior regions of the midgut and, also in the anterior hindgut in the case of Na+ [97]. The two
2.3.4 Metal ions
Metal ions such as copper, iron and zinc are essential micronutrients required for the correct folding and activity of a broad range of enzymes. The contribution of the intestine to metal homeostasis has not been extensively investigated but midgut regions, the Cu cell and Fe regions, are the most proposed sites of metal ion absorption. The Cu cell region turns bright luminescent orange upon Cu ingestion due to the fixation of copper by metallothionein [101, 102], and appears to be an important site of accumulation of ingested radioactively labeled Cu [101, 103]. The Fe cell region in R4a stains by Prussian blue and also accumulates exogenously administered radioactive Fe [101, 104]. Many studies have confirmed the roles for the Cu/Fe regions by exploring the molecular machinery involved in the intestinal uptake, intracellular trafficking, and efflux of metal ions.
2.3.5 Transit and excretion
Nutrient extraction and utilization may get affected by the passage of food along the alimentary canal and by its subsequent excretion. Transport of food to travel the entire length of the digestive tract takes less than 1 hour [105] in flies. As suggested, the amount of food retained in the crop is much larger in starved flies then refed flies than in flies fed
3. ENS and GBA
The ENS in humans, equivalent to GBA in flies is a part of the peripheral nervous system (PNS) that governs the running of the neurons which influence the GI. It is exploited nowadays in flies to understand more about how the two organs affect one another and lead to decisions regarding appetite, feeding mechanisms, taste preference and how to deal with hunger and satiety. How taste receptors detect different nutrients in the gut remains to be explored.
3.1 Gustatory receptors in gut
3.1.1 Humans
In humans, G-protein coupled receptors (GPCR) are involved in detection of five common tastes- sweet, salty, bitter, sour and umami. T1R family of taste receptors determine sweet and umami flavor. T2R family includes the bitter receptors [110]. Sweet taste receptors (T1R family) are found in intestinal tract as well as EE cells [111]. T1R1, T1R2, T1R3 and α-gustducin are expressed in the stomach, intestine and colon of humans and mice [112]. Cells of duodenal villi show co-localization of T1R1, T1R3 and α-gustuducin. Mammalian Gustducin protein is involved in bitter and sweet taste signaling and detection [113, 114]. α-subunit of gustducin is expressed in gastric cells and may play a role in nutrient detection [115, 116, 117] in rats and mice. Present in the mucosal lining of mammals and taste cells of the epithelium suggesting its possible role in taste uncovering on exposure to luminal contents [118]. Expression of T2R receptors in mouse GI tract including mT2R119 and mT2R108 has been looked into. mT2R119 expression is found in gastric and intestinal tissues, tongue and liver [115] like mT2R134 [118]. mT2R108, mT2R138 are present in the fundus, antrum, duodenum and tongue (not in liver) [119].
It has been found that sweet taste receptors stimulated in rat intestine influence and increase glucose absorption [120] through GLUT (glucose transporter) [121]. Presence of sugar in the diet galvanizes ECs into action to release hormones which in turn activate SGLT (Na+ / glucose cotransporter) [122, 123]. Similar results are shown in sheep where sugar receptor / sensor present on the luminal membrane stimulates SGLT1 via cAMP and G-protein dependent pathway [124]. Equine T1R2 (homologous to cows and pigs) is expressed on the luminal membrane of EE cells in the small intestine. In response to increased sugars, T1R2 along with T1R3, stimulates SGLT1 and enhances the ability of gut to absorb more glucose [125]. Analysis of in vitro line of ECs suggest that T1R2 and T1R3 detect sweet taste and exposure to sucralose increases release of hormones such as GIP (Gastric Inhibitory Polypeptide) and GLP (Glucagon like peptide) which further activate glucose activation and metabolism. An inhibitor of these receptors inhibits glucose metabolism suggesting these receptors present in the gut alter feeding mechanisms and post-ingestion decisions [120, 121].
3.1.2 Flies
Gustatory system in
3.2 Gut-brain Neural Circuits
3.2.1 Humans
Nutrient signaling and sensing are fundamental processes that animals including humans and flies undergo [131]. Proper coordination and communication between gut and brain is necessary to regulate metabolic homeostasis and physiology in all animals. In this regard, many research groups have shown the role of enteric neurons and endocrine signals as important mediators of these processes. The way the enteric nervous system communicates to the brain via neural circuits is a multifaceted question and poorly explored. In mammals such as humans, alterations in neuropeptides and brain – gut hormone levels can derail people otherwise on the path to a healthy life. These changes can also lead to diseases such as neurodegenerative diseases, metabolic syndrome and diabetes [132].
Gluconeogenesis is a biochemical pathway by which animals make sugars from non-carbohydrate precursors and sources [133]. It is used to regulate homeostasis and a stable internal state in post – fed state [134]. Studies in rats showed that stimulation of intestinal gluconeogenesis (IGN) sends a signal from sodium – glucose co- transporters present at the intestinal mucosa to the brain, initiating a neural gut – brain axis [135, 136, 137]. Diets rich in protein [138, 139, 140] and fiber [141] promote IGN stressing on the importance of nutrient sensing for initiating several gut – brain axis [137]. It has been found that μ – opoid receptors (MOR) regulate IGN. These receptors (present in the nerves in the portal vein wall) react to neuropeptides to stimulate a gut – brain neural circuit that affects IGN, hunger and satiety mechanisms [141]. Further analysis of MOR deficient mice shows the role of MORs in regulating food intake, referred as “reward” system [142, 143]. Analyses of MOR-knockouts (MOR-KO) demonstrate how they play a role in managing satiety effects of alimentary proteins, through a neural gut-brain circuit [140].
Vagus nerve (VN; pneumogastric nerve) is the longest cranial nerve [144] in humans which runs from the medulla oblongata in brain to colon in GI [145]. It innervates other structures as well such as larynx, pharynx, heart and lungs thus affects digestive, cardiovascular and respiratory system – all at one [146]. Vagal efferent send down signals from the brain to gut, which accounts for about 10% – 20% of all the nerve fibers. Remaining 80% is accounted for by the vagal afferents carrying information from the gut to the brain [147]. Vagal sensory neurons in the GI keep an eye on stomach volume and luminal contents through different neural circuits [148]. VN contains and branches into several sensory neurons (~2300 in mouse) that further innervate and render support and supply to other internal organs. A variety of sensory neurons, one side facing the brainstem and the terminal one facing the organ such as GI [149] have been revealed. Free terminals of vagal afferents are rooted within lamina propria of intestinal villi [148]. Some mammalian models like in cat and rat, it has been shown how these sensory neurons detect different nutrients in diets with the help of unambiguous and explicit fibers [150, 151, 152]. Vagal afferent endings in the intestine express several mechanosensitive as well as chemical receptors [153]. Glucagon- like peptide 1 (GLP1) is a gut hormone receptor that intercedes the nutrient sensing mechanism via VN [154]. GLP1R (GLP1 receptor) is present in many cells [155]. Agonists for GLP1R show how it affects brain further proving its presence in both, gut and brain [156]. Another receptor of vagal afferents, GPR65 near the intestinal villi, plays a role in nutrient detection drawing attention to how these sensory neurons are a part of the gut – brain axis [157, 158]. It detects serotonin and impact gut motility [147]. Such receptors detect several hormones present in the gut, like choleocystokinin (CKK), ghrelin and leptin which play a role in the regulation of hunger and satiety [159, 160, 161]. Because of its role in gut motility and mobility, VN and its afferent neurons present in the gut play a role in Intestinal Bowel Syndrome (IBS) [162] and new treatment plans around the same are being looked at in rat [162] and mice [158] models.
To take the findings in vagal nerves forward, nerves allowing communication of cNST (caudal nucleus of the solitary tract) with gut were focused on. Information about sugar detection to cNST via gut – brain axis is a topic of research nowadays. In live mice, it is noticed that glucose detection by cNST is robust and VN transaction silences that activation [163]. Nodose ganglion of vagus nerve when silenced prevents the sugar preference of cNST [163] suggesting the presence of a physical gut – brain axis. It has been shown that inactivation of sugar-activated cSNT prevents the mice to choose sugar from water or an artificial sweetener [163]. This study specifically calls attention to how organisms have paths for detecting nutrient signals, sensing them in the diet and also have circuitries to carry forward the signals and communicate with the rest of the body, purposely the brain.
3.2.2 Drosophila
With the help of several markers and reporter genes, it is found that
In the anterior portion of adult and larval midgut, serotonin positive neurites and various neuropeptides including Akh, Dh44, Myosuppressin, and possibly Allatostatin C and FMRFamide (or an FMRFamide-like peptide such as the NPY-like neuropeptide short neuropeptide F [sNPF]) [131, 168, 172, 180, 181, 182] have been described suggesting chemical diversity of enteric innervation. Four serotonergic neurons are found to innervate the enteric nervous system in fly larva. These neurons project from the antennal nerve (AN) near the SEZ and extend throughout the ENs [14]. These projections end at the anterior region of the midgut and are primarily around the proventriculus region and the foregut. These innervations are considered structurally analogous to the mammalian VN because of similar projections from the brain to the different structures of the foregut. It is yet to be seen if there are functional similarities as well [172].
Pigment- dispersing factor (Pdf), Ion transport peptide, and Proctolin positive neurites have been reported in the larval and adult hindgut [169, 183, 184, 185, 186]. All three innervated regions receive insulinergic innervation from the CNS; the
Figure 3.
Innervation of adult
Functional studies of insect innervation have primarily focused on the control of peristalsis and peptide hormone secretion so far. Studies of peristaltic regulation in flies have primarily concerned the effects of neuropeptides (Allatostatins, Myosuppressin, or Drosulfakinins) on
Recently epithelial roles for gut-innervating neurons e.g. role in the control of fluid balance have been revealed by a semi-automated analysis of defecation behavior in adult flies, providing quantitative readouts for food intake, fluid/ion balance, and intestinal transit [14, 193]. The HGN1 neurons (2–5 CNS neurons located in the posterior segments of the abdominal ganglion) innervate the hindgut and the rectum (Figure 3), with a subset of their neurites projecting through the visceral muscles to reach the underlying epithelium [14]. HGN1 neuronal silencing experiments resulted in increased defecation rate. These neurons are shown to be required for the post-mating changes in intestinal fluid retention due to their epithelial innervation. It has been established that HGN1 neurons and the Pdf hindgut-innervating neurons have their direct action on the hindgut and anal sphincter muscles [192]. A role for gut-innervating neurons in the maintenance of epithelial turnover has also been suggested by the finding of anatomical proximity between enteric neurites in the posterior midgut and adult somatic intestinal progenitors, and the reduced ISC to EC differentiation resulting from downregulating Hedgehog (Hh) signaling (albeit pan-neuronally) [194]. The more anterior innervation of the proventriculus may also play a role in maintaining gut permeability. This is inferred from the finding that inactivation of a relatively broad subset of neurons, including a subset of anterior midgut-innervating neurons results in an abnormal proventricular structure, increased permeability of the epithelial barrier, and increased susceptibility to oral bacterial infection: all suggestive of defects in the production of peritrophic matrix [175].
In adult flies, inactivation of insulin-producing neurons results in contrasting effects on the hyperphagic response triggered by nutrient scarcity. Silencing of the insulin-producing cells of the brain PI that innervates the anterior midgut lowered this response, whereas silencing of the hindgut-innervating Ilp7 neurons increased it, and also resulted in higher circulating glucose [14, 195]. Not much is known about the importance of sparse sensory innervation of the intestine. One remarkable exception are the pharyngeal taste neurons.
Post-ingestive sensory feedback from the gut has been assumed to inhibit feeding based on work in other insects for example severing the recurrent nerve or the medial abdominal nerve, which transmit information from the gut to the brain, results in overconsumption in blowflies [199]. Work done in flies lends support to this idea; whereas severing the medial abdominal nerve did not disturb food consumption, severing the recurrent nerve elevated consumption of sucrose but not water or bitter tasting solutions [200]. The existence of neuronal stretch receptors on the gut that monitor the volume of ingested food is supported by both neurophysiological and anatomical data in numerous other insects [4, 80, 199, 201]. However, the existence and molecular nature of these receptors in
4. Enteroendocrine hormones, neuropeptides and signals
4.1 Humans
The gut–brain transmission systems involve both circulation-based endocrine-like and neuronal communication routes [205]. Neuropeptides (transmitters or neurosecretory) act as messenger molecules of enteric, sensory, autonomic and central neurons. Several peptide families have been found in both brain and gut. They act as neuropeptides and/or gut hormones and have significantly contributed to the understanding of gut brain interaction. Central and peripheral neurons together with EE cells in the GI tract and other endocrinologically active cells produce variety of peptides [206, 207, 208, 209, 210, 211] including hormones peptide YY (PYY) and pancreatic polypeptide (PP); neuropeptide Y (NPY), on the other hand [212]. These and other peptide families represented by glucagon-like peptide (GLP), ghrelin, cholecystokinin (CCK), corticotropin-releasing factor (CRF), leptin, osteocalcin and insulin (the last three are extra intestinal endocrine peptides) act on specific and genetically related groups of receptors that are expressed by distinct cells in the periphery and CNS. For their functional roles, endocrine peptides and neuropeptides are relevant for the regulation of digestion, control of food intake, metabolic homeostasis, and the impact of GI signals on sensation, emotion, affect, and cognition. Disturbances of the gut microbiota–brain axis result in changes of the expression and activity of many neuropeptides and their receptors in the CNS. Neuropeptides are therefore important secondary messengers of gut microbes in cerebral neuro circuitries that mediate the alterations in brain function and behavior that take place in response to changes in the GI microbial community [212]. Together it is emerging that neuropeptide systems such as NPY, CRF, ghrelin, and brain-derived neurotrophic factor (BDNF) play a particular role in the cerebral manifestations of gut microbiota perturbations.
In rats, choleocystokinin and glutamate neuro-epithelial circuit makes the communication between the brain and intestinal lumen possible stressing on the importance of a physical gut – brain axis [213]. It has been proposed that the physical connection between vagal nodose neurons and EE cells is present in rats which leads to regulation of gastrointestinal functions [213]. Intragastric nutrient infusion and optical readings of AgRP (Agouti-related protein) neurons in live and awake mice [214] suggest AgRP neurons affect long term homesostasis and energy balance of the body and do not get altered by minute changes in nutrient levels [215]. AgRP neurons get inhibited by high levels of satiation signals such as CCK and PYY (peptide YY). Receptors for both CCK and PYY are expressed by vagal afferent neurons’ terminals that innervate GI [159] suggesting a possibility of a physical connection between the gut and brain carrying the message from one point to another [216].
4.2 Drosophila hormones and neuropeptides
Apart from using neurons, fly intestine can also communicate with other organs through systemic signals. Intestinal physiology is modulated by both extrinsic hormonal signals (emanating from endocrine glands, neuroendocrine structures, or organs such as the fat body) and by its own peptide hormones, produced by EE cells. In turn, gut-derived signals such as EE cell-derived peptide hormones can have long-range effects on other internal organs. EE cells accounts for 5–10% of midgut epithelial cells in flies compared to 0.4–0.6% in the mammalian small intestine [217, 218, 219]. Majority of them express peptide hormones, often more than one and with regional stereotypy [219, 220, 221, 222]. The developmental program of EE cells shares similarities with that of neurons, probably reflecting a common phylogenetic origin [223, 224, 225]. Consistent with this idea, all known EE peptide hormones (exception insect CCHamides) [226] are also produced by the brain. Acting through these hormones, EE cells may play “neural-like” roles in regulating intestinal physiology and conveying intestinal as well as nutritional state to other cell types or organs. These roles are particularly prominent in the midgut given the relatively sparse innervation of midgut region.
A role for EE cells on muscle peristalsis has been suggested by the finding that ablation of Diuretic hormone 31 (Dh31)-expressing EE cells or Dh31 downregulation both reduce muscle peristalsis in the larval anterior midgut, which may function as a valve to minimize mixing of acidified and non-acidified food in the acidic region of the midgut [227]. Adult EE cells produce Bursicon. Signaling through the Bursicon/DLGR2 receptor in visceral muscle, represses the production of the visceral muscle-derived mitogen Vein and, consequently, ISC proliferation. Another study found in
A high-sugar diet leads to increased midgut EE cell number and enhanced production of EE-derived Activin ligand (Activin-β not Daw) [230] suggesting systemic roles for EE-derived peptide hormones. Mirroring the activin-mediated fat-to-gut signaling involved in sucrose repression, midgut-derived Activin-β binds to the TGF-β receptor Baboon in fat cells which, in turn, leads to enhancement of Akh signaling it the fat body and consequent hyperglycemia [230]. CCHamides are insect hormones [231, 232] and their expression is promoted by nutrient availability and sites of expression include the gut EE cells, a subset of central neurons and, possibly, the fat body [226, 233, 234]. Their receptors are expressed in the nervous system including the insulin-producing neurons, and are absent from the gut [226, 234]. Although not strictly gut-derived, a new peptide hormone produced not by EE cells, but by an adjacent secretory gland may have provided the most compelling example to date of gut-to-brain communication. Indeed, Limostatin (Lst) peptide is produced by the
In animals with a vascular system, peptides secreted from EE cells can enter the bloodstream and reach tissues at a considerable distance, ranging from other cells in the digestive tract to brain centers regulating appetite [236]. Nutrient availability can also affect the number of EE cells; signaling through the nuclear hormone receptor Hr96, dietary lipids control EE differentiation during the first few days of adult life, providing another way to couple nutrient availability with tissue architecture and physiology [237]. Modulation of intestinal physiology by systemic signals has also been looked into [220, 221]. Control of epithelial turnover by insulin-like peptides or JH (juvenile hormone), and the coupling of dietary availability of sugars with EC digestive enzyme production via the fat body-derived Activin ligand Daw are some examples. The actions of the diuretic peptide Leucokinin (Lk), secreted into the circulation from CNS-derived nerves that terminate at the abdominal wall [14, 238, 239] is an another example. Downregulation of either this peptide or its receptor leads to abnormal excreta and extreme fluid retention that can rupture the abdominal wall [14]. Finally, a link between energy balance, intestinal permeability, and immunity has been suggested by the finding that sNPF is a target of the Crtc/CREB energy sensing pathway, and functions to maintain epithelial barrier integrity acting through its receptor in ECs [240]. Although the precise source of sNPF remains to be established, tissue-specific genetic and expression data points to roles in neurosecretory cells [240], consistent with roles as a neuroendocrine hormone or in gut-innervating neurons. Gut can also produce long-range signals to affect the physiology of other organs, for example by production of the signaling protein Hh by larval EC. Circulating Hh regulates developmental timing by controlling ecdysteroid production in the prothoracic gland, and is required for mobilization of fat body TAG stores during starvation [241].
Other functions of brain-gut peptides and hormones include detection and utilization of nutrients during hunger, stress or normal conditions. Diuretic hormone 44 (Dh44), a homolog of the mammalian corticotropin –releasing -hormone (CRH) activate by nutritive sugars. Disturbed activity of Dh44 neurons leads to fail to select nutritive sugars [131]. These neurons localized to PI in adult brain, counterpart of mammalian hypothalamus [242] are filled with neurosecretory cells [131]. Dh44 conveys information from Dh44 neurons to Dh44 receptor R1 neurons in the brain and R2 cells in the gut suggesting requirement for nutrient selection. Artificial activation of these neurons causes rapid PER and it has been suggested that Dh44 is necessary and sufficient for gut motility and excretion in flies [131]. Both Dh44 neurons and the gut-innervating insulin-producing neurons of the PI are innervated by Hugin-producing neurons that suppress food intake and induce locomotion, providing a possible link between food-related behaviors and intestinal physiology [243]. It is seen later that Dh44+ neurons rapidly activate during amino acid feeding and are a direct sensor of dietary amino acids [244].
Fly gut peptide Dromyosuppressin [181] expresses in the number of cells in central nervous system (CNS) in adult stage, extending into the rectum, near the anus; part of the adult gut. Their immunoreactive fibers also project into the crop and show expression of Dromyosuppressin [172] and crop abundantly expresses Dromyosuppressin receptors (Dromyosuppressin receptor I) [220]. The effects of neuropeptide on neural regulation of crop motility and contractions have been shown [245]. Serotonergic neurons have also been shown to regulate insulin producing cells (IPC) located in the PI of the adult brain of the fly [246]. DILP2, 3 and 5 express particularly in midgut [187, 220, 247] and extend their axons to proventriculus, crop and corpora cardiaca [248]. DILP2 is particularly involved in carbohydrate metabolism [249]. Decreased levels of DILP2 affects stored trehalose as well [248]. IPC knockdown flies show increased glycogen storage, high levels of circulating triglycerides and extended lifespan [248].
Mammalian neuropeptide NPY (Neuropeptide – Y- precursor) has an invertebrate homologous peptide called NPF due to characteristic C- terminal F residue [250]. NPF has been shown to co- localize within midgut cells in
Other neuropeptides include Allatostatin characterized into three kinds namely Allatostatin A/B/C [258, 259, 260]. The endocrine cells producing Allatostatin are found in the posterior midgut and is innervated by axons from thoracico-abdominal ganglion [220]. Its receptors, DAR1 and DAR2 are predominantly located in the central nervous system and gastrointestinal tract (including crop, midgut and hindgut) respectively [261, 262]. Allatostain is also present throughout midgut. Another major peptide is PDF. It is expressed in central nervous system [263] and its neurons are also found in thoracico-abdominal ganglion [184]. Axons from these neurons innervate midgut and hindgut and in the crop. These neuropeptides are closely associated to circadian rhythm [264] and locomotor activity as well.
5. Drosophila gut brain axis: role in health and disease
Various studies have suggested that intestinal health has a significant impact on neurodegeneration. Specifically, dysregulation of GBA cross talk has been associated with metabolic syndrome [265, 266] and psychiatric disorders. GBA is largely mediated by CNS, ENS and intestinal microbiota. With its much simpler gut microbiota (1–30 taxa) as opposed to vertebrates intestine (1–500 taxa) [267],
5.1 Metabolic disorders
Most genes and pathways that play crucial roles in metabolic diseases are conserved between flies and humans [268]. Diet-induced obesity in flies is associated with many of the pathophysiological consequences found in humans, including hyperglycemia, insulin resistance, cardiac arrhythmia and fibrosis, reduced longevity [269, 270] and nephrosis [271]. The gut is crucial for peripheral body fat storage and serves as a major site of dietary lipid absorption and absorbs other dietary macronutrients (sugars, proteins and fats). It also metabolizes both glucose and lipids into metabolic intermediates, which after loading into hemolymph get used in other tissues and organs. In flies, lipoprotein complexes containing apolipophorins carry sterols and diacylglycerols from the gut to other tissues [81]. Fly lipoproteins also contain Hh, a cholesterol-linked, gut-derived ligand that binds the transmembrane receptor Patched on fat body target cells to promote lipolysis during larval starvation [241, 272]. The human anti-obesity drug orlistat, a gastric lipase inhibitor, has been shown to reduce body fat accumulation in adult flies [56]. Supporting a crucial role for lipolysis, midgut lipid accumulation and global fat storage are reduced by the insulin signaling pathway inhibitor Foxo in enterocytes, via reducing the expression of
The gut microbiota and its metabolism plays an important role in modulation of fat storage in the fly. As seen in humans, fly gut is enriched in
5.2 Neurodegenerative diseases
5.3 Intestinal dysbiosis and infections
5.4 Gut-cancer model
6. Conclusion and outlook
Recently
In the recent past, data from the intestinal mechanisms found in flies have been shown to be active in mammals, and may therefore become relevant in the context of human pathologies including diabetes, obesity, neurodegenerative diseases, gastrointestinal cancers, or aging. Parallel findings of communication between gut and brain via neuropeptides in
Developing new techniques and behavioral assays can help us explore physiological drives: what is the gut function to maintain the overall health of the animal. It would be interesting to find out the key intestinal sensors. How the physical association between gut and brain via neural micro circuits regulate decisions regarding nutrition, hunger and satiety is under question. Better “holistic” and quantitative methods, integration of spatial and temporal information about genetic events more comprehensively are required, so that cause and effect can be uncoupled in a physiological context [306]. We anticipate and hope that fly models of intestinal pathology, in addition to uncovering newly identified genes (chemosensory and others) and basic biology mechanisms will emphasize the most conserved aspects of human intestinal biology. As a result, fly will contribute to translational research investigating drug effects, and microbial and host genetic component analyses, leading to biological findings that are broadly applicable to human health and disease.
Funding declaration
This work is supported by Wellcome Trust/DBT India Alliance Fellowship (grant number IA/I/15/2/502074) awarded to PK.
Criteria for authorship
ZS, SK and PK all substantially contributed to the conception and design of the work. Everybody participated in drafting and revising the work, made the figures, wrote the chapter and approved the final version for publication.
Abbreviations
| ENS | Enteric nervous system |
| GI | Gastrointestinal |
| GBA | Gut brain axis |
| CNS | Central nervous system |
| ISCs | Intestinal stem cells |
| ECs | Enterocytes |
| EE | Enteroendocrine |
| EBs | Enteroblasts |
| TA | Transient amplifying |
| AMP-5′ | Adenosine monophosphate |
| AMPK- 5’ | AMP activated kinase |
| TAG | Triacylglyceride |
| Sug | Sugarbabe |
| CD36 | Cluster of differentiation 36 |
| Npc1 | Niemann-Pick C1 |
| SREBPs | Sterol regulatory element-binding proteins |
| Akh | Adipokinetic hormone |
| RNAi | RNA interference |
| Kcc | Kazachoc |
| SIET | Scanning ion-selective electrode technique |
| Dh44 | Diuretic hormone 44 |
| GPCR | G-protein coupled receptors |
| PNS | Peripheral nervous system |
| GLUT | Glucose transporter |
| SGLT | Na+/glucose cotransporter |
| GIP | Gastric inhibitory polypeptide |
| GLP | Glucagon like peptide |
| TK | Tachykinin |
| MOR | μ – opoid receptors |
| IGN | Intestinal gluconeogenesis |
| MOR-KO | MOR-knockouts |
| cNST | Caudal nucleus of the solitary tract |
| GLP1 | Glucagon- like peptide 1 |
| sNPF | Short neuropeptide F |
| AN | Antennal nerve |
| Ilp7 | Insulin-like peptide 7 |
| VN | Vagus nerve |
| IBS | Intestinal bowel syndrome |
| HGN1 | Hindgut neuron1 |
| Hh | Hedgehog |
| Poxn | Pox-neuro |
| Dh31 | Diuretic hormone 31 |
| Lst | Limostatin |
| Lk | Leucokinin |
| JH | Juvenile hormone |
| Crtc/CREB | cAMP-regulated transcriptional co-activator/ cyclic AMP-responsive element-binding protein |
| SEZ | Suboesophageal zone |
| CRH | Corticotropin –releasing -hormone |
| PI | Pars intercerebralis |
| IPC | Insulin producing cells |
| DILP | Drosophila insulin- like- peptide |
| 5-HT1A-5 | Hydroxy tryptamine /serotonin |
| AKH | Adipokinetic hormone |
| PYY | Peptide YY |
| AgRP | Agouti-related protein |
| CCK | Choleocystokinin |
| PP | Pancreatic polypeptide |
| NPY | Neuropeptide Y |
| Pigment- dispersing factor | |
| CRF | Corticotropin-releasing factor |
| BDNF | Brain-derived neurotrophic factor |
| SCFAs | Short-chain fatty acid |
| AMP | Anti-microbial peptide |
| ROS | Reactive oxygen species |
| DUOX | Dual oxidase |
| IMD | Immune deficiency |
| APC | Adenomatous polyposis coli gene |
| CRC | Colorectal cancer |
References
- 1.
Thompson DG, Malagelada JR. Guts and their motions (gastrointestinal motility in health and disease). J Clin Gastroenterol. 1981;3 Suppl 1:81-87 - 2.
Edgecomb RS, Harth CE, Schneiderman AM. Regulation of feeding behavior in adult Drosophila melanogaster varies with feeding regime and nutritional state. J Exp Biol. 1994;197:215-35 - 3.
Demerec M. Biology of Drosophila . Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. 1994:425-41 - 4.
Stoffolano JG, Jr., Haselton AT. The adult Dipteran crop: a unique and overlooked organ. Annu Rev Entomol. 2013;58:205-25 - 5.
King DG. Cellular organization and peritrophic membrane formation in the cardia (proventriculus) of Drosophila melanogaster . J Morphol. 1988;196(3):253-82 - 6.
Tzou P, Ohresser S, Ferrandon D, Capovilla M, Reichhart JM, Lemaitre B, et al. Tissue-specific inducible expression of antimicrobial peptide genes in Drosophila surface epithelia. Immunity. 2000;13(5):737-48 - 7.
Demerec M. (Editor). Biology of Drosophila . John Wiley & Sons, Inc, New York. 1950 - 8.
Douglas A. E. ebCRF, Simpson S. J., Douglas A. E. The alimentary canal in The Insects: Structure and Function. Cambridge University Press, Cambridge. 2013:46-80 - 9.
Chen J, Sayadian AC, Lowe N, Lovegrove HE, St Johnston D. An alternative mode of epithelial polarity in the Drosophila midgut. PLoS Biol. 2018;16(10):e3000041 - 10.
Hegedus D, Erlandson M, Gillott C, Toprak U. New insights into peritrophic matrix synthesis, architecture, and function. Annu Rev Entomol. 2009;54:285-302 - 11.
Murakami R, Shigenaga A, Matsumoto A, Yamaoka I, Tanimura T. Novel tissue units of regional differentiation in the gut epithelium of Drosopbila, as revealed by P-element-mediated detection of enhancer. Rouxs Arch Dev Biol. 1994;203(5):243-9 - 12.
Buchon N, Osman D, David FP, Fang HY, Boquete JP, Deplancke B, et al. Morphological and molecular characterization of adult midgut compartmentalization in Drosophila . Cell Rep. 2013;3(5):1725-38 - 13.
Marianes A, Spradling AC. Physiological and stem cell compartmentalization within the Drosophila midgut. Elife. 2013;2:e00886 - 14.
Cognigni P, Bailey AP, Miguel-Aliaga I. Enteric neurons and systemic signals couple nutritional and reproductive status with intestinal homeostasis. Cell Metab. 2011;13(1):92-104 - 15.
Linneweber GA, Jacobson J, Busch KE, Hudry B, Christov CP, Dormann D, et al. Neuronal control of metabolism through nutrient-dependent modulation of tracheal branching. Cell. 2014;156(1-2):69-83 - 16.
Strand M, Micchelli CA. Quiescent gastric stem cells maintain the adult Drosophila stomach. Proc Natl Acad Sci U S A. 2011;108(43):17696-701 - 17.
Strand M, Micchelli CA. Regional control of Drosophila gut stem cell proliferation: EGF establishes GSSC proliferative set point & controls emergence from quiescence. PLoS One. 2013;8(11):e80608 - 18.
Sandborn EB, Duclos S, Messier PE, Roberge JJ. Atypical intestinal striated muscle in Drosophila melanogaster . J Ultrastruct Res. 1967;18(5):695-702 - 19.
Pitsouli C, Apidianakis Y, Perrimon N. Homeostasis in infected epithelia: stem cells take the lead. Cell Host Microbe. 2009;6(4):301-7 - 20.
Rubin DC. Intestinal morphogenesis. Curr Opin Gastroenterol. 2007;23(2):111-4 - 21.
Kedinger M, Simon-Assmann P, Haffen K. Growth and differentiation of intestinal endodermal cells in a coculture system. Gut. 1987;28 Suppl:237-41 - 22.
Tepass U, Hartenstein V. Epithelium formation in the Drosophila midgut depends on the interaction of endoderm and mesoderm. Development. 1994;120(3):579-90 - 23.
Crosnier C, Stamataki D, Lewis J. Organizing cell renewal in the intestine: stem cells, signals and combinatorial control. Nat Rev Genet. 2006;7(5):349-59 - 24.
Shanbhag S, Tripathi S. Epithelial ultrastructure and cellular mechanisms of acid and base transport in the Drosophila midgut. J Exp Biol. 2009;212(Pt 11):1731-44 - 25.
Baumann O. Posterior midgut epithelial cells differ in their organization of the membrane skeleton from other drosophila epithelia. Exp Cell Res. 2001;270(2):176-87 - 26.
Gartner LP. Submicroscopic morphology of the adult drosophila midgut. J Baltimore Coll Dent Surg. 1970;25(2):64-76 - 27.
Gooday GW. Aggressive and defensive roles for chitinases. EXS. 1999;87:157-69 - 28.
Vodovar N, Vinals M, Liehl P, Basset A, Degrouard J, Spellman P, et al. Drosophila host defense after oral infection by an entomopathogenic Pseudomonas species. Proc Natl Acad Sci U S A. 2005;102(32):11414-9 - 29.
Sengupta N, MacDonald TT. The role of matrix metalloproteinases in stromal/epithelial interactions in the gut. Physiology (Bethesda). 2007;22:401-9 - 30.
Jiang H, Edgar BA. EGFR signaling regulates the proliferation of Drosophila adult midgut progenitors. Development. 2009;136(3):483-93 - 31.
Kvietys PR, Granger DN. Physiology and pathophysiology of the colonic circulation. Clin Gastroenterol. 1986;15(4):967-83 - 32.
Rhee SH, Pothoulakis C, Mayer EA. Principles and clinical implications of the brain-gut-enteric microbiota axis. Nat Rev Gastroenterol Hepatol. 2009;6(5):306-14 - 33.
Komuro T, Hashimoto Y. Three-dimensional structure of the rat intestinal wall (mucosa and submucosa). Arch Histol Cytol. 1990;53(1):1-21 - 34.
Barker N, van de Wetering M, Clevers H. The intestinal stem cell. Genes Dev. 2008;22(14):1856-64 - 35.
Barker N, van Es JH, Kuipers J, Kujala P, van den Born M, Cozijnsen M, et al. Identification of stem cells in small intestine and colon by marker gene Lgr5. Nature. 2007;449(7165):1003-7 - 36.
Ohlstein B, Spradling A. The adult Drosophila posterior midgut is maintained by pluripotent stem cells. Nature. 2006;439(7075):470-4 - 37.
Takashima S, Mkrtchyan M, Younossi-Hartenstein A, Merriam JR, Hartenstein V. The behaviour of Drosophila adult hindgut stem cells is controlled by Wnt and Hh signalling. Nature. 2008;454(7204):651-5 - 38.
Apidianakis Y, Rahme LG. Drosophila melanogaster as a model for human intestinal infection and pathology. Dis Model Mech. 2011;4(1):21-30 - 39.
Martini E, Krug SM, Siegmund B, Neurath MF, Becker C. Mend Your Fences: The Epithelial Barrier and its Relationship With Mucosal Immunity in Inflammatory Bowel Disease. Cell Mol Gastroenterol Hepatol. 2017;4(1):33-46 - 40.
Kopp ZA, Jain U, Van Limbergen J, Stadnyk AW. Do antimicrobial peptides and complement collaborate in the intestinal mucosa? Front Immunol. 2015;6:17 - 41.
Buchon N, Broderick NA, Poidevin M, Pradervand S, Lemaitre B. Drosophila intestinal response to bacterial infection: activation of host defense and stem cell proliferation. Cell Host Microbe. 2009;5(2):200-11 - 42.
Scoville DH, Sato T, He XC, Li L. Current view: intestinal stem cells and signaling. Gastroenterology. 2008;134(3):849-64 - 43.
Amcheslavsky A, Jiang J, Ip YT. Tissue damage-induced intestinal stem cell division in Drosophila . Cell Stem Cell. 2009;4(1):49-61 - 44.
Buchon N, Broderick NA, Chakrabarti S, Lemaitre B. Invasive and indigenous microbiota impact intestinal stem cell activity through multiple pathways in Drosophila . Genes Dev. 2009;23(19):2333-44 - 45.
Cronin SJ, Nehme NT, Limmer S, Liegeois S, Pospisilik JA, Schramek D, et al. Genome-wide RNAi screen identifies genes involved in intestinal pathogenic bacterial infection. Science. 2009;325(5938):340-3 - 46.
Abraham I, Doane WW. Genetic regulation of tissue-specific expression of amylase structural genes in Drosophila melanogaster . Proc Natl Acad Sci U S A. 1978;75(9):4446-50 - 47.
Dutta D, Buchon N, Xiang J, Edgar BA. Regional Cell Specific RNA Expression Profiling of FACS Isolated Drosophila Intestinal Cell Populations. Curr Protoc Stem Cell Biol. 2015;34:2F 1-2F 14 - 48.
Lehane M. J. BPF. Biology of the Insect Midgut. Chapman & Hall, London. 1996 - 49.
Amcheslavsky A, Song W, Li Q , Nie Y, Bragatto I, Ferrandon D, et al. Enteroendocrine cells support intestinal stem-cell-mediated homeostasis in Drosophila . Cell Rep. 2014;9(1):32-9 - 50.
Wigglesworth V. B. The Principles of Insect Physiology. Chapman and Hall Ltd, London. 1972 - 51.
Clissold FJ, Tedder BJ, Conigrave AD, Simpson SJ. The gastrointestinal tract as a nutrient-balancing organ. Proc Biol Sci. 2010;277(1688):1751-9 - 52.
Chng WA, Sleiman MSB, Schupfer F, Lemaitre B. Transforming growth factor beta/activin signaling functions as a sugar-sensing feedback loop to regulate digestive enzyme expression. Cell Rep. 2014;9(1):336-48 - 53.
Mattila J, Havula E, Suominen E, Teesalu M, Surakka I, Hynynen R, et al. Mondo-Mlx Mediates Organismal Sugar Sensing through the Gli-Similar Transcription Factor Sugarbabe. Cell Rep. 2015;13(2):350-64 - 54.
Bujold M, Gopalakrishnan A, Nally E, King-Jones K. Nuclear receptor DHR96 acts as a sentinel for low cholesterol concentrations in Drosophila melanogaster . Mol Cell Biol. 2010;30(3):793-805 - 55.
Horner MA, Pardee K, Liu S, King-Jones K, Lajoie G, Edwards A, et al. The Drosophila DHR96 nuclear receptor binds cholesterol and regulates cholesterol homeostasis. Genes Dev. 2009;23(23):2711-6 - 56.
Sieber MH, Thummel CS. The DHR96 nuclear receptor controls triacylglycerol homeostasis in Drosophila . Cell Metab. 2009;10(6):481-90 - 57.
Sieber MH, Thummel CS. Coordination of triacylglycerol and cholesterol homeostasis by DHR96 and the Drosophila LipA homolog magro. Cell Metab. 2012;15(1):122-7 - 58.
Karpac J, Biteau B, Jasper H. Misregulation of an adaptive metabolic response contributes to the age-related disruption of lipid homeostasis in Drosophila . Cell Rep. 2013;4(6):1250-61 - 59.
Miguel-Aliaga I. Nerveless and gutsy: intestinal nutrient sensing from invertebrates to humans. Semin Cell Dev Biol. 2012;23(6):614-20 - 60.
Caccia S, Leonardi MG, Casartelli M, Grimaldi A, de Eguileor M, Pennacchio F, et al. Nutrient absorption by Aphidius ervi larvae. J Insect Physiol. 2005;51(11):1183-92 - 61.
Caccia S, Casartelli M, Grimaldi A, Losa E, de Eguileor M, Pennacchio F, et al. Unexpected similarity of intestinal sugar absorption by SGLT1 and apical GLUT2 in an insect (Aphidius ervi, Hymenoptera) and mammals. Am J Physiol Regul Integr Comp Physiol. 2007;292(6):R2284-91 - 62.
Price DR, Wilkinson HS, Gatehouse JA. Functional expression and characterisation of a gut facilitative glucose transporter, NlHT1, from the phloem-feeding insect Nilaparvata lugens (rice brown planthopper). Insect Biochem Mol Biol. 2007;37(11):1138-48 - 63.
Price DR, Tibbles K, Shigenobu S, Smertenko A, Russell CW, Douglas AE, et al. Sugar transporters of the major facilitator superfamily in aphids; from gene prediction to functional characterization. Insect Mol Biol. 2010;19 Suppl 2:97-112 - 64.
Bifano TD, Alegria TG, Terra WR. Transporters involved in glucose and water absorption in the Dysdercus peruvianus (Hemiptera: Pyrrhocoridae) anterior midgut. Comp Biochem Physiol B Biochem Mol Biol. 2010;157(1):1-9 - 65.
Escher SA, Rasmuson-Lestander A. The Drosophila glucose transporter gene: cDNA sequence, phylogenetic comparisons, analysis of functional sites and secondary structures. Hereditas. 1999;130(2):95-103 - 66.
Artero RD, Terol-Alcayde J, Paricio N, Ring J, Bargues M, Torres A, et al. saliva, a new Drosophila gene expressed in the embryonic salivary glands with homologues in plants and vertebrates. Mech Dev. 1998;75(1-2):159-62 - 67.
Baker RF, Leach KA, Braun DM. SWEET as sugar: new sucrose effluxers in plants. Mol Plant. 2012;5(4):766-8 - 68.
Meyer H, Vitavska O, Wieczorek H. Identification of an animal sucrose transporter. J Cell Sci. 2011;124(Pt 12):1984-91 - 69.
Kanamori Y, Saito A, Hagiwara-Komoda Y, Tanaka D, Mitsumasu K, Kikuta S, et al. The trehalose transporter 1 gene sequence is conserved in insects and encodes proteins with different kinetic properties involved in trehalose import into peripheral tissues. Insect Biochem Mol Biol. 2010;40(1):30-7 - 70.
Vitavska O, Wieczorek H. The SLC45 gene family of putative sugar transporters. Mol Aspects Med. 2013;34(2-3):655-60 - 71.
Boudko DY. Molecular basis of essential amino acid transport from studies of insect nutrient amino acid transporters of the SLC6 family (NAT-SLC6). J Insect Physiol. 2012;58(4):433-49 - 72.
Colombani J, Raisin S, Pantalacci S, Radimerski T, Montagne J, Leopold P. A nutrient sensor mechanism controls Drosophila growth. Cell. 2003;114(6):739-49 - 73.
Martin JF, Hersperger E, Simcox A, Shearn A. minidiscs encodes a putative amino acid transporter subunit required non-autonomously for imaginal cell proliferation. Mech Dev. 2000;92(2):155-67 - 74.
Goberdhan DC, Meredith D, Boyd CA, Wilson C. PAT-related amino acid transporters regulate growth via a novel mechanism that does not require bulk transport of amino acids. Development. 2005;132(10):2365-75 - 75.
Miller MM, Popova LB, Meleshkevitch EA, Tran PV, Boudko DY. The invertebrate B(0) system transporter, D. melanogaster NAT1, has unique d-amino acid affinity and mediates gut and brain functions. Insect Biochem Mol Biol. 2008;38(10):923-31 - 76.
Reynolds B, Roversi P, Laynes R, Kazi S, Boyd CA, Goberdhan DC. Drosophila expresses a CD98 transporter with an evolutionarily conserved structure and amino acid-transport properties. Biochem J. 2009;420(3):363-72 - 77.
Roman G, Meller V, Wu KH, Davis RL. The opt1 gene of Drosophila melanogaster encodes a proton-dependent dipeptide transporter. Am J Physiol. 1998;275(3):C857-69 - 78.
Capo F, Chaduli D, Viallat-Lieutaud A, Charroux B, Royet J. Oligopeptide Transporters of the SLC15 Family Are Dispensable for Peptidoglycan Sensing and Transport in Drosophila . J Innate Immun. 2017;9(5):483-92 - 79.
Thimgan MS, Berg JS, Stuart AE. Comparative sequence analysis and tissue localization of members of the SLC6 family of transporters in adult Drosophila melanogaster . J Exp Biol. 2006;209(Pt 17):3383-404 - 80.
Chapman R. F. The Insects. Structure and Function. Cambridge University Press, New York. 2013 - 81.
Palm W, Sampaio JL, Brankatschk M, Carvalho M, Mahmoud A, Shevchenko A, et al. Lipoproteins in Drosophila melanogaster --assembly, function, and influence on tissue lipid composition. PLoS Genet. 2012;8(7):e1002828 - 82.
Huang X, Warren JT, Buchanan J, Gilbert LI, Scott MP. Drosophila Niemann-Pick type C-2 genes control sterol homeostasis and steroid biosynthesis: a model of human neurodegenerative disease. Development. 2007;134(20):3733-42 - 83.
Voght SP, Fluegel ML, Andrews LA, Pallanck LJ. Drosophila NPC1b promotes an early step in sterol absorption from the midgut epithelium. Cell Metab. 2007;5(3):195-205 - 84.
Chakrabarti S, Poidevin M, Lemaitre B. The Drosophila MAPK p38c regulates oxidative stress and lipid homeostasis in the intestine. PLoS Genet. 2014;10(9):e1004659 - 85.
Song W, Veenstra JA, Perrimon N. Control of lipid metabolism by tachykinin in Drosophila . Cell Rep. 2014;9(1):40-7 - 86.
Reiff T, Jacobson J, Cognigni P, Antonello Z, Ballesta E, Tan KJ, et al. Endocrine remodelling of the adult intestine sustains reproduction in Drosophila . Elife. 2015;4:e06930 - 87.
Dubreuil RR, Frankel J, Wang P, Howrylak J, Kappil M, Grushko TA. Mutations of alpha spectrin and labial block cuprophilic cell differentiation and acid secretion in the middle midgut of Drosophila larvae. Dev Biol. 1998;194(1):1-11 - 88.
Overend G, Luo Y, Henderson L, Douglas AE, Davies SA, Dow JA. Molecular mechanism and functional significance of acid generation in the Drosophila midgut. Sci Rep. 2016;6:27242 - 89.
Dubreuil RR. Copper cells and stomach acid secretion in the Drosophila midgut. Int J Biochem Cell Biol. 2004;36(5):745-52 - 90.
Li H, Qi Y, Jasper H. Preventing Age-Related Decline of Gut Compartmentalization Limits Microbiota Dysbiosis and Extends Lifespan. Cell Host Microbe. 2016;19(2):240-53 - 91.
Filippov V, Aimanova K, Gill SS. Expression of an Aedes aegypti cation-chloride cotransporter and itsDrosophila homologues. Insect Mol Biol. 2003;12(4):319-31 - 92.
Sun Q , Tian E, Turner RJ, Ten Hagen KG. Developmental and functional studies of the SLC12 gene family members from Drosophila melanogaster . Am J Physiol Cell Physiol. 2010;298(1):C26-37 - 93.
Brenner R, Atkinson NS. Calcium-activated potassium channel gene expression in the midgut of Drosophila . Comp Biochem Physiol B Biochem Mol Biol. 1997;118(2):411-20 - 94.
Feingold D, Starc T, O’Donnell MJ, Nilson L, Dent JA. The orphan pentameric ligand-gated ion channel pHCl-2 is gated by pH and regulates fluid secretion in Drosophila Malpighian tubules. J Exp Biol. 2016;219(Pt 17):2629-38 - 95.
Remnant EJ, Williams A, Lumb C, Yang YT, Chan J, Duchene S, et al. Evolution, Expression, and Function of Nonneuronal Ligand-Gated Chloride Channels in Drosophila melanogaster . G3 (Bethesda). 2016;6(7):2003-12 - 96.
Dubreuil RR, Das A, Base C, Mazock GH. The Drosophila Anion Exchanger (DAE) lacks a detectable interaction with the spectrin cytoskeleton. J Negat Results Biomed. 2010;9:5 - 97.
Naikkhwah W, O’Donnell MJ. Phenotypic plasticity in response to dietary salt stress: Na+ and K+ transport by the gut of Drosophila melanogaster larvae. J Exp Biol. 2012;215(Pt 3):461-70 - 98.
Chintapalli VR, Wang J, Dow JA. Using FlyAtlas to identify better Drosophila melanogaster models of human disease. Nat Genet. 2007;39(6):715-20 - 99.
Chintapalli VR, Kato A, Henderson L, Hirata T, Woods DJ, Overend G, et al. Transport proteins NHA1 and NHA2 are essential for survival, but have distinct transport modalities. Proc Natl Acad Sci U S A. 2015;112(37):11720-5 - 100.
Day JP, Wan S, Allan AK, Kean L, Davies SA, Gray JV, et al. Identification of two partners from the bacterial Kef exchanger family for the apical plasma membrane V-ATPase of Metazoa. J Cell Sci. 2008;121(Pt 15):2612-9 - 101.
Poulson D. F. BLE. Organization and function of the inorganic constituents of nuclei. Exp Cell Res. 1952;2:161-80 - 102.
McNulty M, Puljung M, Jefford G, Dubreuil RR. Evidence that a copper-metallothionein complex is responsible for fluorescence in acid-secreting cells of the Drosophila stomach. Cell Tissue Res. 2001;304(3):383-9 - 103.
Lauverjat S, Ballan-Dufrancais C, Wegnez M. Detoxification of cadmium. Ultrastructural study and electron-probe microanalysis of the midgut in a cadmium-resistant strain of Drosophila melanogaster . Biol Met. 1989;2(2):97-107 - 104.
Missirlis F, Kosmidis S, Brody T, Mavrakis M, Holmberg S, Odenwald WF, et al. Homeostatic mechanisms for iron storage revealed by genetic manipulations and live imaging of Drosophila ferritin. Genetics. 2007;177(1):89-100 - 105.
Wong R, Piper MDW, Blanc E, Partridge L. Pitfalls of measuring feeding rate in the fruit fly Drosophila melanogaster . Nat Methods. 2008;5(3):214-5 - 106.
Urquhart-Cronish M, Sokolowski MB. Gene-environment interplay in Drosophila melanogaster : chronic nutritional deprivation in larval life affects adult fecal output. J Insect Physiol. 2014;69:95-100 - 107.
Apger-McGlaughon J, Wolfner MF. Post-mating change in excretion by mated Drosophila melanogaster females is a long-term response that depends on sex peptide and sperm. J Insect Physiol. 2013;59(10):1024-30 - 108.
Buchanan RL, Benzer S. Defective glia in the Drosophila brain degeneration mutant drop-dead. Neuron. 1993;10(5):839-50 - 109.
Peller CR, Bacon EM, Bucheger JA, Blumenthal EM. Defective gut function in drop-dead mutant Drosophila . J Insect Physiol. 2009;55(9):834-9 - 110.
Li X, Staszewski L, Xu H, Durick K, Zoller M, Adler E. Human receptors for sweet and umami taste. Proc Natl Acad Sci U S A. 2002;99(7):4692-6 - 111.
Dyer J, Salmon KS, Zibrik L, Shirazi-Beechey SP. Expression of sweet taste receptors of the T1R family in the intestinal tract and enteroendocrine cells. Biochem Soc Trans. 2005;33(Pt 1):302-5 - 112.
Bezencon C, le Coutre J, Damak S. Taste-signaling proteins are coexpressed in solitary intestinal epithelial cells. Chem Senses. 2007;32(1):41-9 - 113.
Wong GT, Gannon KS, Margolskee RF. Transduction of bitter and sweet taste by gustducin. Nature. 1996;381(6585):796-800 - 114.
Ueda T, Ugawa S, Yamamura H, Imaizumi Y, Shimada S. Functional interaction between T2R taste receptors and G-protein alpha subunits expressed in taste receptor cells. J Neurosci. 2003;23(19):7376-80 - 115.
Wu SV, Rozengurt N, Yang M, Young SH, Sinnett-Smith J, Rozengurt E. Expression of bitter taste receptors of the T2R family in the gastrointestinal tract and enteroendocrine STC-1 cells. Proc Natl Acad Sci U S A. 2002;99(4):2392-7 - 116.
Hofer D, Puschel B, Drenckhahn D. Taste receptor-like cells in the rat gut identified by expression of alpha-gustducin. Proc Natl Acad Sci U S A. 1996;93(13):6631-4 - 117.
Hofer D, Drenckhahn D. Identification of the taste cell G-protein, alpha-gustducin, in brush cells of the rat pancreatic duct system. Histochem Cell Biol. 1998;110(3):303-9 - 118.
Wu SV, Chen MC, Rozengurt E. Genomic organization, expression, and function of bitter taste receptors (T2R) in mouse and rat. Physiol Genomics. 2005;22(2):139-49 - 119.
Rozengurt E. Taste receptors in the gastrointestinal tract. I. Bitter taste receptors and alpha-gustducin in the mammalian gut. Am J Physiol Gastrointest Liver Physiol. 2006;291(2):G171-7 - 120.
Mace OJ, Affleck J, Patel N, Kellett GL. Sweet taste receptors in rat small intestine stimulate glucose absorption through apical GLUT2. J Physiol. 2007;582(Pt 1):379-92 - 121.
Egan JM, Margolskee RF. Taste cells of the gut and gastrointestinal chemosensation. Mol Interv. 2008;8(2):78-81 - 122.
Margolskee RF, Dyer J, Kokrashvili Z, Salmon KS, Ilegems E, Daly K, et al. T1R3 and gustducin in gut sense sugars to regulate expression of Na+−glucose cotransporter 1. Proc Natl Acad Sci U S A. 2007;104(38):15075-80 - 123.
Au A, Gupta A, Schembri P, Cheeseman CI. Rapid insertion of GLUT2 into the rat jejunal brush-border membrane promoted by glucagon-like peptide 2. Biochem J. 2002;367(Pt 1):247-54 - 124.
Dyer J, Vayro S, King TP, Shirazi-Beechey SP. Glucose sensing in the intestinal epithelium. Eur J Biochem. 2003;270(16):3377-88 - 125.
Daly K, Al-Rammahi M, Arora DK, Moran AW, Proudman CJ, Ninomiya Y, et al. Expression of sweet receptor components in equine small intestine: relevance to intestinal glucose transport. Am J Physiol Regul Integr Comp Physiol. 2012;303(2):R199-208 - 126.
Park JH, Kwon JY. Heterogeneous expression of Drosophila gustatory receptors in enteroendocrine cells. PLoS One. 2011;6(12):e29022 - 127.
Park JH, Chen J, Jang S, Ahn TJ, Kang K, Choi MS, et al. A subset of enteroendocrine cells is activated by amino acids in the Drosophila midgut. FEBS Lett. 2016;590(4):493-500 - 128.
Palamiuc L, Noble T, Witham E, Ratanpal H, Vaughan M, Srinivasan S. A tachykinin-like neuroendocrine signalling axis couples central serotonin action and nutrient sensing with peripheral lipid metabolism. Nat Commun. 2017;8:14237 - 129.
Mace OJ, Schindler M, Patel S. The regulation of K- and L-cell activity by GLUT2 and the calcium-sensing receptor CasR in rat small intestine. J Physiol. 2012;590(12):2917-36 - 130.
Depoortere I. Taste receptors of the gut: emerging roles in health and disease. Gut. 2014;63(1):179-90 - 131.
Dus M, Lai JS, Gunapala KM, Min S, Tayler TD, Hergarden AC, et al. Nutrient Sensor in the Brain Directs the Action of the Brain-Gut Axis in Drosophila . Neuron. 2015;87(1):139-51 - 132.
Sandoval D, Cota D, Seeley RJ. The integrative role of CNS fuel-sensing mechanisms in energy balance and glucose regulation. Annu Rev Physiol. 2008;70:513-35 - 133.
Miyamoto T, Amrein H. Gluconeogenesis: An ancient biochemical pathway with a new twist. Fly (Austin). 2017;11(3):218-23 - 134.
Mithieux G, Andreelli F, Magnan C. Intestinal gluconeogenesis: key signal of central control of energy and glucose homeostasis. Curr Opin Clin Nutr Metab Care. 2009;12(4):419-23 - 135.
Delaere F, Duchampt A, Mounien L, Seyer P, Duraffourd C, Zitoun C, et al. The role of sodium-coupled glucose co-transporter 3 in the satiety effect of portal glucose sensing. Mol Metab. 2012;2(1):47-53 - 136.
Delaere F, Akaoka H, De Vadder F, Duchampt A, Mithieux G. Portal glucose influences the sensory, cortical and reward systems in rats. Eur J Neurosci. 2013;38(10):3476-86 - 137.
Soty M, Penhoat A, Amigo-Correig M, Vinera J, Sardella A, Vullin-Bouilloux F, et al. A gut-brain neural circuit controlled by intestinal gluconeogenesis is crucial in metabolic health. Mol Metab. 2015;4(2):106-17 - 138.
Mithieux G, Misery P, Magnan C, Pillot B, Gautier-Stein A, Bernard C, et al. Portal sensing of intestinal gluconeogenesis is a mechanistic link in the diminution of food intake induced by diet protein. Cell Metab. 2005;2(5):321-9 - 139.
Pillot B, Soty M, Gautier-Stein A, Zitoun C, Mithieux G. Protein feeding promotes redistribution of endogenous glucose production to the kidney and potentiates its suppression by insulin. Endocrinology. 2009;150(2):616-24 - 140.
Duraffourd C, De Vadder F, Goncalves D, Delaere F, Penhoat A, Brusset B, et al. Mu-opioid receptors and dietary protein stimulate a gut-brain neural circuitry limiting food intake. Cell. 2012;150(2):377-88 - 141.
De Vadder F, Kovatcheva-Datchary P, Goncalves D, Vinera J, Zitoun C, Duchampt A, et al. Microbiota-generated metabolites promote metabolic benefits via gut-brain neural circuits. Cell. 2014;156(1-2):84-96 - 142.
Glass MJ, Billington CJ, Levine AS. Opioids and food intake: distributed functional neural pathways? Neuropeptides. 1999;33(5):360-8 - 143.
Shin AC, Pistell PJ, Phifer CB, Berthoud HR. Reversible suppression of food reward behavior by chronic mu-opioid receptor antagonism in the nucleus accumbens. Neuroscience. 2010;170(2):580-8 - 144.
Howland RH. Vagus Nerve Stimulation. Curr Behav Neurosci Rep. 2014;1(2):64-73 - 145.
Krahl SE. Vagus nerve stimulation for epilepsy: A review of the peripheral mechanisms. Surg Neurol Int. 2012;3(Suppl 1):S47-52 - 146.
George MS, Sackeim HA, Rush AJ, Marangell LB, Nahas Z, Husain MM, et al. Vagus nerve stimulation: a new tool for brain research and therapy. Biol Psychiatry. 2000;47(4):287-95 - 147.
Breit S, Kupferberg A, Rogler G, Hasler G. Vagus Nerve as Modulator of the Brain-Gut Axis in Psychiatric and Inflammatory Disorders. Front Psychiatry. 2018;9:44 - 148.
Brookes SJ, Spencer NJ, Costa M, Zagorodnyuk VP. Extrinsic primary afferent signalling in the gut. Nat Rev Gastroenterol Hepatol. 2013;10(5):286-96 - 149.
Berthoud HR, Blackshaw LA, Brookes SJ, Grundy D. Neuroanatomy of extrinsic afferents supplying the gastrointestinal tract. Neurogastroenterol Motil. 2004;16 Suppl 1:28-33 - 150.
Mei N. Vagal glucoreceptors in the small intestine of the cat. J Physiol. 1978;282:485-506 - 151.
Jeanningros R. Vagal unitary responses to intestinal amino acid infusions in the anesthetized cat: a putative signal for protein induced satiety. Physiol Behav. 1982;28(1):9-21 - 152.
Lal S, Kirkup AJ, Brunsden AM, Thompson DG, Grundy D. Vagal afferent responses to fatty acids of different chain length in the rat. Am J Physiol Gastrointest Liver Physiol. 2001;281(4):G907-15 - 153.
Berthoud HR. Vagal and hormonal gut-brain communication: from satiation to satisfaction. Neurogastroenterol Motil. 2008;20 Suppl 1:64-72 - 154.
Holst JJ. The physiology of glucagon-like peptide 1. Physiol Rev. 2007;87(4):1409-39 - 155.
Thorens B. Expression cloning of the pancreatic beta cell receptor for the gluco-incretin hormone glucagon-like peptide 1. Proc Natl Acad Sci U S A. 1992;89(18):8641-5 - 156.
Hayes MR, De Jonghe BC, Kanoski SE. Role of the glucagon-like-peptide-1 receptor in the control of energy balance. Physiol Behav. 2010;100(5):503-10 - 157.
Williams EK, Chang RB, Strochlic DE, Umans BD, Lowell BB, Liberles SD. Sensory Neurons that Detect Stretch and Nutrients in the Digestive System. Cell. 2016;166(1):209-21 - 158.
Bonaz B, Bazin T, Pellissier S. The Vagus Nerve at the Interface of the Microbiota-Gut-Brain Axis. Front Neurosci. 2018;12:49 - 159.
Berthoud HR. The vagus nerve, food intake and obesity. Regul Pept. 2008;149(1-3):15-25 - 160.
Badman MK, Flier JS. The gut and energy balance: visceral allies in the obesity wars. Science. 2005;307(5717):1909-14 - 161.
Owyang C, Heldsinger A. Vagal control of satiety and hormonal regulation of appetite. J Neurogastroenterol Motil. 2011;17(4):338-48 - 162.
Zurowski D, Nowak L, Wordliczek J, Dobrogowski J, Thor PJ. Effects of vagus nerve stimulation in visceral pain model. Folia Med Cracov. 2012;52(1-2):57-69 - 163.
Tan HE, Sisti AC, Jin H, Vignovich M, Villavicencio M, Tsang KS, et al. The gut-brain axis mediates sugar preference. Nature. 2020;580(7804):511-6 - 164.
Hartenstein V, Tepass U, Gruszynski-Defeo E. Embryonic development of the stomatogastric nervous system in Drosophila . J Comp Neurol. 1994;350(3):367-81 - 165.
Gonzalez-Gaitan M, Jackle H. Invagination centers within the Drosophila stomatogastric nervous system anlage are positioned by Notch-mediated signaling which is spatially controlled through wingless. Development. 1995;121(8):2313-25 - 166.
Pankratz MJ, Hoch M. Control of epithelial morphogenesis by cell signaling and integrin molecules in the Drosophila foregut. Development. 1995;121(6):1885-98 - 167.
Spiess R, Schoofs A, Heinzel HG. Anatomy of the stomatogastric nervous system associated with the foregut in Drosophila melanogaster andCalliphora vicina third instar larvae. J Morphol. 2008;269(3):272-82 - 168.
Lee G, Park JH. Hemolymph sugar homeostasis and starvation-induced hyperactivity affected by genetic manipulations of the adipokinetic hormone-encoding gene in Drosophila melanogaster . Genetics. 2004;167(1):311-23 - 169.
Miguel-Aliaga I, Thor S. Segment-specific prevention of pioneer neuron apoptosis by cell-autonomous, postmitotic Hox gene activity. Development. 2004;131(24):6093-105 - 170.
Miguel-Aliaga I, Thor S, Gould AP. Postmitotic specification of Drosophila insulinergic neurons from pioneer neurons. PLoS Biol. 2008;6(3):e58 - 171.
Talsma AD, Christov CP, Terriente-Felix A, Linneweber GA, Perea D, Wayland M, et al. Remote control of renal physiology by the intestinal neuropeptide pigment-dispersing factor in Drosophila . Proc Natl Acad Sci U S A. 2012;109(30):12177-82 - 172.
Schoofs A, Huckesfeld S, Surendran S, Pankratz MJ. Serotonergic pathways in the Drosophila larval enteric nervous system. J Insect Physiol. 2014;69:118-25 - 173.
Myers L, Perera H, Alvarado MG, Kidd T. The Drosophila Ret gene functions in the stomatogastric nervous system with the Maverick TGFbeta ligand and the Gfrl co-receptor. Development. 2018;145(3) - 174.
Perea D, Guiu J, Hudry B, Konstantinidou C, Milona A, Hadjieconomou D, et al. Ret receptor tyrosine kinase sustains proliferation and tissue maturation in intestinal epithelia. EMBO J. 2017;36(20):3029-45 - 175.
Kenmoku H, Ishikawa H, Ote M, Kuraishi T, Kurata S. A subset of neurons controls the permeability of the peritrophic matrix and midgut structure in Drosophila adults. J Exp Biol. 2016;219(Pt 15):2331-9 - 176.
Stocker RF. The organization of the chemosensory system in Drosophila melanogaster : a review. Cell Tissue Res. 1994;275(1):3-26 - 177.
Wang Z, Singhvi A, Kong P, Scott K. Taste representations in the Drosophila brain. Cell. 2004;117(7):981-91 - 178.
Marella S, Fischler W, Kong P, Asgarian S, Rueckert E, Scott K. Imaging taste responses in the fly brain reveals a functional map of taste category and behavior. Neuron. 2006;49(2):285-95 - 179.
Ito K, Shinomiya K, Ito M, Armstrong JD, Boyan G, Hartenstein V, et al. A systematic nomenclature for the insect brain. Neuron. 2014;81(4):755-65 - 180.
Budnik V, Wu CF, White K. Altered branching of serotonin-containing neurons in Drosophila mutants unable to synthesize serotonin and dopamine. J Neurosci. 1989;9(8):2866-77 - 181.
McCormick J, Nichols R. Spatial and temporal expression identify dromyosuppressin as a brain-gut peptide in Drosophila melanogaster . J Comp Neurol. 1993;338(2):278-88 - 182.
Nichols R, Bendena WG, Tobe SS. Myotropic peptides in Drosophila melanogaster and the genes that encode them. J Neurogenet. 2002;16(1):1-28 - 183.
Anderson MS, Halpern ME, Keshishian H. Identification of the neuropeptide transmitter proctolin in Drosophila larvae: characterization of muscle fiber-specific neuromuscular endings. J Neurosci. 1988;8(1):242-55 - 184.
Nassel DR, Shiga S, Mohrherr CJ, Rao KR. Pigment-dispersing hormone-like peptide in the nervous system of the flies Phormia and Drosophila : immunocytochemistry and partial characterization. J Comp Neurol. 1993;331(2):183-98 - 185.
Dircksen H, Tesfai LK, Albus C, Nassel DR. Ion transport peptide splice forms in central and peripheral neurons throughout postembryogenesis of Drosophila melanogaster . J Comp Neurol. 2008;509(1):23-41 - 186.
Dircksen H. Insect ion transport peptides are derived from alternatively spliced genes and differentially expressed in the central and peripheral nervous system. J Exp Biol. 2009;212(Pt 3):401-12 - 187.
Cao C, Brown MR. Localization of an insulin-like peptide in brains of two flies. Cell Tissue Res. 2001;304(2):317-21 - 188.
Kaminski S, Orlowski E, Berry K, Nichols R. The effects of three Drosophila melanogaster myotropins on the frequency of foregut contractions differ. J Neurogenet. 2002;16(2):125-34 - 189.
Price MD, Merte J, Nichols R, Koladich PM, Tobe SS, Bendena WG. Drosophila melanogaster flatline encodes a myotropin orthologue to Manduca sexta allatostatin. Peptides. 2002;23(4):787-94 - 190.
Palmer GC, Tran T, Duttlinger A, Nichols R. The drosulfakinin 0 (DSK 0) peptide encoded in the conserved Dsk gene affects adult Drosophila melanogaster crop contractions. J Insect Physiol. 2007;53(11):1125-33 - 191.
Vanderveken M, O’Donnell MJ. Effects of diuretic hormone 31, drosokinin, and allatostatin A on transepithelial K(+) transport and contraction frequency in the midgut and hindgut of larval Drosophila melanogaster . Arch Insect Biochem Physiol. 2014;85(2):76-93 - 192.
Zhang W, Yan Z, Li B, Jan LY, Jan YN. Identification of motor neurons and a mechanosensitive sensory neuron in the defecation circuitry of Drosophila larvae. Elife. 2014;3 - 193.
Wayland MT, Defaye A, Rocha J, Jayaram SA, Royet J, Miguel-Aliaga I, et al. Spotting the differences: probing host/microbiota interactions with a dedicated software tool for the analysis of faecal outputs in Drosophila . J Insect Physiol. 2014;69:126-35 - 194.
Han H, Pan C, Liu C, Lv X, Yang X, Xiong Y, et al. Gut-neuron interaction via Hh signaling regulates intestinal progenitor cell differentiation in Drosophila . Cell Discov. 2015;1:15006 - 195.
Olds WH, Xu T. Regulation of food intake by mechanosensory ion channels in enteric neurons. Elife. 2014;3 - 196.
Galindo K, Smith DP. A large family of divergent Drosophila odorant-binding proteins expressed in gustatory and olfactory sensilla. Genetics. 2001;159(3):1059-72 - 197.
LeDue EE, Chen YC, Jung AY, Dahanukar A, Gordon MD. Pharyngeal sense organs drive robust sugar consumption in Drosophila . Nat Commun. 2015;6:6667 - 198.
Yapici N, Cohn R, Schusterreiter C, Ruta V, Vosshall LB. A Taste Circuit that Regulates Ingestion by Integrating Food and Hunger Signals. Cell. 2016;165(3):715-29 - 199.
Dethier V. G. GA, 1967 HyperphagiaintheblowflyJExpBiol - 200.
Pool AH, Kvello P, Mann K, Cheung SK, Gordon MD, Wang L, et al. Four GABAergic interneurons impose feeding restraint in Drosophila . Neuron. 2014;83(1):164-77 - 201.
Belzer W. R. Recurrent nerve inhibition of protein feeding in the blowfly Phormia regina . Physiol Entomol. 1978;3:259-63 - 202.
Miyamoto T, Slone J, Song X, Amrein H. A fructose receptor functions as a nutrient sensor in the Drosophila brain. Cell. 2012;151(5):1113-25 - 203.
Mishra D, Miyamoto T, Rezenom YH, Broussard A, Yavuz A, Slone J, et al. The molecular basis of sugar sensing in Drosophila larvae. Curr Biol. 2013;23(15):1466-71 - 204.
Miyamoto T, Amrein H. Diverse roles for the Drosophila fructose sensor Gr43a. Fly (Austin). 2014;8(1):19-25 - 205.
Holzer P, Farzi A, Hassan AM, Zenz G, Jacan A, Reichmann F. Visceral Inflammation and Immune Activation Stress the Brain. Front Immunol. 2017;8:1613 - 206.
Burbach JP. Neuropeptides from concept to online database www.neuropeptides.nl . Eur J Pharmacol. 2010;626(1):27-48 - 207.
Dockray GJ. Gastrointestinal hormones and the dialogue between gut and brain. J Physiol. 2014;592(14):2927-41 - 208.
Holzer P, Farzi A. Neuropeptides and the microbiota-gut-brain axis. Adv Exp Med Biol. 2014;817:195-219 - 209.
Kastin AJ. Handbook of biologically active peptides (2nd ed.). New York, NY: Academic Press. 2013 - 210.
Lach G, Schellekens H, Dinan TG, Cryan JF. Anxiety, Depression, and the Microbiome: A Role for Gut Peptides. Neurotherapeutics. 2018;15(1):36-59 - 211.
Strand FL. Neuropeptides: Regulators of physiological processes. Cambridge, MA: MIT Press1999 - 212.
Holzer P. Neuropeptides, Microbiota, and Behavior. Int Rev Neurobiol. 2016;131:67-89 - 213.
Kaelberer MM, Buchanan KL, Klein ME, Barth BB, Montoya MM, Shen X, et al. A gut-brain neural circuit for nutrient sensory transduction. Science. 2018;361(6408) - 214.
Beutler LR, Chen Y, Ahn JS, Lin YC, Essner RA, Knight ZA. Dynamics of Gut-Brain Communication Underlying Hunger. Neuron. 2017;96(2):461-75 e5 - 215.
Cummings DE, Overduin J. Gastrointestinal regulation of food intake. J Clin Invest. 2007;117(1):13-23 - 216.
Koda S, Date Y, Murakami N, Shimbara T, Hanada T, Toshinai K, et al. The role of the vagal nerve in peripheral PYY3-36-induced feeding reduction in rats. Endocrinology. 2005;146(5):2369-75 - 217.
Cheng H, Leblond CP. Origin, differentiation and renewal of the four main epithelial cell types in the mouse small intestine. III. Entero-endocrine cells. Am J Anat. 1974;141(4):503-19 - 218.
Micchelli CA, Perrimon N. Evidence that stem cells reside in the adult Drosophila midgut epithelium. Nature. 2006;439(7075):475-9 - 219.
Beehler-Evans R, Micchelli CA. Generation of enteroendocrine cell diversity in midgut stem cell lineages. Development. 2015;142(4):654-64 - 220.
Veenstra JA, Agricola HJ, Sellami A. Regulatory peptides in fruit fly midgut. Cell Tissue Res. 2008;334(3):499-516 - 221.
Veenstra JA. Peptidergic paracrine and endocrine cells in the midgut of the fruit fly maggot. Cell Tissue Res. 2009;336(2):309-23 - 222.
Reiher W, Shirras C, Kahnt J, Baumeister S, Isaac RE, Wegener C. Peptidomics and peptide hormone processing in the Drosophila midgut. J Proteome Res. 2011;10(4):1881-92 - 223.
Hartenstein V. The neuroendocrine system of invertebrates: a developmental and evolutionary perspective. J Endocrinol. 2006;190(3):555-70 - 224.
Hartenstein V, Takashima S, Adams KL. Conserved genetic pathways controlling the development of the diffuse endocrine system in vertebrates and Drosophila . Gen Comp Endocrinol. 2010;166(3):462-9 - 225.
Hartenstein V, Takashima S, Hartenstein P, Asanad S, Asanad K. bHLH proneural genes as cell fate determinants of entero-endocrine cells, an evolutionarily conserved lineage sharing a common root with sensory neurons. Dev Biol. 2017;431(1):36-47 - 226.
Li S, Torre-Muruzabal T, Sogaard KC, Ren GR, Hauser F, Engelsen SM, et al. Expression patterns of the Drosophila neuropeptide CCHamide-2 and its receptor may suggest hormonal signaling from the gut to the brain. PLoS One. 2013;8(10):e76131 - 227.
LaJeunesse DR, Johnson B, Presnell JS, Catignas KK, Zapotoczny G. Peristalsis in the junction region of the Drosophila larval midgut is modulated by DH31 expressing enteroendocrine cells. BMC Physiol. 2010;10:14 - 228.
O’Brien LE, Soliman SS, Li X, Bilder D. Altered modes of stem cell division drive adaptive intestinal growth. Cell. 2011;147(3):603-14 - 229.
Li J, Song J, Zaytseva YY, Liu Y, Rychahou P, Jiang K, et al. An obligatory role for neurotensin in high-fat-diet-induced obesity. Nature. 2016;533(7603):411-5 - 230.
Song W, Cheng D, Hong S, Sappe B, Hu Y, Wei N, et al. Midgut-Derived Activin Regulates Glucagon-like Action in the Fat Body and Glycemic Control. Cell Metab. 2017;25(2):386-99 - 231.
Roller L, Yamanaka N, Watanabe K, Daubnerova I, Zitnan D, Kataoka H, et al. The unique evolution of neuropeptide genes in the silkworm Bombyx mori . Insect Biochem Mol Biol. 2008;38(12):1147-57 - 232.
Hansen KK, Hauser F, Williamson M, Weber SB, Grimmelikhuijzen CJ. The Drosophila genes CG14593 and CG30106 code for G-protein-coupled receptors specifically activated by the neuropeptides CCHamide-1 and CCHamide-2. Biochem Biophys Res Commun. 2011;404(1):184-9 - 233.
Ren GR, Hauser F, Rewitz KF, Kondo S, Engelbrecht AF, Didriksen AK, et al. CCHamide-2 Is an Orexigenic Brain-Gut Peptide in Drosophila . PLoS One. 2015;10(7):e0133017 - 234.
Sano H, Nakamura A, Texada MJ, Truman JW, Ishimoto H, Kamikouchi A, et al. The Nutrient-Responsive Hormone CCHamide-2 Controls Growth by Regulating Insulin-like Peptides in the Brain of Drosophila melanogaster . PLoS Genet. 2015;11(5):e1005209 - 235.
Alfa RW, Park S, Skelly KR, Poffenberger G, Jain N, Gu X, et al. Suppression of insulin production and secretion by a decretin hormone. Cell Metab. 2015;21(2):323-34 - 236.
Clemmensen C, Muller TD, Woods SC, Berthoud HR, Seeley RJ, Tschop MH. Gut-Brain Cross-Talk in Metabolic Control. Cell. 2017;168(5):758-74 - 237.
Obniski R, Sieber M, Spradling AC. Dietary Lipids Modulate Notch Signaling and Influence Adult Intestinal Development and Metabolism in Drosophila . Dev Cell. 2018;47(1):98-111 e5 - 238.
O’Donnell MJ, Dow JA, Huesmann GR, Tublitz NJ, Maddrell SH. Separate control of anion and cation transport in malpighian tubules of Drosophila melanogaster . J Exp Biol. 1996;199(Pt 5):1163-75 - 239.
Radford JC, Davies SA, Dow JA. Systematic G-protein-coupled receptor analysis in Drosophila melanogaster identifies a leucokinin receptor with novel roles. J Biol Chem. 2002;277(41):38810-7 - 240.
Shen R, Wang B, Giribaldi MG, Ayres J, Thomas JB, Montminy M. Neuronal energy-sensing pathway promotes energy balance by modulating disease tolerance. Proc Natl Acad Sci U S A. 2016;113(23):E3307-14 - 241.
Rodenfels J, Lavrynenko O, Ayciriex S, Sampaio JL, Carvalho M, Shevchenko A, et al. Production of systemically circulating Hedgehog by the intestine couples nutrition to growth and development. Genes Dev. 2014;28(23):2636-51 - 242.
de Velasco B, Erclik T, Shy D, Sclafani J, Lipshitz H, McInnes R, et al. Specification and development of the pars intercerebralis and pars lateralis, neuroendocrine command centers in the Drosophila brain. Dev Biol. 2007;302(1):309-23 - 243.
Schoofs A, Huckesfeld S, Schlegel P, Miroschnikow A, Peters M, Zeymer M, et al. Selection of motor programs for suppressing food intake and inducing locomotion in the Drosophila brain. PLoS Biol. 2014;12(6):e1001893 - 244.
Yang Z, Huang R, Fu X, Wang G, Qi W, Mao D, et al. A post-ingestive amino acid sensor promotes food consumption in Drosophila . Cell Res. 2018;28(10):1013-25 - 245.
Richer S, Stoffolano JG, Jr., Yin CM, Nichols R. Innervation of dromyosuppressin (DMS) immunoreactive processes and effect of DMS and benzethonium chloride on the Phormia regina (Meigen) crop. J Comp Neurol. 2000;421(1):136-42 - 246.
Luo J, Lushchak OV, Goergen P, Williams MJ, Nassel DR. Drosophila insulin-producing cells are differentially modulated by serotonin and octopamine receptors and affect social behavior. PLoS One. 2014;9(6):e99732 - 247.
Ikeya T, Galic M, Belawat P, Nairz K, Hafen E. Nutrient-dependent expression of insulin-like peptides from neuroendocrine cells in the CNS contributes to growth regulation in Drosophila . Curr Biol. 2002;12(15):1293-300 - 248.
Nassel DR, Kubrak OI, Liu Y, Luo J, Lushchak OV. Factors that regulate insulin producing cells and their output in Drosophila . Front Physiol. 2013;4:252 - 249.
Kannan K, Fridell YW. Functional implications of Drosophila insulin-like peptides in metabolism, aging, and dietary restriction. Front Physiol. 2013;4:288 - 250.
Brown MR, Crim JW, Arata RC, Cai HN, Chun C, Shen P. Identification of a Drosophila brain-gut peptide related to the neuropeptide Y family. Peptides. 1999;20(9):1035-42 - 251.
Fadda M, Hasakiogullari I, Temmerman L, Beets I, Zels S, Schoofs L. Regulation of Feeding and Metabolism by Neuropeptide F and Short Neuropeptide F in Invertebrates. Front Endocrinol (Lausanne). 2019;10:64 - 252.
Nassel DR, Wegener C. A comparative review of short and long neuropeptide F signaling in invertebrates: Any similarities to vertebrate neuropeptide Y signaling? Peptides. 2011;32(6):1335-55 - 253.
Mertens I, Meeusen T, Huybrechts R, De Loof A, Schoofs L. Characterization of the short neuropeptide F receptor from Drosophila melanogaster . Biochem Biophys Res Commun. 2002;297(5):1140-8 - 254.
Lee KS, Kwon OY, Lee JH, Kwon K, Min KJ, Jung SA, et al. Drosophila short neuropeptide F signalling regulates growth by ERK-mediated insulin signalling. Nat Cell Biol. 2008;10(4):468-75 - 255.
Lee KS, You KH, Choo JK, Han YM, Yu K. Drosophila short neuropeptide F regulates food intake and body size. J Biol Chem. 2004;279(49):50781-9 - 256.
Findeisen M, Rathmann D, Beck-Sickinger AG. Structure-activity studies of RFamide peptides reveal subtype-selective activation of neuropeptide FF1 and FF2 receptors. ChemMedChem. 2011;6(6):1081-93 - 257.
Bechtold DA, Luckman SM. The role of RFamide peptides in feeding. J Endocrinol. 2007;192(1):3-15 - 258.
Woodhead AP, Stay B, Seidel SL, Khan MA, Tobe SS. Primary structure of four allatostatins: neuropeptide inhibitors of juvenile hormone synthesis. Proc Natl Acad Sci U S A. 1989;86(15):5997-6001 - 259.
Lorenz MW, Kellner R, Hoffmann KH. A family of neuropeptides that inhibit juvenile hormone biosynthesis in the cricket, Gryllus bimaculatus. J Biol Chem. 1995;270(36):21103-8 - 260.
Kramer SJ, Toschi A, Miller CA, Kataoka H, Quistad GB, Li JP, et al. Identification of an allatostatin from the tobacco hornworm Manduca sexta. Proc Natl Acad Sci U S A. 1991;88(21):9458-62 - 261.
Birgul N, Weise C, Kreienkamp HJ, Richter D. Reverse physiology in drosophila : identification of a novel allatostatin-like neuropeptide and its cognate receptor structurally related to the mammalian somatostatin/galanin/opioid receptor family. EMBO J. 1999;18(21):5892-900 - 262.
Lenz C, Williamson M, Grimmelikhuijzen CJ. Molecular cloning and genomic organization of an allatostatin preprohormone from Drosophila melanogaster . Biochem Biophys Res Commun. 2000;273(3):1126-31 - 263.
Renn SC, Park JH, Rosbash M, Hall JC, Taghert PH. A pdf neuropeptide gene mutation and ablation of PDF neurons each cause severe abnormalities of behavioral circadian rhythms in Drosophila . Cell. 1999;99(7):791-802 - 264.
Shafer OT, Yao Z. Pigment-Dispersing Factor Signaling and Circadian Rhythms in Insect Locomotor Activity. Curr Opin Insect Sci. 2014;1:73-80 - 265.
de Lartigue G, de La Serre CB, Raybould HE. Vagal afferent neurons in high fat diet-induced obesity; intestinal microflora, gut inflammation and cholecystokinin. Physiol Behav. 2011;105(1):100-5 - 266.
Grasset E, Puel A, Charpentier J, Collet X, Christensen JE, Terce F, et al. A Specific Gut Microbiota Dysbiosis of Type 2 Diabetic Mice Induces GLP-1 Resistance through an Enteric NO-Dependent and Gut-Brain Axis Mechanism. Cell Metab. 2017;25(5):1075-90 e5 - 267.
Broderick NA, Lemaitre B. Gut-associated microbes of Drosophila melanogaster . Gut Microbes. 2012;3(4):307-21 - 268.
Reiter LT, Potocki L, Chien S, Gribskov M, Bier E. A systematic analysis of human disease-associated gene sequences in Drosophila melanogaster . Genome Res. 2001;11(6):1114-25 - 269.
Birse RT, Choi J, Reardon K, Rodriguez J, Graham S, Diop S, et al. High-fat-diet-induced obesity and heart dysfunction are regulated by the TOR pathway in Drosophila . Cell Metab. 2010;12(5):533-44 - 270.
Na J, Musselman LP, Pendse J, Baranski TJ, Bodmer R, Ocorr K, et al. A Drosophila model of high sugar diet-induced cardiomyopathy. PLoS Genet. 2013;9(1):e1003175 - 271.
Na J, Sweetwyne MT, Park AS, Susztak K, Cagan RL. Diet-Induced Podocyte Dysfunction in Drosophila and Mammals. Cell Rep. 2015;12(4):636-47 - 272.
Palm W, Swierczynska MM, Kumari V, Ehrhart-Bornstein M, Bornstein SR, Eaton S. Secretion and signaling activities of lipoprotein-associated hedgehog and non-sterol-modified hedgehog in flies and mammals. PLoS Biol. 2013;11(3):e1001505 - 273.
Lin WS, Huang CW, Song YS, Yen JH, Kuo PC, Yeh SR, et al. Reduced Gut Acidity Induces an Obese-Like Phenotype in Drosophila melanogaster and in Mice. PLoS One. 2015;10(10):e0139722 - 274.
Wong AC, Dobson AJ, Douglas AE. Gut microbiota dictates the metabolic response of Drosophila to diet. J Exp Biol. 2014;217(Pt 11):1894-901 - 275.
Newell PD, Douglas AE. Interspecies interactions determine the impact of the gut microbiota on nutrient allocation in Drosophila melanogaster . Appl Environ Microbiol. 2014;80(2):788-96 - 276.
Whon TW, Shin NR, Jung MJ, Hyun DW, Kim HS, Kim PS, et al. Conditionally Pathogenic Gut Microbes Promote Larval Growth by Increasing Redox-Dependent Fat Storage in High-Sugar Diet-Fed Drosophila . Antioxid Redox Signal. 2017;27(16):1361-80 - 277.
Huang JH, Douglas AE. Consumption of dietary sugar by gut bacteria determines Drosophila lipid content. Biol Lett. 2015;11(9):20150469 - 278.
Iadanza MG, Jackson MP, Hewitt EW, Ranson NA, Radford SE. A new era for understanding amyloid structures and disease. Nat Rev Mol Cell Biol. 2018;19(12):755-73 - 279.
Llorens F, Thune K, Andres-Benito P, Tahir W, Ansoleaga B, Hernandez-Ortega K, et al. MicroRNA Expression in the Locus Coeruleus, Entorhinal Cortex, and Hippocampus at Early and Middle Stages of Braak Neurofibrillary Tangle Pathology. J Mol Neurosci. 2017;63(2):206-15 - 280.
Vangay P, Ward T, Gerber JS, Knights D. Antibiotics, pediatric dysbiosis, and disease. Cell Host Microbe. 2015;17(5):553-64 - 281.
Muegge BD, Kuczynski J, Knights D, Clemente JC, Gonzalez A, Fontana L, et al. Diet drives convergence in gut microbiome functions across mammalian phylogeny and within humans. Science. 2011;332(6032):970-4 - 282.
Delzenne NM, Neyrinck AM, Cani PD. Modulation of the gut microbiota by nutrients with prebiotic properties: consequences for host health in the context of obesity and metabolic syndrome. Microb Cell Fact. 2011;10 Suppl 1:S10 - 283.
Tilg H, Moschen AR. Microbiota and diabetes: an evolving relationship. Gut. 2014;63(9):1513-21 - 284.
Rosenfeld CS. Microbiome Disturbances and Autism Spectrum Disorders. Drug Metab Dispos. 2015;43(10):1557-71 - 285.
Galloway S, Jian L, Johnsen R, Chew S, Mamo JC. beta-amyloid or its precursor protein is found in epithelial cells of the small intestine and is stimulated by high-fat feeding. J Nutr Biochem. 2007;18(4):279-84 - 286.
Galloway S, Takechi R, Pallebage-Gamarallage MM, Dhaliwal SS, Mamo JC. Amyloid-beta colocalizes with apolipoprotein B in absorptive cells of the small intestine. Lipids Health Dis. 2009;8:46 - 287.
Nam KN, Mounier A, Wolfe CM, Fitz NF, Carter AY, Castranio EL, et al. Effect of high fat diet on phenotype, brain transcriptome and lipidome in Alzheimer’s model mice. Sci Rep. 2017;7(1):4307 - 288.
Lin B, Hasegawa Y, Takane K, Koibuchi N, Cao C, Kim-Mitsuyama S. High-Fat-Diet Intake Enhances Cerebral Amyloid Angiopathy and Cognitive Impairment in a Mouse Model of Alzheimer’s Disease, Independently of Metabolic Disorders. J Am Heart Assoc. 2016;5(6) - 289.
Goedert M. NEURODEGENERATION. Alzheimer’s and Parkinson’s diseases: The prion concept in relation to assembled Abeta, tau, and alpha-synuclein. Science. 2015;349(6248):1255555 - 290.
Ano Y, Sakudo A, Nakayama H, Onodera T. Uptake and dynamics of infectious prion protein in the intestine. Protein Pept Lett. 2009;16(3):247-55 - 291.
Yoshioka H, Takeda T, Higuchi K, Ohshio G, Miyake T, Sugiyama T, et al. Immunohistochemical examination of Peyer’s patches in senescence-accelerated mice. Autoimmunity. 1990;8(1):25-35 - 292.
Kimura I, Inoue D, Maeda T, Hara T, Ichimura A, Miyauchi S, et al. Short-chain fatty acids and ketones directly regulate sympathetic nervous system via G protein-coupled receptor 41 (GPR41). Proc Natl Acad Sci U S A. 2011;108(19):8030-5 - 293.
Tan FHP, Liu G, Lau SA, Jaafar MH, Park YH, Azzam G, et al. Lactobacillus probiotics improved the gut microbiota profile of a Drosophila melanogaster Alzheimer’s disease model and alleviated neurodegeneration in the eye. Benef Microbes. 2020;11(1):79-89 - 294.
Feltzin V, Wan KH, Celniker SE, Bonini NM. Role and impact of the gut microbiota in a Drosophila model for parkinsonism. 2019:718825 - 295.
Scheperjans F, Aho V, Pereira PA, Koskinen K, Paulin L, Pekkonen E, et al. Gut microbiota are related to Parkinson’s disease and clinical phenotype. Mov Disord. 2015;30(3):350-8 - 296.
Chen K, Luan X, Liu Q , Wang J, Chang X, Snijders AM, et al. Drosophila Histone Demethylase KDM5 Regulates Social Behavior through Immune Control and Gut Microbiota Maintenance. Cell Host Microbe. 2019;25(4):537-52 e8 - 297.
Ryu JH, Kim SH, Lee HY, Bai JY, Nam YD, Bae JW, et al. Innate immune homeostasis by the homeobox gene caudal and commensal-gut mutualism in Drosophila . Science. 2008;319(5864):777-82 - 298.
Ha EM, Oh CT, Bae YS, Lee WJ. A direct role for dual oxidase in Drosophila gut immunity. Science. 2005;310(5749):847-50 - 299.
Ha EM, Oh CT, Ryu JH, Bae YS, Kang SW, Jang IH, et al. An antioxidant system required for host protection against gut infection in Drosophila . Dev Cell. 2005;8(1):125-32 - 300.
Yu X, Waltzer L, Bienz M. A new Drosophila APC homologue associated with adhesive zones of epithelial cells. Nat Cell Biol. 1999;1(3):144-51 - 301.
Tian A, Benchabane H, Ahmed Y. Wingless/Wnt Signaling in Intestinal Development, Homeostasis, Regeneration and Tumorigenesis: A Drosophila Perspective. J Dev Biol. 2018;6(2) - 302.
Cordero JB, Stefanatos RK, Myant K, Vidal M, Sansom OJ. Non-autonomous crosstalk between the Jak/Stat and Egfr pathways mediates Apc1-driven intestinal stem cell hyperplasia in the Drosophila adult midgut. Development. 2012;139(24):4524-35 - 303.
Cordero JB, Ridgway RA, Valeri N, Nixon C, Frame MC, Muller WJ, et al. c-Src drives intestinal regeneration and transformation. EMBO J. 2014;33(13):1474-91 - 304.
Christofi T, Apidianakis Y. Ras-oncogenic Drosophila hindgut but not midgut cells use an inflammation-like program to disseminate to distant sites. Gut Microbes. 2013;4(1):54-9 - 305.
Ghosh S, Tibbit C, Liu JL. Effective knockdown of Drosophila long non-coding RNAs by CRISPR interference. Nucleic Acids Res. 2016;44(9):e84 - 306.
Miguel-Aliaga I, Jasper H, Lemaitre B. Anatomy and Physiology of the Digestive Tract of Drosophila melanogaster . Genetics. 2018;210(2):357-96