In the current study, we aimed to understand how neurotrophic factors affect ENS neuronal plasticity and function. Our data revealed that RET transcripts were co‐expressed with ChAT in enteric ganglia. To understand the function of enteric RET, a potent, selective, and GI‐restricted small molecule inhibitor of RET, GSK3179106, was employed. 12 We demonstrate that RET inhibition had profound effects on intestinal epithelial permeability, carbachol‐induced secretion, and neostigmine‐induced motility. Our data suggest that RET kinase activity in the adult gut functions as part of intrinsic neural pathways regulating gut mucosal and smooth muscle function.

Neurotrophic factor, glial cell line‐derived neurotrophic factor (GDNF), and its receptor, rearranged during transfection (RET), are essential for enteric nervous system (ENS) development and may also dynamically maintain the adult ENS by contributing to synapse formation, signal transmission, and neuronal plasticity. 1 - 4 The critical role of RET in the development of the ENS has been well established with the identification of genetic loss‐of‐function mutations in Hirschsprung patients who suffer from colonic intestinal pseudo‐obstruction due to variable lengths of bowel aganglionosis, often in association with loss‐of‐function mutations in RET. 5 This data combined with knockout mouse models that recapitulate human Hirschsprung's disease has provided insight into the role of RET in the development of the ENS. 5 - 9 Although the role of RET during the development of the ENS has been well established, the role of RET in the survival and plasticity of neurons in the adult ENS is not as well defined even though RET continues to be expressed throughout adulthood. 10 Recent data implicate RET in the maintenance and plasticity of the adult ENS. Indeed, neurons within the submucosal and myenteric plexus of the adult human colon express RET and the co‐receptors GFRα1 and GFRα2 while GDNF is expressed in the muscularis mucosa, circular and longitudinal muscle tissue. 10 Mechanistically, RET signaling is associated with the maturation of presynaptic axon terminals through regulating the expression of presynaptic proteins. 1 - 3 Systemic administration of GDNF in adult rodents significantly increases submucosal neuron density in both the small intestine and the colon. 11

Data for ex vivo assays are analyzed using GraphPad Prism software (version 7.0, GraphPad Software, Inc., La Jolla, California) and presented as scatter plots with mean ± standard deviation. A repeated‐measure two‐way ANOVA with a Bonferroni's post hoc test or a one‐way ANOVA with Dunnett's post hoc test was used to compare individual means between groups. P < 0.05 was considered statistically significant. For Ussing chambers or muscle bath experiments, n = an individual piece of tissue, with 1‐2 tissues used per rat. Chambers were randomly assigned to treatment. Group sizes were based on similar studies.

Resected normal human full‐thickness colon tissue was obtained with patient's consent through The MT Group, Van Nuys, CA, and in accordance with GSK's guidance for the use of human biological samples. Rat colons and mouse DRG were obtained at the end of an in‐life study. All animal studies were conducted in accordance with the GSK Policy on the Care, Welfare and Treatment of Laboratory Animals and were reviewed by the University of Oklahoma Health Sciences Center Institutional Animal Care and Use Committee (IACUC). RNAscope reagents, Discovery mRNA RED pretreatment kit (Cat# 760‐237), Discovery mRNA RED Detection kit (Cat# 760‐234), Discovery mRNA RED Probe Amplification kit (Cat# 760‐236), and mRNA RED Pretreatment A/B (Cat# 253‐4893/4), were all purchased from Ventana. Rn RET (Cat# 452358), Rn PPIB (Cat# 313921‐C2), Hs RET (Cat# 424878), Hs PPIB (Cat# 313901), Hs ChAT (Cat# 450678‐C2), RNAscope 2.5 VS Detection Reagents–RED (Cat# 322260), RNAscope 2.5 LS Green Accessory Pack, (Cat# 322550), RNAscope 2.5 VS Sample Prep. Reagents (Cat# 322220), and RNAscope VS Ventana Accessory Kit (Cat# 320630) were all purchased from ACD. Immunohistochemistry reagents CC1 (Cat# 950‐224), Rabbit anti‐Synaptophysin (clone MRQ‐40, Cat# 7604595), Anti‐Rb NP (Cat# 760‐4817), Anti‐NP AP (Cat# 760‐4827), and Discovery Yellow (Cat# 760‐239) were all purchased from Ventana.

Rats were treated orally BID for 3.5 days with either vehicle, 0.5% hydroxypropyl methylcellulose (HPMC)/0.1% Tween 80, or GSK3179106 prepared as a suspension in vehicle at 2.5 mL/kg prior to fecal pellet output assessment. One hour after the last oral dose of vehicle or GSK3179106 and immediately prior to fecal pellet output assessment, an intraperitoneal (ip) dose of vehicle, 100% propylene glycol, or neostigmine (0.1 mg/kg) was administered to all animals. Immediately following neostigmine administration, lower GI transit was assessed by measuring fecal pellet output. Rats were placed into empty cages for 2.5 hours, and the expelled fecal pellets were collected and counted every 30 minutes.

Following animal euthanasia, the colon was rapidly isolated, removed, and placed into ice‐cold oxygenated Krebs buffer. Krebs buffer prepared as a solution as follows: 120 mmol/L NaCl, 6 mmol/L KCl, 1.2 mmol/L MgCl 2 •6H 2 O, 1.2 mmol/L NaH 2 PO 4 , 2.5 mmol/L CaCl 2 •2H 2 O, 14.4 mmol/L NaHCO 3 , 11.5 mmol/L glucose. Colonic smooth muscle was separated from the mucosa by blunt dissection, oriented in a circular direction, and cut into strips (2 × 9 mm). Strips were mounted vertically in 10 mL organ baths (Radnoti LLC, Monrovia, CA, USA) filled with aerated Krebs solution, maintained at 37°C, and were allowed to equilibrate for 1 hour at an optimum tension of 0.75 g before recording contractility. Fifteen minutes prior to the first stimulation, GSK3179106 was dissolved in DMSO and diluted in Krebs buffer to obtain a final organ bath concentration of either 10, 100, or 1000 nmol/L. Electrical stimulation was delivered by a Grass stimulator (Grass Instruments, Quincy, MA, USA) and applied by pairs of platinum zigzag electrodes (Radnoti Glass Technology, Monrovia, CA, USA) lying parallel to the muscle strips. Contractility was recorded during a 10‐second electrical stimulation at frequencies of 8, 16, 32, and 64 Hz (pulse duration 0.5 ms, train duration 10 seconds, and voltage 60 V) with a 1‐minute rest period between each stimulation. Contractility was expressed as change in force (ΔForce (g/cm 2 )).

Rats were brought to the laboratory between 8:00 and 10:00 AM and euthanized by decapitation under isoflurane anesthesia. In an Ussing chamber system (World Precision Instruments, Sarasota, FL, USA), EFS‐stimulated active ion transport was measured in isolated colonic mucosa preparations bathed in oxygenated Krebs buffer, consistently maintained at 37°C, (concentrations in mmol/L: 120 NaCl, 6.0 KCl, 1.2 MgCl 2 •6H 2 O, 1.2 NaH 2 PO 4 , 2.5 CaCl 2 •2H 2 O, 14.4 NaHCO 3 , 11.5 glucose) in the presence of control (100 µL DMSO) or GSK3179106 (10‐1000 nmol/L) in the luminal side of the chambers. All experiments were conducted at the same time each day to ensure experimental reproducibility and to reduce experimental variability. Under voltage clamped conditions, to nullify the voltage potential difference (PD), a 5‐volt electrical field was applied using stimulus isolation unit Grass SIU5 (Auto‐med Inc, Warwick, RI) at stimulation frequencies of 2, 4, 8, 16, 32, and 64 Hz to the bath for 5 seconds with a 1‐minute rest period between each stimulation and the short circuit current ( I SC ) was recorded at each frequency. Following the first EFS series, the compound or control Krebs buffer was drained from the chambers. Fresh oxygenated Krebs solution was added to the chambers, and the tissue was allowed to equilibrate for an additional 20 minutes. After equilibration, control or GSK3179106 was added to the luminal side of the chambers and a second series of EFS was performed as described above. Tissues that received control prior to the 1st EFS series received compound prior to the 2nd EFS series and vice versa. Active ion transport was expressed as Δ I SC (μA/cm 2 ). For ligand‐stimulated preparations, an EC 50 of carbachol (3.8 μmol/L), bethanechol (42.1 μmol/L), lubiprostone (2.6 μmol/L), linaclotide (2.0 μmol/L), vasoactive intestinal peptide (430 nmol/L), or serotonin (3.2 μmol/L) ligand was used.

RET in situ hybridization by RNAscope followed by sequential immunostaining against synaptophysin was achieved on an autostainer Ventana Discovery Ultra (Ventana, Tucson, AZ, USA). Software version 4 was used for RNAscope, and Universal protocol was used for synaptophysin staining. On the day of the experiment, formalin‐fixed and paraffin‐embedded (FFPE) slide sections, cut at 4.0 m, were transferred to Superfrost slides (Thermo Fisher Scientific, Waltham, MA, USA), air‐dried for 1 hour, and baked in a 60°C oven for 1 hour prior to loading onto the autostainer. Pretreatments selected in the protocol for RNAscope included slide baking, H 2 O 2 blocking (pretreatment 1, ACD, Newark, CA, USA), epitope retrieval (pretreatment 2, ACD), and protease digestion (pretreatment 3, ACD) were performed. Following pretreatments, slide sections were incubated with Hs RET, Hs PPIB, Hs ChAT, Rn RET, or Rn PPIB probe (ACD) at 43°C for 2 hours. Signal amplification was achieved using dispensing kits (Cat #760‐236, Ventana) filled with AMP reagents 1‐7 from ACD (Cat# 322260, Newark, CA, USA). Detection was achieved using Discovery mRNA RED Detection kit (Ventana). Sequential staining with anti‐synaptophysin antibody followed with an initial antigen retrieval with Tris‐based (EDTA) buffer solution, CC1 (Ventana). Rabbit anti‐human synaptophysin, clone MRQ‐40 (Ventana), or IgG isotype control (Invitrogen, Waltham, MA, USA) was incubated for 32 minutes at 36°C followed by detection using anti–Rb NP (Ventana), anti–NP AP (Ventana), and Discovery Yellow (Ventana) kits. Slide sections were counterstained with hematoxylin (ACD), dehydrated, and coverslipped. Images were captured using a Nanozoomer slide scanner (Hamamatsu, Bridgewater, NJ, USA).

Male Sprague‐Dawley rats (225‐250 g, ~7‐8 weeks old) were purchased from Charles River Laboratories (Wilmington, MA). Upon receipt, animals were housed 2 per cage in standard shoebox cages on Sanichip bedding, with standard rat chow and water available, ab libitum . Rats were housed on a 12:12 light:dark cycle (lights on at 6:00 AM) under controlled temperature (23 ± 2°C) and humidity (50% ± 20%). All animal studies were conducted in accordance with the GSK Policy on the Care, Welfare and Treatment of Laboratory Animals, and use was approved by the University of Oklahoma Health Sciences Center Institutional Animal Care and Use Committee (Approval #15‐071). In the current study, female rats were excluded due to estrus cycle effects on GI function that remain incompletely understood.

Ex vivo assays suggest that RET plays a functional role in ENS signaling and may specifically influence the transmission of cholinergic signals. Furthermore, cholinergic stimulation is known to induce GI prokinetic effects in both experimental models and human patients. 17 - 19 The acetylcholinesterase inhibitor, neostigmine, increases local concentrations of acetylcholine at the neuronal synapse by inhibiting the catabolism of acetylcholine, resulting in the potentiation of cholinergic signaling and increased GI motility. In a preclinical rat model, the administration of neostigmine induces an increase in lower GI motility as measured by an increase in fecal pellet excretion. 18 This model was utilized to determine the effect of RET inhibition on neostigmine‐induced colonic motility. A single intraperitoneal injection of neostigmine alone significantly increased the excretion of fecal pellets, whereas GSK3179106 alone had no effect on normal lower GI motility (Figure 6 ). However, oral administration of the RET inhibitor, GSK3179106, attenuated (44% inhibition; F (3,26) = 19.45, P < 0.0001, one‐way ANOVA) neostigmine‐induced lower GI motility in comparison with rats dosed with neostigmine alone suggesting a functional role for RET in cholinergic control of colonic motility.

To understand the effect of RET inhibition on colonic smooth muscle function, contractility of the colon was examined in an ex vivo organ bath. While treatment of colonic muscle preparations with EFS‐induced frequency‐dependent colonic muscle contractions, there were no significant effects of the drug treatment at any frequency ( F (3,55) = 0.829, P = 0.483 at 8 Hz; F (3,55) = 0.561, P = 0.643 at 16 Hz; F (3,55) = 2.697, P = 0.055 at 32 Hz; F (3,55) = 2.000, P = 0.125 at 64 Hz, one‐way ANOVA). The addition of the RET inhibitor, GSK3179106, had a modest effect (9%–36% inhibition) on EFS muscle contractions at the highest concentration (1000 nmol/L; P = 0.026 at 32 Hz; P = 0.047 at 64 Hz, one‐way ANOVA) (Figure 5 ) suggesting only a minor role for RET inhibition in the attenuation of EFS‐induced muscle contraction.

RET inhibition effect on active ion transport induced by bethanechol, lubiprostone, linaclotide, vasoactive intestinal peptide, and serotonin. GSK3179106 at 10, 100, and 1000 nmol/L had no effect on active ion transport in colonic mucosa preparations when stimulated with (A) bethanechol (EC 50 =42.1 μmol/L, n = 8/group), (B) lubiprostone (EC 50 =2.6 μmol/L, n = 8/group), (C) linaclotide (EC 50 =2.0 μmol/L, n = 8/group), (D) vasoactive intestinal peptide (EC 50 =430 nmol/L, n = 7‐8/group), or (E) serotonin (EC 50 =3.2 μmol/L, n = 5‐8/group) compared to vehicle. Representative traces of the effect of vehicle and 100 nmol/L GSK3179106 on the increase in Isc in response to each of the secretagogues are adjacent to the graphed data

RET inhibition decreases the response of colonic mucosa to electrical field stimulated or carbachol‐induced active ion transport. GSK3179106 at (A) 10 ( n = 10), (B) 100 ( n = 10), or (C) 1000 nmol/L ( n = 10) significantly decreased the change in I sc when stimulated with EFS from 2‐64 Hz. * P < 0.05, ** P < 0.01, **** P < 0.0001 Control compared to GSK3179106, repeated‐measure two‐way ANOVA, Bonferroni post‐test. (D) GSK3179106 at both 100 and 1000 nmol/L inhibits carbachol‐induced change in I sc ( n = 8, all groups). (E) Representative trace of the inhibitory effect of GSK3179106 on the carbachol‐induced increased in Isc. *** P = 0.0005, **** P < 0.0001 compared to vehicle, one‐way ANOVA followed by a Bonferroni post‐test

The ENS regulates multiple functions of the GI tract including peristalsis and secretion. 15 , 16 To determine the effect of RET inhibition on GI mucosal secretion, colonic mucosal preparations were examined in an ex vivo organ bath Ussing chamber system for an effect on active ion transport assessed via electrophysiological measurements of short circuit current (Isc). GSK3179106, at concentrations up to 1000 nmol/L, decreased (17%‐40% inhibition) active ion transport in response to electrical field stimulation (EFS) (main effect of GSK317106 treatment: 10 nmol/L: F (1,9) = 10.34, P = 0.011; 100 nmol/L: F (1,9) = 15.07, P = 0.004; 1000 nmol/L: F (1,9) = 27.01, P = 0.0001; two‐way ANOVA) (Figure 3 A–C). To understand the type of neurons that were specifically being stimulated by EFS, the non‐selective acetylcholine receptor agonist, carbachol, was used to stimulate cholinergic signaling. Similar to the findings with EFS‐induced response, GSK3179106 was able to decrease (4%‐37% inhibition; F (3,28) = 16.00, P < 0.0001, one‐way ANOVA) carbachol‐induced active ion transport (Figure 3 D). In contrast, RET inhibition had no effect when bethanechol, a muscarinic receptor‐selective acetylcholine receptor agonist, was used to stimulate active ion transport ( F (3,28) = 0.982, P = 0.415, one‐way ANOVA) (Figure 4 A). RET inhibition, moreover, had no effect on active ion transport induced by lubiprostone, a fatty acid agonist of chloride channel protein 2 ( F (3,28) = 0.884, P = 0.462, one‐way ANOVA) (Figure 4 B), linaclotide ( F (3,28) = 0.009, P = 0.999, one‐way ANOVA) (Figure 4 C), a peptide agonist of guanylate cyclase, when applied to the luminal side of the tissue, or vasoactive intestinal peptide (VIP) ( F (3,27) = 0.025, P = 0.995, one‐way ANOVA) (Figure 4 D), serotonin (5‐HT) when applied to the basolateral side of intestinal preparations ( F (3,25) = 0.622, P = 0.607, one‐way ANOVA) (Figure 4 E), thereby excluding a functional role for RET with cells expressing receptors for these various secretagogues.

RET is expressed in neurons of the submucosal and myenteric plexus of human and rat colon. Representative images of human (A) or rat (C) submucosal and human (B) or rat (D) myenteric ganglion incubated with antisense probe to human RET mRNA (red, A and B) or rat RET mRNA (red, C and D) and stained with an antibody against synaptophysin (yellow, A‐D). Scale bar =50 μm for A and B or 100 μm for C and D

RET expression in the adult peripheral nervous system has been described in the ENS within myenteric and submucosal ganglia. 10 , 13 , 14 To determine the location of RET‐expressing neurons in adult colon, human and rat colon tissues were probed for transcripts encoding RET and counterstained with an antibody to synaptophysin to label neurons. A subset of neuronal cell bodies within the submucosal and myenteric ganglion expressed both RET transcripts and synaptophysin in human (Figure 1 A,B) and rat (Figure 1 C,D) colons. Furthermore, RET transcript‐expressing cell bodies frequently co‐expressed ChAT defining them as cholinergic (Figure 2 A,B).

4 DISCUSSION

In the current study, we sought to explore a role for RET in adult ENS. We demonstrated that RET transcripts are expressed in mature ENS submucosal and myenteric ganglia, and using a selective RET inhibitor, we found that RET is functionally involved in the transmission of cholinergic secretomotor activity and lower GI motility. These findings provide a link to the identity and function of RET‐expressing neurons in the adult ENS and how RET inhibition may be of therapeutic value for the treatment of intestinal disorders associated with increased secretion and accelerated GI transit such as IBS‐D.

Recent work has identified the location of RET, GFRα1, and GFRα2 expression in neurons of both the submucosal plexus and the myenteric plexus of adult human colon.10 This study confirms and extends these findings by identifying RET‐expressing neurons within the submucosal and myenteric ganglia with the use of a specific antisense RNA probe for human or rat RET transcripts. A subset of submucosal and myenteric ganglia exhibit high levels of RET expression as evidenced by the RET antisense probe labeling of these cells. Moreover, in a subset of neurons the co‐expression of ChAT identified these RET‐expressing neurons as cholinergic.

To understand the functional implications of RET inhibition within the bowel, we used GSK3179106, a potent, selective, reversible, and GI‐restricted RET inhibitor that is equally potent against the human and rat RET enzymes.12 It is active in multiple cell‐based assays of functional RET activity and is highly selective for RET as demonstrated by in vitro kinase assays and proteomic profiling in colon lysates. However, while we are confident that RET is the likely target for the observed findings utilizing GSK3179106 based on our findings and published literature, we cannot rule out the potential effects GSK3179106 may have on DDR1 and DDR2 as well as other kinases at high concentrations. Moreover, GSK3179106 is minimally absorbed through the GI tract when dosed orally resulting in low systemic bioavailability and negligible brain penetration.12 These characteristics make GSK3179106 a useful tool for studying the effects of RET inhibition within the ENS without affecting systemic or central targets. Furthermore, GI restriction potentially provides for a higher therapeutic index and a better safety profile for a drug to treat intestinal disorders by eliminating side effects known to occur with systemically available kinase inhibitors.20

In the current study, ex vivo studies of the secretomotor reflex were conducted. The secretomotor reflex is regulated by multiple neurons and their stimulation results in the release of either VIP and/or acetylcholine from secretomotor neurons that induce enterocytes to secrete Cl‐ into the lumen.15, 21-23 Inhibition of RET in rat colon mucosa preparations was found to decrease electrically induced ion transport indicative of an inhibitory effect on the secretomotor reflex. Confirmation of neuronal involvement was made by the demonstration that the RET inhibitory effect was observed in the presence of a cholinergic agonist, carbachol. As carbachol non‐selectively stimulates both nicotinic and muscarinic receptors, it remains possible that RET inhibition affects only a subset of cholinergic responsive cells.24-26 Interestingly, RET inhibition had no effect on bethanechol (a muscarinic selective agonist)‐stimulated ion transport indicating that RET is unlikely to be expressed within cells expressing muscarinic receptors. M 1 and M 3 receptors have been localized to the mucosal epithelium, suggesting that bethanechol directly stimulates enterocytes.27 Similarly, RET inhibition had no effect on secretagogue‐induced ion transport using serotonin, VIP, lubiprostone, and linaclotide, which may indicate that secretagogue‐responsive cells do not express RET. In fact, enterocytes did not express RET transcripts (data not shown); however, intestinal enterocytes have been shown to express receptors for the secretagogues assayed, likely indicating a direct stimulatory effect of the secretagogues on enterocytes.27-31 Taken together, these findings suggest RET inhibition may affect the function of neurons expressing presynaptic nicotinic receptors.32 Thus, the demonstration of RET inhibition on cholinergic neuron function may indicate the potential to affect other cholinergic GI functions such as the peristaltic reflex.15

The ENS coordinates muscle contraction and relaxation to enable GI peristalsis. Cholinergic stimulation of enteric smooth muscle drives proximal contractions that result in a GI prokinetic effect.17-19 To understand the effect of RET inhibition on colonic smooth muscle function, colonic smooth muscle contractility was examined in an ex vivo organ bath. Although ENS control of muscle contractility has been well established, RET inhibition had a modest inhibitory effect on smooth muscle contractility and only at high concentrations of RET inhibitor tested (1 μmol/L) suggesting a rather minor role for RET inhibition in the attenuation of EFS‐induced muscle contraction and likely may represent an off‐target effect given the high concentration of inhibitor at which the effect was observed. In contrast, in an in vivo model of neostigmine‐stimulated lower GI transit, RET inhibition attenuated the induced prokinetic effect demonstrating that RET inhibition can functionally attenuate increased neural motor activity controlling proximal smooth muscle contractions.