Source of dopamine in gastric juice and luminal dopamine‐induced duodenal bicarbonate secretion via apical dopamine D₂ receptors

Abstract

Background and Purpose

Dopamine protects the duodenal mucosa. Here we have investigated the source of dopamine in gastric juice and the mechanism underlying the effects of luminal dopamine on duodenal bicarbonate secretion (DBS) in rodents.

Experimental Approach

Immunofluorescence, UPLC‐MS/MS, gastric incubation and perfusion were used to detect gastric‐derived dopamine. Immunofluorescence and RT‐PCR were used to examine the expression of dopamine receptors in the duodenal mucosa. Real‐time pH titration and pH_i measurement were performed to investigate DBS.

Key Results

H⁺‐K⁺‐ATPase was co‐localized with tyrosine hydroxylase and dopamine transporters in gastric parietal cells. Dopamine was increased in in vivo gastric perfusate after intravenous infusion of histamine and in gastric mucosa incubated, in vitro, with bethanechol chloride or tyrosine. D₂ receptors were the most abundant dopamine receptors in rat duodenum, mainly distributed on the apical membrane of epithelial cells. Luminal dopamine increased DBS in a concentration‐dependent manner, an effect mimicked by a D₂ receptor agonist quinpirole and inhibited by the D₂ receptor antagonist L741,626, in vivo D₂ receptor siRNA and in D₂ receptor ^−/− mice. Dopamine and quinpirole raised the duodenal enterocyte pH_i. Quinpirole‐evoked DBS and PI3K/Akt activity were inhibited by calcium chelator BAPTA‐AM or in D₂ receptor^−/− mice.

Conclusion and Implications

Dopamine in the gastric juice is derived from parietal cells and is secreted along with gastric acid. On arrival in the duodenal lumen, dopamine increased DBS via an apical D₂ receptor‐ and calcium‐dependent pathway. Our data provide novel insights into the protective effects of dopamine on the duodenal mucosa.

Abbreviations

cAMP

cyclic AMP

CFTR

cystic fibrosis transmembrane conductance regulator

DBS

duodenal bicarbonate secretion

DDC

DOPA decarboxylase

I_SC

short‐circuit current

K‐HS

Krebs–Henseleit solution

TTX

tetrodotoxin

UPLC‐MS/MS

ultra‐performance LC tandem MS

What is already known

• Anti‐dopaminergic drugs induce ulcers, and dopamine receptor agonists prevent the formation of gastroduodenal mucosal ulcers.

• Duodenal bicarbonate secretion (DBS) is the most important defence mechanism against gastric acid‐peptic injury.

What this study adds

• Gastric parietal cells can release dopamine along with gastric acid secretion.

• Dopamine in duodenal lumen dose‐dependently increased DBS via an apical D₂ receptor‐ and calcium‐dependent pathway.

What is the clinical significance

• Gastric‐derived dopamine protects the duodenal mucosa when gastric acid secretion increases.

• Dopamine precursors or D₂ receptor agonists, given orally, could provide a therapeutic approach to ulcers.

1. INTRODUCTION

Duodenal bicarbonate secretion (DBS) is currently accepted as the most important defence mechanism against gastric acid‐peptic injury (Isenberg, Selling, Hogan, & Koss, 1987; Sjoblom, Jedstedt, & Flemstrom, 2001; Yin et al., 2018). Decreased DBS contributes to the pathogenesis of duodenal ulcers (Wen et al., 2018).

Dopamine is an important neurotransmitter and modulator in the gastrointestinal epithelium (Li et al., 2019; Yang et al., 2017). Patients with dopaminergic deficiency suffer from a higher incidence of peptic ulcers (Altschuler, 1996), whereas dopaminergic hyperfunction is associated with lower incidence of ulcers (Ozdemir et al., 2007). The available evidence suggests that dopamine may contribute to the preservation of the gastrointestinal mucosa. Dopamine receptors are widely present in the gastrointestinal tract (Al‐Jahmany, Schultheiss, & Diener, 2004) and can be grouped into two families, D₁‐like and D₂‐like receptors. The D₁‐like receptor family includes D₁ and D₅ receptors, which elevate intracellular cytosolic cAMP levels by activating adenylate cyclase. The D₂‐like family includes D₂, D₃, and D₄ receptors, all of which decrease intracellular cAMP and/or modulate the Ca²⁺ and phospholipase C pathway (Pivonello et al., 2007). Activation of D₂ receptors decreases the severity of ulcerative colitis through attenuation of vascular permeability (Tolstanova et al., 2015) and modulates innate immunity by affecting the transcription of pro‐inflammatory cytokines (Shao et al., 2013). Intravenous infusion of dopamine or the D₁‐like receptor agonist SKF‐38393 stimulated in vivo DBS (Flemstrom, Safsten, & Jedstedt, 1993) and inhibited the formation of duodenal ulcers (Desai et al., 2008), while D₂‐like receptor antagonists ameliorated small intestinal ulceration by activating α7 nicotinic acetylcholine receptors (Yasuda et al., 2011). However, many discrepancies remain in the field.

The source of the dopamine found in the upper gastrointestinal tract is still unclear. Co‐location of tyrosine hydroxylase (the rate‐limiting enzyme for dopamine synthesis) and the proton pump (a unique feature of parietal cells) in the gastric mucosa suggests that parietal cells might produce dopamine (Eisenhofer et al., 1997). Epithelial HCO₃ ⁻ secretion is under the control of several cellular signalling pathways, most prominently the cAMP and Ca²⁺ signalling pathways (Jung & Lee, 2014). We previously reported that dopamine receptors are widely distributed in the duodenal mucosa and involved in the regulation of ion transport (Feng et al., 2013) and epithelial permeability (Feng et al., 2017). However, it remains unknown whether the gastric juice contains dopamine derived from parietal cells and whether dopamine stimulates DBS when it arrives at the duodenal lumen, after gastric emptying.

We have tested the possibility that gastric juice contains dopamine derived from gastric parietal cells, that dopamine enters the duodenal lumen along with the gastric chyme, and that the luminal dopamine in the duodenum stimulates DBS by combining with dopamine receptors on the duodenal epithelia. This study provides new experimental and theoretical evidence for the regulation of duodenal mucosal barrier function by dopamine and may contribute to the development of novel potential drug targets.

2. METHODS

2.1. Animals and tissue preparation

2.1.1. Ethics statement

All animal care and experimental procedures were performed in accordance with the guidelines established by the National Institutes of Health (NIH, USA). All experimental operations were approved by the Animal Care and Use Committee of Capital Medical University, Beijing, China. Animal studies are reported in compliance with the ARRIVE guidelines (Kilkenny, Browne, Cuthill, Emerson, & Altman, 2010; McGrath & Lilley, 2015) and with the recommendations made by the British Journal of Pharmacology. All efforts were made to minimize animal suffering, and the minimum number of animals necessary to produce reliable scientific data was used.

2.1.2. Animal care

Sprague–Dawley rats (RRID:RGD_728193, male, 16 weeks old, 210–240 g) and C57BL/6J mice (RRID:IMSR_JAX:000664, male, 8 weeks old, 20–25 g) were purchased from the Laboratory Animal Services Center of Capital Medical University. D₂ receptor knockout (D₂R^−/−) mice (RRID:IMSR_JAX:003190, male, 8 weeks old, 20–25 g, C57BL/6J background, stock number 003190) were provided by the Institute of Laboratory Animal Sciences, Chinese Academy of Medical Science. PCR was used for genotyping. The verified primers used in the present study were as follows:

Common Reaction A, 5′‐TGA TGA CTG GGA ATG TTG GTG TGC‐3′;

• Reverse Reaction A, 5′‐CTC CCC AGA GTT GTG GCA AAA GG‐3′;

• Reverse Reaction A, 5′‐AGG ATT GGG AAG ACA ATA GCA G‐3′.

The expected PCR products were as follows: mutant (D₂R^−/−) = 329 bp; heterozygote (D₂R^+/−) = 329 bp and 221 bp; wild type (WT) = 221 bp. All animals were housed in an animal care facility at a temperature of 23°C under a light–dark cycle of 12:12 h and given free access to standard rodent laboratory chow and water.

2.1.3. Tissue preparation

All animals were killed by excessive anaesthesia with isopentane. The abdominal wall was opened, and the duodenum was quickly removed and immersed in Ringer's solution for DBS measurement or in Krebs–Henseleit solution (K‐HS) for other experiments. The duodenum was then divided into two segments. Each segment (1 cm long) was cut longitudinally along the mesenteric border, washed free of its luminal contents, and pinned mucosal side down in a Sylgard‐lined petri dish containing ice‐cold oxygenated Ringer's solution or K‐HS solution. The serosa and muscularis were stripped away with fine forceps to obtain the duodenal mucosa preparations (including mucosa and some submucosal tissues).

2.2. Solutions

The K‐HS contained (in mM): NaCl 117, KCl 4.7, NaHCO₃ 24.8, KH₂PO₄ 1.2, MgCl₂·6H₂O 1.2, CaCl₂·2H₂O 2.5, and glucose 11.1. The solution was bubbled with a gas mixture of 95% O₂ and 5% CO₂, and NaOH or HCl was added to maintain the pH of the solution at 7.4. A 0.9% NaCl solution was used as HCO₃ ⁻‐free solution and gassed with 100% O₂. The Ringer's solution contained (in mM): NaCl 145.5, KCl 4, CaCl₂·2H₂O 1.2, and indomethacin 0.05.

2.3. In vivo interfering RNA (RNAi) technique

siSTABLE modified siRNA was purchased from Thermo Scientific (Lafayette, CO, USA). The siSTABLE‐Stability Enhanced siRNA was converted to the 2′‐hydroxyl annealed salted, dialysed, and sterile‐filtered duplex. The sequences used for D₂R silencing were as follows:

• Sense 5′‐GUG CAU GGC UGU AUC ACG AUU‐3′,

• Antisense 5′‐PUC GUG AUA CAG CCA UGC ACU U‐3′.

The scrambled siRNA used as the negative control has no target sequence in mice. The sequences of the scrambled siRNA were as follows:

• Sense 5′‐UGG UUU ACA UGU CGA CUA AUU‐3′,

• Antisense 5′‐PUU AGU CGA CAU GUA AAC CAU U‐3′.

D₂R siRNA or scrambled siRNA and Entranster™ in vivo complex were prepared according to the manufacturer's protocol (Engreen Biosystem Co, Ltd, Beijing, China). The siRNA was diluted in Entranster™, and 10% glucose was then added to the solution at room temperature. The complex (400 μl) containing 100‐μg D₂R or scrambled siRNA was randomly injected into the tail veins of C57BL/6J mice by the investigators who were blinded to group assignment. Three days later, the mice were killed for experiments.

2.4. Immunohistochemistry

Segments of rat stomach or duodenum were embedded in optimum cutting temperature medium (McCormick, St. Louis, MO, USA) and immediately frozen in liquid nitrogen. The experimental details of the immunohistochemistry procedures conformed to the BJP guidelines (Alexander et al., 2018). Sections (6 μm) were cut at −20°C in a cryostat microtome (Leica CM1850; Leica Microsystems, St. Gallen, Switzerland) and thaw‐mounted onto polylysine‐coated frost plus slides. The sections were fixed for 15 min in cold acetone and then washed (3 × 5 min) in 0.3% Triton X‐100 phosphate buffer solution to eliminate residual fixative. After blocking with 10% normal goat serum (Sigma‐Aldrich) for 30 min at room temperature, the sections were incubated overnight at 4°C with primary antibody (Table 1) diluted in 1% BSA. After washing in phosphate buffer solution (3 × 5 min), the sections were incubated with the secondary antibody (Table 2) diluted in 1% BSA for 1 h at room temperature. Nuclei were stained with DAPI for 5 min. The negative control was treated in the same manner with omission of the primary antibody. After the final wash, the sections were coverslipped and observed under a laser scanning confocal microscope (FV1000; Olympus, Tokyo, Japan).

TABLE 1.

Primary antibodies used in the study

		Dilution
Antigen	Host species	IF	WB	Source/Catalog no.	RRIDs
D2R	Rabbit	1:100	1:200	abcam/ab21218	AB_2277536
H+‐K+‐ATPase	Mouse	1:200	N/A	Novus/NB300‐583	AB_10130553
H+‐K+‐ATPase	Mouse	1:4,000	N/A	abcam/ab2866	AB_303367
TH	Mouse	1:50	N/A	Sigma/T1299	AB_477560
DDC	Rabbit	1:200	N/A	Santa/sc‐99203	AB_2088975
PI3K	Rabbit	N/A	1:300	CST/4292	AB_329869
p‐PI3K	Rabbit	N/A	1:200	CST/4228	AB_659940
GAPDH	Rabbit	N/A	1:10000	Sigma/G9545	AB_796208

Abbreviations: D₂R, dopamine D₂ receptor; DDC, DOPA decarboxylase; IF, immunofluorescence; N/A, not applicable; PI3K, phosphoinositide 3‐kinase; p‐PI3K, phosphorylation of phosphoinositide 3‐kinase; WB, western blot.

TABLE 2.

Secondary antibodies used in the study

Secondary	Conjugation	Dilution	Source/Catalog no.	RRIDs
IF
Donkey anti‐mouse IgG	Alexa Fluor 488	1:1,000	Invitrogen/A21202	AB_141607
Donkey anti‐rabbit IgG	Alexa Fluor 488	1:1,000	Invitrogen/A21206	AB_2535792
Donkey anti‐rabbit IgG	Alexa Fluor 594	1:1,000	Invitrogen/A21207	AB_141637
Donkey anti‐mouse IgG	Alexa Fluor 594	1:1,000	Invitrogen/A21203	AB_141633
WB
Donkey anti‐rabbit IgG	IRDye800	1:10,000	Rockland/611‐132‐122	AB_220152

Abbreviations: IF, immunofluorescence; WB, western blot.

2.5. Measurement of dopamine by ultra‐performance LC tandem MS (UPLC‐MS/MS)

The dopamine content of the gastric perfusion and gastric incubated fluid was measured by UPLC‐MS/MS analysis (Zhang et al., 2015). Each sample was ultrasonically dissociated in a mixture of acetonitrile/methanol/formic acid (750:250:2, vol/vol/vol) for 2 min and centrifuged at 16000x g for 15 min at 4°C. The supernatant was evaporated to dryness and redissolved in reconstitution solvent; after another round of centrifugation at 1000x g and 4°C for 5 min, the supernatant was used immediately in UPLC‐MS/MS analysis (Key Laboratory of Radiopharmaceuticals, Ministry of Education, College of Chemistry, Beijing Normal University).

2.6. In vivo gastric perfusion

The procedure used to conduct in vivo gastric perfusion has been described in detail in previous reports (Brage et al., 1986; Zhang, Lu, Li, Wen, & Yang, 2019). In brief, rats were deprived of food for 24 h but allowed free access to water and then anaesthetized using isopentane inhalation. The stomach was intubated to facilitate luminal perfusion using one tube via the oesophagus and another via the duodenum. The anteroduodenal cannula was led outside the abdominal wall. The stomach cavity was perfused with warm 0.9% NaCl (6 ml·h⁻¹) by a peristaltic pump (Watson‐Marlow Ltd, Falmouth, England). After a 1‐h stabilization period, the pH of the effluent perfusate from the stomach was continuously recorded by means of a glass electrode coupled to a pH meter (TIM485 pH‐Stat Titrator, Radiometer‐Analytical SAS, Lyon, France). When the gastric acid output reached a steady state, the pH was recorded for 30 min as a measure of baseline acid secretion. Then, the response induced by intravenous infusion of histamine (2 mg·kg⁻¹·h⁻¹) was monitored. The pH recorded during 30 min of infusion was considered to represent stimulated acid and dopamine secretion. The dopamine content of the gastric perfusate was measured by UPLC‐MS/MS.

2.7. In vitro incubation of gastric mucosa preparations

In the in vitro gastric incubation experiment, rat gastric corpus mucosa preparations (Wang et al., 2012) were divided into four equal parts and incubated in Eppendorf tubes with 1.5‐ml K‐HS solution, which was maintained at 37°C and continuously gassed with 95% O₂/5% CO₂. The preparations were routinely treated with indomethacin (10 μM), a COX inhibitor, and TTX (1 μM), a neuronal Na⁺ channel blocker, to abolish the effects of endogenous PG and of remnants of the enteric nervous system, respectively. After a 30‐min equilibration period, the samples were subjected to a 30‐min period of drug stimulation. The tissue and the fluid content in the Eppendorf tubes were then carefully collected and analysed by UPLC‐MS/MS.

2.8. RNA extraction and real‐time RT‐PCR

Total RNA was extracted from rat duodenal mucosa preparations using TRIzol reagent (Invitrogen, Carlsbad, CA, USA). The cDNA was synthesized using the Superscript First‐Strand Synthesis System for RT‐PCR (Invitrogen). The mRNA encoding dopamine receptors in the samples was quantified by RT‐PCR as described previously (Wang et al., 2012).

2.9. Measurement of duodenal bicarbonate (HCO ₃ ⁻) secretion

The duodenal mucosa preparation was mounted between two modified Ussing chambers with an exposed area of 0.5 cm². The mucosal side was bathed in unbuffered HCO₃ ⁻‐free solution (0.9% NaCl solution) circulated by a gas lift with 100% O₂. The serosal side was bathed in K‐HS (pH 7.4) gassed with 95% O₂ and 5% CO₂. Each bath contained 5.0 ml of the respective solution maintained at 37°C by a heated water jacket. The luminal pH was maintained at 7.40 by the continuous infusion of 1‐mM HCl under the automatic control of a pH‐stat system (TIM485 pH‐Stat Titrator). The volume of titrant infused per unit time was used to quantitate HCO₃ ⁻ secretion. Measurements were recorded at 5‐min intervals. The rate of luminal HCO₃ ⁻ secretion was expressed as μmol·cm⁻²·h⁻¹. The duodenal mucosa preparations were routinely treated with indomethacin (10 μM) and TTX (1 μM) on the basolateral side. After a 30‐min period of measurement of baseline parameters, stimulants were added to the apical or basolateral side of the tissue in the Ussing chambers. Changes in DBS during the 30‐min period following the addition of stimulants were determined.

2.10. Fluorescence measurements of intracellular pH (pH _i) and image analysis

The method of measuring pH_i through imaging of villous epithelial cells in the intact murine duodenum has been described previously (Simpson, Gawenis, Walker, Boyle, & Clarke, 2005). Briefly, duodenal villi were isolated and immobilized using Cell‐Tak tissue adhesive. The villi were incubated for 5 min with K‐HS containing 100‐μM 1,4‐DTT to remove mucus and then incubated with 10‐μM BCECF‐AM for 15 min. After washing in K‐HS, three epithelial cells from the middle to upper region of each villus and a total of seven single villi from seven experiments were randomly selected for ratiometric analysis. Changes in pH_i were measured by dual‐excitation wavelength techniques (440 and 488 nm), and the villi were imaged at 535‐nm emission. Ratiometric images were obtained at 20‐s intervals using a camera and processed using Axon Imaging Workbench 2.2 (Axon Instruments, Union City, CA, USA). The 488/440 ratios were converted to pH_i values using a standard curve generated by the K⁺/nigericin technique.

2.11. Western blot analysis

The methods used for western blotting have been previously described (Feng et al., 2017), and the immuno‐related procedures used comply with the recommendations made by the British Journal of Pharmacology (Alexander et al., 2018). Briefly, tissue preparations were homogenized in cold lysis buffer supplemented with protease inhibitors for protein extraction. The protein extracts were centrifuged at 16000x g for 30 min at 4°C to remove cellular debris, and the resulting supernatants were stored at −80°C. The total protein concentration of the supernatants was determined using a Pierce™ BCA Protein Assay Kit (Thermo Scientific, 23225). The samples were dissolved in loading buffer containing 20% bromophenol blue and boiled (95°C) for 5 min prior to electrophoresis (80 V, 35 min; 120 V, 50 min) on a 10% sodium dodecyl sulphate/polyacrylamide gel. The separated proteins were transferred to a nitrocellulose membrane at 4°C (295 mA, 90 min) in transfer buffer; the membrane was then washed for 10 min with Tris‐buffered saline containing Tween 20 and blocked in blocking buffer for 2 h at room temperature. The blocked membrane was incubated with primary antibodies against D₂ receptors (RRID:AB_2277536) (Table 1) at 4°C overnight, washed three times for 10 min with TBST, and incubated with the secondary antibody (Table 2) for 1 h at room temperature. The membrane was then washed three times for 10 min with TBST, blocked for 2 h at room temperature in blocking buffer, and washed again with TBST. The blots were scanned on an Odyssey Infrared Imager (LI‐COR Biosciences, Lincoln, NE, USA) and were analysed using ImageJ software (RRID:SCR_003070) by another investigator who was blinded to group assignment.

2.12. elisa analysis

The duodenal mucosa preparations were placed in an Ussing chamber containing 5 ml of K‐HS gassed with 95% O₂ and 5% CO₂. After 30‐min equilibration, duodenal mucosa preparations were routinely treated with indomethacin and TTX on the basolateral side. All segments were randomly incubated with drugs applied to the apical side for 30 min at 37°C and then immediately flash‐frozen in liquid nitrogen. Intracellular levels of cAMP were determined using an elisa kit according to the manufacturer's instructions (Sigma‐Aldrich). The levels of phosphorylated Akt (pS473) and total Akt (Total) were also measured in a blinded elisa analysis using ab176657 kits (Abcam, Cambridge, UK).

2.13. Data and statistical analysis

The data and statistical analysis comply with the recommendations of the British Journal of Pharmacology on experimental design and analysis in pharmacology. (Curtis, Alexander et al., 2018). The littermates of rodents were equally divided into control and treatment groups by the investigators who were blinded to group assignment. To ensure blinding, the investigators who conducted the experiments, did not analyse the data. Statistics and graphs were generated using GraphPad Prism, version 6.0 (RRID:SCR_002798, GraphPad Software, San Diego, CA, USA). The results were statistically analysed using all the independent values and presented as arithmetic mean ± SEM. The sample sizes were the number of rodents in each group and were determined based on our previous studies, preliminary results and statistical test (n ≥ 5) (Curtis et al., 2018). Allowing for experimental failures, each protocol was repeated in at least seven individual animals. Statistical analysis was undertaken only using the independent values with n ≥ 5. P values >.05 were assumed to denote significant differences. Statistical comparisons between two groups were evaluated using the unpaired Student's t test. Statistical analyses among more than two groups were performed by one‐way or two‐way ANOVA with a post hoc Tukey's test. The drug effect in in vivo gastric perfusion through comparison of before and after drug application was tested by self‐control; in vitro experimental protocols were performed in parallel from separated samples of individual animals. The concentration–response relationships for dopamine‐ and quinpirole‐induced DBS and pH_i were calculated to obtain the EC₅₀ values for these compounds. The concentrations of dopamine and quinpirole were expressed using log‐transformed values.

2.14. Data availability

The authors declare that all data supporting the findings of this study are presented within the paper and its supporting information files. The data that support the findings of this study are available from the corresponding author upon reasonable request.

2.15. Materials

The drugs used in the present study were supplied as indicated: dopamine hydrochloride, indomethacin, dimethyl sulphoxide (DMSO), α‐methyl‐DL‐tyrosine methyl ester hydrochloride (α‐MDL), tyrosine, bethanechol chloride, omeprazole, histamine, 1,4‐DTT, Nigericin sodium salt, forskolin (FSK), R(+)‐SCH‐23390, L741,626, sulpiride hydrochloride, quinpirole, MDL12330A, BAPTA‐AM, LY294002 (Sigma‐Aldrich, St. Louis, MO, USA); tetrodotoxin (TTX) (Kangte Biological Engineering Co., Ltd, Xinghua, Jiangsu, China); BCECF‐AM (Beyotime Biotechnology Co, Ltd, Shanghai, China).

2.16. Nomenclature of targets and ligands

Key protein targets and ligands in this article are hyperlinked to corresponding entries in http://www.guidetopharmacology.org, the common portal for data from the IUPHAR/BPS Guide to PHARMACOLOGY (Harding et al., 2018), and are permanently archived in the Concise Guide to PHARMACOLOGY 2019/20 (Alexander, Christopoulos et al., 2019, Alexander, Fabbro et al., 2019, Alexander, Kelly et al., 2019).

3. RESULTS

3.3. Parietal cells are the main source of dopamine in the gastric mucosa

The UPLC‐MS/MS results showed that the dopamine content of the fluid incubated with the gastric mucosa in vitro was much higher than that of plasma obtained from blood samples. The dopamine content of gastric mucosa was approximately 60% of that found in the substantia nigra (SN) of rats (Figure 1a). Catecholamines, including dopamine, are derived from a common precursor, the amino acid tyrosine. Tyrosine is converted to l‐DOPA by the rate‐limiting enzyme tyrosine hydroxylase (TH), and l‐DOPA is converted to dopamine by DOPA decarboxylase (DDC) (Korner et al., 2019). The immunofluorescence results showed that TH and DDC were co‐localized in cells of the gastric corpus mucosa (Figure 1b). Application of tyrosine induced a concentration‐dependent increase of dopamine in the in vitro gastric mucosa‐incubated fluid. This effect was largely blocked by the TH inhibitor α‐methyl‐DL‐tyrosine methyl ester hydrochloride (α‐MDL) (Figure 1c). As shown in Figure 1d, the parietal cell marker H⁺‐K⁺‐ATPase was co‐localized with the dopaminergic cell markers TH and the dopamine transporter (DAT) in rat gastric corpus mucosa.

FIGURE 1.

Parietal cells are the main source of dopamine in the gastric mucosa. (a) Dopamine content of gastric incubation fluid and plasma and of gastric mucosa and substantia nigra (SN) measured by UPLC‐MS/MS in rats. *P < .05, significantly different from plasma (left) or SN (right); Student's unpaired t test. (b) The labelling immunofluorescence of TH and DDC in the gastric corpus mucosa of rat. Images are representative of three different animals. A nuclear marker, DAPI (blue), was used in the present study. Scale bars: 25 μm. (c) Dopamine content measured by UPLC‐MS/MS after application of tyrosine (.05, .1, and .5 mM) or pretreatment with TH inhibitor α‐methyl‐DL‐tyrosine methyl ester hydrochloride (α‐MDL, .05 mM). *P < 0.05, significantly different from control, ^# P < .05, significantly different from .5‐mM tyrosine; one‐way ANOVA with Tukey's test. (d) The labelling immunofluorescence of H⁺‐K⁺‐ATPase and TH/DAT in the gastric corpus mucosa of rat. Images are representative of three different animals. A nuclear marker, DAPI (blue), was used in the present study. Scale bars: 25 μm. (e) Dopamine content measured by UPLC‐MS/MS after application of bethanechol chloride (10 μM) and omeprazole (10 μM). *P < .05, significantly different from control; one‐way ANOVA with Tukey's test. (f) The pH of gastric perfusate after intravenous (i.v.) infusion of histamine (2 mg·kg⁻¹·h⁻¹). *P < .05, significantly different from control by Student's paired t test. (g) Dopamine content of effluent perfusate measured by UPLC‐MS/MS after intravenous infusion of histamine (2 mg·kg⁻¹·h⁻¹). *P < .05, significantly different from control; paired Student's t test. The data are expressed as mean ± SEM; n = 8 per group

To demonstrate the correlation of gastric acid secretion with dopamine release, gastric mucosa was incubated in vitro with pharmacological agents. Bethanechol chloride, which mimics the effect of vagal stimulation on gastric acid secretion, significantly increased dopamine release, while omeprazole, a proton pump inhibitor, markedly reduced dopamine release (Figure 1e). To determine whether dopamine could be released from gastric parietal cells into the gastric cavity, in vivo gastric perfusion was employed. Intravenous infusion of histamine significantly decreased the pH of the gastric perfusate (Figure 1f); at the same time, the dopamine content of the effluent perfusate was increased (Figure 1g).

3.4. Dopamine increased DBS

Although apical addition of dopamine did not affect the I _sc of rat duodenal mucosa preparations (Feng et al., 2013), enhanced DBS was detected through a real‐time pH titration experiment. The results showed that luminal (apical) but not serosal (basolateral) addition of dopamine (10 μM) produced a sustained upward DBS in the presence of indomethacin and TTX (Figure 2a). DBS was induced in a concentration‐dependent manner by apical application of dopamine (Figure 2b). DBS was also indirectly measured using pH‐sensitive fluorescent probe (BCECF‐AM) microfluorometry of intracellular pH (pH_i). Figure 2d shows the distribution of BCECF‐loaded cells in duodenal villi and the corresponding differential interference images. The results indicate that dopamine significantly increased the steady‐state pH_i in the superficial and upper villous epithelium (Figure 2c,d).

FIGURE 2.

Dopamine increases duodenal HCO₃ ⁻ secretion (DBS). (a) Apical (A) and basolateral (B) addition of dopamine (DA; 10 μM) produced DBS after routine basolateral addition of indomethacin (10 μM) and TTX (1 μM). (b) Apical addition of dopamine (DA; 0.01–100 μM) induced DBS in rats. (c) Dopamine (DA; 10 μM) increased intracellular pH in the chorionic epithelial cells of the rat duodenum. (d) Distribution of the pH‐sensitive microfluorometry probe BCECF‐AM after addition of dopamine (DA; 10 μM). Differential interference contrast (DIC) microscopic image showing the three‐dimensional morphology of a duodenal villus. Images are representative of seven different animals. Scale bars: 100 μm. *P < .05, significantly different from baseline; paired Student's t test. The data are expressed as mean ± SEM; n = 7 per group

3.5. Apical D₂ receptors mediated the dopamine‐induced DBS

To investigate which dopamine receptors mediate luminal dopamine‐induced DBS in the rat duodenum, the mucosa preparations were pretreated with dopamine receptor antagonists prior to apical addition of dopamine. Apical dopamine‐induced DBS was completely inhibited by apical pretreatment with the non‐selective D₂‐like receptor antagonist sulpiride (10 μM) or the specific D₂ receptor antagonist L741,626 (10 μM), while the non‐selective D₁‐like receptor antagonist SCH‐23390 (10 μM) had no effect (Figure 3a,b). Apical application of the D₂‐like receptor agonist quinpirole dose‐dependently mimicked the effect of the dopamine ‐induced increase in DBS (Figure 3c). The result obtained using the pH‐sensitive fluorescent probe BCECF‐AM indicated that quinpirole also significantly enhanced pH_i in a dose‐dependent manner (Figure 3d). At the same time, mRNA transcripts encoding all dopamine receptors were observed in the duodenal mucosa; among these, transcripts encoding D₂ receptors were the most abundant (Figure 3e). The immunofluorescence results showed that D₂ receptor immunoreactivity was principally distributed on the apical sides of intestinal crypts and Brunner glands (Figure 3f).

FIGURE 3.

Apical D₂ receptors (D₂Rs) mediate dopamine‐induced DBS in normal rats. (a) Effects of dopamine receptor (DAR) antagonists on the response to apical addition of dopamine (DA; 10 μM) induced DBS. SCH‐23390 (10 μM) (a non‐selective D₁‐like receptor antagonist), apical addition; Sulpiride (10 μM) (a non‐selective D₂‐like receptor antagonist), apical addition; L741,626 (10 μM) (the specific D₂ receptor antagonist), apical addition; forskolin (FSK; 10 μM) (the positive control for DBS), basolateral addition. (b) Summary of the effects of dopamine receptor antagonists on dopamine‐induced DBS. (c) Changes in DBS after apical addition of the D₂‐like receptor agonist quinpirole (0.01–100 nM). (d) Apical addition of quinpirole (1–100 nM) increased the intracellular pH of chorionic epithelial cells of the rat duodenum. (e) mRNA expression of dopamine receptors in the duodenal epithelium as analysed by real‐time RT‐PCR. β‐Actin was used as an internal control. (f) Haematoxylin–eosin (HE) staining and immunofluorescence of D₂ receptors in the rat duodenum. Images are representative of three different animals. Scale bars: 150 μm (upper) and 50 μm (lower). *P < .05, significantly different from control (a–d) or from D₂ receptors (e), ^# P < .05, significantly different from dopamine (b); one‐way ANOVA with Tukey's test. The data are expressed as mean ± SEM. n = 7 per group

To further determine whether D₂ receptors regulate dopamine‐induced DBS, we conducted a set of experiments in which we used D₂ receptor siRNA knockdown and D₂ receptor genetic knockout mice. Western blotting analysis revealed that D₂ receptor siRNA reduced the protein expression of these receptors in duodenal epithelial cells by 50%, relative to scrambled siRNA (Figure 4a). RT‐PCR was used to differentiate wild‐type (WT), heterozygous (D₂R^+/−), and homozygous (D₂R^−/−) animals (Figure 4b). Apical application of quinpirole significantly increased DBS in control and WT mice but failed to induce DBS in D₂ receptor knockdown mice and in D₂R^−/− mice (Figure 4d,e). In addition, baseline and quinpirole‐induced pH_i enhancement was also significantly decreased in D₂R^−/− mice (Figure 4c,f).

FIGURE 4.

Apical D₂ receptors (D₂R) mediate dopamine‐induced DBS in model mice. (a) Characterization of in vivo D₂ receptor siRNA knockdown mice by western blotting (GAPDH, d‐glyceraldehyde‐3‐phosphate dehydrogenase). (b) Characterization of the D₂ receptor knockout mice by PCR. (c) Distribution of the pH‐sensitive microfluorometry probe BCECF‐AM in the chorionic epithelial cells of D₂ receptor knockout (D₂R^−/−) mice after addition of quinpirole (100 nM). Images are representative of seven pairs of different animals. (d) DBS in the control (scramble siRNA) and in vivo D₂ receptor siRNA knockdown mice after apical addition of quinpirole (10 and 100 nM). (e) DBS in the wild‐type (WT) and D₂R^−/− mice after apical addition of quinpirole (10 and 100 nM). (f) Intracellular pH in the chorionic epithelial cells of D₂R^−/− mice after addition of quinpirole (100 nM). *P < .05, significantly different from control; unpaired Student's t test (a); *P < .05, significantly different from control/WT; paired Student's t test (d, e); *P < .05, significantly different from baseline, ^# P < .05, significantly different from WT; two‐way ANOVA with Tukey's test (f). The data are expressed as mean ± SEM. n = 7 per group

3.6. The calcium/PI3K/Akt pathway is involved in dopamine‐induced DBS

To delineate the intracellular signal pathways that mediate dopamine‐stimulated DBS via D₂ receptors, the adenylate cyclase inhibitor MDL12330A was used. Apical pretreatment of duodenal mucosa preparations with MDL12330A (10 μM) did not affect the induction of DBS by dopamine (10 μM) or the D₂‐like receptor agonist quinpirole (10 nM) (Figure 5a). Intracellular cAMP measurement further confirmed that neither apical addition of dopamine (10 μM) nor addition of quinpirole (100 nM) increased intracellular cAMP levels. In fact, these agents decreased the intracellular cAMP level in the duodenal mucosa (Figure 5b). It has been reported that Ca²⁺ signalling regulates DBS via a novel PI3K/Akt pathway (He et al., 2018). We further investigated the role of the Ca²⁺/PI3K/Akt pathway in dopamine‐induced DBS. Pretreatment of the duodenal mucosa preparations with the intracellular calcium chelator BAPTA‐AM (10 μM, apical addition) completely blocked the effect of dopamine on DBS (Figure 5c). To confirm the role of PI3K in the regulation of DBS via D₂ receptors, the protein levels of PI3K and phospho‐PI3K (p‐PI3K) in duodenal epithelium were measured. The basal protein levels of p‐PI3K/PI3K in control mice were higher than those in D₂R^−/− mice. Apical addition of quinpirole (100 nM) significantly increased the protein levels of p‐PI3K in WT mice but not in D₂R^−/− mice (Figure 5d). We also examined whether quinpirole induces the phosphorylation of Akt, the downstream effector of PI3K. Quinpirole rapidly enhanced Akt phosphorylation levels in duodenal mucosa preparations from WT mice but not in those from D₂R^−/− mice (Figure 5e). Furthermore, both the intracellular calcium chelator BAPTA‐AM (10 μM, apical addition) and the selective PI3K inhibitor LY294002 (20 μM, apical addition) significantly inhibited quinpirole‐stimulated duodenal phosphorylation of Akt in mice (Figure 5f).

FIGURE 5.

The calcium/PI3K/Akt pathway is involved in dopamine‐induced DBS. (a) Effects of the adenylate cyclase inhibitor MDL12330A (10 μM, apical addition) on dopamine‐ (DA; 10 μM, apical addition), or quinpirole‐ (10 nM, apical addition) induced DBS. (b) Intracellular cAMP levels after apical addition of dopamine (DA; 10 μM) or quinpirole (10 nM). (c) Effects of the intracellular calcium chelator BAPTA‐AM (10 μM, apical addition) on dopamine (DA)‐induced DBS (apical addition, 10 μM). (d) Protein levels of PI3K and p‐PI3K in the duodenal mucosa preparations of WT and D₂R^−/− mice after treatment with quinpirole (100 nM, apical addition). (e) The ratio of phosphorylated Akt (S473) to Akt in the duodenal mucosa preparations of WT and D₂R^−/− mice measured by elisa after treatment with quinpirole (100 nM, apical addition). (f) The ratio of phosphorylated Akt (S473) to Akt in the duodenal mucosa preparations after pretreatment with BAPTA‐AM (10 μM, apical addition) and LY294002 (a selective PI3K inhibitor, 20 μM, apical addition). *P < .05, significantly different from vehicle; one‐way ANOVA with Tukey's test (b); *P < .05, significantly different from dimethyl sulphoxide (DMSO) by unpaired Student's t test (c); *P < .05, significantly different from WT, ^# P < .05, significantly different from WT + quinpirole; one‐way ANOVA with Tukey's test (d, e); *P < .05, significantly different from vehicle, ^# P < .05, significantly different from quinpirole; one‐way ANOVA with Tukey's test (f). The data are expressed as mean ± SEM. n = 6–7 per group

4. DISCUSSION

In the present study, we demonstrated that dopamine was derived from gastric parietal cells and released into the gastric juice at the time of gastric acid secretion. When dopamine arrives at the duodenal lumen through the pylorus, luminal dopamine in the duodenum stimulates DBS. This effect is mediated by D₂ receptors located at the apical membrane of the duodenal epithelium through a calcium‐dependent pathway (Figure 6).

FIGURE 6.

Diagram of the working hypothesis of gastric dopamine‐induced duodenal bicarbonate secretion via an apical D₂ receptors‐ and calcium‐dependent pathway. Dopamine (DA) in the gastric juice is derived from parietal cells and is secreted along with gastric acid. Histamine stimulates parietal cells to release additional dopamine, which arrives at the duodenal lumen and binds to D₂ receptors (D₂R) on the duodenal apical epithelium, thereby increasing bicarbonate secretion through a calcium‐dependent pathway

4.1. Source of dopamine in duodenal lumen

Dopamine, as one of the neurotransmitters involved in the regulation of the gastrointestinal system, is not only released from enteric neurons (Tian et al., 2008) but is also produced by immune cells (Levite, 2016), intestinal flora (Wall et al., 2014), gastrointestinal epithelium (Eisenhofer et al., 1997), pancreas (Ustione & Piston, 2012), and from nutritional tyrosine (Korner et al., 2019). The duodenal juice contains a high level of dopamine after sympathetic nerve fibres are eliminated (Ustione & Piston, 2012), but the sources of the dopamine found are unclear. The low level of circulating dopamine in plasma indicates that the blood is not the main contributor of duodenal dopamine (Keller et al., 2004). We therefore postulated that the duodenal dopamine was derived from local sources rather than from blood and searched for possible sources in the GI tract. Enterochromaffin cells in the duodenum are possible sources of dopamine. However, the very scattered distribution of dopamine‐positive enterochromaffin cells in the duodenal epithelium suggests that these cells are not the major source of the dopamine in the duodenal juice (Mezey et al., 1996).

The presence of TH and DAT in H⁺‐K⁺‐ATPase‐positive gastric mucosal cells (Eisenhofer et al., 1997) and the presence of very high dopamine concentrations in the gastric juice after pyloric ligation in rats (Mezey, Eisenhofer, Hansson, Hunyady, & Hoffman, 1998) suggested that dopamine might be produced by gastric parietal cells. Many lines of evidence in the present study support this idea. First, the parietal cells positive for H⁺/K⁺‐ATPase were co‐localized with the dopaminergic markers TH and DDC, suggesting that parietal cells can produce dopamine. Second, gastric mucosa, in vitro, can take up tyrosine and then synthesize and release dopamine into the incubation fluid. Third, in the study of in vitro gastric incubation, application of bethanechol chloride, a muscarinic receptor agonist that increases gastric acid secretion, also enhanced the dopamine content of the incubation fluid. In addition, the proton pump inhibitor omeprazole markedly reduced the dopamine content of the fluid when gastric acid production was suppressed. Similarly, in in vivo experiments, intravenous infusion of histamine increased gastric acid secretion and enhanced the dopamine content of the effluent perfusate. According to the relative volumes of gastric juice (0.23 ± 0.1 ml) (Wang, Yin, Liu, Nie, & Xie, 2018) and 0.9% NaCl (6 ml·h⁻¹) perfused into gastric cavity of rats in the present study, we estimated the concentration of gastric‐derived dopamine was approximately 1.82–4.62 μM, which is consistent with the EC₅₀ (4.52 μM) of dopamine‐induced DBS. The high rate of dopamine release under acidic conditions strongly supports our hypothesis that gastric acid secretion is accompanied by the secretion of gastric‐derived dopamine, which then enters the duodenum with chyme through the pylorus.

4.2. The role of luminal dopamine in DBS

Clinical studies have confirmed that patients with duodenal ulcers have significantly diminished DBS compared with healthy volunteers (Isenberg et al., 1987). Although intravenous dopamine (50 μg·kg⁻¹·h⁻¹) was shown to increase DBS in in vivo experiments (Flemstrom et al., 1993; Josefsson, Bergquist, Ekman, & Tarkowski, 1996), the endogenous source of dopamine and the mechanism underlying dopamine‐induced DBS were still unclear. In the present study, we demonstrate the release of substantial amounts of dopamine into gastric juice by parietal cells and a significant enhancement of DBS by luminal dopamine. In particular, increased gastric H⁺ load significantly enhances dopamine secretion and this dopamine arrives at the duodenal lumen through the pylorus and increases subsequent DBS. This further suggests that dopamine can be synthesized in the rat gastric mucosa and that it may exert a protective effect on the gastrointestinal mucosa. The present study provides important experimental evidence for the possibility of developing clinical therapeutic approaches to duodenal ulcers through oral administration of dopamine precursors or D₂ receptor agonists.

Dopamine suppressed the severity of indomethacin‐induced small intestinal lesions, and the effect was blocked by D₂ receptor antagonists (Miyazawa, Matsumoto, Kato, & Takeuchi, 2003). Low doses of a D₂ receptor antagonist aggravated stress ulcerogenesis, while a D₂ receptor agonist attenuated stress ulcers (Puri, Ray, Chakravarti, & Sen, 1994). In our present study, luminal dopamine also enhanced DBS via apical D₂ receptors, based on the following evidence. First, dopamine treatment of the apical side, but not the basolateral side, of the mucosa increased DBS in a dose‐dependent manner; this suggests that dopamine in the duodenal juice, rather than dopamine in the plasma, has an important protective effect on the duodenal mucosa. Second, dopamine significantly increased pH_i in the upper villous cells, indicating that intracellular HCO₃ ⁻ production, secondary to the intracellular carbonic anhydrase conversion of CO₂ + H₂O in the duodenal epithelial cells and/or HCO₃ ⁻ import by the basolateral NaHCO₃ co‐transporter (NBC) was increased, which would be helpful in increasing HCO₃ ⁻ efflux from duodenal epithelial cells. Third, dopamine‐induced DBS was completely inhibited by the non‐selective D₂‐like receptor antagonist sulpiride and by the specific D₂ receptor antagonist L741,626. Fourth, apical application of the D₂‐like receptor agonist quinpirole mimicked the effect of dopamine on DBS, and this effect was absent in D₂ receptor siRNA and D₂R^−/− mice. Fifth, the fact that mRNA encoding D₂ receptors was present at the highest level and the distinct distribution of the D₂ receptor‐IF signal on the apical side of the rat duodenum provide strong morphological evidence for D₂ receptor‐mediated dopamine‐induced HCO₃ ⁻ production.

Epithelial HCO₃ ⁻ secretion in the digestive tract is mainly controlled by cytosolic cAMP and Ca²⁺ signalling pathways (Jung & Lee, 2014). The role of cAMP signalling in the regulation of HCO₃ ⁻ secretion by epithelial cells has been well characterized (Tuo et al., 2012). However, an inhibitor of adenylate cyclase, MDL12330A, had no effect on dopamine‐ or quinpirole‐induced DBS. Dopamine and quinpirole decreased the intracellular cAMP level in duodenal mucosa preparations, suggesting that the cAMP signalling pathway does not play a major role in dopamine‐induced DBS. Increasing evidence suggests that carbachol‐ and oestrogen‐induced Ca²⁺‐mediated DBS is mediated via the PI3K/Akt pathway (Tuo et al., 2012; Yang et al., 2017). In the present study, the calcium chelator BAPTA‐AM completely blocked dopamine‐induced DBS in a real‐time pH titration experiment. Both BAPTA‐AM and LY294002 significantly inhibited quinpirole‐stimulated Akt phosphorylation, suggesting the involvement of Ca²⁺/PI3K/Akt signalling in dopamine‐induced DBS.

DBS mainly involves the contributions of apical membrane Cl^‐/HCO₃⁻ exchangers and the cystic fibrosis transmembrane conductance regulator (CFTR) (Flemstrom & Isenberg, 2001). Considerable evidence supports the idea that the anion exchangers SLC26A3 (DRA) and SLC26A6 (PAT1) are involved in duodenal Cl⁻/HCO₃ ⁻ exchange. Opposite electrogenic Cl⁻/HCO₃ ⁻ stoichiometries operate in parallel to yield electroneutral HCO₃ ⁻ secretion and Cl⁻ absorption (Walker et al., 2009). PI3K activation, which results in downstream signalling through Akt phosphorylation, has been associated with activation of the SLC26A6 in intestinal epithelial cells (Lissner et al., 2010; Saksena et al., 2008). CFTR, which is usually measured by Isc expression, is also critical for the regulation of HCO₃ ⁻ and Cl⁻ transport in the duodenal epithelium (Tuo et al., 2009). In the present study, apical addition of dopamine induced DBS without changes in Isc expression in rat duodenal mucosa preparations, similar to the results obtained after genetic ablation of SLC26A6 (Tuo et al., 2006). Therefore, the contribution of the Cl⁻/HCO₃ ⁻ exchanger to dopamine‐induced DBS might be higher than that of CFTR.

The alkalinization of the cytosol in duodenal villous epithelial cells after application of dopamine or quinpirole may be induced by H⁺ extrusion and/or HCO₃ ⁻ import. Basolateral Na⁺/H⁺ exchanger type 1 (NHE1) regulates intracellular pH by extrusion of H⁺ into the subepithelial space (Pedersen & Cala, 2004; Praetorius et al., 2000), and PI3K and Akt activation increases NHE1 exchanger activity (H⁺ efflux) (Meima, Webb, Witkowska, & Barber, 2009; Sauvage, Maziere, Fathallah, & Giraud, 2000). Outside the duodenum, activation of the PI3K/Akt pathway leads to the stimulation of NBC in rat ventricular myocytes (Di Carlo et al., 2014).

In addition, oestrogen stimulates DBS in humans, and the expression of oestrogen receptors in duodenal mucosa does not differ between men and women. The basal DBS in adults is significantly higher in women than in men, while the time course and the net peak of 17β‐estradiol stimulated DBS are similar between men and women (Tuo et al., 2011). To avoid the influence of female hormones on DBS, we chose male rodents in these studies, which is a limitation of the present study.

In conclusion, the present study demonstrates that gastric parietal cells can release dopamine following gastric acid secretion. Luminal dopamine increases DBS through apical D₂ receptors in the rodent duodenum, and the Ca²⁺/PI3K/Akt pathway is involved in this process. This study reports that gastric‐derived dopamine regulates DBS, a finding that provides new insight into the modulation and protection of the duodenum by dopamine and suggests a potential drug target for the improvement of the chemical barrier function in the gut.

CONFLICT OF INTEREST

None of the authors have conflicts of interest to declare.

DECLARATION OF TRANSPARENCY AND SCIENTIFIC RIGOUR

This Declaration acknowledges that this paper adheres to the principles for transparent reporting and scientific rigour of preclinical research as stated in the BJP guidelines for Design & Analysis, Immunoblotting and Immunochemistry, and Animal Experimentation, and as recommended by funding agencies, publishers and other organisations engaged with supporting research.

Feng X‐Y, Yan J‐T, Li G‐W, et al. Source of dopamine in gastric juice and luminal dopamine‐induced duodenal bicarbonate secretion via apical dopamine D₂ receptors. Br J Pharmacol. 2020;177:3258–3272. 10.1111/bph.15047