Gastrointestinal hormones contribute to the beneficial effects of Roux-en-Y gastric bypass surgery (RYGB) on glycemic control. Secretin is secreted from duodenal S cells in response to low luminal pH, but it is unknown whether its secretion is altered after RYGB and if secretin contributes to the postoperative improvement in glycemic control. We hypothesized that secretin secretion increases after RYGB as a result of the diversion of nutrients to more distal parts of the small intestine, and thereby affects islet hormone release.
A specific secretin radioimmunoassay was developed, evaluated biochemically, and used to quantify plasma concentrations of secretin in 13 obese individuals before, 1 week after, and 3 months after RYGB. Distribution of secretin and its receptor was assessed by RNA sequencing, mass-spectrometry and in situ hybridization in human and rat tissues. Isolated, perfused rat intestine and pancreas were used to explore the molecular mechanism underlying glucose-induced secretin secretion and to study direct effects of secretin on glucagon, insulin, and somatostatin secretion. Secretin was administered alone or in combination with GLP-1 to non-sedated rats to evaluate effects on glucose regulation.
Plasma postprandial secretin was more than doubled in humans after RYGB (P < 0.001). The distal small intestine harbored secretin expressing cells in both rats and humans. Glucose increased the secretion of secretin in a sodium-glucose cotransporter dependent manner when administered to the distal part but not into the proximal part of the rat small intestine. Secretin stimulated somatostatin secretion (fold change: 1.59, P < 0.05) from the perfused rat pancreas but affected neither insulin (P = 0.2) nor glucagon (P = 0.97) secretion. When administered to rats in vivo, insulin secretion was attenuated and glucagon secretion increased (P = 0.04), while blood glucose peak time was delayed (from 15 to 45 min) and gastric emptying time prolonged (P = 0.004).
Glucose-sensing secretin cells located in the distal part of the small intestine may contribute to increased plasma concentrations observed after RYGB. The metabolic role of the distal S cells warrants further studies.
In the duodenum, enteroendocrine S cells secrete secretin in response to luminal acidification and thereby regulate the intraluminal milieu by stimulating the exocrine pancreas to secrete water and bicarbonate . Secretin, however, also has other actions including the inhibition of gastric emptying [2,3,4] and has been suggested to regulate the secretion of the pancreatic islet hormones [5,6,7,8,9,10].
Gastrointestinal hormones, including glucagon-like peptide-1 (GLP-1), are currently used for the treatment of metabolic diseases and endogenous GLP-1 contributes to the glucose lowering and weight reducing effect of Roux-en-Y gastric bypass (RYGB) [11, 12]. Whereas it is well established that GLP-1 secretion is increased after RYGB in humans , it is unknown if secretin secretion is increased.
Glucose is a powerful stimulus for GLP-1 secretion  and is rapidly absorbed in the upper part of the small intestine, therefore intraluminal concentrations are low in the distal part of the small intestine. We hypothesized that the delivery of glucose to the distal small intestine, as seen after the diversion following RYGB, stimulates secretin secretion by luminal-sensing mechanisms. To address this, we measured plasma secretin before and after RYGB. Furthermore, we isolated and perfused the upper or lower half of the rat small intestine and studied the effect of intraluminal glucose on secretin secretion as well as the molecular sensors involved.
To investigate the potential effects of secretin on blood glucose and secretion of islet hormones, we administered secretin subcutaneously to conscious rats and perfused, in separate experiments, the rat pancreas, testing the effects of secretin on glucagon, insulin, and somatostatin secretion.
Materials and methods
Human studies: Written informed consent was obtained from all study participants, and the study was approved by the Municipal Ethics Committee of Copenhagen in accordance with the Helsinki-II declaration and by the Danish Data Protection Agency, and registered at www.clinicaltrials.gov (NCT01993511).
Animal Studies: Animal studies were conducted with permission from the Danish Animal Experiments Inspectorate, Ministry of Environment and Food of Denmark, permit 2018-15-0201-01397, and in accordance with the EU Directive 2010/63/EU and guidelines of Danish legislation governing animal experimentation (1987), the National Institutes of Health (publication number 85-23), and approved by the local ethical committee (EMED, P18-336).
GLP-1 7–36NH2, rat and human secretin were obtained from Bachem (cat no. 4030663, cat no. 4037181 and cat no. 4031250, Bubendorf, Switzerland). Radioactive labeled rat and human secretin was obtained from Phoenix Pharmaceuticals, Inc (cat no. T-067–06 and T-067-07, CA, USA). Development and evaluation of a secretion radioimmunoassay are described in Supplementary Materials 1.
Mixed-meal tests before and after RYGB surgery in obese subjects
Plasma obtained during a standardized liquid mixed-meal test before, 1 week, and 3 months after RYGB from 13 obese subjects (type 2 diabetes; n = 4, impaired glucose tolerance; n = 3, normal glucose tolerance; n = 6) was analyzed. Glucose tolerance was determined by standard OGTT preoperatively. All samples were from a study by Martinussen et al. as previously described .
Distribution of secretin and GLP-1 along the gastrointestinal tract in humans and rats
Mass-spectrometry based detection was used to assess the distribution of secretin and GLP-1 (for comparison) in human intestinal tissue as described previously . The secretin and GLP-1 profiles are presented as peak area divided by tissue weight.
Whole-wall tissue biopsies (~1 cm) of esophagus, ventricle, duodenum, proximal jejunum, distal ileum, and colon were collected from non-fasted rats (anatomical definitions are listed in Supplementary Table 1). Peptides in tissue biopsies were extracted using trifluoroacetic acid (Supplementary Materials 2), and immunohistochemical staining of secretin-positive cells was performed on paraffin embedded tissue samples using anti-secretin (5585–3), a generous gift from Professor Jan Fahrenkrug (Supplementary Materials 3).
Male Wistar rats (200–250 g) were obtained from Janvier (Le Genest-Saint-Isle, France) and housed two to four rats per cage. Rats were allowed 1 week of acclimatization and kept on a 12 h light/dark cycle with ad libitum access to water and standard chow.
Isolated perfused rat small intestine and pancreas
Perfusion was performed using a single pass perfusion system (UP100, Hugo Sachs Harvard Apparatus, Germany). The rat small intestine and pancreas were surgically isolated (as described in Supplementary Materials 4). Each protocol started with a baseline period followed by the addition of various test substances. Test stimulants used in the perfused intestine included 0.1 M HCl, 0.1 M NaHCO3−, 0.05 M KCl (Sigma-Aldrich, Brøndby, Denmark), 20% (w/v) d-glucose (cat no. G8270, Sigma-Aldrich, Denmark), and 10 mmol/L Phloridzin (cat no. P3449, Sigma-Aldrich, Denmark). Test stimulants used in the perfused pancreas were 1 nmol/L GLP-1 7–36NH2 and both 20 pmol/L and 2 nmol/L secretin.
In vivo experiments in rats
Experiments were carried out on two occasions on fasted rats (300 ± 13 g) just before their nocturnal feeding period (5:00 PM). Rats were divided into weight-matched groups (n = 8/group). At −10 min, 200 μL tail blood was collected into prechilled EDTA-coated capillary tubes (catalog no. 200 K3E, Microvette; Sarstedt, Nümbrecht, Germany) and instantly transferred onto ice. At −5 min, 300 μL test solution was injected subcutaneously. Test solutions were isotonic saline, (negative control) or peptides prepared in isotonic saline: secretin (30 nmol/kg), GLP-1 (30 nmol/kg) or secretin + GLP-1 (both 30 nmol/kg). At 0 min, a bolus of d-glucose (2 g/kg) and acetaminophen (100 mg/kg), prepared in isotonic saline, was given orally. Rats from the same cage received different treatments. Blood was collected at times −10, −5, 5, 15, 30, 45, 60, and 90-min. Glucose was measured immediately after blood collection, while the remainder of the samples were instantly transferred onto ice and centrifuged (1650 × g, 4 °C, 10 min) within half an hour to obtain plasma. Plasma was transferred to Eppendorf tubes, immediately frozen on dry ice and stored at −20 °C until analysis.
Biochemical measurements of perfusion effluents, rat blood and rat plasma
Perfusion effluents: insulin was measured with antiserum (code no. 2006–3) which cross-reacts with rodent insulin I and II . Glucagon was measured with an antibody directed against the C-terminus (code no. 4305) as previously described . Total GLP-1 (the sum of 7–36NH2, 9–36NH2 and potential mid-terminal cleaved fragments) was measured using a C-terminal specific radioimmunoassay targeting amidated forms (code no. 89390) . Somatostatin was measured using a side-viewing antibody (code no. 1758–5), detecting all the bioactive forms of somatostatin [20, 21].
Blood glucose was measured using a glucometer (Accu-Chek Mobile, catalog no. 05874149001; Roche Diagnostics, Mannheim, Germany). Plasma concentrations of glucagon were measured by sandwich ELISA (catalog no. 10-1281-01; Mercodia AB, Uppsala, Sweden) . Plasma concentrations of insulin and C-peptide were measured using ELISAs (catalog no. 10-1250-01 and 10-1172-01; Mercodia AB, Uppsala, Sweden).
RNA sequencing of human islets
Publicly available RNA-seq dataset (from GSE85241, GSE81608 and E-MTAB-5061 [23,24,25]) were obtained. Average reads per kilobase million values for secretin receptor (Sctr) and GLP-1 receptor were uploaded to the Jupyter Notebook (http://jupyter.org/). Data were then log2 transformed and mean expression levels were calculated in alpha, beta, and delta cells, respectively. We excluded individuals with diabetes from these analyses. For further details about the donors, isolation of cells, and RNA-sequencing methods please see the original studies [23,24,25].
In situ hybridization on rat pancreases is described in Supplementary Materials 5.
Clinical samples and in vivo data: plasma concentrations of hormones were evaluated using area under the curve analysis and statistical testing by one-way ANOVA followed by Holm–Sidak multiple comparisons test. A mixed-effect model was applied in order to test the effect of treatment and time on the variables.
Perfusion experiments: data are expressed as mean ± SEM concentrations in venous effluents (pmol/L). Since the perfusion flow was kept constant through the experiments, this mirrors the actual secretion output (which can be calculated by multiplying with the flow rate). To test for statistical significance, mean values within the test period (based on 10 subsequent minutes) were compared with mean values in the baseline period (10 subsequent minutes prior to test stimulant administration) using statistical testing as described above.
Calculations were made using STAT14 (SE), College Station, Texas, USA. For illustrations, GraphPad Prism version 8.0 (GraphPad Software, La Jolla California USA) and Adobe CC software suite were used (San Francisco, CA, USA). P values < 0.05 were considered statistically significant. Data are shown as means ± SEM.
Development of a sensitive secretin assay
We generated titer curves for three different antibodies against human secretin (structure in Fig. 1a) using two radioactive iodine-labeled secretin peptides (termed tracer). Based on binding characteristics, the antibody named 5595–3 was selected for further testing and for the preparation of calibrator curves (Supplementary Fig. 1). By sequencing alignment, we found that human and rat secretin differ at position 14–16 (human: REG vs. rat: QDS) (Fig. 1a) but no species variation was found within the antibody’s epitope (position 18–27) (Fig. 1a). For rat samples, we therefore used the same antibody and tracer. For calibrating purposes, we included the rat isoform of secretin as calibrator control.
Recovery of human secretin added to human pooled plasma was calculated to 71 ± 11% (mean ± SD) with a lower limit of detection of 1 pmol/L and a dynamic range from 2.5 to 80 pmol/L (Supplementary Fig. 1) using solvent phase-extraction (70% ethanol). We did not observe cross-reactivity (not significantly different compared with background) toward CCK, glucagon, glicentin, GLP-1, human insulin, oxyntomodulin, neurotensin, and PYY at concentrations up to 300 pmol/L.
Comparison of commercial ELISAs
The two commercially available ELISAs had low recoveries of human secretin of 15 ± 8% (mean ± SD) in assay buffer. A Bland–Altman analysis (comparing the concentrations measured using the two ELISAs to the in-house developed RIA) showed acceptable (<2 SD) degree of variation for concentrations above 20 pmol/L, however, for physiological plasma concentrations of secretin (<20 pmol/L) the two commercial ELISAs were inadequate (Supplementary Fig. 2).
Having developed a specific and sensitive secretin radioimmunoassay, we next investigated whether postprandial plasma secretin concentrations are elevated after RYGB surgery.
RYGB surgery increases meal-induced secretin release in obese individuals
Fasting plasma concentrations of secretin were not significantly different after comparison with before RYGB (Fig. 1b, P > 0.45), whereas postprandial plasma secretin concentrations in response to a liquid meal test were significantly increased 3 months after compared with before RYGB (P < 0.05), reaching a two- to threefold higher peak value, and total AUCs were approximately doubled (tAUC0–18 min: 569 ± 475 vs. 956 ± 360 pmol/L × min) (Fig. 1c). Concentrations were also increased 1 week after RYGB but to a more modest degree and total AUCs did not differ from preoperative AUCs (P = 0.67).
Distribution of secretin in humans and rats
Using publicly available RNA-seq data from 37 different human tissues  expression profiles of secretin showed a maximum in the duodenum, as expected. However, high levels were also found in the more distal parts of the small intestine (Fig. 1d), consistent with previous reports on secretin distribution in other species [27, 28]. Mass-spectrometry analysis on human gastrointestinal tissue revealed that secretin concentrations were highest in the duodenum as well as in jejunal biopsies (Fig. 1e). Concentrations of GLP-1 7–36NH2 were included as positive controls. GLP-1 concentration was highest in the distal part of the gastrointestinal tract (ileum, colon, and rectum) (Fig. 1e) in line with previous reports .
To examine if the distribution of secretin in rats was similar to humans, we measured the concentration of extractable secretin from esophagus to colon in rats (Supplementary Table 1). Secretin was not detectable in esophagus and the ventricle (Fig. 1f, n = 7) but in the duodenum and in the distal ileum, secretin was found at comparable concentrations ~20–30 pmol/g tissue (Fig. 1f, n = 7). Concentrations of GLP-1 increased from duodenum to colon with the highest concentration in the distal ileum (~60 pmol/g tissue) (Fig. 1f, n = 7) in line with a previous report . Immunohistochemical stainings of secretin (Fig. 1g, h) showed no staining in esophagus, ventricle, and colon and high intensity in proximal jejunum and distal ileum, consistent with the measured extractable concentrations.
Secretin responses from the proximal and distal small intestine using the isolated perfused rat intestine model
Luminal infusion of HCl increased secretin secretion fourfold (Baseline: 3.7 ± 0.9 vs. HCl: 15.6 ± 2.3 pmol/L, P < 0.05, n = 6) (Fig. 2a, b). Following HCl infusion, the intestine was flushed with sodium bicarbonate (0.1 M) to neutralize luminal pH (Fig. 2a). This returned secretin secretion to pre-HCl-stimulatory levels. KCl (0.05 M) was infused intravascularly at the end of the experiment (Fig. 2a). Secretion of secretin increased significantly compared with preceding baseline secretion (Baseline: 5.2 ± 0.8 vs. KCl: 9.1 ± 0.9 pmol/L, P < 0.05, n = 6) (Fig. 2b). Our data therefore suggests that cell depolarization is involved in secretin release from the perfused rat small intestine and furthermore that our perfusion model reflects the physiology of secretin secretion. We therefore decided to evaluate potential differences in HCl and glucose-induced secretin secretion in the proximal and distal small intestine using this model.
Differential sensing of glucose by secretin producing cells in the proximal and distal small intestine
Luminal HCl infusion increased secretin secretion (P < 0.05) from both the proximal and distal half of the small intestine, but to the greatest extent in the distal half (baseline-subtracted values; Proximal. HCl: 7.6 ± 1.5 vs. Distal. HCl: 16.3 ± 2.6 pmol/L, P < 0.05, n = 6) (Fig. 2c, d). Of particular notice, distal, but not proximal, infusion of glucose resulted in a significant (P < 0.05) secretin response (baseline-subtracted values; Proximal. Glucose: 0.48 ± 0.3 vs. Distal. Glucose: 4.3 ± 1 pmol/L, n = 6) (Fig. 2c, d) suggesting that distally located secretin cells are capable of sensing glucose.
GLP-1 secretion was measured as this is a well validated control in this experimental setup [31, 32]. Secretion of GLP-1 in response to both glucose and HCl infusion was likewise significantly higher from the distal small intestine compared with the proximal small intestine (Fig. 2e, f), which is consistent with high extractable concentrations of GLP-1 in the distal small intestine (Fig. 1f).
Glucose-induced secretin secretion from the distal small intestine is mediated by sodium-glucose cotransporter
Inhibition of SGLT with the competitive SGLT1/2 inhibitor, phloridzin, eliminated glucose-stimulated secretin response from the distal rat small intestine (baseline-subtracted values; Glucose: 1.9 ± 0.7 vs. Glucose + Phloridzin: 0 ± 0.2 pmol/L, P < 0.05, n = 6) (Fig. 2g, h), indicating that sodium-coupled glucose absorption is involved in the mechanism of glucose-stimulated secretin secretion from the distal rat small intestine.
Consistent with previous reports [33, 34], phloridzin eliminated glucose-stimulated GLP-1 secretion from the distal rat small intestine (baseline-subtracted values; Glucose: 38.5 ± 10.4 vs. Glucose + Phloridzin: 2.6 ± 1.2 pmol/L, P < 0.05, n = 6) (Fig. 2i, j). HCl was included as a positive control for secretin secretion in the end of each experiment.
Secretin delays blood glucose peak time and increases plasma glucagon in non-sedated rats
Blood glucose concentrations were similar across groups at baseline (P > 0.71). Peak times for blood glucose were prolonged from 15 min (saline group) to 30 min (GLP-1), 45 min (secretin), and 60 min (secretin + GLP-1) (Fig. 3a) but the incremental area under the curve (iAUC0–90 min) was not significantly different between groups (P > 0.77, Fig. 3b). Gastric emptying, reflected by changes in plasma acetaminophen concentrations, was markedly prolonged in secretin-treated rats compared with saline and GLP-1 treated groups (P < 0.05, Fig. 3c, d). GLP-1 had significant effect on plasma acetaminophen at the initial time points compared with saline (timepoint 5 min, P < 0.05).
Plasma concentrations of insulin and C-peptide mirrored blood glucose levels (Fig. 3e–g). However, in rats receiving both secretin and GLP-1, insulin and C-peptide peaked 5 min after the oral glucose gavage at which timepoint no change in blood glucose levels was observed (Fig. 3a). When comparing iAUC0–90 min of insulin and C-peptide, there was a tendency toward a lowering effect of secretin and GLP-1, but neither reached statistical significance (Secretin vs. Saline; Pinsulin = 0.07 and Pc-peptide = 0.09, GLP-1 vs. Saline; Pinsulin = 0.09 and Pc-peptide = 0.55 and Secretin + GLP-1 vs. Saline; Pinsulin = 0.69 and Pc-peptide = 0.44) (Fig. 3f–h).
Glucose administration lowered plasma glucagon concentrations in all groups (Fig. 3i). However, secretin (either alone or in combination with GLP-1) treated rats had a significantly increased plasma concentration of glucagon during the first 5 min after administration (Secretin: 9.7 ± 1.1 vs. Saline: 6.5 ± 1.4 pmol/L, P = 0.04 and Secretin + GLP-1: 9.8 ± 0.7 vs. Saline: 6.5 ± 1.4 pmol/L, P = 0.03) but after 30 min, levels were similar to those in the control group (P > 0.05). Glucagon n-AUCs were lower in the group receiving GLP-1 compared with saline group (P = 0.08) and to the secretin-treated group (P < 0.05) (Fig. 3j).
Pancreatic delta cells express the Sctr and exogenous secretin increases somatostatin secretion from the perfused rat pancreas
To evaluate the potential metabolic effects of increased secretin release, we initially assessed tissue specific enrichment of the Sctr. We found that aside from the known expression of SCTRs in the duodenum, pancreatic tissue contained the Sctr transcript (Fig. 4a). However, since these values represent both expression in exocrine acinar/ductular tissue and islets, we next investigated if Sctr was present in human islets. Sctr was expressed in delta cells and detectable at lower levels in alpha and beta cells (Fig. 4b). Sctr expression were compared with Glp-1r profiles, which, expectedly, showed high expression in the beta cells and, to a lesser extent, in the delta cells (Fig. 4b). To investigate the translational relevance of rat islets to humans in regard to Sctr expression, we used an in situ hybridization approach. The Sctr was primarily found to be expressed in pancreatic delta cells and to a lesser extent in the alpha and beta cells of the rat (Fig. 4c).
To examine whether secretin has an insulinotropic effect on the rat pancreas, we perfused the rat pancreas with 10 mmol/L glucose and administered secretin at a concentration matching post-RYGB concentrations as well as a supra-physiological concentration (20 pmol/L and 2 nmol/L, respectively) (Fig. 4d, e). GLP-1 was infused at the end of the experiment (1 nmol/L) as a positive control for insulin and somatostatin release [35,36,37]. Secretion of insulin continuously increased throughout the experiment independent of secretin infusion (because of the elevated glucose concentration), and the baseline secretion of insulin was therefore calculated as the average secretion before and after secretin stimulation. Insulin did not increase in response to both of the applied concentrations of secretin (Baseline: 325 ± 34 vs. Secretin (20 pmol/L): 365.9 ± 35.2 pmol/L, P = 0.13 and Baseline: 426.8 ± 41.7 vs. Secretin (2 nmol/L): 454 ± 36.8 pmol/L, P = 0.56, n = 6) (Fig. 4e).
Consistent with the expression of the Sctr in rat and human delta cells, secretin infusion significantly increased the secretion of somatostatin (Baseline: 8.3 ± 1.1 vs. Secretin (20 pmol/L): 13.3 ± 1.9 pmol/L, P < 0.05 and Baseline: 9.8 ± 1.6 vs. Secretin (2 nmol/L): 14.9 ± 2.6 pmol/L, P < 0.05, n = 6) (Fig. 4f, g).
Glucagon secretion was low, due to the high glucose concentration in the perfusion buffer, and neither secretin nor GLP-1 infusions led to significant changes in its secretion (P > 0.18, n = 6) (Fig. 4h, i).
Here, we demonstrate that postprandial plasma concentrations of secretin are increased after RYGB surgery in obese individuals. This may result from the anatomical rearrangement of the small intestine which diverts luminal nutrients, like glucose, to more distal sites of the small intestine, where glucose is effectively absorbed and may give rise to increased secretion of some gut hormones that are more abundant in the distal part of the small intestine . Increased villus length after RYGB or accelerated gastric emptying may also contribute to the observed changes in plasma secretin concentrations . In addition, secretin is also produced to some extent in the biliary epithelial cells, and secretin secreted from these cells may influence total plasma secretin concentrations, although the relative importance of biliary secretin in this regard warrants further investigation. Tissue levels of secretin have been reported to be upregulated in a rodent model of RYGB  but postoperative plasma concentrations of secretin have to our knowledge not been reported previously.
Furthermore, we show that secretin, besides in the duodenum, is expressed in the distal small intestine in humans and rats, and using a physiologically relevant model: the perfused rat intestine , we demonstrate that glucose is a strong stimulus for secretin release from the distal but not from the proximal rat small intestine. Using an inhibitor of SGLT1/2 activity (phloridzin)  we found that the molecular mechanism responsible for glucose-induced secretin secretion involves glucose absorption through the SGLT. SGLT-2 inhibition has gained major clinical interest due to its glucose lowering effect and reduced risk of cardiovascular disease . One may therefore speculate that patients treated with nonselective SGLT inhibitors would have an altered secretin secretion profile, similar to what has been reported for GLP-1  but this remains to be explored. Previous attempts to show glucose-induced secretin secretion have been negative [7, 43,44,45], probably because in these studies glucose was administered to the proximal rather than the distal part of the small intestine.
Glucose-stimulated secretin release from the distal small intestine is probably not of major physiological importance in unoperated humans since glucose is mainly absorbed in the proximal part of the small intestine. However, after intake of large carbohydrate rich meals, some intraluminal glucose may reach more distal parts of the small intestine , and secretion under these circumstances may play a role together with GLP-1 in improving glycemic control by inhibition of gastric emptying. This putative effect is supported by our in vivo data and consistent with previous findings [2,3,4].
Intraluminal acidification in the distal small intestine resulted in a larger secretory secretin response compared with the proximal intestine. The physiological relevance of this finding is not clear. The acidity of the gastric chyme entering the upper part of the small intestine is rapidly neutralized by a mixture of bile, mucosal-, and pancreatic bicarbonate secretion, but the stimulatory effect of low pH in the distal small intestine on secretin-secreting cells may reflect a safety mechanism to reduce gastric emptying. The fact that also GLP-1 secretion was increased, may point to a more general effect, however, where hydrogen ions formed during digestion and perhaps fermentation in the mucosa micromilieu, stimulate ileal endocrine cells to activate the ileal brake. Further studies are required to investigate this new unexpected observation.
The observations regarding the effects of secretin on pancreatic islets are conflicting [6, 7, 47,48,49,50,51]. Our data support that physiological levels of secretin regulate the pancreatic secretion of somatostatin which is consistent with Sctr expression in rat and human delta cells. Although Sctr was also expressed by beta cells, secretin did not affect insulin secretion in the perfused rat pancreas whereas in vivo secretin actually led to the lowering of plasma insulin levels early after an OGTT. The underlying reason for this awaits further investigation but this may be related to prolonged gastric emptying time and to an increased secretion of somatostatin, which through paracrine effects may have overruled potential direct stimulatory effects of secretin on insulin secretion. Glucagon secretion was also not affected but this may be due to the already low levels when the pancreas was perfused in hyperglycemic conditions, thereby restricting the capability to detect further inhibition of alpha cells. However, to our surprise, there was a short-lasting increase in glucagon secretion after secretin injection in vivo, which was uninfluenced by the glucagonostatic effect of GLP-1  and sympathetic stress (as this was not observed in the control group), as observed when similar doses of both peptides were injected. The underlying reason(s) is not clear.
Our study expands the current knowledge regarding secretin physiology and suggest that the RYGB-related increase in plasma secretin concentrations is mediated by nutrients reaching S cells located in the distal small intestine. The physiological role of glucose-sensing S cells warrant further studies but given secretin’s potential effects on islet secretion and whole-body metabolism  it may be speculated that secretin contributes to the metabolic effects of RYGB.
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The study was supported by a grant from the European Research Council (grant no. 695069) to JJH and by an Excellence Emerging Investigator Grant—Endocrinology and Metabolism (NNF19OC0055001) to NJWA.
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The authors declare that they have no conflict of interest.
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Modvig, I.M., Andersen, D.B., Grunddal, K.V. et al. Secretin release after Roux-en-Y gastric bypass reveals a population of glucose-sensitive S cells in distal small intestine. Int J Obes (2020). https://doi.org/10.1038/s41366-020-0541-7
Frontiers in Endocrinology (2020)