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Sensory representation and detection mechanisms of gut osmolality change

Nature volume 602, pages 468–474 (2022)Cite this article

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Abstract

Ingested food and water stimulate sensory systems in the oropharyngeal and gastrointestinal areas before absorption1,2. These sensory signals modulate brain appetite circuits in a feed-forward manner3,4,5. Emerging evidence suggests that osmolality sensing in the gut rapidly inhibits thirst neurons upon water intake. Nevertheless, it remains unclear how peripheral sensory neurons detect visceral osmolality changes, and how they modulate thirst. Here we use optical and electrical recording combined with genetic approaches to visualize osmolality responses from sensory ganglion neurons. Gut hypotonic stimuli activate a dedicated vagal population distinct from mechanical-, hypertonic- or nutrient-sensitive neurons. We demonstrate that hypotonic responses are mediated by vagal afferents innervating the hepatic portal area (HPA), through which most water and nutrients are absorbed. Eliminating sensory inputs from this area selectively abolished hypotonic but not mechanical responses in vagal neurons. Recording from forebrain thirst neurons and behavioural analyses show that HPA-derived osmolality signals are required for feed-forward thirst satiation and drinking termination. Notably, HPA-innervating vagal afferents do not sense osmolality itself. Instead, these responses are mediated partly by vasoactive intestinal peptide secreted after water ingestion. Together, our results reveal visceral hypoosmolality as an important vagal sensory modality, and that intestinal osmolality change is translated into hormonal signals to regulate thirst circuit activity through the HPA pathway.

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Fig. 1: The vagus nerve responds to visceral osmolality changes.
Fig. 2: Visceral hypotonic stimuli activate a dedicated vagal population.
Fig. 3: Vagal sensory inputs from the HPA transmits gut-to-brain osmolality signals.
Fig. 4: HPA vagal afferents are required for pre-absorptive thirst satiation.
Fig. 5: Visceral osmolality sensing recruits VIP–VIPR2 signalling in the HPA.

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Data availability

Additional data that support the finding of this study are available from the corresponding author upon reasonable request. Source data are provided with this paper.

Code availability

The MATLAB code used to perform the imaging analysis is modified from the CalmAn code at https://github.com/flatironinstitute/CaImAn-MATLAB, and is available at https://github.com/Oka-Lab/Imaging-analysis.

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Acknowledgements

We thank the members of the Oka laboratory, S. D. Liberles and C. S. Zuker for helpful discussion and comments; B. Ho and A. Koranne for maintaining and genotyping mouse lines; T. Zhang, W. Han and I. E. Araujo for technical advice on surgical techniques; L. Luebbert for initial imaging analysis; T. Karigo for technical help; X. Chen and V. Gradinaru for advice on virus development and tissue clearing; and J. Parker for sharing the qPCR machine. This work was supported by Startup funds from the President and Provost of California Institute of Technology and the Biology and Biological Engineering Division of California Institute of Technology. Y.O. is also supported by New York Stem Cell Foundation, NIH (R01NS109997, R01NS123918), Alfred P. Sloan Foundation, and Heritage Medical Research Institute. T.I. is supported by the Japan Society for the Promotion of Science.

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Authors and Affiliations

  1. Division of Biology and Biological Engineering, California Institute of Technology, Pasadena, CA, USA

    Takako Ichiki, Tongtong Wang, Ann Kennedy, Allan-Hermann Pool, Haruka Ebisu, David J. Anderson & Yuki Oka

  2. Department of Physiology, Northwestern University, Chicago, IL, USA

    Ann Kennedy

  3. Howard Hughes Medical Institute, California Institute of Technology, Pasadena, CA, USA

    David J. Anderson

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  1. Takako Ichiki
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Contributions

T.I. and Y.O. conceived the research programme and designed experiments. T.I. performed the experiments and analysed the data, with help from T.W. A.K., and D.J.A. wrote the code for the imaging analysis and provided advice on the data analysis. A.-H.P. analysed the single-cell RNA sequencing data. H.E. performed and analysed chemogenetic experiments. T.I. and Y.O. wrote the paper. Y.O. supervised the entire work.

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Correspondence to Yuki Oka.

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Extended data figures and tables

Extended Data Fig. 1 Responses of vagal and thoracic DRG neurons in vivo.

a, Schematic of intestinal infusion during electrophysiological and optical recording. Isotonic (0.9%) saline was continuously perfused using a peristaltic pump. Stimulus was delivered using a solenoid valve at the same flow rate (800 μL/min). b, Integrated vagal nerve responses during stimulation (AUC) were quantified (n = 8 mice for saline and water, and n = 6 for other stimuli). c, Quantification of vagal water response at different flow rates (n = 5 mice). d, Number of WGA-positive cells in DRG (thoracic,T5-lumber,L3) retrogradely tracing from HPA, intestine or stomach. Representative images from 3 mice for each tracing site are shown in the right. Scale bar, 100 μm. e, Schematic diagram of DRG imaging. GRIN lens was placed on a thoracic DRG (T12 or 13). Calcium dynamics of individual neurons was visualized in Slc17a6-Cre;Ai96 mice. f, Response heatmaps to skin pinch, intestinal infusion of 4% acetic acid, water, 1M NaCl and Ensure from individual DRG neurons (ΔF/F(σ), left) and percentage of responding neurons (right, n = 3 mice for acetic acid, water and NaCl, 4 mice for other stimuli). *P < 0.05, **P < 0.01, ***P < 0.001 by Kruskal–Wallis test (with Dunn’s post test). Data are shown as mean ± s.e.m.

Source data

Extended Data Fig. 2 Osmolality and mechanical responses under vagal ganglia imaging.

a, Calcium dynamics during saline (blue), first (red) and second (purple) intestinal water infusion (Slc17a6-Cre;Ai96 mice). Representative images and a heatmap of individual vagal responses from 4 animal are shown. 64% of neurons activated by one trial were also activated by another trial. b, After subdiaphragmatic vagotomy (SVx), responses to intestinal water, 500 mM NaCl infusion, and distension were abolished (left). The number of total responding neurons was quantified (right, Slc17a6-Cre;Ai96, n = 4 mice). c, Heatmap of individual vagal activity during short- (20 s) and long- (2 min) term of intestinal water infusion (Slc17a6-Cre;Ai96, n = 3 mice). d, Osmotic-stimuli and mechanical distension activated largely non-overlapping neurons. Venn diagrams indicate the number of responding neurons (right, Slc17a6-Cre;Ai96, n = 3 mice). e, Representative vagal ganglia responses to water infusion and distension in the intestine (red) and stomach (green, n = 4 mice). f, Total number of vagal neurons responding to the indicated stimuli in the intestine (red, n = 4 mice) and stomach (green, n = 4 mice). Scale bar, 200 μm. Data are presented as mean ± s.e.m.

Source data

Extended Data Fig. 3 Transcriptomic analyses of the vagal ganglia.

a, Vagal ganglia contain 27 transcriptomic neuron types shown in a t-SNE embedding with color-coded cell identity. b, Violin plot of log-normalized expression of marker genes in vagal ganglia, including Tac1 and Nts with maximum counts per million (max CPM) above. t-SNE embedding for Nts (c) and Tac1 (d) shows log-normalized expression in vagal neuron types. These data are reanalyzed and plotted from a previous study (Prescott et al, 2020)24. e, Two-color RNA in situ hybridization for validating Cre expression in Tac1-Cre lines. 91.2% of Tac1-Cre expression matched endogenous gene expression. Scale bar, 20 µm.

Extended Data Fig. 4 Functional manipulation of TAC1+ and NTS+ vagal neurons.

a, TAC1+ neurons that responded to intestinal water or 500 mM NaCl are shown (Tac1-Cre;Ai162D, n = 4 mice). Same datasets from Fig. 3a are shown. b, Chemogenetic activation of TAC1+ vagal neurons significantly inhibited water intake after deprivation. AAV-hSyn-DIO-hM3D(Gq)-mCherry was injected into the vagal ganglia of Nts-Cre, Tac1-Cre, Slc17a6-Cre, or wild type (WT) mice. Water intake after 24-h water deprivation, 300 mM sucrose or chow consumption after 24-h food deprivation were measured after intraperitoneal PBS (vehicle) or clozapine N-oxide (CNO) administration (n = 7, 7, 5, and 4 mice (water and sucrose), and 5, 5, 4, and 4 mice (chow) were analyzed for Nts-Cre, Tac1-Cre, Slc17a6-Cre and WT animals, respectively.). c, Loss-of-function of TAC1+ vagal neurons. Cre-dependent taCasp3-encoding AAV was injected into the vagal ganglia of Tac1-Cre mice (left, n = 6) or WT mice (right, n = 6). After 2 weeks of recovery period, water intake was measured under water-deprived state. *P < 0.05, **P < 0.01, ***P < 0.001, ns, not significant by two-way ANOVA (Šídák multiple comparisons) or two-tailed paired t-test. Data are presented as mean ± s.e.m.

Source data

Extended Data Fig. 5 Anatomical projection patterns of TAC1+ and NTS+ vagal sensory neurons.

Whole-mount cleared tissue or sliced brain sections showing vagal terminals (AAV-Flex-tdTomato, red) and cell bodies (counterstained with DAPI or NeuN for Vagal ganglia and DRG, blue). Enteric neurons are labeled by Fluoro-Gold (green). A magnified image of stomach IGLE in a Slc17a6-Cre mouse is presented (bottom, left). The expression patterns in Slc17a6-Cre, Tac1-Cre and Nts-Cre mice are summarized (bottom, right, n = 3 animals for each mouse line). NTS, nucleus of the solitary tract; CG-SMG, celiac ganglion and superior mesenteric ganglion complex. Scale bar, 200 μm.

Extended Data Fig. 6 Anatomical projection patterns of vagal sensory neurons in the liver.

a, Whole-mount staining of innervation of SLC17A6+, TAC1+, NTS+ vagal populations in the hepatic area (n = 2 animals for each mouse line). Sensory terminals were visualized by AAV-Flex-tdTomato (red). Scale bar, 100 μm. b, Cell bodies of vagal sensory neurons were retrogradely labeled by WGA 555 (magenta) from the different liver parts (#2, #4, or all parts except #2 and #4, referred from a), and by WGA 647 (green) from HPV. The vagal innervation to the hilus and HPV mostly overlapped due to their proximity (84%), while the majority of the liver lacked innervation except a limited area in the left lobe (n = 3 mice). Scale bar, 200 μm. c, Hepatic branch denervation with intact liver branch abolished intestinal hypoosmotic responses (Slc17a6-Cre;Ai96, n = 3 mice).

Extended Data Fig. 7 Hepatic-branch-specific vagal denervation and its physiological significance.

a, Heatmap of individual vagal activity during long-term intestinal water infusion before and after HVx (Slc17a6-Cre;Ai96, n = 3 mice). b, Distinct vagal populations project to different organs. Whole-mount vagal sensory neurons retrogradely labeled by WGA 555 (magenta) and WGA 647 (green) from the same organ (HPA vs HPA) or different organs (HPA vs intestine, HPA vs stomach). Percentage of overlapping neurons is quantified (right, n = 3 mice). c, Validation of hepatic-branch-specific denervation. Vagal sensory neurons were retrogradely labeled by WGA 555 from intestine (magenta) and WGA 647 from HPA (green). After HPA denervation, the number of labeled neurons from HPA was drastically reduced (n = 3 mice). d, Reversible inhibition of water responses by a local anesthesia to HPA. Vagal responses to intestinal water infusion before and after application of 5% lidocaine (left, Slc17a6-Cre;Ai96, n = 4 mice). Responses recovered after the drug wore off. The number of the responded cells and mean ΔF/F(σ) values in neurons responded after 100 min are shown in the bottom. By contrast, the same anesthesia treatment did not affect intestinal distension responses (right, Slc17a6-Cre;Ai96, n = 3 mice). e, Duodenum and pancreas denervation using 85% phenol did not affect vagal activity during intestinal water infusion (left, Slc17a6-Cre;Ai96, n = 2 mice). Denervation was verified by injecting WGA 555 to pancreas (magenta) and WGA 647 to duodenum (green). After denervation, the number of retrogradely labeled neurons was drastically reduced (right, n = 3 mice). Scale bar, 200 μm. *P < 0.05, ***P < 0.001, ns, not significant by two-way or one-way ANOVA (Šídák multiple comparisons) or by Friedman test (Dunn’s multiple comparisons). Data are presented as mean ± s.e.m.

Source data

Extended Data Fig. 8 Functional and anatomical analyses of the hepatic vagal branch.

a, Vagal responses to intestinal water stimuli were abolished after HPA denervation using 85% phenol (Slc17a6-Cre;Ai96, n = 4 mice). b and c, Vagal response heatmaps to 500 mM NaCl or 300 mM glucose stimuli before and after HVx (b, n = 3 mice), or after injecting WGA 647 into the HPA (c, n = 3 mice). d, A minor population of intestine-innervating neurons (labeled by WGA 647) responded to intestinal acute water stimulus (Slc17a6-Cre;Ai96, n = 3 mice).

Extended Data Fig. 9 The effects of vagal manipulation on nutrient consumption and SFO neuron activity.

a, Water intake after 24-h deprivation during a 20-min session after HVx (n = 9 mice for sham and HVx, left) and HPA denervation using 85% phenol (n = 4 for sham, n = 7 mice for denervation, right). The same dataset from Fig. 4a for HVx experiments is presented. b, No change was observed for daily ad libitum water intake in sham (grey) and HPA denervation (red) animals (n = 5 mice). c, The vagus nerve does not contribute to water absorption. Plasma osmolality was measured in water-deprived mice after intestinal water infusion with or without SVx (n = 3 for sham, n = 3 for water-deprived SVx mice, n = 4 for water-infused SVx mice). d, Water (left) or food (normal chow, middle) intake after 24-h deprivation during a 20-min (water) or 1-h (food) session. Consistent with previous studies17,43, no difference was observed between sham (grey) and SVx (red) mice, or pre- and post-operation (n = 5 and 11 mice for pre- and post-op water intake measurement, n = 5 mice for food consumption measurement). Spontaneous daily water intake under sated states was not affected (right, n = 5 mice). e, Representative stomach and intestine samples after SVx (left) or HVx (right), with sham as a comparison. Stomach weight was significantly increased in the SVx animals but not in the HVx animals (n = 5 mice for sham, n = 6 mice for SVx and HVx). f, A diagram of vagal sensory circuits transmitting thirst satiation signals. g, Simultaneous photometry recording from SFONOS1 neurons and chemogenetic stimulation of vagal ganglion neurons. AAV-hSyn-hM3D(Gq)-mCherry was injected into the vagal ganglia while SFONOS1 neurons were infected with AAV-Flex-GCaMP6s in Nos1-Cre animals. h, Calcium dynamics of thirst neurons after PBS (vehicle) or CNO administration (n = 5 mice). Chemogenetic activation of vagal ganglia induced strong feed-forward inhibition in thirst neurons in sated- (left, red trace) and water-deprived condition (right, red trace). AUC was quantified between 300 to 600 s after CNO injection. *P < 0.05, **P < 0.01, ***P < 0.001, ns, not significant by two-way ANOVA (Šídák multiple comparisons), two-tailed Mann-Whitney U-test, or two-tailed paired t-test. Data are presented as mean ± s.e.m.

Source data

Extended Data Fig. 10 Contribution of hormones to HPA-innervating vagal responses and fluid balance.

a, Representative responses of vagal neurons to hormones infused into the HPV before (red) and after (green) HVx (Slc17a6-Cre;Ai96 mice, n = 3 mice). Scale bar, 200 μm. b, Vagal responses to intestinal water infusion and HPV GLP1 infusion (n = 3 mice). c, Vagal responses to a near-physiological concentration of VIP and intestinal hypoosmotic stimuli (n = 3 mice). d, VIP concentration in the HPV plasma was measured by ELISA from water-deprived (W.D.) animals and water-repleted animals with 10 min or 20 min water access (left, n = 9 mice for W.D., n = 8 mice for +water 10 min, and n = 6 mice for +water 20 min). No change in the systemic VIP level was observed in the retro-orbital sinus after water consumption (right, n = 9 mice for W.D., n = 8 mice for +water 10 min). e, After VIPR antagonist application, water and VIP dual-responding neurons were suppressed. f, qPCR analysis of VIPR1 and VIPR2 expression in the HPA, liver, duodenum, and colon. The same dataset from Fig. 5e is presented. *P < 0.05 by one-way ANOVA (Šídák multiple comparisons) or two-tailed unpaired t-test. Data are presented as mean ± s.e.m.

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Ichiki, T., Wang, T., Kennedy, A. et al. Sensory representation and detection mechanisms of gut osmolality change. Nature 602, 468–474 (2022). https://doi.org/10.1038/s41586-021-04359-5

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