High ambient temperature activates a neural circuit for gut glucose uptake in male mice

Abstract

Heat stress triggers defensive energy demands to prevent cellular damage. Using male mice, we found that exposure to high ambient temperature (HAT) induces an increase in intestinal glucose absorption mediated by sodium/glucose cotransporter 1 (SGLT1). Glucose supplementation alleviated HAT-induced physiological damage in mice. Mechanistically, we found that blocking intestinal vagal motor nerve transmission eliminated HAT’s stimulatory effects on intestinal glucose absorption. This suggests that cholinergic motor neurons originating from the dorsal motor nucleus of the vagus (DMV^ChAT) play a crucial role. Transneuronal tracing revealed an ascending pathway from glutamatergic neurons in the medial preoptic area (mPOA^Glu) to the vagus nerve, with corticotropin-releasing factor neurons in the paraventricular nucleus of the hypothalamus (PVN^CRF) acting as a key relay. Inhibiting key neurons in the mPOA^Glu → PVN^CRF → DMV^ChAT circuit mimicked the effects of vagus blockage on HAT-induced intestinal glucose absorption. Our findings establish the brain-vagal-gut pathway as an interoceptive circuit for adapting to HAT challenges.

Researchers report that exposure to high ambient temperature triggers a brain–vagus–gut neural circuit that boosts intestinal glucose uptake via sodium/glucose cotransporter 1 in male mice.

Introduction

Stressful life events often trigger carbohydrate craving^1,2. A sudden rise in ambient temperature leading to elevated body temperature significantly increases the risk of heatstroke, especially for the elderly and children³. Numerous studies have shown that heat exposure damages cellular structures (including the cytoskeleton, endoplasmic reticulum, and mitochondria) and disrupts normal physiological functions (such as DNA transcription and translation, protein synthesis, and folding). In severe cases, it can be life-threatening^4,5. Fortunately, the body can adapt to high-temperature environments through physical cooling and metabolic regulation⁶. During high ambient temperature (HAT) exposure, enhanced cardiopulmonary function promotes respiration and blood circulation, thereby improving heat dissipation^7,8. This process triggers increased synthesis of cellular heat shock proteins (HSPs), which maintain normal cellular protein synthesis⁹ and inhibit excessive heat production in brown adipose tissue¹⁰. These protective responses increase cellular ATP demand⁵. At the same time, because digestion and oxidation of protein and fat can generate substantial heat (diet-induced thermogenesis), limiting their utilization can be advantageous under heat stress^11,12. In this context, selectively increasing intestinal glucose absorption provides a rapid energy supply to support heat-defense programs while minimizing additional heat production.

Maintaining these physiological processes requires substantial energy, primarily in the form of ATP⁵. Glucose serves as a quick and efficient energy source, generating less metabolic heat during ATP conversion than fatty acid oxidation¹³. Evidence has accumulated to support glucose’s essential role in metabolic adaptation to HAT exposure. Heat exposure significantly increases skeletal muscle glucose absorption and metabolism, thereby promoting HSP72 synthesis¹⁴. An oral glucose tolerance test on healthy young males revealed elevated blood glucose levels following heat exposure^15,16. Moreover, glucose nutritional intervention enhances the body’s water retention and absorption under high-temperature conditions, highlighting glucose metabolism’s positive role in heat adaptation¹⁷.

Glucose from food is primarily absorbed in the small intestine, especially in its upper portion (duodenum and proximal jejunum)¹⁸. The intestinal glucose absorption process is heavily influenced by parasympathetic input, mainly originating from the cholinergic dorsal motor nucleus of the vagus (DMV^ChAT) in the brainstem. Activation of the vagal motor nerve significantly enhances intestinal motility and absorption efficiency¹⁹. Interestingly, the DMV^ChAT neurons receive signals from corticotropin-releasing factor neurons in the paraventricular nucleus of the hypothalamus (PVN^CRF)²⁰, which in turn receive input from medial preoptic area (mPOA) neuron²¹. The mPOA is well known as a temperature-control center, deeply involved in warm sensing and mediating downstream effector behaviors⁶. These findings suggest that HAT exposure may trigger changes in vagal motor nerve input to the gut, potentially influencing glucose absorption and contributing to HAT-induced enhanced glucose metabolism.

In the current study, we first confirmed that the HAT paradigm increases intestinal glucose absorption and circulating blood glucose levels. We then used a combination of pharmacological blockade and genetic deletion of intestinal sodium/glucose cotransporter 1 (SGLT1) to determine if intestinal glucose absorption is necessary for HAT’s stimulatory effects on blood glucose levels. Furthermore, we investigated whether intestinal vagal motor nerve transmission mediates HAT’s stimulatory effects on intestinal glucose absorption. Lastly, we employed transneuronal tracing and designer receptors exclusively activated by designer drugs (DREADD) to assess whether HAT exposure activates intestinal glucose absorption through the mPOA→PVN^CRF → DMV^ChAT circuit.

Results

HAT exposure increases intestinal glucose uptake

To study the impact of HAT exposure on blood glucose levels, male C57BL/6 J mice were fasted for 12 h and placed in a 37 °C environment, then allowed to eat freely for 2 h (Fig. 1a). Consistent with the known appetite-suppressing effects of HAT exposure²², we observed that 30 and 60 min of HAT exposure tended to decrease food intake, with a significant reduction at 120 min compared to mice kept at room temperature (RT, Fig. S1a). Interestingly, despite the reduced food intake after HAT exposure, blood glucose levels in mice rose significantly (Fig. 1b), suggesting HAT-induced enhancement of glucose metabolism. Additionally, we also measured changes in food intake and body weight under chronic HAT exposure, and the results showed that, consistent with acute HAT stimulation, food intake and weight gain were significantly reduced (Fig. S1b, c).

Fig. 1. HAT upregulates SGLT1 expression in the proximal small intestine to enhance intestinal glucose uptake.

a Strategy of HAT treatment (RT: room temperature, HAT: high ambient temperature). b Blood glucose levels in male C57BL/6 J mice during the first two hours of refeeding under RT or HAT conditions, following a 12-hour overnight fast (RT n = 8 mice; HAT n = 8 mice). 15 min: HAT vs. RT *P = 0.0109; 30 min: HAT vs. RT ***P = 0.000002. c Plasma ¹³C₆ content changes in male C57BL/6 J mice measured 30 min after D-[¹³C₆] glucose gavage under RT or HAT conditions, following a 12-hour overnight fast (RT n = 6 mice; HAT n = 6 mice). HAT vs. RT ***P = 0.000002. d HAT exposure protocol for continuously fasted mice. e Blood glucose levels in male C57BL/6 J mice during 90 min of extended fasting under RT or HAT conditions after a 12-hour overnight fast (RT n = 4 mice; HAT n = 4 mice). 15 min: HAT vs. RT *P = 0.03837; 30 min: HAT vs. RT **P = 0.009256; 45 min: HAT vs. RT *P = 0.018127. f Relative expression of SGLT1 mRNA in the proximal small intestine of male C57BL/6 J mice after 30 min of refeeding under RT or HAT conditions, following a 12-hour overnight fast (RT n = 6 mice; HAT n = 6 mice). HAT vs. RT ***P = 0.0003. g, h. Representative images of immunofluorescence staining for SGLT1 protein in the proximal small intestine of male C57BL/6 J mice after 30 min of refeeding under RT or HAT conditions, following a 12-hour overnight fast (Scale bars, 200 μm. RT n = 6 mice; HAT n = 6 mice). HAT vs. RT ***P = 0.0003. i Representative images of western blotting for SGLT1 protein in the proximal small intestine of male C57BL/6 J mice after 30 min of refeeding under RT or HAT conditions, following a 12-hour overnight fast. j Expression levels of SGLT1 protein in the proximal small intestine of male C57BL/6 J mice after 30 min of refeeding under RT or HAT conditions, following a 12-hour overnight fast (RT n = 6 mice; HAT n = 6 mice). HAT vs. RT ***P = 0.00007. k Changes in plasma ¹³C₆ content in male proximal small intestine SGLT1-specific knockdown mice 30 min after D-[¹³C₆] glucose gavage under RT or HAT conditions, following a 12-hour overnight fast (SGLT1^WT&flox/++RT n = 4 mice; SGLT1^WT&flox/++HAT n = 4 mice; SGLT1 ^flox/flox + RT n = 4 mice; SGLT1 ^flox/flox + HAT n = 4 mice). SGLT1^WT&flox/++HAT vs. SGLT1^WT&flox/++RT *P = 0.0372; SGLT1 ^flox/flox + RT vs. SGLT1^WT&flox/++RT *P = 0.0349; SGLT1 ^flox/flox + HAT vs. SGLT1^flox/flox + RT ^nsP = 0.8714. All data analyzed using an unpaired two-tailed student’s t-test. Data are shown as mean ± SEM; *P < 0.05, **P < 0.01, ***P < 0.001. Source data are provided as a Source Data file.

To explore the potential sources of HAT exposure-induced increases in blood glucose, we investigated its effects on endogenous glucose production pathways. We found that during refeeding, HAT had no significant effect on the mRNA expression of liver gluconeogenesis and glycogen metabolism-related enzymes (Fig. S1d). In contrast, HAT promoted the expression of GK, a key enzyme in glycolysis (Fig. S1e). It also did not alter serum insulin (in refed mice) (Fig. S1f) and corticosterone (in fasting or refed mice) (Fig. S1g) levels. Moreover, even when the HPA axis was blocked by an intraperitoneal injection of antalarmin, a CRF receptor 1 (CRFR1) antagonist²³, HAT still elevated blood glucose levels during refeeding (Fig. S1h). These results suggest that HAT increases blood glucose levels through an alternative pathway, independent of gluconeogenesis or the HPA axis.

Unlike mice that resumed feeding, male C57BL/6 J mice that were fasted overnight for 12 hours and continued fasting under RT or HAT conditions (Fig. 1d) showed significantly decreased blood glucose levels with HAT exposure (Fig. 1e), this may be related to the enhanced metabolism of the body in HAT. However, when these fasted mice received the same dose of 5% glucose via oral gavage every 30 minutes, the HAT exposure group maintained higher blood glucose levels (Fig. S1i). These results suggest that HAT enhanced intestinal glucose absorption in mice. To confirm this hypothesis, we used plasma ¹³C₆ levels after oral gavage of isotope ¹³C₆-labeled glucose (D-[¹³C₆] glucose), the most direct indicator of intestinal glucose absorption. Fig. S1j–k demonstrates that our chromatography-mass spectrometry method for detecting ¹³C₆ is feasible. More importantly, mice exposed to HAT exhibited significantly elevated plasma ¹³C₆ levels (Fig. 1c). These findings demonstrate that HAT exposure enhanced intestinal glucose absorption, likely contributing to HAT-induced elevation in blood glucose levels.

Enhanced intestinal glucose absorption mediates HAT’s stimulatory effects on blood glucose levels

To explore how HAT affects nutrient uptake in the proximal small intestine, we examined mRNA and protein expression levels of key proteins involved in glucose, amino acid, and fatty acid absorption. HAT significantly increased the expression of SGLT1 (Fig. 1f–j) but not GLUT2 (Fig. S2c), this may be SGLT1 is the main glucose transporter in the proximal small intestine^24,25, also had no effect on proteins related to amino acid and fatty acid absorption (Fig. S2a, b). This suggests HAT exposure specifically stimulates glucose absorption via SGLT1, but not amino acid or fatty acid absorption. Notably, HAT did not affect the mRNA expression of tight junction-related proteins in the proximal small intestinal mucosa (Fig. S3e), which was consistent with the HE staining of the proximal small intestinal histology and the electron microscopy of epithelial cells (Fig. S3c, d), indicating that intestinal mucosal barrier function and permeability remained intact. Additionally, we evaluated the effect of HAT on gut motility and found that HAT does not affect the transit time of the entire intestine (Fig. S3f). However, interestingly, HAT significantly increased the activity of the disaccharidase maltase in the intestine without affecting sucrase activity (Fig. S3g, h), which may be related to the fact that polysaccharides in food are primarily broken down into maltose²⁶. Furthermore, HAT did not affect the polysaccharidases, proteases, and lipases released by the pancreas (Fig. S3i–k), suggesting that HAT primarily promoted the entire process of glucose digestion and absorption in the proximal small intestine.

To test whether increased glucose absorption is necessary for HAT’s stimulatory effects on blood glucose, we blocked intestinal glucose absorption by orally administering the SGLT1 blocker phlorizin to male C57BL/6 J mice. After blocking SGLT1, HAT exposure no longer increased blood glucose levels (Fig. S2d). These pharmacological blocking experiments support the idea that HAT increases blood glucose levels by enhancing intestinal glucose uptake through increased SGLT1 expression. To avoid potential nonspecific effects of pharmacological SGLT1 inhibition, we adopted a second transgenic strategy to generate intestine-specific SGLT1 knockout mice. Specifically, we injected the AAV2/9-CMV-Cre-EGFP-pA virus into the proximal small intestine of male SGLT1^flox/flox mice, SGLT1^flox/+ mice, and wild-type littermates (Fig. S2e). Four weeks after viral infection, genomic allele-specific PCR revealed truncated DNA fragments in the proximal small intestine tissue of SGLT1^flox/flox mice (Fig. S2f). Additionally, these mice showed significantly reduced SGLT1 protein expression compared to SGLT1^flox/+ or WT mice (Fig. S2g, h). This intestinal SGLT1 knockdown led to decreased body weight without changing food intake, likely due to reduced intestinal glucose absorption (Fig. S2i, j). Consistent with the pharmacological blockade results, genetic knockdown of intestinal SGLT1 significantly reduced both HAT-induced increases in blood glucose levels and plasma ¹³C₆ content after D-[¹³C₆] glucose gavage (Fig. 1k and Fig. S2k). These data indicate that HAT can promote SGLT1 expression, thereby enhancing intestinal glucose absorption and increasing blood glucose levels.

Oral glucose intake alleviates HAT-induced damage

A sudden rise in ambient temperature causes cell injury and alters organelle structure, disrupting their function. To defend against this, cells use ATP to produce HSPs, which help mitigate HAT-induced damage⁵. Based on observed increases in intestinal glucose absorption due to HAT exposure, we hypothesize that circulating glucose plays an essential role in the metabolic adaptation and defensive responses to HAT. To verify this hypothesis, we examined the mRNA expression levels of ATP-generation-related enzymes and ATP content in the livers of HAT-treated mice following glucose gavage. Male C57BL/6 J mice that underwent a 12-hour overnight fast followed by 5% glucose gavage (Glu group) significantly increased the expression of ATP generation-related genes (DLAT and PDK2) and ATP content, likely due to improved energy availability after glucose administration under fasting conditions (Fig. 2a, b). But compared to the RT+Glu or HAT group, followed by 5% glucose gavage under HAT conditions, exhibited significantly upregulated expression of DLAT, PDK2 or CS (Fig. 2a) and ATP content (Fig. 2b) in livers. Moreover, this phenomenon is even more pronounced compared to the RT group.

Fig. 2. Glucose supplementation improves adaptation to HAT.

a Relative expression of four representative ATP-generating genes—DLAT (Dihydrolipoamide Acetyltransferase), PDK1 (Pyruvate Dehydrogenase Kinase 1), PDK2 (Pyruvate Dehydrogenase Kinase 2), and CS (Citrate Synthase)—in the livers of male C57BL/6 J mice after a 12-hour overnight fast, followed by saline (Sal) or 5% glucose (Glu) gavage and 30 min of treatment under RT and HAT conditions (RT+Sal n = 6 mice, RT+Glu n = 6 mice, HAT+Sal n = 6 mice, HAT+Glu n = 6 mice). b ATP content in the livers of male C57BL/6 J mice after a 12-hour fast, followed by saline or 5% glucose gavage, and 30 min of exposure to RT or HAT conditions (RT+Sal n = 6 mice, RT+Glu n = 5 mice, HAT+Sal n = 6 mice, HAT+Glu n = 6 mice). c Relative expression of heat shock protein mRNAs in the liver of male C57BL/6 J mice after a 12-hour overnight fast, followed by saline or glucose gavage and 30 min of treatment under RT and HAT conditions (RT+Sal n = 6 mice, RT+Glu n = 6 mice, HAT+Sal n = 6 mice, HAT+Glu n = 6 mice). d Representative transmission electron microscopy images of liver sections from male C57BL/6 J mice. Images were taken after a 12-hour overnight fast, gavage with saline or glucose (500 mg/kg), and 2 h of treatment under RT and HAT conditions. Blue arrows indicate lipid droplets, yellow arrows indicate mitochondria, and red arrows indicate endoplasmic reticulum (Scale bars: 10 or 2 μm). e Quantification on the average area of mitochondria in hepatocytes (RT+Sal n = 3 mice, RT+Glu n = 3 mice, HAT+Sal i = 3 mice, HAT+Glu n = 3 mice). f Statistics on the average number of lipid droplets in hepatocytes (RT+Sal n = 3 mice, RT+Glu n = 3 mice, HAT+Sal n = 3 mice, HAT+Glu n = 3 mice). g Heat shock responses in male C57BL/6 J mice fasted overnight for 12 h, followed by gavage with saline or glucose solution (500 mg/kg) and exposure to 42 °C. All data analyzed using an unpaired two-tailed student’s t-test. Data are shown as mean ± SEM; *P < 0.05, **P < 0.01, ***P < 0.001, compared with the RT+Sal group; ^#P < 0.05, ^##P < 0.01, ^###P < 0.001, compared with the HAT+Glu group. Source data are provided as a Source Data file.

Next, we investigated HAT’s effects on liver organelle structure and whether oral glucose intake could alleviate HAT-induced damage. As shown in previous studies²⁷, HAT caused hepatocyte mitochondria to shrink, damaged the endoplasmic reticulum, and increased liver lipid droplets (Fig. 2d–f). Lipid droplet balance is crucial for cellular antioxidant defense²⁸. We hypothesize that HAT increases lipid droplets as a protective mechanism to shield vulnerable unsaturated fatty acid triglycerides from further oxidation, thereby maintaining lipid balance. Supporting this hypothesis, oral glucose supplementation reduced HAT-induced liver organelle damage (Fig. 2d, e). This reduction was associated with further increases in hepatocyte lipid droplets (Fig. 2f) and HSP mRNA expression compared to mice exposed to HAT without glucose supplementation (Fig. 2c). In addition, oral glucose gavage under RT conditions did not affect HSP expression and liver organelle structure (Fig. 2c–f). It should be noted that, compared to liver mitochondrial damage, HAT also causes damage to the mitochondria of intestinal epithelial cells, but this did not reach statistical significance (0.05 < p < 0.1) (Fig. S3a, b). Moreover, glucose supplementation significantly reduced the rate of heat shock in mice at an extreme temperature of 42 °C (Fig. 2g). These findings indicate that glucose metabolism plays an essential role in metabolic adaptation to HAT exposure.

HAT activates vagal motor nerve to improve intestinal glucose absorption

The intestinal glucose absorption process is heavily influenced by parasympathetic input originating from DMV^ChAT neurons in the brainstem²⁹, which is potentially connected to the temperature control center mPOA^6,20,21. HAT exposure may trigger changes in vagal motor nerve input to the gut, thereby modulating glucose absorption. To directly test the mediating role of parasympathetic vagal input, we generated a mouse model with blocked neural transmission between the central nervous system (CNS) and intestine through subdiaphragmatic vagotomy (sdVx, Fig. 3a). Consistent with previous studies³⁰, the gastric volume of mice increased significantly after sdVx (Fig. S4a). To further validate the specificity of denervation, we injected the retrograde transsynaptic PRV-CAG-EGFP virus into the myenteric plexus of the proximal small intestine. Seven days post-infection, EGFP-positive neurons were observed in the sensory dorsal root ganglion (DRG), but not in the DMV (Fig. S4a), collectively confirming the success of sdVx.

Fig. 3. HAT increases intestinal glucose uptake via vagal motor nerve.

a,b Schematic diagram of subdiaphragmatic vagotomy surgery in male C57BL/6 J mice. Plasma ¹³C₆ content measured in sham or subdiaphragmatic vagotomy (sdVx) mice 30 min after D-[¹³C₆] glucose gavage under RT or HAT conditions, following a 12-hour overnight fast (RT+sham n = 5 mice, HAT+sham n = 5 mice, RT+sdVx n = 5 mice, HAT+sdVx n = 5 mice). HAT+sham vs. RT+sham **P = 0.0097; RT+sdVx vs. RT+sham ^nsP = 0.1394; HAT+sdVx vs. RT+sdVx ^nsP = 0.9021. c,d Schematic diagram of intraperitoneal atropine injection in male C57BL/6 J mice. Plasma ¹³C₆ content in mice 30 min after D-[¹³C₆] glucose gavage, following an intraperitoneal injection of saline or atropine (Atr) and a 12-hour overnight fast, under RT or HAT conditions (RT+Sal n = 5 mice, HAT+Sal n = 5 mice, RT+Atr n = 6 mice, HAT+ Atr n = 6 mice). HAT+Sal vs. RT+Sal **P = 0.0016; RT+Atr vs. RT+Sal ^nsP = 0.6795; HAT+ Atr vs. RT+Atr ^nsP = 0.6717. e,f Representative images of AAV-PHP.eB-DIO-hM4D(Gi)-mCherry injected in the proximal small intestine of male ChAT-cre mice (Scale bars, 100 μm). In male ChAT-cre mice fasted overnight for 12 h, plasma ¹³C₆ levels were measured 30 min after D-[¹³C₆] glucose gavage under RT and HAT conditions. These measurements followed an intraperitoneal injection of saline or Clozapine N-oxide (CNO, 1 mg/kg) administered 30 min before the gavage (Scale bars: 100 μm, RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 5 mice, HAT + CNO n = 5 mice). HAT+Sal vs. RT+Sal *P = 0.0383; RT + CNO vs. RT+Sal ^nsP = 0.9981; HAT + CNO vs. RT + CNO ^nsP = 0.0861. All data analyzed using an unpaired two-tailed student’s t-test. Data are shown as mean ± SEM; *P < 0.05, **P <0.01. Source data are provided as a Source Data file.

Consistent with the observations in male C57BL/6 J naïve mice, the sham control male C57BL/6 J mice exposed to HAT during refeeding showed increased blood glucose levels and intestinal glucose absorption compared to sham control male C57BL/6 J mice kept at RT (Figs. S4c and 3b). Conversely, in sdVx male C57BL/6 J mice, HAT not only failed to increase blood glucose levels but also reduced them further (Fig. S4c). Interestingly, this phenomenon occurred without significant changes in food intake (Fig. S4b). This is potentially due to the increased energy demand during HAT exposure, with unchanged intestinal glucose absorption (Fig. 3b). These findings support the mediating role of vagal motor input for the stimulatory effects of HAT on blood glucose levels. It should be noted that SDVx denervates both the afferent and efferent vagal nerves from the CNS to the gut³¹. The blockage of HAT-induced blood glucose elevation by sdVx might be due to the interaction between the intestine and CNS mediated by the vagal sensory nerve.

To rule out this possibility, we pharmacologically inhibited vagal motor nerve transmission by intraperitoneally injecting atropine (Fig. 3c), an acetylcholine M receptor blocker³². Notably, atropine treatment inhibited food intake under RT conditions (Fig. S4d), reducing it to a minimal amount. We considered that the observed effects might be due to fasting rather than blockage of intestinal glucose absorption. To address this concern, we gavaged mice with an equivalent dose of glucose while simultaneously injecting atropine intraperitoneally. As shown in Fig. S4e, HAT not only failed to increase blood glucose but also caused a significant decrease. Subsequently, we performed D-[¹³C₆] glucose gavage in atropine-treated mice. The results confirmed that HAT did not promote intestinal glucose absorption (Fig. 3d). These findings suggest that vagal motor input plays an essential role in the HAT-induced increases in intestinal glucose absorption and blood glucose levels.

To avoid potential nonspecific effects of systemic pharmacologic blockade of acetylcholine M receptors, we developed a chemogenetic mouse model to selectively inhibit intestinal vagal transmission. Vagal motor preganglionic DMV^ChAT neurons are known to activate cholinergic postganglionic myenteric neurons, which release acetylcholine to enhance intestinal function²⁹. We targeted these cholinergic postganglionic myenteric neurons in the intestinal tract to block vagal transmission. Specifically, we injected cre-dependent AAV-PHP.eB-hSyn-DIO-hM4D(Gi)-mCherry virus into the proximal small intestine of male cholinergic-specific Cre mice (ChAT-Cre). This virus contains a PHP.eB capsid that enables it to cross the intestinal mucosal barrier and a double-floxed inverted open-reading frame (DIO) that induces long-term hM4D(Gi)-mCherry expression exclusively in ChAT-Cre positive myenteric neurons. Three weeks after the injection, the intestinal myenteric plexus showed abundant expression of hM4D(Gi)-mCherry (Fig. 3e), confirming the sufficient infection.

Saline or CNO was injected intraperitoneally to create a control group and an intestinal vagal motor postganglionic neuron inhibition group, respectively. Notably, saline-injected control mice exposed to HAT showed similar increases in blood glucose and intestinal glucose absorption as observed in naïve male C57BL/6 J mice. However, inhibiting intestinal vagal motor postganglionic neurons through CNO injection blocked these stimulatory effects induced by HAT exposure (Fig. 3f and Fig. S4f), supporting a mediating role of vagal motor input. By combining evidence from surgical denervation, pharmacological blockade, and chemogenetic inhibition, we conclude that vagal motor input mediates HAT’s stimulatory effects on intestinal glucose absorption and subsequent blood glucose increases. This mechanism is likely crucial for metabolic adaptation and defensive responses to HAT exposure.

mPOA → PVN → DMV^ChAT→gut neural circuit

Our findings demonstrate that the vagal motor nerve functions as an efferent component, facilitating HAT’s stimulatory effects on intestinal glucose absorption. The next logical questions are: What is the source of the upstream input, and what type of neural population detects ambient temperature changes and transmits signals to the vagal motor nerve? To identify the upstream neural populations sending signals to the intestine, we injected the transsynaptic retrograde tracing PRV-CAG-EGFP virus into multiple points of the proximal small intestine (Fig. 4a). In line with the well-known classic parasympathetic vagal efferent circuit, we found EGFP-labeled neurons in the DMV (Fig. S5a), the main location of preganglionic parasympathetic vagus neuron cell bodies. We also observed EGFP signals in the PVN, mPOA, rostral ventrolateral medulla (RVL), lateral paragigantocellular nucleus (LPGi), central amygdala (CeA), medial globus pallidus (MGP), and parasubthalamic nucleus (PSTh) (Fig. S5a). These findings suggest potential anatomical connections between these central nuclei and the proximal small intestine.

Fig. 4. The mPOA → PVN → DMV^ChAT circuit is connected to the gut.

a Schematic diagram of the tracking strategy for transsynaptic retrograde tracing using pseudorabies virus (PRV) injection in the proximal small intestine. b Representative pictures of EGFP expressing neurons in DMV, PVN and mPOA nucleus on days 3, 5, and 7 after PRV-CAG-EGFP virus injection in the proximal small intestine (Scale bars, 200 μm). c Number of EGFP expressing neurons in DMV, PVN and mPOA nucleus after days 3, 5, and 7 of PRV virus infection (3d n = 6 mice, 5d n = 4 mice, 7d n = 3 mice). Data analyzed using an unpaired two-tailed student’s t-test. DMV: 5d vs. 3d ***P = 0.00001; 7d vs. 5 d *P = 0.04; PVN: 5d vs. 3d ***P = 0.00001; 7d vs.5d **P = 0.0011; mPOA: 5d vs. 3d **P = 0.006082; 7d vs.5d **P = 0.0013. d Ratio of the number of EGFP expressing neurons in PVN and mPOA nucleus to the number of GFP neurons in DMV after days 3, 5, and 7 of PRV virus infection (3d n = 6 mice, 5 d n = 4 mice, 7 d n = 3 mice). Data analyzed using an unpaired two-tailed student’s t-test. PVN: 7 d vs.5 d ***P = 0.000138; mPOA: 5 d vs. 3 d ***P = 0.0006; 7 d vs.5 d ***P = 0.000482. e Schematic diagram of the monosynaptic retrograde rabies virus (RV) tracing strategy for PVN→DMV^ChAT circuit. f Representative images of viral expression in the injection site DMV and the upstream PVN nucleus with direct projections (Scale bar, DMV: 100 μm, PVH and mPOA :200 μm). g Schematic diagram of the monosynaptic retrograde RV tracing strategy for mPOA→PVN→DMV^ChAT circuit. h Representative images of viral expression in the injection site DMV and the upstream PVN and mPOA nucleus with direct projections (Scale bar, 200 μm). i Schematic diagram of the in vivo fiber photometry strategy for the mPOA→PVN → DMV circuit (Scale bar, 100 μm). j Activity changes in the PVN nucleus of male C57BL/6 J mice in response to saline or 5% glucose gavage under RT conditions (Sal n = 4 mice, Glu n = 4 mice). Paired Student’s t-test. Glu vs. Sal ^nsP = 0.5108. k Activity changes in the PVN nucleus of male C57BL/6 J mice under RT and HAT conditions following a 12-hour overnight fast (RT n = 4 mice, HAT n = 4 mice). Paired Student’s t-test. HAT vs. RT *P = 0.0496. l Activity changes in the PVN nucleus of male C57BL/6 J mice unde RT and HAT conditions following glucose gavage (RT n = 4 mice, HAT n = 4 mice). Paired Student’s t-test. HAT vs. RT *P = 0.0176. All data are shown as mean ± SEM; *P < 0.05, **P < 0.01.***P < 0.001 Source data are provided as a Source Data file.

To establish a functional link between HAT exposure and these central nuclei, we performed whole-brain c-Fos immunofluorescence staining after HAT exposure. We found significantly upregulated c-Fos expression in the DMV, PVN, and mPOA, but not in the other PRV-EGFP positive central nuclei (Fig. S5a, b), in HAT-exposed male C57BL/6 J mice compared to those kept at RT. This suggests these three nuclei may play a major role in HAT-induced blood glucose elevation.

Additionally, the number of EGFP-positive neurons in these three nuclei gradually increased with viral infection duration (Fig. 4b, c). Compared with day 3 post-injection, the mPOA increased only on day 7, while the PVN increased on both days 5 and 7, with day 7 higher than day 5. The DMV increased on both days 5 and 7 relative to day 3, with similar neuron counts on these two days. These findings suggest a potential connectivity relationship: the mPOA is likely upstream of the PVN, which is potentially upstream of the DMV. Supporting this, the proportion of infected neurons in the mPOA and PVN relative to DMV-infected neurons also increased over time (Fig. 4d). This time-dependent anatomical progression supports the potential connectivity between the mPOA, PVN, DMV, and intestine.

To further establish this connection, we performed an opposite transsynaptic anterograde tracing by injecting the HSV-tdTomato virus into the mPOA. In line with our retrograde labeling results, we observed tdTomato-labeled neurons in the mPOA, PVN, and DMV (Fig. S5c). We also found tdTomato-positive cells in the intestinal villi and myenteric plexus of the proximal small intestine. These infected intestinal cells likely include vagal motor postganglionic neurons, enteric neurons, and some endocrine cells (Fig. S5c), given their synapse-like connections with intestinal nerves^33,34. This evidence strongly supports the existence of a neural connection between the central nuclei (DMV, PVN, and mPOA) and the proximal small intestine. We also utilized Fiber Photometry to detect the responses of the mPOA→PVH → DMV circuit to temperature and intestinal glucose (Fig. 4i). We found that at room temperature, oral glucose administration did not affect its activity (Fig. 4j), but under fasting or oral glucose conditions, when ambient temperature increased from 25 °C to 37 °C, the circuit was significantly activated (Fig. 4k, l). This indicates that the mPOA→PVH → DMV circuit does not depend on feedback from intestinal glucose signals but rather is regulated by ambient temperature.

To investigate whether PVN neurons provide monosynaptic inputs to DMV^ChAT neurons, we employed a rabies-mediated retrograde tracing system. This system comprised an EnvA-pseudotyped, glycoprotein-deleted Rabies-EGFP (RV-EnvA-SAD-ΔG-Cre-EGFP) and Cre-dependent helper AAV viruses: AAV2/9-CAG-DIO-TVA-mCherry and AAV2/9-CAG-FLEX-oG. The EnvA-pseudotyped rabies virus only infects neurons with the expression of the cellular receptor for subgroup A avian leukosis viruses (TVA)³⁵. Moreover, this rabies virus lacks the envelope glycoprotein (oG) necessary for trans-synaptic retrograde spreading through axon terminals³⁵. We implemented this system by first delivering AAV2/9-CAG-DIO-TVA-mCherry and AAV2/9-CAG-FLEX-oG into the DMV of ChAT-Cre mice, inducing TVA and oG expression exclusively in DMV^ChAT neurons. Three weeks later, we introduced RV-EnvA-SAD-ΔG-Cre-EGFP into the same DMV brain region (Fig. 4e). The selective expression of TVA-mCherry in DMV^ChAT neurons allows for rabies infection of these neurons, while oG expression enables subsequent retrograde transsynaptic Rabies-EGFP labeling of upstream neurons. We observed successful TVA-mCherry expression in the DMV^ChAT neurons, along with Rabies-EGFP expression in the PVN, but not in the mPOA (Fig. 4f). These findings confirm that PVN neurons provide direct, monosynaptic inputs to the DMV^ChAT neural population.

Next, to further investigate this circuit, we expanded our viral injection protocol based on the previous experiments. We first injected AAV2/9-CAG-DIO-TVA-mCherry and AAV2/9-CAG-FLEX-oG into the DMV of ChAT-Cre mice, inducing expression of TVA and oG exclusively in DMV^ChAT neurons. Three weeks later, we injected RV-EnvA-SAD-ΔG-Cre-EGFP into the same DMV region and supplemented it with Cre-dependent helper virus AAV2/9-CAG-FLEX-oG in the PVN (Fig. 4g). The EnvA-pseudotyped rabies virus injected into the DMV carried Cre, allowing PVN neurons projecting to DMV^ChAT to express oG protein. This enabled subsequent retrograde transsynaptic Rabies-EGFP labeling of upstream neurons. Through this carefully designed series of viral tracing protocols, we detected Rabies-EGFP signals in the mPOA nucleus (Fig. 4h), confirming the existence of monosynaptic connections in the mPOA→PVN→DMV^ChAT circuit. Combined with the PRV retrograde tracing and HSV anterograde tracing results, these findings suggest that the mPOA→PVN→DMV^ChAT circuit might regulate intestinal function via projections from DMV^ChAT neurons to the proximal small intestine.

Building on previous studies demonstrating that DMV^ChAT neurons receive signals from PVN^CRF20, we further investigated whether PVN^CRF serves as a crucial relay between mPOA and DMV^ChAT neurons. Specifically, we injected a Cre-dependent anterograde transsynaptic AAV2/1-EF1α-DIO-FLP virus and locally expressed AAV2/9-EF1α-EGFP virus into the mPOA of C57BL/6 J mice. Concurrently, we injected a CRF promoter-driving AAV2/9-CRF-Cre-EGFP virus and a Flp-dependent AAV2/9-hsyn-fDIO-eNpHR3.0-mCherry virus into the PVN (Fig. S5d). The exclusive expression of EGFP in the mPOA confirmed the accurate injection of both AAV2/1-EF1α-DIO-FLP and AAV2/9-EF1α-EGFP (Fig. S5e).

As previously reported³⁶, anterograde transneuronal transport of AAV2/1-EF1α-DIO-FLP enabled Cre-dependent expression of Flp in the PVN. Specifically, the first PVN-targeted virus, AAV2/9-CRF-Cre-EGFP (green), selectively induced Flp recombinase expression in PVN^CRF neurons receiving input from mPOA. The second PVN-targeted virus, AAV2/9-hsyn-fDIO-eNpHR3.0-mCherry, enabled Flp-dependent expression of mCherry specifically in these mPOA-recipient PVN^CRF neurons (red), filling their soma, dendrites, and axons. The mCherry⁺ neurons co-localized with EGFP expression in these PVN^CRF neurons (Fig. S5f), confirming the infection’s specificity. We observed mCherry-labeled fibers in the DMV (Fig. S5g), indicating that mPOA-recipient PVN^CRF neurons project to the DMV. This anatomical evidence further supports the existence of an mPOA→PVN^CRF → DMV^ChAT circuit.

HAT enhances intestinal glucose uptake through DMV^ChAT, PVN^CRF, and mPOA^Glu neurons

Previous studies have shown that warm-responsive neurons in the mPOA are primarily glutamatergic³⁷, suggesting a potential role for mPOA glutamatergic (mPOA^Glu) neurons in HAT-induced responses. To test the relevance of the mPOA^Glu → PVN^CRF → DMV^ChAT neural circuit in the physiological response to HAT exposure, we injected three different promoter viruses into male Rosa26-LSL-tdTomato mice: AAV2/9-ChAT-Cre-P2A-WPRE-hGH into the DMV, AAV2/9-CRF-Cre-P2A-WPRE-hGH into the PVN, and AAV2/9-Vglut2-Cre-P2A-WPRE-hGH into the mPOA. To confirm that this strategy successfully and specifically induced tdTomato expression in DMV^ChAT, PVN^CRF, and mPOA^Glu neurons, respectively. We injected AAV2/9-ChAT/CRF/Vglut2-Cre, AAV-DIO-GFP, and AAV-fDIO-mCherry viruses into the DMV/PVN/mPOA nuclei of ChAT/CRF/Vglut2-Flp mice. Fluorescence expression results showed that neurons co-expressing both Cre and Flp (yellow) accounted for more than 80% of the total neurons expressing Cre (green) or Flp (red) (Fig. S6a–c), indicating that these promoters have high specificity. We then investigated whether HAT exposure activates these neurons (Fig. 5a). Compared to overnight-fasted mice refed at RT, those refed for 30 minutes during HAT exposure showed a significantly higher co-localization rate of c-Fos with tdTomato-positive neurons in DMV^ChAT, PVN^CRF, and mPOA^Glu neurons (Fig. 5b and Fig. S6d–f). This suggests that HAT exposure enhances the excitability of these neuronal populations, potentially playing a crucial role in HAT-induced increases in intestinal glucose absorption.

Fig. 5. HAT-induced intestinal glucose uptake is mediated by DMV^ChAT, PVN^CRF and mPOA^Glu neurons.

a Schematic diagram depicting the injection of three distinct viruses into male Rosa26-LSL-tdTomato mice: AAV2/9-ChAT-Cre-P2A-WPRE-hGH into the DMV, AAV2/9-CRF-Cre-P2A-WPRE-hGH into the PVN, and AAV2/9-Vglut2-Cre-P2A-WPRE-hGH into the mPOA. b Statistical graphs for figure a. ChAT: n = 6 mice, HAT vs. RT ***P <0.00001; CRF: n = 6 mice, ***P < 0.00001; Vglut2: n = 6 mice, ***P < 0.00001. c Plasma ¹³C₆ content was measured in male ChAT-Cre mice expressing viral activation of the PVN→DMV^ChAT circuit. Measurements were taken 30 min after D-[¹³C₆] glucose gavage under RT and HAT conditions, following a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg) (RT+Sal n = 6 mice, RT + CNO n = 6 mice). RT+Sal vs. RT + CNO ***P = 0.0005. d After a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg), male ChAT-cre mice expressing hM4D(Gi)-mCherry in DMV^ChAT neurons received D-[¹³C₆] glucose via gavage. Plasma ¹³C₆ content was then measured after 30 min of treatment under RT or HAT conditions (RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal *P = 0.011, HAT + CNO vs. RT + CNO ^nsP = 0.9843. e After a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg), male CRF-cre mice expressing hM4D(Gi)-mCherry in PVN^CRF neurons received D-[¹³C₆] glucose via gavage. Plasma ¹³C₆ content was then measured after 30 min of treatment under RT or HAT conditions (RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal *P = 0.0122, HAT + CNO vs. RT + CNO ^nsP = 0.9299. f After a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg), male Vglut2-cre mice expressing hM4D(Gi)-mCherry in mPOA^Glu neurons received D-[¹³C₆] glucose via gavage. Plasma ¹³C₆ content was then measured after 30 min of treatment under RT or HAT conditions (RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 5 mice). HAT+Sal vs. RT+Sal **P = 0.0044, HAT + CNO vs. RT + CNO ^nsP = 0.1147. All data analyzed using an unpaired two-tailed student’s t-test. Data are shown as mean ± SEM; *P < 0.05, **P < 0.01,***P < 0.001. Source data are provided as a Source Data file.

In addition, recent studies have identified multiple subtypes of temperature-sensitive neurons in the POA that participate in regulating heat acclimatization (e.g., LepR and BDNF neurons)^38,39. To preliminarily investigate whether these neurons are involved in the heat-induced enhancement of intestinal glucose absorption, we similarly injected PRV-GFP into the proximal small intestine and examined POA neuronal subtypes that have synaptic connections with the intestine by fluorescent colocalization. The results showed that retrogradely labeled PRV-GFP from the intestine was mainly expressed in the middle and caudal parts of the POA, whereas BDNF neurons (immunofluorescence) were primarily located in the rostral region (Fig. S6g). Likewise, GFP-labeled neurons and tdTomato-labeled LepR neurons (LepR-Cre::LSL-tdTomato mice) showed no colocalization in the POA (Fig. S6g). Therefore, in subsequent work, we will focus on the role of mPOA^Glu neurons in heat-promoted intestinal glucose absorption.

DMV^ChAT, as the origin site of vagal motor preganglionic neurons, primarily participates in regulating gastrointestinal motility. However, recent studies have found that it can regulate intestinal fat absorption⁴⁰. Therefore, we first injected Cre-dependent AAV2/9-DIO-hM3D-mcherry virus bilaterally into the DMV of ChAT-Cre mice (Fig. S7a), to investigate whether activation of DMV^ChAT enhances glucose absorption. Intraperitoneal injection of CNO significantly increased c-Fos expression in DMV^ChAT neurons (Fig. S7a, b), and also significantly increased SGLT1 expression in epithelial cells of the proximal small intestine (Fig. S7c, d). However, surprisingly, it significantly decreased intestinal transit time and glucose absorption (Fig. S7e, f), which may be similar to heat exposure in young rats, enhancing intestinal motility and thus reducing glucose absorption efficiency⁴¹. Instead, we injected an anterograde trans-synaptic monosynaptic AAV-DIO-Flp virus into the PVN of ChAT-Cre mice and injected AAV-fDIO-hM3D-GFP into the DMV to selectively activate the PVN→DMV^ChAT circuit (Fig. S7g, h). Intraperitoneal injection of CNO not only similarly activated DMV^ChAT neurons and increased SGLT1 expression in the proximal small intestine (Fig. S7i, j), but also significantly enhanced glucose absorption (Fig. 5c), further confirming that distinct neuronal subtypes in the DMV differentially regulate intestinal function. We then investigated whether chemogenetic inhibition of these neurons counteracts the effects of HAT. Specifically, we bilaterally injected Cre-dependent inhibitory AAV2/9-hSyn-DIO-hM4Di-mCherry into the DMV of ChAT-Cre, PVN of CRF-Cre, or mPOA of Vglut2-Cre mice, respectively (Fig. 5d–f). The hM4Di-mCherry expression was exclusively induced in DMV^ChAT, PVN^CRF, and mPOA^Glu neurons (Fig. S8a, d, g). Compared to saline-injected controls, CNO injections effectively blocked the HAT-induced increases in neural excitability (Fig. S8b, e, h). Notably, chemogenetic inhibition of any of the three neural populations blocked the HAT-induced increases in blood glucose and intestinal glucose absorption (Fig. S8c, f, i and Fig. 5d–f). The inhibition of mPOA^Glu neurons even reversed HAT’s stimulatory effects, leading to inhibitory effects on blood glucose (Fig. S8i). These findings demonstrate that DMV^ChAT, PVN^CRF, and mPOA^Glu activation are necessary for HAT’s stimulatory effects on intestinal glucose absorption and blood glucose levels.

HAT enhances intestinal glucose uptake through the mPOA^Glu → PVN^CRF → DMV^ChAT neural circuit

To further establish a functional link between the mPOA^Glu → PVN^CRF → DMV^ChAT circuit and HAT-induced responses, we used a viral intersection approach to selectively block the neural circuit’s activation. We first generated a mouse model with the mPOA^Glu → PVN circuit selectively inhibited. This was achieved by bilaterally injecting AAV2/9-Vglut2-Cre and Flp-dependent AAV2/9-hsyn-fDIO-hM4D(Gi)-mCherry into the upstream mPOA, and retrograde non-transsynaptic AAV2/retro-EF1α-DIO-FLP and locally expressing AAV2/9-EF1α-EGFP into the downstream PVN of male C57BL6/J mice (Fig. 6a). The exclusive expression of EGFP in the PVN confirmed the accurate injection of both AAV2/retro-EF1α-DIO-FLP and AAV2/9-EF1α-EGFP (Fig. S9a). The first mPOA-targeted AAV2/9-Vglut2-Cre virus induced Cre recombinase expression in mPOA^Glu neurons, filling their soma, dendrites, and axons, including those in the PVN. Retrograde axonal transport of the PVN-targeted AAV2/retro-EF1α-DIO-FLP further enabled Cre-dependent expression of Flp in the mPOA^Glu neurons projecting to PVN. The second mPOA-targeted virus, AAV2/9-hsyn-fDIO-hM4D(Gi)-mCherry, then induced Flp-dependent expression of hM4D(Gi)-mCherry specifically in these PVN-projecting mPOA^Glu neurons (red, Fig. S9b). In these mPOA^Glu → PVN circuit inhibition mice, compared to saline-injected controls, CNO injections effectively blocked the HAT-induced increases in neural excitability (Fig. S9c). Notably, chemogenetic inhibition of the mPOA^Glu → PVN circuit effectively blocked the HAT-induced increases in blood glucose and intestinal glucose absorption (Fig. 6b and Fig. S9d).

Fig. 6. The mPOA^Glu → PVN^CRF→DMV^ChATcircuit is involved in HAT-induced intestinal glucose uptake.

a Viral injection strategy for specifically inhibiting mPOA^Glu → PVN neurons (Viral injection n = 24 mice). b Plasma ¹³C₆ content was measured in male C57BL6/J mice expressing viral inhibition of the mPOA^Glu → PVN circuit. Measurements were taken 30 min after D-[¹³C₆] glucose gavage under RT and HAT conditions, following a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg, RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal ***P = 0.0005, HAT + CNO vs. RT + CNO ^nsP = 0.8806. c Viral injection strategy for specifically inhibiting mPOA→PVN^CRF neurons (Viral injection n = 24 mice). d Plasma ¹³C₆ content was measured in male C57BL6/J mice expressing viral inhibition of the mPOA→PVN^CRF circuit. Measurements were taken 30 min after D-[¹³C₆] glucose gavage under RT and HAT conditions, following a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg, RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 5 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal **P = 0.0046, HAT + CNO vs. RT + CNO ^nsP = 0.4217. e Viral injection strategy for specifically inhibiting PVN→DMV^ChAT neurons (Viral injection n = 24 mice). f Plasma ¹³C₆ content was measured in male C57BL6/J mice expressing viral inhibition of the PVN→DMV^ChAT circuit. Measurements were taken 30 min after D-[¹³C₆] glucose gavage under RT and HAT conditions, following a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg, RT+Sal n = 5 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal **P = 0.0071, HAT + CNO vs. RT + CNO ^nsP = 0.1655. g Viral injection strategy for specifically inhibiting mPOA→PVN → DMV neurons (Viral injection n = 24 mice). h Plasma ¹³C₆ content was measured in male C57BL6/J mice expressing viral inhibition of the mPOA→PVN → DMV circuit. Measurements were taken 30 min after D-[¹³C₆] glucose gavage under RT and HAT conditions, following a 12-hour overnight fast and intraperitoneal injection of saline or CNO (1 mg/kg, RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal **P = 0.0045, HAT + CNO vs. RT + CNO ^nsP = 0.6614. i Representative immunofluorescence staining images of SGLT1 protein in the proximal small intestine of male C57BL6/J mice with virus-mediated inhibition of the mPOA→PVN → DMV circuit. Images were captured following a 12-hour overnight fast, intraperitoneal injection of saline or CNO (1 mg/kg), and 30 min of refeeding under RT and HAT conditions (Scale bars, 200 μm. RT+Sal n = 6 mice, HAT+Sal n = 6 mice, RT + CNO n = 6 mice, HAT + CNO n = 6 mice). HAT+Sal vs. RT+Sal **P = 0.0043, HAT + CNO vs. RT + CNO ^nsP = 0.5604. All data analyzed using an unpaired two-tailed student’s t-test. Data are shown as mean ± SEM; **P <0.01,***P <0.001. Source data are provided as a Source Data file.

We then targeted mPOA-recipient PVN^CRF neurons, or the mPOA→PVN^CRF circuit. We used a strategy similar to that described in Fig. S4e–h, but substituted the PVN-targeted viruses. Instead of AAV2/9-CRF-Cre-EGFP and AAV2/9-hsyn-fDIO-eNpHR3.0-mCherry, we used AAV2/9-CRF-Cre and AAV2/9-hsyn-fDIO-hM4D(Gi)-mCherry (Fig. 6c). This approach induced EGFP expression in the mPOA, confirming accurate injection (Fig. S10a). The expression of hM4Di-mCherry in the downstream PVN validated the successful expression of inhibitory DREADD in PVN^CRF neurons receiving projections from upstream mPOA neurons (Fig. S10b). Notably, we observed responses similar to those seen in PVN-projecting mPOA^Glu inhibitory mice. These included blunted HFA-induced increases in c-Fos expression (Fig. S10b, c), blood glucose levels (Fig. S10d), and intestinal glucose absorption (Fig. 6d). We also observed similar results when targeting PVN-recipient DMV^ChAT neurons or the PVN→DMV^ChAT circuit using a comparable strategy (Fig. 6e, f and Fig. S11a–d). The findings from these circuit-specific inhibitions provide strong evidence for the critical role of the mPOA^Glu → PVN^CRF → DMV^ChAT pathway. However, it is important to note that these manipulations primarily focus on single synaptic connections, which may not be sufficient to draw conclusions about the entire multi-synaptic pathway.

We next employed both anterograde and retrograde strategies to target relay PVN neurons that receive innervation from upstream mPOA and project to downstream DMV. Specifically, we injected anterograde transsynaptic AAV2/1-EF1a-DIO-FLP and locally expressing AAV2/9-EF1α-EGFP into the mPOA, Flp-dependent AAV2/9-hsyn-fDIO-hM4D(Gi)-mCherry into the PVN, and retrograde non-transsynaptic AAV2/retro-hsyn-Cre and locally expressing AAV2/9-EF1α-EGFP into the DMV (Fig. 6g). EGFP expression in the mPOA and DMV confirmed accurate injection (Fig. S12a). The retrograde axonal transport of the DMV-targeted AAV2/retro-hsyn-Cre enabled Cre recombinase expression in PVN neurons projecting to DMV. The anterograde transneuronal transport of AAV2/1-EF1a-DIO-FLP from mPOA to PVN enabled Cre-dependent Flp expression in DMV-projecting and mPOA-recipient PVN neurons, which subsequently induced Flp-dependent hM4Di-mCherry in these neurons (Fig. S12b). Consistent with results from inhibition of single neural populations or circuits, the chemogenetic inhibition of DMV-projecting and mPOA-recipient PVN neurons blunted HFA-induced increases in c-Fos expression (Fig. S12b, c), blood glucose levels (Fig. S12d), intestinal glucose absorption (Fig. 6h), and SGLT1 protein expression in the proximal small intestine (Fig. 6i). In summary, our findings strongly suggest that the CNS enhances intestinal glucose uptake through the mPOA^Glu → PVN^CRF → DMV^ChAT circuit in response to HAT exposure.

Discussion

This study investigates how the CNS adapts to HAT by enhancing intestinal glucose uptake through a brain-gut axis. Our findings reveal that HAT activates glutamatergic neurons in the mPOA, a crucial region for temperature regulation. This activation stimulates vagal motor nerves via the mPOA^Glu → PVN^CRF → DMV^ChAT circuit, leading to increased expression of the glucose transporter SGLT1 in the proximal small intestine. We propose that this reflex is homeostatic: heat stress raises cellular ATP demand for protective programs, but increasing reliance on protein and fat can increase diet-induced thermogenesis^11,12. Preferentially boosting glucose absorption can therefore provide rapid energy while limiting additional heat production¹³.

At first glance, this mechanism appears paradoxical because HAT reduces overall food intake²² (Fig. S1a), which would be expected to reduce energy availability. We propose that the nervous system partially compensates for reduced intake by prioritizing carbohydrate digestion and absorption through specific vagal motor circuits. Consistent with this selective prioritization, HAT increased SGLT1 expression but did not alter the expression of proteins related to amino acid or fatty acid absorption (Fig. S2a, b). This may occur because animals continuously adapt to their environment by evaluating which foods are urgently needed and safe under different physiological states. For example, during the recovery period after catabolism, high protein intake is restricted⁴².

This idea is consistent with “labeled line” organization of DMV output, in which distinct DMV neuron subtypes regulate specific digestive or absorptive processes by targeting different enteric neurons⁴³. For instance, GABRA1-expressing DMV neurons can significantly enhance intestinal fat absorption⁴⁰, suggesting that the body could enhance glucose digestion and absorption through specific neural circuits. Therefore, the reduction in food intake caused by HAT may, in a compensatory response, trigger enhanced intestinal glucose absorption to alleviate HAT-induced harm.

Heat stress triggers defensive energy demands to protect against cellular damage, requiring increased energy for protection^44,45. Our first intriguing observation was that circulating glucose, the primary energy source for the CNS⁴⁶, increased in mice exposed to HAT after a 12-hour fast and a 30-minute refeeding period. Interestingly, food intake remained similar between HAT and RT groups within the first hour of refeeding. We first investigated whether this increase was due to stress-induced activation of the HPA axis. However, blocking the HPA axis with a CRFR1 antagonist did not prevent the HAT-induced blood glucose elevation. Moreover, HAT neither affected serum corticosterone levels nor liver glycogen-metabolism related protein expression, suggesting alternative mechanisms. These findings align with previous research showing that HAT does not affect the HPA axis⁴⁷ and that glucocorticoids lose their ability to trigger glucose release from hepatocytes at high temperatures⁴⁸. Additionally, during fasting, hepatic glycogenolysis serves as the primary source of blood glucose⁴⁹. The unchanged liver glycogen metabolism may be attributed to 12-hour overnight fasting protocol. This extended fast likely depleted stored energy, potentially exhausting liver glycogen reserves over time⁵⁰. Like gluconeogenesis and hepatic glycogen metabolism, serum insulin levels in refed mice showed no changes. While we examined multiple key factors related to glucose metabolism, circulating glucose is influenced by many factors, making it difficult to rule out the potential impact of endogenous glucose metabolism under HAT conditions. However, it is important to note that exogenous isotope-labeled glucose uptake is the gold standard for detecting intestinal glucose absorption. Our data definitively confirm that HAT enhances intestinal glucose absorption.

Prior research has demonstrated that prolonged exposure of mice to a 38 °C environment (two hours daily for two weeks) compromises their intestinal barrier and enhances intestinal permeability⁵¹. To evaluate the impact of a 30-minute HAT exposure on intestinal function, we analyzed the tight junction in the proximal small intestine tissue⁵². Notably, in contrast to previous findings, our study revealed that HAT did not significantly affect the intestinal barrier. This variance may be attributed to two primary factors: the potential benefits of appropriate fasting on intestinal function recover⁵³, and the relatively short duration and moderate temperature of heat exposure in our protocol, which may not have been sufficient to cause intestinal dysfunction.

Our research revealed that HAT specifically increases the expression of the glucose transporter SGLT1 in the proximal small intestine, without significantly affecting amino acid or fatty acid transport. This preferential glucose absorption may be an adaptive response to meet the increased energy demands during heat stress. Unlike amino acid or fatty acid, glucose is readily used for ATP generation^54,55, crucial for producing HSPs that protect against heat-induced cellular damage⁵. The higher plasma levels of isotope-labeled ¹³C₆ glucose in HAT-exposed mice after gavage, alongside the consistently lower blood glucose levels in fasting mice under HAT conditions compared to the RT group, further support this hypothesis. This finding aligns with the common shift towards increased carbohydrate use and decreased fat use during heat stress^2,56–59, suggesting a physiological basis for this preferential glucose absorption. However, this contrasts with findings from a previous study, which observed that HAT increased intestinal motility and decreased glucose absorption in young rats (weighing between 40-60 g)⁴¹. We hypothesize that this discrepancy might be due to young mice having immature intestinal function, as HAT can significantly increase the gastric emptying rate in young rats, but does not affect adult rats⁶⁰. This is consistent with our showing that HAT does not affect intestinal transit function in adult mice, or young animals lacking the physiological adaptations necessary to quickly adjust to elevated environmental temperatures⁴¹.

The investigation into the effects of HAT exposure on the liver, a crucial metabolic orga⁶¹, further revealed significant benefits of glucose supplementation. HAT damaged hepatocyte organelles, triggering defensive responses, including increased HSP expression and lipid droplet content in hepatocytes. Pre-administration of glucose mitigated HAT-induced damage to hepatocyte organelles. Notably, glucose supplementation also enhanced the protective response by further increasing HSP expression and lipid droplet content. The increase in lipid droplets, which serve as storage units for neutral lipids and components of the cellular antioxidant system²⁸, suggests that glucose intake may improve hepatocytes’ ability to resist oxidative stress under HAT conditions. These findings highlight glucose supplementation’s potential as a protective strategy against HAT-induced liver damage. Additionally, HAT-induced cellular damage can lead to severe heatstroke and death⁶². In our study, under extreme high-temperature exposure (42 °C), mice without glucose supplementation exhibited a significantly higher rate of heat shock within 100 minutes compared to mice supplemented with 500 mg/kg glucose, further supporting glucose’s essential role in metabolic defense against HAT exposure. Moreover, we found that HAT also causes damage to the mitochondria of intestinal epithelial cells, but this did not reach statistical significance (0.05 < p < 0.1). This may be the liver, as the organ with the most vigorous metabolism, experiences liver injury as a typical complication of HAT stress, which is usually the direct cause of death in heat stroke patients⁶³ and can exhibit organelle damage even under acute high-temperature exposure.

We next investigated the potential contribution of vagal motor neurons, which play a major regulatory role in intestinal nutrient digestion and absorption^33,64. Evidence from three distinct mouse models consistently supports the mediating role of vagal motor input in HAT-induced increases in intestinal glucose absorption. The first mouse model employed sdVx, which denervates both afferent and efferent vagus nerves communicating with the gut^65,66. This procedure increased stomach volume due to impaired gastrointestinal motility, validating successful denervation³⁰. Notably, sdVx prevented the HAT-induced increase in intestinal glucose absorption, supporting the essential role of vagal input. Interestingly, sdVx also led to lower blood glucose levels during HAT exposure. We speculated that, under HAT conditions and with reduced intestinal glucose supply, the body might need to oxidize more stored glucose to maintain normal cellular functions. This observation aligns well with the observation that HAT reduces blood glucose levels in fasted mice. Therefore, increased respiration in organisms exposed to HAT may serve not only to increase heat dissipation but also to meet elevated oxygen consumption demands.

To differentiate the roles of the afferent and efferent vagus nerves, we developed a second chemical-blockade mouse model. We systemically injected atropine, a selective antagonist of acetylcholine M-type receptors⁶⁷, to inhibit postganglionic cholinergic transmission of vagal motor nerves in the intestine²⁹. Initially, atropine treatment combined with fast-refeeding experiments yielded inconclusive results because its significant appetite-suppressing effects confounded our investigation of intestinal glucose absorption. However, direct oral gavage of an equivalent amount of glucose, coupled with atropine treatment, produced results comparable to those observed in sdVx mice. These findings provide compelling evidence for the critical role of vagal motor neurons in modulating intestinal glucose absorption.

It is important to note that systemic chemical blockage affects multiple organs beyond the intestines, as vagal motor nerves extensively innervate various visceral organ⁴⁰. To further improve specificity, we generated a third mouse model with selective chemogenetic inhibition of intestinal cholinergic postganglionic neurons. We injected AAV-DIO-hM4D(Gi)-mCherry at multiple points into the proximal small intestine of ChAT-Cre mice. After testing various viral serotypes, only the blood-brain barrier-crossing serotype PHP.eB showed good infection efficiency in neurons within the intestinal myenteric plexus. This approach specifically inhibited postganglionic vagal motor neurons. The results showed that inhibiting these neurons prevented the HAT-induced increase in blood glucose levels. Collectively, the results from all three mouse models provide strong evidence that vagal motor input mediates HAT-induced increases in intestinal glucose absorption.

Our research then focused on identifying the CNS circuit responsible for regulating intestinal glucose uptake under HAT conditions. Using viral tracing and c-Fos mapping techniques, we discovered that the mPOA, PVN, and DMV not only project to the proximal small intestine but also exhibit increased activity under HAT stimulation. These findings suggest a potential role for these three nuclei in HAT-induced brain-gut communication. This hypothesis aligns with the well-known role of mPOA in ambient temperature sensing and metabolic adaptation to heat stress⁶. Studies have shown that prolonged exposure to high (but non-lethal) temperatures enhances both the activity of leptin receptor-expressing neurons in the anteroventral preoptic area (VMPO^LepR) ³⁸and the expression of brain-derived neurotrophic factor (BDNF) in the medial preoptic area (MPO)³⁹, thereby improving heat tolerance in mice. Although we did not detect a neural circuit connection between POA^BDNF or POA^LepR neurons and the proximal small intestine by fluorescence colocalization, the long-distance trans-synaptic transport of PRV likely results in labeling only a subset of neurons connected to the PRV-GFP injection site in the proximal small intestine. Therefore, the current labeling may not fully capture the complete circuitry. Future work will be required to systematically investigate how these neurons interact with the known gut–brain circuits. Notably, the PVN, composed primarily of autonomic and endocrine neurons, significantly contributes to visceral regulation⁶⁸, while the DMV houses the preganglionic neurons of the vagal motor nerve⁴³. Lyu et al.‘s study, DMV activation promoted fat absorption and weight gain⁴⁰, Although sdVx and atropine did not affect intestinal glucose absorption, this may be because the intestine, as the second brain, is capable of self-regulation in the absence of central control⁶⁹. However, when we activated DMV^ChAT, it also affected intestinal glucose absorption, except that activation of the entire DMV^ChAT significantly inhibited glucose absorption, which may be similar to HAT exposure in young rats, reducing intestinal transit time, thereby decreasing glucose absorption efficiency⁴¹. Collectively, this indicates that DMV^ChAT not only regulates intestinal motility but also plays an important role in nutrient absorption. However, our experiment showed that when male mice fasted overnight for 12 h and were refed for 30 min under HAT conditions, their excited DMV^ChAT neurons enhanced intestinal glucose absorption while intestinal fatty acid absorption and intestinal transit time remained unchanged. This may stem from different subtypes of DMV neurons exerting regulatory control over the digestion and absorption of different nutrients by targeting distinct enteric neurons⁴³. We believe this selective nutrient absorption represents an adaptive response to HAT. It is worth noting that with respect to gut motility, although we measured total intestinal transit time, we did not directly assess gastric emptying rate. Future work incorporating direct assessments of gastric emptying will be valuable for further clarifying the mechanism.

Further investigation revealed mPOA→PVN → DMV neural circuit. We demonstrated its existence through sophisticated tracing experiments, including multisynaptic PRV retrograde tracing and monosynaptic retrograde rabies virus systems. In our PRV retrograde tracing experiment, we observed a gradual increase in PRV-EGFP-positive neurons in the mPOA and PVN as infection time increased. This aligns with PRV’s characteristic bottom-up infection of neural circuits over time⁷⁰, suggesting a potential network hierarchy: the mPOA is likely upstream of the PVN, which is potentially upstream of the DMV. The subsequent monosynaptic retrograde rabies tracing confirmed the connection between PVN neurons and DMV^ChAT neurons. Importantly, adding oG, a key glycoprotein for transsynaptic transmission⁷¹, to these Rabies EGFP-infected DMV^ChAT-projecting PVN neurons enabled further monosynaptic tracing to mPOA neurons. These findings align well with studies showing that PVN^CRF neurons modulate intestinal motility via DMV^ChAT72 and that HAT-induced anorexia is regulated by mPOA via the PVN nucleus²². Our research offers anatomical evidence of monosynaptic connections between mPOA→PVN→DMV^ChAT. More importantly, through Fiber Photometry recordings, it was discovered that the mPOA→PVH → DMV circuit does not depend on feedback from intestinal glucose signals but is instead regulated by ambient temperature. As the ambient temperature rises, this circuit is activated to enhance glucose absorption.

To investigate the specific neuron types in this circuit, we analyzed cell-type-specific c-Fos expression under HAT conditions and used targeted chemogenetic inhibition. We identified three crucial neural populations: mPOA^Glu, PVN^CRF, and DMV^ChAT. Chemogenetic inhibition of distinct neural populations, single circuits, or multisynaptic circuits effectively suppressed the HAT-induced enhancement of intestinal glucose absorption, confirming the importance of these circuits in this adaptive response. It should be noted that warm-responsive neurons in mPOA are mainly glutamatergic neurons (such as those that express the peptides Pituitary Adenylate Cyclase-Activating Polypeptide (PACAP), BDNF, or the Quadruple RFamide peptide (QRFP); and LepR, estrogen (ERα), or prostaglandin E2 (EP3R))³⁷, PVN^CRF play an important role in the specific sensing of intestinal glucose⁷³, and DMV^ChAT can significantly improve the absorption of fat under the HFD, suggesting that DMV^ChAT neurons can selectively absorb intestinal nutrients according to the state of the body⁴⁰.

Previous studies have highlighted significant metabolic and behavioral functions of individual neural populations and circuits. For example, the mPOA^Glu is linked to temperature-induced metabolic adaptation⁶ and stress-induced anxiety⁷⁴.HAT primarily suppresses food intake through the mPOA^Glu → PVN circuit²². Moreover, PVN^CRF neurons not only perform traditional neuroendocrine functions but also regulate visceral activities via the HPA axis and autonomic nervous system^20,75. While PVN^CRF inhibits DMV^ChAT-induced gastric motility disorders in a gastric dilation model, it can also activate downstream neurons through glutamate and CRF release⁷⁶. Consequently, PVN^CRF’s regulation of DMV^ChAT may vary depending on the type of neurotransmitter released. To date, no research has linked these three neural population groups or demonstrated the crucial role of this multisynaptic circuit in intestinal glucose absorption. Our study provides empirical evidence for this brain-gut circuit in metabolic adaptation to HAT exposure.

In conclusion, our research elucidates an adaptive mechanism that increases intestinal glucose absorption in response to elevated ambient temperatures. This suggests that increased glucose intake might help prevent HAT-induced cellular damage and heat stroke. Nevertheless, it remains unclear whether HAT directly promotes glucose absorption through the vagus motor nerve or indirectly facilitates absorption by enhancing digestion. Additional research is required to fully elucidate the complexities of this mechanism, such as the specific neurotransmitters and the exact nature of neuronal connections in this circuit.

Methods

Animals

The mice in this experiment were housed in a controlled indoor environment with a constant temperature of 22 °C–23 °C, humidity of 60%, and a 12-hour light/dark cycle. They were provided with adequate standardized pellet feed suitable for mouse growth and clean drinking water. Animal care and experimental procedures at South China Agricultural University adhered to “The Instructive Notions regarding Caring for Laboratory Animals” issued by the Ministry of Science and Technology of the People’s Republic of China. These procedures were approved by the Animal Subjects Committee of South China Agricultural University (SYXK2022-0136). All mice used in the experiments were healthy and over 7 weeks old. It should be noted that, because female mice may introduce confounding factors such as the estrous cycle, most of the in vivo experiments in this study were conducted in male mice to improve reproducibility and consistency. Only in the anatomical (neural circuits) studies did we use both male and female mice, and we observed no sex-specific differences. The specific strains of mice used are listed below: C57BL/6 J (JAX, 000664), ChAT-IRES-Cre (JAX, 006410), CRF-IRES-Cre (JAX, 012704), Vglut2-IRES-Cre (JAX, 016963), Rosa26-LSL-tdTomato mice (JAX, 007914), SGLT1^flox/+ (Cyagen, CKOCMP-20537-Slc5a1-B6J-VA), Vglut2-Flpo (JAX, 030212), CRF-Flpo (JAX, 031559), ChAT-Flpo (JAX, 036281), LepR-Cre::LSL-tdTomato mice were kindly provided by Dr. Liu Yi from Southern Medical University.

Heat treatment and blood glucose measurement

Male mice were fasted for 12 h prior to exposure to HAT. After measuring baseline blood glucose, the mice were placed in an artificial climate chamber (37 °C, 60% relative humidity [RH])) and allowed to resume feeding. Tail vein blood glucose was measured at 15, 30, 60 and 120 min using a Yuwell blood glucose meter (model 580). For the fasted group, mice were placed in the climate chamber but not allowed to resume feeding, with blood glucose measured at 15, 30, 45, 60, 75 and 90 min. In the additional glucose supplementation experiment, mice were gavaged with a 500 m g/kg glucose solution every 30 min, and blood glucose was measured at 15, 30, 60, 90 and 120 min. The RT control group was maintained in an environment at 25 °C and 60% humidity.

Growth performance of mice under chronic heat stress

Male C57BL/6 J mice were randomly allocated into two groups based on their initial body weight using a randomized complete block design: an RT group (25 °C, 60% RH) and a chronic HAT group (37 °C, 60% RH). All mice were individually housed in controlled-environment chambers under a 12-h light/12-h dark cycle. Mice in the HAT group were subjected to high-temperature treatment for eight consecutive hours daily during the fixed period of 09:00–17:00, while maintaining the same environmental conditions as the RT group during the remaining time. Throughout the experiment, feed intake was monitored daily at 08:00 by measuring the weight of the feed provided and the residual feed to calculate the 24 h consumption, with fresh feed replenished immediately thereafter. Body weight was measured every two days at 08:30. All mice had ad libitum access to water throughout the experimental period.

D-[¹³C₆] glucose assay

Stable isotope-labeled glucose (D-[¹³C₆] glucose) was detected by liquid chromatography tandem mass spectrometry (LC-MS/MS) to determine glucose uptake in mice⁷⁷. The detailed sample preparation is as follows: After a 12-hour fast, male C57BL/6 J mice were gavaged with 500 mg/kg of D-[¹³C₆] glucose and then placed in artificial climate chambers set to HAT and RT conditions. Thirty minutes after heat treatment, blood was collected from the retro-orbital venous plexus of mice into clean centrifuge tubes. Plasma samples were centrifuged at 2500 × g for 15 min at 4 °C. Subsequently, 200 µL of plasma was thoroughly mixed with 400 µL of acetonitrile, vortexed for 30 s, and centrifuged at 12,000 × g for 10 min at 4 °C to precipitate plasma proteins. The supernatant was transferred to a new centrifuge tube, dried in a vacuum freeze-dryer, and the dried extract was dissolved in 100 µL of water. A series of standard solutions with gradient concentrations was prepared by diluting the standard with water for subsequent analysis.

For derivatization, 20 µl of the above sample was mixed with 100 µl of derivatization reagent (containing 0.385 g Butyl Aminobenzoate, 0.625 g Sodium Cyanoborohydride, 4.63 ml Methanol, and 0.375 ml Glacial Acetic Acid). The mixture was heated at 80 °C for 1 h. After the reaction, the mixture was transferred to a solid phase extraction (SPE) column, which was pre-activated with methanol, pretreated with water, and equilibrated with acetonitrile-water (5:95 v/v). The sample was then washed twice with acetonitrile-water (5:95 v/v) and eluted with acetonitrile-water (30:70 v/v). 5 µL of the eluate was thoroughly mixed with 100 µL of acetonitrile-water (30:70 v/v) and filtered through a 0.22 µm nylon membrane (Agilent ValueLab), preparing it for LC-MS/MS analysis.

Equipped with an LC analysis was performed via an ACQUITY UPLC ultra-high performance liquid chromatograph (Waters, USA) using a XBridge BEH C₁₈ XP column (100 × 3.0 mm, 2.5 μm) (Waters, USA). The mobile phases consisted of (A) acetonitrile and (B) 5 mmol L − 1 ammonium acetate aqueous solution, with a linear gradient elution at a flow rate of 0.4 mL min−1. The gradient program was as follows: 0.0–1.0 min 30–50% A, 1.0–3.0 min 50–70% A, 3.0–4.5 min 70–80% A, 4.5–5.0 min 80–100% A, 5.0–5.1 min 100–30% A, 5.1–6.0 min 30% A, with an injection volume of 3 μL. MS/MS analysis was operated on a Triple Quad^TM 5500 mass spectrometer system (AB SCIEX, USA) in positive electrospray ionization mode (ESI⁺) applied with a multiple reaction monitoring (MRM) mode. The transitions (m/z) used for quantification and qualification were 364.2 → 290.1 (collision energy of 10 eV) and 364.2 → 207.1 (collision energy of 10 eV), respectively. The retention time was 3.48 min. The capillary voltage was set at 5.5 kV, with a drying gas flow rate of 1000 L/h, temperature of 150 °C, and atomizing gas pressure of 50 psi. Plasma samples from each mouse were tested once. Data acquisition and processing were performed using the MultiQuant algorithm from MultiQuant 3.0.2 (Analyst; AB SCIEX). Figure S1h demonstrates that D-[¹³C₆] glucose derivatives exhibit a single sharp peak at their retention time without any interfering peaks. This confirms our method’s excellent retention and separation capabilities for D-[¹³C₆] glucose derivatives.

A linear standard curve was created by plotting the mass spectrometric response (peak area) of D-[¹³C₆] glucose derivatives against different standard concentrations (Fig. S1g). This standard curve was then employed to quantify the ¹³C₆ content in plasma samples.

Corticosterone measurement

After 12 h of fasting, male mice were divided into three groups: one group was refed and placed in a HAT environment, another group continued fasting and was placed in a HAT environment, while the control group was kept at room temperature. After 30 min, mice were immediately anesthetized with isoflurane. Blood was collected by enucleation and left at room temperature for two hours. The samples were then centrifuged at 3000 × g for 10 min to isolate the serum for subsequent assays. Serum corticosterone levels were determined using an enzyme-linked immunosorbent assay kit according to the manufacturer’s instructions.

Quantitative real-time PCR

Gene expression analysis of proximal small intestine and liver samples was performed using qPCR. Briefly, total RNA was extracted from target tissues using TRIzol reagent (TaKaRa) and quantified using a Bio Photometer 6131 spectrophotometer (Eppendorf). The mRNA levels of target genes were measured using a Bio-Rad qPCR instrument with SYBR qPCR Mix (TaKaRa). Relative expression of target genes was calculated using the 2^−ΔΔCt method, with β-actin serving as the reference gene for data normalization. Primer (Sangon Biotech) sequences are detailed in the primer information table provided in the Supplementary Information Table 1.

Protein extraction and Western blot assay

Total protein was extracted from proximal small intestine tissue using radioimmunoprecipitation (RIPA) buffer (Beyotime, P0013B) containing protease and phosphatase inhibitor cocktail (Beyotime, P1045). Protein concentration was determined using the BCA assay kit (Thermo Fisher Scientific). The total protein was then separated by SDS-PAGE and transferred to a PVDF membrane (Millipore). Membranes were blocked with blocking buffer (Tris-buffered saline containing 0.05% Tween-20 and 5% non-fat milk) and incubated with primary antibodies against SGLT1 and GLUT2 at 4 °C overnight, followed by incubation with an HRP-conjugated secondary antibody (Bioworld) for 1 h at room temperature. Proteins on the membranes were detected using a FluorChem M Fluorescent Imaging System (Protein Simple), and band density was analyzed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). Antibody details are listed in the antibody information table provided in Supplementary Table 2.

Experiment to antagonize CRFR1

Male C57BL/6 J mice, fasted for 12 h, received intraperitoneal injections of antalarmin (30 mg/kg) dissolved in saline. After 30 min, the mice were placed in an artificial climate chamber to measure fasting blood glucose. Following HAT and RT treatments, feeding was resumed. Blood glucose levels were then measured at 15, 30, 60 and 120 min post-treatment.

Experiment to inhibit SGLT1

Male C57BL/6 J mice, fasted for 12 h, received phlorizin dissolved in saline via gavage at a concentration of 100 mg/kg. Thirty minutes later, the mice were placed in an artificial climate chamber to measure their fasting blood glucose. Subsequently, they underwent HAT and RT treatments and were allowed to resume feeding. Blood glucose levels were then measured at 15, 30, 60 and 120 min.

Transmission electron microscopy observation and analysis

To qualitatively visualize the morphological responses of liver and proximal small intestine tissues to HAT treatment, we used transmission electron microscopy to observe hepatocytes 30 min after 12 h of fasting, followed by resumed feeding in mice under HAT or RT conditions (n = 3 per group). Briefly, 30 min post-HAT, tissue was excised and immediately fixed using 2% glutaraldehyde in 0.1 M sodium citrate buffer (pH 7.4) solution (Servicebio, G1102). This was followed by fixation with 1% osmium tetroxide at room temperature in the dark for 2 hours. The tissue was then rinsed and dehydrated through an ethanol gradient, followed by two propylene oxide exchanges. Samples were embedded in 812 resin and polymerized in a 60 °C oven for 48 h. Subsequently, sample blocks were trimmed and cut into 60–80 nm thin sections, which were collected on 150-mesh square formvar-coated copper grids. The grids were stained with 2% uranyl acetate saturated alcohol solution in the dark for 8 min, followed by 2.6% lead citrate solution (avoiding carbon dioxide) for 8 min. After rinsing three times with ultrapure water and drying overnight at room temperature, the samples were observed under a transmission electron microscope (HITACHI, HT7800/HT7700) for image capture and analysis.

Heat shock rate experiment under extreme environmental temperatures

After a 12-hour fast, male C57BL/6 J mice were administered 200 µl of either saline or 5% glucose solution via gavage. The mice were then placed in an artificial climate chamber set at 42 °C and 60% humidity, with food access restored. The cages were shaken every 20 min, and mice that did not move within 2 min were recorded as having heat shock. The final heat shock rate was calculated after 100 minutes.

Determination of intestinal digestive enzyme activities

Digestive enzyme activities in the proximal small intestine were measured using commercial assay kits. Prior to the experiment, all mice were fasted overnight for 12 h with free access to water. After fasting, the mice were randomly divided into two groups and placed in either an RT or HAT environment, followed by refeeding. At 30 min post-refeeding, the mice were immediately anesthetized and sacrificed. Proximal small intestine tissues were collected for subsequent analysis. All digestive enzyme activities were determined using commercial kits purchased from Nanjing Jiancheng Bioengineering Institute (Nanjing, China), strictly following the manufacturers’ instructions.

Measurement of hepatic ATP content

Animals and tissue preparation: After a 12-hour overnight fast, mice were randomly divided into three groups: the RT group, the HAT group, and the HAT+Glu group (administered an equal volume of glucose solution via gavage at a dose of 500 mg/kg body weight and immediately subjected to HAT conditions). Following 30 min of treatment in each respective group, the mice were euthanized under isoflurane anesthesia. Livers were rapidly dissected, rinsed to remove residual blood and connective tissue, snap-frozen in liquid nitrogen, and subsequently stored at −80 °C until further analysis.

Sample preparation: all sample preparation procedures were performed at low temperature. Liver tissues were removed from the −80 °C freezer and weighed quickly under liquid nitrogen protection. Tissues were immediately placed into a mortar pre-cooled to 4 °C, and pre-chilled 0.4 mol/L perchloric acid (HClO₄) was added at a ratio of 1:5 (g: mL). The mixture was thoroughly homogenized on ice until completely lysed. The homogenate was transferred to a pre-cooled centrifuge tube and centrifuged at 3500 r/min for 10 min at 4 °C. The supernatant was collected into a new pre-cooled centrifuge tube, and an equal volume of 1 mol/L dipotassium hydrogen phosphate (K₂HPO₄) solution was slowly added. The pH was adjusted to 6.5 by inversion mixing, and the mixture was allowed to stand at room temperature for 10 min. After a second centrifugation at 3500 r/min for 10 min at 4 °C to remove precipitates, the supernatant was filtered through a 0.22 μm organic phase filter membrane. The resulting filtrate was collected as the test sample, stored at 4 °C in the dark, and analyzed within 12 h.

HPLC analysis was performed on an Agilent 1260 series system (Agilent Technologies, USA) at room temperature (<25 °C). The mobile phase consisted of 50 mmol/L potassium phosphate buffer (pH 6.5) delivered at a flow rate of 1 mL/min. The detection wavelength was set at 254 nm, and the injection volume was 20 μL. Quantification was achieved using an external standard method.

Intestine motility measurements

For measurement of total intestinal transit time, mice were given an oral gavage of 6% carmine red dissolved in 0.5% methylcellulose (mixed with sterile 0.9% saline). Observation was conducted every 10 min. Total intestinal transit time was defined as the interval from oral gavage to the excretion of the first fecal pellet containing carmine red.

Subdiaphragmatic vagotomy

The male C57BL/6 J mice were anesthetized using a mixture of xylazine (10 mg/kg) and ketamine (100 mg/kg) diluted in sterile saline. Then, the abdominal cavity was opened to fully expose the liver and stomach. The liver was gently moved aside to prevent injury. Using forceps, the esophagus was carefully pulled out, revealing the ventral branch of the subphrenic vagus nerve. This nerve was lifted with sharp forceps, and a 2–3 mm section was cut. Next, the esophagus was gently flipped, and the surrounding fat tissue was separated to expose the dorsal branch of the subphrenic vagus nerve. Then, a 2–3 mm segment of this nerve was removed using sharp forceps. Finally, the muscle and skin layers were sutured sequentially with sterile sutures, and iodine was applied to the wound to prevent infection. The mice were allowed to recover for one week before any experimental treatments were conducted.

Cerebral stereotaxic injection surgery

Male mice were anesthetized with a mixture of xylazine (10 mg/kg) and ketamine (100 mg/kg) diluted in sterile saline, then secured in a single-arm stereotaxic frame. After exposing the skull, a small hole was drilled at the target nucleus location using a cranial drill. The virus was then injected using a microsyringe at a rate of 30 nL/min. The syringe was left in place for 5 min after injection before being withdrawn.

The DMV was injected bilaterally with 200 nl of AAV virus (coordinates relative to bregma: ML: ±0.29, AP: −7.56, DV: −3.47). The PVN received bilateral injections of 200 nl AAV virus (coordinates relative to bregma: ML: ±0.18, AP: −0.94, DV: −4.78). The mPOA was injected bilaterally with 200 nl of AAV virus (coordinates relative to bregma: ML: ±0.16, AP: 0.26, DV: −5.29). All mice underwent a three-week recovery period to ensure they were in optimal condition before experimental procedures began.

For chemogenetic experiments, AAV2/9-hSyn-DIO-hM3D(Gq)-mCherry-WPRE-pA (Shanghai Taitool Bioscience) was bilaterally injected into the DMV of ChAT-IRES-Cre mice, AAV2/9-hSyn-DIO-hM4D(Gi)-mCherry-WPRE-pA (Shanghai Taitool Bioscience) was bilaterally injected into the DMV of ChAT-IRES-Cre mice, the PVN of CRF-IRES-Cre mice, and the mPOA of Vglut2-IRES-Cre mice.

To validate mPOA^Glu → PVN connectivity, male C57BL/6 J mice received bilateral injections of AAV2/9-Vglut2-CRE-WPRE-hGH polyA and AAV/2/9-hsyn-fDIO-hM4D(Gi)-mCherry-WPRE-hGH polyA (both from BrainVTA) in the mPOA. The PVN was injected bilaterally with AAV2retro-EF1α-DIO-FLP-WPRE-hGH pA (BrainVTA) and AAV2/9-EF1α-EGFP-WPRE-pA (Shanghai Taitool Bioscience), with the latter serving to label the injection site.

The specificity of each promoter was confirmed using a Cre-Flp intersectional strategy. AAV2/9-ChAT-Cre was injected into the DMV of ChAT-Flp mice, along with AAV-DIO-GFP and AAV-fDIO-mCherry (both from BrainVTA). By the same token, AAV2/9-CRF-Cre and AAV2/9-Vglut2-Cre were delivered to the PVN (in CRF-Flp mice) and mPOA (in Vglut2-Flp mice), respectively, using the identical cocktail of reporter viruses.

For mPOA→PVN^CRF validation, male C57BL/6 J mice received bilateral PVN injections of AAV2/9-CRF-CRE-WPRE-hGH polyA and AAV/2/9-hsyn-fDIO-hM4D(Gi)-mCherry-WPRE-hGH polyA (both from BrainVTA). The mPOA was injected bilaterally with AAV2/1-EF1α-DIO-FLP-WPRE-hGH pA (BrainVTA) and AAV2/9-EF1α-EGFP-WPRE-pA (Shanghai Taitool Bioscience), with the latter marking the injection site.

To validate the PVN → DMV connectivity, two independent viral injection experiments were performed. In the first experiment, male ChAT-Cre mice received bilateral injections of AAV2/9-hsyn-fDIO-hM3D(Gq)-EGFP-WPRE-hGH polyA (BrainVTA) into the DMV, followed by bilateral PVN infusion of AAV2/1-EF1α-DIO-FLP-WPRE-hGH pA (BrainVTA). In the second experiment, male C57BL/6 J mice were bilaterally injected in the DMV with a mixture of AAV2/9-ChAT-CRE-WPRE-hGH polyA and AAV2/9-hsyn-fDIO-hM4D(Gi)-mCherry-WPRE-hGH polyA (both from BrainVTA). For injection localization, AAV2/1-EF1α-DIO-FLP-WPRE-hGH pA (BrainVTA) combined with AAV2/9-EF1α-EGFP-WPRE-pA (Shanghai Taitool Bioscience) was bilaterally delivered into the PVN, in which the EGFP-containing virus was used to label the injection site.

For mPOA→PVN → DMV validation, male C57BL/6 J mice received the following injections: bilaterally in the mPOA, AAV2/1-EF1α-DIO-FLP-WPRE-hGH pA (BrainVTA) and AAV2/9-EF1α-EGFP-WPRE-pA (Shanghai Taitool Bioscience); bilaterally in the PVN, AAV/2/9-hsyn-fDIO-hM4D(Gi)-mCherry-WPRE-hGH polyA (BrainVTA); and bilaterally in the DMV, AAV2/1-hsyn-CRE-WPRE-hGH-polyA (BrainVTA) and AAV2/9-EF1α-EGFP-WPRE-pA (Shanghai Taitool Bioscience). The AAV2/9-EF1α-EGFP-WPRE-pA injections were used to label the injection sites.

Anterograde-tracing

C57BL/6 J mice were bilaterally injected with 50 nL of HSV-tdTomato in the mPOA. After injection, the mice were returned to their cages. Three days later, the mice were perfused, and their brains and duodena were removed for observation of viral expression.

Peripheral organs viral injections

The viral constructs were loaded into a Nanofil™ 36 G beveled needle. PRV-CAG-EGFP (BrainVTA) virus was injected into the lamina propria of the proximal small intestine of C57BL/6 J mice. Multiple injections of 20 nL each were administered, totaling 200 nL per mouse. AAVPHP.eB-hSyn-DIO-hM4D(Gi)-mCherry (Addgene) virus was injected into the lamina propria of the proximal small intestine of ChAT-Cre mice. Multiple 50 nL injections were given, totaling 1 μL per mouse.

For the SGLT1 knockout assay in the proximal small intestine, AAV2/9-CMV-Cre-pA viruses were injected at multiple points into the lamina propria of male SGLT1^flox/flox and male SGLT1^WT&flox+ mice. Each injection was 50 nL, with a total of 1 μL administered throughout the proximal small intestine.

Retrograde transsynaptic tracing

AAV2/9-CAG-DIO-TVA-mCherry-WPRE-pA and AAV2/9-CAG-FLEX-oG-WPRE-pA hybrid viral vectors (both from Shanghai Taitool Bioscience) were injected bilaterally into the DMV of ChAT-Cre mice. Two weeks later, rabies virus EnvA-SAD-B19(delta)G-Cre-EGFP (Shanghai Taitool Bioscience) was injected at the same location, while AAV2/9-CAG-FLEX-oG-WPRE-pA was injected into the PVN. One week after viral expression, the animals were euthanized, and brain tissue sections were prepared for further analysis.

Immunofluorescence

Male mice were anesthetized with isoflurane and rapidly perfused with sterile saline until blood was completely flushed out. They were then perfused with 4% paraformaldehyde until body stiffness indicated complete fixation. Brain tissues were carefully removed and placed in 4% paraformaldehyde overnight. The tissues underwent gradient dehydration in sucrose solutions (10, 20, 30%) and were cut into 30 μm sections using a flat slide slicer for subsequent immunofluorescence detection.

For c-Fos staining, sections were washed with 0.1 M PBS containing 0.1% Triton-X100 (PBST) for 30 min, then blocked with 0.3% PBST containing 5% goat serum for 1 h at room temperature. Sections were incubated with anti-c-Fos antibody overnight at 4 °C. After washing with 0.1% PBST for 30 min, sections were incubated with fluorescent secondary antibodies for 2 h at room temperature. Following a final 30-minute wash with 0.1% PBST, sections were sealed with quench-resistant sealing agent. Protein expression was observed using fluorescence microscopy.

For SGLT1 immunofluorescence, proximal small intestine tissue from mice was cut into 10 μm sections using a frozen microtome. Sections were blocked with 0.3% PBST containing 5% goat serum for 1 h, then incubated with anti-SGLT1 antibody overnight at 4 °C. Sections were rinsed with 0.1% PBST for 30 min, incubated with fluorescent secondary antibody goat anti-rabbit 488 for 2 h at room temperature, and rinsed again with 0.1% PBST for 30 min. Finally, sections were treated with an anti-quenching blocking agent, and protein expression was observed using fluorescence microscopy. Antibody details are listed in the antibody information table provided in Supplementary Table 2.

Retrograde viral tracing injection and neuronal colocalization analysis

To identify POA neuronal subtypes with synaptic connections to the small intestine, retrograde transneuronal tracing was performed using pseudorabies virus (PRV) carrying green fluorescent protein (GFP). Adult male mice were anesthetized with isoflurane, and the abdominal cavity was gently opened under sterile conditions. For viral tracing, the PRV-GFP virus was repeatedly injected into the submucosa of the proximal small intestine at multiple injection sites. After viral injection, the abdominal wall was sutured layer by layer, and the mice were placed on a constant-temperature heating pad until full recovery. To ensure sufficient retrograde viral transport and neuronal labeling, all mice were housed under standard feeding conditions for the same survival period.

Subsequently, mice were deeply anesthetized and transcardially perfused with 0.9% saline, followed by 4% paraformaldehyde. Whole brain tissues were dissected, post-fixed, and dehydrated in gradient sucrose solutions. Continuous coronal brain sections containing the POA were sliced using a freezing microtome. For BDNF neuronal immunofluorescence staining, brain sections were incubated with a primary antibody against BDNF (abcam, 1:1000 dilution), followed by the corresponding fluorescent secondary antibody. LepR-expressing neurons were visualized using LepR-Cre::LSL-tdTomato transgenic mice, in which LepR-positive neurons were specifically labeled with tdTomato red fluorescence.

All brain slices were imaged using a laser scanning confocal microscope. The distribution and colocalization of PRV-GFP-labeled neurons, BDNF-immunopositive neurons, and tdTomato-positive LepR neurons in the rostral, middle, and caudal regions of the POA were systematically observed and analyzed. Neuronal colocalization was evaluated by fluorescence overlapping signals. All image analyses were performed blindly using professional image analysis software.

In vivo fiber photometry experiments

Male C57BL/6 J mice were used for in vivo fiber photometry Ca²⁺ imaging. Mice received stereotaxic injections of AAV2/1-DIO-flp into the mPOA, AAV2/9-fDIO-GCaMP6s into the PVN, and AAV2/retro-hsyn-Cre into the DMV, followed by optic fiber implantation targeting the PVN nucleus. We utilized a fiber photometry system (Doric Lenses) to monitor Ca²⁺ signals in PVN nucleus populations. Prior to the commencement of each testing session, mice were acclimated to the behavioral chamber for a minimum of 15 min.

Statistical analysis

All statistical analyses and graph construction were performed using GraphPad Prism 9.00 (GraphPad Software, Inc., La Jolla, CA, USA). Comparisons were made by paired or unpaired two-tailed Student’s t-tests, 2-way ANOVA followed by Tukey’s multiple comparison post-hoc tests. Error bars in graphs represent mean ± standard error of the mean (SEM).

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

R.L. was the main contributor to the conduct of the study, data collection and analysis, data interpretation and manuscript writing. M.L. provided assistance in the brain stereotaxic injection experiment. Z.Z. was responsible for conducting the plasma ¹³C₆ content detection test. Q.Z. and M.C. provided assistance in the subdiaphragmatic vagotomy experiment. J.L., L.P., A.S., Q.Z., R.Q., Z.L., R.W., S.W., L.W. and G.S. contributed to the conduct of the study. P.X., Q.J., and C.Z. contributed to study design, data interpretation, and manuscript writing.

Peer review

Peer review information

Nature Communications thanks Sung-Yon Kim, Wei Shen and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Funding

This study was supported by the National Natural Science Foundation of China (32522099, 32102626 and U21A20245-3 to C.Z.; 32372968 and 32430098 to Q.J.; 32272954 to G.S.), the Guangdong Natural Science Funds for Distinguished Young Scholars (2023B1515020095 to C.Z.), the National Key Research and Development Program of China (2022YFD1300401 to Q.J.), and the Specific University Discipline Construction Project (2023B10564001 and 2023B10564003 to C.Z.).

Data availability

All data supporting the findings of this study are available within the paper and its Supplementary Information. Source data are provided with this paper.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-75522-7.