Regulation of Energy and Glucose Homeostasis by the Nucleus of the Solitary Tract and the Area Postrema

Article information

Endocrinol Metab. 2024;39(4):559-568

Publication date (electronic) : 2024 August 1

doi : https://doi.org/10.3803/EnM.2024.2025

Kyla Bruce ¹^,²^,^*, Ameth N. Garrido ²^,³^,^*, Song-Yang Zhang ², Tony K.T. Lam^,¹^,²^,³^,⁴^,⁵

¹Institute of Medical Science, University of Toronto, Toronto, ON, Canada

²Toronto General Hospital Research Institute, University Health Network (UHN), Toronto, ON, Canada

³Department of Physiology, University of Toronto, Toronto, ON, Canada

⁴Department of Medicine, Medicine, University of Toronto, Toronto, ON, Canada

⁵Banting and Best Diabetes Center, University of Toronto, Toronto, ON, Canada

Corresponding author: Tony K.T. Lam Toronto General Hospital Research Institute, University Health Network (UHN), MaRS Center, 101 College Street, TMDT 10-705, Toronto, ON M5G 1L7, Canada Tel: +1-4165817880, Fax: +1-4165817880, E-mail: tonykt.lam@uhn.ca

*These authors contributed equally to this work.

Received 2024 April 30; Revised 2024 May 29; Accepted 2024 June 7.

Abstract

The central nervous system regulates feeding, weight and glucose homeostasis in rodents and humans, but the site-specific mechanisms remain unclear. The dorsal vagal complex in the brainstem that contains the nucleus of the solitary tract (NTS) and area postrema (AP) emerges as a regulatory center that impacts energy and glucose balance by monitoring hormonal and nutrient changes. However, the specific mechanistic metabolic roles of the NTS and AP remain elusive. This mini-review highlights methods to study their distinct roles and recent findings on their metabolic differences and similarities of growth differentiation factor 15 (GDF15) action and glucose sensing in the NTS and AP. In summary, future research aims to characterize hormonal and glucose sensing mechanisms in the AP and/or NTS carries potential to unveil novel targets that lower weight and glucose levels in obesity and diabetes.

Keywords: Area postrema; Solitary nucleus; GDF15; Glucose; Feeding; Energy metabolism; Glucose metabolism; Stereotaxic surgery

INTRODUCTION

The central nervous system is critical in maintaining metabolic homeostasis by integrating immediate and long-term nutritional-dependent signals related to energy intake, usage, storage, and glucose metabolism [1-3]. Specifically, the hypothalamus has been established to detect changes in circulating hormones such as insulin [4-7], leptin [8,9], glucagon [10,11], and glucagon-like peptide-1 (GLP1) [12,13], as well as nutrients, including glucose [14-16], fatty acids [4,17], and amino acids [18,19] to regulate feeding, weight, and glucose homeostasis.

Recently, an extra-hypothalamic region within the brainstem termed the dorsal vagal complex (DVC) has emerged as a significant regulator of energy and glucose metabolism [20-22]. The DVC consists of the nucleus of the solitary tract (NTS), the area postrema (AP), and the dorsal motor nucleus of the vagus. Upon refeeding, neurons in the DVC are activated [23], and the DVC neurocircuitry integrates hypothalamic and gastrointestinal vagal afferent circuits to control feeding and glucose homeostasis [22,24-29]. Direct sensing of the DVC may regulate metabolic homeostasis as well. For example, intranasal administration of leptin into rodents reaches the DVC and hypothalamus [30], while in humans, intranasal leptin or insulin administration reduces body weight [31,32] and glucose production [33,34]. Specifically, direct infusions of insulin, glucagon, or glycine into the DVC of rats suppress appetite and hepatic glucose production [26,35-38]. However, leptin infusions in the NTS, but not in the AP, lower feeding [39], while activation of the GLP1 receptor (GLP1R) in the NTS suppresses appetite, but with minimal effects in the AP [40-43].

Thus, there are still considerable gaps in our understanding of how the NTS and AP distinctively or jointly influence metabolism. This mini-review explores methods used for NTS and AP-specific studies and examines recent findings on the metabolic effects of growth differentiation factor 15 (GDF15) and glucose in the NTS and/or AP. Ultimately, we advocate for future research to employ AP—or NTS-specific approaches to enhance our understanding of central metabolic regulation beyond the hypothalamus.

BRAIN GDF15 SENSING

GDF15 is a hormone from the transforming growth factor-β superfamily [44]. GDF15 is expressed in the kidney, liver, small intestine, adipose tissue, and muscle with the kidney and liver having the highest expression [45-47]. Injection of GDF15 lowers feeding and weight in high-fat fed rodents [48-51], while increasing energy expenditure [52,53], glucose tolerance [48,49], and insulin sensitivity [48,54].

GDF15 receptor, GDNF family receptor α-like (Gfral), is mostly expressed in the AP and NTS of the hindbrain [48-54]. Infusion of GDF15 activates Gfral-expressing cells within the AP and NTS [50], while AP and NTS lesions block the anorectic effect of peripheral GDF15 treatment [55,56]. Direct GDF15 administration into the lateral ventricle lowers feeding and weight in rodents as well [51,55]. In parallel, GDF15 improves glucose tolerance and insulin sensitivity, independent of changes in food intake and weight [54,57], and these effects were negated in Gfral knockout mice [54]. Although these collective findings suggest that the NTS and/or AP are the likely sites of GDF15 action, the direct impact of GDF15 on the AP versus the NTS remains elusive.

NTS or AP site specificity

A 26-gauge bilateral guide cannula or a 26-gauge single guide cannula can be inserted into the NTS or AP of rats, respectively, using stereotaxic surgeries to clarify their role in GDF15 metabolic regulation. The NTS is located within the DVC at the following coordinates in approximately 300 g male Sprague-Dawley rats: 0.0 mm on the occipital crest, ±0.4 mm lateral to the midline and 7.9 mm below the skull surface (Fig. 1A) [38]. Conversely, the AP of rats can be targeted via the following coordinates: 0.3 mm posterior to the occipital crest, 0.0 mm lateral to the midline, and 7.9 mm below the cranial surface (Fig. 1A) [58]. Of note, the NTS of 9- to 11-week-old mice can be stereotaxically targeted with bilateral steel guide cannulae positioned 2.0 mm above the NTS using a cannula holding bar in a 10-degree rostrocaudal angle at the following coordinates relative to the occipital suture: 0.5 mm anterior, 3 mm ventral, and ±0.4 mm lateral to the midline [20,59,60]. However, to the best of our knowledge, no study has implanted a chronic cannula into the AP of mice. For viral infections, the obex in mice,a landmark of the DVC may be used to target the NTS and AP. Viral infusions using the following coordinates relative to the obex: 0.2 mm posterior, ±0.2 mm lateral, and 0.2 mm ventral have been used to inject 100 nL of virus at a rate of 5 to 30 nL/min into the NTS of mice to virally and selectively infect the NTS [59,61,62]. Viral infusions directly into the obex of mice will target the AP using 35 nL of virus at a 2 nL/sec infusion rate [63-65].

Fig. 1.

Surgical targeting and metabolic impacts of the area postrema (AP) and nucleus of the solitary tract (NTS). (A) Schematic of rat skull and brain anatomy to provide reference for NTS and AP stereotaxic surgery (top). Images show coronal sections of the rat brain with the AP and NTS highlighted, with corresponding surgical coordinates (bottom). (B) Schematic detailing the specific metabolic effects of growth differentiation factor 15 (GDF15) and glucose sensing in the AP and NTS. In the AP, GDF15’s feeding and weight-lowering effects partly stem from the GDNF family receptor α-like (Gfral)-calcitonin gene-related peptide (CGRP) pathway. These effects may not be dependent on glucagon-like peptide-1 receptor (Glp1r) activation, despite the co-expression of Glp1r with Gfral in AP cells. GDF15-Gfral axis in the AP enhances glucose tolerance and insulin sensitivity and inhibits hepatic glucose production as well. Lastly, Gfral-expressing cells in the AP and leptin receptor (Lepr)-expressing cells in the NTS synergistically reduce feeding and weight. In the NTS, Glp1r and Lepr activation lower feeding and weight. With respect to glucose sensing, both the NTS and AP share a glucose transporter 1 (GLUT1) and pyruvate kinase-dependent glucose sensing mechanism that regulates peripheral glucose metabolism. Images adapted from BioRender. PBN, parabrachial nucleus; PDH, pyruvate dehydrogenase; DCA, dichloroacetate; HFD, high-fat diet.

Infusing GDF15 into the NTS has minimal effects on food intake and weight gain in rats after 23 to 24 hours of refeeding [45,66], while the same dose of GDF15 administered into the AP markedly decreases food intake and weight gain, highlighting a significant contrast in the response elicited by these two regions [45,66]. To validate cannula placement in the AP and NTS in rats, bromophenol blue injection was used [45,58]. Alternatively, radioactive tracer [3-³H] glucose may be infused into the NTS or AP to measure radioactivity in anatomical locations of interest [38,67].

To study the biological role of Gfral expressing neurons, Gfral-Cre mice were crossed with a Cre-inducible hM3DqTg background to express activating designer receptors exclusively activated by designer drugs (DREADD) in Gfral neurons, which are activated by clozapine N-oxide (CNO) [66]. CNO activates Gfral expressing neurons which are mostly in the AP but not the NTS of mice [66], consistent with the fact that Gfral is mainly expressed in the AP and not the NTS of rats [45]. More importantly, CNO activates Gfral expressing neurons to lower feeding and weight [66], strongly suggesting that AP Gfral expressing neurons mediate GDF15 action to lower feeding and weight. GDF15’s role in the central regulation of weight holds potential as a target for anti-obesity treatments but is complicated by its associated side effects, such as nausea and/or emesis [46,68,69].

Chemogenetic activation of Gfral-expressing neurons (mainly expressed in the AP), activates calcitonin gene related peptide (CGRP) neurons to induce nausea [66], while CGRP neurons in the parabrachial nucleus are sufficient to cause visceral malaise [70]. Conversely, silencing CGRP parabrachial neurons by injecting Cre-dependent AAV with doubled floxed tetanus toxin:green fluorescent protein (GFP) into Calca^Cre:GFP mice negates GDF15-induced nausea [66]. Together with the fact that activation of Gfral expressing neurons selectively in the AP induces nausea [63], these findings collectively suggest that GDF15-induced weight loss is linked to nausea, as also evidenced by increased kaolin consumption induced by GDF15 [68,69,71]. Thus, the appetite-suppressing effects of GDF15 through AP Gfral-expressing cells may involve triggering malaise via an AP-Gfral-CGRP pathway (Fig. 1B).

A subset of cells in the AP, but not the NTS, co-express Gfral with the Glp1r [63,72], which also regulates weight. Interestingly, the effectiveness of GDF15 in reducing food intake remains intact in Glp1r-null mice, while Glp1r agonist liraglutide remains effective in GDF15- and Gfral-null mice [73]. It would appear Glp1r agonism and GDF15 inhibit food intake and body weight through different mechanisms [74]. Of note, combining GDF15 with Glp1r agonists in normal rodents further reduces food intake and weight loss without increasing malaise in rats [74]. Despite the co-expression of Glp1r with Gfral in AP neurons, most Glp1r-expressing cells do not co-express Gfral [63,72], indicating that GDF15 and Glp1r agonists do not target identical neuronal populations in the AP. Together, this suggests that GDF15 and Glp1r act through distinct anorectic neural circuits (Fig. 1B).

GDF15 and leptin interact as leptin-deficient ob/ob mice and wild-type mice treated with leptin receptor antagonists are less responsive to GDF15 leading to diminished weight loss and adiposity reduction [75]. Co-administration of subcutaneous leptin and GDF15 amplified weight and adiposity reduction by reducing food intake compared to the GDF15 treatment alone [75]. Although leptin receptor and Gfral are not co-expressed on the same neurons, synaptic connections between synaptophysinleptin receptor and Gfral-expressing neurons are identified between the NTS and AP using immunofluorescence (Fig. 1B) [75]. The interconnection of GDF15 and leptin action in the NTS and/or AP warrants future investigation.

Glucose regulation

GDF15 infusion into the AP improves glucose tolerance and insulin sensitivity, leading to decreased hepatic glucose production [76]. These effects occur independently of any changes in food intake and weight, signifying a broader metabolic impact of GDF15 [76]. Subcutaneous administration of GDF15 in rats fed high-fat diets also exhibit decreased hepatic glucose production and increased insulin sensitivity in the liver, brown and white adipose tissue [54]. These changes occurred through increased β-adrenergic receptor signaling mediated by Gfral, and were independent of weight loss (Fig. 1B) [54]. In Gfral global knockouts and pharmacological inhibition of β-adrenergic receptors, the GDF15 insulin-sensitizing effect was nullified [54].

Knocking down Gfral, specifically in the AP, by infusing 2 μL of lentivirus harboring shRNA of Gfral over 15 minutes [45], disrupted glucose tolerance in the presence of GDF15 administration [76]. To verify gene knockdown specificity, reverse transcription polymerase chain reaction (qRT-PCR) assays were done to assess transcriptional levels of the targeted gene in cannulated and adjacent tissues [58,76]. If the infusion rate is too high, such as 3 μL of virus infused over 30 seconds [38], leakage into adjacent structures will result in non-specific knockdown of genes in the rest of the DVC when targeting only the AP [45]. Disrupted glucose tolerance in AP Gfral knockdown rats suggests that Gfral in the AP is essential for mediating the glucoregulatory and metabolic effects of GDF15.

Metformin-induced glucose, feeding, and weight regulation via brainstem GDF15 signaling

Metformin lowers plasma glucose, feeding and weight in rodents and humans [77-81]. Metformin was first associated with elevated plasma GDF15 levels in an epidemiological study [82] and was later confirmed in a randomized control trial, establishing a link between metformin-induced weight loss and elevated GDF15 levels [83]. The metformin-induced rise in plasma GDF15 results in reduced food intake, weight gain, and increased glucose tolerance and insulin sensitivity in high-fat fed rodents through activation of Gfral [76,83,84]. Metformin increases glucose tolerance and plasma GDF15 levels in high-fat fed rats within 2 hours of upper small intestinal infusion, independent of GDF15’s role in insulin secretion and weight changes [76]. However, AP-specific knockdown of Gfral expression negates the glucoregulatory and food intake-lowering effects of upper small intestinal metformin infusion, indicating that the GDF15-Gfral signaling axis is necessary for metformin to increase glucose tolerance and weight loss in rats fed a high-fat diet [76]. Contrasting reports highlight that metformin’s glucoregulatory effects may not always depend on GDF15. Coll et al. [83] found no significant glucose tolerance difference between GDF15-null and wildtype mice after a single metformin dose. However, Coll et al. [83] used a high dosage of metformin (300 mg/kg), potentially activating GDF15-independent pathways to promote glucose tolerance. This suggests a need for further research to elucidate the GDF15-Gfral signaling-dependent and independent pathways mediating metformin’s glucoregulatory actions. Collectively, these studies reveal that the weight-reducing and possible glucose-regulating effects of metformin are mediated through the GDF15-Gfral signaling pathway in the AP.

BRAIN GLUCOSE SENSING

Glucose sensing in the brain maintains whole-body glucose homeostasis [85], but site-specific mechanisms remain unclear. Glucose uptake in the brain [86,87] is facilitated by the expression of glucose transporter genes across multiple cell types: endothelial cells forming the blood-brain barrier (glucose transporter 1 [GLUT1]), astrocytes (GLUT1), and neurons (GLUT2/3) [88]. The extent and requirement for neuronal glucose metabolism remains unclear. The reliance on glucose as the sole energy source seems insufficient for maintaining energy stability, especially under high metabolic demand or limited oxygen availability. Supporting this idea, inhibiting glucose uptake in hippocampal neurons by deleting GLUT3 increases pyruvate-to-lactate conversion as a potential compensatory response [86]. The brain utilizes alternative fuels, such as lactate, produced through aerobic glycolysis by astrocytes and is referred to as the astrocyte-neuron lactate shuttle [89,90]. In response to excitatory neuronal activity, the uptake of glutamate by astrocytes via Na+-cotransport stimulates Na+-K+-ATPase activity, releasing lactate into the extracellular space. Neurons can uptake lactate through monocarboxylate transporters [91] and convert it to pyruvate by lactate dehydrogenase (LDH) isoform B for fuel [92].

Lactate infusion mirrors the hypothalamic glucose-induced lowering effect in rats and mice on plasma glucose and insulin levels as well as hepatic glucose production [14]. Blocking lactate-to-pyruvate conversion through LDH chemical inhibitor, oxamate, or lentiviral-mediated knockdown of astrocytic LDHA expression [93] impairs the glucose-lowering response of hypothalamic glucose/lactate [14,93]. These findings highlight the in vivo relevance of the astrocyte-neuron lactate shuttle hypothesis in hypothalamic regulation of glucose homeostasis. Not all areas of the brain preferentially utilize lactate; the cortex directly uptakes glucose via neurons during increased neuronal activity [87]. Nonetheless, impairments in glucose transporters are linked to disrupted central glucose sensing and metabolic disorders. In uncontrolled diabetic rats, sustained hyperglycemia impairs hypothalamic glucose sensing via a reduction in hypothalamic glial GLUT1 expression [94] while downstream lactate sensing remains intact [15]. Hypothalamic glucose, but not lactate sensing, fails to lower glucose production in high-fat fed rats as well [15,93], although the role of hypothalamic GLUT1 remains unclear.

To assess whether glucose sensing in the DVC regulates glucose homeostasis, rats underwent a pancreatic euglycemic clamp with glucose infused directly into the DVC [14]. Like hypothalamic glucose infusion [14], glucose infusion into the DVC of chow rats required more exogenous glucose to prevent hypoglycemia [67]. This glucose-lowering effect was due to a decrease in hepatic glucose production rather than an increase in glucose uptake [67].

To examine whether the astrocyte-neuronal lactate shuttle mediates glucose sensing in the DVC, like in the hypothalamus [14], lactate was first infused in the DVC to see whether it recapitulates the glucoregulatory effect of glucose. In contrast to the hypothalamus, DVC lactate infusion fails to alter the glucose infusion rate and glucose production during the pancreatic clamps [67]. Additionally, chemical inhibition of LDH in the DVC fails to disrupt the glucoregulatory effects of DVC glucose infusion, suggesting that lactate metabolism is neither sufficient and necessary for glucose sensing in the DVC to regulate hepatic glucose production. These findings highlight the DVC detects changes in glucose to regulate glucose homeostasis, but the glucose sensing mechanisms engaged in the DVC is different than the hypothalamus.

NTS and AP glucose sensing mechanism

Given that hypoglycemia induces cFOS (a proto-oncogene) expression in NTS and AP neurons [95,96], and AP lesions attenuate hyperphagic response to glucoprivation [97], both the NTS and/or AP within the DVC could sense glucose to regulate peripheral glucose metabolism. To investigate the specific contributions and mechanisms of glucose sensing in the NTS and AP, site-specific brain infusions were performed as described above for GDF15 studies. Bromophenol blue dye was infused through NTS or AP cannulas to confirm accurate delivery.

Direct glucose infusion into the NTS or AP (like DVC) lowers hepatic glucose production in chow-fed rats during clamps [58]. To investigate the role of GLUT1 (i.e., glucose uptake) as well as pyruvate kinase (i.e., pyruvate formation via glycolysis) within the NTS and/or AP for glucose sensing, site-specific, lentiviral-mediated knockdown of GLUT1 and pyruvate kinase was performed. This involved injecting 3 μL of lentivirus via bilateral cannulae over 10 minutes into the NTS or 2 μL of lentivirus over 15 minutes in the AP, respectively [58]. Knocking down GLUT1 or pyruvate kinase expression in either the NTS or AP negated the glucose-regulating effects of glucose infusion into these regions, preventing the reduction in hepatic glucose production [58]. These findings indicate that glucose uptake and subsequent metabolism to pyruvate are necessary for glucose sensing in the NTS and AP to regulate hepatic glucose production. Importantly, high-fat feeding impairs NTS and AP glucose sensing in association with a reduction in GLUT1 expression, though pyruvate kinase levels remained unaffected [58]. Interestingly, administration of dichloroacetate, an agonist of pyruvate dehydrogenase (that converts pyruvate to acetyl-coenzyme A), into the NTS or AP mimicked the glucose-lowering effect of glucose sensing in chow-fed rats [58]. However, dichloroacetate in both NTS and AP failed to reduce glucose production in rats fed a high-fat diet, suggesting that high-fat feeding not only disrupt GLUT1-mediated glucose uptake but also downstream pyruvate metabolism in the NTS and AP that affected glucose sensing (Fig. 1B) [58]. Taken together, both the NTS and AP rely on a GLUT1 and pyruvate kinase-dependent glucose sensing mechanism to decrease glucose production, thereby highlighting the critical role of the NTS and AP in glucose sensing for whole-body glucose metabolic regulation.

CONCLUSIONS

Although the hypothalamus is an established center that senses changes in circulating hormones and nutrients to maintain whole-body metabolic homeostasis, the DVC containing the NTS and AP has received much attention recently that impacts energy and glucose homeostasis as well. Herein, we highlight the AP and NTS-targeted techniques used to characterize GDF15 action and glucose sensing in these specific regions. Both the NTS and AP are essential for glucose sensing, which is dependent on both GLUT1 and pyruvate kinase expression. Differences in function between the NTS and AP arise in the context of GDF15. GDF15 seems to specifically exert its anorectic and glucoregulatory effects through its receptor Gfral in the AP. Future studies aim to advance mechanistic knowledge in understanding how hormones and nutrients activate respective signals in the NTS and/or AP to regulate weight and glucose levels could unveil therapeutic targets to lower weight and glucose levels in obesity and diabetes.

Notes

CONFLICTS OF INTEREST

No potential conflict of interest relevant to this article was reported.

Acknowledgements

Kyla Bruce is supported by the Banting and Best Diabetes Center Graduate Scholarship and Ontario Graduate Scholarship. Ameth N. Garrido is supported by the University of Toronto MD/PhD program. Song-Yang Zhang is supported by a Canadian Institutes of Health Research (CIHR) post-doctoral fellowship. Tony K.T. Lam is supported by CIHR grants (PJT 189957, PJT 183901, MRT-168045) and holds the Canada Research Chair (Tier 1) in Diabetes and Obesity at the Toronto General Hospital Research Institute & University of Toronto.

References

1. Myers MG Jr, Olson DP. Central nervous system control of metabolism. Nature 2012;491:357–63.

2. Woods SC. The control of food intake: behavioral versus molecular perspectives. Cell Metab 2009;9:489–98.

3. Deem JD, Tingley D, Bjerregaard AM, Secher A, Chan O, Uzo C, et al. Identification of hypothalamic glucoregulatory neurons that sense and respond to changes in glycemia. Diabetes 2023;72:1207–13.

4. Obici S, Zhang BB, Karkanias G, Rossetti L. Hypothalamic insulin signaling is required for inhibition of glucose production. Nat Med 2002;8:1376–82.

5. Pocai A, Lam TK, Gutierrez-Juarez R, Obici S, Schwartz GJ, Bryan J, et al. Hypothalamic K(ATP) channels control hepatic glucose production. Nature 2005;434:1026–31.

6. Clegg DJ, Riedy CA, Smith KA, Benoit SC, Woods SC. Differential sensitivity to central leptin and insulin in male and female rats. Diabetes 2003;52:682–7.

7. Brown LM, Clegg DJ, Benoit SC, Woods SC. Intraventricular insulin and leptin reduce food intake and body weight in C57BL/6J mice. Physiol Behav 2006;89:687–91.

8. Buettner C, Pocai A, Muse ED, Etgen AM, Myers MG Jr, Rossetti L. Critical role of STAT3 in leptin’s metabolic actions. Cell Metab 2006;4:49–60.

9. Morton GJ, Schwartz MW. Leptin and the central nervous system control of glucose metabolism. Physiol Rev 2011;91:389–411.

10. Mighiu PI, Yue JT, Filippi BM, Abraham MA, Chari M, Lam CK, et al. Hypothalamic glucagon signaling inhibits hepatic glucose production. Nat Med 2013;19:766–72.

11. Abraham MA, Yue JT, LaPierre MP, Rutter GA, Light PE, Filippi BM, et al. Hypothalamic glucagon signals through the KATP channels to regulate glucose production. Mol Metab 2013;3:202–8.

12. Sandoval DA, Bagnol D, Woods SC, D’Alessio DA, Seeley RJ. Arcuate glucagon-like peptide 1 receptors regulate glucose homeostasis but not food intake. Diabetes 2008;57:2046–54.

13. Taher J, Baker CL, Cuizon C, Masoudpour H, Zhang R, Farr S, et al. GLP-1 receptor agonism ameliorates hepatic VLDL overproduction and de novo lipogenesis in insulin resistance. Mol Metab 2014;3:823–33.

14. Lam TK, Gutierrez-Juarez R, Pocai A, Rossetti L. Regulation of blood glucose by hypothalamic pyruvate metabolism. Science 2005;309:943–7.

15. Chari M, Lam CK, Wang PY, Lam TK. Activation of central lactate metabolism lowers glucose production in uncontrolled diabetes and diet-induced insulin resistance. Diabetes 2008;57:836–40.

16. Parton LE, Ye CP, Coppari R, Enriori PJ, Choi B, Zhang CY, et al. Glucose sensing by POMC neurons regulates glucose homeostasis and is impaired in obesity. Nature 2007;449:228–32.

17. Pocai A, Lam TK, Obici S, Gutierrez-Juarez R, Muse ED, Arduini A, et al. Restoration of hypothalamic lipid sensing normalizes energy and glucose homeostasis in overfed rats. J Clin Invest 2006;116:1081–91.

18. Arrieta-Cruz I, Su Y, Knight CM, Lam TK, Gutierrez-Juarez R. Evidence for a role of proline and hypothalamic astrocytes in the regulation of glucose metabolism in rats. Diabetes 2013;62:1152–8.

19. Su Y, Lam TK, He W, Pocai A, Bryan J, Aguilar-Bryan L, et al. Hypothalamic leucine metabolism regulates liver glucose production. Diabetes 2012;61:85–93.

20. Blouet C, Schwartz GJ. Brainstem nutrient sensing in the nucleus of the solitary tract inhibits feeding. Cell Metab 2012;16:579–87.

21. Grill HJ, Hayes MR. Hindbrain neurons as an essential hub in the neuroanatomically distributed control of energy balance. Cell Metab 2012;16:296–309.

22. Cheng W, Gordian D, Ludwig MQ, Pers TH, Seeley RJ, Myers MG Jr. Hindbrain circuits in the control of eating behaviour and energy balance. Nat Metab 2022;4:826–35.

23. Wu Q, Lemus MB, Stark R, Bayliss JA, Reichenbach A, Lockie SH, et al. The temporal pattern of cfos activation in hypothalamic, cortical, and brainstem nuclei in response to fasting and refeeding in male mice. Endocrinology 2014;155:840–53.

24. Duca FA, Waise TM, Peppler WT, Lam TK. The metabolic impact of small intestinal nutrient sensing. Nat Commun 2021;12:903.

25. Waise TM, Dranse HJ, Lam TK. The metabolic role of vagal afferent innervation. Nat Rev Gastroenterol Hepatol 2018;15:625–36.

26. Yue JT, Abraham MA, Bauer PV, LaPierre MP, Wang P, Duca FA, et al. Inhibition of glycine transporter-1 in the dorsal vagal complex improves metabolic homeostasis in diabetes and obesity. Nat Commun 2016;7:13501.

27. Duca FA, Bauer PV, Hamr SC, Lam TK. Glucoregulatory relevance of small intestinal nutrient sensing in physiology, bariatric surgery, and pharmacology. Cell Metab 2015;22:367–80.

28. Yue JT, Abraham MA, LaPierre MP, Mighiu PI, Light PE, Filippi BM, et al. A fatty acid-dependent hypothalamic-DVC neurocircuitry that regulates hepatic secretion of triglyceride-rich lipoproteins. Nat Commun 2015;6:5970.

29. Lam CK, Chari M, Rutter GA, Lam TK. Hypothalamic nutrient sensing activates a forebrain-hindbrain neuronal circuit to regulate glucose production in vivo. Diabetes 2011;60:107–13.

30. Hackl MT, Furnsinn C, Schuh CM, Krssak M, Carli F, Guerra S, et al. Brain leptin reduces liver lipids by increasing hepatic triglyceride secretion and lowering lipogenesis. Nat Commun 2019;10:2717.

31. Benedict C, Kern W, Schultes B, Born J, Hallschmid M. Differential sensitivity of men and women to anorexigenic and memory-improving effects of intranasal insulin. J Clin Endocrinol Metab 2008;93:1339–44.

32. Frank-Podlech S, von Schnurbein J, Veit R, Heni M, Machann J, Heinze JM, et al. Leptin replacement reestablishes brain insulin action in the hypothalamus in congenital leptin deficiency. Diabetes Care 2018;41:907–10.

33. Dash S, Xiao C, Morgantini C, Koulajian K, Lewis GF. Intranasal insulin suppresses endogenous glucose production in humans compared with placebo in the presence of similar venous insulin concentrations. Diabetes 2015;64:766–74.

34. Heni M, Wagner R, Kullmann S, Gancheva S, Roden M, Peter A, et al. Hypothalamic and striatal insulin action suppresses endogenous glucose production and may stimulate glucose uptake during hyperinsulinemia in lean but not in overweight men. Diabetes 2017;66:1797–806.

35. Filippi BM, Bassiri A, Abraham MA, Duca FA, Yue JT, Lam TK. Insulin signals through the dorsal vagal complex to regulate energy balance. Diabetes 2014;63:892–9.

36. Filippi BM, Yang CS, Tang C, Lam TK. Insulin activates Erk1/2 signaling in the dorsal vagal complex to inhibit glucose production. Cell Metab 2012;16:500–10.

37. LaPierre MP, Abraham MA, Yue JT, Filippi BM, Lam TK. Glucagon signalling in the dorsal vagal complex is sufficient and necessary for high-protein feeding to regulate glucose homeostasis in vivo. EMBO Rep 2015;16:1299–307.

38. Lam CK, Chari M, Su BB, Cheung GW, Kokorovic A, Yang CS, et al. Activation of N-methyl-D-aspartate (NMDA) receptors in the dorsal vagal complex lowers glucose production. J Biol Chem 2010;285:21913–21.

39. Kanoski SE, Zhao S, Guarnieri DJ, DiLeone RJ, Yan J, De Jonghe BC, et al. Endogenous leptin receptor signaling in the medial nucleus tractus solitarius affects meal size and potentiates intestinal satiation signals. Am J Physiol Endocrinol Metab 2012;303:E496–503.

40. Hayes MR, Skibicka KP, Grill HJ. Caudal brainstem processing is sufficient for behavioral, sympathetic, and parasympathetic responses driven by peripheral and hindbrain glucagon-like-peptide-1 receptor stimulation. Endocrinology 2008;149:4059–68.

41. Hayes MR, Leichner TM, Zhao S, Lee GS, Chowansky A, Zimmer D, et al. Intracellular signals mediating the food intake-suppressive effects of hindbrain glucagon-like peptide-1 receptor activation. Cell Metab 2011;13:320–30.

42. Alhadeff AL, Mergler BD, Zimmer DJ, Turner CA, Reiner DJ, Schmidt HD, et al. Endogenous glucagon-like peptide-1 receptor signaling in the nucleus tractus solitarius is required for food intake control. Neuropsychopharmacology 2017;42:1471–9.

43. Alhadeff AL, Grill HJ. Hindbrain nucleus tractus solitarius glucagon-like peptide-1 receptor signaling reduces appetitive and motivational aspects of feeding. Am J Physiol Regul Integr Comp Physiol 2014;307:R465–70.

44. Bootcov MR, Bauskin AR, Valenzuela SM, Moore AG, Bansal M, He XY, et al. MIC-1, a novel macrophage inhibitory cytokine, is a divergent member of the TGF-beta superfamily. Proc Natl Acad Sci U S A 1997;94:11514–9.

45. Zhang SY, Bruce K, Danaei Z, Li RJ, Barros DR, Kuah R, et al. Metformin triggers a kidney GDF15-dependent area postrema axis to regulate food intake and body weight. Cell Metab 2023;35:875–86.

46. Patel S, Alvarez-Guaita A, Melvin A, Rimmington D, Dattilo A, Miedzybrodzka EL, et al. GDF15 provides an endocrine signal of nutritional stress in mice and humans. Cell Metab 2019;29:707–18.

47. Campderros L, Moure R, Cairo M, Gavalda-Navarro A, Quesada-Lopez T, Cereijo R, et al. Brown adipocytes secrete GDF15 in response to thermogenic activation. Obesity (Silver Spring) 2019;27:1606–16.

48. Mullican SE, Lin-Schmidt X, Chin CN, Chavez JA, Furman JL, Armstrong AA, et al. GFRAL is the receptor for GDF15 and the ligand promotes weight loss in mice and nonhuman primates. Nat Med 2017;23:1150–7.

49. Emmerson PJ, Wang F, Du Y, Liu Q, Pickard RT, Gonciarz MD, et al. The metabolic effects of GDF15 are mediated by the orphan receptor GFRAL. Nat Med 2017;23:1215–9.

50. Hsu JY, Crawley S, Chen M, Ayupova DA, Lindhout DA, Higbee J, et al. Non-homeostatic body weight regulation through a brainstem-restricted receptor for GDF15. Nature 2017;550:255–9.

51. Yang L, Chang CC, Sun Z, Madsen D, Zhu H, Padkjaer SB, et al. GFRAL is the receptor for GDF15 and is required for the anti-obesity effects of the ligand. Nat Med 2017;23:1158–66.

52. Suriben R, Chen M, Higbee J, Oeffinger J, Ventura R, Li B, et al. Antibody-mediated inhibition of GDF15-GFRAL activity reverses cancer cachexia in mice. Nat Med 2020;26:1264–70.

53. Wang D, Townsend LK, DesOrmeaux GJ, Frangos SM, Batchuluun B, Dumont L, et al. GDF15 promotes weight loss by enhancing energy expenditure in muscle. Nature 2023;619:143–50.

54. Sjoberg KA, Sigvardsen CM, Alvarado-Diaz A, Andersen NR, Larance M, Seeley RJ, et al. GDF15 increases insulin action in the liver and adipose tissue via a β-adrenergic receptor-mediated mechanism. Cell Metab 2023;35:1327–40.

55. Tsai VW, Manandhar R, Jorgensen SB, Lee-Ng KK, Zhang HP, Marquis CP, et al. The anorectic actions of the TGFβ cytokine MIC-1/GDF15 require an intact brainstem area postrema and nucleus of the solitary tract. PLoS One 2014;9e100370.

56. Borner T, Arnold M, Ruud J, Breit SN, Langhans W, Lutz TA, et al. Anorexia-cachexia syndrome in hepatoma tumour-bearing rats requires the area postrema but not vagal afferents and is paralleled by increased MIC-1/GDF15. J Cachexia Sarcopenia Muscle 2017;8:417–27.

57. Tsai VW, Zhang HP, Manandhar R, Lee-Ng KK, Lebhar H, Marquis CP, et al. Treatment with the TGF-b superfamily cytokine MIC-1/GDF15 reduces the adiposity and corrects the metabolic dysfunction of mice with diet-induced obesity. Int J Obes (Lond) 2018;42:561–71.

58. Li RJ, Chiu JF, Bruce K, Zhang SY, Barros DR, Yue JT, et al. A glucose-sensing mechanism with glucose transporter 1 and pyruvate kinase in the area postrema regulates hepatic glucose production in rats. J Biol Chem 2023;299:104633.

59. Cheng W, Gonzalez I, Pan W, Tsang AH, Adams J, Ndoka E, et al. Calcitonin receptor neurons in the mouse nucleus tractus solitarius control energy balance via the non-aversive suppression of feeding. Cell Metab 2020;31:301–12.

60. Cavanaugh AR, Schwartz GJ, Blouet C. High-fat feeding impairs nutrient sensing and gut brain integration in the caudomedial nucleus of the solitary tract in mice. PLoS One 2015;10e0118888.

61. Cheng W, Ndoka E, Hutch C, Roelofs K, MacKinnon A, Khoury B, et al. Leptin receptor-expressing nucleus tractus solitarius neurons suppress food intake independently of GLP1 in mice. JCI Insight 2020;5e134359.

62. Qiu W, Hutch CR, Wang Y, Wloszek J, Rucker RA, Myers MG, et al. Multiple NTS neuron populations cumulatively suppress food intake. Elife 2023;12e85640.

63. Zhang C, Kaye JA, Cai Z, Wang Y, Prescott SL, Liberles SD. Area postrema cell types that mediate nausea-associated behaviors. Neuron 2021;109:461–72.

64. Zhang C, Vincelette LK, Reimann F, Liberles SD. A brainstem circuit for nausea suppression. Cell Rep 2022;39:110953.

65. Fulton S, Horn CC, Zhang C. Characterizing a new tool to manipulate area postrema GLP1R+ neurons across species. Physiol Behav 2024;276:114474.

66. Sabatini PV, Frikke-Schmidt H, Arthurs J, Gordian D, Patel A, Rupp AC, et al. GFRAL-expressing neurons suppress food intake via aversive pathways. Proc Natl Acad Sci U S A 2021;118e2021357118.

67. Li RJ, Batchuluun B, Zhang SY, Abraham MA, Wang B, Lim YM, et al. Nutrient infusion in the dorsal vagal complex controls hepatic lipid and glucose metabolism in rats. iScience 2021;24:102366.

68. Borner T, Shaulson ED, Ghidewon MY, Barnett AB, Horn CC, Doyle RP, et al. GDF15 induces anorexia through nausea and emesis. Cell Metab 2020;31:351–62.

69. Worth AA, Shoop R, Tye K, Feetham CH, D’Agostino G, Dodd GT, et al. The cytokine GDF15 signals through a population of brainstem cholecystokinin neurons to mediate anorectic signalling. Elife 2020;9e55164.

70. Carter ME, Han S, Palmiter RD. Parabrachial calcitonin gene-related peptide neurons mediate conditioned taste aversion. J Neurosci 2015;35:4582–6.

71. Borner T, Workinger JL, Tinsley IC, Fortin SM, Stein LM, Chepurny OG, et al. Corrination of a GLP-1 receptor agonist for glycemic control without emesis. Cell Rep 2020;31:107768.

72. Ludwig MQ, Cheng W, Gordian D, Lee J, Paulsen SJ, Hansen SN, et al. A genetic map of the mouse dorsal vagal complex and its role in obesity. Nat Metab 2021;3:530–45.

73. Frikke-Schmidt H, Hultman K, Galaske JW, Jorgensen SB, Myers MG Jr, Seeley RJ. GDF15 acts synergistically with liraglutide but is not necessary for the weight loss induced by bariatric surgery in mice. Mol Metab 2019;21:13–21.

74. Ghidewon M, Wald HS, McKnight AD, De Jonghe BC, Breen DM, Alhadeff AL, et al. Growth differentiation factor 15 (GDF15) and semaglutide inhibit food intake and body weight through largely distinct, additive mechanisms. Diabetes Obes Metab 2022;24:1010–20.

75. Breit SN, Manandhar R, Zhang HP, Lee-Ng M, Brown DA, Tsai VW. GDF15 enhances body weight and adiposity reduction in obese mice by leveraging the leptin pathway. Cell Metab 2023;35:1341–55.

76. Zhang SY, Danaei Z, Bruce K, Chiu JF, Lam TK. Acute activation of GFRAL in the area postrema contributes to glucose regulation independent of weight. Diabetes 2024;73:426–33.

77. Diabetes Prevention Program Research Group. The 10-year cost-effectiveness of lifestyle intervention or metformin for diabetes prevention: an intent-to-treat analysis of the DPP/DPPOS. Diabetes Care 2012;35:723–30.

78. Lee A, Morley JE. Metformin decreases food consumption and induces weight loss in subjects with obesity with type II non-insulin-dependent diabetes. Obes Res 1998;6:47–53.

79. Knowler WC, Barrett-Connor E, Fowler SE, Hamman RF, Lachin JM, Walker EA, et al. Reduction in the incidence of type 2 diabetes with lifestyle intervention or metformin. N Engl J Med 2002;346:393–403.

80. Lachin JM, Christophi CA, Edelstein SL, Ehrmann DA, Hamman RF, Kahn SE, et al. Factors associated with diabetes onset during metformin versus placebo therapy in the diabetes prevention program. Diabetes 2007;56:1153–9.

81. Diabetes Prevention Program Research Group, Knowler WC, Fowler SE, Hamman RF, Christophi CA, Hoffman HJ, et al. 10-Year follow-up of diabetes incidence and weight loss in the Diabetes Prevention Program Outcomes Study. Lancet 2009;374:1677–86.

82. Gerstein HC, Pare G, Hess S, Ford RJ, Sjaarda J, Raman K, et al. Growth differentiation factor 15 as a novel biomarker for metformin. Diabetes Care 2017;40:280–3.

83. Coll AP, Chen M, Taskar P, Rimmington D, Patel S, Tadross JA, et al. GDF15 mediates the effects of metformin on body weight and energy balance. Nature 2020;578:444–8.

84. Day EA, Ford RJ, Smith BK, Mohammadi-Shemirani P, Morrow MR, Gutgesell RM, et al. Metformin-induced increases in GDF15 are important for suppressing appetite and promoting weight loss. Nat Metab 2019;1:1202–8.

85. Mirzadeh Z, Faber CL, Schwartz MW. Central nervous system control of glucose homeostasis: a therapeutic target for type 2 diabetes? Annu Rev Pharmacol Toxicol 2022;62:55–84.

86. Li H, Guglielmetti C, Sei YJ, Zilberter M, Le Page LM, Shields L, et al. Neurons require glucose uptake and glycolysis in vivo. Cell Rep 2023;42:112335.

87. Lundgaard I, Li B, Xie L, Kang H, Sanggaard S, Haswell JD, et al. Direct neuronal glucose uptake heralds activity-dependent increases in cerebral metabolism. Nat Commun 2015;6:6807.

88. Maher F. Immunolocalization of GLUT1 and GLUT3 glucose transporters in primary cultured neurons and glia. J Neurosci Res 1995;42:459–69.

89. Pellerin L, Magistretti PJ. Glutamate uptake into astrocytes stimulates aerobic glycolysis: a mechanism coupling neuronal activity to glucose utilization. Proc Natl Acad Sci U S A 1994;91:10625–9.

90. Magistretti PJ, Pellerin L. Cellular bases of brain energy metabolism and their relevance to functional brain imaging: evidence for a prominent role of astrocytes. Cereb Cortex 1996;6:50–61.

91. Bergersen LH. Is lactate food for neurons?: comparison of monocarboxylate transporter subtypes in brain and muscle. Neuroscience 2007;145:11–9.

92. Brooks GA, Dubouchaud H, Brown M, Sicurello JP, Butz CE. Role of mitochondrial lactate dehydrogenase and lactate oxidation in the intracellular lactate shuttle. Proc Natl Acad Sci U S A 1999;96:1129–34.

93. Abraham MA, Rasti M, Bauer PV, Lam TK. Leptin enhances hypothalamic lactate dehydrogenase A (LDHA)-dependent glucose sensing to lower glucose production in high-fat-fed rats. J Biol Chem 2018;293:4159–66.

94. Chari M, Yang CS, Lam CK, Lee K, Mighiu P, Kokorovic A, et al. Glucose transporter-1 in the hypothalamic glial cells mediates glucose sensing to regulate glucose production in vivo. Diabetes 2011;60:1901–6.

95. Bohland M, Matveyenko AV, Saberi M, Khan AM, Watts AG, Donovan CM. Activation of hindbrain neurons is mediated by portal-mesenteric vein glucosensors during slow-onset hypoglycemia. Diabetes 2014;63:2866–75.

96. Ritter S, Dinh TT. 2-Mercaptoacetate and 2-deoxy-D-glucose induce Fos-like immunoreactivity in rat brain. Brain Res 1994;641:111–20.

97. Ozawa Y, Arima H, Banno R, Ito Y, Goto M, Morishita Y, et al. Lesion of area postrema attenuated hyperphagic responses to glucoprivation, but not transcriptional activation of the neuropeptide Y gene in rats. Neuroreport 2012;23:673–5.

Article information Continued

This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.