Neural Circuits between Nodose Ganglion and Pulmonary Neuroendocrine Cells Regulate Lung Inflammatory Responses
Jie Chen, Shitao Xie, Zhekai Lin, Caiqi Zhao, Rujia Tao, Yingying Ma, Xiaoyan Chen, Renlan Wu, Qingjian Han, Pengfei Sui, Sheng Wang, Hongbin Ji, Hai Song, Xiaoming Zhang, Yangang Sun, Yuanlin Song, Xiao Su
TL;DR
The study reveals a neural circuit in the lungs that detects harmful substances and amplifies inflammation through interactions between nerve cells and specialized lung cells.
Contribution
The discovery of a feed-forward neural-epithelial circuit involving TRPA1+αCGRP+ sensory neurons and PNECs in regulating lung inflammation.
Findings
Vagal sensory endings synapse with PNECs and detect bacterial endotoxins via TRPA1.
TRPA1 detection triggers αCGRP production in nodose ganglia, which stimulates PNEC activation and proliferation.
This neural circuit amplifies endotoxin-induced lung inflammation through a feed-forward loop.
Abstract
The lungs interface directly with the external environment, exposing them to airborne pathogens like endotoxins. We investigated whether the vagus nerve, which innervates the lungs‐detects such pathogens. Using transcriptomics, tissue clearance imaging, electrophysiology, and cell‐specific knockout models, we discovered that vagal sensory endings synapse with pulmonary neuroendocrine cells (PNECs). These nerve endings detect bacterial endotoxins primarily through the pain receptor TRPA1, not via Toll‐like receptor 4 (TLR4). This detection triggers electrical excitation in vagal neurons and upregulates neuropeptide (e.g., αCGRP) production in the nodose ganglia. Released αCGRP then acts back on PNECs, stimulating their neuropeptide synthesis and proliferation. This creates a feed‐forward loop that amplifies endotoxin‐induced lung inflammation. Our findings reveal a critical neural…
Genes, proteins, chemicals, diseases, species, mutations and cell lines named across the full text — each resolved to its canonical identifier and authoritative record.
Click any figure to enlarge with its caption.
FIGURE 1
FIGURE 2
FIGURE 3
FIGURE 4
FIGURE 5
FIGURE 6
FIGURE 7- —NSFC10.13039/501100001809
- —Science and Technology Commission of Shanghai Municipality10.13039/501100003399
- —China Postdoctoral Science Foundation10.13039/501100002858
- —Key Technologies Research and Development Program10.13039/501100012165
Peer Reviews
No public reviews on file for this paper yet. If you reviewed it on a platform where reviews are public (OpenReview, ICLR, NeurIPS, ICML), you can paste yours below so the community can read it here.
Videos
No videos yet. Explain this paper in a talk, walkthrough, or lecture? Add one.
Taxonomy
TopicsVagus Nerve Stimulation Research · Respiratory and Cough-Related Research · Neuroscience of respiration and sleep
Introduction
1
The jugular‐nodose ganglia contain the sensory peripheral neurons of the vagus nerve, connecting visceral organs (i.e., lungs) to the medulla oblongata [1, 2, 3, 4], and transmitting peripheral pathogenic information to the brain [5, 6]. In brain regions, caudal nucleus of the solitary tract (cNST) [7] and the paraventricular hypothalamic nucleus (PVN) [8] are activated to modulate the inflammatory response. The vagal modulation mechanism of inflammation in the lungs is referred to as the parasympathetic inflammatory reflex [6, 9]. Regarding the efferent arm of this reflex, we have previously reported that the vagal‐α7nAChR (nicotinic acetylcholine receptor) signaling pathway plays a role in regulating lung injury, viral replication, and stem cell repair [10, 11, 12, 13]. However, the afferent arm of this reflex has not been thoroughly studied.
As we know, the vagal sensory neurons form the jugular (Prdm12 ^+^) and nodose (Phox2b ^+^) ganglia residing in the same nerve sheath [1]. The nodose ganglia innervate the distal airways of lung, while the jugular ganglia innervate the proximal airway [14]. Although TLR4 has been reported to detect lipopolysaccharide (LPS) exposed in lung TRPV1^+^ sensory neurons [8], it remains unclear whether lung‐innervated nodose sensory neurons express TLR4, which contributes to LPS perception in the lungs. In addition, LPS can activate suprarenal and celiac ganglia neurons and upregulate NPY expression by which affects the proliferation and activation of splenic lymphocytes [15]. Similarly, we need to understand whether vagal sensory neurons can respond to lung LPS and regulate lung inflammation by increasing the expression of their neuropeptides (e.g., CGRP).
The lungs are an organ that is open to the outside environment. The air inhaled contains various pathogens (e.g., endotoxins). Neuro‐epithelial crosstalk is crucial for sensing and regulating pulmonary inflammation. In the respiratory tract, sensory neurons serve as both first‐order and second‐order sensory neurons. They directly sense LPS from Gram‐negative bacteria using TRPA1, TRPV1, and TLR4, mediating acute neurogenic inflammation, pain, and sickness behavior [8, 16, 17]. Additionally, they act as second sensory neurons along with respiratory epithelial cells such as pulmonary neuroendocrine cells (PNECs), brush cells, solitary chemoreceptor cells, and taste bud cells, collectively identifying pathogens [1, 18, 19, 20, 21, 22]. These epithelial cells sense airway signals and transmit them to sensory neurons. PNECs, in particular, participate in lung recognition processes, including responses to hypoxia, hypercarbia, mechanical stretch, allergens, water, and extracellular traps [22, 23, 24, 25]. It has been reported that PNECs can produce CGRP and γ‐Aminobutyric acid (GABA) to regulate pulmonary immune responses [20, 24, 26]. Many studies have shown that PNECs receive projections from various vagal sensory neurons, including P2ry1 and Pvalb [1, 27, 28]. However, it remains unclear whether and how vagal sensory neurons (i.e., nodose αCGRP^+^ neurons) feedforward regulate CGRP production in PNECs and trigger lung inflammation.
The preganglionic synapse of GABAergic neurons is finely regulated by GABA receptor signaling [29]. In this study, we hypothesize that αCGRP^+^ nodose sensory neurons couple with PNECs to form a synapse. In response to the pulmonary LPS challenge, PNECs are activated to produce CGRP and GABA. PNEC‐derived CGRP can amplify lung inflammation by increasing the production of proinflammatory cytokine and neutrophil infiltration, and GABA can modulate preganglionic GABA receptors in αCGRP^+^ nodose sensory neurons to limit the release of αCGRP. Through this feedback loop, CGRP production in PNECs will be fine‐tuned. Overall, we sought to elucidate the neural circuits between the nodose ganglia and the PNECs, and explore their role in regulating the inflammatory response.
To address these issues, we employed transcriptome analysis, tissue clearance imaging, electroexcitability recording, gene and cell‐specific knockout, and other advanced methods. We found that pulmonary LPS challenge increased vagal excitability and CGRP expression in nodose sensory neurons via TRPA1 rather than TLR4. Deletion of Myd88 in Phox2b ^+^ nodose sensory neurons reduced LPS‐triggered vagal excitability and lung inflammation. A large number of nodose sensory neurons were TRPA1^+^αCGRP^+^ neurons, which innervated PNECs and alveoli. Deletion of Trpa1 or Calca in the vagal ganglia reduced CGRP expression in PNECs, the proliferation of PNECs, and lung inflammation. Ablation of PNECs reduced LPS‐induced lung inflammation, and lung GABA feedback modulated CGRP expression in nodose sensory neurons. Thus, we have established a nodose ganglion‐pulmonary neuroepithelial circuit that regulates the inflammatory response in the lungs.
Results
2
The Sensory Neurons in Nodose Ganglion via MYD88 Sense Intratracheal LPS and Promote Lung Inflammation
2.1
The majority of sensory neurons that project to the lungs originate from the nodose ganglia (NG) and the dorsal root ganglia (DRG). To distinguish between these two neuronal populations, we utilized *Phox2b^Cre^
- mice, which selectively label NG neurons. By crossing *Phox2b^Cre^
- mice with Cre‐dependent *Rosa26^tdTomato^
- reporter mice, we observed that *Phox2b^tdTomato+^
- signals were specifically expressed in the NG, with no detectable expression in the DRG (Figure S1A,B). To trace NG neurons that project to the lungs and respond to LPS stimulation, we injected Fast Blue—a fluorescent retrograde tracer—into the lungs (Figure S1A). A substantial number of Fast Blue^+^ signals were detected in Phox2b^tdTomato+^ NG neurons (Figure S1A), whereas only a small number were observed in the T2 DRG (Figure S1B). We also utilized optogenetic reporter mice by crossing *Phox2b^Cre^
- mice with Cre‐dependent ChR2‐EYFP reporter mice. Approximately 80% of the Fast Blue^+^ neurons projecting to the lungs originate from the NG. About 6% of the neurons in the NG project to the lungs and are responsible for regulating various physiological and pathological processes in the lung (Figure S1C). These data further elucidate the differences in pulmonary neural projections between NG and JG, with NG providing the majority of sensory innervation to the lungs.
Vagal sensory neurons are capable of detecting pathogens, with significant research focused on LPS from Gram‐negative bacteria, a model commonly used to induce acute lung injury. Previous studies have shown that intraperitoneal injection of LPS can be recognized by the vagus nerve, leading to the generation of specific neural electrical signals [30]. However, it remains unclear whether the nerve endings of vagal sensory neurons can directly sense LPS in the lungs. To investigate whether lung LPS stimulation can trigger a rapid vagal nerve response, we developed a specialized method for delivering LPS directly to the lungs, thereby bypassing potential interference from the upper respiratory tract. After administering either PBS or LPS, we placed electrodes on the cervical vagus nerve to record changes in its excitability (Figure 1A). We observed that both LPS and PBS stimulation resulted in an increase in vagal firing rates. The vagal excitation induced by PBS stimulation diminished within 1 min, while LPS stimulation triggered more intense and sustained nerve firing. Further analysis revealed that both the normalized vagal compound action potentials and the area under the curve were significantly increased in the LPS‐challenged group compared to the PBS‐challenged group (Figure 1B,C).
The sensory neurons in nodose ganglion via MYD88 sense intratracheal LPS and promote lung inflammation. (A,B) In anesthetized wild‐type mice, lung LPS stimulation was performed via endotracheal intubation, while recording the electrical activity of the cervical vagus nerve. Vagal responses to stimuli within the lung, including baseline (PBS, gray traces) and LPS (5 mg/kg, red traces) (n = 5–6 mice/group). (C) Quantification of peak responses to LPS (n = 5–6 mice/group). (D) Vagal responses to LPS stimuli within the lung in Myd88Phox2b mice and Myd88fl/fl mice. (n = 3–4 mice/group). (E) Vagal responses to baseline (PBS) and LPS (5mg/kg), including whether nodose neurons are Myd88 knockout (n = 3–4 mice/group). (F) Quantification of peak responses to LPS in Myd88Phox2b mice and Myd88fl/fl mice (n = 3–4 mice/group). (G) LPS induced a lower level of TNF‐α in Myd88Phox2b mice compared with Myd88fl/fl mice in BALF (n = 3–6 mice/group). (H,I) LPS induced a lower level of TNFα in F4/80+ macrophages in the BALF of Myd88Phox2b mice compared with Myd88fl/fl mice (n = 3 mice/group). (J) Injection of AAV‐Myd88 into the vagal ganglion of Myd88Phox2b mice restored Myd88 expression. Following LPS challenge, the TNF‐α level in the bronchoalveolar lavage fluid (BALF) of Myd88Phox2b mice with Myd88 reconstitution was significantly higher than that of the control group (n = 8 mice/group). Student's t‐test in (C,J). One‐way ANOVA in (F,G,I). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.
MYD88 is a key adaptor molecule in the Toll‐like receptor signaling pathway and plays a critical role in pathogen recognition. It is broadly expressed in sensory neurons of the NG [3]. To specifically delete Myd88 in NG sensory neurons, we crossed *Myd88^fl/fl^
- mice with *Phox2b^Cre^
- mice, generating *Myd88^Phox2b^
- mice (Figure 1D). By recording vagal nerve excitability, we found that the peak of LPS‐induced compound action potentials observed in *Myd88^Phox2b^
- mice was abolished after Myd88 gene deletion (Figure 1E). Furthermore, in *Myd88^Phox2b^
- mice challenged with LPS, the area under the compound action potential curve was significantly reduced compared to the *Myd88^fl/fl^
- control group challenged with LPS (Figure 1F). These findings suggest that sensory neurons in the NG detect pulmonary LPS through MYD88‐dependent signaling, thereby initiating vagal nerve activation.
To investigate the role of sensory neurons in LPS‐induced lung inflammation, we established a 1‐h pulmonary LPS challenge model in mice (Figure S2A). One hour after LPS administration, we observed a significant increase in TNF‐α levels in bronchoalveolar lavage fluid (BALF) (Figure S2B), elevated TNF‐α expression in F4/80^+^ cells (Figure S2C–E), and pronounced infiltration of polymorphonuclear neutrophils (PMNs) in the lung tissue (Figure S2F–H). To further examine the regulatory role of MYD88‐dependent vagal nerve activation in this model, we compared LPS‐challenged *MyD88^fl/fl^
- mice and *MyD88^Phox2b^
- mice. *MyD88^Phox2b^
- mice exhibited significantly reduced TNF‐α levels in BALF and decreased TNF‐α expression in F4/80^+^ macrophages (Figure 1G–I). Additionally, the expression levels of Il6, Cxcl1, and Ccl22 were significantly lower in LPS‐challenged *MyD88^Phox2b^
- mice than in the control group (Figure S3a,b). These findings indicate that sensory neurons in the NG detect pulmonary LPS via MYD88 signaling and play a crucial role in initiating the lung inflammatory response.
Next, we investigated whether the altered pulmonary inflammatory response observed in *Myd88^Phox2b^
- conditional knockout mice was due to potential defects in neural development. We first assessed pulmonary sensory innervation by examining the distribution of CGRP‐positive sensory nerve fibers, as CGRP‐expressing sensory neurons are a key subtype involved in immunomodulation. Our results showed that Myd88 conditional knockout did not affect the pattern of CGRP‐positive fiber projections in the lung, including those targeting alveolar regions and pulmonary neuroendocrine cells (PNECs) (Figure S3C). Furthermore, we performed a functional rescue experiment by selectively delivering AAV‐Myd88 into the nodose ganglion of *Myd88^Phox2b^
- mice. Restoring MYD88 expression in the nodose ganglion significantly enhanced the pulmonary inflammatory response to LPS compared to control mice (Figure 1J). These findings indicate that MYD88 in the nodose ganglion plays a critical role in sensing LPS and regulating lung inflammation.
Pulmonary LPS‐Induced Vagal Responses Rely on TRPA1+ Rather Than TLR4+ Nodose Sensory Neurons
2.2
The TLR4 signaling pathway is a well‐established mechanism for the recognition of LPS. To determine whether NG sensory neurons directly sense pulmonary LPS via TLR4, we assessed vagal nerve excitability in TLR4 knockout (*TLR4^KO^ *) mice and wild‐type controls following pulmonary LPS challenge. Although TNF‐α levels in bronchoalveolar lavage fluid (BALF) were significantly reduced in LPS‐treated *TLR4^KO^
- mice compared to PBS‐treated controls (Figure S4A), there were no significant differences in vagal compound action potentials (CAPs) or firing rates between the two groups following LPS stimulation (Figure S4B–D). Furthermore, we established an in vivo calcium imaging platform to monitor LPS‐induced neural activity in nodose ganglion neurons. AAV9‐CMV‐GCaMP6s was injected into the nodose ganglia, enabling GCaMP6s expression in all sensory neurons. Using a miniscope imaging system, we recorded real‐time vagal responses to pulmonary LPS stimulation. Remarkably, LPS‐induced calcium signals in nodose sensory neurons persisted in *TLR4^KO^
- mice (Figure S4E,F). Collectively, these findings suggest that pulmonary LPS‐induced vagal responses do not rely on TLR4‐expressing sensory neurons in the nodose ganglion.
In addition to TLR4, it has been reported that LPS can directly activate TRPA1 and TRPV1 expressed in neurons [16, 17]. By reanalyzing the single‐cell RNA sequencing datasets (GSE192987) from Chang's group [31], we found that the Trpa1 and Myd88 genes are highly expressed in *Prdm12^+^
- jugular and *Phox2b^+^
- nodose sensory neurons (Figure S4G,H). However, *Tlr4^+^
- sensory neurons account for 10.6% of the jugular ganglia and only 1.8% of the nodose ganglia (Figure S4I). Notably, 53% of the *Trpa1^+^
- lung‐innervating neurons project to the lungs, which is significantly higher than the 1.8% of *Tlr4^+^
- neurons (Figure S4J). *Trpa1^+^
- lung‐innervating neurons express lower levels of Tlr4, suggesting that TRPA1 is more likely to serve as a sensor for LPS in the pulmonary environment. Therefore, we examined LPS‐induced calcium signaling in nodose sensory neurons from *Trpa1^KO^
- mice. The results showed that LPS‐induced calcium responses were significantly attenuated in Trpa1‐deficient mice (Figure S4E,F).
The single‐cell RNA sequencing dataset (GSE192987) labels vagal sensory neurons projecting to the lungs using QZ2 [31]. In QZ2‐labeled nodose ganglion sensory neurons projecting to the lungs, Myd88, a key molecule involved in LPS sensing, was widely expressed. Furthermore, Myd88 co‐localized with Trpa1, which is also considered a molecular sensor for LPS (Figure S5A). We then investigated whether TRPA1 activation enhances MyD88 oligomerization to facilitate signal transduction. We transfected isolated neurons with a Myd88‐EGFP plasmid and observed a pronounced aggregation effect of MYD88‐EGFP, characterized by spontaneous formation of MYD88 oligomers (Figure S5B). We hypothesized that this phenomenon might be related to the high expression of Trpa1 in vagal sensory neurons, which promotes MyD88 self‐oligomerization. To test this hypothesis, we selected the Neuro‐2a neuronal cell line, which does not express Trpa1, for further study (Figure S5C). We first generated a Neuro‐2a cell line stably expressing MyD88‐GFP (Neuro‐2a‐MyD88‐GFP) (Figure S5D), and then overexpressed TRPA1 in these cells. Unlike vagal sensory neurons, Neuro‐2a cells did not show obvious spontaneous MYD88 aggregation. However, overexpression of TRPA1 increased the basal level of MYD88 aggregation. Upon LPS stimulation, both the number and size of MyD88‐GFP puncta were significantly increased in Neuro‐2a‐MyD88‐GFP cells overexpressing TRPA1 (Figure S5E,F).
We assessed the in vivo role of TRPA1 in vagal signaling induced by pulmonary LPS by selectively knocking out TRPA1 in the NG. We injected AAV‐CMV‐Cas9 and AAV‐sgTrpa1 adenoviruses targeting TRPA1 expression into the right jugular/nodose complex of mice, successfully knocking out Trpa1 expression within the ganglion (Figure 2A). In *NG^Trpa1KO^
- mice, the vagal response to pulmonary LPS stimulation was significantly diminished (Figure 2B,C). Regarding the regulation of pulmonary inflammation, *NG^Trpa1KO^
- mice exhibited significantly reduced TNF‐α expression in the bronchoalveolar lavage fluid (BALF) compared to controls (Figure 2D). Additionally, the TNF expression in F4/80 cells from the BALF was also notably decreased (Figure 2E,F). Collectively, these results suggest that vagal sensory neurons can utilize TRPA1 to sense and respond to pulmonary LPS, thereby contributing to the promotion of pulmonary inflammation in the physiological environment of the lungs.
*αCGRP+Trpa1+ nodose sensory neurons projecting to the lungs respond to pulmonary LPS stimulation. (A) Injection of AAV‐sgTrpa1 into the right nodose ganglion to generate NGTrpa1KO mice. (B,C) Vagal responses to LPS stimuli within the lung in NGTrpa1KO mice and Ctrl mice, including PBS for Ctrl mice (gray traces), LPS for Ctrl mice (red traces), and LPS for Trpa1KO mice (blue traces) (n = 5–7 mice/group). (m) Quantification of average responses to LPS in NGTrpa1KO mice and Ctrl mice (n = 3 mice/group). (D) LPS induced a lower level of TNF‐α in NGTrpa1KO mice compared with Ctrl mice in BALF (n = 4–7 mice/group). (E and F) LPS induced a lower level of TNF‐α in F4/80+ macrophages in the BALF of NGTrpa1KO mice compared with Ctrl mice (n = 4–7 mice/group). (G) Schematic diagram of RNA‐seq analysis using isolated vagal ganglia (jugular‐nodose complex) following pulmonary LPS administration. (H) RNA‐seq volcano plot of vagal ganglion treated or not treated with LPS (5 mg/kg) for 1 h. The neurotransmitter‐related and immune‐related genes are labeled. (n = 4 mice/group). (I) Analysis of CGRP gene expression in vagal ganglion upon LPS treatment via RNA‐seq. (n = 4 mice/group). (J) LPS induced a larger increase of CGRP levels in plasma (n = 8–16 mice/group). (K) Flow cytometric analysis of CGRP expression intensity in the vagal ganglion. (L) Using Fast Blue retrograde tracing in Phox2bChR2‐EYFP optogenetic reporter mice, we determined the proportion of CGRP‐positive neurons among nodose ganglion sensory neurons projecting to the lung. (M) The proportion of TRPA1‐positive sensory neurons among vagal αCGRP‐positive sensory neurons projecting to the lung in CalcatdTomato mice. (N) The expression of Calca and Trpa1 genes in QZ2+ neurons, which are vagal neurons projecting to the lungs [31]. (O) The distribution of vagal Trpa1/Calca double‐positive neurons, Trpa1‐positive neurons, and Calca‐positive neurons projecting to the lungs and other locations. (P) The expression levels of Trpa1 and Calca in neurons projecting to the lungs and other locations. (Q) The expression levels of Trpa1 and Calca in Trpa1/Calca double‐positive neurons, Trpa1
- neurons, and Calca
- neurons that project to the lungs. (R) CGRP mean fluorescence intensity in the nodose ganglion of NGTrpa1KO mice and Ctrl mice. (n = 4–7 mice/group). Scale bars: 200 µm in (R), 100µm in (A). Student's t‐test in (I,J,P,Q). One‐way ANOVA in (C,D,F,R). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.*
NG Sensory Neurons Upregulate αCGRP Expression in Response to Lung LPS Challenge
2.3
During the development of acute lung injury (ALI) and acute respiratory distress syndrome (ARDS), vagal sensory neurons rapidly transition into a nerve injury–like state, resulting in altered vagal excitability [32]. To investigate how early neural sensing modulates immune responses, we performed bulk RNA sequencing analysis on vagal ganglia isolated 1 h after pulmonary LPS challenge (Figure 2G; Figure S6A). A total of 886 differentially expressed genes were identified, with 463 genes significantly upregulated and 423 genes significantly downregulated (Figure S6B). These differentially expressed genes were visualized in a heatmap, and Gene Ontology (GO) biological process (GO‐BP) enrichment analysis was conducted (Figure S6C,D). The enriched pathways included regulation of nervous system processes, detection of external stimuli, regulation of sensory perception, detection of temperature stimuli, and sensory perception of pain (Figure S6D). These pathways suggest that vagal sensory neurons can recognize and respond to pulmonary LPS challenge. Furthermore, the differentially expressed genes were also significantly enriched in pathways related to the regulation of epithelial cell proliferation (Figure S6D), indicating that vagal sensory neurons may exert neuro‐immune regulatory effects by modulating lung epithelial cells in response to pulmonary LPS stimulation.
Through volcano plot analysis, we identified that the neuropeptide genes Calca (encoding α‐CGRP) and Calcb (encoding β‐CGRP) were among the most significantly upregulated genes in the vagal ganglia of LPS‐challenged mice (Figure 2H,I). Consistent with this, plasma CGRP levels in LPS‐challenged mice were also significantly elevated compared to the control group (Figure 2J). Flow cytometry and tissue clearing imaging analyses further revealed that CGRP expression was higher in the vagal sensory neurons of the LPS‐challenged group (Figure 2K). To further validate the effect of LPS on CGRP expression in the vagal ganglion, we performed serial sectioning of the vagal ganglia followed by immunofluorescence staining for CGRP. Consistent with flow cytometry results, LPS treatment significantly enhanced the fluorescence intensity of CGRP in the vagal ganglia (Figure S7A). We also used Fast Blue to label sensory neurons projecting to the lungs in order to determine whether the LPS‐induced upregulation of CGRP expression in the vagal ganglia is restricted to CGRP^+^ neurons that innervate the lungs. The results showed that CGRP expression was not limited to lung‐projecting CGRP^+^ neurons, but was also observed in other CGRP^+^ neurons that do not project to the lungs (Figure S7B). We speculate that this phenomenon may be attributed to inter‐neuronal signaling within the ganglia or to reflexive activation mediated by central nervous system nuclei.
Given that Calca expression is higher than Calcb, and the number of Calca ^+^ vagal sensory neurons projecting to the lungs is significantly greater than that of *Calcb^+^
- neurons, we focused our analysis on α‐CGRP‐positive sensory neurons (Figure S4G,H). Additionally, the differentially expressed gene Calca was enriched in GO biological process (GO‐BP) pathways, including detection of temperature stimuli, feeding behavior, sensory perception of pain, detection of abiotic stimuli, and detection of external stimuli, with four of these pathways specifically related to stimulus detection (Figure S6D).
Through GO analysis, we identified that differentially expressed genes related to “response to stimulus,” “nervous system,” “axon,” and “synapse” were significantly enriched (Figure S8A,B). We further examined synaptic pathways and found notable changes in genes associated with “regulation of dopamine secretion,” “dopamine receptor signaling,” “sensory perception,” “response to stimulus,” “GABAergic synapses,” and “glutamatergic synapses” (Figure S8C,D). These results suggest that the pulmonary LPS challenge may promote synapse formation in various groups of sensory neurons within the vagal ganglia. Using methods and code from Zanos et al. [33], we conducted dimensionality reduction and unsupervised classification analysis on the vagal nerve electrical signals induced by pulmonary LPS to further explore their characteristics and patterns. Our analysis revealed five clusters of responders that consistently responded to the pulmonary LPS challenge, each characterized by distinct waveforms (Figure S8E,G). These findings highlight the critical role of different clusters of vagal sensory neurons in sensing and responding to the pulmonary LPS challenge.
Based on the data showing that *Trpa1^+^
- vagal sensory neurons respond to LPS, we hypothesize that activation of *Trpa1^+^
- sensory neurons in the vagal ganglion upregulates αCGRP to mediate the response to LPS‐induced lung inflammation. To test this hypothesis, we first confirmed whether αCGRP^+^ sensory neurons project to the lungs. By isolating the vagal ganglia from mice that received an intratracheal injection of Fast Blue, we found that 3.92% of the cells were Fast Blue–positive neurons, indicating their projections to the lungs. Among these lung‐projecting neurons, approximately 30% were CGRP‐positive, and more than 90% originated from the nodose ganglion (Figure 2L). To further investigate the co‐expression of αCGRP and TRPA1 in sensory neurons projecting to the lungs, we crossed *Calca^CreERT2^
- mice with Cre‐dependent Rosa26‐tdTomato mice (*αCGRP^tdTomato^
- mice) to specifically label αCGRP‐expressing neurons that project to the lungs. Through neuroanatomical tracing, we observed that approximately 76.2% of αCGRP^+^ sensory neurons projecting to the lungs were TRPA1‐positive neurons (Figure 2M).
To elucidate the characteristics of nodose *αCGRP^+^Trpa1^+^
- neurons, we reanalyzed the projection‐seq data of vagal neurons conducted by Chang and colleagues [31]. We found distinct expression patterns of *Trpa1^+^
- and Calca ^+^ in vagal neurons projecting to different visceral organs (Figure 2N; Figure S8H). This suggests that different types of sensory neurons project to distinct locations, perform various functions, and generate different electrical signals [33]. Single‐cell RNA sequencing analysis (GSE192987) [32] revealed that QZ2‐labeled vagal sensory neurons project to the lungs. The composition of these neurons includes *Trpa1^+^Calca^+^
- (39.5%), Calca ^+^ (20.4%), Trpa1 ^+^ (13.5%), and other types (26.5%) (Figure 2O). Trpa1 ^+^ Calca ^+^ double‐positive neurons accounted for 39.5% of the lung‐projecting sensory neurons, a proportion significantly higher than that observed in other visceral organs such as the esophagus, stomach, duodenum, colon, heart, and pancreas (Figure 2O). Notably, the majority of these lung‐innervating Trpa1 ^+^ Calca ^+^ neurons originated from the nodose ganglion, comprising up to 86.7% (Figure 2N). Not only was the proportion higher in lung‐projecting neurons, but the expression levels of Trpa1 and Calca were also elevated compared to other tissues (Figure 2O). Further analysis of sensory neurons projecting to the lungs revealed significantly higher expression levels of Calca and Trpa1 in double‐positive neurons compared to single‐positive neurons (Figure 2P,Q). This suggests that nodose *Trpa1^+^Calca^+^
- sensory neurons are more likely to sense LPS and release αCGRP to regulate LPS‐induced pulmonary inflammation.
To confirm the direct role of TRPA1 in mediating CGRP production, we generated a mouse model with TRPA1 specifically deleted in the vagal ganglion (*NG^Trpa1KO^
- mice). In control mice, LPS stimulation led to a significant upregulation of CGRP expression in the NG. In contrast, *NG^Trpa1KO^
- mice showed no increase in CGRP expression following LPS administration (Figure 2R).
αCGRP+ Nodose Sensory Neurons Densely Innervate NEBs
2.4
In the lungs, nodose sensory neurons are primarily distributed beneath the large airways (Figure 3A,B), projecting to the airway bifurcations where they converge into neuroepithelial bodies (NEBs) composed of PNECs. We examined the distribution of αCGRP sensory neurons in the lungs of *αCGRP^tdTomato^
- mice and observed that αCGRP^+^ sensory neurons densely projected to PNECs (Figure 3C,D; Video S1). To confirm the contribution of nodose ganglia, we crossed *Phox2b^Cre^
- mice with Cre‐dependent Rosa26‐tdTomato mice and co‐labeled with CGRP to mark nodose sensory neurons in the lungs. While some nodose sensory neurons project into the interior of NEBs, CGRP‐positive nodose ganglion sensory neurons are located beneath NEBs and do not penetrate into the NEB interiors (Figure 3E,F; Video S2). We conducted a statistical analysis of the nerve fibers surrounding the NEBs and observed that approximately 40% of sensory neurons projected to the NEBs were CGRP^+^ neurons (Figure 3F). Additionally, we observed that CGRP^+^ nodose sensory neurons projected to the alveoli, accounting for 41% of all nodose sensory neurons projecting to alveoli (Figure 3G,H).
αCGRP+Trpa1+ nodose sensory neurons innervate PNECs, promoting excessive CGRP production in PNECs (A,B) Anatomical mapping of vagal sensory neurons within the lung using Phox2btdTomato mice. (C,D) In CalcatdTomato mice, detect the projection of CGRP‐positive neurons onto PNEC cells. (E) In Phox2btdTomato mice, two‐color immunohistochemistry for CGRP (green) and Phox2b‐tdTomato (red) was performed to label CGRP‐positive nodose neurons and PNECs at airway bifurcations. (F) The proportion of CGRP‐positive sensory neurons among the nodose sensory neurons projecting to NEBs. (G) In Phox2btdTomato mice, two‐color immunohistochemistry for CGRP (green) and Phox2b‐tdTomato (red) was conducted to identify CGRP‐positive nodose neurons within the alveolar region. (H) The proportion of CGRP‐positive sensory neurons among the nodose sensory neurons projecting to the alveolar region. (I) Vagal sensory neuronal transport under LPS stimulation was analyzed using RNAseq databases of the vagal ganglion. (J) 19 transport genes in vagal ganglia, along with their enriched GO pathways, were induced by pulmonary LPS stimulation. (K) Administration of LPS or αCGRP+LPS into the lungs of mice. (L) Gating strategies for CGRP+EpCAM+NCAM+CD45− PNEC used in flow cytometry. (M,N) Comparison of the mean CGRP fluorescence intensity in CGRP+EpCAM+NCAM+ cells within lung tissue between the αCGRP‐treated group and the LPS‐treated group. Scale bars: 500µm in (A).30 µm in the upper part of (E). 20 µm in (D). 5 µm in the lower part of (E,G). One‐way ANOVA in (N). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.
LPS Challenge Enhances Vagal Axonal Transport and Promotes CGRP‐Induced CGRP Expression in PNECs
2.5
To investigate whether LPS challenge influences vagal sensory neuronal transport, we analyzed RNA‐seq data from the vagal ganglion (PRJNA1191398). We found that LPS challenged in the mouse lungs increased 19 transport genes in the vagal ganglia. Among the 19 genes, Rab27b, Sybu, Nefl, and Kif5b were associated with anterograde axonal transport. Syt7 (synaptotagmin 7) and Lrrk2 have been reported to mediate neuropeptide release or secretion in neurons [34, 35] (Figure 3I,J).
Activation of αCGRP^+^ sensory neurons and the subsequent release of αCGRP function not only as neurotransmission but also as a neuroimmune regulatory mechanism, contributing to various immune and inflammatory responses—particularly within the lungs and airways [4, 24, 26]. Pulmonary neuroendocrine cells (PNECs), a specialized epithelial cell type, serve as the principal source of CGRP in the lung. These cells co‐express the epithelial marker EpCAM and the neural cell adhesion molecule NCAM1 (Figure S9A) [1]. To identify PNECs in the lung, we performed flow cytometry to detect CGRP^+^EpCAM^+^NCAM1^+^CD45^−^ cells. Our analysis revealed that approximately 80%–90% of EpCAM^+^NCAM1^+^CD45^−^ cells were CGRP‐positive (Figure 3L). To determine whether αCGRP modulates CGRP expression in PNECs, we intratracheally administered αCGRP to mice, followed by LPS stimulation (Figure 3K). LPS treatment enhances CGRP expression intensity in PNECs, and this expression is further increased upon exogenous αCGRP stimulation (Figure 3M,N). To further investigate whether LPS enhances αCGRP‐induced CGRP expression in PNECs, we utilized an in vitro model using the SCLC #1 cell line—a well‐established surrogate for PNECs [36]—and treated the cells with both αCGRP and LPS. Consistent with in vivo findings, CGRP levels were significantly elevated both intracellularly and extracellularly in the combined treatment group compared to cells treated with LPS alone (Figure S9B–D). It is well‐established that CGRP signaling promotes cAMP/PKA pathway activation [37], which in turn plays a critical role in upregulating CGRP expression [38]. In the absence of αCGRP, treatment of PNEC cells with the PKA inhibitor H‐89 significantly reduced CGRP levels in the culture supernatant (Figure S9E,F). Unexpectedly, however, in the presence of αCGRP, H‐89 treatment led to a marked increase in CGRP levels in both the supernatant and cell lysate compared to the αCGRP‐only group (Figure S9E–H). To further dissect the role of PKA in CGRP synthesis, we used lentiviral vectors to deliver shRNA specifically targeting Prkaca in PNEC cells. Knockdown of Prkaca resulted in a significant increase in the expression of CGRP‐related genes (Calca and Calcb) compared to controls (Figure S9I). Moreover, αCGRP stimulation further enhanced CGRP protein expression in Prkaca‐silenced cells (Figure S9J). Collectively, these findings suggest that the regulatory role of PKA in modulating CGRP expression in PNECs may diverge from previously reported mechanisms [39, 40, 41].
Nodose TRPA1+αCGRP+ Sensory Neurons Induce CGRP Expression in PNECs and Amplify Lung Inflammation
2.6
Next, we examined whether αCGRP^+^ nodose sensory neurons regulate CGRP expression in PNECs. We injected diphtheria toxin (DT) into the vagal ganglion of *Calca^CreERT2^Rosa26^DTR^
- mice to specifically ablate αCGRP‐expressing neurons in the vagal ganglion (referred to as *NG^CalcaABLATE^
- mice). Age‐matched Cre‐negative littermates that also received DT served as controls (Figure 4A). As expected, αCGRP^+^ neurons in the vagal ganglion were effectively ablated (Figure 4B). Notably, the structural integrity of neuroepithelial bodies (NEBs) in the lungs of *NG^CalcaABLATE^
- mice remained unchanged (Figure 4C). However, flow cytometric analysis revealed that in *NG^CalcaABLATE^
- mice, the intensity of CGRP expression within the CGRP^+^EpCAM^+^NCAM1^+^ lung epithelial population was significantly reduced, and LPS was no longer able to induce upregulation of CGRP expression in PNECs (Figure 4D–F). In addition, LPS injury led to a decrease in the total number of CD45^−^EpCAM^+^NCAM1^+^ epithelial cells (Figure 4G), and TNF expression in F4/80^+^ macrophages isolated from bronchoalveolar lavage fluid (BALF) was also diminished in *NG^CalcaABLATE^
- mice relative to controls (Figure 4H,I).
Nodose αCGRP+ sensory neurons deficiency results in reduced CGRP expression levels in PNECs and reduced lung inflammation. (A) Diphtheria Toxin (DT) was injected into the nodose ganglia in CalcaCreERT2; loxP‐DTR (NGCalcaABLATE ) mice and loxP‐DTR (Ctrl) mice. (B,C) CGRP immunostaining (green) in the nodose/jugular complex (B) and lung (C) of CalcaCreERT2; loxP‐DTR (NGCalcaABLATE ) mice and loxP‐DTR (Ctrl) mice following NG injection, with DAPI nuclear counterstaining. (D)The gating strategy of CGRP+EPCAM+NCAM+ PNECs from NGCalcaABLATE mice and Ctrl mice following LPS stimulation. (E,F) Comparison of the mean CGRP fluorescence intensity in EpCAM+NCAM+ cells in lung tissue from NGCalcaABLATE mice and Ctrl mice following LPS stimulation. (n = 3–14 mice/group) (G) The proportion of CGRP+EPCAM+NCAM+ cells among all CD45− cells in lung tissue from NGCalcaABLATE mice and Ctrl mice following LPS stimulation. (n = 3–7 mice/group) (H) The number of PNECs per 5105 CD45− cells in lung tissue from NGCalcaABLATE mice and Ctrl mice following LPS stimulation. (n = 3–7 mice/group) (I) LPS induced a lower level of TNFα in F4/80+ macrophages in the BALF of NGCalcaABLATE mice compared with Ctrl mice (n = 3–7 mice/group). (J) The injections of AAV‐sgCalca‐Calcb and AAV‐sgScramble into the jugular/nodose complex are performed to generate NGCGRPKO mice and Ctrl mice, respectively.CGRP immunostaining (green) in the nodose/jugular complex of NGCGRPKO mice and Ctrl mice following NG injection. (K,L) LPS induced a lower level of TNFα in F4/80+ macrophages in the BALF of NGCGRPKO mice compared with Ctrl mice (n = 3–7 mice/group). (M–O) Comparison of the mean CGRP fluorescence intensity in EpCAM+NCAM+ cells in lung tissue from NGCGRPKO mice and Ctrl mice following LPS stimulation. (n = 3–14 mice/group) Scale bars: 100 µm in (C,J). 50 µm in (B). One‐way ANOVA in (F,G,I,L,O). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.*
Vagal‐Derived CGRP Amplifies LPS‐Induced Inflammation and Activates PNECs
2.7
Ablation of CGRP‐positive vagal sensory neurons can also deplete other neuropeptides within these neurons. To specifically investigate the role of CGRP, we injected AAV‐CMV‐Cas9 together with AAV‐sgCalca‐Calcb targeting the Calca/Calcb genes, or a control AAV‐scramble virus, into the jugular/nodose complex of mice. This approach successfully generated mice with CGRP deficiency in the vagal ganglia (*NG^CGRPKO^ *) and corresponding control mice (Ctrl) (Figure 4J). Consistent with the results from ablation of αCGRP‐positive vagal neurons, *NG^CGRPKO^
- mice exhibited significantly reduced TNF‐α expression in F4/80‐positive macrophages from BALF (Figure 4K,L). Moreover, CGRP expression in pulmonary neuroendocrine cells (PNECs) was also significantly lower in *NG^CGRPKO^
- mice compared to controls (Figure 4M–O). These findings indicate that vagal‐derived CGRP plays a critical role in amplifying LPS‐induced inflammation and activating PNECs.
Nodose TRPA1+αCGRP+ Sensory Neurons Regulate the Proliferation of PNECs
2.8
To further validate these findings, we performed similar experiments in *NG^Trpa1KO^
- mice, in which Trpa1 is selectively deleted in the vagal ganglion (Figure 5A). Consistent with the results observed in *NG^Trpa1KO^
- mice, both the number of CGRP^+^ cells and the expression level of CGRP within the CGRP^+^EpCAM^+^NCAM1^+^ lung epithelial population were significantly decreased in LPS‐challenged *NG^Trpa1KO^
- mice compared to controls (Figure 5B–D). These findings suggest that TRPA1^+^ neurons in the NG innervate PNECs and modulate their CGRP expression in response to LPS stimulation. Collectively, our data indicate that TRPA1^+^αCGRP^+^ neurons in the NG likely serve as a major source of αCGRP and may enhance CGRP expression in PNECs.
Nodose Trpa1+αCGRP+ sensory neurons are necessary for PNECs proliferation. (A) The injections of AAV‐sgTrpa1 and AAV‐sgScramble into the right jugular/nodose complex are performed to generate NGTrpa1KO mice and Ctrl mice, respectively. (B) The gating strategy of CGRP+EPCAM+NCAM+ PNECs from NGTrpa1KO mice and Ctrl mice. (C) The proportion of highly CGRP‐expressing PNECs among all CGRP+EPCAM+NCAM+ cells in lung tissue from NGTrpa1KO mice and Ctrl mice (n = 5 mice/group). (D) Comparison of CGRP mean fluorescence intensity in CGRP+EpCAM+NCAM+ cells in lung tissue from NGTrpa1KO mice and Ctrl mice (n = 5 mice/group). (E) The number of PNECs per 5105 CD45− cells in lung tissue from NGTrpa1KO mice and Ctrl mice following LPS stimulation (n = 5 mice/group). (F) Diphtheria Toxin (DT) was injected into the nodose ganglia in CalcaCreERT2; loxP‐DTR (NGCalcaABLATE ) mice and loxP‐DTR (Ctrl) mice. Three weeks after DT injection, a 7‐day LPS stimulation model was established. (G,H) During the 7 days of LPS stimulation, NGCalcaABLATE mice exhibited lower levels of TNF‐α in BALF compared to Ctrl mice (n = 3 mice/group). (I) The mortality rate in NGCalcaABLATE mice and Ctrl mice during 7 days of LPS stimulation. (n = 10 mice/group). (J) The proportion of highly CGRP‐expressing PNECs among all CGRP+EPCAM+NCAM+ cells in lung tissue from NGCalcaABLATE mice and Ctrl mice following 7 days of LPS stimulation (n = 3–4 mice/group). (K) The number of PNECs per 5105 CD45− cells in lung tissue from NGCalcaABLATE mice and Ctrl mice following 7 days of LPS stimulation (n = 3–4 mice/group). (L,M) The proportion of Ki67+ PNECs among all CGRP+EPCAM+NCAM+ cells in lung tissue from NGCalcaABLATE mice and Ctrl mice following 7 days of LPS stimulation (n = 3‐4 mice/group). Student's t‐test in (c–e). One‐way ANOVA in (h,j,k,m). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.
In the RNA sequencing dataset (PRJNA1191398) of the NG following LPS stimulation, differential pathway analysis revealed significant enrichment in pathways associated with the “regulation of epithelial cell proliferation.” This observation suggests that TRPA1^+^αCGRP^+^ sensory neurons in the NG may contribute to the regulation of PNEC proliferation. In *NG^Trpa1KO^
- mice, in which TRPA1 is selectively deleted in the vagal ganglion, we observed a significant reduction in the number of CGRP^+^EpCAM^+^NCAM1^+^ PNECs compared to control mice (Figure 5E). To further examine the role of αCGRP released from activated TRPA1^+^αCGRP^+^ sensory neurons in the NG on PNEC proliferation, we utilized *NG^CalcaABLATE^
- mice, in which αCGRP^+^ sensory neurons in the NG are selectively ablated, and subjected them to a 7‐day LPS treatment protocol (Figure 5F). Consistent with the findings from the 1‐h LPS stimulation model, TNF‐α levels in the BALF of *NG^CalcaABLATE^
- mice were significantly lower than those of control mice (Figure 5G,H). Additionally, selective ablation of αCGRP^+^ sensory neurons in the vagal ganglion significantly reduced LPS‐induced mortality (Figure 5I). With respect to PNECs, ablation of αCGRP^+^ neurons led to a significant reduction in CGRP expression levels (Figure 5J), accompanied by a decrease in PNEC number and impaired regenerative capacity compared to controls (Figure 5K). Furthermore, Ki67 immunostaining revealed a significantly lower proportion of proliferating Ki67^+^ cells within the CGRP^+^EpCAM^+^NCAM1^+^ population in *NG^CalcaABLATE^
- mice (Figure 5L,M). Together, these findings demonstrate that activation of TRPA1^+^αCGRP^+^ sensory neurons in the NG and the subsequent release of αCGRP not only regulate CGRP expression in PNECs but also influence their abundance and proliferative potential, thereby playing a pivotal role in modulating LPS‐induced pulmonary inflammation.
PNECs Amplify LPS‐Induced Lung Inflammation
2.9
To investigate whether PNECs contribute to LPS‐induced lung inflammation, we generated *Ascl1^ABLATE^
- mice by crossing *Ascl1^CreERT2^
- mice with Cre‐dependent Rosa26‐DTR mice, followed by intraperitoneal injection of diphtheria toxin (DT) to selectively ablate PNECs (Figure 6A). Immunofluorescence confirmed the effective depletion of CGRP^+^ neuroepithelial bodies (NEBs) in these mice (Figure 6A), along with a significant reduction in EpCAM^+^NCAM1^+^ lung epithelial cells compared to control mice (Figure 6B,C). We next recorded vagal nerve responses to LPS in both *Ascl1^ABLATE^
- and control mice. PNEC ablation did not alter the electrophysiological response of the vagus nerve to LPS, further supporting our previous observation that the vagus nerve senses and responds to pulmonary LPS directly via TRPA1 (Figure 6D,E). Upon LPS challenge, *Ascl1^ABLATE^
- mice exhibited significantly attenuated lung inflammation, as indicated by significantly lower levels of TNF‐α in bronchoalveolar lavage fluid (BALF) and a reduction in TNF‐α^+^F4/80^+^ macrophages (Figure 6F–H). Given the central role of polymorphonuclear neutrophils (PMNs) in pulmonary inflammation [42], we performed flow cytometric analysis and observed a significant decrease in the number of Ly6G^high^Ly6C^low^CD11b^+^ PMNs in the lungs of LPS‐treated *Ascl1^ABLATE^
- mice compared to controls (Figure 6I,J). Consistently, the expression of inflammatory genes Il6 and Ccl2 was also significantly reduced in the lungs of *Ascl1^ABLATE^
- mice (Figure 6K). Furthermore, PNEC ablation significantly improved survival in response to LPS‐induced lung injury (Figure 6L).
PNECs deficiency alleviated LPS‐induced pulmonary inflammation. (A) Sequential intraperitoneal injections of tamoxifen and diphtheria toxin were used to genetically ablate PNECs in Ascl1CreERT2; loxP‐DTR (Ascl1ABLATE ) mice and loxP‐DTR (Ctrl) mice. (B,C) The proportion of EPCAM+NCAM+ cells among CD45‐ cells was assessed in Ascl1ABLATE mice and Ctrl mice. (n = 5 mice/group). (D) Vagal responses to LPS stimuli within the lung in Ascl1ABLATE mice and Ctrl mice. (n = 3–7 mice/group) (E) Quantification of maximum responses to LPS in Trpa1KO mice and Ctrl mice (n = 3 mice/group). (F) Following LPS stimulation, the level of TNF‐α in the BALF of Ascl1ABLATE mice was lower compared to that in Ctrl mice (n =4–5 mice/group). (G,H) Following LPS stimulation, Ascl1ABLATE mice exhibited a lower level of TNF‐α in F4/80+ macrophages in the BALF compared to Ctrl mice (n = 4‐5 mice/group). (I) The proportion of Ly6GhighLy6Clow cells among CD11B+ cells in lung tissue was assessed in Ascl1ABLATE mice and Ctrl mice following LPS stimulation (n = 4–5 mice/group). (J) The proportion of Ly6GhighLy6ClowCD11b+ cells (PMNs) among live cells in lung tissue was assessed in Ascl1ABLATE mice and Ctrl mice following LPS stimulation (n = 4 mice/group). (K) Il6 and Ccl2 expression levels in lung tissue were assessed in Ascl1ABLATE mice and Ctrl mice following LPS stimulation. (n = 6–10 mice/group) (L) The mortality rate in Ascl1ABLATE mice and Ctrl mice during 7 days of LPS stimulation. (n = 10 mice/group). Scale bars:100 µm in (a). Student's t‐test in (c,e). One‐way ANOVA in (f,h,j,k). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.
To further investigate the role of αCGRP in LPS‐induced lung inflammation, we intratracheally instilled αCGRP into mouse lungs, followed by LPS treatment. The αCGRP + LPS group exhibited a higher number of pulmonary Ly6G^high^Ly6C^low^CD11b^+^ PMNs compared to the LPS‐only group (Figure S10A–C). Moreover, in vitro treatment of bone marrow–derived macrophages (iBMDMs) with αCGRP prior to LPS exposure further enhanced the LPS‐induced expression of the chemokines Cxcl10 and Cxcl1 (Figure S10D,E). Together, these findings strongly support the conclusion that PNECs and their secreted product, CGRP, promote LPS‐induced lung inflammation.
GABA Feedback Regulates the Production of CGRP in PNECs and Vagal Sensory Neurons
2.10
PNECs are main GABA‐producing cells in the lung, and GAD1 and ABAT are key enzymes for GABA metabolism [43]. GAD1 mediates the synthesis of GABA, while ABAT controls its breakdown [44]. The GABAergic synapse and receptor clustering are regulated by SHISA7, PHF24, LHFPL4, DBI, NRXN1, and NLGN2 [45]. ACBP (Acyl‐CoA Binding Protein, a polypeptide encoded by Dbi) binds to GABA receptors and regulates GABA inhibition [46]. By analyzing our vagal ganglion and lung RNAseq databases, we found that lung Abat expression was downregulated under LPS challenge, indicating that GABA is increased in PNECs. Concurrently, Shisa7, Phf24, Nrxn1, Usp46 were downregulated in PNECs and upregulated in vagal ganglia. Dbi expression was shown in the opposite direction (Figure 7A). These findings indicate that GABA receptors are activated in vagal ganglia and inhibited in PNECs under LPS challenge. Furthermore, we analyzed vagal ganglia GSE192987 [31] and PNEC [1] scRNAseq databases and found that Gabbr1/2 and Gabrb3 were expressed by lung‐innervated Trpa1 ^+^ *Calca^+^
- neurons and Gabrb1/3 and Gabbr2 were expressed in PNECs [1] (Figure S11). As such, we hypothesize that PNEC‐derived GABA can inhibit the excitability of *Trpa1^+^Calca^+^
- neurons and CGRP synthesis in PNECs (Figure 7B).
GABA feedback regulates the production of CGRP in PNECs and vagal sensory neurons. (A) Analyze the expression of genes related to GABAergic synapses, GABA metabolism, and its receptors in the RNAseq databases of LPS‐stimulated lungs and vagal ganglia. (B) A schematic of the hypothesis that GABA derived from PNECs can inhibit the excitability of Trpa1+Calca+ neurons and CGRP synthesis in PNECs. (C) Measurement of Calcb gene expression in SCLC cells stimulated with LPS and GABA. (D) Measurement of Gabrb3 and Calca gene expression in Gabrb3‐knockdown PNEC cells and control cells after GABA stimulation. (E) The levels of CGRP in the vagal ganglia were measured after GABA and LPS stimulation via endotracheal intubation. (F) Experimental timeline of primary vagal sensory neurons stimulation with GABA and LPS. (G) The levels of CGRP in primary vagal sensory neurons were measured following GABA and LPS stimulation. (H) Experimental timeline of primary vagal sensory neurons stimulation with GABAr antagonist, GABA, and LPS. (I,J) The levels of CGRP in primary vagal sensory neurons were measured following treatment with GABAA receptor antagonist SR‐95531 (SR) and GABAB receptor antagonist CGP52432 (CGP). One‐way ANOVA in (c,d). Mean ± SEM. ns, not significant. * p < 0.05, ** p < 0.01, and *** p < 0.001.
To test this hypothesis, we pretreated the PNEC cell line with GABA, followed by LPS stimulation. The results showed that LPS significantly upregulated Calcb expression, while GABA pretreatment markedly attenuated this effect (Figure 7C). Given that Gabrb3 is highly expressed in PNECs [1], we knocked down this gene and found that Calca expression was significantly upregulated in the *Gabrb3‐*silenced group (Figure 7D). These findings suggest that GABA may inhibit CGRP expression in PNECs through GABRB3. To further investigate whether GABA affects CGRP expression in the vagal ganglia, we delivered GABA intratracheally into the lungs of mice [24], followed by LPS challenge. Under LPS stimulation, we found that GABA treatment enhanced CGRP expression in vagal sensory neurons (Figure 7F,G). This result contradicts our initial hypothesis that GABA suppresses the activity of Trpa1 ^+^ Calca ^+^ sensory neurons. To further test this mechanism, we treated isolated vagal sensory neurons with GABA and LPS in vitro and added specific GABA receptor antagonists: SR‐95531 (a GABA_A_ receptor antagonist) and CGP52432 (a GABA_B_ receptor antagonist) [47, 48] (Figure 7H). We found that blockade of the GABAA receptor had no significant effect on CGRP expression in neurons treated with GABA + LPS. In contrast, blockade of the GABA_B_ receptor significantly reduced the number of CGRP^+^ neurons in the GABA + LPS group (Figure 7H–J). These results suggest that GABA_B_ receptor blockade may promote CGRP release from sensory neurons, thereby lowering intracellular CGRP levels and reducing signals detected by flow cytometric staining. Together with our finding that GABA treatment elevates intracellular CGRP levels in sensory neurons, these data support a model wherein GABAB receptor activation inhibits CGRP secretion, resulting in its intracellular retention within vagal sensory neurons. These findings suggest that GABA regulates the CGRP release from vagal sensory neurons.
Discussion
3
In this study, we identified that αCGRP^+^TRPA1^+^ nodose sensory neurons innervated PNECs and utilized TRPA1 to sense LPS, increasing their αCGRP expression. αCGRP derived from sensory neurons acts on PNECs, not only promoting their CGRP expression but also regulating their proliferation, thereby amplifying pulmonary inflammation responses. GABA (as a major product of PNECs) feedback regulated CGRP production in nodose sensory neurons. Thus, we have uncovered that αCGRP^+^TRPA1^+^ nodose sensory neurons collaborate with PNECs to form neural circuits that regulate LPS‐induced lung inflammation (Figure S10F).
During the development of acute lung injury (ALI) and acute respiratory distress syndrome (ARDS), vagal sensory neurons rapidly enter a nerve injury‐like state, leading to altered vagal excitability [32]. It has been demonstrated that LPS can exert fast excitatory actions via TRPA1 in neurons, mediating acute neurogenic inflammation and pain [16]. Considering the slow activation of traditional immune signaling pathways [49], a plausible pattern of recognition is coupling TLR4 receptors with ion channels, such as TLR4‐TRPV1, which may explain rapid neuronal activation [50]. However, antagonism of TLR4 could not completely inhibit LPS‐induced neuronal activation, such as calcium influx [16, 51]. In a recent study, bacterial LPS can activate the TLR4 receptor on TRPV1^+^ vagal sensory neurons, transmitting signals to the paraventricular hypothalamic nuclei (PVN), leading to the activation of corticotropin‐releasing hormone neurons and inducing sickness behavior [8]. The vagus nerve response to LPS was assessed by activating the TLR4 receptor, which can increase neuronal sensitivity to capsaicin [8]. Another in vitro neuronal calcium signaling recording experiment found that TRPA1^+^ neurons in the nodose ganglion can directly respond to LPS stimulation without relying on TLR4[16]. However, in an in vivo intestinal neuronal calcium signaling recording experiment, LPS did not directly activate vagal sensory neurons [7]. This issue remains controversial: whether vagal sensory neurons can directly sense LPS in the pulmonary environment in vivo, and which receptor (TLR4 or TRPA1) mediates LPS recognition, is still unclear. In our study, pulmonary LPS‐induced vagal excitability was eliminated by deletion of Trpa1 in the nodose ganglia (Figure 2B,C), but not by systemic deletion of Tlr4 (Figure S4A–F). By analyzing scRNAseq datasets of sensory neurons projecting to the lungs [31], we found that few Tlr4 ^+^ sensory neurons were presented in QZ2 labeled lung‐innervated nodose sensory neurons, and Trpa1 was 20‐fold more expressed than Tlr4 (Figure S4J). We ligated the middle of the trachea and inserted a catheter to inject LPS into the distal region of the lung. This is done to prevent LPS from regurgitating into the laryngopharynx to activate sensory neurons in the jugular ganglia, where TLR4 is expressed 10‐fold higher than in nodular ganglia (Figure S4I). It is estimated that less than 12% of vagal Tlr4‐positive neurons were reported to be located in the nodose ganglia, and more than 50% of vagal Tlr4‐positive neurons were located in the jugular ganglia [52]. Jin et al. discovered that LPS cannot directly activate vagal sensory neurons projecting to the gut but instead activates them through inflammatory cytokines [7]. Indeed, immune cell‐derived inflammatory cytokines are important factors in neuronal activation and can also promote the release of CGRP from neurons. However, the time required for immune signaling pathways is much longer than that for neuronal signaling. Moreover, the inflammatory cytokines induced by LPS are complex and diverse, including both pro‐inflammatory and anti‐inflammatory factors, which activate different populations of sensory neurons. Additionally, we found that, unlike vagal sensory neurons projecting to the gut, those projecting to the lungs exhibit higher expression of TRPA1 and co‐express elevated levels of αCGRP (Figure 2L–Q). These features confer unique functions in pathogen recognition and immune regulation to vagal neurons projecting to the lungs. Thus, we conclude that TRPA1, rather than TLR4 in the nodose sensory neurons senses LPS and couples with MYD88 to increase vagal excitability. This notion is supported by LPS stimulation induces MYD88 aggregation in Trpa1‐overexpressed neurons (Figure S5D–F) and deletion of Myd88 in nodose sensory neurons (*Phox2b^+^ *) abolishes LPS‐triggered vagal excitability (Figure 1D–F). We further introduced Myd88‐EGFP constructs into vagal sensory neurons and found that, at baseline, MYD88 aggregation was significantly more pronounced in vagal sensory neurons than in Trpa1‐negative Neuro‐2a cells. However, the precise mechanism of interaction between TRPA1 and MYD88 within vagal neurons remains to be elucidated.
Different vagal sensory neurons project to various visceral organs [31]. It remains unclear whether vagal sensory neurons, such as those expressing TRPA1 and αCGRP, which project to different organs, have similar inflammatory regulatory functions. A recent study on the vagal‐cNST inflammatory circuit revealed that different vagal sensory neurons can respond to different inflammatory signals. TRPA1^+^ neurons respond to anti‐inflammatory factor IL‐10, while CGRP^+^ neurons respond to pro‐inflammatory factor IL‐6 and IL‐1𝛽, thereby suppressing systemic inflammatory responses [7]. This anti‐inflammatory effect originates from the activation of sensory neurons, whereby signals are transmitted to DBH^+^ neurons in the cNST, which mediate inflammatory regulatory reflexes, such as through the cholinergic anti‐inflammatory pathway [9]. In our study, we propose that TRPA1 expressed in αCGRP^+^ sensory neurons can directly sense LPS and make neurons synthesize αCGRP to influence PNECs. Through analyzing the database by Chang and colleagues [31], we found that vagal sensory neurons projected to the lungs are very different from those projecting to abdominal organs (Figure 2O,P). Compared with vagal sensory neurons that innervate the esophagus, stomach, duodenum, colon, and pancreas, those innervating the lungs show the highest proportion of Trpa1^+^Calca^+^ double‐positive cells (Figure 2O; Figure S8H). Notably, single Calca^+^ sensory neurons innervating the lungs constitute only a very small fraction (Figure 2O,P). Deletion of Trpa1 or Calca in sensory neurons had the same effect on CGRP expression in PNECs and lung inflammatory responses, suggesting that *Trpa1^+^Calca^+^
- sensory neurons, rather than single *Trpa1^+^
- or *Calca^+^
- sensory neurons, can mitigate their effects on PNECs (Figure 5).
Vagal nodose sensory neurons project various types of sensory terminals at the NEBs, among which Pvalb ^+^ neurons are responsible for responding to airway closure [1], but the functions of nodose TRPA1^+^αCGRP^+^ neurons remain largely unresolved. We observed that in larger airway, αCGRP^+^ neurons of the NG project to PNECs. The nerve endings of these neurons are primarily located beneath the airway epithelium, especially projecting below the NEB structure, rather than extending internally into the NEBs, suggesting that these αCGRP^+^ nodose sensory neurons may not contact with LPS. However, in the distant airway, CGRP^+^ nerve fibers can extend into the alveolar space, indicating that nerve endings in these areas may sense LPS (Figure 3).
One important finding in this study is that nodose *TRPA1^+^αCGRP^+^
- neurons innervate PNECs and release αCGRP to promote CGRP production in PNECs in response to LPS challenge. PNECs are one of the epithelial cell types lining large and small airways that form a tiny cellular population (0.4% of total airway epithelial cells or 0.01% of all lung cells) [53]. NEBs frequently populate diametrically opposed positions to the bifurcation points of branching airways [53]. For defense against bacterial invasion, *Trpa1^+^Calca^+^
- nodose sensory neurons can sense LPS to produce CGRP and amplify its effects via PNECs. We used EpCAM^+^NCAM1^+^CD45^−^ cells to define PNECs, and 80%–90% of cells are CGRP^+^ in this cell population. In our study, deletion of Trpa1 or Calca in sensory neurons reduced CGRP expression in PNECs from LPS‐challenge mice. Our study supports the activation of TRPA1^+^αCGRP^+^ nodose sensory neurons as a mechanism to amplify LPS‐induced lung inflammation. However, several reports suggest that activation of nociceptor sensory neurons can suppress lung inflammation [54, 55, 56]. Nociceptors are a diverse group of sensory neurons that include various subtypes capable of releasing CGRP, SP, VIP, and other neuronal mediators [49, 57]. Notably, CGRP^+^ sensory neurons are predominantly localized in the jugular ganglion [57], while CGRP^+^ neurons in the nodose ganglion are often overlooked. We found that over 90% of the CGRP^+^ sensory neurons projecting to the lungs originate from the nodose ganglion (Figure 2L). CGRP can be further divided into αCGRP (encoded by Calca) and βCGRP (encoded by Calcb), but previous studies have often referred to them collectively as “CGRP.” This distinction is crucial, as the expression patterns of these genes differ significantly in vagal sensory neurons (Figure S4G,H). For instance, βCGRP^+^ vagal sensory neurons have been shown to suppress group 2 innate lymphoid cell function and allergic airway inflammation [56]. However, the role of nodose CGRP^+^ sensory neurons, particularly αCGRP, in LPS‐induced pneumonia remains unclear. Most of these studies utilized Trpv1 ^Cre^ and Nav1.8 ^Cre^ mice to investigate sensory neuron functions. TRPV1 and Nav1.8 are markers of nociceptive neurons, encompassing a broad range of neuron types, not limited to αCGRP‐positive neurons. Additionally, our study focuses on αCGRP. Furthermore, using *Nav1.8^Cre^/Rosa26^DTA^
- mice results in the absence of systemic nociceptive neurons from birth or systemic depletion of nociceptive neurons via intraperitoneal injection of diphtheria toxin (DTA). In our study, we used *Calca^CreERT2^/Rosa26^DTR^
- mice and achieved specific ablation of αCGRP^+^ neurons within the nodose ganglion through localized injection of DTA directly into the nodose ganglion. Due to differences in the target cells, sensory neurons may exert different regulatory effects. Our focus is on αCGRP^+^ neurons targeting PNECs. PNECs themselves contain various bioactive substances that, together with CGRP, collectively regulate lung inflammatory responses.
At the same time, compared to PNEC cell lines treated with αCGRP alone, both protein and transcriptional levels of CGRP were increased in the LPS + αCGRP treatment group, suggesting that LPS‐induced signaling is essential for the αCGRP‐enhanced CGRP expression in PNECs. Unlike previous reports [39, 40, 41], PKA signaling may negatively regulate CGRP expression in PNECs, as evidenced by higher CGRP levels in cell lysates and supernatants in the H89 (PKA inhibitor) + αCGRP treatment group than in the αCGRP treatment group (Figure S9). Whether PKA signaling is involved in LPS stimulation‐enhanced CGRP expression is worthy of further investigation. As previously reported, the release of CGRP from respiratory vagal afferents is critical to bacterial clearance in lungs [55].
Taken together, we have identified a population of Trpa1 ^+^ Calca ^+^ sensory neurons that project to the lungs, detect intrapulmonary LPS, trigger vagal electrical excitations, and interact with PNECs through CGRP. Meanwhile, PNECs also release GABA, providing negative feedback inhibition on vagal CGRP. Therefore, we have established a nodose ganglion‐pulmonary neuroepithelial circuit that senses and regulates lung inflammatory responses.
Experimental Section/Methods
4
Mice
4.1
C57BL/6J mice were purchased from Charles River Laboratories. Tlr4 knockout (*Tlr4^−/−^ *) mice, Trpa1 knockout (*Trpa1^−/−^ *) mice Rosa26‐stop(flox)‐DTR and Ai9 strain Rosa26‐stop(flox)‐tdTomato reporter mice, as well as Ai32 (Rosa26‐LSL‐ChR2/EYFP) mice, were all obtained from The Jackson Laboratory. *Phox2b^Cre^
- mice were provided by S. Wang (Hebei Medical University). *MyD88^flox^
- mice were provided by X. Zhang (Shanghai Institute of Immunology, CAS). *Ascl1^CreERT2^
- mice were provided by P. Sui (Shanghai Institute of Biochemistry and Cell Biology, CAS). *Calca^CreERT2^
- mice were provided by H. Song (Zhejiang University). *Trpa1^−/−^
- mice were provided by Q. Han (Fudan University).
For labeling of nodose ganglion sensory neurons, *Phox2b^Cre/Cre^
- mice were crossed with *Rosa26^stop(flox)‐tdTomato^
- mice to generate nodose ganglion neuron lineage‐labeled mice (*Phox2b^tdTomato^ *). For nodose ganglion‐specific MyD88 deletion experiments, *MyD88^fl/fl^_Phox2b^Cre/−^
- mice were crossed with *MyD88^fl/fl^
- mice to generate mice with nodose ganglion‐specific MyD88 deletion (MyD88^Phox2b^) or control (*MyD88^fl/fl^ *) littermates. For Tlr4 knockout mouse experiments, Tlr4^+/−^ heterozygous mice were paired to produce wild‐type *Tlr4^+/+^
- (Ctrl mice) and Tlr4 knockout *Tlr4^−/−^
- (*Tlr4^KO^ *) mice. For Calca lineage neuron depletion experiments, *Calca^CreERT2/−^ *; *Rosa26^stop(flox)‐DTR/stop(flox)‐DTR^
- mice were crossed with *Rosa26^stop(flox)‐DTR/ stop(flox)‐DTR^
- mice to generate Calca lineage neuron‐depleted mice (*NG^CalcaABLATE^ *) and control *CalcaCreERT2‐/‐; Rosa26^stop(flox)‐DTR/stop(flox)‐DTR^
- (Ctrl) littermates. Then, DT was injected into the nodose ganglion to achieve the depletion of Calca lineage neurons. For vagal ganglion‐specific TRPA1 deletion experiments, we used C57BL/6J mice and injected AAV‐sgTrpa1 (*NG^Trpa1KO^
- mice) or AAV‐Ctrl (Ctrl mice) into the nodose ganglion. For the experiment of PNEC ablation, *Ascl1Cre^ERT2/−^; Rosa26^stop(flox)‐DTR/stop(flox)‐DTR^
- mice were crossed with *Rosa26^stop(flox)‐DTR/stop(flox)‐DTR^
- mice to generate Ascl1 lineage‐ablated mice (*Ascl1^CreERT2/−^; Rosa26^stop(flox)‐DTR/stop(flox)‐DTR^
- mice or *Ascl1^ABLATE^
- mice) and control littermates (*Ascl1^CreERT2^‐/‐; Rosa26^stop(flox)‐DTR/stop(flox)‐DTR^
- or Ctrl mice).
All animal experiments were carried out in compliance with animal care standards and received approval from the Committees on Animal Research at the Shanghai Institute of Immunity and Infection, Chinese Academy of Sciences, under approval number A2021032. These mice were bred and maintained in an SPF animal facility at Shanghai Institute of Immunity and infection. The mice were housed with free access to food (ad libitum) and water in 12‐h dark/light cycle. Both male and female mice were used in all experiments. Anesthesia was induced with an intraperitoneal (ip) injection of Avertin (1.25%). The number of mice used is as indicated in Figure legends; 3 or more mice were used per experimental condition.
Mouse Animal Models
4.2
The mice were anesthetized by inhalation of isoflurane. They were intratracheally administered either lipopolysaccharide (LPS; 5 mg/kg, Sigma, L2880) dissolved in PBS or an equal volume of PBS as a control. Similarly, CGRP (20pg, Phoenix pep, 015‐09) or PBS was intratracheally delivered.
Cells
4.3
Cell cultures were maintained in a humidified atmosphere at 37°C with 5% CO_2_. SCLC cells were cultured as previously described [36]. Briefly, cells were cultured in RPMI 1640 medium supplemented with 10% FBS (Gibco) and 1% penicillin/streptomycin (Hyclone). After observing the cell status and confluency, cells were harvested by centrifugation at 800 rpm for 5 min following 1‐2 days of culture. The supernatant was discarded, and the cell pellet was resuspended in 2 mL of fresh medium. Cells were then gently pipetted multiple times to dissociate cell aggregates and obtain a single‐cell suspension, which was used for subsequent experiments.
Neuro‐2a (ATCC CCL‐131) cells were cultured in EMEM medium supplemented with 10% FBS, and 1% penicillin/streptomycin. Trypsin‐EDTA solution (Gibco) was added to obtain a cell suspension for subsequent experiments.
Viruses
4.4
rAAV vectors were generated in 293T cells by transfecting packaging plasmids (pHelper, pRep/Cap) and the ITR‐containing vector plasmid in a 1:1:1 molar ratio. The supernatant was collected at 72 and 120 h after transfection, and the cells were collected at 120 h after transfection. Viral particles were concentrated by iodixanol density gradient centrifugation at 288 000 × g for 3 h at 10°C. AAV titers were quantified by qPCR [58]. Primer sequences for qPCR are listed in the table. For lentivirus production, HEK293 cells in a 10 cm dish were transfected with 4.5 µg of the psPAX2 plasmid (Addgene#12260), 1.5 µg of the pMD2.G plasmid (Addgene#12259), and 6 µg of the vector plasmid. Lentiviruses were harvested from the supernatant at 48 and 72 h after transfection.
Plasmids Construction
4.5
To label MyD88 in live cells [59], mouse MyD88 was modified with EGFP at the C‐terminus using overlap PCR. The MyD88‐GFP fragment was then cloned into the lentiviral backbone pWPI‐IRES‐puro. To overexpress Trpa1, mouse Trpa1 was fused with IRES‐mCherry and inserted into a plasmid driven by the CMV promoter (CMV‐Trpa1‐IRES‐mCherry). The AAV plasmids for CRISPR guidance were modified from pX601‐saCas9 (Addgene#61591). In brief, the saCas9 fragment was fused with the hsyn promoter, WPRE, and bGH (poly A) to generate the hsyn‐saCas9‐WPRE‐bGH fragment. This fragment was then inserted between the two ITRs of the pX601 donor plasmid to generate pXN‐hsyn‐saCas9. For the sgRNA vector, saCas9 in pX601 was replaced with mCherry. sgRNA oligos were designed using CRISPick (https://portals.broadinstitute.org/gppx/crispick/public) and cloned into the U6‐sgRNA scaffold. The U6‐sgRNA units were then amplified by PCR and cloned into the pX601‐mCherry backbone using Gibson assembly, as described previously [60].
PNEC Ablation
4.6
For the experiment of PNEC ablation, *Ascl1^ABLATE^
- mice and Ctrl mice were intraperitoneally injected with 100 µL of tamoxifen (Sigma, T5648) (20 mg/mL in corn oil) for 5 consecutive days. Experiments were performed 7 days after the last administration. Mice were intraperitoneally injected with 200 ng of diphtheria toxin (Sigma, D0564) in 100 µL of PBS once daily for four consecutive days.
Bronchoalveolar Lavage in Mice
4.7
Collect bronchoalveolar lavage fluid (BALF) as previously described [61]. Mice were euthanized under avertin anesthesia. The neck skin was cut, and blunt dissection was performed with forceps to expose the trachea. A small incision was made on the trachea, and care was taken not to cut it completely. A 1 mL syringe was used to aspirate 1 mL of PBS, and then replaced with a blunt needle. The needle was inserted through the small incision into the trachea. A surgical knot was made with a surgical thread to prevent liquid leakage. The lavage was repeated three times. Cells in the lung lavage fluid were obtained by centrifugation, and the supernatant was collected to measure protein and inflammatory cytokine concentrations.
Cytokine Measurement
4.8
The collected procedures for Bronchoalveolar lavage (BALF) were described above. TNF‐α concentrations were analyzed in the BALF using the TNF‐α ELISA Kit (Biolegend, 430904) following the manufacturer's instructions.
Isolation of Mouse Lung Primary Cells
4.9
The mice were euthanized, and the lungs were dissected and dissociated as previously described [62]. After euthanasia, the mice were fixed on a surgical board. The chest cavity of the mice was opened, and the heart was exposed. The left atrium was excised, and the heart was perfused. A 10 mL disposable syringe was inserted into the right ventricle from the apex of the heart, and 10 mL of chilled (4°C) PBS solution was slowly and evenly injected to remove the blood from the body. Subsequently, 1 mL of tissue digestion solution—comprising 1 mg/mL collagenase (Sigma, C9891) and 2.2 mg/mL dispase (Roche, 47801700)—was instilled into the lungs via the trachea. After tying off the trachea, the lung tissue was removed and placed in a 15 mL centrifuge tube containing 1 mL of tissue digestion solution, and digested in a shaking water bath at 37°C for 30 min. After digestion, the lungs were carefully aspirated using a Pasteur pipette to obtain a single‐cell suspension of whole lung cells. The lung cells were washed, subjected to red blood cell lysis (Biolegend, 420301), and filtered through a 70 µm nylon mesh (BD). The lung cells were resuspended in RPMI 1640 medium (10% FBS and 1% penicillin/streptomycin), counted, plated in cell culture plates, and incubated for 24 h before further experiments.
Antibodies
4.10
The primary antibodies and the secondary antibodies used in the procedure include: Rabbit anti‐mouse TRPA1 (Proteintech, 19124‐1‐AP), Rabbit anti‐mouse CGRP‐I+CGRP‐II(Abcam, b283568), Rat anti‐mouse NCAM1 BV421 (BD, 78094), Rat anti‐mouse NCAM1 BV605 (BD, 809220), Rat anti‐mouse CD11b PEcy7 (Biolegend, 101216), Rat anti‐mouse F4/80 BV421 (BD, 565411), Rat anti‐mouse Ly6G AF488 (Biolegend, 127626), Rat anti‐mouse EpCAM PE (Biolegend, 118205), Rat anti‐mouse EpCAM APC (Biolegend, 118213), Rat anti‐mouse CD45 FITC (Biolegend, 123107), Rat anti‐mouse CD45 APCcy7 (Biolegend, 103116), Rat anti‐mouse Ly6C APC (Biolegend, 128016), Rat anti‐mouse Ly6C BV421 (Biolegend, 128031), Rat anti‐mouse TNF‐α PE (Biolegend, 506306), Rat anti‐mouse NCAM1 APC (Abcam, ab237383), Donkey anti‐rabbit IgG AF647 (Biolegend, 406414), Goat anti‐rabbit IgG AF700 (Invitrogen, A‐21038), Goat anti‐Rabbit CoraLite488 (Proteintech, SA00013‐2), Goat anti‐rabbit Cy3 (Servicebio, GB21303), Rabbit anti‐GAPDH (CST, 5174), Rabbit anti‐β‐Actin (CST, 4967), Goat anti‐rabbit IgG HRP(Jackson Immuno, 111‐035‐144).
Flow Cytometry
4.11
The detailed procedures were described previously [61]. The isolation of lung cells for flow cytometry followed the previously described steps. For flow cytometry staining of lung cells, the following sequential steps were performed: viability staining, surface staining, and intracellular staining. 0.1 µL of Zombie Yellow or Zombie NIR (Biolegend, 423106) was added to each sample to assess cell viability. The samples were then incubated at room temperature for 30 min, followed by two washes with FACS buffer and centrifugation at 1500 rpm. After preincubating for 15 min with anti‐mouse CD16/32 antibodies (Biolegend, 101320), single‐cell suspensions were stained for 30 min on ice with surface antibodies at a 1:100 dilution in FACS buffer, then washed twice. For intracellular staining of TNF‐α^+^ (BioLegend, 506306) and CGRP^+^ (Abcam, ab283568) cells, cells were fixed and permeabilized using the Fixation/Permeabilization Kit (BD, 554723) for 20 min at 500 × g, followed by incubation with intracellular antibodies (1:100 dilution) in Permeabilization Buffer for 30 min, and then washed twice. For CGRP staining, indirect labeling was performed using the secondary antibody Goat anti‐rabbit AF700 (Invitrogen, A‐21038). The cells were washed twice and resuspended in FACS buffer for analysis by flow cytometry. Debris and aggregates were excluded, and live cells were analyzed using an LSRFortessa flow cytometer (BD Biosciences). Data were analyzed using FlowJo software (Tree Star Inc).
The isolation of neurons from the vagal ganglia followed the same procedures as described earlier for sensory neuron isolation and culture. The staining process was similar to that of lung cells, including viability, surface, and intracellular staining, with a modification of the centrifugation speed to 800g.
RNA Isolation, Reverse Transcription, and RT‐PCR
4.12
Total RNA was isolated from cells or homogenized lungs and DRG using TRIzol reagent (Invitrogen, 15596018) according to the manufacturer's instructions. RNA was quantified and reverse‐transcribed to generate cDNA using a reverse transcriptase kit (Yeasen, 11141ES60), which was followed by quantitative real‐time polymerase chain reaction analysis. RT‐PCR was performed using the QuantStudioc 6 Flex system (ABI) with SYBRGreen Mix (Yeasen, 11203ES08). The fluorescence values were measured by the instrument to determine the cycle threshold (Ct). The 2−ΔΔCT method was used to analyze the target gene expression, and the Gapdh mRNA levels were used as an internal control for normalization in each sample. The primers used in the procedure include: Gapdh:5′‐CCCACTAACATCAAATGGGG‐3′(forward) and 5′‐CCTTCCACAATGCCAAAGTT‐3′(reverse); Cxcl1: 5′‐ACTGCACCCAAACCGAAGTC‐3′(forward) and 5′‐TGGGGACACCTTTTAGCATCTT ‐3′(reverse); Ccl2: 5′‐ TTAAAAACCTGGATCGGAACCAA‐3′(forward) and 5′‐ GCATTAGCTTCAGATTTACGGGT‐3′(reverse); Il6: 5′‐CCCCAATTTCCAATGCTCTCC‐3′(forward) and 5′‐CGCACTAGGTTTGCCGAGTA‐3′(reverse); Calca: 5′‐CCTTTCCTGGTTGTCAGCATCTTG‐3′(forward) and 5′‐CTGGGCTGCTTTCCAAGATTGAC‐3′(reverse); Calcb: 5′‐CTCTCAGCACGATATGGGTCC‐3′(forward) and 5′ ‐GCAAGAGATGTTTTTCCTGGTCG‐3′(reverse);
Immunofluorescence
4.13
Animals were euthanized by intravenous injection of Avertin (1.25%). The chest cavity of the animals was then opened for standard transcardial perfusion with PBS buffer for 5–10 min at room temperature until the blood was washed out, followed by perfusion with 4% paraformaldehyde (PFA) for 10–20 min (Servicebio, G1101). The lungs were dissected and post‐fixed overnight at 4 °C, while the DRG and nodose ganglia were fixed for 1 h at 4°C. These tissues were then incubated overnight at 4°C in 30% sucrose/PBS (Sigma, V900116), embedded in Tissue‐Tek OCT compound (Sakura), and frozen using liquid nitrogen. Tissues were sectioned into 10 to 30 µm slices using a cryostat. The slices were blocked with blocking solution (5% BSA, 0.1% Triton X‐100 in PBS) at room temperature for 1 h, followed by overnight incubation with primary antibodies at 4°C. After washing, the sections were incubated with secondary antibodies for 1 h. Following washing with PBS, the slides were mounted using DAPI mounting medium (Abcam,104139).
For cell experiments, cells were plated on coverslips. After stimulation, the culture medium was discarded, and cells were washed twice with PBS. Cells were fixed with 500 µL of 4% PFA (Servicebio) for 20 min. After two washes with PBS, cells were permeabilized with 0.1% Triton X‐100 for 15 min at room temperature. Blocking was performed with blocking solution (5% FBS, 0.1% Triton X‐100 in PBS) for 20 min. Cells were then incubated with primary antibodies overnight in a humidified chamber. After washing, cells were incubated with secondary antibodies at room temperature for 60 min. Following three washes with PBS, cell nuclei were stained with DAPI, and coverslips were mounted (Abcam, 104139).
Detecting Myd88 Oligomerization
4.14
We first transfected Neuro‐2a cells with Myd88‐GFP to establish the Neuro‐2a‐MyD88‐GFP cell line. Next, we transfected the Neuro‐2a‐MyD88‐GFP cells with Trpa1‐IRES‐mCherry or a control plasmid, Cas9‐IRES‐mCherry. After 30 min of LPS stimulation, we collected samples from Neuro‐2a cells overexpressing Trpa1 (*Trpa1^ov^ * ^er^) and control cells (Ctrl) and acquired data using fluorescence microscopy. We then used HALO software (Indica labs) to automatically count the number of MyD88‐GFP puncta larger than 0.5 µm^2^ per individual cell.
Ganglia Clearing by CUBIC
4.15
Advanced CUBIC protocols for tissue clearing were introduced as described previously [63]. Briefly, mice were anesthetized and transcardially perfused with cold PBS to remove the blood from the organs as much as possible. Subsequently, they were perfused with 4% PFA. The ganglia were dissected and fixed in 4% PFA, with shaking at 4°C for 1 h. After washing with PBS three times, the samples were immersed in Reagent‐1 diluted with 1/2 water (25 wt.% urea, 25 wt.% Quadrol, 15 wt.% Triton X‐100) and shaken at 37°C for 2 h. The 1/2‐diluted Reagent‐1 was discarded. Reagent‐1 was replaced, and primary antibodies were added, then gently shaken at 37°C for 1 day. After that, the samples were incubated overnight with secondary antibodies in Reagent‐1. The samples were degassed using a vacuum desiccator. Then, the samples were immersed in Reagent‐2 diluted with approximately 1/2 PBS (25 wt.% urea, 50 wt.% sucrose, 10 wt.% triethanolamine), shaken at 37°C for 2 h, and then treated in Reagent‐2 for one day. Clear ganglia were imaged using a confocal microscope (Olympus).
Vagus Nerve Recording
4.16
The vagus nerve recordings were performed as described previously [33]. Briefly, mice were induced under anesthesia with 2.5% isoflurane in 100% air at a flow rate of 0.9 L/min for 3 min. Anesthesia was then maintained with 2% isoflurane during surgery. Subsequently, the mice were maintained under 1.75% isoflurane and placed on a heating pad to keep body temperature at approximately 37°C for vagus nerve signal recording. To isolate the cervical vagus nerve, the neck area was cleaned with iodine, and a midline cervical incision was made from the level of the larynx to the sternum. Submandibular salivary glands and the muscles on the trachea were separated to expose the trachea. A small dosing tube was inserted into the lung by perforating the trachea with a 29G needle and placing the dosing tube approximately 7 mm deep into the trachea. The right cervical vagus nerve was carefully separated from the artery and desheathed by gently removing the thin connective tissue surrounding the nerve under magnification. The vagus nerve was then placed on two hook electrodes with mineral oil to insulate the nerve and the surrounding tissue. A ground electrode was inserted into the mouse's tail.
The PowerLab 4/26 data‐acquisition system (PowerLab; ADINSTRUMENTS, Inc.) was used to record signals from the vagus nerve. Recordings were sampled at 40 kHz, filtered at 120 Hz, and amplified 100 times to ensure the signals were stable enough for long‐term recording. Following the acquisition of baseline activity (10 min), 10 µL of LPS (10 mg/mL) or PBS (as a control) was injected into the lung via the dosing tube, and recordings continued for 20 min post‐injection. The analysis process was previously described [33]. In brief, the signal in mat format was exported from LabChart (LabChart; ADINSTRUMENTS, Inc.) and analyzed in MATLAB. The signals were decomposed into individual spikes, and the action potentials were detected using a threshold determined by a smallest of constant false‐alarm rate (SO CFAR) filter. The t‐SNE method was used to perform dimensionality reduction of all spike data, and the DBSCAN method was used to perform cluster analysis of all spikes. Then, all spikes were clustered into different groups based on their waveforms. All significant waveforms were counted, and their distribution over time was analyzed after normalization.
Calcium Imaging and Analysis of the Jugular‐Nodose Ganglia
4.17
AAV9‐CMV‐GCaMP6s was injected into the jugular‐nodose ganglion of a 4‐week‐old mouse to express GCaMP6s in the neurons of the ganglion. After a 4‐week incubation period, the animals underwent a calcium signal recording procedure. Mice were administered isoflurane anesthesia via a mask with an airflow rate of 0.8 L/min at a 2% isoflurane concentration. They were placed on a heating pad to maintain body temperature. To isolate the nodose ganglia, the neck area was disinfected with iodine, and a midline cervical incision was made from the level of the larynx to the sternum. Initially, the vagal nerve was isolated, and a dosing tube was inserted into the trachea, as previously described. Subsequently, two blunt hooks were used to detach the muscles and expose the jugular‐nodose ganglia. A pair of micro dissecting spring scissors was then used to excise the nodose on the rostral side. Once separated, PBS was used to keep the nerve and ganglia moist. The isolated ganglia were placed on a platform made from a coverslip, held in place by a magnetic stand. The miniscope was also held by a magnetic stand with a lifting rod to adjust the distance between the lens and the ganglia [64]. The miniscope was adjusted to capture 20 images per second (20 fps), and recording was initiated. After approximately 3 min of baseline ganglia recording, 25 µL of LPS (10 mg/mL) or PBS was injected into the lung through the dosing tube. Recording was continued for about 5–7 min post‐stimulation.
All raw video data were analyzed using the open‐source analysis pipeline named Calcium Imaging Analysis (CaImAn) in Matlab [65]. The fluorescence intensity of the region of interest in the videos was extracted using ImageJ. The first 3 min of the videos were considered as the baseline for GCaMP6s. Fold change calculations based on the baseline fluorescence intensity were performed, and the data was ultimately plotted in the form of a heatmap.
Genetic Ablation of αCGRP+ Neurons in NG
4.18
5‐week‐old *DTR^CalcaCreERT2^
- mice or DTR ^fl/fl^ mice (as controls) were intraperitoneally injected with 100 µL of tamoxifen (Sigma) (20 mg/mL in corn oil) for 5 consecutive days. After two weeks, 20 ng DT in 120 nL PBS was injected into nodose ganglion with nanoinjector (Drummond Scientific Company). Mice were anesthetized with 2.5% isoflurane in air and maintained at 2.0% isoflurane throughout the surgery (RWD). The vagal ganglion was exposed after a midline incision in the neck. Waiting for 2 weeks recovery, the mice were performed experiments.
AAV Microinjection and AAV‐Guided Trpa1 Knockout in JNC
4.19
To knock out Trpa1 in JNC neurons, we generated pX601‐mCherry(sgTrpa1), which contains two U6‐sgRNA units targeting Trpa1 (sgTrpa1‐1: GCTTAACGATGTCAGTGGCTCC; sgTrpa1‐2: GTATACCGGAAGTAGTGATATT). The control plasmid contains two empty U6‐sgRNA units (sg‐empty: GGAGACCACGGCAGGTCTCAG). These vectors were packaged into AAV‐PHP.S serotype with hsyn‐driven saCas9 (pXN‐hsyn‐saCas9) to generate AAV‐Cas9 and AAV‐sgRNA. The AAV was diluted to 10^12^ vg/mL after titration. Five‐week‐old mice were anesthetized with isoflurane, and their limbs were immobilized. A 1 cm incision was made in the ventral neck area skin. Under a dissecting microscope, the fascia and adipose tissue were bluntly dissected, taking care to avoid damaging surrounding blood vessels, to expose the vagal ganglia clearly. Using a Nanoject III Injector (Drummond), 120 nL of AAV‐Cas9 and 120 nL of AAV‐sgRNA was slowly injected into the vagal ganglia. The injector needle was left in place for an additional 5 min before slow withdrawal, allowing viral particles to diffuse and be absorbed at the injection site.
AAV Microinjection and AAV‐Guided CGRP Knockout in JNC
4.20
To knock out Calca/Calcb in JNC neurons, we generated pX601‐mCherry(sgCalca/Calcb), which contains two U6‐sgRNA units targeting Calca/Calcb (sgCalca/Calcb‐1:CACCGGCAGTGTTGCAGGATCTCTT/AAACAAGAGATCCTGCAACACTGCC; Calca/Calcb‐2: CACCGCTGAGCAGATCAGGAGGTGTG/AAACCACACCTCCTGATCTGCTCAGC). The control plasmid contains two empty U6‐sgRNA units (sg‐empty: GGAGACCACGGCAGGTCTCAG). These vectors were packaged into AAV‐PHP.S serotype with hsyn‐driven saCas9 (pXN‐hsyn‐saCas9) to generate AAV‐Cas9 and AAV‐sgRNA. The AAV was diluted to 10^12^ vg/mL after titration. Five‐week‐old mice were anesthetized with isoflurane and their limbs were immobilized. A 1 cm incision was made in the ventral neck area skin. Under a dissecting microscope, the fascia and adipose tissue were bluntly dissected, taking care to avoid damaging surrounding blood vessels, to expose the vagal ganglia clearly. Using a Nanoject III Injector (Drummond), 120 nL of AAV‐Cas9 and 120 nL of AAV‐sgRNA was slowly injected into the vagal ganglia. The injector needle was left in place for an additional 5 min before slow withdrawal, allowing viral particles to diffuse and be absorbed at the injection site.
Short Hairpin RNA (shRNA)–Mediated RNA Interference
4.21
Short hairpin RNA (shRNA)‐mediated RNA interference was employed, with shRNA sequences designed based on the target gene and synthesized as corresponding stem‐loop primers (Sangon Biotech). These primers were annealed and inserted into the pLKO.1 plasmid between the EcoRI and NheI restriction sites (CST). The shRNA pLKO.1 construct was introduced into competent E. coli DH5α (Tsingke) for plasmid amplification and extraction. The resulting plasmid was then co‐transfected with VSV‐G and Δ8.9 plasmids into 293T cells. After culturing, the produced lentivirus was used to infect PNEC cells, and the infected cells were selected with puromycin to establish stable cell lines. The knockdown assay primer sequences were Scramble CAACAAGATGAAGAGCACCAA; Prkaca CGAGTAACTTTGACGACTATG.
CGRP Release of SCLC Cultures and Plasma
4.22
SCLC cells were cultured in a 12‐well plate (2 × 10^5^ cells per well) following the above‐mentioned method for one day. The cells were stimulated with 100 nm CGRP (Phoenix pep) for 30 min. After stimulation, the media was aspirated, and the cells were washed three times with fresh media. The CGRP concentration in 100 µL of culture supernatant was measured using a CGRP EIA Kit (Cayman Chemicals).
For CGRP release in plasma, blood was collected from mice and centrifuged at 500 × g for 10 min. 100 µL of plasma was collected, and the CGRP concentration was measured using a CGRP EIA Kit (Cayman Chemicals).
Western Blot
4.23
The samples were collected by scraping as described above. After quantification by the BCA protein assay (Thermo Scientific, 23225), equal amounts of protein were mixed with loading buffer and boiled at 100°C in a water bath for 10 min. The prepared protein samples were loaded onto SDS‐PAGE gels for electrophoresis. Initially, electrophoresis was performed at 80 V for 30 min for gel concentration, followed by adjusting the voltage to 120 V until the ideal separation was achieved. For protein transfer, a wet transfer method was employed to transfer the proteins from the gel to a PVDF membrane (Millipore, IPVH00010). The transfer was conducted at 200 mA for 60 min. After transfer, the membrane was blocked with 5% skim milk (BD, 232100) at room temperature for 1 h, followed by three washes with TBST. Subsequently, the membrane was incubated with the primary antibody overnight at 4°C. After the primary antibody incubation, the membrane was incubated with the secondary antibody, which was an HRP‐conjugated antibody (Jackson Immuno), at room temperature for 1 h. Following the secondary antibody incubation, the membrane was washed three times with TBST, each for 10 min, and then subjected to chromogenic detection using an ECL substrate (Yeasen, 36208). Data were collected using a Tanon chemiluminescence imaging system. The bands were analysed and quantitated using Image J software.
Single‐Cell Isolation of Vagal Sensory Neurons
4.24
Vagal sensory neurons were isolated as previously described [66]. Briefly, Ganglion Dissociation Solution (GaDS) was prepared by combining Advanced DMEM/F12, Glutamine, HEPES, N2 supplement, B27, and NGF. Mice were anesthetized with avertin via intraperitoneal injection. Following euthanasia, the vagal ganglia were dissected out, and two local cuts were made to facilitate dissociation. The ganglia were transferred into a centrifuge tube (ganglia from 3 mice were combined into one sample to ensure flow cytometric analysis). The ganglia were washed three times with PBS, allowed to settle at the bottom of the tube, and then the PBS was removed. Next, 1 mL of GaDS and 22 µL of digestion enzymes (Roche, 5401054001; 2.5 mg/mL) were added to the tube. The tube was then incubated at 37°C for 30 min with gentle shaking every 15 min and tapping it lightly. After digestion, the GaDS solution was removed, and the ganglia were washed twice with 1 mL of DPBS. Subsequently, 200 µL of GaDS was added to the tube, and the solution was pipetted up and down until no intact tissue pieces were visible, while avoiding bubble formation. The dissociated cells were passed through a 70 µm cell strainer. Carefully, 300 µL of cells were layered onto the percoll density gradient and centrifuged at 2900 × g for 10 min. After centrifugation, the top 700 µL of solution containing a large amount of cell debris was removed and discarded. Then, 700 µL of fresh GaDS was added to the remaining cell‐containing solution, and the solution was pipetted up and down several times to mix. After centrifuging at 2900 × g for 15 min, the supernatant was carefully removed and discarded, leaving the cell pellet. Finally, the cells were resuspended in 300 µL of GaDS, and the sample was kept on ice for further experiments.
Retrograde Tracing of the Vagal Subgroups Projecting to the Lung
4.25
Fast Blue Retrograde Tracing was performed as previously described [66]. Briefly, anesthetize mice with an intraperitoneal injection of avertin. Place the anesthetized mouse on a 45° inclined platform with a heating pad and suspend the mouse by its incisors on a fixed line on the platform to maintain a supine position. Position a cold light source above the trachea. Gently grasp the mouse's tongue with flat‐tipped forceps and slightly move the tongue to expose the translucent opening of the trachea. A 22 G blood collection needle was inserted into the trachea to a depth of 0.5–1 cm, then withdrawn. Subsequently, 40 µL of Fast Blue solution (0.4 mm; Polysciences, 17740‐2) was slowly instilled through the needle. With each breath, the mouse will slowly inhale the Fast Blue solution into its lungs. To prevent the dripping material from overflowing from the trachea, keep the blood collection needle in place for 10 s before withdrawing it, and continue to suspend the mouse on the inclined surface for at least 1 min. If respiratory arrest occurs, chest massage can be performed for cardiopulmonary resuscitation. After the procedure, place the mouse in a supine position with its head raised on the heating pad for recovery. Once the mouse has regained mobility, return it to its original cage. Three days later, the Fast Blue tracer will have reached the ganglia, enabling further experiments such as neural isolation or immunofluorescence. The dye will remain visible for up to 8 weeks.
RNA‐seq Library Preparation, Sequencing, and Analysis
4.26
After euthanizing the mice, the vagal ganglia were dissected and isolated. RNA from the vagal ganglia was extracted using The RNeasy Mini Kit (Qiagen, 74106). The ganglia were placed in a micro glass homogenizer containing Buffer RLT and carefully homogenized to release the RNA. Subsequently, RNA was extracted following the protocol outlined in the RNeasy Mini Kit manual. The purity and concentration of the RNA were measured, with the following standards: RIN > 7.0, 28S/18S > 1.0, m > 20 ng. For samples containing less than 4 µL of RNA, Oligo‐dT Primer was added, and the first‐strand cDNA was synthesized using the SMART amplification technology. The second‐strand cDNA synthesis was performed using KAPA HiFi HotStart DNA Polymerase. The synthesized double‐stranded cDNA was purified using Agencourt AMPure XP magnetic beads, and the quality of the amplification product was assessed using an Agilent 2100 chip. Qualified amplification products were subjected to library construction using the Nextera XT DNA Library Preparation Kit. The qualified libraries were transformed into single‐stranded circular molecules under certain conditions. The single‐stranded circular DNA molecules were replicated by rolling circle replication, forming DNA nanoballs (DNBs) containing over 200 copies of DNA. The obtained DNBs were loaded into the mesh holes on the high‐density DNA nanochip using high‐density DNA nanoball array technology. Sequencing was performed using the combined probe anchor sequencing (cPAS) technology, with a sequencing read length of PE100. Library construction and sequencing were conducted at BGI (Beijing Genomics Institute). Data analysis was performed using R and BGI platform.
Statistical Analysis
4.27
All statistical analyses were performed using R and GraphPad Prism 8.0. Summary data were compared for statistical differences using t‐tests or one‐way ANOVA. Statistical data were presented as means ± SEM. If no statistical significance was observed, “ns” was used. If there was a statistically significant difference, ^^ p < 0.05 indicates a significant difference; ^^ p < 0.01 indicates a highly significant difference; ^^ p < 0.001 indicates an extremely significant difference.
Author Contributions
J.C. and X.S. designed the experiments. J.C. and X.S. contributed to the conceptualization. J.C., S.X., Z.L., C.Z., R.T., C.Z., X.C., Y.Z., Y.M., and R.W. contributed to methodology. J.C., S.X., Z.L., and C.Z. carried out the investigation. J.C., S.X., and S.X. performed formal analyses. J.C. and X.S. contributed to Visualization and funding acquisition. Q.H., H.S., S.W., X.Z., P.S., H.J., Y.S., and X.S. provided resources. X.S. contributed to the supervision. J.C. was responsible for writing the original draft. X.S. was responsible for writing, reviewing, and editing. All authors reviewed the final version of the manuscript and agreed with its content and submission.
Funding
This work was supported by NSFC programs (82241042, 82495200/02, 82400103); National Key Research and Development Program of China (No.2022YFC2304702); Science and Technology Commission of Shanghai Municipality (20DZ2261200); China Postdoctoral Science Foundation (2024M750522).
Conflicts of Interest
The authors declare no conflicts of interest.
Supporting information
Supporting File 1: advs71699‐sup‐0001‐SuppMat.docx.
Supporting File 2: advs71699‐sup‐0002‐VideoS1.mp4.
Supporting File 3: advs71699‐sup‐0003‐VideoS2.avi.
The reference list from the paper itself. Each links out to its DOI / PubMed record.
- 1M. S. Schappe , P. A. Brinn , N. R. Joshi , et al., “A Vagal Reflex Evoked by Airway Closure,” Nature 627 (2024): 830–838.38448588 10.1038/s 41586-024-07144-2PMC 10972749 · doi ↗ · pubmed ↗
- 2W. Han and I. E. de Araujo , “Dissection and Surgical Approaches to the Mouse Jugular‐Nodose Ganglia,” STAR Protocols 2 (2021): 100474.33997807 10.1016/j.xpro.2021.100474 PMC 8091130 · doi ↗ · pubmed ↗
- 3L. Bai , S. Mesgarzadeh , K. S. Ramesh , et al., “Genetic Identification of Vagal Sensory Neurons That Control Feeding,” Cell 179 (2019): 1129–1143.e 23.31730854 10.1016/j.cell.2019.10.031PMC 6916730 · doi ↗ · pubmed ↗
- 4J. Chen , X. Lai , Y. Song , and X. Su , “Neuroimmune Recognition and Regulation in the Respiratory System,” European Respiratory Review: An Official Journal of the European Respiratory Society 33 (2024), 240008.38925790 10.1183/16000617.0008-2024 PMC 11216688 · doi ↗ · pubmed ↗
- 5S. L. Prescott and S. D. Liberles , “Internal Senses of the Vagus Nerve,” Neuron 110 (2022): 579–599.35051375 10.1016/j.neuron.2021.12.020PMC 8857038 · doi ↗ · pubmed ↗
- 6Y. Huang , C. Zhao , and X. Su , “Neuroimmune Regulation of Lung Infection and Inflammation,” QJM: An International Journal of Medicine 112 (2019): 483–487.30016504 10.1093/qjmed/hcy 154 · doi ↗ · pubmed ↗
- 7H. Jin , M. Li , E. Jeong , F. Castro‐Martinez , and C. S. Zuker , “A Body–Brain Circuit that Regulates Body Inflammatory Responses,” Nature 630 (2024): 695–703.38692285 10.1038/s 41586-024-07469-y PMC 11186780 · doi ↗ · pubmed ↗
- 8E. Granton , L. Brown , M. Defaye , et al., “Biofilm Exopolysaccharides Alter Sensory‐Neuron‐Mediated Sickness During Lung Infection,” Cell 187 (2024): 1874–1888.e 14.38518773 10.1016/j.cell.2024.03.001 · doi ↗ · pubmed ↗