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date: 02 December 2021

Development of Lung Innervationfree

Development of Lung Innervationfree

  • Talita de Melo e Silva, Talita de Melo e SilvaDepartment of Pathology, The Ohio State University
  • Catherine Miriam CzeislerCatherine Miriam CzeislerDepartment of Pathology, The Ohio State University
  •  and José Javier OteroJosé Javier OteroDepartment of Pathology, The Ohio State University


Breathing is essential for survival and is precisely regulated by the nervous system. From a neuroanatomical perspective, the respiratory tract is innervated by afferent and efferent autonomic nerves, which regulate aspects of airway function and ensure appropriate tissue oxygenation. The general concepts of how the peripheral nervous system (PNS) develops as it relates to lung function are reviewed. The vagus (cranial nerve X), a mixed motor and sensory nerve, supplies parasympathetic and sensory fibers to the airways. During development, preganglionic visceromotor efferent neurons of the cranial nerves arise in the hindbrain basal plate and later migrate dorsally through the neuroepithelium.

The neural crest is a migratory and multipotent embryonic cell population that develops at the dorsal portion of the neural tube, which delaminates from the neuroepithelium to enter distinct pathways, forming various derivatives, among which include the peripheral nervous system. Neural crest cells emerging from the vagal region migrate into the ventral foregut and give rise to intrinsic ganglia in the respiratory tract that are innervated from the vagus and send out postganglionic fibers.

The lung is innervated by sympathetic nerves derived from the upper thoracic and cervical ganglia. The sympathetic preganglionic neurons are derived from trunk neural crest cells that migrate, forming two chains of sympathetic ganglia referred to as the lateral vertebral sympathetic chains. Neural crest cells that migrate along defined pathways to generate sympathetic ganglia also derivate the dorsal root ganglia that send somatosensory afferent innervations to the respiratory tract.


  • Neuroendocrine and Autonomic Systems


The cardiorespiratory organs represent an integrated interoceptive system whose proper function permits life in vertebrates. Significant insights into this integrated unit have been generated from understanding the neural control of respiratory rhythm, frequency, and tidal volume, all of which ultimately are regulated through neural control of diaphragmatic contraction. However, diaphragmatic contraction is only one phase of respiration, the others being principally airflow and blood perfusion through lung tissue. Airflow and blood perfusion modulation occur through neural stimulation of smooth muscle circumferentially localized around bronchioles and arterioles. The respiratory tract innervations mediate basic physiological functions in respiratory defense and breathing control.

Neuroanatomy of Lung Innervation

Vagal Efferents and Sensory Afferents Innervating the Respiratory Tract

The respiratory tract is innervated by efferent and afferent autonomic nerves, which regulate many aspects of airway function (see Figure 1 for schematics). Early in development, the neural progenitor cells at the rostral region of the neural tube expand and develop into swellings, forming the primary brain vesicles: forebrain (prosencephalon), midbrain (mesencephalon), and hindbrain (rhombencephalon). The most caudal region of the neural tube elongates to form the spinal cord (Stiles & Jernigan, 2010; Swanson, 2000). The cranial motor neuron cell bodies reside in the midbrain and hindbrain, and their axons extend through the cranial nerves to control head and neck muscles. The vagus (cranial nerve X) is a mixed motor and sensory nerve that contains A, B, and C fibers (so named based on their extent of myelination, with A being the most myelinated and C being predominantly unmyelinated, as defined by Erlanger and Gasser) and innervates organs in the thorax and abdomen. This nerve supplies parasympathetic and sensory fibers to the airways (Yuan & Silberstein, 2016).

Figure 1. Schematic of brainstem innervation of lung. Visceral afferents (left, red circuit) are sensory fibers whose cell bodies are located in the nodose and jugular ganglia and that project to the nucleus of the solitary tract (NTS). NTS neurons project to multiple brainstem, midbrain, and forebrain neurons to modulate central control of respiration. The efferent pathway (right side, blue circuit) has cell bodies originating in the dorsal motor nucleus of the vagus nerve (DMNV) and nucleus ambigus (NA). These project through the vagus nerve, with some fibers entering the lung, whereas others synapse onto postganglionic fibers located on the trachea and bronchus (green, postganglionic parasympathetic fibers.).

The parasympathetic innervations to the respiratory tract are composed of preganglionic neurons, also referred to as extrinsic neurons, whose cell bodies are located outside of the respiratory tract in the medulla, and postganglionic neurons, known as intrinsic pulmonary neurons, whose cell bodies reside in the trachea and bronchi, where they cluster to form ganglia (Larsell, 1922; Tollet et al., 2001). The vagus nerve innervates airway smooth muscles (ASMs) and neuroendocrine bodies (NEBs) in the lung epithelium (Adriaensen et al., 1998; Van Lommel et al., 1998; Wasano, 1977; Weichselbaum et al., 1996). Stimulation of vagal efferents provides postganglionic parasympathetic input and leads to bronchoconstriction (Nadel, 1974; Roddie, 1970).

Vagal sensory innervation from neurons of the nodose and jugular ganglia conducts afferent signals arising from the lung and airways to the nucleus tractus solitarii (NTS) in the medulla that can adjust autonomic neural regulation to the lung through the parasympathetic or sympathetic systems (Widdicombe, 1981). Kupari et al. (2019) identified the diversity of neuronal types of jugular and nodose ganglion complex through interrogation of single-cell RNA sequencing. They identified fundamental differences between neurons of these ganglia, among them that the activation of jugular neurons generates viscerosensory functions. However, the nodose neurons are associated with the sensory vagus nerve and the activation of the autonomic nervous system (Kupari et al., 2019). Other important functions of these neurons include initiating defense reflexes that protect the lung from particulates and chemical irritants (Coleridge, 1986; Paintal, 1973).

The afferent fibers are classified in stretch-sensitive mechanosensors based on adaptation to sustained lung distension. These include rapidly adapting receptors (RARs) that respond rapidly to sustained lung distension and slowly adapting receptors characterized by slower responses to lung distension. These stretch-sensitive fibers are located in the intrapulmonary tissues and can be activated by lung inflation. RARs also respond to lung deflation. The depth and rate of breathing and also lung volume and compliance can determine the pattern of action potential discharge of these fibers.

Vagal C fibers also provide afferent signals and innervate the entire respiratory tract, from the lung parenchyma to large conducting airways. Based on their anatomic distribution, vagal C fibers are classified as bronchial and pulmonary, and the cell bodies reside at the nodose and jugular ganglia. The bronchopulmonary C fibers are sensitive to mechanical and chemical stimuli and consist of unmyelinated fibers with slower rate conduction of action potentials than the stretch-sensitive mechanosensors. The stimulation of these afferents can elicit reflex responses such as bronchoconstriction and submucosal gland secretion and change the rate and depth of breathing (Coleridge & Coleridge, 1984; Lee & Pisarri, 2001; Lee et al., 2003).

In 2015, Chang and collaborators classified sensory neurons in the vagus nerve based on receptor expression and effects on breathing. They identified a population of vagus nerve afferents that express purinergic receptor type 1 (P2ry1) and are largely fast-conducting A fibers that innervate the neuroepithelial bodies. In addition, they showed that stimulation of these neurons by optogenetic approach acutely silences respiration. The other neuronal population identified expresses neuropeptide Y receptor type 2 (Npy2r) and is largely slow-conducting C fibers that innervate the alveoli-containing respiratory region of the lung and, when stimulated, cause rapid shallow breathing (Chang et al., 2015).

Sympathetic Innervation

The sympathetic preganglionic neurons have their cell bodies in the thoracic and lumbar regions of the spinal cord and reach the sympathetic chain of ganglia by passing through the ventral roots (see Figure 2 for schematic). Sympathetic neurons are derived from trunk neural crest cells that migrate ventrally, lying adjacent to the dorsal surface of the aorta at approximately E10.5 in the rat (Cochard et al., 1979; Rubin, 1985a; Yntema & Hammond, 1947). The migrating cells form the two chains of sympathetic ganglia on either side of the vertebral column, ultimately forming the lateral vertebral sympathetic chains (Rubin, 1985b). The presence and distribution of adrenergic fibers innervating the bronchial smooth muscle differ between species. Nevertheless, beta-adrenergic receptors are abundantly expressed on airway smooth muscle, and activation of these receptors by circulating catecholamines mediates bronchodilation (Doidge & Satchell, 1982; El-Bermani, 1978). The adrenergic stimulation of the airway vasculature induces constriction (Matran, 1991).

Figure 2. Schematic of spinal cord innervation of the lung. Sensory fibers (left, red circuit), composed of pseudo-unipolar neurons, possess cell bodies in the dorsal root ganglia (DRG). Sympathetic fibers exit the spinal cord in the ventral root and synapse onto the sympathetic chain (postganglionic sympathetic neurons), which then project to the lung.

Sensory Innervation

In addition to the vagal sensory innervations, the respiratory tract receives somatosensory afferent innervation from dorsal root ganglia (DRG) (Springall et al., 1987). The neural crest cells that migrate along defined pathways to generate sympathetic ganglia also derivate the DRG. DRG are pseudo-unipolar neurons having two branches that act as a single axon connected by a cell body. Pulmonary DRG cell bodies are located between the levels of T1 and T6. These fibers innervate the lungs and bronchi and are primarily responsible for the transduction of sensory information from the periphery to the central nervous system (CNS) to maintain the homeostasis regulating the depth and rate of breathing, bronchomotor tone, and airway secretion (Haberberger et al., 2019; Lee & Yu, 2014).

Other unique components of the pulmonary autonomic nervous system are pathways within the airways and lung, which are neither adrenergic nor cholinergic, the so-called nonadrenergic, noncholinergic (NANC) system. NANC transmissions are mediated from classic parasympathetic or sympathetic nerves, use different molecules and neuropeptides as neurotransmitters, and modulate airway smooth muscle tone either through relaxation by inhibitory or contraction by excitatory NANC activity. The bronchodilation mediated by inhibitory NANC (i-NANC) has been demonstrated in vivo. Diamond and O‘Donnell showed in 1980 that under muscarinic receptor blockade, stimulation of efferent vagal nerves induces a pronounced bronchodilatation in cats. Bronchodilation was not affected by beta- and alpha-adrenergic blockade, supporting the idea of a nonadrenergic inhibitory system in the pulmonary airways of the cat. They also showed that bronchodilatation was abolished by autonomic ganglionic blockade with hexamethonium, indicating that NANC nerves have pre- and postganglionic fibers (Diamond & O‘Donnell, 1980). The i-NANC neurotransmission is mediated due to the release of peptides such as vasoactive intestinal peptide and peptide histidine methionine (Barnes, 1986; Carstairs & Barnes, 1986; Palmer et al., 1986).

Excitatory NANC mechanisms produce bronchoconstrictor responses. In guinea pigs, vagus nerve stimulation elicits a NANC bronchoconstrictor in the presence of atropine due to the release of peptide neurotransmitters from a subpopulation of nonmyelinated sensory C fibers in the smooth muscle layer of the respiratory tract (Lundberg & Saria, 1982). The principal peptides for this process are neurokinins and calcitonin gene-related peptide (Boni et al., 1995; Lundberg et al., 1983). NANC innervation plays an important role in bronchomotor tone maintenance and regulation by effects resulting from specific neuropeptides or by regulating the autonomic tone in airway smooth muscle.

Cranial Motor Neurons and Vagus Nerve Development

The rhombencephalon arranges into rhombomeres, morphologically discernable as segmental swellings. Each segment can be identified by unique regulatory gene expression patterns that define a specific embryonic progenitor domain. There is an organized correlation between the rhombomeres, their specific neuronal populations, and their axon projections (Astick et al., 2014; Lumsden & Keynes, 1989; Lumsden et al., 1991). Motoneurons of the branchiomotor series, including the trigeminal (V), facial (VII), glossopharyngeal (IX), and vagus (X), have bimeric origins. For instance, V derives from r2-r3, VII derives from r4-r5, IX derives from r6-r7, and X derives from r7-r8 (reviewed by Bass & Baker, 1997). These branchiomotor neurons are derived from Nkx2.2-expressing neural progenitors (Alzate-Correa et al., 2021; Jarrar et al., 2015). The vagus motor axons supply parasympathetic and sensory innervations to the airways (Yuan & Silberstein, 2016). This occurs respectively through visceral motor (VM) and visceral sensory (VS) fibers. Early in development, preganglionic, visceromotor, and branchiomotor efferent neurons of the cranial nerves arise in hindbrain basal plate and later migrate dorsally through the neuroepithelium (reviewed by Guthrie, 2007).

Compelling evidence suggests that neural crest (NC) cells emerge through continuous induction at the neural plate border (Baker & Bronner-Fraser, 1997). These migratory embryonic cells develop at the dorsal portion of the neural tube and delaminate from the neuroepithelium to enter distinct migratory pathways throughout the embryo to form a wide range of derivatives, including the peripheral nervous system (PNS) (Baker & Bronner-Fraser, 1997). At mouse embryonic period 11 (E11), the vagus nerve and associated neural crest cells are already projecting into the lung buds. At this period, nerve fibers may be found extending from the vagus to the trachea, bronchi, and pulmonary arteries. Tollet et al. (2002) showed that by E13, it is possible to observe intrinsic ganglia in paratrachial tissues, and that by E14, ganglia at the junction of bronchi, with full connections to the vagus network, can be observed.

In relation to the embryonic origin of vagal sensory complex, the jugular ganglia are derived from neural crest and nodose ganglia derived from placodes (D’Amico-Martel & Noden, 1983; Nassenstein et al., 2010). Transient placodes are thickened columnar epithelia in the cephalic region of vertebrate embryos. During the development, vagal or nodose epibranchial placode develops above the third branchial cleft, generating neurons in the nodose ganglia (Baker & Bronner-Fraser, 1997; Lee et al., 2001).

Neural Crest Cells

The three primary germ layers include ectoderm, mesoderm, and endoderm and are generated during gastrulation, resulting a structure known as the trilaminar embryonic disk. The ectoderm ultimately develops into two critical structures: the surface ectoderm (which will generate placodes, dermis, and skin), and the neural ectoderm (which will generate the nervous system). The neural ectoderm, also known as neuroepithelium or neural plate, develops in the midline of the trilaminar embryonic disk and extends nearly the entire axis of the embryo. It can be visualized as swellings that emerge, which then fuse dorsally across the midline to generate the neural tube (a process known as neurulation). Neural crest cell generation occurs at the dorsal most part of the embryo at the interface of the neuroepithelium and surface ectoderm, known as the neural plate border. Once formed, neural crest cells undergo an epithelial–mesenchymal transition, delaminate from the neuroepithelium, and begin their migration to distinct locations in the embryo. Although evidence of conserved molecular mechanisms of neural crest cell delamination between avians and mammals is unknown, it is presumed that a combination of intrinsic DNA regulatory proteins, known as transcription factors with extracellular morphogens and growth factors, are required for neural crest induction. Many elegant studies have shown that the intrinsic innervation of the lung is derived from neural crest (Burns & Delalande, 2005; Burns et al., 2008; Freem et al., 2010; Langsdorf et al., 2011). The intrinsic neurons of the lung show their cell bodies placed along the smooth muscle strip and the major bronchi. These cuboidal cell bodies can be identified at low magnification in stereomicroscopes and predominate around the tracheal bifurcation (Weigand & Myers, 2010). During the development of the respiratory tract, vagal neural crest cells migrate into the ventral foregut to give rise to neural tissue (Langsdorf et al., 2011).

Vagal Neural Crest Cells and the Development of Lung Intrinsic Innervation

The respiratory tract, trachea, and lungs are derived from endoderm and develop as outgrowths of the foregut. The primitive foregut divides into the esophagus and trachea. In the respiratory region, a pair of primary lung buds initiates a process of growth into the surrounding mesenchyme, forming tubules that extend and branch to give rise to the bronchial tree (Schittny, 2017; Spooner & Wessells, 1970; Ten Have-Opbroek, 1981). A beautiful manuscript published by Tollet et al. in 2001 provides insight into the development and growth of the fetal mouse lung. They showed that the lung buds begin to evaginate from the foregut at E9.5 and that at this period, the lung consists of the future trachea and two main stem epithelial tubules, with buds developing laterally (Tollet et al., 2001). The vagal neural crest cells, in contrast, initiate their development at the neural tube–somite boundary and utilize, for at least part of their journey, the vagus nerve as a key migratory pathway to reach their end organs (Sadaghiani & Thiebaud, 1987).

Following migratory exit from the vagus nerve, it has been postulated that neural crest migrate in a rostro-caudal direction within the developing foregut simultaneously with the outgrowth of lung buds (Burns & Delalande, 2005; Burns & Le Douarin, 2001; Le Douarin & Teillet, 1973). These cells later form ganglia and send out postganglionic fibers that follow the tubules to the base of the epithelial buds to innervate the airway smooth muscle in chick (HH20), mouse (E10.5), and human (E8–10) (Burns & Delalande, 2005; Burns et al., 2008; Freem et al., 2010; Sparrow & Lamb, 2003; Sparrow et al., 1999; Tollet et al., 2001). Tollet et al. (2001) evaluated the structural organization of nerves and smooth muscle in relation to the branching epithelial tubules of the primordial mouse lung. Migrating neural crest cells can be identified by E11 in fetal lung using the neural crest marker p75NTR (the low-affinity neurotrophin receptor), neuronal marker PGP 9.5 (protein gene product 9.5), and synaptic vesicle protein (synapsin) antibodies. They observed vagus processes positive for p75NTR entering to the lobes of the lung, demonstrating that neural crest cells are migrating from the vagus nerve into the lung as early as E11, the earliest stage examined in their study (Tollet et al., 2001).

In 2010, Freem and collaborators also showed neural crest cell development within the lungs of mouse embryos. Through generation of a double transgenic mouse line constitutively expressing yellow fluorescent protein (YFP) in all neural crest cells and their progeny, Freem and colleagues confirmed neural crest cell development in the embryonic lung. These data show that primitive lung buds have begun to form at E10.5, and immunostaining using anti-green fluorescence protein showed that within the foregut, the majority of neural crest cells were linked with the esophagus, although YFP cells were also present within the trachea and primitive lung buds. At E11.5, a large part of the YFP-positive cells was within the lung buds, and afterward, at E12.5 and E13.5, the colonization of the lung buds by neural crest cells continues, where they noticed that YFP-positive cells frequently occur in close association with primitive branching bronchi and epithelial tubules. Association between neural crest cell-derived tissue and the airways continues as the lungs develop and mature up to birth, when YFP-positive neural crest cells combine to form ganglia that lie underneath the airway epithelium (Freem et al., 2010). Experiments performed in chick embryo showed neural crest cells in the lung buds earlier during the emergence of branches of the vagus nerve within the lungs (Burns & Delalande, 2005). The authors suggested that neural crest cells may subsequently enter from the vagus nerve into the lung and contribute to the intrinsic pulmonary neuronal population.

The airway smooth muscles (ASM) also seem to be involved with the vagus nerve branches extending to the developing trachea and lung buds as part from the onset of lung development (Sparrow et al., 1999; Tollet et al., 2002). Double staining immunohistochemistry to PGP 9.5 and α‎-actin performed by Tollet et al. (2001) revealed the distribution of nerves and smooth muscle at E11, showing prominent vagal nerves and ASM covering the trachea and epithelial tubules to the base of the growing lung buds. The same experiments demonstrated that the nerve fibers extend from the vagus to the trachea, pulmonary arteries, and lobar bronchi. Examining the mouse embryo at E12, they noticed two nerve trunks emerged from the ganglia, sending out fine fibers. At E13 on the trachea, they showed a network of nerve trunks and ganglia connected to the vagus and also the vagus nerve contributing to many neural processes to the hilum of the lung (Tollet et al., 2001).

In order to better describe vagal neural crest cell migratory behavior, in 2011, Kuo and Erickson used the chick in ovo model to delineate pathways taken by the neural crest cells from each somite level from the time they emigrate from the neural tube until they arrive at their final destination. They determined the subpopulations of vagal neural crest cells along the anterior and posterior axes that populate the heart and enteric nervous system of the gut by focally electroporating plasmids expressing Green Fluorescent Protein into the neural tube at individual somite levels, and the migration of the labeled neural crest cells was analyzed in older embryos. They observed aggregation of cells next to the neural tube, which have been referred to as the ganglionic crest of the cranial nerve IX and X (Kuo & Erickson, 2011). They also showed that neural crest cells from somite-level 1-2 are the ones that reach the anterior foregut (esophagus and lung bud region), extending posteriorly to the site where the laryngotracheal groove has split into the esophagus and the lung buds (Kuo & Erickson, 2011).

More specific considerations were made by Espinosa-Medina et al. (2017) in a recent study focused on murine vagal neural crest. They showed that neural crest-derived cells adjacent to somites 1-2 give rise to Schwann cells that migrate along the descending vagus nerve and form autonomic ganglia at their final destination. These data were focused on examining enteric ganglia in the walls of esophagus, but based on other experimental models, one cannot exclude the possibility that this also occurs for the respiratory tract (Espinosa-Medina et al., 2017).

Trunk Neural Crest Cells and the Development of Sympathetic and Sensory Innervations to the Lungs

Sympathetic and sensory neurons send projections to the airways and trunk neural crest cells give rise to these structures: the sympathetic ganglia and the dorsal root ganglia (DRG) (Yntema & Hammond, 1947). Pathways taken by neural crest cells to generate the sympathetic neural derivates are influenced by the surrounding structures, particularly somites (Lehmann, 1927). The somites are metameric mesodermal structures located lateral to the neural tube (Bronner-Fraser, 1986). Neural crest cells begin to migrate within and enter the anterior half of the sclerotome (Bronner-Fraser, 1986; Rickmann et al., 1985). Experiments in chick embryos using HNK-1 antibody to label neural crest cells during early development showed that crest cells migrate ventrally around the somites and then go either side of the neural tube (Loring & Erickson, 1987; Rickmann et al., 1985). After the ventral migration, neural crest cells follow a dorsolateral pathway, migrating between the ectoderm and dermomyotome (Erickson et al., 1992).

Cell fates depend on the migratory pathway. Cells that populate the sclerotome form the sensory neurons and glia of the DRG, whereas those that reach and pass through the sclerotome contribute to sympathetic ganglia (Teillet et al., 1987). Trunk neural crest cells that move ventrally between the neural tube accumulate dorsolateral to the dorsal aorta and give rise to the sympathetic ganglia (Kirby & Gilmore, 1976). Those neural crest cells that follow a dorsolateral pathway give rise to melanocytes (Erickson & Goins, 1995).

Kasemeier-Kulesa and collaborators (2005) investigated the cellular dynamics that mediate DRG and sympathetic ganglia formation by confocal microscopy in chicken embryos. They showed that during the migratory route, trunk neural crest cells are highly motile and dynamically interact with other cells. Furthermore, the cells migrate collectively, forming chain-like arrangements that extend from the DRG to the sympathetic ganglia. Interestingly, they also showed that at early stages, neural crest cells can reverse their direction of migration and switch locations between developing the DRG and sympathetic ganglia. However, later migrating cells are incapable of rerouting their trajectory once they reach their fate to be the DRG or sympathetic ganglia (Kasemeier-Kulesa et al., 2005). Seminal studies have demonstrated that neural crest cell-derived sensory neurons become fate specified soon after delamination from the neuroepithelium through a mechanism mediated by Wnt/β‎-catenin signaling (Hari et al., 2002; Lee et al., 2004). The β‎-catenin is important to migratory neural crest cells aggregate to form DRG and produce a wave of sensory neurogenesis (Hari et al., 2002).

Extrinsic and Intrinsic Signals that Generate Peripheral Autonomic Neurons

Sympathetic Neurons

Support for an extrinsic growth factor and morphogen role in neural crest development was first generated by Cohen (1972). In these studies, the authors prevented migrating crest cells from reaching the future site of the sympathetic ganglia. Tissue ventral to the neural tube was excluded from a graft of embryonic trunk and placed on the chorioallantoic membrane. They noticed that the sympathetic elements had the ability to differentiate in whole embryonic trunks, showing that the differentiation of sympathetic neuroblasts does not require migration to their final site (Cohen, 1972).

Experiments in which the notochord and neural tube were microsurgically excised showed that these structures are required for migrating neural crest cells to differentiate into sympathetic neurons. After the removal of notochord, in the presence of neural tube the peripheral nervous system develops close to normal the dorsal root ganglia (DRG), and sympathetic ganglia were developed and were segmentally distributed. However, the neural tube removal affected the pattern of crest cell development deeply (Teillet & Le Douarin, 1983).

Neural tube-derived factors are members of the transforming growth factor β‎ (TGF-β‎) family, whereas the dorsal aorta supplies crucial extrinsic instruction for the generation of sympathetic neurons of the sympathetic trunk through the secretion of bone morphogenetic proteins (BMPs) (Howard & Gershon, 1993; Shah et al., 1996). BMP signaling can be antagonized by treatment with Noggin, an extracellular protein with high regulated expression during development that binds to BMPs, thereby preventing BMP–BMP–receptor interactions (Ali & Brazil, 2014). Implantation of Noggin-releasing beads at the dorsal aorta resulted in the loss of tyrosine hydroxylase and dopamine beta hydroxylase, two key enzymes required in the biosynthesis of the sympathetic transcription factor noradrenaline (Schneider et al., 1999). Neural crest cell cultures exposed to BMPs in vitro also induce neuradrenergic fates (Varley & Maxwell, 1996; Wu & Howard, 2001). Aggregation of the neural crest cells around the dorsal aorta is correlated with expression of the basic helix–loop–helix DNA binding protein achaete-scute homologue (CASH in avians, MASH in mice, and HASH in humans [Guillemot & Joyner, 1993; Johnson et al., 1990]).

Generation of the sympathetic chain requires the function of the MASH and the homeodomain proteins Phox2b and Phox2a (Lo et al., 1997, 1998; Morin et al., 1997; Pattyn et al., 1999; Stanke et al., 2004). Of note, interactions between bHLH and homeodomain-containing proteins are thought to regulate neurotransmitter phenotype, as evidenced by interactions between Phox2a and Mash1 in generation of the noradrenergic phenotype in sympathetic cells (Xu et al., 2003).

Parasympathetic Neurons

The dynamic expression of transcriptional networks and neurogenic genes involved in parasympathetic differentiation during early development was evaluated by Burns and Delaland in 2005. Using orthotopic quail-chick interspecies grafting to label specific populations of neural crest cells, they showed that glia and neurons from ganglia are derived from vagal neural crest, express Sox10, a member of the SRY-like HMG box family of transcription factors, and receptor components of the Ret and endothelin receptor-B (EDNRB) signaling pathways (Burns & Delalande, 2005). Furthermore, they showed that those neural cells have the same developmental origin as the neurons and glial cells that compose the enteric nervous system. In situ hybridization experiments performed on chick embryo sections at E9 revealed Sox10 expressing cells distributed throughout the lung buds and at the periphery of ganglia limited to glial cells, both in the foregut and lungs. At the same period, the expression of the EDNRB receptor was similar to Sox10 and was present in the vagus nerve and its branches and at the periphery of lung ganglia. Ret expression at E9 was observed within lung bud ganglia, presumptively present at neurons (Burns & Delalande, 2005).

Glial cell-derived neurotrophic factor (GDNF) plays an important role during development and maintenance of different populations of central and peripheral neurons. This growth factor belongs to a family that includes neurturin, artemin, and persephin (Airaksinen & Saarma, 2002; Baloh et al., 2000). The receptor complex for GDNF family ligands requires the receptor subunit glycosyl-phosphatidyl inositol (GPI)-anchored coreceptor (GFRα‎) specialized in ligand binding and the receptor tyrosine kinase Ret involved in transmembrane signaling forming a heterocomplex necessary for GDNF signaling (Durbec et al., 1996; Jing et al., 1996).

Through in situ hybridization and immunohistochemistry were performed in mice embryos, Langsdorf and collaborators showed that neurturin is expressed in the trachea and bronchi and that the respiratory neural crest cells express both GFRα‎1 and GFRα‎2 (high-affinity receptors for GDNF and neurturin). Taking advantage of a neurturin knockout embryo, they evaluated intrinsic neurogenesis and observed that although neurturin is expressed in the trachea and bronchi, the absence of it is not sufficient to disrupt airway intrinsic innervation. In addition, Ret knockout embryos, with signaling activity compromised for all GDNF family members, reduces the number of intrinsic neurons in both the trachea and primary bronchi. The data showed that GDNF family signaling, through the Ret receptor, likely plays a role in the formation of intrinsic neurons in the respiratory tract (Langsdorf et al., 2011).

Other evidence shows that the ligand for the Ret receptor, GDNF, also contributes as an important chemoattractant for neural crest cells within the lung. Experiments demonstrating that GDNF-impregnated agarose beads, placed near the proximal part of the mouse lobe grown in in vitro organ culture, attracted neuronal precursors and influenced the direction of neurite extension (Tollet et al., 2002).

Disruption of Autonomic Function in Pulmonary Disease

Preterm infants with gestational ages as low as 23 weeks are prone to bronchopulmonary dysplasia (BPD). In the United States, BPD, a severe inflammatory lung disease, affects over 14,000 preterm infants (Stoll et al., 2010; Van Marter, 2009;). Among these patients, babies weighing 0.5–1 kg suffer the highest BPD incidence, estimated at 35%–68% (Fanaroff et al., 1998; Schmidt et al., 2012). Diminished lung function results in intervention with mechanical ventilation and oxygen supplementation, both of which induce BPD (Speer, 2006) as well as other morbidities that present to adulthood (Aukland et al., 2009; Halvorsen et al., 2004). Pulmonary hypertension (PH) secondary to BPD (BPD-PH) eventually burdens the right ventricle, resulting in limited cardiac output. Some 15%–30% of BPD patients suffer PH (Bhat et al., 2012), and BPD with PH (BPD-PH) patients show only 50% survival after 2 years (Khemani et al., 2007).

PH risk factors overlap with BPD and systemic inflammation risk factors, which include low gestational age, prolonged mechanical ventilation, and oxygen dependency, among other effects (Khemani et al., 2007; Taglauer et al., 2018). Furthermore, up to 46% of infants born before 26 weeks gestational age will develop sepsis (Stoll et al., 2002). A total of 16% of preterm babies suffer septicemia, of which 55%–65% experience apnea as a presenting symptom (Fanaroff et al., 1998; Lim et al., 2012). These apneic episodes induce hypoxia, further exacerbating hypoxic ischemic encephalopathy and periventricular leukomalacia injuries.

Significant strides in the understanding of BPD have been made from deriving basic fundamental knowledge of lung alveolar development with lung vascular development. For instance, alveolarization and vascularization represent synchronous developmental events (Alvira, 2016). By disrupting lung development, BPD reduces alveoli and thus overall lung volume. This reduces the cross-sectional area of the pulmonary vascular bed, resulting in increased pulmonary arterial blood pressure (Farquhar & Fitzgerald, 2010; Khemani et al., 2007; Stenmark & Abman, 2005). Right ventricular hypertrophy and remodeling initiates to compensate for the increased afterload of the pulmonary vasculature (Berkelhamer et al., 2013; Kim, 2010; Sehgal et al., 2016).

It is noted that lack of effective therapies for BPD and its associated disorders represents a major institutional focus for lung disease (McEvoy et al., 2014). Many studies have demonstrated links between BPD models and brain neurodevelopmental pathology (Poon et al., 2016). However, developmental neuropathology independent of BPD-related failure to thrive has been difficult to incisively test. Nevertheless, defective interoception as associated with BDP induces brain damage.

In addition to the well-documented vascular-based disease mechanisms for PH, a growing body of evidence suggests that marked autonomic dysfunction occurs in PH sympathetic, parasympathetic, and sensory nerve fibers innervating the pulmonary vessels. For instance, sympathetic α‎-adrenoreceptor agonism increases vascular resistance. Pulmonary baroreceptors activate noradrenergic fibers in the pulmonary artery and proximal airway segments (Barthelemy et al., 1996). Peripheral chemoreceptors respond to decreased arterial PO2 levels by increasing sympathetic nerve stimulation. Cholinergic-mediated relaxation of pulmonary arteries is mediated by parasympathetic activation via vagal stimulation, resulting in increased sympathetic nerve activity in PAH patients. Multiple pieces of evidence exist in the literature that raise autonomic dysfunction as a comorbidity of BPD-PH. For instance, Nootens et al. (1995) showed that in 32 patients with PH, plasma norepinephrine concentration was associated with poor estimated 5-year survival (Nootens et al., 1995). Furthermore, Chen et al. (2013) found reversal of pulmonary arterial changes in animals that had undergone pulmonary artery denervation (PADN).


  • Adriaensen, D., Timmermans, J. P., Brouns, I., Berthoud, H. R., Neuhuber, W. L., & Scheuermann, D. W. (1998). Pulmonary intraepithelial vagal nodose afferent nerve terminals are confined to neuroepithelial bodies: An anterograde tracing and confocal microscopy study in adult rats. Cell and Tissue Research, 293, 395–405.
  • Airaksinen, M. S., & Saarma, M. (2002). The GDNF family: Signalling, biological functions and therapeutic value. Nature, 3, 383–394.
  • Ali, I. H., & Brazil, D. P. (2014). Bone morphogenetic proteins and their antagonists: Current and emerging clinical uses. British Journal of Pharmacology, 171, 3620–3632.
  • Alvira, C. M. (2016). Aberrant pulmonary vascular growth and remodeling in bronchopulmonary dysplasia. Frontiers in Medicine (Lausanne), 3, 21.
  • Alzate-Correa, D., Liu, J., Jones, M., Silva, T. M., Alves, M. J., Burke, E., Zuñiga, J., Kaya, B., Zaza, G., Aslan, M. T., Blackburn, J., Shimada, M. Y., Fernandes-Junior, S. A., Baer, L. A., Stanford, K. I., Kempton, A., Smith, S., Szujewski, C. C., Silbaugh, A., . . . Otero, J. J. (2021). Neonatal apneic phenotype in a murine congenital central hypoventilation syndrome model is induced through non-cell autonomous developmental mechanisms. Brain Pathology, 31(1), 84–102.
  • Astick, M., Tubby, K., Mubarak, W. M., Guthrie, S., & Price, S. R. (2014). Central topography of cranial motor nuclei controlled by differential cadherin expression. Current Biology, 24, 2541–2547.
  • Aukland, S. M., Rosendahl, K., Owens, C. M., Fosse, K. R., Eide, G. E., & Halvorsen, T. (2009). Neonatal bronchopulmonary dysplasia predicts abnormal pulmonary HRCT scans in long-term survivors of extreme preterm birth. Thorax, 64, 405–410.
  • Baker, C. V., & Bronner-Fraser, M. (1997). The origins of the neural crest. Part I: Embryonic induction. Mechanisms of Development, 69, 3–11.
  • Baloh, R. H., Enomoto, H., Johnson, E. M. Jr., & Milbrandt, J. (2000). The GDNF family ligands and receptors - implications for neural development. Current Opinion in Neurobiology, 10, 103–110.
  • Barnes, P. J. (1986). Non-adrenergic non-cholinergic neural control of human airways. Archives Internationales de Pharmacodynamie et de Thérapie, 280, 208–228.
  • Barthelemy, P., Sabeur, G., & Jammes, Y. (1996). Assessment of an airway-to-pulmonary circulation reflex in cats. Neuroscience Letters, 211, 89–92.
  • Bass, A. H., & Baker, R. (1997). Phenotypic specification of hindbrain rhombomeres and the origins of rhythmic circuits in vertebrates. Brain, Behavior and Evolution, 50(Suppl. 1), 3–16.
  • Berkelhamer, S. K., Mestan, K. K., & Steinhorn, R. H. (2013). Pulmonary hypertension in bronchopulmonary dysplasia. Seminars in Perinatology, 37, 124–131.
  • Bhat, R., Salas, A. A., Foster, C., Carlo, W. A., & Ambalavanan, N. (2012). Prospective analysis of pulmonary hypertension in extremely low birth weight infants. Pediatrics, 129, e682–e689.
  • Boni, P., Ballati, L., & Evangelista, S. (1995). Tachykinin NK1 and NK2 receptors mediate the non-cholinergic bronchospastic response to capsaicin and vagal stimulation in guinea-pigs. Journal of Autonomic Pharmacology, 15, 49–54.
  • Bronner-Fraser, M. (1986). Analysis of the early stages of trunk neural crest migration in avian embryos using monoclonal antibody HNK-1. Developmental Biology, 115, 44–55.
  • Burns, A. J., & Delalande, J. M. (2005). Neural crest cell origin for intrinsic ganglia of the developing chicken lung. Developmental Biology, 277, 63–79.
  • Burns, A. J., & Le Douarin, N. M. (2001). Enteric nervous system development: Analysis of the selective developmental potentialities of vagal and sacral neural crest cells using quail-chick chimeras. The Anatomical Record, 262, 16–28.
  • Burns, A. J., Thapar, N., & Barlow, A. J. (2008). Development of the neural crest-derived intrinsic innervation of the human lung. American Journal of Respiratory Cell and Molecular Biology, 38, 269–275.
  • Carstairs, J. R., & Barnes, P. J. (1986). Visualization of vasoactive intestinal peptide receptors in human and guinea pig lung. Journal of Pharmacology and Experimental Therapeutics, 239, 249–255.
  • Chang, R. B., Strochlic, D. E., Williams, E. K., Umans, B. D., & Liberles, S. D. (2015). Vagal sensory neuron subtypes that differentially control breathing. Cell, 161, 622–633.
  • Chen, S. L., Zhang, Y. J., Zhou, L., Xie, D. J., Zhang, F. F., Jia, H. B., Wong, S. S., & Kwan, T. W. (2013). Percutaneous pulmonary artery denervation completely abolishes experimental pulmonary arterial hypertension in vivo. EuroIntervention, 9, 269–276.
  • Cochard, P., Goldstein, M., & Black, I. B. (1979). Initial development of the noradrenergic phenotype in autonomic neuroblasts of the rat embryo in vivo. Developmental Biology, 71, 100–114.
  • Cohen, A. M. (1972). Factors directing the expression of sympathetic nerve traits in cells of neural crest origin. Journal of Experimental Zoology, 179, 167–182.
  • Coleridge, H. M. (1986). Reflexes evoked from tracheobronchial tree and lungs. In N. S. Cherniack & J. G. Widdicombe (Eds.), Handbook of physiology: A critical, comprehensive presentation of physiologic knowledge and concepts. The respiratory system (Vol. 2, pp. 395–430). American Physiological Society.
  • Coleridge, J. C., & Coleridge, H. M. (1984). Afferent vagal C fibre innervation of the lungs and airways and its functional significance. Reviews of Physiology, Biochemistry and Pharmacology, 99, 1–110.
  • D’Amico-Martel, A., & Noden, D. M. (1983). Contributions of placodal and neural crest cells to avian cranial peripheral ganglia. The American Journal of Anatomy, 166, 445–468.
  • Diamond, L., & O‘Donnell, M. (1980). A nonadrenergic vagal inhibitory pathway to feline airways. Science, 208, 185–188.
  • Doidge, J. M., & Satchell, D. G. (1982). Adrenergic and non-adrenergic inhibitory nerves in mammalian airways. Journal of the Autonomic Nervous System, 5, 83–99.
  • Durbec, P., Marcos-Gutierrez, C. V., Kilkenny, C., Grigoriou, M., Wartiowaara, K., Suvanto, P., Smith, D., Ponder, B., Costantini, F., Saarma, M., Sariola, H., & Pachnis, V. (1996). GDNF signalling through the Ret receptor tyrosine kinase. Nature, 381, 789–793.
  • El-Bermani, A. W. (1978). Pulmonary noradrenergic innervation of rat and monkey: A comparative study. Thorax, 33, 167–174.
  • Erickson, C. A., Duong, T. D., & Tosney, K. W. (1992). Descriptive and experimental analysis of the dispersion of neural crest cells along the dorsolateral path and their entry into ectoderm in the chick embryo. Developmental Biology, 151, 251–272.
  • Erickson, C. A., & Goins, T. L. (1995). Avian neural crest cells can migrate in the dorsolateral path only if they are specified as melanocytes. Development, 121, 915–924.
  • Espinosa-Medina, I., Jevans, B., Boismoreau, F., Chettouh, Z., Enomoto, H., Müller, T., Birchmeier, C., Burns, A. J., & Brunet, J. F. (2017). Dual origin of enteric neurons in vagal Schwann cell precursors and the sympathetic neural crest. Proceedings of the National Academy of Sciences of the United States of America, 114, 11980–11985.
  • Fanaroff, A. A., Korones, S. B., Wright, L. L., Verter, J., Poland, R. L., Bauer, C. R., Tyson, J. E., Philips, J. B., III, Edwards, W., Lucey, J. F., Catz, C. S., Shankaran, S., & Oh, W. (1998). Incidence, presenting features, risk factors and significance of late onset septicemia in very low birth weight infants. The Pediatric Infectious Disease Journal, 17, 593–598.
  • Farquhar, M., & Fitzgerald, D. A. (2010). Pulmonary hypertension in chronic neonatal lung disease. Paediatric Respiratory Reviews, 11, 149–153.
  • Freem, L. J., Escot, S., Tannahill, D., Druckenbrod, N. R., Thapar, N., & Burns, A. J. (2010). The intrinsic innervation of the lung is derived from neural crest cells as shown by optical projection tomography in Wnt1-Cre YFP reporter mice. Journal of Anatomy, 217, 651–664.
  • Guillemot, F., & Joyner, A. L. (1993). Dynamic expression of the murine Achaete-Scute homologue Mash-1 in the developing nervous system. Mechanisms of Development, 42, 171–185.
  • Guthrie, S. (2007). Patterning and axon guidance of cranial motor neurons. Nature Reviews Neuroscience, 8, 859–871.
  • Haberberger, R. V., Barry, C., Dominguez, N., & Matusica, D. (2019). Human dorsal root ganglia. Frontiers in Cellular Neuroscience, 13, 271.
  • Halvorsen, T., Skadberg, B. T., Eide, G. E., Roksund, O. D., Carlsen, K. H., & Bakke, P. (2004). Pulmonary outcome in adolescents of extreme preterm birth: A regional cohort study. Acta Paediatrica, 93, 1294–1300.
  • Hari, L., Brault, V., Kleber, M., Lee, H. Y., Ille, F., Leimeroth, R., Paratore, C., Suter, U., Kemler, R., & Sommer, L. (2002). Lineage-specific requirements of beta-catenin in neural crest development. Journal of Cell Biology, 159, 867–880.
  • Howard, M. J., & Gershon, M. D. (1993). Role of growth factors in catecholaminergic expression by neural crest cells: In vitro effects of transforming growth factor beta 1. Developmental Dynamics, 196, 1–10.
  • Jarrar, W., Dias, J. M., Ericson, J., Arnold, H. H., & Holz, A. (2015). Nkx2.2 and Nkx2.9 are the key regulators to determine cell fate of branchial and visceral motor neurons in caudal hindbrain. PLOS One, 10, e0124408.
  • Jing, S., Wen, D., Yu, Y., Holst, P. L., Luo, Y., Fang, M., Tamir, R., Antonio, L., Hu, Z., Cupples, R., Louis, J. C., Hu, S., Altrock, B. W., & Fox, G. M. (1996). GDNF–induced activation of the Ret protein tyrosine kinase is mediated by GDNFR-a, a novel receptor for GDNF. Cell, 85, 1113–1124.
  • Johnson, J. E., Birren, S. J., & Anderson, D. J. (1990). Two rat homologues of Drosophila achaete-scute specifically expressed in neuronal precursors. Nature, 346, 858–861.
  • Kasemeier-Kulesa, J. C., Kulesa, P. M., & Lefcort, F. (2005). Imaging neural crest cell dynamics during formation of dorsal root ganglia and sympathetic ganglia. Development, 132, 235–245.
  • Khemani, E., McElhinney, D. B., Rhein, L., Andrade, O., Lacro, R. V., Thomas, K. C., & Mullen, M. P. (2007). Pulmonary artery hypertension in formerly premature infants with bronchopulmonary dysplasia: Clinical features and outcomes in the surfactant era. Pediatrics, 120, 1260–1269.
  • Kim, G. B. (2010). Pulmonary hypertension in infants with bronchopulmonary dysplasia. Korean Journal of Pediatrics, 53, 688–693.
  • Kirby, M. L., & Gilmore, S. A. (1976). A correlative histofluorescence and light microscopic study of the formation of the sympathetic trunks in chick embryos. The Anatomical Record, 186, 437–449.
  • Kuo, B. R., & Erickson, C. A. (2011). Vagal neural crest cell migratory behavior: A transition between the cranial and trunk crest. Developmental Dynamics, 240, 2084–2100.
  • Kupari, J., Häring, M., Agirre, E., Castelo-Branco, G., & Emfors, P. (2019). An atlas of vagal sensory neurons and their molecular specialization. Cell Reports, 27, 2508–2523.
  • Langsdorf, A., Radzikinas, K., Kroten, A., Jain, S., & Ai, X. (2011). Neural crest cell origin and signals for intrinsic neurogenesis in the mammalian respiratory tract. American Journal of Respiratory Cell and Molecular Biology, 44, 293–301.
  • Larsell, O. (1922). The ganglia, plexuses, and nerve-terminations of the mammalian lung and pleura pulmonalis. The Journal of Comparative Neurology, 35(1), 97–132.
  • Le Douarin, N. M., & Teillet, M. A. (1973). The migration of neural crest cells to the wall of the digestive tract in avian embryo. Journal of Embryology and Experimental Morphology, 30, 31–48.
  • Lee, H. Y., Kleber, M., Hari, L., Brault, V., Suter, U., Taketo, M. M., Kemler, R., & Sommer, L. (2004). Instructive role of Wnt/beta-catenin in sensory fate specification in neural crest stem cells. Science, 303, 1020–1023.
  • Lee, L. Y., & Pisarri, T. E. (2001). Afferent properties and reflex functions of bronchopulmonary C-fibers. Respiration Physiology, 125, 47–65.
  • Lee, L. Y., Shuei Lin, Y., Gu, Q., Chung, E., & Ho, C. Y. (2003). Functional morphology and physiological properties of bronchopulmonary C-fiber afferents. The Anatomical Record, Part A: Discoveries in Molecular Cellular and Evolutionary Biology, 270, 17–24.
  • Lee, L. Y., & Yu, J. (2014). Sensory nerves in lung and airways. Comprehensive Physiology, 4, 287–324.
  • Lehmann, F. E. (1927). Further studies on the morphogenetic role of the somites in the development of the nervous system of amphibians: The differentiation and arrangement of the spinal ganglia in Pleurodeles waltli. Journal of Experimental Zoology, 49, 93–131.
  • Lim, W. H., Lien, R., Huang, Y. C., Chiang, M. C., Fu, R. H., Chu, S. M., Hsu, J. F., & Yang, P. H. (2012). Prevalence and pathogen distribution of neonatal sepsis among very-low-birth-weight infants. Pediatrics and Neonatology, 53, 228–234.
  • Lo, L., Sommer, L., & Anderson, D. J. (1997). MASH1 maintains competence for BMP2-induced neuronal differentiation in post-migratory neural crest cells. Current Biology, 7, 440–450.
  • Lo, L., Tiveron, M. C., & Anderson, D. J. (1998). MASH1 activates expression of the paired homeodomain transcription factor Phox2a, and couples pan-neuronal and subtype-specific components of autonomic neuronal identity. Development, 125, 609–620.
  • Loring, J. F., & Erickson, C. A. (1987). Neural crest cell migratory pathways in the trunk of the chick embryo. Developmental Biology, 121, 220–236.
  • Lumsden, A., & Keynes, R. (1989). Segmental patterns of neuronal development in the chick hindbrain. Nature, 337, 424–428.
  • Lumsden, A., Sprawson, N., & Graham, A. (1991). Segmental origin and migration of neural crest cells in the hindbrain region of the chick embryo. Development, 113, 1281–1291.
  • Lundberg, J. M., & Saria, A. (1982). Bronchial smooth muscle contraction induced by stimulation of capsaicin-sensitive sensory neurons. Acta Physiologica Scandinavica, 116, 473–476.
  • Lundberg, J. M., Saria, A., Brodin, E., Rosell, S., & Folkers, K. (1983). A substance P antagonist inhibits vagally induced increase in vascular permeability and bronchial smooth muscle contraction in the guinea pig. Proceedings of the National Academy of Sciences of the United States of America, 80, 1120–1124.
  • Matran, R. (1991). Neural control of lower airway vasculature: Involvement of classical transmitters and neuropeptides. Acta Physiologica Scandinavica, 601(Suppl.), 1–54.
  • McEvoy, C. T., Jain, L., Schmidt, B., Abman, S., Bancalari, E., & Aschner, J. L. (2014). Bronchopulmonary dysplasia: NHLBI workshop on the primary prevention of chronic lung diseases. Annals of the American Thoracic Society, 11(Suppl. 3), S146–S153.
  • Morin, X., Cremer, H., Hirsch, M. R., Kapur, R. P., Goridis, C., & Brunet, J. F. (1997). Defects in sensory and autonomic ganglia and absence of locus coeruleus in mice deficient for the homeobox gene Phox2a. Neuron, 18, 411–423.
  • Nadel, J. A. (1974). Parasympathetic nervous control of airway smooth muscle. Annals of the New York Academy of Sciences, 221, 99–102.
  • Nassenstein, C., 1,2, Taylor-Clark, T. E., Myers, A. C., Ru, F., Nandigama, R., Bettner, W., Undem, B. J. (2010) Phenotypic distinctions between neural crest and placodal derived vagal C-fibres in mouse lungs, Journal of Physiology, 588.23, 4769–4783.
  • Nootens, M., Kaufmann, E., Rector, T., Toher, C., Judd, D., Francis, G. S., & Rich, S. (1995). Neurohormonal activation in patients with right ventricular failure from pulmonary hypertension: Relation to hemodynamic variables and endothelin levels. Journal of the American College of Cardiology, 26, 1581–1585.
  • Paintal, A. S. (1973). Vagal sensory receptors and their reflex effects. Physiological Reviews, 53, 159–227.
  • Palmer, J. B., Cuss, F. M., & Barnes, P. J. (1986). VIP and PHM and their role in nonadrenergic inhibitory responses in isolated human airways. Journal of Applied Physiology, 61, 1322–1328.
  • Pattyn, A., Morin, X., Cremer, H., Goridis, C., & Brunet, J. F. (1999). The homeobox gene Phox2b is essential for the development of autonomic neural crest derivatives. Nature, 399, 366–370.
  • Poon, A. W., Ma, E. X., Vadivel, A., Jung, S., Khoja, Z., Stephens, L., Thebaud, B., & Wintermark, P. (2016). Impact of bronchopulmonary dysplasia on brain and retina. Biology Open, 5, 475–483.
  • Rickmann, M., Fawcett, J. W., & Keynes, R. J. (1985). The migration of neural crest cells and the growth of motor axons through the rostral half of the chick somite. Journal of Embryology and Experimental Morphology, 90, 437–455.
  • Roddie, I. C. (1970). Modern views on physiology. XXIV. Autonomic nervous system. Practitioner, 205, 828–834.
  • Rubin, E. (1985a). Development of the rat superior cervical ganglion: Ganglion cell maturation. The Journal of Neuroscience, 5, 673–684.
  • Rubin, E. (1985b). Development of the rat superior cervical ganglion: Ingrowth of preganglionic axons. The Journal of Neuroscience, 5, 685–696.
  • Sadaghiani, B., & Thiebaud, C. H. (1987). Neural crest development in the Xenopus laevis embryo, studied by interspecific transplantation and scanning electron microscopy. Developmental Biology, 124, 91–110.
  • Schittny, J. C. (2017). Development of the lung. Cell and Tissue Research, 367, 427–444.
  • Schmidt, B., Anderson, P. J., Doyle, L. W., Dewey, D., Grunau, R. E., Asztalos, E. V., Davis, P. G., Tin, W., Moddemann, D., Solimano, A., Ohlsson, A., Barrington, K. J., Roberts, R. S., & Caffeine for Apnea of Prematurity Trial I. (2012). Survival without disability to age 5 years after neonatal caffeine therapy for apnea of prematurity. JAMA, 307, 275–282.
  • Schneider, C., Wicht, H., Enderich, J., Wegner, M., & Rohrer, H. (1999). Bone morphogenetic proteins are required in vivo for the generation of sympathetic neurons. Neuron, 24, 861–870.
  • Sehgal, A., Malikiwi, A., Paul, E., Tan, K., & Menahem, S. (2016). Systemic arterial stiffness in infants with bronchopulmonary dysplasia: Potential cause of systemic hypertension. Journal of Perinatology, 36, 564–569.
  • Shah, N. M., Groves, A. K., & Anderson, D. J. (1996). Alternative neural crest cell fates are instructively promoted by TGFbeta superfamily members. Cell, 85, 331–343.
  • Sparrow, M. P., & Lamb, J. P. (2003). Ontogeny of airway smooth muscle: structure, innervation, myogenesis and function in the fetal lung. Respiratory Physiology & Neurobiology, 137, 361–372.
  • Sparrow, M. P., Weichselbaum, M., & McCray, P. B. (1999). Development of the innervation and airway smooth muscle in human fetal lung. American Journal of Respiratory Cell and Molecular Biology, 20, 550–560.
  • Speer, C. P. (2006). Pulmonary inflammation and bronchopulmonary dysplasia. Journal of Perinatology, 26(Suppl. 1), S57–S64.
  • Spooner, B. S., & Wessells, N. K. (1970). Mammalian lung development: Interactions in primordium formation and bronchial morphogenesis. Journal of Experimental Zoology, 175, 445–454.
  • Springall, D. R., Cadieux, A., Oliveira, H., Su, H., Royston, D., & Polak, J. M. (1987). Retrograde tracing shows that CGRP-immunoreactive nerves of rat trachea and lung originate from vagal and dorsal root ganglia. Journal of the Autonomic Nervous System, 20, 155–166.
  • Stanke, M., Stubbusch, J., & Rohrer, H. (2004). Interaction of Mash1 and Phox2b in sympathetic neuron development. Molecular and Cellular Neuroscience, 25, 374–382.
  • Stenmark, K. R., & Abman, S. H. (2005). Lung vascular development: Implications for the pathogenesis of bronchopulmonary dysplasia. Annual Review of Physiology, 67, 623–661.
  • Stiles, J., & Jernigan, T. L. (2010). The basics of brain development. Neuropsychology Review, 20, 327–348.
  • Stoll, B. J., Hansen, N., Fanaroff, A. A., Wright, L. L., Carlo, W. A., Ehrenkranz, R. A., Lemons, J. A., Donovan, E. F., Stark, A. R., Tyson, J. E., Oh, W., Bauer, C. R., Korones, S. B., Shankaran, S., Laptook, A. R., Stevenson, D. K., Papile, L. A., & Poole, W. K. (2002). Late-onset sepsis in very low birth weight neonates: The experience of the NICHD Neonatal Research Network. Pediatrics, 110, 285–291.
  • Stoll. B. J., Hansen, N. I., Bell, E. F., Shankaran, S., Laptook, A. R., Walsh, M. C., Hale, E. C., Newman, N. S., Schibler, K., Carlo, W. A., Kennedy, K. A., Poindexter, B. B., Finer, N. N., Ehrenkranz, R. A., Duara, S., Sanchez, P. J., O‘Shea, T. M., Goldberg, R. N., Van Meurs, K. P. . . . Eunice Kennedy Shriver National Institute of Child Health & Human Development Neonatal Research Network. (2010). Neonatal outcomes of extremely preterm infants from the NICHD Neonatal Research Network. Pediatrics, 126, 443–456.
  • Swanson, L. W. (2000). What is the brain? Trends in Neuroscience, 23, 519–527.
  • Taglauer, E., Abman, S. H., & Keller, R. L. (2018). Recent advances in antenatal factors predisposing to bronchopulmonary dysplasia. Seminars in Perinatology, 42, 413–424.
  • Teillet, M. A., Kalcheim, C., & Le Douarin, N. M. (1987). Formation of the dorsal root ganglia in the avian embryo: Segmental origin and migratory behavior of neural crest progenitor cells. Developmental Biology, 120, 329–347.
  • Teillet, M. A., & Le Douarin, N. M. (1983). Consequences of neural tube and notochord excision on the development of the peripheral nervous system in the chick embryo. Developmental Biology, 98, 192–211.
  • Ten Have-Opbroek, A. A. (1981). The development of the lung in mammals: An analysis of concepts and findings. American Journal of Anatomy, 162, 201–219.
  • Tollet, J., Everett, A. W., & Sparrow, M. P. (2001). Spatial and temporal distribution of nerves, ganglia, and smooth muscle during the early pseudoglandular stage of fetal mouse lung development. Developmental Dynamics, 221, 48–60.
  • Tollet, J., Everett, A. W., & Sparrow, M. P. (2002). Development of neural tissue and airway smooth muscle in fetal mouse lung explants: A role for glial-derived neurotrophic factor in lung innervation. American Journal of Respiratory Cell and Molecular Biology, 26, 420–429.
  • Van Lommel, A., Lauweryns, J. M., & Berthoud, H. R. (1998). Pulmonary neuroepithelial bodies are innervated by vagal afferent nerves: an investigation with in vivo anterograde DiI tracing and confocal microscopy. Anatomy and Embryology (Berlin), 197, 325–330.
  • Van Marter, L. J. (2009). Epidemiology of bronchopulmonary dysplasia. Seminars in Fetal & Neonatal Medicine, 14, 358–366.
  • Varley, J. E., & Maxwell, G. D. (1996). BMP-2 and BMP-4, but not BMP-6, increase the number of adrenergic cells which develop in quail trunk neural crest cultures. Experimental Neurology, 140, 84–94.
  • Wasano, K. (1977). Neuro-epithelial bodies in the lung of the rat and the mouse. Archivum Histologicum Japonicum, 40(Suppl.), 207–219.
  • Weichselbaum, M., Everett, A. W., & Sparrow, M. P. (1996). Mapping the innervation of the bronchial tree in fetal and postnatal pig lung using antibodies to PGP 9.5 and SV2. American Journal of Respiratory Cell and Molecular Biology, 15, 703–710.
  • Weigand, L. A., & Myers, A. C. (2010). Synaptic and membrane properties of parasympathetic ganglionic neurons innervating mouse trachea and bronchi. American Journal of Physiology: Lung Cellular and Molecular Physiology, 298, L593–L599.
  • Widdicombe, J. (1981). Nervous receptors in the respiratory tree and lungs. In T. Hornbeing (Ed.), Lung biology in health and disease: Regulation of breathing (Chapter 6). CRC Press.
  • Wu, X., & Howard, M. J. (2001). Two signal transduction pathways involved in the catecholaminergic differentiation of avian neural crest-derived cells in vitro. Molecular and Cellular Neuroscience, 18, 394–406.
  • Xu, H., Firulli, A. B., Zhang, X., & Howard, M. J. (2003). HAND2 synergistically enhances transcription of dopamine-beta-hydroxylase in the presence of Phox2a. Developmental Biology, 262, 183–193.
  • Yntema, C. L., & Hammond, W. S. (1947). The development of the autonomic nervous system. Biological Reviews of the Cambridge Philosophical Society, 22, 344–359.
  • Yuan, H., & Silberstein, S. D. (2016). Vagus nerve and vagus nerve stimulation: A comprehensive review. Part I. Headache, 56, 71–78.
  • Zhuo, H., Ichikawa, H., & Helke, C. J. (1997). Neurochemistry of the nodose ganglion. Progress in Neurobiology, 52, 79–107.