The carotid bodies are small glands found in either side of our neck, near the carotid artery. When the level of oxygen in our blood drops, specialized cells in the carotid bodies signal to the brain to increase our heart rate and make us breathe more rapidly and deeply. As a result, more oxygen is delivered to our cells.

Fish have similar oxygen-sensitive cells in their gills, known as neuroepithelial cells, that detect changes in the oxygen levels in the surrounding water and their blood. It has been suggested that after our vertebrate (back-boned animal) ancestors moved onto land, the neuroepithelial cells in their gills eventually evolved to form the carotid bodies. Knowing whether this is true would allow researchers to better understand how our ancestors were able to adapt to an obligate air-breathing lifestyle on land.

If the carotid body did evolve from ancestral neuroepithelial cells, we would expect that they would both develop from the same kind of cells in the embryo. Carotid body cells develop from a group of cells called neural crest cells, which give rise to many tissues, including nerve cells. Hockman et al. have now investigated whether neuroepithelial cells also develop from neural crest cells.

Hockman et al. labelled the neural crest cells in the embryos of zebrafish, frogs and lampreys using techniques such as injecting the cells with fluorescent dye or genetically modifying the cells to make fluorescent proteins. Unexpectedly, the neuroepithelial cells that developed in the gills of these embryos did not contain these fluorescent labels, meaning that they did not develop from the neural crest cells. The patterns of gene activity found in the developing neuroepithelial cells were also different from those in the carotid body. Further investigation revealed that neuroepithelial cells develop from the lining of the mouth and gills and may be related to a similar population of oxygen-sensitive cells found in the lungs.

Overall, it appears that the carotid body did not evolve from ancestral neuroepithelial cells. However, Hockman et al. did find some cells near blood vessels in the gills of zebrafish that had developed from neural crest cells. Equivalent cells in our ancestors could therefore be the cells that evolved into carotid bodies. A first test of this theory will be to determine whether or not these cells are oxygen-sensitive.

During hypoxia in vertebrates, respiratory reflexes such as hyperventilation are triggered by neurotransmitter release from hypoxia-sensitive serotonergic cells associated with pharyngeal arch arteries, as well as in the lungs and/or gills (reviewed by López-Barneo et al., 2016; Cutz et al., 2013; Jonz et al., 2016). In amniotes, these are the ‘glomus cells’ of the carotid body, located at the bifurcation of the common carotid artery (reviewed by Nurse, 2014; López-Barneo et al., 2016), and the ‘pulmonary neuroendocrine cells’ (PNECs) of lung airway epithelia, found either as solitary, flask-shaped cells, or collected into ‘neuroepithelial bodies’ (NEBs), preferentially located at airway branch points (reviewed by Cutz et al., 2013). Glomus cells respond to hypoxia in arterial blood by releasing stored neurotransmitters including acetylcholine, ATP, the catecholamine dopamine, and serotonin (the latter two most likely acting as autocrine/paracrine neuromodulators) (reviewed by Nurse and Piskuric, 2013; Nurse, 2014). These excite afferent terminals of the carotid sinus nerve (a branch of the glossopharyngeal nerve, arising from neurons in the petrosal ganglion) in mammals (reviewed by Nurse and Piskuric, 2013; Nurse, 2014), and of the vagal nerve (arising from neurons in the nodose ganglion) in birds (Kameda, 2002). The afferent nerves relay signals to the nucleus of the solitary tract within the hindbrain, to elicit respiratory reflex responses such as hyperventilation (reviewed by Teppema and Dahan, 2010). PNECs respond to hypoxia by releasing stored serotonin and various neuropeptides onto vagal afferents, and are thought to act as hypoxia-sensitive airway sensors (reviewed by Cutz et al., 2013; also see Branchfield et al., 2016). PNECs also provide an important stem-cell niche for regenerating the airway epithelium after injury (Reynolds et al., 2000; Guha et al., 2012; Song et al., 2012) and were recently shown to be the predominant cells of origin for small cell lung cancer (Park et al., 2011; Sutherland et al., 2011; Song et al., 2012).

The evolution of glomus cells was critical for the transition from aquatic life, where externally facing hypoxia-sensors are essential for monitoring the variable oxygen levels in water, to fully terrestrial life, where reflex responses to variations in internal oxygen levels are more important, given the stability of oxygen levels in air (Burleson and Milsom, 2003; Milsom and Burleson, 2007). However, the evolutionary history of glomus cells - and, indeed, PNECs - is uncertain. One commonly suggested hypothesis (e.g., Milsom and Burleson, 2007; Hempleman and Warburton, 2013; Jonz et al., 2016) is that glomus cells are homologous to the chemosensory ‘neuroepithelial cells’ (NECs) of fish gills. These were originally identified within the primary epithelium of the gills in various teleosts and a shark, as innervated cells (isolated or clustered) containing dense-cored vesicles; formaldehyde-induced fluorescence revealed the presence of biogenic amines, identified as serotonin, while electron microscopy following exposure of trout to acute hypoxia revealed fewer vesicles, which appeared degranulated (Dunel-Erb et al., 1982). In vitro patch-clamp studies on NECs isolated from zebrafish and catfish gills (identified by neutral red, a vital dye that stains monoamine-containing cells including serotonergic PNECs; Youngson et al., 1993), confirmed that teleost gill NECs are hypoxia-sensitive (Jonz et al., 2004; Burleson et al., 2006). The conventional marker for teleost gill NECs is serotonin: although non-serotonergic gill NECs were identified in adult zebrafish by immunoreactivity for synaptic vesicle glycoprotein 2, these may represent immature NECs (Jonz and Nurse, 2003; Jonz et al., 2004). NECs are found near efferent gill arteries and on the basal lamina of the gill epithelia, facing the flow of water, hence can detect hypoxia and other stimuli in either blood or external water (reviewed by Jonz et al., 2016). Like glomus cells (reviewed by Nurse, 2014; López-Barneo et al., 2016), zebrafish gill NECs also respond to acid hypercapnia (increased CO₂/H⁺) (López-Barneo et al., 1988; Buckler, 1997; Jonz et al., 2004; Qin et al., 2010). The putative shared evolutionary ancestry of glomus cells and gill NECs (e.g. Milsom and Burleson, 2007; Hempleman and Warburton, 2013; Jonz et al., 2016) is supported by many similarities: both are associated with pharyngeal arch arteries (the carotid body develops in association with the third pharyngeal arch artery, which will form the carotid artery), provided with afferent innervation by glossopharyngeal and/or vagal nerves, and have background ('leak') K⁺ currents that are inhibited by hypoxia, resulting in membrane depolarization, activation of voltage-gated Ca²⁺ channels, and neurotransmitter release (López-Barneo et al., 1988; Buckler, 1997; Jonz et al., 2004; Qin et al., 2010).

If glomus cells and gill NECs evolved from the same ancestral cell population, they should share a common embryonic origin. Glomus cells are neural crest-derived, as demonstrated in birds by quail-chick neural fold grafts (Le Douarin et al., 1972; Pearse et al., 1973), and in mouse by Wnt1-Cre genetic lineage-tracing (Pardal et al., 2007), but the embryonic origin of NECs is unknown. Lack of immunoreactivity for the HNK1 antibody, which labels migrating neural crest cells in many but not all vertebrates, has been reported for NECs (Porteus et al., 2013, 2014). However, the carbohydrate epitope recognized by the HNK1 antibody (Voshol et al., 1996) is borne by multiple glycoproteins and glycolipids, and gene or antigen expression in itself cannot indicate lineage. An alternative to the hypothesis that gill NECs and glomus cells evolved from a common ancestral cell population is that NECs share ancestry with PNECs, to which they were originally likened (Dunel-Erb et al., 1982). Hypoxia-sensing by PNECs, as in NECs and glomus cells, involves inhibition of a K⁺ current by hypoxia (reviewed by Cutz et al., 2013; Nurse, 2014; López-Barneo et al., 2016; Jonz et al., 2016). In contrast to the neural crest origin of glomus cells (Le Douarin et al., 1972; Pearse et al., 1973; Pardal et al., 2007), PNECs have an intrinsic pulmonary epithelial origin: the first experimental support for this was provided by a tritiated thymidine labeling study of hamster lung development (Hoyt et al., 1990), later confirmed by mouse genetic lineage-tracing studies using Id2-CreER^T2, Shh-Cre, Nkx2.1-Cre, and Sox9-Cre driver lines that showed a common origin for all lung airway epithelial cell types, including PNECs (Rawlins et al., 2009; Song et al., 2012; Kuo and Krasnow, 2015).

Here, we demonstrate that neural crest cells do not contribute to gill NECs: instead, these are endoderm-derived. This refutes the hypothesis that glomus cells and gill NECs evolved from a common ancestral cell population, and instead supports an evolutionary relationship between NECs and PNECs, whose endodermal origin we confirm in mouse. We also show that the transcription factor Phox2b, which is required for glomus cell development (Dauger et al., 2003), is not expressed by gill NECs (or by PNECs), arguing against the possibility of cell-type homology between glomus cells and gill NECs via activation of the same genetic network. Finally, we report the discovery of neural crest-derived chromaffin (catecholaminergic) cells associated with blood vessels in the pharyngeal arches of juvenile zebrafish, which we speculate could share an evolutionary ancestry with glomus cells. Given these results, we propose a new model for the evolution of hypoxia-sensitive cells during the transition to terrestrial life.

Carotid body glomus cells develop from the neural crest (Le Douarin et al., 1972; Pearse et al., 1973; Pardal et al., 2007), while PNECs differentiate in situ within pulmonary airway epithelia (Hoyt et al., 1990; Rawlins et al., 2009; Song et al., 2012; Kuo and Krasnow, 2015). We aimed to shed light on the evolutionary origins of these amniote hypoxia-sensitive cell types by determining the embryonic origin of NECs, the hypoxia-sensitive cells of anamniote gills.

Zebrafish NECs are not neural crest-derived

In developing zebrafish, gill NECs were previously identified as serotonin (5-HT)-immunoreactive cells in gill filaments from 5-dpf, which are innervated by 7-dpf (Jonz and Nurse, 2005). We investigated any neural crest contribution to gill NECs via genetic lineage-tracing, using a collection of transgenic zebrafish lines with different cis-regulatory sequences driving Cre and subsequent lineage reporter expression in neural crest-derived cells, as well as via neural crest-deficient zebrafish embryos. Lineage-tracing using different Cre driver and reporter lines enabled us to control for false negatives potentially arising from variable promoter activity in, or incomplete labeling of, some neural crest cells in either neural crest driver or reporter lines.

In larval and juvenile zebrafish, we identified NECs in the gill filaments by serotonin immunoreactivity, as previously reported (Jonz and Nurse, 2005); we also found similar innervated serotonergic cells scattered in the orobranchial epithelium (putative NECs). NECs in the gill filaments, and putative NECs in the orobranchial epithelium, were unlabeled in larvae/metamorphic juveniles with sox10 cis-regulatory sequences driving Cre and resulting lineage reporter expression [Tg(-28.5sox10:cre);Tg(ef1a:loxP-DsRed-loxP-EGFP) (Kague et al., 2012); Tg(-4.9sox10:creER^T2);Tg(βactin:loxP-SuperStop-loxP-DsRed) (Mongera et al., 2013)], even when nearby neural crest derivatives such as branchial arch cartilages/mesenchyme and/or gill pillar cells (Mongera et al., 2013) were reporter-positive (Figure 1a–f'; 183 serotonergic cells located near such reporter-positive cells [≥6 per fish] were counted across 11 larvae/metamorphic juveniles: n = 8 for −28.5sox10; n = 3 for −4.9sox10). Gill NECs, and putative NECs in the orobranchial epithelium, were also unlabeled in Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) (Mosimann et al., 2011; Kaufman et al., 2016) larvae/metamorphic juveniles, even when nearby neural crest-derived cells in branchial arch cartilages and/or gill filament mesenchyme were mCherry-positive (Figure 1g-h'; 166 serotonergic cells located near such mCherry-positive cells [≥6 per fish] were counted across 10 larvae/metamorphic juveniles). We ruled out the possibility that lack of lineage reporter expression in NECs was a false-negative result arising from inactivity in NECs of the promoters driving the Cre-switchable reporter cassettes, by confirming that NECs expressed the native unrecombined reporter in both transgenic lines [Tg(-28.5sox10:cre);Tg(ef1a:loxP-DsRed-loxP-EGFP) and Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry)] (Figure 1—figure supplement 1).

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Finally, we analyzed tfap2a^mob;foxd3^mos zebrafish, which lack neural crest derivatives (Barrallo-Gimeno et al., 2004; Montero-Balaguer et al., 2006; Wang et al., 2011). At 7-dpf, tfap2a^mob;foxd3^mos mutants lacked pigment cells and lower jaw structures (n = 8; Figure 2a). In the absence of the neural crest-derived pharyngeal endoskeleton, pharyngeal arches and gills are hard to recognize, but putative NECs, visualized here as serotonergic cells associated with HNK1 epitope-immunoreactive neurites (Metcalfe et al., 1990), were still present in the orobranchial epithelium (Figure 2b,c) (and occasionally could also be identified ventral to the orobranchial cavity, where the pharyngeal arches would be located; Figure 2d,e). We counted all putative NECs in the orobranchial region of three tfap2a^mob;foxd3^mos larvae (294 putative NECs counted in total) and three wild-type siblings (244 putative NECs counted in total): there was no change in mean number (mean/embryo ± s.d.: 81.3 ± 24.0 for wild-type larvae, n = 3; 98.0 ± 49.7 for tfap2a^mob;foxd3^mos larvae, n = 3; p=0.63, unpaired two-tailed Student’s t-test) (Figure 2f–i).

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Putative NECs have also been identified as serotonergic cells in the skin of embryonic zebrafish (Jonz and Nurse, 2006; Coccimiglio and Jonz, 2012) and of adult mangrove killifish, which respire through the skin as well as the gills (Regan et al., 2011). Although they have not been shown directly (e.g., by patch-clamp experiments) to be hypoxia-sensitive, the putative NECs in killifish increase in area in response to hypoxia (Regan et al., 2011), while in zebrafish, hypoxia decreased or delayed, and hyperoxia accelerated, the normal decline in number of these cells seen with increasing age (Coccimiglio and Jonz, 2012). In zebrafish, these serotonergic cells are most abundant at 3-dpf, and most evident scattered in the skin over the eyes, yolk-sac and tail (Coccimiglio and Jonz, 2012). Whole-mount immunostaining of Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) embryos at 3-dpf for serotonin and mCherry revealed the expected pattern of scattered serotonergic cells in the epidermis, but none was mCherry-positive (i.e., neural crest-derived) (Figure 2—figure supplement 1 shows a sample three-dimensional rendering for the eye; for the associated z-stack movies, see Videos 1 and 2). We quantified this in the eye, where the corneal endothelium is neural crest-derived, as previously reported for chicken (Noden, 1978; Johnston et al., 1979), mouse (with a minor mesodermal contribution also noted; Gage et al., 2005) and Xenopus (Hu et al., 2013). At 3-dpf, the zebrafish cornea comprises an outer corneal epithelium, a thin acellular collagenous stroma, and a monolayer of flattened corneal endothelial cells (Soules and Link, 2005; Zhao et al., 2006; Akhtar et al., 2008). Across 13 embryos, we counted 328 serotonergic cells in the epidermis over the eye, all of which were mCherry-negative, and 219 nearby mCherry-positive (i.e., neural crest-derived) corneal endothelial cells. Thus, the putative NECs in the skin of embryonic zebrafish are not neural crest-derived.

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Taken together, these results show that in zebrafish (a ray-finned fish), the neural crest does not contribute to gill NECs (the previously proposed homologs of glomus cells), or to putative NECs in the orobranchial epithelium and embryonic epidermis.

Putative NECs in Xenopus and lamprey are not neural crest-derived

Innervated serotonergic cells in the internal gills of Xenopus tadpoles are the proposed homologs of teleost gill NECs (Saltys et al., 2006). We detected scattered serotonergic cells in gill filaments from stage 43 (Figure 3a,b), and in the orobranchial epithelium from stage 41 (Figure 3a,c). To determine if these putative NECs are neural crest-derived, we unilaterally grafted neural folds from GFP-labeled donors to unlabeled hosts (Figure 3d). GFP-labeled neural crest cells migrated away from the neural tube and contributed to branchial arch cartilages and mesenchyme, but putative NECs in the gill filaments and orobranchial epithelium were GFP-negative (n = 14; Figure 3e–m’).

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In the sea lamprey, the gills are arranged in pairs within the orobranchial cavity, supported by an interbranchial septum (Figure 4a–c). Serotonergic cells in lamprey gills are proposed to correspond to the gill NECs of jawed fishes (Barreiro-Iglesias et al., 2009). We detected putative NECs from embryonic day (E)18.5, in clusters on the medial edges of the gills, intimately associated with HNK1 epitope-immunoreactive neurites (Figure 4d). Scattered serotonergic cells were also found in the epithelium lining the roof and floor of the orobranchial cavity. To determine any neural crest cell contribution, the vital lipophilic dye DiI was injected into E5 vagal neural folds and the embryos followed to E19.0 (Figure 4e–n). DiI-labeled neural crest cells migrated into the branchial arches and contributed to the branchial arch basket, as expected (McCauley and Bronner-Fraser, 2003), but putative NECs in the gills and orobranchial epithelia, despite being near DiI-labeled cells, were unlabeled (n = 19; Figure 4i,j,n; Figure 4—figure supplement 1).

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These results show that putative NECs in the internal gills and orobranchial epithelium of a frog (i.e., a lobe-finned tetrapod) and the sea lamprey (a jawless fish) are not neural crest-derived.

Gill NECs are endoderm-derived, like PNECs

Overall, our data show that the neural crest does not contribute to gill NECs in zebrafish, or to their presumed homologues in Xenopus and lamprey gills (or to the putative NECs identified in the orobranchial epithelium of all three species). Hence, glomus cells and gill NECs cannot have evolved from the same ancestral cell population. In zebrafish, Xenopus and the little skate (a cartilaginous fish), vital dye fate-mapping experiments have shown that the gills and orobranchial cavity are lined with an epithelium derived mostly from endoderm (Warga and Nüsslein-Volhard, 1999; Chalmers and Slack, 2000; Gillis and Tidswell, 2017), suggesting endoderm as an alternative origin for NECs. Indeed, in one of the first descriptions of fish gill NECs (Dunel-Erb et al., 1982), they were likened to the PNECs of amniotes, which share a common embryonic origin with other airway epithelial cells in rodents (Hoyt et al., 1990; Rawlins et al., 2009; Song et al., 2012; Kuo and Krasnow, 2015). We demonstrated the endodermal origin of PNECs in the mouse by lineage-tracing using the Sox17^2A-iCre driver line (in which all endoderm-derived lineages, as well as vascular endothelial cells and the hematopoietic system, express Cre; Engert et al., 2009) crossed to the R26R^lacZ or R26R^tdTomato reporter lines (Soriano, 1999; Madisen et al., 2010) (Figure 5a–e’’’). We also confirmed the recent exclusion by Wnt1-Cre lineage-tracing (Danielian et al., 1998) of a neural crest contribution to mouse PNECs (Kuo and Krasnow, 2015) (Figure 5—figure supplement 1a–b’). Similarly, we saw no neural crest contribution to PNECs in the chicken lung after labeling the premigratory neural crest by neural fold grafting from GFP-transgenic donor embryos (McGrew et al., 2008) (Figure 5—figure supplement 1c–e’).

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To test the hypothesis that NECs, like PNECs, are endoderm-derived, we first attempted to label the pharyngeal endoderm of Xenopus embryos at stage 14 via focal DiI injections into anterior endoderm (Chalmers and Slack, 2000) (Figure 5f–i). Only three embryos with endoderm-specific DiI labeling survived to stage 45 for analysis. The DiI labeling was very sparse, but in one embryo, 15 serotonergic cells in the orobranchial epithelium (putative NECs) were DiI-labeled, supporting an endodermal origin (Figure 5g–i’’).

Since the direct labeling approach in Xenopus proved to be technically challenging, we used genetic lineage-tracing of endodermal sox17 expression in zebrafish, inducing gastrulation-stage Cre expression and recombination in embryos from crosses between a sox17:creER^T2 zebrafish driver line [Tg(sox17:creER^T2;cmlc2:DsRed); Joseph J. Lancman, Keith P. Gates, and P. Duc S. Dong, personal communication, March, 2017] and the switchable reporter line Tg(-3.5ubi:loxP-GFP-loxP-mCherry) (Mosimann et al., 2011) (Figure 5j–n’’). At 8-dpf, mCherry expression was seen in both gill NECs and putative NECs in the orobranchial epithelium: 147/331 serotonergic cells counted across six larvae (≥36 cells counted per fish) were mCherry-positive (Figure 5n–n”). [The relatively low labeling efficiency likely results from a lack of optimization of the 4-OHT dose for the Tg(sox17:creER^T2;cmlc2:DsRed) driver in combination with this particular switchable reporter line (Mosimann et al., 2011).] These data demonstrate an endodermal origin in zebrafish for gill NECs, and also for putative NECs in the orobranchial epithelium. (Our genetic lineage-tracing data also confirm the endodermal origin of zebrafish gill filament epithelium, previously reported from vital dye fate-mapping experiments; Warga and Nüsslein-Volhard, 1999.)

Taken together, these results reveal the endodermal origin of gill NECs and putative NECs in the orobranchial epithelium. This supports the shared evolutionary ancestry of NECs with endoderm-derived PNECs, rather than neural crest-derived glomus cells.

NECs do not express Phox2b, which is essential for glomus cell development

It is formally possible that, despite their different embryonic origins, glomus cells and NECs could still be homologous cell types through activation of the same genetic network. Although the molecular basis of NEC development has not been investigated, the basic helix-loop-helix transcription factor Ascl1 (Mash1) is required for the formation of both PNECs (Ito et al., 2000) and glomus cells (Kameda, 2005). The homeodomain transcription factor Phox2b is also essential for glomus cell development (Dauger et al., 2003). However, we were unable to detect Phox2b-positive cells in embryonic zebrafish gills or orobranchial epithelium at 5-dpf (n = 4) or 7-dpf (n = 3), although Phox2b was expressed by a subset of cells in the hindbrain, as expected (Coppola et al., 2012) (Figure 6a–f’). Similarly, we could not detect any Phox2-positive cells in sea lamprey gills or orobranchial epithelia at E16 or E18 (n = 6), although Phox2 was expressed in the epibranchial ganglia, and in patches of ectoderm and subjacent mesenchyme ventral to the epibranchial ganglia (Figure 6g–j’), in the same position as the hypobranchial placodes and associated ganglia identified in Xenopus (Schlosser, 2003).

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We also found that PNECs lack Phox2b expression. In mouse embryos at E16.5, when PNECs can be identified by Ascl1 expression (serotonin is only expressed in a few PNECs at this stage), Phox2b was not seen in PNECs, despite expression in nearby intrinsic pulmonary ganglia (n = 2; Figure 6k–l’). Similarly, in chicken embryos at E12-E13.5, when scattered PNECs can be identified in the lung epithelium by serotonin immunoreactivity, Phox2b expression was not seen in the lung epithelium either by in situ hybridization or by immunostaining, although it could be detected in nearby intrinsic pulmonary ganglia (n = 3; Figure 6m,m’).

Overall, these data show that NECs (and PNECs) do not activate the same genetic network as glomus cells, since they lack expression of a transcription factor, Phox2b, which is essential for glomus cell development (Dauger et al., 2003). Hence, NECs and glomus cells cannot be homologous cell types.

Candidate glomus cell homologues in fish: neural crest-derived chromaffin cells associated with pharyngeal arch blood vessels

Since NECs are endoderm-derived, we reasoned that neural crest-derived glomus cells must have evolved independently. Glomus cells are catecholaminergic, like the neural crest-derived chromaffin cells of the adrenal gland, which are also hypoxia-sensitive (reviewed by López-Barneo et al., 2016). Intriguingly, in lampreys, catecholaminergic (chromium salt-staining, i.e., ‘chromaffin’) cells were reported a century ago in association with large blood vessels not only in the trunk, but also as far rostrally as the second branchial arch, ‘in the walls of the segmental veins as these run round the notochord’ (Giacomini, 1902; Gaskell, 1912). This suggested to us the possibility that glomus cells could have evolved from catecholaminergic cells associated with blood vessels in anamniote pharyngeal arches, if such cells are neural crest-derived.

We first used immunostaining for the catecholaminergic marker tyrosine hydroxylase and the neurite-marker acetylated tubulin to confirm the existence of catecholaminergic cells, at least some of which may be innervated, in the walls of the anterior cardinal veins in the gill arches of ammocoete-stage sea lamprey (Figure 7a–c). It would be difficult to test whether the gill arch catecholaminergic cells are neural crest-derived: even if DiI-labeled embryos could be raised to ammocoete stages, the DiI-labeling would likely be very sparse. We reasoned that if present in lamprey, such cells might also be present in the zebrafish. We sectioned metamorphic juveniles and identified similar clusters of catecholaminergic cells in close association with blood vessels in the gill arches (Figure 7d–e’’’). To our knowledge, this is the first demonstration of the existence of catecholaminergic cells associated with gill arch blood vessels in a jawed anamniote. Importantly, these catecholaminergic cells were mCherry-positive, i.e., neural crest-derived, in Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) metamorphic juveniles (Figure 7e–e’’’) (51 such catecholaminergic cells were mCherry-positive across five juveniles [≥5 counted per fish]). This discovery suggests a new, speculative hypothesis for carotid body evolution, namely that it evolved in the amniote lineage via the aggregation of neural crest-derived catecholaminergic cells that were already associated with pharyngeal arch blood vessels in anamniotes.

Figure 7

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Author details

1. Dorit Hockman

   1. Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, United Kingdom
   2. Weatherall Institute of Molecular Medicine, University of Oxford, Oxford, United Kingdom
   3. Department of Molecular and Cell Biology, University of Cape Town, Cape Town, South Africa

   Contribution

   DH, conceived and designed the experiments with CVHB, performed all experiments except the Xenopus DiI injections reported in Figure 5e-i' (GS), the chicken neural fold grafts reported in Figure 5-figure supplement 1c-e' (AJB), the whole-mount zebrafish immunostaining reported in Figure 5-figure supplement 2 (KPG) and the chicken Phox2b immunostaining reported in Figure 6m,m' (BJ), analyzed all the results except the zebrafish skin NEC data (KPG), prepared all the figures and wrote the manuscript with CVHB

   Competing interests

   The authors declare that no competing interests exist.

2. Alan J Burns

   1. Stem Cells and Regenerative Medicine, UCL Great Ormond Street Institute of Child Health, London, United Kingdom
   2. Department of Clinical Genetics, Erasmus Medical Center, Rotterdam, The Netherlands

   Present address

   Gastrointestinal Drug Discovery Unit, Takeda Pharmaceuticals United States, Inc., Cambridge, United States

   Contribution

   AJB, performed the chicken neural fold grafts reported in Figure 5-figure supplement 1c-e', provided Wnt1:cre;R26R:YFP mouse embryos and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

3. Gerhard Schlosser

   School of Natural Sciences, National University of Ireland, Galway, Ireland

   Contribution

   GS, performed the Xenopus DiI injections reported in Figure 5e-i' and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

4. Keith P Gates

   Human Genetics Program, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, United States

   Contribution

   KPG, performed, imaged and analyzed the whole-mount zebrafish immunostaining data reported in Figure 5-figure supplement 2, provided (with JJL and PDSD) 4-OHT-treated Tg(sox17:creER^T2;cmlc2:DsRed);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) zebrafish larvae, and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

5. Benjamin Jevans

   Stem Cells and Regenerative Medicine, UCL Great Ormond Street Institute of Child Health, London, United Kingdom

   Contribution

   BJ, performed and imaged the chicken Phox2b immunostaining reported in Figure 6m,m', and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

6. Alessandro Mongera

   Department of Genetics, Max-Planck Institut für Entwicklungsbiologie, Tübingen, Germany

   Present address

   Department of Mechanical Engineering, California NanoSystem Institute, University of California, Santa Barbara, Santa Barbara, United States

   Contribution

   AM, provided 4-OHT-treated Tg(-4.9sox10:creER^T2);Tg(βactin:loxP-SuperStop-loxP-DsRed) zebrafish larvae and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

7. Shannon Fisher

   Department of Cell and Developmental Biology, University of Pennsylvania, Philadelphia, United States

   Present address

   Department of Pharmacology & Experimental Therapeutics, Boston University School of Medicine, Boston, United States

   Contribution

   SF, provided Tg(-28.5sox10:cre);Tg(ef1a:loxP-DsRed-loxP-EGFP) zebrafish larvae and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

8. Gokhan Unlu

   Division of Genetic Medicine, Department of Medicine, Vanderbilt University Medical Center, Nashville, United States

   Contribution

   GU, provided (with EWK) tfap2a^mob;foxd3^mos zebrafish larvae and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

9. Ela W Knapik

   Division of Genetic Medicine, Department of Medicine, Vanderbilt University Medical Center, Nashville, United States

   Contribution

   EWK, provided (with GU) tfap2a^mob;foxd3^mos zebrafish larvae and commented on the manuscript

   Competing interests

   The authors declare that no competing interests exist.

10. Charles K Kaufman

    Children’s Hospital Boston, Howard Hughes Medical Institute, Harvard Medical School, Boston, United States

    Present address

    1. Department of Medicine, Washington University School of Medicine, St. Louis, United States
    2. Division of Oncology, Washington University School of Medicine, St. Louis, United States
    3. Department of Developmental Biology, Washington University School of Medicine, St. Louis, United States

    Contribution

    CKK, provided (with CM and LIZ) 4-OHT-treated Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) zebrafish larvae and juveniles, and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

11. Christian Mosimann

    Children’s Hospital Boston, Howard Hughes Medical Institute, Harvard Medical School, Boston, United States

    Present address

    Institute of Molecular Life Sciences, University of Zürich, Zürich, Switzerland

    Contribution

    CM, provided (with CKK and LIZ) 4-OHT-treated Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) zebrafish larvae and juveniles, and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

12. Leonard I Zon

    Children’s Hospital Boston, Howard Hughes Medical Institute, Harvard Medical School, Boston, United States

    Contribution

    LIZ, provided (with CKK and CM) 4-OHT-treated Tg(crestin:creER^T2);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) zebrafish larvae and juveniles, and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

    "This ORCID iD identifies the author of this article:" 0000-0003-0860-926X

13. Joseph J Lancman

    Human Genetics Program, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, United States

    Contribution

    JJL, provided (with KPG and PDSD) 4-OHT-treated Tg(sox17:creER^T2;cmlc2:DsRed);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) zebrafish larvae, and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

14. P Duc S Dong

    Human Genetics Program, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, United States

    Contribution

    PDSD, provided (with KPG and JJL) 4-OHT-treated Tg(sox17:creER^T2;cmlc2:DsRed);Tg(-3.5ubi:loxP-GFP-loxP-mCherry) zebrafish larvae, and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

15. Heiko Lickert

    Institute of Diabetes and Regeneration Research, Helmholtz Zentrum München, Neuherberg, Germany

    Contribution

    HL, provided access to the Sox17^2A-iCre mouse line and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

16. Abigail S Tucker

    Department of Craniofacial Development and Stem Cell Biology, King’s College London, London, United Kingdom

    Contribution

    AST, provided Sox17^2A-iCre/+;R26^R/+ mouse embryos and cryosections of Sox17^2A-iCre/+;R26R^tdTomatomouse embryos, and commented on the manuscript

    Competing interests

    The authors declare that no competing interests exist.

17. Clare V H Baker

    Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, United Kingdom

    Contribution

    CVHB, conceived and designed the experiments with DH and wrote the manuscript with DH

    For correspondence

    cvhb1@cam.ac.uk

    Competing interests

    The authors declare that no competing interests exist.

    "This ORCID iD identifies the author of this article:" 0000-0002-4434-3107

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