Xavier AL , Fontaine R , Bloch S , Affaticati P , Jenett A , Demarque M , Vernier P , Yamamoto K .

	Figure 1. Open in figure viewer Monoaminergic CSF-c cells of chicken, Xenopus, and zebrafish in relation to the organization of the hypothalamic ventricle. Schematic drawings of sagittal sections of chicken, Xenopus, and zebrafish brains are shown in the top panels, and for each species, micrographs of the red squared areas are shown below (a–c for chicken, d–f for Xenopus, and g–i for zebrafish). DAPI staining (gray) delineates the recesses of the hypothalamic ventricle. Confocal images (Z-projection = 10 µm) with higher magnification obtained from the area depicted in (a) (dashed green square) show 5-HT+ CSF-c cells (green) aligned along the dorsal side of the hypothalamic recess (b; inset at higher magnification). TH immunoreactive cells (orange) are not observed within the PVO (arrowhead), but are abundant in the area dorsocaudal to it (c; asterisk). In the Xenopus sagittal section close to the midline, PVO (d; arrowhead) is observed at the anterior edge of the large ventricle (v). The PVO is visualized with 5-HT+ CSF-c cells (e; green; inset at higher magnification). TH immunoreactive cells (orange) are observed dorsal to the PVO (f; asterisk). In zebrafish, three CSF-c cell populations (locations indicated by arrowheads in g) are located around two hypothalamic recesses. The two anterior CSF-c cell populations are located in front of and around the lateral recess (LR), while the posterior population surrounds the posterior recess (PR). Higher magnification of the squared area in (g) is shown in (h) and (i) (Z-projection = 10 µm). CSF-c cells revealed by the expression of GFP in the enhancer trap transgenic line ETvmat2:GFP (vmat2-GFP; green inset) are lined along the ventricular zone (h). The white inset in (h) shows the 5-HT labeling in the same area (the image is taken from a different sample). TH immunoreactive cells (orange) are found dorsal to the LR (i; asterisk). D = dorsal; Die = diencephalon; Hyp = hypothalamus; LR = lateral recess; PR = posterior recess; PVO = paraventricular organ; R = rostral; v = ventricle. Scale bar = 200 µm in (a–g); 50 µm in (h, i)
	Figure 2. Open in figure viewer 5-HT+ CSF-c cells in the Xenopus PVO. The laterally extended hypothalamic recess (lateral recess; LR) is visualized with DAPI staining (magenta) from a frontal section (midline to the left). CSF-c cells immunolabeled for 5-HT (green) are located medially in the rostral hypothalamus and laterally in the caudal hypothalamus. (a) depicts both DAPI and 5-HT stainings, while (b) shows 5-HT only (same picture). In the projection of confocal images (15 µm), the rostromedial and caudolateral CSF-c cells look continuous. Scale bar = 50 µm
	Figure 6. Open in figure viewer Localization of th1, th2, and TH immunoreactivity in the PVO of Xenopus. Confocal images obtained from a frontal section of the Xenopus PVO are shown (Z projections = 10 µm). Counterstaining with DAPI (gray) shows that th2 (magenta) is expressed in the region corresponding to PVO (a). A cell population displaying strong th1 signal (green) is found dorsolaterally to the PVO (asterisks in b), which overlaps with TH immunoreactivity (orange; asterisks in c). Higher magnification of the area delimited by a dashed square in (c) is shown in (d–f). Faint TH immunoreactivity is also found in CSF-c cells in the Xenopus PVO. TH+ labeling is observed in the soma and in the processes bathing the ventricle of CSF-contacting cells (arrowheads in d–f). A few CSF-contacting cells expressing th2 are also TH immunoreactive (arrows in d–f). D = dorsal; L = lateral. Scale bar = 50 µm in (a) (applies to a, b, c); 100 µm in (d) (applies to d, e, f)
	Figure 7. Colocalization of TH2 and TPH1 in the PVO of chicken and Xenopus. Frontal sections obtained from the PVO of chicken (a–d) and Xenopus (e–l) are shown. TH2 (a; magenta) and TPH1 (b; green) transcripts are observed in the same region of the chicken PVO, and some CSF-c cells coexpress both genes (c, d; white). The area depicted by a dashed rectangle in (c) is shown in (d1–d3) at higher magnification. Optical sectioning by confocal microscopy shows that TH2 (d2) and TPH1 (d3) are expressed in the soma (arrowheads) and in the processes of CSF-contacting cells (arrow). Orthogonal views of optical sections confirm the overlap of TH2 and TPH1 signals. In Xenopus, th2 (e, i; magenta) and tph1 (f, j; green) are expressed both in the rostromedial (e–h) and the caudolateral (i–l) parts of the PVO. Some CSF-c cells coexpress both genes (g, k). The coexpression is confirmed by orthogonal views of optical sections (h1) of the area depicted by a dashed rectangle in (g), showing that th2 (h2) and tph1 (h3) are colocalized in the same cells (arrowheads). D = dorsal; L = lateral; LR = lateral recess; v = ventricle. Scale bar = 100 µm in (a) (applies to a, b, c) and in (e) (applies to e, f, g); 50 µm in (i) (applies to i, j, k, l)
	Figure 10. Comparison of HuC/D immunoreactivity in the anterior PVO cell populations in chicken, Xenopus, and zebrafish. Frontal sections of the anterior PVO of chicken (a–e), Xenopus (f–j), and zebrafish (k–o) are shown. The HuC/D immunoreactivity (dark blue) is absent in the PVO, where monoaminergic CSF-c cells are located (dashed lines), in chicken (a–d), Xenopus (f–i), and zebrafish (k–n). The areas within dashed squares in (d), (i), and (n) are shown at higher magnification in (e), (j), and (o), respectively. TH2+/TPH1+ CSF-c cells (white in e; arrows) are not HuC/D+ (e; arrowheads) in the chicken PVO. In the Xenopus PVO as well, monoaminergic 5-HT+/TH+ CSF-c cells (j; arrows) are not immunolabeled by HuC/D, whereas TH+ adjacent to the PVO are HuC/D+ (j; arrowheads). Similarly, CSF-c cells that express GFP in the enhancer trap transgenic line ETvmat2:GFP (vmat2-GFP; cyan) are HuC/D- (n). HuC/D+ cells (o; arrowheads) are adjacent to vmat2-GFP cells (o; arrow) whose process bathes the ventricle (o; curved arrowhead). D = dorsal; L = lateral; v = ventricle. Scale bar = 50 µm in (a) (applies to a, b, c, d); 50 µm in (f) (applies to f, g, h, i); 50 µm in (k) (applies to k, l, m, n)
	Figure 12. Open in figure viewerDownload Powerpoint slide Comparison of monoaminergic cell populations in different groups of Osteichthyes. Some comparable monoaminergic cell groups are plotted on schematic drawings of brain sections of mouse, chicken, Xenopus, and zebrafish. The sagittal plane is close to the midline (rostral to the left). The diamonds represent CSF-c cells, and the circles represent non-CSF-c cells. TH2/TPH1-expressing CSF-c cells (red diamonds) are commonly found along the hypothalamic recess throughout vertebrates, while placental mammals have lost the CSF-c cells in the hypothalamic region. Below the sagittal section of chicken, Xenopus, and zebrafish, frontal sections around the hypothalamic recess (corresponding to the level of gray lines; a,b,c) are shown. In chicken and Xenopus, there is only one hypothalamic recess (named either 3rd ventricle or lateral recess; LR), while in zebrafish, there is an additional recess (posterior recess; PR). Note that in Xenopus, the caudally located CSF-c cells are not observable in the sagittal section close to the midline, because it is located at the lateral end of the LR. The A11-like TH1-expressing cell population (blue dots; non-CSF-c cells) projecting to the spinal cord is commonly found dorsolateral to the CSF-c cells in all the vertebrate groups. More caudally, the A9/10-like DA cell population projecting to the telencephalon (green dots) is found in tetrapods, while this cell population is lacking in teleosts. L = lateral recess; M = mesencephalon; P = prosencephalon; PR = posterior recess; R = rhombencephalon
	Figure 3. Organization of the two hypothalamic recesses in zebrafish. The hypothalamic area of adult (a–e; sagittal view) and embryonic (f, g; horizontal view) zebrafish brains are shown. The anterior is to the left in all figures. 3D visualization of the images of (a) and (b) are shown in the Supporting Information Figures S1 and S2 respectively. (a) The three vmat2‐GFP CSF‐c cells (green; PVO, IN, and Hc) are organized around the two hypothalamic recesses, namely the lateral recess (LR) and the posterior recess (PR). The CSF‐c cells (green) are visualized using an enhancer trap vmat2:GFP zebrafish line, and the ventricular zone (magenta) is reconstructed from ventricular surfaces delineated by DiI staining. (b) Immunolabeling for vmat2‐GFP (green) and TH (orange). Prominent TH‐immunopositive cells (orange) are found dorsal to the CSF‐c cells populations (green): PVO, IN, and Hc, which are alternatively named PVOa, PVOi, and PVOp. (c–e) Adult hypothalamic recesses double labeled for vmat2‐GFP and ZO‐1. The areas delimited by dashed rectangles in (c) are shown in (d) and (e), which demonstrate the end feet of the process of CSF‐c cells bathing in the ventricle. (f, g) The ZO‐1 immunolabeling in the developing brain (48 hr postfertilization), demonstrating that the LR and PR are separate ventricular extension from early embryonic stages. (g) shows 3D semiautomatic segmentation of the ZO‐1 immunolabeling in the LR and PR shown in (f). The right hemisphere and the midline are visualized in green, while the left hemisphere is visualized in white, to highlight the intricate ventricular organization. C = caudal; D = dorsal; Hc = caudal zone of periventricular hypothalamus; IN = intermediate nucleus of hypothalamus; LR = lateral recess; PR = posterior recess; PVO = paraventricular organ; PVOa = anterior paraventricular organ; PVOi = intermediate paraventricular organ; PVOp = posterior paraventricular organ; R = rostral. Scale bars = 50 µm in (c) and (f); 10 µm in (d) and (e)
	Figure 4. Localization of TH1, TH2, and TH immunoreactivity in the PVO of chicken. Representative micrographs of chicken brains in sagittal sections (a–e) and frontal sections (f–h) are shown. Abundant labeling of TH2 transcripts is found all along the hypothalamic recess (a; magenta). In the inset of (a), the expression of TH2 by PVO CSF‐c cells is shown at higher magnification. Double labeling with TH1 (green) shows that there is no overlap for the expression of the two genes in the PVO (b). Confocal images (Z projections = 10 µm) of the squared area in (b) are shown in (c–e). TH2 transcripts are localized along the PVO, in the area delimited by dashed lines (c). In contrast, TH1 is absent in the PVO (d), as are TH immunoreactive cells (e; orange). The micrograph of the frontal section (ventricular zone to the left) also shows that the PVO contains TH2 transcripts (f; magenta), but not TH1 (g; green) nor TH immunoreactive cells (h; orange). TH1 expressing cells and TH immunoreactive cells are observed dorsally to the PVO (asterisks in b–h). D = dorsal; Hyp = hypothalamus; Inf = infundibulum; L = lateral; R = rostral; Tel = telencephalon; TeO = optic tectum; v = ventricle. Scale bars = 1 mm in (a) (applies to a, b); 200 µm in (c) (applies to c, d, e); 100 µm in (f) (applies to f, g, h)
	Figure 5. Double labeling for TH2 nonfluorescent in situ hybridization and TH immunofluorescence in chicken. A sagittal section of chicken brain showing TH2 transcripts (purple) and TH immunoreactivity (green) in the hypothalamic (a, b) and mesencephalic (c, d) areas. TH2 transcripts are abundant in the PVO (a; dashed area) and scarce in a cell population corresponding to A11 (a; asterisk). In contrast, TH immunoreactive cells are abundant in the A11 cell group (b; asterisk), while they are absent in the PVO (b; dashed area). The mesencephalic TH immunoreactive cells also express TH2 (c; arrows), but compared to the TH immunoreactivity (d; arrows), the TH2 + signal is very low. D = dorsal; Hyp = hypothalamus; Inf = infundibulum; M = mesencephalon; R = rostral. Scale bar = 200 µm in (a) (applies to a, b, c, d)
	Figure 8. Colocalization of th2 and tph1a in the three CSF‐c cell populations of zebrafish. Frontal sections of the adult zebrafish brain showing the three CSF‐c cell populations, PVO (a–c), IN (d–f), and Hc (g–i), which are also known as PVOa, PVOi, and PVOp, respectively. Single confocal planes show that th2 (magenta) and tph1a (green) are found in the same cell population, and some cells coexpress both of them (arrows). Scale bar = 20 µm
	Figure 9. Colocalization of DA and 5‐HT in the PVO cells of zebrafish. Frontal sections of the anterior PVO (PVOa) in adult zebrafish demonstrate that CSF‐c cells are immunoreactive to DA (magenta; a, d) and to 5‐HT (green; b, e). The areas in dashed rectangles in (a), (b), and (c) are shown at higher magnification (Z‐projection = 5 µm) in (d), (e), and (f) respectively. Both monoamines were observed in a few cell bodies (arrow in f) and endfeet (arrowhead in f). In teleosts, some of the DA+ CSF‐c cells (g) are also immunoreactive for TH (h; orange). Higher magnification images of the PVO are shown in (i–l). In CSF‐c cells, intense DA immunoreactivity is present in the cell soma (j; arrows), processes, and the endfeet contacting the ventricle (k; arrowheads). In contrast, intense TH immunoreactivity is mostly observed in the soma and processes (l; arrows), but not in the endfeet. D = dorsal; L = lateral; v = ventricle. Scale bar = 50 µm in (a) (applies to a, b, c); 200 µm in (g) (applies to g, h); 100 µm in (i) (applies to i, j, k, l)
	Figure 11. HuC/D immunoreactivity in the caudal CSF‐c cell populations in zebrafish. Frontal sections of the zebrafish IN (a and b) and Hc (c and d), with the midline to the left. Gray represents DAPI staining. The tph1a + CSF‐c cells (green) are negative for HuC/D (magenta). Scale bar = 50 µm (applies to a, b, c, d)

Affaticati, Identification of the optic recess region as a morphogenetic entity in the zebrafish forebrain. 2016, Pubmed

Affaticati, Identification of the optic recess region as a morphogenetic entity in the zebrafish forebrain. 2016, Pubmed
                     Alunni, Notch3 signaling gates cell cycle entry and limits neural stem cell amplification in the adult pallium. 2013, Pubmed
                     Bellipanni, Cloning of two tryptophan hydroxylase genes expressed in the diencephalon of the developing zebrafish brain. 2016, Pubmed
                     Biran, Role of developmental factors in hypothalamic function. 2015, Pubmed
                     Björklund, Evidence for a major spinal cord projection from the diencephalic A11 dopamine cell group in the rat using transmitter-specific fluorescent retrograde tracing. 1980, Pubmed
                     Bosco, Development of hypothalamic serotoninergic neurons requires Fgf signalling via the ETS-domain transcription factor Etv5b. 2013, Pubmed
                     Chen, Complementary developmental expression of the two tyrosine hydroxylase transcripts in zebrafish. 2010, Pubmed
                     Chung, Structural and molecular interrogation of intact biological systems. 2013, Pubmed
                     Corio, Distribution of catecholaminergic and serotoninergic systems in forebrain and midbrain of the newt, Triturus alpestris (Urodela). 1992, Pubmed
                     Ekström, Distribution of dopamine-immunoreactive neuronal perikarya and fibres in the brain of a teleost, Gasterosteus aculeatus L. comparison with tyrosine hydroxylase- and dopamine-beta-hydroxylase-immunoreactive neurons. 1990, Pubmed
                     Filippi, vglut2 and gad expression reveal distinct patterns of dual GABAergic versus glutamatergic cotransmitter phenotypes of dopaminergic and noradrenergic neurons in the zebrafish brain. 2015, Pubmed
                     Filippi, Expression of the paralogous tyrosine hydroxylase encoding genes th1 and th2 reveals the full complement of dopaminergic and noradrenergic neurons in zebrafish larval and juvenile brain. 2010, Pubmed
                     Fontaine, Dopamine inhibits reproduction in female zebrafish (Danio rerio) via three pituitary D2 receptor subtypes. 2013, Pubmed
                     Fontaine, Dopaminergic Neurons Controlling Anterior Pituitary Functions: Anatomy and Ontogenesis in Zebrafish. 2015, Pubmed
                     Gaspar, Probing the diversity of serotonin neurons. 2012, Pubmed
                     Hirunagi, Immunocytochemical demonstration of serotonin-immunoreactive cerebrospinal fluid-contacting neurons in the paraventricular organ of pigeons and domestic chickens. 1992, Pubmed
                     Hornby, Localization of immunoreactive tyrosine hydroxylase in the goldfish brain. 1987, Pubmed
                     Kang, Dopamine-melatonin neurons in the avian hypothalamus controlling seasonal reproduction. 2008, Pubmed
                     Kang, Melanopsin expression in dopamine-melatonin neurons of the premammillary nucleus of the hypothalamus and seasonal reproduction in birds. 2011, Pubmed
                     Kaslin, Comparative anatomy of the histaminergic and other aminergic systems in zebrafish (Danio rerio). 2002, Pubmed
                     Kawakami, A transposon-mediated gene trap approach identifies developmentally regulated genes in zebrafish. 2004, Pubmed
                     Kotrschal, Distribution of aminergic neurons in the brain of the sterlet, Acipenser ruthenus (Chondrostei, Actinopterygii). 1985, Pubmed
                     López, Organization of the serotonergic system in the central nervous system of two basal actinopterygian fishes: the Cladistians Polypterus senegalus and Erpetoichthys calabaricus. 2014, Pubmed
                     Mayrhofer, A novel brain tumour model in zebrafish reveals the role of YAP activation in MAPK- and PI3K-induced malignant growth. 2017, Pubmed
                     McPherson, Motor Behavior Mediated by Continuously Generated Dopaminergic Neurons in the Zebrafish Hypothalamus Recovers after Cell Ablation. 2016, Pubmed
                     Medina, Development of catecholamine systems in the brain of the lizard Gallotia galloti. 1995, Pubmed
                     Meek, Distribution of dopamine immunoreactivity in the brain of the mormyrid teleost Gnathonemus petersii. 1989, Pubmed
                     Meek, Distribution of serotonin in the brain of the mormyrid teleost Gnathonemus petersii. 1989, Pubmed
                     Meek, Distribution of noradrenaline-immunoreactivity in the brain of the mormyrid teleost Gnathonemus petersii. 1993, Pubmed
                     Meneghelli, Distribution of tryptophan hydroxylase-immunoreactive neurons in the brainstem and diencephalon of the pigeon (Columba livia). 2009, Pubmed
                     Nakane, A mammalian neural tissue opsin (Opsin 5) is a deep brain photoreceptor in birds. 2010, Pubmed , Xenbase
                     Neary, Nuclear organization of the bullfrog diencephalon. 1983, Pubmed
                     Okano, A hierarchy of Hu RNA binding proteins in developing and adult neurons. 1997, Pubmed
                     Parent, The monoamine-containing neurons in the brain of the garfish, Lepisosteus osseus. 1983, Pubmed
                     Parent, The organization of monoamine-containing neurons in the brain of the sunfish (Lepomis gibbosus) as revealed by fluorescence microscopy. 1979, Pubmed
                     Puelles, Forebrain gene expression domains and the evolving prosomeric model. 2003, Pubmed
                     Puelles, Expression patterns of homeobox and other putative regulatory genes in the embryonic mouse forebrain suggest a neuromeric organization. 1994, Pubmed
                     Ren, Zebrafish tyrosine hydroxylase 2 gene encodes tryptophan hydroxylase. 2013, Pubmed
                     Rink, The teleostean (zebrafish) dopaminergic system ascending to the subpallium (striatum) is located in the basal diencephalon (posterior tuberculum). 2001, Pubmed
                     Semenova, The tyrosine hydroxylase 2 (TH2) system in zebrafish brain and stress activation of hypothalamic cells. 2015, Pubmed
                     Skagerberg, Origin and termination of the diencephalo-spinal dopamine system in the rat. 1983, Pubmed
                     Smeets, Are putative dopamine-accumulating cell bodies in the hypothalamic periventricular organ a primitive brain character of non-mammalian vertebrates? 1990, Pubmed
                     Sánchez-Camacho, Descending supraspinal pathways in amphibians. II. Distribution and origin of the catecholaminergic innervation of the spinal cord. 2001, Pubmed , Xenbase
                     Tay, Comprehensive catecholaminergic projectome analysis reveals single-neuron integration of zebrafish ascending and descending dopaminergic systems. 2019, Pubmed
                     Treweek, Whole-body tissue stabilization and selective extractions via tissue-hydrogel hybrids for high-resolution intact circuit mapping and phenotyping. 2016, Pubmed
                     Ueda, Immunohistochemical demonstration of the serotonin neuron system in the central nervous system of the bullfrog, Rana catesbeiana. 1984, Pubmed
                     Vigh, Nonvisual photoreceptors of the deep brain, pineal organs and retina. 2002, Pubmed
                     Vígh, The system of cerebrospinal fluid-contacting neurons. Its supposed role in the nonsynaptic signal transmission of the brain. 2004, Pubmed
                     Wakamatsu, Sequential expression and role of Hu RNA-binding proteins during neurogenesis. 1997, Pubmed
                     Walther, Synthesis of serotonin by a second tryptophan hydroxylase isoform. 2003, Pubmed
                     Walther, A unique central tryptophan hydroxylase isoform. 2003, Pubmed
                     Wen, Visualization of monoaminergic neurons and neurotoxicity of MPTP in live transgenic zebrafish. 2008, Pubmed
                     Xavier, Comparative analysis of monoaminergic cerebrospinal fluid-contacting cells in Osteichthyes (bony vertebrates). 2017, Pubmed , Xenbase
                     Yamamoto, Two tyrosine hydroxylase genes in vertebrates New dopaminergic territories revealed in the zebrafish brain. 2010, Pubmed
                     Yamamoto, Differential expression of dopaminergic cell markers in the adult zebrafish forebrain. 2011, Pubmed
                     Yamamoto, The evolution of dopamine systems in chordates. 2011, Pubmed
                     Yoshida, Monoaminergic neurons in the brain of goldfish as observed by immunohistochemical techniques. 1983, Pubmed
                     Zhang, Establishment of a neuroepithelial barrier by Claudin5a is essential for zebrafish brain ventricular lumen expansion. 2010, Pubmed