Reproduction requires the adequate integration of var-ious sensory cues, including odors and/or visual stimuliassociated, for example, with feathers, fur, and egg pat-terns; auditory stimuli such as songs in birds; or tactilestimuli specific to reproduction (e.g., Murphy and Schnei-der, 1970; Konishi, 1973; Keverne et al., 1983; Ball andBalthazart, 2002; Blaustein and Erskine, 2002; Hull et al.,2002). The period of reproduction is often correlated withchanges in sensory acuity that are potentially controlledby estrogens and that are a prerequisite for the adequateintegration of reproductive sensory cues. For instance, infemale canaries, estrogens enhance the response to a lightmechanical stimulation of the ventral brood patch, a pro-cess that is required for the successful construction of thenest and adequate incubation of the eggs (Hinde andSteel, 1964; Hutchison et al., 1967). In female rats, estro-gens facilitate lordosis behavior by enlarging the sensoryfield of the pudendal nerve, which contributes to an en-hanced perception of the male-derived lordosis-inducing
stimulus (Komisaruk et al., 1972; Kow and Pfaff, 1973).Estrogens also regulate the perception of male olfactorysignals that facilitate lordosis behavior in rodents (Chabliet al., 1985). In addition to experiments showing an inter-action between sex steroids, sensory acuity and reproduc-tion, there is a wealth of evidence indicating that sexsteroids potentially modulate most, if not all, sensory mo-
Grant sponsor: National Institute of Mental Health; Grant number:NIMH50388. Grant sponsor: Belgian Fonds de la Recherche FondamentaleCollective; Grant number: 2.4555.01; Grant sponsor: Government of theFrench Community of Belgium; Grant number: ARC99/04-241.
*Correspondence to: Henry C. Evrard, Boston University, Department ofBiology, 5 Cummington Street, Boston MA 02215.E-mail: [email protected]
Received 6 June 2003; Revised 18 December 2003; Accepted 26 Decem-ber 2003
DOI 10.1002/cne.20068Published online in Wiley InterScience (www.interscience.wiley.com).
THE JOURNAL OF COMPARATIVE NEUROLOGY 473:194–212 (2004)
dalities, including nociception (e.g., Bereiter and Barker,1980; Klimek, 1985; Duval et al., 1996; Amandusson et al.,1999; Bradshaw et al., 2000; Gintzler and Liu, 2001; Pajotet al., 2003; Evrard and Balthazart, 2004; Ji et al., 2003),olfaction (Hara, 1967; Deems et al., 1991; Hummel et al.,1991; Guillot and Chapouthier, 1996), light touch sensi-tivity (Burris et al., 1991; Romanzi et al., 2001), taste(Zucker et al., 1972; Mascarenhas et al., 1992; Than et al.,1994; Alberti-Fidanza et al., 1998; Li and Hay, 2000),audition (Cox, 1980; Yovanof and Feng, 1983; Elkind-Hirsch et al., 1992; Coleman et al., 1994; Caruso et al.,2000), and baro- and chemoreception (Tatsumi et al.,1997; Mohamed et al., 1999; Minson et al., 2000).
Numerous sensory nuclei in the brain and spinal cordcontain estrogen receptors (e.g., Martinez-Vargas et al.,1976; Sar and Stumpf, 1977; Simerly et al., 1990; Aman-dusson et al., 1995; Shughrue et al., 1997; VanderHorst etal., 1997; Simonian et al., 1998; Evrard and Balthazart,2000; Evrard and Balthazart, 2002) and, therefore, couldplay a role in the estrogen-dependent changes in sensoryacuity. However, the origin of the estrogens potentiallyregulating sensory functions by a direct action in sensorynuclei has never been carefully analyzed, but it has beenimplicitly assumed that these estrogens are from a go-nadal origin. During studies on the control of nociceptionat the spinal level, we recently demonstrated the presenceof numerous estrogen synthase (aromatase) -immuno-reactive (ARO-ir) neurons throughout the entire rostro-caudal extent of the spinal dorsal (sensory) horn in maleand female Japanese quail and in rats (Evrard et al., 2000;Evrard and Balthazart, 2001). In addition, we demon-strated that, in quail, systemic and intrathecal injectionsof an aromatase inhibitor decrease markedly responsive-ness to a noxious thermal stimulus (Evrard and Balthaz-art, 2003). On this basis, we wondered whether, in paral-lel, estrogens could be also produced in sensory nuclei ofthe hindbrain and affect the functioning of other sensorymodalities. In the present work, through the use immu-nocytochemistry, we assessed the presence of aromatasein the sensory nuclei of the hindbrain in male Japanesequail, a useful model for the study of neural aromatizationin which aromatase immunocytochemistry reliably re-flects aromatase activity (AA) and the distribution of aro-matase mRNA (Balthazart and Ball, 1998a,b).
MATERIALS AND METHODS
Seven adult male Japanese quail (Coturnix japonica; 6weeks of age, weighing approximately 200 g) purchasedfrom a local breeder (C. Dujardin, Liernu, Belgium) wereindividually housed in cages under a long day photoperiod(16 hours of light and 8 hours in dark) at a temperatureranging from 18 to 22°C. Food and water were alwaysprovided ad libitum. All experimental procedures com-plied with the Belgian laws on Protection and Welfare ofAnimals and the International Guiding Principles for Bio-medical Research Involving Animals published by theCouncil for International Organizations of Medical Sci-ences. The protocols were approved by the Ethical Com-mittee for the Use of Experimental Animals at the Uni-versity of Liege.
All subjects were injected intravenously with heparin(20 mg/ml; H-7005, Sigma-Aldrich, St. Louis, MO) anddeeply anesthetized with an intramuscular injection ofHypnodil (50 mg/kg body weight; Janssen Pharmaceutica,
Beerse, Belgium). They were perfused through the heartwith approximately 500 ml of saline solution (9%; 0.15 M)followed by approximately 500 ml of fixative (3% parafor-maldehyde, 15% of water saturated with picric acid, and0.1% glutaraldehyde in 0.1 M phosphate buffer, pH 7.2).The brain was immediately dissected out of the skull andplaced overnight in a 20% sucrose solution in 0.1 M phos-phate buffer. Brains were then frozen on powdered dry iceand stored in a freezer at �76°C until used.
Transverse sections were cut at 40 �m thickness andcollected in 0.05 M Tris buffer containing 0.125 M NaCl(TBS, pH 7.2). Alternate sections were stained by immu-nocytochemistry for aromatase (all subjects), serotonin or5-hydroxytryptamine (5HT; n � 3), tyrosine hydroxylase(TH; n � 3), or by toluidine blue to visualize Nissl bodies(all subjects). Staining for TH and 5HT was used to ascer-tain the location of the catecholaminergic and serotonin-ergic nuclei.
Transverse sections assigned to the coloration of Nisslbodies were stained by toluidine blue as described before(Evrard et al., 2000). For immunocytochemistry, sectionswere pretreated for 20 minutes at room temperature with0.6% hydrogen peroxide in TBS and then washed threetimes in TBS containing 0.1% Triton X-100 (TBST). Sec-tions were incubated 48 hours at 4°C with the primaryantibody that was either a polyclonal affinity-purified an-tibody raised in rabbit against quail recombinant aro-matase (anti-ARO; diluted at 1/500 in TBST; see Foidartet al., 1995, for description of the preparation of thisantibody and its validation and specificity for use in quail),a monoclonal antibody raised in mouse against TH (1/1,000; anti-TH; Incstar, Stillwater, MN), or a polyclonalantibody raised in rabbit against 5HT (1/50,000; anti-5HT; Incstar).
Sections were rinsed five times in TBST and incubatedfor 90 minutes at room temperature in a solution contain-ing either biotinylated goat anti-rabbit IgG for aromataseand 5HT staining (1/400; DAKO A/S, Glostrup, Denmark)or goat anti-mouse IgG for TH staining (1/400; DAKOA/S). Sections were again rinsed five times in TBST andthen reacted for 90 minutes at room temperature withTBST containing either a peroxidase–anti-peroxidasecomplex (1/1,000; DAKO A/S for TH and 5HT) or anavidin–biotin complex (VECTASTAIN Elite ABC Kit, Vec-tor Laboratories, Inc., Burlingame, CA, for aromatase).After five more washes in TBST, the bound peroxidasewas revealed by immersing sections for 10 minutes in asolution of 3,3�diaminobenzidine tetrahydrochloride(Sigma, D-5637; 20 mg in 50 ml of TBST containing 20 �lof hydrogen peroxide at 30%). Finally, sections weremounted in a gelatin medium and cover-slipped.
All sections were observed and some were digitized at300 DPI with a Leica DMBR microscope equipped with a3-CCD camera (Sony DXC-950P) connected to a digitalvideo recorder (Sony DKR-700P). Images were stored on140 Mb Sony data disks and transferred to a Macintoshcomputer. Images generated from sections treated withantibodies were recovered in the Adobe Photoshop pro-gram (Adobe Systems, Inc.). Figures including micropho-tographs were prepared from this material with no alter-ation except for the adjustments in contrast and densityperformed to obtain matching panels within a same plate.Images generated from serial 40-�m-thick Nissl-stainedsections were recovered in the program Canvas 5.0.2(Deneba Systems, Inc.) and then used to prepare sche-
195ESTROGEN SYNTHESIS IN THE HINDBRAIN
matic drawings of the hindbrain at different rostrocaudallevels (Fig. 1). Cell groups were recognized based on theircytoarchitecture and with the help of atlases and treatisesof avian neuroanatomy (e.g., Kuenzel and Masson, 1988;Hartwig, 1993; Dubbeldam, 1994). Sections stained for5HT or TH allowed the exact localization of catecholamin-ergic cell groups (e.g., nuclei of the solitary tract, centralgray, locus ceruleus) and of the serotoninergic raphe sub-nuclei as described in previous studies of the brain in quail(Cozzi et al., 1991; Bailhache and Balthazart, 1993) andother avian species (Parent, 1981; Berk, 1991; Berk et al.,1993; Reiner et al., 1994; Toledo et al., 1995). It must benoticed that the avian brain nomenclature is now in aprocess of profound revision, and some of the terms usedin the present study perhaps will be changed soon (AvianBrain Nomenclature Exchange Web site: http://jarvis.neuro.duke.edu/nomen/index.html or at http://www.avianbrain.org/). Once the transverse schemes of thehindbrain were completed, sections immunostained foraromatase were closely observed under the microscope.The distribution of ARO-ir cell bodies and fibers wascharted on the schemes (dots and grayed areas, respec-tively). Four levels of grayness were associated with fourincreasing degrees of density of ARO-ir fibers that werequalitatively assessed on an ordinal scale (Fig. 1).
RESULTS
General observations
The cytoarchitecture of the hindbrain in quail, as re-vealed by toluidine blue, closely corresponded to the orga-nization described in other birds and, to some extent, inmammals (Dubbeldam, 1994; Butler and Hodos, 1996).The distributions of the 5HT and TH matched those pre-viously reported in quail (Cozzi et al., 1991; Bailhache andBalthazart, 1993) and other birds (Parent, 1981; Berk,1991; Berk et al., 1993; Reiner et al., 1994; Toledo et al.,1995). They are, therefore, not re-described in detail.
The ARO-ir structures displayed a typical neuronalmorphology characterized by immunoreactive perikaryawith immunonegative round nuclei often associated withimmunoreactive fibers bearing punctate structures (e.g.,Fig. 2D). The expression of aromatase immunoreactivityappeared to occur in the cytoplasm and also at the level of(or close to) the cell membrane, but the image resolutionmade it impossible to determine the exact location of thesignal (e.g., internal vs. external side of the cell mem-brane). Isolated immunoreactive fibers with punctatestructures were also frequently observed (e.g., Fig. 4H).ARO-ir perikarya were usually clustered and allowed thedelimitation of specific cytoarchitectonic areas. Isolatedfibers formed scattered to dense networks that were eitherconfined, like perikarya, within specific cytoarchitectonicstructures or were spreading through important volumesof the hindbrain overlapping with several nuclei. In thefollowing paragraphs, the distribution of the ARO-ir neu-rons is presented in detail in a caudal to rostral order thatfollows the order also used in a recent review on the avianfunctional neuroanatomy that was used as the main ref-erence in our work (Dubbeldam, 1994).The different cellgroups were considered in functional systems. This distri-bution is illustrated by schematic drawings (Fig. 1) andrepresentative photomicrographs of transverse sections(Figs. 2–4). These observations are also summarized in
Table 1. Besides the review prepared by Dubbeldam(1994), other bibliographical references on the neuroanat-omy and functions of the main groups of nuclei are citedafter their first mention.
Somatosensory system
Sensory trigeminal system. Aromatase immunore-activity was found to be broadly though heterogeneouslydistributed throughout the entire rostrocaudal extent ofthe sensory trigeminal system (P 4.2 to A 2.1; Dubbeldamand Karten, 1978; Wild and Zeigler, 1996; Dubbeldam,1998). The most numerous and intensely stained ARO-irneurons in the caudal hindbrain were circumscribedwithin the limits of the caudal subnucleus (nTTDc; P 4.2to 2.8, Fig. 2A,C,E,F). This subnucleus is divided into fourto five laminae. ARO-ir structures occupied mainly lami-nae I and III. They were less dense in lamina II and absentin lamina IV. Caudally, the labeling extended withoutdiscontinuity in the dorsal horn of the spinal cord aspreviously described (Evrard et al., 2000). Rostrally, thelimit of the caudal subnucleus was clearly defined by theend of its laminar organization and, coincidentally, by thedisappearance of ARO-ir somata (P 2.8). Although ARO-irfibers remained present, they were rare all along the restof the descending trigeminal tract (TTD; P 4.2 to 0.8, Figs.2I, 3A); a network of scarce positive fibers heterogeneouslyfilled the entire interpolar (nTDDi; P 2.8 to 2.4) and oral(nTDDo; P 1.6 to 0.8) subnuclei. In the three caudal nuclei,no ARO-ir fibers were seen exiting the trigeminal systemand extending in adjacent loci such as the adjacent retic-ular formation, which is known to receive inputs from thecaudal and interpolar subnuclei (Dubbeldam, 1998). Theentering trigeminal nerve itself (NV; A 0.2 to 0.4) did notcontain any immunoreactivity. In the ascending part ofthe trigeminal system, the principal trigeminal nucleus(nPrV; A 0.4 to 1.2) contained only a small amount ofARO-ir fibers and was surrounded by a ring of denseimmunoreactive processes that overlapped with the tri-geminal ascending tract (TTA; A 0.2 to 0.4, Fig. 3G) and,to some extent with the superior vestibular nucleus (VeS;A 0.4). The mesencephalic trigeminal nuclei (nVM; Fig. 1,A 2.3 to 4.4; Hummel et al., 1988; Scott et al., 1994)expressed long transverse ARO-ir fibers with punctatestructures.
Dorsal column nuclei and external cuneate nucleus.
Another conspicuous group of ARO-ir soma occupied theventral position of the external cuneate nucleus (P 4.2-3.0;CE; Fig. 2A,B; Dubbeldam, 1984) and seemed to be therostral continuation a group of ARO-ir neurons previouslyobserved in the spinal cervical nucleus (Evrard et al.,2003). At the same rostrocaudal level, and mediodorsally,a few stained fibers were also observed in the cuneate-gracilis or dorsal column nuclei (GC; P 4.2 3.0; these twonuclei are not distinguishable in birds; Wild, 1989;Necker, 1991).
Viscerosensory system
With the exception of a limited area located at itscaudal end (the posterior dorsomedial subnucleus; mDp;P 2.6), the whole solitary tract nucleus was covered byARO-ir fibers associated with varicosities (nTS, cnTS; P4.2 to 1.6, Figs. 2A,D,J, 3B–D; Krol and Dubbeldam,1979; Katz and Karten, 1983; Gentle, 1984; Dubbeldam,1994). Two groups of ARO-ir soma were present in thesolitary tract nucleus. At the levels P 4.2 to 3.2 (the
196 H.C. EVRARD ET AL.
Figure 1
197ESTROGEN SYNTHESIS IN THE HINDBRAIN
Figure 1 (Continued)
Figure 1 (Continued)
same level as the caudal trigeminal subnucleus andexternal cuneate nucleus), the first group consisted ofsmall round-shaped neurons that send coarse stainedfibers transversally toward the mediodorsal part of thetrigeminal spinal nucleus (cnTS; Fig. 2A,D). More ros-trally (P 2.0, Fig. 3C,D), larger multipolar ARO-ir cellswere confined in a particular area of the solitary tract
nucleus that may correspond to the rostral part of thedorsomedial subnuclei (mDa) described by Katz andKarten (1983). These cells displayed only short immu-nopositive processes. Dorsal to the solitary tract nu-cleus, the area postrema (Apa; P 2.6) was also denselyinnervated by aromatase fibers forming a kind of sheatharound the area postrema.
Figure 1 (Continued)
200 H.C. EVRARD ET AL.
Fig. 1. Schematic drawings of transverse 40-�m-thick sectionsthrough the hindbrain in Japanese quail (from the spinocerebralintersection to the occulomotor nerve). Black or white large dots areused to illustrate the general distribution of aromatase-immunoreactive perikarya. The number of dots has been adjusted toprovide a rough qualitative estimate of the amount of immunoreactivecells per surface unit. The different gray tones correspond to differentdensities of ARO-ir fibers per surface unit with the darker colorscorresponding to the denser networks of immunoreactive fibers. Theapproximate correspondence between gray levels and amount ofaromatase-immunoreactive fibers per surface unit is illustrated at thebottom of the first panel of the Figure 1. Bundles of hatched lines in
RF are used for areas containing homogeneously distributed, lightlylabeled aromatase-immunoreactive fibers. Sections are presented in acaudal to rostral order. The sections are located as being anterior (A)or posterior (P) to a reference plane in the quail brain (0.0), whichcorresponds to the reference plane 0.0 used by Kuenzel and Masson(1988) in the chicken atlas. The rostrocaudal coordinates used herealso refer to the corresponding plates in the chicken atlas and, there-fore, do not correspond to actual millimeters in the smaller quailbrain. For abbreviations, see Table 1. Magnification has been adjustedin the successive pages to allow a detailed representation of all im-munoreactive structures (see scale bars at the bottom right of eachpage). Scale bar � 250 �m in P4.2–A1.6; 500 �m in A2.2–A4.0.
Fig. 2. Digital photomicrographs of 40-�m-thick transversal sec-tions at different rostrocaudal levels through the quail hindbrain,illustrating the distribution of aromatase-immunoreactive (ARO-ir)neuronal structures. A: Laminae I–III of the caudal nucleus of thedescending trigeminal tract, the extern cuneate nucleus (CE), and thecaudal solitary tract nucleus (cnTS). B–D: Higher magnifications of A.In C, the double arrowheads point to ARO-ir cell bodies embedded inlamina I ARO-ir fibers tangle. E: Caudal/interpolar nucleus of the
descending trigeminal tract (nTTDc/i). F–H: Higher magnifications ofE. I–L: Interpolar nucleus of the descending trigeminal tract (nTTDi;I), solitary tract nucleus (nTS; J; arrows point to ARO-ir punctatestructures), raphe pallidus (Rp; K), and obscurus (Ro; L). For otherabbreviations, see Table 1. Scale bar � 150 �m in L (applies to I–L);200 �m for A; 75 �m for B,H; 100 �m for C,D,G; 350 �m for E; 35 �mfor F.
Fig. 3. Digital photomicrographs of 40-�m-thick transversal sec-tions at different rostrocaudal levels through the quail hindbrain,illustrating the distribution of aromatase-immunoreactive (ARO-ir)neuronal structures. A: Lateroventral part of the oral nucleus of thedescending trigeminal tract. B,C: Solitary tract nucleus. D,E; Highermagnifications of C (mediodorsal [D, mDa] and medioventral [E, mVa]
tiers of the solitary tract nucleus). F–H: reticular formation (F; RF),ascending trigeminal tract (G; TTA), and central gray (or periaque-ductal gray; H; GCt). I: Caudal locus ceruleus (LoC) and dorsolateralraphe (Rdl). J: A higher magnification of I. For other abbreviations,see Table 1. Scale bar � 85 �m in J (applies to D,E,J); 100 �m forA,F,H; 350 �m for C,I; 200 �m for G.
203ESTROGEN SYNTHESIS IN THE HINDBRAIN
Fig. 4. Digital photomicrographs of 40-�m-thick transversal sec-tions through the quail hindbrain, illustrating the distribution ofaromatase-immunoreactive (ARO-ir) neuronal structures. A–C: Dif-ferent strata of the optic lobe. D–F: dorsal (Rd) and dorsolateral raphe(Rdl), area of the subcerulean nuclei (SCv), and linearis caudalis (LC).
G: Dorsal (SCd) and ventral (SCv) subceruleus nuclei. H,I: Dorsal andlateral part of the rostral hindbrain. J: Lateral parabrachial nucleus(PBl). For other abbreviations, see Table 1. Scale bar � 85 �m in J;100 �m for A–C, G–I; 350 �m for D; 500 �m for E; 200 �m for F.
TABLE 1. Abbreviations and Aromatase-Immunoreactivity Distribution in the Hindbrain1
Cochlear system. With the exception of ARO-ir fiberscovering the dorsal lateral lemniscal nucleus (LLd; A 2.2to 2.5) and surrounding the intercollicular nucleus (ICo; A2.3 to 4.4), no immunostaining was detected in the co-chlear system and its related nuclei (P 2.0 to A 4.4; Arendsand Zeigler, 1986; Akesson et al., 1987; Ball et al., 1989;Dubbeldam, 1994; Kubke and Carr, 2000). It has to bestressed that the ARO-ir fibers seen in lateral lemniscalnucleus and around the intercollicular nucleus are in factpart of a very large and dense network of ARO-ir fiberscovering almost entirely the dorsolateral part of the ros-tral hindbrain (from A 1.2 to 4.4; photomicrographs of thisnetwork are shown in Fig. 4H,I).
rior olive nucleus. Besides the presence of a few ARO-irfibers in the dorsal vestibular nucleus (VeD; P 2.6 to 2.4)and in the rostral to the superior vestibular nucleus (VeS;A 0.4), no immunoreactivity was found in the vestibularsystem (Wold, 1978a,b; Kuenzel and Masson, 1988; Dub-beldam, 1994).
Optic lobes and related nuclei
In contrast with their absence in the isthmic nuclei,dense networks of ARO-ir fibers with varicosities wereheterogeneously distributed in various strata of the optictectum (A 1.2 to 4.4, Fig. 4A–C; Reiner and Karten, 1982;Kuenzel and Masson, 1988; Gunturkun, 2000). In trans-verse sections, ARO-ir fibers preferentially displayed anorientation parallel to the plane of section in most of theoptic lobe (Fig. 4A,B) except in two oval areas located atthe dorsomedial and ventromedial ends of the most super-ficial layer of the tectum (see asterisks in Fig. 1, A1.2 toA2.2; Fig. 4C). ARO-ir fibers were also present in some
parts of the isthmo-optic area (IO; A 2.2 to 2.5; Gun-turkun, 2000). These fibers were embedded into the largeARO-ir fibers network of the aforementioned laterodorsalpart of the hindbrain.
Integrating nuclei
Premotor nuclei of the reticular formation. In thecharts of Figure 1, we highlighted most of the reticularformation with light gray hatches corresponding to thevery scarce but relatively homogeneous distribution ofARO-ir fibers with varicosities throughout the reticularformation (Fig. 3F).
Serotoninergic and catecholaminergic systems.
The 5HT-ir raphe subnuclei could be separated in a ven-trocaudal (P 3.0 to A 0.4) and a dorsorostral (A 0.4 to 2.5)group, in agreement with previous studies in quail (Cozziet al., 1991) and, to some extent, in mammals (Parent,1981). Both groups contain conspicuous populations of5HT-ir cells (not shown here in detail, see, however, Fig.4). These groups are similar to those described by Parentin a comparative study of the serotoninergic system inbirds and mammals (Parent, 1981). Based on this compar-ative study, we tentatively named in quail the differentgroups of 5HT-ir cells according to the mammalian no-menclature (Paxinos and Watson, 1986). Additional stud-ies, however, will be necessary to ascertain homologiesdiscussed by Parent.
Dense to scattered ARO-ir fibers were found all alongthe raphe subnuclei (see Fig. 1). In addition, five groups ofARO-ir somata were observed. The most caudal group waslocated between the two bundles of the mediolateral fas-cicle (FLM; P 2.8, Fig. 2K,L). The second one was made ofonly a few small immunoreactive cells that were locatedmedially on the floor of the hindbrain at level A 1.6. The
TABLE 1. (continued)
ARO-irfibers
ARO-irsomata
Ro raphe obscurus �� ��Rp raphe pallidus �� ��RPam nucleus reticularis paramedianus �RPgc nucleus paragigantocellularis �Ru nucleus ruber (red nucleus) �RxVM radix mesencephalica nervi trigeminiSAC stratum album centrale ��SCbd tractus spinocerebellaris dorsalisSCd nucleus subcoeruleus, pars dorsalis ��� ���SCv nucleus subcoeruleus, pars ventralis ��� ���SFGS stratum fibrosum et griseum superficiale ���SGC stratum griseum centrale ��SGP stratum griseum periventriculare ����SLu nucleus semilunarisSN substantia nigra ��SO stratum opticumSS nucleus supraspinalisTa nucleus tangentiallisTD nucleus tegmenti dorsalis ���� ����TIO tractus isthmo-opticus ���TTA tractus nervi trigemini ascendens ��/���TTD tractus nervi trigemini descendens ��/���TV nucleus tegmenti ventralis ��� ���VeD nucleus vestibularis descendens or Deiter’s nucleus �VeL nucleus vestibularis lateralis or Deiter’s nucleusVeM nucleus vestibularis medialis or nucleus triangularisVeS nucleus vestibularis superior �Vla nucleus tractus solitarius, ventralis lateralis, pars anteriorVT ventriculus tecti mesencephaliVTA area tegmenti ventralis ���
1List of abbreviations used and summary of the distribution of aromatase- immunoreactive (ARO-ir) fibers and perikarya in the hindbrain in male Japanese quail. The numberof crosses (1 to 3) provides a qualitative estimate of the number of immunoreactive structures (fibers or somata) per unit surface. Crosses in parentheses for FLM indicate thatthese cells are more likely to be in between bundles of the FLM rather than in the FLM itself.
206 H.C. EVRARD ET AL.
third group consisted also of a few ARO-ir cells located inthe raphe linearis caudalis (LC; A 2.2 to 2.3; Fig. 4E). Thefourth group (the most conspicuous group of ARO-irperikarya in the rostral hindbrain) consisted in a largewing-shaped nucleus (at levels A 1.2 to 2.5; Fig. 4D,E). Inthis group, large ARO-ir cells were found around and, tosome extent, in the mediolateral fascicle (FLM, A 1.6) andoverlapping with the ventral and dorsal tegmental nuclei(TV and TD). At levels A 1.2 to 2.5, a fifth group of smallerARO-ir perikarya was observed in the vicinity of the ven-tral and dorsal subceruleus nuclei (SCv and SCd; Fig.4F,G). Throughout their entire rostrocaudal extent, themorphology of the fourth and fifth groups of ARO-ir cellswas found to be very similar to the morphology of the5HT-ir cells observed on parallel sections (Fig. 5), suggest-ing that these two neurochemical markers could be colo-calized.
As shown previously for serotoninergic cells (see Pan-zica et al., 1991), ARO-ir cells overlapped with TH-ir cellgroups. They were found, indeed, in the ventral part of thelocus ceruleus (LoC, A 1.2) and central gray (GCt, A 1.6 to2.2) as well as in the median part of the dorsal and ventralsubceruleus nuclei (A 1.2 to 2.5). However, double labelingof the same sections for aromatase and TH revealed that,in fact, these ARO-ir perikarya stand at the border of theTH-ir cell groups of the locus and central gray rather thanbeing part of them (Evrard H.C. and Balthazart J., un-published data). In the subceruleus groups, ARO-ir cellswere actually intermingled with TH-ir cells but werenever colocalized in a same cell (Evrard H.C. and Baltha-zart J., unpublished data). Most TH-ir cells groups were inaddition densely innervated by ARO-ir fibers throughouttheir entire rostrocaudal extent. Both the substantia nigra
(SN; A 2.3 to 2.5) and the ventral area of Tsai (AVT; A 3.4)were included in a large network of ARO-ir fibers thatcovered most of the hindbrain at this rostrocaudal level.This ventral network of ARO-ir fibers, however, was lessdense than at more dorsal corresponding levels. The in-terpeduncular nucleus was devoid of staining (IP; A.3.6).
Parabrachial system. The parabrachial nuclei areinvolved in various functions related for example to noci-ception, cardiovascular function, and homeostasis (Jac-coby et al., 1999; Millan, 1999; Saleh et al., 2002). In theNissl-stained sections, two parabrachial cell groups (PBvand PBl) were clearly identified ventral and lateral to thebrachium conjunctivum (BC; A 1.2 to 1.6; for a compara-tive study on the parabrachial system, see Petrovicky andKolesarova, 1989). The entire parabrachial system wascovered with ARO-ir fibers (PBv and PBl; A 1.2 to 1.6). Inaddition, the lateral parabrachial nucleus (PBl) containedARO-ir somata (Fig. 4J).
Other integrating nuclei. The nucleus interpeduncu-laris (IP; A.3.4) was almost entirely devoid of ARO-irstructures. However, numerous ARO-ir fibers without ap-parent orientation (like TH-ir fibers) were found abovethis nucleus and in between the two branches of nerve III.This area has not been named or described to our knowl-edge. The nucleus ruber (red nucleus, Ru, A 3.4/4.4), likethe reticular formation, contained few ARO-ir fibers withvaricosities.
Other areas
Besides sensory and integrating nuclei, numerous areasrelated to specific functions were found completely devoidof staining (e.g., motor nuclei) or covered by scatterednetworks of fibers (e.g., median and spinal lemniscal path-ways). These areas are listed in the Table 1.
DISCUSSION
This study demonstrates the presence of abundantARO-ir neuronal structures in numerous sensory and in-tegrating nuclei of the hindbrain in adult male quail.Scattered fibers were also found in the premotor nuclei ofthe reticular formation. Previous studies had identifiedARO-ir neurons in the spinal dorsal horn and forebrain inquail (Foidart et al., 1995; Evrard et al., 2000). Thepresent results thus complete our knowledge of the distri-bution of aromatase in this species. It has to be noted that,like in the spinal cord, no ARO-ir structures were found inthe motor area in the hindbrain.
Comparison with other species
There is, to this date, no comprehensive study of thedistribution of aromatase in the spinal cord or hindbrainof any species other than Japanese quail. The fragmen-tary data already available, however, in most cases, areconsistent with the present findings and hint at a goodphyletic conservation of the expression of aromatase insensory and integrating nuclei. In other avian studies,aromatase mRNA has been detected in the solitary tractnucleus (Metzdorf et al., 1999), in the spinal dorsal horn(Fusani et al., 2002) and in a wing-shaped cells group thathas also been shown to express 5HT (Cozzi et al., 1991;Shen et al., 1995; in the present study: dorsolateral anddorsal raphe subnuclei). In quail and in pied flycatchers(Ficedula hypoleuca), AA has been detected in both theoptic lobes and the rest of the hindbrain, although no
Fig. 5. A,B: Digital photomicrographs of adjacent 40-�m-thicktransversal sections immunolabeled for aromatase (A) or serotonin(B), illustrating the coexistence of these antigens in sections in thequail hindbrain (level A2.3). Arrows point to some of the areas whereboth aromatase- and serotonin-immunoreactive structures were ob-served. For abbreviations, see Table 1. Scale bar � 250 �m.
207ESTROGEN SYNTHESIS IN THE HINDBRAIN
microdissection was carried out to locate more preciselythe enzymatically active structures within these largebrain areas (Balthazart et al., 1998; Foidart et al., 1998).In rats, aromatase immunoreactivity has been detected inall sensory nuclei of neonates (Horvath and Wikler, 1999)and evidence from estrogen microdialysis also suggeststhe presence of aromatase in the adult parabrachial nu-cleus (Saleh et al., 2002; see below). In addition, previouswork in our laboratory has demonstrated in rats the pres-ence of ARO-ir structures in spinal regions and hindbrainsensory and integrating nuclei homologues to thearomatase-expressing nuclei identified in Japanese quail,including the parabrachial nuclei (Evrard et al., 2001;Evrard H.C., Harada N., Balthazart J., unpublished data).Finally, although the expression of AA in hindbrain neu-rons appears similar in amniotes and, to some extent, invarious teleost species (Timmers et al., 1987; Callard etal., 1988; Mayer et al., 1991; Gelinas and Callard, 1993;Schlinger et al., 1999), other fish species (i.e., plainfinmidshipman fish and rainbow trout) differ significantly byexpressing aromatase in glial cells rather than in neuronsand, at least in the midshipman fish, by expressing suchglial aromatase in motor rather than sensory nuclei (For-lano, 2001; Menuet et al., 2003; Forlano, personal commu-nication).
Functional implications
The broad distribution of aromatase in the sensoryhindbrain indicates that the presence of aromatase in thedorsal horns of the spinal cord of adult quail is not aunique feature of this structure (Evrard et al., 2000) butrepresents a more common trait of sensory nuclei. Aro-matase immunoreactivity has been observed also in theolfactory bulb of canaries, ring doves (Metzdorf et al.,1999), quail (Evrard H.C. and Balthazart J., unpublisheddata), and fishes (Chiang, 2001). These observations allowthe extension of the notion of “spinal aromatase” to thebroader and new concept of “sensory aromatase.” As ex-plained in the introductory section, the regulation of sen-sory function by estrogens produced in the sensory nucleicould play an important role during reproduction.
Besides sensory nuclei, the present study also demon-strates, for the first time in vertebrates, the presence ofARO-ir structures in the parabrachial nucleus, periaque-ductal gray, raphe, substantia nigra, and cerulean nuclei.Aromatase located in these integrating nuclei could alsoplay a role during reproduction. For instance, the controlof the motor aspects of male sexual behavior that dependon the action of estrogens among others in the periaque-ductal gray could rely, at least in part, on locally producedestrogens (Absil et al., 2001; Murphy and Hoffman, 2001).
Furthermore, it is interesting to note that most hind-brain nuclei expressing aromatase are also involved in theregulation of nociception and, more broadly, of homeosta-sis (Millan, 1999; Craig, 2002). Indeed, the caudal trigem-inal (as well as the spinal lamina I), solitary tract, para-brachial, cerulean, and periaqueductal nuclei that containaromatase have been shown to represent an importantcluster of nuclei in the pain pathway and, more broadly, inthe homeostasis system. It would be interesting to inves-tigate further whether the juxtaposed aromatase expres-sion and pain/homeostatic system are developmentallyand functionally interdependent. From a clinical perspec-tive, the study of hindbrain aromatase could bring rele-vant information regarding estrogen-dependent discom-
fort such as migraine (involving caudal trigeminalsubnucleus; Silberstein, 2000; Pardutz et al., 2002), dys-pareunia (involving possibly the solitary tract area receiv-ing inputs from female genitalia through the vagal nerve;Meana and Binik, 1994; Collins et al., 1999; Komisarukand Whipple, 2000), temporal mandibular disorder (Fill-ingim, 2000), excessive nausea (involving a.o. the areapostrema; Jarnfelt-Samisioe, 1987), Parkinson’s disease(involving the dopaminergic neurons of the substantianigra; Leranth et al., 2000). Moreover, in humans, thehindbrain homeostatic system is assumed to be an impor-tant relay in the neural circuitry mediating the processesof self-recognition and emotion because it integrates infor-mation concerning the state of the body (Craig, 2002).
Subcellular distribution of aromatase
The identification of the origin of the numerous isolatedhindbrain ARO-ir fibers will require piecemeal tract trac-ings combined with aromatase immunocytochemistry. Byusing this method, Absil et al. (2001) demonstrated thatthe quail GCt (avian homologue of the periaqueductalgray) receives numerous ARO-ir afferents from the lateralpreoptic area so that part of the ARO-ir fibers presentlyobserved in the GCt may arise from this area (Absil et al.,2001). The presence of aromatase in fibers and punctatestructures (presumptive synaptic boutons) suggests thatestrogens could be produced and released at sites fairlydistant from the ARO-ir perikarya. This idea is supportedby the detection of AA in the optic lobes (100 fmol/mgprotein/hr; Balthazart et al., 1998) where immunocyto-chemistry identified ARO-ir fibers and punctate struc-tures but no immunoreactive perikarya. The synaptic na-ture of the ARO-ir punctate structures is supported by theprevious detection of AA in synaptosomal fractions andthe detection by electron microscopy of aromatase immu-noreactivity in presynaptic boutons in addition to theperikarya (Steimer, 1988; Schlinger and Callard, 1989;Naftolin et al., 1996). In the perikarya, the exact subcel-lular distribution of aromatase has still to be confirmed,but it seems that the enzyme mainly resides in the mem-brane of the endoplasmic reticulum (e.g., Naftolin et al.,1996). In the boutons, the immunoreactive material isessentially associated to the surface of clear vesicles butdoes not seem to bear any relationship with the neuronalmembrane (Naftolin et al., 1996). The functional charac-teristics of the cell organite(s) in which estrogens areproduced and presumably from which they are releasedstill has to be investigated. For instance, whether theARO-ir vesicles in boutons derive from perikaryal reticu-lum, how they arrive in the terminals, and whether pro-cesses other than simple diffusion are associated withestrogens release (e.g., by means of synaptic vesicle-likeexocytose) remain to be investigated. In both theperikarya and the boutons, estrogens could be formedfrom circulating androgens that enter the central nervoussystem (CNS) and then the cells through diffusion or,alternatively, from androgens directly produced withinthe CNS (e.g., Mensah-Nyagan et al., 1996; Steckelbroecket al., 1999; Matsunaga et al., 2002). Once formed, estro-gens produced in the perikarya could interact with estro-gen receptors and DNA in the nucleus of the cell wherethey are produced or diffuse through cell membranes andbind to nuclear receptors in other cells as suggested pre-viously for spinally produced estrogens (Evrard and Bal-thazart, 2003). Estrogens produced in boutons could sim-
208 H.C. EVRARD ET AL.
ilarly diffuse locally and reach adjacent cells expressingnuclear estrogen receptors and induce thereby genomicchanges in their environment. However, because, in addi-tion to their action at the genomic level, estrogens havebeen shown also to change rapidly the excitability of neu-rons through the interaction with specific membrane sites(Kelly and Ronnekleiv, 2002), estrogens produced in theperikarya and in the presynaptic boutons could addition-ally activate fast nongenomic mechanisms in an autocrineand/or paracrine manner. In particular, the presence ofaromatase in the proximity of the synaptic cleft is as-sumed to lead to the establishment of high synaptic con-centrations of estrogens that may be necessary to exertfast nongenomic action at the postsynaptic membranelevel through mechanisms that are similar to those usedby neurosteroids (e.g., Naftolin et al., 1996; Baulieu et al.,2000; Ball and Balthazart, 2002). Finally, it must also bementioned that, besides estrogen synthesis, aromatasehas been shown to catalyze other reactions such as the 1�-and 2�-hydroxylations of androgens, the 2-hydroxylationof estrogens, and the methylation of various substrates,including dopamine or cocaine (Osawa et al., 1993, 1994).This finding could provide alternative nongenomic modesof action for the products of AA.
Control of hindbrain aromatase activity
AA is regulated either through relatively slow controlsof the transcription of the aromatase gene Cyp19 or bymore rapid phosphorylations/dephosphorylations of thearomatase protein (Balthazart and Ball, 1998b; Balthaz-art et al., 2003). Cyp19 consists of 10 exons, including atissue-specific first untranslated exon that contains a va-riety of promoters (Simpson et al., 1994). In the preopticarea, the transcription of Cyp19 and, as a consequence,the levels of AA are dramatically increased by testoster-one (Harada et al., 1992; Roselli and Resko, 1993). Incontrast, in the spinal cord, this steroid does not alter AAnor the number of ARO-ir cells (Evrard et al., 2000).Similarly, AA in the hindbrain could be independent of thelevels of circulating steroids. Although this possibility re-mains to be rigorously assessed, it is already supported byseveral indirect data. For example, the fluctuation ofplasma estrogen concentration does not alter the estrogenconcentration in the parabrachial nucleus of female ratsbut a peripheral stimulation of the vagal nerve elicits arapid local release of estrogens in the absence of change inadjacent structures (Saleh et al., 2002). In addition, in thepied flycatcher (Ficedula hypoleuca), while AA decreasesin the preoptic area after the seasonal drop of testosteroneplasma concentration, this enzymatic activity does notchange in other brain areas, including the optic lobes andthe remaining hindbrain (Foidart et al., 1998). AA insen-sitive to steroids has been also detected in the rodentlimbic system (Roselli and Resko, 1993) and in the oscinetelencephalon (Schlinger and Arnold, 1991; Saldanha etal., 2000). Taken together, these results suggest that thegenomic control of AA by androgens and estrogens may bespecific to the preoptic area and parts of the limbic systemand that the control of neural AA is a rather heteroge-neous process. Further investigations are now necessaryto identify the major mechanisms controlling AA in areaswhere transcription of the enzyme is not controlled bysteroid. In the sensory nuclei, as we previously suggestedfor the spinal cord (Evrard et al., 2003), AA could bemodulated by substances released from local afferent in-
puts, including primary afferent and supraspinal fibers(see the example of substance P release in the spinal cordin Evrard et al., 2003). It would be of particular interest toassess whether specific sensory stimuli induce or reduceAA in sensory nuclei through genomic control but alsothrough faster control mechanisms involving aromatasephosphorylation (see Discussion in Evrard et al., 2003).For example, while normal sensory cues might inhibit ornot affect aromatase, reproductively relevant sensory cuesmight activate aromatase and, thereby, influence the pro-cessing of subsequent sensory cues in a way that favorsreproduction.
CONCLUSION
In conclusion, the present study demonstrates the pres-ence of the estrogen-synthesizing enzyme in broad areasof the quail hindbrain, in particular in the sensory andintegrating areas. These data suggest that estrogens arelocally produced in broad areas where they could exertsignificant effects on a variety of brain functions, includ-ing the regulation of various sensory modalities and theintegration of different types of inputs and outputs. Thesefindings also allow extending the concept of spinal sensoryaromatase to the much broader idea of a sensory aro-matase that would be implicated in the modulation ofmost if not all sensory inputs.
ACKNOWLEDGMENTS
We thank Profs. J.L. Dubbeldam (University of Leiden,The Netherlands) and G.C. Panzica (University of Turin,Italy) for their useful comments on an earlier version ofthe diagrams of the present article.
LITERATURE CITED
Absil P, Riters LV, Balthazart J. 2001. Preoptic aromatase cells project tothe mesencephalic central gray in the male Japanese quail (Coturnixjaponica). Horm Behav 40:369–383.
Akesson TR, Lanerolle NC, Cheng MF. 1987. Ascending vocalization path-ways in the female ring dove: projections of the nucleus intercollicu-laris. Exp Neurol 95:34–43.
Alberti-Fidanza A, Fruttini D, Servili M. 1998. Gustatory and food habitchanges during the menstrual cycle. Int J Vitam Nutr Res 68:159–153.
Amandusson A, Hermanson O, Blomqvist A. 1995. Estrogen receptor-likeimmunoreactivity in the medullary and spinal dorsal horn of the fe-male rat. Neurosci Lett 196:25–28.
Amandusson A, Hallbeck M, Hallbeck A-L, Hermanson O, Blomqvist A.1999. Estrogen-induced alterations of spinal cord enkephalin geneexpression. Pain 83:243–248.
Arends JJ, Zeigler HP. 1986. Anatomical identification of an auditorypathway from a nucleus of the lateral lemniscus system to the frontaltelencephalon (nucleus basalis) in the pigeon. Brain Res 398:375–381.
Bailhache T, Balthazart J. 1993. The catecholaminergic system of the quailbrain: immunocytochemical studies of dopamine �-hydroxylase andtyrosine hydroxylase. J Comp Neurol 329:230–256.
Ball GF, Balthazart J. 2002. Neuroendocrine mechanisms regulating re-productive cycles and reproductive behavior in birds. In: Pfaff DW,Arnold AP, Etgen AM, Fahrbach SE, Rubin RT, editors. Brain, hor-mones and behavior. San Diego: Academic Press. p 545–586.
Ball GF, Foidart A, Balthazart J. 1989. A dorsomedial subdivision withinthe nucleus intercollicularis identified in the Japanese quail (Coturnixcoturnix japonica) by means of alpha 2-adrenergic receptor autoradiog-raphy and estrogen receptor immunohistochemistry. Cell Tissue Res257:123–128.
Balthazart J, Ball GF. 1998a. The Japanese quail as a model for theinvestigation of steroid-catecholamine interactions mediating appeti-
209ESTROGEN SYNTHESIS IN THE HINDBRAIN
tive and consummatory aspects of male sexual behavior. Annu Rev SexRes 9:96–176.
Balthazart J, Ball GF. 1998b. New insights into the regulation and func-tion of brain estrogen synthase (aromatase). Trends Neurosci 21:243–249.
Balthazart J, Foidart A, Baillien M, Harada N, Ball GF. 1998. Anatomicalrelationships between aromatase and tyrosine hydroxylase in the quailbrain: double-label immunocytochemical studies. J Comp Neurol 391:214–226.
Balthazart J, Baillien M, Charlier TD, Ball GF. 2003. Calcium-dependentphosphorylation processes control brain aromatase in quail. Eur J Neu-rosci 17:1591–1606.
Baulieu E-E, Schumacher M, Robel P. 2000. Neurosteroids: a new regula-tory function in the nervous system. New York: Blackwell.
Bereiter DA, Barker DJ. 1980. Hormone-induced enlargement of receptivefields in trigeminal mechanoreceptive neurons. I. Time course, hor-mone, sex and modality specificity. Brain Res 184:395–410.
Berk ML. 1991. Distribution and hypothalamic projection of tyrosine-hydroxylase containing neurons of the nucleus of the solitary tract inthe pigeon. J Comp Neurol 312:391–403.
Berk ML, Smith SE, Mullins LA. 1993. Distribution, parabrachial regionprojection, and coexistence of neuropeptide and catecholamine cells ofthe nucleus of the solitary tract in the pigeon. J Comp Neurol 327:416–441.
Blaustein JD, Erskine MS. 2002. Feminine sexual behavior: cellular inte-gration of hormonal and afferent information in the rodent forebrain.In: Pfaff DW, Arnold AP, Etgen AM, Fahrbach SE, Rubin RT, editors.Brain, hormones and behavior. San Diego: Academic Press. p 326–456.
Bradshaw H, Miller J, Ling Q, Malsnee K, Ruda MA. 2000. Sex differencesand phases of the estrous cycle alter the response of spinal cord dynor-phin neurons to peripheral inflammation and hyperalgesia. Pain 85:93–99.
Burris AS, Gracely RH, Carter CS, Sherins RJ, Davidson JM. 1991. Tes-tosterone therapy is associated with reduced tactile sensitivity in hu-man males. Horm Behav 25:195–205.
Butler AB, Hodos W. 1996. Comparative vertebrate neuroanatomy: evolu-tion and adaptation. New York: Wiley-Liss, Inc.
Callard GV, Specker JL, Knapp J, Nishioka RS, Bern HA. 1988. Aromataseis concentrated in the proximal pars distalis of tilapia pituitary. GenComp Endcrinol 71:70–79.
Caruso S, Cianci A, Grasso D, Agnello C, Galvani F, Maiolino L, Serra A.2000. Auditory brainstem response in postmenopausal women treatedwith hormone replacement therapy: a pilot study. Menopause 7:178–183.
Chabli A, Schaeffer C, Samama B, Aron C. 1985. Hormonal control of theperception of the olfactory signals which facilitate lordosis behavior inthe male rat. Physiol Behav 35:729–734.
Chiang EF. 2001. Two Cyp19 (P450 Aromatase) Genes on DuplicatedZebrafish Chromosomes Are Expressed in Ovary or Brain. Mol BiolEvol 18:542–550.
Coleman JR, Campbell D, Cooper WA, Welsh MG, Myer J. 1994. Auditorybrainstem responses after ovariectomy and estrogen replacement inrat. Hear Res 80:209–215.
Collins JJ, Lin CE, Berthoud HR, Papka RE. 1999. Vagal afferent from theuterus and cervix provide direct connections to the brainstem. CellTissue Res 295:43–54.
Cox JR. 1980. Hormonal influence on auditory function. Hear Res 1:219–222.
Cozzi B, Viglietti-Panzica C, Aste N, Panzica GC. 1991. The serotoninergicsystem in the brain of the Japanese quail. Cell Tissue Res 263:271–284.
Craig AD. 2002. How do you feel? Interoception: the sense of the physio-logical condition of the body. Nat Rev Neurosci 3:655–666.
Deems DA, Doty RL, Settle RG, Moore-Gillon V, Shaman P, Mester AF,Kimmelman CP, Brightman VJ, Snow JBJ. 1991. Smell and tastedisorders, a study of 750 patients from the University of PennsylvaniaSmell and Taste Center. Arch Otolaryngol Head Neck Surg 117:519–528.
Dubbeldam JL. 1984. Afferent connections of nervus facialis and nervusglossopharyngeus in the pigeon (Columba livia) and their role in feed-ing behavior. Brain Behav Evol 24:47–57.
Dubbeldam JL. 1994. Birds. In: Nieuwenhuys R, ten Donkelaar HJ, Ni-cholson C, editors. The central nervous system of vertebrates. NewYork: Springer. p 1525–1636.
Dubbeldam JL. 1998. The sensory trigeminal system in birds: input, orga-
nization and effects of peripheral damage. A review. Arch PhysiolBiochem 106:338–345.
Dubbeldam JL, Karten HJ. 1978. The trigeminal system in the pigeon(Columba livia). I. Projections of the gasserian ganglion. J Comp Neu-rol 180:661–678.
Duval P, Lenoir V, Moussaoui S, Garret C, Kerdelhue B. 1996. SubstanceP and neurokinin A variations throughout the rat estrous cycle; com-parison with ovariectomized and male rats: II. Trigeminal nucleus andcervical spinal cord. J Neurosci Res 45:610–616.
Elkind-Hirsch KE, Wallace E, Stach BA, Jerger JF. 1992. Cyclic steroidreplacement alters auditory brainstem responses in young women withpremature ovarian failure. Hear Res 64:93–98.
Evrard HC, Balthazart J. 2000. Localization of estrogen-synthase andestrogen receptor immunoreactive cells in the descending nucleus ofthe trigeminal nerve in Japanese quail. Eur J Neurosci 12(Suppl 11):147.
Evrard HC, Balthazart J. 2002. Localization of estrogen receptors in thesensory and motor areas of the spinal cord in Japanese quail (Coturnixjaponica). J Neuroendocrinol 14:894–903.
Evrard HC, Balthazart J. 2003b. Long term and short term aromataseinhibitions result in slow and rapid alteraion of pain. 2nd InternationalConference on Steroids and Nervous System. Torino. p 122.
Evrard HC, Balthazart J. 2004. Aromatization of androgens into estrogensreduces response latency to an aversive thermal stimulus in malequail. Horm Behav (in press).
Evrard H, Baillien M, Foidart A, Absil P, Harada N, Balthazart J. 2000.Localization and controls of aromatase in the quail spinal cord. J CompNeurol 423:552–564.
Evrard HC, Harada N, Balthazart J. 2001. Localization of estrogen-synthase (aromatase) in the rat spinal cord. 31st Annual Meeting of theSociety for Neuroscience. San Diego. p 897.
Evrard HC, Harada N, Balthazart J. 2003. Specific innervation of aro-matase cells by substance P containing fibers in the dorsal horn of thespinal cord in Japanese quail. J Comp Neurol 465:309–318.
Fillingim RB. 2000. Sex, gender, and pain. Seattle: IASP Press.Foidart A, Reid J, Absil P, Yoshimura N, Harada N, Balthazart J. 1995.
Critical re-examination of the distribution of aromatase-immunoreactive cells in the quail forebrain using antibodies raisedagainst human placental aromatase and against the recombinantquail, mouse or human enzyme. J Chem Neuroanat 8:267–282.
Foidart A, Silverin B, Baillien M, Harada N, Balthazart J. 1998. Neuro-anatomical distribution and variations across the reproductive cycle ofaromatase activity and aromatase-immunoreactive cells in the piedflycatcher (Ficedula hypoleuca). Horm Behav 33:180–196.
Forlano PM. 2001. Anatomical distribution and cellular basis for highlevels of aromatase activity in the brain of teleost fish: aromataseenzyme and mRNA expression identify glial source. J Neurosci 21:8943–8955.
Fusani L, Schultz JD, Canoine V, Donaldson Z, Reinman DR, SchlingerBA. 2002. Androgen control of a complex avian courtship behavior. In:Orlando: Society for Neuroscience. Program No. 781.784.
Gelinas D, Callard GV. 1993. Immunocytochemical and biochemical evi-dence for aromatase in neurons of the retina, optic tectum and retino-tectal pathways in goldfish. J Neuroendocrinol 5:635–641.
Gentle MJ. 1984. The chorda tympani and taste in chicken. J Comp PhysiolA 130:455–463.
Gintzler AR, Liu N-J. 2001. The maternal spinal cord: biochemical andphysiological correlates of steroid-activated antinociceptive processes.Prog Brain Res 133:83–97.
Guillot PV, Chapouthier G. 1996. Olfaction, GABAergic neurotransmissionin the olfactory bulb, and intermale aggression in mice: modulation bysteroids. Behav Genet 26:497–504.
Gunturkun O. 2000. Vision. In: Whittow GC, editor. Sturkie’s avian phys-iology. Fifth edition. San Diego: Academic Press. p 1–120.
Hara TJ. 1967. Electrophysiological studies of the olfactory system of thegoldfish, Carassius auratus L. 3. Effects of sex hormones on olfactoryactivity. Comp Biochem Physiol 22:109–115.
Harada N, Yamada K, Foidart A, Balthazart J. 1992. Regulation of aro-matase cytochrome P-450 (estrogen synthetase) transcripts in the quailbrain by testosterone. Brain Res Mol Brain Res 15:19–26.
Hartwig H-G. 1993. The central nervous system of birds: a study of func-tional morphology. In: Avian biology. London: Academic Press. p 1–118.
Hinde RA, Steel EA. 1964. Effect of exogenous hormones on the tactilesensitivity of the canary brood patch. J Endocrinol 30:355–359.
210 H.C. EVRARD ET AL.
Horvath TI, Wikler KC. 1999. Aromatase in developing sensory systems ofthe rat brain. J Neuroendocrinol 11:77–84.
Hull EM, Meisel RL, Sachs BD. 2002. Male sexual behavior. In: Pfaff DW,Arnold AP, Etgen AM, Fahrbach SE, Rubin RT, editors. Brain, hor-mones and behavior. San Diego: Academic Press. p 3–137.
Hummel G, Pohlschmidt J, Goller H. 1988. Cytoarchitecture and ultra-structure of the mesencephalon trigeminal nucleus of the domesticchicken. J Hirnforsch 29:695–700.
Hummel T, Gollish R, Wildt K, Kobal G. 1991. Changes in olfactoryperception during the menstrual cycle. Experientia 47:712–715.
Hutchison RE, Hinde RA, Steel E. 1967. The effects of oestrogen, proges-terone and prolactin on brood patch formation in ovariectomized ca-naries. J Endocrinol 39:379–385.
Jaccoby S, Koike TI, Cornett LE. 1999. c-fos expression in the forebrainand brainstem of White Leghorn hens following osmotic and cardiovas-cular challenges. Cell Tissue Res 297:229–239.
Jarnfelt-Samisioe A. 1987. Nausea and vomiting in pregnancy: a review.Obstet Gynecol Surv 42:422–427.
Ji Y, Murphy AZ, Traub RJ. 2003. Estrogen modulates the visceromotorreflex and responses of spinal dorsal horn neurons to colorectal stim-ulation in the rat. J Neurosci 23:3908–3915.
Katz DM, Karten HJ. 1983. Visceral representation within the nucleus ofthe tractus solitarius in the pigeon, Columba livia. J Comp Neurol218:42–73.
Kelly MJ, Ronnekleiv OK. 2002. Rapid membrane effects of estrogen in thecentral nervous system. In: Pfaff DW, Arnold AP, Etgen AM, FahrbachSE, Rubin RT, editors. Brain, hormones and behavior. San Diego:Academic Press. p 361–380.
Keverne EB, Levy F, Poindron P, Lindsay DR. 1983. Vaginal stimulation:an important determinant of maternal bonding in sheep. Science 219:81–83.
Klimek A. 1985. Use of testosterone in the treatment of cluster headache.Eur Neurol 24:53–56.
Komisaruk BR, Whipple B. 2000. How does vaginal stimulation producepleasure, pain, and analgesia? In: Fillingim RB, editor. Sex, gender,and pain. Seattle: IASP Press. p 109–134.
Komisaruk BR, Adler NT, Hutchison J. 1972. Genital sensory field: en-largement by estrogens treatment in female rats. Science 178:1295–1298.
Konishi M. 1973. Genital sensory field. Science 181:365–366.Kow LM, Pfaff DW. 1973. Effects of estrogen treatment on the size of
receptive field and response threshold of pudendal nerve in the femalerat. Neuroendocrinology 13:299–313.
Krol CPM, Dubbeldam JL. 1979. On the innervation of taste buds by the N.facialis in the mallard, Anas platyrhynchos L. Neth J Zool 29:467–477.
Kubke MF, Carr CE. 2000. Development of the auditory brainstem inbirds: comparison between barn owls and chickens. Hear Res 147:1–20.
Kuenzel WJ, Masson M. 1988. A stereotaxic atlas of the brain of the chick(Gallus domesticus). Baltimore: The Johns Hopkins University Press.
Leranth C, Roth RH, Elswoth JD, Naftolin F, Horvath TL, Redmond DE.2000. Estrogen is essential in maintaining nigrostriatal dopamine neu-rons in primates: implication for Parkinson’s disease and memory.J Neurosci 20:8604–8609.
Li Z, Hay M. 2000. 17-beta-estradiol modulation of area postrema potas-sium currents. J Neurophysiol 84:1385–1391.
Martinez-Vargas MC, Stumpf WE, Sar M. 1976. Anatomical distribution ofestrogens target cells in the avian CNS: a comparison with the mam-malian CNS. J Comp Neurol 167:83–104.
Mascarenhas JF, Borker AS, Venkatesh P. 1992. Differential role of ovar-ian hormones for taste preferences in rats. Indian J Physiol Pharmacol36:244–246.
Matsunaga M, Ukena K, Tsutsui K. 2002. Androgen biosynthesis in thequail brain. Brain Res 948:180–185.
Mayer I, Borg B, Berglund I, Lambert JG. 1991. Effect of castration andandrogen treatment on aromatase in the brain of mature male Atlanticsalmon (Salmo salar L.) parr. Gen Comp Endcrinol 82:86–92.
Meana M, Binik YM, 1994. Painful coitus: a review of female dyspareunia.J Nerv Ment Dis 82:264–272.
Mensah-Nyagan AM, Feuilloley M, Do-Rego JL, Marcual A, Lange C,Tonon MC, Pelletier G, Vaudry H. 1996. Localization of 17beta-hydroxysteroid dehydrogenase and characterization of testosterone inthe brain of the male frog. Proc Natl Acad Sci U S A 93:1423–1428.
Menuet A, Anglade I, Le Guevel R, Pellegrini E, Pakdel F, Kah O. 2003.Distribution of aromatase mRNA and protein in the brain and pituitary
of female rainbow trout: comparison with estrogen receptor alpha.J Comp Neurol 462:180–193.
Metzdorf R, Gahr M, Fusani L. 1999. Distribution of aromatase, estrogenreceptor, and androgen receptor mRNA in the forebrain of songbirdsand nonsongbirds. J Comp Neurol 407:115–129.
Millan MJ. 1999. The induction of pain: an integrative review. Prog Neu-robiol 57:1–164.
Minson CT, Hallwill JR, Young TM, Joyner MJ. 2000. Influence of themenstrual cycle on sympathetic activity, baroreflex sensitivity, andvascular transduction in young women. Circulation 101:862–862.
Mohamed MK, El-Mass MK, Rahman KK. 1999. Estrogen enhancement ofbaroreflex sensitivity is centrally mediated. Am J Physiol 276:R1030–R1037.
Murphy AZ, Hoffman GE. 2001. Distribution of gonadal steroid receptor-containing neurons in the preoptic periaqueductal gray-brainstempathway: a potential circuit for the initiation of male sexual behavior.J Comp Neurol 438:191–212.
Murphy MR, Schneider GE. 1970. Olfactory bulb removal eliminates mat-ing behavior in the male golden hamster. Science 167:302–304.
Naftolin F, Horvath TL, Jakab RL, Leranth C, Harada N, Balthazart J.1996. Aromatase immunoreactivity in axon terminals of the vertebratebrain. An immunocytochemical study on quail, rat, monkey and humantissues. Neuroendocrinology 63:149–155.
Necker R. 1991. Cells of origin of avian postsynaptic dorsal column path-ways. Neurosci Lett 126:91–93.
Osawa Y, Higashiyama T, Shimizu Y, Yarborough C. 1993. Multiple func-tions of aromatase and the active site structure; aromatase is theplacental estrogen 2-hydroxylase. J Steroid Biochem Mol Biol 44:469–480.
Osawa Y, Higashiyama T, Yarborough C. 1994. Diverse functions of aro-matase cytochrome P450: catecholestrogen synthesis, cocainN-demethylation, and other selective drug metabolism. In: LechnerMC, editor. Cytochrome P450. 8th International Conference. Paris:John Libbey Eurotext. p 893–896.
Pajot J, Ressot C, Ngom I, Woda A. 2003. Gonadectomy induces site specificdifferences in nociception between male and female rats. Pain 104:367–373.
Panzica GC, Guglielmone R, Cozzi B, Aste N, Viglietti-Panzica C. 1991.Distribution of catecholamines and serotonin in the brain of the Japa-nese quail. A histochemical and immunocytochemical study. Gen CompEndocrinol 28:298–290.
Pardutz A, Multon S, Malgrange B, Parducz A, Vecsei L, Schoenen J. 2002.Effect of systemic nitroglycerin on CGRP and 5-HT afferents to ratcaudal spinal trigeminal nucleus and its modulation by estrogen. EurJ Neurosci 15:1803–1809.
Parent A. 1981. Comparative anatomy of the serotoninergic systems.J Physiol 77:147–156.
Paxinos G, Watson C. 1986. The rat brain in stereotaxic coordinates. SanDiego: Academic Press.
Petrovicky P, Kolesarova D. 1989. Parabrachial nuclear complex. A com-parative study of its cytoarchitecture in birds and some mammalsincluding man. J Hirnforsch 30:539–550.
Reiner A, Karten HJ. 1982. Laminar distribution of the cells of origin of thedescending tectofugal pathways in the pigeon (Columba livia). J CompNeurol 204:165–187.
Reiner A, Karle EJ, Anderson KD, Medina L. 1994. Catecholaminergicperikarya and fibers in the avian nervous system. In: Smeets WJAJ,Reiner A, editors. Phylogeny and development of catecholamine sys-tems in the CNS of vertebrates. Cambridge UK: Cambridge UniversityPress. p 135–181.
Romanzi LJ, Groutz A, Feroz F, Blaivas JG. 2001. Evaluation of femalegenitalia sensitivity to pressure/touch: a preliminary prospective studyusing Semmes-Weinstein monofilaments. Urology 57:1145–1150.
Roselli CE, Resko JA. 1993. Aromatase activity in the rat brain: hormonalregulation and sex differences. J Steroid Biochem Mol Biol 44:499–508.
Saldanha CJ, Tuerk MJ, Kim YH, Fernandes AO, Arnold AP, SchlingerBA. 2000. Distribution and regulation of telencephalic aromatase ex-pression in the zebra finch revealed with a specific antibody. J CompNeurol 423:619–630.
Saleh TM, Saleh MC, Deacon CL, Chisholm A. 2002. 17beta-Estradiolrelease in the parabrachial nucleus of the rat evoked by visceral affer-ent activation. Mol Cell Endocrinol 186:101–110.
Sar M, Stumpf WE. 1977. Androgen concentration in motor neurons ofcranial nerves and spinal cord. Science 197:77–79.
211ESTROGEN SYNTHESIS IN THE HINDBRAIN
Schlinger BA, Arnold AP. 1991. Brain is a major site of estrogen synthesisin a male songbird. Neurobiol 88:4191–4194.
Schlinger BA, Callard GV. 1989. Localization of aromatase in synaptoso-mal and microsomal subfractions of quail (Coturnix coturnix japonica)brain. Neuroendocrinology 49:434–441.
Schlinger BA, Greco C, Bass AH. 1999. Aromatase activity in the hindbrainvocal control region of a teleost fish: divergence among males withalternative reproductive tactics. Proc R Soc Lond 266:131–136.
Scott SA, Dinowitz S, Terhaar K, Sherlock D, Campbell MA, Levine D.1994. Cytochemical characteristics of neurons in the trigeminal mes-encephalic nucleus of hatchling chicks. J Comp Neurol 350:302–310.
Shen P, Schlinger BA, Campagnoni AT, Arnold AP. 1995. An atlas ofaromatase mRNA expression in the zebra finch brain. J Comp Neurol360:172–184.
Shughrue PJ, Lane MV, Merchenthaler I. 1997. Comparative distributionof estrogen receptor-� and -� mRNA in the rat central nervous system.J Comp Neurol 388:507–525.
Silberstein SD. 2000. Sex hormones and headache. Rev Neurol 156(Suppl4):4S30–4S41.
Simerly RB, Chang C, Muramatsu M, Swanson LW. 1990. Distribution ofandrogen and estrogen receptor mRNA-containing cells in rat brain: anin situ hybridization study. J Comp Neurol 294:76–95.
Simonian SX, Delaleu B, Caraty A, Herbison AE. 1998. Estrogen receptorexpression in brainstem noradrenergic neurons of the sheep. Neuroen-docrinology 67:392–402.
Simpson ER, Mahendroo MS, Means GD, Kilgore MW, Hinshelwood MM,Graham-Lorence S, Amarneh B, Ito Y, Fisher CR, Michael MD, et al.1994. Aromatase cytochrome P450, the enzyme responsible for estro-gen biosynthesis. Endocr Rev 15:342–355.
Steckelbroeck S, Stoffel-Wagner B, Reichelt R, Schramm J, BidlingmaierF, Siekmann L, Klingmuller D. 1999. Characterization of 17beta-hydroxysteroid dehydrogenase activity in brain tissue: testosteroneformation in the human temporal lobe. J Neuroendocrinol 11:457–464.
Steimer T. 1988. Aromatase activity in rat brain synaptosomes. Is an
enzyme associated with the neuronal cell membrane involved in medi-ating non-genomic effects of androgens. Eur J Neurosci Suppl 1988:9.
Tatsumi K, Pickett CK, Jacoby CR, Weil JV, Moore LG. 1997. Role ofendogenous female hormones in hypoxic chemosensitivity. J ApplPhysiol 83:1706–1710.
Than TT, Delay ER, Maier ME. 1994. Sucrose threshold variation duringthe menstrual cycle. Physiol Behav 56:237–239.
Timmers RJ, Lambert JG, Peute J, Vullings HG, van Oordt PG. 1987.Localization of aromatase in the brain of the male African catfish,Clarias gariepinus (Burchell), by microdissection and biochemical iden-tification. J Comp Neurol 258:368–377.
Toledo CA, Hamassaki-Britto DE, Britto LR. 1995. Serotonergic afferentsof the pigeon accessory optic nucleus. Brain Res 705:341–344.
VanderHorst VGJM, Meijer E, Schasfoort FC, Van Leeuwen FW, HolstegeG. 1997. Estrogen receptor-immunoreactive neurons in the lumbosa-cral cord projecting to the periaqueductal gray in the ovariectomizedfemale cat. Neurosci Lett 236:25–28.
Wild JM. 1989. Avian somatosensory system: II. Ascending projections ofthe dorsal column and external cuneate nuclei in the pigeon. J CompNeurol 287:1–18.
Wild JM, Zeigler HP. 1996. Central projections and somatotopic organisa-tion of trigeminal primary afferents in pigeon (Columba livia). J CompNeurol 368:136–152.
Wold JE. 1978a. The vestibular nuclei in the domestic hen (Gallus domes-ticus) III. Ascending projections to the mesencephalic eye motor nuclei.J Comp Neurol 179:393–405.
Wold JE. 1978b. The vestibular nuclei in the domestic hen (Gallus domes-ticus). IV. The projection to the spinal cord. Brain Behav Evol 15:41–62.
Yovanof S, Feng AS. 1983. Effects of estradiol on auditory evoked re-sponses from the frog’s auditory midbrain. Neurosci Lett 36:291–297.
Zucker I, Wade GN, Ziegler R. 1972. Sexual and hormonal influences oneating, taste preferences, and body weight of hamsters. Physiol Behav8:101–111.
