Organization of the cholinergic systems in the brain of two lungfishes, Protopterus dolloi and Neoceratodus forsteri

ORIGINAL ARTICLE

Organization of the cholinergic systems in the brain of twolungfishes, Protopterus dolloi and Neoceratodus forsteri

Jesus M. Lopez • Laura Domınguez •

Ruth Morona • R. Glenn Northcutt •

Agustın Gonzalez

Received: 21 June 2011 / Accepted: 23 July 2011 / Published online: 9 August 2011

� Springer-Verlag 2011

Abstract Lungfishes (dipnoans) are currently considered

the closest living relatives of tetrapods. The organization of

the cholinergic systems in the brain has been carefully

analyzed in most vertebrate groups, and major shared

characteristics have been described, although traits partic-

ular to each vertebrate class have also been found. In the

present study, we provide the first detailed information on

the distribution of cholinergic cell bodies and fibers in the

central nervous system in two representative species of

lungfishes, the African lungfish (Protopterus dolloi) and

the Australian lungfish (Neoceratodus forsteri), as revealed

by immunohistochemistry against the enzyme choline

acetyltransferase (ChAT). Distinct groups of ChAT

immunoreactive (ChAT-ir) cells were observed in the basal

telencephalon, habenula, isthmic nucleus, laterodorsal

tegmental nucleus, cranial nerve motor nuclei, and the

motor column of the spinal cord, and these groups seem to

be highly conserved among vertebrates. In lungfishes, the

presence of a cholinergic cell group in the thalamus and the

absence of ChAT-ir cells in the tectum are variable traits,

unique to this group and appearing several times during

evolution. Other characters were observed exclusively in

Neoceratodus, such as the presence of cholinergic cells in

the suprachiasmatic nucleus, the pretectal region and the

superior raphe nucleus. Cholinergic fibers were found in

the medial pallium, basal telencephalon, thalamus and

prethalamus, optic tectum and interpeduncular nucleus.

Comparison of these results with those from other classes

of vertebrates, including a segmental analysis to correlate

cell populations, reveals that the cholinergic systems in

lungfishes largely resemble those of amphibians and other

tetrapods.

Keywords Acetylcholine � Immunohistochemistry �Basal forebrain � Motor nuclei � Segmentation � Lungfish �Brain evolution

Abbreviations

a Cerebellar auricle

ac Anterior commissure

al Adenohypophysis

Am Amygdaloid complex

BST Bed nucleus of the stria terminalis

Cb Cerebellum

cc Central canal

CeA Central amygdala

DCN Dorsal column nucleus

DF Dorsal funiculus

DH Dorsal hypothalamus

dh Dorsal horn of spinal cord

do Dorsal octavolateral nucleus

Dp Dorsal pallium

flm Fasciculus longitudinalis medialis

fr Fasciculus retroflexus

Hb Habenula

ig Internal granular layer of the olfactory bulb

inf Infundibulum

Ip Interpeduncular nucleus

Ipn Interpeduncular neuropil

J. M. Lopez � L. Domınguez � R. Morona �R. G. Northcutt � A. Gonzalez (&)

Department of Cell Biology, Faculty of Biology,

University Complutense, 28040 Madrid, Spain

e-mail: [email protected]

R. G. Northcutt

Laboratory of Comparative Neurobiology, Scripps Institution

of Oceanography and Department of Neurosciences,

School of Medicine, University of California,

San Diego, La Jolla, CA, USA

123

Brain Struct Funct (2012) 217:549–576

DOI 10.1007/s00429-011-0341-x

Is Nucleus isthmi

IT Intermediate tuberal nucleus of the hypothalamus

LA Lateral amygdala

Lc Locus coeruleus

LDT Laterodorsal tegmental nucleus

LF Lateral funiculus

lfb Lateral forebrain bundle

Lp Lateral pallium

Ls Lateral septum

MC Mauthner cell

MCa Mauthner cell axon

MeA Medial amygdala

meV Mesencephalic trigeminal nucleus

ml Mitral layer of the olfactory bulb

mo Medial octavolateral nucleus

Mp Medial pallium

Ms Medial septum

nl Neural lobe of the hypophysis

NPv Nucleus of the periventricular organ

Nsol Nucleus of the solitary tract

nso Spino-occipital nerves

nsp1 First spinal nerve

nIII Oculomotor nerve

nIV Trochlear nerve

nV Trigeminal nerve

nVI Abducens nerve

nVII Facial nerve

nVIII Octaval nerve

nIX Glossopharyngeal nerve

nX Vagal nerve

oc Optic chiasm

OT Optic tectum

PA Pallidum

p 1–3 Prosomeres 1–3

pc Posterior commissure

PGPS Preganglionic parasympathetic cells

PMg Magnocellular preoptic nucleus

PO Preoptic region

PPT Pedunculopontine tegmental nucleus

PT Pretectal region

PTh Prethalamus

r 0–8 Rhombomeres 0–8

Rai Inferior raphe nucleus

Ras Superior raphe nucleus

Ri Inferior reticular nucleus

Rm Median reticular nucleus

Rs Superior reticular nucleus

SC Suprachiasmatic nucleus

sco Subcommissural organ

Smn Somatomotor neurons of spinal cord

so Spino-occipital motor nucleus

sol Solitary tract

SRN Superior reticular nucleus

Str Striatum

Th Thalamus

Tegm Mesencephalic tegmentum

Tor Torus semicircularis

TP Nucleus of the posterior tubercle

v Ventricle

VF Ventral funiculus

VH Ventral hypothalamus

vh Ventral horn of spinal cord

III Oculomotor nucleus

IV Trochlear nuclus

Vm Trigeminal motor nucleus

Vp Principal trigeminal sensory nucleus

VI Abducens nucleus

VIIm Facial motor nucleus

VIIIv Octaval ventral zone

IXm Glossopharyngeal motor nucleus

Xm Vagal motor nucleus

Introduction

Numerous studies have demonstrated that the cholinergic

systems in the central nervous system are important to

functions such as modulation of behavior, learning and

memory, the sleep-wakefulness cycle, and in superior

cognitive functions like generating of conscious experi-

ences (Vanderwolf 1987; Woolf 1991; Reiner and Fibiger

1995; Levin and Simon 1998; Perry et al. 1999; van der

Zee and Luiten 1999). The definitive anatomical mapping

of the central cholinergic cell groups and pathways became

possible only when antibodies to the enzyme choline ace-

tyltransferase (ChAT) were available. In fact, the neuro-

transmitter acetylcholine is one-step synthesized from

choline by ChAT in the cytoplasm of cholinergic neurons

and, thus, this enzyme is a reliable and specific marker of

cholinergic cells and fibers, which appear intimately rela-

ted to the distribution of acetylcholine (Kimura et al. 1981;

Crawford et al. 1982; Eckenstein and Thoenen 1983, Wa-

iner et al. 1984).

Many mapping studies have reported the distribution of

ChAT immunoreactive cells and fibers in mammalian

brains (Kimura et al. 1981; Mesulam et al. 1984; Houser

et al. 1985; Satoh and Fibiger 1985; Vincent and Reiner

1987; Maley et al. 1988; Mufson and Cunningham 1988;

Tago et al. 1989; St-Jacques et al. 1996; Ichikawa et al.

1997; Manger et al. 2002; Varga et al. 2003; Motts et al.

2008; Gravett et al. 2009) but ChAT immunohistochem-

istry has also been used in numerous comparative studies to

describe the organization of the cholinergic systems in the

550 Brain Struct Funct (2012) 217:549–576

123

brains of other vertebrate groups (lampreys, Pombal et al.

1999, 2001; elasmobranchs, Anadon et al. 2000; chond-

rosteans, Adrio et al. 2000; teleosts, Ekstrom 1987;

Brantley and Bass 1988; Molist et al. 1993; Perez et al.

2000; Clemente et al. 2004; Kaslin et al. 2004; Mueller

et al. 2004; Giraldez-Perez et al. 2009; amphibians, Marın

et al. 1997; Gonzalez et al. 2002a; reptiles, Mufson et al.

1984; Brauth et al. 1985; Hoogland and Vermeulen-Van

der Zee 1990; Medina et al. 1993; Powers and Reiner 1993;

birds, Sorenson et al. 1989; Medina and Reiner 1994). In

all these studies, the data revealed not only common fea-

tures shared by all vertebrates but also particular characters

specific to the cholinergic system of each group.

The lungfishes (dipnoans) are a group of sarcopterygian

fishes that are of extraordinary interest in investigating

evolution, including the evolution of certain neurochemical

features. This ancient group was derived from a lobe-finned

fish (Sarcopterygii) ancestor that diverged from the other

Osteichthyes in the lower Devonian, more than 390 million

years ago (Carroll 1988). Extant species belong to the in-

fraclass Dipnoi, and are segregated into two orders: Cera-

todontiformes and Lepidosireniformes. The first order has

one extant species, the Australian lungfish Neoceratodus

forsteri, whereas the second order is represented by five

extant species in two genera, the South American lungfish

Lepidosiren paradoxa, and four African lungfish species of

the genus Protopterus (P. aethiopicus, P. amphibius,

P. annectens, and P. dolloi). These orders have a mono-

phyletic origin but have been separated for at least 120

million years (Carroll 1988) when the super continent

Pangaea broke up and led to the geographic isolation of the

Australian lungfish from the South American and African

species.

The evolutionary history of lungfishes has been disputed

since their discovery because this group shares many fea-

tures with other fishes and many features with tetrapods

(Rosen et al. 1981). They occupy an interesting evolu-

tionary niche, as they diverged from the vertebrate lineage

after the divergence of most other fish lineages, such as that

leading to the teleosts, but prior to the divergence of the

amphibians. In addition, molecular phylogenetic studies

favor the lungfishes as the closest living relatives of tet-

rapods (Meyer and Wilson 1990; Meyer and Dolven 1992;

Hedges et al. 1993; Zardoya et al. 1998, Tohyama et al.

2000; Brinkmann et al. 2004a, b; Hallstrom and Janke

2009).

In spite of their important phylogenetic position, the

cholinergic system in the brain of lungfishes has not been

previously investigated. Therefore, the main goal of the

present study was to provide the first detailed information

on the localization of ChAT immunoreactive (ChAT-ir)

cell bodies and fibers in the brain of representative species

of the two orders of dipnoans, an African lungfish

(Protopterus dolloi) and the Australian lungfish (Neocer-

atodus forsteri). Furthermore, we have compared our

results with those from other vertebrate classes to shed

light on the evolution of the cholinergic systems and extend

what is known about this neurotransmitter systems. In

addition, we combined our ChAT immunohistochemistry

with that for tyrosine hydroxylase (TH, the first and rate-

limiting enzyme for catecholamine synthesis) and calbin-

din-D28k (CB). These experiments were added because the

neuroanatomy of lungfishes is not well documented, and

the combined localization of ChAT-ir structures and TH-ir

and CB-ir structures was useful in clarifying the actual

position of certain cell groups.

Materials and methods

A total of ten juvenile African lungfish, Protopterus dolloi,

and five juvenile Australian lungfish Neoceratodus forsteri

were used in this study. Although the sex of all animals was

not established, males and females of both species were

studied. The African lungfish were purchased from com-

mercial suppliers (PezyCia, Madrid, Spain), and the Aus-

tralian lungfish were obtained from Jindalee International

Pty Limited in Milton, Queensland, a approved breeder and

exporter. All animals were maintained in aquaria at

24–288C under natural light conditions. The original

research reported herein was performed according to the

regulations and laws established by the European Union

(86/609/EEC) and Spain (Royal Decree 1201/2005) and in

conformation with standards established by the Institu-

tional Animal Care and Use Committee at the University of

California, San Diego for the care and handling of animals

in research.

The animals were deeply anesthetized by immersion in

0.01% tricaine methanesulfonate solution (MS222, Sandoz

Basel, SW; pH 7.3) and perfused transcardially with

physiological saline followed by 200 ml of cold 4% para-

formaldehyde in 0.1 M phosphate buffer (PB, pH 7.4). The

brain and the spinal cord were removed from the skulls and

kept in the same fixative for 2–3 h. Subsequently, they

were immersed in a solution of 30% sucrose in PB for

4–6 h at 4�C until they sank, then embedded in a solution

of 20% gelatin with 30% sucrose in PB, and stored for 6 h

in a 3.7% formaldehyde solution at 4�C. The brains were

cut on a freezing microtome at 40 lm in the transverse or

sagittal plane, and sections were collected and rinsed in

cold PB.

ChAT immunohistochemistry

The free-floating sections were rinsed twice in PB, treated

with 1% H2O2 in PB for 15 min to reduce endogenous

Brain Struct Funct (2012) 217:549–576 551

123

peroxidase activity, rinsed again three times in PB and

processed by the peroxidase antiperoxidase (PAP) method

(Sternberger 1979). This included a first incubation of the

sections in a goat anti-ChAT serum (Chemicon, Temecula,

CA, USA; AB144P) diluted 1:100 in PB containing 0.5%

Triton X-100 (PBS-T), 15% normal rabbit serum (NRS),

and 2% bovine serum albumin (BSA), for 48 h at 4�C.

Subsequently, the sections were rinsed three times in PB

for 10 min and incubated for 60 min at room temperature

in rabbit anti-goat serum (Chemicon) diluted 1:50. After

rising again three times for 10 min each, the sections were

incubated for 90 min in goat PAP complex (diluted 1:500;

Chemicon). Secondary antiserum and PAP complex were

diluted in PB containing 0.5% Triton X-100, 15% NRS and

2% BSA. Finally, the sections were rinsed three times for

10 min each in PB and subsequently stained in 0.5 mg/ml

3,30-diaminobenzidine (DAB; Vector SK4100) intensified

with nickel (Adams 1981), 0.01% H2O2 in PB for 3–5 min.

Transverse and sagittal series were mounted on glass slides

with 0.25% gelatin in 0.1 M Tris–HCl buffer (TB, pH 7.6)

and, after dehydration, coverslipped with Entellan (Merck,

Darmstadt, Germany). Some sections were counterstained

with cresyl violet to facilitate analysis of the results.

Double ChAT and tyrosine hydroxylase (TH)

or calbindin-D28k (CB) immunohistochemistry

A procedure based on immunohistofluorescence was used

as follows: (1) first incubation for 72 h at 4�C in a mixture

of goat anti-ChAT (diluted 1:100) and mouse anti-TH

(diluted 1:1,000; Immunostar, USA; code P22941) or

rabbit anti-CB (diluted 1:1,000; Swant, Bellinzona, Swit-

zerland; cat. number CB-38a); (2) second incubation for

90 min at room temperature in a mixture of secondary

antisera: donkey anti-goat Alexa 594 (red fluorescence;

diluted 1:300; Molecular Probes, Denmark) and chicken

anti-mouse Alexa 488 (green fluorescence; diluted 1:300;

Molecular Probes) or FITC-conjugated chicken anti-rabbit

(green fluorescence; diluted 1:100; Chemicon). After

rinsing three times in PB, the sections were mounted on

glass slides and coverslipped with Vectashield (Vector,

Burlingame, CA).

Western blotting analysis

Two animals of each lungfish species were anesthetized in

MS222, and the brains were quickly removed and

mechanically homogenized in an equal volume of cold

buffer (5 mM EDTA, 20 mM Tris, pH 7.4, 150 mM NaCl,

10% glycerol, 1% Nonidet P40; Roche, Mannheim, Ger-

many) supplemented with protease and phosphatase

inhibitors (50 lg/ml phenylmethyl-sulfonyl fluoride,

10 lg/ml aprotinin, 25 lg/ml leupeptin, and 100 nM

orthovanadate; all from Sigma). Samples of the superna-

tants containing in each case 50 lg of protein were applied

in each lane of a 12% polyacrylamide gel (#161-0801, Bio-

Rad, Hercules, CA) and separated by sodium dodecyl

sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)

with a Mini-Protean system (Bio-Rad). The samples of rat

and Xenopus brain and molecular weight standards (Pre-

cision Plus Protein Dual color Standards, Bio-Rad) were

run in other lanes. The separated samples in the gel were

transferred to nitrocellulose membrane (Bio-Rad). Non-

specific binding sites were blocked by incubation overnight

in Tris–HCl buffer (TBS) containing 0.1% Tween-20

(TBST) and 5% nonfat milk at 4�C. The blots were then

incubated for 24 h at 4�C in primary antibody dilution (as

for immunohistochemistry). After rinsing in TBS, the blots

were incubated in horseradish peroxidase-coupled sec-

ondary goat anti-mouse or goat anti-rabbit antisera (Jack-

son ImmunoResearch, West Grove, PA; diluted 1:15,000)

for 2 h at room temperature. Immunoreactive bands were

detected using an enhanced chemiluminescence system

(Super Signal West Pico Chemiluminescent Substrate,

Pierce, Thermo Scientific, Rockford, IL). Photographs

were taken after applying an autoradiographic film to the

membrane, in the darkness, for 1–4 min.

Controls and specificity of the antibodies

There were two general controls for the immunohisto-

chemical reaction: (1) staining of selected sections with

preimmune rabbit serum; and (2) omission of the primary

and/or the secondary antibody was omitted. In all these

negative controls, the immunostaining was eliminated. In

addition, Western blot analysis using brain extract from

Protopterus dolloi and Neoceratodus forsteri showed that

all the antibodies we used labeled a band (Fig. 1), and, with

slight variations, that band corresponded well with the

bands labeled in the rat and Xenopus lanes used as refer-

ences (see also Morona et al. 2011).

The specificity of the three antibodies used has been

assessed by the commercial companies who manufactured

them (Table 1). The specificity of the ChAT antiserum was

analyzed by immunoblot (and Western blot) performed in

rat, guinea pig, and rabbit, in which a band in the range of

68–70 kDa was always observed (see manufacturer’s data

sheet). In addition, Western blot analysis of protein extracts

from the brains of dogfish, sturgeon, trout, and diverse

amphibians showed the presence of similar bands of

68–72 kDa (Anadon et al. 2000; Morona and Gonzalez

2009). The band observed in the Western blots from the

two lungfish brains (Fig. 1a; Morona et al. 2011) corre-

sponds to that from rat and Xenopus brain extracts at the

expected molecular weight in relation to the published

nucleotide sequence for rat ChAT (NCBI accession number


123

XM_001061520). Furthermore, the staining with this

antibody colocalizes with the mRNA distribution of the

same enzyme by in situ hybridization histochemistry (Oh

et al. 1992). In addition, we used the specific peptide

(Chemicon, catalog number AG220) for blocking the

staining of the AB144P antibody, following the manufac-

ture’s protocol (2–5 lg/mL, incubated with the antibody

for 3 h at 4�C prior to use). The specificity of the TH

antibody was corroborated by Western blot analysis in rat,

mouse, ferret, cat, and Aplysia (see specification data sheet;

ImmunoStar), diverse amphibians (Morona and Gonzalez

2009), and the turtle Pseudemys scripta (Moreno et al.

2010) in which it selectively labels a single band at

approximately 62 kDa. The Western blot performed with

brain extracts from the two lungfishes (Morona et al. 2011)

detected a single band at the same molecular weight (about

62 kDa) as that of the major product detected in rat and

Xenopus brain extract (Fig. 1b). In the case of the poly-

clonal antibody anti-CB used in this study, it was shown in

Western blot that it labeled a single band between 28 and

29 kDa in lanes with brain extracts of four different

amphibian species (Morona and Gonzalez 2008), and a

similar band was observed in the lanes of rat and lungfish

brain extracts (Fig. 1c). Finally, it should also be noted that

these ChAT, TH and CB antibodies have recently been

used in immunohistochemical studies of the brain in

lungfishes and yielded results comparable to those from

many other vertebrates (Gonzalez and Northcutt 2009;

Gonzalez et al. 2010).

Evaluation and presentation of the results

The distribution of ChAT-ir cell bodies and fibers in the

brains of Protopterus and Neoceratodus was carefully

analyzed and the pattern of labeling was charted in repre-

sentative transverse sections at different brain levels

(Figs. 2, 3). Drawings were made by means of camera

lucida in which the sections counterstained with cresyl

violet facilitated the interpretation of the localization of the

labeled structures. The sections were analyzed with an

Olympus BX51 microscope equipped with a digital camera

(Olympus DP70). Contrast and brightness were adjusted in

Adobe Photoshop CS3 (Adobe Systems, San Jose, CA),

and photos were mounted on figures in Canvas 11 (ACS

Systems International).

The nomenclature used is essentially the same followed

in previous recent studies of lungfish brains (Gonzalez and

Northcutt 2009; Northcutt 2009, 2011).

Results

The antibody against ChAT used in the present study

revealed patterns of immunoreactivity that were constant

from animal to animal in both species, and no indications

Fig. 1 Western blot identification of protein bands recognized in the

two lungfishes for the antibodies used: a goat anti-ChAT, b mouse

anti-TH, and c rabbit anti-CB. A single band is seen in each of the

lines corresponding to the extracts from the lungfish brains, which are

compared with the bands stained in each case for brain extracts from

Xenopus and rats. Molecular weight standards are indicated on the left

Table 1 Antibodies used in the present study

Name Immunogen Commercial supplier MW (KDa) Dilution

ChAT Whole human placental enzyme Polyclonal goat anti-ChAT

Chemicon; catalogue reference: AB 144P

68 1:100

TH TH purified from rat PC12 cells Monoclonal Mouse-anti-TH

ImmunoStar, Inc. Catalogue reference: 22941

62 1:1,000

CB Recombinant rat calbindin-D28k Polyclonal rabbit-anti-CB. Swant; catalogue reference: CB-38a 38 1:1,000


123

Fig. 2 Diagrams of transverse sections (a–u) through the brain of

Protopterus dolloi at the rostrocaudal levels indicated in the lateral

view of the brain. ChAT-ir cell bodies (large dots) and fibers (small

dots, wavy lines) are represented in the right half of each section.

Faintly labeled cells are drawn empty. Scale bars 500 lm


123

of sex differences were observed. In both transverse and

sagittal sections, labeling of neuronal cell bodies and fibers

was observed in every major brain division. A summary of

the interspecific variations in the distribution of cholinergic

cells and fibers in the brains of the two lungfishes is shown

in Table 2. The general brain distribution of ChAT-ir

neurons and fibers in Protopterus dolloi and Neoceratodus

forsteri is illustrated in a series of selected transverse

sections in Figs. 2 and 3, respectively. Widespread

immunoreactive structures are shown in selected photo-

micrographs in Figs. 4, 5, 6 and 7 and will be described in

the following sections from rostral to caudal levels. The

sections double-labeled for ChAT and TH, or ChAT and

CB, were mainly used to corroborate the topographical

position of certain cell groups in the brainstem, and

selected images are shown in Fig. 8.

Forebrain

The olfactory bulbs of Protopterus were devoid of ChAT-ir

elements. Only in Neoceratodus were some immuno-

reactive fibers and terminals seen in the mitral and

internal plexiform layers of the olfactory bulb (Fig. 3a).

In both species, the pallium also lacked ChAT-ir cells,

although some immunoreactive fine fibers and varicosi-

ties were seen in the medial zone of the medial pallium

(Figs. 2a, b, 3b). In comparison, the subpallium was rich

in ChAT-ir elements (Figs. 2b–d, 3b–d). A conspicuous

telencephalic group of cholinergic cells was located in

the ventrocaudal telencephalon, forming the basal fore-

brain cholinergic system of lungfishes. This cell group

was distributed in the vicinity of the recently described

region of the bed nucleus of the stria terminalis and the

pallidum (Gonzalez and Northcutt 2009) and was con-

stituted by small round cells with long lateral and ven-

trolateral processes (Figs. 2c, d, 3d, 4a, b). In the rostral

part of this group, some cells were seen laterally within

the ventral striatal region and medially in the lateral

septum (Figs. 2b, 3c, 4c, d). The number of cells in this

basal cholinergic group was higher at the level of the

anterior commissure, and caudally it reached the level of

rostral preoptic area (Figs. 2c, d, 3e, 4e).

The amygdaloid complex in both species was devoid of

cholinergic cells, and only moderate numbers of labeled

fibers were found, mainly in the central amygdala

(Figs. 2c, 3d). The fibers and terminal-like structures

Fig. 2 continued


123

Fig. 3 Diagrams of transverse sections (a–v) through the brain of

Neoceratodus forsteri at the rostrocaudal levels indicated in the lateral

view of the brain. ChAT-ir cell bodies (large dots) and fibers (small

dots, wavy lines) are represented in the right half of each section.

Faintly labeled cells are drawn empty. Scale bars 500 lm


123

observed in the telencephalon probably belong to cholin-

ergic cells intrinsic to the telencephalon.

In the preoptic area, a small group of dispersed and very

weak ChAT-ir cells were seen in periventricular positions

and showed large perikarya and a few processes directed

ventrolaterally (Figs. 2e, 3f). Because of their size and

morphology, these cells probably belong to the neurose-

cretory magnocellular preoptic nucleus. Additionally, in

Neoceratodus, small cholinergic cells were observed in the

preoptic area and frequently showed processes that con-

tacted the cerebrospinal fluid (CSF) (Figs. 3e, f, 4e).

The suprachiasmatic nucleus of Protopterus was devoid

of ChAT-ir cells and distinct bundles of nonvaricose

ChAT-ir fibers decussate in the supraoptic commissure and

course parallel to the optic tract (Fig. 2f). These fibers also

were observed in Neoceratodus, in which a notable group

of weakly stained cells were seen in the suprachiasmatic

nucleus (Figs. 3g, 4f). Many of these cells showed a long

thin process contacting the CSF and another process

directed laterally or ventrolaterally.

The basal hypothalamus of Protopterus was devoid of

ChAT-ir cells, whereas two groups of cholinergic cells

were seen in the hypothalamic region of Neoceratodus

(Fig. 3h, i): one in the dorsal part of the nucleus of the

periventricular organ (Fig. 4g) and one in the intermediate

tuberal nucleus (formerly considered the ventral hypo-

thalamus; Fig. 4h). Both groups showed rounded cells with

long CSF-contacting processes. In addition, a number of

thin, varicose ChAT-ir fibers were seen coursing along the

infundibular region (Figs. 2g–i, 3h, i) and reaching the

neural lobe of the hypophysis further caudally (Figs. 2j, k,

3i); these fibers were most likely part of the fiber tract from

the magnocellular preoptic cells.

In the epithalamus, the habenula showed a substantial

population of weakly stained ChAT-ir cells, formed by

small, round closely packed neurons restricted mainly to

the medial habenular region in Protopterus (Fig. 2e, f). The

number of cholinergic cells and the intensity of the

immunoreactivity were both higher in Neoceratodus,

where these cells occupied medial and lateral positions

Fig. 3 continued


123

(Figs. 3f, g, 4i). Labeled axons originating in these cells

were seen to follow a caudoventral path, thus forming the

fasciculus retroflexus. In Neoceratodus this tract appeared

to be divided into fasciculi (Figs. 3h, i), which reached the

ventral mesencephalic surface on both sides and converged

in the large interpeduncular neuropil at the isthmic level

(Figs. 2e–m, 3h–o).

Along the rostrocaudal extent of the thalamus (formerly

termed the dorsal thalamus), a sizable group of cholinergic

cells was observed near the ventricle in the most dorsal part

of this region (Figs. 2f, g, 4j). These cells were small in

size, round shaped and with very long processes directed

laterally. In Neoceratodus, this cholinergic group was

clearly more reduced and limited to the caudal part of the

thalamus (Fig. 3h). A notable plexus of ChAT-ir fibers and

terminals was formed in the lateral part of the thalamus in

both species, particularly in Neoceratodus (Figs. 3f, g, 4k),

and labeled fiber tracts were also observed in the superficial

Table 2 Comparative

localization and relative

abundance of ChAT-ir cells and

fibers in CNS of the species of

lungfishes studied

C immunoreactive cell bodies,

F immunoreactive fibers

?, low density; ??, moderate

density; ???, high density; -,

no immunoreactive cell bodies

or fibers

Protopterus dolloi Neoceratodus forsteri

C F C F

Forebrain

Olfactory bulb 2 2 2 ?

Pallium 2 ? 2 ?

Striatum ? ? ? ?

Lateral septum ?? ?? ?? ??

Basal telencephalon ??? ?? ??? ??

Amygdaloid complex 2 ? 2 ?

Preoptic area ? ?? ?? ??

Suprachiasmatic nucleus 2 ?? ?? ??

Tuberal hypothalamus 2 ?? ?? ??

Hypophysis 2 ?? 2 ?

Habenula ?? ?? ??? ??

Thalamus ??? ?? ? ???

Prethalamus 2 ?? 2 ???

Pretectum 2 ?? ? ???

Midbrain

Optic tectum 2 ?? 2 ???

Mesencephalic tegmentum 2 ?? 2 ??

III ?? ?? ??? ??

Hindbrain

IV ?? ?? ??? ??

Isthmic nucleus ?? ?? ?? ??

Interpeduncular nucleus 2 ?? 2 ??

Laterodorsal tegmental nucleus ??? ?? ??? ??

Cerebellum 2 2 2 ??

Superior reticular nucleus 2 ? ?? ??

Vm ??? ?? ??? ??

Superior raphe nucleus 2 ? ?? ??

VI ?? ? ?? ?

Middle reticular nucleus 2 ? ? ??

Medial octavolateral nucleus ? ? ? ??

VIIm-IXm ??? ?? ??? ??

Solitary tract nucleus ? ? ? ??

Dorsal column nucleus ? ? ? ??

Xm ??? ?? ??? ??

Inferior reticular nucleus ? ?? ? ??

Spino-occipital motor nucleus ??? ?? ??? ??

Spinal cord ??? ? ??? ??


123

zone of the prethalamus (formerly the ventral thalamus)

(Figs. 3f–h, 4l).

Only a moderate plexus of ChAT-ir fibers and varicos-

ities was observed in the superficial zone of the pretectal

region in Protopterus (Fig. 2h), whereas there was a small

group of round cholinergic cells with long lateral processes

that arborized in the superficial zone in the pretectal region

of Neoceratodus (Fig. 3i).

Fig. 4 Photomicrographs of transverse sections through the forebrain

of Protopterus dolloi (a, c, j) and Neoceratodus forsteri (b, d–i, k,

l) illustrating ChAT-ir cell bodies and fibers in the ventrocaudal

telencephalon (a, b), striatum and lateral septum (c, d), preoptic area

(e; arrows point to CSF-contacting cell processes), suprachiasmatic

nucleus (f), nucleus of the periventricular organ and intermediate

tuberal nucleus of the hypothalamus (g, h; arrows point to CSF-

contacting cell processes), habenula (i), thalamus (j, k; arrows in

k point to weakly stained cells) and prethalamus (l). (a and j are

higher magnifications of the framed area shown in a panoramic view

on the upper left side). Scale bars 100 lm


123

Fig. 5 Photomicrographs of transverse and sagittal (d, m) sections

through the brainstem of Protopterus dolloi (a, e, i, k) and

Neoceratodus forsteri (b–d, f–h, j, l, m) illustrating ChAT-ir cells

and fibers in the optic tectum (a, b), mesencephalic tegmentum (c),

oculomotor and trochlear nuclei (d–g; asterisk in d marks the gap

between the oculomotor and trochlear nuclei, whereas in f marks

lightly stained cells of the possible Edinger–Westphal nucleus),

isthmic and laterodorsal tegmental nuclei (h–m; arrowheads in h and

i point to lightly stained cell; arrows in c, j and m mark the fiber tract

from isthmic and laterodorsal tegmental nuclei through the mesen-

cephalon). Scale bars 500 lm (h, m), 200 lm (d–g, i, j, l), 100 lm

(a–c), 50 lm (k)


123

Fig. 6 Photomicrographs of transverse and sagittal (b) sections

through the rostral and middle rhombencephalon of Protopterusdolloi (c, e, f, h, l) and Neoceratodus forsteri (a, b, d, g, i–k, m,

n) illustrating ChAT-ir cells and fibers in the pretrigeminal reticular

zone (a), trigeminal motor nucleus (b–d), the level of entrance of the

facial motor nerve and the course of the axons toward the midline

(arrows) (e), level of the Mauthner cell and rostral facial nucleus (f),superior raphe nucleus (g), abducens nucleus (h–j; h is a higher

magnification of the framed area), facial and glossopharyngeal nuclei

(k–n; arrows in k and m point to axons from the facial motoneurons).

Scale bars 200 lm (b, c, f), 100 lm (a, d, e, g, i, k–n), 50 lm (h)


123

Fig. 7 Photomicrographs of transverse and sagittal (a, k) sections

through the caudal rhombencephalon and spinal cord of Protopterusdolloi (b, d, f, g, l) and Neoceratodus forsteri (a, c, e, h–k, m, n)

illustrating ChAT-ir cells and fibers in the vagal motor nucleus (a–c),

nucleus of the solitary tract (b, c), dorsal column nucleus (d, e), spino-

occipital motor nucleus (f–i; g is a higher magnification of the framedarea), lateral reticular cell near the obex (k), and the spinal cord (l–n;

arrows mark the small cholinergic cells of the possible sympathetic

autonomic system). Arrows in a and k point to vagal motor rootlets.

Scale bars 200 lm (a, b, l), 100 lm (c, e, f, g–j, m, n), 50 lm (d, k)


123

Midbrain

No ChAT-ir cells were observed in the optic tectum in

either of the two species studied. In Protopterus, scattered

varicose fibers and terminals were sparsely distributed in

the tectum, predominantly in the superficial layers

(Figs. 2i–l, 5a). In Neoceratodus, this innervation was

more pronounced and not only limited to the superficial

Fig. 8 Photomicrographs showing, in the same sections, staining for

ChAT (red fluorescence) and TH (a–c, g) or CB (d–f, h–j) (greenfluorescence). The relationship between the distinct cell populations is

illustrated for the oculomotor nucleus (a, b), and trochlear nucleus

(b–d). Both nuclei are separated by the prominent catecholaminergic

group in the ventral mesencephalic tegmentum (a–c). The trochlear

nucleus is followed caudally by the CB positive cell population of the

interpeduncular nucleus (d). The cholinergic cells of the laterodorsal

tegmental nucleus are intermingled with small CB immunopositive

cells (e, f) and caudally are limited by the scarce and large TH

immunoreactive cells of the locus coeruleus (g). The large and faintly

immunoreactive cholinergic cells of the rostral pole of the trigeminal

motor column are generally immunoreactive for CB (h–j). Scale bars500 lm (a, e), 200 lm (b, c, f, g), 100 lm (h–j)


123

layers but also included the deeper periventricular layers

(Figs. 3j–m, 5b). There was no clear laminar organization

of these ChAT-ir fibers in the tectum.

In the dorsal tegmentum, thick ChAT-ir fibers from the

cholinergic groups of the isthmic region (Fig. 5m) were

observed in a tract coursing rostrally and dorsally (Figs. 2j, k,

3j, k, 5c), toward the diencephalon and, to a lesser extent,

the tectum.

In the ventral mesencephalic tegmentum, the oculomo-

tor neurons were intensely ChAT-ir (Figs. 2j, 3j, 5d–f). The

oculomotor nucleus in both species was formed by a

population of medium-sized cells (more numerous in

Neoceratodus than in Protopterus) grouped in the peri-

ventricular gray of the rostral mesencephalon (Figs. 5d,

8a–c). The dendrites of the oculomotor neurons arborized

profusely toward the ventrolateral aspect of the tegmentum

(Fig. 5e, f). The axons of most oculomotor neurons col-

lected at the ventrolateral aspect of the nucleus and exited

the brain ventrolaterally in the oculomotor nerve (Figs. 3j,

5e, f). However, one group of axons was consistently seen

to cross ventrally and exit in the contralateral nerve (arrows

in Fig. 5e, f). The oculomotor nucleus extended rostro-

caudally over a distance of 250–300 lm, and was located

just rostral to a prominent catecholaminergic group in the

ventral mesencephalic tegmentum that labeled intensely for

TH (Fig. 8a, b). In general, differences in neuronal size

were noted in the oculomotor neurons, but only Neocer-

atodus showed small cholinergic cells situated dorsal to the

large oculomotor neurons (asterisk in Fig. 5f). These cells

showed less immunoreactivity and could belong to the

Edinger–Westphal nucleus.

Hindbrain

The hindbrain showed the most conspicuous cholinergic

cell groups in both species, mainly due to the presence of

motor nuclei of the cranial nerves, although some addi-

tional ChAT-ir cells were also seen in several places in the

rhombencephalon.

In the basal plate of the isthmic region, caudal to the

oculomotor nucleus, the trochlear nucleus was seen to form

a column of cholinergic cells in a ventromedial position

(Figs. 2k, 3k, l, 5d, g). These neurons were the same size as

those of the oculomotor nucleus but had shorter dendritic

trees, which were directed mainly ventrolaterally (Fig. 5g).

Moreover, their axons were seen to collect into several

fascicles that coursed dorsolaterally in the tegmentum

(Figs. 3k, l, 5g) and, immediately rostral to the cerebellum,

crossed to the contralateral side to leave the brain dorsally,

(Fig. 5h, i). The trochlear nucleus extended rostrocaudally

for about 200 lm and, as seen in sagittal section, a large

gap existed between it and the oculomotor nucleus

(Fig. 5d); this gap was primarily occupied by the

catecholaminergic cells that form the potential lungfish

homologue of the substantia nigra/ventral tegmental area

(Reiner and Northcutt 1987; Gonzalez and Northcutt 2009)

(Fig. 8a, b). The caudal limit of the trochlear nucleus was

clearly marked by a CB-ir population of neurons in the

interpeduncular nucleus (Fig. 8d). A notable plexus of

immunoreactive nerve terminals was also seen in the

neuropil of the interpeduncular nucleus (Figs. 2k–m, 3k–o,

5g, h).

Dorsal and caudal to the trochlear nucleus two different

ChAT-ir cell groups were found (Figs. 2l, 3l, m, 5h, m).

The first was formed by a sparse population of small and

weakly stained cells, located dorsolaterally in the teg-

mentum. These cells showed short, laterally directed pro-

cesses (Figs. 2l, 3l, 5i, j). The second group comprised

more numerous medium-sized and intensely ChAT-ir

neurons, located ventromedial to the first group (Fig. 5j, m)

and just rostral to the cells of the locus coeruleus, as

revealed by double labeling with TH (Fig. 8g). These

cholinergic cells showed long dendrites that arborized

ventrolaterally (Figs. 2l, 3m, 5j–l) and intermingled with

small CB-ir cells, with some cells being double labeled

(Fig. 8e, f). Because of their position and cholinergic nat-

ure, we have designated these dorsolaterally and ventro-

medially located cell groups the putative nucleus isthmi

and laterodorsal tegmental nucleus, respectively. Axons

from these nuclei were seen to form the dorsorostrally

directed thick tract observed through the mesencephalon

(Figs. 5c, j, m, 8e, f). In addition, some cholinergic cells

(primarily in Neoceratodus), lying medial to the cells of the

isthmic nucleus and close to the ventricular surface, were

weakly ChAT-ir and formed a separate cell population

(Figs. 3l, 5h, i).

The cerebellum in both species was devoid of ChAT-ir

cells, although some cholinergic fibers were seen in the

granule cell layer in Neoceratodus (Fig. 3n). Also in

Neoceratodus only, a prominent pretrigeminal group of

medium-sized, bipolar cholinergic cells was detected in the

superior reticular nucleus, extending to the rostral pole of

trigeminal motor nucleus (Figs. 3n, 6a). Some of these

cells had very long processes that crossed the midline to the

contralateral reticular formation.

Caudal to the laterodorsal tegmental nucleus, and sep-

arated from it by about 200 lm, the large trigeminal motor

nucleus was intensely labeled (Figs. 2m, 3o, 5m). This

nucleus is located in the ventrolateral periventricular gray

and forms a long column (about 1,800 lm long) extending

up to the level of entrance of the facial motor root

(Fig. 6b). Rostral and caudal parts of the trigeminal motor

nucleus could be distinguished. The rostral part consisted

primarily of large neurons with round somata and weak

ChAT immunoreactivity (Fig. 6b, c). Unlike the rest of

trigeminal motor neurons, these large neurons were


123

double-labeled for CB (Fig. 8h–j). The caudal part of the

trigeminal motor nucleus comprised smaller neurons that

were intensely ChAT-ir (Fig. 6b, d). Axons of the tri-

geminal motoneurons were seen to course in a ventro-

lateral direction and exit the brainstem in the ventral

one-fourth of the trigeminal root (Figs. 2m, 3o, 6c, d).

Medial to the trigeminal motor nucleus, a discrete group

of weakly immunoreactive cholinergic cells were

observed in the superior raphe nucleus in Neoceratodus

only (Fig. 3o, 6g). These small rounded cells had pro-

cesses directed ventrally.

The caudal pole of the trigeminal nucleus coincided

with the level of the entrance for the facial nerve, just

beneath the octaval nerve root (Figs. 2n, 3p, 6b, e, f). The

axons of the facial nerve were seen to follow a peculiar

course, progressing from the nerve root medially and then

caudally above the Mauthner axon and the medial longi-

tudinal fascicle (Figs. 2n–p, 3p–s, 6e, f, h–k, m, n). At this

level, bilaterally, the giant perikarya of the Mauthner cells

were not ChAT-ir and the few immunoreactive cells

present likely represent a minor rostral component of the

facial motor nucleus (Fig. 6f).

Approximately 200 lm caudal to the Mauthner cells, a

small group of scattered ChAT-ir cells was seen ventral to

the bundle of facial axons in a medial positions and

adjacent to the medial longitudinal fascicle (Figs. 2o, 3q,

6h–j). These medium-sized cells represented the sparse

population of neurons of the abducens nucleus. They

possessed long ventral dendritic arborizations and their

axons were seen to collect in the ventral aspect of the

rhombencephalon and exit the brain ventrally in the

abducens nerve (Figs. 2o, 6j). No accessory abducens

nucleus was detected in the adjacent lateral region. Lat-

eral to the abducens motoneurons, a discrete group of

weakly ChAT-ir cells was observed in the median retic-

ular nucleus in Neoceratodus only (Figs. 3q, 6j). These

large cells possessed long processes directed ventrally and

extending caudally up to the level of the facial motor

nucleus (Figs. 3r, 6k).

At the same levels as the trigeminal and abducens motor

nuclei, small round and weakly labeled cells were seen in

the medial octavolateral nucleus (Figs. 2n, o, 3o, 6l), and

there was notable innervation of the dorsal and medial

octavolateral nuclei, more abundant in Neoceratodus than

in Protopterus (Figs. 3o–q).

The facial axons coursing caudally above the medial

longitudinal fascicle were seen to turn laterally and reach

their cell bodies of origin in the facial motor nucleus

(Figs. 2p, 3r, s, 6k, m). The morphology of these cells

resembled that of the trigeminal neurons, with long den-

dritic arborizations directed ventrolaterally (Fig. 6k, m).

The facial motor nucleus appeared as a caudal continuation

of the abducens nucleus, but in the ventrolateral reticular

formation. Its caudal pole could not be distinguished

because it overlapped with the motoneurons of the glos-

sopharyngeal motor nucleus in the ventrolateral gray

(Figs. 3s, 6n). The morphology of the glossopharyngeal

motoneurons was similar to that of the facial motoneurons

with axons directed laterally (Figs. 2q, 6n). The absence of

medially directed axons served to distinguish the glosso-

pharyngeal motoneurons from the facial motoneurons

(Fig. 3s). Also the facial column in Neoceratodus appeared

longer than that in Protopterus, and in Neoceratodus only,

some small cholinergic cells were observed dorsolateral to

the large facial motoneurons (Fig. 6m). These cells showed

less immunoreactivity and could represent preganglionic

parasympathetic cells.

Starting at the level of the caudal pole of the glosso-

pharyngeal motor nucleus and extending caudally to the

obex region, a population of small ChAT-ir cells at the

ventrolateral margin of the solitary tract was identified as

part of the nucleus of the solitary tract (Figs. 2q–s, 3s, t, 7b,

c). Dorsal to the solitary tract, at caudal rhombencephalic

levels, small ChAT-ir cells were seen in the dorsal column

nucleus (Figs. 2r, 3s, 7d, e).

Immediately caudal to the glossopharyngeal motor

nucleus, the vagal motor nucleus was intensely labeled and

formed a long column in the ventrolateral reticular zone

that extended caudally to the obex into the upper spinal

cord (Figs. 2r–t, 3t, u, 7a, b). The vagal motoneurons

possessed long lateral dendritic arborizations, and their

axons collected in the lateral aspect of the caudal rhomb-

encephalon to exit the brain laterally in several rootlets

(Figs. 2r–t, 3t, 7a, b, f, k). The preganglionic parasympa-

thetic neurons associated with the facial, glossopharyngeal

and vagal motor nuclei were not clearly distinguishable

with ChAT immunohistochemistry in Protopterus,

although the small and weakly ChAT-ir cells observed in

these nuclei in Neoceratodus could represent this auto-

nomic component of the cholinergic system (Figs. 6m, n,

7a, c).

Ventromedial to the vagal motor nucleus and close to

the midline, a conspicuous group of strongly labeled cells

was seen to form the spino-occipital nucleus (Figs. 2r–t, 3t,

u, 7f–j). These large neurons showed very long dendritic

arborizations, ventrally or ventrolaterally directed, and

their axons were observed to collect in the ventral aspect of

the caudal rhombencephalon, exiting ssthe brain ventrally

in the spino-occipital nerves (Figs. 2r–t, 3t, u, 7f–j). The

spino-occipital nucleus comprised a long column of neu-

rons that were continuous with the somatic motoneurons of

the spinal cord. Occasionally, some small, round, weakly

labeled cells were observed in the inferior reticular

nucleus, between the vagal and spino-occipital nuclei in

caudal rhombencephalic levels close to the obex (Figs. 3t,

7h–j). Some larger cells were also seen occasionally in


123

lateral reticular zones, near the axons of the spino-occipital

nucleus (Fig. 7k).

Spinal cord

In the upper segments of the spinal cord, ChAT-ir moto-

neurons were seen to occupy the ventrolateral margin of

the gray matter within the ventral horn (Figs. 2u, 3v, 7l,

m). These large cells possessed profuse dendritic arbor-

izations that almost entirely filled the ventrolateral white

matter, and their axons collected to exit the brain ventrally

in the ventral spinal roots (Figs. 3v, 7l, m). In the inter-

mediate gray zone of the rostral spinal cord, a tiny group of

small ChAT-ir cells likely corresponded to the caudal

continuation of the vagal motor nucleus (Fig. 7l). No

cholinergic cells were observed in the dorsal gray field of

the spinal cord.

The presence of this group of large spinal motoneurons

in the ventral horn was a feature observable throughout the

rostrocaudal extent of the spinal cord. At midlevels, cor-

responding to the thoracic spinal cord, some small ChAT-ir

cells were in the intermediate gray zone might belong to

the sympathetic autonomic system (asterisk in Fig. 7n). In

addition, the presence of ChAT-ir fibers in the dorsal and

lateral funiculi was especially abundant in Neoceratodus

(Figs. 3v, 7m).

Discussion

The present study provides the first detailed description of

the organization of the cholinergic systems in the brain of

two representative species of dipnoans. We used the same

ChAT immunohistochemical techniques used extensively

in similar studies of different vertebrates classes, thus

allowing a direct comparison of the results obtained.

These comparisons with other vertebrate groups are

shown in Table 3 and highlight both conserved and var-

iable features of cholinergic systems across phylogeny;

for specific features of each vertebrate class, references

are indicated. In the following sections we will discuss

the general organization of the cholinergic systems in

lungfishes and variations with that in other vertebrates, in

order to define primitive versus derived features of these

systems. Finally, we have made an attempt to frame the

cholinergic cell populations in lungfishes within the cur-

rent segmental interpretation of the brain (Fig. 9), because

this approach has been followed for representatives of

many vertebrate classes and allows a direct comparison of

topological relationships between homologous ChAT-ir

groups across vertebrates (Medina et al. 1993; Medina

and Reiner 1994; Marın et al. 1997; Anadon et al. 2000;

Pombal et al. 2001; Gonzalez et al. 2002a; Mueller et al.

2004).

Localization of ChAT-ir elements in the forebrain

of lungfishes: comparative aspects

The large and histologically well-differentiated olfactory

bulbs in lungfishes were devoid of ChAT-ir cells in the

species studied, and only some fibers were detected in the

mitral and internal plexiform layers, exclusively in Neo-

ceratodus. Among other vertebrates, cholinergic fibers

have been reported in the olfactory bulb only in zebrafish

(Edwards et al. 2007), in anuran and urodele amphibians

(Marın et al. 1997), and in macaque monkeys (Porteros

et al. 2007). The absence of cholinergic cells in the

olfactory bulbs seems to be a shared character among

vertebrates, with the exception of the lesser spotted dog-

fish, where there is one ChAT-ir cell population around the

olfactory glomeruli and another in the granular layer of the

olfactory bulb (Anadon et al. 2000). The presence of these

ChAT-ir populations appears to be a derived feature for at

least this family of elasmobranchs.

No ChAT-ir cells were observed in the pallial regions in

either lungfish studied. Similarly, cholinergic cells have not

been detected in the pallium of any anamniote studied with

the exception of the dorsal pallium of the lesser spotted

dogfish (Anadon et al. 2000) and the dorsal part of the

dorsal telencephalic area (pallium) of the rainbow trout and

brown trout (Perez et al. 2000). With the exception of the

lizard Gallotia (Medina et al. 1993), no ChAT-ir cells have

been found in the cortex of birds and reptiles (Mufson et al.

1984; Brauth et al. 1985; Hoogland and Vermeulen-Van

der Zee 1990; Powers and Reiner 1993; Medina and Reiner

1994). In mammals, cholinergic cells have been detected in

the cortex of rats (Houser et al. 1983; Ichikawa and Hirata

1986; Parnavelas et al. 1986; Ichikawa et al. 1997) and

mice (Mufson and Cunningham 1988; Consonni et al.

2009) but not dogs, cats, guinea pigs, other rodents,

monotremes (Kimura et al. 1981; Vincent and Reiner 1987;

Maley et al. 1988; St-Jacques et al. 1996; Manger et al.

2002; Bhagwandin et al. 2006) or adult primates including

humans (Mesulam et al. 1984; Satoh and Fibiger 1985;

Mesulam and Geula 1988; Geula et al. 1993; Alonso and

Amaral 1995). Therefore, the presence of cholinergic cells

in the pallium/cortex of vertebrates is is most likely not a

primitive feature for vertebrates.

Only a moderate plexus of ChAT-ir fibers was observed

in the medial pallium of the lungfishes studied. This

innervation is similar to that observed in amphibians (Marın

et al. 1997; Gonzalez et al. 2002a) and likely arises from the

septal cholinergic cells (present results; Northcutt and

Westhoff 2011), as was observed in anuran amphibians,


123

Table 3 Summary of the distribution of ChAT-ir cells in different nuclei of the central nervous system of the major vertebrate groups

Lampreys Elasmobranchs Chondrosteans Teleosts Lung

fishes

Amphibians Reptiles Birds Mammals

Forebrain

Olfactory bulb - ? - - - - - - -

Pallium/cortex - ? - ± - - ± – ±

Striatum ? - - - ? ± ? ? ?

Basal telencephalon - - - ± ? ? ? ? ?

Magnocellular preoptic

nucleus

? ? ? ? ? ? - - -

Hypothalamus ? ? ? ? ± ? ? ? ±

Habenula ? ? ? ? ? ? ? ? ?

Thalamus - - ? ? ? - - ? ±

Pretectum ? ? - ± ± - - ? -

Brainstem

Optic tectum 2 2 2 1 2 – 2 1 –Isthmic/parabigeminal

Nucleus

? ?? ? ? ? ? ? ? ±

Cerebellum ? - ± - - - - ±

Non-motor nuclei in the upper

rhombencephalon: LDT/PPT/

SRN

? ?? ? ? ? ? ? ? ?

Reticular formation ? ? - ? ? ? ? ? ?

Octaval (octavolatel) area - ? ? ± ? ? ? ? ?

Cranial nerve motor nuclei ? ? ? ? ? ? ? ? ?

Spinal cord

Spinal motor column ? ? ? ? ? ? ? ? ?

?, Presence of ChAT-ir cells; -, absence of ChAT-ir cells; ±, presence of ChAT-ir cells only in some species of the group; ??, A clear

equivalence between cholinergic groups has not established

Lampreys, Pombal et al. (1999, 2001); elasmobranchs, Anadon et al. (2000); chondrosteans, Adrio et al. (2000); teleosts, Ekstrom (1987);

Brantley and Bass (1988); Molist et al. (1993); Perez et al. (2000); Clemente et al. (2004); Mueller et al. (2004); Giraldez-Perez et al. (2009);

amphibians, Marın et al. (1997); Gonzalez et al. (2002a); reptiles, Mufson et al. (1984); Brauth et al. (1985); Hoogland and Vermeulen-Van der

Zee (1990); Medina et al. 1993; Powers and Reiner (1993); birds, Sorenson et al. (1989); Medina and Reiner (1994); mammals, Kimura et al.

(1981); Mesulam et al. (1984); Houser et al. 1985; Satoh and Fibiger (1985); Vincent and Reiner (1987); Maley et al. (1988); Mufson and

Cunningham (1988); Tago et al. (1989); St-Jacques et al. (1996); Ichikawa et al. (1997); Manger et al. (2002); Varga et al. (2003); Motts et al.

(2008); Gravett et al. (2009)

Fig. 9 Schematic drawing summarizing the distribution of the main cholinergic cell groups in Neoceratodus forsteri according to a segmental

interpretation of the brain


123

where a forerunner of the septohippocampal system was

demonstrated (Gonzalez and Lopez 2002).

In the basal portion of the telencephalon (subpallium) a

population of numerous ChAT-ir cells has been observed in

lungfishes. These cells are distributed over a wide region,

including the caudal lateral septum, the caudoventral por-

tion of the striatum, and, principally, in areas close to the

recently characterized bed nucleus of the stria terminalis

and the pallidum based on their expression of the tran-

scription factor Nkx2.1 (Gonzalez and Northcutt 2009;

Northcutt 2009). This cell population represents the basal

forebrain cholinergic system (BFCS), which in mammals

forms a longitudinal column of neurons distributed in

association with the medial septum, vertical and horizontal

limbs of the diagonal band of Broca, ventral pallidum,

preoptic area, substantia innominata, and magnocellular

basal nucleus (Semba 2004). The BFCS in birds and rep-

tiles resembles that in mammals in terms of location and

cortical projections (Mufson et al. 1984; Brauth et al. 1985;

Hoogland and Vermeulen-Van der Zee 1990; Medina et al.

1993; Powers and Reiner 1993; Medina and Reiner 1994).

Observations in amphibians suggest that there are cholin-

ergic neurons in those regions of the basal telencephalon

that correspond to the location of the BFCS in mammals,

and that these neurons project to the medial pallium (Marın

et al. 1997, Gonzalez and Lopez 2002; Sanchez-Camacho

et al. 2006). Surprisingly, the BFCS of lungfishes is formed

by a higher population of cells than in anurans and urodeles

and it resembles more closely the situation found in

gymnophionans (Gonzalez et al. 2002a). Furthermore, in

teleosts cholinergic neurons in ventral telencephalic

regions are considered to be homologous to the BFCS in

tetrapods (Brantley and Bass 1988; Perez et al. 2000;

Kaslin et al. 2004; Mueller et al. 2004) and, in particular,

the cholinergic cells located in the lateral nucleus of area

ventralis were compared to the nucleus basalis of Meynert

(Mueller et al. 2004). In contrast, in sturgeons, dogfishes

and lampreys, the basal telencephalon is devoid of cho-

linergic neurons (Adrio et al. 2000; Anadon et al. 2000;

Pombal et al. 2001). Therefore, it was suggested that the

ancestors of modern teleosts likely had a BFCS (Semba

2004) but, since only advanced teleosts have been exam-

ined for ChAT expression, and the status in the basal act-

inopterygians is unknown, it cannot be concluded that a

BFCS first evolved in teleosts.

The striatal cholinergic cells observed in lungfishes

deserve special mention. In general, no ChAT-ir cells have

been seen in the striatum of fishes (Ekstrom 1987; Brantley

and Bass 1988; Molist et al. 1993; Adrio et al. 2000;

Anadon et al. 2000; Perez et al. 2000; Mueller et al. 2004;

Giraldez-Perez et al. 2009), the exception being those

reported in lampreys, in the region homologous to the

striatum (Pombal et al. 1997, 2001). In contrast, the

presence of cholinergic cells in the striatum of amniotes is

a very conservative feature (Marın et al. 1998; Reiner et al.

1998). In mammals, these cholinergic cells are interneu-

rons of local circuits, with cell bodies larger than the

projection neurons (Kasa 1986; Woolf 1991), and similar

observations have been reported in reptiles and birds

(Medina et al. 1993; Henselmans and Wouterlood 1994;

Medina and Reiner 1994). Striatal cholinergic neurons are

also present in some anuran amphibians (Marın et al. 1997)

and, especially, in gymnophionan amphibians (Gonzalez

et al. 2002a). Cholinergic cells do not originate in striatal

regions during development but reach this location through

tangential migration from their place of origin in the cau-

domedial telencephalon (Marın and Rubenstein 2003).

Interestingly, a similar region in the caudal telencephalon

of lungfishes has been recently characterized molecularly

(Gonzalez and Northcutt 2009), and tangential migration

toward the striatum was suggested to occur as in amphib-

ians (Moreno et al. 2008).

In the preoptic region, a few faintly labeled ChAT-ir

cells were seen in the magnocellular preoptic nucleus of

lungfishes. These large neurons are probably neurosecre-

tory cells projecting to the neural lobe of the hypophysis, as

has been observed in the magnocellular preoptic nucleus

and some neurons of the hypothalamic tuberal nucleus in

all anamniotes, from lampreys to amphibians (Marın et al.

1997; Adrio et al. 2000; Anadon et al. 2000; Perez et al.

2000; Pombal et al. 2001; Gonzalez et al. 2002a; Rodrı-

guez-Moldes et al. 2002; Mueller et al. 2004). This feature

is in contrast to the situation in amniotes, where cholinergic

cells are absent from the neurosecretory nuclei in the pre-

optic region, although ChAT-ir cells have been described

among or adjacent to the neurosecretory cells of the

supraoptic nucleus (Mason et al. 1983; Tago et al. 1987;

Medina et al. 1993; Powers and Reiner 1993; Medina and

Reiner 1994; Ichikawa et al. 1997).

No other ChAT-ir cells were seen in the hypothalamus

of Protopterus, whereas a prominent group of cholinergic

cells was observed in the suprachiasmatic nucleus of

Neoceratodus, as were two additional groups in the dorsal

part of the nucleus of the periventricular organ and the

intermediate tuberal nucleus. In other groups of fishes,

cholinergic cells have been observed in the paraventricular

nucleus of lampreys (Pombal et al. 2001) and the tuberal

hypothalamus of sharks, sturgeons and teleosts (Ekstrom

1987; Adrio et al. 2000; Anadon et al. 2000; Perez et al.

2000; Mueller et al. 2004). In anurans and urodeles, ChAT-

ir cells were detected in the suprachiasmatic region (Marın

et al. 1997) and, additionally, in the infundibular hypo-

thalamus in Xenopus laevis and in the gymnophionan

Dermophis mexicanus (Marın et al. 1997; Gonzalez et al.

2002a). Cholinergic neurons have been described in the

infundibular hypothalamus and in the suprachiasmatic


123

region in most amniotes (Mason et al. 1983; Tago et al.

1987; Medina et al. 1993; Powers and Reiner 1993; Medina

and Reiner 1994; Ichikawa et al. 1997; Gravett et al. 2009),

with the exception of monotremes where no ChAT-ir cells

were reported in the hypothalamus (Manger et al. 2002).

Consequently, the existence of cholinergic hypothalamic

cells in Neoceratodus can be considered a primitive and

conserved feature of the vertebrate cholinergic systems,

which was secondarily lost in Protopterus as in other

groups.

Within the epithalamus, faintly labeled cells were seen

in the habenula in the two lungfish species, and the course

of the habenulo-interpeduncular tract (fasciculus retro-

flexus) was clearly visible across the diencephalon and up

to the interpeduncular neuropil. This feature has been

reported in all amniotes studied (Houser et al. 1983;

Mesulam et al. 1984; Satoh and Fibiger 1985; Vincent and

Reiner 1987; Maley et al. 1988; Sorenson et al. 1989; Tago

et al. 1989; Medina et al. 1993; Powers and Reiner 1993;

Medina and Reiner 1994; St-Jacques et al. 1996; Ichikawa

et al. 1997; Gravett et al. 2009), as well as in all anamniotes

studied (Marın et al. 1997; Adrio et al. 2000; Anadon et al.

2000; Perez et al. 2000; Pombal et al. 2001; Gonzalez et al.

2002a; Rodrıguez-Moldes et al. 2002; Mueller et al. 2004;

Giraldez-Perez et al. 2009). This confirms the importance

of the cholinergic nature in this part of the dorsal dience-

phalic conduction system, a highly conserved pathway that

is likely a primitive feature for vertebrates (Bianco and

Wilson 2009).

The thalamus (formerly termed the dorsal thalamus;

Puelles and Rubenstein 2003) of lungfishes possess a

prominent group of ChAT-ir cells (more reduced in Neo-

ceratodus), and numerous ChAT-ir fibers are also distrib-

uted in the thalamus and prethalamus (formerly termed the

ventral thalamus). Cholinergic cells in the thalamus have

been reported in chondrostean and teleost fishes (Adrio

et al. 2000; Perez et al. 2000; Rodrıguez-Moldes et al.

2002; Mueller et al. 2004; Giraldez-Perez et al. 2009), but

not in lampreys or elasmobranches (Anadon et al. 2000;

Pombal et al. 2001), amphibians (Marın et al. 1997; Gon-

zalez et al. 2002a), or reptiles (Medina et al. 1993; Powers

and Reiner 1993). In addition, ChAT-ir cells have been

described in the thalamus of birds (Medina and Reiner

1994) and some species of mammals (Rico and Cavada

1998; Gravett et al. 2009). These data indicate that the

presence of cholinergic cells in the thalamus of vertebrates

is a variable character, and that it has likely appeared

several times during the evolution. Similarly, another such

trait is the presence of cholinergic cells in the pretectum.

Protopterus, like chondrosteans, amphibians, reptiles and

mammals does not contain ChAT-ir cells in this brain

structure (Kasa 1986; Woolf 1991; Medina et al. 1993;

Powers and Reiner 1993; Marın et al. 1997; Adrio et al.

2000; Gonzalez et al. 2002a; present study), whereas

Neoceratodus, lampreys, elasmobranches, and some tele-

osts and birds do (Ekstrom 1987; Sorenson et al. 1989;

Medina and Reiner 1994; Anadon et al. 2000; Perez et al.

2000; Pombal et al. 2001; present results). In the thalamus

and pretectum of lungfishes, the localization of ChAT-ir

fibers, which were more abundant in Neoceratodus,

matches those regions identified as centers of primary

retinofugal projections (Northcutt 1980), suggesting a

cholinergic influence on these retinorecipient areas. This

localization has also been described in amphibians (Marın

et al. 1997; Gonzalez et al. 2002a) and in some amniotes

(Woolf 1991; Medina and Smeets 1992; Medina et al.

1993; Medina and Reiner 1994).

Localization of ChAT-ir elements in the brainstem

of lungfishes: comparative aspects

The optic tectum of lungfishes does not contain cholinergic

neurons, and only ChAT-ir fibers and terminals were seen,

mainly in the superficial fiber layers. The absence of tectal

cholinergic cells is shared by lampreys, elasmobranch and

chondrostean fishes (Adrio et al. 2000; Anadon et al. 2000;

Pombal et al. 2001), anuran and urodele amphibians (De-

san et al. 1987; Marın et al. 1997), reptiles (Brauth et al.

1985; Medina et al. 1993; Powers and Reiner 1993) and

most mammalian species (Mesulam et al. 1984; Satoh and

Fibiger 1985; Mizukawa et al. 1986; Maley et al. 1988;

Gravett et al. 2009). In contrast, tectal ChAT-ir cells are

reported to be abundant in all teleosts studied (Tumosa

et al. 1986; Ekstrom 1987; Zottoli et al. 1987; Brantley and

Bass 1988; Molist et al. 1993; Perez et al. 2000; Mueller

et al. 2004; Giraldez-Perez et al. 2009) as well as in

gymnophionan amphibians (Gonzalez et al. 2002a), birds

(Sorenson et al. 1989; Medina and Reiner 1994), and in the

superior colliculus of some mammals (Vincent and Reiner

1987; Tago et al. 1989; Motts et al. 2008). These results

suggest that the presence of cholinergic cells in the tectum

is a variable feature, which has arisen several times during

evolution.

The optic tectum of lungfishes is moderately or densely

innervated by cholinergic fibers. Similar variability has

been described in amphibians (Marın et al. 1997; Gonzalez

et al. 2002a), and the source of this innervation could be in

the cholinergic groups of the upper rhombencephalon,

especially the isthmic nucleus, as has been demonstrated in

amphibians (Marın and Gonzalez 1999). Therefore, ace-

tylcholine may influence retinal afferents by modulating

synaptic function in the optic tectum (Sargent et al. 1989;

King 1990; King and Schmidt 1991).

In the isthmic region of the two lungfishes studied, a

group of small ChAT-ir cells was interpreted as the nucleus

isthmi. In other nonmammalian vertebrates studied, a


123

nucleus isthmi has been characterized by the cholinergic

nature of its cells and by its reciprocal connections with the

optic tectum (chondrosteans, Adrio et al. 2000; teleosts,

Ekstrom 1987; Brantley and Bass 1988; Zottoli et al. 1988;

Perez et al. 2000; Mueller et al. 2004; Pushchina and

Karpenko 2007; Giraldez-Perez et al. 2009; amphibians,

Marın et al. 1997; Wiggers 1998; Marın and Gonzalez

1999; reptiles, Brauth et al. 1985; Medina and Smeets

1992; Medina et al. 1993; Powers and Reiner 1993; birds,

Sorenson et al. 1989; Bagnoli et al. 1992; Medina and

Reiner 1994); a putative cholinergic group in lampreys has

been proposed as the homologue of the nucleus isthmi in

gnathostomes (Pombal et al. 2001). This nucleus has also

been proposed as the homologue of the parabigeminal

nucleus in mammals, which contains cholinergic cells and

projects to the superior colliculus (Beninato and Spencer

1986; Mufson et al. 1986; Vincent and Reiner 1987; Tago

et al. 1989; Woolf 1991; Gravett et al. 2009). The excep-

tions among mammals are monotremes, the laboratory

shrew, and microbats, where no ChAT-ir cells were seen in

this nucleus (Manger et al. 2002; Maseko and Manger

2007). Even with these exceptions, the existence of a

cholinergic isthmic/parabigeminal nucleus appears to be a

primitive characteristic in the brain of vertebrates, which

has been generally conserved during the evolution.

Medial to the cholinergic isthmic nucleus, some ChAT-

ir cells were seen close to the ventricular surface in the

lungfishes in this study. These cells could be homologous

to the secondary gustatory (visceral) nucleus described in

teleosts (Zottoli et al. 1988; Molist et al. 1993; Perez et al.

2000; Mueller et al. 2004; Giraldez-Perez et al. 2009). This

nucleus receives projections from the general viscerosen-

sory region of the medulla (Finger and Kanwal 1992). An

ascending secondary visceral tract could be traced rostrally

from the nucleus of the solitary tract and may terminate in

a secondary gustatory nucleus identified in Lepidosiren

(Nieuwenhuys 1998) and Neoceratodus (Holmgren and

van der Horst 1925), although more specific hodological

studies are needed to resolve the identification of this

isthmic cholinergic group in lungfishes.

Ventrocaudal to the nucleus isthmi, a conspicuous group

of intensely labeled ChAT-ir cells had processes that were

seen to course both rostrally and caudally in the tegmentum

of lungfishes. This group was similar in localization and

cholinergic nature to the laterodorsal tegmental nucleus

defined in amphibians (Marın et al. 1997; Gonzalez et al.

2002a), and its homology is further supported by experi-

ments demonstrating that these cells produce nitric oxide

(unpublished own observations), as has been reported in

amphibians (Gonzalez et al. 1996; Munoz et al. 1996;

Gonzalez et al. 2002b) and amniotes (Alonso et al. 2000).

In the lungfishes studied, an additional cholinergic group

potentially homologous to the pedunculopontine tegmental

nucleus was not identified. This is in line with data from

other anamniotes, with the exception of anuran amphibians

(Marın et al. 1997). In teleosts, the cholinergic cells of the

superior reticular nucleus might be homologous to the

amniote pedunculopontine/laterodorsal tegmental system

in amniotes, based on connections to the optic tectum

(Grover and Sharma 1981; Perez et al. 2000), or to the

subpallium (Rink and Wullimann 2002, 2004; Wullimann

and Rink 2002), as has been proposed in amphibians

(Marın and Gonzalez 1999; Sanchez-Camacho et al. 2006)

and mammals (Woolf and Butcher 1986; Woolf et al.

1990). However, it is not clear if this single cell group in

anamniotes represents the pedunculopontine nucleus or the

laterodorsal tegmental nucleus (or both) in amniotes. Tract

tracing studies in lungfishes are needed to clarify hod-

ological features of the cholinergic cell groups in the upper

rhombencephalon before proposing further homologies.

The cerebellum of lungfishes is completely devoid of

ChAT-ir cells, and only some immunoreactive fibers were

seen in Neoceratodus. This is the situation in the majority

of vertebrate groups studied, with the exception of elas-

mobranchs (Anadon et al. 2000) and some species of tel-

eosts (Brantley and Bass 1988; Giraldez-Perez et al. 2009)

and mammals (Ikeda et al. 1991). Therefore, the presence

of ChAT-ir cells in the cerebellum is a peculiar feature

limited to a few species of vertebrates and is likely derived

in these taxa, rather than being primitive for vertebrates.

Moderate cholinergic innervation has been described in the

cerebellum of amphibian, with the possible sources being

the inferior reticular nucleus, the octaval area, and the

dorsal column nucleus (Marın et al. 1997).

The presence of ChAT-ir cells in the reticular formation

of lungfishes is especially marked in Neoceratodus. These

reticular cholinergic cells have been reported in all other

vertebrates studied, with the exception of chondrosteans

(Adrio et al. 2000) and many of these cells were assumed

to be reticulospinal neurons (Anadon et al. 2000; Perez

et al. 2000). Among the reticular cells, the Mauthner cells

were not ChAT-ir in the two lungfishes studied, as is also

the case in teleosts (Ekstrom 1987; Brantley and Bass

1988; Molist et al. 1993; Perez et al. 2000; Mueller et al.

2004; Giraldez-Perez et al. 2009) and amphibians (Marın

et al. 1997; Gonzalez et al. 2002a) with the exceptions of a

urodele, Pleurodeles waltl (Marın et al. 1997), and an

anuran, Xenopus laevis, during development (Lopez et al.

2002). A peculiarity observed in Neoceratodus, but not in

Protopterus, was the presence of the small group of

ChAT-ir cells in the superior raphe nucleus surrounding

the medial longitudinal fascicle dorsally. This cell group

has also been observed in some teleosts, such as two

species of trout (Perez et al. 2000), but not others as

zebrafish (Mueller et al. 2004) and goldfish (Giraldez-

Perez et al. 2009).


123

Cells immunoreactive for ChAT were present in the

medial octavolateral nucleus of the two lungfishes studied.

This nucleus receives acoustic and vestibular information

and is the main target of projections from the mechanore-

ceptive lateral line nerves (Nieuwenhuys 1998; Northcutt

2011). Cholinergic cells were also detected in cells of the

octavolateral system of elasmobranchs (Anadon et al.

2000), chondrosteans (Adrio et al. 2000), some species of

teleosts (Perez et al. 2000; Clemente et al. 2004; Mueller

et al. 2004; Giraldez-Perez et al. 2009), amphibians

(Gonzalez et al. 1993; Marın et al. 1997; Gonzalez et al.

2002a), and amniotes (reptiles, Medina et al. 1993; Powers

and Reiner 1993; birds, Medina and Reiner 1994; mam-

mals, Satoh and Fibiger 1985; Vincent and Reiner 1987;

Maley et al. 1988; Tago et al. 1989; Ichikawa et al. 1997;

Motts et al. 2008).

Two small groups of cholinergic cells were seen arranged

around the solitary tract in the lungfishes studied. These

populations likely represent the nucleus of the solitary tract

and the dorsal column nucleus, respectively, as in amphibi-

ans (Munoz et al. 1995; Marın et al. 1997; Gonzalez et al.

2002a) and other vertebrates, such as lampreys (Pombal et al.

2001) and birds and mammals (Armstrong et al. 1988; Tago

et al. 1989; Ruggiero et al. 1990; Medina and Reiner 1994;

Lan et al. 1995; Motts et al. 2008), whereas the homologous

nuclei in reptiles do not appear to contain cholinergic neu-

rons (Medina et al. 1993; Powers and Reiner 1993).

In closing, we will focus on the hindbrain the motor

nuclei that innervate the muscles of the head and upper part

of the trunk, as they showed very intense ChAT immuno-

reactivity. The organization of the ocular motor nuclei

observed in the present study was consistent with that

revealed by tract tracing techniques in the same lepido-

sirenid species, Protopterus dolloi (von Bartheld 1992). In

that study, three groups of oculomotor neurons were

identified, based on the muscles innervated and whether

this innervation was ipsilateral or contralateral. In our

study, ChAT immunohistochemistry did not clearly dif-

ferentiate three oculomotor subdivisions, although both

ipsilaterally projecting neurons and contralaterally pro-

jecting neurons were observed. We also corroborated in

both species the presence of a small trochlear nucleus in the

isthmic tegmentum, just caudal to the strongly catechol-

aminergic cell group in the mesencephalon and rostral to

the interpeduncular nucleus, which was highlighted by our

double labeling techniques. Caudal to the facial nerve root,

and medial in the basal plate, we observed a small abdu-

cens motor nucleus, slightly caudal to the Mauthner cells,

as described by von Bartheld (1992), and the ChAT

immunohistochemistry did not evidence an accessory

abducens nucleus. In general, we observed a better devel-

oped organization of the ocular motor nuclei in Neocerat-

odus than in Protopterus, as might be expected from the

fact that Neoceratodus has relatively well-developed eyes,

in contrast to lepidosirenid lungfishes where the eyes are

more reduced.

The motor system comprises neurons that are intensely

ChAT-ir in the branchiomeric motor centers of the tri-

geminal, facial, glossopharyngeal, and vagal nerves (clas-

sically considered the special visceral motor nuclei), which

innervate the jaw musculature and muscles of the brachial

arches. The relatively large number of trigeminal motor

neurons in lungfishes no doubt reflects their well-developed

jaw musculature. The motoneurons of the parasympathetic

nervous system, associated with the facial, glossopharyn-

geal and vagal motoneurons, are difficult to identify in

lungfishes, although the small cholinergic cells located at

the lateral aspect of these nuclei, more obvious in Neo-

ceratodus than in Protopterus, might represent the visceral

motor component of the autonomic nervous system, as

reported in amphibians (Marın et al. 1997). Finally, close

to the obex and extending into the spinal cord, the large

spino-occipitalis somatic motoneurons that innervate hyp-

obrachial muscles of the head to open the lower jaw,

elevate the floor of the mouth, and extend the gills

(Nieuwenhuys 1998), form the most intense ChATir cell

group in the caudal rhombencephalon.

The somatomotor neurons in the ventral horn of the

spinal cord in both species studied were strongly ChAT-ir

and were seen to form a long column throughout the ro-

strocaudal extent of this structure. Similar observations

have been reported in all vertebrates studied. Brachial and

lumbar enlargements were not found in our lungfishes and

lateral subgroups of motoneurons were never observed in

the ventral horn. Smaller ChAT-ir cells were present in the

intermediate gray in the middle levels of the spinal cord

and may correspond to the sympathetic cells described in

amphibians (Munoz et al. 2000; Gonzalez et al. 2002a), but

tracing studies are needed to confirm this.

Segmental organization of cholinergic cell groups

in the brain of lungfishes

A segmental approach to comparative neuroanatomy is

useful in revealing topological relationships within verte-

brate brains, especially when comparing neurotransmitter

systems (Smeets and Gonzalez 2000). The segmental par-

adigm is based on numerous studies that demonstrate a

segmental pattern of organization in vertebrate brains

(Lumsden and Keynes 1989; Bulfone et al. 1993; Figdor

and Stern 1993; Puelles and Rubenstein 1993; Puelles and

Rubenstein 2003 Rubenstein et al. 1994; Puelles et al.

1996). The number and organization of the neuromeres

(brain segments) are therefore a constant characteristic in

all vertebrates (Lumsden and Keynes 1989; Puelles 1995).

This model proposes that a developing vertebrate brain


123

consists of nine rhombomeres in the hindbrain (r0–r8;

Gilland and Baker 1993; Marın and Puelles 1995; Fritzsch

1998; Cambronero and Puelles 2000; Aroca and Puelles

2005; Straka et al. 2006), whereas the midbrain consists of

only one mesomere (Diaz et al. 2000). The forebrain is

formed by the diencephalon, which has three prosomeres

(p1–p3) and the secondary prosencephalon (telencephalon

plus hypothalamus), which is not subdivided into proso-

meres (Puelles and Rubenstein 2003).

The cholinergic systems of lampreys, elasmobranchs,

amphibians, reptiles, and birds have all been analyzed within

the context of this segmental model (Medina et al. 1993;

Medina and Reiner 1994; Marın et al. 1997; Anadon et al.

2000; Pombal et al. 2001; Gonzalez et al. 2002a). We will

now summarize our results following a similar approach in

order to clarify comparisons among vertebrate groups.

Secondary prosencephalon

The septal, striatal, and basal cholinergic cells are located

in the telencephalon and develop from the evaginated

secondary prosencephalon, the rostralmost region of the

brain. The ChAT-ir cells in the anterior preoptic area, the

neurosecretory magnocellular nucleus, the suprachiasmatic

nucleus, the nucleus of the periventricular organ and the

tuberal region (the last three present only in Neoceratodus)

belong to the nonevaginated secondary prosencephalon.

All these populations are situated in the alar plate of this

protosegment, except for the nucleus of the periventricular

organ and the intermediate tuberal nucleus in Neocerato-

dus, which are located in the basal plate.

Diencephalon

Only two cholinergic cell groups in Protopterus and three

in Neoceratodus were found to be diencephalic. The first

group is situated dorsally in p2 and corresponds to the

habenula; the second one is also present in the alar plate of

p2 and corresponds to the cholinergic population of the

thalamus. The third group, present only in Neoceratodus, is

located in the alar plate of p3 and corresponds to the small

cholinergic cell group in the pretectum.

Mesencephalon

Only one population of cholinergic cells was found in the

basal plate of the single mesencephalic neuromere that

corresponds to the oculomotor nucleus.

Isthmus

The isthmic region comprises a single curved segment, named

rhombomere 0 (r0). In the alar plate are located the cholinergic

populations of the isthmic and the possible secondary gusta-

tory (visceral) nuclei. The basal plate of this segment contains

the cholinergic cells of the trochlear nucleus.

Hindbrain

This region is one of the most conserved in vertebrate

brains. It contains most of the cholinergic groups of the

motor nuclei and also nonmotor cell components. In the

rostral alar plate of the rhombomere 1, there is the con-

spicuous ChAT-ir group of the laterodorsal tegmental

nucleus. In addition, in Neoceratodus only, the caudal basal

plate of r1 and the rostral part of r2, house the pretrige-

minal cholinergic cells of the superior reticular nucleus and

the ChAT-ir cells of the superior raphe area. Rhombomeres

2 and 3 contain the large trigeminal motor nucleus. The

boundary between r3 and r4 is marked by the exit of axons

of the facial nerve. The large Mauthner cells are contained

in r4, which lacks motor neurons (except for a few cells

representing a rostral component of the facial nucleus).

This situation was also described in gymnophionans

(Gonzalez et al. 2002a) but not in lampreys, anuran

amphibians, and birds where the facial motor nucleus lies

in r4, just in front of the nerve entrance (Medina and Reiner

1994; Marın et al. 1997; Pombal et al. 2001). In lungfishes,

as in gymnophionans (Gonzalez et al. 2002a), the facial

motoneurons are located caudally in r6, and their axons

cross r5 into r4 above the medial longitudinal fascicle, then

bend laterally (genus facialis) to exit the brain in r4, as in

all vertebrates. In elasmobranchs, reptiles, and mammals,

the facial motoneurons also migrate caudally into r6 during

development (Medina et al. 1993; Anadon et al. 2000).

Therefore, both migrated and non-migrated components of

the facial motor nucleus are represented among vertebrate

classes or even within a single class, as described in

amphibians (Marın et al. 1997; Gonzalez et al. 2002a).

The dispersed motoneurons of the abducens nucleus are

located in rhombomere 5, as in mammals, whereas in

reptiles and birds this nucleus spans r5 and r6 (Medina

et al. 1993; Medina and Reiner 1994). The situation

observed in lampreys is very special, as the abducens

motoneurons extend through r4 to r6, and their axons exit

the brain in r2 (Pombal et al. 2001). The cholinergic cells

of the median reticular nucleus observed in Neoceratodus

lie in rhombomeres 5 and 6.

The glossopharyngeal motoneurons form a continuous

column with the facial motoneurons, with some overlap,

extend into rhombomere 7; they are then continuous with

the vagal motoneurons, which occupy rhombomere 8 and

the rostral extension of the spinal cord. The spino-occipital

motor nucleus also lies in rhombomere 8 and forms a

continuous column with the spinal motoneurons. The small

cholinergic cells observed in the inferior reticular nucleus,


123

between the vagal and spino-occipitalis motoneurons also

lie in r8. Finally, the non-motor cholinergic cells of the

nucleus of the solitary tract and dorsal column nucleus are

located in the alar plate of the rhombomeres 7 and 8.

Conclusion

The distribution of the cholinergic systems in the brain of

the two lungfishes studied shows many features shared with

tetrapods, especially amphibians. Our work does not sup-

port alternative views that considered lungfishes equally

related to actinopterygians and crossopterygians; rather it

provides substantial anatomical evidence to corroborate the

idea that living lungfishes are close relatives to tetrapods.

Acknowledgments We thank Mary Sue Northcutt for help with

many phases of the research and manuscript preparation. This

research was supported by the Spanish Ministry of Science and

Technology (Grant number: BFU2006-01014/BFI), by the US

National Science Foundation (IBN-0919077), and private funding.

Conflict of interest The authors declare that they have no conflict

of interest.

References

Adams JC (1981) Heavy metal intensification of DAB-based HRP

reaction product. J Histochem Cytochem 29:775

Adrio F, Anadon R, Rodrıguez-Moldes I (2000) Distribution of

choline acetyltransferase (ChAT) immunoreactivity in the cen-

tral nervous system of a chondrostean, the siberian sturgeon

(Acipenser baeri). J Comp Neurol 426:602–621

Alonso JR, Amaral DG (1995) Cholinergic innervation of the primate

hippocampal formation. I. Distribution of choline acetyltrans-

ferase immunoreactivity in the Macaca fascicularis and Macacamulatta monkeys. J Comp Neurol 355:135–170

Alonso JR, Arevalo R, Wereuaga E, Porteros A, Brinon JG, Aijon J

(2000) Comparative and developmental neuroanatomical aspects

of the NO system. In: Steinbusch HWM, De Vente J, Vincent SR

(eds) Functional neuroanatomy of the nitric oxide system.

Elsevier, Amsterdam, pp 51–109

Anadon R, Molist P, Rodrıguez-Moldes I, Lopez JM, Quintela I,

Cervino MC, Barja P, Gonzalez A (2000) Distribution of choline

acetyltransferase immunoreactivity in the brain of an elasmo-

branch, the lesser spotted dogfish (Scyliorhinus canicula).

J Comp Neurol 420:139–170

Armstrong DM, Rotler A, Hersh LB, Pickel VM (1988) Localization

of choline acetyltransferase in perikarya and dendrites within the

nuclei of the solitary tracts. J Neurosci Res 20:279–290

Aroca P, Puelles L (2005) Postulated boundaries and differential fate

in the developing rostral hindbrain. Brain Res Brain Res Rev

49:179–190

Bagnoli P, Fontanesi G, Alesci R, Erichsen JT (1992) Distribution of

neuropeptide Y, substance P, and choline acetyltransferase in the

developing visual system of the pigeon and effects of unilateral

retina removal. J Comp Neurol 318:392–414

Beninato M, Spencer RF (1986) A cholinergic projection to the rat

superior colliculus demonstrated by retrograde transport of

horseradish peroxidase and choline acetyltransferase immuno-

histochemistry. J Comp Neurol 253:525–538

Bhagwandin A, Fuxe K, Manger PR (2006) Choline acetyltransferase

immunoreactive cortical interneurons do not occur in all rodents:

a study of the phylogenetic occurrence of this neural character-

istic. J Chem Neuroanat 32:208–216

Bianco IH, Wilson SW (2009) The habenular nuclei: a conserved

asymmetric relay station in the vertebrate brain. Philos Trans R

Soc Lond B Biol Sci 364:1005–1020

Brantley RK, Bass AH (1988) Cholinergic neurons in the brain of a

teleost fish (Porichthys notatus) located with a monoclonal

antibody to choline acetyltransferase. J Comp Neurol 275:87–105

Brauth SE, Kitt CA, Price DL, Wainer BH (1985) Cholinergic

neurons in the telencephalon of the reptile Caiman crocodilus.

Neurosci Lett 58:235–240

Brinkmann H, Denk A, Zitzler J, Joss JJ, Meyer A (2004a) Complete

mitochondrial genome sequences of the South american and the

Australian lungfish: testing of the phylogenetic performance of

mitochondrial data sets for phylogenetic problems in tetrapod

relationships. J Mol Evol 59:834–848

Brinkmann H, Venkatesh B, Brenner S, Meyer A (2004b) Nuclear

protein-coding genes support lungfish and not the coelacanth as

the closest living relatives of land vertebrates. Proc Natl Acad

Sci USA 101:4900–4905

Bulfone A, Puelles L, Porteus MH, Frohman MA, Martin GR,

Rubenstein JL (1993) Spatially restricted expression of Dlx-1,

Dlx-2 (Tes-1), Gbx-2, and Wnt-3 in the embryonic day 12.5

mouse forebrain defines potential transverse and longitudinal

segmental boundaries. J Neurosci 13:3155–3172

Cambronero F, Puelles L (2000) Rostrocaudal nuclear relationships in

the avian medulla oblongata: a fate map with quail chick

chimeras. J Comp Neurol 427:522–545

Carroll RL (1988) Vertebrate paleontology and evolution. Freeman

Press, New York

Clemente D, Porteros A, Weruaga E, Alonso JR, Arenzana FJ, Aijon

J, Arevalo R (2004) Cholinergic elements in the zebrafish central

nervous system: Histochemical and immunohistochemical anal-

ysis. J Comp Neurol 474:75–107

Consonni S, Leone S, Becchetti A, Amadeo A (2009) Developmental

and neurochemical features of cholinergic neurons in the murine

cerebral cortex. BMC Neurosci 10:18

Crawford GD, Correa L, Salvaterra PM (1982) Interaction of

monoclonal antibodies with mammalian choline acetyltransfer-

ase. Proc Natl Acad Sci USA 79:7031–7035

Desan PH, Gruberg ER, Grewell KM, Eckenstein F (1987) Cholin-

ergic innervation of the optic tectum in the frog Rana pipiens.

Brain Res 413:344–349

Diaz C, Yanes C, Trujillo CM, Puelles L (2000) Cytoarchitectonic

subdivisions in the subtectal midbrain of the lizard Gallotiagalloti. J Neurocytol 29:569–593

Eckenstein F, Thoenen H (1983) Cholinergic neurons in the rat

cerebral cortex demonstrated by immunohistochemical localiza-

tion of choline acetyltransferase. Neurosci Lett 36:211–215

Edwards JG, Greig A, Sakata Y, Elkin D, Michel WC (2007)

Cholinergic innervation of the zebrafish olfactory bulb. J Comp

Neurol 504:631–645

Ekstrom P (1987) Distribution of choline acetyltransferase-immunor-

eactive neurons in the brain of a cyprinid teleost (Phoxinusphoxinus L.). J Comp Neurol 256:494–515

Figdor MC, Stern CD (1993) Segmental organization of embryonic

diencephalon. Nature 363:630–6344

Finger TE, Kanwal JS (1992) Ascending general visceral pathways

within the brainstems of two teleost fishes: Ictalurus punctatusand Carassius auratus. J Comp Neurol 320:509–520

Fritzsch B (1998) Of mice and genes: evolution of vertebrate brain

development. Brain Behav Evol 52:207–217


123

Geula C, Schatz CR, Mesulam MM (1993) Differential localization of

NADPH-diaphorase and calbindin-D28k within the cholinergic

neurons of the basal forebrain, striatum and brainstem in the rat,

monkey, baboon and human. Neuroscience 54:461–476

Gilland E, Baker R (1993) Conservation of neuroepithelial and

mesodermal segments in the embryonic vertebrate head. Acta

Anat (Basel) 148:110–123

Giraldez-Perez RM, Gaytan SP, Torres B, Pasaro R (2009) Co-

localization of nitric oxide synthase and choline acetyltransfer-

ase in the brain of the goldfish (Carassius auratus). J Chem

Neuroanat 37:1–17

Gonzalez A, Lopez JM (2002) A forerunner of septohippocampal

cholinergic system is present in amphibians. Neurosci Lett

327:111–114

Gonzalez A, Northcutt RG (2009) An immunohistochemical approach

to lungfish telencephalic organization. Brain Behav Evol

74:43–55

Gonzalez A, Meredith GE, Roberts BL (1993) Choline acetyltrans-

ferase immunoreactive neurons innervating labyrinthine and

lateral line sense organs in amphibians. J Comp Neurol

332:258–268

Gonzalez A, Munoz A, Munoz M, Marın O, Arevalo R, Porteros A,

Alonso JR (1996) Nitric oxide synthase in the brain of a urodele

amphibian (Pleurodeles waltl) and its relation to catecholamin-

ergic neuronal structures. Brain Res 727:49–64

Gonzalez A, Lopez JM, Sanchez-Camacho C, Marın O (2002a)

Localization of choline acetyltransferase (ChAT) immunoreac-

tivity in the brain of a caecilian amphibian, Dermophismexicanus (Amphibia: Gymnophiona). J Comp Neurol 448:

249–267

Gonzalez A, Moreno N, Lopez JM (2002b) Distribution of NADPH-

diaphorase/nitric oxide synthase in the brain of the caecilian

Dermophis mexicanus (amphibia: gymnophiona): comparative

aspects in amphibians. Brain Behav Evol 60:80–100

Gonzalez A, Morona R, Lopez JM, Moreno N, Northcutt RG (2010)

Lungfishes, like tetrapods, possess a vomeronasal system. Front

Neuroanat 4:130. doi: 10.3389/fnana.2010.00130

Gravett N, Bhagwandin A, Fuxe K, Manger PR (2009) Nuclear

organization and morphology of cholinergic, putative catechol-

aminergic and serotonergic neurons in the brain of the rock

hyrax, Procavia capensis. J Chem Neuroanat 38:57–74

Grover BG, Sharma SC (1981) Organization of extrinsic tectal

connections in Goldfish (Carassius auratus). J Comp Neurol

196:471–488

Hallstrom BM, Janke A (2009) Gnathostome phylogenomics utilizing

lungfish EST sequences. Mol Biol Evol 26:463–471

Hedges SB, Hass CA, Maxson LR (1993) Relations of fish and

tetrapods. Nature 363:501–502

Henselmans JM, Wouterlood FG (1994) Light and electron micro-

scopic characterization of cholinergic and dopaminergic struc-

tures in the striatal complex and the dorsal ventricular ridge of

the lizard Gekko gecko. J Comp Neurol 345:69–83

Holmgren N, van der Horst CJ (1925) Contribution to the morphology

of the brain of Ceratodus. Acta Zoologica 6:59–165

Hoogland PV, Vermeulen-Van der Zee E (1990) Distribution of

choline acetyltransferase immunoreactivity in the telencephalon

of the lizard Gekko gecko. Brain Behav Evol 36:378–390

Houser CR, Crawford GD, Barber RP, Salvaterra PM, Vaughn JE

(1983) Organization and morphological characteristics of cho-

linergic neurons: an immunocytochemical study with a mono-

clonal antibody to choline acetyltransferase. Brain Res 266:97–

119

Houser CR, Crawford GD, Salvaterra PM, Vaughn JE (1985)

Immunocytochemical localization of choline acetyltransferase

in rat cerebral cortex: a study of cholinergic neurons and

synapses. J Comp Neurol 234:17–34

Ichikawa T, Hirata Y (1986) Organization of choline acetyltransfer-

ase-containing structures in the forebrain of the rat. J Neurosci

6:281–292

Ichikawa T, Ajiki K, Matsuura J, Misawa H (1997) Localization of

two cholinergic markers, choline acetyltransferase and vesicular

acetylcholine transporter in the central nervous system of the rat:

in situ hybridization histochemistry and immunohistochemistry.

J Chem Neuroanat 13:23–39

Ikeda M, Houtani T, Ueyama T, Sugimoto T (1991) Choline

acetyltransferase immunoreactivity in the cat cerebellum. Neu-

roscience 45:671–690

Kasa P (1986) The cholinergic systems in brain and spinal cord. Prog

Neurobiol 26:211–272

Kaslin J, Nystedt JM, Ostergard M, Peitsaro N, Panula P (2004) The

orexin/hypocretin system in zebrafish is connected to the

aminergic and cholinergic systems. J Neurosci 24:2678–2689

Kimura H, McGeer PL, Peng JH, McGeer EG (1981) The central

cholinergic system studied by choline acetyltransferase immu-

nohistochemistry in the cat. J Comp Neurol 200:151–201

King WM (1990) Nicotinic depolarization of optic nerve terminals

augments synaptic transmission. Brain Res 527:150–154

King WM, Schmidt JT (1991) A cholinergic circuit intrinsic to optic

tectum modulates retinotectal transmission via presynaptic

nicotinic receptors. Ann N Y Acad Sci 627:363–367

Lan CT, Wen CY, Tan CK, Ling EA, Shieh JY (1995) Multiple

origins of cerebellar cholinergic afferents from the lower

brainstem in the gerbil. J Anat 186:549–561

Levin ED, Simon BB (1998) Nicotinic acetylcholine involvement in

cognitive function in animals. Psychopharmacology 138:217–

230

Lopez JM, Smeets WJ, Gonzalez A (2002) Choline acetyltransferase

immunoreactivity in the developing brain of Xenopus laevis.


Lumsden A, Keynes R (1989) Segmental patterns of neuronal

development in the chick hindbrain. Nature 337:424–428

Maley BE, Frick ML, Levey AI, Wainer BH, Elde RP (1988)

Immunohistochemistry of choline acetyltransferase in the guinea

pig brain. Neurosci Lett 84:137–142

Manger PR, Fahringer HM, Pettigrew JD, Siegel JM (2002) The

distribution and morphological characteristics of cholinergic

cells in the brain of monotremes as revealed by ChAT

immunohistochemistry. Brain Behav Evol 60:275–297

Marın O, Gonzalez A (1999) Origin of tectal cholinergic projections

in amphibians: a combined study of choline acetyltransferase

immunohistochemistry and retrograde transport of dextran

amines. Vis Neurosci 16:271–283

Marın F, Puelles L (1995) Morphological fate of rhombomeres in

quail/chick chimeras: a segmental analysis of hindbrain nuclei.

Eur J Neurosci 7:1714–1738

Marın O, Rubenstein JL (2003) Cell migration in the forebrain. Annu

Rev Neurosci 26:441–483

Marın O, Smeets WJ, Gonzalez A (1997) Distribution of choline

acetyltransferase immunoreactivity in the brain of anuran (Ranaperezi, Xenopus laevis) and urodele (Pleurodeles waltl) amphib-

ians. J Comp Neurol 382:499–534

Marın O, Smeets WJ, Gonzalez A (1998) Evolution of the basal

ganglia in tetrapods: a new perspective based on recent studies in

amphibians. Trends Neurosci 21:487–494

Maseko BC, Manger PR (2007) Distribution and morphology of

cholinergic, catecholaminergic and serotonergic neurons in the

brain of Schreiber’s long-fingered bat, Miniopterus schreibersii.J Chem Neuroanat 34:80–94

Mason WT, Ho YW, Eckenstein F, Hatton GI (1983) Mapping of

cholinergic neurons associated with rat supraoptic nucleus:

combined immunocytochemical and histochemical identifica-

tion. Brain Res Bull 11:617–626


123

http://dx.doi.org/10.3389/fnana.2010.00130

Medina L, Reiner A (1994) Distribution of choline acetyltransferase

immunoreactivity in the pigeon brain. J Comp Neurol

342:497–537

Medina L, Smeets WJ (1992) Cholinergic, monoaminergic and

peptidergic innervation of the primary visual centers in the brain

of the lizards Gekko gecko and Gallotia galloti. Brain Behav

Evol 40:157–181

Medina L, Smeets WJ, Hoogland PV, Puelles L (1993) Distribution of

choline acetyltransferase immunoreactivity in the brain of the

lizard Gallotia galloti. J Comp Neurol 331:261–285

Mesulam MM, Geula C (1988) Nucleus basalis (Ch4) and cortical

cholinergic innervation in the human brain: observations based

on the distribution of acetylcholinesterase and choline acetyl-

transferase. J Comp Neurol 275:216–240

Mesulam MM, Mufson EJ, Levey AI, Wainer BH (1984) Atlas of

cholinergic neurons in the forebrain and upper brainstem of the

macaque based on monoclonal choline acetyltransferase immu-

nohistochemistry and acetylcholinesterase histochemistry. Neu-

roscience 12:669–686

Meyer A, Dolven SI (1992) Molecules, fossils, and the origin of

tetrapods. J Mol Evol 35:102–113

Meyer A, Wilson AC (1990) Origin of tetrapods inferred from their

mitochondrial DNA affiliation to lungfish. J Mol Evol 31:359–364

Mizukawa K, McGeer PL, Tago H, Peng JH, McGeer EG, Kimura H

(1986) The cholinergic system of the human hindbrain studied

by choline acetyltransferase immunohistochemistry and acetyl-

cholinesterase histochemistry. Brain Res 379:39–55

Molist P, Maslam S, Velzing E, Roberts BL (1993) The organization

of cholinergic neurons in the mesencephalon of the eel, Anguillaanguilla, as determined by choline acetyltransferase immuno-

histochemistry and acetylcholinesterase enzyme histochemistry.

Cell Tissue Res 271:555–566

Moreno N, Gonzalez A, Retaux S (2008) Evidences for tangential

migrations in Xenopus telencephalon: developmental patterns

and cell tracking experiments. Dev Neurobiol 68:504–520

Moreno N, Morona R, Lopez JM, Gonzalez A (2010) Subdivisions of

the turtle Pseudemys scripta subpallium based on the expression

of regulatory genes and neuronal markers. J Comp Neurol

518:4877–4902

Morona R, Gonzalez A (2008) Calbindin-D28k and calretinin

expression in the forebrain of anuran and urodele amphibians:

further support for newly identified subdivisions. J Comp Neurol

511:187–220

Morona R, Gonzalez A (2009) Immunohistochemical localization of

calbindin-D28k and calretinin in the brainstem of anuran and

urodele amphibians. J Comp Neurol 515:503–537

Morona R, Northcutt RG, Gonzalez A (2011) Immunohistochemical

localization of calbindin D28k and calretinin in the retina of two

lungfishes, Protopterus dolloi and Neoceratodus forsteri: colo-

calization with choline acetyltransferase and tyrosine hydroxy-

lase. Brain Res 1368:28–43

Motts SD, Slusarczyk AS, Sowick CS, Schofield BR (2008)

Distribution of cholinergic cells in guinea pig brainstem.

Neuroscience 154:186–195

Mueller T, Vernier P, Wullimann MF (2004) The adult central

nervous cholinergic system of a neurogenetic model animal, the

zebrafish Danio rerio. Brain Res 1011:156–169

Mufson EJ, Cunningham MG (1988) Observations on choline

acetyltransferase containing structures in the CD-1 mouse brain.

Neurosci Lett 84:7–12

Mufson EJ, Desan PH, Mesulam MM, Wainer BH, Levey AI (1984)

Choline acetyltransferase-like immunoreactivity in the forebrain

of the red-eared pond turtle (Pseudemys scripta elegans). Brain

Res 323:103–108

Mufson EJ, Martin TL, Mash DC, Wainer BH, Mesulam MM (1986)

Cholinergic projections from the parabigeminal nucleus (Ch8) to

the superior colliculus in the mouse: a combined analysis of

horseradish peroxidase transport and choline acetyltransferase

immunohistochemistry. Brain Res 370:144–148

Munoz A, Munoz M, Gonzalez A, Ten Donkelaar HJ (1995) Anurandorsal column nucleus: organization, immunohistochemical

characterization, and fiber connections in Rana perezi and

Xenopus laevis. J Comp Neurol 363:197–220

Munoz M, Munoz A, Marın O, Alonso JR, Arevalo R, Porteros A,

Gonzalez A (1996) Topographical distribution of NADPH-

diaphorase activity in the central nervous system of the frog,

Rana perezi. J Comp Neurol 367:54–69

Munoz M, Marın O, Gonzalez A (2000) Localization of NADPH

diaphorase/nitric oxide synthase and choline acetyltransferase in the

spinal cord of the frog, Rana perezi. J Comp Neurol 419:451–470

Nieuwenhuys R (1998) Lungfishes. In: Nieuwenhuys R, Ten

Donkelaar HJ, Nicholson C (eds) The central nervous system

of vertebrates. Springer, Berlin, pp 939–1006

Northcutt RG (1980) Retinal projections in the Australian lungfish.

Brain Res 185:85–90

Northcutt RG (2009) Telencephalic organization in the spotted

African Lungfish, Protopterus dolloi: a new cytological model.

Brain Behav Evol 73:59–80

Northcutt RG (2011) The central nervous system of lungfishes. In:

Jørgensen JM, Joss J (eds) Biology of Lungfishes. Science

Publishers, Enfield, pp 393–446

Northcutt RG, Westhoff G (2011) Connections of the medial

telencephalic wall in the spotted African Lungfish. Brain Behav

Evol 77:14–32

Oh JD, Woolf NJ, Roghani A, Edwards RH, Butcher LL (1992)

Cholinergic neurons in the rat central nervous system demon-

strated by in situ hybridization of choline acetyltransferase

mRNA. Neuroscience 47:807–822

Parnavelas JG, Kelly W, Franke E, Eckenstein F (1986) Cholinergic

neurons and fibres in the rat visual cortex. J Neurocytol

15:329–336

Perez SE, Yanez J, Marın O, Anadon R, Gonzalez A, Rodrıguez-

Moldes I (2000) Distribution of choline acetyltransferase

(ChAT) immunoreactivity in the brain of the adult trout and

tract-tracing observations on the connections of the nuclei of the

isthmus. J Comp Neurol 428:450–474

Perry E, Walker M, Grace J, Perry R (1999) Acetylcholine in mind: a

neurotransmitter correlate of consciousness? Trends Neurosci

22:273–280

Pombal MA, El Manira A, Grillner S (1997) Afferents of the lamprey

striatum with special reference to the dopaminergic system: a

combined tracing and immunohistochemical study. J Comp

Neurol 386:71–91

Pombal MA, Marın O, Gonzalez A (1999) Choline acetyltransferase

immunoreactivity in the hypothalamoneurohypophysial system

of the lamprey. Eur J Morphol 37:103–106

Pombal MA, Marın O, Gonzalez A (2001) Distribution of choline

acetyltransferase-immunoreactive structures in the lamprey

brain. J Comp Neurol 431:105–126

Porteros A, Gomez C, Valero J, Calvo-Baltanas F, Alonso JR (2007)

Chemical organization of the macaque monkey olfactory bulb:

III. Distribution of cholinergic markers. J Comp Neurol

501:854–865

Powers AS, Reiner A (1993) The distribution of cholinergic neurons

in the central nervous system of turtles. Brain Behav Evol

41:326–345

Puelles L (1995) A segmental morphological paradigm for under-

standing vertebrate forebrains. Brain Behav Evol 46:319–337

Puelles L, Rubenstein JL (1993) Expression patterns of homeobox

and other putative regulatory genes in the embryonic mouseforebrain suggest a neuromeric organization. Trends Neurosci

16:472–479


123

Puelles L, Rubenstein JL (2003) Forebrain gene expression domains

and the evolving prosomeric model. Trends Neurosci

26:469–476

Puelles L, Milan FJ, Martınez-de-la-Torre M (1996) A segmental map

of architectonic subdivisions in the diencephalon of the frog

Rana perezi: acetylcholinesterase-histochemical observations.


Pushchina EV, Karpenko AA (2007) Distribution of cholineacetyl-

transferase histochemistry in isthmus and medulla of Onchoryn-chus masu. Tract-tracing observation on the ascending meso-

pontine cholinergic system. Tsitologiia 49:581–593

Reiner PB, Fibiger HC (1995) Psychopharmacology: the fourth

generation of progress. In: Bloom FE, Kupfer DJ (eds) Func-

tional heterogeneity of central cholinergic systems. Raven, New

York, pp 147–153

Reiner A, Northcutt RG (1987) An immunohistochemical study of the

telencephalon of the African lungfish, Protopterus annectens.


Reiner A, Medina L, Veenman CL (1998) Structural and functional

evolution of the basal ganglia in vertebrates. Brain Res Brain Res

Rev 28:235–285

Rico B, Cavada C (1998) A population of cholinergic neurons is

present in the macaque monkey thalamus. Eur J Neurosci

10:2346–2352

Rink E, Wullimann MF (2002) Connections of the ventral telenceph-

alon and tyrosine hydroxylase distribution in the zebrafish brain

(Danio rerio) lead to identification of an ascending dopaminer-

gic system in a teleost. Brain Res Bull 57:385–387

Rink E, Wullimann MF (2004) Connections of the ventral telenceph-

alon (subpallium) in the zebrafish (Danio rerio). Brain Res

1011:206–220

Rodrıguez-Moldes I, Molist P, Adrio F, Pombal MA, Yanez SE,

Mandado M, Marın O, Lopez JM, Gonzalez A, Anadon R (2002)

Organization of cholinergic systems in the brain of different fish

groups: a comparative analysis. Brain Res Bull 57:331–334

Rosen DE, Forey PL, Gardiner BG, Patterson C (1981) Lungfishes,

tetrapods, paleontology, and plesiomorphy. Bull Am Mus Nat

Hist 167:159–276

Rubenstein JL, Martınez S, Shimamura K, Puelles L (1994) The

embryonic vertebrate forebrain: the prosomeric model. Science

266:578–580

Ruggiero DA, Giuliano R, Anwar M, Stornetta R, Reis DJ (1990)

Anatomical substrates of cholinergic-autonomic regulation in the

rat. J Comp Neurol 292:1–53

Sanchez-Camacho C, Lopez JM, Gonzalez A (2006) Basal forebrain

cholinergic system of the anuran amphibian Rana perezi:evidence for a shared organization pattern with amniotes.


Sargent PB, Pike SH, Nadel DB, Lindstrom JM (1989) Nicotinic

acetylcholine receptor-like molecules in the retina, retinotectal

pathway, and optic tectum of the frog. J Neurosci 9:565–573

Satoh K, Fibiger HC (1985) Distribution of central cholinergic

neurons in the baboon (Papio papio). I. General morphology.


Semba K (2004) Phylogenetic and ontogenetic aspects of the basal

forebrain cholinergic neurons and their innervation of the

cerebral cortex. Prog Brain Res 145:3–43

Smeets WJ, Gonzalez A (2000) Catecholamine systems in the brain of

vertebrates: new perspectives through a comparative approach.

Brain Res Brain Res Rev 33:308–379

Sorenson EM, Parkinson D, Dahl JL, Chiappinelli VA (1989)

Immunohistochemical localization of choline acetyltransferase

in the chicken mesencephalon. J Comp Neurol 281:641–657

Sternberger LA (1979) Immunocytochemistry. Wiley, New York

St-Jacques R, Gorczyca W, Mohr G, Schipper HM (1996) Mapping of

the basal forebrain cholinergic system of the dog: a choline

acetyltransferase immunohistochemical study. J Comp Neurol

366:717–725

Straka H, Baker R, Gilland E (2006) Preservation of segmental

hindbrain organization in adult frogs. J Comp Neurol 494:228–245

Tago H, McGeer PL, Bruce G, Hersh LB (1987) Distribution of

choline acetyltransferase-containing neurons of the hypothala-

mus. Brain Res 415:49–62

Tago H, McGeer PL, McGeer EG, Akiyama H, Hersh LB (1989)

Distribution of choline acetyltransferase immunopositive struc-

tures in the rat brainstem. Brain Res 495:271–297

Tohyama Y, Ichimiya T, Kasama-Yoshida H, Cao Y, Hasegawa M,

Kojima H, Tamai Y, Kurihara T (2000) Phylogenetic relation of

lungfish indicated by the amino acid sequence of myelin DM20.

Brain Res Mol Brain Res 80:256–259

Tumosa N, Stell WK, Johnson CD, Epstein ML (1986) Putative

cholinergic interneurons in the optic tectum of goldfish. Brain

Res 370:365–369

van der Zee EA, Luiten PG (1999) Muscarinic acetylcholine receptors

in the hippocampus, neocortex and amygdala: a review of

immunocytochemical localization in relation to learning and

memory. Prog Neurobiol 58:409–471

Vanderwolf CH (1987) Near-total loss of ‘learning’ and ‘memory’ as

a result of combined cholinergic and serotonergic blockade in

the rat. Behav Brain Res 23:43–57

Varga C, Hartig W, Grosche J, Keijser J, Luiten PG, Seeger J, Brauer

K, Harkany T (2003) Rabbit forebrain cholinergic system:

morphological characterization of nuclei and distribution of

cholinergic terminals in the cerebral cortex and hippocampus.


Vincent SR, Reiner PB (1987) The immunohistochemical localization

of choline acetyltransferase in the cat brain. Brain Res Bull

18:371–415

von Bartheld CS (1992) Oculomotor and sensory mesencephalic

trigeminal neurons in lungfishes: phylogenetic implications.


Wainer BH, Levey AI, Mufson EJ, Mesulam MM (1984) Cholinergic

systems in mammalian brain identified with antibodies against

choline acetyltransferase. Neurochem Int 6:163–182

Wiggers W (1998) Isthmotectal connections in plethodontid sala-

manders. J Comp Neurol 395:262–272

Woolf NJ (1991) Cholinergic systems in mammalian brain and spinal

cord. Prog Neurobiol 37:475–524

Woolf NJ, Butcher LL (1986) Cholinergic systems in the rat brain:

III. Projections from the pontomesencephalic tegmentum to the

thalamus, tectum, basal ganglia, and basal forebrain. Brain Res

Bull 16:603–637

Woolf NJ, Harrison JB, Buchwald JS (1990) Cholinergic neurons of

the feline pontomesencephalon. II. Ascending anatomical pro-

jections. Brain Res 520:55–72

Wullimann MF, Rink E (2002) The teleostean forebrain: a compar-

ative and developmental view based on early proliferation, Pax6

activity and catecholaminergic organization. Brain Res Bull

57:363–370

Zardoya R, Cao Y, Hasegawa M, Meyer A (1998) Searching for the

closest living relative(s) of tetrapods through evolutionary

analyses of mitochondrial and nuclear data. Mol Biol Evol

15:506–517

Zottoli SJ, Rhodes KJ, Mufson EJ (1987) Comparison of acetylcho-

linesterase and choline acetyltransferase staining patterns in the

optic tectum of the goldfish Carassius auratus. A histochemical

and immunocytochemical analysis. Brain Behav Evol

30:143–159

Zottoli SJ, Rhodes KJ, Corrodi JG, Mufson EJ (1988) Putative

cholinergic projections from the nucleus isthmi and the nucleus

reticularis mesencephali to the optic tectum in the goldfish

(Carassius auratus). J Comp Neurol 273:385–398


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Documents

Organization of the cholinergic systems in the brain of two lungfishes, Protopterus dolloi and Neoceratodus forsteri