neuroendocrinology of fish metamorphosis and puberty

Special Issue, pp. 55-68 (2007)
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NEUROENDOCRINOLOGY OF FISH
METAMORPHOSIS AND PUBERTY: EVOLUTIONARY
AND ECOPHYSIOLOGICAL PERSPECTIVES
Sylvie Dufour* and Karine Rousseau*
Key words: puberty, metamorphosis, vertebrates, teleosts, neuroendocrinology, thyroid hormones, steroids, silvering, smoltification,
ABSTRACT
Metamorphosis and puberty are two major events of the postembryonic development in Vertebrates. Based on some examples from
fish species, we review the definition, role and regulation of these
events, analyze their common and different features, as well as their
impact on the evolution and diversity of life cycles.
The term of puberty, firstly defined in humans, has been subsequently extended to the first acquisition of the capacity to reproduce
in all mammalian and non-mammalian vertebrates as well as in
invertebrates. By definition, puberty occurs only once in the life
cycle. However, some similarities may be found with other events,
such as annual re-activation of the reproductive function in seasonal
breeders or sex-change in adults, as observed in some fish species.
Metamorphosis allows the transition from one developmental
stage in a specific environment to the next stage in a different
environment, and includes a migration between the two habitats.
Metamorphosis corresponds to drastic changes in body shape, physiology and behavior, and, unlike puberty, is encountered only in
some phyla/species. In Vertebrates, the most described metamorphosis is the transformation in Amphibians of the aquatic larva
(tadpole) into the terrestrial juvenile. Larval metamorphosis is also
encountered in some other Vertebrates, such as lampreys and some
teleosts (Elopomorphes and Pleuronectiformes). Less drastic
morphological, physiological and behavioral changes occur in juveniles of some migratory teleosts. This is the case of smoltification
in salmons and silvering in eels, which are referred to as “secondary
metamorphoses”.
Investigations on the regulation of puberty and metamorphoses
in Vertebrates reveal the crucial roles of the neuroendocrine axes. In
all Vertebrates, puberty is triggered by the activation of the gonadotropic axis, constituted of brain neuropeptide (gonadotropin-releasing
hormone, GnRH), pituitary glycoprotein hormones (gonadotropins:
luteinizing hormone, LH and follicle stimulating hormone, FSH) and
gonadal steroids. Sex steroids induce the morpho-physiological and
behavioral transformations characteristic of puberty.
Metamorphosis in Amphibians is triggered by the thyrotropic
axis, constituted of brain neuropeptide corticotropin-releasing horAuthor for Correspondence: Sylvie Dufour.
E-mail: [email protected]
*UMR 5178 CNRS Biology of Marine Organisms and Ecosystems, National
Museum of Natural History (MNHN), National Center for Scientific
Research (CNRS), University Pierre & Marie Curie (Paris VI), Paris,
France.
mone (CRH), instead of thyrotropin-releasing hormone, (TRH), pituitary glycoprotein hormone (thyrotropin, TSH) and thyroid hormones
(TH: thyroxine, T4 and triiodothyronine, T3), which play a key-role
in the induction of morpho-physiological and behavioral changes. A
similar control is suggested for larval metamorphosis in teleosts.
Studies on smoltification also indicate an important role of
thyroid hormones in secondary metamorphoses in teleosts, even
though other hormones such as growth hormone (GH) and corticosteroids may be of prime importance. In contrast, recent investigations in
the eel reveal that the gonadotropic axis, and ultimately sex steroids
would be the major triggering control of silvering. The similarities of
the morpho-physiological and behavioral changes between the two
species indicate remarkable evolutionary convergences in the morphogenetic roles and target tissues of TH and sex steroids for the
induction of secondary metamorphoses. In all cases, the possible
synergistic role of cortisol is highlighted.
Comparison of puberty and metamorphosis may also favor our
understanding of the internal and environmental triggering signals of
these postembryonic developmental events. In teleosts, the large
plasticity in the occurrence and timing of metamorphosis and puberty,
which contributes to the high diversity of fish life cycles, may provide
new and relevant models to such investigations.
INTRODUCTION: DEFINITION OF
PUBERTY AND METAMORPHOSIS
Metamorphosis and puberty are two major events
of the postembryonic development.
1. Puberty and related events
The term of puberty, firstly defined in humans, has
been subsequently extended to the first acquisition of
the capacity to reproduce in all mammalian and nonmammalian vertebrates as well as in invertebrates.
By definition, puberty occurs only once in the life
cycle after a certain period of juvenile growth phase.
However, some similarities may be found with other
physiological events, such as the annual re-activation of
the reproductive function, as observed in seasonal breeders [43, 143]. Furthermore, in the remarkable case of
fish species that are able to change sex during their life
cycle [4], the new ability to reproduce as a member of
the opposite sex could also be considered as a “second
puberty”.
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Special Issue (2007)
2. Metamorphosis and related events
Metamorphosis allows the transition from one developmental stage in a specific environment (ecophase
1 in habitat 1) to the next stage in a quite different
environment (ecophase 2 in habitat 2), and normally
includes the occurrence of a migratory phase between
the two habitats. Metamorphosis corresponds to drastic
changes in body shape, physiology and behavior, and,
unlike puberty, is encountered only in some phyla/
species. In Vertebrates, the classical and most described model for metamorphosis is the one of the
Anuran Amphibians, which transforms the aquatic larva
(tadpole) into a terrestrial, adult-like shaped, juvenile.
Larval metamorphosis, also called “primary metamorphosis” is encountered in some other vertebrates, such
as in lampreys (Agnathans) and teleosts (Elopomorphes,
Pleuronectiformes) [157].
Other developmental events, which also encompass morphological, physiological and behavioral
(migratory) changes, even though less drastic than for
the larval metamorphosis, are encountered in some migratory teleosts [116]. This is the case of smoltification
in salmons and silvering in eels. These developmental
events occur after some period of juvenile phase and
prepare the fish to the transition between the continental
habitat and the oceanic one. They are traditionally
referred to as “secondary metamorphoses”.
Birth in mammals or hatching in oviparous
Vertebrates, which are abrupt transitions from egg/
maternal to outside environment, may also present some
common features with metamorphoses.
Investigations on the regulation of puberty and
metamorphoses in Vertebrates reveal the key roles of
the brain-pituitary neuroendocrine axes.
NEUROENDOCRINE CONTROL OF PUBERTY
1. Neuroendocrine control of puberty in mammals
Puberty in mammals is clearly characterized by an
activation of the gonadotropic axis [31, 127]. This neuroendocrine axis is constituted of a brain neuropeptide,
the gonadotropin-releasing hormone (GnRH), which
stimulate synthesis and release of pituitary glycoprotein hormones, the gonadotropins, luteinizing hormone
(LH) and follicle stimulating hormone (FSH), which act
on the gonads to activate gametogenesis and
steroidogenesis. Sex steroids (androgens and estrogens)
act as potent morphogenic hormones on the peripheral
target tissues, inducing various morpho-physiological
and behavioral changes, characteristics of the puberty
(secondary sexual characters) [115, 127]. In addition,
sex steroids exert positive and negative feedbacks on
the brain-pituitary axis, allowing a regulatory cross talk
between central and peripheral components of the gonadotropic axis.
The pubertal activation of the gonadotropic axis
occurs after a certain duration of juvenile phase (also
called “infancy”), which allows the organism to reach
certain age, size, energy stores sufficient enough to
ensure the success of reproduction [7, 38, 39, 60, 68].
The duration of this phase depends on genetic, internal
and environmental factors. It can be short (for instance
in the mice) or long (as in humans) depending on the life
cycle strategies. In seasonal breeders (for instance in
sheep), a seasonal reactivation of the gonadotropic axis
occurs every year after puberty and presents many
common features with puberty itself [143]. Thus, alternate phases of inhibition and re-activation of the gonadotropic axis are at the basis of the annual cycles of
reproduction.
Recent studies showed that a peptide named
kisspeptin (or metastin), product of the Kiss-1 gene,
played a major role in the onset of puberty in mammals
[12, 95, 122, 130]. In 2003, three groups described the
effects of knock-out [40, 123] and mutation [21] of
GPR54, which is kisspeptin receptor. They observed
that when GPR54 is absent or mutated, mice or humans
were unable to undergo puberty, because of small gonads,
and low concentrations of sexual steroids and
gonadotropins. In 2005, Messager and collaborators
showed that in mice lacking GPR54, the anatomy and
localisation of GnRH neurons, as well as GnRH concentration in the brain, remained unchanged. These results
suggested that there was no problem in GnRH synthesis
in these mice, but that GnRH release was blocked.
Hypothalamic expression of kisspeptin and its receptor
increase dramatically at puberty and is modulated by
sex steroids (rat: [97]; mouse: [130]; rhesus monkey:
[124]). Recently, studies demonstrated that GPR54
receptor was expressed within GnRH neurons in mammals (mouse: [86]; ovins: [108]). Kisspeptin injections
to different animal models can induce the release of
GnRH release, as well as FSH and LH, whereas administration of antibodies against kisspeptin block reproductive function, even when puberty has been initiated
[25, 44, 82, 144]. Concomitant injection of kisspeptin
and GnRH antagonist blocks the stimulatory effect of
the peptide on FSH and LH release [124], and kisspeptin
has no direct effect in vitro on FSH and LH release
[144]. All these data show that at puberty, kisppeptin,
which brain expression is increased, acts via GPR54
receptor directly on GnRH neurons in order to induced
GnRH release which then stimulates the pituitary production of LH and FSH . In a seasonal model, the Syrian
hamster, it was shown that melatonin impacted on Kiss1 expression to control reproduction and that Kiss-1
S. Dufour & K. Rousseau: Neuroendocrinology of Fish Metamorphosis and Puberty: Evolutionary and Ecophysiological Perspectives
expression was significantly higher in hamsters kept in
long-day as compared to short-day [111]. These data
suggest that photoperiod, via melatonin, modulates Kiss1 neurons to drive the reproductive axis in seasonal
breeders [111].
2. Neuroendocrine control of puberty in teleosts
This scheme (GnRH/LH-FSH/sex steroids) is
largely conserved among Vertebrates [101], even though
additional controls may occur such as the dopaminergic
inhibition of gonadotropin production in some teleosts
[29, 104]. Indeed, pioneer works from Peter and collaborators on goldfish using hypothalamic lesions demonstrated the existence of a GRIF (gonadotropin release-inhibiting factor) [105-107] Subsequent studies
using agonists or antagonists in vivo [9, 10], primary
culture of pituitary cells in vitro [11] and immunocytochemistry [64-66] provided evidences that GRIF was
dopamine (DA). An inhibitory role of DA on the control
of LH has been evidenced in many adult teleosts at the
time of ovulation and spermiation (catfish: [16, 145];
coho salmon: [146]; rainbow trout: [76, 118]; common
carp: [75]; tilapia: [154]). However, DA does not play
an inhibitory role in all adult teleosts (Atlantic croaker:
[13]; gilthead seabream: [161]).
Concerning the early stages of gametogenesis and
the control of puberty, up to now, the possible involvement of DA has only been studied in a few species. In
juvenile striped bass [55] and red seabream [72], data
refuted a role for DA in the prepubertal control of
gonadotropins, as GnRH alone was able to trigger precocious puberty. In contrast in European eel, only a
triple treatment with testosterone, GnRH agonist and
pimozide (DA-receptor antagonist) could induce increases in LH synthesis and release, indicating that
removal of DA inhibition is required in prepubertal eel
for triggering GnRH-stimulated LH synthesis and release [148]. A recent study in the grey mullet has
demonstrated that D2 type receptor expressions in the
brain and in the pituitary were high at the early and
intermediate stages of puberty [98], when inhibition of
the reproductive function by DA is particularly pronounced [1].
In male tilapia, GPR54 mRNA was found to be colocalized within all three GnRH neuron subtypes [103],
and the number of neurons expressing GPR54, as well
as the level of GPR54 expression, increased with gonadal maturation [103]. In cobia Rachycentron canadum,
concomitant expression patterns of GPR54 and GnRH
mRNAs were reported during different stages of larval
and juvenile developments [92]. Moreover, an increase
in GPR54 was observed during early puberty [92], as in
pubertal rats [97] and rhesus monkey [124]. Similarly,
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in female grey mullet (Mugil cephalus), significantly
high levels of GPR54 mRNAs were demonstrated in
brain at the early stage of puberty that subsequently
decreased as puberty advanced [98].
NEUROENDOCRINE CONTROL OF LARVAL
METAMORPHOSIS
1. Neuroendocrine control of amphibian metamorphosis
Early data on the regulation of metamorphosis
came from Amphibians and demonstrated the key-role
of a surge in thyroid hormones (TH: thyroxine, T4 and
triiodothyronine, T3), in the induction of the many
morpho-physiological and behavioral changes characteristics of the larval metamorphosis. The role of the
thyroid gland in the control of larval metamorphosis
was first demonstrated by Gudernatsch in 1912 after he
observed the acceleration of the tadpole transformation
into frogs when feeding them with thyroid gland extracts.
Inversely, Allen [2] was able to completely prevent
metamorphosis by thyroidectomy. As the thyrotropic
axis is activated, a series of sequential morphological
transformations occur. An early change is the growth
and differential of the limbs, which in the absence of
hormone, still form but will not progress beyond the bud
stage. The final morphological change, tail resorption,
occurs when the level of TH is highest at the climax of
metamorphosis [67, 142].
Thyrotropin (TSH), a pituitary glycoprotein
hormone, belonging to the same family as gonadotropins,
controls the production of TH in Amphibians [80], as
classically shown in mammals [85]. Early studies of
hypophysectomy and immunization demonstrated that
TSH played a central role in amphibian metamorphosis
[26, 32]. Indeed, Dodd and Dodd [26] showed that the
negative effect of hypophysectomy prior to metamorphosis could be reversed by treatment with mammalian
TSH. Furthermore, passive immunization of tadpoles
with an antiserum to bovine TSH prevented spontaneous metamorphosis [32]. Morphological and biochemical changes observed during metamorphosis, such as
complete regression of tail and gills, de novo formation
of bone, visual pigment transformation or functional
differentiation of liver, can be induced by TH [126,
141]. In contrast to the situation in mammals in which
the brain peptide discovered for its stimulatory control
on TSH is TRH (for Thyrotropin Releasing Hormone)
[93], the brain neurohormone responsible for the activation of TSH production during amphibian metamorphosis, is corticotropin-releasing hormone (CRH) and
not TRH [23]. In fact, in amphibians, the production
and release of TSH by the pituitary appears to be regulated by different neuropeptides according to the life
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stage. Indeed, in premetamorphic amphibians, stimulation of the pituitary-thyroid axis is only responsive to
CRH-like peptides and unresponsive to all other neurohormones tested, including TRH [22, 24]. Thus, CRH is
the thyrotropin-releasing factor during the induction of
metamorphosis in tadpoles, and the stimulatory action
of TRH on TSH secretion develops after metamorphic
climax [23, 142].
Although TH is the only obligatory signal for the
initiation and completion of amphibian metamorphosis,
other hormones can modulate the onset and progression
of metamorphosis [142]. These include glucocorticoids
and prolactin (PRL), which can accelerate and prevents
TH-induced metamorphosis, respectively [70, 140, 151].
In Amphibians, the importance of CRH in metamorphosis is reinforced by its traditional and evolutionary
conserved role in the corticotropic axis itself (activation
of pituitary corticotropin (ACTH) and adrenal cortisol).
Indeed, cortisol has been shown to act in synergy with
TH for the induction of various metamorphosis-related
morphogenetic changes.
2. Neuroendocrine control of larval metamorphosis in
lamprey
Lampreys, which are extant representatives of some
of the oldest known vertebrates, the jawless fish or
Agnatha [35, 36], also show a larval metamorphosis,
first described by Muller [94], in their life cycle [158].
During this metamorphosis, there are major changes in
external and internal features, and among them: final
development of the eye; total regression of the larval
kidney, replaced by an adult one; transformation of the
epithelium in the intestine, gills and endostyle; development of teeth and tongue needed for adult feeding
[156, 157].
Opposite to the situation observed in amphibians,
larval (ammocoete) metamorphosis in lampreys is characterized by a sharp drop in T4 and T3 plasma levels
(sea lamprey, Petromyzon marinus: [152, 159]; sea
lamprey and Lampetra lamottenii: [77]; southern hemisphere lamprey, Geotria australis: [74], whereas concentrations of thyroid hormones during their larval life
are among the highest recorded in any vertebrate [77].
Accordingly, immersion of ammocoetes in potassium perchlorate (KClO4) (which inhibits iodide uptake
and TH synthesis by the thyroid: [8] resulted in precocious metamorphosis (Lampetra planeri: [55, 134];
Lampetra reissneri: [136]). Moreover, both T4 and T3
treatment can block KClO4-induced metamorphosis [81,
158] and T3 treatment can inhibit spontaneous metamorphosis [160]. However, the use of propythiouracil
(PTU), another inhibitor of TH synthesis, was unable to
induce metamorphosis in the southern hemisphere
lamprey, despite the decline in serum levels of T4 and
T3 [74]. In addition, artificial maintenance of serum
concentrations of thyroid hormones in immediately
premetamorphic lampreys did not block metamorphosis
in all individuals [158].
Studies on deiodinases and TH receptors gave
interesting data, which may help to understand the
involvement of TH in lamprey metamorphosis. Indeed,
there is a shift in monodeiodinase pathways between
larval and adult life in lampreys that may account for the
decline in serum levels of TH at the beginning of the
metamorphosis [30]. In addition, a reduction in the
capacity of T3 nuclear receptors in hepatocytes following larval life may reflect the importance of this hormone to the larval phases of growth and metamorphosis
[78].
All these data, even if still controversial, suggest
that, lamprey metamorphosis is unlike any other vertebrate metamorphosis in that TH appear to be antagonistic (inhibitory) to the process. Indeed, in contrast to
amphibians, in lampreys, induction of metamorphosis
seems to be driven by a drop of TH [158].
These two opposite examples show that, during
the evolution of vertebrates, the role of thyroid hormones in the endocrinology of larval metamorphosis
may have differed dramatically, being possibly inhibitory in lampreys and stimulatory in amphibians.
3. Neuroendocrine control of larval metamorphosis in
teleosts
Typical larval metamorphosis in teleost fish is
restricted to Anguilliformes, Elopiformes, Notacanthiformes and Pleuronectiformes [158]. The two groups,
Anguilliformes and Pleuronectiformes, represent one
of the most ancient and one of the most recent groups of
teleosts, respectively. This suggests that larval metamorphosis may have been acquired independently by
these two groups during teleost evolution. The alternative hypothesis would be that larval metamorphosis
could have been lost in most other teleost groups. The
most spectacular (and studied) larval metamorphoses in
teleosts are the metamorphosis from leptocephalus larva
to glass eels and the flatfish metamorphosis.
(1) Anguilliformes
Early studies on leptocephali of Anguilla anguilla
showed increased thyroid gland development [96] and
activation [128] during metamorphosis, a result later
confirmed in Conger myriaster [71, 153]. During early
metamorphosis, T4 body content increases about sixfold in C. myriaster, and then decreases as metamorphosis progresses; T3 body content increases gradually in
S. Dufour & K. Rousseau: Neuroendocrinology of Fish Metamorphosis and Puberty: Evolutionary and Ecophysiological Perspectives
early metamorphosis and then increases abruptly (about
13-fold) toward the end of the period [153]. In
agreement, histological evidence revealed an activation
of the thyroid gland [153].
Exogenous thyroid hormone has been shown to
stimulate metamorphosic changes in leptocephali of
Conger myriaster [69] and of Anguilla anguilla [149],
confirming the major role of TH in the induction of
elopomorph larval metamorphosis.
(2) Pleuronectiformes (flatfish)
Hormone assays in flatfish showed a surge in TH
concentration during metamorphosis with a peak around
the time of metamorphic climax [17, 50]; Japanese
flounder, Paralichthys olivaceus: [90, 139]). In agreement with the activation of the thyroid function during
larval metamorphosis, histological study of the pituitary also showed an activation of TSH cells (plaice,
Pleuronectes platessa: [128]; Japanese flounder: [87]).
Accordingly, early works by Miwa and Inui [61,
88] in flounder (Paralichthys olivaceus) reported that
exogenous TH could induce metamorphosis (eye
migration, settling behavior and length of second dorsal
fin ray) with the production of a miniature of the normally metamorphosed juvenile, while thiourea treatment arrested the metamorphic process of the fish.
Subsequent experimental studies by various authors
have demonstrated that TH treatment was able to induce
the many morphological, physiological and behavioral
changes, characteristics of flatfish metamorphosis, such
as shift in erythrocyte populations [89], histological and
biochemical changes in muscle [153], development of
gastric glands [57, 91, 131], changes of gill mitochondria-rich cells from larval to juvenile form [121] and
bone remodelling for eye relocation [100, 132]. Also,
[62] showed that injection of bovine TSH into flounder
larvae increased tissue concentrations of T4 and accelerated the metamorphic process, such as shortening of
the second fin ray and eye migration.
These data demonstrated that the thyrotropic axis
(TSH-T4/T3) would be the main axis controlling metamorphosis in flatfish. However, nothing is yet known
on the brain neurohormones potentially involved in this
activation.
Beside the thyrotropic axis, cortisol was shown to
synergize with thyroid hormones, while sex steroids
and prolactin exhibited an antagonist effect (for review:
[20]; Japanese flounder: [18, 19]), in agreement with
data in amphibians. Changes in tissue cortisol concentrations closely parallel those of thyroid hormones,
except that cortisol peaks a few days earlier than T4
(Japanese flounder: [17]). Both PRL and growth hormone (GH) expression increased gradually but steadily
59
during metamorphosis and showed a dramatic rise in
post-climax fish [20]. In contrast, tissue levels of
estradiol and testosterone remain low and do not show
marked change during metamorphosis [16].
These data demonstrated the involvement of thyroid hormones as major triggers of metamorphosis in
eels as in flatfish. Further studies are clearly needed to
investigate the potential synergistic or antagonistic roles
of other hormones as well as to determine the brainpituitary control of thyroid function.
A few data in adult teleosts suggest also a role for
CRH and/or TRH in the control of TSH, with variations
possibly depending on species or physiological status
(coho salmon: [73]; European eel: [109]). Further
studies are clearly needed to investigate which brain
neuromediator is specifically implicated in the triggering of larval metamorphosis in teleosts.
In conclusion, these data indicate that thyroid
hormones play a key role in the induction of larval
metamorphoses in teleosts, as well as in amphibians,
while in lampreys TH would be inhibitory. This suggests that the stimulatory role of TH in metamorphosis
may have been acquired in a common ancestor of teleosts (actinopterygian lineage) and amphibians
(sarcopterygian lineage) posteriorly to the emergence
of agnathans. An alternative hypothesis is that the
stimulatory role of TH in larval metamorphosis could
have been acquired independently in amphibians and in
teleosts.
NEUROENDOCRINE CONTROL OF
SECONDARY METAMORPHOSES
Secondary metamorphoses have been described in
some diadromous migratory teleosts. They prepare the
fish to the river downstream migration, to the transfer
from fresh to seawater and finally to the oceanic
migration.
1. Neuroendocrine control of smoltification
In teleosts, a well-known example of “secondary
metamorphosis” is provided by smoltification in
salmons. This transformation from parr to smolt, which
occurs in the river, encompasses various morphological
(silvering of the body color), physiological
(osmoregulation, vision) and behavioral (rheotaxism)
changes, and preadapts the smolt to its future oceanic
growth ecophase [5]. According to the crucial role of
smoltification in the salmon capacity to adapt to seawater and thus in the success of its biological cycle and
aquaculture, many investigations have been performed
on the endocrine control of smoltification.
A number of endocrine investigations emphasized
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Special Issue (2007)
the role of TH in these changes. Thyroid involvement in
smoltification was originally suggested by Hoar [51]
who observed histological activation of thyroid tissue
of the Atlantic salmon (Salmo salar). The availability
of radioimmunassay (RIA) procedures has enabled different groups to demonstrate T4 and T3 surges during
the smoltification process [52, 53]. Moreover, the
administration of exogenous TH to juvenile parr-status
salmonids results in morphological and physiological
changes, which are consistent with the parr-smolt transformation [27, 34, 49, 84, 113, 135].
Many other studies revealed the key-role in
smoltification of other hormones, such as growth hormone (GH) and cortisol. Hypertrophy and hyperplasia
of somatotrophs [27] and a rise in plasma GH have been
observed during smoltification (coho salmon: [137, 138,
155]; Atlantic salmon: [110]). Moreover, administration of GH clearly improves hypo-osmoregulatory ability and seawater survival of parr (for review: [5, 27]). In
addition, GH treatment also induces other smoltificationrelated changes, such as condition factor and skin pigmentation [27]. Variations of plasma levels of cortisol
suggest a potential synergistic role during smoltification
[133, 150]. Data suggest that cortisol could also play a
significant stimulatory role in osmoregulation. Indeed,
prolonged cortisol treatment in pre-smolt coho salmon
caused an increase in Na+/K+-ATPase activity, while
treatment of smolts had no effect [112].
This is leading to a complex scheme of the pituitary control of smoltification. Moreover, neuroendocrine investigations on the brain components of the
control of smoltification are still lacking.
2. Neuroendocrine control of silvering
Another exemple of “secondary metamorphosis”
is provided by another migratory teleost, the eel.
Silvering, which transforms the yellow eel into the
silver eel, shares many similarities with smoltification,
such as change in body color, preparation to osmoregulation in seawater, and downstream migratory behavior.
Because of these analogies, it had been classically
assumed that silvering and smoltification would be
under a similar endocrine control. However, our recent
studies have contributed to reveal striking discrepancies,
with a potential key-role of the gonadotropic axis in the
induction of silvering. Indeed, a significant increase in
FSH( mRNA level could be observed during the early
stages of silvering and may correspond to the first
appearance of lipid vesicles in oocytes (endogenous
vitellogenesis) [3]. This increase in FSHβ is followed
by a sharp increase in LHβ mRNA levels at the late
stages, which may be related to the beginning of exogenous vitellogenesis [3]. At the peripheral level, sig-
nificant increases in plasma levels of sex steroids
(oestradiol, testosterone and the teleost specific andogen,
11-ketotestosterone) have been measured between yellow and silver stages (A. australis and A. dieffenbachii:
[3, 120]; A. anguilla: [79]; A. rostrata: [14]; A. japonica:
[47]). This control strongly differs from smoltification,
which is in contrast inhibited by sex steroids.
Experimental data using exogenous sex steroids
are in agreement with the involvement of the gonadotropic axis in the induction of silvering, as treatment
with androgens can induce increases of eye diameter [3,
114] and of skin thickness [114], regression of digestive
tract [3, 114] in yellow eels, and amplification of silvering parameters in silver eels (eye diameter: [6, 102];
skin thickness and darkening: [102]; regression of the
digestive tract: [148]).
In contrast, measurement of pituitary TSH mRNAs
and plasma levels of TH during silvering shows no
change in TSH and T3, and a moderate increase in T4
(Anguilla anguilla: [3]; Anguilla japonica: [48]).
Similarly, no increases were observed in GH plasma
levels, pituitary content neither in GH mRNA levels
throughout silvering in European eel [3].
This discrepancy is likely related to the position of
silvering versus smoltification in the migratory fish life
cycle, silvering corresponding to the initiation of the
reproductive phase in the ocean, while smoltification
prepares the fish to the growth phase. The similarities
of the morpho-physiological and behavioral changes
between the two species indicate remarkable evolutionary convergences in the morphogenetic roles and target
tissues of TH and sex steroids for the induction of
secondary metamorphoses. Furthermore our ongoing
studies suggest a synergistic role of glucocorticosteroids
on sex steroid-induction of silvering parameters [56,
119], a synergism recalling that observed with TH during larval and possibly also secondary metamorphosis.
It is of great interest to note that while smoltification and silvering share many similarities in term
of morphological changes, the endocrinology of these
two secondary metamorphoses drastically differs, with
the major involvement of different neuroendocrine
axes, the thyrotropic/somatotropic one for
smoltification and the gonadotropic one for silvering.
This suggests that secondary metamorphoses may have
been acquired independently, via different endocrine
mechanisms, during teleost evolution. The convergence between some morphological (skin silvering,
eye size and pigments), metabolic and behavioural
changes reflects that the control of the same peripheral
target organs (skin, eye, muscle...) and target genes is
exerted by different hormonal receptors (thyroid hormone receptors in salmon versus androgen receptors in
the eel).
S. Dufour & K. Rousseau: Neuroendocrinology of Fish Metamorphosis and Puberty: Evolutionary and Ecophysiological Perspectives
CONCLUSIONS: COMPARATIVE
NEUROENDOCRINOLOGY OF PUBERTY AND
METAMORPHOSIS.
1. Neuroendocrine “crises”
Metamorphosis and puberty are triggered by transient activations of neuroendocrine axes namely of the
thyrotropic axis for classical larval metamorphosis and
the gonadotropic axis for puberty. Beside this classical
scheme, more complex situations are revealed. As
discussed above, other axes, such as the somatotropic
and corticotropic axes, may also play a key-synergistic
role in the induction of all morphological, physiological
and behavioral changes characteristic of metamorphosis and puberty. Furthermore, distinction between metamorphosis and puberty vanishes when studying a traditionally so-called “secondary metamorphosis” such as
silvering in the eel.
Common features are the transient and large activation of brain-pituitary-peripheral neuroendocrine axes
(“neuroendocrine crises”) during which classical regulations such as homeostatic maintenance of hormones
levels and negative feedbacks may be overruled. Thus,
in teleosts, strong positive feedbacks by sex steroids on
the brain and pituitary have been evidenced that are
largely amplifying the activation of the gonadotropic
axis at puberty. In other cases, a reset of negative
feedbacks to another threshold level may occur.
The “neuroendocrine crises” also reflect the keyrole of the brain in the timing and coordination of these
developmental events. Internal and environmental cues
(triggering signals) are integrated at the brain level
leading to the activation of specific neuroendocrine
axes. Coordinated implication of neuroendocrine axes
may result from common brain control. Such a case is
exemplified by the common role of CRH in the
corticotropic axis and thyrotropic axis at metamorphosis in amphibians and also possibly in fish. Interrelationships between neuroendocrine axes may also result
from interaction by peripheral hormones. For instance
we could demonstrate stimulatory roles of cortisol
(corticotropic axis) and IGF (somatotropic axis) in the
pubertal simulation of LH (gonadotropic axis) in the eel
[58, 59, 117].
61
of the next phase (migration/reproduction) [125]. Thus,
metabolic signals such as insulin-growth factors or the
more recently discovered hormone, leptin and ghrelin,
could likely be involved in the triggering of both types
of events (mammals: [15, 33, 41, 147]; non-mammals:
[101, 125]). Comparative studies would help deciphering the brain networks implied in the integration of the
internal and environmental signals of puberty/
metamorphosis, as well as the pathways of the activation of the gonadotropic /thyrotropic and corticotropic
axes, respectively. In teleosts, the large plasticity in the
occurrence and timing of metamorphosis and puberty,
which contributes to the high diversity of fish life
cycles, may provide new and relevant models to such
investigations.
3. Threats on metamorphosis and puberty: Environmental and endocrine disruptions
The crucial morphogenetic roles of TH and steroids at the time of metamorphosis and puberty, as well
as their role on peripheral and central (feedback actions
on the brain and pituitary) target tissues, make the
organism particularly vulnerable to the endocrine disrupting effects of xenobiotics. Furthermore, aquatic
species are specially concerned by the increasing number of endocrine disruptors, currently accumulating in
fresh as well as in marine hydrosystems. Indeed, the
aquatic existence means that the animal is bathed constantly in a solution containing pollutants, and that
uptake of chemicals readily occurs via the gills and skin,
in addition to the diet. Impacts of contaminants are
various and range from subtle changes in the physiology
and sexual behavior to permanently altered sexual differentiation and impairment of fertility (wild freshwater
fish: [63]; marine fish: [83]; invertebrates: [99]; aquatic
mammals: [37]; amphibians: [42]; reptiles: [46]). In
fish, the three major neuroendocrine axes involved in
the control of puberty and metamorphoses: HPG
(hypothalamo-pituitary-gonads), HPT (hypothalamopituitary-thyroids) and HPI (hypothalamo-pituitaryinterrenals) are potentially affected by endocrine
disruptors (for reviews: freshwater fish: [63]; marine
fish: [83]).
REFERENCES
2. Triggering signals
Comparison between puberty and metamorphosis
may also favor our understanding of the triggering
signals of these postembryonic developmental events.
For instance, metamorphoses as well as puberty or even
sex change, should occur only when body size and
energy stores are sufficient enough to allow the success
1. Aizen, J., Meiri, I., Tzchori, I., Levavi-Sivan, B., and
Rosenfeld, H., “Enhancing Spawning in the Grey Mullet (Mugil cephalus) by Removal of Dopaminergic
Inhibition,” General and Comparative Endocrinology,
Vol. 142, pp. 212-221 (2005).
2. Allen, B.M., “Extirpation Experiments in Rana Pipiens
Larva,” Science, Vol. 44, pp. 755-757 (1916).
62
Special Issue (2007)
3. Aroua, S., Schmitz, M., Baloche, S., Vidal, B., Rousseau,
K., and Dufour, S., “Endocrine Evidence That Silvering,
a Secondary Metamorphosis in the Eel, Is a Pubertal
Rather Than a Metamorphic Event,” Neuroendocrinology, Vol. 82, pp. 221-232 (2005).
4. Baroiller, J.F., Guiguen, Y., and Fostier, A., “Endocrine
and Environmental Aspects of Sex Differentiation in
Fish,” Cellular and Molecular Life Sciences, Vol. 55,
pp. 910-931 (1999).
5. Boeuf, G., “Salmonid Smolting: a Pre-adaptation to the
Oceanic Environment,” In Rankin, J.C. and Jensen, F.
B. (eds), Fish Ecophysiology, Chapman and Hall,
London, pp. 105-135 (1993).
6. Boëtius, I. and Larsen, L.O., “Effects of Testosterone
on Eye Size and Spermiation in Silver Eels, Anguilla
anguilla,” General and Comparative Endocrinology,
Vol. 82, pp. 238 (1991).
7. Bronson, F.H., “Puberty and Energy Reserves: a Walk
on the Wild Side,” In Walken, K. and Schneider, J.E.
(eds), Reproduction in Context, MIT Press, Cambridge,
MA, pp. 15-33 (2000).
8. Carrasco, N., “Iodide Transport in the Thyroid Gland,”
Biochimica et Biophysica Acta, Vol. 1154, pp. 65-82
(1992).
9. Chang, J.P. and Peter, R.E., “Effects of Dopamine on
Gonadotropin Release in Female Goldfish, Carassius
auratus,” Neuroendocrinology, Vol. 36, pp. 351-357
(1983).
10. Chang, J.P., Peter, R.E., Nahorniak, C.S., and
Sokolowska, M., “Effects of Catecholaminergic Agonists and Antagonists on Serum Gonadotropin Concentrations and Ovulation in Goldfish: Evidence for
Specificity of Dopamine Inhibition of Gonadotropin
Secretion,” General and Comparative Endocrinology,
Vol. 55, pp. 351-360 (1984).
11. Chang, J.P., Yu, K.L., Wong, A.O., and Peter, R.E.,
“Differential Actions of Dopamine Receptor Subtypes
on Gonadotropin and Growth Hormone Release in vitro
in Goldfish,” Neuroendocrinology, Vol. 51, pp.
664-674 (1990).
12. Colledge, W.H., “GPR54 and Puberty,” Trends
Endocrinol and Metab, Vol. 15, pp. 448-453 (2004).
13. Copeland, P.A. and Thomas, P., “Control of Gonadotropin Release in the Atlantic Croaker (Micropogonias
undulatus): Evidence for Lack of Dopaminergic
Inhibition,” General and Comparative Endocrinology,
Vol. 74, pp. 474-483 (1989).
14. Cottrill, R.A., McKinley, R.S., Van der Kraak, G.,
Dutil, J.-D., Reid, K.B., and McGrath, K.J., “Plasma
Non-esterified Fatty Acid Profiles and 17b-Oestradiol
Levels of Juvenile Immature and Maturing Adult American Eels in the St Lawrence River,” Journal of Fish
Biology, Vol. 59, pp. 364-379 (2001).
15. Daftary, S.S. and Gore, A.C., “IGF-1 in the Brain as a
Regulator of Reproductive Neuroendocrine Function,”
Experimental Biology and Medicine, Vol. 230, pp. 292306 (2005).
16. de Leeuw, R., Goos, H.J., and Van Oordt, P.G., “The
Dopaminergic Inhibition of the Gonadotropin-Releasing Hormone-Induced Gonadotropine Release: an in
vitro Study with Fragments and Cell Suspensions from
Pituitaries of the African Catfish, Clarias gariepinus
(Burchell),” General and Comparative Endocrinology,
Vol. 63, pp. 171-177 (1986).
17. de Jesus, E.G., Hirano, T., and Inui, Y., “Changes in
Cortisol and Thyroid Hormone Concentrations during
Early Development and Metamorphosis in the Japanese
Flounder, Paralichthys olivaceus,” General and Comparative Endocrinology, Vol. 82, pp. 369-376 (1991).
18. de Jesus, E.G., Hirano, T., and Inui, Y., “Gonadal
Steroids Delay Spontaneous Flounder Metamorphosis
and Inhibit T3-Induced Fin Ray Shortening in vitro,”
Zoological Science, Vol. 9, pp. 633-638 (1992).
19. de Jesus, E.G., Hirano, T., and Inui, Y., “Flounder
Metamorphosis: Its Regulation by Various Hormones,”
Fish Physiology and Biochemistry, Vol. 11, pp. 323328 (1993).
20. de Jesus, E.G., Hirano, T., and Inui, Y., “The
Antimetamorphic Effect of Prolactin in the Japanese
Flounder,” General and Comparative Endocrinology,
Vol. 93, pp. 44-50 (1993).
21. de Roux, N., Genin, E., Carel, J.C., Matsuda, F.,
Chaussain, J.L., and Milgrom, E., “Hypogonadotropic
Hypogonadism due to Loss of Function of the KiSS1Derived Peptide Receptor GPR54,” Proceedings of the
National Academy of Science U.S.A., Vol. 100, pp.
10972-6 (2003).
22. Denver, R.J., “Acceleration of Anuran Amphibian
Metamorphosis by Corticotropin Releasing Hormonelike Peptides,” General and Comparative
Endocrinology, Vol. 91, pp. 38-51 (1993).
23. Denver, R.J., “Evolution of the Corticotropin-Releasing Hormone Signaling System and its Role in Stressinduced Phenotypic Plasticity,” Annals of the New York
Academy of Sciences, Vol. 897, pp. 46-53 (1999).
24. Denver, R.J. and Licht, P., “Neuropeptide Stimulation
of Thyrotropin Secretion in the Larval Bullfrog: Evidence for a Common Regulator of Thyroid and
Interregnal Activity during Metamorphosis,” Journal
of Experimental Zoology, Vol. 252, pp. 101-104 (1989).
25. Dhillo, W.S., Chaudhri, O.B., Patterson, M., Thompson,
E.L., Murphy, K.G., Badman, M.K., McGowan, B.M.,
Amber, V., Patel, S., Ghatei, M.A., and Bloom, S.R.,
“Kisspeptin-54 Stimulates the Hypothalamic-Pituitary
Gonadal Axis in Human Males,” Journal of Clinical
Endocrinology and Metabolism, Vol. 90, pp. 66096615 (2005).
26. Dodd, M.H.I. and Dodd, J.M., “The Biology of
S. Dufour & K. Rousseau: Neuroendocrinology of Fish Metamorphosis and Puberty: Evolutionary and Ecophysiological Perspectives
Metamorphosis,” In Lofts, E. (eds), Physiology of
Amphibia, Academic Press, New York, Vol. 3, pp 467599 (1976).
27. Donaldson, E., Fagerlund, U., Higgs, D., and McBride,
J., “Hormonal Enhancement of Growth,” In Hoar, W.
S., Randall, D.J. and Brett, J.R. (eds), Fish Physiology,
Academic Press, New York, Vol. VIII,.pp. 456-597
(1979).
28. Dufour, S., Lopez, E., Le Menn, F., Le Belle, N.,
Baloche, S., and Fontaine, Y.A., “Stimulation of Gonadotropin Release and of Ovarian Development, by the
Administration of a Gonadoliberin Agonist and of
Dopamine Antagonists, in Female Silver Eel Pretreated
with Estradiol,” General and Comparative Endocrinology, Vol. 70, pp. 20-30 (1988).
29. Dufour, S., Weltzien, F.A., Sébert, M.E., Le Belle, N.,
Vidal, B., Vernier, P., and Pasqualini, C., “Dopaminergic Inhibition of Reproduction in Teleost Fishes: Ecophysiological and Evolutionary Implications,” Annals
of the New York Academy of Sciences, Vol. 1040, pp. 921 (2005).
30. Eales, J.G., Holmes, J.A., McLeese, J.M., and Youson,
J.H., “Thyroid Hormone Deiodination in Various Tissues of Larval and Upstream-Migrant Sea Lampreys,
Petromyzon marinus,” General and Comparative
Endocrinology, Vol. 106,pp. 202-210 (1997).
31. Ebling, F.J.P., “The Neuroendocrine Timing of Puberty,”
Reproduction, Vol. 129, pp. 675-683 (2005).
32. Eddy, L. and Lipner, H., “Amphibian Metamorphosis:
Role of Thyrotropin-Like Hormone,” General and Comparative Endocrinology, Vol. 29, pp. 333-336 (1976).
33. Fernandez-Fernandez, R., Martini, A.C., Navarro, V.
M., Castellano, J.M., Dieguez, C., Aguilar, E., Pinilla,
L., and Tena-Sempere, M., “Novel Signals for the
Integration of Energy Balance and Reproduction,”
Molecular and Cellular Endocrinology, Vol. 254-255,
pp. 127-132 (2006).
34. Fontaine, M., “Physiological Mechanisms in the Migration of Marine and Amphihaline Fish,” Advances in
Marine Biology, Vol. 13, pp. 241-255 (1975).
35. Forey, P. and Janvier, P., “Agnathans and the Origin of
Jawed Vertebrates,” Nature, Vol. 361, pp. 129-134
(1993).
36. Forey, P. and Janvier, P., “Evolution of the Early
Vertebrates,” American Scientist, Vol. 82, pp. 554-565
(1994).
37. Fossi, M.C. and Marsili, L., “Effects of Endocrine
Disrupors in Aquatic Mammals,” Pure and Applied
Chemistry, Vol. 75, pp. 2235-2247 (2003).
38. Frisch, R. and McArthur, J., “Menstrual Cycles: Fatness as a Determinant of Minimum Weight for Height
Necessary for Their Maintenance or Onset,” Science,
Vol. 185, pp. 949-951 (1974).
39. Frisch, R. and Revelle, R., “Height and Weight at
63
Menarche: a Hypothesis of Critical Body Weights and
Adolescent Events,” Science, Vol. 169, pp. 397-399
(1970).
40. Funes, S., Hedrick, J.A., Vassileva, G., Markowitz, L.,
Abbondanzo, S., Golovko, A., Yang, S., Monsma, F.J.,
and Gustafson, E.L., “The KiSS-1 Receptor GPR54 is
Essential for the Development of the Murine Reproductive System,” Biochemical and Biophysical Research
Communications, Vol. 312, pp. 1357-63 (2003).
41. Garcia, M.C., Lopez, M., Alavarez, C.V., Casanueva,
F., Tena-Sempere, M. and Dieguez, C., “Role of Ghrelin
in Reproduction,” Reproduction, Vol. 133, pp. 531-540
(2007).
42. Gardiner, D., Ndayibagira, A., Grun, F., and Blumberg,
B., “Deformed Frogs and Environmental Retinoids,”
Pure and Applied Chemistry, Vol. 75, pp. 2263-2273
(2003).
43. Gerlach, T. and Aurich, J.E., “Regulation of Seasonal
Reproductive Activity in the Stallion, Ram and Hamster,” Animal Reproduction Science, Vol. 58, pp. 197213 (2000).
44. Gottsch, M.L., Cunningham, M.J., Smith, J.T., Popa,
Sm., Acohido, B.V., Crowley, W.F., Seminara, S.,
Clifton, D.K., and Steiner, R.A., “A Role for Kisspeptins
in the Regulation of Gonadotropin Secretion in the
Mouse,” Endocrinology, Vol. 145, pp. 4073-4077
(2004).
45. Gudernatsch, J.F., “Feeding Experiments on Tadpoles.
I. The Influence of Specific Organs Given as Food on
Growth and Differentiation. A Contribution to the
Knowledge of Organs with Internal Secretion,” Archiv
für Entwicklungsmechanik der Organismen, Vol. 35,
pp. 457-483 (1912).
46. Guillette, L.J. and Iguchi, T., “Contaminant-Induced
Endocrine and Reproductive Alterations in Reptiles,”
Pure and Applied Chemistry, Vol. 75, pp. 2275-2286
(2003).
47. Han, Y.-S., Liao, I.-C., Tzeng, W.-N., Huang, Y.-S.,
and Yu, J. Y.-L., “Serum Estradiol-17β and Testosterone Levels during Silvering in Wild Japanese Eel
Anguilla Japonica,” Comparative Biochemistry and
Physiology, Vol. 136B, pp. 913-920 (2003).
48. Han, Y.-S., Liao, I.-C., Tzeng, W.-N., and Yu, J. Y.-L.,
“Cloning of the cDNA for Thyroid Stimulating Hormone β Subunit and Changes in Activity of the Pituitary-thyroid Axis during Silvering of the Japanese
Eel,” Journal of Molecular Endocrinology, Vol. 32, pp.
179-194 (2004).
49. Higgs, D.A., Fagerland, U.H.M., Eales, J.G., McBride,
R.E., “Application of Thyroid and Steroid Hormones as
Anabolic Agents in Fish Culture,” Comparative Biochemistry and Physiology, Vol. 73B, pp. 143-176 (1982).
50. Hiroi, J., Sakakura, Y., Tagawa, M., Seikai, T., and
Tanaka, M., “Developmental Changes in Low-Salinity
64
Special Issue (2007)
Tolerance and Responses of Prolactin, Cortisol, and
Thyroid Hormones to Low-Salinity Environment in
Larvae and Juveniles of Japanese Flounder, Paralichthys
olivaceus,” Zoological Science, Vol. 14, pp. 987-992
(1997).
51. Hoar, W.S., “The Thyroid Gland of the Atlantic
Salmon,” Journal of Morphology, Vol. 65, pp. 257-295
(1939).
52. Hoar, W.S., “Smolt Transformation: Evolution,
Behaviour and Physiology,” Journal of the Fisheries
Research Board of Canada, pp. 1233-1252 (1976).
53. Hoar, W.S., “The Physiology of Smolting Salmonids,”
In Hoar, W.S. and Randall, D.J. (eds), Fish Physiology,
Academic Press, New York, Vol. 11B, pp. 275-343
(1988).
54. Hoheisel, G. and Sterba, G., “Uber die Wirkung von
Kaliumperchlorat (KClO4) auf Ammocoeten von
Lampetra planeri Bloch,” Zeitschrift Fur MikroskopischAnatomische Forschung, Vol. 70, pp. 490-516 (1963).
55. Holland, M.C., Hassin, S., and Zohar, Y., “Effects of
Long-Term Testosterone, Gonadotropin-Releasing
Hormone Agonist, and Pimozide Treatments on Gonadotropin II levels and Ovarian Development in Juvenile
Female Striped bass (Morone saxatilis),” Biology
Reproduction, Vol. 59, pp. 1153-1162 (1998).
56. Huang, Y.S., “Rôle des Stéroides Sexuels et des Hormones Métaboliques dans le Contrôle Direct
Hypophysaire de l’hormone Gonadotrope (GtH-II) chez
l’anguille Européenne, Anguilla anguilla,” PhD thesis,
University Paris VI (1998).
57. Huang, L., Schreiber, A.M., Soffientino, B., Bengtson,
D.A., and Specker, J.L., “Metamorphosis of Summer
Flounder (Paralichthys dentatus): Thyroid Status and
the Timing of Gastric Gland Formation,” Journal of
Experimental Zoology, Vol. 280, pp. 413-420 (1998).
58. Huang, Y.S., Rousseau, K., Le Belle, N., Vidal, B.,
Burzawa-Gérard, E., Marchelidon, J., and Dufour, S.,
“Insulin-Like Growth Factor-I Stimulates Gonadotropin Production from Eel Pituitary Cells: a Possible
Metabolic Signal for Induction of Puberty,” Journal of
Endocrinology, Vol. 159, pp. 43-52 (1998).
59. Huang, Y.S., Rousseau, K., Sbaihi, M., Le Belle, N.,
Schmitz, M., Dufour, S., “Cortisol Selectively Stimulates Pituitary Gonadotropin β-Subunit in a Primitive
Teleost, Anguilla anguilla,” Endocrinology, Vol. 140,
pp. 1228-1235 (1999).
60. I’Anson, H., Foster, D.L., Foxcroft, G.R., and Booth, P.
J., “Nutrition and Reproduction,” Oxford. Reviews of
Reproductive Biology, Vol. 13, pp. 239-311 (1991).
61. Inui, Y. and Miwa. S., “Thyroid Hormone Induces
Metamorphosis of Flounder Larvae,” General and Comparative Endocrinology, Vol. 60, pp. 450-454 (1985).
62. Inui, Y., Tagawa, M., Miwa, S., and Hirano, T., “Effects
of Bovine TSH on the Tissue Thyroxine Level and
Metamorphosis in Prometamorphic Flounder Larvae,”
General and Comparative Endocrinology, Vol. 74, pp.
406-410 (1989).
63. Jobling, S. and Tyler, C.R., “Endocrine Disruption in
Wild Freshwater Fish,” Pure Applied Chemistry, Vol.
75, pp. 2219-2234 (2003).
64. Kah, O., Chambolle, P., Thibault, J., and Geffard, M.,
“Existence of Dopaminergic Neurons in the Preopric
Region of the Goldfish,” Neuroscience Letters, Vol. 48,
pp.293-298 (1984).
65. Kah, O., Dubourg, P., Onteniente, B., Geffard, M., and
Calas, A., “The Dopaminergic Innervation of the Goldfish Pituitary. An Immunocytochemical Study at the
Electron-Microscope Level using Antibodies Against
Dopamine,” Cell and Tissue Research, Vol. 244, pp.
577-582 (1986).
66. Kah, O., Dulka, J.G., Dubourg, P., Thibault, J., and
Peter, R.E., “Neuroanatomical Substrate for the Inhibition of Gonadotrophin Secretion in Goldfish: Existence
of a Dopaminergic Preoptico-Hypophyseal Pathway,”
Neuroendocrinology, Vol. 45, pp. 451-458 (1987).
67. Kanamori, A. and Brown, D.D., “The Analysis of
Complex Developmental Programmes: Amphibian
Metamorphosis,” Genes to Cells, Vol. 1, pp. 429-435
(1996).
68. Kennedy, G.C. and Mitra, J., “Body Weight and Food
Intake as Initiating Factors for Puberty in the Rat,”
Journal of Physiology, Vol. 166, pp. 408-418 (1963).
69. Kitajima, C., Sato, T., and Kawanishi, M., “On the
Effect of Thyroxine to Promote the Metamorphosis of
a Conger Eel-Preliminary Report,” Bulletin of the Japanese Society of Scientific Fisheries, Vol. 33, pp. 919922 (1967).
70. Kikuyama, S., Kawamura, K., Tanaka, S., and
Yamamoto, K., “Aspects of Amphibian Metamorphosis, Hormonal Control,” International Review of
Cytology, Vol. 145, pp. 105-148 (1993).
71. Kubota, S.S., “Studies on the Ecology, Growth and
Metamorphosis in Conger Eel, Conger myriaster
(Brevoort),” Journal of Faculty of Fish Prefectural
University of Mie, Vol. 5, pp. 190-370 (1961).
72. Kumakura, N., Okuzawa, K., Gen, K., and Kagawa, H.,
“Effects of Gonadotropin-Releasing Hormone Agonist
and Dopamine Antagonist on Hypothalamus-Pituitarygonadal Axis of Prepubertal Female Eed Seabream
(Pagrus major),” General and Comparative Endocrinology, Vol. 131, pp. 264-273 (2003).
73. Larsen, D.A., Swanson, P., Dickey, J.T., Rivier, J., and
Dickhoff, W.W., “In Vitro Thyrotropin-Releasing Activity of Corticotropin-Releasing Hormone-Family
Peptides in Coho Salmon, Oncorhynchus kisutch,”
General and Comparative Endocrinology, Vol. 109,
pp. 276-285 (1998).
74. Leatherland, J.F., Hilliard, R.W., Macey, D.J., and
S. Dufour & K. Rousseau: Neuroendocrinology of Fish Metamorphosis and Puberty: Evolutionary and Ecophysiological Perspectives
Potter, I.C., “Changes in Serum Thyroxine and Triiodothyronine Concentrations during Metamorphosis
of the Southern Hemisphere Lamprey Geotria australis,
and the Effect of Propylthiouracil, Triiodothyronine
and Environmental Temperature on Serum Thyroid
Hormone Concentrations of Ammocoetes,” Fish Physiology and Biochemistry, Vol. 8, pp. 167-177 (1990).
75. Lin, H.R., Van der Kraak, G., Zhou, X.J., Liang, J.Y.,
Peter, R.E., Rivier, J.E., and Vale, W.W., “Effects of
[D-Arg6, Trp7, Leu8, Pro9NEt]-Luteinizing HormoneReleasing Hormone (sGnRH-a) and [D-Ala6, Pro9NEt]Luteinizing Hormone-Releasing Hormone (LHRH-a),
in Combination with Pimozide or Domperidone, on
Gonadotropin Release and Ovulation in the Chinese
Loach and Common Carp,” General and Comparative
Endocrinology, Vol. 69, pp. 31-40 (1998).
76. Linard, B., Bennami, S., and Saligaut, C., “Involvement
of Estradiol in a Catecholamine Inhibitory Tone of
Gonadotropin Release in Rainbow Trout (Oncorhynchus
mykiss),” General and Comparative Endocrinology,
Vol. 99, pp. 192-196 (1995).
77. Lintlop, S.P. and Youson, J.H., “Concentration of
Triidothyronine in the Sera of the Sea Lamprey,
Petromyzon Marinus, and the Brook Lamprey, Lampetra
Lamottenii at Various Phases of Their Life Cycle,”
General and Comparative Endocrinology, Vol. 49, pp.
187-194 (1983).
78. Lintlop, S.P. and Youson, J.H., “Binding of Triiodothyronine to Hepatocyte Nuclei from Sea Lamprey,
Petromyzon marinus L., at Various Stages of the Life
Cycle,” General and Comparative Endocrinology, Vol.
49, pp. 428-436 (1983).
79. Lokman, P.M., Vermeulen, G.J., Lambert, J.G.D., and
Young, G., “Gonad Histology and Plasma Steroid Profiles in Wild New Zealand Freshwater Eels (Anguilla
dieffenbachii and A. australis) before and at the Onset
of the Natural Spawning Migration. I. Females,” Fish
Physiology and Biochemistry, Vol. 19, pp. 325-338
(1998).
80. Manzon, R.G. and Denver, R.J., “Regulation of Pituitary Thyrotropin Gene Expression during Xenopus
Metamorphosis: Negative Feedback is Functional
Throughout Metamorphosis,” Journal of Endocrinology, Vol. 182, pp. 273-285 (2004).
81. Manzon, R.G. and Youson, J.H., “The Effects of Exogenous Thyroxine (T4) or Triiodothyronine (T3) in the
Presence or Absence of Potassium Perchlorate, on the
Incidence of Metamorphosis and on Serum T4 and T3
Concentrations in Larval Sea Lampreys (Petromyzon
marinus L.).” General and Comparative Endocrinology,
Vol. 106, pp. 211-220 (1997).
82. Matsui, H., Takatsu, Y., Kumano, S., Matsumoto, H.,
and Ohtaki, T., “Peripheral Administration of Metastin
Induces Marked Gonadotropin Release and Ovulation
65
in the Rat,” Biochemical and Biophysical Research
Communications, Vol. 320, pp. 383-388 (2004).
83. Matthiessen, P., “Endocrine Disruption in Marine
Fish,” Pure Applied Chemistry, Vol. 75, pp. 2249-2261
(2003).
84. McBride, J.R., Higgs, D.A., Fagerlund, U.H.M.,
Buckley, J.T., “Thyroid Hormones and Steroid
Hormones: Potential for Control of Growth and
Smoltification of Salmonids,” Aquaculture, Vol. 28,
pp. 201-210 (1982).
85. McNabb, F.M.A., Thyroid Hormones, Englewood
Cliffs, Prentice Hall, pp. 283 (1992).
86. Messager, S., Chatzidaki, E.E., Ma, D., Hendrick, A.G.,
Zahn, D., Dixon, J., Thresher, R.R., Malinge, I., Lomet,
D., Carlton, M.B., Colledge, W.H., Caraty, A., and
Aparicio, S.A., “Kisspeptin Directly Stimulates Gonadotropin-Releasing Hormone Release Via G ProteinCoupled Receptor 54,” Proceedings of the National
Academy of Science U.S.A., Vol. 102, pp. 1761-6 (2005).
87. Miwa, S. and Inui, Y., “Histological Changes in the
Pituitary-Thryoid Axis during Spontaneous and Artificially-Induced Metamorphosis of Larvae of the Flounder Paralichtys olivaceus,” Cell and Tissue Research,
Vol. 249, pp.117-123 (1987).
88. Miwa, S. and Inui, Y., “Effects of Various doses of
Thyroxine and Triidothyronine on the Metamorphosis
of Flounder (Paralichtys olivaceus),” General and Comparative Endocrinology, Vol. 67, pp. 356-363 (1987).
89. Miwa, S. and Inui, Y., “Thyroid Hormone Stimulates
the Shift of Erythrocyte Populations during Metamorphosis of the Flounder,” Journal of Experimental
Zoology, Vol. 259, pp. 222-228 (1991).
90. Miwa, S., Tagawa, M., Inui, Y., and Hirano, T., “Thyroxine Surge in Metamorphosing Flounder Larvae,”
General and Comparative Endocrinology, Vol. 70, pp.
158-163 (1988).
91. Miwa, S., Yamano, K., and Inui, Y., “Thyroid Hormone
Stimulates Gastric Development in Flounder Larvae
during Metamorphosis,” Journal of Experimental
Zoology, Vol. 261, pp. 424-430 (1992).
92. Mohamed, J.S., Benninghoff, A.D., Holt, G.J., and
Khan, I.A., “Developmental Expression of the G Protein-Coupled Receptor 54 and Three GnRH mRNAs in
the Teleost Fish Cobia,” Journal of Molecular
Endocrinology, Vol. 38, pp.235-244 (2007).
93. Morley, J.E., “Neuroendocrine Control of Thyrotropin
Secretion,” Endocrine Reviews, Vol. 2, pp. 396-436
(1981).
94. Müller, A., “On the Development of Lampreys,” Annals and Magazine of Natural History, Vol. 18, pp. 298301 (1856).
95. Murphy, K.G., “Kisspeptins: Regulators of Metastasis
in the Hypothalamic-Pituitary-Gonadal Axis,” Journal
of Neuroendocrinol, Vol. 17, pp. 519-525 (2005).
66
Special Issue (2007)
96. Murr, E. and Sklower, A., “Untersuchungen über die
Inkretorischen Organe der Fische. I. Das Verhalten der
Schilddrüse in der Metamorphose des Aales,” Zeitschr
Vergleichende Physiologie, Vol. 7, pp. 279-288 (1928).
97. Navarro, V.M., Castellano, J.M., Fernandez-Fernandez,
R., Barreiro, M.L., Roa, J., Sanchez-Criado, J.E.,
Aguilar, E., Dieguez, C., Pinilla, L., and Tena-Sempere,
M., “Developmental and Hormonally Regulated Messenger Ribonucleic Acid Expression of KiSS-1 and its
Putative Receptor, GPR54, in Rat Hypothalamus and
Potent Luteinizing Hormone-Releasing Activity of
KiSS-1 Peptide,” Endocrinology, Vol. 145, pp.
4565-74 (2004).
98. Nocillado, J.N., Levavi-Sivan, B., Carrick, F., Elizur,
A., “Temporal Expression of G-Protein-Coupled Receptor 54 (GPR54), Gonadotropin-Releasing Hormones
(GnRH), and Dopamine Receptor D2 (drd2) in Pubertal
Female Grey Mullet, Mugil cephalus,” General and
Comparative Endocrinology, Vol. 150, pp. 278-287
(2007).
99. Oehlmann, J. and Schulte-Oehlmann, U., “Endocrine
Disruption in Invertebrates,” Pure Applied Chemistry,
Vol. 75, pp. 2207-2218 (2003).
100. Okada, N., Tanaka, M., and Tagawa, M., “Bone Development during Metamorphosis of the Japanese Flounder (Paralichthys olivaceus): Differential Responses to
Thyroid Hormone,” In Browman, H.I. and Skiftesvik,
A.B., (eds), The Big Fish Bang: Proceedings of the 26th
Annual Larval Fish Conference, Institute of Marine
Research, Bergen, Norway (2003).
101. Okuzawa, K., “Puberty in Teleosts,” Fish Physiology
and Biochemistry, Vol. 26, pp. 31-41 (2002).
102. Olivereau, M. and Olivereau, J., “Effects of 17 αMethyltestosterone on the Skin and Gonads of Freshwater Male Silver Eels,” General and Comparative
Endocrinology, Vol. 57, pp. 64-71 (1985).
103. Parhar, I.S., Ogawa, S., and Sakuma, Y., “Laser-Captured Single Digoxigenin-Labeled Neurons of Gonadotropin-Releasing Hormone Types Reveal a Novel G
Protein-Coupled Receptor (Gpr54) during Maturation
in Cichlid Fish,” Endocrinology, Vol. 145, pp.
3613-3618 (2004).
104. Pasqualini, C., Vidal, B., Le Belle, N., Sbaihi, M.,
Weltzien, F.-A., Vernier, P., Zohar, Y., and Dufour, S.,
“Un Contre-Pouvoir au Contrôle de la Reproduction
Par la GnRH chez Les Poisons Téléostéens: l’Inhibition
Dopaminergique. Rôle Ancestral et Conservation
Différentielle Chez les Vertébrés? Journal de la Société
de Biologie, Vol. 198, pp. 61-67 (2004).
105. Peter, R.E. and Crim, L.W., “Hypothalamic Lesions of
Goldfish: Effects on Gonadal Recrudescence and Gonadotropin Secretion,” Aun Biol Anim Biochim Biophys,
Vol. 18, pp. 819-823 (1978).
106. Peter, R.E. and Paulencu, C.R., “Involvement of the
Preoptic Region in the Gonadotropin Release-Inhibition in the Goldfish,” Neuroendocrinology, Vol. 31,
133-141 (1980).
107. Peter, R.E., Crim, L.W., Goos, H.J.Th., and Crim, J.W.,
“Lesioning Studies on the Gravid Female Goldfish:
Neuroendocrine Regulation of Ovulation,” General
and Comparative Endocrinology, Vol. 35, pp. 391-401
(1978).
108. Pompolo, S., Pereira, A., Estrada, K.M., and Clarke, I.
J., “Colocalization of Kisspeptin and GonadotropinReleasing Hormone in the Ovine Brain,” Endocrinology,
Vol. 147, pp. 804-810 (2006).
109. Pradet-Balade, B., “Evolution de la Regulation de la
Fonction Thyréotrope: Étude chez les Téléostéens,”
PhD Thesis University Paris XI (1998).
110. Prunet, P., Boeuf, G., Bolton, J.P., and Young, G.,
“Smoltification and Seawater Adaptation in Atlantic
Salmon (Salmo salar): Plasma Prolactin, Growth Hormone and Thyroid Hormones,” General and Comparative Endocrinology, Vol. 74, pp. 355-364 (1989).
111. Revel, F.G., Ansel, L., Klosen, P., Saboureau, M.,
Pévet, P., Mikkelsen, J.D., and Simmoneaux, V.,
“Kisspeptin: A Key Link to Seasonal Breeding,”
Reviews in Endocrine and Metabolic Disorders, Vol. 8,
pp. 57-65 (2007).
112. Richman, N.H., de Diaz, S.T., Nishioka, R.S., and Bern,
H.A., “Developmental Study of Coho Gill Functional
Morphology and the Effects of Cortisol,” Aquaculture,
Vol. 45, pp. 386-387 (1985).
113. Robertson, O.G., “Production of Silvery Smolt Stage in
Rainbow Trout by Intramuscular Injection of Mammalian Thyroid Extract and Thyrotropic Hormone,” Journal of Experimental Zoology, Vol. 110, pp. 337-355
(1949).
114. Rohr, D.H., Lokman, P.M., Davie, P.S., and Young, G.,
“11-Ketotestosterone Induces Silvering-Related
Changes in Immature Female Short-Finned Eels,
Anguilla australis,” Comparative Biochemistry and
Physiology, Vol. 130A, pp. 701-714 (2001).
115. Romeo, R.D., “Puberty: a Period of Both Organizational and Activational Effects of Steroid Hormones on
Neurobehavioural Development,” Journal of
Neuroendocrinol, Vol. 15, pp. 1185-1192 (2003).
116. Rousseau, K. and Dufour, S., “Endocrinology of Migratory Fish Life Cycle in Special Environments: the
Role of Metamorphoses,” In Sébert, P., Onyango, D.
W., and Kapoor, B.G. (eds), Fish Life in Special
Environments, Science Publishers, UK, pp. 193-231
(2007). (in press)
117. Rousseau, K., Huang, Y.S., Le Belle, N., Vidal, B.,
Marchelidon, J., Epelbaum, J., and Dufour, S., “Longterm Inhibitory Effects of Somatostatin and InsulinLike Growth Factor 1 on Growth Hormone Release by
Serum-Free Primary Culture of Pituitary Cells from
S. Dufour & K. Rousseau: Neuroendocrinology of Fish Metamorphosis and Puberty: Evolutionary and Ecophysiological Perspectives
European eel (Anguilla anguilla),” Neuroendocrinology,
Vol. 67, pp. 301-309 (1998).
118. Saligaut, C., Linard, B., Breton, B., Anglade, I.,
Bailhache, T., Kah, O., and Jego, P., “Brain Aminergic
Systems in Salmonids and Other Teleosts in Relation to
Steroid Feedback and Gonadotropin Release,”
Aquaculture, Vol. 177, pp. 13-20 (1999).
119. Sbaihi, M., “Interaction des Stéroïdes Sexuels et du
Cortisol dans le Contrôle de la Reproduction et du
métabolisme Calcique Chez un Téléostéen Migrateur,
l’anguille (Anguilla anguilla),” PhD Thesis University
Paris VI (2001).
120. Sbaihi, M., Fouchereau-Peron, M., Meunier, F., Elie,
P., Mayer, I., Burzawa-Gérard, E., Vidal, B., and Dufour,
S., “Reproductive Biology of the Conger Eel from the
South Coast of Brittany, France and Comparison with
the European Eel,” Journal of Fish Biology, Vol. 59, pp.
302-318 (2001).
121. Schreiber, A.M. and Specker, J.L., “Metamorphosis in
the Summer Flounder, Paralichthys dentatus: Thyroidal Status Influences Gill Mitochondria-Rich Cells,”
General and Comparative Endocrinology, Vol. 117,
pp. 238-250 (2000).
122. Seminara, S.B., “We All Remember our First Kiss:
Kisspeptin and the Male Gonadal Axis,” Journal of
Clinical Endocrinology and Metabolism, Vol. 90, pp.
6738-6740 (2005).
123. Seminara, S.B., Messager, S., Chatzidaki, E.E., Thresher,
R.R., Acierno, J.S. Jr., Shagoury, J.K., Bo-Abbas, Y.,
Kuohung, W., Schwinof, K.M., Hendrick, A.G., Zahn,
D., Dixon, J., Kaiser, U.B., Slaugenhaupt, S.A., Gusella,
J.F., O’Rahilly, S., Carlton, M.B., Crowley, W.F. Jr.,
Aparicio, S.A., and Colledge, W.H., “The GPR54 Gene
as a Regulator of Puberty,” The New England Journal
of Medicine, Vol. 349, pp. 1614-27 (2003).
124. Shahab, M., Mastronardi, C., Seminara, S.B., Crowley,
W.F., Ojeda, S.R., and Plant, T.M., “Increased Hypothalamic GPR54 Signaling: a Potential Mechanism for
Initiation of Puberty in Primates,” Proceedings of the
National Academy of Science U.S.A., Vol. 102, pp.
2129-34 (2005).
125. Sheridan, M.A. and Kao, Y.-H., “Regulation of Metamorphosis-Associated Changes in the Lipid Metabolism of Selected Vertebrates,” American Zoologist,
Vol. 38, pp. 350-368 (1998).
126. Shi, Y.-B., Amphibian Metamorphosis, From Morphology to Molecular Biology. John Wiley, New York
(1999).
127. Sisk, C.L. and Foster, D.L., “The Neural Basis of
Puberty and Adolescence,” Nature Neuroscience, Vol.
7, pp.1040-1047 (2004).
128. Sklower, A., “Die Bedeutung der Schilddruse Fur die
Metamorphose des Aales und der Plattfische,” Forsch
Fortschr Dtsch Wiss, Vol. 6, pp. 435-436 (1930).
67
129. Smith, J.T., Clifton, D.K., and Steiner, R.A., “Regulation of the Neuroendocrine Reproductive Axis by
Kisspeptin-GPR54 Signaling,” Reproduction, Vol. 131,
pp. 623-630 (2006).
130. Smith, J.T., Dungan, H.M., Stoll, E.A., Gottsch, M.L.,
Braun, R.E., Eacker, S.M., Clifton, D.K., and Steiner,
R.A., “Differential Regulation of Kiss-1 mRNA Expression by Sex Steroids in the Brain of the Male
Mouse,” Endocrinology, Vol. 146, pp. 2976-2984
(2005).
131. Soffientino, B. and Specker, J.L., “Metamorphosis of
Summer Flounder, Paralichthys dentatus: Cell Proliferation and Differentiation of the Gastric Mucosa and
Developmental Effects of Altered Thyroidal Status,”
Journal of Experimental Zoology, Vol. 290, pp. 31-40
(2001).
132. Solbakken, J.S., Norberg, B., Watanabe, K., and Pittman,
K., “Thyroxine as a Mediator of Metamorphosis of
Atlantic Halibut, Hippoglossus hippoglossus,” Environmental Biology of Fishes, Vol. 1-2, pp. 53-65 (1999).
133. Specker, J.L., “Interrenal Function and Smoltification,” Aquaculture, Vol. 28, pp. 59-66 (1982).
134. Sterba, G. and Schneider, J., “Zur Wirkung von Kalliumperchlorat (KClO4) auf Ammocoeten,” Naturwissenschaften, Vol. 48, pp. 485-486 (1961).
135. Sullivan, C.V., Darling, D.S., and Dickhoff, W.W.,
“Effects of Triiodothyronine and Propylthiouracil on
Thyroid Function and Smoltification of Coho Salmon
(Oncorhynhcus kisutch),” Fish Physiology and
Biochemistry, Vol. 4, pp. 121-135 (1987).
136. Suzuki, S., “Induction of Metamorphosis and Thyroid
Function in the Larval Lamprey,” In Mederios-Neto, G.
and Gaitan, E. (eds), Frontiers in Thyroidology, Plenum,
New York, Vol. 1, pp. 667-670 (1986).
137. Sweeting, R.M. and McKeown, B.A., “Changes in
Plasma Growth Hormone and Various Metabolic Factors during Smoltification of Coho Salmon,
Oncorhynchus kisutch,” Aquaculture, Vol. 82, pp. 279295 (1989).
138. Sweeting, R.M., Wagner, G.F., and McKeown, B.A.,
“Changes in Plasma Glucose, Amino Acid, Nitrogen
and Growth Hormone during Smoltification and Seawater Adaptation in Coho Salmon, Oncorhynchus
kisutch,” Aquaculture, Vol. 45, pp. 185-197 (1985).
139. Tagawa, M., Miwa, S., Inui, Y., de Jesus, E.G., and
Hirano, T., “Changes in Thyroid Hormone Concentrations during Early Development and Metamorphosis of
the Flounder, Paralichthys olivaceus,” Zoological
science, Vol. 7, pp. 93-96 (1990).
140. Tata, J.R., “Hormonal Interplay and Thyroid Hormone
Receptor Expression during Amphibian Metamorphosis,” In Gilbert, L.I., Tata, J.R., and Atkinson, B.G.
(eds), Metamorphosis. Postembryonic Reprogramming
of Gene Expression in Amphibian and Insect Cells,
68
Special Issue (2007)
Academic Press, San Diego (1996).
141. Tata, J.R., Hormonal Signalling and Postembryonic
Development, Springer, Berlin (1998).
142. Tata, J.R., “Amphibian Metamorphosis as a Model for
the Developmental Actions of Thyroid Hormone,”
Molecular and Cellular Endocrinology, Vol. 246, pp.
10-20 (2006).
143. Thiéry, J.C., Chemineau, P., Hernandez, X., Migaud,
M., and Malpaux, B., “Neuroendocrine Interactions
and Seasonality,” Domestic Animal Endocrinology,
Vol. 23, pp. 87-100 (2002).
144. Thompson, E.L., Patterson, M., Murphy, K.G., Smith,
K.L., Dhillo, W.S., Todd, J.F., Ghatei, M.A., and Bloom,
S.R., “Central and Peripheral Administration of
Kisspeptin-10 Stimulates the Hypothalamic-PituitaryGonadal Axis,” Journal of Neuroendocrinol, Vol. 16,
pp. 850-8 (2004).
145. Van Asselt, L.A., Goos, H.J., Smit-van Dijk, W.,
Speetjens, P.A.M., Van Oordt, P.G., “Evidence for the
Involvement of D2 Receptors in the Dopaminergic
Inhibition of Gonadotropin Release in the African
Catfish, Clarias gariepinus,” Aquaculture, Vol. 72, pp.
369-378 (1988).
146. Van der Kraak, G., Donaldson, E.M., and Chang, J.P.,
Dopamine Involvement in the Regulation of Gonadotropin Release in Coho Salmon,” Canadian Journal of
Zoology, Vol. 64, pp. 1245-1248 (1986).
147. Veldhuis, J.D., Roemmich, J.N., Richmond, E.J., and
Bowers, C.Y., “Somatotropic and Gonadotropic Axes
Linkages in Infancy, Childhood and the Puberty-Adult
Transition,” Endocrine Reviews, Vol. 27, pp. 101-140
(2006).
148. Vidal, B., Pasqualini, C., Le Belle, N., Holland, M.C.,
Sbaihi, M., Vernier, P., Zohar, Y., and Dufour, S.,
“Dopamine Inhibits Luteinizing Hormone Synthesis
and Release in the Juvenile European Eel: a Neuroendocrine Lock for the Onset of Puberty,” Biology
Reproduction, 71, pp. 1491-500 (2004).
149. Vilter, V., “Action de la Thyroxine Sur la Metamorphose Larvaire de l’Anguille,” Comptes Rendus des
Seances de la Societe de Biologie et des Ses Filiales,
Vol. 140, pp. 783-785 (1946).
150. Virtanen, E. and Soivio, A., “The Patterns of T3, T4,
Cortisol and Na+-K+ ATPase during Smoltification of
Hatchery Reared Salmo salar and Comparison with
wild Smolts,” Aquaculture, Vol. 45, pp. 97-109 (1985).
151. White, B.A. and Nicoll, C.S., “Hormonal Control of
Amphibian Metamorphosis,” In Gilvert, L.I. and
Frieden, E. (eds), Metamorphosis, Plenum Press: New
York, pp. 363-396 (1981).
152. Wright, G.M. and Youson, J.H., “Serum Thyroxine
Concentrations in Larval and Metamorphosing Anadromous Sea Lamprey, Petromyzon marinus L,” Journal
of Experimental Zoology, Vol. 202, pp. 27-32 (1977).
153. Yamano, K., Tagawa, M., de Jesus, E.G., Hirano, T.,
Miwa, S., and Inui, Y., “Changes in Whole Body
Concentrations of Thyroid Hormones and Cortisol in
Metamorphosing Conger Eel,” Journal of Comparative
Physiology B, Vol. 161, pp. 371-375 (1991).
154. Yaron, Z., Gur, G., Melamed, P., Rosenfeld, H., Elizur,
A., and Levavi-Sivan, B., “Regulation of Fish
Gonadotropins,” International Review of Cytology, Vol.
225, pp. 131-185 (2003).
155. Young, G., Björnsson, B.T., Prunet, P., Lin, R.J., and
Bern, H.A., “Smoltification and Seawater Adaptation
in Coho Salmon (Oncorhynchus kisutch): Plasma
Prolactin, Growth Hormone, Thyroid Hormone and
Cortisol,” General and Comparative Endocrinology,
Vol. 74, pp. 335-345 (1989).
156. Youson, J.H., “The Morphology and Physiology of
Lamprey Metamorphosis,” Canadian Journal of Fisheries and Aquatic Sciences, Vol.37, pp. 687-710 (1980).
157. Youson, J.H., “First Metamorphosis,” In Hoar, W.S.
and Randall, D.J. (eds), Fish Physiology, Vol. XI,
Physiology of Developing Fish, Part B, Viviparity and
Posthatching Juveniles, Academic Press, San Diego, pp
135-196 (1988).
158. Youson, J.H., “Is Lamprey Metamorphosis Regulated
by Thyroid Hormones?” American Zoologist, Vol. 37,
pp. 441-460 (1997).
159. Youson, J.H., Plisetskaya, E.M., and Leatherland, J.F.,
“Concentrations of Insulin and Thyroid Hormones in
Serum of Landlocked Sea Lampreys (Petromyzon
marinus) of Three Larval Year Classes, Larvae Exposed to Two Temperature Regimes, and Both during
and after Metamorphosis,” General and Comparative
Endocrinology, Vol.94, pp. 294-304 (1994).
160. Youson, J.H., Manzon, R.G., Peck, B.J., and Holmes, J.
A., “Effects of Exogenous Thyroxine (T4) and Triiodothyronine (T3) on Spontaneous Metamorphosis
and Serum T4 and T3 Levels in Immediately Premetamorphic Sea Lampreys, Petromyzon marinus,” Journal
of Experimental Zoology, Vol. 279, pp. 145-155 (1997).
161. Zohar, Y., Harel, M., Hassin, S., and Tandler, A.,
“Broodstock Management and Manipulation of Spawning in the Gilthead Seabream, Sparus Aurata,” In
Bromage, N. and Roberts, R.J. (eds), Broodstock Management and Egg and Larval Quality, Blackwell Scientific Press, London, pp. 94-117.