Journal of Reproduction and Development, Vol. 54, No. 1, 2008, 19112
Secretion of Inhibin in Female Japanese Quails (Coturnix japonica) from
Hatch to Sexual Maturity
Manila SEDQYAR1,2), Qiang WENG2,3), Gen WATANABE1,2), Mohamed M.M. KANDIEL2,4),
Sinji TAKAHASHI5), Akira K SUZUKI6), Shinji TANEDA6) and Kazuyoshi TAYA1,2)
1)Department of Basic Veterinary Science, The United Graduate School of Veterinary Sciences, Gifu University, Gifu 501-
1193, 2)Laboratory of Veterinary Physiology, Department of Veterinary Medicine, Faculty of Agriculture, Tokyo University
of Agriculture and Technology, Tokyo 183-8509, Japan, 3)College of Biological Science and Technology, Beijing Forestry
University, Beijing 100083, P.R. China, 4)Department of Theriogenology, Faculty of Veterinary Medicine, Benha University,
Benha, Egypt, 5)Ecological Effect Research Team, Dioxin and Environmental Endocrine Disrupter Research Project and
6)Environmental Nanotoxicology Section, Research Center for Environmental Risk, National Institute for Environmental
Studies, Ibaraki 305-8506, Japan
Abstract. To clarify the cellular source and secretory pattern of inhibin in the Japanese quail during follicular
development, the plasma concentrations of immunoreactive (ir) inhibin were measured from 1 to 7 weeks after
hatching. Localization of the inhibin/activin α, βA and βB subunts was investigated by immunohistochemistry. To
monitor development of the pituitary and ovarian functions, the plasma luteinizing hormone (LH) and progesterone
concentrations were also measured. Ovarian weight increased gradually until 6 weeks of age and then abruptly
increased at 7 weeks of age just at the onset of egg production. Plasma concentrations of LH increased significantly at 6
weeks of age. The plasma concentrations of ir-inhibin and progesterone and the pituitary contents of LH also increased
significantly at 7 weeks of age. Immunohistochemically, the inhibin/activin α, βA and βB subunts were localized in the
granulosa cells of all follicles during different stages of development from 1 to 7 weeks after hatching. The inhibin α, βA
and βB subunts were also found in the interstitial cells but not theca cells of all follicles. These results demonstrated that
the plasma concentrations of ir-inhibin of the female Japanese quails rose with ovarian development. The
immunohistochemical results suggested that granulosa and interstitial cells are the major source of ovarian inhibins in
female Japanese quails.
Key words: Development, Female quail, Inhibin, Luteinizing hormone (LH), Progesterone
(J. Reprod. Dev. 54: 52–57, 2008)
nhibins and activins are structurally related dimeric gonadal
proteins with the ability to regulate follicle-stimulating hor-
mone (FSH) secretion from pituitary glands in mammals [1, 2].
Inhibin consists of an α subunit linked by a disulfide bridge to one
of the 2 highly homologous β subunits (βA and βB) to form inhibin
A (α and βA) or inhibin B (α and βB) . Apart from their action
on FSH secretion, inhibins and activins have been shown to exert
paracrine/autocrine effects within the gonads [4, 5] and other tis-
sues . In mammalian females, the roles played by inhibin/
activin in modulating FSH secretion by the pituitary gland in vivo
and in vitro have been well documented. Inhibins selectively sup-
press [7–12] while activins stimulate [11–15] the release of FSH.
Previous studies have shown that the avian ovary also produces
inhibins, which plays important roles in the regulation of pituitary
FSH [16–21]. However, the roles of inhibins have received little
attention in quail. As a laboratory animal, the female Japanese
quail has been extensively used in reproductive research, because
of its adaptability to battery breeding cages, small body size, early
sexual maturation, short generation interval, regular egg laying and
high egg production. However, studies on the reproductive endo-
crinology of quails are scarce. Specifically, no research results
have been published concerning the secretion of inhibins in female
quails. In order to clarify the possible endocrine interaction of the
plasma concentrations of immunoreactive (ir) inhibins, the local-
ization of inhibin subunits was investigated in female Japanese
quails from hatching to sexual maturity. The aim of the present
study was to elucidate the relationship between inhibin secretion
and ovarian function during sexual maturation of the Japanese quail
from hatching to sexual maturity.
Materials and Methods
Female Japanese quails (Coturnix japonica) were selected from
low antibody response (L) lines for use in this study from hatching
to sexual maturity [22, 23]. Fertilized eggs were incubated using a
Showa Incubator Laboratory (Trade Mark Showa Furanki, Urawa,
Japan) under regular conditions, such as temperature (38.7 C),
humidity (55 ± 10%) and turning (once every hour), that were con-
trolled daily. One day before hatching (at day 16 of incubation),
candling was performed. The birds were provided with food
(Kanematsu quail diet; Kanematsu Agri-tech, Ibaraki, Japan) and
water and were allowed to feed ad libitum. They were housed in
metal cages in a controlled environment (lights on 0500–1900 h;
Accepted for publication: October 3, 2007
Published online: November 13, 2007
Correspondence: G. Watanabe (e-mail: Gen@cc.tuat.ac.jp)
53INHIBIN IN FEMALE JAPANESE QUAILS
temperature, 23 ± 2 C; humidity 55 ± 10%; air exchanged 20 times
hourly). This study was conducted in accordance with the Guiding
Principles in the Use of Animals in Toxicology and was approved
by the Animal Care and Use Committee of the Japanese National
Institute for Environmental Studies.
Birds were decapitated weekly and blood samples were col-
lected into heparinized tubes between 1000 h and 1200 h. The
blood samples were centrifuged at 4 C for 15 min at 1700 g. The
plasma was separated and stored at –20 C until it was assayed for
ir-inhibin, LH and progesterone by specific radioimmunoassay
(RIA). After dissection, the chicks were sexed, and their ovaries
were removed, weighed and fixed for 24 h in 4% paraformaldehyde
(Sigma-Aldrich Chemical, St. Louis, MO, USA) in 0.05 M PBS
(pH 7.4) for immunohistochemical examination.
RIA of LH, ir-inhibin and progesterone
Luteinizing hormone (LH) concentrations were measured with a
USDA-ARS RIA kit (Beltsville, MD, USA) for chicken LH. The
antiserum used was anti-avian LH (HAC-CH27-01 RBP75). The
hormone for iodination was chicken USDA-cLH-I-3. The results
were expressed in terms of USDA-cLH-K-3. The sensitivity of the
assay for LH was 15.6 pg/tube, and the intra- and interassay coeffi-
cients of variation were 5.2 and 11.2%, respectively. USDA-cLH-
I-3 and USDA-cLH-K-3 were kindly provided by Dr. John A.
Proudman, Biotechnology and Germplasm Laboratory, Animal and
Natural Resources Institute, Beltsville, MD, USA . The antise-
rum against avian LH was kindly provided by the Biosignal
Research Center, Institute for Molecular and Cellular Regulation,
Gunma University, Gunma, Japan . Plasma concentrations of
ir-inhibin were measured as described previously . The iodi-
nated preparation used was 32 kDa bovine inhibin, and the
antiserum used was rabbit antiserum against bovine inhibin
(TNDH-1). The results were expressed in terms of 32 kDa bovine
inhibin. Serial dilutions of plasma from female and male quails
were compared with the standard curve for 32 kDa bovine inhibin.
All curves obtained from the serially diluted samples were parallel
to the standard curve, indicating that it was possible to measure the
concentration of ir-inhibin in the peripheral plasma of the quails
using the present RIA system. The sensitivity of the assay for ir-
inhibin was 19.5 pg/tube of purified bovine 32 kDa inhibin, and the
intra- and interassay coefficients of variation were 8.8 and 14.4%,
respectively. The concentrations of progesterone were determined
by a double-antibody RIA system with125I-labeled radioligands as
described previously . The antiserum against progesterone
(GDN 337) was kindly provided by Dr. G.D. Niswender (Colorado
State University, Fort Collins, CO, USA). The sensitivity of the
assay for progesterone was 2.5 pg/tube, and the intra- and interas-
say coefficients of variation were 6.3 and 7.2%, respectively.
The ovaries were dehydrated through a series of graded concen-
trations of ethanol and xylene, embedded in paraffin, sectioned
serially at 4 µm, mounted on glass slides coated with 3-aminopro-
pyltriethoxysilane (APS; Sigma Diagnostics, St. Louis, MO, USA)
and dried overnight at 37 C.
The serial sections of ovaries were incubated with 10% normal
goat serum to reduce background staining caused by the second
antibody. The sections were then incubated with primary antibod-
ies raised against chicken inhibin α subunit (1-25)-NIe-Tyr, cyclic
inhibin βA (81-113)-NH2 (#305-24-D) and cyclic inhibin βB (80-
112)-NH2 (#305-25-D) for 12 h at room temperature. The antibody
against chicken inhibin α subunit was kindly provided by Dr. Pat
Johnson (Animal Physiology, Cornell University, NY, USA). The
antibodies for inhibin/activin (βA and βB) were kindly provided by
Dr. W. Vale (Salk Institute for Biological Studies, La Jolla, CA,
USA). The sections were then incubated with a second antibody,
goat anti-rabbit IgG conjugated with biotin and peroxidase with
avidin, using a rabbit ExtrAvidin staining kit (Sigma). This was
followed by visualizing with 30 mg 3,3-diaminobenzidine (Wako,
Osaka, Japan) solution in 150 ml of 0.05mol Tris-HCL 1-1 buffer
(pH 7.6) and 30 µl H2O2. Finally, the reacted sections were coun-
terstained with hematoxylin solution (Merck, Tokyo, Japan). The
control sections were treated with normal rabbit serum (Sigma)
instead of the primary antisera. The specificity of antibodies
against the inhibin α, βA and βB subunits was not examined using
neutralized antibodies instead of primary antibodies in the present
Mean values (± SEM) were calculated and analyzed using one-
way ANOVA. Duncan’s multiple-range test was used for detection
of significant differences using the SAS computer software pack-
age. A value of P<0.05 was considered to be statistically
Body and ovarian weight
The body and ovarian weights of the female Japanese quails
from hatching to sexual maturity are shown in Fig. 1. Body weight
increased gradually and reached a maximum at 7 weeks of age (Fig.
1A). On the other hand, ovarian weight did not show any signifi-
cant changes by 6 weeks of age, but there was an abrupt increase at
7 weeks of age (Fig. 1B).
Plasma concentrations of LH, ir-inhibin and progesterone
The plasma concentrations of LH did not change until 5 weeks
of age. They then increased significantly at 6 weeks of age and
declined at 7 weeks of age (Fig. 2A).The plasma concentrations of
ir-inhibin remained at a low level until 6 weeks of age and then
abruptly increased at 7 weeks of age (Fig. 2B). There were non-
significant fluctuations in the plasma concentrations of progester-
one until 6 weeks of age. A significant increase in the plasma
concentration of progesterone was observed at 7 weeks of age (Fig.
2C). The pituitary contents of LH began to increase at 6 weeks of
age and showed a significant increase at 7 weeks of age (Fig. 3).
SEDQYAR et al.
Immunolocalization of inhibin/activin subunits
Immunolocalization of the inhibin α, βA and βB subunits in the
Japanese quail ovaries is shown in Fig. 4. Positive staining for
inhibin α-subunit was clearly seen in the granulosa cells of ovarian
follicles from 1–7 weeks post-hatching. However, immunostaining
of theca interna cells was weak or negative in all follicular catego-
ries. Immunostaining for inhibin βA and βB was found in the same
cells as inhibin α subunit. Although the theca cell layer showed
lower affinity to the inhibin α, βA and βB subunits, some cells
showed positive staining for the inhibin βA subunit. The intensity
of immunostaining for inhibin α and βB was usually stronger than
that for βA in all stages of follicles. Positive staining of the number
of interstitial cells for all the inhibin subunits increased and tended
to surround the health large follicle. No immunostaining was
detected in control sections in which normal rabbit serum was sub-
stituted in place of the primary antibody (data not shown).
The present study demonstrated that developmental changes in
plasma ir-inhibin, LH and progesterone occurred in the female
quails from hatching to sexual maturity and that the plasma concen-
trations of ir-inhibin and progesterone were significantly increased
at 7 weeks of age. These results showed that ovarian activity are
accompanied by developmental changes in circulating ir-inhibin
Fig. 1.Changes in the body (A) and ovarian (B) weights of the
female Japanese quails from 1 to 7 weeks after hatching.
Values are means ± SEM (n=5–12). Within the same panel,
values without common superscripts are significantly
Fig. 2.Changes in the concentrations of LH (A), immunoreactive
(ir) inhibin (B) and progesterone (C) in the female Japanese
quails from 1 to 7 weeks after hatching. Values are means ±
SEM (n=5–12). Within the same panel, values without
common superscripts are significantly different (P<0.05).
Fig. 3.Changes in the pituitary content of LH in the female Japanese
quails from 1 to 7 weeks after hatching. Values are means ±
SEM (n=5–12). Within the same panel, values without
common superscripts are significantly different (P<0.05).
55INHIBIN IN FEMALE JAPANESE QUAILS
and progesterone. In addition, our immunohistochemical results
demonstrated that positive staining for the inhibin α, βA and βB
subunits is observed in granulosa and interstitial cells, whereas it is
not clear in the theca cells of female quails from hatching to sexual
maturity. These results suggest that the granulosa and interstitial
cells of female quails may secrete bioactive inhibins from hatching
to sexual maturity and that inhibins may play an important role in
the follicular development of Japanese quails. The developmental
changes in inhibins in female Japanese quails were observed for the
first time in the present study, and these observations showed that
Fig. 4.Immunohistochemical localization of the inhibin/activin subunits in the Japanese quail ovary from 1 to 7 weeks after hatching.
Immunostaining of the inhibin α, βA and βB subunits was observed in the granulosa and interstitial cells of all follicles at all ages from 1 to 7
weeks of age. Results at 1 week (Ai-A iii), 2 weeks (Bi-B iii), 3 weeks (C i-C iii), 4 weeks (Di-D iii), 5 weeks (Ei-E iii), 6 weeks (F i-F iii) and
7 weeks (G i-G iii) of age are shown. F1 follicles are shown from 5 to 7 weeks of age. The scale bar represents 100 µm. G: granulosa cells.
I: interstitial cells.
SEDQYAR et al.
plasma ir-inhibins were significantly increased at 7 weeks of age.
In other avian species, such as chickens [18–21, 28, 29] and ducks
[30, 31], similar developmental changes in circulating ir-inhibin
concentrations have been observed in accordance with the ovarian
activity. Previous studies in female chickens have shown that
inhibins are involved in regulation of their reproductive function
[18, 19]. Ir-inhibins in plasma have been shown to increase during
sexual maturation , and it has been reported that the ir-inhibin
content of granulosa cells and secretion into culture medium is cor-
related with the plasma levels . Yang et al.  reported that
the rise in ir-inhibin is correlated with age at sexual maturity in
female ducks, while a progressive increase in steroid hormones
may be consistent with a progressive increase in the steroidogenic
processes of the ovary. In the present study, the developmental
changes in circulating ir-inhibins parallelled progesterone and
exhibited higher values at 7 weeks of age. Supporting the present
study, previous studies have shown similar developmental profiles
for ir-inhibin and progesterone that reportedly correspond to Japa-
nese quails. In hens, circulating progesterone concentrations are
positively correlated with those of plasma inhibin A and ir-inhibin,
and the concentrations of inhibin A, ir-inhibin and progesterone
increase progressively prior to the onset of laying . Therefore,
our results are in agreement with the proposals that the maturing
ovary and, in particular, the large preovulatory follicles that are
more steroidogenically active  are primary sources of ir-inhibin
and progesterone .
Gonadal inhibin has been implicated as a negative regulator of
FSH secretion in the anterior pituitary gland with little or no effect
on LH [35, 36]. In this study, the plasma LH levels significantly
increased at 6 weeks of age, but declined at 7 weeks age, indicating
that the increased levels of LH stimulate maturation of large folli-
cles. Similar observations have also been reported in the chicken;
the plasma LH levels start to rise steeply between 16 and 19 weeks
of age, and then decrease at sexual maturity . The present
results suggest that the suppression of plasma LH in female quails
at 7 weeks of age maybe due to a high level of plasma progesterone
through the negative feedback regulation of gonadotropin releasing
hormone (GnRH). These results also agree with opinions
expressed in a review concerning birds, that gonadal steroids have
the ability to feedback at the hypothalamus and pituitary level to
control chicken GnRH receptor gene expression . Identifica-
tion of the sites of expression and production of inhibin subunit
messenger RNA and protein is critical to understanding the biology
of inhibins. In the present study, the inhibin α and inhibin/activin
βA and βB subunits were expressed in the granulosa cells of all fol-
licles during different stages of development from 1 to 7 weeks
after hatching, indicating that granulosa cells may secrete dimeric
and bioactive inhibins during the follicular development of Japa-
nese quails. Immunolocalization of the inhibin/activin subunits in
the granulosa cells of the quail ovary is in agreement with previous
studies showing that the granulosa cells of chicken [18, 19] and
duck [30, 31] ovaries are the major source of inhibin. In hens, the
greatest amount of mRNA expression for the inhibin/activin βA
subunit is found in the F1 follicle, while the inhibin α subunit is
expressed more abundantly than the inhibin βA subunit in large pre-
ovulatory follicles. Furthermore, the greatest amount of mRNA
expression for the inhibin βB subunit is found in the small yellow
developmental follicles. This suggests that the βA subunit, either as
inhibin A or as activin A, may be critical for ovulatory events,
while the inhibin/activin βB subunit may play an important role
during early follicular development . In the present study, the
expressions of the inhibin α, βA and βB subunits were similar in the
quail ovarian follicles during the period from hatching to sexual
maturity. This may explain the species difference in ovarian secre-
tion of inhibin during the prepubertal period. These differences
may even be found between breeds, such as in some mammals.
The expression levels of inhibin α and βA mRNA have been shown
to be correlated with the status of follicular development, whereas
inhibin βA mRNA is predominanrly expressed in large antral folli-
cles [39–41]. The levels of inhibin α subunit mRNA have been
shown to further increase during developmental maturation of folli-
cles and to dramatically decrease in preovulatory follicles
following the proestrous gonadotropin surges . This type of
difference may extend to the period after maturity in hens ,
whereas there is a differential level of inhibin expression between
broiler breeders and layers. The present study demonstrated that
granulosa cells are primarily the source of secretion of inhibin in
the quail ovary. Moreover, interstitial cells are an additional source
of inhibin and may share in control of the final stages of follicular
development. These results provide new physiological evidence
concerning follicular development of Japanese quails, particulary
that ovarian interstitial cells have the ability to synthesize inhibins.
The presence of inhibin subunits in ovarian interstitial cells could
imply a potential function for inhibin or activin in the quail ovary.
Supporting the present study, previous studies in the golden ham-
ster clearly demonstrated that both the proteins and mRNA of the
inhibin α and inhibin/activin βA subunits were found in ovarian
interstitial cells and that its expression was induced by LH surge
[44–46]. In summary, the present results demonstrated that hor-
monal changes depend on age, the rise in ir-inhibin is correlated
with ovarian activity and age at sexual maturity and ovarian granu-
losa cells and interstitial cells are the main sources of inhibin
secretion in the female Japanese quail from hatching to sexual
We thank Dr. G D Niswender, Animal Reproduction and Bio-
technology Laboratory, Colorado State University (Fort Collins,
CO, USA), for providing antiserum to progesterone (GDN 337);
Dr. J A Proudman, USDA-ARS, Biotechnology and Germplasm
Laboratory (Beltsville, MD, USA), for LH radioimmunoassay
materials; Dr. P Johnson (Animal Physiology, Cornell University,
NY, USA) for providing antiserum to chicken inhibin α subunit;
Dr. W Vale (Salk Institute for Biological Studies, La Jolla, CA,
USA) for providing anti-cyclic inhibin βA (81-113)-NH2 (#305-24-
D) and cyclic inhibin βB (80-112)-NH2 (#305-25-D); and the Bio-
signal Research Center, Institute for Molecular and Cellular
Regulation, Gunma university, Gunma, Japan, for providing antise-
rum against chicken LH. This study was supported in part by
Grants-in-Aid for Scientific Research (Basic research B-18310044,
P06445) and Japan-Thailand joint research from the Japan Society
57INHIBIN IN FEMALE JAPANESE QUAILS
for the Promotion of Science and a Grant-in-Aid from the National
Natural Science Foundation of China (NSFC) (No. 30670261).
Vale W, Rivier J, Vaughan J, McClintock R, Corrigan A, Woo W, Karr D, Spiess J.
Purification and characterization of an FSH releasing protein from porcine ovarian fol-
licular fluid. Nature 1986; 321: 776–779.
Vale W, Rivier C, Hsueh A. Chemical and biological characterization of the inhibin
family of protein hormones. Recent Prog Horm Res 1988; 44: 1–34.
Tanaka Y, Taniyama H, Tsunoda N, Shinbo H, Nagamine N, Nambo Y, Nagata S,
Watanabe G, Herath CB, Groome NP, Taya K. The testis as a major source of circulat-
ing inhibins in the male equine fetus during the second half of gestation. J Androl 2002;
Lin T, Calkins H, Morris PL, Vale WW, Bardin CW. Regulation of Leydig cell func-
tion in primary culture by inhibin and activin. Endocrinology 1989; 125: 2134–2150.
Chen CC. Editorial: Inhibin and activin as paracrine/autocrine factors. Endocrinology
1993; 132: 4–5.
Spencer SJ, Rabinovici J, Jaffe B. Human recombinant activin-A inhibits proliferation
of human fetal adrenal cells in vitro. J Clin Endocrinol Metab 1990; 71: 1678–1680.
Findlay JK, Robertson DM, Clarke IJ. Influences of dose and route of administration
of bovine follicular fluid and the suppressive effect of purified bovine inhibin (Mr
31000) on plasma FSH concentration in ovariectomized ewes. J Reprod Fertil 1987; 80:
Mercer JE, Clements JA, Funder JW, Clarke IJ. Rapid and specific lowering of pitu-
itary FSHb mRNA levels by inhibin. Mol Cell Endocrinol 1987; 53: 251–254.
Ling N, Ueno N, Ying S-Y, Esch F, Shimasaki S, Hotta M, Cuevas M, Guillemin R.
Inhibins and activins. Vitam Horm 1988; 44: 1–46.
Rivier C, Schwall R, Maso A, Burton L, Vaughan J, Vale W. Effects of recombinant
inhibin on luteinizing hormone and follicle-stimulating hormone secretion in the rat.
Endocrinology 1991; 128: 1548–1554.
Carroll RS, Kowash PM, Lofgren JA, Schwall RH, Chin WW. In vivo regulation of
FSH synthesis by inhibin and activin. Endocrinology 1991; 129: 3299–3304.
Ying S-Y. Inhibins, activins, and follistatins: gonadal proteins modulating the secre-
tion of follicle stimulating hormone. Endocr Rev 1988; 9: 267–293.
Ling N, Ying S-Y, Ueno N, Shimasaki S, Esch F, Hotta M, Guillemin R. Pituitary
FSH is released by a heterodimer of the β-subunits from the two forms of inhibin.
Nature 1986; 321: 779–782.
Schwall RH, Nikolics K, Szonyi E, Gorman C, Mason AJ. Recombinant expression
and characterization of human recombinant activin-A. Mol Endocrinol 1988; 2: 1237–
Muttukrishna S, Knight PG. Inverse effects of activin and inhibin on the synthesis
and secretion of FSH and LH by ovine pituitary cells in vitro. J Mol Endocrinol 1991; 6:
Akashiba H, Taya K, Sasamoto S. Secretion of inhibin by chicken granulosa in vitro.
Poult Sci 1988; 67: 1625–1631.
Tsonis CG, Sharp PJ, McNeilly AS. Inhibin bioactivity and pituitary cell mitogenic
activity from cultured chicken ovarian granulosa and thecal/stromal cells. J Endocrinol
1988; 116: 293–299.
Vanmontfort D, Rombauts L, Decuypere E, Verhoeven G. Source of immunoreactive
inhibin in the chicken ovary. Biol Reprod 1992; 47: 977–983.
Vanmontfort D, Berghman LR, Rombauts L, Verhoeven G, Decuypere E. Develop-
mental changes in immunoreactive inhibin and FSH in plasma of chickens from hatch
to sexual maturity. Br Poult Sci 1995; 36: 779–790.
Johnson PA, Brooks C, Wang S-Y, Chen C-C. Plasma concentrations of immunoreac-
tive inhibin and gonadotrophins following removal of ovarian follicles in the domestic
hen. Biol Reprod 1993; 49: 1026–1031.
Johnson PA, Brooks C. Developmental profile of plasma inhibin and gonadotrophins
from hatch to sexual maturity in male and female chickens. Gen Comp Endocrinol 1996;
Inooka S, Takahashi S, Takahashi H, Mizuma Y. Immunological traits in generations
7 to 12 of two lines of Japanese quail selected for high or low antibody response to
Newcastle disease virus. Poult Sci 1984; 63: 1298–1302.
Takahashi S, Inooka S, Mizuma Y. Selective breeding for high and low antibody
responses to inactivated Newcastle disease virus in Japanese quails. Poult Sci 1984; 63:
Krishnan KA, Proudman JA, Bahr JM. Purification and partial characterization of iso-
forms of luteinizing hormone from the chicken pituitary gland. Comp Biochem Physiol
Biochem Mol Biol 1994; 108: 253–264.
Hattori M, Wakabayashi K. Isoelectric focusing and gel filtration studies on the heter-
ogeneity of avian pituitary luteinizing hormone. Gen Comp Endocrinol 1979; 39: 215–
Hamada T, Watanabe G, Kokuho T, Taya K, Sasamoto S, Hasegawa Y, Miyamoto K,
Igarashi M. Radioimmunoassay of inhibin in various mammals. J Endocrinol 1989; 122:
Taya K, Watanabe G, Sasamoto S. Radioimmunoassay for progesterone, testosterone
and estradiol-17 β 125I-iodohistamine radioligands. J Anim Reprod 1985; 31: 186–197.
Davis AJ, Johnson PA. Expression pattern of messenger ribonucleic acid for follistatin
and the inhibin/activin subunits during follicular and testicular development in gal-
lus domesticus. Biol Reprod 1998; 59: 271–277.
Chen CC, Johnson PA. Expression of inhibin alpha and inhibin/activin beta A sub-
units in the granulosa layer of the large preovulatory follicles of the hen. Biol Reprod
1996; 55: 450–454.
Yang PX, Arai KY, Jin WZ, Watanabe G, Groome NP, Taya K. Preovulatory follicles
in the ovary as the source of circulating inhibin in the duck. Gen Comp Endocrinol 2001;
Yang P, Medan MS, Watanabe G, Taya K. Developmental changes of plasma inhibin,
gonadotropins, steroid hormones, and thyroid hormones in male and female Shao
ducks. Gen Comp Endocrinol 2005; 143: 161–167.
Vanmontfort D, Room G, Bruggeman V, Rombauts L, Berghman LR, Verhoeven G,
Decuypere E. Ovarian and extragonadal sources of immunoreactive inhibin in the
chicken; effects of dexamethasone. Gen Comp Endocrinol 1997; 105: 333–343.
Lovell TM, Knight PG, Groome NP, Gladwell RT. Changes in plasma inhibin A lev-
els during sexual maturation in the female chicken and the effects of active immuniza-
tion against inhibin alpha-subunit on reproductive hormone profiles and ovarian
function. Biol Reprod 2001; 64: 188–196.
Furr BJ, Smith GK. Effects on antisera against gonadal steroids on ovulation in the
hen Gallus domesticus. J Endocrinol 1975; 66: 303–304.
Woodruff TK, Mather JP. Inhibin, activin and the female reproductive axis. Annu Rev
Physiol 1995; 57: 219–244.
Knight PG. Roles of inhibins, activins and follistatin in the female reproductive sys-
tem. Front Neuroendocrinol 1996; 17: 476–509.
Sharp PJ. A comparison of variations in plasma luteinizing hormone concentrations in
male and female domestic chickens (Gallus domesticus) from hatch to sexual maturity.
J Endocrinol 1975; 67: 211–223.
Gregoy Y, Bedecarrats MS, Daniel G. Gonadotropin releasing hormones and their
receptors in avian species. J Poultry Sci 2006; 43: 199–214.
Torney AH, Hodgson YM, Forage R, de Kretser DM. Cellular localization of inhibin
mRNA in the bovine ovary by in-situ hybridization. J Reprod Fertil 1989; 86: 391–399.
Schwall RH, Mason AJ, Wilcox JN, Bassett SG, Zeleznik AJ. Localization of inhibin/
activin subunit mRNAs within the primate ovary. Mol Endocrinol 1990; 4: 75–79.
Fraser HM, Lunn SF, Cowen GM, Saunders PT. Localization of inhibin/activin sub-
unit mRNAs during the luteal phase in the primate ovary. J Mol Endocrinol 1993; 10:
Woodruff TK, D’Agostino J, Schwartz NB, Mayo KE. Dynamic changes in inhibin
messenger RNAs in rat ovarian follicles during the reproductive cycle. Science 1988;
Safi M, Buys N, Onagbesan OM, Vleugels B, Decuypere E. Quantification of
inhibin/activin alpha and betaA subunit messenger ribonucleic acid by competitive
reverse transcription-polymerase chain reaction in chicken granulosa cells during fol-
licular development. Biol Reprod 1998; 59: 1047–1054.
Kishi H, Ohshima K, Itoh M, Tsukada J, Arai KY, Nakano S, Watanabe G, Taya K.
Changes in expression of inhibin subunits in the cyclic golden hamster (Mesocricetus
auratus) and the regulation of inhibin alpha subunit production by luteinizing hor-
mone. Zoolog Sci 2002; 19: 225–232.
Shi Z, Jin W, Watanabe G, Suzuki AK, Takahashi S, Taya K. Expression of nerve
growth factor (NGF), and its receptors trkA and p75 in ovaries of the cyclic golden
hamster (Mesocricetus auratus) and the regulation of their production by luteinizing
hormone. J Reprod Dev 2004; 50: 605–611.
Arai KY, Kishi H, Onodera S, Jin W, Watanabe G, Suzuki AK, Takahashi S,
Kamada T, Nishiyama T, Taya K. Cyclic changes in messenger RNAs encoding
inhibin/activin subunits in the ovary of the golden hamster (Mesocricetus auratus). J
Endocrinol 2005; 185: 561–575.