AN ABSTRACT OF THE THESIS OF
Kelly Lynn Pinckard for the degree of Master of Science in Animal Science
presented on May 8. 1998. Title: Influence of Castration and Estrogen Replacement on
Sexual Behavior in Heterosexual. Male-Oriented and Asexual Rams.
Abstract Approved:
Redacted for Privacy
Fredrick Stormshak
An experiment was conducted to determine whether exogenous estradio1-1713
(E2) could restore sexual behavior in castrated rams. The experimental protocol
consisted of three sequential 6 wk periods during which rams were studied while: 1)
intact, 2) bilaterally castrated, and 3) implanted s.c. with E2 (two 7.6 cm silastic implants
[3.12 mm ID, 4.65 mm OD] packed with 309±16 mg E2). On the basis of previously
observed sexual behavior, rams were classified as heterosexual (n=7), male-oriented
(MORS; n=7), or asexual (n=7) and were subjected to 30 min sexual behavior tests
every 2 wk during the ensuing 18 wk investigation. Rams were observed for mounts and
ejaculations utilizing a test arena containing two ovariectomized progesterone-primed
E,-treated ewes and two intact males secured in stanchions. One week prior to each test
session, all subjects were isolated to prevent any sexual contact with pen mates.
Behavioral data were analyzed utilizing the signed lest. Asexual rams consistently
showed no sexual behavior and therefore were not evaluated statistically. Jugular blood
was collected at the beginning and the end of the 18 wk and testicular venous (n=21) and
arterial (n=8) bloods were collected during surgery before removal of the testes.
Systemic sera were quantified for estrone (E1) and E, using radioimmunoassay (RIA)
and the resulting data were analyzed statistically using ANOVA. Testicular sera were
quantified for oxytocin (OT) concentration using RIA and the resulting data were
analyzed by t-test and ANOVA. Mounting behavior declined after castration for MORS
(p<0.05) and heterosexual rams (p>0.10). Castration reduced the number of
ejaculations/intromissions by heterosexual rams (p<0.05), but not by MORS (p>0.10).
Treatment of castrated rams with exogenous E2 failed to restore sexual behavior of
MORS or heterosexual rams (p>0.10) and did not stimulate sexual behavior in asexual
rams. There were no marked differences (p>0.10) among ram groups with regard to
serum concentrations of E1 or E2 prior to castration (overall mean ±SE, 12.8±0.7 and
7.6±0.5 pgm1-1, respectively), nor any difference (p>0.10) in systemic concentration of
E1 or E2 among ram groups after rams were implanted with E2 (overall mean±SE,
9.7±0.7 and 9.0±0.7 pgm1-1, respectively). Systemic concentrations of E2 after
implantation of the steroid did not differ from those present while rams were intact
(p>0.10); however, mean E1 concentrations were reduced after castration with estrogen
replacement compared to levels present in the intact animal (p<0.02). Serum testicular
venous and arterial concentrations of OT were low and did not differ within or between
rams. These results suggest that restoration of E2 concentrations to physiological levels
in castrated adult rams (regardless of sexual orientation) cannot maintain or reestablish
sexual behaviors to levels observed prior to castration.
K,
Copyright by Kelly Lynn Pinckard
May 8, 1998
All Rights Reserved
Influence of Castration and Estrogen Replacement on Sexual Behavior in Heterosexual,
Male-Oriented and Asexual Rams
by
Kelly Lynn Pinckard
A THESIS
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Master of Science
Completed May 8, 1998
Commencement June 1998
Master of Science thesis of Kelly Lynn Pinckard presented on May 8, 1998
APPROVED:
Redacted for Privacy
Major Professor, representing Animal Science
Redacted for Privacy
Head of Department o Animal Sciences
Redacted for Privacy
Dea
I understand that my thesis will become part of the permanent collection of Oregon State
University libraries. My signature below authorizes release of my thesis to any reader
upon request.
Redacted for Privacy
Kelly Lynn Pinckard, Author
ACKNOWLEDGMENTS
When looking back at things you go through during the educational process,
sometimes it's hard to see the silver lining of the clouds that seem to follow you closely
as things come to a finish. Sacrifices can be great, but make you a better person through
the things you absorb along the way. I think it only appropriate to acknowledge the
many who have guided me through this process, helped me in a multitude of countless
ways and to mention loved ones that have sometimes been neglected, but unselfishly
stood by me when times got tough---all f o r the pursuit of m y goal .
. .
First and foremost, I wish to thank God for keeping me sane and listening to me
when I'm positive nobody else would want to. Thank you to my parents, for gently
urging me to keep going and supporting my every decision (even when they thought I
was out of my mind), and for raising my daughter, providing her with family stability,
loving her as if she was their own and instilling in her the intellect, integrity and unique
qualities that make her the gem that she truly is. Thanks to my sister Stacy and brother­
in-law Dave, who have always been straight up with me telling me exactly what was on
their minds, giving me their sometimes brutally honest opinion in all matters, and simply
for making the time to listen to me rant and rave when I needed someone to take the
time to care. Thanks to my new husband Timothy "Future Boy" Hazzard, who had no
idea from the moment he asked me "
.
.
. so are you married?" to the time he said "I do"
what he was getting into. He has displayed an unbelievable amount of patience, stood by
me when I didn't think I could make it and had the courage to keep supporting me by
kind words of "you can do this" no matter how somber my mood. And thanks to my
first baby, Alexa, she is truly a gift from God and manages to remain cheery, intelligent
and steadfast, despite the fact that she has lived the majority of her young life with an
absentee mom. Although I can never replace the time lost, I hope to build a wonderful
new relationship with her and be the kind of mother she has only dreamed about---one
that is always there for her. To all of my family, I could never express in words the
extent of my thanks and the depth of my gratitude.
I am also greatly thankful to those in Idaho without whom, this project could not
have been completed: Dr. John Stellflug, lead technician Mark Williams, supporting
technicians Verne LaVoie and Bill Gardner. Other collaborators I wish to express my
thanks to include Dr. Anne Perkins, Dr. Charles Roselli, Dr. James Fitzgerald, Dr. John
Resko, Henry Stadelman for his time and assistance with assay techniques and Dr. Dave
Thomas for advice on statistical analyses. I would also like to acknowledge Dr. Gordon
Niswender for the generous provision of P4 antiserum and Dr. Dieter Schams for the
donation of the OT antiserum, important little necessities required for successful
laboratory work. I would sincerely like to thank my committee members, Dr. Fredrick
Stormshak, Dr. Victor L. Hsu, Dr. Alfred Menino, Dr. Jerry Heidel and Dr. Lloyd
Swanson-all for taking the time out of their busy schedules to challenge me and test my
knowledge while watching me sweat it out!
In closing, I wish to thank Dr. Fredrick "Stormy" Stormshak, for his guidance
and implanting in me a "graphic understanding" of a higher work ethic .
.
.
and a special
note of thanks to his trusty dog Mike, who gave many seasons of service bringing the
animals in from the pastures (come ran or shine) so that I might not have to fall on my
face in the mud, waving a stick as the heifers or sheep stared on in blank delight!
TABLE OF CONTENTS
Page
REVIEW OF THE LITERATURE
MALE REPRODUCTIVE PHYSIOLOGY: AN OVERVIEW
ARCHITECTURE OF THE TESTES
Seminiferous Tubules and the Seminiferous Epithelium
The Sertoli Cell
The Leydig Cell
Steroidogenesis
SPERMATOGENESIS
Spermatocytogenesis
Spermiation
Cycle of the Seminiferous Epithelium and the Spermatogenic
Wave
Seminal Transport and Storage
HYPOTHALMO-HYPOPHYSEAL AXIS REGULATION OF THE
TESTES
Hypothalamic/Pituitary Hormones
Gonadotropin-Releasing. Hormone
Follicle-Stimulating Hormone and Luteinizing Hormone
Neuropeptides: Oxytocin and Vasopressin
Oxytocin
Vasopressin
Hormonal Control Through Feedback Mechanisms
RAM SEXUAL BEHAVIOR
Seasonal Influences
Genetic Basis for Homosexual Behavior
Aromatase: Relationship to Differentiation and Behavior
Sexual Differentiation
The Aromatase Enzyme Complex
Aromatase, Sexual Behavior and Sex Steroids
"Critical Periods" For Sexual Differentiation and
Relationship to Aromatase
STATEMENT OF THE PROBLEM
THE PRODUCER'S STANDPOINT
THE SCIENTIFIC STANDPOINT
1
5
5
6
6
8
8
10
10
11
12
12
13
14
15
16
18
20
20
22
23
24
24
28
29
32
36
36
37
TABLE OF CONTENTS (Continued)
Page
INFLUENCE OF CASTRATION AND ESTROGEN
REPLACEMENT ON SEXUAL BEHAVIOR IN
HETEROSEXUAL, MALE-ORIENTEDAND ASEXUAL
RAMS
38
INTRODUCTION
38
MATERIALS AND METHODS
Rams
Teaser Animals
Experimental Treatments
Preparation of Estradiol -17(3 Implants
Hormone Analyses
Serum Oxytocin Extraction and Radioimmunoassay
Steroid Extraction
Estradiol Radioimmunoassay
Estrone Radioimmunoassay
Statistical Analysis
39
39
40
40
42
46
47
48
RESULTS
48
DISCUSSION AND CONCLUSION
51
43
43
45
BIBLIOGRAPHY
59
APPENDIX
70
EFFECTS OF IN VIVO ESTRADIOL AND IN VITRO LH
TREATMENT ON PROGESTERONE SYNTHESIS BY OVINE
CORPORA LUTEA
71
LIST OF FIGURES
Figure
Page
1.
Biochemical steroidogenic pathway in testicular tissue.
9
2.
Mean (±SE) mounting behavior observed in heterosexual, MORS and
asexual rams while intact, after castration and after castration with E2
replacement.
49
3.
Mean (±SE) testicular venous OT concentrations in heterosexual,
MORS and asexual rams.
52
4.
Mean (±SE) testicular venous and arterial OT concentrations in rams.
53
LIST OF TABLES
Table
Page
1.
Mean (±SE) ejaculations of heterosexual, MORS and asexual rams
while intact, after castration and while castrated with E2 therapy.
50
2.
Mean (±SE) serum concentrations of estradiol and estrone (pgm11) in
heterosexual, MORS and asexual rams while intact and after castration
with E2 replacement.
54
INFLUENCE OF CASTRATION AND ESTROGEN
REPLACEMENT ON SEXUAL BEHAVIOR IN HETEROSEXUAL,
MALE-ORIENTED AND ASEXUAL RAMS
REVIEW OF THE LITERATURE
MALE REPRODUCTIVE PHYSIOLOGY: AN OVERVIEW
ARCHITECTURE OF THE TESTES
Testes are located exterior to the abdominal cavity in most mammalian species.
Extraordinary exceptions are species such as the elephant, which possesses testes inside
the abdominal cavity positioned caudoventrally to the kidneys, and sea mammals that
also have internally located testes. But in some mammalian species such as the bull, ram
and boar, the testes descend into the scrotum prenatally and are enveloped by an
extension of the peritoneum as they pass through the inguinal canal. The bull, ram and
billy goat have testes that are pendulous and vertically oriented, while stallion testes are
more horizontally oriented and only slightly pendulous. In contrast, the boar possesses
testes that are somewhat vertical in orientation but that protrude from the animal in the
perinea! region.
Testes of the ram are cradled within multiple muscle and tissue layers that are
covered by the protective outer scrotal skin. The scrotum is a layered structure that
serves to protect the testes from physical injury and aids in the maintenance of a
temperature 4-6° lower than the core of the body, which is requisite for viable sperm
production. The outermost layer, the skin, is in itself composed of three layers: the
epidermis, the dermis, and the hypodermis. A notable aspect of the epidermal layer is
2
that it possesses hair or wool and sebaceous glands that assist in thermoregulation. The
tunica dartos is the next scrotal layer primarily comprised of smooth muscle which gives
rise to the scrotal septum or raphe that lies between the two testes, confining each in its
respective compartment. In the case of a "fight or flight" response, the tunica dartos
retains the capacity for sustained contractions in order to bring the testes closer to the
body cavity for protection (Guyton and Hall, 1996). During non-stress responses, in
conjunction with contraction and wrinkling of the scrotal skin, the tunica dartos aids in
homeostatic thermoregulation when exposed to cold environments; while in response to
hot weather, the scrotal skin has the capacity to stretch, distancing the testes from the
body to maintain a lower temperature and thereby maintaining the integrity of testicular
sperm production (Hafez, 1993). Deeper layers surrounding the testes, the tunica
vaginalis parietal and tunica vaginalis viscera, are derivatives from a double layer of
peritoneum arising during early prenatal testicular descent. The coating in closest
contact, protecting the testes, is the whitish connective tissue capsule called the tunica
albuginea.
Testicular tissues are supplied by an intricate network of arteries and veins that
converge at the dorsal apex of the testes forming a complex structure known as the
pampiniform plexus. The function of this structure lends beautifully to its design in that
the intertwined vessels serve as a countercurrent thermoregulating mechanism meant to
contribute to the maintenance of an optimum temperature in the testes of 4-6° lower than
core body temperature. The veins leaving the testes contained in the complex serve to
cool arterial blood flow entering the testes while the arteries warm venous blood entering
3
the body. The spermatic cord which contains the nerves that innervate the testes, the
pampiniform plexus, lymphatic vessels, the testicular artery and veins, and the vas
deferens basically serves as a conduit between the testes and the abdominal cavity.
Interior of the testes can be broadly separated into two types of tissue: 1) the
parenchymal tissue composed primarily of seminiferous tubules, and 2) the stroma or
connective interstitial tissue support layer. The parenchyma is intersected at the central
core of the testes by a dense fibrous mass of connective tissue called the mediastinum.
Thinner layers of this connective tissue called septa radiate out from the mediastinum
creating compartmentalized lobular regions of concentrated, highly convoluted
seminiferous tubules. Both ends of each seminiferous tubule open into a duct called the
rete testes that converge within the mediastinum and then ascend upward to the dorsal
apical region of the testes to form the efferent ducts. Efferent ducts ultimately merge
and form one duct that ends and empties into the beginning or caput epididymis.
The caput (head), corpus (body), and cauda (tail) are all portions of the
epididymis that collectively have five main functions. The epididymis transports sperm
and is thought to concentrate it to a certain extent, and it secretes proteins that are
beneficial to sperm survival and contribute to seminal volume, and perhaps most
importantly, the cauda is a site for maturation and storage of sperm. When sperm are
voided into seminiferous tubule lumen they travel through the rete testes to the efferent
ducts. From this point, sperm enter the epididymis, traverse its length reaching the
storage site in the cauda in approximately 7 days in the bull (Amann and Schanbacher,
1983), 12 days in the boar (Swierstra, 1968), and 16 days in the ram (Amann, 1981).
4
Maturation as well as storage actually take place exclusively in this last section of the
epididymis.
Located at the terminal end of the epididymis, the vas deferens is a duct that
primarily serves the purpose of moving sperm from the site of storage in the cauda to the
pelvic urethra. The vas deferens extends from the ventral or distal aspect of the testes
and follows a path back up along the epididymis to the proximal pole where it joins the
spermatic cord complex. This structure then continues upward into the abdominal cavity
until it enlarges into the ampulla of the vas deferens at the junction with the prostate
gland and empties into the pelvic urethra. The ampulla serves as a transient, but minor
storage site for spermatozoa in some species, while the prostate gland contributes a
milky alkaline conglomeration of citrate ion, Ca', phosphate ion, clotting enzymes and
profibrinolysin to the seminal plasma. On either side of the prostate gland are bilateral
paired seminal vesicles that secrete a fructose, citric acid, prostaglandins, fibrinogen and
mucus laden milieu, which is the greatest contributor to the volume of the seminal
plasma and empties into the pelvic urethra. Near the dorsal portion of the pelvic urethra
are bilateral bulbourethral glands or Cowper's glands that are typically embedded deep
within local muscles that surround the urethra. Along the length of the urethra,
multitudes of minute urethal glands secrete mucus into the lumen of the urethra. The
accessory glands mentioned, the seminal vesicular glands, prostate gland, Cowper's
gland and urethal glands are not present in all mammalian males. Some species have
developed evolutionarily with a lack of one or more of these glands without impediment
to their overall reproductive function.
5
Seminiferous Tubules and the Seminiferous Epithelium
Seminiferous tubules possess a lining of epithelium consisting mainly of two
distinct cellular populations: the Sertoli cells and the germ cells. From the basement
membrane or basal lamina to the lumen, the seminiferous epithelium is unique in that it
consists of four to five germ cell layers that are the progenitors of sperm cells.
The Sertoli Cell
Within the seminiferous tubules, somatic Sertoli cells, acting as nurse or
sustentacular cells, extend from the basal lamina to the lumen and play an active role in
modification during the dynamic metamorphic evolution from germ cell to spermatid.
The term "germ cells" refer to spermatogonia closest to the basement membrane that are
of the least advanced stage in development and can be seen wedged between adjacent
Sertoli cells. Tight junction associations between neighboring Sertoli cells form the basis
for a division between diploid primordial germ cells along the basal lamina and haploid
spermatocytes toward the luminal side of the junctions. Sertoli cells provide the only
communication link traversing this compartmental division. Capillary walls of interstitial
vessels, the seminiferous tubule basal lamina and myoid layer along with Sertoli cell tight
junctions contribute to this exclusive compartmentalizing mechanism called the blood-
testis barrier, through which only small lipid compounds can pass (Setchell, 1980). The
blood-testis barrier was probably an evolutionary development established to protect
haploid spermatocytes within the seminiferous tubules from autoimmune destruction
because male systems tend to perceive haploid cells as foreign entities (Setchell, 1980).
If the blood-testis barrier was not intact, it would most certainly render the male infertile
6
and perhaps damage the seminiferous tubules themselves. Follicle-stimulating hormone
secreted from the adenohypophysis, in combination with testicular testosterone,
stimulates Serto li cells to produce critical secretory proteins such as androgen binding
protein (ABP) and inhibin (Amann and Schanbacher, 1983). Additionally, they
synthesize estrogen via aromatization of testosterone and play a role in facilitation of
spermatid cell transformation to spermatozoa, all of which will be addressed in more
detail later.
The Leydig Cell
Leydig cells are large polyhedral-shaped cells that possess multiple lipid-filled
vacuoles and vast networks of smooth endoplasmic reticulum (Hooker, 1944). In
response to adenohypophyseal pituitary secretion of luteinizing hormone, Leydig cells
synthesize great quantities of testosterone and small amounts of progesterone (Zirkin et
al., 1980). To facilitate the distribution of testosterone, Leydig cells are clustered around
blood and lymphatic vessels in the interstitial space of the testes, exterior to the
seminiferous tubules. Because spermatogenesis is androgen dependent, there exists an
intimate relationship between the required concentrations of testosterone and the
production and maturation of healthy viable sperm.
Steroidogenesis
Steroids constitute a class of lipids derived from perhydrocyclopentano­
phenanthrene, a cyclic four-ringed organic compound. Steroids are synthesized from
abundant stores of cholesterol sequestered within the endoplasmic reticulum of
7
steroidogenic cells, which are primarily located in the gonads and adrenal cortex as well
as the placenta in the pregnant female. In general, cholesterol is shuttled to the inner
mitochondrial membrane by a transport protein designated StAR, an acronym for
steroidogenic acute regulatory protein (Lin et al., 1995). There, cholesterol
instantaneously undergoes two hydroxylation reactions and is subjected to enzymatic
side chain cleavage by a cytochrome P450 side chain cleavage complex (Simpson et al.,
1993). At this initial stage, the molecule is no longer cholesterol, but has become
pregnenolone, an intermediate enroute from cholesterol to all other known steroid
compounds. Pregnenolone diffuses out of the mitochondria where it is acted upon by
3J3- hydroxysteroid dehydrogenase, converting it to progesterone. Within cells of the
zona glomerulosa of the adrenal cortex, cytosolic progesterone is subjected to a series of
three hydroxylation reactions, followed by enzymatic dehydrogenation producing the
mineralocorticoid aldosterone, which functions to regulate ion balance by promoting
reabsorption of Na4, Cl" and HCO3 at the level of the kidney. Progesterone undergoes a
series of different hydroxylation reactions in the zona fasciculata cells of the adrenal
cortex, which creates glucocorticoids, cortisol and corticosterone (Short, 1960), that act
to promote gluconeogenesis, and in pharmacological doses, suppress inflammation. In
male gonads, Leydig cells progressively convert progesterone to androstenedione via
hydroxylation that precedes a lyase reaction that splits a carbon-carbon double bond.
This androgen is further processed by a hydroxylase to produce testosterone. In cells
possessing the enzyme aromatase, androstenedione can be converted to the estrogenic
hormone estrone, and testosterone can be aromatized to estradio1-1713 (Ryan et al.,
8
1972; Figure 1). It is important to point out that steroids cannot be stored in the body;
therefore, they are continually synthesized as needed.
SPERMATOGENESIS
Spermatocytogenesis
By definition, spermatogenesis is simply the processes involved in producing
sperm cells or spermatozoa. Spermatogenesis is composed of a 1) spermatocytogenic
phase, which is the first phase of development of primitive sperm cells through a series of
mitotic and meiotic divisions to produce a haploid cell and 2) a spermiogenic phase, that
encompasses the differentiation of the nucleus from the cytoplasm in the haploid cell to
form a spermatozoan.
Diploid undifferentiated spermatogonia (AO adjacent to the basal lamina can
either divide to form additional A, cells through mitotic division or progress and divide
to form Al spermatogonia. From the AI stage, the cell becomes committed to a series of
divisions and transitions from an Al through A2, A3 and A4 cell stage to an intermediate
(In) spermatogonia. Progression continues to a B1 and B, stage then to the primary
spermatocyte (1°). From this point, the first meiotic division creates a secondary haploid
spermatocyte (2°).
Subsequently, a second round of meiosis forms the spermatid. In
the bull, this process takes approximately 13.5 days from the initial Ao cell stage to the
formation of an AI cell. Another 13.5 days must pass for the AI cell to progress through
multiple divisions creating the 1° cell. Finally, the transition from the 1° cell to the 2°
cell stage requires an additional 13.5 days (Amann and Schanbacher, 1983). Notably,
9
CHOLES TEROL
cytochronv P450scc
PRE GNE NOL ONE
3,Q hydrcaysteroidl
dehydrogenase
PROGES TERONE
17a -hydroxyl asel
17 a - HYDROXYPROGES TERONE
17, 20-lyase
ANDROS TENEDI ONE..
Ns. TESTOSTERONE
17/3-hydroxysteroid dohydrogenase
aronutasel
aronutase
I 33-hydroxysteroid dehydrogenase
ES TRONE
ES TRADI OL- 1 7 13
Figure 1. Biochemical steroidogenic pathway in testicular tissue.
10
the 2° stage is extremely brief in that it lasts only 2 to 3 hours. During this gradual
progression, Sertoli cells slowly phagocytize residual cytoplasmic droplets on spermatids
so that only a small residual body remains until maturity is reached within the cauda
epididymis.
The next phase, spermiogenesis, consists of four sub-phases called the Golgi,
cap, acrosomal and maturation phase that morphologically transform the spermatid to a
spermatozoa without undergoing actual cell divisions. The head of the spermatozoan
possesses a condensed nucleus with a scant cytoplasm and thin surface cell membrane.
The anterior portion of the spermatozoan head is known as the acrosome region
primarily derived from the Golgi apparatus and saturated with proteolytic enzymes (i.e.
hyaluronidase). The flagellum or tail of the cell is composed of microtubules and the
proximal region is encompassed by a mitochondria! sheath that supplies the ATP energy
substratum for motility.
Spermiation
Spermiation is simply the process that occurs when the spermatozoa are voided
into the lumen of the seminiferous tubules, which are then free for transport to the
epididymis for storage.
Cycle of the Seminiferous Epithelium and the Spermatogenic Wave
Upon close examination of cross sections of the seminiferous tubules, different
organizations of sperm cells, at various stages (of which there are 8 to 12) of
spermatogenesis, are arranged in a fixed hierarchy from the basement membrane to the
11
lumen. This organized structure is referred to as the cycle of the seminiferous epithelium
and in sum, it describes the arrangement of sperm cells as they divide, develop and
migrate to the lumen of the seminiferous tubules. This cycle, made up of different stages
of developmental cell layers, varies by individual animal and by species. Along the length
of the seminiferous tubules, longitudinally, there exists orientational changes in the stages
of the cycle of the seminiferous epithelium. For example, a given section of a tubule at
one stage in development, when examined in comparison to neighboring areas of tubule,
appears to have adjacent regions in either a stage of development preceding its own
present stage or just following it with regard to cellular development and cycle of the
seminiferous epithelium. These sequential stage changes along the length of the
seminiferous tubules are collectively called the sperrnatogenic wave.
Seminal Transport and Storage
Spermatozoa and seminal fluid released from the seminiferous tubules travel
through the rete testes up through the efferent ducts where they enter the epididymis.
While traversing the caput, the spermatozoa are immotile and do not possess any
fertilization capabilities. At this time, a proximal cytoplasmic droplet can be observed
near the head of spermatozoa indicative of the level of cellular immaturity. As the cells
enter the corpus, cytoplasmic droplets translocate down to the tail of the sperm, they
gain some motility and begin to possess a low level of fertility. Once in the cauda, the
cytoplasmic droplet is positioned on the distal portion of the tail or commonly falls off
Here the spermatozoa gain most of their motility, exhibit a high degree of fertility, and
remain in storage until mobilization during ejaculation.
12
HYPOTHALMO-HYPOPHYSEAL AXIS REGULATION OF THE TESTES
Hypothalamic/Pituitary Hormones
In concert with the nervous system, the endocrine system coordinates and
controls physiological functions including metabolism, growth and reproduction. Deep
at the root of this "harmonious interplay" lie two portions of the brain designated the
hypothalamus and the pituitary gland. The hypothalamus acts as an integration center of
the body as it receives sensory impulses and chemical messages and relays them in the
form of chemical responses at the level of the pituitary and bodily fluids. Hypothalamic
and pituitary chemical signals are then sent to peripherally located target tissues and
organs to elicit tissue-specific responses. Anatomically, the hypothalamus is an area of
the diencephalon that creates the floor of the third ventricle of the brain. This region
encompasses the optic chiasm, tuber cinereum, mammillary bodies, median eminence and
various nuclei sequestered within. The hypothalamus functions to regulate the
autonomic nervous system and influence hormonal secretion of the pituitary gland
through the synthesis and release of peptides and biogenic amines transported locally via
the hypothalmo-hypophyseal portal system that traverses the median eminence (Green
and Harris, 1946). Located ventrally with respect to the hypothalamus and protected
within a bony sheath known as the sella turcica, the pituitary gland is composed of the
adenohypophysis (pars distalis or anterior lobe), and the neurohypophysis (pars nervosa
or posterior lobe), the pars intermedia (the intermediate lobe), and the pars tuberalis.
During fetal development, the adenohypophysis arises from an area of the roof of the
embryonic oral ectoderm termed Rathke's Pouch that extends upward to meet the
13
neurohypophysis (Cunningham, 1992). In contrast, the neurohypophysis is derived from
a downward extension of an outpocketing of neural ectoderm from the floor of the third
ventricle (Cunningham, 1992).
Generally, hormone production in the hypothalamus includes a number of
releasing hormones that act at the level of the adenohypophysis (Green and Harris,
1946). For instance, thyrotropin-releasing hormone, corticotropin-releasing hormone
and gonadotropin-releasing hormone (GnRH) cause the release of thyroid stimulating
hormone, adrenocorticotropin-stimulating hormone and gonadotropins, respectively.
From a reproductive standpoint, the hormones of interest and critical importance are
GnRH and gonadotropins, which will be primarily emphasized in this discussion.
Gonadotropin-Releasing Hormone
A decapeptide consisting of 10 amino acids, GnRH or gonadotropin-releasing
hormone, as the name implies, acts to cause the release of gonadotropic hormones.
Multiple nuclei primarily located within the hypothalamus retain the capacity to
synthesize GnRH. The stimuli for synthesis and release of GnRH are generated in a
diffusely related network of neurons within the hypothalamus that sense hormonal
change and/or are affected by feedback mechanisms.
For instance, upon neuronal
stimulation by norepinephrine in conjunction with an intraneuronal Ca' influx, the
hormone is released in a pulsatile manner and moved from the site of origin to the
median eminence where it enters the hypothalmo-hypophyseal portal system through
which it reaches the adenohypophysis (Norris, 1997). Gonadotropin-releasing hormone
mediates its "releasing" effects when bound to a Gq-protein coupled GnRH receptor
14
present on the plasma membrane of gonadotropes in the adenohypophysis. Binding of
GnRH to its receptor results in activation of phospholipase Ca and initiation of the
phosphoinositide cascade during which there is an increase in intracellular Ca'
mobilization and protein kinase C activation (Norris, 1997). As a result, the
gonadotropins follicle-stimulating hormone (FSH) and luteinizing hormone (LH) are
synthesized and released by the adenohypophysis into the general systemic circulation to
act on the gonads.
Follicle-Stimulating Hormone and Luteinizing Hormone
Both FSH and LH are glycoprotein hormones, possessing an a and 13 subunit
structure linked together noncovalently, and with the a subunits of both being identical,
each hormone has a unique 3 subunit that confers specificity (Boothby et al.,
1981).
Follicle-stimulating hormone receptors have been associated with testicular Sertoli cells
which, when exposed to FSH, may have effects at the level of ongoing spermatogenic
processes in the testes (Schanbacher, 1979). Additionally, evidence exists to support the
supposition that Sertoli cells secrete small amounts of estrogens (Huggins and Moulder,
1945),
which may be attributed to aromatase activity in those cells. As previously
mentioned, FSH activates the Sertoli cells to synthesize and release inhibin (Ying,
1988),
which acts to inhibit further pituitary FSH secretion. Under the influence of FSH, Sertoli
cells are also stimulated to synthesize and secrete ABP, an essential systemic carrier
protein found in the blood and seminiferous fluids that binds the androgen testosterone
(Sanborn et al., 1975). At the same time, mediated through LH membrane receptors on
Leydig cells, LH stimulates the synthesis and secretion of testosterone, an androgen that
15
plays an essential role in that it is required for healthy sperm production (Norris, 1997).
When exposed to ewes in estrus, sexually active heterosexual rams typically experience
increased plasma LH concentrations (D'Occhio et al., 1983; Schanbacher et al., 1987;
Perkins et al., 1995), and subsequent increased testicular testosterone synthesis and
secretion (D'Occhio et al., 1983). However, studies by Perkins and Fitzgerald (1992),
as well as Perkins et al. (1995), have demonstrated that rams displaying homosexual
tendencies do not experience the endocrine LH surge in response to sexual activity
characteristic of heterosexual rams. The investigators hypothesized the lack of endocrine
response in rams during sexual stimulation, in conjunction with social or behavioral
effects learned during the rearing process and improper prenatal hypothalamic hormonal
priming of sexual centers within, might be responsible for male-oriented sexual behavior
in rams.
Neuropeptides: Oxytocin and Vasopressin
The neurohypophysis is also a vital player in the release of two peptide hormones
in the mammalian system, namely oxytocin (OT) and vasopressin (AVP). The actual
synthetic site of both OT and AVP is located in the magnocellular neurons in the
supraoptic nuclei (SON) and paraventricular nuclei (PVN) in the hypothalamus
(Cunningham, 1992). Axonal termini from these neurons extend down into the
neurohypophysis proper where OT and AVP are stored until nervous impulses or chemi­
osmotic changes stimulate cellular depolarization of the membrane to expedite an influx
of Ca`' and thus facilitate the release of OT and AVP.
16
acytocin
Commonly, when one thinks of the impact OT has on the animal body, one thinks
of the many roles the hormone plays in the female system with regard to smooth muscle
contractions in oviduct gamete transport, in the uterus facilitating parturition, the milk
ejection reflex, and uterine involution. Surprisingly, OT has been identified in males of
many species and may have implications revolving around penile erection (Melis et al.,
1986), smooth muscle contractility in the vas deferens during ejaculation (Fjellstrom et
al.,1968; Knight and Lindsay, 1970; Voglmayr, 1975; Carmichael et al., 1987; Murphy et
al., 1987), as well as having a role in steroidogenesis (Nicholson et al., 1991; Frayne and
Nicholson, 1995) and reproductive behaviors (Melin and Kihlstrom, 1963; Arletti et al.,
1985; Stoneham et al., 1985; Hughes et al., 1987; see review Carter, 1992). For
example, exogenous OT microinjected into the brain of the male rat at the level of the
PVN or the hippocampus induced multiple spontaneous penile erections and increased
the incidence of yawning, a characteristic masculine sexual behavior in rats, during a 1
hour observation period following treatment (Melis et al
1986). Hughes et al. (1987)
examined the concentration of OT in cerebral spinal fluid following sexual stimulation in
mature male rats. These researchers reported a significant increase in cerebral spinal
fluid OT concentrations double that of basal concentrations 5 minutes after ejaculation,
and three times above basal levels 20 minutes post-ejaculation (9 fmolm1-1 vs. 18
fmolm1-1 and 27 fmolm1-1, respectively). Further, Hillegaart and coworkers (1998)
investigated systemic OT concentrations of male rats in response to sexual stimulation.
Results of this study indicate that only sexually naive male rats have a significant increase
in serum OT concentrations immediately after interaction with receptive female rats
17
compared to controls (that received no sexual exposure) or sexually-stimulated
experienced male rats. Oxytocinergic effects and interaction with neurological brain
signals mediating sexual responses are indicative that this neurohormone plays a role in
the male with regard to erection and other aspects of sexual behavior. A later study,
utilizing chronic systemic administration of OT, appeared to reduce testicular and
circulating concentrations of testosterone, while 5a-dihydrotestosterone (DHT)
concentrations seemed to be higher compared with that of control male rats (Nicholson
et al., 1991). These same investigators also observed a reduction in FSH while LH was
not affected. Additionally, epididymal sperm numbers and the number of Leydig cells in
the interstitium, remained unchanged in response to chronic OT treatment. Thus, in
vivo, OT treatment appeared to have the capacity to modify the steroidogenic processes
in the testes. Fjellstrom et al. (1968) demonstrated that male rabbits, in response to
exogenous OT treatment, exhibited a significant increase in accessory sex gland fluid
production. Concomitantly, the researchers collected semen which showed an increased
concentration of spermatozoa in the ejaculate presumably due to enhanced smooth
muscle responsiveness and contractions of genital ducts facilitating spermatozoa release
from the seminiferous epithelium. And another investigation by Knight and Lindsey
(1970) identified an increase in seminal plasma volume and number of spermatozoa per
ejaculate in rams receiving short-term applications of exogenous OT. However, the
investigators also discovered that long-term treatment with OT had adverse effects on
spermatogenesis, which was evident by the dramatic increase in numbers of abnormal
sperm cells present in ram ejaculates. The possibility that substantial OT is synthesized
18
and secreted from the testes and the possibility that OT exerts paracrine or autocrine
effects in the ram are still under investigation.
Vasopressin
Another important neurohypophyseal hormone, AVP, functions in a regulatory
capacity in the homeostatic maintenance of many body systems. Vasopressin plays a role
in regulating blood pressure and glucose concentrations, and regulates homeostatic ion
and water balance within the body (acting as an antidiuretic agent). Further, AVP can
also act as a vasoconstrictive agent, and in times of bodily stress or trauma it acts at the
level of the adrenal cortex by stimulating the release of stress-related hormones. An
alternate role for AVP has been proposed in that it may modulate male sexual behavior
in the rat (Moltz, 1990). With this in mind, recent results of research conducted by
Swaab et al. (1995), suggest that the number of AVP neurons in the hypothalamic
suprachiasmatic nucleus (SCN) of the rat may be influenced by prenatal and perinatal
hormone exposure. In this study, male rats were treated with the aromatase inhibitor
1,4,6-androstatriene-3,17-dione (ATD), which blocks aromatization of testosterone to
estradiol, prenatally or during both the prenatal and perinatal period. In comparison to
control rats and rats that received only prenatal treatment (which preferred to mate with
the opposite sex), a 'bisexuality' of partner preference was observed in subjects that
were treated both pre- and perinatally. Investigators also observed a 65 % greater
concentration of AVP neurons in the SCN of 'bisexual' rats compared to controls and
prenatally-treated rats. These data may be a reflection of altered development during the
postnatal "critical period" of sexual differentiation of dimorphic centers in the rat brain.
19
Similarly, Bakker and colleagues (1993) observed that male rats neonatally treated with
ATD demonstrated a biphasic sexual preference as they preferred male rats when tested
in the dark phase and female rats when tested during the light phase of a 12:12 hour
light:dark cycle. This appeared to be indicative of an involvement of the "biological
clock" functions of the SCN in the regulation of sexual orientation and behavior in male
rats.
Further evidence for the role of the AVP containing neurons in the SCN and
association with sexual orientation has been documented from studies conducted with
the human brain following postmortem examination. Swaab and Hofman (1990)
reported that the SCN in homosexual men was approximately 1.7 times as large with 2.1
times as many cells as that of heterosexual men. It is hypothesized that observations
from studies on the AVP-containing regions such as the SCN and the significance with
regard to sexual orientation may be caused by the differential interaction of testosterone,
aromatase, estrogens and hormone receptors in the developing fetal or neonatal brain
(Swaab and Hofman, 1995). It is important to note that in the studies investigating
differences in the SCN of heterosexual men, women and homosexual men, that the
homosexual men died of complications from acquired immune deficiency syndrome,
otherwise known as AIDS. Therefore, it is uncertain whether the differences in the SCN
observed were an artifact that developed as a result of AIDS or due to sexual orientation
of the individual.
20
Hormonal Control through Feedback Systems
The main mechanism of communication between hormonally governed systems in
the body is through feedback regulation. Chemical substances or hormones secreted into
bodily fluids elicit a response from the target tissue or cell. This response is the basis for
feedback mechanisms. Such terms as ultra-short loop, short loop, long loop and ultralong loop, as well as positive and negative feedback refer to the distance in the body, the
sensitivity and response of the effector tissues. With regard to the hypothalamus and the
pituitary gland in the brain, the correspondence works through "downward" flow by way
of the hypothalamo-hypophyseal portal system (Wislocki and King, 1936) and/or by
retrograde flow back to the hypothalamus (Bergland and Page, 1978). This illustrates an
ultra-short loop at work in close proximity to the site of hormonal release with the site of
action. Hormones can also function through both positive and negative feedback.
Generally, stimulatory positive feedback occurs when a hormone acts on a site of release
and signals an increase in the synthesis of a hormone. Conversely, when concentrations
of a particular hormone are extremely high, the target tissue can respond by diminishing
the secretion of hormones, thus illustrating action through a negative feedback response.
RAM SEXUAL BEHAVIOR
Reproduction in the domestic ovine species is typically facilitated by the
evolution of sexual behavior in response to sexual stimuli in post-pubertal animals. In
particular, sexual behavior of the mature heterosexual ram entails a series of behavioral
patterns peremptory to and including copulation, directed toward the opposite sex.
21
Ultimately, the goal of expressed sexual behavior concludes with the union of the male
and female, which may result in the fusion of gametes, perpetuation of genes and
propagation of the species through offspring. Typical ram exhibitions of courtship and
copulatory behaviors include the sniffing and licking of the perineal area and urine of the
female, lending periodically to the Flehmen response, (a curling of the upper lip of the
male), as a mechanism of odor delivery to the chemosensory olfactory systems in the
vomeronasal organ and the brain (Lindsay, 1965; Ladewig and Hart, 1980). Upon
determination of reproductive readiness of the female to breed, the ram continues his
display with vocalizations, foreleg kicks, mounts and intromission, terminating after
ejaculation by a dismount and refractory period (Perkins and Fitzgerald, 1992).
Male-orientation or homosexual behavior in male animals is not uncommon.
Observations in many other species, both wild and domestic have described mounting
and copulatory behaviors between same sex individuals (see review Dagg, 1984). It is
thought that homosexuality may serve as a mechanism of developing a social hierarchical
structure and for the assertion of dominance within specie groups as has been observed
in wild flocks of Mountain sheep by Geist (1971). Alternatively, it has been postulated
that perhaps mounting behavior aids in the development of correct posterior orientation
for copulation (Silver and Price, 1986). However, observations made by Perkins and
Fitzgerald (1992), strayed from these patterns and hypotheses in that homosexual or
male-oriented rams (MORS) did not practice behaviors with the intent to assert
dominance, establish hierarchy or to discern proper rear orientation. Behavior of MORS
appears much the same as observed in heterosexual rams, with one major distinction:
MORS select and seek out other males specifically out of personal preference and desire
22
for a sexual partner. The documented discovery and research conducted on the
phenomenon of male-orientation in rams is in its infancy at best.
However, described
along with MORS in 1992, was the discovery and classification of asexual rams that
display no sexual behavior or desires whatsoever. To date, little is known, or has been
investigated with regard to these animals.
Seasonal Influences
Although sheep are considered "seasonal short-day" breeders, rams maintain the
capacity to breed year round (Karsch et al., 1984). The "season" of the ram could be
considered that time during which fertility and libido crescendo to a peak.
Conventionally, these periods of increased fertility and libido correspond to the late
summer through late winter reproductive cyclicity of the ewe (Schanbacher and Ford,
1979). A capitulation of physiological factors that correspond to a decrease in fertility,
and are inversely correlated to an increase in photoperiod, affect rams. These negative
effects comprise reduced scrotal circumference, decreased testicular weight and
seminiferous tubule diameters, and flaws or interruptions in the spermatogenic process
with a significant subsequent drop in sperm production (Dacheux et al., 1981). Further,
there is a concomitant diminished gonadotropic activity, which accounts for a wane in
libido (Borg et al., 1992) along with reduced testosterone secretion (Schanbacher and
Ford, 1979).
23
Genetic Basis for Homosexual Behavior
Many questions and much controversy has been raised over whether there is a
genetic link associated with homosexual behavior. With regard to preliminary studies on
fruit flies (Drosophila melanogaster), clear evidence has come to light supporting the
supposition of ties to homosexuality and the genome as has been revealed by Ryner and
colleagues (1996), as well as Hing and Carlson (1996). The, fruitless gene in Drosophila
has been identified and determined to be one that plays a role in the sexual preference of
male-oriented courtship behavior among male flies expressing the gene. Interestingly,
researchers hypothesize that the fruitless gene dictates neuronal functions within the
brain that assist in the coordination and execution of masculine courtship behavior;
however, these flies do not mate or court females but direct their affections exclusively
toward other males. Thus, males carrying the fruitless gene are "fruitless" or referenced
as sterile (Ryner et al., 1996). Male-male courtship behavior has also been observed in
Drosophila possessing the white gene. Investigations aimed at examining the significance
of this gene are similar to the previous discussed fruitless gene in that it appears that
expression of the white gene causes some sort of malfunction in sensory information
processing, which is represented by homosexual courting behavior in male flies. In
addition, study of the white gene also showed a correlation between homosexual
behavior due to the white gene and age. The incidence of male-male courtship occurred
at higher levels in flies between the ages of 1 and 4 weeks, compared to those less than a
week in age (Hing and Carlson, 1996).
Among humans, homosexuality is a very sensitive and controversial subject;
however, it has become apparent, according to research conducted by Hamer et al.
24
(1993), that there is evidence of a genetic linkage to male homosexuality. In a study
utilizing data from 114 families of homosexual men, it was determined that there was an
increased rate of same-sex orientation found in maternal uncles and male cousins of
homosexual men compared to their paternal relatives. This finding hinted at a
relationship to sex-linked transmission as a possibility in some cases of homosexuality.
The authors then focused on 40 of the families in the investigation that were selected on
the basis of the containment of two homosexual brothers and no paternal homosexual
relatives in the family history, to examine a genetic link. Upon DNA analysis, a high
correlation between a marker on the distal end of the long arm of the X chromosome
(designated Xq28), and male homosexual orientation was observed. The investigators
concluded that this correlation was indicative of a genetic influence on homosexuality in
some men.
The revelation of the genetic link to homosexuality in species as widely diverse as
those described in Drosophila to humans leaves the scientific door open to many
questions of applicability across other specie lines. Future continued research will need
to be conducted to confirm the validity of the genetic basis for homosexual behavior.
Aromatase: Relationship to Differentiation and Behavior
Sexual Differentiation
Chromosomal sex in mammalian species is determined by the presence or absence
of a "Y" chromosome in the animals genome. In the heterogametic genetic male the
presence of the Y chromosome results in the undifferentiated genital ridge tissue
25
developing into male gonads during embryonic development. Consequent production of
Miillerian Inhibiting Substance (MIS) from early Sefton cells in the evolving male gonad
inhibits intrinsic female reproductive tract tissues from emerging (Jost et al., 1973; Josso
et al., 1976; Josso et al., 1979; Donahoe et al., 1987). Somewhat like the reproductive
tract and gonads, the fetal brain also undergoes differentiation in response to gonadal
hormones (Wilson et al., 1981). Upon exposure of the fetal brain to androgens produced
by the early prenatal gonads, the male brain undergoes a transformation called
defeminization, which entails the loss of female behaviors such as lordosis and
proceptivity and likewise a loss of feminine neuroendocrine secretory patterns
(MacLusky and Naftolin, 1981). The first reported sexual dimorphism in the brain of a
vertebrate species, presumably due to the effects of steroidaily induced dimorphic
differentiation, was attributed to Nottebohm and Arnold (1976). These researchers
evaluated adult songbirds of finch and canary species for structural brain differences
between male and female conspecifics. Investigatory results revealed brain regions that
specifically controlled song patterns to be of greater volume in males compared with
females, evidence that sexual dimorphism does indeed exist in the avian brain. More
specifically, research has determined that the medial preoptic area (MPOA) within the
hypothalamus acts as the dimorphic sexual center, responsible in part for subsequent
reproductive masculine and feminine development and behavior. Assertion of masculine
"behavioral effects" are thought to be mediated by means of aromatization of androgens
to estrogens in some species (Beyer et al., 1976). Lesions created in the MPOA tend to
terminate male sexual behaviors in birds (Balthazart et al., 1992), rats (Giantonio et al.,
1970, Ginton and Merari, 1977), cats (Hart et al., 1973), and monkeys (Slimp et al.,
26
1978). As hypothalamic growth occurs, aromatase activity in the MPOA, stimulated by
the presence of gonadally derived androgens, progressively converts androgens to
estrogens further transforming the brain through masculinization or augmentation of
masculine patterns of gonadotropin secretion and the induction of male behavioral
characteristics such as courting and mounting behavior, which are expressed after
puberty. Sexual dimorphism in the MPOA has also been observed with regard to the
human brain and sexual orientation in men. LeVay (1991) described a region of the
human brain designated INAH 3 (the "interstitial nuclei of the anterior hypothalamus
region 3", analogous to the MPOA in other species), that was evaluated for volumetric
differences in heterosexual women, heterosexual and homosexual men during
postmortem examination. The investigation determined that this region was twice as
large in heterosexual men (0.12±0.01 mm3) than that of both homosexual men and
heterosexual women (0.05±0.01 mm3 and 0.06±0.02mm3, respectively). The researcher
concluded that INAH 3 exhibited dimorphism with regard to homosexual orientation in
men. However, the possibility of confounding results due to the fact that all of the
homosexual subjects in the study had died of AIDS could not be ruled out, regardless of
the fact that a comparison between the few heterosexual men and all homosexual men
that died of AIDS revealed the same result. In a different postmortem study conducted
by Allen and Gorski (1992), the size of the anterior commissure (a fiber tract in the
midsagittal area of the hypothalamus) of the human was evaluated in heterosexual
women, heterosexual and homosexual men. Results of this research showed the anterior
commissure in homosexual men to be 18% larger than heterosexual women and 34%
larger than heterosexual men. From these data, it was deduced that dimorphic structures
27
and function of the human brain must be affected by factors operating during prenatal
fetal development that differentiate anatomical features which have implications on
sexually orientated behavior. Yet, another question remains in light of the results of
these studies regarding the observed size differences of regions of the human brain
between heterosexual- and homosexually oriented men...are the differences the result of
sexual orientation or the cause of it?
Another key player from the standpoint of the female brain and sexual
differentiation is a liver protein which acts to protect the female fetus from androgenizing
effects of hormones. Genetic female fetuses are protected from their own gonadal
production of estradiol by an endogenous transient circulatory protein called a­
fetoprotein (AFP). Although present in both male and female fetuses in diminishing
concentrations until around the time of birth, or shortly thereafter, AFP binds systemic
estradiol with high affinity (Raynaud et al., 1971; Uriel et al., 1972). This serves to
provide a molecular barrier such that tissues sensitive to hormonal differentiating effects
of estradiol in the fetus (i.e., the hypothalamus) are unaffected because estradiol is unable
to bind its receptor or exert any effects while tightly bound to AFP. In the male fetal
brain, testosterone crosses the blood-brain barrier and at the level of hypothalamus and is
locally converted to estradiol via aromatase. The resulting free estradiol has
autocrine/paracrine differential action in the central nervous system (Plapinger and
McEwen, 1978). Thus, male genetics dictate that after puberty, an animal will behave
and perform like a male if proper hormonal defeminization and masculinization have
occurred in !tier° during "critical periods" of sexual differentiation. To reiterate this turn
of events, as was so eloquently supported by experimental evidence in the guinea pig and
28
stated by Young et al. (1964) "...during fetal morphogenesis, androgens exert a
fundamental influence on the organization of the soma, determining whether sexual
reactions brought to expression in the adult will be masculine or feminine in character."
The Aromatase Enzyme Complex
Aromatization occurs in placental tissue (Thompson and Siiteri, 1974; Kellis and
Vickery, 1987), fetal hypothalamic and limbic tissues (Ryan et al., 1972), fetal liver
(Slaunwhite et al., 1965), fetal lung, thymus, kidney and skin (Schindler et al., 1975),
fetal (Attal, 1969) and adult gonads (Payne et al., 1976), adult kidney, adrenal glands,
and to a very limited extent in adult liver (Friedan et al., 1968; Longcope et al., 1969),
adult adipose and muscle (Longcope et al., 1976, 1978; Gray et al., 1979) and has been
identified in the mandibular bone of the rat (Vittek et al., 1974), in addition to adult
hypothalamic and limbic tissues. Within the adult rat brain, aromatase has been identified
in localized discrete regions including the following: the hypothalamic medial and
periventricular preoptic area, medial and cortical amygdala, bed nucleus of the stria
terminalis, diencephalon, suprachiasmatic nucleus, anterior and periventricular anterior
hypothalamus, ventromedial nucleus, arcuate nucleus, median eminence, lateral preoptic
nucleus, supraoptic nucleus, dorsomedial nucleus and lateral hypothalamus. Notably, the
highest activity has been associated with the MPOA and amygdala (Roselli and Resko,
1987).
The enzyme complex (aromatase) responsible for progressive conversion of
androgens to estrogens is localized within the endoplasmic reticulum of tissues that are
responsive to this class of steroids. Two components comprise the complex, namely a 1)
29
cytochrome P450. component (Mendelson et al 1985; Kellis and Vickery, 1987),
which has been identified as a CYP19 gene product (Nebert et al., 1991) and 2) an
ubiquitous flavoprotein NADPH cytochrome P450 reductase which acts to transfer H+
reducing equivalents to any cytochrome P450 with which it comes in contact (Griffen
and Ojeda, 1996). Aromatase expression appears to be regulated by testosterone
exposure (Rose lli and Resko, 1987) and tissue-specific promoters at the transcriptional
level (Simpson et al., 1993). Aromatase, when up-regulated, evokes a series of complex
reactions that result in the binding of cytochrome P450., to an androgenic substrate.
The reaction series entails sequential hydroxylation, oxidation and removal of the C-19
carbon followed by aromatization of the A ring of the steroid. These reactions require 3
moles of NADPH and 3 moles of oxygen for the conversion of each mole of androgen
metabolized (Griffen and Ojeda, 1996).
Aromatase, Sexual Behavior and Sex Steroids
An example of the implications of aromatase, the interaction with hormones and
its effects on sexual behavior in avian species can be seen in extensive research
conducted on the Japanese quail. Aromatase-immunoreactive neurons localized in the
sexually dimorphic MPOA of the male bird encompass specific critical regions for
testosterone action presumed to be required for the activation of male sexual behavior
(Balthazart et al., 1992). Both intracerebral testosterone implants centralized in the
vicinity of the MPOA with adjunct subcutaneous systemic testosterone implants, not
only stimulated sexual behavior in castrated birds to levels demonstrated in pre-castrates,
but appeared to induce an up-regulation of neurons possessing aromatase activity rather
30
than the overall shrinkage of the region observed in control capons. Ablation of the
MPOA via electrolytic lesioning successfully quelled male sexual behavior; however, if
complete lesioning was not established, only partial suppression of behavior could be
achieved dependent on the degree of tissue destruction (Balthazart et al., 1992).
Another study investigated the action of testosterone treatment in conjunction with the
aromatase inhibitor ATD. Results demonstrated that testosterone had the capacity to
restore male sexual behavior in castrated birds until concurrent administration of ATD,
afterwhich, sexual behavior completely ceased (Watson and Adkins-Regan, 1989).
Therefore, these observations documented for quail lend support to the hypothesis that
testosterone acts within the dimorphic nucleus to activate male copulatory behavior
through its aromatization to estrogen.
In contrast, Schumacher and Balthazart (1986) revealed that the female
hypothalamus in the Japanese quail does not retain the capacity to convert testosterone
into behaviorally active metabolites such as estradiol and the non-aromatizable androgen
DHT to the same extent as occurs in the male; thus, the female is rendered insensitive to
the sexual behavior effects of testosterone inherent in males. Additionally, researchers
discovered that aromatase is present in smaller quantities in the female brain in
comparison to that of males. From the evidence of their study, the authors postulated
that these facts are a reflection of a cascade effect, magnified by the presence of higher
systemic concentrations of testosterone in the male which, in turn, acts to induce
significant aromatase activity in the brain.
An induction of male sexual behavior was also achieved in castrated mature mice
when injected with either testosterone (Luttge and Hall, 1973), estradiol benzoate, or
31
DHT plus estradiol benzoate (Wallis and Luttge, 1975); however, DHT treatment alone
was ineffective (Luttge and Hall, 1973). Similar results were observed in the male
hamster because testosterone (Wood and Newman, 1995), and estradiol but not DHT
stimulated masculine sexual behavior (Wood, 1996). In the hamster, these steroid
treatments were applied as intracranial implants within either the amygdala or the
MPOA. Implants in either region were equally capable of eliciting the same behavioral
response; therefore, testosterone may facilitate behavior by means of aromatization to
estrogen through multiple brain regions in the male hamster. Unlike most other
mammalian species, when implanted intracranially and subdermally with DHT, sexual
responses are restored in castrated male guinea pigs (Butera and Czaja, 1989).
However, neither DHT nor estradiol stimulates male sexual behavior in amphibians
(Deviche and Moore, 1988) or pigs (Levis and Ford, 1989), suggesting that perhaps in
these species and others that react similarly to DHT or estradiol treatment, testosterone
is the hormone responsible for the promotion of some aspects of masculine behavior. In
light of this, it is important to note that both androgen and estrogen synergism function
to maintain the full complement of male sexual behavior in pigs (Levis and Ford, 1989),
but not in amphibians (Deviche and Moore, 1988).
In non-human primates, subcutaneous injections of testosterone propionate
maintained the full spectrum of male sexual behavior, but behavior declined after
cessation of testosterone propionate treatment and subsequent administration of estradiol
benzoate (Michael et al., 1990). Further, estrogen treatment alone failed to restore
sexual behavior in castrated Rhesus macaques (Michael et al., 1990). In spite of the
findings in the adult, estrogens are readily synthesized by the Rhesus monkey fetal brain
32
as was determined by Rose lli and Resko (1986). In concert with estrogen production,
the highest levels of aromatase activity were located in the MPOA of the fetal brain.
Results of this investigation demonstrated that aromatase activity in the fetal brain was as
much as ten times greater than levels measured in adult monkeys. These investigators
concluded that this activity was not regulated by the concentrations of circulating
androgens in the fetus (Roselli and Resko, 1986) as was subsequently found to be the
situation for the adult system (Roselli and Resko, 1989). Although aromatase appears to
be regulated by androgen concentrations in the adult monkey, brain concentrations of
5a-reductase, the enzyme that converts testosterone to DHT, remain static and are
thought to be independent of systemic androgen concentrations (Roselli and Resko,
1989). Hence, the presence of functional androgen receptors in the brain of the fetal
Rhesus monkey suggests that perhaps aromatization is not necessary for sexual
differentiation of reproductive behaviors in nonhuman primates and that androgens may
act directly by binding to their receptor (Connolly et al., 1994a).
"Critical Periods" For Sexual Differentiation and Relationship to Aromatase
It is hypothesized that "critical periods" exist during fetal or perinatal
development during which the brain must be exposed to androgens for an animal to
mature physiologically and behaviorally into a male at puberty. The designated "critical
periods" have been defined for few species and those that have been reported appear to
vary among species. For example, the "critical period" for the guinea pig appears to be
between 30 and 37 days of gestation (Connolly and Resko, 1994), during which time the
brain appears to possess high concentrations of aromatase (Connolly et al., 19946).
33
Hypothalamic MPOA differentiation may be complete before 70 days gestation in the
sheep (Attal, 1969), and interestingly, prior to masculine differentiation of the fetal brain,
the "critical period" for prenatal development of external genitalia of sheep has been
reported to be a confined window between 40 and 50 days gestation (Clarke et al.,
1976). Additionally, Wood et al. (1995) conducted an experiment to determine if the
centers that control tonic and surge LH release could be masculinized through the
androgenization of female lambs in utero. In the process of this study, results revealed
that female lambs born to ewes treated daily with testosterone cypionate from either 30
through 51 or 30 through 86 days gestation developed masculine external genitalia,
while those born to ewes treated 65 through 86 days gestation developed as normal
phenotypic females. However, the researchers were unable to unequivocally masculinize
the tonic and surge patterns of LH release, but they hypothesized that the "critical
periods" of androgen exposure necessary might be independent from those observed for
differentiation of genitalia and sexual centers within the brain. The "critical period" for
the Rhesus monkey has been narrowed down to 35-50 days gestation, a time of peak
fetal testicular testosterone production (Resko et al., 1980). Nevertheless, to test the
primate "critical period" hypothesis and the effects on male sexual behavior in another
primate species, intramuscular injections of testosterone propionate were administered
and appeared to restore sexual behavior in mature marmoset monkeys castrated
neonatally. However, these data may differ from observations for that of the Rhesus
monkey because a specific "critical period" has not been defined for the marmoset
species and it was not stated at what point during gestation these animals underwent
orchidectomy (Dixson, 1993). Perhaps the subjects under investigation were castrated
34
after the neonatal period of androgen sensitivity, and thus retained the capacity to
respond to androgen treatment after puberty. Alternatively, the "critical period" of
sexual differentiation may simply be unique for different primate species. In contrast, the
"critical period" for the rat appears to be primarily prior to a 5 day postnatal time frame
(Barraclough, 1961; Goy et al., 1962; Grady and Phoenix, 1963). Neonatal female rats
treated with a single androgen injection on or before 5 days of age were found to be
acyclic and sterile at maturity (Barraclough, 1961). Conversely, newborn male pups
castrated postnatally before the 5th day after birth and concomitantly receiving estrogen
plus progesterone treatment, tended to display feminine sexual behaviors after puberty
(Goy et al., 1962; Grady and Phoenix, 1963). Similarly, estrogen-treated mature male
pigs that were castrated within 48 hours after birth displayed characteristic female sexual
behavior and sought out adjacently penned mature intact male pigs rather than adjoining
pens containing female pigs (Ford, 1983). These data suggest that in the rat and pig,
steroidal androgens may play a role in the defeminizing process during a developmental
postnatal period rather than a "critical period" of prenatal origin.
Resko et al. (1996) reported that sexual orientation of the ram might also be
contingent upon exposure of the MPOA to aromatized androgens during the "critical
periods" of fetal development in sheep. As suggested from their study, the MPOAs in
male-oriented individuals were found to be low in aromatase activity when compared
with those from rams that exhibited more male-typic behavior directed toward females.
Reduced concentrations of aromatase, in conjunction with a decreased capacity for
testicular production of testosterone in rams displaying male-orientation, raises questions
as to whether the dimorphic centers of the hypothalamus were exposed to necessary
35
levels of androgens and estrogens required for defeminization and masculinization of the
brain during the fetal development of these subjects.
36
STATEMENT OF THE PROBLEM
THE PRODUCER'S STANDPOINT
Among domesticated livestock species, courtship usually results in the union of a
male and female to produce offspring and profit for those that own them. However,
within the ovine species exists a subclass of animals, male members that have been
observed in partaking in same-sex mating practices for reasons other that simple male-
male societal dominance related interactions (Adkins-Regan, 1988). Rams designated
male-oriented (MORS) have been classified by a method of sexual preference testing
whereby the ram is given a mating selection of stanchioned females in estrus or males
(Perkins et al, 1992b). When faced with partner choices, the MORS seek out males and
exclusively engage in courtship rituals followed by attempts at copulation with males.
Following introduction at the commercial level, MORS actively seek out other rams as
sexual partners not only neglecting their duties as male progenitors, but sometimes
causing disruption to heterosexual rams' completion of normal copulatory practices.
This has the potential for tremendous economic loss to producers with regard to open
ewes and reduced lamb crop if they possess even a single male-oriented individual
(Perkins et al., 1992a). Hence, infertility does not lie within the female, a problem that is
typically remedied by culling the few non-productive females in the flock; infertility
becomes seriously compounded, for a multiplicity of females, due to the unwillingness of
the male to mate with anything other than other rams within the flock.
37
THE SCIENTIFIC STANDPOINT
Investigation of the phenomenon of sexual orientation in sheep is in its infancy at
best. To date, it is hypothesized that MORS have a reduced capacity for testosterone
synthesis in the testes, lower aromatase activity in the brain (Resko et al., 1996), and
little or no secretory pattern of pulsatile LH release from the pituitary compared to
heterosexual rams (Fitzgerald and Perkins, 1994; Perkins et al., 1995), all of which are
thought to be influencial in the sexual orientation of these animals. In addition, sexual
behavior of homosexual rams is also thought to be associated with a decreased
concentration of estradiol-receptors, specifically in the amygdala of the hypothalamus
(Alexander et al., 1993; Perkins et al., 1995). Another aspect of ram fertility that needs
to be addressed, described along with MORS by Perkins and Fitzgerald in 1992, is the
little investigated asexual ram that never displays any sexual behaviors. Currently, no
research has included asexual rams as a model of study. Thus, the aims of this study are
multifaceted stemming from the necessity to expand the body of knowledge surrounding
male-oriented and asexual rams and to better physiologically and endocrinologically
characterize aspects encompassing sexuality and fertility in rams. And in light of this, it
is feasible to investigate the effects of physiological estradiol replacement to the
castrated ram as a means of altering sexual behavior, thus possibly re-orienting and/or
restoring breeding capacity specifically in MORS and asexual rams.
38
INFLUENCE OF CASTRATION AND ESTROGEN
REPLACEMENT ON SEXUAL BEHAVIOR IN HETEROSEXUAL,
MALE-ORIENTED AND ASEXUAL RAMS
INTRODUCTION
In mammalian species, reproductive success is dependent upon aspects that play
a role in sexual behavior including the endocrine and nervous systems working
synchonously. As far back as the origin of neonatal organization of reproductive and
neurological tissues, "critical periods" have been defined for many species during which
hormonal exposure determines masculine or feminine destiny of differentiating tissues of
the reproductive tract, external anatomy and fetal brain.
Male-oriented or homosexual behavior in male animals seems to be relatively
widespread among species and has been well documented (Dagg, 1984); however,
expression of male-oriented behavior in some sheep has been shown to be for reasons
other than establishment of hierarchical social structure or the development of proper
posterior orientation for copulation. Male-oriented rams (MORS) and asexual rams
were originally described by Perkins and Fitzgerald in 1992. Rams classified as "male­
oriented" exclusively sought out other males for sexual partners even in the presence of
females in estrus, while asexual rams did not display any sexual behavior at all. To date
little is known about asexual rams and a handful of studies have focused on MORS.
Researchers investigating MORS have characterized them thus far as: lacking the
endocrine LH surge in response to sexual activity characteristic of heterosexual rams
(Perkins and Fitzgerald, 1992; Perkins et al., 1995); possessing lower estradiol receptors
39
in the amygdala (Alexander et al., 1993; Perkins et al , 1995); lacking the capacity to
synthesize testicular testosterone to the same extent as heterosexual rams, and lower
aromatic activity in the medial preoptic area (MPOA) of the hypothalamus compared
with heterosexual rams (Resko et al., 1996). This ultimately results in lower
concentrations of estradiol and estrone in male-oriented individuals (Resko et al., 1996).
It has been established that oxytocin (OT) is present in the testes of many
mammalian species including the sheep (see review, Wathes, 1984) and may play a role
in reproductive behavior (see review, Carter, 1992). Oxytocin has also been identified in
the ovine epididymis and been implicated in facilitating spermatozoa development and
function (Knickerbocker et al., 1988; Nicholson et al., 1991). Yet the question remains
as to whether the testes of the ram are a synthetic site for OT.
Therefore, the present study was designed to determine if the testes of the ram
synthesized OT, and to determine whether estradiol replacement could restore, reorient
or stimulate sexual behavior in castrated heterosexual, MORS and asexual rams.
MATERIALS AND METHODS
Rams
Twenty-one mature rams previously tested for sexual behavior as described by
Perkins and Fitzgerald (1992) were classified as 1) heterosexual (n=7), 2) male-oriented
(n=7) or 3) asexual (n=7). All subjects were housed in outdoor pens in their respective
groups until 1 wk prior to behavioral testing At this time, animals were individually
isolated in indoor pens under simulated natural seasonal lighting conditions. Rams were
40
fed approximately 1.4-1.8 kg long stem harhead-Iday -1 and had ad libitum access to a
mineral salt block and water. All experimental procedures and surgeries were performed
in accordance with the Institutional Animal Care and Use Committee.
Teaser Animals
Four randomly selected mature ovariectomized (OVX) ewes were selected from
a pool of 15 OVX ewes, 14 days prior to each behavioral preference test. Upon
selection, each ewe received a vaginal progesterone pessary that remained in place for 13
days. Pessaries were removed on day 13 and each ewe received an injection of estradiol
to induce standing behavioral estrus for teasing purposes. Four mature intact rams were
also employed as stimulus teasers during behavioral preference tests. Hence, two OVX
progesterone-primed estrogen-treated ewes and two intact rams were secured in
stanchions within the test arena and utilized as stimulus animals during the behavioral
testing periods.
Experimental Treatments
The investigation period of this experiment was 18 wk in duration and consisted
of three sequential 6 wk periods when rams were: 1) intact, 2) bilaterally castrated and 3)
implanted s.c. with estradiol -17P (E). During the intact period, rams were tested for
behavior every 2 wk for a total of three observations per animal. One wk prior to sexual
preference testing, rams were isolated in individual pens to prevent any sexual contact
with pen mates. Each behavioral test session consisted of a 30 min observation period
during which the test subject entered the an arena with unfettered access to two OVX
41
progesterone-primed, estrogen-treated ewes and two intact rams secured in stanchions.
Observations with regard to sexual behavior (mounts and ejaculations/intromissions) of
the test subjects and the sex of the stimulus animal toward which the behavior was
directed were recorded.
The second phase of the experiment was conducted after all rams were bilaterally
castrated. Prior to castration, a jugular blood sample was drawn from each ram by
venipuncture, collected in a 5 ml vacutainer tube and allowed to clot for 2 h at room
temperature. All samples were then stored at 4° C overnight, centrifuged at 2000 x g the
following day and sera were harvested and stored at -20° C to be later extracted and
analyzed by radioimmunoassay for estrogen concentration. For the castration procedure,
rams were anesthetized with a jugular injection of 0.5 ml diazipam and 2.25 ml ketamine.
Testes were exposed prior to orchidectomy through bilateral proximal scrotal incisions at
which time testicular venous blood, proximal to the pampiniform plexus (n=21), as well
as arterial blood distal to the plexus (n=8) were collected via venipuncture into 5 ml
vacutainer collection tubes. Samples were allowed to clot for 2 h at room temperature,
then stored at 4° C overnight. The following day, samples were centrifuged as described
above, sera harvested and stored at -20° C until extracted and subjected to
radioimmunoassay for OT. During recovery from surgery, all rams were given 1 g oral
phenylbutazone, and an i.m. injection of both 6 ml LA-200 and 5 ml vitamin K. Rams
were then returned to their respective groups in outdoor pens. Sexual behavioral testing
of the rams commenced as previously described every 2 wk for a total of three
observations per animal at which time, all rams received s.c. E, implants. Rams were
implanted with E, in the axillary region of the foreleg following guidelines implemented
42
by Karsh et al. (1980). Following a general cleansing of the region, skin was scrubbed
with 70% ethanol, then spritzed with a betadine solution. The incision site was liberally
injected s.c. with 0.25 ml increments of lidocaine and an approximately 2-3 cm incision
was made with surgical scissors. Hemostats were inserted into the incision and opened
to create a subcutaneous pouch into which two 7.6-cm silastic implants packed with
crystalline E2, (presoaked for I h in 70% ethanol), were placed. Incisions were closed
with surgical wound clips and sprayed topically with Furasone. Rams were then
returned to their respective groups and tested for sexual behaviors as described above
every 2 wk for a total of three observations per animal. At the conclusion of this final 6
wk period, jugular blood samples were collected into 5 ml vacutainer collection tubes by
venipuncture from each ram. Blood samples were allowed to clot for 2 h at room
temperature, stored overnight at 4° C. The following day, samples were centrifuged as
described above, sera harvested and stored at -20° C until extracted and analyzed for
estrogen concentration.
Preparation of Estradio1-1713 Implants
Plasma estrogen concentrations based on those observed by Resko et al. (1996)
were achieved in castrated rams by the s.c. implantation of silastic capsules containing
crystalline E2. These capsules were prepared by a modification of procedures described
by Karsh et al. (1973). Briefly, one end of a 7.6-cm length of Silastic Medical-Grade
Tubing (3.12 mm ID, 4.65 mm OD; Dow Corning, Midland, MI) was sealed with a 3.0­
mm plug of Silastic Medical Adhesive (Silicone type A; Dow Corning). Approximately
309 ± I6 mg (7.0-cm length column of hormone) crystalline E, (Sigma Chemical Co., St.
43
Louis, MO) was packed into the tube, and the open end was sealed as described above.
The implants were soaked overnight in tap water to equalize the amount of hormone in
the walls of the capsule, then dried and stored in an air tight container at 4° C until
utilized for implantation.
Hormone Analyses
Serum Oxytocin Extraction and Radioimmunoassay
Oxytocin in the testicular serum samples was extracted and quantified as
described by Orwig et al. (1994) as adapted from Abdelgadir et al. (1987) and Schams
(1983), by manual separation through Sep-Pak' Plus C18 cartridges (Waters
Chromatography Division, Millipore Corp., Milford, MA). Briefly, approximately 4000
cpm.100
[3H] -OT (specific activity of 48.5 CimM-1; Dupont NEN', Boston, MA)
was added to 1 ml aliquots of every sample and vortexed briefly for 10 sec. Samples
were then added to a prewetted (2 ml methanol) and rinsed (5 ml ddH20) column
apparatus consisting of a 10 ml syringe with a Sep-Pak" Plus C18 cartridge attached and
manually driven through the column at a slow rate. Following the sample, 1 ml of 0.05
M phosphate buffer, then 4 ml 1.5% acetic acid were slowly pushed through the column.
The final fractions, 2 ml tetrahydrofuran hormone laden extracts, were captured in 12 x
75 mm glass borosilicate test tubes and evaporated in a 37° C water bath under air.
Once dry, samples were reconstituted in I ml assay buffer (0.05 M sodium phosphate, 50
mM EDTA, 0.5 mgm1-1 gelatin) and a 100
aliquot from each sample removed and
counted for extraction efficiency. The remaining portion of the extracts was parafilmed
44
and stored overnight at 4° C for use in the OT radioimmunoassay the next day. Mean
extraction efficiency for testicular sera OT was 53.3 ± 6.0%.
Radioimmunoassay was performed for OT using OT standards (0.12 to 32.0
pgtube1) in assay buffer (described above). Oxytocin standards (100 p.1tube') in
triplicate or extracted samples in duplicate (100 µltube') were pipetted into 12 x 75 mm
glass borosilicate tubes followed by the addition of 300 pl of assay buffer to total count
and non-specific binding tubes, and 100 pl of assay buffer to all other tubes. One
hundred microliters per tube OT antibody (generously provided by Dr. Dieter Schams,
Technical University of Munich, Freising-Weihenstephan, Germany) were added to
standard and sample tubes at a final dilution of 1:2000. Tubes were vortexed briefly for
20 sec, parafilmed and covered with aluminum foil, then incubated for 24 h at 4° C.
After incubation, 100 p.1 of 6000 cpm-100 pl [I'25] -OT (specific activity of 2200
CimM-1; Dupont NEN', Boston, MA) was quickly added to each assay tube. Tubes
were then vortexed briefly for 10 sec, reparafimed and foiled, and incubated for 48 h at
4°C. Following incubation, 400 pi dextran-coated charcoal [0.66% wv-1 washed neutral
norit charcoal (Sigma), 0.066% wv1 Dextran T-70 (Pharmacia, Uppsala, Sweden)] in
phosphate-buffered saline (PBS; 0.01 M NaPO4 pH 7.0, 0.14 M NaCI, 1:10,000
thimerosol) was added rapidly to all other tubes while in an ice bath. Tubes were
vortexed briefly for 10 sec, incubated at 4° C for 15 min, then centrifuged at 2540 x g for
15 min. Five hundred microliters of supernatant from each tube were removed and
placed into 12 x 75 mm glass borosilicate tubes and counted on a Beckman 5500 gamma
45
counter for 4 min each. Intra- and interassay coefficients of variation were 3.9% and
17.9±0.8%, respectively. Sensitivity of the OT assay was 0.125 pgtube1.
Steroid Extraction
Steroids were extracted from 1 ml of serum diluted 1:1 with ddH,0 utilizing 6-7
ml diethyl ether by manual inversion of samples for 2 min. Samples were centrifuged at
1000 x g at 4° C for 15 min, then snap frozen in a methanol-dry ice bath. Ether extracts
were quickly poured into 12 x 75 mm borosilicate tubes and evaporated on a heating
block at 37° C under air. Tubes were reconcentrated two times with 200 p.1 aliquots of
diethyl ether and redried as stated above. Extracts were reconstituted with 100 p.1
hexane:benzene:methanol combination solvent (62:20:13), then subsequently subjected
to chromatographic separation on glass columns [14 x 0.92 cm ID] packed with 1 g of
Sephadex LH-20 (Pharmacia, Inc., Piscataway, N.J.). After application of the hormone
extracts to the columns, the columns were rinsed with an additional 100 pl of the
combination solvent. Steroids were separated on the columns with the
hexane:benzene:methanol combination solvent mentioned above. Estrone fractions (3 ml
collected between 6 and 9 ml of added eluent) and estradiol fractions (5 ml collected
between 10 and 15 ml added eluent) were collected. Extracts were evaporated as
previously described and reconstituted with 1 ml of ethanol. The estradiol and estrone
antibodies used for radioimmunoassay were produced as described by Resko et al.
(1975). The extraction efliciences for estradiol and estrone were 65.0% and 78.7%,
respectively. Estradiol and estrone fractions were immediately assayed for hormone
content.
46
Estradiol Radioimmunoassay
Radioimmunoassay as described by Resko et al. (1980) was performed for
estradiol in a single assay using estradiol standards (2.5 to 150 pgtube1) in ethanol.
Ethanol standards (100 pl-tube1) in triplicate or extracted samples (9001_1J-tube) were
pipetted into 12 x 75 mm glass tubes and evaporated on a heating block at 37° C under
air. When samples were dry, 100 ill of 0.1% phosphate-buffered saline with gelatin
(GPBS), was added to the total count and non-specific binding tubes. Standards and
sample tubes received 100 ill of estradiol antibody in 0.1% GPBS then 100 pi of 4500
cpm.1001A1-1 [3H]- estradiol (specific activity of 95.3 CimM-1, Dupont NEN', Boston,
MA) in 0.1% GPBS were added to all tubes. Tubes were then briefly vortexed and
incubated overnight at 4° C. The following day, in an ice bath, 100 p1 of cold 0.5%
GPBS was added to all tubes. An additional 1 ml of cold 0.5% GPBS added to total
count tubes and 1 ml dextran-charcoal [2.5 21-1 washed neutral norit charcoal (Fisher
Scientific, Pittsburgh, PA), 0.25 g1-1 Dextran T-70 (Pharmacia, Uppsala, Sweden)] in
PBS were added rapidly to all other tubes. Tubes were vortexed and incubated at 4° C
for 15 min, then centrifuged at 2000 x g for 10 min. The supernatant from each tube
was decanted into a 7 ml mini-polypropylene scintillation vial containing 3 ml of Packard
Ultima Gold scintillation cocktail. Vials were counted in a Packard 460 beta-liquid
scintillation counter. The intraassay coefficient of variation was approximately 0.63%
(n=1 assay). Sensitivity of the estradiol assay was 2.5 pgtube-1.
47
Estrone Radioimmunoassay
Radioimmunoassay as described by Resko et al. (1980) was performed for
estrone in a single assay using estrone standards (5.0 to 150 pgtube1) in ethanol.
Ethanol standards (100 µltube"') in triplicate or extracted samples (900 pltube1) were
pipetted into 12 x 75 mm glass tubes and evaporated on a heating block at 37° C under
air. When samples were dry, 100 .tl of 0.1% GPBS, was added to the total count and
non-specific binding tubes. Standards and sample tubes received 100 111 of estrone
antibody in 0.1% GPBS then 100 1A1 of 4500 cpm-1001A11 [3H]- estrone (specific activity
of 85 CimM-1, Dupont NEN', Boston, MA) in 0.1% GPBS were added to all tubes.
Tubes were then briefly vortexed and incubated overnight at 4° C. The following day, in
an ice bath, 100 µl of cold 0.5% GPBS was added to all tubes. An additional 1 ml cold
0.5% GPBS was added to the total count tubes and 1 ml dextran-charcoal [2.5 g1-'
washed neutral norit charcoal (Fisher Scientific, Pittsburgh, PA), 0.25 g1-1 Dextran T-70
(Pharmacia, Uppsala, Sweden)] were added rapidly to all other tubes. Tubes were
vortexed and incubated at 4°C for 15 min, then centrifuged at 2000 x g for 10 m. The
supernatant from each tube was decanted into a 7 ml mini-polypropylene scintillation vial
containing 3 ml of Packard Ultima Gold scintillation cocktail. Vials were counted in a
Packard 460 beta-liquid scintillation counter. The intraassay coefficient of variation was
approximately 6.0% (n=1 assay). Sensitivity of the estrone assay was 5.0 pgtube-1.
48
Statistical Analysis
Behavioral data for mounts and ejaculations were analyzed utilizing the sign test
(SAS 6.12, 1996). Asexual rams consistently showed no sexual behavior and therefore
were not evaluated statistically.
Differences among mean testicular venous sera OT concentrations between
heterosexual, MORS and asexual rams were determined by one-way ANOVA (n=21).
Mean testicular venous serum OT concentration in comparison to mean testicular arterial
serum OT concentration was statistically evaluated for difference by paired t-test (n=8).
Differences among serum estrone and estradiol concentrations in intact and
castrated rams implanted with E2 were analyzed by repeated measures ANOVA using the
general linear model procedures of SAS. Sources of variation for the statistical analysis
of data for each hormone were Group (heterosexual, MORS, asexual), Treatment
(intact, castrated plus E,) and Group x Treatment. Group differences among mean sera
concentrations of estrone and estradiol during the intact period and after castration plus
exogenous E, replacement were determined by the contrast procedure of SAS.
RESULTS
Mounting behavior declined after castration for MORS (p<0.05) and
heterosexual rams (p>0.10; Figure 2). Castration reduced the number of
ejaculations/intromissions by heterosexual rams (p<0.05), but not by MORS (p>0.10;
Table 1). Asexual rams consistently displayed no sexual behaviors over the entire
duration of the investigation.
49
60
'!,
50
HETEROS
MORS
ASEXUAL
40
30
20
10
Intact
Castrated Castrated
+ E2
Figure 2. Mean (±SE) mounting behavior observed in heterosexual, MORS and asexual
rams while intact, after castration and after castration with E2 replacement.
*Castrated and castrated plus E2 vs. intact (p<0.05).
50
Table 1. Mean (±SE) ejaculations of heterosexual, MORS and asexual rams while intact,
after castration and while castrated with E2 therapy.
Treatment Period
RAM GROUP
Intact
Castrated
Castrated + E2
Heterosexual
14.5±1.1
4.3±1.1*
2.8±0.5*
MORS
1.9±0.83
0.14±0.14
0.0±0.0
Asexual
0.0±0.0
0.0±0.0
0.0±0.0
*Means differ from means of rams while intact (p<0.05)
51
Mean testicular venous sera OT concentrations were not significantly different
among heterosexual, MORS and asexual rams (117.6±11.7, 128.6±21.1 and 102.9±20.1
pgrn11, respectively; p>0.05; n=21; Figure 3). There was no evidence that the testes of
the ram synthesized OT because the overall mean testicular venous serum OT
concentration (96.3±9.4 pgm1-1; n=8) did not differ significantly from overall mean
testicular arterial serum concentration of OT (106.3±17.5 pgm14; p>0.25; n=8; Figure
4).
Endogenous serum estrone and estradiol concentrations were not significantly
different among intact heterosexual, MORS and asexual rams (p>0.10; Table 2).
Analysis of serum estrone and estradiol concentrations following castration and
exogenous E2 replacement, likewise, did not differ significantly among heterosexual,
MORS and asexual rams (p>0.50 and p>0.30, respectively; Table 2). Statistically,
treatment (intact or post-castration with E, replacement) had no effect on serum
concentrations of estradiol among groups before or after castration with exogenous E2
therapy indicating that exogenous hormone replacement resulted in systemic levels of the
steroid comparable to physiological levels observed in the intact animals. However,
mean serum concentrations of estrone were significantly lower in castrated rams
administered exogenous E2 than in the same animals before castration (p<0.02; Table 2).
DISCUSSION AND CONCLUSION
Typically following castration, animal sexual desire is quelled and a reduction in
libido is experienced due to the lack of the stimulatory effects of testosterone at the level
52
0111.,
gr
E
160
atm
140
uc 120
2
'cm)
100
0
80
u)
60
c
40
cu
a
as
20
0
Heterosexuals MORS
Asexual
Ram Type
Figure 3. Mean (±SE) testicular venous OT concentrations in heterosexual, MORS and
asexual rams.
53
140
120
0.6)
u
aS.
0
100
80­
51
60
0
40­
x
C
as
o
E
20
0
Venous
Arterial
Blood Source
Figure 4. Mean (±SE) testicular venous and arterial OT concentrations in rams.
54
Table 2. Mean (±SE) serum concentrations of estradiol and estrone (pgm1-1) in
heterosexual, MORS and asexual rams while intact and after castration with E2
replacement.
1 (pmi -)
Estraulm
Estrone tprmi -)
RAM
GROUP
Intact
Heterosexual
8.48±0.79
10.4±1.3
14.1±1.7
11.1±1.3*
MORS
7.57±0.65
8.24±0.67
12.8±2.0
9.14±1.7*
Asexual
6.67±0.77
8.32±1.1
11.6±0.93
8.83±0.73*
Castrated
Intact
+E2
Castrated
+E2
* Differs from means for intact rams (p<0.02)
55
of the hypothalamus. In the male, testosterone secreted by the testes is aromatized to
estradiol in the hypothalamus, or more specifically the MPOA, which is thought to
facilitate the maintenance of sexual behavior in some male mammalian and avian species
(Beyer et al., 1976; Balthazart et al., 1992). Hence, it is not surprising that mounting
behavior was significantly reduced in MORS after castration and that heterosexual rams
showed a similar declining trend in mounting behavior. Had the interval between
castration and exogenous E2 replacement been extended, the reduction in mounting
behavior witnessed during behavioral testing may have been more significant.
Conversely, it has also been shown that with an increase in the interval between
castration and exogenous hormone replacement, there is an inverse relationship to the
responsiveness of the system to hormonal restoration (Clark et al., 1995); consequently,
it was felt the 6 wk period between castration and E2 replacement would have been
sufficient to observe the effects of endogenous hormone removal and also to observe
more immediate effects of exogenous E2 on sexual behavior in rams. With regard to
ejaculation, castration was suffficient to drastically reduce the "ejaculatory" capacity of
heterosexual rams. On the other hand, ejaculatory intromissions achieved by MORS did
not appear significantly reduced following castration due to the fact that MORS rarely
experienced ejaculation prior to castration. This was due, in part, to the angle and
anatomy of MORS' preferred sexual partner, the teaser ram, being non-condusive to
intromission, thus precluding successful ejaculatory attempts by MORS the majority of
the time. Accordingly, little difference could be observed with regard to MORS
ejaculatory intromissions during the intact, castrated or castrated plus E, periods.
56
Although OT has been implicated as a hormone playing a role in sexual behavior
(see review Carter, 1992), there was no significant difference among ram groups in
testicular serum concentrations of OT; therefore, there was no evidence to suggest that
changes in production of OT contributed to sexual orientation in rams. It has been
suggested by Nicholson and Jenkin (1995), that the testes of the ram might be a site of
OT synthesis. Hence, testicular venous and arterial OT concentrations were evaluated in
eight of the animals to determine if the testes of the ram synthesized OT. It was
expected that if the testes of the ram synthesized OT, testicular venous serum
concentrations of OT would exceed OT concentrations in testicular arterial serum.
However, there was no significant difference between OT concentrations in the testicular
venous and arterial sera in the rams, thus providing evidence that the testes of the ram
does not synthesize OT. However, testicular synthesis of OT cannot be completely ruled
out because testicular venous and arterial bloods were sampled on the day after sexual
behavioral testing during surgical castration. It is possible that testicular venous and
arterial bloods sampled immediately following sexual stimulation might have revealed
that OT is synthesized by the testes of the ram as a direct response to sexual stimuli.
Systemic concentrations of estradiol were not significantly different among
groups of intact rams. This was unexpected due to the fact that Resko et al. (1996)
recently detected a significant difference in systemic concentrations of estradiol between
heterosexual rams compared to MORS (15 pgm1-1 vs. 8 pgml
respectively). The
overall mean estradiol concentration of the intact animals we investigated was 7.57±0.52
pgm1-1, comparable to levels measured by Resko et al. (1996) for MORS. When
exposed to ewes in estrus, sexually active heterosexual rams typically experience
57
increased plasma LH concentrations (D'Occhio et al., 1983; Schanbacher et al., 1987;
Perkins et al., 1995), and subsequent increased testicular testosterone synthesis and
secretion (D'Occhio et al., 1983). However, studies by Perkins and Fitzgerald (1992),
as well as Perkins et al. (1995), have demonstrated that rams displaying homosexual
tendencies do not experience the LH surge in response to sexual activity characteristic of
heterosexual rams. Therefore, MORS lack the capacity to synthesize testosterone to the
same extent as heterosexual rams. This ultimately results in lower concentrations of
testosterone, less testosterone available to upregulate aromatase activity, and thus a
reduced capacity to aromatize the available testosterone to estradiol. Perhaps, if blood
samples were collected immediately after sexual stimulation there might have been a
significant difference in estradiol concentrations observed between heterosexual rams
and MORS. Such a difference might be anticipated, because heterosexuals would have
experienced an LH surge that would have stimulated a cascade effect on the pathway to
increased systemic concentrations of estradiol.
Analysis of serum estradiol concentrations after castration with E2 replacement
did not result in a significant Group x Treatment interaction suggesting that sufficient E2
had been administered to the rams to mimic the concentrations of estradiol normally
present in the intact rams. In spite of physiological E2 replacement in castrated rams, the
treatment was not sufficient to restore, reorient, or in the case of asexual rams, to
stimulate sexual behavior. It is possible that adequate quantities of E2 at the cellular level
in the brain were not available to facilitate or promote appropriate sexual behavior in the
ram. Further research, including physiological E, implantation directly into the MPOA
or pharmacological systemic administration of E-, will have to be conducted to determine
58
whether exogenous E2 can restore, reorient or stimulate sexual behaviors in castrated
heterosexual, MORS and asexual rams.
Interestingly, systemic estrone concentrations did not differ significantly among
groups while rams were intact or after castration and treatment with E2. However,
systemic concentrations of estrone differed significantly within group between treatments
(intact vs. castrated plus E2). Concentrations of this estrogen were reduced in all ram
groups after castration and E2 replacement compared with the same rams while intact.
The observed differences in serum concentration of estrone may be due to removal of the
testes, which eliminated endogenous androgens that ordinarily give rise to estrone.
In conclusion, these results suggest that restoration of E2 concentrations in the
systemic circulation of adult rams (regardless of sexual orientation) to physiological
amounts cannot maintain or reestablish sexual behaviors to levels observed prior to
castration. Furthermore, this treatment does not seem to affect sexual orientation.
Additionally, these data suggest that the testes is not a source of OT and that this
nonapeptide does not have a major effect in promoting sexual behavior of rams.
59
BIBLIOGRAPHY
Abdelgadir, SE, LV Swanson, JE Oldfield and F Stormshak. 1987. Prostaglandin F2­
induced release of oxytocin from bovine corpora lutea in vitro. Biol Reprod
37:550-557.
Adkins-Regan, E. 1988. Sex hormones and sexual orientation in animals.
Psychobiology 16:335-347.
Alexander, BM, A Perkins, EA Van Kirk, GE Moss and JA Fitzgerald. 1993.
Hypothalamic and hypophyseal receptors for estradiol in high and low sexually
performing rams. Horm Behav 27:296-307.
Allen, LS and RA Gorski. 1992. Sexual orientation and the size of the anterior
commissure in the human brain. Proc Natl Acad Sci USA 89:7199-7202.
Amann, RP. 1981. A critical review of methods for evaluation of spermatogenesis from
seminal characteristics. J Androl 2:37-58.
Amann, RP and BD Schanbacher. 1983. Physiology of male reproduction. J Anim Sci
57(Suppl 2):380-403.
Arletti, R, C Bazzani, M Castelli and A Bertolini. 1985. Oxytocin improves male
copulatory performance in rats. Horm Behav 19:14-20.
Attal, J. 1969. Levels of testosterone, androstenedione, estrone and estradiol -17(3 in the
testes of fetal sheep. Endocrinology 85:280-289.
Bakker, J, J van Ophemert and AK Slob. 1993. Organization of partner prefence and
sexual behavior and its nocturnal rhymicity in male rats. Behav Neurosci
107:1049-1058.
Balthazart, J, C Surlemont and N Harada. 1992. Aromatase as a cellular marker of
testosterone action in the preoptic area. Physiol Behav 51:395-409.
Barraclough, CA. 1961. Production of anovulatory, sterile rats by single injections of
testosterone propionate. Endocrinology 68:62-67.
Bergland, RM and RB Page. 1978. Can the pituitary secrete directly to the brain?
(Affirmative anatomical evidence). Endocrinology 102:1325 -1338.
Beyer, C, G Morali, K Larsson and P SOdersten. 1976. Steroid regulation of sexual
behavior. J Steroid Biochem 7:1171-1176.
60
Boothby, M, RW Ruddon, C Anderson, D McWilliams and I Boime. 1981. A single
gonadotropin a-subunit gene in normal tissue and tumor derived cell lines. J Biol
Chem 256:5121-5127.
Borg, KE, KL Esbenshade and BH Johnson. 1992. Effects of sexual experience, season
and mating stimuli on endocrine concentrations in the adult ram. Horm Behav
26:87-109.
Butera, PC and JA Czaja. 1989. Effects of intracranial implants of dihydrotestosterone
on the reproductive physiology and behavior of male guinea pigs. Horm Behav
23:424-431.
Carmichael, MS, R Humbert, J Dixon, G Palmisano, W Greenleaf and JM Davidson.
1987. Plasma oxytocin increases in the hyman sexual response. J Clin
Endocrinol Metab 64:27-31.
Carter, CS. 1992. Oxytocin and sexual behavior. Neurosci Biobehav Rev 16:131-144.
Clark, JT, PE Micevych, V Panossian and AK Keaton. 1995. Testosterone-induced
copulatory behavior is affected by the postcastration interval. Neurosci Biobehav
Rev 19:369-376.
Clarke, IJ, RJ Scaramuzzi and RV Short. 1976. Effects of testosterone implants in
pregnant female ewes on their female offsprina. J Ernbryol Exp Morphol 36:87­
99.
Connolly, PB, JVA Choate and JA Resko. 1994a. Effects of exogenous androgen on
brain androgen receptors of the fetal Rhesus monkey. Neuroendocrinology
59:271-271.
Connolly, PB and JA Resko. 1994. Prenatal testosterone differentiates brain regions
controlling gonadotropin release in guinea pigs. Biol Reprod 51:125-130.
Connolly, PB, CE Roselli and JA Resko. 1994b. Aromatase activity in developing
guinea pig brain: ontogeny and effects of exogenous androgens. Biol Reprod
50:436-441.
Cunningham, JG. 1992. Textbook of Veterinary Physiology. W.B. Saunders Company,
Philadelphia. Pp 379-381.
Dacheux, JL, C Pisselet, MR Blanc, MT Hochereau-de-Reviers and M Courot. 1981.
Seasonal variations in rete testis fluid secretion and sperm production in different
breeds of ram. J Reprod Fertil 61:363-371.
61
Dagg, Al. 1984. Homosexual behavior and female-male mounting in mammals-a first
survey. Mammal Rev 14:155-185.
Deviche, P and FL Moore. 1988. Steroidal control of sexual behavior in the roughskinned newt (Taricha granulosa): effects of testosterone, estradiol, and
dihydrotestosterone. Horm Behav 22:26-34.
Dixson, AF. 1993. Effects of testosterone propionate upon the sexual and aggressive
behavior of adult male marmosets (Calliihrix jacchus) castrated as neonates.
Horm Behav 27:216-230.
D'Occhio, MJ, BD Schanbacher and JE Kinder. 1983. Androgenic and oestrogenic
steroid participation in feedback control of luteinizing hormone secretion in male
sheep. Acta Endocrinol (Copenh) 102:499-504.
Donahoe, PK, RL Gate, DT MacLaughlin, J Epstein, AF Fuller, M Takahashi, JP
Coughlin, EG Ninfa and LA Taylor. 1987. Mullerian inhibiting substance: gene
structure and mechanism of action of a fetal regressor. Rec Prog Horm Res
43:431-476.
Fitzgerald, JA and A Perkins. 1994. Effect of morphine and naloxone on LH response
and sexual behavior in rams (Ovis aries). Domest Anim Endocrinol 11:271-279.
FjellstrOm, D, JE Kihlstrom and P Melin. 1968. The effect of synthetic oxytocin upon
seminal characteristics and sexual behavior in male rabbits. J Reprod Fertil
17:207-209.
Ford, JJ. 1983. Postnatal differentiation of sexual preference in male pigs. Horm Behav
17:152-162.
Frayne, J and RD Nicholson. 1995. Effect of oxytocin on testosterone production by
isolated rat Leydig cells is mediated via specific oxytocin receptor. Biol Reprod
52:1268-1273.
Friedan, EH, JK Patkin and M Mills. 1968. Effects of follicle-stimulating hormone
(FSH) upon steroid aromatization in vitro. Proc Soc Exp Biol Med 129:606­
609.
Geist, V. 1971. Mountain sheep: A Study in Behavior and Evolution. University of
Chicago Press, Chicago.
Giantonio, GW, NL Lund and AA Gerall. 1970. Effect of diencephalic and
rhinencephalic lesions on the male rat's sexual behavior. J Comp Physiol Psycho!
73 38-46
62
Ginton, A and A Merari. 1977. Long range effects of MPOA lesion on mating behavior
in the male rat. Brain Res 120:158-163.
Goy, RW, CH Phoenix and WC Young. 1962. A critical period for the suppression of
behavioral receptivity in adult female rats by early treatment with androgen.
Anat Rec 142:307.
Grady, KL and CH Phoenix. 1963. Hormonal determinants of mating behavior; the
display of feminine behavior by adult male rats castrated neonatally. Amer Zool
3:482-483.
Gray, JM, AA Nunez, LI Siegel and GN Wade. 1979. Effects of testosterone on body
weight and adipose tissue: role of aromatization. Physiol Behav 23:465-469.
Green, JD and GW Harris. 1946. The neurovascular link between the neurohypophysis
and adenohypophysis. J Endocrinol 5:136-149.
Griffen, JE and SR Ojeda. 1996. Textbook of Endocrine Physiology. 3rd Edition.
Oxford University Press, New York. Pp 167, 207.
Guyton, AC and JE Hall. 1996. Textbook of Medical Physiology. 9th Edition. WB
Saunders Company, Philadelphia. Pp 1008.
Hafez, ESE. 1996. Reproduction in Farm Animals. 6th Edition. Lea & Febiger,
Philadelphia. Pp 7.
Hamer, DH, S Hu, VL Magnuson, N Hu and AML Pattatucci. 1993. A linkage
between DNA markers on the X chromosome and male sexual orientation.
Science 261:321-327.
Hart, BL, CM Haugen and DM Peterson. 1973. Effects of medial preoptic-anterior
hypothalamic lesions on mating behavior of male cats. Brain Res 54:177-191.
Hillegaart, V, P Alster, K Uvnas-Moberg and S Ahlenius. 1998. Sexual motivation
promotes oxytocin secretion in male rats. Peptides 19:39-45.
Hing, ALY and JR Carlson. 1996. Male-male courtship behavior induced by ectopic
expression of the Drosophila white gene: role of sensory function and age. J
Neurobiol 30:454-464.
Hooker, CW. 1944. The postnatal history and function of the interstitial cells of the
testis of the bull. Amer J Anat 74:1-37.
Huggins, C and PV Moulder. 1945. Estrogen production by the Sertoli cell tumors of
the testes. Cancer Res 5:510-514.
63
Hughes, AM, BJ Everitt, SL Lightman and K Todd. 1987. Oxytocin in the central
nervous system and sexual behavior in male rats. Brain Res 414:133-137.
Josso, N, JY Picard and D Tran. 1976. The anti-Mtillerian hormone. Rec Prog Horm
Res 33:117-167.
Josso, N, JY Picard, JL Dacheux and M Courot. 1979. Detection of anti-Mullerian
activity in boar rete testis fluid. J Reprod Fertil 57:397-400.
Jost, A, B Vigier, J Prepin and JP Perchellet. 1973. Studies on sex differentiation in
mammals. Rec Prog Horm Res 29:1-41.
Karsch, FJ, EL Bittman, DL Foster, RL Goodman, SJ Legan and JE Robinson. 1984.
Neuroendocrine basis of seasonal reproduction. Rec Prog Horm Res 40:185­
232.
Karsch, FJ, DJ Dierschke, RF Weick, T Yamaji, J Hotchkiss and E Knobil. 1973.
Positive and negative feedback control by estrogen of luteinizing hormone
secretion in the Rhesus monkey. Endocrinology 92:799-804.
Karsch, FJ, SJ Legan, KD Ryan and DL Foster. 1980. Importance of estradiol and
progesterone in regulating LF1 secretion and estrous behavior during the sheep
estrous cycle. Biol Reprod 23:404-413.
Kellis, JT and LE Vickery. 1987. Purification and characterization of human placental
aromatase cytochrome P-450. J Biol Chem 262:4413-4420.
Knickerbocker, JJ, HR Sawyer, RP Amann, FR Tekpetey and GD Niswender. 1988.
Evidence for the presence of oxytocin in the ovine epididymis. Biol Reprod
39:391-397.
Knight, TW and DR Lindsay. 1970. Short- and long-term effects of oxytocin on quality
and quantity of semen in rams. J Reprod Fertil 21:523-529.
Ladewig, J and BL Hart. 1980. Flehmen and vomeronasal organ function in male goats.
Physiol Behav 24:1067-1071.
LeVay, S. 1991. A difference in hypothalamic structure between heterosexual and
homosexual men. Science 253:1034-1037.
Levis, DG and JJ Ford. 1989. The influence of androgenic and estrogenic hormones on
sexual behavior in castrated adult male pigs. Horm Behav 23:393-411.
64
Lin, D, T Sugawara, JF Strauss III, BJ Clark, DM Stocco, P Saenger, A Rogol and WL
Miller. 1995. Role of steroidogenic acute regulatory protein in adrenal and
gonadal steroidogenesis. Science 267:1828-1831.
Lindsay, DR. 1965. The importance of olfactory stimuli in the mating behavior of the
ram. Anim Behav 13:75-78.
Longcope, C, T Kato and R Horton. 1969. Conversion of blood androgens to
estrogens in normal adult men and women. J Clin Invest 48:2191-2201.
Longcope, C, JH Pratt, SH Schneider and SE Fineberg. 1976. In vivo studies on the
metabolism of estrogens by muscle and adipose tissue in normal males. J Clin
Endocrinol Metab 43:1134-1145.
Longcope, C, JH Pratt, SH Schneider and SE Fineburg. 1978. Aromatization of
androgens by muscle and adipose tissue in vivo. J Clin Endocrinol Metab
46:146-152.
Luttge, WG and NR Hall. 1973. Differential effectiveness of testosterone and its
metabolites in the induction of male sexual behavior in two strains of albino mice.
Horm Behav 4:31-43.
MacLusky, NJ and F Naftolin. 1981. Sexual differentiation of the central nervous
system. Science 211:1294-1302.
Melin, P and JE Kihlstrom. 1963. Influence of oxytocin on sexual behavior in male
rabbits. Endocrinology 73:433-435.
Melis, MR, A Ariolas and GL Gessa. 1986. Oxytocin-induced penile erection and
yawning: site of action in the brain. Brain Res 398:259-265.
Mendelson, CR, EE Wright, CT Evans, JC Porter and ER Simpson. 1985. Preparation
and characterization of polyclonal and monoclonal antibodies against human
aromatase cytochrome P-450 and their use in its purification. Archs Biochem
Biophys 243:480-491.
Michael, RP, D Zumpe and RW Bonsall. 1990. Estradiol administration and the sexual
activity of castrated male Rhesus monkeys (Macaca miilatta). Horm Behav
24:71-88.
Moltz, H. 1990. E-series prostaglandins and arginine vasopressin in the modulation of
male sexual behavior. Neurosci Biobehav Rev 14:109-115.
65
Murphy, MR, JR Seckl, S Burton, SA Check ley and SL Lightman. 1987. Changes in
oxytocin and vasopressin secretion during sexual activity in men. J Clin
Endocrinol Metab 65:738-741.
Neubert, DW, DR Nelson, MJ Coon, RW Estabrook, R Feyereisen, Y Fujii-Kuriyama,
FJ Gonzalez, FP Guengerich, IC Gunsalus, EF Johnson, JC Loper, R Sato, MR
Waterman and DJ Waxman. 1991. The P450 superfamily: update on new
sequences, gene mapping and recommended nomenclature. DNA 10:1-14.
Nicholson, I-ID, SEF Guldenaar, GJ Boer and BT Pickering. 1991. Testicular oxytocin:
effects of intratesticular oxytocin in the rat. J Endocrinol 130:231-238.
Nicholson, I-ID and L Jenkin. 1995. Oxytocin and prostatic function. In: R Ivell and
JA Russell, Eds. Oxytocin. Cellular and Molecular Approaches in Medicine and
Research. Plenum Press, New York. Advances in Experimental Medicine and
Biology 395:530.
Norris, DO. 1997. Vertebrate Endocrinology. 3rd Edition. Academic Press, San
Diego. Pp 146-149, 380.
Nottebohm, F and AP Arnold. 1976. Sexual dimorphism in vocal control areas of the
songbird brain. Science 194:211-213.
Orwig, KE, JE Bertrand, B-R Ou, NE Forsberg and F Stormshak. 1994. Involvement
of protein kinase-C, calpains, and calpastatin in prostaglandin Fla- induced
oxytocin secretion from the bovine corpus luteum. Endocrinology 134:78-83.
Payne, AH, RP Kelch, SS Musich and ME Halpern. 1976. Intratesticular site of
aromatization in the human. J Clin Endocrinol Metab 42:1081-1087.
Perkins, A and JA Fitzgerald. 1992. Luteinizing hormone, testosterone, and behavioral
response of male-oriented rams to estrous ewes and rams. J Anim Sci 70:1787­
1794.
Perkins, A, JA Fitzgerald and GE Moss. 1995. A comparison of LH secretion and brain
estradiol receptors in heterosexual and homosexual rams and female sheep.
Horm Behav 29:31-41.
Perkins, A, JA Fitzgerald and EO Price. 1992a. Sexual performance of rams in serving
capacity tests predicts success in pen breeding. J Anim Sci 70:2722-2725.
Perkins, A, JA Fitzgerald and EO Price. 1992b. Luteinizing hormone and testosterone
response of sexually active and inactive rams. J Anim Sci 70:2086 -2093.
66
Plapinger, L and BS Mc Ewen. 1978. Gonadal steroid-brain interactions in sexual
differentiation. In: Hutchison, JB, Ed. Biological Determinants of Sexual
Behavior. John Wiley & Sons Ltd, New York. Pp 199-202.
Raynaud, JP, C Mercier-Bodard and EE Baulieu. 1971. Rat estrogen binding plasma
protein (EBS). Steroids 18:767-788.
Resko, JA, WE Ellinwood, LM Pasztor and AE Buhl. 1980. Sex steroid in the
umbilical circulation of fetal Rhesus monkeys from the time of gonadal
differentiation. J Clin Endocrinol Metab 50:900-905.
Resko, JA, A Perkins, CE Roselli, JA Fitzgerald, JVA Choate and F Stormshak. 1996.
Endocrine correlates of partner preference behavior in rams. Biol Reprod
55:120-126.
Resko, JA, JG Ploem and HL Stadelman. 1975. Estrogens in fetal and maternal plasma
of the Rhesus monkey. Endocrinology 97:425-430.
Roselli, CE and JA Resko. 1986. Effects of gonadectomy and androgen treatment on
aromatase activity in the fetal monkey brain. Biol Reprod 35:106-112.
Roselli, CE and JA Resko. 1987. The distribution and regulation of aromatase activity
in the central nervous system. Steroids 50:495-508.
Roselli, CE and JA Resko. 1989. Testosterone regulates aromatase activity in discrete
brain areas of male Rhesus macaques. Biol Reprod 40:929-934.
Ryan, KJ, F Naftolin, V Reddy, F Flores and Z Petro. 1972. Estrogen formation in the
brain. Amer J Obstet Gynecol 114:454-460.
Ryner, LC, SF Goodwin, DH Castrillion, A Anand, A Villella, BS Baker, JC Hall, BJ
Taylor and SA Wasserman. 1996. Control of male sexual behavior and sexual
orientation in Drosophila by the fruitless gene. Cell 87:1079-1089.
Sanborn, BM, JS Elkington, A Steinberger and E Steinberger. 1975. Androgen binding
in the testis: in vitro production of androgen binding protein (ABP) by Sertoli
cell cultures and measurement of nuclear bound androgen by a nuclear exchange
assay. Curr Top Mol Endocrinol 2:293-309.
SAS (Version 6.12). 1996. SAS Institute, Cary, NC.
Schams, D. 1983. Oxytocin determination by radioimmunoassay Ill. Improvement to
subpicogram sensitivity and application to blood levels in cyclic cattle. Acta
Endocrinol (Copenh) 103:180-183.
.
67
Schanbacher, BD. 1979. Relationship of in vitro gonadotropin binding to bovine testes
and the onset of spermatogenesis. J Anim Sci 48:591-597.
Schanbacher, BD and JJ Ford. 1979. Photoperiodic regulation of ovine
spermatogenesis: relationship to serum hormones. Biol Reprod 20:719-726.
Schanbacher, BD, MP Johnson and DJ Tindall. 1987. Androgenic regulation of
luteinizing hormone secretion: relationship to androgen binding in sheep
pituitary. Biol Reprod 36:340-350.
Schindler, AE. 1975. Steroid metabolism of fetal tissues. II. Conversion of
androstenedione to estrone. Amer J Obstet Gynecol 123:265-268.
Schumacher, M and J Balthazart. 1986. Testosterone-induced brain aromatase is
sexually dimorphic. Brain Res 370:285-293.
Setchell, BP. 1980. The functional significance of the blood-testis barrier. J Androl
1:3-10.
Short, RV. 1960. The biosynthesis and secretion of adrenocortical steroids. In: Clark,
F and JK Grant, Eds. Biochem Soc Symp No 18. Cambridge University Press,
Cambridge. Pp 59.
Silver, GV and E0 Price. 1986. Effects of individual vs. group rearing on the sexual
behavior of prepubertal beef bulls: mount orientation and sexual responsiveness.
Appl Anim Behav Sci 15:287-294.
Simpson, ER, MS Mahendroo, GD Means, MW Kilgore, CJ Corbin and CR Mendelson.
1993. Tissue-specific promotors regulate aromatase cytochrome P450
expression. J Steroid Biochem Mol Biol 44:321-330.
Slaunwhite, WR, MA Karsay, A Hollmer, AA Sandberg and K Niswander. 1965. Fetal
liver as an endocrine tissue. Steroids (Suppl 2):21 1 -221.
Slimp, JC, BL Hart and RW Goy. 1978. Heterosexual, autosexual and social behavior
of adult male Rhesus monkeys with preoptic-anterior hypothalamic lesions.
Brain Res 142:105-122.
Stoneham, MD, BJ Everitt, S Hansen, SL Lightman and K Todd. 1985. Oxytocin and
sexual behavior in the male rat and rabbit. J Endocrinol 107:97-106.
Swaab, DF and MA Hofman. 1990. An enlarged suprachiasmatic nucleus in
homosexual men. Brain Res 537:141-148.
68
Swaab, DF and MA Hofman. 1995. Sexual differentiation of the h hypothalamus in
relation to g sexual orientation. Trends Neurosci 18:264-270.
Swaab, DF, AK Slob, EJ Houtsmuller, T Brand and JN Zhou. 1995. Increased number
of vasopressin neurons in the suprachiasmatic nucleus (SCN) of 'bisexual' adult
male rats following perinatal treatment with the aromatase blocker ATD. Dev
Brain Res 85:273-279.
Swierstra, EE. 1968. Cytology and duration of the cycle of the seminiferous epithelium
of the boar; duration of spermatazoa transit through the epididymis. Anat Rec
161:171-185.
Thompson Jr, ER and PK Siiteri. 1974. Utilization of oxygen and reduced nicotinamide
adenine dinucleotide phosphate by human placental microsomes during
aromatization of androstenedione. J Biol Chem 249:5364-5368.
Uriel, J, B de Nechaud and M Dupiers. 1972. Estrogen-binding properties of rat,
mouse, and man fetospecific serum proteins. Demonstration by immuno­
autoradiographic methods. Biochem Biophys Res Commun 46:1175-1180.
Vittek, J, K Altman, GG Gorgen and AL Southren. 1974. The metabolism of 7-a-3H­
testosterone by rat mandibular bone. Endocrinology 94:325-329.
Vogelmayr, JK. 1975. Output of spermatozoa and fluid by the testis of the ram and its
response to oxytocin. J Reprod Fertil 43:119-122.
Wallis, CJ and WG Luttge. 1975. Maintenance of male sexual behavior by combined
treatment with oestrogen and dihydrotestosterone in CD-1 mice. J Endocrinol
66:257-262.
Wathes, DC. 1984. Possible actions of gonadal oxytocin and vasopressin. J Reprod
Fertil 71:315-345.
Watson, JT and E Adkins-Regan. 1989. Testosterone implanted in the preoptic area of
male Japanese quail must be aromatized to activate copulation. Horm Behav
23:432-447.
Wilson, JD, FW George and JE Griffin. 1981. The hormonal control of sexual
development. Science 211:1278-1284.
Wislocki, GB and LS King. 1936. The permeability of the hypophysis and
hypothalamus to vital dyes, with a study of the hypophyseal vascular supply.
Amer .1 Anat 58:421.
69
Wood, RI. 1996. Estradiol, but not dihydrotestosterone, in the medial amygdala
facilitates male hamster sex behavior. Physiol Behav 59:833-841.
Wood, RI, V Mehta, CG Herbosa and DL Foster. 1995. Prenatal testosterone
differentially masculinizes tonic and surge modes of luteinizing hormone
secretion in the developing sheep. Neuroendocrinology 62:238-247.
Wood, RI and SW Newman. 1995. The medial amygdaloid nucleus and medial preoptic
area mediate steroidal control of sexual behavior in the male Syrian hamster.
Horm Behav 29:338-353.
Ying, SY. 1988. Inhibins, activins and follistatins: gonadal proteins modulating the
secretion of follicle-stimulating hormone. Endocrine Rev 9:267-293.
Young, WC, RW Goy and CH Phoenix. 1964. Hormones and sexual behavior: broad
relationships between the gonadal homones and sexual behavior. Science
143:212 -218.
Zirken, BR, LL Ewing, N Kromann and RC Cochran. 1980. Testosterone secretion by
rat, rabbit, guinea pig, dog and hamster testes in vitro: correlation with Leydig
cell ultrastructure. Endocrinology 106:1867-1874.
70
APPENDIX
71
EFFECTS OF IN VIVO ESTRADIOL AND IN VITRO LH
TREATMENT ON PROGESTERONE SYNTHESIS BY OVINE
CORPORA LUTEA
ABSTRACT
An experiment was conducted to examine in vitro progesterone production by
corpora lutea maintained by chronic treatment of ewes with estradiol -1713 (E2). Ten
mature ewes were assigned randomly in equal numbers to a treatment and control group.
Ewes were injected s.c. with 2501..tg E2days' or vehicle (corn oil; CO) on days 4 through
14 of the estrous cycle (day 0 = estrus). Corpora lutea were removed on day 15 of the
cycle, weighed and sliced. Aliquots of luteal tissue (97.8±0.8 mg) were placed into
flasks containing 2 ml of Ham's F-12 medium and to which was added the following
treatments in duplicate: 1) 5 ill saline (unincubated control; uninc), 2) 5 1..t1 saline
(incubated control; inc) and 3) 5 ill saline containing ovine LH (incubated+LH; inc+LH)
for a final concentration of 20.0 ngm11. Corpora lutea of E2-treated ewes were larger
and contained more progesterone than luteal tissue of control ewes (uninc, E2-treated
1.60±0.21 vs. controls 0.24±0.13 ngmg tissue; p<0.0005). Incubation alone or in
combination with LH significantly increased progesterone synthesis in corpora lutea from
E2-treated ewes (p<0.04) compared to controls (estradiol: inc, 0.67±0.19, inc+LH,
0.50±0.18 ngmg tissue; controls: inc, 0.09±0.05, inc+LH, 0.15±0.12 ngmg tissue).
There was no overall effect of in vitro treatment with LH on progesterone production
by corpora lutea from E2-treated ewes (p>0.10) or controls (p>0.10). In vivo estradiol
72
treatment of ewes beginning early in the luteal phase of the cycle greatly enhanced the
production of progesterone in the day 15 corpus luteum in vitro, but incubation in the
presence of LH had no effect on progesterone production by corpora lutea from control
or E2-treated animals.
INTRODUCTION
Exogenous estradio1-1713 (E2) has been shown by Piper and Foote (1968) to
prolong the life span of the ovine corpus luteum if administered daily beginning early in
the luteal phase of estrous cycle. Further research conducted by Hazzard and Stormshak
(1997) confirmed the luteotropic effects of chronically administered E2 in the ewe.
These investigators found that ewes chronically treated daily with E2 from day 4 of the
estrous cycle until day 24 post-estrus (day 0 = estrus), caused corpora lutea to be
maintained and prolonged the interestrous interval. Corpora lutea of E7-treated ewes
were heavier on day 15 than those of controls.
Luteinizing hormone (LH) has been demonstrated to be luteotropic in the ewe
(Karsch et al., 1971). Luteinizing hormone is hypothesized to be responsible for the
initial increase in post-ovulatory progesterone synthesis (Niswender and Nett, 1994)
through the facilitation of increased enzymatic conversion of cholesterol to pregnenlone
(Rodgers et al., 1986, 1987; Lauber et al., 1991). Following ovulation and complete
establishment of the formation of the corpus luteum, two distinct cellular types,
differentiated by size are present in the corpus luteum. These cells have been so
identified as small and large luteal cells (Koos and Hansel, 1981) Receptors for LH
73
(Fitz et al., 1982), as well as LH receptor mRNA (Smith et al., 1996) have been
associated with small luteal cells, while large luteal cells appear to be deficient in both
(Fitz et al., 1982; Nelson et al., 1992). Although large luteal cells are the primary source
of basal luteal progesterone production (Fitz et al., 1982; Alila et al., 1988; Nelson et al.,
1992), small luteal cells are responsible for progesterone production under the influence
of LH in vivo (Niswender et al., 1985), as well as in vitro (Rodgers et al., 1983).
The objective of the present study was to evaluate in vitro production of
progesterone by corpora lutea maintained as a consequence of chronic E2 treatment of
ewes beginning early during the estrous cycle.
MATERIALS AND METHODS
Animals
Ten mature ewes were checked twice daily for behavioral estrus with a
vasectomized ram. After at least one estrous cycle of normal duration (15.5-18.0 days),
ewes were assigned randomly to treatment groups. All experimental procedures and
surgeries were performed in accordance with Institutional Animal Care and Use
Committee guidelines.
Experimental Protocol
Ewes were assigned randomly in equal numbers to one of two groups (n=5 each)
receiving 1) corn oil (CO) or 2) 250 tg of E2 (Sigma Chemical Co., St. Louis, MO)
dissolved in 2 ml of corn oil. Ewes in both groups were injected s.c. daily from day 4 of
74
the cycle (day 0=estrus) until day 14. On day 15, ewes were anesthetized by i.v.
injection of 5% sodium pentothal and anesthesia was maintained by halothane-oxygen
inhalation. Ewes were subjected to a midventral laparotomy using aseptic conditions to
expose the ovaries. The uterus and ovaries were exteriorized, all corpora lutea were
counted, enucleated and then transported to the laboratory in sterile, phenol red-free
Ham's F-12 medium (Nutrient Mixture F-12 [Ham], Gibco Laboratories, Inc., Grand
Island, NY) containing 14 mM sodium bicarbonate, 24 mM HEPES and 30 pim1-1
gentamicin (Gibco), pH 7.3.
For determination of progesterone synthesis, corpora lutea were sliced (0.3 mm
thickness), and 97.8±0.8 mg of sliced tissue was aliquoted to six 10 ml Erlenmeyer
flasks, each containing 2 ml of incubation medium. Incubation medium consisted of
Ham's F-12 (as described above) plus 5 ii.gm1-1 insulin, 5 1.1.gm1-1 transferrin and 5
ngtn1-1 selenium (ITS, Sigma Chemical Co., St. Louis, MO). Flasks were designated as
follows: two unincubated controls, two incubated controls and two incubated flasks
containing ovine LH dissolved in 5.0 ml of sterile saline (final concentration 20.0 ngml"
1).
Flasks were gassed with 95% 02-5% CO2 for 6 sec each and capped with silicone
stoppers. Two milliliters of cold ethanol were immediately added to the two unincubated
control flasks to preclude further progesterone synthesis. Contents and two additional
milliliters of ethanol used to rinse the flask were transferred to 20 ml glass scintillation
vials and capped. The remaining flasks were incubated for 2 h at 37° C in a Dubnoff
shaking water bath. After incubation, cold ethanol was added to these flasks to
terminate incubation. Tissue plus medium samples were transferred, rinsed and stored as
75
previously mentioned at -20° C until extraction and determination of progesterone
content by radioimmunoassay.
Progesterone Extraction
Tissue plus medium were extracted and assayed for progesterone using
techniques described by Bertrand and Stormshak (1996) as modified from the procedure
of Koligian and Stormshak (1976). After addition of 100 ill of 20,000 cpm-100
[3H]- progesterone (specific activity of 92 CimM-1, Dupont NEN', Boston, MA) to the
sample for determination of extraction efficiency, each sample was placed in a Duall
ground glass homogenizer. The sample storage vial was rinsed three times with 2 ml
aliquots of ethanol which were added to the homogenizer, then the sample was
homogenized. Homogenates were filtered into 100 ml round bottom flasks through
Whatman No. 1 filter paper cradled in a glass funnel. The glass homogenizer and pestle
were rinsed four times with 2 ml aliquots of ethanol and these rinses were filtered. The
filter paper was rinsed five times with 2 ml aliquots of ethanol, after which the glass
funnel and inner neck of the flask were rinsed with ethanol. Flasks were roto-evaporated
at 45° C until samples were nearly dry, resuspended in 3 ml distilled de-ionized water
and vortexed for 30 sec. Samples were incubated at room temperature for 30 min, then
extracted with 20 ml benzene:hexane (1:2) by vortexing for 2 min. Following storage
overnight at -20° C, the organic phase was decanted into a 20 ml scintillation vial and
evaporated under air. Extract residues were respended in 20 ml ethanol and a 1 ml
aliquot was removed to determine extraction efficiency. Mean extraction efficiency was
76
61.0±7.2%; however, each sample was corrected for its own unique extraction
efficiency.
Progesterone Radioimmunoassay
Radioimmunoassay was performed using progesterone standards ranging from 5
to 2000 pg.100 Ill' in ethanol. Ethanol standards (100 1.1.1 in triplicate) or diluted sample
(100 IA in duplicate) were pipetted into 12 x 75 mm borosilicate glass test tubes and
evaporated under air. When tubes were dry, 1001.11 of phosphate-buffered saline with
gelatin (GPBS; 0.01 M NaPO4 pH 7.0, 0.14 M NaCI, 1:10,000 thimerosol, 0.1% gelatin)
were then added to the total count and non-specific binding tubes. Anti-progesterone11-BSA (generously donated by Dr. Gordon Niswender, Colorado State University, Fort
Collins, CO.), in 100 p.1 GPBS was added to each standard and sample tube at a 1:2400
dilution. Tubes were vortexed and incubated at room temperature for 30 min followed
by the addition of 20,000 cpm [3H]-progesterone (92 CimM-1, Dupont NEI\r, Boston,
MA) in 100 pi GPBS to each tube. Tubes were then vortexed for 10 sec and incubated
overnight at 4° C. The following day, 1 ml of ice-cold GPBS was added to the total
count tubes, and 1 ml of ice-cold dextran-charcoal (2.5 g1-1 washed norit charcoal
[Sigma], 0.25
Dextran T-70 [Pharmacia, Uppsala, Sweden]) in GPBS was added
rapidly to all other tubes. Tubes were vortexed and incubated at 4° C for 15 min, then
centrifuged at 2540 x g for 10 min. The supernatant from each tube was decanted into a
20 ml glass scintillation vial and 6 ml of Ecolume scintillation cocktail (ICN
Pharmaceutical Inc., Irvine, CA) were added. Samples were counted in a Beckman LS­
77
6000 liquid scintillation counter. Standard curves were plotted and unknown
concentrations determined using the RIA AID computer program (Robert Maciel
Associates, Inc., Arlington, MA). Intra- and interassay coefficients of variation were
determined using a 75 pg-100 IA1-1 sample in ethanol (12 tubes per assay), and were 6.2
and 11.0%, respectively (n=14 assays). Sensitivity of the progesterone assay was 0.05
ngtube
Statistical Analysis
Data on weights of corpora lutea were analyzed by analsis of covariance using
number of corpora lutea as the independent variable. Effect of in vivo estradiol
treatment on day 15 luteal progesterone concentrations (unincubated control samples)
was tested for significance by t-test. Data on mean progesterone concentrations in
medium and tissue of duplicate samples subjected to incubation alone or in the presence
of LH were analyzed as a factorially arranged split-plot design using ANOVA. The
luteal concentrations of progesterone in each unincubated control sample was subtracted
from respective luteal concentrations of steroid after incubation alone or incubation in
the presence of LH. In vivo treatment (Estradiol), in vitro treatment (LH) and Estradiol
x LH interaction were included as sources of variation.
RESULTS
On day 15, corpora lutea of E,- treated ewes weighed more than those of controls
but the difference in weight was not significant statistically (mean±SE, controls,
78
332.2±32.0 vs. E2-treated, 392.3±12.8 mg; p>0.10). Corpora lutea of E2-treated ewes
contained more progesterone on day 15 than luteal tissue of control ewes (unincubated
controls: E2-treated, 1.60±0.21 vs. controls, 0.24±0.13 ngmg tissue; p<0.0005).
Incubation alone or in combination with in vitro LH administration significantly
increased progesterone synthesis in corpora lutea from E2-treated ewes (p<0.04)
compared to controls (E2-treated: incubated, 0.67±0.19, incubated with LH, 0.50±0.18
ngmg tissue vs. controls: incubated, 0.09±0.05, incubated with LH, 0.15±0.12 ngmg
tissue; Figure 1). However, addition of LH to the incubation medium failed to affect
progesterone production by corpora lutea from E,-treated ewes (p>0.10) or controls
(p>0. 10).
DISCUSSION AND CONCLUSION
Exogenous estradiol -17J3 (E2) given early in the luteal phase (beginning day 4
after estrus) has been shown to prolong the life span of the ovine corpus luteum in vivo
(Piper and Foote, 1968, 1970; Hazzard and Stormshak, 1997). Therefore, it was not
surprising that day 15 corpora lutea from E2-treated ewes contained more progesterone
than those of control ewes. Incubation of luteal tissue alone resulted in a significant
increase in progesterone production by luteal tissue from E 2-treated ewes. However,
addition of LH to the incubation medium failed to increase progesterone by luteal tissue
from control or E,-treated ewes. The fact that corpora lutea of control ewes were
regressing by day 15 can serve to explain the lack of response of this luteal tissue to
synthesize progesterone in response to incubation or incubation in combination with LH.
79
7CD
0.9
az;
co
=1 Corn Oil
0.8
Estradiol
0.7
QJ 0.6
CD
0.5 ­
0.4 ­
CD
0.3 ­
07
0.2
2
a.
0.0
a)
Incubated
Incubated + LH
In Vitro Treatment
Figure I. Mean (±SE) in vitro concentrations of progesterone (ngmg tissue') in luteal
tissue incubated alone and in the presence of LH from E2-treated and control (corn oil)
ewes.
*Means denoted with an asterisk differed from respective means for control ewes
(p<0.04)
80
Additionally, progesterone production has been shown to be suppressed in regressing
ovine corpora lutea as a result of the ability of prostaglandin F2c, to reduce LH-activated
adenylate cyclase activity and augment phosphodiesterase activity (Aguda et al., 1984).
Consequently, the reduction in cAMP contributes to the diminished capacity of luteal
cells to synthesize progesterone during the regression process. Interestingly,
progesterone synthesized in vitro by day 15 corpora lutea from E2-treated ewes in this
study were only a fraction of the progesterone synthesized in vitro by midluteal ovine
corpora lutea (Slayen and Stormshak, 1990). Although corpora lutea in E2-treated ewes
were larger than those of control ewes, it is obvious that their functional capabilities
were not on par with those of corpora lutea examined during the midluteal phase of the
cycle.
It is well established that the corpus luteum is composed of two different
steriodogenic cell-types: small luteal cells and large luteal cells (Koos and Hansel, 1981).
Both cell types have the capacity to synthesize progesterone, but large luteal cells
synthesize greater quantities of the hormone at both the basal level (Fitz et al., 1982;
Alila et al., 1988; Nelson et al., 1992) and during the luteal phase (Niswender et al.,
1985). Luteal progesterone synthesis has been shown to be dependent on LH in small
luteal cells, but LH-independent in large luteal cells (Fitz et al., 1982; Nelson et al.,
1992). Unexpectedly, there was no increase in progesterone concentration with in vitro
LH treatment of luteal tissue from E2-treated ewes. Although chronic E2-treatment of
ewes can prolong luteal life span it is conceivable that the maintained corpora lutea
consisted of a different ratio of functional small to large luteal cell population than
normally present during the midluteal phase of the cycle. Perhaps the small luteal cells
81
had already begun to regress, and therefore would be unresponsive to in vitro LH
treatment, while large luteal cells remained in a state of basal progesterone
production/secretion. Histological examination of the proportional composition of small
to large luteal cells in maintained corpora lutea of ewes chronically treated with E2 has
not yet been investigated.
In light of the fact that small ovine luteal cells possess LH receptors (Fitz et al.,
1982) and LH receptor mRNA (Smith et al., 1996), while large luteal cells remain largely
unresponsive to LH (Hoyer et al., 1984), it is possible that chronic E2-treatment of ewes
in this study resulted in the down-regulation of LH receptors on the small luteal cells.
This possibility is supported by the data of Kirchick and Birnbaumer (1983) who found
that pharmacological administration of E2 to female rabbits reduced luteal adenylyl cylase
activity and LH receptors. Similarly, human male patients given pharmacological
estrogen treatment have also been reported to experience a reduction in testicular LH
receptors (Huhtaniemi et al., 1980). Hence, the potential down-regulating action of E2­
treatment may have been reflected by the failure of corpora lutea from E2-treated ewes to
respond to in vitro LH with an increase in progesterone synthesis.
In conclusion, chronic in vivo E2 treatment, initiated early in the luteal phase of
the estrous cycle greatly enhanced the production of progesterone in the day 15 ovine
corpus luteum in vitro. However, in vitro exposure of luteal tissue of control and E2­
treated ewes to LH failed to induce additional synthesis of progesterone.
82
BIBLIOGRAPHY
Agudo, LS, WL Zahler and MF Smith. 1984. Effect of prostanglandin Fla on the adenyl
cyclase and phosphodiesterase activity of ovine corpora lutea. J Anim Sci
58:955-962.
Alila, HW, RA Corradino and W Hansel. 1988. A comparison of the effects of
cyclooxygenase prostanoids on progesterone production by small and large
bovine luteal cells. Prostaglandins 36:259-270.
Bertrand, JE and F Stormshak. 1996. In vivo and in vitro responses of the bovine
corpus luteum after exposure to exogenous gonadotropin-releasing hormone and
prostaglandin F2 Endocrine 4:165-173.
Fitz, TA, MH Mayan, HR Sawyer and GD Niswender. 1982. Characterization of two
steroidogenic cell types in the ovine corpus luteum. Biol Reprod 27:703-711.
Hazzard, TM and F Stormshak. 1997. Down-regulation of oxytocin receptors and
secretion of prostaglandin Fla after chronic treatment of ewes with estradiol -17(3.
Biol Reprod 56:1576-1581.
Hoyer, PB, TA Fitz and GD Niswender. 1984. Hormone independent activation of
adenyl cyclase in large steroidogenic ovine luteal cell does not result in increased
progesterone secretion. Endocrinology 114:604-608.
Huhtaniemi, I, P Leinonen, GL Hammond and R Vihko. 1980. Effect of oestrogen
treatment on testicular LI-I/HCG receptors and endogenous steroids in prostatic
cancer patients. Clin Endocrinol (Oxf) 13:561-568.
Karsch, FJ, JF Roche, JW Noveroske, DL Foster, HW Norton and AV Nalbandov.
1971. Prolonged maintenance of the corpus luteum of the ewe by continous
infusion of luteinizing hormone. Biol Reprod 4:129-136.
Kirchick, HJ and L Birnbaumer. 1983. Effects of estradiol treatment on rabbit luteal
adenylyl cyclase: loss of luteinizing hormone receptors and attenuation of the
regulatory N component activity. Endocrinology 113:1629-1637.
Koligian, KB and F Stormshak. 1976. Progesterone synthesis by ovine fetal cotyledons
in vitro. J Anim Sci 42:439-443.
Koos, RD and W Hansel. 1981. The large and samll cells of the bovine corpus luteum:
ultrastructure and functional differences. In: NB Schwartz and M HunzickerDunn, Eds. Dynamics of Ovarian Function. Raven Press Ltd, New York. Pp
197-203.
83
Lauber, ME, MR Waterman and ER Simpson. 1991. Expression of genes encoding
steroidogenic enzymes in the bovine corpus luteum. J Reprod Fertil Suppl
43:57-64.
Nelson, SE, MP McLean, PG Jayatilak and G Gibori. 1992. Isolation, characterization,
and culture of cell subpopulations froming the pregnant rat corpus luteum.
Endocrinology 130:954-966.
Niswender, GD and TM Nett. 1994. The corpus luteum and its control in infraprimate
species. In: E Knobil and JD Neill, Eds. The Physiology of Reproduction. 2nd
Edition. Raven Press Ltd, New York. Pp 781-816.
Niswender, GD, RH Schwall, TA Fitz, CE Farin and HR Sawyer. 1985. Regulation of
luteal function in domestic ruminants: new concepts. Rec Prog Horm Res
41:101-151.
Piper, EL and WC Foote. 1968. Ovulation and corpus luteum maintenance in ewes
treated with 170-oestradiol. J Reprod Fertil 16:253-259.
Piper, EL and WC Foote. 1970. The effect of 17f3-estradiol on corpus luteum function
in sheep. Biol Reprod 2:48-52.
Rodgers, RJ, JD O'Shea and JK Findlay. 1983. Progesterone production in vitro by
small and large ovine luteal cells. J Reprod Fertil 69:113-124.
Rodgers, RJ, MR Waterman and ER Simpson. 1986. Cytochromes P-450sce, P-450s17
adrenoxin, and reduced nicotanamide adenine dinucleotide phosphate­
cytochrome P-450 reductase in bovine follicles and corpora lutea.
Endocrinology 118:1366-1374.
Rodgers, RJ, MR Waterman and ER Simpson. 1987. Levels of messenger ribonucleic
acid encoding cholesterol side-chain cleavage cytochrome P450, 17a­
hydroxylase cytochrome P450, adrenodoxin, and low density lipoprotein receptor
in bovine follicles and corpora lutea throughout the ovarian cycle. Mol
Endocrinol 1:274-279.
Slayden, OV and F Stormshak. 1990. In vivo and in vitro effects of a cyclopropenoid
fatty acid on ovine corpus luteum function. Endocrinology 127:3166-3171.
Smith, GW, PC Gentry, RM Roberts and MF Smith. 1996. Ontogeny and regulation of
luteinizing hormone receptor messenger ribonucleic acid within the ovine corpus
luteum. Biol Reprod 54:76-83.