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Influence of insulin-like growth factor-1, steroids, and nitrate on reproduction in amphibians

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Influence of insulin-like growth factor-1, steroids, and nitrate on reproduction in amphibians
Creator:
Barbeau, Tamatha R., 1970
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English
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ix, 169 leaves : ill. ; 29 cm.

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Subjects / Keywords:
Amphibians ( jstor )
Frogs ( jstor )
Hormones ( jstor )
Human growth ( jstor )
Nitrates ( jstor )
Nitrites ( jstor )
Ovaries ( jstor )
Oviducts ( jstor )
Plasmas ( jstor )
Steroids ( jstor )
Dissertations, Academic -- Zoology -- UF ( lcsh )
Frogs -- Reproduction -- Endocrine aspects ( lcsh )
Zoology thesis, Ph. D ( lcsh )
The Everglades ( local )
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bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

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Thesis:
Thesis (Ph. D. )--University of Florida, 2004.
Bibliography:
Includes bibliographical references.
General Note:
Printout.
General Note:
Vita.
Statement of Responsibility:
by Tamatha R. Barbeau.

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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
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INFLUENCE OF INSULIN-LIKE GROWTH FACTOR-i, STEROIDS, AND NITRATE ON
REPRODUCTION IN AMPHIBIANS

















By

TAMATHA R. BARBEAU


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2004































Copyright 2004

By

Tamatha R. Barbeau















ACKNOWLEDGMENTS

I thank my advisor, Louis J. Guillette, whose encouragement, guidance, and motivation

were invaluable during this work. His expertise and enthusiasm for research provided the

foundation for this work, and his passion for teaching these skills to others has molded my own

desire to pursue an academic career. I owe much gratitude to my committee members (William

Buhi, Lauren Chapman, David Evans, and Harvey Lillywhite) for their advice and insightful

conversations throughout this study. As physiologists David and William made indelible

impressions on me and provided invaluable insights and ideas for my research. Lou, Harvey, and

Lauren have been my committee members, mentors, and friends during both my M.S. and Ph.D.

degrees. Collectively, they have made the greatest contributions to my professional development,

academic philosophies, research skills, scientific curiosity, and perspectives on life.

This research has been supported by grants from SIGMA XI Grants in Aid of Research,

The Brian Riewald Memorial Fund (UF), and Declining Amphibians Population Task Force. All

frogs were used in compliance with and supervision of the Institutional Animal Care and Use

Committee at the University of Florida (IACUC #Z023 and #Z095).

For their generous laboratory support, training, and camaraderie, I thank my friends and

colleagues (Dieldrich Bermudez, Teresa Bryan, Thea Edwards, Mark Gunderson, Iske Larkin,

Matthew Milnes, and Brandon Moore). I extend additional thanks to Colin Chapman, Ginger

Clark, and Douglas Levey for their help in providing me with laboratory techniques and space to

conduct my research. Many friends and colleagues at the University of Florida provided valuable

insights and assistance with various aspects of my work (namely Keith Choe, Martin Cohn,

Franklin Percival, and Kent Vliet). I am especially grateful for the help of Loretta Azzinario,

Jason Bridge, Arika Brown, Tim Buhi, Leo Choe, Brandy Cunningham, Lauren Farrar, David

i i1








Iglesias, Kapila Karakota, Caroline Keicher, Dana LaKam, Axel Lucca, Courtney Marler, Amy

McGreane, Pamela Moses, Amanda Mulligan, Reshma Patel, Sonia Parikh, Wilhelmina Randtke,

Maria Samuel, Catherine Vallance, and Kyu Mee Yo.

My husband, Greg Pryor, continues to be the most important and supportive person in my

academic development and in life. His encouragement, guidance, and sense of humor have been

the one constant, throughout the triumphs and tribulations of my graduate work. I look forward to

sharing many more adventures with him in the future. I am grateful to my mother, for being

patient when I brought snakes and frogs home as a child; and to my father, for taking me on

camping trips in the Northern Adirondacks. These events inspired my appreciation and curiosity

for ecosystems and animals of all kinds.















TABLE OF CONTENTS


page

ACKNOWLEDGMENTS................................................................................ ii

ABSTRACT ................................................................................................................................. viii

CHAPTER

I INTRODUCTION: INFLUENCE OF STEROIDS, INSULIN-LIKE GROWTH
FACTOR-1, AND NITRATE ON REPRODUCTION IN AMPHIBIANS ................................ 1

Reproductive Steroids and Amphibian Reproduction ................................................................. 1
Insulin-Like Growth Factor- I ................................................................................................ 3
Aquatic N itrate and Amphibian Reproduction ........................................................................ 7
Research Objectives .................................................................................................................... 9
Physiology and Evolution ................................................................................................. 11
Conservation ...................................................................................................................... 12

2 THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS OF NITRATE (IN VIVO) ON PLASMA STEROIDS AND
INSULIN-LIKE GROWTH FACTOR-I, ON OVARIAN STEROID SYNTHESIS,
AND ON OVIDUCT GROWTH IN THE AFRICAN CLAWED FROG (Xenopus
laevis) ........................................................................................................................................ 15

Introduction ............................................................................................................................... 15
M aterials and M ethods .............................................................................................................. 18
Animals and Samples ................................................................................................... 18
Nitrate Study Design ..................................................................................................... 18
Steroid Radioimmunoassay (RIA) Procedures ............................................................. 20
Insulin-Like Growth Factor-I (IGF-1) RIA Procedures .............................................. 21
Biochemical RIA Validations ..................................................................................... 21
Statistics ............................................................................................................................ 22
Results ....................................................................................................................................... 22
Tissue W eights .................................................................................................................. 22
Follicle Diameters ....................................................................................................... 22
Plasma Steroid Concentrations ..................................................................................... 23
Plasma IGF- 1 Concentrations ..................................................................................... 23
Ovarian Follicle Steroid Concentrations (Ex Vivo) ..................................................... 23
Discussion ................................................................................................................................. 24









3 SEASONAL CHANGES IN INSULIN-LIKE GROWTH FACTOR-1, STEROIDS,
AND REPRODUCTIVE TISSUES IN PIG FROGS (Rana grylio) ................................... 36

Introduction ............................................................................................................................... 36
M aterials and M ethods .............................................................................................................. 39
Water Parameters, Animal Captures, and Sample Collections ................................... 39
Steroid Radioimmunoassay (RIA) Biochemical Validation ........................................ 42
Steroid RIA Procedures .............................................................................................. 43
Insulin-like Growth Factor-I (IGF-1) RIA Biochemical Validation ............................ 44
IGF-1 RIA Procedures ................................................................................................ 45
Statistics ............................................................................................................................ 46
Results ....................................................................................................................................... 47
Seasonal Environmental Param eters ............................................................................ 47
Seasonal Tissue M ass and Ovarian M aturation ......................................................... 47
Seasonal Plasm a Steroid and IGF- I Concentrations ................................................... 48
Correlations: Plasma Steroids, Tissue Mass, and Environmental Parameters ............. 49
Discussion ................................................................................................................................. 50

4 THE EFFECTS OF INSULIN-LIKE GROWTH FACTOR-i AND ESTRADIOL
IMPLANTS (IN VIVO) ON OVIDUCT MORPHOLOGY, AND ON PLASMA
HORMONES IN BULLFROGS (Rana catesbeiana) .......................................................... 70

Introduction ............................................................................................................................... 70
M aterials and M ethods .............................................................................................................. 72
Ovariectom y ...................................................................................................................... 73
Horm one Im plants ....................................................................................................... 74
Tissue Sampling ................................................................................................................ 75
Steroid Radioimmunoassay (RIA) Biochemical Validation ....................................... 76
Steroid RIA Procedures ................................................................................................. 77
Insulin-Like Growth Factor-i (IGF-1) RIA Biochemical Validation .......................... 78
IGF-I RIA Procedures ................................................................................................ 78
Statistics ............................................................................................................................ 79
Results ....................................................................................................................................... 80
Biochem ical RIA Validations ..................................................................................... 80
Tissue W eights .................................................................................................................. 80
Oviduct M orphom etrics .............................................................................................. 80
Plasm a Steroid and IGF- 1 Concentrations ................................................................... 81
D iscussion ................................................................................................................................. 82

5 OVARIAN STEROIDOGENESIS (IN VITRO) IN PIG FROGS (Rana grylio) AFTER
EXPOSURE TO ENVIRONMENTALLY RELEVANT CONCENTRATIONS OF
NITRA TE AND NITRITE ...................................................................................................... 102

Introduction ............................................................................................................................. 102
M aterials and M ethods ............................................................................................................ 106
Collection of Anim als ..................................................................................................... 106
Ovarian Follicle Culture (In Vitro) .................................................................................. 106
Steroid Radioimmunoassay (RIA) Procedures and Validations ..................................... 107
Statistics .......................................................................................................................... 108
Results ..................................................................................................................................... 108
Discussion ............................................................................................................................... 109









6 THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS ON NITRATE ON OVIDUCTAL MORPHOLOGY
MORPHOLOGY AND PLASMA STEROIDS AND INSULIN-LIKE GROWTH
FACTOR-1 IN BULLFROGS (Rana catesbeiana) ................................................................ 118

Introduction ............................................................................................................................. 118
M aterials and M ethods ............................................................................................................ 121
A n im als ........................................................................................................................... 12 1
Nitrate Treatments ........................................................................................................... 122
Steroid Radioimmunoassay (RIA) Procedures and Validations ..................................... 124
Insulin-Like Growth Factor-i (IGF-1) RIA Procedures and Validations ....................... 125
Statistics .......................................................................................................................... 127
Results ..................................................................................................................................... 128
Oviduct W eights .............................................................................................................. 128
Plasma Steroid and IGF-1 Concentrations ...................................................................... 128
Oviduct M orphometrics .................................................................................................. 128
Discussion ............................................................................................................................... 129

7 CONCLUSIONS ..................................................................................................................... 141

Seasonal Plasma Steroids and IGF- 1, and Reproductive Tissue Growth ................................ 141
The Effects of IGF- 1, E2, and Nitrate on Oviduct Growth ...................................................... 143
Nitrate Exposure (In Vivo and In Vitro): Effects on Steroidogenesis ...................................... 147
Nitrate Exposure and Plasma IGF-1 ........................................................................................ 150

RE FERE NCES ............................................................................................................................. 152

BIOGRAPHICAL SKETCH ........................................................................................................ 169















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

INFLUENCE OF INSULIN-LIKE GROWTH FACTOR-1, STEROIDS, AND
NITRATE ON REPRODUCTION IN AMPHIBIANS

By

Tamatha R. Barbeau

August 2003

Chair: Louis J. Guillette
Major Department: Zoology

My goal was to examine the influence of insulin-like growth factor-i (IGF- 1), 17-03

estradiol (E2), testosterone (T), and nitrate exposure on various aspects of reproduction in frogs.

To accomplish this, I investigated seasonal changes in plasma IGF-1, E2, and T concentrations in

a wild population of Rana grylio. I also determined the importance of steroid and growth factor

hormones in reproductive physiology by examining ovariectomized Rana catesbeiana for

changes in plasma IGF-1, E2, and T concentrations, and changes in oviduct morphology after

treatment with known doses of IGF-1, E2, and epidermal growth factor (EGF). Finally, I

examined three aquatic frogs species (Xenopus laevis, R. grylio, and R catesbeiana) for the

effects of nitrate exposure on changes in plasma IGF- 1, E2, and T concentrations, and on oviduct

morphology.

I have demonstrated that plasma IGF- 1, E2, and T concentrations (and reproductive tissue

growth) exhibit a clear seasonal pattern of changes that overlap with changes in environmental

variables, such that reproductive condition is optimized to match favorable environmental

temperatures. I also demonstrated that E2 is a potent stimulator of oviduct growth, while EGF and








IGF- 1 do not induce oviductal growth in R. catesbeiana. I also provide the first evidence that

exposure to environmentally relevant concentrations of nitrate alters endocrine hormones in

Xenopus laevis, R. grylio, and R. catesbeiana. Furthermore, IGF-1 and steroid hormone

concentrations are altered with exposure to nitrate at concentrations deemed safe for human

drinking water by the US EPA (10 mg/L). In vivo exposure of X laevis (for 7 continuous days) to

nitrate concentrations below 50 mg/L significantly increased plasma IGF- I concentrations, and

inhibited ovarian E2 and T synthesis. In vitro incubation of ovarian tissue (from wild-caught R.

grylio) with nitrate concentrations between 0.17 and 33.00 mg/L nitrate (and between 0.20 and

40.60 mg/L nitrite) inhibited E2 and T synthesis after 3 hours of exposure. Lastly, in vivo

exposure of R. catesbeiana to nitrate concentrations between 1.65 and 16.50 mg/L increased

plasma IGF-1, E2, and T concentrations; and caused oviductal atrophy. These findings

demonstrate that exposure to nitrate at extremely low concentrations causes endocrine disruption

in frogs.














CHAPTER 1
INTRODUCTION: INFLUENCE OF STEROIDS, INSULIN-LIKE GROWTH
FACTOR-1, AND AQUATIC NITRATE ON REPRODUCTION IN AMPHIBIANS

Reproductive Steroids and Amphibian Reproduction

Amphibians display some of the most diverse reproductive modes compared to other

vertebrates. Most amphibians exhibit the ancestral reproductive mode, and are restricted to water

to oviposit and fertilize eggs externally. Some species are terrestrial breeders, have internal

fertilization, and either oviposit eggs on land or retain them within the oviducts for all or part of

the embryonic developmental period. Finally, some amphibians oviposit terrestrial eggs from

which offspring hatch, bypass the free-living tadpole stage, and undergo direct development to

emerge as fully developed froglets (Wake and Dickie, 1998).

The reproductive system in amphibians is characterized by cyclic changes in growth and

function that are modulated by hypothalamic releasing hormones, pituitary gonadotropins, and

gonadal steroids. The process of steroid synthesis or steroidogenesis is regulated primarily by the

hypothalamic-pituitary-gonadal axis (Licht, 1970, 1979; Licht et al., 1983). Gonadotropin-

releasing hormone (GnRH) is secreted by the hypothalamus (in response to internal or

environmental cues) and stimulates the anterior pituitary to release luteinizing hormone (LH) and

follicle stimulating hormone (FSH) into the bloodstream. These gonadotropins stimulate gonadal

steroidogenesis and gametogenesis. The principal gonadal steroids are progesterone (P4), estradiol

1703 (E2), and testosterone (T). Theca interna cells within the ovary synthesize T in response to

LH stimulation. In response to FSH stimulation, ovarian granulosa cells synthesize aromatase,






















Stimulation




C


Ovarian Cell


E


Inhibition




D


Theca Interna


Figure 1-1. Regulation of gonadal steroidogenesis. (A) Hypothalamic gonadotropin releasing
hormone (GnRH) induces pituitary secretion of luteinizing hormone (LH) and follicle
stimulating hormone (FSH) (B). LH stimulates theca interna cells (C) to synthesize
testosterone (T). FSH stimulates granulosa cell to produce aromatase enzymes (D),
which convert T into estrogen (E2). Steroidogenesis induces inhibin release from the
gonad, which inhibits further hypothalamic and pituitary stimulation (E). F.
Circulating steroids are also metabolized and cleared from the bloodstream by the
liver.

an enzyme that converts T into E2 (Figure 1-1). These steroids induce gonadal release of inhibin,

a hormone that inhibits hypothalamic-pituitary stimulation of further steroid synthesis. Gonadal

steroids exert an autocrine or paracrine action, by influencing localized tissues; or function as

endocrine hormones when released into the bloodstream, to affect distant target tissues. Within

steroid-responsive tissues, T and E2 bind to and activate cytosolic or nuclear receptors and form a

steroid-receptor complex. This complex binds to a hormone-response element on DNA to









stimulate or inhibit transcription, protein synthesis, and tissue growth (Segars and Driggers,

2002). Through this process, E2 and T regulate normal development of secondary sexual

characteristics, regulate growth of steroid-responsive tissues, and regulate reproductive function

(Guidice, 1999).

In amphibians, E2 is essential for oocyte development and maturation within ovarian

follicles (Dumont, 1971; Fortune, 1983). Gonadal E2 and T regulate many aspects of reproductive

function, such as oviduct growth and secretions (Licht et al., 1983; Norris, 1997). The oviduct is a

vital structure for reproductive function in oviparous vertebrates, including amphibians (Giudice,

1992; Wake and Dickie, 1998). After ovulation from the ovaries, mature oocytes travel through

the oviduct to the cloaca and are expelled into the environment. In addition to providing physical

transport, the oviduct synthesizes and secretes proteins and other substances that nourish and

encapsulate the ova, and also aid in fertilization (Low et al., 1976; Buhi et al., 1997).

Insulin-Like Growth Factor-1

It has become increasingly apparent that reproductive function and physiology are

regulated by steroid-signaling pathways, and also by other pathways involving insulin-like

growth factor- I (IGF- 1). Originally called somatomedin C, IGF- 1 is a polypeptide hormone that

is structurally similar to IGF-II and proinsulin, and likely originated early in vertebrate evolution.

IGF-1 is part of the growth factor system, which consists of a family of proteins that function in

regulating many cellular processes (including cell proliferation, differentiation, and apoptosis) in

virtually all tissues. (LeRoith et al., 2001a,b). Thus, IGF-1 is important for normal growth and

function of reproductive tissues, and also for somatic tissues. Accordingly, the role of IGF- 1 in

growth of reproductive and somatic tissues has been examined in a variety of vertebrates

including mammals, fish, birds, and reptiles (Girbau et al., 1987; Murphy and Ghahary, 1990; De

Pablo et al., 1990; Serrano et al., 1990; Simmen et al., 1990; Scavo, 1991; Kapur et al., 1992; Cox

and Guillette, 1993; Tang et al., 1994; Guillette et al., 1996; Buhi et al., 2000; Qu et al., 2000;

Allan et al., 2001). Although IGF-1 has been identified in the plasma and tissues of some









amphibians, the role of this peptide hormone in tissue growth and function in these animals

remains unclear, and requires further study (Daughaday et al., 1985; Pancak-Roessler and Lee,

1990).

Traditionally, IGF- 1 was thought to influence tissue growth primarily by mediating the

effects of growth hormone (GH). This physiological function of IGF-1 is the basis of the original

somatomedin hypothesis (LeRoith et al., 2001 b). More recently, IGF-l has been found to play an

important role in growth and differentiation of reproductive tissues (independent of GH), by

mediating the mitogenic effects of E2 (Girbau et al., 1987; Murphy and Ghahary, 1990; Cox,

1994).

In the presence of E2, IGF- I has been shown to mediate growth of E2-sensitive

reproductive tissues like the oviduct (Mead et al., 1981; Murphy and Murphy, 1994). Research

indicates that the growth effects of IGF- 1 does not require E2 but requires only the presence of the

1713 estradiol alpha receptor (ERax) in reproductive tissues (Klotz et al., 2000). This is supported

by findings of an E2-like growth response in the oviduct of ovariectomized animals treated with

IGF-1 (Cox, 1994). These findings demonstrate that IGF-1 potentiates E2-induced growth and

also stimulates E2-independent tissue growth.

In addition to mediating the growth effects of reproductive steroids, IGF- 1 has also been

shown to regulate intraovarian steroid synthesis in mammals (Adashi et al., 1991; Guidice, 1992;

Adashi, 1993). Decreased E2 expression increases ovarian IGF- 1 expression. Ovarian IGF- I

stimulates synthesis of E2 and P4, and increases aromatization of androgens into E2 (Adashi et al.,

1991). Additionally, the ovaries and oviduct synthesize and secrete IGF-1 in response to GH,

FSH, E2, and other hormones. Based on these findings, the list of factors that regulate (or are

influenced by) the IGF- 1 system has been expanded to include reproductive steroids.

Research spanning nearly 50 years has defined many components of the surprisingly

complex IGF- 1 system (Le Roith et al., 2001 b). In all vertebrates examined, the liver synthesizes

and secretes most of the circulating concentrations of IGF- 1. Hypothalamic release of growth








hormone-releasing hormone (GHRH) stimulates the pituitary to secrete growth hormone (GH)

into the bloodstream. In response to GH stimulation, the liver synthesizes and secretes IGF-1 into

the bloodstream. Hepatic IGF-1 can affect peripheral tissues in a paracrine or autocrine manner;

or it can be transported through the bloodstream, bound to IGF binding proteins (IGFBPs), as an

endocrine hormone that mediates growth and apoptosis of distant target tissues. After reaching its

target tissue, IGF-1 interacts with a transmembrane cell-surface IGF-1 receptor (IGF- IR) where it

is released from its binding protein to initiate a cellular response. Excess circulating IGF-1 is then

filtered and degraded by the kidneys (LeRoith et al., 2003). In this manner, IGF-1 mediates GH-

induced cellular proliferation. This endocrine-signaling pathway is the basis of the original

somatomedin hypothesis. However, recent research suggests that the somatomedin hypothesis

should be revised. IGF- 1 has been shown to have many GH-independent effects on regulating

tissue growth. Additionally, non-hepatic tissues (including the ovaries and oviduct) are now

known to synthesize and secrete IGF-1 (LeRoith et al., 2001 b).

The extracellular functional components of the IGF- 1 system include IGF-1, IGFBPs,

and IGF- 1R. Expression of IGF- 1 can be stimulated by various factors including growth

hormone, E2, T, P4, FSH, glucose, insulin, and thyrotropin; whereas, IGF-1 expression can be

inhibited by somatostatin, LH, cortisol, and interferon. Six known IGFBPs can bind with IGF- I

to modulate cellular effects. The IGFBPs that regulate the cellular effects of IGF- 1 include

IGFBPs 1, 3, 4, and 5. The other binding proteins (IGFBPs 2 and 6) specifically regulate the

effects of IGF-2 on embryonic development. The IGFBPs prevent IGF-1 degradation during

circulation, transport IGF- 1 to target tissues, and regulate binding of IGF- 1 to IGF- 1 R. Like IGF-

1, IGFBPs can be stimulated or inhibited by various factors. Another functional component of the

IGF-1 system is the IGF-IR. The IGF-1R is a tyrosine kinase, transmembrane receptor found on

virtually every tissue type, and it mediates a majority of IGF-1 actions on cell growth. There is an

IGF-2 receptor, but it is highly specific for IGF-2 and functions mostly in mediating embryonic

development. Expression of the IGF- 1 R can be stimulated by a variety of factors including E2,









A IGF-I


Figure 1-2. Binding of IGF-1 with the IGF- IR, initiates phosphorylation of intracellular proteins
in a signaling cascade that leads to a cellular response. Activation of adaptor proteins
includes the mitogen activated protein kinase (MAPK) pathway, the
phosphatidylinositol 3-kinase (IP-3K) pathway and its secondary messengers IP3,
DAG, and Ca'. Briefly, binding of IGF-1 (A) to the receptor (B) results in
autophosphorylation of the intracellular 13-subunit of the receptor (C). This then
activates intracellular adaptor proteins, insulin receptor substrate (IRS) and Shc, to
bind with the receptor and become phosphorylated. If adaptor protein Shc is
activated, it forms a complex with SOS to activate Raf. Activation of Raf
phosphorylates protein kinase MEK and leads to phosphorylation (D) of mitogen-
activated protein kinase (MAPK). This activates transcription factors (TF) that bind
to nuclear DNA (F) to elicit a cellular response (G). If the adaptor protein IRS is
activated (H), a sequence of phosphorylations involving protein subunits p85 and
p1 10 will activate the IP-3K pathway (I). Activation of IP-3K pathway
phosphorylates the conversion of phosphoinositol bisphosphate (PIP2) to the second
messengers (J) inositol trisphosphate (IP3) and (K) diacyglycerol (DAG). Each of
these second messengers can induce cellular responses (L) either by activating the
Ca*/calmodulin complex by IP3 or by the activation of phosphokinase C (PKC) by
DAG.

FSH, LH, and oncogenes. Conversely, IGF-1, P4, and tumor suppressors can inhibit expression of

the IGF- 1 R.

The functional IGF- 1 system also has intracellular functional components that become

activated by binding of IGF- 1 to the IGF- 1 R. Once bound, the IGF- 1 R becomes phosphorylated,

and a variety of intracellular proteins and second messengers are involved in a signaling cascade








that leads to a cellular response (Fig 1-2). Thus, each of the functional components of the IGF-I

system can be regulated by complex extracellular and intracellular factors.

Aquatic Nitrate and Amphibian Reproduction

In the past few years, there has been increased global concern over contamination of

water by anthropogenic sources of nitrate. Nitrate is an anionic form of nitrogen that infiltrates

watersheds in agricultural and urban environments, and reaches harmful concentrations largely

due to human activities. In agricultural areas, watersheds are polluted with nitrate from

unregulated run-off of nitrogen-based fertilizers and run-off of animal wastes. In urban areas,

nitrates contaminate watersheds primarily through runoff of industrial and wastewater effluent

from treatment plants and of fertilizers applied to lawns and golf courses (Rouse et al., 1999). The

application of fertilizers in close proximity to watersheds during the spring frequently results in

an overwhelming nitrate "pulse" that overlaps the breeding season of many amphibians.

Unfortunately, most studies of the effects of nitrate on amphibians report nitrate concentrations

differently, making comparisons and interpretation of these studies extremely difficult. For

consistency throughout this dissertation, nitrate is reported as equivalent to nitrate-as-nitrogen

(N03-N). This represents the concentration of nitrogen present in a given concentration of nitrate.

Additionally, equivalent measures of nitrate are provided here to facilitate comparison among

other nitrate and nitrite studies (Table 1-1).

Most studies on the effects of nitrate on amphibians have addressed toxicological rather

than sublethal concentrations (Rouse et al., 1999). Most of these studies also focused on juvenile

amphibian stages rather than on adults (Table 1-2). The impact of nitrate exposure on mammalian

steroidogenesis has been examined and described in a few studies. Nitrate exposure has been

shown to inhibit androgen synthesis in rodents in vivo and also in Mouse Leydig tumor cells in

vitro (Panesar, 1999; Panesar and Chan, 2000). One mechanism for altered steroid expression (in

vivo) by nitrates involves enzyme-dependent synthesis of nitric oxide (NO) (Panesar and Chan,

2000). The NO is synthesized from an L-arginine precursor by nitric oxide synthase (NOS)









enzymes (Kleinert et al., 1995; Mayer and Hemmenns, 1997). In addition to NOS-dependent NO

formation, non-enzymatic synthesis of NO can also occur through acidic reduction of nitrite

(izuka et al., 1999; Zweier et al., 1995, 1999; Modin et al., 2001). Cosby et al. (2003) reported

that hemoglobin functions as a nitrite reductase contributing to enzyme-independent NO

synthesis. Furthermore, Zweier et al., (1999) reported that enzyme-independent NO formation is

associated with cellular damage and loss of organ function. Regardless of the mechanisms by

which it is produced, NO is thought to regulate many physiological processes. Within the gonad,

NO can inhibit steroidogenesis by binding to the heme (iron-containing) groups located on the

enzymes of the cytochrome P450 superfamily necessary for steroid synthesis, like 3p-

dehydroxysteroid dehydrogenase (303-HSD). (Van Voorhis et al., 1994; Panesar and Chan, 2000).

The IGF- I counteracts the effects of NO by increasing ovarian E2 synthesis (Van Voorhis et al.,

1994; Van Voorhis et al., 1995; Srivastava et al., 1998; Inigues et al., 2001; Les Dees et al.,

2001). Within the mitochondria of cells, the enzymes P450,,, and 313-HSD convert free

cholesterol into P4 (the precursor for T and E2). Steroid enzyme pathways disrupted by NO can

inhibit P4 and downstream androgen synthesis (Panesar and Chan, 2000). If P4 and T synthesis are

inhibited by nitrate, then less androgen is available for aromatase enzymes to synthesize into E2,

and estrogen concentrations would be altered. Despite findings of endocrine disruption by nitrate

in mammals, no study has examined whether nitrate disrupts endocrine function in adult,

reproductive amphibians.

Since E2 and IGF- 1 interact to regulate growth-related responses in reproductive tissues,

it is plausible that alteration of E2 expression by nitrates might also influence IGF- 1 and oviduct

growth (perhaps through a NO-dependent pathway). In humans, intraovarian IGF- 1 expression

increases in response to elevated intraovarian NO. The IGF- 1 apparently counteracts the

inhibitory effects of NO on steroids, by stimulating increased expression of aromatase enzymes,

StAR protein, P4, and E2. Thus, increased intraovarian IGF-1 in response to NO might be a









compensatory response, functioning to amplify steroid synthesis that has been compromised

(Schams et al., 1988; Erickson et al., 1989; Adashi, 1993; Samaras et al., 1996; Iniguez et al.,

2001; Les Dees et al., 2001). These studies indicate that IGF- 1 plays a vital role in steroid

synthesis and regulation, and possibly functions through an NO-dependent pathway. The

mechanism by which steroids and IGF- I interact to stimulate growth of reproductive tissue

remains enigmatic and requires further study. In addition, the influence of nitrate exposure on

reproductive physiology of anurans remains unknown.

Organic nitrate and nitrite are normally present in aquatic habitats, in low concentrations,

due to bacterial breakdown of organic matter and accumulation of biological wastes. In addition

to contributions from natural sources, anthropogenic sources of nitrate and nitrite can

compromise water quality even further. Unusually high concentrations of nitrate and nitrite can

accumulate in aquatic habitats that receive runoff of agricultural fertilizers and animal wastes.

Aquatic nitrate and nitrite contamination might provide a biological signal to frogs that water

quality is unsuitable for reproduction. High nitrate and nitrite concentrations might repress

physiological changes that stimulate reproductive condition of frogs. Contamination of aquatic

habitats with nitrate and nitrite has been shown to be detrimental to survival of anuran eggs and

tadpoles (Table 1-2), and amphibian populations are reportedly declining in some agricultural

areas (Berger, 1989).

Research Objectives

One goal of my study was to gain a better understanding of the interaction of IGF- 1 with

E2 -dependent and independent growth of reproductive tissues in aquatic amphibians. Although

IGF- I is important for cell growth and differentiation, abnormally high concentrations of plasma

IGF- 1 are associated with abnormal growth of reproductive tissues; and with cancer of the breast,

ovaries, uterus, endometrium, and prostate (LeRoith et al., 1995a,b; Grimberg and Cohen, 1999;

van Dessel et al., 1999; Werner and Le Roith, 2000; Smith et al., 2000). The IGF-1 and IGF-1R

can protect cells from apoptosis; but in some mammals, over-expression of these receptors









induces ligand-dependent tumor formation. Over-expression of IGF-1 R can be induced by up-

regulation of IGF-1 expression in response to growth hormone (GH)-, E2-, and ERa-dependent

pathways (Kaleko et al., 1990). Additionally, uterine IGF- 1 and IGF- I R up-regulation (along

with increased uterine epithelial cell growth) occurs in ovariectomized rodents in response to

synthetic estrogens (DES and bisphenol A) and phytoestrogens (Klotz et al., 2000). From these

findings, I hypothesized that endocrine disrupting contaminants (EDCs) could affect the IGF-1

system. In a variety of vertebrates EDCs have been shown to alter reproduction. Much research

has focused on the interaction of EDCs with steroid hormones and their receptors (Rooney and

Guillette, 2000). Unfortunately, the effect of EDCs on the IGF- 1 system has received surprisingly

little scientific scrutiny (Backlin and Bergman. 1995; Backlin, et al., 1998). Thus, another goal of

my study was to determine whether nitrate and nitrite (known to induce developmental

abnormalities in amphibians and reproductive abnormalities in other vertebrates) can alter

concentrations of IGF- 1 and steroid hormones and alter growth of reproductive tissues in

amphibians.

The effect of nitrate on synthesis of IGF- 1 and steroids remains an important topic for

investigation. Growing evidence indicates that nitrate exposure stimulates NO synthesis in body

tissues. Furthermore, increased NO expression in gonadal tissues affects steroid and IGF- I

expression. Thus, nitrate exposure might influence IGF- 1 synthesis, similar to steroids, through

an NO-dependent or independent pathway.

Finally, my study examined adult anurans for seasonal changes in IGF- 1 and steroid

concentrations, and in reproductive tissues. Seasonal patterns of change in plasma IGF- 1 and

steroid hormone concentrations, and in growth of reproductive tissues, are reported for alligators

and turtles (Crain et al., 1995; Guillette et al., 1996). In anurans, seasonal changes in plasma IGF-

1 have been reported for the Woodhouse toad, Bufo woodhousei (Pancak-Roessler and Lee,

1990). Thus, I expected that plasma IGF- 1 and steroid concentrations, and growth of reproductive








tissues, would exhibit a seasonal pattern of change in response to endogenous stimulation and

environmental cues.

In addition to addressing the goals mentioned above, findings from my study also have

more general applications for studies of amphibian physiology, evolution, and conservation.

Physiology and Evolution

Physiological regulation of the IGF-1 system has been examined in mammals and

reptiles. The IGF- 1 has been shown to regulate gonadal steroid synthesis, to stimulate oviductal

growth, and to exhibit seasonal cyclicity in mammals and reptiles. Recent research on reptiles and

mammals demonstrates that IGF- 1 potentiates E2-induced growth of reproductive tissues like the

oviduct. Even in the absence of endogenous E2, IGF- 1 stimulates significant oviduct growth.

Thus, the role of growth factors in reptilian and mammalian reproduction is more important than

previously recognized. Additionally, seasonal cycles of increased plasma steroid concentrations

and increased reproductive tissue growth overlap with increases in plasma IGF- I in reptiles and

mammals (Crain et al., 1995; Guillette et al., 1996; Webster et al., 2001). These findings indicate

that IGF- 1 is associated with reproductive activity, and is responsive to changes in reproductive

parameters and environmental cues.

In amphibians, the presence of IGF- I has been documented; but the physiological

processes that regulate this system remain largely under-investigated. If the amphibian oviduct

responds to IGF (similar to mammals and reptiles), then IGF- I regulation of reproductive tissues

represents an early evolutionary phenomenon. However, if the amphibian oviduct is unresponsive

to IGF-1 stimulation, then IGF-induced oviduct growth might represent a relatively recent

development in reptiles and mammals. Seasonal changes in plasma IGF- 1 concentration have

been described for B. woodhousei, but it remains unknown if changes in IGF- I parallel

reproductive parameters in this or other amphibian species (Pankcak-Roessler and Lee, 1990).

My study provides the first description of how endogenous steroids, environmental factors, and

reproductive cyclicity influence the IGF-1 system in amphibians.









Conservation

Amphibian populations in some agricultural areas are declining, and frogs have been

found with dramatic deformities. The factors responsible for these declines and deformities are

hard to identify, but might include runoff of nitrogenous fertilizers from agricultural land into

watersheds where amphibians live and reproduce. Mammals drinking nitrate- and nitrite-

contaminated water exhibit decreased gonadal steroid synthesis after only relatively brief

exposure periods. Despite findings of abnormal growth and metamorphosis in tadpoles exposed

to nitrate, no study has investigated whether nitrate alters endocrine function in juvenile or adult

amphibians. Furthermore, most studies focus on the effects of lethal rather than sublethal

concentration of nitrate on amphibians. My study examined the effects of sublethal

concentrations of nitrate and nitrite on plasma steroids, gonadal steroid synthesis, and growth of

reproductive tissues in amphibians. If nitrate or nitrite exposure alters endocrine function,

specifically reproductive steroids, then these contaminants should be considered as an important

factor to consider in amphibian reproduction and population declines.










Table 1-1. The molecular formula weight (MFW) of sodium nitrate (NaNO3) and sodium nitrite (NaNO2), the MFW percent of sodium (Na),
nitrate (NO3), nitrite (NO2), and nitrogen (N), and the equivalent concentrations of nitrate, nitrite, nitrate as nitrogen (N03-N), and
nitrite as nitrogen (N02-N) as milligrams per liter (mg/L) and millimolar (mM) of solution.
Nitrate as Nitrate as
Percent Percent Percent Nitrate Nitrate
sodium nitrate nitrogen NaNO3 N03 nitrogen NO3 nitrogen
(Na) (N0) (N) (mg) NO3) (N03-N) (mM) (N03-N)
(mg/L) (mg/L) (mM)
0 0.00 0.00 0.00 0.00
Sodium 1 0.73 0.17 0.009 0.002
nitrate 10 7.30 1.65 0.01 0.003
27.0 % 73.0 % 16.5 % 40 29.20 6.60 0.34 0.08
(NaNO3) 100 73.00 16.50 0.86 0.19
FW 84.99 150 109.50 24.75 1.29 0.29
g 200 146.00 33.00 1.72 0.39
300 219.00 49.50 2.58 0.58
Nitrite as Nitrite as
Percent Percent Percent Nitrite Nitroge Nitrite Nitrogen
sodium nitrite nitrogen NaNO2 NO2 nitrogen NO2 nitrogen
(Na) (NO) (N) (mg/L) (mg/L) (mM) (mM)

0 0.00 0.00 0.00 0.00
Sodium 1 0.67 0.20 0.01 0.003
nitrite 10 6.67 2.03 0.10 0.03
33.3 % 66.7 % 20.3 % 40 26.68 8.12 2.90 0.12
(NaNO2) 100 66.70 20.30 0.39 0.29
FW68.99g 150 100.05 30.45 0.97 0.44
200 133.40 40.60 1.45 0.59
300 200.10 60.90 2.90 0.88











Table 1-2. A comparison among amphibians of the effects of nitrate and nitrite
Species Stage Treatment End Point
Ambystoma gracile Larvae 0.78-25 mg/L nitrate Decreased feeding & activity, bent tails, edema


Bufo americanus

B. boreas

B. bufo
Hyla regilla

Litoria caerulea
Pseudacris triseriata

Rana aurora

R. cascadae

R. catesbeiana

R. pipiens
R. pipiens

R. clamitans

R. pretiosa

R. temporaria
R. temporaria


Tadpoles

Tadpoles

Tadpoles
Tadpoles

Tadpoles
Tadpoles

Tadpoles

Tadpoles

Tadpoles

Tadpoles
Tadpoles

Tadpoles

Tadpoles

Tadpoles
Adults


4 mg/L nitrite
Acute: 13.6-39.3 mg/L nitrate
Chronic: 2-10 mg/L nitrate
0.78-25 mg/L nitrate
4 mg/L nitrite
385 mg/L
0.78-25 mg/L nitrate
4 mg/L nitrite
9-22.6 mg/L nitrate
Acute: 13.6-39.3 mg/L nitrate
Chronic: 2-10 mg/L nitrate
0.78-25 mg/L nitrate
4 mg/L nitrite
3.5 mg/L nitrate

9-26 mg/L nitrate

9-26 mg/L nitrate
Acute: 13.6-39.3 mg/L nitrate
Chronic 2-10 mg/L nitrate
Acute: 13.6-39.3 mg/L nitrate
Chronic: 2-10 mg/L nitrate
0.78-25 mg/L nitrate
4 mg/L nitrite
5 mg/L nitrate
3.6-6.9 g/m2 nitrate on
substrate


LC50 < 15 days
LC50 96 h, decreased swimming and feeding
Bent tail, edema, head and digestive deformities
Decreased feeding & activity, bent tails, edema
LC50 < 15 days
LC50 96 h
Decreased feeding & activity, bent tails, edema
LC50 < 15 days
Decreased growth rates, behavior abnormalities, increased
mortality
LC50 96 h, decreased swimming and feeding, bent tail
Edema, head and digestive deformities
Decreased feeding & activity, bent tails, edema
LC50 < 15 days
Decreased rates metamorphosis at earlier stage development

Decreased white blood cells and hemoglobin

Decreased white blood cells and hemoglobin
LC50 96 h, decreased swimming and feeding, bent tail
Edema, head and digestive deformities
LC50 96 h, decreased swimming and feeding, bent tail
Edema, head and digestive deformities
decreased feeding & activity, bent tails, edema
LC50 < 15 days
Decreased growth rates and decreased size at
metamorphosis
Increased toxicity and mortality


Reference


Marco and Blaustein, 1999

Hecnar, 1995

Marco and Blaustein, 1999

Xu and Oldham, 1997

Marco and Blaustein, 1999

Baker and Waights, 1994

Hecnar, 1995

Marco and Blaustein, 1999

Marco and Blaustein, 1999

Dappen, 1983

Dappen, 1983

Hecnar, 1995

Hecnar, 1995

Marco and Blaustein, 1999

Johansson et al., 2001

Oldham et al., 1997














CHAPTER 2
THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS OF NITRATE (IN VIVO) ON PLASMA STEROIDS AND INSULIN-
LIKE GROWTH FACTOR-i, ON OVARIAN STEROID SYNTHESIS, AND ON OVIDUCT
GROWTH IN THE AFRICAN CLAWED FROG (Xenopus laevis)

Introduction

During the last few years there has been increased global concern over contamination of

water by anthropogenic sources of nitrates. Nitrate is among the most stable, water-soluble ionic

forms of nitrogen persistent in aquatic habitats. Nitrate contaminates watersheds in agricultural

and urban environments, reaching harmful concentrations largely due to human activities. In

agricultural areas, nitrate contaminates watersheds primarily through poorly regulated runoff of

nitrogen-based fertilizers and animal wastes from farms. In urban areas, nitrate contaminates

watersheds primarily through release of industrial and wastewater effluent from treatment plants,

runoff of fertilizers applied to lawns and golf courses, and air pollution from the burning of fossil

fuels (Pucket, 1995; Rouse et al., 1999). In temperate North America, concentrations of aquatic

nitrate are highest between the fall and spring when reduced ion uptake by agricultural plants

increases soil nitrate loads leaching from the ground (Hallberg, 1989; Nolen and Stoner, 1995;

Nolen et al., 1995, 1997). Additionally, fertilizers applied in close proximity to watersheds,

coupled with spring rainstorms, contributes to an overwhelming aquatic nitrate pulse that

frequently exceeds 100 mg/L and overlaps the breeding season of many amphibians (Rouse et al.,

1999). Many studies on the effects of nitrate on amphibians have addressed the effects of

toxicological rather than sublethal doses on growth, skeletal, and tissue deformities in juvenile

amphibians (Cooke, 1981; Baker and Waights, 1993; Hecnar, 1995; Watt and Oldham, 1995;

Oldham et al., 1997; Xu and Oldham, 1997; March and Blaustein, 1998; Marco et al., 1999;








Johansson et al., 2001; Chapter 1, Table 1-2). Surprisingly, few studies have investigated effects

of exposure to sublethal nitrate concentrations on adult, reproductive frogs.

There is mounting evidence that nitrate interferes with steroid-signaling pathways.

Panesar and Chan (Panesar, 1999; Panesar and Chan, 2000) demonstrated that administration of

nitrate and nitrite inhibits testosterone (T) synthesis (in vitro and in vivo) in rodents. Once nitrate

enters the body, through consumption or absorption across skin surfaces, it can be converted into

nitrite by endogenous microbial activity in the mouth or gastrointestinal tract (Fried, 1991;

Doblander and Lackner, 1996). Nitrite can be converted into N-nitrosoamines, which are

carcinogens in laboratory animals and in humans (National Academy of Sciences, 1981; Tricker

and Preussmann. 1991; US EPA, 1995). One proposed mechanism for altered steroid expression

by nitrates involves enzyme-dependent synthesis of nitric oxide (NO) (Panesar and Chan, 2000).

The NO is synthesized (in vivo) from an L-arginine precursor by nitric oxide synthase (NOS)

enzymes (Kleinert et al., 1995; Mayer and Hemmenns, 1997). In addition to NOS-dependent NO

formation, non-enzymatic synthesis of NO can also occur through acidic reduction of nitrite

(lizuka et al., 1999; Zweier et al., 1995, 1999; Modin et al., 2001). Cosby et al. (2003) reported

that hemoglobin functions as a nitrite reductase contributing to enzyme-independent NO

synthesis. Regardless of the mechanisms by which it is produced, NO is thought to regulate many

physiological processes. Zweier et al. (1999) reported that enzyme-independent NO formation is

associated with cellular damage and loss of organ function. Panesar and Chan (2000) proposed

that, in steroidogenic tissues, NO binds to the heme groups inherent to mitochondrial cytochrome

P450 enzymes, such as those involved in side-chain cleavage (P450..): the rate-limiting step in

steroid synthesis. The NO can inhibit other P450 enzymes, such as 303-dehydroxysteroid

dehydrogenase (3f-HSD) involved in androgen synthesis; and P450 aromatase (Snyder et al.,

1996) involved in aromatization of androgens to estrogens. Collectively, these P450 enzymes are









necessary for conversion of free cholesterol into progesterone (P4): the steroid precursor for T and

17f3-estradiol (E2).

Various isoforms of NOS are found within the ovary and other steroidogenic tissues in

vertebrates (Szabo and Thiemermann. 1995; Van Voorhis et al., 1995; Srivastava et al., 1997).

Disruption of these enzymes by NO might inhibit P4 synthesis, which would decrease or prevent

downstream T synthesis. Inhibition of gonadal T synthesis likely reduces the T available for

aromatase conversion to E2 and would contribute to decreased overall gonadal E2 synthesis. This

speculation is supported by studies in mammals demonstrating that increased NOS activity and

NO concentrations are associated with decreased ovarian E2 synthesis (VanVoorhis et al., 1994,

1995; Jablonka-Shariff and Olsen, 1997; Srivastava et al., 1997; Dees et al., 2000).

Relatively few studies have reported the impact of nitrate on steroidogenesis, but no

study has investigated the effect of nitrate exposure on of insulin-like growth factor-I (IGF- 1) in

vertebrates. Insulin-like growth factor-I is a potent growth-stimulating hormone that regulates

bone and skeletal muscle growth, limb bud emergence, reproductive and somatic tissue growth,

steroidogenesis, and other physiological functions (Daughaday and Rotwein, 1989; Erickson et

al., 1989; Adashi, 1993; Hiney et al., 1996; Olsen et al., 1996; Dees et al., 1998; Kaliman et al.,

1999; Allen et al., 2001). Thus, IGF-1 is a relevant hormone to examine in the cases of amphibian

skeletal deformities, sex ratio reversal, and reproductive abnormalities. Abnormal expression of

IGF- I is associated with altered growth and function of reproductive tissues in vertebrates.

Increased concentrations of plasma IGF- 1 in humans is positively correlated with cancer of the

endometrium, breast, prostate, skin, pancreas, lung, and colon (Cohen et al., 1991; Lippman,

1993; Papa et al., 1993; LeRoith et al., 1993, 1995; Werner and LeRoith, 1996; Cascinu et al.,

1997; Mantzoros et al., 1997; Stoll, 1997). Despite these reports, IGF- 1 can have beneficial

effects on tissue growth and function. For example, IGF-1 also mediates growth of E2- sensitive

reproductive tissues. In addition to this, IGF- 1 regulates gonadal steroid expression. Intraovarian

IGF-I expression counteracts NO-induced steroid inhibition by increasing aromatase activity and









stimulating E2 synthesis (Daughaday and Rotwein, 1989; Erickson et al., 1989; Adashi, 1993;

Hiney et al., 1996; Olsen et al., 1996; Dees et al., 1998). Furthermore, evidence indicates that NO

stimulates ovarian IGF-l expression (Dees et al., 1998). Thus, NO interacts with the IGF-1

system and influences expression of steroids and also their actions in reproductive tissues.

Based on the aforementioned studies, I hypothesized that nitrate alters concentrations of

steroids and IGF-1, and alters oviduct growth in a model frog species, Xenopus laevis. My study

tested this hypothesis using environmentally relevant concentrations of nitrate.

Materials and Methods

Animals and Samples

Adult female X laevis were purchased from Xenopus Express (Plant City, Florida). This

species is entirely aquatic, and thus would remain in constant exposure to administered

treatments. Frogs were maintained under a 12-h light/dark cycle in 38 L tanks with 19 L of static-

flow, dechlorinated water at 23C (pH 7.0 7.4), with ammonia and nitrite content below 1.0

mg/L as confirmed by daily water measurements. Animals were fed spirulina pellets (Aquatic

Ecosystems, Orlando, FL) every other day for the duration of the experiment. All procedures

were performed with approval of the University of Florida Institute of Animal Care and Use

Committee (IACUC Permit #Z023). Pregnant mare serum gonadotropin (PMSG) and human

chorionic gonadotropin (hCG) were obtained from Sigma-Aldrich (St. Louis, MO), and sodium

nitrate (99% purity) was obtained from Fisher Scientific (Orlando, FL).

Nitrate Study Design

Treatment groups were divided into control (0 mg/L), 150 mg/L, and 300 mg/L sodium

nitrate; respectively equivalent to 0, 24.75, and 49.50 mg/L nitrate-as-nitrogen (N03-N). Nitrate

as nitrogen represents the concentration of nitrogen present in a given concentration of sodium

nitrate administered (Chapter 1, Table 1-1). For the remainder of this chapter, nitrate will refer to

N03-N.








The frogs were randomly assigned to each of 3 replicate tanks per treatment for a total

sample size of 12 frogs per treatment. No significant differences in mass were detected (ANOVA;

P > 0.05) or snout-vent-length (SVL; ANOVA; P > 0.05) among frogs in each treatment group.

After a 1-week acclimation period, frogs were injected into the dorsal lymph sac with 50 IU of

PMSG, followed 3 days later by an injection of 750 IU hCG. These treatments stimulated

ovulation and formation of new ovarian follicles within 6 weeks (Dumont, 1971; Fortune and

Tsang, 1981; Fortune, 1983). This procedure synchronized the size and maturation of new

follicles before nitrate exposure, and minimized possible variation in gonadal steroid synthesis

among frogs in response to treatment.

After 6-weeks, frogs were exposed to nitrate applied to tank water for 7 consecutive days.

Every 24 h, water was changed, and fresh water with nitrate was added. After 7 days, the frogs

were anesthetized with MS-222 (1.5% 3-aminobenzoic acid ethyl-ether, Aquatic Ecosystems,

Orlando, FL). Blood was collected by cardiac puncture using heparinized syringes, placed into

heparin vacutainer tubes, and centrifuged (2500xG) for 15 min; and plasma was stored at -70'C

for E2, T, and IGF- 1 radioimmunoassay (RIA) analysis. The ovaries were removed and weighed,

and follicles were dissected for a culture study (ex vivo). Follicles of specific maturation stages

were chosen: stage 4 follicles synthesize E2, and stage 5 and 6 follicles synthesize T (Fortune and

Tsang, 1981; Fortune, 1983;). From each frog, 33 follicles, each of stages 4, 5, and 6 were

incubated in 35x 10 mm sterile culture dishes, in duplicate, at 230C with 2 mL of sterile, phenol-

free culture media (IL M199 HBSS, 3.4 mL 200 mM L-glutamine, 5.96 g/L HEPES, 0.35 g/L

sodium bicarbonate, 8.0 mL 0.1 mM IBMX, pH 6.9; Sigma-Aldrich, St. Louis, MO) for both 5

and 10 h. Follicles were incubated at the same temperature Incubation temperature was selected

based on the water temperature maintained in the tanks holding X laevis. After incubation,

culture media was decanted, flash-frozen, and stored at -70'C for E2 and T RIA. The diameter of

the remaining, uncultured follicles was measured with a dissecting microscope and an ocular









micrometer. For each follicle stage, 5 follicles (un-cultured) were measured in each frog from 0

mg/L (control, N = 8), 24.75 mg/L nitrate (N = 10), and 49.50 mg/L nitrate (N = 8) treatment

groups. Sample sizes of frogs were uneven among treatment groups for follicle measurements

because for some frogs, all of the follicles were incubated in the culture study. Ovary, liver, and

oviduct weights were recorded to compare post-treatment tissue weights among groups.

Steroid Radioimmunoassay (RIA) Procedures

RIAs were performed for E2 and T (Guillette et al., 1994; Guillette et al., 1996) on culture

media and on plasma samples using validated procedures. Duplicate media samples or plasma (50

gL for E2 T) were extracted twice with ethyl-ether, air-dried, and reconstituted in borate buffer

(0.05 M; pH 8.0). Antibody (Endocrine Sciences) was added at a final concentration of 1:55,000

for E2 and of 1:25,000 for T. Radiolabeled steroid ([2,4,6,7,16,17-3H] estradiol at I mCi/mL;

[1,2,6,7-3H] and testosterone at 1 mCi/mL; Amersham Int., Arlington Heights, IL) was added at

12,000 cpm per 100 gL for a final assay volume of 500 gL. Interassay variance tubes were

prepared from two separate pools of media and of plasma for E2 and T. Standards for E2 and T

were prepared in duplicate at 0, 1.56, 3.13, 6.25, 12.5, 25, 50, 100, 200, 400, and 800 pg/tube.

Assay tubes were vortexed and incubated overnight at 4C.

Bound-free separation was performed using a mixture of 5.0% charcoal to 0.5% dextran,

pulse-vortexing, and centrifuging tubes (1500g, 4C, 30 min). Supernatant was added to 5 mL of

scintillation cocktail, and counted. Media intraassay and interassay variance averaged 2.50% and

3.70% for E2, and 4.20% and 8.38% for T, respectively. Plasma intraassay variance for E2 and T

averaged 4.20% and 4.60%, respectively. Plasma E2 and T samples were run in a single assay and

interassay variances are not reported.

Validation of the steroid assays included media and plasma dilutions (50, 100, and 200

gL for E2 and 20, 50 and 100 j.L for T) compared with E2 and T standards.








Insulin-Like Growth Factor-1 RIA Procedures

The IGF-1 RIA was performed as described by Crain et al. (1995). The National

Hormone and Pituitary Program (Torrance, CA 90509) supplied human recombinant IGF-1

standard (9.76 to 2500 pg/tube) and human IGF- I antisera (Lot # AFP4892898, 1:400,000 final

dilution). The antiserum had less than 1.0% cross-reactivity with human IGF-II. lodinated IGF- I

label (IGF- 11125 sp act 2000 Ci/mmol; 16,000 cpm/tube) and Amerlex-M donkey anti-rabbit

secondary antibody (RPN5 10) were obtained from Amersham International (Arlington Heights,

IL).

For each treatment group, plasma was pooled (8, 16, 24, and 36 p.L aliquots in borate

buffer) for validation using plasma dilutions (equivalent to 1.9, 3.9, 5.7, and 8.3 J.L plasma) that

were compared with IGF-1 standard. Plasma validation and experimental samples (20 gL) were

acid-ethanol extracted and IGF-1 RIA performed (Crain et al., 1995). Validation samples were

run in one assay with intraassay variance averaging 3.10%.

Biochemical RIA Validations

Plasma dilutions and internal standards were parallel to E2 standards (ANCOVA; F

0.48; P = 0.52 and F = 0.35; P = 57, Fig. 2-A) and recovery of E2 after extraction was 81.0%.

Plasma dilutions and internal standards were parallel to T standards exhibited parallel

displacement (ANCOVA; F = 0.12, P = 0.33 and F = 1.18, P = 0.31, Fig. 2-1B) and recovery of T

after extraction was 93.8%. Plasma dilutions and IGF-1 standards exhibited parallel displacement

curves (ANCOVA; F = 0.08; P = 0.79, Fig. 2-1C) and recovery of IGF-1 after extraction was

78.0%.

Media dilutions and E2 standards gave parallel displacement curves (ANCOVA; F

1.05; P = 0.37, Fig. 2-2A). Recovery of E2 after media extraction was 91.5% and all sample

values were corrected for loss using this value. Media dilutions and T standards gave parallel

displacement curves (ANCOVA; F = 0.60; P = 0.48, Fig. 2-2B). Recovery of T after media









extraction averaged 98.9% and all sample values were corrected for this loss. For subsequent

steroid and IGF- 1 analyses, all sample values were corrected for respective losses.

Statistics

Ovary, oviduct, and liver wet mass were compared among treatment groups with body

mass as a covariate using ANCOVA, followed by LSD post-hoc contrasts. Concentrations of E2,

T, and IGF- I were estimated from raw data using Microplate Manager software (Microplate

Manager III, BioRad Laboratories, Inc., Hercules, CA, 1988). Statistical analyses were performed

using SPSS software (v. 10, SPSS Inc., Chicago, IL, 1999) with C = 0.05. ANCOVA was used to

validate plasma and media samples and to determine if plasma IGF- 1 concentrations were

correlated to body mass. Concentrations of E2, T, and IGF- 1 among replicate tanks within each

treatment group were compared using one-way ANOVA. Where no significant difference existed

among replicate tanks within treatment groups, mean E2, T, and IGF-1 concentrations were

compared among treatment groups with one-way ANOVA. Ovarian follicle diameters were

compared, separately according to stage, among treatment groups with one-way ANOVA.

Following one-way ANOVA analyses Scheffe post-hoc contrasts were used. Tamhane post-hoc

contrasts were used where variances were unequal among groups for plasma IGF- 1

concentrations.

Results

Tissue Weights

Tissue weights were not different among treatment groups for ovary (ANCOVA; F =

0.57, P = 0.57), oviduct (ANCOVA; F = 0.28, P = 0.76), and liver (ANCOVA; F = 1.13, P =

0.34).

Follicle Diameters

Diameter of stage 4 follicles was larger (ANOVA; P = 0.01, Fig. 2-3A) in frogs exposed

to 24.75 mg/L and 49.50 mg/L nitrate relative to frogs exposed to 0 mg/L. Mean diameter was

smaller in stage 5 (ANOVA; P = 0.04, Fig. 2-3B) and stage 6 follicles (ANOVA; P = 0.005, Fig.








2-3C) in frogs exposed to 49.50 mg/L nitrate compared to frogs exposed to 24.75 mg/L nitrate

and 0 mg/L.

Plasma Steroid Concentrations

Analyses revealed no significant difference (P > 0.05) in E2 or T concentrations among

replicate tanks; thus data from frogs in replicate tanks was combined per treatment group. Plasma

E2 was not significantly different among treatment groups (ANOVA; P = 0.08). Plasma T did not

differ among treatment groups (ANOVA; P = 0.70).

Plasma IGF-1 Concentrations

Analyses revealed no significant difference in IGF- 1 concentrations among replicate

tanks; thus data from frogs in replicate tanks was combined per treatment group. Plasma IGF- 1

concentrations were significantly higher in frogs exposed to 24.75 mg/L and 49.50 mg/L nitrate

relative to the control frogs (ANOVA; P = 0.007, Fig. 2-4). Plasma IGF-1 was not significantly

correlated to body mass (ANOVA; RW = 0.12, P > 0.05).

Ovarian Follicle Steroid Concentrations (Ex Vivo)

Statistical analyses revealed no significant difference in mean E2 or T (P > 0.05)

concentrations among replicates for each treatment group; thus, data from frogs in replicate tanks

was combined per treatment group. After 5 h, media E2 concentrations were significantly lower

for ovarian follicles of frogs exposed to 49.50 mg/L nitrate compared to the other treatment

groups (ANOVA; P < 0.00 1, Fig. 2-5A). However, after 10 h, media E2 concentrations were

significantly lower for ovarian follicles of frogs exposed to both the 24.75 mg/L and 49.50 mg/L

nitrate relative to the controls (ANOVA; P < 0.001, Fig. 2-5B).

After 5 h, media T concentrations were similar among treatment groups (ANOVA; P >

0.05, Fig. 2-6A). However, after 10 h, media T concentrations were significantly lower for

ovarian follicles from frogs exposed to 24.75 mg/L and 49.50 mg/L nitrate relative to the control

group (ANOVA; P < 0.001, Fig. 2-6B).









Discussion

This study has shown that exposure of X laevis to sublethal doses of aquatic nitrates at

environmentally relevant concentrations (24.75 mg/L and 49.50 mg/L) is associated with

endocrine disruption of E2, T, and IGF-1. This study raises new and troubling questions regarding

the effects of nitrates on endocrine function. No other study has examined the effects of exposure

to sublethal concentrations of nitrate on adult anurans, despite reports of altered growth, behavior,

and mobility in tadpoles at similarly low (1 40 mg/L) nitrate concentrations (Baker and Waights,

1994; Hecnar, 1995; Xu and Oldham, 1997; Marco and Blaustein, 1999 Johansson et al., 2001.

Over the past 30 years amphibian populations have declined in various regions of the

world (Wake, 1991; McCoy, 1994), especially in agricultural landscapes (Dappen, 1983; Berger,

1989; de Solla et al., 2002). Alteration of aquatic habitats is considered a primary contributor to

these declines (Blaustein and Wake, 1990; Carey and Bryant, 1995). Altered endocrine function

in frogs has been associated with exposure to sublethal concentrations of various contaminants

(Mohanty-Hejmadi and Dutta, 1981; Carey and Bryant, 1995; Reeder et al., 1998; Kloas et al.,

1999; Hayes et al., 2002). In agricultural and urban areas, contamination of aquatic habitats by

anthropogenic sources of nitrate poses a serious threat to wildlife and humans. Approximately 72

million tons of nitrogen-based fertilizers are used worldwide and, combined with release of

industrial nitrogenous wastes, are likely responsible for increased nitrate contamination reported

in surface waters, aquifers, and drinking water (Rouse et al., 1999). Most studies examining the

effects of sublethal nitrate concentrations on frogs have focused on juvenile stages from egg

through metamorphosing tadpole. There is an absence of research examining the effects of

sublethal nitrate concentrations on the endocrine profile of adult, reproductive frogs.

Panesar and Chan (2000) reported inhibition of T synthesis (in vitro) in rodents after

exposure to nitrate. Within body tissues, various isoforms of NOS enzymes are capable of

converting nitrates into NO (VanVoorhis et al., 1994, 1995; Srivastava et al., 1997; Olsen et al.,

1996; Jablonka-Shariff and Olson, 1997). In addition, acidic reduction and hemoglobin have been








shown to mediate non-enzymatic NO formation from nitrite (in vivo) (Zweier et al., 1995, 1999;

Modin et al., 2001; Cosby et al. 2003). Many studies have shown that NO inhibits E2 and T

synthesis in rodents, humans, and cows (VanVoorhis et al., 1994; Wang and Marsden, 1995;

Basini et al., 1998; Omura, 1999). Panesar and Chan (2000) proposed a mechanism (based on a

synthesis of their work and that of other researchers) involving formation of NO. Nitrate and

nitrite can be converted to NO within steroidogenic cells, and the NO inhibits steroidogenic P450

enzymes necessary for conversion of free cholesterol to steroid precursors. In addition to

inhibiting P450 enzymes, NO has also been shown to inhibit steroid-acute regulatory protein

(StAR) protein expression. During steroidogenesis, StAR protein is essential for transporting free

cholesterol to the inner mitochondrial membrane (Wang and Marsden, 1995). I propose a similar

nitrate-associated steroid inhibition, possibly involving NO formation, occurred within ovarian

follicles of X laevis. This steroid inhibition also and includes downstream inhibition of E2

synthesis and stimulation of IGF-1.

In X laevis exposed to nitrate ovarian steroid synthesis was inhibited (ex vivo) while

plasma steroid concentrations (in vivo) were unaffected. These findings indicate that different

mechanisms were involved in regulating ex vivo versus in vivo steroids in nitrate-exposed frogs. It

is possible plasma steroid concentrations were unchanged due to compensatory responses of the

hypothalamic-pituitary-gonadal (HPG) axis (Chapter 1, Fig. 1-1). Inhibition of steroid synthesis

at the gonad level might have signaled a compensatory hypothalamic release of gonadotropin-

releasing hormone (GnRH) causing pituitary release of luteinizing hormone (LH) and follicle-

stimulating hormone (FSH) into the blood. Increased plasma LH/FSH concentrations would

stimulate ovarian synthesis of T and E2, which could have contributed to normal circulating

plasma steroid concentrations. In this study, ovarian ex vivo follicle steroid synthesis was

recorded without measuring corresponding plasma gonadotropins. It is unknown if plasma steroid

concentrations in nitrate-exposed frogs were maintained at levels similar to control frogs by

compensation by the HPG axis. It is unlikely that compensatory responses of the HPG axis to








stimulate steroidogenesis by the gonads would influence plasma steroid concentrations because

ovarian steroid synthesis was shown to be inhibited in nitrate-exposed frogs. Thus, gonadotropins

would not be effective in stimulating steroid synthesis in nitrate-exposed frogs when

steroidogenesis is inhibited at the level of the gonad. Therefore, another explanation must be

considered.

The liver is the main organ for degradation of nitrate, and degradation of nitrate can

elevate hepatic NO concentrations. Continuous administration of nitrate has been shown to

increase hepatic NO synthesis and inhibit hepatic P450 enzymes activity (Minamiyama et al.,

2004). Hepatic P450 enzymes are necessary for metabolism and excretion of circulating steroids.

Thus, hepatic nitrate degradation can lead to NO formation and inhibition of hepatic P450 steroid

metabolic enzymes. Reduced hepatic steroid metabolism could cause stasis or even augmentation

of circulating steroid concentrations.

I propose a mechanism for the increase in plasma IGF- I concentrations (in vivo)

observed in nitrate-exposed X laevis. Nitrate, once consumed or absorbed across skin surfaces,

can be converted by microbial activity in the mouth and gastrointestinal tract to nitrite. Nitrite has

been shown to stimulate hypothalamic NO formation and increase hypothalamic secretion of

growth hormone-releasing hormone GHRH and pituitary release of growth hormone (GH) (de

Caceres et al., 2003). Thus, the hypothalamic-pituitary-hepatic (HPH) axis regulates circulating

IGF- 1 concentrations, and this axis is influenced by nitrite and NO exposure (Fig. 2-7). Further

research will be necessary to confirm the validity of this proposed pathway.

In addition to the liver, the ovary also produces IGF-1, although in relatively smaller

quantities (Adashi, 1993). Stimulation of the ovary by pituitary FSH results in decreased

synthesis of IGF-1-binding proteins and increased intraovarian IGF-1 synthesis and availability.

Intraovarian IGF- I might have an autocrine and endocrine effect of ovarian steroid synthesis

(Grimes et al., 1992; Adashi, 1993; Basini et al., 1998). Increased IGF-1 has been shown to

increase intraovarian aromatase activity and E2 synthesis (Erickson et al., 1989; Monnieaux and









Pisselet, 1992; Adashi, 1993; Samaras et al., 1994; Samaras et al., 1996). However, plasma IGF-l

concentrations increased and ovarian E2 concentrations decreased in X laevis upon nitrate

exposure. Perhaps the in vivo nitrate exposure period of 7 days was too brief to observe a

compensatory increase in ovarian steroid synthesis with IGF- 1 stimulation.

Plasma IGF-1 binding proteins (IGF-BP) play an important role in regulating the

availability of IGF-1 to and within tissues. In this study, IGF-BP in the plasma and the ovaries

were not measured, so the availability of increased plasma IGF-1 in nitrate-exposed animals

merits investigation. If plasma IGF- 1 increased in conjunction with a decrease in tissue IGF I -BP,

then there might be an increase in IGF-1 utilization and growth response by tissues. In this study,

there was no difference detected in ovary, oviduct, or liver tissue mass among nitrate treatment

groups. This might indicate either that the increased circulating IGF-1 was not stimulating a

growth response in these tissues or that circulating IGF- 1 was bound to IGF-BP and unavailable

for tissue uptake. Although no difference in total ovary weights was detected among treatment

groups, follicle diameter varied among groups. The diameter of E2-producing follicles (stage 4)

were larger in nitrate-exposed frogs compared to control, which might reflect a growth response

to increased IGF- 1 exposure or compensatory tissue growth in response to declining E2 levels.

The diameter of T-producing follicles (stage 5 and 6) was smaller in frogs exposed to 49.50 mg/L

nitrate compared to follicles of frogs exposed to 24.75 mg/L and 0 mg/L. This could reflect either

the absence of IGF-1 uptake by these follicles or an absence of a growth-response to IGF-1.

This study raises new and troubling questions regarding the effects of nitrates on

endocrine function in vertebrates. Chemical alteration of aquatic habitats is considered a foremost

contributor to the declines and deformities reported for amphibian populations (Carey and Bryant,

1995; Wake, 1998; Hayes et al., 2002). Amphibians exposed to various contaminants, even at

sublethal concentrations, exhibit malformations, reproductive abnormalities, sex ratio reversal,

male feminization, and altered endocrine function (Reeder et al., 1998; Kloas et al., 1999; Hayes

et al., 2002). The nitrate-associated endocrine disruption in X laevis might differ from other








anuran species due to the interspecific variation in physiological response to nitrates (Chapter 1,

Table 1-2). Further research is necessary to determine whether nitrate alters steroid and IGF-1

hormones in other anuran species, and to describe the range of sublethal nitrate concentrations

capable of endocrine disruption. The nitrate concentrations used in this study were relevant to

environmental concentrations measured in North American ground and surface water (Rouse et

al., 1999; Nolen and Stoner. 1995). However, it would be valuable to ascertain if even lower

nitrate concentrations have a similar endocrine disrupting capacity in frogs.

More research is needed to elucidate the mechanism by which nitrate inhibits steroid

synthesis and increases circulating IGF- 1 concentrations in amphibians and in other animals.

Thus far, most reports of steroid inhibition by nitrate have focused on steroid synthesis and

regulation exclusively at the gonad level. It is important to consider both upstream and

downstream steroid regulation. Upstream regulation would changes in hypothalamic and pituitary

hormone secretions in response to in vivo nitrate exposure. The important hormones to examine

include hypothalamic GnRH and GHRH, and pituitary LH, FSH, and GH. Pituitary LH and FSH

function in stimulating gonadal steroid synthesis and GH stimulates hepatic IGF- 1 synthesis.

Downstream regulation would include hepatic degradation and clearance of circulating steroids,

and secretion of IGF-1. In addition to these topics, it is important to determine whether amphibian

gonadal tissue contains NOS enzymes capable of synthesizing NO. It has been already been

established that the amphibian brain contains NOS capable of generating NO (McLean et al.,

2001; Gonzalez et al., 2002; McLean and Bilar, 2002). Furthermore, it is necessary to understand

how nitrate exposure of frogs regulates intracellular expression of NOS, NO, steroidogenic

enzymes, and steroid regulatory proteins. Lastly, since nitrate exposure is associated with changes

in circulating IGF- 1 concentrations, in addition to steroid synthesis, is vital to understand how

these hormones collectively influence the reproductive physiology of amphibians.









0 0 0 mp
0
o a


E2 (pg)


0.
o0*


T (pg)


o Standard
Internal Standards
0. Plasma Dilutions

0*
0 0
M 0 0


100


1000


Standard
Internal Standards
Plasma Dilutions


O *
0


100


1000


0


100
80
60
40
20
0


100


o Standard
* Plasma Dilutions


1000


10000


IGF-1 (pg)
Figure 2-1. Biochemical validation of Xenopus laevis plasma. A. estradiol RIA internal standards
(ANCOVA; F = 0.35; P = 0.57) and plasma dilutions (ANCOVA; F = 1.86; P =
0.23) were parallel to the standard curve. B. testosterone RIA. Internal standards
(ANCOVA; F = 1.18; P = 0.31) and plasma dilutions (ANCOVA; F = 0.12; P
0.33) were parallel to the standard curve. C. IGF-1 RIA. plasma dilutions
(ANCOVA; F = 1.05; P = 0.37) were parallel to the standard curve.


100
80
60
40
20
0


B 100
~80
- 60
~40
~20
0



















E2 (pg)


B 100


0ov
60

.,40
.20


o Standard
* Media Dilutions


0


100


1000


o Standard
* Media Dilutions


100


1000


Figure 2-2. Biochemical validation of Xenopus laevis media. A. estradiol RIA media dilutions
(ANCOVA; F = 0.08; P = 0.79) were parallel to the standard curve. B. testosterone
RIA. Media dilutions (ANCOVA; F = 0.60; P = 0.48) were parallel to the standard
curve.


100
80
60


0


.0


0-*


T (pg)










A Stage 4
1.04
L bb
O0.98 a

E 0.92

0.86
0 o
~ 0.80


B 1.40 Stage 5
BL 1.30 a
ia
:t 1.30 ia b

z 1.20 1


1.10


S1.00

C Stage 6
1.30 a

= 1.24 T
b
2 1.18

1.12

-- 1.06
*o 1.00

'i en en -T N
Nitrate (mg/b)
Figure 2-3. Diameter of ovarian follicles of stages 4, 5, and 6 in Xenopus laevis exposed in vivo
for 7 days to 0, 24.75, and 49.50 mg/L nitrate. Data presented as means SEM.
Different letters above bars indicate significant differences for: A. stage four
(ANOVA; P = 0.01), B. stage five (ANOVA; P = 0.040, and C. stage six (ANOVA;
P = 0.005).












60 b b

50
40Oa






o 0 0 2,0 0
o o 0 0 0 r- 0 0 0 W)


Nitrate (mg/L)
Figure 2-4. Plasma insulin-like-growth factor-I (IGF- 1) in Xenopus Iaevis after 7 days of in vivo
exposure to 0, 24.75, and 49.50 mg/L nitrate. Data presented as means SEM.
Numbers within bars indicate sample sizes and different letters above bars indicate
significant differences (ANOVA; P = 0.007).









5h


A 350
300
250
200
P 150
V 100
50
S 0~



B 350
300 -i
-, 250
200
'U 150
100

.'U 50
40


10 h


b







o... 0 0 C0 0 0 0. 0)
o 0 0> 0 0 rl 0 0 0 0 W)
0In0 kl 6 tf 6 V C
M I IRT I-


Nitrate (mg/L)

Figure 2-5. Media 170-estradiol concentrations in culture media from incubated ovarian follicles
of Xenopus laevis after 7 days in vivo exposure to 0, 24.75, and 49.50 mg/L nitrate.
Data presented as means SEM for A. 5 h and B. 10 h of incubation. Numbers within
bars indicate sample sizes and different letters above bars indicate significant
differences for B. (ANOVA; P < 0.001).


a
b





0 I 0 W 0 C> 0 C)C
0 In 0 0 ci ci cl> kn~


$11


i











A 800 a

600

400

-V

0
o. 0


5h


o 0 0 0 LO 0 0 0 0 0
o 0 0 0 N-. 0 0 0 0 '0


B 2000 1 a


10 h


" 1600

,1200

0800

400

0-


o 0 0 0 0 LO 0 0 0 0) 0
o C o 0 0 N%- 0 0 0 0 U.
o '0 0 u'0 0 0 1'0 0 L' 0


Nitrate (mg/L)

Figure 2-6. Media testosterone concentrations in culture media from incubated ovarian follicles of
Xenopus laevis after 7 days in vivo exposure to 0, 24.75, and 49.50 mg/L nitrate. Data
presented as means SEM. Numbers within bars indicate sample sizes and different
letters above bars indicate significant differences for A. 5 h (ANOVA; P = 0.007)
and B. 10 h (ANOVA; P < 0.001) of incubation.










B.E

4

4-


Nitrite Nitrate D.


A.
Nitrate

4
Nirte


t
Nitrate


C.


Figure 2-7. Diagram of mechanism for nitrate-associated inhibition of steroidogenesis and
increased plasma IGF-1. Only ovarian testosterone, (T), estradiol 17P (E2), and
plasma T, E2, and insulin-like growth factor-I (IGF- 1) were measured in Xenopus
laevis exposed (in vivo) for 7 days to 0, 24.75, and 49.50 mg/L nitrate (N03-N). Other
parameters are adapted from other studies (Licht 1984; Panesar and Chan, 2000; de
Caceres et al., 2003; Minamiyama et al., 2004). A. Ovarian steroid synthesis is
inhibited by nitric oxide (NO) formation from nitrate and nitrite. The NO inhibits
cytochrome P450 steroidogenic enzymes. NO might also inhibit steroid-acute
regulatory (StAR) protein which escorts free cholesterol into the mitochondria.
Inhibition of these enzymes reduces progesterone (P4) synthesis, and reduces T
available for aromatization (Arom)to E2. B. Decreased steroid synthesis could signal
compensatory hypothalamic secretion of gonadotropin-releasing hormone (GnRH)
pituitary secretion of luteinizing hormone (LH) and follicle stimulating hormone
(FSH). C. Hepatic nitrate metabolism can cause NO inhibition of P450 enzymes
involved in hepatic steroid metabolism and clearance resulting in augmented
circulating steroid concentrations. D. Nitrate and nitrite could cause hypothalamic NO
formation, which can stimulate secretion of growth hormone-releasing hormone
(GHRH) and secretion of pituitary growth hormone (GH). The GH stimulates liver
IGF-1 synthesis and secretion into the blood.














CHAPTER 3
SEASONAL CHANGES IN INSULIN-LIKE GROWTH FACTOR-I, STEROIDS, AND
REPRODUCTIVE TISSUES IN PIG FROGS (Rana grylio)

Introduction

The sex steroids, 17p-estradiol (E2) and testosterone (T), regulate virtually every facet of

reproduction, and in ectotherms these hormones are responsive to changes in temperature, pH,

and photoperiod among other environmental factors (Licht, 1970; Feder and Burggren, 1992;

Norris, 1997; Kim et al., 1998).

Only one comprehensive profile of the pattern of seasonal changes in circulating steroid

concentrations and changes in gonadal growth and maturation has been reported for a population

of wild bullfrogs, Rana catesbeiana (Licht et al., 1983). Female R. catesbeiana exhibited a

seasonal pattern of changes in plasma concentrations of gonadotropins and steroids, and in

relative weights of reproductive tissues that indicate reproductive and non-reproductive periods.

Reproductive period is here defined as physiological conditions that are optimal for reproduction.

such as elevated plasma concentrations of the reproductive steroids E2, T, and progesterone (P4),

and also by elevated weights of reproductive tissues such as the ovaries and oviducts.

Reproductive condition of the frogs was also discerned by elevated plasma concentrations of the

gonadotropins luteinizing hormone (LH), follicle stimulating hormone (FSH). In R. catesbeiana,

E2, T, and progesterone (P4) concentrations were greatest in the reproductive period between May

and July. The non-reproductive period of frogs is defined here as the physiological condition

marked by decreased plasma concentrations of steroids and gonadotropins, and decreased weights

of reproductive tissues. Licht et al. (1983) reported that plasma steroid concentrations and

gonadal-somatic index (GSI) declined sharply after July and remained depressed between August

and February, indicating the frogs were in non-reproductive condition. Similarly, plasma LH and








FSH concentrations declined precipitously by July of both years (Licht et al., 1983). Increased

plasma steroid concentrations were likely stimulated by the elevated plasma gonadotropin

observed. Elevated concentrations of LH stimulate gonadal steroidogenesis whereas elevated

concentrations of FSH stimulate increased ovarian mass or gonadal somatic index (GSI) in

females during the reproductive period (Licht, 1970, 1979; Norris, 1997). Plasma T

concentrations in females greatly exceeded that of E2 at all times, and plasma T concentrations

were highly correlated with ovarian developmental stage. The relatively high T concentrations

might serve as a circulating androgen pool for synthesis of E2 by aromatase activity in peripheral

tissues such as the brain, fat and skin, and even the oviduct (Follett and Redshaw, 1968). Plasma

androgen pools also might serve functions unrelated to E2 synthesis. For example, it has been

reported that T synthesized by Xenopus ovaries might function to stimulate oocyte development

directly through androgen receptors (Lutz et al., 2001).

In addition to steroid hormones, insulin-like growth factor-I (IGF-1) regulates many

aspects of reproduction including gonadal function and steroidogenesis (Adashi et al., 1991;

Hammond et al., 1991). The presence of IGF- 1 has been identified in representative animals from

all vertebrate classes and includes humans, cows, rodents, birds, alligators, turtles, fish, and

amphibians (Daughaday et al., 1985; Pancak-Roessler and Lee, 1990; Crain et al., 1995; Guillette

et al., 1996; Le Roith et al., 2001a,b; Table 3-1). IGF-1 is a polypeptide hormone that stimulates

cell growth in somatic and reproductive tissues and orchestrates many aspects of development,

metabolism, and steroidogenesis (LeRoth et al., 2001 a,b). In response to pituitary growth

hormone (GH), the liver secretes IGF-1 into circulation complexed to IGF-1 binding proteins

(IGF- 1BPs). IGF- I interacts with IGF- I receptors located on tissues throughout body. Although

initially described as an intermediate of GH action on skeletal muscle growth, more recently IGF-

I has been recognized as hormonal regulator of many GH-independent cellular processes (Butler

and Le Roith, 2001; Le Roith et al., 2001a). Recent research has shown that IGF-1 synthesized

within endometrial and ovarian tissue functions as a paracrine and autocrine hormone (Adashi,








1993). Studies in mammals demonstrate that increased intraovarian IGF- I increases ovarian P4

and E2 synthesis, as well as aromatase, and steroid-acute regulatory protein (StAR) protein

expression (Adashi et al., 1991; Adashi, 1993; Samaras et al., 1994, 1996; Devoto et al., 1999).

Aromatase is a cytochrome P450 enzyme necessary for converting androgens into estrogens, and

StAR proteins assist the entry of free cholesterol into the mitochondria to initiate steroid synthesis

in steroidogenic tissues. Collectively, these findings demonstrate that IGF- 1 is an important

regulator of gonadal steroids. Other studies have shown that intraovarian IGF- 1 regulates

selection of dominant follicles for ovulation in mammals (Adashi et al., 1991; Giudice, 1999).

These studies indicate that IGF- 1 also plays a vital role in steroid synthesis, regulation, and

gonadal function.

Although the IGF- I system has been described in mammals, comparatively few studies

have examined this system in non-mammalian vertebrates (Table 3-1). Oviparous vertebrates are

intriguing models for examining the role of IGF- 1 in reproduction because they lack a prolonged

period of maternal and fetal chemical and nutritive interaction during embryonic development.

Nutrients and growth promoting substances, like IGF- 1, must be sequestered into eggs before

oviposition and fertilization (Guillette et al., 1996). In a turtles, geckos, and alligators, the

presence of plasma IGF- I has been confirmed and demonstrated to play an important role in

mediating reproduction (Daughaday et al., 1985; Cox and Guillette, 1995; Crain et al., 1995a,b;

Guillette et al., 1996). Cox and Guillette (1995) demonstrated that ovariectomized (lacking

endogenous E2) geckos, exhibited an estrogen-like proliferation of oviductal tissue in response to

treatment with IGF- I implants. Additionally, plasma IGF- 1 concentrations vary according to

season and stages of reproductive maturation in female alligators and turtles (Crain et al., 1995;

Guillette et al., 1996). These studies indicate IGF-1 plays a more important role in the growth of

reproductive tissues than previously realized.

Unfortunately, the importance of IGF- 1 in amphibian reproduction and growth remains

largely under-investigated (Daughaday et al., 1985; Pancak-Roessler and Lee, 1990; Table 3-1).








Only one study reported seasonal changes of plasma IGF- I concentrations in a wild population of

Bufo woodhousei (Pancak-Roessler and Lee, 1990). Although this study was limited to a 10-

month profile, it was evident that IGF-I concentrations peaked during the reproductive period

between May and June and decreased during the non-reproductive period between August and

December (Pancak-Roessler and Lee, 1990).

No study has provided a simultaneous examination of seasonal changes in plasma steroid

and IGF- 1 concentrations, and in gonadal growth in a wild population of frogs. In order to

understand the functional relationships among reproductive steroids, IGF-1, and reproductive

tissues in frogs, it is essential to describe how these parameters fluctuate naturally under the

influence of temporal and environmental factors. The objective of the following study was to

document changes in concentrations of plasma IGF- I and reproductive steroids (E2 and T), and in

gonadal tissues in conjunction with environmental factors for a population of wild female Pig

frogs (Rana grylio) in a north-central Florida lake. Rana grylio were chosen for this study

because they were abundant, they were relatively easy to acquire year-round, and they are the

largest ranid frogs in Florida, which made them ideal for the tissue and blood collections required

in this study. Additionally, R. grylio are closely related to bullfrogs (R. catesbeiana), a species for

which documented seasonal profiles of E2 and T served as a reference for this study (Castellani,

1958; Licht et al., 1983). Finally, the seasonal trends of gonadal maturation and breeding activity

for R. grylio have been well-established (Ligas, 1960; Lamb, 1983). The seasonal pattern of

changes in IGF-1 and sex steroids of wild-caught R. grylio established in this study serve as an

ecologically relevant reference for comparison with findings presented in other chapters.

Materials and Methods

Water Parameters, Animal Captures and Sample Collections

From April of 2002 to July of 2003, 6 20 adult female R. grylio were collected during

the fourth week of each month from Orange Lake (Lat. 290 27'853'N, Long. 82' 11.380'W), in

Alachua County, Florida (Fig 3-1). In October of 2002, frogs were not collected due to rain and








lightening storms encountered on the lake during 3 separate collection attempts. Animals were

collected by hand from an airboat between 10 pm and 12 am. Captured frogs were transported, in

covered buckets with a small amount of water, to the Dept. of Zoology where they were housed

for less then 12 h in 38 L tanks with 19 L of dechlorinated water before examination.

Ligas (1960) reported that environmental factors such as rainfall, air temperature, and

water temperature influence reproductive condition of R grylio in the Everglades; therefore, these

same parameters were measured at the collection site. Water temperature and pH were measured

using a Myron L Ultrameter (model 6P, Carlsbad, CA 92009). Monthly rainfall and air

temperature data from Orange Lake were recorded by Weather Station Number 02741536 and

were kindly provided by David Clapp of the USGS and National Weather Service.

Additionally, water samples from the collection site were examined for nitrate and nitrite

concentrations. Low precipitation combined with low water levels during the first 4 months of

this study might have contributed to slightly eutrophic conditions within the collection site.

Nitrate in known to interfere with gonadal steroidogenesis (Panesar and Chan, 2000) and with

amphibian reproduction (Rouse et al., 1999). Thus, nitrate was an important parameter to measure

when documenting plasma steroids of frogs collected at this site. Water nitrate and nitrite

concentrations were measured, with the generous assistance of Thea Edwards, using an auto-

analyzer (Technicon auto-analyzer II with colorimeter, Bran+Luebbe Inc., Chicago, (888)917-

PUMP) equipped with a copper-cadmium reductor column. Methods for use are given in

Bran+Luebbe method number US-158-71 C, which is equivalent to EPA method 353.2. The auto-

analyzer has a detection limit of 0.43 g.g/L with a detection range of 0-400 ;.tg/L of nitrate as

nitrogen. Samples are diluted in distilled water to fall within the detection range. Prior to analysis,

samples are filtered through a 1 micron glass fiber filter, collected in new or acid-washed

containers, and frozen (1 month) prior to measurement. Samples were quantified on a standard

curve created with each batch of water samples.








Previous studies on wild-caught bullfrogs reported that increasing duration of captivity

significantly decreased plasma hormones (Licht et al., 1983). Thus, a pilot study was performed

to determine the influence of duration of captivity on plasma hormone concentrations in R. grylio.

Blood samples were collected from frogs at 0, 6, 12, and 24 h post-capture. Blood samples were

drawn from frogs immediately after capture and then at time intervals afterwards while being

contained in covered buckets holding a small amount of lake water. No significant changes were

detected in concentrations of plasma E2, T, and IGF- I over the 24 h period (Fig. 3-2). For

consistency in all subsequent procedures, blood and tissue samples were collected from frogs

within 12 h of capture. All animal procedures were performed in accordance with regulations

specified by University of Florida, Institute of Animal Care and Use Committee (Permit #Z095)

and a valid freshwater fishing license issued to T.R. Barbeau during the years of 2002 and 2003

as required by the State of Florida.

The frogs were anesthetized with MS-222 (1.5% 3-aminobenzoic acid ethyl-ether,

Aquatic Ecosystems, Orlando, FL), snout-vent length (SVL) and body mass were recorded, and

blood samples were obtained via cardiac puncture with heparinized syringe and needle. Blood

samples were centrifuged and resultant plasma frozen (-70'C) for E2, T, and IGF- 1

radioimmunoassay (RIA) analyses. Frogs were then euthanized by dissection through the spinal

cord followed by pithing.

The gonadal-fat bodies, liver, ovaries, and oviducts were removed from each frog and

weighed. Fat bodies were examined because they are an important energy reservoir that can be

metabolized to provide energy for growth of reproductive tissues before (and throughout) the

reproductive period. The liver was examined because it is the primary site for synthesis of plasma

IGF-1, vitellogenin, and other substances vital for reproduction in oviparous ectotherms (Crain et

al. 1995; Guillette et al., 1996). Ovarian maturation was categorized as either regressed (stage 1),

yellow (stage 2), black (stage 3), or mature "black and white" (stage 4) based predominantly on

the stages of follicular development described by Ligas (1960). Briefly, regressed ovaries were









small (< 0.75 mm diameter), yellow, and contained no visible follicles. Yellow ovaries were also

small but contained yellow follicles up to 0.75 mm in diameter. Black ovaries were medium to

large and contained mostly black follicles 1.0 1.25 mm in diameter. Black ovaries can mature

within a relatively brief time to stage 4 ovaries. Lastly, mature ovaries were large, composed of

highly polarized follicles 1.25 2.0 mm in diameter, and had a sharp delineation of light and dark

colors indicating a vegetal and animal hemispheres. Mature ovaries contained oocytes ready for

ovulation and fertilization (Fig. 3-3).

Small cross-sections of the ampulla region of the oviducts were fixed in 4%

paraformaldehyde (4C; 48 h) followed by rinse and storage in 75% ethanol for subsequent

histological analyses. The ampulla region, or middle portion of the oviduct, was examined

because it was the longest, most convoluted, and most visually distinct region (Wake and

Dickie, 1998). The ampulla region contains more glands and has a greater secretory activity than

other oviductal regions. The oviduct samples were dehydrated in a graded series of ethanol

changes, embedded in paraffin, serially cross-sectioned on a rotary microtome (7 Rm), stained

with modified Masson's staining procedure, and examined using light microscopy. To ascertain

oviductal proliferation, an ocular micrometer was used to make 10 morphological measurements

on 5 tissue sections, for a total of 50 measures per frog. The following oviductal parameters were

measured: epithelial cell height, endometrial thickness, endometrial gland height, and endometrial

gland width. Gland height and width measurements were used to calculate gland surface area

(4im2).

Steroid Radioimmunoassay (RIA) Biochemical Validation

Validation samples were obtained by creating plasma pools using aliquots from

individual frogs collected. Two methods were used to validate the E2 and T RIA: internal

standards and plasma dilutions. One half of the plasma pool, for use with internal standards, was

mixed with Norit charcoal (10 mL plasma to I g charcoal ratio; 4C; 24 h) to strip steroid









hormones from the plasma. The solution was then centrifuged (3000 rpm; 1200xG; 45 min) and

the resultant supernatant decanted. Separate, duplicate aliquots of stripped plasma (25 pL) were

added to tubes and spiked with 100 pL of assay buffer containing 1.56, 3.13, 6.25, 12.5, 25, 50,

100,200, 400, 800 pg E2 or T hormone. These tubes were extracted twice with ethyl-ether, air-

dried, and reconstituted in 100 pL borate buffer (100 pL; 0.05 M; pH 8.0).

For plasma dilutions, 6.25, 12.5, 25, 50, and 100 jtL plasma was added to different tubes.

Appropriate volumes of borate buffer were added to bring the final sample volume of each tube

up to 200 jtL. Samples were extracted twice with ethyl-ether, air-dried, and reconstituted with

100 ptL borate buffer. Resultant samples for both internal standards and plasma dilutions were

examined by the RIA procedure described below.

Plasma extraction efficiencies were determined by adding 100 1L tritiated E2 and T

(15,000 cpm) to 100 giL of pooled plasma samples, extracting twice with ethyl-ether, air-drying,

and adding 500 jtL scintillation fluid to tubes, and reading samples on a Beckman LS 5801

scintillation counter to determine the tritiated hormone remaining. The extraction efficiencies for

E2 and T samples were 90.0% and 91.4%, respectively. Supernatant (500 j.L) was added to 5 mL

of scintillation fluid, and counted on a Beckman scintillation counter. Plasma validation samples

were run in one assay with intraassay variance for E2 and T averaging 1.53% and 1.23%,

respectively. Plasma interassay variance for E2 and T averaged 6.99% and 3.27%, respectively.

Steroid RIA Procedures

RIAs were performed for E2 and T on plasma samples collected before surgery and after

treatments. For E2 samples, 25 p.L of plasma was used and for T samples, 6.25 gtL of plasma was

used. For T RIA, 50 ptL of plasma samples were diluted with 200 pL of borate buffer, and 25 pL

of this dilution (6.25 gL plasma equivalents) were used as samples in the RIA. These volumes

were selected for analysis based on RIA volume determinations conducted on these samples

previously. Briefly, duplicates of plasma samples were extracted twice with ethyl-ether, air-dried,








and reconstituted in borate buffer. To each tube, bovine serum albumin (Fraction V; Fisher

Scientific) in 100 [tL of borate buffer was added to reduce nonspecific binding at a final

concentration of 0.15% for T and 0.19% for E2. Antibody (Endocrine Sciences) was then added in

200 jiL of borate buffer at a final concentration of 1:25,000 for T and 1:55,000 for E2. Finally,

radiolabeled steroid ([2,4,6,7,16,17-3H] 1713-estradiol at 1 mCi/mL; [1,2,6,7-3H] testosterone at I

mCi/ml; Amersham Int., Arlington Heights, IL) was added at 12,000 cpm per 100 JLL for a final

assay volume of 500 gtL. Interassay variance tubes were similarly prepared from 2 separate

plasma pools for E2 and T. Standards for both E2 and T were prepared in duplicate at 0, 1.56, 3.13,

6.25, 12.5, 25, 50, 100, 200, 400, and 800 pg/tube. Assay tubes were vortexed for 1 min and

incubated at 4C overnight.

Bound-free separation was performed by adding 500 gL of a mixture of 5% charcoal to

0.5% dextran, pulse-vortexing, and centrifuging tubes (1500g, 4C, 30 min). Supernatant (500

jiL) was added to 3 mL of scintillation fluid, and counted on a Beckman scintillation counter.

Plasma samples were run in 3 assays with intraassay variance for E2 and T averaging 3.35% and

4.99%, respectively. Plasma interassay variance for E2 and T averaged 3.97% and 6.99%,

respectively.

Insulin-like Growth Factor-1 (IGF-1) RIA Biochemical Validation

Pooled plasma samples (200 gL) were extracted in polypropylene tubes with acid-ethanol

(12.5% 2 N HCI, 87.5% ethanol; 400 giL) to dissociate IGF binding proteins from the IGF-1

molecules and to precipitate globular proteins as per Daughaday et al. (1980) and Crain et al.

(1995). After 30 min incubation (23C) and 10 min centrifugation (2500xG; 4C), the supernatant

was aliquoted to produce plasma equivalents of 12.5, 25, 50, 100, and 200 iL. Volume of the

plasma dilutions was brought to 200 gtL with acid-ethanol before air-drying. Plasma dilutions

were compared with 0, 39, 156, 313, 625, 1000, 1250, 2500 pg of human recombinant IGF-1








standard (National Hormone and Pituitary Program, Torrance, CA 90509). Validation samples

were examined by IGF RIA procedures as described for experimental sample analyses below.

Plasma extraction efficiencies were determined by adding 100 IL iodinated IGF-1

(15,000 cpm) to 100 ;.L of pooled plasma samples, extracting with acid-ethanol, air-drying, and

reading samples on a Beckman 5500B gamma counter to determine the iodinated hormone

remaining. The extraction efficiency of plasma was 78.0% and all sample concentrations were

corrected for this loss. Validation of plasma dilutions was accomplished in one assay having an

intraassay variance of 2.3%. Internal standards and plasma dilutions were parallel to the standard

curve for E2 (ANCOVA; F = 0.24, P = 0.63 and F = 2.89, P = 0.15, Fig. 3-4A), and T RIA

(ANCOVA; F = 0.001, P = 0.99 and F = 0.013, P = 0.92, Fig. 3-4B).

IGF-1 RIA Procedures

IGF-1 RIA was performed as described by Crain et al. (1995) and Guillette et al. (1994).

The National Hormone and Pituitary Program (Torrance, CA 90509) supplied human

recombinant IGF- 1 standard (9.76 to 2500 pg/tube) and human IGF- 1 antisera (Lot #

AFP4892898, 1:400,000 final dilution). The antiserum had less than 1.0% cross-reactivity with

human IGF-II. Amersham International (Arlington Heights, IL) supplied iodinated IGF- I label

(IGF-1"25 sp act 2000 Ci/mmol; 16,000 cpm/tube) and Amerlex-M donkey anti-rabbit secondary

antibody (code RPN510, 500 IiL/tube). Buffer reagents were purchased from Fisher Chemical

Co. (Pittsburgh, PA). Briefly, 20 p.L plasma samples were aliquoted into polypropylene tubes,

extracted with 400uL acid-ethanol, and incubated 30 min prior to centrifugation (2500xG; 40C;

10 min). For each sample, supernatant (100 p1) was pipetted into duplicate polypropylene tubes

and air-dried. IGF- I standards were prepared in duplicate with 100 gtL of known concentrations

of human recombinant IGF- 1 standard (ranging from 9 2500 pg/tube), and 300 L RIA buffer

(200 mg/L protamine sulfate, 4.14 g/L sodium phosphate monobasic, 0.05% TWEEN 20, 0.02%

sodium azide, 3.72 g/L EDTA) added to each tube. Air-dried samples were reconstituted with 350








jiL RIA buffer and vortexed. To each sample was added 50 pgL IGF-1 antibody (human IGF-1

antisera, UB3-189) at a 1:10,000 final dilution. After adding 100 p.L of iodinated IGF-1 label

(I12-IGF-1), with -15,000 CPM, samples were vortexed and incubated (40 C) overnight. Bound-

free separation of IGF-l was accomplished by incubating samples for 10 min with 500 giL of

secondary antibody (Amerlex-M donkey anti-rabbit secondary antibody, code RPN.5 10 obtained

from Amersham International) at a final dilution of 1:10,000. Sample tubes were centrifuged

(2500xG; 40C; 10 min) to separate the secondary antibody, which is bound to the primary

antibody and ligand. The supernatant was decanted and the pellet counted on a Beckman 5500B

gamma counter. Plasma samples were run in 3 assays having an average intraassay variance of

3.65% and an interassay variance of 4.63%. Plasma dilutions were parallel to the standard curve

for IGF-1 RIA (ANCOVA; F = 0.67, P = 0.43; Fig. 3-4C).

Statistics

Tissue mass is typically highly correlated to body mass; thus, tissue weights were

compared among months using ANCOVA, with body mass as a covariate, followed by Fishers

Protected LSD post hoc. Data were presented as adjusted mean mass (mg) SEM. Pair-wise

monthly comparisons, of mature and immature ovary stages, was performed with non-parametric

chi-square analyses. Concentrations of E2, T, and IGF- I were estimated from raw data using the

commercially available Microplate Manager software (Microplate Manager III, BioRad

Laboratories, Inc., Hercules, CA, 1988). For RIA validation of pooled plasma dilutions and

internal standards, hormone concentrations were log 1 0-transformed prior to testing for

homogeneity of slopes with standard curves by ANCOVA. Hormone concentrations of E2, T, and

IGF-1 were compared among months with one-way ANOVA followed by SNK post hoc

contrasts. Tamhane post-hoc contrasts were used where variances were unequal among months

for IGF-1 concentrations. The relationships between plasma hormones, tissue weights, air and

water temperature, and rainfall were tested using Pearson's correlation analysis. Statistical









analyses were performed using SPSS software (v. 10, SPSS Inc., Chicago, IL, 1999) with t =

0.05.

Results

Seasonal Environmental Parameters

Elevated air temperatures on Orange Lake between March and July of both years

overlapped with the reproductive period of R. grylio, as determined by patterns of peak plasma E2

and T concentrations described below. Conversely, decreased air temperatures overlapped with

the non-reproductive period between November and February (Fig. 3-5). High levels of

precipitation between June and September of 2002 overlapped the reproductive period season but

rainfall fluctuated considerably throughout 2003. Water temperature, pH, and nitrate and nitrite

ion concentrations were recorded between December of 2002 and July of 2003. Water

temperature was low between December and January, and showed a steady increase in February

that continued through the 2003 reproductive season (Fig. 3-5). Water pH between December and

May ranged from 6.5 to 6.8 and between June and July ranged from 5.7 to 6.0. Aquatic nitrate

and nitrate concentrations remained below 1 mg/L throughout the 2003 season. Generally, peak

reproductive condition, as determined by reproductive tissue weights and plasma E2 and T

concentrations, was considered to occur between April and July of 2002 and between March and

May of 2003, indicating that reproductive condition in R. grylio occurred during different months

over the 15 month study.

Seasonal Tissue Mass and Ovarian Maturation

Fat body weights exhibited seasonal variation with the greatest weights occurring during

June of 2002 and during January and March of 2003. The lowest fat body weights occurred

between July and December of 2002 and between April and July of 2003 (Fig. 3-6A). Liver

weights, which varied comparatively less with season, were greatest in April and March, and

lowest between September and December of 2002 (Fig. 3-6B). Oviductal weights were greatest

between April and July of 2002 and in May of 2003, whereas lowest weights occurred between








August of 2002 and March of 2003. Oviductal weights were also low between June and July of

2003 (Fig. 3-6C). Ovarian weights (GSI) were greatest in June of 2002, intermediate in May of

2002 and between March and May of 2003, and were lowest in April and between July and

December of 2002, in addition to in June and July of 2003 (Fig. 3-6D).

A distinct seasonal pattern of ovarian maturation stages was observed in R. grylio (Fig. 3-

3, 3-7). Frogs with black ovaries were considered to be in reproductive condition. Thus, frogs

having either black or mature ovaries were considered reproductively mature for analyses.

Conversely, frogs having either regressed or yellow ovaries, indicative of immature ovarian

follicles, were considered reproductively immature for analyses. A greater percentage of frogs

collected during the reproductive period had reproductively mature ovaries (Fig. 3-10). On

average, approximately 80% of the females examined during the reproductive period had

reproductively mature ovaries. In contrast, between 50% 80% of frogs collected during the non-

reproductive period (August December) had reproductively immature ovaries (Fig. 3-7) but not

all females collected had regressed ovaries this period.

Seasonal Plasma Steroid and IGF-1 Concentrations

Plasma E2 concentrations were elevated during the reproductive period of both years

compared to the non-reproductive period. Additionally, E2 concentrations were higher in 2002

than in 2003 (Fig. 3-8A). Plasma T concentrations were elevated during the reproductive period

of both years compared to the non-reproductive period. However, the period of elevated T

concentrations was of slightly shorter duration in 2002 than in 2003 (Fig. 3-8B). Plasma IGF-1

concentrations exhibited a pattern opposite that of T between reproductive periods. Plasma IGF- 1

concentrations were increased during both reproductive period compared to the non-reproductive

period. However, IGF- 1 concentrations were elevated for more months in 2003 than in 2002

(Figure 3-8C). The variation in environmental factors associated with variation in plasma

hormone concentrations between the two reproductive periods indicate that reproductive

physiology of R. grylio is influenced by environmental factors.








Throughout the season, plasma E2 concentrations were comparatively lower than plasma

concentrations of T and IGF-1. Plasma E2 and IGF- 1 concentrations peaked during similar

months but plasma E2 concentrations declined precipitously after the reproductive period whereas

plasma IGF- 1 declined less sharply and remained elevated slightly longer. A peak in plasma T

concentration occurred slightly prior to increases in plasma concentrations of E2 and IGF- 1 (Fig.

3-9).

Peaks in ovarian and oviductal weights generally corresponded to elevated plasma T and

E2 concentrations, but they paralleled plasma T concentrations more closely (Fig. 3-15). During

the non-reproductive period, plasma steroids, plasma IGF-1, and reproductive tissue weights were

lower than in the reproductive period. Plasma IGF-1 concentrations appeared to peak in the latter

months of the reproductive period after steroid concentrations and the weights of reproductive

tissues began to decline (Fig. 3-10). Liver and fat body weights increased before, or in association

with, peaks in plasma T concentrations and much earlier than peaks in E2 and IGF-1

concentrations (Fig. 3-11).

Correlations: Plasma Steroids, Tissue Mass, and Environmental Parameters

For some months, the sample number of frogs collected was small; thus, analyses were

focused on correlations among means for all months. A strong positive correlation was detected

between plasma E2 and T concentrations (r2 = 0.67; P = 0.006) but not between concentrations of

E2 and IGF- 1 and not between IGF- I and T. Plasma E2 concentration correlated strongly to

oviductal weights (r2 = 0.84; P < 0.0001), to ovarian weights (r2 = 0.51; P < 0.05), and to water

temperature (r2 = 0.71; P = 0.05). Plasma T concentrations correlated strongly with ovarian (r2

0.85; P < 0.0001) and oviductal (r2 = 0.76; P = 0.001) weights but not to environmental

parameters. Plasma IGF-1 concentrations were negatively correlated to air temperature (r2 = 0.55;

P = 0.03) and positively correlated with fat body weights (r2 = 0.62; P = 0.02) but not correlated

to other parameters. Additionally, correlations were detected between ovarian and oviduct (r2 =

0.59; P = 0.02), or fat body masses (r2 = 0.57; P = 0.03), and between oviductal and liver weights








(r2 = 0.71; P = 0.003). Finally, fat body weight was correlated with water temperature (r2 = 0.75;

P = 0.03; Table 3-2).

Discussion

Relatively few studies have described a pattern of seasonal changes in the reproductive

tissues obtained from wild populations of ranid frogs (Ligas, 1960; Licht et al., 1983; Kim et al.,

1998). Previous studies of reproductive cyclicity in R grylio have been limited to ovarian

maturation, male calling behavior, sexual dimorphism, and observations of amplexus (Lygas,

1960; Lamb, 1983; Wood et al., 1998). Licht et al. (1983) reported that ovarian and oviductal

weights in R. catesbeiana increased during the reproductive season, between May and July,

declined sharply in August, and remained reduced through October in a population from central

California.

In R. grylio from Orange Lake, increased ovarian and oviductal weights and plasma

steroid concentrations clearly define the months during which peak reproductive condition

occurred during this study. Analysis of ovarian maturation stages revealed that the largest

percentage of frogs had mature ovaries during the reproductive period between April and July of

2002 and between March and July in 2003. During the months of the non-reproductive period, a

greater percentage of frogs had regressed ovaries. A similar pattern of ovarian maturation was

described for RI grylio in the Okefenokee Swamp of Georgia and in the Everglades of South

Florida (Ligas, 1960; Lamb, 1983).

The reproductive period occurred during slightly different months over the 15 month

study, lasting between April and July of 2002 but only between March and June of 2003. Our

data indicate that reproductive condition of R. grylio, as in other amphibians studied previously,

is responsive to changing environmental conditions. The reproductive period generally

overlapped with the months of high water and air temperature, and of high rainfall during both

years. Also, concentrations of plasma IGF- I increased with rising air temperature. Reproductive

tissue weights were greatest during periods of elevated air and water temperature, but no








significant correlations were observed between tissue weights and air or water temperature.

However, fat body weights decreased with increasing air temperature. These data lend support to

the theory that fat bodies are an energy reserve that are metabolized at the onset of warmer

weather to fuel rapid growth of reproductive tissues for breeding activity. Additionally, fat body

weights were positively correlated to ovary weights. Saidapur and Hoque (1995) reported similar

findings for Rana tigrina in India where decreasing fat body weights corresponded to increased

egg production, and both fat body weights and egg production were correlated to increasing air

temperature. In R. grylio from the Florida Everglades, reproductive activity is reportedly

suppressed during periods of low air and water temperature, and cease entirely during periods of

drought (Lygas 1960). The reproductive period of these frogs occurred primarily between March

and September and the non-reproductive period extends from October to February. Unlike frogs

from Orange Lake, R grylio from the Everglades appear to have an extended reproductive period

based on the mature ovarian tissue late into the season. This is supported by observations of

calling behavior by males, which also extends late into the season. However, it was unknown if

female R. grylio in the Everglades were actively mating and ovipositing eggs during these times,

so the extended reproductive period is speculative. This temporal variation in reproductive period

according to season could be attributed geographic differences, extended months of warm

temperatures in the summer, and milder temperatures during the winter in the Everglades

compared to north-central regions of Florida. Licht et al. (1983) attributed a similar temporal

variation in reproductive periods of bullfrogs according to geographic location. Also, in the

Okefinokee Swamp of Georgia, male R. grylio continue vocalizing between March and

September but peak reproductive condition of females occurs during June and July (Wright,

1932; Wright and Wright, 1949). These studies indicate that reproductive periods for R. grylio are

associated with localized environmental changes and also with geographic location.

The increase in relative liver weights of R. grylio,just prior to the onset of the

reproductive season, is indicative of increased hepatic biochemical or secretory activity. The liver









synthesizes many proteins that regulate metabolism, growth, reproduction, and development. One

of these proteins, vitellogenin, is a precursor of egg yolk in oviparous ectotherms and provides

valuable nutritive and energetic support for developing embryos (Carnevali et al., 1995; Sumpter

and Jobling, 1995; Guillette et al., 1996; Palmer and Guillette, 1998). Estrogen produced by

mature ovaries stimulates vitellogenesis in the liver. Vitellogenin is a yolk precursor protein that

is transported through the plasma to the ovaries where it accumulates within developing ova

(Licht, 1979)(Palmer et al., 1998; Sumpter and Jobling, 1995). In female alligators vitellogenesis

is accompanied by an elevation of plasma IGF-1 concentrations during the reproductive period.

IGF-1 has also been detected in the egg albumin of birds and reptiles, suggesting that this

hormone plays a role in embryonic growth and development (Cox and Guillette, 1993; Guillette

et al., 1996). In R. grylio, liver weights were elevated before peaks in plasma IGF-I

concentrations, and might reflect hepatic synthesis of proteins that function in reproduction

(Palmer et al., 1998; LeRoith et al., 2001 b). In oviparous ectotherms, plasma IGF-1 is also

influenced by nutritional status and feeding activity (Crain et al., 1995). In this study, plasma

IGF-1 concentrations were negatively correlated to decreasing fat body weights and positively

correlated to increasing air temperature. Accordingly, in R grylio, the correlation between plasma

IGF-1 concentrations and fat body weights might reflect metabolism of stored fat (during the

warmer months of the spring and summer months) to provide energy for growth of reproductive

tissues. In contrast, decreased concentrations of plasma IGF- 1 during the non-reproductive period

might reflect a decline in feeding behavior and in fat metabolism in female R. grylio.

Seasonal changes in concentrations of plasma steroids and IGF-1, in association with

changes in reproductive tissues, are largely undescribed for anurans. Seasonal changes in

reproductive tissues, and in concentrations of plasma steroids and plasma gonadotropins had been

described in ranid frogs from temperate North American and in India (Licht et al. 1983, Kim et

al., 1998). In California R. catesbeiana, plasma E2 and T concentrations generally peaked

between April and June; a pattern similar to that shown for plasma steroids in R. grylio. Plasma








steroid concentrations in R. grylio were most similar (1-4 ng/mL for E2 and 20-80 ng/mL for T) to

those measured in bullfrogs within 12 h of capture (Licht et al., 1983). Although plasma E2 and T

concentrations in R. grylio were positively correlated to each other, only plasma T concentrations

were correlated to ovarian and oviductal weights. This observation conflicts with findings in

mammals but indicates it might be common among non-mammalian vertebrates. In ectotherms,

androgens might play an important role in regulating reproductive condition. Amphibian ovarian

follicles synthesize and secrete large quantities of androgens during ovarian maturation (Fortune

and Tsang, 1981; Fortune, J.E. 1983; Lutz et al., 2001). Androgens might be aromatized to

estrogens in peripheral tissues such as the brain, fat and skin (Follett and Redshaw, 1968). The

oviduct might also be a site of peripheral aromatization of androgens and be a target for androgen

activity. The oviduct of oviparous species synthesizes huge quantities of protein (perhaps in

response to androgen stimulation) for use as secondary or tertiary egg coatings such as in anuran

egg jellies (Maack et al., 1985; Olsen and Chandler, 1999; Arranz and Cabada, 2000; Jesu-Anter

and Carroll, 2001). Similar to R. grylio, female R. catesbeiana also exhibited greater T than E2

plasma concentrations indicating that this pattern might be prevalent among ranids (Licht et al.,

1983). Rana grylio exhibited peak ovarian and oviductal weights during similar months as

reported for bullfrogs (Licht et al., 1983). However, plasma steroid concentrations in R. grylio did

not decrease significantly 24 h after capture as reported for R. catesbeiana (Licht et al., 1983). In

R. catesbeiana, increases in ovarian and oviductal mass closely paralleled increases in plasma

gonadotropins and steroids (Licht et al., 1983). Plasma LH and FSH were not measured in R.

grylio and it remains unknown whether plasma gonadotropins increased before elevations in

plasma steroid concentrations or tissue mass. In future studies, it would be valuable to examine

changes in plasma gonadotropins with respect to steroids to better understand the reproductive

cycle of R grylio.

Before this study on R. grylio, seasonal changes in plasma IGF- 1 concentration were

reported for only one other anuran species, Bufo woodhousei (Pancak-Roessler and Lee, 1990).








Plasma IGF-1 concentrations in B. woodhousei peaked (1 ng/ml) in July and declined sharply

thereafter. In R. grylio, plasma IGF- 1 concentrations peaked between May and July and declined

after August. In R. grylio a peak in plasma IGF-1 concentrations occurred later in the season

compared to Bufo woodhousei and is likely due to several factors including geographical

variation, interspecific differences, and differences in IGF-1 RIA methods.

In reptiles, seasonal changes in plasma IGF- 1 concentrations have been described for

alligators and turtles. In loggerhead sea turtles, elevated plasma IGF-1 concentrations occurred

between April and June and were associated with reproductive activity and increased feeding

behavior of female turtles during these months (Crain et al., 1995). Guillette et al. (1996)

examined reproductive tissues, and plasma steroids and plasma IGF-1 concentrations, and their

respective associations with reproductive condition in alligators. In female alligators, plasma

IGF- 1 concentrations increased in June and were associated with gravidity. Elevated plasma E2

and P4 concentrations were associated with peak vitellogenesis, and also preceded gravidity and

peaks in plasma steroid concentrations. Seasonal patterns of plasma IGF-1 concentrations were

not examined simultaneously with changes in plasma steroids concentrations; therefore, it is

unknown how these hormones change (with respect to each other) seasonally in alligators

(Guillette et al., 1996). In a separate study, alligators collected from the same locality exhibited a

peak in plasma E2, T, and P4 concentrations in May (Guillette et al., 1997). Thus, alligators are

similar to R. grylio in that elevated plasma steroid concentrations precede peaks in plasma IGF-1.

In mammals, IGF-1 expression is associated various aspects of reproduction including

ovarian maturation, follicular atresia, selection of dominant follicles, and regulation of gonadal

steroidogenesis. Increasing concentrations of IGF-1 can be synthesized in reproductive tissues or

the in liver, and can be transported directly to offspring in utero. In contrast to mammals,

oviparous animals must provide growth-promoting substances to eggs prior to oviposition

(Palmer and Guillette, 1991; Guillette et al., 1996). Accordingly, IGF-1 has been detected in the





55


yolks of chicken eggs, the albumen of alligator eggs, and the oviductal glands of geckos and

alligators (Scavo et al., 1989; Guillette and Williams, 1991; Cox and Guillette, 1993; Cox, 1994).

In conclusion, this study provides the first evidence that IGF- 1 is present in the plasma of

R. grylio. Plasma IGF-1 concentrations were correlated with several environmental factors and

exhibited a clear pattern of change with reproductive period, and with reproductive steroid

concentrations and weights of reproductive tissues. Although the role of IGF- I in anurans

requires further study, this study has provided valuable information for understanding the

association of IGF-1 with reproductive physiology in R. grylio.









Table 3-1. Comparison of plasma insulin-like growth factor- I (IGF-1) concentrations among
mammalian, avian, reptilian, and amphibian species.
IGF-I
(nG/L I Reference
(ng/mL)
Mammalian
Rat
Juvenile males 574.0 a
Human
Adult females 261.0 a
(Reproductive status unknown)
Cow
Adult females 182.0 a
(Reproductive status unknown)
Avian
Chicken
Juveniles (8-week) 42.0 a
Reptilian
Red-eared slider turtle
Juvenile males 17.0 a
Loggerhead sea turtle
Reproductive 7.5 b
Non-reproductive 3.0 b
American alligator
Reproductive 16.0 c
Non-reproductive 5.0 c
Amphibian
African Clawed frog
Non-reproductive females 3.0 d
American toad
Reproductive males 4.0 d
Non-reproductive males 0.5
Bullfrog
Non-reproductive female 1.0 d
Marine toad
Non-reproductive female 1.0 d
Pig frog
Reproductive females 22.0 This Study
Non-reproductive females 10.0 This Study
a Daughaday et al., 1985
b Crain et al., 1995
c Guillette et al., 1996
d Pancak-Roessler and Lee, 1990
Note For all studies shown, plasma IGF-1 was measured, after acid-extraction
of IGF- 1 binding proteins, by radioimmunoassay.









Table 3-2. Correlations among body mass, snout vent length (SVL), hormone concentrations,
tissues weights of Rana grylio, and environmental parameters.
Pearson Correlation P-value R2 Relationship


Body Mass and E2
Body Mass and T
Body Mass and IGF- 1
Snout-Vent-Length and E2
Snout-Vent-Length and T
Snout-Vent-Length and IGF-1


Plasma E2 and T
Plasma E2 and IGF-1
Plasma E2 and Ovary Weight
Plasma E2 and Oviduct Weight
Plasma E2 and Liver Weight
Plasma E2 and Fat Body Weight
Plasma E2 and Air Temperature
Plasma E2 and Water Temperature
Plasma E2 and Rainfall

Plasma T and IGF-1
Plasma T and Ovary Weight
Plasma T and Oviduct Weight
Plasma T and Liver Weight
Plasma T and Fat Body Weight
Plasma T and Air Temperature
Plasma T and Water Temperature
Plasma T and Rainfall

Plasma IGF- 1 and Ovary Weight
Plasma IGF- 1 and Oviduct Weight
Plasma IGF-1 and Liver Weight
Plasma IGF- 1 and Fat Body Weight
Plasma IGF-1 and Air Temperature
Plasma IGF- I and Water Temperature
Plasma IGF- 1 and Rainfall

Ovary and Oviduct Weight
Ovary and Liver Weight
Ovary and Fat Body Weight
Oviduct and Liver Weight
Oviduct and Fat Body Weight
Liver and Fat Body Weight


0.27
0.06
0.94
0.31
0.09
0.58


0.006 **
0.28
0.05
< 0.0001 **
0.18
0.88
0.07
0.05
0.77

0.64
< 0.0001 **
0.001 **
0.06
0.16
0.20
0.09
0.25

0.81
0.67
0.35
0.02 *
0.03 *
0.12
0.48

0.02 *
0.25
0.03 *
0.003 **
0.27
0.13


0.67

0.51
0.84



0.71



0.85
0.80










0.62
0.55



0.59

0.57
0.71








Table 3-2. Continued.
Pearson Correlation P-value R2 Relationship

Air Temperature and Ovary Weight 0.84
Air Temperature and Oviduct Weight 0.37
Air Temperature and Liver Weight 0.70
Air Temperature and Fat Body Weight 0.08
Water Temperature and Ovary Weight 0.45
Water Temperature and Oviduct Weight 0.07
Water Temperature and Liver Weight 0.59
Water Temperature Fat Body Weight 0.03 0.75
Rainfall and Ovary Weight 0.55
Rainfall and Oviduct Weight 0.88
Rainfall and Liver Weight 0.37
Rainfall and Fat Body Weight 0.84
Significant
** Highly Significant
+ positive correlation
negative correlation












A
N


3km


Figure


%-,uto tiUt sute ior itana grylo, indicated by asterisk, on Orange Lake in Alachua
county, Florida (Latitude 29027.853'N, Longitude 82011.380'W) between April,
2002 and July, 2003. Image created by T. Barbeau.









20000 A E2
18000 mT i60000
18000 M UT
0 IGF-1 50
'16000 50000 ,

14000
f 14000
1200 I 400002
"12000

10000 130000
+008000 '
6000 I20000
4000
20005 5 10000
2 0"
2000 OO
5 5 5 4
.. .. + -- .. I 0
0 6 12 24
Time (h)
Figure 3-2. Plasma 17f-estradiol (E2), testosterone (T), and insulin-like growth factor-I (IGF-1)
concentrations in Rana grylio at 0, 6, 12, and 24 h post-capture. Plasma samples for
each time interval were collected from different frogs. Data presented as means +
SEM. Letters within axes represent sample size at each time interval. No significant
differences detected among time intervals for each hormone (ANOVA; E2 P = 0.40;T
P = 0.83; IGF-1 P = 0.42).










Regressed



Yellow





Black







Mature






2 cm

Figure 3-3. Staging of Rana grylio ovaries in progression from least to most mature stages. A.
regressed (stage 1), B. yellow (stage 2), C. black (stage 3), and D. black and white
(stage 4) ovaries.









A 100Q
80
60
~ 40!
~ 20-

1


.0


0*


0 Standard
* Plasma Dilutions
0 Internal Standards


0 0
a-m


100


1000


-3-


10000


Estradiol (pg)


0 0


0 Standard
Plasma Dilutions
06 Internal Standards


0.


100


1000


10000


Testosterone (pg)


0 0
0


0 Standard
* Plasma Dilutions


0


00


100
IGF-I (pg)


1000


10000


Figure 3-4. Biochemical validation of Rana grylio plasma for RIA. A. For 1713-estradiol RIA
internal standard and plasma dilution curves were parallel to the standard curve
(ANCOVA; F = 0.24, P = 0.63 and F = 2.89, P = 0.15). B. For testosterone RIA
internal standard and plasma dilution curves were parallel to the standard curve
(ANCOVA; F = 0.00 1, P = 0.99 and F = 0.01, P = 0.92). C. For insulin-like growth
factor-I (IGF- 1) RIA the plasma dilution curve was parallel to the standard curve
(ANCOVA; F = 1.05, P = 0.33).


B 100
80
0
S60

X 40
~20


100
80
60
40
20
0










E Air High Air Low 0 Water A Rainfall
6o0 30

Vso~~~ **l***25
Q 50-
20
24o
4) 15

S30
4 10
20' A A A
0A A5
10 A A A A A tO


... 4 ----+ ..' .. -- f ... 1-+ -- --- : .;.. -
A M J A S O N D J F M A M J J
2002 2003
Month
Figure 3-5. Monthly changes in rainfall, water temperature, and high and low air temperature at
collection site on Orange Lake, Florida. Data presented as means per month.










Non-Reproductive


Reproductive


ab


abc abc

C


1.0-

0.8-
A 0.6-
04-

0.2-

00-
B.
4.0

E 3.0


.2.0

r 1.0


0.0
C2.

.3.0
5
S2.5-
U
2.0

.~1.5-
61.0-
aI .


D. Reproductive Non-Reproductive Reproductive
2.5

ZV,-I a b
2,0
bT
11.5 b bed b b Tbcd
T' cd d bc d
1. c d
l c d
0.5
40 1.0i

0.0
A M J J A S O N D J F M A M J J
2002 2003
Month

Figure 3-6. Seasonal change in fat body weights in Rana grylio during the reproductive and non-
reproductive periods. Data presented as means SEM. Numbers within bars indicate
sample size and different letters above bars indicate significantly different means A.
for fat bodies (ANCOVA; F = 2.78, P = 0.002), B. for liver (ANCOVA; F = 4.90, P
< 0.001), C. for oviducts (ANCOVA; F = 1.55, P < 0.001), and D. ovaries
(ANCOVA; F = 4.27, P < 0.001).


Reproductive
a


Non-Reproductive Reproductive



ab

bed ab
d bcd
cd T bed bcd bed











m Yellow U Black IM Mature


ab ab a


A M
2002


b d d













J A S O


N D J F M
2003
Month


Figure 3-7. Seasonal changes in ovarian maturation stages of Rana grylio. Data presented as
percentage of frogs exhibiting immature, yellow, black, and mature ovary stages, out
of total frogs, from the total collected that month. Different letters above bars
indicate significantly different percentages as determined by Mann Whitney U
pairwise contrasts (P < 0.05).


100

80

60

,40

S20


A MJ


10 Immature


a b a b


d b b c












A. Reproductive Non-Reproductive Reproductive
5000 0
a a a


A M J J


A S O N D


FM A M J


Reproductive Non-Reproductive Reproductive
4, A a -- rN


45000
40000


30000
2 25000
2 20000 cd c






0
A M J J
C. Reproductive

40

35


d d d d
d d



A S O N D J F M A M J J
Non-Rqroductive Reproductive
a


Ca T c
cd C
cd cd cd cdd




A M J J A S O N D J F M A
2002 2003


M J J


Month
Figure 3-8. Seasonal change in plasma hormones in Rana grylio during the reproductive and non-
reproductive periods. Data presented as means SEM. Numbers within bars indicate
sample size while different letters above bars indicate significantly different means
for A. 1713-estradiol (ANOVA; P < 0.001), B. testosterone (ANOVA; P < 0.001), and
C. insulin-like growth factor-I (IGF-1, ANOVA; P < 0.001).











45000 -


40000 .


I 35000 -


Ipi 30000 -

o 25000


'20000-


15000

10000


5000


Reproductive Non- Renroductive Reproductive


# i+


A


U U


A M J J A
2002


o N


Month


J F M A M J
2003


Figure 3-9. Seasonal changes in plasma 1713-estradiol (E2), testosterone (T), and insulin-like
growth factor-i (IGF-1) in Rana grylio during the reproductive and non-reproductive
periods. Data presented as means SEM.


- 6000



-5000



-4000 w



3000 -



2000



1000










50000 -


6b 40000 -


-30000 -


o 20000


GEM 10000


0-

(-4
W-10000-


-20000


Reproductive

A
/A mA/


E 0


U 0

AA

U.


Non-Reproductive


A Ax


II~LJL


II


A M J J A S
2002


O N D J F M A M J J
2002
Month


Figure 3-10. Seasonal changes in plasma 1713-estradiol (E2), testosterone (T), and insulin-like
growth factor- I (IGF- 1), and of ovary and oviduct weights in Rana grylio during the
reproductive and non-reproductive periods. Steroid data presented as means and
tissue data presented as means SEM.


Reproductive


A
A

Emi
*


.i


- 175
(j2

- 150


- 125 ,


- 100 f



- 75 I
0
-50 "


-25


-0


C im. .


I !










50000 Reproductive Non-Reproductive Reproductive 60

1i 40000 A -U

30000-A
45
S 30000 -


H 20000 -
(( m maa 30
OU


10000 0

0 AO

S15 t
00




20000 It i"




A M J J A S N D J F M A M J J
2002 Month 2003

Figure 3-11. Relative seasonal changes in plasma 173-estradiol (E2), testosterone (T), insulin-like
growth factor-I (IGF- 1), and of liver and fat body weights Rana grylio during the
reproductive and non-reproductive periods. Steroid data presented as means and
tissue data presented as means SEM.














CHAPTER 4
THE EFFECTS OF INSULIN-LIKE GROWTH FACTOR-I AND ESTRADIOL IMPLANTS
(IN VIVO) ON OVIDUCT MORPHOLOGY, AND ON PLASMA HORMONES IN
BULLFROGS (Rana catesbeiana)


Introduction

The regulation of oviduct growth and function by endocrine hormones has been described

for mammals and some reptiles but comparatively little is known for amphibians (Christiansen,

1973; Mead et al., 1981; Murphy and Ghahary, 1990; Cox and Guillette, 1993; Buhi et al., 1999;

Girling et al., 2000). Most studies of frog reproduction have focused on variation in plasma

concentrations of steroid hormones and ovarian maturation, with little attention to regulation of

oviductal structure (Licht et al., 1984; Wake and Dickie, 1998). For oviparous animals, the

oviduct is vital for reproduction because it synthesizes and secretes important substances that

nourish and encapsulate ovulated oocytes. The female bullfrog (Rana catesbeiana) can oviposit

as many as 40 to 80 thousand eggs at one breeding event (Norris, 1997). Without the provision of

oviductal secretions, oocytes could not be fertilized successfully nor could they develop into

normal embryos (Low et al., 1976; Buhi et al., 1997; Buhi et al., 1999; Olsen and Chandler,

1999). The amphibian oviduct includes four major structural and functional regions: the

infundibulum, the atrium, the ampulla, and the ovisac (Uribe et al., 1989). The infundibulum is

the anterior-most region of the oviduct and receives mature oocytes ovulated from the ovaries.

Distal to the infundibulum is the atrium a narrow aglandular region that precedes the ampulla.

The ampulla region is longest portion of the oviduct and contains numerous glands within the

endometrial layer (Wake and Dickie, 1998). The glands within the ampulla region are

biochemically active and secrete a variety of substances that are incorporated into mature oocytes

as they traverse the oviduct (Uribe et al., 1989). The last region of the oviduct is the ovisac or








uterus that leads to the cloaca. The narrow and aglandular ovisac is the final site from which

oocytes are deposited from the reproductive tract into the environment.

Oviductal growth occurs primarily in response to stimulation by elevated E2

concentrations of E2, of ovarian origin, and involves proliferation of epithelial and endometrial

cells (Christiansen, 1973; Mead et al., 1981; Cox, 1994). In amphibians, the major reproductive

steroids progesterone (P4), testosterone (T), and 17p-estradiol (E2), are produced and secreted by

the ovary, in response to pituitary follicle stimulating hormone (FSH) and luteinizing hormone

(LH) (Licht, 1979; Chapter 3). The principal steroid that regulates structure and function of the

oviduct is reported to be E2. In addition to E2, polypeptide growth factors have been shown to

elicit a growth response in the reptilian and mammalian oviduct (Cox and Guillette, 1994;

Stevenson et al., 1994; Tang et al., 1994; Richards et al., 1997). Growing evidence demonstrates

that autocrine and paracrine sources of epidermal growth factor (EGF) and insulin-like growth

factor-I (IGF- 1) are potent hormonal mitogens that mediate E2-induced oviduct growth. In

mammals, these growth factors induce oviduct growth in the absence of endogenous E2, and

induce an even greater growth in the presence of E2 compared to either hormone administered

alone (Nelson, 1991; Murphy and Murphy, 1994). These findings indicate a hormonal synergy

between E2 and IGF- I in the stimulation of oviduct growth.

Cox and Guillette (1994) reported that ovariectomized geckos exhibit oviductal growth in

response to EGF and IGF- 1 even in the absence of stimulation by endogenous E2. However,

neither EGF nor IGF- I stimulation induced oviduct growth similar to that observed with E2 alone.

Unfortunately, the effect of simultaneous treatment with E2 and growth factors on oviduct growth

was not examined in this study and it remains unknown if an E2 and IGF-1 synergy exists for

these animals.

In this study, I examined the effects of controlled doses of steroid hormone (E2), and

peptide hormones (IGF- 1 and EGF) on oviduct growth in adult, female bullfrogs (Rana

catesbeiana). The objective of this study was to determine whether the oviducts in R. catesbeiana









exhibited a growth response with exposure to E2 or to growth factors administered separately or

in combination. I predicted that ovariectomized R. catesbeiana treated with either E2 or growth

factors (EGF and IGF-1) would exhibit oviduct growth. Additionally, I predicted that oviduct

growth would be greater in frogs treated simultaneously with E2 and IGF- I than with either

hormone alone.

Materials and Methods

Adult female Rana catesbeiana (N = 65) were purchased (Charles D. Sullivan Co. Inc.,

TN). They were maintained under a 12-h diurnal light/dark cycle in 38 L tanks with 19 L of

static, dechlorinated water at 26C. They were fed crickets every other day throughout the

experiment. Frogs were randomly assigned to each of the following treatment groups: E2 (N= 10),

IGF-1 (N=10), EGF (N=10), E2 /IGF-1 (N=10), placebo (N-10), and sham (N=5). The use of

sham frogs is explained below. Weight and snout vent length (SVL) of frogs were recorded and

no significant difference was detected in mass (ANOVA; P = 0.84) or SVL (ANOVA; P = 0.85)

of frogs among treatment groups. Numbered stainless steel tags were applied to the webbing of

the hind foot of frogs for individual identification. Animals were maintained and experiments

were performed as approved by the Institute for Animal Care and Use Committee (IACUC

project #Z095).

Sham animals are frogs that were subjected to identical anesthesia and surgical

procedures that ovariectomized frogs were subjected to (explained below) with the exception that

the ovaries were not removed and they received no hormone treatment. Thus, sham frogs were

intact frogs that were included to account for an effect of the surgical procedure itself on

physiological responses of the frogs.

For this study, R. catesbeiana were chosen in lieu of X laevis and 1R grylio that were

examined in earlier chapters. Previous attempts to maintain wild-caught R. grylio in captivity

demonstrated that this species exhibited considerable stress and was considered inappropriate for

a long-term, surgical study. In contrast to R. grylio, R. catesbeiana were very adaptable to








captivity and exhibited less stress. Previous attempts to ovariectomize X laevis were largely

unsuccessful while R. catesbeiana responded optimally to the ovariectomy procedures with low

mortality and fast recovery. Additionally, R. catesbeiana are large-bodied frogs and it was easier

to collect blood samples of greater volume than in X laevis and R grylio. Lastly, R catesbeiana

is closely related to R. grylio and was considered a relevant and appropriate substitution for this

study.

Ovariectomy

After a 2-week acclimation period, frogs were anesthetized with MS-222 (1.5% 3-

aminobenzoic acid ethyl ether, Aquatic Ecosystems, Orlando, FL), and ovariectomy was

performed. The ovaries were removed to ensure that endogenous hormones did not conflict with

or obscure effects observed in response to experimental treatments. Conducting more than six

surgeries per day would have compromised my ability to carefully conduct surgeries and oversee

post-operative recovery of individuals. Thus, ovariectomy was performed each day, for 10

consecutive days, on one individual selected from each of the six treatment groups.

Under sterile conditions, a 2.5 cm right paramedial incision was made through the skin

and muscle layers into the abdominal cavity, and the left and right ovaries were excised through

this single incision. Hemostasis of the mesovarium (vascular tissue supporting the ovaries) was

accomplished by a series of double-ligatures of the vessels using 5-0, monofilament nylon suture

material (Fig. 4-1). The incision layers (peritoneal, muscle, and skin) were closed with a single

interrupted pattern using the same suture material. For each female, the mass of excised ovaries

was recorded and reproductive status of the female was determined by visual inspection of

ovarian follicle maturation according to Dumont (1971). This procedure was performed to

confirm that females were reproductively similar at the onset of the experiment to minimize

variation in responses to subsequent treatments.

Pre-ovariectomy blood samples were collected by cardiac puncture to determine whether

plasma E2, T, and IGF- I concentrations were similar among females at the start of the








experiment. No more than 1.0% of total blood volume estimated per body mass was taken from

frogs (Mader, 1996; Wright, 2001). Blood samples were stored in heparin vacutainer tubes,

centrifuged, and subsequent plasma was stored (-70'C) for radioimmunoassay (RIA) analyses. If

blood could not be collected within two cardiac punctures attempts were ceased to avoid potential

injury to the frog. Blood could not be sampled from all individuals prior to surgery; therefore,

sample sizes for pre-ovariectomy blood samples were as follows: E2 (N= 8), IGF-1 (N=9), EGF

(N=7), E2 /IGF- 1 (N=9), placebo (N=8), and sham (N=5).

Post-ovariectomy frogs were placed in recovery tanks containing benzalkonium chloride

(antibiotic) dissolved in 1 liter of water for 48 h. Afterwards, recovered frogs were returned to

their tanks. Frogs were allowed a 3-week recovery period during which the surgical sites were

closely monitored for signs of inflammation or infection. Although most frogs experienced no

post-operative complications, five frogs failed to recover from the ovariectomy. Therefore, the

final sample sizes for post-ovariectomy treatment groups were as follows: E2 (N= 10), IGF- 1

(N=10), EGF (N=8), E2/IGF-1 (N=9), placebo (N=8), and sham (N=5). No further losses

occurred, and these sample sizes were maintained for each treatment group for the remainder of

the study.

Hormone Implants

After the 3-week recovery period, all frogs exhibited complete healing of the

ovariectomy incision site (Fig. 4-2). Treatments were administered by surgical insertion of an

intra-abdominal, 21-day release treatment pellet (Innovative Research of America, Sarasota, FL)

containing either of the following dosages: E2 (420 jig), IGF-1 (10 jig), EGF (10 jig), E2/IGF-1

(420 jig E2 and 10 jig IGF- I), and placebo (10 jig vehicle pellet). Surgical procedures for the

treatment pellet implantation were similar to those described for ovariectomy with respect to

incision and abdominal closure; however, an 1.0 cm left paramedial incision was made. Pellets

were inserted into the abdominal cavity midway between the left and right oviducts (Fig. 4-3).








The E2 treatment served as a positive control, while the placebo treatment served as a negative

control. The EGF, IGF-1, and E2/IGF-I treatments were experimental. For simultaneous

treatment with E2/IGF-1, one pellet of each hormone was inserted 2.54 cm apart from each other

within the abdomen. The hormone concentrations administered were physiologically relevant and

chosen based on a literature review of similar studies in which treatments were given for

durations ranging from 7 20 days to elicit a tissue response (Redshaw et al., 1968; Follet and

Redshaw 1968; Fortune 1981; Cox 1994; Crain et al. 1995). After 18 days of treatment, the frogs

were euthanized and examined as described below.

Tissue Sampling

The response of R. catesbeiana to 18 days of treatment was determined by measuring the

following parameters: weights of tissues (liver and oviduct), oviductal growth (macroscopic and

microscopic), and plasma concentrations of E2, T, and IGF-1. After treatment, frogs were

anesthetized with MS-222, and blood samples were collected. Frogs were euthanized by

dissection through the spinal cord followed by pithing. Plasma samples (post-treatment) were

frozen (-70'C) for RIA analyses. The liver and oviducts were removed from each frogs and

weighed for comparison of wet tissue mass among treatment groups. Cross-sectional samples of

oviducts were fixed in 4% paraformaldehyde (4C; 48 h) followed by rinse and storage in 75%

ethanol for subsequent histological analyses. The oviducts were dehydrated in a graded series of

ethanol changes, embedded in paraffin, serially sectioned on a rotary microtome (7 gim), stained

with modified Masson's staining procedure, and examined microscopically. To evaluate oviductal

growth, an ocular micrometer was used to record 10 morphological measurements on 5 tissue

sections per frog (for a total of 50 measures) for each of the following oviductal parameters:

epithelial cell height, endometrial layer thickness, endometrial gland height, and endometrial

gland width. The gland height and width measurements were used to calculate cross-sectional

gland surface area.









Steroid Radioimmunoassay (RIA) Biochemical Validation

Validation samples were obtained by pooling plasma aliquots from each individual. Two

methods were used to biochemically validate the E2 and T RIA: internal standards and plasma

dilutions. One half of the plasma pool, for use with internal standards, was mixed with Norit

charcoal (10 mL plasma: I g charcoal; 4C; 24 h) to strip steroid hormones from the plasma. The

solution was then centrifuged (3000 rpm; 1200xG; 45 min) and the resultant supernatant

decanted. Separate aliquots of stripped plasma (25 pL) were added to 10 tubes and spiked with

100 gtL of 1.56, 3.13, 6.25, 12.5, 25, 50, 100, 200, 400, 800 pg cold E2 or T hormone. These tubes

were extracted twice with ethyl-ether, air-dried, and reconstituted in 100 ptL of borate buffer (100

p.L; 0.05 M; pH 8.0).

For plasma dilutions, 6.25, 12.5, 25, 50, 100, and 200 pI plasma was added to 6 tubes.

Appropriate volumes of borate buffer were added to each tube to bring the final sample volume to

200 jiL. Samples were extracted twice with ethyl-ether, air-dried, and reconstituted with 100 pL

of borate buffer. Resultant samples for both internal standards and plasma dilutions were

examined by the RIA procedure described below.

Plasma extraction efficiencies were determined by adding 100 j.L tritiated E2 and T

(15,000 cpm) to 100 gL of pooled plasma samples, twice extracting with ethyl-ether, air-drying,

adding 500 gL scintillation fluid to tubes, and reading samples on a Beckman LS 5801

scintillation counter to determine the tritiated hormone remaining. The extraction efficiencies for

E2 and T samples were 93.9% and 87.9%, respectively. Supernatant (500 jiL) was added to 5 mL

of scintillation fluid, and counted on a Beckman scintillation counter. Plasma intraassay variance

for E2 and T validation RIAs for averaged 1.53% and 1.23%, respectively. Plasma interassay

variance for E2 and T averaged 2.87% and 4.88%, respectively.









Steroid RIA Procedures

RIAs were performed for E2 and T on plasma samples collected both pre-ovariectomy

and post-treatment. For pre-ovariectomy E2 samples, 50 tL of plasma was used, and for post-

treatment samples, 50 gL of plasma was used for E2, E2/IGF-1, and sham samples and 300 gL

plasma used for IGF-1, EGF, and placebo samples. For pre-ovariectomy T samples, 30 ptL of

plasma was used while for post-treatment samples, 50 gtL of plasma was used for E2, E2/IGF-1,

and sham samples and 300 p.L plasma used for IGF-1, EGF, and placebo samples. These volumes

were selected for analysis based on RIA volume determinations conducted on these samples

previously. Briefly, duplicates of plasma samples were twice extracted with ethyl ether, air-dried,

and reconstituted in borate buffer. To each tube, bovine serum albumin (Fraction V; Fisher

Scientific) in 100 pL of borate buffer was added to reduce nonspecific binding at a final

concentration of 0.15% for T and 0.19% for E2. Antibody (Endocrine Sciences) was then added to

200 gtL of borate buffer for a final concentration of 1:25,000 for T and 1:55,000 for E2. Finally,

radiolabeled steroid ([2,4,6,7,16,17-3H] E2 at 1 mCi/mL; [1,2,6,7-3H] T at I mCi/mL; Amersham

Int., Arlington Heights, IL) was added at 12,000 cpm per 100 jtL for a final assay volume of 500

jiL. Interassay variance tubes were similarly prepared from two separate plasma pools for E2 and

T. Standards for both E2and T were prepared in duplicate at 0, 1.56, 3.13, 6.25, 12.5, 25, 50, 100,

200, 400, and 800 pg/tube. Assay tubes were vortexed for 1 min and incubated overnight at 4C.

Bound-free separation was performed by adding 500 ptL of a mixture of 5.0% charcoal to 0.5%

dextran, pulse-vortexing, and centrifuging tubes (1500g, 4C, 30 min). Supernatant (500 IiL) was

added to 5 mL of scintillation fluid, and counted on a Beckman scintillation counter. Plasma

intraassay variance for E2 and T averaged 2.87% and 4.93%, respectively. Plasma interassay

variance for E2 and T averaged 7.64% and 5.25%, respectively.








Insulin-Like Growth Factor-1 (IGF-1) RIA Biochemical Validation

From each treatment group, plasma (200 j.L) was pooled for validation, and was

extracted in polypropylene tubes with acid-ethanol (12.5% 2 N HCI, 87.5% ethanol; 800 gtL) to

dissociate IGF binding proteins from the IGF- 1 molecules and to precipitate globular proteins as

per Daughaday et al. (1980) and Crain et al. (1995). After 30 min incubation (room temperature)

and 1 0-min centrifugation (2500xG; 4C), the supernatant was aliquoted to produce plasma

equivalents of 12.5, 25, 50, 100, and 200 pL. Plasma dilution volumes were brought to 200 JtL

with acid-ethanol prior to air-drying. Plasma dilutions were compared with 0, 39, 156, 313, 625,

1000, 1250, 2500 pg of human recombinant IGF-1 standard (National Hormone and Pituitary

Program, Torrance, CA 90509). Validation samples were examined by IGF RIA procedures as

described for experimental sample analyses below. Plasma extraction efficiencies were

determined by adding 100 gL iodinated IGF-1 (15,000 cpm) to 100 ViL of pooled plasma

samples, extracting with acid-ethanol, air-drying, and reading samples on a Beckman 5500B

gamma counter to determine the iodinated hormone remaining. The extraction efficiency of

plasma was 77.0% and all sample concentrations were corrected for this loss. Validation of

plasma dilutions was accomplished in one assay having an intraassay variance of 2.27%.

IGF-1 RIA Procedures

IGF-1 RIA was performed as described by Crain et al. (1995) and Guillette et al. (1994).

The National Hormone and Pituitary Program (Torrance, CA 90509) supplied human

recombinant IGF- 1 standard (9.76 to 2500 pg/tube) and human IGF- 1 antisera (Lot #

AFP4892898, 1:400,000 final dilution). The antiserum had less than 1.0% cross-reactivity with

human IGF-II. lodinated IGF-1 label (IGF-1"25 sp act 2000 Ci/mmol; 16,000 cpm/tube) and

Amerlex-M donkey anti-rabbit secondary antibody (code RPN510, 500 giL/tube) were supplied

through from Amersham International (Arlington Heights, IL). Buffer reagents were purchased

from Fisher Chemical Co. (Pittsburgh, PA). Briefly, samples were aliquoted into polypropylene









tubes, extracted with 400 gL acid-ethanol, and incubated 30 min prior to centrifugation (2500xG;

4'C; 10 min). For each sample, supernatant (100 gL) was pipetted into duplicate polypropylene

tubes and air-dried. IGF- I standards were prepared in duplicate with 100 pL of known

concentrations of human recombinant IGF-1 standard (ranging from 9 2500 pg/tube), and 300

jiL RIA buffer (200 mg/L protamine sulfate, 4.14 g/L sodium phosphate monobasic, 0.05%

TWEEN 20, 0.02% sodium azide, 3.72 g/L EDTA) added to each tube. Air-dried samples were

reconstituted with 350 p.L RIA buffer and vortexed. To each sample was added 50 pL IGF-1

antibody (human IGF-l antisera, UB3-189; 1:10,000 final dilution). After adding 100 tL of

iodinated IGF-1 label (I12'-IGF-1; 15,000 cpm) samples were vortexed and incubated (4'C)

overnight. Separation of bound and free IGF- 1 was accomplished by incubating samples for 10

min with 500 jtL of secondary (20) antibody (Amerlex-M donkey anti-rabbit secondary antibody,

code RPN.5 10, Amersham International; 1: 10,000 final dilution). Sample tubes were centrifuged

(2500xG; 4C; 10 min) to separate the secondary antibody, which is bound to the primary

antibody and ligand. The supernatant was decanted and the pellet counted on a Beckman 5500B

gamma counter. Pre-ovariectomy and post-treatment plasma samples were run in two assays

having an average intraassay variance of 5.76% and an interassay variance of 4.19%.

Statistics

Wet tissue mass (mg) of liver, oviduct, and ovary were compared among treatment

groups using ANCOVA, with body mass as a covariate, followed by LSD post-hoc tests. Data are

presented as adjusted means (mg) SEM. The oviductal growth parameters were compared

among treatment groups with ANOVA followed by Fishers Protected LSD post-hoc test. Log

transformation of the data was performed in order to achieve homogeneity of variances prior to

ANOVA. Plasma concentrations of E2, T, and IGF-1 were estimated from raw data using the

commercially available Microplate Manager software (Microplate Manager III, BioRad

Laboratories, Inc., Hercules, CA, 1988). For RIA validation of pooled plasma dilutions and









internal standards, hormone concentrations were log 1 0-transformed prior to testing for

homogeneity of slopes with standard curves by ANCOVA. Plasma concentrations of E2, T, and

IGF- I were compared among treatment groups with one-way ANOVA followed by Scheffe post

hoc. Tamhane post hoc was used where variances were unequal among groups. Statistical

analyses were performed using SPSS software (v. 10, SPSS Inc., Chicago, IL, 1999) with c =

0.05.

Results

Biochemical RIA Validations

Internal standards and plasma dilutions were parallel to the standard curve for E2

(ANCOVA; F = 0.13, P = 0.73 and ANCOVA; F = 0.01, P = 0.91, Fig. 4-4A), and T RIA

(ANCOVA; F = 0.0001, P = 0.99; Fig. 4-4B). Plasma dilutions were parallel to the standard

curve for IGF-I RIA (ANCOVA; F = 0.01, P = 0.90; Fig. 4-4C).

Tissue Weights

At the time of the ovariectomy surgeries, excised ovaries contained predominantly

mature, highly polarized follicles exhibiting clear demarcation between animal and vegetal

hemispheres. Several of the sham females had slightly immature ovaries with mostly yellow and

some vitellogenic follicles. Ovary mass did not vary significantly among treatment groups

(ANCOVA; F = 0.16, P = 0.16).

Liver mass was not significantly different among treatment groups (ANCOVA; F = 0.48,

P = 0.79). Oviduct mass was greatest in frogs given E2 and E2/IGF- 1 compared to those given

placebo, EGF, IGF-1, and sham treatments (ANCOVA; F = 5.14, P = 0.001; Fig. 4-5).

Oviduct Morphometrics

The wall of the oviduct is composed of three distinct morphological regions: the

endometrium lined with a lumenal epithelium, the myometrium, and the outer serosa layers. The

lumen, or central region of the oviduct, receives secretions synthesized by the epithelial cells and

endometrial glands. The epithelial layer forms a continuous boundary surrounding the oviductal









lumen. The endometrial layer, lying internally to the lumenal epithelium, contained secretory

glands, connective tissue, and capillaries. The muscle layer, or myometrium, is composed of

smooth muscle and forms a continuous external boundary around the endometrium. The outer

covering of the oviduct is a relatively thin layer of connective tissue, the serosa. Representative

sections from oviducts of frogs under each treatment group are shown (Fig. 4-6, 4-7).

Regardless of treatment group, the oviducts exhibited a ciliated epithelial layer composed

primarily of cuboidal cells with darkly staining, basal nuclei. The epithelial cell layer appeared

more convoluted in frogs given placebo (Fig. 4-6A), EGF (Fig. 4-6B), IGF-1 (Fig. 4-6C), and

sham (Fig. 4-7C) treatments. For frogs given E2 (Fig. 4-7A) and simultaneous E2/IGF-1 (Fig. 4-

7B) treatment, the epithelial cell layer exhibited little or no convolution, and formed a fairly

straight, continuous boundary around the lumen. Epithelial cell height was greater in E2 and

simultaneous E2/IGF-1 treated frogs compared to controls and other treatment groups (Fig. 4-8A).

Endometrial layer thickness (Fig. 4-8B) and surface area (Fig. 4-8C) were greater for

frogs given E2 and simultaneous E2/IGF treatments compared to other groups, and no difference

in growth as noted between these two treatment groups. The endometrial layer in frogs receiving

these treatments contained numerous large and densely arranged glands. Often the gland height

extended the entire width of the endometrium. Cells containing abundant cytoplasm, darkly

staining nuclei, and a central gland lumen comprised the glands, which also had a duct opening

onto the lumenal epithelium. The oviductal glands in frogs receiving placebo, EGF, IGF-1, and

sham treatments were much smaller in height, width, and total surface area. The endometrial

layer of these frogs was much reduced and connective tissue occupied more relative endometrial

space than did the glands.

Plasma Steroid and IGF-1 Concentrations

Pre-ovariectomy plasma hormones were similar in females among treatment groups for

E2 (P = 0.08), T (P = 0.40), and IGF-1 (P = 0.30). Collectively these data indicate that the

females were in a similar reproductive stage, and had similar plasma steroid and IGF-1









concentrations before treatments. Thus, their responses to the treatments are unlikely to have been

obscured by pre-ovariectomy differences in these parameters.

Plasma E2 concentrations were decreased significantly after treatment with placebo (P <

0.001), EGF (P = 0.001), or IGF-1 (P < 0.001) but were similar to pre-ovariectomy

concentrations for E2, E2 /IGF-1, and sham treatment groups (Fig. 4-9). After treatments, plasma

E2 concentrations were greater in E2, E2/IGF-1, and sham female compared to placebo, EGF, and

IGF-l treatment groups (P < 0.001; Fig. 4-10).

Compared to pre-ovariectomy samples, plasma T concentrations were decreased after

placebo (P = 0.02), EGF (P = 0.01), IGF-1 (P = 0.002), E2 (P = 0.01), and E2/IGF-1 (P = 0.03)

treatment, but not for sham treatment (P > 0.05; Fig. 4-11). Following treatments, plasma T was

higher for only the sham group (P < 0.001; Fig. 4-12).

Compared to pre-ovariectomy samples, plasma IGF- 1 was significantly increased in frogs

given IGF-1 (P = 0.0005), E2 (P < 0.001), and E2/IGF-I (t-test; P < 0.001) but lower for placebo,

EGF, and sham females (t-test; P > 0.05; Fig. 4-13). After treatment, plasma IGF-I was higher in

IGF-1, E2, and E2/IGF-I females than in placebo, EGF, and sham females (ANOVA; P < 0.001;

Fig. 4-14).

Ovariectomized frogs given placebo, EGF, and IGF- 1 exhibited lower post-treatment

steroid concentrations compared to pre-ovariectomy concentrations, and verify that the

ovariectomy surgeries were successful in removing endogenous ovarian steroid sources. In

addition, similar pre-ovariectomy and post-treatment plasma E2 concentrations for E2 and E2/IGF

treated frogs, and similar pre-ovariectomy and post-treatment plasma IGF- 1 concentrations for

IGF and E2/IGF-I treated frogs indicate that E2 and IGF-1 treatments were delivered effectively

at physiologically relevant concentrations.

Discussion

Results from this study confirm that E2 stimulates oviductal growth in R. catesbeiana.

Treatment with growth factors, placebo, and sham produced no oviduct growth. Lastly, treatment









with combined E2/IGF failed to stimulate a greater oviduct growth than was observed with E2

treatment alone. In contrast to reptiles and mammals examined using similar technique, R.

catesbeiana did not exhibit an oviductal growth response with EGF or IGF-1 treatment, nor did

they exhibit a synergistic growth response to E2/IGF-l treatment. Although E2 and IGF-1 are not

synergistic in stimulation of oviduct growth in R. catesbeiana, both hormones might still be

required for oviductal growth.

There are several possible explanations for the absence of an oviduct growth response in

R. catesbeiana to growth factor (or to combined E2/IGF- 1) treatment. First, the IGF- 1 treatment

doses might have been insufficient to elicit an oviductal growth response in R. catesbeiana.

Future studies should investigate what doses of IGF- 1 are capable of stimulating oviduct growth

in ovariectomized R catesbeiana. However, it is unlikely that the IGF- 1 dose was insufficient

because frogs given IGF- 1 exhibited greater post-treatment than pre-ovariectomy plasma IGF- 1

concentrations. It is possible that the oviduct must first be "primed" with E2-stimulation before

IGF- 1 exposure to become sensitive to the effects of IGF-1. This priming of oviductal tissue

might involve E2-alpha receptors (ERa) and IGF- I receptors (IGF- 1 R) upregulation. Klotz et al.

(2000) demonstrated that ERa is required for IGF-1 to induce a cellular response. Additionally,

Clark et al. (1997) demonstrated that E2 stimulates proliferative responses of reproductive tissues

by upregulating IGF- 1 R expression, which increases tissue response to circulating IGF-1. These

findings imply that an increase in circulatory E2 concentrations can sensitize receptor-dependent

tissue growth IGF- 1 stimulation without necessarily requiring an increase in circulating IGF- 1

concentrations. As a second explanation, we must consider that circulating steroids can arise from

non-gonadal sources such as the adrenal glands (Norris, 1997). As a third explanation, sensitivity

to these growth factors represents a relatively recent evolutionary change in reptilian and

mammalian oviductal physiology. It is important to recognize that findings reported in this study

might be exclusive to R. catesbeiana. There are likely interspecific differences in hormonal









regulation of oviduct growth among amphibians. Accordingly, more amphibian species should be

examined, using similar techniques, before we can fully understand how amphibian oviduct

growth is regulated by interactions of steroids and growth factors.

It is interesting to note that oviduct growth in sham frogs was not similar to growth in E2-

and in E2/IGF-1 treated frogs. Sham frogs were expected to exhibit oviduct growth, similar to E2-

treated frogs but greater than that of placebo frogs, because their intact ovaries would continue to

synthesize and secrete E2 throughout the study. There are several possible explanations for these

unexpected findings. First, it is possible that the implants in E2- and E2/IGF-treated frogs

contained E2 concentrations higher than is typically found in R. catesbeiana. E2 doses were

determined based on studies of E2 necessary to elicit oviduct growth in Xenopus laevis (Follett

and Redshaw, 1967; Redshaw et al., 1968) and in reptiles (Cox, 1994). Thus, these doses might

have been comparatively high for R. catesbeiana. However, this hypothesis seems unlikely

because pre-surgery and post-treatment plasma E2 concentrations were similar for E2- and

E2/IGF- 1-treated frogs. A second explanation is that sham frogs were different from E2- and

E2/IGF- 1-treated frogs with respect to pre-surgery and post-treatment plasma IGF- I

concentrations. In E2- and E2/IGF-1 -treated frogs, plasma IGF-1 concentrations increased after

treatment compared to pre-ovariectomy levels. In sham frogs, however, post-treatment plasma

IGF-1 concentrations did not increase relative to pre-surgery levels. In E2- and E2/IGF- 1-treated

frogs, the increase in plasma IGF-1 could have stimulated increased IGF- I R expression in

oviductal tissues, making them more sensitive to E2- and IGF- I stimulation. As mentioned

previously, IGF-1 R does interact, or exhibit "cross-talk" with the ERx in stimulating oviduct

growth (Klotz et al., 2002). Since plasma IGF-1 concentrations in sham frogs did not change

during the experiment, it is possible that oviductal IGF- 1 R expression also remained unchanged,

making oviductal tissue comparatively less sensitive to E2- or IGF-1-induced stimulation of

growth. Future studies should examine oviduct growth not only in response to steroid and growth

factors hormones, but in also in response to changes in ERa and IGF- I R activity to better








understand the role of these receptors in mediating the effects of E2 and IGF-1 on oviduct growth.

Finally, there might have also been an implant effect on oviduct growth. With or without

hormones, the implant might have elicited oviductal hypertrophy due to an irritation response of

the frogs to implant "foreign body" within the abdomen; this implant effect would have been

absent in sham frogs.

Changes in oviductal mass associated with seasonal changes in plasma steroids have been

described for a wild population of R. catesbeiana (Licht et al., 1984). However, more research is

necessary to understand the mechanism by which E2 induces a growth response in target tissues

of amphibians. In mammals, ovarian steroids induced a complex suite of morphological,

physiological, and biochemical changes in the oviduct (Buhi et al., 1997). Estrogen-induced

oviduct growth relies upon activation of genes that modulate expression of growth factors and

their receptors (Murphy and Murphy, 1994; Cox and Guillette, 1994). Estrogen stimulates DNA

synthesis and mitosis of epithelial cells, and increases uterine IGF- I and IGF- 1 R gene expression.

In uterine cells, IGF-1 induces DNA synthesis similar to E2-stimulation. Thus, activation of the

growth factor signaling systems by E2 is an important part of uterine growth and proliferation in

mammals (Klotz et al., 2002; Segars and Driggers, 2002; Driggers and Segars, 2002). It remains

unknown whether E2-induced oviduct growth in amphibians occurs through activation of these

growth factors and their receptors, release of IGF- 1 from IGF- 1 binding proteins, or by another

mechanism not yet identified.

No study has comprehensively examined the effects of both growth factors and steroids

on oviduct morphology in amphibians. However, both IGF- 1 and EGF have been associated with

oviduct growth in mammals and reptiles (Cox and Guillette, 1994; Murphy, 1990; DiAugustine et

al., 1988). In ovariectomized geckos, IGF-1 and EGF stimulates moderate growth of the oviduct

in the absence of E2 indicating these growth factors play an important role in reptilian

reproduction. Also in reptiles, EGF and IGF-1 are known to stimulate oviduct growth directly,

although the exact mechanism for proliferation in not known. In amphibians, IGF- 1 and IGF- 1









binding proteins have been identified in the plasma but the location of IGF- 1, IGF- I binding

proteins, and IGF- 1 receptors in reproductive tissues of amphibians have not been examined.

Location and activity of IGF-1BPs in oviductal tissue, in addition to their interaction with

circulating IGF- 1 in amphibians, is necessary to understand how these binding proteins regulate

IGF-1 activity and oviduct growth.

As expected, plasma IGF- 1 concentrations increased with IGF- I and simultaneous

E2/IGF- 1 treatment. However, an unexpected increase in plasma IGF-1 was observed in females

treated exclusively with E2. An interesting endocrine pathway can be described from these

findings. The liver is the primary site for synthesis of IGF- 1 found in the plasma. Perhaps E2

stimulated liver IGF-1 synthesis directly, or E2-stimulated increased pituitary growth hormone

release that, in turn, stimulated liver IGF-1 synthesis. Another possible source for the increased

plasma IGF- 1 in E2-treated females is the oviduct. There are an increasing number of studies that

have identified non-hepatic sources of IGF- 1 and examined their role in mediating tissue growth.

Numerous studies in reptiles and mammals have shown that the oviduct synthesizes IGF- 1 (Cox

and Guillette, 1993; Cox, 1994; Le Roith et al., 2001; Driggers and Segars, 2002; Klotz et al.,

2002; Segars and Driggers, 2002). As part of separate study not described here, IGF-1

immunoreactivity has been detected in the oviduct of R catesbeiana using immunocytochemistry

(T. Barbeau, unpub. obs.). There is some evidence that oviduct-derived IGF- I affects the oviduct

itself in an autocrine manner or affects nearby tissues in a paracrine manner. Whether the oviduct

can secrete and contribute significantly to plasma concentrations of IGF-1 remains unknown and

is an intriguing area for future research.

In summary, treatment of ovariectomized R. catesbeiana with exogenous E2 and resulted

in increased plasma IGF- 1 concentrations. This finding indicates that E2 interacts with the IGF- I

system in amphibians. It remains unknown if increased hepatic or oviductal IGF- 1 synthesis and

secretion contributed to the increase in plasma IGF- 1 concentrations observed in E2-treated

females. The mechanism by which E2 stimulates increased plasma IGF- I concentrations in frogs,






87


and the localization of non-hepatic sources of IGF-1 synthesis in amphibians is poorly understood

and requires further investigation.
































Figure 4-1. Exteriorized right ovary during ovariectomy surgery in Rana catesbeiana. Both the
left and right ovaries were removed from a 2.5 cm, right paramedial incision into the
abdominal cavity.























sietorn
site


are 4-2. Healed right-paramedical incision site visible three weeks after ovariectomy. Also
shown is the site of the left paramedial incision in which pellet implants were placed
into the abdominal cavity of Rana catesbeiana.


































I .. .. w V% kkl
Figure 4-3. Location of intra-abdominal treatment pellet positioned over the left oviduct at time
of final dissection, after completion of 18 days of treatment in Rana catesbeiana.










o Standard
* Plasma Dilutions
* Internal Standards


0 *o0


100(


100 -

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40
20


0 Standard
Plasma Dilutions
* N Internal Standards


0*
oA o


1000


10000


* Standard
* Plasma Dilutions


0 0
0


1000


10000


IGF-I (pg)
Figure 4-4. Biochemical validation of Rana catesbeiana plasma. A. 17f3-estradiol RIA internal
standards (ANCOVA; F = 0.13; P = 0.73) and plasma dilutions (ANCOVA; F =
0.01; P = 0.91) were parallel to the standard curve. B. testosterone RIA internal
standards (ANCOVA; F = 0.0001; P = 0.99) and plasma dilutions (ANCOVA; F
0.0.08, P = 0.79) were parallel to the standard curve. C. insulin-like growth factor-1
(IGF- 1) RIA .the plasma dilutions curve was parallel to the standard curve
(ANCOVA; F = 0.014; P = 0.91).


0 0


10000


100


Estradiol (pg)


B 100
80
-60
40
20


100


Testosterone (pg)


C 100
80
~60~
40
S20


00


100




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INFLUENCE OF INSULIN-LIKE GROWTH FACTOR-1, STEROIDS, AND NITRATE ON
REPRODUCTION IN AMPHIBIANS
By
TAMATHA R. BARBEAU
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
2004

Copyright 2004
By
Tamatha R. Barbeau

ACKNOWLEDGMENTS
I thank my advisor, Louis J. Guillette, whose encouragement, guidance, and motivation
were invaluable during this work. His expertise and enthusiasm for research provided the
foundation for this work, and his passion for teaching these skills to others has molded my own
desire to pursue an academic career. I owe much gratitude to my committee members (William
Buhi, Lauren Chapman, David Evans, and Harvey Lillywhite) for their advice and insightful
conversations throughout this study. As physiologists David and William made indelible
impressions on me and provided invaluable insights and ideas for my research. Lou, Harvey, and
Lauren have been my committee members, mentors, and friends during both my M.S. and Ph.D.
degrees. Collectively, they have made the greatest contributions to my professional development,
academic philosophies, research skills, scientific curiosity, and perspectives on life.
This research has been supported by grants from SIGMA XI Grants in Aid of Research,
The Brian Riewald Memorial Fund (UF), and Declining Amphibians Population Task Force. All
frogs were used in compliance with and supervision of the Institutional Animal Care and Use
Committee at the University of Florida (IACUC #Z023 and #Z095).
For their generous laboratory support, training, and camaraderie, I thank my friends and
colleagues (Dieldrich Bermudez, Teresa Bryan, Thea Edwards, Mark Gunderson, Iske Larkin,
Matthew Milnes, and Brandon Moore). I extend additional thanks to Colin Chapman, Ginger
Clark, and Douglas Levey for their help in providing me with laboratory techniques and space to
conduct my research. Many friends and colleagues at the University of Florida provided valuable
insights and assistance with various aspects of my work (namely Keith Choe, Martin Cohn,
Franklin Percival, and Kent Vliet). I am especially grateful for the help of Loretta Azzinario,
Jason Bridge, Arika Brown, Tim Buhi, Leo Choe, Brandy Cunningham, Lauren Farrar, David
iii

Iglesias, Kapila Karakota, Caroline Keicher, Dana LaKam, Axel Lucca, Courtney Marler, Amy
McGreane, Pamela Moses, Amanda Mulligan, Reshma Patel, Sonia Parikh, Wilhelmina Randtke,
Maria Samuel, Catherine Vallance, and Kyu Mee Yo.
My husband, Greg Pryor, continues to be the most important and supportive person in my
academic development and in life. His encouragement, guidance, and sense of humor have been
the one constant, throughout the triumphs and tribulations of my graduate work. I look forward to
sharing many more adventures with him in the future. I am grateful to my mother, for being
patient when I brought snakes and frogs home as a child; and to my father, for taking me on
camping trips in the Northern Adirondacks. These events inspired my appreciation and curiosity
for ecosystems and animals of all kinds.
IV

TABLE OF CONTENTS
page
ACKNOWLEDGMENTS iii
ABSTRACT viii
CHAPTER
1 INTRODUCTION: INFLUENCE OF STEROIDS, INSULIN-LIKE GROWTH
FACTOR-1, AND NITRATE ON REPRODUCTION IN AMPHIBIANS 1
Reproductive Steroids and Amphibian Reproduction 1
Insulin-Like Growth Factor-1 3
Aquatic Nitrate and Amphibian Reproduction 7
Research Objectives 9
Physiology and Evolution 11
Conservation 12
2 THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS OF NITRATE (IN VIVO) ON PLASMA STEROIDS AND
INSULIN-LIKE GROWTH FACTOR-1, ON OVARIAN STEROID SYNTHESIS,
AND ON OVIDUCT GROWTH IN THE AFRICAN CLAWED FROG (Xenopus
laevis) 15
Introduction 15
Materials and Methods 18
Animals and Samples 18
Nitrate Study Design 18
Steroid Radioimmunoassay (RIA) Procedures 20
Insulin-Like Growth Factor-1 (IGF-1) RIA Procedures 21
Biochemical RIA Validations 21
Statistics 22
Results 22
Tissue Weights 22
Follicle Diameters 22
Plasma Steroid Concentrations 23
Plasma IGF-1 Concentrations 23
Ovarian Follicle Steroid Concentrations (Ex Vivo) 23
Discussion 24
v

3 SEASONAL CHANGES IN INSULIN-LIKE GROWTH FACTOR-1, STEROIDS,
AND REPRODUCTIVE TISSUES IN PIG FROGS {Rana grylio) 36
Introduction 36
Materials and Methods 39
Water Parameters, Animal Captures, and Sample Collections 39
Steroid Radioimmunoassay (RIA) Biochemical Validation 42
Steroid RIA Procedures 43
Insulin-like Growth Factor-1 (IGF-1) RIA Biochemical Validation 44
IGF-1 RIA Procedures 45
Statistics 46
Results 47
Seasonal Environmental Parameters 47
Seasonal Tissue Mass and Ovarian Maturation 47
Seasonal Plasma Steroid and IGF-1 Concentrations 48
Correlations: Plasma Steroids, Tissue Mass, and Environmental Parameters 49
Discussion 50
4 THE EFFECTS OF INSULIN-LIKE GROWTH FACTOR-1 AND ESTRADIOL
IMPLANTS {IN VIVO) ON OVIDUCT MORPHOLOGY, AND ON PLASMA
HORMONES IN BULLFROGS {Rana catesbeiana) 70
Introduction 70
Materials and Methods 72
Ovariectomy 73
Hormone Implants 74
Tissue Sampling 75
Steroid Radioimmunoassay (RIA) Biochemical Validation 76
Steroid RIA Procedures 77
Insulin-Like Growth Factor-1 (IGF-1) RIA Biochemical Validation 78
IGF-1 RIA Procedures 78
Statistics 79
Results 80
Biochemical RIA Validations 80
Tissue Weights 80
Oviduct Morphometries 80
Plasma Steroid and IGF-1 Concentrations 81
Discussion 82
5 OVARIAN STEROIDOGENESIS {IN VITRO) IN PIG FROGS {Rana grylio) AFTER
EXPOSURE TO ENVIRONMENTALLY RELEVANT CONCENTRATIONS OF
NITRATE AND NITRITE 102
Introduction 102
Materials and Methods 106
Collection of Animals 106
Ovarian Follicle Culture {In Vitro) 106
Steroid Radioimmunoassay (RIA) Procedures and Validations 107
Statistics 108
Results 108
Discussion 109
vi

6 THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS ON NITRATE ON OVIDUCTAL MORPHOLOGY
MORPHOLOGY AND PLASMA STEROIDS AND INSULIN-LIKE GROWTH
FACTOR-1 IN BULLFROGS (Rana catesbeiand) 118
Introduction 118
Materials and Methods 121
Animals 121
Nitrate Treatments 122
Steroid Radioimmunoassay (RIA) Procedures and Validations 124
Insulin-Like Growth Factor-1 (IGF-1) RIA Procedures and Validations 125
Statistics 127
Results 128
Oviduct Weights 128
Plasma Steroid and IGF-1 Concentrations 128
Oviduct Morphometries 128
Discussion 129
7 CONCLUSIONS 141
Seasonal Plasma Steroids and IGF-1, and Reproductive Tissue Growth 141
The Effects of IGF-1, E2, and Nitrate on Oviduct Growth 143
Nitrate Exposure (In Vivo and In Vitro)'. Effects on Steroidogenesis 147
Nitrate Exposure and Plasma IGF-1 150
REFERENCES 152
BIOGRAPHICAL SKETCH 169
VII

Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
INFLUENCE OF INSULIN-LIKE GROWTH FACTOR-1, STEROIDS, AND
NITRATE ON REPRODUCTION IN AMPHIBIANS
By
Tamatha R. Barbeau
August 2003
Chair: Louis J. Guillette
Major Department: Zoology
My goal was to examine the influence of insulin-like growth factor-1 (IGF-1), 17-P
estradiol (E2), testosterone (T), and nitrate exposure on various aspects of reproduction in frogs.
To accomplish this, I investigated seasonal changes in plasma IGF-1, E2, and T concentrations in
a wild population of Rana grylio. I also determined the importance of steroid and growth factor
hormones in reproductive physiology by examining ovariectomized Rana catesbeiana for
changes in plasma IGF-1, E2, and T concentrations, and changes in oviduct morphology after
treatment with known doses of IGF-1, E2, and epidermal growth factor (EGF). Finally, I
examined three aquatic frogs species (Xenopus laevis, R. grylio, and R. catesbeiana) for the
effects of nitrate exposure on changes in plasma IGF-1, E2, and T concentrations, and on oviduct
morphology.
I have demonstrated that plasma IGF-1, E2, and T concentrations (and reproductive tissue
growth) exhibit a clear seasonal pattern of changes that overlap with changes in environmental
variables, such that reproductive condition is optimized to match favorable environmental
temperatures. I also demonstrated that E2 is a potent stimulator of oviduct growth, while EGF and

IGF-1 do not induce oviductal growth in R. catesbeiana. I also provide the first evidence that
exposure to environmentally relevant concentrations of nitrate alters endocrine hormones in
Xenopus laevis, R grylio, and R catesbeiana. Furthermore, IGF-1 and steroid hormone
concentrations are altered with exposure to nitrate at concentrations deemed safe for human
drinking water by the US EPA (10 mg/L). In vivo exposure of X. laevis (for 7 continuous days) to
nitrate concentrations below 50 mg/L significantly increased plasma IGF-1 concentrations, and
inhibited ovarian E2 and T synthesis. In vitro incubation of ovarian tissue (from wild-caught R
grylio) with nitrate concentrations between 0.17 and 33.00 mg/L nitrate (and between 0.20 and
40.60 mg/L nitrite) inhibited E2 and T synthesis after 3 hours of exposure. Lastly, in vivo
exposure of R. catesbeiana to nitrate concentrations between 1.65 and 16.50 mg/L increased
plasma IGF-1, E2, and T concentrations; and caused oviductal atrophy. These findings
demonstrate that exposure to nitrate at extremely low concentrations causes endocrine disruption
in frogs.
IX

CHAPTER 1
INTRODUCTION: INFLUENCE OF STEROIDS, INSULIN-LIKE GROWTH
FACTOR-1, AND AQUATIC NITRATE ON REPRODUCTION IN AMPHIBIANS
Reproductive Steroids and Amphibian Reproduction
Amphibians display some of the most diverse reproductive modes compared to other
vertebrates. Most amphibians exhibit the ancestral reproductive mode, and are restricted to water
to oviposit and fertilize eggs externally. Some species are terrestrial breeders, have internal
fertilization, and either oviposit eggs on land or retain them within the oviducts for all or part of
the embryonic developmental period. Finally, some amphibians oviposit terrestrial eggs from
which offspring hatch, bypass the free-living tadpole stage, and undergo direct development to
emerge as fully developed froglets (Wake and Dickie, 1998).
The reproductive system in amphibians is characterized by cyclic changes in growth and
function that are modulated by hypothalamic releasing hormones, pituitary gonadotropins, and
gonadal steroids. The process of steroid synthesis or steroidogenesis is regulated primarily by the
hypothalamic-pituitary-gonadal axis (Licht, 1970, 1979; Licht et al., 1983). Gonadotropin¬
releasing hormone (GnRH) is secreted by the hypothalamus (in response to internal or
environmental cues) and stimulates the anterior pituitary to release luteinizing hormone (LH) and
follicle stimulating hormone (FSH) into the bloodstream. These gonadotropins stimulate gonadal
steroidogenesis and gametogenesis. The principal gonadal steroids are progesterone (P4), estradiol
17P (E2), and testosterone (T). Theca interna cells within the ovary synthesize T in response to
LH stimulation. In response to FSH stimulation, ovarian granulosa cells synthesize aromatase,

2
Hypothalamus
GnRH
Inhibition
Stimulation
Ovarian Cells
Inhibin
Theca Interna
Granulosa
Figure 1-1. Regulation of gonadal steroidogenesis. (A) Hypothalamic gonadotropin releasing
hormone (GnRH) induces pituitary secretion of luteinizing hormone (LH) and follicle
stimulating hormone (FSH) (B). LH stimulates theca interna cells (C) to synthesize
testosterone (T). FSH stimulates granulosa cell to produce aromatase enzymes (D),
which convert T into estrogen (E2). Steroidogenesis induces inhibin release from the
gonad, which inhibits further hypothalamic and pituitary stimulation (E). F.
Circulating steroids are also metabolized and cleared from the bloodstream by the
liver.
an enzyme that converts T into E2 (Figure 1-1). These steroids induce gonadal release of inhibin,
a hormone that inhibits hypothalamic-pituitary stimulation of further steroid synthesis. Gonadal
steroids exert an autocrine or paracrine action, by influencing localized tissues; or function as
endocrine hormones when released into the bloodstream, to affect distant target tissues. Within
steroid-responsive tissues, T and E2 bind to and activate cytosolic or nuclear receptors and form a
steroid-receptor complex. This complex binds to a hormone-response element on DNA to

3
stimulate or inhibit transcription, protein synthesis, and tissue growth (Segars and Driggers,
2002). Through this process, E2 and T regulate normal development of secondary sexual
characteristics, regulate growth of steroid-responsive tissues, and regulate reproductive function
(Guidice, 1999).
In amphibians, E2 is essential for oocyte development and maturation within ovarian
follicles (Dumont, 1971; Fortune, 1983). Gonadal E2 and T regulate many aspects of reproductive
function, such as oviduct growth and secretions (Licht et al., 1983; Norris, 1997). The oviduct is a
vital structure for reproductive function in oviparous vertebrates, including amphibians (Giudice,
1992; Wake and Dickie, 1998). After ovulation from the ovaries, mature oocytes travel through
the oviduct to the cloaca and are expelled into the environment. In addition to providing physical
transport, the oviduct synthesizes and secretes proteins and other substances that nourish and
encapsulate the ova, and also aid in fertilization (Low et al., 1976; Buhi et al., 1997).
Insulin-Like Growth Factor-1
It has become increasingly apparent that reproductive function and physiology are
regulated by steroid-signaling pathways, and also by other pathways involving insulin-like
growth factor-1 (IGF-1). Originally called somatomedin C, IGF-1 is a polypeptide hormone that
is structurally similar to IGF-II and proinsulin, and likely originated early in vertebrate evolution.
IGF-1 is part of the growth factor system, which consists of a family of proteins that function in
regulating many cellular processes (including cell proliferation, differentiation, and apoptosis) in
virtually all tissues. (LeRoith et al., 2001a,b). Thus, IGF-1 is important for normal growth and
function of reproductive tissues, and also for somatic tissues. Accordingly, the role of IGF-1 in
growth of reproductive and somatic tissues has been examined in a variety of vertebrates
including mammals, fish, birds, and reptiles (Girbau et al., 1987; Murphy and Ghahary, 1990; De
Pablo et al., 1990; Serrano et al., 1990; Simmen et al., 1990; Scavo, 1991; Kapur et al., 1992; Cox
and Guillette, 1993; Tang et al., 1994; Guillette et al., 1996; Buhi et al., 2000; Qu et al., 2000;
Allan et al., 2001). Although IGF-1 has been identified in the plasma and tissues of some

4
amphibians, the role of this peptide hormone in tissue growth and function in these animals
remains unclear, and requires further study (Daughaday et al., 1985; Pancak-Roessler and Lee,
1990).
Traditionally, IGF-1 was thought to influence tissue growth primarily by mediating the
effects of growth hormone (GH). This physiological function of IGF-1 is the basis of the original
somatomedin hypothesis (LeRoith et al., 2001b). More recently, IGF-1 has been found to play an
important role in growth and differentiation of reproductive tissues (independent of GH), by
mediating the mitogenic effects of E2 (Girbau et al., 1987; Murphy and Ghahary, 1990; Cox,
1994).
In the presence of E2, IGF-1 has been shown to mediate growth of E2-sensitive
reproductive tissues like the oviduct (Mead et al., 1981; Murphy and Murphy, 1994). Research
indicates that the growth effects of IGF-1 does not require E2 but requires only the presence of the
17(3 estradiol alpha receptor (ERa) in reproductive tissues (Klotz et al., 2000). This is supported
by findings of an E2-like growth response in the oviduct of ovariectomized animals treated with
IGF-1 (Cox, 1994). These findings demonstrate that IGF-1 potentiates E2-induced growth and
also stimulates E2-independent tissue growth.
In addition to mediating the growth effects of reproductive steroids, IGF-1 has also been
shown to regulate intraovarian steroid synthesis in mammals (Adashi et al., 1991; Guidice, 1992;
Adashi, 1993). Decreased E2 expression increases ovarian IGF-1 expression. Ovarian IGF-1
stimulates synthesis of E2 and P4, and increases aromatization of androgens into E2 (Adashi et al.,
1991). Additionally, the ovaries and oviduct synthesize and secrete IGF-1 in response to GH,
FSH, E2, and other hormones. Based on these findings, the list of factors that regulate (or are
influenced by) the IGF-1 system has been expanded to include reproductive steroids.
Research spanning nearly 50 years has defined many components of the surprisingly
complex IGF-1 system (Le Roith et al., 2001b). In all vertebrates examined, the liver synthesizes
and secretes most of the circulating concentrations of IGF-1. Hypothalamic release of growth

5
hormone-releasing hormone (GHRH) stimulates the pituitary to secrete growth hormone (GH)
into the bloodstream. In response to GH stimulation, the liver synthesizes and secretes IGF-1 into
the bloodstream. Hepatic IGF-1 can affect peripheral tissues in a paracrine or autocrine manner;
or it can be transported through the bloodstream, bound to IGF binding proteins (IGFBPs), as an
endocrine hormone that mediates growth and apoptosis of distant target tissues. After reaching its
target tissue, IGF-1 interacts with a transmembrane cell-surface IGF-1 receptor (IGF-1R) where it
is released from its binding protein to initiate a cellular response. Excess circulating IGF-1 is then
filtered and degraded by the kidneys (LeRoith et al., 2003). In this manner, IGF-1 mediates GH-
induced cellular proliferation. This endocrine-signaling pathway is the basis of the original
somatomedin hypothesis. However, recent research suggests that the somatomedin hypothesis
should be revised. IGF-1 has been shown to have many GH-independent effects on regulating
tissue growth. Additionally, non-hepatic tissues (including the ovaries and oviduct) are now
known to synthesize and secrete IGF-1 (LeRoith et al., 2001b).
The extracellular functional components of the IGF-1 system include IGF-1, IGFBPs,
and IGF-1R. Expression of IGF-1 can be stimulated by various factors including growth
hormone, E2, T, P4, FSH, glucose, insulin, and thyrotropin; whereas, IGF-1 expression can be
inhibited by somatostatin, LH, cortisol, and interferon. Six known IGFBPs can bind with IGF-1
to modulate cellular effects. The IGFBPs that regulate the cellular effects of IGF-1 include
IGFBPs 1, 3, 4, and 5. The other binding proteins (IGFBPs 2 and 6) specifically regulate the
effects of IGF-2 on embryonic development. The IGFBPs prevent IGF-1 degradation during
circulation, transport IGF-1 to target tissues, and regulate binding of IGF-1 to IGF-1R. Like IGF-
1, IGFBPs can be stimulated or inhibited by various factors. Another functional component of the
IGF-1 system is the IGF-1R. The IGF-1R is a tyrosine kinase, transmembrane receptor found on
virtually every tissue type, and it mediates a majority of IGF-1 actions on cell growth. There is an
IGF-2 receptor, but it is highly specific for IGF-2 and functions mostly in mediating embryonic
development. Expression of the IGF-1 R can be stimulated by a variety of factors including E2,

6
A IGF-1
Figure 1-2. Binding of IGF-1 with the IGF-1R, initiates phosphorylation of intracellular proteins
in a signaling cascade that leads to a cellular response. Activation of adaptor proteins
includes the mitogen activated protein kinase (MAPK) pathway, the
phosphatidylinositol 3-kinase (IP-3K) pathway and its secondary messengers IP3,
DAG, and Ca*. Briefly, binding of IGF-1 (A) to the receptor (B) results in
autophosphorylation of the intracellular P-subunit of the receptor (C). This then
activates intracellular adaptor proteins, insulin receptor substrate (IRS) and She, to
bind with the receptor and become phosphorylated. If adaptor protein She is
activated, it forms a complex with SOS to activate Raf. Activation of Raf
phosphorylates protein kinase MEK and leads to phosphorylation (D) of mitogen-
activated protein kinase (MAPK). This activates transcription factors (TF) that bind
to nuclear DNA (F) to elicit a cellular response (G). If the adaptor protein IRS is
activated (H), a sequence of phosphorylations involving protein subunits p85 and
pi 10 will activate the IP-3K pathway (I). Activation of IP-3K pathway
phosphorylates the conversion of phosphoinositol bisphosphate (PIP2) to the second
messengers (J) inositol trisphosphate (IP3) and (K) diacyglycerol (DAG). Each of
these second messengers can induce cellular responses (L) either by activating the
CaVcalmodulin complex by IP3 or by the activation of phosphokinase C (PKC) by
DAG.
FSH, LH, and oncogenes. Conversely, IGF-1, P4, and tumor suppressors can inhibit expression of
the IGF-1R.
The functional IGF-1 system also has intracellular functional components that become
activated by binding of IGF-1 to the IGF-1R. Once bound, the IGF-1R becomes phosphorylated,
and a variety of intracellular proteins and second messengers are involved in a signaling cascade

7
that leads to a cellular response (Fig 1-2). Thus, each of the functional components of the IGF-1
system can be regulated by complex extracellular and intracellular factors.
Aquatic Nitrate and Amphibian Reproduction
In the past few years, there has been increased global concern over contamination of
water by anthropogenic sources of nitrate. Nitrate is an anionic form of nitrogen that infiltrates
watersheds in agricultural and urban environments, and reaches harmful concentrations largely
due to human activities. In agricultural areas, watersheds are polluted with nitrate from
unregulated run-off of nitrogen-based fertilizers and run-off of animal wastes. In urban areas,
nitrates contaminate watersheds primarily through runoff of industrial and wastewater effluent
from treatment plants and of fertilizers applied to lawns and golf courses (Rouse et al., 1999). The
application of fertilizers in close proximity to watersheds during the spring frequently results in
an overwhelming nitrate “pulse” that overlaps the breeding season of many amphibians.
Unfortunately, most studies of the effects of nitrate on amphibians report nitrate concentrations
differently, making comparisons and interpretation of these studies extremely difficult. For
consistency throughout this dissertation, nitrate is reported as equivalent to nitrate-as-nitrogen
(N03-N). This represents the concentration of nitrogen present in a given concentration of nitrate.
Additionally, equivalent measures of nitrate are provided here to facilitate comparison among
other nitrate and nitrite studies (Table 1-1).
Most studies on the effects of nitrate on amphibians have addressed toxicological rather
than sublethal concentrations (Rouse et al., 1999). Most of these studies also focused on juvenile
amphibian stages rather than on adults (Table 1-2). The impact of nitrate exposure on mammalian
steroidogenesis has been examined and described in a few studies. Nitrate exposure has been
shown to inhibit androgen synthesis in rodents in vivo and also in Mouse Leydig tumor cells in
vitro (Panesar, 1999; Panesar and Chan, 2000). One mechanism for altered steroid expression (in
vivo) by nitrates involves enzyme-dependent synthesis of nitric oxide (NO) (Panesar and Chan,
2000). The NO is synthesized from an L-arginine precursor by nitric oxide synthase (NOS)

8
enzymes (Kleinert et al., 1995; Mayer and Hemmenns, 1997). In addition to NOS-dependent NO
formation, non-enzymatic synthesis of NO can also occur through acidic reduction of nitrite
(Iizuka et al., 1999; Zweier et al., 1995, 1999; Modin et al., 2001). Cosby et al. (2003) reported
that hemoglobin functions as a nitrite reductase contributing to enzyme-independent NO
synthesis. Furthermore, Zweier et al., (1999) reported that enzyme-independent NO formation is
associated with cellular damage and loss of organ function. Regardless of the mechanisms by
which it is produced, NO is thought to regulate many physiological processes. Within the gonad,
NO can inhibit steroidogenesis by binding to the heme (iron-containing) groups located on the
enzymes of the cytochrome P450 superfamily necessary for steroid synthesis, like 30-
dehydroxysteroid dehydrogenase (30-HSD). (Van Voorhis et al., 1994; Panesar and Chan, 2000).
The IGF-1 counteracts the effects of NO by increasing ovarian E2 synthesis (Van Voorhis et al.,
1994; Van Voorhis et al., 1995; Srivastava et al., 1998; Inigues et al., 2001; Les Dees et al.,
2001). Within the mitochondria of cells, the enzymes P450scC and 30-HSD convert free
cholesterol into P4 (the precursor for T and E2). Steroid enzyme pathways disrupted by NO can
inhibit P4 and downstream androgen synthesis (Panesar and Chan, 2000). If P4 and T synthesis are
inhibited by nitrate, then less androgen is available for aromatase enzymes to synthesize into E2,
and estrogen concentrations would be altered. Despite findings of endocrine disruption by nitrate
in mammals, no study has examined whether nitrate disrupts endocrine function in adult,
reproductive amphibians.
Since E2 and IGF-1 interact to regulate growth-related responses in reproductive tissues,
it is plausible that alteration of E2 expression by nitrates might also influence IGF-1 and oviduct
growth (perhaps through a NO-dependent pathway). In humans, intraovarian IGF-1 expression
increases in response to elevated intraovarian NO. The IGF-1 apparently counteracts the
inhibitory effects of NO on steroids, by stimulating increased expression of aromatase enzymes,
StAR protein, P4, and E2. Thus, increased intraovarian IGF-1 in response to NO might be a

9
compensatory response, functioning to amplify steroid synthesis that has been compromised
(Schams et al., 1988; Erickson et al., 1989; Adashi, 1993; Samaras et al., 1996; Iniguez et al.,
2001; Les Dees et al., 2001). These studies indicate that IGF-1 plays a vital role in steroid
synthesis and regulation, and possibly functions through an NO-dependent pathway. The
mechanism by which steroids and IGF-1 interact to stimulate growth of reproductive tissue
remains enigmatic and requires further study. In addition, the influence of nitrate exposure on
reproductive physiology of anurans remains unknown.
Organic nitrate and nitrite are normally present in aquatic habitats, in low concentrations,
due to bacterial breakdown of organic matter and accumulation of biological wastes. In addition
to contributions from natural sources, anthropogenic sources of nitrate and nitrite can
compromise water quality even further. Unusually high concentrations of nitrate and nitrite can
accumulate in aquatic habitats that receive runoff of agricultural fertilizers and animal wastes.
Aquatic nitrate and nitrite contamination might provide a biological signal to frogs that water
quality is unsuitable for reproduction. High nitrate and nitrite concentrations might repress
physiological changes that stimulate reproductive condition of frogs. Contamination of aquatic
habitats with nitrate and nitrite has been shown to be detrimental to survival of anuran eggs and
tadpoles (Table 1-2), and amphibian populations are reportedly declining in some agricultural
areas (Berger, 1989).
Research Objectives
One goal of my study was to gain a better understanding of the interaction of IGF-1 with
E2 -dependent and independent growth of reproductive tissues in aquatic amphibians. Although
IGF-1 is important for cell growth and differentiation, abnormally high concentrations of plasma
IGF-1 are associated with abnormal growth of reproductive tissues; and with cancer of the breast,
ovaries, uterus, endometrium, and prostate (LeRoith et al., 1995a,b; Grimberg and Cohen, 1999;
van Dessel et al., 1999; Werner and Le Roith, 2000; Smith et al., 2000). The IGF-1 and IGF-1R
can protect cells from apoptosis; but in some mammals, over-expression of these receptors

10
induces ligand-dependent tumor formation. Over-expression of IGF-1R can be induced by up-
regulation of IGF-1 expression in response to growth hormone (GH)-, E2-, and ERa-dependent
pathways (Kaleko et al., 1990). Additionally, uterine IGF-1 and IGF-1R up-regulation (along
with increased uterine epithelial cell growth) occurs in ovariectomized rodents in response to
synthetic estrogens (DES and bisphenol A) and phytoestrogens (Klotz et al., 2000). From these
findings, I hypothesized that endocrine disrupting contaminants (EDCs) could affect the IGF-1
system. In a variety of vertebrates EDCs have been shown to alter reproduction. Much research
has focused on the interaction of EDCs with steroid hormones and their receptors (Rooney and
Guillette, 2000). Unfortunately, the effect of EDCs on the IGF-1 system has received surprisingly
little scientific scrutiny (Backlin and Bergman. 1995; Backlin, et al., 1998). Thus, another goal of
my study was to determine whether nitrate and nitrite (known to induce developmental
abnormalities in amphibians and reproductive abnormalities in other vertebrates) can alter
concentrations of IGF-1 and steroid hormones and alter growth of reproductive tissues in
amphibians.
The effect of nitrate on synthesis of IGF-1 and steroids remains an important topic for
investigation. Growing evidence indicates that nitrate exposure stimulates NO synthesis in body
tissues. Furthermore, increased NO expression in gonadal tissues affects steroid and IGF-1
expression. Thus, nitrate exposure might influence IGF-1 synthesis, similar to steroids, through
an NO-dependent or independent pathway.
Finally, my study examined adult anurans for seasonal changes in IGF-1 and steroid
concentrations, and in reproductive tissues. Seasonal patterns of change in plasma IGF-1 and
steroid hormone concentrations, and in growth of reproductive tissues, are reported for alligators
and turtles (Crain et al., 1995; Guillette et al., 1996). In anurans, seasonal changes in plasma IGF-
1 have been reported for the Woodhouse toad, Bufo woodhousei (Pancak-Roessler and Lee,
1990). Thus, I expected that plasma IGF-1 and steroid concentrations, and growth of reproductive

11
tissues, would exhibit a seasonal pattern of change in response to endogenous stimulation and
environmental cues.
In addition to addressing the goals mentioned above, findings from my study also have
more general applications for studies of amphibian physiology, evolution, and conservation.
Physiology and Evolution
Physiological regulation of the IGF-1 system has been examined in mammals and
reptiles. The IGF-1 has been shown to regulate gonadal steroid synthesis, to stimulate oviductal
growth, and to exhibit seasonal cyclicity in mammals and reptiles. Recent research on reptiles and
mammals demonstrates that IGF-1 potentiates E2-induced growth of reproductive tissues like the
oviduct. Even in the absence of endogenous E2, IGF-1 stimulates significant oviduct growth.
Thus, the role of growth factors in reptilian and mammalian reproduction is more important than
previously recognized. Additionally, seasonal cycles of increased plasma steroid concentrations
and increased reproductive tissue growth overlap with increases in plasma IGF-1 in reptiles and
mammals (Crain et al., 1995; Guillette et al., 1996; Webster et al., 2001). These findings indicate
that IGF-1 is associated with reproductive activity, and is responsive to changes in reproductive
parameters and environmental cues.
In amphibians, the presence of IGF-1 has been documented; but the physiological
processes that regulate this system remain largely under-investigated. If the amphibian oviduct
responds to IGF (similar to mammals and reptiles), then IGF-1 regulation of reproductive tissues
represents an early evolutionary phenomenon. However, if the amphibian oviduct is unresponsive
to IGF-1 stimulation, then IGF-induced oviduct growth might represent a relatively recent
development in reptiles and mammals. Seasonal changes in plasma IGF-1 concentration have
been described for B. woodhousei, but it remains unknown if changes in IGF-1 parallel
reproductive parameters in this or other amphibian species (Pankcak-Roessler and Lee, 1990).
My study provides the first description of how endogenous steroids, environmental factors, and
reproductive cyclicity influence the IGF-1 system in amphibians.

12
Conservation
Amphibian populations in some agricultural areas are declining, and frogs have been
found with dramatic deformities. The factors responsible for these declines and deformities are
hard to identify, but might include runoff of nitrogenous fertilizers from agricultural land into
watersheds where amphibians live and reproduce. Mammals drinking nitrate- and nitrite-
contaminated water exhibit decreased gonadal steroid synthesis after only relatively brief
exposure periods. Despite findings of abnormal growth and metamorphosis in tadpoles exposed
to nitrate, no study has investigated whether nitrate alters endocrine function in juvenile or adult
amphibians. Furthermore, most studies focus on the effects of lethal rather than sublethal
concentration of nitrate on amphibians. My study examined the effects of sublethal
concentrations of nitrate and nitrite on plasma steroids, gonadal steroid synthesis, and growth of
reproductive tissues in amphibians. If nitrate or nitrite exposure alters endocrine function,
specifically reproductive steroids, then these contaminants should be considered as an important
factor to consider in amphibian reproduction and population declines.

Table 1-1. The molecular formula weight (MFW) of sodium nitrate (NaN03) and sodium nitrite (NaN02), the MFW percent of sodium (Na),
nitrate (NO3), nitrite (NO2), and nitrogen (N), and the equivalent concentrations of nitrate, nitrite, nitrate as nitrogen (NO3-N), and
Percent
sodium
(Na)
Percent
nitrate
(NO3)
Percent
nitrogen
(N)
NaN03
(mg/L)
Nitrate
N03
(mg/L)
Nitrate as
nitrogen
(NO3-N)
(mg/L)
Nitrate
N03
(mM)
Nitrate as
nitrogen
(NO3-N)
(mM)
0
0.00
0.00
0.00
0.00
Sodium
1
0.73
0.17
0.009
0.002
nitrate
10
7.30
1.65
0.01
0.003
27.0 %
73.0%
16.5 %
40
29.20
6.60
0.34
0.08
(NaN03)
100
73.00
16.50
0.86
0.19
FW 84.99 g
150
109.50
24.75
1.29
0.29
200
146.00
33.00
1.72
0.39
300
219.00
49.50
2.58
0.58
Percent
sodium
(Na)
Percent
nitrite
(N02)
Percent
nitrogen
(N)
NaN02
(mg/L)
Nitrite
no2
(mg/L)
Nitrite as
nitrogen
(N02-N)
(mg/L)
Nitrite
no2
(mM)
Nitrite as
nitrogen
(N02-N)
(mM)
Sodium
nitrite
(NaN02)
FW 68.99 g
0
0.00
0.00
0.00
0.00
1
0.67
0.20
0.01
0.003
10
6.67
2.03
0.10
0.03
40
26.68
8.12
2.90
0.12
100
66.70
20.30
0.39
0.29
150
100.05
30.45
0.97
0.44
200
133.40
40.60
1.45
0.59
300
200.10
60.90
2.90
0.88
33.3 %
66.7 %
20.3 %

Table 1-2. A comparison among amphibians of the effects of nitrate and nitrite
Species Stage Treatment End Point
Ambystoma gracile Larvae 0.78-25 mg/L nitrate Decreased feeding & activity, bent tails, edema
4 mg/L nitrite LC50 < 15 days
Bufo americanus
Tadpoles
Acute: 13.6-39.3 mg/L nitrate
Chronic: 2-10 mg/L nitrate
B. bóreas
Tadpoles
0.78-25 mg/L nitrate
4 mg/L nitrite
B. bufo
Tadpoles
385 mg/L
Hyla regilla
Tadpoles
0.78-25 mg/L nitrate
4 mg/L nitrite
Litoria caerulea
Tadpoles
9-22.6 mg/L nitrate
Pseudacris triseriata
Tadpoles
Acute: 13.6-39.3 mg/L nitrate
Chronic: 2-10 mg/L nitrate
Rana aurora
Tadpoles
0.78-25 mg/L nitrate
4 mg/L nitrite
R. cascadae
Tadpoles
3.5 mg/L nitrate
R. catesbeiana
Tadpoles
9-26 mg/L nitrate
R. pipiens
Tadpoles
9-26 mg/L nitrate
R. pipiens
Tadpoles
Acute: 13.6-39.3 mg/L nitrate
Chronic - 2-10 mg/L nitrate
R. clamitans
Tadpoles
Acute: 13.6-39.3 mg/L nitrate
Chronic: 2-10 mg/L nitrate
R. pretiosa
Tadpoles
0.78-25 mg/L nitrate
4 mg/L nitrite
R. temporaria
Tadpoles
5 mg/L nitrate
R. temporaria
Adults
3.6-6.9 g/m2 nitrate on
substrate
LC50 96 h, decreased swimming and feeding
Bent tail, edema, head and digestive deformities
Decreased feeding & activity, bent tails, edema
LC50 < 15 days
LC50 96 h
Decreased feeding & activity, bent tails, edema
LC50 < 15 days
Decreased growth rates, behavior abnormalities, increased
mortality
LC50 96 h, decreased swimming and feeding, bent tail
Edema, head and digestive deformities
Decreased feeding & activity, bent tails, edema
LC50 < 15 days
Decreased rates metamorphosis at earlier stage development
Decreased white blood cells and hemoglobin
Decreased white blood cells and hemoglobin
LC50 96 h, decreased swimming and feeding, bent tail
Edema, head and digestive deformities
LC5o 96 h, decreased swimming and feeding, bent tail
Edema, head and digestive deformities
decreased feeding & activity, bent tails, edema
LC50 < 15 days
Decreased growth rates and decreased size at
metamorphosis
Increased toxicity and mortality
Reference
Marco and Blaustein, 1999
Hecnar, 1995
Marco and Blaustein, 1999
Xu and Oldham, 1997
Marco and Blaustein, 1999
Baker and Waights, 1994
Hecnar, 1995
Marco and Blaustein, 1999
Marco and Blaustein, 1999
Dappen, 1983
Dappen, 1983
Hecnar, 1995
Hecnar, 1995
Marco and Blaustein, 1999
Johansson et al., 2001
Oldham et al., 1997

CHAPTER 2
THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS OF NITRATE (IN VIVO) ON PLASMA STEROIDS AND INSULIN¬
LIKE GROWTH FACTOR-1, ON OVARIAN STEROID SYNTHESIS, AND ON OVIDUCT
GROWTH IN THE AFRICAN CLAWED FROG (.Xenopus laevis)
Introduction
During the last few years there has been increased global concern over contamination of
water by anthropogenic sources of nitrates. Nitrate is among the most stable, water-soluble ionic
forms of nitrogen persistent in aquatic habitats. Nitrate contaminates watersheds in agricultural
and urban environments, reaching harmful concentrations largely due to human activities. In
agricultural areas, nitrate contaminates watersheds primarily through poorly regulated runoff of
nitrogen-based fertilizers and animal wastes from farms. In urban areas, nitrate contaminates
watersheds primarily through release of industrial and wastewater effluent from treatment plants,
runoff of fertilizers applied to lawns and golf courses, and air pollution from the burning of fossil
fuels (Pucket, 1995; Rouse et al., 1999). In temperate North America, concentrations of aquatic
nitrate are highest between the fall and spring when reduced ion uptake by agricultural plants
increases soil nitrate loads leaching from the ground (Hallberg, 1989; Nolen and Stoner, 1995;
Nolen et al., 1995, 1997). Additionally, fertilizers applied in close proximity to watersheds,
coupled with spring rainstorms, contributes to an overwhelming aquatic nitrate pulse that
frequently exceeds 100 mg/L and overlaps the breeding season of many amphibians (Rouse et al.,
1999). Many studies on the effects of nitrate on amphibians have addressed the effects of
toxicological rather than sublethal doses on growth, skeletal, and tissue deformities in juvenile
amphibians (Cooke, 1981; Baker and Waights, 1993; Hecnar, 1995; Watt and Oldham, 1995;
Oldham et al., 1997; Xu and Oldham, 1997; March and Blaustein, 1998; Marco et al., 1999;
15

16
Johansson et al., 2001; Chapter 1, Table 1-2). Surprisingly, few studies have investigated effects
of exposure to sublethal nitrate concentrations on adult, reproductive frogs.
There is mounting evidence that nitrate interferes with steroid-signaling pathways.
Panesar and Chan (Panesar, 1999; Panesar and Chan, 2000) demonstrated that administration of
nitrate and nitrite inhibits testosterone (T) synthesis (in vitro and in vivo) in rodents. Once nitrate
enters the body, through consumption or absorption across skin surfaces, it can be converted into
nitrite by endogenous microbial activity in the mouth or gastrointestinal tract (Fried, 1991;
Doblander and Lackner, 1996). Nitrite can be converted into N-nitrosoamines, which are
carcinogens in laboratory animals and in humans (National Academy of Sciences, 1981; Tricker
and Preussmann. 1991; US EPA, 1995). One proposed mechanism for altered steroid expression
by nitrates involves enzyme-dependent synthesis of nitric oxide (NO) (Panesar and Chan, 2000).
The NO is synthesized (in vivo) from an L-arginine precursor by nitric oxide synthase (NOS)
enzymes (Kleinert et al., 1995; Mayer and Hemmenns, 1997). In addition to NOS-dependent NO
formation, non-enzymatic synthesis of NO can also occur through acidic reduction of nitrite
(Iizuka et al., 1999; Zweier et al., 1995, 1999; Modin et al., 2001). Cosby et al. (2003) reported
that hemoglobin functions as a nitrite reductase contributing to enzyme-independent NO
synthesis. Regardless of the mechanisms by which it is produced, NO is thought to regulate many
physiological processes. Zweier et al. (1999) reported that enzyme-independent NO formation is
associated with cellular damage and loss of organ function. Panesar and Chan (2000) proposed
that, in steroidogenic tissues, NO binds to the heme groups inherent to mitochondrial cytochrome
P450 enzymes, such as those involved in side-chain cleavage (P450scc): the rate-limiting step in
steroid synthesis. The NO can inhibit other P450 enzymes, such as 3P-dehydroxysteroid
dehydrogenase (3P-HSD) involved in androgen synthesis; and P450 aromatase (Snyder et al.,
1996) involved in aromatization of androgens to estrogens. Collectively, these P450 enzymes are

17
necessary for conversion of free cholesterol into progesterone (P4): the steroid precursor for T and
17P-estradiol (E2).
Various isoforms of NOS are found within the ovary and other steroidogenic tissues in
vertebrates (Szabo and Thiemermann. 1995; Van Voorhis et al., 1995; Srivastava et al., 1997).
Disruption of these enzymes by NO might inhibit P4 synthesis, which would decrease or prevent
downstream T synthesis. Inhibition of gonadal T synthesis likely reduces the T available for
aromatase conversion to E2 and would contribute to decreased overall gonadal E2 synthesis. This
speculation is supported by studies in mammals demonstrating that increased NOS activity and
NO concentrations are associated with decreased ovarian E2 synthesis (VanVoorhis et al., 1994,
1995; Jablonka-Shariff and Olsen, 1997; Srivastava et al., 1997; Dees et al., 2000).
Relatively few studies have reported the impact of nitrate on steroidogenesis, but no
study has investigated the effect of nitrate exposure on of insulin-like growth factor-1 (IGF-1) in
vertebrates. Insulin-like growth factor-1 is a potent growth-stimulating hormone that regulates
bone and skeletal muscle growth, limb bud emergence, reproductive and somatic tissue growth,
steroidogenesis, and other physiological functions (Daughaday and Rotwein, 1989; Erickson et
al., 1989; Adashi, 1993; Hiney et al., 1996; Olsen et al., 1996; Dees et al., 1998; Kaliman et al.,
1999; Allen et al., 2001). Thus, IGF-1 is a relevant hormone to examine in the cases of amphibian
skeletal deformities, sex ratio reversal, and reproductive abnormalities. Abnormal expression of
IGF-1 is associated with altered growth and function of reproductive tissues in vertebrates.
Increased concentrations of plasma IGF-1 in humans is positively correlated with cancer of the
endometrium, breast, prostate, skin, pancreas, lung, and colon (Cohen et al., 1991; Lippman,
1993; Papa et al., 1993; LeRoith et al., 1993, 1995; Werner and LeRoith, 1996; Cascinu et al.,
1997; Mantzoros et al., 1997; Stoll, 1997). Despite these reports, IGF-1 can have beneficial
effects on tissue growth and function. For example, IGF-1 also mediates growth of E2- sensitive
reproductive tissues. In addition to this, IGF-1 regulates gonadal steroid expression. Intraovarian
IGF-1 expression counteracts NO-induced steroid inhibition by increasing aromatase activity and

18
stimulating E2 synthesis (Daughaday and Rotwein, 1989; Erickson et al., 1989; Adashi, 1993;
Hiney et al., 1996; Olsen et al., 1996; Dees et al., 1998). Furthermore, evidence indicates that NO
stimulates ovarian IGF-1 expression (Dees et al., 1998). Thus, NO interacts with the IGF-1
system and influences expression of steroids and also their actions in reproductive tissues.
Based on the aforementioned studies, I hypothesized that nitrate alters concentrations of
steroids and IGF-1, and alters oviduct growth in a model frog species, Xenopus laevis. My study
tested this hypothesis using environmentally relevant concentrations of nitrate.
Materials and Methods
Animals and Samples
Adult female X. laevis were purchased from Xenopus Express (Plant City, Florida). This
species is entirely aquatic, and thus would remain in constant exposure to administered
treatments. Frogs were maintained under a 12-h light/dark cycle in 38 L tanks with 19 L of static-
flow, dechlorinated water at 23°C (pH 7.0 - 7.4), with ammonia and nitrite content below 1.0
mg/L as confirmed by daily water measurements. Animals were fed spirulina pellets (Aquatic
Ecosystems, Orlando, FL) every other day for the duration of the experiment. All procedures
were performed with approval of the University of Florida Institute of Animal Care and Use
Committee (IACUC Permit #Z023). Pregnant mare serum gonadotropin (PMSG) and human
chorionic gonadotropin (hCG) were obtained from Sigma-Aldrich (St. Louis, MO), and sodium
nitrate (99% purity) was obtained from Fisher Scientific (Orlando, FL).
Nitrate Study Design
Treatment groups were divided into control (0 mg/L), 150 mg/L, and 300 mg/L sodium
nitrate; respectively equivalent to 0, 24.75, and 49.50 mg/L nitrate-as-nitrogen (N03-N). Nitrate
as nitrogen represents the concentration of nitrogen present in a given concentration of sodium
nitrate administered (Chapter 1, Table 1-1). For the remainder of this chapter, nitrate will refer to
NOj-N.

19
The frogs were randomly assigned to each of 3 replicate tanks per treatment for a total
sample size of 12 frogs per treatment. No significant differences in mass were detected (ANOVA;
P > 0.05) or snout-vent-length (SVL; ANOVA; P > 0.05) among frogs in each treatment group.
After a 1-week acclimation period, frogs were injected into the dorsal lymph sac with 50 IU of
PMSG, followed 3 days later by an injection of 750 IU hCG. These treatments stimulated
ovulation and formation of new ovarian follicles within 6 weeks (Dumont, 1971; Fortune and
Tsang, 1981; Fortune, 1983). This procedure synchronized the size and maturation of new
follicles before nitrate exposure, and minimized possible variation in gonadal steroid synthesis
among frogs in response to treatment.
After 6-weeks, frogs were exposed to nitrate applied to tank water for 7 consecutive days.
Every 24 h, water was changed, and fresh water with nitrate was added. After 7 days, the frogs
were anesthetized with MS-222 (1.5% 3-aminobenzoic acid ethyl-ether, Aquatic Ecosystems,
Orlando, FL). Blood was collected by cardiac puncture using heparinized syringes, placed into
heparin vacutainer tubes, and centrifuged (2500xG) for 15 min; and plasma was stored at -70°C
for E2, T, and IGF-1 radioimmunoassay (RIA) analysis. The ovaries were removed and weighed,
and follicles were dissected for a culture study (ex vivo). Follicles of specific maturation stages
were chosen: stage 4 follicles synthesize E2, and stage 5 and 6 follicles synthesize T (Fortune and
Tsang, 1981; Fortune, 1983;). From each frog, 33 follicles, each of stages 4, 5, and 6 were
incubated in 35x10 mm sterile culture dishes, in duplicate, at 23°C with 2 mL of sterile, phenol-
free culture media (1L M199 HBSS, 3.4 mL 200 mM L-glutamine, 5.96 g/L HEPES, 0.35 g/L
sodium bicarbonate, 8.0 mL 0.1 mM IBMX, pH 6.9; Sigma-Aldrich, St. Louis, MO) for both 5
and 10 h. Follicles were incubated at the same temperature Incubation temperature was selected
based on the water temperature maintained in the tanks holding X. laevis. After incubation,
culture media was decanted, flash-frozen, and stored at -70°C for E2 and T RIA. The diameter of
the remaining, uncultured follicles was measured with a dissecting microscope and an ocular

20
micrometer. For each follicle stage, 5 follicles (un-cultured) were measured in each frog from 0
mg/L (control, N = 8), 24.75 mg/L nitrate (N = 10), and 49.50 mg/L nitrate (N = 8) treatment
groups. Sample sizes of frogs were uneven among treatment groups for follicle measurements
because for some frogs, all of the follicles were incubated in the culture study. Ovary, liver, and
oviduct weights were recorded to compare post-treatment tissue weights among groups.
Steroid Radioimmunoassay (RIA) Procedures
RIAs were performed for E2 and T (Guillette et al., 1994; Guillette et al., 1996) on culture
media and on plasma samples using validated procedures. Duplicate media samples or plasma (50
pL for E2 T) were extracted twice with ethyl-ether, air-dried, and reconstituted in borate buffer
(0.05 M; pH 8.0). Antibody (Endocrine Sciences) was added at a final concentration of 1:55,000
for E2 and of 1:25,000 for T. Radiolabeled steroid ([2,4,6,7,16,17-3H] estradiol at 1 mCi/mL;
[1,2,6,7-3H] and testosterone at 1 mCi/mL; Amersham Int., Arlington Heights, IL) was added at
12,000 cpm per 100 pL for a final assay volume of 500 pL. Interassay variance tubes were
prepared from two separate pools of media and of plasma for E2 and T. Standards for E2 and T
were prepared in duplicate at 0, 1.56, 3.13, 6.25, 12.5, 25, 50, 100, 200, 400, and 800 pg/tube.
Assay tubes were vortexed and incubated overnight at 4°C.
Bound-free separation was performed using a mixture of 5.0% charcoal to 0.5% dextran,
pulse-vortexing, and centrifuging tubes (1500g, 4°C, 30 min). Supernatant was added to 5 mL of
scintillation cocktail, and counted. Media intraassay and interassay variance averaged 2.50% and
3.70% for E2, and 4.20% and 8.38% for T, respectively. Plasma intraassay variance for E2 and T
averaged 4.20% and 4.60%, respectively. Plasma E2 and T samples were run in a single assay and
interassay variances are not reported.
Validation of the steroid assays included media and plasma dilutions (50, 100, and 200
pL for E2 and 20, 50 and 100 pL for T) compared with E2 and T standards.

21
Insulin-Like Growth Factor-1 RIA Procedures
The IGF-1 RIA was performed as described by Crain et al. (1995). The National
Hormone and Pituitary Program (Torrance, CA 90509) supplied human recombinant IGF-1
standard (9.76 to 2500 pg/tube) and human IGF-1 antisera (Lot # AFP4892898, 1:400,000 final
dilution). The antiserum had less than 1.0% cross-reactivity with human IGF-II. Iodinated IGF-1
label (IGF-11125 sp act 2000 Ci/mmol; 16,000 cpm/tube) and Amerlex-M donkey anti-rabbit
secondary antibody (RPN510) were obtained from Amersham International (Arlington Heights,
IL).
For each treatment group, plasma was pooled (8, 16, 24, and 36 p.L aliquots in borate
buffer) for validation using plasma dilutions (equivalent to 1.9, 3.9, 5.7, and 8.3 pL plasma) that
were compared with IGF-1 standard. Plasma validation and experimental samples (20 pL) were
acid-ethanol extracted and IGF-1 RIA performed (Crain et al., 1995). Validation samples were
run in one assay with intraassay variance averaging 3.10%.
Biochemical RIA Validations
Plasma dilutions and internal standards were parallel to E2 standards (ANCOVA; F =
0.48; P = 0.52 and F = 0.35; P = 57, Fig. 2-1 A) and recovery of E2 after extraction was 81.0%.
Plasma dilutions and internal standards were parallel to T standards exhibited parallel
displacement (ANCOVA; F = 0.12, P = 0.33 and F = 1.18, P = 0.31, Fig. 2-1B) and recovery of T
after extraction was 93.8%. Plasma dilutions and IGF-1 standards exhibited parallel displacement
curves (ANCOVA; F = 0.08; P = 0.79, Fig. 2-1C) and recovery of IGF-1 after extraction was
78.0%.
Media dilutions and E2 standards gave parallel displacement curves (ANCOVA; F =
1.05; P = 0.37, Fig. 2-2A). Recovery of E2 after media extraction was 91.5% and all sample
values were corrected for loss using this value. Media dilutions and T standards gave parallel
displacement curves (ANCOVA; F = 0.60; P = 0.48, Fig. 2-2B). Recovery of T after media

22
extraction averaged 98.9% and all sample values were corrected for this loss. For subsequent
steroid and IGF-1 analyses, all sample values were corrected for respective losses.
Statistics
Ovary, oviduct, and liver wet mass were compared among treatment groups with body
mass as a covariate using ANCOVA, followed by LSD post-hoc contrasts. Concentrations of E2,
T, and IGF-1 were estimated from raw data using Microplate Manager software (Microplate
Manager III, BioRad Laboratories, Inc., Hercules, CA, 1988). Statistical analyses were performed
using SPSS software (v. 10, SPSS Inc., Chicago, IL, 1999) with a = 0.05. ANCOVA was used to
validate plasma and media samples and to determine if plasma IGF-1 concentrations were
correlated to body mass. Concentrations of E2, T, and IGF-1 among replicate tanks within each
treatment group were compared using one-way ANOVA. Where no significant difference existed
among replicate tanks within treatment groups, mean E2, T, and IGF-1 concentrations were
compared among treatment groups with one-way ANOVA. Ovarian follicle diameters were
compared, separately according to stage, among treatment groups with one-way ANOVA.
Following one-way ANOVA analyses Scheffe post-hoc contrasts were used. Tamhane post-hoc
contrasts were used where variances were unequal among groups for plasma IGF-1
concentrations.
Results
Tissue Weights
Tissue weights were not different among treatment groups for ovary (ANCOVA; F =
0.57, P = 0.57), oviduct (ANCOVA; F = 0.28, P = 0.76), and liver (ANCOVA; F = 1.13, P =
0.34).
Follicle Diameters
Diameter of stage 4 follicles was larger (ANOVA; P = 0.01, Fig. 2-3 A) in frogs exposed
to 24.75 mg/L and 49.50 mg/L nitrate relative to frogs exposed to 0 mg/L. Mean diameter was
smaller in stage 5 (ANOVA; P = 0.04, Fig. 2-3B) and stage 6 follicles (ANOVA; P = 0.005, Fig.

23
2-3C) in frogs exposed to 49.50 mg/L nitrate compared to frogs exposed to 24.75 mg/L nitrate
and 0 mg/L.
Plasma Steroid Concentrations
Analyses revealed no significant difference (P > 0.05) in E2 or T concentrations among
replicate tanks; thus data from frogs in replicate tanks was combined per treatment group. Plasma
E2 was not significantly different among treatment groups (ANOVA; P = 0.08). Plasma T did not
differ among treatment groups (ANOVA; P = 0.70).
Plasma IGF-1 Concentrations
Analyses revealed no significant difference in IGF-1 concentrations among replicate
tanks; thus data from frogs in replicate tanks was combined per treatment group. Plasma IGF-1
concentrations were significantly higher in frogs exposed to 24.75 mg/L and 49.50 mg/L nitrate
relative to the control frogs (ANOVA; P = 0.007, Fig. 2-4). Plasma IGF-1 was not significantly
correlated to body mass (ANOVA; R2 = 0.12, P > 0.05).
Ovarian Follicle Steroid Concentrations (Ex Vivo)
Statistical analyses revealed no significant difference in mean E2 or T (P > 0.05)
concentrations among replicates for each treatment group; thus, data from frogs in replicate tanks
was combined per treatment group. After 5 h, media E2 concentrations were significantly lower
for ovarian follicles of frogs exposed to 49.50 mg/L nitrate compared to the other treatment
groups (ANOVA; P < 0.001, Fig. 2-5A). However, after 10 h, media E2 concentrations were
significantly lower for ovarian follicles of frogs exposed to both the 24.75 mg/L and 49.50 mg/L
nitrate relative to the controls (ANOVA; P < 0.001, Fig. 2-5B).
After 5 h, media T concentrations were similar among treatment groups (ANOVA; P >
0.05, Fig. 2-6A). However, after 10 h, media T concentrations were significantly lower for
ovarian follicles from frogs exposed to 24.75 mg/L and 49.50 mg/L nitrate relative to the control
group (ANOVA; P < 0.001, Fig. 2-6B).

24
Discussion
This study has shown that exposure of A! laevis to sublethal doses of aquatic nitrates at
environmentally relevant concentrations (24.75 mg/L and 49.50 mg/L) is associated with
endocrine disruption of E2, T, and IGF-1. This study raises new and troubling questions regarding
the effects of nitrates on endocrine function. No other study has examined the effects of exposure
to sublethal concentrations of nitrate on adult anurans, despite reports of altered growth, behavior,
and mobility in tadpoles at similarly low (1-40 mg/L) nitrate concentrations (Baker and Waights,
1994; Hecnar, 1995; Xu and Oldham, 1997; Marco and Blaustein, 1999 Johansson et a!., 2001.
Over the past 30 years amphibian populations have declined in various regions of the
world (Wake, 1991; McCoy, 1994), especially in agricultural landscapes (Dappen, 1983; Berger,
1989; de Solía et al., 2002). Alteration of aquatic habitats is considered a primary contributor to
these declines (Blaustein and Wake, 1990; Carey and Bryant, 1995). Altered endocrine function
in frogs has been associated with exposure to sublethal concentrations of various contaminants
(Mohanty-Hejmadi and Dutta, 1981; Carey and Bryant, 1995; Reeder et al., 1998; Kloas et al.,
1999; Hayes et al., 2002). In agricultural and urban areas, contamination of aquatic habitats by
anthropogenic sources of nitrate poses a serious threat to wildlife and humans. Approximately 72
million tons of nitrogen-based fertilizers are used worldwide and, combined with release of
industrial nitrogenous wastes, are likely responsible for increased nitrate contamination reported
in surface waters, aquifers, and drinking water (Rouse et al., 1999). Most studies examining the
effects of sublethal nitrate concentrations on frogs have focused on juvenile stages from egg
through metamorphosing tadpole. There is an absence of research examining the effects of
sublethal nitrate concentrations on the endocrine profile of adult, reproductive frogs.
Panesar and Chan (2000) reported inhibition of T synthesis (in vitro) in rodents after
exposure to nitrate. Within body tissues, various isoforms of NOS enzymes are capable of
converting nitrates into NO (VanVoorhis et al., 1994, 1995; Srivastava et al., 1997; Olsen et al.,
1996; Jablonka-Shariff and Olson, 1997). In addition, acidic reduction and hemoglobin have been

25
shown to mediate non-enzymatic NO formation from nitrite (in vivo) (Zweier et al., 1995, 1999;
Modin et al., 2001; Cosby et al. 2003). Many studies have shown that NO inhibits E2 and T
synthesis in rodents, humans, and cows (VanVoorhis et al., 1994; Wang and Marsden, 1995;
Basini et al., 1998; Omura, 1999). Panesar and Chan (2000) proposed a mechanism (based on a
synthesis of their work and that of other researchers) involving formation of NO. Nitrate and
nitrite can be converted to NO within steroidogenic cells, and the NO inhibits steroidogenic P450
enzymes necessary for conversion of free cholesterol to steroid precursors. In addition to
inhibiting P450 enzymes, NO has also been shown to inhibit steroid-acute regulatory protein
(StAR) protein expression. During steroidogenesis, StAR protein is essential for transporting free
cholesterol to the inner mitochondrial membrane (Wang and Marsden, 1995). I propose a similar
nitrate-associated steroid inhibition, possibly involving NO formation, occurred within ovarian
follicles ofX laevis. This steroid inhibition also and includes downstream inhibition of E2
synthesis and stimulation of IGF-1.
In X. laevis exposed to nitrate ovarian steroid synthesis was inhibited (ex vivo) while
plasma steroid concentrations (in vivo) were unaffected. These findings indicate that different
mechanisms were involved in regulating ex vivo versus in vivo steroids in nitrate-exposed frogs. It
is possible plasma steroid concentrations were unchanged due to compensatory responses of the
hypothalamic-pituitary-gonadal (HPG) axis (Chapter 1, Fig. 1-1). Inhibition of steroid synthesis
at the gonad level might have signaled a compensatory hypothalamic release of gonadotropin¬
releasing hormone (GnRH) causing pituitary release of luteinizing hormone (LH) and follicle-
stimulating hormone (FSH) into the blood. Increased plasma LH/FSH concentrations would
stimulate ovarian synthesis of T and E2, which could have contributed to normal circulating
plasma steroid concentrations. In this study, ovarian ex vivo follicle steroid synthesis was
recorded without measuring corresponding plasma gonadotropins. It is unknown if plasma steroid
concentrations in nitrate-exposed frogs were maintained at levels similar to control frogs by
compensation by the HPG axis. It is unlikely that compensatory responses of the HPG axis to

26
stimulate steroidogenesis by the gonads would influence plasma steroid concentrations because
ovarian steroid synthesis was shown to be inhibited in nitrate-exposed frogs. Thus, gonadotropins
would not be effective in stimulating steroid synthesis in nitrate-exposed frogs when
steroidogenesis is inhibited at the level of the gonad. Therefore, another explanation must be
considered.
The liver is the main organ for degradation of nitrate, and degradation of nitrate can
elevate hepatic NO concentrations. Continuous administration of nitrate has been shown to
increase hepatic NO synthesis and inhibit hepatic P450 enzymes activity (Minamiyama et al.,
2004). Hepatic P450 enzymes are necessary for metabolism and excretion of circulating steroids.
Thus, hepatic nitrate degradation can lead to NO formation and inhibition of hepatic P450 steroid
metabolic enzymes. Reduced hepatic steroid metabolism could cause stasis or even augmentation
of circulating steroid concentrations.
I propose a mechanism for the increase in plasma IGF-1 concentrations (in vivo)
observed in nitrate-exposed X. laevis. Nitrate, once consumed or absorbed across skin surfaces,
can be converted by microbial activity in the mouth and gastrointestinal tract to nitrite. Nitrite has
been shown to stimulate hypothalamic NO formation and increase hypothalamic secretion of
growth hormone-releasing hormone GHRH and pituitary release of growth hormone (GH) (de
Caceres et al., 2003). Thus, the hypothalamic-pituitary-hepatic (HPH) axis regulates circulating
IGF-1 concentrations, and this axis is influenced by nitrite and NO exposure (Fig. 2-7). Further
research will be necessary to confirm the validity of this proposed pathway.
In addition to the liver, the ovary also produces IGF-1, although in relatively smaller
quantities (Adashi, 1993). Stimulation of the ovary by pituitary FSH results in decreased
synthesis of IGF-1-binding proteins and increased intraovarian IGF-1 synthesis and availability.
Intraovarian IGF-1 might have an autocrine and endocrine effect of ovarian steroid synthesis
(Grimes et al., 1992; Adashi, 1993; Basini et al., 1998). Increased IGF-1 has been shown to
increase intraovarian aromatase activity and E2 synthesis (Erickson et al., 1989; Monnieaux and

27
Pisselet, 1992; Adashi, 1993; Samaras et al., 1994; Samaras et al., 1996). However, plasma IGF-1
concentrations increased and ovarian E2 concentrations decreased in X. laevis upon nitrate
exposure. Perhaps the in vivo nitrate exposure period of 7 days was too brief to observe a
compensatory increase in ovarian steroid synthesis with IGF-1 stimulation.
Plasma IGF-1 binding proteins (IGF-BP) play an important role in regulating the
availability of IGF-1 to and within tissues. In this study, IGF-BP in the plasma and the ovaries
were not measured, so the availability of increased plasma IGF-1 in nitrate-exposed animals
merits investigation. If plasma IGF-1 increased in conjunction with a decrease in tissue IGF1-BP,
then there might be an increase in IGF-1 utilization and growth response by tissues. In this study,
there was no difference detected in ovary, oviduct, or liver tissue mass among nitrate treatment
groups. This might indicate either that the increased circulating IGF-1 was not stimulating a
growth response in these tissues or that circulating IGF-1 was bound to IGF-BP and unavailable
for tissue uptake. Although no difference in total ovary weights was detected among treatment
groups, follicle diameter varied among groups. The diameter of E2-producing follicles (stage 4)
were larger in nitrate-exposed frogs compared to control, which might reflect a growth response
to increased IGF-1 exposure or compensatory tissue growth in response to declining E2 levels.
The diameter of T-producing follicles (stage 5 and 6) was smaller in frogs exposed to 49.50 mg/L
nitrate compared to follicles of frogs exposed to 24.75 mg/L and 0 mg/L. This could reflect either
the absence of IGF-1 uptake by these follicles or an absence of a growth-response to IGF-1.
This study raises new and troubling questions regarding the effects of nitrates on
endocrine function in vertebrates. Chemical alteration of aquatic habitats is considered a foremost
contributor to the declines and deformities reported for amphibian populations (Carey and Bryant,
1995; Wake, 1998; Hayes et al., 2002). Amphibians exposed to various contaminants, even at
sublethal concentrations, exhibit malformations, reproductive abnormalities, sex ratio reversal,
male feminization, and altered endocrine function (Reeder et al., 1998; Kloas et al., 1999; Hayes
et al., 2002). The nitrate-associated endocrine disruption in X. laevis might differ from other

28
anuran species due to the interspecific variation in physiological response to nitrates (Chapter 1,
Table 1-2). Further research is necessary to determine whether nitrate alters steroid and IGF-1
hormones in other anuran species, and to describe the range of sublethal nitrate concentrations
capable of endocrine disruption. The nitrate concentrations used in this study were relevant to
environmental concentrations measured in North American ground and surface water (Rouse et
al., 1999; Nolen and Stoner. 1995). Fiowever, it would be valuable to ascertain if even lower
nitrate concentrations have a similar endocrine disrupting capacity in frogs.
More research is needed to elucidate the mechanism by which nitrate inhibits steroid
synthesis and increases circulating IGF-1 concentrations in amphibians and in other animals.
Thus far, most reports of steroid inhibition by nitrate have focused on steroid synthesis and
regulation exclusively at the gonad level. It is important to consider both upstream and
downstream steroid regulation. Upstream regulation would changes in hypothalamic and pituitary
hormone secretions in response to in vivo nitrate exposure. The important hormones to examine
include hypothalamic GnRH and GHRH, and pituitary LH, FSH, and GH. Pituitary LH and FSH
function in stimulating gonadal steroid synthesis and GH stimulates hepatic IGF-1 synthesis.
Downstream regulation would include hepatic degradation and clearance of circulating steroids,
and secretion of IGF-1. In addition to these topics, it is important to determine whether amphibian
gonadal tissue contains NOS enzymes capable of synthesizing NO. It has been already been
established that the amphibian brain contains NOS capable of generating NO (McLean et al.,
2001; Gonzalez et al., 2002; McLean and Bilar, 2002). Furthermore, it is necessary to understand
how nitrate exposure of frogs regulates intracellular expression of NOS, NO, steroidogenic
enzymes, and steroid regulatory proteins. Lastly, since nitrate exposure is associated with changes
in circulating IGF-1 concentrations, in addition to steroid synthesis, is vital to understand how
these hormones collectively influence the reproductive physiology of amphibians.

(B/Bo) X 100 O (B/Bo) X 100 W (B/Bo) X 100
29
A 100 o o o
80
60
40
20
0
° Standard
o â–  Internal Standards
■ o ♦ 4 Plasma Dilutions
o*
1
10 100
E2 (pg)
1000
100
80
60
40
20
0
° Standard
â–  Internal Standards
♦ Plasma Dilutions
-On
10 100
T (pg)
1000
100
80
60
40
20
0
«D
° Standard
♦ Plasma Dilutions
o
1
10 100 1000 10000
IGF-1 (pg)
Figure 2-1. Biochemical validation ofXenopus laevis plasma. A. estradiol RIA internal standards
(ANCOVA; F = 0.35; P = 0.57) and plasma dilutions (ANCOVA; F = 1.86; P =
0.23) were parallel to the standard curve. B. testosterone RIA. Internal standards
(ANCOVA; F = 1.18; P = 0.31) and plasma dilutions (ANCOVA; F = 0.12; P =
0.33) were parallel to the standard curve. C. IGF-1 RIA. plasma dilutions
(ANCOVA; F = 1.05; P = 0.37) were parallel to the standard curve.

(B/Bo) X 100 a (B/Bo) X 100
30
A 100 o
80
60
40
20
0
° Standard
♦ Media Dilutions
o
O o
1
10 100
E2 (pg)
1000
100
80
60
40
20
0
° Standard
o
^ ♦ Media Dilutions
*>
o
o
1 10 100 1000
T (pg)
Figure 2-2. Biochemical validation of Xenopus laevis media. A. estradiol R1A media dilutions
(ANCOVA; F = 0.08; P = 0.79) were parallel to the standard curve. B. testosterone
RIA. Media dilutions (ANCOVA; F = 0.60; P = 0.48) were parallel to the standard
curve.

Follicle Diameter (^m) O Follicle Diameter ( ^m) 03 Follicle Diameter ( ^m) >
31
Stage 4
1.40
Stage 5
a
1.30
1.24
1.18
1.12
1.06
1.00
o o o
o o o
o o
o
o
o
o
o
CN
r-
Tt-’
o
o
o
ro
Nitrate (mg/L)
Figure 2-3. Diameter of ovarian follicles of stages 4, 5, and 6 in Xenopus laevis exposed in vivo
for 7 days to 0, 24.75, and 49.50 mg/L nitrate. Data presented as means ± SEM.
Different letters above bars indicate significant differences for: A. stage four
(ANOVA; P = 0.01), B. stage five (ANOVA; P = 0.040, and C. stage six (ANOVA;
P = 0.005).

IGF-1 (ng/mL)
32
60
50
40
30
20
10
0
10
nh
12
12
o
o
o
O
o
in
o
o
o
o
o
q
q
q
q
O;
O
q
o
in
o
in
o
n
d
d
in
d
ON
» <
(N
(N
m
Nitrate (mg/L)
Figure 2-4. Plasma insulin-like-growth factor-1 (IGF-1) in Xenopus laevis after 7 days of in vivo
exposure to 0, 24.75, and 49.50 mg/L nitrate. Data presented as means ± SEM.
Numbers within bars indicate sample sizes and different letters above bars indicate
significant differences (ANOVA; P = 0.007).

Media Estradiol (pg/mL) ro Media Estradiol (pg/mL) >
33
350
300
250
200
150
100
50
0
350
300
250
200
150
100
50
0
11
o
o
a
o
o
in
11
5 h
a
A
12
12
o
o
o
o
>n
o
o
o
o
o
>n
o
o
o
(N
in
â– 't
o
o
o
en
o
o
l/i
en
o
q
o
b
-i-
11
o
o
o
in
o
©
©
o
o
q
q
q
t -
q
o
q
q
m
©
in
©
o
m
©
in
Os
10 h
m
m
rt-
11
o
q
in
â–  o
»n
Os
Nitrate (mg/L)
Figure 2-5. Media 17P-estradiol concentrations in culture media from incubated ovarian follicles
of Xenopus laevis after 7 days in vivo exposure to 0, 24.75, and 49.50 mg/L nitrate.
Data presented as means ± SEM for A. 5 h and B. 10 h of incubation. Numbers within
bars indicate sample sizes and different letters above bars indicate significant
differences for B. (ANOVA; P < 0.001).

Testosterone (pg/mL) 00 Testosterone (pg/mL)
34
A
800
600
400
200
0
o
o
o
o
o
in
o
o
o
o
o
o
o
o
o
o
N-
o
o
o
o
in
d
in
d
in
d
â– sf
d
in
d
in
d
T—
T—
(N
CM
00
CO
â– sj-
2000
1600
1200
800
400
0
11
10 h
12
11
o
o
o
o
o
in
O
o
o
o
o
o
o
o
o
o
O
o
o
o
in
d
in
d
in
d
â– St
d
in
d
in
d
T—
■*—
CM
CM
00
00
â– sr
Nitrate (mg/L)
Figure 2-6. Media testosterone concentrations in culture media from incubated ovarian follicles of
Xenopus laevis after 7 days in vivo exposure to 0, 24.75, and 49.50 mg/L nitrate. Data
presented as means ± SEM. Numbers within bars indicate sample sizes and different
letters above bars indicate significant differences for A. 5 h (ANOVA; P = 0.007)
and B. 10 h (ANOVA; P < 0.001) of incubation.

35
Hypothalamus
NO
Nitrite Nitrate
Figure 2-7. Diagram of mechanism for nitrate-associated inhibition of steroidogenesis and
increased plasma IGF-1. Only ovarian testosterone, (T), estradiol 17P (E2), and
plasma T, E2, and insulin-like growth factor-1 (IGF-1) were measured in Xenopus
laevis exposed (in vivo) for 7 days to 0, 24.75, and 49.50 mg/L nitrate (NO3-N). Other
parameters are adapted from other studies (Licht 1984; Panesar and Chan, 2000; de
Caceres et al., 2003; Minamiyama et al., 2004). A. Ovarian steroid synthesis is
inhibited by nitric oxide (NO) formation from nitrate and nitrite. The NO inhibits
cytochrome P450 steroidogenic enzymes. NO might also inhibit steroid-acute
regulatory (StAR) protein which escorts free cholesterol into the mitochondria.
Inhibition of these enzymes reduces progesterone (P4) synthesis, and reduces T
available for aromatization (Arom)to E2. B. Decreased steroid synthesis could signal
compensatory hypothalamic secretion of gonadotropin-releasing hormone (GnRH)
pituitary secretion of luteinizing hormone (LH) and follicle stimulating hormone
(FSH). C. Hepatic nitrate metabolism can cause NO inhibition of P450 enzymes
involved in hepatic steroid metabolism and clearance resulting in augmented
circulating steroid concentrations. D. Nitrate and nitrite could cause hypothalamic NO
formation, which can stimulate secretion of growth hormone-releasing hormone
(GHRH) and secretion of pituitary growth hormone (GH). The GH stimulates liver
IGF-1 synthesis and secretion into the blood.

CHAPTER 3
SEASONAL CHANGES IN INSULIN-LIKE GROWTH FACTOR-1, STEROIDS, AND
REPRODUCTIVE TISSUES IN PIG FROGS (Rana grylio)
Introduction
The sex steroids, 17p-estradiol (E2) and testosterone (T), regulate virtually every facet of
reproduction, and in ectotherms these hormones are responsive to changes in temperature, pH,
and photoperiod among other environmental factors (Licht, 1970; Feder and Burggren, 1992;
Norris, 1997; Kim et al„ 1998).
Only one comprehensive profile of the pattern of seasonal changes in circulating steroid
concentrations and changes in gonadal growth and maturation has been reported for a population
of wild bullfrogs, Rana catesbeiana (Licht et al., 1983). Female R. catesbeiana exhibited a
seasonal pattern of changes in plasma concentrations of gonadotropins and steroids, and in
relative weights of reproductive tissues that indicate reproductive and non-reproductive periods.
Reproductive period is here defined as physiological conditions that are optimal for reproduction,
such as elevated plasma concentrations of the reproductive steroids E2, T, and progesterone (P4),
and also by elevated weights of reproductive tissues such as the ovaries and oviducts.
Reproductive condition of the frogs was also discerned by elevated plasma concentrations of the
gonadotropins luteinizing hormone (LH), follicle stimulating hormone (FSH). In R. catesbeiana,
E2, T, and progesterone (P4) concentrations were greatest in the reproductive period between May
and July. The non-reproductive period of frogs is defined here as the physiological condition
marked by decreased plasma concentrations of steroids and gonadotropins, and decreased weights
of reproductive tissues. Licht et al. (1983) reported that plasma steroid concentrations and
gonadal-somatic index (GSI) declined sharply after July and remained depressed between August
and February, indicating the frogs were in non-reproductive condition. Similarly, plasma LH and
36

37
FSH concentrations declined precipitously by July of both years (Licht et al., 1983). Increased
plasma steroid concentrations were likely stimulated by the elevated plasma gonadotropin
observed. Elevated concentrations of LH stimulate gonadal steroidogenesis whereas elevated
concentrations of FSH stimulate increased ovarian mass or gonadal somatic index (GSI) in
females during the reproductive period (Licht, 1970, 1979; Norris, 1997). Plasma T
concentrations in females greatly exceeded that of E2 at all times, and plasma T concentrations
were highly correlated with ovarian developmental stage. The relatively high T concentrations
might serve as a circulating androgen pool for synthesis of E2 by aromatase activity in peripheral
tissues such as the brain, fat and skin, and even the oviduct (Follett and Redshaw, 1968). Plasma
androgen pools also might serve functions unrelated to E2 synthesis. For example, it has been
reported that T synthesized by Xenopus ovaries might function to stimulate oocyte development
directly through androgen receptors (Lutz et al., 2001).
In addition to steroid hormones, insulin-like growth factor-1 (IGF-1) regulates many
aspects of reproduction including gonadal function and steroidogenesis (Adashi et al., 1991;
Hammond et al., 1991). The presence of IGF-1 has been identified in representative animals from
all vertebrate classes and includes humans, cows, rodents, birds, alligators, turtles, fish, and
amphibians (Daughaday et al., 1985; Pancak-Roessler and Lee, 1990; Crain et al., 1995; Guillette
et al., 1996; Le Roith et al., 2001a,b; Table 3-1). IGF-1 is a polypeptide hormone that stimulates
cell growth in somatic and reproductive tissues and orchestrates many aspects of development,
metabolism, and steroidogenesis (LeRoth et al., 2001a,b). In response to pituitary growth
hormone (GH), the liver secretes IGF-1 into circulation complexed to IGF-1 binding proteins
(IGF-lBPs). IGF-1 interacts with IGF-1 receptors located on tissues throughout body. Although
initially described as an intermediate of GH action on skeletal muscle growth, more recently IGF-
1 has been recognized as hormonal regulator of many GH-independent cellular processes (Butler
and Le Roith, 2001; Le Roith et al., 2001a). Recent research has shown that IGF-1 synthesized
within endometrial and ovarian tissue functions as a paracrine and autocrine hormone (Adashi,

38
1993). Studies in mammals demonstrate that increased intraovarian IGF-1 increases ovarian P4
and E2 synthesis, as well as aromatase, and steroid-acute regulatory protein (StAR) protein
expression (Adashi et al., 1991; Adashi, 1993; Samaras et al., 1994, 1996; Devoto et al., 1999).
Aromatase is a cytochrome P450 enzyme necessary for converting androgens into estrogens, and
StAR proteins assist the entry of free cholesterol into the mitochondria to initiate steroid synthesis
in steroidogenic tissues. Collectively, these findings demonstrate that IGF-1 is an important
regulator of gonadal steroids. Other studies have shown that intraovarian IGF-1 regulates
selection of dominant follicles for ovulation in mammals (Adashi et al., 1991; Giudice, 1999).
These studies indicate that IGF-1 also plays a vital role in steroid synthesis, regulation, and
gonadal function.
Although the IGF-1 system has been described in mammals, comparatively few studies
have examined this system in non-mammalian vertebrates (Table 3-1). Oviparous vertebrates are
intriguing models for examining the role of IGF-1 in reproduction because they lack a prolonged
period of maternal and fetal chemical and nutritive interaction during embryonic development.
Nutrients and growth promoting substances, like IGF-1, must be sequestered into eggs before
oviposition and fertilization (Guillette et al., 1996). In a turtles, geckos, and alligators, the
presence of plasma IGF-1 has been confirmed and demonstrated to play an important role in
mediating reproduction (Daughaday et al., 1985; Cox and Guillette, 1995; Crain et al., 1995a,b;
Guillette et al., 1996). Cox and Guillette (1995) demonstrated that ovariectomized (lacking
endogenous E2) geckos, exhibited an estrogen-like proliferation of oviductal tissue in response to
treatment with IGF-1 implants. Additionally, plasma IGF-1 concentrations vary according to
season and stages of reproductive maturation in female alligators and turtles (Crain et al., 1995;
Guillette et al., 1996). These studies indicate IGF-1 plays a more important role in the growth of
reproductive tissues than previously realized.
Unfortunately, the importance of IGF-1 in amphibian reproduction and growth remains
largely under-investigated (Daughaday et al., 1985; Pancak-Roessler and Lee, 1990; Table 3-1).

39
Only one study reported seasonal changes of plasma IGF-1 concentrations in a wild population of
Bufo woodhousei (Pancak-Roessler and Lee, 1990). Although this study was limited to a 10-
month profile, it was evident that IGF-1 concentrations peaked during the reproductive period
between May and June and decreased during the non-reproductive period between August and
December (Pancak-Roessler and Lee, 1990).
No study has provided a simultaneous examination of seasonal changes in plasma steroid
and IGF-1 concentrations, and in gonadal growth in a wild population of frogs. In order to
understand the functional relationships among reproductive steroids, IGF-1, and reproductive
tissues in frogs, it is essential to describe how these parameters fluctuate naturally under the
influence of temporal and environmental factors. The objective of the following study was to
document changes in concentrations of plasma IGF-1 and reproductive steroids (E2 and T), and in
gonadal tissues in conjunction with environmental factors for a population of wild female Pig
frogs (Rana grylio) in a north-central Florida lake. Rana grylio were chosen for this study
because they were abundant, they were relatively easy to acquire year-round, and they are the
largest ranid frogs in Florida, which made them ideal for the tissue and blood collections required
in this study. Additionally, R. grylio are closely related to bullfrogs (R. catesbeiana), a species for
which documented seasonal profiles of E2 and T served as a reference for this study (Castellani,
1958; Licht et al., 1983). Finally, the seasonal trends of gonadal maturation and breeding activity
for R. grylio have been well-established (Ligas, 1960; Lamb, 1983). The seasonal pattern of
changes in IGF-1 and sex steroids of wild-caught R. grylio established in this study serve as an
ecologically relevant reference for comparison with findings presented in other chapters.
Materials and Methods
Water Parameters, Animal Captures and Sample Collections
From April of 2002 to July of 2003, 6-20 adult female R. grylio were collected during
the fourth week of each month from Orange Lake (Lat. 29° 27°853’N, Long. 82° 11.380’W), in
Alachua County, Florida (Fig 3-1). In October of 2002, frogs were not collected due to rain and

40
lightening storms encountered on the lake during 3 separate collection attempts. Animals were
collected by hand from an airboat between 10 pm and 12 am. Captured frogs were transported, in
covered buckets with a small amount of water, to the Dept, of Zoology where they were housed
for less then 12 h in 38 L tanks with 19 L of dechlorinated water before examination.
Ligas (1960) reported that environmental factors such as rainfall, air temperature, and
water temperature influence reproductive condition of R. grylio in the Everglades; therefore, these
same parameters were measured at the collection site. Water temperature and pH were measured
using a Myron L Ultrameter (model 6P, Carlsbad, CA 92009). Monthly rainfall and air
temperature data from Orange Lake were recorded by Weather Station Number 02741536 and
were kindly provided by David Clapp of the USGS and National Weather Service.
Additionally, water samples from the collection site were examined for nitrate and nitrite
concentrations. Low precipitation combined with low water levels during the first 4 months of
this study might have contributed to slightly eutrophic conditions within the collection site.
Nitrate in known to interfere with gonadal steroidogenesis (Panesar and Chan, 2000) and with
amphibian reproduction (Rouse et al., 1999). Thus, nitrate was an important parameter to measure
when documenting plasma steroids of frogs collected at this site. Water nitrate and nitrite
concentrations were measured, with the generous assistance of Thea Edwards, using an auto¬
analyzer (Technicon auto-analyzer II with colorimeter, Bran+Luebbe Inc., Chicago, (888)917-
PUMP) equipped with a copper-cadmium reductor column. Methods for use are given in
Bran+Luebbe method number US-158-71 C, which is equivalent to EPA method 353.2. The auto¬
analyzer has a detection limit of 0.43 pg/L with a detection range of 0-400 pg/L of nitrate as
nitrogen. Samples are diluted in distilled water to fall within the detection range. Prior to analysis,
samples are filtered through a 1 micron glass fiber filter, collected in new or acid-washed
containers, and frozen (1 month) prior to measurement. Samples were quantified on a standard
curve created with each batch of water samples.

41
Previous studies on wild-caught bullfrogs reported that increasing duration of captivity
significantly decreased plasma hormones (Licht et al., 1983). Thus, a pilot study was performed
to determine the influence of duration of captivity on plasma hormone concentrations in R. grylio.
Blood samples were collected from frogs at 0, 6, 12, and 24 h post-capture. Blood samples were
drawn from frogs immediately after capture and then at time intervals afterwards while being
contained in covered buckets holding a small amount of lake water. No significant changes were
detected in concentrations of plasma E2, T, and IGF-1 over the 24 h period (Fig. 3-2). For
consistency in all subsequent procedures, blood and tissue samples were collected from frogs
within 12 h of capture. All animal procedures were performed in accordance with regulations
specified by University of Florida, Institute of Animal Care and Use Committee (Permit #Z095)
and a valid freshwater fishing license issued to T.R. Barbeau during the years of 2002 and 2003
as required by the State of Florida.
The frogs were anesthetized with MS-222 (1.5% 3-aminobenzoic acid ethyl-ether,
Aquatic Ecosystems, Orlando, FL), snout-vent length (SVL) and body mass were recorded, and
blood samples were obtained via cardiac puncture with heparinized syringe and needle. Blood
samples were centrifuged and resultant plasma frozen (-70°C) for E2, T, and IGF-1
radioimmunoassay (RIA) analyses. Frogs were then euthanized by dissection through the spinal
cord followed by pithing.
The gonadal-fat bodies, liver, ovaries, and oviducts were removed from each frog and
weighed. Fat bodies were examined because they are an important energy reservoir that can be
metabolized to provide energy for growth of reproductive tissues before (and throughout) the
reproductive period. The liver was examined because it is the primary site for synthesis of plasma
IGF-1, vitellogenin, and other substances vital for reproduction in oviparous ectotherms (Crain et
al. 1995; Guillette et al., 1996). Ovarian maturation was categorized as either regressed (stage 1),
yellow (stage 2), black (stage 3), or mature “black and white” (stage 4) based predominantly on
the stages of follicular development described by Ligas (1960). Briefly, regressed ovaries were

42
small (< 0.75 mm diameter), yellow, and contained no visible follicles. Yellow ovaries were also
small but contained yellow follicles up to 0.75 mm in diameter. Black ovaries were medium to
large and contained mostly black follicles 1.0 - 1.25 mm in diameter. Black ovaries can mature
within a relatively brief time to stage 4 ovaries. Lastly, mature ovaries were large, composed of
highly polarized follicles 1.25 - 2.0 mm in diameter, and had a sharp delineation of light and dark
colors indicating a vegetal and animal hemispheres. Mature ovaries contained oocytes ready for
ovulation and fertilization (Fig. 3-3).
Small cross-sections of the ampulla region of the oviducts were fixed in 4%
paraformaldehyde (4°C; 48 h) followed by rinse and storage in 75% ethanol for subsequent
histological analyses. The ampulla region, or middle portion of the oviduct, was examined
because it was the longest, most convoluted, and most visually distinct region (Wake and
Dickie, 1998). The ampulla region contains more glands and has a greater secretory activity than
other oviductal regions. The oviduct samples were dehydrated in a graded series of ethanol
changes, embedded in paraffin, serially cross-sectioned on a rotary microtome (7 pm), stained
with modified Masson’s staining procedure, and examined using light microscopy. To ascertain
oviductal proliferation, an ocular micrometer was used to make 10 morphological measurements
on 5 tissue sections, for a total of 50 measures per frog. The following oviductal parameters were
measured: epithelial cell height, endometrial thickness, endometrial gland height, and endometrial
gland width. Gland height and width measurements were used to calculate gland surface area
(uni2).
Steroid Radioimmunoassay (RIA) Biochemical Validation
Validation samples were obtained by creating plasma pools using aliquots from
individual frogs collected. Two methods were used to validate the E2 and T RIA: internal
standards and plasma dilutions. One half of the plasma pool, for use with internal standards, was
mixed with Norit charcoal (10 mL plasma to lg charcoal ratio; 4°C; 24 h) to strip steroid

43
hormones from the plasma. The solution was then centrifuged (3000 rpm; 1200xG; 45 min) and
the resultant supernatant decanted. Separate, duplicate aliquots of stripped plasma (25 pL) were
added to tubes and spiked with 100 pL of assay buffer containing 1.56,3.13, 6.25, 12.5,25,50,
100,200, 400, 800 pg E2 or T hormone. These tubes were extracted twice with ethyl-ether, air-
dried, and reconstituted in 100 pL borate buffer (100 pL; 0.05 M; pH 8.0).
For plasma dilutions, 6.25, 12.5, 25, 50, and 100 pL plasma was added to different tubes.
Appropriate volumes of borate buffer were added to bring the final sample volume of each tube
up to 200 pL. Samples were extracted twice with ethyl-ether, air-dried, and reconstituted with
100 pL borate buffer. Resultant samples for both internal standards and plasma dilutions were
examined by the RIA procedure described below.
Plasma extraction efficiencies were determined by adding 100 pL tritiated E2 and T
(15,000 cpm) to 100 pL of pooled plasma samples, extracting twice with ethyl-ether, air-drying,
and adding 500 pL scintillation fluid to tubes, and reading samples on a Beckman LS 5801
scintillation counter to determine the tritiated hormone remaining. The extraction efficiencies for
E2and T samples were 90.0% and 91.4%, respectively. Supernatant (500 pL) was added to 5 mL
of scintillation fluid, and counted on a Beckman scintillation counter. Plasma validation samples
were run in one assay with intraassay variance for E2 and T averaging 1.53% and 1.23%,
respectively. Plasma interassay variance for E2 and T averaged 6.99% and 3.27%, respectively.
Steroid RIA Procedures
RIAs were performed for E2 and T on plasma samples collected before surgery and after
treatments. For E2 samples, 25 pL of plasma was used and for T samples, 6.25 pL of plasma was
used. For T RIA, 50 pL of plasma samples were diluted with 200 pL of borate buffer, and 25 pL
of this dilution (6.25 pL plasma equivalents) were used as samples in the RIA. These volumes
were selected for analysis based on RIA volume determinations conducted on these samples
previously. Briefly, duplicates of plasma samples were extracted twice with ethyl-ether, air-dried,

44
and reconstituted in borate buffer. To each tube, bovine serum albumin (Fraction V; Fisher
Scientific) in 100 pL of borate buffer was added to reduce nonspecific binding at a final
concentration of 0.15% for T and 0.19% for E2. Antibody (Endocrine Sciences) was then added in
200 pL of borate buffer at a final concentration of 1:25,000 for T and 1:55,000 for E2. Finally,
radiolabeled steroid ([2,4,6,7,16,17-3H] 17p-estradiol at 1 mCi/mL; [1,2,6,7-3H] testosterone at 1
mCi/ml; Amersham Int., Arlington Heights, IL) was added at 12,000 cpm per 100 pL for a final
assay volume of 500 pL. Interassay variance tubes were similarly prepared from 2 separate
plasma pools for E2 and T. Standards for both E2 and T were prepared in duplicate at 0, 1.56, 3.13,
6.25, 12.5, 25, 50, 100, 200, 400, and 800 pg/tube. Assay tubes were vortexed for 1 min and
incubated at 4°C overnight.
Bound-free separation was performed by adding 500 pL of a mixture of 5% charcoal to
0.5% dextran, pulse-vortexing, and centrifuging tubes (1500g, 4°C, 30 min). Supernatant (500
pL) was added to 3 mL of scintillation fluid, and counted on a Beckman scintillation counter.
Plasma samples were run in 3 assays with intraassay variance for E2 and T averaging 3.35% and
4.99%, respectively. Plasma interassay variance for E2 and T averaged 3.97% and 6.99%,
respectively.
Insulin-like Growth Factor-1 (IGF-1) RIA Biochemical Validation
Pooled plasma samples (200 pL) were extracted in polypropylene tubes with acid-ethanol
(12.5% 2 N HC1, 87.5% ethanol; 400 pL) to dissociate IGF binding proteins from the IGF-1
molecules and to precipitate globular proteins as per Daughaday et al. (1980) and Crain et al.
(1995). After 30 min incubation (23°C) and 10 min centrifugation (2500xG; 4°C), the supernatant
was aliquoted to produce plasma equivalents of 12.5, 25, 50, 100, and 200 pL. Volume of the
plasma dilutions was brought to 200 pL with acid-ethanol before air-drying. Plasma dilutions
were compared with 0, 39, 156, 313, 625, 1000, 1250, 2500 pg of human recombinant IGF-1

45
standard (National Hormone and Pituitary Program, Torrance, CA 90509). Validation samples
were examined by IGF RIA procedures as described for experimental sample analyses below.
Plasma extraction efficiencies were determined by adding 100 pL iodinated IGF-1
(15,000 cpm) to 100 pL of pooled plasma samples, extracting with acid-ethanol, air-drying, and
reading samples on a Beckman 5500B gamma counter to determine the iodinated hormone
remaining. The extraction efficiency of plasma was 78.0% and all sample concentrations were
corrected for this loss. Validation of plasma dilutions was accomplished in one assay having an
intraassay variance of 2.3%. Internal standards and plasma dilutions were parallel to the standard
curve for E2 (ANCOVA; F = 0.24, P = 0.63 and F = 2.89, P = 0.15, Fig. 3-4A), and T RIA
(ANCOVA; F = 0.001, P = 0.99 and F = 0.013, P = 0.92, Fig. 3-4B).
IGF-1 RIA Procedures
IGF-1 RIA was performed as described by Crain et al. (1995) and Guillette et al. (1994).
The National Hormone and Pituitary Program (Torrance, CA 90509) supplied human
recombinant IGF-1 standard (9.76 to 2500 pg/tube) and human IGF-1 antisera (Lot #
AFP4892898, 1:400,000 final dilution). The antiserum had less than 1.0% cross-reactivity with
human IGF-II. Amersham International (Arlington Heights, IL) supplied iodinated IGF-1 label
(IGF-11125 sp act 2000 Ci/mmol; 16,000 cpm/tube) and Amerlex-M donkey anti-rabbit secondary
antibody (code RPN510, 500 pL/tube). Buffer reagents were purchased from Fisher Chemical
Co. (Pittsburgh, PA). Briefly, 20 pL plasma samples were aliquoted into polypropylene tubes,
extracted with 400uL acid-ethanol, and incubated 30 min prior to centrifugation (2500xG; 4°C;
10 min). For each sample, supernatant (100 pL) was pipetted into duplicate polypropylene tubes
and air-dried. IGF-1 standards were prepared in duplicate with 100 pL of known concentrations
of human recombinant IGF-1 standard (ranging from 9 - 2500 pg/tube), and 300 L RIA buffer
(200 mg/L protamine sulfate, 4.14 g/L sodium phosphate monobasic, 0.05% TWEEN 20, 0.02%
sodium azide, 3.72 g/L EDTA) added to each tube. Air-dried samples were reconstituted with 350

46
pL RIA buffer and vortexed. To each sample was added 50 pL IGF-1 antibody (human IGF-1
antisera, UB3-189) at a 1:10,000 final dilution. After adding 100 pL of iodinated IGF-1 label
(I125-IGF-1), with -15,000 CPM, samples were vortexed and incubated (4° C) overnight. Bound-
free separation of IGF-1 was accomplished by incubating samples for 10 min with 500 pL of
secondary antibody (Amerlex-M donkey anti-rabbit secondary antibody, code RPN.510 obtained
from Amersham International) at a final dilution of 1:10,000. Sample tubes were centrifuged
(2500xG; 4°C; 10 min) to separate the secondary antibody, which is bound to the primary
antibody and ligand. The supernatant was decanted and the pellet counted on a Beckman 5500B
gamma counter. Plasma samples were run in 3 assays having an average intraassay variance of
3.65% and an interassay variance of 4.63%. Plasma dilutions were parallel to the standard curve
for IGF-1 RIA (ANCOVA; F = 0.67, P = 0.43; Fig. 3-4C).
Statistics
Tissue mass is typically highly correlated to body mass; thus, tissue weights were
compared among months using ANCOVA, with body mass as a covariate, followed by Fishers
Protected LSD post hoc. Data were presented as adjusted mean mass (mg) ± SEM. Pair-wise
monthly comparisons, of mature and immature ovary stages, was performed with non-parametric
chi-square analyses. Concentrations of E2, T, and IGF-1 were estimated from raw data using the
commercially available Microplate Manager software (Microplate Manager III, BioRad
Laboratories, Inc., Hercules, CA, 1988). For RIA validation of pooled plasma dilutions and
internal standards, hormone concentrations were loglO-transformed prior to testing for
homogeneity of slopes with standard curves by ANCOVA. Hormone concentrations of E2, T, and
IGF-1 were compared among months with one-way ANOVA followed by SNK post hoc
contrasts. Tamhane post-hoc contrasts were used where variances were unequal among months
for IGF-1 concentrations. The relationships between plasma hormones, tissue weights, air and
water temperature, and rainfall were tested using Pearson’s correlation analysis. Statistical

47
analyses were performed using SPSS software (v. 10, SPSS Inc., Chicago, IL, 1999) with a =
0.05.
Results
Seasonal Environmental Parameters
Elevated air temperatures on Orange Lake between March and July of both years
overlapped with the reproductive period of R. grylio, as determined by patterns of peak plasma E2
and T concentrations described below. Conversely, decreased air temperatures overlapped with
the non-reproductive period between November and February (Fig. 3-5). High levels of
precipitation between June and September of 2002 overlapped the reproductive period season but
rainfall fluctuated considerably throughout 2003. Water temperature, pH, and nitrate and nitrite
ion concentrations were recorded between December of 2002 and July of 2003. Water
temperature was low between December and January, and showed a steady increase in February
that continued through the 2003 reproductive season (Fig. 3-5). Water pH between December and
May ranged from 6.5 to 6.8 and between June and July ranged from 5.7 to 6.0. Aquatic nitrate
and nitrate concentrations remained below 1 mg/L throughout the 2003 season. Generally, peak
reproductive condition, as determined by reproductive tissue weights and plasma E2 and T
concentrations, was considered to occur between April and July of 2002 and between March and
May of 2003, indicating that reproductive condition in R. grylio occurred during different months
over the 15 month study.
Seasonal Tissue Mass and Ovarian Maturation
Fat body weights exhibited seasonal variation with the greatest weights occurring during
June of 2002 and during January and March of 2003. The lowest fat body weights occurred
between July and December of 2002 and between April and July of 2003 (Fig. 3-6A). Liver
weights, which varied comparatively less with season, were greatest in April and March, and
lowest between September and December of 2002 (Fig. 3-6B). Oviductal weights were greatest
between April and July of 2002 and in May of 2003, whereas lowest weights occurred between

48
August of 2002 and March of 2003. Oviductal weights were also low between June and July of
2003 (Fig. 3-6C). Ovarian weights (GSI) were greatest in June of 2002, intermediate in May of
2002 and between March and May of 2003, and were lowest in April and between July and
December of 2002, in addition to in June and July of 2003 (Fig. 3-6D).
A distinct seasonal pattern of ovarian maturation stages was observed in R. grylio (Fig. 3-
3, 3-7). Frogs with black ovaries were considered to be in reproductive condition. Thus, frogs
having either black or mature ovaries were considered reproductively mature for analyses.
Conversely, frogs having either regressed or yellow ovaries, indicative of immature ovarian
follicles, were considered reproductively immature for analyses. A greater percentage of frogs
collected during the reproductive period had reproductively mature ovaries (Fig. 3-10). On
average, approximately 80% of the females examined during the reproductive period had
reproductively mature ovaries. In contrast, between 50% - 80% of frogs collected during the non-
reproductive period (August - December) had reproductively immature ovaries (Fig. 3-7) but not
all females collected had regressed ovaries this period.
Seasonal Plasma Steroid and IGF-1 Concentrations
Plasma E2 concentrations were elevated during the reproductive period of both years
compared to the non-reproductive period. Additionally, E2 concentrations were higher in 2002
than in 2003 (Fig. 3-8A). Plasma T concentrations were elevated during the reproductive period
of both years compared to the non-reproductive period. However, the period of elevated T
concentrations was of slightly shorter duration in 2002 than in 2003 (Fig. 3-8B). Plasma IGF-1
concentrations exhibited a pattern opposite that of T between reproductive periods. Plasma IGF-1
concentrations were increased during both reproductive period compared to the non-reproductive
period. However, IGF-1 concentrations were elevated for more months in 2003 than in 2002
(Figure 3-8C). The variation in environmental factors associated with variation in plasma
hormone concentrations between the two reproductive periods indicate that reproductive
physiology of R. grylio is influenced by environmental factors.

49
Throughout the season, plasma E2 concentrations were comparatively lower than plasma
concentrations of T and IGF-1. Plasma E2 and IGF-1 concentrations peaked during similar
months but plasma E2 concentrations declined precipitously after the reproductive period whereas
plasma IGF-1 declined less sharply and remained elevated slightly longer. A peak in plasma T
concentration occurred slightly prior to increases in plasma concentrations of E2 and IGF-1 (Fig.
3-9).
Peaks in ovarian and oviductal weights generally corresponded to elevated plasma T and
E2 concentrations, but they paralleled plasma T concentrations more closely (Fig. 3-15). During
the non-reproductive period, plasma steroids, plasma IGF-1, and reproductive tissue weights were
lower than in the reproductive period. Plasma IGF-1 concentrations appeared to peak in the latter
months of the reproductive period after steroid concentrations and the weights of reproductive
tissues began to decline (Fig. 3-10). Liver and fat body weights increased before, or in association
with, peaks in plasma T concentrations and much earlier than peaks in E2 and IGF-1
concentrations (Fig. 3-11).
Correlations: Plasma Steroids, Tissue Mass, and Environmental Parameters
For some months, the sample number of frogs collected was small; thus, analyses were
focused on correlations among means for all months. A strong positive correlation was detected
between plasma E2 and T concentrations (r2 = 0.67; P = 0.006) but not between concentrations of
E2 and IGF-1 and not between IGF-1 and T. Plasma E2 concentration correlated strongly to
oviductal weights (r2 = 0.84; P < 0.0001), to ovarian weights (r2 = 0.51; P < 0.05), and to water
temperature (r2 = 0.71; P = 0.05). Plasma T concentrations correlated strongly with ovarian (r2 =
0.85; P < 0.0001) and oviductal (r2 = 0.76; P = 0.001) weights but not to environmental
parameters. Plasma IGF-1 concentrations were negatively correlated to air temperature (r2 = 0.55;
P = 0.03) and positively correlated with fat body weights (r2 = 0.62; P = 0.02) but not correlated
to other parameters. Additionally, correlations were detected between ovarian and oviduct (r2 =
0.59; P = 0.02), or fat body masses (r2 = 0.57; P = 0.03), and between oviductal and liver weights

50
(r2 = 0.71; P = 0.003). Finally, fat body weight was correlated with water temperature (r2 = 0.75;
P = 0.03; Table 3-2).
Discussion
Relatively few studies have described a pattern of seasonal changes in the reproductive
tissues obtained from wild populations of ranid frogs (Ligas, 1960; Licht et al., 1983; Kim et al.,
1998). Previous studies of reproductive cyclicity in R. grylio have been limited to ovarian
maturation, male calling behavior, sexual dimorphism, and observations of amplexus (Lygas,
1960; Lamb, 1983; Wood et al., 1998). Licht et al. (1983) reported that ovarian and oviductal
weights in R. catesbeiana increased during the reproductive season, between May and July,
declined sharply in August, and remained reduced through October in a population from central
California.
In R. grylio from Orange Lake, increased ovarian and oviductal weights and plasma
steroid concentrations clearly define the months during which peak reproductive condition
occurred during this study. Analysis of ovarian maturation stages revealed that the largest
percentage of frogs had mature ovaries during the reproductive period between April and July of
2002 and between March and July in 2003. During the months of the non-reproductive period, a
greater percentage of frogs had regressed ovaries. A similar pattern of ovarian maturation was
described for R. grylio in the Okefenokee Swamp of Georgia and in the Everglades of South
Florida (Ligas, 1960; Lamb, 1983).
The reproductive period occurred during slightly different months over the 15 month
study, lasting between April and July of 2002 but only between March and June of 2003. Our
data indicate that reproductive condition of R. grylio, as in other amphibians studied previously,
is responsive to changing environmental conditions. The reproductive period generally
overlapped with the months of high water and air temperature, and of high rainfall during both
years. Also, concentrations of plasma IGF-1 increased with rising air temperature. Reproductive
tissue weights were greatest during periods of elevated air and water temperature, but no

51
significant correlations were observed between tissue weights and air or water temperature.
However, fat body weights decreased with increasing air temperature. These data lend support to
the theory that fat bodies are an energy reserve that are metabolized at the onset of warmer
weather to fuel rapid growth of reproductive tissues for breeding activity. Additionally, fat body
weights were positively correlated to ovary weights. Saidapur and Hoque (1995) reported similar
findings for Rana tigrina in India where decreasing fat body weights corresponded to increased
egg production, and both fat body weights and egg production were correlated to increasing air
temperature. In R. grylio from the Florida Everglades, reproductive activity is reportedly
suppressed during periods of low air and water temperature, and cease entirely during periods of
drought (Lygas 1960). The reproductive period of these frogs occurred primarily between March
and September and the non-reproductive period extends from October to February. Unlike frogs
from Orange Lake, R grylio from the Everglades appear to have an extended reproductive period
based on the mature ovarian tissue late into the season. This is supported by observations of
calling behavior by males, which also extends late into the season. However, it was unknown if
female R. grylio in the Everglades were actively mating and ovipositing eggs during these times,
so the extended reproductive period is speculative. This temporal variation in reproductive period
according to season could be attributed geographic differences, extended months of warm
temperatures in the summer, and milder temperatures during the winter in the Everglades
compared to north-central regions of Florida. Licht et al. (1983) attributed a similar temporal
variation in reproductive periods of bullfrogs according to geographic location. Also, in the
Okefmokee Swamp of Georgia, male R. grylio continue vocalizing between March and
September but peak reproductive condition of females occurs during June and July (Wright,
1932; Wright and Wright, 1949). These studies indicate that reproductive periods for R. grylio are
associated with localized environmental changes and also with geographic location.
The increase in relative liver weights of R. grylio, just prior to the onset of the
reproductive season, is indicative of increased hepatic biochemical or secretory activity. The liver

52
synthesizes many proteins that regulate metabolism, growth, reproduction, and development. One
of these proteins, vitellogenin, is a precursor of egg yolk in oviparous ectotherms and provides
valuable nutritive and energetic support for developing embryos (Camevali et al., 1995; Sumpter
and Jobling, 1995; Guillette et al., 1996; Palmer and Guillette, 1998). Estrogen produced by
mature ovaries stimulates vitellogenesis in the liver. Vitellogenin is a yolk precursor protein that
is transported through the plasma to the ovaries where it accumulates within developing ova
(Licht, 1979)(Palmer et al., 1998; Sumpter and Jobling, 1995). In female alligators vitellogenesis
is accompanied by an elevation of plasma IGF-1 concentrations during the reproductive period.
IGF-1 has also been detected in the egg albumin of birds and reptiles, suggesting that this
hormone plays a role in embryonic growth and development (Cox and Guillette, 1993; Guillette
et al., 1996). In R. grylio, liver weights were elevated before peaks in plasma IGF-1
concentrations, and might reflect hepatic synthesis of proteins that function in reproduction
(Palmer et al., 1998; LeRoith et al., 2001b). In oviparous ectotherms, plasma IGF-1 is also
influenced by nutritional status and feeding activity (Crain et al., 1995). In this study, plasma
IGF-1 concentrations were negatively correlated to decreasing fat body weights and positively
correlated to increasing air temperature. Accordingly, in R. grylio, the correlation between plasma
IGF-1 concentrations and fat body weights might reflect metabolism of stored fat (during the
warmer months of the spring and summer months) to provide energy for growth of reproductive
tissues. In contrast, decreased concentrations of plasma IGF-1 during the non-reproductive period
might reflect a decline in feeding behavior and in fat metabolism in female R. grylio.
Seasonal changes in concentrations of plasma steroids and IGF-1, in association with
changes in reproductive tissues, are largely undescribed for anurans. Seasonal changes in
reproductive tissues, and in concentrations of plasma steroids and plasma gonadotropins had been
described in ranid frogs from temperate North American and in India (Licht et al. 1983, Kim et
al., 1998). In California R. catesbeiana, plasma E2 and T concentrations generally peaked
between April and June; a pattern similar to that shown for plasma steroids in R. grylio. Plasma

53
steroid concentrations in R. grylio were most similar (1-4 ng/mL for E2 and 20-80 ng/mL for T) to
those measured in bullfrogs within 12 h of capture (Licht et al., 1983). Although plasma E2 and T
concentrations in R. grylio were positively correlated to each other, only plasma T concentrations
were correlated to ovarian and oviductal weights. This observation conflicts with findings in
mammals but indicates it might be common among non-mammalian vertebrates. In ectotherms,
androgens might play an important role in regulating reproductive condition. Amphibian ovarian
follicles synthesize and secrete large quantities of androgens during ovarian maturation (Fortune
and Tsang, 1981; Fortune, J.E. 1983; Lutz et al., 2001). Androgens might be aromatized to
estrogens in peripheral tissues such as the brain, fat and skin (Follett and Redshaw, 1968). The
oviduct might also be a site of peripheral aromatization of androgens and be a target for androgen
activity. The oviduct of oviparous species synthesizes huge quantities of protein (perhaps in
response to androgen stimulation) for use as secondary or tertiary egg coatings such as in anuran
egg jellies (Maack et al., 1985; Olsen and Chandler, 1999; Arranz and Cabada, 2000; Jesu-Anter
and Carroll, 2001). Similar to R. grylio, female R. catesbeiana also exhibited greater T than E2
plasma concentrations indicating that this pattern might be prevalent among ranids (Licht et al.,
1983). Rana grylio exhibited peak ovarian and oviductal weights during similar months as
reported for bullfrogs (Licht et al., 1983). However, plasma steroid concentrations in R. grylio did
not decrease significantly 24 h after capture as reported for R. catesbeiana (Licht et al., 1983). In
R. catesbeiana, increases in ovarian and oviductal mass closely paralleled increases in plasma
gonadotropins and steroids (Licht et al., 1983). Plasma LH and FSH were not measured in R.
grylio and it remains unknown whether plasma gonadotropins increased before elevations in
plasma steroid concentrations or tissue mass. In future studies, it would be valuable to examine
changes in plasma gonadotropins with respect to steroids to better understand the reproductive
cycle of R grylio.
Before this study on R. grylio, seasonal changes in plasma IGF-1 concentration were
reported for only one other anuran species, Bufo woodhousei (Pancak-Roessler and Lee, 1990).

54
Plasma IGF-1 concentrations in B. woodhousei peaked (1 ng/ml) in July and declined sharply
thereafter. In R. grylio, plasma IGF-1 concentrations peaked between May and July and declined
after August. In R. grylio a peak in plasma IGF-1 concentrations occurred later in the season
compared to Bufo woodhousei and is likely due to several factors including geographical
variation, interspecific differences, and differences in IGF-1 RIA methods.
In reptiles, seasonal changes in plasma IGF-1 concentrations have been described for
alligators and turtles. In loggerhead sea turtles, elevated plasma IGF-1 concentrations occurred
between April and June and were associated with reproductive activity and increased feeding
behavior of female turtles during these months (Crain et al., 1995). Guillette et al. (1996)
examined reproductive tissues, and plasma steroids and plasma IGF-1 concentrations, and their
respective associations with reproductive condition in alligators. In female alligators, plasma
IGF-1 concentrations increased in June and were associated with gravidity. Elevated plasma E2
and P4 concentrations were associated with peak vitellogenesis, and also preceded gravidity and
peaks in plasma steroid concentrations. Seasonal patterns of plasma IGF-1 concentrations were
not examined simultaneously with changes in plasma steroids concentrations; therefore, it is
unknown how these hormones change (with respect to each other) seasonally in alligators
(Guillette et al., 1996). In a separate study, alligators collected from the same locality exhibited a
peak in plasma E2, T, and P4 concentrations in May (Guillette et al., 1997). Thus, alligators are
similar to R. grylio in that elevated plasma steroid concentrations precede peaks in plasma IGF-1.
In mammals, IGF-1 expression is associated various aspects of reproduction including
ovarian maturation, follicular atresia, selection of dominant follicles, and regulation of gonadal
steroidogenesis. Increasing concentrations of IGF-1 can be synthesized in reproductive tissues or
the in liver, and can be transported directly to offspring in útero. In contrast to mammals,
oviparous animals must provide growth-promoting substances to eggs prior to oviposition
(Palmer and Guillette, 1991; Guillette et al., 1996). Accordingly, IGF-1 has been detected in the

55
yolks of chicken eggs, the albumen of alligator eggs, and the oviductal glands of geckos and
alligators (Scavo et al., 1989; Guillette and Williams, 1991; Cox and Guillette, 1993; Cox, 1994).
In conclusion, this study provides the first evidence that IGF-1 is present in the plasma of
R. grylio. Plasma IGF-1 concentrations were correlated with several environmental factors and
exhibited a clear pattern of change with reproductive period, and with reproductive steroid
concentrations and weights of reproductive tissues. Although the role of IGF-1 in anurans
requires further study, this study has provided valuable information for understanding the
association of IGF-1 with reproductive physiology in R. grylio.

56
Table 3-1. Comparison of plasma insulin-like growth factor-1 (IGF-1) concentrations among
mammalian, avian, reptilian, and amphibian species.
IGF-1
(ng/mL)
Reference
Mammalian
Rat
Juvenile males
574.0
a
Human
Adult females
261.0
a
(Reproductive status unknown)
Cow
Adult females
182.0
a
(Reproductive status unknown)
Avian
Chicken
Juveniles (8-week)
42.0
a
Reptilian
Red-eared slider turtle
Juvenile males
17.0
a
Loggerhead sea turtle
Reproductive
7.5
b
Non-reproductive
3.0
b
American alligator
Reproductive
16.0
c
N on-reproductive
5.0
c
Amphibian
African Clawed frog
Non-reproductive females
3.0
d
American toad
Reproductive males
4.0
d
Non-reproductive males
0.5
Bullfrog
Non-reproductive female
1.0
d
Marine toad
Non-reproductive female
1.0
d
Pig frog
Reproductive females
22.0
This Study
Non-reproductive females
10.0
This Study
a - Daughaday et al., 1985
b - Crain et al., 1995
c - Guillette et al., 1996
d - Pancak-Roessler and Lee, 1990
Note - For all studies shown, plasma IGF-1 was measured, after acid-extraction
of IGF-1 binding proteins, by radioimmunoassay.

57
Table 3-2. Correlations among body mass, snout vent length (SVL), hormone concentrations,
tissues weights of Rana grylio, and environmental parameters.
Pearson Correlation P-value R1 Relationship
Body Mass and E2
0.27
Body Mass and T
0.06
Body Mass and IGF-1
0.94
Snout-Vent-Length and E2
0.31
Snout-Vent-Length and T
0.09
Snout-Vent-Length and IGF-1
0.58
Plasma E2 and T
0.006 **
0.67
+
Plasma E2 and IGF-1
0.28
Plasma E2 and Ovary Weight
0.05
0.51
+
Plasma E2 and Oviduct Weight
<0.0001 **
0.84
+
Plasma E2 and Liver Weight
0.18
Plasma E2 and Fat Body Weight
0.88
Plasma E2 and Air Temperature
0.07
Plasma E2 and Water Temperature
0.05
0.71
+
Plasma E2 and Rainfall
0.77
Plasma T and IGF-1
0.64
Plasma T and Ovary Weight
<0.0001 **
0.85
+
Plasma T and Oviduct Weight
0.001 **
0.80
+
Plasma T and Liver Weight
0.06
Plasma T and Fat Body Weight
0.16
Plasma T and Air Temperature
0.20
Plasma T and Water Temperature
0.09
Plasma T and Rainfall
0.25
Plasma IGF-1 and Ovary Weight
0.81
Plasma IGF-1 and Oviduct Weight
0.67
Plasma IGF-1 and Liver Weight
0.35
Plasma IGF-1 and Fat Body Weight
0.02 *
0.62
—
Plasma IGF-1 and Air Temperature
0.03 *
0.55
+
Plasma IGF-1 and Water Temperature
0.12
Plasma IGF-1 and Rainfall
0.48
Ovary and Oviduct Weight
0.02 *
0.59
+
Ovary and Liver Weight
0.25
Ovary and Fat Body Weight
0.03 *
0.57
+
Oviduct and Liver Weight
0.003 **
0.71
+
Oviduct and Fat Body Weight
0.27
Liver and Fat Body Weight
0.13

58
Table 3-2. Continued.
Pearson Correlation
P-value
R2 Relationship
Air Temperature and Ovary Weight
0.84
Air Temperature and Oviduct Weight
0.37
Air Temperature and Liver Weight
0.70
Air Temperature and Fat Body Weight
0.08
Water Temperature and Ovary Weight
0.45
Water Temperature and Oviduct Weight
0.07
Water Temperature and Liver Weight
0.59
Water Temperature Fat Body Weight
0.03 *
0.75
Rainfall and Ovary Weight
0.55
Rainfall and Oviduct Weight
0.88
Rainfall and Liver Weight
0.37
Rainfall and Fat Body Weight
0.84
* Significant
** Highly Significant
+ positive correlation
- negative correlation

59
:igure 3-1. Collection site for Rana grylio, indicated by asterisk, on Orange Lake in Alachua
county, Florida (Latitude 29°27.853’N, Longitude 82°11.380’W) between April,
2002 and July, 2003. Image created by T. Barbeau.

Plasma E2 and T (pg/mL)
60
20000
18000
16000
14000
12000
10000
8000
6000
4000
2000
0
A E2
â–  T
o IGF-1
i
í
ó
5
í
5
4
60000
50000
40000
Vi
3
ss
o
30000 2
20000
10000
"O
ora
3
r
0 6 12 24
Time (h)
Figure 3-2. Plasma 17P-estradiol (E2), testosterone (T), and insulin-like growth factor-1 (IGF-1)
concentrations in Rana grylio at 0, 6, 12, and 24 h post-capture. Plasma samples for
each time interval were collected from different frogs. Data presented as means ±
SEM. Letters within axes represent sample size at each time interval. No significant
differences detected among time intervals for each hormone (ANOVA; E2 P = 0.40;T
P = 0.83; IGF-1 P = 0.42).

Maturation Stages
61
Regressed
Yellow
Black
Mature
2 cm
Figure 3-3. Staging of Rana grylio ovaries in progression from least to most mature stages. A.
regressed (stage 1), B. yellow (stage 2), C. black (stage 3), and D. black and white
(stage 4) ovaries.

62
A
100 ^
80
o
60
X
40
59
20
0
B 100
80
° o
° Standard
♦ Plasma Dilutions
â–  Internal Standards
o o
o
o
60
X 4o
o
i
PQ
20
0
1
10 100 1000 10000
Estradiol (pg)
° Standard
o ♦ Plasma Dilutions
â–  Internal Standards
•o
10 100 1000
Testosterone (pg)
10000
C
100
O
80
X
60
o'
40
55
CO
20
0
o
♦
° o
♦ .
o
° Standard
♦ Plasma Dilutions
o
♦ o
♦ o
o
T
10
100 1000 10000
IGF-1 (pg)
Figure 3-4. Biochemical validation of Rana grylio plasma for RIA. A. For 17P-estradiol RIA
internal standard and plasma dilution curves were parallel to the standard curve
(ANCOVA; F = 0.24, P = 0.63 and F = 2.89, P = 0.15). B. For testosterone RIA
internal standard and plasma dilution curves were parallel to the standard curve
(ANCOVA; F = 0.001, P = 0.99 and F = 0.01, P = 0.92). C. For insulin-like growth
factor-1 (IGF-1) RIA the plasma dilution curve was parallel to the standard curve
(ANCOVA; F = 1.05, P = 0.33).

63
30
25
20
15
10
5
0
-5
Month
Figure 3-5. Monthly changes in rainfall, water temperature, and high and low air temperature at
collection site on Orange Lake, Florida. Data presented as means per month.
Mean Rainfall (in)

Mean Ovary Mass (mg) ‘ Mean Oviduct Mass (mg) * Mean Liver Mass (mg) * Mean Fat Body Mass (mg)
64
Reproductive Non-Reproductive Reproductive
— - — ' „ —
Month
Figure 3-6. Seasonal change in fat body weights in Rana grylio during the reproductive and non¬
productive periods. Data presented as means ± SEM. Numbers within bars indicate
sample size and different letters above bars indicate significantly different means A.
for fat bodies (ANCOVA; F = 2.78, P = 0.002), B. for liver (ANCOVA; F = 4.90, P
< 0.001), C. for oviducts (ANCOVA; F = 1.55, P < 0.001), and D. ovaries
(ANCOVA; F = 4.27, P < 0.001).

% Total Frogs per Month
65
â–¡ Immature E3 Yellow
Black IS Mature
AMJ J ASONDJ FMAMJ J
2002 2003
Month
Figure 3-7. Seasonal changes in ovarian maturation stages of Rana grylio. Data presented as
percentage of frogs exhibiting immature, yellow, black, and mature ovary stages, out
of total frogs, from the total collected that month. Different letters above bars
indicate significantly different percentages as determined by Mann Whitney U
pairwise contrasts (P < 0.05).

Mean Testosterone (pg/mL) QJ Mean Estradiol (pg/mL)
66
A.
Reproductive
Non-Reproductive
5000
4500
4000
3500 ;
3000
2500 -j
2000 1
1500 -
1000
500
o I
Reproductive
-*â– 
45000 -i
40000
35000
30000 i
25000
20000 cd
15000 r-U
10000
be
5000
0
13
18
19
c.
M J J
Reproductive
d d
m cd
A S
d
t\
a I6
O N D
Non-Reproductive
10
be
be
10
12
10
M
A M J J
Reproductive
40
35 j
Í 30
5 25
U 20
ab
cd
e
«
u
5
cd
nh
7
A
2002
18
19
be
cd
cd cd
cd
10
10
12
abc
a be
A
O N
Month
D J F
2003
M
M
Figure 3-8. Seasonal change in plasma hormones in Rana grylio during the reproductive and non-
reproductive periods. Data presented as means ± SEM. Numbers within bars indicate
sample size while different letters above bars indicate significantly different means
for A. 17P-estradiol (ANOVA; P < 0.001), B. testosterone (ANOVA; P < 0.001), and
C. insulin-like growth factor-1 (IGF-1, ANOVA; P < 0.001).

67
2002 2003
Month
Figure 3-9. Seasonal changes in plasma 17p-estradiol (E2), testosterone (T), and insulin-like
growth factor-1 (IGF-1) in Rana grylio during the reproductive and non-reproductive
periods. Data presented as means ± SEM.
Plasma Estradiol (pg/mL)

Plasma Hormone
68
2002 2002
Month
Figure 3-10. Seasonal changes in plasma 17P-estradiol (E2), testosterone (T), and insulin-like
growth factor-1 (IGF-1), and of ovary and oviduct weights in Rana grylio during the
reproductive and non-reproductive periods. Steroid data presented as means and
tissue data presented as means ± SEM.

Plasma Hormone
69
Figure 3-11. Relative seasonal changes in plasma 17P-estradiol (E2), testosterone (T), insulin-like
growth factor-1 (IGF-1), and of liver and fat body weights Rana grylio during the
reproductive and non-reproductive periods. Steroid data presented as means and
tissue data presented as means ± SEM.
Tissue mass (mg) > 1 Liver ««mu Fat Bodies

CHAPTER 4
THE EFFECTS OF INSULIN-LIKE GROWTH FACTOR-1 AND ESTRADIOL IMPLANTS
(IN VIVO) ON OVIDUCT MORPHOLOGY, AND ON PLASMA HORMONES IN
BULLFROGS (Rana catesbeiana)
Introduction
The regulation of oviduct growth and function by endocrine hormones has been described
for mammals and some reptiles but comparatively little is known for amphibians (Christiansen,
1973; Mead et al., 1981; Murphy and Ghahary, 1990; Cox and Guillette, 1993; Buhi et al., 1999;
Girling et al., 2000). Most studies of frog reproduction have focused on variation in plasma
concentrations of steroid hormones and ovarian maturation, with little attention to regulation of
oviductal structure (Licht et al., 1984; Wake and Dickie, 1998). For oviparous animals, the
oviduct is vital for reproduction because it synthesizes and secretes important substances that
nourish and encapsulate ovulated oocytes. The female bullfrog (Rana catesbeiana) can oviposit
as many as 40 to 80 thousand eggs at one breeding event (Norris, 1997). Without the provision of
oviductal secretions, oocytes could not be fertilized successfully nor could they develop into
normal embryos (Low et al., 1976; Buhi et al., 1997; Buhi et al., 1999; Olsen and Chandler,
1999). The amphibian oviduct includes four major structural and functional regions: the
infundibulum, the atrium, the ampulla, and the ovisac (Uribe et al., 1989). The infundibulum is
the anterior-most region of the oviduct and receives mature oocytes ovulated from the ovaries.
Distal to the infundibulum is the atrium - a narrow aglandular region that precedes the ampulla.
The ampulla region is longest portion of the oviduct and contains numerous glands within the
endometrial layer (Wake and Dickie, 1998). The glands within the ampulla region are
biochemically active and secrete a variety of substances that are incorporated into mature oocytes
as they traverse the oviduct (Uribe et al., 1989). The last region of the oviduct is the ovisac or
70

71
uterus that leads to the cloaca. The narrow and aglandular ovisac is the final site from which
oocytes are deposited from the reproductive tract into the environment.
Oviductal growth occurs primarily in response to stimulation by elevated E2
concentrations of E2, of ovarian origin, and involves proliferation of epithelial and endometrial
cells (Christiansen, 1973; Mead et al., 1981; Cox, 1994). In amphibians, the major reproductive
steroids progesterone (P4), testosterone (T), and 17P-estradiol (E2), are produced and secreted by
the ovary, in response to pituitary follicle stimulating hormone (FSH) and luteinizing hormone
(LH) (Licht, 1979; Chapter 3). The principal steroid that regulates structure and function of the
oviduct is reported to be E2. In addition to E2, polypeptide growth factors have been shown to
elicit a growth response in the reptilian and mammalian oviduct (Cox and Guillette, 1994;
Stevenson et al., 1994; Tang et al., 1994; Richards et al., 1997). Growing evidence demonstrates
that autocrine and paracrine sources of epidermal growth factor (EGF) and insulin-like growth
factor-1 (IGF-1) are potent hormonal mitogens that mediate E2-induced oviduct growth. In
mammals, these growth factors induce oviduct growth in the absence of endogenous E2, and
induce an even greater growth in the presence of E2 compared to either hormone administered
alone (Nelson, 1991; Murphy and Murphy, 1994). These findings indicate a hormonal synergy
between E2 and IGF-1 in the stimulation of oviduct growth.
Cox and Guillette (1994) reported that ovariectomized geckos exhibit oviductal growth in
response to EGF and IGF-1 even in the absence of stimulation by endogenous E2. However,
neither EGF nor IGF-1 stimulation induced oviduct growth similar to that observed with E2 alone.
Unfortunately, the effect of simultaneous treatment with E2 and growth factors on oviduct growth
was not examined in this study and it remains unknown if an E2 and IGF-1 synergy exists for
these animals.
In this study, I examined the effects of controlled doses of steroid hormone (E2), and
peptide hormones (IGF-1 and EGF) on oviduct growth in adult, female bullfrogs (Rana
catesbeiana). The objective of this study was to determine whether the oviducts in R. catesbeiana

72
exhibited a growth response with exposure to E2 or to growth factors administered separately or
in combination. I predicted that ovariectomized R. catesbeiana treated with either E2 or growth
factors (EGF and IGF-1) would exhibit oviduct growth. Additionally, I predicted that oviduct
growth would be greater in frogs treated simultaneously with E2 and IGF-1 than with either
hormone alone.
Materials and Methods
Adult female Rana catesbeiana (N = 65) were purchased (Charles D. Sullivan Co. Inc.,
TN). They were maintained under a 12-h diurnal light/dark cycle in 38 L tanks with 19 L of
static, dechlorinated water at 26°C. They were fed crickets every other day throughout the
experiment. Frogs were randomly assigned to each of the following treatment groups: E2 (N=10),
IGF-1 (N=10), EGF (N=10), E2 /IGF-1 (N=10), placebo (N-10), and sham (N=5). The use of
sham frogs is explained below. Weight and snout vent length (SVL) of frogs were recorded and
no significant difference was detected in mass (ANOVA; P = 0.84) or SVL (ANOVA; P = 0.85)
of frogs among treatment groups. Numbered stainless steel tags were applied to the webbing of
the hind foot of frogs for individual identification. Animals were maintained and experiments
were performed as approved by the Institute for Animal Care and Use Committee (IACUC
project #Z095).
Sham animals are frogs that were subjected to identical anesthesia and surgical
procedures that ovariectomized frogs were subjected to (explained below) with the exception that
the ovaries were not removed and they received no hormone treatment. Thus, sham frogs were
intact frogs that were included to account for an effect of the surgical procedure itself on
physiological responses of the frogs.
For this study, R. catesbeiana were chosen in lieu of A! laevis and R. grylio that were
examined in earlier chapters. Previous attempts to maintain wild-caught R. grylio in captivity
demonstrated that this species exhibited considerable stress and was considered inappropriate for
a long-term, surgical study. In contrast to R. grylio, R. catesbeiana were very adaptable to

73
captivity and exhibited less stress. Previous attempts to ovariectomize X. laevis were largely
unsuccessful while R. catesbeiana responded optimally to the ovariectomy procedures with low
mortality and fast recovery. Additionally, R. catesbeiana are large-bodied frogs and it was easier
to collect blood samples of greater volume than in X. laevis and R. grylio. Lastly, R. catesbeiana
is closely related to R. grylio and was considered a relevant and appropriate substitution for this
study.
Ovariectomy
After a 2-week acclimation period, frogs were anesthetized with MS-222 (1.5% 3-
aminobenzoic acid ethyl ether, Aquatic Ecosystems, Orlando, FL), and ovariectomy was
performed. The ovaries were removed to ensure that endogenous hormones did not conflict with
or obscure effects observed in response to experimental treatments. Conducting more than six
surgeries per day would have compromised my ability to carefully conduct surgeries and oversee
post-operative recovery of individuals. Thus, ovariectomy was performed each day, for 10
consecutive days, on one individual selected from each of the six treatment groups.
Under sterile conditions, a 2.5 cm right paramedial incision was made through the skin
and muscle layers into the abdominal cavity, and the left and right ovaries were excised through
this single incision. Hemostasis of the mesovarium (vascular tissue supporting the ovaries) was
accomplished by a series of double-ligatures of the vessels using 5-0, monofilament nylon suture
material (Fig. 4-1). The incision layers (peritoneal, muscle, and skin) were closed with a single
interrupted pattern using the same suture material. For each female, the mass of excised ovaries
was recorded and reproductive status of the female was determined by visual inspection of
ovarian follicle maturation according to Dumont (1971). This procedure was performed to
confirm that females were reproductively similar at the onset of the experiment to minimize
variation in responses to subsequent treatments.
Pre-ovariectomy blood samples were collected by cardiac puncture to determine whether
plasma E2, T, and IGF-1 concentrations were similar among females at the start of the

74
experiment. No more than 1.0% of total blood volume estimated per body mass was taken from
frogs (Mader, 1996; Wright, 2001). Blood samples were stored in heparin vacutainer tubes,
centrifuged, and subsequent plasma was stored (-70°C) for radioimmunoassay (RIA) analyses. If
blood could not be collected within two cardiac punctures attempts were ceased to avoid potential
injury to the frog. Blood could not be sampled from all individuals prior to surgery; therefore,
sample sizes for pre-ovariectomy blood samples were as follows: E2 (N= 8), IGF-1 (N=9), EGF
(N=7), E2 /IGF-1 (N=9), placebo (N=8), and sham (N=5).
Post-ovariectomy frogs were placed in recovery tanks containing benzalkonium chloride
(antibiotic) dissolved in 1 liter of water for 48 h. Afterwards, recovered frogs were returned to
their tanks. Frogs were allowed a 3-week recovery period during which the surgical sites were
closely monitored for signs of inflammation or infection. Although most frogs experienced no
post-operative complications, five frogs failed to recover from the ovariectomy. Therefore, the
final sample sizes for post-ovariectomy treatment groups were as follows: E2 (N=10), IGF-1
(N=10), EGF (N=8), E2/IGF-1 (N=9), placebo (N=8), and sham (N=5). No further losses
occurred, and these sample sizes were maintained for each treatment group for the remainder of
the study.
Hormone Implants
After the 3-week recovery period, all frogs exhibited complete healing of the
ovariectomy incision site (Fig. 4-2). Treatments were administered by surgical insertion of an
intra-abdominal, 21-day release treatment pellet (Innovative Research of America, Sarasota, FL)
containing either of the following dosages: E2 (420 pg), IGF-1 (10 pg), EGF (10 pg), E2/IGF-1
(420 pg E2 and 10 pg IGF-1), and placebo (10 pg vehicle pellet). Surgical procedures for the
treatment pellet implantation were similar to those described for ovariectomy with respect to
incision and abdominal closure; however, an 1.0 cm left paramedial incision was made. Pellets
were inserted into the abdominal cavity midway between the left and right oviducts (Fig. 4-3).

75
The E2 treatment served as a positive control, while the placebo treatment served as a negative
control. The EGF, IGF-1, and E2/IGF-1 treatments were experimental. For simultaneous
treatment with E2/IGF-1, one pellet of each hormone was inserted 2.54 cm apart from each other
within the abdomen. The hormone concentrations administered were physiologically relevant and
chosen based on a literature review of similar studies in which treatments were given for
durations ranging from 7-20 days to elicit a tissue response (Redshaw et al., 1968; Follet and
Redshaw 1968; Fortune 1981; Cox 1994; Crain et al. 1995). After 18 days of treatment, the frogs
were euthanized and examined as described below.
Tissue Sampling
The response of R. catesbeiana to 18 days of treatment was determined by measuring the
following parameters: weights of tissues (liver and oviduct), oviductal growth (macroscopic and
microscopic), and plasma concentrations of E2, T, and IGF-1. After treatment, frogs were
anesthetized with MS-222, and blood samples were collected. Frogs were euthanized by
dissection through the spinal cord followed by pithing. Plasma samples (post-treatment) were
frozen (-70°C) for RIA analyses. The liver and oviducts were removed from each frogs and
weighed for comparison of wet tissue mass among treatment groups. Cross-sectional samples of
oviducts were fixed in 4% paraformaldehyde (4°C; 48 h) followed by rinse and storage in 75%
ethanol for subsequent histological analyses. The oviducts were dehydrated in a graded series of
ethanol changes, embedded in paraffin, serially sectioned on a rotary microtome (7 pm), stained
with modified Masson’s staining procedure, and examined microscopically. To evaluate oviductal
growth, an ocular micrometer was used to record 10 morphological measurements on 5 tissue
sections per frog (for a total of 50 measures) for each of the following oviductal parameters:
epithelial cell height, endometrial layer thickness, endometrial gland height, and endometrial
gland width. The gland height and width measurements were used to calculate cross-sectional
gland surface area.

76
Steroid Radioimmunoassay (RIA) Biochemical Validation
Validation samples were obtained by pooling plasma aliquots from each individual. Two
methods were used to biochemically validate the E2 and T RIA: internal standards and plasma
dilutions. One half of the plasma pool, for use with internal standards, was mixed with Norit
charcoal (10 mL plasma: 1 g charcoal; 4°C; 24 h) to strip steroid hormones from the plasma. The
solution was then centrifuged (3000 rpm; 1200xG; 45 min) and the resultant supernatant
decanted. Separate aliquots of stripped plasma (25 pL) were added to 10 tubes and spiked with
100 pL of 1.56, 3.13, 6.25, 12.5, 25, 50, 100, 200, 400, 800 pg cold E2 or T hormone. These tubes
were extracted twice with ethyl-ether, air-dried, and reconstituted in 100 pL of borate buffer (100
pL; 0.05 M; pH 8.0).
For plasma dilutions, 6.25, 12.5, 25, 50, 100, and 200 pL plasma was added to 6 tubes.
Appropriate volumes of borate buffer were added to each tube to bring the final sample volume to
200 pL. Samples were extracted twice with ethyl-ether, air-dried, and reconstituted with 100 pL
of borate buffer. Resultant samples for both internal standards and plasma dilutions were
examined by the RIA procedure described below.
Plasma extraction efficiencies were determined by adding 100 pL tritiated E2 and T
(15,000 cpm) to 100 pL of pooled plasma samples, twice extracting with ethyl-ether, air-drying,
adding 500 pL scintillation fluid to tubes, and reading samples on a Beckman LS 5801
scintillation counter to determine the tritiated hormone remaining. The extraction efficiencies for
E2and T samples were 93.9% and 87.9%, respectively. Supernatant (500 pL) was added to 5 mL
of scintillation fluid, and counted on a Beckman scintillation counter. Plasma intraassay variance
for E2 and T validation RIAs for averaged 1.53% and 1.23%, respectively. Plasma interassay
variance for E2 and T averaged 2.87% and 4.88%, respectively.

77
Steroid RIA Procedures
RIAs were performed for E2 and T on plasma samples collected both pre-ovariectomy
and post-treatment. For pre-ovariectomy E2 samples, 50 pL of plasma was used, and for post¬
treatment samples, 50 pL of plasma was used for E2, E2/IGF-1, and sham samples and 300 pL
plasma used for 1GF-1, EGF, and placebo samples. For pre-ovariectomy T samples, 30 pL of
plasma was used while for post-treatment samples, 50 pL of plasma was used for E2, E2/IGF-1,
and sham samples and 300 pL plasma used for IGF-1, EGF, and placebo samples. These volumes
were selected for analysis based on RIA volume determinations conducted on these samples
previously. Briefly, duplicates of plasma samples were twice extracted with ethyl ether, air-dried,
and reconstituted in borate buffer. To each tube, bovine serum albumin (Fraction V; Fisher
Scientific) in 100 pL of borate buffer was added to reduce nonspecific binding at a final
concentration of 0.15% for T and 0.19% for E2. Antibody (Endocrine Sciences) was then added to
200 pL of borate buffer for a final concentration of 1:25,000 for T and 1:55,000 for E2. Finally,
radiolabeled steroid ([2,4,6,7,16,17-3H] E2 at 1 mCi/mL; [1,2,6,7-3H] T at 1 mCi/mL; Amersham
Int., Arlington Heights, IL) was added at 12,000 cpm per 100 pL for a final assay volume of 500
pL. Interassay variance tubes were similarly prepared from two separate plasma pools for E2 and
T. Standards for both E2and T were prepared in duplicate at 0, 1.56, 3.13, 6.25, 12.5, 25, 50, 100,
200, 400, and 800 pg/tube. Assay tubes were vortexed for 1 min and incubated overnight at 4°C.
Bound-free separation was performed by adding 500 pL of a mixture of 5.0% charcoal to 0.5%
dextran, pulse-vortexing, and centrifuging tubes (1500g, 4°C, 30 min). Supernatant (500 pL) was
added to 5 mL of scintillation fluid, and counted on a Beckman scintillation counter. Plasma
intraassay variance for E2 and T averaged 2.87% and 4.93%, respectively. Plasma interassay
variance for E2 and T averaged 7.64% and 5.25%, respectively.

78
Insulin-Like Growth Factor-1 (IGF-1) RIA Biochemical Validation
From each treatment group, plasma (200 pL) was pooled for validation, and was
extracted in polypropylene tubes with acid-ethanol (12.5% 2 N HC1, 87.5% ethanol; 800 pL) to
dissociate IGF binding proteins from the IGF-1 molecules and to precipitate globular proteins as
per Daughaday et al. (1980) and Crain et al. (1995). After 30 min incubation (room temperature)
and 10-min centrifugation (2500xG; 4°C), the supernatant was aliquoted to produce plasma
equivalents of 12.5, 25, 50, 100, and 200 pL. Plasma dilution volumes were brought to 200 pL
with acid-ethanol prior to air-drying. Plasma dilutions were compared with 0, 39, 156, 313, 625,
1000, 1250, 2500 pg of human recombinant IGF-1 standard (National Hormone and Pituitary
Program, Torrance, CA 90509). Validation samples were examined by IGF RIA procedures as
described for experimental sample analyses below. Plasma extraction efficiencies were
determined by adding 100 pL iodinated IGF-1 (15,000 cpm) to 100 pL of pooled plasma
samples, extracting with acid-ethanol, air-drying, and reading samples on a Beckman 5500B
gamma counter to determine the iodinated hormone remaining. The extraction efficiency of
plasma was 77.0% and all sample concentrations were corrected for this loss. Validation of
plasma dilutions was accomplished in one assay having an intraassay variance of 2.27%.
IGF-1 RIA Procedures
IGF-1 RIA was performed as described by Crain et al. (1995) and Guillette et al. (1994).
The National Hormone and Pituitary Program (Torrance, CA 90509) supplied human
recombinant IGF-1 standard (9.76 to 2500 pg/tube) and human IGF-1 antisera (Lot #
AFP4892898, 1:400,000 final dilution). The antiserum had less than 1.0% cross-reactivity with
human IGF-II. Iodinated IGF-1 label (IGF-11125 sp act 2000 Ci/mmol; 16,000 cpm/tube) and
Amerlex-M donkey anti-rabbit secondary antibody (code RPN510, 500 pL/tube) were supplied
through from Amersham International (Arlington Heights, IL). Buffer reagents were purchased
from Fisher Chemical Co. (Pittsburgh, PA). Briefly, samples were aliquoted into polypropylene

79
tubes, extracted with 400 pL acid-ethanol, and incubated 30 min prior to centrifugation (2500xG;
4°C; 10 min). For each sample, supernatant (100 pL) was pipetted into duplicate polypropylene
tubes and air-dried. IGF-1 standards were prepared in duplicate with 100 pL of known
concentrations of human recombinant IGF-1 standard (ranging from 9 - 2500 pg/tube), and 300
pL RIA buffer (200 mg/L protamine sulfate, 4.14 g/L sodium phosphate monobasic, 0.05%
TWEEN 20, 0.02% sodium azide, 3.72 g/L EDTA) added to each tube. Air-dried samples were
reconstituted with 350 pL RIA buffer and vortexed. To each sample was added 50 pL IGF-1
antibody (human IGF-1 antisera, UB3-189; 1:10,000 final dilution). After adding 100 pL of
iodinated IGF-1 label (I125-IGF-1; 15,000 cpm) samples were vortexed and incubated (4°C)
overnight. Separation of bound and free IGF-1 was accomplished by incubating samples for 10
min with 500 pL of secondary (2°) antibody (Amerlex-M donkey anti-rabbit secondary antibody,
code RPN.510, Amersham International; 1:10,000 final dilution). Sample tubes were centrifuged
(2500xG; 4°C; 10 min) to separate the secondary antibody, which is bound to the primary
antibody and ligand. The supernatant was decanted and the pellet counted on a Beckman 5500B
gamma counter. Pre-ovariectomy and post-treatment plasma samples were run in two assays
having an average intraassay variance of 5.76% and an interassay variance of 4.19%.
Statistics
Wet tissue mass (mg) of liver, oviduct, and ovary were compared among treatment
groups using ANCOVA, with body mass as a covariate, followed by LSD post-hoc tests. Data are
presented as adjusted means (mg) ± SEM. The oviductal growth parameters were compared
among treatment groups with ANOVA followed by Fishers Protected LSD post-hoc test. Log
transformation of the data was performed in order to achieve homogeneity of variances prior to
ANOVA. Plasma concentrations of E2, T, and IGF-1 were estimated from raw data using the
commercially available Microplate Manager software (Microplate Manager III, BioRad
Laboratories, Inc., Hercules, CA, 1988). For RIA validation of pooled plasma dilutions and

80
internal standards, hormone concentrations were loglO-transformed prior to testing for
homogeneity of slopes with standard curves by ANCOVA. Plasma concentrations of E2, T, and
IGF-1 were compared among treatment groups with one-way ANOVA followed by Scheffe post
hoc. Tamhane post hoc was used where variances were unequal among groups. Statistical
analyses were performed using SPSS software (v. 10, SPSS Inc., Chicago, IL, 1999) with a =
0.05.
Results
Biochemical RIA Validations
Internal standards and plasma dilutions were parallel to the standard curve for E2
(ANCOVA; F = 0.13, P = 0.73 and ANCOVA; F = 0.01, P = 0.91, Fig. 4-4A), and T RIA
(ANCOVA; F = 0.0001, P = 0.99; Fig. 4-4B). Plasma dilutions were parallel to the standard
curve for IGF-1 RIA (ANCOVA; F = 0.01, P = 0.90; Fig. 4-4C).
Tissue Weights
At the time of the ovariectomy surgeries, excised ovaries contained predominantly
mature, highly polarized follicles exhibiting clear demarcation between animal and vegetal
hemispheres. Several of the sham females had slightly immature ovaries with mostly yellow and
some vitellogenic follicles. Ovary mass did not vary significantly among treatment groups
(ANCOVA; F = 0.16, P = 0.16).
Liver mass was not significantly different among treatment groups (ANCOVA; F = 0.48,
P = 0.79). Oviduct mass was greatest in frogs given E2 and E2/IGF-I compared to those given
placebo, EGF, IGF-1, and sham treatments (ANCOVA; F = 5.14, P = 0.001; Fig. 4-5).
Oviduct Morphometries
The wall of the oviduct is composed of three distinct morphological regions: the
endometrium lined with a lumenal epithelium, the myometrium, and the outer serosa layers. The
lumen, or central region of the oviduct, receives secretions synthesized by the epithelial cells and
endometrial glands. The epithelial layer forms a continuous boundary surrounding the oviductal

81
lumen. The endometrial layer, lying internally to the lumenal epithelium, contained secretory
glands, connective tissue, and capillaries. The muscle layer, or myometrium, is composed of
smooth muscle and forms a continuous external boundary around the endometrium. The outer
covering of the oviduct is a relatively thin layer of connective tissue, the serosa. Representative
sections from oviducts of frogs under each treatment group are shown (Fig. 4-6, 4-7).
Regardless of treatment group, the oviducts exhibited a ciliated epithelial layer composed
primarily of cuboidal cells with darkly staining, basal nuclei. The epithelial cell layer appeared
more convoluted in frogs given placebo (Fig. 4-6A), EGF (Fig. 4-6B), IGF-1 (Fig. 4-6C), and
sham (Fig. 4-7C) treatments. For frogs given E2 (Fig. 4-7A) and simultaneous E2/IGF-1 (Fig. 4-
7B) treatment, the epithelial cell layer exhibited little or no convolution, and formed a fairly
straight, continuous boundary around the lumen. Epithelial cell height was greater in E2 and
simultaneous E2/1GF-1 treated frogs compared to controls and other treatment groups (Fig. 4-8A).
Endometrial layer thickness (Fig. 4-8B) and surface area (Fig. 4-8C) were greater for
frogs given E2 and simultaneous E2/IGF treatments compared to other groups, and no difference
in growth as noted between these two treatment groups. The endometrial layer in frogs receiving
these treatments contained numerous large and densely arranged glands. Often the gland height
extended the entire width of the endometrium. Cells containing abundant cytoplasm, darkly
staining nuclei, and a central gland lumen comprised the glands, which also had a duct opening
onto the lumenal epithelium. The oviductal glands in frogs receiving placebo, EGF, IGF-1, and
sham treatments were much smaller in height, width, and total surface area. The endometrial
layer of these frogs was much reduced and connective tissue occupied more relative endometrial
space than did the glands.
Plasma Steroid and IGF-1 Concentrations
Pre-ovariectomy plasma hormones were similar in females among treatment groups for
E2 (P = 0.08), T (P = 0.40), and IGF-1 (P = 0.30). Collectively these data indicate that the
females were in a similar reproductive stage, and had similar plasma steroid and IGF-1

82
concentrations before treatments. Thus, their responses to the treatments are unlikely to have been
obscured by pre-ovariectomy differences in these parameters.
Plasma E2 concentrations were decreased significantly after treatment with placebo (P <
0.001), EGF (P = 0.001), or IGF-1 (P < 0.001) but were similar to pre-ovariectomy
concentrations for E2, E2 /IGF-1, and sham treatment groups (Fig. 4-9). After treatments, plasma
E2 concentrations were greater in E2, E2/IGF-1, and sham female compared to placebo, EGF, and
IGF-1 treatment groups (P < 0.001; Fig. 4-10).
Compared to pre-ovariectomy samples, plasma T concentrations were decreased after
placebo (P = 0.02), EGF (P = 0.01), IGF-1 (P = 0.002), E2 (P = 0.01), and E2/IGF-1 (P = 0.03)
treatment, but not for sham treatment (P > 0.05; Fig. 4-11). Following treatments, plasma T was
higher for only the sham group (P < 0.001; Fig. 4-12).
Compared to pre-ovariectomy samples, plasma IGF-1 was significantly increased in frogs
given IGF-1 (P = 0.0005), E2 (P < 0.001), and E2/IGF-1 (t-test; P < 0.001) but lower for placebo,
EGF, and sham females (t-test; P > 0.05; Fig. 4-13). After treatment, plasma IGF-1 was higher in
IGF-1, E2, and E2/IGF-1 females than in placebo, EGF, and sham females (ANOVA; P < 0.001;
Fig. 4-14).
Ovariectomized frogs given placebo, EGF, and IGF-1 exhibited lower post-treatment
steroid concentrations compared to pre-ovariectomy concentrations, and verify that the
ovariectomy surgeries were successful in removing endogenous ovarian steroid sources. In
addition, similar pre-ovariectomy and post-treatment plasma E2 concentrations for E2 and E2/IGF
treated frogs, and similar pre-ovariectomy and post-treatment plasma IGF-1 concentrations for
IGF and E2/IGF-1 treated frogs indicate that E2 and IGF-1 treatments were delivered effectively
at physiologically relevant concentrations.
Discussion
Results from this study confirm that E2 stimulates oviductal growth in R. catesbeiana.
Treatment with growth factors, placebo, and sham produced no oviduct growth. Lastly, treatment

83
with combined E2/IGF failed to stimulate a greater oviduct growth than was observed with E2
treatment alone. In contrast to reptiles and mammals examined using similar technique, R.
catesbeiana did not exhibit an oviductal growth response with EGF or IGF-1 treatment, nor did
they exhibit a synergistic growth response to E2/IGF-1 treatment. Although E2 and IGF-1 are not
synergistic in stimulation of oviduct growth in R. catesbeiana, both hormones might still be
required for oviductal growth.
There are several possible explanations for the absence of an oviduct growth response in
R. catesbeiana to growth factor (or to combined E2/IGF-1) treatment. First, the IGF-1 treatment
doses might have been insufficient to elicit an oviductal growth response in R. catesbeiana.
Future studies should investigate what doses of IGF-1 are capable of stimulating oviduct growth
in ovariectomized R. catesbeiana. However, it is unlikely that the IGF-1 dose was insufficient
because frogs given IGF-1 exhibited greater post-treatment than pre-ovariectomy plasma IGF-1
concentrations. It is possible that the oviduct must first be "primed" with E2-stimulation before
IGF-1 exposure to become sensitive to the effects of IGF-1. This priming of oviductal tissue
might involve E2-alpha receptors (ERa) and IGF-1 receptors (IGF-1R) upregulation. Klotz et al.
(2000) demonstrated that ERa is required for IGF-1 to induce a cellular response. Additionally,
Clark et al. (1997) demonstrated that E2 stimulates proliferative responses of reproductive tissues
by upregulating IGF-1R expression, which increases tissue response to circulating IGF-1. These
findings imply that an increase in circulatory E2 concentrations can sensitize receptor-dependent
tissue growth IGF-1 stimulation without necessarily requiring an increase in circulating IGF-1
concentrations. As a second explanation, we must consider that circulating steroids can arise from
non-gonadal sources such as the adrenal glands (Norris, 1997). As a third explanation, sensitivity
to these growth factors represents a relatively recent evolutionary change in reptilian and
mammalian oviductal physiology. It is important to recognize that findings reported in this study
might be exclusive to R. catesbeiana. There are likely interspecific differences in hormonal

84
regulation of oviduct growth among amphibians. Accordingly, more amphibian species should be
examined, using similar techniques, before we can fully understand how amphibian oviduct
growth is regulated by interactions of steroids and growth factors.
It is interesting to note that oviduct growth in sham frogs was not similar to growth in E2-
and in E2/IGF-1 treated frogs. Sham frogs were expected to exhibit oviduct growth, similar to E2-
treated frogs but greater than that of placebo frogs, because their intact ovaries would continue to
synthesize and secrete E2 throughout the study. There are several possible explanations for these
unexpected findings. First, it is possible that the implants in E2- and E2/IGF-treated frogs
contained E2 concentrations higher than is typically found in R. catesbeiana. E2 doses were
determined based on studies of E2 necessary to elicit oviduct growth in Xenopus laevis (Follett
and Redshaw, 1967; Redshaw et al., 1968) and in reptiles (Cox, 1994). Thus, these doses might
have been comparatively high for R. catesbeiana. However, this hypothesis seems unlikely
because pre-surgery and post-treatment plasma E2 concentrations were similar for E2- and
E2/IGF-1-treated frogs. A second explanation is that sham frogs were different from E2- and
E2/IGF-1 -treated frogs with respect to pre-surgery and post-treatment plasma IGF-1
concentrations. In E2- and E2/IGF-1-treated frogs, plasma IGF-1 concentrations increased after
treatment compared to pre-ovariectomy levels. In sham frogs, however, post-treatment plasma
IGF-1 concentrations did not increase relative to pre-surgery levels. In E2- and E2/IGF-1 -treated
frogs, the increase in plasma IGF-1 could have stimulated increased IGF-1R expression in
oviductal tissues, making them more sensitive to E2- and IGF-1 stimulation. As mentioned
previously, IGF-1 R does interact, or exhibit "cross-talk" with the ERa in stimulating oviduct
growth (Klotz et al., 2002). Since plasma IGF-1 concentrations in sham frogs did not change
during the experiment, it is possible that oviductal IGF-1 R expression also remained unchanged,
making oviductal tissue comparatively less sensitive to E2- or IGF-1-induced stimulation of
growth. Future studies should examine oviduct growth not only in response to steroid and growth
factors hormones, but in also in response to changes in ERa and IGF-1R activity to better

85
understand the role of these receptors in mediating the effects of E2 and IGF-1 on oviduct growth.
Finally, there might have also been an implant effect on oviduct growth. With or without
hormones, the implant might have elicited oviductal hypertrophy due to an irritation response of
the frogs to implant "foreign body" within the abdomen; this implant effect would have been
absent in sham frogs.
Changes in oviductal mass associated with seasonal changes in plasma steroids have been
described for a wild population of R. catesbeiana (Licht et al., 1984). However, more research is
necessary to understand the mechanism by which E2 induces a growth response in target tissues
of amphibians. In mammals, ovarian steroids induced a complex suite of morphological,
physiological, and biochemical changes in the oviduct (Buhi et al., 1997). Estrogen-induced
oviduct growth relies upon activation of genes that modulate expression of growth factors and
their receptors (Murphy and Murphy, 1994; Cox and Guillette, 1994). Estrogen stimulates DNA
synthesis and mitosis of epithelial cells, and increases uterine IGF-1 and IGF-1R gene expression.
In uterine cells, IGF-1 induces DNA synthesis similar to E2-stimulation. Thus, activation of the
growth factor signaling systems by E2 is an important part of uterine growth and proliferation in
mammals (Klotz et al., 2002; Segars and Driggers, 2002; Driggers and Segars, 2002). It remains
unknown whether E2-induced oviduct growth in amphibians occurs through activation of these
growth factors and their receptors, release of IGF-1 from IGF-1 binding proteins, or by another
mechanism not yet identified.
No study has comprehensively examined the effects of both growth factors and steroids
on oviduct morphology in amphibians. However, both IGF-1 and EGF have been associated with
oviduct growth in mammals and reptiles (Cox and Guillette, 1994; Murphy, 1990; DiAugustine et
al., 1988). In ovariectomized geckos, IGF-1 and EGF stimulates moderate growth of the oviduct
in the absence of E2 indicating these growth factors play an important role in reptilian
reproduction. Also in reptiles, EGF and IGF-1 are known to stimulate oviduct growth directly,
although the exact mechanism for proliferation in not known. In amphibians, IGF-1 and IGF-1

86
binding proteins have been identified in the plasma but the location of IGF-1, IGF-1 binding
proteins, and IGF-1 receptors in reproductive tissues of amphibians have not been examined.
Location and activity of IGF-1 BPs in oviductal tissue, in addition to their interaction with
circulating IGF-1 in amphibians, is necessary to understand how these binding proteins regulate
IGF-1 activity and oviduct growth.
As expected, plasma IGF-1 concentrations increased with IGF-1 and simultaneous
E2/IGF-I treatment. However, an unexpected increase in plasma IGF-1 was observed in females
treated exclusively with E2. An interesting endocrine pathway can be described from these
findings. The liver is the primary site for synthesis of IGF-1 found in the plasma. Perhaps E2
stimulated liver IGF-1 synthesis directly, or E2-stimulated increased pituitary growth hormone
release that, in turn, stimulated liver IGF-1 synthesis. Another possible source for the increased
plasma IGF-1 in E2-treated females is the oviduct. There are an increasing number of studies that
have identified non-hepatic sources of IGF-1 and examined their role in mediating tissue growth.
Numerous studies in reptiles and mammals have shown that the oviduct synthesizes IGF-1 (Cox
and Guillette, 1993; Cox, 1994; Le Roith et al., 2001; Driggers and Segars, 2002; Klotz et al.,
2002; Segars and Driggers, 2002). As part of separate study not described here, IGF-1
immunoreactivity has been detected in the oviduct of R. catesbeiana using immunocytochemistry
(T. Barbeau, unpub. obs.). There is some evidence that oviduct-derived IGF-1 affects the oviduct
itself in an autocrine manner or affects nearby tissues in a paracrine manner. Whether the oviduct
can secrete and contribute significantly to plasma concentrations of IGF-1 remains unknown and
is an intriguing area for future research.
In summary, treatment of ovariectomized R. catesbeiana with exogenous E2 and resulted
in increased plasma IGF-1 concentrations. This finding indicates that E2 interacts with the IGF-1
system in amphibians. It remains unknown if increased hepatic or oviductal IGF-1 synthesis and
secretion contributed to the increase in plasma IGF-1 concentrations observed in E2-treated
females. The mechanism by which E2 stimulates increased plasma IGF-1 concentrations in frogs,

87
and the localization of non-hepatic sources of IGF-1 synthesis in amphibians is poorly understood
and requires further investigation.

88
Figure 4-1. Exteriorized right ovary during ovariectomy surgery in Rana catesbeiana. Both the
left and right ovaries were removed from a 2.5 cm, right paramedial incision into the
abdominal cavity.

89
Figure 4-2. Healed right-paramedical incision site visible three weeks after ovariectomy. Also
shown is the site of the left paramedial incision in which pellet implants were placed
into the abdominal cavity of Rana catesbeiana.

90
12 cm
wl* * * *
*
N . *'Pellet A
implant
t)viduct-
' * 1
• *«
. f .-*• * ,
Figure 4-3. Location of intra-abdominal treatment pellet positioned over the left oviduct at time
of final dissection, after completion of 18 days of treatment in Rana catesbeianaL

(B/Bo) X 100 O (B/Bo) X 100 ® (B/Bo) X 100
91
A 100 ~9 o
80
60
40
20
0
° Standard
♦ Plasma Dilutions
â–  Internal Standards
-+Q» o.
100
80
60
40
20
0
o o
10 100 1000 10000
Estradiol (pg)
° Standard
o ^ ♦ Plasma Dilutions
â–  â–  Internal Standards
% 0 *0 â– 
10 100 1000 10000
Testosterone (pg)
100
80
60
40 -
20
0
° Standard
♦ Plasma Dilutions
I
1
1 10 100 1000 10000
IGF-1 (pg)
Figure 4-4. Biochemical validation of Rana catesbeiana plasma. A. 17(3-estradiol RIA internal
standards (ANCOVA; F = 0.13; P = 0.73) and plasma dilutions (ANCOVA; F =
0.01; P = 0.91) were parallel to the standard curve. B. testosterone RIA internal
standards (ANCOVA; F = 0.0001; P = 0.99) and plasma dilutions (ANCOVA; F =
0.0.08, P = 0.79) were parallel to the standard curve. C. insulin-like growth factor-1
(IGF-1) RIA .the plasma dilutions curve was parallel to the standard curve
(ANCOVA; F = 0.014; P = 0.91).

Mean Oviduct Wet Mass (mg)
92
10000
8000
6000
4000
2000
8
10
10
Placebo EGF
IGF
E2
E2/IGF Sham
Treatment
Figure 4-5. Oviduct weights in Rana catesbeiana after 18 days of placebo (control), epidermal
growth factor (EGF), insulin-like growth factor-1 (IGF-1), 17P-estradiol (E2),
combined E2/IGF-1, or sham treatment Numbers within bars indicate sample size per
treatment while letters above bars indicate significant differences (ANCOVA; F =
5.14, P < 0.001).

93
Figure 4-6. Light microscopy pictures of oviduct morphology for Rana catesbeiana after 18 days
of A. placebo (control), B. epidermal growth factor (EGF), and C. insulin-like growth
factor-1 (IGF-1) treatment. Endo = endometrial layer; Epi = epithelium; G = gland; L
= lumen.

94
of A. 17-P estradiol (E2), B. E2 and insulin-like growth factor-1 (E2/IGF), and C.
sham treatment. Endo = endometrial layer; Epi = epithelium; G = gland; L = lumen.

Gland Surface Area (pm ) O Endometrial Layer Thickness (pm)OJ Epithelial Cell Height (pm) >
95
Placebo EGF IGF E2 E2&IGF Sham
Treatment
Figure 4-8. Oviduct morphology measurements for Rana cates be iana after 18 days of placebo
(control), epidermal growth factor (EGF), insulin-like growth factor-1 (IGF-1), 17(3-
estradiol (E2), combined E2/IGF-I, and sham treatment. Data presented as means ±
SEM. Different letters above bars indicate significant differences for A. epithelial cell
height (ANOVA; P < 0.001), B. endometrial layer thickness ((ANOVA; P < 0.001),
and C. gland surface area (ANOVA; P < 0.001).

Plasma Estradiol (pg/mL)
96
P = 0.23
Treatment
Figure 4-9. Comparison of plasma 17p-estradiol (E2) concentrations for Rana catesbeiana before
and after 18 days of placebo (control), epidermal growth factor (EGF), insulin-like
growth factor-1 (IGF-1), E2, combined E2/IGF-1, or sham treatment. Data presented
as means ± SEM. Numbers within bars indicate sample size per treatment. Parallel
lines above bars, and letters below bars indicate before (B) and after (A) contrasted
pairs. Asterisks and P-values above bars indicate significant differences as
determined by paired t-tests.

Plasma Estradiol (pg/mL)
97
800
700
600
500
400
300
200
100
0
Placebo
b b
EGF IGF
5
Sham
Treatment
Figure 4-10. Plasma 17p-estradiol (E2) concentrations for Rana catesbeiana after 18 days of
placebo (control), epidermal growth factor (EGF), insulin-like growth factor-1 (IGF-
1), E2, combined E2/IGF-1, or sham treatment. Data presented as means ± SEM.
Numbers within bars indicate sample size per treatment. Different letters above bars
indicate significant differences among treatment groups (ANOVA; P < 0.001).

Plasma Testosterone (pg/mL)
98
4000
3500
3000
2500
2000
1500
1000
500
0
P = 0.023
4 4
8
*
rfn
P = 0.012 P = 0.002
4 4 4 4
*
m
9
E3Í
P = 0.014 P = 0.029
4 4 4 4
8
B AB Ag A B A
Placebo EGF IGF E2
Treatment
JL
m
B A
E2/IGF
P = 0.24
4 4
B A
Sham
Figure 4-11. Comparison of plasma testosterone concentrations for Rana catesbeiana before and
after 18 days of placebo (control), epidermal growth factor (EGF), insulin-like
growth factor-1 (IGF-1), 17|3-estradiol (E2), combined E2/IGF-1, or sham treatment.
Data presented as means ± SEM. Numbers within bars indicate sample size per
treatment. Parallel lines above bars and letters below bars indicate before (B) and
after (A) contrasted pairs. Asterisks and P-values above bars indicate significant
differences as determined by paired t-tests.

Plasma Testosterone (pg/mL)
99
2000
1800
1600
1400
1200
1000
800
600
400
200
0
a
É
8
m
JL
Placebo EGF
* m m
IGF E2
Treatment
E2/IGF Sham
Figure 4-12. Plasma testosterone concentrations for Rana catesbeiana after 18 days of placebo
(control), epidermal growth factor (EGF), IGF-1, 17P-estradiol (E2), combined
E2/IGF-1, or sham treatment. Data presented as means ± SEM. Numbers within bars
indicate sample size per treatment. Different letters above bars indicate significant
differences among treatment groups (ANOVA; P < 0.001).

Plasma IGF-1 (ng/mL)
100
25 -i
P = 0.0005
♦ ♦
*
P < 0.001
♦ ♦
P < 0.001
♦ ♦
Placebo EGF IGF E2 E2/IGF Sham
Treatment
Figure 4-13. Comparison of plasma insulin-like growth factor-1 (IGF-1) concentrations for Rana
catesbeiana before and after 18 days of placebo (control), epidermal growth factor
(EGF), IGF-1, 17P-estradiol (E2), combined E2/IGF-1, or sham treatment. Data
presented as means ± SEM. Numbers within bars indicate sample size per treatment.
Parallel lines above bars, and letters below bars indicate before (B) and after (A)
contrasted pairs. Asterisks and P-values above bars indicate significant differences as
determined by paired t-tests.

Plasma IGF-1 (ng/mL)
101
Treatment
Figure 4-14. Plasma insulin-like growth factor-1 (IGF-1) concentrations for Rana catesbeiana
after 18 days of placebo (control), epidermal growth factor (EGF), IGF-1, 170-
estradiol (E2), combined E2/IGF-I, or sham treatment. Data presented as means ±
SEM. Numbers within bars indicate sample size per treatment while letters above
bars indicate significant differences among treatment groups (ANOVA; P < 0.001).

CHAPTER 5
OVARIAN STEROIDOGENESIS (IN VITRO) IN PIG FROGS (Rana grylio) AFTER
EXPOSURE TO ENVIRONMENTALLY RELEVANT CONCENTRATIONS OF NITRATE
AND NITRITE
Introduction
In recent decades, dramatic declines of amphibian populations have been documented
worldwide. Exposure to man-made environmental contaminants is considered an important
contributor to these declines (Blaustein and Wake, 1990; Carey and Bryant, 1995; Hayes et al.,
2002). Recently, nitrate contamination of watersheds has received increasing scientific scrutiny.
Studies indicate that nitrogenous fertilizers contribute to the decline of some amphibian
populations within agricultural landscapes (Berger, 1989; Hecnar, 1995; Baker, 1993; Oldham et
al., 1997; Marco et al., 1999). Unfortunately, more research is needed to understand the harmful
effects of nitrate contamination on amphibian populations.
Nitrate is a natural part of biochemical cycling within terrestrial and aquatic ecosystems.
Nitrate is an anionic form of nitrogen produced by bacterial nitrification. Atmospheric nitrogen
(N2) is fixed by symbiotic bacteria within the root nodules of legumous plants, and by free-living
bacteria in soil and water into ammonia. Ammonia is then converted by nitrifying bacteria
(Nitrosomonas sp.) into nitrite and by Nitrobacter sp. into nitrate (Painter, 1975). Nitrate is the
most stable ionic form of nitrogen that can be assimilated by plants or converted back into nitrite
and nitrogen by denitrifying bacteria. Excessive amounts of nitrite and nitrate can overwhelm the
recycling capacity of nitrifying and denitrifying bacteria, resulting in elevated concentrations of
nitrite and nitrate within the environment.
Sources of excessive nitrogen input into the environment include runoff of nitrogenous
fertilizers, runoff of industrial effluent, leakage of human and animal wastes, and atmospheric
102

103
pollution. Each year, over 10 million tons of nitrogenous fertilizers are used in the US, and over
72 million tons are used globally (International Fertilizer Industry Association, 1993). In the
United States, use of nitrogenous fertilizers increased 20-fold between 1945-1993 (Pucket, 1995)
and currently exceeds 10 million tons. The application of fertilizers in close proximity to
watersheds during the spring often overlaps the breeding season of many amphibians. An
estimated 10-60 % of the nitrogen from fertilizer application remains unused by crops and leaches
into groundwater or washes into surface water as nitrate with rainfall (Ministry of Agriculture,
Fisheries, and Food, 1993). Animal manure contributed approximately 6 million tons of nitrogen
input to the environment. Leakage of human wastes from septic tanks and poorly maintained
sewage systems contributed a small percentage to nitrogen input. Finally, 3 million tons of
nitrogen from atmospheric sources including burning of fossil fuels, automobile exhaust, and
industrial emissions contributed to soil and water nitrogen input (Pucket, 1995). Nitrogen from
these sources is carried by rainfall into surface waters, leached from the soil into ground water,
and deposited into major subterranean aquifers. Since nitrate values are commonly reported in the
literature as nitrate in nitrogen (N03-N), for the remainder of this chapter nitrate refers to NO3-N
(Chapter 1, Table 1-1).
In streams and lakes of North American, nitrate concentrations range from 2 and 40 mg/L
and can persist for long time periods (Rouse et al., 1999). Globally, concentrations of nitrate in
aquatic ecosystems surrounding agricultural and urban landscapes can exceed 100 mg/L (Chilton,
and Foster, 1991; Fried, 1991). The US Geological Survey National Water-Quality Assessment
(NAWQA) Program conducted a 3-year study of 50 prominent hydrologic systems within the
United States to determine the concentration of nitrates in groundwater. Nitrate was detected in
71% of the groundwater sites examined. Shallow groundwater beneath agricultural land had
median nitrate concentrations of 3.4 mg/L nitrate. Of 21 sites where nitrate was examined, 13
sites exceeded the US EPA maximum contaminant limit (MCL) of 10 mg/L nitrate (Nolan and
Stoner, 1995).

104
Nitrate contamination in aquatic ecosystems and in drinking water poses serious health
consequences for wildlife and humans. Nitrate and nitrite enter the body by consumption of food
and water, by crossing the gill epithelia in aquatic organisms (e.g., fishes and tadpoles), and by
absorption through the skin. Nitrate and nitrite are transported against concentration gradients by
substituting for chloride in the bicarbonate-chloride exchange in normal osmoregulatory and
respiratory functions (Lee and Pritchard, 1985; Doblander and Lackner, 1996; Jensen, 1996;
Evans, 1999; Panesar, 1999). These anions can accumulate within extracellular fluid and tissues.
Bacteria within the mouth and gastrointestinal tract convert nitrate into nitrite and N-
nitrosamines. Nitrites are highly toxic and induce methhemoglobinemia, especially in children
exposed to concentrations in drinking water greater than 40 mg/L (US EPA, 1995). Nitrite
oxidizes the iron within the heme subunits of hemoglobin converting it to methhemoglobin.
Methhemoglobin has a reduced ability to transport oxygen and results in cyanosis. For this
reason, the phrase "blue-baby syndrome" is used to describe the condition of infants that develop
methhemoglobinemia from drinking formula made with nitrate-contaminated water (US EPA,
1995). In over 40 animal species examined, including primates, N-nitrosamines have been shown
to be carcinogenic (Weyer et al., 1986). For these reasons, the United States Environmental
Protection Agency (US EPA) has recommended a strict MCL of 10 mg/L for nitrate and 1.0 mg/L
for nitrite in drinking water to (US EPA, 1995). The recommended limit for nitrate in aquatic
ecosystems is 90 mg/L nitrate for freshwater fishes and is based on a lethal exposure values.
Unfortunately these limits are often exceeded due to human activity in agricultural and urban
areas.
Most studies that report the effects of nitrate on amphibians have focused on lethal
concentrations that induce easily observed physiological toxic responses resulting in mortality.
Few studies have examined the effects of exposure to sublethal concentrations of nitrate on
amphibians (Rouse et al., 1999). Sublethal exposure to contaminants can disrupt the endocrine
system in amphibians resulting in altered reproduction, abnormal growth, or deformities (Cooke,

105
1981; Mohanty-Hejmadi and Dutta, 1981; Reeder et al., 1998; Kloas et al., 1999). Studies that
report sublethal effects of nitrate exposure on amphibians have examined only tadpole life-stages
(Table 1-2). Exposure to nitrate concentrations ranging from 2.5 - 100 mg/L nitrate induced
sublethal effects in tadpoles such as reduced feeding activity, nostral and tail deformities, and
erratic swimming behavior. For some species, exposure of tadpoles to nitrate concentrations as
low as 0.78 - 6.0 mg/L nitrite is lethal after 15 days of exposure (Chapter 1, Table 1-2). Variable
responses of amphibians to lethal and sublethal concentrations indicate species differences in
susceptibility to perturbation by nitrate, and effects vary according to life-stages under which
exposure occurs (Marco et al, 1999; De Solía et al., 2002).
Recent research has shown that exposure to relatively high concentrations of nitrate and
nitrite inhibits synthesis (in vivo and in vitro) of testosterone (T) in rodents (Panesar, 1999;
Panesar and Chan, 2000). Mature bulls consuming nitrate-contaminated feed for 30 days also
exhibited decreased plasma T concentrations (Zraly et al., 1997). In alligators, in vivo exposure to
low nitrate concentrations (1.65 mg/L nitrite) increased plasma steroid concentrations but in vitro
exposure of testes to nitrate (1.65 mg/L nitrate) results in decreased T synthesis (Guillette and
Gunderson, unpubl. data). This variable pattern of endocrine disruption associated with in vitro
versus in vivo nitrate exposure indicates that the effects of nitrate depend on route of exposure.
Despite findings of nitrate-induced steroid hormone inhibition in these animals, no study has
examined whether nitrite and nitrate act as endocrine-disruptors in amphibians.
In this study, 1 examined wild-caught Pig frogs (Rana grylio) for steroid hormones, 17(3-
estradiol (E2) and T, synthesized by the ovaries (in vitro) in response to nitrate and nitrite
exposure at environmentally relevant concentrations. Based on the studies already described, I
predicted that nitrate and nitrite inhibits synthesis of steroid hormones.

106
Materials and Methods
Collection of Animals
All work was conducted under the approval of the University of Florida Institute of
Animal Care and Use Committee (IACUC project #Z023). During the breeding season of 2003, a
total of 10 female R. grylio were collected from Orange Lake, Alachua County, Florida. Five
females were collected separately in both June and July. Frogs were transported to the
Department of Zoology and anesthetized, and necropsied, within 12 h of capture. The ovaries
were harvested, weighed, and individual ovarian follicles were dissected for an in vitro culture
study. Ovarian follicles of specific maturation stages 4, 5, and 6 were selected for culture because
they synthesize E2and T (Fortune, 1981).
Ovarian Follicle Culture In Vitro
Preliminary in vitro ovarian follicle cultures demonstrated that 3 h of culture was the
optimum time for measuring steroid concentrations synthesized by R. grylio ovarian follicles (P =
0.007, Figure 5-1). From each frog, 33 follicles each of stages 4, 5, and 6 were incubated under
ambient air conditions in 35x10 mm sterile culture dishes at 29°C with 3 mL of sterile, phenol-
free culture media (1L M199 HBSS, 3.4 mL 200 mM L-glutamine, 5.96 g/L HEPES, 0.35 g/L
sodium bicarbonate, 8.0 mL 0.1 mM IBMX, pH 6.9; Sigma-Aldrich, St. Louis, MO). For each of
the 10 females, 99 follicles were cultured for 3 h under each of the following treatment groups: no
nitrate or nitrite (control), 0.17, 1.65, 6.60, 16.50, and 33.00 mg/L nitrate, and 0.20, 2.03, 8.12,
20.30, and 40.60 mg/L nitrite for a total of 11 treatment groups.
The control treatment group represents steroid synthesis in the absence of nitrate and
nitrite whereas the remaining treatments represent follicular steroid synthesis in the presence of
varying concentrations of nitrate or nitrite. After 3 h of incubation, culture media was decanted,
flash-frozen, and stored at -70°C for RIA analyses of E2 and T concentrations among treatment
groups.

107
Steroid Radioimmunoassay (RIA) Procedures and Validations
RIAs were performed for T and E2 on culture media using validated procedures.
Duplicate media samples (50 pL for T and E2) were twice extracted with ethyl-ether, air-dried,
and reconstituted in borate buffer (0.05 M; pH 8.0). Antibody (Endocrine Sciences) was added at
a final concentration of 1:25,000 for T and 1:55,000 for E2. Radiolabeled steroid ([2,4,6,7,16,17-
3H] estradiol at 1 mCi/mL; [ 1,2,6,7-3H] and testosterone at 1 mCi/mL; Amersham Int., Arlington
Heights, IL) was added at 12,000 cpm per 100 pL for a final assay volume of 500 pL. Interassay
variance tubes were prepared from two separate pools of media and plasma for T and E2.
Standards for T and E2 were prepared in duplicate at 0, 1.56, 3.13, 6.25, 12.5, 25, 50, 100, 200,
400, and 800 pg/tube. Assay tubes were vortexed and incubated overnight at 4°C.
Bound-free separation was performed adding to each tube 500 pL of solution of 5.0%
charcoal to 0.5% dextran, pulse-vortexing, and centrifuging tubes (1500g, 4°C, 30 min).
Supernatant was added to 5.0 mL of scintillation cocktail, and counted on a Beckman scintillation
counter. Media RIA intraassay and interassay variance for E2 averaged 3.31% and 0.987%,
respectively while for T averaged 3.12% and 1.94%, respectively.
Validation of the steroid assay was accomplished with media dilutions (25, 50, 100, 200,
and 300 pL for E2 and 6.25, 25, 50, 100, and 200 pL for T) and with internal standards created by
spiking stripped media samples with E2 and T standards (1.06, 3.125, 6.25, 12.5, 25, 50, 100, 200,
400, and 800 pg per tube). Internal standard and media dilution curves were compared to E2 and
T standard curves to verily homogeneity of slopes. Validation RIA intraassay and interassay
variance for E2 averaged 3.51% and 4.67%, respectively. Validation RIA intrassay and interassay
variance for T averaged 3.00% and 5.58%, respectively.
Internal standards and media dilutions for E2 were parallel to the standard curve (F =
0.36, P = 0.56 and F = 0.002, P = 0.97; Fig. 5-2A). Additionally, internal standards and media
dilutions for T were parallel to the standard curve (F = 0.009, P = 0.93 and F = 0.15, P = 0.71;

108
Fig. 5-2B). Average recovery of E2 was 92.2% and of T was 88.0% after media extractions. Final
media hormone concentrations were compensated to reflect these losses.
Statistics
Concentrations of T and E2 in cultures were estimated from raw data using Microplate
Manager software (Microplate Manager III, BioRad Laboratories, Inc., Hercules, CA, 1988).
Because separate groups of females were collected in June and July, unpaired t-tests were used to
compare hormone concentrations between months, for each treatment group, to verity steroid
synthesis was similar for females collected during both months. Hormone concentrations were
compared among treatments using one-way repeat measures ANOVA followed by Fishers
Protected LSD post-hoc. For RIA validation of pooled media dilutions and internal standards (T
and E2), hormone concentrations were loglO-transformed prior to testing for homogeneity of
slopes with standard curves by ANCOVA. Statistical analyses were performed using SPSS
software (v. 10, SPSS Inc., Chicago, IL, 1999) with a = 0.05.
Results
Analyses revealed similar patterns in the hormonal responses among the treatment groups
for E2 and T (Fig. 5-3). No significant differences in hormone concentrations, per treatment
group, or between months were detected (Table 5-1). Therefore, for comparisons of in vitro
ovarian steroid synthesis among treatment groups, analyses were conducted with a final sample
size of 10 females per treatment group.
After 3 h culture, ovarian E2 synthesis was significantly reduced at all concentrations of
nitrate and nitrite (ANOVA; P < 0.001, Figure 5-4). A similar pattern was observed of reduced
ovarian T synthesis was observed among treatment groups (ANOVA; P <0.001, Figure 5-5). No
clear dose-response pattern in steroid synthesis was observed and hormone synthesis was
similarly inhibited by all nitrate and nitrate concentrations.

109
Discussion
This study provides the first report of nitrate- and nitrite-induced endocrine disruption in
the cultured ovaries of adult, wild-caught frogs. Perhaps most concerning is the finding of
decreased in vitro ovarian steroidogenesis after exposure to sublethal, environmentally relevant
concentrations of nitrate and nitrite; the concentrations used here are commonly detected in the
surface and groundwater of the United States, and also in human drinking water. Decreased
ovarian steroidogenesis (in vitro) in R. grylio was similar to that reported for A", laevis females
exposed to nitrate (in vivo) at comparatively higher concentrations (Chapter 2).
Ovarian steroid synthesis did not exhibit a linear decrease in concentrations, a typical
dose-response curve, with exposure to increasing concentrations of nitrate and nitrite. At 0.17
mg/L nitrate and 0.2 mg/L nitrite, steroid synthesis was already clearly reduced compared to
control treatment but similar to other nitrate and nitrite treatments. This finding indicates that
even lower concentrations of nitrate and nitrite should be examined in future studies. Lower
concentrations might help determine the point at which nitrate and nitrite exposure begins to
inhibit steroid synthesis (in vitro). Conversely, exposure of ovarian follicles to concentrations as
high as 100 or 200 mg/L nitrate and nitrite might demonstrate the point at which steroid synthesis
ceases entirely.
In mammals and reptiles, nitrate and nitrate alter steroidogenesis both in vivo and in vitro.
Panesar and Chan (2000) reported that male rodents given nitrate in drinking water (50 mg/L)
exhibited a marked reduction in plasma T. In vitro culture of mouse Leydig tumor cells with
nitrate and nitrate (1000 mg/L) similarly decreased T synthesis. Adult bulls treated with nitrate in
their feed exhibited decreased plasma T concentrations after 30 days of exposure. In male
alligators, culture of testes with 1.65 mg/L nitrate is associated with decreased T synthesis (in
vitro). However, nitrate exposure of alligators (in vivo), via an i.v. injection, results in increased
plasma steroid concentrations indicating that steroid responsiveness is dependent on route of
exposure (Guillette and Gunderson, unpubl. data). In amphibians, both adult female X. laevis and

110
R. grylio ovarian steroid synthesis is decreased with nitrate exposure. It is unknown if in vivo
exposure of R. grylio results in decreased ovarian synthesis and plasma steroid concentrations
similar to in vitro findings reported here. However, the difficulty in maintaining R. grylio in
captivity for extended time periods makes this kind of comparative study difficult.
A mechanism by which nitrate and nitrite interferes with steroid synthesis has been
proposed for mammals by Panesar and Chan (2000). This mechanism has been modified to
reflect similar pathways of steroid inhibition described for the ovarian follicle of X. laevis
(Chapter 2). Briefly, nitrate reacts with sulfhhydryl groups to create nitric oxide (NO) in most
body tissues (Mayer and Hemmenns, 1997; Panesar and Chan, 2000; Iniguez et al., 2001). Nitric
oxide is a biochemically reactive gas that easily diffuses into tissues and can induce a variety of
cellular responses including vasodilation, smooth muscle relaxation, and steroid regulation
(McDonald and Murad, 1995; Lincolm et al., 1995; Panesar and Chan, 2000; Bastían et al.,
2002). Panesar and Chan (2000) proposed that nitrate either consumed or absorbed into the body
is converted into nitrite by bacteria within the mouth and gastrointestinal tract. Nitrite diffuses
into cells and is converted into NO by activity of nitric oxide synthase (NOS) enzymes found
within most body tissues (Forstermann et al., 1995; Szabo and Thiemermann, 1995; Wang and
Marsden, 1995). Both nitrite and NO can interfere with the iron containing proteins found in
hemoglobin and in heme subunits of cytochrome P450 enzymes (Jensen, 2003). Free cholesterol
is imported, by steroid acute regulatory protein (StAR protein), into the endoplasmic reticulum
and mitochondria of steroidogenic tissue, where cholesterol is converted into progesterone (P4) by
P450-side-chain cleavage (P450scc) and 3P-hydroxysteroid dehydrogenase (3P-HSD) enzymes.
From P4, T is synthesized and then can aromatized into E2 (Chapter 2, Figure 2-7). Research
indicates that NO interferes with or blocks the activity of StAR, 3p-HSD, and P450 enzymes,
thereby inhibiting steroid synthesis at multiple points in the synthetic pathway. Indeed, NO has
been shown to decrease synthesis of E2 and expression of aromatase enzymes in the ovarian

Ill
granulosa cells of humans, rodents, cows, and pigs (VanVoorhis et al., 1994; Olson et al., 1996;
Srivastava et al., 1997; Basini et al., 1998). Thus, nitrate and nitrite have been shown to alter
steroidogenesis in mammals, reptiles, and now in amphibians. The common mechanism for
steroid inhibition in these animals likely involves NO interference of steroidogenic enzymes and
proteins within the gonad.
The present study examined only in vitro gonadal E2 and T synthesis. Progesterone, NOS,
NR, and NO were not analyzed in the culture media or gonadal tissues. Elevated levels of NOS,
NR, and NO in culture media and tissues exposed to nitrate and nitrite compared to control
samples would indicate increased synthesis or activity of these NO-generating agents. For
example, it would be valuable in future studies to examine how NO agonists and inhibitors, given
in conjunction with nitrate and nitrite treatment, affect ovarian steroid synthesis (in vitro). If
steroid synthesis in the presence of nitrate and nitrite remained elevated with the addition of NO
inhibitors, yet is suppressed in the presence of NO donors, this would imply that NO activity is
the causal agent in steroid inhibition.
More research is necessary to evaluate the long-term reproductive impacts of exposure of
amphibians and other vertebrates to nitrate and nitrite. There is already considerable evidence that
nitrate and nitrite exposure decreases reproductive hormone concentrations in a variety of
animals. The discovery of nitrate- and nitrite-induced steroid inhibition in frogs is concerning
because it occurs at concentrations deemed acceptable for human drinking water by current US
EPA standards. Impaired reproductive tissue function must now be considered a potential threat
to reproduction for wild frogs exposed to low concentrations of environmental nitrate and nitrite.
Further studies are needed to determine whether disruption of reproductive hormones by long¬
term exposure to nitrate and nitrite is a prominent factor in the declines in populations of wild
frogs.

112
Table 5-1. Comparisons of in vitro ovarian testosterone and 17P-estradiol concentrations in
culture media with Rana grylio ovarian follicles incubated for 3 h in the absence (0
mg/L) or presence of nitrate and nitrite. For each treatment group, steroid
concentrations were compared between June and July females using T-tests. Steroid
concentrations in table presented as means ± SEM. Significance values in table show
no significant (P > 0.05) differences between June and July means for each treatment
group. (SPSS V. 10.1 (a = 0.05).
June (N = 5)
July (N = 5)
Hormone
Treatment
Mean ± SEM
Mean ± SEM
P-value
0 mg/L
1011.5 ± 67
783.9 ±68
0.25
0.172 mg/L nitrate
451.8 ± 56
369.1 ±32
0.24
1.65 mg/L nitrate
483.0+ 100
478.1 ±117
0.98
6.60 mg/L nitrate
408.4 ±26
364.1 ±21
0.24
16.50 mg/L nitrate
436.5 + 79
387.1 + 59
0.62
Testosterone
33.00 mg/L nitrate
466.9 + 49
399.0 ± 72
0.46
(pg/mL)
0.20 mg/L nitrite
388.61 + 62
421.2 ± 61
0.72
2.03 mg/L nitrite
455.0 ±81
470.3 ± 73
0.89
8.12 mg/L nitrite
414.8 ±52
397.7 ± 43
0.81
20.30 mg/L nitrite
388.8 ±71
515.7 ± 104
0.35
40.60 mg/L nitrite
404.2 ± 65
542.9 ± 127
0.36
0 mg/Lm
165.5 ± 19
200.2 ±19
0.15
0.172 mg/L nitrate
135.3 ±9
118.1 ± 14
0.59
1.65 mg/L nitrate
150.1 ±20
99.9 ± 20
0.13
6.60 mg/L nitrate
148.0 ± 17
93.8 ± 17
0.74
16.50 mg/L nitrate
103.9 ± 12
91.1 + 12
0.52
Estradiol
33.00 mg/L nitrate
121.1 ± 18
108.1 ± 18
0.44
(pg/mL)
0.20 mg/L nitrite
119.1 ±21
111.8 ± 21
0.81
2.03 mg/L nitrite
120.5 ± 17
79.9 ± 19
0.08
8.12 mg/L nitrite
117.3 ± 15
116.7 ± 15
0.99
20.30 mg/L nitrite
104.5 ± 13
91.0 ± 13
0.44
40.60 mg/L nitrite
106.4 ± 15
92.2 ± 15
0.40

Testosterone (pg/mL)
113
3900
3300
2700
2100
1500
0
o
‘ i
1 1 1 1 1
2 4 6 8 10
Time (h)
Figure 5-1. Time trial for in vitro testosterone concentrations synthesized from female Rana
grylio ovarian follicles cultured for time intervals of 3, 5, 7, and 10 h. Data presented
as means ± SEM. Numbers within axis indicate sample sizes and different letters
above the bars indicate significantly different means (ANOVA; P = 0.007).

(B/Bo) X 100 Cd (B/Bo) X 100
114
A
100 o
80
60
40
20
0
o
%
♦
&
♦
° Standard
♦ Plasma Dilutions
â–  Internal Standards
100
80
60
40
20
0
1 10 100 1000 10000
Estradiol (pg)
° Standard
♦ Plasma Dilutions
â–  Internal Standards
T
1
1 10 100 1000 10000
Testosterone (pg)
Figure 5-2. Biochemical validation of media from Rana grylio ovarian follicle culture. A For
17P-estradiol RIA internal standards and plasma dilution curves were parallel to the
estradiol standard curve (ANCOVA; F = 0.36, P = 0.56 and F = 0.0.002, P = 0.97). B.
For testosterone RIA internal standards and plasma dilution curves were parallel to the
testosterone standard curve (ANCOVA; F = 0.009, P = 0.93 and F = 0.05, P = 0.71).

Testosterone (pg/mL) W Media Estradiol (pg/mL)
250
200
150
100
50
0
15
Q June â–¡ July
Í
^íl Í-
¿IT í J
r- — oo^orNioN^omunooooooooomoor^oooo
—«c0‘OV©00O~^rnu~>v0c0t©C'J»OO04,'3'00'OOCN^}-~00r*‘)V©v©
O O o O O — — — ^rov©cov©cooOO~— c-iro'oooc-jotoo
—« m *—■ (N Nitrate
Nitrite
1200
1000 f|
â–¡ June â–¡ July
800
600
400
200
0 41
5
ir
£
JL
Í* f
i1
o r- — oo
o *—■ m «o
o o o o o
O 04 On 'O m »0
OO O ; en «o vo
oooooomooiN
«ooc'i^oovooc'}^ —
vo Nitrate
Nitrite
Treatment (mg/L)
Figure 5-3. A. In vitro 17P-estradiol and B. testosterone concentrations in culture media from Rana grylio ovarian follicles incubated for 3 h in the
absence (O mg/L) or presence of nitrate and nitrite. Data from female R. grylio collected during June and July are presented as means ±
SEM.

Media Estradiol (pg/mL)
250
a
Nitrate
Treatment
Nitrite
Figure 5-4. In vitro 17p-estradiol concentrations in culture media from Rana grylio ovarian follicles incubated for 3 h in the absence (O mg/L) or
presence of nitrate and nitrite. Data presented as means ± SEM. Numbers within bars indicate sample sizes and different letters above
bars indicate significantly different means (ANOVA; P< 0.001).

Media Testosterone (pg/mL)
1200
a
Nitrate Nitrite
Treatment
Figure 5-5. In vitro testosterone concentrations in culture media from Rana grylio ovarian follicles incubated for 3 h in the absence (O mg/L) or
presence of nitrate and nitrite. Data presented as means ± SEM. Numbers within bars indicate sample sizes and different letters above
bars indicate significantly different means (ANOVA; P< 0.001).

CHAPTER 6
THE EFFECTS OF EXPOSURE TO ENVIRONMENTALLY RELEVANT
CONCENTRATIONS ON NITRATE ON OVIDUCTAL MORPHOLOGY AND PLASMA
STEROIDS AND INSULIN-LIKE GROWTH FACTOR-1 IN BULLFROGS {Rana catesbeiana)
Introduction
Nitrate is a widespread contaminant of surface and ground waters worldwide (Halberg,
1989; Nolan and Stoner, 1995; Pucket, 1995). Nitrate is an anionic form of nitrogen that is
produced in soil and water by bacterial nitrification of nitrogen gas, ammonia, and nitrite within
soil and water (Painter, 1975). Of these forms of nitrogen, nitrate is the most water-soluble and
usable by plants. Excessive quantities of nitrate within soil and water often remain largely unused
by plants resulting in accumulation in the environment. Although nitrate is less toxic than
ammonia and nitrite, it is relatively stable in the environment and can persist in high
concentrations for decades (Nolan and Stoner, 1995). Nitrate concentrations in ground and
surface waters of the United States have been rising for the past 20 years (Nolan and Stoner,
1995). Nitrates infiltrate ground and surface waters primarily from runoff of inorganic fertilizer,
runoff of animal wastes from pastures and feedlots, and leakage of improperly contained human
sewage (Nolan and Stoner, 1995; Berndt et al., 1998). Nitrate values are commonly reported in
the literature as nitrate-as-nitrogen (N03-N), and this is how nitrate is reported for the remainder
of this chapter.
Nitrate contamination poses serious health consequences for wildlife and humans alike.
When nitrate enters the body, either through consumption or absorption across epithelial surfaces,
bacteria within the mouth and gastrointestinal tract convert it into harmful nitrite and N-
nitrosamines. Nitrite is highly toxic and induces methhemoglobinemia while N-nitrosoamines are
known to cause cancer in over 40 animal species examined. Based on these studies, the US EPA
established a strict maximum contaminant limit (MCL) for nitrate in human drinking water of 10
118

119
mg/L nitrate. However, wells examined in and around agricultural landscapes frequently exceed
this concentration (Nolan and Stoner, 1995). For example, people drinking water from nitrate-
contaminated wells exhibit increased incidence of bladder, brain, and colon cancer, and also Non-
Hodgkin's lymphoma (Ward et al., 1996, 2000; Weyer et al., 2000; Gulis et al., 2002).
In recent decades, researchers have been concerned over the deformities and the apparent
declines in populations of amphibians worldwide (McCoy, 1994; Pounds and Crump, 1994;
Laurance et al., 1996; Licht, 1995). Exposure of amphibians to man-made environmental
contaminants is considered a prominent contributor to incidents of declines and deformities
(Carey and Bryant, 1995; Carey et al., 1999; Hayes et al., 2000). Exposure to contaminants
disrupts the endocrine system in amphibians resulting in altered reproduction, abnormal growth,
or deformities (Chapter 1, Table 5-1). Research indicates that nitrogenous fertilizers contribute to
the decline of some amphibian populations located in agricultural lands (Berger, 1989; Hecnar,
1995; Baker, 1993; Oldham et al., 1997). Unfortunately, most studies have addressed the effects
of toxicological rather than sublethal concentrations of nitrates on amphibians. Amphibians
exposed to nitrate concentrations ranging from 2.5 - 100 mg/L exhibit sublethal responses that
are subtle compared to responses to toxic concentrations (Chapter 1, Table 1-2). Tadpoles of the
Common toad (Bufo bufó) exposed to 50 - 100 mg/L nitrate exhibited delayed metamorphosis
compared to unexposed tadpoles. Additionally, exposed toads exhibited unusual swimming
movement, nostral deformities, and absent or deformed limbs (Xu et al., 1997). Cascades frog
(Rana cascadae) tadpoles exposed to 3.5 mg/L nitrite also had delayed time to metamorphosis
compared to controls (Marco and Blaustein, 1995). For tadpoles of some other anuran species,
exposure to similarly sublethal concentrations is lethal. The variation in responses of tadpoles to
nitrate exposure (at similar concentrations) indicates that there are species differences in
susceptibility to perturbation by nitrate (Marco et al, 1999; De Solía et al., 2002).
Nitrate exposure has been shown to inhibit testosterone (T) synthesis in rodents (Panesar
and Chan, 2000), and bulls (Zraly et al., 1997). Alligators exposed to nitrate at low concentrations

120
(1.65 mg/L) exhibited increased plasma steroid concentrations (in vivo) but testes exposed to
directly to nitrate (in vitro) exhibited decreased testosterone synthesis (Guillette and Gunderson,
unpubl. data). Thus, steroid synthesis in response to nitrate exposure can result in variable
responses depending on the route of exposure. A mechanism for the disruption of steroidogenesis
by nitrates has been proposed by Panesar and Chan (2000), and has been expanded by other
researchers. Many body tissues contain nitric oxide synthase (NOS) enzymes that can convert
nitrate into nitrite into NO (Forstermann et al., 1995; Szabo and Thiemermann, 1995; Wang and
Marsden, 1995). In mammalian ovaries NO has also been shown to inhibit E2 synthesis, decrease
aromatase activity, and increase insulin-like growth factor 1 (IGF-1) synthesis (Erickson et al.,
1989; Adashi, 1993; Van Voorhis et al., 1994; Olson et al., 1996; Samaras et al., 1996). Thus,
nitrates, via conversion into NO within body tissues, could influence steroid and IGF-1 synthesis
and action.
Insulin-like growth factor 1 (IGF-1), a polypeptide hormone, plays an integral role in
normal reproduction, cell growth, tissue differentiation, and embryo development in amphibians
as in reptiles, birds, and mammals (Pancak-Roessler and Lee, 1989; Smith et al., 2000; Werner
and LeRoith, 2000; Allen et al., 2001). Few researchers have examined whether environmental
contaminants, like nitrate, alters the synthesis or action of IGF-1, despite the importance of this
hormone in normal vertebrate development and growth. Abnormally high expression of IGF-1
mRNA is associated with abnormal limb bud development and emergence in mammals and birds,
and with cancer formation in somatic and reproductive tissues of humans and other vertebrates
(Backlin et al., 1998; Allen et al., 2000; Grimberg and Cohen, 2000; Klotz et al., 2000). Increased
IGF-1 expression, induced by estrogen exposure, induces fetal malformation, abnormal
proliferation of uterine tissues, and formation of reproductive lesions in aquatic mammals
(Backlin and Bergman, 1995; Backlin et al., 1998). Evidence presented in Chapter 4
demonstrated that E2-treated ovariectomized R. catesbeiana exhibited increased plasma IGF-1
concentrations. Thus, E2 and IGF-1 apparently have interrelated functions. Because nitrate has

121
been shown to alter E2, it is probable that nitrate can also alter IGF-1. Despite findings of skeletal
and tissue deformities in tadpoles after exposure to fertilizers (Jones et al., 1997; Lutz and Kloas,
1999), no study has examined whether nitrate alters reproductive steroids and IGF-1 in
amphibians. Evidence presented within earlier chapters (Chapter 2 and 5) indicates that exposure
to sublethal concentrations of nitrate alter reproductive tissues, increases plasma IGF-1
concentrations while decreasing ovarian steroid synthesis. Findings of nitrate-associated
alteration of IGF-1 hormone and reproductive steroids might introduce a novel mechanism by
which contaminants alter reproduction and growth, and contribute further insight into factors
implicated in the apparent global decline of amphibians.
In this study, I examined the effects of sublethal concentrations of nitrate, administered in
vivo to adult bullfrogs (Rana catesbeiana), on reproductive tissue, plasma steroid (E2 and T)
concentrations, and plasma IGF-1 concentration. Based on the aforementioned studies, I
hypothesized that exposure of adult bullfrogs (Rana catesbeiana) to environmentally relevant
concentrations of nitrate alters reproductive tissues, and plasma concentrations of sex steroids (E2
and T) and IGF-1.
Materials and Methods
Animals
All work was conducted under IACUC approved guidelines (project #Z023). For this
study, R, catesbeiana were chosen in lieu of A. laevis and R. grylio, which were examined in
earlier chapters. Previous attempts to maintain wild-caught R. grylio in captivity demonstrated
that they were highly stressed in captivity and were inappropriate subjects for the 10-day study
outlined here. In contrast to R. grylio, R catesbeiana were very adaptable and exhibited no
observable stress in captivity. Additionally, R. catesbeiana are large-bodied frogs, which made it
easier to collect a greater blood sample volume than in X. laevis and R. grylio. Oviduct growth
response to normal physiological concentrations of E2 and IGF-1 had already been established for
R. catesbeiana (Chapter 4). Finally, normal concentrations of plasma steroids and IGF-1 had

122
already been determined for R. catesbeiana in (Chapter 4) and by Licht et al. (1984), and these
studies served as references.
Adult Rana catesbeiana (30 females and 18 males) were purchased (Charles Sullivan Co.
Inc., TN), maintained in 38 L tanks with 19 L of static flow, dechlorinated water at 26°C under a
12-h diurnal light/dark cycle. Tank water was changed and frogs were fed crickets every other
day for the duration of the experiment. Frogs were randomly assigned to nitrate treatment groups.
Each treatment group of either six female or six male frogs was separated into two replicate tanks
each containing three frogs to avoid pseudo-replication. Frogs were maintained under these
conditions for a 3-week acclimation period before the onset of the experiment.
Nitrate Treatments
Sodium nitrate (NaN03) was dissolved in deionized water to produce nitrate as nitrogen
(N03-N) treatment concentrations of 0, 0.17, 1.65, 6.60, and 16.50 mg/L for females and 0, 1.65,
and 6.60 mg/L for males. These concentrations are environmentally relevant and also include
concentrations allowed in human drinking water according to standards established by the US
EPA.
Rather than dissolving nitrate into tank water for whole-body exposure of R. catesbeiana,
treatments were delivered as intra-abdominal injections. Unlike the fully aquatic Xenopus laevis
(chapter 1), ranid frogs are semi-aquatic. Laboratory observations of R. catesbeiana have shown
that they spend prolonged periods of time out of the water and display considerable stress (escape
behavior and nostral abrasions) when denied access to basking platforms (T. Barbeau, pers. obs.).
Whole-body exposure of frogs to aqueous nitrate would require removal of basking platforms to
maintain continuous exposure to the treatments. Preventing R. catesbeiana from engaging in
normal basking behavior could make it difficult to distinguish a stress response from other
physiological responses to experimental treatments. If basking platforms were not removed from
the treatment tanks, basking frogs would remain unexposed to aqueous nitrate treatments for
variable time periods, making it impossible to determine their treatment times. To avoid these

123
confounding variables, nitrate treatments were dissolved in sterile, deionized water and delivered
via intra-abdominal injections (0.5 ml) into the ventral paramedial surface as described by Wright
and Whitaker (2001).
Before initiating the main experiment, 3 extra bullfrogs were given intra-abdominal
nitrate injections (16.50 mg/L) daily for 1 week to confirm that the highest concentration and the
delivery route were not toxic or lethal. Frogs exhibited normal behavior and feeding during the
treatment period, and also for 2 weeks after treatments. Based on these preliminary findings, the
nitrate concentrations and delivery route chosen were deemed safe for R. catesbeiana in this
experiment.
Frogs were injected with nitrate every other day, for a total of 5 injections. This treatment
regime was considered an acute exposure due to the direct route of administration and the relative
brevity of treatment length. Intravenous injections of nitrate (at similar concentration), given to
adult alligators have demonstrated that such brief exposure times are sufficient to elicit an
endocrine response (Gunderson and Guillette, unpubl. obs). After treatment, frogs were
anesthetized with MS-222 (1.5% 3-aminobenzoic acid ethyl ether, Aquatic Ecosystems, Orlando,
FL), blood samples were collected via cardiac puncture, and the frogs were euthanized by
dissection through the spinal cord followed by pithing. Plasma was frozen (-70°C) for E2, T, and
IGF-1 radioimmunoassay (RIA) analyses. The ovaries, oviducts, liver, spleen, and fat bodies
were removed from females, and the testes, liver, spleen, and fat bodies were removed from
males, and tissues were weighed to determine wet mass among treatment groups. Cross sections
of oviducts (ampulla region) were fixed in 4% paraformaldehyde (4°C; 48 h) followed by rinse
and storage in 75% ethanol for subsequent histological analyses. The oviducts were dehydrated in
a graded series of ethanol changes, embedded in paraffin, serially sectioned on a rotary
microtome (7 pm), stained with hematoxylin and eosin, and examined using light microscopy. To
ascertain oviductal proliferation among treatment groups, an ocular micrometer was used to make

124
10 morphological measurements on five tissue sections per frog, for each of the following
oviductal parameters: epithelial cell height, endometrial thickness, endometrial gland height, and
endometrial gland width. Endometrial gland height and width measurements were used to
calculated total gland surface area.
Steroid Radioimmunoassay (RIA) Procedures and Validations
Validation of steroid RIA was accomplished with pooled plasma dilutions for females
and males (25, 50, 100, 200, and 300 pL for E2 and 6.25, 25, 50, 100, and 200 pL for T), and
borate buffer was used to bring all samples up to 200 pL total volume. For internal standards, 100
pL aliquots of steroid-stripped plasma were spiked with standards for E2 and T (1.06, 3.125, 6.25,
12.5, 25, 50, 100, 200, 400, and 800 pg/tube). Plasma dilution curves and internal standard curves
were compared with E2 and T standard curves. Before analyses, hormone concentrations were
loglO-transformed and ANCOVA was used to verify homogeneity of slopes with the standard
curves for plasma dilutions and for internal standards.
Internal standards and plasma dilutions were parallel to the standard curve for E2 in
females (ANCOVA; F = 0.03, P = 0.73 and F = 0.01, P = 0.91, Fig. 6-1 A) and males (ANCOVA;
F = 0.001, P = 0.99 and F = 1.86, P = 0.22, Fig. 6-2A), and for T in females (ANCOVA; F =
0.001, P = 0.99 and F = 0.08, P = 0.79; Fig. 6-1B) and males (ANCOVA; F = 0.003, P = 0.96 and
F = 0.37, P = 0.56, Fig. 6-2B). For females and males, average recoveries after plasma extractions
for E2 were 93.9% and 85.0%, for T were 87.9% and 94.1%, respectively. Final media hormone
concentrations were changed to reflect these losses. Finally, E2 and T validation RIA intraassay
variance averaged 2.87% and 4.88%; and RIA samples for E2 and T were each run in 2 assays
having an average intraassay variance of 1.53% and 1.23%, and interassay variance of 7.84% and
2.56%, respectively.
RIAs were performed for E2 and T on plasma samples using validated procedures. Based
on RIA validations, 50, 6.25, and 50 pL of female plasma were used while 300, 12.5, and 50 pL

125
of male plasma were used for E2, T, and IGF-1 analyses, respectively. Duplicate plasma samples
were extracted twice with ethyl-ether, air-dried, and reconstituted in 100 uL borate buffer (0.05
M; pH 8.0). Antibody (Endocrine Sciences) was added at a final concentration of 1:25,000 for T
and 1:55,000 for E2. Radiolabeled steroid ([2,4,6,7,16,17-3H] estradiol at 1 mCi/ml; [1,2,6,7-3H]
testosterone at 1 mCi/mL; Amersham Int., Arlington Heights, IL) was added at 12,000 cpm per
100 pL for a final assay volume of 500 pL. Interassay variance tubes were prepared from two
separate pools of media and plasma for T and E2. Standards for T and E2 were prepared in
duplicate at 0, 1.06, 3.13, 6.25, 12.5, 25, 50, 100, 200, 400, and 800 pg/tube. Assay tubes were
vortexed and incubated at 4°C for 24 h. Bound-free separation was performed using a mixture of
5% charcoal and 0.5% dextran, pulse-vortexing, and centrifuging tubes (1500g, 4°C, 30 min).
Supernatant was added to 5 mL of scintillation cocktail, and counted on a Beckman scintillation
counter. For E2 and T RIA, male and female plasma samples were analyzed within a single assay
with an average intraassay variance of 4.02% and 5.42%, respectively.
Insulin-Like Growth Factor-1 (IGF-1) RIA Procedures and Validations
Validation of IGF-1 RIA was accomplished with 300 pL of pooled plasma dilutions for
females and males. Plasma was extracted in polypropylene tubes with acid-ethanol (12.5% 2 N
HC1, 87.5% ethanol; 800 pL) to dissociate IGF binding proteins from the IGF-1 molecules and to
precipitate globular proteins (Daughaday et al., 1980; Crain et al., 1995). After 30 min incubation
(room temperature) and 10-min centrifugation (2500xG; 4°C), the supernatant was aliquoted to
produce plasma equivalents of 12.5, 25, 50, 100, 200, and 300 pL. Volume of the plasma
dilutions was brought to 200 pL with acid-ethanol prior to air-drying. Plasma dilutions were
compared with 0, 39, 156, 313, 625, 1000, 1250, 2500 pg of human recombinant IGF-1 standard
(National Hormone and Pituitary Program, Torrance, CA 90509). Validation samples were
examined by IGF RIA procedures as described for experimental analyses below.

126
Plasma extraction efficiencies were determined by adding 100 pL iodinated IGF-1
(15,000 cpm) to 100 pL of pooled plasma samples, extracting with acid-ethanol, air-drying, and
reading samples on a Beckman 5500B gamma counter to determine the iodinated hormone
remaining. The extraction efficiency of plasma was 77.0%, and all sample concentrations were
corrected for this loss. Validation of plasma dilutions for IGF-1 was accomplished in a single
assay having an intraassay variance of 2.27%.
Plasma dilutions were parallel to the standard curve for IGF-1 in females (ANCOVA; F =
0.01, P = 0.91, Fig. 6-1C) and males (ANCOVA; F = 5.14, P = 0.10, Fig. 6-2C). For IGF-1
validation RIA and sample RIA were each run in a single assay with an average intraassay
variance of 2.32% and 4.18%, respectively. For IGF-1, average recoveries after plasma
extractions for females and males were 78.0% and 76.0%, respectively.
IGF-1 RIA was performed as described by Crain et al. (1995) and Guillette et al. (1994).
The National Hormone and Pituitary Program (Torrance, CA 90509) supplied human
recombinant IGF-1 standard (9.76 to 2500 pg/tube), and human IGF-1 antisera (Lot #
AFP4892898, 1:400,000 final dilution). The antiserum had less than 1.0% cross-reactivity with
human IGF-II. Iodinated IGF-1 label (IGF-11125 sp act 2000 Ci/mmol; 16,000 cpm/tube) and
Amerlex-M donkey anti-rabbit secondary antibody (code RPN510, 500 pl/tube) were supplied
through from Amersham International (Arlington Heights, IL). Buffer reagents were purchased
from Fisher Chemical Co. (Pittsburgh, PA). Briefly, samples were aliquoted into polypropylene
tubes, extracted with 400uL acid-ethanol, and incubated 30 min prior to centrifugation (2500xG;
4°C; 10 min). For each sample, supernatant (100 p.L) was pipetted into duplicate polypropylene
tubes and air dried. IGF-1 standards were prepared in duplicate with 100 pL of known
concentrations of human recombinant IGF-1 standard (ranging from 9 - 2500 pg/tube), and 300
pL RIA buffer (200 mg/L protamine sulfate, 4.14 g/L sodium phosphate monobasic, 0.05%
TWEEN 20, 0.02% sodium azide, 3.72 g/L EDTA) added to each tube. Air-dried samples were

127
reconstituted with 350 p.L RIA buffer and vortexed. To each sample, 50 pL IGF-1 antibody
(human IGF-1 antisera, UB3-189) was added at a 1:10,000 final dilution. After adding 100 pL of
iodinated IGF-1 label (II25-IGF-1), with -15,000 CPM, samples were vortexed and incubated
overnight at 4°C. Separation of bound and free IGF-1 was accomplished by incubating samples
for 10 min with 500 pL of secondary antibody (Amerlex-M donkey anti-rabbit secondary
antibody, code RPN.510 obtained from Amersham International) at a final dilution of 1:10,000.
Sample tubes were centrifuged (2500xG; 4°C; 10 min) to separate the secondary antibody, which
is bound to the primary antibody and ligand. The supernatant was decanted and the pellet counted
on a Beckman 5500B gamma counter. Female and male plasma samples were run in one assay
having an average intraassay variance of 2.32%.
Statistics
Concentrations of E2 and T were estimated from raw data using Microplate Manager
software (Microplate Manager III, BioRad Laboratories, Inc., Hercules, CA, 1988). Unpaired,
two-tailed t-tests revealed no significant differences (P > 0.05) in mean hormone concentration or
tissue mass between females and males from replicate tanks per treatment group. Additionally, no
significant differences were detected in female oviduct morphology parameters between replicate
groups (P > 0.05). Thus, for final analyses, parameter measurements from females and males in
replicate treatment tanks were lumped together for a total sample size of six frogs per treatment
group.
Hormone concentrations among treatment groups were analyzed by ANOVA followed by
Fishers Protected LSD contrasts. For E2 concentrations in females and males, and for T
concentrations in females, variances were unequal among treatment groups; therefore, ANOVA
was followed by Tamhane contrasts. Oviduct, liver, spleen, and fat body masses among treatment
groups were analyzed by ANCOVA using body mass as a covariate. Lastly, morphological tissue
measurements of oviducts were analyzed by ANOVA followed by Tamhane contrasts to account

128
for unequal variance among treatment groups. Statistical analyses were performed using SPSS
software (v. 10, SPSS Inc., Chicago, IL, 1999) with a = 0.05.
Results
Oviduct Weights
For females, there was no significant difference in wet tissue mass, with body weight as a
covariate, among treatment groups for liver (ANCOVA; F = 1.25, P = 0.32), ovaries (ANCOVA;
F = 0.53, P = 0.72), oviducts (ANCOVA; F = 0.76, P = 0.56), or fat bodies (ANCOVA; F = 0.54,
P = 0.70). For males, there were no significant differences in wet tissue mass among groups for
liver (ANCOVA; F = 1.73, P = 0.22), testes (ANCOVA; F = 1.34, P = 0.29), or fat bodies
(ANCOVA; F = 3.68, P = 0.06).
Plasma Steroid and IGF-1 Concentrations
Plasma E2 and IGF-1 concentrations were increased in females exposed to 1.65, 6.60, and
16.50 mg/L nitrate compared to females in control and 0.17 mg/L groups (ANOVA; P < 0.001,
Fig. 6-3A,C). Plasma T concentrations in all females exposed to nitrate (0.17, 1.65, 6.60, and
16.50 mg/L) were higher than in control females (ANOVA; P < 0.001, Fig. 6-3B). Rather than a
typical dose-response curve, plasma concentrations of E2 and T displayed a non-monotonic dose
response curve. For both steroids, and initial increase in plasma steroids was seen at 1.65
followed by a decrease at 6.60 mg/L and an increase at 16.50 mg/L. Plasma E2 was elevated in
male frogs exposed to 1.65 mg/L but not to 0 mg/L and 6.60 mg/L nitrate. Plasma T was similar
in males among treatment groups. In males, plasma IGF-1 was higher with exposure to both 1.65
mg/L and 6.60 mg/L nitrate compared to control males (ANOVA; P < 0.001, Fig. 6-4).
Oviduct Morphometries
The epithelial layer bordering the oviductal lumen is a continuous and convoluted layer
of ciliated, cuboidal cells with darkly staining basal nuclei and an apical brush-border of cilia

129
projecting into the lumenal space. The epithelial cilia are relatively short (< 15 pm), uniform, and
numerous.
The epithelial cell heights were lowest in females exposed to 6.60 mg/L, and greatest in
those exposed to 0.17 mg/L and 1.65 mg/L (Fig. 6-5 A). However, epithelial cell heights were
similar in females exposed to 0 and 16.50 mg/L nitrate. Endometrial thickness was significantly
lower in females exposed to all nitrate doses with the lowest measures in the 16.50 mg/L group
(Fig. 6-5B). Overall, endometrial gland area (pm2) was lower in females exposed to nitrate (Fig.
6-5C).
Discussion
In recent decades, high concentrations of nitrate in watersheds of North American might
reflect an increase in nitrogen input from many sources including runoff of nitrogenous fertilizers.
Increased nitrogen contamination of aquatic ecosystems poses severe health consequences to
domestic and wild animals, and perhaps even humans. Rodents exhibit altered steroidogenesis,
both in vivo and in vitro, when exposed to nitrate (Panesar and Chan, 2000). Rodents consuming
8.25 mg/L nitrate-as-as-nitrogen in their drinking water, for 4 weeks, exhibited decreased plasma
T concentrations (in vivo). Testes of rodents incubated for 1 h with 1726 mg/L nitrate (in vitro)
exhibited decreased T synthesis (Panesar and Chan, 2000). Nitrate exposure also inhibits steroid
synthesis in other mammals, including humans (Wang and Marsden, 1995) and bulls (Zraly et al.,
1997). Bulls consuming potassium nitrate (100 - 250 g) in their feed for 30 days exhibited a
decreased testicular T synthesis (in vitro), in response to gonadotropin stimulation (Zraly et al.,
1997). The proposed mechanism for nitrate-associated endocrine disruption steroid involves the
following complex pathway. Nitrate is converted into nitrite by bacteria within the mouth and
gastrointestinal tract (Painter, 1975; Mayer, 1997). Nitrite is then converted into NO by various
isoforms of nitric oxide synthase (NOS) found in body tissues (Kukovetz et al., 1987; Kleinert et
al., 1995; Ellis et al., 1998). The NO is a highly diffusible compound that can enter most cells and

130
tissues to induce a variety of physiological effects. NO reacts readily with metals like iron and
iron-sulfur centers within heme-containing enzymes, such as all the enzymes of the P450
superfamily. Most interactions of NO result in inhibition of these enzymes, with the exception of
guanylate cyclase, that is stimulated by NO (McDonald and Murad, 1995). The mitochondrial
enzymes P450 side-chain cleavage (P450scc) and 3P-hydroxysteroid dehydrogenase (3P-HSD)
contain heme subgroups that are inhibited by NO (Delaforge et al., 1995; Del Punta et al., 1996).
These enzymes are necessary to convert free cholesterol within the mitochondria into
progesterone (P4), the steroid precursor to T. After T is synthesized it is converted into E2 by
aromatase enzymes (Chapter 1, Fig. 1-1). Additionally, NO interferes with steroidogenic acute
regulatory (StAR) protein necessary for transport of free cholesterol into the mitochondria (Wang
and Marsden, 1995; Stocco, 1999). Thus, NO likely disrupts steroidogenesis by inactivation of
P450 and 3P-HSD enzymes and interference with StAR protein (VanVoorhis et al., 1994, 1995;
Panesar, 1999; Stocco, 1999; Panesar and Chan, 2000).
Nitric oxide inhibits steroid synthesis and aromatase gene expression within the gonads
of humans, rodents, pigs, and cows (Adashi, 1993; VanVoorhis et al., 1994; Olsen et al., 1996;
Snyder et al., 1996; Srivastava et al., 1997; Basini et al., 1998). Despite findings of nitrate-
associated endocrine disruption in other animals, no previous study examines whether nitrate
alters endocrine function in amphibians. Most studies on the effects of sublethal and lethal nitrate
concentrations on amphibians have focused on tadpoles. Nitrate exposure reportedly alters
growth, development, feeding, swimming, metamorphosis, and mortality rates of tadpoles both in
the field and the laboratory. Only one study reports the effects of lethal nitrate concentrations on
adult amphibians (Oldham et al., 1997). No published investigations have reported the effects of
exposure to sublethal nitrate concentrations on endocrine function in amphibians.
This study presents the first findings of an association between altered plasma steroids
and IGF-1 and oviductal atrophy in R. catesbeiana after acute exposure to sublethal nitrate

131
concentrations. Furthermore, nitrate exposure is associated with endocrine disruption in R.
catesbeiana at concentrations that are environmentally relevant and lower than the MCL allowed
in drinking water. In female R. catesbeiana, plasma E2 and T concentrations did not exhibit a
typical dose-response increase with increasing nitrate concentrations. Rather, the steroid response
more closely resembled an inverted U-shaped non-monotonic dose response (NMDR) for nitrate
concentrations below 16.50 mg/L. E2 and T concentrations are increased at 1.65 mg/L nitrate but
then decreased with a slightly higher nitrate concentration of 6.60 mg/L. At 16.50 mg/L, E2 and T
concentrations increase again. A similar NMDR was seen for male R. catesbeiana at similar
nitrate concentrations. The NMDR curve is commonly seen with studies of endocrine disruption
and reflects greater sensitivity of steroid receptors to endocrine-disrupting contaminants (EDCs)
present at extremely low concentrations. At extremely low EDC concentrations, many steroid
receptors remain unoccupied and inactive. Only slight increases in the EDC concentrations can
dramatically increase receptor binding up to 100 percent. Afterwards, any further increases in
EDC concentrations cannot lead to a greater steroid response because all the steroid receptors are
bound (vom Saal et al., 1997; Welshons et al., 2003).
In mammals nitrate exposure (in vivo) is associated with decreased plasma and gonadal
steroid concentrations (Panesar and Chan, 2000; Zraly et al., 1997). In X. laevis, nitrate exposure
(in vivo) is associated with a significant decrease in gonadal steroid synthesis (Chapter 2). These
findings in X. laevis were similar to steroid responses of mammals exposed to nitrate through
consumption of nitrate in water (Panesar and Chan, 2000) and feed (Zraly et al., 1997). Rana
grylio ovarian follicles exposed to nitrate and nitrite also exhibited decreased steroid synthesis (in
vitro). The decreased steroid concentrations (ex vivo and in vitro) reported for A. laevis and R.
grylio (Chapters 2 and 5) conflict with the findings of increased plasma steroid concentrations in
R. catesbeiana with nitrate exposure (in vivo). In this study, the use of intra-abdominal nitrate
injections might have influenced the steroid hormone response by a mechanism other than that
seen with exposure of the gonad (in vitro) or ingestion of nitrate (in vivo). For example, studies in

132
alligators have shown that nitrate exposure (via an intravenous injection) stimulated increased
plasma steroid concentrations; however, direct incubation of the testes with nitrate (in vitro)
causes decreased gonadal steroid synthesis (Guillette and Gunderson, unpubl. obs.). Thus, the
route of nitrate exposure apparently influences steroid responses.
The effects of in vivo nitrate exposure likely involve a integrated physiological response
that is highly complex compared to the responses of isolated tissues cultured with nitrate (in
vitro). Circulating hormone concentrations can change in response to signals from the
hypothalamic-pituitary-gonadal (HPG) axis, in response to interaction with plasma binding
proteins, and also in response to steroid metabolism and clearance by the liver (Chapter 1, Fig. 1-
1). Gonadal steroid synthesis (in vitro or ex vivo) was demonstrated in earlier studies to decrease
with nitrate exposure; however, nitrate exposure can influence plasma steroid concentrations (in
vivo) by affecting other organs. A severe depression of liver cytochrome P450 enzyme activity
could decrease steroid metabolism and clearance, and contribute to elevated plasma steroid
concentrations (Fig. 2-7). One recent study has shown that rats administered organic nitrate
exhibit decreased activity of hepatic P450 enzymes within 24 h of exposure (Minamiyama et al.,
2004). Zraly et al. (1997) reported that bulls fed potassium nitrate exhibited increased
concentration of plasma bile acids, decreased progesterone metabolism, and abnormal liver
morphology that collectively signify impaired liver function in response to nitrate exposure.
These findings also support a pathway, proposed here, for the increase in plasma steroid and IGF-
1 concentrations observed in R. catesbeiana in response to nitrate exposure (in vivo). The liver
degrades the majority of nitrate consumed and circulating in the body. Nitrate exposure has been
shown to decrease activity of hepatic cytochrome P450 enzymes, perhaps via formation of NO.
Since hepatic P450 enzymes are necessary to metabolize and clear steroid from circulation,
inhibition of these enzymes with nitrate exposure can contribute to elevated concentrations of
circulating steroid. A hepatic response to nitrate exposure is further supported by findings
reported by de Caceres et al. (2003). Nitrate and nitrite exposure can influence NO formation in

133
the hypothalamus. NO stimulates hypothalamic synthesis of growth hormone-releasing hormone
(GHRH) causing pituitary secretion of growth hormone (GH) into the bloodstream (de Caceres et
al., 2003). Circulating GH stimulates the liver to synthesize and secreted IGF-1 into the
bloodstream. In this manner, nitrate exposure could activate the hypothalamic-pituitary-hepatic
(HPH) axis to influence plasma IGF-1 concentrations (Chapter 2, Fig. 2-7).
In future studies the liver should be examined, in addition to the gonadal steroidogenesis
and plasma hormone concentrations, with different routes of nitrate exposure because studies
reported here indicate that the route of nitrate exposure, as well as the endpoint measured, will
influence the perceived endocrine response.
The increase in plasma IGF-1 concentrations in R. catesbeiana with nitrate exposure
might occur in response to apoptosis within oviductal cells. The stimulatory effect of IGF-1 on
cell growth can counteract apoptosis in reproductive tissues (Pondérate et al., 2000). Increased
plasma IGF-1 concentrations might also occur in response to an initial inhibitory effect of nitrate
on ovarian steroid synthesis. In mammalian ovaries, IGF-1 can also counteract the inhibitory
effects of NO on steroid synthesis by stimulating increased P4, E2, aromatase, and steroidogenic
acute regulatory (StAR) protein synthesis (Devoto et al., 1999; LaVoie et al., 1999; Sekar et al.,
2000; Iniguez et al., 2001; Les Dees et al., 2001). Thus it is possible that plasma IGF-1
concentrations increased with nitrate exposure to counteract decreased gonadal steroid synthesis.
Over the 10-day study, elevated plasma IGF-1 concentrations in response to nitrate might have
been sufficient to restore and even elevate plasma steroid concentrations during this period. Thus,
it is possible that the increase in plasma IGF-1 concentration with nitrate exposure was a
compensatory response to increased gonadal NO synthesis (Chapter 2, Fig. 2-7). However, NO
and steroid synthesis within the gonads of R. catesbeiana was not measured in this study, and this
pathway remains purely speculative.
Nitrate exposure of female R. catesbeiana is associated with atrophy of the oviductal
endometrial layer and gland surface area. Nitrate has been shown to induce apoptosis or cell death

134
in a variety of tissue types including the tail of metamorphosing tadpoles, and the brain, testis,
and gastric mucosa of mammals (Kashiwagi et al., 1999; Peltola et al., 2001; Pant and Srivastava,
2002; Tari et al., 2003). The proposed mechanism for nitrate-induced apoptosis involves, again,
NO activity. NO can target nearly all cell components including proteins, carbohydrates, lipids,
and nucleic acids. NO can induce cell death by inhibiting heme-containing enzymes that are vital
in cell growth including energy metabolism and the synthesis of purines and pyrimidines needed
for nucleic acid production (Lincoln et al., 1995). NO can also induce a cytotoxic response in
tissues by reacting with superoxides to form the free radical peroxynitrite. Peroxynitrite a highly
destructive compound that damages cells and exacerbates inflammatory responses (Bastian et al.,
2002). In R. catesbeiana, the observed atrophy of the endometrium and glands in response to
nitrate exposure might have occurred through NO-inhibition of heme-containing enzymes
involved in DNA synthesis or through a cytotoxic production of peroxynitrite. It must be
mentioned that nitrite, by itself, is highly toxic and might have induced oviductal atrophy in
response to the direct injection of nitrate into the abdominal cavity. However, the effect of nitrite
exposure on amphibian oviductal growth is unknown.
Further studies are necessary to elucidate the pathway by which nitrate interferes with
oviduct morphology and with steroid and IGF-1 hormones in amphibians. It is unknown whether
nitrate-associated oviductal atrophy and endocrine disruption in R. catesbeiana occurs through a
NO-dependent pathway. First, it is necessary to determine whether NOS and NO are found within
amphibian reproductive tissues, and whether they change expression with nitrate exposure. The
use of NO donors and inhibitors might uncover how nitrate alters gonadal steroid synthesis in
amphibians. If steroid synthesis is inhibited in gonadal tissue exposed simultaneously to nitrate
and NO agonists, but steroid synthesis is restored with the addition of NO inhibitors, then nitrate-
associated endocrine disruption in amphibians likely operates through a NO-dependent pathway.
However, if gonadal steroid synthesis with nitrate exposure is unaffected by the addition of NO
donors and inhibitors, a different pathway of nitrate-associated endocrine disruption likely occurs

135
in amphibians. Similarly, if NO expression is increased in atrophied oviductal tissue exposed to
nitrate but is decreased in unexposed tissue, then a NO-pathway for apoptosis exists in amphibian
oviducts.
There is another possible pathway for nitrate-associated endocrine disruption that has
scarcely received attention and should be investigated further. Nitrate exposure might not alter
steroids and IGF-1 solely through direct gonadal effects, but indirectly through effects on the
hypothalamus or the pituitary. The pituitary, brain, and spinal cord contain neuronal NOS (nNOS)
and the central nervous system also contains brain constitutive NOS (bNOS) (McDonald and
Murad, 1995). Nitrate exposure might upregulate NOS enzymes thereby stimulating NO
synthesis within the brain. Subsequently, NO might alter hypothalamic gonadotropin-releasing
hormone (GnRH) and growth hormone releasing hormone (GHRH) secretion and pituitary
gonadotropin (LH and FSH) and growth hormone (GH). The tropic hormones FSH and LH
influence gonadal steroid synthesis, and GH influences hepatic and ovarian IGF-1 synthesis
(Chapter 2, Fig. 2-7). This pathway remains an intriguing topic for further investigation.

ooi x (oa/a) « ooi x (°a/a) 136
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Figure 6-1. Biochemical validation of female Rana catesbeiana plasma for R1A. A. For 1713-
estradiol internal standards and plasma dilution curves were parallel to the estradiol
standard curve (ANCOVA; F = 0.03, P = 0.73; and F = 0.01, P = 0.91). B. For
testosterone internal standards and plasma dilution curves were parallel to the
testosterone standard curve (ANCOVA; F = 0.001, P = 0.79; and F = 0.08, P = 0.79).
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standard curve (ANCOVA; F = 0.001, P = 0.99; and F = 1.86, P = 0.22). B. For
testosterone internal standards and plasma dilution curves were parallel to the
testosterone standard curve (ANCOVA; F = 0.003, P = 0.96; and F = 0.37, P = 0.56).
C. For insulin-like growth factor-1 (IGF-1) the plasma dilution curve was parallel to
the IGF-1 standard curve (ANCOVA; F = 5.14, P = 0.10).

IGF-1 (pg/mL) O Testosterone (pg/mL) * Estradio, (pg/mL)
138
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Nitrate (mg/L)
Figure 6-3. Mean plasma hormone concentrations of female Rana catesbeiana after nitrate (N03-
N) treatments. Data presented as means + SEM. Different letters above bars indicate
significant differences for A. 17(3-estradiol (ANOVA; N = 6, P < 0.001), B.
testosterone (ANOVA; N = 6, P < 0.001), and C. insulin-like growth factor-1 (IGF-
1) (ANOVA; N = 6, P < 0.001).

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139
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Nitrate (mg/L)
Figure 6-4. Mean plasma hormone concentrations of male Rana catesbeiana after nitrate (NO3-N)
treatment. Data presented as means ± SEM. Different letters above bars indicate
significant differences for A. 17P-estradiol (E2) (ANOVA; N = 6, P < 0.001) and C.
insulin-like growth factor-1 (IGF-1) (ANOVA; N = 6, P = 0.03). B. No significant
differences were detected in plasma testosterone (T) among treatment groups
(ANOVA; 7* = 0.41)

140
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NO
Nitrate (mg/L)
Figure 6-5. Oviduct morphology measurements for female Rana catesbeiana after nitrate
treatment. Data presented as means ± SEM. Different letters above bars represent
significant differences among means for A. epithelial cell height (ANOVA; P <
0.001), B. endometrial layer thickness (ANOVA; P < 0.001), and C. gland surface
area (ANOVA; P < 0.001).

CHAPTER 7
CONCLUSIONS
Seasonal Plasma Steroids and IGF-1, and Reproductive Tissue Growth
Plasma steroid and IGF-1 concentrations, in addition to reproductive tissue growth,
exhibited a clear pattern of seasonal changes in adult, female R. grylio. Growth of reproductive
tissues (ovaries and oviducts) was positively correlated with increasing plasma E2 and T
concentrations. The seasonal pattern of changes in reproductive condition in R. grylio varied only
slightly with published reports for other temperate-breeding ranid frogs (Licht et al., 1983; Moore
and Deviche, 1988; Pancharatna and Saidapur, 1992). Licht et al. (1983) reported for R.
catesbeiana that gonadal-somatic index (GSI) and oviductal weights, in addition to plasma
steroid and gonadotropin concentrations, increased between May and July, and declined in
August. In R. grylio from Alachua, county, Florida, seasonal changes in GSI and ovarian
maturation in R. grylio closely match descriptions of seasonal breeding condition and behavior of
R. grylio in Southern Georgia (Lamb, 1984; Wright, 1932), other regions of North-Central
Florida (Wood et al., 1998), and in the Everglades of South Florida (Ligas, 1960). In South
Florida (Lygas, 1960) and in South Georgia (Lamb, 1984), GSI and ovarian maturation in R.
grylio increased between April and July, and regressed between August and March. Collectively,
these findings indicate little variation in seasonal reproductive condition for R. grylio among
these geographic locations.
Very little is known about seasonal changes in circulating IGF-1 concentrations in
amphibians. A seasonal pattern of plasma IGF-1 concentrations has been reported only for Bufo
woodhousei (Pancak-Roessler and Lee, 1990). In B. woodhousei, plasma IGF-1 concentrations
peaked during the reproductive period (May and June) and decreased during the non-reproductive
period (August and December). In R. grylio, plasma IGF-1 concentrations exhibited a similar
141

142
pattern of seasonal change. Additionally, plasma IGF-1 concentrations in R. grylio were highly
correlated to air temperature and fat body weights. As air temperature increased and fat body
weights decreased, plasma IGF-1 concentrations increased. Presumably, fat stores were depleted
due to increased fat metabolism in the liver, which can provide valuable energy to fuel tissue
growth and gamete maturation during the breeding period. In ectotherms, food intake and
metabolic processes typically increase with increasing internal and environmental temperatures
(Lillywhite et al., 1973; Larson, 1992; Rome et al., 1992). Hepatic fat metabolism was not
measured in R. grylio during this study; therefore this remains a speculative but interesting topic
for further research. Nutritional status is known to influence plasma IGF-1 in ectotherms (Crain et
al., 1994; Thissen et al., 1994), suggesting the prediction that plasma IGF-1 concentrations would
increase with declining fat body weights in R. grylio. Overall, these findings suggest that
environmental cues and nutritional status regulate IGF-1 in frogs.
Months of increased plasma IGF-1 overlapped with months of increased ovarian and
oviductal weights, and also with increased plasma E2 and T concentrations. However, no
significant correlation was detected between IGF-1 and steroid concentrations, or between IGF-1
and ovarian and oviductal weights. The association between plasma steroids and IGF-, and the
role of these hormones in regulating growth of reproductive tissues is still unclear. Although
synergy between E2 and IGF-1 stimulates greater oviductal growth than either hormone does
alone in mammals (Murphy and Murphy, 1994) it is unknown whether these hormones stimulate
oviductal growth similarly in R. grylio. During the reproductive periods, synthesis of IGF-1 in R.
grylio might be elevated for incorporation of this growth factor into mature eggs, and for
stimulation of oviductal secretions that function in egg transport and subsequent fertilization
(Low et al., 1976; Guillette et al., 1996; Olsen and Chandler, 1999). Unlike IGF-1, plasma steroid
concentrations showed a strong positive correlation with reproductive tissue weights in R. grylio.
The pattern of increased oviductal growth with increasing plasma steroid concentrations

143
described for R. grylio matches the increase in oviduct growth in R. catesbeiana with E2
treatments reported in Chapter 4.
The Effects of IGF-1, E2, and Nitrate on Oviduct Growth
In Chapter 4, ovariectomized R. catesbeiana were examined for the effects of growth
factors, E2, and combined E2/IGF-1 on various parameters of oviductal growth: oviductal weight,
epithelial cell height, endometrial thickness, and endometrial gland surface-area. In summary, E2-
treated R. catesbeiana exhibited greater oviductal growth compared to IGF-1- and EGF-treated
frogs, indicating that IGF-1 and EGF alone did not stimulate oviductal growth of these
parameters. Additionally, E2/IGF-1-treated frogs did not exhibit greater oviductal growth, for
most parameters, than did frogs given E2 alone, indicating thatE2/IGF-l did not act synergistically
to stimulate greater oviductal growth compared to either treatment alone. The only exception to
this pattern was E2/IGF-1-treated frogs had a greater oviductal epithelial cell height compared to
all other treatments, indicating synergy between the hormones.
The absence of a growth response in the oviductal endometrial layer and glands to either
EGF or IGF-1 alone, and the absence of a synergistic growth response to E2/IGF-1 treatment
distinguished R. catesbeiana from reptilian and mammalian species examined to date using
similar techniques. Some reptiles and mammals exhibit oviductal growth in response to treatment
with IGF-1 or EGF alone (Murphy and Ghahary. 1990; Cox and Guillette, 1994). Additionally,
oviductal growth in these mammals is greater when treated with combined E2/IGF-1 than with
either growth factor alone, indicating hormonal synergy of E2 and IGF-1 (Murphy and Murphy,
1994). Although plasma IGF-1 and EGF have been identified in several amphibian species
(Daughaday et al., 1985; Pancak-Roessler and Lee, 1990), the oviduct of R. catesbeiana
apparently does not proliferate significantly in response to stimulation by these hormones.
Several factors may explain the absence of an oviductal growth response to either growth factor,
or combined E2/IGF-1, in R catesbeiana. First, the IGF-1 doses selected for this study may have
been insufficient to elicit an oviductal growth response in R. catesbeiana. Future studies should

144
investigate several doses of IGF-1, both higher and lower, to determine if any dose is capable of
stimulating oviductal growth in ovariectomized R. catesbeiana. Further, the delivery system may
need to be examined - we used implanted pellets, which have worked in other species, but studies
examining injections could also provide important information. Second, it is possible that the
oviduct must first be "primed" by E2-exposure prior to IGF-1 exposure to become sensitive to the
effects of IGF-1 (Figure 7-1). This priming of oviductal tissue could involve upregulation of
estrogen receptors, such as ERa, and IGF-1 receptors (IGF-1 R). This hypothesis is supported by
studies showing that ERa must be present for IGF-1 to induce effects (Klotz et al., 2000).
Additionally, Clark et al. (1997) demonstrated that E2 stimulates a proliferative response in
reproductive tissues in vitro by upregulating IGF-1R expression, which increases the tissue’s
response to circulating IGF-1. Based on these findings, it is possible that an increase in
circulatory E2 increases receptor-dependent tissue sensitivity to the growth-promoting effects of
IGF-1 without necessarily requiring an increase in circulatory IGF-1 (Figure 7-1). This seems to
be supported by the increased oviductal epithelial cell height with E2/IGF-1-treated R.
catesbeiana.
Another possible explanation is that oviductal sensitivity to these growth factors
represents a relatively recent evolutionary change in reptilian and mammalian reproductive
physiology. However, more amphibian species should be examined, using similar techniques,
before this conclusion can be verified. Findings from this study might be exclusive to R.
catesbeiana, and interspecific differences in hormonal-induced oviductal growth might be
prevalent among amphibians.
It is interesting to note that oviductal growth in sham frogs was not similar to growth in
E2- and E2/IGF-1 treated frogs. It was assumed that the sham frogs would exhibit oviductal
growth, similar to E2-treated frogs but greater than that of placebo frogs, because their intact

145
ovaries would continue to synthesize and secrete E2 throughout the study. There are several
possible explanations for these findings.
It is possible that the implants in E2- and E2/IGF-treated frogs contained E2
concentrations higher than is typically found in R. catesbeiana. E2 doses were determined based
on studies of E2 necessary to elicit oviductal growth in Xenopus laevis (Follett and Redshaw,
1967; Redshaw et al., 1968) and in reptiles (Cox, 1994). Thus, these doses might have been
comparatively high for R. catesbeiana. Unfortunately, prior to this study, there were no published
references for physiologically relevant doses of E2 capable of stimulating oviductal growth in
ovariectomized R. catesbeiana. Regardless, this explanation still seems less probable than the
ones that follow.
Another explanation for relatively low oviductal growth in sham frogs was that they were
different from E2- and E2/IGF-1-treated frogs with respect to pre-surgery and post-treatment
plasma IGF-1 concentrations. In E2- and E2/IGF-1-treated frogs, plasma IGF-1 concentrations
increased after treatment compared to pre-ovariectomy levels. In sham frogs, however, plasma
IGF-1 concentrations after sham-treatment remained similar to pre-surgery concentrations. In E2-
and E2/IGF-1 -treated frogs, the increase in plasma IGF-1 could have stimulated increased IGF-1R
and expression in oviductal tissues, making them more sensitive to E2- and IGF-1-induced
growth. As mentioned above, IGF-1 R does interact, or exhibit "cross-talk" with the ERa in
stimulating oviduct growth (Klotz et al., 2002). There was no change in mean plasma IGF-1
concentrations in sham frogs before and after the experiment. With no change in IGF-1
concentrations, it is possible that oviductal IGF-1R expression also remained unchanged, or

Circulation
146
Hypothalamus
NO
Nitrate
G.
Figure 7-1. Regulation of gonadal steroid synthesis and metabolism, and of hepatic IGF-1
synthesis. A Hypothalamic gonadotropin releasing hormone (GnRH) induces
pituitary luteinizing hormone (LH) and follicle stimulating hormone (FSH) secretion.
B. LH stimulates testosterone (T) synthesis from progesterone (P4) precursor. FSH
stimulates aromatase enzymes (arom) to convert T into estrogen (E2). C. Steroids (E2
& T) enter circulation and can inhibit GHRH and GnRH release, and D. be
metabolized and removed from circulation by liver P450 enzymes. E. Hypothalamic
growth hormone -releasing hormone (GHRH) stimulates pituitary growth hormone
(GH) secretion. GH stimulates liver insulin-like growth factor-1 (IGF-1) synthesis
and secretion. F. Nitrate metabolized by the liver can cause NO formation, which can
inhibit hepatic P450 steroid metabolizing enzymes, preventing steroid metabolism
and clearance from circulation. G. Nitrate and nitrite stimulation formation of
hypothalamic NO GHRH release, and pituitary GH, which stimulates the liver to
secrete IGF-1 into circulation, stimulating GHRH H. Nitrate and nitrite might inhibit
oviduct growth through conversion to nitric oxide (NO), which can induce apoptosis
in tissue.

147
baseline, reducing ERa "cross-talk", and making oviductal tissue comparatively less sensitive to
E2- or IGF-1-induced stimulation of growth (Figure 7-1). Future studies should examine oviduct
growth in response to steroid and growth factors hormones and also in response to ERa and IGF-
1R expression to better understand their possible interactions.
Lastly, there might have also been an "implant effect" on oviductal growth. With or
without hormones, the implant might have elicited oviductal hypertrophy due to an irritation
response to a "foreign body" within the abdomen. In summary, much more research is necessary
to discern the potential influence of these variables on oviduct growth in R. catesbeiana.
It is interesting to report that nitrate exposure of R. catesbeiana was associated with
decreased oviductal endometrial layer thickness and gland surface-area. It is possible nitrate
exposure increased endogenous nitrite levels and nitric oxide (NO) formation in the oviduct. NO
is known to induce apoptosis, or programmed cell death. NO can induce apoptosis through
inhibition of heme-containing enzymes involved in cell growth (Lincoln et al., 1995). An increase
in oviductal NO could also lead to formation of peroxynitrite, a cytotoxic compound that causes
cellular damage and inflammation (Bastían et ah, 2002).
Nitrate Exposure (In Vivo and In Vitro): Effects on Steroidogenesis
Nitrate exposure has been reported to inhibit in vivo and in vitro steroidogenesis in
mammals (Panesar and Chan, 2000). Nitrate within the body can be readily converted to nitrite,
and visa versa, by endogenous microbial activity and oxidation (Jensen, 1995; Doblander and
Lackner, 1996, 1997; Jensen, 2003). Within the liver and other tissues, nitrite is then metabolized
into nitric oxide (NO) (Doblander and Lackner, 1996). NO is known to inhibit steroidogenesis by
binding to the heme groups of cytochrome P450 (CYP) enzymes (Figure 7-1). The steroid
synthetic enzymes include side-chain cleavage (CYP11), 17(3-hydroxylase (CYP 17), and
aromatase (CYP 19). Side-chain cleavage is the major rate-limiting step in the steroidogenic
pathway (Jensen, 2003). Studies in fish and turtles indicate that similar steroidogenic enzymes

148
occur in mammalian and non-mammalian vertebrates (Sakai et al., 1992; Takahashi et al., 1993;
Jeyasuria et ah, 1994; Omura, 1999).
In this dissertation, different responses of frogs were observed with exposure to in vitro
and in vivo nitrate. Xenopus laevis exposed to in vivo nitrate for seven days exhibited no
difference in plasma steroid concentrations among treatments while ex vivo ovarian steroid
concentrations, synthesized by isolated tissue, were significantly decreased in nitrate-exposed
frogs. Similar to ex vivo steroidogenesis observed inX. laevis, in vitro steroidogenesis in R. grylio
ovaries exhibited decreased steroid synthesis when exposed to nitrate (Chapter 5).
The basis for the normal in vivo steroid concentrations but decreased ovarian ex vivo
steroid concentrations in X. laevis remains unclear but likely involves endocrine feedback of the
hypothalamic-pituitary-gonadal (HPG) axis (Figure 7-1). Despite the inhibitory effects of nitrate,
normal in vivo steroid concentrations were likely maintained by compensatory signaling through
the HPG axis. If nitrate exposure inhibited ovarian steroid synthesis in R. catesbeiana, a decrease
in circulating steroids or inhibin could have stimulated the hypothalamus to secrete gonadotropin¬
releasing hormone (GnRH) and the pituitary to secrete plasma gonadotropin (FSH and LH). FSH
and LH would stimulate continued gonadal steroidogenesis to compensate for the initial
inhibition, which could have contributed to normal plasma steroid concentrations. Compensatory
in vivo responses are reported for human patients that develop nitrate tolerance with continuous
treatment of organic nitrate like nitroglycerin or isosorbide dinitrate (Gori and Parker, 2002;
Parker, 2004). In contrast to the in vivo model, ex vivo and in vitro steroidogenesis involved
culture of ovarian tissue that was physically isolated from hypothalamic-pituitary regulation. In
isolated ovarian tissue, nitrate-induced inhibition of steroidogenesis could not be counteracted by
compensatory release of hypothalamic GnRH and pituitary gonadotropins (Figure 7-1). Plasma
gonadotropins and hypothalamic GnRH were not measured in the in vivo study ofX. laevis; thus,
this theory cannot be confirmed.

149
Another possible mechanism for the increased circulating steroid concentrations in
response to nitrate could involve regulation at the level of the hypothalamic-pituitary-hepatic
(HPH) axis (Figure 7-1). In mammals, plasma steroid concentrations have also been reported to
increase with nitrate exposure due to inhibition of hepatic-steroid degradation (Waxman, 1992;
Sivapathasundaram et al., 2003). The liver is the main site for nitrate catabolism where large
amounts of NO are produced (Lalka et al., 1993). Hepatic NO can inhibit P450 enzymes involved
in steroid metabolism and elimination (Waxman, 1992). Minamiyama et al. (2004) reported for
rats that administration of organic nitrate decreased hepatic cytochrome P450 activity within 24
hr of exposure. Other xenochemicals also alter the expression of hepatic P450 enzymes involved
in steroid metabolism and elimination (Waxman, 1992). Minamiyama et al. (2004) reported for
rats that administration of organic nitrate decreased hepatic cytochrome P450 activity within 24
hr of exposure. Other xenochemicals also alter the expression of hepatic P450 enzymes
(Waxman, 1999). Therefore, despite a depression of steroid synthesis in the gonad, as seen in X.
laevis and R. grylio, a severe nitrate-induced depression of P450 activity in the liver could have
depressed steroid clearance in R. catesbeiana and plasma steroid concentrations were actually
augmented (Figure 7-1).
Ovarian ex vivo steroid inhibition in X. laevis was similar to ovarian in vitro steroid
inhibition in R. grylio in that a dose-dependent steroid response curve was not observed with
nitrate exposure. The absence of a dose-dependent decrease in steroidogenesis with increasing
nitrate exposure in these two studies offers several explanations. For one, the difference between
the lowest and highest nitrate concentration might have been insufficient to produce a differential
response in steroid synthesis. For ex vivo culture of X. laevis ovaries, frogs were exposed to
nitrate concentrations of 24.75 and 49.50 mg/L NO3-N, whereas, in vitro culture of R. grylio
ovaries involved nitrate exposure ranging in concentration from 0.17 - 33.00 mg/L NO3-N. Thus,
the lowest and highest nitrate concentrations appeared equally capable of inhibiting steroid
synthesis. For R. grylio, it is also possible that the steroid-inhibition threshold is actually much

150
lower than the lowest nitrate (0.17 mg/L) and nitrite (0.20 mg/L) concentration examined. Future
studies should include a wider range of nitrate and nitrite concentrations; for example, ranging
from 25 pg/L and 1000 mg/L. Reports of amphibians exposed to atrazine have shown that
extremely low concentrations (25 pg/L) are capable of inducing a 10-fold decrease in steroid
synthesis (Hayes et al., 2002). Similar to the sensitivity of amphibian gonads to disruption by
atrazine, ovarian steroidogenesis is apparently sensitive to endocrine disruption by extremely low
concentrations of nitrate.
Nitrate Exposure and Plasma IGF-1
The investigations in this dissertation provide the first evidence that nitrate exposure of
amphibians to environmentally relevant nitrate concentrations, ranging from 1.65 - 49.50 mg/L, is
associated with increased plasma IGF-1 concentrations inX. laevis and R. catesbeiana. The
physiological mechanism regulating nitrate-associated increases in circulating IGF-1 is unclear
but a hypothesis has been proposed (Chapter 2).
There is increasing evidence that IGF-1 functions as a regulator of steroid function and
secretion (Hammond et al., 1991; Adashi, 1993; Devoto et al., 1999; Driggers and Segars, 2002).
Increased circulating IGF-1 concentrations might counteract nitrate-induced steroid inhibition
(Figure 7-1). Intraovarian NO formation, in response to nitrate exposure, might increase
expression of IGF-1. Increased intraovarian IGF-1 has been shown to increase expression of P450
aromatase enzyme (CYP19) and E2, and also regulates estrogen-induced growth of reproductive
tissues (Daughaday and Rotwein, 1989; Erickson et al., 1989; Monnieaux and Pisselet, 1992;
Adashi, 1993; Samaras et al., 1994; Hiney et al., 1996; Olsen et al., 1996; Samaras et al., 1996;
Dees et al., 1998). NO also might influence IGF-1 at the level of the hypothalamic-pituitary-
hepatic axis. NO participates in the regulation of GH secretion at level of the HPH axis, thereby
increasing hepatic IGF-1 synthesis and circulating IGF-1 concentrations (de Caceres et al., 2003;
Figure 7-1). NO not only decreases ovarian P450 aromatase enzyme and E2 expression but also

151
increases IGF-1 mRNA (Srivastava et al., 1997; Dees et al., 2000). Thus, the increase in
circulating IGF-1 concentrations might derive from non-hepatic sources, including the ovaries.
In summary, increasing evidence suggests that steroids should not be the exclusive
endpoint for evaluating endocrine-disruption by nitrate. These endpoints should be expanded
include steroid receptors, steroidogenic enzymes, growth factors and their receptors, steroid and
IGF-1 binding proteins, and hepatic cytochrome P450 enzymes. Understanding the role of nitrate,
steroids, and IGF-1 in normal and abnormal reproduction in amphibians and other animals will be
largely dependent upon which endpoints are examined.

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BIOGRAPHICAL SKETCH
Tamatha R. Barbeau was bom in Plattsburgh, New York, on 28 September 1970. Many
of her summers were spent camping with her family in the NY Adirondacks and in Canada. There
she spent much of her time fishing, studying animals, and exploring the northern woodlands. At
Canton State University of NY, she studied veterinary science and earned her associates degree
and a veterinary technician license in 1990. At Oswego State University of NY, she majored in
biology and earned a baccalaureate in 1992. After completing her undergraduate studies, she
worked as the head technician in a veterinary hospital for 5 years. A growing desire to work with
a wide diversity of animals in their natural environments inspired her to change careers and enter
graduate school. She joined the Department of Zoology at the University of Florida in 1997 and,
working with Dr. Harvey B. Lillywhite, was awarded a Master of Science degree in 2000.
169

I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation
for the degree of Doctor of Philosophy.
Distinguished Professor of Zoology
I certify that I have read this study and that in my o]
standards of scholarly presentation and is fully adequate, f
for the degree of Doctor of Philosophy.
n it conforms to acceptable
cope ancTquaifiy^as a dissertation
William C. Buhi
Professor of Biochemistry and Molecular Biology
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation
for the degree of Doctor of Philosophy.
Professor of Zoology
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation
for the degree of Doctor of Philosophy.
David H. Evans
Professor of Zoology
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation
for the degree of Doctor of Philosophy.
Hafvqy/B. Liltywhite
Professor of Zoology
This dissertation was submitted to the Graduate Faculty of the Department of Zoology in
the College of Liberal Arts and Sciences and to the Graduate School and was accepted as partial
fulfillment of the requirements for the degree of Doctor of Philosophy.
August 2004
Dean, Graduate School