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

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Title:
Influence of insulin-like growth factor-1, steroids, and nitrate on reproduction in amphibians
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ix, 169 leaves : ill. ; 29 cm.
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Barbeau, Tamatha R., 1970
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Frogs -- Reproduction -- Endocrine aspects   ( lcsh )
Zoology thesis, Ph. D   ( lcsh )
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Thesis (Ph. D. )--University of Florida, 2004.
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Includes bibliographical references.
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Printout.
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Vita.
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by Tamatha R. Barbeau.

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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 -

0
60
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