Iowa State University Capstones, Theses and Retrospective Theses and Dissertations Dissertations

2007 The peroxisome proliferator-activated receptor in ovarian biology Nicole Shivonn Tinfo Iowa State University

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This Dissertation is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Retrospective Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. The peroxisome proliferator -activated receptor in ovarian biology

by

Nicole Shivonn Tinfo

A thesis submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Animal Physiology (R eproductive Physiology)

Program of Study Committee: Carolyn Komar, Major Professor Richard Robson Steven Hopkins Leo Timms Jill Pruetz

Iowa State University

Ames, Iowa

2007 UMI Number: 3259475

UMI Microform 3259475 Copyright 2007 by ProQuest Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.

ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, MI 48106-1346 ii

In dedication to my parents, Stephanie and Eugene Tinfo, and my fiancé, Bryan Clark. Your strength, and inspiration have provided me with the encouragement to succeed. Thank you.

iii

TABLE OF CONTENTS ABSTRACT v CHAPTER 1. GENERAL INTRODUCTION 1 INTRODUCTION 1 DISSERTATION ORGANIZATION 2 CHAPTER 2. LITERATURE REVIEW 3 THE OVARY 3 THE RODENT (RAT AND MOUSE) ESTROUS CYCL E 8 THE BOVINE ESTROUS CYCLE 9 PEROXISOME PROLIFERATOR ACTIVATED RECEPTORS 12 REFERENCES 17 CHAPTER 3. POTENTIAL ROLE FOR PEROXISOME PROLIFERATOR -ACTIVATED RECEPTOR γ IN REGULATING LUTEAL LIFESPAN IN THE RAT 30 ABSTRACT 30 INTRODUCTION 31 METHODS AND MATERIALS 33 RESULTS 38 DISCUSSION 40 ACKNOWLEDGEMENTS 44 REFERENCES 44 CHAPTER 4. DEVELOPMENT OF MODELS TO FACILITATE INVESTIGATION OF OVARIAN GENES REGULATED BY THE TRANSCRIPTION FACTOR, PEROXISOME PROLIFERATOR - ACTIVATE RECEPTOR γ 58 ABSTRACT 58 INTRODUCTION 58 METHODS AND MATERIALS 60 RESULTS 66 DISCUSSION 68 REFERENCES 71 CHAPTER 5. EFFECT OF GENISTEIN ON ESTRADIOL PRODUCTION BY BOVINE FOLLICLES 81 ABSTRACT 81 INTRODUCTION 81 METHODS AND MATERIALS 83 RESULTS 86 DISCUSSION 87 ACKNOWLEDGEMENTS 89 REFERENCES 89 iv CHAPTER 6. GENERAL CONCLUSIONS 98 DISCUSSION AND CONCLUSIONS 98 REFERENCES 101 ACKNOWLEDGEMENTS 103 v

ABSTRACT

Understanding how the ovary functions in a normal, cyclic pattern is necessary in order to identify what leads to female sub - and infertility. In order to improve fertility and reproductive success, it is important to understand how genes are regulated in the ovary.

Peroxisome proliferat or-activated receptor γ (PPAR γ) has been detected in the ovary of several species. Previous studies have found that PPAR is involved in steroidogenesis and can influence female fertility . However, the genes it regulates in the ovary are not known.

The studies desc ribed herein were designed to investigate the role of PPAR γ in ovarian function. The effect of PPAR γ on luteal steroid production and luteal cell survival was investigated. These experiments demonstrated that PPAR γ did not play a role in the production a nd/or breakdown of progesterone, the main hormone produced by the corpus luteum. It was also found that the localization of PPAR γ in the corpus luteum was not due to the presence of macrophages. However, it was determined that PPAR may play a role in lu teal cell survival by regulating an important gene involved in cell survival. In vitro and in vivo models were created in which the expression of PPAR γ was selectively reduced in the ovary. The in vivo model allowed for identification of candidate genes regulated by PPAR .

Finally, the impact of an environmental agonist of PPAR γ on follicular estradiol production was also studied. Genistein has been found to bind to and activate PPAR γ. Experiments were performed that suggest that genistein may not affe ct the basal production of estradiol but may alter response to gonadotropin stimulation. Collectively, these studies were able to identify candidate ovarian genes regulated by PPAR and possible role(s) of PPAR γ in the ovary. The knowledge of the genes re gulated by PPAR will allow researchers to further investigate how regulation of these genes by PPAR is involved in the functioning of the ovary. 1

CHAPTER 1 . GENERAL INTRODUCTION

INTRODUCTION Understanding how the ovary functions in a normal, cyclic patte rn is necessary to identify what goes awry in cases of female sub - and infertility. There are many genes involved in regulating the ovarian cycle. One family of transcription factors, the peroxisome proliferator -activated receptors (PPARs), has been loca ted in the ovary of several species. Very little is known regarding what genes the PPARs target or how they function in ovarian tissue.

There are three isotypes of PPARs: α, β, and γ. The expression of PPAR α and β remains steady throughout follicular development. PPAR γ is the only isotype that is differentially expressed in granulosa cells during the ovarian cycle. The expression of PPA R is down -regulated in response to the lutenizing hormone ( LH ) surge, which triggers

ovulation [1, 2] . PPAR γ is expressed primarily in granulosa cells of developing follicles but is also located in luteal tissue of the naturally cycling rat [2, 3] and bovine [4, 5] . It has been

reported that female mice with disrupted expression of PPAR γ were either infertile or subfertile [6] . These findings indicate that PPAR γ may play a critical role in ovarian function and female fertility.

The studies described herein were designed to investigate the role of PPAR γ in ovarian function. The effect of PPAR γ on luteal steroid production was investigated, as well as a possible role for PPAR γ in luteal cell survival. In vitro and in vivo models were created in which the expression of PPAR γ was selectively reduced in granulosa cells to facilitate

investigating ovarian genes that are regulated b y PPAR γ. Finally, the impact of an environmental agonist of PPAR γ, genistein, on follicular estradiol production was studied. Collectively, these studies were able to identify candidate ovarian genes regulated by PPAR

and possible role(s) of PPAR γ in th e ovary. 2

DISSERTATION ORGANIZ ATION This dissertation, written by the author, is one of the requirements for a doctoral degree in Animal Physiology. The dissertation is presented in six chapters. Chapter 2 provides a literature review of ovarian functio n, and the structure and function of PPAR γ. The following chapters describe the experiments performed to investigate the function of

PPAR γ in the ovary. Chapter 3 specifically discusses the role of PPAR γ in luteal tissue of the naturally cycling rat. The process of developing models of red uced expression of PPAR γ in the ovary is presented in chapter 4. Chapter 5 details a project that investigated the effects of genistein, a phytoestrogen and PPAR γ agonist, on estradiol production by bovine follicles. The dissertation concludes with a gen eral discussion of the results from these experiments and the possible roles of PPAR γ in the ovary. 3

CHAPTER 2. LITERATURE REVIEW The following literature review focuses on an overview of the ovary and ovarian function. This is followed by discussion of the rat/mouse and bovine ovarian cycles because these two animal models were utilized in the experiments described in this dissertation.

Following the literature review on the ovary and ovarian cycles, PPAR γ is discussed because it was the focus of the e xperiments described in this dissertation due to its potential to impact ovarian biology. THE OVARY The ovary is comprised of three main regions, the hilus, cortex, and medulla (Figure 2.1 A, B, C). Blood and lymphatic vessels, as well as nerves, enter th e ovary at the hilus (Figure 2.1 A). The majority of those vessels and nerves are housed inside the ovary in the medulla (Figure 2.1 B). Follicles at different stages of development are located in the cortex of the ovary (Figure 2.1 C). All follicles co ntain an ooc yte, and depending on the stage of follicular development, have one or two types of cells capable o f producing steroid hormones - granulosa (Figure 2.1 D) and theca cells (Figure 2.1 E). After ovulation, these two steroidogenic cell types are transformed into luteal cells through a process referred to as luteinization (Figure 2.1 F). The Ovarian Cycle The ovarian, or estrous cycle , is defined as the interval between periods of sexual receptivity in the non -pregnant female. The ovarian cycle occurs in a cyclic pattern in the sexually mature female and is broken up into two phases: the follicular phase and the luteal phase. The follicular phase is characterized by the final stages of follicular growth, and ovulation – release of the oocyte. T he main hormone associated with the follicular phase, estradiol, is produced by the developing follicle(s). After ovulation, the luteinization of follicular cells to form a corpus luteum initiates the luteal phase. The corpus luteum produces progesterone , a hormone needed for the establishment and maintenance of 4

pregnancy. The length of the ovarian cycle varies among species. On average, the ovarian cycle is 4 or 5 days in rats/mice and 21 days in cattle. Follicular and Luteal Development Follicular d evelopment occurs in stages. The most immature follicle is a primordial follicle. A mature follicle is called a Graafian, or preovulatory, follicle. During fetal life, follicular development begins when primordial germ cells develop into oocytes (review ed by [7] ). Primordial follicles consist of an oocyte surrounded by a single layer of flattened epithelial cells (Figure 2.2 A). In most mammals, primordial follicles form either before, or in the first few days after birth. The next step in folliculogenesis is the development of primary follicles. In primary follicles, the oocyte is g rowing and the single layer of flattened epithelial cells surrounding the oocyte develops into cuboidal granulosa cells (Figure 2.2 B). The granulosa cells proliferate and the development of a second layer of granulosa cells results in a secondary follicl e (Figure 2.2 C; reviewed by [8] ). The oocyte in a secondary follicle continue s to grow and produces the zona pellucida. The zona pellucida is a glycoprotein -rich substance that forms a protective coat around the oocyte (reviewed by [8] ). Granulosa cells located next to the zona pellucida extend protoplasmatic processes into the oocyte. These protoplasmatic processes provide nutrition for the oocyte, which influences oocyte development (reviewed by [9] ). During the secondary stage of follicular development, the stromal cells that surround the follicle diff erentiate into theca cells. A basement membrane, referred to as the basal lamina, separates the theca and granulosa cells. The basal lamina prevents blood vessels from passing into the granulosa cell compartment (reviewed by [8] ). As the follicles continue to grow, an antrum, or fluid filled cavity develops. Follicles are referre d to as antral follicles once this antrum forms. The antrum contains fluid produced by the granulosa cells (reviewed by [8] ). As the antrum develops, the oocyte becomes displaced from the center of the follicle and the proliferation of granulosa cells decreases. 5

A mound, or hillock of granulosa cells, connects the oocyte to the f ollicle wall; these granulosa cells are referred to as cumulus cells. The oocyte and cumulus cells together comprise the cumulus oocyte complex. As the production of antral fluid increases, the follicle increases in size and is referred to as a Graafian follicle (Figure 2.2 D). The oocyte in a Graafian follicle has reached its maximum size. At ovulation, the cumulus oocyte complex is released from the follicle (reviewed by [8] ). The follicular phase of the estrous cycle encompasses the final stages of antral follicular growth, and release of the oocyte at ovulation. Not all folli cles will develop into Graffian follicles. Most follicles will go through atresia. Atresia occurs via apoptosis, a highly organized form of cell death, and can occur at any time during follicular development (reviewed by [10] ). After ovulation, the granulosa and the ca cells remaining in the ovary will involute and form a corpus luteum (reviewed by [11] ). The corpus luteum consists of large and small luteal cells that are derived from the granulosa and theca cells, respectively. These luteal cells produce progesterone, which is required for the establishment and maintenance of pregnancy. Hormones of the Ovarian Cycle The development of follicles and corpora lutea is controlled by communication between the hypothalamus, pituitary, and ovary. This r elationship between the hypothalamus, pituitary, and ovary is essential for estradiol production during the follicular phase, ovulation, and subsequent production of progesterone during the luteal phase. Hormones produced by the hypothalamus influence t he production and release of hormones from the anterior pituitary, which in turn regulate ovarian hormone production. One hormone secreted by the hypothalamus, gonadotropin releasing hormone (GnRH), stimulates the release of the gonadotropins, follicle st imulating hormone (FSH) and luteinizing hormone (LH), from the anterior pituitary gland (reviewed by [12] ). FSH 6

stimulates follicles to grow and produce estradiol. LH plays a role in the final stages of antral follicular development and induces ovulation and formation of the corpus luteum (reviewed by [12] ). Cooperation between theca and granulosa cells is necessary for the production of estradiol by the developing follicles . Theca cells contain LH receptors, while gra nulosa cells contain FSH receptors. As antral follicles develop, granulosa cells also develop LH receptors. LH released from the anterior pituitary binds to its receptor on theca cells, which results in the conversion of cholesterol into progesterone thr ough the actions of the enzymes

P450 side -chain cleavage and 3 β-hydroxysteroid dehydrogenase (Figure 2.3). Theca cells also have the enzyme, 17 α-hydroxylase, which converts progesterone into androgens (Figure 2.3; reviewed by [13] ). The androgens produced by the theca cells diffuse across the basal lamina into granulosa cells. Granulosa cells express the enzyme P450 aromatase, which converts androgens into estradiol (Figure 2.3; reviewed by [13] ). Estradiol from growing follicles stimulates the hypothalam us to release more GnRH, which results in an increase in LH and FSH release. The stimulation of GnRH by estradiol occurs in a positive feedback manner . However, when estradiol levels are low estradiol exerts negative feedback on GnRH release (discussed b elow). As the follicle grows, production of estradiol increases. Once estradiol reaches a threshold level, a large quantity of LH and FSH is released in

response to GnRH. The large quantity of LH and FSH is referred to as the LH surge, due to the fact t hat more LH is released than FSH (reviewed by [12] ). The LH surge is followed by ovulation in most species. After ovulation, the granulosa and theca cells remaining in the ovary become luteal cells. This transition is termed luteinizati on (reviewed by [11] ). The granulosa cells become large luteal cells, while the theca cells become small luteal cells. During luteinization, there is a decrease in estrogen production and an increase in progesterone production (reviewed by [11] ). Both the large and small luteal cells produce progesterone. Large luteal cells also 7

produce oxytocin. Oxytocin is also released from the posterior pituitary and b oth pituitary and luteal oxytocin are involved in luteal regre ssion (discussed below) . The main role of the corpus luteum is to produce progesterone. The primary target organs for progesterone are the hypothalamus, uterus, and mammary gland. Progesterone causes a decrease in hypothalamic release of GnRH. This re sults in lower LH and FSH release. The lower production of LH and FSH stimulates low levels of estradiol production. Low levels of estradiol cause a decrease in GnRH release (negative feedback). Due to the low levels of GnRH, the levels of LH and estrad iol needed for ovulation can not occur , and therefore, ovulation can not occur. In order for ovulation to occur, progesterone level s must be low so as not to interfere with the release of GnRH from the hypothalamus. If a female is not successfully mated then the corpus luteum will regress, reducing levels of circulating progesterone. There are two phases to luteal regression. The first is functional luteolysis, which involves a reduction in progesterone production. Structural luteolysis, regression of the luteal tissue, follows functional luteolysis (reviewed by [11] ). Luteolysis is believed to occur in response to down -regulation of the progesterone receptor. The constant high level of progesterone produced by the corpus luteum du ring the luteal phase, desensitizes its own receptor. This down -regulation of the progesterone

receptors allows for increased release of GnRH. The increase in GnRH results in the release of more LH and FSH, and stimulation of follicular cells to produce estradiol [14] . Estradiol stimulates the hypothalamus and pituitary, an d increases GnRH release. This results in greater estradiol production from growing follicles. Associated with declining progesterone production during functional luteolysis is a decrease in blood flow to the corpus luteum, depriving luteal tissue of nut rients and substrates for steroidogenesis. Macrophages and lymphocytes in the corpus luteum increase during luteolysis and aid in phagocytosis of the luteal cells. 8

THE RODENT (RAT AND MOUSE) ESTROUS CYCLE Rats/mice are litter -bearing animals. They ovul ate multiple follicles and are referred to as polyovular species. The rat/mouse estrous cycle is 4 or 5 days. There are four phases of the rat/mouse estrous cycle, each one lasting approximately 24 hours: proestrus, estrus, metestrus, and diestrus. The follicular phase of the rodent estrous cycle occurs during proestrus and estrus. Metestrus and diestrus make up the luteal phase. Prolactin is released from the anterior pituitary and is an important hormone involved in luteal formation and regression i n the rat. It is secreted in low, basal amounts for the majority of the cycle. Follicular growth and peak estradiol production occur during proestrous (Figure 2.4 C). Peak estradiol production occurs on the afternoon of proestrus, which causes an increa se in prolactin release (Figure 2.4 B; reviewed by Freeman 1994). The LH surge occurs on proestrus, in the evening (Figure 2.4 D). Estrus is the time of sexual receptivity, mating, and ovulation. LH, FSH, and estradiol levels decrease following the LH surge and remain relatively low through the day of estrus (Figure 2.4 D, E, and C). The beginning of corpora lutea formation and progesterone production takes place during metestrus (Figure 2.4 A). By diestrus, luteal progesterone levels have decreased an d the

inactive metabolite of progesterone, 20 α-hydroxyprogesterone, is secreted (reviewed by [15] ). The enzyme, 20 α-hydroxysteroid dehydrogenase, converts progesterone into 20 α- hydroxyprogesterone (reviewed by [15] ), and is located in luteal tissue. Due to the fact that rats/mice ovulate multiple oocytes, they also form several corpora lutea. T hese corpora lutea remain in the ovary for 2 -3 cycles. Therefore, the cycling rat/mouse ovary contains new corpora lutea formed from the most recent ovulation, and old corpora lutea from the past two or three cycles. The newly formed corpora lutea are no t considered fully functional due to the fact that progesterone is produced for only one to two 9

days. In order for a rat/mouse to have fully functional corpora lutea, cervical stimulation by a male or artificial means must take place. Cervical stimula tion results in two daily surges of prolactin, which extends luteal production of progesterone. If the mating results in pregnancy, corpora lutea will be maintained for 20 -22 days. If it is an infertile mating, the rat/mouse is said to be pseudopregnant, and the corpora lutea will be maintained and produce progesterone for 12 -14 days (reviewed by [16] ). Prolactin not only plays a role in the development of functional corpora lutea, but it is also involved in luteolysis. An important process in luteal regression is the remova l of old luteal cells by macrophages through the process of phagocytosis. Prolactin has been associated with increased movement of monocytes and macrophages into the regressing corpus luteum [17 -19] . The surge of prolactin during proestrus initiates luteal regression by stimulating the migration of macrophages into the corpus luteum. Multiple cycles are needed for complete luteal demise, because in a non -pregnant rat prolactin is only released once during the cycle [17] . THE BOVINE ESTROUS C YCLE The bovine estrous cycle differs from that of the rodent in several ways. First, the bovine is a monovular species, typically ovulating one oocyte per cycle. Second , the bovine has a functional luteal phase in which the corpus luteum is formed, produces progesterone, and regresses if the female is not successfully mated. The third difference is that prolactin does not appear to be involved in luteal formation or fun ction. The final difference is that during the luteal phase, two or three waves of follicular development and follicular atresia occur. The maintenance and/or regression of a single follicle depend on the hormonal environment and will be discussed below. Only after progesterone levels have decreased due to luteolysis will a follicle ovulate (reviewed by [11] ). 10

The estrous cycle in cattle is also divided into four phases, proestrus, estrus, metestrus and diestrus. Proestrus is th e period prior to estrus; estrus is the period of sexual receptivity and these two phases make up the follicular phase. The periods of metestrus and diestrus correspond to the luteal phase. There are five stages of follicular growth during each follicul ar wave: emergence, recruitment, selection, dominance, and atresia (Figure 2.5). A follicular wave begins with the emergence of a cohort of small, antral follicles over one to several days. Recruitment refers to the time that the continued growth and dev elopment of the cohort of follicles requires FSH (Figure 2.5 R). At the time of recruitment, antral follicles grow from 5 mm to 8 or 9 mm. The granulosa cells in these antral follicles express mRNA for the FSH receptor, P450 side chain cleavage, 3 β-hydro xysteroid dehydrogenase, and P450 aromatase (reviewed by [20] ). Theca cells express mRNA for the LH receptor, P450 side chain cleavage, 17 α-hydroxylase, and 3 β-hydroxysteroid dehydrogenase (reviewed by [21] ). Therefore, those follicles recruited for further growth are producing estradiol. Those follicles not recr uited for continued growth go through atresia. During follicle selection, one or two follicles from the cohort of recruited growing follicles is/are chosen for further development, while the rest of the follicles go through atresia (Figure 2.5 S; reviewed by [22] ). At the time of selection, the follicles start producing the glycoprotein hormone, inhibin. Inhibin suppresses FSH synthesis and release from the anterior pituitary (reviewed by [23] ). The granulosa cells in the selected follicles also express mRNA for the LH receptor and greater levels of mRNA for 3 β-hydroxysteroid dehydrogenase (reviewed by [21] ). The expression of LH receptors in the granulosa cells and an increase in 3 β-hydroxysteroid dehydrogenase allow for increased estradiol production. Estradiol enhances aromatase activity, promotes expression of the LH receptor, and increases sensitivity of granulosa cells to LH and FSH [13] . The beginning of the 11

difference in size between selected follicles is referred to as deviation [24] . The first follicle

to acquire LH receptors in its granulosa cells and an increase in 3β-hydroxysteroid dehydrogenase is the follicle that will become dominant, the morphologically larger follicle that continue s to grow and can be ovulated under appropriate hormonal conditions . The remaining follicle(s), subordinate follicle(s), grows at a slower rate and then regresses (reviewed by [25, 26] ). Early in the follicular wave, th e future dominant follicle may be slightly larger than others in the cohort, since emergence occurs over one to several days. The earlier emergence gives the dominant follicle a growth advantage over later emerging follicles, predisposing it to become dom inant (reviewed by [23] ). The acquisition of LH receptors in the gr anulosa cells of the future dominant follicle allows it to be able to grow in the low FSH environment caused by inhibin release , because it can respond to both LH and FSH (reviewed by [25] ). If progesterone levels are low, the dominant follicle continues to grow, the granulosa cells increase expre ssion of mRNA for P450 side chain cleavage, 3 β-hydroxysteroid dehydrogenase, and P450 aromatase (reviewed by [23] ). Gonadotropin receptors also increase in granulosa cells. The preovulatory dominant follicle becomes dependent on LH for continued growth and estradiol synthesis. Androgens are immediately converted to estradiol in the preovulatory dominant follicle [27] . This increase in estradiol production stimulates the hypothalamus to release a large quantity of GnRH, and subsequently LH, resulting in the LH surge which triggers ovulation (Figure 2.5 D). However, if progesterone levels are high during the follicular wave, such as those characteristic of the luteal phase discussed previously, ovulation does not occu r due to the suppressive effects of progesterone on GnRH release. The dominant follicle will then go through atresia which results in a change in the hormonal environment and another follicular wave occurs. The bovine corpus luteum will be maintained for approximately 16 days before it regresses (luteolysis). In th e bovine, the major luteolytic hormone is prostaglandin F 2α. 12

Prostaglandin F 2α is produced by the uterus and its release is stimulated by oxytocin. The down regulation of the progesterone receptor, as discussed previously, results in increased estradiol production. Estrogen stimulates the posterior pituitary to release oxytocin and to increase oxytocin receptors in the uterus. Pituitary oxytocin binds to its receptor in the uterus and causes low levels of prostaglandin F 2α to be released from the uterus. These low levels

of prostaglandin F 2α initiate the rel ease of luteal oxytocin. Luteal oxytocin augments the

release of uterine prostaglandin F 2α. High levels of prostaglandin F 2α inhibit progesterone secretion and cause functional luteolysis (reviewed by [14] ). Structural luteolysis follows, as described previously. After luteolysi s, the dominant follicle of the follicular wa ve can be ovulated. PEROXISOME PROLIFERA TOR -ACTIVATED RECEPTORS The peroxisome proliferator -activated receptors (PPARs) are transcription factors that are members of the nuclear receptor superfamily. The first PPAR was identified in 1990, by Isseman and G reen, and was suggested to play a role in triglyceride and cholesterol homeostasis [28] . Two other family members were later identified. The initial isotype was named PPAR α, and the other isotypes identified as PPAR β and PPAR γ. The PPARs have been detected in a wide range of tissues from many species. The PPARs are ligand -activated tra nscription factors that control gene expression by binding to peroxisome proliferator response elements (PPREs) in the promoter region of target genes (reviewed by [29] ). The PPARs form a heterodimer with the retinoid X receptors (RXR) and bind to the DNA response element (reviewed by [30] ). Four functional domains, characteristic of steroid hormone receptors, have been identifi ed in all three PPAR isoforms (reviewed by [31] ). The A/B domain contains an activation function element (reviewed by [31]). The C domain is the DNA binding domain (reviewed by [31]). The D domain is the hinge region and plays a role in receptor dimerization, as well as serving as a docking site for co -factors (reviewed by [31, 32] ). The 13

E/F domain contains the ligand -binding domain and is the site of hetero -dimerization (reviewed by [29] ). The PPARs associate with cofactors that either repress ( corepressors) or activate (coactivators) transcription (Figure 2.6; [29] ). Corepressors have histone deacetylase activity. Silenc ing mediator for retinoid and thyroid hormone receptors and nuclear receptor corepressor are two corepressors known to interact with the PPARs (reviewed by [33, 34] ). Coactivators have histone acetylase activity, which causes disruption of the nucleosome. Two examples of coactivators for the PPARs are cAMP response element binding protein and steroid receptor coactivator -1. These coactivators allow for chromatin decondensation near the regulatory region of a gene. A second coactivator complex , vitamin D receptor interacting protein /thryroid receptor accessory proteins , can then connect the PPAR:RXR complex to the general transcription factors, which results in initiation of transcription ( [35] , reviewed by [29] ). The ability of ligands to bind to the PPARs differs from other nuclear receptors. The first difference is that activation of the PPARs can occur through a wide range of li gands (reviewed by [36] ). Secondly, ligand recognition between the three isotypes overlaps, with some ligands binding to one o r more of the PPARs with different affinities (reviewed by

[36] ). Finally, the PPARs can accommodate several types of ligands, both synthetic and natural. There are both endogenous and exogenous ligands for the PPAR s. Common endogenous ligands that bind to the PPARs are eicosanoids a nd fatty acids (saturated and unsaturated; [36] ). There are also exogenous ligands that activate the PPARs. Certain phytoestrogens, such as genistein, have been found to bind to and activate PPAR γ [37, 38] . The thiazolidinediones (TZDs), drugs commonly administered to patients with type II diabetes, such as rosiglitazone and ciglitazone, are the most notable synthetic compounds that 14

can activate PPAR γ [39] . The clinical use of PPAR ligands warr ent investigating the role of this transcription factor in the ovary, t o determine how these therapies may affect female fertility. PPAR γ and Ovarian Biology All three PPARs have been identified in ovarian tissue. The expression of PPAR α

and β remains steady throughout follicular development and the ovarian cycle in the rat , while the expression of PPAR is differentially expressed during the ovarian cycle (reviewed by [40] ). Therefore, only the role of PPAR in ovarian biology will be di scussed. PPAR γ has been identified in ovarian tissue from several species, including humans [41] , pigs [42] , sheep [43] , mice [6] , rats [44, 45] , cattle [4, 5, 46] , zebrafish [47] , and Xenopu s laevis [48] . PPAR is expressed primarily in granulosa cells, and is different ially expressed during the estrous cycle. PPAR is located in granulosa cells of developing follicles and its expression is inversely related to that of the LH receptor [49] . In both the rat and sheep, the expression of PPAR in granulosa c ells is down -regulated in response to the

LH surge [1, 2] . These results indicate that a d ecline in PPAR γ may be needed for ovulation and/or luteinization to occur.

PPAR γ has been found to have various effects on ovarian steroid production. Studies investigating the role of PPAR γ in granulosa cell steroidogenesis have yielded conflicting resu lts. Agonists of PPAR γ stimulated progesterone and estradiol secretion from cultured granulosa cells from rats [2] . In sheep, progesterone production by granulosa c ells was also stimulated by agonists of PPAR [1] . Aromatase is the enzyme responsible for the conversion of androgens into estradiol. Komar and Curry (2003) found no correlation between expression of mRNA for PPAR γ and that for aromatase, either before or after the LH surge in Spraque -Dawley rats [49] . However, with a different strain of rat, Lovekamp - Swan et al . (2003) found that PPAR suppressed aromatase activity in granulosa cells [50] . A decrease in estradiol production due to suppression of aromatase activity was also seen in 15

human granulosa –luteal cells [51] . These studies show that PPAR plays a role in ovarian steroid production; however, its exact role has not been determined. PPAR has also been detected in luteal tissue. In the rat, luteal tissue from past

ovulations contained greater expression of mRNA for PPAR γ tha n luteal tissue from the most recent ovulation [3] . This is in contrast to the bovine corpus luteum, where PPAR γ

declines between the early and mid -luteal phase [4] . In bovine luteal tissue, PPAR γ has been found to be involved in progesterone production [4] . Lohrke et al . (1998) demonstrated that

two PPAR agonists, ciglitazone and PGJ 2, stimulated progesterone production from mid - phase luteal cells over a 24 -hour culture period. Treatment of human granulosa -lutein cells

with PGJ 2 had no effect on basal progesterone production, but inhibited LH -stimulated progesterone production [52] . Taken together, these data suggest that PPAR γ is an important player in luteal steroidogenesis.

Studies have determined that PPAR γ is active in the ovary and that there is endogenous ligand activity [1, 50, 53] . Sheep and ra t granulosa cells were transfected with a reporter construct driven by PPAR γ responsive elements [1, 50] . With or wit hout the addition of an agonist, there was an increase in reporter activity [1, 49, 52]. The increase in reporter activity without addition of a ligand indicates that there is an endogenous ligand for

PPAR γ in these cells. A study by Cui et al . (2002) dem onstrated the importance of PPAR in female fertility [6] . Using cre/loxP technology, Cui et al . (2002) produced a mouse with PPAR γ disrupted in the mammary epithelium, B - and T -cells, and the ovary. Cui et al . (2002)

demonstrated that female transgenic mice with the expression of PPAR disrupted in the ovary are sub - or infertile [6] . Because the expression of PPAR was not disrupted in the uterus of these transgenic animals, the authors concluded that the lesion causing the sub - and infertility resided in the ovary. 16

Cui et al. (2002) reported that on the day of estrus, circulating concentrations of progesterone were 50% lower in animals with PPAR disrupted in the ovary compared with controls. However, the difference was not significant, most likely due to the small sample size (n=4). Because progesteron e from the corpus luteum and estrogen from the ovary are needed for implantation, it is possible that the sub - and infertility in these mice results from ovarian dysfunction. This indicates a role for PPAR γ in proper ovarian function, affecting female fer tility.

Additional evidence supporting a critical role for PPAR γ in ovarian biology stem from findings in other systems. For example, PPAR has been found to play a role in angiogenesis (reviewed in [54] ), inflammation, and cell cycle control (reviewed in [29] ). These processes are important in normal ovarian function. Investigating how PPAR functions in these other processes may indicate the role(s) of PPAR in the ovary. Genes in the ovary that are involved in angiogenesis, inflammation and cell viability are vascular endothelial growth factor, cyclooxygenase -2 and bcl -2 respectively. These genes contain a peroxisome proliferator response elements (PPRE) in their promoter regions [55, 56] . A PPRE in the promoter region of these genes indicate that PP AR may regulate one or more of these genes, thereby playing a role in various processes associated with normal ovarian function .

As stated previously, PPAR is located in ovarian tissue and many studies indicate that PPAR plays an important role in ova rian function and fertility. The following projects were performed to determine the role(s) of PPAR in steroidogenesis, and to develop models of reduced expression of PPAR in the ovary. These models serve as a way to determine what genes may be regulat ed by PPAR . Collectively, these studies were able to identify candidate ovarian genes regulated by PPAR and possible role(s) of PPAR γ in ovarian steroid production . 17

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27. Hendriksen PJM, Gadella BM, Vos PLAM, Mullaart E, Kruip TAM, Dielemann SJ. Follicular Dynamics Around the Recruitment of the First Follicular Wave in the Cow. Biology of Reproduction 2003; 69: 2036 -2044. 28. Issemenn I, Green S. Activation of a M ember of the Steroid Hormone RSuperfamily by Peroxisome Proliferators. Nature 1990; 347: 645 -649. 29. Berger J, Moller DE. The Mechanisms of Action of PPARs. Annual Review of Medicine 2002; 53: 409 -435. 30. Kliewer Steven A, Xu Eric H, Lambert Millard H, W illson Timothy M. Peroxisome Proliferator -Activated Receptors: From Genes to Physiology. The Endocrine Society 2001: 239 -263. 31. Kota Bhavani P, Huang Tom H -W, Roufogalis Basil D. An Overview on Biological Mechanisms of PPARs. Pharmacological Research 200 5; 51: 85 -94. 32. Gelman L, Michalik L, Desvergne B, Wahli W. Kinase Signaling Cascades that Modulate Peroxisome Proliferator -Activated Receptors. Current Opinion in Cell Biology 2005; 17: 216 -222. 33. Escher P, Wahli W. Peroxisome Proliferator -Activated R eceptors: Insight into Multiple Cellular Functions. Mutation Research 2000; 448: 121 -138. 34. Miard S, Fajas L. Atypical Transcriptional Regulators and Cofactors of PPAR .

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38. Shen P, Liu MH, Ng TY, Chan YH, Yong EL. Differential Effects of Isoflavones, from Astragalus Membranaceus and Pueraria Thomsonii, on the Activation of

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Enha nces Progesterone Biosynthesis in Primary Cultures of Porcine Theca Cells. Biology of Reproduction 2002; 66: 190 -198. 44. Braissant O, Foufelle F, Scotto C, Dauca Michel and Wahli W. Differential Expression of Peroxisome Proliferator -Activated Receptors (P PARs): Tissue Distribution of PPAR -Alpha -Beta and Gamma in the Adult Rat. Endocrinology 1996; 137: 354 -366. 45. Komar CM, Berndtson AK, Evans AC, Fortune JE. Decline in Circulating Estradiol During the Periovulatory Period is Correlated with Decreases in E stradiol and 22

Androgen, and in Messenger RNA for p450 Aromatase and p450 17 α-hydroxylase, in Bovine Preovulatory Follicles. Biol Reprod. 2001; 64: 1797 -1805. 46. Mohan M, Malayer JR, Geisert RD, Morgan GL. Expression Patterns of Retinoid X Receptors, Retina ldehyde Dehydrogenase, and Peroxisome Proliferator Activated Receptor Gamma in Bovine Preattachment Embryos. Biology of Reproduction 2002; 66: 692 -700. 47. Ibabe A, Herrero A, Cajaraville MP. Modulation of Peroxisome Proliferator -

Activated Receptors (PPARs ) by PPAR α- and PPAR Specific Ligands and by 17 β- Estradiol in Isolated Zebrafish Hepatocytes. Toxicology in Vitro 2005; 19: 725 -735. 48. Dreyer C, Ellinger -Ziegelbauer H. Retinoic Acid Receptors and Nuclear Orphan Receptors in the Development of Xenopus laevis. In ternational Journal of Developmental Biology 1996; 40: 255 -262. 49. Komar Carolyn M, Curry Thomas E, Jr. Inverse Relationship Between the Expression of Messanger Ribonucleic Acid for Peroxisome Proliferator -Activated Receptor Gamma and P450 Side Chain Clea vage in the Rat Ovary. Biology of Reproduction 2003; 69: 549 -555. 50. Lovekamp -Swan T, Jetten Anton M, Davis Barbara J. Dual Activation of PPAR Alpha and PPAR Gamma by Mono -(2 -ethylhexyl) Phthalate in Rat Ovarian

Granulosa Cells. Molecular and Cellular End ocrinology 2003; 201: 133 -141. 51. Mu YM, Yanase T, Nishi Y, Waseda N, Oda T, Tanaka A, Takayanagi R, Nawata H. Insulin Sensitizer, Troglitazone, Directly Inhibits Aromatase Activity in Human Ovarian Granulosa Cells. Biochem.Biophys.Res.Commun. 2000; 271: 710 -713. 52. Willis DS, White J, Brosen J, Frank S. Effect of 15 -Deoxy -delta (12, 14) - Prostaglandin J2 (PGJ2) A Peroxisome Proliferator Activating Receptor Gamma (PPAR ) Ligand on Human Ovarian Steroidogenesis. The Endocrine Society 1999; P3 -247. 23

53. Lovekamp -Swan T, Chaffin CL. The Peroxisome Proliferator -Activated Receptor Ligand Troglitazone Induces Apoptosis and p53 in Rat Granulosa Cells. Molecular and Cellular E ndocrinology 2005; 233: 15 -24. 54. Margeli A, Kouraklis G, Theocharis S. Perxisome Proliferator Activated Receptor - Gamma (PPAR ) Ligands and Angiogenesis. Angiogenesis 2003; 6: 165 -169. 55. Butts BD, Tran NL, Briehl MM. Identification of a Functional Perox isome Proliferator Activated Receptor Response Element in the 3' Untranslated Region of the Human bcl -2 Gene. International Journal of Oncology 2004; 24: 1305 -1310. 56. Lemay DG, Hwang DH. Genome -Wide Identification of Peroxisome Proliferator Response Elem ents Using Integrated Computational Genomics. Journal of Lipid Research 2006; 47: 1583 -1587. 24

Figure 2 .1. Schematic illustration of the ovary showing the primary structures and their sequences of development. A) hilus, B) medulla, C) cortex, D) granulosa cells, E) theca cells, F) corpus luteum. Adapted from Senger 1999. 25

Figure 2.2. Schematic representation of the development of the ovarian follicle. A) primordial follicle, B) primary follicle , C) secondary follicle, D) Graafian follicle. Adapted from Griffin & Ojeda 2000. 26

CHOLESTEROL P450 side chain cleavag e THECA CELL PREGNENOLONE LH 3β-hydroxysteroid dehyrogenase PROGESTERONE 17 α-hydroxylase

ANDROGENS BASAL LAMINA

LH GRANULOSA ANDROGENS CELL FSH Aromatase

ESTRADIOL

Figure 2.3. Schematic representation of hormone production in theca and granulosa cells.

The cells of th e theca interna contain receptors for LH. Thecal cells produce androgens

(androstenedione or testosterone) through the actions of P450 side chain cleavage, 3 β- hydroxsteroid dehydrogenase and 17 α- hydroxylase. These androgens then diffuse across the basa l lamina into granulosa cells, which contain FSH receptors, and later during development, LH receptors (see text for details). 27

Figure 2.4 . Concentrations of progesterone (A), prolactin (B), estradiol (C), LH (D) and FSH (E) i n peripheral plasma throughout each day of the 4 -day estrous cycle of the rat. Adapted from Smith et al., 1975. 28

Figure 2.5. Follicular waves during the estrous cycle in cattle. Two or three waves of growth occur during the estrous cycle. Taken from Se nger 1999. 29

Figure 2.6. Gene transcription mechanisms of the PPARs. PPARs interacts with corepressors that inhibit gene transcription. PPARs can also interact with coactivators that contain histone acetylase activity and facilitate transcription of various genes. Adapted from Kota et al . 2005. 30

CHAPTER 3. POTENTIAL ROLE FOR PEROXISOME PROLIFERATOR -ACTIVATED RECEPTOR γ IN REGULATING LUTEAL LIFESPAN IN THE RAT A Paper Published in Reproduction 1 NS Tinfo 2, and CM Komar 3

ABSTRACT Peroxisome proliferator -activated receptor (PPAR ) has been shown to stimulate progesterone production by bovine luteal cells. We previously reported greater expression of PPAR in old compared to new luteal tissue in the rat. The following studi es were conducted to determine the role of PPAR in rat corpora lutea (CL) and test the hypothesis that PPAR plays a role in the metabolism of progesterone and/or luteal lifespan. Ovaries were removed from naturally cycling rats throughout the estrous cyc le, and pseudopregnant rats. Messenger RNA for PPAR and P -450 side -chain cleavage (SCC) was localized in luteal tissue by in situ hybridization, and protein corresponding to PPAR and macrophages identified by immunohistochemistry. Luteal tissue was cultu red with agonists (ciglitazone, PGJ2) or an antagonist (GW -9662) of PPAR . Progesterone was measured in media by RIA and levels of mRNA for 20 α-HSD and bcl -2 were measured in luteal tissue after culture by RT -PCR. An inverse relationship existed between th e expression of mRNA for SCC and PPAR . There was no effect of PPAR agonists or the antagonist on luteal progesterone production in vitro, or levels of mRNA for 20 α-HSD. PPAR protein was localized to the nuclei of luteal cells

1 Reprinted with permission of Reproduction , Tinfo &Komar, 2007, 133, 187 -96

© Soceity for Reproduction and Fertility (2007).

2 Primary researcher an d author

3 Author for correspondance 31

and did not correspond wit h the presence of macrophages. In new CLs ciglitazone decreased mRNA for bcl -2 on proestrus, estrus, and metestrus. Interestingly, GW -9662 also decreased mRNA for bcl -2 on proestrus and diestrus in old and new CLs, and metestrus in new CL. These data indi cate that PPAR is not a major player in luteal progesterone production or metabolism, but may be involved in regulating luteal lifespan. INTRODUCTION The peroxisome proliferator -activated receptors (PPARs) are transcription factors and members of the nuclear receptor s uperfamily. There are three isotypes of PPARs:

PPAR α, PPAR β/δ, and PPAR γ. Ovarian tissue from several species, including humans (Lambe & Tugwood 1996) , pigs (Schoppe et al. 2002) , sheep (Froment et al. 2003) , mice (Cui et al. 2002) , rats (Braissant et al. 1996, Komar et al . 2001) and cattle (Sundvold et al .

1997, Lohrke et al. 1998) express PPAR γ. Although this transcripti on factor has been detected in the ovaries of several species, to date very little is known regarding the role(s)

of PPAR γ in ovarian function. Previous work from our laboratory, using the rat as a model, has shown that

PPAR γ is expressed primarily in gr anulosa cells and is down -regulated in response to the LH surge (Komar et al. 2001) . PPAR γ was also identified in luteal tissue of the naturally cycling rat (Komar & Curry 2002) . Interestingly, the expression of mRNA for PPAR γ was higher in luteal tissue from previous ovulations compared with lut eal tissue forming from the most recent ovulation (Komar & Curry 2002) .

PPAR γ is involved in processes that are critical to normal ovarian function such as angiogenesis (Margeli et al. 2003) , inflammation, and cell cycle control (Berger & Moller

2002) , indicating that PPAR γ may be an important player regulating ovarian gene expression. For example, cell death via apoptosis occurs in both ovari an follicles and corpora lutea (CL). In the ovary, the anti -apoptotic gene, bcl -2, has been found to play a 32

role in cell survival. The gene encoding bcl -2 contains a peroxisome proliferator response

element (Butts et al. 2004) , and therefore could be directly regulated by PPAR γ. Another poss ible role for PPAR γ in the ovary may be in the process of luteolysis. The expression of PPAR γ is induced during the differentiation of monocytes into macrophages, and it is expressed in activated macrophages (Ricote et al . 1998, Tontonoz et al . 1998, Cunard et al. 20 02) . Macrophages accumulate in regressing CL in the rat from late proestrus to early estrus (Gaytan et al. 1998) . These macrophages are thought to aid in the removal of luteal tissue by phagocytosis (Niswender et al. 2000) . Therefore,

PPAR γ may play a role in luteolysis via its presence in activated macrophages. Alternatively, PPAR γ may be involved in luteal steroid production. In cattle, activation of PPAR γ has been reported to stimulate progesterone production by the C L (Lohrke et al. 1998) . Luteal tiss ue collected during the midphase of the bovine estrous cycle and

cultured with an agonist of PPAR γ, prostaglandin J 2 (PGJ 2), secreted more progesterone than control tissue (Lohrke et al. 1998) .

The objective of the current study was to determine the role of PPAR γ in the rat CL.

Given the ability of PPAR γ agonists to affect progesterone production by bovine luteal cells coupled with its expression pattern in the rat CL, we hypothesized that PPAR γ plays a role in luteal progesterone metabolism. Alternatively, due to its higher expression in old versus new

CL, and the fact that PPAR γ is expressed in macrophages which play a role in luteolysis, we also investigated the potential involvement of PPAR γ in regulating the life cycle of the CL. Both the naturally cycling and pseudopregnant rat were used as models to assess the impact

of PPAR γ on progesterone production at various physiological stages. The naturally cycling rat allowed for investigating t he role of PPAR γ in both newly forming luteal tissue derived from the most recent ovulation, and regressing luteal tissue present on the ovary from past ovulations. Because fully functional CL do not develop in cycling rats, the pseudopregnant rat was use d to investigate the role of PPAR γ in functioning luteal tissue. 33

METHODS AND MATERIA LS Animals

All animal procedures were approved by the Iowa State University’s Animal Care and Use Committee. Adult, female Sprague -Dawley rats were obtained from Harla n (Indianapolis, IN), and kept on a 14L:10D light:dark cycle. Vaginal smears were taken daily to monitor the estrous cycle. Animals exhibiting 3 normal estrous cycles were used in the following experiments. Ovaries were collected from naturally cyclin g rats to investigate the relationship between PPAR γ and luteal progesterone production. In addition, the association of PPAR γ and macrophages in luteal tissue was also investigated to determine if macrophages were the

cells expressing PPAR γ within the CL. Cycling rats were sacrificed on each day of the estrous cycle (proestrus, estrus, metestrus, and diestrus), between 09:00 and 10:00 am. Ovaries were removed, cleaned of adnexa and frozen until processed for in -situ hybridization, or fixed in 4% paraform aldehyde for immunohistochemical analysis, as described below. Luteal tissue was also established in culture as described below, to investigate how agonists and an antagonist of PPAR γ impact progesterone production and catabolism.

The expression and funct ion of PPAR γ was also investigated in luteal tissue from pseudopregnant rats. Rats were mated with a vasectomized male and sacrificed 10 -16 days later (n=4 animals). Luteal tissue was dissected from one ovary from each animal and cultured in vitro as des cribed below. The second ovary from each animal was fixed in 4% paraformaldehyde and embedded in paraffin for immunohistochemical analysis. In -situ hybridization

Frozen ovarian tissue collected from rats on the days of proestrus, estrus, metestrus, and diestrus (n= 4 animals/day) were s erially sectioned at 10 µM. Consecutive tissue 34

sections were used for localization of mRNA for P -450 side -chain cleavage (SCC) and

PPAR γ. Tissue sections were fixed in 4% paraformaldehyde and dehydrated by passing through a series of increasing ethanol s olutions. Slides were placed in 2 mg/ml glycine in phosphate buffered saline (PBS; pH=7.2), rinsed in PBS, followed by an incubation in 1.5% triethanolamine buffer with 0.25% acetic anhydride. Slides were subsequently treated with 2x standard saline citr ate (SSC; 0.149 M sodium chloride, 14.9 mM sodium citrate, pH= 7.0) and dehydrated by passing through a series of decreasing concentrations of ethanol.

Sense and antisense riboprobes for PPAR and SCC (plasmids containing the cDNA for PPAR and SCC were generously provided by Dr. Walter Walhi, Universite de Lausanne, Lausanne, Switzerland and Dr. J. S. Richards, Baylor College of Medicine, Houston, TX, re spectively) were synthesized using a MAXISCRIPT kit (Ambion, Inc. Austin, TX) and -

35 S-UTP (10 µmCi/ml; ICN Biomedical, Inc., Irvine, CA). Tissues were hybridized with radiolabeled probe (1 x 10 6 cpm) in 100 µl hybridization buffer (4 M NaCl, 1 M Tris, 0 .5 M EDTA, 50x Denhardts, 25 mg/ml yeast tRNA, 10 mg/ml polyadenylic acid). After hybridization, slides were incubated in RNase solution 1 (2x SSC, 50% formamide, 0.1% - mercaptoethanol), followed by incubation in RNase solution 2 (0.5 M NaCl, 10 mM Tris, 20

µg/ml RNase A). Slides were incubated in RNase solution 1 a second time, after which they were incubated in 0.1x SSC, 1% -mercaptoethanol. Slides were dehydrated by rinsing with 30% ethanol containing 0.6 M NaCl, 60% ethanol containing 0.6 M NaCl, 8 0%, 95%, and 100% ethanol. The slides were air dried, dipped in Kodak NTB2 emulsion and exposed at

4°C for 1 week (SCC) or 4 weeks (PPAR ). Slides were developed before being counterstained with hematoxylin. At least one slide was processed with the sen se riboprobe and two slides with the antisense riboprobe (4 tissue sections/slide) per animal. Tissue Cultu re

Luteal tissue from ovaries collected throughout the estrous cycle was dissected from the ovary using 18 gauge needles. CLs from the current c ycle (new) were differentiated from 35

CLs from previous cycles (old) by assessing the degree of vascularization (Fig. 3.1 ). New CLs were well vascularized, whereas CLs present on the ovary from past cycles were less vascularized (Bassette 1943) . There was no attempt to discern the differences betw een old CL s regarding how may cycles they ha d been present on the ovary. Individual CLs were hemisected and cultured in 500 µl of defined media (DMEM: F12, 0.01% sodium pyruvate, 0.22% sodium bicarbonate, 1% bovine serum albumin, 0.125% gentamicin, and insulin - transferrin -sodium selenite media supplement; pH= 7.2). In one ex periment, CLs from the

day of estrus (n=5 animals) were cultured in the presence or absence of the PPAR γ agonists,

ciglitazone (65 µM) or PGJ 2 (25 µM). Agents were added at the time tissues were established in culture and treatments were set up in duplica te. In a second experiment, luteal tissue from the days of proestrus, estrus, metestrus, and diestrus (n=4 animals/day of the

cycle) were cultured with ciglitazone (65 µM) or the PPAR γ antagonist GW -9662 (1 µM). The antag onist , GW -9662 was used because i t has previously been shown that there is

endogenous activity of PPAR γ in the ovary (Froment et al. 2003, Lovekamp -Swan & Chaffin

2005) . Luteal tissue from pseudopregnant rats was also cultured with PGJ 2 (25 µM) or GW -

9662 (1 µM). Tissues were incubated for 24 hours at 37°C in 5% CO 2: 95% air. At the end of culture, media were collected and stored at -20°C until analyzed by RIA for progesterone. Tissues were frozen and stored until used for RNA and DNA trizol extraction. Concentrations of progesterone were corrected by amount of DNA/wel l. Samples of total RNA were analyzed by formaldehyde/agarose gel electrophoresis to assess tissue viability at the end of culture. RIA

Concentrations of progesterone in conditioned media were determined as described previously (Komar et al. 2001) . Th e intra - and inter -assay coefficients of variation were 2.3% and 13.9% respectively. 36

PCR

Total RNA was extracted from cultured tissues using Trizol reagent. Complementary DNA was synthesized using SuperScript II Reverse Transcriptase and oligo (dT) r andom pri mers (Invitrogen). The cDNA obtained was analyzed by PCR to determine the expression of 20 α - hydroxysteriod dehydrogenase (20 α-HSD) and bcl -2 in luteal tissue. S16 and GAPDH were also analyzed and used as internal controls (Ko et al. 1999, Mendoza -

Rodriguez et al . 200 3) . Primers utilized for 20 α-HSD (Sugino et al. 1997) , bcl -2, GAPDH (Mendoza -Rodriguez et al. 2003) and S16 (Ko et al. 1999) were published previously.

PCR reactions (25µl) for 20 α-HSD and S16 consisted of 2.5 mM MgCl 2, 1X PCR buffer (200 mM Tris -HCl, 500 mM KCl), 2 µM of forward and reverse primers for both genes, 200 µM dNTPs, 1.0 unit of Taq polymerase, 10x bovine serum albumin (BSA) and 50 ng of cDNA. All amplifications were carried out for 30 cycles with denaturation at 95°C for 2 minutes, annealing at 65°C for 1 minute, and extension at 72°C for 1 minute per cycle. PCR reactions (25 µl) for bcl -2 and GAPDH consisted of 20 mM Tris -HCl (pH 8.3),

50 mM KCl, 1.0 mM MgCl 2, 0.2 mM dNTPs, 0.5 µM of forward and reverse primers for both bcl -2 and GAPDH, 2.5 units of Taq DNA polymerase and 50 ng of cDNA for GAPDH, and 500 ng of cDNA for bcl -2. All amplifications were carried out for 35 cycles, with denatur ation at 95°C for 5 minutes, annealing at 60°C for 1 minute, and extension at 72°C for 5 minute per cycle. Samples were analyzed in duplicate, and the analysis repeated three times. PCR products were separated on a 2% agarose gel. Densitometric analys is of the resulting bands was conducted using the Spot Denso program (Fluorche m, Alpha Innotech). Relative levels of 20 α-HSD and bcl -2 were determined in relation to S16 and GAPDH, respectively, per sample. 37

Immunohistochemistry

Paraffin embedded ovarian tissue was serially sectioned at 5 µm. Consecutive sections were processed for immunodetection of PPAR γ and macrophages. Paraffin sections were dewaxed in xylene and rehydrated through a series of rinses in decreasing ethanol solutions. Antigen retrieval was performed by placing slides in 0.01M Tris (pH=8.5) in a

microwave on high heat for 10 minutes. After cooling, slides were placed in 1.5% H 2O2 in methanol to quench endogenous peroxidase activity. After rinsing with PBS (pH=7.2), slides were blocked with 10% normal horse (macrophage) or goat (PPAR γ) serum. Slides were the n incubated with either a monoclonal antibody against rat monocytes/macrophages (ED1 clone, Chemicon, Temecula, CA) at a 1:200 dilution for 1 hour at 37°C, or an antibody against PPAR γ (E8, Santa Cruz, CA) at a 1:50 dilution overnight at 4°C. Slides were rinsed with PBS and subsequently incubated with a species specific biotinylated secondary antibody at a 1:200 dilution for macrophage detection (Vector Lab Inc, Burlingame CA) or 1:50 dilution for PPAR γ detection (Amersham). Slides were rinsed in PBS and treated with an avidin -peroxidase complex (ABC kit; Vector Lab Inc). To visualize the immunocomplex, 3, 3’ -diaminobenzidine tetrahydrochloride (DAB) was used. Slides processed for immunodetection of macrophages were counterstained with hematoxylin. All slides were mounted with Permount. For detection of both macrophages and PPAR γ, a minimum of one control slide (blocking serum used in place of primary antibody) and two treated slides were examined per rat (4 ovarian sections/slide). Statistical analys is

Differences in levels of mRNA for 20 α-HSD and bcl -2 were analyzed by ANOVA (SAS Version 8.2, SAS Institute, Cary, NC) followed by the Student Newman Kuels test for comparisons of multiple groups when appropriate (JMP 5.1). Concentrations of progestero ne 38

in conditioned media from old and new CL were log transformed and analyzed

independently by ANOVA. A p -value ≤ 0.05 was considered significant. RESULTS To investigate the relationship between PPAR γ and progesterone production, mRNAs for SCC and PPAR γ were localized in luteal tissue from naturally cycling rats by in -situ hybridization. New and old CLs were differentiated histologically, with new CLs containing more luteal cells and less stromal elements than old CLs (Malven & Sawyer 1966, Gaytan 1997, Simpson 2001) . An inverse re lationship was observed between the expression of

mRNA for SCC and PPAR γ. The expression of mRNA for SCC was greater in newly forming luteal tissue compared with that observed in old luteal tissue on all days of the cycle

(Fig. 3. 2). In contrast, the expression of mRNA for PPAR γ was greate r in old luteal tissue compared to newly forming luteal tissue (Fig. 3. 3). The expression of mRNA for PPAR γ appeared in a punctate pattern in some luteal tissue, most notably on the day of diestrus (Fig. 3. 3 j).

Because of the inverse relationship betw een mRNA for SCC and PPAR γ in luteal tissue with PPAR γ being expressed at a higher level in old CL versus new CL, we investigated the potential of PPAR γ to influence the metabolism of progesterone. Levels of mRNA for 20 α-HSD, the enzyme responsible for th e breakdown of progesterone into its inactive metabolite, were measured in luteal tissue collected on the day of estrus and cultured

with ciglitazone or PGJ 2. Messenger RNA for 20 α-HSD was expressed in new and old CL, but expression did not change in resp onse to treatment with either PPAR γ agonist (Fig. 3. 4). Luteal tissue from the days of proestrus, estrus, metestrus, and diestrus was cultured

in vitro to determine how activation or inhibition of PPAR γ would impact progesterone production throughout the e strous cycle. Old and new CLs were separated and cultured with

ciglitazone (65 µM) or GW -9662 (1 µM). There was no effect of activating or inhibiting PPAR γ on progesterone production by new or old CLs collected throughout the estrous cycle 39

(Fig. 3. 5 a & b). However, a dministration of GW -9662 to old CL on proestrus tended to increase progesterone prod uction compared to controls (p = 0.06 , Fig. 3. 5 b). The secretion of progesterone in vitro by fully functional luteal tissue collected from pseudopregnant r ats

was also not affected by treatment with either PGJ 2 or GW -9662 (data not shown). PPAR γ and macrophages were immunolocalized in serial tissue sections of ovaries collected throughout the cycle and during pseudopregnancy to determine if the expression of

PPAR γ in CLs corresponded to macrophages within the tissue. Positive staining for E D1 (macrophages) was observed in luteal tissue from cycling (Fig. 3. 6 a-d) and pseudopregnant (Fig. 3. 6 e) rats. Luteal tissue on the day of diestrus contained low levels of labeling for ED1

(Fig. 3. 6 d) relative to the other days of the cycle. Protein c orresponding to PPAR γ was detected in the nuclei of luteal cells on all days of the cycle, albeit at a lower level than in granulosa cells of developing follicles (Fig. 3. 7 a). Protein corresponding to PPAR γ was also observed in nuclei of luteal cells during pseudopregna ncy at a relatively low level (data not shown).

The role of PPAR γ in luteal cell survival was investigated by measuring levels of mRNA for bcl -2 in tissue collected throughout the cycle and cultured with ciglitazone or GW -9662. In newly forming luteal t issue collected on all days of the cycle except diestrus, treatment with ciglitazone reduced levels of mRNA for bcl -2 compared to controls (Fig. 3. 8 a, p < 0.05, within day of the cycle). However, the administration of ciglitazone to old CL in vitro resul ted in a decrease in levels of mRNA for bcl -2 only when the tissue was collected on the day of estrus (Fig. 3. 8 b, p < 0.05, within the day of the cycle). Interestingly, treating luteal tissue in vitro with the PPAR γ antagonist, GW -9662, also reduced levels of mRNA for bcl -2 in both new and old CL collected on the days of proestrus and diestrus, as well as new CL collected on metestrus (Fig. 3. 8 a & b, p < 0.05, within day of the cycle). 40

DISCUSSION PP AR γ has been detected in ovarian tissue from several species. However, at this time, it is unknown how this transcription factor impacts ovarian function. Research conducted by Cui et al . (2002) demonstrated that in transgenic mice with the expression of

PPAR γ disrupted in the ovary, the females were sub - or infertile (Cui et al. 2002) . Because the ex pression of PPAR γ was not disrupted in the uterus of these transgenic animals, the authors concluded that the lesion causing the sub - and infertility resided in the ovary. These authors reported that on the day of estrus, circulating concentrations of pro gesterone were

50% lower in animals with PPAR γ disrupted in the ovary compared with controls, although the difference was not significant most likely due to the small sample size (n=4). In cattle, agonists of PPAR γ were found to stimulate progesterone pro duction from mid -phase luteal cells over a 24 hour culture period (Lohrke et al. 1998) . Treatment of human granulosa - lutein cells with PGJ 2 had no effect on basal progesterone production, but inhibited LH - stimulated progesterone production (Willis et al . 1999). Taken together, these data s uggest

that PPAR γ may influence the ability of luteal tissue to produce progesterone, which is necessary for the establishment and maintenance of pregnancy.

The data presented here illustrate that PPAR γ is not a major player in progesterone production in the rat CL. In luteal tissue collected throughout the estrous cycle, as well as from pseudopregnant rats, there was no significant effect of PPAR γ agonists or the antagonist on progesterone production. Our data also show that there is an inverse relatio nship between the expression of PPAR γ in luteal tissue and the ability of that tissue to produce progesterone - as determined by levels of mRNA for SCC. These data agree with a previously published report on the gonadotropin -treated immature rat where an inverse relationship between the expression of mRNA for PPAR γ and SCC in granulosa cells was detected during the periovulatory period (Komar & Curry 2003). These findings indicate that down -regulation of PPAR γ may be important for the change in gene expre ssion involved in luteinization and 41

the switch in steroid production from estrogen to progesterone in the rat. The finding in the

current study that an antagonist of PPAR γ tended to increase progesterone production by cultured luteal tissue further suppor ts the hypothesis that down regulation of PPAR γ may be important for periovulatory progesterone production in the rat. These findings differ from those reported by Lohrke et al . (1998) and could be the result of differences in culture methods and/or speci es differences in luteal development and function. The current study utilized hemisected luteal tissue in vitro , whereas Lohrke et al. (1998) cultured dispersed luteal cells (Lohrke et al. 1998) . Separation of large and small luteal cells alters steroidogenesis. Dispersed luteal cells pro duced lower amounts of hormones in vitro compared to intact tissue from the same CL ( Pate & Nephew 1988, Jaroszewski et al. 2003) . Therefore, the use of intact luteal tissue provides a more accurate model of in vivo luteal function. In cycling rodents, mating or cervical stimulation is necessary for the development of fully functional corpora lutea. In contrast, after ovulation in cattle, luteal tissue develops regardless of whether the female is mated. The hormonal environment of the luteal phase also differs between the rat and cow. In the rat the hormone prolactin is ne eded for luteal development and demise (Freeman et al . 2000), while prolactin does not play a role in luteal function in cattle (Niswender et al . 2000). Another difference between the bovine and rat is that the rat ovary contains CLs from past ovul ations (old CLs) as well as CLs from the most recent ovulation (new CLs). Multiple cycles are needed for the complete regression of old CLs (Freeman et al . 2000). This is in contrast to the bovine, whose luteal tissue develops from the most recent ovulat ion and regresses in the open animal before the next estrous cycle. In the rat, the secretion of progesterone from newly forming luteal tissue increases on the evening of proestrus. By diestrus, there is an increase in the activity of 20 α-HSD, the enzyme responsible for converting progesterone into its inactive metabolite, 20 α-hydroxyprogesterone (Diaz et al . 2002). A decrease in progesterone may trigger the increase in expression of 20 α-HSD, 42

leading to progesterone catabolism (Stocco et al . 2001). In a study by Yoshida et al. (1999), the expression of 20 α-HSD was greater in new luteal tissue collected at the end of pseudopregnancy compared with that at the beginning (Yoshida et al. 1999) . This expression pattern of 20 α-HSD most likely reflects the fact that at the beginning of pseudopregnancy, luteal tissue would secrete progesterone which is needed for the establishment and maintenance of pregnancy, whereas at the end of pseudopregnancy a decline in progesterone would correspond with parturition at the end of gestation. In the current study, agonists of

PPAR γ had no effect on the expression of mRNA for 20 α-HSD in either new or old luteal tissue. These findings indicate that just as PPA Rγ is not a major player in progesterone production, it also does not affect the breakdown of progesterone via regulating the expression of mRNA for 20 α-HSD. Regression of the CL involves functional (decrease in progesterone production), and structural c hanges (regression of luteal tissue, apoptosis, and macrophage infiltration) to the tissue. Accumulation of macrophages in CL has been found to occur from the evening of proestrus to the morning of estrus in the rat (Gaytan et al . 1998). The current stud y examined the expression of macrophages in both the naturally cycling and pseudopregant rat models. In the cycling rat, prolactin is necessary for the establishment of the macrophage population in new luteal tissue as well as for the influx of monocytes and macrophages into regressing CLs (Gaytan et al . 1997). Immunodetection of macrophages and PPAR γ in serial sections of ovarian tissue demonstrated that they are localized primarily to different areas within luteal tissue. Macrophages were situated most ly around luteal cells, while PPAR γ was detected in the nuclei of luteal cells. Therefore, the higher expression of PPAR γ in old luteal tissue compared to newly forming luteal tissue is not due solely to the presence of macrophages within the tissue. The lack of correlation between PPAR γ and macrophages may be due to the fact that the antibody used in the current study recognizes both monocytes 43

and macrophages. PPAR γ is expressed in activated macrophages and therefore would not colocalize with monocytes . Apoptosis, or programmed cell death, plays a role in luteal regression. Apoptosis is modulated by a number of regulatory genes, such as bcl -2. Bcl -2 is a member of the B -cell leukemia/lymphoma family, and plays a role in cell survival in the ov ary. In the human and rodent, protein corresponding to bcl -2 has been detected in the corpus luteum and granulosa cells (Richards 1994, Rodger et al . 1998, Leng et al . 2001, Kim & Tilly 2004). As mentioned previously, the human gene encoding bcl -2 contai ns a peroxisome proliferator response element (Butts et al . 2004). Because bcl -2 and PPAR γ are located in the same tissues within the ovary, PPAR γ may affect ovarian cell survival via regulating the expression of bcl -2. In support of that hypothesis, a s tudy using cultured granulosa cells

showed that treatment with the PPAR γ agonist, troglitazone, decreased protein levels for bcl - 2 (Lovekamp -Swan & Chaffin 2005). In the current study, levels of mRNA for bcl -2 in old

CL were reduced by treatment with a PP AR γ agonist when the tissue was collected on the day of estrus, and by treatment with a PPAR γ antagonist when tissue was collected on proestrus and diestrus. This reduction in mRNA for bcl -2 in response to both an agonist and antagonist of PPAR γ was also seen in new CL collected on proestrus and metestrus. These apparent incongruous findings of both an agonist and antagonist of PPAR γ inhibiting expression of mRNA for bcl -2 may be due to PPAR γ -dependent and -independent effects of these agents. Ciglitazo ne is a member of the thiazolidinedione (TZD) family of drugs. Troglitazone, another TZD, has been shown to have cellular effects that are not mediated by activation of PPAR γ (Chawla et al . 2001), and reportedly can have direct effects on ovarian cells (G asic et al. 2001). GW -9662 has also been shown to act as a PPAR γ agonist in a normal human mammary epithelial cell line (Allred & Kilgore 2005). Further study is warranted to determine how activation/inhibition of PPAR γ impacts luteal cell survival. 44

In conclusion, our data suggest that PPAR γ does not play a significant role in progesterone production in the naturally cycling or pseudopregnant rat. The expression pattern of PPAR γ is inversely related to the ability of luteal tissue to produce progester one, and agonists of PPAR γ had no effect on the ability of luteal tissue to secrete or breakdown progesterone. PPAR γ is located in the nucleus of luteal cells where it may influence luteal cell survival via regulating the expression of bcl -2. Future stud ies are needed to further investigate the role of PPAR γ in luteal cell apoptosis and how its varying expression is regulated during the lifespan of the CL. ACKNOWLEDGEMENTS The authors would like to thank Dr. Walter Wahli for the plasmid containing th e

cDNA for PPAR γ, and Dr. J. S. Richards for the SCC plasmid. We would also like to thank Dr. S. Lamont for use of the PCR machine and Dr. Steven Lonergan and Dr. Elizabeth Huff - Lonergan for use of the Alpha Tech Imager. REFERENCES Allred C & Kilgore MW 2 005 Selective activation of PPAR γ in breast, colon, and lung cancer cell lines. Molecular and Cellular Endocrinology. 235 21 -29.

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OCL

* NCL * a b

Figure 3.1 . Image of an ovary collected on the day of metestrus from a naturally cycling rat. New luteal tissue is more vascularized than old luteal tissue (a). Arrows ( →) denote new luteal tissue, asterisk (*) denotes old luteal tissue. Histological image of old and new luteal tissue (b). OCL, old luteal tissue; NCL, new luteal tissue. 51

Figure 3.2. Localization of mRNA correspo nding to SCC in ovarian tissue collected from rats on the day of proestrus ( a, b and c), estrus ( d, e and f), metestrus ( g, h and i), and diestrus ( j, k and l). Tissue sections (10 µM) were hybridized with 35 S-labeled sense and antisense riboprobes as des cribed in the Materials and Methods . The first column represents darkfield images of ovarian tissue sections labeled with an antisense riboprobe ( a, d, g, j ) with the corresponding brightfield image in the second column ( b, e, h, k) . Darkfield images of tissue hybridized with a sense riboprobe appear in the third column ( c, f, i, l ). GC = Granulosa

Cells, OCL = old luteal tissue, NCL = new luteal tissue. Original magnification = 100X. 52

Figure 3.3. Localization of mRNA corresponding to PPAR γ in ovarian tissue collected from rats on the day of proestrus ( a, b and c), estrus ( d, e and f), metestrus ( g, h and i), and diestrus ( j, k and l). Panels a and b contain an inset image of a NCL from another area of the ovary for comparison. Tissue sections (10 µM) were hybridized with 35 S-labeled sense and antisense riboprobes as described in the Materials and Methods . The first column represents darkfield images of ovarian tissue sections labeled with an antisense riboprobe ( a, d, g, j ) with the corresponding br ightfield image in the second column ( b, e, h, k). Darkfield images of tissue hybridized with a sense riboprobe appear in the third column ( c, f, i, l ). GC = Granulosa Cells, OCL = old luteal tissue, NCL = new luteal tissue. Original magnification = 100 X. 53

Figure 3.4. Relati ve levels of mRNA (mean ± S.E.M.) for 20 α-HSD in rat luteal tissue on the day of estrus as described in the Materials and Methods. The image depicts

PCR products for 20 α-HSD and S16 with corresponding levels of mRNA for 20 α-HSD in

new and old CL cultured alone (control) or with ciglitazone (65 µM) or PGJ 2 (25 µM). OCL, old luteal tissue, NCL, new luteal tissue. 54

Figure 3.5. Progesterone production (mean ± S.E.M.) by new (a) and old (b) luteal tissue collected from naturally cycling rats on all days of the cycle and cultured alone

(control) or with ciglitazone (65 µM) or GW -9662 (1 µM). 55

Figure 3.6. Immunodectection of macrophages (brown color) in sections of ovarian tissue collected from rats on proestrus ( a), estrus ( b), metestrus ( c), diestrus ( d) and during pseudopregnancy ( e). Tissue section from an ovary collected on the day o f proestrus with normal horse serum used in place of the primary antibody as a control ( f).

Original magnification = 50X. 56 a b

GC GC c d

Figure 3.7. Immunodectection of PPAR γ (a and c) and macrophages (b and d) in consecutive tissue sections of new (a and b) and old (c and d) CL in ovaries collected from rats on proestrus. Original magnification =100X. Arrows indicate labeling for

PPAR γ in the nuclei of luteal cells. GC = Granulosa Cells. 57

Figure 3.8. Relative levels of mRNA for bcl -2 and GAPDH in rat luteal tiss ue collected throughout the cycle as described in the Materials and Methods . ( a) PCR products for bcl -2 and GAPDH with corresponding levels of mRNA for bcl -2 in new CL cultured alone (control) or with ciglitazone (65µM) or GW -9662 (1µM). ( b) PCR products for bcl -2 and GAPDH with corresponding levels of mRNA for bcl -2 in old CL cultured alone (control) or with ciglitazone (65µM) or GW -9662 (1µM). Bars with different superscripts denote significant differences within the day of the cycle (p <

0.05). 58

CHAPTER 4. DEVELOPMENT OF MODELS TO FACILITATE INVESTIGATION OF OVARIAN GENES REGULATED BY THE TRANSCRIPTION FACTOR, PEROXISOME PROLIFERATOR -ACTIVATE RECEPTOR γ

NS Tinfo, LJ Luense, P Liu, and CM Komar

ABSTRACT

Peroxisome proliferator -activated receptor γ (PPAR γ) is a transcription factor t hat has been found to be involved in many processes such as steroidogenesis, angiogenesis, and tissue remodeling. These processes are critical for normal ovarian function. PPAR γ has been detected in granulosa, theca, and luteal cells. Previous studies h ave identified that

PPAR γ is differentially expressed in granulosa cells in the ovary during the ovarian cycle. However, the roles of PPAR γ in the ovary have not been determined. Previous attempts to create a PPAR γ knock -out mouse resulted in an embryon ic lethal phenotype, making it difficult to investigate the role of PPAR γ in the ovary. Therefore, to determine the role(s) of PPAR γ in ovarian biology, cre/loxP technology was used to create two models, one in vivo and one in vitro , with reduced expressi on of PPAR γ specifically in ovarian tissue. These two models allowed for the discovery of candidate genes regulated by

PPAR γ and will allow for further study of this transcription factor in ovarian biology. INTRODUCTION Peroxisome proliferator -activated r eceptors (PPARs) are transcription factors and

members of the nuclear receptor superfamily [1] . There are three isotypes of PPARs: α, β, and γ. The PPARs have been found to be involved in many processes, such as steroidogenesis, angiogenesis, and tissue remodeling. These processes are critical for normal ovarian function. All three PPAR isotypes have been identif ied in the ovary [2] , but only 59

PPAR γ is differentially expressed. Certain ligands of PPAR , the thiazolidinedions, are commonly administered to patients with type II diabetes and polycystic ovarian syndrome [3] . Since the role(s) of PPAR in the ovary has not been determined, it is not known how activation of PPAR by these drugs is affecting female fertility.

PPAR γ has been detected in ovarian tissue from humans [4] , pigs [5] , sheep [6] , mice [7] , rats [8] , and cattl e [9] . This transcription factor is found primarily in granulosa cells of

developing follicles. PPAR γ has also been detected in luteal and thecal tissue, but at a lower level than in granulosa cells [2, 6, 8, 10] . Along with the localization of PPAR γ in the ovary, previous studies have also demonstrated that agonists of PPAR γ can affect steroidogenesis (reviewed in [11] ). The inf luence of PPAR γ on steroid production varies within and between species , suggesting that the endocrine environment of the animal influences how PPAR affects ovarian function .

PPAR γ is present in the ovary and it has also been found to be active in ovari an cells [6, 12, 13] . Sheep and rat granulosa cells were transfected with a reporter construct driven by PPAR γ responsive elements [6, 13] . With or without addition of an agonist, there was an increase in reporter activity [6, 12, 13] . The increase in reporter activity in the absence of exogenous ligand indicate d that PPA Rγ is active in granulosa cells and that an endogenous ligand is present in these cells.

Several studies have indicated that P PAR plays an important role in ovarian biology. Previous research on mice with PPAR disrupted in mammary epithelium, B - and T -cells, and the ovary resulted in female mice being either sub - or infertile [7] . This study illustrates the importance of PPAR in fertility. PPAR has also been shown to be involved in processes other than steroid production that are critical for normal ovarian function. Two of these processes are angiogenesis and inflammation. Angiogenesis, the formation of blood vessels, is required for follicular growth and formation/function of corpora lutea (reviewed in [14] ). PPAR has been found to have varying effects on angiogenesis (reviewed in [15] ). 60

The effect of PPAR γ agonists on angiogenesis in the ovary has not been determined. The promoter region of the rate limiting enzyme in prostaglandin production, cyclooxygenase -2 (COX -2), contains a response element for the PPARs [16] . Ovulation and luteolysis have been likened to an inflammatory response [17] . Prostaglandins are involved in ovulation and luteolysis, and are major regulators of inflammation [18] . Prostaglandins have also been shown to be endo genous ligands for the PPAR [19] . T he fact that prostagla ndins are ligands for PPAR , and that the promoter region of COX -2 contains a response element for the

PPARs, suggest that PPAR γ may regulate COX -2 expression in ovarian cells, and thereby play a role in ovulation and/or luteolysis.

The exact role(s) of PPAR in the ovary has not been determined. Studying PPAR γ in the ovary has been difficult due to the fact that knocking out PPAR γ in the genome results in an embryonic lethal phenotype. There is a need for a n experimental model that will allow for the s tudy of this transcription factor in ovarian function. Therefore, t wo models, one in

vivo and one in vitro , of reduced expression of PPAR γ in the ovary were developed. These model s utilized cre/loxP technology to reduce expression of PPAR γ in granulosa c ells. The in vivo model has been used to identify candidate genes regulated by PPAR . Future studies will investigate how the regulation of these genes affects normal ovarian function. METHODS AND MATERIAL S

Development of an in vivo model to study the role of PPAR in the ovary Cre/loxP technology allows for the production of mice with reduced expression of a particular gene in a tissue selective manner. Genetic engineering is used to generate mice expressing cre recombinase , a bacterial enzyme that recognizes and cuts at loxP sites. The use of specific promoters to drive expression of cre recombinase results in this enzyme being active in a tissue specific manner. Genetic engineering is also used to create mice with loxP sites flanking a gene of in terest. By crossing these two lines of mice, the tissue selective 61

expression and activity of cre recominbase will recombine the gene containing the loxP sites, resulting in tissue selective reduction in expression of that gene. Generation of mice with re duced expression of PPAR γ in the ovary. All animal procedures were approved by the Iowa State University Animal Care and Use Committee. Two different lines of mice were mated to produce animals with reduced expression of

PPAR γ in the ovary. One mouse li ne, developed by Jamin et al. (2002), had cre recombinase inserted into the gene encoding anti -Mullerian hormone receptor 2 (Amhr2) (Amhr2 cre/wt ; generously provided by Dr. Richard R. Behringer). Amhr2 is expressed solely in the gonads [20] and, in females, specifically in granulosa cells [21] . Mice with loxP sites inserted on

either side of exon 2 of PPAR , were generated by Akiyama et al. (2002) (PPAR γfl/fl , kindly donated by Dr. Frank J Gonzalez). Homozygous PPAR γfl/fl mice were crossed with mice heterozygous for insertion of cre recombinase into the Amhr2 gene (Amhr2 cre/wt ). The resulting F1 progeny w ere genotyped and mice carrying the allele for cre recombinase

(PPAR γfl/wt , Amhr2 cre/wt ) were mated to founder PPAR γfl/fl mice to obtain animals homozygous for the floxed PPAR γ allele and heterozygous for cre recombinase (PPAR γfl/fl , Amhr2 cre/wt ). The res ulting progeny (PPAR γfl/fl , Amhr2 cre/wt ) were backcrossed to the PPAR γfl/fl founder line for six generations to minimize genetic variation. DNA Extraction and Genotyping. Tail biopsies, approximately 0.5 cm long, were collected from 9 to 11 day old mice. Tissues were dissolved in 465 µl of tail extraction buffer (50 mM Tris pH 7.5, 100mM EDTA, 0.5% SDS, and 650 mg/µl proteinase K) by incubating overnight at 55°C. The following day, 250 µl of dissolved tissue solution were added to an equal amount of phen ol:chloroform:isoamyl alcohol (25:24:1; Invitrogen), mixed vigorously, and centrifuged at 15,000 rpm for 5 minutes at 4°C. The supernatant was removed and extracted twice with phenol:chloroform. Following the final extraction, the aqueous portion was rem oved and added to 1 ml of cold 100% ethyl alcohol to precipitate the DNA. Samples were centrifuged at 15,000 rpm for 15 minutes at 4°C prior to supernatant 62

aspiration and rinsing of the pelleted DNA with room temperature 70% ethyl alcohol. The DNA was th en vortexed and centrifuged at 15,000 rpm for 5 minutes at 4°C, the ethyl alcohol was discarded, and the pellets allowed to air dry for 5 to 10 minutes. The pelleted DNA was resuspended by adding 100 µl of Tris -Cl EDTA (TE) and incubating at 37°C overnigh t. Concentrations of DNA were quantified by spectrophotometry. Samples were diluted to a concentration of 10 ng/µl and stored at -70°C for future use.

Genotypes were determined by PCR at both the PPAR γ and Amhr2 loci. The following primers were used to amplify the wild type, floxed, and null alleles for PPAR γ: F, 5’ -ctc caa tgt tct caa act tac -3’, Rev1, 5’ -gat gag tca tgt aag ttg acc -3’, Rev2, 5’ -gta ttc tat ggc ttc cag tgc -3’ [7] . PCR reactions consisted of 2.5 mM MgCl 2, 1X PCR buffer, 10 µM of each primer, 10 mM dNTPs, 10X bovine serum albumin, 1 U Taq polymerase (Bioline, Boston), and 50 ng DNA in 25 µl reactions. For amplification, 35 cycles were used at 94°C for 30 seconds, 60°C for 30 seconds, and 72°C for 90 seconds, with an initial denaturation at 94°C for 2 minutes and a final extension for 5 minutes at 72°C. Insertion of cre recombinase into the gene encoding Amhr2 was determined by amplification of genomic DNA with the primers: cre1, 5’ -gcg gtc tgg cag taa aaa cta tc -3’, cre2, 5’gtg aaa cag cat tgc tgt cac tt -3’ (sequences from Jackson Laboratories, Bar Harbor, ME). Amplified PCR products were separated by electrophoresis through a 2% agarose gel. PCR products representing wild -type alleles of PPAR γ p roduced a 225 bp fragment, flox -flanked PPAR γ alleles a 275 bp fragment, and the recombined, null PPAR γ allele produced a 400 bp fragment. The primers for cre recombinase produced a 100 bp fragment, and no amplified product when only Amhr2 wild -type allel es were present. Interleukin -2 (Il -2) was amplified as an internal control (Il -2F, 5’ -cta ggc cac aga att gaa afa tct -3’, Il -2R, 5’ -gta ggt gga aat tct agc atc atc c - 3’, 324 bp, Jackson Laboratories). An internal control was used to verify success of the assay. If no band was produced for the Amhr2 allele or internal control, this indicated failure of the PCR reaction. However, if a fragment corresponding to the internal control was 63

generated, but not one for Amhr2, the reaction was successful, but the mice did not have cre recombinase inserted into the Amhr2 allele.

To validate that the expression of PPAR γ was reduced in the ovary of the PPAR Amhr2 mice, PCR was performed to compare levels of the floxed and null alleles of PPAR γ. Animals homozygous f or the floxed PPAR allele and hetero - or homozygous for cre recombinase (PPAR γfl/fl , Amhr2 cre /-) had the ability to produce both non -functional, recombined null PPAR , as well as functional, floxed PPAR alleles. Animals that did not express cre recombin ase , PPAR γfl/ fl , Amhr2 wt /wt only produced the functional floxed PPAR γ allel e. Microarray Analysis. Microarray analysis was used to detect candidate genes

regulated by PPAR . Mice homozygous for the floxed PPAR γ allele and hetero - or homogygous for cre recombinase (PPAR γfl/fl , Amhr2 cre /-; n=3 ), and mice without cre recombinase (PPAR γfl/fl , Amhr2 wt/wt ; n=3 ) were injected with 5 IU of PMSG on day 23 of age to stimulate follicular development. Ovaries were collected 48 hours later and placed in a sucrose/E DTA media (25 mM HEPES, DMEM: F12, 0.01% sodium pyruvate, 0.22% sodium bicarbonate, 0.125% gentamicin, insulin -transferrin -sodium selenite media

supplement, 0.5 M sucrose, and 10 mM EDTA; pH= 7.2) at 37°C in 5% CO 2: 95% air for 20 minutes. Ovaries were th en rinsed and placed in defined media (25 mM HEPES, DMEM: F12, 0.01% sodium pyruvate, 0.22% sodium bicarbonate, 1% bovine serum albumin, 0.125% gentamicin, and insulin -transferrin -sodium selenite media supplement; pH= 7.2). Twenty -six gauge needles were u sed to repeatedly puncture the ovaries, releasing the granulosa cells. The m edia were collected and the granulosa cells collected by centrifugation at 700 g for 4 minutes. DNA and RNA were extracted from granulosa cells using Trizol reagent, following ma nufacturer’s instructions. PCR was performed to detect the level of expression of the PPAR alleles . RNA quality and subsequent microarry analysis w as performed at the Iowa State University Gene Chip Facility. Briefly, the RNA quality was assessed using an 64

Agilient 2100 bioanalyzer. After RNA analysis, the RNA was used to produce cDNA. The cDNA was amplified and biotinylated. The biotin -cDNA was used to hybridize to Affymetrix mouse 430 2. 0 arrays (1 array per mouse, n= 6 arrays). This array contains 45,000 probe sets to analyze the expression level of 40,000 genes. After hybridizati on, the arrays were washed and scanned to determine the level of gene expression for the 40,000 genes. Statistical Analysis. Data were analyzed in collaboration with Dr. Peng Liu at the Iowa State University Statistics Department. Original microarray d ata contained genes that were absent, present and marginal. Absent referred to genes that did not hybridize at all, present referred to genes that hybridized and marginal referred to genes that had some weak hybridization. When a gene was absent from all six arrays, the gene was removed from the database . This narrowed down the list of genes for statistical analysis. The microarray data were then analyzed by a 2 -sample student t -test. A p -value of ≤ 0.01 was considered significant. Development of an in vitro model to study the role of PPAR in the ovary

Adenoviral technology was utilized to reduce the expression of PPAR γ in cultured granulosa cells. An adenovirus will infect dividing cells and is a powerful tool to introduce a gene(s) into cells. The adenoviruses used to infect the granulosa cells contain the gene for green fluorescent protein (GFP; control adenovirus : Ad -CMV -eGFP ), or cre recombinase in addition to GFP (cre adenovirus : Ad5 -CMVCr e-GFP ). Granulosa cells from PPAR fl/fl mice were placed in culture with either the control adenovirus, or the cre adenovirus. GFP allowed for visualization of cells that were successfully infected with the adenoviruses.

Optimization of transfection efficiency Cell culture. Collection of granulosa cells from PPAR γfl/fl , Amhr2 wt/wt mice was performed as described above. The cell pellet was resuspended in 2 ml of media (25 mM HEPES, DMEM: F12, 0.01% sodium pyruvate, 0.22% sodium bicarbonate, 0.125% 65

gentamicin, insulin -transferrin -sodium selenite media supplement and 1% fetal bovine serum; pH= 7.2) and the number of granulosa cells counted with a hemocytometer. Cells

were plated in media containing 1% fetal bovine serum at 37°C in 5% CO 2: 95% air for 24 hours, to allow the cells to attac h to the wells . All experiments performed to optimize transfection efficiency were performed after the initial 24 hours of culture. Experiments were performed to identify media type, adenovirus concentration, plating density, and incubation time that wo uld support efficient and effective infection of granulosa cells. The effect of fetal bovine serum was tested because the manufacturer of the adenovirus (Baylor Vector Development Laboratory, Houston TX) suggested using media with fetal bovine serum. How ever, fetal bovine serum has been reported to interfere with adenoviral transfection efficiency (personal communication , Dr. Boerboom). Therefore, the control (Ad -CMV -eGFP) or cre (Ad5 -CMVCre -GFP) adenovirus was added to the cells in media with or without fetal bovine serum . The following concentrations of each adenovirus were tested: 5, 15, 50, 100, 500, 1000, 2000, 2500, 3000 particles/cell . Plating density was studied to optimize the growth of granulosa cells. Cells were plated at 100,000 or 200,000 cells/well. The concentrations of adenovirus and number of cells plated were chosen based on personal communication with Dr. Boerboom and the manufacturer of the adenovirus.

Cells were cultured with the adenovirus for 24 or 48 hours at 37°C in 5% CO 2: 95 % air. At the defined experimental end point, 24 or 48 hours of culture, granulosa cells were analyzed under a Nikon phase contrast microscope to observe GFP expression to determine the percentage of cells that were infected by the adenovirus. Granulosa cells were subsequently collected and stored at -20°C until use d for reverse transcription PCR (RT - PCR) to determine level s of mRNA for PPAR . RT -PCR. Total RNA was extracted from cultured granulosa cells using Trizol reagent, following the manufacturer’s instructions. Complementary DNA was synthesized using SuperScript II Reverse Transcriptase and oligo (dT) primers 66

(Invitrogen). The cDNA obtained was analyzed by PCR to determine the relative levels

of PPAR and S16 (internal control) in cultured granulosa cells. Primers used to amplify S16 were published previously [22] . PPAR primers were as follows: PPAR forward, 5’ - gat gaa taa aga tgg agt cct cat ctc - 3’, PPAR reverse, 5’ - ccg cag gct ttt gag gaa -3’. The

PCR products generated from the PPAR primers were sequenced at the Iowa State DNA facility to verify amplification of the desired gene.

RT -PCR reactions (25µl) for PPAR and S16 consisted of 1.25 mM MgCl 2, 1X PCR buffer (200 mM Tris -HCl, 500 mM KCl), 1 µM of forward and reverse primers for both genes, 200 µM dNTPs, 2.0 unit of Taq polymerase, 10x bovine serum albumin (BS A), and

100 ng of cDNA for PPAR and 10 ng of cDNA for S16. All amplifications were carried out for 35 cycles with denaturation at 95°C for 1 minute, annealing at 52°C for 1 minute, and extension at 72°C for 1 minute per cycle. The concentrations of cDNA used for RT -PCR reactions were within the linear range of the assay. The linear range was determined by assessing the amount of product produced under the conditions described above when increasing concentrations of cDNA were used in the reaction. Plott ing the output versus input indicated the concentration range of input cDNA that yielded a corresponding increase in output. RESULTS To explore the function of PPAR γ in the ovary, an in vivo model was developed with the expression of PPAR γ specifically reduced in granulosa cells. Mice with loxP sequences

flanking exon 2 of PPAR γ (PPAR γfl/fl , referred to as the flox line) were mated to mice containing cre recombinase (referred to as the cre line). This mating resulted in the crox line of mice. Due to the recombination activity of cre recombinase in the ovaries of crox mice

(PPAR γfl/fl , Amhr2 cre/ -) a shortened, non -functional, or null product , of the PPAR γ gene may be produced, as well as and a functional ( flox ) band. The degree to which the expression of

PPAR γ was reduced in the ovary of crox females was determined by comparing the amount 67

of ovarian genomic DNA encoding the floxed alle le for PPAR γ to that encoding the PPAR γ null -allele (Figure 4. 1) . The flox animals do not express cre recombinase (PPAR fl/fl ,

Amhr2 wt/wt ) and therefore, produce only a functional , floxed PPAR γ allele (Figure 4. 1). All crox mice had approximately a 50% reduc tion in the expression of PPAR . In order to investigate what ovarian genes are regulated by PPAR , a microarray experiment was performed to determine those genes differentially expressed in granulosa cells with PPAR unaltered compared to granulosa c ells with reduced expression of PPAR . Granulosa cells were collected from six animals (Figure 4.2; n=3: PPAR fl/fl , Amhr2 cre/wt , n=3: PPAR fl/fl , Amhr2 wt/wt ) and RNA was extracted and analyzed by microarray. Initial analysis of the microarray data found the genes in Table 1 to be significantly up -regulated by PPAR , and the genes in Table 2 to be significantly down -regulated by PPAR . An in vitro model of reduced expression of PPAR in the ovary will also allow for investigating the roles of PPAR in ov arian biology. Granulosa cells from flox mice were

cultured with an adenovirus containing cre recombinase to reduce the expression of PPAR γ in vitro . Experiments were conducted to establish conditions for optimal transfection of the

granulosa cells, asse ssed by reduced expression of mRNA for PPAR γ. The absence of fetal bovine serum in the culture media reduced the level of mRNA for PPAR as determined by PCR. When four different concentrations of adenovirus were added to the granulosa cells in cultur e with and without fetal bovine serum, only one concentration of adenovirus decreased levels of mRNA for PPAR . However, when fetal bovine serum was removed from the media three of the four concentrations of adenovirus

were able to decrease levels of mRNA for PPAR . Therefore, media without fetal bovine serum were used for the remaining experiments. Experiments were conducted to determine the amount of adenovirus that would result in the most efficient transfection of granulosa cells. The most consiste nt reduction of mRNA for PPAR was found when cells were cultured with either 1000 or 2500 viral particles per 68

cell (pts/cell). Treatment with 1000 and 2500 virual particles per cell resulted in approximately a 25% to 50% reduction in levels of mRNA for P PAR as compared to the corresponding control. While treatment with the other concentrations (5, 15, 50, 100, 500, 3000 particles/cell) of adenovirus resulted in either no change in the levels of mRNA for

PPAR , an increase in levels of mRNA for PPAR or approximately a 10% decrease in levels of mRNA for PPAR . Therefore, the remaining experiments were performed using these two concentrations of adenovirus. Plating density was investigated to optimize growth of granulosa cells. The granulosa cells were plated at two different concentrations: 100,000 and 200,000 granulosa cells per well . These concentrations of cells per well were chosen based on suggestions from the manufacturers of the adenovirus and from personal communication with Dr. Boerboom who ha d previously used this adenovirus with granulosa cells in culture. Using 1.0 x 10 5 cells per well , n either adenovirus concentration was able to decrease expression of PPAR γ (Figure 4.3 a). As seen in Figure 4.3 , a cell density of 200,000 cells/well resulted in both concentrations of adenovirus decreasing the expression of mRNA for PPAR (Figure 4.3 b). Overall, t he best transfection efficiency and most consistent reduction of mRNA for PPAR were observed using 2.0 x 10 5 cells per well and 1000 adenoviral particles per cell. Finally, the length of time the cells were incubated with the adenovirus was

investigated . The greatest reduction of mRNA for PPA Rγ was found after the initial 24 hours of culture with the adenovirus (Figure 4.4 ). Incubation with the adenovirus for 48 hours resulted in either no change ( 1000 pts/cell) or an increase in mRNA for PPAR γ (2500 pts/cell; Figure 4.4 ). DISCUSSION PPAR γ is present in ovarian tissue. Using cre/loxP technology, Cui et al. (2002) produced a mouse line with PPAR γ disrupted in mammary epithelium, B - and T -cells, and in the ovary. This study reported that females with reduced expression of PPAR γ were either 69

sub - or infertile [7] . Because the expression of PPAR was not disrupted in the uterus of these transgenic animals, the authors concluded that the lesion causing the sub - and infertility resided in the ovary. Becaus e progesterone and estrogen production from the ovary are needed for implantation, it is possible that the sub - and infertility in these mice resulted from ovarian dysfunction. Study of gene function has been greatly advanced by the development of knock -out animals lacking a specific gene of interest. An animal model in which the ex pression of

PPAR γ is knocked -out would facilitate investigating the role of this transcription factor in ovarian biology. Unfortunately, deletion of PPAR γ results in embryonic death [23] . Investigati ng the cause of the leth ality determined that PPAR γ is required for placental development . Therefore, the current experiments were undertaken to d evelop models of reduced expression of PPAR γ in the ovary to investigate its role (s) in the ovary . Cre/loxP technology was utilized to create models in which PPAR γ was selectively reduced in the ovary. This was possible due to the creation of an Amhr2 -cre mouse [24] . Amhr2 is only expressed in granulosa cells. PPAR is also primarily localized to granulosa cells. Insertion of cre recombinase into the Amhr2 gene (Amhr2 -cre mouse) allows for cre recombinase to only be expressed in granulosa cells. Mice with loxP insertions flanking PPAR were previously generated an d the mating of these two mouse lines enable granulosa - cell -specific reduction of PPAR γ. One strategy to determine the function of PPAR γ in ovarian biology is to identify potential genes that are regulated by PPAR γ. The creation of the in vivo model des cribed above allowed for the detection of these candidate genes. Initial analysis of the microarry data obtained from the in vivo model indicated genes possibly regulated by PPAR γ in granulosa cells. Further statistical analysis will be performed to inve stigate false discovery rates (FDR), s ince it is possible that some of the significant genes were false positive and that some of the non -significant genes were false negatives . The FDR will be set at 10% to 70

determine which genes were not false positives or negatives . With the knowledge of these genes, future studies can verify if these genes are truly regulated by PPAR γ in the ovary and how this regulation influences ovarian function. The in vivo and in vitro models created may allow investigators to resolve conflicting data on the roles of PPAR γ in processes such as steroidoge nesis, angiogenesis, and inflammation. For example, Komar and Curry (2003) found there to be no correlation between mRNA expression for PPAR γ and aromatase, the enzyme responsible for the conversion of androgens into estradiol, either before or after the LH surge in Spraque - Dawley rats [25] . Using a different strain of rat, Lovekamp -Swan et al . (2003) found that PPAR decreased estradiol production by suppressing aromatase activity [13] . The same conclusion was reached using human granulosa -lutein cells [10] . These studies indicate that

PPAR is involved in steroidogenesis . However , its exact role has not been determined. Using the in vitro model described herein, it is now possible to manipulate granulosa cells with and without PPAR present and measure hormo nal responses. The role of PPAR in fertility can also be explored with the new in vivo model. Since PPAR is selectively reduced in the ovary, experiments can be performed to look at time to pregnancy, pregnancy length, litter size and more. This in v ivo model should allow future researchers to focus on how PPAR is involved in fertility. Many studies have localized PPAR γ in ovarian tissue but the genes that are regulated by PPAR γ are still unknown. Using the models de scribed above, researchers can be gin to explore which genes are regulated in the ovary by PPAR γ and which processes in ovarian function require PPAR . Using the in vivo model describe d in this manuscript the candidate genes that PPAR may regulate have been identified. Future research c an focus on validating if these genes are regulated by PPAR in the ovary and how this regulation impacts ovarian biology. 71

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Gamma in Ovarian Folliculogenesis in the Sheep. Biology of Reproduction 2003; 69: 1665 -1674. 7. Cui Y, Miyoshi K, Claudio E, Siebenlist UK, Gonzalez FJ, Flaws J, Wagner KU, Hennighausen L. Loss of the Peroxisome Proliferation -Activated Receptor Gamma (PPAR ) Does Not Affect Mammary Development and Propensity for Tumor Formation but Leads to Reduced Fertility. Journal of.Biological Chemistry 2002; 277: 17830 -17835. 72

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PCR Product Length 400 bp

275 bp

PPAR γ fl/fl , PPAR γ fl/fl , Amhr2 wt/wt Amhr2 cre/wt

Animal Genotype

Figure 4. 1. Amplification of PPAR γ from genom ic DNA extracted from ovarian tissue. Tissues were collected and analyzed by PCR as described in the Methods and

Materials . Amplification of the floxed PPAR γ allele (fl) produced a 275 bp DNA fragment (flox). Due to the recombination activity of cre rec ombinase (cre), a non - functional (null) product of PPAR γ (400 bp) was produced, as well as a 275 bp functional (flox ) band. 76

PCR Product Length

400 bp

275 bp

PPAR fl/fl , Amhr2 cre/wt PPAR fl/fl , Amhr2 wt/wt Animal Genotype

Figure 4.2 . Amplification of PPAR from genomic DNA extracted from ovarian tissue from PPAR fl/fl , Amhr2 wt/wt (n=3) and PPAR fl/fl , Amhr2 cre/wt (n=3) mice. Tissues were collected and analyzed by PCR as described in the Methods and Materials . Amplification of the floxed PPAR allele produced a 275 bp fragment (flox). Due to the recombination activity of cre recombinase (cre) in the ovaries of

the mice, a non -functional (null) product of PPAR γ (400 bp) was produced, as well as a 275 bp functional ( flox ) band . 77

a

r 0.75 o f A N a R m m 0.5 f m o a s g l e R v A e l P 0.25 P e v i t a l e

R 0 c 1000 cre 1000 c2500 cre 2500 Adenovirus concentration (particles/cell)

b

r 1.5 o f A N R a m m f m o a s g

l 0.75 e R v A e l P P e v i t a l e

R 0 c 1000 cre 1000 c 2500 cre 2500 Adenovirus concentration (particles/cell)

Figure 4.3. Effect of cell density and adenovirus concentration (particles per cell) on the level s of mRNA for PPAR γ. Relative levels of mRNA for PPAR γ with 100,000 ( a), or 200,000 ( b) granulosa cells per well in the presence of 1000 or 2500 par ticles/cell of adenovirus cultured for 24 hours as described in the Methods and Materials . c= control adenovirus, cre = cre adenovirus. 78

A 2.5 N R a m m f m o a s g

l 24 hours e R 1.25 v

A 48 hours e l P P e r v i t o f a l

e 0 R c 1000 cre 1000 c 2500 cre 2500 Adenovirus concentration (particles/cell)

Figure 4.4. The effect of length of culture , 24 or 48 hours , on levels of mRNA for

PPAR γ. Relative levels of mRNA for PPAR γ with 200,000 granulosa cells /well cultured with 1000 or 2500 particles/cell of adenovirus, over 2 4 or 48 hours, as described in the Methods and Materials . c= control adenovirus, cre= cre adenovirus. 79

TABLE 1. Ovarian genes up -regulated by PPAR

N-Acetyltransferase 2 Interleukin Enhanc er Binding Factor

Guanylate Cyclase Activator 1A Tripartite Motif Protein 8

Synaptotagmin -like Protein 3 -b Neuropilin -2

Growth Hormone Inducible Serine Threonine Kinase Receptor Transmembrane Protein Associated Protein

Cartilage Derived Retinoic Acid S ensitive SH2 -B PH Domain Containing Signal Protein Mediator 1

Ras -GTPase -Activating Protein Cyclic Nucleotide -Gated Channel Beta

Truncated Class I MHC Receptor Furmin BP -1

Calmodulin 2 F-box and WD -40 Domain Protein 13

Class 1 Major Histocompatibility Antigen Seryl -Aminoacyl -tRNA Synthetase 2

Bladder Cancer Associated Protein Putative Mannosyltransferase

Nuclear Factor of Activated T -Cells Acetyl CoA Transferase -like Protein

Guanosine Diphosphate Dissociation Huntingtin Associated Protein Inhibitor 3

Paraoxonase 2 Amyloid Beta Precursor -like Protein 80

TABLE 2. Ovarian genes down -regulated by PPAR γ

F-box only Protein 32 Mercaptopyruvate Sulfurtransferase

Diazepam Binding Inhibitor Procollagen Type V, alpha -3

Protein Tyrosine Phosphatase, Receptor Tumor Associated Calcium Signal Type K Transdurcer 1

ATPase, Na+ K+ Transporting B eta 1 Calcium Calmodulin -Dependent Protein Polypeptide Kinase 1

Calcineurin Inhibitory Protein ZAK1 -4 Granzyme A

J Domain Protein 1 Guanylate Nucleotide Binding Protein 3

Procollagen, Type IX, Alpha 3 Phosphatidylinositol 3 -Kinase

T-Cell Lymphomia Inva sion and Protein Tyrosine Phosphatase Receptor Metastasis 1 Type E

Metal Response Element Binding Apoptosis Inhibitor 1 Transcription Factor 1

Adenosine Kinase Serum Inducible Kinase

Protein Inhibitor of Activated STAT 1 ATP -Binding Cassette, sub -family C

T-cell Receptor Beta -Chain Protocadherin 8

Kruppel -like Factor 9 81

CHAPTER 5. EFFECT OF GENISTEIN ON ESTRADIOL PRODUCTION BY BOVINE FOLLICLES

Submitted for Publication in Reproductive Toxicology NS Tinfo, LJ Luense, SM Hopkins, and CM Komar

ABS TRACT Genistein, a phytoestrogen present in food commonly found in human and animal diets, can affect various aspects of ovarian biology. We investigated the effect of genistein on follicular estradiol production. Ovaries were removed from heifers (n=3) w hen the largest follicle measured approximately 8 mm. The largest and second largest follicles were dissected from the ovary, and pieces of follicle wall placed in culture with vehicle, FSH (1 ng/ml), genistein (10 µM), FSH + genistein, or vehicle or genis tein (10 µM), respectively. There was no effect of treatment on estradiol production by cultured follicular tissue from the largest or second largest follicle. However, there was a decrease in levels of mRNA for aromatase in tissue from the largest folli cle treated with genistein + FSH, compared to that treated with FSH alone (p ≤0.05). These data suggest that genistein may not affect the basal production of estradiol, but may alter response to gonadotropin stimulation. INTRODUCTION Phytoestrogens are bio logically active plant substances with a chemical structure similar to that of estradiol [1] . There are three main classes of phytoestrogens: isoflavones, coumestans, and lignans [2] . Isoflavones and coumestans are contained in legumes, which are common components of human and animal diets. Soybean is an abundant source of isoflavones [3] , and o ne of the most abundant isoflavones in soybean is genistein [4] . Animals, such as cattle and sheep, are exposed to soy phytoestrogens via grazing and silage [5] . At least 25% of the infant formula marketed in the 82

United States is comprised of soy based products [6] . Manufacturers are adding soy protein to food as a result of the FDA -supported claim concerning their cardiovascular protec tion. Recent health trends have also spurred the consumption of soy protein in adult diets with increased intake of foods such as tofu and soy milk. The use of genistein as a nutraceutical has also exploded in recent years. Supplements containing genist ein are being studied for treating ailments such as cancer [7] , menopausal sympto ms, and osteoporosis (reported in [8] ). Within the plant, genistein is an inact ive glucoside. Once genistein is ingested it is converted into a heterocyclic phenol [2] . This heterocyclic phenol is structurally similar to estrogen and capable of inducing estrogenic and/or anti -estrogenic effects [2, 9] . In cattle, the specific heterocyclic phenol generated from genistein, p -ethyl -phenol, has been detected in the blood for many ho urs after consumption of feed containing soybean [10] . Genistein may affect ovarian biology by modulating the activity of second messenger systems. Specifically, genistein inhibits tyrosine kinases, important intracellular second messengers involved in steroidogenesis. Inhibi tion of tyrosine kinases could lead to a decrease in ovarian hormone production [11] . Along with the ability of genistein to impact hormone production, it may also affect the activity of estradiol. Production of estrogen by granulosa cells is required fo r the LH surge and ovulation. Estrogen receptor β (ER β) has been identified in several ovarian cell types, primarily granulosa cells [12 -14] . Genistein binds with a greate r affinity to ER β than ER α [15] . Genistein competes with endogenous estrogen for binding to ER β. Genistein has both estrogenic and anti -estrogenic a ctivity [2, 4] . The ef fect of genistein being either estrogenic or anti -estrogenic , depends on the tissue type, ER, and concentration of circulating endogenous estrogens [16, 17] . Genistein could potentially regulate ovarian gene expression not only by interacting with ERs, but also through binding to peroxisome proliferator -activated receptor γ (PPAR γ) 83

[18] . Like ERs, PPAR γ is a member of the steroid recepto r superfamily, and is expressed in ovarian tissue of several species, including the mouse, rat, cow, sheep, pig, and human [19] .

Genistein binds and activates PPAR γ [18, 20] . Dang et al. (2003) found that when genistein activates PPAR , there is a decrease in the activity of estrogen . This effect of genistein is dependent on the concentration of genistein, and the levels of ER and PPAR within the tissue [20] . The decrease in estrogenic activity could be due to the fact that PPAR s compete with endogenous estrogens for binding to estrogen response elements [21, 22] . Activation of PPAR can also influence various processes such as angiogenesis, inflammation, and steroidogenesis (reviewed in [14]). These processes are critical for normal ovarian function.

Therefore, activation of PPAR γ by genis tein may also affect reproduction by regulating several processes involved in normal ovarian function. Since humans and animals are exposed to genistein via the diet, and because this phytoestrogen may affect ovarian function, the following experiments w ere performed to investigate how genistein may influence follicular estradiol production. We hypothesized that exposure to genistein would decrease follicular production of estradiol. METHODS AND MATERIAL S Animals

All animal procedures were approved by th e Iowa State University’s Animal Care and Use Committee. Fifteen to eighteen month old, cyclic Angus heifers (n=3) were monitored by transrectal ultrasonography to follow follicular development, ovulation, and growth and regression of the corpus luteum (C L). Two perpendicular diameters of ovarian structures larger than 5 mm were recorded. Diagrams were made of the relative positions of follicles and CL to monitor follicular growth and luteal formation/regression. Day 1 of the cycle was defined as the fi rst day the ovulatory -sized follicle was no longer observed. To synchronize the day of ovulation, two heifers received 25 mg of Lutalyse (Pharmacia, 84

Peapack, NJ) to regress the CL, followed thirty -six hours later by an injection of a GnRH analog (Cystorel in; 100 µg) to induce an LH surge. The remaining heifer cycled naturally. Before removal of the ovaries by flank incision, the heifers received an “L” block with 2% lidocaine. Ovaries were removed from all heifers when a follicle from the first follicu lar wave post -ovulation measured approximately 8 mm. This size was chosen because it is at this relative developmental stage when deviation occurs between dominant and subordinate follicles [23 ]. After removal, ovaries were immediately placed in ice -cold

transfer medium [minimum essential medium (Gibco, Carlsbad, CA), 2.2% NaHCO 3, penicillin (50 U/ml), streptomycin (50 µg/ml)] and transported to the laboratory. The t wo largest follicles were dissected from ovaries within 30 minutes of removal from the heifer. Follicular fluid was collected and frozen at -20°C until analyzed for estradiol by RIA. Follicular tissue was cultured in vitro as described below. Tissue Cu lture

The largest and second largest follicles were hemisected and cut into pieces. Tissues (n=three pieces per well) were cultured in 500 µl of media [minimum essential medium,

2.2% NaHCO 3, 25 mM HEPES, penicillin (50 U/ml), streptomycin (50 µg/ml), tran sferrin (5 µg/ml), cortisol (40 ng/ml), and insulin (1 µg/ml)], with the following treatments: largest follicle – vehicle (methanol; control), FSH (1 ng/ml), genistein (10 µM), or FSH + genistein; second largest follicle – vehicle or genistein (10 µM). Th e concentration of FSH used in this study has been found to support steroid production typical of the follicular phase in vivo [24 ]. The dose of genistein was chosen because it had been previously found to elicit a

physiological effect in ovarian cells [2 5]. Tissues were incubated at 37°C in a 5% CO 2/95% air environment for 24 hours. After culture, media were collected and stored at -20°C until analyzed for estradiol by RIA. Tissues were weighed and frozen at -70°C until future analysis. 85

Radioimmunoassay

Concentrations of estradiol in follicular fluid and media were determined by RIA (DPC, Los Angeles, CA), according to the manufacturer’s directions. The concentrations of estradiol in media were corrected by the weight of the cultured follicular tissue in each well. The inter - and intra -assay coefficients of variation were 4.49% and 1.57%, respectively. Reverse Transcription -PCR

Total RNA was extracted from cultured tissues using Trizol reagen t (Invitrogen), and

quantified with a spectrophotometer. Complementary DNA was synthesized in a 20 µl reaction using SuperScript II Reverse Transcriptase and Oligo (dT) primers (Invitrogen). The cDNA obtained was analyzed by RT -PCR to determine levels of mRNA for aromatase in follicular tissue. The ribosomal protein, 18S, and GAPDH were used as internal controls. Primers utilized for aromatase [2 6], 18S [2 7], and GAPDH [2 8] were published previously. RT -PCR reactions (25 µl) for aromatase , 18S, and GAPDH, consisted of 1.0 mM

MgCl 2, 1X PCR buffer (190 mM NH 4SO 4, 670 mM Tris, pH 8.8; 0.1% Tween 20), 200 µM dNTPs, and 2.5 unit of Taq polymerase. Concentrations of fo rward and reverse primers, and cDNA used for PCR reactions were as follows; aromatase: 1 µM of primers and 50 or 500 ng cDNA, 18S: 0.025 µM primers and 75 ng cDNA, and GAPDH: 0.065 µM primers and 10 ng cDNA. All amplifications were carried out for 35 cycl es with denaturation at 95°C for 1 minute, annealing at 60°C for 1 minute, and extension at 72°C for 1 minute per cycle, and another elongation at 72° C for 5 minutes. Samples were analyzed in duplicate and the analysis repeated three times. RT -PCR produ cts were separated on a 2% agarose gel. Densitometric analysis of the resulting bands was conducted using the Spot Denso program (Fluorchem, Alpha Innotech). Relative levels of aromatase were determined per sample as a ratio with 18S or GAPDH. 86

Statis tical analysis

Data were tested for normalcy and, if needed, log transformed before analysis. Concentrations of estradiol were analyzed with one - way ANOVA with the SAS 9.1 statistical package (SAS Institute, Cary, NC). A Student’s T -test was used to ana lyze the effect of treatment on levels of mRNA for aromatase. A p -value of • 0.05 was considered significant. RESULTS The first wave of follicular development post -ovulation was recorded for each heifer via transrectal ultrasonography. On the day o f surgery, the follicles were measured after removal from the ovary. The ovaries were collected from the first wave of the cycle post - ovulation, the largest and second largest follicles were removed, and follicular fluid was collected. The concentration of estradiol in follicular fluid collected from the largest follicle from two of the three heifers was at least 8 times greater than that measured in follicular fluid from the second largest follicle (data not shown). However, estradiol concentrations in follicular fluid from the second largest follicle from the third heifer was undetectable (assay sensitivity = 0.8 pg /ml). Production of estradiol by tissue collected from the largest follicle from each heifer is shown in Figure 5. 1. There was a great d eal of variability in response to in vitro treatments between tissue collected from different animals. This variability could not be accounted for by pre -surgical hormonal regimen to synchronize ovulation. Estradiol secretion by tissue from the largest f ollicle collected from two heifers, #261 and #206, decreased in response to treatment with genistein or FSH (Fig. 5. 1 a & b). In contrast, the administration of FSH or genistein to tissue collected from the largest follicle from the remaining heifer resul ted in a doubling, or greater than four -fold increase in estradiol secretion, respectively (#305; Fig. 5. 1 c). Overall, there was no significant effect of any treatment on estradiol production by tissue from the largest follicle (p > 0.05; Fig. 5. 2). 87

Va riability in response to treatment with genistein was also observed in cultured tissue from the second largest follicle collected from different animals. There was an increase in estradiol production by follicular tissue from 2 of the 3 heifers (data not shown). However, tissue from the third heifer produced less estradiol in response to treatment with genistein (data not shown). When the estradiol values from all three heifers were combined, treatment with genistein did not significantly affect estradio l production ( p ≤ 0.05; Fig. 5. 3). After culture, RNA was isolated from follicular tissue, and levels of mRNA for aromatase measured by RT -PCR. Levels of mRNA for aromatase were significantly lower in tissue from the largest follicle after treatm ent with genistein and FSH, compared to levels in tissue treated with FSH alone (p ≤ 0.05; Fig. 5. 4). T reat ment with genistein alone had no effect on levels of mRNA for aromatase. DISCUSSION In the 1940s, there was an increase in the number of cases of both permanent and temporary infertility occurring in sheep grazing on Trifolium subterraneum , a cultivar of subterranean clover [5] . These cases of infertility stimulated interest in the estrogenic effects of plant phytoestrogens on the female reproductive system. There is incr easing evidence that phytoestrogens can affect fertility in both animals and humans (reviewed in [2] ). Therefore, this study was undertaken to investigate the effect of the phytoestrogen genistein, on the ability of ovarian follicles to produce estradiol, using the cow as a model.

The current study demonstrated that genistein, at 10 µM, did not affect the production of estradiol by the largest or second largest follicle from the first wave of follicles developing post -ovulation. However, levels of mRNA f or aromatase decreased when tissue from the largest follicle was cultured with genistein and FSH compared to treatment with FSH alone. Haynes -Johnson et al . (1999) previously reported that genistein inhibited FSH -stimulated estradiol production by rat gra nulosa cells [29] . Taken together, these findings suggest that 88

genistein interferes with gonadotropin -stimulated estradiol production, but not basal production. Genistein is a weak inhibitor of aromatase. In ovarian tissue from trout, genistein was found to inhibit aromatase activity, but w as not as potent an inhibitor as other phytoestrogens [30] . One reason genistein is a weak inhibitor of aromatase may be related to its structure [31, 32] . Genistein has a 4’ -hydroxyphenyl group at carbon 3, which greatly reduces the ability of genistein to bind to aromatase. Le Bail et al . (2000) found that the structur e of genistein lends itself to be a better 3 β –hydroxysteroid dehydrogenase and/or 17 β– hydroxysteroid d ehydrogenase inhibitor [32] . Therefore, the lack of an effect of treatment with gen istein alone on aromatase expression, as seen in the current study , could be due to the structure of genistein. The dose and/or length of exposure to genistein may also influence the ability of this phytoestrogen to impact cellular function. Experiments have shown that in human granulosa -luteal cells exposed to genistein for 48 hours, there was a decrease in mRNA for aromatase, aromatase activity [25] , and estradiol production [25, 33] . In the study performed by Rice et al . (2006), both 10 µM and 50 µM of genistein were able to decrease aromatase activity and estradiol production from human granulosa -luteal cells. However, only a 50 µM dose was able to decrease estradiol production in hu man granulosa -luteal cells in the study by

Whitehead et al . (2002). Furthermore, a 4 hour incubation of human granulosa -luteal cells

with 50 µM genistein, resulted in a decrease in estradiol production in the presence of estrone [33] . When human granulosa -lu teal cells were cultured for 24 hours, genistein (50 µM) significantly decreased estradiol production, but only in the presence of androstenedione and

estrone [33] . The dose of genistein used in the current experiment (10 µM) had been shown previously to decr ease activity and mRNA for aromatase over 48 hours of culture [25] . The lack of an effect of genistein on secretion of estradiol observed in the current experiment may 89

be due to the fact that follicular tissue was studied, not luteinizing cells, and that the tissues were cultured for a shorter period of time. The current data demonstrate that treatment with genistein, in combination with FSH, decreased mRNA for aromatase in follicular tissue from the largest follicle when compared to levels in tissue treated with FSH alone. However, genistein did not inhibit estradiol pro duction by this tissue. This apparent dichotomy between levels of mRNA and estradiol production may be due to a time delay between the reduction in mRNA for aromatase and the loss of aromatase activity, or greater stability of aromatase protein compared t o mRNA. Due to the observed decrease in levels of mRNA of aromatase, it is possible that exposure to a low dose of genistein over a short period may alter an important step in ovarian hormone production. Genistein may not affect the basal production of e stradiol, but may alter the response to gonadotropin stimulation. Further studies are needed to determine how genistein affects follicular, as well as oocyte development, and the time course of response after exposure and the time needed for recovery afte r withdrawal. ACKNOWLEDGEMENTS The authors would like to thank Dr. Robert Cushman, Dr. Philip Bridges, and Dr. Leo Timms for critically reading the manuscript and their insightful comments. We would also like to thank Dr. Steven Lonergan and Dr. Elizab eth Huff -Lonergan for use of the Alpha Tech Imager. REFERENCES 1. Usui T . Pharmaceutical prospects of phytoestrogens. Endocr J . 2006; 53 : 7-20. 2. Rosselli M, Reinhart K, Imthurn B et al. Cellular and biochemical mechanisms by which e nvironmental oestrogens influence reproductive function. Hum Reprod Update . 2000; 6: 332 -350. 3. You L . Phytoestrogen genistein and its pharmacological interactions with synthetic endocrine -active compounds. Curr Pharm Des .2004; 10 : 2749 -2757 , 2004 . 90

4. Ada ms NR . Detection of the effects of phytoestrogens on sheep and cattle. J Anim Sci . 1995; 73 :1509 -1515 . 5. Lundh T . Metabolism of estrogenic isoflavones in domestic animals. Exp Biol Med . 1995; 208 : 33 -39. 6. American Academy of Pediatrics: Committee on N utrition. Soy protein -based formulas: recommendations for use in infant feeding. Pediatrics . 1998 ; 101: 148 - 153. 7. Ravindranath MH, Muthugounder S, Presser N, Viswanathan S. Anticancer therapeutic potential of soy isoflavone, genistein . Advances in Experi mental Medicine and Biology . 2004; 546 :121 -165. 8. Green NS FT, Kelly JW. Genistein, a natural product from soy, is a potent inhibitor of transthyretin amyloidosis . Proceedings of the National Academy of Sciences . 2005; 102 :14545 -14550. 9. Dixon RA, Ferrei ra D . Genistein. Phytochemistry . 2002; 60 : 205 -211. 10 . Woclawek -Potocka I, Acosta TJ, Korzekwa A et al. Phytoestrogens modulate prostaglandin production in bovine endometrium . cell type specificity and intracellular mechanisms. Exp Biol Med . 2005; 230 : 32 6-333. 11 . Akiyama T, Ishida J, Nakagawa S et al. Genistein, a specific inhibitor of tyrosine -

specific protein kinases. J Biol Chem . 1987; 262 : 5592 -5595. 12 . Rosenfeld CS, Yuan X, Manikkam M et al. Cloning, sequencing, and localization of

bovine estrogen receptor -β within the ovarian follicle. Biol Reprod . 1999; 60: 691 - 697. 13 . D'Haeseleer M, Van Poucke M, Van den Broeck W . Cell -specific localization of estrogen receptor beta (ESR2) mRNA within various bovine ovarian cell types using in situ hybridization . Anat Histol Embryolo . 2005; 34: 265 -272. 91

14. Enmark E, Pelto -Huikko M, Grandien K, Lagercrantz S, Lagercrantz J, Fried G, Nordenskjold M, Gustafsson J -A. Human Estrogen Receptor {beta} -Gene Structure, Chromosomal Localization, and Expression Pattern. J Clin Endocrin Met. 1997; 82: 4258 -4265. 15. Barkhem T, Carlsson B, Nilsson Y et al. Differential response of estrogen receptor alpha and estrogen receptor beta to partial estrogen agonists/ antagonists. Mol Pharm acol. 1998; 54: 105 -112. 16. Brzezinski A, De bi A . Phytoestrogens: the "natural" selective estrogen receptor modulators? Eur J Obstet Gynecol Reprod Biol . 1999; 85 : 47 -51. 17. Ososki AL, Kennelly EJ: Phytoestrogens : a review of the present state of research. Phytother Res . 2003; 17: 845 -869. 18. Shen P, Liu MH, Ng TY et al. Differential effects of isoflavones, from astragalus

membranaceus and pueraria thomsonii, on the activation of PPAR α, PPAR , and adipocyte differentiation in vitro. J Nutr . 2006; 136: 899 -905. 19. Komar CM . Peroxisome proliferator -activated receptors (PPARs) and ovarian function -implications for regulating steroidogenesis, differentiation, and tissue remodeling. Reprod Biol Endocrinol . 2005; 3. 20 . Dang ZC, Audinot V, Papapoulos SE et al. Peroxisome proliferator -activated

receptor g amma (PPARgamma ) as a molecular target for the soy phytoestrogen genistein. J Biol Chem . 2003; 278 : 962 -967. 21 . Nunez SB, Medin JA, Braissant O et al. Retinoid X receptor and peroxisome proliferator -activated receptor activate an estrogen responsive gene independent of the estrogen receptor. Mol Cell Endocrinol . 1997; 127 : 27 -40. 22 . Keller H, Givel F, Perroud M et al. Signaling cross -talk between peroxisome proliferator -activated receptor/retinoid X receptor and estrogen receptor through estrogen respons e elements. Mol Endocrinol . 1995; 9 : 794 -804. 92

23 . Ginther OJ, Wiltbank MC, Fricke PM et al. Selection of the dominant follicle in cattle. Biol Reprod . 1996; 55 : 1187 -1194. 24 . Berndtson AK, Vincent SE, Fortune JE . Low and high concentrations of gonadotrop ins differentially regulate hormone production by theca interna and granulosa cells from bovine preovulatory follicles. Biol Reprod . 1995; 52 : 1334 - 1342. 25. Rice S, Mason HD, Whitehead SA . Phytoestrogens and their low dose combinations inhibit mRNA expres sion and activity of aromatase in human granulosa -luteal cells. J Steroid Biochem Mol Biol . 2006; 101 : 216 -225.

26. Okuda K, Uenoyama Y, Berisha B et al. Estradiol -17 β is produced in bovine corpus luteum. Biol Reprod . 2006; 6 5: 1634 -1639. 27. Tian XC, Be rndtson AK, Fortune JE . Changes in levels of messenger ribonucleic acid for cytochrome P450 side -chain cleavage and 3 beta -hydroxysteroid dehydrogenase during prostaglandin F2 alpha -induced luteolysis in cattle. Biol Reprod . 1994; 50 : 349 -356. 28. Tsai SJ, Wiltbank MC, Bodensteiner KJ . Distinct mechanisms regulate induction of messenger ribonucleic acid for prostaglandin (PG) G/H synthase -2, PGE (EP3) receptor, and PGF2 alpha receptor in bovine preovulatory follicles. Endocrinology .

1996; 137 : 3348 -3355. 29 . Haynes -Johnson D, Lai MT, Campen C et al. Diverse effects of tyrosine kinase inhibitors on follicle -stimulating hormone -stimulated estradiol and progesterone production from rat granulosa cells in serum -containing medium and serum -free medium containing epidermal growth factor. Biol Reprod . 1999; 61 : 147 -153. 30 . Pelissero C, Lenczowski MJP, Chinzi D et al. Effects of flavonoids on aromatase activity, an in vitro study. J Steroid Biochem Mol Biol . 1996; 57 : 215 -223. 93

31 . Kao Y -C, Zhou C, Sherman M et al. Molecular basis of the inhibition of human aromatase ( estrogen synthetase) by flavone and isoflavone phytoestrognes . A site - directed mutagenesis study. Environ Health Perspect .1998; 10 6: 85 -92. 32 . Le Bail JC, Champavier Y, Chulia AJ et al. Effects of phyt oestrogens on aromatase,

3β and 17 β-hydroxysteroid dehydrogenase activities and human breast cancer cells. Life Sci .2000; 66 : 1281 -1291. 33 . Whitehead SA, Cross JE, Burden C et al. Acute and chronic effects of genistein, tyrphostin and lavendustin A on ste roid synthesis in luteinized human granulosa cells. Hum Reprod . 2002; 17 : 589 -594. 94

a b # 261 # 206 ) l

m 274.5 30 26.5 / 300 g p 5 p 0 g 3 1 / x

m 150 15 ( 84.0 l l 10.2 9.3 ) 8.2 o 47.8 48.5 i d a r

t 0 s

0 E C FSH Gen FSH +

E C FSH Gen FSH + Gen Gen s

t Treatment r

a Treatment d i o l ( x 1 0 c # 305 ) l 237.8 m

/ 250 g p 2 0 1

x 125 ( l 48.2 o i

d 6.5 13.6 a r

t 0 s

E C FSH Gen FSH + Gen Treatment

Figure 5.1. Secretion of estradiol (pg/ml) by tissue collected from the largest follicle of the first wave of the c ycle (n= 3) from individual animals . Follicular tissue was cultured as described in the Methods and Materials with methanol ( control ), FSH (1 ng/ml), genistein

(10 µM) or FSH and genistein. C = control, Gen= genistein . 95

200 p 5 g / m 100 l ) E s t r

a 0 d i

o C FSH Gen FSH + Gen l (

x Treatment 1 0

Figu re 5.2. Secretion of estradiol (pg/ml) by tissue collected from the largest follicle (n= 3) of the first wave of the cycle . Data are presented as mean ± SEM. Follicular tissue was cultured as described in the Methods and Materials with the following trea tments: methanol ( control ), FSH (1 ng/ml), genistein (10 µM) or FSH and genistein. C= control, Gen = genistein . 96

26 p g 3 / m l

) 13 E s

t 0 r a

d C Gen i o l (

x Treatment 1 0

Figure 5.3. Secretion of estradiol (pg/ml) by tissue collected from the second largest follicle (n=3) from the first wave of the cycle . Data are presented as mean ± SEM. Follicular tissue was cultured as described in the Methods and Materials with the following treatments: methanol (control) or genistein (10 µM). C= control, Gen= genistein . 97

180 a

120 ab ab

b

60

R 0 e l a t C FSH Gen FSH+Gen a r t o i v m Treatment e a l t e a v s e

Fie gure 5.4. Relative l evel s of mRNA for aromatase in tissue from the largest follicle l s ( % o

of the first fof llicular wave after 24 hours of culture. Data are presented as mean ± SEM. c m o n R t

Follicular tisN sue was cultured as described in the Methods and Materials with the following r o A l ) f treatments: mo ethanol ( control ), FSH (1 ng/ml), genistein (10 µM) or FSH and genistein. C= r control (methanol), Gen= genistein . Bars with different superscripts are significantly different (p ≤ 0.05). 98

CHAPTER 6. GENERAL CONCLUSIONS

DISCUSSION AND CONCL USIONS In or der to improve fertility and reproductive success, it is important to understand how genes are regulated in the ovary. One of the most frequent causes of infertility in women is polycystic ovarian syndrome (PCOS). PCOS is characterized by formation of cy sts in the ovaries, chronic absence of ovulation, and clinic signs of hyperandrogenism. Treatment with insulin sensitizers increases fertility in these patients. One class of drugs, the thiazolidinediones (TZDs) , are insulin -sensitizing drugs that are co mmonly administered to patients with type II diabetes and PCOS (reviewed by [1] ). TZDs are also ligands for

PPAR γ. PPAR was first located in the rat ovary in 1996, and has since been found in the ovary of several species. To date, the role(s) of PPAR γ in the ovary has not been determined. Understanding the role(s) of PPAR in the ovary will allow researchers to determine how activating PPAR via these drugs , or other ligands , affects ovarian function and female fertility.

PPAR γ is involved in processes in the body such as steroidogenesis, angiogenesis , and inflammation , which are important for normal ovarian function. The roles of PPAR γ in thes e processes indicate possible ways that PPAR γ may influence ovarian biology. The projects described in this dissertation were conducted to investigate the role(s) of PPAR γ in the ovary. The first study investigated how PPAR influences progesterone produ ction/breakdown and if PPAR is involved in luteal demise in the rat. Experiments

were also performed that investigated the effects of a PPAR γ agonist , genistein, on estradiol production in the bovine. M odels of reduced expression of PPAR γ in granulosa c ells were also developed as a means of investigating genes regulated by PPAR γ in the ovary . 99

Collectively , these studies gave insight into the role(s) PPAR γ does and does not play in the ovary. PPAR has been localized in luteal tissue from several speci es. In the rat, luteal tissue from past ovulations contained greater expression of mRNA for PPAR than luteal tissue from the most recent ovulation [2] . This is in contrast to the bovine corpus luteum, where PPAR decreases between the early and mid -luteal phase [3] . Lohrke et al. (2002) investigated the role of PPAR in the bovine corpus luteum, and concluded that PPAR is involved in prog esterone production. However, the function(s) of PPAR in the rodent corpora lutea was not known. This dissertation describes the experiments that were the first to report that PPAR does not play a role in the production or catabolism of progesterone in rat corpora lutea. Further investigations into the role(s) of PPAR in rat luteal tissue also found that PPAR is not involved in luteal demise via macrophages. Interestingly, experiments did demonstrate that PPAR may play a role in cell survival via r egulation of the bcl -2 gene. Since a peroxisome proliferator response element has previously been located in the human bcl -2 gene [4] , it is possible that PPAR regulates the expression of bcl -2. Future studies are needed to determine how PPAR influences the expression of bcl -2 in the rodent corpora lutea.

Activation of PPAR has been found to have varying effects on estradiol producti on.

In human granulosa -luteal cells and gonadotropin -primed rat granulosa cells, PPAR was able to decrease the expression of mRNA for aromatase [5, 6] . However, other research ha s shown that there is no correlation between the expression of mRNAs for PPAR and aromatase in granulosa cells during folliculogenesis, or the periovulatory period [7] . The study described in this dissertation investigated if one agonist of PPAR , genistein, influenced estradiol production and /or aromatase expression from bovine follicles. These preliminary studies demonstrated that follicles exposed to FSH and genistein had significantly lower levels of mRNA for aromatase than those treate d with FSH alone . 100

Therefore, it was concluded that genistein does not affect the basal production of estradiol but may alter the response to gonadotropin -stimulation . Future studies are needed to look at longer incubation times with genistein and more do ses of genistein to determine the dose range and length of time it takes to minimally and maximally affect aromatase expression

and production of estradiol. Since activation of PPAR has been found to have varying effects on steroidogenesis between specie s, investigating the effect of genistein on the ability of bovine follicles to produce estradiol may not answer the question of how exposure to this phytoestrogen will affect other animals exposed to genistein. Therefore, since phytoestogen exposure also affects sheep, and PPAR is found in ovarian tissue from sheep, experiments could also be performed that investigate the effects of PPAR on steroid production from sheep ovarian cells. The production of two models of reduced expression of PPAR in th e ovary allows for research into the role(s) of PPAR γ in fertility. These models are essential to investigate

PPAR in ovarian function and fertility because a whole genome knockout of PPAR results in embryonic death [8] . The development of the in vivo model allow ed for the identification of candidate genes that may be regulated by PPAR . Identification of these genes was the first step in determining how PPAR affects ovarian function. Further experiments can be performe d to determine the genes that are regulated by PPAR during the reproductive cycle.

The in vitro model will also allow for investigation into how certain ligands, for example phytoestrogens, impact ovarian biology. The in vivo model also allows for studi es to be performed on the role(s) of PPAR in fertility. These models can be used by future researchers to further investigate the function of PPAR γ in ovarian biology . These studies were the first to demonstrate that in the naturally cycling rat, PPA R does not play a role in progesterone production or catabolis m. However, a possible role for PPAR in luteal cell survival, via activation of bcl -2, was found. Using the in vivo model described, t hese studies were also able to identify candidate ovaria n genes regulated by 101

PPAR . The development of both in vivo and in vitro models can be used to further explore the function(s) of PPAR in the ovary and how PPAR affects ovarian biology . REFERENCES 1. Froment P, Gizard F, Defever D, Staels B, Dupont J, Monget P. Peroxisome Proliferator -Activated Receptors in Reproductive Tissues: from Gametogenesis to Parturition. Journal of Endocrinology 2006; 189: 199 -209. 2. Komar CM, Curry TE, Jr. Localization and Expression of Messenger RNAs for the Peroxisome Proliferator -Activated Receptors in Ovarian Tissue from Naturally Cycling and Pseudopregnant Rats. Biology of Reproduction 2002; 66: 1531 -1539. 3. Lohrke B, Viergutz T, Shahi SK, Pohland R, Wollenhaupt K, Goldammer T, Walzel H, Kanitz W. Det ection and Functional Characterisation of the Transcription Factor Peroxisome Proliferator -Activated Receptor Gamma in Lutein Cells. Journal of Endocrinology 1998; 159: 429 -439. 4. Butts BD, Tran NL, Briehl MM. Identification of a Functional Peroxisome Pro liferator -Activated Receptor Response Element in the 3' Untranslated Region of the Human bcl -2 Gene. International Journal of Oncology 2004; 24: 1305 -1310. 5. Lovekamp -Swan T, Jetten Anton M, Davis Barbara J. Dual Activation of PPAR Alpha and PPAR Gamma by Mono -(2 -ethylhexyl) Phthalate in Rat Ovarian

Granulosa Cells. Molecular and Cellular Endocrinology 2003; 201: 133 -141. 6. Mu YM, Yanase T, Nishi Y, Waseda N, Oda T, Tanaka A, Takayanagi R, Nawata H. Insulin Sensitizer, Troglitazone, Directly Inhibits Arom atase Activity in Human Ovarian Granulosa Cells. Biochem.Biophys.Res.Commun. 2000; 271: 710 -713. 7. Komar Carolyn M, Curry Thomas E, Jr. Inverse Relationship Between the Expression of Messanger Ribonucleic Acid for Peroxisome Proliferator -Activated Recepto r gamma and P450 Side Chain Cleavage in the Rat Ovary. Biology of Reproduction 2003; 69: 549 -555. 102

8. Barak Y, Nelson MC, Ong ES, Jones YZ, Ruiz -Lozano P, Chien KR, Koder A, Evans RM. PPAR Is Required for Placental, Cardiac, and Adipose Tissue Development. Molecular Cell 1999; 4: 585 -595. 103

ACKNOWLEDGEMENTS

I would like to thank my major professo r Dr. Carolyn M. Komar, for the opportunity to study under her guidance. Her patience, experience and generosity have been instrumental in my professional devel opment. I would also like to extend my gratitude to my committee members Dr. Leo Timms, Dr. Steven Hopkins, Dr. Richard Robson and Dr. Jill Pruetz for their kind advice, and constructive criticism throughout my degree. It is also imperative to thank Dr. Howard Tyler , as he was an unforgetable mentor and friend who always had his door open for countless conversations and words of encouragement . Most importantly I would like to thank my family, my mother and father Stephanie and Eugene Tinfo, my sister Jes sica, and grandparents Bobcia, Big Pop and Grammie. My family stood by me and was available for support and guidance no matter what day or time it was. I would also like to thank my fiancé Bryan Clark, whose constant love and support gave me strength on a daily basis. Finally, I would like to thank my pets, Boe Brady and Chica, whose unconditional love was given to me everyday regardless of my mood or time constraints. I am very appreciative to all my friends and family who supported me through the past five years. I know it has been a long road and I could not have done it without all of you.