The Role of Gnrh-II and Its Receptor in Testicular Function Amy Desaulniers University of Nebraska - Lincoln, Amy.Desaulniers@Huskers.Unl.Edu

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THE ROLE OF GnRH-II AND ITS RECEPTOR IN TESTICULAR FUNCTION

Amy T. Desaulniers

A THESIS

Presented to the Faculty of

The Graduate College at the University of Nebraska

In Partial Fulfillment of Requirements

For the Degree of Master of Science

Major: Animal Science

Under the Supervision of Professor Brett R. White

Lincoln, Nebraska

August, 2013

THE ROLE OF GnRH-II AND ITS RECEPTOR IN TESTICULAR FUNCTION

Amy Teresa Desaulniers

University of Nebraska, 2013

Advisor: Brett R. White

The second mammalian isoform of GnRH (GnRH-II) has been linked to regulation of cell proliferation, feed intake, and the interaction between energy balance and reproductive behavior. In contrast to the native form of GnRH (GnRH-I), GnRH-II is an inefficient modulator of gonadotropin secretion. Unlike many species, a functional receptor (GnRHR-II) specific to this ligand has been discovered in the pig that may be directly involved in testosterone production. Therefore, our objective was to identify the role of GnRH-II and its receptor in testicular function. First, there was 6-fold more

GnRHR-II protein in the testis than anterior pituitary gland of boars. GnRH-II levels determined by ELISA were highest in testis (1,321 pg/ml), intermediate in pituitary (393 pg/ml) and lowest in hypothalamus (220 pg/ml) tissue homogenates.

Immunohistochemistry indicated that the GnRHR-II was located on the plasma membrane of Leydig cells within the interstitium as well as germ cells. Second, immunocytochemistry demonstrated that GnRHR-IIs were localized to the neck region of spermatozoa. Consistent with this, GnRH-II was also detected in seminal plasma (225 pg/ml). Third, testicular tissue explants exposed to GnRH-II secreted testosterone similarly to hCG treated tissues, without influencing GnRHR-II or LH receptor protein

levels. Finally, 4 in vivo experiments were performed on boars surgically fitted with jugular cannulae. In Exp. 1, testosterone concentrations were elevated following treatment with either a GnRH-I (D-ala6 GnRH-I) or GnRH-II (D-ala6 GnRH-II) agonist, although GnRH-II induced changes in LH levels were not correlated with testosterone production. In Exp. 2, treatment with a GnRH-I (SB-75) or GnRH-II (Trp-1) antagonist suppressed testosterone, but not LH, secretion. In Exp. 3, testosterone concentrations increased after treatment with SB-75 followed by infusion of either GnRH-I or -II, despite the inability of GnRH-II to induce LH secretion. In Exp. 4, intratesticular injection with GnRH-I, GnRH-II or saline resulted in GnRH-II-induced stimulation of testosterone production without increasing LH levels. Thus, GnRH-II may directly impact testosterone production via binding to the GnRHR-II on Leydig cells, bypassing

LH production by the anterior pituitary gland.

Acknowledgements

First of all, I would like to thank Dr. Brett White for taking me on as a graduate student and exposing me to a wonderful balance of both basic and applied research. We spent many hours bleeding boars together, sometimes at 2 AM, which would not have been possible without his dedication to research. He also gave me great freedom to design and conduct experiments, something not many Master’s students are lucky enough to do. I am still astounded by how much I learned in these last 3 years and cannot wait to see what the next chapter brings here at UNL. I also would like to thank Drs. Andrea Cupp and Jennifer Wood for their support in completing this degree. I learned a tremendous amount in their classrooms, yes, but I also benefited from their guidance in everything from laboratory skills to presenting. They have also been extremely accommodating, always willing to meet with me despite their busy schedules. Thus, I am very grateful to have had them on my committee and look forward to working with them in the future. I also received a great deal of guidance from Drs. Clay Lents and Joe Ford. They were instrumental in performing the boar trials and gave a lot of excellent feedback on the data. Michelle McManus was also wonderful for running RIAs out at USMARC. When I first started in the lab, I knew next to nothing. So, I can safely say that Rebecca Cederberg taught me almost all my laboratory skills, and with such patience! She has also become a wonderful friend over the last 3 years, always willing to help me no matter the situation. We spent couples hours troubleshooting lab techniques together and I am so thankful to have had her guidance! I can still remember the look on her face, however, when I ripped a page out of my lab notebook to use as scrap paper….I will never do that again! If you have never met Ginger Mills, you should do so right away! She is such a great person and friend, and I am so lucky to have her on our team at UNL. The very first day I started, she took me under her wing, and I feel like I have never left! In addition to the fantastic support, I have also learned a great deal from her about my favorite livestock species and hers…pigs! Also, she was essential in our bleeding trials and always helped with a great attitude. Next, I would like to thank my parents for being such big fans these last 3 years. They have been tremendously supportive of my goals in animal reproduction research and have been there every step of the way to cheer me on. Almost every day I called my mom to update her on the day’s events. She listened to every gory detail and was so interested in my research, remembering what I told her and asking questions, even if she didn’t understand the area fully. And don’t worry mom…someday I will finish school and will move closer to you!

Also, I would be remiss if I didn’t mention my wonderful boyfriend, Jake Schweitzer. He has been so great along this whole process, even through spans when we never saw each other because I was writing my thesis till midnight or bleeding pigs at 2 AM. On nights like that, he was my number one dog sitter to my boxer, Watson (aka my baby)…and for that I am SOOOO grateful. Moreover, he is always willing to listen to me blather on about testes and boar sperm…or at least I think he is listening!! A lot of this data would not have been possible without the help of several people in the lab. Bill Pohlmeier taught me how to perform IHC and immunofluorescence. Scott Kurz taught me how to do RIAs. I also had a lot of technical assistance from fellow students like John Harms, Chanho Lee and Amy Voss, Daniel Chia and Sara Stauffer. Dr. Kim Varnold and Dr. Adam Summers were also my premier statistics helpers! Finally, I have to talk about the awesome friends I made at UNL: Adam Summers, Bill Pohlmeier, Kristin Beede, Theresa Johnson, Kim Varnold, Justine Stevenson, Kevin Sargent, Michelle Semler, Renee McFee and Becky Vraspir….you guys really made Lincoln feel like home and I appreciate your friendship! vi

TABLE OF CONTENTS

Page

LIST OF FIGURES ...... viii LIST OF TABLES ...... xi

CHAPTER I: Introduction Introduction ...... 1

CHAPTER II: Literature Review Gonadotropin-Releasing Hormone I Structure ...... 6 Reproductive Function ...... 8 Gonadotropin-Releasing Hormone Receptor I Structure ...... 10 Cell Signaling...... 13 Internalization ...... 16 Tissue Expression ...... 17 GnRH-I and its Receptor in the Testis ...... 18 Role in Cancer ...... 22 Gonadotropin-Releasing Hormone II Structure ...... 23 Tissue Distribution ...... 24 Reproductive Function ...... 25 Role in Behavior ...... 28 Role in Nutrition ...... 29 Role in Cancer...... 30 Gonadotropin-Releasing Hormone Receptor II Structure ...... 32 Species Distribution ...... 33 Ligand Interaction ...... 38 Cell Signaling...... 41 Tissue Expression ...... 42 GnRH-II and its Receptor in the Boar ...... 44 Reproductive Physiology of the Boar Testosterone Production...... 49 Spermatogenesis ...... 52 Differences between Chinese Meishan and White Crossbred Boars ...... 57 Specific Antagonists of GnRH Isoforms SB-75 ...... 58 Trptorelix-1 ...... 60

vii

CHAPTER III: Materials and Methods Localization of the GnRHR-II Tissue Collection ...... 63 Embedding and Sectioning ...... 63 Immunohistochemistry ...... 64 Immunohistochemical Validation ...... 65 Protein Extraction ...... 66 Western Blot ...... 68 Semen Collection ...... 71 Protein Extraction ...... 71 Western Blot ...... 72 Immunocytochemistry ...... 72 ELISA ...... 73 Testicular Explant Materials ...... 75 Treatment ...... 75 Protein Extraction ...... 77 Western Blot ...... 77 Radioimmunoassay ...... 77 Boar Trials Animals ...... 79 Experiment 1: GnRH Agonist Trial ...... 79 Animals ...... 81 Experiment 2: GnRH Antagonist Trial ...... 81 Experiment 3: SB-75 + GnRH Agonist Trial ...... 82 Experiment 4: Intratesticular GnRH Injection Trial ...... 85 Radioimmunoassay ...... 87 Statistical Analysis ...... 88

CHAPTER IV: The Role of GnRH-II and its Receptor in the Testes of Boars Abstract ...... 89 Introduction ...... 91 Materials and Methods ...... 94 Results ...... 104 Discussion ...... 126

APPENDIX I ...... 134

Literature Cited ...... 139

viii

LIST OF FIGURES Page

Figure 1.1. The relative change in plasma LH and testosterone concentrations in white crossbred (WC) and Chinese Meishan (MS) boars after administration of a specific GnRHR-I antagonist, SB-75 ...... 5

Figure 2.1. Conformation of GnRH-I when bound to the GnRHR-I ...... 9

Figure 2.2. A graphical depiction of monkey GnRH-I receptor compared to the human GnRH-II receptor ...... 12

Figure 2.3. Diagram of cell signaling pathways activated following GnRH-I binding to GnRHR-I ...... 14

Figure 2.4. Diagram of coding exons for GnRHR-II in the human, sheep, cow, pig and rat...... 37

Figure 2.5. A graphical depiction of the porcine 5- and 7-transmembrane GnRH-II receptor isoforms ...... 39

Figure 2.6. Tissue expression of GnRHR-II mRNA in marmoset tissues ...... 43

Figure 2.7. Plasma testosterone levels in barrows, boars or boars immunized against cGnRH-II or lGnRH-III...... 45

Figure 2.8. Plasma levels of FSH or LH in barrows, boars or boars immunized against cGnRH-II or lGnRH-III...... 46

Figure 2.9. The effect of increasing LH levels on testosterone concentrations of primary Leydig cell cultures from control boars or boars immunized against cGnRH-II or lGnRH-III ...... 47

Figure 2.10. The effect of increasing hCG levels, in the presence or absence of SB-75, on testosterone production from testicular explant media following 3 h of culture ...... 48

Figure 2.11. Photomicrograph of Leydig cells within the interstitium of the boar testis . 50

Figure 2.12. The Δ4 and Δ5 pathways of testosterone production...... 53

Figure 2.13. Graphical depiction of spermatogenesis ...... 55

Figure 3.1. Immunohistochemistry of boar testes using multiple concentrations of an antibody directed against the GnRHR-II (Negative control, 1:200, 1:100, 1:50, 1:25, 1:10, respectively) ...... 67

Figure 3.2. Schematic indicating time of pretreatment (PRE; 0 or 1 µM GnRH-II), treatment (TRT; 0 or 1 I.U. hCG), media collection (hash mark) and tissue collection for testicular explants ...... 76

Figure 3.3. Schematic of boar GnRH agonist trial indicating treatment (TRT) and time of blood sample (hash mark) ...... 80

Figure 3.4. Schematic of GnRH boar antagonist trial indicating treatment (TRT) and time of blood sample (hash mark) ...... 83

Figure 3.5. Schematic of SB-75 + GnRH agonist trial indicating SB-75 administration, treatment (TRT) and time of blood sample (hash mark) ...... 84

Figure 3.6. Schematic of boar intratesticular GnRH injection trial indicating treatment (TRT) and time of blood sample (hash mark) ...... 86

Figure 4.1. Representative immunohistochemistry image of boar testes (n = 7) using an antibody directed against GnRHR-II (Panels B-D) ...... 105

Figure 4.2. Representative Western blot of porcine anterior pituitary gland and testis tissue (n = 5) using an antibody directed against GnRHR-II (Panel A)...... 106

Figure 4.3. GnRH-II levels in the hypothalamus, anterior pituitary gland and testis of boars ...... 108

Figure 4.4. Representative Western blot of boar testis and purified spermatozoa protein (n = 5) using an antibody directed against GnRHR-II (Panel A) ... 109

Figure 4.5. Representative confocal microscopy images of boar spermatozoa (n = 5) using an antibody directed against GnRHR-II labeled with an Alexa Fluor® 488 secondary antibody (Panels A-C; green)...... 111

Figure 4.6. Relative testosterone levels secreted by testicular explant cultures (n = 7) pretreated with or without 1 µM GnRH-II followed by treatment with vehicle or 1 I.U. hCG ...... 112

Figure 4.7. Representative Western blot of porcine explant cultures (n = 7) using an antibody directed against GnRHR-II (Panel A) and LHR (Panel B) ...... 114

Figure 4.8. Relative levels of LH and testosterone after intravenous administration of a specific GnRH-I or GnRH-II agonist...... 115

Figure 4.9. Relative levels of LH and testosterone after intramuscular administration of a specific GnRH-I (SB-75) or GnRH-II antagonist (Trp-1) ...... 117

Figure 4.10. Trend lines for relative changes in LH and testosterone concentrations following suppression with a specific GnRH-I (SB-75) or GnRH-II (Trp-1) antagonist ...... 118

Figure 4.11. Relative levels of LH and testosterone after intramuscular administration of SB-75 and intravenous treatment with a specific GnRH-I or GnRH-II agonist...... 122

Figure 4.12. Relative levels of LH and testosterone after intratesticular administration of saline (control), GnRH-I agonist or GnRH-II agonist ...... 124

LIST OF TABLES Page

TABLE 2.1. COMPARISON OF GnRH ISOFORMS IN VERTEBRATES ...... 7

TABLE 2.2. COMPARISON OF RELATIVE BINDING AFFINITIES AND SIGNALING MOLECULES BETWEEN THE MARMOSET GnRHR-II AND HUMAN GnRHR-I ...... 40

TABLE 2.3. STEROIDS IN BOAR TESTICULAR VEIN BLOOD ...... 51

TABLE 2.4. MEAN INOSITOL PHOSPHATE IC50 VALUES IN GH3 CELLS OVEREXPRESSING EITHER GnRHR-I or GnRHR-II FOLLOWING TREATMENT WITH SB-75 OR TRPTORELIX-1 IN THE PRESENCE OF 1 nM GnRH-II ...... 62

TABLE 4.1 MEAN FOLD CHANGE FOR LH CONCENTRATIONS ± SEM FOLLOWING TREATMENT WITH A GnRH-I ANTAGONIST (SB-75) AND SUBSEQUENT INFUSION OF AGONISTS FOR GnRH-I OR -II ...... 121 1

CHAPTER I

INTRODUCTION

Boar subfertility is a growing constraint of pork production (Dyck et al., 2011) and can impact industry profitability by reducing sow reproductive efficiency and increasing costs to boar studs. First, utilizing semen from subfertile boars can impact sow productivity as poor semen quality is detrimental to conception rates. For example, approximately 11.6 million sows farrowed in 2010 (USDA-NASS, 2010). Based on an estimated feed cost of $40.42 per sow from farrowing to weaning, a 5% reduction in conception rates could lead to $19.9 million spent feeding non-productive sows.

Similarly, it is easy to predict the direct impact of decreasing average litter size, especially since 93% of the pig inventory is on farms raising at least 1,000 pigs (Levis,

2000). Thus, even a 0.2 decrease in number of piglets born alive could result in a

$99,000 loss to a producer raising 10,000 sows. Second, male subfertility also results in economic losses within a boar stud. Subfertile boars likely contribute to fewer doses of semen sold per ejaculate compared to fertile boars, although input costs remain the same.

Approximately 80% of swine producers utilize artificial insemination (AI), requiring 30 million doses of boar semen annually (Kuster and Althouse, 2008). If the average cost per dose is $6.50, even a 1% increase in the number of available doses would provide an additional $2 million per year to US boar studs. 2

The population of subfertile boars is growing despite the use of screening methods to detect suboptimal samples (Dyck et al., 2011). The use of these conventional semen analyses (morphology, motility, concentration and seminal volume) is common in boar studs (Amann et al., 1995). Although these methods are correlated with fertility

(Flowers, 1997; Xu et al., 1998), many authors suggest that these parameters do not reflect the true fertility of a boar (Correa et al., 1997; Brahmkshtri et al., 1999).

Therefore, compensation techniques have been developed in the industry to mask subfertile boars, despite passing standard semen screening (Dyck et al., 2011).

Commonly, semen collections are pooled from multiple sires and AI doses packaged in excess of 3 billion sperm cells (Dyck et al., 2011). These techniques may yield acceptable conception rates but drastically reduce the efficiency of a stud operation. Thus, there is a critical need to better understand testicular function in the boar.

Recently, a new mammalian form of GnRH (GnRH-II) was discovered.

Originally isolated in the chicken (Miyamoto et al., 1984), this decapeptide differs from native GnRH (GnRH-I) by 3 amino acids and is expressed in almost every tissue (Kasten et al., 1996; Millar, 2003). GnRH-II can function through the GnRH-I receptor to activate signal transduction and it also binds its own specific receptor (GnRHR-II).

Originally cloned and sequenced in catfish (Tensen et al., 1997), the GnRHR-II is also present in mammals and like GnRHR-I, is a 7-transmembrane, G-protein coupled receptor (Neill et al., 2001; Millar et al., 2001). However, unlike the GnRHR-I, this receptor contains an intracytoplasmic tail, characteristic of rapid receptor internalization

(Ronacher et al., 2004). Moreover, not all species maintain the coding sequence to 3 produce a functional GnRHR-II due to frameshift mutations and premature stop codons

(Morgan et al., 2003; Gault et al., 2004a; Stewart et al., 2009). Interestingly, the sequence for a functional receptor has been discovered in shrews, old world monkeys and swine

(Stewart et al., 2009). In these species, the function of GnRH-II and its receptor is unclear

(Neill et al., 2004) but evidence suggests that GnRH-II is not an efficient modulator of gonadotropin secretion (Okada et al., 2003). GnRH-II and its receptor have been linked to feed intake (Kauffman and Rissman, 2004b), as well as the interaction between energy balance and sexual behavior in females (Kauffman and Rissman, 2004a; Kauffman et al.,

2005; Kauffman et al., 2006). Further, GnRH-II may regulate cell proliferation and apoptosis in peripheral reproductive tissues (Chou et al., 2002). In addition, new evidence suggests that GnRH-II might also be involved in male reproductive function.

Bowen et al. (2006) discovered that boars immunized against GnRH-II displayed reduced levels of testosterone independent of alterations in gonadotropin secretion.

Similarly, primary Leydig cell cultures from GnRH-II immunized boars secreted less testosterone compared to control boars upon LH challenge. Additionally, boars treated with a specific GnRH-I antagonist (SB-75) daily for 96 h displayed suppressed testosterone levels for the duration of treatment, whereas LH levels returned to basal after only 36 h of treatment (Fig. 1.1; Zanella et al., 2000). Consistent with this, testicular explant cultures treated with human chorionic gonadotropin in the presence of SB-75 produced less testosterone compared to controls (Zanella et al., 2000). Given that

GnRHR-I has never been detected in the boar testis, Zanella et al. (2000) postulated that a different mechanism must be controlling testosterone production directly at the testis. 4

Differential regulation of testosterone secretion could lead to advancements in human fertility treatments, contraceptives, cancer research or pharmacological agents.

Moreover, the discovery of autocrine or paracrine mechanisms for testosterone regulation may aid the swine industry as well. If GnRH-II or its receptor are correlated with testosterone production, novel screening methods could be utilized in boar studs to improve reproductive efficiency. Eliminating infertile or subfertile boars from the herd early will reduce the economic losses associated with rearing and training.

Implementation of these new technologies could lead to enhanced productivity and profitability of pork producers.

Relative change in change LH Relative

Relative change in T in change Relative

Time (h) Figure 1.1. The relative change in plasma LH and testosterone concentrations in white crossbred (WC) and Chinese Meishan (MS) boars after administration of a specific GnRHR-I antagonist, SB-75 (arrows). Levels of LH returned to normal 36 h after the initial treatment, whereas testosterone concentrations remained suppressed for 96 h. From Zanella et al. (2000). 6

CHAPTER II

LITERATURE REVIEW

Gonadotropin-Releasing Hormone I

Structure. Gonadotropin-releasing hormone (GnRH) has been conserved throughout 500 million years of evolution (Millar, 2005) and has been discovered in every vertebrate class examined (Tsai and Zhang, 2008). For almost 30 years, GnRH was thought to be a novel decapeptide (Schally, 1973), however, 23 other forms of GnRH have been identified since then (Millar, 2005), all with 10 amino acids and at least a 50% sequence identity (Dubois et al., 2002). In vertebrates, 3 forms are the most common

(Table 2.1; Millar, 2005). The classical form of GnRH has been designated GnRH-I, as it was the first form to be discovered in mammalian hypothalamic tissues (Baba et al.,

1971; Matsuo et al., 1971; Schally et al., 1971). Two isoforms, GnRH-II and GnRH-III, were later isolated in the chicken (Miyamoto et al., 1984) and lamprey (Sower et al.,

1993), respectively. Today, it is well known that these 2 isoforms are also prevalent in vertebrates (Millar, 2005). GnRH-II is present in many mammalian species, but to date

GnRH-III has not been identified in mammals (White et al., 1998).

GnRH-I is a polypeptide consisting of 10 amino acids: pGlu-His-Trp-Ser-Tyr-

Gly-Leu-Arg-Pro-Gly-NH2 (Baba et al., 1971; Matsuo et al., 1971; Schally et al., 1971).

The amino acid in the eighth position is considered to be the most variable (followed by

5, 6 and 7) among all isoforms of GnRH. However, Arg must be in position 8 for 7

TABLE 2.1. COMPARISON OF GnRH ISOFORMS IN VERTEBRATES. Isoforms Amino Acid Sequencea

GnRH-I pGlu-His-Trp-Ser-Tyr-Gly-Leu-Arg-Pro-Gly-NH2

GnRH-II pGlu-His-Trp-Ser-His-Gly-Trp-Tyr-Pro-Gly-NH2

GnRH-III pGlu-His-Trp-Ser-His-Asp-Trp-Lys-Pro-Gly-NH2 a Amino acids in bold are different from the classic mammalian decapeptide, GnRH-I. 8 gonadotropin production in the pituitary (Karten and Rivier, 1986; Sealfon et al., 1997).

The NH2- and COOH-termini of GnRH-I have been identically conserved through evolution, indicating absolute necessity for biological function (Millar, 2005). The residues on the NH2-terminus are involved in receptor activation and binding. While the

COOH-terminus is only involved in receptor binding. The conformation of GnRH-I is considered a β-II type turn involving amino acids 5-8 (Fig. 2.1; Karten and Rivier, 1986;

Sealfon et al., 1997). Substitutions with D-amino acids in the NH2- terminus result in antagonist production whereas substitutions at residue 6 (Gly) will enhance binding affinity by stabilizing the conformation and therefore, increasing the half-life (Millar,

2002).

Reproductive Function. GnRH-I may have first functioned in cell to cell communication for reproduction in simple invertebrates (Millar, 2005). Indeed, GnRHs have been found in the neural tissue of marine filter feeders and they function to activate the gonads through release into the bloodstream (Powell et al., 1996). Interestingly, this occurs without an organized pituitary gland, suggesting regulation of reproduction by the pituitary, as within mammals, came later in evolution (Millar, 2005).

Mammalian production of GnRH-I occurs in approximately 1,000 specialized neurons originating from the preoptic area and mediobasal hypothalamus of the brain. A precursor polypeptide is enzymatically cleaved and packaged in secretory granules which are then transported through axons, terminating within the medium eminence (Seeburg et al., 1987; Fink, 1988). Newly synthesized GnRH-I is secreted in a pulsatile manner 9

Figure 2.1. Conformation of GnRH-I when bound to the GnRHR-I. The NH2-terminus is involved in receptor binding and activation, whereas the COOH-terminus is only needed for receptor binding. D-amino acid substitution of residues in the NH2-terminus will result in the production of an antagonist, whereas substitutions for Gly6 will produce an agonist, marked by increased binding affinity and half-life. From Millar et al. (2005). 10

(every 30 - 120 min) into the hypothalamic-hypophyseal portal system, for transport to the anterior pituitary gland (Fink, 1988). However, clearance of GnRH-I in blood occurs quickly, with a half-life of approximately 2 - 8 min (Pimstone et al., 1977).

Once GnRH-I reaches the anterior pituitary gland, the GnRH-I receptor (GnRHR-

I) is upregulated in gonadotrope cells to aid in ligand binding for signal transduction

(Lahlou et al., 1987). Next, gonadotropin production and secretion is initiated through the transcription and translation of genes encoding the common α-glycoprotein and the distinct FSHβ- and LHβ-subunits (Conn and Crowley, 1991; Stojilkovic and Catt,

1995). Heterodimerization of the common α-subunit and specific β-subunits result in the glycoprotein hormones, follicle stimulating hormone (FSH) or luteinizing hormone (LH;

Conn and Crowley, 1991; Stojilkovic and Catt, 1995). A pulse of LH occurs after every pulse of GnRH-I, however, synthesis and secretion of FSH occurs in a more constitutive manner (Millar, 2005). Studies demonstrated that LHβ-subunit production is favored when pulses of GnRH-I are frequent (1/min) whereas the FSHβ-subunit is preferentially produced when pulses occur every 120 min (Kaiser et al., 1997). While GnRH-I stimulates gonadotropin production, constant GnRH-I stimulation will reduce gonadotropin secretion due to GnRHR-I downregulation and desensitization of gonadotrope cells (Millar et al., 1987; Casper, 1991; Conn and Crowley, 1991).

Gonadotropin Releasing Hormone Receptor I

Structure. The mouse GnRHR-I was first cloned in the early 1990’s (Tsutsumi et al., 1992). Classified as a 7-transmembrane (TM), G-protein coupled receptor (GPCR), 11 it is a member of the rhodopsin-like GPCR superfamily (Fig. 2.2; Cheng and Leung,

2005). Although it is among the smallest GPCRs ever identified at 327 amino acids

(Brothers et al., 2002), this receptor has stringent requirements for ligand conformation

(Millar et al., 1987). Like most GPCRs, the NH2-terminal domain is followed by 7 putative, α-helical TM domains connected by 3 extracellular loop (ECL) domains and 3 intracellular loop (ICL) domains. The ECL are thought to be involved in ligand binding and the TM domains are associated with the conformational change needed for receptor activation (Millar, 2005).

Most GPCRs have intracytoplasmic tails which are associated with rapid desensitization and internalization (Pawson et al., 2003b). In the GnRHR-I, the carboxyl tail is truncated at the plasma membrane after the seventh TM domain (Eidne et al.,

1992). Millar (2005) hypothesized that the absence of the carboxyl-terminal tail, must be due to a recent beneficial adaption. Although there is no defined tail, the carboxyl- terminal amino acids of TM 7 are important for G-protein coupling, the intracellular mechanism which initiates signal transduction. Since the last 4 residues are conserved in all mammals, they are perceived to be essential to receptor signaling. For example, a mutant GnRHR-I (Ser326Tyr) decreased G-protein coupling, and the mutation of all terminal residues resulted in no effector coupling. The ICL within the GnRHR-I are important for G-protein coupling to the receptor. Both ICL 2 and ICL 3 are thought to couple with the G-protein subunit, Gq/11, whereas ICL1 is involved in Gs signaling (Millar and Pawson, 2004). 12

I receptor. From I Neill receptor.

II II receptor compared to the human GnRH

Graphical Graphical depiction of the monkey GnRH

2. 2.

Figure 2. Figure (2004). al. et 13

Cell Signaling. Gonadotropes make up only 5-10% of the cell population within the anterior pituitary gland but are widely dispersed (Naor and Huhtaniemi, 2012).

Although they are not localized in 1 region of the pituitary, they coordinate together to create pulses of gonadotropin secretion (Krsmanovic et al., 1993). In fact, even gonadotropes dissociated from adjacent cells in culture will secrete gonadotropins in a synchronized fashion (Krsmanovic et al., 1993). Binding of GnRH-I to its cognate receptor is a basic ligand-receptor interaction. Binding stimulates intracellular signaling by initiating receptor activation and conformational changes (Wu et al., 2009). Upon binding, GDP is exchanged for GTP in heterotrimeric G-protein complexes. This causes the dissociation of the subunit Gα from Gβγ, resulting in GTP hydrolysis by GTPase activity. Formation of the Gα-GDPβγ complex resumes and the receptor returns to its inactive conformation, however signal transduction has begun (De Lean et al., 1980). In the pituitary, the predominate G-protein associated with gonadotropin secretion is Gq/11

(Fig. 2.3; Ando et al., 2001; Ferris et al., 2007; Grafer et al., 2009). The hydrolysis of

GTP activates the G-proteins, Gq/G11, thereby increasing the activity of phospholipase C

(PLC) which hydrolyses phosphoinositide, generating inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG). Increasing levels of intracellular IP3 bind to receptors on the endoplasmic reticulum (ER), resulting in sequestered Ca2+ release. Meanwhile, DAG increases intracellular Ca2+ by opening voltage-gated calcium channels in the plasma membrane of the cell and activates protein kinase C (PKC; Stojilkovic and Catt, 1995).

PKC mediates mitogen activated protein kinase (MAPK) activation, including 14

Figure 2.3. Diagram of cell signaling pathways activated following GnRH-I binding to GnRHR-I. When GnRH-I and its receptor bind, Gs/αβγ is activated; causing the dissociation of Gs/α that activates adenylate cyclase (AC). Adenylate cyclase converts ATP to cAMP which subsequently activates protein kinase A (PKA) and cAMP response element binding protein (CREB). Alternatively, Gq/11α stimulates phospholipase C (PLC) which converts phosphoinositol 4,5-bisphosphate (PIP2) into diacylglycerol (DAG) and inositol 1,4,5-trisphosphate (IP3). Calcium is released from extracellular and intracellular endoplasmic reticulum (ER) stores by DAG and IP3 stimulation. Increased calcium binds calmodulin (CaM) and activated calmodulin kinase (CaMK). Protein kinase C (PKC) is stimulated by Ca2+ and DAG. Multiple mitogen activated protein kinase (MAPK) pathways are activated by PKC including extracellular signal-regulated kinase (ERK), c- Jun N-terminal kinase (JNK), p38 MAPK, and big MAPK (BMK), resulting in the recruitment of transcription factors (i.e., Jun and ELK) that stimulate LH and FSH gene expression. Adapted from Naor et al. (2000). 15 extracellular signal-regulated kinase 1/2 (ERK1/2), c-Jun N-terminal kinase (JNK), p38

MAPK, and ERK5 (big MAPK; BMK). MAPKs will then translocate to the nucleus to phosphorylate transcription factors that support gonadotropin gene expression (Naor and Huhtaniemi, 2012), resulting in the synthesis and secretion of LH and FSH.

Interestingly, certain gonadotropes are capable of secreting both LH and FSH (bi- hormonal), whereas others secrete gonadotropins in a mono-hormonal fashion (Childs,

1983).

While Gq/11 is the principal G-protein activated, GnRHR-I can also utilize other G- proteins. Signaling via Gs has been demonstrated in gonadotropes as well as other tissues

(Ando et al., 2001; Ferris et al., 2007; Grafer et al., 2009). Once ligand binding occurs, the Gα-subunit will dissociate and activate the enzyme adenylate cyclase (AC), which is responsible for the conversion of ATP to cAMP. As cAMP levels rise in the cell, they bind regulatory subunits on protein kinase A (PKA), causing the release of catalytic subunits responsible for direct protein activation via phosphorylation or protein synthesis.

Protein synthesis is stimulated when cAMP response element-binding protein (CREB) binds to cAMP response elements (CRE) within the promoter of a gene within the nucleus of gonadotropes (Cheng and Leung, 2001).

This receptor has also been shown to signal through Gi, although the mechanism is still poorly understood. This G-protein acts very differently from both Gs and Gq/11, rather than activating signal transduction, Gi acts as an inhibitory modulator of cAMP production. Specifically, Gi inhibits the activation of AC, preventing the production of cAMP. For example, when rat gonadotropes were treated with pertussis toxin, 16 intracellular IP3 levels were significantly reduced, indicating the activation of the Gi subunit (Hawes et al., 1993). Moreover, it has been demonstrated that GnRHR-I couples to Gi in many cancerous tumors including ovarian, uterine (Imai et al., 1996), and prostate (Limonta et al., 1999).

While it is apparent that GnRHR-I can signal through many different G-proteins, the mechanism behind G-protein selectivity is unclear. Interestingly, GnRHR-I appears to be in a different conformation for activation of Gi versus Gs and Gq/11. This conformational selectivity of GnRHR-I may be due to many factors, including the nature of the ligand (Millar, 2005), cell type, cell cycle stage or the mere availability of Gi machinery within the cell (Millar et al., 2004). Additionally, GnRHR-I has allosteric sites thought to cause alterations in conformation (Millar et al., 2004). In fact, some researchers postulated that the conformational selectivity of GnRHR-I in a particular cell- type or cellular environment can dictate the type of G-protein utilized due to allosteric sites alone (Naor and Huhtaniemi, 2012). Interestingly, GnRHR-I also activated multiple signaling cascades through Gq/11 alone in αT3 cells (Grosse et al., 2000).

Internalization. Classically, GPCRs are internalized by GPCR kinases, β-arrestin and clathrin-coated pits (Millar et al., 2004). After receptor activation and the generation of free Gβγ-subunits, GPCR kinases phosphorylate the receptor at specific serine and threonine residues (De Camilli et al., 1995). Phosphorylation enhances the binding of the protein, β-arrestin, thought to be the signal for internalization. Next, the GTPase, dynamin, initiates formation of a clathrin-coated pit from an invagination of the plasma 17 membrane (De Camilli et al., 1995). The specialized vesicle surrounds and engulfs the receptor, removing it from the plasma membrane. The clathrin is degraded in the cytoplasm of the cell and the vesicle fuses with an endosome (Sorkin and Von Zastrow,

2002). Here, the path of the internalized receptor can vary. Receptors can either be sequestered in endosomes, recycled back to the plasma membrane or transported to lysosomes for proteolysis (Sorkin and Von Zastrow, 2002). Although GPCRs, like

GnRHR-I, are most commonly internalized by clathrin-coated pits, they can also be internalized by smooth, non-coated membranes, vesicles or caveolae, independent of β- arrestin or dynamin (Raposo et al., 1989; Anderson, 1998; Claing et al., 2000).

Tissue Expression. Once considered a solely hypothalamic hormone (Schally et al., 1971), later evidence suggests GnRH-I is also produced in low levels (White et al.,

1998) elsewhere in the body (Gibbons et al., 1975). GnRH-I is widely distributed outside of the brain (Barry, 1979; Sharpe et al., 1982). However, its size, short half-life and relatively low concentration make an extra-pituitary endocrine function infeasible (Nett et al., 1974; Pimstone et al., 1977). Thus, many have hypothesized that GnRH-I is produced locally (Sharpe and Cooper, 1982a). Indeed, studies suggest that extrapituitary GnRH-I was produced in an autocrine or paracrine manner (Millar, 2005; Lin et al., 2010).

Further, Millar (2002) suggested that wide tissue distribution indicates many reproductive and non-reproductive functions have yet to be identified for GnRH-I. To date, GnRH-I has been found in the kidney, pancreas, lymph system, adrenal, pancreas, retina, the 18 immune system and extrahypothalamic brain, as well as reproductive tissues like the gonad, uterus, placenta, breast and prostate (Cheng and Leung, 2005; Millar, 2005).

In situ hybridization revealed GnRH-I mRNA in granulosa cells of primary, secondary and tertiary follicles in the rat ovary (Clayton et al., 1992; Whitelaw et al.,

1995). Further, GnRH-I mRNA was detected in human placental (JEG) cells (Dong et al., 1996) and rat gonads (Bahk et al., 1995). In addition, ligand binding sites were discovered on granulosa and luteal cells through radioligand binding assays (Clayton et al., 1979; Harwood et al., 1980). The localization of GnRH-I and its receptor to the gonads and other reproductive tissues of mammals indicates a specific function (Millar,

2003). Interestingly, GnRH-I is also present in non-mammalian vertebrates suggesting the development of an early reproductive function, possibly before the development of the hypothalamic-pituitary-gonadal (HPG) axis (Millar, 2005).

GnRH-I and its Receptor in the Testis. Evidence for the presence of a testicular

GnRHR-I was first discovered in the late 1970s, when LH receptor mRNA levels increased in response to GnRH-I administration in hypophysectomized rats (Hsueh and Erickson, 1979). Since then, GnRHR-I has been detected in the testis of many species including the human (Clayton et al., 1980; Bahk et al., 1995), alpaca (Zerani et al., 2011), mouse (Bull et al., 2000) frog (Pierantoni et al., 1984; Minucci et al., 1986), monkey (Sharpe and Fraser, 1980) and fish (Pati and Habibi, 1993). Interestingly, the boar (Zanella et al., 2000) and bull (Kakar et al., 1992) lack testicular GnRHR-Is entirely. 19

GnRH-I mRNA has been discovered in adult human testicular tissue, specifically in seminiferous tubules, Sertoli cells and some spermatogenic cells. GnRHR-I mRNA was also found in germ cells of mice and rats (Bull et al., 2000). Competitive binding assays revealed GnRHR-I mRNA within the interstitial compartment of the testis in humans and rats, including the Leydig cells (Clayton et al., 1980; Bahk et al., 1995).

Immunohistochemistry (IHC) and Western blot analyses indicated GnRHR-I was in the cytoplasm of alpaca Leydig cells (Zerani et al., 2011). Since Sertoli cells of several species can produce GnRH-I (Sharpe and Fraser, 1980; Verhoeven and Cailleau, 1985;

Sharpe and Cooper, 1987; Saint Pol et al., 1988) and Leydig cells express the GnRHR-I

(Iwashita and Catt, 1985; Jegou et al., 1985; Nikula and Huhtaniemi, 1988; Petersson et al., 1989; Reinhart et al., 1992; Kakar et al., 1992), it was hypothesized that GnRH-I can act in a paracrine manner within the testis (Bahk et al., 1995).

Evidence suggests that GnRH-I has a direct effect on the testis (Lin et al., 2010).

For example, rats injected with GnRH-I directly into the testis experienced an immediate increase in serum testosterone concentration although LH levels were unaffected (Sharpe et al., 1983). Moreover, short term treatment of GnRH-I on rat primary Leydig cell cultures increased testosterone production (Sharpe and Cooper, 1982b; Browning et al.,

1983; Molcho et al., 1984). However, long term exposure to GnRH-I decreased testosterone production after human chorionic gonadotropin (hCG) challenge, suggesting the downregulation of the GnRHR-I. Further, administration of GnRH-I to pre-pubertal mice increased GnRHR-I mRNA levels in the testes and increased IHC staining for the 20

GnRHR-I in Leydig cells (Anjum et al., 2012). These effects were also correlated with increased expression of the steroidogenic enzyme P450scc.

Interestingly, the scope of action for GnRH-I in the testis is not limited to steroidogenic mediation. For instance, treatment with a GnRH-I agonist (Buserelin) directly increased testosterone production and spermatogonial mitosis in frogs (Pierantoni et al., 1984; Minucci et al., 1986), suggesting that GnRH-I might also impact spermatogenesis. Subsequent research appears to substantiate these claims as high levels of GnRH-I and GnRHR-I mRNA were present in the testes of infertile men (Lin et al.,

2008). Transcript levels were also correlated with low serum testosterone concentrations and testicular volume. Therefore, the authors postulated that increased mRNA transcripts for GnRH-I, and GnRHR-I could be indicative of spermatogenic dysfunction, possibly due to a compensatory increase in gene transcription (Lin et al., 2008). While testosterone is required for proper spermatogenesis, recent research suggests that testosterone can be suppressive to proliferation of spermatogonia (Udagawa et al., 2002). Interestingly, treatment with a GnRH-I antagonist (SB-75) reduced intra-testicular testosterone levels and increased the regeneration of spermatogenesis by stimulating spermatogonial proliferation in irradiated (Shuttlesworth et al., 2000), cryptorchid (Udagawa et al.,

2002), and chemically castrated (Meistrich, 1999) rats, as well as mutant, spermatogonial depleted mice (Matsumiya et al., 1999).

In addition to the regulation of steroidogenesis and spermatogenesis, there is also evidence that supports a role for GnRH-I in fertilization. For example, ~350-fold more human spermatozoa bound the zona pellucida of an oocyte when first exposed to GnRH-I 21

(Morales, 1998), indicating that spermatozoa may interact with GnRH-I as they travel through the male and female reproductive tract, increasing the fertilization capacity. This effect was completely blocked by a GnRH-I antagonist (Morales et al., 1999). Similarly,

GnRHR-I has been found on mouse spermatozoa, also indicating that the interaction between testicular GnRH-I and GnRHR-I may be an important regulator of spermatogenesis (Anjum et al., 2012).

Localized testicular GnRH-I may also be involved in development of puberty and reproductive senescence of the male mouse. It is well recognized that puberty is achieved when GnRH-I neurons are stimulated to begin secretion and reproductive senescence is marked by drastic reductions in testosterone production (Chandolia et al., 1997).

Interestingly, FSH and LH levels increased at senescence, suggesting a primary testicular defect initiates reductions in testosterone (Zwart et al., 1996). Indeed, Anjum and coworkers (2012) discovered that increased staining for the testicular GnRHR-I coincided with the onset of puberty in the mouse whereas reproductive senescence coincided with a reduction in GnRHR-I staining in the testis, followed by a compensatory increase in

GnRH-I. The authors proposed that this phenomenon may work synergistically with regulation of puberty and senescence at the level of the hypothalamus (Anjum et al.,

2012). In conclusion, it appears that GnRH-I and its receptor regulate testicular function in 2 ways, indirectly through alterations in gonadotropin secretion and directly at the level of the testis (Botte et al., 1999).

Role in Cancer. Classically, GnRH-I has been used to inhibit androgen dependent cancer growth (Engel and Schally, 2007). Both GnRH-I agonists and antagonists are useful treatments as they either downregulate or block the GnRH-I receptor in the hypothalamus, reducing androgen levels (Engel and Schally, 2007). In prostate cancer, androgens are vital mitogenic signals for normal cell proliferation, but can also be the cause of an imbalance in cell division (Cook and Sheridan, 2000).

Indeed, the instances of prostate cancer in castrated males are rare (Huggins and Hodges,

2002). In early stages of prostate cancer, androgen deprivation is suppressive to cancer growth, however, an androgen independent growth stage will develop over time (Navarro et al., 2002).

Interestingly, GnRH-I can also have a direct effect on prostate cancer cell growth without working through the HPG axis, and thus, is also a potent androgen independent inhibitor of cancer growth (Moretti et al., 1996). For example, the invasiveness of prostate cancer cells was reduced when treated with GnRH-I, through activation of Gi and inhibition of epidermal growth factor (Moretti et al., 1996). Moreover, insulin-like growth factor-induced cell proliferation was mitigated by GnRH-I agonist treatment

(Zoladex; Moretti et al., 1996; Marelli et al., 1999). Similarly, in vitro proliferation of endometrial, ovarian and breast cancer cells was diminished after treatment with GnRH-I agonists (Emons et al., 2003) and antagonists (Emons et al., 1993a; Emons et al., 1993b;

Irmer et al., 1995; Emons et al., 1997; Grundker et al., 2002b). Interestingly, breast cancer cell proliferation was reduced whereas cell apoptosis was increased after treatment 23 with GnRH-I (Hong et al., 2012). Indeed, GnRH-I analogues can inhibit tumor growth in both an anti-proliferative and pro-apoptotic manner (Cheung et al., 2006).

Apoptosis naturally occurs in healthy cells in response to toxic stimuli or developmental signals and hallmarks include DNA fragmentation, cell shrinkage, plasma membrane blebbing and formation of apoptotic bodies (Jin and El-Deiry, 2005; Elmore,

2007). Treatment with GnRH-I induced apoptosis in rat granulosa cells and human myometrium cells in vivo and in vitro (Yano et al., 1997; Wang et al., 2002). Similarly, pro-apoptotic signals increased in human cancer cell lines (benign prostate hyperplasia and choriocarcinoma) treated with GnRH-I via activation of Gi (Maudsley et al., 2004).

In addition, GnRH-I has apoptotic effects on human granulosa cells (Leung et al., 2003), activating the proteolytic caspase-dependent extrinsic signaling cascade rather than the intrinsic pathway that involves Bcl-2 members (Hong et al., 2008). Interestingly, treatment of rat granulosa cells with a GnRH-I agonist (leuprolide acetate) involved some

Bcl-2 members, suggesting the mechanism initiating apoptosis may be species-specific

(Parborell et al., 2008). Induction of apoptosis by GnRH-I was also shown in human breast cancer cell lines through stress-activated p38 MAPK, loss of mitochondria membrane potential and activation of caspase-3 (Grundker et al., 2010).

Gonadotropin-Releasing Hormone II

Structure. The second isoform of GnRH, GnRH-II, was originally identified in the chicken hypothalamus (Miyamoto et al., 1984). Since then, it has been found in animals of every vertebrate class, from lower orders, such as bony fish, to complex 24 mammals, like humans (Fernald and White, 1999), suggesting it may be the most ancient form of GnRH (Millar et al., 2004). In these animals, the structure of GnRH-II remains entirely conserved, indicating high selection pressure and thus, a critical evolutionary function (Millar and King, 1987). Therefore, complete conservation has persisted despite 500 million years of evolution (Fernald and White, 1999). GnRH-II is also a decapeptide (Table 2.1), differing from GnRH-I by only 3 amino acids (His5,Trp7,Tyr8;

Miyamoto et al., 1984), giving it a 70% sequence identity to GnRH-I (White et al., 1998).

Moreover, GnRH-II has a β-turn conformation similar to GnRH-I. However, GnRH-II exists in a preconfigured conformation, unlike GnRH-I. Thus, GnRH-II does not require extensive conformational changes for receptor activation (Pfleger et al., 2002; Millar,

2003). This indicates GnRH-II has increased stability over GnRH-I, approximately 6- fold higher (Siler-Khodr and Grayson, 2001), which may be reflected in its diverse tissue distribution (Pawson et al., 2003b).

Tissue Distribution. Like GnRH-I, GnRH-II is present in the brain, specifically the pre-optic and medio-basal hypothalamic areas, the midbrain and limbic structures

(Sealfon et al., 1997; Millar et al., 2001; Millar, 2002). However, GnRH-II was only scarcely found in the hypothalamic regions known to regulate gonadotropin secretion

(Rissman et al., 1995). In primates and humans, GnRH-II has also been isolated in the caudate nucleus, hippocampus, amygdala and the peripheral nervous system (Lescheid et al., 1997; Urbanski et al., 1999). Similar to GnRH-I, GnRH-II was also found outside of the brain, potentially due to autocrine or paracrine production as determined by gene 25 expression analysis (Cheon et al., 2001; Kang et al., 2001; Millar, 2003). Interestingly,

GnRH-II is expressed at the highest level outside the brain (White et al., 1998). GnRH-II mRNA has been detected in the kidney, bone marrow, prostate (White et al., 1998), testis

(An et al., 2008), placenta (Siler-Khodr and Grayson, 2001), uterine endometrium

(Cheon et al., 2001), and mammary gland (Chen et al., 2002), as well as ovarian epithelial (Choi et al., 2001) and granulosa cells (Kang et al., 2001). Although GnRH-I is also present in many extra-hypothalamic tissues, GnRH-II is considered to be more ubiquitously expressed (Millar et al., 2004). Given the diverse tissue distribution of

GnRH-II, many have hypothesized different roles for this ligand.

Role in Reproduction. Since this peptide was first localized to the brain, it was originally speculated that GnRH-II functions like GnRH-I in gonadotropin release

(Rissman et al., 1995). Indeed, early studies with mammalian pituitary cultures found that high doses of GnRH-II could stimulate LH release, albeit at about 2% the efficacy of

GnRH-I (Millar and King, 1983; Millar et al., 1986). Additionally, infusions of GnRH-

II could stimulate ovulation, but with only ~10% the potency of GnRH-I (Rissman et al.,

1995). Given that GnRH-II was a less efficient stimulator of gonadotropin secretion, it was postulated that its action was mediated through a receptor for which it has a low affinity (Schneider and Rissman, 2008). It is known that GnRH-II has its own specific receptor (GnRHR-II) but it can also bind the GnRHR-I with low affinity (Millar, 2003), potentially indicating why GnRH-II had a lower potency than GnRH-I in these experiments. This is supported by the results of later experiments on sheep (Gault and 26

Lincoln, 2002), musk shrews (Kauffman et al., 2005) and primates (Densmore and Urbanski, 2003; Okada et al., 2003). When given a GnRH-I antagonist (SB-75) prior to administration of GnRH-II, the modest stimulation of gonadotropin secretion or ovulation by GnRH-II was completely attenuated (Densmore and Urbanski, 2003; Okada et al., 2003; Kauffman et al., 2005). Moreover, primary porcine pituitary cultures released LH and FSH in response to GnRH-II, although this effect was mitigated with

SB-75 pre-treatment (Neill et al., 2002a). Thus, it is now generally accepted that only high doses of GnRH-II can stimulate modest gonadotropin secretion through the

GnRHR-I (Densmore and Urbanski, 2003; Okada et al., 2003; Schneider and Rissman,

2008). Therefore, researchers began examining other roles for GnRH-II in mammals.

Given the distribution of GnRH-II in reproductive tissues, many have examined its role in extra-pituitary reproductive function. Research suggests that GnRH-II may have a specific role in ovarian steroidogenesis and implantation/placentation. For example, GnRH-II modulated the action of gonadotropins on the ovary by down- regulating mRNA for the LH and FSH receptors (Kang et al., 2001). Moreover, GnRH-II was detected in the ovary of the baboon, with exogenous administration reducing progesterone production from cultured granulosa cells (Siler-Khodr et al., 2003).

Additionally, studies have shown that GnRH-II is produced in the human placenta and may affect placentation as exogenous administration of GnRH-II to placental explants elicited the production of hCG (Siler-Khodr and Grayson, 2001). Furthermore, GnRH-II was found in the uterine endometrium at all phases of the menstrual cycle with expression increasing during the secretory phase, indicative of a role in implantation 27

(Cheon et al., 2001). Also, invasion of trophoblast cells in the placenta was stimulated by

GnRH-II (Chou et al., 2002; Chou et al., 2003b).

There is also evidence that GnRH-II may impact male reproduction, specifically steroidogenesis and spermatogenesis in the testis. For example, testicular GnRH-II mRNA levels in the black porgy fish increased at maturity (An et al., 2008). In African catfish, GnRH-II either impaired or had no effect on LH stimulated androgen production by the testes (Schulz et al., 1997). However, GnRH-II treatment (5 µg/kg BW) alone raised plasma LH concentrations. Moreover, boars immunized against GnRH-II had reduced serum testosterone production compared to control boars, although LH levels remained unchanged (Bowen et al., 2006). Furthermore, primary Leydig cell cultures from GnRH-II immunized and control animals demonstrated that GnRH-II immunized boars produced less testosterone in response to LH. Additionally, testicular GnRH-II transcripts were increased in azoospermic men, corresponding with elevated intratesticular testosterone levels and increased expression of CYP11A1 and HSD3B2 (Lin et al., 2008), suggesting the increase in transcripts may be compensatory. Consistent with this, GnRH-II transcripts were detected in human sperm by in situ hybridization, suggesting a role for GnRH-II in spermatogenesis (van Biljon et al., 2002). In contrast, a study in mice concluded that GnRH-II did not impact spermatogenesis (Khan et al.,

2007). However, mice lack both endogenous GnRH-II and its receptor (Stewart et al.,

2009), therefore, the actions of GnRH-II in this study were likely due to the interaction of

GnRH-II with the GnRHR-I.

Role in Behavior. Given the neural location of GnRH-II, many early papers hypothesized that it may be important in animal behavior (Muske et al., 1994; Rissman et al., 1995; King and Millar, 1995). The first evidence demonstrating a role for GnRH-II in female reproductive behavior was in 1997 (Maney et al., 1997). Investigators found that GnRH-II, but not GnRH-I, infusions into the brain increased female sparrow receptivity behavior to male song. Later, the effects of GnRH-II were examined in feed restricted and ad libitum fed, female musk shrews. Feed restricted musk shrews displayed fewer sexual behaviors and had reduced GnRH-II mRNA levels in the brain

(Kauffman et al., 2006). Moreover, central infusions of GnRH-II restored the sexual behavior of feed restricted, but not ad libitum fed, females within minutes. Conversely,

GnRH-I only stimulated behaviors in ad libitum fed animals (Temple et al., 2003a).

Interestingly, GnRH-II mRNA levels were not lowered in feed restricted males and sexual behaviors were not altered by feed restriction or GnRH-II administration

(Kauffman et al., 2006). This is potentially due to the sexually dimorphic expression pattern of GnRH-II in the brain of musk shrews or the level of feed restriction (Schneider and Rissman, 2008). In a subsequent experiment on female musk shrews, restoration of sexual behaviors in feed restricted animals was not altered by use of a GnRHR-I antagonist, suggesting the effect was likely mediated by the GnRHR-II (Kauffman et al.,

2005). This is in agreement with later studies on female marmoset monkeys indicating that GnRH-II and a compound that acts as both a GnRH-II agonist and GnRHR-I antagonist (135-18) increased proceptive (sexual solicitation) behaviors, regardless of nutritional plane (Barnett et al., 2006). The results of these studies demonstrates that 29

GnRH-II influences female reproductive behavior through the GnRHR-II, not GnRHR-I.

Moreover, GnRH-II may be acting as a permissive neurotransmitter to reproductive behaviors based on nutritional status in some species (Temple et al., 2003b; Kauffman et al., 2005).

Role in Nutrition. Since the stimulatory effect of GnRH-II on reproductive behavior was only observed after feed restriction in some species, GnRH-II was examined for a role in nutritional balance. As previously indicated in musk shrews, feed restriction reduced reproductive behaviors and GnRH-II mRNA levels within the ventromedial nucleus midbrain, midbrain-hindbrain boundary, and periaqueductal gray areas of the brain. After only 90 min of ad libitum feeding post-restriction, sexual behaviors and GnRH-II mRNA levels in the ventromedial nucleus midbrain and midbrain-hindbrain boundary rebounded, whereas mRNA levels in the periaqueductal gray area remained suppressed (Kauffman et al., 2006), suggesting that energy balance can regulate both transcription and translation of the GnRH-II gene (Schneider and Rissman, 2008). In underfed musk shrews, intracerebroventricular infusions of

GnRH-II reduced feed intake (33%). Similarly, this was also demonstrated in ad libitum fed musk shrews, only to a lesser extent (28%; Kauffman and Rissman, 2004b).

Regardless of nutritional plane, altered feed intake was acute, beginning 90 min after

GnRH-II infusion and persisting for 3 h (Kauffman and Rissman, 2004b). This effect was thought to be mediated solely through the GnRHR-II as inhibition of the GnRHR-I with a GnRH-I antagonist did not prevent reductions in feed intake (Kauffman et al., 30

2005). Therefore, if energy balance is low, GnRH-II production reduces, inhibiting reproductive behaviors and increasing feed intake. In contrast, if energy is abundant, increased GnRH-II production will favor mating (Kauffman and Rissman, 2004a).

Role in Cancer. It is widely accepted that continuous administration of GnRH-I inhibits the proliferation of reproductive tumor cells, typically through Gi activation

(Grundker et al., 2002a; Grundker and Emons, 2003; Limonta et al., 2003). Recent evidence suggests that GnRH-II may also function to regulate cell proliferation and migration in a variety of reproductive cancers (Grundker and Emons, 2003). Indeed,

GnRH-I and -II mRNA were overexpressed in cancer cells and inhibited ribosomal phosphoproteins, which are needed for proper protein translation (Chen et al., 2002).

Interestingly, administration of exogenous GnRH-II exerted a more potent antiproliferative effect on human breast cancer, endometrial and ovarian cells than GnRH-I

(Grundker et al., 2002a). GnRH-II also more potently mitigates the pro-angiogenic effect of vascular endothelial growth factor (VEGF) in human ectopic stromal cells than GnRH-

I, reducing VEGF production in a dose-dependent manner (Huang et al., 2010).

Moreover, in malignant tissues, endogenous GnRH-II up-regulated matrix metalloprotenases (MPP), key regulators of tumor cell invasion (Ling Poon et al., 2011).

Additionally, treatment with a GnRH-II agonist reduced cell proliferation by inhibiting mitogenic signal transduction via activation of phosphotyrosine phosphate (Grundker et al., 2001). Further, treatment with D-Lys6 GnRH-II inhibited the mitogenic effects of epidermal growth factor (Eicke et al., 2006). In many cases, these effects were not 31 mediated through the GnRHR-I, suggesting a role for GnRHR-II in humans (Grundker et al., 2004a; Maiti et al., 2005; Eicke et al., 2006).

In addition to anti-proliferative actions, GnRH-II analogues may also exert pro- apoptotic effects on cancer cells. Treatment with a GnRH-II antagonist inhibited growth of human cancer cell xenotransplants in nude mice, as well as induced apoptosis in human endometrial and ovarian cancer cells in vitro by the loss of mitochondrial membrane potential and activation of caspase-3 (Fister et al., 2007). Later studies demonstrated that treatment of human granulosa cells with GnRH-II can increase

TUNEL-positive cells via the caspase-dependent intrinsic pathway alone (Hong et al.,

2012), whereas the apoptotic effects of GnRH-I are mediated through both the caspase- dependent and Bcl-2 family-dependent intrinsic pathways (Parborell et al., 2008).

Further, GnRH-II was implicated in initiating apoptosis, disrupting the activity of IGF-1 signaling by attenuation of Akt phosphorylation (Hong et al., 2012). In addition to an anti-proliferative and pro-apoptotic effect, GnRH-II might also regulate cellular degradation. For example, human prostate cancer cells (PC3) treated with a GnRH-II antagonist (Trptorelix-1) displayed increased mitochondrial dysfunction (increased reactive oxygen species and decreased membrane potential) as well as autophagosome formation (Kim et al., 2009). Additionally, Western blot analysis of cell lysates revealed a decrease in Akt phosphorylation as well as increased c-Jun phosphorylation, additional hallmarks of cell autophagy. In conclusion, these data indicate that GnRH-II may have a significant role in cell cycle regulation and GnRH-II analogues could be useful cancer therapies in the future. 32

Gonadotropin-Releasing Hormone Receptor II

Structure. Originally cloned in African catfish (Tensen et al., 1997), GnRHR-II has recently been discovered in mammals (Neill et al., 2001; Millar et al., 2001). While the gene for the human GnRHR-I is on chromosome 4 (Kakar and Neill, 1995), an additional GnRH receptor-like sequence was found on chromosome 1, showing the highest sequence identity (58%) to the fish GnRHR-II. Thus, it was identified as the human GnRHR-II gene (Neill, 2002). The GnRHR-II gene has 3 exons (van Biljon et al.,

2002): exon 1 (920 bp) codes for 4 TM domains, exon 2 (210 bp) contains the sequence for TM domain 5 and exon 3 (420 bp) codes for TM domains 6 and 7 (Neill, 2002). In addition, another truncated GnRHR-II gene has also been identified on chromosome 14

(Neill, 2002), containing only exons 2 and 3 (van Biljon et al., 2002).

The full-length GnRHR-II is 379 amino acids (Fig. 2.2; Tensen et al., 1997) and has only 40% homology to the GnRHR-I (Neill et al., 2001). In early studies, GnRH-II treatment of GnRHR-II transfected cells increased IP3 production, indicating functionality through Gqα coupling (Neill et al., 2001). In contrast, GnRH-I treatment elicited ~400-fold less IP3 production, demonstrating that GnRHR-II is highly specific to

GnRH-II. Later studies determined that residues in positions 7 and 8 of GnRH-II are critical for the selectivity of the GnRHR-II (Wang et al., 2003). Similar to GnRHR-I, this receptor is also a 7-TM, GPCR (Tensen et al., 1997). However, the GnRHR-II has

Asp/Asp (N/D) microdomains in TM regions 3 and 7 (Tensen et al., 1997), whereas the

GnRHR-I has Asn/Asp in TM regions 2 and 7, thought to be important for receptor 33 activation (Flanagan et al., 1999). Moreover, in the GnRHR-II, the LSD/EP (Leu-Ser-

Asp/Glu-Pro) amino acid sequence of ECL 3, which is important for ligand selectivity of the GnRHR-I, is replaced with VPPS (Val-Pro-Pro-Ser). This indicates the sequence

VPPS in ECL 3 is important for the selectivity of GnRH-II, as all other known ligand binding sites are conserved (Troskie et al., 1998).

Unlike GnRHR-I, the GnRHR-II maintains a C-terminal, intracytoplasmic tail

(Tensen et al., 1997), indicative of differential coupling for signal transduction (Millar,

2003) and suggesting rapid receptor internalization and desensitization (Heding et al.,

1998; McArdle et al., 2002). Specifically, GnRHR-II internalization rates were dependent on serine residues 338 and 339. Additionally, the tail may aid in receptor expression and regulation given that the addition of the 51 amino acid tail to the rat

GnRHR-I in transfected cells elevated receptor expression 5-fold and augmented IP production (Lin et al., 1998). The process of internalization also differs between the 2 receptors, with internalization occurring through clathrin-coated pits, caveolae and β- arrestin for GnRHR-I, but independent of β-arrestin for GnRHR-II (Pawson et al.,

2003a).

Species Distribution. Since the discovery of the GnRHR-II gene in fish (Tensen et al., 1997), the sequence has been cloned in other non-mammalian vertebrates like birds

(Shimizu and Bedecarrats, 2006), amphibians (Troskie et al., 1997) and reptiles (Troskie et al., 1998). The GnRHR-II gene has also been found in mammals including the human

(White et al., 1998), pig, cow, chimpanzee (Morgan et al., 2003), sheep (Gault et al., 34

2004b), shrew, African elephant, horse, dolphin, rabbit, guinea pig, squirrel, dog and cat

(Stewart et al., 2009), as well as old world monkeys such as the rhesus, marmoset,

African green and macaque (Millar et al., 2001; Neill et al., 2001). In contrast, the

GnRHR-II gene is not present in the genome of the mouse and only a fragment of exon 1 remains in the rat (Stewart et al., 2009). While the gene is present in many species, some species are unable to produce a functional receptor due to coding errors.

In humans and chimpanzees, the GnRHR-II gene contains a frameshift mutation at codon 10 in exon 1, as well as a premature stop codon in exon 2, which would result in the production of a receptor fragment (Morgan et al., 2003). Although this gene is transcriptionally active, (Pawson et al., 2003), its functionality is controversial (Neill,

2002; Millar et al., 2004). The fragmented protein has been termed the ‘GnRHR-II reliquum,’ spanning the cytoplasmic end of the 5-TM domain to the carboxyl-terminus of the full-length receptor (Pawson et al., 2005). Interestingly, the sequence of the GnRHR-

II reliquum is 80% homologous to the full-length, marmoset GnRHR-II sequence, suggesting it may maintain functionality (Millar et al., 1999). Indeed, there has recently been strong evidence for the functionality of this receptor in humans (Neill et al., 2004).

For example, the GnRHR-II reliquum inhibited production of the GnRHR-I at the level of the nucleus and golgi apparatus (Pawson et al., 2005). Further, in human cancer cells with reduced GnRHR-I numbers as a result of gene knock-down procedures, GnRH-II retains the ability to inhibit cell proliferation (Grundker et al., 2002a; Grundker et al.,

2004b). Moreover, a GnRH-II receptor-like antigen has been found in human placenta and cancer cell lines (Eicke et al., 2005). Further, GnRH-I and -II have differing effects 35 in human decidual stromal cell primary cultures, with GnRH-I increasing and GnRH-II decreasing mRNA and protein expression levels of PAI-1. Like other studies, use of a

GnRH-I antagonist mitigated the effects of GnRH-I, but not GnRH-II (Chou et al.,

2003a).

Given the previous evidence, many hypotheses have arisen for how the GnRHR-II may retain functionality including: 1) counteractive shifts in the reading frame, 2) recoding of stop codons, 3) alternative splicing, 4) alternative protein translation or 5) function via association with the GnRHR-I (Neill et al., 2004). First, a corrective shift in the reading frame, albeit rare, has been demonstrated in mammals and allows for the production of a full-length protein (Namy et al., 2004). The use of an alternative start codon has also been considered, given a similar phenomenon occurs in the translation of the African green monkey GnRHR-II (Neill et al., 2001). Further, a readthrough of the premature stop codon, translating selenocysteine or another amino acid, could allow for the translation of a functional human GnRHR-II (Morgan et al., 2003; Millar, 2003; Neill et al., 2004) and has been shown in eukaryotes (Robinson and Cooley, 1997; Bertram et al., 2001; Namy et al., 2004). Moreover, alternative splicing in the pig (Neill et al.,

2002a) and human (Neill, 2002) produces a 5-TM receptor since the reading frame disruption near the N-terminus is spliced out (Neill et al., 2004). The premature stop codon, however, is still present in the 5-TM transcript, but readthroughs of stop codons are relatively common (Namy et al., 2004). Thus a 5-TM protein may still be translated

(Neill et al., 2004) and functional (Ling et al., 1999a). Additionally, fragments of both the 5- and 7-TM isoforms may reassociate after translation to form a functional protein 36

(Neill et al., 2004). This indeed has been demonstrated with other GPCRs, (Schoneberg et al., 1995; Gudermann et al., 1997). Finally, GnRHR-II fragments from either isoform may interact with the GnRHR-I to produce a functional receptor within humans (Neill et al., 2004). This hypothesis is supported by the work of Pawson and coworkers (2005), finding that GnRHR-II appears to inhibit protein expression of the GnRHR-I.

Like the human, other species also have disruptions in translation of the GnRHR-

II gene (Fig. 2.4). The bovine GnRHR-II gene is non-functional due to frameshift mutations in all 3 exons, in addition to premature stop codons in exons 2 and 3 (Morgan et al., 2006). In sheep, the GnRHR-II gene contains a premature stop codon in exon 1 and a 51-bp deletion in exon 2, preventing the translation of a full-length protein (Gault et al., 2004b). Other gene deletions or mutations disrupt the GnRHR-II gene in the elephant, squirrel, guinea pig, rabbit, horse, cat, dog and dolphin, preventing the production of a functional receptor (Stewart et al., 2009). In contrast, genes without coding errors were found in the orangutan, African green monkey, macaque, marmoset, shrew, kangaroo rat, elephant (Morgan et al., 2003) and pig (Neill et al., 2002a; Neill et al., 2002b; Morgan et al., 2003). Interestingly, the only 4 species known to produce both

GnRH-II and the GnRHR-II are the macaque, marmoset, shrew and pig (Morgan et al.,

2003; Millar, 2005). Thus, the pig is an excellent model to study the interaction between

GnRH-II and its receptor.

Using cDNA from porcine pituitaries, the pig GnRHR-II was cloned and sequenced, showing 90% homology with the monkey GnRHR-II (Neill et al., 2002a;

Neill et al., 2004). Like other species, the porcine GnRHR-II is a 7-TM GPCR with 379 37

Figure 2.4. Diagram of coding exons for GnRHR-II in the human, sheep, cow, pig and rat. Premature stop codons (black boxes) and frame shift mutations (arrows) prevent translation of a full-length receptor in the human, sheep and cow. The rat only maintains a remnant of exon 1. In the pig, 3 exons code for a 7-TM receptor and a 5-TM isoform can be formed by alternative splicing in exon 1. Adapted from Brauer (2009) 38 amino acids, a glycosylated N-terminal extracellular sequence, and a C-terminal, intracellular tail (Neill et al., 2002a). This receptor was found to be functional as GnRH-

II stimulation of COS cells overexpressing the GnRHR-II resulted in IP3 production with an EC50 of 0.5 nM compared to 220 nM for GnRH-I (Neill et al., 2002b). Interestingly, another porcine sequence had 91% homology with a segment of exon 2 in the monkey

GnRHR-II (Neill et al., 2002a). Indeed, a 5-TM receptor was also found in porcine pituitaries, derived from alternative splicing of the 7-TM receptor (Fig. 2.5; Neill et al.,

2002b). Specifically, alternative splicing occurs in exon 1, therefore TM domains 1 and

2 are absent and the extracellular N-terminus couples directly to TM domain 3 (Neill et al., 2004). Interestingly, a 5-TM GnRHR-II has also been described in humans (Neill,

2002). While its functionality has yet to be determined, there is precedent for functional

5-TM, GPCRs (Ling et al., 1999b; Perron et al., 2005; Bokaei et al., 2006; Sanchez et al.,

2012).

Ligand Interaction. Interestingly, GnRH-II can also bind to the GnRHR-I, resulting in ~10% the activity of GnRH-I (Neill, 2002). While GnRH-I can also activate the GnRHR-II, it binds with 100-fold less affinity than GnRH-II (Table 2.2(Millar, 2003).

Therefore, it has been postulated that the actions of GnRH-II may have been inappropriately ascribed to GnRH-I in previous studies (Neill et al., 2004). Moreover, early evidence on the function of GnRH-II may have been misleading, given the issue of receptor binding was not addressed (Neill et al., 2004). For example, administration of

GnRH-II potently induced gonadotropin secretion in primates (Lescheid et al., 1997). 39

II II receptor isoforms. From Neill et al.

transmembrane transmembrane (TM) GnRH

and and 7

Graphical Graphical depiction of the porcine 5

Figure 2.5. Figure (2004). 40

TABLE 2.2. COMPARISON OF RELATIVE BINDING AFFINITIES AND SIGNALING MOLECULES BETWEEN THE MARMOSET GnRHR-II AND HUMAN GnRHR-Ia. Marmoset Human Binding and Coupling GnRHR-II GnRHR-I Binding

GnRH-I 0.02 1.00 GnRH-II 1.00 0.1 GnRH III 0.1 0.1 Coupling Gq/11 + + Ca2+ + + PKC + + ERK1/2 + (protracted) + (transient) p38 + nil JNK nil nil c-Src nil + Receptor internalization Rapid slow Rapid desensitization + nil a Adapted from Millar (2003). 41

However, recent evidence indicates that GnRH-II elicits increases in gonadotropin production through the GnRHR-I in vivo (Neill, 2002) and in vitro (Okada et al., 2003).

Further, the effects of GnRH-I and -II were mitigated with the use of a GnRH-I antagonist (SB-75) in human ovarian cancer cells overexpressing the GnRHR-I (Kim et al., 2006a). Additionally, it has been suggested that the presence of GnRH-II, but not a functional GnRHR-II, in many species may indicate that the GnRHR-I has adopted a mediary role for the actions of GnRH-II through alternative active conformations and signal transduction pathways (Millar et al., 2004).

Cell Signaling. Like GnRHR-I, GnRHR-II couples to Gq/11 to initiate calcium influx and the activation of PKC (Table 2.2; Kang et al., 2000; Millar, 2003; Liu et al.,

2009). Interestingly, GnRHR-II mobilizes intracellular calcium stores without stimulating

IP3 activity; instead, it activates the ryanodine receptor (Maiti et al., 2005). After activation of PKC, GnRH-I and -II differentially stimulate MAPKs (Millar et al., 2001).

In COS-7 cells overexpressing GnRHR-II, GnRH-I transiently activates ERK1/2 and c-

Src, whereas GnRH-II activates ERK1/2 in a prolonged manner as well as p38 MAPK

(Millar et al., 2001; Millar, 2003). This was also demonstrated in OVCAR-3 cells, where

ERK1/2 and p38 MAPK facilitated the effects of GnRH-II (Kim et al., 2004; Kim et al.,

2006b). In human cancer cells, GnRH-II-induced antiproliferative effects were also linked to signaling through ERK1/2 (Kim et al., 2005). In fact, use of PD98059, an inhibitor of the ERK1/2 pathway, reversed GnRH-II activation of ERK1/2. However, neither GnRH-I or -II activates JNK in COS-7 cells overexpressing GnRHR-II (Millar et 42 al., 2001) or OVCAR-3 cells (Kim et al., 2005). Interestingly, the GnRHR-I can initiate different cell signaling pathways based on the analogue, dose, duration of treatment and cell type (Millar et al., 2004). This also appears to be true for the GnRHR-II as GnRH-II altered MMP expression via JNK rather than ERK1/2 in human cytotrophoblasts (Liu et al., 2009). Moreover, GnRH-II stimulated cell proliferation, migration and remodeling in a prostate cancer cell line (TSU-Pr1) but was inhibitory to another (DU-145), suggesting differential signal transduction in alternate cell types (Enomoto et al., 2006). This was also evident in 2 ovarian cancer cell lines, where GnRH-II increased invasion of

OVCAR-3 but not SKOV-3 cells (Chen et al., 2007).

Tissue Expression. Similar to GnRH-II, GnRHR-II is widely expressed throughout the body (Fig. 2.6; Millar et al., 2001). In fact, GnRHR-II mRNA was found in every human tissue tested including brain, pituitary, heart, stomach, intestine, kidney, spleen, skeletal muscle, thymus, lung, liver, pancreas, adrenal, thyroid, placenta, uterus, ovary, breast, prostate and testis (Neill et al., 2001). Additionally, studies utilizing marmoset tissues demonstrated that GnRHR-II mRNA is also present in seminal vesicles, the epididymis and bladder, as well as the hypothalamus and many other areas of the brain (Millar et al., 2001). Relative to GnRHR-II mRNA amounts in the pituitary, expression levels were lowest in the bladder and highest in the testis. Interestingly, testicular GnRHR-II transcripts were increased in infertile men (Lin et al., 2008) and were also found in human sperm and post-meiotic germ cells (van Biljon et al., 2002). 43

sues. Values are expressed as a relative sues. Values fold change over

II mRNA in II mRNA marmoset tis

Tissue expression Tissue of GnRHR

Figure 2.6. Figure mRNA amounts in the pituitary. From Millar Millar al. From et (2001). pituitary. the in amounts mRNA 44

GnRHR-II mRNA has also been detected in many human reproductive cancer cells, such as ovarian (Grundker et al., 2004a), endometrial (Eicke et al., 2006), breast (Chen et al.,

2002) and prostate (Kim et al., 2009). Therefore, like its ligand, GnRHR-II appears to be ubiquitously expressed (Millar et al., 2001).

GnRH-II and its Receptor in Boars. Recent evidence suggests that GnRH-II and its receptor may have a direct role in regulating testicular function in boars. For example, boars immunized against cGnRH-II displayed reduced levels of testosterone compared to control boars (Fig. 2.7; Bowen et al., 2006). Interestingly, LH production in cGnRH-II immunized animals was not different from control boars (Fig. 2.8). Since testosterone is classically regulated by alteration in LH pulses, these results indicate immunization against cGnRH-II impacts testosterone directly at the testis, bypassing the

HPG axis. Consistent with this, primary Leydig cell cultures from GnRH-II immunized boars produced less testosterone after LH challenge than cultures from control boars (Fig.

2.9). In an experiment, boars treated with a GnRH-I antagonist (SB-75; 10 µg/kg BW) daily for 4 d demonstrated suppressed testosterone production throughout treatment, whereas LH levels returned to basal 36 h after the initial treatment (Fig. 1.1). Moreover, testicular explant cultures challenged with hCG in the presence of SB-75 (250 ng/ml) produced less testosterone compared to cultures treated with hCG alone (Fig. 2.10).

These results indicate that LH is not the sole regulator of testosterone production in the boar. Since the GnRHR-I could not be detected in the boar testis (Zanella et al., 2000), it has been postulated that a different mechanism may be responsible for localized 45

Figure 2.7. Plasma testosterone levels in barrows, boars or boars immunized against cGnRH-II or lGnRH-III. Arrows indicate when the booster (B) immunization was administered. Prior to the booster immunization shown (6 wk), animals were also given primary (0 wk) and booster (3 wk) immunizations. From Bowen et al. (2006). 46

1 2 3 4 5 6 7 8 9 10 11 12 13 14 Time (wk) Figure 2.8. Plasma levels of FSH (Panel A) or LH (Panel B) in barrows, boars or boars immunized against cGnRH-II or lGnRH-III. Arrows indicate when either the primary (P) or booster (B) immunizations were administered. From Bowen et al. (2006). 47

Figure 2.9. The effect of increasing LH levels on testosterone concentrations of primary Leydig cell cultures from control boars or boars immunized against cGnRH- a-d II or lGnRH-III. Bars without common letters differ significantly (P < 0.05). From Bowen et al. (2006). 48

Figure 2.10. The effect of increasing hCG levels, in the presence or absence of SB- 75, on testosterone production from testicular explant media following 3 h of culture. Inclusion of SB-75 reduced levels of testosterone compared to the control (P < 0.01). From Zanella et al. (2000). 49 testosterone production. Interestingly, SB-75 can bind the GnRHR-II at high doses, albeit with less potency (Maiti et al., 2003). Given the presence of testicular GnRHR-II mRNA in many species (Millar et al., 2001; Neill et al., 2001; van Biljon et al., 2002; An et al.,

2008), it is plausible that GnRH-II and its receptor may directly regulate testosterone production in the boar.

Reproductive Physiology of the Boar

Testosterone Production. The pig is unique among livestock species with the largest interstitial compartment and highest abundance of Leydig cells within the testis, suggesting a high capacity for steroid production (Fig. 2.11; Fawcett et al., 1973). Indeed, numerous steroids have been detected in venous testicular blood of boars (Table 2.3).

However, the most prominent steroid hormone produced in the testis of boars is of course testosterone (Raeside et al., 2006). Testosterone is produced by the Leydig cells in direct response to LH stimulation (Odell et al., 1974). Briefly, LH secreted from the anterior pituitary gland travels through the blood to bind and activate LH receptors on Leydig cells (Senger, 2012). LH receptors are membrane-bound, GPCRs that function through the G-protein, Gs. Activation causes a conformational change, stimulating a cascade of intracellular events including: 1) activation of AC, catalyzing the conversion of ATP to cAMP; 2) activation of regulatory subunits for protein kinases, causing the dissociation of the catalytic subunits; and 3) induction of intracellular cholesterol availability, transcription of genes needed for steroidogenesis and phosphorylation of proteins already 50

Figure 2.11. Photomicrograph of Leydig cells within the interstitium of the boar testis. Numerous blood vessels are present but little connective tissue or lymphatics. From Fawcett et al. (1973). 51

TABLE 2.3. STEROIDS IN BOAR TESTICULAR VEIN BLOODa. C19 Testosterone Androstenedione Dehydroepiandrosterone Dehydroepiandrosterone sulfate Epiandrosterone sulphate 5-Androstene-3β, 17β-diol sulphate 5α-Androstene-3β, 17β-diol sulphate 11β-Hydroxytestosterone 11β-Hydroxyandrostenedione 19-Hydroxytestosterone 19- Hydroxyandrostenedione 5α-Androst-16-en-3α-ol sulphate 5α-Androst-16-en-3β-ol sulphate 5α-Androst-16-en-3-one 5,16-Androstadien-3β-ol sulphate C18 19-Nortestosterone Oestradiol-17β sulphate Oestrone sulphate a Adapted from Raeside et al. (2006). 52 present in the cell by protein kinases. The resulting products include steroid-acute regulatory protein (StAR), which delivers cholesterol to the mitochondria, as well as steroidogenic enzymes (Hunzicker-Dunn and Birnbaumer, 1985).

The initial step in steroidogenesis occurs in the mitochondria where the side-chain of cholesterol is cleaved by P450 side-chain cleavage (P450scc or CYP11A1) enzyme, resulting in pregnenolone. Steroidogenesis can then proceed through 1 of 2 metabolic pathways, Δ4 or Δ5 (Fig. 2.12). The pathway utilized is determined based on enzymatic competition between 17α-hydroxylase (P450c17; CYP17A1) and 3β-hydroxysteroid dehydrogenase (3β-HSD) for pregnenolone, as well as availability of Δ4/Δ5 intermediates and the relative substrate (Conley and Bird, 1997). Interestingly, humans, primates, sheep and cattle can only utilize the Δ5 pathway, whereas mice, rats and pigs can utilize either the Δ4 or Δ5 pathway (Conley and Bird, 1997). For example, in COS-1 cells expressing porcine P450c17, active intermediates of both Δ4 and Δ5 were present

(Conley, 1992). In the boar, testosterone metabolism proceeds via the Δ5 route of steroidogenesis (Gower, 1972). Therefore, pregnenolone is oxidized to 17α- hydroxypregnenolone (17OH-P5) by P450C17 (CYP17A1) and then further cleaved to dehydroepiandrosterone (DHEA). DHEA is dehydrogenated via 3β-HSD for conversion into androstenedione. Finally, androstenedione is converted into testosterone by 17β- hydroxysteroid dehydrogenase (17β-HSD; Brooks and Pearson, 1986; Cooke, 1991).

Spermatogenesis. Within the seminiferous tubules are Sertoli cells, joined together by tight, occluding and gap junctions (Osman and Plöen, 1978). In the boar, 53

Δ4 Pathway Cholesterol Δ5 Pathway

P450scc

P5 3β-HSD CYP17A1

P4 17OH-P5

CYP17A1 CYP17A1

17OH-P4 DHEA

CYP17A1 3β-HSD

17β-HSD

Figure 2.12. The Δ4 and Δ5 pathways of testosterone production. Enzymes: P450 side-chain cleavage (P450 ), Cytochrome P450 17α-hydroxylase (CYP17A1), 3β- scc hydroxysteroid dehydrogenase (3β-HSD) 17β-hydroxysteroid dehydrogenase (17β- HSD). Products: Pregnenolone (P ), Progesterone (P ), 17α-hydroxyprogesterone 5 4 (17OH-P ), 17α-hydroxypregnenolone (17OH-P ), dehydroepiandrosterone (DHEA), 4 5 androstenedione (A ) and testosterone (T). Adapted from Conley and Bird (1997). 4 54

Sertoli cells comprise ~25% of the volume density of each tubule (Franca et al., 2005).

Generally in mammals, the lower the proportion of Sertoli cells within the seminiferous epithelium, the greater the Sertoli cell efficiency and therefore, the higher the daily sperm production (Russell and de Franca, 1995). Sertoli cells encase developing germ cells and aid in spermatogenesis by providing a variety of functions including: 1) protection by compartmentalization, 2) secretion of fluids, proteins and growth factors, 3) phagocytosis of degenerating germ cells and excess cytoplasm from developing germ cells, and 4) coordinated release of spermatozoa into the lumen of seminiferous tubules (Franca et al.,

2005). Additionally, hormones produced by both Sertoli and Leydig cells help regulate the spermatogenic process, such as testosterone, FSH, estrogen and inhibin (Franca et al.,

2005). Similar to all mammals, the spermatogenic process in pigs is comprised of 3 phases: spermatogonial (proliferation), spermatocytary (meiosis) and spermiogenic

(differentiation; Fig. 2.13; Pelliniemi, 1975; Eddy, 2002). Each phase lasts approximately

2 wk, therefore, the total duration of spermatogenesis is about 40 d in the boar (Franca and Cardoso, 1998).

As with other mammals, germ cells in the boar are arranged in segmental stages, or 1 stage per tubular cross-section (Franca et al., 2005). Boars have 4 types of spermatogonia: undifferentiated type A, differentiated type A, intermediate and type B

(Frankenhuis et al., 1982), with type B spermatogonia displaying the largest amount of visible chromatin (Frankenhuis et al., 1982). In boars, like rodents, the undifferentiated type A spermatogonia can be further subdivided into 3 categories based on their topographical arrangement: single (As), paired (Apr) and aligned (Aal; Frankenhuis et al., 55

Graphical Graphical depiction of spermatogenesis. Numbers indicate the number of cells produced by each

Figure Figure 2.13. (2012). Senger division. From cellular 56

1982). Moreover, As spermatogonia are thought to be true spermatogonial stem cells

(Frankenhuis et al., 1982). In the beginning of each spermatogenic cycle, spermatogonial stem cells (2n) divide into the different types of spermatogonia (mitosis). The number of divisions is species-specific and in the boar there are 6 spermatogonial generations (A1-

A4, I, B; de Rooij and Russell, 2000). Several substances influence spermatogonial proliferation, differentiation and apoptosis, such as glial cell line-derived neurotrophic factor, stem cell factor, retinoic acids, p53, Bax, and Bcl-xL (de Rooij and Russell,

2000). Moreover, significant germ cell loss (atresia) takes place during this phase in the pig, such that only 2-3 spermatozoa out of a possible 10 are produced from each differentiated type A spermatogonia. However, this loss is thought to ensure an optimal germ cell to Sertoli cell ratio during proliferation (de Rooij and Russell, 2000).

Throughout the cycle, cells of the same type undergo synchronous division in the same areas of the seminiferous tubule (Berndtson, 1977). Although the cause of this synchrony is unknown, the cytoplasmic bridges joining germ cells in the same stage of development are potentially involved (Fawcett et al., 1959).

The spermatocyte stage (meiosis) is characterized by the start of meiotic divisions of type B spermatogonia (Frankenhuis et al., 1982). Apoptosis is common during meiosis, with losses up to 15% in the boar, likely due to chromosomal defects (Franca and Cardoso, 1998). The division from a primary to secondary spermatocyte ensues next, forming a haploid, round spermatid. At this stage, Sertoli cells are still supporting the spermatids. In fact, in the boar, each Sertoli cell can support 12-30 spermatids at a time

(Franca et al., 2005). Haploid, round spermatids now undergo differentiation, which 57 includes 4 steps: the golgi, cap, acrosomal and maturation phases (Berndtson, 1977;

Senger, 2012). During these phases, nuclear and cytoplasmic structures undergo intricate morphological, biochemical and physiological modifications, indicating that elongation of the haploid spermatid is governed by transcriptional and translational mechanisms

(Steger, 1999). Once differentiated, the spermatozoa are released from the Sertoli cells

(spermiation) which is potentially mediated through cross-talk between cell types (Osman and Plöen, 1978) as well as testosterone and growth factors (Parvinen and Ventela,

1999).

Differences Between Chinese Meishan and White Crossbred Boars. Chinese

Meishan (MS) pigs were imported into the United States in 1989 for reproduction research as they are a highly prolific breed, averaging 3-5 more pigs per litter than the conventional, white crossbred (WC) pigs typically utilized in industry (Haley et al.,

1995). In addition, MS boars reach puberty at about 56-84 d of age, considerably earlier than WC males (120-180 d). In addition, testicular weight is significantly lower in MS

(30-60 g) versus WC (160-240 g) animals at puberty. At maturity, MS maintain a lighter body weight and smaller testicular mass, about half the size of WC. Furthermore, mature

MS boars produce approximately half the number of spermatozoa per ejaculate compared to WC males (Borg et al., 1993). Interestingly, the sperm production per gram of testicular tissue does not differ between MS and WC boars (Okwun et al., 1996b). This is due to differences in the testis composition of MS animals. Within the testis, MS boars have fewer Sertoli cells; however, each Sertoli cell produces twice as many spermatozoa 58 compared to WC animals (Okwun et al., 1996a). Meishan boars also have fewer Leydig cells than WC, although their size is dramatically larger, 2,855 ± 124 µm3 compared to

3 960 ± 49 µm , respectively (Lunstra et al., 1997).

In addition to physiological differences, MS males also have altered endocrine profiles at maturity. Compared to conventional breeds, MS pigs have higher overall gonadotropin and androgen levels (Lunstra et al., 1997). For example, MS boars have 8- fold higher FSH levels compared to Duroc animals (Borg et al., 1993). Like WC pigs, gonadotropins increase in response to GnRH-I treatment, however, levels were sustained longer in MS than WC boars, suggesting an expanded secretory phase following GnRH-I treatment or slower clearance from the body (Wise et al., 1996). In the same study, testosterone levels were higher in MS compared to WC boars, but post-castration testosterone levels decreased similarly. Moreover, the administration of testosterone propionate or estradiol cypionate caused a similar rate of reduction in gonadotropin secretion for both breeds; however, the hormone levels of MS pigs rebounded from suppression earlier than WC boars (Wise et al., 1996). This indicates that the HPG axis in the Meishan boar functions normally, only with higher overall hormone levels (Lunstra et al., 1997), thought to be due to differential pituitary function (Wise et al., 1996).

Specific Antagonists of GnRH Isoforms

SB-75. Cetrorelix or SB-75 is a highly specific GnRH-I antagonist (Bokser et al.,

1990). Indeed, SB-75 has been shown to have ~ 20-fold greater affinity for the GnRHR-I than GnRH-I in a mouse fibroma cell line transfected with GnRHR-I (Beckers et al., 59

1995). The 4 amino acid substitutions in this highly modified decapeptide are credited with enhancing its binding affinity for GnRHR-I and thus, its function (Bokser et al.,

1990). In general, antagonists function through competition with endogenous hormones, occupying receptors to prevent the binding and action of the native hormone (Millar,

2005). Receptor binding assays revealed that SB-75 reduced GnRHR-I numbers via desensitization and down-regulation (Halmos et al., 1996). In rats, 100 µg of SB-75 caused receptor down-regulation in 2 h, reaching a maximal reduction at 6 h (Halmos and Schally, 2002). When SB-75 was given prior to GnRH-I agonist administration, production of LH was depressed or even ablated, primarily due to minimal available binding sites in the pituitary for GnRH-I (Pinski et al., 1992).

Through both occupancy and down-regulation, SB-75 has a vast effect on LH and subsequent testosterone secretion. Indeed, a single infusion of SB-75 (10 µg/kg BW) to castrated rats reduced LH concentrations 80% by 4 h, with levels rebounding after 24 h.

Higher doses were found to further extend suppression (Bokser et al., 1991). In monkeys treated with various doses of SB-75, the serum LH concentration nadir was achieved at

12 h post-injection and persisted for 96 h (Weinbauer and Nieschlag, 1993). Daily treatment with SB-75 (400 µg/kg BW) for 14 d suppressed testosterone production to a level similar to castration (Weinbauer and Nieschlag, 1993). Moreover, extended daily suppression (7 wk) of intact male monkeys resulted in constant suppression of both LH and testosterone. As a result, testicular volume was reduced and animals became azoospermic, although animals regained normal fertility after cessation of treatment

(Weinbauer et al., 1994). 60

SB-75 has also been examined in human trials. Healthy males given a single subcutaneous injection of SB-75 (1, 2 or 5 mg) displayed maximal testosterone suppression after 8-12 h (73, 80 and 91% reduction, respectively). Serum testosterone concentrations returned to basal after 24 h in the 1 and 2 mg treatment groups, whereas testosterone levels did not rebound until 48 h following treatment with 5 mg of SB-75

(Behre et al., 1992). Multiple doses of SB-75 (0.25 - 10 mg) were also given to healthy males over 7, 8 or 14 d. Suppression of testosterone was achieved for all treatment levels, however, only the highest dose (10 mg) maintained testosterone reduction for the duration of the experiment. Lower doses allowed testosterone to rebound after only 2-4 d of treatment, despite continued SB-75 administration (Behre et al., 1994). In subsequent studies, investigators determined that an elevated initial dose (10 mg) followed by maintenance treatments (1 mg) suppressed testosterone secretion throughout treatment

(Behre et al., 1997). In conclusion, SB-75 is a potent GnRH-I antagonist that can sufficiently suppress LH release and subsequent testosterone synthesis in a dose- and time-dependent manner.

Trptorelix-1. A specific GnRH-II antagonist [Ac-D2Nal - (4 Cl) D-Phe-D-2Pal-

Ser-Tyr-D-Cit-Trp-Tyr-Pro-D-AlaNH3], called Trptorelix-1 (Trp-1), has recently been developed (Seong and Cheon, 2007). It was designed based on the structure of SB-75 by substituting amino acids 7 (Trp for Leu) and 8 (Tyr for Arg; Seong and Cheon, 2007), which is critical to selectivity for the GnRHR-II over GnRHR-I (Wang et al., 2003). GH3 cells transiently transfected with bullfrog GnRHR-II or rat GnRHR-I were pretreated 61 with GnRH-I prior to SB-75 or Trp-1 treatment. Both antagonists decreased GnRH- induced IP production in each cell line. However, Trp-1 was 50-fold more potent than

SB-75 at inhibiting GnRHR-II activity. Conversely, SB-75 was 30-fold more effective at reducing GnRHR-I activity than Trp-1 (Table 2.4). Moreover, membrane-binding assays revealed that SB-75 exhibited a 12-fold higher binding affinity for GnRHR-I than Trp-1

(Maiti et al., 2003). Similarly, CV-1 cells transfected with African green monkey

GnRHR-II were pretreated with GnRH-II prior to Trp-1 or SB-75 treatment (Wang et al.,

2003). Trp-1, but not SB-75, completely blocked activation of the GnRHR-II. These data suggest that Trp-1 is highly selective for GnRHR-II and unlike SB-75, effectively inhibits

GnRHR-II function. 62

TABLE 2.4. MEAN INOSITOL PHOSPHATE IC50 VALUES IN GH3 CELLS OVEREXPRESSING EITHER GnRHR-I or GnRHR-II FOLLOWING TREATMENT WITH SB-75 OR TRPTORELIX-1 IN THE PRESENCE OF 1 nM GnRH-IIa. Mean IC50 values (± SEM) for Inositol Phosphateb

Receptor Trptorelix-1 SB-75

GnRHR-I -9.27 ± 0.07 (0.03) -10.74 ± 0.12

GnRHR-II -6.05 ± 0.17 Partial Inhibition a Adapted from Maiti et al. (2003). b The relative fold change to SB-75 activity is indicated in parentheses.

CHAPTER III

MATERIALS AND METHODS

Localization of the GnRHR-II

Tissue Collection. Animal trials were conducted in accordance with the U.S.

Meat Animal Research Center (USMARC; Clay Center, Nebraska) Animal Care and Use

Committee. Mature Chinese Meishan boars (n = 5) were housed individually at

USMARC and fed 6 lb of feed daily with water available ad libitum. At the USMARC slaughter facility, boars (15-17 wk of age) were rendered unconscious by CO2 inhalation, exsanguinated and the testes were collected. Next, the head was removed and a lateral cranial incision was made, revealing the hypothalamus and anterior pituitary gland within the brain for excision. Tissues were snap frozen in liquid nitrogen and stored at -80°C for later protein extraction. Additional sections were collected for immunohistochemistry

(IHC) by submerging in 4% paraformaldehyde. Samples were maintained on ice for transport and stored at 4°C overnight. The following morning, samples were placed in cassettes (Fischer Scientific; Pittsburgh, PA) and stored in 70% ethanol at room temperature (RT) until further processing.

Embedding and Sectioning. For tissue embedding, cassettes containing testicular samples were removed from 70% ethanol and rinsed in 100% ethanol 3 times (1 h each) to remove water and further dehydrate the tissue. Cassettes were then rinsed 3 64 times (1 h each) in CitriSolv (Fischer Scientific). In the case of both the ethanol and

CitriSolv, the last wash was always in fresh solution. Next, cassettes were placed into paraffin overnight in a TISSUE-TEK III embedding machine (Miles Scientific, Sahura

Fine Technical Company, Naperville, IL). The following morning, tissues were removed with forceps and placed in metal forms. Fresh, hot paraffin was poured over the tissue and the back of the cassette was laid into the mold. After chilling for 2 h, the cooled paraffin could be removed from the mold and stored at RT.

After embedding, the tissue was sectioned by first cutting the paraffin mold into a trapezoid. Next, 5 µm sections were cut on a Spencer “820” Microtome (American

Optical Company, Buffalo, NY) and heated in a 37°C water bath to smooth the paraffin.

Approximately 6 sections were then moved onto an UltraStick slide (Gold Seal Products,

Portsmouth, NH) and dried on a hot plate (37°C) overnight. After drying, the slides were stored in a slide box until further processing.

Immunohistochemistry. Deparaffinization was achieved by washing slides twice through CitriSolv and twice through 100 % ethanol, for 10 min each. Thereafter, slides were rinsed for 5 min through 95%, 70%, 50% and 30% ethanol, respectively. Heat induced epitope retrieval was achieved by boiling slides in 0.01 M sodium citrate for 15 min. Slides were then allowed to cool for 1 h and rinsed 3 times in 1X Tris-buffered saline (TBS; 25 mM Tris, 150 mM NaCl, 2 mM KCl) for 5 min. After rinsing, individual tissue sections were isolated with a PapPen (Research Products International Corp.,

Fredericksburg, VA). Next, the VECTASTAIN ABC-AP kit and its reagents (Vector 65

Laboratories, Burlingame, CA) were used per the manufacturer’s instructions. Briefly, slides were blocked with 10% rabbit serum for 20 min, rinsed briefly in 1X TBS and excess fluid was wicked away with filter paper. Next, the primary antibody, anti-

GnRHR-II [goat polyclonal, 1:500 (SC- 162889; Santa Cruz Biotechnology, Inc.; Santa

Cruz, CA)] was diluted (1:100) in 1X TBS and incubated on sections for 30 min in a humidified chamber. After incubation, sections were rinsed for 5 min in 1X TBS. Then, slides were incubated with the diluted (1:200) biotinylated, anti-goat IgG (H+L) secondary antibody provided. Again, slides were rinsed in 1X TBS for 5 min. Next, sections were incubated with the ABC-AP reagent, diluted 1:200. Slides were again rinsed in 1X TBS for 5 min before the final step, incubation in the alkaline phosphatase substrate solution for 30 min. Slides were washed for a final 5 min in tap water. Excess water was wicked away and slides were mounted as follows. Approximately 1 drop per section of plain Tris-buffered 80% glycerol (warmed to RT) was applied and the sections were cover slipped (Fischer Scientific). Slides were sealed with clear nail polish and images were obtained using an Olympus (Center Valley, PA) microscope (BX51) and camera (DP71).

Immunohistochemical Validation. The use of the GnRHR-II antibody [goat polyclonal, 1:50 (SC- 162889; Santa Cruz Biotechnology)] in IHC was validated prior to data collection. It was determined by protein BLAST (accession number: AS68622) that the epitope for this antibody maintained 85% homology for the sus scrofa GnRHR-II, specifically the 7-TM isoform. This was also verified by the antibody manufacturer. 66

Further, several concentrations of antibody were examined to test specificity on boar testis. The concentration curve was as follows: negative control, 1:200, 1:100, 1:50, 1:25 and 1:10 (Fig. 3.1). Staining was immediately apparent in the 1:200 dilutions and became more intense with higher concentrations. The concentration 1:50 was selected because it was the most dilute concentration exhibiting plasma membrane specific staining on

Leydig cells.

Protein extraction. Protein was extracted from testis and anterior pituitary gland samples by homogenizing tissue in 1 ml of RIPA buffer (20 mM Tris, 137 mM NaCl,

10% glycerol, 1% NP40, 0.1% SDS, 0.5% deoxycholic acid, 2 mM EDTA, 1mM PMSF,

1% protease inhibitor cocktail and 1% phosphatase inhibitor cocktail) with a Biospec

Tissue Tearor (Bartlesville, OK) until completely homogenized. Then, cells were further lysed by use of a Dounce Homogenizer (Wheaton Inc., Millville, NJ) 3 times. Samples were incubated for 30 min on ice before centrifugation (12,000 x g) at 4ºC using a

Beckman (Palo Alto, CA) TJ-6 centrifuge. Samples were aliquoted into fresh, pre-chilled tubes and stored at -80ºC.

The protein concentrations of the testicular lysates were quantified using a bicinchoninic acid (BCA) assay (Pierce Biotechnology, Rockford, IL). The standard curve was comprised of 9 standards, diluted per manufacturer’s instructions. The unknowns were compared to the standard curve to determine a relative protein concentration. Briefly, 25 µl of standards or unknowns were pipetted onto a microplate

(Sarstedt, Inc., Newton, NC) in duplicate. Working reagent (200 µl) was pipetted into 67

tes tes using multiple concentrations of an antibody directed against the

Immunohistochemistry Immunohistochemistry of boar tes

II (Negative control, 1:200, 1:100, 1:50, 1:25, 1:10, respectively). All images shown are shown 20x at are images All magnification. 1:10, control, respectively). 1:200, (Negative 1:50, 1:100, 1:25, II

Figure Figure 3.1. GnRHR 68 each well of the plate. The reagent was prepared by mixing 50 parts BCA Reagent A to 1 part BCA Reagent B for a final ratio of 50:1, Reagent A:B. The samples were mixed well on a plate shaker for 30 s, then covered due to light sensitivity and incubated at 37ºC for

30 min. Next, the microplate was allowed to cool and light absorbance was measured at

562 nm on a Spectra Max 250 plate reader (Molecular Devices, Sunnyvale, CA).

Protein samples were then prepared for use in Western blot analyses.

Dithiothreitol (DTT) was mixed with 4X loading dye [LD; 2% Tris (pH 6.8), 28% glycerol, 20% SDS and a fleck of Orange G] to a ratio of 10:50 (DTT:LD). This mixture was then added to the protein. Samples were vortexed and centrifuged (12,000 x g) for 2 min. Next, samples were boiled for 5 min at 100ºC, vortexed and centrifuged again briefly. Samples were then aliquoted and stored at -80ºC until use.

Western blot. The gel was prepared by assembling BIO-RAD glass plates

(Hercules, CA) into a BIO-RAD casting system. First, acrylamide (10%) was mixed with

4X Tris-Cl/SDS (pH 8.8; 0.5M Tris-Cl, 0.4% SDS) and 3.5 ml millipore water in a 50 ml conical tube. Then, 0.105% ammonium persulfate and 0.75% TEMED were added and the solution was inverted slowly to prevent bubble formation. Next, the solution was poured into the plates to an approximate height of 4 cm. A thin layer of isobutanol- saturated water was applied directly on top of the aqueous gel layer to allow for complete polymerization. After approximately 15 min, the gel had polymerized completely and the isobutanol-saturated water was removed and the gel was rinsed with deionized water 3 times. The water was wicked away with filter paper and the stacking gel was then poured. 69

The stacking gel consisted of 4% acrylamide, 4X Tris-Cl/SDS (pH 6.8; 1.5mM Tris-Cl and 0.4% SDS) and 1.85 ml millipore water (Millipore Corporation, Billerica, MA).

Then, 0.10% ammonium persulfate and 0.833% TEMED were added and the solution was inverted slowly to prevent bubbles and poured on the separating gel until level with the top of the plate. A BIO-RAD 15-well comb was then added and the gel polymerized for 15 min. After complete polymerization, running buffer (25 mM Tris, 192 mM glycine and 0.1% SDS) was added to the center and sides of the BIO-RAD gel box. Next,

4 µl of SDS-PAGE ladder (Thermo Scientific, Waltham, MA) was loaded for comparison of band sizes. Then, 20 µg of protein was loaded into each well. The gel was then run for approximately 1 h (150 V, 30 mA) until good band separation was achieved.

The gel was removed from the glass plates and placed in transfer buffer (48 mM

Tris Base, 39 mM glycine, 0.0375% SDS, and 20% methanol) for 10 min. At the same time, the wells and dye front were removed from the gel and its dimensions were measured. Next, 4 pieces of filter paper (0.34 mm thick) and 1 piece of polyvinylidene fluoride membrane (PVDF; Millipore Corporation, Billerica, MA) were cut to the exact size of the gel. The filter paper (2 pieces) were soaked in transfer buffer and stacked directly on top of each other on an Owl Separation System (Thermo Scientific), followed by the gel, PVDF membrane (pre-wetted in 100% methanol) and 2 more pieces of filter paper. A roller was used between each layer to smooth the stack and remove air bubbles that could interfere with the transfer. Transfer buffer was added to the trough of the transfer apparatus, and excess moisture was removed from the plate and stack. The protein transfer ran for 1.5 h (200 mA, 20 V). 70

After transfer, the blot was rinsed in millipore water and nonspecific binding was blocked by incubation of the membrane in Odyssey Blocking Buffer (Licor Biosciences,

Lincoln, NE) for 1 h at RT. Then, blotting membranes were incubated with a goat polyclonal primary antibody directed against GnRHR-II (1:1000; SC-162889; Santa Cruz

Biotechnology), diluted in Odyssey Blocking Buffer with 0.05% Tween-20. Blots were left to incubate shaking at 4ºC overnight. After washing 4 times (5 min each) in 1X TBS-

T [TBS (20 nM Tris base, 137 mM NaCl, pH 7.6) and 0.001% Tween-20], blotting membranes were incubated with a secondary donkey anti-goat antibody (1:1600; Alexa

Fluor® 680; Invitrogen; Carlsbad, CA) diluted in Odyssey Blocking buffer for 1 h at RT.

The membranes were then washed 4 times in 1X TBS-T and 4 times in 1X TBS. Blotting results were detected with the Odyssey Scanner and analyzed using Odyssey image software (Licor Biosciences).

After scanning, membranes were stripped with NewBlot stripping buffer (Licor

Biosciences) for 30 min at RT. Membranes were then blocked for 15 min and reprobed with the primary antibody, β-actin (1:2000; SC-1516; Santa Cruz Biotechnology), for 1 h at RT. After washing 4 times in 1X TBS-T, blotting membranes were incubated in light- tight boxes with the secondary donkey anti-goat (1:1600; Alexa Fluor® 680; Invitrogen) diluted in Odyssey Blocking buffer for 1 h at RT. The membranes were then washed 4 times in 1X TBS-T and 4 times in 1X TBS. Again, fluorescence was detected with the

Odyssey Scanner and analyzed using Odyssey image software. β-actin served as the internal control for Western blot analyses. Samples were analyzed, normalized to β-actin and then expressed as a fold change. 71

Semen Collection. Semen was collected from mature white crossbred boars

(UNL Index line; n = 5) using the gloved hand method. Semen was transferred to a 50 ml conical tube and centrifuged (800 x g) for 10 min at RT. Seminal plasma was removed and stored at -20°C for later analysis. Then, the sperm pellet was layered on a discontinuous Percoll gradient of a 90% Percoll solution [90% Percoll, 10% Tyrode’s stock (40 mM KCL, 2.92 mM, 0.05% NaH2PO4, 0.02% Hepes buffer, pH 7.3), 1.97 mM

CaCl2, 0.394 mM, 0.002% lactic acid, 0.002% NaHC03] and a 45% Percoll solution

[50% of the 90% Percoll solution and 50% TL sperm solution (100 mM NaCl, 3.1 mM

KCl, 25.0 mM NaHC03, 0.29 mM NaH2Po4H2O, 26 mM Na lactate, 2.107 mM

CaCl22H20, 0.4 mM MgCl6H20 and 10 mM Hepes; pH 7.3)]. Approximately 2 ml of the

45% solution was layered on 2 ml of the 90% solution in a 15 ml conical tube. Then, 1 ml of the sperm pellet was deposited on top and the tubes were centrifuged (800 x g) for 30 min at RT. The Percoll gradients were removed and the live sperm were isolated from the pellet. Purified spermatozoa were diluted in 1X phosphate buffered saline (PBS; 1.36 M

NaCl, 1.08 M KCl, 0.1 M Na2HPO4 and 0.017 M KH2PO4) and air-dried on an UltraStick slide for later immunocytochemistry. Purified sperm were also flash frozen for protein extraction.

Protein extraction. Samples were lysed by shearing through a 21 gauge needle with 200 µl RIPA buffer about 6-7 times. Samples were then passed through a 27 gauge needle 5 times, vortexed and allowed to incubate on ice for 45 min. Thereafter, protein 72 was isolated from cellular debris by centrifugation (12,000 x g) at 4ºC using a Beckman

TJ-6 centrifuge. Samples were aliquoted into fresh, pre-chilled tubes and stored at -80ºC.

Protein quantification and Western blot sample preparation was performed as described above.

Western blot. Western blots of live sperm were performed as previously described. However, the blot was not reprobed for β-actin. The primary antibody used was directed against GnRHR-II [goat polyclonal, 1:1000 (SC-162889; Santa Cruz

Biotechnology)]. The secondary antibody was donkey anti-goat (1:1600; Alexa Fluor®

680; Invitrogen).

Immunocytochemistry. After air-drying, spermatozoa were fixed with 4% paraformaldehyde for 10 min at RT in a humidified chamber. Then, slides were washed twice with 1X TBS for 5 min each. Next, slides were exposed to UV light for 30 min to quench autofluorescence. Cells were permeabilized with 1X TBS-T (0.5% Tween-20) for

10 min at RT. Slides were washed 3 times with 1X TBS for 5 min each. Non-specific binding was then blocked by incubation with 10% normal rabbit serum diluted in TBS-T

(0.045% Tween-20) for 30 min at RT. Next, slides were treated with a polyclonal goat primary antibody directed against the GnRHR-II (sc-162889; Santa Cruz Biotechnology).

This antibody was diluted 1:50 with 10% normal serum in TBS-T (0.045% Tween-20) and slides incubated overnight at 4°C. The following morning, slides were washed with

TBS-T (0.05% Tween-20) 3 times for 5 min each. Then, cells were incubated with a 73 rabbit anti-goat Alexa Fluor® 488 secondary antibody diluted 1:400 in 10% normal serum

(TBS plus 0.045% Tween-20) for 1 h at RT in a dark, moist chamber. Finally, slides were rinsed 3 times with TBS for 5 min each in the dark. Then, slides were mounted with

ProLong Gold Antifade Reagent (Invitrogen), cover slipped and sealed with clear nail polish. Slides were imaged with an inverted (Olympus IX 81) confocal microscope using

FITC filter sets.

ELISA. Isolated seminal plasma as well as testis, anterior pituitary gland and hypothalamus homogenates were assayed for a precursor form of GnRH-II by a commercially available enzyme-linked immunosorbent assay (ELISA) kit, specific to the pig (Cat #: CSE-EL009636PI; Cusabio; Hubei, China). Seminal plasma samples were from mature, white crossbred boars (UNL Index line). Testis, anterior pituitary gland and hypothalamus samples were collected from mature, Chinese Meishan boars, snap frozen and stored at -80°C. Tissue was homogenized per manufacturer’s instructions. Briefly,

100 mg of tissue was rinsed with 1X PBS and then homogenized in 1 ml of 1X PBS, and stored overnight at -20°C. After thawing and freezing again (to break the cell membranes), the samples were centrifuged at (5,000 x g) for 5 min at 4°C. The supernatant was removed and frozen at -80°C until assay. Homogenate samples were thawed fully and centrifuged briefly again prior to use in the assay.

The ELISA was performed according to the manufacturer’s instructions. Prior to assay, all reagents and samples were allowed to warm for 30 min and a 1X wash buffer was prepared. First, the standard curve was added to the plate by pipetting 50 µl of each 74 standard into each well (0, 40, 160, 640, 2,000 and 8,000 pg/ml), in duplicate. An additional standard was also added (20 pg/ml). Two wells were left blank to serve as non-specific binding (NSB) wells. Then, 50 µl of the unknown samples (seminal plasma or tissue homogenates) were added, in duplicate. Immediately following, 50 µl of the provided horseradish peroxidase-conjugate was added to each well (except the NSB wells), followed by 50 µl of the antibody. The plate was shaken gently to mix and then incubated at 37°C for 1 h. Next, each well was aspirated and washed with 200 µl of the

1X wash buffer solution using a multi-channel pipette. The wash buffer was left for 10 s before subsequent aspiration. This was repeated a total of 3 times, ensuring all liquid was decanted between each wash. After the last wash, the plate was inverted and blotted on filter paper to remove any remaining wash buffer. Next, 50 µl of substrate A was added to each well, followed by 50 µl of Substrate B. The plate was then mixed gently on a plate shaker and allowed to incubated in the dark at 37°C for 15 min. Then, 50 µl of the

Stop Solution was added to each well. The reagents were mixed by gentle tapping on the plate. The optical density was immediately read using a Wallac 1420 Victor2 microplate reader (Perkin Elmer; Waltham, MA) set to 450 nm. The data were interpolated based on the standard curve using a four parameter logistic (4-PL) curve fit per the manufacturer's instructions. The minimum sensitivity of the assay was 20 pg/ml. The intra-assay coefficient of variation between pooled samples was 10%.

Testicular Explants 75

Materials. Chinese Meishan boars (n = 7) were transported to the USMARC slaughter facility the morning of collection. Animals were rendered unconscious by inhalation of 100% CO2 and were humanely euthanized by exsanguination. Whole testes were collected, weighed and a random section of the parenchyma was removed and minced finely. Approximately 400 mg of minced tissue was incubated in 50 ml conical tubes (Corning Inc., Corning, NY) with 5 ml of TC199 (with Earle's salts and NaHCO3;

Sigma-Aldrich; St. Louis, MO). The media was buffered with a final concentration of 25 mM Hepes (N-2-hydroxy-ethyl-piperazine-N-2ethane sulfonic acid). Tissue explants were pre-incubated for approximately 2.5 h during transit. Tissue was exposed to a 95%

O2 and 5% C02 atmosphere, maintained at 37ºC and disrupted every 10 min to prevent tissue aggregation. Upon return to UNL, the media was decanted and replaced with fresh

TC199 (25 mM Hepes), and explants were randomly assigned to treatment.

Treatments. At time 0, explant cultures were exposed to either vehicle (water) or

1 µM GnRH-II (Bachem Inc., Torrance, CA). Administration of vehicle or GnRH-II at this time point was considered a pretreatment. Media was collected (250 µl) 1 h after pretreatment from all cultures, then explants were treated. Treatments were either vehicle

(water) or 1 I.U. human chorionic gonadotropin (hCG; Sigma-Aldrich). Therefore, all combinations (pretreatment + treatment) resulted in 4 different treatments: control

(vehicle + vehicle), hCG alone (vehicle + hCG), GnRH-II pretreatment without hCG

(GnRH-II + vehicle) or GnRH-II pretreatment with hCG (GnRH-II + hCG). After treatment, media was collected 120, 180 and 240 min later (Fig. 3.2). Media was 76

PRE TRT

Minutes 0 60 120 180 240

Tissue Collection Figure 3.2. Schematic indicating time of pretreatment (PRE; 0 or 1 µM GnRH-II), treatment (TRT; 0 or 1 I.U. hCG), media collection (hash mark) and tissue collection for testicular explants. 77 immediately frozen after each collection and subsequently stored at -20ºC until further processing. Tissue was snap frozen in liquid nitrogen and stored at -80°C until protein extraction and Western blot analyses.

Protein Extraction. Testis protein was extracted by homogenizing tissue in 1 ml

RIPA buffer with a Biospec Tissue Tearor until completely homogenized. Then, cells were further lysed by use of a Dounce Homogenizer (Wheaton, Inc.) 3 times. Samples were incubated for 30 min on ice before centrifugation (12,000 x g) at 4ºC using a

Beckman TJ-6 centrifuge. Samples were aliquoted into fresh, pre-chilled tubes and stored at -80ºC. Protein concentration was then quantified by BCA assay and samples were processed for use in Western blots, as described previously.

Western Blot. Western blots were performed as described previously. However, for these testis samples, 2 different primary antibodies were used. Either GnRHR-II [goat polyclonal, (1:1000; SC-162889; Santa Cruz Biotechnology)] or LH receptor [rabbit polyclonal (1:300; SC-25828; Santa Cruz Biotechnology)]. The secondary antibodies used were either donkey anti-goat (1:1600; Alexa Fluor® 680; Invitrogen) or goat anti- rabbit (1:1600; Alexa Flour® 800; Invitrogen). As described previously, blots were stripped and reprobed for β-actin to serve as an internal loading control.

Radioimmunoassay. Total testosterone levels were determined using a Total

Testosterone Coat-a-Count RIA kit (Siemens Healthcare Diagnostics, Los Angeles, CA) 78 in accordance with the manufacturer’s instructions. This kit has been previously validated in the pig (Oskam et al., 2010; Ashworth et al., 2011) and prior to use in this trial was examined for parallelism in measuring testis explant media. The minimum sensitivity of the assay was 0.16 ng/ml and the intraassay coefficient of variation was 8%. Briefly, 50

µl of zero calibrator (0 ng/ml testosterone) was added to 2 (duplicate) uncoated 12x75 mm polypropylene tube non-specific binding (NSB) tubes and 2 Total Testosterone Ab-

Coated tubes for maximum binding. Then, 50 µl of each remaining calibrator (B-F; 0.16,

0.96, 4.14, 7.92 and 16.85 ng/ml testosterone, respectively) were pipetted into Total

Testosterone Ab-Coated tubes (Specific binding; Bø) taking care to pipette directly into the bottom of all tubes. Next, media samples were diluted 1:5 with Siemens zero calibrator solution (as per manufacturer’s instructions). This dilution was used based on previous trials. Then, 1 ml of testosterone radioactively labeled with I125 was added to every tube and tubes were covered with parafilm, vortexed for 30 s and incubated in a

37°C waterbath for 3 h. The NSB tubes containing only 1 ml 125I Total Testosterone were set aside for total counts. Following incubation, liquid was decanted thoroughly (except the total counts tubes) and allowed to drain for 2-3 min. Finally, samples were counted using an APEX Automativ gamma counter (ICN Micromedia Systems, Horsham, PA) for

1 min. Percent binding was then determined using the following equation: [(Bø-

NSB)/TC)]*100 = % binding. Data were analyzed and fit to the standard curve using

Graphpad PRISM (San Diego, CA) to determine relative testosterone levels.

Boar Trials 79

Animals. The following animal trial was conducted in accordance with the

USMARC Animal Care and Use of Committee. Mature Chinese Meishan boars were housed individually at USMARC and fed 6 lb of feed daily with water available ad libitum. Mature boars were randomly assigned to treatments. The experiment was performed over a 2 wk time period in a cross-over design, such that each boar received each treatment once, 1 wk apart. Catheterization of boars was conducted according to

Ford and Maurer (1978) at USMARC (Appendix I), approximately 4-5 d prior to initiation of the trial.

Experiment 1: GnRH Agonist Trial. On the day of treatment, blood samples (10 ml) were collected every 20 min for 2 h to obtain a baseline hormonal profile. Then, boars (n = 5/treatment) were given I.V. infusions of either a GnRH-I agonist (D-ala6

GnRH-I; 150 ng/kg BW) or GnRH-II agonist (D-ala6 GnRH-II; 150 ng/kg BW), both dissolved in sterile saline. Blood samples were drawn every 10 min for 30 min and every

20 min for 270 min thereafter (Fig. 3.3). During blood sampling, catheters were kept clear by injecting 2 ml of 3.5% sodium citrate prior to blood draws. Once blood was evident upon drawback, a heparinized vacutainer was utilized to collect 10 ml of blood.

Tubes were inverted slowly to prevent clotting and red blood cell lysis. Following blood collection, 3 ml of heparinized saline was injected and the catheter cap was cleaned with

100% ethanol before being replaced. Blood samples remained at 4°C until centrifugation

(12,000 x g) and plasma was collected and stored at -20°C until analysis. 80

TRT

Minutes -100 -80 -60 -40 -20 0 10 20 30 50 70 90 110 130 150 170 190 210 230 250 270

Figure 3.3. Schematic of boar GnRH agonist trial indicating treatment (TRT) and time of blood sample (hash mark). At 0 min, blood was collected prior to administration of treatment. 81

Testosterone and LH levels were measured by radioimmunoassay (RIA) in Dr. Clay

Lents’ laboratory at USMARC, according to Ford et al. (2001) and Kesner et al. (1987), respectively. Minimum sensitivities were 0.1 ng/ml and 0.1 ng/ml for LH and testosterone, respectively, and corresponding intraassay coefficients of variation for pooled samples averaged 7.3 and 9%, respectively.

Animals. All remaining animal trials were conducted at UNL in accordance with the UNL’s Institutional Animal Care and Use Committee (IACUC). Mature white cross bred boars (UNL Index line) were housed individually with ad libitum access to water and fed approximately 5 lb of feed daily. At least 3 days prior to trial initiation, boars were cannulated in the jugular vein according to either Barb et al. (1982) or Ford and

Maurer (1978).

Experiment 2: GnRH Antagonist Trial. Boars were randomly assigned to be treated with either a specific GnRH-I antagonist (SB-75; 5 µg biologically active compound/kg BW; UNL Protein Core; n = 9), or a specific GnRH-II antagonist,

Trptorelix (Trp-1; 5 µg/kg BW; AnyGen, Gwangju, Korea; n = 10). The experiment was performed over a 2 wk time period in a cross-over design, such that each boar received each treatment once, 1 wk apart. SB-75 was dissolved in 0.9% sterile saline and Trp-1 was dissolved in solution per manufacturer’s instructions. Briefly, Trp-1 was first dissolved in 5% dimethylacetamide, then mixed with 40% propylene glycol and finally diluted in 55% millipore water. 82

Blood (9 ml) was drawn 30 min and immediately prior to treatment. Treatments were then given I.M. in the neck. Each animal also received the vehicle of the opposite treatment. Next, blood sampling occurred every 30 min for 3 h (30, 60, 90, 120, 150, 180 min), at 6 h and at 12 h. Thereafter, blood was taken every 12 h for 48 h (Fig. 3.4).

During blood sampling, catheters were kept clear by injecting 5 ml of 3.5% sodium citrate prior to blood draws. Once blood was evident upon drawback, 9 ml was collected and the cannula was flushed again with 3 ml heparinized saline. The catheter cap was cleaned with 100% ethanol before being replaced. The S-Monvette serum blood tubes

(Sarstedt, Nümbrecht, Germany) utilized were inverted slowly to distribute silicate beads, aiding in clotting and preventing red blood cell lysis. Blood samples remained at 4°C for

1-24 h until centrifugation (2,000 x g), at which time serum was collected and stored at -

20°C until RIA analysis.

Experiment 3: SB-75 + GnRH Agonists Trial. First, blood (9 ml) was collected

20 min, 10 min and immediately prior to treatment. Then, each boar was treated with the

GnRH-I antagonist, SB-75 (5 µg biologically active compound/kg BW; UNL Protein

Core). Approximately 7.5 h later, blood was collected to confirm the activity of SB-75.

Then, boars (n = 5) were given I.V infusions of either a GnRH-I agonist (D-ala6 GnRH-I;

150 ng/kg BW; Bachem) or the GnRH-II agonist (D-ala6 GnRH-II; 150 ng/kg BW;

Anygen). Blood samples were then drawn every 10 min for 30 min and then every 20 min for 4 h (Fig. 3.5). During blood sampling, catheters were kept clear by injecting 2 ml of 3.5% sodium citrate prior to blood draws. Once blood was evident upon drawback, S- 83

TRT

Hour - 0.5 0 0.5 1 1.5 2 2.5 3 6 12 24 36 48

Figure 3.4. Schematic of GnRH boar antagonist trial indicating treatment (TRT) and time of blood sample (hash mark). At 0 h, blood was collected prior to administration of treatment. 84

TRT

Minute s -470 -460 -450 0 10 20 30 60 80 100 120 150 170 190 210 230 250 270

Figure 3.5. Schematic of SB-75 + GnRH agonist trial indicating SB-75 administration, treatment (TRT) and time of blood sample (hash mark). At 0 min, blood was collected prior to administration of GnRH analogues. 85

Monvette serum blood tubes (Sarstedt) were utilized to collect 9 ml of blood. Tubes were inverted following blood collection, 3 ml of sodium citrate was injected and the catheter cap was cleaned with 100% ethanol before being replaced. Blood samples remained at

4°C until centrifugation (12,000 x g) and serum was collected and stored at -20°C until

RIA analysis. This experiment was a crossover design such that each animal received each treatment, 1 week apart.

Experiment 4: Intratesticular GnRH Injection Trial. Approximately 9 ml of blood was collected 20 min, 10 min and immediately prior to I.V. administration of the anesthetic, sodium pentothal (3 mg/lb BW). The use of pentothal as an anesthetic has been previously shown not to alter LH or testosterone concentrations in gilts and barrows

(Clapper, 2008). Once the animal was sedated, either a GnRH-I agonist (D-ala6 GnRH-I;

Bachem), a GnRH-II agonist (D-ala6 GnRH-II; Anygen) or a saline control was immediately injected transscrotally. Each testis was injected 3 times (ventral to dorsal, approximately 2 in apart), ~1 ½ in into the parenchyma with a 20 gauge, 1 ½ in hypodermic needle. The GnRH-I (50 ng) or GnRH-II (50 ng) agonists were dissolved in

1 ml of sterile saline, therefore accumulating to a total dose of 300 ng per animal.

Thereafter, blood was collected every 10 min for 40 min and then every 20 min for 240 min (Fig. 3.6). During blood sampling, catheters were kept clear by injecting 2 ml of

3.5% sodium citrate prior to blood draws. Once blood was evident upon drawback, S-

Monvette serum blood tubes (Sarstedt) were utilized to collect 9 ml of blood. Tubes were inverted slowly to aid in clotting and red blood cell lysis. Following blood collection, 3 86

TRT

Minutes - 20 -10 0 10 20 30 40 60 80 120 180 200 220 240

Figure 3.6. Schematic of boar intratesticular GnRH injection trial indicating treatment (TRT) and time of blood sample (hash mark). At 0 min, blood was collected prior to administration of treatment. 87 ml of sodium citrate was injected and the catheter cap was cleaned with 100% ethanol before being replaced. Blood samples remained at 4°C until centrifugation (12,000 x g) and serum was collected and stored at -20°C until RIA analysis. This experiment was a crossover design such that each animal received each treatment 4 days apart.

Radioimmunoassay. After completion of the experiment 2, 3 and 4, testosterone concentrations were determined using a Total Testosterone Coat-a-Count RIA kit

(Siemens Healthcare Diagnostics) in accordance with the manufacturer’s instructions.

This kit has been validated in the pig (Oskam et al., 2010; Ashworth et al., 2011) and parallelism was confirmed previously in our lab.

Briefly, 50 µl of zero calibrator (0 ng/ml testosterone) was added to 2 (duplicate) uncoated 12 x 75 mm polypropylene, non-specific binding (NSB) tubes and 2 Total

Testosterone Ab-Coated tubes for maximum binding. Then, 50 µl of each remaining calibrator (B-F; 0.16, 0.96, 4.14, 7.92 and 16.85 ng/ml testosterone, respectively) were pipetted into Total Testosterone Ab-Coated tubes (Specific binding; Bø), taking care into pipette directly to the bottom of all tubes. Two additional standards were added to increase the accuracy of the curve (0.08 and 0.04 ng/ml). After pipetting samples into each tube, 1 ml of testosterone radioactively labeled with I125 was added to every tube and tubes were covered with parafilm, vortexed for 30 s and incubated in a 37°C waterbath for 3 h. The NSB tubes containing only 1 ml 125I labeled testosterone were set aside for total counts. Following incubation, liquid was decanted thoroughly (except the total counts tubes) and allowed to drain for 2-3 min. Finally, samples were counted using 88 an APEX Automativ Gamma counter (ICN Micromedia Systems) for 1 min. The minimum sensitivity of the assay was 0.04 ng/ml and the intraassay coefficient of variation was 10.2%. Percent binding was then determined using the following equation:

[(Bø-NSB)/TC)]*100 = % binding. Data were analyzed and fit to the standard curve using Graphpad PRISM to determine relative testosterone levels.

Relative LH values were determined by Dr. Clay Lents’ laboratory at USMARC according to Kesner et al. (1987). The minimum sensitivity of the assay was 0.1 ng/ml

LH and the intraassay coefficient of variation was 10%.

Statistical Analysis

Data were analyzed using the Statistical Analysis System (SAS; Cary, NC).

Western blot data were confirmed to be normally distributed and analyzed using the

GLM procedure with replication as a random effect. Tissue explant RIA data was analyzed by the MIXED procedure with replication as a random effect. Statistical evaluation for all in vivo boar trials was performed using the MIXED procedure of SAS with time as a repeated measure. Day of treatment and assay were added as random effects. The Shapiro-Wilk test for normality indicated that means for RIA data from explant and boar trials were not normally distributed, thus, these means were logarithmically transformed and compared using least-significant differences (P < 0.05).

Non-transformed means, however, were presented in the results sections (Chapter IV) for comparison.

CHAPTER IV

The role of GnRH-II and its receptor in testicular function

Abstract

The highly conserved, second mammalian isoform of GnRH (GnRH-II) has been linked to regulation of cell proliferation, feed intake, and the interaction between energy balance and reproductive behavior. In contrast to the native form of GnRH (GnRH-I), GnRH-II is an inefficient modulator of gonadotropin secretion. Unlike many species, a functional receptor (GnRHR-II) specific to this ligand has been discovered in the pig that may be directly involved in testosterone production. Therefore, the objective of this study was to identify the role of GnRH-II and its receptor in testicular function. First, Western blot analysis of tissues from mature, Chinese Meishan (MS) boars revealed 6-fold more

GnRHR-II protein in the testis than anterior pituitary gland. GnRH-II levels determined by ELISA were highest in testis (1,321 ± 52.5 pg/ml), intermediate in pituitary (393 ±

58.7 pg/ml) and lowest in hypothalamus (220 ± 67.7 pg/ml) tissue homogenates.

Immunohistochemistry of testicular tissue indicated that the GnRHR-II was located on the plasma membrane of Leydig cells within the interstitium as well as germ cells.

Second, immunocytochemistry demonstrated that GnRHR-IIs were localized to the neck region of purified spermatozoa from mature, white crossbred (WC) boars. Consistent with this, GnRH-II was also detected in seminal plasma (225 ± 27.3 pg/ml) via ELISA. 90

Third, testicular tissue explants from MS boars exposed to 1 µM GnRH-II secreted testosterone similarly to hCG treated (1 I.U.) tissues, without influencing GnRHR-II or

LH receptor protein levels. Finally, 4 in vivo experiments were performed on MS or white crossbred (WC) boars surgically fitted with jugular cannulae. In Exp. 1, plasma testosterone concentrations were elevated following I.V. treatment of MS males with either a GnRH-I (D-ala6 GnRH-I) or GnRH-II (D-ala6 GnRH-II) agonist (150 ng/kg BW), although GnRH-II induced changes in LH levels were not correlated with testosterone production. In Exp. 2, I.M. treatment of WC males with a GnRH-I (SB-75; 5 µg biologically active compound/kg BW) or GnRH-II (Trp-1; 5 µg/kg BW) antagonist suppressed testosterone, but not LH, secretion. In Exp. 3, serum testosterone concentrations increased in WC males after I.M. treatment with SB-75 followed by I.V. infusion of either GnRH-I or -II agonist, despite the inability of GnRH-II to induce LH secretion. In Exp. 4, intratesticular injection of WC males with GnRH-I, GnRH-II or saline resulted in GnRH-II-induced stimulation of testosterone production without increasing LH levels. Taken together, these data suggest that GnRH-II directly impacts testosterone production via binding to the GnRHR-II on Leydig cells, bypassing LH production by the anterior pituitary gland.

Introduction

Boar subfertility is a source of economic loss often overlooked (Foxcroft et al.,

2008) since the swine industry has adopted techniques to compensate for subfertility.

Commonly, semen collections are pooled from multiple sires and packaged in excess of 3 billion sperm cells per dose (Dyck et al., 2011). Traditionally, semen samples undergo conventional analyses (semen volume, concentration, morphology and progressive motility) immediately after collection to determine fertility (Amann et al., 1995). While these parameters are useful for identifying infertile boars (Flowers, 1997; Xu et al.,

1998), they may not truly indicate the reproductive status of a boar (Correa et al., 1997;

Brahmkshtri et al., 1999). Therefore, more screening procedures are needed to identify subfertile boars to improve reproductive efficiency.

The classical form of GnRH (GnRH-I) is a key regulator of testosterone production through stimulation of luteinizing hormone (LH) secretion from the anterior pituitary gland. GnRH-I and its receptor (GnRHR-I) have also been detected in the testes of males; specifically, GnRHR-I has been detected in Leydig cells (Iwashita and Catt,

1985; Jegou et al., 1985; Nikula and Huhtaniemi, 1988; Petersson et al., 1989; Reinhart et al., 1992; Kakar et al., 1992), whereas GnRH-I may be produced by Sertoli cells

(Sharpe and Fraser, 1980; Verhoeven and Cailleau, 1985; Sharpe and Cooper, 1987;

Saint Pol et al., 1988). Moreover, the interaction between GnRH-I and its receptor in the testis have been implicated in localized regulation of testosterone production, independent of LH (Sharpe et al., 1983; Verhoeven and Cailleau, 1985; Jegou et al., 92

1985; Petersson et al., 1989; Bahk et al., 1995; Lin et al., 2010; Anjum et al., 2012). For example, rats injected with GnRH-I directly into the testis experienced an immediate increase in serum testosterone concentration, although LH levels were unaffected (Sharpe et al., 1983). Further, short-term GnRH-I treatment of rat primary Leydig cell cultures increased testosterone production (Sharpe and Cooper, 1982b), whereas long-term exposure decreased testosterone secretion, suggesting downregulation of the GnRHR-I.

Recently, a new mammalian form of GnRH (GnRH-II) was discovered, differing from GnRH-I by 3 amino acids and expressed in almost every tissue (Kasten et al., 1996;

Millar, 2003). GnRH-II also maintains its own specific receptor (GnRHR-II) and, like

GnRHR-I, is a 7-transmembrane (TM), G-protein coupled receptor (GPCR; Neill et al.,

2001; Millar et al., 2001). Interestingly, not all species produce a functional GnRHR-II due to coding errors (Stewart et al., 2009). However, the sequence for a functional receptor has been discovered in shrews, old world monkeys and swine (Stewart et al.,

2009). In these species, the function of GnRH-II and its receptor is unclear (Neill et al.,

2004), but evidence suggests GnRH-II is not an efficient modulator of gonadotropin secretion (Okada et al., 2003). Instead, GnRH-II and its receptor have been linked to feed intake (Kauffman and Rissman, 2004b), cell proliferation in peripheral reproductive tissues (Chou et al., 2002), and the interaction between energy balance and sexual behavior in females (Kauffman and Rissman, 2004a). In addition, new evidence suggests that GnRH-II might also be involved in male reproductive function.

Bowen et al. (2006) discovered that boars immunized against GnRH-II displayed reduced levels of testosterone, independent of alterations in gonadotropin secretion. 93

Similarly, primary Leydig cell cultures from GnRH-II immunized boars secreted less testosterone compared to control boars upon LH challenge. Additionally, boars treated with a specific GnRH-I antagonist (SB-75; cetrorelix) daily for 4 d displayed suppressed testosterone levels for the duration of treatment, whereas LH levels returned to basal after only 36 h of treatment (Zanella et al., 2000). Moreover, testicular explant cultures treated with human chorionic gonadotropin (hCG) in the presence of SB-75 produced less testosterone compared to controls.

Given that GnRHR-I has never been detected in the boar testis (Zanella et al.,

2000), our hypothesis is that the interaction between GnRH-II and its receptor can directly stimulate testosterone secretion from the boar testis via an autocrine or paracrine mechanism. Here, we demonstrate that GnRH-II and its receptor are highly abundant in the testis compared to the anterior pituitary gland, localized to the plasma membrane of

Leydig cells and germ cells. Furthermore, GnRH-II levels were highest in the testis, intermediate in the anterior pituitary gland and lowest in the hypothalamus. Testis explants treated with GnRH-II alone secreted testosterone to a similar level as hCG treated tissues, without upregulating the LH receptor. Moreover, boars challenged with

GnRH-I and -II secreted testosterone similarly, however, GnRH-II stimulated testosterone levels were not correlated with LH concentrations. Additionally, administration of a GnRH-I antagonist prior to GnRH-II treatment prevented LH, but not testosterone, production. Consistent with this, GnRH-II injected directly into the testis stimulated testosterone, but not LH, production. Finally, the GnRHR-II was localized to the neck region of spermatozoa, corresponding to GnRH-II presence in seminal plasma. 94

Materials and Methods

Localization of GnRH-II and GnRHR-II

Tissue Collection. Animal trials were conducted at the U.S. Meat Animal

Research Center (USMARC; Clay Center, Nebraska) in accordance with the Animal Care and Use Committee. Mature Chinese Meishan boars (n = 5) were housed individually at

USMARC and fed 6 lb of feed daily with water available ad libitum. Boars (15-17 wk of age) were euthanized and the hypothalamus, anterior pituitary gland and testes were collected. Tissues were either snap frozen in liquid nitrogen and stored at -80°C or fixed in 4% paraformaldehyde for immunohistochemistry (IHC).

Immunohistochemistry. Tissue isolated from the testes was dehydrated with

100% ethanol, cleared with CitriSolv (Fischer Scientific, Pittsburgh, PA), paraffin embedded, sectioned (5 µm) and mounted on UltraStick slides (Gold Seal Products,

Portsmouth, NH). Epitope retrieval was achieved by boiling slides containing sections of testicular tissue in 0.01 M sodium citrate for 15 min. Non-specific binding was blocked with 10% rabbit serum for 20 min and slides were incubated for 30 min at room temperature (RT) with the primary antibody, anti-GnRHR-II (goat polyclonal; SC-

162889; Santa Cruz Biotechnology, Santa Cruz, CA), diluted (1:100) in 1X TBS (25 mM

Tris, 150 mM NaCl, 2 mM KCl). Next, alkaline phosphatase detection was performed using the VECTASTAIN ABC-AP kit (Vector Laboratories, Burlingame, CA) per manufacturer’s instructions. Finally, slides were mounted with Tris-buffered 80% 95 glycerol and imaged using an Olympus (Center Valley, PA) microscope (BX51) and camera (DP71).

Semen Collection. Semen was collected from mature, white crossbred boars

(UNL Index line; n = 5) using a gloved hand method. Semen was centrifuged (800 x g) for 10 min at RT and seminal plasma was removed and stored at -20°C for later analysis.

Then, 1 ml of the semen pellet was layered on a discontinuous Percoll gradient containing 2 ml of a 90% Percoll solution [90% Percoll, 10% Tyrode’s stock (40 mM

KCL, 2.92 mM, 0.05% NaH2PO4, 0.02% Hepes buffer, pH 7.3), 1.97 mM CaCl2, 0.394 mM, 0.002% lactic acid, 0.002% NaHC03] and 2 ml of a 45% Percoll solution (50% of the 90% Percoll solution and 50% TL sperm solution; 100 mM NaCl, 3.1 mM KCl, 25.0 mM NaHC03, 0.29 mM NaH2Po4H2O, 26 mM Na lactate, 2.107 mM CaCl22H20, 0.4 mM

MgCl6H20 and 10 mM Hepes; pH 7.3) layered on top of the 90% Percoll solution in a 15 ml centrifuge tube. The tubes were centrifuged (800 x g) for 30 min at RT and purified spermatozoa were collected from the pellet. Samples were then diluted in 1X phosphate buffered saline (PBS; 1.36 M NaCl, 1.08 M KCl, 0.1 M Na2HPO4 and 0.017 M KH2PO4) and either air-dried on UltraStick slides (Gold Seal Products) for later immunocytochemistry or flash frozen.

ELISA. Isolated seminal plasma as well as testis, anterior pituitary gland and hypothalamus homogenates were assayed for a precursor form of GnRH-II with a commercially available enzyme-linked immunosorbent assay (ELISA) kit, specific to the 96 pig (Cat. # CSE-EL009636PI; Cusabio; Hubei, China). The ELISA was performed according to the manufacturer’s instructions. Briefly, a standard curve (0, 20, 40, 160,

640, 2,000 and 8,000 pg/ml) was added to the plate, followed by the unknown samples

(seminal plasma or tissue homogenates). Next, horseradish peroxidase-conjugate was added to each well, followed by the primary antibody. The plate was mixed and incubated at 37°C for 1 h. Subsequently, each well was aspirated and washed with 1X

Wash Buffer Solution and then Substrate A was added to each well, followed by

Substrate B. The plate was then mixed and incubated in the dark at 37°C for 15 min.

Finally, the Stop Solution was added to each well, the plate was mixed and the optical density was immediately read using a Wallac 1420 Victor2 microplate reader (Perkin

Elmer; Waltham, MA) at 450 nm. The data were interpolated based on the standard curve using a 4 parameter logistic (4-PL) curve fit per manufacturer’s instructions. The minimum sensitivity of the assay was 20 pg/ml. The intraassay coefficient of variation was 10%.

Immunocytochemistry. After air-drying, spermatozoa were fixed with 4% paraformaldehyde for 10 min at RT in a humidified chamber. Then, slides were exposed to UV light for 30 min to quench autofluorescence. Cells were permeabilized with 1X

TBS-T (0.5% Tween-20) for 10 min at RT. Non-specific binding was blocked by incubation with 10% normal rabbit serum diluted in TBS-T (0.045% Tween-20) for 30 min at RT. Cells were incubated in a humidified chamber at 4°C overnight with a polyclonal goat primary antibody directed against GnRHR-II (1:50; SC-162889; Santa 97

Cruz Biotechnology), diluted with 10% normal serum in TBS-T (0.045% Tween-20).

Next, slides were incubated with a rabbit anti-goat Alexa Fluor® 488 secondary antibody

(Invitrogen, Carlsbad, CA) diluted 1:400 in 10% normal serum for 1 h at RT. After all steps, slides were washed with 1X TBS-T. Finally, slides were rinsed with TBS and mounted with ProLong Gold Antifade Reagent (Invitrogen), and imaged with an inverted confocal microscope (Olympus IX 81) using FITC filter sets.

Testicular Explants

Materials. Chinese Meishan boars (n = 7) were euthanized and whole testes were collected. Approximately 400 mg of finely minced tissue was incubated in 5 ml of

TC199 (with Earle's salts and NaHCO3; Sigma-Aldrich, St. Louis, MO) buffered with a final concentration of 25 mM Hepes. Tissue was exposed to a 95% O2 and 5% CO2 atmosphere and maintained at 37ºC for 1 h. This preincubation was necessary as testosterone is released from the tissue during dissection and can mask hormone induced testosterone secretion (Allrich et al., 1983). Then, the media was decanted and replaced with fresh TC199 and explants were randomly assigned to a treatment.

Treatments. At time 0, explant cultures were exposed to either vehicle (water) or

1 µM GnRH-II (Bachem Inc., Torrance, CA) as a pretreatment. Media (250 µl) was collected 1 h after pretreatment from all cultures and explants were treated with either vehicle (water) or 1 I.U. human chorionic gonadotropin (hCG; Sigma-Aldrich).

Therefore, 4 combinations (pretreatment + treatment) were tested: control (vehicle + 98 vehicle), hCG alone (vehicle + hCG), GnRH-II pretreatment without hCG (GnRH-II + vehicle) or GnRH-II pretreatment with hCG (GnRH-II + hCG). After treatment, media

(250 µl) was collected 120, 180 and 240 min later. Media was immediately frozen after each collection and subsequently stored at -20ºC, whereas tissue was snap frozen and stored at -80°C.

Western Blot. Protein was extracted from explant, testis and anterior pituitary gland tissues by homogenization in 1 ml of RIPA buffer (20 mM Tris, 137 mM NaCl,

10% glycerol, 1% NP40, 0.1% SDS, 0.5% deoxycholic acid, 2 mM EDTA, 1mM PMSF,

1% protease inhibitor cocktail and 1% phosphatase inhibitor cocktail) per 100 mg of tissue. Spermatozoa protein was extracted by homogenizing samples (100 µl) in 200 µl of

RIPA buffer. The protein concentration of the lysates was quantified using a bicinchoninic acid (BCA) assay (Pierce Biotechnology, Rockford, IL) per manufacturer’s instructions. Dithiothreitol (DTT) was mixed with 4X loading dye [LD; 2% Tris (pH

6.8), 28% Glycerol, 20% SDS, a fleck of Orange G] to a ratio of 10:50 (DTT:LD) and combined with the extracted protein 1:3 (LD:protein).

Proteins were separated using SDS-PAGE (10%) and transferred to polyvinylidene difluoride (PVDF) membrane. After transfer, non-specific binding was blocked by incubation in Odyssey Blocking Buffer (Licor Biosciences, Lincoln, NE) for

1 h at RT. Then, membranes were probed with a goat polyclonal primary antibody directed against GnRHR-II (1:1000; SC-162889; Santa Cruz Biotechnology), diluted in

Odyssey Blocking Buffer with 0.05% Tween-20 shaking at 4ºC overnight. Next, 99 membranes were incubated with a secondary donkey anti-goat antibody (1:1600; Alexa

Fluor® 680; Invitrogen) in Odyssey Block for 1 h at RT. Between each step, the membranes were washed in TBS-T. On separate Western blots, protein from explant tissue was also probed with an antibody directed against the LH receptor (LHR; rabbit polyclonal; 1:300; SC-25828; Santa Cruz Biotechnology) followed by a secondary goat anti-rabbit antibody (1:1600; Alexa Flour® 800; Invitrogen). For all blots, bands were visualized and quantitated with an Odyssey scanner and image software (Licor

Biosciences). Membranes were then stripped with NewBlot stripping buffer (Licor

Biosciences) for 30 min at 37°C and reprobed with β-actin to serve as the internal control for Western blot analyses. Samples were quantitated, normalized to β-actin and then expressed as a fold change. The blots containing protein from purified spermatozoa were not reprobed with β-actin as comparisons across samples were examined.

Boar Trials

Animals. Experiment 1 was conducted at USMARC in accordance with the

Animal Care and Use Committee. Mature, Chinese Meishan boars were housed individually and fed 6 lb of feed daily with water available ad libitum. Jugular catheterization of boars was conducted according to Ford and Maurer (1978), approximately 4-5 d prior to initiation of the trial. Experiments 2, 3 and 4 were performed at the UNL Animal Science Building. All animal procedures were conducted according to the UNL Institutional Animal Care and Use Committee. Mature, white crossbred boars were housed individually and fed 5 lb of feed daily with water available ad libitum. At 100 least 3 d before trial initiation, boars were cannulated in the jugular vein according to either Barb et al. (1982) or Ford and Maurer (1978).

Materials. D-ala6 GnRH-I (Bachem Inc.) and D-ala6 GnRH-II (Anygen;

Gwangju, Korea) were both dissolved in 0.9% sterile saline. SB-75 was synthesized by the UNL Protein Core (Lincoln, NE) and dissolved in 0.9% sterile saline. Trptorelix-1

(Trp-1; Anygen) was dissolved per manufacturer’s instructions. Briefly, Trp-1 was first dissolved in 5% dimethylacetamide, mixed with 40% propylene glycol and finally, diluted in 55% millipore water.

Experiment 1: GnRH Agonist Trial. Prior to treatment, blood samples were collected every 20 min for 2 h to obtain a baseline hormonal profile. Next, boars (n = 5) were given I.V. infusions of either a GnRH-I (D-ala6 GnRH-I; 150 ng/kg BW) or GnRH-

II (D-ala6 GnRH-II; 150 ng/kg BW) agonist. Blood samples were drawn every 10 min for

30 min and every 20 min for 270 min thereafter. Blood samples remained at 4°C until centrifugation (12,000 x g) and plasma was collected and stored at -20°C until radioimmunoassay (RIA) analysis. The experiment was a cross-over design such that each boar received each treatment once, 1 wk apart.

Experiment 2: GnRH Antagonist Trial. Prior to treatment, blood was drawn 30 min and immediately prior to treatment. Treatments, either a specific GnRH-I (SB-75; 5

µg biologically active compound/kg BW; n = 9) or GnRH-II (Trptorelix; Trp-1; 5 µg/kg; 101 n = 10) antagonist, were given I.M. in the neck. Each animal also received the vehicle of the opposite treatment. Next, blood sampling occurred every 30 min for 3 h (30, 60, 90,

120, 150 and 180 min) and at 6, 12, 24, 36 and 48 h. Blood samples remained at 4°C until centrifugation (2,000 x g) and serum was collected and stored at -20°C until RIA analysis. The experiment was a cross-over design such that each boar received each treatment once, 1 wk apart.

Experiment 3: SB-75 and GnRH Agonist Trial. First, blood was collected 20 min, 10 min and immediately prior to treatment. Next, each boar was treated I.M. with the GnRH-I antagonist, SB-75 (5 µg biologically active compound/kg BW).

Approximately 7.5 h later, blood was collected and boars were given I.V. infusions of either a GnRH-I (D-ala6 GnRH-I; 150 ng/kg BW; n = 5) or GnRH-II (D-ala6 GnRH-II;

150 ng/kg BW; n = 5) agonist. Blood samples were drawn every 10 min for 30 min and then every 20 min for 270 min. Blood samples remained at 4°C until centrifugation

(12,000 x g) and serum was collected and stored at -20°C until RIA analysis. This experiment was a crossover design such that each animal received each treatment once, 1 wk apart.

Experiment 4: Intratesticular GnRH Injection Trial. Blood was collected 20 min, 10 min and immediately prior to I.V. administration of the anesthetic, sodium pentothal (3 mg/lb BW). The use of pentothal as an anesthetic has been previously shown not to effect LH or testosterone concentrations in gilts and barrows (Clapper, 2008). Once 102 the animal was sedated, either a GnRH-I agonist (D-ala6 GnRH-I; n = 6), GnRH-II agonist (D-ala6 GnRH-II; n = 6) or saline control (n = 6) was immediately injected transscrotally. Each testis was injected 3 times (ventral to dorsal, approximately 2 in apart), approximately 1 ½ in into the parenchyma with a 20 gauge, 1 ½ in hypodermic needle. The GnRH-I (50 ng) or GnRH-II (50 ng) agonists were dissolved in 1 ml of sterile saline, therefore accumulating to a total dose of 300 ng per animal. Thereafter, blood was collected every 10 min for 40 min and every 20 min for 240 min. Blood samples remained at 4°C until centrifugation (12,000 x g) and serum was collected and stored at -20°C until RIA analysis. This experiment was a crossover design such that each animal received each treatment once, 4 d apart.

Radioimmunoassay. Total testosterone concentrations from explant media were determined using a Total Testosterone Coat-a-Count RIA kit (Siemens Healthcare

Diagnostics, Los Angeles, CA) in accordance with the manufacturer’s instructions. This kit has been previously validated in the pig (Oskam et al., 2010; Ashworth et al., 2011) and parallelism was confirmed in our lab. The minimum sensitivity of the assay was 0.16 ng/ml and the intraassay coefficient of variation was 8%. Testicular explant media samples were diluted 1:5 with a zero calibrator solution (Siemens Healthcare

Diagnostics).

Plasma samples from Experiment 1 were assayed for testosterone at USMARC in

Dr. Clay Lent’s laboratory according to Ford et al. (2001). The minimum sensitivity of the assay was 0.1 ng/ml and the intraassay coefficient of variation between pooled 103 samples averaged 9%. In Experiments 2, 3 and 4, total testosterone concentrations of boar serum were measured at UNL using a Total Testosterone Coat-a-Count RIA kit (Siemens

Healthcare Diagnostics) per manufacturer’s instructions. However, 2 additional standards were added to increase the sensitivity of the curve (0.04 and 0.08 ng/ml). Data were analyzed and fit to the standard curve using Graphpad PRISM (San Diego, CA) to determine relative testosterone concentrations. The minimum sensitivity of the assay was

0.04 ng/ml and the intraassay coefficient of variation was 10%. LH concentrations for all trials were determined by Dr. Clay Lent’s laboratory at USMARC according to Kesner et al. (1987). The minimum sensitivity of the assay was 0.1 ng/ml LH and the intraassay coefficient of variation for pooled samples was 10%.

Statistical Analysis. All data were analyzed using the Statistical Analysis

System (SAS; Cary, NC). Western blot data were confirmed to be normally distributed and analyzed using the GLM procedure with replication as a random effect. Tissue explant RIA data was analyzed by the MIXED procedure with replication as a random effect. Statistical evaluations for Experiments 1, 2, 3 and 4 were performed using the

MIXED procedure with time as a repeated measure. Day of treatment and assay were added as random effects. The Shapiro-Wilk test for normality indicated that means for

RIA data from explant and boar trials were not normally distributed, thus, the means were logarithmically transformed and compared using least-significant differences. Non- transformed means, however, were presented in the Results section for comparison.

104

Results

GnRHR-II is present on the plasma membrane of Leydig cells and germ cells within the testis. Since our laboratory previously detected GnRHR-II in the boar testis (data not shown), we wanted to determine the location of the GnRHR-II within the testis.

Following IHC of porcine testicular tissue with an antibody directed against GnRHR-II, positive staining was detected in the interstitial compartment, as well as within the seminiferous tubules (Fig. 4.1; Panels B-D). Within the interstitium, staining appeared to be concentrated on the plasma membrane of Leydig cells. Transmembrane staining was also visible within the seminiferous tubules and appeared most intense on germ cells. In contrast, there was no signal detected in control sections incubated without the primary antibody (Fig. 4.1; Panel A). The presence of GnRHR-II in the testis was confirmed via

Western blots using an anti-GnRHR-II antibody. Strikingly, quantification of relative band density from Western blots indicated that the testis maintained 6-fold higher

GnRHR-II protein levels than the anterior pituitary gland (P < 0.0001; Fig. 4.2; Panel A and B).

GnRH-II is present in the hypothalamus, anterior pituitary gland and testis.

Given the abundance of GnRHR-II protein within the testis, we examined tissues for the presence of its ligand. An ELISA specific for a precursor form of GnRH-II in pigs was used to determine GnRH-II levels in tissue homogenates from the hypothalamus, anterior pituitary gland and testis. Although GnRH-II was detected in homogenates from all 3 105

II II

ncubated without the primary antibody. primary the ncubatedwithout

Representative immunohistochemistry image of boar testes (n = 7) using an antibody directed against GnRHR

D). Panel A section a i represents A Panel D).

Figure Figure 4.1. B (Panels 106

A Anterior B 8

Pituitary Testis Actin - β 6 *

4 GnRHR-II 60 kDa

2 Normalized Normalized to β-Actin 42 kDa 0 Anterior Pituitary Testis

Figure 4.2. Representative Western blot of porcine anterior pituitary gland and testis tissue (n = 5) using an antibody directed against GnRHR-II (Panel A). Quantification of Western blots revealed a difference in GnRHR-II protein levels between the tissue types (* P < 0.0001; Panel B). 107 tissues, levels were significantly different among tissue types (Fig. 4.3). Concentrations were highest in the testis (1,321 ± 52.5 pg/ml; P < 0.001), intermediate in the anterior pituitary gland (393 ± 58.7 pg/ml; P < 0.0001) and lowest in the hypothalamus (220 ±

67.7 pg/ml; P < 0.0001). Therefore, the abundance of GnRH-II in testicular homogenates as well as localization of GnRHR-II on the plasma membrane of Leydig and germ cells reflect potential autocrine or paracrine mechanisms controlling testosterone production and/or spermatogenesis.

GnRHR-II is localized to the neck region of the spermatozoa tail, corresponding to

GnRH-II presence in seminal plasma. After our discovery that GnRHR-IIs were present on the plasma membrane of germ cells, we screened mature boar spermatozoa for

GnRHR-II. Western blot analysis of protein from purified boar spermatozoa indicated the presence of GnRHR-II (Fig. 4.4; Panel A). Inclusion of testicular protein was used as a

GnRHR-II positive control based on the results described above. Interestingly, the size of

GnRHR-II protein bands differed between testis (60 kDa) and spermatozoa (54 kDa).

Indeed, receptors in sperm can be post-translationally modified differently than other tissues, resulting in alternative band sizes (Lee et al., 2000). Consistent with these results, GnRH-II was also detected in the seminal plasma of 7 boars (Fig. 4.4; Panel B).

Using an ELISA for a precursor form of porcine GnRH-II, concentrations ranged from

124 to 332 pg/ml, with a mean of 225 ± 27.3 pg/ml. In point of fact, these values were similar to those detected in hypothalamic tissue homogenates (220 ± 67.7 pg/ml).

Moreover, we utilized immunocytochemistry with an antibody directed against GnRHR- 108

1600

1400 c /ml)

1200

(

1000

800

600

Concentration b II II - 400 a

GnRH 200

0 Hypothalamus Anterior Pituitary Testis Figure 4.3. GnRH-II levels in the hypothalamus, anterior pituitary gland and testis of a,b,c boars. Bars with alternate superscripts differ (P < 0.01). 109

A B

400

/ml)

pg ( 300 Testis Sperm 200 60 kDa

54 kDa Concentration II II - 100

GnRHR-II GnRH 0 1 2 3 4 5 6 7 Boar Figure 4.4. Representative Western blot of boar testis and purified spermatozoa protein (n = 5) using an antibody directed against GnRHR-II (Panel A). Concentration of GnRH-II present in the seminal plasma of boars (n = 7; Panel B). 110

II on live sperm separated by a discontinuous Percoll gradient. Staining was present on spermatozoa, localizing to the neck region posterior of the head and at the anterior portion of the midpiece (Fig. 4.5; Panels A-C). In contrast, no fluorescent signal was detected in secondary only controls (Fig 4.5; Panel D-F). Therefore, the interaction between GnRH-II in seminal plasma and its receptor on sperm may be important to motility or tail development.

GnRH-II stimulates testosterone production similar to hCG, without upregulating

LH receptors in testicular explant cultures. In addition to possible roles in spermatogenesis, we also examined the effect of GnRH-II on steroidogenesis in the testis.

Media was collected from testicular explant cultures pretreated for 1 h with or without 1

µM GnRH-II followed by treatment with vehicle or 1 I.U. hCG 1 h later. Samples collected at 60, 120, 180 and 240 min after pretreatment (GnRH-II or vehicle) were assayed for testosterone production by RIA. Treatment values were first normalized to basal testosterone levels and expressed as a fold change over the control. There was no effect of time or treatment x time interaction for fold change in testosterone concentration

(P > 0.05). However, there was a significant effect of treatment (P < 0.001), with all treatments increasing testosterone over the control treatment (Fig. 4.6). Markedly, the

GnRH-II pretreatment + no hCG treatment increased testosterone production similar to explants that were not pretreated with GnRH-II but treated with hCG (P > 0.05).

Moreover, Western blot analyses were conducted on testicular explant tissue collected after 180 h of culture using antibodies directed against either GnRHR-II or LHR. There 111

Figure 4.5. Representative confocal microscopy images of boar spermatozoa (n = 5) ® using an antibody directed against GnRHR-II labeled with an Alexa Fluor 488 secondary antibody (Panels A-C; green). Panels D-F represent secondary only control images. 112

No GnRH-II Pretreatment GnRH-II Pretreatment

3 b 2.5 b b

1.5 a

Fold Change Fold 1

0.5

0 Vehicle hCG Vehicle hCG Figure 4.6. Relative testosterone levels secreted by testicular explant cultures (n = 7) pretreated with or without 1 µM GnRH-II followed by treatment with vehicle or 1 I.U. a,b hCG. Bars with alternate superscripts differ (P < 0.001). 113 was no difference in the levels of GnRHR-II or LHR protein between treatments (P >

0.05; Fig. 4.7), suggesting that GnRH-II mediated testosterone production without modulating the LH receptor.

GnRH-I and -II promote differential secretion of LH, but not testosterone. After establishing that GnRH-II stimulated testosterone production in testicular explant cultures, we examined whether GnRH-II would have a similar effect in vivo. Plasma collected before and after treatment (10 - 270 min) of mature, Chinese Meishan boars with either D-ala6 GnRH-I or D-ala6 GnRH-II was assayed for LH and testosterone concentrations. Mean values were expressed as a fold change over an average of the pretreatment values. There was a treatment x time interaction (P < 0.0001) for fold change in LH concentrations (Fig. 4.8). Within 10 min after administration of GnRH-I,

LH concentrations increased 2-fold over pretreatment levels, reaching maximal levels (4- fold) by 130 min, and remained elevated (3-fold) for the duration of sampling (270 min;

Fig. 4.8). Similarly, relative LH levels increased 2-fold within 10 min following GnRH-II treatment, however, unlike the response to GnRH-I, LH concentrations steadily declined to pretreatment values by 110 min (Fig. 4.8). GnRH-II treated animals had lower LH concentrations than males treated with GnRH-I at 210, 230, 250 and 270 min (P < 0.05).

In addition, LH levels tended to be lower in GnRH-II treated animals compared to those in the GnRH-I treatment group at 170 and 190 min (P < 0.10).

Interestingly, there was no treatment x time interaction for fold change in testosterone concentrations (P > 0.05; Fig 4.8). Although there was a significant effect of 114

A No GnRH-II GnRH-II B No GnRH-II GnRH-II

Pretreatment Pretreatment Pretreatment Pretreatment

Vehicle hCG Vehicle hCG Vehicle hCG Vehicle hCG GnRHR-II 60 kDa LHR 65 kDa β-Actin 42 kDa β-Actin 42 kDa

C D

No GnRH-II Pretreatment GnRH-II Pretreatment

No GnRH-II Pretreatment GnRH-II Pretreatment 2.5

1.5 1.5 1

1 Fold Fold Change

Fold Change Fold 0.5 0.5

0 0 Vehicle hCG Vehicle hCG Vehicle hCG Vehicle hCG

Figure 4.7. Representative Western blot of porcine explant cultures (n = 7) using an antibody directed against GnRHR-II (Panel A) and LHR (Panel B). Quantification of Western blots for GnRHR-II (Panel C) and LHR (Panel D) revealed that there were no differences between treatments (P > 0.05). 115

D-ala⁶ GnRH-I 7 D-ala⁶ GnRH-II LH * * * 6 * * * * * * * 5 * * * * 4 *

3 trt P < 0.0001 time P < 0.0001

FoldChange 2 trt*time P < 0.0001 * * 1 * * * *

-1

10 20 30 50 70 90

Pre

110 130 150 170 190 210 230 250 270

4 Testosterone

trt P = 0.2491 time P < 0.0001

Change 2

trt*time P = 0.8659 Fold 1

10 20 30 50 70 90

Pre

110 130 150 170 190 210 230 250 270

Time (min)

Figure 4.8. Relative levels of LH and testosterone after intravenous administration of a specific GnRH-I or GnRH-II agonist. After pretreatment (Pre) sampling, treatments were administered (arrow). The mean ± SEM values are expressed as a fold change over the pretreatment mean. *Different from pretreatment concentrations (P < 0.05; GnRH-I, above line; GnRH-II, below line).

116 time (P < 0.0001), GnRH-I and -II induced testosterone levels did not differ from pretreatment values at any specific time point (P > 0.05). Both treatments stimulated a similar overall change in testosterone production (P > 0.05). Therefore, testosterone secretion was promoted similarly in GnRH-I and -II treatment groups, despite reduced

LH secretion in GnRH-II treated males.

Testosterone, but not LH, is suppressed in boars treated with SB-75 or Trp-

1. Since the GnRH-II and -II agonists stimulated testosterone production similarly but differed in LH responsiveness, we next utilized a GnRH-II antagonist to determine if testosterone could be suppressed without reducing LH secretion. Serum collected before and after treatment (10 min - 48 h) of mature, white crossbred boars with either a specific

GnRH-I (SB-75) or GnRH-II (Trp-1) antagonist was assayed for LH and testosterone concentrations. Mean values were expressed as a fold change over an average of the pretreatment values. There was no treatment x time interaction for fold change in LH levels (P = 0.13), although there was an effect of both treatment (P < 0.01) and time (P <

0.0001; Fig. 4.9). Trp-1-induced LH levels did not differ from pretreatment levels at any time point (P > 0.05). Similarly, SB-75-induced relative LH levels did not differ from pretreatment values at any specific time point (P > 0.05). Overall, Trp-1 stimulated an increase in LH secretion compared to SB-75, 1.12 ± 0.08 versus 1.01 ± 0.082, respectively (P < 0.001). An overall increase in LH secretion from animals in the Trp-1 treatment group appears to result from a differential response to treatment after 6 h, compared to SB-75 treated animals (Fig. 4.10). 117

2.5 SB-75

LH Trp-1

2.0

1.5 trt P = 0.0015 time P < 0.0001

1.0 trt*time P < 0.1323 FoldChange

0.5

0.0

1 2 3 6

12 24 36 48

0.5 1.5 2.5 Pre

2.0 Testosterone

1.5

trt P < 0.0001 1.0 time P < 0.0001 * trt*time P < 0.0001

0.5 FoldChange

0.0 * * *

-0.5

1 2 3 6

12 24 36 48

0.5 1.5 2.5 Pre Time (h)

Figure 4.9. Relative levels of LH and testosterone after intramuscular administration of a specific GnRH-I (SB-75) or GnRH-II antagonist (Trp-1). After pretreatment (Pre) sampling, treatments were administered (arrow). The mean ± SEM values are expressed as a fold change over the pretreatment mean. *Different from pretreatment concentrations (P < 0.05; Trp-1, above line; SB-75, below line).

118

Trp-1 2 LH SB-75

1.5

1 FoldChange 0.5

0 6 12 24 36 48

2 Testosterone

1.5

1 FoldChange

0.5

0 6 12 24 36 48

Time (h)

Figure 4.10. Trend lines for relative changes in LH and testosterone concentrations following suppression with a specific GnRH-I (SB-75) or GnRH-II (Trp-1) antagonist. The mean values are expressed as a fold change over the pretreatment mean. 119

There was an effect of treatment, time and treatment x time interaction (P <

0.0001) for fold change in testosterone concentration (Fig. 4.9). After treatment with SB-

75, serum testosterone levels were significantly reduced at 3, 6 and 12 h compared to pretreatment values. After administration of Trp-1, relative serum testosterone levels significantly dropped below pretreatment levels at 6 h and returned to basal levels by 12 h. Testosterone values for Trp-1 treated animals were higher than boars treated with SB-

75 at 12 and 24 h (P < 0.05). In addition, testosterone concentration in Trp-1 treated boars tended to be higher than animals in the SB-75 treatment group at 6 h (P < 0.10).

Interestingly, relative testosterone levels returned from suppression similarly between

SB-75 and Trp-1 treated boars (Fig. 4.10). However, the trend for LH concentration to return from suppression was similar to testosterone in animals treated with SB-75, but not boars in the Trp-1 treatment group (Fig. 4.10). Thus, the increase in testosterone levels following Trp-1 suppression occurred without a corresponding increase in LH secretion.

Antagonizing the GnRHR-I prevents GnRH-II-induced LH secretion. Given

GnRH-II can function through the GnRHR-I, we developed a trial to evaluate the effect of GnRH-I and -II on LH and testosterone production following blockage of GnRHR-Is.

Serum collected before and after treatment (10 - 270 min) of mature, white crossbred boars with SB-75 prior to infusion of either D-ala6 GnRH-I or D-ala6 GnRH-II was assayed for LH and testosterone concentrations. Mean values were expressed as a fold change over an average of the pretreatment values. We observed a treatment x time effect for fold change in LH concentration (P < 0.0001), as well as significant effects of time (P 120

< 0.0001) and treatment (P < 0.0001; Fig. 4.11). After administration of SB-75, LH levels dropped numerically in both treatment groups, but were not significantly different from the pretreatment LH concentrations (P > 0.05). After treatment with GnRH-I, LH levels rose significantly above pretreatment levels within 10 min and remained elevated for 90 min. After treatment with GnRH-II, serum LH concentrations tended to surpass pretreatment hormone levels at 10 min (P = 0.07) but not at any other time point (P >

0.05). At 90 min, relative LH levels tended to be lower in GnRH-II treated animals than pretreatment concentrations (P = 0.06) and were significantly lower at 190, 210 and 250 min (P < 0.05). Compared to LH concentrations after SB-75 treatment, GnRH-I induced

LH levels were higher from 10 to 170 min (P < 0.05). Serum LH concentrations resulting from GnRH-II treatment rose above post-SB-75 levels at 10 min and remained elevated through 50 min (P < 0.05). Fold change in serum LH levels of GnRH-II treated animals was lower than GnRH-I treated boars at 10, 20, 30, 50, 70, 90, 110, 150, 170 and

190 min (Table 4.1). At 130 min, there was a tendency for lower LH levels in GnRH-II treated boars (P = 0.09; Table 4.1).

We observed an effect of treatment (P < 0.0001) and time (P < 0.0001) but not a treatment x time interaction (P = 0.74) for fold change in testosterone concentrations

(Fig. 4.11). After treatment with SB-75, testosterone levels for both GnRH-I and -II treatment groups dropped significantly below pretreatment levels, remaining suppressed for 20 min before returning to basal levels (P < 0.05; Fig. 4.11). Compared to post-SB-75 levels, testosterone concentrations were significantly elevated at 50, 70, 90, 110, 130,

150, 170, 190, 210, 230 and 250 min following GnRH-I stimulation. Further, there was a 121

TABLE 4.1. MEAN FOLD CHANGE FOR LH CONCENTRATIONS ± SEM FOLLOWING TREATMENT WITH A GnRH-I ANTAGONIST (SB-75) AND SUBSEQUENT INFUSION OF AGONISTS FOR GnRH-I OR -IIa. Time Post-Treatment D-ala6 GnRH-I D-ala6 GnRH-II Significance (min)

10 2.95 ± 0.23 1.64 ± 0.25 P = 0.0228

20 3.22 ± 0.31 1.56 ± 0.25 P = 0.0041

30 2.80 ± 0.25 1.29 ± 0.24 P < 0.0001

50 2.57 ± 0.23 0.97 ± 0.24 P < 0.0001

70 2.17 ± 0.23 0.76 ± 0.25 P < 0.0001

90 1.91 ± 0.25 0.64 ± 0.25 P < 0.0001

110 1.68 ± 0.25 0.79 ± 0.27 P = 0.0009

130 1.49 ± 0.23 0.87 ± 0.27 P = 0.0889

150 1.33 ± 0.25 0.59 ± 0.31 P = 0.0141

170 1.43 ± 0.31 0.69 ± 0.25 P = 0.0420

190 1.01 ± 0.27 0.58 ± 0.23 P = 0.0251

210 0.93 ± 0.25 0.54 ± 0.25 P = 0.1469

230 0.93 ± 0.23 0.57 ± 0.27 P = 0.9630

250 0.74 ± 0.31 0.51 ± 0.25 P = 0.2733

270 0.99 ± 0.25 0.64 ± 0.31 P = 0.6012 a Values are expressed as a fold change over the pretreatment levels.

122

4 D-ala⁶ GnRH-I LH * D-ala⁶ GnRH-II * * * 3

* * 2 trt P < 0.0001 time P < 0.0001

trt*time P < 0.0001 FoldChange

* * *

10 20 30 50 70 90

110 130 150 170 190 210 230 250 270

Pre Post

4 Testosterone

2 trt P < 0.0001 time P < 0.0001 trt*time P = 0.7417 1 * * FoldChange *

0 * * *

-1

10 20 30 50 70 90

Pre

110 130 150 170 190 210 230 250 270 Post Time (min)

Figure 4.11. Relative levels of LH and testosterone after intramuscular administration of SB-75 and intravenous treatment with a specific GnRH-I or GnRH-II agonist. After pretreatment (Pre) sampling, SB-75 was administered prior to blood collection (black arrow). Approximately 7.5 h later, blood was collected and treatments were administered (Post; gray arrow). The mean ± SEM values are expressed as a fold change over the pretreatment mean. *Different from pretreatment concentrations (P < 0.05; GnRH-I, above line; GnRH-II, below line).

123 tendency for increased testosterone levels at 270 min (P = 0.08). Following GnRH-II treatment, relative testosterone levels were increased at 50, 70, 90, 110, 130, 150 and 170 min (P < 0.05) compared to post-SB-75 hormone concentrations,. In addition, there was a tendency for elevated testosterone levels at 190 min (P = 0.08). Overall, GnRH-I stimulated a higher level of relative testosterone production, (1.16 ± 0.28) compared to

GnRH-II (0. 73 ± 0.28; P < 0.05). In summary, treatment with a GnRH-I antagonist prevented GnRH-II, but not GnRH-I, induced LH secretion.

Intratesticular injection with GnRH-II increases serum testosterone, but not

LH production. Finally, we designed an experiment to test whether direct administration of GnRH-II into the testis would elicit differential results than systemic delivery. Serum collected before and after intratesticular treatment (10 - 240 min) of mature, white crossbred boars with either saline (control), D-ala6 GnRH-I or D-ala6

GnRH-II was assayed for LH and testosterone concentrations. Mean values were expressed as a fold change over an average of the pretreatment values. There was not a significant treatment x time interaction (P = 0.63) or an effect of time (P = 0.48) for fold change in LH concentrations (Fig. 4.12). However, there was an effect of treatment (P <

0.0001). GnRH-I induced an increase in LH secretion (1.46 ± 0.16) compared to control animals (1.13 ± 0.17), whereas GnRH-II treatment had no effect (P > 0.05; Fig. 4.12).

Similarly, there was no effect of time (P = 0.93) or treatment by time interaction (P =

0.11) for testosterone fold changes. Like LH, GnRH-I stimulated significantly higher testosterone production (1.74 ± 0.22) than GnRH-II (1.15 ± 0.22) or saline (0.67 ± 0.22). 124

2 LH

1.5

a a trt P < 0.0001 time P = 0.4831 trt*time P = 0.6296

Foldchange 1

0.5 Control GnRH-I GnRH-II

2 Testosterone b

1.5

c trt P < 0.0001 time P = 0.9280 trt*time P = 0.1079

FoldChange 1 a

0.5 Control GnRH-I GnRH-II

Figure 4.12. Relative levels of LH and testosterone after intratesticular administration of saline (control), GnRH-I agonist or GnRH-II agonist. The mean ± SEM values are expressed as a fold change over an average of the pretreatment mean. There was a significant treatment effect but no effects of time or treatment x time interaction for either hormone. a,b,c Bars with alternate superscripts differ (P < 0.01). 125

Notably, the GnRH-II stimulated testosterone secretion was also higher (P < 0.001) than controls (Fig. 4.12). In conclusion, intratesticular injections of GnRH-II stimulated testosterone production in the absence of LH synthesis.

126

Discussion

This study is the first to elucidate a specific function for GnRH-II and its receptor in the testis of boars. First, we discovered 6-fold more GnRHR-II protein in the testis compared to the anterior pituitary gland. Similarly, Millar et al. (2001) demonstrated that the marmoset testis expressed the highest level of GnRHR-II mRNA compared to any other tissue, including the anterior pituitary gland. In the current study, GnRH-II was detected in tissue homogenates of the hypothalamus, anterior pituitary gland and testis of the boar. Indeed, GnRH-II mRNA has been detected in the hypothalamus (Miyamoto et al., 1984; Lescheid et al., 1997), pituitary (Mongiat et al., 2006) and testis (Mongiat et al.,

2006; An et al., 2008) of other species. However, this is the first report of GnRH-II in pig tissues. Interestingly, GnRH-II levels were significantly elevated in the testis compared to the other tissue types, suggesting a specific biological role for local production of GnRH-II.

Immunohistochemistry revealed the presence of GnRHR-II on the plasma membrane of Leydig cells. The presence of GnRHR-IIs on Leydig cells has implications for a direct role in steroidogenesis. Indeed, boars immunized against GnRH-II displayed reduced testosterone concentrations, whereas LH levels were unaffected (Bowen et al.,

2006). Similarly, primary Leydig cell cultures from GnRH-II immunized boars failed to produce testosterone similar to control animals following LH challenge. Moreover, treatment with the specific GnRH-I antagonist, SB-75, mitigated the response of porcine

Leydig cells to hCG stimulation (Zanella et al., 2000). Given that boars lack the testicular 127

GnRHR-I, the discovery of the GnRHR-II on porcine Leydig cells may further explain the results of these studies.

Concurrently, we identified positive staining for the GnRHR-II on germ cells in the boar testis via IHC and confirmed this discovery by Western blot. Interestingly, the band size differed between testis (60 kDa) and spermatozoa (54 kDa) samples. While

Eicke et al. (2005) also detected marmoset GnRHR-II protein at 54 kDa in the ovary, this variation in band size may be due to alternative post-translational modifications in different cell types. For example, the GnRHR-I has been shown to have a molecular mass of 50-60 kDa in the anterior pituitary gland, whereas the GnRHR-I present on mature sperm has a band size of 45 kDa (Lee et al., 2000). Specifically, glycosylation events have been reported in the sperm maturation process for numerous species including boars (Topfer-Petersen et al., 1990). Therefore, the alternative band size we detected may indicate differential modification of the GnRHR-II on mature sperm.

In the current study, the GnRHR-II was localized on spermatozoa via immunocytochemistry. Interestingly, the signal was detected in the neck region of the tail, just posterior to the head and at the anterior portion of the midpiece. While the precise function of the GnRHR-II in this region is still unknown, its anatomical location indicates that it could be important for a number of sperm functions including fertilization, motility, the acrosome reaction and capacitation. For example, the GnRHR-

II may interact with the sperm centrosome, which is required for fertilization of the oocyte and embryonic development in many species, like humans (Sathananthan et al.,

1996), sheep (Crozet, 1990), and pigs (Szollosi and Hunter, 1973). In addition to well 128 characterized roles in motility (Buffone et al., 2012), the tail may act as a sensory cilium

(Bloodgood, 2010), detecting alterations in the environment and initiating signals that could influence the acrosome reaction (Buffone et al., 2012). Moreover, calcium mobilization is a key contributor to sperm capacitation and intracellular stores are released from the redundant nuclear envelope, located in the neck region of sperm (Ho and Suarez, 2003). In fact, an inositol 1,4,5-trisphosphate (IP3) receptor-gated calcium store is released from the base of the flagellum prior to hypermotility, however, the receptor mediating this action has not yet been identified (Ho and Suarez, 2003).

Interestingly, our laboratory also detected GnRH-II in the seminal plasma of boars, further indicative of a role for the GnRHR-II on sperm. While the presence of GnRH-II in seminal plasma has not been examined previously, GnRHR-II mRNA has been detected in the human and marmoset uterus, oviduct and the ovary (Neill et al., 2001; Millar et al.,

2001). Thus, sows and gilts may maintain functional GnRHR-IIs within the reproductive tract that could interact with GnRH-II in the seminal plasma of boars.

Here we also demonstrated that porcine testicular explants have the capacity to elicit testosterone production. This finding is in agreement with many previous studies in other species, as well as the pig (Stewart and Raeside, 1976; Zanella et al., 2000; Oatley et al., 2004; Lambrot et al., 2006; Hallmark et al., 2007). We observed that short-term treatment with GnRH-II stimulated testosterone secretion to the same level as hCG, indicating that GnRH-II is an effective stimulator of testosterone synthesis directly at the testis. However, treatment with GnRH-II prior to hCG did not elicit a synergistic effect on testosterone synthesis, suggesting that GnRH-II is not acting through the LHR. This 129 was confirmed by Western blot analysis, as the quantity of LHR protein remained consistent across treatments. However, it is important to note that tissues were collected 3 h after GnRH-II treatment in our study. In rats, upregulation of the LHR required between 3 and 6 h after a single dose of hCG (1-10 µg; Dufau et al., 1984). Thus, LHR upregulation may have occurred with an extended culture period.

In the present study, boars challenged with GnRH-I and -II demonstrated differential LH secretion over time. It is well known that GnRH-I stimulates LH secretion in the boar (Wise et al., 1996). Additionally, high doses of GnRH-II can stimulate low levels of LH production in other species (Millar and King, 1983; Millar et al., 1986).

However, this effect is likely mediated through interaction with the GnRHR-I, as administration of a GnRH-I antagonist attenuates GnRH-II induced gonadotropin secretion (Densmore and Urbanski, 2003; Okada et al., 2003; Kauffman et al., 2005). In the current study, LH is produced but markedly reduced in GnRH-II treated boars, suggesting GnRH-II may have activated the GnRHR-I in the anterior pituitary gland.

Despite reduced LH levels, testosterone concentrations in boars treated with GnRH-II mirrored levels seen in GnRH-I treated animals. This discrepancy indicates that GnRH-II may mediate its action directly at the testis.

Next, boars were treated with a specific GnRH-I (SB-75) or GnRH-II (Trp-1) antagonist. In fact, this is the first in vivo examination of Trp-1 on the HPG axis.

Compared to pretreatment levels, LH was not significantly suppressed in either treatment group. Although, SB-75 induced, LH suppression has been demonstrated in boars at doses of 5 (Wise et al., 2000), 10 (Zanella et al., 2000) and 20 µg/kg BW (Wise et al., 130

2000), recent experimental (Horvath et al., 2004) and clinical (Gonzalez-Barcena et al.,

1994; Comaru-Schally et al., 1998; Drewa and Chlosta, 2009) trials demonstrated that low doses of SB-75 may cause only partial suppression of the HPG axis (Rick et al.,

2011). Therefore, the lack of SB-75 induced, LH suppression demonstrated in this trial may be a reflection of the dose utilized or differing experimental conditions. The effects of Trp-1 on LH secretion have not been previously examined, but in the current study

Trp-1 treatment did not impact LH production. This finding may support the hypothesis that GnRHR-II is not involved in gonadotropin secretion in the boar, as suggested in other species (Densmore and Urbanski, 2003; Okada et al., 2003; Schneider and Rissman, 2008).

In contrast, SB-75 reduced testosterone production at 3, 6 and 12 h post- treatment, consistent with reports that maximal testosterone suppression was achieved at

8-12 h following SB-75 treatment (Behre et al., 1992; Klingmuller et al., 1993).

Interestingly, reduced testosterone levels at these time points were not correlated with

LH. Perhaps SB-75 elicited its effects at the level of the testis, given that it can bind the

GnRHR-II with low affinity (Maiti et al., 2003). Although suppression of testosterone release only occurred at 6 h post-treatment, Trp-1 mediated this reduction without affecting LH. Furthermore, the patterns of LH and testosterone return from suppression were similar in SB-75 treated animals, whereas testosterone levels increase post-Trp-1 suppression without a corresponding increase in LH. This differential action of Trp-1 compared to SB-75 may suggest it mitigates testosterone secretion via alternative mechanisms. Perhaps GnRHR-II mediates testosterone synthesis in a constitutive manner 131 in the testis and antagonizing basal production only moderately reduces serum testosterone concentrations.

In the current report, we also pretreated boars with SB-75 prior to administration of GnRH-I or -II agonists. After GnRH-I treatment, SB-75 was unable to suppress LH or testosterone release. This discovery may be related to competition for binding sites between SB-75 and GnRH-I. SB-75 functions, in part, through occupancy of the

GnRHR-I (Halmos et al., 1996), therefore a bolus of competing high affinity ligand

(GnRH-I) may have displaced SB-75 from the GnRHR-I (Ron-El et al., 2000; Fauser et al., 2002). In fact, this phenomenon is utilized frequently in women to prevent ovarian hyperstimulation syndrome during assisted reproduction procedures (Olivennes et al.,

1996). Women pretreated with SB-75 experienced reduced LH levels, although administration of GnRH-I increased LH levels significantly in all 30 patients within 30 min (Felberbaum et al., 1995). After infusion of GnRH-II, relative LH levels did not increase above pretreatment concentrations, suggesting that GnRH-II, unlike GnRH-I, was unable to displace SB-75 from the GnRHR-I. Indeed, GnRH-II has 10-fold lower affinity for the GnRHR-I than GnRH-I (Neill, 2002) and therefore, presumably lacks the binding affinity needed to displace SB-75. Interestingly, testosterone was produced in a manner consistent with GnRH-I treatment, suggesting testosterone production occurred independent of LH. This finding is in agreement with other studies who demonstrated testosterone can be regulated in the boar without affecting LH secretion (Zanella et al.,

2000; Bowen et al., 2006). 132

In our final experiment, we injected saline, GnRH-I or GnRH-II directly into the testis of sedated boars. Here, we demonstrated that intratesticular administration of

GnRH-I can stimulate LH and testosterone secretion, suggesting that GnRH-I was able to travel through the blood to the anterior pituitary gland. However, a much lower dose of each agonist was utilized in this trial compared to Experiment 1 (300 ng/boar versus 150 ng/kg BW, respectively). Consistent with this, intratesticular injection of rats (which have testicular GnRHR-Is; Sharpe and Cooper, 1987) with 1 ng of a GnRH-I agonist stimulated testosterone, but not LH secretion, whereas a 10 ng injection of GnRH-I significantly elevated both LH and testosterone (Sharpe et al., 1983). These results indicated that lower doses of GnRH-I can stimulate steroidogenesis independent of LH, however, higher doses can be transported through the blood to the anterior pituitary gland. In our experiment, GnRH-II did not stimulate LH release, even though it presumably traveled through the blood like GnRH-I. Despite this, GnRH-II was able to stimulate testosterone synthesis, suggesting it had a direct testicular effect.

Taken together, these data indicate that, as in other species, the predominant function of GnRH-II and its receptor in the boar does not appear to be gonadotropin secretion. The interaction between GnRH-II and the GnRHR-II appears to elicit testosterone synthesis in an autocrine or paracrine manner, without production of LH by the anterior pituitary gland. However, further studies utilizing hypophysectomized boars might be necessary to completely delineate the action of GnRH-I versus GnRH-II.

Physiologically, it appears GnRH-II may play a role in constitutive regulation of testosterone production in the testis. This effect may be required to support 133 spermatogenesis synergistically with the pulsatile action of LH. Moreover, the discovery of GnRHR-II on boar spermatozoa as well as GnRH-II in seminal plasma may reflect a role in spermatogenesis or sperm function.

In conclusion, this study is the first demonstration of a specific function for the

GnRHR-II in the testis. Differential regulation of testosterone secretion could lead to advancements in human fertility treatments, contraceptives, cancer research or pharmacological agents. Moreover, novel screening methods could be utilized in boar studs to improve reproductive efficiency. Eliminating infertile or subfertile boars from the herd early will reduce the economic losses associated with rearing and training.

Furthermore, the discovery of GnRHR-II on sperm and GnRH-II in seminal plasma may potentially lead to the development of innovative strategies to improve conception rates on the sow farm. Implementation of these new technologies could lead to enhanced productivity and profitability of pork producers.

134

Appendix I

Boar Cannulation Procedures

Preparation. Prior to Cannulation, 1.5 m of tygon microbore tubing (ID 0.050 in,

OD 0.090 in, Wall 0.020 in, formulation S-54-HL) was cut and pretreated with tridodecylmethylammonium chloride (TDMAC; Polysciences, Inc., Warrington, PA).

Briefly, about 10 ml of TDMAC was poured into a 100 ml beaker secured with a ring stand. Care was taken to constantly keep the TDMAC covered with Saran™ wrap as it evaporates quickly (do not perform in hood). Next, about 30 cm of the tubing was submerged and a 10 ml syringe was used to draw up TDMAC into the entire cannula for

2 min. Finally, the TDMAC was flushed out with 100 ml of air and allowed to dry for 2 d. After drying, the cannula (at the end that was submerged in TDMAC) was well marked with sharpie at 18, 24 and 30 cm and placed into a gas sterilization bag with an adapter

(18 gauge needle with 2/3 of the needle cut off with a Dremel tool) and stopper (1 ml syringe cut off and melted at 0.3 ml) attached loosely. Cannula packs were then gas sterilized at the University of Nebraska Medical Center sterile processing lab (Omaha,

NE).

Non-Surgical Cannulation. Boars were non-surgically cannulated according to

Barb et al. (1982). Boars were snared by use of both a rope and mechanical snare. The neck was exposed and cleaned with Betadine spray. Next, a sterilized 11 gauge needle 135 with vet hub was inserted into the neck until blood was apparent on drawback with a 20 ml luerlock syringe. Then, the syringe was removed and the sterile cannula was threaded through the needle into the jugular vein. Testing the cannula for blood occurred 3 times as threading progressed (at approximately 18, 24 and 30 cm) using a 10 ml syringe filled with 3.5% sodium citrate. Once fully inserted, the needle was removed, the adapter was secured to the tubing with small, thin strips of duct tape and the cannula was flushed again with 5 ml of a sterile heparinized saline solution (0.9% NaCl, 500 units heparin, 1% benzyl alcohol, 5,000 units penicillin; pH 7.4) and capped. Next, the cannula was coiled up and inserted into a 3.5 x 4 in duct tape pouch. Finally, the cannula and pouch would be secured to the animal by wrapping the chest area twice around with 4 in Elastikon adhesive tape followed by 4 in duct tape. Banamine (2 mg/kg BW) and procaine penicillin (300,000 units/lb BW) were given intramuscularly to minimize pain and inflammation. After insertion, blood clotting was prevented by daily catheter infusions with approximately 10 ml of 3.5% sodium citrate and 3 ml of heparinized saline solution.

Surgical Cannulation. Boars were surgically cannulated according to Ford and Maurer (1978). Briefly, boars were sedated with 5 ml of a telazol, atropine, rompun and ketamine (TARK; 0.2 mg/kg atropine, 0.8 mg/kg rompun, 3.1 mg/kg ketamine and

0.8 mg/kg telazol) mixture, I.M. in the neck. Then, animals were given an additional 3 ml

TARK (0.1 mg/kg atropine, 0.5 mg/kg rompun, 2.1 mg/kg ketamine and 0.5 mg/kg telazol) intravenously via an ear vein. Once anesthetized, the animal was placed onto the surgery cart in a dorsal recumbent position and the surgical site (ventral cervical area) 136 was prepared by removing hair with clippers, performing a Betadine scrub and shaving with a razor. The site was then disinfected with 3 Betadine scrubs and water removed with 70% isopropyl rubbing alcohol swabs. The front feet were covered with plastic O.B. gloves and restrained with rope to each side of the animal. Nose tubes were inserted into each nostril and secured before administration of the inhalant anesthetic isoflurane (3.5%) and oxygen (0.7 L/min).

Hereafter, surgical cannulation was performed according to Ford and Maurer,

(1978). Using sterile gloves, a 1 cm skin incision was made approximately 4-8 cm anterior to the manubrium sterni and 4-5 cm lateral to the left side of the midline, exposing the cervical musculature and subcutaneous tissue. Blunt dissection was performed until the approximate location of the external jugular vein was palpable. The vein was then punctured with an 11 gauge, 3 in needle (short beveled, thin walled) with vet hub attached to an empty 3 ml slip tip syringe. Once blood was evident upon drawback, the needle was removed and a 1.5 m sterilized catheter filled with 3.5% sodium citrate was inserted slowly into the vein. During insertion, the presence of blood was confirmed 3 times to ensure adequate catheter function using a 10 ml syringe filled with 3.5% sodium citrate attached to the catheter and a 17 gauge female luer adapter.

After insertion, the animal was moved to a right lateral recumbent position on the cart, exposing the back. Care was taken to hold the catheter in place during the move.

The left ventral side of the neck and down to the scapula was clipped of hair. The site was then washed with Betadine and cleared with 70% alcohol swabs. Next, a 30 cm long stainless steel cannula containing a trocar was inserted into the previous incision adjacent 137 to the catheter. It was then passed dorsally through the lateral musculature to a midway point between the neck and back. The trocar was exteriorized using another 1 cm incision. Next, the trocar was removed, leaving the hollow stainless steel cannula. The loose end of the catheter was then passed through the cannula, and the cannula was removed from the tissue. The process was repeated from the midpoint of the neck to the back, such that the free end of the catheter was exteriorized in a final location anterior to the scapula. During the cannulation process, the catheter was continually tested to ensure function was retained. The 1st and 2nd incisions were then sutured closed with non- dissolvable suture.

Next, protective tubing (termed a collar) was placed around the catheter to anchor it to the animal. The collar was made by inverting a 2.54 cm of amber rubber tubing (ID:

3.2 mm, OD: 6.4 mm) back on itself and secured together with 20 cm of suture. Once the collar was placed around the catheter, the animals head was flexed towards the sternum and the sutures from the collar were secured to the animal directly over the last incision, using a surgical needle. The collar was then adhered to the catheter with a few drops of cyclohexanone at both openings of the collar. Next, the collar-catheter complex was further adhered to the animal at the exteriorization point with aquarium silicone

(containing acetic acid; Marineland, Blacksburg, VA) to prevent bacterial contamination of the incision. The catheter was checked once more, flushed with 5 ml of 3.5% sodium citrate and 5 ml of heparinized saline and stoppered. The catheter was secured further by adhering 4 in Elastikon adhesive strips to the animal with hip tag cement. Strips were adhered ~1 cm to the right and left of the catheter (12 in strips), as well as 1 cm from the 138 top and bottom (7 in strips) of the catheter. One final strip was placed over the catheter, taking care to keep the tag cement on the tape surrounding the catheter. Finally, each animal was injected with (I.M.) with 3 ml of Polyflex (8.8 mg/kg; Fort Dodge Animal

Health, Fort Dodge, IA) and 2 ml of Banamine (2 mg/kg; Schering-Plough Animal

Health Corp., Union, NJ).

139

Literature Cited

Allrich, R. D., R. K. Christenson, J. J. Ford, and D. R. Zimmerman. 1983. Pubertal development of the boar: Age-related changes in testicular morphology and in vitro production of testosterone and estradiol-17 beta. Biol. Reprod. 28(4): 902-909.

Amann, R., D. Katz, and C. Wang. 1995. What is Semen? How does Semen Analysis Assist in Understanding the Reproductive Status of the Male? Pages 22-29 in the Handbook of Andrology. Robaire, B., J. Pryor, and J. Trasler, eds. American Society of Andrology, Schaumburg, IL.

An, K. W., E. R. Nelson, H. R. Habibi, and C. Y. Choi. 2008. Molecular characterization and expression of three GnRH forms mRNA during gonad sex-change process, and effect of GnRHa on GTH subunits mRNA in the protandrous black porgy (acanthopagrus schlegeli). Gen. Comp. Endocrinol. 159(1): 38-45.

Anderson, R. G. 1998. The caveolae membrane system. Annu. Rev. Biochem. 67: 199- 225.

Ando, H., C. L. Hew, and A. Urano. 2001. Signal transduction pathways and transcription factors involved in the gonadotropin-releasing hormone-stimulated gonadotropin subunit gene expression. Comp. Biochem. Physiol. B. Biochem. Mol. Biol. 129(2-3): 525-532.

Anjum, S., A. Krishna, R. Sridaran, and K. Tsutsui. 2012. Localization of gonadotropin- releasing hormone (GnRH), gonadotropin-inhibitory hormone (GnIH), kisspeptin and GnRH receptor and their possible roles in testicular activities from birth to senescence in mice. J. Exp. Zool. A. Ecol. Genet. Physiol. 317(10): 630-644.

Baba, Y., H. Matsuo, and A. V. Schally. 1971. Structure of the porcine LH- and FSH- releasing hormone. II. confirmation of the proposed structure by conventional sequential analyses. Biochem. Biophys. Res. Commun. 44(2): 459-463.

Bahk, J. Y., J. S. Hyun, S. H. Chung, H. Lee, M. O. Kim, B. H. Lee et al. 1995. Stage specific identification of the expression of GnRH mRNA and localization of the GnRH receptor in mature rat and adult human testis. J. Urol. 154(5): 1958-1961.

Barb, C. R., R. R. Kraeling, G. B. Rampacek, E. S. Fonda, and T. E. Kiser. 1982. Inhibition of ovulation and LH secretion in the gilt after treatment with ACTH or hydrocortisone. J. Reprod. Fertil. 64(1): 85-92.

Barnett, D. K., T. M. Bunnell, R. P. Millar, and D. H. Abbott. 2006. Gonadotropin- releasing hormone II stimulates female sexual behavior in marmoset monkeys. Endocrinology. 147(1): 615-623. 140

Barry, J. 1979. Immunohistochemistry of luteinizing hormone-releasing hormone- producing neurons of the vertebrates. Int. Rev. Cytol. 60: 179-221.

Beckers, T., K. Marheineke, H. Reilander, and P. Hilgard. 1995. Selection and characterization of mammalian cell lines with stable over-expression of human pituitary receptors for gonadoliberin. Eur. J. Biochem. 231(3): 535-543.

Behre, H. M., A. Bockers, A. Schlingheider, and E. Nieschlag. 1994. Sustained suppression of serum LH, FSH and testosterone and increase of high-density lipoprotein cholesterol by daily injections of the GnRH antagonist cetrorelix over 8 days in normal men. Clin. Endocrinol. (Oxf). 40(2): 241-248.

Behre, H. M., B. Klein, E. Steinmeyer, G. P. McGregor, K. Voigt, and E. Nieschlag. 1992. Effective suppression of luteinizing hormone and testosterone by single doses of the new gonadotropin-releasing hormone antagonist cetrorelix (SB-75) in normal men. J. Clin. Endocrinol. Metab. 75(2): 393-398.

Behre, H. M., S. Kliesch, G. Puhse, T. Reissmann, and E. Nieschlag. 1997. High loading and low maintenance doses of a gonadotropin-releasing hormone antagonist effectively suppress serum luteinizing hormone, follicle-stimulating hormone, and testosterone in normal men. J. Clin. Endocrinol. Metab. 82(5): 1403-1408.

Berndtson, W. E. 1977. Methods for quantifying mammalian spermatogenesis: A review. J. Anim. Sci. 44(5): 818-833.

Bertram, G., S. Innes, O. Minella, J. Richardson, and I. Stansfield. 2001. Endless possibilities: Translation termination and stop codon recognition. Microbiology. 147(Pt 2): 255-269.

Bloodgood, R. A. 2010. Sensory reception is an attribute of both primary cilia and motile cilia. J. Cell. Sci. 123(Pt 4): 505-509.

Bokaei, P. B., X. Z. Ma, B. Byczynski, J. Keller, D. Sakac, S. Fahim et al. 2006. Identification and characterization of five-transmembrane isoforms of human vasoactive intestinal peptide and pituitary adenylate cyclase-activating polypeptide receptors. Genomics. 88(6): 791-800.

Bokser, L., S. Bajusz, K. Groot, and A. V. Schally. 1990. Prolonged inhibition of luteinizing hormone and testosterone levels in male rats with the luteinizing hormone-releasing hormone antagonist SB-75. Proc. Natl. Acad. Sci. U. S. A. 87(18): 7100-7104.

Bokser, L., G. Srkalovic, K. Szepeshazi, and A. V. Schally. 1991. Recovery of pituitary- gonadal function in male and female rats after prolonged administration of a potent 141

antagonist of luteinizing hormone-releasing hormone (SB-75). Neuroendocrinology. 54(2): 136-145.

Borg, K. E., D. D. Lunstra, and R. K. Christenson. 1993. Semen characteristics, testicular size, and reproductive hormone concentrations in mature duroc, meishan, fengjing, and minzhu boars. Biol. Reprod. 49(3): 515-521.

Botte, M. C., Y. Lerrant, A. Lozach, A. Berault, R. Counis, and M. L. Kottler. 1999. LH down-regulates gonadotropin-releasing hormone (GnRH) receptor, but not GnRH, mRNA levels in the rat testis. J. Endocrinol. 162(3): 409-415.

Bowen, A., S. Khan, L. Berghman, J. D. Kirby, R. P. Wettemann, and J. A. Vizcarra. 2006. Immunization of pigs against chicken gonadotropin-releasing hormone-II and lamprey gonadotropin-releasing hormone-III: Effects on gonadotropin secretion and testicular function. J. Anim. Sci. 84(11): 2990-2999.

Brahmkshtri, B. P., M. J. Edwin, M. C. John, A. M. Nainar, and A. R. Krishnan. 1999. Relative efficacy of conventional sperm parameters and sperm penetration bioassay to assess bull fertility in vitro. Anim. Reprod. Sci. 54(3): 159-168.

Brauer, V.M. 2009. Transcriptional Regulation of the Porcine Type II GnRH Receptor Gene. Thesis. University of Nebraska-Lincoln.

Brooks, R. I., and A. M. Pearson. 1986. Steroid hormone pathways in the pig, with special emphasis on boar odor: A review. J. Anim. Sci. 62(3): 632-645.

Brothers, S. P., J. A. Janovick, G. Maya-Nunez, A. Cornea, X. B. Han, and P. M. Conn. 2002. Conserved mammalian gonadotropin-releasing hormone receptor carboxyl terminal amino acids regulate ligand binding, effector coupling and internalization. Mol. Cell. Endocrinol. 190(1-2): 19-27.

Browning, J. Y., R. D'Agata, A. Steinberger, H. E. Grotjan Jr, and E. Steinberger. 1983. Biphasic effect of gonadotropin-releasing hormone and its agonist analog (HOE766) on in vitro testosterone production by purified rat leydig cells. Endocrinology. 113(3): 985-991.

Buffone, M. G., T. W. Ijiri, W. Cao, T. Merdiushev, H. K. Aghajanian, and G. L. Gerton. 2012. Heads or tails? structural events and molecular mechanisms that promote mammalian sperm acrosomal exocytosis and motility. Mol. Reprod. Dev. 79(1): 4- 18.

Bull, P., P. Morales, C. Huyser, T. Socias, and E. A. Castellon. 2000. Expression of GnRH receptor in mouse and rat testicular germ cells. Mol. Hum. Reprod. 6(7): 582- 586. 142

Casper, R. F. 1991. Clinical uses of gonadotropin-releasing hormone analogues. CMAJ. 144(2): 153-158.

Chandolia, R. K., A. Honaramooz, B. C. Omeke, R. Pierson, A. P. Beard, and N. C. Rawlings. 1997. Assessment of development of the testes and accessory glands by ultrasonography in bull calves and associated endocrine changes. Theriogenology. 48(1): 119-132.

Chen, A., E. Kaganovsky, S. Rahimipour, N. Ben-Aroya, E. Okon, and Y. Koch. 2002. Two forms of gonadotropin-releasing hormone (GnRH) are expressed in human breast tissue and overexpressed in breast cancer: A putative mechanism for the antiproliferative effect of GnRH by down-regulation of acidic ribosomal phosphoproteins P1 and P2. Cancer Res. 62(4): 1036-1044.

Chen, C. L., L. W. Cheung, M. T. Lau, J. H. Choi, N. Auersperg, H. S. Wang et al. 2007. Differential role of gonadotropin-releasing hormone on human ovarian epithelial cancer cell invasion. Endocrine. 31(3): 311-320.

Cheng, C. K., and P. C. Leung. 2005. Molecular biology of gonadotropin-releasing hormone (GnRH)-I, GnRH-II, and their receptors in humans. Endocr. Rev. 26(2): 283-306.

Cheng, K. W., and P. C. Leung. 2001. Human gonadotropin-releasing hormone receptor gene transcription: Up-regulation by 3',5'-cyclic adenosine monophosphate/protein kinase A pathway. Mol. Cell. Endocrinol. 181(1-2): 15-26.

Cheon, K. W., H. S. Lee, I. S. Parhar, and I. S. Kang. 2001. Expression of the second isoform of gonadotrophin-releasing hormone (GnRH-II) in human endometrium throughout the menstrual cycle. Mol. Hum. Reprod. 7(5): 447-452.

Cheung, L. W., P. C. Leung, and A. S. Wong. 2006. Gonadotropin-releasing hormone promotes ovarian cancer cell invasiveness through c-jun NH2-terminal kinase- mediated activation of matrix metalloproteinase (MMP)-2 and MMP-9. Cancer Res. 66(22): 10902-10910.

Childs, G. V. 1983. Application of dual pre-embedding stains for gonadotropins to pituitary cell monolayers with avidin-biotin (ABC) and peroxidase-antiperoxidase (PAP) complexes: Light microscopic studies. Biotechnic & Histochemistry. 58(5): 281-289.

Choi, K. C., N. Auersperg, and P. C. Leung. 2001. Expression and antiproliferative effect of a second form of gonadotropin-releasing hormone in normal and neoplastic ovarian surface epithelial cells. J. Clin. Endocrinol. Metab. 86(10): 5075-5078. 143

Chou, C. S., C. D. MacCalman, and P. C. Leung. 2003a. Differential effects of gonadotropin-releasing hormone I and II on the urokinase-type plasminogen activator/plasminogen activator inhibitor system in human decidual stromal cells in vitro. J. Clin. Endocrinol. Metab. 88(8): 3806-3815.

Chou, C. S., H. Zhu, C. D. MacCalman, and P. C. Leung. 2003b. Regulatory effects of gonadotropin-releasing hormone (GnRH) I and GnRH II on the levels of matrix metalloproteinase (MMP)-2, MMP-9, and tissue inhibitor of metalloproteinases-1 in primary cultures of human extravillous cytotrophoblasts. J. Clin. Endocrinol. Metab. 88(10): 4781-4790.

Chou, C. S., H. Zhu, E. Shalev, C. D. MacCalman, and P. C. Leung. 2002. The effects of gonadotropin-releasing hormone (GnRH) I and GnRH II on the urokinase-type plasminogen activator/plasminogen activator inhibitor system in human extravillous cytotrophoblasts in vitro. J. Clin. Endocrinol. Metab. 87(12): 5594-5603.

Claing, A., S. J. Perry, M. Achiriloaie, J. K. Walker, J. P. Albanesi, R. J. Lefkowitz et al. 2000. Multiple endocytic pathways of G protein-coupled receptors delineated by GIT1 sensitivity. Proc. Natl. Acad. Sci. U. S. A. 97(3): 1119-1124.

Clapper, J. A. 2008. Effects of two different anaesthetics on serum concentrations of cortisol and luteinizing hormone in barrows and gilts. Lab. Anim. 42(1): 83-91.

Clayton, R. N., L. Eccleston, F. Gossard, J. C. Thalbard, and G. Morel. 1992. Rat granulosa cells express the gonadotrophin-releasing hormone gene: Evidence from in-situ hybridization histochemistry. J. Mol. Endocrinol. 9(3): 189-195.

Clayton, R. N., M. Katikineni, V. Chan, M. L. Dufau, and K. J. Catt. 1980. Direct inhibition of testicular function by gonadotropin-releasing hormone: Mediation by specific gonadotropin-releasing hormone receptors in interstitial cells. Proc. Natl. Acad. Sci. U. S. A. 77(8): 4459-4463.

Clayton, R. N., R. A. Shakespear, J. A. Duncan, J. C. Marshall, P. J. Munson, and D. Rodbard. 1979. Radioiodinated nondegradable gonadotropin-releasing hormone analogs: New probes for the investigation of pituitary gonadotropin-releasing hormone receptors. Endocrinology. 105(6): 1369-1376.

Comaru-Schally, A. M., W. Brannan, A. V. Schally, M. Colcolough, and M. Monga. 1998. Efficacy and safety of luteinizing hormone-releasing hormone antagonist cetrorelix in the treatment of symptomatic benign prostatic hyperplasia. J. Clin. Endocrinol. Metab. 83(11): 3826-3831. 144

Conley, A. 1992. Characterization of cDNA encoding porcine testicular cytochrome p450 17alpha-hydroxylase. Proceedings of the 74th Annual Meeting of the Endocrine Society. : 1501. (Abstr.)

Conley, A. J., and I. M. Bird. 1997. The role of cytochrome P450 17 alpha-hydroxylase and 3 beta-hydroxysteroid dehydrogenase in the integration of gonadal and adrenal steroidogenesis via the delta 5 and delta 4 pathways of steroidogenesis in mammals. Biol. Reprod. 56(4): 789-799.

Conn, P. M., and W. F. Crowley Jr. 1991. Gonadotropin-releasing hormone and its analogues. N. Engl. J. Med. 324(2): 93-103.

Cook, T., and W. P. Sheridan. 2000. Development of GnRH antagonists for prostate cancer: New approaches to treatment. Oncologist. 5(2): 162-168.

Cooke, G. M. 1991. Steroidogenesis in interstitial cells and microsomal fraction of immature pig testes. J. Reprod. Fertil. 91(1): 175-185.

Correa, J. R., M. M. Pace, and P. M. Zavos. 1997. Relationships among frozen-thawed sperm characteristics assessed via the routine semen analysis, sperm functional tests and fertility of bulls in an artificial insemination program. Theriogenology. 48(5): 721-731.

Crozet, N. 1990. Behavior of the sperm centriole during sheep oocyte fertilization. Eur. J. Cell Biol. 53(2): 326-332.

De Camilli, P., K. Takei, and P. S. McPherson. 1995. The function of dynamin in endocytosis. Curr. Opin. Neurobiol. 5(5): 559-565.

De Lean, A., J. M. Stadel, and R. J. Lefkowitz. 1980. A ternary complex model explains the agonist-specific binding properties of the adenylate cyclase-coupled beta- adrenergic receptor. J. Biol. Chem. 255(15): 7108-7117. de Rooij, D. G., and L. D. Russell. 2000. All you wanted to know about spermatogonia but were afraid to ask. J. Androl. 21(6): 776-798.

Densmore, V. S., and H. F. Urbanski. 2003. Relative effect of gonadotropin-releasing hormone (GnRH)-I and GnRH-II on gonadotropin release. J. Clin. Endocrinol. Metab. 88(5): 2126-2134.

Dong, K. W., Z. G. Chen, K. W. Cheng, and K. L. Yu. 1996. Evidence for estrogen receptor-mediated regulation of human gonadotropin-releasing hormone promoter activity in human placental cells. Mol. Cell. Endocrinol. 117(2): 241-246. 145

Drewa, T., and P. Chlosta. 2009. Re: Frans debruyne, arkadij A. gres, dmitrii L. arustamov. placebo-controlled dose-ranging phase 2 study of subcutaneously administered LHRH antagonist cetrorelix in patients with symptomatic benign prostatic hyperplasia. eur urol 2008;54:170-80. Eur. Urol. 55(2): e36-7; author reply e38-9.

Dubois, E. A., M. A. Zandbergen, J. Peute, and H. J. Goos. 2002. Evolutionary development of three gonadotropin-releasing hormone (GnRH) systems in vertebrates. Brain Res. Bull. 57(3-4): 413-418.

Dufau, M. L., C. A. Winters, M. Hattori, D. Aquilano, J. L. Baranao, K. Nozu et al. 1984. Hormonal regulation of androgen production by the leydig cell. J. Steroid Biochem. 20(1): 161-173.

Dyck, M., G. Foxcroft, S. Novak, A. Ruiz-Sanchez, J. Patterson, and W. Dixon. 2011. Biological markers of boar fertility. Reproduction in Domestic Animals. 46: 55-58.

Eddy, E. M. 2002. Male germ cell gene expression. Recent Prog. Horm. Res. 57: 103- 128.

Eicke, N., A. R. Gunthert, G. Emons, and C. Grundker. 2006. GnRH-II agonist [D- Lys6]GnRH-II inhibits the EGF-induced mitogenic signal transduction in human endometrial and ovarian cancer cells. Int. J. Oncol. 29(5): 1223-1229.

Eicke, N., A. R. Gunthert, V. Viereck, D. Siebold, M. Behe, T. Becker et al. 2005. GnRH-II receptor-like antigenicity in human placenta and in cancers of the human reproductive organs. Eur. J. Endocrinol. 153(4): 605-612.

Eidne, K. A., R. E. Sellar, G. Couper, L. Anderson, and P. L. Taylor. 1992. Molecular cloning and characterisation of the rat pituitary gonadotropin-releasing hormone (GnRH) receptor. Mol. Cell. Endocrinol. 90(1): R5-9.

Elmore, S. 2007. Apoptosis: A review of programmed cell death. Toxicol. Pathol. 35(4): 495-516.

Emons, G., C. Grundker, A. R. Gunthert, S. Westphalen, J. Kavanagh, and C. Verschraegen. 2003. GnRH antagonists in the treatment of gynecological and breast cancers. Endocr. Relat. Cancer. 10(2): 291-299.

Emons, G., O. Ortmann, M. Becker, G. Irmer, B. Springer, R. Laun et al. 1993a. High affinity binding and direct antiproliferative effects of LHRH analogues in human ovarian cancer cell lines. Cancer Res. 53(22): 5439-5446. 146

Emons, G., O. Ortmann, K. D. Schulz, and A. V. Schally. 1997. Growth-inhibitory actions of analogues of luteinizing hormone releasing hormone on tumor cells. Trends Endocrinol. Metab. 8(9): 355-362.

Emons, G., B. Schroder, O. Ortmann, S. Westphalen, K. D. Schulz, and A. V. Schally. 1993b. High affinity binding and direct antiproliferative effects of luteinizing hormone-releasing hormone analogs in human endometrial cancer cell lines. J. Clin. Endocrinol. Metab. 77(6): 1458-1464.

Engel, J. B., and A. V. Schally. 2007. Drug insight: Clinical use of agonists and antagonists of luteinizing-hormone-releasing hormone. Nat. Clin. Pract. Endocrinol. Metab. 3(2): 157-167.

Enomoto, M., M. Utsumi, and M. K. Park. 2006. Gonadotropin-releasing hormone induces actin cytoskeleton remodeling and affects cell migration in a cell-type- specific manner in TSU-Pr1 and DU145 cells. Endocrinology. 147(1): 530-542.

Fauser, B. C., D. de Jong, F. Olivennes, H. Wramsby, C. Tay, J. Itskovitz-Eldor et al. 2002. Endocrine profiles after triggering of final oocyte maturation with GnRH agonist after cotreatment with the GnRH antagonist ganirelix during ovarian hyperstimulation for in vitro fertilization. J. Clin. Endocrinol. Metab. 87(2): 709- 715.

Fawcett, D. W., S. ITO, and D. Slautterback. 1959. The occurrence of intercellular bridges in groups of cells exhibiting synchronous differentiation. J. Biophys. Biochem. Cytol. 5(3): 453-460.

Felberbaum, R. E., T. Reissmann, W. Kupker, O. Bauer, S. al Hasani, C. Diedrich et al. 1995. Preserved pituitary response under ovarian stimulation with HMG and GnRH antagonists (cetrorelix) in women with tubal infertility. Eur. J. Obstet. Gynecol. Reprod. Biol. 61(2): 151-155.

Fernald, R. D., and R. B. White. 1999. Gonadotropin-releasing hormone genes: Phylogeny, structure, and functions. Front. Neuroendocrinol. 20(3): 224-240.

Ferris, H. A., H. E. Walsh, J. Stevens, P. C. Fallest, and M. A. Shupnik. 2007. Luteinizing hormone beta promoter stimulation by adenylyl cyclase and cooperation with gonadotropin-releasing hormone 1 in transgenic mice and LBetaT2 cells. Biol. Reprod. 77(6): 1073-1080.

Fink, G. 1988. Oestrogen and progesterone interactions in the control of gonadotrophin and prolactin secretion. J. Steroid Biochem. 30(1-6): 169-178. 147

Fister, S., A. R. Gunthert, G. Emons, and C. Grundker. 2007. Gonadotropin-releasing hormone type II antagonists induce apoptotic cell death in human endometrial and ovarian cancer cells in vitro and in vivo. Cancer Res. 67(4): 1750-1756.

Flanagan, C. A., W. Zhou, L. Chi, T. Yuen, V. Rodic, D. Robertson et al. 1999. The functional microdomain in transmembrane helices 2 and 7 regulates expression, activation, and coupling pathways of the gonadotropin-releasing hormone receptor. J. Biol. Chem. 274(41): 28880-28886.

Flowers, W. L. 1997. Management of boars for efficient semen production. J. Reprod. Fertil. Suppl. 52: 67-78.

Ford, J. J., and R. R. Maurer. 1978. Simple technique for chronic venous catheterization of swine. Lab. Anim. Sci. 28(5): 615-618.

Ford, J. J., T. H. Wise, D. D. Lunstra, and G. A. Rohrer. 2001. Interrelationships of porcine X and Y chromosomes with pituitary gonadotropins and testicular size. Biol. Reprod. 65(3): 906-912.

Foxcroft, G. R., M. K. Dyck, A. Ruiz-Sanchez, S. Novak, and W. T. Dixon. 2008. Identifying useable semen. Theriogenology. 70(8): 1324-1336.

Franca, L. R., G. F. Avelar, and F. F. Almeida. 2005. Spermatogenesis and sperm transit through the epididymis in mammals with emphasis on pigs. Theriogenology. 63(2): 300-318.

Franca, L. R., and F. M. Cardoso. 1998. Duration of spermatogenesis and sperm transit time through the epididymis in the piau boar. Tissue Cell. 30(5): 573-582.

Frankenhuis, M. T., M. F. Kramer, and D. De Rooij. 1982. Spermatogenesis in the boar. Veterinary Quarterly. 4: 57-61.

Gault, P. M., K. Morgan, A. J. Pawson, R. P. Millar, and G. A. Lincoln. 2004a. Sheep exhibit novel variations in the organization of the mammalian type II gonadotropin- releasing hormone receptor gene. Endocrinology. 145(5): 2362-2374.

Gault, P. M., K. Morgan, A. J. Pawson, R. P. Millar, and G. A. Lincoln. 2004b. Sheep exhibit novel variations in the organization of the mammalian type II gonadotropin- releasing hormone receptor gene. Endocrinology. 145(5): 2362-2374.

Gibbons, J. M.,Jr, M. Mitnick, and V. Chieffo. 1975. In vitro biosynthesis of TSH- and LH-releasing factors by the human placenta. Am. J. Obstet. Gynecol. 121(1): 127- 131. 148

Gonzalez-Barcena, D., M. Vadillo-Buenfil, F. Gomez-Orta, M. Fuentes Garcia, I. Cardenas-Cornejo, A. Graef-Sanchez et al. 1994. Responses to the antagonistic analog of LH-RH (SB-75, cetrorelix) in patients with benign prostatic hyperplasia and prostatic cancer. Prostate. 24(2): 84-92.

Gower, D. B. 1972. The Biosynthesis and Metabolism of Androgens and Other C19 Steroids. Pages 170-206 in Biochemistry of the Steroid Hormones. Makin, H. J., ed. Blackwell Scientific Publications, Oxford.

Grafer, C. M., R. Thomas, L. Lambrakos, I. Montoya, S. White, and L. M. Halvorson. 2009. GnRH stimulates expression of PACAP in the pituitary gonadotropes via both the PKA and PKC signaling systems. Mol. Endocrinol. 23(7): 1022-1032.

Grosse, R., A. Schmid, T. Schoneberg, A. Herrlich, P. Muhn, G. Schultz et al. 2000. Gonadotropin-releasing hormone receptor initiates multiple signaling pathways by exclusively coupling to G(q/11) proteins. J. Biol. Chem. 275(13): 9193-9200.

Grundker, C., and G. Emons. 2003. Role of gonadotropin-releasing hormone (GnRH) in ovarian cancer. Reprod. Biol. Endocrinol. 1: 65.

Grundker, C., C. Fost, S. Fister, N. Nolte, A. R. Gunthert, and G. Emons. 2010. Gonadotropin-releasing hormone type II antagonist induces apoptosis in MCF-7 and triple-negative MDA-MB-231 human breast cancer cells in vitro and in vivo. Breast Cancer Res. 12(4): R49.

Grundker, C., A. R. Gunthert, R. P. Millar, and G. Emons. 2002a. Expression of gonadotropin-releasing hormone II (GnRH-II) receptor in human endometrial and ovarian cancer cells and effects of GnRH-II on tumor cell proliferation. J. Clin. Endocrinol. Metab. 87(3): 1427-1430.

Grundker, C., A. R. Gunthert, S. Westphalen, and G. Emons. 2002b. Biology of the gonadotropin-releasing hormone system in gynecological cancers. Eur. J. Endocrinol. 146(1): 1-14.

Grundker, C., L. Schlotawa, V. Viereck, N. Eicke, A. Horst, B. Kairies et al. 2004a. Antiproliferative effects of the GnRH antagonist cetrorelix and of GnRH-II on human endometrial and ovarian cancer cells are not mediated through the GnRH type I receptor. Eur. J. Endocrinol. 151(1): 141-149.

Grundker, C., L. Schlotawa, V. Viereck, N. Eicke, A. Horst, B. Kairies et al. 2004b. Antiproliferative effects of the GnRH antagonist cetrorelix and of GnRH-II on human endometrial and ovarian cancer cells are not mediated through the GnRH type I receptor. Eur. J. Endocrinol. 151(1): 141-149. 149

Grundker, C., P. Volker, and G. Emons. 2001. Antiproliferative signaling of luteinizing hormone-releasing hormone in human endometrial and ovarian cancer cells through G protein alpha(I)-mediated activation of phosphotyrosine phosphatase. Endocrinology. 142(6): 2369-2380.

Gudermann, T., F. Kalkbrenner, E. Dippel, K. L. Laugwitz, and G. Schultz. 1997. Specificity and complexity of receptor-G-protein interaction. Adv. Second Messenger Phosphoprotein Res. 31: 253-262.

Haley, C., G. Lee, and M. Ritchie. 1995. Comparative reproductive performance in meishan and large white pigs and their crosses. Animal Sciences. 60: 259-267.

Hallmark, N., M. Walker, C. McKinnell, I. K. Mahood, H. Scott, R. Bayne et al. 2007. Effects of monobutyl and di(n-butyl) phthalate in vitro on steroidogenesis and leydig cell aggregation in fetal testis explants from the rat: Comparison with effects in vivo in the fetal rat and neonatal marmoset and in vitro in the human. Environ. Health Perspect. 115(3): 390-396.

Halmos, G., and A. V. Schally. 2002. Changes in subcellular distribution of pituitary receptors for luteinizing hormone-releasing hormone (LH-RH) after treatment with the LH-RH antagonist cetrorelix. Proc. Natl. Acad. Sci. U. S. A. 99(2): 961-965.

Halmos, G., A. V. Schally, J. Pinski, M. Vadillo-Buenfil, and K. Groot. 1996. Down- regulation of pituitary receptors for luteinizing hormone-releasing hormone (LH- RH) in rats by LH-RH antagonist cetrorelix. Proc. Natl. Acad. Sci. U. S. A. 93(6): 2398-2402.

Harwood, J. P., R. N. Clayton, and K. J. Catt. 1980. Ovarian gonadotropin-releasing hormone receptors. I. properties and inhibition of luteal cell function. Endocrinology. 107(2): 407-413.

Hawes, B. E., S. Barnes, and P. M. Conn. 1993. Cholera toxin and pertussis toxin provoke differential effects on luteinizing hormone release, inositol phosphate production, and gonadotropin-releasing hormone (GnRH) receptor binding in the gonadotrope: Evidence for multiple guanyl nucleotide binding proteins in GnRH action. Endocrinology. 132(5): 2124-2130.

Heding, A., M. Vrecl, J. Bogerd, A. McGregor, R. Sellar, P. L. Taylor et al. 1998. Gonadotropin-releasing hormone receptors with intracellular carboxyl-terminal tails undergo acute desensitization of total inositol phosphate production and exhibit accelerated internalization kinetics. J. Biol. Chem. 273(19): 11472-11477. 150

Ho, H. C., and S. S. Suarez. 2003. Characterization of the intracellular calcium store at the base of the sperm flagellum that regulates hyperactivated motility. Biol. Reprod. 68(5): 1590-1596.

Hong, I. S., A. P. Cheung, and P. C. Leung. 2008. Gonadotropin-releasing hormones I and II induce apoptosis in human granulosa cells. J. Clin. Endocrinol. Metab. 93(8): 3179-3185.

Hong, I. S., C. Klausen, A. P. Cheung, and P. C. Leung. 2012. Gonadotropin-releasing hormone-I or -II interacts with IGF-I/akt but not connexin 43 in human granulosa cell apoptosis. J. Clin. Endocrinol. Metab. 97(2): 525-534.

Horvath, J. E., G. L. Toller, A. V. Schally, A. M. Bajo, and K. Groot. 2004. Effect of long-term treatment with low doses of the LHRH antagonist cetrorelix on pituitary receptors for LHRH and gonadal axis in male and female rats. Proc. Natl. Acad. Sci. U. S. A. 101(14): 4996-5001.

Hsueh, A. J., and G. F. Erickson. 1979. Extra-pituitary inhibition of testicular function by luteinising hormone releasing hormone. Nature. 281(5726): 66-67.

Huang, F., Q. Liu, H. Wang, and Y. Zou. 2010. Effect of GnRH II and GnRH I on secretion of VEGF by eutopic and ectopic endometrial stromal cells of endometriosis patients. Zhong Nan Da Xue Xue Bao Yi Xue Ban. 35(5): 409-418.

Huggins, C., and C. V. Hodges. 2002. Studies on prostatic cancer: I. the effect of castration, of estrogen and of androgen injection on serum phosphatases in metastatic carcinoma of the prostate. 1941. J. Urol. 168(1): 9-12.

Hunzicker-Dunn, M., and L. Birnbaumer. 1985. The Involvement of Adenylyl Cyclase and Cyclic AMP-Dependent Protein Kinases in Luteinizing Hormone Actions. Pages 57-134 in Luteinizing Hormone Actions and Receptors. CRC Press, Inc, Boca Raton.

Imai, A., H. Takagi, S. Horibe, T. Fuseya, and T. Tamaya. 1996. Coupling of gonadotropin-releasing hormone receptor to gi protein in human reproductive tract tumors. J. Clin. Endocrinol. Metab. 81(9): 3249-3253.

Irmer, G., C. Burger, R. Muller, O. Ortmann, U. Peter, S. S. Kakar et al. 1995. Expression of the messenger RNAs for luteinizing hormone-releasing hormone (LHRH) and its receptor in human ovarian epithelial carcinoma. Cancer Res. 55(4): 817-822.

Iwashita, M., and K. J. Catt. 1985. Photoaffinity labeling of pituitary and gonadal receptors for gonadotropin-releasing hormone. Endocrinology. 117(2): 738-746. 151

Jegou, B., I. Brekke, O. Naess, P. Torjesen, and V. Hansson. 1985. Properties and regulation of GnRH receptors in the anterior pituitary and the testis of the rat: Different response of leydig cell LH and GnRH receptors to hormonal treatments. Arch. Androl. 14(2-3): 161-170.

Jin, Z., and W. S. El-Deiry. 2005. Overview of cell death signaling pathways. Cancer. Biol. Ther. 4(2): 139-163.

Kaiser, U. B., P. M. Conn, and W. W. Chin. 1997. Studies of gonadotropin-releasing hormone (GnRH) action using GnRH receptor-expressing pituitary cell lines. Endocr. Rev. 18(1): 46-70.

Kakar, S. S., L. C. Musgrove, D. C. Devor, J. C. Sellers, and J. D. Neill. 1992. Cloning, sequencing, and expression of human gonadotropin releasing hormone (GnRH) receptor. Biochem. Biophys. Res. Commun. 189(1): 289-295.

Kakar, S. S., and J. D. Neill. 1995. The human gonadotropin-releasing hormone receptor gene (GNRHR) maps to chromosome band 4q13. Cytogenet. Cell Genet. 70(3-4): 211-214.

Kang, S. K., C. J. Tai, K. W. Cheng, and P. C. Leung. 2000. Gonadotropin-releasing hormone activates mitogen-activated protein kinase in human ovarian and placental cells. Mol. Cell. Endocrinol. 170(1-2): 143-151.

Kang, S. K., C. J. Tai, P. S. Nathwani, and P. C. Leung. 2001. Differential regulation of two forms of gonadotropin-releasing hormone messenger ribonucleic acid in human granulosa-luteal cells. Endocrinology. 142(1): 182-192.

Karten, M. J., and J. E. Rivier. 1986. Gonadotropin-releasing hormone analog design. structure-function studies toward the development of agonists and antagonists: Rationale and perspective. Endocr. Rev. 7(1): 44-66.

Kasten, T. L., S. A. White, T. T. Norton, C. T. Bond, J. P. Adelman, and R. D. Fernald. 1996. Characterization of two new preproGnRH mRNAs in the tree shrew: First direct evidence for mesencephalic GnRH gene expression in a placental mammal. Gen. Comp. Endocrinol. 104(1): 7-19.

Kauffman, A. S., K. Bojkowska, A. Wills, and E. F. Rissman. 2006. Gonadotropin- releasing hormone-II messenger ribonucleic acid and protein content in the mammalian brain are modulated by food intake. Endocrinology. 147(11): 5069- 5077. 152

Kauffman, A. S., and E. F. Rissman. 2004a. A critical role for the evolutionarily conserved gonadotropin-releasing hormone II: Mediation of energy status and female sexual behavior. Endocrinology. 145(8): 3639-3646.

Kauffman, A. S., and E. F. Rissman. 2004b. The evolutionarily conserved gonadotropin- releasing hormone II modifies food intake. Endocrinology. 145(2): 686-691.

Kauffman, A. S., A. Wills, R. P. Millar, and E. F. Rissman. 2005. Evidence that the type- 2 gonadotrophin-releasing hormone (GnRH) receptor mediates the behavioural effects of GnRH-II on feeding and reproduction in musk shrews. J. Neuroendocrinol. 17(8): 489-497.

Kesner, J. S., R. R. Kraeling, G. B. Rampacek, and B. Johnson. 1987. Absence of an estradiol-induced surge of luteinizing hormone in pigs receiving unvarying pulsatile gonadotropin-releasing hormone stimulation. Endocrinology. 121(5): 1862-1869.

Khan, M. A., M. Prevost, M. M. Waterston, M. J. Harvey, and V. A. Ferro. 2007. Effect of immunisation against gonadotrophin releasing hormone isoforms (mammalian GnRH-I, chicken GnRH-II and lamprey GnRH-III) on murine spermatogenesis. Vaccine. 25(11): 2051-2063.

Kim, D. K., J. S. Yang, K. Maiti, J. I. Hwang, K. Kim, D. Seen et al. 2009. A gonadotropin-releasing hormone-II antagonist induces autophagy of prostate cancer cells. Cancer Res. 69(3): 923-931.

Kim, K. Y., K. C. Choi, N. Auersperg, and P. C. Leung. 2006a. Mechanism of gonadotropin-releasing hormone (GnRH)-I and -II-induced cell growth inhibition in ovarian cancer cells: Role of the GnRH-I receptor and protein kinase C pathway. Endocr. Relat. Cancer. 13(1): 211-220.

Kim, K. Y., K. C. Choi, N. Auersperg, and P. C. Leung. 2006b. Mechanism of gonadotropin-releasing hormone (GnRH)-I and -II-induced cell growth inhibition in ovarian cancer cells: Role of the GnRH-I receptor and protein kinase C pathway. Endocr. Relat. Cancer. 13(1): 211-220.

Kim, K. Y., K. C. Choi, S. H. Park, N. Auersperg, and P. C. Leung. 2005. Extracellular signal-regulated protein kinase, but not c-jun N-terminal kinase, is activated by type II gonadotropin-releasing hormone involved in the inhibition of ovarian cancer cell proliferation. J. Clin. Endocrinol. Metab. 90(3): 1670-1677.

Kim, K. Y., K. C. Choi, S. H. Park, C. S. Chou, N. Auersperg, and P. C. Leung. 2004. Type II gonadotropin-releasing hormone stimulates p38 mitogen-activated protein kinase and apoptosis in ovarian cancer cells. J. Clin. Endocrinol. Metab. 89(6): 3020-3026. 153

King, J. A., and R. P. Millar. 1995. Evolutionary aspects of gonadotropin-releasing hormone and its receptor. Cell. Mol. Neurobiol. 15(1): 5-23.

Klingmuller, D., M. Schepke, C. Enzweiler, and F. Bidlingmaier. 1993. Hormonal responses to the new potent GnRH antagonist cetrorelix. Acta Endocrinol. (Copenh). 128(1): 15-18.

Krsmanovic, L. Z., S. S. Stojilkovic, L. M. Mertz, M. Tomic, and K. J. Catt. 1993. Expression of gonadotropin-releasing hormone receptors and autocrine regulation of neuropeptide release in immortalized hypothalamic neurons. Proc. Natl. Acad. Sci. U. S. A. 90(9): 3908-3912.

Kuster, C. E., and G. C. Althouse. 2008. Update on current boar stud practices. Proceedings of Boar Stud Managers Conference III. : 152-155.

Lahlou, N., M. Roger, J. L. Chaussain, M. C. Feinstein, C. Sultan, J. E. Toublanc et al. 1987. Gonadotropin and alpha-subunit secretion during long term pituitary suppression by D-Trp6-luteinizing hormone-releasing hormone microcapsules as treatment of precocious puberty. J. Clin. Endocrinol. Metab. 65(5): 946-953.

Lambrot, R., H. Coffigny, C. Pairault, A. C. Donnadieu, R. Frydman, R. Habert et al. 2006. Use of organ culture to study the human fetal testis development: Effect of retinoic acid. J. Clin. Endocrinol. Metab. 91(7): 2696-2703.

Lee, C. Y., J. Ho, S. N. Chow, K. Yasojima, C. Schwab, and P. L. McGeer. 2000. Immunoidentification of gonadotropin releasing hormone receptor in human sperm, pituitary and cancer cells. Am. J. Reprod. Immunol. 44(3): 170-177.

Lescheid, D. W., E. Terasawa, L. A. Abler, H. F. Urbanski, C. M. Warby, R. P. Millar et al. 1997. A second form of gonadotropin-releasing hormone (GnRH) with characteristics of chicken GnRH-II is present in the primate brain. Endocrinology. 138(12): 5618-5629.

Leung, P. C., C. K. Cheng, and X. M. Zhu. 2003. Multi-factorial role of GnRH-I and GnRH-II in the human ovary. Mol. Cell. Endocrinol. 202(1-2): 145-153.

Levis, D. 2000. Liquid boar semen production: Current extender technology and where do we go from here. Boar Semen Preservation. IV: 121-128.

Limonta, P., R. M. Moretti, M. M. Marelli, D. Dondi, M. Parenti, and M. Motta. 1999. The luteinizing hormone-releasing hormone receptor in human prostate cancer cells: Messenger ribonucleic acid expression, molecular size, and signal transduction pathway. Endocrinology. 140(11): 5250-5256. 154

Limonta, P., R. M. Moretti, M. Montagnani Marelli, and M. Motta. 2003. The biology of gonadotropin hormone-releasing hormone: Role in the control of tumor growth and progression in humans. Front. Neuroendocrinol. 24(4): 279-295.

Lin, C. J., G. C. Wu, M. F. Lee, E. L. Lau, S. Dufour, and C. F. Chang. 2010. Regulation of two forms of gonadotropin-releasing hormone receptor gene expression in the protandrous black porgy fish, acanthopagrus schlegeli. Mol. Cell. Endocrinol. 323(2): 137-146.

Lin, X., J. A. Janovick, S. Brothers, M. Blomenrohr, J. Bogerd, and P. M. Conn. 1998. Addition of catfish gonadotropin-releasing hormone (GnRH) receptor intracellular carboxyl-terminal tail to rat GnRH receptor alters receptor expression and regulation. Mol. Endocrinol. 12(2): 161-171.

Lin, Y. M., S. L. Poon, J. H. Choi, J. S. Lin, P. C. Leung, and B. M. Huang. 2008. Transcripts of testicular gonadotropin-releasing hormone, steroidogenic enzymes, and intratesticular testosterone levels in infertile men. Fertil. Steril. 90(5): 1761- 1768.

Ling Poon, S., M. T. Lau, G. L. Hammond, and P. C. Leung. 2011. Gonadotropin- releasing hormone-II increases membrane type I metalloproteinase production via beta-catenin signaling in ovarian cancer cells. Endocrinology. 152(3): 764-772.

Ling, K., P. Wang, J. Zhao, Y. L. Wu, Z. J. Cheng, G. X. Wu et al. 1999a. Five- transmembrane domains appear sufficient for a G protein-coupled receptor: Functional five-transmembrane domain chemokine receptors. Proc. Natl. Acad. Sci. U. S. A. 96(14): 7922-7927.

Ling, K., P. Wang, J. Zhao, Y. L. Wu, Z. J. Cheng, G. X. Wu et al. 1999b. Five- transmembrane domains appear sufficient for a G protein-coupled receptor: Functional five-transmembrane domain chemokine receptors. Proc. Natl. Acad. Sci. U. S. A. 96(14): 7922-7927.

Liu, J., C. D. Maccalman, Y. L. Wang, and P. C. Leung. 2009. Promotion of human trophoblasts invasion by gonadotropin-releasing hormone (GnRH) I and GnRH II via distinct signaling pathways. Mol. Endocrinol. 23(7): 1014-1021.

Lunstra, D. D., J. J. Ford, J. Klindt, and T. H. Wise. 1997. Physiology of the meishan boar. J. Reprod. Fertil. Suppl. 52: 181-193.

Maiti, K., J. H. Li, A. F. Wang, S. Acharjee, W. P. Kim, W. B. Im et al. 2003. GnRH-II analogs for selective activation and inhibition of non-mammalian and type-II mammalian GnRH receptors. Mol. Cells. 16(2): 173-179. 155

Maiti, K., D. Y. Oh, J. S. Moon, S. Acharjee, J. H. Li, D. G. Bai et al. 2005. Differential effects of gonadotropin-releasing hormone (GnRH)-I and GnRH-II on prostate cancer cell signaling and death. J. Clin. Endocrinol. Metab. 90(7): 4287-4298.

Maney, D. L., R. D. Richardson, and J. C. Wingfield. 1997. Central administration of chicken gonadotropin-releasing hormone-II enhances courtship behavior in a female sparrow. Horm. Behav. 32(1): 11-18.

Marelli, M. M., R. M. Moretti, D. Dondi, M. Motta, and P. Limonta. 1999. Luteinizing hormone-releasing hormone agonists interfere with the mitogenic activity of the insulin-like growth factor system in androgen-independent prostate cancer cells. Endocrinology. 140(1): 329-334.

Matsumiya, K., M. L. Meistrich, G. Shetty, K. Dohmae, A. Tohda, A. Okuyama et al. 1999. Stimulation of spermatogonial differentiation in juvenile spermatogonial depletion (jsd) mutant mice by gonadotropin-releasing hormone antagonist treatment. Endocrinology. 140(10): 4912-4915.

Matsuo, H., Y. Baba, R. M. Nair, A. Arimura, and A. V. Schally. 1971. Structure of the porcine LH- and FSH-releasing hormone. I. the proposed amino acid sequence. Biochem. Biophys. Res. Commun. 43(6): 1334-1339.

Maudsley, S., L. Davidson, A. J. Pawson, R. Chan, R. Lopez de Maturana, and R. P. Millar. 2004. Gonadotropin-releasing hormone (GnRH) antagonists promote proapoptotic signaling in peripheral reproductive tumor cells by activating a galphai- coupling state of the type I GnRH receptor. Cancer Res. 64(20): 7533-7544.

McArdle, C. A., J. Franklin, L. Green, and J. N. Hislop. 2002. Signalling, cycling and desensitisation of gonadotrophin-releasing hormone receptors. J. Endocrinol. 173(1): 1-11.

Meistrich, M. L. 1999. Restoration of spermatogenesis by hormone treatment after cytotoxic therapy. Acta Paediatr. Suppl. 88(433): 19-22.

Millar, R. 2002. Gonadotropin-releasing hormones and their receptors. Reproductive Medicine: Molecular Cellular and Genetic Fundamentals.Lancaster, UK: Parthenon Publishing. : 199-224.

Millar, R., D. Conklin, C. Lofton-Day, E. Hutchinson, B. Troskie, N. Illing et al. 1999. A novel human GnRH receptor homolog gene: Abundant and wide tissue distribution of the antisense transcript. J. Endocrinol. 162(1): 117-126. 156

Millar, R., S. Lowe, D. Conklin, A. Pawson, S. Maudsley, B. Troskie et al. 2001. A novel mammalian receptor for the evolutionarily conserved type II GnRH. Proc. Natl. Acad. Sci. U. S. A. 98(17): 9636-9641.

Millar, R. P. 2005. GnRHs and GnRH receptors. Anim. Reprod. Sci. 88(1-2): 5-28.

Millar, R. P. 2003. GnRH II and type II GnRH receptors. Trends Endocrinol. Metab. 14(1): 35-43.

Millar, R. P., and J. A. King. 1987. Structural and functional evolution of gonadotropin- releasing hormone. Int. Rev. Cytol. 106: 149-182.

Millar, R. P., and J. A. King. 1983. Synthesis, luteinizing hormone-releasing activity, and receptor binding of chicken hypothalamic luteinizing hormone-releasing hormone. Endocrinology. 113(4): 1364-1369.

Millar, R. P., J. A. King, J. S. Davidson, and R. C. Milton. 1987. Gonadotrophin- releasing hormone--diversity of functions and clinical applications. S. Afr. Med. J. 72(11): 748-755.

Millar, R. P., Z. L. Lu, A. J. Pawson, C. A. Flanagan, K. Morgan, and S. R. Maudsley. 2004. Gonadotropin-releasing hormone receptors. Endocr. Rev. 25(2): 235-275.

Millar, R. P., R. C. Milton, B. K. Follett, and J. A. King. 1986. Receptor binding and gonadotropin-releasing activity of a novel chicken gonadotropin-releasing hormone ([His5, Trp7, Tyr8]GnRH) and a D-Arg6 analog. Endocrinology. 119(1): 224-231.

Millar, R. P., and A. J. Pawson. 2004. Outside-in and inside-out signaling: The new concept that selectivity of ligand binding at the gonadotropin-releasing hormone receptor is modulated by the intracellular environment. Endocrinology. 145(8): 3590-3593.

Minucci, S., L. Di Matteo, R. Pierantoni, B. Varriale, R. K. Rastogi, and G. Chieffi. 1986. In vivo and in vitro stimulatory effect of a gonadotropin-releasing hormone analog (HOE 766) on spermatogonial multiplication in the frog, rana esculenta. Endocrinology. 119(2): 731-736.

Miyamoto, K., Y. Hasegawa, M. Nomura, M. Igarashi, K. Kangawa, and H. Matsuo. 1984. Identification of the second gonadotropin-releasing hormone in chicken hypothalamus: Evidence that gonadotropin secretion is probably controlled by two distinct gonadotropin-releasing hormones in avian species. Proc. Natl. Acad. Sci. U. S. A. 81(12): 3874-3878. 157

Molcho, J., Y. Eli, H. Zakut, and Z. Naor. 1984. Stimulation of prostaglandin E and testosterone production in rat interstitial cells by a gonadotropin-releasing hormone agonist. Endocrinology. 114(6): 2382-2387.

Mongiat, L. A., M. O. Fernandez, V. A. Lux-Lantos, L. G. Guilgur, G. M. Somoza, and C. Libertun. 2006. Experimental data supporting the expression of the highly conserved GnRH-II in the brain and pituitary gland of rats. Regul. Pept. 136(1-3): 50-57.

Morales, P. 1998. Gonadotropin-releasing hormone increases ability of the spermatozoa to bind to the human zona pellucida. Biol. Reprod. 59(2): 426-430.

Morales, P., B. Kerr, C. Oliva, E. Pizarro, and M. Kong. 1999. Gonadotrophin-releasing hormone antagonists inhibit sperm binding to the human zona pellucida. Hum. Reprod. 14(8): 2069-2074.

Moretti, R. M., M. M. Marelli, D. Dondi, A. Poletti, L. Martini, M. Motta et al. 1996. Luteinizing hormone-releasing hormone agonists interfere with the stimulatory actions of epidermal growth factor in human prostatic cancer cell lines, LNCaP and DU 145. J. Clin. Endocrinol. Metab. 81(11): 3930-3937.

Morgan, K., D. Conklin, A. J. Pawson, R. Sellar, T. R. Ott, and R. P. Millar. 2003. A transcriptionally active human type II gonadotropin-releasing hormone receptor gene homolog overlaps two genes in the antisense orientation on chromosome 1q.12. Endocrinology. 144(2): 423-436.

Morgan, K., R. Sellar, A. J. Pawson, Z. L. Lu, and R. P. Millar. 2006. Bovine and ovine gonadotropin-releasing hormone (GnRH)-II ligand precursors and type II GnRH receptor genes are functionally inactivated. Endocrinology. 147(11): 5041-5051.

Muske, L. E., J. A. King, F. L. Moore, and R. P. Millar. 1994. Gonadotropin-releasing hormones in microdissected brain regions of an amphibian: Concentration and anatomical distribution of immunoreactive mammalian GnRH and chicken GnRH II. Regul. Pept. 54(2-3): 373-384.

Namy, O., J. P. Rousset, S. Napthine, and I. Brierley. 2004. Reprogrammed genetic decoding in cellular gene expression. Mol. Cell. 13(2): 157-168.

Naor, Z., and I. Huhtaniemi. 2012. Interactions of the GnRH receptor with heterotrimeric G proteins. Front. Neuroendocrinol.

Navarro, D., O. P. Luzardo, L. Fernandez, N. Chesa, and B. N. Diaz-Chico. 2002. Transition to androgen-independence in prostate cancer. J. Steroid Biochem. Mol. Biol. 81(3): 191-201. 158

Neill, J., L. Duck, and L. Musgrove. 2002a. GnRH II receptor is encoded in genomes of human, monkey, and pig but not mouse. Program of the 84th annual meeting of the endocrine society, san francisco, CA,

Neill, J., L. Duck, and L. Musgrove. 2002b. Potential regulatory role for GnRH II in gonadotropin secretion: Molecular characterization of a GnRH II receptor in the pig pituitary. Proc soc neurosci, abstract,

Neill, J. D. 2002. GnRH and GnRH receptor genes in the human genome. Endocrinology. 143(3): 737-743.

Neill, J. D., L. C. Musgrove, and L. W. Duck. 2004. Newly recognized GnRH receptors: Function and relative role. Trends Endocrinol. Metab. 15(8): 383-392.

Neill, J. D., L. W. Duck, J. C. Sellers, and L. C. Musgrove. 2001. A gonadotropin- releasing hormone (GnRH) receptor specific for GnRH II in primates. Biochem. Biophys. Res. Commun. 282(4): 1012-1018.

Nett, T. M., A. M. Akbar, and G. D. Niswender. 1974. Serum levels of luteinizing hormone and gonadotropin-releasing hormone in cycling, castrated and anestrous ewes. Endocrinology. 94(3): 713-718.

Nikula, H., and I. Huhtaniemi. 1988. Gonadotropin-releasing hormone agonist activates protein kinase C in rat leydig cells. Mol. Cell. Endocrinol. 55(1): 53-59.

Oatley, J. M., D. M. de Avila, J. J. Reeves, and D. J. McLean. 2004. Testis tissue explant culture supports survival and proliferation of bovine spermatogonial stem cells. Biol. Reprod. 70(3): 625-631.

Odell, W. D., R. S. Swerdloff, J. Bain, F. Wollesen, and P. K. Grover. 1974. The effect of sexual maturation on testicular response to LH stimulation of testosterone secretion in the intact rat. Endocrinology. 95(5): 1380-1384.

Okada, Y., A. Murota-Kawano, S. S. Kakar, and S. J. Winters. 2003. Evidence that gonadotropin-releasing hormone (GnRH) II stimulates luteinizing hormone and follicle-stimulating hormone secretion from monkey pituitary cultures by activating the GnRH I receptor. Biol. Reprod. 69(4): 1356-1361.

Okwun, O. E., G. Igboeli, J. J. Ford, D. D. Lunstra, and L. Johnson. 1996a. Number and function of sertoli cells, number and yield of spermatogonia, and daily sperm production in three breeds of boar. J. Reprod. Fertil. 107(1): 137-149. 159

Okwun, O. E., G. Igboeli, D. D. Lunstra, J. J. Ford, and L. Johnson. 1996b. Testicular composition, number of A spermatogonia, germ cell ratios, and number of spermatids in three different breeds of boars. J. Androl. 17(3): 301-309.

Olivennes, F., R. Fanchin, P. Bouchard, J. Taieb, and R. Frydman. 1996. Triggering of ovulation by a gonadotropin-releasing hormone (GnRH) agonist in patients pretreated with a GnRH antagonist. Fertil. Steril. 66(1): 151-153.

Osman, D., and L. Plöen. 1978. The ultrastructure of sertoli cells in the boar.1: 162-179.

Parborell, F., G. Irusta, A. Rodriguez Celin, and M. Tesone. 2008. Regulation of ovarian angiogenesis and apoptosis by GnRH-I analogs. Mol. Reprod. Dev. 75(4): 623-631.

Parvinen, M., and S. Ventela. 1999. Local regulation of spermatogenesis: A living cell approach. Hum. Fertil. (Camb). 2(2): 138-142.

Pati, D., and H. R. Habibi. 1993. Gonadotropin-releasing hormone (GnRH) binding characteristics in the testis of goldfish (carassius auratus). J. Exp. Zool. 267(2): 155- 163.

Pawson, A. J., S. Maudsley, K. Morgan, L. Davidson, Z. Naor, and R. P. Millar. 2005. Inhibition of human type i gonadotropin-releasing hormone receptor (GnRHR) function by expression of a human type II GnRHR gene fragment. Endocrinology. 146(6): 2639-2649.

Pawson, A. J., S. R. Maudsley, J. Lopes, A. A. Katz, Y. M. Sun, J. S. Davidson et al. 2003a. Multiple determinants for rapid agonist-induced internalization of a nonmammalian gonadotropin-releasing hormone receptor: A putative palmitoylation site and threonine doublet within the carboxyl-terminal tail are critical. Endocrinology. 144(9): 3860-3871.

Pawson, A. J., K. Morgan, S. R. Maudsley, and R. P. Millar. 2003b. Type II gonadotrophin-releasing hormone (GnRH-II) in reproductive biology. Reproduction. 126(3): 271-278.

Pelliniemi, L. J. 1975. Ultrastructure of the early ovary and testis in pig embryos. Am. J. Anat. 144(1): 89-111.

Perron, A., P. Sarret, L. Gendron, T. Stroh, and A. Beaudet. 2005. Identification and functional characterization of a 5-transmembrane domain variant isoform of the NTS2 neurotensin receptor in rat central nervous system. J. Biol. Chem. 280(11): 10219-10227. 160

Petersson, F., G. Emons, and M. Hammar. 1989. Testicular GnRH-receptors and direct effects of a GnRH-agonist on human testicular steroidogenesis. Scand. J. Urol. Nephrol. 23(3): 161-164.

Pfleger, K. D., J. Bogerd, and R. P. Millar. 2002. Conformational constraint of mammalian, chicken, and salmon GnRHs, but not GnRH II, enhances binding at mammalian and nonmammalian receptors: Evidence for preconfiguration of GnRH II. Mol. Endocrinol. 16(9): 2155-2162.

Pierantoni, R., S. Fasano, L. Di Matteo, S. Minucci, B. Varriale, and G. Chieffi. 1984. Stimulatory effect of a GnRH agonist (buserelin) in in vitro and in vivo testosterone production by the frog (rana esculenta) testis. Mol. Cell. Endocrinol. 38(2-3): 215- 219.

Pimstone, B., S. Epstein, S. M. Hamilton, D. LeRoith, and S. Hendricks. 1977. Metabolic clearance and plasma half disappearance time of exogenous gonadotropin releasing hormone in normal subjects and in patients with liver disease and chronic renal failure. J. Clin. Endocrinol. Metab. 44(2): 356-360.

Pinski, J., T. Yano, G. Miller, and A. V. Schally. 1992. Blockade of the LH response induced by the agonist D-trp-6-LHRH in rats by a highly potent LH-RH antagonist SB-75. Prostate. 20(3): 213-224.

Powell, J. F., S. M. Reska-Skinner, M. O. Prakash, W. H. Fischer, M. Park, J. E. Rivier et al. 1996. Two new forms of gonadotropin-releasing hormone in a protochordate and the evolutionary implications. Proc. Natl. Acad. Sci. U. S. A. 93(19): 10461-10464.

Raeside, J. I., H. L. Christie, R. L. Renaud, and P. A. Sinclair. 2006. The boar testis: The most versatile steroid producing organ known. Soc. Reprod. Fertil. Suppl. 62: 85-97.

Raposo, G., I. Dunia, C. Delavier-Klutchko, S. Kaveri, A. D. Strosberg, and E. L. Benedetti. 1989. Internalization of beta-adrenergic receptor in A431 cells involves non-coated vesicles. Eur. J. Cell Biol. 50(2): 340-352.

Reinhart, J., L. M. Mertz, and K. J. Catt. 1992. Molecular cloning and expression of cDNA encoding the murine gonadotropin-releasing hormone receptor. J. Biol. Chem. 267(30): 21281-21284.

Rick, F. G., A. V. Schally, N. L. Block, G. Halmos, R. Perez, J. B. Fernandez et al. 2011. LHRH antagonist cetrorelix reduces prostate size and gene expression of proinflammatory cytokines and growth factors in a rat model of benign prostatic hyperplasia. Prostate. 71(7): 736-747. 161

Rissman, E. F., V. E. Alones, C. B. Craig-Veit, and J. R. Millam. 1995. Distribution of chicken-II gonadotropin-releasing hormone in mammalian brain. J. Comp. Neurol. 357(4): 524-531.

Robinson, D. N., and L. Cooley. 1997. Examination of the function of two kelch proteins generated by stop codon suppression. Development. 124(7): 1405-1417.

Ronacher, K., N. Matsiliza, N. Nkwanyana, A. J. Pawson, T. Adam, C. A. Flanagan et al. 2004. Serine residues 338 and 339 in the carboxyl-terminal tail of the type II gonadotropin-releasing hormone receptor are critical for beta-arrestin-independent internalization. Endocrinology. 145(10): 4480-4488.

Ron-El, R., A. Raziel, M. Schachter, D. Strassburger, E. Kasterstein, and S. Friedler. 2000. Induction of ovulation after gnRH antagonists. Hum. Reprod. Update. 6(4): 318-321.

Russell, L. D., and L. R. de Franca. 1995. Building a testis. Tissue Cell. 27(2): 129-147.

Saint Pol, P., E. Hermand, and G. Tramu. 1988. Paracrine factors in adult rat testis gonadotrophin control of opioids and LHRH like peptide. Andrologia. 20(2): 173- 181.

Sanchez, C., C. Escrieut, P. Clerc, V. Gigoux, B. Waser, J. C. Reubi et al. 2012. Characterization of a novel five-transmembrane domain cholecystokinin-2 receptor splice variant identified in human tumors. Mol. Cell. Endocrinol. 349(2): 170-179.

Sathananthan, A. H., S. S. Ratnam, S. C. Ng, J. J. Tarin, L. Gianaroli, and A. Trounson. 1996. The sperm centriole: Its inheritance, replication and perpetuation in early human embryos. Hum. Reprod. 11(2): 345-356.

Schally, A. V. 1973. Hypothalamic LH and FSH releasing hormone; physiological and clinical studies. Clin. Sci. 44(2): 1P passim.

Schally, A. V., A. Arimura, Y. Baba, R. M. Nair, H. Matsuo, T. W. Redding et al. 1971. Isolation and properties of the FSH and LH-releasing hormone. Biochem. Biophys. Res. Commun. 43(2): 393-399.

Schneider, J. S., and E. F. Rissman. 2008. Gonadotropin-releasing hormone II: A multi- purpose neuropeptide. Integr. Comp. Biol. 48(5): 588-595.

Schoneberg, T., J. Liu, and J. Wess. 1995. Plasma membrane localization and functional rescue of truncated forms of a G protein-coupled receptor. J. Biol. Chem. 270(30): 18000-18006. 162

Schulz, R. W., F. van der Wind, C. Janssen-Dommerholt, J. Peute, C. C. Mylonas, Y. Zohar et al. 1997. Modulation of testicular androgen production in adolescent african catfish (clarias gariepinus). Gen. Comp. Endocrinol. 108(1): 56-66.

Sealfon, S. C., H. Weinstein, and R. P. Millar. 1997. Molecular mechanisms of ligand interaction with the gonadotropin-releasing hormone receptor. Endocr. Rev. 18(2): 180-205.

Seeburg, P. H., A. J. Mason, T. A. Stewart, and K. Nikolics. 1987. The mammalian GnRH gene and its pivotal role in reproduction. Recent Prog. Horm. Res. 43: 69-98.

Senger, P. L. 2012. Pathways to pregnancy and parturition. Currecnt Concepts, Inc.

Seong, J., and J. Cheon. 2007. Use of the gonadotropin-releasing hormone II antagonist and its analogues
.(2,007,052,854)

Sharpe, R. M., and I. Cooper. 1987. Comparison of the effects on purified leydig cells of four hormones (oxytocin, vasopressin, opiates and LHRH) with suggested paracrine roles in the testis. J. Endocrinol. 113(1): 89-96.

Sharpe, R. M., and I. Cooper. 1982a. Stimulatory effect of LHRH and its agonists on leydig cell steroidogenesis in vitro. Mol. Cell. Endocrinol. 26(1-2): 141-150.

Sharpe, R. M., and I. Cooper. 1982b. Variation in the steroidogenic responsiveness of isolated rat leydig cells. J. Reprod. Fertil. 65(2): 475-481.

Sharpe, R. M., D. G. Doogan, and I. Cooper. 1983. Direct effects of a luteinizing hormone-releasing hormone agonist on intratesticular levels of testosterone and interstitial fluid formation in intact male rats. Endocrinology. 113(4): 1306-1313.

Sharpe, R. M., and H. M. Fraser. 1980. HCG stimulation of testicular LHRH-like activity. Nature. 287(5783): 642-643.

Sharpe, R. M., H. M. Fraser, I. Cooper, and F. F. Rommerts. 1982. The secretion, measurement, and function of a testicular LHRH-like factor. Ann. N. Y. Acad. Sci. 383: 272-294.

Shimizu, M., and G. Y. Bedecarrats. 2006. Identification of a novel pituitary-specific chicken gonadotropin-releasing hormone receptor and its splice variants. Biol. Reprod. 75(5): 800-808.

Shuttlesworth, G. A., D. G. de Rooij, I. Huhtaniemi, T. Reissmann, L. D. Russell, G. Shetty et al. 2000. Enhancement of A spermatogonial proliferation and 163

differentiation in irradiated rats by gonadotropin-releasing hormone antagonist administration. Endocrinology. 141(1): 37-49.

Siler-Khodr, T. M., and M. Grayson. 2001. Action of chicken II GnRH on the human placenta. J. Clin. Endocrinol. Metab. 86(2): 804-810.

Siler-Khodr, T. M., M. Grayson, and C. A. Eddy. 2003. Action of gonadotropin-releasing hormone II on the baboon ovary. Biol. Reprod. 68(4): 1150-1156.

Sorkin, A., and M. Von Zastrow. 2002. Signal transduction and endocytosis: Close encounters of many kinds. Nat. Rev. Mol. Cell Biol. 3(8): 600-614.

Sower, S. A., Y. C. Chiang, S. Lovas, and J. M. Conlon. 1993. Primary structure and biological activity of a third gonadotropin-releasing hormone from lamprey brain. Endocrinology. 132(3): 1125-1131.

Steger, K. 1999. Transcriptional and translational regulation of gene expression in haploid spermatids. Anat. Embryol. (Berl). 199(6): 471-487.

Stewart, A. J., A. A. Katz, R. P. Millar, and K. Morgan. 2009. Retention and silencing of prepro-GnRH-II and type II GnRH receptor genes in mammals. Neuroendocrinology. 90(4): 416-432.

Stewart, D. W., and J. I. Raeside. 1976. Testosterone secretion by the early fetal pig testes in organ culture. Biol. Reprod. 15(1): 25-28.

Stojilkovic, S. S., and K. J. Catt. 1995. Expression and signal transduction pathways of gonadotropin-releasing hormone receptors. Recent Prog. Horm. Res. 50: 161-205.

Szollosi, D., and R. H. Hunter. 1973. Ultrastructural aspects of fertilization in the domestic pig: Sperm penetration and pronucleus formation. J. Anat. 116(Pt 2): 181- 206.

Temple, J. L., R. P. Millar, and E. F. Rissman. 2003a. An evolutionarily conserved form of gonadotropin-releasing hormone coordinates energy and reproductive behavior. Endocrinology. 144(1): 13-19.

Temple, J. L., R. P. Millar, and E. F. Rissman. 2003b. An evolutionarily conserved form of gonadotropin-releasing hormone coordinates energy and reproductive behavior. Endocrinology. 144(1): 13-19.

Tensen, C., K. Okuzawa, M. Blomenröhr, F. Rebersi, R. Leurs, J. Bogerd et al. 1997. Distinct efficacies for two endogenous ligands on a single cognate gonadoliberin receptor. European Journal of Biochemistry. 243(1-2): 134-140. 164

Topfer-Petersen, E., J. Calvete, W. Schafer, and A. Henschen. 1990. Complete localization of the disulfide bridges and glycosylation sites in boar sperm acrosin. FEBS Lett. 275(1-2): 139-142.

Troskie, B., N. Illing, E. Rumbak, Y. M. Sun, J. Hapgood, S. Sealfon et al. 1998. Identification of three putative GnRH receptor subtypes in vertebrates. Gen. Comp. Endocrinol. 112(3): 296-302.

Troskie, B., J. A. King, R. P. Millar, Y. Y. Peng, J. Kim, H. Figueras et al. 1997. Chicken GnRH II-like peptides and a GnRH receptor selective for chicken GnRH II in amphibian sympathetic ganglia. Neuroendocrinology. 65(6): 396-402.

Tsai, P. S., and L. Zhang. 2008. The emergence and loss of gonadotropin-releasing hormone in protostomes: Orthology, phylogeny, structure, and function. Biol. Reprod. 79(5): 798-805.

Tsutsumi, M., W. Zhou, R. P. Millar, P. L. Mellon, J. L. Roberts, C. A. Flanagan et al. 1992. Cloning and functional expression of a mouse gonadotropin-releasing hormone receptor. Mol. Endocrinol. 6(7): 1163-1169.

Udagawa, K., M. Takeda, M. Hosaka, Y. Kubota, and T. Ogawa. 2002. Recovery of spermatogenesis by high dose gonadotropin-releasing hormone analogue treatment in rat cryptorchid testis after orchiopexy. J. Urol. 168(3): 1279-1283.

Urbanski, H. F., R. B. White, R. D. Fernald, S. G. Kohama, V. T. Garyfallou, and V. S. Densmore. 1999. Regional expression of mRNA encoding a second form of gonadotropin-releasing hormone in the macaque brain. Endocrinology. 140(4): 1945-1948.

USDA-NASS. 2010. NASS. National Agricultural Statistic Service. van Biljon, W., S. Wykes, S. Scherer, S. A. Krawetz, and J. Hapgood. 2002. Type II gonadotropin-releasing hormone receptor transcripts in human sperm. Biol. Reprod. 67(6): 1741-1749.

Verhoeven, G., and J. Cailleau. 1985. A factor in spent media from sertoli-cell-enriched cultures that stimulates steroidogenesis in leydig cells. Mol. Cell. Endocrinol. 40(1): 57-68.

Wang, A. F., J. H. Li, K. Maiti, W. P. Kim, H. M. Kang, J. Y. Seong et al. 2003. Preferential ligand selectivity of the monkey type-II gonadotropin-releasing hormone (GnRH) receptor for GnRH-2 and its analogs. Mol. Cell. Endocrinol. 209(1-2): 33- 42. 165

Wang, Y., H. Matsuo, O. Kurachi, and T. Maruo. 2002. Down-regulation of proliferation and up-regulation of apoptosis by gonadotropin-releasing hormone agonist in cultured uterine leiomyoma cells. Eur. J. Endocrinol. 146(3): 447-456.

Weinbauer, G. F., and E. Nieschlag. 1993. Evaluation of the antigonadotropic activity of different GnRH antagonists in the non-human primate. Andrologia. 25: 141-147.

Weinbauer, G. F., A. Limberger, H. M. Behre, and E. Nieschlag. 1994. Can testosterone alone maintain the gonadotrophin-releasing hormone antagonist-induced suppression of spermatogenesis in the non-human primate? J. Endocrinol. 142(3): 485-495.

White, R. B., J. A. Eisen, T. L. Kasten, and R. D. Fernald. 1998. Second gene for gonadotropin-releasing hormone in humans. Proc. Natl. Acad. Sci. U. S. A. 95(1): 305-309.

Whitelaw, P. F., K. A. Eidne, R. Sellar, C. D. Smyth, and S. G. Hillier. 1995. Gonadotropin-releasing hormone receptor messenger ribonucleic acid expression in rat ovary. Endocrinology. 136(1): 172-179.

Wise, T., D. D. Lunstra, and J. J. Ford. 1996. Differential pituitary and gonadal function of chinese meishan and european white composite boars: Effects of gonadotropin- releasing hormone stimulation, castration, and steroidal feedback. Biol. Reprod. 54(1): 146-153.

Wise, T., E. L. Zanella, D. D. Lunstra, and J. J. Ford. 2000. Relationships of gonadotropins, testosterone, and cortisol in response to GnRH and GnRH antagonist in boars selected for high and low follicle-stimulating hormone levels. J. Anim. Sci. 78(6): 1577-1590.

Wu, H. M., H. S. Wang, H. Y. Huang, Y. K. Soong, C. D. MacCalman, and P. C. Leung. 2009. GnRH signaling in intrauterine tissues. Reproduction. 137(5): 769-777.

Xu, X., S. Pommier, T. Arbov, B. Hutchings, W. Sotto, and G. R. Foxcroft. 1998. In vitro maturation and fertilization techniques for assessment of semen quality and boar fertility. J. Anim. Sci. 76(12): 3079-3089.

Yano, T., N. Yano, H. Matsumi, Y. Morita, O. Tsutsumi, A. V. Schally et al. 1997. Effect of luteinizing hormone-releasing hormone analogs on the rat ovarian follicle development. Horm. Res. 48 Suppl 3: 35-41.

Zanella, E. L., D. D. Lunstra, T. H. Wise, J. E. Kinder, and J. J. Ford. 2000. GnRH antagonist inhibition of gonadotropin and steroid secretion in boars in vivo and steroid production in vitro. J. Anim. Sci. 78(6): 1591-1597. 166

Zerani, M., G. Catone, L. Quassinti, E. Maccari, M. Bramucci, A. Gobbetti et al. 2011. In vitro effects of gonadotropin-releasing hormone (GnRH) on leydig cells of adult alpaca (lama pacos) testis: GnRH receptor immunolocalization, testosterone and prostaglandin synthesis, and cyclooxygenase activities. Domest. Anim. Endocrinol. 40(1): 51-59.

Zwart, A. D., R. J. Urban, W. D. Odell, and J. D. Veldhuis. 1996. Contrasts in the gonadotropin-releasing hormone dose-response relationships for luteinizing hormone, follicle-stimulating hormone and alpha-subunit release in young versus older men: Appraisal with high-specificity immunoradiometric assay and deconvolution analysis. Eur J Endocrinol. 135(4): 399-406.