Anti-Müllerian Hormone in Stallions and Mares: Physiological Variations, Clinical Applications, and Molecular Aspects

University of Kentucky UKnowledge

Theses and Dissertations--Veterinary Science Veterinary Science

2014

ANTI-MÜLLERIAN HORMONE IN STALLIONS AND MARES: PHYSIOLOGICAL VARIATIONS, CLINICAL APPLICATIONS, AND MOLECULAR ASPECTS

Anthony N.J. Claes University of Kentucky, [email protected]

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Recommended Citation Claes, Anthony N.J., "ANTI-MÜLLERIAN HORMONE IN STALLIONS AND MARES: PHYSIOLOGICAL VARIATIONS, CLINICAL APPLICATIONS, AND MOLECULAR ASPECTS" (2014). Theses and Dissertations-- Veterinary Science. 18. https://uknowledge.uky.edu/gluck_etds/18

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REVIEW, APPROVAL AND ACCEPTANCE

The document mentioned above has been reviewed and accepted by the student’s advisor, on behalf of the advisory committee, and by the Director of Graduate Studies (DGS), on behalf of the program; we verify that this is the final, approved version of the student’s thesis including all changes required by the advisory committee. The undersigned agree to abide by the statements above.

Anthony N.J. Claes, Student

Dr. Barry A. Ball, Major Professor

Dr. Daniel K. Howe, Director of Graduate Studies

ANTI-MÜLLERIAN HORMONE IN STALLIONS AND MARES: PHYSIOLOGICAL VARIATIONS, CLINICAL APPLICATIONS, AND MOLECULAR ASPECTS

DISSERTATION

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the College of Agriculture, Food and Environment at the University of Kentucky

By Anthony N.J. Claes

Lexington, Kentucky

Director: Dr. Barry A. Ball, Professor of Equine Reproduction

Lexington, Kentucky

2014

ABSTRACT OF DISSERTATION

ANTI-MÜLLERIAN HORMONE IN STALLIONS AND MARES: PHYSIOLOGICAL VARIATIONS, CLINICAL APPLICATIONS, AND MOLECULAR ASPECTS

Anti-Müllerian hormone (AMH) is a homodimeric glycoprotein that is best known for its role in regression of the Müllerian duct in the male fetus. Accumulating evidence indicates that AMH also has an important role during different physiological processes after birth. In contrast to other species, relatively little is known about AMH in the horse. In chapter one, developmental and seasonal changes in serum AMH concentrations in male horses were determined, and the use of AMH for determination of retained cryptorchid testes was established. In chapter two, the interrelationship between plasma AMH concentrations, antral follicle counts (AFC), and age in mares was evaluated. Molecular and hormonal changes in the equine follicle with regard to variations in antral follicle count and follicular development were examined in chapter three. In chapter four, the effect of AFC on age-related changes in follicular parameters in light-type horse mares was examined. Peripheral AMH concentrations were significantly higher in prepubertal colts than in postpubertal stallions and varied with season in mature stallions with higher concentrations during the physiological breeding season. Furthermore, serum AMH concentrations were significantly higher in cryptorchid stallions compared to intact stallions or geldings. Circulating AMH concentrations varied widely amongst mares of the same age while the repeatability of AMH was high within and between estrous cycles. More importantly, AMH concentrations were positively associated with AFC, and this relationship increased with mare age. In addition, variations in AMH concentrations or AFC were associated with molecular differences in granulosa cells of growing follicles, and the expression of AMH and genes co-expressed with AMH in the equine follicle as well as intrafollicular AMH concentrations decreased during follicular development. Finally, the inter-ovulatory interval and length of the follicular phase is increased in aged mares with low AFC. In conclusion, AMH is a useful biomarker for cryptorchidism in stallions and ovarian reserve in mares. Furthermore, follicular function was interrelated to AFC or AMH based upon molecular differences in growing follicles, while age-related changes in follicular parameters are linked to differences in AFC.

KEYWORDS: anti-Müllerian hormone, stallion, mare, physiology, endocrinology

Anthony N.J. Claes Student’s Signature

August 4, 2014 Date

ANTI-MÜLLERIAN HORMONE IN STALLIONS AND MARES: PHYSIOLOGICAL VARIATIONS, CLINICAL APPLICATIONS, AND MOLECULAR ASPECTS

Anthony N.J. Claes

Dr. Barry A. Ball Director of Dissertation

Dr. Daniel K. Howe Director of Graduate Studies

August 4, 2014

ACKNOWLEDGEMENTS

First and foremost, I would like to thank Dr. Barry Ball who has been a great mentor for me over the past eight years. Most of my clinical and scientific knowledge that I obtained throughout those years is linked one way or another to you. I am grateful for your patience, endless support and excellent scientific guidance. I appreciate all the constructive criticism, suggestions and corrections that you made in order to improve the quality of my manuscripts or presentations. Dr. Ball always pushed me to take everything a step further and taught me to think like a scientist. It is an honor that I have learned from one the best and I will always appreciate everything that you have done for me.

Now, the time has come to stand on my own feet. Nevertheless, I will never forget what you have done for me. Without you, I would never have accomplished what I have done so far. Dr. Ball, thank you so much.

I would also like to thank Drs. Troedsson, Squires and Curry for being members of my committee. It was a privilege to have all of you in my committee. All your comments, ideas and suggestions on a variety of different topics were well appreciated. Furthermore,

I would like to thank my fellow students Alejandro Esteller-Vico, Elizabeth Woodward,

Gabriel Davolli, Carleigh Fedorka, Sydney Hughes, Julian Kalmar, and Jessalyn Walters for all their support. They were always happy to help with the animal experiments even if it was early in the morning. Another person that I would like to thank is Dr. Kirsten

Scoggin. She taught me all the different laboratory techniques which will be important for my future career. Her advice and experience with molecular techniques during the past three years was invaluable to me. I would also like to thank the entire farm crew, especially farm manager Kevin Gallagher. He was always very supportive and tried to

iii

accommodate me in any way possible during the clinical part of my experiments. Also a special thank you to Heidi Ball; she always embraced Charlotte and I as her own family.

We will miss the good conversations during all the parties at your house. Thank you as well Diane Furry for your help getting this dissertation into the right format. Last but not least I would like to thank my lovely wife, Charlotte. She is the most positive person that

I know in my entire life and supports me in all my endeavors. Without her, it would not have been that easy.

TABLE OF CONTENTS

ACKNOWLEDGEMENTS ...... iii LIST OF TABLES ...... viii LIST OF FIGURES ...... ix Chapter One: Literature Review ...... 1 1.1 Anti-Müllerian hormone ...... 1 1.1.1 The discovery of AMH ...... 1 1.1.2 Anti-Müllerian hormone: the gene...... 2 1.1.3 Anti-Müllerian hormone: the protein ...... 3 1.2 The immunoexpression of AMH in the equine testicular and ovarian tissue ...... 4 1.3 Changes in circulating concentrations of AMH in males and females ...... 7 1.4 The regulation of AMH ...... 9 1.4.1 The regulation of AMH in males ...... 9 1.4.2 The regulation of AMH in females ...... 10 1.5 Biological activity of AMH ...... 11 1.6 Pre- and postnatal biological functions of AMH in males and females ...... 12 1.6.1 Sexual differentiation: regression of the Müllerian ducts ...... 12 1.6.2 Folliculogenesis ...... 13 1.6.3 Inhibition of Leydig cell differentiation ...... 14 1.7 Molecular aspects of AMH signaling ...... 14 Chapter Two: Serum anti-Müllerian hormone concentrations in stallions: Developmental changes, seasonal variation, and differences between intact stallions, cryptorchid stallions and geldings ...... 16 2.1 Introduction ...... 16 2.2 Materials and methods ...... 18 2.2.1 Validation of anti-Müllerian hormone and testosterone assay ...... 18 2.2.2 Serum anti-Müllerian hormone concentrations in neonatal colts and fillies, prepubertal colts and postpubertal stallions, and serum testosterone concentrations in prepubertal colts and postpubertal stallions ...... 19 2.2.3 Seasonal variations in AMH and testosterone concentrations of intact mature stallions ...... 19 2.2.4 Serum anti-Müllerian hormone and testosterone concentrations in geldings, cryptorchid stallions, and intact stallions ...... 20

2.2.5 Detection of anti-Müllerian hormone in seminal plasma of a stallion and determination of seminal plasma AMH concentrations in intact stallions ...... 21 2.3 Statistical analysis ...... 21 2.4 Results ...... 22 2.4.1 Validation of the anti-Müllerian hormone assay for stallions ...... 22 2.4.2 Serum anti-Müllerian hormone concentrations in neonatal colts and fillies, prepubertal colts and postpubertal stallions, and serum testosterone concentrations in prepubertal colts and postpubertal stallions ...... 24 2.4.3 Seasonal concentrations of anti-Müllerian hormone and testosterone in intact stallions ...... 25 2.4.4 Serum anti-Müllerian hormone and testosterone concentrations in geldings, cryptorchid stallions and intact stallions ...... 27 2.4.5 Detection of anti-Müllerian hormone in seminal plasma of a stallion and determination of seminal plasma AMH concentrations in intact stallions ...... 29 2.5 Discussion and Conclusion ...... 31 2.6 Acknowledgments...... 35 Chapter Three: The interrelationship between anti-Müllerian hormone, ovarian follicular populations, and age in mares ...... 36 3.1 Introduction ...... 36 3.2 Materials and Methods ...... 38 3.2.1 Experimental design ...... 38 3.2.2 Determination of antral follicle counts ...... 38 3.2.3 Measurement of plasma anti-Müllerian hormone concentrations ...... 39 3.3 Statistical analysis ...... 41 3.4 Results ...... 42 3.4.1 Antral follicle count and plasma AMH concentrations in mares of different age groups, and the repeatability within and across estrous cycles ...... 42 3.4.2 Relationship between AFC and plasma AMH concentrations with regard to age and follicle size ...... 45 3.5 Discussion and Conclusion ...... 47 3.6 Acknowledgements ...... 50 Chapter Four: Molecular and hormonal differences in the equine follicle in relation to variations in antral follicle count and stage of follicular development ...... 51 4.1 Introduction ...... 51 4.2 Materials and Methods ...... 53 4.2.1 Animal tissue collection ...... 53 4.2.2 Histology ...... 54

4.2.3 RNA extraction and Real-time Reverse Transcriptase Polymerase Chain Reaction (qRT-PCR) ...... 55 4.2.4 Immunohistochemistry ...... 61 4.2.5 Measurement of estradiol and AMH concentrations in follicular fluid of growing and dominant follicles ...... 62 4.3 Statistical analysis ...... 62 4.4 Results ...... 64 4.4.1 Molecular and hormonal differences in growing and dominant equine follicles in relation to variations in AFC and plasma AMH concentrations...... 64 4.4.2 Identification of genes co-expressed with AMH in granulosa cells of growing and dominant follicles ...... 66 4.4.3 Molecular and hormonal differences in the equine follicle in relation to the stage of follicular development...... 68 4.5 Discussion and conclusion ...... 72 4.6 Acknowledgements ...... 77 Chapter Five: The influence of age, antral follicle count and diestrous ovulations on estrous cycle characteristics of mares ...... 78 5.1 Introduction ...... 78 5.2 Materials and Methods ...... 80 5.2.1 Animal procedures ...... 80 5.2.2 Follicular parameters and dynamics ...... 81 5.2.3 Determination of plasma progesterone and FSH concentrations ...... 82 5.2.4 Determination of primordial follicle counts ...... 83 5.3 Statistical analysis ...... 83 5.4 Results ...... 85 5.4.1 Follicular parameters and dynamics ...... 85 5.4.2 Plasma progesterone and FSH concentrations ...... 89 5.4.3 Primordial follicle counts ...... 92 5.5 Discussion and conclusion ...... 94 5.6 Acknowledgments...... 99 Chapter Six: General conclusions ...... 100 References ...... 105 Vita ...... 117

vii

LIST OF TABLES

Table 4.1 Gene, tissue type: granulosa versus theca cells, NCBI reference sequence of the transcript, primer sequence of the forward and reverse primer, and PCR primer efficiency...... 58

Table 4.2 Pearson’s correlations between the expression of AMH mRNA and the expression of genes involved in follicular development (AMHR2, ESR1, ESR2,

FSHR, LHCGR, IGF, INHA, INHBA) and steroidogenic genes (CYP19A1) in granulosa cells of growing and dominant follicles...... 67

Table 4.3 Anti-Müllerian hormone and estradiol concentrations in follicular fluid of growing and dominant follicles...... 71

Table 5.1 Follicular parameters and estrous cycle characteristics in young, middle-aged and aged mares...... 87

Table 5.2 Follicular parameters of mares with a one-wave estrous cycle

(no diestrous ovulation) and a two-wave estrous cycle with a secondary ovulatory follicular wave (diestrous ovulation)...... 89

viii

LIST OF FIGURES

Figure 1.1 The immunoexpression of AMH in testicular tissue of equine fetuses, prepubertal colts and post-pubertal stallions ...... 5

Figure 1.2 The immunoexpression of AMH in equine primordial, preantral, and antral follicles...... 7

Figure 2.1 Parallelism plot for the anti-Müllerian hormone assay ...... 23

Figure 2.2 Biological validation of the anti-Müllerian hormone assay ...... 24

Figure 2.3 Serum testosterone and AMH concentrations in prepubertal colts and postpubertal stallions ...... 25

Figure 2.4 Seasonal variation in AMH and testosterone concentrations in intact stallions ...... 26

Figure 2.5 Mean serum AMH concentration in cryptorchid stallions, intact stallions, and geldings ...... 28

Figure 2.6 Mean serum AMH concentration in cryptorchid stallions without a scrotal testis, cryptorchid stallions with a scrotal testis, and intact stallions ...... 29

Figure 2.7 Western blot analysis of anti-Müllerian hormone in seminal plasma from a stallion under non-reducing and reducing conditions ...... 30

Figure 3.1 Parallelism between dilution curves and the standard curve of AMH ...... 40

Figure 3.2 Box plots showing antral follicle count in young, middle-aged and old mares ...... 43

Figure 3.3 Box plots of plasma AMH concentrations in young, middle-aged and old mares ...... 44

Figure 3.4 Correlation between AFC and plasma AMH concentration independent of mare age ...... 46

Figure 3.5 Correlations between AFC and plasma AMH concentration within different age groups ...... 47

Figure 4.1 Pearson’s correlations between AFC and the expression of genes involved in follicular development (AMH, AMHR2, ESR1, ESR2, FSHR, LHCGR, IGF1,

INHBA, INHA) and steroidogenesis (CYP19A1) in granulosa cells of growing follicles ...... 65

Figure 4.2 Pearson’s correlations between plasma AMH concentrations and the expression of genes involved in follicular development (AMH, AMHR2, ESR1,

ESR2, FSHR, LHCGR, IGF1, INHBA, INHA) and steroidogenesis (CYP19A1) in granulosa cells of growing follicles ...... 66

Figure 4.3 The expression of selected genes in granulosa cells of growing follicles prior to follicle deviation and dominant follicles after follicle deviation ...... 69

Figure 4.4 The immunoexpression of AMH and P450arom in granulosa cells of growing and dominant follicles ...... 70

Figure 5.1 Plasma progesterone concentrations across two consecutive estrous cycles in young, middle-aged, and aged mares ...... 90

Figure 5.2 Mean plasma FSH concentrations during one estrous cycle in a subset of young, middle-aged, and aged mares ...... 91

Figure 5.3 Mean plasma progesterone concentrations in mares with and without a diestrous ovulation ...... 92

Figure 5.4 A histological image of a primordial follicle in equine ovarian tissue ...... 93

Chapter One: Literature Review

Anti-Müllerian hormone (AMH) belongs to one of the most important reproductive hormones because of its role in sexual differentiation during fetal development.

Accumulating evidence indicates that AMH also has several important physiological functions after birth. This review will cover some general aspects of AMH including its discovery, gene and protein structure, expression pattern, secretion, regulation as well as the biological activity and function of AMH throughout the reproductive life of males and females. The primary goal of this review was to focus on AMH in the horse; nevertheless, some of the described information is derived from other species including but not limited to cattle, humans, and rodents.

1.1 Anti-Müllerian hormone

1.1.1 The discovery of AMH

The discovery of anti-Müllerian hormone dates back to the middle of last century when

Alfred Jost, a French physiologist, was interested in the process of sexual differentiation.

Initially, he showed that when fetal gonads were removed in utero, the Müllerian ducts developed while the Wolffian ducts regressed [1]. Consequently, it was assumed that testosterone played a key role in sexual differentiation, but this was rejected because administration of androgens to female fetuses resulted in differentiation of the Wolffian ducts while the Müllerian ducts failed to regress. However, when small pieces of testicular tissue were grafted in close proximity to the ovary, regression of Müllerian

ducts was observed indicating that the fetal testis must have an important role in sexual differentiation. Therefore, Alfred Jost concluded that the fetal testis does not only produces androgens but also secretes another substance which induces the regression of the Müllerian ducts [2]. Although the nature of this substance remained unidentified, he named it ‘hormone inhibitrice’ which later became better known as Müllerian inhibiting substance, Müllerian inhibiting factor or anti-Müllerian hormone [3].

1.1.2 Anti-Müllerian hormone: the gene

Cate et al. cloned the bovine and human AMH gene in 1986. The human AMH gene is

2.75 kbp in size, contains five exons, and has a guanine-cytosine content of approximately 70% [4]. The biological activity of the protein is encoded by the fifth and last exon of the gene [5]. The location of the AMH gene is different between species. The

AMH gene is located on chromosome 19 in humans [6], chromosome 10 in mice [7], and chromosome 7 in cattle [8] and in horses (NCBI: gene ID 102148318). Mutations in the

AMH gene can result in persistent Müllerian duct syndrome (PMDS), an autosomal recessive condition in males in which the Müllerian ducts fail to regress [9]. However, not all patients with PMDS have a mutation in the AMH gene; this condition can also arise from a mutation in the AMH receptor gene. Differentiating a mutation in the gene or gene receptor can be accomplished by measuring circulating AMH concentrations.

Patients with mutations in receptor gene are characterized by normal circulating AMH concentrations while patients with mutations in the AMH gene usually have undetectable

AMH concentrations. Although the majority of the patients with PMDS have mutations

in either the AMH gene or receptor, a small percentage of patients (15%) have another underlying cause that has not yet been identified [10].

1.1.3 Anti-Müllerian hormone: the protein

Although the existence of a substance ‘hormone inhibitrice’ was already implied by

Alfred Jost in 1953, specific information about this substance remained unknown for quite a while. In one of the first studies on AMH, it was indicated that this substance was a large molecule, such as polypeptide, as it was not able pass through a membrane which allowed the passage of androgens [11]. More in depth research showed that this substance was actually a glycoprotein [12, 13] that contained disulfide bridges as smaller fragments were obtained when preparations of fetal testes were examined using SDS-page under reducing conditions [13]. As the methods of AMH purification improved, more detailed information was obtained about the physical properties of this glycoprotein [14] as well as its biochemical composition with respect to amino acids and carbohydrates [4, 15].

The protein AMH is derived from a precursor which varies slightly in length among different species (human, bovine, rat, and mice) ranging from 553 (rat) to 575 (bovine) amino acids [3]. The equine precursor is similar to the length of the bovine precursor with 573 amino acids (unpublished data Drs. Ball and Conley; NCBI: GenBank

AEA11205.1). The amino acid sequence of the carboxyl terminal portion of the precursor is not only highly conserved across different species but also amongst other members of the transforming growth factor  family. This is in contrast to the amino acid sequence in the amino-terminal region which is less conserved between different species [3].

Furthermore, the bovine AMH precursor starts with a signal sequence (16-17 amino acids) followed by a pro-sequence (7-8 amino acids). However, both sequences are cleaved from the precursor before secretion [4]. Finally, dimerization of the remaining monomer (70 kDa) results in the formation of the glycoprotein AMH which is approximately 140 kDa [5].

An important characteristic of this 140 kDa AMH glycoprotein is that cleavage is not required in order to be biologically active. Nevertheless, in vitro cleavage of AMH using plasmin also results in a biologically active product which contains a 25 kDa and 110 kDa fragment representing the carboxyl and amino terminal dimer, respectively.

Furthermore, sequencing of those fragments revealed that cleavage occurs at the monobasic cleavage site between the amino acid arginine and serine; 109 amino acids upstream of the carboxyl-terminus [16]. Interestingly, the biological activity of AMH is localized in carboxyl terminal fragment whereas the amino terminal fragment is biologically inactive [17]. Nevertheless, even though this amino terminal fragment is biologically inactive by itself, it supports the activity of the carboxyl terminal fragment as more complete regression of Müllerian ducts is observed when both fragments are included in organ culture media compared to only the carboxyl terminal fragment [18].

1.2 The immunoexpression of AMH in the equine testicular and ovarian tissue

The immunoexpression of AMH in testicular tissue of stallions with regards to testicular development is well characterized using immunohistochemistry [19]. Similar to other species [20], the expression of AMH is localized to the cytoplasm of the Sertoli cells. In

addition, the expression of AMH is strongly influenced by testicular development as indicated by the prominent differences in AMH immunolabelling in fetal, pre-pubertal, post-pubertal equine testicular tissue. As expected, AMH is strongly expressed in the seminiferous tubules of the fetal testis. Furthermore, AMH is also expressed in Sertoli cells of prepubertal colts but less than the fetal stage. Finally, the immunoexpression of

AMH disappears during pubertal development as no AMH immunoexpression is present in testicular tissue of stallions older than 3 years of age [19] (Figure 1.1).

Figure 1.1 The immunoexpression of AMH in testicular tissue of equine fetuses, prepubertal colts and postpubertal stallions (from left to right). Strong AMH immunolabelling (arrow) is present within the seminiferous tubules of equine fetal testicular tissue. AMH expression can also be observed within the presumptive Sertoli cells (arrowhead) of prepubertal testicular tissue but with reduced intensity. Finally, no

AMH immunoexpression can be detected within the Sertoli cells of seminiferous tubules of postpubertal stallions (modified from Ball et al.[19])

The immunohistochemical expression of AMH has also been examined in the equine ovary with regard to follicular type and size. The expression of AMH in the equine ovary is confined to the cytoplasm of the granulosa cells and appears to be largely influenced by follicular development. No AMH labelling can be detected in the flattened granulosa cell layer of primordial follicles. The first sign of AMH expression is within the cuboidal granulosa cells of the primary follicle and the expression of AMH increases as the number of granulosa cells layers increased (Figure 1.2) [21]. A recent study examined the expression of AMH in growing follicles (15-20 mm) prior to follicle selection and in dominant follicles. Strong AMH immunostaining was detected in the small growing follicles while only faint AMH labelling was present in dominant follicles. This suggests that AMH is downregulated in the immediate time period around follicle selection.

Another factor that appears to have a large influence on the expression of AMH in equine ovarian follicles is follicular viability; only weak immunoexpression of AMH is present in granulosa cells of atretic follicles. Lastly, the equine corpus luteum does not exhibit any AMH immunostaining [21].

Figure 1.2 The immunoexpression of AMH in equine primordial, preantral, and antral follicles (from left to right). No AMH expression is present within primordial follicles

(left image) while mild expression of AMH (arrow) can be detected within the granulosa cells of a primary follicle (middle image). The expression of AMH increases with follicular development; granulosa cells of antral follicles (right image) appear to display stronger AMH labelling (arrowhead) (modified from Ball et al. [21]).

1.3 Changes in circulating concentrations of AMH in males and females

Before describing the regulation of AMH in testicular and ovarian tissue, the reader should have some understanding about the secretion pattern of AMH throughout the reproductive life of males and females. The synthesis of AMH in males is initiated early in fetal life after differentiation of the gonad. The production of AMH by the fetal Sertoli cells is of crucial importance in the process of sexual differentiation as it induces regression of the Müllerian ducts [22]. Concentrations of AMH in cord blood of male human fetuses appear to be high during mid-gestation but declines during the latter part of pregnancy [23, 24]. After birth, serum AMH concentrations increases and reaches a peak within the first year of life [25, 26]. Although a slight decrease in AMH concentrations is observed after one year of age, concentrations of AMH remain high

until puberty [25, 26]. The pubertal period is characterized by a rapid decrease in serum

AMH concentrations during the first stage of puberty [27]. As a consequence, only very low AMH concentrations are present in adult males [25]. Nonetheless, there is still some variation in AMH concentrations among adult men which is largely determined by genetic factors [28].

The secretion pattern of AMH is distinctively different in women than in men. Although

AMH expression can be detected in granulosa cells of the fetal ovary during late gestation [29], no AMH is present in the fetal circulation [24, 30]. Furthermore, AMH remains undetectable in cord blood in approximately 50 to 80% of the female newborns

[30, 31]. Based on a combination of different studies [30-32], it appears that circulating

AMH concentrations temporarily increases immediately after birth up to three months of age [31]. This temporary increase in AMH concentration appears to coincide with a minipuberty period [33]. Another increase in AMH concentration is detected at 4 years of age [31] and around puberty [32]. After puberty, AMH concentrations remain relatively stable until the age of 25 [31, 32] after which time AMH starts to decline. A more rapid decline in AMH concentrations is observed once women reached their 40’s [32]. Finally,

AMH becomes undetectable in the majority of women at the age of 50 or older [31]. It is well documented that the onset of menopause in women can be reasonably predicted using circulating concentrations of AMH [34, 35]. Interestingly, it appears that the factor age only explains 34% of the variation in circulating human AMH concentrations [32].

1.4 The regulation of AMH

1.4.1 The regulation of AMH in males

It is absolutely mandatory that the testicular expression of AMH is tightly regulated during early fetal life to ensure normal sexual differentiation. The promoter region of the

AMH gene contains binding sites for several different transcription factors including SF1

(Steroidogenic factor-1) [36], SOX9 [37], and GATA [38] which regulate the expression of AMH in the fetal testis. Binding of those transcription factors to the promoter region initiates the transcription of AMH. Furthermore, synergistic actions have been detected between the different transcription factors [37]. In addition to those transcription factors,

WT-1 (Wilms’ Tumor 1) and DAX-1 are also involved in the regulation of AMH even though they do not bind to the promoter region of AMH. The expression of AMH is stimulated through a synergistic effect of WT-1 and SF-1 while DAX-1 inhibits this synergistic interaction [39].

As indicated earlier, the postnatal expression of AMH in testicular tissue as well as the secretion of AMH declines at puberty. This decline in AMH at puberty is associated with anatomical, hormonal and cellular changes which are thought to be involved in the regulation of AMH. First, the formation of the blood-testis at puberty appears to redirect the secretion of AMH from the basal compartment of the seminiferous tubule into the lumen with a decrease in circulating AMH and an increase in seminal plasma AMH as result [22, 40, 41]. Second, the decrease in AMH concentrations at puberty coincides with an increase in testosterone concentrations [42]. This inverse relationship between testosterone and AMH [42] along with an increased AMH expression in testicular tissue

in the absence of androgen receptors suggests that testosterone has down-regulatory effect on AMH but requires the presence of androgen receptors [43, 44]. The last factor that appears to be associated with a decrease in AMH expression at puberty is the process of meiosis. The expression of AMH in seminiferous tubules appears to be largely dependent whether or not meiosis is initiated. Once meiosis is initiated, the expression of

AMH in the seminiferous tubule becomes no longer detectable [45, 46].

1.4.2 The regulation of AMH in females

The regulation of AMH in the follicle is not well understood primarily because of the contradictory results between different studies. Nonetheless, the expression of AMH as well as the secretion of AMH in follicular fluid is well characterized during different stages of follicular development. The expression of AMH in granulosa cells and the concentration of AMH in follicular fluid increases during follicular development until follicle selection while a rapid decrease in AMH expression and follicular fluid concentrations are observed after selection of the dominant follicle [47]. Bone morphogenetic proteins (BMPs) have been identified by several studies as having a stimulatory effect on the expression of AMH in granulosa cells [48, 49]. In addition to

BMPs, FSH plays a role in regulation of AMH. The effect of FSH on the mRNA expression of AMH in vitro appears to be different in cattle than in women. In cattle,

FSH has a down-regulatory effect on the expression of AMH mRNA in granulosa cells of medium sized follicles (5-10mm) as well as on the secretion of AMH by small and medium sized follicles [49]. In contrast, the AMH mRNA expression in human granulosa

cells is positively influenced by FSH [50]. Besides FSH, estradiol also appears to be involved in regulation of AMH. Still, there is some confusion over the regulation of estradiol and AMH. Two different studies have shown that AMH down-regulates the expression of aromatase in granulosa cells [51, 52] whereas a recent study showed that increasing concentrations of estradiol induces a reduction in the mRNA expression of

AMH via estrogen receptor- [53].

1.5 Biological activity of AMH

Two different bioassays based upon different in vitro organ culture systems have been described to examine the biological activity of AMH [54, 55]. The first method was based on a pioneering study which revealed that the Müllerian ducts regressed if they were cultured in vitro with the male fetal gonad in rats [56]. This organ culture system was later used and modified into a biological assay by Donahoe [54]. Briefly, the

Müllerian and Wolffian ducts were removed from a 14.5 day old fetal rat, separated from the fetal gonads, cultured in vitro with the sample of interest, embedded in paraffin, and examined histologically. The degree of regression of the Müllerian duct is evaluated on a scale from 0 to 5 with grade 5 representing complete regression of the Müllerian ducts

[54]. Interestingly, it has been shown that this method can be used to test biological material of many different species including the horse [21, 57]. This could potentially be explained by the fact that the biological activity of AMH is localized to the carboxyl terminus which appears to be highly conserved across different species. A second method that has been used to assess the biological activity of AMH in a sample involves the

measurement of aromatase activity in an ovary culture system which contains a fetal ovary of a 16 day old rat. An important advantage of this assay is that it provides quantitative data which can be obtained using lower concentrations of AMH [55].

1.6 Pre- and postnatal biological functions of AMH in males and females

1.6.1 Sexual differentiation: regression of the Müllerian ducts

Without any doubt, the most important biological function of AMH is regression of the

Müllerian ducts in the male fetus. Prior to sexual differentiation, the fetus contains an undifferentiated gonad along with a pair of Wolffian and Müllerian ducts. In the presence of the SRY gene, the undifferentiated gonad develops into a testis which begins to produce AMH and testosterone [58]. The interaction of AMH with the AMH receptor type II induces a cascade of events that result in the regression of the Müllerian ducts

[59]. In addition, the production of testosterone by the fetal Leydig cells causes the

Wolffian ducts to differentiate into the epididymis, vas deferens, and seminal vesicles. In the absence of the SRY gene, the undifferentiated gonad develops into a fetal ovary. In contrast to the testis, the fetal ovary does not produce AMH or testosterone. As result, the

Müllerian ducts develop into the fallopian tube, uterus, cervix and cranial end of the vagina while the Wolffian ducts regress.

As the gestation length differs between species, the period during gestation when the regression of the Müllerian ducts is initiated and completed differs. The regression of the

Müllerian ducts is completed by gestational day 17 in rats [54], day 46 in dogs [60] and

by day 64 in humans [61]. It is unknown when regression of the Müllerian duct is initiated or even completed in the equine male fetus.

1.6.2 Folliculogenesis

Although regression of the Mullerian duct is a critical feature of AMH during fetal development, the role of AMH in folliculogenesis is emerging as an important function.

The initial studies examining the function of AMH during folliculogenesis made use of

AMH knock-out mice [62]. First, it was demonstrated that the depletion of the follicular pool occurred earlier in AMH knock-out mice than in wild-type mice. This rapid decline in primordial follicles in AMH knock out mice was attributed to an increased rate of follicular recruitment. Based on these results, it was concluded that the recruitment of primordial follicles is inhibited by AMH [62, 63]. The negative influence of AMH on the recruitment of primordial follicles was also examined by assessing follicular growth in the ovary of newborn mice after ovaries were cultured in vitro with or without AMH.

Indeed, the number of growing follicles was significantly lower in ovaries cultured with

AMH than without AMH [64]. Furthermore, as the number of growing follicles was increased in AMH knock-out mice, the possibility exists that the growth of FSH-sensitive follicles was also reduced by AMH [62, 63]. A subsequent study by the same laboratory clearly showed that the growth of FSH-sensitive preantral follicles in mice is inhibited by

AMH [65]. Likewise, Pellet et al showed that granulosa cells of growing follicles become less sensitive to FSH in the presence of AMH [66].

1.6.3 Inhibition of Leydig cell differentiation

As already expected by the relatively high concentrations of AMH in males prior to puberty, AMH also has an important role during the postnatal period. The first indication that AMH has an influence on Leydig cell differentiation was provided by Behringer who showed that testicular tissue in AMH deficient mice is characterized by Leydig cells hyperplasia [67]. Anti-Müllerian hormone, acting through the AMH receptor on the

Leydig cells, does not only inhibit the differentiation of Leydig cells but also decreases the steroidogenic activity of Leydig cells. More specifically, AMH appears to have a down-regulatory effect on the mRNA expression of cytochrome P450 17α- hydroxylase/C17-20 lyase in Leydig cells [68]. In accordance to this study, Rouiller-

Fabre et al. demonstrated that testicular steroidogenesis is inhibited by

AMH [69].

1.7 Molecular aspects of AMH signaling

Two types of transmembrane serine/threonine kinase receptors are involved in the signal transduction of AMH, namely AMH receptor type 2 (AMHR2) and AMH receptor type 1.

In contrast to AMHR2, it appears that the AMH receptor type 1 is shared among some members of TGF-β family. As signal transduction of AMH is mediated through smad 1,

5, and 8, it is likely that AMH shares its type 1 receptor with BMPs which have three different types of type 1 receptors: ALK2, ALK3, and ALK6. Furthermore, one or more of these receptors forms a receptor complex with AMHR2 when AMH binds to AMHR2.

After the type 1 receptor is activated, smad 1, 5, 8 are phosphorylated. Subsequently, the

phosphorylated smad proteins bind to smad 4 and the complex is transported to the cell nucleus to initiate transcription of the target gene [5].

Chapter Two: Serum anti-Müllerian hormone concentrations in stallions:

Developmental changes, seasonal variation, and differences between intact stallions,

cryptorchid stallions and geldings

Theriogenology 2013, 79 (9), 1229-35

2.1 Introduction

Anti-Müllerian hormone (AMH) or Müllerian inhibiting substance is a homodimeric disulfide-linked glycoprotein hormone that belongs to the transforming growth factor-β superfamily. In males, anti-Müllerian hormone is secreted by the Sertoli cells early in fetal life where it induces the regression of the Müllerian ducts. Even though its crucial role in sexual differentiation is accomplished in utero, the secretion of human AMH persists after birth [3, 22]. The postnatal secretion of human anti-Müllerian hormone is characterized by high concentrations of AMH during the prepubertal period followed by a significant decrease at the onset of puberty [25, 27, 70]. Previous studies have described the expression of AMH in the fetal, prepubertal, postpubertal and cryptorchid equine testis by immunohistochemistry. The expression of AMH is intense in the Sertoli cells of the fetal equine testis, decreases in the prepubertal testis with faint to no immunoexpression in the testis of adult stallions [19, 71]. Changes in serum AMH concentrations in intact stallions and whether they relate to sexual development have not yet been reported.

It appears reasonable that season might have an effect on peripheral AMH concentrations in stallions as numerous endocrine parameters change with season. The secretion of

gonadotropins, testicular steroids, and proteins is increased during the breeding season, which reflects an altered activity of hypothalamic-pituitary axis [72]. To date, only one study in Siberian hamsters suggests a seasonal influence on AMH concentrations as significantly higher serum AMH concentrations are observed in female hamsters when they are exposed to long day lengths compared to short day lengths [73]. The presence of a pronounced seasonal effect in stallions, as described above, would indicate that the stallion could serve as a good model to examine the effect of season on serum AMH concentrations.

Diagnostically, anti-Mullerian hormone is also better known for its ability to distinguish cryptorchidism from anorchia early in prepubertal boys. Because AMH is a Sertoli-cell derived product, measurable amounts of AMH are indicative of the presence of testicular tissue [30, 74-76]. The availability of a single, reliable serological test could be of additional value in the diagnosis of equine cryptorchidism. Ball et al. [19] demonstrated that the expression of AMH by Sertoli cells in the equine cryptorchid testis persists beyond puberty which suggests that the measurement of serum AMH might have some clinical utility in the diagnosis of cryptorchidism. To date, no evidence is available to support this hypothesis.

The objectives of the study reported here were to (i) validate a commercially available heterologous enzyme immunoassay for determination of anti-Müllerian hormone concentrations in the stallion (ii) determine serum anti-Müllerian hormone concentrations in neonatal colts and fillies, prepubertal colts , and postpubertal stallions, as well as serum testosterone concentrations in prepubertal colts and postpubertal

stallions (iii) determine seasonal variations in AMH and testosterone concentrations of intact mature stallions (iv) determine serum AMH and testosterone concentrations in geldings, cryptorchid stallions, and intact stallions, and (v) determine seminal plasma anti-Müllerian hormone concentrations in intact mature stallions.

2.2 Materials and methods

2.2.1 Validation of anti-Müllerian hormone and testosterone assay

Serum concentrations of anti-Müllerian hormone were measured using a commercially available heterologous sandwich enzyme immunoassay (Active AMH-ELISA;

Diagnostic Systems Laboratories, Webster, TX, USA) per manufacturer’s directions.

Final concentrations of AMH were measured in a dual wavelength mode (450/600 nm) using a microplate spectrophotometer (BioTek Powerwave HT, Winooski, VT, USA). To assess dilutional parallelism, serial dilutions of serum from 2 intact stallions were assayed, and the results obtained were compared to the standard curve. To validate the assay biologically, serum AMH concentrations were measured daily from 3 intact stallions immediately before and until 7 days after castration, and the decay in serum AMH was monitored over time. Furthermore, the biological half-life of AMH was determined by using the following equation: t1/2 = ln (2) / k where ln represents the natural logarithm, and k the decay constant. Serum testosterone concentrations were measured by radioimmunoassay, as previously described by Stabenfeldt et al. [77].

2.2.2 Serum anti-Müllerian hormone concentrations in neonatal colts and fillies, prepubertal colts and postpubertal stallions, and serum testosterone concentrations in prepubertal colts and postpubertal stallions

To characterize circulating AMH concentrations in neonates of different gender and in the stallion at different developmental stages, serum samples were collected from newborn fillies (n=5, < 3 days of age), newborn colts (n=8, < 2 days of age), prepubertal colts (n=8, 13-15 months of age) and postpubertal stallions (n=15, > 2 years of age). All serum samples were collected during the breeding season from horses boarded at a local

Thoroughbred stud farm or presented to Veterinary Medical Teaching Hospital (VMTH) at the University of California, Davis (UC Davis). All serum samples were stored at -20C° until assayed for AMH. Serum samples from prepubertal colts and postpubertal stallions were also assayed for testosterone. All animal research was conducted under an

Institutional Animal Use and Care committee protocol approved at UC Davis.

2.2.3 Seasonal variations in AMH and testosterone concentrations of intact mature stallions

Serum samples were collected between 8 and 9 AM on the 14th day of each month from

September 1st, 2010 till August 31st, 2011. Serum samples were stored at -20°C until assayed for AMH and testosterone. A total of nine intact mature stallions (age range: 8-

22 years of age) were used, including six Quarter Horse stallions, one Thoroughbred, one

Selle Français, and one Appaloosa stallion. All stallions were used in a teaching program and housed at either the Center for Equine Health (n=5) or at the Department of Animal

Science (n=4) at UC Davis. All stallions were fed alfalfa hay and had access to water ad libitum.

2.2.4 Serum anti-Müllerian hormone and testosterone concentrations in geldings, cryptorchid stallions, and intact stallions

Serum samples were utilized from accessions submitted to the Clinical Endocrinology

Laboratory and clinical cases admitted to the VMTH at the School of Veterinary

Medicine, UC Davis. A total of 97 serum samples were included and classified as geldings (n= 41), cryptorchid stallions (n= 41) or intact stallions (n= 15) according to criteria described below. The cryptorchid stallion group included horses with at least one retained testis regardless the status of the other testis (removed, retained, descended) and was subdivided into cryptorchid stallions with a scrotal testis (n=8) and without a scrotal testis (n=33). Only horses 2 years of age or older were included in this part of the study.

Serum samples from the Clinical Endocrinology Laboratory (n=70) were classified solely according to their history and serum testosterone concentrations. A history of either; castrated male, castrated male with stallion-like behavior , absence of scrotal testes or cryptorchid suspect was required in order to be included in the study. Final classification of those serum samples was based on reference ranges for serum testosterone concentrations provided by the Clinical Endocrinology Laboratory at UC Davis (gelding:

< 50 pg/mL; cryptorchid stallion: > 100 pg/mL). Classification of horses (n=27) presented to VMTH was based on a combination of history, clinical findings, testosterone concentrations, and/or surgery reports.

2.2.5 Detection of anti-Müllerian hormone in seminal plasma of a stallion and determination of seminal plasma AMH concentrations in intact stallions

Semen was collected from a stallion of known fertility using an artificial vagina

(Missouri Model) with a built-in filter. A small aliquot of raw semen was centrifuged

(1,500 x g) at 4C° for 5 min, and the supernatant was collected. Approximately 5.4 µg of seminal plasma protein was separated on 12% sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and electroblotted onto polyvinylidene fluoride (PVDF) membranes (Immobilon P, Millipore Corp., Bedford, MA) under non-reducing and reducing conditions. For analyses in reducing conditions, proteins were heated in the presence of β-mercaptoethanol. AMH protein was detected with polyclonal antisera against human recombinant AMH (1:2,500, courtesy of Dr. D. T. MacLaughlin, Boston,

MA, USA). Immunoreactive proteins were detected with HRP-linked IgG (1:10,000) and luminol reagents with electrochemiluminescence (Amersham, Arlington Heights, IL). To determine seminal plasma AMH concentrations, semen was collected from 7 additional stallions in April and processed under the same conditions as described above. The supernatant was stored at -20C° until analyzed for AMH and protein concentration.

Immediately after the semen collection, serum samples were collected from each stallion and stored at -20C° until assayed for AMH concentration.

2.3 Statistical analysis

Linear regression analysis was conducted to determine the biological half-life of AMH.

The Mann-Whitney U test was used to compare AMH concentrations between neonatal colts and fillies because of the small sample size. The Student’s t-test was used to 21

compare serum AMH and testosterone concentrations between prepubertal colts and postpubertal stallions. Because of the skewed distribution of the data, AMH and testosterone concentrations were log transformed, and a one-way analysis of variance was used to compare AMH and testosterone concentrations between intact stallions, cryptorchid stallions, and geldings. Tukey’s test was used to identify significant differences between individual means. For the seasonal data, testosterone concentrations were log transformed. A mixed model was used to compare monthly changes in serum

AMH and testosterone concentrations with stallion as a random effect and month as a fixed effect and differences between individual means were compared using Tukey’s test.

The correlation between serum AMH and testosterone concentrations in the seasonal data was determined by Pearson correlation coefficient. Because of the small sample size, the

Wilcoxon signed-rank test was used to examine differences between serum and seminal plasma AMH concentrations. The correlation between serum and seminal plasma AMH concentrations was determined by a nonparametric correlation (Spearman’s ρ). All statistical analyses were conducted with JMP (JMP, Version 9.0; SAS Institute; Cary, NC,

USA).

2.4 Results

2.4.1 Validation of the anti-Müllerian hormone assay for stallions

The curves of the serially diluted serum from 2 stallions were parallel to and within the range of the standard curve for the AMH assay (Figure 2.1). The intra- and inter-assay coefficients of variation for the AMH assay were 3.1 % and 7.8%, respectively.

Biological validation of the assay was demonstrated by the exponential decline of serum anti-Müllerian hormone concentrations in all 3 stallions after castration

(Figure 2.2). The biological half-life of AMH was 1.5 days.

Figure 2.1 Parallelism plot for the anti-Müllerian hormone assay. The curves of the serially diluted serum samples from two intact stallions are parallel to the standard curve.

Figure 2.2 Biological validation of the anti-Müllerian hormone assay. Plot natural logarithm of the mean AMH concentration of 3 stallions against time (day post castration). Data are expressed as mean ± s.e.m.

2.4.2 Serum anti-Müllerian hormone concentrations in neonatal colts and fillies, prepubertal colts and postpubertal stallions, and serum testosterone concentrations in prepubertal colts and postpubertal stallions

Serum AMH concentrations were higher (P<0.005) in neonatal colts (26.4 ± 2.1 ng/mL) than in neonatal fillies (1.2 ± 0.46 ng/mL). Furthermore, concentrations of AMH were higher (P<0.02) in prepubertal colts than in postpubertal stallions. In contrast, serum concentrations of testosterone were lower (P<0.0001) in prepubertal colts compared to postpubertal stallions (Figure 2.3).

Figure 2.3 Serum testosterone (black bar) and AMH (gray bar) concentrations in prepubertal colts and postpubertal stallions. Asterisks above the bars indicate statistical significance between different age groups (P<0.05). Data are expressed as mean ± s.e.m.

2.4.3 Seasonal concentrations of anti-Müllerian hormone and testosterone in intact stallions

Overall mean serum AMH concentrations across all stallions and months was 19.8 ± 0.65 ng/mL. There was an effect of time (P< 0.001) on serum AMH concentrations with significantly higher concentrations in May and June than from September till February

(Figure 4). Serum AMH concentrations peaked in May and reached a nadir in November with a 1.7 fold difference between peak and nadir concentrations. In contrast, serum testosterone concentrations varied more widely (Figure 2.4), and the overall mean concentration of testosterone was 988.4 ± 55.5 pg/mL. Similar to AMH concentrations,

there was an effect of time (P<0.002) on serum testosterone concentrations. Serum testosterone concentrations were significantly higher in April, May, and July than in

October. Furthermore, a weak but significant positive correlation (r = 0.27: P<0.006) was observed between serum AMH and testosterone concentrations.

Figure 2.4 Seasonal variation in AMH (solid line) and testosterone (dashed line) concentrations in intact stallions (n=9). Serum samples were collected at a monthly interval. Within AMH and testosterone, means with different superscripts (a,b) differ

(P<0.05). Data are expressed as mean ± s.e.m.

2.4.4 Serum anti-Müllerian hormone and testosterone concentrations in geldings, cryptorchid stallions and intact stallions

Mean serum AMH concentration was higher (P<0.004) in cryptorchid stallions than in intact stallions or geldings (Figure 2.5). In addition, mean serum AMH concentration was higher (P<0.0001) in intact stallions compared to geldings. Serum AMH concentrations in geldings were at or below the limit of detection for the assay (Figure 2.5). Serum AMH concentrations in cryptorchid stallions without a scrotal testis were higher (P<0.007) than serum AMH concentrations in intact stallions. Serum AMH concentrations tended

(P<0.1) to be higher in cryptorchid stallions without a scrotal testis than in cryptorchid stallions with a scrotal testis. Finally, no significant differences in serum AMH concentrations were observed between cryptorchid stallions with a scrotal testis and intact stallions (Figure 2.6).

As expected, serum concentrations of testosterone were higher (P<0.0001) in intact stallions (905.4 ± 110.9 pg/mL) compared to cryptorchid stallions (448.3 ± 40.0 pg/mL) or geldings (22.7 ± 1.3 pg/mL). Serum concentrations of testosterone were also higher

(P<0.0001) in cryptorchid stallions than in geldings.

Figure 2.5 Mean serum AMH concentration in cryptorchid stallions (n=41), intact stallions (n=15), and geldings (n=41). Different letters above the bars represent statistically significant differences (P <0.05). Data are expressed as mean ± s.e.m.

Figure 2.6 Mean serum AMH concentration in cryptorchid stallions without a scrotal testis (n=33), cryptorchid stallions with a scrotal testis (n=8), and intact stallions (n=15).

Different letters above the bars represent statistically significant differences (P <0.05).

Data are expressed as mean ± s.e.m.

2.4.5 Detection of anti-Müllerian hormone in seminal plasma of a stallion and determination of seminal plasma AMH concentrations in intact stallions

Immunoblotting of seminal plasma under non-reducing conditions revealed two immunoreactive proteins, one at 61 kDa which represents the monomer and another at

120 kDa which represents the dimer. Under reducing conditions, a less prominent band was observed at 53 kDa and a prominent band around 61 kDa (Figure 2.7).

No significant differences were observed between seminal plasma AMH concentrations

(49.1 ± 26.7 ng/mL) and their corresponding serum AMH concentrations (18.4 ± 2.7 ng/mL), and there was no significant correlation

(ρ= -0.19, P = 0.65) between serum and seminal plasma AMH concentrations. Mean seminal plasma AMH was 4.9 ± 2.45 ng/mg seminal plasma protein.

Figure 2.7 Western blot analysis of anti-Müllerian hormone in seminal plasma from a stallion under non-reducing and reducing conditions. The arrow represents the dimer at

120 kDa whereas the arrow head represents the monomer.

2.5 Discussion and Conclusion

To the authors’ knowledge, this is the first study to report serum anti-Müllerian hormone concentrations in neonatal colts and fillies, prepubertal colts and postpubertal stallions, and changes associated with season or cryptorchidism. The results of our study indicate that serum AMH concentrations are significantly higher in neonatal colts and prior to puberty, and that concentrations of AMH vary with season in intact mature stallions.

Moreover, serum anti-Müllerian hormone can serve as a biological marker for the presence of testicular tissue with higher serum AMH concentrations in cryptorchid stallions than in intact stallions.

The higher concentrations of AMH in neonatal colts compared to neonatal fillies are not entirely unexpected as similar sex-related differences in AMH concentrations are observed in cattle and humans [78, 79]. The secretion of AMH is elevated prior to puberty as demonstrated by higher concentrations of AMH in prepubertal colts compared to postpubertal stallions. This finding is consistent with the expression of AMH in fixed tissue specimens [19, 71]. In addition, significant differences in serum AMH concentrations between pre- and postpubertal stallions appear to be consistent among other mammalian species, such as cattle [78, 80] and sheep [81]. Because longitudinal sampling was not conducted in the present study, the exact period when serum AMH concentrations started to decline could not be determined. However, serum AMH in human males decreases at the onset of puberty with a rather rapid decline during the early stages of puberty [27].

The decrease in human AMH concentrations during puberty coincides with some important anatomical, cellular and endocrine changes including the formation of the blood-testis barrier, meiosis and increasing testosterone concentrations [82]. Similar changes have been observed at the molecular level during pubertal development of the stallion. Connexin 43, a gap junctional protein, is more strongly expressed between the

Sertoli cells of the seminiferous tubules after puberty which may reflect changes associated with the formation of the blood-testis barrier. Moreover, a negative correlation was present between the stage of development of the seminiferous tubule and the expression of AMH [71]. Finally, our results indicate that serum AMH concentrations decrease while testosterone concentrations increase during the transition from the prepubertal to postpubertal period. These data are consistent with the negative association between serum AMH and testosterone concentrations as reported to occur during puberty in humans [42]. However, whether all of the above described changes play an important role in the down regulation of AMH in the stallion still remains to be determined.

As expected, serum testosterone concentrations were higher in postpubertal stallions than in prepubertal colts. Significant differences in testosterone concentrations between prepubertal and postpubertal stallions are most likely due to an increase in testosterone concentrations during pubertal development. Puberty is characterized by slowly increasing testosterone concentrations beginning around 68 weeks of age with a rapid increase in testosterone around 75-80 weeks of age [83, 84].

Besides differences in AMH concentrations between pre- and postpubertal horses, there was also a significant seasonal effect on serum AMH concentrations in intact mature stallions. The increase in AMH concentrations during the spring and summer coincide with increases in gonadotropins and steroid hormones as previously reported in stallions

[72]. Therefore, seasonal variations of AMH in intact stallions might reflect a Sertoli cell response to an increased gonadotropic drive. Alternatively, seasonal changes in AMH concentrations in stallions could reflect the number of Sertoli cells in the testis. The population of Sertoli cells in stallions is variable as indicated by increasing numbers of

Sertoli cells during the breeding season [85-87] with the largest number present in May and June [88]. This corresponds well with the peak of serum concentrations of AMH observed here.

Several studies have shown that serum testosterone concentrations vary with season, with significantly higher concentrations during the breeding season [89-91]. This seasonal variation in testosterone concentrations was confirmed by the present study despite the wide variation in testosterone concentrations throughout the year. We speculate that environmental cues might be responsible for the wide variability in testosterone concentrations between different months.

In addition to seasonal changes, serum AMH concentrations were also elevated in cryptorchids, suggesting that AMH concentrations may be useful in the diagnosis of cryptorchidism. Moreover, serum concentrations of AMH in geldings were at or below the limit of detection of the assay suggesting that the testis is the sole source of

circulating AMH in the stallion whereas testosterone may be synthesized by the adrenal cortex in addition to the testis [92]. Therefore, serum AMH concentrations might be more specific for the presence of testicular tissue than serum testosterone. The present study was not able to determine the sensitivity or specificity of AMH for diagnosis of cryptorchidism in the stallion because the exact gonadal status of serum samples derived from the Clinical Endocrinology Laboratory was unknown. However, earlier work in prepubertal boys reported that anti-Müllerian hormone appears to be more sensitive and specific than testosterone in the diagnosis of anorchia [74]. Interestingly, serum AMH concentrations were higher in cryptorchid stallions than in intact stallions. Recently,

Almeida et al. characterized some molecular differences between the cryptorchid and descended equine testis [93]. The cryptorchid testis resembles in many characteristics the prepubertal testis as demonstrated by an increased expression of AMH and a reduced expression of connexin 43. However, further studies into the molecular regulation of

AMH in normal and cryptorchid testes are needed to better understand the genesis of elevated AMH levels in cryptorchid versus intact stallions of similar ages.

Apart from measurable serum AMH concentrations, we were also able to detect and measure AMH concentrations in seminal plasma from intact stallions. It has been speculated that the formation of the blood-testis barrier during puberty results in a shift in the direction of AMH secretion from the basal compartment into the circulation towards the seminiferous tubule lumen into seminiferous fluid [22]. As a result, concentrations of human AMH in seminal plasma exceed serum AMH concentrations [22, 40, 41].

However, the present study did not detect significant differences or even a positive

correlation between seminal plasma and serum AMH concentrations in the stallion; seminal plasma AMH concentrations varied widely among different stallions. Even though AMH is present in seminal plasma of stallions, the biological function of AMH in seminal plasma still needs to be determined.

In conclusion, our results indicate that serum AMH concentrations are significantly higher in neonatal colts than in fillies, and higher in prepubertal colts compared to postpubertal stallions. Besides sex and developmental-related differences, there is a seasonal variation in AMH concentrations among intact mature stallions with highest

AMH concentrations during the physiological breeding season. Finally, serum AMH concentrations are higher in cryptorchid stallions than either intact stallions or geldings.

Therefore, anti-Müllerian hormone appears to be a useful biomarker for the presence of testicular tissue and can serve as a valuable alternative in the endocrine diagnosis of cryptorchid or monorchid horses.

2.6 Acknowledgments

Financial support for this research project was provided by the John P. Hughes

Endowment, Albert G. Clay Endowment and Center for Equine Health at UC Davis.

Chapter Three: The interrelationship between anti-Müllerian hormone, ovarian

follicular populations, and age in mares

(Equine Veterinary Journal, Accepted)

3.1 Introduction

Anti-Müllerian hormone (AMH) is a granulosa-cell derived hormone that plays a major regulatory role in female reproduction during folliculogenesis [94]. The recruitment of primordial follicles [62] as well as the growth of FSH-responsive follicles [65, 66] is inhibited by AMH. In cows, serum AMH concentrations vary widely among individuals of similar age and are positively related to the number of antral follicles, better known as antral follicle count (AFC) [95]. Furthermore, in cattle and in mice, concentrations of

AMH as well as AFC reflect the number of primordial follicles, and both AFC and AMH serve as biomarkers for follicular reserve [95, 96]. As women age, the size of the follicular pool declines and eventually becomes depleted at menopause [97]. However, the age when menopause occurs varies widely among women with corresponding differences in reproductive lifespan [98]. Among several other factors [99], differences in the number of primordial follicles in the ovary at birth are thought to contribute to variation in the reproductive lifespan of females. As AMH is strongly correlated with the number of primordial follicles [96], and given that the number of primordial follicles varies widely within females of similar age [95], circulating AMH concentrations appear to be a better reflection of reproductive age than calendar age [100].

In contrast to cattle and women, relatively little is known about AMH in the mare. A previous study in mares showed that the immunoexpression of AMH is restricted to granulosa cells of the equine ovary and varies with follicular development [21]. With the exception of primordial and atretic follicles, immunohistochemical labeling for AMH can be observed in equine granulosa cells of pre-antral and small antral follicles with increasing intensity as the number of granulosa cell layers increases. In contrast, the expression of AMH decreases in granulosa cells of ovarian follicles larger than 30 mm

[21]. Anti-Müllerian hormone can be measured in the peripheral circulation of mares using a heterologous enzyme-linked immunosorbent assay (ELISA) [101]. Although serum AMH concentrations do not change significantly throughout the estrous cycle of mares, substantial variation exists between individual mares [101]. Vernunft et al. showed that plasma AMH concentrations were significantly higher in mares with a high number of growing follicles than in mares with a low number of growing follicles [102].

As described earlier, reproductive aging is associated with decreasing concentrations of

AMH in mice [96]. Although menopause does not occur in horses, older mares undergo ovarian senescence which is characterized initially by prolonged follicular phases and ultimately leads to cessation of ovulation and follicular growth [103]. Interestingly, ovarian senescence occurs in less than 50% of mares older than 24 years of age [104].

This indicates considerable age differences at the time of follicular depletion which, in turn, might be represented by differences in AMH concentrations between older mares based upon their remaining follicular population. Therefore, the objectives of this study were (i) to determine AFC and plasma AMH concentrations in mares of different ages and the repeatability within and across estrous cycles, and (ii) to determine the

relationship between plasma AMH concentrations and AFC with regard to mare age and follicle size.

3.2 Materials and Methods

3.2.1 Experimental design

All animal procedures in this experiment were performed in compliance with the animal use protocol approved by the Institutional Animal Care and Use Committee at the

University of Kentucky (#2010-067). A total of 45 light-horse mares of mixed breeds were included in this study. Mares, kept in pastures, had ad libitum access to grass hay and water. Mares were classified by age into 3 groups: young (3-8 y, n=10), middle-aged

(9-18 y, n=16), and old (19-27 y, n=19) mares with a mean age of 5.2, 14.2, and 23 years, respectively. Mares were examined for follicular activity by transrectal palpation and ultrasonography (Sonoscape S8, Universal Medical Systems Inc, Bedford Hills, NY) from mid-June to early September. Examinations were performed over two to three interovulatory intervals on alternate days from ovulation (day 0) till day 14, and then daily until ovulation. On day 12 of the first, day 6 and 12 of the second, and day 6 of the third estrous cycle, antral follicle counts were determined in combination with the determination of plasma AMH concentrations.

3.2.2 Determination of antral follicle counts

Transrectal ultrasonography was used to determine AFC which corresponds to the total number of antral follicles detected with ultrasound. Mares were sedated with 200 mg xylazine hydrochloride intravenously (AnaSed®,Llyod laboratories, Shenandoah, IA)

because AFC determination required mares to stand in stocks for long periods of time

(range: 5-50 min). The ovary was examined systematically with ultrasound starting at the ovarian pedicle and ending at the ovarian ligament. Sequential ovarian sections were examined and several cross-sectional image planes of the ovary were frozen for detailed examination. The largest diameter of all antral follicles as well as the location of each follicle was recorded on an “ovarian map”. The location of each follicle in relation to larger follicles as well as real time ultrasonography between the different sections was used to ensure that follicles were accurately assessed for size and number. In addition, antral follicles were classified by size into different groups (2-5, 6-10, 11-15, 16-20, 21-

25, 26-30, 31-35, 36-40, >40 mm). To avoid confusion with other anatomic structures, only antral follicles with a diameter equal to or larger than 2 mm were recorded. The ability to accurately determine the number of follicles equal to or larger than 2 mm using ultrasonography has been previously validated by Ginther and Pierson [105]. All mares were examined by the same individual during the entire study.

3.2.3 Measurement of plasma anti-Müllerian hormone concentrations

Blood samples were collected into heparinized tubes by venipuncture between 8 and 9

AM and kept on ice until centrifugation. Plasma recovered after centrifugation (1200 x g for 10 min) was divided into aliquots and stored at -20°C until analyzed for AMH concentrations. Plasma AMH concentrations were determined using a commercially available ELISA (MOFA Global AMH ELISA, Verona, WI) per the manufacturer’s instructions. According to the manufacturer, this assay was developed to measure AMH concentrations in equine serum and plasma samples. The authors examined serial

dilutions of plasma samples from 2 mares and assessed parallelism between the dilution curves and the standard curve (Figure 3.1). A microplate spectrophotometer (BioTek

Powerwave HT, Winooski, VT) was used to read the absorbance at 450 and 630 nm. The intra- and inter-assay coefficient of variation for anti-Müllerian hormone concentrations was 6.2%, and 15.8%, respectively. The limit of sensitivity of the AMH assay was 0.017 ng/mL.

Figure 3.1 Parallelism between dilution curves and standard curve of AMH. Serial dilutions (neat, 1:2, 1:4, and 1:8) of plasma samples from two different mares were examined and the percent binding (B/Bo in which Bo= maximum binding) of each diluted sample was determined. Parallelism between the curve derived from serial dilutions and the standard curve was assessed. The dilution curve of both plasma samples was parallel to the standard curve.

3.3 Statistical analysis

For each mare, AFC and plasma AMH concentration from all estrous cycles were averaged, and mean values were used for subsequent analysis. Differences in AFC and plasma AMH concentrations between different age groups were determined using the

Kruskal-Wallis test followed by the Wilcoxon test for multiple comparisons to identify which age groups differ from each other. A Wilcoxon signed rank test was used to detect differences in AFC between the left and right ovary. The repeatability of AFC and plasma AMH concentrations within and between estrous cycles was assessed using

Spearman’s rank correlation (ρ) and the Cronbach’s alpha coefficient (α). The relationship between AFC, plasma AMH concentrations and age was examined using a mixed model with AMH concentration as a covariate, age group, interaction of AMH by age group, estrous cycle, estrous cycle day and the interaction of estrous cycle day by estrous cycle as fixed effects, and mare as a random effect. The variable AFC was square root transformed to satisfy the assumptions of the mixed model. Correlations between

AFC and AMH concentrations independent of age and within different age groups were determined using the nonparametric Spearman’s rank correlation (ρ). A mixed model with mare as random effect was used to determine the relationship between plasma AMH concentrations and the number of antral follicles within different follicle size groups. The level of significance was set at P<0.05. Statistical analyses were carried out using JMP

(JMP, V.10.0; SAS Institute; Cary, North Carolina, USA)

3.4 Results

3.4.1 Antral follicle count and plasma AMH concentrations in mares of different age groups, and the repeatability within and across estrous cycles

Compared to old mares (median: 13.6 follicles), antral follicle count was higher in young

(P<0.007) (median: 25.5 follicles) and middle-aged (P<0.0006) (median: 27.8 follicles) mares, but no difference (P=0.4) in AFC was observed between young and middle-aged mares (Figure 3.2). Furthermore, AFC was higher (P<0.03) for the left ovary than for the right ovary in mares across all age groups. Plasma AMH concentrations varied widely within different age groups (Figure 3.3). Although no significant differences were observed between young (median: 0.29 ng/mL) and middle-aged mares (median: 0.47 ng/mL), plasma AMH concentrations were lower (P<0.008) in old mares (median: 0.21 ng/mL) compared to middle-aged mares

(Figure 3.3).

The repeatability of AFC and plasma AMH concentrations (P<0.0001) was high within

(AFC: ρ=0.96; AMH: ρ=0.91) and between (AFC: ρ=0.87-0.95; AMH: ρ=0.78-0.85) estrous cycles. Similarly, the overall Cronbach’s alpha coefficient was high (AMH:

α=0.95; AFC: α=0.99) which indicates that AFC and AMH concentrations are consistent within and between estrous cycles.

Figure 3.2 Box plots showing antral follicle count (AFC) in young (3-8 y), middle-aged

(9-18y) and old (19-27y) mares. The box represents the 25% quartile, median, and 75% quartile, whereas the lower whiskers represent the 25% quartile – 1.5*(interquartile range) and the upper whisker represents the 75% quartile + 1.5*(interquartile range).

Outliers are indicated by individual dots. Antral follicle count was significantly higher in young and middle-aged mares than in old mares. Different letters above the box plots indicate statistically significant differences. Significance was set at P<0.05.

Figure 3.3 Box plots of plasma anti-Müllerian hormone (AMH) concentrations in young

(3-8y), middle-aged (9-18y) and old (19-27y) mares. The box represents the 25% quartile, median, and 75% quartile, whereas the lower whiskers represent the 25% quartile – 1.5*(interquartile range) and the upper whisker represents the 75% quartile +

1.5*(interquartile range). The individual dot indicates an outlier. Plasma AMH concentrations were lower (P<0.008) in older mares than in middle-aged mares. Different letters above the box plots indicate statistically significant differences. Significance was set at P<0.05.

3.4.2 Relationship between AFC and plasma AMH concentrations with regard to age and follicle size

Analysis of the mixed model (P<0.0001, r2=0.97) indicated that AFC was significantly related to the concentration of AMH (P=0.0004) and age group (P=0.003), as well as the interaction of AMH by age group (P=0.003). Estrous cycle (P=0.7), estrous cycle day

(P=0.8) and the interaction of estrous cycle day by estrous cycle (P=0.9) were not related to AFC. Across all mares, AFC was positively (ρ=0.77) correlated (P<0.0001) with the concentration of AMH (Figure 3.4), and this relationship was influenced by age group as shown by the higher correlation coefficient in older (ρ=0.86; P<0.0001) mares than in middle-aged (ρ=0.60; P=0.01) mares (Figure 3.5). In contrast, the correlation (ρ=0.40) between AFC and AMH was not significant in young mares (Figure 3.5).

Finally, analysis of statistical model (P<0.0001, r2=0.89) showed that plasma AMH concentrations were positively associated (P < 0.05) with the number of antral follicles for follicles between 6 and 20 mm in diameter; whereas, the number of follicles between

2 and 5 mm (P=0.2) or greater than 20 mm (P>0.2) were not associated with plasma

AMH concentrations.

Figure 3.4 Correlation between AFC and plasma AMH concentration independent of mare age. The black dots represent mean AFC and AMH concentrations. Antral follicle count is positively correlated (ρ=0.77, P<0.0001) with plasma AMH concentrations.

Figure 3.5 Correlations between AFC and plasma AMH concentration within different age groups (shown from left to right: young (3-8y), middle-aged (9-18y) and old (19-27y) mares). The black dots represent the mean AFC and AMH concentrations. The Spearman correlation coefficient was higher in older (ρ=0.86, P<0.0001) mares than in middle-aged

(ρ=0.60, P=0.01), while no significant correlation (ρ=0.40, P<0.4) was detected in young mares.

3.5 Discussion and Conclusion

To the authors’ knowledge, this is the first report that describes the interrelationship between AFC, peripheral AMH concentration and age in mares. Our results indicate that antral follicle count was strongly correlated with plasma AMH concentrations in old mares and that plasma AMH concentrations were significantly related to the number of small antral follicles between 6 and 20 mm in diameter. Furthermore, antral follicle count

was significantly lower in older mares than in young and middle-aged mares and plasma

AMH concentrations were significantly lower in old mares than in middle-aged mares.

The positive correlation between AFC and AMH concentrations has also been described in cattle [95], mice [96], and women [106-108], and likely reflects the follicular origin of anti-Müllerian hormone. Contrary to studies in other species, the present study examined the relationship between AFC and AMH across different age groups. Interestingly, the correlation between AFC and AMH differed by mare age with a strong correlation in old mares, a moderate correlation in middle-aged mares, and no significant correlation between AFC and AMH in young mares. Whether this finding reflects a biological difference for the horse compared to other species reported to date remains to be determined.

For the mare, circulating AMH concentrations were associated with the number of antral follicles between 5 and 21 mm in diameter. In women, approximately 60% of circulating human AMH is derived from antral follicles between 5 to 8 mm, which corresponds to follicles up to the time of follicular deviation in women [47]. The comparable follicular diameter in mares is approximately 11-17 mm [109]. Circulating concentrations of AMH in mares were not significantly influenced by the number of antral follicles larger than 20 mm. This might be explained by a limited number of follicles larger than 20 mm compared to other follicle size classes. However, more importantly, it might suggest that the secretion of AMH by follicles larger than 20 mm subsided around follicle deviation, a process initiated when the two largest follicles reach a diameter of 19.0 and 22.5 mm

[110]. Indeed, it has been shown in women that the expression of AMH as well as follicular fluid concentrations of AMH decreases rapidly after follicle selection [47].

The lower number of antral follicles in older mares, as observed in the study, likely reflects an age-related reduction in follicular reserve. Similarly, Ginther et al. showed that number of follicles smaller than 20 mm per follicular wave was significantly lower in older mares compared to young mares [111]. Furthermore, the wide variation in antral follicle counts within age groups suggests inherent differences in the population of ovarian follicles among mares. Indeed, the number of primordial follicles in young mares was estimated to range from 5600 to 75000 [112]. Data in mice indicate that the number of growing and primordial follicles is strongly correlated to each other as well as to AMH and that both parameters decrease with maternal age [96]. This finding is supported in cattle as AFC is strongly associated with concentrations of AMH and primordial follicle count [95]. Based on these results together with the significant correlation between AFC and AMH in middle-aged and older mares, we speculate that anti-Müllerian hormone can provide valuable information in the assessment of ovarian reserve in middle-aged and older mares. The ability to assess ovarian reserve could have some clinical utility in mares experiencing early signs of ovarian senescence such as prolonged or even erratic interovulatory intervals. As in sheep [113] and in women [114], plasma AMH concentrations were highly repeatable within and between consecutive estrous cycles.

This, together with the relatively long biological half-life [115], suggests that AMH is a consistent and useful biological marker of ovarian function.

Finally, data in cattle, ewes and goats suggest that variations in AMH concentrations reflect differences in fertility [116-118]. Anti-Müllerian hormone concentrations are positively associated with the response to superovulatory agents [119] and the number embryos recovered from superovulated cows [120, 121] and does [118]. Furthermore, first service conception rate is positively related to the prepubertal AMH concentrations in ewes [117]. In women, concentrations of AMH have been used to predict the success of assisted reproductive technology procedures [122-124]. However, AMH does not appear to be a good predictor in younger women [125, 126]. Whether these findings can be extrapolated to mares still remains to be determined.

In summary, a positive correlation exists between AFC and plasma AMH concentration which is influenced by age as indicated by a strong and moderate correlation in old and middle-aged mares, respectively. Finally, circulating AMH concentrations are primarily influenced by the number of antral follicles between 6 and 20 mm in diameter.

3.6 Acknowledgements

Funding for this project was provided by the Albert G. Clay Endowment of the

University of Kentucky. The authors would like to thank MOFA Global for providing the

AMH ELISA. We would also like to thank Dr. E.Woodward, C. Fedorka, and Dr. G.

Davolli for their assistance with the clinical part of this research project.

Chapter Four: Molecular and hormonal differences in the equine follicle in relation

to variations in antral follicle count and stage of follicular development

4.1 Introduction

Folliculogenesis is a complex and dynamic process regulated by various factors and hormones signaling via autocrine, paracrine, and endocrine pathways. One of the hormones that plays an important role in this process is anti-Müllerian hormone (AMH), a glycoprotein that is synthesized by the granulosa cells of the ovary and secreted into the peripheral circulation [3]. Circulating AMH concentrations vary widely between mares

[101], and this wide variability in AMH is mainly attributed to differences in antral follicle counts (AFC) indicating the potential usefulness of AMH as marker for ovarian reserve [127]. In cattle, increasing amount of data indicate that differences in the number of antral follicles are not only a reflection of ovarian reserve but are also related to follicular function [128-130]. Foremost, distinct differences in the mRNA expression of steroidogenic enzymes and intra-follicular steroid concentrations have been observed in cows with low and high antral follicle counts [129, 130]. More specifically, the mRNA expression of aromatase in granulosa cells as well as follicular fluid estradiol concentrations is increased in growing follicles of cows with a low number of antral follicles [130] while the mRNA expression of 17α-hydroxylase in theca cells of growing follicles is decreased in cows with low AFC [129]. Secondly, it has been shown that the mRNA expression of the FSH receptor (FSHR) in granulosa cells as well as the responsiveness of granulosa cells to FSH is reduced in cows with low AFC compared to

cows with high AFC [128]. To the authors’ knowledge, follicular function in mares has never been linked to AFC or to circulating AMH concentrations.

Equally important, but not well documented, are molecular and hormonal changes in the equine follicle during follicular development with regards to AMH. In women, concentrations of AMH in follicular fluid decrease with follicular development [47, 131], and follicular AMH concentrations are negatively correlated with the mRNA expression of aromatase and intra-follicular estradiol concentrations [47, 132]. Furthermore, in vitro studies demonstrated that AMH reduces the expression of aromatase in granulosa cells

[51, 52]. A recent study, however, revealed that increasing estradiol concentrations reduces the expression of AMH mRNA through the action of estradiol via estrogen receptor beta (ESR2) [53]. Thus, there is still some confusion regarding the interaction of

AMH, estradiol, and aromatase in the follicle. Follicle-stimulating hormone acting through the FSHR also plays a role in the regulation of AMH; however, it is unclear whether FSH has a stimulatory or inhibitory effect on the expression or synthesis of

AMH [49, 50]. Although several regulatory factors for AMH have been identified in other species, data regarding the regulation of AMH in the equine follicle are lacking.

Identifying genes that are co-expressed with AMH in the equine follicle and genes that are differentially regulated during follicular development will provide insight regarding the regulation of AMH in the equine follicle. Therefore, the objectives of this study were

(i) to examine molecular and hormonal differences in growing and dominant equine follicles in relation to variations in AFC and plasma AMH concentrations (ii) to identify genes co-expressed with AMH in granulosa cells of growing and dominant follicles, and

(iii) to examine molecular and hormonal differences in the equine follicle in relation to the stage of follicular development.

4.2 Materials and Methods

4.2.1 Animal tissue collection

All animal procedures were approved by the Institutional Animal Care and Use

Committee (Protocol: #2010-067) of the University of Kentucky. Cyclic mares (n=30) of mixed breeds (mean (± sem) age: 15.6 ± 1.4 years; age range: 3-27 years) were examined by transrectal ultrasonography. Beginning at ovulation, two dimensions (width and height) of the six largest follicles were recorded every other day to track follicular waves.

When follicular growth was identified, ultrasonographic examinations were performed daily until euthanasia and tissue collection. A single blood sample was collected six days after ovulation for determination of plasma AMH concentrations, and AFC was determined as previously described [127]. Mares were euthanized either prior to follicle deviation (n=17 mares) when follicular growth was identified and the largest growing follicle, as identified by ultrasonography, reached a diameter between 15 and 20 mm or after follicle deviation during estrus (n=13 mares) when endometrial edema was present and the dominant follicle reached a diameter of 35 mm.

Ovaries were processed using a protocol similar to that described by Evans et al. [133].

Briefly, ovaries were removed immediately after euthanasia and kept on ice. The oviduct and connective soft tissue was dissected away from the ovary, and the ovary was rinsed with ice cold PBS. Depending on when the mare was euthanized, either small, growing

follicles (n = 1 to 4) or the dominant follicle (n = 1) was identified using a previously constructed ovarian map to confirm localization and identity of the follicle(s). Follicular fluid was aspirated with a 25G needle, snap frozen in liquid nitrogen, and stored at -80°C until processing. The follicle was bisected using a scalpel blade and a small section (full thickness) of intact follicular wall was retrieved and fixed in formalin for histology and immunohistochemistry. The theca-basement membrane-granulosa cell layer of the remaining ovarian follicle was peeled away and transferred to a Petri dish containing cold

RNAlater® (Invitrogen, Carlsbad, CA). Granulosa cells were scraped off the basement membrane and theca layer using a cell scraper (Fisher Scientific, Waltham, MA). The stripped granulosa cells in RNAlater® (Invitrogen) were aspirated, transferred to a cryovial, and stored at 5°C for 24 h before subsequent storage at -80°C. The remaining tissue, basement membrane and theca cell layer, was transferred to another cryovial containing RNAlater®, and stored under similar conditions. The combination of the basement membrane and theca cell layer will be referred as “theca cells” for the remainder of the study.

4.2.2 Histology

Formalin-fixed intact follicular wall was embedded in paraffin and paraffin blocks were sectioned (6 µm) for histology and immunohistochemistry. Sections were mounted on microscope slides, heat fixed (37 °C for 6 hours), and stained with hematoxylin and eosin for histology. Slides were examined (400x) with bright-field microscopy (Zeiss Axio

Imager Upright Microscope, Carl Zeiss MicroImaging GmbH, Göttingen, Germany).

Viable follicles were distinguished from atretic follicles using the criteria described by

Kenney et al. [134]. Follicles with signs of atresia were excluded from the study. One viable follicle per mare was identified from which granulosa and theca cells were used for further analyses.

4.2.3 RNA extraction and Real-time Reverse Transcriptase Polymerase Chain

Reaction (qRT-PCR)

Granulosa and theca cells stored in RNAlater® were thawed on ice. Extraction of total

RNA from granulosa cells and theca cells was performed using TRIzol® reagent

(Invitrogen). For granulosa cells, RNAlater® was diluted (1:1) with phosphate-buffered saline (PBS) to decrease its viscosity prior to centrifugation (2000 x g; 5 min).

Immediately after centrifugation, the supernatant was removed and 1 mL of TRIzol® reagent was added to the granulosa cell pellet. Theca cells were removed from RNA later® and transferred to a 5-mL cryovial tube that contained 1 mL of TRIzol® reagent.

All subsequent steps performed are described in the TRIzol protocol provided by the manufacturer. The purity of total RNA isolated from granulosa and theca cells was analyzed using the Nanodrop 2000 spectrophotometer (Thermo Scientific). All RNA samples were diluted with RNase-free water to a total volume of 150 µL, and precipitated by adding 150 µL of isopropanol and 15 µL of sodium acetate (pH=6.1) for at least 24 h.

Following precipitation, RNA samples were centrifuged (12,000 x g; 10 min), washed with 75% ethanol, centrifuged (12,000 x g; 5 min), air dried, and re-suspended with

RNase-free water. The purity of RNA was re-evaluated with the Nanodrop 2000

spectrophotometer. Only RNA samples with a 260/280 and 260/230 ratio between 1.7 and 2.3 were included in the study.

To digest contaminating DNA, 2 µg of total RNA was DNase-treated using a DNA- free™ Kit (Invitrogen). Subsequently, 2 µg of DNase-treated total RNA was reverse transcribed to cDNA using TaqMan® Reverse transcription reagents (Invitrogen) which contains a mix of RNase-free water, 10X Taqman RT Buffer, 25mM Magnesium

Chloride, deoxyNTPs mixture (2.5mM), random hexamers (50 µM), RNase inhibitor

(20/L), and Multiscribe™ Reverse transcriptase (50 U/µl). Thermal cycler conditions of the reverse transcription reaction were: 25°C for 10 min, 48°C for 30 min, and 95°C for

5 min.

Real-time PCR was used to examine the expression of glyceraldehyde-3-phosphate dehydrogenase (GAPDH), actin beta (ACTB), anti-Müllerian hormone (AMH), anti-

Müllerian hormone receptor type 2 (AMHR2), estrogen receptor 1 (ESR1), estrogen receptor 2 (ESR2), follitropin receptor (FSHR), luteinizing hormone/choriogonadotropin receptor (LHCGR), insulin-like growth factor 1 (IGF1), inhibin alpha (INHA), inhibin beta A (INHBA), and clone A17 cytochrome P450 aromatase (CYP19A1) in granulosa cells of growing and dominant follicles. The expression of GAPDH, ACTB, LHCGR, cytochrome P-450 17 alpha-hydroxylase/C17,20-lyase (CYP17A1), and steroidogenic acute regulatory protein (STAR) was examined in theca cells of growing and dominant follicles. Primers were designed using the primer design tool of the National Center for

Biotechnology Information (NCBI), and their corresponding sequences are shown in table 4.1. Real-time PCR of duplicate samples was performed using the 7900HT Fast real-Time PCR System (Applied Biosystems). Reactions contained a combination of

cDNA (25 ng), primers, and a SYBR Green Master Mix (Invitrogen). Cycle parameters of polymerase chain reaction were: 95°C for 10 min, followed by 40 cycles of 95°C for

15 sec and 60°C for 1 min, and then a dissociation step of 95°C for 15 sec. Melting curves for each sample were assessed to evaluate the specificity of the reaction.

Individual primer efficiencies for all the samples were obtained using LinRegPCR [135].

The ΔCT for each gene of interest was calculated by subtracting the CT of the housekeeping gene from the CT of the gene of interest. In a preliminary experiment, the expression stability of 4 different housekeeping genes (GAPDH, ACTB, glucuronidase beta, and beta-2-microglobulin) in granulosa and theca cells was examined using

Normfinder software [4]. The combination of GAPDH and ACTB was identified as the most stable reference transcripts (data not shown). Data analysis was performed using the

-ΔΔCt 2 method as described by Livak and Schmittgen [136]. The average ΔCT of all dominant follicles was used as a calibrator. Gene expression data are presented as RQ

(relative quantification) values. Finally, all obtained PCR products were purified using the GeneJET PCR Purification Kit (Thermo Scientific), and the size of the purified PCR products was confirmed using gel electrophoresis (4% agarose gel).

Table 4.1 Gene, tissue type: granulosa (G) versus theca (T) cells, NCBI reference sequence of the transcript, primer sequence of the

forward and reverse primer, and PCR primer efficiency.

Gene G /T NCBI reference sequence transcript Primer sequence of the forward and reverse primer PCR efficiency

GAPDH G, T NM_001163856.1 Forward: 5’ - AGAAGGAGAAAGGCCCTCAG - 3’ 2.03

Reverse: 5’ - GGAAACTGTGGAGGTCAGGA - 3’

ACTB G, T NM_001081838.1 Forward: 5’ - CGACATCCGTAAGGACCTGT - 3’ 1.99

Reverse: 5’ - CAGGGCTGTGATCTCCTTCT - 3’

AMH G JF330269.1 Forward: 5’ - CCATCTGGAGGAGCCAAC- 3’ 1.97

Reverse: 5’ - CCCGTGACAGTGACCTCAG - 3’

AMHR2 G XM_005611400 Forward: 5’ - CTAGACTTGCGGCCTGACA- 3’ 1.96

Reverse: 5’ - GCCCAGCTCTGCCTCATA - 3’

ESR1 G NM_001081772.1 Forward: 5’ - TCCATGGAGCACCCAGGAAAGC- 3’ 1.99

Reverse: 5’ - CGGAGCCGAGATGACGTAGCC- 3’

Table 4.1 (continued)

ESR2 G XM_001915519.2 Forward: 5’ - CAGTATTCCCAGCAGTGCCA- 3’ 1.91

Reverse: 5’ - GCCACAACACATTTGGGCTT- 3’

FSHR G S70150.1 Forward: 5’ - GCCATGGCATTGCTGACTGA- 3’ 2.01

Reverse: 5’ - ATCTTACAGGTGTGCCAGGAA- 3’

LHCGR G,T AY464091.1 Forward: 5’ - GGCACACCATCACCTATGCT- 3’ 1.89

Reverse: 5’ - AATTGCTGACACCCACAAGG- 3’

IGF1 G NM_001082498.2 Forward: 5’ - TTTCAACAAGCCCACAGGGT- 3’ 1.99

Reverse: 5’ - TCCAGCCTCCTCAGATCACA- 3’

INHA G NM_001081910.1 Forward: 5’ - AGAGCTGGACCGGGAAATTG- 3’ 2.05

Reverse: 5’ - CAAACGCCTGACTCCAGGAT- 3’

INHBA G NM_001081909.1 Forward: 5’ - GAGGATGACATCGGCAGGAG- 3’ 2.05

Reverse: 5’ - CGACAGGTCACTGCCTTCTT- 3’

Table 4.1 (continued)

CYP19A1 G AF031520.1 Forward: 5’ - CCACATCATGAAACACGATCA - 3’ 2.08

Reverse: 5’ - TACTGCAACCCAAATGTGCT- 3’

CYP17A1 T D30688.1 Forward: 5’ - GCATGCTGGACTTACTGATCC- 3’ 2.01

Reverse: 5’ - CTGGGCCAGTGTTGTTATTG- 3’

STAR T NM_001081800.1 Forward: 5’ - AGGAGAACCAGCAGGCAAAC- 3’ 1.98

Reverse: 5’ - TGCGTTCCACAAGCTCTTCA- 3’

4.2.4 Immunohistochemistry

The differential expression of two genes (AMH, CYP19A1) in growing and dominant follicles identified by qRT-PCR was further confirmed with immunohistochemistry. The immunoexpression of AMH and P450arom was evaluated in randomly chosen growing

(n=4) and dominant (n=4) follicles using an anti-human AMH antibody (1:100, goat polyclonal SC-6886, Santa Cruz Biotechnology Inc., Santa Cruz, CA [21]) and an anti- human P450 arom (1:500, rabbit polyclonal generously provided by Dr. Nobuhiro

Harada, Fujita Health University, School of Medicine, Aichi, Japan [137, 138]). All primary antibodies were diluted with Bond Primary Antibody Diluent (Leica

Biosystems). The Leica BOND-MAX system (Leica Microsystems, Buffalo Grove, IL), a fully automated system, was used to process all the slides. Briefly, paraffin sectioned slides were dewaxed and rehydrated prior to heat-induced antigen retrieval using an

EDTA-based (pH 8.9) proprietary solution (Leica Biosystems). Subsequently, all slides underwent a series of incubation steps using reagents from the Bond Polymer Refine

Detection kit: 3% hydrogen peroxide for 5 min, primary antibody for 15 min, blocking reagent for 8 min, HRP-labeled polymer for 8 min, and diaminobenzidine substrate for 10 min. Each incubation step was followed by washing step using a 1X wash solution derived by diluting the Bond Wash Solution 10X concentrate with distilled water.

Negative controls were obtained by omitting the primary antibody.

4.2.5 Measurement of estradiol and AMH concentrations in follicular fluid of growing and dominant follicles

Concentrations of estradiol in follicular fluid were measured with a solid-phase, competitive chemiluminescent enzyme immunoassay (Immulite® 1000 system, Siemens

Healthcare Diagnostics, Los Angeles, CA, USA). Follicular fluid samples were diluted with estradiol sample diluent (LE2Z, Diagnostics Products Corporation, Los Angeles,

CA, USA) to a ratio of 1:320 – 1:640 for growing follicles and 1:10,000 for dominant follicles. The sensitivity of the immunoassay was 15 pg/mL, and the intra-assay coefficient of variation was 15%. Concentrations of AMH in follicular fluid and plasma were measured with an ELISA (MOFA Global Equine AMH ELISA, Verona, WI, USA) according to the manufacturer’s instructions. The AMH assay buffer was used to dilute follicular fluid of growing (1:30) and preovulatory follicles (neat to 1:10). The intra-assay coefficient of variation for follicular fluid AMH was 12.3 %. Finally, serial dilutions of follicular fluid were analyzed for AMH and estradiol to assess dilutional parallelism with the respective standard curves. Concentrations of AMH and estradiol were linear within the analytic range of the assay and proportional to the dilution factor.

4.3 Statistical analysis

A multiple linear regression model including AFC or plasma AMH concentration, follicle type (growing versus dominant follicle), mare age, and the interaction of mare age by follicle type was used to examine whether differences in intrafollicular AMH concentrations, estradiol concentrations, and mRNA expression of selected genes

(table 4.1) in granulosa cells or theca cells were associated with AFC or plasma AMH concentrations. The Pearson correlation coefficient was used to examine mutual correlations between AFC, plasma AMH concentrations, and the expression of selected genes (table 4.1) in granulosa cells of growing and dominant follicles. Within each follicle type, the relationship between follicular diameter and AFC or plasma AMH concentrations as well correlations between the expression of AMH mRNA and the expression of other selected genes in granulosa cells were also examined using the

Pearson correlation coefficient. Differences in the expression of selected genes in granulosa and theca cells between growing and dominant follicles were assessed using an unpaired t-test. A one-way analysis of variance with a student’s t-test as post hoc test was used to detect differences between plasma AMH concentrations, follicular AMH concentrations in growing follicles, and follicular AMH concentrations in dominant follicles. The unpaired t-test was also used to detect differences in follicular fluid estradiol concentrations between growing and dominant follicles. Correlations between intrafollicular AMH concentrations, intrafollicular estradiol concentrations, the mRNA expression of AMH and the mRNA expression of CYP19A1 in granulosa cells were examined using the Pearson correlation coefficient. Where needed, variables were square root or log transformed to satisfy the assumptions of the employed statistical test but untransformed mean data (± sem) are presented. Statistical analyses were performed using JMP (JMP version 10.0, SAS Institute; Cary, North Carolina), and statistical significance was set at P<0.05.

4.4 Results

4.4.1 Molecular and hormonal differences in growing and dominant equine follicles in relation to variations in AFC and plasma AMH concentrations.

Antral follicle counts were positively correlated (P<0.001, r=0.79) with plasma AMH concentrations. Analysis of the overall multiple linear regression model including both growing and dominant follicles, revealed that the expression of AMH, AMHR2, ESR2, and INHA in granulosa cells was significantly associated with AFC and plasma AMH concentrations. The expression of FSHR in granulosa cells was significantly associated with plasma AMH concentrations, and there was a tendency (P=0.10) for an association between FSHR and AFC. The expression of ESR1 in granulosa cells was significantly associated with plasma AMH concentrations but not with AFC. In granulosa cells, there were no significant associations between the expression of IGF1, INHBA, and CYP19A1 and AFC or plasma AMH concentrations. Mare age was significantly associated with the expression of AMHR2 and ESR1 in granulosa cells but the interaction of mare age by follicle type was not significant. The expression of AMHR2 and ESR1 in granulosa cells increased with mare age. The expression of LHCGR, CYP17A1, and STAR in theca cells were not significantly associated with AFC or plasma AMH concentrations. In addition, neither estradiol nor AMH concentrations in follicular fluid were significantly associated with AFC or plasma AMH concentrations.

Within growing follicles, the expression of AMH, AMHR2, ESR2, and INHA in granulosa cells was significantly correlated with AFC (Figure 4.1) and plasma AMH concentrations

(Figure 4.2). In addition, the expression of ESR1 and FSHR was significantly associated

with plasma AMH concentrations and had a tendency to be correlated with AFC. In contrast to growing follicles, no significant correlations were observed in dominant follicles (data not shown). Within each follicle type, there was no relationship between follicular diameter and AFC or plasma AMH concentration.

Figure 4.1 Pearson’s correlations (r) between AFC and the expression of genes involved in follicular development (AMH, AMHR2, ESR1, ESR2, FSHR, LHCGR, IGF1,

INHBA, INHA) and steroidogenesis (CYP19A1) in granulosa cells of growing follicles.

The expression of genes is shown as RQ values. Note that the Y-axis scale is different for the expression of AMH, and therefore is plotted individually.

Figure 4.2 Pearson’s correlations (r) between plasma AMH concentrations and the expression of genes involved in follicular development (AMH, AMHR2, ESR1, ESR2,

FSHR, LHCGR, IGF1, INHBA, INHA) and steroidogenesis (CYP19A1) in granulosa cells of growing follicles. The expression of genes is shown as RQ values. Note that the

Y-axis scale is different for the expression of AMH, and therefore is plotted individually.

4.4.2 Identification of genes co-expressed with AMH in granulosa cells of growing and dominant follicles

Correlations between the expression of AMH and the expression of genes involved in follicular development (AMHR2, ESR1, ESR2, FSHR, LHCGR, IGF1, INHA, INHBA) and steroidogenesis (CYP19A1) were examined in granulosa cells of growing and dominant follicles (Table 4.2). In granulosa cells of growing follicles, the expression of

AMH was strongly correlated with the expression of AMHR2 and INHA. In addition, a moderate to strong correlation was observed between the expression of AMH and the expression of ESR2 and FSHR. In contrast, no significant correlations were observed in

granulosa cells of dominant follicles with the exception that the expression of AMH had a tendency to be moderately correlated with the expression of ESR2.

Table 4.2 Pearson’s correlations (r) between the expression of AMH mRNA and the expression of genes involved in follicular development (AMHR2, ESR1, ESR2, FSHR,

LHCGR, IGF, INHA, INHBA) and steroidogenic genes (CYP19A1) in granulosa cells of growing and dominant follicles.

Gene Gene Granulosa cells of Granulosa cells of

transcript transcript growing follicles dominant follicles

Correlation P-value Correlation P-value

AMH AMHR2 r=0.76 P=0.0004 r=0.45 P=0.12

AMH ESR1 r=0.37 P=0.14 r=0.42 P=0.15

AMH ESR2 r=0.66 P=0.004 r=0.52 P=0.07

AMH FSHR r=0.63 P=0.006 r=0.30 P=0.3

AMH LHCGR r=-0.36 P=0.2 r=0.34 P=0.3

AMH IGF1 r=-0.27 P=0.3 r=0.22 P=0.5

AMH INHA r=0.70 P=0.002 r=-0.06 P=0.9

AMH INHBA r=0.01 P=0.9 r=0.20 P=0.5

AMH CYP19A1 r=-0.01 P=0.9 r=0.22 P=0.5

4.4.3 Molecular and hormonal differences in the equine follicle in relation to the stage of follicular development.

Differences in the expression of selected genes in granulosa and theca cells between growing and dominant follicles were evaluated. The expression of AMH and several genes co-expressed with AMH (AMHR2, ESR2, and FSHR) was significantly greater in granulosa cells of growing follicles than in dominant follicles whereas the expression of

LHCGR, IGF1, INHA, and CYP19A1 was significantly greater in granulosa cells of dominant follicles (Figure 4.3). Likewise, the expression of CYP17A1 and STAR in theca cells was significantly higher in dominant follicles than in growing follicles. No significant differences were detected in the expression of LHCGR in theca cells between growing and dominant follicles.

Figure 4.3 The expression (RQ) of selected genes in granulosa cells of growing follicles

(G) prior to follicle deviation and dominant follicles (D) after follicle deviation. Asterisks above the bars indicate differences (P<0.05) in expression between growing and dominant follicles. Data are presented as mean ± sem. Note that the Y-axis scale is different for the expression of AMH.

Two differentially expressed genes (AMH and CYP19A1) in granulosa cells of growing and dominant follicles identified by qRT-PCR were examined using immunohistochemistry (Figure 4.4). The immunoexpression of AMH and P450arom was localized to the cytoplasm of the granulosa cells of growing and dominant follicles.

AMH immunostaining was more intense in granulosa cells of growing follicles whereas only a faint immunoexpression was detected in granulosa cells of dominant follicles. In contrast, only slight immunostaining for P450arom was detected in granulosa cells of growing follicles whereas a more intense immunolabelling for P450arom was observed in granulosa cells of dominant follicles.

Figure 4.4 The immunoexpression of AMH and P450arom in granulosa cells of growing and dominant follicles. Strong AMH immunolabelling is observed in granulosa cells of growing follicles (A) whereas only faint expression is detected in granulosa cells of dominant follicles (B). In contrast, only slight P450arom immunostaining was detected in granulosa cells of growing follicles (C) while the expression of P450arom was more intense in granulosa cells of dominant follicles (D). The small inset figure in A and C shows the negative control for that particular antibody.

Finally, concentrations of AMH were significantly higher in follicular fluid than in circulating plasma, and follicular fluid AMH concentrations were significantly higher in growing than in dominant follicles. In contrast, concentrations of estradiol in follicular fluid were significantly higher in dominant follicles than in growing follicles (Table 4.3).

During follicular development, follicular fluid AMH concentrations were positively correlated with the expression of AMH mRNA (r=0.80, P<0.0001), and follicular estradiol concentrations were positively correlated with the expression of CYP19A1 mRNA (r=0.84, P<0.0001). In addition, there was a strong negative correlation (r=-0.84,

P<0.0001) between AMH and estradiol concentrations in follicular fluid. Furthermore, follicular fluid AMH concentrations were negatively associated with the expression of

CYP19A1 mRNA (r=-0.77, P<0.0001), and follicular fluid estradiol concentrations were negative associated with the expression of AMH mRNA (r=-0.76, P<0.0001).

Table 4.3 Anti-Müllerian hormone and estradiol concentrations in follicular fluid of growing and dominant follicles. Data are presented are mean ± sem. Statistically significant differences (P<0.001) are indicated by the different letters.

Growing follicles Dominant follicles

(n=17) (n=13)

Anti-Müllerian hormone 155.7 ± 20.8 ng/mLa 13.5 ± 1.9 ng/mLb

Estradiol 242.0 ± 54.0 ng/mLa 2424.6 ± 217.5 ng/mLb

4.5 Discussion and conclusion

To our knowledge, this is the first study to examine molecular and hormonal differences in the equine follicle in relation to AFC, plasma AMH concentrations, and follicular development. Our results demonstrated that AFC and plasma AMH concentrations were highly associated and in turn were associated with molecular differences in the growing equine follicle prior to selection of the presumptive dominant follicle. Furthermore, several genes (AMHR2, FSHR, and ESR2) were co-expressed with AMH in the growing equine follicle prior to follicle selection. The expression of these genes decreased during follicular development coincident with a decrease in mRNA and protein expression of

AMH in the follicle. In contrast, intrafollicular estradiol concentrations increased and were inversely correlated with intrafollicular AMH concentrations during follicular development.

It is well established in cows that variations in AFC are associated with differences in ovarian function and reproductive success [116, 139]. Ovarian response to exogenous stimulation is reduced [140], and cows with low AFC have significantly lower fertility rates which may be associated with an increased incidence of aneuploidy in oocytes

[141]. In cows with low AFC, the expression of aromatase in granulosa cells as well as intrafollicular estradiol concentrations are increased [130] despite a lower expression of

CYP17A1 in theca cells of growing follicles [129]. In addition, the expression of AMH and FSHR is lower in granulosa cells in cows with low AFC. A reduced expression of

AMH might be a reflection of a diminished oocyte quality [130] while the lower expression of FSHR is associated with a reduced responsiveness to FSH in cattle and 72

women [128, 142]. It is hypothesized that chronically elevated concentrations of circulating FSH in cows with low AFC may induce refractoriness at the level of the follicle [128]. In contrast to reports in cattle, the present study reveals that variations in

AFC in mares were not accompanied by molecular and hormonal differences related to steroidogenesis in growing follicles. Nevertheless, low AFC in the mare appears similar to women and cows in regards to a reduced expression of AMH and FSHR. The expression of the AMH receptor (AMHR2) was also lower in granulosa cells of growing follicles from mares with low AFC or low AMH concentrations. Although little is known about the autocrine / paracrine regulation of follicular development by AMH, a reduced expression of AMHR2 might indicate that the AMH signaling pathway is compromised in mares with low AFC. Additionally, the expression of mRNA for estrogen-receptor β

(ESR2) is also reduced in mares with low AFC which may adversely affect proliferation and differentiation of granulosa cells and subsequent folliculogenesis in mares with low

AFC. Based upon these observations, low AFC in the mare is associated with molecular differences in the equine follicle which could have an adverse impact on follicular function; however, observed changes in aromatase expression and follicular fluid estradiol concentrations in mares with low AFC were not observed in the present study.

Besides the relationship between AFC, plasma AMH concentrations and molecular changes in the growing follicle, a number of genes (AMHR2, FSHR, ESR2) known to be involved in the regulation of AMH were co-expressed with AMH in the growing equine follicle prior to follicle selection. The co-expression of AMH and AMHR2 might indicate that autocrine / paracrine signaling plays a role in the regulation of AMH in the equine

follicle. Also FSH, acting through the FSHR, has an important role in the regulation of

AMH even though there is controversy whether it inhibits or stimulates the production of

AMH. In cattle, FSH had an inhibitory effect on the secretion of AMH in cultured bovine granulosa cells [49] while in humans, FSH stimulated the transcription of AMH [50]. In mares, circulating FSH concentrations peak when the diameter of largest growing follicle is approximately 13 mm and decline thereafter resulting in lower FSH concentrations during the later stages of follicular development [143]. A similar pattern is observed in the expression of AMH and FSHR in granulosa cells of the equine follicle; the expression of AMH and FSHR was higher in the small growing follicle than in the dominant follicle.

Therefore, it is likely that increased FSH concentrations are positively associated with expression of AMH in the equine follicle which could indicate that FSH has a stimulatory effect on the expression of AMH in the equine follicle. In addition to FSH, estradiol appears to regulate AMH expression in the human follicle with a down-regulation of

AMH via estrogen receptor β [53]. Even though the effect of estradiol on AMH in the equine follicle remains to be determined, our study showed that the mRNA and protein expression of AMH in granulosa cells and intrafollicular AMH concentrations decreased during follicular development concomitant with an increase in intrafollicular estradiol concentrations. Furthermore, the expression of ESR2 in granulosa cells was correlated with the expression of AMH in growing follicles and had a tendency to be correlated with the expression of AMH in dominant equine follicles. Therefore, we hypothesize that increasing concentrations of estradiol act on the ERβ during follicular development with subsequent down-regulation of AMH in granulosa cells of the equine follicle.

A number of transcripts examined in this study demonstrated differential regulation during follicular development as indicated by the significant differences between growing and dominant follicles. The expression of ESR2 was significantly lower in dominant compared to growing follicles consistent with data in women [144] and in rats showing that the expression of ESR2 is down-regulated after the LH surge [145]. Although we did not measure peripheral concentrations of LH in this study, the relatively long LH surge in mares [146] combined with an increased expression of LHCGR in granulosa cells of the dominant follicle led us believe that the LH surge was already initiated in our study.

Similar to ESR2, the expression of FSHR was lower in dominant follicles than in growing follicles. This lower expression of FSHR in dominant follicles likely coincides with lower circulating FSH concentrations during the later stages of follicular development

[147]. On the other hand, the expression of IGF1, INHA, and CYP19A1 is increased in dominant follicles, and likely coincides with increased intrafollicular concentrations of estradiol, IGF1, and INHA as intra-follicular estradiol, IGF1, and INHA begin to increase in the future dominant follicle immediately before follicle selection [148]. The importance of IGF1 during the process of follicle deviation has been well described in the mare. At the time of follicle deviation, an increase in IGF1 is observed in the dominant follicle but not in subordinate follicles [148]. Ablation of the largest follicle at the time follicle deviation results an increase in intra-follicular IGF1 concentrations in the second largest follicle which subsequently becomes the dominant follicle [149]. Furthermore, when IGF1 is injected in the second largest follicle around the time of follicle selection, this injected follicle will develop into the dominant follicle [150]. Thus, the expression of

several genes in the equine follicles is strongly influenced by the stage of follicular development.

Finally, in accordance to differences in the gene and protein expression of AMH between growing and dominant follicles, concentrations of AMH in follicular fluid were significantly higher in small growing follicles than in dominant follicles. This result supports a previous study showing that peripheral AMH concentrations are primarily associated with the number of follicles between 6 and 20 mm [127]. It also indicates that concentrations of AMH in follicular fluid decrease with increasing follicular size. Indeed,

AMH concentrations are negatively correlated with follicular diameter and become undetectable in the equine preovulatory follicle [151]. The high follicular AMH concentrations in growing follicles immediately prior to follicle selection compared to the very low concentrations in dominant follicles suggests that follicular AMH concentrations decline rather rapidly after follicle deviation. Studies in women have shown that follicular AMH concentrations do not decrease linearly throughout follicular development; a prominent and sharp decrease in AMH concentrations in follicular fluid is observed when the dominant follicle is selected from a cohort of recruited growing follicles [47]. The sharp decline in follicular AMH concentrations further suggests that

AMH might play an important role in follicle selection in the horse.

In conclusion, our study demonstrates that variations in AFC and plasma AMH concentrations are associated with molecular differences in the equine follicle which, in

turn, may have an impact on follicular function. In addition, the expression of AMH and genes co-expressed with AMH in the equine follicle as well as intrafollicular AMH concentrations decrease during follicular development while follicular estradiol concentrations are inversely related to follicular AMH concentrations.

4.6 Acknowledgements

The authors would like to thank Alexandra Betancourt for her technical assistance with the qRT-PCR. Funding for this project was provided by the Albert G. Clay endowment of the University of Kentucky and by MOFA Global. Dr. Troedsson, a co-author, is professionally connected as a consultant to MOFA Global.

Chapter Five: The influence of age, antral follicle count and diestrous ovulations on

estrous cycle characteristics of mares

5.1 Introduction

Aging is an important physiological process that has a negative impact on the reproductive system [152]. Reproductive aging in mares is associated with a decreased fertility that has been associated with a variety of factors including a delayed uterine clearance [153], reduced oocyte quality [154], and a higher rate of early embryonic loss

[155]. Nevertheless, older mares are often bred because of their genetic value and/or performance and not for their fertility. Therefore, it is not uncommon to encounter aged broodmares in equine practice. To optimize the reproductive management of these older mares, veterinarians practicing reproductive medicine should understand which, how, and when reproductive parameters change with age. Reproductive aging in pony mares is characterized by prolonged follicular phases, longer inter-ovulatory intervals and increased circulating FSH concentrations [103, 156, 157]. Compared to light horse mares, pony mares have longer inter-ovulatory intervals [158], fewer multiple ovulations, and fewer diestrous ovulations [159]. Because of differences in reproductive parameters between pony and light-horse mares, extrapolation of findings related to effects of reproductive aging from the existing studies in ponies to horse mares may not always be possible.

Aging in mares is also associated with a decrease in the population of antral follicles

[111, 157]. Nevertheless, the number of antral follicles varies widely between aged mares

[160]. As large individual differences in follicular populations exist within the same age 78

group, it is important to distinguish reproductive age from chronological age [98, 161].

Therefore, it is possible that some of the age-related changes in follicular parameters are linked to differences in antral follicle counts (AFC). The number of primordial follicles, representing the ovarian reserve, decreases with mare age due to a high turnover of follicles during each follicular wave. The follicular pool eventually becomes depleted which results in ovulation failure and cessation of estrous cycles [162]; however, the age at which ovarian senescence occurs varies widely between older mares [104] suggesting that there are inherent differences in the number of primordial follicles between mares of the same age along with an age-related decline in primordial follicles,

Mare age is not the only factor that has an impact on different parameters of the estrous cycle. The occurrence of major secondary follicular wave is also associated with changes such as a prolonged inter-ovulatory interval and a delay in the emergence of the primary follicular wave. [163, 164]. A major secondary wave is characterized by the development of a large diestrous follicle which either ovulates (diestrous ovulation), luteinizes, or regresses [143]. The incidence of diestrous ovulations is extremely low in pony mares [159] but can be as high as 21% in Thoroughbred mares [165]. It is generally accepted that when a diestrous ovulation precedes luteolysis it can result in a prolonged luteal phase (>30 days) as the immature corpus luteum has a reduced sensitivity to endometrial PGF2α. However, the majority of the diestrous ovulations occur within the first 10 days after ovulation [166], and therefore, are not associated with prolonged luteal phases. To the authors’ knowledge, relative little has been published about the effect of diestrous ovulations on estrous cycle characteristics in light horse mares. Therefore, the

objectives of this study were (i) to determine age-related differences in follicular dynamics, endocrine profiles, and primordial follicles (ii) to examine the influence of antral follicle count on age-related changes in follicular parameters, and (iii) to examine the effect of diestrous ovulations on follicular parameters and progesterone concentrations.

5.2 Materials and Methods

5.2.1 Animal procedures

All animal procedures were approved by the Institutional Animal Care and Use

Committee of the University of Kentucky (#2010-067). Mares of mixed breeds, weighing approximately 450-650 kg, were assigned to one of the following groups: young (n=10,

3-8 yr), middle-aged (n=16, 9-18 yr), or aged (n=19, 19-27 yr) mares. This study was part of a larger experiment that examined the relationship between anti-Müllerian hormone, antral follicle count, and age in mares [160]. Briefly, mares were examined using transrectal ultrasonography during two consecutive estrous cycles. Ultrasound examinations were conducted every-other day for 14 days after ovulation and then daily until the subsequent ovulation. Follicular growth was monitored by recording two dimensions (width and height) of the six largest follicles and the location of all recorded follicles was drawn on an ovarian map. On day 12 of each estrous cycle, antral follicle count (AFC) was determined as previously described [160]. Mares were classified as low, medium, and high AFC based upon the interquartile ranges for AFC. Mares with a detectable corpus luteum and turgid uterine tone that did not return to estrus by day 22

post ovulation were given prostaglandin F2α (Lutalyse®, dinoprost tromethamine, 5 mg intramuscularly) and data from that estrous cycle were excluded from the analysis. Daily blood samples were collected by jugular venipuncture into heparinized tubes between 8 and 9 AM. Plasma samples obtained after centrifugation (1200 x g, 10 min) were stored at -20°C until analysis for progesterone and follicle stimulating hormone (FSH). After completion of the study, ovaries were removed from six young mares (age range: 3-8 yr) and four aged (age range: 19-26 yr) mares for determination of the number of primordial follicles. The ovary was separated from the oviduct and mesovarium, and sectioned into

0.8-cm transverse slices starting at the cranial pole of the ovary. All ovarian slices were submerged and stored in formalin until further processing.

5.2.2 Follicular parameters and dynamics

The following reproductive parameters were determined: inter-ovulatory interval, length of the luteal and follicular phase, day of luteolysis, day of follicle deviation, diameter of the dominant follicle at deviation, growth rate of the emerging and dominant follicle, diameter of the pre-ovulatory follicle two days (d-2) and one day (d-1) prior to ovulation

(day of ovulation = d0), number of major follicular waves per estrous cycle, number of diestrous ovulations and number of estrous ovulations. The inter-ovulatory interval was the time between two consecutive ovulations each associated with a different estrous period [143]. The length of the luteal and follicular phase was determined by calculating the number of days from ovulation to luteolysis, and between luteolysis and subsequent ovulation, respectively. The last day of the estrous cycle with a progesterone

concentration greater than 1 ng/mL was assigned as the day when luteolysis occurred

[167]. The day of deviation was determined as described by Gastal et al. [168]. The growth rate of the dominant follicle was determined for two different time periods: from deviation until two days prior to ovulation (dev. to d-2), and from two days prior to ovulation to one day prior to ovulation (d-2 to d-1). The growth rate of dominant follicle

(from dev. to d-2) was calculated by dividing the difference between the diameter of the pre-ovulatory follicle two days prior to ovulation and the diameter of dominant follicle at deviation by the number of days between those two time points. The growth rate of the emerging follicle was determined by calculating the mean growth of the largest growing follicle from the time when follicular growth was identified until deviation. A major follicular wave was identified when the largest growing follicle reached a diameter of at least 28 mm [143].

5.2.3 Determination of plasma progesterone and FSH concentrations

Plasma progesterone concentrations were determined in all mares on day 0, 2, 4, 6, 8, 10, and 12 and daily around the anticipated time of luteolysis during both estrous cycles.

Concentrations of progesterone were measured in duplicate using an enzyme-linked immunosorbent assay, as described and validated by Munro et al. [169]. The limit of sensitivity of the progesterone assay was 0.12 ng/mL. The intra- and inter-assay coefficient of variation for progesterone concentrations was 7.5%, and 13.1%, respectively.

Plasma FSH concentrations were measured on alternating days in a subset of young

(n=4), middle-aged (n=4), and aged (n=6) mares with a single major ovulatory wave during one estrous cycle. Plasma FSH concentrations were determined using a radioimmunoassay as described in detail by Yoon et al. [170]. Data were normalized to the mean inter-ovulatory interval of all mares using a method described by Carnevale et al.[156]. The sensitivity of the FSH assay was 0.5 ng/mL. The intra- and inter-assay coefficient of variation was 2.6 and 6.5 %, respectively.

5.2.4 Determination of primordial follicle counts

Determination of primordial follicle counts was performed by selecting the middle section of each ovary for sampling. From this section a one-by-one cm tissue section was removed from the center of the ovarian section and embedded in paraffin. Paraffin embedded tissues were sectioned (8 µm) using a rotary microtome, and each 40th section was mounted on a microscope slide. All slides were stained with hematoxylin and eosin.

Bright field microscopy (200 x magnification) was used to examine stained slides for the presence of primordial follicles which were only counted if the nucleus of the oocyte was present in the section.

5.3 Statistical analysis

Three aged mares were excluded from the analysis because the inter-ovulatory interval was longer than 30 days (> 3 standard deviations away from the mean inter-ovulatory

interval). Furthermore, one middle-aged and one aged mare were excluded from the analysis because intra-uterine fluid was present during diestrus resulting in premature luteolysis. The influence of age on the inter-ovulatory interval, length of the luteal and follicular phase, as well as the day of luteolysis and deviation was examined using a mixed model with mare as a random effect and age group, number of major follicular waves, and estrous cycle number as fixed effects. A mixed model was also used to examine the effect of age group on the growth rate of the emerging follicle, the growth rate of dominant follicle, the diameter of the dominant follicle at deviation, and the diameter of pre-ovulatory follicle. Within different age groups, differences in the growth rate between the emerging follicle and dominant follicle, and differences in the growth rate of the dominant follicle with respect to the days prior to ovulation (from dev. to d-2 versus from d-2 to d-1) was examined during each estrous cycle using a paired t-test. A chi-square test was used to examine the effect of age on the number of major follicular waves and ovulations for each estrous cycle. The influence of AFC on the inter-ovulatory interval, length of the follicular phase, and day of deviation was determined using a mixed model with mare as random effect and AFC group, age, the interaction of age by

AFC group, number of major follicular waves and estrous cycle number as fixed effects.

As a different estrous cycle without a diestrus ovulation from the same mare was available for each estrous cycle with a diestrus ovulation, a paired t-test was used to compare the inter-ovulatory interval, follicular and luteal phase duration, and the day of deviation between estrous cycles with and without a diestrous ovulation. A mixed model with estrous cycle day, number of corpora lutea, age group, and the interaction of estrous cycle day by age group as fixed effects and mare as random effect was used to evaluate

the influence of age group on plasma progesterone concentrations during both estrous cycles and a Fisher’s protected least-significant difference (LSD) used for post hoc analysis. The influence of age group on plasma FSH concentrations was determined using a mixed model with estrous cycle day, age group, and the interaction of estrous cycle day by age group as fixed effects and mare as random effect. A mixed model with estrous cycle day, follicular wave (diestrous ovulation versus no diestrous ovulation), and the interaction of estrous cycle day by follicular wave as fixed effects and mare as random effect was also used to compare progesterone concentrations between estrous cycles with and without a diestrous ovulation and Fisher’s protected LSD was used as post hoc test. Differences in primordial follicle counts in the ovarian section from young and aged mares were examined using the Wilcoxon signed rank test. All statistical analyses were performed using JMP version 10.0 (SAS Institute; Cary, North Carolina).

5.4 Results

5.4.1 Follicular parameters and dynamics

5.4.1.1 Effect of age on estrous cycle parameters

Inter-ovulatory intervals were significantly longer in aged mares (P<0.04) than in young mares (Table 5.1). Likewise, the length of the follicular phase was longer and the day of follicle deviation occurred later (P<0.02) in aged mares than in young or middle-aged mares while no significant differences were observed in the length of luteal phase between young, middle-aged or aged mares. The diameter of the dominant follicle at deviation was not significantly different in young, middle-aged or aged mares.

Prostaglandins were administrated to two middle-aged and five aged mares at day 22 post-ovulation because they did not return to estrus. Progesterone concentration in all those mares was greater than 1 ng/mL at the time of prostaglandinF2α administration.

Diestrous ovulations were associated with prolonged luteal phases (22 days) in the two middle-aged mares but diestrous ovulations did not occur in any of the five aged mares.

The number of major follicular waves per estrous cycle was not significantly associated with age group during the first estrous cycle; however, two follicular waves were less frequently observed in aged mares than in young or middle-aged (P<0.05) mares during the second estrous cycle. The number of ovulations and the growth rate of the emerging and dominant follicles (from dev. to d-2, and from d-2 to d-1) were not significantly different between young, middle-aged or aged mares. Within each age group, the growth rate of emerging follicles was not significantly different than the growth rate of the dominant follicle (dev. to d-2) whereas the growth rate of the dominant follicle (dev. to d-

2) was higher (P<0.05) than the growth rate of the dominant follicle on the last day prior to ovulation (d-2 to d-1). Finally, the diameter of the pre-ovulatory follicle had a tendency (P<0.08) to be smaller in aged mares than in middle-aged mares.

Table 5.1 Follicular parameters and estrous cycle characteristics in young, middle-aged and aged mares. Data are expressed as mean ± s.e.m. Different letters indicate statistical significant differences. Statistical significance was set at P<0.05.

Age group

Reproductive parameters Young Middle-aged Aged

(3-8 yrs) (9-18 yrs) (19-27 yrs)

Inter-ovulatory interval (days) 20.9 ± 0.5 a 21.6 ± 0.4 a,b 23.0 ± 0.8 b

Luteal phase (days) 15.6 ± 0.4 a 15.7 ± 0.3 a 16.4 ± 0.4 a

Follicular phase (days) 5.3 ± 0.3 a 5.8 ± 0.4 a 7.2 ± 0.5 b

Day of deviation 14.2 ± 0.6 a 13.1 ± 0.8 a 16.7 ± 0.7 b

Follicle size (mm) at deviation 24.5 ± 0.5 a 24.0 ± 0.7 a 23.7 ± 0.5 a

Growth rate (mm/day) 3.0 ± 0.2 a 3.0 ± 0.1 a 2.6 ± 0.2 a emerging follicle

Growth rate (mm/day)

dominant follicle 3.0 ± 0.2 a 2.8 ± 0.2 a 2.9 ± 0.2 a

(dev. to d-2)

Growth rate (mm/day)

dominant follicle 1.6 ± 0.3 a 0.8 ± 0.4 a 1.5 ± 0.4 a

(d-2 to d-1)

Diameter (mm) of the pre- 39.4 ± 0.7 a 40.6 ± 0.6 a 37.8 ± 0.7 a ovulatory follicle (d-1)

5.4.1.2 Effect of antral follicle count on age-related changes in estrous cycle parameters

Mare age (P<0.0004), the number of major follicular waves (P<0.007), and the interaction of age by AFC group (P<0.008) had an influence on the inter-ovulatory interval as well as the length of follicular phase but not on the day of deviation. As mare age increased, the increase in the inter-ovulatory interval and the length of the follicular phase was significantly associated with the number of antral follicles; mares with a low number of antral follicles had a longer inter-ovulatory interval and a longer follicular phase than mares with a medium or high number of antral follicles.

5.4.1.3 Effect of a diestrous ovulations on estrous cycle parameters

The overall incidence of a two-wave estrous cycle was 22.8 %. The incidence of a major secondary ovulatory wave (characterized by the occurrence of a diestrous ovulation)

(15.2%) was twice as common as major secondary anovulatory wave (7.6%). Irrespective of whether the secondary follicular wave is ovulatory or anovulatory, mares with a two- wave estrous cycle had a longer inter-ovulatory interval (P=0.003) as well as longer luteal

(P<0.04) and follicular phases (P<0.02) compared to mares with a one-wave estrous cycle. Furthermore, the day of deviation (P<0.005) occurred later in mares with a two- wave estrous cycle than in mares with a one-wave estrous cycle. Two-wave estrous cycles with a secondary ovulatory follicular wave (Table 5.2) had a longer inter- ovulatory interval (P<0.001) and follicular (P<0.02) and luteal phase (P<0.0001) while the day of deviation occurred later (P<0.0009).

Table 5.2 Follicular parameters of mares with a one-wave estrous cycle (no diestrous ovulation) and a two-wave estrous cycle with a secondary ovulatory follicular wave

(diestrous ovulation). Data are presented as mean ± s.e.m. Different letters indicate statistical significant differences (P<0.05)

Reproductive No diestrous ovulation Diestrous ovulation

parameter

Inter-ovulatory interval 19.3 ± 0.4 a 22.7 ± 0.5 b

(days)

Follicular phase (days) 4.2 ± 0.4 a 6.0 ± 0.6 b

Luteal phase (days) 15.1 ± 0.3 a 16.7 ± 0.5 b

Day of deviation (day) 10.2 ± 1.0 a 15.4 ± 0.5 b

5.4.2 Plasma progesterone and FSH concentrations

5.4.2.1 Effect of age

During the first estrous cycle, plasma progesterone concentrations were related to estrous cycle day (P<0.0001), number of corpora lutea (P<0.0001), and age group (P=0.08) while the interaction of estrous cycle day by age group was not significant. Mares with two corpora lutea had higher progesterone concentrations than mares with one corpus luteum.

Furthermore, plasma progesterone concentrations had a tendency to be higher in middle- aged mares than in young mares (Figure 5.1). During the second estrous cycle, plasma progesterone concentrations were significantly related to estrous cycle day (P<0.0001), number of corpora lutea (P<0.0001), age group (P=0.02), and the interaction of estrous

cycle day by age group (P=0.02). Plasma progesterone concentrations were significantly higher on day 6, 8, and day 10 in aged mares compared to young and middle-aged mares

(Figure 1). Although plasma FSH concentrations appeared to be higher in aged mares during the follicular phase (Figure 5.2), neither age group nor the interaction of age group by estrous cycle day was significant.

Figure 5.1 Plasma progesterone concentrations across two consecutive estrous cycles in young (dotted line), middle-aged (dashed line), and aged (solid line) mares. Plasma progesterone concentrations during the first estrous cycle are shown on the left, while the image on the right represents plasma progesterone concentrations during the second estrous cycle. Data are presented as mean ± s.e.m. Asterisks indicate statistical significant differences (P<0.05).

Figure 5.2 Mean plasma FSH concentrations during one estrous cycle in a subset of young (n=4), middle-aged (n=4), and aged mares (n=6). The solid, dashed, and dotted line represent old mares, middle-aged mares, and young mares, respectively. Data are presented as mean ± s.e.m.

5.4.2.2 Effect of a diestrous ovulation on peripheral progesterone concentration

Plasma progesterone concentrations were significantly associated with the day of the estrous cycle (P<0.0001) as well as the interaction of estrous cycle day by the number of follicular waves (P<0.0001) although the progesterone concentrations were not associated with the number of follicular waves alone. Plasma progesterone concentrations were

significantly higher on day 10, 12, 14, and 16 in cycles with a diestrous ovulation than in cycles without a diestrous ovulation (Figure 5.3).

Figure 5.3 Mean plasma progesterone concentrations in mares with (solid line) and without a diestrus ovulation. Data are presented as mean ± s.e.m. Asteriks indicate statistical significant differences (P<0.05)

5.4.3 Primordial follicle counts

Primordial follicles in ovarian tissue were characterized histologically by a single layer of flattened granulosa cells surrounding an oocyte. In several cases, a dark stained nucleolus can be detected within nucleus of oocyte (Figure 5.4). The number of primordial follicles varied widely within young (median: 227, range: 19-700) and aged mares (median: 0, range: 0-46), and the number of primordial follicles was higher (P<0.02) in young mares 92

than in aged mares. No primordial follicles were detected in 3 of the 4 aged mares even though all mares included in this study were cyclic mares.

Figure 5.4 A histological image of a primordial follicle in equine ovarian tissue. The primordial follicle consist of a single layer of flattened granulosa cells (arrowhead) surrounding a oocyte which contains a nucleus. The darkly stained nucleolus (arrow) is visible at the periphery of the nucleus.

5.5 Discussion and conclusion

To the authors’ knowledge, this is the first report to evaluate age-related changes in estrous cycle parameters in light horse mares in relationship with putative ovarian reserve based upon antral follicle counts. As noted in a number of other mammals, the number of primordial follicles not only decreases with age but also varies widely within the same age group [95, 171]. This difference likely reflects differences in ovarian reserve that appear to be established early in life and which ultimately affect reproductive aging variably across individuals.

In general, age-associated changes in estrous cycle parameters in light horse mares in the current study were similar to those reported previously in pony mares [111, 156, 157].

Prolonged inter-ovulatory intervals in aged mares were attributed to a longer follicular phase and not to changes in luteal phase length. The day of follicle deviation was later in older mares likely due to a delay in the emergence of the major primary follicular wave

[172]. The growth rate of the emerging or dominant follicle between young, middle-aged and aged mares was similar although aged mares had a tendency to ovulate smaller follicles than middle-aged mares. This observation is consistent with a clinical study showing that older Thoroughbred mares ovulate smaller follicles [173]. Although mean luteal phase length was not different across age groups, five aged and two middle-aged mares had a prolonged luteal phase. In two middle-aged mares, prolonged luteal phase was associated with a diestrous ovulation with subsequent maintenance of an immature corpus luteum. In aged mares, diestrous ovulations were not identified and the cause of prolonged maintenance of the CL was not apparent. Neely et al. demonstrated that

luteolysis can be compromised in mares with chronic endometritis resulting in a prolonged luteal phase [174], and this may have been the cause in the older mares in the current study.

Although there were clear age-related effects on estrous cycle parameters, these data also demonstrate an effect of AFC on follicular dynamics. As mare age increased, mares with low AFC had longer inter-ovulatory intervals and longer follicular phases than mares with medium or high AFC. This observation suggests that follicular reserve in older mares may influence changes in follicular phase parameters and argues that reproductive age and calendar age are distinct variables in the mare. Differences between reproductive and calendar age are well described in women [98, 161]. The age at which menopause varies widely among women [175]. This is thought to be the result of inherent differences in the number of primordial follicles at birth as well as variations in the rate of follicular depletion throughout life [98]. One of determining factors of ovarian senescence in aged broodmares is the remaining number of primordial follicles (follicular reserve) in the ovary. The lower number of primordial follicles in the ovarian tissue of aged mares indicates that the ovarian reserve of mares decreases with mare age, and supports the results of a recent study demonstrating that primordial follicle counts decreases with mare age [176]. Irrespective of the age-related decline in primordial follicles, the number of primordial follicles varied widely between mares of the same age group. This wide range in the number of primordial follicles is in accordance with an earlier study by Driancourt et al. [112] and suggests the presence of inherent differences in the population of primordial follicles among mares. Consequently,

this finding could potentially explain differences in mare age at onset of ovarian senescence and highlights the necessity of differentiating the reproductive age of a mare from its chronological age.

Based upon the methodology used in the current study, it was not possible to provide a robust estimate for the number of primordial follicles remaining in the ovary of mares across different age groups. The use of an ovarian biopsy, as in the current study, has been proposed as a method to estimate follicular reserve in mares [177] however, the methodology employed proved difficult to reliably detect primordial follicles in the ovary from older mares. The distribution of primordial follicles in the ovary of older mares has not been characterized and available sampling techniques would require that the entire ovary be examined in order to provide a reliable estimate of follicular reserve in the older mare. A valuable alternative to determine the size of the follicular pool is stereology

[178], a technique that employs systemic random sampling in which each part of ovary has an equal chance to be included. Still, the usefulness of this technique in counting primordial follicles has not been described in the light horse mares.

Besides age-related differences in the number of primordial follicles, reproductive aging in light horse mares was also associated with changes in luteal activity Differences in progesterone concentrations were detected between young, middle-aged and aged mares in contrast to a study in pony mares [156]. Concentrations of progesterone were significantly higher in aged mares during the second estrous cycle. Vanderwall et al. also

measured progesterone concentrations during two consecutive estrous cycles in young and aged mares and, likewise, found that aged mares had higher progesterone concentrations during the latter part of diestrus during one of two estrous cycles [179].

However, without knowing regulatory factors that influence progesterone production in the mare, it is difficult to explain why aged mares have higher progesterone concentrations.

In contrast to peripheral progesterone concentrations, circulating FSH concentrations were not significantly influenced by age even though FSH concentrations appeared higher in aged mares during the follicular phase. Studies in pony mares have shown that increased FSH concentrations are primarily observed in mares with an inter-ovulatory interval close to or longer than 30 days or in mares that have reached reproductive senescence when estrous cycles have ceased [103]. Mares with more advanced signs of ovarian senescence were excluded from our study, and this could explain why FSH concentrations were not significantly higher in the older age group. Based on the FSH data in aged pony mares and the association between longer inter-ovulatory intervals and low AFC in older mares detected in our study, we hypothesize that older mares with low

AFC have significantly higher FSH concentrations than older mares with medium or high

AFC. In fact, an inverse relationship between AFC and circulating FSH concentrations in cattle is supportive of this hypothesis [180]. Thus, taking all the data into account, monitoring changes in follicular parameters might be a more sensitive method to detect early ovarian senescence than measuring circulating FSH concentrations.

Finally, the occurrence of a major secondary follicular wave, particularly if accompanied by a diestrous ovulation, significantly increased the length of both the follicular and luteal phase of the estrous cycle in this study. The longer luteal phase in estrous cycles with a diestrous ovulation might indicate that the process of luteolysis is delayed. Although the overall progesterone concentration was not significantly different between estrous cycles with and without a diestrous ovulation, progesterone concentrations were significantly higher immediately prior to and around the expected time of luteolysis. As a diestrous ovulation usually occurs during the early to mid-luteal phase, it will take several days before the progesterone production from this immature corpus is markedly increased.

Therefore, it is not surprisingly that progesterone concentrations are only higher during the latter part of the luteal phase in estrous cycles with a diestrous ovulation.

In conclusion, changes in follicular parameters in light horse mares with reproductive aging are, to a large extent, comparable with changes noted previously in pony mares.

This study is the first to note, however, that antral follicle count is closely linked to age- related changes in follicular parameters. In addition to the wide variation in the number of primordial follicles between mares of the same age, a decrease in the ovarian reserve is present in older mares. As changes in follicular parameters occur prior to changes in the endocrine profiles, monitoring follicular parameters might be a more sensitive method to detect early stages of ovarian senescence than measuring circulating hormone concentrations. Finally, estrous cycles with diestrous ovulations are characterized by a longer inter-ovulatory interval in which both the luteal and follicular phase are prolonged.

5.6 Acknowledgments

This experiment was supported by the Albert G. Clay Endowment of the University of

Kentucky. The authors’ would like to thank Jessalyn Bridges, Carleigh Fedorka, Dr.

Alejandro Esteller-Vico, and Lillian E. Sybley for their help with either the clinical or laboratory part of this study.

Chapter Six: General conclusions

Research presented in this dissertation examined physiological variations and clinical applications of anti-Mullerian hormone (AMH) in the stallion and mare. In addition, molecular and hormonal changes in the equine follicle with regards to antral follicle count (AFC) and follicular development as well as age-related changes in follicular parameters and ovarian reserve were evaluated.

The methodological and biological validation of a commercially available human AMH assay for the horse was based upon dilutional parallelism and the disappearance of AMH after castration. Validation of this heterologous assay was an important step to reliably assess variations in serum AMH concentrations in male horses with regards to pubertal development, season, and gonadal status. As in humans, concentrations of AMH were higher in pre-pubertal colts than in post-pubertal stallions. This finding suggests that

AMH declines during puberty but the underlying mechanism of this decline was not determined. Although AMH concentrations were higher prior to puberty, concentrations of AMH remain relative high in intact mature stallions compared to humans. This unique feature could indicate that AMH plays an important role in testicular function or steroidogenesis in the postpubertal stallion. The seasonal variations in AMH in intact mature stallions support this hypothesis. Moreover, an increase in AMH concentrations during the breeding season likely coincides with an increased production of spermatozoa.

Therefore, concentrations of AMH could be a reflection of either Sertoli cell number or function. Nonetheless, whether AMH can be used to monitor the process of spermatogenesis in intact, mature stallions still remains to be determined. On the other

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hand, the usefulness of AMH as biomarker for the presence of testicular tissue was clearly documented. Diagnosis of cryptorchidism can be a challenge in equine practice especially after a questionable cryptorchid castration or in case of inconclusive endocrine test result. Therefore, the availability of an additional reliable marker for cryptorchidism can be useful for equine practitioners. As for any diagnostic assay, knowing the sensitivity and specificity of the assay is imperative. Unfortunately, although sensitivity could be determined in the current study, we were unable to establish conclusive information concerning specificity. Nonetheless, the Sertoli cell specific origin of AMH as well as the higher concentrations of AMH in cryptorchid stallions than intact stallions suggests that AMH is particularly valuable as diagnostic marker for cryptorchidism.

The clinical utility of AMH was not limited to diagnosing cryptorchidism in stallions.

Anti-Müllerian hormone also appears to be a useful biomarker for ovarian reserve in middle-aged and aged mares as indicated by the moderate to strong correlation between

AMH and AFC. The wide variation in AMH between mares of the same age likely attributes to differences in follicular reserve between mares while the lower concentrations of AMH in aged mares support an age-related decline in ovarian reserve.

Furthermore, the high repeatability of AMH within and across estrous cycles renders

AMH suitable as biomarker for ovarian reserve. On the other hand, the inability to detect a significant relationship between AFC and AMH in young mares was unexpected. It is unclear if small sample size could have influenced the relationship between AFC and

AMH in young mares or if this represents a biological difference between young and middle-aged or aged mares. To date, AMH has been compared to AFC which represents

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the total number of follicles detectable using ultrasonography. Still, it is likely that a large proportion of follicles, more specifically atretic follicles, do not significantly contribute to circulating AMH concentrations. Therefore, we hypothesize that circulating AMH concentrations are also a reflection of the number of follicles recruited during each follicular wave. However, without aspirating all antral follicles, it would be difficult to test this hypothesis as newly recruited follicles are overlapped by larger, regressing follicles. Irrespective, AMH appears to be a useful predictor of follicular reserve in middle-aged and aged mares.

Variations in AFC and circulating AMH concentrations in mares are not only a reflection of ovarian reserve but also appear to be related to follicular function as distinct molecular differences were detected in growing follicles of mares with low AFC or low AMH concentrations. Studies in other species have linked those molecular differences to impaired ovarian function. More specifically, it has been shown that, in association with a reduced expression of FSHR, the ovarian response to FSH is reduced in cows with low

AFC, and the lower expression of AMH is thought to be concomitant to a reduced oocyte quality. In view of these data, the possibility exists that follicular function and oocyte quality are impaired in mares with low AFC or low AMH.

Besides the relationship between AFC and molecular differences in the equine follicle, follicular development was also associated with molecular and hormonal changes in the equine follicle. The expression of AMH and a number of genes co-expressed with AMH, known to play a role in the regulation of AMH, decreased during follicular development.

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In accordance with molecular changes during follicular development, the synthesis of

AMH in the equine follicle also changed in relation to the stage of follicular development.

Circulating AMH concentrations were primarily associated with the numbers of antral follicles between 6 and 20 mm, suggesting that the secretion of AMH appears to subside around the time of follicle deviation. Indeed, concentrations of AMH in follicular fluid were high immediately prior to follicle selection and decreased to low concentrations in dominant follicles. So, it appears that follicular fluid AMH concentrations decline rather rapidly after follicle deviation. This sharp decline in AMH could indicate that AMH plays a role in follicle deviation in mares.

Finally, throughout the several different experiments, a large amount of evidence was gathered indicating the reproductive age of a mare must be distinguished from the chronological age. The large variation in AMH and AFC between old mares suggests that there are considerable differences in follicular populations between old mares. Likewise, the number of primordial follicles varied widely between mares of the same age which also implies that there are inherent differences in the size of the follicular pool between mares. Lastly, age-related changes in follicular parameters were closely linked to differences in AFC. Taking all of these data into consideration, we conclude that the variability in reproductive aging of mares is attributed to individual differences in follicular populations. Therefore, measuring the concentration of AMH as well as determining the number of antral follicle follicles should be an integral part in the reproductive examination of older mares as it provides valuable information about the

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process of reproductive aging and could help veterinarians to predict the reproductive lifespan of mares.

In conclusion, AMH can serve as a biomarker for cryptorchidism in stallions and ovarian reserve in mares. In addition, impaired ovarian function as well as age-related changes in follicular parameters in mares is linked to a low number of antral follicles.

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References

[1] Jost A. Recherches sur la differentiation sexuelle de l’embryon de lapin. III. Role des gonades foetales dans la differenciation sexuelle somatique. Arch Anat Microsc Morph Exp. 1947;36:271-315. [2] Jost A. Problems of fetal endocrinology: the gonadal and hypophyseal hormones. Recent progress in hormone research. 1953;8:379-418. [3] Josso N, Cate RL, Picard JY, Vigier B, di Clemente N, Wilson C, et al. Anti- mullerian hormone: the Jost factor. Recent progress in hormone research. 1993;48:1-59. [4] Cate RL, Mattaliano RJ, Hession C, Tizard R, Farber NM, Cheung A, et al. Isolation of the bovine and human genes for Mullerian inhibiting substance and expression of the human gene in animal cells. Cell. 1986;45:685-98. [5] Josso N, Clemente N. Transduction pathway of anti-Mullerian hormone, a sex- specific member of the TGF-beta family. Trends in endocrinology and metabolism. 2003;14:91-7. [6] Cohen-Haguenauer O, Picard JY, Mattei MG, Serero S, Nguyen VC, de Tand MF, et al. Mapping of the gene for anti-mullerian hormone to the short arm of human chromosome 19. Cytogenetics and cell genetics. 1987;44:2-6. [7] King TR, Lee BK, Behringer RR, Eicher EM. Mapping anti-mullerian hormone (Amh) and related sequences in the mouse: identification of a new region of homology between MMU10 and HSA19p. Genomics. 1991;11:273-83. [8] Gao Q, Womack JE. A genetic map of bovine chromosome 7 with an interspecific hybrid backcross panel. Mammalian genome. 1997;8:258-61. [9] Knebelmann B, Boussin L, Guerrier D, Legeai L, Kahn A, Josso N, et al. Anti- Mullerian hormone Bruxelles: a nonsense mutation associated with the persistent Mullerian duct syndrome. Proceedings of the National Academy of Sciences of the United States of America. 1991;88:3767-71. [10] Josso N, Belville C, di Clemente N, Picard JY. AMH and AMH receptor defects in persistent Mullerian duct syndrome. Human reproduction update. 2005;11:351-6. [11] Josso N. Permeability of membranes to the Müllerian-Inhibiting Substance Synthesized by the human fetal testis in vitro: A clue to its biochemical nature. The Journal of Clinical Endocrinology & Metabolism. 1972;34:265-70. [12] Budzik GP, Swann DA, Hayashi A, Donahoe PK. Enhanced purification of Mullerian Inhibiting Substance by lectin affinity-chromatography. Cell. 1980;21:909-15. [13] Picard JY, Tran D, Josso N. Biosynthesis of labeled anti-Mullerian hormone by fetal testes - Evidence for glycoprotein nature of hormone and for its disulfide-bonded structure. Molecular and cellular endocrinology. 1978;12:17-30. [14] Picard JY, Josso N. Purification of testicular anti-Mullerian hormone allowing direct visualization of the pure glycoprotein and determination of yield and purification factor. Molecular and cellular endocrinology. 1984;34:23-9. [15] Picard JY, Goulut C, Bourrillon R, Josso N. Biochemical-analysis of bovine testicular anti-Mullerian hormone. Febs Lett. 1986;195:73-6. [16] Pepinsky RB, Sinclair LK, Chow EP, Mattaliano RJ, Manganaro TF, Donahoe PK, et al. Proteolytic processing of mullerian inhibiting substance produces a transforming growth factor-beta-like fragment. The Journal of biological chemistry. 1988;263:18961-4.

105

[17] MacLaughlin DT, Hudson PL, Graciano AL, Kenneally MK, Ragin RC, Manganaro TF, et al. Mullerian duct regression and antiproliferative bioactivities of mullerian inhibiting substance reside in its carboxy-terminal domain. Endocrinology.1992;131: 291-6. [18] Wilson CA, di Clemente N, Ehrenfels C, Pepinsky RB, Josso N, Vigier B, et al. Mullerian inhibiting substance requires its N-terminal domain for maintenance of biological activity, a novel finding within the transforming growth factor-beta superfamily. Mol Endocrinol. 1993;7:247-57. [19] Ball BA, Conley AJ, Grundy SA, Sabeur K, Liu IK. Expression of anti-Mullerian hormone (AMH) in the equine testis. Theriogenology. 2008;69:624-31. [20] Tran D, Picard JY, Campargue J, Josso N. Immunocytochemical detection of anti- mullerian hormone in Sertoli cells of various mammalian species including human. The journal of histochemistry and cytochemistry : official journal of the Histochemistry Society. 1987;35:733-43. [21] Ball BA, Conley AJ, MacLaughlin DT, Grundy SA, Sabeur K, Liu IK. Expression of anti-Mullerian hormone (AMH) in equine granulosa-cell tumors and in normal equine ovaries. Theriogenology. 2008;70:968-77. [22] Rey R, Lukas-Croisier C, Lasala C, Bedecarras P. AMH/MIS: what we know already about the gene, the protein and its regulation. Molecular and cellular endocrinology. 2003;211:21-31. [23] Schwindt B, Doyle LW, Hutson JM. Serum levels of mullerian inhibiting substance in preterm and term male neonates. The Journal of urology. 1997;158:610-2. [24] Josso N, Lamarre I, Picard JY, Berta P, Davies N, Morichon N, et al. Anti-mullerian hormone in early human development. Early human development. 1993;33:91-9. [25] Aksglaede L, Sorensen K, Boas M, Mouritsen A, Hagen CP, Jensen RB, et al. Changes in anti-Mullerian hormone (AMH) throughout the life span: a population-based study of 1027 healthy males from birth (cord blood) to the age of 69 years. The Journal of clinical endocrinology and metabolism. 2010;95:5357-64. [26] Baker ML, Metcalfe SA, Hutson JM. Serum levels of mullerian inhibiting substance in boys from birth to 18 years, as determined by enzyme immunoassay. The Journal of clinical endocrinology and metabolism. 1990;70:11-5. [27] Baker ML, Hutson JM. Serum levels of mullerian inhibiting substance in boys throughout puberty and in the first two years of life. The Journal of clinical endocrinology and metabolism. 1993;76:245-7. [28] Pietilainen KH, Kaprio J, Vaaralahti K, Rissanen A, Raivio T. Circulating anti- Mullerian hormone levels in adult men are under a strong genetic influence. The Journal of clinical endocrinology and metabolism. 2012;97:E161-4. [29] Rajpert-De Meyts E, Jorgensen N, Graem N, Muller J, Cate RL, Skakkebaek NE. Expression of anti-Mullerian hormone during normal and pathological gonadal development: association with differentiation of Sertoli and granulosa cells. The Journal of clinical endocrinology and metabolism. 1999;84:3836-44. [30] Guibourdenche J, Lucidarme N, Chevenne D, Rigal O, Nicolas M, Luton D, et al. Anti-Mullerian hormone levels in serum from human foetuses and children: pattern and clinical interest. Molecular and cellular endocrinology. 2003;211:55-63.

106

[31] Hagen CP, Aksglaede L, Sorensen K, Main KM, Boas M, Cleemann L, et al. Serum levels of anti-Mullerian hormone as a marker of ovarian function in 926 healthy females from birth to adulthood and in 172 Turner syndrome patients. The Journal of clinical endocrinology and metabolism. 2010;95:5003-10. [32] Kelsey TW, Wright P, Nelson SM, Anderson RA, Wallace WH. A validated model of serum anti-mullerian hormone from conception to menopause. PloS one. 2011;6:e22024. [33] Chellakooty M, Schmidt IM, Haavisto AM, Boisen KA, Damgaard IN, Mau C, et al. Inhibin A, inhibin B, follicle-stimulating hormone, luteinizing hormone, estradiol, and sex hormone-binding globulin levels in 473 healthy infant girls. The Journal of clinical endocrinology and metabolism. 2003;88:3515-20. [34] Tehrani FR, Shakeri N, Solaymani-Dodaran M, Azizi F. Predicting age at menopause from serum antimullerian hormone concentration. Menopause . 2011;18: 766-70. [35] van Rooij IA, Tonkelaar I, Broekmans FJ, Looman CW, Scheffer GJ, de Jong FH, et al. Anti-mullerian hormone is a promising predictor for the occurrence of the menopausal transition. Menopause . 2004;11:601-6. [36] Shen WH, Moore CC, Ikeda Y, Parker KL, Ingraham HA. Nuclear receptor steroidogenic factor 1 regulates the mullerian inhibiting substance gene: a link to the sex determination cascade. Cell. 1994;77:651-61. [37] De Santa Barbara P, Bonneaud N, Boizet B, Desclozeaux M, Moniot B, Sudbeck P, et al. Direct interaction of SRY-related protein SOX9 and steroidogenic factor 1 regulates transcription of the human anti-Mullerian hormone gene. Molecular and cellular biology. 1998;18:6653-65. [38] Viger RS, Mertineit C, Trasler JM, Nemer M. Transcription factor GATA-4 is expressed in a sexually dimorphic pattern during mouse gonadal development and is a potent activator of the Mullerian inhibiting substance promoter. Development. 1998;125:2665-75. [39] Nachtigal MW, Hirokawa Y, Enyeart-VanHouten DL, Flanagan JN, Hammer GD, Ingraham HA. Wilms' tumor 1 and Dax-1 modulate the orphan nuclear receptor SF-1 in sex-specific gene expression. Cell. 1998;93:445-54. [40] Fenichel P, Rey R, Poggioli S, Donzeau M, Chevallier D, Pointis G. Anti-Mullerian hormone as a seminal marker for spermatogenesis in non-obstructive azoospermia. Hum Reprod. 1999;14:2020-4. [41] Fujisawa M, Yamasaki T, Okada H, Kamidono S. The significance of anti-Mullerian hormone concentration in seminal plasma for spermatogenesis. Hum Reprod. 2002;17:968-70. [42] Rey R, Lordereau-Richard I, Carel JC, Barbet P, Cate RL, Roger M, et al. Anti- mullerian hormone and testosterone serum levels are inversely during normal and precocious pubertal development. The Journal of clinical endocrinology and metabolism. 1993;77:1220-6. [43] Boukari K, Meduri G, Brailly-Tabard S, Guibourdenche J, Ciampi ML, Massin N, et al. Lack of androgen receptor expression in Sertoli cells accounts for the absence of anti- Mullerian hormone repression during early human testis development. The Journal of clinical endocrinology and metabolism. 2009;94:1818-25.

107

[44] Rey RA, Musse M, Venara M, Chemes HE. Ontogeny of the androgen receptor expression in the fetal and postnatal testis: its relevance on Sertoli cell maturation and the onset of adult spermatogenesis. Microscopy research and technique. 2009;72:787-95. [45] Rey R, AlAttar L, Louis F, Jaubert F, Barbet P, NihoulFekete C, et al. Testicular dysgenesis does not affect expression of anti-Mullerian hormone by Sertoli cells in premeiotic seminiferous tubules. Am J Pathol. 1996;148:1689-98. [46] Hirobe S, He WW, Lee MM, Donahoe PK. Mullerian inhibiting substance messenger ribonucleic acid expression in granulosa and Sertoli cells coincides with their mitotic activity. Endocrinology. 1992;131:854-62. [47] Jeppesen JV, Anderson RA, Kelsey TW, Christiansen SL, Kristensen SG, Jayaprakasan K, et al. Which follicles make the most anti-Mllerian hormone in humans? Evidence for an abrupt decline in AMH production at the time of follicle selection. Molecular human reproduction. 2013;19:519-27. [48] Ogura-Nose S, Yoshino O, Osuga Y, Shi J, Hiroi H, Yano T, et al. Anti-Mullerian hormone (AMH) is induced by bone morphogenetic protein (BMP) cytokines in human granulosa cells. European journal of obstetrics, gynecology, and reproductive biology. 2012;164:44-7. [49] Rico C, Medigue C, Fabre S, Jarrier P, Bontoux M, Clement F, et al. Regulation of anti-Mullerian hormone production in the cow: a multiscale study at endocrine, ovarian, follicular, and granulosa cell levels. Biology of reproduction. 2011;84:560-71. [50] Taieb J, Grynberg M, Pierre A, Arouche N, Massart P, Belville C, et al. FSH and its second messenger cAMP stimulate the transcription of human anti-Mullerian hormone in cultured granulosa cells. Mol Endocrinol. 2011;25:645-55. [51] Grossman MP, Nakajima ST, Fallat ME, Siow Y. Mullerian-inhibiting substance inhibits cytochrome P450 aromatase activity in human granulosa lutein cell culture. Fertility and sterility. 2008;89:1364-70. [52] Di clemente N, Goxe B, Remy JJ, Cate R, Josso N, Vigier B, et al. Inhibitory effect of AMH upon the expression of aromatase and LH receptors by cultured granulosa cells of rat and porcine immature ovaries. Endocrine. 1994;2:553-8. [53] Grynberg M, Pierre A, Rey R, Leclerc A, Arouche N, Hesters L, et al. Differential regulation of ovarian anti-Mullerian hormone (AMH) by estradiol through alpha- and beta-estrogen receptors. J Clin Endocr Metab. 2012;97:E1649-E57. [54] Donahoe PK, Ito Y, Hendren WH. Graded organ-culture assay for detection of Mullerian Inhibiting Substance. J Surg Res. 1977;23:141-8. [55] Diclemente N, Ghaffari S, Pepinsky RB, Pieau C, Josso N, Cate RL, et al. A Quantitative and interspecific test for biological-activity of anti-Mullerian hormone - the fetal ovary aromatase assay. Development. 1992;114:721-7. [56] Picon R. Action of rat fetal testis on development of Mullerian ducts in vitro. Arch Anat Microsc Mo. 1969;58:1-&. [57] Josso N, Picard JY, Rey R, di Clemente N. Testicular anti-Mullerian hormone: history, genetics, regulation and clinical applications. Pediatric endocrinology reviews : PER. 2006;3:347-58. [58] Nef S, Parada LF. Hormones in male sexual development. Gene Dev. 2000;14: 3075-86. [59] Jenny A V. AMH signaling: from receptor to target gene. Molecular and cellular endocrinology. 2003;211:65-73.

108

[60] Meyerswallen VN, Manganaro TF, Kuroda T, Concannon PW, Maclaughlin DT, Donahoe PK. The critical period for Mullerian duct regression in the dog embryo. Biology of reproduction. 1991;45:626-33. [61] Taguchi O, Cunha GR, Lawrence WD, Robboy SJ. Timing and irreversibility of Mullerian duct inhibition in the embryonic reproductive tract of the human male. Developmental biology. 1984;106:394-8. [62] Durlinger ALL, Kramer P, Karels B, de Jong FH, Uilenbroek JTJ, Grootegoed JA, et al. Control of primordial follicle recruitment by anti-Mullerian hormone in the mouse ovary. Endocrinology. 1999;140:5789-96. [63] Durlinger ALL, Visser JA, Themmen APN. Regulation of ovarian function: the role of anti-Mullerian hormone. Reproduction. 2002;124:601-9. [64] Durlinger AL, Gruijters MJ, Kramer P, Karels B, Ingraham HA, Nachtigal MW, et al. Anti-Mullerian hormone inhibits initiation of primordial follicle growth in the mouse ovary. Endocrinology. 2002;143:1076-84. [65] Durlinger ALL, Gruijters MJG, Kramer P, Karels B, Kumar TR, Matzuk MM, et al. Anti-Mullerian hormone attenuates the effects of FSH on follicle development in the mouse ovary. Endocrinology. 2001;142:4891-9. [66] Pellatt L, Rice S, Dilaver N, Heshri A, Galea R, Brincat M, et al. Anti-Mullerian hormone reduces follicle sensitivity to follicle-stimulating hormone in human granulosa cells. Fertility and sterility. 2011;96:1246-51. [67] Behringer RR, Finegold MJ, Cate RL. Mullerian-inhibiting substance function during mammalian sexual development. Cell. 1994;79:415-25. [68] Racine C, Rey R, Forest MG, Louis F, Ferre A, Huhtaniemi I, et al. Receptors for anti-mullerian hormone on Leydig cells are responsible for its effects on steroidogenesis and cell differentiation. Proceedings of the National Academy of Sciences of the United States of America. 1998;95:594-9. [69] Rouiller-Fabre V, Carmona S, Merhi RA, Cate R, Habert R, Vigier B. Effect of anti- Mullerian hormone on Sertoli and Leydig cell functions in fetal and immature rats. Endocrinology. 1998;139:1213-20. [70] Lee MM, Donahoe PK, Hasegawa T, Silverman B, Crist GB, Best S, et al. Mullerian inhibiting substance in humans: normal levels from infancy to adulthood. The Journal of clinical endocrinology and metabolism. 1996;81:571-6. [71] Almeida J, Conley AJ, Mathewson L, Ball BA. Expression of anti-Müllerian hormone, cyclin-dependent kinase inhibitor (CDKN1B), androgen receptor, and connexin 43 in equine testes during puberty. Theriogenology. 2012;77:847-57. [72] Taya K, Nagata S, Tsunoda N, Nagamine N, Tanaka Y, Nagaoka K, et al. Testicular secretion of inhibin in stallions. Journal of reproduction and fertility Supplement. 2000;56:43-50. [73] Kabithe EW, Place NJ. Photoperiod-dependent modulation of anti-Mullerian hormone in female Siberian hamsters, Phodopus sungorus. Reproduction. 2008;135: 335-42. [74] Lee MM, Donahoe PK, Silverman BL, Hasegawa T, Hasegawa Y, Gustafson ML, et al. Measurements of serum mullerian inhibiting substance in the evaluation of children with nonpalpable gonads. The New England journal of medicine. 1997;336:1480-6.

109

[75] Lee MM, Misra M, Donahoe PK, MacLaughlin DT. MIS/AMH in the assessment of cryptorchidism and intersex conditions. Molecular and cellular endocrinology. 2003;211:91-8. [76] Misra M, MacLaughlin DT, Donahoe PK, Lee MM. Measurement of Mullerian inhibiting substance facilitates management of boys with microphallus and cryptorchidism. The Journal of clinical endocrinology and metabolism. 2002;87: 3598-602. [77] Stabenfeldt GH, Hughes JP, Kennedy PC, Meagher DM, Neely DP. Clinical findings, pathological changes and endocrinological secretory patterns in mares with ovarian tumours. Journal of reproduction and fertility Supplement. 1979:277-85. [78] Rota A, Ballarin C, Vigier B, Cozzi B, Rey R. Age dependent changes in plasma anti-Mullerian hormone concentrations in the bovine male, female, and freemartin from birth to puberty: relationship between testosterone production and influence on sex differentiation. General and comparative endocrinology. 2002;129:39-44. [79] Hudson PL, Dougas I, Donahoe PK, Cate RL, Epstein J, Pepinsky RB, et al. An immunoassay to detect human mullerian inhibiting substance in males and females during normal development. The Journal of clinical endocrinology and metabolism. 1990;70:16-22. [80] Vigier B, Tran D, du Mesnil du Buisson F, Heyman Y, Josso N. Use of monoclonal antibody techniques to study the ontogeny of bovine anti-Mullerian hormone. Journal of reproduction and fertility. 1983;69:207-14. [81] Cazorla O, Seck M, Pisselet C, Perreau C, Saumande J, Fontaine J, et al. Anti- Mullerian hormone (AMH) secretion in prepubertal and adult rams. Journal of reproduction and fertility. 1998;112:259-66. [82] Al-Attar L, Noel K, Dutertre M, Belville C, Forest MG, Burgoyne PS, et al. Hormonal and cellular regulation of Sertoli cell anti-Mullerian hormone production in the postnatal mouse. The Journal of clinical investigation. 1997;100:1335-43. [83] Clay CM, Clay JN. Endocrine and testicular changes associated with season, artificial photoperiod, and the peri-pubertal period in stallions. The Veterinary clinics of North America Equine practice. 1992;8:31-56. [84] Naden J, Amann RP, Squires EL. Testicular growth, hormone concentrations, seminal characteristics and sexual behaviour in stallions. Journal of reproduction and fertility. 1990;88:167-76. [85] Johnson L, Nguyen HB. Annual cycle of the Sertoli cell population in adult stallions. Journal of reproduction and fertility. 1986;76:311-6. [86] Johnson L, Varner DD, Tatum ME, Scrutchfield WL. Season but not age affects Sertoli cell number in adult stallions. Biology of reproduction. 1991;45:404-10. [87] Johnson L, Thompson DL, Jr. Age-related and seasonal variation in the Sertoli cell population, daily sperm production and serum concentrations of follicle-stimulating hormone, luteinizing hormone and testosterone in stallions. Biology of reproduction. 1983;29:777-89. [88] Johnson L, Tatum ME. Temporal appearance of seasonal changes in numbers of Sertoli cells, Leydig cells, and germ cells in stallions. Biology of reproduction. 1989;40:994-9.

110

[89] Altinsaat C, Uner AG, Sulu N, Ergun A. Seasonal variations in serum concentrations of melatonin, testosterone, and progesterone in Arabian horse. Ankara Univ Vet Fak. 2009;56:19-24. [90] Berndtson WE, Pickett BW, Nett TM. Reproductive physiology of the stallion. IV. Seasonal changes in the testosterone concentration of peripheral plasma. Journal of reproduction and fertility. 1974;39:115-8. [91] Raeside JI. Seasonal-Changes in the Concentration of Estrogens and Testosterone in the Plasma of the Stallion. Animal Reproduction Science. 1979;1:205-12. [92] Morganti M, Conley AJ, Vico AE, Munro CJ, Ball BA. Stallion-like behavior in mares: What is the role of the adrenal glands? In Proceedings, 56th Annual Convention of the American Association of Equine Practitioners 2010;56:314. [93] Almeida J, Conley AJ, Ball BA. Expression of anti-Müllerian hormone, CDKN1B, connexin 43, androgen receptor and steroidogenic enzymes in the equine cryptorchid testis. Equine veterinary journal. 2013;45:538-45 [94] Visser JA, Themmen AP. Anti-Mullerian hormone and folliculogenesis. Molecular and cellular endocrinology. 2005;234:81-6. [95] Ireland JL, Scheetz D, Jimenez-Krassel F, Themmen AP, Ward F, Lonergan P, et al. Antral follicle count reliably predicts number of morphologically healthy oocytes and follicles in ovaries of young adult cattle. Biology of reproduction. 2008;79: 1219-25. [96] Kevenaar ME, Meerasahib MF, Kramer P, van de Lang-Born BM, de Jong FH, Groome NP, et al. Serum anti-mullerian hormone levels reflect the size of the primordial follicle pool in mice. Endocrinology. 2006;147:3228-34. [97] Faddy MJ, Gosden RG. A model conforming the decline in follicle numbers to the age of menopause in women. Human Reproduction. 1996;11:1484-6. [98] Broekmans FJ, Knauff EAH, Velde ERT, Macklon NS, Fauser BC. Female reproductive ageing: current knowledge and future trends. Trends Endocrin Met. 2007;18:58-65. [99] Gold EB, Bromberger J, Crawford S, Samuels S, Greendale GA, Harlow SD, et al. Factors associated with age at natural menopause in a multiethnic sample of midlife women. American journal of epidemiology. 2001;153:865-74. [100] van Disseldorp J, Faddy MJ, Themmen AP, de Jong FH, Peeters PH, van der Schouw YT, et al. Relationship of serum antimullerian hormone concentration to age at menopause. The Journal of clinical endocrinology and metabolism. 2008;93:2129-34. [101] Almeida J, Ball BA, Conley AJ, Place NJ, Liu IK, Scholtz EL, et al. Biological and clinical significance of anti-Mullerian hormone determination in blood serum of the mare. Theriogenology. 2011;76:1393-403. [102] Vernunft A, Schneider F, Tuchscherer A, Becker F, Kanitz W. Anti-Muellerian Hormone (AMH) can help to predict follicular growth in mares. Pferdeheilkunde. 2011;27:293-6. [103] Carnevale EM, Bergfelt DR, Ginther OJ. Follicular activity and concentrations of FSH and LH associated with senescence in mares. Anim Reprod Sci. 1994;35:231-46. [104] Vanderwall DK, Woods GL. Age-related subfertility in the mare. Proc 36th Ann Conv Am Assoc Equine Pract. 1991:85-9. [105] Ginther OJ, Pierson RA. Ultrasonic anatomy of equine ovaries. Theriogenology. 1984;21:471-83.

111

[106] Nardo LG, Christodoulou D, Gould D, Roberts SA, Fitzgerald CT, Laing I. Anti- Mullerian hormone levels and antral follicle count in women enrolled in in vitro fertilization cycles: Relationship to lifestyle factors, chronological age and reproductive history. Gynecological Endocrinology. 2007;23:486-93. [107] Li HWR, Lee VCY, Lau EYL, Yeung WSB, Ho PC, Ng EHY. Role of Baseline Antral Follicle Count and Anti-Mullerian Hormone in Prediction of Cumulative Live Birth in the First In Vitro Fertilisation Cycle: A Retrospective Cohort Analysis. PloS one. 2013;8:e61095. [108] Hansen KR, Hodnett GM, Knowlton N, Craig LB. Correlation of ovarian reserve tests with histologically determined primordial follicle number. Fertility and sterility. 2011;95:170-5. [109] Ginther OJ, Gastal EL, Gastal MO, Bergfelt DR, Baerwald AR, Pierson RA. Comparative study of the dynamics of follicular waves in mares and women. Biology of reproduction. 2004;71:1195-201. [110] Ginther OJ, Beg MA, Donadeu FX, Bergfelt DR. Mechanism of follicle deviation in monovular farm species. Anim Reprod Sci. 2003;78:239-57. [111] Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Beg MA. Age-related dynamics of follicles and hormones during an induced ovulatory follicular wave in mares. Theriogenology. 2009;71:780-8. [112] Driancourt MA, Paris A, Roux C, Mariana JC, Palmer E. Ovarian follicular populations in pony and saddle-type mares. Reproduction, nutrition, development. 1982;22:1035-47. [113] Lahoz B, Alabart JL, Cocero MJ, Monniaux D, Echegoyen E, Sanchez P, et al. Anti-Mullerian hormone concentration in sheep and its dependence of age and independence of BMP15 genotype: an endocrine predictor to select the best donors for embryo biotechnologies. Theriogenology. 2014;81:347-57. [114] Hehenkamp WJ, Looman CW, Themmen AP, de Jong FH, Te Velde ER, Broekmans FJ. Anti-Mullerian hormone levels in the spontaneous menstrual cycle do not show substantial fluctuation. The Journal of clinical endocrinology and metabolism. 2006;91:4057-63. [115] Claes A, Ball BA, Almeida J, Corbin CJ, Conley AJ. Serum anti-Mullerian hormone concentrations in stallions: developmental changes, seasonal variation, and differences between intact stallions, cryptorchid stallions, and geldings. Theriogenology. 2013;79:1229-35. [116] Ireland JJ, Smith GW, Scheetz D, Jimenez-Krassel F, Folger JK, Ireland JL, et al. Does size matter in females? An overview of the impact of the high variation in the ovarian reserve on ovarian function and fertility, utility of anti-Mullerian hormone as a diagnostic marker for fertility and causes of variation in the ovarian reserve in cattle. Reproduction, fertility, and development. 2011;23:1-14. [117] Lahoz B, Alabart JL, Monniaux D, Mermillod P, Folch J. Anti-Mullerian hormone plasma concentration in prepubertal ewe lambs as a predictor of their fertility at a young age. BMC veterinary research. 2012;8:118. [118] Monniaux D, Baril G, Laine AL, Jarrier P, Poulin N, Cognie J, et al. Anti- Mullerian hormone as a predictive endocrine marker for embryo production in the goat. Reproduction. 2011;142:845-54.

112

[119] Rico C, Fabre S, Medigue C, di Clemente N, Clement F, Bontoux M, et al. Anti- mullerian hormone is an endocrine marker of ovarian gonadotropin-responsive follicles and can help to predict superovulatory responses in the cow. Biology of reproduction. 2009;80:50-9. [120] Monniaux D, Barbey S, Rico C, Fabre S, Gallard Y, Larroque H. Anti-Mullerian hormone: a predictive marker of embryo production in cattle? Reprod Fert Develop. 2010;22:1083-91. [121] Rico C, Drouilhet L, Salvetti P, Dalbies-Tran R, Jarrier P, Touze JL, et al. Determination of anti-Mullerian hormone concentrations in blood as a tool to select Holstein donor cows for embryo production: from the laboratory to the farm. Reprod Fert Develop. 2012;24:932-44. [122] Broer SL, Mol BW, Hendriks D, Broekmans FJ. The role of antimullerian hormone in prediction of outcome after IVF: comparison with the antral follicle count. Fertility and sterility. 2009;91:705-14. [123] Choi MH, Yoo JH, Kim HO, Cha SH, Park CW, Yang KM, et al. Serum anti- Mullerian hormone levels as a predictor of the ovarian response and IVF outcomes. Clinical and experimental reproductive medicine. 2011;38:153-8. [124] Wu CH, Chen YC, Wu HH, Yang JG, Chang YJ, Tsai HD. Serum anti-Mullerian hormone predicts ovarian response and cycle outcome in IVF patients. J Assist Reprod Gen. 2009;26:383-9. [125] Himabindu Y, Gopinathan KK, Pandey AK, Sriharibabu M. Correlation of age and antimullerian hormone in assisted reproductive technology program outcome. Indian journal of physiology and pharmacology. 2013;57:9-15. [126] Wang JG, Douglas NC, Nakhuda GS, Choi JM, Park SJ, Thornton MH, et al. The association between anti-Mullerian hormone and IVF pregnancy outcomes is influenced by age. Reproductive biomedicine online. 2010;21:757-61. [127] Claes A, Ball BA, Scoggin KE, Esteller-Vico A, Kalmar JJ, Conley AJ, et al. The interrelationship between anti-Müllerian hormone, ovarian follicular populations, and age in mares. Equine veterinary journal. 2014. [128] Scheetz D, Folger JK, Smith GW, Ireland JJ. Granulosa cells are refractory to FSH action in individuals with a low antral follicle count. Reproduction, Fertility and Development. 2011;24:327-36. [129] Mossa F, Jimenez-Krassel F, Folger JK, Ireland JL, Smith GW, Lonergan P, et al. Evidence that high variation in antral follicle count during follicular waves is linked to alterations in ovarian androgen production in cattle. Reproduction. 2010;140:713-20. [130] Ireland JJ, Zielak-Steciwko AE, Jimenez-Krassel F, Folger J, Bettegowda A, Scheetz D, et al. Variation in the ovarian reserve is linked to alterations in intrafollicular estradiol production and ovarian biomarkers of follicular differentiation and oocyte quality in cattle. Biology of reproduction. 2009;80:954-64. [131] Andersen CY, Schmidt KT, Kristensen SG, Rosendahl M, Byskov AG, Ernst E. Concentrations of AMH and inhibin-B in relation to follicular diameter in normal human small antral follicles. Human Reproduction. 2010;25:1282-7. [132] Nielsen ME, Rasmussen IA, Fukuda M, Westergaard LG, Andersen CY. Concentrations of anti-Mullerian hormone in fluid from small human antral follicles show a negative correlation with CYP19 mRNA expression in the corresponding granulosa cells. Molecular human reproduction. 2010;16:637-43.

113

[133] Evans AC, Ireland JL, Winn ME, Lonergan P, Smith GW, Coussens PM, et al. Identification of genes involved in apoptosis and dominant follicle development during follicular waves in cattle. Biology of reproduction. 2004;70:1475-84. [134] Kenney RM, Condon W, Ganjam VK, Channing C. Morphological and biochemical correlates of equine ovarian follicles as a function of their state of viability or atresia. Journal of reproduction and fertility Supplement. 1979:163-71. [135] Ruijter JM, Ramakers C, Hoogaars WM, Karlen Y, Bakker O, van den Hoff MJ, et al. Amplification efficiency: linking baseline and bias in the analysis of quantitative PCR data. Nucleic acids research. 2009;37:e45. [136] Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 2001;25:402-8. [137] Conley AJ, Corbin CJ, Hinshelwood MM, Liu Z, Simpson ER, Ford JJ, et al. Functional aromatase expression in porcine adrenal gland and testis. Biology of reproduction. 1996;54:497-505. [138] Walters KW, Corbin CJ, Anderson GB, Roser JF, Conley AJ. Tissue-specific localization of cytochrome P450 aromatase in the equine embryo by in situ hybridization and immunocytochemistry. Biology of reproduction. 2000;62:1141-5. [139] Mossa F, Walsh SW, Butler ST, Berry DP, Carter F, Lonergan P, et al. Low numbers of ovarian follicles ≥ 3 mm in diameter are associated with low fertility in dairy cows. Journal of dairy science. 2012;95:2355-61. [140] Ireland JJ, Ward F, Jimenez-Krassel F, Ireland JL, Smith GW, Lonergan P, et al. Follicle numbers are highly repeatable within individual animals but are inversely correlated with FSH concentrations and the proportion of good-quality embryos after ovarian stimulation in cattle. Hum Reprod. 2007;22:1687-95. [141] Luciano AM, Franciosi F, Lodde V, Tessaro I, Corbani D, Modina SC, et al. Oocytes isolated from dairy cows with reduced ovarian reserve have a high frequency of aneuploidy and alterations in the localization of progesterone receptor membrane component 1 and aurora kinase B. Biology of reproduction. 2013;88:58. [142] Cai J, Lou HY, Dong MY, Lu XE, Zhu YM, Gao HJ, et al. Poor ovarian response to gonadotropin stimulation is associated with low expression of follicle-stimulating hormone receptor in granulosa cells. Fertility and sterility. 2007;87:1350-6. [143] Ginther OJ, Beg MA, Gastal MO, Gastal EL. Follicle dynamics and selection in mares. Anim Reprod. 2004;1:45-63. [144] Jakimiuk AJ, Weitsman SR, Yen HW, Bogusiewicz M, Magoffin DA. Estrogen receptor alpha and beta expression in theca and granulosa cells from women with polycystic ovary syndrome. The Journal of clinical endocrinology and metabolism. 2002;87:5532-8. [145] Byers M, Kuiper GG, Gustafsson JA, Park-Sarge OK. Estrogen receptor-beta mRNA expression in rat ovary: down-regulation by gonadotropins. Mol Endocrinol. 1997;11:172-82. [146] Ginther OJ. Reproductive Biology of the Mare: Basic and Applied Aspects: Equiservices, Cross Plains, WI; 1992. [147] Bergfelt DR, Ginther OJ. Relationships between FSH surges and follicular waves during the estrous cycle in Mares. Theriogenology. 1993;39:781-96. [148] Donadeu FX, Ginther OJ. Changes in concentrations of follicular fluid factors during follicle selection in mares. Biology of reproduction. 2002;66:1111-8.

114

[149] Ginther OJ, Meira C, Beg MA, Bergfelt DR. Follicle and endocrine dynamics during experimental follicle deviation in mares. Biology of reproduction. 2002;67:862-7. [150] Ginther OJ, Gastal EL, Gastal MO, Checura CM, Beg MA. Dose-response study of intrafollicular injection of insulin-like growth factor-I on follicular fluid factors and follicle dominance in mares. Biology of reproduction. 2004;70:1063-9. [151] Vernunft A, Lohrke B, Tuchscherer A, Weitzel JM, Viergutz T. Concentrations of Anti-Muellerian-Hormone in follicular fluid from antral follicles and their relation to functional characteristics of follicle development in mares. Berl Munch Tierarztl. 2013;126:77-82. [152] Pellicer A, Simon C, Remohi J. Effects of aging on the female reproductive system. Hum Reprod. 1995;10:77-83. [153] Carnevale EM, Ginther OJ. Relationships of age to uterine function and reproductive efficiency in mares. Theriogenology. 1992;37:1101-15. [154] Carnevale EM, Uson M, Bozzola JJ, King SS, Schmitt SJ, Gates HD. Comparison of oocytes from young and old mares with light and electron microscopy. Theriogenology. 1999;51:299 [155] Ball BA, Little TV, Hillman RB, Woods GL. Pregnancy rates at Days 2 and 14 and estimated embryonic loss rates prior to day 14 in normal and subfertile mares. Theriogenology. 1986;26:611-9. [156] Carnevale EM, Bergfelt DR, Ginther OJ. Aging effects on follicular activity and concentrations of FSH, LH, and progesterone in mares. Anim Reprod Sci. 1993;31: 287-99. [157] Ginther OJ, Gastal MO, Gastal EL, Jacob JC, Siddiqui MAR, Beg MA. Effects of age on follicle and hormone dynamics during the oestrous cycle in mares. Reproduction, Fertility and Development. 2008;20:955-63. [158] Miller KF, Berg SL, Sharp DC, Ginther OJ. Concentrations of circulating gonadotropins during various reproductive states in mares. Biology of reproduction. 1980;22:744-50. [159] Ginther OJ. Reproductive biology of the mare. Second Edition. 1992. [160] Claes A, Ball BA, Scoggin KE, Esteller-Vico A, Kalmar JJ, Conley AJ, et al. The interrelationship between anti-Mullerian hormone, ovarian follicular populations, and age in mares. Equine Veterinary Journal. 2014. [161] Alviggi C, Humaidan P, Howles CM, Tredway D, Hillier SG. Biological versus chronological ovarian age: implications for assisted reproductive technology. Reproductive biology and endocrinology : RB&E. 2009;7:101. [162] Carnevale EM. The mare model for follicular maturation and reproductive aging in the woman. Theriogenology. 2008;69:23-30. [163] Buratini J, Rosa e Silva AA, Barros CM, Papa FO, Caldas MC, Meira C. Follicular dynamics in Mangalarga mares. Equine veterinary journal Supplement. 1997:7-11. [164] Ginther OJ. Major and minor follicular waves during the equine estrous cycle. J Equine Vet Sci. 1993;13:18-25. [165] Watson ED, Mcdonnell AM, Cuddeford D. Characteristics of Cyclicity in Maiden Thoroughbred Mares in the United-Kingdom. Vet Rec. 1994;135:104-6. [166] Newcombe JR. The incidence of ovulation during the luteal during the luteal phase from day 4 to day 20 in pregnant and non pregnant mares. J Equine Vet Sci. 1997;17:120-2.

115

[167] Ginther OJ, Beg MA. Dynamics of circulating progesterone concentrations before and during luteolysis: a comparison between cattle and horses. Biology of reproduction. 2012;86:170. [168] Gastal EL, Gastal MO, Bergfelt DR, Ginther OJ. Role of diameter differences among follicles in selection of a future dominant follicle in mares. Biology of reproduction. 1997;57:1320-7. [169] Munro C, Stabenfeldt G. Development of a microtitre plate enzyme immunoassay for the determination of progesterone. The Journal of endocrinology. 1984;101:41-9. [170] Yoon MJ, Bolme I, Colgin M, Niswender KD, King SS, Alvarenga M, et al. The efficacy of a single chain recombinant equine luteinizing hormone (reLH) in mares: Induction of ovulation, hormone profiles, and inter-ovulatory intervals. Domest Anim Endocrin. 2007;33:470-9. [171] Erickson BH. Development and senescence of the postnatal bovine ovary. Journal of animal science. 1966;25:800-5. [172] Ginther OJ, Carnevale EM, Bergfelt DR. Delay in emergence of the ovulatory follicular wave in old mares. J Equine Vet Sci. 1993;13:75-9. [173] Morel MC, Newcombe JR, Hayward K. Factors affecting pre-ovulatory follicle diameter in the mare: the effect of mare age, season and presence of other ovulatory follicles (multiple ovulation). Theriogenology. 2010;74:1241-7. [174] Neely DP, Kindahl H, Stabenfeldt GH, Edqvist LE, Hughes JP. Prostaglandin release patterns in the mare: physiological, pathophysiological, and therapeutic responses. Journal of reproduction and fertility Supplement. 1979:181-9. [175] Thomas F, Renaud F, Benefice E, De Meeus T, Guegan JF. International variability of ages at menarche and menopause: Patterns and main determinants. Hum Biol. 2001;73:271-90. [176] Haag KT, Magalhaes-Padilha DM, Fonseca GR, Wischral A, Gastal MO, King SS, et al. Quantification, morphology, and viability of equine preantral follicles obtained via the Biopsy Pick-Up method. Theriogenology. 2013;79:599-609. [177] Haag KT, Magalhaes-Padilha DM, Fonseca GR, Wischral A, Gastal MO, King SS, et al. Equine preantral follicles obtained via the Biopsy Pick-Up method: histological evaluation and validation of a mechanical isolation technique. Theriogenology. 2013;79:735-43. [178] Charleston JS, Hansen KR, Thyer AC, Charleston LB, Gougeon A, Siebert JR, et al. Estimating human ovarian non-growing follicle number: the application of modern stereology techniques to an old problem. Hum Reprod. 2007;22:2103-10. [179] Vanderwall DK, Woods GL, Freeman DA, Weber JA, Rock RW, Tester DF. Ovarian follicles, ovulations and progesterone concentrations in aged versus young mares. Theriogenology. 1993;40:21-32. [180] Burns DS, Jimenez-Krassel F, Ireland JL, Knight PG, Ireland JJ. Numbers of antral follicles during follicular waves in cattle: evidence for high variation among animals, very high repeatability in individuals, and an inverse association with serum follicle- stimulating hormone concentrations. Biology of reproduction. 2005;73:54-62.

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Vita

Name: Anthony Claes

Place of Birth: Antwerp, Belgium

Educational institutions attended and degrees

2011-Present University of Kentucky, Department of Veterinary Science, Lexington,

Kentucky

2006-2008 University of California, School of Veterinary Medicine, Davis: Board

certification American College of Theriogenologists

2004 University of Ghent, School of Veterinary Medicine, Belgium: DVM

(Cum Laude)

Professional positions

Sep ‘10 – Aug’11 University of California, Davis, Veterinary MedicalTeaching

Hospital, School of Veterinary Medicine, Department of

Population Health and Reproduction

2010 – August 2010 Veterinary Medical Teaching Hospital, School of Veterinary

Medicine, University of California, Davis

2009 – 2010 Private Practice (Equine), Belgium, Europe

2008 – 2009 Scone Veterinary Hospital, Australia

2006 – 2008 Veterinary Medical Teaching Hospital, School of Veterinary

Medicine, University of California, Davis

Residency Equine Reproduction

2005 – 2006 Equine Clinic Bears – VDL Stud (van de Lageweg), The

Netherlands

117

2004 – 2005 Large Animal Mixed Practice, Germany

Veterinarian

Professional publications

Claes A, Ball BA, Scoggin KE, Esteller-Vico A, Kalmar JJ, Conley AJ, Squires EL,

Troedsson MHT. The interrelationship between anti-Müllerian hormone, ovarian follicular populations, and age in mares. Equine Vet J: In review

Canisso IF, Ball BA, Erdal E, Claes A, Scoggin KE, McDowell K, Williams N, Dorton

A, Wolfsdorf K, Squires EL, Troedsson MHT. Experimental induction of nocardioform placentitis (Crosiela equi) in mares. Equine Vet J; early view

Claes A, Ball BA, Corbin CJ, Conley AJ. Anti-Müllerian hormone as a diagnostic marker for equine cryptorchidism in three cases with equivocal testosterone concentrations.

Journal of Equine Vet Science 2014, 34, 442-445

Meyers-Brown GA, McCue PM, Troedsson MHT, Klein C, Zent W, Ferris FA, Lindholm

ARG, Scofield DB, Claes A, Morganti M, Colgin MA, Wetzel RL, Peters AR, Roser JF.

Induction of ovulation in seasonally anestrous mares under ambient lights using recombinant equine FSH (reFSH). Theriogenology 2013; 80: 456-462.

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Claes A, Ball BA, Corbin CJ, Conley AJ. Age and season affect serum testosterone concentrations in cryptorchid stallions. Vet Rec 2013. Aug 17-24; 173(7):168. doi:

10.1136/vr.101706. Epub 2013 Jun 27.

Claes A, Ball BA, Almeida J, Corbin CJ, Conley AJ. Serum anti-Müllerian hormone concentrations in stallions: Developmental changes, seasonal variation, and differences between intact stallions, cryptorchid stallions, and geldings. Theriogenology 2013; 79:

1229-1235.

Claes A, Ball BA, Liu IKM, Vaughan B, Highland MA, Brown JA. Uterine B-cell lymphoma in a mare. Equine Vet Edu; early view

Claes A, Ball BA, Brown JA, Kass PH. Evaluation of risk factors, management, and outcome associated with rectal tears in horses: 99 Cases (1985-2006). J Am Vet Med

Assoc 2008; 233: 1605-9.

Brown JA, O’Brien MA, Hodder ADJ, Peterson T, Claes A, Liu IKM, Ball BA.

Unilateral testicular mastocytoma in a Peruvian Paso stallion. Equine Vet Edu 2008; 20:

172-5.

119