Associations between Canine Male Reproductive Parameters and

Serum Vitamin D and Prolactin Concentrations

by

Adria Julianne Kukk

A Thesis presented to The University of Guelph

In partial fulfillment of requirements for the degree of Doctor of Veterinary Science in Population Medicine

Guelph, Ontario, Canada

¤ Adria Julianne Kukk, December, 2011

ABSTRACT

ASSOCIATIONS BETWEEN CANINE MALE REPRODUCTIVE PARAMETERS

AND SERUM VITAMIN D AND PROLACTIN CONCENTRATIONS

Adria Julianne Kukk Advisor: University of Guelph, 2011 Professor C.J. Gartley

Maintaining reproductive health and diagnosing and treating conditions of infertility in stud dogs is important in canine theriogenology. However, there is still a great deal to be learned about reproductive physiology and factors that affect reproductive organs and semen quality in dogs. This thesis is an investigation of two factors in the male dog; serum 25-hydroxy Vitamin D

(25OHVD) and prolactin (PRL) concentrations, and their possible associations with benign prostatic hyperplasia (BPH), prostate volume and/or sperm morphology and motility characteristics.

28 (Vitamin D Study) and 29 (28 plus one for the Prolactin study) client dogs of various breeds from the Ontario Veterinary College and Graham Animal

Hospital in Southwestern Ontario, Canada were enrolled in the study from March to December 2009. Of these dogs 22 were successfully collected for semen.

BPH was diagnosed using prostate volume measured by ultrasound, as well as clinical signs including blood in the ejaculate. Semen analysis was performed using manual microscopic techniques for morphology and computer assisted sperm analysis equipment for motility.

In the vitamin D study, no associations were found between BPH and serum 25OHVD concentrations. In contrast, several sperm motility (motility, progressive motility, beat cross frequency (BCF), distance average path (DAP), curvilinear distance (DCL), linear distance (DSL), average path velocity (VAP), curvilinear velocity (VCL) and straight line velocity (VSL), amplitude lateral head displacement (ALH) and average orientation change (AOC)) and morphology characteristics (percentage normal sperm, head defects and detached heads) had desirable outcomes with 25OHVD concentrations between 120-180 nmol/l.

Using bivariable analysis, positive associations were observed with 25OHVD and some semen quality characteristics from 4 to 8 years of age (motility, progressive motility, BCF, DCL, VCL, ALH, AOC) and at transformed prostate volumes smaller than or equal to 4.5 (motility, progressive motility, DCL, VCL, and normal morphology) while negative associations of these semen parameters were found at ages greater than 8 years and transformed prostate volumes greater than or equal to 5.5. Head defects were negatively associated with 25OHVD. Vitamin D may have an impact on spermatogenesis and normal sperm physiology that warrants further research.

The prolactin study showed no statistically significant associations between serum PRL and BPH and serum PRL and sperm motility characteristics.

However, two sperm morphology characteristics (percentage proximal droplets and percentage midpiece defects) had significant negative associations with PRL concentrations. Age interaction with PRL was also a factor in the percentage of midpiece defects with desirable outcomes associated at 4 years of age compared with older ages. Overall, undesirable outcomes occurred at PRL concentrations less than 2.5 ng/ml. In conclusion, both 25OHVD and PRL may have important roles in spermatogenesis and normal sperm physiology in the dog.

ACKNOWLEDGEMENTS

The past few years have been an exciting and important part of my life. I would not have been able to complete this great undertaking without the support and knowledge of a great many people. First I would like to thank my advisor

Dr. Cathy Gartley for her wealth of knowledge in the field of theriogenology, for her mentorship, her kindness and lastly, for her sense of humour to make me laugh even on the greyest of days. Also, I would like to thank members of my advisory committee: Drs. Tracey Chenier, Andria Jones, Stephanie Nykamp and

Heather Chalmers for their patience and expertise in guiding me through my research and for their critical advice in pushing me to create a body of work I can be proud of. For his statistical expertise, I am grateful to William Sears whose help was invaluable in the analysis of my data.

For their friendship, support and encouragement during my D.V.Sc. programme I would like to thank Drs. Rasa Levstein and Cyril Stephen, our veterinary technicians Jim Rahn and Karen DiCaro, as well as my fellow graduate students Drs. Mariana Diel De Amorim and Leslie Gonzalez. I would also like to thank Dr. Walter Johnson for his encouragement and for sharing his knowledge of bovine theriogenology. A special thank-you to Sally, Linda, Julie and Karla in the Population Medicine office, and Mary Elliot for helping me with the bureaucratic aspects of graduate studies and for making my life that much easier.

To Dr. Carol Graham, all the staff at Graham Animal Hospital, and to all our clients who agreed to enroll their dogs in this study, I extend a warm

v appreciation for their participation. Without them there wouldn’t have been any data to analyse. I wish to also thank the Pet Trust for providing the funding to make this study possible.

Lastly, I would like to thank two very special people in my life, my mother

Eva and my husband Anthony (Toncsi) for the much needed emotional support during my graduate programme. Their patience and unending belief in me gave me the strength I needed to achieve my goals.

vi

DECLARATION OF WORK PERFORMED

I declare that with the exception of the items listed below, all work presented in this thesis was performed by me.

The Animal Health Laboratory, University of Guelph, performed urinalysis and culture and sensitivities for urine, prostatic fluid and semen.

The Endocrinology Division of the Diagnostic Center of Population and Animal

Health, Michigan State University, performed the radioimmunoassay for Vitamin

D analysis.

The New Animal Science Laboratory, Washington State University, performed the enzyme-linked immunosorbent Assay for Prolactin analysis.

vii

TABLE OF CONTENTS

CHAPTER ONE ………………………………………………………………… 1 INTRODUCTION, RESEARCH OBJECTIVES AND LITERATURE REVIEW …………………………………………...... 1 1. INTRODUCTION …………………………………………………….. 1 2. RESEARCH OBJECTIVES..………………………………………… 3 3. LITERATURE REVIEW.……………………………………………… 4 3.1 PHYSIOLOGY OF CANINE SPERMATOGENESIS AND METHODS OF SEMEN ANALYSIS………………………… 4 3.1.1 Reproductive Physiology of the Testes and the Role of Hormones and Local Factors In Male Reproduction….. 4 3.1.2 Spermatogenesis ………………………………………. 13 3.1.3 Factors Affecting Semen Quality.…………………… 16 3.1.4 Methods of Semen Evaluation……………………… 17 3.2 THE CANINE PROSTATE GLAND ……………………….. 31 3.2.1 Anatomy and Physiology of the Prostate Gland ……... 31 3.2.2 Pathophysiology of Benign Prostatic Hyperplasia…….. 33 3.2.3 Conventional Treatment of Benign Prostatic Hyperplasia ……………………………………………….. 39 3.2.4 Ultrasonography of the Prostate.……………………….. 42 3.3 VITAMIN D AND ITS ROLE IN MALE REPRODUCTION.. 44 3.3.1 Vitamin D Metabolism ………………………………… 45 3.3.2 Vitamin D and the Prostate …………………………… 49 3.3.3 Role of Vitamin D in Sperm Production ……………… 54 3.4 PROLACTIN AND ITS ROLE IN MALE REPRODUCTION.. 56 3.4.1 Prolactin and Ovarian Physiology.……………………. 56 3.4.2 Prolactin and Male Physiology.………………………. 58 3.4.2.1 Prolactin and the Prostate.…………………………. 58 3.4.2.2 Prolactin, Male Fertility and Semen Quality ……… 60

viii References ………………………………………………………… 67

CHAPTER TWO …………………………………………………………… 81 VITAMIN D, BENIGN PROSTATIC HYPERPLASIA, PROSTATE VOLUME AND SEMEN PARAMETERS IN THE DOG………… 81 Abstract.……………………………………………………………... 81 1. Introduction……………………………………………………….. 82 2. Materials and Methods.………………………………………….. 84 3. Results.……………………………………………………………. 92 4. Discussion and Conclusions.…………………………………… 98 References ………………………………………………………….. 143

CHAPTER THREE.…………………………………………………………… 147 PROLACTIN, BENIGN PROSTATIC HYPERPLASIA, PROSTATE VOLUME AND SEMEN PARAMETERS IN THE DOG.. 147 Abstract.………………………………………………………………... 147 1. Introduction …………………………………………………………. 148 2. Materials and Methods ……………………………………………. 151 3. Results ……………………………………………………………… 155 4. Discussion and Conclusions ……………………………………… 157 References.…………………………………………………………….. 178

CHAPTER FOUR …………………………………………………………….. 180 SUMMARY AND CONCLUSIONS ………………………………… 180 References……………………………………………………………. 184 APPENDICES ……………………………………………………………….. 185

ix

LIST OF TABLES

Table 1.1 Selected Physical and Chemical Analyses of Prostatic Fluid in Dogs with Healthy Prostates.…………………………………………… 33

Table 1.2 Genes influenced by Vitamin D receptor ligands and their effects 48

Table 1.3 Prolactin values in Dogs (ELISA) ……………………………………65

Table 2.1 Technical Parameters for SpermVision™ CASA …………………111

Table 2.2 Classification scheme for determination of the presence of BPH by ultrasonography in 29 dogs, March-December 2009, Ontario, Canada ………………………………………………………….112

Table 2.3 Characteristics of the study population of 28 dogs, in Ontario, Canada, March-December 2009.……………………………………….113

Table 2.4 Characteristics of the study population of 22 dogs from whom semen was collected in Ontario, Canada March-December 2009…………...114

Table 2.5 Breed, Vitamin D concentration, age and fertility data on 22 dogs from whom semen was collected in Ontario, Canada, 2009…………115

Table 2.6 Mean serum Vitamin D concentration, with respect to type of diet, with associated test-statistic, p-value and confidence intervals………116

Table 2.7 Mean serum Vitamin D concentration, with respect to time of year of sampling, with associated test-statistic, p-value and confidence intervals……………………………………………………………………..117

Table 2.8 Mean serum Vitamin D concentration, with respect to number of hours spent outdoors at time of year of sampling, with associated test-statistic, p-value and confidence intervals…………………………………………118

Table 2.9 Mean serum Vitamin D concentration, with respect to BPH status, and associated test-statistic, p-value and confidence intervals……….119

Table 2.10 Univariable simple linear regression analyses of sperm motility parameters with serum 25-hydroxy vitamin D as explanatory variable, with associated coefficient of determination (R2), test statistic and p-value…………………………………………………………………120

x Table 2.11 Non-significant bivariable linear regressions of motility with explanatory variable regression coefficients (where applicable), coefficients of determination for the overall model (R2), test statistics, and the associated p-values..……………………………………………………………………121

Table 2.12 Three bivariable linear regressions of morphology parameters and associated coefficients of determination (R2), test-statistic and p-values………………………………………………………………..123

Table 3.1 Technical Parameters for SpermVision™ CASA …………………..166

Table 3.2 Classification scheme for determination of the presence of Benign Prostatic Hyperplasia (BPH) by ultrasonography in 29 dogs, March-December 2009, Ontario, Canada.………………………………167

Table 3.3 Breed, prolactin concentration, age and fertility data on 22 dogs from whom semen was collected in Ontario, Canada, 2009………….168

Table 3.4 Mean serum prolactin concentrations and confidence intervals with respect to benign prostatic hyperplasia status………………………….169

Table 3.5 Univariable analyses using simple linear regression modeling of semen motility parameters and prolactin with coefficients of determination (R2) of the overall model, test statistic and associated p-value…………………………………………………………………...... 170

Table 3.6 Three bivariable linear regressions of motility parameters with coefficients of determination of overall model (R2), test statistics, and associated p-values…………………………………...... 171

Table 3.7 Univariable analyses of sperm morphology parameters and PRL with coefficients of determination (R2) of the overall model, test-statistic and associated p-values……………….………………………………………173

Table 3.8 Three bivariable linear regressions of morphology parameters, coefficients of determination (R2) of the overall model, test statistics and associated p-values…………………………………………………… 174

Table 3.9 Three non-significant bivariable linear regressions of proximal droplets with variable coefficients where PRL slope coefficients were significant, coefficients of determination for the overall model (R2) and associated p-values………………………………………………………………… 175

xi

LIST OF FIGURES

Figure 2.1 Predicted sperm motility as a response to normal serum 25OHVD range in dogs…………………………………………124

Figure 2.2 Progressive motility as a response to normal serum 25OHVD range in dogs………………………………………………...125

Figure 2.3 Predicted sperm motility in response to normal serum 25OHVD range at multiple ages……………………………………….126

Figure 2.4 Predicted sperm motility in response to normal serum 25OHVD range at various transformed prostate volumes…………..127

Figure 2.5 Predicted progressive motility in response to normal serum 25OHVD range at various ages………………………………..128

Figure 2.6 Predicted progressive motility in response to normal serum 25OHVD range at various transformed prostate volumes…..129

Figure 2.7 Predicted BCF in response to normal serum 25OHVD range at various ages……………………………………………………………130

Figure 2.8 Predicted DCL in response to normal serum 25OHVD range at various ages…………………………………………………………….131

Figure 2.9 Predicted DCL in response to normal serum 25OHVD range at various transformed prostate volumes……………………………….132

Figure 2.10 Predicted VCL in response to normal serum 25OHVD range at various ages……………………………………………………………..133

Figure 2.11 Predicted VCL in response to normal serum 25OHVD range at various transformed prostate volumes………………………………..134

Figure 2.12 Predicted DSL in response to normal serum 25OHVD range at various transformed prostate volumes……………… ……………….135

Figure 2.13 Predicted ALH in response to normal serum 25OHVD range at various ages……………………………………………………………..136

Figure 2.14 Predicted AOC in response to normal serum 25OHVD range at various ages……………………………………………………………..137

xii Figure 2.15 Predicted Normal sperm in response to normal serum 25OHVD range…………………………………………………………138

Figure 2.16 Predicted Loose Heads in response to normal serum 25OHVD range…………………………………………………………139

Figure 2.17 Predicted Normal sperm in response to normal serum 25OHVD range at various ages………………………………………140

Figure 2.18 Predicted Normal sperm in response to normal serum 25OHVD range at various transformed prostate volumes…………141

Figure 2.19 Predicted head defects in response to normal serum 25OHVD range at various transformed prostatic volumes…………142

Figure 3.1 Predicted percentage of proximal droplets in response to serum PRL ………………………………………………………………176

Figure 3.2 Predicted percentage of midpiece defects in response to serum PRL……………………………………………………………….177

xiii LIST OF ABBREVIATIONS:

ABP Androgen binding protein IL Interleukin ALH Amplitude lateral head IVF In vitro fertilization diplacement AOC Average orientation change LH Luteinizing hormone AR Androgen receptor LIN Linearity BCF Beat cross frequency MMP Matrix metalloprotease BPH Benign prostatic hyperplasia NF Necrosis factor BTB Blood-testis barrier NRR Non-return rates Ca Calcium P Phosphorus CASA Computer assisted sperm PR Progesterone receptor analysis CBC Complete blood count PRL Prolactin CNRR Corrected non-return rates PTH Parathyroid hormone DAP Mean distance PVN Periventricular nucleus DCL Curvilinear distance RXR Retinoid X receptor DHT Dihydrotestosterone SCN Suprachiasmatic nucleus DSL Straight line distance SE Seminiferous epithelium E2 Estradiol STR Straightness coefficient ER Estrogen receptor TGFE Transforming growth factor beta FGF Fibroblast growth factor TNFD Tumor necrosis factor alpha FGF-7 Keratinocyte growth factor VAP Mean velocity FITC Fluorescein isothiocyanate VCL Curvilinear velocity FSH Follicle stimulating hormone VDR Vitamin D receptor GH Growth hormone VDRE Vitamin D response element GnRH Gonadotrophin releasing VSL Straight line velocity hormone HPTA Hypothalamic-pituitary-testes WOB Wobble coefficient axis HPTH Hypothalamus 3EHSDH 3-beta hydroxysteroid dehydrogenase IGF Insulin-like growth factor 25OHVD 25-hydroxy Vitamin D

xiv CHAPTER ONE

INTRODUCTION, RESEARCH OBJECTIVES AND LITERATURE

REVIEW

1. INTRODUCTION

Prostatic diseases impact both general and reproductive health and are influenced by androgens produced by the testes. Ironically, the very hormone that is necessary for sperm production and male behaviours, such as libido, can also have a negative impact on fertility and reproduction. Benign prostatic hyperplasia (BPH) is the most common age-related condition in intact male dogs.

The condition affects approximately 80% of sexually intact dogs over five years old and is characterized by an enlarged prostate as a result of both hyperplasia and hypertrophy of prostatic epithelial cells [1, 2]. BPH may affect fertility, especially the ability to cryopreserve semen in valuable stud dogs, however, it also increases the susceptibility of the prostate to infection via ascension of normal bacterial flora into an abnormal prostate and consequent prostatitis [3].

Surgical castration is the treatment of choice but represents a severe financial and genetic loss for dog breeders. Unlike human males with the similar condition, an enlarged prostate is not associated with impairment of urination but can eventually lead to signs of hematuria, constipation and discomfort [2, 3].

Treatment with estrogens is known to decrease prostate size, however, severe side effects such as squamous metaplasia of the prostate, thrombocytopenia,

1 leukopenia and fatal aplastic anemia discourage their use. Human approved treatments used off-label in veterinary medicine are often cost prohibitive. There is also very little published research on specific effects on physiological factors related to spermatogenesis and semen quality in the dog. Recent human research suggests that both Vitamin D and prolactin (PRL) have an influence on the size of the human prostate and semen quality [4-7]. These findings have suggested that investigation into prostate size and any possible association with

Vitamin D and/or prolactin may aid development of new treatments for BPH and poor semen quality in the dog.

2 2. RESEARCH OBJECTIVES

The overall goal of this study was to investigate and identify contributing etiologies of BPH that could lead to new prevention and treatment strategies.

A. Vitamin D study

I. Determine whether an association exists between serum

concentrations of 25-hydroxyvitamin D and BPH in male dogs.

II. Determine whether an association exists between serum

concentrations of 25-dihydroxyvitamin D and semen quality in

dogs.

B. Prolactin Study

I. Determine whether an association exists between serum

prolactin concentrations and BPH in male dogs.

II. Determine whether an association exists between

serum prolactin concentrations and semen quality in male dogs.

3 3. LITERATURE REVIEW

3.1 PHYSIOLOGY OF CANINE SPERMATOGENESIS AND METHODS OF

SEMEN ANALYSIS

3.1.1 Reproductive Physiology of the Testes and the Role of Hormones and

Local Factors In Male Reproduction

Male reproductive physiology controls the processes by which spermatozoa are produced, transported and deposited into the female tract for subsequent fertilization of the oocyte. Any interference with these processes, whether physical, biological or chemical, can consequently affect fertility.

Spermatogenesis is controlled by the hypothalamic-pituitary-testicular axis, which will be discussed in detail. The remainder of the reproductive tract, including epididymides, deferent ducts, urethra, prostate gland, and penis is responsible for the maturation, storage and/or transport of spermatozoa and seminal fluid to be deposited in the female during estrus. Although male physiology shares many similarities across different mammalian species, the dog has its own set of unique variations.

The testes are found retroperitoneally and contained within the externally located scrotum. The external location of the testes is crucial in maintaining optimal temperatures for sperm production, as the process of spermatogenesis is sensitive to the relatively high internal body temperature [8]. Thermoregulation is

4 further controlled through counter-current heat exchange between the arterial and venous blood supplying the testes, via the complicated network of vessels that form the pampiniform plexus [8]. The activity of scrotal sebaceous glands, the thinness and lack of hair of the scrotal skin, and most importantly the mechanical repositioning of the testis, with relation to the body wall, via the cremaster muscle located within the spermatic cord and dartos of the scrotum, further contributes to thermoregulation [8].

The testes contain three main cell types that are responsible for sperm production: germ cells or spermatogonia, Sertoli and Leydig cells. The latter two cell types are under hormonal control via the hypothalamic-pituitary-testicular axis (HPTA). Gonadotrophin releasing hormone (GnRH) from the hypothalamus, stimulates the secretion of pituitary luteinizing hormone (LH) and follicle stimulating hormone (FSH). These hormones act on Leydig and Sertoli cells, respectively, stimulating production of testosterone from Leydig cells and synthesis of androgen binding protein (ABP) and aromatization of testosterone to estrogen in Sertoli cells [9]. Testosterone released into the peripheral circulation is converted to dihydrotestosterone (DHT) through 5D-reductase activity in a variety of tissues, such as the prostate and hair follicles, and is responsible for many of the secondary sex characteristics, especially during male embryonic development [8, 9]. Negative feedback of the HPTA is mediated via both testosterone and estrogen that act directly on the hypothalamus. Inhibin, another hormone product of Sertoli cells, also acts via negative feedback on the pituitary

5 to limit FSH release. In this way a fine-tuned control of spermatogenesis is achieved [9].

The Sertoli cell is responsible for the support and protection of developing spermatozoa, by supplying the necessary nutrients and protecting the haploid and antigenically foreign spermatocytes, spermatids and spermatozoa from immune attack by the male’s defenses [10]. This barrier, known as the blood- testis barrier (BTB), is crucial for the survival of the developing spermatozoa that reside within the intercellular spaces of adjacent Sertoli cells and which are separated from each other in their different developmental stages and the extracellular environment via intercellular tight-cell junctions between Sertoli cells

[10, 11]. Testosterone is responsible for the adhesion between the developing spermatocyte and Sertoli cell, maintaining the integrity of the BTB through epidermal growth factor and its receptor [12]. The Leydig cells, also known as interstitial cells, are separated from the Sertoli cells and spermatogonia by a basement membrane and are supplied and supported by connective tissue, lymphatics and blood vessels with which they have close contact [8]. The specialized needs and functions of these differing testicular cell types mirror the morphological and histological arrangement of the testes.

The Leydig cells, connective tissues and vessels are part of the interstitial compartment of the testicular parenchyma, and as previously mentioned are separated from Sertoli and germ cells by a basement membrane. The seminiferous epithelium (SE) is present on the opposite side of the basement membrane and is divided into three compartments: basal, deep and peripheral

6 adluminal [8, 13]. The importance of these compartments will be further explained in the section on spermatogenesis. The seminiferous epithelium forms a convoluted network of tubules within the microscopic lobules of the testicular parenchyma that eventually join with rete tubules in the centre of the testis to form the macroscopic structure called the mediastinum [8]. This is the major collection region for spermatozoa within the testes, before they are further transported through the efferent ductules to the head of the epididymis where the final stages of spermatozoal maturation take place.

In addition to the long feedback loop with respect to the HPTA, certain local factors have attracted attention in regulation of Sertoli and Leydig cell function. These include, but are not exclusive to the inflammatory cytokines tumor necrosis factor alpha (TNFD) and interleukins (IL), and growth factors such as growth hormone (GH), transforming growth factor beta (TGFE), and insulin- like growth factor (IGF) [12, 14]. Unfortunately, studies relating to these local regulating factors of spermatogenesis are lacking in dogs.

In vitro immunohistochemical studies with testicular tissue have isolated

TNFD (a 50kDa homotrimer originally isolated from endotoxin-stimulated macrophages that caused necrosis in tumor tissue) from round and elongated spermatids, pachytene spermatocytes (see section on spermatogenesis), interstitial macrophages in mice [15], and in Sertoli cells in rats [16]. Its receptor

(TNFD-R) has been shown to be present in both Sertoli cells in pigs [17] and

Leydig cells in the rat [18]. In porcine Sertoli cell culture under FSH stimulation, an up-regulation of TNFD -R in Sertoli cells has been reported [17] and suggests

7 a role of FSH in TNFD sensitization in Sertoli cells. Apoptosis of germ cells through FSH mediated expression of TNFD has also been shown [19]. In Leydig cells, TNFD inhibits steroidogenesis via the nuclear factor-NB (NF-NB) pathway in mouse cell culture to block the enzymes P450scc, p450c17 and 3E- hydroxysteroid dehydrogenase (3EHSD) that are necessary in the multi-stage conversion of cholesterol to testosterone [18]. Up-regulation of the androgen receptor (AR) in rat Sertoli cells through the same NF-NB pathway in DNA-protein binding studies [20] is yet another effect of TNFD in the possible regulation of spermatogenesis and the interaction between Sertoli, Leydig and germ cells.

Of particular importance, TNFD has been shown to down-regulate certain proteins involved in maintaining the BTB, suggesting a restructuring of tight junctions to maintain spermatocytes in their separate developmental states as they move through to the luminal surface of the seminiferous epithelium (SE)

[12]. This is achieved in part by enhancing plasminogen activator/plasminogen activator inhibitor system present in Sertoli and germ cells to induce phagocytosis of residual bodies by the Sertoli cells during spermiation [12, 21,{Liu, 2007

#740}]. Induction of matrix metalloproteases (MMPs) necessary in endocytosis and degradation of tight junction proteins causes reversible increases in the permeability of tight junctions in the blood-testes barrier (BTB) [16]. Much of the data on TNFD has been compiled from studies examining rodent and swine testicular tissue in culture, and it is yet unknown whether similar mechanisms of action exist in the dog and/or how these factors may interact in vivo at physiological levels in this species.

8 Li et al (2003) studied the function of TNFD in in vivo studies in rats [23].

Using a recombinant human TNFD at a dose of 0.5 micrograms per testis

(determined previously to be within physiological levels by the criteria that effects were reversible and non-cytotoxic to Sertoli cells [16]) and an acute high dose of

2 micrograms per testis, immunohistochemical, immunoblotting and electron microscopic techniques were used to identify a multitude of tight junction proteins and observe seminiferous SE structure. The contralateral testis was used as a vehicle-treated or non-treated control. Tissue samples were collected at time 0,

7 hours – 3 days, 3-5 days, 5-8 days and 14-60 days post-treatment. Decreases in occludin and zona occludins-1 along with disruption of actin filament structure related to tight junctions were found by day 3 for both treatment doses.

Complete recovery of the blood-testis barrier with germ cell repopulation was complete by days 14-60 post-treatment. Increases in intercellular spaces between Sertoli cells and thinning of the SE, damage to the tubules, the presence of spermatids and spermatocytes within the SE lumen and depletion in numbers of these cell types by day 5 were consistent with tight junction disruption. Functional BTB integrity was also determined by testing diffusion of fluorescent dye (FITC) administered systemically. FITC was found in the lumen of the SE while in control testes FITC remained confined behind the BTB. By day

8, occludin and zona occludin-1 levels, as well as FITC permeability, returned to normal showing no difference among treated, vehicle and non-treated controls.

In summary, although greater than normal physiological doses of TNFD result in dramatic disruption of Sertoli cell tight junctions, the presence of small amounts

9 of TNFD in physiological normal testis suggests it is needed for normal function of the BTB. It is possible that TNFD helps in physiological reorganizing of the

Sertoli cell tight junctions to allow normal progress of developing spermatocytes through the SE for eventual release of spermatids into the SE lumen. It is important to note that only the results of the acute high dose treatment with TNFD were reported. Although physiological doses of TNFD were investigated and found to be significant and similar to the results obtained using supraphysiological doses, it is unclear why more detailed reporting of these results was absent.

The transforming growth factor beta (TGFE) family consists of three isoforms and all three are expressed in a developmental and stage dependent manner by early spermatids, pachytene spermatocytes and Sertoli cells, as well as Leydig and myoid cells in the rat and porcine testes [24]. Along with TNFD, they can disrupt the BTB in a reversible manner by degradation, endocytosis and reorganization of tight junction transmembrane proteins to allow movement of developing spermatocytes through the seminferous epithelium [12, 24]. Xia et al

(2009) used similar methodology to the study by Li et al (2003) and produced similar outcomes with respect to functional permeability of the BTB when exposed to TGFE3 [25]. However, they went one step further using an in vitro endocytosis assay to measure amounts of endocytosed tight junction proteins

(occludin, junction adhesion molecule-A and N-cadherin) over time in Sertoli cells when treated with endogenous amounts of TGFE3, TNFD and germ cell culture medium. The study found increases in the endocytosed proteins by 10 minutes

10 post-treatment when compared with controls. This effect was reversed by 180 minutes post-treatment. It is apparent that cross-talk between the developing sperm cells and Sertoli cells is necessary for the gradual breakdown of these tight junctions as spermatocytes make their journey across the SE to the lumen for eventual release. The current literature supplies strong evidence for this paracrine communication between developing germ cells, Sertoli and Leydig cells and their regulatory roles in tight junction physiology.

Interleukin-1, traditionally known for its role in inflammation and produced from macrophages, has been discovered to play a part in normal physiological processes, and more specifically, spermatogenesis [26]. The 17kDa IL-1D subtype has been studied in detail and determined to have multifunctional properties including acting as a growth factor for Sertoli cells [27] and spermatogonia [28], stimulating production of other cytokines such as IL-6

(associated with apoptosis of germ cells [29]) and activin A (part of the TGFE family) in Sertoli cells, and regulation of steroid production by Leydig cells [26].

Interleukin 1D is produced by Sertoli cells and shown to be dependent on the presence of germ cells [30]. Colon et al (2005) were able to confirm previous studies relating IL-1D with increased Leydig cell steroidogenesis [31] with addition of IL-1D to cultured immature rat Leydig cells where they observed a 4.3 fold increase in testosterone and a 2.86 fold increase in DHT [32] concentrations.

This stimulatory effect on androgen production is only seen in immature rat cell culture while the opposite effect is seen in mature Sertoli cells and may be the primary mechanism necessary for testicular maturation in pubescence [32].

11 Indeed, Svechnikov et al (2001) studied age related effects of IL-1D in 40 and 80 day old rats with suppression of testosterone secretion in the adult Sertoli cell culture [31], while Colon et al (2005) reported IL-1D action on increasing testosterone and DHT concentrations is potentiated by GH and IGF-1 in immature rat cell cultures [32]. As with TNFD and TGFE, IL-1D also plays a role in the regulation of the BTB through recycling of tight junction proteins within

Sertoli cells [26]. Sarkar et al (2008) reported that although steady-state levels of tight junction proteins were not affected by IL-1D treatment of cultured Sertoli cells, a loss of F-actin filament structure, identified using immunofluorescence staining, was observed starting at day 8 and persisted until day 45 [33]. In addition, diffuse localization of tight junction proteins (such as occludin and junction adhesion molecule-1) away from the BTB and increased permeability as assessed using the FITC technique described previously were significant findings and were not seen in the non-treated controls [33]. Although similar results were obtained to TNFD and TGFE studies of tight junction physiology, IL-1D actions were delayed and not readily reversible in the rat [33].

The growth hormones GH and IGF-1 have also been associated with spermatogenesis and semen quality. Their main effects appear to involve survival and functioning of Leydig cells and are produced locally by Leydig and

Sertoli cells [14]. GH stimulates testosterone secretion by Leydig cells in rats

[34] and IGF-1 stimulates testosterone secretion in rat Leydig cells [35] and increases the number of LH-receptors (LH-R) present in porcine Leydig cells in culture [36]. Their effects are more dramatic in immature compared with adult

12 Leydig cell culture in both species [35, 36]. A recent in vitro study by Yoon et al

(2011) in horses however, found no effect of IGF-1 alone or in combination with

LH in prepubertal horses although a synergistic increase in testosterone was observed in post-pubertal equine Leydig cell culture with LH than with LH alone

[37]. This indicates a less important role for IGF-1 during sexual development compared with post-pubertal steroidogenesis in this species and is supported by previous study [38]. It is important to note that incubation with IGF-1 was much shorter in the equine study (24 hours) compared with a three day and 48 hour incubation period in the rat and pig studies, respectively [35, 36] and may not have been long enough to elicit a significant change. As with the other local factors studied with respect to spermatogenesis, there is little or no information as to their presence or role in spermatogenesis in the dog. Further research within this area is needed if spermatogenesis in the canine is to be understood and how other factors may impact the finely tuned control of reproductive physiology in this species.

3.1.2 Spermatogenesis

Spermatogenesis is the process by which the primary germ cells are transformed into the antigenically and chromosomally distinct spermatozoa that are capable of fertilizing an oocyte. The process of spermatogenesis is commonly broken down into several distinct stages: mitosis, meiosis, spermiogenesis and spermiation [39, 40]. During mitosis primitive type A spermatogonia in the basal compartment actively replenish themselves [39], to

13 overcome the consequences of normal aging and apoptosis. In this way, there is a continuous supply of primary germ cells to maintain sperm production, accounting for the male’s ability to produce sperm throughout his lifetime. Some of these type A spermatogonia are selected through mechanisms involving testosterone, retinoic acid and stem cell factors, and Dazl, Sox3 and UTP14 genes to become type B spermatogonia [41, 42]. Type B spermatogonia continue to undergo mitosis and their last mitotic replication results in the primary spermatocytes. At this transitional or preleptotene meiotic stage the primary spermatocyte is present in the basal compartment [39].

Meiosis begins with the leptotene stage and at this time the primary spermatocyte traverses the BTB or junctional complex (adherins and tight junctions) between adjacent Sertoli cells and comes to lie in the deep adluminal compartment [9, 39]. Meiosis I proceeds similarly to mitosis to produce the diploid secondary spermatocyte (2n) and subsequently meiosis II, which occurs behind the BTB, results in haploid round spermatids (1n) that are now antigenically distinct from the other cells in the body. In this way four spermatids are produced from one primary spermatocyte.

Spermiogenesis is divided into four phases: Golgi, cap, acrosome and maturation. In the Golgi phase, the Golgi tubules manufacture the enzymes that will be necessary for the development of the acrosome system and dictate its position over the sperm nucleus [39]. Capping is the next process by which the acrosome comes into contact with the nuclear envelope and covers the nucleus over one third of its length. Following ‘capping’ the acrosomal phase continues

14 with the migration of the acrosomal system over the ventral surface of the elongating spermatid [39]. The last phase of spermiogenesis is less dramatic as the acrosome thins and its migration ends as it covers the majority of the nucleus. The nuclear material continues to condense and excess cytoplasm is removed from the cell and phagocytosed by the Sertoli cell. Also during this stage the centriole elongates to become the tail and is oriented towards the lumen. Spermiation is the final stage by which the Sertoli cell releases the spermatid and excess cytoplasm and organelles are expelled from the spermatid

[39] resulting in a spermatozoon.

The timing of spermatogenesis has been described by the cell associations or stages of the spermatogenic cycle (i.e. the time from beginning of spermatogenesis until spermatid release from the associated Sertoli cell). Since each Sertoli cell has many stages of developing spermatocytes, patterns of cellular development are recognized and referred to as cell stages. In the dog, one cell cycle contains eight stages and is 13.6 days in length [10, 42]. Total duration of spermatogenesis is 61.2 days and is equivalent to 4.5 spermatogenic cycles [42, 43].

The final stages of sperm maturation occur in the epididymis and are also under the control of androgens. In the epididymis sperm acquire motility and fertilization capacity through epididymal cell production of glycoproteins and transpeptidases (antioxidant regulation) that promote condensation of DNA. In the dog this process lasts approximately 10 days [44]. During transport from the head to the tail of the epididymis the last remnants of cytoplasm travel from the

15 proximal to the distal portion of the sperm flagellum for release prior to final maturation.

3.1.3 Factors Affecting Semen Quality

There are several factors that can affect semen quality in otherwise healthy dogs. These include inbreeding, age of dog, breed, collection technique and frequency of collection. In a study by Santos et al (2006), a knobbed acrosome defect and poor fertility was observed in four closely related Miniature

Schnauzer dogs where coefficients of inbreeding were 5.3, 10.5, 2.5 and 19.8%

[45]. England et al (2010) did not see any significant correlations between parents (n=24) and offspring (n=24) with respect to the five semen parameters studied (percentages of normal motility, percentage morphologically normal sperm, sperm concentration, total sperm output or total number of live morphologically normal sperm in a group of normal dogs with no inbreeding present). Moderate narrow sense heritability (h2) was found for percentage normal motility (h2= 0.57) and percentage live morphologically normal sperm

(h2=0.21), and total number of live morphologically normal sperm (h2=0.47) suggesting that breeding stock should be chosen with these parameters in mind.

These two studies suggest that genetics may play a part in semen quality potential. Age is also thought to impact semen quality with young (< one year of age) [46] and old dogs having poorer semen quality due to sexual immaturity and degenerative changes in the testes (testicular neoplasia and decreased testosterone production) [46, 47], respectively. However, Peters et al (2000)

16 were unable to find differences in spermatogenic changes in the testis with respect to age in otherwise healthy dogs [48] . Breed effects on semen quality mainly concern total sperm output and varies greatly with size of the testes, meaning that larger dogs with larger testes produce larger number of sperm [44].

Semen quality can be negatively affected by the absence of a bitch in heat and unfamiliar and stressful environments that can influence libido [44]. Frequency of collection in the dog has been shown to result in optimal semen quality when done once every two to five days [49]. Collection approximately one hour apart results in a 70% decrease in total sperm numbers [50]. Although, no literature exists on the effect of sexual abstinence on the quality of semen collected, decreased sperm motility and lower numbers of morphologically normal sperm may be present due to sperm cell senescence. As many factors have a role to play in semen quality it is important to minimize the effects of those that can manipulated and taking them into consideration when performing semen evaluation and investigating possible causes of altered semen quality in the dog, as well as in other species.

3.1.4 Methods of Semen Evaluation

Several methods of semen analysis are available, ranging from simple light microscopy to computer assisted semen analysis (CASA) techniques using various staining techniques to ultrastructural evaluation using electron microscopy [51-53]. The three main properties of sperm commonly used for semen evaluation are concentration, motility and morphology. Although

17 evaluation of these qualities does not give a complete picture of the fertilization potential in every scenario, it can provide an adequate screening technique for sperm defects. However, to the average veterinarian in reproductive practice, light microscopy remains the most common technique when evaluating canine semen [54, 55]. For research purposes, computer assisted techniques have the advantage of standardization of evaluation, objectivity, repeatability, accuracy and precision that cannot be achieved otherwise [56].

Determination of Sperm Concentration.

Measurement of sperm concentration in itself is not a measure of sperm quality as ejaculate volume is dependent on the accessory sex gland secretion, which in turn is influenced by time of year of collection, duration and intensity of teasing, and frequency of collection in species such as the horse [57, 58]. In the dog, semen volume results from prostatic secretions and depends chiefly on the collector and separation of the second fraction from the rest of the ejaculate with total volumes ranging from one to 80 millilitres and concentrations ranging from 4 to 400 million per millilitre [51]. England et al (1999) collected semen from a mixed group of 65 dogs twice, approximately one hour apart and compared the volumes of the second fraction [50]. First ejaculate second fraction volumes averaged 1.9ml with a standard deviation of 1.3ml. They reported a significant decrease in second ejaculate second fraction volume (mean=1.7ml and standard deviation=1.2ml) as well as concentration and total sperm numbers in the study population, with German Shepherd Dogs having a significantly higher second

18 collection volume, concentration and total sperm numbers than other breeds.

Wildt et al (1982) also observed a decrease in second fraction volume in inbred dogs [59] although this finding was not statistically significant. Both these studies showed that volume of the ejaculate can be highly variable without an impact on reproductive parameters. However, by measuring volume and concentration, total sperm numbers can be determined by multiplying the two values together.

Total sperm numbers are dependent on scrotal width and grams of testicular tissue [44], which explains the discrepancy in sperm output between large and small dogs. Ultimately, total sperm number is one measure of testicular and reproductive function in the male.

Traditionally, sperm concentration is determined using a counting chamber with a grid network i.e. Neubauer or Improved Neubauer chamber, standard dilution of the semen sample and the use of a dilution media that kills or immobilizes sperm (Unopette™, Becton-Dickinson) [54, 55]. Other electronic counters such as densimeters and computerized assisted semen counters (within

CASA systems) have been developed, however, all are standardized using the traditional hemocytometer method. Schafer-Somi and Aurich (2007) compared concentrations of dog semen measured by hemocytometer, SpermaCue® and

SpermVision® analyser and found that they are all highly correlated with a p- value < 0.01 with no significant difference between the methods studied or with dilution using different extenders [60]. The ease of use of the latter methods mentioned has allowed for quick and easy determination of sperm numbers in large semen handling and processing facilities, where speed is essential.

19 However, the gold standard remains the hemocytometer method. A review of the physics behind sample flow in capillary loaded slides in CASA systems compared to hemocytometer methods explains differences in concentration measurements due to the Segre-Silberberg effect [61]. In the shallower and longer chambers used in CASA (20 µm) compared to the hemocytometer (100 µm) different velocity gradients due to capillary action are created and result in increased sperm at a defined distance from the chamber walls. It is then necessary for a compensation factor to be applied for higher correlation between the two methods (r2=0.936 compared with r2=0.984) [62]. This should be taken into consideration when using CASA for sperm concentration measurements.

Evaluation of Sperm Morphology.

Sperm morphology has been used as a measure of reproductive competency and fertility in multiple species [63, 64] and normal sperm percentages in an ejaculate should range in excess of 80% [51]. A 2008

Swedish retrospective study by Al-Makhzoomi et al (2008) investigated semen collected (107 ejaculates) and fertility history of 12 proven bulls used for artificial insemination over a period of 22 months of progeny testing [65]. Fifty-six day non-return rates (NRR) were corrected (CNRR) for several factors such as season, inseminator, estrous control program, breed and parity and used as a measure of bull fertility. The investigators found percentage of normal morphology was positively correlated to fertility (CNRR) (r=0.22, p<0.05).

Percentage of head abnormalities was negatively associated with CNRR (r=-

20 0.23, p<0.05), pear-shaped head defects were negatively associated with NRR and CNRR (r=-0.55, p<0.05), loose sperm heads with NRR and CNRR (r=-0.32, p<0.01), and sperm heads of variable size and NRR (r=-0.27, p<0.05). In the same study, spermatozoa with double-folded tails were also negatively associated with NRR (-0.21, p<0.05). Other defects associated with infertility include those located in the midpiece as well as the presence of proximal droplets [66]. Enciso et al (2011) were also able to establish a correlation between major sperm defects in bulls (double forms, pear-shaped heads, narrow base of head, small abnormal heads, free heads, tail stump, whip tail, proximal droplets, and midpiece ‘Dag’ defects) and DNA damage by validating a sperm chromatic dispersion test and comparing it to sperm morphology assessed by eosin-aniline blue staining techniques under light microscopy [67]. Assessment of sperm morphology therefore can be considered a marker for reproductive potential and DNA integrity.

An early study in a group of 42 bitches bred to 67 stud dogs (34 different breeds), compared number of pregnant versus non-pregnant bitches inseminated with differing percentages of normal sperm (50%, 55%, 60%, 65% and 70%) using the &2 statistic and the greatest strength of association (I coefficient) to determine the limit beyond which fertility was adversely affected [68]. Those dogs that had greater than or equal to 60% normal sperm (using both light and electron microscopic techniques and the sperm morphology criteria as described by Blom (1973) [69]) had better pregnancy rates than those with less than 60% normal sperm (&2=4.54, I coefficient = 0.42)) after correcting for those bitches

21 proven to be infertile (n=4) [68]. As well, percentage of head defects was highest in the lower fertility group (<60% normal sperm) in addition tail defects, proximal droplets and other midpiece defects were also associated with this group.

However, the statistical significance of the individual sperm defects is invalidated as the investigators decided (in a non-repeatable way) which abnormalities were most significant in the case where multiple abnormalities were present or which was the most predominant defect amongst those of equal of significance. In further support of morphological assessment of sperm in dogs, Rijsselaere et al

(2007) took breeding histories of dogs and divided them into three groups: fertile

(produced at least one litter in the three months leading up to the study), subfertile (low pregnancy rates of <50% and/or dogs were unable to fertilize bitches which became pregnant when mated to other dogs) or not used for breeding yet [70]. Fresh semen was collected and stained using eosin-nigrosin staining technique and compared to the breeding histories obtained. Fertile dogs had significantly (p<0.05) higher percentages of normal sperm (meanrSD, 63.3% r 28.5) than subfertile (meanr SD, 29.4% r 29.9) dogs. Difficulty in assessing fertility in dogs is challenging due to the limited number of breedings for dogs compared with other species such as cattle and that conception failure is often due to poor breeding management in the bitch [71]. Therefore the arbitrary nature of the categorization of fertility from this study may not be an accurate representation, although the results are consistent with those found in cattle.

Pena et al (2007) showed significantly decreased zona binding, compared with the control dog, in a Basque Shepherd dog that failed to impregnate three

22 bitches of which two became subsequently pregnant after being mated with another male [72]. Of the 98% abnormal sperm found on semen analysis, 88% of these had proximal droplets and transmission electron microscopy revealed no head, acrosomal or neck defects. This last study specifically pinpoints an individual defect associated with decreased fertility as measured by a fertility assay. The information from multiple species including dogs supports the assessment of sperm morphology as a measure of reproductive potential and fertility in the male.

Sperm morphology is traditionally evaluated by identifying defects related to head, midpiece and tail region under light microscopy using special staining techniques and/or phase contrast under high power (1000X magnification) oil immersion. Among the stains commonly used are modified Wright’s-Giemsa stains (DiffQuik£ or Protocol¥) and eosin-nigrosin, the latter also known as a vital/acrosome differential stain. In eosin-nigrosin stained specimens, live, normal spermatozoa are unstained and appear white against a dark purple background with a well defined smooth apical ridge indicating a non-reacted normal acrosome while those with acrosomal damage appear darker or pink with the apical ridge absent. Other useful vital stains use combinations, such as;

Trypan blue and Giemsa or Trypan blue, Bismark Brown and Rose Bengal stains

[52]. These staining techniques are more time consuming to use, however, they are useful in detecting specific acrosomal defects. The use of these stains is limited to evaluation of fresh semen, as many of the cryopreservatives used in fresh-chilled and frozen semen have been shown to be incompatible with each

23 other [52]. Morphometric analysis using CASA is most beneficial to evaluation of the sperm head; mainly the dimensions of length (L) and width (W), and measures of head perimeter (p). From these measures the area (A), ellipticity

(L/W UXJRVLW\ ʌA/P2), elongation [(L - W)/(L + W @DQGUHJXODULW\ ʌLW/4A) of the sperm head can be determined [73, 74]. However, normal reference ranges with standardized classification systems have not yet been established based on these parameters.

Fluorescein staining techniques may be used in conjunction with CASA.

The advantages of these techniques are that 1) cryopreserved semen can be evaluated without interference from the media used in preservation of samples,

2) proportions of live-dead sperm can be determined, 3) acrosomal integrity, capacitation status, mitochondrial function, chromosome fragmentation and DNA content can be determined [52]. These characteristics of sperm are detected through fluorescein staining used in conjunction with fluorescence microscopy and/or flow cytometry. The disadvantage of these systems is the cost of equipment and necessity for standardization and validation, so they are not routinely used in clinical practice [56].

Morphological defects can be grouped as primary or secondary, referring to whether it is a defect of spermatogenesis or occurs during epididymal maturation and transport, respectively, or after ejaculation due to environmental or processing effects. Primary sperm defects that are associated with poor fertility include midpiece defects. Proximal droplets associated with poor fertility can be considered primary or secondary depending on whether maturation of

24 sperm or hidden midpiece defects are the source of the abnormality [52]. This particular classification, however, does not always determine the severity of the defect and whether it affects the fertility of the whole ejaculate.

The concept of compensable versus non-compensable sperm defects has become a more acceptable form of classification. It is based on the notion that by increasing numbers of normal sperm within an ejaculate, the chances for fertilization are increased [66]. For example, increasing the total number of normal sperm in the ejaculate could compensate for those sperm defects that do not permit binding and/or penetration of the zona pellucida. This would lead to increased numbers of normal sperm that can fertilize ova, thereby increasing the chances of fertilization. On the other hand, defects that allow for normal fertilization of the ovum but have their impact on embryo development would be considered non-compensable. This is due to the fact that the percent chance of fertilization of these types of defects does not change with increasing sperm numbers.

CASA and Sperm Motility.

Computer assisted sperm analysis (CASA) is becoming the gold standard in semen analysis since the digitization of data allows for increased objectivity, precision and accuracy of the parameters measured and the potential for decreasing inter- and intra-laboratory variation through validation and standardization of analysis [56]. Briefly, CASA consists of a microscope connected with a video camera with a video frame grabber card (50-60

25 frames/sec) that inputs data as pixels and data is then digitized into a computer with specialized software to measure concentration, motility and/or morphology of sperm. Most CASA systems use phase contrast/dark field microscopy that illuminates sperm as white against a black background making it possible to identify the centroid (determined by Fourier analysis as the centre of gravity of an object [75]) of the sperm head (pixel dimensions can be set for different species with identification of a tail) for tracking purposes, while other systems use fluorescent dyes (concentration and morphology studies) for identifying sperm.

Software algorithms, through identification of the centroid, identifies successive fields by predicting the probability of the zone within which the sperm is likely to move, determines the minimum number of centroids needed for analysis within a given timeframe, determines the minimum distance between video frames that indicates the sperm is moving and determines the number of forward video frames to be looked at to find a missing centroid [76]. In this way the CASA system is able to track and record data from multiple individual sperm trajectories.

Although not without issues such as agreement among research laboratories regarding the validation and standardization of parameters used in the software programs of different systems, CASA still holds promise in both human and veterinary clinical practice. The parameters of concentration, motility and morphology can be assessed; however, there is little information to be found in the literature on validated computerized morphological assessment of sperm in veterinary research [56, 77]. Using any kind of CASA system requires that the

26 sample be diluted. Iguer-Ouada (2001) and Verstegen (2002) found that motility parameters were not affected between the concentrations studied (50-300 X

106/ml), however, accuracy and repeatability of the measures were affected at concentrations below 50 X 106/ml [56, 77, 78].

Some systems have been validated for dog semen such as the Hamilton-

Thorne sperm analyzer [78] and Sperm Vision® [60]. It has been shown that type of specimen chamber, temperature, frame rate, number of fields analyzed, sperm concentration (see above) and semen handling techniques and extenders can all have an effect on the measurements of concentration and motility [56, 60,

77]. It is recommended that a minimum number of 100 cells should be analyzed for results to be valid, especially in oligospermic samples, as larger numbers of cells analysed reduces the coefficient of variation and increases the precision of the results [56]. High numbers of sperm per examined field can interfere with estimations due to the affect of collisions, sperm exiting the field, motion in the wake of moving spermatozoa and the ability of the system to differentiate and register individual cells. Misidentification of foreign particles or debris can also occur especially when using semen extenders with egg yolk.

The main motion parameters measured in most CASA systems include the following: 1) Straight line distance (DSL, µm), 2) Curvilinear distance (DCL,

µm), 3) Mean distance (DAP, µm), 4) Curvilinear velocity (VCL µm/s) which is the instantaneous velocity of the spermatozoa along its path, 5) Linear velocity (VSL

µm/s) the velocity of the spermatozoa along its straight line path, 6) Mean velocity (VAP µm/s), 7) Straightness (STR%), a ratio of the VSL/VAP which gives

27 the straightness of the path taken as a percentage of the optical straightness

(100%), 8) Linearity (LIN = VSL/VCL) which corresponds to how close the cell travels along to the straight line path, 9) Wobble coefficient (WOB=VAP/VCL) or the oscillation of the curvilinear path along the mean path, 10) Amplitude of lateral head displacement (ALH µm), 11) Average orientation change of sperm head (AOC, degrees), and 12) Beat cross frequency (BCF Hz) or the frequency with which the sperm head crosses its average pathway [60]. All these parameters give a detailed description of sperm motion and are ultimately used as tools in measuring fertility potential and reproductive health in the male and assessing the sperm motion necessary to travel through the female tract to reach the oocyte and fertilize it.

In a study by Farrell et al (1987) of 11 artificial insemination (AI) bulls ranging in age between 6 and 11 years, sperm motility parameters from 44 total ejaculates (211 956 total services) measured by CASA were compared to 59- day non-return rates in cows and adjusted for whether semen was used in heifers or cows or whether the females were enrolled in Dairy Herd Information or not

[79]. The results obtained showed significant positive associations between fertility and the motility parameters of total motility, progressive motility, ALH,

VAP, VCL, VSL, BCF, LIN using multiple linear regression modeling and achieved r2 values ranging from 0.34 to 0.98. It is important to note that non- return rates between herds were also accounted for in order to increase the accuracy of the estimates of fertility. The high r2 estimates from this study were produced using combinations of motility parameters while only total motility in

28 univariable analysis with fertility had significant correlation alone. It is apparent that fertility is dependent on the collective nature of these parameters in this species.

Love et al (2011) looked at several fertility measures in the stallion

(percent pregnant per season, percent pregnant per first cycle and percent pregnant per cycle) and motility parameters (total motility, progressive motility,

VAP and VSL) and found significant correlations with total motility and progressive motility with all three fertility parameters [80], however, significance of other motility parameters varied amongst the three fertility parameters measured. The most sensitive measure of fertility in the stallion in this study was determined to be percent pregnant per first cycle and percent pregnant per cycle when compared to seasonal pregnancy rate. This study in particular shows the importance of definitions of fertility and the difficulty in assessing outcomes from

CASA motility parameters. Care should be taken when interpreting those findings and correlating them across studies and species.

CASA has importance in in vitro fertilization (IVF) and embryo transfer as a tool to overcome subfertility in humans. A study looking at differences in sperm motility between good (fertilization rate >50%) and poor (fertilization rate d 50%) fertilization rates found that significant associations exist between these two groups and the motility measurements of ALH, VCL, and VSL [81] but not BCF,

VAP, LIN, and STR. Earlier, Aitken et al (1985) attempted to determine which characteristics of sperm motion allowed it to penetrate cervical mucus. The study measured the motion characteristics of sperm and determined bovine cervical

29 mucous sperm concentrations after incubation with a known volume and concentration of sperm. The authors determined that a large ALH was necessary for cervical mucus penetration and movement in the bovine [82]. These findings were further supported by studies showing that in human IVF, larger ALH means were correlated with higher fertilization success rates and were attributed to an ability to penetrate the cumulus oophorus and zona pellucida of the oocyte [83,

84]. ALH would appear to be necessary to both in vivo movement through the barriers encountered in the female reproductive tract and through the barriers encountered with the oocyte in IVF. In the dog, it has been found that most of motility parameters measured by CASA had correlation to fertility except for BCF,

LIN, and STR measured by the Hamilton-Thorne semen CASA [70]. Fertile dogs in this instance were defined as those dogs having produced one litter in the three months leading up to the study and sub-fertile if pregnancy rates were below 50%. A significant correlation (r=0.44) between percentage of morphologically normal sperm and percentage progressive motility has been shown in the dog [85].

In general the motility parameters that have consistently been associated with fertility across species include total and progressive motility, ALH, and VSL and may explain why these parameters are most often studied with relation to fertility. However, it is important to note that in most studies multiple interactions with other motility parameters are found, suggesting that fertility is correlated to combinations of motion characteristics necessary for sperm to travel to the female oocyte. In this way CASA results should be interpreted according to

30 whether natural breeding/insemination or assisted reproductive techniques are to be used. Although CASA measures motility in an artificial environment it does not measure how motility is affected within the female reproductive tract, nor does it evaluate the genetic competency or fertilizing ability of sperm. Although, correlated with fertility, motility measured by CASA is merely one piece of a complex puzzle and cannot account for the multitude of factors affecting fertility of a male.

3.2 THE CANINE PROSTATE GLAND

3.2.1 Anatomy and Physiology of the Prostate Gland

The dog has only one accessory sex gland, the prostate gland. As a result it contributes over 97% of the ejaculate fluid [1]. The prostate is positioned retroperitoneally in the pelvis and surrounds the urethra near the neck of the bladder. The rectum is located dorsally and the symphysis pubis ventrally to the gland. Its craniocaudal limits are variable and dependent on the size of the gland; which increases normally with age, and can therefore be present in a completely pelvic position or abdominally [86]. It has a symmetrically bilobed structure with a fibrous tissue capsule. It is supplied by the prostatic artery, a branch of the internal pudendal artery and drains via the prostatic and urethral veins. Nervous innervation from the parasympathetic branches of the pelvic nerve stimulates glandular secretion while sympathetic innervation causes contraction of smooth muscle and emptying of the gland through multiple

31 ductules into the pelvic urethra, on the ventral aspect of the gland, near the openings of the vasa deferens in the colliculus seminalis [8, 86].

Histologically the prostate is separated into stromal and glandular parts.

The stroma is made up of a network of delicate fibrous connective tissue strands, smooth muscle, blood vessels and nerves that are continuous with the capsule, extend throughout the substance of the gland and coalesce near the urethra.

The glandular part consists of columnar epithelial cells and is organized into tubuloalveolar lobules that empty through a duct system into the urethra through several small openings on the colliculus seminalis, as previously described [8,

86].

Prostatic secretions contain components that nourish and maintain sperm such as fructose, as well as other constituents that provide a mechanical transport medium through which sperm travel through the female reproductive tract. In addition, prostatic secretions have anti-bacterial properties [87].

Selected prostatic components and their normal ranges are listed in Table 1.1.

The main protein secreted by the prostate is canine prostate-specific esterase which accounts for 90% of the total protein secretion by this gland [88]. Although this enzyme differs in amounts between normal and diseased prostatic tissue it cannot be used to differentiate among major prostatic diseases [88]. It is still unknown what role this enzyme plays in male dog reproduction.

32 Table 1.1. Selected Physical and Chemical Analyses of Prostatic Fluid in Dogs with Healthy Prostates

Mean r SD (range) Sample Size pH 6.2 r 0.3 (5.5-7.1) n = 43 Specific gravity 1.018 r 0.005 (1.008-1.028) n = 40 Cholesterol mg/dl 27 r 17.0 (8.0-73.0) n = 29 Zinc (Pg/ml) 62.3 r 35.3 (10.3-120.6) n = 20 Copper (Pg/ml) 7.1 r 4.8 (1.3-19.5) n = 20 Iron (Pg/ml) 0.7 r 0.5 (0-1.6) n = 20 Magnesium (Pg/ml) 16.4 r 9.5 (3.4-40.0) n = 20 Data adapted from Branam et al [89]

The prostate is an androgen-dependent organ as evidenced by complete atrophy of the gland following castration [86]. It is dependent mainly on the steroid hormone dihydrotestosterone (DHT) which is converted from testosterone by the enzyme 5D-reductase produced within the gland. In fact, re-growth of the atrophied gland can be achieved via supplementation with a combination of exogenous estradiol (E2) and DHT [90]. DHT is also the main hormone responsible for the differentiation and development of the gland from the urogenital sinus during embryogenesis [8].

3.2.2 Pathophysiology of Benign Prostatic Hyperplasia (BPH)

BPH, as the name implies, is a benign enlargement of the prostate gland that is due to hyperplasia, as well as hypertrophy, of mainly prostatic epithelial cells in the dog. There appears to be no breed predilection for BPH [91]. Early

33 studies of Beagle dogs have shown maturity of the prostate, according to histological pattern of developed alveoli and maximum size, occurs by 1.5 years of age with a remarkable three-fold increase in prostatic size compared with immature dogs. One study suggested that starting as early as 2-3 years of age and by 4 years of age >40% of dogs have histological evidence of BPH with 80% and 100% of Beagles having confirmed BPH by 4 and 7 years of age, respectively [1]. However, this study was limited to Beagles and may not have relevance in other breeds. A complex version of BPH has also been described in which dilation of ducts and the presence of cysts and inflammatory cells occur with degeneration of epithelial cells [1]. Although the same study also correlated a decline in secretory volume with age and state of the prostate, many factors play a role in ejaculate volume that can be difficult to control for. A definite correlation between prostate size, age and weight has been proven in multiple studies [47, 92-94].

Androgen:estrogen ratio appears to play a significant role in the development of BPH. This theory is supported by reported declines in serum testosterone and DHT concentrations while 17E-estradiol concentrations remain unchanged [1]. Using histology to identify immature, normal and BPH tissue

(defined as having both hyperplastic and hypertrophic elements) and measuring individually pooled serum hormone levels using radioimmunoassay in 42 beagles

(8 months to 9 years of age) over three weeks, Brendler et al (1983) were able to compare mean serum hormone levels amongst the three prostatic groups. They also compared mean testosterone and E2 with age and found a general trend of

34 decreasing testosterone (with no changes in mean E2) that followed histological changes and weight increases in the prostate suggesting altered androgen metabolism in prostatic hyperplasia/hypertrophy. It is important to note that statistical analysis was not undertaken in this study and significance of these results is unknown. Experimentally induced BPH requires the administration of estradiol and DHT, rather than DHT alone [95]. Castrated and control dogs were divided into groups receiving different hormone treatments given three days a week for a total of 40 weeks after prostate size regressed at 4 weeks post- surgery. Testosterone and DHT alone did not produce BPH while E2 alone caused a squamous metaplasia of the gland. In contrast, the combination of E2 and DHT but not E2 and testosterone caused hyperplastic and hypertrophic changes in the prostate. This information further suggests androgen:estrogen ratio to be of significance. It is likely that although DHT and testosterone serum levels have been shown to be decreased in dogs with BPH, they are still necessary in the pathophysiology of BPH. This may be due to increased uptake and utilization of testosterone and DHT in prostatic tissue causing a decrease in peripheral serum concentrations of these hormones.

A study investigating enzymes of steroid hormone metabolism within the prostate in normal young (age range 17-40 years) and aged men (age range 60-

80 years) showed that aged men with or without BPH had decreased prostatic tissue enzyme activity of 3D(E) hydroxysteroid dehydrogenase (HSDH) compared with normal young men [96]. The reduction in enzymatic activity suggests that altered androgen to estrogen ratios found in BPH are due to decreased

35 degradation of stromally-produced dihydrotestosterone in BPH, rather than an increase in DHT production, which might occur through conversion from testosterone under the influence of increased 5D-reductase activity. The mechanisms which lead to alterations in HSDH activity in BPH have not been elucidated.

It has been suggested that lack of degradation of DHT mediated by estradiol correlates with an abundance of estrogen receptors (ER) present in healthy and hyperplastic prostatic stromal and epithelial tissues [97, 98].

Compared to normal prostatic tissue there is a significant decrease in the percentage of ERs in hyperplastic and neoplastic canine prostates [98]. These findings suggest a more indirect role of estrogens in the pathology of BPH. It has been postulated that free radical production by increases in relative estrogen concentrations cause tissue damage by which DHT induces abnormal prostatic growth [99]. Interestingly, progesterone receptors (PR) were also identified for the first time in diseased conditions of the prostate suggesting as yet an unknown role of progesterone in BPH development [98].

Similarly to the testes, biological effects of testosterone and estrogen in hyperplastic prostatic tissue may be mediated through several growth factors

[100, 101]. The most important of these growth factors include fibroblast growth factors (FGF) of which keratinocyte growth factor (FGF-7) appears to be the most potent mitogenic factor in hyperplastic tissue [102]. IL, IGF and TGFE appear to be important in BPH [100]. These autocrine mediators between stromal and

36 epithelial tissues are considered to be responsible for the increase in prostatic growth seen in BPH.

Using ELISA and comparing results from normal (peripheral and transition zones) and hyperplastic human prostatic tissue, Ropiquet et al (1999) were able to assess whether over-expression of FGF-2 and FGF-7 was present in hyperplastic tissue [103]. They found a significant increase in both FGFs compared to normal prostatic tissue. In addition, they investigated the effect of

FGF-7 on epithelial and FGF-2 on stromal growth in prostatic tissue based on results from other studies showing specificity of these FGFs for distinct cell populations within the prostate [104, 105]. The results showed significant increases in total epithelial cell numbers when treated with FGF-7 compared with controls as well as significant increases in stromal cell numbers when treated with FGF-2. ILs have been shown to stimulate both these growth factors in culture. IL-1D was observed to be produced by prostatic epithelial cells and stimulates FGF-7 production in stromal cells [106], while IL-8, also produced by prostatic epithelial cells, stimulates FGF-2 in stromal cell culture [107]. These two growth factors appear to play a significant role in the pathogenesis of hyperplasia in humans where both stromal and epithelial cells show abnormal growth. Increases in and irritation of smooth muscle in BPH, as a result of inflammation, are important in the lower urinary tract symptoms (caused by increased urethral pressure) that are seen in men [108]. This clinical sign, however, is not one normally seen in the dog and therefore stromal components may not have the same importance. In the dog epithelial cells are the main cell

37 type involved in the pathophysiology of BPH, although there is a minimal stromal component suggesting a greater role of FGF-7 and not FGF-2 in pathogenesis of

BPH in the dog.

IGFs, and perhaps more specifically, IGF binding proteins (IGFBP) seem to work in conjunction with TGFE in regulation, or lack thereof, in BPH. TGFE has been localized mainly in prostatic epithelial cells and expressed in larger amounts in BPH [109]. Cohen et al (2000) were able to show a 15-fold increase in IGFBP3 in normal stromal cells compared with a two-fold increase in BPH when treated with TGFE [110]. These finding were correlated to a 68% growth inhibition in normal prostatic culture compared to 26% in the BPH culture [111].

IGFs have also been localized in prostatic tissue and found to be up to ten-fold higher in stromal cells from BPH tissue compared with normal tissue [112].

Therefore, growth of tissue in BPH is most likely mediated through multi-factor complex processes where there are not only increases in growth factors but also in inhibitory factors that interfere with normal apoptotic processes. The result is an overall increase in the size of the gland.

Unlike BPH in men, BPH in the dog involves predominantly glandular rather than stromal components [113]. Growth of the prostate proceeds in a peripherally diffuse pattern in the dogs and does not interfere with urinary tract function [1]. Mechanical occlusion of the prostatic urethra occurs in men where localized nodular stromal proliferation in the so-called transition zone near the urethral sphincter causes compression of the urethra and impaired urination

[114]. This is exacerbated by the fixed nature of the prostate. Blood in the

38 ejaculate is the most common clinical sign of BPH in dogs and the outwardly expanding gland rarely causes compression of the rectum and constipation.

More commonly as the non-fixed gland increases in size and weight it assumes a more abdominal position, especially when the bladder is full, and in some instances can be difficult to palpate per rectum.

3.2.3 Conventional Treatment of Benign Prostatic Hyperplasia

There are at present several treatment modalities for BPH, however, side effects and/or relapses post-treatment are common [86]. Due to the benign nature of the condition, treatment is only sought or recommended where clinical signs are present. The treatments for BPH in the dog centre on the hormonal component of the condition. Ultimately, there is down-regulation of testosterone and/or DHT production as these are the major pathophysiological components of the condition. Castration, whether by physical or chemical means, is the most dramatic recourse for complete remission and involution of the gland by eliminating the main source of androgens. This solution however, is not considered to be desirable in the valuable breeding animal or where the risks of surgery/anesthesia far outweigh the benefits. Pharmacological treatments have focused on three modes of action: 1) 5D reductase inhibitors (decrease DHT in the prostate), 2) androgen receptor antagonism and 3) GnRH modulators [86,

91].

The azasteroid Finasteride, a 5D reductase inhibitor, has become the most widely used drug for treatment of BPH in dogs for several reasons. In a double-

39 blinded drug trial, nine dogs with BPH confirmed by having clinical signs of blood in the ejaculate or constipation, were treated with finasteride (five dogs, 5 mg/day

PO for 16 weeks) or placebo (four dogs, 5 mg powdered sugar in gelatin capsule/day PO for 16 weeks) [115]. Using both ultrasound (prostate volume) and radiological (prostate diameter as a percentage of the distance from the sacral promontory to the pubis) measurements of prostate size at 8 and 16 weeks of treatment, the investigators observed a 24% decrease in prostate diameter and a 41% decrease in prostate volume by 8 weeks of treatment that did not differ significantly by the end of the 16 week treatment period. Control dogs experienced no significant changes in prostate diameter or volume at either the 8 or 16-week treatment periods. Serum DHT decreased by 50% in the treatment group by 8 weeks and remained unchanged through the completion of the study while serum testosterone did not change. No changes in semen quality were noted except for a decrease in semen volume that was not significant. It was noted that the five finasteride treated dogs bred successfully during and after finasteride treatment. None of the dogs showed changes in libido or clinical signs of blood in the ejaculate or constipation. All clinical signs i.e. blood in the ejaculate and constipation used to diagnose BPH in individual dogs were alleviated within four and one week(s) after the beginning of treatment, respectively. The successful treatment and alleviation of BPH and its clinical signs in this drug trial, in combination with no evidence of side effects relating to semen quality, fertility and libido have made finasteride the drug of choice in treating BPH in dogs.

40 Treatments involving androgen receptor antagonism have also shown efficacy in decreasing prostate size in BPH. Osaterone and other androgen receptor antagonists have been shown to decrease prostate size in dogs [116].

A clinical trial examined 142 dogs that were treated with either osaterone or delmadinone for 180 days. Osaterone had a significantly greater decrease in prostate volume (38%) compared with delmadinone (28%) with maximum reduction in volume by 14 days of treatment for osaterone. Complete remission of clinical symptoms of BPH occurred in 50% of the dogs by day 14 with 83% remissions by the end of the study period. Major side effects included increased appetite and changes in behaviour in a small percentage of the dogs. In another study by Tsutsui et al (2001) examined semen parameters of beagle dogs during treatment with osaterone and found no significant differences in sperm count, morphology, motility, semen volume or pH [117]. Significant decreases in serum testosterone and LH were observed during the treatment period, however these levels were maintained during treatment and no adverse side effects were observed. Although osaterone has shown good clinical efficacy with little or no side effects it is not currently available as a treatment option in North America.

Progestins, with antiandrogen action, such as delmadinone have also shown similar efficacy in reduction in prostate volume [116]. Side effects of delmadinone treatment include inhibition of secretion of adrenocorticotropic hormone at the hypothalamic-pituitary level, changes in maturation of epididymal spermatozoa, increases in appetite [116], as well as detrimental effects on sperm morphology, motility and sperm numbers at higher doses [91]. These effects do

41 not make it desirable when other pharmacologically safe alternatives are available.

GnRH agonists, such as deslorelin, act by initially causing an increase in

GnRH secretion with subsequent increases in LH/FSH and testosterone. After the initial increase in hormone up-regulation, a negative feedback inhibition of

GnRH ensues causing decreases in LH/FSH and testosterone. Although, deslorelin causes a decrease in prostate volume [118], major side effects on fertility through down-regulation of hormones regulating reproduction in the male does not make it a desirable option in the breeding dog. At the present time no

GnRH antagonists for BPH treatment are commercially available in Canada or the USA.

In summary, only those drugs that provide a decrease in prostate volume and clinical signs of BPH without affecting fertility are desired in the stud dog.

The effects and side effects of the drug treatments mentioned here are reversible with discontinuation of treatment. This holds true for the decrease in prostate volume and therefore long-term administration is required to maintain a desirable prostate size as well as fertility in the stud dog. This has led to the use of azasteroids with inhibitory action on 5D-reductase activity to be the drug of choice in breeding animals.

3.2.4 Ultrasonography of the Prostate

Traditional methods of determining size, symmetry and quality of the prostate gland are subjective, such as digital palpation per rectum, as well as

42 semi-quantitative means using radiography [91, 119]. However, with the introduction of, and advancements in, ultrasonography, an efficient and non- invasive tool for determining prostate size, symmetry and lesions has become available [93, 120, 121]. Due to the pelvic position of the prostate in human males, ultrasonography is performed using a specialized transrectal probe to evaluate prostate volume. Ultrasonographic measurements are highly correlated to volumes measured in cadaver specimens in humans [120].

Although 95% accuracy can be achieved using planimetric measurements with sequential cross-sectional images at 4 mm apart, this is considered far too time- consuming and tedious in a clinical veterinary setting. Acceptable accuracy can be attained by using length, width and height of the gland and using a simple elliptoid equation to calculate volume [120, 122].

In the veterinary setting transrectal ultrasound is not commonly used in small animal evaluation thus specific ultrasound probes for this purpose do not exist. Therefore, ultrasonography of pelvic structures is performed mainly using a transabdominal approach, and is common practice for prostatic evaluation in dogs. As previously described, the canine prostate can have a pelvic and/or abdominal position due to lack of pelvic fixation as seen in men. The greatest difficulty in evaluation of the completely pelvic prostate in dogs is the positioning of the ultrasound probe, which must be angled obliquely on the abdomen towards the pelvic inlet [94, 121]. This makes achieving a true transverse cross section of the gland difficult. However, relatively accurate estimations of prostate size can be achieved as the canine prostate can be manipulated via digital

43 manipulation per rectum without causing distress in the patient [93]. There is little information in the literature on ultrasonography of the canine prostate with respect to correlation of ultrasound measurements versus actual prostate volume. However, both Atalan et al (1999) and Kamolpatana et al (2000) found a high correlation between estimated prostate volume and actual volume measured through caliper measurement of length, width and height using a modified formula for the volume of an ellipse (R = 0.76) or through the use of water displacement technique (R2=0.94) of the dissected prostate glands [94,

121]. As a result, it has been shown that ultrasonography is a fairly accurate method for evaluation of prostate volume and for monitoring size changes during treatment for prostate disease [121] and more specifically BPH.

3.3 VITAMIN D AND ITS ROLE IN MALE REPRODUCTION

Currently, the non-traditional roles of Vitamin D have become important in cancer research due to its ability to affect the normal life cycle of cell populations within multiple tissues ensuring that natural cell death, or apoptosis, occurs within these populations [123]. Although BPH is not categorized as a cancerous process, it shares similarities in that cell proliferation exceeds that of cell death leading to an increase in both the number and size of prostatic cells. This unchecked rate of growth, albeit a non-fatal one, has an effect on susceptibility to infection [91], as well as a secondary impact on reproduction. Treatment can be expensive, consequent prostatitis a risk, and the inability to use assisted

44 reproductive techniques, such as semen freezing [124], a concern in valuable breeding animals.

3.3.1 Vitamin D Metabolism

The traditional role of Vitamin D in calcium and phosphate homeostasis

(Ca:P) and bone metabolism was first elucidated by Edward Mellanby in 1918 while researching rickets in children [125]. Unlike humans, dogs do not synthesize Vitamin D through the action of ultraviolet radiation on the skin [126,

127]. Therefore, dogs rely heavily upon dietary intake of Vitamin D. The majority of commercial dog foods meet, and more frequently exceed, the minimum daily requirements for this vitamin (500-5000 IU Vitamin D/kg food/day) set out by the

National Research Council and Association of American Feed Control Officials

[126]. In some cases commercial dog foods come close to the maximum allowable amounts beyond which toxicity occurs. Toxicity may manifest as cardiac arrhythmias, stiff joints and renal disease [128].

Vitamin D is a fat-soluble vitamin, and is ingested from the diet as Vitamin

D3 from animal sources such as liver and fish oils. Hydroxylation to the more active form, 25-hydroxycholecalciferol (25OHD3), occurs via 25-hydroxylation in the liver, both directly in response to decreased blood calcium and phosphorous concentrations and indirectly through an increase in Parathyroid Hormone (PTH) when decreased blood calcium concentrations occur [126]. Further conversion of

25OHD3, occurs via renal 1D-hydroxylase activity to 1,25(OH)2D3, also known as calcitriol. Calcitriol works directly to increase intestinal absorption and renal

45 reabsorption of calcium and phosphorous and finally to dissolve both components from bone reserves [126]. Perhaps the most important and significant role of Vitamin D through this pathway is the prevention of rickets, a condition affecting growth in the young that leads to softening and deformation of developing bone tissue [125]. Alternatively, excess serum Ca and P triggers hydroxylation by the kidney to 24,25(OH)2D3, resulting in deposition of Ca and P in bone [125].

In recent years, emphasis on Vitamin D research has grown owing to its association with decreased cancer mortality risk [125, 129, 130]. Garland was the first to show an association between decreased risk of colorectal cancer and geographic latitude [130, 131]. This stimulated further study into the anti-cancer effects of Vitamin D. As a result, the non-traditional role of Vitamin D has come to light; mainly its anti-proliferative effects and its role as a mediator of cell differentiation and apoptosis in multiple tissues [132, 133]. Identification of

1,25(OH)2D3 and the enzyme responsible for its formation, 1D-hydroxylase, ubiquitously throughout human tissues, including the prostate gland [134] encouraged further research into the role of Vitamin D. These tissues include human and rat kidney, stomach, large intestinal epithelium, mammary gland, ovary, prostate gland, and pancreatic duct cells [135]. However, studies are lacking with respect to these extra tissue effects of Vitamin D in the dog.

Tissue concentrations of 1,25(OH)2D3 are dependent on the circulating serum 25OHVD3. Deficiency, or more importantly insufficiency, in circulating

25OHD3 results in a lack of Vitamin D effects in individual tissues through

46 decreased binding to the Vitamin D receptor (VDR) [135]. Consequently, there is a break in the chain of inhibition or induction of transcription of specific genes related to regulation of the cell cycle (proliferation, invasiveness, metastatic potential, differentiation and apoptosis) and angiogenesis, all important factors in cancer physiology [123].

The VDR is a cytosolic receptor that binds with calcitriol to form a complex that further binds with the retinoid X receptor (RXR) to form a heterodimer. The formation of the heterodimer is obligatory for translocation into the nucleus and binding to the VDR response elements (VDRE) of certain genes [133]. The genes in question can respond in three different ways: 1) bind to promoter regions of certain genes, 2) bind to negative VDREs (nVDRE) or 3) antagonize other transcription factors. As a result, multiple actions involving genes with protein actions involved in bone remodeling, calcium binding, metabolism, adhesion, anti-proliferation and differentiation may be achieved, with emphasis on those that are anti-inflammatory and anti-proliferative in nature. The VDR regulates numerous gene sequences. For instance, up-regulating gene transcription of osteocalcin or down-regulating gene transcription of PTH [132] stimulates osteoblasts to increase bone formation or allows for increased renal elimination of Ca and P, respectively. The VDR-RXR heterodimer can also act independently by directly inhibiting transcription factors such as the two nuclear factors (NF): NF-AP and NF-NB. The regulatory role of Vitamin D on certain genes and their effects are summarized in Table 1.2.

47 Table 1.2 Genes influenced by Vitamin D receptor ligands and their effects.

Gene Protein Anti- Anti- negVDRE Function Inflammation Proliferation Osteocalcin Bone matrix protein Osteopontin Bone matrix protein RANKL Bone IL-2 EGF-R PTH remodeling CA II Bone IL-12 c-myc PTHrP remodeling Calbindin-9k Calcium TNF-D K16 Rel B binding 24- Metabolism IFN-J hydroxylase mCYP3A1 Metabolism GM-CSF mCYP3A11 Metabolism hCYP3A4 Metabolism E3 integrin Adhesion P21 Anti- Proliferation Involucrin Differentiation PLCJ1 Differentiation IGFBP-3 Anti- Differentiation Adapted from Nagpal et al 2005[133]

LEGEND IL = Interleukin TNF = Tumor Necrosis Factor IFN = Interferon GM-CSF = Granulocyte Macrophage Colony Stimulating Factor IGFBP = Insulin-like Growth Factor Binding Protein PLC = Phospholipase C EGF-R = Epidermal Growth Factor Receptor

48 The VDR has also been identified in multiple human tissues such as epithelial cells of the epidermis, hair follicles, the female reproductive tract, mammary gland, colon and lung; endocrine cells of the thyroid, pancreas and ovary; in cardiac muscle cells; adipocytes and in cancer cell lines [132]. More importantly, with regards to male reproduction, it has been found in rat and human testicular tissue, seminiferous tubules and individual spermatozoa, along with its associated activation enzymes [136-141]. VDR has also been isolated and identified in rat and human prostatic tissue [139, 142, 143]. With regards to female reproductive health, female calcitriol deficient mice experience hypoplasia of the uterus and ovaries [144].

3.3.2 Vitamin D and the Prostate

Prostate cancer in men has been given high priority in the research literature [4]. Recent cancer statistics reveal that prostate cancer is the second leading diagnosed cancer in men and its mortality rate is second only to lung cancer [145]. An American report in 2000 showed three out of four men had chronic symptoms of BPH in their seventh decade while 6.5 million out of 27 million between the ages of 50 to 79 years sought treatment of symptoms [146].

The first strong epidemiological evidence linking Vitamin D to cancer risk, through a negative association with sunlight exposure, was found by Garland et al while looking at colorectal cancer in people [147]. In 1990, Schwartz and

Hulka (1990) hypothesized a similar relationship between prostate cancer and geographical location showing a negative correlation to amount of ultraviolet

49 radiation exposure [148]. Hanchette and Schwartz (1992) later found evidence to support this hypothesis while examining prostate cancer risk and geographical distribution patterns in the ethnic white male population of the United States of

America [130]. This discovery resulted in extensive study of Vitamin D and its role in human prostatic disease mainly prostatic carcinoma and later BPH [4].

In humans normal, cancerous and hyperplastic prostatic tissue have been shown to have the VDR, 1D hydroxylase activity and 1,25(OH)2D3 intracellularly

[142]. Indeed, human prostate tissue itself, independent of kidney 1-hydroxylase activity, can synthesize its own intracellularly active 1,25(OH)2D3 from 25OHD3

[134]. However, in one human prostatic cancer cell line it was revealed that 1D hydroxylase activity was absent and no anti-proliferative effects of Vitamin D were noted. Furthermore, this study showed concentrations of 1,25(OH)2D3 in normal and BPH tissue to be comparable to kidney tissue [134], suggesting an adaptation might exist in some prostatic carcinomas to avoid the anti-proliferative effects of Vitamin D by removal of this enzyme.

Prostatic carcinoma is relatively rare in both intact and neutered male dogs with approximately 4% of the male dog population having subclinical evidence of carcinoma according to the South African study by Mukaratira et al

(2007) [149]. BPH in this species has been accepted as having a similar prevalence rate to that of men; greater than 80% in intact male dogs over the age of four years in a review by Memon (2007) [150]. In an early study by Brendler,

BPH was confirmed histologically in 88% of beagles over the age of six years [1].

Atalan et al (1999) found 36/60 (60%) of the dogs in an ultrasound study of

50 prostate volume to have histological BPH while the rest of the dogs were diagnosed by histology with normal prostates (17/60 or 28%), prostatic neoplasia

(4/60 or 7%) and prostatitis (3/60 or 5%) [94]. However, there is a lack of prevalence studies using histology to diagnose BPH in the general intact dog population and prevalence is based heavily on the findings in early beagle studies and anecdotal evidence [151]. The limitation of prevalence studies is due to the lack of invasive techniques used to definitively confirm non-clinical BPH.

The benign nature of the condition does not justify the costs or risks involved.

There is also a lack of defined prostate dimension parameters for different bodyweights and ages when using ultrasonography to measure prostate size as is present in BPH in men.

Vitamin D studies involving the prostate have concentrated on the effect of the vitamin in cancerous tissue. Due to the risk of hypercalcemic effects that occur with high doses of naturally occurring Vitamin D, several experimental

Vitamin D agonists have been developed which demonstrate high affinity for the

VDR in prostatic tissue without this potential side effect. These studies have shown some promise in reducing PSA concentrations in early clinical trials [134,

152]. However, the effects of Vitamin D on BPH have only more recently attracted attention [4, 153-155].

As stated earlier, BPH is an androgen dependent condition and Vitamin D is thought to work downstream from the androgen receptor (AR) to inhibit production of certain growth factors and pro-inflammatory cytokines [4, 133].

Crescioli et al (2003) studied cultured rat BPH cells using the experimental

51 Vitamin D analogue BXL-353 [153]. They found that, in the presence of 10 nmol/l testosterone, increasing doses of BXL-353 caused a dose-dependent reduction in the number of BPH cells, as a percentage of pre-treatment numbers, after 48 hours incubation. In the same study, BXL-353 AR receptor binding and its ability to convert testosterone to DHT were also studied to determine whether the mechanism of action was through the AR or 5D-reductase activity, respectively. Finasteride-treated cell cultures were used as controls. BXL-353 demonstrated an inability to bind to the AR in the presence R1881 (a synthetic androgen), through a competitive binding assay. No changes in testosterone and DHT concentrations from prostatic cell cultures were observed for the

Vitamin D agonist while finasteride-treated controls maintained previously recorded 50% inhibitory concentration for both isoforms of 5D-reductase. These results indicate an androgen-independent mechanism for reduced cellular growth and cellular apoptosis in BPH cells.

FGF-7 and IGF are implicated in the mechanism by which Vitamin D exerts its action in BPH cells. Crescioli et al (2000) looked at FGF-7 action on human BPH cells in culture [156]. First, they tested FGF-7 treatment alone on

BPH cell culture and found increased cell proliferation that proceeded in a dose- dependent manner. Next they treated these cell cultures with both BXL-353 and native Vitamin D and found an equal and significant decrease in the number of

BPH cells in the FGF-7 treated cultures for both the Vitamin D analogue and

Vitamin D. With this evidence, BXL-353 was further studied to determine whether an effect on FGF-7-receptor signaling was present. They showed a

52 significant reduction in tyrosine phosphorylation of the FGF-7-receptor induced by FGF-7 in culture, thereby inhibiting primary pathways in the stimulation of gene expression relating to cell proliferation. This same group also investigated the effect of the analogue on IGF-1 mediated cell proliferation and found that

BXL-353 was able to achieve this in a dose dependent manner [157]. Therefore, action of Vitamin D and its analogues appear to be mediated through the pathways down-stream of androgen action and target the paracrine factors related to cell growth in prostatic tissue. Crescioli et al (2004) found similar results with the experimental Vitamin D analogue BXL-628 (elocalcitol), yet had less hypercalcemic effect than other Vitamin D analogues [154]. In pre-clinical trials with elocalcitol, Adorini et al (2007) used a small group of male beagle dogs

(n=4) to study the effects of 5P/kg/day of elocalcitol for 9 months [158]. Prostate weights, expressed as a percentage of prostate weight to bodyweight, were reduced in this group, however these values did not reach statistical significance.

This may have been due to the low number of dogs used in the study. Of note, serum calcium levels did not differ between pre-treatment and treatment values.

To date, no other canine studies related to Vitamin D analogues or their effects on BPH are available.

Inflammation has been shown to be a factor in human BPH and may explain many of the lower urinary tract symptoms experienced in men. In canine studies to date, BPH is considered to be without the inflammatory component present in the human form of the condition [1, 151, 159]. The similarities of BPH between men and dogs in the spontaneous and cellular proliferation observed in

53 both has been the reason for the dog prostate being used as a primary model to study in human pre-clinical pharmaceutical research. With the roles of Vitamin D being elucidated in human research and the potential for treatment of the condition of BPH being one of these roles, research into Vitamin D and BPH in dogs is reasonable and warranted.

3.3.3 Role of Vitamin D in Sperm Production

The effects of Vitamin D on semen quantity and quality characteristics and thus its possible role in male fertility have been studied. Early research in rats showed that Vitamin D deficiency in male rats greatly affected their ability to impregnate females [136]. The researchers found an absence of sperm in vaginal swabs after mating in Vitamin D deficient rats, and a lower fertility rate even in those where vaginal sperm were found after mating (45% pregnancy rate), compared with their Vitamin D supplemented controls (73% pregnancy rate). Vitamin D deficient male rats also had reduced testicular and epididymal sperm count along with lower concentrations of Sertoli cell testicular glutamyl transpeptidase activity from whole testes [160]. Histologically, decreased Leydig cell numbers and degenerative changes in the germinal epithelium were noted in

Vitamin D rats compared to supplemented controls [160].

Further confirmation of a role for Vitamin D within testicular tissue occurred with the identification of a nuclear VDR in the Sertoli cells [140]. Of note, however, was that sperm count and sperm motility were dramatically reduced in VDR null mice when compared with controls. Histologically, thinning

54 of the seminiferous epithelium, dilated seminiferous tubules and decreased or infrequent spermatogenesis was noted in the testes of VDR null mice.

Interestingly, those VDR null mice that had calcium supplementation and normal serum calcium concentrations did not show signs of decreased fertility.

In human males the VDR has been identified in the head of sperm cells but is lacking in the neck and tail region [138]. Cholesterol efflux, a priming event in the phosphorylation of proteins leading to human sperm capacitation, was increased in the presence of calcitriol [161]. Also, an increase in phosphorylation of tyrosine and threonine suggests that the VDR has a role in capacitation and survival of sperm [143, 161]. Other researchers were able to identify the VDR and the enzymes of Vitamin D metabolism in the human testis, epididymis, prostate, and seminal vesicles in varying degrees [139].

The VDR gene is highly conserved among humans, mice and rats [162].

In particular, exons 1e, 1a and 1d of the human VDR gene are conserved in multiple species, including the dog [163]. Although differences amongst species exist it is reasonable to assume that the VDR function coded for in these areas also conserved. Therefore presence of the VDR in similar tissues across species may be associated with similar functions. Taken together, these studies suggest an important role for Vitamin D in reproductive development and fertility in mice, rats and humans. Although Vitamin D has not been investigated in dogs, there is sufficient evidence to warrant research in this area.

55 3.4 PROLACTIN AND ITS ROLE IN MALE REPRODUCTION

Literature exists on the role of prolactin (PRL) in the female of multiple species including women, rats and the bitch [5, 164] however PRL has only recently attracted some interest in the literature regarding the male dog [165-

168]. The physiological role of PRL and its specific target organs have not been fully identified in any species. Hyperprolactinemia, caused by a prolactinoma or the use of some medications such as specific serotonin reuptake inhibitors

(SSRIs), is associated with symptoms of decreased libido and infertility in human males [5, 139, 169].

3.4.1 Prolactin and Ovarian Physiology

Prolactin is a 23 kDa pituitary peptide hormone related to GH and placental lactogen and produced by the lactotroph cells of the anterior pituitary

[5]. Just as with the other pituitary hormones like LH and FSH, study of PRL in the male may prove that similar characteristics related to its regulation and action are conserved not just among species but also between the two sexes. Prolactin secretion is pulsatile in women and pregnant rats, following a diurnal pattern, and thought to be regulated by the influence of the light/dark cycle on the regulatory suprachiasmatic nucleus (SCN) of the brain [5]. Although pulsatile, a 24-hour circadian rhythm is absent in the bitch [168]. The pattern of secretion in the dog follows an ultradian and circannual rhythm of secretion with higher concentrations occurring during periods of increased light [168]. The SCN influences the

56 hypothalamus (HTH) by stimulating the release of dopamine (DA) – the major inhibitory factor of PRL release. Alternatively, oxytocin acting on the periventricular nucleus (PVN) has been shown to be a major stimulus for PRL release. Nerve fibres from the SCN have connections with both the HTH and

PVN oxytocin-containing neurons in rats [170]. Interestingly, in pregnant women

PRL is also produced in extra-pituitary sites, mainly by the decidua, while in bitches there is no evidence of a source other than the pituitary [171, 172].

With respect to comparative physiology between the ovary and testes it is important to note that PRL action in the ovary has been linked to influencing enzymes involved in steroid hormone conversion causing decreases in E2 and increasing P4 production in the granulosa cells through down-regulation of 20D- hydroxysteroid dehydrogenase and up-regulation of LH-R in these cells to maintain the corpus luteum. In two studies of fertility in female PRL and PRL receptor (PRL-R) knock-out mice, researchers found that the PRL knock-out group experienced normal cycles and ovulatory patterns but could not maintain pregnancy [173] while the PRL-R knock-out mice had reduced ovulation, fertilization and arrest of pre-implantation development [174]. It is the mechanisms of PRL action at the cellular level where parallels may exist between female and male physiology showing similar patterns in Leydig cells and/or

Sertoli cells.

57 3.4.2 Prolactin and Male Physiology

3.4.2.1 Prolactin and the Prostate

Considerable evidence exists for a role of PRL in prostate physiology.

Human and rat prostatic epithelial tissue expresses both PRL and its receptor

PRLR [175, 176]. Early studies on the canine prostate detected intracellular endogenous PRL and binding sites for exogenous PRL within prostatic epithelial cells [177]. PRL has been shown to be a necessary component for prostatic epithelial growth and survival in human and rat tissue culture [5, 178]. Original studies in mice found that hyperprolactinemia increased in the weight of the prostate gland, by nine to twenty times, as well as histological evidence of prostatic hyperplasia [179]. Both in vivo and in vitro studies showed PRL affected growth and differentiation of the prostate [180] and sensitized the prostatic epithelial cells to androgen effects. This was thought to occur due to synergism between testosterone and PRL producing an increase in 5D-reductase activity [181]. These same changes were not noted in canine studies. Increased cell proliferation was seen only in those cultures supplemented with bovine and dog serum alone and not with any steroid hormones or PRL [182]. However, clinical studies of 5D-reductase inhibitors have shown a dramatic effect in size reduction of the canine prostate through decreased DHT concentrations [115].

Helmerich et al (1976) showed that pretreatment with PRL significantly decreased prostatic tissue concentrations of DHT and subsequently increased tissue testosterone [183]. To determine a PRL effect on prostatic androgen

58 metabolism investigators used recently (24 hours) castrated dogs and divided them into three groups: 1) control, 2) pre-treatment for 3 days with 500 IU PRL intramuscularly and 3) pre-treatment with 5 mg bromocriptine per os for 3 days.

All dogs were treated at 96 hours post-castration with tritiated testosterone (H3-

T), sacrificed one-hour post H3-T treatment and prostate tissue was harvested and measured for testosterone and DHT concentrations. PRL pretreatment significantly increased testosterone concentration over DHT when compared with controls. Pretreatment with bromocriptine (a dopamine agonist), significantly increased prostatic DHT and decreased testosterone compared with controls.

Dopamine agonists are compounds that bind to dopamine receptors and cause increased or enhanced dopamine effects and therefore PRL secretion is decreased due to the inhibitory action of dopaminergic stimulation. As a result, the study showed that PRL, through some mechanism, decreases conversion of testosterone to DHT; the hormone known to be primarily responsible for hyperplastic changes in the prostate.

Robertson et al (2003) determined that the castration-induced regression of the prostate in PRLR knockout mice was greater than in the normal controls, suggesting that PRL and testosterone act together in development of the ventral lobe of the prostate. The authors suggested PRL has a regulating effect of prostate development under normal physiological conditions of testosterone

[184]. In PRLR null mice the ventral half of the prostate was 20% heavier compared to controls. However, the ratio of epithelial cells to stroma within the dorsal lobes was decreased in PRLR null mice. This difference in size between

59 PRL null and control mice disappeared at one year of age suggesting there is a transient affect of PRL during prostate development.

Hyperprolactinemia due to prolactinomas in men has not been shown to increase prostate size [185]. Although there was no correlation between testosterone and prostate size in patients with prolactinoma, they had significantly smaller prostates and lower testosterone concentrations than their normal age-matched controls, suggesting lowered androgen levels in these patients contributes to a decrease in prostate size. This effect is most likely PRL- mediated inhibition of androgen production [186], however further study is warranted.

Conflicting findings on the role of PRL in male prostate physiology are apparent in the literature. This may be attributed to dose-dependent effects, chronicity of diseases, as seen with other pituitary hormones such as GnRH agonists or species-specific differences in prostate physiology. The PRL feedback loop appears to have increased delays when compared with other hormones [170, 187]. Nevertheless, PRL does appear to have a role in prostatic physiology. Studies of prostatic conditions such as BPH and cancer in dogs should therefore include PRL as a hormone of interest.

3.4.2.2 Prolactin, Male Fertility and Semen Quality

Hyperprolactinemia is associated with decreased semen quality and libido in both rodents and men [185, 186, 188-191] and, depending on the severity of the hyperprolactinemia and chronicity of the condition, hypogonadism can be

60 present. The main effects on semen quality include oligozoospermia, asthenozoospermia and teratozoospermia in these species [185, 186, 188-191].

Many of these conditions can be corrected using dopamine agonists or PRL antagonists (see review by De Rosa et al (2003) [192]). This impaired reproductive function in pathological conditions of PRL does not explain normal reproductive function, yet it has aided in its understanding. A study by Shafik et al (1994) used supra-physiological doses of ovine PRL (600Pg/kg/week IM for 6 months) to induce sterility in dogs [193]. By the end of the six-month study all dogs were azoospermic. At three months declines in total motility (<70%) and normal morphology (<60%) were observed, however, no other semen parameters were studied. It is important to note that by three months after cessation of treatments semen numbers, motility and normal morphology returned to pre-treatment values. This study looked at contraceptive effects of

PRL in the dog, so it is unknown whether naturally occurring hyperprolactinemia in the dog causes infertility and whether native canine PRL has a similar effect, however, these results are similar to those in men experiencing hyperprolactinemia.

In contrast to the female, PRL deficiency appears to have no effect on male fertility. In a study by Steger et al (1998), mature (2-3 months old) male

PRL null and control mice were individually housed for three weeks with two mature virgin females [194]. Fertility was measured by percentage of live litters produced from each group. From the control group 17/18 (94%) females delivered live litters and 32/33 (97%) delivered litters from the PRL null group

61 while litter size did not differ between the two groups (9.68 ± 0.39 and 10.06 ±

0.63 for PRL null and control males, respectively). It is unknown whether any compensable defects in semen quality were present that might have been overcome through increased normal sperm numbers via increased number of matings. Long-term effects of PRL deficiency from this study were also not accounted for. PRL deficiency may also impact fertility rates where issues of subfertility in the female are in question, however it is possible that PRL effects enhance but are not necessary to normal reproductive physiology in the male.

A recent review by Gill-Sharma (2009) details the feedback mechanisms and action of PRL in the male in both the rat and human [186]. PRL has both a short and long feedback loop involving the HPTA and the hypophyseal-pituitary- gonadal axis and appears to be complex.

Although there appear to be multiple regulators of PRL in the female the only regulator in the male of any importance to date appears to be dopamine.

Dopamine, a neuropeptide hormone, is a potent inhibitor of PRL secretion and is hypothalamic in origin. PRL down-regulates its own secretion by stimulating dopaminergic neurons to inhibit its own release. PRL primarily acts directly on the testis, chiefly the interstitial cells of Leydig [195]. It up-regulates and increases affinity of LH receptors in Leydig cells [195], causing an increase in testosterone secretion, although a direct role in increasing serum testosterone is still controversial [196, 197]. Testosterone produced by Leydig cells then acts on

Sertoli cells to produce E2 and increases production of androgen binding proteins

62 (ABP) [198]. Testosterone then acts via hypothalamic (dopamine) and pituitary routes to inhibit PRL secretion.

Stimulators of PRL secretion include the following components: Serotonin,

ȕHQGRUShin, Met enkephalin, Leu enkephalin, thyrotropin-releasing hormone

(TRH), GnRH, substance P, E2, epidermal growth factor, fibroblast growth factor, cholecystokinin, angiotensin II, and prolactin-releasing peptides (PrRPs) by direct action at the pituitary level [199-208]. The main hormone of PRL up-regulation, however, appears to be E2 through aromatization of testosterone. Estradiol acts directly on lactotrophs but also has a negative feedback on GnRH causing reduced LH and FSH secretion. In addition, PRL also has an inhibitory effect on

GnRH[209]. The presence of high concentrations of PRL and E2 seen in hyperprolactinemia is thought to be the mechanism by which serum testosterone,

LH, and FSH are lowered in these conditions. At physiological concentrations E2 feedback does not affect testosterone concentrations mainly due to PRL action on Leydig cells.

Prolactin reference ranges and ultradian and circannual patterns of PRL secretion have been published for the male dog [165, 167, 168, 210], although a difference in the type of assay used in these studies should be noted. The ranges published by Corrada et al (2006), using homologous enzyme immunometric assay [167], are summarized in Table 1.3. Of note, Beagle dogs had, on average, much higher PRL concentrations than both the crossbred dogs and German Shepherd Dogs sampled in this study. This same group was able to show a circannual variation in PRL concentrations with an association of higher

63 concentrations with increased daylight hours (November, December and January compared to May, June, July in the southern hemisphere). Kreeger et al (1992) recorded nadir during the fall months of October and November [210], but taking latitude into account, both studies saw similar patterns with respect to hours of daylight.

Although mean PRL concentration measured using radioimmunoassay

(RIA) did not differ widely among breeds in individuals with normospermia,

Urhausen et al (2009) saw a significant difference between these values in Fox

Terriers compared with Great Danes, with the latter being lower [165]. It was difficult to draw breed specific differences, however, due to the inbreeding of the

Fox Terriers enrolled in the study. A sharp increase in PRL concentrations was observed after thyroid stimulating hormone (TSH) stimulation in this same study.

Mean values of PRL remained within the range specified by Corrada et al (2006), yet it is important to note that sampling was only done once prior to TSH stimulation [165]. Therefore normal PRL fluctuation was not accounted for.

Although semen quality parameters are known to be adversely affected in human males with hyperprolactinemia, the study by Koivisto et al (2009) determined that semen parameters and libido remained unchanged after induced short-term hyperprolactinemia [211]. In this study, six male beagle dogs between the ages of 12 months to two years were divided into five three-week treatment periods: 1) pre-treatment, 2) metoclopramide 2mg/kg/ q8h po, 3) cabergoline 5 Pg/kg q24h po, 4) post-treatment period 1 and 5) post-treatment period 2. Fasting blood samples were collected in the morning twice weekly and hormone analysis was

64 done using RIA. PRL levels significantly increased with metoclopramide treatment (mean ± SD: 6.5 ± 1.6 ng/ml (p < 0.05)), a known stimulator of PRL secretion through dopamine antagonism, compared with pre-treatment (mean ±

SD: 4.5 ± 1.1 ng/ml) values. Cabergoline treatment resulted in a decrease in

PRL concentration (mean ± SD: 3.0 ± 0.6 ng/ml) compared with pre-treatment values (p < 0.05). Post-treatment values did not differ from pre-treatment ones

(mean ± SD 4.4 ± 0.8 ng/ml). It is important to note that although the investigators described the mean elevation in PRL as mild hyperprolactinemia it does not appear to be significantly different from the high range of normal determined previously (6.0 ng/ml). Indeed, mild hyperprolactinemia in men is defined as PRL levels in excess of 20 ng/ml [188], although it is difficult to extrapolate these values among species. It is also unknown whether a three- week period of induced hyperprolactinemia may accurately describe possible long-term effects of PRL on semen quality.

Table 1.3 Prolactin values in Dogs (ELISA)

Range (Mean r SE) (ng/ml) 0 - 6.0 (2.7 r 0.2) Mean Baseline r SE (ng/ml) 1.4 r 0.6

Pulse frequency (peaks/6hr) 1-2

Pulse duration (mins) (Mean r SE) 15-75 (45 r11)

Pulse amplitude (ng/ml) (Mean r SE) 1.7-2.4 (1.7 r0.4)

Adapted from Corrada et al 2006[167]

65

It appears that the few studies of PRL and fertility conducted in male dogs seem to correlate well with the findings of most human and rat studies. The presence of PRL and its receptors in prostate tissue in all species, influences of

PRL on prostatic androgen metabolism and semen parameters in states of excessive hyperprolactinemia appear to be conserved. However, the in vivo and cellular mechanisms of PRL physiology in the prostate and in spermatozoal development, especially in the dog, require further investigation.

66

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80 CHAPTER TWO

VITAMIN D, BENIGN PROSTATIC HYPERPLASIA, PROSTATE VOLUME AND

SEMEN PARAMETERS IN THE DOG.

Prepared for publication in Theriogenology

Abstract

The role of Vitamin D in normal skeletal metabolism is well-known; however, other functions of this vitamin in multiple physiological processes, including reproduction, are still being discovered. The aims of this observational study were to determine whether Vitamin D is associated with benign prostatic hyperplasia (BPH), prostate volume and/or semen quality characteristics in the dog.

Using a convenience sample of healthy dogs, associations between serum

25-hydroxy Vitamin D (25OHVD) and a) BPH, b) prostate volume (n=28) or c) sperm motility and morphology (n=22) were examined. Using the Student t-test no difference in 25OHVD between BPH positive dogs and normal dogs was observed (p=0.59). Bivariable analyses controlling for BPH, revealed no significant associations between Vitamin D and prostate volume (p=0.51).

Significant associations between Vitamin D and semen quality including motility

(R2=0.55, p=0.0008), progressive motility (R2=0.51, p=0.002), beat cross frequency (BCF; R2=0.28, p=0.01), distance average path (DAP; R2= 0.38, p=0.003), curvilinear distance (DCL; R2=0.25, p=0.02), linear distance (DSL;

R2=0.31, p=0.01), average path velocity (VAP; R2=0.35, p=0.005), curvilinear velocity (VCL; R2=0.25, p=0.02), straight line velocity (VSL; R2=0.27, p=0.02),

81 morphologically normal sperm (R2=0.43,p=0.01) and detached heads (R2=0.31, p=0.01) were found using univariable analysis. 25OHVD, with an age interaction term, was significantly associated with the following sperm motility parameters in bivariable analyses: motility (R2 = 0.57, p=0.0019), progressive motility (R2=0.57, p=0.002), BCF (R2=0.47, p=0.003), DCL (R2=0.51, p=0.006), VCL (R2=0.51, p=0.006), amplitude of lateral head displacement (ALH; R2=0.42, p=0.02) and average orientation change (AOC; R2=0.38, p=0.04), and normal sperm morphology (R2=0.49, p=0.01). Similarly, 25OHVD and prostatic volume interaction were associated with motility (R2=0.72, p<0.0001), progressive motility

(R2=0.60, p=0.001), DCL (R2=0.53, p=0.01), VCL (R2=0.52, p=0.01), normal morphology (R2=0.52, p=0.01), and head defects (R2=0.39, p=0.04).

25OHVD was associated with several sperm motility and morphology parameters which may suggest a role of 25OHVD on spermatogenesis and sperm function. Further study of the possible role(s) of Vitamin D in spermatogenesis, sperm function and sperm physiology is warranted.

Keywords; Vitamin D, semen, prostate, dog, benign prostatic hyperplasia

1. Introduction

Vitamin D has an important role in calcium and phosphorous metabolism and is integral to normal bone physiology. In severe deficiencies of Vitamin D in the very young, incomplete mineralization of developing or maturing bone leads to malformation of the skeleton manifested as rickets. In adults the same deficiency results in brittle and fragile bones leading to fractures. Research is

82 beginning to suggest a larger role for this vitamin in many different physiological systems, including reproduction [1-3].

Vitamin D is essential in inducing or inhibiting transcription of numerous genes that influence cell proliferation, invasiveness, angiogenesis, metastatic potential, differentiation and apoptosis [4]. The use of experimental Vitamin D analogues specific to the Vitamin D receptor (VDR) in human prostatic tissue has resulted in a significant decrease in prostatic volume in men with benign prostatic hyperplasia [5]. Vitamin D and its receptor have also been identified in human and rat prostatic tissue, testes and sperm [6-9]. In experimentally induced

Vitamin D deficiency in rats, a decrease in Sertoli cell numbers, degenerative changes in the seminiferous tubules and decreased sperm motility have been observed, while Vitamin D deficiency in men has been associated with increased mortality from prostate and other cancers[10, 11].

In dogs much is known about normal prostate function and physiology as well as spermatogenesis and sperm function. The effect of fat-soluble vitamins on reproduction in the stud dogs however, is largely unstudied. The purpose of this study was to examine whether associations exist between 25-hydroxy

Vitamin D (25OHVD) and BPH, 25OHVD and prostate volume, and 25OHVD and several sperm motility and morphology characteristics.

83 2. Materials and Methods

2.1 Case Subjects

A convenience sample of 28 healthy intact male dogs of various breeds

(total 23) and fertility histories, with or without indication of prostate disease, were obtained between the months of March and December of 2009. Dogs were recruited from cases presented to the Theriogenology Service at the Ontario

Veterinary College and from a private clinic in Southwestern Ontario with a large breeder clientele. All clients were required to fill out a consent form (Appendix 1) and brief questionnaire (Appendix 2) outlining their dogs’ fertility history, diet and amount of time their dogs spent outdoors.

All dogs underwent a complete male breeding soundness examination; including semen collection and analysis and ultrasonographic examination of the prostate gland. After completion of the examination, 6 ml of whole blood was collected via cephalic venipuncture; 2mls was immediately placed into a lavender top EDTA vial (Vacutainer™ Becton-Dickinson) for complete blood count (CBC) analysis (Animal Health Laboratory, Ontario Veterinary College). The remainder was placed in a red top vial (Vacutainer™ 366430 Becton-Dickinson) and allowed to clot for 20 minutes before centrifugation and collection of serum. Serum was then frozen and stored at -80°C until analysis. Finally, urine was collected mid- stream during micturition into a clean vessel for urinalysis. All dogs were examined and evaluated by the same operator.

84 2.2 Semen collection, dilution and staining.

Twenty-two of the 28 dogs (78.6%) enrolled for the study had semen successfully collected once with manual stimulation using a latex artificial vagina attached to a prewarmed sterile glass tube. Semen was divided during ejaculation into sperm-rich (2nd) and prostatic (3rd) fractions. Semen concentration was determined using the Unopette® - Neubauer system (Becton-Dickinson,

Rutherford, NJ) as previously described [12]. Sperm-rich semen was diluted 1:1 up to 3:1 with Canipro™ Chill 5 semen extender (Minitube Canada, ON) and chilled according to product specifications (Appendix 3) and either transported or stored between 5-8ºC for 1-3 hours prior to computer assisted semen assessment (CASA) evaluation. Morphological analysis was undertaken using eosin-nigrosin staining techniques[12] at 1000X magnification under oil (Nikon

Eclipse 50i, Japan) using the classification system of the Society for

Theriogenology (SFT) (Appendix 4). Semen collection and specimen preparation were performed by the same operator. A small amount of fresh undiluted unextended sperm-rich semen was submitted to the Animal Health Laboratory at the Ontario Veterinary College for aerobic bacterial culture and sensitivity testing.

2.3 Motility Analysis

Chilled extended semen samples were slowly warmed to room temperature over a 10-minute period, and placed on a prewarmed stage of the microscope (Olympus BX41 (U-Spt, Japan) prior to evaluation. Samples were given one minute to equilibrate in the slide chamber and then evaluated using the

85 SpermVision™ CASA system (Minitube, America) with the technical settings for canine semen as described by Schäfer-Somi and Aurich[13]. The present study used 50-300 cells per field and used a Leja® Standard Count chamber (Leja,

Netherlands) of 2 µl capacity. Table 2.1 details the technical settings used in the

CASA.

The following parameters were assessed using SpermVision™ analysis

(for a complete description of terms see Schäfer-Somi and Aurich, 2007): (1) total motility expressed as the percentage of spermatozoa with curvilinear velocity

(VCL) >15 µm/s [14], (2) progressive motility (%), (3) amplitude of lateral head displacement (ALH, µm), (4) average orientation change (AOC, degrees) where sperm <7 are considered immotile [13], (5) frequency of head displacement or beat cross frequency (BCF, Hz), (6) mean distance travelled (DAP, µm), (7) average velocity (VAP, µm/s), (8) curved line distance (DCL, µm), (9) curvilinear velocity (VCL, µm/s), (10) straight line distance (DSL, µm), (11) linear velocity

(VSL, µm/s), (12) mean straightness coefficient (STR = VSL/VAP X100, %), (13) linear coefficient (LIN = VSL/VCL X 100, %), and (14) wobble coefficient (WOB =

VAP/VCL X 100, %), and progressive fast spermatozoa (PF), which is defined by minimal STR for progressive fast sperm = 90% [15]. CASA analysis was performed by the same individual.

2.4 Prostate Examination

The prostate gland of all dogs was evaluated by the same individual after semen collection by several methods, including transabdominal ultrasonography,

86 digital palpation per rectum and cytological examination. Culture and sensitivity of prostatic fluid was performed by the Animal Health Laboratory, Ontario

Veterinary College, Guelph, Ontario.

2.5 Ultrasonography of the Prostate

Dogs were either placed in dorsal recumbency in a V-shaped trough to stabilize them for examination, or were examined in standing position depending on patient compliance and size. Using B-mode ultrasonography, measurements of length, width and height of the prostate gland were taken similarly as described previously [16, 17] using a MyLab® 5 Portable Ultrasound (Universal Ultrasound,

NY) with a 5-8 MHz curved linear array transducer. The prostate of each dog was measured three times in transverse and longitudinal sections and the average of each measurement was used in the analyses of prostate size. Prostate size was calculated using the volume formula described by Kamolpatana et al (2000) (Vm

= [1/2.6 (length x width x depth)] +1.8] cm3)[17]. Symmetry of the gland, as well as any lesions present were noted and measured. A BPH case was defined as a patient with a symmetrical prostate and at least one of the parameters of height, length and/or width being larger than the set maximum values based on the clinical experience of the evaluator (see Table 2.2) and the presence of cysts on ultrasonographic examination; one of the clinical signs of hemospermia, rectal tenesmus, and dysuria; and/or blood in the prostatic fraction on cytological examination [18].

87 2.6 Vitamin D Analysis

Vitamin D analysis was performed by the Diagnostic Center for Population and Animal Health, Michigan State University using a quantitative radioimmunoassay (RIA) for 25-hydroxyvitamin D (DiaSorin, Stillwater MN) validated for canines with normal ranges between 60 and 215 nmol/l. All samples were analysed in a single batch.

The 25OHVD assay consisted of a two-step procedure. Rapid extraction of 25OHVD and other hydroxylated metabolites from serum of plasma was accomplished with acetonitrile. Following extraction, the treated samples were then assayed using an equilibrium RIA procedure. The serum sample, antibody to

25OH-VitD and 125I-labelled 25OHVD tracer were incubated for 90 minutes at 20-

25°C. Phase separation was accomplished after a 20 minute incubation at 20-

25°C with a second antibody precipitating complex. Buffer was added after incubation and prior to centrifugation to aid in reducing nonspecific binding.

Sensitivity of the assay was 3.7 nmol/l with an inter- and intra-assay coefficient of variation of 11% and 10%, respectively, with cross-reactivities as follows: 100% for 25-OH-D2, 25-OH-D3, 24,25-(OH)2-D2, 24,25-(OH)2-D3, 25,26-(OH)2-D2, and 25,26-(OH)2-D3, 11% cross-reactivity to 1,25-(OH)2-D2, and 1,25-(OH)2-D3, and 0.8% cross-reactivity to ergo- and cholecalciferol.

2.7 Urinalysis

Urinalysis consisted of a urine test strip (Chemstrip 9¥, Roche

Diagnostics, Quebec), microscopic examination of urine sediment, determination

88 of urine specific gravity and bacterial culture and sensitivity (Animal Health

Laboratory, Guelph, ON) in order to rule out cystitis.

2.8 Statistical Analyses

2.8.1 Overall Statistical Analyses

SAS version 9.2 software (SAS Institute Inc. Cary, NC, USA) was used for all statistical analyses. The GLM procedure was used to analyze the univariable and multivariable associations between the explanatory variables and outcomes

(see next sections for details). Due to the small sample size and the limited degrees of freedom available, only two explanatory variables were used in any model to discourage over-fitting of the model. Models were generated using backwards elimination in the case of multivariable analyses. The significance level was set to p<0.05 for all analyses, and 95% confidence intervals were constructed.

The UNIVARIATE procedure was performed to assess residuals and

ANOVA assumptions in order to determine the fit of the model and whether transformation of the data was needed to achieve normality. Four tests were used to test normality: Shapiro-Wilk, Cramér-von Mises, Kolmogorov-Smirnov,

Anderson-Darling tests [19]. Residuals were plotted against the predicted values and explanatory variables used in the model. Such plots and tests for normality may reveal outliers, unequal variances or other problems with the assumptions and may suggest the need for a data transform [19, 20]. Data transformation included a logit transform (logit of outcome = log((r+k)/(n+k)); r=responding cells,

89 n=number of cell counted, k=small bias correction term = 0.25) in cases of variables with a percentage value.

Using the Grubbs’ test for outliers [21], any result where the residual value was greater than 2.58 times the standard deviation of the residuals (one-tailed test p<0.05) were labeled as outliers and the models were rerun to determine how both normality of the data and the model changed with removal of these points

(Appendix 5).

2.8.2 Study Population

Descriptive statistics including means, standard deviations (SD), and standard errors (SE) were used to analyze the characteristics of the study population. Ages of dogs were simplified using half-year increments as not all dogs had exact birth dates. Time of year was divided into Summer (March 20-

September 21) and Winter (September 22-March 19) according to the vernal and autumnal equinoxes in the northern hemisphere, to account for the influence of natural ultraviolet radiation on vitamin D production via the skin. The Student’s t- test was used to test whether there was a difference in mean serum 25OHVD concentrations between dogs of the two seasonal groups. Amount of time spent outdoors was grouped according to whether the dogs spent greater than or less than five hours per day during the season of sampling and differences between groups was analyzed using a one-way ANOVA. Diets were categorized according to whether a commercial, raw or a combined diet was given to account

90 for other vitamin D sources. Differences in mean serum 25OHVD concentrations amongst the different diets were analyzed using one-way ANOVA.

2.8.3 BPH and Prostatic Volume

The Student’s t-test was used to test whether there was a difference in mean serum 25OHVD concentrations between dogs with and without BPH.

Univariable analysis described in section 2.8.1 included the explanatory variable serum 25OHVD and prostate volume as the outcome variable. Analysis of prostate volume was analytically controlled for BPH status by including it in the model. Age and bodyweight were also tested separately as explanatory variables with serum 25OHVD as the outcome variable. In keeping with previous studies of prostate volume [16, 17] bivariable regression model of prostate volume with age and bodyweight as explanatory variables was tested.

2.8.4 Sperm Motility

Univariable analysis of sperm motility described in section 2.8.1 included total motility, progressive motility, ALH, AOC, BCF, DAP, VAP, DCL, VCL, DSL,

VSL, STR, LIN, and WOB, as outcome variables with serum 25OHVD as the explanatory variable. All sperm motility parameters were included in bivariable analyses. Three bivariable regression models were created for each motility parameter; the first model included age and serum 25OHVD, the second included bodyweight and serum 25OHVD, and the third included prostate volume and serum 25OHVD as explanatory variables.

91 2.8.5 Sperm Morphology

Univariable analysis of semen morphology separately modeled normal sperm, head defects, midpiece defects, tail defects, loose heads, proximal droplets and distal droplets as outcome variables with serum 25OHVD as the explanatory variable. All sperm morphology parameters were included in bivariable analyses. Three bivariable regression models were created for each sperm motility parameter as the first model included age and serum 25OHVD, the second included bodyweight and serum 25OHVD, and the third included prostate volume and serum 25OHVD as explanatory variables.

3. Results

3.1 Vitamin D and the Study Population

Serum vitamin D concentrations of subjects ranged from 70 to 203 nmol/l with a mean and SD of 140.5 r 37.1 nmol/l. BPH was found in 82.1% (23/28) of dogs studied. Ages of dogs ranged from 2 -11 years with a mean and SD of 6 r 2 years. Bodyweight of dogs ranged from 9.2-78.2Kg with a mean and SD of 35.4 r 15.8Kg. Tables 2.3 to 2.5 describe the other characteristics of the study population of dogs. No significant associations between mean serum 25OHVD concentrations and types of diet (p=0.79), time of year of sampling (p=0.54) and number of hours spent outdoors in the summer and in the winter (p=0.85) were observed (Tables 2.6 - 2.8). CBC and urinalyses of all dogs enrolled in the study were within normal limits. Cultures and sensitivities of urine, prostatic fluid and

92 semen in dogs that gave semen samples, were all negative. All dogs were healthy having neither illness nor receiving any medications. Fertility histories of the study population consisted of dogs that had never been bred or had not been bred within the year prior to evaluation. Body condition of the majority of dogs was considered ideal except for two dogs (Golden Retriever, Mastiff Cross) that were categorized as slightly overweight.

3.2 BPH and Prostate Volume

There was no significant association between mean serum 25OHVD concentration and BPH status (p=0.59) (Table 2.9). Univariable analysis revealed no significant association between 25OHVD concentration and prostate volume

(R2=0.02, p=0.51). Controlling analytically for BPH in the previous model did not change the results obtained. There was no significant association with serum

25OHVD as a response to age (R2=0.02, p=0.52) or bodyweight (R2=0.0004, p=0.92). There was significant association in bivariable analysis of prostate volume with age and bodyweight (log2Volume=0.13*Age + 0.02*Bodyweight,

R2=0.20, F-value=3.33, p=0.05) without interaction. Overall there was an increase in prostatic volume with increasing age and bodyweight. Using three research hounds examined twice one day apart, intra- and inter-assay coefficients of variation on prostatic measurements and their 95% confidence intervals were

3.6% (2%, 4.6%) and 21.8% (17.3%, 29.7%) for prostatic length, 4.8% (4%, 6%) and 19.7% (16.5%, 26.4%) for prostatic width, and 4.3% (3.5%, 5.4%) and 21.1%

(16.7%, 28.7%), respectively.

93 3.3 Sperm Motility

Univariable analyses of both total motility (R2=0.55, p=0.0008) and progressive motility (R2=0.51, p=0.002) had significant association with serum

25OHVD alone (Figures 2.1 and 2.2). For both motility outcomes, the pattern of association was an increase in the motility parameter at 25OHVD concentrations between 70 and 100 nmol/l, little or no effect on motility at 25OHVD concentrations between 100 and 180 nmol/l and a slight decrease in motility at

25OHVD concentrations above 180 nmol/l (all 25OHVD concentrations stated here are within the normal range for healthy dogs).

Significant univariable associations between 25OHVD and BCF, DAP,

VAP, DCL, VCL, DSL and VSL were also found (Table 2.10); specifically each had a simple linear positive association with increasing serum 25OHVD. No significant associations were found among semen motility parameters ALH, AOC,

LIN, STR, and WOB and the explanatory variable serum 25OHVD (Table 2.10).

Bivariable regression analysis revealed significant interaction of age

(R2=0.71, p=0.0001 and R2=0.57, p=0.002) and prostatic volume (R2=0.73, p=0.0001, and R2=0.60, p=0.0012), but not bodyweight, with serum 25OHVD on both sperm motility and progressive motility, respectively. Figures 2.3 – 2.6 depict these relationships. At younger ages and smaller prostatic volumes, there is a noticeable effect of increasing concentrations of serum 25OHVD on increasing motility and progressive motility at 25OHVD concentrations between

70 and 140 nmol/l. At older ages (i.e.10 years) and larger transformed prostate volumes (i.e. 5.5) an opposite effect of serum 25OHVD on motility is observed

94 with a decrease in total motility parameters with increasing serum 25OHVD.

Progressive motility follows a similar pattern to total motility with increasing

25OHVD and prostate volume however, there is an increase in progressive motility with increasing 25OHVD at all ages with less dramatic results between 8 and 10 years of age.

Using bivariable analysis, BCF had a significant association with serum

25OHVD and age of dog as explanatory variables (R2=0.47, p=0.003) but not with serum 25OHVD and bodyweight nor with serum 25OHVD and prostate volume.

BCF increased slightly with age and increasing serum 25OHVD (Figure 2.7). For both response variables DCL and VCL, significant interactions were found between the explanatory variables serum 25OHVD and age, and serum 25OHVD and prostatic volume but not with serum 25OHVD and bodyweight. Specifically, at younger ages DCL increased with increasing serum 25OHVD but by 8 years of age there was little effect of increasing serum 25OHVD on DCL. At 10 years of age, increasing serum 25OHVD was associated with decreasing DCL (Figure

2.8). Smaller prostate volumes were associated with an increase in DCL with increasing serum 25OHVD, however, at a larger transformed prostate volume of

5.5 the reverse is true with a decrease in DCL as a response to increasing serum

25OHVD (Figure 2.9). VCL responded to increasing serum 25OHVD in the same manner as DCL with respect to both age and prostate volume (Figures 2.10-2-

11). A significant association between DSL and prostate volume was found

(R2=0.46, F-value=4.81, p=0.001) (Figure 2.12). Although VSL did not have a

95 significant association with prostate volume it did approach significance (R2=0.28,

F-value=3.42, p=0.06) (Table 2.10).

There was a significant association of each of ALH and AOC with serum

25OHVD and age as explanatory variables but not with serum 25OHVD and bodyweight nor with serum 25OHVD and prostate volume (Figures 2.13 and

2.14). There was an increase in both ALH and AOC with increasing age and increasing serum 25OHVD at younger ages and a negative association between these motility parameters and increasing serum 25OHVD at older ages. There were no other significant bivariable associations between the remaining motility parameters DAP, VAP, VSL, STR, LIN, WOB with serum 25OHVD and age, serum 25OHVD and bodyweight and serum 25OHVD and prostate volume (Table

2.11).

3.4 Outlier Removals

Removal of outliers, as previously defined, within this set of data influenced two of the significant associations observed (Appendix 5). Sperm total motility and Vitamin D associations at various prostate volumes changed significantly with respect to their slope coefficients with removal of one outlier

(Dog 6, a 5-year old Golden Retriever that had never sired a litter even after multiple breedings). The overall pattern of association was similar to those found with the outlier maintained although the magnitude of those associations was decreased. Secondly, removal of the same outlier with respect to the associations between progressive motility and 25OHVD alone became insignificant (p=0.21).

96 Although, the removal of outliers in large data sets may be advised we maintained the data due to the small sample size and possibility of removing valuable information from the analysis.

3.5 Sperm Morphology

The relationship between 25OHVD and morphologically normal sperm was quadratic, predicting an increase in percent normal morphology with increasing 25OHVD concentrations at low to mid-normal range of normal serum

25OHVD (see Figure 2.15). However, at high normal values of serum 25OHVD this effect was lost and a negative effect on normal morphology was observed.

A significant decrease in the percentage of loose heads can be seen with increasing serum 25OHVD (see Figure 2.16). No other significant associations with 25OHVD using univariable analyses were found with the remaining morphology characteristics including midpiece (R2=0.03, F-value=0.52, p=0.48) and tail (R2=0.002, F-value=0.05, p=0.82) defects, proximal (R2=0.006, F- value=0.11, p=0.74) and distal (R2=0.02, F-value=0.34, p=0.56) droplets. There was a tendency towards significance with head defects and 25OHVD (R2=0.17, F- value=3.61, p=0.07).

Bivariable analyses explained greater variation than univariable analysis between normal sperm and 25OHVD. Both age and serum 25OHVD and prostatic volume and serum 25OHVD used in modeling the response of normal sperm were significant (Table 2.12 and Figures 2.17-2.18). With increasing age and prostatic volume, response to 25OHVD was greatest at younger ages and

97 smaller prostate volumes, while at the age of 10 years and transformed prostate volume of 5.5 there was a decrease in percentage of normal sperm with increasing serum 25OHVD. Prostate volume and 25OHVD interacted and were significantly associated with percentage of head defects (Table 2.12 and Figure

2.19). The percentage of head defects decreased with increasing 25OHVD at small prostate volumes, but increased at transformed prostate volume 5.5. The bivariable regression models for the remaining sperm morphology characteristics were non-significant (Table 2.12).

4. Discussion and Conclusions

Many of the motility parameters studied have strong correlations to fertility in many species [22-24] and the associations of Vitamin D with these parameters have importance in the reproductive health and potential of the stud dog. Among these parameters total motility, progressive motility, ALH, VAP, VCL, VSL, BCF,

LIN have shown high correlation with AI fertility in multivariable analysis in bulls

[22] while ALH, VCL, and VSL in in vitro human studies have shown correlation with high fertilization rates [23]. In the dog, it has been found that most of motility parameters measured by CASA had correlation to fertility except for BCF, LIN, and STR [24]. Therefore fertility potential, as measured by CASA, and associations with 25OHVD in the dog can provide new insight into reproductive health in the stud dog.

Serum 25OHVD had significant positive associations with several motility parameters in this study in both univariable and bivariable analysis. Total and progressive motilities in univariable analysis showed almost identical patterns of

98 association with 25OHVD and are in agreement with human studies [25].

25OHVD had positive associations with total and progressive motilities in the low to mid-range of normal (70-180 nmol/l), and a negative effect at high normal

(>180 nmol/l) concentrations, with the most desirable outcomes in the mid-range of normal 25OHVD (120-180 nmol/l). Blomberg-Jensen et al (2011) found similar associations, using a 25OHVD squared term with human sperm, although their associations were smaller than in our study and a negative effect at higher concentrations of 25OHVD was not seen [25]. Due to possible interspecies differences in 25OHVD concentrations, it is difficult to determine whether the higher 25OHVD concentrations observed in the human study are comparable to the concentrations observed in our canine population and could a similar effect have been seen at even higher 25OHVD concentrations (nearer the upper limit) in men. In addition, the human study analyzed total motility only and found that in vitro addition of the active Vitamin D tissue form, 1,25-dihydroxyvitamin D

(1,25diOHVD), to mature sperm caused a dose-dependent increase in total motility, but after a certain concentration (10-7 M) a negative effect was seen.

This finding is supported by studies of sperm survival where a similar biphasic dose-response curve to 1,25diOHVD was found [6]. This could in part be due to exceeding Ca2+ stores and exhausting the intracellular Ca2+ influx and ligand binding and activation of the protein kinase C system responsible for sperm motility [26]. Blomberg-Jensen et al (2011) also found that addition of

1,25diOHVD caused increased intracellular Ca2+ concentrations in ejaculated spermatozoa compared with controls. They confirmed that Vitamin D action was

99 through a non-genomic VDR [25]. The results from these studies suggest that the Vitamin D associated increases in sperm motility and progressive motility we observed may work independently of VDR mediated gene transcription.

Total and progressive motilities in bivariable analysis with both 25OHVD and age, and 25OHVD and prostatic volume showed an increase in both motility parameters at younger ages (4-8 years of age) and smaller transformed prostate volumes (log2volume = 3.5 - 4.5) and a decrease at older ages (10 years of age) and larger transformed prostate volumes (log2volume=5.5). Age effects on motility parameters have not been previously found in the dog [24] although they have been found in men [27]. Multivariable effects were not investigated in these studies and the effect of age and 25OHVD with age-25OHVD interaction observed in our study suggest that age may directly influence Vitamin D physiology. In a study that investigated age-related changes in Vitamin D physiology in rat duodenum, an age effect was observed [28]. Measuring both protein kinase C (PKC) and Ca2+ uptake by cultured duodenal cells from young

(three month-old) and aged (22-24 month-old) rats in the presence and absence of 1,25diOHVD, aged rats showed higher basal PKC concentrations than younger rats while basal Ca2+ uptake didn’t differ between the two groups. With addition of 1,25diOHVD there was a significant increase in both PKC activity and

Ca2+ uptake in the cells from the young rats compared with the aged rats.

Specifically, percentage of PKC activity with 1,25diOHVD treatment was acutely increased in younger rats while in aged rats there was an acute decrease in percentage PKC activity returning to normal by 10 and 6 minutes in the young

100 and aged cells, respectively. The results obtained from the rat study suggest ageing may have a similar effect on sperm total and progressive motilities through Ca2+/PKC signaling pathways and the patterns of this activity observed appear to mimic those seen in the motility parameters with 25OHVD of our study.

The results of the rat study also suggest that non-genomic Vitamin D metabolism is impaired with ageing although it is unknown what effects nuclear Vitamin D action may have. The association of 25OHVD and prostate volume on motility parameters may be more difficult to explain. Age and bodyweight together make up the components that influence prostate volume in the dog in this study as well as in others [17]. While bodyweight and 25OHVD were not found to have an association with either total or progressive motility, the opposite was true for age and 25OHVD and therefore suggests that the age component of prostate volume may account for the association found. To further support this hypothesis, clinical BPH with blood in the ejaculate has not been found to have association with semen quality and fertility [18, 29], although lysis of red blood cells in thawed frozen semen samples causes detrimental effects on sperm motility and viability

[30]. Studies to determine whether Vitamin D and the VDR are present in the testes and sperm in the dog and what physiological role Vitamin D plays in sperm motility are warranted to further detail the associations found in this study.

In univariable analysis of BCF, DAP, DCL VAP, VCL and VSL, each motility parameter followed a simple positive linear association with serum

25OHVD. It is unknown whether increases in BCF with increasing 25OHVD in the dog have clinical relevance, as this parameter did not correlate to fertility in

101 this species [24]. DAP and VAP, as well as, DCL and VCL are related parameters (increases in distance travelled per examination time and the increase in distance travelled per second) and it is therefore not surprising that both sets of parameters saw similar increases with increasing 25OHVD concentrations. It is also important to note that VCL is a component of defining both total and progressive motilities via CASA and therefore its significance is not surprising. Increases in VSL were also significant with increasing 25OHVD concentrations and although DSL association with 25OHVD approached significance (p=0.06), it was not statistically significant possibly due to a lack of statistical power due to a small sample size. Fertility has correlated well with these parameters in dogs [24] and improvements in average, straight and curvilinear distances and velocities of sperm may be translated as the ability of sperm to traverse the female reproductive tract efficiently. No other published studies relating Vitamin D to these motility parameters exist at present. The mechanism of Vitamin D action on these parameters may occur during crucial development of the midpiece (mitochondrial/axoneme function) and/or tail

(axoneme) regions of spermatids in the testis. Identification of the VDR in the midpiece of immature human spermatozoa was found [31] indicating a possible role of Vitamin D in the development and maturation of sperm in this species and encourages further study in the dog to determine if the same holds true. A recent study has shown VDR to be present in the mitochondria of human platelets [32] suggesting that the VDR localization in the midpiece of human spermatozoa may be specific to the mitochondria. The presence of the VDR in the tail regions of

102 mature sperm suggests that Vitamin D may influence axoneme structure and this hypothesis is supported by studies that have shown Vitamin D deficiency during fetal development in rats alters protein structure of certain cytoskeletal elements, specifically E-tubulins, in rat brain cells [33]. These E-tubulin proteins are also present only in the midpiece and tail regions of human sperm [34]. If Vitamin D action also holds true for maintaining normal axonemic structure in the sperm tail, sperm propulsion may therefore be affected. All developmental stages of sperm have been shown to express the VDR and 25OHVD hydroxylase enzymes in rats and men, as well as Sertoli and Leydig cells suggesting that Vitamin D has a role, directly and/or indirectly on sperm maturation. Calcium signaling through cytosolic VDR may also be the trigger to fuel mechanisms involved in increasing distance travelled and velocity of sperm, however a negative effect at high normal 25OHVD concentrations was not observed as with total and progressive motilities suggesting that this concentration limited mechanism may not be involved.

There was an increase in BCF with increasing age and 25OHVD in bivariable analysis. It is uncertain what this association may mean to age-related male dog fertility and 25OHVD. There were increases in DCL and VCL at younger ages (4-8 years of age) and smaller prostate volumes (log2volume=3.5-

4.5) while both decreased at older ages (10 years) and larger prostate volumes

(log2volume=5.5) with increasing 25OHVD. Similarly to total and progressive motility, age-related effects on DCL and VCL may be dependent on Ca2+/PKC physiology in the mitochondria of the sperm midpiece. It is also possible

103 however, that age-related influences on nuclear Vitamin D effects with respect to structural protein metabolism may also be involved. As mentioned previously E- tubulins are necessary for axoneme structure in the midpiece and tail regions of spermatozoa. In a study investigating age-related structural effects in human fibroblasts, both actin and E-tubulin proteins were decreased in senescent compared with fetal fibroblast culture [35]. F-actin is a structural protein component of the sperm flagellum and cleavage of F-actin from this structure has been shown to hamper sperm motility in guinea pigs [36]. In addition, 25OHVD action on sperm structure in ageing may occur through impaired inhibition of nuclear factor NB (NFNB) signaling – known to be a target of nuclear Vitamin D signaling (see review by Nagpal [37]). NFNB has been suggested to be a mediator through which actin and tubulin decrease and vimentin increases in senescent human fibroblast cells [35]. Whether a similar mechanism exists in developing spermatocytes remains unknown. DSL also increased at smaller transformed prostate volumes and decreased less dramatically at larger transformed prostate volumes with increasing 25OHVD. Although no affect of age was seen it is possible that the age component of prostate volume is still a factor in DSL outcome with 25OHVD. The small sample size and resulting statistical power of the study may not have been high enough to detect age effects with 25OHVD. Although, our study found the same positive association of prostate volume to bodyweight and age as in previous studies [16, 38, 39], it is possible that bodyweight and age are not the factors of prostate volume responsible for DSL outcomes with increasing 25OHVD. It is perhaps possible

104 that altered Vitamin D physiology of the enlarged prostate exists. BPH has not been associated with infertility in the dog, however only a few components of prostatic fluid have been identified in previous studies [40]. Men with BPH and chronic pelvic pain syndrome have been shown to have higher concentrations of

IL-8 [41] and the VDR agonist elocalcitol has been shown to decrease these concentrations in seminal plasma [42]. Whether, seminal plasma components are altered in BPH in the dog is unknown and the potential influence of Vitamin D on prostatic secretion and possible semen effects on ejaculated sperm is worth further study.

Both ALH and AOC increased with increasing 25OHVD at younger ages

(4-6 years) and decreased at older ages (8-10 years). Both parameters are positively correlated with fertility in dogs [24]. In human sperm, ALH is associated with fertilizing potential by overcoming the barriers of cervical mucus and the peri-oocyte envelope before fertilization of the oocyte can take place [43-

45] and increases in this motility parameter suggest a positive role for Vitamin D in male fertility. Measuring AOC determines whether sperm during analysis are being passively moved. Sperm with an AOC level of <9.5 have been defined as being immotile and/or dead [13]. Therefore increases in this parameter indicate that sperm are able to change direction according to changes and/or obstacles in their path or are more sensitive and more likely to respond to those obstacles. It is likely that age-related 25OHVD effects of these parameters are mediated through the same energy and structural mechanisms as the motility parameters already mentioned.

105 Normal sperm morphology followed a quadratic pattern similarly to total and progressive motilities. As with the motility parameters, a positive effect of

25OHVD on normal morphology in the low to mid-range of normal 25OHVD concentrations (70-160 nmol) and a negative effect in the high end of the range

(>160 nmol/l) were observed. Desirable normal morphology (>80%) was observed in the 140-180 nmol/l range. As normal spermatogenesis logically results in production of morphologically normal sperm it is possible that Vitamin D may exert an effect at the level of spermatocytes, Sertoli and Leydig cells in the testis of the dog; this is supported by studies in rats that have also shown reduced fertility and degenerative changes in testis and spermatogonia in

Vitamin D deficiency [2]. Certain local factors such as cytokines and growth factors are known to have a part in regulation of spermatogenesis [46]. Among these factors interleukins (ILs), insulin-like growth factors (IGFs), and tumor necrosis factor- alpha (TNFD) action are influenced by Vitamin D in other tissues and more specifically, Vitamin D has been shown to negate the effects of these factors (see review by Nagpal [37]). The inflammatory cytokines IL, IGF and

TNFD are thought to be involved in normal tight-junction complex recycling, however, they can cause disruption of the tight-junction adhesions and consequently compromise the BTB at greater than physiological normal concentrations [47-49]. It is possible that maintenance of these tight-junctions is mediated through Vitamin D and may therefore have a protective function of the developing sperm from inflammatory insult. Vitamin D has also been shown to influence enzymes such as phospholipase C (PLC) that are associated with

106 cellular differentiation, while stimulating insulin-like growth factor binding protein 3

(IGFBP-3) secretion to bind and inhibit insulin-like growth factor (IGF) [50].

Perhaps there is a dose-dependent effect of Vitamin D in the testes and germ cells that causes IGF inhibition at both high and low concentrations yet switches to stimulate PLC action at physiologically optimal concentrations. The percentage of detached heads decreased with increasing concentrations of

25OHVD indicating a possible role for Vitamin D in the development of the head midpiece/tail connection similarly to the prior discussion of normal morphology and supports our hypothesis for a spermatogenic role of Vitamin D. Although no other morphological defects of sperm were associated with 25OHVD it is possible that subtler defects as assessed by scanning or transmission electron microscopy may have revealed a different outcome with respect to 25OHVD.

Increases in percentage of morphologically normal sperm were associated with increasing 25OHVD at younger ages (4-8 years) and smaller transformed prostate volumes (log2volume=3.5-4.5) and decreased with increasing 25OHVD at older ages (10 years) and larger transformed prostate volumes

(log2volume=5.5). Age and prostate volume influences were most likely due to the possible age effect on spermatogenesis mentioned previously in the section on total and progressive motilities. Whether this positive sperm morphological trait occurs through genomic and/or non-genomic Vitamin D effects remains to be elucidated. Similarly, percentage of head defects decreased at smaller transformed prostate volumes (log2volume=3.5-4.5) and increased at larger transformed prostate volumes (log2volume=5.5). Whether the reason was due to

107 a masked age effect on Vitamin D physiology during spermatogenesis and/or an influence of Vitamin D associated changes in prostatic fluid composition causing acrosomal or membrane damage to spermatozoa necessitates further research.

There were a large number of dog breeds compared with the number of individuals in our study and therefore we were unable to account for breed effect or breed trends in this study. Nevertheless, the information obtained on Vitamin

D in this population of dogs has the potential to open new avenues of research and possible treatment for cases of poor or sub-normal semen characteristics in the dog. It is important to note that the majority of dogs enrolled in the study were sexually rested for at least one year prior to examination and few had been used for breeding therefore breeding histories were not analyzed.

Although many forms of Vitamin D exist, 25OHVD most accurately represents the physiological Vitamin D reserves due to its long half-life of 2-3 weeks and also because the first hydroxylation step in its formation is unregulated; dependent only on substrate supply [51]. It is this supply of

25OHVD that feeds the conversion to the more active forms of Vitamin D such as

1,25diOHVD in specific tissues such as the prostate and testes. Therefore,

25OHVD was considered to be the rate-limiting step in the enzymatic cascade in

Vitamin D action within these tissues and was thus the chosen substrate to be assayed [51]. It is also the main circulating form of Vitamin D in the dog.

Day length and ultraviolet (UV) light exposure in the Northern Hemisphere, as determined by time of year of sampling had no effect on 25OHVD concentrations in dogs. These data further support other studies determining

108 that dogs are unable to synthesize Vitamin D through the action of UV radiation on the skin [52, 53]. Vitamin D concentrations were also not influenced by type of diet, whether raw or commercial indicating appropriate Vitamin D intake by study subjects was not dependent on type of diet and all diets were able to meet the daily requirements of the study population.

Experimental trials with Vitamin D analogues in human cases of BPH have shown a significant shrinkage of enlarged prostates [5, 54, 55]. This study showed that 25OHVD concentrations were not associated with prostate size or

BPH in the study population of dogs. This lack of significance may be due to the sample size being too small to detect any differences in the population. BPH was also not confirmed using histology in this study. Instead, the presence of BPH was defined according to clinical signs, predetermined cut-off points for prostatic dimensions on ultrasound examination, and elimination of other pathologies.

Controlling for BPH in prostate volume analysis was done to determine whether enlargement of the prostate was associated with Vitamin D concentrations independent of our diagnostic criteria for BPH. Future studies using larger sample sizes and histological confirmation of BPH are warranted and may reveal different results.

Adiponectin, a marker for body mass index, has been negatively associated with Vitamin D concentrations in humans [56]. More specifically, obese patients with lowered adiponectin concentrations have the strongest associations with Vitamin D concentrations bordering on deficiency [56].

Adiponectin has also been correlated to body condition scoring in dogs [57].

109 Therefore a possible influence of body condition on Vitamin D concentrations might exist in the canine. In our study population there were only two dogs

(Golden Retriever and Mastiff cross) that were considered slightly overweight with the majority of the population falling into the ideal body condition category and were not considered to be Vitamin D deficient (104 and 115 nmol/l 25OHVD, respectively). Further study looking at Vitamin D status, body condition score and adiponectin concentrations in a wide range of dogs would be worthwhile in understanding Vitamin D physiology in this species. It is important to mention that, statistically, a great many comparisons were made in our study and the risk for Type I error (false positives) is increased with the large amount of associations found to be significant. Unfortunately, there is no reliable method for correcting for this issue and identifying this type of error in the present study.

Methods that have been suggested i.e. Bonferroni come with their own risk of increasing Type II error (false negatives).

In conclusion, serum 25OHVD concentrations between 120-180 nmol/l were associated with desirable semen characteristics that are connected to dog fertility. A possible new tool in diagnosing and treating abnormalities in the spermiogram is an exciting concept. Further research aimed at identifying

Vitamin D and/or its receptor in reproductive tissues and the mechanisms by which they exert their effects would be beneficial to increasing our understanding of Vitamin D in spermatogenesis and reproductive health in the dog.

110 Table 2.1 Technical Parameters for SpermVision™ CASA

Parameter Depth of chamber 20um Light adjustment 96-104 Volume per Chamber 2ul Temperature 37°C Sperm concentration Variable Area of sperm heads 20-60 um2 Number of fields or Cell number 7 or 1000 Number cells/field 50-200 Frame rate 60/sec Total motility VCL > 15 Progressively Fast VCL >15, STR>0.9 Linear STR >0.9, LIN>5 Immotile AOC <7, DSL <3 Local motility DSL <6 Hyperactive VCL>118, ALH>6.5, LIN<0.5 Non-Linear LIN<=0.5, STR<=0.9 Curvilinear DAP/Radius >=3, LIN<0.5 Incubation time 2 mins Modified from Schäfer-Somi and Aurich, 2007[13]

111 Table 2.2 Classification scheme for determination of the presence of BPH by ultrasonography in 28 dogs, March-December 2009, Ontario, Canada

Prostatic Measurement Bodyweight (Maximum of Length, Width or Height) > 3.0 cm ” 20 kg > 3.5 cm !WR” 40 kg > 4.0 cm >WR” 60 kg > 4.5 cm !WR” 80 kg Classification based on arbitrary cut-points.

112 Table 2.3 Characteristics of the study population of 28 dogs, in Ontario, Canada March-December 2009. Variable Mean SD SEM Min Max Age (years) 6.02 2.05 0.38 2.00 11.00 Body Weight (kgs) 35.36 15.84 2.94 9.20 78.20 Vitamin D (nmol/l) 140.50 37.05 5.72 72.00 203.00 .

*SD=standard deviation SEM=standard error of the mean Min = Minimum value Max = Maximum value

113 Table 2.4 Characteristics of the study population of 22 dogs from which semen was collected in Ontario, Canada March-December 2009. Variable Mean SD SEM Min Max Age (years) 6.25 2.09 0.45 4.00 11.00 Body Weight (kgs) 35.47 14.71 3.14 9.20 71.30 Vitamin D (nmol/l) 141.86 35.03 7.47 72.00 189.00 Total sperm (106) 524.26 324.89 69.27 33.00 1000.00 Motility (%) 89.1 15.74 3.36 27.00 98.60 Progressive Motility (%) 80.19 18.57 3.96 13.30 95.70 Normal sperm (%) 61.62 29.58 6.45 0 95.00 Head Defects (%) 10.76 19.97 4.36 0 78.00 Midpiece Defects (%) 8.19 5.90 1.29 0 20.00 Tail Defects (%) 3.38 4.43 0.97 0 16.00 Loose Heads (%) 2.00 3.86 0.84 0 16.00 Proximal Droplets (%) 12.00 18.57 4.05 0 61.00 Distal Droplets (%) 1.67 1.62 0.35 0 5.00 DCL (um) 65.12 15.71 3.43 29.78 94.83 DAP (um) 30.77 5.73 1.25 16.60 40.50 DSL (um) 21.76 4.56 1.00 13.68 33.00 VCL (um/s) 144.30 35.23 7.69 67.64 208.70 VAP (um/s) 68.67 13.44 2.93 38.72 92.77 VSL (um/s) 48.65 10.90 2.38 32.11 75.48 LIN (%) 34.19 7.06 1.54 24 50 STR (%) 70.52 7.69 1.68 57 84 WOB (%) 48.05 5.19 1.13 39 60 BCF (Hz) 21.83 1.95 0.43 18.17 26.53 ALH (um) 5.37 1.16 0.25 2.95 7.36 AOC 17.69 3.75 0.82 11.80 24.99 *SD=standard deviation SEM=standard error of the mean Min = Minimum value Max = Maximum value

114 Table 2.5 Breed, Vitamin D concentration, age and fertility data on 22 dogs from which semen was collected in Ontario, Canada, 2009

Dog Breed Vitamin Age No. No. No. D (years) Litters Breedings Years (nmol/l) /year Total 1 Shiloh Shepherd 95 5.5 7 1-2 5 2 Labrador Retriever 113 5 0 0 0 3 Great Pyrenees 182 4.5 2 <1 2 4 Australian Shepherd 102 5.5 0 0 0 5 Newfoundland 189 4.5 0 0 0 6 Golden Retriever 72 5 0 Multiple Unknown 7 Golden Retriever 104 9 0 0 0 8 Whippet 179 11 6 <1 7 9 Cavalier King 117 9 15 2-3 7 Charles Spaniel 10 Belgian Shepherd 186 8.5 0 0 0 11 Labrador Retriever 97 8 3 0.5 6 12 Chesapeake Bay 175 6.5 0 0 0 Retriever 13 Boxer 163 6 0 0 0 14 Labrador Retriever 158 5.5 6 1-2 4 15 Whippet 152 5 0 0 0 16 Mastiff Cross 115 7 0 1 0 17 Golden Retriever 170 4.5 0 0 0 18 Shiloh Shepherd 177 5.5 0 0 0 19 German Shepherd 146 4 0 0 0 20 Welsh Cardigan 128 10 1 3 Unknown Blue Corgi 21 Akita 147 4 2 1 2 22 Samoyed 154 4 0 0 0 No.=number

115 Table 2.6 Mean serum Vitamin D concentration, with respect to type of diet, with associated test-statistic, p-value and confidence intervals.

Type of Diet Mean Serum F-value p-value 95% Confidence Vitamin D (nmol/l) Interval Lower Upper Limit Limit Raw 145.2 0.24 0.79 95.8 194.6 Commercial 140.5 123.4 157.5 Combined 154.0 100.4 207.6

116 Table 2.7 Mean serum Vitamin D concentration, with respect to time of year of sampling, with associated test-statistic, p-value and confidence intervals.

Time of Year Mean Serum t-value p-value 95% Confidence Vitamin D (nmol/l) Interval Lower Upper Limit Limit Summer 139.6 -0.63 0.54 119.8 159.5 Winter 148.1 127.5 168.7

117 Table 2.8 Mean serum Vitamin D concentration, with respect to number of hours spent outdoors at time of year of sampling, with associated test-statistic, p-value and confidence intervals.

Time of Year Mean Serum F- p-value 95% Confidence and number of Vitamin D value Interval hours spent (nmol/l) outdoors Lower Upper Limit Limit Summer • 137.3 0.263 0.85 127.1 147.6 hours Summer < 5 146.8 132.1 161.5 hours Winter • 155.3 141.7 168.7 hours Winter < 5 146.7 130.5 162.8 hours

118 Table 2.9 Mean serum Vitamin D concentration, with respect to BPH status, and associated test-statistic, p-value and confidence intervals.

Prostate Status Mean Serum t-value p-value 95% Confidence 25OHVD (nmol/l) Interval Lower Upper Limit Limit BPH 145.0 0.55 0.59 130.4 159.5 Normal 135.4 81.7 189.1

119 Table 2.10 Univariable simple linear regression analyses of sperm motility parameters with serum 25-hydroxy vitamin D as explanatory variable, with associated coefficient of determination (R2), test statistic and p-value.

Motility Parameter R2 value F-value p-value Total Motility (%) 0.55 10.88 0.0008* Progressive Motility (%) 0.51 9.32 0.002* ALH 0.01 2.99 0.10 AOC 0.11 2.45 0.13 BCF (Hz) 0.28 7.23 0.01* DAP (µm) 0.38 11.61 0.003* VAP (µm/s) 0.35 10.35 0.005* DCL (µm) 0.25 6.28 0.02* VCL (µm/s) 0.25 6.50 0.02* DSL (µm) 0.31 8.53 0.01* VSL (µm/s) 0.27 7.16 0.01* STR 0.00 0.03 0.87 LIN 0.01 0.11 0.75 WOB 0.01 0.18 0.68 ,QGLFDWHVVLJQLILFDQWDVVRFLDWLRQ S”

120 Table 2.11 Non-significant bivariable linear regressions of motility with explanatory variable regression coefficients (where applicable), coefficients of determination for the overall model (R2), test statistics, and the associated p- values.

Explanatory Variable Model Fit Outcome Explanatory Regression p- R2- F- p- Variable Variable Coefficient value value value value Overall Model Motility 25OHVD 0.02 0.002 Bodyweight -0.01 0.35 0.43 6.65 0.01 Progressive 25OHVD 0.02 0.005 Motility Bodyweight -0.01 0.36 0.38 5.48 0.01 ALH (model 1) 25OHVD - - Bodyweight - - 0.14 1.14 0.26 ALH (model 2) 25OHVD - - Prostate Volume - - 0.14 1.14 0.26 AOC (model 1) 25OHVD - - Bodyweight - - 0.11 1.16 0.33 AOC (model 2) 25OHVD - - Prostate Volume - - 0.13 1.33 0.29 BCF (model 1) 25OHVD 0.03 0.02 Bodyweight -0.01 0.59 0.29 3.63 0.05 BCF (model 2) 25OHVD 0.03 0.01 Prostate Volume 0.75 0.15 0.35 4.95 0.02 DAP (model 1) 25OHVD 0.10 0.004 Age -0.31 0.55 0.39 5.80 0.01 DAP (model 2) 25OHVD 0.10 0.004 Bodyweight 0.04 0.60 0.39 5.72 0.01 DAP (model 3) 25OHVD 0.10 0.004 Prostate Volume 0.56 0.70 0.38 5.62 0.01 DCL 25OHVD - - Bodyweight - - 0.28 3.41 0.06 DSL (model 1) 25OHVD 0.07 0.01 Age 0.34 0.43 0.33 4.51 0.03 DSL (model 2) 25OHVD 0.07 0.01 Bodyweight -0.03 0.64 0.32 4.21 0.03

121

Table 2.11 (continued) Non-significant bivariable linear regressions of motility with explanatory variable regression coefficients (where applicable), coefficients of determination for the overall model (R2), test statistics, and the associated p- values.

Explanatory Variable Model Fit Outcome Explanatory Regression p- R2- F- p- Variable Variable Coefficient value value value value Overall Model VAP (model 1) 25OHVD 0.23 0.006 Age -0.74 0.56 0.37 5.18 0.02 VAP (model 2) 25OHVD 0.23 0.006 Bodyweight 0.06 0.72 0.38 5.00 0.02 VAP (model 3) 25OHVD 0.23 0.006 Prostate Volume 1.17 0.74 0.36 4.99 0.02 VCL (model 1) 25OHVD 0.23 0.006 Bodyweight 0.33 0.49 0.36 5.00 0.02 VSL (model 1) 25OHVD 0.17 002 Age 0.70 0.51 0.29 3.70 0.05 VSL (model 2) 25OHVD 0.17 0.02 Bodyweight -0.07 0.61 0.28 3.57 0.05 VSL (model 3) 25OHVD - - Prostate Volume - - 0.28 3.42 0.06 STR (model 1) 25OHVD - - Age - - 0.25 2.87 0.08 STR (model 2) 25OHVD - - Bodyweight - - 0.12 1.20 0.32 STR (model 3) 25OHVD - - Prostate Volume - - 0.00 0.03 0.97 LIN (model 1) 25OHVD - - Age - - 0.27 3.13 0.07 LIN (model 2) 25OHVD - - Bodyweight - - 0.10 0.94 0.41 LIN (model 3) 25OHVD - - Prostate Volume - - 0.01 0.05 0.95 WOB (model 1) 25OHVD - - Age - - 0.21 2.31 0.13 WOB (model 2) 25OHVD - - Bodyweight - - 0.06 0.51 0.61 WOB (model 3) 25OHVD - - Prostate Volume - - 0.01 0.09 0.92

122 Table 2.12 Three bivariable linear regressions of morphology parameters and associated coefficients of determination (R2), test-statistic and p-values

Response (%) Explanatory R2-value F-value p-value of Variable with Overall 25OHVD in the Model model Normal Age 0.49 5.05 0.01* Bodyweight 0.28 3.38 0.06 Prostate Volume 0.52 5.77 0.01*

Head Defects Age 0.17 1.78 0.20 Bodyweight 0.16 1.70 0.21 Prostate Volume 0.39 3.47 0.04*

Midpiece Defects Age 0.07 0.66 0.53 Bodyweight 0.03 0.28 0.76 Prostate Volume 0.03 0.25 0.78

Proximal Droplets Age 0.17 1.74 0.21 Bodyweight 0.10 0.94 0.41 Prostate Volume 0.01 0.06 0.95

Distal Droplets Age 0.24 2.70 0.10 Bodyweight 0.04 0.39 0.69 Prostate Volume 0.20 2.14 0.15

Tail Defects Age 0.02 0.19 0.83 Bodyweight 0.00 0.03 0.96 Prostate Volume 0.03 0.22 0.81

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123

Figure 2.1 Predicted sperm motility as a response to normal serum 25OHVD range in dogs. Back-transformation of multiple linear regression model logitMotility=-7.64+0.135VD-0.0004VD2 (R2=0.55, p=0.0008). VD=serum 25- hydroxy vitamin D. CI= 95%Confidence Interval. Predicted Motility (%), Lower CI, Upper CI

124

Figure 2.2 Progressive motility as a response to normal serum 25OHVD range in dogs. Back-transformation of multiple linear regression model logitPMotility=- 7.92+0.128VD-0.0004VD2(R2=0.51, p=0.002). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. Predicted Progressive Motility (%), Lower CI, Upper CI.

125

Figure 2.3 Predicted sperm motility in response to normal serum 25OHVD range at multiple ages. Back-transformation of multiple linear regression model logitMotility=-46.19 +5.66A+9.72lnVD-1.12A*lnVD (R2 = 0.57, p=0.0019). VD=serum 25- hydroxy vitamin D. CI= 95% Confidence Interval. A=Age Predicted Motility (%), Lower CI, Upper CI.

126

Figure 2.4 Predicted sperm motility in response to normal serum 25OHVD range at various transformed prostate volumes. Back-transformation of multiple regression model logitMotility=-16.7 +3.69Vol +0.14VD-0.03Vol*VD (R2=0.72, p<0.0001). VD=serum 25-hydroxy vitamin D. CI= 95% Confidence Interval. Vol=Prostate Volume. Predicted Motility (%), Lower CI, Upper CI.

127

Figure 2.5 Predicted progressive motility in response to normal serum 25OHVD range at various ages. Back- transformation of multiple regression model logitPMotility=-8.1+1.11A +0.06VD-0.007A*VD (R2=0.57, p=0.002). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age. Predicted Progressive Motility (%), Lower CI, Upper CI.

128

Figure 2.6 Predicted progressive motility in response to normal serum 25OHVD range at various transformed prostate volumes. Back-transformation of multiple regression model logitPMotility=-14.2 +3.0Vol +0.11VD -002Vol*VD (R2=0.60, p=0.001). VD=serum 25-hydroxy vitamin D. CI= 95% Confidence Interval. Vol=Prostate Volume. Predicted Progressive Motility (%), Lower CI, Upper CI.

129

Figure. 2.7 Predicted BCF in response to normal serum 25OHVD range at various ages. Multiple regression model BCF=14.9+0.422A +0.03VD (R2=0.47, p=0.003). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age Predicted BCF (%), Lower CI, Upper CI.

130

Figure. 2.8 Predicted DCL in response to normal serum 25OHVD range at various ages. Multiple regression model DCL=-56.5 +0.9VD+14.5A-0.11VD*A (R2=0.51, p=0.006). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age Predicted DCL (um), Lower CI, Upper CI.

131

Figure 2.9 Predicted DCL in response to normal serum 25OHVD range at various transformed prostate volumes. Back- transformation of multiple regression model DCL=-1790.1+488.7Vol+305.7lnVD -65.9Vol*lnVD-18.8Vol2 (R2=0.53, p=0.01). VD=serum 25-hydroxy vitamin D. CI= 95% Confidence Interval. Vol=Prostate Volume. Predicted DCL (um), Lower CI, Upper CI.

132

Figure 2.10 Predicted VCL in response to normal serum 25OHVD range at various ages. Multiple regression model VCL=-129.7+2.0VD+32.5A -0.11VD*A (R2=0.51, p=0.006). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age. Predicted VCL (um/s), Lower CI, Upper CI.

133

Figure 2.11 Predicted VCL in response to normal serum 25OHVD range at various transformed prostate volumes. Multiple regression model VCL=-1405.2+525.2Vol+5.6VD -1.2Vol*VD-40.79Vol2 (R2=0.52, p=0.01). VD=serum 25- hydroxy vitamin D. CI=95% Confidence Interval. Vol=Prostate Volume. Predicted VCL (um/s), Lower CI, Upper CI.

134

Figure 2.12 Predicted DSL in response to normal serum 25OHVD range at various transformed prostate volumes. Multiple regression model DSL=-34.1+10.3Vol+0.403VD-0.075Vol*VD (R2= 0.46, p=0.01). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. Vol=Prostate Volume. Predicted DSL (um), Lower CI, Upper CI.

135

Figure 2.13 Predicted ALH in response to normal serum 25OHVD range at various ages. Multiple regression model ALH=-3.1+1.1*A+0.06VD-0.01A*VD (R2=0.42, p=0.02). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age. Predicted ALH (um), Lower CI, Upper CI.

136

Figure 2.14 Predicted AOC in response to normal serum 25OHVD range at various ages. Multiple regression model AOC=-6.6+3.1*A+0.2VD-0.02A*VD (R2=0.38, p=0.04). VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age. Predicted AOC (degrees), Lower CI, Upper CI

137

Figure 2.15 Predicted Normal sperm in response to normal serum 25OHVD range Back-transformation model logitNormal=-17.79+0.246VD-0.0008VD2 (R2=0.43, p=0.01). Predicted Normal Sperm (%), Lower CI, Upper CI. VD=serum 25-hydroxy vitamin D. CI= 95% Confidence Interval.

138

Figure 2.16 Predicted Loose Heads in response to normal serum 25OHVD range. Back-transformation of logitLooseHeads=-1.62+0.22VD (R2=0.31, p=0.01). Predicted Loose Heads (%), Lower CI, Upper CI. VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval.

139

Figure 2.17 Predicted Normal sperm in response to normal serum 25OHVD range at various ages. Back-transformation of multiple regression model logitNormal=-17.59+2.09A +0.12VD-0.01A*VD (R2=0.49, p=0.01). Predicted Normal Sperm (%), Lower CI, Upper CI. VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. A=Age.

140

Figure 2.18 Predicted Normal sperm in response to normal serum 25OHVD range at various transformed prostate volumes. Back-transformation of multiple regression logitNormal=-29.2+5.77Vol +0.21VD-0.04Vol*VD (R2=0.52, p=0.01 Predicted Normal Sperm (%), Lower CI, Upper CI. VD=serum 25-hydroxy vitamin D. CI= 95% Confidence Interval. Vol=Prostate Volume.

141

Figure 2.19 Predicted head defects in response to normal serum 25OHVD range at various transformed prostatic volumes. Back-transformation of multiple logitHead=24.5-5.55Vol-0.19VD+0.04Vol*VD (R2=0.39, p=0.04). Predicted Head Defects (%), Lower CI, Upper CI. VD=serum 25-hydroxy vitamin D. CI=95% Confidence Interval. Vol=Prostate Volume.

142 References

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144 [33] L. Almeras, et al., "Developmental vitamin D deficiency alters brain protein expression in the adult rat: implications for neuropsychiatric disorders," Proteomics, vol. 7, pp. 769-80, Mar 2007. [34] K. Kadam, et al., "Spatial distribution of actin and tubulin in human sperm nuclear matrix-intermediate filament whole mounts-a new paradigm," Microsc Res Tech, vol. 70, pp. 589-98, Jul 2007. [35] K. Nishio and A. Inoue, "Senescence-associated alterations of cytoskeleton: extraordinary production of vimentin that anchors cytoplasmic p53 in senescent human fibroblasts," Histochem Cell Biol, vol. 123, pp. 263-73, Mar 2005. [36] Y. Azamar, et al., "F-actin involvement in guinea pig sperm motility," Mol Reprod Dev, vol. 74, pp. 312-20, Mar 2007. [37] S. Nagpal, et al., "Noncalcemic actions of vitamin D receptor ligands," Endocr Rev, vol. 26, pp. 662-87, Aug 2005. [38] S. J. Berry, et al., "Effects of aging on prostate growth in beagles," Am J Physiol, vol. 250, pp. R1039-46, Jun 1986. [39] C. B. Brendler, et al., "Spontaneous benign prostatic hyperplasia in the beagle. Age-associated changes in serum hormone levels, and the morphology and secretory function of the canine prostate," J Clin Invest, vol. 71, pp. 1114-23, May 1983. [40] J. E. Branam, et al., "Selected physical and chemical characteristics of prostatic fluid collected by ejaculation from healthy dogs and from dogs with bacterial prostatitis," Am J Vet Res, vol. 45, pp. 825-9, Apr 1984. [41] G. Penna, et al., "Seminal plasma cytokines and chemokines in prostate inflammation: interleukin 8 as a predictive biomarker in chronic prostatitis/chronic pelvic pain syndrome and benign prostatic hyperplasia," Eur Urol, vol. 51, pp. 524-33; discussion 533, Feb 2007. [42] G. Penna, et al., "The vitamin D receptor agonist elocalcitol inhibits IL-8- dependent benign prostatic hyperplasia stromal cell proliferation and inflammatory response by targeting the RhoA/Rho kinase and NF-kappaB pathways," Prostate, vol. 69, pp. 480-93, Apr 1 2009. [43] R. J. Aitken, "Diagnostic value of the zona-free hamster oocyte penetration test and sperm movement characteristics in oligozoospermia," Int J Androl, vol. 8, pp. 348-56, Oct 1985. [44] C. Jeulin, et al., "Sperm factors related to failure of human in-vitro fertilization," J Reprod Fertil, vol. 76, pp. 735-44, Mar 1986. [45] T. A. Bongso, et al., "Effect of sperm motility on human in vitro fertilization," Arch Androl, vol. 22, pp. 185-90, 1989. [46] C. Y. Cheng, et al., "Regulation of spermatogenesis in the microenvironment of the seminiferous epithelium: new insights and advances," Mol Cell Endocrinol, vol. 315, pp. 49-56, Feb 5 2010. [47] P. P. Lie, et al., "The biology of interleukin-1: emerging concepts in the regulation of the actin cytoskeleton and cell junction dynamics," Cell Mol Life Sci, Jul 9 2011.

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146 CHAPTER THREE

PROLACTIN, BENIGN PROSTATIC HYPERPLASIA,

PROSTATE VOLUME AND SEMEN PARAMETERS IN THE DOG.

Abstract

Prolactin (PRL) is well known for its role in pregnancy and lactation in the bitch, however; the role of PRL in male dogs has not been fully characterized.

The aim of this observational study was to determine whether PRL is associated with benign prostatic hyperplasia (BPH), prostate volume and semen quality characteristics in stud dogs.

A convenience sample of 29 healthy dogs consisting of 24 different breeds ranging in age from two to 11 years was studied to determine whether an association exists between serum PRL concentrations and BPH. Of the dogs that successfully gave semen samples (n=22; 17 breeds; age range: 4 to 11 years), associations between PRL and multiple characteristics of semen motility and morphology were also investigated. Student’s t-test revealed no difference in serum PRL concentrations between dogs with and without BPH (t-value=0.87, p=0.39). Bivariable linear regression analysis revealed no associations between prostate volume and PRL when controlling for BPH (R2=0.04, p=0.31). No significant associations were found between PRL and any of the 14 sperm motility parameters, as determined by computer assisted motility analysis

(CASA). Two sperm morphology parameters had significant associations with

PRL: percentage proximal droplets (R2=0.23, p=0.03) and percentage midpiece

147 defects with an age interaction (R2=0.42, p=0.03). Serum PRL concentrations less than 2.5ng/ml were associated with higher percentages of midpiece defects and proximal cytoplasmic droplets in canine sperm.

Serum PRL concentration ranges in this study of normal healthy dogs

(range: undetectable to 28.24 ng/ml) were higher than previously reported values

(range: undetectable to 6.0 ng/ml) using the same validated ELISA kit. In conclusion, PRL concentrations had no effect on BPH or sperm motility parameters, but had a limited negative effect on sperm morphological defects, namely percentage proximal droplets and midpiece defects.

Keywords: prolactin, prostate, semen, dog, benign prostatic hyperplasia

Abbreviations: Amplitude of lateral head displacement (ALH), Average orientation change (AOC), Beat cross frequency (BCF), Mean distance (DAP),

Curvilinear distance (DCL), Straight line distance (DSL), Linearity (LIN =

VCL/VSL), Straightness (STR), Mean velocity (VAP), Curvilinear velocity (VCL),

Linear velocity (VSL µm/s), Wobble (WOB=VAP/VCL).

1. Introduction

Prolactin (PRL) is a 199 amino acid (23kDa) protein hormone related to both growth hormone and placental lactogen [1]. It is well known for its role in pregnancy/pseudopregnancy and lactation in bitches [2]. Other non-traditional physiological roles in vertebrate species are still being elucidated including water and electrolyte balance, growth and development, endocrinology and metabolism, brain function and behaviour, and immunoregulation [3]. Although

148 its role in reproduction and fertility in men has been studied in relation to pathological conditions such as prostatic atrophy and teratozoospermia [4, 5], little is known about the role of canine PRL in the reproductive physiology of the stud dog.

The PRL feedback loop in male rodents and humans has been shown to work in the following manner. Prolactin is produced stored and released in the lactotroph cells of the anterior pituitary similarly to luteinizing hormone (LH) and follicle stimulating hormone (FSH). PRL upregulates LH receptors in Leydig cells, thereby stimulating increased secretion of testosterone [1]. Increased testosterone stimulates estradiol-17E (E2) production by Sertoli cells which in turn acts on the pituitary lactotroph cells in a positive feedback loop to secrete more

PRL. A direct negative feedback mechanism is accomplished through E2 and testosterone inhibition of GnRH secretion in the hypothalamus and LH/FSH secretion in the pituitary. Indirectly, GnRH inhibition of LH/FSH secretion also down-regulates testosterone secretion [6].

In the rat, prolactin has been suggested to work synergistically with testosterone to increase 5D-reductase activity in both in vitro and in vivo studies, and induces growth, differentiation, and hyperplastic changes of the prostate [7-

9]. PRL is considered to have stimulatory effects on prostate growth and development as its receptors are present in fibromuscular and epithelial cells, as well as in focal glandular hyperplastic cells, in experimentally induced BPH [10-

12]. PRL and its binding sites have been identified in the canine prostate.

However, PRL differs in its action by down-regulation of 5D-reductase activity,

149 resulting in a decrease in prostatic DHT concentrations in canine prostatic cells studied in vitro [10]. This suggests a possible role of PRL in atrophy or decreased size of the prostate by acting on intraprostatic DHT concentrations in the dog.

The effect of experimentally induced hyperprolactinemia in dogs has been studied. Shafik et al (1994) used high doses (600ug/kg/week) of ovine PRL on a population of male beagles to determine whether any contraceptive effects could be shown [13]. Their treatment resulted in a negative impact on both semen morphology and motility after two weeks of treatment with degenerative changes of the seminiferous tubules and complete azoospermia by six months of treatment [13]. Although treatment of dogs with high doses of a non-canine PRL can be useful as a tool for contraception, it does not provide information on the normal reproductive physiology of PRL in the dog nor on possible naturally occurring PRL related reproductive pathologies.

Semen quality studies in hyperprolactinemic rats and men reported oligozoospermia, asthenozoospermia and teratozoospermia [4, 5, 14]. In conditions causing hyperprolactinemia, increased E2 results in low testosterone concentrations and clinical manifestations of hypogonadism and decreased semen quality [6]. In the dog, published long-term studies of the effects of spontaneous or induced hyperprolactinemia with canine PRL on semen quality do not exist. Short-term hyperprolactinemia using dopamine antagonists did not alter semen characteristics, while short-term hypoprolactinemia using dopamine agonists resulted in slight alterations on one sperm motion characteristic (VSL)

150 as assessed by CASA [15]. Clearly, minimal data exists on the role of PRL in sperm and prostate physiology in dogs.

The purpose of this study was to investigate the associations of PRL with

BPH, prostate volume and semen quality characteristics in stud dogs. We hypothesized that high serum PRL concentrations would be negatively associated with BPH, prostate volume and semen quality [16].

2. Materials and Methods

2.1 Case Subjects

A convenience sample of 29 healthy intact male dogs belonging to 24 breeds with varying fertility histories, with or without indication of prostate disease was obtained between March and December of 2009. Dogs were recruited from cases presented to the Theriogenology Service at the Ontario Veterinary College

(Guelph, Ontario) and from a small private clinic in Southwestern Ontario with a large breeder clientele. All clients were required to fill out a consent form

(Appendix 1) and brief questionnaire (Appendix 2) outlining their dogs’ fertility history, diet and amount of time their dogs spent outdoors. All dogs underwent a complete male breeding soundness examination, blood and urine collection as described in the Materials and Methods section in Chapter Two.

2.2 Semen collection, dilution and staining.

Twenty-two of the 29 dogs (75.9%) enrolled in the study had semen collected. Semen processing was completed according to the Materials and

151 Methods section of Chapter Two.

2.3 Motility Analysis

All semen motility analysis was undertaken according to the protocols set out in the Materials and Methods section of Chapter Two. Technical settings for computer assisted semen analysis (CASA) were programmed according to

Schafer-Somi and Aurich (2009) (Table 3.1) [17].

2.4 Prostate Examination

The prostate gland of all dogs was evaluated after semen collection by several methods, as described in the Materials and Methods section in Chapter

Two.

2.5 Ultrasonography of the Prostate

Ultrasonography of the prostate was accomplished according the Materials and Methods section in Chapter Two. Table 3.2 describes the criteria by which

BPH was diagnosed according to ultrasound measurements of the prostate.

2.6 Prolactin Assay

Serum samples were batched and shipped overnight on dry ice to the

Endocrinology Laboratory, Washington State University. Prolactin concentrations were measured using a commercially available validated ELISA for canine prolactin [16] according to manufacturer instructions (Milenia Canine prolactin

152 (MKVCP-1), Milenia Biotec distributed by Alpco Immunoassays, Salem NH). As samples were batched and run in the same assay no inter-assay variation was present. The intra-assay coefficient of variation (CV) was 5.2% with a sensitivity of 0.4 ng/ml. The lowest PRL concentration standard used in the assay was 2.5 ng/ml.

2.7 Urinalysis

Urinalysis was completed according to the protocols set out in the

Materials and Methods section in Chapter Two.

2.8 Statistical Analyses

2.8.1 Overall Statistical Analysis

Statistical analyses of the data from individual dogs was performed using

SAS version 9.2 software (SAS Institute Inc. Cary, NC, USA). All statistical analyses of both univariable and bivariable associations were completed using the general linear model (GLM) procedure as described in the Material and

Methods of Chapter Two. No more than two explanatory variables were used in any analysis due to the small sample size to avoid over-fitting the data to the statistical models. Assumptions of normality were tested using the UNIVARIATE procedure, as described in Chapter Two, and data transformations were used when necessary. A logit transformation was used in the case of percentage values and a small bias correction term added in order to accommodate zero values (Logit outcome variable = log((r+k)/(n-r+k)); r= number of responding

153 cells, n=number of cells counted, k= bias correction term=0.25). In cases where transformations were not successful in normalizing the data, the Monte Carlo exact non-parametric test was used to determine if simple univariable correlations were significant. P-YDOXHVRI”ZHUHFRQVLGHUHGVWDWLVWLFDOO\VLJQLILFDQWIRUDOO analyses, and 95% confidence limits were used. The Grubbs’ test for outliers was used on the residuals[18] and models were rerun with removal of these points as described in the Materials and Methods in Chapter Two.

2.8.2 Study Population

Descriptive statistics of the characteristics of the study population of dogs were completed as per the Materials and Methods of Chapter Two.

2.8.3 BPH and Prostate Volume

The Student’s t-test was used to detect whether there was a difference in mean prolactin concentrations (PRL) between dogs with and without BPH.

Univariable analysis was also completed with age and bodyweight as explanatory variables with PRL as the response variable. Bivariable analysis was performed using the explanatory variable PRL and the outcome variable prostate volume, controlling for benign prostatic hyperplasia analytically by including it in the model.

2.8.4 Sperm Motility

Univariable and bivariable analyses of semen motility as previously described included the following outcome variables: total motility, progressive

154 motility, ALH, AOC, BCF, DAP, DCL, DSL, VAP, VCL, VSL, LIN, STR, and WOB.

PRL, age, bodyweight and prostatic volume were each tested as explanatory variables. Bivariable analyses included the explanatory variable combinations of

PRL and age, PRL and prostate volume, and PRL and bodyweight.

2.8.5 Sperm Morphology

Univariable and bivariable analyses of semen morphology included the following outcome variables: normal sperm, head defects, midpiece defects, tail defects, loose heads, proximal droplets and distal droplets. PRL, age and bodyweight were each tested as explanatory variables. Bivariable analyses included the explanatory variable combinations of PRL and age, PRL and prostate volume and PRL and bodyweight.

3. Results

3.1 Study Population

The mean serum prolactin concentration of the study population (n=29) was 4.19 ng/ml r 1.16 ng/ml, and concentrations ranged between undetectable to

28.24 ng/ml. The mean serum prolactin concentration of part of the population from whom semen was collected (n=22) was 5.23 ng/ml ± 1.46 ng/ml. All other characteristics of the study population of dogs can be found in Table 3.3 and in

Chapter Two (Table 2.4).

155 3.2 BPH and Prostate Volume

The mean serum PRL concentration did not significantly differ between dogs with BPH and dogs without (t-value=0.87, p=0.39), the details of which are shown in Table 3.4. No significant univariable associations were observed between PRL and age (R2=0.04, p=0.31) or bodyweight (R2=0.01, p=0.54).

There was also no significant association between serum PRL and prostate volume when controlling for BPH in the model (R2=0.04, p=0.31).

3.3 Sperm Motility

No significant associations between PRL and semen motility characteristics of total motility, progressive motility, ALH, AOC, BCF, DAP, DCL,

DSL, LIN, STR, VAP, VCL, VSL, and WOB were found using univariable simple linear regression analysis (Table 3.5) or bivariable analyses (Table 3.6).

3.2 Semen Morphology

PRL was significantly associated with proximal droplets in univariable analyses (Table 3.7, Figure 3.1). Overall there was a decrease in proximal droplets with increasing PRL concentrations. No other significant univariable associations between PRL and sperm morphology parameters were found (Table

3.7).

The final bivariable model of midpiece defects was significant with PRL and age

(Table 3.8, Figure 3.2). Overall, there was a decrease in midpiece defects with increasing PRL concentrations. Percentages of midpiece defects were lowest at

156 4 years of age and highest at 6 years of age with a decrease in midpiece defects from 8 to 10 years of age. No other bivariable analyses showed significant associations with serum PRL (Table 3.8). The overall bivariable models with proximal droplets as a response to PRL were significant, although the regression slope of the second variable were not (Table 3.9). This result indicated a strong influence of PRL on the analysis and in these cases the model was further simplified to its univariable form.

3.3 Outlier Removals

Outliers were identified and removed from the model of normal sperm and

PRL. The results of this can be viewed in Appendix 5. Removal of three outliers

(dogs 6, 1, and 15) revealed associations between normal sperm morphology and PRL (R2=0.36, p=0.01), normal sperm morphology and PRL with age

(R2=0.66, p=0.0005), and normal sperm morphology and PRL with prostate volume (R2= 0.52, p=0.006) (see Appendix 5). Univariable analysis, with these outliers omitted, between normal sperm morphology and PRL showed an increasing percentage of normal sperm with higher concentrations of PRL within the range of the study population. Dogs with PRL values less than 2.5 ng/ml had much lower percentages of normal sperm. In bivariable analyses, there was a similar pattern of increasing percentage of normal sperm with increasing prolactin, however, at older ages and higher prostate volumes there was a lower percentage of normal sperm compared to dogs at younger ages and smaller prostate volumes. Although, the removal of outliers in large data sets may be

157 advised we maintained the data due to the small sample size and possibility of removing valuable information from the analysis.

4. Discussion and Conclusions

The physiology and role of prolactin in the stud dog is poorly understood.

The purpose of this observational study was to determine whether PRL is associated with BPH and prostate volume and associated with sperm motility and morphology characteristics in intact male dogs of various breeds.

A significant unconditional effect of PRL on proximal droplets was observed, with a higher percentage of proximal droplets (20%; 95% CI 3.5%-

62%) at serum PRL concentrations below 2.5ng/ml and a small yet significant decrease in proximal droplet percentages at PRL concentrations at or above 2.5 ng/ml. Proximal droplets, which represent excess cytoplasm in the final maturation process, move distally from the proximal to distal midpiece region before being released from the epididymis. Prolactin concentrations were also significantly associated with midpiece defects of sperm, but this effect was shown to be dependent on the age of the dog. An increase in percentage of midpiece defects was seen with PRL concentrations below 2.5 ng/ml throughout all age groups with significant decreases in those defects at higher PRL concentrations.

Although the general trend in all age groups shows a small yet significant decrease in midpiece defects as PRL concentrations increase over 2.5ng/ml, it is unclear why middle-aged dogs (6 years) have the largest percentages of midpiece defects at PRL concentrations less than 2.5 ng/ml and older dogs (10

158 years) have the lowest. It was initially thought that one Mastiff cross (Dog 16) in the study with bilateral interstitial cell tumours and a high percentage of midpiece defects (20%) was influencing the results. However, removal of this dog from the data set did not alter the results of the analysis. These results relating to proximal droplets and midpiece defects suggest that PRL may play a role in epididymal sperm maturation and/or spermatogenesis and that an age dependent effect on midpiece defects is apparent, although no other evidence exists to support this hypothesis in the dog.

PRL, and its receptor (PRLR), have been identified in seminiferous tubules, developing spermatozoa and epididymal tissue in men and rats [19-21] and in the testicular tissue of rams and pigs [22, 23], although it has yet to be studied in the dog. PRL has also been shown to increase rat testicular tissue testosterone levels in vitro by increasing LH receptors in Leydig cells [24]. It is reasonable to assume then that a similar PRL function and PRLR localization in the dog exists and may have relevance with the results of our study. This previous research in humans and other species also supports the hypothesis that normal PRL levels may be necessary for normal development of sperm both at the level of spermatogenesis within the seminiferous tubules and maturation of sperm within the epididymis itself. Alternatively, PRL may act indirectly on sperm by potentiating or modifying testosterone action during spermatogenesis and maturation through action on either Sertoli cells and/or epididymal tissue through the androgen receptors located in these areas. The latter hypothesis may also explain why reports of hypoprolactinemia on fertility do not exist in men and why

159 fertility rates in male PRL knock-out mice do not show alterations in pregnancy rates [25]. Although our study detected increased proximal droplets and midpiece defects at PRL concentrations less than 2.5 ng/ml, the amount of normal sperm was still in excess of 70% and therefore a perceptible impact on fertility would be unlikely.

Outlier removals (Appendix 5) showed similar results with respect to normal sperm and PRL. Again, dogs with PRL values less than 2.5 ng/ml had substantially lower percentages of normal sperm once these outliers were removed. This suggests that these PRL values may result in defects of spermatogenesis as previously discussed. Ultimately, a lack of evidence for other causes of decreased morphologically normal sperm to support removal of apparent statistical outliers, as well as a small sample size in this study, were factors in maintaining them in the analysis.

Serum PRL concentrations in excess of 13 ng/ml in men were associated with infertility and poor sperm motility and morphology [4]. Serum concentrations of PRL in excess of 20 ng/ml in men is considered hyperprolactinemia [26].

However, the results of the current study suggest that concentrations as high as

28.24 ng/ml are within the normal reproductive range for dogs as no adverse effects on semen quality were observed at this concentration. Concentrations below 2.5ng/ml could be considered hypoprolactinemic as impacts on sperm quality were observed. Hypoprolactinemia has been associated with metabolic syndrome in humans [27]. None of our study subjects had a history of diabetes or presented as obese, however it is possible that subclinical cases may have been

160 present. Thyrotrophin releasing hormone (TRH) is a stimulator of PRL secretion by lactotroph cells of the pituitary. The absence of clinical signs of hypogonadism or lethargy, alopecia and skin disorders due to hypothyroidism suggest that the highest values for PRL obtained in our study were within normal limits. Prolactinoma, or naturally- or pharmacologically-induced hyperprolactinemia are rare occurrences in dogs, therefore research about hyperprolactinemia and its pathology outside of pregnancy and lactation in dogs is scarce. Overall, the absence of significance in the remaining sperm motility and morphology parameters may be due to the small sample size in this study.

In order to increase the power and sensitivity of the analysis, a sample size of 88 or greater dogs would be needed. A long-term study to recruit adequate numbers may resolve this issue. We did not perform a serum biochemical analysis as part of our evaluation of the study dogs and therefore were not able to evaluate other conditions including metabolic disease.

Use of the Milenia Canine Prolactin ELISA was chosen as it has recently been validated for dogs [16] with a published normal reference range

(undetectable to 6.0ng/ml) and is a commercial assay that would be readily available to practising clinicians. The choice was therefore made with practical considerations in mind. Although RIA is the gold standard for research purposes, any clinically significant findings may not be transferable in dealing with clinical cases of infertility in patients. The high end of the range for PRL differed from the high end of the range in our study (6.0 ng/ml compared with 28.24 ng/ml).

Although Corrada et al found one individual had higher PRL concentrations

161 compared to the other dogs in that study [16], five dogs from our population (one each of Samoyed, Labrador Retriever, German Shepherd Dog, Golden Retriever, and Welsh Cardigan Blue Corgi) had similar PRL concentrations to the excluded individual. These differences between studies may be due to the differences in sample size (n=65 compared to n=20) and over-representation of Beagles and

German Shepherd Dogs in the reference range study [16]. Study of a larger population including many breeds, sizes and ages would clarify whether these high values are truly normal in the overall intact canine male population. We could then define hyper- and hypoprolactinemia and establish breed-specific normal ranges for serum PRL concentrations to aid in the diagnosis and treatment of reproductive conditions.

No significant associations between PRL and sperm motility characteristics were observed in our study. This is in contrast to the work of

Koivisto et al, (2009), who found that short-term experimentally induced hypoprolactinemia with cabergoline was associated with decreased VSL [15].

The discrepancies between studies may be explained by several factors.

Koivisto’s study population consisted of only beagles and did not examine other breeds. Beagles were shown to have higher serum concentrations of PRL in one study [16] while in another study Fox Terriers were shown to have significantly lower PRL concentrations [28], therefore breed effects are relevant. The dogs in

Koivisto’s study were aged 12 months to 2 years compared to our study group of

2 to 11 years. It is possible that our study was able to find age related effects due to including dogs of broader ages. Koivisto’s study evaluated the effects of

162 experimentally induced hypoprolactinemia using treatment with cabergoline for 3 weeks, while our study represented a one-time observation of semen quality and

PRL levels. A single measurement may not reflect the overall PRL status of our dogs. Conversely, the cabergoline treatment may not have been long enough to elicit chronic morphological effects as seen in our study. Methods used to measure PRL also differed in the two studies, although ranges and means were comparable with our validated ELISA to RIA studies. In addition, our small sample size may not have provided enough power to the study to detect changes in VSL in our study population. Not all dogs examined were able to provide semen samples for the study. The absence of an estrous bitch during semen collection and the sexual inexperience of some of the study subjects lead to failure of adequate sexual stimulation in these cases.

No significant associations between PRL concentrations and BPH or prostate volume were found in the present study. It is important to note that PRL release is pulsatile in nature and the lack of multiple blood sampling during the study may not have given us an accurate representation of overall PRL concentrations. Further investigation by increasing the number and frequency of subject sampling and increasing the number of dogs enrolled in a study of this nature is recommended.

PRL has been shown to decrease in vitro prostatic DHT production in the dog

[10] and may have a role to in prostatic pathology. Whether the mechanism involved in decreasing prostatic DHT is by direct action of PRL on prostatic epithelial cells or due to a chronic negative feedback of PRL on testosterone

163 secretion is unknown and cannot be determined by the results obtained here.

The diagnosis of BPH in this study used the generally accepted clinical signs of enlarged prostatic dimensions by digital palpation per rectum, a predetermined classification of enlarged prostate with respect to bodyweight, symmetry of the gland on ultrasound examination and/or the presence of blood in the ejaculate.

BPH is most accurately determined using histology as the gold standard, and some cases reported here might not have had histological evidence of BPH.

Further research using prostatic biopsy is recommended and may reveal associations between PRL and BPH. Biopsy and/or fine needle aspirate of the prostate in our study would have been an unwarranted risk in otherwise healthy client dogs. We were also unable to control for bodyweight and age simultaneously in evaluating prostate volume with PRL concentrations.

In rats, PRL has shown delayed long-term feedback on spermatogenesis and the prostate [29, 30]. Long-term hyperprolactinemia results in decreased serum testosterone concentrations and teratozoospermia in men [4], so an effect of PRL on certain aspects of sperm development related to testosterone concentrations in dogs may be plausible. Analyzing concentrations of other hormones such as testosterone, estradiol (as a modifier of PRL secretion), thyroid stimulating hormone (as an index of TRH concentration), and LH/FSH in our study population, may add to furthering our understanding of PRL in its role in reproductive physiology. By further investigating and identifying PRL and/or its receptor in canine testes and sperm cells, one may better understand the specifics and localization of PRL effects on the dog.

164 In conclusion, low serum PRL concentration was associated with higher percentages of proximal droplets and midpiece defects; the latter also increased with age at PRL concentrations less than 2.5 ng/ml. No effects of serum PRL concentration on BPH, prostate volume or other semen parameters related to motility and morphology were observed. PRL appears to have minimal reproductive effects in the dog and may not have equal importance in reproductive pathologies seen in other species, such as man, however; further study on a larger scale is required. The information obtained indicates that in some cases of sub-fertility related to defects of the sperm midpiece and proximal droplets, maintenance of PRL concentrations above 2.5 ng/ml may be desirable to improve reproductive potential in individual dogs.

165 Table 3.1 Technical Parameters for SpermVision™ Computer Assisted Semen Analysis used on samples from 29 dogs, March-December 2009, Ontario, Canada

Parameter Depth of chamber 20um Light adjustment 96-104 Volume per Chamber 2ul Temperature 37°C Sperm concentration Variable Area of sperm heads 20-60 um2 Number of fields or Cell number 7 or 1000 Number cells/field 50-200 Frame rate 60/sec Total motility VCL > 15 Progressively Fast VCL >15, STR>0.9 Linear STR >0.9, LIN >5 Immotile AOC <7, DSL <3 Local motility DSL <6 Hyperactive VCL>118, ALH>6.5, LIN<0.5 Non-Linear LIN<=0.5, STR<=0.9 Curvilinear DAP/Radius >=3, LIN<0.5 Incubation time 2 mins Modified from Schäfer-Somi and Aurich, 2007[17]

166 Table 3.2 Classification scheme for determination of the presence of benign prostatic hyperplasia by ultrasonography in 29 dogs, March-December 2009, Ontario, Canada

Prostatic Measurement Bodyweight Maximum length, width or height > 3.0 cm ” 20 kg > 3.5 cm >20 to ” 40 kg > 4.0 cm !WR” 60 kg > 4.5 cm !WR” 80 kg Classification based on arbitrary cut-points.

167 Table 3.3 Breed, prolactin concentration, age and fertility data on 22 dogs from which semen was collected in Ontario, Canada, 2009

Dog Breed Prolactin Age No. No. No. (ng/ml) (years) Litters Breedings Years /year Total 1 Shiloh Shepherd 4.14 5.5 7 1-2 5 2 Labrador Retriever 0.98 5 0 0 0 3 Great Pyrenees 0.92 4.5 2 <1 2 4 Australian 2.02 5.5 0 0 0 Shepherd 5 Newfoundland 4.57 4.5 0 0 0 6 Golden Retriever 5.68 5 0 Multiple Unknown 7 Golden Retriever 0 9 0 0 0 8 Whippet 0 11 6 <1 7 9 Cavalier King 0.11 9 15 2-3 7 Charles Spaniel 10 Belgian Shepherd 0.31 8.5 0 0 0 11 Labrador Retriever 17.47 8 3 0.5 6 12 Chesapeake Bay 6.56 6.5 0 0 0 Retriever 13 Boxer 2.07 6 0 0 0 14 Labrador Retriever 3.96 5.5 6 1-2 4 15 Whippet 3.46 5 0 0 0 16 Mastiff Cross 2.07 7 0 1 0 17 Golden Retriever 11.49 4.5 0 0 0 18 Shiloh Shepherd 1.59 5.5 0 0 0 19 German Shepherd 9.89 4 0 0 0 20 Welsh Cardigan 9.59 10 1 3 Unknown Blue Corgi 21 Akita 0 4 2 1 2 22 Samoyed 28.24 4 0 0 0

168 Table 3.4 Mean serum PRL concentrations and confidence intervals with respect to Benign Prostatic Hyperplasia (BPH) status

Prostate Status Mean serum t- p-value 95% Confidence Interval PRL (ng/l) Value Lower Limit Upper Limit BPH 4.4 0.87 0.39 1.6 7.3 Non-hyperplastic 3.0 2.6 5.5

169 Table 3.5 Univariable analyses using simple linear regression modeling of semen motility parameters and PRL with coefficients of determination (R2) of the overall model, test statistic and associated p-values.

Semen Parameter R2-value F-statistic p-value Total Motility (%) 0.01 0.23 0.64 Progressive Motility (%) 0.03 0.64 0.43 ALH (um) 0.02 0.42 0.52 AOC 0.05 0.91 0.35 BCF (Hz) 0.08 1.70 0.21 DAP (um) 0.01 0.10 0.75 DCL (um) 0.04 0.79 0.38 DSL (um) 0.00 0.00 0.99 VAP (um/s) 0.01 0.11 0.75 VCL (um/s) 0.04 0.77 0.39 VSL (um/s) 0.00 0.00 0.97 STR 0.01 0.28 0.60 LIN 0.05 0.95 0.34 WOB 0.11 2.26 0.15

170 Table 3.6 Three bivariable linear regressions of motility parameters with coefficients of determination of overall model (R2), test statistics, and associated p-values.

Response Explanatory R2- F-Statistic p-value of Variables with value Linear Model PRL in the model Total Motility Age 0.01 0.12 0.89 Bodyweight 0.03 0.29 0.75 Prostate Volume 0.01 0.11 0.90 Progressive Motility Age 0.03 0.33 0.73 Bodyweight 0.05 0.48 0.62 Prostate Volume 0.05 0.33 0.73 ALH Age 0.10 1.00 0.39 Bodyweight 0.02 0.22 0.80 Prostate Volume 0.02 0.21 0.81 AOC Age 0.13 1.36 0.28 Bodyweight 0.05 0.43 0.66 Prostate Volume 0.06 0.57 0.58 BCF Age 0.20 2.30 0.13 Bodyweight 0.08 0.82 0.46 Prostate Volume 0.16 1.66 0.22 DAP Age 0.02 0.17 0.84 Bodyweight 0.02 0.18 0.84 Prostate Volume 0.01 0.09 0.91 DCL Age 0.10 1.02 0.38 Bodyweight 0.07 0.64 0.54 Prostate Volume 0.04 0.40 0.68

171 Table 3.6 (continued) Three bivariable linear regressions of motility parameters with coefficients of determination of overall model (R2), test statistics, and associated p-values.

Response Explanatory R2- F-Statistic p-value of Variables with value Linear Model PRL in the model DSL Age 0.02 0.19 0.84 Bodyweight 0.00 0.04 0.96 Prostate Volume 0.00 0.02 0.98 VAP Age 0.02 0.17 0.85 Bodyweight 0.01 0.13 0.88 Prostate Volume 0.01 0.09 0.92 VCL Age 0.10 0.98 0.39 Bodyweight 0.06 0.57 0.58 Prostate Volume 0.04 0.39 0.68 VSL Age 0.02 0.14 0.87 Bodyweight 0.01 0.05 0.95 Prostate Volume 0.00 0.01 0.99 STR Age 0.25 2.88 0.08 Bodyweight 0.12 1.26 0.31 Prostate Volume 0.02 0.15 0.85 LIN Age 0.27 3.15 0.07 Bodyweight 0.13 1.25 0.31 Prostate Volume 0.05 0.46 0.64 WOB Age 0.25 2.81 0.09 Bodyweight 0.14 1.41 0.27 Prostate Volume 0.11 1.08 0.36

172 Table 3.7 Univariable analyses of sperm morphology parameters and PRL with coefficients of determination (R2) of the overall model, test-statistic and associated p-values.

Morphology R2-value F-Statistic P-value Parameter (%) Normal Sperm 0.00 0.00 0.99 Head Defects 0.00 0.01 0.93 Midpiece Defects 0.11 2.32 0.14 Tail Defects 0.07 1.36 0.26 Loose Heads 0.09 1.78 0.20 Proximal Droplets 0.23 5.50 0.03* Distal Droplets 0.005 0.08 0.78 * Indicates significant p-YDOXH S”

173 Table 3.8 Three bivariable linear regressions of morphology parameters, coefficients of determination (R2) of the overall model, test statistics and associated p-values

Response (%) Explanatory R2-value F-Statistic p-value Variables with PRL in the model Normal Sperm Age 0.00 0.01 0.99 Bodyweight 0.00 0.00 1.00 Prostate Volume 0.03 0.27 0.77

Head Defects Age 0.01 0.11 0.90 Bodyweight 0.00 0.01 0.99 Prostate Volume 0.03 0.31 0.74

Loose Heads Age 0.17 1.77 0.20 Bodyweight 0.10 0.98 0.40 Prostate Volume 0.12 1.14 0.34

Tail Defects Age 0.12 1.12 0.32 Bodyweight 0.07 0.65 0.54 Prostate Volume 0.10 0.96 0.40

Distal Droplets Age 0.24 2.69 0.10 Bodyweight 0.03 0.25 0.78 Prostate Volume 0.19 1.96 0.17

Midpiece Defects Age 0.42 3.89 0.03* Bodyweight 0.13 1.24 0.31 Prostate Volume 0.12 1.11 0.35 * Indicates significant p-value S”

174 Table 3.9 Three non-significant bivariable linear regressions of proximal droplets with variable coefficients where PRL slope coefficients were significant, coefficients of determination for the overall model (R2) and associated p-values.

Explanatory Variable Model Fit Model of Explanatory Regression p- R2- F- p- Proximal Variable Coefficient value value statistic value Droplets for Overall Model Model 1 PRL -0.49 0.08 Age 0.28 0.23 0.30 3.61 0.05 Model 2 PRL -0.55 0.05 Bodyweight -0.03 0.31 0.28 3.30 0.06 Model 3 PRL -0.61 0.04 Prostate -0.09 0.88 0.24 2.61 0.10 Volume

175

Fig 3.1 Predicted percentage of proximal droplets in response to serum PRL. Back-transformation of the linear regression model logitProxDrop=-2.94- 0.60lnPRL (R2=0.23, p=0.03). Predicted Proximal Droplets (%), Lower CI, Upper CI. CI= 95% Confidence Interval.

176

Fig 3.2 Predicted percentage of midpiece defects in response to serum PRL. Back-transformation of the linear regression model logitMidpiece= -8.24+1.93A-0.41lnPRL- 0.15A2 (R2=0.42, p=0.03). Predicted Midpiece Defects (%), Lower CI, Upper CI. CI= 95% Confidence Interval, A=Age

177 References

[1] A. Bachelot and N. Binart, "Reproductive role of prolactin," Reproduction, vol. 133, pp. 361-9, Feb 2007. [2] C. Gobello, et al., "A review of canine pseudocyesis," Reprod Domest Anim, vol. 36, pp. 283-8, Dec 2001. [3] C. Bole-Feysot, et al., "Prolactin (PRL) and its receptor: actions, signal transduction pathways and phenotypes observed in PRL receptor knockout mice," Endocr Rev, vol. 19, pp. 225-68, Jun 1998. [4] G. Merino, et al., "Hyperprolactinemia in men with asthenozoospermia, oligozoospermia, or azoospermia," Arch Androl, vol. 38, pp. 201-6, May- Jun 1997. [5] A. Colao, et al., "Outcome of cabergoline treatment in men with prolactinoma: effects of a 24-month treatment on prolactin levels, tumor mass, recovery of pituitary function, and semen analysis," J Clin Endocrinol Metab, vol. 89, pp. 1704-11, Apr 2004. [6] M. K. Gill-Sharma, "Prolactin and male fertility: the long and short feedback regulation," Int J Endocrinol, vol. 2009, p. 687259, 2009. [7] E. Reiter, et al., "Effects of pituitary hormones on the prostate," Prostate, vol. 38, pp. 159-65, Feb 1 1999. [8] K. E. Lane, et al., "Suppression of testosterone and estradiol-17beta- induced dysplasia in the dorsolateral prostate of Noble rats by bromocriptine," Carcinogenesis, vol. 18, pp. 1505-10, Aug 1997. [9] L. Tangbanluekal and C. L. Robinette, "Prolactin mediates estradiol- induced inflammation in the lateral prostate of Wistar rats," Endocrinology, vol. 132, pp. 2407-16, Jun 1993. [10] S. Chevalier, et al., "Nonsteroidal serum factors involved in the regulation of the proliferation of canine prostatic epithelial cells in culture," Prostate, vol. 5, pp. 503-12, 1984. [11] M. F. El Etreby and A. T. Mahrous, "Immunocytochemical technique for detection of prolactin (PRL) and growth hormone (GH) in hyperplastic and neoplastic lesions of dog prostate and mammary gland," Histochemistry, vol. 64, pp. 279-86, 1979. [12] D. Helmerich and J. E. Altwein, "Effect of prolactin and the anit-prolactin bromocriptin on the testosterone uptake and metabolism in androgen- sensitive and insensitive canine organs," Urol Res, vol. 4, pp. 101-5, Nov 10 1976. [13] A. Shafik, "Prolactin injection, a new contraceptive method: experimental study," Contraception, vol. 50, pp. 191-9, Aug 1994. [14] M. Laszczynska, et al., "Evaluation of spermatozoa of the rat in hyperprolactinaemia induced by metoclopramide," Andrologia, vol. 24, pp. 101-8, Mar-Apr 1992. [15] M. B. Koivisto, et al., "Effects of short-term hyper- and hypoprolactinaemia on hormones of the pituitary, gonad and -thyroid axis and on semen

178 quality in male Beagles," Reprod Domest Anim, vol. 44 Suppl 2, pp. 320-5, Jul 2009. [16] Y. Corrada, et al., "Prolactin reference range and pulsatility in male dogs," Theriogenology, vol. 66, pp. 1599-602, Oct 2006. [17] S. Schafer-Somi and C. Aurich, "Use of a new computer-assisted sperm analyzer for the assessment of motility and viability of dog spermatozoa and evaluation of four different semen extenders for predilution," Anim Reprod Sci, vol. 102, pp. 1-13, Nov 2007. [18] F. Grubbs, "Procedures for Detecting Observations in Samples," Technometrics, vol. 11, pp. 1-21, 1969. [19] A. Ouhtit, et al., "Visualization of gene expression of short and long forms of prolactin receptor in rat reproductive tissues," Biol Reprod, vol. 49, pp. 528-36, Sep 1993. [20] W. M. Hair, et al., "Prolactin receptor expression in human testis and accessory tissues: localization and function," Mol Hum Reprod, vol. 8, pp. 606-11, Jul 2002. [21] E. Hondo, et al., "Prolactin receptor expression in rat spermatogenic cells," Biol Reprod, vol. 52, pp. 1284-90, Jun 1995. [22] H. N. Jabbour and G. A. Lincoln, "Prolactin receptor expression in the testis of the ram: localisation, functional activation and the influence of gonadotrophins," Mol Cell Endocrinol, vol. 148, pp. 151-61, Feb 25 1999. [23] P. Guillaumot, et al., "Sertoli cells as potential targets of prolactin action in the testis," Mol Cell Endocrinol, vol. 122, pp. 199-206, Sep 18 1996. [24] K. Purvis, et al., "Prolactin and Leydig cell responsiveness to LH/hCG in the rat," Arch Androl, vol. 3, pp. 219-30, Nov 1979. [25] R. W. Steger, et al., "Neuroendocrine and reproductive functions in male mice with targeted disruption of the prolactin gene," Endocrinology, vol. 139, pp. 3691-5, Sep 1998. [26] A. R. Hoffman, et al., "Patient guide to hyperprolactinemia diagnosis and treatment," J Clin Endocrinol Metab, vol. 96, pp. 35A-6A, Feb 2011. [27] G. Corona, et al., "Hypoprolactinemia: a new clinical syndrome in patients with sexual dysfunction," J Sex Med, vol. 6, pp. 1457-66, May 2009. [28] C. Urhausen, et al., "Concentrations of prolactin, LH, testosterone, TSH and thyroxine in normospermic dogs of different breeds," Reprod Domest Anim, vol. 44 Suppl 2, pp. 279-82, Jul 2009. [29] M. De Rosa, et al., "The treatment with cabergoline for 24 month normalizes the quality of seminal fluid in hyperprolactinaemic males," Clin Endocrinol (Oxf), vol. 64, pp. 307-13, Mar 2006. [30] M. De Rosa, et al., "Six months of treatment with cabergoline restores sexual potency in hyperprolactinemic males: an open longitudinal study monitoring nocturnal penile tumescence," J Clin Endocrinol Metab, vol. 89, pp. 621-5, Feb 2004.

179 CHAPTER FOUR

SUMMARY AND CONCLUSIONS

This thesis investigated possible associations between serum concentrations of Vitamin D and prolactin (PRL) and reproductive characteristics related to prostate volume, sperm morphology and sperm motility in male dogs.

The first part of this research determined whether an association existed between serum 25OHVD and BPH, as well as between 25OHVD and prostate volume controlling analytically for BPH. Possible associations between serum

25OHVD and a) semen motility as evaluated by CASA and b) microscopically determined sperm morphology were also studied. The second part of the research investigated the same reproductive traits for associations with serum

PRL concentrations.

In both the Vitamin D and PRL studies, we were not able to show any associations between Vitamin D or PRL with BPH, nor were there associations between Vitamin D or PRL and prostate volume controlling for BPH analytically.

The low power of the study due to the small sample size may not have permitted detection of small differences. The diagnosis of BPH was determined based on transrectal palpation per digitum, ultrasound dimensions of the prostate, and/or blood present in the third fraction of the ejaculate. However, a lack of histological diagnosis of BPH as a gold standard may have led to a misdiagnosis of BPH, thus introducing a misclassification bias. One must also consider that prostate physiology may differ from that in men, although experimental Vitamin D

180 analogues decreased prostate size in a small number of beagle dogs in a preliminary drug trial[1]. Increasing the number of dogs studied would allow for more accurate findings and possibly result in a different outcome with regards to

BPH and prostate health.

Vitamin D was shown to have associations with several measures of semen motility and morphology. We determined that serum 25OHVD concentrations for optimal semen quality characteristics ranged from 120-180 nmol/l using a validated and commercially available ELISA for dogs. A negative effect on semen quality could be observed below or above these concentrations, although this effect was lessened at concentrations at the high range of normal and was associated with smaller decreases in semen quality. In the human and rat, Vitamin D and its receptor have been identified in sperm cells in all stages of development and testicular tissue, chiefly in Sertoli cells [2-4]. It is possible that

Vitamin D is necessary for normal and healthy development of sperm in all species, including dogs, and is translated into improvements in normal morphology and consequently, normal and improved motility at optimal concentrations.

Prostate volume is correlated with age and both age and prostate volume were factors that influenced Vitamin D in its semen quality associations. It is possible that Vitamin D exerts a toxic effect at older ages or that older animals are unable to use Vitamin D in the same way as younger animals. One rat study showed a decrease in intestinal Vitamin D action in older compared with younger rats and may suggest impaired Vitamin D metabolism in older animals [5].

181 Perhaps this is also true for the ageing dog with respect to Vitamin D and semen quality. Isolation and identification of Vitamin D and its receptor in the reproductive tissues and sperm cells of the dog at various ages may shed some more light on the roles and mechanisms of Vitamin D, and may provide explanations for the age related effects.

The second half of this thesis looked at serum PRL concentrations and semen quality characteristics. Using the same validated ELISA our serum PRL concentrations ranges were greater than previously published (non-detectable to

28.24 ng/ml compared with non-detectable to 6.0 ng/ml [6]). Prolactin concentrations were associated with sperm midpiece defects as well as proximal droplets, an observation not previously published. Fewer midpiece defects were found when serum PRL concentrations were greater than 2.5 ng/ml. This new information suggests that PRL concentrations less than 2.5 ng/ml may be defined as hypoprolactinemia due to the increase in midpiece defects and proximal droplets at these levels. Evidence of hyperprolactinemia was not observed in this study.

It is important to note that PRL does undergo periodic fluctuation in physiological concentrations and that sampling in this study was undertaken at different times of the day and during different times of the year. The lack of associations with the majority of semen parameters may be a result of small sample size or due to the restriction of having only one PRL measurement per subject. Taking multiple measurements and controlling for the time of year of

182 sampling may impact serum PRL concentrations, and consequently, semen quality.

Prolactin has been associated with semen pathologies in cases of chronic hyperprolactinemia in men through long-term negative feedback loops that cause decreases in circulating testosterone concentrations. Commercial validated tests for PRL in the dog have only recently become available to the veterinary community. Although, hyperprolactinemia has not been previously diagnosed in dogs it is still possible this condition may occur and have an impact on fertility in the male dog. Further research is needed to determine whether situations of

PRL excess occur naturally in the dog and what role that might play in male dog infertility.

In conclusion, this study has revealed new associations with respect to semen quality characteristics and Vitamin D and PRL in the stud dog. This is an exciting avenue of research that promises to have important clinical ramifications in the treatment of canine fertility issues related to semen morphology and motility characteristics. We hope that these preliminary studies help to further supplement the arsenal of tools available to optimize reproductive health in the dog.

183 References:

[1] L. Adorini, et al., "Vitamin D receptor agonists target static, dynamic, and inflammatory components of benign prostatic hyperplasia," Ann N Y Acad Sci, vol. 1193, pp. 146-52, Apr 2010. [2] S. Aquila, et al., "Human sperm anatomy: ultrastructural localization of 1alpha,25-dihydroxyvitamin D receptor and its possible role in the human male gamete," J Anat, vol. 213, pp. 555-64, Nov 2008. [3] J. Merke, et al., "Nuclear testicular 1,25-dihydroxyvitamin D3 receptors in Sertoli cells and seminiferous tubules of adult rodents," Biochem Biophys Res Commun, vol. 127, pp. 303-9, Feb 28 1985. [4] M. Blomberg Jensen, et al., "Vitamin D receptor and vitamin D metabolizing enzymes are expressed in the human male reproductive tract," Hum Reprod, vol. 25, pp. 1303-11, May 2010. [5] G. Balogh, et al., "Influence of age on 1,25(OH)2-vitamin D3 activation of protein kinase C in rat duodenum," Mol Cell Endocrinol, vol. 129, pp. 127- 33, May 16 1997. [6] Y. Corrada, et al., "Prolactin reference range and pulsatility in male dogs," Theriogenology, vol. 66, pp. 1599-602, Oct 2006.

184

APPENDICES

185 Appendix 1. Investigation of serum vitamin D and prolactin levels in benign prostatic hyperplasia and semen quality in intact male dogs. Client Consent Form.

“Investigation of serum vitamin D and prolactin levels in benign prostatic hyperplasia and semen quality in intact male dogs”

CLIENT CONSENT FORM

Benign Prostatic Hyperplasia (BPH) is a non-cancerous enlargement of the prostate, associated with poor semen quality, and affects approximately 80% of intact male dogs. Recent research into prostate disease in men has shown that vitamin D is associated with a decrease in the occurrence of this condition and better overall prostate health. Prolactin, a reproductive hormone most commonly associated with females during pregnancy and lactation, also has a role in prostate health. In increase in serum prolactin levels in men has been shown to negatively affect fertility and potentiate hyperplasia of the prostate. Interestingly, the prostate gland is similar in both human and canine species. Our objectives are to determine: 1) whether an association exists between BPH and circulating vitamin D and prolactin levels and 2) to provide information to the canine breeding industry regarding the possible effects of vitamin D and prolactin on canine male fertility and prostate health. We request that you complete a brief questionnaire to obtain background information on breeding history and management. Semen collection will be undertaken, after which, both a physical and ultrasound examination will be performed. A blood and urine sample will also be taken for research purposes. The procedures being used in this study are considered neither painful nor invasive. The costs for the procedures and testing will be assumed by the project. All results will be kept in strictest confidence.

We appreciate your time and consideration.

Dr. Cathy Gartley, Dr. Adria Kukk

I agree to participate in the study. Ƒ

Patient Name______Patient ID______

Signature of Owner/Authorized Agent______Date______

Signature of Clinician ______Date______

186 Appendix 2. Investigation of serum vitamin D and prolactin levels in benign prostatic hyperplasia and semen quality in intact male dogs. Client Questionnaire.

Questionnaire

The following questions address necessary background information for our study on the role of vitamin D and prolactin on benign prostatic hyperplasia and fertility in intact male dogs. Please answer all of the questions to the best of your ability.

1. Client name: ______

2. Dog’s name, breed, and age: ______

Part 1: Management

3. What sort of diet is your dog on? Please state the brand name if using a commercial diet and any regular treats or ‘goodies’ they receive daily. Please include the approximate amounts.

i) ______

ii) ______

iii) ______

4. How is your dog housed? For example, does he live in a kennel situation with outdoor runs, does he live indoors with the family, or a combination of both?

______

______

5. How many hours a day, approximately, does your dog spend out-doors? For example leash walks, in the backyard, in outdoor runs etc.

In the summer: ______

In the winter: ______

Page 1 of 2

187

6. Is your dog on any supplements and, if so, what are they?

i) ______

ii) ______

iii) ______

Part 2: Medical history

7. History of past illness (Type/year/duration)

i) ______

ii) ______

iii) ______

8. Is your dog on any medication(s) and, if so, what are they?

i)______

ii)______

Part 3: Breeding History

9. How many litters has you dog sired?

Per year: ______

Per career: ______

10. Average size of litter? ______

11. Approximately how many breedings/year? Please include fresh chilled/frozen semen inseminations if applicable. ______

12. How many years has he been at stud in total? ______

Page 2 of 2

188 Appendix 3 Minitube Chill 5 extender Protocol (Excerpt).

Protocol for CaniPRO Chill 5 Culture Medium for Canine Semen (13574/0105)

Extender preparation: Warm CaniPRO Chill 5 to room temperature.

Semen extension: x Use only the sperm-rich fraction of the ejaculate (2nd fraction) for insemination. Do not include the first fraction of the ejaculate (clear prostatic fraction before sperm rich fraction) or the third fraction (prostatic fraction post-sperm rich fraction) of the ejaculate in the sample because the quality of the semen may decrease. x After collection evaluate semen quality (Concentration, Motility, Morphology). Dilute one part of semen in three to five parts of extenders. (ie: 1ml in 3-5ml of x CaniPRO Chill 5) base on initial semen concentration. x Semen can be preserved for up to 5 days at 4°C (39°F) with preservation of a minimum of 70% initial motility. Avoid changes in temperature during preservation by placing the tube with the extended semen in small water- bath. x At time of insemination warm semen to room temperature and perform a motility evaluation (See insemination instructions below).

For purpose of shipping chilled semen x Store diluted semen at 4°C (39°F) for at least 2 hours before packaging and shipping. x Perform a motility evaluation before shipping and retain a control aliquot for future evaluation. x If needed to reach ideal insemination volume (See insemination instructions below) add some extender in extra 15ml tube in Canine Transport Box.

Canine Semen Concentration (106) Ratio of Semen to Extender Per ML

250 – 750 1 to 3

750 – 1.25 1 to 4

Above 1.25 1 to 5

189 Appendix 4 SFT Guidelines (Excerpt)

GUIDELINES FOR CANINE BREEDING SOUNDNESS EXAMINATION

Original authors: Original reviewers: B.J. Purswell, Virginia Tech P.N.S. Olson, University of Minnesota G.C. Althouse, Iowa State University S.D. Johnston, University of Minnesota M.V. Root, University of Minnesota L.E. Evans, Iowa State University

Current reviewer: M.V. Root Kustritz, University of Minnesota

Other changes: G.C. Althouse is now at University of Pennsylvania, P.N.S. Olson is with the Morris Animal Foundation, and S.D. Johnston is at Western College of Health Sciences.

Morphology

Morphology most often is assessed by staining the semen sample and observing the cells under 1000X magnification (oil immersion). Morphology may be assessed without staining using a phase contrast microscope after fixing the sample with formol-buffered saline at a dilution of 1:9. Different sample preparations cause artifacts that will be seen as morphologic abnormalities; percentage morphologically normal spermatozoa should be fairly consistent regardless of method [1]. The two most common stains used are a rapid Wright’s Giemsa stain (DiffQuik, Baxter Healthcare, Miami FL) and eosin/nigrosin stain (SFT stain, Lane Manufacturing, Denver CO).

To stain with rapid Wright’s Giemsa stain, place one drop of semen on a slide and smear out as for a blood smear. Allow to air dry. Immerse the slide into each of the three solutions in the same order as used for any type of cytology, allowing the slide to sit in each solution for five minutes. Rinse the slide completely and let it air dry before evaluation under oil immersion.

To stain with eosin/nigrosin, place one drop of semen and a similar-sized drop of stain on one end of a glass slide. Gently mix the two together with a pusher slide, and draw out as a thin film similar to a blood smear. Allow to air dry as quickly as possible; placing the slide on a hot plate (37oC) and blowing on the slide will hasten the drying process and help prevent staining artifacts [2].

Spermatozoa stained with rapid Wright’s Giemsa stain appear purple on a clear background. Normal spermatozoa stained with eosin/nigrosin stain appear white against a black or violet background. Eosin/nigrosin stain is taken up by spermatozoa that have abnormal plasma membranes and so appear pink against a dark background. It might be assumed that such spermatozoa are non-viable

190 but no significant correlation between “live-dead” ratio and fertility has been described in dogs.

Under oil immersion, examine and count at least 100 spermatozoa. The number of normal spermatozoa in 100 is the percentage morphologically normal spermatozoa. Total normal for the sample is calculated by multiplying total number of spermatozoa in the ejaculate by percentage morphologically normal. Commonly accepted values are greater than 80% morphologically normal spermatozoa and greater than 200 million total normal spermatozoa in an ejaculate. Abnormal spermatozoa may be classified as having primary defects (those that occur during spermatogenesis, including defects in head shape, bent midpiece, persistent proximal cytoplasmic droplet, and doubling of any portion of the spermatozoon) or secondary defects (those that occur during epididymal maturation or slide preparation, including detached heads, persistent distal cytoplasmic droplets, and bent tails). Correlation between specific defects and fertility in dogs is poorly defined.

References

[1] Root Kustritz MV, Johnston SD, Olson PN and Root TK: The effect of stains and investigators on assessment of morphology of canine spermatozoa. J Amer Anim Hosp Assoc 1998;34:348-352.

[2] Shaffer HE, Almquist JO. Vital staining of bovine spermatozoa with a eosin- aniline blue staining mixture. J Dairy Sci 1948;3:677-678.

191 Appendix 5. Outlier Removal Statistics

Predicted sperm motility in response to normal serum 25OHVD range at various transformed prostate volumes with removal of dog 6. Back-transformation of multiple regression model logitMotility=-11.7 +2.73Vol +0.11VD-0.02Vol*VD (R2=0.56, p=0.003). CI= 95% Confidence Interval. Vol=Prostate Volume. Predicted Motility (%), Lower CI, Upper CI

192 Table. Outlier removal of dog 6. Explanatory variable with vitamin D (VD) in the model and corresponding coefficient of determination (R2) and p-value. Resulting analysis revealed no significant associations between serum 25OHVD and progressive motility.

Response Explanatory Variable with R2-value p-value of Linear Vitamin D in the model Model

Progressive VD2 0.18 0.21 Motility

193

Outlier removal of dog 6, 1 and 15. Predicted normal sperm and serum PRL. Back transformation of linear regression model logitNormal=0.26 + 0.004lnPRL (R2=0.36, p=0.01). Predicted Normal Sperm (%), Lower CI, Upper CI. CI= 95% confidence interval.

194

Predicted normal sperm in response to serum PRL range at various transformed prostate volumes with removal of dogs 6, 1 and 15. Back-transformation of multiple regression model logitNormal=3.32 -0.52Vol + 0.32lnPRL (R2=0.52, p=0.006). CI= 95% Confidence Interval. Vol=Transformed Prostate Volume. Predicted Normal Sperm (%), Lower CI, Upper CI.

195

Predicted normal sperm in response to normal serum PRL range at various ages with removal of dogs 6, 1 and 15. Back- transformation of multiple regression model logitNormal=2.66 + 0.25lnPRL - 0.25Age (R2=0.66, p=0.0005). CI=Confidence Interval. PRL=prolactin. Predicted Normal Sperm (%), Lower CI, Upper CI

196