The Formation of Androstenone Conjugates in Testis Tissue of the Mature Boar

THE FORMATION OF ANDROSTENONE CONJUGATES IN TESTIS TISSUE OF THE MATURE BOAR

by JILLIAN EVE DESNOYER

A Thesis Presented to the University of Guelph

In partial fulfillment of requirements for the degree of Master of Science in Animal and Poultry Science (Toxicology)

Guelph, Ontario, Canada

ABSTRACT

THE FORMATION OF ANDROSTENONE CONJUGATES IN TESTIS TISSUE OF THE

MATURE BOAR.

Jillian Eve Desnoyer Advisor: University of Guelph, 2011 Dr. E.J. Squires

The accumulation of androstenone in the fat of mature boars results in boar taint; the conjugation of androstenone would decrease this important meat quality problem by decreasing the accumulation and increasing the excretion of androstenone. Leydig cells and testis microsomes from mature boars were incubated with radiolabeled pregnenolone, and the free and conjugated metabolites were examined by HPLC. Sulfated androstenone with a mass of 367 m/z was directly identified by MS, with a novel tentative structure of 3-keto-4- sulfoxy-androstenone. Addition of enolase to the microsomal incubations increased the formation of 3-keto-4-sulfoxy-androstenone. Overexpression of SULT2A1 in HEK cells resulted in the sulfoconjugation of dehydroepiandrosterone, but not androstenone, suggesting that SULT2A1 may not be involved in sulfoconjugation of androstenone. This thesis describes the novel direct characterization of androstenone sulfate and the importance of enolase in its formation. The relevance to boar taint metabolism is discussed.

ACKNOWLEDGEMENTS

To Dr. Jim Squires: Thank you for your boundless breadth of knowledge, understanding and encouragement throughout the completion of this thesis. Your patience and willingness to deal with all of my self-proclaimed “stupid questions” was unparalleled to any other during my education. Delving feet first into this project would have been far more intimidating if it weren’t for you. I appreciate it more than I can possibly say. To my advisory committee Dr. Gregoy Bedecarrats and Dr. Jim Raeside – thank you for your continued support and help throughout the process of researching and writing this thesis. Without the knowledge that you were there to help, I wouldn’t have been able to proceed with the utmost confidence. Dr. Raeside, your incredible talents at pouring Percoll gradients are, well, incredible as is your amazing knowledge of everything related to reproductive endocrinology.

To Yanping Lou: you are the technical superhero of our lab. Your ability to fix the HPLC when all seemed lost – on a regular basis – is incredible. Without your teaching, technical support and constantly smiling face, our lab wouldn’t function and wouldn’t be nearly as great. To Heather Christie: thank you for getting up early with me to isolate those pesky Leydig cells and for being THE woman to see for all things related to steroidal conjugates. I loved our endless talks and I look forward to continuing them for years to come. To Dr. Jim Atkinson: thank you for your teasing and reassurance throughout the process of both my undergraduate degree and this thesis. I cannot possibly say how much our chats both meant to and helped me.

To my lab mates both past and present: Matt, thank you for putting up with my endless questions related to transfection and cloning and for all of my random trivia tidbits that interrupted you daily. I’m sure you’ll miss me when the lab is completely quiet. Kim, thank you for everything. I missed your presence greatly this last year and, when you’re all grown up and back from Indiana, I look forward to pestering you to play Munchkin.

To Andy, thank you. I would never have made it through this without you. Thank you for sitting through constant repeats of the same presentations, listening to me wax poetic about androstenone sulfate and for getting excited with me when I found something – anything – exciting. Thank you for taking care of our little furry animals while I delved headfirst into the most trying educational triumph of my life - thus far. Also, to Ella, Flint and Bailey: ruff ruff meow meow ruff barrooo. This translates roughly to “thank you for the constant cuddles and love”.

To my family and friends – without your support and encouragement, this wouldn’t have been possible. Mom, your biweekly phone calls and feigned excitement over a project you barely understood was awesome. I can’t believe I finally did it and I know I couldn’t do it without you. Dad, thank you for believing in me, even when it wasn’t said aloud, I know you knew I could do it. Thank you to my friends who sat through my presentations, giving helpful and great advice.

Not only has this project taught me more about endocrinology and molecular biology than I could ever have imagined, it has taught me more than that. It has taught me that my love of research is boundless and that, more importantly, I can do it. It is through the support, patience and encouragement of the above people that I can truly say I am a Mistress of Science.

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TABLE OF CONTENTS

THE FORMATION OF ANDROSTENONE CONJUGATES IN TESTIS TISSUE FROM THE MATURE BOAR...... i LIST OF FIGURES...... vii LIST OF TABLES...... x LIST OF ABBREVIATIONS...... xii CHAPTER I: REVIEW OF THE LITERATURE...... 1 1.1 Boar Taint and Pig Production...... 1 1.1.1 Introduction...... 1 1.1.2 Current Solutions...... 3 1.1.2.1 Carcass Measurements/On-Line Testing...... 3 1.1.2.2 Semen Sexing...... 4 1.1.2.3 Immunocastration...... 5 1.1.2.4 Other GnRH Agonists ...... 7 1.2. Steroidogenesis, 16-Androstene Steroids and Steroid Circulation...... 9 1.2.1 Location of Steroidogenesis in the Boar...... 9 1.2.2 Steroidogenesis Throughout the Lifetime of the Pig ...... 10 1.2.3 Enterohepatic Circulation of Steroids ...... 12 1.2.4 Binding of Steroid Hormones in the Blood and Saliva ...... 14 1.2.4.1 Steroid Hormone Movement in the Blood ...... 14 1.2.4.2 Steroid Binding in Boar Saliva...... 16 1.2.5 Steroids and Target Tissues Including Adipose Tissue ...... 17 1.3 Androstenone Metabolism ...... 19 1.3.1 Androstenone Synthesis...... 19 1.3.2 Androstenone Degradation and Excretion...... 22 1.3.3 Androstenone Sulfate ...... 23 1.4 Enzymes Involved in Conjugation...... 25 1.4.1 Sulfotransferases...... 25 1.4.1.1 Sulfotransferase 2A1 ...... 27 1.4.1.2 Sulfotransferase 2B1 ...... 28 1.4.2 Steroid Sulfatase ...... 29

1.4.2 Enolases...... 30 1.4.3 UDP-glucuronosyl transferases...... 32 1.5 Rationale for Experiment...... 34 CHAPTER II: HYPOTHESIS AND OBJECTIVES...... 36 2.1. Hypothesis and Research Objectives ...... 36 2.1.2 Hypothesis...... 36 2.2.1 Objective One: Discovery of Androstenone Sulfate in Leydig Cells...... 36 2.2.2 Objective Two: Characterisation of the Role of Enolase in Androstenone Sulfate Formation...... 36 2.2.3 Objective Three: Identification of Sulfotransferase Involved ...... 37 CHAPTER III: IDENTIFICATION OF ANDROSTENONE SULFATE...... 38 3.1 Introduction...... 38 3.2 Methods ...... 42 3.2.1 Animals ...... 42 3.2.2 Leydig Cell Isolation ...... 42 3.2.3 Leydig Cell Incubation...... 43 3.2.4 Steroid Conjugate Separation...... 44 3.2.5 Initial HPLC Steroid Analysis...... 45 3.2.6 Conjugate Peak Analysis and Identification...... 46 3.2.7 Mass Spectrometry ...... 47 3.2.8 Statistical Analysis ...... 48 3.3 Results ...... 48 3.3.1 Conjugation Capacity of Leydig Cells...... 48 3.3.2 Analysis of Conjugated Fraction ...... 51 3.3.6 Conjugation Over Time: 8 Hour vs 24 Hour Incubations...... 54 3.3.3 Discovery and Identification of Androstenone Sulfate ...... 55 3.3.4 Androstenone Sulfate Identification...... 56 3.3.5 Androstenone Sulfate vs Androstenone Production in Incubated Leydig Cells ...... 60 3.3.6 Conjugated Steroid Production ...... 60 3.3.7 Free Steroid Profiles for Individuals...... 61 3.4 Discussion...... 62 CHAPTER IV: CHARACTERIZATION OF THE ROLE OF ENOLASE IN ANDROSTENONE CONJUGATION...... 72 v

4.1 Introduction...... 72 4.2 Methods ...... 73 4.2.1 Chemicals and Materials...... 73 4.2.2 Testes Microsome Preparation ...... 73 4.2.3 Enolase and Pregnenolone Microsomal Incubations...... 74 4.2.4 Steroid Extraction from Microsomal Incubations...... 75 4.2.5 Sep-Pak Chromatography and HPLC Analysis...... 75 4.3 Results ...... 76 4.3.1 Optimum Pregnenolone Incubation Time ...... 76 4.3.2 Conjugate Fractions from Microsome Incubations...... 78 4.3.2 Free Fractions ...... 83 4.4 Discussion...... 86 CHAPTER V: CHARACTERIZATION OF THE ROLE OF SULT2A1 IN ANDROSTENONE CONJUGATION...... 89 5.1 Introduction...... 89 5.2 Materials and Methods...... 92 5.2.1 Chemicals Used...... 92 5.2.2 Cloning of Sulfotransferase...... 92 5.2.3 Sulfotransferase Expression in HEK Cells...... 93 5.2.4 Cell Lysate Assays...... 94 5.2.5 HPLC and Mass Spectrometry of Steroids ...... 95 5.2.5 HPLC and Mass Spectrometry of Steroids ...... 95 5.3 Results ...... 96 5.2.1 Conjugate Fraction Analysis ...... 96 5.2.2 Mass Spectrometry ...... 98 5.2.3 Free Fraction Analysis...... 99 5.4 Discussion...... 100 CHAPTER VI: FUTURE CONSIDERATIONS AND THESIS CONLUCIONS ...... 102 6.1 Discussions and Future Considerations...... 102 6.2 Thesis Conclusions...... 111 APPENDIX...... 113 References...... 116

LIST OF FIGURES

Figure 1.1: The steroidogenic pathway resulting in the formation of androstenone. A. cytochrome P450 side chain cleavage; b. Andien-β synthetase; c. Δ4,5 isomerase/3β-dehydrogenase; d. 5α-reductase; e. 3β-hydroxysteroid dehydrogenase; f. 3α-hydroxysteroid dehydrogenase ...... 21 Figure 1.2: The sulfation pathway for the sulfation of androstenol. SULT: sulfotransferase. PAP: 3’-phosphodenosine-5’-phosphate...... 24 Figure 1.3: The proposed mechanism for the formation of androstenone sulfate, by the enolation of its’ ketone group. ENOL: enolase; SULT: sulfotransferase...... 24 Figure 3.1: Relative radioactive content of samples following separation of conjugate and free fractions by Sep-Pak Chromatography from 8 hour Leydig cell incubations. The remainder minor amount of the percentage of radioactivity of samples was accounted for in the sample and wash fraction (data not shown)...... 51

Figure 3.2a: Example of HPLC Protocol A output for conjugate fraction. This graph shows the large peak at 3.5, 7 and 15 minutes. The earliest peak encompasses the coelution of the aforementioned early peaks corresponding to estrone glucuronide, E1S, DHEA glucuronide and DHEAS...... 53

Figure 3.2b: The same graph as Figure 2a but showing the smaller peaks, located at 7, 8, 15 and 21 minutes. In this individual, the peak at 7 minutes corresponded to DHEA glucuronide elution whereas the peak at 8 minutes corresponded to DHEAS elution. The small peak at 21 minutes corresponds to the expected elution of androstenone sulfate...... 54 Figure 3.3: Example graph of solvolyzed peak at 21 minutes, showing a glucuronide conjugate at approximately 3.5 minutes and free androstenone at 21 minutes...... 56 Figure 3.4: The formation of 3-keto-4-sulfoxy-androstenone from androstenone. Enolase (A.) converts the 3-keto group found in androstenone to a 3-enol intermediate. Sulfotransferase (B) attaches the sulfate group to C4, resulting in the double bond between C3 and C4 to convert the C3 oxygen back to a keto group. The 3-sulfoxy-androstenone with a mass of 352 was not found by mass spectrometry...... 58 Figure 3.5: Tandem mass spectrometry output for peak found at 18-21 minutes identified with a mass of 367.1 m/z. This could be an ionized form of the compound found at 368.1597 m/z, which corresponds directly to the predicted mass of 3-keto-4-sulfoxy-androstenone. The peak with a mass of 96.9623 (97

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m/z) corresponds to a sulfate group, whereas the remaining peaks could be due to the degradation of ANS...... 59 Figure 3.6: Scheme 2 adapted from Yi et al. (2006). Removal of sulfate group from carbon, resulting in a C=C bond and a charged sulfate group with a mass of 97 m/z. This is what is theorized to occur at the 4-carbon of androstenone, resulting in the formation of androstenone after the removal of the sulfate group...... 69

Figure 4.1: Unmetabolized radiolabeled pregnenolone as indicated by peak at 6.8 minutes from the free fraction. Peaks found at 3.54 and 4.37 minutes correspond to estrone and DHEA respectively. Peaks found at 14 minutes and 16 minutes correspond to 3β-androstenol and androstenone respectively...... 77 Figure 4.2: Optimum pregnenolone metabolism to 16-androstenes occurred after 5 minute pre-incubation with pregnenolone, 20 minute incubation with NADPH and 20 minute incubation with PAPS and enolase. Peaks found at 13 minutes, 14 minutes and 16 minutes corresponded to 3α-androstenol, 3β-androstenol and androstenone respectively...... 78

Figure 4.3a. Conjugate fraction output (counts per minute (CPM)) from HPLC Protocol A showing a peak at 2 to 4 minutes, indicative of steroid glucuronides and DHEAS and E1S and a peak at 21-22 minutes, indicative of androstenone sulfate. This incubation was from individual 8 and was done without the addition of enolase...... 79 Figure 4.3b. Conjugate fraction output (counts per minute (CPM)) from HPLC Protocol A showing a peak at 2 to 4 minutes, indicative of steroid glucuronides and DHEAS and E1S and a peak at 21-22 minutes, indicative of androstenone sulfate. In this case, the peak at 18 minutes was determined to be the same compound as the compound in the 21-22 minute peak. This incubation was from individual 8 and was done in the presence of enolase. Note the much larger androstenone sulfate peak compared to Figure 4.3a...... 80 Figure 4.3c. Graph of total counts per minute (CPM) for a testis microsome incubation from individual 8 that did not contain enolase. Peaks at 5-6 minutes were found to be unmetabolised pregnenolone whereas the two, smaller peaks were found to be, in this run only, DHEAS and E1S. Very small peaks found at 15 and 16 minutes were 3β-androstenol and 3α-androstenone and the large peak at 21-22 minutes was androstenone sulfate...... 82

Figure 4.4: Androstene production from microsome incubations with radiolabeled pregnenolone with enolase from individual pig 8...... 84

Figure 4.5: Androstene production from microsome incubations with radiolabeled pregnenolone and without enolase from individual pig 8...... 84

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Figure 5.1a: HPLC output from Protocol A (conjugate fraction) for SULT2A1 transfected cells incubated with PAPS and NADPH, following correction for background peaks. The 3 minute peak corresponds to DHEAS...... 97

Figure 5.1b: HPLC output for Protocol A (conjugated) for control transfected cells incubated with PAPS and NADPH following correction for background peaks, showing a very small level of endogenous sulfotransferase activity from the HEK cells...... 97

LIST OF TABLES Table 3.1: Conjugation capacities of Leydig cells incubated with [7-3H(N)]- pregnenolone reported as a mean (n=3), +/- SE...... 50

Table 3.2: Comparison of percentage of total radiolabelled material in the conjugated and free fractions of Leydig cell incubations at two time points of three individuals...... 55 Table 3.3: Comparison table of HPLC output from pig 8, showing no differences in compound elution times or relevant steroidal content between 8 hour and 24 hour incubations. The peak at 4 minutes corresponds to steroid hormones other than our compound of interest, androstenone, and thus the large difference between time points is not relevant. The peak at 7 minutes corresponds to unmetabolized pregnenolone. Although pregnenolone is present in both samples, the size of the peak does not differ widely compared to the other peaks...... 55 Table 3.4: Amount of free and conjugated androstenone as a percentage of total radiolabeled pregnenolone added (%)...... 60 Table 3.5: Amount of conjugated steroid hormones produced from the addition of radiolabeled pregnenolone as indicated by the percentage conversion of pregnenolone (%). E1G = estrone glucuronide, DHEAG= DHEA glucuronide, E1S = estrone sulfate, DHEAS= DHEA sulfate, BAS=3β-androstenol, AAS=3α- androstenol, ANS=androstenone sulfate...... 61 Table 3.6: Concentrations of steroids (in pM) and percentage of total 16-androstene production in peaks attributed to five 16-androstene steroids from 8 hour Leydig cell incubations...... 62 Table 4.2: The amount of conjugated metabolite found in testicular microsomes from 13 individual pigs as determined by percent conversion of radiolabeled pregnenolone (%). E+ percentages are from incubations with enolase and E- percentages are from incubations without enolase. E1S=estrone sulfate, DHS=DHEA sulfate, BAS=3β-androstenol sulfate, AAS=3α-androstenol sulfate and ANS=androstenone sulfate...... 82 Table 4.3: Summary of important pregnenolone metabolites produced by testicular microsomes for four pigs. The amount of each metabolite is depicted as a percentage of the radiolabeled material present for each HPLC injection. The percentage does not add up to 100% due to disregarding background peaks. 3α=3α-androstenol, 3β=3β-androstenol, AN=androstenone, ANE=androstadienone, ANL=androstadienol, E1=estrone, DH=DHEA...... 85

Table 4.3: Comparison between 16-androstene production and estrone and DHEA production in testis microsome incubations (%)...... 86 Table 4.4: Conjugated and free androstenone production. Conjugate androstenone is represented as a percentage of pregnenolone conversion from testis microsome incubations whereas free androstenone is represented as a percentage of total radiolabeled material injected into the HPLC column...... 86 Table 5.1: DHEAS formation in transfected and control HEK cells as calculated as a percentage of DHEA added (%). For the incubations with the additional cofactors, the first column is without subtraction of the blank samples whereas the second column takes into account the blank samples...... 98 Table 5.2: Androstenone concentration following treatments of HEK cell lysates....99

LIST OF ABBREVIATIONS AAS - 3α-androstenol sulfate ACN – Acetonitrile

Androstadienol/ANL – 5,16-androstadien-3β-ol Androstadienone/ANE – androsta-4,16,-dien-3-one Androstenol (3α, 3β) - α/β-androst-16-en-3-ol Androstenone/AN - 5α-androst-16-en-3-one

ANS – Androstenone sulfate

APPI – atmospheric pressure photoionization BAS – 3β-androstenol sulfate

CaCo-2 – Heterogeneous human epithelial colorectal adenocarcinoma cells DHEAG – DHEA glucuronide DHEAS/DHS – dehydroepiandrosterone sulfate DMEM – Dulbecco’s Modified Eagle Medium

DNAse – Deoxyribonuclease E1G – Estrone glucuronide E1S – Estrone sulfate

EDTA - Ethylenediaminetetraacetic acid ELISA – Enzyme-linked immunosorbant assay ENOL – Enolase ESI – electron spray ionization

FBS – Fetal bovine serum FGly – α-formylglycine

GC-MS – Gas chromatography-mass spectrometry

GnRH - Gonadotrophin releasing hormone

GST – glucocorticoid sulfotransferase

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HEK293 – Human embryonic kidney cells 293 HepG2 – Human liver carcinoma cell line

HPLC – High Performance Liquid Chromatography HSD (3α, 3β) – hydroxysteroid dehydrogenase LB – Lysogeny broth LC-MS – Liquid chromatography-mass spectrometry

LH - Lutenizing hormone MALDI – matrix assisted laser desorption ionization MS/MS – Tandem mass spectrometry

NADPH - Nicotinamide adenine dinucleotide phosphate NCBI – National Center for Biotechnology Information OATP – Organic anion-transporting polypeptide PAPS – 3’-phoshoadenosine 5’-phosphosulfate

PBS – Phosphate-Buffered Salines PCR – Polymerase chain reaction

PMSF – phenylmethanesulfonyl fluoride Pregnenolone - 3β-hydroxypregn-5-en-20-one

Progesterone - pregn-4-ene-3,20-dione QToF – Quantitative Time of Flight

SHBG/SBP – Sex-hormone binding globulin

STS – Steroid Sulfatase SULT – Sulfotransferase SULT1A1 – phenol sulfotransferase

SULT1E – estrogen sulfotransferase SULT2 – hydroxysteroid sulfotransferase SULT2A1 – Sulfotransferase 2A1

SULT2B1 – Sulfotransferase 2B1

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Tris-HCl - tris(hydroxymethyl)aminomethane hydroxychloride UDP – Uridine diphosphate

UGT – UDP-glucuronosyl transferase

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CHAPTER I: REVIEW OF THE LITERATURE

1.1 Boar Taint and Pig Production

1.1.1 Introduction

In modern pig production, male piglets are castrated between 2 and 4 days of age. Reasons for performing this procedure include reduced aggression, ease of management and the prevention of boar taint (Thun et al., 2006). However, traditional methods of castration without anaesthetic have been shown to cause acute pain and stress, as evidenced by observable behavioural changes and a measureable increase in pain-associated hormones for 4 days post-castration (Hay et al., 2003). Additionally, Norway banned surgical castration in 2009, encouraging worldwide research into non-surgical alternatives (Fredriksen and Nafstad, 2006).

With the growing trend of welfare-friendly animal agriculture, the elimination of surgical castration in piglets is a worthwhile goal.

The main reason for castration is the reduction of boar taint, a distasteful odour and flavour in the meat and fat of intact male pigs. There are two compounds that have been associated with this off-flavour: androstenone and skatole, both of which accumulate particularly well in fat. These compounds are synthesized throughout the lifetime of pigs but synthesis and accumulation in fat increases dramatically after puberty and can be above threshold levels as early as 110 days of age (Aldal et al., 2005). If androstenone concentrations in the plasma of the boar exceed 15 ng/ml, free androstenone tends to accumulate in the fat more readily (as

reviewed by Andresen, 2006), causing a distinct odour to emanate from cooked pork products.

Skatole on its own may not cause boar taint, but the odour associated with androstenone is intensified when skatole is present (Bonneau, 1982). Interestingly, skatole is mostly associated with the off odour whereas the combination of skatole and androstenone accounts for the off-flavour in sensory panels (Bonneau et al.,

2000). Both skatole and androstenone have been shown to have a high correlation with boar taint levels. However, there are other compounds of relevance such as indole (Babol et al., 1996a) that need to be considered.

There are several breeds of pig used in pig production, including Hampshire,

Landrace, Yorkshire, Duroc, Pietrain and Meishan. The latter breed, Meishan, is generally used in Chinese agriculture and is coveted for its larger litter size.

Unfortunately, Meishan boars also have some of the highest levels of boar taint compared to other breeds used in production. This trait is potentially due to their high levels of luteinizing hormone (LH) and androgens throughout their lifetime as well as their propensity for large, numerous Leydig cells within the testes (Lunstra et al., 1997).

Of the breeds used in Western pig production, Hampshire boars have the lowest boar taint as determined by human sensory panels whereas the Landrace boars have the lowest concentration of skatole and androstenone associated steroids, as measured by utilizing colorimetric testing of carcasses (Xue et al., 1996).

The lower androstenone concentration in Landrace boars may be due to increased

clearance of sex steroids as compared to Duroc boars (Oskam et al., 2010). Duroc boars had the highest measurable levels of 16-androstenes and androgens in saliva and fat (Xue et al., 1996), in addition to achieving sexual maturity more quickly than

Landrace boars (Oskam et al., 2010).

As carcass steroid profiles can vary from breed to breed, it is prudent to examine the impact of genetics on formation of boar taint. Theoretically, if the genes responsible for boar taint are identified in these low or high taint breeds, long-term breeding programs may result in the elimination of taint altogether. However, there may be some shorter term solutions for boar taint that are available to the industry now.

1.1.2 Current Solutions

Current solutions to boar taint include: on-line tests for skatole developed by

Danish industry experts, carcass measurements, semen sexing by flow cytometry, active immunization against GnRH and the use of other GnRH agonists.

1.1.2.1 Carcass Measurements/On-Line Testing

The level of boar taint compounds in a carcass can be predicted and detected at the time of slaughter using a combination of hormone levels and bulbo-urethral gland and testis size (Zamaratskaia et al., 2005). Levels of 16-androstene steroids can be measured from the salivary gland and fat – although levels in the salivary gland have been shown to be more effective at predicting boar taint than levels in fat using colorimetric analysis (Babol et al., 1996a). It has also been suggested that

sorting carcasses based on skatole levels would be the most appropriate post- slaughter method of decreasing boar taint (Bonneau et al., 2000).

Because bulbo-urethral gland and testis weight and length are indicative of sex steroid hormone synthesis, these measurements can be a good indicator of low fat concentrations of androstenone as they identify immature animals (Bonneau and

Russeil, 1985, Xue et al., 1996).

Although these post-slaughter methods seem to be effective at preventing boar taint flavour in meat from getting to the consumer, they can be both costly

(hormone assays, discarded carcasses) and time consuming (physical measurements) with a high degree of variation in the relationship between these measurements and the accumulation of boar taint compounds. Therefore, the use of these methods as the only prevention of boar taint may not be ideal.

1.1.2.2 Semen Sexing

Another method of decreasing boar taint is to eliminate the rearing of males altogether in pig production. Separation of semen based on weight by flow cytometry is an option that has been explored recently (Thun et al., 2006). Sperm containing the X chromosome-only is used to impregnate sows by artificial insemination thus producing only female litters of piglets. Unfortunately, female pigs have decreased feed conversion efficiency (Thun et al., 2006), thereby increasing the cost to the producer. Currently, this method is prohibitively expensive and there is the possibility of decreased sperm viability due to physical damage by flow cytometry.

1.1.2.3 Immunocastration

The principle behind immunocastration is to prevent boar taint by limiting the release of the hormones that initiate sex steroid synthesis. Gonadotrophin

Release Hormone (GnRH) from the hypothalamus is released and travels to the pituitary gland where it initiates the release of Luteinizing Hormone (LH). LH then initiates the production of male sex hormones including androstenone from the

Leydig cells of the testis (Squires, 2003). Injection of a vaccine, for example

Improvac™ from Pfizer, causes the body to create antibodies against GnRH, preventing GnRH from initiating the release of LH, thereby stopping the production of androstenone (Zamaratskaia and Squires, 2008). This lack of stimulation for steroidogenesis results in decreased testicular size, and impaired pituitary gland and Leydig cell function (Awoniyi et al., 1988), resulting in a decrease of the release of androstenone into the circulation.

One advantage to immunocastration is that growth performance and feed conversion of immunocastrated boars is superior to that of barrows – although not on par with intact males (as reviewed by Millet et al., 2011). Additionally, barrows tend to have fattier carcasses than both intact males, immunocastrates and female pigs (Gispert et al., 2010). Overall, although it is preferable from a feed conversion and carcass quality perspective to raise intact males; immunocastrates and females are acceptable and both perform better than surgically castrated males.

Immunocastration has been shown to be as effective as surgical castration at reducing aggression and boar taint levels (Zamaratskaia and Squires, 2008).

Another advantage of immunocastration is its inherent potential for reversal should a boar have the appropriate characteristics to become a member of the breeding herd. A recent study has examined the long term effects of GnRH immunization on

Leydig cell function and found that, although testicular function returned quickly, steroid synthesis had a highly variable return that was determined to be unrelated to antibody titre in the blood during immunization (Claus et al., 2008). Although the authors did not discuss semen and sperm quality, if the testicular function returns readily enough, the lag in the steroid synthesis should not have a large impact on the reproductive refractory period following immunization.

In addition to the popular GnRH immunization, there has been some research done on immunizing boars against androstenone itself (Williamson and Patterson,

1982). The authors compared one-time immunization and twice-delivered immunization against androstenone. The twice-immunized animals showed decreased androstenone accumulation in the fat compared to both control and once- immunized boars. However, these results were not as dramatic as the GnRH- immunized boars in previous studies. Although vaccination against androstenone did not impair growth nor change carcass composition, the combination of the lacklustre results of this study and the potentially prohibitive cost of the vaccine and implementation resulted in poor reception of this idea.

For all the positives of immunocastration, the practice has yet to become main stream in international pig production facilities. Vaccinating all intact boars in the finisher wing is expensive and time consuming (Thun et al., 2006) – particularly

as the vaccine must be injected twice over the span of 2 months and cannot be injected within 4 weeks of slaughter (Zamaratskaia and Squires, 2008). There is some suggestion that a new vaccine utilizing GnRH and a different conjugate may decrease the potential for residues in the meat although this was only briefly mentioned by the authors (Turkstra et al., 2011). In addition to the cost, the combination of attention to detail required in order to maintain this vaccination schedule and consumer concerns of residues makes this solution impractical for most pig production systems.

1.1.2.4 Other GnRH Agonists

In addition to immunocastration, there has been other research on the depression of GnRH activity utilizing a GnRH agonist and its impact on steroid production. Kauffold et al. (2010) examined Suprelorin, an implant containing a long-acting GnRH analogue (deslorin) that is currently used as male contraception in dogs. From previous studies in dogs, it is known that deslorelin decreases sperm production and testes size, although the implant’s impact on steroid production had yet to be studied. The authors measured steroid hormones, including estrone, estradiol, testosterone and their associated sulfate conjugates and compared these levels to castrated and uncastrated control boars. The authors found that deslorelin treatment suppressed testicular function, steroidogenesis and reproductive abilities in 80% (4/5) of boars treated. The discrepancy in physiological reactivity in the remaining boar was attributed to variation in individual response to the analogue dose.

The use of deslorin in particular as a potential treatment for boar taint is very new and, thus, further research is required into androstenone-specific steroid profiles, carcass composition, time of implantation, potential residues in the meat and production qualities like feed conversion (Kauffold et al., 2010).

In addition to deslorin, the effects of another GnRH agonist called leuprolide

(Lupron depot) had been previously examined in prepubertal boars (Sinclair et al.,

2001a). The authors examined the impact of long-acting leuprolide treatment on the prepubertal testicular development of cross-bred boars and its impact on steroid hormone profiles in the plasma. As expected, treatment with leuprolide decreased testicular steroid hormone concentrations in both the plasma and the salivary gland and delayed testicular development. However, hormone concentrations and testicular development rebounded without further leuprolide treatment once the animals reached puberty. The boars also showed no difference in growth rate near puberty when compared to control animals. The resumption of testicular development and steroid hormone levels at puberty suggests that animals should be treated with leuprolide closer to puberty as opposed to at a young age in order to alleviate the extra cost and labour associated with multiple injections.

Much like other GnRH agonists and immunocastration, concerns about consumer safety, carcass composition and costs are important when considering the use of leuprolide.

1.2. Steroidogenesis, 16-Androstene Steroids and Steroid

Circulation

As briefly discussed above, the regulation of steroidogenesis begins in the hypothalamus, resulting in the release of GnRH, which acts on the anterior lobe of the pituitary gland, resulting in the release of LH. LH acts directly on the adrenal glands, testes and, to a lesser extent, salivary glands of the boar (Katkov et al., 1972,

Preslock, 1980), resulting in the production of androgens from cholesterol. It is unknown whether androstenone acts on GnRH and gonadotropin excretion in a feedback loop (Andresen, 2006).

1.2.1 Location of Steroidogenesis in the Boar

Upon stimulation from LH, Leydig cells begin to convert cholesterol in the mitochondria into pregnenolone, the main precursor for eventual 16-androstene production. 16-androstene steroids include androstenone, α- and β-androstenol and androstadienone and their associated conjugates. It is thought that further pregnenolone conversion occurs within the endoplasmic reticulum of the Leydig cell

(Preslock, 1980, Cooke and Gower, 1977). Other androgen steroidogenesis occurs in the interstitial cells, with some production occurring in the seminiferous tubules of the testes in pigs (Cooke and Gower, 1977). Other researchers have examined the overall 16-androstene content of the testes and determined that there is a high amount of both free and sulfoconjugated 16-androstenes produced (Vihko and

Ruokonen, 1974), although the identity of these conjugated steroids was not studied by these authors. Sinclair (2004) found that porcine testes have the capacity to

conjugate androstenone, 3α-androstenol and 3β-androstenol, thus identifying the conjugated steroids described by Vihko and Ruokonen (1974).

In addition to the testes, the submaxillary salivary gland of the pig acts as a secondary steroidogenesis location. Boars utilize the combination of their saliva and a chomping motion to release steroid hormones –including androstenone- into the environment, encouraging a reproductive response such as lordosis from receptive females. The salivary gland is capable of producing androstenone from other 16- androstenes, although it is missing the enzymes required to complete the pathway from pregnenolone (Katkov et al., 1972). Booth (1986) studied the correlation between salivary, plasma and fat 16-androstenes in addition to other carcass parameters including bulbourethral gland weight and testes weight. The author found plasma free androstenone was well correlated with androstenone in fat as well as 16-androstene content of the saliva. Booth (1986) also determined that there is a conjugated androstene fraction in the plasma – although this was not further elucidated.

1.2.2 Steroidogenesis Throughout the Lifetime of the Pig

Steroid profiles of pigs change dramatically over the lifetime of the animal – especially in reference to the 16-androstene steroids. These differences result in differential gene expression, encouraging the growth of certain cells and not others.

Steroid hormones have a vast, important effect on the growth of an animal, beginning in the womb.

Pigs are an interesting species with regards to Leydig cell maturation and development throughout their lifetime. Boars have a triphasic pattern of prepubertal Leydig cell development, unlike humans and other mammals that have a biphasic development. Pigs have one foetal Leydig cell growth phase, one perinatal and one post-natal phase that completes around 21 days of age. The final phase of

Leydig cell development begins at approximately 90 days of age, as the boar begins pubertal development (as reviewed by Lunstra et al., 1986). Leydig cell development is largely attributed to stimulation by gonadotrophin releasing hormone (GnRH), as opposed to other gene expression, both in the womb and for the duration of puberty (Allrich, et al., 1983, Moran et al., 2002). In terms of steroidal production, Leydig cell production of testosterone is highest over 130 days of age (Allrich et al., 1983) and both sulfoconjugated and unconjugated testosterone is also found in the highest concentrations compared to other steroids during foetal

Leydig cells development (as reviewed by Raeside et al., 2006). Interestingly, work by Sinclair et al (2001a) suggests that proper pubertal testicular development relies mostly on the late foetal phases of Leydig cell growth as opposed to the perinatal phase of Leydig cell growth.

Although pigs have several very distinct pre-pubertal periods of Leydig cell development, Kwan et al (1985) found that foetal testes microsomes from pigs could not produce 16-androstenes as these cells appear to lack the required enzymes. Therefore, there is limited qualitative steroid production in the pig’s early

Leydig cell development – although steroid production does increase sharply in the final foetal stage of Leydig cell development (Kwan et al., 1985). As expected, it was

also found that 16-androstenes are more readily produced as boars begin sexual maturity, as these steroids are biologically significant for reproduction by stimulating steroidogenesis and the development of secondary sexual characteristics (Kwan et al., 1985).

1.2.3 Enterohepatic Circulation of Steroids

In order for the body to maximize the utilization of steroids, some recycling must occur. This recycling is also known as enterohepatic circulation and begins with the biliary excretion of steroids into the small intestine and continues with further intestinal metabolism by the gut microflora or the intestinal wall itself. The next step of enterohepatic circulation involves the absorption of steroids into the enterocytes of the intestinal wall and their subsequent movement into portal venous blood or the lymphatic system (Hellman et al., 1956). Steroids from the portal vein can re-enter the liver or be utilized elsewhere in the body. Enterohepatic circulation can have a dramatic impact on whole body steroid hormone levels

(Aldercrutz et al., 1979).

Bile acids are formed from circulating cholesterol and are very soluble due to their ionized micellar forms (Black, 1988). This micellar form enables them to bind to steroids and other solutes, resulting in biliary excretion. Enterohepatic circulation of bile acids is approximately 90% efficient (Hofmann, 1984), with the majority of bile acids passively absorbed in the duodenum and actively reabsorbed in the ileum of the small intestine (Hofmann, 1984, Black, 1988). The remaining

10% of bile acids undergo microbial degradation in the large intestine and are either

excreted in the feces or reabsorbed in the colon (as reviewed by Hofmann, 1984,

Black, 1988, Juste et al, 1988). Simple biliary excretion of steroids requires a molecular weight of at most 500 kDa. In order for general enterohepatic circulation to occur, compounds are require to have a molecular weight between 300 and 600; therefore, enterohepatic circulation of biliary-excreted steroids tend to involve conjugated steroids with groups such as sulfates and glucuronides (Sandberg et al.,

1967, Aldercrutz et al., 1979, Black, 1988). Steroid hormones can be further metabolised with each additional enterohepatic cycle, including conjugation and deconjugation (Eriksson, 1971).

Once excreted into the gut, conjugated biliary steroids are deconjugated by gut microflora. Intestinal microflora have been found to contain sulfatases that can hydrolyze steroids with sulfate groups conjugated to 3α- and 3β-hydroxyl groups

(MacDonald et al., 1983), similar to those found in some pig steroid hormones.

Although there have been limited studies in enterohepatic circulation of steroids in pigs, the studies that have been performed have shown promising results.

Dziuk et al (1999) fed pigs estrogen, progesterone and testosterone and examined steroid levels in the bloodstream, liver and gut over time. The authors found that all three steroid hormones are able to undergo enterohepatic circulation.

They also found that steroid levels may remain high for a number of days following steroid administration – a result that is explained by the recirculation of steroids via enterohepatic circulation. Additional research by Markovich (2001) found that steroid sulfates are absorbed into the portal venous system in the ileum and

jejunum in pigs. These results suggest that steroid hormones with similar characteristics as estrogen, progesterone and testosterone and their conjugates may also undergo enterohepatic circulation.

1.2.4 Binding of Steroid Hormones in the Blood and Saliva

1.2.4.1 Steroid Hormone Movement in the Blood

In order for hydrophobic steroids to move readily through the blood, they must be bound to proteins in the serum. As such, only small amounts of unbound steroid hormones are found in the plasma (Anderson, 1974). Steroid hormones travelling in the blood must be reversibly bound to these binding proteins in order to readily separate once the target tissue is reached. In general, two types of non- covalent bonds are formed between the steroid hormones and the binding proteins.

Non-polar interactions, such as van der Waals forces form the first low-energy interaction between lipophilic groups in the steroids. The second type of bond utilizes the interaction between steroidal oxygens and binding protein hydrogen groups to form hydrogen bonds. Both of these interactions are low energy, allowing the steroid hormones to rapidly dissociate (Burton and Westphal, 1972). Not only do these proteins enable steroid hormones to readily travel through the blood, binding proteins also act to prevent degradation and other chemical alterations

(Burton and Westphal, 1972).

Albumin is generally considered the most promiscuous of plasma binding proteins as it binds many different compounds. As such, albumin is found in high concentrations within the blood (Burton and Westphal, 1972). However, albumin

has a low affinity for steroid binding, especially in humans. Humans have a sex- hormone binding globulin (SHBG or, in some literature, SBP) that is found in low concentrations in the blood but has a much higher affinity for sex steroid hormones than albumin (Burton and Westphal, 1972). SHBG is produced in the liver and is the primary transporter of testosterone (Anderson, 1974).

SHBG was first discovered in 1958 and the first group to purify SHBG in humans was Mickelson and Petra (1975). Since then, SHBG has been found in the serum of many species including rabbits, chickens, rats and a variety of primates

(Renoir et al., 1980). However, pigs do not have SHBG in plasma or follicular fluid

(Cook et al., 1977), leading researchers to look for similar binding proteins in the blood of boars, particularly in reference to androstenone transportation. Although this binding protein has yet to be found (Zamaratskaia et al., 2008), it is important to understand the characteristics of SHBG in humans.

SHBG consists of two subunits, with a hydrophobic pocket where steroid binding takes place (Strelchyonok and Avvakumav, 1990, Grishkovskaya et al.,

2000). A homodimer is formed of the two subunits, which are considered laminin G- like domains with additional carbohydrate side chains (Grishkovskaya et al., 2000).

The laminin-like scaffolding of SHBG acts to create the hydrophobic binding pocket.

If a similar protein is found that explains the transportation of androstenone and its associated metabolites in boars, research programs could concentrate on genetic selection to increase its levels. A higher concentration of transporting

binding proteins could result in a larger clearance in androstenone from the body, thereby decreasing levels of androstenone in the fat.

1.2.4.2 Steroid Binding in Boar Saliva

There is one family of steroid binding proteins in pigs that is well characterised: the salivary pheromaxeins and lipocalins. Lipocalins are found in the saliva of boars and bind well to steroid hormones (Marchese et al., 1997).

Pheromaxein is found exclusively in pigs and is produced in the submaxillary gland.

It is not found in the adipose tissue or the blood (Booth and von Glos, 1991). In order to be produced, pheromaxein requires mature testes, as it binds with high affinity to the 16-androstene steroids – particularly androstenone (Booth and

White. 1988, Babol et al., 1996b). Any correlations to differences in androstenone accumulation in the fat are liable to be artefacts, as pheromaxein production in the salivary glands is related to overall body content of steroids and not the other way around.

Pheromaxein is made up of alpha and beta isomers, although the percentage of each isomer differs between breeds. The β-isomer is the most likely isomer to be found in the domesticated breeds compared to the wild breeds (Booth and White,

1988). Pheromaxein is also a relatively stable compound and begins to degrade in air between 21 and 37°C, approximately 72 hours after release. Although this stability in air may seem short, pheromaxein has a more prolonged activity in saliva at temperatures closer to that of porcine body temperature (Booth 1987).

In addition to pheromaxeins and lipocalins there is some evidence that salivary microorganisms also contribute to 16-androstene steroids in the mouth of the boar. After approximately 168 hours in saliva, 3α-androstenol is converted to androstenone and on to 3β-androstenol whereas 3β-androstenol is not metabolised further. This activity was attributed to microorganisms in the saliva as opposed to additional activities performed by pheromaxein or other endogenous binding proteins (Booth 1987).

1.2.5 Steroids and Target Tissues Including Adipose Tissue

Once steroids and their conjugates travelling through the bloodstream reach their target tissues, they must be taken up by the tissue. One of the target tissues for steroids, especially androstenone, is adipose tissue including adipocytes. Although hydrophobic steroids like androstenone and other free steroids may enter cells by passive diffusion or by using receptors, hydrophilic steroids such as conjugated steroids require a more active uptake into the cell. One such transporting peptide is the organic anion-transporting polypeptide (OATP). OATP and its role in the movement of conjugated steroids has been well characterised in a variety of tissues including liver, intestine, adipose tissue and the testis of man (Bossuyt et al., 1995, as reviewed by Hagenbuch and Meier, 2003 Konig et al., 2004, Mikkaichi et al., 2004, and Valle et al., 2006). OATPs act on compounds with similar characteristics as those that are excreted into the bile (Hagenbuch and Meier, 2003).

OATPs have twelve transmembrane domains that enable the transportation of a variety of compounds including sulfate steroid hormones such as

dehydroepiandrosterone sulfate (DHEAS). DHEAS has a high affinity for a number of different types OATP including OATP1A2 (OATP-A), OATP2B1 (OATP-B), OATP2A1

(OATP-D) and OATP4A1 (OATP-E) (Kullack-Ublick et al, 1998, as reviewed by

Hagenbuch and Meier, 2003, Konig et al., 2004 and Valle et al., 2006). In addition to being involved in the movement of steroids from the blood to target tissue, there is some evidence that members of the OATP1 family are found in the intestine and are involved in enterohepatic circulation (Hagenbuch and Meier, 2003).

Adipose tissue has recently been examined as having characteristics similar to that of an intracrine organ. These intracrine abilities include metabolism of steroids and control of steroidogenesis both locally and on a whole animal level. One of the ways that these steroids and their conjugates can enter into adipose tissue, including adipocytes and pre-adipocytes, is through the action of OATPs. OATPs that have been identified in adipose tissue include OATP2B1, OATP2A1 and OATP4A1

(Valle et al., 2006).

Conjugated steroids are brought into the cell by OATPs and are deconjugated through the action of steroid sulfatase (Valle et al., 2006). The newly-free steroids can be further metabolised using endogenous 5α-reductase, aromatase, 3β- hydroxysteroid dehydrogenase, and UDP-glucuronosyl transferase activity (Zhang et al., 2009, Belanger et al., 2003a, Belanger et al., 2003b, Meseguer et al., 2002). The activity of these enzymes results in the metabolism of androgens and estrogens, including testosterone, DHEA, estrogen, progesterone and androstenedione. These compounds are both metabolised and readily accumulated in the fat, resulting in an

additional storage pool for steroid hormones other than the traditional endocrine organs (Feher and Bodrogi, 1982). Although androgens and other steroid hormones are found in adipose tissue, there is some evidence that androgens may begin the lipolytic cascade, resulting in the release of stored steroid hormones (as reviewed by Blouin et al., 2009). The contradictive activity of androgens on adipose tissue as both a site of stored compounds and a precipitator of lipolytic action emphasizes the complex relationship that steroid hormones have within the body.

1.3 Androstenone Metabolism

1.3.1 Androstenone Synthesis

Androstenone is synthesized in the Leydig cells of the testes from pregnenolone in a stepwise conversion involving numerous enzymes (Figure 1.1).

Pregnenolone is first converted into 5,16-androstadien-3β-ol (androstadienol) by andien-β synthetase. From there, 5,16-androstadien-3β-ol is converted to 4,16- androstadienone by a group of hydroxysteroid dehydrogenase enzymes (HSD) – specifically 3β-HSD (Dufort et al., 2001). 4,16-androstadienone is then converted into androstenone by the enzyme 5α-reductase. Androstenone is subsequently released into the spermatic vein for circulation around the body where, due to its hydrophobicity, it accumulates well in fat (Brooks and Pearson, 1986).

There is some evidence of other androgens and steroids acting on this system and preventing the conversion of pregnenolone to androstadienol. Brophy and Gower

(1974) found that treatment of boar testis microsomes with pregnanedione

inhibited subsequent pregnenolone metabolism, thereby potentially decreasing androstenone production. However, further search within the literature did not result in discovery of further studies examining this phenomenon.

1.3.2 Androstenone Degradation and Excretion

Androstenone is degraded in the liver and salivary gland to α- or β- androstenol by the 3α- and 3β-HSD enzymes respectively (Dufort et al., 2001,

Sinclair et al., 2005). Androstenols formed in the liver are metabolised into easily excretable conjugated compounds (Figure 1.2). Conjugated androstenols, such as androstenol sulfate and androstenol glucuronide, are excreted in the urine and bile, particularly in young boars (Bonneau, 1982). Unconjugated androstenone and androstenol are the forms and metabolites of androstenone most easily accumulated in fat; therefore, conjugation is an especially important step in the prevention of boar taint (Sinclair and Squires, 2005).

Androstenol is conjugated by one of two enzymes: sulfotransferases (SULTs) or glucuronosyl transferases (UGTs) (Sinclair et al., 2004). Decreased sulfotransferase expression in the liver has been associated with increased androstenone accumulation in the fat (Sinclair et al., 2004, Sinclair and Squires,

2005). Of the sulfotransferase enzymes involved, hydroxysteroid sulfotransferases

(HSTs or SULTs) are likely to be the main contributor to androstenol sulfoconjugation in the liver and are particularly important as their activity level determines the amount of androstenone in circulation (Sinclair et al., 2004).

The reverse of sulfoconjugation, deconjugation, also occurs in the pig (as reviewed by Hobkirk, 1985). Deconjugation results in the removal of the sulfate group by sulfohydrolase or sulfatase enzymes. Sulfatases are also widely distributed in mammalian tissues (Munroe and Chang, 1987), and fifteen human sulfatases have

been described as having de-sulfation action on substrates varying from heparin and chondroitin to steroids (as reviewed by Hanson et al., 2004). Sulfatase activity has not been studied in relation to the metabolism of boar taint compounds.

However, sulfatases may have a biological role in the deconjugation of compounds known to be conjugated in the mature boar and, thus, should be further studied.

1.3.3 Androstenone Sulfate

Previously, 5α-androstenone was thought to be only found in the testis as a free steroid in the plasma and testis (as reviewed by Brooks and Pearson, 1986).

However, recent findings suggest that androstenone may also be conjugated in the testis (Sinclair and Squires, 2005), enabling androstenone to be excreted more readily and causing an increase in androstenone sulfate levels in the blood. Older works have also pointed to steroid sulfates being readily formed in the testis

(Raeside, 1969, 1971, Ruokonen, 1978). Sinclair et al. (2005) purified porcine hepatocytes and subsequently incubated the cells with a variety of androstenone metabolites and precursor compounds. The conjugated (sulfated or glucuronidated) fractions were separated from free fractions using a Sep-Pak® chromatography column and free metabolites were identified using gas chromatography followed by mass spectrometry (GC-MS). Conjugated fractions were deconjugated and metabolites were identified using the same GC-MS protocol as used to identify free metabolites. 5α-androstenone was found in the deconjugated fraction, providing indirect evidence that androstenone may be sulfated in the liver.

Figure 1.2: The sulfation pathway for the sulfation of androstenol. SULT: sulfotransferase. PAP: 3’- phosphodenosine-5’-phosphate.

Figure 1.3: The proposed mechanism for the formation of androstenone sulfate, by the enolation of its’ ketone group. ENOL: enolase; SULT: sulfotransferase.

In a related paper, Sinclair and Squires (2005) examined sulfotransferase activity and its relation to androstenone concentrations in plasma and adipose tissue. Boars were selected for maturity based on androstenone concentration in the blood. Steroid fractions were identified using HPLC, followed by GC-MS. The results of this study indicated that the majority of 5α-androstenone in the peripheral plasma was found in the conjugate fraction, suggesting that androstenone is, in fact, conjugated in intact male pigs.

However, when examining the structure of androstenone, it appears that androstenone is not easily sulfated as it lacks the hydroxyl group traditionally required for sulfotransferase activity. The enzyme family that is responsible for transforming 3-keto groups to 3-enol groups capable of sulfation is enolase (as reviewed by Pancholi, 2001) . This enzyme may be responsible for enabling the formation of androstenone sulfate (Sinclair and Squires, 2005). Therefore, it has been suggested that androstenone sulfate can be formed only through the synergistic action of sulfotransferase and enolase enzymes (Figure 1.3).

1.4 Enzymes Involved in Conjugation

1.4.1 Sulfotransferases

Sulfotransferases have been extensively studied and characterised in humans

(as reviewed by Hobkirk, 1985, Negishi et al., 2001, and Gamage et al., 2006).

Through sulfation, or sulfonation, these enzymes can act as a potential bioactivation

step for xenobiotics, or as a step in the excretory pathway from the body (Gamage et al., 2006). The sulfation of steroids specifically acts to aide in the movement and distribution of the steroids throughout the body (Hobkirk, 1985). These enzymes are present in the cytosol and the membrane fractions of cells and consist of α- and

β- pleated sheets, with the β-sheets as the primary site of action. The β-pleated sheets are highly conserved among species due to their role in binding the primary sulfate donator, 3’-phosphoadenosine 5’-phosphosulfate (PAPS) whereas the substrate binding sites differ wildly among species (Negishi et al., 2001, Gamage et al., 2006). PAPS is readily found in the cytoplasm of cells and is the main sulfate donor within the cell. There are four types of cytosolic sulfotransferases found in humans: hydroxysteroid sulfotransferase (SULT2s), estrogen sulfotransferase

(SULT1Es), glucocorticoid sulfotransferase (GST), and phenol sulfotransferase

(SULT1As). Of the four cytosolic sulfotransferases, SULT2s, SULT1Es, and the

SULT1As family have been studied in relation to androgen sulfation in many tissues including Leydig cells (Hobkirk, 1985, Hobkirk et al., 1989, Miki et al., 2002, Gamage et al., 2006).

Although there is high variability in the activity and number of subfamilies of sulfotransferases among species, there are a number of homologues to human sulfotransferases that have been studied in pigs. SULT1A, SULT1E, SULT2A1 and

SULT2B1 are just a few of the sulfotransferases characterized in porcine cells, mainly through the use of DHEA and pregnenolone and their associated sulfates

(Sinclair et al., 2005, Sinclair and Squires, 2005, Sinclair et al., 2006, Moe et al.,

2007a, Panelle-Riera et al., 2008). DHEA, DHEA sulfate, pregnenolone and

pregnenolone sulfate are used to assay sulfotransferase activity because they are readily sulfated, well characterized in other species and are readily available as radiolabeled compounds. Baranczyk-Kuzma et al. (1989) studied phenol sulfotransferase (SULT1A1) activity in the boar and the bull. The authors found that, unlike bull testes, boar tissue contains two types of SULT1A: one that is thermostable and the other, thermolabile. The presence of a thermolabile SULT1A in boars was confirmed by Sinclair (2002). This is just one example of the vast variety of sulfotransferase activity among species.

Of the sulfotransferases thought to be involved in androstenone metabolism,

SULT2A1 and SULT2B1 have been most actively studied – particularly when comparing activity in individuals and populations with high and low boar taint compound concentrations.

1.4.1.1 Sulfotransferase 2A1

It has been shown that sulfotransferase 2A1 has extensive activity in numerous pathways including steroid hormone metabolism, neurotransmitter metabolism, bile acid excretion, drug metabolism and xenobiotic metabolism

(Sinclair et al., 2006). In humans, sulfotransferases of the SULT2A family, particularly SULT2A1, have been found to be expressed in the adrenal glands, ovaries and other steroidogenic tissue (Gamage et al., 2006, Lindsay et al., 2008), with the exception of human Leydig cells (Miller and Auchus, 2011). In a recent review of cytosolic sulfotransferases, human SULT2A1 was most commonly identified as being responsible for the sulfation of hydroxysteroids, including DHEA,

and pregnenolone (Gamage et al., 2006), although SULT2A1 does not sulfate cholesterol in the adrenal glands (Miller and Auchus, 2011).

In pigs, conjugation attributed to SULT2A1 activity has been associated with increased sulfated androstenone concentrations in the plasma and, therefore, a decrease in the accumulation of androstenone in fat (Sinclair et al., 2006). This may be due to high androstenone levels causing an increase in SULT2A1 expression as the body attempts to excrete excess hormone (Moe et al., 2007a). Based on the results of porcine studies on SULT2A1 activity and results from the human-specific studies above on function and expression locations, SULT2A1 is most likely to be responsible for the formation of androstenone sulfate.

1.4.1.2 Sulfotransferase 2B1

The study of SULT2B1 in humans has yielded some interesting results. Of the two forms of SULT2B1 found in humans, only SULT2B1b has been found in male reproductive tissues (Geese and Raftogianis, 2001), whereas androstenone is thought to be sulfated in male pigs. SULT2B1a was only found in human ovary, fetal brain and colon tissue (Geese and Raftogianis, 2001). Human SULT2B1 is also highly stereo-selective to 3β-hydroxysteroids, possibly due to the location of their hydroxyl groups on the 16- and 17-carbons groups (Geese and Raftogianis, 2001), a feature androstenone lacks.

Although, based on human distribution and activity, SULT2B1 is unlikely to be involved in androstenone sulfate formation, this sulfotransferase has recently been studied for its potential impact on androstenone metabolism. SULT2B1 studies

have particularly emphasized gene expression, although SULT2B1 has yet to be characterised in porcine tissue. Moe et al. (2007) found that SULT2B1 gene expression was up-regulated in the testis of high-androstenone boars. In contrast, a study done by Moe et al. (2007a), SULT2B1 activity in the testis was high in low- androstenone boars and hepatic SULT2B1 activity was low in high androstenone boars; however, these results were associated with breed differences.

Panella-Riera et al. (2008) studied the impact of steroids on SULT2B1 activity in porcine hepatocytes. The authors examined SULT2B1 activity over time and in the presence or absence of androstenone, testosterone and estrone. The maximum SULT2B1 expression as measured by Western blotting was after 24 hours of incubation and only in the presence of testosterone and estrone; activity was not affected by androstenone. Although the study was done using hepatocytes and not testis-derived cells, the results suggest that SULT2B1, in the liver especially, may not be responsible for androstenone sulfate formation as SULT2B1 activity is not affected by androstenone.

1.4.2 Steroid Sulfatase

In order to gain a complete picture of androstenone sulfate formation and subsequent metabolism, enzymes that may be responsible for the deconjugation of androstenone sulfate back to androstenone must be examined. The enzyme group most commonly responsible for steroid sulfate deconjugation are sulfatase enzymes.

Sulfatases are generally five hundred to six hundred amino acids long and catalyze the hydrolysis of sulfated compounds – i.e. androstenol sulfate to

androstenol. The active site of these enzymes involves a polar amino acid residue and a metal cation, which together results in sulfate cleavage (Hanson et al., 2004).

As there are many different types of sulfatases, it is important to be able to differentiate amongst them. Hanson et al. (2004) state that sulfatases can be differentiated by the α-formylglycine (FGly) residue within the active site, a feature that increases enzyme specificity. The characteristics of the FGly residue is unique to specific types of sulfatases.

As the compounds of interest in boar taint metabolism are steroids, the sulfatase of interest is steroid sulfatase (STS). STS is a membrane-bound sulfatase, primarily found in the rough endoplasmic reticulum of the cell, with an unknown number of isoforms (Hanson et al., 2004). STS activity has been described in many human tissues, including ovary, adrenal glands, placenta, prostate, skin, brain and kidney (Reed et al., 2005). Miki et al (2002) found that STS activity is detectable, albeit minimally, in human Leydig cells when compared to other human cells including liver and adrenal gland cells, indicating that STS may be responsible for the deconjugation of steroids in the testis. The potential for STS to be involved in steroid deconjugation in the testis, combined with the lack of information in the literature regarding STS activity in the boar, makes the study of STS activity in boars a valuable investigation.

1.4.2 Enolases

Enolase is involved in the conversion of a ketone group to an enol, facilitating conjugation of substrates. The enolase family includes three dimeric isoenzymes: α-

enolase, β-enolase and γ-enolase. α-enolase is found in many tissues throughout the body, β-enolase is associated with muscle tissue and γ-enolase is involved in neuroendocrine and neuronal tissues (Pancholi, 2001). Enolase must form a dimer in order to be active (Chorazyczewski et al., 1987). However, enolase can also form a functional heterodimer, including αγ, αβ and γβ subunit combinations (Pearce et al.,

1976). The entire enolase family in humans is encoded by three highly conserved gene loci ENO1, ENO2 and ENO3 (Pearce et al., 1976). Due to the findings that enolase is highly conserved among species, Giallongo et al. (1990) suggested that today’s enolase genes evolved from a single gene. The conservation of the enolase sequence allows researchers to more readily extrapolate the human enolase gene loci to the corresponding potential pig loci.

All enolases are considered dimeric metalloenzymes as a metal ion is integral to their function (Reed et al., 1996). As a metalloenzyme, enolase activity requires at least one metal ion, generally magnesium (Mg2+). Generally, enolase requires two magnesium molecules in order to achieve the two-step enzymatic reaction. The first magnesium acts as a “conformational” site wherein the substrate attaches in preparation for the ketone to enol conversion. The second magnesium acts as a catalyst for the reaction (Pancholi, 2001).

Enolases are most famous for their involvement in the final step of glycolysis

– the conversion of 2-phosphoglycerate to phosphoenolpyruvate, which is the aforementioned conversion of a ketone to an enol. However, enolase functions in additional pathways in man, depending on pathology, physiology and stage of the cell cycle (Pancholi, 2001). As one can imagine, enolase has a strong role in cellular

development and recovery of hypoxic cells due to their role in anaerobic respiration

(Pancholi, 2001). Enolase’s characteristic conversion reaction of the 3-keto group of androstenone to a 3-enol group is necessary for the sulfation of androstenone, and, as such, must be also found in the Leydig cells of the testes to facilitate this conversion.

Enolase has been well characterized in human tissues, including identifying that a neuron (γ) enolase-like isozyme is found in Leydig cells of the human male

(Schulze et al., 1991). Although enolase has been characterised in porcine brain, kidney and muscle (Gorisch et al., 1999, Farrar and Deal, 1995, Oh and Brewer,

1973), porcine enolase has yet to be characterised directly from the testis.

1.4.3 UDP-glucuronosyl transferases

UDP-glucuronosyl transferases (UGTs) are endogenous enzymes found in many different organisms and many different tissues. UGTs are responsible for the glucuronidation of a variety of compounds including bilirubin, xenobiotics, and steroid hormones, including the C19 steroids. Glucuronidation involves the transfer of a glucuronosyl group from UDP-glucuronic acid to hydrophobic compounds termed aglycones (as reviewed by Barbier and Belanger, 2008, Guillemette, 2003).

In general, glucuronidation increases the excretion of aglycones from the body by changing their polarity from non-polar to very polar. This polarity results in the aglycones’ being more readily drawn into the urine and gut (Belanger et al., 2003b,

Guillemette, 2003). UGT activity has not been well characterised in the pig.

Although UGTs are widely expressed in the body, the highest expression levels are found in sources of entry and exit from the body and major metabolic

organs such as the skin, intestine and the liver. UGT proteins are between 50 and 57 kDa in size and are composed of two main families in humans: UGT1 and UGT2, with the vast majority of enzymes being termed UGT1A and UGT2B (Barbier and

Belanger, 2008, Guillemette, 2003). These families are encoded by two specific genes found in different genomic locations. In humans, UGT1 gene expression is from a single gene locus on chromosome 2. The majority of UGT1A enzymes are found within the liver whereas UGT2B are found both in the liver and other extrahepatic tissues. The UGT1A family is most famously responsible for the glucuronidation of bilirubin (UGT1A1) and deactivation of some drugs (UGT1A6,

UGT1A7) (King et al., 2000) whereas members of the UGT2B family have wide variety of functions. The highly varied identity and expression of UGT2B enzymes is due to differences in the gene structure between members of the subfamilies.

UGT2Bs tend to differ due to high variation in the N-terminus of the gene and high similarity in the C-terminus of the gene encoding UGT2B. The functional differences between UGT2Bs are great enough that if one member of the UGT2B subfamily is knocked out, other members are unable to fill their role (Guillemette, 2003).

Some groups of the UGT2B family in humans have been identified as being responsible for steroid hormone glucuronidation including UGT2B7, UGT2B15 and

UGT2B17 (Coffman et al., 1998, King et al., 2000, Turgeon et al., 2001). UGT2B expression levels in tissues may actually increase with levels of C19 steroids that have been reduced by 5α-reductase enzymes, such as androsterone (Belanger et al.,

1991). This is indicative that if androstenone acts as an aglycone, the UGT2B family

is more likely to be responsible for the formation of androstenone glucuronide than the UGT1A family.

Of the UGT2B subfamily, UGT2B7, 2B15 and 2B17 have been more widely studied. UGT2B7 has been identified as being responsible for androsterone glucuronidation as well as other androgens, estrogens and catechol estrogens (Gall et al., 1999, Coffman et al., 1998, Turgeon et al., 2001). UGT2B7 is expressed in a wide variety of tissues including liver, intestine, kidney and prostate basal cells

(Turgeon et al., 2001, Belanger et al., 2003b). In humans, there are two isoforms responsible for the wide variety of aglycones that are targeted by UGT2B7 (Coffman et al., 1998). Although UGT2B15 and UGT2B17 are expressed at lower levels than

UGT2B7, these UGTs are expressed in testis tissue and adipocytes (Turgeon et al.,

2001), two tissues that are more relevant to androstenone glucuronidation than other tissues. UGT2B15 and UGT2B17 differ slightly in their affinity for steroids, with UGT2B15 having a slightly higher affinity for steroid glucuronidation in humans (King et al., 2000). Additionally, UGT2B15 and 2B17 activities are used as markers for steroid production from the prostate of humans in order to identify prostate dysfunction (Barbier and Belanger, 2008). The fact that two UGT2Bs are used as markers for dysfunction of a steroid producing organ is very indicative of their role in steroid hormone homeostasis.

1.5 Rationale for Experiment

It is well established that androstenone is an important component of boar taint. Based on recent research, a novel aspect of androstenone metabolism – the

sulfation of androstenone – may have an impact on boar taint levels. However, this evidence is indirect, as androstenone sulfate has yet to be specifically identified and characterised. In order to fully establish the production of androstenone sulfate in the boar, further research is required. Finding direct evidence of androstenone sulfate and characterizing the potential enzymes involved will provide another piece of the androstenone metabolic puzzle. The characterisation of this novel aspect of androstenone metabolism will help further our understanding of the development of boar taint and, in the future, may help develop on-site testing methods to identify high-boar taint boars.

CHAPTER II: HYPOTHESIS AND OBJECTIVES

2.1. Hypothesis and Research Objectives 2.1.2 Hypothesis

In the mature boar, androstenone is found in both the free and sulfoconjugated forms. Leydig cells produce sulfoconjugated androstenone through the synergistic action of enolase and sulfotransferase.

2.2.1 Objective One: Discovery of Androstenone Sulfate in Leydig Cells

Objective 1: To confirm or deny the presence of androstenone sulfate in the

Leydig cell of the mature boar. The production of androstenone sulfate in the Leydig cell was examined using radiolabeled pregnenolone and non-radioactive pregnenolone and the identity of androstenone sulfate was confirmed using mass spectrometry.

2.2.2 Objective Two: Characterisation of the Role of Enolase in

Androstenone Sulfate Formation

Objective 2: To examine the role of enolase in the formation of androstenone sulfate. As it is theorized that enolase is required for the 3-keto to 3- enol conversion that is necessary for the production of androstenone sulfate, it was imperative to further examine the role of this enzyme. Testis microsome incubations with required co-factors and with and without enolase were performed, followed by HPLC analysis of the conjugate fractions from the same incubations. 36

2.2.3 Objective Three: Identification of Sulfotransferase Involved

Objective 3: To examine the relationship between sulfotransferase 2A1

(SULT2A1) expression and androstenone sulfate production in the pig and to determine whether SULT2A1 is responsible for the sulfation of androstenone.

SULT2A1 was expressed in human embryonic kidney cells and incubated with the necessary precursors and enzymes required for androstenone sulfate production.

Sulfated androstenone levels were examined using HPLC-MS/MS.

CHAPTER III: IDENTIFICATION OF ANDROSTENONE SULFATE

3.1 Introduction

The off-odour and flavour termed boar taint is caused by the production of androstenone from the testes and skatole from the gut microflora of the uncastrated boar (Gower, 1972, Bonneau, 1982). Androstenone was first identified as being partially responsible for boar taint in 1968 (Patterson, 1968). As such, the metabolism and excretion of androstenone has been heavily researched in subsequent years. There have been many attempts to isolate and quantify free androstenone and its various metabolic products, ranging from the use of enzyme- linked immunosorbent assays (ELISAs) (e.g. Sinclair et al., 2001) and high performance liquid chromatography (HPLC) (e.g. Chen et al., 2007), to gas chromatography (GC) and mass spectrometry (MS) (e.g. Sinclair and Squires, 2005).

Earlier works using radioimmunoassay showed promise; however, the antibodies used tend to cross-react with other 16-androstene metabolites (Bicknell et al.,

1976). Later works combined the incubation of androstenone with HEK293 cells over-expressing enzymes including 5α-reductase and 3β-HSD with newer, more sophisticated HPLC-MS detection methods (Dufort et al., 2001). The combination of

LC-MS results in higher sensitivity for compounds of interest.

Chen et al (2007) compared four analytical methods commonly used to examine androstenone and its metabolites: HPLC, GC, ELISA and GC in conjunction with MS. HPLC followed by MS analysis proved to be the most effective at

characterizing androstenone concentration whereas, in results echoing earlier work by Bicknell et al (1976), ELISAs were found to overestimate androstenone due to cross-reactivity of the antibodies. Recent work by Sinclair and Squires (2005) combined HPLC and GC-MS to examine the presence of conjugated and unconjugated 16-androstenes in Leydig cells and plasma. The conjugated fraction was hydrolyzed and both fractions were analyzed using HPLC. The HPLC profile of the deconjugated fraction showed the presence of androstenone. These profiles indicated, indirectly, that androstenone was found in the conjugated fraction, suggesting that androstenone is indeed conjugated in the boar.

Conjugated androstenone would be more polar than free androstenone, thereby preventing its accumulation in the fat and increasing its excretion from the body. Of the many types of conjugation, sulfation and glucuronidation impact most on steroid hormone homeostasis in humans (Gamage et al., 2006, Guillemette,

2003). Sulfation readily occurs with other steroids including DHEA and estrone, and

DHEA sulfate (DHEAS) plays a large role in steroid hormone accumulation in the fat in both men and women. It is theorized that DHEAS is converted into other sex steroids including estrogens in adipose tissue, resulting in their accumulation. The significance of this conversion as opposed to accumulation of the steroids themselves is due to the high concentration of DHEAS found in the plasma compared to other hormones (Valle et al., 2006).

Additionally, previous work has indicated that steroid sulfates, including

DHEAS and estrone sulfate (E1S) are readily formed in the testis (Ruokonen, 1978,

as reviewed by Raeside et al., 2006). As sulfates are involved in the excretion of steroids from the body, it is important to note their direct production in the boar testes. Steroid sulfates formed in the testes of the boar include α- and β-androstenol, androstadienol, the androstanediols (3α-17α-, 3α-17β-, 3β-17α-, and 3β-17β-), and the androstenediols (3α-17β-, 3β-17α-, and 3β-17β-) (as reviewed by Raeside et al.,

2006). In particular, estrogens including E1S are a major component of urinary steroid excretion from the boar (Raeside, 1983).

However, in order for androstenone sulfation or glucuronidation to occur in the boar, androstenone first requires the conversion of its 3-ketone group to a 3- enol group (Sinclair and Squires, 2005). The enzyme thought to be responsible for this is enolase, a member of the glycolytic enzymes well characterized in humans

(Pancholi, 2001). Once this conversion occurs and the 3-enol intermediate is formed, the action of sulfation enzymes such as sulfotransferases is thought to result in the formation of androstenone sulfate.

Previous studies have utilized many methods of separating conjugate and free steroid fractions from one another. Some studies have used a combination of chemically sulfated forms of the compound of interest and thin layer chromatography to differentiate conjugated and unconjugated fractions (Raeside and Howells, 1971, Hobkirk et al, 1989). Other, more recent, studies have utilized various Sep-Pak® chromatography protocols to separate the sulfated and free fractions, using diethyl ether and methanol (Sinclair and Squires, 2005, Raeside et

al., 1997, Sinclair et al., 2006) or differing concentrations of methanol (Tuomola et al., 1997, Zamaratskaia et al, 2007).

The analysis of conjugated androstenone has yet to be performed directly by mass spectrometry. Technical aspects of this analysis that must be considered include the ionization mode (negative versus positive) and methods of sample ionization. Of the many ionization methods used in organic material analysis, electron spray ionization (ESI), atmospheric pressure photoionization (APPI), and matrix assisted laser desorption ionization (MALDI) are most commonly used

(Mitamura et al., 2003). The main difference between these techniques is their optimization for specific compound characteristics. For example, ESI is most commonly used with polar molecules ranging from 100 Da to 1,000,000 Da in mass.

ESI is sometimes modified to nanospray ionization, a lower flow rate version of ESI.

Nanospray is a more sensitive technique to maximize ionization of molecules injected into the mass spectrometer (Ashcroft, 2011), resulting in more efficient ionization. Additionally, each sample can be examined in positive and negative ionization modes. Positive ionization is optimal for compounds likely to gain a proton (i.e. amines) whereas negative ionization is optimal for compounds likely to lose a proton (i.e. acids) (Downard, 2004).

The goal of this research was to confirm the presence of sulfated androstenone produced from Leydig cells after incubation with pregnenolone. This was achieved through the separation of conjugate steroids from free steroids utilizing Sep-Pak® Chromatography followed by separation and purification of

individual conjugated steroids by HPLC. Purified conjugates were collected and analyzed by mass spectrometry to identify androstenone sulfate.

3.2 Methods

3.2.1 Animals

Mature, intact Yorkshire boars between 160 and 180 days of age were acquired from the Arkell Research Station at the University of Guelph. Testes were collected immediately following euthanasia by exsanguination.

3.2.2 Leydig Cell Isolation

Leydig cells were isolated as previously described with some modifications

(Raeside and Renaud, 1983, Sinclair et al., 2005). Briefly, testes were decapsulated, sliced and rinsed with 1:10 dilution of Pen-Strep:water (Gibco®) to prevent bacterial contamination. Testis tissue was digested at 37 degrees C for between 40 and 60 minutes in digestion media containing 1 mg/mL collagenase (Type 1A from

Sigma-Aldrich Ltd), 0.05 mg/mL DNAse (bovine pancreas from Sigma-Aldrich Ltd), and 0.05 mg/mL trypsin inhibitor (Type 1-S from Sigma-Aldrich Ltd) in 100 mL media A (sodium bicarbonate (Fisher Scientific) powdered T199, L-Glutamine

(culture tested), bovine serum albumin and D-glucose (all from Sigma-Aldrich)).

Cells were separated from connective tissue by filtering using two different grade

Nylon mesh (150 um pore size followed by 75 um pore size). The filtrate was then

3x diluted with media and centrifuged at 1500 g for 10 minutes at 4°C. The pellet

was reconstituted with media and carefully layered on a Percoll (Sigma-Aldrich Ltd) gradient, in ascending order, of 60%, 40%, 34%, 26% and 21% Percoll:media.

Leydig cells were removed from the 60% and 40% Percoll gradient interface and cell identity was confirmed by microscopy. Cells were then 3x diluted with media to remove Percoll and centrifuged at 500 g for 10 minutes at 4°C. The cells were re-suspended with media and ready for incubation. Cell yield varied amongst individuals from approximately 56 million cells/ml to 75 million cells/ml, as counted by hemocytometer.

3.2.3 Leydig Cell Incubation

Leydig cells were incubated at a concentration of 2 million cells/ml media with 1.8 uCi/ml [7-3H(N)]-pregnenolone and 0.02 mM unlabelled pregnenolone to a final volume of 10 ml. Radiolabelled pregnenolone was used in order to detect metabolites that would not be detected by fluorescence or UV absorption. Cells were incubated at 37°C in a shaking water bath with 5% CO2 in air for 8 to 24 hours. Cell incubations of 8 hours were deemed optimal for androstenone sulfate production by previous research done in our lab (Sinclair and Squires, 2004). Cells were collected by centrifugation at 3000 g at 25°C for 10 minutes and stored at -20°C. The cell pellet was extracted with TRI Reagent (Sigma-Aldrich Inc) at 0.1 ml: 2 million cells and kept frozen at -70°C until further use. The supernatants were analyzed further for content of free and conjugated steroids.

Twenty-four hour incubations were done to compare production of metabolites and conjugation over time. These incubations were done using plated

Leydig cells to ensure cell survival. A total of 20 million cells (5 million cells per plate) were suspended in media A and left to adhere to plates for 4 hours. Media and free-floating cells were removed by gentle suction and media containing 1.8 uCi/ml of [7-3H(N)]-pregnenolone and 0.02 mM cold pregnenolone were added.

Supernatant containing metabolites was removed after 24 hours and kept at -20°C until further use. Cells were scraped off of plates and vortexed well with TRI

Reagent at 0.1ml:2 million cells.

3.2.4 Steroid Conjugate Separation

Incubation media was separated into conjugate and free metabolite fractions using solid phase extraction Sep-Pak® (Waters Corp. tC18, 500 mg) optimized for androgen recovery as utilized by Zamaratskaia et al. (2007) and Tuomola et al

(1997). The column was first primed using 5 mL methanol, followed by 5 mL Milli-Q water. Three milliliters of sample was diluted with 5 mL of Milli-Q water and loaded on a Sep-Pak® cartridge. Sample was slowly passed through the cartridge using very gentle vacuum pressure. The column was then washed with 5 mL Milli-Q water.

The conjugated fraction was slowly eluted using 5 mL of 47% methanol followed by the elution of the free fraction using 5 mL of 100% methanol. Sample, wash, conjugated and free fractions were all collected in glass test tubes. Radioactivity was measured by liquid scintillation counting of 100 ul aliquot of each fraction in 5 ml scintillation fluid (Ecolite; ICN Pharmaceuticals Inc.) using a Beckman LS6000Sc β-

Counter. The counts per minute (CPM) from the different Sep-Pak fractions were used to determine the average conjugation by the Leydig cells. Fractions were then sealed tightly and kept at 4°C until further use.

3.2.5 Initial HPLC Steroid Analysis

Conjugated and free fractions from Sep-Pak® separations were dried down using Savant Speed-Vac Concentrator (Thermo-Scientific) and re-suspended in 33% and 85% acetonitrile respectively (HPLC-grade acetonitrile from Fisher-Scientific).

Any samples where re-suspension was difficult were re-suspended using 100% acetonitrile.

The conjugate fraction was analyzed using Reverse-Phase High Performance

Liquid Chromatography. One hundred microliters of samples was injected in two separate runs into Agilent HPLC Chemstation (series 1100, Hewlett-Packard

Industries) running a gradient of 33% ACN:66% H2O to 90% ACN:10% H2O over 40 minutes (HPLC Protocol A) . A Phenomenex Prodigy 5u (250 x 4.60 mm,

Phenomenex Industries) column was used. Fractions for both HPLC runs were collected every 30 seconds and 1/10th of each fraction was measured for radioactivity by scintillation counter to determine metabolite separation. Detected peaks were then pooled and saved at 4°C for further analysis. For all HPLC runs on this system, ultraviolet (UV) data were collected for the following wavelengths: 210,

230, 250, 254, and 280 nm. It is not expected that androstenone sulfate would be detected in any UV range. Conjugated standards were analyzed by UV. Standards included estrone glucuronide, estrone sulfate (E1S), DHEA glucuronide, DHEAS, pregnenolone glucuronide, pregnenolone, free androstenone and free 3α- and 3β- androstenol. These standards eluted at 2.5, 3, 4, 5, 7, 8, 34, 35 and 36 minutes respectively (see Appendix).

The free fraction was analyzed using HPLC (Protocol B). The HPLC system used was a SpectraPhysics Spectra system FL2000 with a SP8800 pump, an SP8780 autosampler and a SP2490 integrator. Two hundred microliters of samples were injected onto a Phenomenex Prodigy 5u HPLC column (250 x 4.60 mm) running an isocratic program of 85%ACN:15% H2O at 1.0 ml/min for 25 minutes. Radioactivity was measured on-line using a Canberra-Packard 500TR flow scintillation analyzer.

Metabolite peaks were tentatively identified using known standards of 16- androstenes and estrones. Estrone, DHEA, androstadienone, androstadienol, 3α- and 3β-androstenol and androstenone reference standards eluted at 3, 4, 12, 14, 17,

19 and 21 minutes in the initial standards.

3.2.6 Conjugate Peak Analysis and Identification

The peaks found in the conjugate fraction of HPLC were further analyzed to determine their identity. Isolated peaks were dried down using a Speed-Vac concentrator and solvolyzed by incubating with 5 mL of 1/100 v/v trifluoroacetic acid/ethyl acetate at 40°C for 18 hours. This technique chemically removes sulfate groups from compounds of interest. Samples were dried by a nitrogen stream and re-suspended in 85% ACN. These samples were then run on HPLC using the free fraction protocol B and metabolite peaks were identified by comparison to known standards.

In order to confirm that these peaks were sulfated as opposed to glucuronidated, sample replicates were treated by solvolysis only (a), solvolysis followed by incubation with β-glucuronidase (b) and β-glucuronidase incubation

only (c). β-glucuronidase incubations consisted of the combination of the sample,

500 ul of 0.5M sodium acetate buffer (pH 5.0) and 1250 Units of β-glucuronidase

(Sigma, type B-1 from bovine liver) in a screw top tube and incubated for 18 hours in a water bath at 37°C. Samples were then examined using HPLC Protocol B for free samples.

3.2.7 Mass Spectrometry

The identities of unknown conjugate peaks were confirmed using Waters

Micromass Global Ultima High Resolution QToF, an Electrospray Triple Quadropole with Time of Flight Detection with a typical resolution of less than 5 ppm and mass accuracy of approximately 5500 m/z. Samples were analyzed in negative mode by nanospray at approximately 200 nL/min using Proxeon nanospray emitters with a

360 um x 75 um taper tip with borosilicate.

Approximately 10 uL of each sample was pipetted into glass emitters and the nanospray tip broken such that the flow rate was between 200-1000 nL/min. The xyz position of the emitter was adjusted for maximum intensity. The data was collected in full scan mode from 50 to 3000 m/z in order to search for the metabolites of interest. The intensity was optimized by adjusting the capillary voltage in order to allow individual spectral counts of up to 200 counts per second.

This ensured proper peak shape, width and accuracy.

In order to confirm the ionization efficiency of using positive versus negative mode, standards were analyzed in both modes, resulting in negative ionization proving to be the best option. The success of the use of negative ionization mode is

reflective of the literature that suggests that this mode of analysis is best for samples with groups that are ready proton donators such as acids and compounds with a high number of loosely-bound hydrogen atoms (Rauh, 2009). The ability of sulfated compounds to donate protons is well established.

3.2.8 Statistical Analysis

Statistical analysis was performed using SAS/STAT version 9.2 (SAS Institute

Inc.). The differences in the percentage of steroidal conjugates as indicated by conjugated fraction and conjugated steroidal content (uM) between 8 hours and 24 hours were analyzed using a T-Test for statistical significance.

3.3 Results

3.3.1 Conjugation Capacity of Leydig Cells

When radioactivity was counted in the Leydig cell incubation supernatant following Sep-Pak® chromatography separation, a general trend was seen. The majority of radioactivity was present in the conjugate fraction - between 51% and

89% - with conjugation capacity of the cells and conjugation percentage varying between individuals (Figure 3.1). The amount of radioactivity found in the free fraction also varied amongst individuals, with a range between 7.0% and 41.0% of total radioactivity measured. The amount of radioactivity found in both the flow- through (sample) and the water wash fractions of Sep-Pak® chromatography separation was relatively small, with the sum of radioactivity for both fractions ranging from 0.9% in individual 5 to 11.8% in individual 3 (Appendix 1a and b).

The conjugation capacity indicated the steroidal conjugation potential of cells. These capacities also varied amongst individuals and ranged from 42.8 mmol conjugated steroids/million cells in the youngest individual to 177.5 mmol/million cells in another individual (Table 3.1). Free steroid production was also examined.

Certain individuals, such as individual 5, had more efficient conjugate conversion of free steroids as they had the lowest free steroid content compared to other individuals but did not have proportionally the lowest conjugation capacity. It is unlikely that this difference in most individuals is due to the loss of radiolabeled material in the flow-through and water wash fractions as, with the exception of individual 3, the total percentage of radiolabeled material lost was very small.

Additionally, individual 11 had the highest amount of free steroid content but not the highest conjugation capacity, indicating that the conjugation in this individual was not very efficient. As Figure 3.1 and Table 3.1 suggests, there was a wide variation in conjugation capacity of cells between individuals.

Individual Total Conjugation Capacity Total Free Steroid (mmol/million cells) Production (mmol/million cells)

Pig 1 42.8±3.7 23.2±2.8

Pig 2 49.3±1.4 16.5±2.4

Pig 3 110.3±4.4 22.5±4.4

Pig 4 125.5±2.1 20.3±2.0

Pig 5 132.7±1.9 16.0±1.9

Pig 6 123.2±4.0 24.3±4.0

Pig 7 106.6±4.1 34.2±4.2

Pig 8 165.0±1.9 26.1±1.9

Pig 9 177.5±2.8 15.1±2.7

Pig 10 132.9±5.7 52.9±6.0

Pig 11 113.5±5.0 70.2±5.7

Pig 12 170.3±1.6 19.5±1.9

Pig 13 164.0±2.4 25.8±2.6

Table 3.1: Conjugation capacities of Leydig cells incubated with [7-3H(N)]-pregnenolone reported as a mean (n=3), +/- SE.

100.00

90.00 Pig 1 80.00 Pig 2 Pig 3 70.00 Pig 4 60.00 Pig 5 50.00 Pig 6 Pig 7 40.00 Pig 9 30.00 Pig 10 20.00 Pig 11 Pig 12 10.00 Pig 13 0.00 Conjugate Free

Figure 3.1: Relative radioactive content of samples following separation of conjugate and free fractions by Sep-Pak Chromatography from 8 hour Leydig cell incubations. The remaining minor amount of the percentage of radioactivity of samples was accounted for in the sample and wash fraction (Appendix 1 a and b). 3.3.2 Analysis of Conjugated Fraction

Conjugated metabolites, as separated by Sep-Pak® chromatography, were analyzed by HPLC. As there were no standards available for androstenone sulfate with which to optimize this HPLC method, samples from the first six individuals were used. While optimizing the HPLC for our compound of interest, it was discovered that a gradient of 33%ACN:67% H2O to 90% ACN:10%H2O at a flow rate of 1.0 ml/min over 40 minutes, followed by a wash of 100% ACN for 12 minutes was needed. This is the aforementioned HPLC Protocol A. Because this method of HPLC separation was only used for individuals seven to thirteen, the earlier individuals

(numbers one through six) were not included in further metabolite testing.

With HPLC Protocol A, peaks were consistently found at 2, 3.5, 4, 5, 7, 15-16,

18 and 21 minutes (Figures 3.2a and 3.2b). The earlier peaks were identified, using standards in addition to peak solvolysis followed by HPLC protocol B analysis, as estrone glucuronide, E1S, DHEA glucuronide and DHEAS (Appendix 2a and 2b).

Additionally, the peak at 15-16 minutes was found to be the coelution of 3β- androstenol and 3α-androstenol.

The combined radiolabeled material from peaks between 18 and 21 minutes did not correspond to any UV output. Due to its ineffective UV absorbance, this peak was further examined as a potential androstenone conjugate using MS. Additionally, in many replicates, this peak began at 18 minutes and ended at approximately 21 minutes. If a sample had a peak at 21 minutes, it did not have a peak at 18 minutes, lending to the idea that the peaks 18 and 21 minutes were one in the same. As such, peaks at 18 and 21 minutes were isolated for further analysis.

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Figure 3.2b: The same graph as Figure 2a with output from HPLC Protocol A but showing the smaller peaks, located at 7, 8, 15 and 21 minutes. In this individual, the peak at 7 minutes corresponded to DHEA glucuronide elution whereas the peak at 8 minutes corresponded to DHEAS elution. The small peak at 21 minutes corresponds to the expected elution of androstenone sulfate. 3.3.6 Conjugation Over Time: 8 Hour vs 24 Hour Incubations

Three individuals were selected to examine the differences in conjugation between 8 hour and 24 hour incubations. These individuals were selected at random and were pigs eight, ten and eleven. Sep-Pak® chromatography did not indicate any statistical difference in conjugation between these time points (Table

3.2). HPLC separation also did not show any major differences in metabolite formation (Table 3.3), including the presence of unmetabolized pregnenolone.

Conjugated Fraction (%) Free Fraction (%)

Pig 8 - 8 Hour 80.5 15.2

Pig 8 - 24 Hour 68.0 26.5

Pig 10 - 8 Hour 65.5 27.5

Pig 10 - 24 Hour 83.2 7.8

Pig 11 – 8 Hour 51.8 41.4

Pig 11 – 24 Hour 48.5 49.1

Table 3.2: Comparison of percentage of total radiolabeled material in the conjugated and free fractions of Leydig cell incubations at two time points of three individuals.

8 Hour Incubation (uM) 24 Hour Incubation (uM)

Peak at 4 minutes 0.33 2.15

Peak at 5 minutes 1.09 1.48

Peak at 7 minutes 0.92 1.09

Peak at 10 minutes 0.12* 0.09*

Peak at 16 minutes 0.02 0.02

Peak at 21 minutes 0.01** 0.01**

Table 3.3: Comparison table of HPLC output from pig 8, showing no differences in compound elution times or relevant steroidal content between 8 hour and 24 hour incubations. The peak at 4 minutes corresponds to steroid hormones other than our compound of interest, androstenone, and thus the large difference between time points is not relevant. The peak at 7 minutes corresponds to unmetabolized pregnenolone. Although pregnenolone is present in both samples, the size of the peak does not differ widely compared to the other peaks. * and ** indicates a statistically significant difference in steroidal conjugate production between time points.

3.3.3 Discovery and Identification of Androstenone Sulfate

The peaks found previously at 18 and 21 minutes were solvolyzed overnight and run on the HPLC protocol B for free androstenone. The solvolyzed sample for this peak regularly showed a peak at 4.5 minutes and a peak at the unconjugated androstenone standard of 20 minutes (Figure 3.3). The identity of the peak found at

21 minutes from HPLC Protocol A of the conjugated fraction was thus determined to be sulfated androstenone. The identity of androstenone sulfate was further examined using mass spectrometry. The peak found at 3.5 minutes during this process was further treated with β-glucuronidase and found to be a background glucuronide of unknown identity and origin and is very unlikely to be androstenone glucuronide.

Pig 5: Solvolyzed Peak at 21 Minutes 120.00 100.00 80.00 60.00 40.00 20.00 0.00 0.00 1.50 3.00 4.50 6.00 7.50 9.00 10.50 12.00 13.50 15.00 16.50 18.00 19.50 21.00 22.50 24.00 25.50 27.00 28.50 30.00 32.00 33.50

Figure 3.3: Example graph of solvolyzed peak at 21 minutes from HPLC Protocol B, showing a glucuronide conjugate at approximately 4.5 minutes and free androstenone at 21 minutes. 3.3.4 Androstenone Sulfate Identification

Peaks from HPLC Protocol A from two individuals (9 and 13) were analyzed by mass spectrometry to identify compounds that eluted at the anticipated elution time for androstenone sulfate. Standards of glucuronidated DHEA, E1 and pregnenolone, DHEAS, E1S, 3α-androstenol, 3β-androstenol, androsterone, androstenone and androstanol were all examined by MS in order to aid in the identification of peaks from samples.

Based on predicted chemical formula and structure, the expected mass of androstenone sulfate was 352-353 m/z (Figure 3.4). Compounds matching this mass were not found in any peaks that eluted between 12 and 22 minutes from

HPLC Protocol A from individuals 9 and 13. However, a sulfated compound corresponding by mass and degradation pattern to 3-keto-4-sulfoxy-androstenone was identified with a mass of 368 m/z. Although the mass spectrometric output of this compound did not show a peak at the mass of the androstenone or any other standards run, the sulfate group is clearly seen with a mass of 96.96 (or 97) m/z

(Figure 3.5). This lack of androstenone in the degradation pattern could be due to the impact of the extra oxygen found on this predicted 3-keto-4-sulfoxy- androstenone. This is believed to be the sulfated form of androstenone as the removal of the sulfate group with a mass of 97 m/z would result in the formation of androstenone as opposed to 4-hydroxyandrostenone. Although the compound with a mass of 287.1649 corresponds to a predicted hydroxylated androstenone, removal of a sulfate group from this position would have a mass of 80 m/z, corresponding to the removal of a SO3 group. The theoretical formation of 3-keto-4-sulfoxy- androstenone (henceforth known as androstenone sulfate or ANS) is summarized in

Figure 3.4.

Figure 3.4: The formation of 3-keto-4-sulfoxy-androstenone from androstenone. Enolase (A.) converts the 3-keto group found in androstenone to a 3-enol intermediate. Sulfotransferase (B) attaches the sulfate group to C4, resulting in the double bond between C3 and C4 to convert the C3 oxygen back to a keto group. The 3-sulfoxy-androstenone with a mass of 352 was not found by mass spectrometry.

Figure 3.5: Tandem mass spectrometry output for peak found at 18-21 minutes identified with a mass of 367.1 m/z. This could be an ionized form of the compound found at 368.1597 m/z, which corresponds directly to the predicted mass of 3-keto-4-sulfoxy-androstenone. The peak with a mass of 96.9623 (97 m/z) corresponds to a sulfate group, whereas the remaining peaks could be due to the degradation of ANS.

3.3.5 Androstenone Sulfate vs Androstenone Production in Incubated

Leydig Cells

For individuals seven through thirteen, the conjugate fractions and free fractions from Sep-Pak® chromatography were run on HPLC Protocol A and

Protocol B, respectively, and androstenone sulfate production was compared to free androstenone production within the individuals. Although androstenone sulfate production was less than 1% of total radiolabeled pregnenolone added, it was still detectable in all individuals (Table 3.4). For nearly all individuals, with the exception of individual 7, conjugated androstenone content was higher than free androstenone content in the Leydig cells.

Individual Free Conjugated

Androstenone (%) Androstenone (%)

Pig 7 0.003 0.001

Pig 8 0.002 0.021

Pig 9 0.005 0.007

Pig 10 0.006 0.041

Pig 11 0.018 0.120

Pig 12 0.036 0.135

Pig 13 0.021 0.055

Table 3.4: Amount of free and conjugated androstenone as a percentage of total radiolabeled pregnenolone added (%).

3.3.6 Conjugated Steroid Production

The amount of each conjugated steroid produced as a percentage of pregnenolone converted to other steroids is summarized in Table 3.5. Briefly, androstenone sulfate production was highest in individual 12, with a percentage

conversion of 0.14% and lowest in individuals 7 and 9 at 0.01%. Similarly, individual 11 had the highest percentage conversion to the androstenol sulfates at

0.41% and 1.03% for 3α-androstenol sulfate and 3β-androstenol sulfate respectively; the lowest percent conversion to the androstenol sulfates were found in individual 9, with 0.002% and 0.01% for 3α-androstenol sulfate and 3β- androstenol sulfate respectively. With the exception of individual 11, the formation of glucuronidated DHEA and estrone was higher than the formation of sulfated

DHEA and estrone. This is unexpected as previous research indicates that the formation of glucuronidated steroids is significantly lower than the sulfation of steroid content in Leydig cells (Sinclair, 2004).

Pig-7 Pig-8 Pig-9 Pig-10 Pig-11 Pig-12 Pig-13

E1G 1.040 0.970 2.100 14.800 6.770 3.360 4.450

DHEAG 0.270 1.650 9.050 1.550 1.980 6.470 3.610

E1S 0.050 0.530 0.560 0.550 1.110 1.330 2.590

DHEAS 0.010 0.280 0.110 0.150 3.000 2.220 1.960

BAS 0.010 0.020 0.010 0.170 1.030 0.300 0.190

AAS 0.004 0.020 0.002 0.020 0.410 0.300 0.190

ANS 0.001 0.020 0.010 0.080 0.120 0.140 0.060

Table 3.5: Amount of conjugated steroid hormones produced from the addition of radiolabeled pregnenolone as indicated by the percentage conversion of pregnenolone (%). E1G = estrone glucuronide, DHEAG= DHEA glucuronide, E1S = estrone sulfate, DHEAS= DHEA sulfate, BAS=3β- androstenol, AAS=3α-androstenol, ANS=androstenone sulfate.

3.3.7 Free Steroid Profiles for Individuals

16-Androstene steroidal profiles were compared between individuals. While radiolabeled material attributed to androstadienone was generally the highest,

relative amounts of 3β-androstenol, 3α-androstenol and androstenone were variable. In pig 7, androstenone was higher than the androstenols whereas in pigs 8,

9, 10 and 13, androstenone was the least abundant (Table 3.6).

Pig Androstadienone Androstadienol 3β- 3α- Androstenone Androstenol Androstenol

pM % pM % pM % pM % pM %

7 40.1 46.2 40.7 46.8 1.7 1.96 1.3 1.5 3.1 3.6

8 100.5 45.3 40.2 14.6 50.8 22.9 10.3 4.6 20.1 9.1

9 310.4 34.4 410.8 45.5 50.6 5.61 70.1 7.8 61.0 6.8

10 700.6 46.9 600.7 40.2 80.8 5.41 50.4 3.4 60.8 4.1

11 1910.0 52.5 530.1 14.6 530.2 14.6 430.6 11.9 220.7 6.1

12 1460.0 52.7 430.7 15.5 450.4 16.3 110.8 4.0 350.3 12.6

13 4160.0 57.0 990.1 13.6 1470.0 20.1 440.9 6.0 240.8 3.3

Table 3.6: Concentrations of steroids (in pM) and percentage of total 16-androstene production in peaks attributed to five 16-androstene steroids from 8 hour Leydig cell incubations.

3.4 Discussion and Conclusions

The production of unconjugated or free androstenone has been extensively studied in many breeds and ages of boars, particularly when related to its release into the plasma and accumulation in the fat (e.g. Bonneau, 1982, Booth, 1982, Aldal et al., 2006). Whole body androstenone concentration is highest in the boar after puberty and can reach threshold levels in the fat as early as 110 days of age (Aldal et al., 2006). Threshold levels in this case were determined using consumer sensory panels for taint detection. As the animals used in this study had already achieved

sexual maturity based on their ages, androstenone production should have reached its peak.

Saat et al. (1972) characterized testicular concentrations of 3α- and 3β- androstenols and androstenone production when mature boars were injected with radiolabeled pregnenolone. The authors found that the radioactivity from these compounds in the testis was as follows: androstenone (236 dpm/g), 3α-androstenol

(534 dpm/g) and 3β-androstenol (876 dpm/g). These results follow a similar pattern of metabolism to that of this study, with the highest amount of radioactivity corresponding to 3β-androstenol followed by 3α-androstenol and, finally, androstenone – with the exception of two individuals where androstenone production was higher than both 3α- and 3β-androstenol. Unfortunately, as these authors did not analyze pregnenolone metabolism in Leydig cells themselves, it is not possible to directly compare the numeric values found by Saat et al (1972) and those reported in this chapter.

Sinclair (2005) examined the concentration of free 16-androstenes produced over time in Leydig cell incubations under very similar conditions to those discussed in this chapter. The concentrations of free steroids seen in the author’s experiment are higher than those found in this experiment, potentially due to the use of a lower amount of radioactive material and higher amount of unlabeled material compared to the current study.

As can be expected, differences in capacity for steroidal conjugation exist between individuals, as seen by the range of conjugation capacity of isolated Leydig

cells (Table 3.1). It would be expected that breed differences would also be seen, had other breeds of boars been used for Leydig cell incubations. These differences may be due to genetic variations in steroidal metabolizing enzymes (Moe et al.,

2007). The examination of androstenone sulfate production in the Leydig cells of other pig breeds would be necessary to completely characterize the role of androstenone sulfate in boar taint metabolism.

In addition to overall conjugation of steroids by Leydig cells incubated for 8 hours with pregnenolone, the conjugation of steroids by Leydig cells incubated for

24 hours was examined. Initially, 8 hour Leydig cell incubations were performed due to research by Sinclair (2005), indicating that maximum steroidal conjugation occurred at 8 hours. Although the additional 16 hours of incubation did not change the amount of radiolabeled material in the conjugate fraction of most individuals, it did result in an apparent increase in conjugation in individual 10. The differences in optimal conjugation time between individuals could be due to altered steroidal conjugation enzyme activity.

Although overall conjugation capacity is important to examine, the characterization of the type of conjugation occurring within the cells is equally important. When examining the proportion of glucuronidated estrone and DHEA compared to sulfated estrone and DHEA, an unexpected trend was noticed (Table

3.5). Previous research has indicated that, in the Leydig cells of the boar, sulfated steroids were responsible for 95% of the total conjugate fraction, with the remaining 5% attributed to other forms of conjugation including glucuronidated

steroids (Sinclair, 2004). Additionally, UDP-glycosyltransferase enzymes (UGT) involved in the glucuronidation of steroid hormones are found in highest abundance in metabolic organs such as the liver as opposed to primary steroidogenic tissues such as the testes (Guillemette, 2003). However, with the exception of individual 11, estrone and DHEA glucuronides were more abundant than their sulfated counterparts. This could be due to an overestimation of glucuronide content as the

HPLC protocol was optimized to ensure the coelution of both estrone and DHEA conjugates. This coelution made it difficult to calculate the relative percentage pregnenolone conversion to E1S, DHEAS, DHEAG and E1G. In order to quantify the production of these conjugates, an HPLC protocol optimized for estrone and DHEA conjugate analysis should be used – however, as this was not the primary purpose of this study, the protocol used was not ideal for this analysis.

One of the main purposes of examining two time points in Leydig cell incubations was to determine whether pregnenolone was completely metabolized to its metabolites, including androstenone. Although there was some pregnenolone remaining at 8 hours, this pregnenolone concentration did not differ between 8 hours and 24 hours, indicating that pregnenolone is effectively metabolized by 8 hours (Table 3.5). The more effectively pregnenolone is metabolized, the more likely it is that androstenone and its associated conjugates are formed. As pregnenolone was effectively metabolized at 8 hours, Leydig cell incubations from the remaining individuals were performed for 8 hours only.

When analyzing the HPLC runs of conjugate fraction, a pattern was noticed.

Due to its inefficient UV absorbance, androstenone conjugates were not expected to be detected by UV. Therefore, it was important to use radiolabeled material in order to visualize any androstenone conjugate peaks. A peak was noted at 18 minutes to approximately 21 minutes in the radiolabeled-visualized HPLC output but was not found in any of the UV-visualized HPLC output of the same sample. It was theorized that this peak was an androstenone conjugate and this provided a basis for choosing specific peaks for further analysis.

An additional method of examining conjugate identity by the removal of sulfate groups by solvolysis was used. Following deconjugation, it was determined that a peak found at 21 minutes in HPLC Protocol A was in fact androstenone sulfate. The aforementioned glucuronide peak at 3.5 minutes in the HPLC Protocol B analysis (Figure 3.3) is unlikely to be androstenone glucuronide as its elution would occur prior to the elution of androstenone sulfate and thus would not be present in the 21 minute peak found in HPLC Protocol A. Additionally, it is unlikely that androstenone glucuronide is produced in a quantity that is easily quantitatively measured due to the low steroid glucuronide conjugation in the Leydig cell (Sinclair,

2004).

Upon examination of HPLC output from time points between 17 minutes and

21 minutes by mass spectrometry, it was determined that a sulfated form of androstenone was eluted at 18 minutes. This elution time corresponds with peaks taken from radiolabeled samples where the peak at 18 minutes effectively extended

to 21 minutes. It was found that these peaks were one in the same and contained androstenone sulfate.

It was originally expected that androstenone was sulfated at the 3-C position as opposed to the 4-C position. This expectation resulted in the projected mass of androstenone sulfate falling between 352 and 353 m/z by mass spectrometry.

However, no compounds corresponding to this mass were found in any of the samples examined. The compound found at an elution time of 18 minutes with a mass of 369 m/z was indicative of a 4-C sulfated form of androstenone, resulting from the reaction between the enol intermediate and a sulfate group as described in

Figure 3.6. The formation of the enol intermediate from a keto group is well established (Morrison and Boyd, 1966) and results from the movement of the double bond from the 3-keto-position oxygen to form a double bond between the 3- and 4-carbons catalysed by the enolase enzyme. This double bond movement results in the exposure of a single bonded, charged oxygen at the 3-position. In this case, in order for the sulfation to occur, the sulfotransferase targets the available 4- position carbon, resulting in the removal of the hydrogen and the addition of the sulfate group. The C=C bond is moved back to the 3-C position, resulting in the formation of a 3-keto group and sulfated androstenone named 3-keto-4-sulfoxy- androstenone.

When examining sulfated compounds, it is important to know the mass of the sulfate group, which can change depending on how it is attached to the compound of interest. This has been extensively studied by Yi et al. (2006). The authors of this

paper attempted to group sulfated compounds into four groups based on their structure as well as the mass of the removed sulfate group as indicated by MS/MS.

Enols and alcohols, such as the originally projected androstenone sulfate generally have a sulfate group with a mass of 80 m/z as determined by MS/MS. The sulfate group, in this case, is an uncharged SO3 molecule. This occurs through the same mechanism as the originally projected androstenone sulfate – with the sulfate group attaching to the enol or alcohol O (Figure 3.4). However, according to their Scheme

2, which is reserved for special-case alcohols and enolated compounds, sulfate ions can produce a characteristic ion with a weight of 97 m/z, as we found in 3-keto-4-

– sulfoxy-androstenone. The sulfate ion in this case is a charged SO4 group. This is formed through the addition of the sulfate group to a double bonded C, found in benzene structures. The double bonded C of this Scheme 2 (Figure 3.6) is similar to that found in the projected enol intermediate of androstenone, prior to the addition of the sulfate group. The differentiation in sulfate ion mass indicates that the sulfate group in our androstenone sulfate is not attached to the 3-keto group and is thus attached either to the 2-C or 4-C position, depending on where the double bond from enolation occurs. We theorize that it occurs at the 4-C position, resulting in the

3-keto-4-sulfoxy-androstenone. Additionally, it is unlikely that the removal of the 4- sulfoxy group would result in the formation of 3-hydroxyl-androstenone as the 3- enol state is unstable and short lived (Morrison and Boyd, 1969). It is much more likely that the 3-enol will convert back to the more kinetically favourable 3-keto group, resulting in androstenone.

Figure 3.6: Scheme 2 adapted from Yi et al. (2006). Removal of sulfate group from carbon, resulting in a C=C bond and a charged sulfate group with a mass of 97 m/z. This is what is theorized to occur at the 4-carbon of androstenone, resulting in the formation of androstenone after the removal of the sulfate group. Mass spectrometric analysis of androstenone found in fat and plasma has been recently studied (Magard and Berg, 1995, Tuomola et al., 1998, Chen et al.,

2010). All MS analyses of these compounds has been preceded by isolation of steroids by either HPLC or, in the case of fat, packed column supercritical fluid chromatography. In this study, HPLC preceded MS analysis - although the samples were not run directly on an LC-MS system. The HPLC protocol allowed for the separation and isolation of conjugated metabolites for further analysis. Although there are published analyses of de-conjugated and free androstenone fractions (i.e.

Sinclair and Squires, 2005, Chen et al., 2010), direct MS analysis of conjugated androstenone had not yet been done.

In terms of MS ionization methodology, electron spray ionization (ESI) is more often used for steroid analysis when compared to atmospheric pressure photoionization (APPI), although APPI is more sensitive for the analysis of multiple steroids by LC-MS (Mitamura et al., 2003, Rauh, 2009). ESI-MS is also frequently used in conjugated steroid analysis, particularly for E1S and DHEAS analysis (Diaz-

Cruz et al., 2003, Mitamura et al., 2003). For the purposes of this study, nanospray

ESI was determined to be of adequate sensitivity as multiple steroids were not present in each individual fraction sample and the compounds of interest were conjugated.

Rauh (2009) suggests that steroids in human pediatric medicine are more effectively analyzed in positive ionization mode, although this is not true of every steroid. As the present study did not examine common steroids found in pediatric medicine but rather porcine sex steroids, standards were analyzed in both positive and negative ionization modes. Negative mode was determined to be the most appropriate analytical mode. The use of nanospray tips acted to increase the efficiency of initial ionization, thereby increasing the likelihood of all compounds in a sample being effectively analyzed.

As androstenone is capable of sulfoconjugation, androstenone likely undergoes other forms of conjugation, including glucuronidation. This was examined utilizing β-glucuronidase treatment. Although deconjugated glucuronide fractions retained little radioactivity, it is possible that the material was lost during the multiple drying phases of analysis. Every effort was made to ensure dissolving of samples after every drying period; however, this was not always possible. In order to further characterize androstenone glucuronidation, the samples should not be completely dried when possible prior to dissolving in solvents such as acetonitrile.

As such, an important source of discrepancy amongst individual steroidal analyses was the difficulty found during dissolving of steroid hormones following complete drying of fractions. Whether following Sep-Pak® chromatography or deconjugation

protocols, it was difficult to completely dissolve all dried samples, even in solutions as effective at dissolving steroids as acetonitrile. This resulted in the potential for some steroids remaining in the test tubes and not being analyzed by HPLC. This difficulty could selectively affect less polar steroids as they would not dissolve as readily in solution compared to more polar steroids. Due to this difficulty, although androstenone sulfate was not found by solvolysis in every individual, it is possible that this is a result of a difficulty in resuspension of samples.

Sinclair and Squires (2005) lay excellent ground work for the discovery and identification of sulfated androstenone upon which this chapter elaborated.

Although initially unexpected, the identification of a 3-keto-4-sulfoxy- form of androstenone by mass spectrometry confirmed indirect evidence of androstenone conjugation found by other authors (i.e. Sinclair and Squires, 2005, Chen et al.,

2010). As it has been proven that androstenone is capable of conjugation, it is now necessary to identify the enzymes involved, including the role of enolase and sulfotransferases such as SULT2A1 and SULT2B1 and to investigate the role of androstenone sulfate on a whole animal physiological level.

CHAPTER IV: CHARACTERIZATION OF THE ROLE OF ENOLASE IN

ANDROSTENONE CONJUGATION

4.1 Introduction Microsomes are a useful tool to study metabolism in cell types of specific origin, including the testis and liver. Free enzymes derived from the fragmentation of the endoplasmic reticulum and other membrane-bound enzymes are present in microsomes. These enzyme-containing vesicles are used often to study cytochrome

P450 metabolism in a variety of cell types (Voet and Voet, 2004).

Among the enzymes found in testicular microsomes, andien-β synthase and other cytochrome P450s directly involved in the conversion of pregnenolone to its

16-androstene metabolites have been characterised (Samuels et al., 1975, Suhara et al., 1984, Squires, 1989). However, in order for these catalytic reactions to occur, the addition of the reduced form of the coenzyme nicotinamide adenine dinucleotide phosphate (NADPH) and any additional cofactors or coenzymes are necessary as the microsomes do not contain high levels of cofactors necessary for metabolic function

(Voet and Voet, 2004). The activity of the few membrane-bound conjugating enzymes in porcine testicular microsomes has yet to be studied.

In order for androstenone to be sulfated, the conversion of its 3-keto group to a 3-enol group is necessary. This conversion is theorized to occur due to the action of the metalloenzyme enolase (Pancholi, 2001, Sinclair et al., 2005). The mammalian enolase family consists of three isozymes: α-enolase, β-enolase and γ-

7 2

enolase. Enolase activity is found in many tissues throughout the body, although β- enolase and γ-enolase are associated with muscular and neuronal tissues respectively. Alpha-enolase is found throughout the body, including the testes

(Pancholi, 2001). Enolase activity has been well-characterized in rats and humans

(Schulze et al., 1991, as reviewed by Pancholi, 2001). Although enolase has been characterised in porcine brain, kidney and muscle (Gorisch et al., 1999, Farrar and

Deal, 1995, Oh and Brewer, 1973), porcine enolase has yet to be characterised directly from the testis. Porcine-derived enolase is not commercially available for scientific research.

The purpose of this study was to determine the role of enolase in the formation of androstenone sulfate from pregnenolone by testis microsomes.

4.2 Methods

4.2.1 Chemicals and Materials

Unless otherwise stated, all chemicals and materials were purchased from

Sigma-Aldrich Canada.

4.2.2 Testes Microsome Preparation

Frozen whole testes from individuals from Chapter 3 were removed from

-70°C freezer and allowed to thaw slowly to room temperature. Three grams of testes tissue were added to 10 mL of Buffer A consisting of 50 mM Tris-HCl, 100 mM potassium chloride, 10 mM EDTA and 10 uM PMSF at pH 7.5. Tissue was

homogenized using a Polytron Kinemetica GmBH (Brinkmann Instruments, Canada) homogenizer. Homogenized testis was centrifuged for 20 minutes at 20,000 g at 4°C.

The supernatant was then transferred to a pre-cooled centrifuge tube and centrifuged again under vacuum at 4°C for 60 min at 100,000 x g. The supernatant from this step was carefully discarded and the pellet containing testes microsomes was gently suspended in 3 mL of Buffer B at pH 7.4 containing 50 mM Tris-HCl, 0.1 mM EDTA, 10 mM potassium phosphate and 20% glycerol and stored at -70° until further use. Microsomal protein was measured using the BioRad Protein Assay.

4.2.3 Enolase and Pregnenolone Microsomal Incubations

Three different incubation times: 5 minutes, 10 minutes and 20 minutes were tested for the production of pregnenolone metabolites, including androstenone. Ten mg of testis microsomal protein was incubated in 10 mM

NADPH, 10 mM PAPS, 10 uL of 1 mM [7-3H(N)]-pregnenolone (0.1 mCi/mL) and 118

Units of baker’s yeast enolase in 1 mL of Buffer B. Testis microsomes, pregnenolone and Buffer B were pre-incubated for 5 minutes in a 37°C water bath before the addition of NADPH, PAPS and enolase. There were also control incubations without the addition of enolase.

The optimum time for androstenone production from testis microsomes was determined to be 20 minutes with NADPH and androstenone incubation. However, in order to maximize androstenone sulfate production, subsequent incubation protocols were examined. After an initial pre-incubation of the microsomes with

androstenone of 5 minutes, NADPH was added for a 20 minute incubation followed by the addition of PAPS and enolase for either 5 minute or 20 minute incubations.

4.2.4 Steroid Extraction from Microsomal Incubations

In order to stop the metabolism, samples were vortexed well for 1 minute with 3 mL of fresh methylene chloride and centrifuged at room temperature for 15 minutes at 1500 RPM. The bottom layer of the solution was carefully removed by pipette and transferred to a clean tube. The remaining solution was re-extracted using 2 mL of methylene chloride and centrifuged again. The bottom layer of this extraction was added to the previous bottom layer and extracts were dried under nitrogen stream in a warm sand bath and resuspended using 200 uL of methanol.

These extracts were separated into free and conjugate fractions by Sep-Pak®

Chromatography.

4.2.5 Sep-Pak Chromatography and HPLC Analysis

Metabolite extracts were diluted with 5 mL of MilliQ water. Sep-Pak® C18 cartridges (Waters Corp USA) were primed under gentle vacuum using 5 mL of methanol followed by 5 mL MilliQ water. Samples were placed on cartridges and eluted under gentle vacuum followed by a wash with 5 mL MilliQ water. Two different concentrations of methanol were used to separate conjugate versus free metabolites: the first, 47% methanol, resulted in the elution of the conjugate fraction of the metabolites. The second, 100% methanol, resulted in the elution of the free fraction of the metabolites. These fractions were dried down using a gentle nitrogen stream in a warm sand bath and re-suspended using 230 uL 100%

acetonitrile. Further analysis of both conjugate and free fractions was performed using HPLC as previous described in Chapter 3.

4.3 Results

4.3.1 Optimum Pregnenolone Incubation Time

The optimal pregnenolone incubation time was determined using the free fraction methodology from Chapter 3 in order to determine whether pregnenolone was converted efficiently into the 16-androstenes.

Radiolabeled pregnenolone remained following the first incubation of 5 minute pre-incubation followed by 5 minute incubation with NADPH, PAPS, pregnenolone and enolase as indicated by the presence of free pregnenolone at 6.81 minutes (Figure 4.1). Radiolabeled pregnenolone was mostly metabolised by testis microsomes after the following protocol: 5 minutes pre-incubation with pregnenolone, 20 minute NADPH incubation and, finally, 20 minute PAPS and enolase incubation (Figure 4.2).

Figure 4.2: Optimum pregnenolone metabolism to 16-androstenes occurred after 5 minute pre- incubation with pregnenolone, 20 minute incubation with NADPH and 20 minute incubation with PAPS and enolase. Peaks found at 13 minutes, 14 minutes and 16 minutes corresponded to 3α- androstenol, 3β-androstenol and androstenone respectively. 4.3.2 Conjugate Fractions from Microsome Incubations

Conjugate fractions were analyzed by HPLC Protocol A from Chapter 3 for the presence of androstenone sulfate. These fractions were taken from control incubations – without enolase – and incubations with enolase from four randomly chosen individuals (pigs 5, 7, 8, 12). In half of the individuals (pigs 5 and 8), the production of androstenone sulfate was dramatically increased with the addition of enolase (Figures 4.3a and 4.3b).

Individual 8 - Total CPM - ENOL- 16000

14000

12000

10000

8000

6000

4000

2000

0 4.50 9.00 0.00 1.50 3.00 6.00 7.50 10.50 12.00 13.50 15.00 16.50 18.00 19.50 21.00 22.50 24.00 25.50 27.00 28.50 30.00 31.50 33.00 34.50 36.00 37.50 39.00

Individual 8 - Total CPM ENOL+ 16000

14000

12000

10000

8000

6000

4000

2000

0 4.50 9.00 0.00 1.50 3.00 6.00 7.50 10.50 12.00 13.50 15.00 16.50 18.00 19.50 21.00 22.50 24.00 25.50 27.00 28.50 30.00 31.50 33.00 34.50 36.00 37.50 39.00

Figure 4.3b. Conjugate fraction output (counts per minute (CPM)) from HPLC Protocol A showing a peak at 2 to 4 minutes, indicative of steroid glucuronides and DHEAS and E1S and a peak at 21-22 minutes, indicative of androstenone sulfate. In this case, the peak at 18 minutes was determined to be the same compound as the compound in the 21-22 minute peak. This incubation was from individual 8 and was done in the presence of enolase. Note the much larger androstenone sulfate peak compared to Figure 4.3a.

Conjugate fractions were also analyzed to identify pregnenolone metabolites.

On average, incubations without enolase had the highest peaks of radioactivity between 2 and 4 minutes, indicating the presence of steroidal glucuronides and sulfates as determined by comparison to standards for estrone and DHEA glucuronide, and DHEAS and E1S (Figure 4.3a or b). However, incubations with the addition of enolase tended to have the highest radioactivity found between 21 and

22 minutes, corresponding to the presence of androstenone sulfate. Additionally, individuals had peaks at 15 minutes, and 16 minutes, corresponding to standards indicating the presence of 3β-androstenol sulfate, and 3α-androstenol sulfate,

respectively (Figure 4.3c). The relative metabolite production from all incubations with and without enolase is summarized in Table 4.2.

The individual with the highest change in pregnenolone conversion to androstenone sulfate was individual 8, with an 6.54-fold increase in androstenone sulfate production with the addition of enolase. Similarly, the individual in which the addition of enolase resulted in the least difference in androstenone sulfate was individual 12, with a 0.89-fold increase – which is not statistically significant as a decrease. Interestingly, this individual also had the highest formation of androstenone sulfate from the Leydig cell incubations from the previous chapter – although this same pattern was not seen in the individual with the highest response to enolase treatment – the production of androstenone sulfate in individual 8 from

Chapter 3 was average compared to the other individuals.

Individual 8- CPM - ENOL- 2000 1800 1600 1400 1200 1000 800 600 400 200 0 0.00 1.50 3.00 4.50 6.00 7.50 9.00 10.50 12.00 13.50 15.00 16.50 18.00 19.50 21.00 22.50 24.00 25.50 27.00 28.50 30.00 31.50 33.00 34.50 36.00 37.50 39.00

Figure 4.3c. Graph of total counts per minute (CPM) for a testis microsome incubation from individual 8 that did not contain enolase. Peaks at 5-6 minutes were found to be unmetabolized pregnenolone whereas the two, smaller peaks were found to be, in this run only, DHEAS and E1S. Very small peaks found at 15 and 16 minutes were 3β-androstenol and 3α-androstenone and the large peak at 21-22 minutes was androstenone sulfate.

Pig-5 Pig-7 Pig-8 Pig-12

Metabolite E+ E- E+ E- E+ E- E+ E-

E1S 0.09 0.04 0.15 0.07 0.11 0.05 0.06 0.18

DHS 0.06 0.01 0.11 0.05 0.05 0.05 0.05 0.11

AAS 0.01 0.03 0.02 0.03 0.01 0.00 0.02 0.05

BAS 0.04 0.06 0.02 0.06 0.01 0.01 0.04 0.07

ANS 0.63 0.32 0.48 0.49 1.70 0.26 0.89 1.00

Increase in 1.97-fold 0.98-fold 6.54-fold 0.89-fold ANS with E+ Table 4.2: The amount of conjugated metabolite found in testicular microsomes from 13 individual pigs as determined by percent conversion of radiolabeled pregnenolone (%). E+ percentages are from incubations with enolase and E- percentages are from incubations without enolase. E1S=estrone sulfate, DHS=DHEA sulfate, BAS=3β-androstenol sulfate, AAS=3α-androstenol sulfate and ANS=androstenone sulfate.

4.3.2 Free Fractions

The amount of free 16-androstenes produced by microsomes from all individuals was also examined using HPLC Protocol B from Chapter 3. Although certain individuals were more effective at producing androstenone, all individuals produced 3α-androstenol (13 minute), 3β-androstenol (14 minute) and androstenone (16 minute) in quantifiable quantities (Figures 4.4, 4.5). The pregnenolone metabolite production for individuals 5, 7, 8 and 12 is summarized in

Table 4.3. Androstenone production was highest from testicular microsomes produced from individual 8 at 6.7% of total radiolabeled material injected onto the

HPLC column and was lowest in individual 12 at 4.6% of total radiolabeled material injected onto the HPLC column. The data for individual 5 for the incubations without enolase was not available due to an HPLC error.

Figure 4.4: Androstene production from microsome incubations with radiolabeled pregnenolone with enolase from individual pig 8.

Figure 4.5: Androstene production from microsome incubations with radiolabeled pregnenolone and without enolase from individual pig 8.

Metabolite Pig-5 Pig-7 Pig-8 Pig-12 E+ E- E+ E- E+ E- E+ E- 3α 4.6 4.0 4.3 2.3 2.2 2.8 2.6

3β 29.1 27.7 30.8 35.6 30.9 25.0 31.3

N/A AN 4.9 5.1 5.9 6.7 5.6 4.6 5.4

ANE 2.1 0.9 4.3 3.1 3.5 0.8 4.3

ANL 4.6 2.3 1.9 0.1 0.6 4.3 0.6

E1 33.7 26.3 30.7 16.0 23.2 35.1 27.8

DH 15.2 10.7 13.3 33.0 30.5 21.5 23.6

Table 4.3: Summary of important pregnenolone metabolites produced by testicular microsomes for four pigs. The amount of each metabolite is depicted as a percentage of the radiolabeled material present for each HPLC injection. The percentage does not add up to 100% due to disregarding background peaks. 3α=3α-androstenol, 3β=3β-androstenol, AN=androstenone, ANE=androstadienone, ANL=androstadienol, E1=estrone, DH=DHEA.

As seen in Table 4.3, the percentage of 16-androstenes produced compared to other steroids are approximately the same and do not differ widely between individuals. The lowest 16-androstene content was found in number 12 which also had the highest production of other steroids. This could be due to pregnenolone conversion in this individual diverted from 16-androstene production. The individual with the highest 16-androstene production was number 8 with 47.7%

16-androstene content. The individual with the lowest production of other steroids was number 7 with 37.0% - this individual also had the second lowest 16- androstene production, indicating that overall steroid conversion from pregnenolone in number 7 may be reduced.

Conjugated and free androstenone content was compared between individuals as indicated in Table 4.4. Conjugated androstenone is represented as a percentage of total pregnenolone in the incubation whereas free androstenone is represented as a percentage of total radiolabeled material injected into the HPLC.

This was done due to differing HPLC protocols. Individual variation between androstenone metabolism is obvious here – number 7 had the lowest conjugate androstenone production and, as expected, one of the highest free androstenone production.

Pig-5 Pig-7 Pig-8 Pig-12

E+ E- E+ E- E+ E- E+ E-

16- 45.3 N/A 39.9 47.2 47.7 42.7 37.5 44.1 androstenes

E1+DHEA 48.9 37.0 44.0 49.0 53.7 56.6 51.4

Table 4.3: Comparison between 16-androstene production and estrone and DHEA production in testis microsome incubations (%).

Individual Conjugated Androstenone Free Androstenone 5 0.6 4.9 7 0.5 5.1 8 1.7 6.7 12 0.9 4.6 Table 4.4: Conjugated and free androstenone production. Conjugate androstenone is represented as a percentage of pregnenolone conversion from testis microsome incubations whereas free androstenone is represented as a percentage of total radiolabeled material injected into the HPLC column.

4.4 Discussion and Conclusions

The formation of androstenone sulfate had yet to be elucidated in testicular microsomes. As enolase is thought to be required to form androstenone sulfate

(Sinclair et al., 2005), it was important to characterize this role.

Although the presence of 3-keto-4-sulfoxy-androstenone was never confirmed directly by mass spectrometry from these incubations, its presence was extrapolated from previous results in Chapter 3. Not surprisingly, all individuals did not have equal androstenone sulfate production. This could be due to individual variation in androstenone production, enolase activity or due to differences in sulfation from sulfotransferases (Butt, 1975) – or all three. In order to differentiate among the three, further research on many individuals is required. Additionally, the

vast majority of conjugating enzymes are cytosolic and are thus not present in large quantities in microsomal fractions (Weinshilboum et al., 1997). The low conjugation of androstenone could be simply due to individual variation in quantity of cytosolic sulfotransferases resulting in even smaller amounts of these enzymes in microsomal fractions. Future research should examine the addition of cytosolic sulfotransferases to these microsomes.

The findings that the production of androstenone sulfate greatly increases – by a 6.54-fold increase in one individual - with the addition of enolase confirms the importance of enolase in the formation of androstenone sulfate. However, this relationship was seen only in half of the individuals in which testes microsome incubations were performed. This variation could be due to individual differences in enolase activity levels in the testis. For example, individuals with high background enolase activity – close to the saturation point for androstenone sulfate production – would have a lower response to additional enolase compared to individuals with low background enolase activity.

Leydig cell incubations have been directly shown to produce androstenone sulfate in Chapter 3 whereas the direct production of androstenone sulfate by hepatocytes has yet to be studied. Previous research by Sinclair (2004) has shown that the liver has lower quantities of sulfoconjugated 16-androstenes compared to the Leydig cells of the same boars. The author theorized that the liver may have decreased enolase activity, resulting in a lack of the formation of 3-keto to 3-enol tautomerism. Future research may examine whether porcine hepatocytes do in fact

contain decreased enolase activity, thereby decreasing its capacity to form sulfoconjugated androstenone. It is very important to characterize the role of the liver in the conjugation of androstenone as the liver is widely known as the organ that is largely responsible for the conjugation of other hormones such as DHEAS

(Pancholi, 2001).

Androstenone sulfate was formed in all individuals with enolase treatment.

The increased formation of androstenone sulfate with the addition of enolase highlights the importance of enolase in this reaction. This also suggests that microbial yeast enolase could be effective at the imperative conversion of the 3-keto bond of androstenone to a 3-enol intermediate, allowing androstenone to be conjugated. The use of this commercially available enolase has great potential for further production of androstenone sulfate using in vitro cloning systems such as those found in the next chapter – Chapter 5.

CHAPTER V: CHARACTERIZATION OF THE ROLE OF SULT2A1 IN

ANDROSTENONE CONJUGATION

5.1 Introduction

Steroid hormones are known to be conjugated in many different species (as reviewed by Gamage et al., 2006, Guillemette, 2003). One of several reasons for conjugation includes altering a hormone’s polarity in order to aid in circulation in the blood and subsequent excretion from the body. Without steroidal conjugation, hydrophobic steroids require binding proteins in the blood in order to travel in the circulation (Anderson, 1974) and are more likely to accumulate in adipose tissue

(Feher and Bodrogi, 1982). Steroidal conjugates travel readily within the blood and more readily enter the urine and bile than their unconjugated counterparts. Pigs are no exception to this. Steroid conjugates are more readily excreted from the boar in the urine and bile, thereby preventing their accumulation in the fat (Bonneau,

1982).

Of the possible conjugation reactions, sulfation of androstenone by sulfotransferases (SULTs) has been briefly, indirectly studied (Sinclair et al., 2006).

The action of sulfotransferases, along with 3’-phosphoadenosine 5’-phosphosulfate

(PAPS), involves the transfer of a sulfate group from the donor (PAPS) to the recipient compound – in this case, androstenone (as reviewed by Gamage et al.,

2006). Of the SULTs in the human body, the hydroxysteroid sulfotransferases

(SULT2 family) are most readily involved in the sulfation of androgens, including

dihydroepiandrosterone (DHEA) and their activity has been characterized in many sites including the Leydig cells of the testes (Hobkirk et al., 1989, Miki et al., 2002 and Gamage et al., 2006).

Based on work in humans, SULT2A1 and SULT2B1 have both been identified as being extensively involved in the sulfation of hydroxysteroids, including androgens (Geese et al., 2001, Gamage et al., 2006, Lindsay et al., 2008,) and, therefore, have the potential to be involved in androstenone sulfation. As SULT2A1 and SULT2B1 have been characterized in humans, some studies have looked into the role of SULT2A1 and SULT2B1 in androstenone metabolism – specifically in androstenone accumulation in fat (Sinclair et al., 2006, Moe et al., 2007a). These studies have suggested that both sulfotransferases impact androstenone distribution in boars. However, Moe et al (2007a) found conflicting results in the impact of SULT2B1 expression on the distribution of androstenone in adipocyte deposits in the body due to breed differences. Additionally, the authors utilized goat anti-human SULT2B1 antibodies to examine gene expression as opposed to porcine- derived antibodies. This is likely due to the fact that there is limited SULT2B1 gene information available for the pig. In another study published by the same lead author in the same year (Moe et al., 2007), genetic sequences of a wide variety of genes were examined using microarrays. The authors found SULT2B1 to be of some significance in androstenone metabolism; however, again, the authors used human

SULT2B1 to screen a porcine database. Unpublished preliminary research from our laboratory indicates that porcine SULT2B1 is more homologous to bovine SULT2B1

than human SULT2B1. Therefore, the impact of SULT2B1 on androstenone metabolism requires substantial further study.

In order to quantify the role of SULT2A1 and SULT2B1 in the formation of androstenone sulfate, these sulfotransferases can be cloned and expressed in a standard cell line. Previous work by our laboratory using Human Embryonic Kidney

(HEK293) cells has determined that these cells, when transfected with the appropriate genes coding for enzymes involved in the metabolism of pregnenolone, are able to readily synthesize DHEA, an androgen that utilizes similar metabolic enzymes to those for the formation of androstenone (Billen and Squires, 2009).

HEK cells are generally used due to their low levels of endogenous enzymes (as reviewed by Panter et al., 2005). In particular, enzymes involved in androgen metabolism, including 5α-reductase and 17βHSD, have been identified at low levels

(Panter et al., 2005, Quinkler et al., 2003). Although 17βHSD is not directly involved in androstenone metabolism, 5α-reductase is involved in the conversion of androstadienone to androstenone (Brooks and Pearson, 1986). However, the presence of endogenous sulfotransferase activity in HEK cells has yet to be studied.

Other cell lines such as HepG2 and CaCo-2 cells should not be used in studies examining sulfotransferase activity as human SULT activity is readily inducible in this cell line (Chen et al., 2008), potentially resulting in interfering background levels of sulfotransferase activity.

The purpose of this study was to determine whether HEK cells transfected with porcine SULT2A1 and incubated with androstenone, enolase, PAPS and NADPH are capable of producing androstenone sulfate.

5.2 Materials and Methods

5.2.1 Chemicals Used

Geneticin®, L-glutamine, penicillin-streptomycin, sodium pyruvate, non- essential amino acids for cell media were purchased from Gibco® (Invitrogen™,

Burlington, Ontario) whereas DMEM was from Lonza Walkersville Inc. (Maryland,

USA) and FBS was purchased from PAA Laboratories (Etobicoke, Ontario). For the cell lysis buffer, Tris HCl, Triton X-100, PMSF, pepstatin A, EDTA and N- ethylmaleimide were all purchased from Sigma-Aldrich® Co (Oakville, Ontario).

Enzymes required for cell lysis incubations including PAPS, enolase from baker’s yeast, and β-NADPH were also purchased from Sigma-Aldrich® Co.

5.2.2 Cloning of Sulfotransferase

SULT2A1 was amplified using the following forward and reverse primers: 5’-

CACGAGGCGCAAAGAACT-3’ and 5’-TTGCCATGGGAACAGCACTT-3’. The primers were designed from the NCBI reference sequence for SULT2A1 (NM_001037150.1).

Using Platinum Taq DNA Polymerase High Fidelity (Invitrogen™) enzyme, porcine

SULT2A1 cDNA sequence was amplified using a T3 Thermocycler (Biometra®) with

37 cycles of denaturing for 1 minute at 94°C, annealing for 1 minute at 63°C, and extending for 2 minutes at 72°C followed by a final extension of 10 minutes at 72°C.

Following amplification, the SULT2A1 sequence was inserted into a pcDNA3.1

TOPO® TA plasmid (Invitrogen) as per the manufacturer’s instructions. Plasmids were then transformed into chemically competent E. coli provided in the TOPO® kit and plated on pre-warmed ampicillin-selective LB agar plates and left to grow overnight at 37°C. Surviving E. coli colonies were “miniprepped” using the Gen Elute

Plasmid Miniprep Kit from Sigma-Aldrich® Co.; colonies were sent for sequencing to ensure the sequence inserted correctly. On colonies in which the sequence was correct, maxipreps were performed using the QIAGEN Plasmid Maxi Kit (BioBar at

University of Guelph) and plasmids were stored at -20°C until further use.

The amplification of SULT2B1 utilizing the published human SULT2B1 sequence (NM_004605.2) and proven PCR techniques (Sinclair, 2004) was unsuccessful and was therefore not estimated in this thesis.

5.2.3 Sulfotransferase Expression in HEK Cells

HEK293T cells (ATCC) were grown at 37°in T-75 cm2 flasks in warmed

DMEM media containing 10% FBS, 1% L-glutamine, 1% penicillin-streptomycin, 1% sodium pyruvate, 1% Non-essential amino acids and 1% Geneticin®.

When cells were ready to be plated, media was removed and discarded. Cells were rinsed with pre-warmed PBS (Sigma-Aldrich® Co.) and, gently removed by suction. In order to detach cells from flasks, 500 ul of pre-warmed trypsin was added and cells were removed, diluted with media containing FBS and counted.

Approximately 4.5 million cells were plated in 10 cm petri dishes with a total of 10 mL of media and incubated at 37°C for 24 hours.

In order to transfect cells with SULT2A1 plasmids, media from plated cells was removed and refreshed. SULT2A1 plasmids were transfected into HEK cells using Fugene® HD Transfection Reagent (Promega Corp., USA) as per manufacturer’s instructions, utilizing a ratio of 2:1 Fugene:DNA. Cells were allowed to transfect for 48 hours at 37°C before harvested and lysed for further use. In order to confirm HEK expression of SULT2A1, Western blotting for the V5-His tagged protein was performed as previously described (Billen and Squires, 2009).

5.2.4 Cell Lysate Assays

Following SULT2A1 transfection, HEK cells were harvested by first removing and saving the media using suction. Cell lysate assay was adapted from Rong et al.

(2001). Cells were rinsed with 3 mL of PBS and then 500 uL of cell Lysis buffer containing 50 mM Tris HCl, 1% Triton X-100, 1mM PMSF, 10 ug/mL pepstatin A, 2 mM EDTA and 2mM N-ethylmaleimide was added to each plate and a cell scraper was used to harvest cells. Cell lysates were transferred to a 1.5 mL plastic centrifuge tube, incubated at 4°C for 30 minutes, and centrifuged at 295 g for 15 minutes at room temperature and the supernatant was collected. The supernatant containing cell lysates used for further assay.

In order to form androstenone sulfate, cell lysate was incubated with the appropriate precursors. Cell lysis supernatant was measured for total protein content using BioRad Protein Assay (Bio-Rad Laboratories Inc.). One mg of protein was incubated with 0.1 mL each of 1 mM unlabeled androstenone, 10 mM PAPS, 10 mM NADPH and 130 Units of enolase enzyme from yeast in 50 mM Tris-HCl, 10mM

potassium phosphate, 0.1 mM EDTA and 20% glycerol, with a pH of 7.4 and incubated at 37°C for 30 minutes. The use of 130 Units of enolase was chosen as the average enolase activity in porcine tissue is 86 Units/mg protein (Gorsich et al.,

1999, Chorazyczewski et al, 1987) and so the use of 130 Units provides almost double the theoretical endogenous amount of enolase in porcine tissue. Thirty minutes was chosen to maximize androstenone sulfate production based on previous tests in our laboratory. Testing to ensure that the SULT2A1 protein was expressed and functional involved incubation as described above; however, rather than incubating with 1 mM unlabeled androstenone, incubations were performed with 3H-DHEA at a concentration of 0.1 mCi/mL.

Androstenone sulfate was collected and concentrated from the incubations using Sep-Pak® Chromatography as described previously in Chapters 3 and 4.

5.2.5 HPLC and Mass Spectrometry of Steroids

Metabolite separation and identification was performed as previously described in Chapter 3.

5.2.5 Statistical Analysis

Statistical analysis was performed using SAS/STAT version 9.2 (SAS Institute

Inc.). The difference in the percentage of DHEAS produced as a percentage of total radiolabeled DHEAS added between transfected (SULT2A1+) and control

(SULT2A1-) incubations was analyzed using a T-Test for statistical significance.

5.3 Results

5.2.1 Conjugate Fraction Analysis

As the HEK cells were not incubated with radiolabeled androstenone, the conjugated 16-androstenes were only visualized using UV at the wavelengths described in Chapter 3. However, HEK cells do not contain the appropriate enzymes required to convert androstenone to androstenol and, as such, only conjugated androstenone should be present. Sulfated androstenone was examined by mass spectrometry.

SULT2A1 is well established as being responsible for the sulfoconjugation of

DHEA to DHEAS (Gamage et al., 2006). In order to confirm that transfected SULT2A1 was functional, incubations were performed with SULT2A1+ and SULT2A1- HEK cells with and without the presence of the co-factors required for sulfation of DHEA

– PAPS and NADPH (Figure 5.1a, 5.1b). Incubations performed with cells transfected with SULT2A1 and incubated in the presence of PAPS and NADPH resulted in a 9.3– fold increase in DHEAS formation when compared to control, SULT2A1- cells incubated with the same co-factors (Table 5.1). Additionally, control samples transfected with an empty plasmid show a very small level of endogenous sulfotransferase activity, although these levels are negligible. These results indicate that the transfected SULT2A1 is functional.

SULT2A1+/PAPS,NADPH+ After Blank Correction 40000 35000 30000 25000 20000 15000 10000 5000 0 0.00 1.50 3.00 4.50 6.00 7.50 9.00 10.50 12.00 13.50 15.00 16.50 18.00 19.50 21.00 22.50 24.00 25.50 27.00 28.50 30.00 31.50 33.00 34.50 36.00 37.50 39.00

Figure 5.1a: HPLC output from Protocol A (conjugate fraction) for SULT2A1 transfected cells incubated with DHEA, PAPS and NADPH, following correction for background peaks. The 3 minute peak corresponds to DHEAS.

SULT2A1-/PAPS, NADPH+ After Blank Correction 40000 35000 30000 25000 20000 15000 10000 5000 0 0.00 1.50 3.00 4.50 6.00 7.50 9.00 10.50 12.00 13.50 15.00 16.50 18.00 19.50 21.00 22.50 24.00 25.50 27.00 28.50 30.00 31.50 33.00 34.50 36.00 37.50 39.00

Figure 5.1b: HPLC output for Protocol A (conjugated) for control transfected cells incubated with DHEA, PAPS and NADPH following correction for background peaks, showing a very small level of endogenous sulfotransferase activity from the HEK cells.

Incubation DHEAS Formation (%) Conjugate Fraction Overall (%)

SULT2A1+/PAPS, 4.1* 3.1** 36.9 NADPH+

SULT2A1-/PAPS, NADPH+ 1.0* 0.3** 12.0

SULT2A1+/Blank 1.0 13.0

SULT2A1-/Blank 0.6 11.3

Table 5.1: DHEAS formation in transfected and control HEK cells as calculated as a percentage of DHEA added (%). For the incubations with the additional cofactors, the first column is without subtraction of the blank samples whereas the second column takes into account the blank samples. * and ** indicates statistical significance in the production of DHEAS between SULT2A1+ and SULT2A1- cell incubations.

5.2.2 Mass Spectrometry

Unfortunately, the only method to directly confirm the production of androstenone sulfate from these experiments is by mass spectrometry. As such, peaks found by UV and other fractions present at approximately the same elution time as androstenone sulfate (as found in Chapter 3) were examined by mass spectrometry.

None of the peaks examined by mass spectrometry contained compounds matching the mass of androstenone sulfate. Additionally, fractions from HPLC runs for all samples collected between 12 and 22 minutes at 1-minute intervals were subjected to mass spectrometry. Again, none of these fractions contained compounds matching the mass of androstenone sulfate. However, this does not necessarily mean that androstenone sulfate was not formed. There is the potential that androstenone sulfate was produced in such low quantities that, in the process of sample preparation and ionization into MS, the androstenone sulfate was simply

lost. Additionally, the method of detection could be not sensitive enough to detect such small quantities of androstenone sulfate.

5.2.3 Free Fraction Analysis

The free fractions were examined in order to compare the amount of free androstenone added to the incubations with the free androstenone found in the free fractions after incubations. The concentration of androstenone for each treatment before and after incubation is summarized in Table 5.2. As free androstenone is visible by UV at high concentrations, free androstenone concentration was determined by peak area from UV data. There is little concentration difference in free androstenone resulting from the different treatments, with the lowest concentration found in the SULT+/ENOL- and SULT-/ENOL- incubations at 0.032 mM and the highest found in the SULT+/ENOL+ and SULT-/ENOL+ incubation with a concentration of 0.037 mM. Control incubations (SULT-/ENOL+, SULT-/ENOL-) showed similar androstenone concentrations compared to incubations with transfected HEK cells.

Treatment Androstenone Concentration

Added to incubations 0.1 mM

SULT+/ENOL+ 0.037 mM

SULT+/ENOL- 0.032 mM

SULT-/ENOL+ 0.037 mM

SULT-/ENOL- 0.032 mM

Table 5.2: Androstenone concentration following treatments of HEK cell lysates.

5.4 Discussion and Conclusions

Although each incubation contained the enzymes thought to be responsible for the formation of androstenone sulfate, it was not found in any of the treatments as indicated by mass spectrometry. This could be due to porcine SULT2A1 not being responsible for 3-keto-4-sulfoxy-androstenone formation or that the mass spectrometry methods required more sensitivity. One method of improving the sensitivity of these experiments would be to perform them with radiolabeled androstenone – a compound that was not available at the time of the study.

There has been some debate in the literature regarding SULT2A1 versus

SULT2B1 being responsible for androstenone sulfation (Sinclair et al, 2005, Moe et al, 2007a). Sinclair et al (2005) demonstrated directly the importance of SULT2A1 and the conjugation of androstenone whereas Moe et al (2007a) took a less direct approach utilizing methods not optimized for swine research in examining SULT2B1 and free androstenone metabolism. Moe et al (2007a) examined SULT2B1 gene expression in hepatocytes and microsomes in high and low androstenone boars from two different breeds using SULT2B1 antibodies for human SULT2B1 expression. This was likely due to a lack of commercially available porcine-specific antibodies – therefore, the use of human SULT2B1 antibodies brings into question the validity of their results. As the experiments in this chapter largely discount

SULT2A1 from being directly involved in androstenone sulfation, it would be prudent for future research to examine additional sulfotransferases, including the various isoforms of SULT2B1.

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Although HEK cell lysate assays of SULT2A1-transfected cells did not show the capacity to sulfate androstenone in this study, future research should not discount the involvement of SULT2A1. Additionally, research should involve adding

SULT2A1 to the microsomal incubations from Chapter 4. As that protocol has already shown the production of androstenone sulfate, the examination of the role of excess SULT2A1 on this production is necessary in order to completely discount the role of SULT2A1 in androstenone sulfate production. This research should also involve the examination of other potential sulfotransferases and consider utilizing an alternative enolase source to continue to characterize the role of SULT2A1.

The role of enolase in androstenone sulfation has been established in

Chapter 4 – however, there may be additional enzymes required for the sulfation or additional cofactors not yet considered that are required for HEK cells to effectively conjugate androstenone. Although testicular microsomes produced androstenone sulfate, these enzymes and cofactors may not be found in the membrane-bound microsomal fractions and may be cytosolic in origin. Studies with complete cells, not just microsomes, should be performed. As the study of androstenone conjugation is a relatively novel field, all enzymes required for androstenone sulfate formation have yet to be identified. Future research should involve further characterization of enzymes involved in androstenone sulfation.

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CHAPTER VI: FUTURE CONSIDERATIONS AND THESIS CONCLUSIONS

6.1 Discussions and Future Considerations

Objective 1: To confirm or deny the presence of androstenone sulfate in the

Leydig cell of the mature boar.

Androstenone production in the mature boar has been well characterized

(e.g. Saat et al., 1972, Bonneau, 1982, Aldal et al., 2006) – however, the direct production of androstenone sulfate had yet to be characterized. Indirect evidence from previous research indicated that androstenone was conjugated in Leydig cells after 8 hour incubations with pregnenolone (Sinclair 2005). This indirect evidence was produced from the deconjugation of entire conjugate fractions from these incubations followed by HPLC metabolite separation to confirm the presence of androstenone. Although this evidence was well established and scientifically sound, the important direct analysis of androstenone sulfate had yet to occur. Direct analysis of androstenone sulfate decreases the potential for artefacts and other contaminants in samples that have the potential to falsely inflate measures of sulfoconjugate content. Therefore, the goal of Chapter 3 was to directly analyze the production of androstenone sulfate from Leydig cell incubations with pregnenolone.

Leydig cell incubations treated with pregnenolone were separated by Sep-

Pak chromatography in order to separate and concentrate the conjugate and free fractions for further analysis. Conjugate fractions were run on a different HPLC

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protocol than free fractions. Conjugate fraction peaks were isolated, chemically deconjugated and their identity was confirmed by analysis using the free HPLC protocol. Androstenone sulfate was found in peaks eluting at approximately 21 minutes - although occasionally, the elution of this peak began at 18 minutes.

In order to confirm the presence of androstenone sulfate, fractions from the conjugate HPLC runs were sent to mass spectrometry. A compound corresponding to a sulfated form of androstenone was found. Although this androstenone sulfate was not at the expected 352-353 m/z mass, it was determined that it was in fact 3- keto-4-sulfoxy-androstenone – that is, the sulfate group was attached at the 4- carbon as opposed to the expected 3-carbon. This compound had a mass of 367-368 m/z. This attachment occurred due to the formation of an enolated androstenone intermediate, resulting in a double bond between 3-C and 4-C as well as an open alcohol group attached to the 3-C. Although the action of enolase could result in this double bond forming between the 2-C and 3-C as opposed to the 3-C and 4-C, it is unknown which of the two forms occurs. For this study, we have assumed that the double bond occurs at the 3-C and 4-C position, thus resulting in the name 3-keto-4- sulfoxy-androstenone. The sulfation of androstenone was expected to happen at the

3-C open alcohol group; however, the MS/MS data proved otherwise.

The sulfated group removed from the androstenone sulfate found had a mass of 97 m/z. According to work done by Yi et al. (2006), this indicates that the sulfate is attached to a double bonded C group as opposed to the expected alcohol group.

The mass of the sulfate group removed from the 3-enol group would have had a

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mass of 80 m/z, due to its SO3 composition. However, the sulfate group had a much

- higher mass – owing to the fact that this group is likely a charged SO4 group resulting from its hydrolysis from the 4-C position. Additionally, the mass of the androstenone sulfate found corresponds to the conservation of the 3-keto group in the androstenone sulfate, further indicating that the sulfation did not occur at the 3- position. The combination of the mass of the actual compound at 367-368 m/z and the mass of the aforementioned sulfate group lend credence to the nomenclature of this compound as 3-keto-4-sulfoxy-androstenone.

The formation of androstenone sulfate requires enzymes that enable it to be conjugated – the same enzymes would be required to form the intermediates for other forms of conjugation including glucuronidation. Glucuronidation involves the addition of a glucuronosyl group from UDP-glucuronic acid to an aglycone, in this case androstenone (Guillemette, 2003). The enzyme group responsible for this is the

UDP-glucuronosyl transferases (UGTs) and have been well characterized in humans.

UGT activity has been associated with the preparation of compounds for removal from the body through the gut and urine (as reviewed by Barbier and Belanger,

2008). UGT expression is highest in the liver and gut as these organs are heavily involved in detoxification and removal of compounds from the body (Guillemette,

2003). However, UGT expression and activity has not been characterized in the pig – as such, future research should involve the search for and potential identification of a glucuronidated form of androstenone as well as the role of different UGTs in its formation.

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The production of androstenone sulfate in the liver has yet to be studied. The potential production of sulfated androstenone in the liver could dramatically change the balance of overall free and conjugated androstenone levels in the boar. As such, future research should examine the potential production of glucuronidated and sulfated androstenone in the liver.

As the direct identification of androstenone sulfate is novel, further characterization of its production and degradation, transportation and distribution throughout the body, interactions with fat tissue and its relationship to free androstenone accumulation throughout the body is required. In addition to the future research discussed above, the relationship between free androstenone in the plasma, free androstenone in the fat and the production of conjugated androstenone from the Leydig cells should also be established.

Although some of the enzymes required in androstenone sulfate formation have been discussed in earlier chapters, including the role of enolase and SULT2A1, other factors can impact its production. Our studies have shown that androstenone sulfate varies amongst individuals and, as such, there is potential for breed differences as well. Previous research has indicated that the impact of conjugation on androstenone metabolism varies by breed of pig (Moe et al., 2007a). This indicates that future research should examine the role of breed differences in the production of androstenone sulfate.

The degradation of androstenone sulfate to androstenone is an important factor in the impact of the compound on boar taint levels. Although we have shown

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that chemical hydrolysis of androstenone sulfate results in androstenone, direct enzymatic hydrolysis of androstenone sulfate has yet to be studied. The enzyme involved in this hydrolysis in vivo is likely steroid sulfatase (STS) as this compound is responsible for the deconjugation of a variety of sulfated steroids including

DHEAS (Hanson et al., 2004). STS has been characterized in human Leydig cells as well as adipocytes, and is known to be involved in the hydrolysis of DHEAS once it is inside adipocytes (Miki et al., 2002, Reed et al., 2005, Valle et al., 2006). The role of

STS in the homeostasis of androstenone sulfate and free androstenone has yet to be established - as such, future research should concentrate on the relationship between STS and sulfated androstenone.

Other than the production and degradation of androstenone sulfate, the transportation, accumulation and excretion from the body must also be examined.

As conjugated steroids are hydrophilic, they are capable of being transported freely in the plasma and bind very weakly to albumin (Johnson and Everitt, 2007).

However, as discussed previously, free steroids require binding to plasma proteins in order to move throughout the body (Anderson, 1974). This binding occurs most readily to albumin or to compound-specific binding proteins as in the case of testosterone movement and the involvement of human sex-hormone binding globulin (SHBG) (Buron and Westphal, 1972). Free androstenone would require these proteins although androstenone-specific proteins have yet to be established.

As the role of these proteins in the movement of conjugated and free androstenone has yet to be studied, their relationship to free androstenone accumulation and androstenone sulfate excretion from the body should be further studied. In addition

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to the transportation of androstenone sulfate throughout the body, the characterization of the relationship between plasma free androstenone and the production of conjugated androstenone from Leydig cells should be a priority in future research. It has been shown that individuals with higher amounts of conjugated androstenone have a correspondingly lower amount of plasma free androstenone, these individuals should have less free androstenone accumulation in the fat and more androstenone excretion from the body (Sinclair et al., 2005). The direct characterization of androstenone sulfate content in the plasma and its relationship to free androstenone in the fat would help confirm this previous research.

In addition to the movement throughout the body, the accumulation of androstenone in the fat should be studied. If androstenone sulfate acts in a similar fashion as excess plasma DHEAS, it can be actively transported into adipocytes by organic anion-transporting polypeptides (OATPs) (as reviewed by Hagenbuch and

Meier, 2003). Once inside the adipocytes, DHEAS is hydrolysed by STS into free

DHEA and further metabolized. Should this occur readily with excess androstenone sulfate, OATPs may have a dramatic impact on the accumulation of free androstenone in the fat. If, once in the adipocyte, androstenone sulfate is hydrolyzed in a manner similar to that of DHEAS, the resulting free androstenone could contribute to the formation of boar taint. If OATPs are required for androstenone sulfate movement into the fat, genetic selection against high-OATP individuals could aide in the decrease of the accumulation of free androstenone in the boar. As such,

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the interaction between androstenone sulfate, OATPs and adipocytes should be a subject for future research.

Although it is well established that steroids can be excreted from the body in the urine (Butt, 1975), there has been some recent research from our laboratory indicating that androstenone undergoes enterohepatic circulation (Jen, 2010). 3- keto-4-sulfoxy-androstenone has a molecular mass of 367-368 m/z, which falls between the 300 and 600 m/z required for enterohepatic circulation (Sandberg et al., 1967, Black, 1988) – therefore, androstenone sulfate is a good candidate for enterohepatic circulation. Future research should work towards directly characterizing the formation of androstenone sulfate from the gut as well as examining the pathway of the excretion of androstenone sulfate from circulation into the gut and urine and, subsequently, from the animal.

The characterization of the formation of androstenone conjugates and their relationship to the metabolism, fat accumulation, enterohepatic circulation and excretion of free androstenone will add much to the breadth of knowledge of androstenone metabolism and, perhaps, aid in the ultimate elimination of boar taint.

Objective 2: To examine the role of enolase in the formation of androstenone sulfate.

Testis microsomes from four individuals were incubated with commercially available yeast-derived enolase, PAPS, NADPH and pregnenolone – all factors thought to be required for the production of androstenone sulfate. Control

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incubations without the presence of enolase were performed in order to elucidate the role of enolase in androstenone sulfate formation.

Androstenone was conjugated in all of the testis microsomes from individuals in the presence of enolase and in very small quantities in control incubations without enolase. An increase in androstenone conjugation compared to control incubations was found – up to 6.54-fold in one individual – in incubations with enolase. This increase is an indication that enolase is required in the formation of androstenone conjugates. As an enol-intermediate is required for androstenone sulfate formation (Sinclair et al., 2005), this impact of enolase was expected. These results also confirm that commercially available yeast-derived enolase is capable of performing the 3-keto to 3-enol conversion with sufficient efficiency for androstenone sulfate formation in testis microsomes.

The direct formation of androstenone sulfate in hepatocytes and liver microsomes has yet to be studied. As sulfoconjugation of androstenone does occur, to a certain degree, in the liver (Sinclair, 2005), enolase expression and activity in the liver should be examined. The role of enolase activity, in the liver especially, would go far beyond the formation of androstenone sulfate and would likely increase the formation of glucuronidated androstenone.

Testis microsomes were found to be very effective at producing androstenone sulfate and the potential use of these to produce androstenone sulfate standards should be examined.

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Objective 3: To examine the relationship between SULT2A1 expression and androstenone sulfate production utilizing HEK293 cells.

Porcine sulfotransferase 2A1 (SULT2A1) was transfected into HEK293 cells in order to overexpress the protein, enabling the study of SULT2A1 and the formation of androstenone sulfate. In order to confirm functional SULT2A1 production, cell lysates were incubated with 3H-DHEA, PAPS and NADPH. SULT2A1 was deemed functional when SULT2A1- control incubations did not yield DHEAS formation whereas SULT2A1+ incubations did. Cell lysates were incubated with enolase, PAPS, NADPH and unlabeled androstenone in order to produce androstenone sulfate. The presence of androstenone sulfate was not found by analysis by HPLC followed by MS.

Although, on the surface, these findings indicate that SULT2A1 plays a limited role in the formation of androstenone sulfate, SULT2A1 should not be discounted from involvement in this pathway. The analysis techniques used to confirm the presence of androstenone sulfate were not as sensitive as those in previous chapters

– indicating that, should androstenone sulfate be present, it may be at low enough levels that it is not quantifiable.

In order to confirm that SULT2A1 itself is not responsible for sulfoconjugation in androstenone, overexpressed SULT2A1 should be added to testis microsome incubations in a similar protocol to that found in Chapter 4. If

SULT2A1 is involved, androstenone sulfate production should be increased compared to controls. Another method of examining this would be to utilize

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sulfotransferase inhibitors like those Sinclair (2005) used to examine SULT2A1 expression and activity. If SULT2A1 is involved, androstenone sulfate production from these microsomes should decrease with the addition of a SULT2A1-specific inhibitor. Additionally, in order to completely discount the function of overexpressed SULT2A1 in androstenone sulfate formation, the same cell lysate incubations as performed in Chapter 5 should be performed with radiolabeled androstenone, as this technique is more sensitive for androstenone-specific metabolism.

If, after all of this, SULT2A1 is determined to not be involved in androstenone sulfoconjugation, other sulfotransferases should be examined. Previous research has discussed the potential for SULT2B1 involvement in androstenone conjugation

(Moe et al., 2007a, Moe et al., 2007). Although these authors did not use porcine- specific antibodies and sequences, this sulfotransferase should not be completely discounted and should be examined in a similar manner to that of SULT2A1 from

Chapter 5.

The further elucidation of the role of specific sulfotransferases as well as porcine specific enolases will help in the complete characterization of the formation of androstenone sulfate.

6.2 Thesis Conclusions

This study has shown that 3-keto-4-sulfoxy-androstenone is formed from pregnenolone in Leydig cells derived from sexually mature boars. The importance of

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enolase in the formation of androstenone sulfate was established, setting the groundwork for future enolase-specific studies. The role of sulfotransferase 2A1 in the formation of androstenone sulfate was considered and it was determined that future research is required. The involvement of this sulfated form of androstenone in androstenone metabolism holds great promise for future research into the prevention of boar taint in mature boars. Further studies should examine androstenone sulfate formation, degradation, transportation and accumulation throughout the body and excretion from the body. This will help elucidate the role of this unique, novel compound in boar taint metabolism – hopefully resulting in decreasing the need for the castration of male piglets, thereby increasing animal welfare and decreasing the cost to the producer.

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APPENDIX

100.00

90.00 Experiment 1 80.00 Experiment 2 70.00 Experiment 3 Experiment 4 60.00 Experiment 5 50.00 Experiment 6 Experiment 7 Percentage 40.00 Experiment 9 30.00 Experiment 10 20.00 Experiment 11 Experiment 12 10.00 Experiment 13 0.00 Sample Wash Conjugate Free

Appendix 1a: Average percentage conjugation of each individual pig including sample (flow-through), free and water wash fractions.

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Individual Sample (%) Wash (%) Conjugate(%) Free (%)

1 3.54±0.2 0.21±0.0 63.03±1.5 33.22±1.5

2 2.87±0.5 0.18±0.1 72.82±3.9 24.13±3.6

3 10.96±1.0 0.74±0.2 83.36±7.5 7.22±7.6

4 1.01±0.0 0.13±0.0 85.01±1.7 13.7±1.6

5 0.81±0.0 0.1±0.0 87.5±1.3 11.58±1.3

6 1.59±0.1 0.17±0.0 82.11±6.0 16.22±6.2

7 5.31±0.14 1.06±0.1 71.05±6.6 22.77±6.7

9 3.12±0.13 0.58±0.1 88.95±2.2 7.35±2.1

10 6.38±1.0 0.59±0.1 65.49±9.4 27.54±10.3

11 6.05±1.7 0.73±0.3 51.81±7.2 41.41±9.2

12 4.80±0.4 0.44±0.0 85.07±0.7 9.7±1.1

13 4.65±0.38 0.38±0.1 82.05±1.7 12.94±1.9

Appendix 1b: Average percentage conjugation of each individual pig including sample (flow-through), free and water wash fractions (n=3, ± S.E.).

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Standards HPLC Protocol A 2500.00

2000.00

Pregnenolone Std 1500.00 Preg Glucuronide DHEA Glucuronide 1000.00 "DHEA Sulfate" Estrone Sulfate

500.00 Estrone Glucuronide

0.00 0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00

Appendix 2a: Retention times for standards for HPLC Protocol A (Conjugate Fraction HPLC Protocol).

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